Liposomes, Part F (Methods in Enzymology, Volume 464)

VOLUME FOUR HUNDRED AND SIXTY-FOUR

METHODS

IN

ENZYMOLOGY Liposomes, Part F

METHODS IN ENZYMOLOGY Editors-in-Chief

JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of Technology Pasadena, California, USA Founding Editors

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

VOLUME FOUR HUNDRED AND SIXTY-FOUR

METHODS

IN

ENZYMOLOGY Liposomes, Part F EDITED BY

¨ ZGU ¨ NES¸ NEJAT DU Department of Microbiology University of the Pacific Arthur A. Dugoni School of Dentistry San Francisco, California, USA

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32 Jamestown Road, London NW1 7BY, UK First edition 2009 Copyright # 2009, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@ elsevier.com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made For information on all Academic Press publications visit our website at elsevierdirect.com ISBN: 978-0-12-374969-7 ISSN: 0076-6879 Printed and bound in United States of America 09 10 11 12 10 9 8 7 6 5 4 3 2 1

CONTENTS

Contributors Preface Volumes in Series

Section I. Bioactive Liposomes 1. Tubular Liposomes with Variable Permeability for Reconstitution of FtsZ Rings

xi xvii xix

1

3

Masaki Osawa and Harold P. Erickson 1. Introduction 2. Reagents 3. Bacterial Expression of Membrane Targeting FtsZ 4. Purification of FtsZ-mts and FtsZ-YFP-mts 5. Renatured Preparation of FtsZ-YFP-mts 6. Tubular Multilamellar Liposome Preparation 7. Permeability of the Multilamellar Liposomes 8. Z-ring Formation in Liposomes 9. A Crude Flow Chamber to Exchange Buffer Outside Liposomes 10. Factors Affecting Z-ring Formation in Liposomes 11. Utility of the Liposomes Beyond FtsZ References

2. Detection and Analysis of Protein Synthesis and RNA Replication in Giant Liposomes

4 4 5 5 7 7 9 11 11 14 16 16

19

Takeshi Sunami, Hiroshi Kita, Kazufumi Hosoda, Tomoaki Matsuura, Hiroaki Suzuki, and Tetsuya Yomo 1. Introduction 2. Methods 3. Analysis of the FACS Data 4. Conclusions Acknowledgments References

20 21 27 28 29 29

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Contents

3. Construction of Cell-Sized Liposomes Encapsulating Actin and Actin-Cross-Linking Proteins

31

Kingo Takiguchi, Ayako Yamada, Makiko Negishi, Makoto Honda, Yohko Tanaka-Takiguchi, and Kenichi Yoshikawa 1. Introduction 2. Experimental Section 3. Morphogenesis of Giant Liposomes Encapsulating Actin and Its Cross-linking Proteins 4. Concluding Remarks Acknowledgments References

4. Reconstitution of Membrane Budding with Unilamellar Vesicles

32 37 42 49 50 50

55

Anna V. Shnyrova and Joshua Zimmerberg 1. Introduction 2. M Protein Purification 3. Evaluation of the Membrane Activity of M Protein Through its Interaction with Intermediate-Sized Unilamellar Liposomes 4. Reconstitution of M-Protein-Driven Membrane Budding on GUVs 5. Concluding Remarks References

Section II. Liposomes and Nanotechnology 5. Detection of Antimycolic Acid Antibodies by Liposomal Biosensors

56 57 58 66 73 73

77

79

Y. Lemmer, S. T. Thanyani, P. J. Vrey, C. H. S. Driver, L. Venter, S. van Wyngaardt, A. M. C. ten Bokum, K. I. Ozoemena, L. A. Pilcher, D. G. Fernig, A. C. Stoltz, H. S. Swai, and J. A. Verschoor 1. Introduction 2. Experimental 3. Conclusion Acknowledgments References

6. Solid Lipid Nanoparticle Formulations: Pharmacokinetic and Biopharmaceutical Aspects in Drug Delivery

80 81 102 102 102

105

Eliana B. Souto and Slavomira Doktorovova´ 1. Introduction 2. Production of SLN

106 107

Contents

3. Pharmacokinetics and Pharmacodynamics 4. Modified Release Profile 5. Biopharmaceutical Aspects of Administration Routes 6. Clinical Pharmacology 7. Concluding Remarks References

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107 112 114 117 121 122

7. Preparation of Complexes of Liposomes with Gold Nanoparticles 131 Chie Kojima, Yusuke Hirano, and Kenji Kono 1. 2. 3. 4. 5. 6. 7.

Introduction Preparation of Complexes of EYPC Liposomes with Au NPs Time-Dependent SPR of the Complexes TEM Analysis of the Complexes DLS Analysis of the Complexes Calcein Release from the Complexes Estimation of Numbers of the Au NP and the Liposome in the Complexes 8. Optimization of Lipid Components of the Complexes 9. Concluding Remarks Acknowledgment References

8. Bio-Nanocapsule–Liposome Conjugates for In Vivo Pinpoint Drug and Gene Delivery

132 134 134 136 137 137 139 140 142 143 144

147

Takeshi Kasuya, Joohee Jung, Rie Kinoshita, Yasumasa Goh, Takashi Matsuzaki, Masumi Iijima, Nobuo Yoshimoto, Katsuyuki Tanizawa, and Shun’ichi Kuroda 1. Introduction 2. First-Generation Bio-Nanocapsules 3. Second-Generation BNCs 4. Retargeting of BNC–LP Conjugates 5. Overexpression of BNCs in S. cerevisiae 6. Conjugation of BNCs with LPs 7. Preparation of Antibody-Displaying BNC–LP Conjugates 8. Preparation of Biotin-Displaying BNC–LP Conjugates 9. Concluding Remarks Acknowledgments References

148 149 150 152 153 155 161 163 163 164 164

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9. Nanoliposomal Dry Powder Formulations

167

Gaurang Patel, Mahavir Chougule, Mandip Singh, and Ambikanandan Misra 1. Introduction 2. Preparation of Nanoliposomal DPFs 3. Physicochemical Characterization of NLDPFs 4. Concluding Remarks References

10. Lanthanide-Loaded Paramagnetic Liposomes as Switchable Magnetically Oriented Nanovesicles

168 169 174 187 188

193

Silvio Aime, Daniela Delli Castelli, and Enzo Terreno 1. 2. 3. 4.

Introduction Paramagnetic Ln(III)-Based Shift Reagents Preparation of Osmotically Shrunken Liposomes NMR Characterization of Magnetically Oriented Nonspherical Liposomes 5. Sample Experiments 6. Concluding Remarks Acknowledgments References

11. Reconstitution of Membrane Proteins in Phospholipid Bilayer Nanodiscs

194 195 197 198 200 208 208 208

211

T. K. Ritchie, Y. V. Grinkova, T. H. Bayburt, I. G. Denisov, J. K. Zolnerciks, W. M. Atkins, and S. G. Sligar 1. Introduction 2. Overview of Nanodisc Technology 3. Reconstitution Considerations 4. Optimizing the Reconstitution for P-glycoprotein Acknowledgments References

12. DNA-Controlled Assembly of Liposomes in Diagnostics

212 212 218 223 228 228

233

Ulla Jakobsen and Stefan Vogel 1. Introduction 2. Probe Design 3. General Description of Materials and Techniques 4. Concluding Remarks Acknowledgment References

234 235 244 247 248 248

Contents

13. Soft Hybrid Nanostructures Composed of Phospholipid Liposomes Decorated with Oligonucleotides

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Martina Banchelli, Francesca Baldelli Bombelli, Debora Berti, and Piero Baglioni 1. Introduction 2. Materials 3. Liposome Preparation and Determination of Lipid Content 4. Incorporation of Oligonucleotides 5. Characterization of the Soft Hybrid Nanostructure 6. Applications of Oligo-Decorated Liposomes 7. Challenges and Perspectives Acknowledgments References

250 251 252 253 256 262 275 276 276

14. Synthesis, Characterization, and Optical Response of Gold Nanoshells Used to Trigger Release from Liposomes

279

Guohui Wu, Alexander Mikhailovsky, Htet A. Khant, and Joseph A. Zasadzinski 1. Introduction 2. Synthesis of HGNs 3. Optimization of HGN Dimensions for Maximum Absorption in the NIR 4. HGN Response to Femtosecond NIR Laser Pulses 5. Coupling HGN to Liposomes 6. Liposome Disruption and CF Release Due to Pulsed Laser Irradiation 7. Mechanism of Triggered Liposome Release 8. Effect of Proximity of HGNs to Liposomes 9. Conclusions Acknowledgments References

15. Complex Nanotube-Liposome Networks

280 283 286 290 294 299 300 303 304 304 304

309

Aldo Jesorka and Owe Orwar 1. Introduction 2. Network Fabrication Protocols 3. Complexity and Topology 4. Internal and Membrane Functionalization 5. Transport Phenomena and Controlled Mixing Procedures 6. Enzymatic Reactions in NVN 7. Concluding Remarks Acknowledgments References

309 310 314 315 318 320 323 323 324

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Contents

16. Bionanotubules Formed from Liposomes

327

Josemar A. Castillo and Mark A. Hayes 1. Introduction 2. Bionanotubule Formation by Applying Electric Fields to Surface-Attached Liposomes 3. Bionanotubular Formation from Liposomes in Solution Using Electric Fields 4. Other Methods of Bionanotubular Formation from Liposomes 5. Concluding Remarks References

17. Engineering Cationic Liposome: siRNA Complexes for In Vitro and In Vivo Delivery

328 329 334 337 339 339

343

Jennifer E. Podesta and Kostas Kostarelos 1. Introduction 2. Cationic Liposome Systems for siRNA Delivery 3. Experimental Methods 4. Troubleshooting 5. Concluding Remarks References Author Index Subject Index

344 345 347 352 353 354 355 363

CONTRIBUTORS

Silvio Aime Department of Chemistry IFM and Molecular Imaging Center, University of Torino, Torino, Italy W. M. Atkins Department of Medicinal Chemistry, University of Washington, Seattle, Washington, USA Piero Baglioni Department of Chemistry and CSGI, University of Florence, Sesto Fiorentino, Florence, Italy Martina Banchelli Department of Chemistry and CSGI, University of Florence, Sesto Fiorentino, Florence, Italy T. H. Bayburt Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois, USA Debora Berti Department of Chemistry and CSGI, University of Florence, Sesto Fiorentino, Florence, Italy Francesca Baldelli Bombelli Department of Chemistry and CSGI, University of Florence, Sesto Fiorentino, Florence, Italy Daniela Delli Castelli Department of Chemistry IFM and Molecular Imaging Center, University of Torino, Torino, Italy Josemar A. Castillo Department of Chemistry and Biochemistry, Arizona State University, Tempe, Arizona, USA Mahavir Chougule Pharmaceutics Department, College of Pharmacy and Pharmaceutical Science, Florida A&M University, Tallahassee, Florida, USA

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Contributors

I. G. Denisov Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois, USA Slavomira Doktorovova´ Department of Pharmaceutical Technology, Faculty of Health Sciences, Fernando Pessoa University, Porto, Portugal C. H. S. Driver Department of Chemistry, University of Pretoria, Pretoria, South Africa Harold P. Erickson Department of Cell Biology, Duke University Medical Center, Durham, North Carolina, USA D. G. Fernig School of Biological Sciences, University of Liverpool, United Kingdom Yasumasa Goh Beacle Inc., ORIC, Haga, Okayama, Japan Y. V. Grinkova Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois, USA Mark A. Hayes Department of Chemistry and Biochemistry, Arizona State University, Tempe, Arizona, USA Yusuke Hirano Graduate School of Engineering, Osaka Prefecture University, Osaka, Japan Makoto Honda Division of Biological Science, Graduate School of Science, Nagoya University, Furo-cho, Chikusa-ku, Nagoya, Japan Current address: Stem Cell and Drug Discovery Institute, Shimogyo-ku, Kyoto, Japan Kazufumi Hosoda Department of Bioinformatics Engineering, Graduate School of Information Science and Technology, Osaka University, Suita, Osaka, Japan Masumi Iijima Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan, and Graduate School of Bioagricultural Sciences, Nagoya University, Chikusa, Nagoya, Japan Ulla Jakobsen Nucleic Acid Center, University of Southern Denmark, Odense, Denmark

Contributors

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Aldo Jesorka Department of Chemical and Biological Engineering, Chalmers University of Technology, Go¨teborg, Sweden Joohee Jung Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan Present address: Institute for Innovative Cancer Research, ASAN Medical Center, Pungnap-2, Songpa, Seoul, Korea Takeshi Kasuya Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan Htet A. Khant Department of Chemical Engineering, University of California, Santa Barbara, California, USA Rie Kinoshita Beacle Inc., ORIC, Haga, Okayama, Japan Hiroshi Kita Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Agency ( JST), Chiyoda-ku, Tokyo, Japan Chie Kojima Nanoscience and Nanotechnology Research Center, Research Institutes for the Twenty First Century, Osaka Prefecture University, Osaka, Japan Kenji Kono Graduate School of Engineering, Osaka Prefecture University, Osaka, Japan Kostas Kostarelos Nanomedicine Laboratory, Centre for Drug Delivery Research, The School of Pharmacy, University of London, London, United Kingdom Shun’ichi Kuroda Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan, and Beacle Inc., ORIC, Haga, Okayama, Japan; Graduate School of Bioagricultural Sciences, Nagoya University, Chikusa, Nagoya, Japan Y. Lemmer Department of Biochemistry, University of Pretoria, Pretoria, South Africa, and Materials Science and Manufacturing, CSIR, Pretoria, South Africa Tomoaki Matsuura Department of Bioinformatics Engineering, Graduate School of Information Science and Technology, Osaka University, Suita, Osaka, Japan

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Contributors

Takashi Matsuzaki Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan Alexander Mikhailovsky Department of Chemistry, University of California, Santa Barbara, California, USA Ambikanandan Misra TIFAC-CORE in NDDS, Pharmacy Department, Faculty of Technology and Engineering, The Maharaja Sayajirao University of Baroda, Kalabhavan, Vadodara, Gujarat, India Makiko Negishi Department of Physics, Graduate School of Science, Kyoto University, Sakyo-ku, Kyoto, Japan Owe Orwar Department of Chemical and Biological Engineering, Chalmers University of Technology, Go¨teborg, Sweden Masaki Osawa Department of Cell Biology, Duke University Medical Center, Durham, North Carolina, USA K. I. Ozoemena Department of Chemistry, University of Pretoria, Pretoria, South Africa Gaurang Patel TIFAC-CORE in NDDS, Pharmacy Department, Faculty of Technology and Engineering, The Maharaja Sayajirao University of Baroda, Kalabhavan, Vadodara, Gujarat, India L. A. Pilcher Department of Chemistry, University of Pretoria, Pretoria, South Africa Jennifer E. Podesta Nanomedicine Laboratory, Centre for Drug Delivery Research, The School of Pharmacy, University of London, London, United Kingdom T. K. Ritchie Department of Medicinal Chemistry, University of Washington, Seattle, Washington, USA Anna V. Shnyrova Laboratory of Cellular and Molecular Biology, Program in Physical Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, Maryland, USA Mandip Singh Pharmaceutics Department, College of Pharmacy and Pharmaceutical Science, Florida A&M University, Tallahassee, Florida, USA

Contributors

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S. G. Sligar Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois, USA Eliana B. Souto Department of Pharmaceutical Technology, Faculty of Health Sciences, Fernando Pessoa University, Porto, Portugal, and Institute of Biotechnology and Bioengineering, Centre of Genetics and Biotechnology, University of Tra´s-os-Montes and Alto Douro (CGB-UTAD/IBB), Vila Real, Portugal A. C. Stoltz Department of Biochemistry, and Department of Infectious Diseases, University of Pretoria, Pretoria, South Africa Takeshi Sunami Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Agency ( JST), Chiyoda-ku, Tokyo, Japan Hiroaki Suzuki Department of Bioinformatics Engineering, Graduate School of Information Science and Technology, Osaka University, Suita, Osaka, Japan H. S. Swai Materials Science and Manufacturing, CSIR, Pretoria, South Africa Kingo Takiguchi Division of Biological Science, Graduate School of Science, Nagoya University, Furo-cho, Chikusa-ku, Nagoya, Japan Yohko Tanaka-Takiguchi Division of Biological Science, Graduate School of Science, Nagoya University, Furo-cho, Chikusa-ku, Nagoya, Japan Katsuyuki Tanizawa Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan A. M. C. ten Bokum Department of Biochemistry, University of Pretoria, Pretoria, South Africa Enzo Terreno Department of Chemistry IFM and Molecular Imaging Center, University of Torino, Torino, Italy S. T. Thanyani Department of Biochemistry, University of Pretoria, Pretoria, South Africa S. van Wyngaardt Department of Biochemistry, University of Pretoria, Pretoria, South Africa

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Contributors

L. Venter Department of Biochemistry, University of Pretoria, Pretoria, South Africa J. A. Verschoor Department of Biochemistry, University of Pretoria, Pretoria, South Africa Stefan Vogel Nucleic Acid Center, University of Southern Denmark, Odense, Denmark P. J. Vrey Department of Biochemistry, University of Pretoria, Pretoria, South Africa Guohui Wu Department of Chemical Engineering, University of California, Santa Barbara, California, USA Ayako Yamada Department of Physics, Graduate School of Science, Kyoto University, Sakyo-ku, Kyoto, Japan Current address: Department of Chemistry, Ecole Normale Superieure, Paris, France Tetsuya Yomo Department of Bioinformatics Engineering, Graduate School of Information Science and Technology, and Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan, and Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Agency ( JST), Chiyoda-ku, Tokyo, Japan Kenichi Yoshikawa Department of Physics, Graduate School of Science, Kyoto University, Sakyo-ku, Kyoto, Japan Nobuo Yoshimoto Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan, and Graduate School of Bioagricultural Sciences, Nagoya University, Chikusa, Nagoya, Japan Joseph A. Zasadzinski Department of Chemical Engineering, University of California, Santa Barbara, California, USA Joshua Zimmerberg Laboratory of Cellular and Molecular Biology, Program in Physical Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, Maryland, USA J. K. Zolnerciks Department of Medicinal Chemistry, University of Washington, Seattle, Washington, USA

PREFACE

Previous Methods in Enzymology volumes on ‘‘Liposomes’’ have described methods of liposome preparation and the physicochemical characterization of liposomes (Volume 367), and the use of liposomes in biochemistry, molecular cell biology (Volume 372), immunology, diagnostics, gene delivery, and gene therapy (Volume 373). Methods involved in the production and application of antibody- or ligand-targeted liposomes, environmentsensitive liposomes, and liposomal oligonucleotides were provided in Volume 387, as were methods for studying the in vivo fate of liposomes. Finally, Volume 391 presented methods in liposomal anticancer, antibacterial, antifungal, and antiviral agents, miscellaneous liposomal therapies and electron microscopy of liposomes. The latter volume also included a short introductory chapter on ‘‘The Origin of Liposomes: Alec Bangham at Babraham.’’ This new volume includes sections focusing on bioactive liposomes and the interface of liposomes and nanotechnology. I hope that these chapters will be helpful to graduate students, postdoctoral fellows, research associates, and established scientists initiating projects on liposomes, or shifting the focus of their research. Although the chapters are not written in ‘‘protocol’’ format, they describe the experimental methods in sufficient detail that can be adopted readily for the reader’s project. In addition, the chapters provide the perspective of the authors on the field, as well as examples of results obtained with the described methods. I would like to thank all the colleagues who graciously contributed to this volume with relatively short notice, Associate Editor Tara Hoey, Editorial Services Manager Delsy Retchagar, and Production Manager Radhakrishnan Lakshmanan of Elsevier for their help in preparing this volume, and Shirley Light, formerly of Academic Press, for her initiation of the ‘‘Liposomes’’ volumes about a decade ago. I would also like to express my gratitude to my supportive and loving family. I dedicate this volume to my wife, Diana, and my curious, creative, playful, and loving children, Avery and Maxine. NEJAT DU¨ZGU¨NES¸

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METHODS IN ENZYMOLOGY

VOLUME I. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME II. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME III. Preparation and Assay of Substrates Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME IV. Special Techniques for the Enzymologist Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME V. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VI. Preparation and Assay of Enzymes (Continued) Preparation and Assay of Substrates Special Techniques Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VII. Cumulative Subject Index Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VIII. Complex Carbohydrates Edited by ELIZABETH F. NEUFELD AND VICTOR GINSBURG VOLUME IX. Carbohydrate Metabolism Edited by WILLIS A. WOOD VOLUME X. Oxidation and Phosphorylation Edited by RONALD W. ESTABROOK AND MAYNARD E. PULLMAN VOLUME XI. Enzyme Structure Edited by C. H. W. HIRS VOLUME XII. Nucleic Acids (Parts A and B) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XIII. Citric Acid Cycle Edited by J. M. LOWENSTEIN VOLUME XIV. Lipids Edited by J. M. LOWENSTEIN VOLUME XV. Steroids and Terpenoids Edited by RAYMOND B. CLAYTON

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VOLUME XVI. Fast Reactions Edited by KENNETH KUSTIN VOLUME XVII. Metabolism of Amino Acids and Amines (Parts A and B) Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME XVIII. Vitamins and Coenzymes (Parts A, B, and C) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME XIX. Proteolytic Enzymes Edited by GERTRUDE E. PERLMANN AND LASZLO LORAND VOLUME XX. Nucleic Acids and Protein Synthesis (Part C) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXI. Nucleic Acids (Part D) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXII. Enzyme Purification and Related Techniques Edited by WILLIAM B. JAKOBY VOLUME XXIII. Photosynthesis (Part A) Edited by ANTHONY SAN PIETRO VOLUME XXIV. Photosynthesis and Nitrogen Fixation (Part B) Edited by ANTHONY SAN PIETRO VOLUME XXV. Enzyme Structure (Part B) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVI. Enzyme Structure (Part C) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVII. Enzyme Structure (Part D) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVIII. Complex Carbohydrates (Part B) Edited by VICTOR GINSBURG VOLUME XXIX. Nucleic Acids and Protein Synthesis (Part E) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXX. Nucleic Acids and Protein Synthesis (Part F) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXXI. Biomembranes (Part A) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXII. Biomembranes (Part B) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXIII. Cumulative Subject Index Volumes I-XXX Edited by MARTHA G. DENNIS AND EDWARD A. DENNIS VOLUME XXXIV. Affinity Techniques (Enzyme Purification: Part B) Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK

Methods in Enzymology

VOLUME XXXV. Lipids (Part B) Edited by JOHN M. LOWENSTEIN VOLUME XXXVI. Hormone Action (Part A: Steroid Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVII. Hormone Action (Part B: Peptide Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVIII. Hormone Action (Part C: Cyclic Nucleotides) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XXXIX. Hormone Action (Part D: Isolated Cells, Tissues, and Organ Systems) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XL. Hormone Action (Part E: Nuclear Structure and Function) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XLI. Carbohydrate Metabolism (Part B) Edited by W. A. WOOD VOLUME XLII. Carbohydrate Metabolism (Part C) Edited by W. A. WOOD VOLUME XLIII. Antibiotics Edited by JOHN H. HASH VOLUME XLIV. Immobilized Enzymes Edited by KLAUS MOSBACH VOLUME XLV. Proteolytic Enzymes (Part B) Edited by LASZLO LORAND VOLUME XLVI. Affinity Labeling Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK VOLUME XLVII. Enzyme Structure (Part E) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLVIII. Enzyme Structure (Part F) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLIX. Enzyme Structure (Part G) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME L. Complex Carbohydrates (Part C) Edited by VICTOR GINSBURG VOLUME LI. Purine and Pyrimidine Nucleotide Metabolism Edited by PATRICIA A. HOFFEE AND MARY ELLEN JONES VOLUME LII. Biomembranes (Part C: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER

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VOLUME LIII. Biomembranes (Part D: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LIV. Biomembranes (Part E: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LV. Biomembranes (Part F: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVI. Biomembranes (Part G: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVII. Bioluminescence and Chemiluminescence Edited by MARLENE A. DELUCA VOLUME LVIII. Cell Culture Edited by WILLIAM B. JAKOBY AND IRA PASTAN VOLUME LIX. Nucleic Acids and Protein Synthesis (Part G) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME LX. Nucleic Acids and Protein Synthesis (Part H) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME 61. Enzyme Structure (Part H) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 62. Vitamins and Coenzymes (Part D) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 63. Enzyme Kinetics and Mechanism (Part A: Initial Rate and Inhibitor Methods) Edited by DANIEL L. PURICH VOLUME 64. Enzyme Kinetics and Mechanism (Part B: Isotopic Probes and Complex Enzyme Systems) Edited by DANIEL L. PURICH VOLUME 65. Nucleic Acids (Part I) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME 66. Vitamins and Coenzymes (Part E) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 67. Vitamins and Coenzymes (Part F) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 68. Recombinant DNA Edited by RAY WU VOLUME 69. Photosynthesis and Nitrogen Fixation (Part C) Edited by ANTHONY SAN PIETRO VOLUME 70. Immunochemical Techniques (Part A) Edited by HELEN VAN VUNAKIS AND JOHN J. LANGONE

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VOLUME 71. Lipids (Part C) Edited by JOHN M. LOWENSTEIN VOLUME 72. Lipids (Part D) Edited by JOHN M. LOWENSTEIN VOLUME 73. Immunochemical Techniques (Part B) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 74. Immunochemical Techniques (Part C) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 75. Cumulative Subject Index Volumes XXXI, XXXII, XXXIV–LX Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 76. Hemoglobins Edited by ERALDO ANTONINI, LUIGI ROSSI-BERNARDI, AND EMILIA CHIANCONE VOLUME 77. Detoxication and Drug Metabolism Edited by WILLIAM B. JAKOBY VOLUME 78. Interferons (Part A) Edited by SIDNEY PESTKA VOLUME 79. Interferons (Part B) Edited by SIDNEY PESTKA VOLUME 80. Proteolytic Enzymes (Part C) Edited by LASZLO LORAND VOLUME 81. Biomembranes (Part H: Visual Pigments and Purple Membranes, I) Edited by LESTER PACKER VOLUME 82. Structural and Contractile Proteins (Part A: Extracellular Matrix) Edited by LEON W. CUNNINGHAM AND DIXIE W. FREDERIKSEN VOLUME 83. Complex Carbohydrates (Part D) Edited by VICTOR GINSBURG VOLUME 84. Immunochemical Techniques (Part D: Selected Immunoassays) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 85. Structural and Contractile Proteins (Part B: The Contractile Apparatus and the Cytoskeleton) Edited by DIXIE W. FREDERIKSEN AND LEON W. CUNNINGHAM VOLUME 86. Prostaglandins and Arachidonate Metabolites Edited by WILLIAM E. M. LANDS AND WILLIAM L. SMITH VOLUME 87. Enzyme Kinetics and Mechanism (Part C: Intermediates, Stereo-chemistry, and Rate Studies) Edited by DANIEL L. PURICH VOLUME 88. Biomembranes (Part I: Visual Pigments and Purple Membranes, II) Edited by LESTER PACKER

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VOLUME 89. Carbohydrate Metabolism (Part D) Edited by WILLIS A. WOOD VOLUME 90. Carbohydrate Metabolism (Part E) Edited by WILLIS A. WOOD VOLUME 91. Enzyme Structure (Part I) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 92. Immunochemical Techniques (Part E: Monoclonal Antibodies and General Immunoassay Methods) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 93. Immunochemical Techniques (Part F: Conventional Antibodies, Fc Receptors, and Cytotoxicity) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 94. Polyamines Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME 95. Cumulative Subject Index Volumes 61–74, 76–80 Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 96. Biomembranes [Part J: Membrane Biogenesis: Assembly and Targeting (General Methods; Eukaryotes)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 97. Biomembranes [Part K: Membrane Biogenesis: Assembly and Targeting (Prokaryotes, Mitochondria, and Chloroplasts)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 98. Biomembranes (Part L: Membrane Biogenesis: Processing and Recycling) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 99. Hormone Action (Part F: Protein Kinases) Edited by JACKIE D. CORBIN AND JOEL G. HARDMAN VOLUME 100. Recombinant DNA (Part B) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 101. Recombinant DNA (Part C) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 102. Hormone Action (Part G: Calmodulin and Calcium-Binding Proteins) Edited by ANTHONY R. MEANS AND BERT W. O’MALLEY VOLUME 103. Hormone Action (Part H: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 104. Enzyme Purification and Related Techniques (Part C) Edited by WILLIAM B. JAKOBY

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VOLUME 105. Oxygen Radicals in Biological Systems Edited by LESTER PACKER VOLUME 106. Posttranslational Modifications (Part A) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 107. Posttranslational Modifications (Part B) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 108. Immunochemical Techniques (Part G: Separation and Characterization of Lymphoid Cells) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 109. Hormone Action (Part I: Peptide Hormones) Edited by LUTZ BIRNBAUMER AND BERT W. O’MALLEY VOLUME 110. Steroids and Isoprenoids (Part A) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 111. Steroids and Isoprenoids (Part B) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 112. Drug and Enzyme Targeting (Part A) Edited by KENNETH J. WIDDER AND RALPH GREEN VOLUME 113. Glutamate, Glutamine, Glutathione, and Related Compounds Edited by ALTON MEISTER VOLUME 114. Diffraction Methods for Biological Macromolecules (Part A) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 115. Diffraction Methods for Biological Macromolecules (Part B) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 116. Immunochemical Techniques (Part H: Effectors and Mediators of Lymphoid Cell Functions) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 117. Enzyme Structure (Part J) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 118. Plant Molecular Biology Edited by ARTHUR WEISSBACH AND HERBERT WEISSBACH VOLUME 119. Interferons (Part C) Edited by SIDNEY PESTKA VOLUME 120. Cumulative Subject Index Volumes 81–94, 96–101 VOLUME 121. Immunochemical Techniques (Part I: Hybridoma Technology and Monoclonal Antibodies) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 122. Vitamins and Coenzymes (Part G) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK

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VOLUME 123. Vitamins and Coenzymes (Part H) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK VOLUME 124. Hormone Action (Part J: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 125. Biomembranes (Part M: Transport in Bacteria, Mitochondria, and Chloroplasts: General Approaches and Transport Systems) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 126. Biomembranes (Part N: Transport in Bacteria, Mitochondria, and Chloroplasts: Protonmotive Force) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 127. Biomembranes (Part O: Protons and Water: Structure and Translocation) Edited by LESTER PACKER VOLUME 128. Plasma Lipoproteins (Part A: Preparation, Structure, and Molecular Biology) Edited by JERE P. SEGREST AND JOHN J. ALBERS VOLUME 129. Plasma Lipoproteins (Part B: Characterization, Cell Biology, and Metabolism) Edited by JOHN J. ALBERS AND JERE P. SEGREST VOLUME 130. Enzyme Structure (Part K) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 131. Enzyme Structure (Part L) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 132. Immunochemical Techniques (Part J: Phagocytosis and Cell-Mediated Cytotoxicity) Edited by GIOVANNI DI SABATO AND JOHANNES EVERSE VOLUME 133. Bioluminescence and Chemiluminescence (Part B) Edited by MARLENE DELUCA AND WILLIAM D. MCELROY VOLUME 134. Structural and Contractile Proteins (Part C: The Contractile Apparatus and the Cytoskeleton) Edited by RICHARD B. VALLEE VOLUME 135. Immobilized Enzymes and Cells (Part B) Edited by KLAUS MOSBACH VOLUME 136. Immobilized Enzymes and Cells (Part C) Edited by KLAUS MOSBACH VOLUME 137. Immobilized Enzymes and Cells (Part D) Edited by KLAUS MOSBACH VOLUME 138. Complex Carbohydrates (Part E) Edited by VICTOR GINSBURG

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VOLUME 139. Cellular Regulators (Part A: Calcium- and Calmodulin-Binding Proteins) Edited by ANTHONY R. MEANS AND P. MICHAEL CONN VOLUME 140. Cumulative Subject Index Volumes 102–119, 121–134 VOLUME 141. Cellular Regulators (Part B: Calcium and Lipids) Edited by P. MICHAEL CONN AND ANTHONY R. MEANS VOLUME 142. Metabolism of Aromatic Amino Acids and Amines Edited by SEYMOUR KAUFMAN VOLUME 143. Sulfur and Sulfur Amino Acids Edited by WILLIAM B. JAKOBY AND OWEN GRIFFITH VOLUME 144. Structural and Contractile Proteins (Part D: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 145. Structural and Contractile Proteins (Part E: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 146. Peptide Growth Factors (Part A) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 147. Peptide Growth Factors (Part B) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 148. Plant Cell Membranes Edited by LESTER PACKER AND ROLAND DOUCE VOLUME 149. Drug and Enzyme Targeting (Part B) Edited by RALPH GREEN AND KENNETH J. WIDDER VOLUME 150. Immunochemical Techniques (Part K: In Vitro Models of B and T Cell Functions and Lymphoid Cell Receptors) Edited by GIOVANNI DI SABATO VOLUME 151. Molecular Genetics of Mammalian Cells Edited by MICHAEL M. GOTTESMAN VOLUME 152. Guide to Molecular Cloning Techniques Edited by SHELBY L. BERGER AND ALAN R. KIMMEL VOLUME 153. Recombinant DNA (Part D) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 154. Recombinant DNA (Part E) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 155. Recombinant DNA (Part F) Edited by RAY WU VOLUME 156. Biomembranes (Part P: ATP-Driven Pumps and Related Transport: The Na, K-Pump) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER

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VOLUME 157. Biomembranes (Part Q: ATP-Driven Pumps and Related Transport: Calcium, Proton, and Potassium Pumps) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 158. Metalloproteins (Part A) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 159. Initiation and Termination of Cyclic Nucleotide Action Edited by JACKIE D. CORBIN AND ROGER A. JOHNSON VOLUME 160. Biomass (Part A: Cellulose and Hemicellulose) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 161. Biomass (Part B: Lignin, Pectin, and Chitin) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 162. Immunochemical Techniques (Part L: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 163. Immunochemical Techniques (Part M: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 164. Ribosomes Edited by HARRY F. NOLLER, JR., AND KIVIE MOLDAVE VOLUME 165. Microbial Toxins: Tools for Enzymology Edited by SIDNEY HARSHMAN VOLUME 166. Branched-Chain Amino Acids Edited by ROBERT HARRIS AND JOHN R. SOKATCH VOLUME 167. Cyanobacteria Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 168. Hormone Action (Part K: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 169. Platelets: Receptors, Adhesion, Secretion (Part A) Edited by JACEK HAWIGER VOLUME 170. Nucleosomes Edited by PAUL M. WASSARMAN AND ROGER D. KORNBERG VOLUME 171. Biomembranes (Part R: Transport Theory: Cells and Model Membranes) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 172. Biomembranes (Part S: Transport: Membrane Isolation and Characterization) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER

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VOLUME 173. Biomembranes [Part T: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 174. Biomembranes [Part U: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 175. Cumulative Subject Index Volumes 135–139, 141–167 VOLUME 176. Nuclear Magnetic Resonance (Part A: Spectral Techniques and Dynamics) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 177. Nuclear Magnetic Resonance (Part B: Structure and Mechanism) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 178. Antibodies, Antigens, and Molecular Mimicry Edited by JOHN J. LANGONE VOLUME 179. Complex Carbohydrates (Part F) Edited by VICTOR GINSBURG VOLUME 180. RNA Processing (Part A: General Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 181. RNA Processing (Part B: Specific Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 182. Guide to Protein Purification Edited by MURRAY P. DEUTSCHER VOLUME 183. Molecular Evolution: Computer Analysis of Protein and Nucleic Acid Sequences Edited by RUSSELL F. DOOLITTLE VOLUME 184. Avidin-Biotin Technology Edited by MEIR WILCHEK AND EDWARD A. BAYER VOLUME 185. Gene Expression Technology Edited by DAVID V. GOEDDEL VOLUME 186. Oxygen Radicals in Biological Systems (Part B: Oxygen Radicals and Antioxidants) Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 187. Arachidonate Related Lipid Mediators Edited by ROBERT C. MURPHY AND FRANK A. FITZPATRICK VOLUME 188. Hydrocarbons and Methylotrophy Edited by MARY E. LIDSTROM VOLUME 189. Retinoids (Part A: Molecular and Metabolic Aspects) Edited by LESTER PACKER

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VOLUME 190. Retinoids (Part B: Cell Differentiation and Clinical Applications) Edited by LESTER PACKER VOLUME 191. Biomembranes (Part V: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 192. Biomembranes (Part W: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 193. Mass Spectrometry Edited by JAMES A. MCCLOSKEY VOLUME 194. Guide to Yeast Genetics and Molecular Biology Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 195. Adenylyl Cyclase, G Proteins, and Guanylyl Cyclase Edited by ROGER A. JOHNSON AND JACKIE D. CORBIN VOLUME 196. Molecular Motors and the Cytoskeleton Edited by RICHARD B. VALLEE VOLUME 197. Phospholipases Edited by EDWARD A. DENNIS VOLUME 198. Peptide Growth Factors (Part C) Edited by DAVID BARNES, J. P. MATHER, AND GORDON H. SATO VOLUME 199. Cumulative Subject Index Volumes 168–174, 176–194 VOLUME 200. Protein Phosphorylation (Part A: Protein Kinases: Assays, Purification, Antibodies, Functional Analysis, Cloning, and Expression) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 201. Protein Phosphorylation (Part B: Analysis of Protein Phosphorylation, Protein Kinase Inhibitors, and Protein Phosphatases) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 202. Molecular Design and Modeling: Concepts and Applications (Part A: Proteins, Peptides, and Enzymes) Edited by JOHN J. LANGONE VOLUME 203. Molecular Design and Modeling: Concepts and Applications (Part B: Antibodies and Antigens, Nucleic Acids, Polysaccharides, and Drugs) Edited by JOHN J. LANGONE VOLUME 204. Bacterial Genetic Systems Edited by JEFFREY H. MILLER VOLUME 205. Metallobiochemistry (Part B: Metallothionein and Related Molecules) Edited by JAMES F. RIORDAN AND BERT L. VALLEE

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VOLUME 206. Cytochrome P450 Edited by MICHAEL R. WATERMAN AND ERIC F. JOHNSON VOLUME 207. Ion Channels Edited by BERNARDO RUDY AND LINDA E. IVERSON VOLUME 208. Protein–DNA Interactions Edited by ROBERT T. SAUER VOLUME 209. Phospholipid Biosynthesis Edited by EDWARD A. DENNIS AND DENNIS E. VANCE VOLUME 210. Numerical Computer Methods Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 211. DNA Structures (Part A: Synthesis and Physical Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 212. DNA Structures (Part B: Chemical and Electrophoretic Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 213. Carotenoids (Part A: Chemistry, Separation, Quantitation, and Antioxidation) Edited by LESTER PACKER VOLUME 214. Carotenoids (Part B: Metabolism, Genetics, and Biosynthesis) Edited by LESTER PACKER VOLUME 215. Platelets: Receptors, Adhesion, Secretion (Part B) Edited by JACEK J. HAWIGER VOLUME 216. Recombinant DNA (Part G) Edited by RAY WU VOLUME 217. Recombinant DNA (Part H) Edited by RAY WU VOLUME 218. Recombinant DNA (Part I) Edited by RAY WU VOLUME 219. Reconstitution of Intracellular Transport Edited by JAMES E. ROTHMAN VOLUME 220. Membrane Fusion Techniques (Part A) Edited by NEJAT DU¨ZGU¨NES, VOLUME 221. Membrane Fusion Techniques (Part B) Edited by NEJAT DU¨ZGU¨NES, VOLUME 222. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part A: Mammalian Blood Coagulation Factors and Inhibitors) Edited by LASZLO LORAND AND KENNETH G. MANN

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VOLUME 223. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part B: Complement Activation, Fibrinolysis, and Nonmammalian Blood Coagulation Factors) Edited by LASZLO LORAND AND KENNETH G. MANN VOLUME 224. Molecular Evolution: Producing the Biochemical Data Edited by ELIZABETH ANNE ZIMMER, THOMAS J. WHITE, REBECCA L. CANN, AND ALLAN C. WILSON VOLUME 225. Guide to Techniques in Mouse Development Edited by PAUL M. WASSARMAN AND MELVIN L. DEPAMPHILIS VOLUME 226. Metallobiochemistry (Part C: Spectroscopic and Physical Methods for Probing Metal Ion Environments in Metalloenzymes and Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 227. Metallobiochemistry (Part D: Physical and Spectroscopic Methods for Probing Metal Ion Environments in Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 228. Aqueous Two-Phase Systems Edited by HARRY WALTER AND GO¨TE JOHANSSON VOLUME 229. Cumulative Subject Index Volumes 195–198, 200–227 VOLUME 230. Guide to Techniques in Glycobiology Edited by WILLIAM J. LENNARZ AND GERALD W. HART VOLUME 231. Hemoglobins (Part B: Biochemical and Analytical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 232. Hemoglobins (Part C: Biophysical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 233. Oxygen Radicals in Biological Systems (Part C) Edited by LESTER PACKER VOLUME 234. Oxygen Radicals in Biological Systems (Part D) Edited by LESTER PACKER VOLUME 235. Bacterial Pathogenesis (Part A: Identification and Regulation of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 236. Bacterial Pathogenesis (Part B: Integration of Pathogenic Bacteria with Host Cells) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 237. Heterotrimeric G Proteins Edited by RAVI IYENGAR VOLUME 238. Heterotrimeric G-Protein Effectors Edited by RAVI IYENGAR

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VOLUME 239. Nuclear Magnetic Resonance (Part C) Edited by THOMAS L. JAMES AND NORMAN J. OPPENHEIMER VOLUME 240. Numerical Computer Methods (Part B) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 241. Retroviral Proteases Edited by LAWRENCE C. KUO AND JULES A. SHAFER VOLUME 242. Neoglycoconjugates (Part A) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 243. Inorganic Microbial Sulfur Metabolism Edited by HARRY D. PECK, JR., AND JEAN LEGALL VOLUME 244. Proteolytic Enzymes: Serine and Cysteine Peptidases Edited by ALAN J. BARRETT VOLUME 245. Extracellular Matrix Components Edited by E. RUOSLAHTI AND E. ENGVALL VOLUME 246. Biochemical Spectroscopy Edited by KENNETH SAUER VOLUME 247. Neoglycoconjugates (Part B: Biomedical Applications) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 248. Proteolytic Enzymes: Aspartic and Metallo Peptidases Edited by ALAN J. BARRETT VOLUME 249. Enzyme Kinetics and Mechanism (Part D: Developments in Enzyme Dynamics) Edited by DANIEL L. PURICH VOLUME 250. Lipid Modifications of Proteins Edited by PATRICK J. CASEY AND JANICE E. BUSS VOLUME 251. Biothiols (Part A: Monothiols and Dithiols, Protein Thiols, and Thiyl Radicals) Edited by LESTER PACKER VOLUME 252. Biothiols (Part B: Glutathione and Thioredoxin; Thiols in Signal Transduction and Gene Regulation) Edited by LESTER PACKER VOLUME 253. Adhesion of Microbial Pathogens Edited by RON J. DOYLE AND ITZHAK OFEK VOLUME 254. Oncogene Techniques Edited by PETER K. VOGT AND INDER M. VERMA VOLUME 255. Small GTPases and Their Regulators (Part A: Ras Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL

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VOLUME 256. Small GTPases and Their Regulators (Part B: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 257. Small GTPases and Their Regulators (Part C: Proteins Involved in Transport) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 258. Redox-Active Amino Acids in Biology Edited by JUDITH P. KLINMAN VOLUME 259. Energetics of Biological Macromolecules Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 260. Mitochondrial Biogenesis and Genetics (Part A) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 261. Nuclear Magnetic Resonance and Nucleic Acids Edited by THOMAS L. JAMES VOLUME 262. DNA Replication Edited by JUDITH L. CAMPBELL VOLUME 263. Plasma Lipoproteins (Part C: Quantitation) Edited by WILLIAM A. BRADLEY, SANDRA H. GIANTURCO, AND JERE P. SEGREST VOLUME 264. Mitochondrial Biogenesis and Genetics (Part B) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 265. Cumulative Subject Index Volumes 228, 230–262 VOLUME 266. Computer Methods for Macromolecular Sequence Analysis Edited by RUSSELL F. DOOLITTLE VOLUME 267. Combinatorial Chemistry Edited by JOHN N. ABELSON VOLUME 268. Nitric Oxide (Part A: Sources and Detection of NO; NO Synthase) Edited by LESTER PACKER VOLUME 269. Nitric Oxide (Part B: Physiological and Pathological Processes) Edited by LESTER PACKER VOLUME 270. High Resolution Separation and Analysis of Biological Macromolecules (Part A: Fundamentals) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 271. High Resolution Separation and Analysis of Biological Macromolecules (Part B: Applications) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 272. Cytochrome P450 (Part B) Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 273. RNA Polymerase and Associated Factors (Part A) Edited by SANKAR ADHYA

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VOLUME 274. RNA Polymerase and Associated Factors (Part B) Edited by SANKAR ADHYA VOLUME 275. Viral Polymerases and Related Proteins Edited by LAWRENCE C. KUO, DAVID B. OLSEN, AND STEVEN S. CARROLL VOLUME 276. Macromolecular Crystallography (Part A) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 277. Macromolecular Crystallography (Part B) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 278. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 279. Vitamins and Coenzymes (Part I) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 280. Vitamins and Coenzymes (Part J) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 281. Vitamins and Coenzymes (Part K) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 282. Vitamins and Coenzymes (Part L) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 283. Cell Cycle Control Edited by WILLIAM G. DUNPHY VOLUME 284. Lipases (Part A: Biotechnology) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 285. Cumulative Subject Index Volumes 263, 264, 266–284, 286–289 VOLUME 286. Lipases (Part B: Enzyme Characterization and Utilization) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 287. Chemokines Edited by RICHARD HORUK VOLUME 288. Chemokine Receptors Edited by RICHARD HORUK VOLUME 289. Solid Phase Peptide Synthesis Edited by GREGG B. FIELDS VOLUME 290. Molecular Chaperones Edited by GEORGE H. LORIMER AND THOMAS BALDWIN VOLUME 291. Caged Compounds Edited by GERARD MARRIOTT VOLUME 292. ABC Transporters: Biochemical, Cellular, and Molecular Aspects Edited by SURESH V. AMBUDKAR AND MICHAEL M. GOTTESMAN

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VOLUME 293. Ion Channels (Part B) Edited by P. MICHAEL CONN VOLUME 294. Ion Channels (Part C) Edited by P. MICHAEL CONN VOLUME 295. Energetics of Biological Macromolecules (Part B) Edited by GARY K. ACKERS AND MICHAEL L. JOHNSON VOLUME 296. Neurotransmitter Transporters Edited by SUSAN G. AMARA VOLUME 297. Photosynthesis: Molecular Biology of Energy Capture Edited by LEE MCINTOSH VOLUME 298. Molecular Motors and the Cytoskeleton (Part B) Edited by RICHARD B. VALLEE VOLUME 299. Oxidants and Antioxidants (Part A) Edited by LESTER PACKER VOLUME 300. Oxidants and Antioxidants (Part B) Edited by LESTER PACKER VOLUME 301. Nitric Oxide: Biological and Antioxidant Activities (Part C) Edited by LESTER PACKER VOLUME 302. Green Fluorescent Protein Edited by P. MICHAEL CONN VOLUME 303. cDNA Preparation and Display Edited by SHERMAN M. WEISSMAN VOLUME 304. Chromatin Edited by PAUL M. WASSARMAN AND ALAN P. WOLFFE VOLUME 305. Bioluminescence and Chemiluminescence (Part C) Edited by THOMAS O. BALDWIN AND MIRIAM M. ZIEGLER VOLUME 306. Expression of Recombinant Genes in Eukaryotic Systems Edited by JOSEPH C. GLORIOSO AND MARTIN C. SCHMIDT VOLUME 307. Confocal Microscopy Edited by P. MICHAEL CONN VOLUME 308. Enzyme Kinetics and Mechanism (Part E: Energetics of Enzyme Catalysis) Edited by DANIEL L. PURICH AND VERN L. SCHRAMM VOLUME 309. Amyloid, Prions, and Other Protein Aggregates Edited by RONALD WETZEL VOLUME 310. Biofilms Edited by RON J. DOYLE

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VOLUME 311. Sphingolipid Metabolism and Cell Signaling (Part A) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 312. Sphingolipid Metabolism and Cell Signaling (Part B) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 313. Antisense Technology (Part A: General Methods, Methods of Delivery, and RNA Studies) Edited by M. IAN PHILLIPS VOLUME 314. Antisense Technology (Part B: Applications) Edited by M. IAN PHILLIPS VOLUME 315. Vertebrate Phototransduction and the Visual Cycle (Part A) Edited by KRZYSZTOF PALCZEWSKI VOLUME 316. Vertebrate Phototransduction and the Visual Cycle (Part B) Edited by KRZYSZTOF PALCZEWSKI VOLUME 317. RNA–Ligand Interactions (Part A: Structural Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 318. RNA–Ligand Interactions (Part B: Molecular Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 319. Singlet Oxygen, UV-A, and Ozone Edited by LESTER PACKER AND HELMUT SIES VOLUME 320. Cumulative Subject Index Volumes 290–319 VOLUME 321. Numerical Computer Methods (Part C) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 322. Apoptosis Edited by JOHN C. REED VOLUME 323. Energetics of Biological Macromolecules (Part C) Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 324. Branched-Chain Amino Acids (Part B) Edited by ROBERT A. HARRIS AND JOHN R. SOKATCH VOLUME 325. Regulators and Effectors of Small GTPases (Part D: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 326. Applications of Chimeric Genes and Hybrid Proteins (Part A: Gene Expression and Protein Purification) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 327. Applications of Chimeric Genes and Hybrid Proteins (Part B: Cell Biology and Physiology) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON

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Tubular Liposomes with Variable Permeability for Reconstitution of FtsZ Rings Masaki Osawa and Harold P. Erickson Contents 1. 2. 3. 4.

Introduction Reagents Bacterial Expression of Membrane Targeting FtsZ Purification of FtsZ-mts and FtsZ-YFP-mts 4.1. For FtsZ-mts 4.2. For FtsZ-YFP-mts 5. Renatured Preparation of FtsZ-YFP-mts 6. Tubular Multilamellar Liposome Preparation 7. Permeability of the Multilamellar Liposomes 8. Z-ring Formation in Liposomes 9. A Crude Flow Chamber to Exchange Buffer Outside Liposomes 10. Factors Affecting Z-ring Formation in Liposomes 11. Utility of the Liposomes Beyond FtsZ References

4 4 5 5 6 6 7 7 9 11 11 14 16 16

Abstract We have developed a system for producing tubular multilamellar liposomes that incorporate the protein FtsZ on the inside. We start with a mixture of spherical multilamellar liposomes with FtsZ initially on the outside. Shearing forces generated by applying a coverslip most likely distort some of the spherical liposomes into a tubular shape, and causes some to leak and incorporate FtsZ inside. We describe protocols for liposome preparation, and for preparing membrane-targeted FtsZ that can assemble contractile Z rings inside the tubular liposomes. We also describe the characterization of the multilamellar liposomes in terms of the permeability or leakiness for a small fluorescent dye and larger protein molecules. These liposomes may be useful for reconstitution of other biological systems. Department of Cell Biology, Duke University Medical Center, Durham, North Carolina, USA Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64001-5

#

2009 Elsevier Inc. All rights reserved.

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1. Introduction FtsZ is a bacterial tubulin homologue that forms a ring structure called the ‘‘Z ring’’ at the division plane in bacteria. The Z ring is anchored to the membrane and constricts to divide the bacteria. FtsZ recruits a dozen other essential division proteins, which are mostly involved in remodeling the peptidoglycan cell wall. We recently succeeded in reconstituting Z rings inside tubular liposomes, and found that they generated a constriction force on the liposome wall (Osawa et al., 2008). The assembly of Z rings and the generation of the constriction force were achieved with FtsZ alone, and did not require any other division protein. This was an important discovery itself for understanding the mechanism of bacterial cell division. Now the liposome system we developed should provide a simple in vitro system for studying molecular details of how FtsZ works. To achieve these results we had to overcome two technical problems. The first problem was to tether FtsZ to the membrane. Pichoff and Lutkenhaus (2005) discovered that the carboxy terminus of FtsZ binds to FtsA, and FtsA has an amphipathic helix at its carboxy terminal that inserts into the membrane. We made an FtsZ that could tether itself to the membrane by fusing an amphipathic helix (membrane targeting sequence: mts) to the carboxy terminus of FtsZ. To visualize the protein, we inserted a yellow fluorescent protein (YFP) before the mts, giving FtsZ-YFP-mts. Here we provide detailed protocols for the purification of FtsZ-YFP-mts. The second problem was how to get the protein inside liposomes. We have succeeded in getting FtsZ-YFP-mts inside spherical unilamellar liposomes using the emulsion method (Noireaux and Libchaber, 2004; Pautot et al., 2003). However, we have never found Z rings assembled in such spherical unilamellar liposomes. Eventually, we discovered a procedure that produced tubular multilamellar liposomes, and incorporated FtsZ-YFP-mts inside, where it formed Z rings. Initially this was a fortunate accident, since the cylindrical geometry was not designed, and the FtsZ was initially on the outside. We have since refined the procedures for producing tubular multilamellar liposomes, and we now understand some aspects of the permeability or leakiness that lets FtsZ inside. We describe here our protocols for producing the tubular liposomes and the tests of permeability.

2. Reagents The following reagents are used in our experiments: 

Column buffer: 50 mM Tris/HCl, pH 7.9, 50 mM KCl, 1 mM EDTA, 10% (v/v) glycerol

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HMKCG buffer: 50 mM HEPES/KOH, pH 7.7, 5 mM MgAc, 300 mM KAc, 50 mM KCl 10% (v/v) glycerol HMK50-350 buffer: 50 mM HEPES/KOH, pH 7.7, 5 mM MgAc, 50–350 mM KAc DOPG:1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (Avanti) Egg PC: phosphatidylcholine (Avanti) HccA: 7-Hydroxycoumarin-3-carboxylic acid (Invitrogen) Teflon disc, 37 mm diameter

3. Bacterial Expression of Membrane Targeting FtsZ The FtsZ-YFP-mts is expressed from a pET-11b expression vector, with FtsZ366-YFP-mts or FtsZ366-mts genes inserted at NdeI/BamHI sites (366 indicates that the FtsZ was truncated there, removing the FtsA-binding C-terminal peptide). The YFP we use is the variety Venus (Nagai et al., 2002), which gave superior results in FtsZ fusions in E. coli (Osawa and Erickson, 2005). The mts used here is the amphipathic helix from E. coli MinD (Szeto et al., 2003). We have not yet tested the amphipathic helix from FtsA, which has three to five additional extra amino acids that extend the amphipathic helix (Pichoff and Lutkenhaus, 2005). The expression vector is transformed into E. coli strain C41 (Miroux and Walker, 1996), which gives better yields of soluble proteins than BL21. After transforming, colonies are selected on an LB (Luria broth) agar plate containing 100 mg/ml ampicillin. A colony is picked and cultured overnight in 50 ml LB media with 100 mg/ml ampicillin at 37  C. Five milliliters of the overnight culture is diluted in 500 ml LB and cultured at 37  C until the optical density at 600 nm reaches 0.8–1.0. Protein expression is induced by addition of 0.5 mM IPTG and at the same time the temperature of the shaker is set to 20  C (our shaker takes 1–2 h to reach 20  C). The cells are cultured overnight and spun down at 3750 rpm for 45 min in a Beckman GPR rotor.

4. Purification of FtsZ-mts and FtsZ-YFP-mts Since FtsZ-mts and FtsZ-YFP-mts are expressed as soluble proteins, we purify them using the same protocol as for wild-type FtsZ. The packed cells are resuspended in a final volume of 20 ml column buffer, and 1 mM phenylmethanesulphonylfluoride (PMSF) and 0.1–0.2 mg/ml

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lysozyme are added. The mixture is then incubated on a rotator at 4  C for 15 min. They are frozen at 80  C overnight or longer. Two cycles of freeze–thaw (fresh 1 mM PMSF is added after each thawing) method are performed. The resultant mixture is sonicated on ice until the viscosity is reduced. We usually sonicate it for three cycles of 20 s, with 1 min cooling intervals. It is then centrifuged at 32,000 rpm for 20 min at 4  C (Beckman 42.1 Ti rotor). The supernatant is collected and ammonium sulfate is added to 30% saturation (3.52 g dry ammonium sulfate to the 20 ml volume). This mixture is incubated for 20 min on ice and again centrifuged at 32,000 rpm for 20 min at 4  C (Beckman 42.1 Ti rotor). The supernatant is discarded and the pellet is resuspended in 10 ml column buffer and passed through a 0.22 mm filter. The protein is purified on an anion exchange column. A 1  10 cm Source Q column (Source 15Q, GE Healthcare) is used. The column is eluted with a 100 ml gradient from 50–500 mM KCl in column buffer.

4.1. For FtsZ-mts FtsZ has very low UV absorbance, so the peak is located by running each fraction on SDS–PAGE. The peak fractions are pooled and dialyzed into HMK350. The protein concentration is determined by the BCA method (Pierce). FtsZ produces 75% as much color as BSA (Lu et al., 1998), so it is necessary to correct for this. Aliquots are frozen and stored at 80  C.

4.2. For FtsZ-YFP-mts After elution from the Source 15Q column, the peak fractions are pooled. There are typically two peaks: a large main peak and a following small peak, and both peaks have an indistinguishable activity. These peaks can be identified by yellow fluorescence and confirmed by SDS–PAGE. They are concentrated using an Amicon Ultra-15 with centrifugation at 5000g. We have noted that incomplete boiling of FtsZ-YFP-MTS with SDS sample buffer generates two bands on the gel. The upper band (68 Kd) results from completely denatured protein and the lower band (60 Kd), which still has yellow fluorescence in the gel, is due to FtsZ-YFP-mts where the YFP is not denatured. The concentration of FtsZ-YFP-mts can be determined from its absorption at 515 nm, using the extinction coefficient for YFP-Venus 92,200 M 1 cm 1.

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Our preferred buffer for FtsZ-YFP-mts is now HMKCG because FtsZ-YFP-mts seems to be more stable, as described below; we now use HMKCG for dialysis, dilution, reaction, and storage buffer.

5. Renatured Preparation of FtsZ-YFP-mts In our previous study (Osawa et al., 2008), we used a renaturing technique to prepare FtsZ366-YFP-mts. We developed this protocol because an early preparation of soluble protein as described above had no activity. We recently found that the soluble protein has full activity, equal to the best fractions of the renatured preparation. Because of the simplicity and much higher yield, we now use the soluble preparation for all work. One curious observation with the renatured FtsZ-YFP-mts was that it lost activity when dialyzed into HMK350. We found that addition of 10% glycerol and 50 mM chloride ion to HMK350 would preserve the activity of the renatured FtsZ-YFP-mts during dialysis. Although this precaution may only be necessary for the renatured FtsZ-YFP-mts, our preferred buffer is now HMKCG, which contains this glycerol and chloride ion.

6. Tubular Multilamellar Liposome Preparation PC and DOPG are dissolved separately in methanol at 100 mg/ml and mixed at a 4:1 ratio (20 ml/5 ml) in a 1.5 ml Eppendorf tube. The mixture is dried with an air current. 250 ml of milliQ water is added to the Eppendorf tube, and the dried lipid is suspended with vigorous vortexing. Many drops (5 ml each, total 250 ml) of the suspension are placed on a 37 mm diameter Teflon disc (Fig. 1.1A) and dried using an air current (Fig. 1.1B). The Teflon disc is placed in a beaker slightly larger than its diameter, and covered with 5 ml reaction buffer. It is then incubated at 37  C overnight to hydrate lipid (Fig. 1.1C). High salt buffers such as HMKCG or HMK350 will give aggregated multilamellar liposomes, which are desired to make tubular liposomes for Z-ring formation. The beaker is gently agitated to generate liposomes (Fig. 1.1D). Too vigorous agitation generates many small liposomes which interfere with observation of Z rings. About 1 ml of the suspension is pipetted out closest to the Teflon, which contains the most concentrated liposomes. This is placed in a 1.5 ml Eppendorf tube and left on a bench for 1 h (Fig. 1.1E). The multilamellar liposomes in HMKCG float to the surface. To obtain concentrated

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A

B

C

Liposome suspension (less than 5 ml)

Teflon disc D

Reaction buffer (HMKCG) E

Multilamellar liposome

F

HMKCG

Figure 1.1 Schematic illustration of the production of multilamellar liposomes. (A) A 37 mm Teflon disc is placed in a small beaker and 250 ml total aqueous suspension of lipid is deposited as many small drops (each drop is less than 5 ml). (B) The drops are dried with an air current and (C) the Teflon disc is covered with 5 ml of HMKCG and incubated overnight at 37  C. (D) The Teflon disc is gently agitated and multilamellar liposomes floated off. (E) One microliter of the most concentrated suspension, nearest the Teflon, is transferred to an Eppendorf tube and left on the bench for 1 h. The liposomes rise to the surface. (F) HMKCG is carefully removed from the bottom, leaving a concentrated suspension of liposomes.

liposomes, buffer from the bottom of the tube is carefully pipetted out, leaving the concentrated top layer (Fig. 1.1F). Liposomes that have a thinner wall (including unilamellar liposomes) can be easily obtained by using HMK100 with the same methods. If the KAc is reduced to 50 mM or less, many unilamellar liposomes will be obtained. Interestingly, liposomes prepared in HMK100 settle to the bottom of the Eppendorf tube over several hours. This is the opposite of multilamellar liposomes in HMKCG, which float to the top (probably because the 10% glycerol increases the density of the solution). To produce tubular liposomes, 5 ml of the aggregated multilamellar liposomes in HMKCG or HMK350 is placed on a glass slide and a coverslip is applied. To make liposomes with Z rings, liposomes are mixed first with 1 mM GTP and 4 mM FtsZ-YFP-mts. In this procedure, tubular liposomes are formed, probably generated by shear force when the suspension is spreading quickly through the narrow space between glasses. The tubular liposomes are always a minority, but with practice they can be found reproducibly. The tubular liposomes are

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more abundant near the edges of the coverslip, perhaps because that is where shear is maximized. We have noticed that the fluorescence background is higher in the center, where the drop is initially deposited. This may reflect FtsZ-YFP-mts binding to the glass. We have not tested whether the concentration of soluble FtsZ-YFP-mts is substantially reduced by binding to the glass.

7. Permeability of the Multilamellar Liposomes To check the permeability of the multilamellar tubular liposomes that were produced by the above procedures, a polar fluorescence dye HccA is used as a small molecule indicator (MW 206.15 kDa) and FtsZ fused with YFP-Venus (FtsZ-YFP) as a large molecule indicator (MW 67 kDa). Neither of these probes can cross the lipid bilayer, so they should report on smaller or larger pores in the liposomes. Also, the FtsZ-YFP is missing the mts, and was given no GTP, so it would not make protofilaments or Z rings. It should just report the existence of pores large enough to admit the globular protein. When we mix the multilamellar liposomes with 1 mM HccA and 32 mM FtsZ-YFP and apply a coverslip, we find tubular liposomes with both HccA and YFP fluorescence. More than 80% of tubular liposomes have FtsZ-YFP inside and even more liposomes have HccA inside, indicating that multilamellar liposomes are leaky at least for some time after addition of the fluorescent probes and application of the coverslip. A simple FRAP (fluorescent recovery after photobleach) assay is used to determine the leakiness of liposomes 30 min after the coverslip is applied. The entire field of view is exposed to UV from the mercury lamp for 20 s, without any filter. This results in complete bleaching of both fluorescent molecules inside and outside the liposomes. HccA fluorescence on the outside recovers over 20–60 s as unbleached probe diffuses into the area from outside. At 20 s HccA has entered about half of the liposomes (Fig. 1.2F), and by 5 min almost all the liposomes contain fluorescent HccA. Diffusion of FtsZ-YFP is slower, requiring 5 min to recover 70% of original fluorescence on the outside. After 45 min about half of the liposomes have YFP fluorescence inside. We conclude that the multilamellar liposomes are almost all permeable or leaky to the small molecule probe, and about half have pores that will pass the larger protein probe. A curious phenomenon is the appearance of a bright ring on the inner layer of the liposome when the inside of the liposome is dark and the outside is bright (Fig. 1.2D and F). When we repeat the FRAP experiment with a confocal microscope, the bright ring is seen only rarely and is also very dim. We suggest that the bright ring may be an optical artifact seen in the wide field fluorescence microscopy.

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A pre

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1 2 3

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Figure 1.2 FRAP assay for checking permeability of multilamellar liposomes. The top row shows FtsZ-YFP (lacking the mts and GTP), the middle row shows HccA fluorescence, and the bottom row shows DIC. The first column shows prebleach images, and the numbers on the other panels show time after bleach. In (D), vesicles 1–3 show little or no recovery of FtsZ-YFP inside, but they do show a thin bright layer of internal fluorescence. The vesicle between 1 and 2 shows substantial recovery of FtsZ-YFP. In (F), (G) and (H) vesicles 1–3 show no recovery of HccA at 20 s, but full recovery at 5 min. The exposure of fluorescence images is kept at the same level from prebleach to postbleach. Notice also the change in shape of several vesicles over 45 min. (B) 50% magnification showing that fluorescence is completely bleached.

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8. Z-ring Formation in Liposomes We have shown previously that FtsZ-YFP-mts can form Z rings inside tubular liposomes when FtsZ-YFP-mts plus GTP was mixed with multilamellar liposomes just before applying the coverslip (Osawa et al., 2008). To examine the leakiness of these liposomes with Z rings inside, we repeated the FRAP assay using 8 mM FtsZ-YFP-mts plus 2 mM GTP instead of 32 mM FtsZ-YFP. Most liposomes that had Z rings before the bleach showed no recovery of fluorescent Z rings, even though HccA fluorescence recovered rapidly (Fig. 1.3). Fluorescent FtsZ-YFP-mts could be seen coating the outside of the liposome but apparently did not enter the liposome, since no fluorescent Z rings were seen. We did, however, find some liposomes that recovered fluorescent Z rings after the bleach (Fig. 1.4). Figure 1.4C shows an intriguing example where HccA fluorescence invades through a hole at the top side of the tubular liposome (arrow in Fig. 1.4F). FtsZ-YFP-mts followed and fluorescent Z rings assembled as the nonbleached FtsZ-YFP-mts exchanged with the bleached (Fig. 1.4C and D), showing that occasional open liposomes can recover FtsZ from the outside and assemble Z rings. Recovery of Z rings with FtsZ-YFP-mts appeared to be less frequent than recovery of fluorescence of FtsZ-YFP. A potentially important difference is that FtsZ-YFPmts had GTP, so it should be mostly assembled into protofilaments 30 subunits long (Chen and Erickson, 2005). FtsZ-YFP was used without GTP so it should be much smaller protein monomers.

9. A Crude Flow Chamber to Exchange Buffer Outside Liposomes For various purposes one might want to change the buffer outside the liposomes, especially knowing that most of the liposomes are permeable to small molecules. The best production of tubular liposomes occurs with a small sample volume, which generates an optimal shear when the coverslip is applied. A 5 ml sample spread over a 4.8 cm2 coverslip gives a liquid layer 10 mm thick. If we try to perfuse buffer into this thin layer the liposomes are subject to high shear that results in severe elongation, loss of Z rings, and loss of liposomes. To minimize these effects, we first apply a 50 ml drop of perfusion buffer to one edge of the coverslip (Fig. 1.5B). We can then drain the liquid by applying a Kimwipe at the opposite edge, while applying a pipette tip or thin rod to prevent the coverslip from slipping. The arrangement shown in Fig. 1.5D, with a thin strip of Kimwipe applied to the coverslip and a large piece farther away, provides an optimal slow flow.

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B 1 min

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Figure 1.3 A liposome containing multiple Z rings (arrowheads) before bleach shows recovery of HccA fluorescence both outside and inside. FtsZ-YFP-mts fluorescence is recovered on the outside surface of the liposome, but no fluorescent Z rings recover inside. The exposure of fluorescence images is kept at the same level from prebleach to postbleach. (D) We did not detect any fluorescent Z rings even when we optimized the contrast in this specific panel.

We tested the system by adding FtsZ-YFP-mts without GTP to multilamellar liposomes. Cylindrical liposomes were found with the protein inside but Z rings could not form without GTP. We then perfused HMKCG containing 1 mM GTP. As the GTP entered the liposomes, Z rings assembled (Fig. 1.5E). An additional advantage of this system is that the fluorescent protein outside the liposome is washed away during the perfusion. This removes the fluorescent protein in solution and binding to the outside surface of liposomes and enhances the contrast of the Z rings inside. Because there is no FtsZ outside to hydrolyze the GTP, the Z rings can last a long time.

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Figure 1.4 A liposome with Z rings inside shows permeability to both FtsZ-YFP-mts and to HccA. (A) Arrowheads indicate Z rings prebleach. (B) YFP fluorescence is completely eliminated on the outside and inside of the liposome at 1 min after bleach. (C) About 12 min later YFP fluorescence has partially recovered on the outside, and fluorescent Z rings have formed on the inside, showing that unbleached FtsZ-YFP-mts has entered the leaky liposome. (D) At 45 min fluorescent Z rings have spread more toward the bottom. (E, F) About 20 s after bleaching, HccA has begun to enter the liposome, apparently through a hole at the top (note the zone of brighter fluorescence in the top 1/5 of the liposome). (G, H) At 12 min HccA fluorescence has fully recovered inside compared to the prebleached image in (E). Note that the FtsZ rings recover only at the top, near the hole, at 12 min. At 45 min they have spread through the top half of the liposome.

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A

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Figure 1.5 Changing buffer by flow/perfusion. (A) The normal preparation has a very thin layer of liquid between the slide and coverslip. (B) When a drop of the new buffer is placed on the side, the coverslip rises and is able to move back and forth. (C) To adsorb the liquid, the slide is tilted and a small piece of Kimwipe is placed on the edge opposite the drop. To prevent the coverslip from moving we used a thin rod or a pipette. (D) To minimize shear we slow the draining by placing a narrow piece of Kimwipe touching the coverslip and a larger piece farther away. (E) As a demonstration of buffer exchange, we prepared liposomes with FtsZ-YFP-mts but no GTP. FtsZ was trapped inside some tubular liposomes but did not form Z rings. We then flowed through a buffer containing GTP but no FtsZ. The GTP entered the liposome and initiated Z-ring assembly. This liposome was apparently leaky to GTP but not to protein, since the FtsZ did not leak out.

10. Factors Affecting Z-ring Formation in Liposomes During months of experimenting with the liposome system, we have made a number of observations that seem to be important for Z-ring formation. 

The shape of the liposome is very important, the cylindrical geometry being optimal for Z-ring assembly. We have rarely seen Z rings inside spherical liposomes. When these have internalized FtsZ-YFP-mts, the protein localizes to small patches or forms arcs that move unstably

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(Fig. 1.6B). One possible interpretation is that when a Z ring forms inside a spherical liposome, the contractile force causes it to collapse to a small patch. Another interpretation is that FtsZ filaments cannot determine the direction to form stable Z rings. We have sometimes observed this situation in which a large round liposome had radially formed FtsZ filaments inside (Fig. 1.6B, liposome on right). With a cylindrical geometry, a contractile force will cause it to form a circle in a plane perpendicular to the axis.  The diameter of the liposome is important. Z-ring assembly is optimal in tubular liposomes less than 2 mm in diameter (bacteria are typically 1 mm in diameter). In larger diameter tubular liposomes, some Z rings are

A

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Figure 1.6 Behavior of FtsZ-YFP-mts in large multilamellar liposomes. (A) In a large tubular liposome (3 mm inside diameter), FtsZ-YFP-mts formed some normal Z rings indicated by two spots of equal intensity directly across from each other (labeled *1). However, many spirals were also formed, indicated by dots spaced irregularly (*2). Note the bright ring at (*3), which appears to have peeled away the inner layer of the multilamellar wall (DIC image on right). Figure 1.6A is reprinted from Osawa et al. (2008) with permission of the publisher. (B) Several incomplete Z rings, spirals or arc structures, (arrowheads) are formed in a large spherical liposome.

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formed, but many structures appear to be helices or spirals rather than closed rings (Fig. 1.6A).  A mixture of neutral (PC) and negatively charged (DOPG) lipids is important. We have obtained Z rings with 2–40% DOPG, but typically use 1:4 DOPG:PC. 100% PC or DOPG, or a 2:3 ratio of DOPG:PC failed to support Z-ring assembly. This is probably because the amphipathic helix has positively charged amino acids on the hydrophilic side, and these need some negatively charged phospholipids in the bilayer to bind to. Interestingly, tubular liposomes produced by E. coli polar lipid (Avanti) did not support Z rings. This particular batch of E. coli lipid that we used may have contained more than 40% of charged lipid.  We have made several attempts to assemble Z rings in unilamellar liposomes, without success. As mentioned above, the spherical shape of unilamellar liposomes may not support Z-ring assembly. Another factor may be the rigidity of the wall. The rigidity of the multilamellar wall may slow constriction and stabilize the Z rings.

11. Utility of the Liposomes Beyond FtsZ The liposome system that we have developed may have uses beyond the study of FtsZ. The tubular liposomes are similar in size to a bacterium, and can range up to the size of a yeast cell. Similar to our reconstitution of Z rings, they may eventually be useful in reconstituting the cytokinetic apparatus of yeast or larger animal cells. More generally, we have demonstrated that most of the liposomes are permeable or leaky to small molecules, while about half are leaky to larger proteins. They therefore constitute small femtoliter chambers in which one can enclose proteins and manipulate the small molecule environment.

REFERENCES Chen, Y., and Erickson, H. P. (2005). Rapid in vitro assembly dynamics and subunit turnover of FtsZ demonstrated by fluorescence resonance energy transfer. J. Biol. Chem. 280, 22549–22554. Lu, C., Stricker, J., and Erickson, H. P. (1998). FtsZ from Escherichia coli, Azotobacter vinelandii, and Thermotoga maritima—Quantitation, GTP hydrolysis, and assembly. Cell Motil. Cytoskel. 40, 71–86. Miroux, B., and Walker, J. E. (1996). Over-production of proteins in Escherichia coli: Mutant hosts that allow synthesis of some membrane proteins and globular proteins at high levels. J. Mol. Biol. 260, 289–298. Nagai, T., Ibata, K., Park, E. S., Kubota, M., Mikoshiba, K., and Miyawaki, A. (2002). A variant of yellow fluorescent protein with fast and efficient maturation for cellbiological applications. Nat. Biotechnol. 20, 87–90.

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Noireaux, V., and Libchaber, A. (2004). A vesicle bioreactor as a step toward an artificial cell assembly. Proc. Natl. Acad. Sci. USA 101, 17669–17674. Osawa, M., and Erickson, H. P. (2005). Probing the domain structure of FtsZ by random truncation and insertion of GFP. Microbiology 151, 4033–4043. Osawa, M., Anderson, D. E., and Erickson, H. P. (2008). Reconstitution of contractile FtsZ rings in liposomes. Science 320, 792–794. Pautot, S., Frisken, B. J., and Weitz, D. A. (2003). Engineering asymmetric vesicles. Proc. Natl. Acad. Sci. USA 100, 10718–10721. Pichoff, S., and Lutkenhaus, J. (2005). Tethering the Z ring to the membrane through a conserved membrane targeting sequence in FtsA. Mol. Microbiol. 55, 1722–1734. Szeto, T. H., Rowland, S. L., Habrukowich, C. L., and King, G. F. (2003). The MinD membrane targeting sequence is a transplantable lipid-binding helix. J. Biol. Chem. 278, 40050–40056.

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C H A P T E R

T W O

Detection and Analysis of Protein Synthesis and RNA Replication in Giant Liposomes Takeshi Sunami,† Hiroshi Kita,† Kazufumi Hosoda,* Tomoaki Matsuura,* Hiroaki Suzuki,* and Tetsuya Yomo*,†,‡ Contents 1. Introduction 2. Methods 2.1. Liposome preparation 2.2. Internal protein synthesis reactions followed by the reaction catalyzed by the synthesized proteins 2.3. Detecting reactions in liposomes by fluorescence-activated cell sorting 3. Analysis of the FACS Data 4. Conclusions Acknowledgments References

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Abstract In living cells, biochemical reaction systems are enclosed in small lipidic compartments. To experimentally simulate various biochemical reactions occurring in extant cells, giant liposomes are used to reconstruct an artificial model cell. We present methods for conducting a protein synthesis reaction, followed by the reaction catalyzed by the synthesized proteins inside liposomes, and for measurement of the in liposome reaction using a fluorescence-activated cell sorter (FACS). These techniques enable us to perform detailed analysis of the biochemical reactions occurring in the microcompartments, and have the potential to reveal the role of compartmentalization in cellular systems.

* {

{

Department of Bioinformatics Engineering, Graduate School of Information Science and Technology, Osaka University, Suita, Osaka, Japan Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Agency ( JST), Chiyoda-ku, Tokyo, Japan Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka, Japan

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64002-7

#

2009 Elsevier Inc. All rights reserved.

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1. Introduction Liposomes are small vesicles composed of a lipid bilayer membrane, which are widely used in both basic and applied research in life sciences (Luisi and Walde, 2000; Torchilin and Weissig, 2003). Especially, giant liposomes (diameter > 1 mm) are of particular interest as a cell model and a microreactor containing biochemical reaction systems in a cell-sized volume (Hanczyc et al., 2003; Kaneko et al., 1998; Noireaux and Libchaber, 2004; Nomura et al., 2003; Tsumoto et al., 2001). Reconstruction of such a cell model, which we call here as an artificial cell, from defined biochemical components, with which the characteristics of the extant cells are expected to be elicited (Luisi et al., 2006; Szostak et al., 2001), is a challenging task for biochemists and chemical engineers. Researchers have been attempting to synthesize artificial cells aiming to identify various aspects of living systems. Attempts to synthesize artificial cells can contribute to a better understanding of the origin of life, as this understanding will provide a physically possible path that could have lead to primitive living cells. Furthermore, experimentally increasing the complexity of the artificial cell, starting from the simplest one, will provide an opportunity to simulate evolutionary processes to the development of more complex organisms, and eventually current organisms (Deamer, 2005; Forster and Church, 2006; Luisi et al., 2006; Szostak et al., 2001). While the experimental construction of artificial cells has been proposed many times over the last several years (Deamer, 2005; Forster and Church, 2006; Luisi et al., 2006; Szostak et al., 2001), progress toward this goal has been proceeding in discrete steps, with researchers assembling elements partially fulfilling the properties of a living system. For example, it was shown to be possible to generate artificial lipid vesicles (liposomes) of the same size as small bacteria from amphiphilic molecules (Bangham and Horne, 1964). Artificial vesicles were also shown to be capable of autocatalytic growth, and to even be able to undergo repeated cycles of growth and division (Hanczyc et al., 2003; Oberholzer et al., 1995; Takakura and Sugawara, 2004). Various types of biological reactions have been successfully performed within the environment provided by liposomes (Fischer et al., 2002; Murtas et al., 2007; Noireaux and Libchaber, 2004; Nomura et al., 2003; Tsumoto et al., 2001; Walde and Ichikawa, 2001; Yu et al., 2001). These studies represent significant steps toward the assembly of an artificial cell. What is common among most of these studies is the usage of giant liposomes (diameter > 1 mm) that are similar in size to extant living cells, which provides an environment that is biologically relevant. Researchers including us have been attempting to encapsulate various biochemical reactions, including the protein synthesis reaction, in giant

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liposomes (Fischer et al., 2002; Murtas et al., 2007; Noireaux and Libchaber, 2004; Nomura et al., 2003; Tsumoto et al., 2001; Walde and Ichikawa, 2001; Yu et al., 2001). For the protein synthesis reaction, the cell-free protein synthesis system consisting of large number of components are used as the material for encapsulation. When encapsulating this system in liposomes, it is necessary that the all components required for the reaction are being encapsulated. If the concentrations of the components are below nM, however, and the size of liposomes is below mm, it is possible that some of the components will be missing. It is often the case that components involved in the protein synthesis reaction are at sub-nM concentrations (Shimizu et al., 2001), and therefore, for the protein synthesis reaction to occur in liposomes, giant liposomes have to be used. Here, we describe the strategy for preparing giant liposomes used for encapsulating the b-glucuronidase synthesis reaction and the RNA replication by self-encoded replicase, both of which utilize the cell-free protein synthesis system. We also describe the methods to detect the reaction in individual liposomes using a fluorescence-activated cell sorter (FACS), and to analyze the data. FACS allows the evaluation of large quantities of individual liposomes, at a rate of more than 20,000 liposomes/s. By the quantitative evaluation of the internal reaction within liposomes, properties of the reactions conducted in a small compartment (Kita et al., 2008), and an interesting property of the internal structure of liposomes can be identified (Hosoda et al., 2008).

2. Methods 2.1. Liposome preparation For encapsulation of the cell-free protein synthesis system, the choice of vesicle formation method is critical, since the concentration and encapsulation efficiency of the number of individual components in such a dense suspension may vary over a wide range (Gregoriadis et al., 1999; Monnard et al., 1997; Pupo et al., 2005; Walde and Ichikawa, 2001). Among the various vesicle formation methods proposed to date, the freeze-dried empty liposomes (FDEL) method (Kikuchi et al., 1999; Kirby and Gregoriadis, 1984; Murtas et al., 2007; Torchilin and Weissig, 2003) is one of the most promising candidates. The FDEL method has long been used as a means of vesicle formation with high entrapment efficiency even for biomolecules (Kirby and Gregoriadis, 1984; Torchilin and Weissig, 2003). In this method, liposomes in suspension are lyophilized (freeze-dried) to form empty lamellae of dry lipid film. Upon rehydration, a suspension containing biochemical molecules permeates into the lamellae, which swell to form material-containing vesicles. This method provides facile, stable, and reproducible formation of lipid

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vesicles containing complex and dense reaction mixtures. It is also important to note that this method is applicable for almost any combination of buffer and components. For the preparation of FDEL membranes, 1-palmitoyl-2-oleoyl-snglycero-3-phosphocholine (POPC; Avanti Polar Lipids, Alabaster, AL), cholesterol (Nacalai Tesque, Kyoto, Japan), and 1,2-distearoyl-sn-glycero3-phosphoethanolamine-n-[methoxy(polyethylene glycol)-5000] (DSPEPEG5000; NOF Corporation, Tokyo, Japan) dissolved in dichloromethane/diethyl ether (1:1, v/v) are mixed at a molar ratio of 58:39:3 and dried using a rotary evaporator, followed by complete removal of solvent in the vacuum chamber. Note that the lipid composition can be altered as desired, depending on the objective of the experiments, while some lipid compositions may strongly inhibit the internal reactions. The dried lipid film is hydrated with Milli-Q water (12 mM lipid), and this suspension is subjected to vortex mixing for 20 s and sonication for 5 s. After passing through a polycarbonate filter with a pore size of 0.4 mm (Nuclepore TrackEtch Membranes; Whatman, Maidstone, Kent, UK), the suspension is dispensed into small aliquots (40 mL each), and freeze-dried overnight (Labconco Corp., Kansas, MO). After purging the tubes with Argon gas, the freeze-dried membranes are stored in a freezer. Figure 2.1A shows the scanning electron microscope image of the freeze-dried liposome membrane. This dried membrane has a highly porous structure in the micrometer scale. To encapsulate the reaction mixture in liposomes, the solution is simply injected over this dried membrane (Fig. 2.1B).

2.2. Internal protein synthesis reactions followed by the reaction catalyzed by the synthesized proteins In practice, cell-free protein synthesis systems have been used to produce functional proteins, such as green fluorescent protein (GFP) (Ishikawa et al., 2004; Sunami et al., 2006; Yu et al., 2001), T7 RNA polymerase (Ishikawa et al., 2004), Qb RNA replicase, b-galactosidase (Kita et al., 2008), and b-glucuronidase (Hosoda et al., 2008) in liposomes. Among these, we describe the synthesis of b-glucuronidase (Fig. 2.2A) (Hosoda et al., 2008) and Qb replicase (Fig. 2.2B) (Kita et al., 2008). Qb replicase has been used to construct a ‘‘self-encoding system,’’ which is described in detail below. 2.2.1. b-Glucuronidase synthesis A plasmid encoding b-glucuronidase (pET-uidA) (Hosoda et al., 2008) is mixed with the cell-free protein synthesis system (PURESYSTEM classic II (Post Genome, Tokyo, Japan); Shimizu et al., 2001), 50 mM 5-(pentafluorobenzoylamino) fluorescein di-b-D-glucuronide (PFB-FDGlcU; Invitrogen, Carlsbad, CA), and a red fluorescent protein (allophycocyanin, hereafter APC, Molecular Probes, USA) at a high concentration (500 nM final

A

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Figure 2.1 Microscopic pictures of liposomes prepared by the FDEL method. (A) Scanning electron microscope (SEM) image of freezedried liposomes. Freeze-dried liposomes are observed in a SEM (Keyence, VE-9800, Japan) at 950
magnification with 1 kV acceleration voltage after evaporation of gold (a few hundred nanometers). (B) Microscopic images of liposomes encapsulating R-PE. Upper images: bright-field observation; lower images: fluorescence observation.

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Figure 2.2 Schematic of the encapsulated reactions. (A) b-Glucuronidase synthesis. The mRNA is transcribed from the DNA template, and b-glucuronidase is translated from the mRNA using the cell-free protein synthesis system within the liposomes. Synthesized b-glucuronidase hydrolyzes the fluorogenic substrate (PFB-FDGlcU) to yield the green fluorophore PFB-fluorescein. Red fluorescent protein (APC) is added as a marker of the internal aqueous volume. (B) RNA replication by self-encoded replicase. Rep(þ)Gal() RNA encodes the Qb replicase b-subunit and an antisense sequence of the b-galactosidase gene (lacZ). Synthesized b-subunit assembles three host proteins of Escherichia coli: ribosomal protein S1, elongation factor Tu (EF-Tu), and Ts (EF-Ts) that are present in the cell-free system to generate active Qb replicase. Upon the replication of the sense strand RNA by self-encoded replicase, the sense sequence of lacZ appears, which then yield the b-galactosidase. Synthesized b-galactosidase then hydrolyzes the fluorogenic substrate (CMFDG) to yield the green fluorophore CM-fluorescein.

concentration). PFB-FDGlcU is a fluorogenic substrate of b-glucuronidase that turns into a fluorescent molecule (PFB-fluorescein) as a product of the enzyme reaction and emits green fluorescence. APC is used as a marker for the internal aqueous volume. The liposomes containing the reaction system are prepared by adding 10 mL of reaction mixture into an aliquot of freezedried liposomes (final lipid concentration 48 mM ). The liposome suspension is diluted 20-fold with the dilution buffer (reaction mixture without DNA, substrate, and APC). Protease (final concentration 1 mg/mL) is also included in the dilution buffer to suppress completely the enzyme reaction that may occur outside of the vesicles. All preparation procedures mentioned above are performed on ice. The reaction is then initiated by incubating at 37  C, and the suspension is time-sampled and subjected to FACS measurements. 2.2.2. RNA replication by self-encoded replicase In all living systems, the genome is replicated by proteins encoded within the genome itself. This universal reaction is essential for evolvability of the system. We previously constructed a simplified system termed a self-encoding system, where the genetic information is replicated by self-encoded replicase

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in liposomes (Kita et al., 2008). The self-encoding system is assembled using RNA, as shown in Fig. 2.2B. The Rep(þ)Gal() RNA encodes the sense strand of the b-subunit of Qb replicase, an RNA-dependent RNA polymerase responsible for replicating the RNA genome of coliphage Qb (Blumenthal and Carmichael, 1979), and an antisense sequence of the bgalactosidase gene (lacZ ). In addition, the RNA is designed to serve as a template for the replication by Qb replicase. When Rep(þ)Gal() RNA is added to the cell-free protein synthesis system, Qb replicase is first synthesized, which then replicates its own template to produce the antistrand RNA. The antistrand RNA also serves as a template of Qb replicase. As the RNA is replicated by its own encoding gene, we termed this as a self-encoding system. The occurrence of the replication reaction can be probed by the production of b-galactosidase, as it is encoded on the antistrand RNA. Synthesized b-galactosidase then hydrolyzes the fluorogenic substrate. In practice, Rep(þ)Gal() RNA is mixed with the cell-free protein synthesis system (PURESYSTEM, customized as described in Kita et al. (2008)), 100 mM fluorogenic substrate 5-chloromethylfluorescein di-b-Dgalactopyranoside (CMFDG; Invitrogen), and R-phycoerythrin (R-PE; Invitrogen) at high concentration (400 nM at a final concentration). CMFDG is a fluorogenic substrate of b-galactosidase that turns into a fluorescent molecule (CM-fluorescein) as a product of the enzyme reaction and emits green fluorescence. R-PE is used as a marker for internal aqueous volume. Note that both R-PE and APC can be used as an internal volume marker. Liposomes containing the reaction system are prepared and initiated essentially the same as the b-glucuronidase synthesis.

2.3. Detecting reactions in liposomes by fluorescence-activated cell sorting Fluorescence-activated cell sorting is used to measure various reactions in individual cells by detecting the emission intensities of multiple fluorescent markers upon laser irradiation. FACS can also be used to measure the internal reactions in liposomes (Kageyama et al., 2007; Taly et al., 2007). For example, green fluorescent products in liposomes can be quantified by measuring the green fluorescence intensity of individual liposomes, whereas the inner aqueous volume of each liposome can be quantified from the intensity of the red fluorescent marker protein encapsulated. As the number of green product molecules and the internal aqueous volume can be simultaneously measured, the concentration of the product molecule in individual liposomes can be estimated. The rate of measurement by FACS is more than 20,000 liposomes/s, significantly faster than that of microscopic observations. Typical results of FACS measurements are shown in Fig. 2.3. Note that FACS can be used not only to measure the internal reaction of individual liposomes but also to sort the liposomes with desired fluorescence

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Figure 2.3 Time course of the reaction analyzed by FACS. Typical results of in liposome (A) b-glucuronidase synthesis and (B) RNA replication by self-encoded replicase. The product (horizontal) and internal aqueous volume (vertical) of each liposome are shown. Dots represent the data of individual liposomes. In all plots, 20,000 data points from a total of 100,000 obtained are shown for clarity. The reactions proceed only in a fraction of the liposomes. This is because not all liposomes carry the plasmid DNA or the reaction efficiency is very low (Hosoda et al., 2008; Kita et al., 2008).

Biochemical Reactions in Giant Liposomes

27

signals, and the sorted samples can be used for further analysis such as microscopic observations (e.g., Hosoda et al., 2008). For FACS measurements, the sampled liposome suspension is diluted by at least 10-fold with sheath solution (FACSFlow; Becton Dickinson, San Jose, CA) to reduce the measurement frequency to smaller than 20,000 events/s. Then, red and green fluorescence signals, reflecting the whole internal aqueous volume and the amount of product, respectively, are measured with a FACS (FACSAria; Becton Dickinson). Typically, 100,000 data are sufficient for each measurement. The time course of the internal reaction can also be measured by analyzing the liposomes at different time points. APC or R-PE, used as a measure for the internal aqueous volume, is excited with a HeNe laser (633 nm) (or semiconductor laser (488 nm) in the case of R-PE), and the emission is detected through a 660  10 nm band-pass filter (or 576  13 nm band-pass filter in the case of R-PE). The number of R-PE molecules is converted from the red fluorescence signal detected (FIR) using the linear relation between the intensity from fluorescent beads carrying a known amount of R-PE (QuantiBRITE PE Quantitation Kit; BD Biosciences Clontech, Palo Alto, CA). The number of APC molecules is converted from the red fluorescence signal detected (FIR) using the linear relation between the intensity from fluorescent beads carrying a known amount of RPE, and the linear relation between fluorescence intensity of R-PE and APC. The quantity of the whole volume (Vw) is calculated by dividing the number of APC or R-PE molecules by the concentration of encapsulated APC (500 nM) or R-PE (400 nM). The final equation used for the conversion is Vw ¼ 0.054 FIR, or Vw ¼ 0.094 FIR, with APC or R-PE, respectively. The number of product molecules is derived from the intensity of the green fluorescence signal (FIG). PFB-fluorescein and CM-fluorescein, the reaction product of b-glucuronidase and b-galactosidase, respectively, are excited with a 488-nm semiconductor laser and the emission is detected through a 530  15 nm band-pass filter. For conversion, the linear relation between the green fluorescence intensities obtained by FACS and the number of product molecules (PFB-fluorescein or CM-fluorescein) is obtained. This is obtained by determining the intensity of a known amount of fluorescent molecules, and the linear relation between the intensity of the fluorescent molecule and the product molecules. The final equation used for conversion of PFB-fluorescein and CM-fluorescein is NG ¼ 330 FIG and NG ¼ 920 FIG, respectively, where NG is the number of product molecules.

3. Analysis of the FACS Data The kinetics of the internal reaction can be obtained from the FACS data. Figure 2.4A shows the histogram (frequency distribution) of the product concentration of RNA replication by self-encoded replicase in

28

Takeshi Sunami et al.

0.1 Frequency

B

0.12 0.08 0.06

0 min 120 min 200 min 275 min

0.04 0.02 0 0.1

1 10 100 CM-fluorescein (mM )

CM-fluorescein (mM )

A

20 15 10

Internal volume (fL) 1.2-4.0: 4.0-13: 13-40: 40-130:

5 0

0 100

200 300 400 Time (min)

500

Figure 2.4 Analysis of the FACS data. (A) Histogram (frequency distribution) of the CM-fluorescein concentration in liposomes with an internal volume of 4–13 fL at different time points. This is obtained from the data shown in Fig. 2.3B. (B) Time course of the reaction in liposomes with different internal volumes.

liposomes with different internal aqueous volumes and at different time points. In the measurement data, liposomes in which reaction proceeded (reacted liposomes) can be distinguished from those remained non-reacted based on the intensity of green fluorescence. The median of the histogram of the reacted liposomes are employed as a representative product concentration in liposomes with a given aqueous volume. The median values are used to trace the representative behavior of the internal reaction. As an example, we describe the strategy to obtain the time course data from the data shown in Fig. 2.4A. First, median values of the product concentration at the end of reaction, where the population of reacted liposomes can be readily distinguished, are obtained for each liposome internal volume (1.2–4.0, 4.0–13, 13–40, 40–130 fL). The rank in the product concentration (rank 1 ¼ highest product concentration) of the liposomes exhibiting the median value is then obtained. To obtain the representative product concentration at intermediate time points, where the distributions of reacted and non-reacted liposomes overlap, the product concentrations of the liposomes of identical rank to those obtained above are subsequently obtained for each liposome internal volume at different reaction times. These give the time courses of the reaction within liposomes shown in Fig. 2.4B.

4. Conclusions The presented method for in liposome protein synthesis followed by the measurement by FACS allows us to assess the kinetics of reactions in liposomes. This method can thus be used to elucidate the reaction dynamics in a microcompartment. In all living cells, cytosolic biochemical reactions are enclosed in a lipidic compartment. The effect of the microcompartment

Biochemical Reactions in Giant Liposomes

29

properties on the internal reaction has not been fully investigated. It may be possible to elucidate this effect with the presented method, for example, by carrying out the reaction in liposomes with different lipid composition and/ or different size. Thus, by providing an environment mimicing that of living cells, and by investigating the reactions inside, it may be possible to reveal the role of compartmentalization in cellular systems.

ACKNOWLEDGMENTS This research was supported in part by ‘‘Special Coordination Funds for Promoting Science and Technology: Yuragi Project’’ and ‘‘Global COE (Centers of Excellence) Program’’ of the Ministry of Education, Culture, Sports, Science, and Technology, Japan.

REFERENCES Bangham, A. D., and Horne, R. W. (1964). Negative staining of phospholipids and their structural modification by surface-active agents as observed in the electron microscope. J. Mol. Biol. 8, 660–668. Blumenthal, T., and Carmichael, G. G. (1979). RNA replication: Function and structure of Qbeta-replicase. Annu. Rev. Biochem. 48, 525–548. Deamer, D. (2005). A giant step towards artificial life? Trends Biotechnol. 23, 336–338. Fischer, A., Franco, A., and Oberholzer, T. (2002). Giant vesicles as microreactors for enzymatic mRNA synthesis. ChemBioChem 3, 409–417. Forster, A. C., and Church, G. M. (2006). Towards synthesis of a minimal cell. Mol. Syst. Biol. 2, 45. Gregoriadis, G., McCormack, B., Obrenovic, M., Saffie, R., Zadi, B., and Perrie, Y. (1999). Vaccine entrapment in liposomes. Methods 19, 156–162. Hanczyc, M. M., Fujikawa, S. M., and Szostak, J. W. (2003). Experimental models of primitive cellular compartments: Encapsulation, growth, and division. Science 302, 618–622. Hosoda, K., Sunami, T., Kazuta, Y., Matsuura, T., Suzuki, H., and Yomo, T. (2008). Quantitative study of the structure of multilamellar giant liposomes as a container of protein synthesis reaction. Langmuir 24, 13540–13548. Ishikawa, K., Sato, K., Shima, Y., Urabe, I., and Yomo, T. (2004). Expression of a cascading genetic network within liposomes. FEBS Lett. 576, 387–390. Kageyama, Y., Toyota, T., Murata, S., and Sugawara, T. (2007). Study on structural changes in supramolecular assemblies composed of amphiphilic nicotinamide and its dihydronicotinamide derivative by flow cytometry. Soft Matter 3, 699–702. Kaneko, T., Itoh, T. J., and Hotani, H. (1998). Morphological transformation of liposomes caused by assembly of encapsulated tubulin and determination of shape by microtubuleassociated proteins (MAPs). J. Mol. Biol. 284, 1671–1681. Kikuchi, H., Suzuki, N., Ebihara, K., Morita, H., Ishii, Y., Kikuchi, A., Sugaya, S., Serikawa, T., and Tanaka, K. (1999). Gene delivery using liposome technology. J. Control. Release 62, 269–277. Kirby, C., and Gregoriadis, G. (1984). Dehydration-rehydration vesicles—A simple method for high-yield drug entrapment in liposomes. Biotechnology 2, 979–984.

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Kita, H., Matsuura, T., Sunami, T., Hosoda, K., Ichihashi, N., Tsukada, K., Urabe, I., and Yomo, T. (2008). Replication of genetic information with self-encoded replicase in liposomes. ChemBioChem 9, 2403–2410. Luisi, P. L., and Walde, P. (2000). Giant Vesicles. John Wiley & Sons Inc., New York. Luisi, P. L., Ferri, F., and Stano, P. (2006). Approaches to semi-synthetic minimal cells: A review. Naturwissenschaften 93, 1–13. Monnard, P. A., Oberholzer, T., and Luisi, P. (1997). Entrapment of nucleic acids in liposomes. Biochim. Biophys. Acta 1329, 39–50. Murtas, G., Kuruma, Y., Bianchini, P., Diaspro, A., and Luisi, P. L. (2007). Protein synthesis in liposomes with a minimal set of enzymes. Biochem. Biophys. Res. Commun. 363, 12–17. Noireaux, V., and Libchaber, A. (2004). A vesicle bioreactor as a step toward an artificial cell assembly. Proc. Natl. Acad. Sci. USA 101, 17669–17674. Nomura, S. M., Tsumoto, K., Hamada, T., Akiyoshi, K., Nakatani, Y., and Yoshikawa, K. (2003). Gene expression within cell-sized lipid vesicles. ChemBioChem 4, 1172–1175. Oberholzer, T., Wick, R., Luisi, P. L., and Biebricher, C. K. (1995). Enzymatic RNA replication in self-reproducing vesicles: An approach to a minimal cell. Biochem. Biophys. Res. Commun. 207, 250–257. Pupo, E., Padron, A., Santana, E., Sotolongo, J., Quintana, D., Duenas, S., Duarte, C., de la Rosa, M. C., and Hardy, E. (2005). Preparation of plasmid DNA-containing liposomes using a high-pressure homogenization-extrusion technique. J. Control. Release 104, 379–396. Shimizu, Y., Inoue, A., Tomari, Y., Suzuki, T., Yokogawa, T., Nishikawa, K., and Ueda, T. (2001). Cell-free translation reconstituted with purified components. Nat. Biotechnol. 19, 751–755. Sunami, T., Sato, K., Matsuura, T., Tsukada, K., Urabe, I., and Yomo, T. (2006). Femtoliter compartment in liposomes for in vitro selection of proteins. Anal. Biochem. 357, 128–136. Szostak, J. W., Bartel, D. P., and Luisi, P. L. (2001). Synthesizing life. Nature 409, 387–390. Takakura, K., and Sugawara, T. (2004). Membrane dynamics of a myelin-like giant multilamellar vesicle applicable to a self-reproducing system. Langmuir 20, 3832–3834. Taly, V., Kelly, B. T., and Griffiths, A. D. (2007). Droplets as microreactors for highthroughput biology. ChemBioChem 8, 263–272. Torchilin, V., and Weissig, V. (2003). Liposomes: A Practical Approach. 2nd ed. Oxford University Press, Oxford. Tsumoto, K., Nomura, S. M., Nakatani, Y., and Yoshikawa, K. (2001). Giant liposome as a biochemical reactor: Transcription of DNA and transportation by laser tweezers. Langmuir 17, 7225–7228. Walde, P., and Ichikawa, S. (2001). Enzymes inside lipid vesicles: Preparation, reactivity and applications. Biomol. Eng. 18, 143–177. Yu, W., Sato, K., Wakabayashi, M., Nakaishi, T., Ko-Mitamura, E. P., Shima, Y., Urabe, I., and Yomo, T. (2001). Synthesis of functional protein in liposome. J. Biosci. Bioeng. 92, 590–593.

C H A P T E R

T H R E E

Construction of Cell-Sized Liposomes Encapsulating Actin and Actin-Cross-linking Proteins Kingo Takiguchi,* Ayako Yamada,†,1 Makiko Negishi,† Makoto Honda,*,2 Yohko Tanaka-Takiguchi,* and Kenichi Yoshikawa† Contents 1. Introduction 1.1. Actin-cross-linking proteins coencapsulated in cell-sized liposomes 1.2. Methodologies to encapsulate actin and actin-cross-linking proteins into giant liposomes 2. Experimental Section 2.1. Preparation of actin and actin-cross-linking proteins for encapsulation 2.2. Natural swelling method for coencapsulation of actin and fascin, a-actinin, filamin, or BBMI into liposomes 2.3. Spontaneous transfer method for coencapsulation of actin and HMM (ActoHMM) into liposomes 3. Morphogenesis of Giant Liposomes Encapsulating Actin and Its Cross-linking Proteins 3.1. G-Actin and its polymerization in liposomes (natural swelling method) 3.2. Actin and fascin (natural swelling method) 3.3. Actin and a-actinin (natural swelling method) 3.4. Actin and filamin (natural swelling method) 3.5. Actin and BBMI (natural swelling method) 3.6. F-Actin (spontaneous transfer method)

* { 1 2

32 33 34 37 37 37 38 42 42 43 43 43 45 45

Division of Biological Science, Graduate School of Science, Nagoya University, Furo-cho, Chikusa-ku, Nagoya, Japan Department of Physics, Graduate School of Science, Kyoto University, Sakyo-ku, Kyoto, Japan Current address: Department of Chemistry, Ecole Normale Superieure, Paris, France Current address: Stem Cell and Drug Discovery Institute, Shimogyo-ku, Kyoto, Japan

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64003-9

#

2009 Elsevier Inc. All rights reserved.

31

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Kingo Takiguchi et al.

3.7. ActoHMM (spontaneous transfer method) 3.8. Actin and S-1 (spontaneous transfer method) 3.9. Mechanism determining liposome morphology 4. Concluding Remarks Acknowledgments References

45 46 46 49 50 50

Abstract To shed light on the mechanism underlying the active morphogenesis of living cells in relation to the organization of internal cytoskeletal networks, the development of new methodologies to construct artificial cell models is crucial. Here, we describe the successful construction of cell-sized liposomes entrapping cytoskeletal proteins. We discuss experimental protocols to prepare giant liposomes encapsulating desired amounts of actin and cross-linking proteins including molecular motor proteins, such as fascin, a-actinin, filamin, myosin-I isolated from brush border (BBMI), and heavy meromyosin (HMM). Subfragment 1 (S-1) is also studied in comparison to HMM, where S-1 and HMM are singleheaded and double-headed derivatives of conventional myosin (myosin-II), respectively. In the absence of cross-linking proteins, actin filaments (F-actin) are distributed homogeneously without any order within the liposomes. In contrast, when actin is encapsulated together with an actin-cross-linking protein, mesh structures emerge that are similar to those in living motile cells. Optical microscopic observations on the active morphological changes of the liposomes are reported.

1. Introduction Living cells and their organelles are compartmentalized by biomembranes in a self-organized manner, and each has a specific shape depending on its function. Cellular morphologies are thought to be determined and maintained by cytoskeletal networks (Hotani et al., 2003; Rodriguez et al., 2003). F-actin is a major component of the cytoskeleton, and is involved in a variety of cellular functions. Such functions include the extension of microspikes from cells (Albrecht-Buehler and Lancaster, 1976), the movement of filopodia in neural growth cones (Mitchison and Cramer, 1996; Mitchison and Kirschner, 1988), the extension or retraction of pseudopods during amoeboid movement (Taylor and Condeelis, 1979), and the contraction of contractile rings during cell division (Schroeder, 1973). F-actins also provide mechanical support, including stress fibers (Byers and Fujikawa, 1982). It is therefore important to study the role of the actin cytoskeleton in relation to membrane morphogenesis. Along this line, artificial model systems using giant liposomes containing actin and its associating proteins have been developed (Ba¨rmann et al., 1992; Cortese et al., 1989; Fygenson

33

Construction of Actin-Encapsulating Liposomes

et al., 1997; Ha¨ckl et al., 1998; Limozin et al., 2003, 2005; Limozin and Sackmann, 2002; Miyata and Hotani, 1992; Miyata et al., 1999; Pontani et al., 2009). In this chapter, we use the term ‘‘cell-sized liposome’’ to indicate giant liposomes with sizes of several to several tens of micrometers, which constitute closed thin membranes of phospholipids in aqueous solution. Cell-sized liposomes are expected to serve as a useful model in the real world, allowing us to make real-time observations using optical microscopy (Bangham, 1995; Cortese et al., 1989; Hotani et al., 2003; Lasic, 1995; Lipowsky, 1991; Miyata et al., 1999).

1.1. Actin-cross-linking proteins coencapsulated in cell-sized liposomes F-actins usually function in vivo in bundles or networks, and they are organized by various actin-binding, especially actin-cross-linking, proteins. Accordingly, we studied the effects of five different actin-cross-linking proteins, fascin, a-actinin, filamin, BBMI, and HMM on the morphogenesis of liposomes caused by actin assembly, by coencapsulating actin and one of those proteins together in liposomes. Actin-cross-linking proteins are characterized by the manner of cross-linking, that is, by the distance and angular flexibility between adjacent cross-linked F-actin molecules, as illustrated in Fig. 3.1. Fascin is a 55 kDa globular protein and is responsible for the tight crosslinking of F-actin to form bundles (Cant et al., 1994; Edwards and Bryan, 1995; Edwards et al., 1995; Otto et al., 1980; Yamashiro-Matsumura and Matsumura, 1985). a-Actinin is a larger protein (about 100 kDa), and Proteins

Fascin

a-actinin

Filamin

BBMI

HMM

S-1

Domain structure

Crosslinking pattern

Figure 3.1 Model of F-actin bundles or networks formed by actin-cross-linking proteins. From left to right, cross-linked F-actins mediated with fascin, a-actinin, filamin, BBMI, or HMM are shown. The upper row shows the molecular structure of each cross-linking protein. The lower row illustrates the arrangement of F-actins crosslinked by each cross-linking protein. S-1 and S-1-associating F-actins are also shown (right). F-actin is depicted as a line, while fascin, a-actinin, filamin, BBMI, HMM, and S-1 are drawn as globular monomers, dumbbell-shaped dimers, V-shaped dimers, single-headed musical note-like structures, double-headed structures, and single-headed structures, respectively.

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antiparallel side-by-side associated homodimers of a-actinin form F-actin bundles with much looser packing (Imamura et al., 1988; Meyer and Aebi, 1990; Mimura and Asano, 1987). Filamin is a protein of approximately 250 kDa with a flexible elongated structure, and end-to-end associated homodimers of filamin are able to mediate flexible and high-angle linking between two adjacent F-actins (Bretscher, 1991; Hartwig and Kwiatkowski, 1991; Hock et al., 1990). The dynamics and mechanical properties of actin networks caused by these actin-cross-linking proteins have been well studied (Collins et al., 1991; Grazi et al., 1994; Hou et al., 1990; Wachsstock et al., 1993, 1994; Xu et al., 1998; Yamashiro et al., 1998). However, the relationship between the actin network organization and the morphogenesis of biological membranes has not been clarified yet. In living cells, the cytoskeleton is thought to be anchored to the membrane through membrane proteins, which probably regulate their own specific membranous morphology, position, and duration. BBMI exhibits a unique property in comparison with the other actin-cross-linking proteins mentioned above (fascin, a-actinin, filamin, or HMM). BBMI consists of a single motor domain (head) and a tail domain, and since the tail domain can bind both F-actin and a lipid membrane, BBMI can play a role as a cross-linker between F-actin and the membrane, as well as between F-actins (Coluccio, 1997). HMM has been frequently studied as a representative of double-headed myosin motors, and it can form and transform bundles and gels of F-actin (Takiguchi, 1991; Tanaka-Takiguchi et al., 2004). On the other hand, S-1 has often been studied as a representative of simple single-headed myosin motors. S-1 has only one actin-binding motor domain and is unable to cross-link F-actins, thus S-1 often has been studied in comparison to HMM.

1.2. Methodologies to encapsulate actin and actin-cross-linking proteins into giant liposomes Through the methodology of natural swelling, which is a method to obtain liposomes by the hydration of dried lipid films, we have successfully prepared liposomes containing monomeric actin (G-actin) and an actincross-linking protein (such as fascin, a-actinin, filamin, or BBMI) in the presence of 30 mM KCl (Honda et al., 1999). As the encapsulated G-actin polymerizes into F-actins, they form bundles or gels depending on the type of coencapsulated actin-cross-linking protein, causing various morphological changes of liposomes. The differences in morphology among the transformed liposomes indicate that the actin-cross-linking proteins determine the liposome shape by organizing their specific actin networks. On the other hand, in living cells, and depending on the cell type and the intracellular location, actin is expressed up to about 300 mM and undertakes its functions with the cooperation of various myosin motor proteins under physiological salt conditions (several mM Mg2þ and several tens of

Construction of Actin-Encapsulating Liposomes

35

mM Kþ) ( Janson et al., 1991; Pollard et al., 2000). However, using the natural swelling method, it is difficult or almost impossible to entrap the desired amounts of F-actin and myosin simultaneously in giant liposomes at physiological salt conditions. This is mainly due to the difficulty in preparing giant liposomes in the presence of salt, especially divalent cations such as Mg2þ, and the difficulty in controlling the amount of macromolecules to be incorporated within the liposomes, because of the passive nature of the encapsulation process. To overcome those problems, we have developed the spontaneous transfer method through an oil/water interface, by adapting water-in-oil phospholipid-coated cell-sized-droplets (W/O droplets) as precursors of giant liposomes (Yamada et al., 2006). Recently, there have been several attempts to employ W/O droplets coated by phospholipids as a model for living cells instead of liposomes (Hase and Yoshikawa, 2006), or as a precursor of liposomes (Hamada et al., 2008; Noireaux and Libchaber, 2004; Pautot et al., 2003a,b; Pontani et al., 2009; Takiguchi et al., 2008; Yamada et al., 2006). The W/O droplets are prepared easily by emulsifying an aqueous solution, together with oil, containing phospholipids. The process enables us to encapsulate biomolecules at a controlled concentration under any salt strength, into cell-sized compartments covered with a monolayer of phospholipids. By taking advantage of this, the behavior of F-actins entrapped in a closed space (Claessens et al., 2006) or conformational changes of actin molecules interacting with a phospholipid membrane in the presence of Mg2þ (Hase and Yoshikawa, 2006) have been studied. In addition, we have succeeded in developing a system for the controlled fusion of two droplets containing a substrate and an enzyme, respectively (Hase et al., 2007). Consequently, methodologies to obtain liposomes by transferring phospholipid-coated W/O droplets from an oil phase to an aqueous phase through their interface using centrifugation (Noireaux and Libchaber, 2004; Pautot et al., 2003a,b; Pontani et al., 2009) or without an external force (the spontaneous transfer method; Hamada et al., 2008; Takiguchi et al., 2008; Yamada et al., 2006) have been developed. With those methodologies, one can obtain liposomes with sizes of 10–100 mm containing desired amounts of molecules. Furthermore, by taking advantage of microfluidic techniques, new methodologies to obtain monodisperse liposomes from monodisperse droplets have been actively studied (Funakoshi et al., 2007; Shum et al., 2008; Stachowiak et al., 2008). Table 3.1 compares the various methodologies used to prepare giant liposomes. Using one of the simplest methods, that is, the spontaneous transfer method, we succeeded in constructing giant liposomes encapsulating 200 mM F-actin in the presence of 5 mM MgCl2 and 50 mM KCl. Moreover, this method enabled us to succeed in encapsulating desired amounts of HMM or S-1 with F-actin into giant liposomes (Takiguchi et al., 2008).

Table 3.1 Methods for giant liposome preparation Natural swellinga

Preparation Required time for encapsulation Yield par observation field Required amount of the sample Control of inner/ outer condition Control of the size Asymmetric membrane Physiological salt concentration Problem of oil contamination a b c d e f g h

Easy Long Low

Electroformationb

Centrifugationc

Spontaneous transferd

Jettinge

Double emulsion f

Easy Comparably long High

Easy Short

Easy Short

Small

Difficult Comparably short Comparably low Large

Difficult Comparably short Comparably low Large

High

Comparably large Difficult

Large

Comparably low Small

Difficult

Easy

Easy

Easy

Easy

Difficult

Difficult

Difficult

Easy

Easy

Difficult

Difficult

Easyg

In principle easy Easyh

Difficult

Difficult

Comparably difficult No

Difficult

Easy

Easy

Easy

Easy

No

Yes

Yes

Yes

Yes

Robust method to prepare liposomes of almost any content, but yield is sometimes low. Multilamellar liposomes tend to be obtained (Bangham et al., 1965). High yield of unilamellar liposomes. A special technique is required for salt concentrations higher than 10 mM (Angelova and Dimitrov, 1986). The principle is similar to that of the spontaneous transfer method. Liposomes are free in an aqueous phase (Noireaux and Libchaber, 2004; Pautot et al., 2003b). Easy encapsulation. Liposomes are anchored to an oil/water interface (Yamada et al., 2006). Microfluidic technique is required. Highly monodisperse liposomes can be obtained (Funakoshi et al., 2007; Stachowiak et al., 2008). Microfluidic technique and use of volatile oil are required. Highly monodisperse liposomes can be obtained (Shum et al., 2008). Pautot et al. (2003a). Hamada et al. (2008).

Construction of Actin-Encapsulating Liposomes

37

2. Experimental Section 2.1. Preparation of actin and actin-cross-linking proteins for encapsulation G-Actin is obtained by incubating dried muscle (acetone powder) prepared from rabbit skeletal muscle with cold water followed by a polymerization– depolymerization cycle (Ebashi and Ebashi, 1965; Higashi-Fujime, 1983). Fascin is isolated from porcine brain extracts using the actin gel method, following DE-52 and hydroxylapatite (Bio-Rad) column chromatographies (Honda et al., 1999; Yamashiro-Matsumura and Matsumura, 1985). a-Actinin is purified from an extract of minced rabbit skeletal muscle that had been incubated overnight at room temperature using a repeating fractionation with ammonium sulfate (Ebashi and Ebashi, 1965). Filamin is isolated from chicken gizzard by a modification (Muguruma et al., 1990) of the method of Molony et al. (1987). BBMI is purified from an extract of homogenized brush borders isolated from chicken intestines using a series of column chromatographies, Sepharose CL-4B, CM sepharose, and monoQ HR5/5 (Collins et al., 1991; Jontes and Milligan, 1997). Skeletal muscle myosin (myosin-II) was obtained from rabbit skeletal muscles (white muscles of the back and hind legs). HMM and S-1 are obtained by digestion of skeletal muscle myosin with chymotrypsin in the presence of 1 mM MgCl2/ 0.5 M KCl and of 1 mM EDTA (ethylenediamine tetraacetic acid)/120 mM KCl, respectively (Tanaka-Takiguchi et al., 2004).

2.2. Natural swelling method for coencapsulation of actin and fascin, a-actinin, filamin, or BBMI into liposomes To obtain giant liposomes encapsulating actin and its cross-linking proteins, dried phospholipid films are hydrated with an aqueous solution containing the proteins (Honda et al., 1999; Hotani and Miyamoto, 1990; Kaneko et al., 1998; Miyata and Hotani, 1992). The phospholipids used are L-aphosphatidylcholine (PC), L-a-phosphatidylethanolamine (PE) and L-aphosphatidylglycerol (PG). Each phospholipid has been isolated from egg yolk and is purchased from Sigma (St. Louis, MO) or Avanti Polar Lipid (Alabaster, AL), and is dissolved in a chloroform/methanol solution (98:2, v/v) at a concentration of 10 mM. Methanol is added to protect lipid molecules from damage resulting from the moisturization of chloroform. The phospholipid solutions are mixed in a glass test tube (total of 200 mg lipid). Typically, 20 ml of the solution is poured into a glass tube (10 mm in diameter and 30 mm in height) which has been cleaned with the chloroform/methanol solution prior to use. The organic solvent of the phospholipid solution is evaporated under a flow of nitrogen gas, and the lipids are

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Kingo Takiguchi et al.

further dried under vacuum for at least 90 min with a vacuum pump (G-20D, ULVAC, Chigasaki, Japan). Note that the phospholipid solutions are stocked at 4  C and are protected from light, and the dried lipid films are stocked in an evacuated dessiccator at room temperature and are also protected from light. To generate liposomes from the film, 40 ml buffer A (5 mM Tris–HCl, pH 8.0, 0.2 mM ATP, 5 mM dithiothreitol, 30 mM KCl) containing various concentrations of G-actin and an actin-cross-linking protein are added to the dried lipid films on ice. The lipid films immediately started swelling to form liposomes containing the proteins, and further swelling is facilitated by occasionally agitating the test tubes by hand. After 30 min, the liposome suspensions are diluted 30-fold with buffer A to prevent the polymerization of the G-actin outside the liposomes. The polymerization of G-actin to form F-actin inside the liposomes is then initiated by raising the temperature to 25  C under dark-field microscopy (BHF, Olympus, Tokyo, Japan) (Hotani, 1984). 2.2.1. Visualization of actin filament (F-actin) To visualize F-actin localization within liposomes, rhodamine-phalloidin (R-415, Molecular Probes, Eugene, OR) in methanol is added to the lipid solution prior to the film preparation. The final concentration of rhodamine-phalloidin in the specimens is about 20 nM. The rhodamine-phalloidin specifically binds F-actin, so that actin filaments become visible by fluorescence microscopy (Miyata and Hotani, 1992). 2.2.2. Visualization of liposome membranes Giant liposomes can be visualized without any labeling by several types of optical microscopes, such as phase contrast, differential interference contrast (DIC), and dark-field. To observe liposome membranes by fluorescence microscopy, a fluorescence-labeled phospholipid is mixed with other phospholipids (about 1:200, w/w) (Honda et al., 1999). NBD- or BODIPYconjugated phospholipids are frequently used as the fluorescence-labeled phospholipid. The fluorescence-labeled liposomes are then observed with fluorescence optics usually assembled into another microscope, that is, a phase contrast, DIC, or dark-field microscope.

2.3. Spontaneous transfer method for coencapsulation of actin and HMM (ActoHMM) into liposomes A schematic representation of the experimental setup is illustrated in Fig. 3.2 (Takiguchi et al., 2008; Yamada et al., 2006).

39

Construction of Actin-Encapsulating Liposomes

Egg PC in mineral oil

A

5 mm

PDMS Buffer

Cover glass

4 mm

Objective lens B

ActoHMM

Oil

Z X

Buffer

Figure 3.2 (A) Experimental setup for construction and observation of actoHMMencapsulating giant liposomes using the spontaneous transfer method. (B) Schematic representation of the transformation from a W/O droplet in the oil phase (left) to a liposome in the aqueous phase (right). F-actin is illustrated with a line, and HMM is illustrated as a double-headed structure. The interface of the droplet is depicted as a lipid monolayer, while a multilayered interface may be generated to some extent in these experiments. The figure is adopted from Takiguchi et al. (2008).

2.3.1. Observation chamber An observation chamber is prepared consisting of a cylindrical hole (ca. 4 mm in diameter) in a poly(dimethylsiloxane) (PDMS) sheet (ca. 5 mm thick) on a glass microscope slide (0.12–0.17 mm thick, Matsunami Glass, Osaka, Japan). To obtain the PDMS sheet, the base solution and a curing agent of Silpot 184 W/C (Dow Corning Toray, Tokyo, Japan) are mixed well at a ratio of 10:1 (w/w) and are poured into a plastic Petri dish to harden. Note that bubbles generated during the mixing disappear spontaneously in about 1 h at room temperature. One can speed the hardening process by baking but the temperature must not be higher than 85  C to avoid melting the Petri dish. It takes about 24 h at room temperature or 2 h at 85  C

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to harden the PDMS. A piece of the PDMS sheet is cut off and a hole with ca. 4 mm diameter is made in it with a leather-craft punch. One can make several holes in each PDMS sheet to compare different conditions. Note that if the diameter is larger than 4 mm, it is difficult to maintain a proper oil/water interface in the chamber because the water phase tends to be a spherical shape due to surface tension, since the PDMS surface is rather hydrophobic. These PDMS sheets are repeatedly used after washing with ethanol or isopropanol. The PDMS surface is cleaned with scotch tape before being set on a glass cover slide, which is cleaned by baking at 500  C for 1 h. The PDMS and the glass surface must be tightly attached. 2.3.2. Oil containing phospholipids Chloroform or a chloroform/methanol solution of PC is prepared at a concentration of 10 mM and poured into a glass test tube. Typically, 25–50 ml of the solution is poured into a glass tube (7 mm in diameter and 30 mm in height) (Maruemu, Osaka, Japan), which has been cleaned with acetone prior to use. The organic solvent of the phospholipid solution is then evaporated under a nitrogen flow and is dried under vacuum for more than 15 min to produce a dry film at the bottom of the glass tube. Typically, 500 ml of mineral oil (Nacalai Tesque, Kyoto, Japan) is then added to the tube and is sealed tightly to avoid moisture prior to ultrasonication for 60 min at 50  C and vortex mixing. For sealing, Parafilm (Pechiney Plastic Packaging, Chicago, IL) alone cannot be used because it melts under these conditions. A combination of a cap from the tube and Parafilm, or Dura Seal (Diversified Biotech, Boston, MA) and Parafilm can be used. Note that the vortex mixing must be performed immediately after ultrasonication to avoid aggregation of the phospholipid. The final concentration of PC is 0.5 or 1.0 mM, and these prepared oils must be used in 1 day. 2.3.3. Oil/water interface in the chamber Firstly, 10 ml of an aqueous phase, that is, the outer solution of liposomes, is placed at the bottom of the observation chamber. Here, we use a solution consisting of 25 mM imidazol-HCl, 5 mM MgCl2, 50 mM KCl, and 10 mM DTT at pH 7.5 (buffer B) and up to 100 mM sucrose, or the concentration of buffer B is increased up to twice instead of adding sucrose to adjust the osmotic pressure to that of the inner solution. It is recommended to touch the glass surface with the pipette tip so that one can fill the chamber from the bottom. The glass surface should be totally wet without air bubbles and the air/water interface must be rather flat and horizontal. Ten microliters of the PC-containing oil is then gently added along the PDMS wall to obtain an oil/water interface covered with

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phospholipid between the upper oil phase and the lower aqueous phase. The observation chamber is then set on the optical microscope. An objective lens with a magnification not higher than 20 times can be employed to focus on the formed interface. 2.3.4. Preparation of W/O droplets and transfer into liposomes One hundred microliters of the PC-containing oil is placed in an Eppendorf tube and 5 ml of the inner solution, that is, the target solution to be encapsulated in liposomes, is then added to the tube. As the inner solution, buffer B containing F-actin or actoHMM is prepared. Immediately after addition of the aqueous solution, the solution is emulsified by pipetting it up and down for about 30 s. Ten microliters of the obtained W/O droplet solution is then placed on the oil phase in the chamber. One can observe that, in the oil phase, the W/O droplets gradually fall down onto the oil/water interface because of gravity. Interestingly, the droplets then spontaneously move through the interface into the aqueous solution keeping their spherical shape. In our experimental conditions, the transferred droplets, or liposomes, are anchored onto the interface (Fig. 3.2). Although it is possible to transfer the liposomes further into the bulk aqueous phase by encapsulating higher density solution than the outer phase (Hamada et al., 2008), or using centrifugation (Pontani et al., 2009), we performed the observations on liposomes anchored to the interface since we could then monitor the full process of the transfer and the subsequent transformation of each specific liposome. As for the transformation of a droplet in oil into a liposome in water, we have already discussