Methods
in
Molecular Biology™
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
Liposomes Methods and Protocols Volume 2: Biological Membrane Models
Edited by
Volkmar Weissig Department of Pharmaceutical Sciences, Midwestern University College of Pharmacy Glendale, Glendale, AZ, USA
Editor Volkmar Weissig Department of Pharmaceutical Sciences Midwestern University College of Pharmacy Glendale Glendale, AZ USA
[email protected]
ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-446-3 e-ISBN 978-1-60761-447-0 DOI 10.1007/978-1-60761-447-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2009933261 © Humana Press, a part of Springer Science+Business Media, LLC 2010 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. Cover illustration: Background art is derived from Figure 6c in Chapter 21 Printed on acid-free paper Humana Press is a part of Springer Science+Business Media (www.springer.com)
Preface Efforts to describe and model the molecular structure of biological membranes go back to the beginning of the last century. In 1917, Langmuir described membranes as a layer of lipids one molecule thick [1]. Eight years later, Gorter and Grendel concluded from their studies that “the phospholipid molecules that formed the cell membrane were arranged in two layers to form a lipid bilayer” [2]. Danielli and Robertson proposed, in 1935, a model in which the bilayer of lipids is sequestered between two monolayers of unfolded proteins [3], and the currently still accepted fluid mosaic model was proposed by Singer and Nicolson in 1972 [4]. Among those landmarks of biomembrane history, a serendipitous observation made by Alex Bangham during the early 1960s deserves undoubtedly a special place. His finding that exposure of dry phospholipids to an excess of water gives rise to lamellar structures [5] has opened versatile experimental access to studying the biophysics and biochemistry of biological phospholipid membranes. Although during the following 4 decades biological membrane models have grown in complexity and functionality [6], liposomes are, besides supported bilayers, membrane nanodiscs, and hybrid membranes, still an indisputably important tool for membrane biophysicists and biochemists. In vol. II of this book, the reader will find detailed methods for the use of liposomes in studying a variety of biochemical and biophysical membrane phenomena concomitant with chapters describing a great palette of state-of-the-art analytical technologies. Moreover, besides providing membrane biophysicists and biochemists with an immeasurably valuable experimental tool, Alex Bangham’s discovery has triggered the launch of an entirely new subdiscipline in pharmaceutical science and technology. His observation that the lamellar structures formed by phospholipids exposed to aqueous buffers are able to sequester small molecules has lead to the development of the colloidal drug delivery concept. Following initial studies of enzyme encapsulation in liposomes as an approach towards the treatment of storage diseases [7, 8], a few years later in two New England Journal of Medicine landmark papers, Gregory Gregoriadis outlined the huge carrier potential of liposomes in biology and medicine [9, 10]. The following 2 decades saw immense efforts in academia and in soon-to-be-founded start-up companies to turn Gregoriadis’ vision into clinical reality. These 20 years of intense work in liposome laboratories around the world finally culminated with the FDA (USA) approval of the first injectable liposomal drug, Doxil, in February of 1995. Today, liposomes present the prototype of all nanoscale drug delivery vectors currently under development. Lessons learned in the history of over 40 years of Liposome Technology should be heeded by new investigators in the emerging field of pharmaceutical and biomedical nanotechnology. Volume I of this book is dedicated to state-of-the-art aspects of developing liposome-based pharmaceutical nanocarriers. All chapters were written by leading experts in their particular fields, and I am extremely grateful to them for having spent parts of their valuable time to contribute to this book. It is my hope that together we have succeeded in providing an essential source of practical
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know-how for every investigator, young and seasoned ones alike, whose research area involves in one way or another phospholipids, glycolipids, and cholesterol. Last but not least, I would like to thank John Walker, the series editor of “Methods in Molecular Biology,” for having invited me to assemble this book and above all for his unlimited guidance and help throughout the whole process. Glendale, AZ
Volkmar Weissig
References 1. Bangham AD, Standish MM, Watkins JC (1965) Diffusion of univalent ions across the lamellae of swollen phospholipids. J Mol Biol 13(1):238–252 2. Chan YH, Boxer SG (2007) Model membrane systems and their applications. Curr Opin Chem Biol 11(6):581–587 3. Danielli JF, Davson H (1935) A contribution to the theory of permeability of thin films. J Cell Comp Physiol 5:495–508 4. Gorter E, Grendel F (1925) On bimolecular layers of lipoids on the chromocytes of the blood. J Exp Med 41:439–443 5. Gregoriadis G (1976) The carrier potential of liposomes in biology and medicine (second of two parts). N Engl J Med 295(14):765–770
6. Gregoriadis G (1976) The carrier potential of liposomes in biology and medicine (first of two parts). N Engl J Med 295(13):704–710 7. Gregoriadis G, Ryman BE (1971) Liposomes as carriers of enzymes or drugs: a new approach to the treatment of storage diseases. Biochem J 124(5):58P 8. Gregoriadis G, Leathwood PD, Ryman BE (1971) Enzyme entrapment in liposomes, FEBS Lett 14(2):95–99 9. Langmuir I (1917) The constitution and structural properties of solids and liquids. II. Liquids. J Am Chem Soc 39:1848–1906 10. Singer SJ, Nicolson GL (1972) The fluid mosaic model of the structure of cell membranes. Science 175(23):720–731
Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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1 Utilization of Liposomes for Studying Drug Transfer and Uptake . . . . . . . . . . . . Alfred Fahr and Xiangli Liu 2 The Use of Liposomes in the Study of Drug Metabolism: A Method to Incorporate the Enzymes of the Cytochrome P450 Monooxygenase System into Phospholipid, Bilayer Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . James R. Reed 3 Use of Liposomes to Study Cellular Osmosensors . . . . . . . . . . . . . . . . . . . . . . . . Reinhard Krämer, Sascha Nicklisch, and Vera Ott 4 Studying Mechanosensitive Ion Channels Using Liposomes . . . . . . . . . . . . . . . . . Boris Martinac, Paul R. Rohde, Andrew R. Battle, Evgeny Petrov, Prithwish Pal, Alexander Fook Weng Foo, Valeria Vásquez, Thuan Huynh, and Anna Kloda 5 Studying Amino Acid Transport Using Liposomes . . . . . . . . . . . . . . . . . . . . . . . . Cesare Indiveri 6 Use of Liposomes for Studying Interactions of Soluble Proteins with Cellular Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chris T. Höfer, Andreas Herrmann, and Peter Müller 7 Liposomal Reconstitution of Monotopic Integral Membrane Proteins . . . . . . . . . Zahra MirAfzali and David L. DeWitt 8 The Reconstitution of Actin Polymerization on Liposomes . . . . . . . . . . . . . . . . . Mark Stamnes and Weidong Xu 9 Electroformation of Giant Unilamellar Vesicles from Native Membranes and Organic Lipid Mixtures for the Study of Lipid Domains under Physiological Ionic-Strength Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . L.-Ruth Montes, Hasna Ahyayauch, Maitane Ibarguren, Jesus Sot, Alicia Alonso, Luis A. Bagatolli, and Felix M. Goñi 10 Visualization of Lipid Domain-Specific Protein Sorting in Giant Unilamellar Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martin Stöckl, Jörg Nikolaus, and Andreas Herrmann 11 Biosynthesis of Proteins Inside Liposomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pasquale Stano, Yutetsu Kuruma, Tereza Pereira de Souza, and Pier Luigi Luisi 12 Study of Respiratory Cytochromes in Liposomes . . . . . . . . . . . . . . . . . . . . . . . . . Iseli L. Nantes, Cintia Kawai, Felipe S. Pessoto, and Katia C.U. Mugnol 13 Use of Liposomes to Evaluate the Role of Membrane Interactions on Antioxidant Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Salette Reis, Marlene Lúcio, Marcela Segundo, and José L.F.C. Lima
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14 Studying Colloidal Aggregation Using Liposomes . . . . . . . . . . . . . . . . . . . . . . . . Juan Sabín, Gerardo Prieto, and Félix Sarmiento 15 Assessment of Liposome–Cell Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jan A.A.M. Kamps 16 Methods to Monitor Liposome Fusion, Permeability, and Interaction with Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nejat Düzgünes¸, Henrique Faneca, and Maria C. Pedroso de Lima 17 The Use of Isothermal Titration Calorimetry to Study Multidrug Transport Proteins in Liposomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David Miller and Paula J. Booth 18 Studying Lipid Organization in Biological Membranes Using Liposomes and EPR Spin Labeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Witold K. Subczynski, Marija Raguz, and Justyna Widomska 19 Membrane Translocation Assayed by Fluorescence Spectroscopy . . . . . . . . . . . . . J. Broecker and S. Keller 20 Interaction of Lipids and Ligands with Nicotinic Acetylcholine Receptor Vesicles Assessed by Electron Paramagnetic Resonance Spectroscopy . . . . . . . . . . Hugo Rubén Arias 21 Environmental Scanning Electron Microscope Imaging of Vesicle Systems . . . . . . Yvonne Perrie, Habib Ali, Daniel J. Kirby, Afzal U.R. Mohammed, Sarah E. McNeil, and Anil Vangala 22 Freeze-Fracture Electron Microscopy on Domains in Lipid Mono- and Bilayer on Nano-Resolution Scale . . . . . . . . . . . . . . . . . . . . . . . . . . . Brigitte Papahadjopoulos-Sternberg 23 Atomic Force Microscopy for the Characterization of Proteoliposomes . . . . . . . . Johannes Sitterberg, Maria Manuela Gaspar, Carsten Ehrhardt, and Udo Bakowsky 24 Method of Simultaneous Analysis of Liposome Components Using HPTLC/FID . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sophia Hatziantoniou and Costas Demetzos 25 Viscometric Analysis of DNA-Lipid Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . Sadao Hirota and Nejat Düzgünes¸ 26 Fluorometric Analysis of Individual Cationic Lipid-DNA Complexes . . . . . . . . . . Edwin Pozharski 27 Fluorescence Resonance Energy Transfer-Based Analysis of Lipoplexes . . . . . . . . . Edwin Pozharski 28 Analysis of Lipoplex Structure and Lipid Phase Changes . . . . . . . . . . . . . . . . . . . Rumiana Koynova 29 Fluorescence Methods for Evaluating Lipoplex-Mediated Gene Delivery . . . . . . . Henrique Faneca, Nejat Düzgünes‚ , and Maria C. Pedroso de Lima 30 FRET Imaging of Cells Transfected with siRNA/Liposome Complexes . . . . . . . . Il-Han Kim, Anne Järve, Markus Hirsch, Roger Fischer, Michael F. Trendelenburg, Ulrich Massing, Karl Rohr, and Mark Helm 31 Spectral Bio-Imaging and Confocal Imaging of the Intracellular Distribution of Lipoplexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sebastian Schneider and Regine Süss
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32 Techniques for Loading Technetium-99m and Rhenium-186/188 Radionuclides into Pre-formed Liposomes for Diagnostic Imaging and Radionuclide Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Beth Goins, Ande Bao, and William T. Phillips 33 Fluorescence Correlation Spectroscopy for the Study of Membrane Dynamics and Organization in Giant Unilamellar Vesicles . . . . . . . . . . . . . . . . . . Ana J. García-Sáez, Dolores C. Carrer, and Petra Schwille 34 Liposome Biodistribution via Europium Complexes . . . . . . . . . . . . . . . . . . . . . . . Nathalie Mignet and Daniel Scherman 35 Biosensor-Based Evaluation of Liposomal Binding Behavior . . . . . . . . . . . . . . . . . Gerd Bendas 36 Use of Liposomes to Study Vesicular Transport . . . . . . . . . . . . . . . . . . . . . . . . . . Kohji Takei, Hiroshi Yamada, and Tadashi Abe Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Contributors Tadashi Abe • Department of Neuroscience, Okayama University Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama, Japan Hasna Ahyayauch • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Habib Ali • School of Life and Health Sciences, Aston University, Birmingham, UK Alicia Alonso • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Hugo Rubén Arias • Department of Pharmaceutical Sciences, College of Pharmacy, Midwestern University, Glendale, AZ, USA Luis A. Bagatolli • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Udo Bakowsky • Department of Pharmaceutical Technology and Biopharmacy, Philipps-Universität Marburg, Marburg, Germany Ande Bao • Department of Radiology, University of Texas Health Science Center, San Antonio, TX, USA Andrew R. Battle • Molecular Biophysics Laboratory, School of Biomedical Sciences and Institute for Molecular Bioscience, The University of Queensland, Brisbane, QLD, Australia Gerd Bendas • Department of Pharmacy, Rheinische Friedrich Wilhelms University Bonn, Bonn, Germany Paula J. Booth • Department of Biochemistry, University of Bristol, Bristol, UK Jana Broecker • Leibniz Institute of Molecular Pharmacology FMP, Berlin, Germany Dolores C. Carrer • BIOTEC, Technische Universität Dresden, Dresden, Germany Costas Demetzos • Department of Pharmaceutical Technology, School of Pharmacy, University of Athens, Athens, Greece David L. DeWitt • Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, USA Nejat Düzgünes¸ • Department of Microbiology, Arthur A. Dugoni School of Dentistry, University of the Pacific, San Francisco, CA, USA Carsten Ehrhardt • School of Pharmacy and Pharmaceutical Sciences, University of Dublin, Trinity College Dublin, Dublin, Ireland Alfred Fahr • Department of Pharmaceutics, Friedrich-Schiller-University, Jena, Germany Henrique Faneca • Faculty of Science and Technology, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal Roger Fischer • German Cancer Research Center (DKFZ), Heidelberg, Germany Alexander Fook Weng Foo • Molecular Biophysics Laboratory, School of Biomedical Sciences and Institute for Molecular Bioscience, The University of Queensland, Brisbane, QLD, Australia Ana J. García-Sáez • BIOTEC, Technische Universität Dresden, Dresden, Germany Maria Manuela Gaspar • Unidade Novas Formas de Agentes Bioactivos, iMed, Faculdade de Farmácia, Universidade de Lisboa, Lisboa, Portugal
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Beth Goins • Department of Radiology, University of Texas Health Science Center, San Antonio, TX, USA Felix M. Goñi • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Sophia Hatziantoniou • Department of Pharmaceutical Technology, School of Pharmacy, University of Athens, Athens, Greece Mark Helm • Department of Chemistry, Institute of Pharmacy and Molecular Biotechnology, University of Heidelberg, Heidelberg, Germany Andreas Herrmann • Mathematisch-Naturwissenschaftliche Fakultät I, Institut für Biologie/Biophysik, Humboldt Universität zu Berlin, Berlin, Germany Sadao Hirota • Department of Material Science, School of Engineering, Tokyo Denki University, Tokyo, Japan Markus Hirsch • Department of Chemistry, Institute of Pharmacy and Molecular Biotechnology, University of Heidelberg, Heidelberg, Germany Chris Höfer • Institut für Biologie/Biophysik, Humboldt Universität zu Berlin, Berlin, Germany Thuan Huynh • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia Maitane Ibarguren • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Cesare Indiveri • Dipartimento di Biologia Cellulare, Università della Calabria, Arcavacata di Rende, CS, Italy Anne Järve • Department of Chemistry, Institute of Pharmacy and Molecular Biotechnology, University of Heidelberg, Heidelberg, Germany Jan A.A.M. Kamps • Laboratory for Endothelial Biomedicine & Vascular Drug Targeting Research, Medical Biology Section, Department Pathology & Medical Biology, University Medical Center Groningen, Groningen, The Netherlands Cintia Kawai • Centro Interdisciplinar de Investigação Bioquímica CIIB, Universidade de Mogi das Cruzes, Mogi das Cruzes, S.P., Brazil Sandro Keller • Leibniz Institute of Molecular Pharmacology FMP, Berlin, Germany Il-Han Kim • Department of Bioinformatics and Functional Genomics, German Cancer Research Center (DKFZ), Institute of Pharmacy and Molecular Biotechnology, University of Heidelberg, Heidelberg, Germany Daniel J. Kirby • School of Life and Health Sciences, Aston University, Birmingham, UK Anna Kloda • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia Rumiana Koynova • Northwestern University, Evanston, IL, USA Reinhard Krämer • Institute of Biochemistry, University of Cologne, Cologne, Germany Yutetsu Kuruma • “Enrico Fermi” Study and Research Center, Rome, Italy José L.F.C. Lima • REQUIMTE, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal Xiangli Liu • Department of Pharmaceutics, Friedrich-Schiller-University, Jena, Germany Marlene Lúcio • REQUIMTE, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal Pier Luigi Luisi • Biology Department, University of RomaTre, Rome, Italy
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Boris Martinac • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia Ulrich Massing • Department of Clinical Research, Tumor Biology Center, Freiburg, Germany Sarah E. McNeil • School of Life and Health Sciences, Aston University, Birmingham, UK Nathalie Mignet • Unité de Pharmacologie Chimique et Génétique; CNRS, UMR 8151, Paris, France; Inserm, U 640, Paris, France; Faculté des Sciences Pharmaceutiques et Biologiques, Université Paris Descartes, Paris, France; ENSCP, Paris, France David Miller • Department of Biochemistry, University of Bristol, Bristol, UK Zahra MirAfzali • Encapsula NanoSciences LLC, 441 Donelson, Pike, Suite 345, Nashville, TN 37214, USA Afzal U. R. Mohammed • School of Life and Health Sciences, Aston University, Birmingham, UK L.-Ruth Montes • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Katia C.U. Mugnol • Centro Interdisciplinar de Investigação Bioquímica CIIB, Universidade de Mogi das Cruzes, Mogi das Cruzes, S.P., Brazil Peter Müller • Institut für Biologie/Biophysik, Humboldt Universität zu Berlin, Berlin, Germany Iseli L. Nantes • Centro Interdisciplinar de Investigação Bioquímica CIIB, Universidade de Mogi das Cruzes, Mogi das Cruzes, S.P., Brazil Sascha Nicklisch • Institute of Biochemistry, University of Cologne, Cologne, Germany Jörg Nikolaus • Mathematisch-Naturwissenschaftliche Fakultät I, Institut für Biologie/Biophysik, Humboldt-Universität zu Berlin, Berlin, Germany Vera Ott • Institute of Biochemistry, University of Cologne, Cologne, Germany Prithwish Pal • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia Brigitte Papahadjopoulos-Sternberg • NanoAnalytical Laboratory, San Francisco, CA, USA Maria C. Pedroso de Lima • Department of Biochemistry, Faculty of Science and Technology, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal Tereza Pereira de Souza • Biology Department, University of RomaTre, Rome, Italy Yvonne Perrie • School of Life and Health Sciences, Aston University, Birmingham, UK Regine Süss • Department of Pharmaceutical Technology and Biopharmacy, AlbertLudwigs University, Freiburg, Germany Felipe S. Pessoto • Centro Interdisciplinar de Investigação Bioquímica CIIB, Universidade de Mogi das Cruzes, Mogi das Cruzes, S.P., Brazil Evgeny Petrov • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia William T. Phillips • Department of Radiology, University of Texas Health Science Center, San Antonio, TX, USA Edwin Pozharski • Department of Pharmaceutical Sciences, University of Maryland, Baltimore, MD, USA
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Gerardo Prieto • Biophysics and Interfaces Group, Department of Applied Physics, Faculty of Physics, University of Santiago de Compostela, Santiago de Compostela, Spain Marija Raguz • Department of Biophysics, Medical College of Wisconsin, Milwaukee, WI, USA James R. Reed • Department of Pharmacology, Louisiana State University Health Science Center, New Orleans, LA, USA Salette Reis • REQUIMTE, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal Paul R. Rohde • Molecular Biophysics Laboratory, School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia Karl Rohr • Department of Bioinformatics and Functional Genomics, German Cancer Research Center (DKFZ), Institute of Pharmacy and Molecular Biotechnology, University of Heidelberg, Heidelberg, Germany Juan Sabín • Biophysics and Interfaces Group, Department of Applied Physics, Faculty of Physics, University of Santiago de Compostela, Santiago de Compostela, Spain Félix Sarmiento • Biophysics and Interfaces Group, Department of Applied Physics, Faculty of Physics, University of Santiago de Compostela, Santiago de Compostela, Spain Daniel Scherman • Unité de Pharmacologie Chimique et Génétique; CNRS, UMR 8151, Paris, France; Inserm, U 640, Paris, France; Faculté des Sciences Pharmaceutiques et Biologiques, Université Paris Descartes, Paris, France; ENSCP, Paris, France Sebastian Schneider • Department of Pharmaceutical Technology and Biopharmacy, Albert-Ludwigs University, Freiburg, Germany Petra Schwille • BIOTEC, Technische Universität Dresden, Dresden, Germany Marcela Segundo • REQUIMTE, Faculdade de Farmácia, Universidade do Porto, Porto, Portugal Johannes Sitterberg • Department of Pharmaceutical Technology and Biopharmacy, Philipps-Universität Marburg, Marburg, Germany Jesus Sot • Unidad de Biofisica (CSIC-UPV/EHU), Leioa, Spain Mark Stamnes • Department of Molecular Physiology & Biophysics, Roy J. and Lucille A. Carver College of Medicine, University of Iowa, Iowa City, IA, USA Pasquale Stano • Biology Department, University of RomaTre, Rome, Italy Martin Stöckl • Mathematisch-Naturwissenschaftliche Fakultät I, Institut für Biologie/Biophysik, Humboldt-Universität zu Berlin, Berlin, Germany Witold K. Subczynski • Department of Biophysics, Medical College of Wisconsin, Milwaukee, WI, USA Kohji Takei • Department of Neuroscience, Okayama University Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama, Japan Michael F. Trendelenburg • German Cancer Research Center (DKFZ), Heidelberg, Germany
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Anil Vangala • School of Pharmacy and Chemistry, Kingston University, London, UK Valeria Vásquez • Biochemistry Department, Gordon Center for Integrative Science, The University of Chicago, Chicago, IL, USA Justyna Widomska • Department of Plant Physiology and Biochemistry, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Krakow, Poland Weidong Xu • Department of Molecular Physiology & Biophysics, Roy J. and Lucille A. Carver College of Medicine, University of Iowa, Iowa City, IA, USA Hiroshi Yamada • Department of Neuroscience, Okayama University Graduate School of Medicine, Dentistry and Pharmaceutical Sciences, Okayama, Japan
Chapter 1 Utilization of Liposomes for Studying Drug Transfer and Uptake Alfred Fahr and Xiangli Liu Abstract On entry into the body of the patient, drugs have to overcome many barriers in order to reach the target. The knowledge of the ability of drugs to cross these barriers, which mostly consist of lipid membranes, is of utmost interest in pharmacy. High values of lipophilicity of a drug might be a good pre-requisite for crossing these barriers. It also led liposomologists to think that highly lipophilic drugs may “stick” in the lipophilic interior of liposomal phospholipid membranes and therefore these liposomes may act as a retard formulation of the lipophilic drug. The presented method here estimates the transfer time of lipophilic drugs between liposomal lipid bilayers. This may help to judge the presumed retardation function of a specific liposomal delivery system for a chosen lipophilic drug. Key words: Membrane transfer, Liposome drug delivery system, Lipophilic drug, Retardation, Mini column method
1. Introduction In the pharmaceutical field, liposomes are commonly known as drug carrier systems, but appeared at first in the scientific community as models of biological membranes (1). Similar to the early use in membrane biophysics, liposomes can also be very useful in pharmaceutical sciences, as the passive diffusion of drugs through physiological barrier membranes (e.g. epithelial cells in the gut) can be estimated with their help by different kinds of experimental setups (2). Another drug-delivery related problem can also be assessed by applying membrane biophysics methods to pharmaceutical V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_1, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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liposomology: the often not verified, but very often cited assumption that a lipophilic drug should bind to a high degree to the lipophilic interior of the liposomal bilayer simply because of its similarities in lipophilicity. Rather, as we had found for several lipophilic compounds, these compounds are transferred quite rapidly to other lipophilic binding places (3), which can also be reasoned by theoretical considerations (4). Methods like dialysis (5) have their value for estimating the thermodynamics of the transfer processes, but for analysing the kinetics in the minute time-range with absolute mass transfer values, other methods should be used. Not for all drugs the elegant method of spin-label measurements (6) can be used to get an estimate of drug retardation. Here we provide a protocol for measuring the transfer of lipophilic drugs between liposomal membranes. The drug is transferred during a mixing process from donor liposomes to acceptor liposomes; the donor liposomes are removed from the sample using a micro ion exchange column method. We describe this method under the assumption, that the drugs are available with a radioactive marker. This very comfortable situation regarding the transfer analysis is typically present in big pharmaceutical companies with their own radiochemistry labs ready for labelling the drugs of interest. A chromophoric group in the drug moiety helps as well in making the analysis simple, especially if this leads to a fluorescence drug. In the author’s experience, a sensitive HPLC-method can also be a sufficient method for analysis, but it may be time-consuming for reliable results. The obtained data can in most cases be easily analysed. By incorporating an un-exchangeable marker, one can not only determine the retention of donor liposomes on the column, but can also validate the whole method.
2. Materials 2.1. Substances
1. Egg phosphatidylcholine (EPC) (Lipoid KG, Ludwigshafen, Germany). Synthetic phosphatidylcholines (e.g. from Avanti Polar Lipids, Alabaster, AL, USA.) are highly recommended. 2. Dicetylphosphate (DCP) (Sigma-Aldrich, Taufkirchen, Germany). 3. Cholesterol and 1-palmitoyl-1,2-oleoyl phosphatidylcholine (POPC) (Genzyme, Liestal, Switzerland or Avanti Polar Lipids). 4. 14C-labelled cholesteryl-oleoyl-ether and 3H-labelled cholesteryl-oleate (GE Healthcare UK Ltd, Buckinghamshire, UK). The lipids were stored in –20°C before use. Typically small aliquots of a delivered batch are stored in the freezer to avoid degradation caused by repeated thaw and freeze.
Utilization of Liposomes for Studying Drug Transfer and Uptake
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5. The 14C labelled cyclosporin A (CyA) was synthesised and kindly provided by the radiochemistry group of Sandoz Pharm Ltd., now Novartis Inc. (Basel, Switzerland). 1–5 are either bought as chloroformic solution or the obtained powder is dissolved in chloroform in order to make stock solutions. These stock solutions should be kept at −20 to −80°C and should not be used after three months of storage. Always allow the frozen stock solution to reach ambient temperature in order to avoid water contamination of the solution which leads to lipid degradation. 6. DEAE Sepharose CL-6B (GE Healthcare Bio-Sciences AB, Uppsala, Sweden) is supplied pre-swollen in 20% ethanol. 7. Tris, saccharose, sodium chloride, and sodium azide (all in the highest purity available) (e.g. Sigma-Aldrich, Taufkirchen, Germany). 8. The two buffers used throughout the experiments are named Buffer A: 145 mM NaCl, 10 mM Tris pH 7.4, and Buffer B: 290 mM Saccharose, 10 mM Trizma pH 7.4, 0.02 % sodium azide. 2.2. Devices
1. Minicolumns were made in the workshop of the University of Perspex® and should preferably have the dimensions as indicated in Fig. 1. 2. All other devices should be available in a standard laboratory for liposomal research and are mentioned in the following text for reference.
3. Methods 3.1. Preparation of Donor Liposomes
1. Egg phosphatidylcholine (EPC), Dicetylphosphate (DCP), and cholesterol are weighed into individual test tubes and dissolved in chloroform to get the concentrations 20 mg/ml for EPC, 10 mg/ml for DCP, and 10 mg/ml for cholesterol. Appropriate volumes of each lipid with a mole ratio of each lipid component 7:1:2 (EPC:DCP:Cholesterol = 7:1:2) and 14 C-CyA (phospholipid:CyA = 300:1) are transferred to a single tube (see Note 1). 2. The mixture is evaporated using a Rotavapor device rotating at slow speed and heated (30°C) and is then subjected to vacuum to form a thin layer of lipid on the wall of the flask (see Note 2). This cyclosporin-lipid layer is hydrated with the appropriate amount of Buffer A (to get a phospholipid concentration of 10 mg/ml) for 2 h using the same Rotavapor
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Fig. 1. Schematic drawing of the device for measuring the transfer of substances between liposomes by the microcolumn technique
device after degassing the buffer and purging it with nitrogen gas. The final milky suspension is vortexed for 30 s at moderate speed, until all lipids have been removed from the glass vessel. This can be easily checked by optical inspection of the glass vessel from the outside. 3. This suspension is extruded 21 times (or more, but always use an uneven number!) through polycarbonate membranes with pore size 100 nm using a commercially available extruder
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(LiposoFast, Avestin Inc., Canada) in order to get homogenously sized liposomes. 4. The resulting liposomes are assessed for size distribution and zeta-potential and stored at +4°C before use. Liposomes formed by this method typically display a diameter range of 110–130 nm and a zeta-potential of −40 to −50 mV. 3.2. Preparation of Donor Blank Liposomes
1. Donor blank liposomes (without any transferrable drug included) are prepared using the same procedures as in Subheading 3.1. 2. The donor blank liposomes are labelled by 14C-cholesteryloleoyl-ether at 1 mCi/ml (see Note 3) as a non-exchangeable lipid marker.
3.3. Preparation of Acceptor Liposomes
1. 1-palmitoyl-1,2-oleoyl phosphatidylcholine (POPC) and cholesterol are weighed into individual test tubes and dissolved in chloroform to get the concentrations 20 mg/ml for POPC and 10 mg/ml for cholesterol. Appropriate volumes of each lipid with a mole ratio of the two lipids (POPC:cholesterol = 8:2) are transferred to a single tube. 3H-labelled cholesteryl-oleate is incorporated into the formulation at 1 mCi/ml. 2. The mixture is evaporated, hydrated with an appropriate amount of buffer A (to get a phospholipid concentration of 10 mg/ml), and extruded following the procedures as described in Subheading 3.1 to get the homogenously sized liposomal suspension. 3. Liposomes display a diameter range of 130–150 nm and zetapotential of ~0 mV.
3.4. Preparation of Liposomes for Saturation
1. 1-palmitoyl-1,2-oleoyl phosphatidylcholine (POPC), and cholesterol are weighed and mixed at a mole ratio of 8:2 following the procedures as described in Subheading 3.3. 2. The mixture is evaporated and hydrated (to get a phospholipid concentration of 10 mg/ml) following the procedures as described in Subheading 3.1 to get the liposomal suspension. 3. 2 ml of the liposomal suspension is transferred to a glass vial. This vial is mounted in a MSE-Soniprep 150 (Zivy AG, Oberwil, Switzerland) ultrasonic disintegrator with a titanium probe. The tip of the titanium probe is positioned 4 mm below the surface of the liposomal suspension. The vial is kept in an ice/water bath during the sonication, and nitrogen gas is used for purging the suspension to avoid oxidation of lipids during sonication. 4. The suspension is sonicated for 60 min (2 times 30 cycles, 1 cycle = 30 s sonicating and 30 s non-sonicating) with a 50% duty cycle using a MSE process timer (Zivy AG, Oberwil, Switzerland). The amplitude of the titanium probe is adjusted to 12 mm. The temperature of the suspension is checked to ensure that it does not rise significantly during sonication.
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5. This method allows the preparation of very small liposomes by introducing high amounts of energy into the phospholipid suspension. The average particle size of liposomes by sonication method is around 30–60 nm. 3.5. Size Determination of Liposomes
1. The size of the liposomes is determined by Dynamic laser light scattering using a Zetasizer Nano ZES3600 (Malvern Instruments Inc., UK). 2. Data are analysed by the Dispersion Technology software version 5.02 (Malvern Instruments Inc., UK).
3.6. Preparation of the Column Filling Gel DEAE-Sepharose CL-6B
1. The necessary amount of the DEAE-Sepharose CL-6B (depending on the number of mini-columns to be filled) is poured in an Erlenmeyer flask of appropriate volume (usual 200 mL size). 2. The gel settles down and the ethanol is carefully and slowly decanted by means of a Pasteur pipette connected via a tube to a water jet vacuum. 3. The gel is washed twice with buffer A in a ratio of 75% settled gel to 25% buffer A and kept in a last washing step with buffer B (1:1, v/v) at 4°C (see Note 4).
3.7. Preparation of the Ion-Exchange Microcolumn
1. The ion-exchange microcolumns were manufactured from Perspex® in the mechanical workshop of the university. The total bed volume of the microcolumn is about 0.5 ml (not very critical). 2. Some glass wool was placed at the bottom of the inner side of the column in order to ease the retaining of the gel in the column. 1.0 ml DEAE-sepharose CL-6B suspension pretreated as described in Subheading 3.6 is filled in the column (see Note 5). 3. The column is eluted with 2 ml buffer B at a speed of 30 drops in 45 s by using a pump (for example LKB Pump P-1, Pharmacia (Uppsala, Sweden)) and packed at the same time. 4. The microcolumns are saturated by applying 20 ml of the saturation liposomes prepared as described in Subheading 3.4 and eluted with 1.5 ml buffer B (see Note 6).
3.8. Validation of the Separation Efficiency of the Microcolumns 3.8.1. Donor Blank Liposomes
1. 10 ml donor blank liposomes prepared as described in Subheading 3.2 are placed on the top surfaces of the saturated microcolumns prepared as described in Subheading 3.7 and eluted with 1.5 ml buffer B at a speed of 30 drops in 45 s by using a pump. 2. The quantity of donor liposomes in the eluate is measured by liquid scintillation counting and the capturing capacity of the microcolumns for donor liposomes is estimated.
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3. The experiment is repeated nine times by using the same columns to check the reproducibility of the experiments. 3.8.2. Acceptor Liposomes
1. 10 ml acceptor liposomes prepared as described in Subheading 3.3 are placed on the top surfaces of the saturated microcolumns prepared as described in Subheading 3.7. 2. The same procedures are performed as described in Subheading 3.8.1 to measure the recovery of the acceptor liposomes in the eluate by liquid scintillation counting and the reproducibility of the experiments.
3.8.3. Mixture of Donor and Acceptor Liposomes
1. Donor blank liposomes prepared as described in Subheading 3.2 and acceptor liposomes prepared as described in 3.3 are mixed in the ratio of 1:10 (v/v). 10 ml of the mixture is placed on the surface of the saturated microcolumns and is eluted with 1.5 ml buffer B (see Note 7). 2. The same procedures are performed as described in Subheading 3.8.1 to measure the capturing capacity of the microcolumns for donor liposomes and the recovery of the acceptor liposomes in the eluate and the reproducibility of the experiments.
3.9. Example: Cyclosporin A (CyA) Transfer Between Liposomes
1. Transfer of CyA between liposomal membranes was studied using the microcolumn prepared as described in Subheading 3.7. Donor liposomes prepared as described in Subheading 3.1 and acceptor liposomes prepared as described in Subheading 3.3 are mixed in a ratio of 1:10 (v/v) at a temperature of 37°C. 2. At certain time points, 10 ml of the mixture is placed on the top surface of the saturated microcolumns and eluted with 1.5 ml buffer B as shown in Fig. 1. 3. The CyA quantity in the eluates was measured by liquid scintillation counting. The transfer kinetics of CyA, an additional model drug (cholesterol) and 3H-cholesteryl-oleoyl-ether as non-transferrable liposome-marker is shown in Fig. 2.
3.10. Liquid Scintillation Counting
The quantity of liposomes and CyA is determined by liquid scintillation counting on a Liquid Scintillation Analyzer Tri-Carb 2800 TR (Perkin Elmer, Herrenberg, Germany).
3.11. Data Analysis
Data analysis is mostly done by applying a simple mono-exponential model to the obtained data: y = y0 × (1 – e–k × t). This describes the data in all simple transfer cases quite well and delivers an understandable value k, which can easily be converted into half times. Of course, this is only a phenomenological description of the transfer processes, but elaborate models studying the mechanisms involved in the transfer from one liposome to the other are out of the scope of this chapter.
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Fig. 2. Transfer of CyA between liposomes. Donor liposomes (EPC:DCP:Chol = 7:1:2) were loaded with 14C-CyA (phospholipid:CyA = 300:1) Acceptor liposomes, composed of POPC:Chol = 8:2, were mixed with donor liposomes in a ratio of 10:1. For comparison, 14C-cholesterol and 3H-cholesteryl-oleoyl-ether transfer was also studied. Curve fits were done using the equation described in Subheading 3.11
4. Notes 1. The lipids and drug used here for liposome preparation are only an example; different lipids and drugs can be used for liposome preparation for different purposes, as long as the donor liposomes are strongly negatively charged and the acceptor liposomes neutral. 2. The experimental conditions (e.g. temperature, organic solvent) depend on lipids and drugs used in the study. The vacuum should be applied gradually in order to form a homogeneous lipid layer. After formation of the lipid layer, it should be kept under vacuum for more time to remove the organic solvent as much as possible. A very good control is the experimenter’s nose. If you are suffering from a cold at the time of the experiment, ask a colleague. 3. Linear calibration between scintillation counting and concentration of the radiolabelled lipid marker should be done to choose the suitable concentration of the lipid marker incorporated into liposome. After successfully establishing the whole procedure, the amount of radioactive tracer used can
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be reduced to the absolute minimum necessary for counting. But do these measurements regularly, especially, if you have not done it for a while. 4. In the washing process, any strong or even violent stirring or shaking of the gel must be avoided. We are not exactly sure, why this influences the quality of the results, but it does! Just treat the gel as a sensitive flower at Valentine’s Day. 5. Great care should be taken to avoid air bubbles to be induced or included in the microcolumn, which will influence the experimental results by delivering erratic results. This can be easily optically checked through the Perspex® material used for making the microcolumns. 6. The optimal amount of saturation liposomes depends on the type of liposomes, so this amount should be checked in different cases. This can be done by applying small amounts of saturation liposomes to the column and observing the eluate by turbidity measurements. A steep increase in turbidity means, that sufficient saturation liposomes have been applied. The saturation of the minicolumns is necessary, as the resin (gel) contains many lipophilic binding places, which could adsorb complete or part of liposomes during elution. There will be a small loss of drug from the acceptor liposomes to the pre-saturated minicolumn gel, but as the donor liposomes are also adsorbed and stay in equilibrium with the acceptor liposomes, the total net transfer is negligible for this type of measurement for the chosen set-up. 7. The buffer used can be pre-cooled to 4°C in order to minimise transfer processes that might occur (see Note 6) during the elution.
Acknowledgment We thank Rene Schaufelberger (Novartis Pharma Inc., Basel) for excellent technical assistance in setting up these procedures.
References 1. Papahadjopoulos D, Bangham AD (1996) Biophysical properties of phospholipids. II. Permeability of phosphatidyl liquid crystal to univalent ions. Biochim Biophys Acta 126:185–188 2. Flaten GE et al (2006) Drug permeability across a phospholipid vesicle-based barrier 2. Characterization of barrier structure, storage stability and stability towards pH changes. Eur J Pharm Sci 28(4):336–43
3. Shabbits JA, Chiu GN, Mayer LD (2002) Development of an in vitro drug release assay that accurately predicts in vivo drug retention for liposome-based delivery systems. J Control Release 84(3):161–70 4. Fahr A et al (2005) Transfer of lipophilic drugs between liposomal membranes and biological interfaces: consequences for drug delivery. Eur J Pharm Sci 26:251–265
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5. Joguparthi V, Xiang TX, Anderson BD (2008) Liposome transport of hydrophobic drugs: gel phase lipid bilayer permeability and partitioning of the lactone form of a hydrophobic camptothecin, DB-67. J Pharm Sci 97(1):400–20
6. Bienveue A et al (1985) Kinetics of phospholipid transfer between liposomes (neutral or negatively charged) and high-density lipoproteins: a spin-label study of early events. Biochim Biophys Acta 835:557–566
Chapter 2 The Use of Liposomes in the Study of Drug Metabolism: A Method to Incorporate the Enzymes of the Cytochrome P450 Monooxygenase System into Phospholipid, Bilayer Vesicles James R. Reed Abstract Although lipids are essential for the optimal activity of the cytochromes P450 monooxygenase system, relatively little is known about the membrane environment in which these enzymes function. One approach used to mimic the structural arrangement of lipids and enzymes within the endoplasmic reticulum is to physically incorporate the cytochromes P450 and their redox partners in a vesicle bilayer of phospholipids. Several methods have been devised for this purpose. This chapter describes a method in which the P450 monooxygenase system is incorporated by first, solubilizing the enzymes and lipid with sodium glycocholate. After the protein and lipid aggregates are dispersed, the detergent is removed by adsorption using BioBeads SM-2 resin which leads to the formation of bilayer vesicles of phospholipid containing incorporated cytochrome P450 and NADPH cytochrome P450 reductase. This procedure requires relatively a short preparation time, provides concentrated reconstituted systems that can be used in a wide range of applications, allows for several enzyme samples to be prepared simultaneously so that different conditions can be compared, and results in minimal loss of active enzyme. Key words: Phospholipid vesicles, Cytochromes P450, Reconstituted systems, Drug metabolism
1. Introduction The cytochromes P450 (P450) represent a ubiquitous gene superfamily comprising a diversity of isoforms that are expressed virtually in every organism in a species-specific pattern and are responsible for most xenobiotic metabolism in vivo (1). Thus the cytochromes P450 play a key role in the oxidation and clearance of most drugs and, in some instances, bioactivate toxins and promutagens to reactive intermediates that bind to cellular macromolecules V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_2, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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which, in turn, may lead to toxicity and/or the initiation of carcinogenesis (2–5). Because of these attributes, metabolism by the cytochromes P450 is relevant to a range of interests, including those of pharmacologists, toxicologists, and cancer researchers. The monooxygenase reactions catalyzed by these enzymes require electrons that can be delivered by various redox partners (6). The NADPH-cytochrome P450 reductase (reductase) is the primary redox partner in vivo and is capable of delivering both electrons needed in the catalytic cycle of the P450. Thus, the minimum enzyme assemblage needed to study monooxygenase reactions by the P450 system includes the reductase and the P450 isoform of interest. Much of the early information regarding the substrate specificity of the individual isoforms has come from studies in which the metabolites are identified after incubating the compound of interest with the active P450 monooxygenase enzyme assemblage and a source of NADPH (7–9). The P450 enzymes involved in xenobiotic metabolism and their redox partners are embedded in the membrane of the endoplasmic reticulum. As discussed in more depth later on, the membrane environment is essential for the functioning of the P450 monooxygenase system. Early attempts, over 30 years ago, to purify and characterize the various isoforms were limited by the dependence of enzyme activity on an unknown, heat-stable factor that was lost during the purification of the proteins. It was later found that the heat-stable factor was microsomal lipid (8, 10). Thus, it was determined that a lipid milieu was essential for the reconstitution of the catalytic activity of the purified enzymes. Subsequent studies have proposed that the lipid serves both as a “scaffold” to properly orient the P450 enzyme and the reductase for functional interaction (11, 12) and an effector that influences catalysis by the P450 enzyme (13). The effector role of phospholipid is evidenced by the modulation of P450 enzymatic activity at lipid:P450 enzyme ratios that are too low to facilitate the formation of liposomes. The “scaffold function” of lipid is ascribed to an additional level of modulation of enzyme activity observed at lipid concentrations at which liposomes form. Most enzymatic studies with the purified P450 enzymes use a short-chain (C-12), non-physiologic lipid, dilaurylphosphatidylcholine (DLPC). The reasons for this choice are the following: (1) the ease of preparation of the reconstituted systems with this lipid and (2) the lipid stimulates metabolism with most of the commonly studied P450 enzymes (14). However, studies have shown that the enzyme-lipid assemblages in the reconstituted systems with DLPC bear little structural similarity to the monolamellar, bilayer arrangement of lipid in the endoplasmic reticulum (15–17). Thus, it is clear that in order to truly appreciate the significance of the “scaffold” effect of the lipid on P450 metabolism, the enzymes must be physically incorporated in a vesicular bilayer of lipid.
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Several methods used to generate these P450 vesicular reconstituted systems (VRS) have been published (18–20). Unfortunately, the methods require detergent, and this makes the procedures considerably more difficult and time-consuming than those using DLPC. However, the methods have been applied extensively and have generated interesting findings regarding the scaffold effect of the lipid bilayer on P450 metabolism (21–25). The approach common to all the methods for the preparation of VRS involves the following sequence of steps: (a) drying of the lipid in order to remove organic solvents that would inactivate or denature the enzymes, (b) detergent treatment to solubilize and disperse the enzymes and the lipids, and (c) a detergent removal procedure which causes the lipids to coalesce and form lipid bilayer vesicles, and in the process, the enzymes are incorporated in the vesicles. The thermodynamics associated with the detergent solubilization of a membrane complicate the ability of the VRS preparation methods to study the relationship between P450 activity and lipid concentrations. As mentioned above, this is an important aspect when evaluating the scaffold effect of lipids on enzymatic activity. The complexity of membrane solubilization has been reviewed in detail previously (26). At low concentrations, detergent binds to lipid and partitions between the lipid and aqueous phases (27, 28). Thus, in the simplest terms at low concentrations, two populations of detergent can be assigned – that bound to lipid and that partitioned into solution as monomers. The ratio of detergent concentrations in the two forms is dependent on the partition coefficient of the detergent for the aqueous and lipid phases. The initial stage of lipid solubilization occurs when the detergent concentration is increased to a point at which the bilayers containing a mixture of detergent and lipid are lysed into mixed micelles (26). Furthermore, it has been shown that this process occurs when the monomeric detergent concentration, in equilibrium with that bound to lipid, approaches or reaches the critical micelle concentration (CMC) of the pure detergent in aqueous solution (26, 27). As the detergent concentration is increased above the threshold required for membrane lysis, the excess detergent binds primarily to the mixed micelles of lipid and detergent, and the monomeric detergent concentration remains relatively constant at the CMC of the detergent. In the process of increasing the detergent concentration above the lysis threshold, the sizes of these micelles are reduced. Upon complete solubilization of the lipid/enzyme assemblage, each component lipid and enzyme is contained in an individual micelle of detergent. Thus, the total concentration of detergent needed to lyse the membrane will depend on both the concentration of the lipid and the extent to which the detergent partitions to the lipid phase. In applying the VRS methods at different concentrations of lipid, it is necessary to first identify the “effective” detergent to
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lipid ratio (which is the ratio of the lipid-bound detergent concentration to the total lipid concentration) under a given set of optimized conditions that result in a desired degree of lipid/enzyme solubilization. The effective detergent to lipid ratio can then be used to calculate the detergent concentration needed to achieve comparable degrees of solubilization at different lipid concentrations. If the total detergent concentration (and not the lipid-bound detergent concentration) is adjusted in proportion to the lipid concentration in the methods for VRS preparation, the final VRS will have an excess detergent concentration (because both the monomeric and the lipid bound detergent concentrations will be scaled up). This in turn, likely will result in a high proportion of inactive enzyme because of the destructive effects of the excess detergent. Alternatively, if the detergent concentration is not adjusted when the lipid concentration is increased dramatically, the lipid will not be sufficiently solubilized to allow for the vesicular incorporation of the enzymes. We have tested two of the published methods, cholate gel filtration and cholate dialysis (17) and have found the active P450 enzyme tends to be extremely labile to the detergent, whereas the reductase tends not to incorporate into the vesicular fraction of the enzyme/lipid assemblage. Furthermore, preparations made using gel filtration are diluted and can be prepared only one at a time, limiting the opportunity to compare different experimental conditions, whereas, the dialysis procedure used to remove detergent is labor intensive and extremely time-consuming (3 × 12 h incubations against 3 L of dialysis buffer). Because of the limitations associated with these common methods for VRS preparation, we developed an improved technique for the preparation of P450 VRS (29). In the course of this work, we found a detergent (sodium glycocholate) that was less destructive to the P450 enzyme and utilized a more rapid way to remove the detergent by adsorption to BioBeads SM2 resin. We found this method to be superior to the two most commonly applied methods because of the following characteristics: (1) samples prepared using this method were also found to contain a higher proportion of the vesicular-incorporated reductase, (2) the detergent removal step could be carried out much more rapidly than the dialysis (2 h vs. 3 days), (3) the generated VRS samples could be easily prepared at relatively high enzyme concentrations (³5 mM), and (4) several samples could be prepared simultaneously thus allowing for the comparison of different conditions used in the reconstitution of the enzymes. We found that the VRS prepared under these conditions had very high catalytic activity relative to the reconstituted systems made using sonicated DLPC (17). The later sections of this chapter describe in detail the VRS preparation method. In addition, this chapter shows an approach that can be used to estimate the
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effective detergent to lipid ratio in order to obtain VRS preparations with comparable levels of enzyme incorporation and activity over a range of lipid concentrations. This VRS method can be used to address general questions about the scaffold effect of the lipid. However, this type of reconstituted system also provides a more favorable structural framework with which to study the interaction of P450 enzymes with the various redox partners. Furthermore, the potential for future interest in the methods to prepare VRS with reductase and P450 is tremendous given the fact that lipids are known to separate into functional domains identified as “rafts” in studies with the plasma membrane (30). Evidence is just starting to accumulate that functional, lipid microdomains may also be present in the endoplasmic reticulum (31) and thus, may be significant in regulating P450 metabolism.
2. Materials 1. 0.5 M Hepes (pH 7.5) 2. 1 M MgCl2 3. 10% (w/v) aqueous sodium glycocholate (Calbiochem La Jolla, CA); membrane phospholipid of choice in chloroform at a concentration of 10 mg/mL. We have routinely used phosphatidylcholine from bovine liver (Avanti Polar Lipids Alabaster, AL). The lipid is both light and air sensitive with a tendency to oxidation of unsaturated acyl chains. The chloroform solutions are stored at –20°C. 4. Concentrated enzyme stock solutions (>10°mM reductase or P450 enzyme, respectively) in 100°mM potassium phosphate (pH 7.4) with 20 % glycerol (enzyme stock solutions are frozen at −80°C). 5. BioBeads SM-2 (Bio-Rad Hercules, CA). 6. 5 mm syringe filter (GE Osmonics, Minnetonka, MN).
3. Methods The general method characterized and described previously (29) results in a 0.5 mL solution of VRS containing 5 mM each of reductase and P450 enzyme and a 500:1 ratio of lipid:P450 (see Note 1). An adaptation is described below the general method which allows for adjustments in the lipid:P450 ratio of the VRS.
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3.1. Preparation of the BioBeads SM-2 for Detergent Removal
This needs to be done at least two days before preparing the VRS. The use of BioBeads SM-2 to remove detergent from biological preparations has been described in depth previously (32). 1. Add 200 mL methanol to 30 g of BioBeads SM-2. 2. Stir for 15 min and collect beads by filtration with a Buchner funnel fitted with standard filter paper. 3. Immediately wash beads with another 500 mL of methanol; then repeat filtration. 4. Immediately wash beads with 1000 mL of water and repeat filtration. 5. Transfer beads to a chromatographic column and slowly wash beads with 2000 mL of water. 6. The moist beads are stored in ultrapure water at 4°C until required. When used, the beads are filtered as described above and added to the preparation containing detergent, lipid, and enzyme (supernatant derived from Step 5 of Subheading 3.3). Beads have been stored up to 3 months, periodically changing the water without noticeable problems in preparing the VRS.
3.2. General Method for Preparation of VRS: Drying the Lipid
1. Dry 1 mg of phospholipid (from a 10 mg/mL solution in chloroform) in a 1.5 mL microfuge tube overnight in a lyophilizer (see Note 2). 2. Release the vacuum on the lyophilizer by filling the chamber with N2 (see Note 3). 3. Add 50 mL of 0.5 M of Hepes (pH 7.5) and 50 mL of 10% sodium glycocholate to the tube containing dried lipid (see Notes 4 and 5).
3.3. Detergent Solubilization of Lipid and enzyme
1. Blow nitrogen over the tube opening of the solution from step 3.2.3 before capping, then bath-sonicate, and periodically vortex the tube until the solution is clear (usually 5 min). 2. While the lipid is being solubilized (step 3.3.1.), add P450 enzyme and reductase (2.5 nmol of each) to a second 1.5 mL microfuge tube and dilute to 482.5 mL with ultrapure water. 3. Add 7.5 mL of 1 M MgCl2 to the tube containing the P450 enzyme and the reductase. 4. Add the solution derived from step B.1 in approximate aliquots of 25 mL (approximately 1/4th the total volume of the solution) to the mixture of the P450 enzyme and the reductase (tube from step 3.3.3.). (See Note 6). Nitrogen is blown over the tube after the addition of each aliquot, and then the tube is capped and gently inverted to mix the enzymes with the detergent/lipid. 5. After the final addition of the solution from step B.1, blow nitrogen over the tube opening, cap the tube, invert several times, and incubate at 4°C for 1 h.
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1. Add 0.25 g of BioBeads SM-2 to the solution from step 5 of Subheading 3.3, blow nitrogen over the tube opening, cap the tube, and rock at 4°C for 2 h. 2. Draw off the solution from the Bio-Beads with a 26.5 gauge needle and filter through a 5 µm syringe filter. 3. The beads are rinsed twice with 0.2 mL aliquots of 50 mM Hepes (pH 7.4) and 15 mM MgCl2. These bead washes are filtered and added to the original bead filtrate. The final filtrate (containing the original filtrate in addition to the two bead washes) is typically around 0.65 mL in volume. 4. The sample is ready for use. An aliquot (0.1 mL) of the solution is routinely taken to determine the recovery of active P450 by determining the amount of enzyme capable of forming a ferrous CO-complex (33). A second 0.1–0.2 mL aliquot is run through a Superose 6 size exclusion column (MWCO 5,000 kDa) to determine the efficiency of enzyme incorporation. In this chromatographic step, it is assumed that the protein and lipid eluting in the void volume of the column represent the components that are incorporated into the bilayer, lipid vesicles. Reductase incorporation and activity is determined in the final preparation and in the fractions from the column by measuring the rate of the reductase-mediated reduction of cytochrome c (34). In addition, the concentration of phospholipid can also be determined in each fraction (35) in order to more accurately determine the lipid:protein ratio of the vesicular fraction.
3.5. Calculation Used to Adjust the Detergent Concentration Needed for Preparation of VRS with Different Lipid Concentrations
In adjusting our VRS method to changing concentrations of lipid, we assume that at the optimized conditions, the lipid bilayer is sufficiently solubilized to allow for the physical incorporation of the enzymes into the PC bilayer vesicles that form as the detergent is removed. If the monomeric concentration of sodium glycocholate in our optimized VRS preparation is equal to the CMC (as predicted by the studies that have examined the solubilization of membranes by detergents (discussed in the Introduction)) we can approximate the concentration of lipid-bound detergent from the total concentration used in the optimized conditions (those described in the protocol). More specifically, with a molecular weight of 488 g/mol, a concentration of 1% sodium glycocholate solution corresponds to 20.5 mM. The CMC of this detergent is 7.1 mM (36). Thus, we assume that the concentration of lipidbound detergent in the mixed detergent-lipid micelles is 13.4 mM (20.5 mM–7.1 mM). This value can be used to calculate the effective detergent to lipid ratio, and at any concentration of lipid, the concentration of detergent needed to achieve the same level of solubilization is then determined by first using the effective detergent to lipid ratio to calculate the amount of lipid-bound detergent and adding this amount to the CMC.
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An example is shown to better explain the calculation involved: Amount of lipid in the defined, optimized condition (those described in the protocol) = 1.25 mmol Amount of lipid-bound detergent = 6.7 mmol (13,400 mM * 0.0005 L) Thus, the effective detergent:lipid ratio = 5.36 mole of detergent/ mole of PC Amount of monomeric detergent in the reconstitution = 3.6 mmol (7,100 mM * 0.0005 L) If one wants to make a reconstituted system with five times the lipid concentration as that in the defined, optimized condition, the calculation is as follows: Amount of PC = 6.25 mmol Amount of bound detergent = 5.36 detergent/PC × 6.25 mmol = 33.5 mmol detergent Amount of monomeric detergent = CMC*volume = 3.6 mmol Sum of monomeric and bound detergent in the 0.5 mL reconstitution = 37.1 mmol Detergent concentration = 74.2 mM = 3.62%. Thus, one should add 3.62% (w/v) sodium glycocholate to solubilize the lipid in a VRS containing 6.25 mmol of phospholipid (see Note 7).
4. Notes 1. We have found that when the lipid concentration is £1250 mM, the bilayer vesicles of lipid do not readily form. Thus, in order to make vesicles with P450: lipid ratios £250:1, the P450 concentration should be lowered (and not the concentration of lipid) from those stated in the general method. 2. We have also prepared the VRS by drying the lipid solution under a stream of N2. The lipid must be dried slowly and for a minimum of 2 h. If the flow of the nitrogen stream is too high, residual chloroform may be “trapped” under a film of lipid. This is apparent if the detergent-solubilized lipid (Step B.1) is cloudy. If this is observed, the enzymes will not incorporate properly in the lipid vesicles of the final preparation. In general, it has been found that the reductase does not incorporate into the vesicle bilayer as readily if the lipid is dried under N2 as compared to the results obtained with lyophilized lipid. 3. Our lab flushes the lyophilizer with anaerobic nitrogen that has run through a heated column of BASF palladium catalyst (BASF RO-20). 4. Solutions are bubbled with N2 (or preferably Argon) for 2–5 min before adding to the lipid.
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5. We have also prepared the VRS using 0.5 M potassium phosphate (pH 7.25) without the MgCl2 and have not observed any significant differences in the quality of the final preparations. 6. Initially, we would add the enzymes from the concentrated stock solutions to the sonicated lipid and detergent solution after the latter was diluted with water. However, it was found that this method resulted in a much higher loss of the active P450. It seems the incremental addition of the detergent is less harmful to the P450. 7. The amount of BioBeads should not be adjusted to remove the excess detergent. The adsorptive capacity of BioBeads has been studied extensively (37), and it has been shown that the beads will also adsorb lipids. Thus, when the beads are scaled up with detergent, the excess lipid is more readily adsorbed and the desired increase in lipid concentration in the VRS is not attained. References 1. Porter TD, Coon MJ (1991) Cytochrome P-450. Multiplicity of isoforms, substrates, and catalytic and regulatory mechanisms. J Biol Chem 266:13469–13472 2. Guengerich FP (2001) Common and uncommon cytochrome P450 reactions related to metabolism and chemical toxicity. Chem Res Toxicol 14:611–650 3. Weng Y, Fang C, Turesky RJ, Behr M, Kaminsky LS, Ding X (2007) Determination of the role of target tissue metabolism in lung carcinogenesis using conditional cytochrome P450 reductasenull mice. Cancer Res 67:7825–7832 4. Iyanagi T (2007) Molecular mechanism of phase I and phase II drug-metabolizing enzymes: implications for detoxification. Int Rev Cytol 260:35–112 5. Rooney PH, Telfer C, McFadyen MC, Melvin WT, Murray GI (2004) The role of cytochrome P450 in cytotoxic bioactivation: future therapeutic directions. Curr Cancer Drug Targets 4:257–265 6. Hannemann F, Bichet A, Ewen KM, Bernhardt R (2007) Cytochrome P450 systems–biological variations of electron transport chains. Biochim Biophys Acta 1770:330–344 7. Guengerich FP (1989) Characterization of human microsomal cytochrome P-450 enzymes. Annu Rev Pharmacol Toxicol 29:241–264 8. West SB, Lu AYH (1972) Reconstituted liver microsomal enzyme system that hydroxylates drugs, other foreign compounds and endogenous substrates. V. Competition between cytochromes P-450 and P-448 for reductase in 3,
9.
10.
11.
12.
13.
14.
15.
4-benzpyrene hydroxylation. Arch Biochem Biophys 153:298–303 Saine SE, Strobel HW (1976) Drug metabolism in liver tumors. Resolution of components and reconstitution of activity. Mol Pharmacol 12:649–657 Strobel HW, Lu AYH, Heidema J, Coon MJ (1970) Phosphatidylcholine requirement in the enzymatic reduction of hemoprotein P-450 and in fatty acid, hydrocarbon, and drug hydroxylation. J Biol Chem 245:4851–4854 Ingelman-Sundberg M (1977) Phospholipids and detergents as effectors in the liver microsomal hydroxylase system. Biochim Biophys Acta 488:225–234 Taniguchi H, Pyerin W (1988) Phospholipid bilayer membranes play decisive roles in the cytochrome P-450-dependent monooxygenase system. J Cancer Res Clin Oncol 114:335–340 Causey KM, Eyer CS, Backes WL (1990) Dual role of phospholipid in the reconstitution of cytochrome P- 450 LM2-dependent activities. Mol Pharmacol 38:134–142 Balvers WG, Boersma MG, Veeger C, Rietjens IM (1993) Kinetics of cytochromes P-450 IA1 and IIB1 in reconstituted systems with dilauroyl- and distearoyl-glycerophosphocholine. Eur J Biochem 215:373–381 Autor AP, Kaschnitz RM, Heidema JK, Coon MJ (1973) Sedimentation and other properties of the reconstituted liver microsomal mixedfunction oxidase system containing cytochrome P-450, reduced triphosphopyridine nucleotidecytochrome P-450 reductase, and phosphatidylcholine. Mol Pharmacol 9:93–104
20
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16. French JS, Guengerich FP, Coon MJ (1980) Interactions of cytochrome P-450, NADPHcytochrome P-450 reductase, phospholipid, and substrate in the reconstituted liver microsomal enzyme system. J Biol Chem 255:4112–4119 17. Reed JR, Kelley RW, Backes WL (2006) An evaluation of methods for the reconstitution of cytochromes P450 and NADPH P450 reductase into lipid vesicles. Drug Metab Dispos 34:660–666 18. Taniguchi H, Imai Y, Iyanagi T, Sato R (1979) Interaction between NADPH-cytochrome P-450 reductase and cytochrome P-450 in the membrane of phosphatidylcholine vesicles. Biochim Biophys Acta 550:341–356 19. Ingelman-Sundberg M, Glaumann H (1980) Incorporation of purified components of the rabbit liver microsomal hydroxylase system into phospholipid vesicles. Biochim Biophys Acta 599:417–435 20. Schwarz D, Gast K, Meyer HW, Lachmann U, Coon MJ, Ruckpaul K (1984) Incorporation of the cytochrome P-450 monooxygenase system into large unilamellar liposomes using octylglucoside, especially for measurements of protein diffusion in membranes. Biochem Biophys Res Commun 121:118–125 21. Ingelman-Sundberg M, Blanck J, Smettan G, Ruckpaul K (1983) Reduction of cytochrome P-450 LM2 by NADPH in reconstituted phospholipid vesicles is dependent on membrane charge. Eur J Biochem 134: 157–162 22. Bosterling B, Trudell JR, Galla HJ (1981) Phospholipid interactions with cytochrome P-450 in reconstituted vesicles. Preference for negatively-charged phosphatidic acid. Biochim Biophys Acta 643:547–556 23. Kawato S, Gut J, Cherry RJ, Winterhalter KH, Richter C (1982) Rotation of cytochrome P-450. I. Investigations of protein-protein interactions of cytochrome P-450 in phospholipid vesicles and liver microsomes. J Biol Chem 257:7023–7029 24. Schwarz D, Pirrwitz J, Ruckpaul K (1982) Rotational diffusion of cytochrome P-450 in the microsomal membrane-evidence for a clusterlike organization from saturation transfer electron paramagnetic resonance spectroscopy. Arch Biochem Biophys 216: 322–328 25. Taniguchi H, Imai Y, Sato R (1987) Proteinprotein and lipid-protein interactions in a reconstituted cytochrome P-450 dependent
26. 27.
28.
29.
30. 31.
32. 33.
34.
35. 36.
37.
microsomal monooxygenase. Biochem 26: 7084–7090 Hjelmeland LM (1990) Solubilization of native membrane proteins. Meth Enzymol 182: 253–264 Jackson ML, Schmidt CF, Lichtenberg D, Litman BJ, Albert AD (1982) Solubilization of phosphatidylcholine bilayers by octyl glucoside. Biochem 21:4576–4582 Bayerl TM, Werner G-D, Sackmann E (1989) Solubilization of DMPC and DPPC vesicles by detergents below their critical midellization concentration: high-sensitivity differential scanning calorimetry, Fourier transform infared spectroscopy and freeze-fracture electron microscopy reveal two interaction sites of detergents in vesicles. Biochim Biophys Acta 984:214–224 Reed JR, Brignac-Huber LM, Backes WL (2008) Physical incorporation of NADPHcytochrome P450 reductase and cytochrome P450 into phospholipid vesicles using glycocholate and Bio-Beads. Drug Metab Dispos 36:582–588 Pike LJ (2004) Lipid rafts: heterogeneity on the high seas. Biochem J 378:281–292 Browman DT, Resek ME, Zajchowski LD, Robbins SM (2006) Erlin-1 and erlin-2 are novel members of the prohibitin family of proteins that define lipid-raft-like domains of the ER. J Cell Sci 119:3149–3160 Holloway PW (1973) A simple procedure for removal of Triton X-100 from protein samples. Anal Biochem 53:304–308 Omura T, Sato R (1964) The carbon monoxidebinding pigment of liver microsomes. I. Evidence for its hemoprotein nature. J Biol Chem 239: 2370–2378 Phillips AH, Langdon RG (1962) Hepatic triphosphopyridine nucleotide-cytochrome c reductase: isolation, characterization, and kinetic studies. J Biol Chem 237:2652–2660 Stewart JC (1980) Colorimetric determination of phospholipids with ammonium ferrothiocyanate. Anal Biochem 104:10–14 Antonian L, Deb S, Spivak W (1990) Critical self-association of bile lipids studied by infrared spectroscopy and viscometry. J Lipid Res 31:947–951 Levy D, Bluzat A, Seigneuret M, Rigaud JL (1990) A systematic study of liposome and proteoliposome reconstitution involving BioBead-mediated Triton×-100 removal. Biochim Biophys Acta 1025:179–190
Chapter 3 Use of Liposomes to Study Cellular Osmosensors Reinhard Krämer, Sascha Nicklisch, and Vera Ott Abstract When cells are exposed to changes in the osmotic pressure of the external medium, they respond with mechanisms of osmoregulation. An increase of the extracellular osmolality leads to the accumulation of internal solutes by biosynthesis or uptake. Particular bacterial transporters act as osmosensors and respond to increased osmotic pressure by catalyzing uptake of compatible solutes. The functions of osmosensing, osmoregulation , and solute transport of these transporters can be analyzed in molecular detail after solubilization, isolation, and reconstitution into phospholipid vesicles. Using this approach, intrinsic functions of osmosensing transporters are studied in a defined hydrophilic (access to both sides of the membrane) and hydrophobic surrounding (phospholipid membrane), and free of putative interacting cofactors and regulatory proteins. Key words: Osmotic stress, Osmosensing, Transport, Reconstitution, Liposome, Proteoliposome, Membrane protein, Phospholipid, Signal transduction
1. Introduction Under steady state conditions, cells maintain a certain ratio of internal versus external osmotic pressure which results in a particular cell turgor. A change in the external osmolality leads to a change in the turgor, and cells have developed sophisticated mechanisms of osmoregulation to respond to this challenge. Changing osmolality is one of the most frequent types of environmental stress for many cells, in particular microbial cells. In order to properly respond to this stress, cells harbor sensory mechanisms which perceive stimuli related to osmotic stress, transduce these stimuli into appropriate intracellular signals, or directly respond to these stimuli by appropriate actions.
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_3, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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In the case of hypo-osmotic stress, the major elements of stimulus response are mechanosensitive channels, which are dealt with in another chapter of this book. For the response to hyperosmotic stress, a major reaction is the activation of transport systems leading to the accumulation of compatible solutes. On the other hand, membrane bound sensory systems for hyperosmotic stress must be available, leading to appropriate responses in the cell on the level of transcription. In vitro systems for the study of membrane bound osmosensing transporters for compatible solutes require the presence of topologically closed vesicles and thus the use of liposomes. Functional reconstitution of transporter proteins in proteoliposomes allows the study of these systems in the absence of any other interfering mechanisms, cofactors or proteins, facilitating experimental access to both sides of the membrane, as well as to the composition of the phospholipid membrane in which the proteins are embedded. On the other hand, proteoliposomes have a number of drawbacks for the study of osmosensing in comparison to intact cells, mainly due to the inherent lack of a cell wall. As a consequence, proteoliposomes are in general more fragile than cells, and they lack turgor pressure. Consequently, osmosensory events related to turgor cannot be studied using proteoliposomes. As a further consequence, the fact that the morphological response of liposomes to hyperosmotic stress is different from cells has to be taken into consideration. Since liposomes behave as osmometers, i.e., they change their volume by water efflux according to the changing external osmotic pressure, and membranes are rigid in terms of their surface dimension; on the other, liposomes do not shrink in size but just invaginate under conditions of hyperosmotic stress to adapt the internal volume. The physical nature of stimuli, perceived by cellular osmosensors, is difficult to define, and in most cases not known (1). There are, however, a number of transport systems (1–3) in which these stimuli have been defined to a significant extent, and one of these model systems, the betaine transporter BetP from the grampositive soil bacterium Corynebacterium glutamicum, is used as an example here. In order to study osmosensing by a transport system, solute transport has to be measured. This necessarily requires some knowledge of transport kinetics and energetics, which, at least at a basic level, can be treated exactly as enzyme kinetics and energetics, and will not be further discussed here. Another basic requirement for the issue dealt with here is the proper handling of membrane proteins, their solubilization, isolation, purification, and, in particular, reconstitution. In this article, the relevant procedures of functional reconstitution of BetP from C. glutamicum will be described; however, no detailed reference to appropriate
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procedures for obtaining solubilized membrane proteins in a functional active state will be made.
2. Materials 2.1. Preparation of Preformed Liposomes for Reconstitution
1. E. coli polar lipid extract (Avanti, Alabaster, USA).
2.2. Reconstitution of BetP Into Preformed Liposomes
1. Reconstitution buffer: 100 mM phosphate buffer, pH 7.5.
2. Liposome buffer: 100 mM phosphate buffer, pH 7.5.
2. Solubilized and purified BetP protein (minimal protein concentration should be around 0.3 mg/ml, detergent concentration about 0.1% dodecyl maltoside (DDM solgrade, Anatrace, Maumee, USA). The protein concentration has to be measured with an appropriate method, e.g., using amido black (4). 3. 20% v/v Triton X-100 solution in water. 4. Spectrophotometer for monitoring liposome titration with detergent. 5. Biobeads as absorbent for detergents (Biobeads SM2, Biorad, Munich, Germany; washing of the beads in methanol/water should be carried out as described by the manufacturer). The beads are stored in water. Directly before the use for absorbing detergent, the “wet beads” are prepared by placing biobeads suspended in water on filter paper, which removes the surplus water. These “wet beads” are used for weighing. 6. Minivial ultracentrifuge for harvesting liposomes. 7. Avanti Mini Extruder System (Avanti, Alabaster, USA). 8. Polycarbonate filters, pore size 400 nm (Nucleopore, Schleicher & Schuell, Dassel, Germany).
2.3. Betaine Transport as a Response of BetP to an Osmotic Shift
1. Rapid filtration unit (FH225V, Hoefer, Holliston, USA). 2. Membrane filters (Nitrocellulose, 0.45 mm pore size; Millipore, Schwalbach, Germany). 3. Radioactively labeled 14C-betaine (synthesized from 14C-choline by the use of choline oxidase)(5). 4. Transport buffer: 20 mM phosphate buffer, pH 7.5, 25 mM NaCl (if necessary, an increased osmolality is adjusted by addition of ionic solutes (e.g., NaCl), or neutral solutes, e.g., proline, if the ionic strength is not be changed significantly), 15 mM 14 C-betaine, and 0.5 mM valinomycin. 5. Washing buffer: 0.1 M LiCl. 6. Szintillation fluid and szintillation counter.
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3. Methods In the following, functional reconstitution of BetP, an osmosensing membrane protein, and the quantification of its response to osmotic stress is described. BetP is a secondary active glycine betaine transporter from the grampositive soil bacterium C. glutamicum (3). This carrier is energetically coupled to the co-transport of two sodium ions and thus driven by the electrochemical sodium potential. The transport activity of BetP is regulated by the actual osmotic stress, and it has been shown that the carrier protein is able to sense the extent of osmotic stress without any additional factors or components (3, 6). Isolation and purification of membrane proteins in a functionally active state requiring optimization of the kind and concentration of detergent used, of the solubilization conditions (ionic strength, type of buffer, pH, temperature, etc.), on the one hand, as well as optimization of the purification procedure (type of tag used, variation of isolation conditions, etc.), on the other. The protocol described here starts with membrane protein(s) in solubilized and functionally active form. The stability of solubilized membrane proteins in terms of functionality is in general a serious problem and needs attention (see Note 1). In order to obtain an appropriate in vitro test system, the solubilized osmosensing membrane protein is reconstituted into liposomes, often prepared from E. coli lipids. There are a large number of different methods for membrane protein reconstitution; the method of choice used in most of the successful procedures recently published is the integration into preformed liposomes developed by Rigaud and colleagues (7, 8). A number of different aspects have to be tested and optimized when trying to obtain an appropriate in vitro system for testing functional properties of a reconstituted membrane protein. The preformed liposomes may differ in the type of lipids used depending on the requirement of the particular membrane protein to be reconstituted (see Note 2). Furthermore, lipid quality is an important issue, which means purity and lipid stability. The latter critically depends on how they are handled and stored (see Note 3). 3.1. Preparation of Preformed Liposomes for Reconstitution
1. Purified lipids dissolved in organic solvent (chloroform/methanol) are mixed. The mixture is evaporated to dryness in a rotary evaporator (30°C; temperature should be above phase transition temperature of the lipids). 2. Traces of solvent are removed overnight by freeze-drying (lyophilization). 3. Liposome buffer (plus 2 mM b-mercaptoethanol) is added at a concentration of 20 mg lipid/ml and lipids are suspended by stirring at room temperature (RT) for about 2 h.
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4. The lipid suspension is flushed with N2 gas, frozen in liquid N2, and stored at −80°C until use. 5. For reconstitution an aliquot of lipids is thawed gently at RT. Thereafter, liposomes are prepared by extrusion through polycarbonate filters (pore size 400 nm, multiple extrusions 17 times). 3.2. Reconstitution of BetP Into Preformed Liposomes
1. The preformed liposomes are diluted fourfold into liposome buffer. 20% v/v Triton X-100 is added stepwise (in 2–4 ml portions, thorough mixing by the use of a pipette) until detergent saturation of liposomes is achieved. Reaching the correct physical state of the liposomes for reconstitution is monitored by measuring optical density at 540 nm (7, 8). Optimal conditions for incorporation of the solubilized protein are obtained right after reaching the maximum value of light scatter at 540 nm. 2. The solubilized protein is then added drop by drop with thorough mixing. The amount of detergent added to the mixture together with the solubilized protein should be small in comparison to the amount of Triton X-100 added before. This mixture is shaken gently for 30 min at RT. 3. For removal of detergent according to the batch-procedure (7, 8), polystyrol beads (biobeads are added according to their absorption capacity for particular detergents (9), e.g., 5 mg and 10 mg of wet beads per mg of triton or DDM, respectively) are added. After shaking for 1 h at RT the same amount of beads is added and the mixture is shaken again for 1 h. The sample is then shaken overnight at 4°C with an additional twofold amount of beads. Subsequently, wet beads (same amount as for the first addition) are again added and shaken for 45 min at 4°C (see Note 4). 4. Finally, the proteoliposomes are separated from the beads with a pipette and washed twice with ice cold liposome buffer by centrifugation (Beckman Optima TLX tabletop ultracentrifuge, 350.000 g, 20 min, 4°C) before being resuspended in liposome buffer at a concentration of about 60 mg/ml lipid. The proteoliposomes are frozen in liquid N2 and stored at −80°C. 5. In order to assess the efficiency of reconstitution, the protein content of each batch of reconstituted protein has to be thoroughly measured by protein determination, e.g., using the amido black assay (4) (see Note 5).
3.3. Betaine Transport as a Response of BetP to an Osmotic Shift
1. For transport measurements, an aliquot of the proteoliposomes is thawed gently at RT, and extruded (polycarbonate filter, pore-size 400 nm) 17 times in a total volume of up to 1 ml, before being concentrated to the original volume by centrifugation (ultracentrifuge for mini vials, 350.000 g, 20 min, 20°C) and resuspended in liposome buffer. 2. In the transport assay the proteoliposomes, supplemented with internal K+ during the preparation in liposome buffer, are rapidly
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diluted into transport buffer containing valinomycin and the labeled substrate, betaine, and thoroughly mixed. In a standard experiment, the reaction is started by pipetting 5 ml of concentrated proteoliposomes into 1000 ml of transport buffer. According to the dilution ratio, this will lead to a K+-diffusion potential of about 140 mV. For measuring the activity of a particular transport protein, an appropriate driving force has to be provided. The driving force for BetP is the electrochemical Na+ potential, consequently, a sufficient amount of Na+ has to be present in the medium, in addition to the membrane potential, supplied here as K+-diffusion potential. 3. For each time-point in the following kinetic assay, 200 ml samples are used. These samples are withdrawn from the transport assay and instantly filtered through membrane filters in a filtration unit. The time intervals chosen depend on the velocity of the reaction under study; experienced experimenters are able to handle time points down to 5 s intervals. The filters are washed immediately twice with about 2 ml of washing buffer each (see Note 6). 4. After finishing the kinetic experiment, the filters are removed with tweezers and transferred into a scintillation vial each, which is filled with scintillation fluid, and counted. 5. The amount of betaine taken up into the proteoliposomes by the reconstituted carrier protein at the chosen time-points is calculated on the basis of the specific radioactivity applied in the transport buffer and the amount of protein present in the 200 ml sample of proteoliposomes. Protein determination should be carried out for each experiment, since variable losses during the extrusion procedure may give rise to variation. 6. Substrate uptake activity is interpreted in terms of transport kinetics, and specific transport rates are calculated (see Note 7). Beside the activity of the reconstituted protein, the result of uptake measurements may depend on a variety of other factors (see Note 8). 7. Testing a transport protein for its osmosensing function will need variation of the medium on both sides of the membrane, with respect to concentration and quality of solutes, e.g., ionic and nonionic solutes, or type of cations and anions, respectively. The solute composition in the interior of the proteoliposomes can be varied during liposome preparation (Step 1 of Subheading 3.3.). The composition of the internal compartment can also be changed after proteoliposome preparation by applying additional freezing and thawing cycles in a medium of changed composition followed by liposome generation using the extrusion procedure. The lipid composition can also be varied during the preparation of the preformed liposomes.
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Additional lipids can be integrated by repeated cycles of freezing and thawing mixtures of (proteo) liposomes with different lipid compositions. 8. It should be noted, that there are two different types of experiments testing the dependence of a membrane inserted protein on variation of osmotic properties. On the one hand, the external osmolality (or solute composition) can be changed at the start of the transport experiment, leading to an osmotic gradient and a subsequent volume change of the proteoliposomes. A variation of solute concentration or composition on both sides of the membrane at the same time, i.e., during proteoliposome preparation, on the other hand, changes the osmotic pressure without introducing an osmotic gradient and without concomitant volume changes (6).
4. Notes 1. The stability of solubilized membrane (carrier) proteins is a serious issue for the success of this type of experiment. The conditions of solubilization, isolation, and storage, if necessary, have to be optimized carefully. Most transport proteins are notoriously unstable in the solubilized state, but, in general, rather stable once properly inserted into a phospholipid membrane. Consequently, they should be purified quickly and reconstituted into proteoliposomes right after purification. 2. Often, selected lipids have to be used for the reconstitution of a particular membrane protein, in order to reach optimum reconstitution efficiency or functionality.. Different membrane proteins have different preference for lipids, with respect to the phospholipid headgroup, the headgroup charge, and the fatty acid composition. Typical lipids to start with are lipids from E. coli membranes, asolectin (soy bean lipids), or egg yolk lipids. Once, a basic activity of the reconstituted protein is observed, variation of lipids will normally improve the results. An obvious strategy to overcome some of these problems is the use of synthetic phospholipids; however, for this, a balanced composition of unsaturated (essential for most membrane proteins) and saturated fatty acids is required. For unknown reasons, effective reconstitution into synthetic lipid mixtures seems to be more difficult compared with natural lipids, e.g., from E. coli. In any case, the fact that proteins in general prefer mixtures of phospholipids, both with respect to the headgroup and the fatty acid composition over the membranes prepared from single compounds has to be taken into account.
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3. The same as stated for handling of the protein is also true for handling of lipids. Always the best lipids available should be used. In general, Avanti is a good choice as a supplier for lipids, however, be careful for variation of lipid quality from batch to batch. After having established an optimized protocol for reconstitution, try to examine different batches of the lipids of choice and then buy a stock of lipids of the optimal batch. In general, if the proteoliposome test system, after having been successfully established, fails to work for unknown reasons, the quality of lipids is a good candidate to think about. For this reason, it is recommended always that part of a lipid batch which was proven to function is stored away, to serve as a control for later experiments. Other reasons for failure, of course, are the functionality of the solubilized protein, and, rather often, the efficiency of detergent removal by adsorbance to biobeads, which directly affects the efficiency of reconstitution and/or the permeability properties of the proteoliposomes. 4. The stability of the lipids used is a serious issue, too. The manufacturer supplies lipids in general sealed under nitrogen gas; it is recommended that the same condition is kept when storing lipids in the lab, preferably at −80°C. Useful recommendations for lipid handling and storage are found in (10–12). 5. Detergent removal is an important step during reconstitution and is prone to a number of difficulties. Complete removal of detergent from the proteoliposomal sample can simply be tested by the absence of foam after shaking. It has to be taken into account that the beads not only absorb detergents, but all amphiphilic (and even hydrophilic) compounds, in general in the order detergent > lipid > protein. It is thus important to limit the amount of biobeads used for absorption of the detergent in order not to lose too much protein. An alternative strategy partially avoiding this problem is to presaturate the biobeads with lipids (7, 8). 6. The efficiency of reconstitution is a complex problem. First, it refers to the amount of protein successfully integrated. This value may vary with the kind of detergent and lipid used, as well as with particular aspects of the reconstitution procedure. It has to be quantified by protein determination in the proteoliposomes, e.g., by the amido black method (4). In general, the efficiency of protein insertion decreases with increasing protein/lipid ratio during reconstitution. Second, reconstitution efficiency also means the fraction of functionally active protein in comparison to the total amount of integrated protein. Only the latter is measured by protein determination. This ratio is rather difficult to determine and in general needs additional methods, e.g., quantification of substrate binding. Third, protein orientation may be an issue. In most cases, the
Use of Liposomes to Study Cellular Osmosensors
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orientation of the reconstituted membrane protein is not known. This, however, may be of crucial significance for interpreting its function in the in proteoliposomal in vitro system, in particular for transport proteins being dependent on oriented driving forces. There are a number of methods available for defining the orientation of a membrane-inserted protein, e.g., the use of antibodies, proteases, and impermeable labeling reagents for specific amino acids. 7. Although seeming apparently simple, fast, and efficient liposome filtration is a crucial step in the whole procedure. It should be taken into account that filtration, as used here, is to a large extent adsorption to the filter material, in view of the pore size of the filters used. Before application to the filtration unit, the membrane filters have to be presoaked in washing buffer, in order to avoid labeled substrate to be soaked into the external ring of the filter which is fixed in the filtration unit. Before running the kinetic experiment, the filters should be briefly soaked by application of vacuum. When applied to the filter during transport kinetics, the proteoliposome sample should quickly be distributed over the entire filter surface. The same holds true for the washing solution, which preferentially should be applied along the walls of the filtration chamber. 8. Transport kinetics has been elaborated in relevant text books. In general, when establishing transport kinetics of a particular carrier protein, a time course including several experimental points (five at least) has to be used in order to find out in which time window the uptake kinetics is reasonably linear with time. Once established, experiments are frequently carried out using two time points only within the linear kinetic phase, because of the requirement of rapid assays and/or frequently because of a limited range of linearity (mainly due to the small size of the proteoliposomes used). In order to provide sufficient experimental reliability, these two point-kinetics have to be carried out in multiple sets, e.g., fivefold. When changing experimental conditions, multiple time point kinetics has to be applied again to verify linearity. It should furthermore be mentioned that the linearity of a transport assay also depends on the amount of substrate taken up during the time course monitored in the kinetic experiment. For this reason, the amount of labeled substrate taken up at the end of the time course should not exceed one third of the added label. This amount, however, may sometimes be much lower, if higher substrate concentrations have to be used because of a relatively low substrate affinity of the transport protein under study. 9. It should be pointed out that the result of the kinetic analysis of a reconstituted transport protein not only depends on the functionality of the protein, and the experimental conditions
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applied (e.g., kind and extent of driving force, pH, ionic strength, kind of lipid provided, etc.), but to a large extent also on the quality of liposomes used. This refers, on the one hand, to the integrity and stability of liposomes, which in the presence of residual detergent, or high amounts of protein integrated, can be an issue. The integrity of the proteoliposomes used can be tested by the incorporation of calceine (13). Another reason for unexpected failure of transport experiments is partial leakiness of the proteoliposomes to small ions, e.g., protons, leading to a quick collapse of the driving force. This will in general not lead to leakiness of the larger solutes, like substrates (or calceine), and is frequently caused by residual amounts of detergent present. Furthermore, the size of the liposomes used is of critical significance. An early deviation of transport kinetics from linearity may indicate the presence of liposomes too small in size for the particular transport experiment. The impact of the liposome size on the shape of the transport kinetics observed is also a major reason for using liposomes sized by the extrusion procedure through polycarbonate filters. All other methods of proteoliposome formation lead to mixtures of vesicles with a broad size distribution. This fact will have a complex influence on the resulting transport kinetics. References 1. Wood JM (1999) Osmosensing by bacteria: signals and membrane-based sensors. Microbiol Mol Biol Rev 63:230–262 2. Poolman B, Spitzer JJ, Wood JM (2004) Bacterial osmosensing: roles of membrane structure and electrostatics in lipid-protein and protein-protein interactions. Biochim Biophys Acta 1666:88–104 3. Morbach S, Krämer R (2003) Impact of transport processes in the osmotic response of Corynebacterium glutamicum. J Biotechnol 104:69–75 4. Schaffner W, Weissmann C (1973) A rapid, sensitive and specific method for the determination of protein in dilute solution. Anal Biochem 56:502–514 5. Landfald B, Strøm AR (1986) Choline-glycine betaine pathway confers a high level of osmotic tolerance in Escherichia coli. J Bacteriol 165:849–55 6. Rübenhagen R, Morbach S, Krämer R (2001) The osmoreactive betaine carrier BetP from Corynebacterium glutamicum is a sensor for cytoplasmic K+. EMBO J 20:5412–5420
7. Rigaud J-L, Levy D (2003) Reconstitution of membrane proteins into liposomes. Methods Enzymol 372:65–86 8. Paternostre MT, Roux M, Rigaud J-L (1988) Mechanism of membrane protein insertion into liposomes during reconstitution procedures involving the use of detergents. Biochemistry 27:2668–2677 9. Rigaud J-L, Mosser G, Lacapere J-J, Olofsson A, Levy D, Ranck J-L (1997) Bio-beads: an efficient strategy for two-dimensional crystallization of membrane proteins. J Struct Biol 118:226–235 10. Zuidam NJ, Crommelin DJ (1995) Chemical hydrolysis of phospholipids. J Pharm Sci 84:1113–9 11. Hernández-Caselles T, Villalaín J, GómezFernández JC (1990) Stability of liposomes on long term storage. J Pharm Pharmacol 42:397–400 12. Torchilin VP, Weissig V (2003) Liposomes, 2nd edn. Oxford University Press, New York 13. Allen TM, Cleland LG (1980) Serum-induced leakage of liposome contents. Biochim Biophys Acta 597:418–426
Chapter 4 Studying Mechanosensitive Ion Channels Using Liposomes Boris Martinac, Paul R. Rohde, Andrew R. Battle, Evgeny Petrov, Prithwish Pal, Alexander Fook Weng Foo, Valeria Vásquez, Thuan Huynh, and Anna Kloda Abstract Mechanosensitive (MS) ion channels are the primary molecular transducers of mechanical force into electrical and/or chemical intracellular signals in living cells. They have been implicated in innumerable mechanosensory physiological processes including touch and pain sensation, hearing, blood pressure control, micturition, cell volume regulation, tissue growth, or cellular turgor control. Much of what we know about the basic physical principles underlying the conversion of mechanical force acting upon membranes of living cells into conformational changes of MS channels comes from studies of MS channels reconstituted into artificial liposomes. Using bacterial MS channels as a model, we have shown by reconstituting these channels into liposomes that there is a close relationship between the physico-chemical properties of the lipid bilayer and structural dynamics bringing about the function of these channels. Key words: MscL, MscS, NMDA, Liposome reconstitution, Patch clamp, EPR spectroscopy, FRET spectroscopy, Confocal microscopy
1. Introduction MS channels present a classical example of ion channels for which the composition and properties of the surrounding lipid matrix are crucial for their function. In a nutshell, the MS channels’ leitmotif is “force from lipids” (1, 2). Studies on prokaryotic (MS) channels over many years have demonstrated that the lipid bilayer transmits the mechanical force directly to this type of MS channels enabling them to detect changes in osmotic forces acting upon these microorganisms (3, 4). Among prokaryotic MS channels, the best characterized channels are MscL and MscS of
V. Weissig, (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_4, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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Escherichia coli. Bacterial MS channels were the first channels shown to sense directly the membrane tension in the lipid bilayer caused by the external mechanical force applied to bacterial cell membrane (5,6). Recent evidence indicates that, similar to prokaryotic MS channels, some MS ion channels found in membranes of a variety of eukaryotic cells, such as TREK-1, TRAAK and TRPC1, can also be gated by the bilayer mechanism (7–10). The glutamate-gated NMDA receptor channel has also been shown to exhibit mechanosensitivity upon reconstitution into artificial liposomes (11). The question of how MS channel proteins detect mechanical stresses in the lipid bilayer of cellular membranes and what makes these channels undergo conformational changes in response to membrane tension has been addressed in several studies examining the physical principles underlying gating of MscL channels by bilayer deformation forces (12–14). In one study, a combination of functional patch-clamp experiments with structural electronparamagnetic resonance (EPR) spectroscopic experiments allowed the drawing of the following conclusions about forces from lipids that affect both MS channel structure and function. First, the importance of the hydrophobic surface match for MscL mechanosensitivity was shown to result from a membrane-tension-induced bilayer thinning, which stabilizes intermediate conformations of MscL leading to channel opening due to a better hydrophobic match of the thinner bilayer with the open channel compared with that of the closed conformation. Second, geometric shape inequalities between lipid molecules asymmetrically distributed between the two leaflets of the lipid bilayer were shown to be critical in triggering the opening of the MscL pore of >25 Å in diameter (12, 13). These results have been confirmed by a FRET spectroscopic study in which asymmetric distribution of lipid molecules between the two leaflets of the lipid bilayer was shown to open MscL (14). Both studies thus indicate that the mechanism of mechanotransduction in MS channels is defined by both local and global asymmetries in the transbilayer pressure profile at the lipid protein interface. The role of bilayer thickness and the concept of hydrophobic mismatch in MscL functionality have further been investigated in studies exploring the effect of static magnetic fields (SMF) of moderate intensity (~400 mT) on the open probability of MscL reconstituted in phospholipid bilayers (15, 16). Results of these studies suggest that due to co-operative superdiamagnetism of phospholipid molecules, SMF most likely cause a rearrangement of the lipids resulting in a change of bilayer thickness. Given that not only ion channels but also other signaling membrane proteins may experience local change in bilayer thickness and/or curvature as a consequence of association with lipid microdomains or chemical modification of the lipid bilayer, the findings from studies of bacterial MS channels may also have implications
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for the functioning of all kinds of signaling membrane proteins including ion channels. The production of the MscL or MscS channel entails the protein expression of its subunit which then readily self-assimilates into the homo-multimeric functional channel within the bacterial membrane, without any post-translational requirements. Subunit expression with a fused purification tag allows simplified isolation of the channel (subunits) from bacterial membrane preparations using commonly available commercial purification supplies. Suitable tag expression vectors are described further below. Two affinity expression tag systems commonly used include charged affinity and specific enzymatic affinity. Charged affinity relies on a gene construct to produce a fusion with six histidine amino acids to either the amino or carboxyl terminal of the protein. This allows the fusion protein to bind to nickel or cobalt chelated to sepharose beads. Capture to beads allows further washing (purification) and final elution with a charged competitor (imidazole) solution. Alternatively for higher purity at lower yields and for a product with no artificial charges, enzymatic affinity may be used. This involves a fusion product with (typically) glutathione-S-transferase (GST) which allows specific tight binding to sepharose beads coupled to the GST target, glutathione. The protein of interest is cleaved off GST with (typically) thrombin via a target site that has been designed between the two fusion products from the onset. It is also possible to design affinity histidine tags that are also cleavable (not described). It should be noted that thrombin cleavage will leave part of its recognition sequence on the final product, and other amino acids from cloning vectors are typically also attached to the final protein, including any (uncleaved) 6 × His sequence.
2. Materials 2.1. MscL Production with GST or 6 × His Affinity Fusion Tags
1. Luria Broth (LB) (10 g/l bacto-tryptone, 5 g/l yeast extract, 10 g/l NaCl) autoclave sterilized.
2.1.1. Materials Common for Both MscL Production Methods
3. Ampicillin dihydrate antibiotic, 100 mg/ml in water, 0.22 mm filter sterilized.
2. Autoclave sterilized 100 ml and ~2,800 ml culture flasks.
4. 1 M Isopropyl b-D-1-thiogalactopyranoside (IPTG) in water, 0.22 mm filter sterilized. 5. Phosphate buffered saline (PBS) (10×): 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4 2H2O and 2.4 g KH2PO4 into 1 L of water. On diluting adjust pH with HCl or NaOH.
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6. Flat bristle brushes (in the order of approximately 5 mm and 8 mm width). 7. Phenylmethanesulphonylfluoride (PMSF), 20 mg/ml in isopropanol. 8. Deoxyribonuclease I (DNase) (e.g., Sigma DN25). 9. n-Dodecyl-b-D-Maltopyranoside (DDM), 200 mM in water (Anatrace D310). 10. Bradford Reagent (Coomassie dye binding protein assay, Protein dye reagent for quantitation). 2.1.2. MscL Production with GST Affinity Fusion Tag
1. AW737KO MscL knock-out strain transformed with N-terminus gluthione-S-transferase tagged MscL (pGEX-2T plasmid, GE) expression construct. 2. GST Wash Buffer: 1× PBS pH 7.4 with 1 mM DDM 3. 10 ml chromatography columns (e.g., Biorad 731–1550) 4. Glutathione sepharose (Bioworld or GE), washed following manufacture’s protocol, but with 1 mM DDM detergent. 5. Thrombin. (GE or Sigma) Dissolved and stored as per supplier’s instructions.
2.1.3. MscL Production with 6×His Affinity Fusion Tag
1. M15 E.coli [with pREP4 plasmid (Qiagen)] and with N-terminus 6×His-tagged MscL (pQE30 plasmid, Qiagen) or C-terminus 6×His-tagged MscL (pQE70 plasmid, Qiagen) expression construct. 2. Kanamycin sulphate antibiotic, 25 mg/ml ml in water, 0.22 mm filter sterilized. 3. Talon metal affinity resin (Clonetech 635502), washed following manufacture’s protocol, but with 1 mM DDM detergent. 4. 20 ml chromatography columns (e.g., Biorad 732–1010) 5. His Wash Buffer, 1× PBS pH 6.0, 1 mM DDM, 5 mM imidazole. 6. His Elution Buffer, 1× PBS pH 6.0, 1 mM DDM, 500 mM imidazole. 7. Millipore Amicon Ultra-15 Ultracel 10 k centrifugal filter device. Washed and used to manufacturer’s instructions.
2.2. MscS Production 6×His Affinity Fusion Tags
1. M15 E. coli [with pREP4 plasmid (Qiagen)] and transformed with pRARE plasmid (Merck), made competent ready for transformation. 2. N-terminus 6 × His-tagged MscS (pQE30 plasmid, Qiagen), or C-terminus 6 × His-tagged MscS (pQE70 plasmid, Qiagen) expression construct. The C-terminus construct gives greater yield.
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3. Luria Broth (LB): 10 g/l bacto-tryptone, 5 g/l yeast extract, 10 g/l NaCl, autoclave sterilized. 4. Autoclave sterilized 100 ml and ~2800 ml culture flasks. 5. Ampicillin dihydrate antibiotic, 100 mg/ml in water, 0.22 µm filter sterilized. 6. Kanamycin sulphate antibiotic, 25 mg/ml ml in water, 0.22 µm filter sterilized. 7. Chloramphenicol antibiotic 25 mg/ml in ethanol. 8. Glycerol. 9. Isopropyl b-D-1-thiogalactopyranoside (IPTG) 1 M in water, 0.22 mm filter sterilized. 10. Flat bristle brushes (in the order of approximately 5 mm and 8 mm width). 11. Phenylmethanesulphonylfluoride (~115 mM) in isopropanol.
(PMSF),
20
mg/ml
12. Deoxyribonuclease I (DNase) (e.g., Sigma DN25). 13. n-Dodecyl-b-D-Maltopyranoside (DDM), 200 mM in water (Anatrace D310). 14. Talon metal affinity resin (Clonetech 635502), washed following manufacture’s protocol, but with 1 mM DDM. 15. 20 ml chromatography columns (e.g., Biorad 732–1010). 16. Phosphate buffered saline (PBS), (10×): 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4 2H2O and 2.4 g KH2PO4 into 1 L of water. On diluting to make 1× adjust pH with HCl or NaOH. 17. Solubilization Buffer: 1× PBS pH 7.5, 1 mM PMSF, 10% v/v glycerol. 18. MscS Wash Buffer 1: 1× PBS pH 7.5, 1 mM DDM, 10% v/v glycerol. 19. MscS Wash Buffer 2: 1× PBS pH 6.0, 1 mM DDM, 10% v/v glycerol, 5 mM imidazole. 20. MscS Elution Buffer: 1× PBS pH 6.0, 1 mM DDM, 10% v/v glycerol, 300 mM imidazole. 21. Millipore Amicon Ultra-15 Ultracel 10 k centrifugal filter device. Washed and used to manufacturer’s instructions. 2.3. NMDA Protein Expression and Purification 2.3.1. Cell Culture and NMDA Receptor Channel Constructs
1. Cell culture used for expression of NMDA receptor NR1 and NR2 subunits. (a) SF9 Spodoptera frugiperda (insect cell culture) (1 or 2 L). (b) XL99, a suspension CHO-K1 (Chinese Hamster Ovary cells) cell line adapted to growth in suspension in serum-free media (1 L).
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2. Growth media (a) Sf-900II (Invitrogen) serum-free media for insect cell culture. (b) CHO-S-SFMII (Invitrogen) media for CHO mammalian cell culture. 3. NMDA receptor channel constructs. (a) 6 × His-tagged NR1a subunit (C-terminus, pENTR/ D-TOPO or pcDNA 3.3-TOPO plasmids). (b) 6 × His-tagged NR2A subunit (C-terminus, pENTR/ D-TOPO or pcDNA 3.3-TOPO plasmids). 2.3.2. Buffers for Purification of NMDA Receptor Channel Proteins
1. Cell Lysis and Solubilization Buffer (100 ml, pH 7.3): 20 mM sodium phosphate 500 mM NaCl, 10 mM imidazole, 3% octyl glucoside (OG), 20 mg/ml DNase protease inhibitor tablet (Roche), 1.25 ml of TALON resin (Clontech) added after solubilization. 2. Wash Buffer (50 ml, pH 7.3): 20 mM sodium phosphate, 500 mM NaCl, 20 mM imidazole, 1.5% OG, protease inhibitor tablet (Roche). 3. Elution Buffer (4 ml, pH 7.3): 20 mM sodium phosphate, 500 mM NaCl, 150 mM imidazole, 1.5% OG, protease inhibitor tablet (Roche). 4. Gel Filtration Buffer (50 ml, pH 7.5): 20 mM sodium phosphate, 300 mM NaCl.
2.4. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE)
1. An electrophoresis vertical mini electrophoresis rig system and if desired compatible pre-cast 12% polyacrylamide denaturing gels of 1 mm thickness, ten well for protein analysis, as well as associated running buffers as per manufacture’s specifications. For manually cast SDS-PAGE gels, the following materials are required. 2. Distilled water (not from a source utilizing ultraviolet light sterilization.) 3. Tris(hydroxymethyl)aminomethane (TRIS), electrophoresis grade. 4. 1.5 M Tris-HCl pH 8.8. 5. 0.5 M Tris-HCl pH 6.8. 6. 10% Sodium Dodecyl Sulfate (Sodium lauryl sulfate) (SDS) 7. 30% Acrylamide/Bis Solution, 37.5:1 (e.g., Biorad 161–0158). The non-polymerized acrylamide monomer solution is a potent neurotoxin hazard and must be handled appropriately. Nonvolatile when supplied as a solution. 8. 10% (in water) Ammonium persulphate (APS), stored in ~500 ml aliquots stored at −20°C. 9. Tetramethylethylenediamine (TEMED).
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10. 5× Elpho Buffer: 15.1 g Tris, 94 g Glycine, double distilled water to 900 ml, mix, add 50 ml 10% SDS, top up to 1 L. 11. 4× Sample Buffer: 1.52 g Tris-HCl, 20 ml glycerol, 2 g SDS, 2 ml 2-mercaptoethanol, 1 mg bromophenol blue. 12. Protein molecular weight marker (suitable is Fermentas #SM0431). 13. Coomassie Stain: 1.25 g (Coomassie) Brilliant Blue R-250, 225 ml methanol, 50 ml glacial acetic acid, double distilled water to 500 ml. 14. Destain (300 ml methanol, 10% glacial acetic acid, water to 500 ml). 2.5. Liposome Reconstitution 2.5.1. Dehydration/ Rehydration (D/R) Method
1. Chloroform, AR grade. 2. Lipid. Azolectin (L-a-Phosphatidylcholine from soybean, Type II-S, Sigma P5638, or Type IV-S, Sigma P3644) or pure lipids (e.g., Phosphatidylcholine (PC) with acyl lengths of 16 to 20, or PC mixture with phosphatidylglycerol (PG) and/or Phosphatidylethanolamine (PE) pure lipids from Avanti Polar Lipids, Inc, Alabaster) 10 mg/ml in chloroform, stored at −20°C. 3. Nitrogen (N2) Industrial Grade Dry Gas supply, fed through a glass pipette for a nitrogen stream. 4. Purified mechanosensitive channel protein, as described within this chapter. 5. Dehydration/Rehydration Buffer Solution (D/R Buffer): 200 mM KCl, 5 mM HEPES (pH 7.2 adjusted with KOH). 6. Bio-Beads SM-2 Adsorbent (Biorad #152–8920) prepared by washing thrice (on shaker/rocker) in methanol for 30 min each. Remove methanol. Rinse briefly in distilled water by shaking three times, followed by two 30 min washes in distilled water. Decant and cover beads with fresh water. Degas by vacuum for 3 h to overnight. Store at 4°C. Before use, blot the required amount of beads with tissue paper to remove excess moisture. 7. Ethanol, AR grade. 8. Glass microscope slides, cleaned with ethanol, and dried. 9. Filter paper or similar absorbent paper/tissue type.
2.5.2. Sucrose Method
In addition to materials 1 to 6 listed above in Subheading 2.5.1, sucrose is also required.
2.6. Patch-Clamp Recording From MS Channels Reconstituted into Liposomes
1. Borosilicate glass calibrated microcapillary tube (e.g Drummond Scientific Co., Broomall, PA, 100 ml Calibrated Pipets [sic] cat: 2-000-100).
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2. Recording Solution: 200 mM KCl, 40 mM MgCl2, 5 mM HEPES (pH 7.2 adjusted with KOH). 3. 0.22 mm syringe filter. 2.7. Confocal Microscopy of Liposome Reconstituted MS Channels
1. Single cysteine mutant expression construct and the resulting protein purified using deoxygenated Wash Buffer and Elution Buffer (see text). 2. Labeling Buffer: Deoxygenated PBS pH 7.5, with 1 mM DDM (n-Dodecyl-b-D-Maltopyranoside (DDM) (Anatrace D310)) 3. 40 mM tris(2-carboxyethyl)phosphine (TCEP) in distilled water. 4. Alexa Fluor® 488 C5-maleimide (AF488) (Invitrogen Molecular Probes A10254). Resuspended in water, separated in 100 mg aliquots which are then lyophilized/desiccated and stored at −20°C in a light sealed container. 5. Optional b-mercaptoethanol. 6. Dialysis cassette, 10 K molecular weight cut off, 3 ml capacity (e.g., Pierce, Thermo Fisher Scientific Inc, # 66380). 7. Dialysis Buffer: PBS pH 7.5 with 0.5% Triton X-100, chilled to 4°C for use. A 10× stock may be preferentially made, adjusted to pH 7.5 on dilution. 8. Confocal microscope with 60× objective lens, and appropriate excitation laser. 9. Cover slips and glass slides.
3. Methods Different constructs of MscL and MscS (GST fusion proteins or 6×His-tagged proteins) have been used for liposome reconstitution. The former is more suitable for testing channel function by the patch-clamp technique while the latter is appropriate for structural studies using EPR or FRET spectroscopy. In the case of NMDA receptor channels a 6×His-tagged protein has been used. The amount of protein required for reconstitution depends on the protein-to-lipid ratio (w/w for patch clamp experiments or mol/mol for structural EPR and FRET experiments), reconstitution method, and the type of experiment. For MscL reconstitution we have also found that the lipid-to-protein ratio depends on which construct is being used. Particularly, for MscS we have found that the lipids commonly used for electrophysiological measurements, such as DOPC or E. coli polar lipids cannot be used for MscS structural determination through spectroscopy, since they promote two-dimensional aggregation, regardless of the expression and purification conditions (20). This has not
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been tested in detail for experiments with MscL or NMDA. Assuming that dehydration/rehydration (D/R) method is used (see further) the number of active channels per liposome patch varies with the protein-to-lipid ratio (e.g., 1:1000 w/w) and is higher when the GST-MscL construct is reconstituted compared to 6 × His-tagged MscL constructs. If a sucrose reconstitution method is used (see further) less protein is required and higher percentage of liposome patches with active channels become available for experiments. Using the Vesicle PrepPro (Nanion Technologies GmbH) (see further) MscL (1/900 w/w protein-to-lipid ratio) can be reconstituted into liposomes in less than three hours total preparation time. Usually 5 micrograms of any of the MS channels described here is sufficient for several liposome preparations using the D/R method. If protein is scarce, to minimize the amount of protein required for reconstitution smaller amounts of lipids (ca. 2 mg) can be used per single liposome prep, which usually gives enough material for two days of patch clamping. 3.1. MscL- GST or His Tag Fusion Protein Purification
This procedure takes several days, thus it is advisable to start at the beginning of a week. Days 2 and 3 may be combined into a single long day.
3.1.1. Day 1: Overnight Starter Culture
1. Add 10 ml glycerol stock of bacterial strain (with expression construct) into a 100 ml culture flask with 12 ml LB and 12 ml 100 mg/ml Ampicillin. For pQE His constructs in M15 strain, also add 12 ml 25 mg/ml kanamycin sulphate. 2. Incubate at 37°C at 240 rpm (for ¾ inch orbit) in an orbital shaker incubator.
3.1.2. Day 2: Growth, Expression, and Cell Collection
1. Add 10 ml of overnight culture into a ~2,800 ml culture flask with 1000 ml LB and 1 ml 100 mg/ml Ampicillin. For pQE 6 × His constructs in M15 strain, also add 1000 ml 25 mg/ml kanamycin sulphate. 2. For aeration, seal the flasks with tissue paper (such as low lint fine task wipers) and incubate at 37°C at 150 rpm (for 2 inch orbit) in an orbital shaker incubator. 3. When optical density of the culture (OD) A600 = 0.8 (occurs in ~2–2.5 h) induce expression with the addition of 1000 ml 1 M IPTG. Also add 500 ml 100 mg/ml ampicillin. (Hint: For multiple samples, hold a considerably faster culture(s) at 4°C until others catch up before adding IPTG etc.). 4. Incubate with shaking for a further 4 h. 5. Centrifuge culture at around 8,000 ´ g for 15 min at 4°C. Discard supernatant. 6. For every 10 g of cell pellet resuspend in 10 ml PBS (with a ~8 mm wide brush). 7. Store the resuspension at −20°C or continue with day two.
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3.1.3. Day 3: Cell Disruption and Membrane Processing
1. Thaw sample and keep on ice. 2. Add roughly a “match head” amount of DNase, and 1/100 volume of 20 mg/ml PMSF. 3. French press (Thermo Electron Corporation) twice at 16,000 PSI. 4. Remove debris, unbroken cells, or inclusion bodies by high speed centrifugation at 8,000 ´ g (k-Factor ~3,600) for 30 min at 4°C. Collect supernatant. 5. Pellet membranes by ultracentrifugation at 235,000 × g (k-Factor ~133) for 3 h at 4°C. Discard supernatant. 6. Suspend pellet in 5 ml PBS with a 5 mm width brush. 7. Add a further 17 ml PBS and 880 ml 200 mM DDM. 8. Solubilize on a roller wheel overnight at 4°C.
3.1.4. Day 4: Clarification and Column Binding and Purification
1. Clarify the solubilization by ultracentrifugation at 10,000 for 20 min at 4°C. Recover supernatant to use. 2. For 6×His fusion constructs add 2 ml prepared cobalt-resin beads. For GST fusion constructs add 1 ml prepared glutathione sepharose beads. 3. Incubate on a rocker or roller for 2 h at room temperature. 4. Pour the cobalt resin suspension into 20 ml chromatography columns. Pour the GST resin suspension into 10 ml chromatography columns. Release bottom caps when resin has settled somewhat after several minutes, and allow all liquid to flow out as discard. 5. Wash the cobalt resin twice with 25 ml His Wash Buffer. Wash GST resin twice with 10 ml GST Wash Buffer. For GST fusion protein, recap column and mix beads by pipetting with 1 ml GST Wash Buffer and transfer to a 2 ml centrifugation tube (see Note 1). Add 20 units of thrombin, and incubate overnight at room temperature. (B) For His tag protein elute with 15 ml His Elution Buffer into/or then transfer to a centrifugal filter device for concentration down to or just under 1 ml. Centrifuge to concentrate. If desired, imidazole can be removed, or reduced, by careful layering (to avoid mixing) with ~10 ml Wash Buffer and concentrated again. Repeat the “reverse decanting” process with another 10 ml Wash Buffer. Protein is ready for analysis and / or for use. Ensure protein is at less concentration than 500 µg / ml to avoid precipitation. Store at 4 °C. 6. Protein concentration can be determined by the Bradford assay at an absorbance of 595 nm, (typically measure 33.3 ml in 1 ml reagent) following the supplier’s instructions, referenced to a standard. Imidazole does not affect the assay.
Studying Mechanosensitive Ion Channels Using Liposomes 3.1.5. Day 5: Protein Collection and Determination of Protein Concentration
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1. For GST fusion protein, centrifuge the beads at 1,000 × g (with slow down ramp) for 5 min. Collect supernatant as purified protein. Store at 4°C. 2. Protein concentration can be determined by the Bradford assay at an absorbance of 595 nm, (typically measure 33.3 ml in 1 ml reagent) following the supplier’s instructions, referenced to a standard. 3. For GST fusion protein, (optional), if poor thrombin cutting is suspected, analyze uncleaved protein present by loading ~15 ml of leftover beads by polyacrylamide gel electrophoresis (PAGE), or by first eluting protein off 15 ml leftover beads with 15 ml of 20 mM glutathione. Collect supernatant for PAGE analysis.
3.2. MscS- His Tag Fusion Protein Purification
This procedure takes several days, thus it is advisable to start at the beginning of a week. Membrane processing is required to be the same day as cell disruption, thus day 2 is a longer than usual work day.
3.2.1. Day 1: Overnight Starter Culture
1. Using sterile technique, on ice, to 50 ml competent M15 cells (with pRARE and pREP4) add 1 ml expression construct, mix, and let stand for 20 min. Heat shock at 42°C for 45 s, and place on ice for 2 min. 2. Add the transformation mixture into a 100 ml culture flask with 12 ml of LB and 12 ml Ampicillin (100 mg/ml- to maintain pQE plasmid), 12 ml kanamycin sulphate (25 mg/ml- to maintain pREP4 plasmid), and 12 ml chloramphenicol (25 mg/ ml-to maintain pRARE plasmid). 3. Incubate at 37°C at 240 rpm (for ¾ inch orbit) in an orbital shaker incubator.
3.2.2. Day 2: Growth, Expression, Cell Disruption, and Membrane Processing
1. Add 10 ml of overnight culture into a ~2,800 ml culture flask containing 1000 ml LB, 1 ml 100 mg/ml ampicillin, 1 ml 25 mg/ ml kanamycin sulphate, and 1 ml 25 mg/ml chloramphenicol. 2. For aeration, seal the flasks with tissue paper (such as low lint fine task wipers) and incubate at 37°C at 150 rpm (for 2 inch orbit) in an orbital shaker incubator. 3. When optical density of the culture (OD) A600 = 1.0 to 1.2 (occurs in ~2–2.5 h), let the culture stand at room temperature without shaking for an hour. 4. Induce expression with the addition of 800 ml 1 M IPTG and 4 ml glycerol (0.4% final). Incubate within shaker as previously, but at 25 °C for 4 h. 5. Centrifuge culture at around 8,000 × g for 15 min at 4°C. Discard supernatant.
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6. For every 10 g of cell pellet resuspend (with a ~8 mm wide brush) in 10 ml PBS with 145 ml 20 mg/ml PMSF. Continue with cell disruption on the same day. 7. Add roughly a “match head” amount of DNase, mix. 8. French press (Thermo Electron Corporation) twice at 16,000 PSI. 9. Remove debris, unbroken cells, or inclusion bodies by high speed centrifugation at 8,000 × g (k-Factor ~3,600) for 22 min at 4°C. Collect supernatant for next step. 10. Pellet membranes by ultracentrifugation at 235,00 × g (k-Factor ~133) for 3 h at 4°C. Discard supernatant. 11. Suspend pellet in 5 ml Solubilization Buffer with a ~5 mm width brush. 12. Transfer suspended pellet to a 50 ml tube. Add 2.24 ml 200 mM DDM. Make up to final volume 50 ml using Solubilization Buffer (8 mM final DDM). 13. Solubilize on a roller wheel overnight at 4°C. 3.2.3. Day 3: Clarification and Column Binding, Protein Purification and Determination of Protein Concentration
1. Clarify the solubilization by ultracentrifugation at 100,000 × g for 20 min at 4°C. Recover supernatant to use. 2. Add 1.5 ml prepared cobalt-resin beads to the supernatant. Incubate on a rocker or roller for 3 h at 4°C. 3. Pour the cobalt resin suspension into a 20 ml chromatography column. Release bottom caps when resin has settled somewhat after several minutes, and allow all liquid to flow out and discard. 4. Wash the cobalt resin twice with 7.5 ml MscS Wash Buffer 1. 5. Wash the cobalt resin with 20 ml MscS Wash Buffer 2. 6. Elute with 4.5 ml MscS Elution Buffer into/or then transfer to a centrifugal filter device for concentration down to or just under 1 ml. Centrifuge to concentrate. Avoid over concentration that will result in precipitation. 7. If desired, imidazole can be removed, or reduced, by careful layering (to avoid mixing) with ~10 ml MscS Wash Buffer 1 and concentrate again as per step 6. Repeat the “reverse decanting” process with another 10 ml of MscS Wash Buffer 1. Protein is ready for analysis and/or for use. 8. Protein concentration can be determined by the Bradford assay measuring absorbance at 595 nm, (typically measure 33.3 ml in 1 ml reagent) following the supplier’s instructions, referenced to a standard. 9. When expressing from E. coli MS-channel knockout cells, ensure the protein concentration is lower than 500 µg/ml, to avoid precipitation. Expression in M15 (pQE32 or pQE70) or BL21-
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Rosetta cells (pET28) appears to allow further concentration of wild type MscS up to 10–15 mg/ml without precipitation. However, for some mutants, it is not recommeded to go above 4–5 mg/ml. Store at 4°C, do not freeze. 3.3. NMDA-6 × His-Tagged Protein Purification 3.3.1. Protein Expression in Sf9 cells
1. Grow Sf9 cells in Sf-900II media with 120 rpm shaking at 27°C. 2. For co-expression of NR1A and NR2A subunits, the cells are co-infected at the same MOI (multiplicity of infection) with both NR1A and NR2A recombinant baculoviruses at 3 × 106 cells/ml, using a MOI of 2–5 plaques forming unit per cell. The optimal time of harvest for the co-expression of NR1A and NR2A subunits is 72 h post infection. 3. Centrifuge the culture at 8,000 × g for 15 min at 4°C. Store the cell pellet at −80°C until required.
3.3.2. Protein Expression in CHO Cells
1. CHO cells are planted in 500 ml flasks at a density 1 × 106 cells/ml in CHO-S-SFMII (serum free, low protein media). 2. Cell culture is incubated at 37°C in a humidified atmosphere of 37°C, 7.5% CO2 in air. Samples are taken daily for determination of viable cell density using tryptan blue exclusion. 3. For transfection, cells are cultured to 80–90% confluence. 4. DNA constructs (1.6 mg/ml of transfection volume) and lipofectamine (1.6 ml) are diluted in 40 ml CHO-S-SFMII media and mixed gently. 5. After 5 min of incubation at room temperature the diluted DNA constructs and lipofectamine are combined and incubated for 20 min at room temperature to allow complex formation to occur. 6. The DNA-Lipofectomine complexes are slowly added to the flask containing CHO-S-SFMII media and gently mixed by swirling the flask. 7. Cells are incubated at 37°C, 7.5% CO2 on shaker at 150 rpm for 24 or 48 h.
3.3.3. Protein Purification
1. For cell lysis, the cell pellet from a 1 L expression is resuspended in 100 ml cell lysis and solubilization buffer without the detergent. 2. Cells are disrupted by sonication using a probe at 75 W for 15 s with 10–20 s cooling periods on ice (3×). 3. The OG concentration is adjusted to 3% after sonication and a sample is placed on rotating wheel for 20 min. 4. The lysed sample is clarified by centrifugation at 18,000 × g for 15 min at 4°C.
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5. The clarified lysate is applied to Talon resin at a ratio 10:1 (e.g., 12.5 ml of lysate is applied to 1.25 ml talon) and placed on a rotating wheel for 20–30 min. 6. Unbound proteins are removed by centrifugation at 700 × g for 2 min at 4°C. 7. After binding, each 1.25 ml talon is washed for 10 min using 50 ml wash buffer. 8. Elution of the protein is performed by transferring 2.5 ml talon to 10 ml disposable column (Biorad, Hercules, CA) and protein is eluted with 4 × 1 ml of Elution Buffer. 9. For gel filtration, 500 ml of Talon purified protein sample is loaded onto a Superdex 75 10/300 GL column (GE Healthcare, Rydalmere NSW) at 0.25 ml/min using gel filtration buffer, and fractions (25 × 0.5 ml) are collected for analysis by Western blot and reconstitution into liposomes. 3.4. SDS-PAGE
Typically, a 12 % SDS PAGE gel is used for gel analysis of both MscL and MscS. 1. Assemble the electrophoresis vertical mini electrophoresis rig system to manufacturer’s instructions and use precast gels as prescribed. If manual gels are to be cast, the following gel recipe is generally suitable for a 1 mm thick mini protein gel: 2. In a disposable tube, make 10 ml 12% Running Gel by mixing together (without undue aeration): 3.35 ml distilled water, 2.5 ml 1.5 M Tris-HCl pH 8.8, 100 ml 10% SDS, 4 ml 30% acrylamide/Bis solution, 50 ml 10% APS, mix, 5 ml TEMED, mix. 3. With a pipette fill the bottom ~3/4 of constructed glass plate structure with the freshly prepared Running Gel before it sets. Usually two gels are prepare with the 10 ml solution, as usually two gels are required for electrophoresis within the rig. 4. Without mixing, carefully drop about 500 ml of water to the top to act as an air barrier. Allow the gel to set, which occurs within about half an hour. 5. In a disposable tube, make 10 ml 12% Stacking Gel by mixing together (without undue aeration): 3 ml distilled water, 1.25 ml 0.5 M Tris-HCl pH 6.8, 50 ml 10% SDS, 650 ml 30% acrylamide/Bis solution, 25 ml 10% APS, mix, 5 ml TEMED mix. 6. Turn gel setting rig upside down to remove water, use lint free tissues to remove remaining water if required. Return rig to normal position. 7. With a pipette, add the Stacking Gel on top of the Running Gel, to the top of the constructed glass plate structure (see Note 2). Place the well-combs into the top to allow sample
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well moulds to set. Place combs without introducing air bubbles. 8. Once the Stacking Gel layer has set, place the gel(s) within the running rig (as per manufacturer’s instructions) and add 1 × Elpho Buffer (at the same time to stop the gel from drying) to the top/full level, so that buffer is above the well layer. The apparatus will have inner and outer buffer reservoirs – both should be full to the same level to halt buffer from possible leak below the gel top and to allow uniform dissipation of heat. Remove well-combs. 3.4.1. Preparation of Protein Samples for PAGE and Electrophoresis
1. To ~20 ml protein sample mix in ¼ volume 4 × Sample Buffer in a microcentrifuge tube. Cap. 2. Heat (denature) sample using a hot block at 95°C for 5 min. Denature protein molecular weight marker only if instructed by manufacturer. (Fermentas #SM0431 requires denaturing.). 3. Spin any condensation down, and load sample(s) and molecular weight marker into separate gel wells (see Note 3). 4. To manufacturer’s instructions, cover rig and run electrophoresis at 100 V (or as appropriate) until the bromophenol blue marker dye is near the end of the gel. 5. Remove gel from glass plate and incubate, covered, with rocking in Coomassie Stain for several hrs to overnight. 6. Remove Stain solution (can be reused), rinse briefly with Destain and incubate covered with rocking in Destain for several hrs to overnight or until sample bands are distinct, and background is down to a desired level. Refresh Destain if saturated with stain. Adding lint free tissues to the Destain solution will sequester excess brilliant blue diffusing from the gel.
3.4.2. Gel Analysis: MscL
Wild-type MscL will appear at a molecular weight position of around 15 kDa, or theoretically higher at 16 kDa depending on how many further extra amino acids are added due to the cloning construct used. For the Fermentas #SM0431 marker, MscL will migrate in between the two smallest marker bands at 18.4 and 14.4 kDa. For reasons not yet defined, MscL often appears as a doublet at its size position. His-tag reliant purifications may yield faint contamination of multiple bands, whilst GST-tag reliant purifications may yield specific contaminants potentially identified as: 37 kDa bovine thrombin 29 kDa of the GST (Glutathione S-transferase) cleaved protein 58.5 kDa of the GST dimer 44 kDa MscL-GST fusion protein
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3.4.3. Gel Analysis: MscS
The MscS subunit is about 31 kDa and will migrate at this size. Despite the denaturing conditions of SDS PAGE, an intense dimer band may exist. If denaturing is suboptimal, trimer and tetramer bands may also be detected. Of notable importance, degraded MscS will yield a band (a sub-band) just slightly smaller than the 31 kDa band, which may be reflected in slightly smaller multimers. Although the sub-band would normally not be present in fresh preparations, its presence, which may become the predominant product, will be reflective of compromised MscS function, and is a detrimental indicator. MscS, which can be stored for several months or to a year or so, should be analyzed on a gel periodically (or by other characterization methods) to detect the presence of the sub-band.
3.4.4. Gel Analysis: NMDA Receptor Channel Proteins
The metal affinity chromatography resulted in several low molecular weight mass bands visible on SDS/PAGE gel indicating nonspecific bindings and/or breakdown products of NR1a and NR2A subunits. Gel filtration chromatography purification is required to separate two main protein bands corresponding to 170 kDa for NR2A subunit and 100 kDa for the NR1a subunit from the contaminants.
3.5. Liposome Reconstitution
We have used several methods for reconstitution of MS channels into artificial liposomes (Fig. 1). Details are described below.
3.5.1. Dehydration/ Rehydration (D/R) Method
This method was the first developed for studies of MS channels by reconstitution into liposomes (17) and has since been the most frequently used method for characterization of biophysical and pharmacological properties of MS channels (7, 11, 13, 18) (Fig. 1a).
3.5.1.1. Preparation of Liposome Vesicles Using Bio-Beads SM2 (BioRad)
1. Rinse glass test-tube with chloroform, discard, and dry with nitrogen stream. 2. Dissolve 20 mg lipid into 1 ml chloroform in the glass tube. 3. Obtain an even lipid film on the surface of the tube by swirling the tube rapidly under a nitrogen stream until all the chloroform has evaporated. 4. Completely remove all chloroform by placing the tube under a high nitrogen stream for at least 15 min. 5. Resuspend lipid(s) in 2 ml of D/R Buffer and vortex for 60 s until cloudy. 6. Bath sonicate within a standard sonicator cleaner for 10–20 min. 7. Within a sealable tube, mix 200 ml (2 mg lipid) of liposome vesicles (from the sonication) with an amount of purified mechanosensitive protein to obtain the desired protein-tolipid wt/wt ratio (see Note 4). Fill to 2 ml with D/R Buffer.
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a D/R buffer
Purified MscL protein Rehydrated lipids in D/R buffer
Sonicate
3 hr rocking/Remove detergent (Biobeads), centrifuge
Dried lipid film
b
Dehydration/ Rehydration
Patch clamp pipette Blister
Pellet
MscL or MscS
0.4 M Sucrose
Oven incubation, 3 hrs at 45 ºC
Reconstituted proteoliposome
3 hr rocking (or overnight rocking)
Patch clamp pipette Lipid cloud
Dried lipid film
c
Reconstituted proteoliposome
O-ring
Purified MscL protein*
NMDAR protein*
Blister
Reconstituted proteoliposome
AC, 3 V, 5 Hz, 2 hours Patch clamp pipette
Electroformation Dried lipid film
GUVs
Blister
Fig. 1. Diagram showing different liposome reconstitution methods used for studies of MS channels (Subheading 3.5.3). (a) Dehydration/rehydration method. (b) Sucrose method. (c) Electric field method (Vesicle PrepPro, Nanion Technologies). (* Please note MscL is added before formation of GUVs, whereas NMDAR protein is added after GUV formation.)
8. Roller mix at room temperature for 1 h. 9. Add 320 mg of tissue blotted Bio-Beads per ml of protein solubilized in 1 mM DDM. (The amount of Bio-Beads used is determined by total detergent content.) The mixture is gently stirred for 3 h. 10. Allow beads to settle to the bottom of the tube, and use the supernatant which contains detergent-free proteoliposomes. 11. Micro ultracentrifuge the supernatant at 250,000 × g for 45 min at 4°C. 12. Remove supernatant completely. Resuspend pellet in 80 ml D/R Buffer by pipetting.
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13. Place 2–4 spots of 20 ml of proteoliposome suspension on a cleaned microscope slide. 14. Place slide(s) in vacuum desiccator for at least 6 h at 4°C. (Dehydration step.). 15. Add 20 ml D/R Buffer to each spot and leave overnight at 4°C within a Petri dish containing filter paper moistened with distilled water. (Rehydration step.) Sample is now ready to transfer into a patch recording chamber. 3.5.2. Sucrose Method
Our laboratory recently developed this reconstitution method (19), which has several advantages over the classical D/R method (Fig 1b). Requiring preparation times of 6 h or less, this method significantly saves time compared with the D/R procedure, which on average requires two days preparation time. It also represents the first highly reproducible method for incorporation of the MscS channel protein, which does not incorporate readily into azolectin liposomes or liposomes made of mixtures of pure lipids. Therefore, this new method has the potential to be used for studies of ion channels that may be difficult to study by liposome reconstitution using D/R method.
3.5.2.1. Day 1
1. Dissolve 10 mg of PC lipid (azolectin) in 1 ml of chloroform. 2. Take 200 mL aliquots and place into small test tubes (12 × 75 mm is ideal). 3. Remove solvent under a stream of N2 whilst swirling tube. 4. After continued drying of the lipid film under a stronger stream of N2 add 2 mL of H2O as a prehydration step (5 min)place drop of H2O directly onto the top of the lipid film; after prehydration the clear film develops opaqueness. 5. To the prehydrated lipids add 1 ml of 0.4 M sucrose and place in an oven or water bath at 45°C for 3 h. After this time the lipid will peel off the surface of the glass. 6. Add required protein amount to achieve a desired protein/lipid ratio. Important: add to bottom of tube, and do not shake. 7. Place on an orbital mixer at approximately 100 rpm and shake for 3 h. 8. Protein has now incorporated and can be used for experiments. However to ensure complete incorporation it is recommended to mix overnight.
3.5.2.2. Day 2
1. The lipids should have formed a cloud floating in the middle of the solution. Add approximately a cm³ of prepared BioBeads, and return to the mixer for 3 h. 2. Add 2 ml of the lipid cloud to recording bath, and they will form very large blisters.
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3.5.3. Electric Field Method (Vesicle PrepPro, Nanion Technologies)
20 ml of 5 mM lipid dissolved in chloroform was dried on the conductive side of an ITO-slide. A rubber O-ring acts as a spacer in between the two slides (Fig 1c).
3.5.3.1. MscL Protein Reconstitution
1. Once dried, add 250 ml of 210 mM D-sorbitol into the confines of the O-ring. 2. Add required protein/lipid ratio (typically 1:900 w/w) 3. The settings used are: Temperature = 23°C Frequency = 5 Hz Amplitude = 3 V 4. Allow to run for 2 h 5. After process is completed, extract GUVs with a 1 ml pipette.
3.6. Patch-Clamp Recording From MS Channels Reconstituted Into Liposomes
The results from studies of MS channels reconstituted into artificial liposomes have been described in numerous publications. Several reviews and original research papers (2, 3, 7, 11, 13, 17, 18, 20–22) highlight major findings of the structural and functional properties of MS channels using liposome reconstitution techniques.
3.6.1. Patch-Clamp Pipettes
1. Recording micropipettes are formed from borosilicate glass microcapillaries by using a pipette puller (e.g., P-87 Flaming/ Brown, Sutter Instrument Co., Novato, CA). Pulled recording pipettes should be ~1 mm in diameter corresponding to a pipette resistance in the range of 3.0–6.0 MW in Recording Solution. 2. To reduce electrical noise, pipette tips can be coated using Sylgard 184 (23) or transparent nail enamel (24) but is not required if single channel current is more than around 15 pA.
3.6.2. Recording
1. A 2–5 ml aliquot of liposomes made using any of the above methods is introduced into the recording chamber filled with the Recording Solution at 22°C. Recording pipettes are back filled with the same recording solution cleaned through a 0.22 mm filter. (Symmetrical solutions are preferentially used for determining single channel conductance of MS channels.). 2. Unilamellar blisters will be visible from several minutes up to 30 min (17) by which time the liposomes will have settled to the bottom of the chamber. The most common causes of “blistering” failure are incorrect pH or temperature of the Recording Solution. 3. Lower recording pipette to a blister and form a gigaohm seal (>1 GW) by applying suction to the patch pipette with a syringe. Suction is halted when a sudden decrease in pipette current occurs. MscL and MscS activities are usually recorded in excised liposomes patches (18, 19) which are obtained by next exposing the pipette tip to air for an instant only. (Fig. 2b.).
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Fig. 2. (a) Confocal imaging of liposomes containing AlexaFluor488-labeled MscS proteins (Subheading 3.7) reconstituted by the Sucrose method at 1:100 (w/w) ratio (Subheading 3.5.2). Images were recorded about 10 min after addition of liposomes to the imaging buffer (same as buffer used for patch-clamp recording) and were obtained by an Olympus FluoView1400 Laser Scanning Microscope (Olympus) using a Nikon 60× water-immersion objective, NA 1.00 (Nikon, Japan), with excitation at 488 nm by an Argon laser (Melles–Griot). (b) Patch-clamp recording of the AF488-MscS liposomes prepared by the same method showing that the channel activity (recorded at pipette voltage VP = +30 mV) is not compromised by labeling. Patch-clamping was performed as per protocols described in Subheading 3.6
NMDA receptor channel activity is recorded after inclusion of agonists in the pipette solution or adding agonists directly to the bath solution (11). 4. Ion currents arising from activation of MS channels using negative pipette pressure are recorded with a patch clamp amplifier (e.g., Axon 1D or Axopatch 200B, Axon Instruments). The suction is monitored with a piezoelectric pressure transducer (Omega Engineering, Stamford, USA). Currents are usually filtered at 2 kHz and digitized at 5 kHz for offline analysis. 5. Single channel recordings can be analyzed using software such as pCLAMP (Axon Instruments) or in-house applications. 6. For NMDA receptor channels the protein is mixed with GUVs at required ratio depending on protein concentration (usually 1:10 or 1:20). 3.7. Confocal Microscopy of Liposome Reconstituted MS Channels
Confocal imaging of fluorescent-tagged MS proteins reconstituted into liposomes is often useful to observe protein expression, clustering phenomena, or for analytical studies such as Föster Resonance Energy Transfer (FRET) studies. MS proteins can be fluorescently tagged either genetically or by generating a single cysteine mutant for site-specific attachment of thiol-reactive probes (Fig. 2a). While an MscL protein tagged genetically with Green Fluorescent Protein has been expressed and characterized (25), this approach is more problematic for MscS due to its large cytoplasmic domain. Here we describe a protocol for confocal imaging of liposome reconstituted MS proteins tagged with the fluorescent dye, Alexa Fluor 488 C5-maleimide (AF488).
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1. Generate expression constructs of single cysteine mutants using site directed mutagenesis. Suitable kit systems include QuickChange (Stratagene) or Phusion (Finnzymes). 2. Express and purify the six histidine-tagged single cysteine mutants using protocols similar to wild-type proteins described in Subheading 3.1, except that the Wash and Elution buffers are previously deoxygenated thoroughly (before addition of DDM to avoid frothing) and supplemented with 10 mM tris(2-carboxyethyl)phosphine (TCEP) to reduce any protein disulfide bonds that may have formed. A suitable method of deoxygenation employs percolating nitrogen gas for 15 min through a fish-tank aerator placed at the bottom a buffer. 3. Set up a labeling reaction consisting of 10 nmole of protein (usually ~300 mg), to a tenfold excess of AF488 (i.e., use a 100 mg prepared aliquot) made up to a final volume of 2 ml with Labeling Buffer (see Notes 5–7). Rock for 2 h at room temperature or overnight at 4°C. 4. Optional: stop reaction using 5 mM b-mercaptoethanol final concentration. 5. Transfer reaction to a dialysis cassette and place in 2 L chilled Dialysis Buffer with stirring at 4°C. Replace the buffer twice, after 2 and 4 h, and then dialyse overnight. 6. Collect protein from dialysis cassette and concentrate if necessary. 7. Reconstitute labeled proteins into azolectin lipids by either the D/R or Sucrose Method (Subheading 3.5) at desired concentrations. For imaging of reconstituted liposomes, protein to lipid ratios of 1:100 to 1:250 (w/w) usually suffices. However, for applications such as FRET, it is advisable to use ratios of 1:20 to 1:50 (w/w). 8. Add liposomes to the buffer used for patch-clamp recordings (Subheading 3.6) and wait about 10 min for blisters to form. 9. Image liposomes using a 60× lens and appropriate laser excitation (488 nm for AF488) and power settings (See Fig. 2a).
4. Notes 1. End of pipette may be cut off for a bigger hole to avoid clogging. Also do not place tip to very bottom of column. Allow room to enable beads to enter the tip. 2. Avoid introducing air bubbles because bubbles would cause mould disruption. 3. Long narrow gel load micropipette tips are available.
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4. Usually, a 1:4000 to 1:6000 protein-to-lipid ratio is used for the purpose of single channel recordings. To obtain a membrane fraction containing many mechanosensitive proteins a 1:100 to 1:200 protein-to-lipid ratio may be used. 5. Handling of Alexa Fluor dye should be done in subdued light. 6. In case the protein is not labeled immediately after purification, it should be pre-treated with a tenfold excess of TCEP for 1 h at room temperature. 7. The pH of the labeling reaction solution should be between 7.2 and 7.5; therefore, it is advisable to ensure that the added protein (previously eluted at pH 6.0) be at an adequate concentration.
Acknowledgments We wish to thank Dr Stephen Hughes for his contribution to studies of magnetic field effects on the MscL channels using liposome reconstitution technique. This research has been supported by grants of the Australian Research Council and the National Health and Medical Research Council of Australia to B. Martinac and A. Kloda. References 1. Kung CA (2005) A possible unifying principle for mechanosensation. Nature 436:647–54 2. Martinac B (2005) Force from lipids: physical principles of gating mechanosensitive channels by mechanical force revealed by chemical manipulation of cellular membranes. The Chemical Educator 10(2):107–14 3. Hamill OP, Martinac B (2001) Molecular basis of mechanotransduction in living cells. Physiol Rev 81:685–740 4. Martinac B (2001) Mechanosensitive channels in prokaryotes. Cell Physiol Biochem 11:61–76 5. Markin VS, Martinac B (1991) Mechanosensitive ion channels as reporters of bilayer expansion. A theoretical model. Biophys J 60:1120–7 6. Martinac B, Adler J, Kung C (1990) Mechanosensitive ion channels of E. coli activated by amphipaths. Nature 348:261–3 7. Maroto R, Raso A, Wood TG, Kurosky A, Martinac B, Hamill OP (2005) TRPC1 forms the stretch-activated cation channel in vertebrate cells. Nat Cell Biol 7:179–85 8. Maingret F, Patel AJ, Lesage F, Lazdunski M, Honoré E (2000) Lysophospholipids open the two-pore domain mechano-gated K+
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channels TREK-1 and TRAAK. J Biol Chem 275:10128–33 Zhang Y, Gao F, Popov VL, Wen JW, Hamill OP (2000) Mechanically gated channel activity in cytoskeleton-deficient plasma membrane blebs and vesicles from Xenopus oocytes. J Physiol 523(Pt 1):117–30 Zhou XL, Batiza AF, Loukin SH, Palmer CP, Kung C, Saimi Y (2003) The transient receptor potential channel on the yeast vacuole is mechanosensitive. Proc Natl Acad Sci USA 100:7105–10 Kloda A, Lua L, Hall R, Adams DJ, Martinac B (2007) Liposome reconstitution and modulation of recombinant N-methyl-D-aspartate receptor channels by membrane stretch. Proc Natl Acad Sci USA 104:1540–5 Perozo E, Cortes DM, Sompornpisut P, Kloda A, Martinac B (2002) Open channel structure of MscL and the gating mechanism of mechanosensitive channels. Nature 418:942–8 Perozo E, Kloda A, Cortes DM, Martinac B (2002) Physical principles underlying the transduction of bilayer deformation forces during mechanosensitive channel gating. Nat Struct Biol 9:696–703
Studying Mechanosensitive Ion Channels Using Liposomes 14. Corry B, Rigby P, Liu ZW, Martinac B (2005) Conformational changes involved in MscL channel gating measured using FRET spectroscopy. Biophys J 89:L49–51 15. Hughes S, El Haj AJ, Dobson J, Martinac B (2005) The influence of static magnetic fields on mechanosensitive ion channel activity in artificial liposomes. Eur Biophys J 34:461–8 16. Petrov E, Martinac B (2007) Modulation of channel activity and gadolinium block of MscL by static magnetic fields. Eur Biophys J 36:95–105 17. Delcour AH, Martinac B, Adler J, Kung C (1989) Modified reconstitution method used in patch-clamp studies of Escherichia coli ion channels. Biophys J 56:631–6 18. Häse CC, Le Dain AC, Martinac B (1995) Purification and functional reconstitution of the recombinant large mechanosensitive ion channel (MscL) of Escherichia coli. J Biol Chem 270:18329–34 19. Battle AR, Petrov E, Pal P, Martinac B (2009) Rapid and improved reconstitution of bac terial mechanosensitive ion channel pro teins MscS and MscL into liposomes using
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a modified sucrose method. FEBS Letters 583: 407–412 20. Vasquez V, Cortes DM, Furukawa H, Perozo E (2007) An optimized purification and reconstitution method for the MscS channel: strategies for spectroscopical analysis. Biochemistry 46:6766–73 21. Sukharev S (2002) Purification of the small mechanosensitive channel of Escherichia coli (MscS): the subunit structure, conduction, and gating characteristics in liposomes. Biophys J 83:290–8 22. Martinac B, Kloda A (2003) Evolutionary origins of mechanosensitive ion channels. Prog Biophys Mol Biol 82:11–24 23. Sakmann B, Neher E (1995) Single-channel recording. Plenum Press, New York and London 24. Martinac B, Buechner M, Delcour AH, Adler J, Kung C (1987) Pressure-sensitive ion channel in Escherichia coli. Proc Natl Acad Sci USA 84:2297–301 25. Norman C, Liu ZW, Rigby P, Raso A, Petrov Y, Martinac B (2005) Visualisation of the mechanosensitive channel of large conductance in bacteria using confocal microscopy. Eur Biophys J 34:396–402
Chapter 5 Studying Amino Acid Transport Using Liposomes Cesare Indiveri Abstract The transport of amino acid across the membranes has great importance in cell metabolism. Specific experimental methodologies are required for measuring the vectorial reactions catalyzed by the membrane transporters. So far, the most widely used technique to study amino acid transport was the measure of amino acid flux in intact cell systems expressing a specific transporter. Some limitations in this procedure are caused by the presence of endogenous transporters and intracellular enzymes and by the inaccessibility of the intracellular compartment. Alternative experimental strategies which allow to reducing the interferences and improving the handling of the internal compartment would be useful to the amino acid transport knowledge. An experimental protocol, which makes use of liposomes to study the transport of amino acid mediated by the glutamine/amino acid (ASCT2) transporter, solubilized from rat kidney brush borders, is described. The procedure is based on the reconstitution of the transporter in liposomes by removal of detergent from mixed micelles of detergent, solubilized protein, and phospholipid. The transport is assayed in the formed proteoliposomes measuring the Na+ dependent uptake of l-[3H]glutamine in antiport with internal l-glutamine. This method allows measuring the transport activity under well controlled experimental conditions and permits performing experiments which cannot be realized in intact cell systems. Key words: Liposomes, Reconstitution, Transport, Membrane, Amino acids, Glutamine
1. Introduction The interest in the study of amino acids transport across the cell membrane increased exponentially in the last decade. This was also due to the relevance of this research field in human physiology and pathology. In mammalian cells the transport of amino acids is carried out by a large number of proteins which are different in primary structure and function. Most of these transport systems have been functionally characterized using intact eukaryotic cell systems in which specific transporters were expressed. Other V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_5, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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transporters, which have been predicted on the basis of gene sequences, are still orphan with respect to the function. The knowledge of the state of the art in this field is clearly and systematically described in a number of reviews (1–8). Even though intact cell experimental models allowed classification of a large number of amino acid transporters, technical limitations of the models made several aspects of the structure, function, and regulation of the transporters still unclear or unknown. Thus, the development of alternative experimental strategies would give benefits to the knowledge of amino acid transport. A suitable model for transport studies is the liposome system, which has often been helpful in defining and clarifying functional properties of membrane transporters of different types. Thanks to this model it is possible to perform experiments which are forbidden in intact cells. An important advantage of reconstituting membrane transporters in liposomes with respect to the study in cell systems is the control of experimental conditions in the internal compartment, which help to determine internal parameters like the Km for substrates or the influence of internal effectors on the transport. In this respect, a reconstitution method which allows the insertion of the protein in the liposomal membrane in the same orientation of the cell membrane is particularly reliable since it mimics the physiological situation, i.e., the correspondence of the intraliposomal and extraliposomal with the intracellular and extracellular sides. Other advantages are the reduction of interferences due to the absence of enzymes which could modify the substrates and the absence of different transporters in the same vesicles that is guaranteed by the higher phospholipids/protein ratio (one transport protein per liposome) with respect to the cell membrane (several transport proteins per cell); this feature has also the effect of prolonging the time course of the substrate uptake, leading to better resolution of the initial transport rate and the kinetics. The liposome model provides the possibility of modifying the lipid composition of the membrane to study the influence of specific lipids on the transport. To get further insights into the transport of amino acids, we have pointed out a procedure of reconstitution of the glutamine/amino acid transporter ASCT2 extracted from apical plasma membranes (brush-borders) of rat kidney where the transport of glutamine is very active being involved in essential physiological functions. The previous studies in intact cells expressing the ASCT2 protein (9–14) showed that it cata lyzes antiport of glutamine with neutral amino acids which is dependent on extra cellular Na+ in kidney, intestine, lung, muscle,
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testis, as well as in nervous tissue. The external Km’s for glutamine, other amino acids, and Na+ were determined and it was proposed that, differently from other neutral amino acid transporters, ASCT2 could transport also the anionic amino acid glutamate. The reconstitution of the glutamine/amino acid transporter in liposomes, besides confirming the previous data, permitted a deeper understanding of the ASCT2 transporter (15, 16). The reconstituted transporter, which is inserted in the liposomal membrane with the same orientation of the cell membrane, is inhibited by externally added Cys and Lys specific reagents. The internal Km’s for the amino acids were determined; their values are 20 times higher than the external ones. The mechanism of the complex transport reaction was found to be random simultaneous. The transporter is regulated by internal (intracellular) ATP. It also catalyzes a glutamate/glutamine antiport mode in which, glutamate, not the zwitterion glutamic acid, is transported with lower Km at acidic pH. The protocol for performing the reconstitution and the transport assay of the glutamine/amino acid transporter ASCT2 from rat kidney is described here. This protocol requires, as starting materials, brush border membranes from rat kidney which are prepared according to a previously described method (17). The transporter is solubilized from the brush borders with the non ionic detergent C12E8 and reconstituted in liposomes. The reconstitution procedure derives from a method which was originally pointed out for mitochondrial transport proteins (18, 19). It is based on the slow removal of detergent from mixed micelles of detergent phospholipid and protein by repeated passages through chromatography columns filled with hydrophobic resin. This leads to the formation of unilamellar phospholipid vesicles with the protein inserted in the membrane. The method has been adapted for the reconstitution of plasma membrane transporters. This goal has been achieved by changing the type of hydrophobic resin used and the critical parameters of the reconstitution like the protein concentration, the detergent/phospholipid ratio,and the number of repeated passages through the chromatography columns (15, 20). The transport is finally assayed by measuring the uptake of radioactive l-glutamine externally added to the proteoliposomes. The transport is active only in the presence of external Na+ and internal unlabeled l-glutamine or other neutral amino acids, since ASCT2 catalyzes an obligatory antiport of neutral amino acids which is dependent on extraliposomal Na+. Using this protocol, several types of experiments can be performed, by changing or adding some components or modifying some steps of the procedure.
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2. Materials All the solutions and suspensions must be prepared in bi-distilled water. 2.1. Preparation of Brush Border Membranes
1. Buffer 1: 0.3 M sucrose, 5 mM EGTA 12 mM Tris-HCl pH 7.4. 2. Buffer 2: 0.15 M sucrose, 2.5 mM EGTA 6 mM Tris-HCl pH 7.4. 3. 50 mM MgCl2.
2.2. Solubilization of the Transporter
1. Brush borders. Aliquots of 50 µl brush borders prepared as described in Subheading 3.1 (containing from 150 to 300 µg protein depending on the preparations) stored at −20°C. 2. Solubilization buffer: 1.95 % C12E8 (octaethylene glycol monododecyl ether from Fluka). The solution can be stored at 0–4°C for one week.
2.3. Reconstitution of the Brush Border Extract
1. Stock buffer pH 7.0: 500 mM Hepes-Tris pH 7.0. Prepare the buffer dissolving Hepes in a volume of water about 70% of that final; then add Tris powder up to pH 7.0 and water up to the final volume. Use this solution to prepare all the diluted solutions. The stock buffer pH 7.0 can be stored at 0–4°C for 4–6 weeks. 2. 100 mM l-glutamine: Prepare fresh and use this solution to prepare all the diluted solutions. 3. Amberlite resin. Swell Amberlite XAD-4 resin (20-50 mesh from Fluka) adding 2 volumes of methanol to about 50 g resin. Mix gently using a glass stick; then leave for about 30 min at room temperature. Remove the supernatant methanol and repeat the procedure from 5 to 8 times until the supernatant methanol remains limpid. Then wash the resin with excess water 5 times (leave the resin in water 15 min among the washings). Store the resin at 0–4°C in 2 volumes of water. The resin can be stored for 4 weeks without adding conservatives. 4. Amberlite columns. Prepare 4 columns of about 0.5 cm internal diameter. Pasteur pipettes whose lower outlets are closed by cotton can be used (see Note 1). Fill the columns with the Amberlite resin up to 2.5 cm height. The Amberlite columns cannot be reused. 5. Amberlite equilibrating buffer. Freshly prepare 15 ml of a solution of 30 mM glutamine, 20 mM Hepes/Tris at pH 7.0 using the 100 mM l-glutamine, and the stock buffer pH 7.0. The concentration of glutamine may vary depending on the experiments (see Subheading 3.4, item 12 and related Notes).
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6. Solution of 10% C12E8 (see also Subheading 2.2, item 2). The solution can bnume stored at 0–4°C for 4 weeks. 7. Preformed liposomes. Prepare the liposomes dissolving 1 g phospholipid (egg yolk phospholipids from Fluka) in a final volume of 10 ml water. Vortex to homogeneity, i.e., until no solid particles are present. Then sonicate 2 ml of the suspension in a glass tube for 2 min in pulse, i.e., 1 s sonication, 1 s intermission using a sonicator (at least 100 W of maximum power equipped with a 3 mm diameter micro tip) set at 40 W output power. The tube must be maintained at low temperature in an ice/water bath during sonication (see Note 2). The preformed liposomes can be stored at 0–4°C for 3–4 days. 2.4. Transport Assay
1. Sephadex resin. Swell about 2 g of Sephadex G-75 (40-120 µm dry bead diameter from Sigma-Aldrich) in excess water overnight. Then, eliminate excess air by vacuum for 15 min. 2. Columns A: Fill 4 columns of 0.7 cm internal diameter up to 15 cm height (Econo-Columns from Bio-Rad are suitable for this purpose) with Sephadex resin. The columns can be reused (see Notes 3 and 4). Store the columns at 0–4°C. 3. Buffer A: 30 mM sucrose and 20 mM Hepes-Tris pH 7.0 (use the stock buffer pH 7.0 solution). Prepare fresh. 4. Columns B: Fill 16 columns of 0.6 cm internal diameter up to 8 cm height (Glass Columns-Reusable from Pierce are suitable for this purpose) with Sephadex resin. The columns can be reused (see Notes 3 and 4). Store the columns at 0–4°C. 5. Buffer B: 50 mM NaCl. The solution can be stored for 1–2 weeks. 6. 100 mM l-glutamine (see Subheading 2.3, item 2). 7. 2 M NaCl can be stored at room temperature for several weeks. 8. Stock buffer pH 7.0 (see Subheading 2.3, item 1) 9. Labeled glutamine. Prepare freshly the solution with 2 µl of 100 mM l-glutamine, 50 µl of 2 M NaCl, 4 µl of stock buffer pH 7.0, 2 µl radioactive l-[3H]glutamine (l-[G-3H]glutamine from GE Healthcare, 1 mCi/ml), and 42 µl water (see Note 5). The concentrations of labeled glutamine may vary depending on the experiment (see Subheading 3.4, item 12 and related Notes). 10. Inhibitor. Freshly prepared solution of 0.8 mM mersalyl (mersalyl acid from Sigma-Aldrich) in 20 mM Hepes/Tris pH 7.0 (use the stock buffer pH 7.0).
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3. Methods The method for the study of the transport mediated by the glutamine/amino acid transporter ASCT2 in proteoliposomes consists of three main steps: the solubilization of brush borders, the reconstitution, and the transport assay. The brush borders are prepared using a procedure previously described by other authors (17) with few modifications: details to carry out this procedure are described in Subheading 3.1. The brush border extract is inherently labile. To obtain reliable results it is important to prepare all the materials needed for the whole procedure before thawing the brush borders for the solubilization step. Initial reconstitution mixtures are then prepared mixing the brush border extract with phospholipids and further detergent to obtain mixed micelles. The mixtures are repeatedly chromatographed on Amberlite XAD-4 columns. After repeated passages through the resin, the detergent is removed; thus, phosholipids and proteins form the proteoliposomes. These vesicles contain, in the internal and external compartments, the solutes added in the initial reconstitution mixture and in the Amberlite equilibrating buffer, i.e., the substrate glutamine and the buffer. The reconstituted proteoliposomes are stable up to 4 h at 0–4°C. The external substrate is removed by size exclusion chromatography on Sephadex G-75, before starting the transport assay. This procedure permits the starting of the transport with a concentration of external labeled glutamine (0.1 mM) lower than the concentration of the internal unlabeled glutamine (30 mM). In the case of antiport systems like the glutamine/amino acid transporter ASCT2, the low external/internal concentration ratio leads to the accumulation of the radioactive substrate (added outside) in the internal compartment of the proteoliposomes. 50 mM Na+ (as NaCl) is added together with 0.1 mM l-[3H]glutamine. The transport reaction is stopped by the inhibitor mersalyl (20 µM) added at different times. In control samples the inhibitor is added together with the l-[3H]glutamine and Na+. The l-[3H]glutamine taken up by these controls represents the unspecific permeability of the proteoliposomes that will be subtracted from the l-[3H] glutamine taken up by the samples. After stopping the transport at various times, the external radioactivity is removed by Sephadex G-75 chromatography and the radioactivity entrapped inside the vesicles is counted. The transport activity is calculated from the entrapped radioactivity data. The method described allows obtaining reliable time courses of the transport process which represent the basis for every type of experiment (see Subheading 3.4, step 12) - to study the kinetics or the dependence of the transport on different types of inhibitors and effectors (specific reagents, nucleotides, ions, phospholipids).
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3.1. Preparation of the Brush-Border Membranes
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1. Cut thin slices (1–2 mm thickness) from kidney cortex (16–20 kidneys from male wistar rats of about 200 g) upto collect about 10 g of material. Put the slices in cold buffer 1 (previously kept at 0°C for 1 h) and wash 3 times to eliminate the blood. 2. Add 60 ml of cold buffer 1 and homogenize in a blender (good quality kitchen blender of about 500 ml) at maximal speed for 3 min. 3. Add cold MgCl2 solution up to reach a final concentration of about 12 mM (see Note 6), mix and keep 15 min at 0°C. 4. Centrifuge the homogenate in 50 ml tubes at 2,600 g in a preparative centrifuge for 15 min at 0°C. Collect together the supernatants. 5. Centrifuge the supernatant in 50 ml tubes at 18,500 g in a preparative centrifuge for 30 min at 0°C. Collect the sediments and resuspend together with 30 ml of cold buffer 2. 6. Homogenize with a 30 ml potter and then add cold MgCl2 solution up to reach a final concentration of about 12 mM (see Note 6), mix and keep 15 min at 0°C. 7. Centrifuge the suspension in 50 ml tubes at 2,600 g in a preparative centrifuge for 15 min at 0°C. Collect together the supernatants. 8. Centrifuge the supernatant in 50 ml tubes at 18,500 g in a preparative centrifuge for 30 min at 0°C. Collect the sediments and resuspend together in 30 ml of cold buffer 2. 9. Homogenize with a 30 ml potter and centrifuge in 50 ml tubes at 18,500 g in a preparative centrifuge for 30 min at 0°C. Collect the sediments and resuspend together in 2 ml of cold buffer 2. 10. Store the preparation at −20°C, in aliquots of 50 µl (150– 300 µg protein) in disposable tubes of 1 or 1.5 ml. The preparation can be stored without loss of activity up to 2 months.
3.2. Solubilization of the Brush-Border Membranes
1. Thaw 50 µl of brush border in ice/water bath. When the membranes are liquid, vortex the tube and add 100 µl of the solubilization buffer. Vortex for 30 s and keep at 0°C for 2 min. 2. Centrifuge the solubilized brush border at about 13,000 g for 4 min at 0°C. Collect the supernatant (brush border extract) in a disposable tube and keep for not more than 10 min, at 0°C until the preparation of the initial reconstitution mixture (Subheading 3.3, item 2). 3. After use (see Subheading 3.3, item 2) store the remaining extract at 0°C for the determination of the protein concentration. 4. The protein concentration in the extract is determined by a standard procedure like the BioRad or the Lowry method (stock solutions from Bio-Rad). This can be done after the transport assay.
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Fig. 1. Columns holders. The photo shows examples of Plexiglas home- made columns holders for Amberlite (left), A (middle) and B (right) columns. Holes of appropriate sizes permit the insertion of the columns and the tubes. In the case of the holder for columns B, a movable support for vials is inserted below the columns. This support can be removed and replaced by a container that will be marked “radioactive” (see Subheading 3.4, step 10). Movable containers for equilibrating or washing the columns are present in the holders for Amberlite and A columns. Note the black sheet beyond the column tips in the left and middle holders to facilitate the view of the turbid drops
3.3. Reconstitution of the Brush Border Extract
1. Equilibrate 4 Amberlite columns applying 3 ml of the Amberlite equilibrating buffer. Columns holders are useful in this and the following sections (Fig. 1 and see Note 4). 2. Prepare 4 initial reconstitution mixtures (see Note 7) in 1.5 ml disposable tube adding the different solutions in the following order: 25 µl of brush border extract, 75 µl of the 10% C12E8, 100 µl of preformed liposomes, 210 µl of 100 mM l-glutamine (final concentration 30 mM), 28 µl of the stock buffer pH 7.0 (final concentration 20 mM), 262 µl water (to reach a final volume of 700 µl); vortex for 30 s. These operations must be performed at 0–4°C, for example in ice/water bath. 3. Apply each reconstitution mixture onto an equilibrated Amberlite column. Discard the first 5 drops and then collect the eluate. 4. Apply the eluate onto the same Amberlite column. Collect the eluate without discarding drops and repeat this passage further 14 times on the same column (16 total passages). These operations must be performed at room temperature. After these passages, active proteoliposomes are formed which are stable at 0°C up to 4 h.
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1. Prepare a sample tube with 44 µl of labeled glutamine and a control tube with 44 µl of labeled glutamine and 22 µl of inhibitor. 2. Prepare 8 inhibition tubes with 2.5 µl of inhibitor. 3. Load 550 µl of each of the 4 proteoliposome samples on each of the 4 columns A and wait for the absorption of the samples into the resin. Gently, but quickly, apply small volumes (200– 400 µl) of the Buffer A on the columns, till the proteoliposomes are in the middle of the column. Then add 10 ml of buffer A. Collect 600 µl of proteoliposomes into disposable 1.5 ml tubes starting from the second turbid droplet (see Note 8 and Fig. 1). 4. Collect 500 µl of proteoliposomes eluted from each of the 4 columns A and mix together in a 2 ml disposable tube. 5. Transfer 814 µl proteoliposomes into both the sample and control tubes and mix quickly with the pipette tip. This represents the start of the transport reaction for both the samples (S) and the controls (C). A delay of 15–30 s can be practised between the two transfers (see also Note 9). 6. After 5, 10, 20, 30, 40, 60, 80, and 100 min, rapidly transfer (in such a way as to mix the proteoliposomes with the inhibitor) 100 µl from the (S) to an inhibition tube obtaining the inhibited sample (IS). At each time quickly apply 100 µl of the (IS) and 100 µl of the (C) on two columns B (see Note 9). Wait for adsorption of the suspension into the resin and elute adding 100 µl, 200 µl, 200 µl, 400 µl, 500 µl, 500 µl of 50 mM NaCl, waiting for the complete adsorption of each aliquot of the solution before adding the following. Discard the first 900 µl eluate and collect the following 1,000 µl into vials of the appropriate size to be analyzed in a b-counter. Also in this step a column holder will be very useful (Fig. 1). 7. In a separate vial (total radioactivity) add 5 µl of the labeled glutamine for the determination of the total radioactivity added to each sample. 8. After addition of at least 4 ml of scintillation cocktail (PicoFluor 40 from PerkinElmer) vortex each vial and count the radioactivity. 9. Regenerate the columns A with 10 ml buffer A and then 50 ml water. Equilibrate the columns applying 20 ml 0.1 % NaN3 (see Note 3). 10. Regenerate the columns B, with 10 ml buffer B and then 50 ml water. Equilibrate the columns applying 10 ml 0.1 % NaN3 (see Note 3). Very important: the solution eluted from the columns B contains radioactivity (3H); it must be collected
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in special containers and treated according to the local regulation concerning the use of radioisotopes (see also Note 5). 11. Calculate the specific transport using the following equation: transportt (nmol/mg protein) = (cpmsample– cpmcontrol)/(SR × mg protein) (see Note 10) where transportt is the specific transport at time t, cpmsample and cpmcontrol are the cpm measured in the (IS) and in the respective (C), SR is the specific radioactivity (cpm/nmol) calculated as: total radioactivity/nmoles of glutamine per sample (see Note 11), and mg protein is the amount of proteins per sample derived from the concentration of the protein added to the initial reconstitution mixture (see Note 12). 12. Experimental data expressed as nmol/mg protein obtained above are interpolated by non linear regression analysis using the first order rate equation: y = A(1 – e– kt) where y are the specific transport values at the different times, A are the nmoles of glutamine taken up at infinite time (extrapolated from the fitting), k is the first order rate constant of the uptake process. The initial rate of the transport process is calculated as k · A (nmol/min/mg protein).
Fig. 2. Time course of l-[3H]glutamine uptake in the proteoliposomes. 0.1 mM l-[3H] glutamine is added at time zero to proteoliposomes containing 30 mM internal glutamine in the presence of 50 mM external NaCl. The transport reaction is stopped at the indicated times using 20 µM mersalyl. In the control samples the inhibitor is added at time zero (see Subheading 3.4). The curve is derived from the interpolation of the data points in a first order rate equation (see Subheadings 3.4, item 11 and 3.4, item 12)
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A non linear regression analysis software should be used (Grafit from Sigma-Aldrich is suitable). From the interpolation of the experimental data in the described equation a time course is obtained like the example reported in Fig. 2. The time course experiment described can be used, with some modifications, to study the effect of inhibitors or activators (see Note 13) and the transport kinetics (see Note 14).
4. Notes 1. Cut the Pasteur pipettes leaving a short tip of about 2 cm. Put the cotton in the upper part of the tip. The elution rate of the column depends mainly on the compression of the cotton. Use medium compression to obtain elution rate of 30–50 µl/s (about 1 drop/s). 2. The glass tube should have a rounded bottom. It is very important to use a Pb free glass tube. If you are not sure about the feature of the tube, then use a plastic 2 ml disposable tube; this may lead to some variability in the liposome preparations. Ensure that the temperature of the suspension in the tube does not exceed 10°C during sonication. 3. The filled columns A and B can be reused, normally for 10 to 20 times. In any case, the columns must be refilled with new swollen resin when the elution rate has evidently decreased. Before refilling with the new resin, the glass columns must be cleaned with common detergents used for dishes and then washed with bidistilled water to completely remove any trace of detergent. 4. The number of Amberlite columns, columns A and columns B depends on the type of the experiment. In the protocol for the time course described here the requirement is 4 Amberlite columns and columns A; 16 columns B. The operator has to consider that: (1) a column A is needed for each Amberlite column; (2) the proteoliposomes eluted from column A are normally sufficient to perform 4–5 transport assay samples; (3) a column B is necessary for each transport assay sample. Column holders are of great help in the experimental procedures requiring the simultaneous elution of several columns (see Fig. 1). 5. The 3H isotope is not particularly hazardous; follow the local regulation concerning the use of radioisotopes. For safety measures, it is sufficient to use disposable gloves and take care not to directly touch the solutions containing radioactive glutamine. To this aim, centrifuge by short spin the tubes which contain radioactive solutions in order to avoid some radioactive drops that may stick to the tube edge or closure.
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6. The volume of the MgCl2 solution (50 mM) required to be added must be calculated on the basis of the volume of the suspended sediment, which may vary in different preparations. The final concentration of MgCl2 (12 mM) is not very critical. 7. In this protocol four identical initial reconstitution mixtures are prepared. However, the number or the composition of initial reconstitution mixtures may change depending on the type of experiment. Two general types can be distinguished: (1) experiments in which compounds or concentrations in the external liposomal space are changing. In this case identical initial reconstitution mixtures are prepared; (2) experiments in which compounds or concentrations in the intraliposomal space are changing. In this case different reconstitution mixtures are required. The Amberlite columns must be equilibrated with different solutions containing the same compounds and buffer of the corresponding reconstitution mixture. 8. To clearly distinguish the turbid droplets place a sheet of black plastic or other material beyond the outlet of the columns (see Fig. 1). 9. To be faster use two different pipettes, one for the samples (Subheading 3.4, item 5) or inhibited samples (Subheading 3.4, item 6),and the other for the controls. A constant delay of 15–30 s can be maintained, after the Subheading 3.4, item 5 procedure (start of the transport reaction), between the two different (S) and (C) samples. The delay among different samples will be very useful in the case of more complex experiments requiring large number of samples (see Notes 13 and 14). 10. Depending on the type and settings, ß-counters will give the radioactivity measurements as cpm or dpm units. Both units can be used equally if the radioactivity of the samples and the specific radioactivity are expressed in the same units. In a well performed experiment, cpmcontrol must be lower than 10% of the respective cpmsample. 11. Calculate the nmoles from the concentration of externally added l-[3H]glutamine in the100 µl sample (each sample in the present experiment contains 10 nmoles of glutamine). 12. The protein content in each sample of 100 µl should correspond to 1/7 of the amount of protein in an initial reconstitution mixture (measured in the extract). However, a dilution of the samples occurs during the chromatography on column A. We have empirically calculated that the amount of protein in the 100 µl sample actually corresponds to 1/12 of that in the initial reconstitution mixture. It has to be stressed that the protein cannot be directly assayed in the proteoliposome samples since lipids interfere with the assay methods. 13. To study the influence of effectors on the transporter on the external side, these molecules must be added to the
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proteoliposomes before the transport assay together with the labeled glutamine in the sample and in the control tubes. To study the influence of effectors on the internal side, the molecules must be included in the initial reconstitution mixture and in the Amerlite equilibrating buffer at the same concentration. In this case, several different initial reconstitution mixtures are needed, each with a different compound or different internal concentration of the effector (see also Note 7). The concentration of each effector should be chosen according to previous data (15, 16). In the case of investigation with novel molecules, a preliminary experiment should be performed in advance with few concentrations chosen in a wide range (for example 0.1, 1, 10, 100 µM). 14. To obtain kinetic constants like Km or Vmax several time courses must be performed at different substrate concentrations and the initial transport rate calculated. In experiments of this type which require several data points, the time course can be reduced to 4 points (5, 10, 30 and 60 min). For reliable constant determinations at least five different glutamine concentrations within an appropriate range must be used and each experiment should be repeated at least three times. In particular, for the determination of external Km, the l-glutamine in the labeled glutamine solution must be varied from 0.05 to 5 mM (final concentration in the proteoliposome samples; the other components do not vary) at fixed internal glutamine concentration of 30 mM. Since the transporter catalyzes an obligatory antiport of glutamine and neutral amino acids, the internal Km can be investigated by following the time course of the uptake of fixed 1 mM l-[3H]glutamine (see above for the labeled glutamine solution) in the presence of various intraliposomal glutamine or other amino acids which are accepted by the ASCT2 transporter (15, 16). The intraliposomal concentrations of the amino acid must be varied from 1 to 30 mM; the concentration of internal glutamine or other amino acids are given by the concentration in the initial reconstitution mixture and in the Amberlite equilibrating buffer (see also Note 7).
Acknowledgments This work was supported by the PRIN (Progetti di Ricerca Scientifica di Rilevante Interesse Nazionale) 2006 grant n. 2006054479 from MiUR (Ministero dell’Università e della Ricerca). The author is indebted to Dr. Francesca Oppedisano, Dr. Lorena Pochini, and Dr. Michele Galluccio for help in preparing the manuscript.
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References 1. Palacin M, Estevez R, Bertran J, Zorzano A (1998) Molecular biology of mammalian plasma membrane amino acid transporters. Physiol Rev 78:969–1054 2. Bode BP (2001) Recent molecular advances in mammalian glutamine transport. J Nutr 131:2475S–2485S 3. Broer S (2002) Adaptation of plasma membrane amino acid transport mechanisms to physiological demands. Pflugers Arch 444: 457–466 4. Mackenzie B, Erickson JD (2004) Sodiumcoupled neutral amino acid (System N/A) transporters of the SLC38 gene family. Pflugers Arch 447:784–795 5. Kanai Y, Hediger MA (2004) The glutamate/ neutral amino acid transporter family SLC1: molecular, physiological and pharmacological aspects. Pflugers Arch 447:469–479 6. Verrey F, Ristic Z, Romeo E, Ramadan T, Makrides V, Dave MH, Wagner CA, Camargo SM (2005) Novel renal amino acid transporters. Annu Rev Physiol 67:557–572 7. McGivan JD, Bungard CI (2007) The transport of glutamine into mammalian cells. Front Biosci 12:874–882 8. Broer S (2008) Amino acid transport across mammalian intestinal and renal epithelia. Physiol Rev 88:249–286 9. Utsunomiya-Tate N, Endou H, Kanai Y (1996) Cloning and functional characterization of a system ASC-like Na+-dependent neutral amino acid transporter. J Biol Chem 271:14883–14890 10. Torres-Zamorano V, Leibach FH, Ganapathy V (1998) Sodium-dependent homo- and hetero-exchange of neutral amino acids mediated by the amino acid transporter ATB. Biochem Biophys Res Commun 245:824–829 11. Broer A, Brookes N, Ganapathy V, Dimmer KS, Wagner CA, Lang F, Broer S (1999) The astroglial ASCT2 amino acid transporter as a mediator of glutamine efflux. J Neurochem 73:2184–2194
12. Dolinska M, Dybel A, Zablocka B, Albrecht J (2003) Glutamine transport in C6 glioma cells shows ASCT2 system characteristics. Neurochem Int 43:501–507 13. Dolinska M, Zablocka B, Sonnewald U, Albrecht J (2004) Glutamine uptake and expression of mRNA’s of glutamine transporting proteins in mouse cerebellar and cerebral cortical astrocytes and neurons. Neurochem Int 44:75–81 14. Lim J, Lorentzen KA, Kistler J, Donaldson PJ (2006) Molecular identification and characterisation of the glycine transporter (GLYT1) and the glutamine/glutamate transporter (ASCT2) in the rat lens. Exp Eye Res 83:447–455 15. Oppedisano F, Pochini L, Galluccio M, Cavarelli M, Indiveri C (2004) Reconstitution into liposomes of the glutamine/amino acid transporter from renal cell plasma membrane: functional characterization, kinetics and activation by nucleotides. Biochim Biophys Acta 1667:122–131 16. Oppedisano F, Pochini L, Galluccio M, Indiveri C (2007) The glutamine/amino acid transporter (ASCT2) reconstituted in liposomes: transport mechanism, regulation by ATP and characterization of the glutamine/glutamate antiport. Biochim Biophys Acta 1768:291–298 17. Biber J, Stieger B, Haase W, Murer H (1981) A high yield preparation for rat kidney brush border membranes. Different behaviour of lysosomal markers. Biochim Biophys Acta 647:169–176 18. Krämer R, Heberger C (1986) Functional reconstitution of carrier proteins by removal of detergent with a hydrophobic ion exchange column. Biochim Biophys Acta 863:289–296 19. Palmieri F, Indiveri C, Bisaccia F, Iacobazzi V (1995) Mitochondrial metabolite carrier proteins: purification, reconstitution, and transport studies. Methods Enzymol 260:349–369 20. Pochini L, Oppedisano F, Indiveri C (2004) Reconstitution into liposomes and functional characterization of the carnitine transporter from renal cell plasma membrane. Biochim Biophys Acta 1661:78–86
Chapter 6 Use of Liposomes for Studying Interactions of Soluble Proteins with Cellular Membranes Chris T. Höfer, Andreas Herrmann, and Peter Müller Abstract Methods are described that have been used for characterizing the interaction of the soluble bovine seminal plasma protein PDC-109 with liposomes. PDC-109 binds to bull sperm cells upon ejaculation and is an important modulating factor of sperm cell maturation. The binding of the protein to sperm cells is mediated via lipids of the sperm plasma membrane. Most of our current knowledge about the molecular mechanisms of PDC-109–membrane interaction has been obtained by studies employing lipid vesicles. The experimental strategy described here can be applied to investigate the interaction of soluble proteins with membranes in order to understand their physiological role. Key words: Protein–membrane interaction, Membrane integrity, Phospholipid, Seminal plasma protein, PDC-109, Fluorescence spectroscopy, ESR spectroscopy
1. Introduction The interaction of soluble proteins with membranes is crucial for the regulation of biological activities. Proteins become recruited to the leaflets of plasma or intracellular membranes by binding to protein receptors or to lipids which results in adhesion and/or insertion of the protein to the membrane. Thereby, the protein and/or membrane structure and dynamics are influenced, finally resulting in a modified biological activity. Some examples for soluble proteins that modulate physiological functions upon binding to membranes are (1) phospholipases influencing phospholipid metabolism (1), (2) annexins which are supposed to play a role in membrane trafficking events such as exocytosis, endocytosis, and cell–cell adhesion (2), (3) bacterial toxins having deleterious impact on cells (3, 4), (4) mammalian seminal plasma proteins V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_6, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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which interact with the plasma membrane of sperm cells modulating the genesis of the cells, and (5) amyloid proteins like a-synuclein which is probably involved in the genesis of Parkinson’s disease (5, 6). With regard to proteins of seminal plasma, a protein family which is characterized by an Fn type II domain has gained special importance. This domain was originally described for the extracellular matrix protein fibronectin, in which it constitutes part of the collagen binding region and is implicated in a variety of extracellular binding events (7). Bull seminal plasma contains various Fn type II proteins, the most prominent representative being PDC-109 (also named BSP-A1/A2) (8). PDC-109 binds to sperm cells upon ejaculation and influences the capacitation process of the cells (9). To understand the physiological role of PDC109 on a molecular level, numerous studies, in particular using liposomes, have been perfomed (for reviews see (10, 11)). It has been found that the interaction with spermatozoa is realized via binding to membrane phospholipids carrying a choline head group, which are phosphatidylcholine (PC) and sphingomyelin (SM). The lipid-binding sites have been characterized from the protein crystal structure (12). Upon binding to vesicles, the protein influences membrane structure and dynamics, in that e.g., (1) the mobility of lipids is reduced, (2) membrane integrity is impaired, and (3) lipids are extracted from the membrane. Here, the application of various methods is described to identify and characterize important aspects of the interaction of PDC-109 with membranes by using liposomes.
2. Materials 2.1. Isolation of PDC-109
1. PDC-109 is purified from the seminal plasma of reproductively active Holstein bulls by a combination of affinity chromatography on heparin-Sepharose and DEAE-Sephadex chromatography as described by Calvete et al. (13). 2. The purity of the protein is verified by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) as well as reverse-phase HPLC, N-terminal sequence amino acid, and mass spectrometric analyses. 3. For the experiments, a 1 mM stock solution of the protein is prepared in Hepes-buffered salt solution (HBS) containing 145 mM NaCl and 5 mM HEPES, pH 7.4.
2.2. Lipids
1. Lipids are obtained from Avanti Polar Lipids (Alabaster, AL), if not stated otherwise and used without further purification. Stock solutions of lipids are prepared in chloroform or chloroform/ methanol (1:1) and stored at −80°C (see Notes 1 and 2).
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2.3. Fluorescence Measurements
1. All fluorescence measurements are done at 37°C using an Aminco Bowman Series 2 spectrofluorometer (SLM-AMINCO, Rochester, NY) with 4 nm slit width for both excitation and emission.
2.4. Electron Spin Resonance (ESR) Measurements
1. The spin-labeled analog of PC, 1-palmitoyl-2-(4-doxylpentanoyl)sn-glycero-3-phosphocholine (SL-PC), is synthesized according to (14). 2. ESR spectra are recorded at 4°C using a Bruker EMX spectrometer (Bruker, Karlsruhe, Germany) with the following parameters: modulation amplitude 1 G, power 20 mW, scan widths 100 G, accumulation nine times.
3. Methods 3.1. Preparation of Large Unilamellar Vesicles (LUVs)
1. Appropriate quantities of the lipid stock solutions (including when necessary fluorescent or spin-labeled lipids) are transferred into a glass tube, and the solvent is removed under a stream of nitrogen and subsequently under vacuum for 1 h. For labeled vesicles, one thereby obtains symmetrically labeled membranes, i.e., the analog is localized on both leaflets. 2. Lipids are resuspended in a small volume of ethanol (giving a final ethanol concentration below 1% (v/v)). HBS is added and the mixture is vortexed to induce the formation of multilamellar vesicles (see Note 3). 3. For the preparation of LUVs, the lipid suspension is subjected to five freeze–thaw cycles and extruded 10 times at 40°C through a 100-nm-diameter polycarbonate filter (Nucleopore GmbH, Tübingen, Germany) using either an extruder (Extruder, Lipex Biomembranes Inc., Vancouver, Canada) or a mini-extruder (Avanti Polar Lipids, Alabaster, AL) (15) (see Note 4).
3.2. Binding of PDC-109 to Liposomes Measured by Flotation Assay
1. As a first approach to characterize membrane binding of a protein, ultracentrifugation techniques can be used to separate the liposome-bound fraction of the protein from the free unbound protein. A variety of methods based upon this principle exists, and the appropriate one has to be chosen according to individual requirements. Either liposomes and bound protein can be pelleted using multilamellar vesicles (16) or sucrose-loaded vesicles (17), or liposomes and bound protein are floated on a sucrose density gradient (18). The flotation technique allows a clear separation of membrane-bound floating proteins especially for studying proteins that tend to aggregate or form large complexes that might be pelleted independently of the binding to liposomes. The assay can
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serve for qualitative as well as quantitative analysis and can be utilized to investigate binding conditions such as pH, ionic strength, and lipid specificity. 2. LUVs are prepared consisting of 2 mM 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 0.5 mol% 1,2-dipalmitoylsn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) (N-NBD-DPPE) (see Notes 3 and 5). 3. Sucrose solutions (75% and 25% w/v) are prepared in HBS. 4. PDC-109 (12 µM) is mixed with different amounts of LUVs in a total volume of 150 µl in 1.5-ml Polyallomer Microfuge tubes for ultracentrifugation (Beckman Instruments Inc., Palo Alto, CA). 5. The solution is mixed thoroughly with 100 µl of 75% sucrose giving a final sucrose concentration of 30%. This bottom fraction is carefully overlayered with 200 µl of 25% sucrose and 50 µl of HBS. 6. The tubes are centrifuged for 1 h at 240,000 g and 4°C using an ultracentrifuge TL-100 and rotor TLA-100.3 (Beckman Instruments Inc., Palo Alto, CA). 7. Immediately after centrifugation, the fractions of the gradient are collected manually (see Note 6). 8. Aliquots of all fractions are analyzed by SDS-PAGE followed by silver staining. 9. The fluorescence of N-NBD-DPPE can be detected in the front part of the gel by scanning the gel before or after silverstaining on a fluorescent image analyzer FLA-3000 (FUJIFILM, Düsseldorf, Germany) with excitation at 473 nm. 10. Flotation of PDC-109 with DOPC-LUVs shows an increase in unbound protein with decreasing lipid concentration (Fig. 1). Above a protein-to-lipid ratio of 1:40, all protein is membrane-bound, whereas at 1:20 part of the protein is unbound. Since only the lipids of the outer leaflet of LUVs are accessible to protein binding, these data indicate a binding stoichiometry of about 10–20 lipid molecules per one protein molecule, which is in agreement with other studies (19). 3.3. Binding of PDC-109 to Liposomes Measured by Intrinsic Protein Fluorescence
1. Upon binding to membranes, protein structure is often modified. Those changes might be followed by measuring the intrinsic protein fluorescence if the protein contains fluorescent amino acids. Among these, tryptophan is most useful because of its comparatively high quantum yield. Since fluorescence is dependent on the local environment of the fluorophore, conformational changes can be monitored at least for those parts of the protein containing fluorescent amino acids. 2. PDC-109 is diluted from the stock solution to a concentration of 2.5 µM into a fluorescence cuvette containing lipid vesicles and
Protein-Membrane Interaction sucrose gradient
0% 25 %
= LUVs
NBD-labeled LUVs + protein
73
NBD fluorescence 1h
Collecting
240 000 g
the fractions
30 % 0% 25% 30%
Fig. 1. Top: Schematic diagram of the flotation assay. Bottom: Flotation assay analysis of PDC-109 binding to lipid vesicles. PDC-109 was floated with DOPC-LUVs in a sucrose step gradient at different protein-to-lipid ratios. After centrifugation, the fractions of the gradient with 30% sucrose (bottom fraction), 25% sucrose (middle fraction), and 0% sucrose (floating liposome fraction) were analyzed by SDS-PAGE, and subsequent silver staining showed a decrease of floating liposomebound protein with decreasing lipid concentration. The fluorescence of the membrane-anchored liposome label N-NBD-DPPE in the gel front was detected on a fluorescent image analyzer, and indicated the presence of LUVs in 0% and 25% fractions if the overall lipid concentration was sufficient. The bottom fraction was devoid of lipids 0.6
fluorescence intensity [a.u.]
3 0.5
2 0.4
1 0.3 0.2 0.1 0.0 300
320
340
360
380
400
420
wave length [nm] Fig. 2. Influence of lipid vesicles on the intrinsic fluorescence of PDC-109. The fluorescence spectra (excitation at 280 nm) of 2.5 µM PDC-109 were recorded at 37°C in the absence of liposomes (curve 1) and in the presence of 25 µM eggPC/ eggPE(2:1)-LUVs (curve 2 ) or eggPC-LUVs (curve 3). Note the shift of the wavelength of the maximum fluorescence intensity upon addition of LUVs
fluorescence spectra are recorded in the range of 300–400 nm (excitation wave length 280 nm) (Note 7). 3. Upon interaction with eggPC-LUVs, the intrinsic fluorescence of PDC-109 is increased and shifted to shorter wave lengths (Fig. 2, curves 1 and 3) (see Note 8). These data indicate a localization of tryptophan residues in a more hydrophobic
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environment in the presence of vesicles which is probably caused by an intercalation of (part of) the protein into the bilayer (see Notes 9 and 10). 4. Mixing of PDC-109 with eggPC/eggPE(2:1)-LUVs results in lower changes of protein fluorescence compared to those of pure PC liposomes (Fig. 2, curve 2). This indicates that PDC-109 has a specificity for phosphorylcholine-containing phospholipids (19, 20). 3.4. Influence of PDC-109 on Membrane Integrity Measured by Calcein Leakage
1. LUVs are loaded with a high concentration of the nonpermeable fluorophore calcein, resulting in low fluorescence intensity due to self-quenching. A perturbation of membrane integrity results in a release of the fluorophore from the vesicles, which can be measured by the increase of fluorescence. 2. LUVs are prepared in HBS containing additionally 70 mM of calcein (Fluka Feinchemikalien, Neu-Ulm, Germany) (see Note 11). 3. Calcein-filled vesicles are separated from bulk calcein using NAP-5 columns (GE Healthcare, Freiburg, Germany) at room temperature and HBS as elution buffer (see Note 12). 4. Calcein-filled LUVs are diluted with HBS in a fluorescence cuvette (giving a final lipid concentration of 10–20 µM, see Note 13) while continuously stirring the solution. The timedependent fluorescence is monitored at 515 nm (excitation wave length 490 nm). After about 30 s, 5 µM PDC-109 is added and maximal leakage is determined by addition of 0.5% (w/v) Triton X-100 after about 230 s. 5. The addition of PDC-109 to eggPC-LUVs induces a release of calcein, indicating that the protein is able to disturb the membrane integrity of these vesicles (Fig. 3) (see Note 14). In contrast, the perturbation of eggPC/eggPE membranes in the presence of PDC-109 is significantly lower as seen from the low calcein leakage, a result again supporting the specificity of PDC-109 for phosphorylcholine-containing lipids.
3.5. Verification of PDC-109 Intercalation into Membrane Bilayer Measured by Förster Resonance Energy Transfer (FRET)
1. In order to see whether PDC-109 intercalates into the hydrophobic lipid phase upon binding to LUVs, FRET was recorded from tryptophan residues of the protein (donor) to the fluorescent lipid 1-hexadecanoyl-2-(1-pyrenedecanoyl)-sn-glycero-3phosphocholine (pyrPC, Invitrogen Ltd, Karlsruhe, Germany). This analog bears the pyrene group (acceptor) in the hydrophobic part of the membrane. A FRET signal strongly indicates a penetration of tryptophan residues into the lipid bilayer since for such signal both fluorophores have to approach below 10 nm. 2. EggPC-LUVs are labeled with 1 mol% pyrPC (see Subheading 3.1). Fluorescence spectra of labeled LUVs (final lipid concentration 20 µM) are recorded between 300 and
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normalized fluorescence
1.0
Triton X-100 0.8
0.6
PDC-109
1
0.4
0.2
2 0.0 0
50
100
150
200
250
time [s] Fig. 3. Influence of PDC-109 on the leakage of calcein from lipid vesicles. The time-dependent fluorescence of calcein was measured at 37°C for LUVs composed of eggPC (curve 1) or eggPC/eggPE (2:1) (curve 2). Leakage was induced by adding 5 µM PDC-109 to the vesicles at time zero, giving a final lipid-to-protein ratio of about 4. After 230 s, Triton X-100 was added (final concentration 0.5 % (w/v)) to obtain complete leakage of calcein, which was set to 1
fluorescence intensity [a.u.]
2.5
8 6
2.0
4 2
1.5
1.0
2
0 300
350
400
450
3 4
1
3
0.5
0.0 300
350
400
450
500
550
wave length [nm] Fig. 4. Förster resonance energy transfer (FRET) from tryptophan residues of PDC-109 to pyrene-labeled PC. eggPC-LUVs were labeled with 1 mol% pyrPC, and the fluorescence spectra of PDC-109 and pyrPC were recorded at 37°C in the absence (curve 1) and in the presence of unlabeled (L/P = 10, curve 2) and labeled PC-LUVs (L/P = 10, curve 3) with excitation at 280 nm as well as in the presence of labeled LUVs (L/P = 10, curve 4) with an excitation at 345 nm, i.e., direct excitation of pyrene. The inset shows the spectra with the entire range of the y-axes
500 nm in the absence and in the presence of 2 µM PDC-109 with excitation at 280 nm (Fig. 4). For comparison, fluorescence spectra of PDC-109 without vesicles and with unlabeled PC-LUVs were also recorded (see Notes 15 and 16).
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3. The spectra reveal that part of the PDC-109 excitation energy is transferred to the pyrene moiety as seen from a decrease in tryptophan fluorescence at around 340 nm and an increase of pyrene (monomer) fluorescence between 380 and 450 nm (Fig. 4, compare unlabeled and labeled LUVs in the presence of protein, curves 2 and 3, respectively). 4. From these data, it can be concluded that upon binding of PDC-109 to lipid membranes (part of) the protein bearing tryptophan residues intercalates into the membrane phase (see Notes 9 and 17). 3.6. Interaction of PDC-109 with Spin-Labeled Lipids
1. ESR spectra of spin-labeled lipids localized in membranes reflect the lipid mobility and are sensitive to lipid–protein interactions (21). 2. EggPC-LUVs are labeled with 2.5 mol% of SL-PC (see Subheading 3.1). 3. Labeled LUVs (2 mM) are mixed with 0.2 mM PDC-109 and, after a 5 min incubation on ice, ESR spectra are recorded at 4°C. 4. Figure 5 (spectrum 1) shows a typical membrane spectrum of SL-PC in eggPC-LUVs, reflecting a partially restricted motion of the analog within the membrane. 5. In the presence of PDC-109, one observes an additionally superimposed spectrum which is especially visible in the region of the low-field peak (Fig. 5, spectrum 2, see arrow).
10 G 1
2
3
Fig. 5. Influence of PDC-109 on the ESR spectrum of spin-labeled membranes. 2 mM eggPC-LUVs were labeled with 0.05 mM SL-PC, and the ESR spectra were recorded at 4°C in the absence (spectrum 1) and in the presence (spectrum 2) of 0.2 mM PDC-109. The immobilized component caused by PDC-109 (see arrow) was extracted by spectra subtraction, giving spectrum 3
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6. The spectral component that originated in the presence of PDC-109 can be obtained by subtracting the ESR spectrum in the absence of protein from that in the presence of protein (Fig. 5, spectrum 3) (see Note 18). This spectrum reflects a decreased mobility of the analog as seen from the peak broadening, indicating that PDC-109 causes an effective restriction of the lipid mobility in PC membranes (22) (see Note 19). 3.7. Summary
The experiments described here allow characterization of the interaction of a soluble protein with membranes. Other approaches partly using liposomes are described in the literature. To study PDC-109, the following methods have been also applied: Fouriertransform infrared (FTIR) spectroscopy, CD spectroscopy, and calorimetric methods (differential scanning calorimetry, isothermal titration calorimetry) (see (10, 11)). The results of those experiments, in combination with in vitro and in vivo studies on the biologically relevant cellular system, allow the elucidation of the physiological role(s) of the respective protein (11, 12).
4. Notes 1. In case of storage for longer times, we prefer to store the lipids in the dry form, i.e., after evaporation of the solvent, at −80°C in order to prevent the lipids from any decomposition. 2. During handling with the organic solutions of lipids, the usage of any plastics should be prevented and solely glassware (tubes, pipette tips etc.) should be used since the solvents may dissolve substances (e.g., softeners) from the plastics. 3. When preparing liposomes, the detaching of lipids from the glass wall and their dissolution in the buffer is, in our opinion, an important point to be considered. Certain lipid species may be incompletely dissolved, resulting in a deviation of the desired lipid composition of liposomes. In general, unsaturated PC species or the often-used eggPC mixture can be easily dissolved by the addition of buffer and subsequent vortexing. However, problems may arise when using other lipid such as PE, SM, or cholesterol (also when used in a mixture with PC species) due to their distinct physicochemical parameters. For example, it is extremely difficult to dissolve pure cholesterol in an aqueous buffer. The dissolution of lipids could be facilitated by either pre-resuspending the dried lipids in a small volume of ethanol and/or by performing all steps of liposome production at higher temperatures. For example, when using SM species (which are characterized by
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long fatty acid chains) we work at 65°C. After preparation of liposomes, lipid concentration of vesicles can be checked by measuring the phospholipid and, if necessary, the cholesterol content (see [23] and references therein). For measuring the lipid composition, the use of thin-layer chromatography or mass spectrometric methods (ESI, MALDI-TOF) is recommended. 4. Basically, different kinds of lipid vesicles can be used: small unilamellar vesicles (SUVs), large unilamellar vesicles (LUVs), or multilamellar vesicles (MLVs). SUVs are prepared by sonication of aqueous lipid dispersion. These vesicles are rapidly prepared. However, owing to their small size (diameter of about 30 nm), they have a large surface curvature questioning their use as a model for a biological membrane. Moreover, SUVs are comparatively unstable and fuse. Therefore, those vesicles should be used shortly after preparation. MLVs can also be prepared easily (see Subheading 3.1). However, due to the presence of more than one bilayer, these vesicles have an undefined accessible surface area. Therefore, use of MLVs could be unfavorable for certain problems. LUVs, although their preparation takes comparatively more effort, are appropriate for investigating numerous aspects of protein–membrane interaction, e.g., their membranes mimick biological membranes with regard to membrane curvature. After preparation, LUVs are supposed to be stable for several days; however, this might be checked. 5. The addition of the fluorescent lipid N-NBD-PE allows (1) visualization of the liposome fractions on a fluorescent image analyzer and (2) estimation of the loss of lipids during the preparation of LUVs by comparing the fluorescence of the lipid solution before and after vesicle preparation. For an accurate quantitative analysis, one has to determine the phospholipid and, if necessary, cholesterol content of LUVs (see Note 3). 6. There are two strategies for collecting the fractions. The three fractions of 250, 150, and 50 µl are collected from the bottom to the top with a Hamilton syringe, permitting a very clean separation of the fractions and restricting the liposomes mainly to the 0% sucrose floating fraction which is suitable for a quantitative analysis. But, if the protein adsorbs to the surface of the tubes as membrane-binding proteins often do, it is preferable to collect the fractions from the top to the bottom, as otherwise the floating fraction can become contaminated with protein sticking to the bottom of the tube. This can be checked by rinsing and boiling the emptied tube with 50 µl sample buffer and subsequent analysis by SDS-PAGE. After collecting the 0% and 25% fractions from the top, the 30% bottom fraction is taken from the bottom of the tube leaving
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a rest of 20 µl in the tube to prevent an incompletely removed lipid film from mixing with the bottom fraction. Therefore, the latter technique is not very suitable for quantitative analysis, but, nevertheless, can qualitatively show whether a protein binds to liposomes. If the utilized lipid concentration is high enough, one can visualize the localization of liposomes in the tube under UV light due to the fluorescence of N-NBDDPPE and estimate whether the floating of liposomes was succesful (Fig. 1). 7. In order to correct fluorescence spectra for the influence of light scattering caused by liposomes, the spectra of lipid vesicles in the absence of protein are also recorded and subtracted from those in the presence of protein. 8. For quantification of spectra, the fluorescence intensity at 333 nm is determined and normalized to the protein fluorescence in the absence of vesicles. By measuring the fluorescence changes at different lipid-to-protein ratios, the stoichiometry of protein lipid interaction can be determined which is about 10 for PDC-109 (19). Moreover, one can also estimate from the spectra the wavelength of the fluorescence maximum which is shifted to shorter wavelength (blue shift) upon change of tryptophan residues to a more hydrophobic environment. 9. If the protein to be investigated contains more than one tryptophan residue (for PDC-109 five residues), the exact determination which residue(s) intercalate(s) into the membrane bilayer is hampered since the measured fluorescence spectrum is the superposition of the spectra of each residue. The stepwise replacement of tryptophan residues by nonfluorescent amino acids using molecular biological approaches and measuring fluorescence spectra of those mutants in the presence of liposomes could allow the determination of the region(s) of the protein that are involved in membrane intercalation. Moreover, measurement of the life-time or quenching of fluorescence may allow distinguishing several tryptophan residues (usually 2–3). 10. In principle, an increase of fluorescence intensity and a blue shift of fluorescence reflect a change of the fluorophore to a more hydrophobic environment. However, changes of fluorescence with opposite tendencies in the presence of liposomes do not argue inevitably against a protein–membrane interaction since tryptophan fluorescence could also be influenced by other interactions, e.g., leading to fluorescence quenching. Moreover, if the protein solely adheres to the membrane surface, it is possible that the intrinsic fluorescence remains unchanged.
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11. For preparing the calcein solution, first add only half of the necessary volume of buffer to calcein, then adjust the pH of the solution with NaOH to pH = 7.5, and finally fill up with buffer to the desired volume. The high calcein concentration (70,000 times the concentration of general fluorescence measurements) causes an extreme contamination of all labware that comes into contact with this solution. Therefore, all labware has to be cleaned thoroughly after contact with calcein solutions protecting other colleagues from undesirable calcein measurements. This especially holds for the extruder. To prevent its expensive cleaning, we used to reserve one mini-extruder in the lab solely for calcein measurements. 12. NAP-5 columns calibrated for the addition of 500 µl solution are used according to the manufacturer’s instructions. If the volume of liposome solution is smaller and has to remain undiluted, one can load the original solution onto the column and allow it to enter the gel bed completely. Subsequently, the eluting buffer is added. The running front containing the liposomes can be easily followed by eye because of the intense color of calcein and the eluant is collected by dropwise sampling. 13. In case LUVs are diluted after running on NAP-5 columns, the final lipid concentration varies resulting in different molar lipid-to-protein ratios in different experiments. Therefore, in order to estimate the exact ratio, the lipid concentration of calcein-loaded LUVs has to be measured after column chromatography (see [23]). 14. The degree of leakage mediated by PDC-109 increases with rising protein concentrations (23). The leakage kinetics is quantified by determining the amount of leakage (L) according to
L=
Ft − F0 Fmax − F0
where F0 and Fmax refer to the initial fluorescence intensity of calcein-filled LUVs before the addition of PDC-109 and the fluorescence intensity after addition of Triton X-100, respectively (see Fig. 3). Ft denotes to the fluorescence intensity reached in the plateau phase after addition of PDC-109 (in the experiment of Fig. 3 after 200 s). Intensities are corrected for dilution due to the addition of PDC-109 and Triton X-100. 15. Since pyrene is sensitive to quenching by oxygen, it is recommended to degas buffers with nitrogen before using. 16. As in our experiments, one may observe some pyrene fluorescence by excitation of a control sample (i.e., labeled LUVs in the absence of protein) at 280 nm. Therefore, this spectrum is subtracted from that measured in the presence of protein to extract the component solely caused by FRET (see Fig. 4).
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17. As a negative control, in order to exclude that FRET occurs from protein dissolved in the bulk buffer onto pyrPC in the membrane, a search was made for FRET between the soluble, tryptophan-containing protein bovine serum albumin and pyrPC labeled LUVs. Upon mixing bovine serum albumin with labeled vesicles, no FRET-dependent pyrene fluorescence was detected at all (24). 18. The extent of immobilization, i.e., the amount of spin-labeled lipids affected by the protein, can be estimated by quantifying the subtraction (22). However, this quantification presumes that (1) solely two fractions of lipids exist in the presence of the protein (i.e., a part of molecules influenced by the protein and the other not influenced) and (2) the exchange rate between these states is negligible in the timescale of ESR. 19. The spin-labeled PC used here is characterized by a short fatty acyl chain bearing the spin moiety at the sn2 position of the glycerol backbone. This feature allows incorporation of the analog also into preformed membranes labeling solely the outer leaflet of LUVs membranes (22). However, those experiments can also be performed by using other spin-labeled phospholipid analogs having different head groups and/or two long fatty acyl chains (long-chain PC analogs available from Avanti Polar Lipids, Alabaster, AL) or spin-labeled steroids (partly available from Sigma-Aldrich Chemie GmbH, Taufkirchen, Germany) in order to characterize the lipid specificity of the protein (22, 25). Adding these analogs during liposome preparation results in symmetrically labeled vesicles (see also Subheading 3.1); however, the labeling of preformed membranes might be difficult.
Acknowledgments The work was supported by the Deutsche Forschungsgemeinschaft (Mu 1017/2). The fruitful cooperation during this project with Edda Töpfer-Petersen is kindly acknowledged. We thank Anja Arbuzova for critical reading of the manuscript. References 1. Winget JM, Pan YH, Bahnson BJ (2006) The interfacial binding surface of phospholipase A2s: PLA2. Biochim Biophys Acta 1761:1260–1269 2. Waisman DM (1995) Annexin II tetramer: structure and function. Mol Cell Biochem 150:301–322
3. Cabiaux V, Wolff C, Ruysschaert JM (1997) Interaction with a lipid membrane: a key step in bacterial toxins virulence. Int J Biol Macromol 21:285–298 4. Palmer M (2004) Cholesterol and the activity of bacterial toxins. FEMS Microbiol Lett 238:281–289
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5. Davidson WS, Jonas A, Clayton DF, George JM (1998) Stabilization of alpha-synuclein secondary structure upon binding to synthetic membranes. J Biol Chem 273:9443–9449 6. Beyer K (2007) Mechanistic aspects of Parkinson’s disease: alpha-synuclein and the biomembrane. Cell Biochem Biophys 47:285–299 7. Skorstengaard K, Thogersen HC, Petersen TE (1984) Complete primary structure of the collagen-binding domain of bovine fibronectin. Eur J Biochem 140:235–243 8. Manjunath P, Sairam MR (1987) Purification and biochemical characterization of three major acidic proteins (BSP-A1, BSP-A2 and BSP-A3) from bovine seminal plasma. Biochem J 241:685–692 9. Therien I, Bleau G, Manjunath P (1995) Phosphatidylcholine-binding proteins of bovine seminal plasma modulate capacitation of spermatozoa by heparin. Biol Reprod 52: 1372–1379 10. Wah DA, Fernández-Tornero C, Sanz L, Romero A, Calvete JJ (2002) Sperm coating mechanism from the 1.8 A crystal structure of PDC-109-phosphorylcholine complex. Structure 10:505–514 11. Manjunath P, Therien I (2002) Role of seminal plasma phospholipid-binding proteins in sperm membrane lipid modification that occurs during capacitation. J Reprod Immunol 53:109–119 12. Ekhlasi-Hundrieser M, Müller P, TöpferPetersen E (2008) Male secretory proteins sperm tools for fertilisation. In: Glander HJ, Paasch U (eds) Biology of male germ cells. Shaker Publisher GmbH, Aachen, Germany 13. Calvete JJ, Varela PF, Sanz L, Romero A, Mann K, Töpfer-Petersen E (1996) A procedure for the large-scale isolation of bovine seminal plasma proteins. Protein Expr Purif 8:48–56 14. Fellmann P, Zachowski A, Devaux PF (1994) Synthesis and use of spin-labeled lipids for studies of the transmembrane movement of phospholipids. Methods Mol Biol 27:161–175 15. Mayer LD, Hope MJ, Cullis RP, Janoff AS (1985) Solute distributions and trapping efficiencies observed in freeze-thawed multilamellar vesicles. Biochim Biophys Acta 817:193–196
16. Heuer K, Arbuzova A, Strauss H, Kofler M, Freund C (2005) The helically extended SH3 domain of the T cell adaptor protein ADAP is a novel lipid interaction domain. J Mol Biol 348:1025–1035 17. Buser CA, McLaughlin S (1998) Ultracentrifugation technique for measuring the binding of peptides and proteins to sucrose-loaded phospholipid vesicles. Methods Mol Biol 84:267–281 18. Bigay J, Casella JF, Drin G, Mesmin B, Antonny B (2005) ArfGAP1 responds to membrane curvature through the folding of a lipid packing sensor motif. EMBO J 24: 2244–2253 19. Desnoyers L, Manjunath P (1992) Major proteins of bovine seminal plasma exhibit novel interactions with phospholipid. J Biol Chem 267:10149–10155 20. Müller P, Erlemann KR, Müller K, Calvete JJ, Töpfer-Petersen E, Marienfeld K, Herrmann A (1998) Biophysical characterization of the interaction of bovine seminal plasma protein PDC-109 with phospholipid vesicles. Eur Biophys J 27:33–41 21. Marsh D, Horvath LI (1998) Structure, dynamics and composition of the lipidprotein interface. Perspectives from spinlabelling. Biochim Biophys Acta 1376: 267–296 22. Greube A, Müller K, Töpfer-Petersen E, Herrmann A, Müller P (2001) Influence of the bovine seminal plasma protein PDC-109 on the physical state of membranes. Biochemistry 40:8326–8334 23. Tannert A, Töpfer-Petersen E, Herrmann A, Müller K, Müller P (2007) The lipid composition modulates the influence of the bovine seminal plasma protein PDC-109 on membrane stability. Biochemistry 46:11621–11629 24. Greube A, Müller K, Töpfer-Petersen E, Herrmann A, Müller P (2004) Interaction of fibronectin type II proteins with membranes: the stallion seminal plasma protein SP-1/2. Biochemistry 43:464–472 25. Müller P, Greube A, Töpfer-Petersen E, Herrmann A (2002) Influence of the bovine seminal plasma protein PDC-109 on cholesterol in the presence of phospholipids. Eur Biophys J 31:438–447
Chapter 7 Liposomal Reconstitution of Monotopic Integral Membrane Proteins Zahra MirAfzali and David L. DeWitt Abstract In spite of considerable progress in the methodology for reconstitution of membrane proteins into the liposomes, a successful reconstitution still appears to be more an art than a science. Reconstitution of membrane proteins into bilayers is required for establishing several aspects of the functions of membrane proteins and lipids and for elaborating models of naturally occurring membranes. Cyclooxygenase (COX)-1 and -2 (also prostaglandin endoperoxide H2 synthase, PGHS-1 and -2) belong to the class of monotopic membrane proteins. Membrane-binding domains of both COX-1 and -2 contain four short, consecutive, amphipathic a-helices (A, B, C, and D). Crystal structures of the COXs indicate that basic, hydrophobic, and aromatic residues in the membrane-binding domain are oriented away from the protein core and form a surface on the enzyme, which has been proposed to interact with the lipid bilayer (1). In this chapter, we describe a fast and efficient method for direct incorporation of COX-1 and -2 isozymes – as models for monotopic integral membrane proteins – into preformed liposomes containing fatty acids without loss of activity. Key words: Monotopic membrane protein, Proteoliposomes, Liposome reconstitution, Direct incorporation, Membrane defect, Incorporating impurities into membranes, Cyclooxygenase enzyme
1. Introduction Monotopic membrane proteins such as cyclooxygenase (COX)-1 and -2 interact only with one leaflet of the lipid bilayer. Detergent removal is a classical method that is used for incorporation of integral membrane proteins into liposomes. However, this technique does not work with monotopic membrane proteins such as COX isozymes. Thus, it seems that a more generally applicable method is needed for this class of proteins. Medium-length-chain V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_7, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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(C12–28) fatty acids, cholates, and cholesterol have been shown to promote the reconstitution of integral membrane proteins into liposomes. Therefore, one of the key features for successful incorporation of solubilized membrane proteins into preformed liposomes appears to be the state of organization of the lipid bilayer. Bilayers conducive to direct incorporation of large membrane proteins are achieved by incorporating “impurities”, such as fatty acids, lysophospholipids, mixtures of structurally different phospholipids, and cholesterol. The organizational defects in a bilayer decreases the activation energy for insertion of proteins and increases the free energy change associated with incorporation of proteins [2]. In this chapter, we explain the step by step process of incorporating a monotopic membrane protein into preformed liposomes using direct incorporation method.
2. Materials 2.1. Expression of Recombinant COX into SF21 Insect Cells
1. Spodoptera frugiperda (SF21) insect cells (Invitrogen Corpo ration, Carlsbad, CA). 2. Recombinant baculovirus expressing His-tagged COX-1 or COX-2 isozymes. 3. Media: HyQ-SFX-Insect serum-free insect cell media (Hyclone, Logan, UT) is supplemented with 0.1% pluronic F-68, 1× lipid concentrate, and 0.2% glucose. 4. Fernbach flasks (Bellso Biotechnology Inc., Vineland, NJ). 5. Innova model 4000 benchtop gyrotory incubator shaker (New Brunswick Scientific Co. Inc, Edison, NJ). 6. Hausser Nageotte Bright-Line Hemacytometer (Thermo Fisher Scientific, Waltham, MA). 7. Zeiss light microscope (Thermo Fisher Scientific, Waltham, MA). 8. Beckman refrigerated floor centrifuge (Beckman-Coulter, Fullerton, CA). 9. DuPont-Sorvall HS-4 centrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 10. Nalgene centrifuge bottles for Sovall HS-4 centrifuge rotor (Thermo Fisher Scientific, Waltham, MA).
2.2. Purification of His-tagged COX from Baculovirus-Infected SF21 Insect Cells
1. Bioneb cell disruption system (Glas-Col, Terre Haute, Indiana). 2. Nitrogen cylinder. 3. Nickel-homogenization buffer: 25 mM NaPO4, 100 mM NaCl, 20 mM imidazole, pH = 7.4.
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4. Nickel-wash buffer: 25 mM NaPO4, 300 mM NaCl, 20 mM imidazole, pH = 7.4. 5. Nickel-elution buffer: 25 mM NaPO4, 100 mM NaCl, 200 mM imidazole, pH = 7.4. 6. Dialysis buffer: 25 mM Tris-HCl, 50 mM KCl, pH 7.4. 7. Fast-flow Ni-NTA Sepharose resin (Qiagen Inc, Valencia, CA). 8. Polypropylene centrifuge tubes 50 ml with caps (Thermo Fisher Scientific, Waltham, MA). 9. Beckman refrigerated floor ultracentrifuge (BeckmanCoulter, Fullerton, CA). 10. Sorvall T865 ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 11. Ultracentrifuge tubes for Sorvall T865 ultracentrifuge rotor (Beckman-Coulter, Fullerton, CA). 12. Slide-A-Lyzer dialysis cassette with 10,000 MWCO (Thermo Fisher Scientific, Waltham, MA). 13. Amcon ultracentrifuge filter device with 3,000 MWCO (Millipore, Billerica, MA). 2.3. Preparation of Unilamellar Liposomes
1. Buchi rotavapor with temperature-controlled water bath connected to a vacuum pump (Buchi Corporation, New Castle, DE). 2. Round-bottom flask (250 ml). 3. 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC) (Avanti Polar Lipids, Alabaster, AL). 4. 1,2-Dioleoyl-sn-Glycero-3-[Phospho-L-Serine] (Sodium Salt) (DOPS) (Avanti Polar Lipids, Alabaster, AL). 5. Avanti mini liposome extruder with heating block (Avanti Polar Lipids, Alabaster, AL). 6. Two gas-tight Hamilton syringes (1 ml) (Hamilton Company, Reno, NV). 7. Nuclepore polycarbonate membrane filter (100 nm pore size, 19 mm) (Thermo Fisher Scientific, Waltham, MA). 8. Liposome buffer: 25 mM Tris–HCl, 50 mM KCl, pH 7.4.
2.4. Incorporation of COX Enzyme into Preformed Liposomes
1. Microfuge tubes (2 ml) (Thermo Fisher Scientific, Waltham, MA).
2.5. Separation of Proteoliposomes from Free Proteins
1. Beckman refrigerated floor ultracentrifuge (BeckmanCoulter, Fullerton, CA).
2. Dry block temperature controlled incubator (Thermo Fisher Scientific, Waltham, MA).
2. Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA).
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3. Ultracentrifuge tubes for Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 4. Syringe and needle (Thermo Fisher Scientific, Waltham, MA). 2.6. Protein Quantification Using Bicinchoninic Acid(BCA)
1. Bicinchoninic acid (BCA) protein assay reagent (Thermo Fisher Scientific, Waltham, MA). 2. Albumin standard (2 µg/µl) (Thermo Fisher Scientific, Waltham, MA). 3. Thermo Fisher UV/vis micro-plate reader (Thermo Fisher Scientific, Waltham, MA). 4. Disposable plates (96-wells) (Thermo Fisher Scientific, Waltham, MA). 5. Incubator oven (37°C) (Thermo Fisher Scientific, Waltham, MA).
2.7. Phosphorous Quantification
1. Required solutions a. 8.9 N H2SO4: 123.5 ml of concentrated H2SO4 is added to 376.5 ml of deionized water. b. Ascorbic acid (10%): 5 g of ascorbic acid is put into a 50-ml volumetric flask. Fifty milliliters of deionized water is added. c. Ammonium molybdate(VI) tetrahydrate (2.5%): 1.25 g of ammonium molybdate(VI) tetrahydrate is put into a 50-ml volumetric flask. Fifty milliliters of deionized water is added. d. Phosphorus standard solution (0.64 mM): 0.1 ml of 32 mM phosphorus standard solution is added to 4.9 ml of deionized water. 2. Metal rack for test tubes (Thermo Fisher Scientific, Waltham, MA). 3. Glass test tubes (Thermo Fisher Scientific, Waltham, MA). 4. Thermo Fisher laboratory oven (Thermo Fisher Scientific, Waltham, MA). 5. Thermo Fisher UV/vis spectrophotometer (Thermo Fisher Scientific, Waltham, MA). 6. Spectrophotometer cuvettes (Thermo Fisher Scientific, Waltham, MA).
2.8. Cyclooxygenase Assay
1. Assay buffer: 0.1 M Tris–HCl, pH 8.0. 2. Required solutions a. Heme in DMSO: 1 ml solution of 8 mM of heme in DMSO is made and kept in the freezer. b. Arachidonic acid buffered solution: 1 ml solution of 100 µM arachidonic acid in assay buffer is made and kept in the refrigerator.
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3. YSI 5300A biological oxygen monitor (YSI, Yellow Springs, OH) 4. Hamilton syringe (10 µl) connected to a long needle (Hamilton Company, Reno, NV). 2.9. Gold Labeling Proteoliposomes for Electron Microscopy
1. Beckman refrigerated floor ultracentrifuge (Beckman-Coulter, Fullerton, CA). 2. Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 3. Ultracentrifuge tubes for Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 4. Syringe and needle (Thermo Fisher Scientific, Waltham, MA).
2.10. Transmission Electron Microscopy on Gold Labeled Proteoliposomes
1. Affinity-purified rabbit anti-COX antibody (custom made). 2. Gold immunoprobe NANO-GOLD-goat anti-rabbit IGGNRF (1.4-nm particles) (Immunoprobe, Yaphank, NY). 3. Beckman refrigerated floor ultracentrifuge (Beckman-Coulter, Fullerton, CA). 4. Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 5. Ultracentrifuge tubes for Sorvall SW 55 Ti ultracentrifuge rotor (Thermo Fisher Scientific, Waltham, MA). 6. Syringe and needle (Thermo Fisher Scientific, Waltham, MA). 7. Formvar (polyvinylformaldehyde)- carbon coated copper grids (G-300 mesh) (Electron Microscopy Sciences Inc, Fort Washington, PA). 8. Whatman filter paper #1 (Thermo Fisher Scientific, Waltham, MA). 9. JOEL transmission electron microscope (JOEL, Tokyo, Japan).
3. Methods Large quantities of COX enzyme are expressed in insect cells using baculovirus system containing the DNA of His-tagged COX enzyme. The purity of COX enzyme that is used for incorporation studies is more than 95%. The enzyme is purified using nickel-affinity chromatography method. In order to use this purification method, the protein is tagged with six histidines. In vitro mutagenesis is used to introduce a six-residue histidine sequence (His-tag) near the amino terminal end of the human COX-1 and -2. These isozymes are expressed using the baculovirus system. The His-tags are located one and two amino acids beyond the signal peptide cleavage sites of COX-1 and COX-2, respectively,
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positions that do not affect their activities or sensitivities to nonsteroidal anti-inflammatory drugs. When expressed in sf-21 cells, the His-tagged enzymes have Km values for arachidonate, and IC50 values for inhibition by nonsteroidal anti-inflammatory drugs that are similar to values reported for the nontagged enzymes. The His-tags allowed purification of the COX isozymes by a simplified protocol involving nickel-affinity chromatography [3]. Unilamellar liposomes containing impurities such as oleic acid are made in buffer and sized to 100 nm by extrusion using polycarbonate membrane filters. Various amounts of enzyme are added to fixed amount of liposomes in order to find the optimum protein-to-lipid ratio. Proteoliposomes (protein-incorporated liposomes) are separated from free liposomes (unincorporated proteins) using ficoll density gradient and ultracentrifugation methods. The incorporated proteins can be observed on the surface of the liposomes using immunogold labeling in combination with high-resolution transmission electron microscopy [4]. 3.1. Expression of Recombinant COX into SF21 Insect Cells
1. Spodoptera frugiperda (SF21) insect cells are added to sterile media in 1 L cultures at 27°C in 2.8-L Frenchback flasks shaken at 120 rpm using Innova model 4000 benchtop gyrotory incubator shaker. 2. The insect cells are counted every 8 h using Hausser Nageotte Bright-Line Hemacytometer and light microscope. 3. When cells reach a density of 1.5–2.0 × 106 cells/ml, baculovirus containing the DNA of COX enzyme is added at a multiplicity of infection of 1.0, and the infection is allowed to proceed for 72–96 h. 4. During the 72–96 h wait period, the cells are counted and lysed every 8 h and assayed for COX activity in order to monitor the progress of protein expression (see Note 1). 5. To harvest the cells, the cells and media are poured into Nalgene centrifuge bottles for Sorvall HS-4 centrifuge rotor. 6. The cells are pelleted at the bottom of the centrifuges tube using the Beckman refrigerated floor centrifuge operating at 10,000× g for 30 min. 7. The cells are washed using phosphate buffered saline (PBS) buffer and pelleted again using centrifugation. 8. The buffer is separated from the pelleted cells. 9. The tubes containing the cells are flash-frozen using liquid nitrogen and stored in a −80°C freezer.
3.2. Purification of His-tagged COX from Baculovirus-Infected SF21 Insect Cells
1. The tubes containing frozen cells are thawed at room temperature. 2. SF21 cell pellets are resuspended in 3 ml/g wet weight of nickel-homogenization buffer containing (1% v/v) of Tween 20 detergent.
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3. The cells are disrupted by Bioneb cell disruption system. The Bioneb is connected to high pressure nitrogen at 150 psi. The cells are passed through the system at least three times in order to achieve 100% disruption of all cells. 4. The cells are checked under the light microscope to make sure that they are completely lysed. 5. After low-speed (10,000× g for 1 h) centrifugation to pellet cell debris, the supernant which contains the solubilized COX is poured into cramp-seal ultracentrifuge tubes. 6. The insoluble material is removed by ultracentrifugation at 100,000× g for 2 h. 7. The solubilized extract is next incubated with fast-flow Ni-NTA resin overnight at 4°C with rocking. 8. The resin is poured into a column and washed with five volumes of nickel-homogenization buffer containing 0.1% Tween 20 detergent. 9. Next the resin is washed with three volumes of nickel-wash buffer containing 0.1% Tween 20. 10. The protein is eluted with nickel-elution buffer containing 0.2% Tween 20. 11. Fractions with high specific COX activity are pooled and concentrated in the Amicon ultracentrifugal filter device with 30,000 MWCO. 12. The purified COX enzyme is injected into a Slide-A-Lyzer (10,000 MWCO) dialysis cassette using a syringe. 13. The dialysis cassette is dialyzed against 2 L dialysis buffer. 14. The dialysis buffer is changed every 6 h for 24 h (see Note 2). 3.3. Preparation of Unilamellar Liposomes
1. Unilamellar liposomes are prepared using a DOPC: DOPS molar ratio of 3:7 and 9.2% (w/w) oleic acid. DOPC, DOPS, and oleic acid are dissolved in 1:1 (v/v) mixture of chloroform and methanol in a round bottom flask. 2. The round bottom flask is dried using a Buchi rotary evaporator connected to a vacuum pump for 4 h. 3. The resulting film is hydrated in a liposome buffer for 2 h. The total concentration of lipid is 38 mM. 4. The liposome dispersion is extruded 20 times through a polycarbonate membrane with 100 nm pore diameter using a handheld mini-extruder (see Note 3). Extruded liposomes form a transparent milky suspension.
3.4. Incorporation of COX Enzyme into Preformed Liposomes
1. The COX protein is added to the preformed liposomes. The concentration of the lipids in the liposome solution is 38 mM. A 1:500 protein: lipid molar ratio is used. 2. The mixture is incubated for 20 min at 37°C.
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3.5. Separation of Proteoliposomes from Free Proteins
1. A 0.5-ml liposome suspension is mixed with 1 ml of 30% (w/v) Ficoll in liposome buffer to give a final concentration of 20% (w/v) Ficoll. 2. The liposome suspension is transferred to a 5-ml ultracentrifuge tube. 3. Three milliliters of 10% (w/v) Ficoll is gently layered on top of the liposome suspension. 4. The swing-out rotor (Beckman SW 50.1) is used for 30 min at 100,000× g at 4°C. 5. The liposomes and proteoliposomes create a band at the interface between the 0% Ficoll and 10% Ficoll layers. A 1-ml syringe is used to isolate the proteoliposomes band from the buffer solution. The activity and concentration of the enzyme in the isolated proteoliposomes band are measured in order to calculate the incorporation efficiency of COX enzyme. 6. The unincorporated protein is retained in the 30% Ficoll layer. The activity and the concentration of the free protein are measured in order to calculate the incorporation efficiency of the COX enzyme.
3.6. Protein Quantification Using Bicinchoninic Acid (BCA)
1. The oven is turned on at 37°C. 2. Seven 1.5-ml tubes 0, 0.2, 0.4, 0.6, 0.8, 1, and 1.2 are labeled for the standard curve. 3. For the standard curve, 0, 10, 20, 30, 40, 50, and 60 µl of (2 µg/µl of albumin stock solution) is added to 100, 90, 80, 70, 60, 50, and 40 µl of water. 4. Samples for standard curve are added in triplicates at a volume of 10 µl to the wells in the 96-well plate. 5. Ten microliters of diluted aliquots of COX samples is added in triplicates to the wells in the 96-well plate. 6. Two hundred microliters of the BCA working reagent is added to each of the wells (some of the wells may turn purple after addition of the working reagent). 7. The plate that is covered by a lid is incubated for 30 min in the 37°C oven. 8. The plate is read at 562 nm. 9. The concentration of COX enzyme is calculated by linear regression of the standard curve.
3.7. Phosphorous Quantification
1. Liposome samples (~ 0.1 µmoles phosphorus) are placed in glass tubes. 2. Samples for standard curve are prepared by adding 0 (0 µmoles), 50 (0.032 µmoles), 100 (0.064 µl), 175 (0.112 µmoles), 250
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(0.160 µmoles), and 350 µl (0.224 µmoles) of phosphorus standard solution to each glass tube. 3. The sample and standard tubes are placed in a metal test tube rack. 4. To each of the tubes, 0.45 ml 8.9 N H2SO4 is added. Each tube is capped with a glass marble. 5. The samples are kept in oven at 200–215°C for 25 min. Temperature must be above 200°C. 6. Tubes are removed from the oven and cooled for 5 min. 7. To each tube, 150 µl H2O2 is added. 8. The tubes are returned to the oven and heated for an additional 30 min. The samples are colorless at this point (see Note 4). Tubes are cooled to ambient temperature. 9. To each tube, 3.9 ml deionized water is added. 10. To each tube, 0.5 ml of ammonium molybdate(VI) tetrahydrate solution is added. 11. The tubes are vortexed for 1 min. 12. To each tube, 0.5 ml ascorbic acid solution is added . 13. The tubes are vortexed for 1 min. 14. The tubes are heated at100°C for 7 min. Each tube is capped with a glass marble. 15. The tubes are cooled to room temperature. 16. The absorbance of each standard and sample is read at 820 nm. 17. The total amount of phosphorous in sample is calculated by linear regression of the standard curve. 3.8. COX Assay
1. To each chamber of YSI 5300A biological oxygen monitor device, 3 ml of assay buffer is added. The buffer is warmed up in to 37°C using the temperature control unit on the device. 2. The buffer inside the chamber is stirred using the magnet inside the chamber. 3. To the chamber, 50 µl of COX protein in added. 4. To the chamber ,5 µl of heme solution in DMSO is added (see Note 5). 5. The oxygen probe is inserted into the chamber. 6. Using a Hamilton syringe, 10 µl of arachidonic acid solution in buffer is added to the chamber. 7. The recorder that is connected to the oxygen probe records the rate of oxygen consumption. 8. The activity of the COX enzyme is calculated on the basis of the oxygen consumption rate.
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3.9. Gold Labeling Proteoliposomes for Electron Microscopy
1. About 100 ml of proteoliposomes is placed into an Eppendorf tube and 10 ml of primary antibody is added. The molar ratio of COX to antibody is 1:4. 2. The mixture is incubated for 30 min in room temperature with occasional shaking. 3. The nonattached primary antibodies are separated from the complex of proteoliposomes–primary antibodies using the Ficoll gradient technique, which was described previously. 4. The liposomes and the complex of proteoliposomes–primary antibodies are separated at the interface between the 0% Ficoll and 10% Ficoll layers. The unattached antibody will remain in the 30% Ficoll layer. 5. The complex of proteoliposomes–primary antibodies is placed into an Eppendorf tube and about 100 ml of gold immunoprobe NANOGOLD-anti rabbit (NRF) (1.4 nm particle attached to affinity-purified Fab fragment, raised in goat, against rabbit IgG (whole molecule)) is added to the tube and incubated for 30 min in room temperature with occasional shaking. 6. The nonattached gold-labeled secondary antibodies are separated from the complex of proteoliposomes–primary antibodies– gold labeled secondary antibodies using the Ficoll gradient method. 7. The liposomes and the complex of proteoliposomes–primary antibodies–gold labeled secondary antibodies are separated at the interface between the 0% Ficoll and 10% Ficoll layers. The unattached gold-labeled secondary antibody will remain in the 30% Ficoll layer.
3.10. Transmission Electron Microscopy on Gold Labeled Proteoliposomes
1. A drop of liposome solution (concentration about 5 mM of lipid) is placed on Formvar (polyvinylformaldehyde)-carbon coated copper grid G-300 mesh, and after 1 min the excess is removed with a Whatman filter paper #1. 2. A thin film is left on the grid and allowed to air dry. One drop of 0.1% solution of uranyl acetate is placed the grid (see Note 6). 3. After 1 min, this drop is again removed with the filter paper, and the resulting stained film is dried in a dust-free place. 4. A JEOL transmission electron microscope operating at 100 kV under vacuum is used to observe the liposomes (Figs. 1 and 2). Fixation and dehydration of liposome for electron microscopy flattens the liposomes and allows one to observe gold labeling on both sides of the liposomes. The average number of gold spots observed in each proteoliposomes is 123 ± 33 (n = 3). Assuming that average proteoliposomes are spheres of 100 nm, and that the two leaflets essentially double the lipid surface area, the total
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Fig. 1. Negative stain transmission electron micrograph of liposomes reconstituted with COX-2 and immunogold labeled with anti-COX-2 antisera (bar = 100 nm, magnification = 200,000). 0.1% uranyl acetate stain was used. Reproduced with permission from (4)
Fig. 2. Negative stain transmission electron micrograph of liposomes reconstituted with COX-2 (bar = 200,000 nm, magnification = 67,000). 1% uranyl acetate stain was used. Reproduced with permission from (4)
surface area of the proteoliposomes is (4p × 500 Å 2) × 2 = 6,280,000 Å2 of phospholipid. DOPC has been calculated to have a surface area of 59 Å2 per molecule. To simplify these calculations, we assumed that DOPS has the same surface area as DOPC. Using these assumptions, it calculated that there are 106,440 molecules of phospholipid per liposome. With our estimated incorporation ratio of 1,000:1 for phospholipid to COX per liposome, this would predict 106 molecules of COX per liposome, close to the 123 average spots observed.
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4. Notes 1. COX-2 is expressed in significantly higher level than COX-1 using the baculovirus system. 2. The buffer is exchanged to a nonphosphate-based buffer. The concentration of lipid in proteoliposomes is measured by phosphorous assay and therefore the buffer solution should be phosphate-free. 3. Liposomes should be extruded at a temperature that is above the liquid-to-gel phase transition temperature (Tc) of the lipid mixture. Attempts to extrude below the Tc will be unsuccessful, as the membrane has a tendency to foul with rigid membranes that cannot pass through the pores. 4. H2O2 is added in order to bleach the brown solution. If any brown color persists, 50 µl H2O2 is added to all cooled tubes and the tubes are heated again for 15 min until they become colorless. 5. During the purification process, the heme is stripped off from COX enzyme. Therefore, heme should be added to the enzyme before assaying the enzyme. 6. Transmission electron microscopy on gold-labeled liposome is carried out using a diluted stain (0.1–0.2 % solution of uranyl acetate). Because the NANOGOLD particles are small, overstaining with uranyl acetate may tend to obscure direct visualization of individual NANOGOLD particles. Therefore a diluted stain is used.
Acknowledgments The authors would like to thank Dr. Alicia Pastor and Mr. Robert Pcionek from Michigan State University Center for Advanced Microscopy for their help with the electron microscopy work.
References 1. Picot D, Loll PJ, Garavito RM (1994) The X-ray crystal structure of the membrane protein prostaglandin H2 synthase-1. Nature 367: 243–249 2. Jain MK, Zakim D (1987) The spontaneous incorporation of proteins into preformed bilayers. Biochim Biophys Acta 906: 33–67
3. Smith T, Leipprandt J, DeWitt D (2000) Purification and characterization of the human recombinant histidine-tagged prostaglandin endo peroxide H synthases-1 and -2. Arch Biochem Biophys 375:195–200 4. Mirafzali Z, Leipprandt J, DeWitt D (2005) Fast, efficient reconstitution of cyclooxygenases into proteoliposomes. Arch Biochem Biophys 443:60–65
Chapter 8 The Reconstitution of Actin Polymerization on Liposomes Mark Stamnes and Weidong Xu Abstract Membrane-associated actin polymerization is of considerable interest due to its role in cell migration and the motility of intracellular organelles. Intensive research efforts are underway to investigate the physiological role of membrane-associated actin as well as the regulation and mechanics of actin assembly. Branched actin polymerization on membranes is catalyzed by the Arp2/3 complex. Signaling events leading to the activation of the guanosine triphosphate (GTP)-binding protein Cdc42 stimulate Arp2/3dependent actin polymerization. We have studied the role of Cdc42 at the Golgi apparatus in part by reconstituting actin polymerization on isolated Golgi membranes and on liposomes. In this manner, we showed that cytosolic proteins are sufficient for actin assembly on a phospholipid bilayer. Here we describe methods for the cell-free reconstitution of membrane-associated actin polymerization using liposomes and brain cytosol. Key words: Liposome, Actin, Arp2/3, Wiscott–Aldrich syndrome protein (WASP), Cdc42
1. Introduction The actin cytoskeleton plays two basic roles connected to cell migration and the motility of intracellular organelles. First, actin microfilaments serve as tracks for myosin-motor-based motility. The best characterized role for an actomyosin complex is in muscle contraction (1). Myosin-based movement along actin is also utilized for vesicle targeting, organelle positioning, and the retraction of the cell’s trailing edge during migration (2). A second role for actin is to provide a propulsive force directly through Arp2/3catalyzed polymerization at a membrane (3). The most dramatic examples of this are the comet-tail type of motility used by endosomes that is also usurped by pathogenic bacteria such as Listeria. Similarly, actin polymerization is used directly to force the extension
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_8, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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of the leading edge membrane during cell migration. The ability to utilize actin polymerization to direct motility requires precise temporal and spatial regulation. A key regulatory mechanism for Arp2/3-dependent actin polymerization involves Cdc42, a member of the Rho-family of guanosine triphosphate (GTP)-binding proteins, and its effectors N-WASP or WAVE (4). GTP-bound Cdc42 causes a conformation change in N-WASP that exposes a C-terminal Arp2/3-binding domain. The actin polymerization activity of Arp2/3 is stimulated when bound to N-WASP. Thus, Cdc42 activation can stimulate actin polymerization at a distinct time and place. In many cell types, Cdc42 is localized to the Golgi apparatus through a binding interaction with the vesicle-coat protein, coatomer (5). There, Cdc42 serves to regulate both actin and microtubule-dependent trafficking events. We have investigated the role of Cdc42 at the Golgi complex, in part through the cell-free reconstitution of Cdc42-regulated Arp2/3dependent actin polymerization on isolated membranes and liposomes (6–10). The ability to reconstitute actin polymerization in vitro has served as a powerful tool for dissecting the mechanisms and regulation of this process at the molecular level. Indeed, the primary catalyst of actin polymerization, the Arp2/3 complex, was discovered through a biochemical purification using cell-free actin polymerization as an assay (11). The reconstitution of actin polymerization on biological and liposomal membranes has revealed details about the regulation and role of actin in the secretory pathway (5). It was recently demonstrated that the vesicle-associated GTP-binding protein ARF1 causes comet-tail-like motility of liposomes (12). We have exploited ARF1 and Cdc42 dependent actin polymerization on liposomes to demonstrate that this process involves the recruitment of Arp2/3 to the membrane (8). Here we provide detailed methods for reconstituting actin polymerization on membranes using brain cytosol and liposomes.
2. Materials 2.1. Preparation of Bovine-Brain Cytosol
1. Brain buffer: 25 mM Tris-HCl, pH 7.4, 320 mM sucrose. 2. Protease inhibitor stock solutions: 10 mg/ml aprotinin in H2O, 10 mg/ml leupeptin in H2O, 1 mM pepstatin A in dimethyl sulfoxide, 50 mM 1,10-phenanthroline in H2O, pH 5.0, 100 mM phenylmethanesulfonyl fluoride (PMSF) in 2-propanol.
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3. Breaking buffer: 25 mM Tris-HCL, pH 8.0, 500 mM KCl, 250 mM sucrose, 2 mM EGTA. Add dithiothreitol (DTT) and protease inhibitors (from the stock solutions) immediately before use to the following final concentrations, 1 mM DTT, 2 mg/ml aprotinin, 0.5 mg/ml leupeptin, 2 mM pepstatin A, 500 mM 1,10-phenanthroline, 1 mM PMSF. 4. Dialysis buffer: 25 mM Tris-HCl, pH 8.0, 50 mM KCl, add 1 mM DTT just prior to use. (see Note 1). 5. Dialysis tubing with 12,00–14,000 molecular weight cut off and 2.8 cm diameter (Membra-Cel®). 2.2. Preparation of Liposomal Membranes
1. Phospholipids: 10 mg/ml phospholipid stock solutions are prepared in chloroform. A bovine liver lipid extract (Avanti Polar Lipids) is an inexpensive source to provide a complex phospholipid bilayer. (see Note 2). 2. Resuspension buffer: 20 mM HEPES, pH 7.2, 150 mM potassium acetate, 250 mM sucrose.
2.3. Actin Polymerization Reactions
1. 5× Reaction buffer: 125 mM HEPES, pH 7.2, 12.5 mM magnesium acetate, 75 mM potassium chloride. 2. 100× ATP regenerating stock solution: 100 mM creatine phosphate, 25 mM UTP, 5 mM ATP. 3. Creatine phosphokinase stock solution: 1,000 units/ml in H 2O. 4. GTPgS solution: 10 mM in H2O. 5. 45% Sucrose/Tris (ST) solution: 10 mM Tris-HCl, pH 7.4, 45% sucrose (weight/weight, i.e. 45 g sucrose plus 55 ml water). 6. 35% ST solution: 10 mM Tris-HCl, pH 7.4, 35% sucrose (weight/weight). 7. 15% ST solution: 10 mM Tris-HCl, pH 7.4, 15% sucrose (weight/weight). 8. TCA stock solution: 100% trichloroacetic acid (weight/volume) in water.
2.4. Characterizing Actin Polymerization by SDS-PAGE
1. Laemmli sample buffer: 100 mM Tris-HCl, pH 6.8, 3% (weight/volume) sodium dodecyl sulfate (SDS), 10% (volume/volume) glycerol, 715 mM-mercaptoethanol, 0.03% (weight/volume) bromophenol blue.
2.5. Characterizing Actin Polymerization by Western Blotting
1. TBS-Tween solution: 20 mM Tris-HCl, pH 7.6, 137 mM NaCl, 0.5% Tween-20 (volume/volume). 2. Blocking solution: 5% non-fat milk powder (weight/volume) in TBS-Tween solution.
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3. Methods The general procedure for the reconstitution of actin polymerization involves incubating the membranes with cytosol under conditions where GTP-binding proteins are active. This may be done by including the non-hydrolyzable GTP analog GTPgS in the reaction. Following the incubation, the liposomes are isolated from the reaction by centrifugation and analyzed by SDS-PAGE and Western blotting. A key to these experiments is obtaining good-quality cytosol to provide a source of actin, Arp2/3, and regulatory proteins. 3.1. Preparation of Bovine-Brain Cytosol
1. Obtain fresh bovine brains from a slaughter house. Two to three brains provide a 100–300 ml of cytosol. The brains should be placed in ice-cold brain buffer as soon as possible and stored in the buffer on ice while in transit. All of the following steps should be performed on ice or in a 4°C cold room. 2. Remove and discard the brain stem and cerebellum. Carefully peel the meninges and blood vessels from the cerebrum using forceps. Weigh the remaining cerebral tissue. 3. Chop the cerebrum into approximately 1 cm pieces using scissors. Immediately place the brain tissue into a small volume of breaking buffer (with protease inhibitors). Once the tissue is fully chopped, add additional breaking buffer such that the final volume of brain tissue plus buffer equals 1.25 L per 500 g tissue. 4. The tissue is homogenized using a blender. Blend the sample 2 times at a high setting for 30 s each. 5. Pour the homogenate into 500 ml centrifuge bottles. Centrifuge for 1 h at 14,000× g in a Sorvall SLA-3000 rotor (Thermo Scientific) or equivalent. 6. Decant the supernatant and pour into polycarbonate or quickseal tubes for Type 45 Ti rotor (Beckman-Coulter). Spin for 90 min at 140,000× g. 7. Collect the supernatant and distribute to several 1 m sections of dialysis tubing. Seal the tubing by tying knots or using clips. Be certain to leave extra space in the tubing as the retentate volume will expand considerably during the dialysis. Dialyze the sample for 4 h in 30 L of dialysis buffer. Switch the tubing to fresh dialysis buffer and dialyze for an additional 4–8 h (see Note 3). 8. Record the volume of the retentate and spin the sample for 90 min at 140,000× g using the Type 45 Ti rotor. Collect the supernatant. 9. The cytosol is concentrated by precipitation with ammonium sulfate. Slowly add 36.5 g of ammonium sulfate per 100 ml
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cytosol while stirring to obtain a final concentration of 60% saturation. Allow the sample to stir for an additional 30 min. 10. Recover the precipitate by centrifugation in 500 ml bottles in the SLA-3000 rotor 30 min. at 14,000× g. Discard the supernatant. Resuspend the pellet in dialysis buffer using one-sixth the volume recorded in step 8 (see Note 4). 11. Dialyze the sample against 30 L dialysis buffer for 4 h. Transfer the dialysis tubing to 30 L fresh dialysis buffer without DTT for an additional 4–8 h (see Note 3). 12. Collect the retentate and centrifuge for 90 min at 140,000× g using the Type 45Ti rotor. Recover the supernatant. 13. Mix the sample and determine the protein concentration of the cytosol using bicinchoninic acid (BCA) protein assay kit (Pierce). A typical preparation has a concentration of ~10 mg/ml. 14. Freeze the cytosol in aliquots using liquid nitrogen. Store the cytosol at −80°C. 3.2. Preparation of Liposomal Membranes
1. Dispense the appropriate volumes of the phospholipid stock solutions into the bottom of a glass round-bottom tube (see Note 5). 2. Evaporate the chloroform by blowing air across the tube or using an Evap-O-Rac (Cole-Parmer). 3. The phospholipids are hydrated by adding resuspension buffer and mixing using a vortex mixer (see Note 6). The final phospholipid concentration should be 2.3 mg/ml. 4. The lipid resuspension is subjected to 10 freeze-thaw cycles using a dry-ice/methanol bath. 5. Unilamellar vesicles with a uniform size can be generated by extrusion through a polycarbonate membrane. Nine passages through a LiposoFast membrane extrusion device (Avestin) containing a membrane with pore diameter of 400 nm works well. 6. The liposomes are best if used immediately, but we have successfully stored them as aliquots at −80°C for several months.
3.3. Actin Polymerization Reactions
1. The reactions are set up in 1.5 ml plastic tubes with a final volume of 200 ml (see Note 7). The final reaction conditions are as follows: 1× reaction buffer, 1× ATP regenerating solution, 8 units/ml creatine phosphokinase, 0.2 mg/ml bovine brain cytosol, 0.23 mg/ml liposomes. Actin polymerization may be stimulated by adding 20 mM GTPgS to activate GTP-binding proteins. 2. Incubate the reactions at 37°C for 20 min. 3. At the end of the incubation, isolate the liposomes by microcentrifugation for 30 min at 15,000 × g.
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4. The liposomes may be separated from non-membrane-associated F-actin by flotation through a sucrose cushion. Begin by resuspending the liposome pellet in 50 ml of 45% ST solution (see Note 7). The sample is placed at the bottom of a 7 × 20 pollyallomer ultracentrifugation tube (Beckman-Coulter). The sample is overlayed with 130 ml 35% ST solution followed by 20 ml 15% ST solution. The sample is subjected to centrifugation for 45 min at 386,000× g using a TLA-100 rotor (Beckman Coulter). 5. After centrifugation, the sample tubes are frozen by careful immersion in liquid nitrogen. Slice the tube with a guillotine or heated razor blade to recover the top 100 ml of the gradient. 6. Precipitate the liposome-associated proteins by addition of trichloroacetic acid to a final concentration of 10%. Recover the precipitate by spinning in a microcentrifuge for 15 min. 7. Wash the precipitate with ice-cold acetone. 3.4. Characterizing Actin Polymerization by SDS-PAGE
1. Resuspend the precipitate using Laemmli sample buffer. 2. Load sample onto a 12% SDS-PAGE Gel (BioRad) and install the gel into an electrophoresis apparatus containing running buffer (BioRad). 3. Carry out electrophoresis at 200 V until the bromophenol blue dye front reaches the bottom of the gel. 4. Liposome-bound proteins can be visualized using Coomassie blue or silver stain as shown in Fig. 1.
Fig. 1. Shown is a Coomassie-blue-stained SDS PAGE gel of liposomes isolated by flotation from incubations carried out with brain cytosol. When the non-hydrolyzable GTP analog GTPgS is included in the reaction, multiple proteins bind to the liposomes. Actin is observed as a prominent band at 42 kD. The actin plus end-binding toxin cytochalasin D partly inhibits actin polymerization. Brefeldin A (BFA), an inhibitor of the GTP-binding protein ARF1, is a potent inhbitor of actin assembly on the liposomes. A peptide p23 that blocks Cdc42 recruitment to the membrane also inhibits actin polymerization on liposomes
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Fig. 2. Shown is a Western blot probed with antibodies against actin and tubulin. Tubulin binding to the liposomes is not affected by the activation of GTP-binding proteins. By contrast, actin levels are greatly increased in the presence of GTPgS. Actin polymerization is inhibited by cytochalasin D (CytoD), brefeldin A (BFA), and the p23 peptide as described in the legend to Fig. 1
3.5. Characterizing Actin Polymerization by Western Blotting
1. Instead of staining the SDS gel, the presence or absence of specific proteins can be assessed by Western blot analysis. 2. Transfer proteins onto a nitrocellulose membrane using an electroblot apparatus containing transfer buffer (BioRad). 3. Block the nitrocellulose by incubating in blocking solution for 1 h at room temperature. 4. Wash the membrane two times with TBS–Tween solution. 5. To detect actin, incubate the membrane in rabbit anti-actin antibody (Sigma) diluted 1/500 in TBS–Tween for 1 h at room temperature (see Note 8). 6. Wash the membrane five times with TBS–Tween and incubate with peroxidase-conjugated anti-rabbit secondary antibodies (BioRad) diluted 1/4000 in TBS–Tween for 30 min. 7. Wash the membrane five times with TBS–Tween and incubate with a chemiluminescence reagent (Pierce). 8. Expose blot to film as shown in Fig. 2.
4. Notes 1. Chill approximately 100 L of water ahead of time. 50 L cylindrical tanks with lids (Nalgene) are convenient for chilling water and carrying out the dialyses. 2. Alternatively, synthetic phospholipids may be mixed to obtain a bilayer with a defined phospholipid composition. 3. The second dialysis tank can be reused for the first dialysis in Subheading 3.1, step 11. 4. The ammonium sulfate pellets may be dislodged from the centrifuge bottles with a pipette and easily resuspended using a 100-ml dounce homogenizer.
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5. A trace amount of fluorescent phospholipid may be added to the preparation so that the recovery of liposomes may be quantified using a fluorescence spectrometer. For this 10 mg/ml NBD-phosphatidylcholine (Avanti Polar Lipids) works well. 6. A bath sonicator may also be used to resuspend the phospholipids. 7. The reaction can easily be scaled up to characterize the properties of less abundant cytokeletal proteins. In this case, controls should be carried out to confirm that liposomes are successfully recovered from larger volume sucrose gradients. 8. Duplicate blots may be probed with other antibodies to characterize the properties of multiple cytoskeleton-related proteins.
Acknowledgments This work was supported by NIH grant RO1 GM068674 (M.S.). The methods for preparing cytosol and liposomes have been adapted from (13, 14). We thank Heidi Hehnly for reading the manuscript.
References 1. Cooke R (2004) The sliding filament model: 1972–2004. J Gen Physiol 123:643–656 2. Wu X, Jung G, Hammer JA III (2000) Functions of unconventional myosins. Curr Opin Cell Biol 12:42–51 3. Theriot JA (2000) The polymerization motor. Traffic 1:19–28 4. Ridley AJ (2006) Rho GTPases and actin dynamics in membrane protrusions and vesicle trafficking. Trends Cell Biol 16:522–529 5. Hehnly H, Stamnes M (2007) Regulating cytoskeleton-based vesicle motility. FEBS Lett 581:2112–2118 6. Fucini RV, Chen JL, Sharma C, Kessels MM, Stamnes M (2002) Golgi vesicle proteins are linked to the assembly of an actin complex defined by mAbp1. Mol Biol Cell 13:621–631 7. Chen JL, Fucini RV, Lacomis L, ErdjumentBromage H, Tempst P, Stamnes M (2005) Coatomer-bound Cdc42 regulates dynein recruitment to COPI vesicles. J Cell Biol 169:383–389
8. Chen JL, Lacomis L, Erdjument-Bromage H, Tempst P, Stamnes M (2004) Cytosol-derived proteins are sufficient for Arp2/3 recruitment and ARF/coatomer-dependent actin polymerization on golgi membranes. FEBS Lett 566:281–286 9. Fucini RV, Navarrete A, Vadakkan C, Lacomis L, Erdjument-Bromage H, Tempst P, Stamnes M (2000) Activated ADP-ribosylation factor assembles distinct pools of actin on golgi membranes. J Biol Chem 275:18824–18829 10. Xu W, Stamnes M (2006) The actin-depolymerizing factor homology and charged/ helical domains of drebrin and mAbp1 direct membrane binding and localization via distinct interactions with actin. J Biol Chem 281: 11826–11833 11. Welch MD, Iwamatsu A, Mitchison TJ (1997) Actin polymerization is induced by Arp2/3 protein complex at the surface of Listeria monocytogenes. Nature 385: 265–269
Actin Polymerization on Liposomes 12. Heuvingh J, Franco M, Chavrier P, Sykes C (2007) ARF1-mediated actin polymerization produces movement of artificial vesicles. Proc Natl Acad Sci U S A 104:16928–16933 13. MacDonald RC, MacDonald RI, Menco BP, Takeshita K, Subbarao NK, Hu LR (1991) Smallvolume extrusion apparatus for preparation of
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large, unilamellar vesicles. Biochim Biophys Acta 1061:297–303 14. Wattenberg BW, Rothman JE (1986) Multiple cytosolic components promote intra-Golgi protein transport. Resolution of a protein acting at a late stage, prior to membrane fusion. J Biol Chem 261:2208–2213
Chapter 9 Electroformation of Giant Unilamellar Vesicles from Native Membranes and Organic Lipid Mixtures for the Study of Lipid Domains under Physiological Ionic-Strength Conditions L.-Ruth Montes, Hasna Ahyayauch, Maitane Ibarguren, Jesus Sot, Alicia Alonso, Luis A. Bagatolli, and Felix M. Goñi Abstract Giant unilamellar vesicles (GUVs) constitute a cell-sized model membrane system that allows direct visualization of particular membrane-related phenomena, such as domain formation, at the level of single vesicles using fluorescence microscopy-related techniques. Currently available protocols for the preparation of GUVs work only at very low salt concentrations, thus precluding experimentation under physiological conditions. In addition, the GUVs thus obtained lack membrane compositional asymmetry. Here we show how to prepare GUVs using a new protocol based on the electroformation method either from native membranes or organic lipid mixtures at physiological ionic strength. Additionally, we describe methods to test whether membrane proteins and glycosphingolipids preserve their natural orientation after electroformation of GUVs composed of native membranes. Key words: Giant unilamellar vesicles (GUVs), Electroformation, Physiological conditions, Biological membranes, Lipid domains
1. Introduction Extensive documentation has appeared in recent years describing the use of giant unilamellar vesicles (GUVs, 20 µm mean diameter) as model systems to study different physical aspects of membranes (lateral structure, mechanical properties), particularly considering the effect of not only lipid–lipid but also lipid–DNA, lipid–peptide, and lipid–protein interactions (1–6). Owing to their size, single vesicles can be directly observed using light microscopy techniques (1, 2). V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_9, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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However, two main drawbacks regarding GUV preparation are still unsolved: (1) low salt concentration (<10 mM NaCl) must be used during the preparation process (higher ionic strengths impair GUV formation) (7) and (2) GUVs lack membrane asymmetry. Interesting methodological advances include the possibility of preparing GUVs at high salt concentrations, provided that negatively charged lipids are present in high proportions in the bilayer composition (8). Also, Baumgart et al. generated giant plasma membrane vesicles by inducing plasma membrane vesiculation or “blebbing” (9). A protocol was also published for the generation of asymmetric GUVs, but in the absence of proteins (10). Recently, on the basis of a protocol described by Pott et al. (11), a method for the electroformation of GUVs (Fig. 1) from either cell membranes (Fig. 2) or lipid extracts (Fig. 3) under physiological ionic-strength conditions was published by our laboratories (12). When the starting materials are red blood cell (RBC) membranes, the resulting giant vesicles keep the membrane asymmetry. The electroformation method and ancillary techniques are described in detail below.
2. Materials 2.1. Preparation of Human Erythrocyte Ghosts
1. Assay buffer: In all experiments, the assay buffer consists of 25 mM HEPES, 150 mM NaCl, pH 7.2. 2. Right-side out (RSO) ghosts hemolysing nonsaline buffer (A): 1.2 mM acetic acid, 4 mM MgSO4, pH 3.2. 3. Sucrose buffer (B): 43% sucrose in assay buffer. Keep at 4°C in the refrigerator. 4. Inside out (IO) ghosts hemolysing solution (C): 5 mM KH2PO4, pH 8.0. 5. IO ghosts membrane resealing buffer (D): 0.5 mM Tris– HCl, 1 mM EDTA, 2 mM DTT, pH 8.5. 6. Glucose buffer (E): 1.03 g/ml glucose in assay buffer (d = 1.03). 7. Buffer F: 10 mM Tris–HCl, 1 mM ehtylenediamide tetraacetic acid (EDTA) pH 7.8. 8. Buffer G: 20 mM HEPES, 0.5 mM dithiothreitol (DTT), 1 mM MgCl2, 150 mM KCl, pH 7.2. 9. Molibdate solution (H): 22 mg (NH4)6Mo7O24⋅4H2O, 143 ml H2SO4 in 1 L dH2O. 10. EDTA anticoagulating tubes for human blood collection (BD Vacutainer Systems, NJ, USA). 11. Three 23-gauge microlance needles (BD Biosciences, NJ, USA).
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12. Cholesterol concentration oxidase/peroxidase measurement kit (BioSystems, Barcelona, Spain). 2.2. Preparation of GUVs
1. DiIC18 stock solution: 1.25 mM DiIC18 in dimethyl sulfoxide (DMSO). Protect from light and store in the refrigerator. 2. Stocks of lipid mixtures in organic solvent: 0.2 mg/ml stock of the desired composition in organic solvent (generally chloroform:methanol [2:1] [vol:vol]) supplemented with 0.2 mol% of DiIC18. Keep in the freezer at −20°C. 3. Electroformation chamber that allows visualization under the microscope (homemade, see Fig. 1).
Fig. 1. Homemade GUV electroformation chamber. The Pt electrodes can be extracted from the chamber to allow sample addition and vacuum treatment. A cover glass allows direct visualization of GUVs under the microscope. The temperature is maintained constant by using a water bath
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4. 24 × 50 mm cover glasses (Menzel-Gläser, Braunschweig, Germany) to couple at the bottom of the visualization chamber. 5. TG330 function generator from Instruments (Cambridgeshire, UK).
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6. F12 water bath (Julabo, Seelbach, Germany) for adjusting vesicle formation temperature. 2.3. Immunostaining of RBC ghost membranes (Membrane Asymmetry Assays)
1. Primary antibodies: Mouse IgG antihuman band III (Sigma, MO, USA). Goat IgG antihuman glycophorin A and blood group A1, A2, A3 antigen mouse monoclonal IgM (Santa Cruz Biotechnologies, CA, USA). 2. Secondary antibodies: Alexa fluor 488 donkey anti-goat IgG and Alexa fluor 633 rabbit anti-mouse IgG (Invitrogen, OR, USA). 3. Alexa fluor 633 protein labeling kit (Invitrogen).
2.4. Fluorescence Microscopy
1. Zeiss-LSM 510 META NLO inverted confocal excitation fluorescence microscope (Carl Zeiss, Jena, Germany). 2. Multitrack mode software (Carl Zeiss). 3. C-Apochromat objective 40×, water immersion, NA 1.2 (Carl Zeiss).
3. Methods 3.1. Preparation of Human Erythrocyte Ghosts
Blood (group A) is collected from healthy donors, placed in EDTA anticoagulating tubes, and washed three times (or until supernatant is clear) with assay buffer by centrifugation at 1,250 × g for 10 min at 4°C. The supernatant is discarded and the erythrocyte-containing pellet is used to form ghosts. Around 5 ml of fresh blood should be enough to obtain an adequate final ghost concentration (see Note 1).
3.1.1. Preparation of RSO Ghosts
RSO ghosts are prepared following the method of Steck and Kant (13), with some modifications: 1. Osmotic shock: In order to produce hemolysis, add 10 volumes of non-saline buffer (A) to the packed RBCs. Mix gently and incubate for 15 min, 4°C, under smooth agitation. After centrifugation at 31,000 × g for 15 min at 4°C, remove the supernatant and wash the pellet two more times (or until the supernatant is clear) in the same buffer to obtain the ghost membranes. 2. Membrane resealing: Add 10 ml isoosmotic assay buffer to the precipitate and incubate for 1 h at 37°C. Centrifuge at 14,000 × g for 10 min and repeat 2–3 washes. Bring the
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resulting membrane suspension to a final volume of around 1.5 ml in assay buffer. 3. Sealed membrane separation by sucrose gradient centrifugation: To separate the fractions (sealed and non-sealed membranes), layer 1.5 ml of membranes over 6 ml sucrose buffer (B) in a centrifuge tube. Centrifuge using a swing-out rotor at 138,000 × g for 60 min at 4°C. Recover the band containing the resealed vesicles (RSO ghosts), which floats on top of the sucrose solution. Harvest and wash the resealed ghosts three more times in assay buffer at 14,000 × g for 10 min. Resuspend in around 1 ml assay buffer. 3.1.2. Preparation of Inside Out Ghosts
IO ghosts preparation is based on the method by Schow and Passow (14) with some modifications: 1. Osmotic shock: Dilute the washed erythrocytes in 20 volumes of hemolysing solution (C). Incubate at room temperature for 5 min to break down the cells, and pellet by centrifugation at 35,000 × g for 20 min. Repeat this wash twice with the same buffer. 2. Membrane resealing: Resuspend hemoglobin-free membranes in 10 ml of membrane resealing buffer (D). Incubate for 45 min at 4°C and for 45 min at 37°C to allow membrane resealing. Centrifuge at 28,000 × g for 30 min, and wash the pellet three times with the same buffer. After the last wash, bring the sample to a final volume of 2 ml and homogenate with a 23-gauge needle for 10–15 times. 3. Sealed membrane separation by glucose barrier centrifugation: To purify the resealed IO ghosts, layer 2 ml of membranes over 6 ml glucose buffer (E) and centrifuge at 100,000 × g for 2 h at 4°C. Collect the vesicles remaining above the barrier, wash once in 20 ml buffer F for 30 min at 40,000 × g, and resuspend in around 1 ml buffer G.
3.1.3. Determination of Lipid Concentration
In order to form GUVs from erythrocyte membranes, ghost suspensions are brought to a final lipid concentration (cholesterol + phospholipids) of ≅2.5 mM (~1 mM final concentration of cholesterol can be measured using a cholesterol concentration measurement kit). Phospholipid concentration of ghosts is determined by measuring the phosphorous content (15, 16) of lipids extracted from the ghost membranes (17).
3.1.3.1. Ghost Lipid Extraction
1. Precipitation of lipoproteic complexes: Take 250 µl of the ghost suspension and add the same volume of cold 0.6 M perchloric acid in 10-ml glass tubes. Centrifuge at 14,000 × g for 15 min, and discard the supernatant. 2. Lipid extraction: Resuspend the precipitate in 2.5 ml of cold chloroform:methanol (2:1, [vol:vol]) and incubate at room temperature for 30 min mixing every 5 min. Add 5 ml of cold
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0.1N HCl, vortex, and centrifuge at 1,700 × g for 20 min. Discard the upper phase and recover the chloroformic lipidcontaining lower phase. Repeat the extraction on the intermediate phase and mix the two chloroformic fractions together. 3.1.3.2. Phospholipid Concentration Determination
1. Prepare the calibration curve from 0 to 25 nmol of NaH2PO4 standard from a 1 mM stock. 2. Add 100 µl of 70% perchloric acid to 200 µl of sample/calibration curve, and let digestion happen at 200°C for 45 min. Mix with 500 µl distilled H2O, 100 µl molibdate solution (H), and 100 µl 10% ascorbic acid. The colorimetric reaction takes place at 100°C in 5–10 min, after which absorbance can be measured in a colorimeter at 820 nm. 3. Calculate ghost phospholipid concentration:
3.2. Preparation of GUVs from Erythrocyte Membranes (G-ghosts) and from Organic Lipid Mixtures under Physiological Conditions 3.2.1. Formation of G-ghosts
[P] × total volume 200 where [P] is the concentration calculated by this protocol and total volume is the volume of the chloroformic phase obtained after lipid extraction. [Ghost phospholipids] =
In order to be able to visualize the GUVs under the microscope, add 1 mol% DiIC18 (from the stock) with respect to total lipids to the ghost solutions before the formation. Deposit a small aliquot (~1 µl) of erythrocyte ghost suspension (RSO or IO) onto each Pt electrode in a special homemade chamber, which allows direct visualization under the microscope (Fig. 1) (18). Cover the sample to avoid light exposure and allow precipitation onto the Pt wires for 5 min at room temperature (see Notes 2 and 3).
3.2.2. Formation of GUVs Composed of Organic Lipid Mixtures
Add 2.5 µl of the appropriate lipid stock on the surface of each Pt electrode and let the solvent traces evaporate by placing the electrodes under high vacuum for at least 2 h.
3.2.3. Electroformation under Physiological Conditions
Place the electrodes on the chamber and add 500 µl of isoosmotic assay buffer. Apply an AC field using a function generator in three main steps, usually at 37°C: 1. Frequency 500 Hz, amplitude 106 mV (35 V/m) for 5 min, 2. Frequency 500 Hz, amplitude 940 mV (313 V/m) for 20 min, 3. Frequency 500 Hz, amplitude 2.61 V (870 V/m) for 90 min. The temperature used for GUV formation should ensure that the membrane under study displays a single fluid phase (above the highest phase transition temperature). Due to the design of the chamber used in vesicle formation (Fig. 1), GUVs can be directly observed under the microscope, avoiding transfer of vesicles to conventional chambered cover glasses (see Notes 4 and 5).
Electroformation of Giant Unilamellar Vesicles
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The extracellular domain of glycophorin A (outer hemimembrane label) is identified with a primary goat IgG antibody. The bound antibody is detected by a secondary Alexa fluor 488-labeled donkey anti-goat IgG antibody. The cytoplasmic domain of band III (inner hemimembrane label) is recognized with a primary mouse IgG antibody and a secondary Alexa fluor 633-labeled rabbit anti-mouse IgG antibody (19). Blood group A antigen mouse monoclonal IgM was labeled with the Alexa fluor 633 protein labeling kit (Fig. 2). 1. Add each primary antibody at a final concentration of 8 µg/ml. 2. Incubate for 30 min at room temperature to allow binding of the antibodies to their specific antigen. 3. Wash three times with physiological buffer (the same buffer used during electroformation). This will reduce considerably the background fluorescent signal. 4. When necessary, add the secondary antibodies, incubate, and wash under the same conditions. Addition of primary and secondary antibodies in two steps avoids agglutination of the immunoglobulins (see Notes 6 and 7).
3.4. Fluorescence Microscopy
An inverted confocal excitation fluorescence microscope is used to visualize giant vesicles in this kind of experiments (see Note 8). Excitation wavelengths: 488, 543, and 633 nm (for Alexa 488, DiIC18, and Alexa 633 labeled antibodies). Filters: The fluorescence signals for Alexa 488, DiIC18, and Alexa 633 can be simultaneously collected using multitrack mode into three (or two, depending on the experiment) different channels using bandpass filters of 520 ± 10, 590 ± 25, and 670 ± 25 nm, respectively. Objective: Use a 40× water immersion, NA 1.2 objective.
4. Notes 1. Sometimes, RBCs break up during the centrifugation washes. This is detectable because the hemoglobin-containing supernatant becomes red, which can be avoided by decreasing the centrifugation force (g) and increasing the washing time. 2. The final volume of DMSO added to a membrane-containing sample must always be less than 2% of the final total volume in order to preserve membrane integrity. Stocks should be concentrated enough to obey this proportion. 3. Whenever working with fluorescent probes, it is convenient to cover and protect the samples from light, and to place the stocks in amber containers in order to preserve the fluorescent properties of the probes.
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4. It is important to control temperature conditions in these experiments: electroformation of GUVs must take place above the gel–fluid transition temperature of the membrane under study. Take also into account that lipid domain formation depends on the temperature at which GUVs are visualized. 5. If electroformation takes place at a high temperature, buffer might evaporate while the experiment is running. Covering the chamber with parafilm might reduce evaporation to keep the volume of the buffer constant. 6. There is a vast variety of fluorescent probes (with different partition properties than DiIC18) which allow visualization of membrane domains from different natures (see e.g., ref. (20)). There are also membrane lipid-based probes available (e.g., NBD-SM) for the study of domain formation due to specific lipid degradation (e.g., by a sphingomyelinase) (21). Take into account the dynamic properties and photobleaching susceptibility when selecting a probe for confocal fluorescence experiments. 7. In this chapter we show how to form giant vesicles from erythrocyte membranes and from some lipidic mixtures of our interest in the study of lipid domains (Figs. 2 and 3). Nevertheless, other lipid mixtures, cell types, or fractionated organelles might be susceptible to GUV formation. Their visualization and study of interaction with other molecules (such as DNA, proteins, lipids) should then be possible. The immunostaining of specific proteins present in each biological membrane can help to assess whether membrane integrity is maintained after electroformation. The experiments shown here seem to demonstrate that electroformation does not affect the presence and orientation of ghost membrane proteins, but similar experiments could be performed on membranes from different nature. 8. Here we electroform GUVs using a high-ionic strength buffer to reproduce physiological conditions. The chamber used allows direct visualization of GUVs, but if this technology is not available, the buffers could be supplemented with glucose/sucrose to facilitate vesicle sedimentation to the bottom of chambered cover glasses (18).
Fig. 3. (continued) low-fluorescence-intensity regions correspond to liquid-ordered lipid domains enriched in SM and Chol. (d) GUVs composed of PI (phosphatidylinositol):SM (1:1) (mol:mol) supplemented with DG (diglyceride); the dark areas represent gel-like SM-enriched domains. (e) GUVs composed of PI:GalCer (1:1) (mol:mol); the low-fluorescenceintensity regions correspond to gel domains enriched in GalCer. (f) GUVs composed of PI:DSPC (distearoyl phoshatidylcholine):DG (1:1:0.3) (mol:mol:mol:mol); the dark areas represent gel-like DSPC-enriched domains. All the images were obtained in assay buffer (i.e., under physiological conditions). The white bars correspond to 10 mm
Fig. 2. (a) Multicolor fluorescence image of RSO G-ghosts in the presence of band III (red, A2) and glycophorin A (magenta, A3) specific immunofluorescence markers (false color representation); the G-ghosts are labeled with DiIC18 lipophilic fluorescent probe (yellow, A1). (b) Multicolor fluorescence image of IO G-ghosts in the presence of band III (red, B2) and glycophorin A (magenta, B3) specific markers. As shown in A, the G-ghosts are labeled with DiIC18 (yellow, B1). (c) and (d) Fluorescence intensity representative images of RSO (c) and IO (d) G-ghosts in the presence of blood group A specific marker (blue); the G-ghosts are labeled with DiIC18 (green). All the images have been obtained in assay buffer (i.e., under physiological conditions). The white bars correspond to 20 mm. From ref. (12), with permission
Fig. 3. Fluorescence images (false color representation) of lipid domains appearing in DiIC18-labeled GUVs. (a) GUVs composed of red blood cell membranes (G-ghosts) after sphingomyelinase addition; the brighter regions correspond to domains enriched in ceramide produced by enzyme activity. (b) GUVs composed of POPC (1-palmitoyl-2oleoyl phosphatidylcholine)/DPPC (dipalmitoyl phosphatidylcholine) (3:2) (mol:mol); the high-fluorescence-intensity areas correspond to DPPC-rich gel phase. (c) GUVs composed of PC:PE:SM:Chol (1:1:1:1) (mol:mol:mol:mol); the
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Acknowledgements This work was supported by FNU, Denmark (272-06-0511), the Danish National Research Foundation, the Spanish Ministerio de Educación y Ciencia (BFU 2005-0695 and BFU 2004-02955), and the University of the Basque Country (GIU06/42). References 1. Bagatolli LA (2006) To see or not to see: Lateral organization of biological membranes and fluorescence microscopy. Biochim Biophys Acta 1758:1541–1556 2. Menger FM, Keiper JS (1998) Chemistry and physics of giant vesicles as biomembrane models. Curr Opin Chem Biol 2:726–732 3. Veatch SL, Keller SL (2005) Seeing spots: Complex phase behavior in simple membranes. Biochim Biophys Acta 1746:172–185 4. Ambroggio EE, Separovic F, Bowie JH, Fidelio GD, Bagatolli LA (2005) Direct visualization of membrane leakage induced by the antibiotic peptides: Maculatin, citropin, and aurein. Biophys J 89:1874–1881 5. Bernardino de la Serna J, Perez-Gil J, Simonsen AC, Bagatolli LA (2004) Cholesterol rules: Direct observation of the coexistence of two fluid phases in native pulmonary surfactant membranes at physiological temperatures. J Biol Chem 279:40715–40722 6. Kahya N, Schwille P (2006) Fluorescence correlation studies of lipid domains in model membranes. Mol Membr Biol 23:29–39 7. Bagatolli LA, Parasassi T, Gratton E (2000) Giant phospholipid vesicles: Comparison among the whole lipid sample characteristics using different preparation methods: A two photon fluorescence microscopy study. Chem Phys Lipids 105:135–147 8. Akashi K, Miyata H, Itoh H, Kinosita K Jr (1996) Preparation of giant liposomes in physiological conditions and their characterization under an optical microscope. Biophys J 71:3242–3250 9. Baumgart T, Hammond AT, Sengupta PS, Hess T, Holowka DA, Baird BA, Webb WW (2007) Large scale fluid/fluid phase separation of proteins and lipids in giant plasma membrane vesicles. Proc Natl Acad Sci USA 104:3165–3170 10. Pautot S, Frisken BJ, Weitz DA (2003) Engineering asymmetric vesicles. Proc Natl Acad Sci USA 100:10718–10721 11. Pott T, Bouvrais H, Méléard P (2008) Giant unilamellar vesicle formation under
12.
13.
14. 15. 16. 17. 18.
19.
20.
21.
physiologically relevant conditions. Chem Phys Lip 154:115–119 Montes LR, Alonso A, Goñi FM, Bagatolli LA (2007) Giant unilamellar vesicles electroformed from native membranes and organic lipid mixtures under physiological conditions. Biophys J 10:3548–3554 Steck TL, Kant JA (1974) Preparation of impermeable ghosts and insideout vesicles from human erythrocyte membranes. Meth Enzymol 31:172–180 Schow G, Passow H (1973) Preparation and properties of human erythrocyte ghosts. Mol Cell Biochem 2:197–217 Fiske HC, Subbarow Y (1925) The colorimetric determination of phosphorus. J Biol Chem 66:375–400 Böttcher C, van Gent C, Fries C (1961) A rapid and sensitive sub-micro phosphorous determination. Anal Chim Acta 1061:297–303 Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37:911–917 Fidorra M, Duelund L, Leidy C, Simonsen AC, Bagatolli LA (2006) Absence of fluidordered/fluid-disordered phase coexistence in ceramide/POPC mixtures containing cholesterol. Biophys J 90:4437–4451 Kaufmann S, Tanaka M (2003) Cell adhesion onto highly curved surfaces: One-step immobilization of human erythrocyte membranes on silica beads. Chemphyschem 4:699–704 Sot J, Ibarguren M, Busto JV, Montes LR, Goñi FM, Alonso A (2008) Cholesterol displacement by ceramide in sphingomyelin-containing liquid-ordered domains, and generation of gel regions in giant lipidic vesicles. FEBS Lett 582:3230–3236 Montes LR, López DJ, Sot J, Bagatolli LA, Stonehouse MJ, Vasil ML, Wu BX, Hannun YA, Goñi FM, Alonso A (2008) Ceramideenriched membrane domains in red blood cells and the mechanism of sphingomyelinaseinduced hot-cold hemolysis. Biochemistry 47:11222–11230.
Chapter 10 Visualization of Lipid Domain-Specific Protein Sorting in Giant Unilamellar Vesicles Martin Stöckl, Jörg Nikolaus, and Andreas Herrmann Abstract Recent studies suggest that phospholipids in the plasma membrane of mammalian cells are not homogenously distributed but may form domains either by lipid–lipid interactions or/and as consequence of lipid– protein interactions. Such lipid compartments may act as protein recruiting platforms which, for example, are essential components of cell signaling pathways. Model membrane systems with a defined lipid composition are ideally suited to study domain-specific interactions of peripheral and integral membrane proteins. Giant unilamellar vesicles (GUVs) offer the opportunity to directly visualize in parallel, both the lateral lipid domains and the interaction sites of proteins using fluorescence microscopy. The application of GUVs is exemplarly illustrated for studying domain-specific interactions of the protein a-synuclein and the domain-specific distribution of synthetic transmembrane peptides. Key words: Giant unilamellar vesicles (GUVs), Fluorescence microscopy, Lipid domain, Raft, a-Synuclein, Transmembrane domain
1. Introduction In recent years, the relevance of lipid domains in plasma membranes of eukaryotic cells serving as a platform for recruitment and enrichment of specific integral and peripheral membrane proteins has become a challenging research focus in cell biology, biochemistry, and biophysics. Those domains may play an important role in cellular uptake mechanisms, signal transduction, and vesicle and virus budding (1–4). Domains are formed by lipid–lipid interactions, but can be also triggered by protein–lipid interactions. However, specific attention has been given to domains based on lipid–lipid interactions. Interaction between cholesterol and glycosphingolipids or saturated phospholipids, in particular phosphatidylcholine, is known to generate liquid ordered (lo) V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_10, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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domains clearly distinct from liquid-disordered (ld) domains (5, 6). The latter are essentially formed by unsaturated phospholipids and characterized by a reduced lipid packing density and membrane thickness with respect to lo domains. Both lipid packing and membrane thickness are important factors for specific redistribution of integral and peripheral proteins to distinct lipid domains. Of course, other factors are feasible. Enrichment of phospholipids with charged headgroups in distinct lipid domains can direct peripheral proteins to those domains via electrostatic interaction (7, 8). In native cells, it is difficult to visualize lipid domains and to study domain-specific enrichment of membrane proteins in the plasma membrane. Indeed, several attempts to image lo domains, the so-called rafts, in the plasma membrane of mammalian cells suggested that they are highly dynamic and very small, in the order of 10–100 nm (9). Giant Unilamellar Vesicles (GUVs) have become an attractive tool to study lipid domains. GUVs of appropriate lipid mixtures have been shown to form lo and ld domains with a size in the micrometer scale (5, 10, 11). These domains can be visualized by various lipid-like fluorophores which preferentially enrich either in the ld or in the lo domain. Such model systems are well suited to study the association of membrane proteins with specific lipid domains. For this purpose, the lateral distribution of fluorescently tagged proteins can be compared with the lipid domain-specific distribution of lipid-like fluorophores. For the protein a-synuclein, which is associated with the development of neurodegenerative diseases as Parkinson’s disease, it has been suggested that interaction with membranes may play a relevant role in pathogenesis. However, a controversy about the membrane binding site of a-synuclein exists (12, 13). An approach based on lipid domainforming GUVs allows direct visualization of potential binding sites of fluorescent-tagged proteins (8). Using the GUV model system, it is also possible to study the lateral distribution of transmembrane domains of integral membrane proteins. Exemplarily, the distribution of the so-called LV peptides, which were designed as a low-complexity model that mimics fusion protein transmembrane domains, will be illustrated here (14).
2. Materials 2.1. Preparation of GUVs on Indium Tin Oxide (ITO) Slides
1. ITO slides are glass slides (50 × 60 mm; 1.1 mm thickness) which are coated on one side with ITO (R £ 20 W/sq) from Präzisions Glas & Optik GmbH (Iserlohn, Germany). 2. Chloroform, methanol, and trifluoroethanol were of spectroscopic grade (Uvasol; Merck KGaA, Darmstadt, Germany).
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Fig. 1. Chamber design based on ITO-coated glass slides
3. Lipids and cholesterol were obtained from Avanti Polar Lipids (Alabaster, AL). 4. Polytetrafluoroethene (PTFE) spacers can be cut from several layers of PTFE film for surface protection. Alternatively, a silicone rubber spacer (thickness: 1 mm) can be used. For a sketch of the chamber assembly, see Fig. 1. PVC-based nonmelting plasticine (FimoTM; Eberhard Faber GmbH, Germany) was used for sealing chambers. 5. Swelling buffer: 250 mM sucrose and as a preservative 15 mM NaN3 in double deionized water. Measure osmolality of buffer (approx. 280 mOsm/kg). (see Note 1) Remove particles by means of filtration (0.22 µm pores). Store at 4°C until use. Note that NaN3 is highly toxic. 2.2. Preparation of GUVs on Titanium Slides
1. Titanium chambers (33 × 50 mm; 2 mm thickness) were made from pure titanium slides from Alfa Aesar GmbH (Karlsruhe, Germany). For a sketch of the chamber assembly see Fig. 2. 2. Parafilm is a product of Pechiney plastic packaging (Chicago, IL). 3. Swelling buffer: 250 mM sucrose and as a preservative 15 mM NaN3 in double deionized water. Measure osmolality of buffer (approx. 280 mOsm/kg) (see Note 1). Remove particles by means of filtration (0.22 µm pores). Store at 4°C until use. Note that NaN3 is highly toxic.
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Fig. 2. Chamber design based on titanium slides
4. PVC based non melting plasticine (FimoTM; Eberhard Faber GmbH, Germany) is used for sealing of filling holes. 2.3. Preparation of GUV Harboring Transmembrane Peptides
2.4. Membrane Binding of aSynuclein Evaluated by Laser Scanning Confocal Microscopy
1. Materials for GUV preparation on ITO or titanium slides correspond to the materials described in Subheading 2.1 (ITO slides) or Subheading 2.2 (titanium slides), respectively. 2. The peptide (amino acid sequence Rh-KKKKWLLVLLVLLVLLVLL VLKKKK-Rh) was synthesized by Boc chemistry (PSL, Heidelberg, Germany by courtesy of Prof. Dieter Langosch, TU Munich). Rhodamine-tagged peptides were made by coupling of a Lys derivative (Fmoc-Lys(Dde)-OH) to the C- and N-termini during synthesis. Reaction of the peptide with 5-(and-6)carboxytetramethylrhodamine succinimidylester (TMR-SE) yielded TMR-labeled peptides (see Notes 2 and 3). 1. Tetramethylrhodamine-6-maleimide was obtained from Molecular Probes (Eugene, OR). 2. Microscopy buffer: 280 mM glucose, 5.8 mM NaH2PO4, and 5.8 mM Na2HPO4 in double deionized water. Measure osmolality of buffer (approx. 300 mOsm/kg). If necessary, adjust by dilution with double deionized water. Remove particles by means of filtration (0.22 µm pores). Store at 4°C until use. 3. Glass coverslips, 24 × 60 mm (thickness: No. 1), were obtained from Carl Roth GmbH (Karlsruhe, Germany). Coverslips were used without prior cleaning.
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3. Methods Using appropriate lipid mixtures, GUVs with laterally segregated lipid domains are readily accessible by the electroformation technique using ITO-coated glass slides developed by Angelova and Dimitrov (15, 16). However, in a recent work it has been shown that the ITO may catalyze the oxidation of unsaturated lipids (17). Chambers made from titanium sheets can be used to circumvent that issue. Yet, no differences were found using the described lipid mixtures. For visualization of the domain structure, specific fluorescent lipid analogues such as C6-NBD-PC (1-palmitoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol4-yl)amino]hexanoyl]-sn-glycero-3-phosphocholine) are included in the lipid mixture. The ld domains in the vesicles show bright green fluorescence as C6-NBD-PC preferentially distributes to those domains, while the lo domains, from which C6-NBD-PC is excluded, appear almost completely dark in fluorescence microscopy. After incubation of a GUV suspension with the fluorescent-labeled proteins, protein–membrane interactions can be qualitatively and quantitatively investigated by measuring the fluorescence level at the vesicle membrane. For the investigation of transmembrane peptides, these hydrophobic peptides can be directly mixed to the lipid mixture before vesicle preparation. 3.1. Preparation of GUVs on ITO Slides
1. Clean two ITO slides (Fig. 1) in three successive steps by washing with detergent, deionized water, and finally ethanol. Rub down with lint-free paper. 2. Prepare lipid stocks by dissolving powdered lipids in chloroform and dilute to a suitable concentration (5 mM). If a turbid solution results (lipids are not completely dissolved), add methanol until the lipid stock is clear. To minimize solvent evaporation, stocks should be stored in tightly sealed glass vials at −20°C until use. 3. For GUV preparation, mix on ice 100 nmol of total lipids in a volume of 50 µl chloroform in a small glass vessel. For detection of domains by fluorescence microscopy, add 1 nmol of C6-NBD-PC. Keep the stocks on ice all the time. See Note 4 for lipid mixtures. 4. Place the slides on a heating stage and heat to approx. 50°C. Pipette 25 µl of the lipid mixture in small droplets on each of the conducting side of the slides. Let the solvent evaporate completely. Check the quality of spotted lipid films (see Note 5). 5. Place the spotted slides in an exsiccator and apply vacuum (<10 mbar) for at least 1 h to evaporate remaining traces of solvent.
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6. Assemble the GUV chamber consisting of the two lipidcoated ITO slides and an additional PTFE or silicone rubber spacer (1 mm thickness). The chamber is held together by two clamps. If PTFE spacers are used, the chamber has to be sealed with nonmelting modeling clay to prevent leakage. 7. Preheat the required amount of swelling buffer (approx. 1 ml per chamber) to 50°C. Fill the whole chamber with the heated buffer using a syringe and needle. Seal the chamber opening with modeling clay. 8. For electroformation of GUVs, heat the chamber to 50°C and apply a sinusoidal alternating voltage (10 Hz) which is stepwise increased from 0.02 to 1.1 V over 48 min and hold this voltage for at least 100 min. To detach the vesicles, incubate for further 30 min with an increased voltage of 1.3 V at a frequency of 4 Hz. 9. Remove the vesicle suspension from the chamber with a syringe and needle. GUVs are stable for a few days when stored at room temperature in the dark. 3.2. Preparation of GUVs on Titanium Slides
1. For cleaning of titanium chambers and preparation of lipid stocks see steps 1–3 of the protocol for GUV preparation on ITO slides. 2. To achieve an insulated layer between the bottom and top titanium slides, heat the bottom slide to approx. 50°C and surface-fuse one layer of Parafilm to the rim of the slide. Cut away excess Parafilm from the edges and the cavity of the slide with a razor blade. 3. Put the slides on a heating stage and heat to approx. 50°C. Pipette 25 µl of the lipid mixture in small droplets on each of the slides. Let the solvent evaporate completely. 4. Put the spotted slides in an exsiccator and apply vacuum (<10 mbar) for at least 1 h to evaporate remaining traces of solvent. 5. For chamber assembly, heat the bottom slide until the Parafilm layer melts and becomes sticky. Press on the top slide firmly to seal the chamber. After cooling to room temperature, insulation should be checked by means of measuring the resistance between the slides with a multimeter. The resistance should be >200 kW. 6. Preheat the required amount of swelling buffer (approx. 1 ml per chamber) to 50°C. Fill the whole chamber with the heated buffer with a pipette. Seal the filling holes with modeling clay. 7. Electroformation is equivalent to the procedure described for ITO slides. GUV suspension again is removed by pipetting.
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1. GUVs harboring transmembrane peptides can be generated using either ITO (see Subheading 3.1) or titanium slides (see Subheading 3.2) following the respective protocol. 2. For preparation of GUV, add 1 nmol of the peptide dissolved in trifluoroethanol and 1 nmol of C6-NBD-PC for the visualization of lipid domains to 100 nmol of total lipids including cholesterol in a total volume of 50 µl chloroform in a small glass vessel. 3. Continue GUV preparation by spotting small droplets of the lipid/peptide mixture according to step 4 (ITO) or step 3 (titanium) of the protocol, respectively (see Note 5). 4. To investigate the lateral distribution of the peptides, suspensions of GUVs containing peptide and C6-NBD-PC are added to microscopy buffer at a ratio of 1:1 to 1:3. Prior to microscopy, GUVs are given some minutes to settle down on a cover slip because of the greater specific density. 5. For microscopy, follow steps 4–6 of the Subheading 3.4 about laser scanning confocal microscopy. NBD fluorescence – enriched in the ld domains – together with the rhodamine fluorescence of the peptides allows a direct visualization of the lateral distribution of the peptides (see Fig. 3). See Note 10 for lipid abbreviations.
Fig. 3. Lateral distribution of transmembrane peptides in GUVs of two lipid mixtures. TMR-labeled synthetic transmembrane peptide (1 mol%) and C6-NBD-PC (1 mol%) were incorporated into GUVs prepared from a nonraft lipid mixture of POPC/DOPE/DOPS (3/1/1 – molar ratio) (left column) and in lipid domain-forming GUVs made from DOPC/SSM/Chol (1/1/1 – molar ratio) (middle and right column). Confocal images of the equatorial plane of the GUVs show C6-NBD-PC fluorescence (top) and fluorescence of TMR-labeled peptide (bottom). C6-NBD-PC as a marker for the ld domain and the peptide are colocalized. Images were taken at 25°C. The scale bar corresponds to 10 mm
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3.4. Membrane Binding of a-Synuclein Evaluated by Laser Scanning Confocal Microscopy
1. Perform covalent labeling of protein with an appropriate reactive fluorophore. Fluorescence of label should not interfere with fluorescence properties of lipid analogues used for domain visualization. Here, a-synuclein was labeled with tetramethylrhodamine-6-maleimide (TMR) (protein labeling was done according to the manufacturer’s protocol). 2. In a reaction tube, mix 75 µl of microscopy buffer (see Note 6) with labeled protein to yield the desired protein concentration in a total volume of 100 µl. Add 25 µl GUV suspension and mix well by pipetting. In case of strong membrane affinity, protein concentrations in the range of tenth of nanomoles per liter are sufficient. 3. For microscopy, pipette the sample on a coverslip (see Note 7). Wait a few minutes for the vesicles to settle down due to their greater specific density. 4. Using a laser scanning confocal microscope, the NBD lipid analogue is ideally excited with the 488-nm line of an Ar-ion laser; for excitation of the TMR fluorescence, a wavelength of 543 nm (He–Ne laser) or 559 nm (diode laser) is suitable (see Note 8). 5. Images of the equatorial focal plane of a single GUV allow a straightforward identification of the lipid domains. In the case of C6-NBD-PC, the ld domains show a bright green fluorescence due to the preferential enrichment of the probe. In contrast, lo or gel phase domains appear as dark regions, since the C6-NBD-PC is excluded (see Fig. 4 (DPPG), Fig. 5 and Note 9). See Note 10 for lipid abbreviations. 6. Binding of the fluorescent-labeled protein to the membrane and – if applicable – specific lipid domains can be directly deduced from the images by comparison with the C6-NBD-PC fluorescence (see Figs. 4 and 5). See Note 10 for lipid abbreviations. 7. For quantification of the amount of membrane bound protein, intensity profiles through the equatorial section of single vesicles can be used. The peak values of TMR fluorescence at the membrane are quite reliable for quantification of the bound protein (see Fig. 6 and Note 9).
4. Notes 1. The amount of sucrose in the swelling buffer is not predefined, but depends on the osmolality of the microscopy buffer. The osmolality of the swelling buffer should be approx. 20 mOsm/ kg lower or at least equal to that of the microscopy buffer, since otherwise the stability of the GUVs can be impaired.
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Fig. 4. Binding of a-synuclein requires negatively charged lipids in liquid disordered phase. Laser scanning microscopy images showing equatorial sections of GUVs incubated with fluorescent labeled a-synuclein (0.1 µM). All GUVs were labeled with 1 mol% C6-NBD-PC. DOPC: DOPC only; All other: 70 mol% DOPC and 30 mol% of indicated anionic lipid. Top image – fluorescence of C6-NBD-PC; Bottom image – fluorescence of TMR-labeled a-synuclein. All images were obtained at 25°C. Some GUVs contain smaller vesicles or lipid debris. The scale bar corresponds to 10 mm
2. The length of the hydrophobic stretch of the transmembrane peptide varies with the number of hydrophobic amino acids in the sequence of the peptide. Choosing lipids with corresponding acyl chain length may help to avoid a mismatch situation in which hydrophobic side chains may become exposed to polar environment, which is energetically unfavorable. 3. The presence of terminal lysines is a common approach to enhance the peptide solubility and to promote a transmembrane insertion into the membrane (18). 4. The lipid mixtures used for GUV preparation can be adapted to meet the requirements of the experiment. GUVs with membranes being solely in the liquid disordered state can be formed from unsaturated (e.g., DOPC) or partly unsaturated (e.g., POPC) phosphatidylcholines (PCs) with different acyl chain lengths. Alternatively, also PC preparations of natural sources
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Fig. 5. Binding of a-synuclein requires negatively charged lipids in liquid disordered phase. Laser scanning microscopy images showing equatorial sections of GUVs incubated with fluorescent labeled a-synuclein (0.1 µM). All GUVs were labeled with 1 mol% C6-NBD-PC. Raft: DOPC/SSM/Chol = 33/33/33; Raft (DOPS): DOPC/SSM/Chol/DOPS = 25/25/25/25; Raft (DSPS): DOPC/SSM/Chol/DSPS = 25/25/25/25. Top image – fluorescence of C6-NBD-PC; Bottom image – fluorescence of TMR-labeled a-synuclein. All images were obtained at 25°C. Some GUVs contain smaller vesicles or lipid debris. The scale bar corresponds to 10 mm
Fig. 6. Quantitative analysis of labeled a-synuclein binding to GUV (DOPC/DOPS = 70/30). (a) In order to measure the fluorescence intensity at the vesicle membrane, a line profile in the equatorial plane of the GUV is selected. (b) The amount of labeled protein bound at the membrane can be measured at the fluorescence peaks found in the profile. (c) Mean intensities of fluorescence at the GUV membrane for different a-synuclein concentrations. Mean values measured from six different GUVs. Bars correspond to the standard error of the mean. The data were fitted according to the scaled particle theory (19)
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(e.g., egg yolk) can be used. To study the effect of anionic lipids (e.g., phosphatidylserine (PS) or phosphatidylglycerol (PG)), rather large fractions of these lipids can be included in the lipid mixture (e.g., 70 mol% of DOPS with 30 mol% DOPC still yield GUVs). As long as these lipids are unsaturated, no phase separation occurs. For GUVs with coexisting ld and lo domains, use a ternary lipid mixtures consisting of unsaturated PC in combination with saturated PC or sphingomyelin and cholesterol. Especially, mixtures of DOPC/SSM/ Chol = 1/1/1 and DOPC/DPPC/Chol = 1/1/1 are often used, and the phase behavior is well studied. An overview (by far not complete) of possible lipid mixtures consisting of zwitterionic lipids has been published, together with the respective phase transition temperatures, by Veatch and Keller (5). 5. The lipid spots on ITO slides should show violet color and concentric ring-like structures. If reproducibly small yellow lumps of lipids on the slides are formed, evaporate the chloroform of the lipid (lipid/peptide) mixture in a nitrogen stream and redissolve the lipids in 50 µl trifluoroethanol. Then, pipette 25 µl of the lipid mixture in one large drop on each of the ITO slides before heating, and subsequently place the spotted slides on a heating stage at approx. 50°C to evaporate the solvent and to form a homogenous lipid film. Normally, this problem can also be solved by the use of titanium chambers instead of ITO slides. 6. The composition of the microscopy buffer can be adjusted to the requirements of the experiment. Instead of glucose, any ionic or nonionic substance can be used as long as it does not disturb the membrane integrity. 7. Due to solvent evaporation, GUVs will be stable only for a limited time. For prolonged periods, instead of coverslips cell culture dishes with a glass bottom can be used. If GUVs tend to spread quickly by forming a flat bilayer at the glass surface, the use of a polylysine-coated support can be advantageous. 8. Microscopy can also be done with an epifluorescence microscope, although a high background from unbound protein or label may result. In each case, the experimental setup should be checked for spectral crosstalk with samples containing only one of the fluorescent dyes. For observation, at least a 40× air objective is adequate, although a 60× oil objective with a high numerical aperture will greatly enhance fluorescence intensity and resolution. 9. As labeling of proteins can significantly change their properties, it must be precluded that the membrane binding of the protein is changed by the introduced label. Therefore, important controls include the incubation of the vesicles with
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free label to prove that it does not bind to the membrane by itself. Additionally, competition experiments with equal amounts of unlabeled and labeled protein should show a reduction of membrane fluorescence of 50% if the binding effectiveness is not changed by the label. 10. Lipids were abbreviated using the following scheme: DO, di-oleoyl-; DP, di-palmitoyl-; DS, di-stearoyl-; PO, 1-palmitoyl2-oleoyl-; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PS, phosphatidylserine; SSM, stearoyl-sphingomyelin; Chol, cholesterol. References 1. Allen JA, Halverson-Tamboli RA, Rasenick MM (2007) Lipid raft microdomains and neurotransmitter signalling. Nat Rev Neurosci 8:128–140 2. Hanzal-Bayer MF, Hancock JF (2007) Lipid rafts and membrane traffic. FEBS Lett 581:2098–2104 3. Lajoie P, Nabi IR (2007) Regulation of raftdependent endocytosis. J Cell Mol Med 11:644–653 4. Nayak DP, Hui EK, Barman S (2004) Assembly and budding of influenza virus. Virus Res 106:147–165 5. Veatch SL, Keller SL (2003) Separation of liquid phases in giant vesicles of ternary mixtures of phospholipids and cholesterol. Biophys J 85:3074–3083 6. Radhakrishnan A, McConnell H (2005) Condensed complexes in vesicles containing cholesterol and phospholipids. Proc Natl Acad Sci U S A 102:12662–12666 7. Lim LH, Pervaiz S (2007) Annexin 1: the new face of an old molecule. Faseb J 21:968–975 8. Stockl M, Fischer P, Wanker E, Herrmann A (2008) Alpha-synuclein selectively binds to anionic phospholipids embedded in liquid-disordered domains. J Mol Biol 375:1394–1404 9. Sengupta P, Baird B, Holowka D (2007) Lipid rafts, fluid/fluid phase separation, and their relevance to plasma membrane structure and function. Semin Cell Dev Biol 18:583–590 10. Korlach J, Schwille P, Webb WW, Feigenson GW (1999) Characterization of lipid bilayer phases by confocal microscopy and fluorescence correlation spectroscopy. Proc Natl Acad Sci U S A 96:8461–8466 11. Dietrich C, Bagatolli LA, Volovyk ZN, Thompson NL, Levi M, Jacobson K, Gratton E
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18.
19.
(2001) Lipid rafts reconstituted in model membranes. Biophys J 80:1417–1428 Fortin DL, Troyer MD, Nakamura K, Kubo S, Anthony MD, Edwards RH (2004) Lipid rafts mediate the synaptic localization of alpha-synuclein. J Neurosci 24:6715–6723 Kubo S, Nemani VM, Chalkley RJ, Anthony MD, Hattori N, Mizuno Y, Edwards RH, Fortin DL (2005) A combinatorial code for the interaction of {alpha}-synuclein with membranes. J Biol Chem 280:31664–31672 Ollesch J, Poschner BC, Nikolaus J, Hofmann MW, Herrmann A, Gerwert K, Langosch D (2008) Secondary structure and distribution of fusogenic LV-peptides in lipid membranes. Eur Biophys J 37:435–445 Angelova MI, Dimitrov DS (1986) Liposome electroformation. Faraday Discuss Chem Soc 81:303–311 Angelova MI, Soleau S, Meleard P, Faucon JF, Bothorel P (1992) Preparation of giant vesicles by external AC electric fields. Kinetics and applications. Prog Colloid Polym Sci 89:127–131 Ayuyan AG, Cohen FS (2006) Lipid peroxides promote large rafts: effects of excitation of probes in fluorescence microscopy and electrochemical reactions during vesicle formation. Biophys J 91:2172–2183 Hesselink RW, Koehorst RB, Nazarov PV, Hemminga MA (2005) Membrane-bound peptides mimicking transmembrane Vph1p helix 7 of yeast V-ATPase: a spectroscopic and polarity mismatch study. Biochim Biophys Acta 1716:137–145 Chatelier RC, Minton AP (1996) Adsorption of globular proteins on locally planar surfaces: models for the effect of excluded surface area and aggregation of adsorbed protein on adsorption equilibria. Biophys J 71:2367–2374
Chapter 11 Biosynthesis of Proteins Inside Liposomes Pasquale Stano, Yutetsu Kuruma, Tereza Pereira de Souza, and Pier Luigi Luisi Abstract Protein expression is the most complex metabolic reaction that has been encapsulated in liposomes, mainly as an intermediate step toward the synthesis of minimal semisynthetic cells. Although there are different experimental approaches to achieving the synthesis of proteins inside liposomes and it is therefore not possible to give a standard recipe, all methods follow a general strategy, which is briefly discussed. On this basis, we provide general indications for designing and realizing protein-expressing liposomes. Our approach for the green fluorescent protein expression inside 200-nm extruded vesicles is described in detail. Key words: Cell-free protein expression, PURESYSTEM, Enhanced green fluorescent protein (EGFP), Minimal cell, Synthetic biology
1. Introduction 1.1. Background
The biosynthesis of functional proteins is, to date, the most complex metabolic network that has been synthetically reconstructed inside lipid vesicles (liposome). The interest and the relevance for such kind of work concern several fundamental aspects of biology and important applications in biotechnology. First, it represents the most advanced step toward the bottomup construction of a semisynthetic minimal cell (1), which is an ambitious goal aiming to understand the very nature of cellular life. This research started about 15 years ago by carrying out molecular biology reactions of increasing complexity inside liposomes (for a review, see ref. (1)). Its final target is the construction of molecular systems that exhibit living-like properties, guided by the theory of autopoiesis (2, 3) also considering that (minimal)
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_11, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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life emerges as a system property as soon as a certain degree of molecular complexity and spatial/functional organizations are reached (4). The theoretical analysis of the minimal number of molecular components required to construct cellular life states that about 200 genes constitute the “minimal genome,” i.e., the genome required for cell survival in a highly permissive environment. According to this view, the environment provides the cell with several low-molecular-weight compounds, whereas the cell synthesizes by itself the characteristic and functional macromolecules coded by the minimal genome. Based on this analysis, it is possible to conceive an experimental research program for the
Fig. 1. Protein synthesis inside liposomes. The whole transcription–translation (T&T) kit, inclusive of an energy regeneration system, is entrapped inside liposomes. The T&T kit, composed of macromolecules (translation factors, ribosomes, tRNAs, aa-tRNA synthases, kinases), and low-molecular-weight compounds (nucleotides, amino acids, salts, phosphate donors) effect the synthesis of functional proteins. Unentrapped components are removed, or their reaction is inhibited by addition of ribonuclease, protease, or EDTA. In advanced models, membrane pores (a-hemolysin (10)) are inserted in order to exchange low-molecular-weight compounds, without releasing macromolecules from the liposome. Drawing adapted from (20), with modifications
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construction of minimal cells of increasing complexity: from simple liposome bioreactors hosting a single enzyme to very complex systems as the whole transcription–translation (T&T) machinery inside liposomes (Fig. 1). Along the roadmap to living (although limping) minimal cells, which are models of primitive cells (1), the expression of one (or more) functional protein(s) inside liposomes has been reported by us and several other groups (5–15). In addition to basic research, the encapsulation of entire metabolic networks inside liposomes has great biotechnological potentiality as therapeutic of diagnostic agents, as discussed by Pohorille and Deamer (16), or as tools for genome replication, modified RNA synthesis, drug discovery, or generation of new microorganism, within the context of the emerging discipline of synthetic biology (17). 1.2. Experimental Strategy
Today, the biosynthesis of proteins inside liposomes is achieved by combining cell-free in vitro protein expression and liposome technology. It should be noted that there is not yet a standard approach to carrying out protein biosynthesis inside liposomes. Different researchers have used different experimental setups, which differ with respect to (i) size and morphology of liposomes, (ii) liposome composition and lipid concentration, (iii) liposome preparation, (iv) nature of cell-free T&T kits and of the external inhibitors, (v) nature of expressed protein(s), and (vi) protein detection method. In addition, only in one case it has been reported that the protein-expressing liposome can exchange matter with the environment (10), realizing a thermodynamically open system; in all other cases, liposomes act essentially as a nonpermeable compartment, i.e., a closed system (5–9, 11–15). The general strategy can be summarized as shown in Fig. 2. The gene of interest downstream to a proper promoter (generally T7, but SP6 has been also used (9)) is mixed together with a T&T kit, composed of cell extracts (18) (homemade or commercially available), or completely reconstituted from purified enzymes, ribosomes, and tRNAs (19, 20). The mixture is kept at low temperature (in ice) in order to prevent the beginning of the reaction, and then liposomes are formed and processed according to specific requirements. Soon after, an inhibitor is added to the liposome suspension. The inhibitor should not permeate the liposome membrane. Alternatively, the external medium may be exchanged with an isotonic buffer. Following proper incubation at 37°C, the synthesis of the protein(s) inside liposomes is then verified analytically (when possible, in real time). Table 1 summarizes the diverse approaches, according to the six (i–vi) points listed above. Vesicles of very different size and morphology have been successfully employed to express proteins (mainly green fluorescent proteins, GFPs) in their inner aqueous
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Fig. 2. General strategy for protein expression inside liposomes. Liposomes are formed in a solution containing a transcription–translation (T&T) kit, composed of cell extracts or reconstituted by purified components (i.e., PURESYSTEM), and a gene of interest, under T7 promoter. The liposome suspension is formed and processed at low temperature to prevent the beginning of the protein synthesis. Then ribonuclease (or protease, or EDTA) is added to the sample in order to inhibit the synthesis of protein by external unentrapped molecules (route a). The inhibitors should not permeate the liposome membrane. Alternatively, the external solution is exchanged by size exclusion chromatography (route b). Compartmentalized protein synthesis will occur within 2–3 h incubation at 37°C
core. Uni- or oligolamellar vesicles with diameter between 200 and 400 nm can be obtained by the extrusion method (21) or by the ethanol injection method (22). Large multilamellar vesicles are generally obtained by hydration of a lipid film or of a liposome cake (prepared by freeze-drying of preformed liposomes) (23), whereas giant vesicles (GVs) can be generated by natural swelling (24) or by the new method of w/o emulsion inversion, developed by Pautot et al. (25) (notice that there are no report on protein expression in GVs formed by the electroswelling method). In order to carry out protein biosynthesis inside vesicles, homemade or commercially available cell extracts have been largely used. After the initial report of Ueda and coworkers (19, 20) and the successive introduction in the market of purified T&T components (i.e., the PURESYSTEM, sold by the Japanese company Post Genome Institute- Co., Ltd., Tokyo), it is foreseeable that this totally reconstituted kit will prevail over the E. coli cell extracts, which actually can be seen as a “black box” of not well-defined molecular composition. In fact, the concentrations of all molecular species in the
VET400
MLV, GV
LUV
GV
MLV, GV
GV
MLV, GV
MLV
1
2
3
4
5
6
7
8
50–200 mM POPC
48 mM POPC, PLPC, SOPC, SLPC, cholesterol, DSPEPEG5000
EggPC
15 mM EggPC, CHOL, DSPE-PEG5000 (1.5:1:0.08)
11 mM DOPC, DOPG (10:1)
1.5 mM POPC
20 mM EggPC, CHOL, DSPE-PEG5000 (1.5:1:0.08)
66.7 mM POPC
(i) Size and (ii) Composition and Entry morphology concentration
PURE SYSTEM; 5.7 mg/ mL RNase
PURE SYSTEM; 7.3 mg/ mL RNase
Liposome cake rehydration
Liposome cake rehydration
Cell extract (Roche); no external inhibitor
Homemade; 12 mg/mL RNase
Cell extracts (Ambion); 50 mg/mL proteinase K
Cell extracts (Promega); 33 mM EDTA
Homemade; 12 mg/mL RNase
Homemade; 35 mM EDTA
(iv) T&T and inhibitors
w/o emulsion droplets inversion
Liposome cake rehydration
Natural swelling
Ethanol injection
Film hydration
Film hydration, FT, Extrusion
(iii) Preparation
Table 1 Summary of protein biosynthesis inside liposomes
EGFP
EGFP
EGFP and alphahemolysin
GFPmut1 and T7RNApol
rsGFP
EGFP
GFPmut1
Poly(Phe)
(v) Expressed protein(s)
Fluorescence: batch fluorimetry and confocal laser microscopy
Fluorescence: flow cytometry
(12)
(11)
(10)
(9)
(8)
(7)
(6)
(5)
Ref.
(continued)
Fluorescence: laser scanning microscopy
Radiolabeling and fluorescence: SDS-PAGE and flow cytometry
Fluorescence: laser scanning microscopy
Fluorescence: batch fluorimetry
Fluorescence: flow cytometry and laser scanning microscopy
Radiolabeling: SDSPAGE
(vi) Detection method
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PURE SYSTEM; 20 mg/mL RNase
Radiolabeling: SDSPAGE
Fluorescence: batch fluorometry
EGFP
GPAT and LPAAT
Radiolabeling: SDSPAGE
(vi) Detection method
GPAT and LPAAT
(v) Expressed protein(s)
(15)
(14)
(13)
Ref.
MLV multilamellar vesicles; GV giant vesicles; VET400 vesicles by extrusion techniques, with diameter 400 nm; LUV large unilamellar vesicles (100–200 nm); LUV-EI large unilamellar vesicles obtained by the ethanol injection method; POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine; egg-PC egg yolk phosphatidylcholine; CHOL cholesterol; DSPE-PEG5000 1,2-distearoyl-sn-glycero-3-phosphatidylethanolamine-poly(ethylenglycol) 5000; DOPC 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine; DOPG 1,2-dioleoyl-sn-glycero-3-phosphatidylglycerol; PLPC 1-palmitoyl-2-linoleoyl-sn-phosphatidylcholine; SOPC 1-stearoyl-2-oleoyl-sn-phosphatidylcholine; SLPC 1-stearoyl-2linoleoyl-sn-phosphatidylcholine; POPE 1-stearoyl-2-oleoyl-sn-phosphatidylethanolamine; POPG 1-stearoyl-2-oleoyl-sn-phosphatidylglycerol; FT freeze-and-thaw; w/o water-in-oil; T&T transcription and translation kit; EDTA ethylendiaminotetraacetic acid; RNase ribonuclease; SEC size exclusion chromatography; poly(Phe) poly(phenylalanine); GFPmut1 green fluorescent protein mutant 1; EGFP enhanced green fluorescent protein; rsGFP red-shifted green fluorescent protein; T7RNApol T7 RNA polymerase; GPAT glycerol-3phosphate acyltransferase; LPAAT lysophosphatidic acid acyltransferase; SDS-PAGE sodium dodecylsulfate-polyacrylamide gel electrophoresis
200 mM POPC, POPE, POPG, cardiolipin (50.8:35.6:11.5:2.1)
Film hydration
MLV
11
Cell extracts (Promega) and PURE SYSTEM; 180 mg/mL RNase or SEC
VET200 and 8.5–15 mM LUV-EI POPC
10
PURE SYSTEM; 20 mg/mL RNase
(iv) T&T and inhibitors
Ethanol injection, extrusion
Film hydration
MLV
9
200 mM POPC
(iii) Preparation
(i) Size and (ii) Composition and Entry morphology concentration
Table 1 (continued)
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PURESYSTEM are known (see Note 1) and individually tunable, at least in principle. Protein synthesis that may occur outside vesicles can be inhibited by the addition of ethylenediaminetetraacetic acid (EDTA) (Mg(II) chelator), or ribonuclease (RNase A, digesting the mRNA), or proteinase K (digesting the enzymes needed to accomplish the T&T processes). Finally, almost all studies refer to the synthesis of GFPs, which is a model protein with easy detection. Notice that GFP is fluorescent due to an intramolecular cyclocondensation between three residues (Ser 65, Tyr 66, Gly 67) and consequent oxidation to give a p-hydroxybenzylideneimidazolinone (26). It is recognized that the appearance of its fluorescence implies a correct fold, so that it can be said that the produced protein is “functional” (26). In addition to GFPs, some studies have focused on the expression of T7 RNA polymerase (Table 1, entry 5) in order to create a two-stage cascading expression network, or of a-hemolysin (Table 1, entry 6), in order to make the liposome membrane porous. All these proteins (GFP, RNA polymerase, a-hemolysin) are water soluble. Only recently, Kuruma et al. (13, 15) reported on the synthesis of membrane proteins inside liposomes (Table 1, entries 9 and 11). 1.3. Choice of the Conditions
From the foregoing discussion, it is evident that there does not exist a unique and specific protocol to carry out protein expression inside liposomes. The choice of the different variables (i–vi) very much depends on the aim of the experiment; moreover, some analytical techniques are possible only for specific cases (e.g., fluorescence-based techniques can be used only when GFPs are expressed). Here we shortly report some general guidelines. (i) Vesicle size and morphology. Unilamellar vesicles should be preferred whenever is possible. Working with unilamellar GVs (e.g., diameter ³10 mm), for example, is advantageous, since it is possible to follow directly the evolution of the system, for example, at morphological level or by employing some fluorescent reporters. On the other hand, GVs must be observed one by one, or in small groups, and the advantage of large number averaging is lost. On the contrary, smaller vesicles (100–200 nm), especially if homogeneously sized, provide a good experimental model, and their preparations are reproducible but suffer the disadvantage of a small internal volume and therefore need highly sensitive methods for assessing the yield of the synthesized protein. Intermediate sized vesicles, e.g., ca. 500–1,000 nm diameter vesicles, are generally oligoor multilamellar and difficult to produce. To the best of our knowledge, there are no standard methods to produce unilamellar vesicles with those sizes. Multilamellar vesicles, although convenient in some applications, are to be considered unsuitable to carry out protein expression in a controlled way for a liposome system.
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(ii) Liposome composition and lipid concentration. It is interesting to notice that protein expression has been carried out mainly in phoshatidylcholine vesicles. This lipid is chemically quite inert (for example, negatively charged lipids which may interact with Mg2+ ions, or positively charged lipids which strongly interact with nucleic acids and are reported to strongly reduce the protein synthesis (5, 27)), readily forms stable vesicles, and ensures good entrapping yields. DOPC and POPC have low phase-transition temperatures −18.3 and −2.5°C in water, respectively, (28), so that they can be used at a low (+4°C) temperature. Liposome composition is an important issue in the case of expressing membrane proteins (15, 29) and when liposomes with special performances are required. (iii) Liposome preparation. This is of course related to point (i). Since enzymes and ribosomes have to be entrapped inside liposomes, preparation protocols, which require the application of harsh conditions or high detergent concentration may lead to poor results. The hydration of lipid film or rehydration of liposome cake (obtained by freeze-drying previously formed liposomes) is the safest procedures. Ethanol injection method is also applicable, but yields are partially reduced because of the presence of alcohol in the final suspension (methanol can also be used: the protein yield is slightly higher, but the liposome size distribution is broader). Extrusion also leads to a reduction of yield (14), whereas mild (bath) sonication and freeze-andthaw are apparently well tolerated by T&T kits. A systematic study on the effect of liposome manipulation on the protein synthesis inside liposomes is still missing, so that the conclusions sketched above must be considered with caution. (iv) Nature of cell-free T&T kits and of the external inhibitors. For the expression of prokaryotes protein, the choice is essentially between cell extracts (homemade or bought from several companies) and the PURESYSTEM. Cell-free kits based on cell extracts generally provide higher yields than PURESYSTEM; their composition, however, is unknown and cannot be considered really functional for the constructive paradigm of synthetic biology. PURESYSTEM offers the additional advantage of easy purification of expressed protein, due to the fact that its components are all His-tagged, and the target protein can be easily isolated by selective removal of His-tagged components (19). Cell extracts are also generally viscous and may affect the liposome preparation steps. RNase A has been widely employed to avoid the occurrence of the reaction outside the liposomes (6, 9, 12), but its effectiveness must be checked in control experiments to take into account the effect of liposomes. Proteinase K has also been used (8). Protein expression can be inhibited by EDTA
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(>30 mM). EDTA does not suffer limitations as in the case of macromolecular inhibitors (RNase A and proteinase K); however, it may provide osmotic stress to liposomes, and its use must be checked with respect to liposome size, morphology, and manipulation/processing. The most elegant way to eliminate external synthesis is to exchange the external solution with an isotonic buffer by size exclusion chromatography (14), see Fig. 2b. (v) Nature of expressed protein(s). This parameter depends on the aim of the experiment. Water-soluble or membrane-associated proteins require different strategies, mainly related to the composition of liposome membrane. In fact, when a membrane protein must be expressed inside vesicles, great care must be taken for the choice of lipids, which will affect the membrane/protein interaction and also the protein activity. Liposome membrane, in a sense, should be considered in these cases as a reactant. When the research aim is not the expression of a specific protein, and the experiment is focused on other aspects of compartmentalized reactions, an easily detectable water-soluble protein must be chosen. (vi) Protein detection method. The common strategy here follows the classical methods developed for cell-free protein expression. The most general detection method involves the incorporation of labeled amino acids in the protein sequence. Very common is the use of [35S]-methionine; when required, however, other 35S-, 14C-, or 3H-labeled amino acids can be employed (see Note 2). The freshly synthesized protein, in all these methods, is concentrated by precipitation (trichloroacetic acid (TCA) or acetone precipitation, see Note 3) and then analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). In the case of radiolabeling, the dried gel is then exposed to autoradiographic film or detected with a phosphor imager. Alternatively to the incorporation of labeling amino acids, the protein can be detected by Western blotting (primary and secondary antibodies are required). Here the synthesized protein is analyzed by SDSPAGE and then transferred to the membrane for primary and secondary antibody binding and immunodetection. These methods are very useful, since they can be opportunely applied to every protein, regardless from their sequence, structure, and function. In particular cases, however, it is possible to directly detect the expressed protein by probing its “activity.” This is possible when the target protein is an enzyme or an intrinsically fluorescent protein, such as in the case of the family of GFPs. If the protein activity is successfully measured, this will provide strong evidence for the proper synthesis and folding of the synthesized protein. The enzymatic assay must
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be properly designed for each case, also considering the compartmentalized nature of the system, and therefore the problem of membrane permeability. This will affect the way of substrate delivery. When GFPs are expressed, on the other hand, the product analysis becomes easier. For this reason, the first studies on the biosynthesis of proteins inside liposomes have been carried out with GFPs as target proteins. In fact, GFPs can be revealed by following their intrinsic fluorescence. GFP fluorescence can be measured by single-wavelength measurements (5, 14), emission spectra measurements (5, 14), laser scanning fluorescence microscopy (6, 8, 10, 12, 14), and flow cytometry (6, 9, 11). Many of these methods can be used to monitor fluorescence signals in real time, providing a facile way to follow the kinetics of protein synthesis. A serious problem in fluorescent measurements of GFP inside liposomes stems from sample turbidity, which gives rise to a scattering-based inner filter effect (IFE) (see Note 4). It is clear that the expression of proteins inside liposomes is still a challenging task and that it is difficult to provide a standard recipe for all cases. Here we describe in detail a protocol that has been successfully developed in our laboratory concerning the expression of enhanced GFP (EGFP) in extruded vesicles. The production of EGFP can be followed flurometrically; however, we also provide the classical protocol to verify its production by SDS-PAGE.
2. Materials 2.1. Cell-Free Protein Expression Kit, Buffer and Reagents
1. Plasmid pWM-T7-EGFP (3,026 bp, see Note 5) (BioTecton, Zurich, Switzerland), encoding egfp gene under T7 promoter dissolved in pure nuclease-free water (1 mg/mL) and stored in proper aliquots at −20°C. 2. PURESYSTEM Classic II kit (Post Genome Institute Co., Ltd., Tokyo, Japan) (see Note 6), must be properly divided in aliquots and stored at −80°C. Avoid freeze–thaw cycles. Do not mix solution A and solution B before use. 3. l-[35S]-Methionine, 1,000 Ci/mmol (GE Healthcare, code SJ1015) (this chemical is needed only when the synthesized protein must be analyzed by SDS-PAGE and autoradiography). 4. Nuclease-free water (produced by Milli-Q apparatus, Millipore). 5. Recombinant EGFP (BD Bioscience, Switzerland), stored at −20°C in the dark.
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6. “Multicore”-like sterile buffer (MC): 125 mM potassium acetate, 25 mM Tris acetate, pH 7.5. Store at 4°C. 7. RNase A, 2 mg/mL (Sigma-Aldrich), in MC buffer; freshly prepared and kept in ice. 8. Sterile Eppendorf-like 1.5-mL vials and 200 mL PCR-like vials (see Note 7). 2.2. Liposome Preparation, Manipulation, and Processing
1. Synthetic POPC, i.e., 1-palmitoyl-2-oleoyl-sn-glycero-3phosphatidylcholine, (Avanti Polar Lipids, Inc., Alabaster, USA) is dissolved in absolute ethanol (500 mM) and stored in aliquots under nitrogen atmosphere at −20°C. 2. Two sterile Teflon-covered magnetic stirring bars; length 5 mm, diameter 2 mm (Sigma-Aldrich, Code Z328839). Wash extensively with ethanol and dry before use. 3. Microsyringe, 10 mL, with a narrow needle (Hamilton). 4. Extruder “Liposo-fast” (Avestin Inc., Ottawa, Canada), equipped by two 250-mL gastight microsyringes (Hamilton). Its components contain glass, Teflon, viton (O-rings), and stainless steel. They are easily cleaned and the entire instrument is autoclavable. 5. Three drain discs, 13 mm (Osmonics, Inc., Article Nr. C32WP01300). 6. Two polycarbonate membranes, 200 nm pore size (Nuclepore Track-Etched Membranes, Whatman, Maidstone, UK).
2.3. SDS-PAGE
1. Stacking buffer (4×): 0.5 M Tris–HCl, pH 6.8, 0.4% (w/v) SDS. Store at room temperature. 2. Separating buffer (3.75×): 1.5 M Tris–HCl, pH 8.8, 0.4% SDS. Store at room temperature. 3. SDS running buffer (10×): 250 mM Tris, 1.92 M glycine, 1% SDS. Store at room temperature. 4. Staining buffer: 0.5% (w/v) Coomassie Brilliant blue R-250 (CBB), 50% (v/v) methanol, 10% (v/v) acetic acid. Store at room temperature. 5. Destaining (fixing) buffer: 50% methanol, 10% acetic Acid. Store at room temperature. 6. SDS loading buffer: 125 mM Tris–HCl. pH 6.8, 20% (v/v) glycerol, 4.6% SDS, 0.006% (w/v) Bromophenol Blue (BPB), 10% (v/v) b-mercaptoethanol (add before use). Store at room temperature. 7. Bisacrylamide/30% acrylamide: 30% (w/v) acrylamide, 0.8% (w/v) N,N¢-methylenebisacrylamide. Store at 4°C. 8. Ammonium peroxodisulfate (APS) (10% (w/v)). Store at 4°C.
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9. N,N,N¢,N¢-Tetramethylethylenediamine (TEMED). Store at 4°C. 10. Acetone, diethyl ether (analytical grade). Important: All chemicals should be dissolved in distilled water. Use gloves when acrylamide is handled, because it is a neurotoxin when unpolymerized and so care should be taken not to receive exposure.
3. Methods 3.1. Preparation of the Extruder
1. Assemble the extruder by placing three drain discs (see Note 8) and two polycarbonate membranes (200 nm pore size) between the two Teflon blocks, so that the two membranes are sandwiched between the three drain discs (i.e., drain disc – membrane – drain disc – membrane – drain disc). Close the extruder by tightening the two external screws. 2. Wash the extruder by passing through the membranes 200 mL MC buffer. 3. Keep the assembled extruder in the cold room at least 30 min before the experiment (see Note 9).
3.2. Preparation of Reaction Mixture and Liposome Formation
All operations described below must be done in the cold room (4°C). All equipmentfor liposome formation/manipulation/processing must be kept in the cold room at least 30 min before the beginning of experiments. 1. Thaw all components required for the transcription–translation reaction in ice (solution A and B of PURESYSTEM, pWMT7-EGFP plasmid). Nuclease-free water, “multi-core”-like buffer, POPC solution in ethanol, and RNase A solution must also be kept in ice. 2. Gently mix 100 mL solution A, 40 mL solution B, and 50 mL nuclease-free water in a sterile Eppendorf-like vial, and keep the resulting mixture in ice. This mixture will provide the positive control sample (protein synthesis inside and outside liposomes) and the test sample (protein synthesis only inside liposomes). Negative control samples (no protein synthesis) are similarly prepared, but 20 mL of RNase A solution is also added at this stage. 3. Add two Teflon-covered followers to the mixture prepared (point 2). 4. Add 4 mL plasmid pWM-T7-EGFP (1 mg/mL) to the mixture (point 3).
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5. Put the Eppendorf-like vial containing 194 mL of T&T/ gene mixture in a small water/ice bath over a magnetic stirrer and stir at about 500 rpm; quickly inject 1.5 mL POPC solution; charge the microsyringe again and inject three further 1.5 mL fractions, for a total of four injections (total 6 mL). To avoid excess shearing force, decrease the stirring rate between injections. The suspension should appear turbid, by the presence of 15 mM POPC (final concentration). The ethanol content is 3% v/v. 6. Put the liposome suspension in ice. 7. Repeat the procedure (points 3–6) for the negative control sample. When the EGFP synthesis has to detected on the SDS-PAGE gel, 4 mL of l-[35S]methionine (1,000 Ci/mmol) must be supplied to the PURESYSTEM reaction mixture (total reaction volume 200 mL), and the amount of pure water decreased to 46 mL. 3.3. Liposome Processing
1. Mount a needle on the 250-mL gastight microsyringe of the extruder and fill it with the liposome suspension. Remove the needle. 2. Insert the syringe into the extruder inlet and extrude the liposomes by passing them 11 times back and forth through the two polycarbonate membranes (see Note 10). Avoid prolonged contact between hands and syringes, in order to avoid heat transfer. 3. Collect the extruded vesicles and make 50 mL aliquots inside PCR-type 200-mL vials. Put the aliquots in ice. (see Note 11). 4. Add 5 mL of cold MC buffer to the positive control samples (protein synthesis inside and outside liposomes). Keep the samples in ice. 5. Add 5 mL of cold RNase A solution to the other fractions (final RNase A concentration 0.18 mg/mL) and mix gently; these will be the test samples (protein synthesis only inside vesicles). Keep the samples in ice. 6. Repeat the procedure (points 1–3) for negative control samples, but make 55 mL aliquots. Keep the samples in ice.
3.4. Real-Time Fluorescence Analysis of EGFP-Producing Liposomes
1. Real-time (RT) PCR instruments can be used as sensitive fluorimeters if the temperature is set constant (37°C). We have successfully employed a RT-PCR Corbett Rotor-Gene 6000. Such an instrument improves reproducibility, enhances sensibility, allows the use of small sample volumes (25 or 50 µL), and provides homogeneous heating. Instrumental settings: fluorescence channel: green (excitation 470 nm, emission 510 nm); gain 5, 7, or 10; cycle setting: 120 s at 37.0 (±0.5)°C; 400 rpm.
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Fig. 3. Analysis of produced EGFP by (a) real-time fluorimetric assay and (b) SDS-PAGE. In (a) the fluorescence versus time curves show that positive control sample (EGFP expression inside and outside liposomes) reaches high fluorescence values, whereas negative control sample (RNase A added before the liposome formation) has low fluorescent values that do not increase in time. The test sample, on the other hand, can express EGFP inside liposomes, since RNase A has been added after liposome formation. In this example, the relative yield of inside-liposome production is about 3% ( 28.5, 1.9, and 0.8 being the values of fluorescence for the three samples). In (b), the classical pattern of SDS-PAGE analysis is shown (M MW markers; P positive control; T test sample; N negative control). The marker has been added for the sake of clarity (after radiolabeling, only expressed proteins can be visualized, and correspondence with markers for MW check should be done in a separate experiment)
2. Insert the 55-mL samples (positive controls, test samples, negative controls) in the RT-PCR machine and record fluorescence for 3–5 h. This will provide the fluorescence versus time curves for each sample. Remove the sample from ice immediately before starting fluorescence recording. 3. Read the asymptotic fluorescence value for the samples, and average values for samples run in duplicate. Calculate the EGFP yield inside liposomes as follows: EGFP relative yield (%) = 100 × (Ftest − Fneg)/(Fpos − Fneg). Typical relative yields range from 1 to 10% (see Fig. 3a). 3.5.Calibration Line for Fluorescence Analysis
1. A calibration line is needed to quantify the production of EGFP in terms of absolute yield (EGFP amount) (for an alternative method, see Note 12). 2. Create five samples (55 mL) by mixing solution A, solution B, water, and non-EGFP coding plasmid of same size as pWM-T7EGFP, preformed 200 nm extruded POPC liposomes, RNase if required, and different amounts of recombinant EGFP (for example: 0, 0.1, 0.3, 0.6, 0.8 mg/mL), so that the composition of the samples is qualitatively and quantitatively similar to real
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experimental samples, and measure their fluorescence on the RT-PCR machine for 10 min. Five straight lines (fluorescence vs. time) should be obtained. Use the fluorescence values to construct a calibration line (fluorescence vs. [EGFP]). 3. Use the calibration line to measure the amount of EGFP produced in the liposome-based experiment. 3.6. SDS-PAGE Analysis
1. In this study, Bio-Rad Mini-PROTEAN Tetra Cell kit is used for the SDS-PAGE analysis, since this kit has been commonly used in molecular biology laboratories. Before composing the kit, two mini glasses are first cleaned up by distilled water and then rinsed by 95% ethanol. After completely dry, the glasses are assembled by using the accessory platform. 2. Mix the separating gel (lower gel) solution as follows. For 10 mL of 12% gel: 4 mL of 30% acrylamide/bisacrylamide, 2.5 mL of 4× separating gel buffer, and 100 mL of 10% APS are mixed, and then distilled water is added up to 10 mL. Ten microliters of TEMED is also supplied and mixed just before pouring into the glasses. The percentage of gel can be varied by adjusting the volumes of acrylamide solution and water. 3. Pour the gel solution into the slit of the composed glasses leaving space for the stacking gel. Isopropanol or distilled water is overlaid on the poured solution to avoid drying the solution surface and to allow formation of a horizontal surface. The gel should polymerize in about 30 min. 4. Pour off the isopropanol or water completely. 5. Mix the stacking gel solution (upper gel) as follows. For 3 mL of 5% gel: 0.5 mL of 30% acrylamide/bisacrylamide, 0.8 mL of 4× stacking gel buffer, and 30 mL of 10% APS are mixed, and then distilled water is added up to 3 mL. Three microliters of TEMED is also supplied and mixed just before pouring into the glasses. 6. Pour the gel solution over the separating gel, and insert the comb. The stacking gel should polymerize within 30 min. 7. Prepare the running buffer by diluting 50 mL of 10× running buffer with 450 mL of water. The diluted buffer should be stirred by a magnetic stirrer bar in a beaker. 8. Once the stacking gel has polymerized, insert the gel assembly unit to the electrophoresis apparatus and supply the running buffer. Carefully remove the comb within the running buffer and wash the wells by pipetting. Check that there is no air at the bottom of the gel. When bubbles are present, remove them by a syringe fitted with thin distorted needle. 9. Before loading the samples (positive control, test sample, and negative control), a pretreatment must be carried out in
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order to avoid the jumbling of the gel pattern due to lipid components. For this, proteins must be precipitated by acetone (see Note 3). Acetone (200 mL) is added to liposomecontaining samples (50 mL) and incubated on ice for 30 min. Then, the samples are centrifuged at 18000 × g for 10 min. The resulting supernatants (acetone) are removed by pipetting, and additional acetone (200 mL) is added again to the precipitates. The suspensions are vortexed and the suspended precipitates are finely dispersed by using bath sonication, and then centrifuged (18000 × g for 10 min). After removal of the supernatants (acetone), 100 mL of diethylether is added to the precipitates. The precipitates are dispersed again by bath sonication and centrifuged (18000 × g for 10 min). After removing the supernatants (diethylether), the resulting precipitates are completely dried by a centrifuge-evaporator for 5 min. Finally, the dried precipitates are dissolved in SDS loading buffer with the appropriate volume (max 20 mL). Through these processes, lipid components can be removed, and the produced radiolabeled [35S]-protein can be safely analyzed by SDS-PAGE. The dissolved samples should be boiled at 95°C for 5 min just before loading into the gel. 10. Carefully load the samples into the wells, which include one well for the prestained molecular weight marker. 11. Complete the assembly of the apparatus and connect to a power supply. Apply a voltage (less than 100 V) until the dye of sample enters the separating gel. After the sample enters the separating gel, the voltage can be increased up to 200 V. Stop the gel running when the dye front arrives at the bottom of gel, and carefully remove the gel from the glasses. 12. If the sample is nonradiolabeled, the gel is processed to stain by staining buffer, and then destained by destaining buffer with a piece of paper towel. If the sample is radiolabeled, incubate the gel with fixing buffer. All these steps are done for 30 min with gentle shaking. 13. Take a photograph of the stained gel (nonradiolabelled). 14. For the radiolabeled gel, the gel is dried up on a filter paper (Whatman) by using a Bio-Rad Gel Dryer, for 30 min at 80°C. 15. The dried gel can be analyzed by an imaging system, such as GS-525 Molecular Imager System (BioRad). Expose the gel to the storage phosphor imaging screen for at least 12 h (this time must be optimized according to the yield of the expressed protein), and then analyze the screen by the IR-laser-scanning acquisition module. A typical pattern is shown in Fig. 3a.
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4. Notes 1. The PURESYSTEM is composed by 10 translation factors (IF1, IF2, IF3, EF-G, EF-Tu, EF-Ts, RF1, RF2, RF3, RRF), 20 aminoacyl-tRNA synthases, T7 RNA polymerase, ribosomes, methionyl-tRNA formyltransferase, creatine kinase, myokinase, nucleoside diphosphate kinase, pyrophosphatase, and a mix of tRNAs (37 macromolecular compounds + tRNAs). In addition, low-molecular-weight compounds are present (ATP, GTP, CTP, UTP, DTT, spermidine, creatine phosphate, 20 amino acids, 10-formyl-5,6,7,8-tetrahydrofolic acid, potassium glutamate, and magnesium acetate) (19, 20). 2. Alternative to radiolabeling, biotinylated amino acids (30) or fluorescently labeled amino acids (31) can be used. In the first case, after SDS-PAGE and blotting, streptavidin-linked enzymes (phosphatase, peroxidase) allow colorimetric or chemiluminescent detection. In the second case, direct visualization of the fluorescent bands in the gel/membrane is possible. 3. Acetone precipitation (followed by diethyl ether rinse) should be preferred to TCA precipitation (12, 13, 15). In fact, acetone effectively solubilizes lipids, which may interfere in SDS-PAGE analysis. TCA precipitation is the classic choice in liposome-free cell-free synthesis. 4. Turbidity-based IFE is not very well described in literature, in contrast to absorbance-based IFE. IFE leads to a reduction of fluorescence signal due to absorption or scattering of the excitation and emitted radiation. In some extreme cases, the IFE may remove completely the fluorescence signal, especially the signal is not very strong. It follows that the quantification of a fluorescent molecule may be difficult in the presence of IFE. To solve this problem, two approaches can be followed. In the first approach, liposomes are solubilized as micelles by addition of a detergent (e.g., sodium cholate). This treatment reduces IFE by reducing the size of particles present in the solution, and therefore by reducing the scattered light. Of course, the effect of such a detergent on the protein fluorescence must be checked in preliminary experiments. In the second approach, a proper calibration line (in the presence of liposomes) must be constructed. The liposomes used to record the calibration line should be similar in average size and dispersity to the liposomes obtained in the real experiment. 5. Detail of the T7 (underlined), linker (italics), and EGFP sequences of pWM-T7-EGFP plasmid: TAATAC GACTCA CTATAG – GGAGAC CACAAC GGTTTC CCTCTA
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GAAATA ATTTTG TTTAAC TTTAAG AAGGAG ATATAC AT – EGFP sequence. Plasmid purity should be carefully checked, as well as the quality of water. Nuclease contamination, e.g., deriving from the plasmid preparation, should be avoided. 6. http://www.postgenome.jp/english/index.html 7. PCR-like vials should be colorless and possibly made by thin plastic, in order to have higher transparency. 8. Cut three drain discs so that their diameter fits well with the head area (limited by the O-rings) on the extruder Teflon blocks. 9. In some cases, low temperature causes shrinkage of the pistons’ tips, giving rise to leakage during extrusion. A preliminary check of the extruder in the cold room is suggested. 10. A possible problem at this stage is leakage from the extruder, due to imperfect assembly of the extruder or to the wear of mechanical parts. Check the extruder (in the cold room) in a preliminary experiment by extruding liposomes at similar concentration. Very small volumes (e.g., 50 mL) cannot be effectively extruded. 11. Label the vials on the top. 12. Use the standard addition method to quantify EGFP in a complex matrix. In this method, small aliquots of pure EGFP (at a known concentration) are added to the sample where EGFP of unknown concentration has to be determined. The addition(s) should provide an increase of EGFP concentration of the same order of magnitude as the unknown EGFP concentration (e.g., if synthesized EGFP is present at an estimated concentration of 0.05 mg/mL, add EGFP aliquots so that the total estimated EGFP concentration becomes 0.1 mg/ mL, 0.2 mg/mL, etc.). Record the fluorescence values and make a plot of recorded fluorescence versus the added EGFP concentration. The x-intercept (a negative number) is the concentration of EGFP in the sample. This method takes into account the interference of the matrix (liposomes, T&T kit, etc.) with the fluorescence signal of EGFP, and do not need a calibration line. Further information can be found in classical analytical chemistry books, for example (32).
Acknowledgements This work was carried out within the SYNTHCELLS project (Approaches to the Bioengineering of Synthetic Minimal Cells, EU Grant #FP6–043359) and further supported by the Human Frontiers Science Program.
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References 1. Luisi PL, Ferri F, Stano P (2006) Approaches to semi-synthetic minimal cells: a review. Naturwissenschaften 93:1–13 2. Varela FJ, Maturana HR (1974) Autopoiesis: the organization of living systems, its characterization and a model. Biosystems 5:187–196 3. Luisi PL (2003) Autopoiesis: a review and a reappraisal. Naturwissenschaften 90:49–59 4. Luisi PL (2006) The emergence of life: from chemical origin to synthetic biology. Cambridge University Press, Cambridge 5. Oberholzer T, Nierhaus KH, Luisi PL (1999) Protein expression in liposomes. Biochem Biophys Res Comm 261:238–241 6. Yu W, Sato K, Wakabayashi M, Nakatshi T, Ko-Mitamura EP, Shima Y, Urabe I, Yomo T (2001) Synthesis of functional protein in liposome. J Biosci Bioeng 92:590–593 7. Oberholzer T, Luisi PL (2002) The use of liposomes for constructing cell models. J Biol Phys 28:733–744 8. Nomura SM, Tsumoto K, Hamada T, Akiyoshi K, Nakatani Y, Yoshikawa K (2003) Gene expression within cell-sized lipid vesicles. ChemBioChem 4:1172–1175 9. Ishikawa K, Sato K, Shima Y, Urabe I, Yomo T (2004) Expression of a cascading genetic network within liposomes. FEBS Lett 576:387–390 10. Noireaux V, Libchaber A (2004) A vesicle bioreactor as a step toward an artificial cell assembly. Proc Natl Acad Sci USA 101:17669–17674 11. Sunami T, Sato K, Matsuura T, Tsukada K, Urabe I, Yomo T (2006) Femtoliter compartment in liposomes for in vitro selection of proteins. Anal Biochem 357:128–136 12. Murtas G, Kuruma Y, Bianchini P, Diaspro A, Luisi PL (2007) Protein synthesis in liposomes with a minimal set of enzymes. Biochem Biophys Res Comm 363:12–17 13. Kuruma Y (2007) Question 7: Biosynthesis of phosphatidic acid in liposome compartments – Toward the self-reproduction of minimal cells. Orig Life Evol Biosph 37:409–413 14. Souza T, Stano P, Luisi PL (2009) The minimal size of liposome-based model cells brings about a remarkably enhanced entrapment and protein synthesis. ChemBioChem 10: 1056–1063 15. Kuruma Y, Stano P, Ueda T, Luisi PL (2009) A synthetic biology approach to the construction of membrane proteins in semi-synthetic minimal cells. Biochim. Biophys. Acta 1788: 567–574 16. Pohorille A, Deamer D (2002) Artificial cells: prospects for biotechnology. Trends Biotechnol 20:123–128
17. Forster AC, Church GM (2006) Towards synthesis of a minimal cell. Mol Syst Biol 2:45 18. Zubay G (1973) In vitro protein systems synthesis of in microbial systems. Annu Rev Genet 7:267–287 19. Shimizu Y, Inoue A, Tomari Y, Suzuki T, Yokogawa T, Nishikawa K, Ueda T (2001) Cell-free translation reconstituted with purified components. Nature Biotechnol 19:751–755 20. Shimizu Y, Kanamori T, Ueda T (2005) Protein synthesis by pure translation systems. Methods 36:299–304 21. Hope MJ, Bally MB, Webb G, Cullis PR (1985) Production of large unilamellar vesicles by a rapid extrusion procedure. Characterization of size distribution, trapped volume and ability to maintain a membrane potential. Biochim Biophys Acta 812:55–65 22. Batzri S, Korn ED (1973) Single bilayer liposomes prepared without sonication. Biochim Biophys Acta 298:1015–1019 23. Kikuchi H, Suzuki N, Ebihara K, Morita H, Ishii Y, Kikuchi A, Sugaya S, Serikawa T, Tanaka K (1999) Gene delivery using liposome technology. J Control Release 62:269–297 24. Nomura S-IM, Yoshikawa Y, Yoshikawa K, Dannenmuller O, Chasserot-Golaz S, Ourisson G, Nakatani Y (2001) Towards proto-cells: “primitive” lipid vesicles encapsulating giant DNA and its histone complex. ChemBioChem 6:457–459 25. Pautot S, Frisken BJ, Weitz DA (2003) Production of unilamellar vesicles using an inverted emulsion. Langmuir 19:2870–2879 26. Tsien RY (1998) The green fluorescent protein. Annu Rev Biochem 67:509–544 27. Tachibana R, Harashima H, Ishida T, Shinohara Y, Hino M, Terada H, Baba Y, Kiwada H (2002) Effect of cationic liposomes in an in vitro transcription and translation system. Biol Pharm Bull 25:529–531 28. Cevc G (ed) (1993) Phospholipids handbook. Marcel Dekker Inc, New York 29. Lee AG (2004) How lipids affect the activities of integral membrane proteins. Biochim Biophys Acta 1666:62–87 30. Kurzchalia TV, Wiedmann M, Breter H, Zimmermann W, Bauschke E, Rapoport TA (1988) tRNA-mediated labeling of proteins with biotin. A nonradioactive method for the detection of cell-free ranslation products. Eur J Biochem 172:663–668 31. PROMEGA Product # L5001, technical NOTES, # TB285 32. Harris DH (2003) Quantitative chemical analysis. W. H. Freeman and Co., New York
Chapter 12 Study of Respiratory Cytochromes in Liposomes Iseli L. Nantes, Cintia Kawai, Felipe S. Pessoto, and Katia C.U. Mugnol Abstract Important findings regarding the structure and function of respiratory cytochromes have been made from the study of these hemeproteins associated to liposomes. These studies contributed to the comprehension of the biological role of these proteins in the electron transfer process, the regulatory mechanisms, the energy transduction mechanisms, the protein sites that interact with mitochondrial membranes and the role played by the non-redox subunits present in the protein complexes of the respiratory chain of eukaryotes. In this chapter, the protocols developed to study cytochrome bc1 activity in liposomes and the binding of cytochrome c to lipid bilayers is presented . The former protocol was developed to study the mechanism of energy transduction related to the topology of the components of bc1 complex in the mitochondrial membrane. These studies were done with purified cytochrome bc1 complexes reconstituted into potassium-loaded vesicles. The latter protocol was developed to study the influence of pH, DpH, and DY on the interaction of cytochrome c with liposomes that mimic the inner mitochondrial membrane. Key words: Cytochromes bc1, Eukaryotic cytochrome c, Respiratory chain, Energy transduction, Liposomes
1. Introduction 1.1. The bc1 Complex
Several studies of bc1 complex incorporated into liposomes can be found in the literature. These studies are concerned with the mechanism of proton translocation in the cytochrome bc1 complex and the function of individual subunits of the enzyme in the energy transduction process. Two different mechanisms have been proposed to explain the coupling of the electron transfer in the bc1 complex with the proton translocation from the matrix to the intermembrane space: the redox loop and the proton pump mechanisms. The redox loop mechanism explains the coupling by the
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_12, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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existence of arranged redox centers that, after reduction, accept protons from the matrix and, after reoxidation, release the protons to the intermembrane space. According to the proton pump mechanism, the reduction and re-oxidation of protein redox centers induce changes in the pKa of their amino acid side chains and their alternate exposure to the internal and external side of the membrane (1–3). The electron transfer pathway through bc1 complex is linked with the coenzyme Q cycle. The coenzyme molecules solved inside the membrane lipid fraction are completely reduced to QH2 by accepting electrons from complex I or II and the high potential b562 of complex III concomitant with the uptake of two protons from the mitochondrial matrix. In the re-oxidation process, QH2 donates one electron to cytochrome c1 via the iron sulfur protein (ISP) center, and one electron to heme b566 that recycles it back to oxidized coenzyme Q via heme b562. Concomitant with the re-oxidation, coenzyme Q releases two protons in the intermembrane space (Scheme 1). According to the originally proposed coenzyme Q cycle (4), in the bc1 complex, approximately 40% of the transmembrane potential is generated by the vectorial transmembrane electron transfer from cytochrome b566 to cytochrome b562 and the remaining 60% has been attributed to the electron transfer from cytochrome b562 to Q localized in the i center of the bc1 complex (5) and to
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the transfer of protons associated with the oxidation and reduction of Q (6, 7). Several studies concerning the mechanism that couples the electron transfer to a transmembrane proton electrochemical potential have been done by using bc1 complex incorporated into liposomes and involve the measurement of the catalytic and the back reaction, i.e., the electron transfer from a component with a more positive redox potential to a component with a more negative potential driven by an electrochemical potential gradient of protons. The study of electron transfer within the purified cytochrome bc1 complex reconstituted in liposomes has several advantages relative to the study with submitochondrial particles because, in the model system, the influence of other components of the electron transfer chain is eliminated, and the effect of Q can be studied by using the bc1 complexes depleted and replenished of Q. The mechanism of energy transduction related to the topology of the components of bc1 complex in the mitochondrial membrane has been studied in purified cytochrome bc1 complexes reconstituted into K+-loaded vesicles. In this model system, the addition of valinomycin to the K+-loaded proteoliposomes in a medium that does not have external KCl promotes the back electron transfer from ascorbate-reduced cytochrome c1 to ISP which, in turn, reduces Q to QH2. The reduced coenzyme then reduces the high potential heme b562. The thermodinamically unfavorable electron transfer from ISP to Q and from the high potential b562 to low potential b566 is coupled with proton uptake from the internal side of the membrane. Thus, in this model, the reversed transport of electrons is closely coupled to the reverse flow of protons related to the Q cycle (8). However, the demonstration that the reduction potential of the protein redox centers are responsive to conformational changes led to the postulation that there are least two redox-linked ionizable groups (9–12) in the b subunit, one closer to the outside and another closer to the inside. These groups have pKa values that vary according to the redox states of both hemes b and, possibly, to the redox states of quinone. More recently, a model compromising between pure “redox-loop” and pure “protonpump” mechanisms has been proposed . According to this model, in the respiratory chain, the one-electron reduction of oxidized coenzyme Q by complex I or complex II leads to the formation the semiquinone intermediate Q•−. This intermediate accepts one electron from b562 and is converted to the reduced form. The reduction of Q•− is accompanied by the uptake of one proton from the matrix and one proton from a redox-linked ionizable group. QH2 is reconverted to semiquinone by donating one electron to b566 and one proton to a redox-linked ionizable group while the second proton is delivered in the positive side of the membrane. The second electron is transferred to ISP and from that to cytochrome c1. The conformational changes promoted by the electron
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H+
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Scheme 2. Coenzyme Q cycle related to bc1 electron transport and the generation of transmembrane potential according to the redox cycle associated to the proton pump mechanism
transfer to b subunit lead to the uptake of a proton from the matrix side by the redox-linked ionizable group concomitant with the electron transfer from b566 to b562 (Scheme 2). 1.2. Respiratory Cytochrome c
Cytochrome c is a mobile electron carrier in the respiratory chain that exhibits also the ability to detach from the inner mitochondrial membrane to integrate the apoptosome in the cytosol and promote apoptosis (13, 14). The cellular oxidative stress and loss of mitochondrial membrane potential promoted as Ca2+-induced permeability transition or photodynamic therapy (PDT), are known inducers of cytochrome c release and apoptosis (15–24). Therefore, the nature and specificity of the interaction of cytochrome c with model membranes have been extensively studied (25–28). The association of cytochrome c with phospholipid membranes involves both electrostatic and hydrophobic interactions (29–31). In model systems, at pH 7.4, the interaction of cytochrome c with acidic phospholipids occurs via site A (19, 32, 33), an electrostatically interacting site constituted of basic residues in cytochrome c, probably Lys72 and Lys73.
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The interaction of cytochrome c with the phospholipid head group leads to the accommodation of one phospholipid acyl chain within a hydrophobic channel of the protein lined by hydrophobic amino acid residues from the surface of the protein to the heme crevice. The insertion of a phospholipid acyl chain in the cytochrome c structure leads to perturbation in the heme iron spin state detected by electron paramagnetic resonance (EPR) (32). These studies have shown a clear correlation between the nature of the lipid acyl chain and the spin states of cytochrome c interacting with different types of lipid membranes. However, on the external surface of the inner mitochondrial membrane, the clusters of positively charged amino acid side chains (site A) are important for the interaction of the cytochrome c redutase and oxidase (13, 14, 34). More recently, the interaction of cytochrome c with mitochondrial mimetic vesicles of PCPECL (1,2-dipalmitoyl-sn-glycero-3-phosphocholine, 1,2-dipalmitoyl-snglycero-3-phosphoethanolamine and heart cardiolipin) was investigated over the 7.4–6.2 pH range by means of turbidimetry and photon correlation spectroscopy. In the presence of cytochrome c, the decrease of pH induced vesicle fusion dependent of protein ionizable groups with a pKa(app) around 7.0. The carbethoxylation of these groups by diethylpirocarbonate impaired cytochrome c-induced vesicle fusion, remaining cytochrome c association to vesicles unaffected. Matrix-Assisted Laser Desorption/Ionization (MALDI) Time-of-Flight (ToF) analysis revealed that Lys22, Lys27, His33 and Lys87 cytochrome c residues were the main targets for carbethoxylation performed at low pH values (<7.5). This site was denominated site L because the high lysine content. The structural analysis of cytochrome c structure shows that these amino acid residues belong to clusters of positively charged amino acids responsible for the lowering of the pKa. Subsequent unpublished studies have revealed that the association of cytochrome c with PCPECL liposomes via site L and monitored by turbidimetry is favored by electric potential difference provided by DpH and DY. Thus, the turbidimetry analysis of these vesicles in the presence of cytochrome c can be used to determine the existence and loss of electric potential difference in these vesicles. Besides, turbidimetry, z potential analysis is also an adequate technique to detect the binding and vesicle fusion promoted by cytochrome c.
2. Materials 2.1. Protocols to Study the Back Electron Transfer in the bc1 Complex
1. 10 l 0.1 M phosphate buffer pH 7.4. 2. 1,300 ml 0.02 M phosphate buffer, pH 7.4. 3. 250 ml borate–phosphate buffer: 0.1 M Na2HPO4 and H3BO3. 4. 800 ml 0.01 M KH2PO4.
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2.1.1. Purification of bc1 Complex from beef heart muscle
1. 50 mM phosphate buffer, pH 7.4, containing 0.25 M sucrose.
2.1.2. Separation of Cytochrome bc1 Complex from SuccinateCytochrome c Reductase
4. 50 mM Tris–HCl buffer, pH 8.0, containing 0.67 M sucrose.
2. 50 mM phosphate–borate buffer, pH 7.8. 3. 50 mM phosphate buffer, pH 7.4. 5. 200 mM stock solution of succinate was prepared in an argon atmosphere in the presence of 10 mM dithiothreitol. 6. Potassium Deoxycholate. 7. Ammonium acetate solution, which was prepared according to Hatefi and Rieske (35) by dissolving 454 g of solid ammonium acetate in 613 ml of H2O. 8. 18.5% ammonium acetate saturation (20 ml of 50% saturated ammonium acetate solution to 100 ml solution). 9. 2 M urea. 10. Sodium cholate.
2.1.3. K+-Loaded Cytochrome bc1 Vesicles
1. 500 mM Stock solution cytochrome c (horse heart, type III) was prepared in deionized water (see Note 1) and the concentration was calculated using molar absorptivity e409 = 106.1 × 103 M/cm (36) and stored at −20°C. 2. Stock solution of valinomycinin ethanol. 3. Stock solution of nigericin in ethanol. 4. Stock solution of antimycin A in ethanol. 5. 48 mM stock solution of safranine in deionized water. 6. 50 mM stock solution of sodium ascorbate in deionized water (see Note 2). For a storage during hours the solution should be saturated with argon (see Note 3). 7. Asolectin (soybean phospholipids) and purified partially according to Sone et al. (37). 8. 10 mM 3-(N-mopholino)propanesulfonic acid (MOPS), pH 7.4, prepared by titration of the solution with NaOH solution. 9. 100 mM KCl. 10. 100 mM NaCl.
2.2. Protocols to Study the Interaction of Respiratory Cytochrome c Sites A and L with Lipid Bilayers (38)
1. Stock solution cytochrome c (horse heart, type III) was prepared in deionized water and the concentration was calculated using molar absorptivity e409 = 106.1 × 103 M/cm (39) and stored at −20°C (see Note 6). 2. HCl, NaOH, NaCl, HClO4, KH2PO4 and ammonium molybdate used to prepare stock solutions at deionized water (conductivity = 18 ohms). 3. HEPES (4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid) buffer prepared at different pH values by the titration of the solution with NaOH and HCl.
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4. Tris base 2-Amino-2-(hydroxymethyl)-1,2-propanediol buffer solution prepared by the titration of the solution with NaOH and HCl. 5. Acetic acid used to prepare acetate buffer. 6. Ascorbic acid and d-glucose used to prepare stock solutions at deionized water (conductivity = 18 ohms). 7. Chloroform used to solve phospholipids stock solutions. 8. Egg phosphatidylcholine(PC), Lipids 1,2-dioleoyl-sn-glycero-3phosphoethanolamine (DOPE) and 1,1¢,2,2¢tetraoleoyl cardiolipin (CL). To prepare the lipid stock solution, each lipid type is weighed and dissolved in chloroform in order to obtain a concentration of 20 mg/ml stored at −20°C. The correct concentration was determined by phosphate dosage. Total phospholipid content is determined by phosphate analysis, using the method of Bartlett (40), with some modifications. 2 ml of each lipid is dried in a pyrex tube and heated in hot block at 120°C for 10 min. In the following, 0.4 ml HClO4 70% is added in a tube of HClO4 and heated at 180°C for 1 h. After this step, the tubes are removed from the hot block and cooled to room temperature. 1.0 ml of water and 0.4 ml of 1.252% ammonium molybdate are added inside each tube and the samples are vortexed. 0.4 ml of fresh ascorbic acid 3% is added, mixed by vortex and then incubated at 90°C in water bath for 10 min. The samples are then cooled to room temperature and absorbance at 797 nm is recorded. Concentrations are determined using a calibration curve obtained with KH2PO4 standard solutions from 0 to 100 nmols. 9. Polycarbonate membrane filter with 100 and 1,000 nm pore. 10. Spectrapor dialyze tubing of 3,500 molecular weight cutoff. 11. PyPC (2-(10-(1-pyrene) decanoyl) phosphatidylcholine).
3. Methods 3.1. Purification of bc1 Complex from Beef Heart Muscle
Step 1: Extraction of succinate-cytochrome c reductase from beef heart muscle 1. The preparation of heart muscle is based on Keilin and Hartree method with modifications done by King (1961) (41). Heart muscle is cleaned of fat and connective tissues, and minced in a power-driven meat grinder. One kilogram of mince is washed with 20 l of tap water in a plastic bucket and efficient mechanical stirring is carried out for 20 min (see Notes 3 and 4). In the paper published in 1961 by King, the author describes the model of the mechanical stirrer, equipped with a three-blade stainless steel propeller of approximately 6 in. across (see Notes 4 and 5).
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2. The mince is pressed and squeezed either by hand in cheese cloth, or by a hydraulic press in canvas (37). Usually, six washings result in a mince as a light yellow, but not pink, preparation. The mince is then used either directly for grinding, or further mixed with 10 l of 0.1 M phosphate buffer, pH 7.4, and stirred mechanically for approximately 1 h. It is finally washed with 20 l of deionized water. 3. Five hundred grams of the hard-pressed mince thus obtained are ground in a mechanical mortar for 2 h with 700 ml of 0.02 M phosphate buffer, pH 7.4 and 200 g of sand (37). 4. The mixture is diluted with another 600 ml of buffer and then centrifuged for 30 min at 600 × g. 5. The upper turbid layer is recentrifuged at 6,000 × g for 90 min. 6. The pellet is dispersed in 250 ml of borate–phosphate buffer, with a Potter-Elvehjem type homogenizer. All operations described are done at room temperature. 7. The turbid supernatant fluid from the first centrifugation is cooled to approximately 2°C by the addition of crushed ice and brought to pH 5.5 with 0.1N acetic acid. 8. The mixture is centrifuged at 600 × g for 15 min in a refrigerated centrifuge. 9. The clear supernatant fraction is discarded and the precipitate is washed with 800 ml 0.01 M KH2PO4 and centrifuged again. The residue is finally suspended in borate–phosphate buffer. The heart muscle preparation is not stable at room temperature in mediums with pH values lower than 6.8. Step 2: Separation of cytochrome bc1 complex and succinate dehydrogenase from the succinate-cytochrome c reductase extracted from beef heart muscle Separation of the bc1 complex and succinate dehydrogenase from succinate-cytochrome c reductase can be achieved essentially according to the method previously described (42) except that a slightly lower pH and a more strictly anaerobic condition should be used in order to get a better recovery of succinate dehydrogenase. 1. Succinate-cytochrome reductase as prepared was dialyzed against 50 mM phosphate buffer, pH 7.4, containing 0.25 M sucrose overnight with two changes of buffer and frozen at −70°C until be used. 2. Succinate-cytochrome reductase, 10 mg/ml, was centrifuged at 170,000 × g for 45 min and the collected precipitate was resuspended in 50 mM phosphate–borate buffer, pH 7.8, to a protein concentration of 15 mg/ml. 3. Then it was incubated with 20 mM succinate in an argon atmosphere in the presence of 1 mM dithiothreitol for 30 min before the pH of the solution was brought to 10 by anaerobic addition of NaOH.
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4. The mixture was further incubated at 0°C for 20 min before being centrifuged at 170,000 × g for 45 min. 5. Succinate dehydrogenase was collected from the supernatant solution and the collected precipitate containing the cytochrome bc1 complex was resuspended and homogenized in 50 mM phosphate buffer, pH 7.4 and then brought to pH 10.3 once more by the addition of 0.1N NaOH. 6. After incubation at 0°C for 20 min, the mixture was centrifuged at 170,000 × g for 40 min. 7. The bc1 complex was collected in the precipitate, and the supernatant solution, containing a small amount of succinate dehydrogenase, was discarded. 8. The precipitate was suspended in 50 mM Tris–HCl buffer, pH 8.0, containing 0.67 M sucrose and kept at –70°C for the isolation of ubiquinone-binding protein and bc1 III complex (ubiquinol-cytochrome c reductase). Step 3: Separation of ubiquinone-binding protein and bc1 complex from ubiquinol-cytochrome c reductase (complex III) 1. The frozen bc1 complex is thawed and protein concentration adjusted to 20 mg/ml with 50 mM Tris–HCl buffer, pH 8.0, containing 0.67 M sucrose. 2. The suspension is solubilized with potassium deoxycholate at a concentration of 0.5 mg per mg of protein, and fractionated with ammonium acetate solution. 3. Ammonium acetate solution is added to the deoxycholatesolubilized bc1 complex, 11 ml of 50% saturated ammonium acetate solution per 100 ml solution, to give 5% ammonium acetate saturation. 4. The mixture is stirred at 0°C for 20 min and then centrifuged at 50,000 × g for 30 min. The supernatant is collected and brought to 10% ammonium acetate saturation (12 ml of ammonium acetate solution to 100 ml of solution). 5. The pellet, which contains ubiquinone-binding protein, is collected by centrifuging at 50,000 × g for 30 min and dissolved in 50 mM Tris–HCl buffer, pH 8.0, containing 0.67 M sucrose. 6. The crude ubiquinone-binding protein preparation can be frozen at −70°C for several weeks without significant loss of activity. 7. The supernatant solution is collected and brought to 18.5% ammonium acetate saturation (see Subheading 2). 8. After stirring for 20 min, the solution is centrifuged at 170,000 × g for 30 min. 9. The pellet which contains purified ubiquinol-cytochrome reductase is dissolved in 50 mM Tris–HCl pH 8.0, containing 0.67 M sucrose, diluted to a protein concentration of approximately 20 mg/ml, and frozen at −70°C.
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10. The ubiquinol-cytochrome c reductase thus obtained can be further purified by ammonium sulfate fractionation in the presence of 2 M urea and 0.5% sodium cholate at a protein concentration about 10 mg/ml. 11. The highly purified ubiquinol-cytochrome c reductase is recovered in the precipitate formed between 55% and 70% ammonium sulfate saturation. 12. Succinate dehydrogenase, ubiquinone-binding protein and bc1 complex can be identified by their enzymatic activities as described by (43). 13. By following the above procedure, a SDS–polyacrylamide gel electrophoresis, of the purified sample shows that the bc1 is free from contamination by succinate dehydrogenase protein. 3.2. Preparation of Vesicles
Vesicles containing cytochrome bc1 vesicles were prepared by the cholate dialysis method (44–46) as follows in presence of potassium buffer. 1. Asolecitin dissolved in chloroform was mixed in the pyrex tube and then evaporated with N2 gas in order to obtain film and kept under reduced pressure for at least 2 h. 2. The lipid film was resuspended at concentration of 40 mg/ml in 10 mM MOPS/NaOH, pH 7.4, 100 mM KCl and 1.5% sodium cholate and then sonicated for 10–15 min at 0˚C.
3.3. Reconstitution of Proteoliposomes
1. The purified bc1 complex isolated from beef heart muscle according to the method describe by Yu and Yu, 1980 (40), was added for final concentration of 5 mM, incubated for 5 min at 4°C. 2. The solution was dialyzed against 400 times volume in 10 mM MOPS/NaOH, pH 7.4 plus 100 mM KCl at 4°C for 24 h with two changes of outer buffered solution. 3. In order to remove the external potassium, the proteoliposomes were dialyzed again against 10 mM MOPS/NaOH, pH 7.4 plus 100 mM NaCl at 4°C for two days with four changes of the dialysis medium.
3.4. Reduction of Cytochrome c Coupled to ValinomycinInduced K+ Diffusion Potential (8)
1. Reduction of cytochrome c1 was done by spectroscopic measurements can be carried out in a UV-visible spectrophotometer equipped with a thermostated cell holder. 2. The catalytic activity of the reconstituted bc1 complex is assayed at 20°C in a medium containing 10 mM MOPS/ NaOH, pH 7.4 and 100 mM KCl, 50 mM cytochrome c, and 25 mM Q2H2. 3. The initial rate of cytochrome c reduction is corrected against non-enzymatic reduction of cytochrome c by ubiquinol in the absence of bc1 vesicles. Proteoliposomes reconstituted
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with the bc1 complex exhibit the oxidation control ratio ranging from 7 to 9 that is determined as the ratio of the enzyme activity in the presence and absence of 0.2 mg of valinomycin plus 2 mg of nigericin. 4. The ability of K+ diffusion potential can be checked spectrophotometrically using safranine as an optical probe for membrane potential (47) as follows. A 5 ml portion of proteoliposomes containing the bc1 complex is added to 1 ml of a medium containing 4.8 mM safranine, 10 mM MOPS/ NaOH, pH 7.4 and 100 mM NaCl. The reaction is initiated by the addition of valinomycin, and the change in the membrane potential is measured by the changes in the absorbance of safranine at 525 nm (48) or at 530–578 nm (4). 5. The reduction of cytochrome c and c1 is determined spectrometrically using the following extinction coefficients: De550 nm = 19.2 mM/cm for cytochrome c and De553−539 nm (reduced−oxidaded) = 17.5 mM/cm for cytochrome c1. (reduced−oxidaded) 6. The response of the reduction level of cytochrome b in the reconstituted bc1 complex to the valinomycin-induced K+ diffusion potential can be monitored spectrophotometrically. In these experiments, cytochrome c1 and iron–sulfur protein in the bc1 complex are reduced with 5 mM sodium ascorbate in a medium in which there is no external KCl. In the following, valinomicin is added and this procedure is accompanied by a rapid increase in absorbance at 562 nm relative to 575 nm, the wavelength chose to monitor the redox state of cytochrome b components. After approximately 3 min, it is observed the maximum reduction level of reduced cytochrome b. 7. The rapid reoxidation of b components can be observed after addition of nigericin that collapses the K+ gradient. 8. The dependence of cytochrome b reduction on the magnitude of the diffusion potential (DY) formed across liposomal membranes induced by valinomycin can be monitored by changing the concentration of KCl in the reaction medium. The DY can be calculated from the Nernst equation DY (mV) = 60 log([K+]in/[K+]out) and assuming that the internal KCl concentration is 100 mM. Table 1 shows the results obtained by measuring the cytochrome b reduction as a function of the membrane potential. Cytochrome b562 was not reduced at potentials below 30 mV, and the reduction and the maximal reduction is attained at more than 210 mV (the initial concentration of external KCl was less than 30 mM). The data were obtained from ref. (8). 9. According to the reported method (49) ubiquinone-10 content of the purified bc1 complex can be determined spectrophotometrically using an extinction coefficient of 12.25 mM/cm of the difference between the oxidized and reduced quinine at 275 nm (50).
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Table 1 Results obtained by measuring the cytochrome b reduction as a function of the membrane potential Potential (mV)
A(562–575)
0
0
30
0
60
0.0005
87
0.0016
120
0.0029
150
0.0033
180
0.0035
205
0.0039
240
0.00375
10. The total protein content can be estimated according to Lowry et al. (51). 3.5. Study the Interaction of Respiratory Cytochrome c Sites A and L with Lipid Bilayers [52]
Step 1: Preparation of PCPECL liposomes 1. Aliquots of lipids dissolved in chloroform and calculated to a final concentration of 5 mM in a final volume of 2 ml, are mixed in the pyrex tube and the solvent evaporated by N2 or argon gas stirring (see Note 7). The aliquot of lipid stock to be added in the pyrex is calculated according to the molar percentage of each lipid in the mixture: 50% PC, 30% PE and 20% CL. 2. After complete evaporation of the solvent, the lipid film was kept under reduced pressure for at least 2 h to eliminate residual solvent. The films can be used immediately after the preparation or stored at −20°C until use (see Notes 7 and 8). 3. The liposomes are prepared by adding 1 ml cold buffer followed by vortex stirring. This procedure renders multilamelar liposomes. Unilamelar liposomes can be obtained by extrusion of hydrated lipid dispersions in a mini-extruder. Samples are subjected at least to 11 passes through one polycarbonate filters (100 or 1,000 nm pore size) installed in tandem (53). After this procedure one more milliliter of buffer is added in the lipid pyrex tube and the previous step is repeated using the same filters. 4. The preparation of liposomes with DpH by varying inside and outside pH at a range of 6.2–8.2 is done by the hydration of the lipid film with 5 mM HEPES buffer.
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5. For inside pH variations (acidic outside) the lipid films are individually hydrated by 5 mM HEPES buffer at the pH values from 6.2 to 8.2. After the procedures described in the item 3 above, the external pH is corrected to 6.2 by HCl titration. 6. For outside pH variations (alkaline inside), one lipid film is hydrated by 5 mM HEPES buffer pH 8.2. After the procedures described in the item 3 above, the external pH is corrected with NaOH solution to the desired pH (from 6.4 to 8.2). Due to the high permeability of lipid bilayers to protons (54), the liposomes should be used for experimental procedures immediately after the preparation. 7. To obtain liposomes with DY, the lipid film is hydrated by 5 mM HEPES buffer pH 6.2 containing NaCl at a concentration range of 50–150 mM. The NaCl outside liposomes is eliminated by exhaustive dialysis against 500 times volume of 5 mM HEPES in presence of d-glucose with equivalent osmolarity (Os/kg). 150 mM NaCl (~0.277 Os/kg) = 266 mM d-glucose (55). The dialysis is performed using Spectrapor tubing (3,500 molecular weight cutoff) previously hydrated for 5 min. Step 2: Binding of cytochrome c to PCPECL liposomes and vesicle fusion 1. Determination of cytochrome c binding to PCPECL liposomes and vesicle fusion by PyPC (2-(10-(1-pyrene) decanoyl) phosphatidylcholine) fluorescence. This technique permits to detect the binding of cytochrome c to PyPC-labeled vesicles because the hemeprotein quenches the fluorescence of pyrene. The vesicle fusion is monitored by the decrease in the excimer/ monomer (E/M) ratio of PyPC (56, 57) present in PCPECL vesicles, which were mixed with unlabeled vesicles in the presence of protein. In conditions, in which the binding occurs predominantly via site A, there is no vesicle fusion and only partial quenching of pyrene fluorescence is observed without significant changes in the E/M ratio. In conditions in which sites A and L of cytochrome c are protonated, the protein binding is accompanied by vesicle fusion and, besides pyrene fluorescence quenching, changes in the E/M ratio are also detected. (a) The preparation of PyPC-labeled liposomes is done as follows. Aliquots of small unilamellar PCPECL (50/30/20% mol) vesicles containing PyPC 7% mol are added to identical unlabelled PCPECL vesicles to a final ratio of 1/10 labeled/unlabeled vesicles. Fluorescence emission spectra are recorded at 30°C using a fluorescence spectrometer. (b) To determine the binding of cytochrome c and vesicle fusion, aliquots of cytochrome c to a final concentration of 0.9 mM are added to 50 mM PCPECL liposomes in 10 mM HEPES pH 6.2 or 7.4 (3 ml) and the fluorescence
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2100
Relative Fluorescence
Monomer
pH 7.4
pH = 6.2 1400
1400
700
0
700
Excimer
400
450
500
400
450
500
0
Wavelength (nm)
Fig. 1. Effect of cytochrome c on the fluorescence emission spectra of PCPECL vesicles containing 7 mol% PyPC. Vesicles (0.05 mM PCPECL; labeled/unlabeled 1:10) incubated at 30 °C in 10 mM HEPES buffer with 0.9 mM cyt c for 378 s at pH 6.2 and pH 7.4. Thin solid line corresponds to the initial time, before addition of cytochrome c and thick solid line corresponds to the final incubation time
is monitored during 10 min with the following parameters: excitation wavelength at 344 nm, scan range = 350 to 550 nm and emission slits = 5 nm. The E/M ratio is calculated by using the fluorescence intensity at 459 nm (excimer) and at 398 nm (monomer). Figure 1 shows an example of the results obtained in conditions in which cytochrome c binds to lipid bilayer (pH = 7.4) and in conditions in which the binding was favored and accompanied by vesicles fusion (pH = 6.2). 3.6. Determination of Cytochrome c Binding to PCPECL Liposomes and Vesicle Fusion by Turbidimetry
1. Turbidity measurements are conducted in a spectrophotometer using quartz cuvettes of 1 cm light path and a slit of 0.5 nm. 2. The kinetics of vesicle fusion is monitored continuously by the increase of the turbidity at 480 nm as a function of time. Vesicles alone, without cytochrome c are used as blank. 3. A typical kinetics is carried out with 1 ml of 0.25 mM liposomes in HEPES buffered water pH 6.2 in the presence of 4.0 mM cytochrome c, at 30°C, during 10 min with 1 s time interval. 4. Because the protonation of cytochrome c site L, the cytochrome c-promoted PCPECL vesicle fusion is favored at acidic pH. Transmembrane potential provided by DpH and DY acts synergistically with external acidic pH to favor the binding of cytochrome c to lipid bilayers. Therefore, the comparative capacity of cytochrome c to fuse negatively charged vesicles is an indicator of the presence of transmembrane potential. An example of these effects is shown in Fig. 2.
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Turbidity (480 nm)
0,8 pHi = 7.4, pHo = 6.2 pHi = pHo = 6.2 + 50 mM NaCl(in) pHi = pHo = 6.2
0,6
0,4
0,2
0,0
0
100
200
300 400 Time (s)
500
600
Fig. 2. Effect of DpH and DY on the turbidity of the medium containing PCPECL vesicles and cytochrome c. Experiments were carried out at 30°C using 0.25 mM lipids containing 50% DPPC, 30% DPPE, and 20% CL in 10 mM HEPES buffer in the conditions indicated in the Figure
0.6
Turbidity (480 nm)
0.5 0.4 0.3 0.2 0.1 0.0 6.0
6.4
6.8 7.2 Outside pH
7.6
8.0
Fig. 3. Effect of outer pH (inside 7.4) on the turbidity at 480 nm of the medium containing PCPECL vesicles. The turbidity was determined 10 min after incubation at 30°C. Experiments were carried out at with 0.25 mM lipids containing 50% DPPC, 30% DPPE, and 20% CL or phosphatidylglycerol
5. The data on turbidity increase (Dturbidity at 480 nm) as a function of external pH (inside pH = 7.4) render a sigmoid curve that is well fitted by Eq. 1 and Fig. 3.
∆turbidity =
lim1 10(pH-pk a) 10(pH-pk a) + 1
(1)
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3.7. Determination of Cytochrome c Binding to PC/PE/CL Liposomes by Mean Zeta-Average Diameters (Dz) and Zeta-Potentials (z)
1. At 25°C, Dz and z are determined by dynamic light scattering and laser-Doppler microelectrophoresis, respectively, using a ZetaPlus-ZetaPotential Analyzer (Brookhaven Instruments Corporation, Holtsville, NY) equipped with a 570 nm laser and dynamic light-scattering at 90° for particle sizing (58). Particle size (Dz) and zeta-potential (z) are the mean values calculated from at least 10 independent measurements with standard deviations lower than 5% from the mean. z is determined, in pure water, from electrophoretic mobility m and the Smoluchowski’s equation: z = mh/e, where h is the medium viscosity and e the medium dielectric constant. 2. For the mean zeta-average diameters (Dz) and zeta-potentials (z) measurements the final lipid concentration can be varied from 0.25 to 1.0 mM. The binding of cytochrome c to negatively charged vesicles is accompanied by increase of potential and in the occurrence of fusion by increase of vesicle mean diameter. In conditions in which the binding of cytochrome c to the negatively charged PCPECL vesicles is favored, the z potential increase to less-negative values and in conditions in which the vesicle fusion is also favored the mean diameter of the vesicles is significantly increased.
4. Notes 1. All solutions necessary for the protocols described in this chapter should be prepared in water that has a resistivity of 18.2 MW cm. This standard is referred to as “deionized water” in this chapter. 2. Ascorbate is susceptible to oxidation and only fresh solution should be used for the reduction of cytochrome c. For storage during hours, the solution should be saturated with argon. 3. The saturation of solutions with argon can be done by purging the gas directly inside the solution during at least 10 min followed by closing the tube or cuvette with a lid septum. 4. The mechanical stirring of tissues (heart muscle in this chapter) should be done under refrigeration that could be warranted by an ice bath. 5. For a better yield and to avoid proteolytic fragmentation of the proteins of the bc1 complex during the extraction procedure, the following inhibitors of proteases should be added to the solutions: 0.1 M aminocaproic acid, 6.5 mM benzamidine, 5.5 mM iodoacetamide and 0.1 mM phenylmethylsuphonyl fluoride (PMSF).
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6. The UV spectrometry should be done by using quartz cuvettes because glass cuvettes impair the passage UV light. 7. The preparation of liposomes could be done by using extrusor, tip or bath sonicator. If the bath sonicator is used, the temperature should be set to 50°C. The formation of unilamelar liposomes can be observed by the decrease of the sample turbidity. If the extrusor is used, it can be heated to facilitate the traffic of the solution. 8. In the preparation of lipid films, the evaporation of chloroform should be done under direct jet of argon or nitrogen concurrently with the manual rotation of the tube maintained in an inclined position relative to the needle. References 1. Trumpower BL (1990) The protonmotive Q cycle. J Biol Chem 265:11409–11412 2. Boyer PD (1993) The binding change mechanism for ATP synthase. Some probabilities and possibilities. Biochim Biophys Acta 1440:215–250 3. Erecinska M, Wilson DF (1982) Regulation of cellular energy metabolism. J Membr Biol 70:1–14 4. Mitchell P (1976) Possible molecular mechanisms of the protonmotive function of cytochrome systems. J Theor Biol 62:327–367 5. Robertson DE, Dutton PL (1988) The nature and magnitude of the charge-separation reactions of ubiquinol cytochrome c2 oxidoreductase. Biochim Biophys Acta 935: 273–291 6. Drachev LA, Kaurov BS, Mamedov MD, Mulkidjanian AY, Semonev AY, Shinkarev VP, Skulachev VP, Verkhovsky MI (1989) Biochim Biophys Acta 973:189–197 7. Konstantinov A, Kunz WS, Kamensky YA (1981) In: Skulachev VP, Hinkle PC (eds) Chemiosmotic proton circuits in biological membranes, Addison-Wesley pp 123–146 8. Miki T, Mikin M, Orii Y (1994) Membrane potential-linked reversed electron transfer in the Beef Heart Cytochrome bcl complex reconstituted into potassium-loaded phospholipid vesicles. J Biol Chem 269:1827–1833 9. West IC, Mitchell P, Rich PR (1988) Electron conduction between b cytochromes of the mitochondrial respiratory chain in the presence of antimycin plus myxothiazol. Biochim Biophys Acta 933:35–41 10. Konstantinov AA, Popova E (1988) In: Papa S (ed) Cytochrome systems: molecular biology and bioenergetics, Plenum Publishing Corp., New York, pp 751–765
11. Kamensky YuA, Artzabanov VYu, Shevchenko DV, Konstantinov AA (1979) Effect of antimycin A on redox-dependent protonation of the b cytochromes of the mitochondrial respiratory chain. Dokl Acad Nauk SSSR 249:994–997 12. Von Jagow G, Link TA, Ohnishi T (1986) Organization and function of cytochrome b and ubiquinone in the cristae membrane of beef heart mitochondria. J Bioenerg Biomembr 18:157–179 13. Kluck RM, BossyWetzel E, Green DR, Newmeyer DD (1997) Science 275:1132–1136 14. Yang J, Liu XS, Bhalla K, Kim CN, Ibrado AM, Cai J, Peng T, Jones DP, Wang X (1997) Science 275:1129–1132 15. Crompton M (1999) The mitochondrial permeability transition pore and its role in cell death. Biochem J 341:233–249 16. Lenaz G (1998) Quinone specificity of complex. Biochim Biophys Acta 1364:207–221 17. De Giorgi F, Lartigue L, Bauer MK, Schubert A, Grimm S, Hanson GT, Remington SJ, Youle RJ, Ichas F (2002) The permeability transition pore signals apoptosis by directing Bax translocation and multimerization. FASEB J 16:607–609 18. Matroule JY, Carthy CM, Granville DJ, Jolois O, Hunt DW, Piette J (2001) Mechanism of colon cancer cell apoptosis mediated by pyropheophorbide-a methylester photosensitization. Oncogene 20:4070–4084 19. Sugawara T, Lewen A, Gasche Y, Yu F, Chan PH (2002) Overexpression of SOD1 protects vulnerable motor neurons after spinal cord injury by attenuating mitochondrial cytochrome c release. FASEB J 16:1997–1999 20. Kowaltowski AJ, Castilho RF, Vercesi AE (2001) Mitochondrial permeability transition and oxidative stress. FEBS Lett 495:12–15
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Nantes et al.
21. Kowaltowski AJ, Vercesi AE, Fiskum G (2000) Bcl-2 prevents mitochondrial permeability transition and cytochrome c release via maintenance of reduced pyridine nucleotides. Cell Death Differ 7:903–910 22. Kowaltowski AJ, Vercesi AE (1999) Mitochondrial damage induced by conditions of oxidative stress. Free Radic Biol Med 26:463–471 23. Barros MH, Netto LE, Kowaltowski AJ (2003) H2O2 generation in Saccharomyces cerevisiae respiratory pet mutants: effect of cytochrome c. Free Radic Biol Med 35:179–188 24. Kowaltowski AJ, Turin J, Indig GL, Vercesi AE (1999) Mitochondrial effects of triarylmethane dyes. J Bioenerg Biomembr 31:581–590 25. Wuthrich K, Aviram I, Schejter A (1971) Structural studies of modified cytochromes c by nuclear magnetic resonance spectroscopy. Biochim Biophys Acta 253:98–103 26. Nicholls P (1974) Cytochrome c binding to enzymes and membranes. Biochim Biophys Acta 346:261–310 27. Pinheiro TJ (1994) The interaction of horse heart cytochrome c with phospholipid bilayers. Structural and dynamic effects. Biochimie 76:489–500 28. Nantes IL, Zucchi MR, Nascimento OR, Faljoni-Alario A (2001) Effect of heme iron valence state on the conformation of cytochrome c and its association with membrane interfaces. A CD and EPR investigation. J Biol Chem 276:153–158 29. Rytömaa M, Kinnunen PK (1995) Reversibility of the binding of cytochrome c to liposomes. Implications for lipid–protein interactions. J Biol Chem 270:3197–3202 30. Tuominen EK, Wallace CJ, Kinnunen PK (2002) Phospholipid–cytochrome c interaction: evidence for the extended lipid anchorage. J Biol Chem 277:8822–8826 31. Zucchi MR, Nascimento OR, Faljoni-Alário A, Prieto T, Nantes IL (2003) Modulation of cytochrome c spin states by lipid acyl chains: a continuous-wave electron paramagnetic resonance (CW-EPR) study of haem iron. Biochem J 370:671–678 32. Rytömaa M, Mustonen P, Kinnunen PKJ (1992) Reversible, nonionic, and pH-dependent association of cytochrome c with cardiolipinphosphatidylcholine liposomes. J Biol Chem 267:22243–22248 33. Rytömaa M, Kinnunen PKJ (1994) Evidence for two distinct acidic phospholipid-binding sites in cytochrome c. J Biol Chem 269:1770–1774 34. Pelletier H, Kraut J (1992) Crystal structure of a complex between electron transfer partners, cytochrome c peroxidase and cytochrome c. Science 258:1748–1755
35. Hatefi Y, Rieske JS (1967) Methods Enzymol 10:225–230 36. Margoliash E, Frohwirt N (1959) Spectrum of horse-heart cytochrome c. Biochem J 71:570–572 37. Sone N, Yoshida M, Hirata H, Kagawa Y (1977) J Biochem (Tokyo) 81:519–528 38. Kawai C, Prado FM, Nunes GLC, Di Mascio P, Carmona-Ribeiro AM, Nantes IL (2005) pH-dependent interaction of cytochrome c with mitochondrial mimetic membranes. The role of an array of positively charged amino acids. J Biol Chem 280:34709–34717 39. Margoliash E, Frohwirt N (1959) Spectrum of horse-heart cytochrome c. Biochem J 71:570–572 40. Bartlett GR (1959) Phosphorus assay in the column chromatography. J Biol Chem 234:466–468 41. King TE (1961) J Biol Chem 236:2342–2346 42. Yu CA, Yu L, King TE (1974) J Biol Chem 249:4905–4910 43. Yu CA, Yu L (1980) Resolution and reconstitution of succinate-cytochrome c reductase: preparations and properties of high purity succinate dehydrogenase and ubiquinol-cytochrome c reductase. Biochim Biophys Acta 591:409–420 44. Leung KH, Hinkle PC (1975) Reconstitution of ion transport and respiratory control in vesicles formed from reduced coenzyme Q-cytochrome c reductase and phospholipids. J Biol Chem 250:8467–8471 45. Beattie DS, Villalobo A (1982) Energy transduction by the reconstituted b-c1 complex from yeast mitochondria. Inhibitory effects of dicyclohexylcarbodiimide. J Biol Chem 257:14745–14752 46. Miki T, Orii Y, Mukohata Y (1987) A mechanism of respiratory control: studies with proteoliposomes containing cytochrome oxidase and bacteriorhodopsin. J Biochem 102: 199–209 47. Gutweniger H, Massari S, Beltrame M, Colonna R, Veronesse P, Ziche B (1977) Biochim Biophys Acta 459:216–224 48. Miki T, Orii Y (1986) Cytochrome c peroxidase activity of bovine heart cytochrome oxidase incorporated in liposomes and generation of membrane potential. J Biochem 100: 735–745 49. Redfeam ER (1967) Isolation and determination of ubiquinone. Methods Enzymol 10:381–384 50. Tsai A-L, Olson JS, And Palmer G (1987) The kinetics of reoxidation of yeast complex III. An evaluation of the Q-cycle. J Biol Chem 262:8677–8684
Study of Respiratory Cytochromes in Liposomes 51. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–275 52. Kawai C, Prado FM, Nunes GLC, Di Mascio P, Carmona-Ribeiro AM, Nantes IL (2005) pH-dependent interaction of cytochrome c with mitochondrial mimetic membranes. The role of an array of positively charged amino acids. J Biol Chem 280:34709–34717 53. Hunter DG, Frisken BJ (1998) Effect of extrusion pressure and lipid properties on the size and polydispersity of lipid vesicles. Biophys J 74:2996–3002 54. Nichols JW, Deamer DW (1980) Net protonhydroxyl permeability of large unilamellar liposomes measured by an acid–base titration technique. Proc Natl Acad Sciences (USA) 77:2038–2042
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55. Handbook of Chemistry and Physics (1983) part II 56. Schenkman S, Araujo PS, Dijkman R, Qina FH, Chaimovich H (1981) Effects of temperature and lipid composition on the serum albumin-induced aggregation and fusion of small unilamellar vesicles. Biochm Biophy Acta 649:633–641 57. Carmona-Ribeiro AM, Yoshida LS, Chaimovich H (1985) Salt effects on the stability of dioctadecyldimethylammonium chloride and sodium dihexadecyl phosphate vesicles. J Phys Chem 89:2928–2933 58. Grabowski EF, Morrison ID (1983) Particle size distributions from analyses of quase-elastic lightscattering data. In: Dahneke B (ed) Measurements of suspended particles by quase-elastic light-scattering, Wiley-Interscience, NY, p 7
Chapter 13 Use of Liposomes to Evaluate the Role of Membrane Interactions on Antioxidant Activity Salette Reis, Marlene Lúcio, Marcela Segundo, and José L.F.C. Lima Abstract Cellular membranes, which contain abundant phospholipids, such as phosphatidylcholine, are major targets subjected to the damage caused by free radicals. Cellular damage due to lipid oxidation is strongly associated with ageing, carcinogenesis and other diseases. In addition, lipid oxidation is an important deteriorative reaction in the processing and storage of lipid-containing foods. Liposomes have been used extensively as biological models for in vitro lipid oxidation studies. The resemblance between the liposomal and membrane bilayer core makes liposomes a very useful tool to investigate the significance of the antioxidant-membrane interactions for antioxidant activity. The antioxidant activity of a compound is strongly influenced by numerous factors including the nature of the lipid substrate, the hydrophilic– lipophilic balance of the antioxidant, the physical and chemical environments of the lipids, and various other interfacial interactions. Thus, compounds that are effective antioxidants in one model system or food matrix may be unsuitable in other systems. This chapter describes fluorescent probes-based methods commonly used for testing antioxidant activity in liposomes and stresses the need to combine antioxidant assays and drug-membrane interaction studies to get a better description of the antioxidants’ profile considering their location in lipid bilayer and their effect on membrane fluidity and consequently provide additional information to that obtained currently from assays performed in aqueous buffer media. Key words: Liposomes, Radical inducers, Peroxyl radicals, Hydroxyl radicals, Antioxidant activity, In vitro assays, Interactions with membranes
1. Introduction Reactive oxygen species (ROS), and reactive nitrogen species (RNS), are generated in living organisms through many pathways. Accumulation of reactive species in aerobic organisms is thought to cause oxidative damage of biological macromolecules such as DNA, proteins, and membrane lipids (1, 2). V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_13, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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In biological systems, the oxidation of membrane and lipoprotein lipids in circulation plays a central role in many pathologic processes (3–6), especially in the context of oxidative stress-related pathologies, including cardiovascular and neurodegenerative diseases (7–9). Furthermore, oxidation of lipids in food is also of great concern, since it leads to fatty acid decomposition and development of undesirable rancid odors and flavors, with a decrease in nutritional value and food safety (10–12). Therefore, there is an increasing interest in the efficacy of antioxidant activity of the many naturally occurring molecules in food and biological systems to minimize potential oxidative damage in vivo and/or to retard the oxidation of easily oxidizable materials in foods. While the antioxidant activity of many synthetic and naturally occurring compounds has been studied in solution and such results are extremely useful to provide an index for the intrinsic antioxidative activity, they bear little relevance to the multicellular systems, in which the tested compounds have to be metabolised and transported to the “reaction sites”. According to Halliwell and Gutteridge (13), the mechanisms of antioxidant action can include: (a) suppressing reactive species formation either by inhibition of enzymes or by chelating trace elements involved in free radical production; (b) scavenging reactive oxygen species; and (c) up-regulating or protecting antioxidant defence. Methods, which involve homogeneous solutions only assess free radicalscavenging activity. However, biological systems are inherently quite heterogeneous (6). There are both hydrophilic and hydrophobic domains in these systems which cannot be reproduced in homogeneous solutions. Consequently, the effectiveness of antioxidants in food or biological systems may depend not only on their chemical interactions with components of the oxidation pathway, but also on their location and partitioning into interfacial regions of the lipid membranes (14). It can thus be inferred that when the antioxidant of biological material is to be evaluated, the type and polarity of the system used as the substrate significantly affects the activity of the antioxidant in study (15, 16) and therefore, biological membranes may be regarded as more relevant substrates for evaluating antioxidant activity in food and in biological systems (17, 18). Examples of lipid-containing substrates for the study of antioxidants include the use of low-density lipoproteins (LDL) in the in vitro LDL oxidation assay, which has become popular for assessing the antioxidant activity of wines (19). Other lipid-containing substrates, such as erythrocytes (20), primary hepatocytes (21), microsomal membrane preparations (22–25) can also be used to evaluate the in vitro antioxidant activity of foods and phenolic compounds.
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In view of the complexity of biological membranes, much of the work described in the literature was performed in simplified model systems able to simulate a membrane environment (26). Model lipid oxidation systems included micelles (27, 28) and mostly lipid vesicles (liposomes). Liposomes, in particular, are relevant to the field of lipid oxidation studies for at least two reasons: (1) they constitute a simple and convenient system where the oxidation process can be reproduced with different levels of complexity allowing the analysis of water- and lipid-soluble antioxidants in the same system (29, 30) and (2) liposomes are currently being used for diagnostic and therapeutic purposes and lipid oxidation should be evaluated since it may impair their stability (29). Several studies indicated that a clear correlation exists between the study of antioxidant potency of various compounds in liposomes used as membrane models and the protective effects of these compounds against oxidation in biological systems (6). However, it is important to note that the possibility of gaining biologically relevant information on the potency of antioxidants in liposomes as membrane model systems cannot be assured without a thorough analysis of the results obtained and experimental conditions used. As a matter of fact, antioxidant effects depend on several factors, including the inducer of oxidation and its concentration; the composition, charge surface and fluidity of the liposomes. Ultimately antioxidant effects depend on the interactions of the compounds in study with the lipid membranes. Regarding these concerns, this chapter describes the use of fluorescent probes commonly used for testing antioxidant activity in liposomes and stresses the need to combine the antioxidant assays and drug-membrane interaction studies to get a better description of the antioxidants’ profile and provide additional information to that obtained currently from assays performed in aqueous buffer media. 1.1. Methods for Assessing Antioxidant Activity
The antioxidant activities of many compounds and biological fluids and extracts have been evaluated in liposome systems by various methods. In general, the oxidation is carried out in the presence and absence of an inducer and the extent of suppression of the rate of oxidation and duration of inhibition produced by antioxidant are measured. Some of the most commonly used methods for testing antioxidant activity measure the extent of oxidation from the oxygen uptake, peroxide formation, conjugated diene formation, formation of a coloured compound between TBA and malonaldehyde, chemiluminescence emission, and disappearance of fluorescence or ESR signal from a reference probe (31). Among the mentioned methods, the assays using molecular probes appear as more convenient avoiding the experimental complexity and the limitations of directly monitoring reaction kinetics of the inhibited autoxidation of lipids in the liposome model used.
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In general, the assays using probes have the following components: (a) a thermal radical generator to give a steady flux of peroxyl radicals in air-saturated solution; (b) a molecular probe (UV or fluorescence) for monitoring reaction progress; (c) antioxidant that competes with probes (substrates in this case) for the radicals and inhibits or retards the probe oxidation and (d) reaction kinetic parameters collected for antioxidant capacity quantification. The difference between the methods using probes lies mostly in the type of inducers used to initiate oxidation and the type of probes used to follow the oxidation progress which, in turn, depend on the aim of the study. Therefore, this chapter purposes the evaluation of antioxidant activity of any compounds using three different fluorescent probes specifically diphenylhexatriene propionic acid (DPH-PA); hexadecanoyl aminofluorescein (HDAF) and fluorescein. The degree of lipid oxidation is indirectly monitored by the oxidation of the probes with a consequent decay in their fluorescence intensity. The probes have different lipophilic properties, and consequently different locations (32–35) and thus, the antioxidant capacity of compounds can be related to the molecular distribution between aqueous and lipid media, and to other components of the interaction of the compounds with lipid membranes, namely, the effect in membrane surface charge; and the effect in membrane physical properties (Fig. 1). From the observation of Fig. 1 it becomes evident that the choice of a certain inducer depends on the aim of the study. If the researcher intends to study lipophilic antioxidants (e.g. a-tocopherol) or follow the oxidation of a lipophilic probe (e.g. DPH-PA) it is reasonable to choose a liposoluble radical initiator. In this way, AMVN is a synthetic azo-compound that dissociates spontaneously to form carbon-centered free radicals in the hydrophobic core of the bilayer which, in turn, produce peroxyl radicals when oxygen is present (6, 36). If the researcher intends to study hydrophilic compounds or follow the oxidation of hydrophilic probe (e.g. fluorescein) then it is preferable to choose a hydrophilic inducer like the commonly used water-soluble radical initiator, AAPH, which generates radicals in the aqueous phase of the liposomes. Metal ions are also used as hydrophilic inducers. However, some water-soluble antioxidants including ascorbic acid can become pro-oxidant in the presence of metal ions, depending on the antioxidant: metal ratio, so it is not considered desirable to use this method of generating radicals for the assessment of natural extracts where the concentration of different antioxidants will vary widely (17). If the studied compounds are soluble in the aqueous media but locate in the polar surface region of the lipid bilayers (e.g. trolox) or the oxidation is followed by a probe located both in polar and hydrophobic regions of the bilayer, it might be helpful
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FLUORESCEIN HO
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Fig. 1. Schematic illustration of the principle of the assays used to evaluate the antioxidant activity against peroxyl radical (ROO•) showing the location of the fluorescent probes fluorescein, hexadecanoyl aminofluorescein (HDAF) and diphenylhexatriene propionic acid (DPH-PA) in a liposome-aqueous media; the generation of the radical in the aqueous and liposome media, and the lipid-water partition of the antioxidant studied
to perform a comparative assay with both liposoluble and hidrosoluble radical initiators. 1.2. Methods for Assessing Membrane Interactions
As it has been previously said, the efficacy of the antioxidants in vivo is determined not only by the reactivity toward a radical, but also by their interaction with membrane lipid bilayers. Several aspects of this interaction are explored in this chapter and include: location and partitioning of the antioxidant into interfacial regions in liposome systems; as well as the antioxidant mobility at the microenvironment determined by membrane physical changes and also interactions between specific membrane surface charges and antioxidants of different charge types (31, 37–39) (Fig. 1).
1.2.1. Assessing Location and Partitioning of the Antioxidant into Interfacial Regions in Liposome Systems
The rate of scavenging depends on the chemical structure of the inhibitor, but it may also be promoted by concentrating the scavenger in the neighbourhood of the target lipid. It is for the latter reason that the precise positioning, orientation and partition of many of the currently available antioxidant compounds within
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the membrane, is also an important factor contributing to their scavenger effectiveness so when both antioxidant and free radical coincide in the same place, the antioxidant effect is greater (38). For a similar reason, the partition coefficient has also great importance because it determines the accessibility and interaction of antioxidants to the lipid radicals within the liposome (15). The type of radicals also determines the rate of scavenging, which will, in turn, be dependent on the antioxidant location. Peroxyl radicals are known to be highly polarized. As a consequence, such radicals formed initially in the hydrophobic region of membranes are expected to diffuse rapidly to the polar head group region near the aqueous phase where a water-soluble antioxidant could terminate them – diffusion-trapping mechanism (37). It is likely that compounds that are mostly located near the surface of membranes, where aqueous peroxyl radicals are easily trapped and more readily accessible, have better antioxidative effect in liposomes. On the other hand,•OH radicals, although are generated in the aqueous phase, diffuse easily across the bilayer and react with acyl chains causing their oxidation. In this case, the efficiency of an antioxidant to interrupt the oxidative sequence will be greater when located at the hydrophobic region of the bilayer, as the acyl chains are (38). The partition coefficient (Kp) can be determined using derivative spectroscopy and fluorescence-quenching techniques, which permit evaluating and comparing the extent of penetration and/ or interaction of antioxidants with membrane phospholipids. Moreover, fluorescence-quenching studies and zeta potential measurements can be used to preview the location of the antioxidants within the bilayer (40). 1.2.2. Assessing Membrane Physical Changes Induced by the Antioxidant
Antioxidants can act either by scavenging free radicals or by modifying their propagation across cell membranes. Several authors have demonstrated that free radical scavenger antioxidants could interact more efficiently with lipid radicals in a disordered lipid bilayer (41, 42). Then, the fluidizing effect of compounds could favor the known antioxidant capability and scavenging characteristics of these compounds (41, 42). Moreover, an agent effective against free radical oxidation may also act by altering the cell membrane in such a way that the free radical chain is unable to propagate efficiently (39). Modifications of membrane biophysical properties elicited by the antioxidants under study can be evaluated by fluorescence measurements of anisotropy (40).
1.2.3. Assessing Interactions Between Membrane Surface and Antioxidant Charge
It is important to consider the possible effect of conditions (e.g. pH) and the polarity of antioxidants as well as the charge type of the target membranes when employing antioxidants for medicinal purposes (37). According to this, zeta-potential measurements
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can be used to evaluate changes in membrane surface and thus obtain more information about the antioxidant’s binding behaviour (40).
2. Materials 2.1. Preparation of Liposomes
1. Egg l-a-phosphatidylcholine (EPC) (Sigma, St. Louis, MO) is stored at −20°C. 2. Hydration buffer: 10 mM Hepes, pH 7.4, 0.1 M NaCl. Store at room temperature (see Notes 1 and 2). 3. Chloroform and methanol. 4. Stainless-steel extruder (Lipex Biomembranes, Vancouver, BC, Canada), equipped with a circulating water bath. 5. Polycarbonate filters of three different sizes: 400, 250, and 100 nm (Nucleopore, Pleasanton, CA). 6. Malvern ZetaSizer 5000 (Malvern Instruments, Malvern, Worcestershire, UK) for quasielastic light scattering analysis.
2.2. Methods for Assessing Antioxidant Activity 2.2.1. Studies Using Fluorescein as a Fluorescent Probe in Aqueous and Liposome Media
1. Phosphate buffer: 75 mM KH2PO4, 0.1 M NaCl. Adjust pH to 7.4 with 10% KOH. Store at room temperature (see Note 2). 2. Fluorescein stock solution is prepared by dissolution of 3.77 mg fluorescein sodium salt (Aldrich, St. Louis, MO) in 25.00 mL of phosphate buffer to achieve a concentration of 0.42 mM and stored in dark conditions at 4°C. The fluorescein stock solution at such conditions can last several months. A fresh fluorescein working solution is prepared daily at 480 nM (studies in liposome media) and at 576 nM (studies in aqueous media) by dilution of the stock solution in the phosphate buffer (see Note 3). 3. The radical inducer 2, 2¢-azobis(2-amidinopropane) dihydrochloride (AAPH) (Fluka, St. Louis, MO) is completely dissolved in the phosphate buffer at 188 mM (510 mg of AAPH in 10.00 mL of buffer when studies are performed in liposome media) and at 36 mM (98 mg of AAPH in 10.00 mL of buffer when studies are performed in aqueous media) and is kept in an ice bath (see Notes 4 and 5). 4. Standard preparation: 0.5 g of Trolox (Carboxy-2, 5, 7, 8-tetramethyl-6-chromanol) (Fluka, St. Louis, MO) is dissolved in 10.00 mL of phosphate buffer (pH 7.4) to give a 200 mM stock solution. The stock solution is diluted with the same phosphate buffer to trolox working solutions in the range of 0–100 mM (see Note 6). 5. Sample preparation: pure compounds are directly dissolved in phosphate buffer (pH 7.4) to give a 200 mM stock solution.
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The stock solution is diluted with the same phosphate buffer to samples working solutions in the range of 0–100 mM. 6. 96-well polystyrene microplates International Inc, Bridgeport, NJ).
and
7. A FL600 microplate fluorescence Instruments, Inc., Winooski, VT). 2.2.2. Studies Using 5-Hexadecanoyl Aminofluorescein as a Fluorescent Probe
covers reader
(VWR (Bio-Tek
1. Phosphate buffer: 75 mM KH2PO4, 0.1 M NaCl. Adjust pH to 7.4 with 10% KOH. Store at room temperature (see Note 2). 2. HDAF stock solution is prepared by dissolution of 1.46 mg of 5-N-hexadecanoyl-aminofluorescein (HDAF; Aldrich, St. Louis, MO) in 10.00 mL of dimethylsulphoxide (DMSO) to achieve a concentration of 0.293 mM and stored in dark conditions at −20°C. 3. The radical inducer 2, 2¢-azobis(2-amidinopropane) dihydrochloride (AAPH) (Fluka, St. Louis, MO) is completely dissolved in the phosphate buffer at 545.5 mM (1.48 g of AAPH in 10.00 mL of buffer) and is kept in an ice bath (see Notes 4 and 5). 4. Standard preparation: 1 g of Trolox (Carboxy-2, 5, 7, 8-tetramethyl-6-chromanol) (Fluka, St. Louis, MO) is dissolved in 10.00 mL of phosphate buffer (pH 7.4) to give a 400 mM stock solution. The stock solution is diluted with the same phosphate buffer to trolox working solutions in the range of 0–200 mM. 5. Sample preparation: pure compounds are directly dissolved in phosphate buffer (pH 7.4) to give a 400 mM stock solution. The stock solution is diluted with the same phosphate buffer to samples working solutions in the range of 0–200 mM. 6. 96-well polystyrene microplates International Inc, Bridgeport, NJ).
and
covers
(VWR
7. A FL600 microplate fluorescence reader (Bio-Tek Instruments, Inc., Winooski, VT). 2.2.3. Studies Using Diphenylhexatriene Propionic Acid as a Fluorescent Probe
1. Hepes buffer: 10 mM Hepes, pH 7.4, 0.1 M NaCl. Store at room temperature (see Note 2). 2. DPH-PA stock solution is prepared by dissolution of 2.92 mg of 3-(p-(6-phenyl)-1,3,5-hexatrienyl)-phenylpropionic acid (DPH-PA; Invitrogen Corporation, Carlsbad, California, USA) in 5.00 mL of tetrahydrofuran (THF) to achieve a concentration of 1.92 mM and stored in dark conditions at −20°C (see Note 7). 3. The radical inducer 2, 2¢-azobis(2-amidinopropane) dihydrochloride (AAPH) (Fluka, St. Louis, MO) is completely dissolved in the phosphate buffer at 67.5 mM (183 mg of AAPH in 10.00 mL of buffer and is kept in an ice bath (see Notes 4 and 5)).
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4. Standard preparation: 1 g of Trolox (Carboxy-2, 5, 7, 8tetramethyl-6-chromanol) (Fluka, St. Louis, MO) is dissolved in 10.00 mL of phosphate buffer (pH 7.4) to give a 400 mM stock solution. The stock solution is diluted with the same phosphate buffer to trolox working solutions in the range of 0–200 mM. 5. Sample preparation: pure compounds were directly dissolved in phosphate buffer (pH 7.4) to give a 1.35 mM stock solution. The stock solution was diluted with the same phosphate buffer to samples working solutions in the range of 0–675 mM. 6. Steady-state spectrofluorimeter (Perkin-Elmer LS 50B). 2.3. Methods for Assessing Membrane Interactions
1. Materials for liposome preparation (see Subheading 2.1) 2. Samples: pure compounds are directly dissolved in Hepes buffer (pH 7.4). 3. DPH stock solution is prepared by dissolution of 5.8 mg of 1,6-diphenyl-1,3,5-hexatriene (DPH; Invitrogen Corporation, Carlsbad, California, USA) in 5.00 mL of tetrahydrofuran (THF) to achieve a concentration of 5 mM and stored in dark conditions at −20°C (see Note 7). 4. UV-VIS spectrophotometer (Perkin-Elmer Lambda 45, Waltham, Massachusetts, USA). 5. Steady-state spectrofluorimeter Waltham, Massachusetts, USA).
(Perkin-Elmer
LS
50B,
6. Malvern ZetaSizer 5000 (Malvern Instruments, Malvern, Worcestershire, UK) for quasielastic light scattering analysis.
3. Methods 3.1. Preparation of Liposomes
1. Liposomes suspension is prepared by dissolving EPC in chloroform/methanol (9:1) (10–20 mg lipid per mL of organic solvent) in a round-bottomed flask. 2. Solvent is removed under reduced pressure on a rotary evaporator with the water bath set at 30°C. N2 is introduced to re-establish atmospheric pressure and the flask covered in aluminium foil. A pump is then used to keep the flask under a vacuum of <0.5 mmHg for at least 3 h. After evaporation, the vacuum is released while introducing N2. A dry thin lipid film is obtained in the round-bottomed flask walls. 3. Hydration of the thin lipid film is made by addition of suitable hydration buffer (see Subheading 2.1) for approximately 30 min at 25°C (see Note 8). The proportion lipid: buffer should be 2.1 mg/mL, which gives a lipid concentration of 3 mM (if the relative molecular mass of EPC is taken as 700).
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4. The flask is vortexed at room temperature (see Note 8) for 20 min, producing a homogeneous white suspension of multilamellar liposomes (MLV). The multilamellar suspension is then shaken while being sonicated for 30 s in an ultrasonic bath. This ensures complete recovery of the lipid from the flask walls (see Note 9). 5. The MLV suspension is transferred to an extrusion device maintained at 25°C by a circulating water bath to form large unilamellar vesicles (LUV) (see Note 8). Prior to extrusion through the final pore size, MLV suspensions are disrupted by pre-extruding the suspension through a larger pore size (typically, five times through 400 nm pore size, then five times through 250 nm pore size) (see Notes 10 and 11). Finally, extrude the lipid suspension ten times through 100 nm pore size (see Note 12). 6. Size distribution of the extruded vesicles should be determined by quasielastic light scattering analysis using a heliumneon laser (633 nm) as a source of incident light, operating at a scattering angle of 90° and a temperature of 25°C. Extrusion through filters with 100 nm pores typically yields large, unilamellar vesicles (LUV) with a mean diameter of 120–140 nm. Mean particle size depends on lipid composition but is quite reproducible from batch to batch. 3.2. Methods for Assessing Antioxidant Activity
The methods for assessing antioxidant activity described in this chapter are based on the decay in probes fluorescence intensity monitored over time. In general, samples, controls, and standard (Trolox of four or five different concentrations for construction of a standard curve) are mixed with the fluorescent probe in aqueous or liposome media and incubated at constant temperature (37°C) before AAPH solution is added to initiate the reaction. As the reaction progresses, the probe is oxidized and fluorescence intensity decreases. In the presence of antioxidant, the fluorescence decay is inhibited. Data obtained are converted to relative fluorescence values, by dividing the fluorescence intensity at a given time by the fluorescence intensity at 0 min. The ability of the compounds studied to act as antioxidants is analysed by determining their IC50 values, which are defined as the concentration (in mM) of each compound required to obtain a ratio defined by (1) equal to 50%: AUCantioxidant/trolox − AUCblank AUCblank
× 100
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where AUC is the area under the curve obtained for the sample and blank assays.
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1. For the assays in aqueous media, a 96 well plate is prepared (maximum well volume = 320 mL) with a reaction mixture (300 mL) containing the following volumes of reagents in potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4): 25 mL of fluorescein working solution (see Subheading 2.1, item 1); 150 mL of samples or standard (trolox) working solutions. Blanks are made with the radical initiator and without the sample or standard replacing sample or standard by 150 mL of potassium phosphate buffer (see Note 13). Controls are made without the radical initiator replacing it by potassium phosphate buffer: 25 mL of fluorescein working solution; 150 mL of samples or standard (trolox) working solutions and 125 mL of potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4) instead of AAPH solution (see Note 14). For the assays in liposome media, a 96 well plate is prepared (maximum well volume = 320 mL) with a reaction mixture (300 mL) containing the following volumes of reagents in potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4): 80 mL of LUV suspension (see Subheading 3.1); 30 mL of fluorescein working solution (see Subheading 2.1, item 1); 150 mL of samples or standard (trolox) working solutions. Blanks are made replacing sample by 150 mL of potassium phosphate buffer (see Note 13). Controls are made without the radical initiator replacing it by potassium phosphate buffer: 80 mL of LUV; 30 mL of fluorescein-working solution; 150 mL of samples or standard (trolox) working solutions and 40 mL of potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4) instead of AAPH solution (see Note 14). 2. The plate was covered with a lid and incubated in the preheated (37°C) microplate reader (Synergy HT, Bio-Tek, Winooski, VT, USA) for 10 min with a 3 min shaking during this time. 3. Incubation is followed by the addition of radical initiator solution (see Subheading 2.1, item 1); to all wells (samples, standard trolox and blanks) except for the wells containing controls: 125 mL of AAPH solution (in assays in aqueous media) or 40 mL of AAPH solution (in assays in liposome media). Thus, the final concentration for each well in aqueous media is: 48 nM of fluorescein; increasing concentrations of samples or standard trolox (0–50 mM) and 15 mM of AAPH. In liposome media, the final concentrations for each well is: 800 mM of LUV suspension; 48 nM of fluorescein; increasing concentrations of samples or standard trolox (0–50 mM) and 25 mM of AAPH. 4. After addition of AAPH, the plate is immediately transferred to the plate reader (see Note 15), and the fluorescence intensity was
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measured every minute for 180 min at previous conditions (pH 7.4, 37°C) with fluorescence filters for an excitation wavelength of 485 ± 20 nm and an emission wavelength of 528 ± 20 nm. The plate reader was controlled by software KC4 3.0. 3.2.2. Studies Using 5-Hexadecanoyl Aminofluorescein as a Fluorescent Probe
1. Liposomes labelled with HDAF probe are prepared in a round-bottomed flask by dissolving EPC in chloroform/ methanol (9:1) (10–20 mg lipid per mL of organic solvent) and adding an aliquot of the probe to the organic phase so that the ratio lipid to probe is 6,000:1 (see Note 16). Typically, 21 mg of EPC and 20 mL of HDAF stock solution are added to the chloroform/methanol (9:1) mixture and dried together under reduced pressure on a rotary evaporator with the water bath set at 30°C as previously described in preparation of liposomes method (see Subheading 3.1). 2. Hydration of the thin lipid film is made by addition of potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4) as hydration buffer (see Subheading 2.1, item 1) for approximately 30 min at 25°C (see Note 8). The proportion lipid: buffer should be 2.1 mg/mL, which gives a lipid concentration of 3 mM (if the relative molecular mass of EPC is taken as 700) and a probe concentration of 0.5 mM. After labelling the lipid with the probe and hydration of the thin lipid film, the preparation of LUV follows the procedure previously described in the preparation of liposomes method (see Subheading 3.1). 3. A 96 well plate is prepared (maximum well volume = 320 mL) with a reaction mixture (300 mL) containing: 80 mL of HDAF labelled LUV and 110 mL of samples or standard (trolox) working solutions. Blanks are made with the radical initiator and without the sample or standard replacing sample or standard by 110 mL of potassium phosphate buffer (see Note 13). Controls are made without the radical initiator replacing it by potassium phosphate buffer: 80 mL of HDAF labelled LUV; 110 mL of samples or standard (trolox) working solutions and 110 mL of potassium phosphate buffer (75 mM, I = 0.1 M, pH = 7.4) instead of AAPH solution (see Note 14). 4. The plate is covered with a lid and incubated in the preheated (37°C) microplate reader (Synergy HT, Bio-Tek, Winooski, VT, USA) for 10 min with a 3 min shaking during this time. 5. Incubation is followed by the addition of 110 mL of radical initiator solution (see Subheading 2.1, item 1); to all wells (samples, standard trolox and blanks) except for the wells containing controls. Thus, the final concentrations for each well in aqueous media are: 800 mM of LUV suspension 0.13 mM of HDAF; increasing concentrations of samples or standard trolox (0–200 mM) and 200 mM of AAPH.
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6. After addition of AAPH, the plate is immediately transferred to the plate reader (see Note 15), and the fluorescence intensity was measured every minute for 360 min at previous conditions (pH 7.4, 37°C) with fluorescence filters for an excitation wavelength of 485 ± 20 nm and an emission wavelength of 521 ± 20 nm. The plate reader was controlled by software KC4 3.0. 3.2.3 Studies Using Diphenylhexatriene Propionic Acid (DPH-PA) as a Fluorescent Probe
1. Liposomes labelled with DPH-PA probe are prepared in a round-bottomed flask by dissolving EPC in chloroform/ methanol (9:1) (10–20 mg lipid per mL of organic solvent) and adding an aliquot of the probe to the organic phase so that the ratio lipid to probe was 300:1 (see Note 16). Typically, 10 mg of EPC and 25 mL of DPH-PA stock solution are added to the chloroform/methanol (9:1) mixture and dried together under reduced pressure on a rotary evaporator with the water bath set at 30°C as previously described in the preparation of liposomes method (see Subheading 3.1). 2. Hydration of the thin lipid film is made by addition of Hepes buffer (10 mM, I = 0.1 M, pH = 7.4) as hydration buffer (see Subheading 2.1, item 1) for approximately 30 min at 25°C (see Note 8). The proportion lipid: buffer should be 1.05 mg/mL. After labelling the lipid with the probe and hydration of the thin lipid film, the preparation of LUV follows the procedure previously described in the preparation of liposomes method (see Subheading 3.1). 3. An aliquot of 1 mL of the LUV suspension, containing 1.5 mM of EPC and 5 mM of DPH-PA, is incubated in a fluorimetric cuvette placed in a thermostatted holder (37°C) for 5 min with continuous stirring and away from light. An aliquot (400 mL) of samples or standard (trolox) working solutions is added to the mixture and incubated for 5 min. Blanks are made with the radical initiator and without the sample or standard replacing sample or standard by 400 mL of Hepes buffer (see Note 13). Controls are made without the radical initiator replacing it by Hepes buffer: 1 mL of DPH-PA labelled LUV; 400 mL of samples or standard (trolox) working solutions and 400 mL of Hepes buffer (10 mM, I = 0.1 M, pH = 7.4) instead of AAPH solution (see Note 14). 4. Incubation is followed by the addition of 400 mL of radical initiator solution (see Subheading 2.1, item 1); to the cuvette (in the assays of samples, standard trolox and blanks) except for the control assays (see Note 17). Thus, the final concentrations for each assay are: 800 mM of LUV suspension 2.7 mM of DPH-PA; increasing concentrations of trolox and samples (0–150 mM) and 15 mM of AAPH.
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5. After addition of AAPH, the fluorescence measurements are immediately started (see Note 15), and the fluorescence intensity is monitored every minute for 60 min at previous conditions (pH 7.4, 37°C) using a steady-state spectrofluorimeter (Perkin-Elmer LS 50B) at excitation and emission wavelengths of 399 and 435 nm, respectively. 3.3. Methods for Assessing Membrane Interactions 3.3.1. Determination of Lipid/Water Partition Coefficient by Derivative Spectrophotometry
The extent of interaction of a solute with a microheterogeneous system is evaluated in a quantitative way from its partition coefficient. The octanol/water biphasic system is traditionally used to evaluate partition coefficients, which are then extrapolated to biomembrane/aqueous phase systems, this being an oversimplification. Instead, it is advisable to use liposomes as biomembranes models and quantify the concentration of the solute in one of the phases (aqueous or lipidic), or both. UV–Vis absorption spectrophotometric techniques are usually based on the change of an absorption parameter upon incorporation of compounds into membranes, which permits the determination of partition coefficient values from the following equation (42, 43): Abs f = Abs a +
(Absm − Absa )K p (L ) Vϕ 1 + K p (L ) V ϕ
(2)
where Absa is the absorbance of the compound in buffer solution without lipid; Absf is the absorbance of the compound in liposome suspension with different lipid concentrations; Absm is the absorbance calculated assuming that all compound is membrane bound; [L] is the lipid concentration and Vf is the lipid molar volume (for EPC, Vf = 0.688 L/mol and the mean molecular weight is 700 (44)). Despite the overall simplicity of (2), its practical application is usually limited to systems with low light scattering background signals. When the direct application of this spectrophotometric method is prevented by high background signals, caused by the presence of liposomes or cells, the problem can be minimized by the use of second derivative spectrophotometry with Absf replaced by Df = (dnAbs)/(dln). The second derivative spectra are thus calculated from the recorded absorption spectra of a compound in the presence of increasing lipid concentration after blank subtraction. Then, the partition coefficient is calculated by fitting (2) to the experimental data (Df versus [L]), using a non-linear regression method, where the adjustable parameter is Kp (44). Second derivative spectrophotometry technique has the additional advantage of analysing the signals originated from both phases (aqueous and lipidic) with no demand for their physical separation, which may be laborious and may result in equilibria perturbation.
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The procedure for the determination of lipid/water partition coefficient of an antioxidant by derivative spectrophotometry, is the following: 1. LUV suspension with a final concentration of 1.5 mM is prepared by the previously described thin lipid film hydration method (see Subheading 3.1) 2. Samples (total volume of 1,800 mL) are prepared in 2 mL Eppendorfs containing in Hepes buffer (Hepes: 10 mM, I = 0.1 M, pH 7.4): 80 mL of compound in study and increasing volumes (0–1,200 mL) of LUV suspension. This gives a series of samples containing a fixed concentration of studied compound (see Note 18) and increasing concentrations of LUV suspension (0–1,000 mM). 3. Corresponding reference suspensions are prepared identically as samples, but without compounds in study which are replaced by Hepes buffer (10 mM, I = 0.1 M, pH 7.4). 4. The resulting suspensions of samples and references are vortexed and incubated in the dark for 1 h. 5. The absorbance spectra of each sample and reference are recorded in a UV spectrometer equipped with a constant temperature cell holder at 25.0 ± 0.1°C in 1-cm path length cuvettes with adequate slits. 3.3.2. Assessing Membrane Location by Fluorescence Quenching
Fluorescence-quenching studies can be used to reveal the membrane location of compounds, which will behave as quenchers of a fluorescent probe inserted in the lipid bilayer (45). The extent of quenching is expressed by the Stern-Volmer equation (45): I0 app − 1 = K SV [Q ] I
(3)
app
where K SV is the apparent Stern-Volmer constant, [Q] is the total quencher concentration, I0 and I are the fluorescence intensity values in the absence and in the presence of the quencher (compound which location is being studied). High values of Stern-Volmer constants mean that the location of the quenching compound is very similar to the location of the fluorescent probe. According to this, it is required that the molecular location of the probe within membranes is known with certainty. This chapter describes the use of DPH probe (see Note 19) to assess membrane location of antioxidants using the following procedure: 1. Liposomes labelled with DPH probe are prepared in a roundbottomed flask by dissolving EPC in chloroform/methanol (9:1) (10–20 mg lipid per mL of organic solvent) and adding an aliquot of the probe to the organic phase so that the ratio lipid to probe was 300:1 (see Note 16). Typically, 10 mg of EPC and 9.5 mL of DPH-PA stock solution are added to the
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chloroform/methanol (9:1) mixture and dried together under reduced pressure on a rotary evaporator with the water bath set at 30°C as previously described in the preparation of liposomes method (see Subheading 3.1). 2. Hydration of the thin lipid film is made by addition of Hepes buffer (10 mM, I = 0.1 M, pH = 7.4) as hydration buffer (see Subheading 2.1, item 1) for approximately 30 min at 25°C (see Note 8). The proportion lipid: buffer should be 1.05 mg/ mL. After labelling the lipid with the probe and hydration of the thin lipid film, the preparation of LUV follows the procedure previously described in the preparation of liposomes method (see Subheading 3.1). The DPH-labelled LUV have a final EPC concentration of 1.5 mM and DPH of 5 mM. 3. Samples (total volume of 1,800 mL) are prepared in 2 mL Eppendorfs containing: 600 mL of DPH-labelled LUV suspension and increasing volumes (0–1,200 mL) of Hepes buffer solutions (Hepes: 10 mM, I = 0.1 M, pH 7.4) of compounds in study. This gives a series of samples containing a fixed concentration of DPH-labelled LUV suspension (500 mM of lipid and 1.7 mM of DPH) and increasing concentrations of studied compounds (see Note 18). 4. Corresponding reference suspension is prepared identically as sample, but without compound in study which is replaced by Hepes buffer (10 mM, I = 0.1 M, pH 7.4). 5. The resulting suspensions of reference and samples are vortexed and incubated in the dark for 1 h. 6. The fluorescent spectra of each reference and sample are recorded in a steady-state fluorescence spectrometer equipped with a constant temperature cell holder at 25.0 ± 0.1°C in 1-cm path length cuvettes with adequate excitation and emission slits. Excitation and emission wavelength is set to 361 and 432 nm, respectively (see Note 20). 3.3.3. Assessing Membrane Physical Changes Induced by the Antioxidant
Steady-state fluorescence anisotropy is a technique that assesses the range of rotational motion of the membrane-associated fluorescent probe, in this case DPH, during the lifetime of its excited electronic state (46). Using this technique, DPH probe is automatically excited with vertically polarized light and resulting fluorescence intensities are recorded with the analyzing polarizer oriented parallel (IVV) and perpendicular (IVH) to the excitation polarizer allowing the determination of steady-state fluorescence anisotropy for labelled samples by the following equation (45): rss =
I VV − GI VH I VV + 2GI VH
(4)
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where G = IHV/IHH is the grating correction factor (45).The greater the extent of probe rotation during its excited state lifetime, the smaller will be the observed fluorescence anisotropy (rss), to the extent that rss = 0 for complete DPH reorientation. The probe rotational motions are tightly coupled to the phospholipid fatty acid chain fluctuations, which provide a measure of membrane “fluidity” (45). To assess membrane physical changes induced by antioxidant, liposomes labelled with DPH probe are incubated with increasing concentrations of the antioxidant tested and the steady-state fluorescence anisotropy is measured. The procedure is the following: 1. Liposomes labelled with DPH probe, reference and samples are prepared in the same way as described for fluorescencequenching measurements method (see Subheading 3.3.2). 2. The resulting suspensions of reference and samples are vortexed and incubated in the dark for 1 h. 3. The steady-state anisotropy of each reference and sample is recorded in a steady-state fluorescence spectrometer equipped with a constant temperature cell holder at 25.0 ± 0.1°C in 1-cm path length cuvettes with adequate excitation and emission slits. Excitation and emission wavelength is set to 361 and 432 nm, respectively (see Note 20). 3.3.4. Assessing Interactions Between Membrane Surface and Antioxidant Charge
1. LUV suspension with a final concentration of 1.5 mM is prepared by the previously described thin lipid film hydration method (see Subheading 3.1). 2. Samples (total volume of 1,800 mL) are prepared in 2 mL Eppendorfs containing in Hepes buffer (10 mM, I = 0.1 M, pH 7.4): 600 mL of LUV suspension and increasing volumes (0–1,200 mL) of Hepes buffer solutions (10 mM, I = 0.1 M, pH 7.4) of compounds in study. This gives a series of samples containing a fixed concentration of LUV suspension (500 mM of EPC) and increasing concentrations of studied compounds (see Note 18). 3. Corresponding reference suspension is prepared identically as samples, but without compounds in study which are replaced by Hepes buffer (10 mM, I = 0.1 M, pH 7.4). 4. The resulting suspensions of samples and references are vortexed and incubated in the dark for 1 h. 5. The zeta-potential (z-potential) values of the membrane vesicles, with and without incorporated compound (samples and reference) are determined at pH 7.4 (Hepes buffer) at 25.0 ± 0.1°C, by quasi-elastic light scattering analysis with a 90° scattering angle. The values for the viscosity and refractive index are taken as 0.890 and 1.330 cP, respectively.
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4. Notes 1. Hepes buffer interferes with fluorescent probes fluorescein and its derivative 5-hexadecanoyl aminofluorescein (HDAF). Whenever using these probes, prepare the liposomes with phosphate buffer: 75 mM KH2PO4 (Merck, Darmstadt, Germany), 0.1 M NaCl. Adjust pH to 7.4 with 10% KOH. 2. All water used for buffer preparations is double-deionised water (conductivity smaller than 0.1 mS/cm). Buffer solutions are typically filtered using standard MF-Millipore membrane filters (0.45 mm) (Millipore, Bedford, MA). 3. The fluorescence of fluorescein is highly pH dependent and thus pH of the solutions must be carefully monitored. 4. AAPH powder should be stored at room temperature in a desiccator. Buy small bottles as it may decline in quality (oxidation induction requires higher concentrations of AAPH) after opening. 5. AAPH is an azo-compound that undergoes spontaneous decomposition producing carbon-centered radicals, which in turn produce peroxyl radicals when oxygen is present, with a rate primarily determined by temperature. Therefore, it is fundamental that the AAPH solution is kept in the ice bath before and after use. AAPH solution should be discarded within 8 h. 6. Trolox has advantages over other active antioxidants (e.g. vitamin E), which are only lipid-soluble. For example, Trolox does not have to be incorporated into the lipid membrane by solvent extraction and co-evaporation methods; it can be added directly to the intact system. This makes it convenient for studies on natural biological systems and for quantitative studies on model systems (47). However, vitamin E can also be used as a lipophilic standard compound and in these cases, vitamin E is co-dissolved with the lipids in chlorophorm: methanol during the preparation of the thin lipid film. 7. Long-term storage of stock solutions of DPH-PA and DPH in THF is not recommended because of possible peroxide formation in that solvent. 8. The lipid suspension should be kept above the phase transition temperature (Tm) of the lipid during hydration and extrusion. Attempts to extrude below the Tm will be unsuccessful as the membrane has a tendency to foul with rigid membranes which cannot pass through the pores. In the case of EPC, it is possible to work at room temperature inasmuch as the transition temperature for EPC ranges
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from −15 to −7°C (29). If other lipids are used, care should be taken to work always with a controlled temperature well above the Tm of the lipid used. 9. In some cases, if the complete recovery of the lipid from the flask walls is difficult, it might be useful to include glass beads while shaking the multilamellar suspension. 10. MLV suspensions are disrupted either by several freeze-thaw cycles or by pre-extruding the suspension through a larger pore size. The later process is preferred to the freeze–thawing procedure. Indeed, liposome bilayers are damaged by internal ice formation during freezing. Damaged bilayers will reassemble due to the “hydrophobic effect” and form new liposomes possibly of a different size, which may involve fusion with other liposomes. Each freeze–thaw cycle would provide an opportunity for this process to occur, both in previously unaffected liposomes and those damaged by a previous freeze–thaw cycle. 11. Extrusion is a technique in which the lipid suspension is forced, under inert (N2) atmosphere, through a polycarbonate filter with a defined pore size to yield LUV having a diameter near the pore size of the filter used. 12. To prevent the fouling of membranes and improve the homogeneity of the size distribution of the final suspension, the use of two-stacked filters is recommended. 13. Care must be taken to ensure homogeneity of each dilution by thorough mixing at each stage of addition of solutions through repeated aspiration and dispensing volumes with micropipette. To avoid human errors, it is advisable to use an eight-channel micropipette. 14. Controls should be prepared before addition of AAPH to reaction mixtures and are important to evaluate the photostability of the fluorescent probe. Fluorescein in controls is exposed to excitation light of the assay (491 nm) in the absence of AAPH over a 180 min period and there should be no showing of any significant fluorescence intensity changes over the time of the assay meaning that fluorescein, under such conditions, is photostable. 15. AAPH is the radical initiator, and the oxidation starts immediately after its addition to reaction mixture. Therefore, this initiator must be the last reagent added and its addition must be as quick as possible following immediate start of the measurement. 16. The ratio of lipid to probe should be always greater than 100:1 to prevent changes in the structure of the liposomes (40).
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17. While stirring, the fluorescence time scan is started simultaneously with the addition of radical initiator solution with a 500 mL syringe through the hole located at the top of the sample compartment. This procedure avoids the exposure of sample to light while ensuring a better homogeneity of the initiator with the sample mixture. 18. The concentrations of studied compounds should be used in the range over which Beer’s law is obeyed. 19. DPH probe was chosen because of its advantageous spectral and structural properties. The high extinction coefficient and fluorescence quantum yield in non-aqueous solvents make it possible to use very small amounts of the probe in biological studies, thus minimizing perturbations and making DPH particularly attractive for obtaining dynamic and structural information about membrane bilayers (48). Additionally, the knowledge of molecular location of DPH makes it a commonly employed probe to explore the membrane hydrocarbon chain region without a direct and significant influence on membrane properties close to the membrane-water interface (34, 49). 20. Fluorescence values are corrected for light scattering contributions by subtraction of intensities from unlabeled samples at the same conditions. These contributions are always negligible (less than 0.5%). References 1. Halliwell B, Gutteridge JMC (1990) Role of free radicals and catalytic metal ions in human disease: an overview. In: Packer L and Glazer AN (eds) Methods in enzymology. Academic, San Diego, NY 2. Davies MJ, Dean RT (1997) The pathology of protein oxidation. In: Radical-mediated protein oxidation. Oxford Science Publications, Oxford, UK 3. Mattson MP (2004) Metal-catalyzed disruption of membrane protein and lipid signaling in the pathogenesis of neurodegenerative disorders. Ann N Y Acad Sci 1012:37–50 4. Poon HF, Calabrese V, Scapagnini G, Butterfield DA (2004) Free radicals: key to brain aging and heme oxygenase as a cellular response to oxidative stress. J Gerontol A Biol Sci Med Sci 59:478–493 5. Schroepfer GJ (2000) Oxysterols: modulators of cholesterol metabolism and other processes. Physiol Rev 80:361–554 6. Schnitzer E, Pinchuk I, Lichtenberg D (2007) Peroxidation of liposomal lipids. Eur Biophys J 36:499–515
7. van Ginkel G, Sevanian A (1994) Lipid peroxidation-induced membrane structural alterations. Methods Enzymol 233:273–288 8. Mason RP, Walter M, Mason P (1997) Effect of oxidative stress on membrane structure: small-angle X-ray diffraction analysis. Free Radic Biol Med 23:419–425 9. Spiteller G (2003) Are lipid peroxidation processes induced by changes in the cell wall structure and how are these processes connected with diseases? Med Hypotheses 60:69–83 10. Al-Ismail KM, Aburjai T (2004) Antioxidant activity of water and alcohol extracts of chamomile flowers, anise seeds and dill seeds. J Sci Food Agric 84:173–178 11. Viljanen K, Kylli P, Kivikari R, Heinonen M (2004) Inhibition of protein and lipid oxidation in liposomes by berry phenolics. J Agric Food Chem 52:7419–7424 12. Diaz M, Decker EA (2004) Antioxidant mechanisms of caseinophosphopeptides and casein hydrolysates and their application in ground beef. J Agric Food Chem 52:8208–8213
Role of Membrane Interactions on Antioxidant Activity 13. Halliwell B, Gutteridge JMC (1998) Free radicals in biology and medicine. Oxford University Press, Oxford, UK 14. Baublis A, Decker EA, Clydesdale FM (2000) Antioxidant effect of aqueous extracts from wheat based ready-to-eat breakfast cereals. Food Chem 68:1–6 15. Hassimotto NMA, Genovese MI, Lajolo FM (2005) Antioxidant activity of dietary fruits, vegetables, and commercial frozen fruit pulps. J Agric Food Chem 53:2928–2935 16. Yi O, Jovel EM, Towers GHN, Wahbe TR, Cho D (2007) Antioxidant and antimicrobial activities of native Rosa sp from British Columbia. Canada Int J Food Sci Nutr 58: 178–189 17. Roberts WG, Gordon MH (2003) Determination of the total antioxidant activity of fruits and vegetables by a liposome assay. J Agric Food Chem 51:1486–1493 1 8. Yen WJ, Chang LW, Duh PD (2005) Antioxidant activity of peanut seed testa and its antioxidative component, ethyl protocatechuate. LWT-Food Sci Technol 38:193–200 19. Frankel EN, Waterhouse AL, Teissedre PL (1995) Principal phenolic phytochemicals in selected California wines and their antioxidant activity in inhibiting oxidation of human lowdensity lipoproteins. J Agric Food Chem 43:890–894 20. Tedesco I, Russo GL, Nazzaro F, Russo M, Palumbo R (2001) Antioxidant effect of red wine anthocyanins in normal and catalaseinactive human erythrocytes. J Nutr Biochem 12:505–511 21. Morel I, Abalea V, Sergent O, Cillard P, Cillard J (1998) Involvement of phenoxyl radical intermediates in lipid antioxidant action of myricetin in iron-treated rat hepatocyte culture. Biochem Pharmacol 55: 1399–1404 22. Daglia M, Papetti A, Gregotti C, Berte F, Gazzani G (2000) In vitro antioxidant and ex vivo protective activities of green and roasted coffee. J Agric Food Chem 48:1449–1454 23. Mora A, Paya M, Rios JL, Alcaraz MJ (1990) Structure-activity-relationships of polymetho xyflavones and other flavonoids as inhibitors of nonenzymatic lipid-peroxidation. Biochem Pharmacol 40:793–797 24. Plumb GW, Chambers SJ, Lambert N, Wanigatunga S, Williamson G (1997) Influence of fruit and vegetable extracts on lipid peroxidation in microsomes containing specific cytochrome P450. Food Chem 60:161–164
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25. van Acker SABE, van den Berg D-J, Tromp MNJL, Griffioen DH, van Bennekom WP, van der Vijgh WJF, Bast A (1996) Structural aspects of antioxidant activity of flavonoids. Free Radic Biol Med 20:331–342 26. de Beer D, Joubert E, Gelderblom WCA, Manley M (2005) Antioxidant activity of South African red and white cultivar wines and selected phenolic compounds: in vitro inhibition of microsomal lipid peroxidation. Food Chem 90:569–577 27. Roginsky V, Barsukova T (2001) Chainbreaking antioxidant capability of some beverages as determined by the clark electrode technique. J Med Food 4:219–229 28. Shi H, Noguchi N, Niki E (1999) Comparative study on dynamics of antioxidative action of alpha-tocopheryl hydroquinone, ubiquinol, and alpha-tocopherol against lipid peroxidation. Free Radic Biol Med 27:334–346 29. Goñi MF, Alonso A (1989) Studies of phospholipid peroxidation in liposomes. In: CRC handbook of free radicals and antioxioxidants in biomedicine. CRC, Boca Raton, FL 30. Murakami M, Yamaguchi T, Takamura H, Matoba T (2002) A comparative study on the various in vitro assays of active oxygen scavenging activity in foods. J Food Sci 67:539–541 31. Niki E, Noguchi N (2000) Evaluation of antioxidant capacity. What capacity is being measured by which method? Life 50:323–329 32. Fernandes E, Costa D, Toste SA, Lima JLFC, Reis S (2004) In vitro scavenging activity for reactive oxygen and nitrogen species by nonsteroidal anti-inflammatory indole, pyrrole, and oxazole derivative drugs. Free Radic Biol Med 37:1895–1905 33. Kachel K, Asuncion-Punzalan E, London E (1998) The location of fluorescent probes with charged groups in model membranes. Biochim Biophys Acta 1374:63–76 34. Kaiser RD, London E (1998) Location of diphenylhexatriene (DPH) and its derivatives within membranes: comparison of different fluorescence quenching analysis of membrane depth. Biochemistry 37:8180–8190 35. Ou B, Hampsch-Woodill M, Prior RL (2001) Development and validation of an improved oxygen radical absorbance capacity assay using fluorescein as the fluorescent probe. J Agric Food Chem 49:4619–4626 36. Broniowska KA, Kirilyuk I, Wisniewska A (2007) Spin-labelled lutein as a new antioxidant in protection against lipid peroxidation. Free Radical Res 41:1053–1060
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Reis et al.
37. Barclay L, Vinqvist MR (1994) Membrane peroxidation: inhibiting effects of water soluble antioxidants on phospholipids of different charge types. Free Radic Biol Med 16:779–788 38. Gutiérrez ME, García AF, Africa de Mandariaga M, Sagrista ML, Casadó FJ, Mora M (2003) Interaction of tocopherols and phenolic compounds with membrane lipid components: evaluation of their antioxidant activity in a liposomal model system. Life Sci 72:2337–2360 39. Saija A, Scalese M, Lanza M, Marzullo D, Bonina F, Castelli F (1995) Flavonoids as antioxidant agents: importance of their interaction with biomembranes. Free Radic Biol Med 19:481–486 40. Lúcio M, Ferreira H, Lima JLFC, Reis S (2007) Use of liposomes to evaluate the role of membrane interactions on antioxidant activity. Anal Chim Acta 597:163–170 41. Brooks P (1998) Use and benefits of nonsteroidal anti-inflammatory drugs. Am J Med 104:9S–13S 42. Kitamura K, Imayoshi N, Goto T, Shiro H, Mano T, Nakai Y (1995) Second derivative spectrophotometric determination of partition coefficients of chlorpromazine and promazine between lecithin bilayer vesicles and water. Anal Chim Acta 304:101–106 43. Kitamura K, Imayoshi N (1992) Secondderivative spectrophotometric determination
44.
45. 46.
47.
48.
49.
of the binding constant between chlorpromazine and -cyclodextrin in aqueous solutions. Anal Sci 8:497–501 Ferreira H, Lúcio M, Castro B, Gameiro P, Lima JLFC, Reis S (2003) Partition and location of nimesulide in EPC liposomes: a spectrophotometric and fluorescence study. Anal Bioanal Chem 377:293–298 Lakowicz JR (1999) Principles of fluorescence spectroscopy. Kluwer Academic/Plenum, New York Lentz BR (1993) Use of fluorescent probes to monitor molecular order and motions within liposome bilayers. Chem Phys Lipids 64:99–116 Barclay LRC, Artz JD, Mowat JJ (1995) Partitioning and antioxidant action of the water-soluble antioxidant, trolox, between the aqueous and lipid phases of phosphatidylcholine membranes – C-14 tracer and product studies. Biochim Biophys Acta 1237:77–85 Wang S, Beechem JM, Gratton E, Glaser M (1991) Orientational distribution of 1, 6-diphenyl- 1, 3, 5-hexatriene in phospholipid vesicles as determined by global analysis of frequency domain fluorimetry data. Biochemistry 30:5565–5512 Repáková J, Holopainen JM, Morrow MR, McDonald MC, Capková P, Vattulainen I (2005) Influence of DPH on the structure and dynamics of a DPPC bilayer. Biophys J 88:3398–3410
Chapter 14 Studying Colloidal Aggregation Using Liposomes Juan Sabín, Gerardo Prieto, and Félix Sarmiento Abstract Colloidal aggregation using liposomes has been studied in this chapter. As criteria of stability, the stability factor, an extension of the DLVO theory of colloidal stability, the fractal dimension of the liposome aggregates and the different regimes of aggregation (RLCA and DLCA) and the temperature have been used. Key words: Liposomes aggregation, Liposomes stability, Stability factor, Extension of DLVO model, RLCA regime of aggregation, DLCA regime of aggregation, Fractal dimension
1. Introduction Liposomes can be characterized as associating colloids, built up of amphoteric lipid molecules that self-assemble in aqueous media into spherical, self-closed structures. They consist of one or several concentric membranes; their size ranges from 20 nm to several micrometers, whereas the thickness of the membranes is around 4 nm (1). Liposomes play an important role in the colloidal science as model membranes and are important in biological, pharmaceutical and medical research because liposomes are the most effective carriers for introducing many different types of agents into cells, and the applications for liposome-based samples and products are extremely wide. Liposome dispersions are not thermodynamically stable (2). The total free energy of a dispersed system can always be lowered by reduction in the interfacial area. This tendency to aggregate is attributable to attractive van der Waals forces. The existence of other interactions between particles determines the stability. The applications of liposomes as nanoscale containers for drugs (3, 4), vaccines (5), enzymes or genetic material (6) require control and prediction of the liposome dispersion stability. This chapter introduces some methods to study the stability in solution. V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_14, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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2. Materials 1. l-a-phosphatidylcholine from egg yolk (EYPC), 2. 1,2-dimiristoyl-sn-glycero-3-phospho-choline (DMPC) 3. 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC). 4. Calcium nitrate 5. Lanthanum nitrate. 6. Methanol 7. Chloroform 8. Polycarbonate membranes. 9. Liposome preparation (see Note 1).
3. Methods 3.1. The Stability Factor (W) as a Criterion of Stability of Liposomes
The stability factor (W) is a kinetic criterion for the stability of a colloidal system (7): W =
kr ks
(1)
where kr describes the maximum coagulation rate (rapid coagulation regime where every collision between particles is effective) and ks is the particular coagulation rate (slow coagulation regime where not every collision results in coagulation). Thus, the inverse of the stability ratio provides a measure of the effectiveness of collisions leading to aggregation or fusion. The rate constants can be obtained by nephelometry. This technique consists of measuring at a low-angle, the static light scattering (see Note 2). The total scattering intensity for a dispersion of identical primary particles with a time-varying distribution sizes is (8):
I (t , q ) = 1 + 2knst I q (0)
(2)
where Iq(0) is the initial intensity of light scattered at angle q, ns is the number of primary particles and k is the rate constant. The scattered light intensity at low angles increases linearly with time, and then an absolute coagulation rate can be obtained from the slope if the number of primary particles is known. Figure 1 shows the results of a typical nephelometry experience. From the slopes of intensity versus time of Fig. 1, W can be calculated by (1). The intersection of the linear fit in the slow
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Fig. 1. Scattered light intensity (arbitrary units) versus time for the study of colloidal stability of phosphatidylcholine liposomes in aqueous media at different La+3 concentrations. [From Sabín J, et al. (2005) Langmuir 21:10968–10975]
Fig. 2. Stability ratio (W) versus La3+ concentration for phosphatidylcholine liposomes in aqueous media. The obtained critical coagulation concentration is 0.3 M. [From Sabín J, et al. (2005) Langmuir 21:10968–10975]
regime and the linear fit in the rapid regime define the critical coagulation concentration (ccc) at which liposome aggregates are formed (Fig. 2).
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3.2. Extension to the Classical DLVO Theory to Study Stability of Liposomes
The Derjaguin-Landau-Verwey-Overbeek (DLVO) theory of colloidal stability is, at the moment, the only quantitative theory of colloidal sciences (9, 10). In their classical approach, the interaction potential between similar colloidal particles in polar (especially, aqueous) solutions is described by two types of forces: attractive van der Waals forces and repulsive double layer forces. Although the DLVO theory has been extensively applied with outstanding success, it has its inherent limitations. The development of new techniques in the determination of the interaction potential between surfaces (atomic force microscopy, for example) has shown agreement with DLVO at separation above a few nanometers but, at smaller separations, a short-range repulsive force appears, often termed “hydration” interaction. In this sense, it can be interesting to compare the experimental results of stability of liposomes with a DLVO extended theory that has, in account, the hydration potential. The van der Waals interaction VA(x) for the case of two spherical shells of equal radius, a, and thickness, d, derived by Hamaker, is given by (11): V A (x ) = −
Aa 1 2 1 A x (x + 2d) − + − ln 12 x + 2d x + d d 6 (x + d)2
(3)
where A is the attractive Hamaker constant, a the initial radius of the liposomes, and x the distance between the two spherical shells. The Hamaker constant can be calculated, for example, using an algorithm-based on a SIMPLEX method, which minimizes the error of the nephelometry data to the function:
V b (u) exp T du 2 (u + 2a) kBT W = ∞ VA b (u) ∫0 (u + 2a)2 exp kBT du
∫
∞
0
(4)
where kB is the Bolztmann constant, T is the absolute temperature, VT represents the total DLVO potential, and b(u) is the hydrodynamic correlation factor, related to retardation on the particles diffusion due to the friction with the medium:
b (u) =
6u 2 + 13u + 2a 2 6u 2 + 4ua
(5)
where u = (x - 2a)/a The expression for the repulsive potential, VR(x) , by area unit has the form (12, 13):
VR (x ) = 2p e 0 er (a + ∆)y o2 exp [−k (x − 2∆)]
(6)
where y° is the surface potential, which coincides with z (zeta potential, see Note 3) for high values, and k is the reciprocal Debye length:
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Fig. 3. DLVO potentials of phosphatidylcholine liposomes as a function of the distance between two liposomes for different La3+ concentrations. [From Sabín J, et al. (2005) Langmuir 21:10968–10975]
κ2 =
( z12 z 2 + z22 z1 )cs N A e2 e 0 e r kB T
(7)
where cs is the concentration, e the elementary charge, z i the valence of the ions, and NA the Avogadro number. Finally, D is the thickness of the Stern layer and it is taken as the hydrated radius of the adsorbed ion. In Fig. 3, it can be seen an example of the DLVO potential for different ion concentrations. The aggregation of liposomes occurs when the repulsive barrier is lower than kBT at the contact surface (hydration surface D= 0.49 nm). Figure 3 shows also that the repulsive barrier is below kBT for ion concentrations larger than 0.27 M. 3.3. Fractal Dimension as a Tool to Analyze the Stability of Liposomes
The large structures that liposomes form when they aggregate produce good models for further analysis. The fractal dimension characterizes how the monomers occupy space in the aggregates and can be calculated measuring the angular dependence of the light scattered by the clusters and following the evolution of the cluster size. The aggregation mechanism can be described by two different regimes: diffusion-limited cluster aggregation (DLCA) and reaction-limited cluster aggregation (RLCA) (see Note 4). Figure 4 shows an example of evolution of size of the clusters of different liposomes along time. As can be seen in this Fig. DMPC shows the typical transition from RLCA to the DLCA aggregation. For concentrations below 0.5 × 10−3 M, the liposomes remain stable with no changes in size. At concentrations between 0.5 × 10−3 and 10−2 M, the growing of clusters is slow and exponential, as in a typical RLCA aggregation
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Fig. 4. Evolution of the size of the clusters of dimyristoylphosphocholine (DMPC), dipalmitoylphsphocholine (DPPC) and egg yolk phosphatidylcholine (EYPC) liposomes along time. The aggregation was induced after starting the measurements by addition of different concentrations of Ca2+. [From Sabín J, et al. (2007) Eur Phys J E 24:201–210]
(see Note 4). For higher concentrations, the increase follows a DLCA regime and no differences between Ca2+ concentrations can be noticed. Clusters of EYPC liposomes reach a maximum size, which depends on the Ca2+ concentration and, after this size is attained, the cluster becomes stable and no other liposomes aggregate to the clusters. DPPC liposomes show an intermediate behavior. In all cases, the fractal dimensions fell in the range from 2.1 to 1.8, corresponding to the two regimes. 3.4. Analyzing the Influence of the Temperature on Stability of Liposomes
The aggregation of liposomes is influenced by temperature. As an example, we show the time evolution of DMPC liposomes when La3+ at fixed concentration is added at different temperatures (Fig. 5).
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Fig. 5. Size evolution as a function of time for DMPC liposomes. The arrows indicates when the addiction of La3+ and solvent, respectively, was realized. [Results not published]
As it can be seen in this Fig. 5, at 25°C, a notable increment of the size is detected, indicating a DLCA regime of aggregation. At approximately 1,500 s, solvent is added and the clusters of liposomes are destroyed, indicating a reversible aggregation. Similar behavior was observed at 37°C although in this case the size of increment is less pronounced (RLCA regime). At 40°C, the liposomes remain stable with a slight decrease in their size. This reduction could be due to osmotic effects.
4. Notes 1. Liposomes are prepared by the thin-film hydration method. A solution of the lipids in chloroform/methanol or chloroform is evaporated in a rotary evaporator to dryness under a stream of nitrogen and the resultant lipid film hydrated with double-distilled, degassed, and deionized water. This mixture is extruded a minimum of five times through polycarbonate filters of the adequate pore size. 2. In a typical experiment, the angle detecting scattered light is around 10°, the time of 120 s, and the flow cell is rectangular with 2 mm path length. The cell must be thoroughly cleaned with chromic acid and rinsed with distilled water. Normally, the liposome stability is studied in the presence of ions.
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Then, equal volumes (for example, 1 mL) of salt and liposome solutions are mixed and introduced into the cell by an automatic mixing device. The dead time is, in general, quite short (14). 3. The zeta potential is calculated from the electrophoretic mobilities, mE by means of the Henry correction of Smoluchoswski equation (15):
mE =
3m E h 1 2e 0 e r f (κa )
where e0 is the permittivity of the vacuum, er is the relative permittivity dielectric constant, a is the particle radius, k is the Debye length, and eta the viscosity of water. The function f(ka) depends on the particle shape. For, k > 1 (case of the liposomes) takes the form:
f (κa) =
3 9 75 − + 2 2 2 2κa 2κ a
Electrophoretic mobilities can be measured with the appropriate instrument. For example, in most of our experiments, a Malvern Instruments Zetamaster 5002 was used, taking the average of five measurements at stationary level. The cell used was a 5 mm × 2 mm rectangular quartz capillarity. The controlled temperature was 25°C. Following Fig. 6 shows an example of the variation of the zeta potential as a function of ion concentration.
Fig. 6. Zeta potential of phosphatidylcholine liposomes as a function of La3+ concentration. [From Sabín J, et al. (2005) Langmuir 21:10968–10975]
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4. In the DLCA model, the cluster´s growth is controlled by diffusion and every collision between particles results in the formation of a permanent contact (16). The fractal dimension is insensitive to the sticking probability. Computer simulations of this model yield a fractal dimension of 1.8 in three dimensions. In the RLCA model (17), the particles are added one at a time at random to sites adjacent to occupied sites. This process produces a relatively compact cluster whose density correlations are independent of distance. The growth is limited by reaction kinetics at the interface of the growing clusters and only a small fraction of particle collisions leads to permanent contact. In this case, the computer simulations yield a fractal dimension of 2.0 in three dimensions. The two models of aggregation lead to two different kinds of clusters. Clusters obtained in the DLCA regime are very open structures because the particles stick where the collisions occur and the probability for branches to collide is higher than to penetrate to the center of the cluster. On the contrary, in the RLCA regime, the particles have to collide with the cluster several times before sticking, increasing the possibility of the particle to reach the center of the cluster. The resulting structure is a compact and dense cluster. Computer simulation also reveals that the size of the clusters grows differently in the DLCA and the RLCA regimes. DLCA clusters grow faster since all collisions are effective, following a power law for the average radius of gyration, R gat 1/df , where t is time and df the fractal dimension. The growth of the RLCA clusters follows an exponential law R gae at, where a is a constant which depends on the sticking probability (P).
Acknowledgments This work was supported by grants-in-aid from the Spanish “Ministerio de Ciencia e Innovación” (Project MAT2008-04722) and the “Xunta de Galicia” (Project INCITE08PXIB20603PR).
References 1. New RRC (1990) Liposomes: a practical approach. IRL, Oxford 2. Lasic DD (1993) Liposomes. From physics to applications. Elsevier, Amsterdam 3. Gregoriadis G (1987) Liposomes as drug carriers. Wiley, New York
4. Devissaguet JP, Puisieux F (1993) Les liposomes: aspects tecnologiques, Biologiques et Pharmacologiques. Les Editions Inserm, Paris 5. Alving C, Richards RL (1983) Immunologic aspects of liposomes. In: Ostro MJ (ed) Liposomes. Marcel Dekker, New York, pp 209–287
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6. Puisieux F (1985) Les liposomes: applications thérapeutiques. Tec and Doc Lavoisier, Paris 7. Hidalgo-Álvarez R, Martín A, Fernández A, Bastos D, Martínez F, de las Nieves FJ (1996) Electrokinetic properties, colloidal stability and aggregation kinetics of polymer colloids. Adv Colloid Interface Sci 67:1–118 8. Lips A, Willis E (1973) Low angle light scattering technique for the study of coagulation. J Chem Soc Faraday Trans 1 69: 1226–1236 9. Derjaguin BV, Landau LD (1941) Theory of the stability of strongly charged lyophobic sols and of the adhesion of strongly charged particles in solution of electrolytes. Acta Physicochim URSS 14:633–662 10. Verwey EJW, Overbeek JThG (1948) Theory of the stability of lyophobic colloids. The interaction of particles having an electric double layer. Elsevier, Amsterdam 11. Tadmor R (2001) The London-van der Waals interaction energy between objects of various geometries. J Phys Condens Matter 13: L195–L202
12. Vincent B, Bijsterbosch H, Lyklema J (1970) Competitive adsorption of ions and neutral molecules in the Stern layer on silver iodide and its effect on colloidal stability. J Colloid Interface Sci 37:171–178 13. Bastos D, de las Nieves FJ (1994) Colloidal stability of sulfonated polystyrene model colloids. Correlation with electrokinetic data. Colloid Polym Sci 272:592–597 14. Molina-Bolívar JA, Galisteo-González F, Hidalgo-Álvarez R (1999) Development of a high sensitivity IgG-latex immunodetection system stabilized by hydration forces. Polym Int 48:685–690 15. Hunter JR (1981) Zeta potential in colloid science: principles and application. Academic, London 16. Meakin P (1983) Formation of fractal clusters and networks by irreversible diffusion-limited aggregation. Phys Rev Lett 51:1119–1122 17. Eden M (1961) Proceedings of the Fourth Berkeley Symposium on Mathematics, Statics and Probability. In: Neyman F (ed) University of California Press, Berkeley, p. 223
Chapter 15 Assessment of Liposome–Cell Interactions Jan A.A.M. Kamps Abstract Liposome–cell interactions have been assessed for over 30 years now by an enormous variety of approaches and methods. In-depth knowledge of liposome–cell interaction is still very relevant since new concepts and applications applying liposomes are being developed every day. This chapter does not aim to give a complete overview on methods to assess liposome–cell interactions but merely gives a handle to approach the assessment of liposome–cell interaction by describing some well-established methods that also allow for modification to adapt them for your specific research questions. Key words: Endocytosis, Fluorescence microscopy, Radioactive liposome markers, Fluorescent liposome markers, Cell binding, Liposome uptake, Cellular processing of liposomes
1. Introduction Liposome–cell interaction is one of the most important features in the research field of liposomes applied as versatile drug carriers, since it largely determines the fate of the compound to be delivered by the liposome. Liposome–cell interaction includes binding, uptake, intracellular homing and processing of liposomes, liposome components and liposomal content. Interactions of liposomes with cells are complex because they involve several distinct mechanisms of interaction and are dependent on many parameters such as the biological environment, cell type and liposome characteristics. Four general mechanisms of liposome–cell interactions can be distinguished: adsorption, (receptor mediated) endocytosis, lipid exchange and fusion (1–4). The nature of the liposome–cell interactions are determined by the type of cell and by liposome characteristics such as composition, size and charge, the presence and characteristics of targeting molecules on the liposome surface and by the biological environment e.g. the presence V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_15, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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of plasma or serum. On top of that, assessment of liposome–cell interactions can be even more complicated because different liposome constituents may interact differently with cells. To study liposome–cell interaction, many liposome markers are available, which can be roughly divided as water-soluble markers and lipid markers, where water-soluble markers can be encapsulated in the interior of the liposomes and lipid markers can be incorporated in the liposome bilayer. Obviously, for following the cellular fate of encapsulated material, one would choose a watersoluble marker, while for the fate of the liposome a stably incorporated and inert lipid label in the bilayer would be preferable. The choice for a specific marker depends primarily on the aim of an experiment and the laboratory facilities available. Both radioactive lipid markers and fluorescent lipid markers have been shown powerful tools to determine liposome–cell interaction and examples of both will be presented. In addition, a combination of more than one marker in a liposome allows for studying the different fates of specific liposome components (5). Different liposome markers will be addressed but it is beyond the scope of this chapter to provide full overview of markers available. The protocols provided deal with in vitro experiments with cultured cells that allow for detailed study of the interaction of liposomes with cells under different conditions, though it should be emphasized that extrapolation to in vivo events can only be made with great carefulness. In a study where we addressed the uptake of liposomes containing phosphatidylserine by liver endothelial cells, we showed massive uptake of these liposomes in primary cultures of these cells, while in vivo, liver endothelial cells did not contribute in liposome uptake at all (6). To study liposome–cell interaction, there are numerous methods available. In this chapter, we will address most common liposome–cell interactions and the methods to experimentally determine binding, uptake and intracellular processing of liposomes. The protocols given have shown in our hands to give reliable information on the nature of the liposome–cell interaction but are by far not the only or even the best methods for a certain research question. However, the methods described here can form a good starting point to study liposome–cell interaction and may offer a rational for further detailed study applying more sophisticated experimental set-ups.
2. Materials 2.1. Liposome Markers
As stated in the introduction, there are many liposomal markers available to assess liposome–cell interaction. The markers listed below we, routinely, use in our laboratory for the applications described in the methods section, but for any of them alternatives
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are commercially available. These markers can be applied for most if not all liposome preparation methods (see Note 1). 1. [3H]cholesteryloleyl ether (GE Healthcare Europe, Diegem, Belgium); Stable and metabolically inert radioactive lipid marker. 2. Cholesteryl[14C]oleate (GE Healthcare Europe, Diegem, Belgium); A lipid marker that is readily hydrolyzed in lysosomes of cells. 3. 1,1¢-dioctadecyl-3,3,3¢,3¢-tetramethyl indocarbocyanine perchlorate (DiI; Molecular Probes, Eugene, OR); Stable fluorescent lipid bilayer marker (exitation: 549 nm; emission: 565 nm) (see Note 2). 4. Calcein (e.g. from Sigma–Aldrich, St. Louis MO, USA); water-soluble fluorophore that is quenched at high (~100 mM) concentration (exitation: 491 nm; emission: 511 nm). 2.2. Cell Culture (General Protocol)
1. Appropriate cell culture medium depending on the cell type (e.g. Dulbecco’s Modified Eagle’s Medium (DMEM), supplemented with 10% fetal bovine serum (FCS)). 2. Cells are grown to ~80% confluency (as determined by microscopy), typically in 12 or 24 well-culture plates or on chamber slides (Lab-Tek, NUNC, Thermo Fischer Scientific) (see Note 3). 3. Before the start of the experiment, cells are washed twice with incubation medium (e.g. DMEM without FCS).
2.3. Liposome–Cell Incubations
1. Appropriate cell culture medium with or without FCS, depending on the nature of the experiment. 2. Stock solutions of labeled liposomes and when applicable stock solutions of other agents/effectors to be added to the cells. 3. Cold (£4ºC) washing buffer, e.g. phosphate-buffered saline (10 mM phosphate buffer, pH 7.4, 150 mM NaCl). 4. (a) 0.1 M NaOH for cell Lysis prior to measurement of radioactivity. (b) Phosphate-buffered saline (PBS): 10 mM phosphate buffer, pH 7.4, 150 mM NaCl, supplemented with 5% FCS or 1% bovine serum albumin (BSA) for fluorescence measurement using fluorescence-activated cell sorting (FACS). 5. (a) Scintillation counter to determine radioactivity. (b) Fluorescence-activated cell sorter.
2.4. Intracellular Processing/ Degradation of Liposomes
1. Liposomes labeled with both [3H]cholesteryloleyl ether and cholesteryl[14C]oleate. 2. Appropriate culture medium, containing an acceptor for 14 C-oleate. Serum proteins (10% FCS) or serum albumin (1–2%) may function as acceptor for 14C-oleate.
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3. Phosphate-buffered saline (PBS): 10 mM phosphate buffer, pH 7.4, 150 mM NaCl 4. 0.1 M NaOH for cell Lysis prior to measurement of radioactivity. 2.5. Fluorescence Microscopy
1. Cells cultured on chamber slides (Lab-Tek, NUNC, Thermo Fischer Scientific) or on microscope cover slips that have been put into a regular 12 or 24 well-culture plate prior to cell seeding. 2. Cold (£4ºC) phosphate-buffered saline or cold culture medium without FCS. 3. Fluorescence microscope with appropriate filter sets.
3. Methods 3.1. Preparation of Labeled Liposomes (General Protocol (see Note 4))
1. lipids (e.g. phospholipids and cholesterol) from stock solutions in chloroform: methanol are mixed in an appropriate molar ratio. (a) When applicable radioactive or fluorescent lipid markers are added to this mixture in an appropriate amount (typically, trace amounts, e.g. for [3H]cholesteryloleyl ether 1 µCi/µmol lipid, for Cholesteryl[14C]oleate 0.4 µCi/µmol lipid, and for DiI 0.5–1 mol% ). 2. The lipids are dried under reduced nitrogen pressure, dissolved in cyclohexane en lyophilized. 3. The lipids are then hydrated in a Hepes buffer (10 mM N-2hydroxyethylpiperazine-N¢-2-ethanesulfonic acid (Hepes), 135 mM NaCl, pH 7.4). (a) When applicable water-soluble markers are dissolved in the hydration buffer, e.g. Calcein, typically at a concentration of ~100 mM. (see Note 5). 4. The liposomes formed can be sized by repeated extrusion through polycarbonate filters (Costar, Cambridge MA, USA), pore size typically 50 nm, using a high-pressure extruder (Lipex, Vancouver, Canada). 5. When a water-soluble marker has been encapsulated, free marker has to be separated from the liposome-associated marker (e.g. by gel permeation chromatography). 6. Of each liposome preparation, phospholipid phosphorous is determined by a phosphate assay after perchloric acid destruction (7), and in case of radioactive liposomes, specific radioactivity of the liposome preparation is calculated after scintillation counting of a liposome sample. 7. Liposomes are routinely stored at 4ºC under argon atmosphere. A tube containing liposomes is carefully gassed with
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argon, allowing air to flush out, after which the tube is closed with an airtight screw cap. When argon is not available, nitrogen gas can be applied (see Note 6). 3.2. Cell Binding and Uptake of [3H] cholesteryloleyl ether Labeled Liposomes
1. Replace the culture medium by serum-free medium and incubate the cells for 1 h at 37°C in a humidified 5% CO2 / 95% air atmosphere (see Note 7). 2. Remove the serum-free medium and add new medium containing appropriate amounts of [3H]cholesteryloleyl ether-labeled liposomes and if necessary, other additions (see Note 8). 3. Incubate cells for the 3 h (see Note 9) at 37°C in a humidified 5% CO2/95% air atmosphere to determine cell association (binding and uptake) or for 3 h at 4°C to determine exclusively cell binding. 4. After the incubation the culture plates are placed on ice and washed six times with cold (£4ºC) phosphate-buffered saline (see Note 10). 5. The cells are lysed in 0.1 M NaOH for 1 h at 37°C or overnight at −20°C and subsequent thawing. 6. The cell-associated 3H radioactivity is determined by liquid scintillation counting of aliquots of the lysed cell suspension and is normalized to the amount of cellular protein as determined according to Lowry (8) using bovine serum albumin as a standard. 7. Cell binding and/or cell uptake of the liposomes is calculated from the specific activity of the liposomes (dpm/µmol lipid) and cellular radioactivity (dpm/mg of cell protein). 8. This procedure distinguishes between binding plus uptake and exclusive binding of liposomes by incubating the cells either at 37ºC or 4ºC. The difference between liposome association at 37ºC and association at 4ºC is a measure for internalization (see Note 11).
3.3. Intracellular Processing/ Degradation of Liposomes
When liposomes are double-labeled with a [3H]cholesteryl ether and the radioactive marker cholesteryl[14C]oleate which is readily hydrolyzed in the lysosomes, it is possible to make an estimation of the degradation of liposomes. Due to rapid release from the cells of [14C]oleate derived from the degraded cholesteryl[14C] oleate and the complete retention of the metabolically inert [3H] cholesteryloleyl ether, the 3H/14C ratio is a sensitive measure for intracellular degradation (9) (see Note 12). 1. Replace the culture medium by serum-free medium and incubate the cells for 1 h at 37°C in a humidified 5% CO2/95% air atmosphere.
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2. Remove the serum-free medium and add new medium containing appropriate amounts of [3H]cholesteryloleyl ether and cholesteryl[14C]oleate-labeled liposomes, an acceptor for [14C] oleate released by the cells, and if necessary other additions. 3. Incubate the cells as described in Subheading 3.2. 4. After the incubation the culture plates are placed on ice and washed six times with cold (£4ºC) phosphate-buffered saline. 5. The cells are lysed in 0.1 M NaOH for 1 h at 37°C or overnight at −20°C and subsequent thawing. 6. The cell-associated 3H and 14C radioactivity are determined by liquid scintillation counting of aliquots of the lysed cell suspension and are normalized to the amount of cellular protein as determined according to Lowry (8) using bovine serum albumin as a standard. 7. Liposome degradation and/or processing by the cells is calculated from the specific 3H and 14C activities of the liposomes (dpm/µmol lipid) and cellular radioactivities (dpm/ mg of cell protein). The 3H/14C ratio is then a measure for cellular degradation/processing. 3.4. Fluorescence Microscopy
1. Replace the culture medium by serum free medium and incubate the cells for 1 h at 37°C in a humidified 5% CO2/95% air atmosphere. 2. Remove the serum-free medium and add new medium containing appropriate amounts of fluorescently labeled liposomes and if necessary other additions. 3. Incubate cells for the appropriate time at 37°C in a humidiffied 5% CO2/95% air atmosphere in the dark, to determine cell binding and uptake. 4. Stop the incubation by careful washing of the cells three times with cold (£4ºC) phosphate- buffered saline and then put some cold medium without FCS on the cells. 5. Transport the chamber slides or culture dishes with cover slips on ice (keep covered from light) to the microscope and in case of chamber slides remove the chambers, cover the cells with a cover slip and immediately observe the cells under the fluorescence microscope. In case of cells cultured on a cover slip, carefully remove the cover slip from the culture well and mount it upside down on an object glass and immediately observe the cells (see Note 13).
3.5 FluorescenceActivated Cell Sorting
Association of fluorescently labeled liposomes with cells can be (semi)quantified using a fluorescence activated cell sorter (10). The incubation protocol is basically the same as with radioactive labeled liposomes (incubate in the dark!).
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1. After incubation in a 24 well-culture plate, stop the incubation by washing the cells two times with cold (£4ºC) phosphatebuffered saline. 2. Harvest the cells using an appropriate method e.g. trypsin treatment and transfer the cells in to FACS tubes that (in the case of trypsine treatment) contain 200 µl FCS. Keep the tubes on ice. 3. Add 3 ml of FACS buffer and wash the cells two times by centrifugation at 1600 rpm for 5 min at 4ºC. 4. Resuspend the cells in an appropriate volume and measure cell fluorescence according to the protocol of the fluorescence-activated cell sorter.
4. Notes 1. It is important to realize when studying liposome cell interaction that most methods rely on certain markers or labeled molecules. In fact, it means that one is studying the fate of the label which is not necessarily similar to the fate of the intact liposome. Therefore, it is needed to confirm data on binding, uptake and processing by other methods and to apply relevant controls. 2. Care should be taken not to expose fluorescent markers or fluorescently labeled liposomes to light. This will decrease fluorescence intensity in time. The examples described here (DiI, Calcein) are relatively insensitive to light. 3. When incubating liposomes with cultured cells, make sure that cell conditions such as medium, atmosphere and cell confluency at the time of incubation are comparable. There can be major differences in the handling of liposomes by cells depending on their confluency, i.e. growth status. 4. Liposomes can be prepared by any method, where lipid markers can be added in appropriate (trace) amounts to lipid mixture prior to hydration of the lipids, while water-soluble markers can be added during the hydration step. 5. The characteristic that calcein is quenched at high concentrations may be used to determine whether liposome content is released intracellularly. Upon intracellular release from the liposomes of the encapsulated content, the calcein will be diluted in the cytoplasm of the cell and emit fluorescence. Double labeling with a stable liposome-associated fluorophore, e.g. DiI, allows to control and/or correcting for differences in liposome uptake.
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6. Stability of markers has to be taken in account when studying the interaction of liposomes with cells. With the liposome markers described in the different protocols we always have routinely performed experiments within 3 weeks of preparation of the liposomes (unless stated otherwise). Although some of the liposome markers that we use showed reproducible results with regard to liposome–cell interaction after storage of the liposomes for 6 months or more we prefer to keep to the term indicated to avoid any artifacts from long-term storage of liposomes. 7. One-hour incubation with serum-free medium prior to incubation with liposomes is performed to exclude influence of serum proteins on binding and uptake of liposomes by cultured cells. However, for some research questions and/or cell types, the presence of serum in the (pre) incubation medium may be preferable. 8. For incubation of liposomes with cultured cells, dilute stock solutions of liposomes directly before the start of the experiment in the culture medium (without FCS) used in the incubation to avoid dilution of the culture medium upon addition of liposomes to the cells. The same holds true for any other addition (inhibitors, stimuli, etc). Always make sure to control for any solvent used in your incubation. 9. Incubation of 3 h is normally sufficient to determine binding and uptake (association); however, duration of incubation depends on specific research question, liposomes and cell type. For determining binding at 4ºC, the incubation should not be longer than 3 h since the quality of the cells often decreases with longer incubations under these conditions. 10. Washing cells prior to assess binding, uptake or processing of liposomes is necessary to get rid of excess (labeled) liposomes. In our hands 5–7 rounds of careful washing suffice. However, make sure to check cells between washing steps microscopically. Some (primary) cells may be very sensitive to washing and detach from their substrate. Efficiency of washing the cells can be determined by measuring the label left in the fluid taken from the cells in the subsequent washing steps. For cells that easily detach upon washing, addition of 1% (W/V) serum albumin to the washing medium may be beneficial. 11. The difference between liposome association at 37ºC and association at 4ºC can be used as a measure for internalization. However, one has to proceed here with caution since binding at 4ºC may not be representative of binding at 37ºC because of dynamic processes such as receptor recycling which do not occur at 4ºC. Differences between the association of liposomes to cells at 37ºC and 4ºC can also be
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accounted for by temperature effects on bilayer mobility which may, as was shown for immunoliposomes, affect the amount of liposomes bound to the cells, even in cases where there is no uptake of liposomes (11). 12. One should take notice that the 3H/14C ratio in the cells is not only dependent on the presence of a fatty acid acceptor in the culture medium such as albumin but also of the nature of the cells used in the experiments. Specialized cells involved in lipid metabolism may not release [14C]oleate to appreciable extents but, as in the case of hepatocytes, further metabolize it to CO2 or for the production of lipoproteins. 13. For fluorescence microscopy in live cells, it is important to establish the time that the cells retain their properties. In our hands, live cells kept at 4ºC should be observed microscopically within 1 hour. One should limit the amount of samples to be examined accordingly. This period has to be established separately for every cell type. If cells are kept too long, you may observe blebbing of cells, membrane deformation and in case of fluorescence microscopy, occurrence of fluorescence in the medium in between the cells. References 1. Pagano RE, Weinstein JN (1978) Interactions of liposomes with mammalian cells. Annu Rev Biophys Bioeng 7:435–468 2. Düzgünes N, Nir S (1999) Mechanisms and kinetics of liposome-cell interactions. Adv Drug Deliv Rev 40(1–2):3–18 3. Kamps JAAM, Scherphof GL (2003) Liposomes in biological systems. In: Torchilin VP, Weissig V (eds) Liposomes, a practical approach, 2nd edn. Oxford University Press, Oxford, pp 267–288 4. Torchilin VP (2005) Recent advances with liposomes as pharmaceutical carriers. Nat Rev Drug Discov 4(2):145–160 5. Yan X, Poelstra K, Scherphof GL, Kamps JA (2004) Role for scavenger receptor B-I in selective transfer of rhodamine-PE from liposomes to cells. Biochem Biophys Res Commun 325(3):908–914 6. Kamps JA, Morselt HW, Scherphof GL (1999) Uptake of liposomes containing phosphatidylserine by liver cells in vivo and by sinusoidal liver cells in primary culture: in vivo–in vitro
7. 8. 9.
10.
11.
differences. Biochem Biophys Res Commun 256(1):57–62 Böttcher CJF, Van Gent CM, Pries C (1961) A rapid and sensitive sub-micro phosphorus determination. Anal Chim Acta 24:203–204 Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193:265–275 Derksen JT, Morselt HW, Scherphof GL (1988) Uptake and processing of immunoglobulin-coated liposomes by subpopulations of rat liver macrophages. Biochim Biophys Acta 971(2):127–136 Kamps JA, Morselt HW, Swart PJ, Meijer DK, Scherphof GL (1997) Massive targeting of liposomes, surface-modified with anionized albumins, to hepatic endothelial cells. Proc Natl Acad Sci U S A 94(21):11681–11685 Koning GA, Morselt HW, Velinova MJ et al (1999) Selective transfer of a lipophilic prodrug of 5-fluorodeoxyuridine from immunoliposomes to colon cancer cells. Biochim Biophys Acta 1420(1–2):153–167
Chapter 16 Methods to Monitor Liposome Fusion, Permeability, and Interaction with Cells Nejat Düzgünes¸, Henrique Faneca, and Maria C. Pedroso de Lima Abstract We describe fluorescence assays for membrane fusion involving the fusion of liposomes with each other and with cultured cells, fluorescence methods to assess liposome uptake by cells and the intracellular delivery of liposome contents, and assays to evaluate liposome membrane permeability. The Tb/DPA and ANTS/DPX assays monitor the intermixing of aqueous contents of liposomes. The NBD-PE/ Rhodamine-PE assay follows the intermixing of liposomal lipids. A variation of this method is suitable for detecting the mixing of the inner monolayers of liposomes. The lipid-mixing assay is also used to study the fusion of cationic liposomes and lipoplexes with cultured cells. The intracellular delivery of liposome contents are monitored, via fluorescence microscopy or flow cytometry, by measuring the release of calcein from the liposome interior, and normalized to cell-associated liposomes quantitated with Rhodamine-PE in the membrane of the same liposomes. The release of liposome contents is monitored by the increase in fluorescence of encapsulated carboxyfluorescein, calcein, or ANTS/DPX, or by the decrease in fluorescence of encapsulated Tb/DPA. Key words: Membrane fusion, Fluorescence microscopy, Fluorescence dequenching, Flow cytometry, Resonance energy transfer
1. Introduction Many studies of liposomes have employed fluorescence techniques. These studies range from the determination of the fluidity of lipid bilayers (using fluorescence anisotropy) and the intracellular fate of endocytosed liposomes (employing fluorescence microscopy), to monitoring the fusion of liposomes with each other and the fusion of viruses with liposomes (via fluorometry). In this chapter,
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_16, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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we will concentrate on fluorescence assays for membrane fusion involving the fusion of liposomes with each other and with cultured cells, fluorescence methods to assess liposome uptake by cells and the intracellular delivery of liposome contents, and assays to evaluate liposome membrane permeability.
2. Materials 2.1. Liposome Preparation 2.1.1. Large Unilamellar Liposomes Prepared by Reverse Phase Evaporation
1. A rotary evaporator (Büchi (Flawil, Switzerland), Labconco (Kansas City, MO), or equivalent) attached to an oil-free vacuum pump (Büchi), with a vacuum gauge, and inlet tubing connected to an argon tank. 2. A bath-type sonicator (Laboratory Supply Co., Hicksville, NY). 3. Vortex mixer. 4. An additional argon tank with tubing connected to a clamped Pasteur pipette. 5. A high-pressure extrusion device (Northern Lipids, Vancouver, Canada, or custom-made) or a syringe extrusion apparatus (Avestin, Toronto, Canada, or Avanti Polar Lipids, Alabaster, AL). 6. Poylcarbonate membranes with defined pore diameter (Nuclepore brand from Whatman, Kent, UK or Florham, NJ; or Poretics/GE Osmonics brand from Fisher Scientific, Waltham, MA). 7. A high-quality glass tube of approx 1 cm diameter, with a “screw-cap” with a Teflon lining at the top, and a larger tube with a fitting enabling attachment to the rotary evaporator and into which the former tube can fit. 8. Teflon tape. 9. Phospholipids, fluorescent lipids and cholesterol (Avanti Polar Lipids, Alabaster, AL or other suppliers). Lipids are kept at −20˚C or −80˚C in glass tubes or bottles, after gently flushing argon over the lipid solution. 10. Beckman Coulter N4 Plus Submicron Particle Sizer (Fullerton, CA).
2.1.2. Large Unilamellar Liposomes Prepared by Extrusion of Multilamellar Liposomes
1. A rotary evaporator (Büchi (Flawil, Switzerland), Labconco (Kansas City, MO), or equivalent) attached to an oil-free vacuum pump (Büchi), with a vacuum gauge, and inlet tubing connected to an argon tank.
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2. Phospholipids, fluorescent lipids and cholesterol (Avanti Polar Lipids, Alabaster, AL or other suppliers). Lipids are kept at −20˚C or −80˚C in glass tubes or bottles, after gently flushing argon over the lipid solution. 3. Vortex mixer. 4. An additional argon tank with tubing connected to a clamped Pasteur pipette 5. A high-pressure extrusion device (Northern Lipids, Vancouver, Canada, or custom-made) or a syringe extrusion apparatus (Avestin, Toronto, Canada, or Avanti Polar Lipids, Alabaster, AL). 6. Polycarbonate membranes with defined pore diameter (Nuclepore, Whatman; or Poretics/GE Osmonics, Fisher Scientific). 7. A high-quality glass tube of approx 1 cm diameter, with a “screw-cap” with a teflon lining at the top, and a larger tube with a fitting enabling attachment to the rotary evaporator and into which the former tube can fit. 8. Teflon tape. 9. Beckman Coulter N4 Plus Submicron Particle Sizer (Fullerton, CA). 2.1.3. Small Unilamellar Liposomes Prepared by Sonication
1. A rotary evaporator (Büchi, Labconco, or equivalent) attached to an oil-free vacuum pump (Büchi), with a vacuum gauge, and inlet tubing connected to an argon tank. 2. A vacuum oven at room temperature. 3. A bath-type sonicator (Laboratory Supply Co., Hicksville, NY). 4. Vortex mixer. 5. An additional argon tank with tubing connected to a clamped Pasteur pipette. 6. A high-quality glass tube of approx 1 cm diameter, with a “screw-cap” with a Teflon lining at the top, and a larger tube with a fitting enabling attachment to the rotary evaporator and into which the former tube can fit. 7. Teflon tape. 8. Phospholipids, fluorescent lipids and cholesterol (Avanti Polar Lipids, Alabaster, AL or other suppliers). Lipids are kept at −20˚C or −80˚C in glass tubes or bottles, after gently flushing argon over the lipid solution. 9. Ultracentrifuge (Beckman or equivalent). 10. Beckman Coulter N4 Plus Submicron Particle Sizer (Fullerton, CA).
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2.1.4. Measurement of the Phospholipid Concentration of Liposomes
1. Heating block set at 160–170˚C, with aluminum block inserts for test tubes of 25 mm diameter. 2. Glass jar pipettor (Oxford) of 1 l capacity, vortex mixer, and electrical boiling water bath into which a metal tube rack with a handle can be inserted. 3. 10 N H2SO4 prepared by adding 100 ml concentrated H2SO4 into water to make up 360 ml of solution, placed in a polypropylene dispenser jar or equivalent container. Care should be taken to wear protective gear (gloves, goggles or face mask) and to use a chemical hood. 4. 9% H2O2 (3 ml of 30% H2O2 is diluted with distilled or purified water to 10 ml final volume). 5. H2O2indicator strips (EM Quant Peroxide Test, EM Science, Gibbstown, NJ). 6. Ammonium molybdate (0.22% in 0.25 N H2SO4; 25 ml of 10 N H2SO4 are added to distilled or purified water, 2.2 g ammonium molybdate are added, and the solution is brought up to 1 L with water). The solution is placed in the Oxford pipettor. 7. ANSA (Fiske) reagent. A 15% NaHSO3 solution is prepared; 250 mg aminonaphtholsulfonic acid and 500 mg Na2SO3 are added to the NaHSO3 solution, bringing the volume up to 100 ml. To dissolve all ingredients, the solution is heated and mixed gently on a stir-plate. Then the solution is stored in the cold. 8. Ascorbate can be used as an alternative to ANSA; 15 g ascorbic acid are dissolved in 100 ml distilled or purified water and stored in the refrigerator.
2.2. Liposome Fusion: Intermixing of Aqueous Contents 2.2.1. The Tb/DPA Assay
1. Buffer A: 100 mM NaCl, 1 mM EDTA, 10 mM TES, pH 7.4 (all chemicals can be obtained from Sigma–Aldrich). All solutions are kept at 4˚C. 2. Buffer B: 100 mM NaCl, 10 mM TES, pH 7.4. 3. A vapor pressure osmometer (Wescor, Logan, Utah). 4. Three 1 × 20 cm chromatography columns (Bio-Rad, Hercules, California) and buffer reservoirs. 5. Sephadex G-75 or G-50 (Pharmacia, Uppsala), pre-swollen in distilled water containing 0.02% sodium azide, and de-gassed, and preferably kept at room temperature. 6. Solution to be encapsulated in large unilamellar “Tb- liposomes”: 2.5 mM TbCl3, 50 mM Na citrate, 10 mM TES, pH 7.4. All solutions are kept at 4˚C. 7. Solution to be encapsulated in large unilamellar “DPAliposomes”: 50 mM Na dipicolinate, 20 mM NaCl, 10 mM TES, pH 7.4.
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8. Solution to be encapsulated in large unilamellar “Tb/DPA liposomes”: A 1:1 mixture of the above two reagents (6 and 7). 9. Solution to be encapsulated in small unilamellar “Tb-liposomes”: 15 mM TbCl3, 150 mM Na citrate, 10 mM TES, pH 7.4. 10. Solution to be encapsulated in small unilamellar “DPAliposomes”: 150 mM Na dipicolinate, 10 mM TES, pH 7.4. 11. Solution to be encapsulated in small unilamellar “Tb/DPA liposomes”: A 1:1 mixture of the reagents in 9 and 10. 12. Polystyrene 15 ml culture tubes. 13. For assay calibration, a solution of 2 mM Na dipicolinate, 10 mM TES, pH 7.4. 14. 10% (w/v) Na cholate (Calbiochem) or 16 mM C 12E8 (octaethyleneglycol-dodecyl ether; Calbiochem). 15. Teflon-coated micro, or castle-type stir-bars. 16. Fluorometer, preferably with a thermal jacket, magnetic stirring, and a pinhole for injection of fusogens into the cuvette. 17. High-pass (>530 nm) cut-off filter (e.g. Corning 3–68). 18. Appropriate phospholipids and/or cholesterol for liposome preparation (Avanti Polar Lipids). 2.2.2. The ANTS/ DPX Assay
1. Solution to be encapsulated in large unilamellar “ANTSliposomes”: 25 mM aminonaphthalene trisulfonic acid (ANTS; molecular probes), 40 mM NaCl, 10 mM TES, pH 7.4. All solutions are kept at 4˚C. The tube containing the ANTS solution is wrapped in aluminum foil to maintain darkness. 2. Solution to be encapsulated in large unilamellar “DPXliposomes”: 90 mM p-xylylene bis(pyridinium) bromide (DPX, Molecular Probes), 10 mM TES, pH 7.4. 3. Solution to be encapsulated in large unilamellar “ANTS/ DPX-liposomes” for calibration of the assay: A 1:1 mixture of the above two solutions. 4. Buffer C: 100 mM NaCl, 0.1 mM EDTA, 10 mM TES, pH 7.4. 5. Vapor pressure osmometer. 6. Sephadex G-75 or G-50 (Pharmacia, Uppsala or Sigma, St. Louis). 7. Three 1 × 20 cm Bio-Rad columns and buffer reservoirs. 8. Fluorometer, preferably with a thermal jacket, magnetic stirring, and a pinhole for injection of fusogens into the cuvette.
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9. Appropriate phospholipids and/or cholesterol for liposome preparation (Avanti Polar Lipids). 2.3. Liposome Fusion: Intermixing of Lipids 2.3.1. The NBD/Rhodamine Assay
1. N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) phosphatidylethanolamine (Avanti Polar Lipids or Invitrogen/Molecular Probes). Lipids are kept at −20˚C or −80˚C in glass tubes or bottles, after gently flushing argon over the lipid solution. 2. N-(lissamine rhodamine B sulfonyl) phosphatidylethano lamine (Avanti Polar Lipids or Invitrogen/Molecular Probes). 3. Appropriate phospholipids and/or cholesterol for liposome preparation (Avanti Polar Lipids). 4. Medium: 100 mM NaCl, 0.1 mM EDTA, 10 mM TES, pH 7.4 (Buffer C). All solutions are kept at 4˚C. 5. Fluorometer
2.3.2. Inner Monolayer Mixing Assay
1. N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) phosphatidylethanolamine (Avanti Polar Lipids). Lipids are kept at −20˚C or −80˚C in glass tubes or bottles, after gently flushing argon over the lipid solution. 2. N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) (Avanti Polar Lipids)
phosphatidylserine
3. DiI(5)C18 (1,1¢-dioctadecyl-3,3,3¢,3¢-tetramethylindodicar bocyanine perchlorate (Invitrogen/Molecular Probes)). 4. Appropriate phospholipids and cholesterol for liposome preparation. 5. Sodium dithionate (stock solution of 200 mM sodium dithionate, 100 mM Tris, pH 10, made shortly before use). 6. Medium: 100 mM NaCl, 0.1 mM EDTA, 10 mM TES, pH 7.4 (Buffer C). 7. Bio-Gel A-30 spin columns (0.8 mL capacity) (Bio-Rad, Richmond, CA). 2.4. Fusion of Liposomes and Lipoplexes with Cultured Cells
1. THP-1 monocytic leukemia cells are available from the UCSF Cell Culture Facility (San Francisco, CA), and other sources including the American Type Culture Collection (Manassas, VA). 2. Culture medium: RPMI-1640 medium containing 25 mM HEPES, 2% sodium bicarbonate, 10% heat-inactivated fetal bovine serum, 100 µg/ml streptomycin and 1 U/ml penicillin (Biochrom KG, Berlin; Invitrogen, Carlsbad, CA). All media are stored at 4˚C, but warmed to 37˚C before addition to cells. 3. Phenol red-free RPMI-1640 containing 25 mM HEPES buffer, kept at 4˚C.
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4. Trypan blue solution (Sigma). 5. The cationic lipid 1,2-dioleoyl-(trimethylammonium) propane (DOTAP). All lipids are stored in glass tubes at −20˚C or −80˚C after gently flushing argon over the lipid solution. 6. Egg phosphatidylcholine (PC) (Avanti). 7. Phosphatidylethanolamine produced by transphosphatidylation of PC (PE) (Avanti). 8. N-(lissamine rhodamine B sulfonyl) phosphatidylethanolamine (Avanti). 9. Plasmid DNA: pCMV.luc. 10. Antimycin A (Calbiochem, La Jolla, CA) stock solution of 100 µg/ml. All solutions should be kept at 4˚C. 11. NaF (Sigma) stock solution of 1 M. 12. NaN3 (Sigma) stock solution of 10% (w/v). 2.5. Intracellular Delivery of Liposome Contents
1. Egg phosphatidylcholine (PC), dioleoylphosphatidylethanolamine (DOPE), lissamine rhodamine B-phosphatidylethanolamine (egg) (Rh-PE), phosphatidylserine (PS), phosphatidylglycerol (PG), distearoylphosphatidylcholine (DSPC) and poly(ethylene glycol) (2000)-distearoylphosphatidylethanolamine (DSPE-PEG) are purchased from Avanti Polar Lipids. All lipids are stored in glass tubes at −20˚C or −80˚C after gently flushing argon over the lipid solution. 2. Cholesteryl hemisuccinate (CHEMS), calcein, MES, TES, EDTA, phorbol 12-myristate 13-acetate, Triton X-100, propidium iodide, bafilomycin A1, NaN3, NaF, and NaCl are obtained from Sigma. Antimycin A is obtained from Calbiochem (La Jolla, CA). Calcein may also be obtained from (Invitrogen/ Molecular Probes). 3. Solution to be encapsulated: 80 mM calcein (Invitrogen/ Molecular Probes), 10 mM TES, pH 8.2, 1 mM EDTA, adjusted to 300 mOsm by the addition of NaCl. 4. For dialysis: TES-buffered saline (TBS, 140 mM NaCl, 10 mM TES, pH 7.4) containing 0.1 mM EDTA. 5. Polycarbonate membranes of 100 nm pore diameter (Costar, Cambridge, MA). 6. LiposoFast extruder (Avestin, Ottawa, Canada). 7. Monocytic human THP-1 cells. 8. RPMI-1640 medium, with and without phenol red (UCSF Cell Culture Facility or Invitrogen). 9. Fetal bovine serum (UCSF Cell Culture Facility or Invitrogen).
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10. Antimycin A (Calbiochem, La Jolla, CA) stock solution of 100 µg/ml. 11. NaF (Sigma) stock solution of 1 M. 12. NaN3 (Sigma) stock solution of 10% (w/v). 13. Cell dissociation buffer (Life Technologies, Inc.) 14. Phosphate-buffered saline (PBS) without calcium and magnesium. 15. Becton Dickinson FACStar Plus flow cytometer (or equivalent). 16. Lab-Tek chambered coverglasses for tissue culture (Nunc, Naperville, IL). 17. Diaphot-inverted fluorescence microscope (Nikon, Melville, NY) and fluorescence image analysis system (Photon Technology International, Birmingham, NJ). 2.6. Liposome Permeability 2.6.1. Release of Carboxyfluorescein or Calcein
1. Some carboxyfluorescein or calcein preparations may contain impurities, which may be eliminated by gel filtration of a concentrated solution in water on Sephadex LH-20. 2. Solution to be encapsulated in large unilamellar liposomes: 50 mM carboxyfluorescein or calcein (Na salt) (Molecular Probes), 10 mM TES, pH 7.4. Solutions of fluorescent probes should be kept in the dark by wrapping aluminum foil over the tube, and stored at 4˚C. 3. Solution to be encapsulated in small unilamellar liposomes: 100 mM carboxyfluorescein or calcein, 10 mM TES, pH 7.4. 4. Buffer C: 100 mM NaCl, 0.1 mM EDTA, 10 mM TES, pH 7.4 5. Triton X-100 (Sigma) stock solution (100×) at 10% (w/v) or C12E8 (Calbiochem) at 80 mM. 6. Fluorometer, preferably with a thermal jacket, magnetic stirring, and a pinhole for injection of membrane-active agents (such as peptides) into the cuvette. 7. High-pass (>530 nm) cut-off filter (e.g. Corning 3–68).
2.6.2. Release of the Tb/ DPA Complex
1. Solution to be encapsulated in large unilamellar “Tb/DPA liposomes”: 1.25 mM TbCl3, 25 mM Na citrate, 25 mM Na dipicolinate, 10 mM NaCl, 10 mM TES, pH 7.4. 2. Solution to be encapsulated in small unilamellar “Tb/DPA liposomes”: 7.5 mM TbCl3, 75 mM Na citrate, 75 mM Na dipicolinate, 10 mM TES, pH 7.4. 3. Buffer A: 100 mM NaCl, 1 mM EDTA, 10 mM TES, pH 7.4. Solutions should be stored at 4˚C. 4. Buffer B: 100 mM NaCl, 10 mM TES, pH 7.4.
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5. Sephadex G-75 or G-50 column equilibrated with Buffer A. 6. Triton X-100 stock solution (100×) at 10% (w/v) or C12E8 (20×) at 16 mM. Detergent solutions are kept at room temperature. 7. Fluorometer, as above. 2.6.3. The Release of ANTS/DPX
1. Solution to be encapsulated in large unilamellar “ANTS/ DPX-liposomes”: 12.5 mM ANTS, 45 mM DPX, 20 mM NaCl, 10 mM TES, pH 7.4. Solutions of fluorescent probes should be kept in the dark by wrapping aluminum foil over the tube, and stored at 4˚C. 2. Buffer C: 100 mM NaCl, 0.1 mM EDTA, 10 mM TES, pH 7.4. Solutions should be stored at 4˚C. 3. Sephadex G-75 or G-50 column equilibrated with Buffer C. 4. Triton X-100 stock solution (100×) at 10% (w/v) or C12E8 (20×) at 16 mM. Detergent solutions are kept at room temperature. 5. Fluorometer, as above.
3. Methods
3.1. Liposome Preparation 3.1.1 Large Unilamellar Liposomes Prepared by Reverse Phase Evaporation
1. Pure or mixed phospholipids in chloroform are measured using gastight Hamilton syringes under a chemical hood, and dispensed into a glass tube. The tube is covered with teflon tape, and then placed inside a larger tube that fits on a rotary evaporator until the lipid dries into a thin film in vacuum. The total lipid is usually 10–20 µmol. 2. A few milliliters of diethyl ether is washed with a similar volume of distilled or purified water in a tightly capped glass tube by gentle shaking, and the mixture is allowed to separate. One milliliter of the ether (the top layer) is removed by means of a glass pipette or syringe and added to the dried phospholipid film, ensuring that the lipid dissolves completely. 3. The buffer to be encapsulated (0.34 ml) is added to the phospholipid solution in ether. A gentle stream of argon gas is flushed over the mixture using a Pasteur pipette, and the tube is sealed with teflon tape and a screw-cap. The mixture is sonicated for 2–5 min, resulting in a stable emulsion. If an emulsion is not formed it is likely that the sonicator is not at an optimal setting. In this case, either the level of the water in the bath or the power supply needs to be adjusted.
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4. The cap is opened in the chemical hood, the tube is sealed again with teflon tape, and placed inside the larger glass tube that fits onto the rotary evaporator. About 1 ml of water is added to the outer tube, both to facilitate thermal contact and to minimize the evaporation of the aqueous solution in the inner tube. The outer tube is immersed in the water bath of the rotary evaporator, kept at 30°C. 5. The ether is evaporated gently in controlled vacuum, at about 350 mm Hg, under constant supervision. The tube is purged occasionally with argon gas attached to the top of the evaporator to maintain the vacuum level and to prevent excessive bubbling. The vacuum is allowed to build up when a stable gel is formed. 6. To break up the gel, the inner glass tube is removed and vortexed vigorously for 5–10 s. The tube is placed again in the outer tube and controlled rotary evaporation is resumed. This step is repeated once or twice, until an aqueous opalescent suspension is formed. An additional 0.66 ml of the encapsulation buffer is added to the suspension and rotary evaporation is continued for an additional 20 min to remove any residual ether (see Note 1). 7. To achieve a uniform size distribution, the liposome suspension is passed several times through polycarbonate membranes of 100 nm pore diameter (or other desired diameter), using a high-pressure or syringe extruder. 8. The average size and the size distribution of the liposomes are assessed by dynamic light scattering in a Beckman Coulter N4 Plus Submicron Particle Sizer (or equivalent instrument). 9. Liposomes are stored at 4˚C after gently flushing the top part of the tube with a stream of argon, and sealing the cap with parafilm. 3.1.2. Large Unilamellar Liposomes Prepared by Extrusion of Multilamellar Liposomes
1. Pure or mixed phospholipids (usually 10–20 µmol) in chloroform are measured using gastight Hamilton syringes under a chemical hood, and dispensed into a glass tube. The tube is covered with teflon tape, and placed inside a larger tube that fits on a rotary evaporator until the lipid dries into a thin film in vacuum. The evaporator is purged with argon to reduce the vacuum, and the inner tube is placed in a vacuum jar or vacuum oven, and kept under high vacuum for at least 2 h to eliminate any residual chloroform. 2. The phospholipid film is hydrated with the buffer to be encapsulated (e.g., the Tb citrate solution), the tube is purged with argon gas and covered with teflon tape, and the cap is closed tightly. The mixture is vortexed for 10 min at room temperature to form multilamellar liposomes. If the lipid has a
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high gel to liquid-crystalline phase transition temperature, it is hydrated with buffer at a higher temperature. The vortex mixing is interrupted every 30 s to place the tube in a water bath at this high temperature for about 15 s. 3. To achieve a uniform size distribution, the liposome suspension is passed 21 times through a polycarbonate membrane of 100 nm pore diameter (or other desired diameter), using a syringe extruder. If the extrusion requires too much force, the multilamellar liposomes should first be passed through a membrane of larger pore diameter (e.g. 800 or 400 nm) (see Note 2). 4. The average size and the size distribution of the liposomes are assessed by dynamic light scattering in a Beckman Coulter N4 Plus Submicron Particle Sizer (or equivalent instrument). 5. Liposomes are stored at 4˚C after gently flushing the top part of the tube with a stream of argon, and sealing the cap with parafilm. 3.1.3. Small Unilamellar Liposomes Prepared by Sonication
1. Ten micromoles of a phospholipid mixture in chloroform are dispensed into the glass tube that is then covered with teflon tape over the top of the tube. The tube is placed in the larger glass tube that fits on the rotary evaporator, and the chloroform is allowed to evaporate, using condensation precautions to trap the solvent. When the lipid forms a dry film, the evaporator is purged with argon to reduce the vacuum. The inner tube is then placed in a vacuum jar or vacuum oven, and is kept under high vacuum for at least 2 h to eliminate any residual chloroform. 2. The dried phospholipid film is hydrated with the buffer to be encapsulated, the tube is purged with argon gas and covered with Teflon tape, and the screw-cap is closed tightly. The mixture is vortexed for 10 min at room temperature. If a lipid with a high gel to liquid-crystalline phase transition temperature is used, the lipid is hydrated with buffer at a temperature above the transition temperature, and the tube is placed in a water bath at this high temperature in between short periods of vortexing. 3. The multilamellar suspension is sonicated in a bath-type sonicator for 0.5–1 h. The level of water in the bath is adjusted such that the water surface breaks up into small droplets under sonication. The top of the liposome suspension is aligned with the level of the water such that an aerosol forms occasionally in the tube. Overheating of the bath should be avoided, by circulating water through the bath or by adding some ice and re-adjusting the level of the water. For lipids with high transition temperatures, the bath should be maintained at a few degrees above this temperature.
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4. To remove any remaining large liposomes, the resulting opalescent suspension is centrifuged at 100,000g for 1 h at 4˚C, preferably in a swinging bucket rotor. The supernatant is removed without disturbing the pellet, and used as the small unilamellar liposome preparation (see Note 3). 5. The average size and the size distribution of the liposomes are assessed by dynamic light scattering in a Beckman Coulter N4 Plus Submicron Particle Sizer (or equivalent instrument). 6. Liposomes are stored at 4˚C after gently flushing the top part of the tube with a stream of argon, and sealing the cap with parafilm. 3.1.4. Measurement of the Phospholipid Concentration of Liposomes
1. Liposome samples are placed in triplicate tubes at an estimated amount of less than 0.1 µmol inorganic phosphate. Phosphate standards are pipetted in triplicate (for example 0.01, 0.05, 0.075, 0.1 µmol inorganic phosphate). Four hundred microliters of 10 N H2SO4 are added to each tube that are then heated for 30 min on the heating block. 2. The tubes are cooled at room temperature and 100 µl H2O2 are added using a pipettor or repeater pipette (e.g. Pipetman or Eppendorf). The tubes are returned to the heating block for 30 min on the heating block. The fumes are tested for the absence of H2O2, using the indicator strips. 3. The tubes are placed in a round metal rack (that will fit eventually into an electric water boiler), 4.6 ml of 0.22% ammonium molybdate reagent are added to the tubes and mixed on a vortex mixer. Two hundred microliters of ANSA (or Fiske) reagent are added to each tube and then mixed on a vortex mixer. Alternatively, 100 µl ascorbate can also be used for this step. 4. The metal rack is lowered into a boiling water bath for 7–10 min, and then cooled. The contents of the tubes are transferred to spectrophotometer cuvettes. Protective latex or vinyl gloves are recommended for this procedure. A spectrophotometer with a sipper accessory is preferable to avoid handling of acid-containing tubes. 5. The absorbance of the solutions in each cuvette is measured at 812 nm or at 660 nm (if the solution is too concentrated). The phosphate content of the sample is assessed from the standard curve (see Note 4).
3.2. Liposome Fusion: Intermixing of Aqueous Contents 3.2.1. The Tb/DPA Acid Assay
1. For large unilamellar liposomes, the osmolality of the solutions inside the liposomes should match that of the medium, and if lower, should be adjusted with the addition of small amounts of NaCl. The Tb-, DPA- and Tb/DPA-liposomes are prepared as described in Subheadings 3.1.1 or 3.1.2. (see Note 5).
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2. The 3 Sephadex G-75 or G-50 columns are equilibrated with Buffer A. The liposomes are passed through separate columns to eliminate the solution outside the liposomes, using Buffer A for elution. It is necessary to include the 1 mM EDTA in Buffer A to prevent the binding of Tb ions to the phospholipids. The first 4 ml after the liposomes are loaded on the gel are discarded, and the next 3 ml are collected in the 15-ml culture tube. Argon gas is flushed over the liposome suspensions for 10 s without causing any significant disruption of the surface of the liquid; then the tubes are capped tightly, sealed with parafilm, and stored in ice. 3. One milliliter of the collected Tb-liposomes is placed on the third Sephadex column equilibrated with Buffer B, and collected as in step 2. Since EDTA would interfere with the formation of the Tb/DPA complex when the vesicles are lysed to obtain 100% fluorescence, this chromatography step replaces the EDTA in the medium, producing the “Tb minus EDTA liposomes.” 4. The lipid concentration of the liposomes is measured in each of the culture tubes by inorganic phosphate analysis (see Subheading 3.1.3). 5. For a fusion assay utilizing 50 µM total lipid concentration, 25 µM Tb-liposomes are mixed with 25 µM DPA-liposomes. To calibrate the fluorescence to “100%” fluorescence; i.e. the maximal fluorescene that can be obtained if all the liposomes formed a mega-liposome, 25 µM of the “Tb minus EDTA liposomes” are placed in a fluorometer cuvette containing the appropriate volume of Buffer B. The amount of buffer should be adjusted to allow for the volumes of the subsequent ingredients. Ten microliters of the 2 mM DPA stock are added to the cuvette (final DPA concentration 20 µM). To lyse the liposomes, 50 µl of either of the detergent stocks are added and the fluorescence is allowed to equilibrate. The final concentrations of the detergents should be 0.5% (w/v) for Na cholate, or 0.8 mM for C12E8. 6. The excitation wavelength of the fluorometer is set at 276– 278 nm. The emission wavelength is set to 545 nm. Intermediate slit widths are used to optimize the intensity, but to minimize scattering artifacts. The high-pass cut-off filter is placed before the emission monochromator, to minimize any contributions from light scattering. Crossed polarizers can also be used to minimize lightscattering contributions. For our experiments we have used SLM 4000, SLM 8000, and Spex Fluorolog fluoro meters. Other instruments with similar light intensity and sensitivity can be used for this assay.
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7. A 1:1 mixture of the Tb-liposomes and the DPA-liposomes is prepared as a stock solution in a 15-ml polystyrene culture tube, at a final lipid concentration that is tenfold higher than that used for the actual assay. For example, a suspension of 500 µM lipid is prepared to achieve a final concentration of 50 µM in the assay. The stock solution (0.1 ml) is diluted into 0.9 ml of Buffer B; this process also dilutes the final EDTA concentration to 0.1 mM. 8. For a fluorometer with a strip-chart recorder, the low fluorescence of this suspension is set to 0% Fmax, using the offset function of the fluorometer. Using the gain function, necessary adjustments are made to the 100% level with the cuvette containing the liposomes lysed in the presence of the DPA. The calibration procedure is repeated after a set of measurements to ensure that the lamp intensity has not changed in the course of the experiment. 9. For a fluorometer with computerized data acquisition, the assay can be calibrated by subtracting the initial level of fluorescence [I(0)] from the data set, and dividing the resulting data set by the numerical difference between the fluorescence intensity of the calibration vesicles [I (•)] and the new 0% level. If the resulting value is multiplied by 100, the percentage value is obtained. Thus, the extent of fusion, F(t), as a percentage of maximal fluorescence, is given by
F (t ) = 100 x [I (t ) - I(0)]/[I (∞) - I(0)] where I(t) is the fluorescence intensity at time t. 10. The assay may be calibrated in an alternative manner also. Here, 50 µM of the Tb/DPA liposomes are considered to have the maximal, 100%, fluorescence. These liposomes represent the fusion product of all the Tb-liposomes and DPA-liposomes in the fusion assay. Of course, these calibration liposomes should have the same size distribution and should be used at the same lipid concentration as the total of the Tb- and DPAliposome populations (25 µM of each). One slight complication of the use of Tb/DPA liposomes is that after they are transferred to room temperature from the storage temperature (0˚C), it takes about 0.5 h for the fluorescence to reach a steady state.
3.2.2. The ANTS/DPX Assay
1. Three batches of large unilamellar liposomes are prepared with the ANTS, DPX, and ANTS/DPX solutions. Small unilamellar liposomes have not produced reliable results with this assay (see Note 6). 2. The liposomes are chromatographed on separate Sephadex G-75 or G-50 columns equilibrated with Buffer C. The first 4 ml, measured from the point where the liposomes are
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allowed to partition into the gel, are discarded as the void volume. The next 3 ml are collected in the polystyrene culture tube. The tube is flushed gently with argon, capped tightly, sealed with parafilm, and stored in ice until use. 3. The phosphate concentration of each of the liposome preparations is measured using the assay described in Subheading 3.1.4. 4. The excitation monochromator of the fluorometer is set to 360 nm, and the emission monochromator to 530 nm, with relatively wide slit widths (10–20 nm). A high-pass filter (e.g. Corning 3–68) is placed in the emission channel to eliminate light scattering artifacts. 5. ANTS-liposomes (25 µM lipid) are placed in the appropriate amount of buffer (1–2 ml, depending on the fluorometer, with “flea” or “castle” stir-bars), and the gain of the fluorometer is adjusted to an arbitrary unit of 100%. The fluorescence of 50 µM ANTS/DPX-liposomes is taken as 0%. The latter liposomes represent the theoretical fusion product of all the ANTS vesicles and all the DPX vesicles (also 25 µM) used in the assay. 6. Putative fusogens, such as divalent cations, protons, or fusogenic peptides are added to the cuvette from a concentrated stock solution under constant stirring. Liposome fusion results in the decrease of fluorescence due to quenching of ANTS by the DPX. 3.3. Liposome Fusion: Intermixing of Lipids 3.3.1. The NBD/Rhodamine Assay
1. “Labeled liposomes” are prepared as described in Subheadings 3.1.1, 3.1.2, or 3.1.3, with 0.8 mole% each of NBD-PE and Rh-PE in the initial chloroform mixture in addition to other phospholipids (see Note 7). 2. “Unlabeled liposomes” of the same lipid composition are prepared without the fluorophores present. 3. “Calibration liposomes”, or “mock fused liposomes,” are made as in 1, but with 0.08 mole% of each fluorophore (see Note 8). 4. The phosphate concentration of each of the liposome preparations is measured (Subheading 3.1.4), and aliquots from the labeled and unlabeled liposomes are mixed at a ratio of 1:9 in 2 mL of Buffer C. For a total lipid concentration of 50 µM, this would be 5 µM-labeled liposomes and 45 µM-unlabeled liposomes. 5. The excitation and emission monochromators are set to 460 nm and 530 nm, respectively. The maximal (100%) fluorescence is set with 50 µM of the calibration liposomes. The residual fluorescence of the mixture of labeled and unlabeled liposomes is set to 0% fluorescence. The percentage of lipid mixing as a function of time is given by M(t) = 100 × [I(t) − I(0)]/ [I(•) − I(0)], where I(t) is the fluorescence intensity at time t,
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I(0) is the residual fluorescence, and I(•) is the maximal fluorescence. 6. Fusion of the liposomes results in an increase in NBD fluorescence as the probes are diluted into the unlabeled liposomes. 7. Maximal fluorescence can also be set by lysing the labeled liposomes at the same concentration to be used in the assay, with a detergent that does not affect the quantum yield of NBD. Commercial preparations of Triton X-100 require a correction factor of 1.4–1.5, while some purified preparations of this detergent may have no inhibitory effect on the fluorescence. Alternative detergents are C12E8 or C12E9 (Calbiochem) 3.3.2. Inner Monolayer Mixing Assay
1. Liposomes containing 0.75 mole% each of N-NBD-PS and either N-Rh-PE or diI(5)C18 (20 mM total phospholipid) are prepared in Buffer C, as described in Subheadings 3.1.1 or 3.1.2 (see Note 9). 2. To a 10 mM liposome suspension on ice, dithionate is added to a final concentration of 80–100 mM, and the mixture incubated for 30–45 min. 3. The dithionate is removed by centrifuging100 µl aliquots through the spin columns. 4. The remainder of the assay is as described in Subheading 3.3.1 to measure the intermixing of the inner monolayers.
3.4. Fusion of Liposomes and Lipoplexes with Cultured Cells
1. THP-1 human monocytic leukemia cells are grown in the growth medium described in Subheading 2.4, at 37˚C and 5% CO2. 2. The cells are harvested by centrifugation at 180 g for 7 min at room temperature, and the pellet is resuspended in phenol red-free RPMI-1640 with HEPES buffer. The cells are centrifuged again. This procedure is then repeated twice. The final pellet of cells is suspended at a concentration of 20 × 106 cells/ml in a polypropylene culture tube and kept on ice until use the same day. Cell viability is determined by mixing a small sample of cells 1:1 with the Trypan blue solution, while counting the cells in a hemacytometer. 3. Large unilamellar liposomes containing the cationic lipid DOTAP alone, or mixed with PE or PC, are prepared first by hydration of a dried lipid film in 150 mM NaCl, 10 mM HEPES, pH 7.4, at a final concentration of 5 mM lipid (see Subheading 3.1.2). This is followed by vortexing under argon to form multilamellar liposomes, and by extrusion through polycarbonate membranes of 100 nm pore diameter. Rh-PE is included in the initial chloroform mixture of lipids at 5 mole%, at which concentration it is self-quenched.
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4. Lipid mixing between the liposomes and cell membrane is monitored continuously in a magnetically stirred fluorometer cuvette maintained at 37˚C or other desired temperatures. Cells (100 µl) are added to the cuvette containing 1.9 ml of 150 mM NaCl, 10 mM HEPES, pH 7.4, and allowed to equilibrate to the set temperature. Liposomes are added to a final concentration of 20 µM. 5. The fluorescence is measured with the excitation wavelength at 568 nm and the emission wavelength at 586 nm. In a Spex fluorometer, the excitation and emission slits are set to 0.5 and 1 mm, respectively. The fluorescence scale is set with the initial fluorescence of liposomes and cells being 0%, and the maximal fluorescence (100%) is obtained by dissolving the liposomes with 0.5% (v/v) Triton X-100. 6. To prevent endocytosis, and hence to measure fusion at the plasma membrane only, the cells are pre-incubated with 1 µg/ml antimycin A, 10 mM NaF and 0.1% (w/v) NaN3 for 30 min at 37˚C. Fusion is monitored in the presence of these inhibitors. 7. To examine the fusion of cationic liposome-DNA complexes with cells, the complexes are prepared at varying lipid nitrogen/DNA phosphate (+/−) ratios right before they are added to the cells in the cuvette. The effect of the maturation of the complexes on fusion can also be investigated by preincubating the complexes for 1–2 h before addition to the cells (see Note 10). 3.5. Intracellular Delivery of Liposome Contents 3.5.1. Flow Cytometric Analysis
1. Large unilamellar liposomes encapsulating 80 mM calcein and containing 1 mole% Rh-PE in their membrane are prepared by extrusion of multilamellar liposomes (Subheading 3.1.2). 2. Monocytic human THP-1 cells are cultured in RPMI 1640 medium supplemented with 10% FBS. The cells are differentiated to macrophage-like cells by adding 160 nM phorbol 12-myristate 13-acetate in 24-well culture plates (106 cells per well). The cells are incubated for 5–6 days, and the culture medium is replaced with fresh medium. 3. The liposomes are added to the cells at a final phospholipid concentration of 100 µM, and incubated for various times at 37°C. The cells are then washed twice with phosphate-buffered saline (PBS) without calcium or magnesium ions. 4. The cells are detached from the plastic by adding 0.5 ml of dissociation buffer and mixed with 0.5 ml of PBS containing divalent cations, 2% FBS and 1 mg/ml propidium iodide. Rhodamine and calcein fluorescence is detected with a Becton Dickinson FACStar Plus flow cytometer (or equivalent),
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controlled by a Hewlett-Packard computer with Lysis II software (Becton Dickinson, San Jose, CA). The samples are analyzed for rhodamine using excitation at 528 nm and emission at 575 nm. Calcein is excited at 488 nm and the fluorescence emission is detected at 520 nm with a 0.1 neutral density filter. For each sample, 10,000 events are recorded. Forward scatter and propidium iodide fluorescence signals are used to gate the cell subset of interest and eliminate debris, dead cells, and cell aggregates. 5. Mean rhodamine fluorescence values reflect the binding and uptake of liposomes by the cells. The mean calcein fluorescence indicates the intracellular dequenching of the dye. The calculated ratio of calcein to rhodamine fluorescence is taken to measure the amount of aqueous marker released intracellularly per cell-associated liposome. The initial calcein to rhodamine fluorescence ratio of liposomes bound to the cells, in the absence of endocytosis, is obtained by incubating the liposomes with the cells at 4°C (see Note 11). 3.5.2. Image Analysis
1. THP-1 cells (105/well) are differentiated for 5–6 days in Lab-Tek chambered coverglasses for tissue culture, obtained from Nunc (Naperville, IL), by adding 160 nM phorbol 12-myristate 13-acetate to the culture medium. Cells are washed with cold RPMI media without phenol red, containing 20 mM HEPES buffer, pH 7.4. 2. Liposomes are added to cells at a final phospholipid concentration of 200 µM and incubated for 1 h at 4°C in phenol red-free RPMI medium with 20 mM HEPES buffer. After this pre-binding step, the cells are washed with cold media and the initial calcein and rhodamine fluorescence images are recorded using the Photon Technology International (PTI) ratio imaging system. To evaluate the kinetics of calcein dequenching, the cold medium is removed, medium at 37°C is added, and the cells are incubated for various times. 3. The cells are washed with cold medium, and the calcein and rhodamine fluorescence images are recorded. Cells are observed in a Nikon Diaphot epi-fluorescence microscope (Melville, NY) using a 100× objective and filters for FITC/ TXRD obtained from Chroma (Rockingham, VT). Averages of 16 snapshots are taken to reduce the background. Ratio images are produced using PTI (or equivalent) software. 4. Histograms of calcein to rhodamine ratio for each cell are determined using a square of 100 × 100 pixels, representing the approximate area of one cell at the magnification used. This provides a measure of calcein dequenching per cell. Average histograms for each liposome composition and time points are then calculated in Excel.
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5. Medians for each average histogram are calculated from cumulative curves. To calculate the growth of the area occupied by the calcein that is dequenched, the highest ratio in the control experiment (incubation at 4°C) is taken as the cut-off. The sum of the pixels with ratios higher than the cutoff ratio, are taken to estimate the dequenched area at various time points (see Note 12). 3.5.3. Fluorometric Measurements
1. THP-1 cells (5 × 106) are differentiated by incubation for 5–6 days in 5–ml of medium containing 160 nM phorbol 12-myristate 13-acetate in 6-well culture plates. The medium is removed and the cells are washed with phenol red-free RPMI medium with 20 mM HEPES buffer. 2. Cells are incubated with liposomes at a final phospholipid concentration of 150–200 µM for different times at 37°C in phenol red-free RPMI medium/20 mM HEPES buffer, and then washed. 3. Two milliliters of dissociation buffer are added, and the cells are incubated for 10 min at 37°C. The cells are detached from plastic with disposable scrapers (Costar Corporation, Cambridge, MA) and transferred into disposable fluorometer cuvettes (Hughes & Hughes Limited, Tonedale, Wellington). 4. Fluorescence measurements are performed in a SPEX Fluorolog 2 fluorometer (SPEX Industries, Inc. Edison, NJ), Perkin Elmer LS50 fluorometer or equivalent instrument. Calcein fluorescence is read at excitation and emission wavelengths of 490 and 520 nm, respectively, using 0.5 mm excitation and 1.0 mm emission slits. Rhodamine fluorescence is measured at excitation and emission wavelengths of 568 and 600 nm, respectively, using 0.5 mm excitation and 1.5 mm emission slits. 5. The sample chamber is adjusted to front-face configuration to prevent cell-scattering effects and is equipped with a magnetic stirrer. The temperature is maintained at 20°C with a thermostatic water circulator (see Note 12).
3.6. Liposome Permeability 3.6.1. Release of Carboxyfluorescein or Calcein
1. Large unilamellar liposomes are prepared using 50 mM carboxyfluorescein or calcein (Na salt) with 10 mM TES, pH 7.4. This concentration is iso-osmotic with Buffer C. 2. Small unilamellar liposomes are prepared using 100 mM carboxyfluorescein or calcein, 10 mM TES, pH 7.4. Larger concentrations of encapsulated material can be used in this case, since these liposomes are not active osmotically. 3. The unencapsulated material is separated by gel filtration on Sephadex G-75, using Buffer C as the elution buffer. 4. The phosphate concentration of the liposomes is measured as in Subheading 3.1.4.
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5. The excitation wavelength is set to 490 nm and the emission wavelength to 530 nm, using 0.5 mm excitation and 1.0 mm emission slits in the Spex fluorometer, and 5 nm and 10 nm slits in the Perkin-Elmer fluorometer. 6. The maximal fluorescence (100%) of a certain concentration of liposome is set by lysing the vesicles with 0.1% (w/v) Triton X-100 or 0.8 mM C12E8. 7. The extent of release, R(t) may also be calculated from R(t) = 100 × [I(t) − I(0)]/[I(•) − I(0)], where I(t) is the fluorescence intensity at time t, I(0) is the initial residual fluorescence of the liposomes before any membrane-active agents are added, and I(•) is the fluorescence obtained when the liposomes are lysed with detergent (see Note 13). 3.6.2. Release of the Tb/DPA Complex
1. Liposomes are prepared using the Tb/DPA solution appropriate for large or small unilamellar liposomes (Subheading 3.1). 2. The lipsomes are passed through the column to eliminate unencapsulated material, using Buffer A for elution. 3. The phosphate concentration of the liposomes is measured as in Subheading 3.1.4. 4. A stock solution is prepared in Buffer B at a lipid concentration tenfold higher than that to be used in the final assay. The stock solution (0.1 ml) is diluted into 0.9 ml of Buffer B, diluting the final EDTA concentration to 0.1 mM. 5. If the liposomes are not affected by the presence of Ca2+ (i.e. they do not contain a high-mole fraction of anionic phospholipids), this cation should be included in the external buffer to facilitate the dissociation of the Tb/DPA complex. 6. The initial fluorescence (excitation at 278 nm, emission at 545 nm, with the 3–68 cutoff filter) of the liposomes is set to 100%. After placing the liposomes into buffer, it is necessary to allow some time for the fluorescence to stabilize. The minimum fluorescence, corresponding to maximal leakage, can be set by lysing the liposomes with detergent (0.1% (w/v) Triton X-100 or 0.8 mM C12E8 (final concentrations)). 7. The extent of release, R(t), may be calculated from R(t) = 100 × [I(0) − I(t)]/[I(0) − I(•)], where I (t) is the fluorescence intensity at time t, I(0) is the initial fluorescence of the Tb/DPA liposomes before any membrane-active agents are added, and I(•) is the fluorescence obtained when all the liposome contents leak. The liposomes are lysed with detergent (see Note 14).
3.6.3. The Release of ANTS/DPX
1. Large unilamellar liposomes are prepared to encapsulate the ANTS/DPX solution. This assay is not suitable for use with small unilamellar liposomes.
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2. The liposomes are passed through a Sephadex column to eliminate unencapsulated material, using Buffer C for elution. 3. The phosphate concentration of the liposomes is measured as in Subheading 2.1.4. 4. The fluorometer is set as follows: Excitation wavelength at 360 nm, the emission wavelength at 530 nm, using relatively wide slit widths (10–20 nm), and a high-pass filter (Corning 3–68) in the emission channel to eliminate light-scattering artifacts. 5. The assay is calibrated to maximal fluorescence by lysing the vesicles with 0.1% (w/v) Triton X-100 or 0.8 mM C12E8 (final concentrations). The extent of release as a function of time can be calculated according to the formula in Subheading 3.6.1. (see Note 15).
4. Notes 1. The reverse-phase evaporation method was first reported by Szoka and Papahadjopoulos [1] and further developed by Düzgünes¸ et al. [2]. 2. Extrusion of multilamellar liposomes to obtain large unilamellar liposomes was developed by Olson et al. [3] and later by Mayer et al. [4]. 3. Small unilamellar liposomes prepared by sonication were first reported by Papahadjopoulos and Miller [5]. The method described here is adapted from Düzgünes¸ et al. [2]. 4. The method for determination of inorganic phosphate described here was first described by Bartlett [6]. The details described here were developed in the Papahadjopoulos laboratory at UCSF by T. Heath and D. Alford. 5. The interaction of Tb and DPA results in the formation of the fluorescent [Tb(DPA)3]3- chelation complex. Internal energy transfer from DPA to Tb results in an increase in the fluorescence intensity of the latter by four orders of magnitude. The reactants are encapsulated in different populations of liposomes. Membrane fusion results in an increase in fluorescence [7, 8]. Contents that are released into the external medium are prevented from interacting by the presence of EDTA in the medium. Divalent cations used to induce fusion in same cases further inhibit the interaction. The presence of citrate in the Tb-liposomes is essential to prevent the interaction of Tb3+ with anionic phospholipids in the liposome
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membrane. If phosphatidic acid is used as an anionic lipid, it is necessary to use a stronger chelator of Tb3+, such as nitrilotriacetic acid. 6. This assay is based on the collisional quenching of ANTS fluorescence by DPX. These reagents are encapsulated in separate populations of liposomes [9]. Membrane fusion results in the intermixing of ANTS and DPX and the quenching of ANTS fluorescence. If the contents are released and diluted into the medium, fluorescence quenching does not occur, since high concentrations of DPX are required for quenching. 7. The lipid-mixing assay measures the dilution of a fluorescence energy donor/acceptor pair from “labeled” liposomes to “unlabeled” liposomes as a result of membrane fusion. The efficiency of resonance energy transfer is then decreased, since the rate and efficiency of the transfer depend on the inverse sixth power of the distance between the two fluorophores (as well as the overlap of the emission spectrum of the donor and the absorption spectrum of the acceptor). Low concentrations of probes, usually less than 1 mole% of total lipid, are used for the assay, minimizing the extent of membrane perturbation. We describe a resonance energy transfer pair that has been used extensively by a number of laboratories [10, 11]. The fluorophores are attached to the headgroups of phosphatidylethanolamine and thus do not cause any appreciable perturbation of bilayer packing. 8. To avoid the use of detergents that may affect NBD fluorescence, liposomes corresponding to the fusion product of the labeled and unlabeled liposomes can be used to calibrate the assay to 100% fusion [12]. 9. The interaction of ions, proteins and other fusogens with the outer monolayer of liposomes containing fluorescent probes may alter the fluorescence intensity or lateral distribution of the probes. If the fluorophores were located only in the inner monolayer of liposomes, the fluorescence would not be affected by ion or protein binding to the liposome surface. In this method, the fluorophores exposed on the outer monolayer of the liposome are reduced with dithionate, thereby eliminating the fluorescence of outer monolayer fluorophores. N-NBD-phosphatidylserine (PS) was found to be more suitable for these experiments than N-NBD-PE, since it is less prone to transbilayer movement following reduction [13]. 10. Results obtained by this method have been described by Pires et al. [14].
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11. This method was first described by Slepushkin et al. [15] to evaluate the ability of sterically stabilized pH-sensitive liposomes to deliver their contents into cultured cells. 12. Results obtained with this method may be found in Simões et al. [16]. 13. The use of liposomal carboxyfluorescein and calcein was described by Weinstein et al. [17] and Allen et al. [18], respectively. The effect of bacterial toxins on liposome membrane permeability, using these fluorophores, was reported by Kayalar and Düzgüneş [19]. 14. The dissociation of the Tb/DPA complex was initially used to measure liposome permeability during membrane fusion [20]. 15. The ANTS/DPX release assay was first developed by Ellens et al. [21].
References 1. Szoka F Jr, Papahadjopoulos D (1978) Procedure for preparation of liposomes with large internal aqueous space and high capture by reverse-phase evaporation. Proc Natl Acad Sci USA 75:4194–4198 2. Düzgüneş N, Wilschut J, Hong K, Fraley R, Perry C, Friend DS, James TL, Papahadjopoulos D (1983) Physicochemical characterization of large unilamellar vesicles prepared by reverse-phase evaporation. Biochim Biophys Acta 732:289–299 3. Olson F, Hunt CA, Szoka FC, Vail WJ, Papahadjopoulos D (1979) Preparation of liposomes of defined size distribution by extrusion through polycarbonate membranes. Biochim Biophys Acta 557:9–23 4. Mayer LD, Hope MJ, Cullis PR (1985) Vesicles of variable sizes produced by a rapid extrusion procedure. Biochim Biophys Acta 858:161–168 5. Papahadjopoulos D, Miller N (1967) Phospholipid model membranes. I. Structural characteristics of hydrated liquid crystals. Biochim Biophys Acta 135:624–638 6. Bartlett GR (1959) Phosphorus assay in column chromatography. J Biol Chem 234:466–468 7. Wilschut J, Düzgüneş N, Fraley R, Papahadjopoulos D (1980) Studies on the mechanism of membrane fusion: Kinetics of Ca2+induced fusion of phosphatidylserine vesicles followed by a new assay for mixing of aqueous vesicle contents. Biochemistry 19:6011–6021
8. Düzgüneş N, Wilschut J (1993) Fusion assays monitoring intermixing of aqueous contents. Methods Enzymol 220:3–15 9. Ellens H, Bentz J, Szoka FC (1985) H+- and Ca2+-induced fusion and destabilization of liposomes. Biochemistry 24:3099–3106 10. Struck DK, Hoekstra D, Pagano RE (1981) Use of resonance energy transfer to monitor membrane fusion. Biochemistry 20:4093–4099 11. Hoekstra D, Düzgüneş N (1993) Lipid mixing assays to determine fusion in liposome systems. Methods Enzymol 220:15–32 12. Düzgüneş N, Allen TM, Fedor J, Papahadjopoulos D (1987) Lipid mixing during membrane aggregation and fusion. Why fusion assays disagree. Biochemistry 26:8435–8442 13. Meers P, Ali S, Erukulla R, Janoff AS (2000) Novel inner monolayer fusion assays reveal differential monolayer mixing associated with cation-dependent membrane fusion. Biochim Biophys Acta 1467:227–243 14. Pires P, Simões S, Nir S, Gaspar R, Düzgüneş N, Pedroso de Lima MC (1999) Interaction of cationic liposomes and their DNA complexes with monocytic leukemia cells. Biochim Biophys Acta 1418:71–84 15. Slepushkin VA, Simões S, Dazin P, Newman MS, Guo LS, Pedroso de Lima MC, Düzgüneş N (1997) Sterically stabilized pH-sensitive liposomes: intracellular delivery of aqueous contents and prolonged circulation in vivo. J Biol Chem 272:2382–2388
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16. Simões S, Slepushkin V, Düzgüneş N, Pedroso de Lima MC (2001) On the mechanisms of internalization and intracellular delivery mediated by pH-sensitive liposomes. Biochim Biophys Acta 1515:23–37 17. Weinstein JN, Yoshikami S, Henkart P, Blumenthal R, Hagins WA (1977) Liposomecell interaction: transfer and intracellular release of a trapped fluorescent marker. Science 195:489–492 18. Allen TM, Cleland LG (1980) Serum-induced leakage of liposome contents. Biochim Biophys Acta 597:418–426
19. Kayalar C, Düzgüneş N (1986) Membrane action of colicin E1: detection by the release of carboxyfluorescein and calcein from liposomes. Biochim Biophys Acta 860:51–56 20. Bentz J, Düzgüneş N, Nir S (1983) Kinetics of divalent cation-induced fusion of phosphatidylserine vesicles: correlation between fusogenic capacities and binding affinities. Biochemistry 22:3320–3330 21. Ellens H, Bentz J, Szoka FC (1984) pHinduced destabilization of phosphatidylethanolamine-containing liposomes: role of bilayer contact. Biochemistry 23:1532–1538
Chapter 17 The Use of Isothermal Titration Calorimetry to Study Multidrug Transport Proteins in Liposomes David Miller and Paula J. Booth Abstract Biophysical measurements of multidrug transporters in vitro can often be of limited relevance to the natural in vivo behavior. In particular, the properties of transporters when removed from their native bilayer and solubilized in detergents or lipids can differ significantly from their in vivo properties, reducing the value of in vitro measurements for the design of antagonists to the transporters. This problem can be addressed by studying the transport protein in liposomes in which the properties of the liposome bilayer are altered through systematic changes in lipid composition. Isothermal titration calorimetry can be used to determine the properties of the lipid-reconstituted protein in bilayers of different lipid compositions as well as to quantify the percentage recovery of functional protein in different lipids. Both these measurements lead to an accurate analysis of substrate binding activity. The approach is illustrated here for the small multidrug transport protein, EmrE from Escherichia coli. The percentage of functional EmrE successfully reconstituted into liposome depends on lipid composition. Differences in ligand binding and subtle differences in the secondary structure also occur in different lipid compositions. Key words: Small multidrug transporters, Ligand binding activity, Dissociation constant, Isothermal titration calorimetry, Reconstitution
1. Introduction Multidrug exporters are of great pharmaceutical interest because of their role in drug resistance of bacterial pathogens and their overexpression in a large number of cancerous cells resulting in resistance to anti-cancer pharmaceuticals. The study of multidrug transporters in vitro is essential for a complete understanding of the mechanism and for the successful design of antagonists to the transporters. The small multidrug transporter EmrE is capable
V. Weissig (ed.), Liposomes: Methods and Protocols, Volume 2: Biological Membrane Models, Methods in Molecular Biology, vol. 606, DOI 10.1007/978-1-60761-447-0_17, © Humana Press, a part of Springer Science+Business Media, LLC 2010
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of recognising and exporting a broad range of hydrophobic cationic substrates from Escherichia Coli (E. coli) (1) and has homologues in a number of bacterial species. The behavior of multidrug transporters in synthetic lipid bilayers is influenced by the bilayer composition and the quantity of functionally incorporated protein. The heterogenous nature of membrane bilayers complicates the detailed biophysical studies of integral membrane proteins. Interactions between the integral membrane proteins and the different regions of the bilayer are complex, with hydrophobic interactions occurring between the protein and the central hydrophobic region of the lipid bilayer, whilst polar and charge interactions exist with the lipid headgroups and aqueous solvent on either side of the bilayer. Furthermore, different lipid compositions, varying in headgroup and aliphatic chain composition, have significant effects on the protein function within the membrane, and there are variations in composition between different cellular compartments and different species (2). Altering the bilayer composition affects the properties of a number of integral membrane proteins, such as changes in the activity of enzymes involved in membrane lipid metabolism (Reviewed in (3, 4)), mechanosensitive channel opening properties (5–7), and the stability of the potassium channel KscA (8, 9). The consequences of lipid composition are not always considered in the structural and functional studies of membrane protein, which are often determined in detergent micelles, or in lipid environments that differ greatly from the native bilayer. The amount of functional transporter incorporated into the lipid bilayers will affect the kinetic parameters determined for transport or activity. We have used an isothermal titration calorimetry (ITC) based assay to quantify the amount of functional protein in synthetic liposomes, reconstituted from a ligand-binding competent, detergent-solubilized state in vitro. The ITC assay accurately quantifies the amount of functional protein from the ligand-binding activity and provides information on the ligandbinding properties in different lipid compositions. Specifically, the dissociation constant, Kd, is determined. In the ITC experiment, ligand is added to the transporter protein reconstituted into liposomes. The amount of heat released in the exothermic binding reaction between ligand and protein is measured for increasing ligand concentrations. This generates a binding isotherm from which Kd can be found by fitting the binding data to a particular binding model. The assay presented here is likely to be applicable for the study of other classes of transporters and receptors, provided a suitable ligand is available.
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2. Materials 2.1. Protein Purification
All chemicals are purchased from Sigma unless otherwise stated. 1. Solubilization buffer: 40 mM Tris HCl, pH 8.2, 100 mM NaCl, 10 mM 2-mercaptoethanol (b-ME), and 4% (w/v) n-Dodecyl-b-D-Malopyranoside (DDM) from Anatrace. All buffers containing b-ME are freshly prepared before use. 2. Ni2+-NTA agarose (QIAGEN) 3. Ni2+ wash buffer 1: 20 mM Tris-HCl, pH 8.3, 400 mM NaCl, 15 mM imidazole, 0.1% w/v DDM, and 5 mM b-ME. 4. Ni2+ wash buffer 2: 20 mM Tris-HCl, pH 8.3, 20 mM NaCl, 15 mM imidazole, 0.1% w/v DDM, and 5 mM 2-mercaptoethanol. 5. Ni2+ elution buffer: 20 mM Tris-HCl, pH 8.3, 25 mM NaCl, 200 mM imidazole, 0.1% w/v DDM, and 5 mM 2-ME. 6. NH4DDM buffer: 15 mM Tris-HCl, pH 7.5, 190 mM NH4Cl, 0.08% (w/v) DDM, and 5 mM 2ME. 7. Denaturing wash buffer: 10 M urea and 5% w/v SDS (173 mM, CMC ~8 mM or ~0.2% w/v) in 20 mM Tris-HCl, pH 8.3. 8. Denaturing elution buffer: 10 M urea, 5% w/v SDS, 20 mM Na Acetate, pH 4.0. 9. Amicon Ultra 50,000 MWCO centrifugal concentrator (Millipore). 10. HiLoad 16/160 Superdex 200 gel filtration column (Amersham).
2.2. EmrE Reconstitution
1. 1,2-Dioleoyl-sn-Glycero-3-Phosphocholine (DOPC), 1,2-Dioleoyl-sn-Glycero-3-[Phospho-rac-(1-glycerol)](DOPG), and 1,2-Dioleoyl-sn-Glycero-3-Phosphoethanolamine (DOPE) (Avanti polar lipids, as powder) prepared as 50 mg/ml stock solutions in either 1:1 chloroform:methanol or cylclohexane. 2. n-Dodecyl-b-D-Malopyranoside (DDM) and Octyl-b-DGlucoside (OG) were purchased from Anatrace and prepared fresh as a 20% (w/v) stock in water. 3. Reconstitution buffer: 15 mM TRIS HCl, pH 7.5 or 8.5, and 190 mM NH4Cl. 4. 1 ml HisTrap columns (Amersham).
2.3. Circular Dichroism
1. Resuspension buffer: 5 mM TRIS HCl, pH 8.0, and 190 mM NH4Cl. 2. Dilution buffer: 20 mM TRIS HCl at pH 7.5 or pH 8.5.
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3. TPP+: tetraphenyphosphonium bromide in 20 mM TRIS HCl, pH 7.5, freshly prepared from a 20 mg/ml stock in water. 4. Ethidium: Ethidium bromide in 20 mM TRIS HCl, pH 8.5, freshly prepared from a 10 mg/ml stock and stored in dark. 2.4. Protein Quantification
1. 1:1 chloroform:methanol solution prepared fresh and stored, wrapped in foil to prevent exposure to light.
3. Methods Efficient reconstitution of membrane protein is essential for ITC and circular dichroism (CD) measurements as large amounts of protein are typically required, and yields of overexpressed membrane protein are often low. Subsequent quantification of reconstituted transporter in lipid bilayers is a prerequisite for accurate activity measurements. The choice of detergent for purification and as a prelude to reconstitution is important as detergents such as DDM, which are very good at stabilising membrane proteins, are very difficult to remove from proteolipid samples, and may interfere with functional assays. Detergents such as OG, although easy to remove from proteolipid samples, do not suitably stabilize EmrE. Detergent choice is particularly important where activity assays involve a pH gradient across the bilayer as this cannot be maintained when residual DDM is present. A pH gradient is required in transport assays of EmrE. However, measurement of ligand binding is not required for the maintenance of a pH gradient, enabling the use of DDM. ITC binding assays provide a quantitative measure of ligand binding through the measurement of Kd. The stoichiometry of binding (n) can also be calculated from the binding isotherm. The binding of two ligands to EmrE is presented: the tight binding TPP+ (small Kd) and the weaker binding ethidium (larger Kd). ITC also has the advantage of determining the quantity of functional protein. Ligand binding to the small multidrug exporters occurs within the transmembrane region, and as ligand binding is a prerequisite for transport, it can be used as a quantitative measure of the amount of functional protein present. A prior knowledge of n enables the amount of functional reconstituted transporter to be determined by ITC (through non-linear curve fitting of the binding data). This is the case for EmrE, where n is known to be 0.5; one ligand binds to an EmrE dimer in DDM (10). The amount of protein in the reconstituted sample can be determined by various other methods. However, these often report on the amount of protein present and not whether it is also functional. A simple method for quantification
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of the amount of protein involving solubilization of reconstituted protein in chloroform:methanol and quantification by A280 using standards prepared in the same solvent is presented. CD is also used to assess the overall secondary structure content of EmrE reconstituted into different lipid compositions. The use of a high intensity UV source with a detector geometry, suitable for collecting scattered light, is essential for accurate CD measurements of liposome containing samples. A synchrotron radiation source with correct detector geometry is the preferred method for collection of CD data of proteolipid samples (11). Scattering of light from samples and failure to collect a significant proportion of the scattered light will result in significant artefacts in data, most notably at shorter wavelengths. For an accurate comparison between different lipid compositions, concentration of inserted protein must be accurately determined (i.e., by chloroform/methanol solubilization). However, a qualitative comparison of a single lipid composition in the presence and absence of a ligand is possible without an accurate protein concentration, provided all the proteins present are functional. 3.1. Sample Preparation: EmrE Overexpression and Purification
1. EmrE-His (cloned into pT7-7) is overexpressed and purified as previously described (12, 13). Briefly, after protein expression, cultures are harvested and disrupted using a cell disrupter. Membranes are isolated by centrifugation, solubilized at 4°C for at least 2 h in solubilising buffer. Solubilized material is diluted 1:1 with water, and insoluble material removed by centrifugation at 35,000× g for 30 min at 4°C. NaCl and imidazole are added to final concentrations of 350 mM and 15 mM, respectively. 2. DDM solubilized protein is subsequently incubated with 1.5 ml of Ni-NTA agarose for 1.5 h at 4°C. Non-specifically bound protein is removed by washing with 20 columns of Ni2+ wash buffer 1, followed by 20 column volumes of Ni2+ wash buffer 2. 3. EmrE is eluted with 10 column volumes of Ni2+ elution buffer. Eluted protein is concentrated to 2 ml using an Amicon Ultra-15 Ultracel 50 k centrifugal concentrator (Millipore) and applied to a HiLoad Superdex 200 gel filtration column (Amersham), equilibrated with NH4DDM buffer. 4. A280 of eluted protein is monitored and fractions collected. Fractions which contain a symmetric protein elution peak, determined to contain EmrE, are collected; fractions outside of this range are discarded, including those which contributed to small shoulders on the EmrE elution peak. Eluted EmrE is concentrated to approximately 1–2 mM as determined by A280 (A280 of 1 mg/ml = 2.56) (10), snap frozen in liquid N2 and stored at −80°C until required.
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3.2. Reconstitution of EmrE in Liposomes
1. Fresh stock solutions of the lipid: DOPC, DOPE, and DOPG at 50 mg/ml are prepared in cyclohexane and mixed to the desired molar ratios, evaporated to a thin film at room temperature and freeze-dried overnight. Dried lipid mixtures are rehydrated to 6.25 mg/ml (for ITC TPP+ binding studies) or 12.5 mg/ml (for ethidium binding) in reconstitution buffer at pH 7.5 or 8.5 for TPP+ or ethidium binding experiments, respectively (the higher pH being required for the weaker binding ethidium substrate). 2. Lipids are stirred for 15 min at room temperature (~22°C) and OG added from a 20% (w/v) stock solution to 0.65% or 1% (w/v) for TPP+ and ethidium binding experiments, respectively, and stirred for an additional hour. Lipid vesicles are extruded to 200 nm diameter by passaging 10 times through a polycarbonate filter (Nucleopore Track-Etch Membrane from Whatman) using a LIPEXTM extruder (Northern Lipids Inc). 3. EmrE is reconstituted to a calculated maximum of 10 µM (for TPP+ experiments) or 50 µM (for ethidium) by rapid mixing of 2.2 ml (for TPP+) or 4.4 ml (for ethidium) of lipid vesicle preparation to purified EmrE, giving a final EmrE concentration of 10 or 25 µM for TPP+ and ethidium, respectively, followed by incubation at room temperature for 30 min. Higher concentrations of EmrE are required for weaker binding ligands (Note 1) 4. Lipid vesicles are diluted 90-fold into the appropriate pH reconstitution buffer, stirred at room temperature for 1 h. 5. Proteoliposomes are recovered by centrifugation at 370,000× g for 50 min, followed by resuspension in 2.2 ml of reconstitution buffer. Reconstitution efficiency varies depending on the lipid composition. Therefore, quantification of the amount of functional protein present in the bilayer is essential for activity measurements of either transport or binding assays.
3.3. Isothermal Titration Calorimetry (ITC)
1. All ITC experiments are performed with a VP-ITC microcalorimeter (MicroCal), using supplied Origin 5.0 software (MicroCal) for data analysis. 2. ITC experiments provide a large amount of information, but are susceptible to a number of artifacts that are not normally observed in other binding assays. It is essential that buffers used in ITC experiments are identical (Notes 2–4). 3. All samples are thoroughly degassed and equilibrated to 5°C below the experimental temperature before use with a Thermovac (MicroCal). 4. Experiments are performed at 25°C, monitoring the change in power to the measurement cell compared to the reference cell.
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The integral over time of the differential power is equal to the thermal energy (DH), and is dependent on the binding reaction monitored in the particular buffer conditions (Note 5). Typically, experiments are performed using 1 initial injection of 7.5 µl of ligand, followed by 17 injections of 15 µl or using 1 initial injection of 10 µl of ligand, followed by 14 injections of 20 µl for TPP+ and ethidium binding experiments, respectively. Ligand concentrations between 20 and 40 µM TPP+ and 100–200 µM ethidium are used depending on the lipid composition and the percentage reconstitution of EmrE. Higher concentrations of ligand are required in experiments with a greater percentage of reconstituted EmrE to ensure saturation of binding sites after addition of all ligands (Note 6). 5. Control experiments to determine the heats of dilution of ligand are performed using lipid vesicles in the absence of EmrE (for TPP+ and ethidium binding experiments) and lipid vesicles containing a non-functional EmrE mutant, E14C (for ethidium binding experiments). Control heats of dilution for ligand-binding experiments were subtracted from either heats of dilution at saturation of EmrE or controls performed with ligand and lipid vesicles containing no protein; these two approaches are often identical. Controls are difficult to subtract from ethidium binding data because of differences in the background ethidium binding properties of liposomes. Control heats of dilution are determined to be linear for ethidium plus liposomes with no protein, and ethidium plus lipid vesicles containing inactive E14C EmrE. Heats of dilution for ethidium binding experiments were determined by subtracting a linear background determined from binding data after saturation. 6. The first data point in ITC binding experiments is always excluded from analysis because of premixing of the ligand solution in the injection syringe tip. After subtraction of a linear correction for control heats of TPP+ and ethidium dilution, binding data are fitted to a single-site binding model using the equations (14): 2 4X t nM t ∆HV o Xt Xt 1 1 1+ + − 1 + + − Q = nM t nKM t 2 nM t nKM t nM t �
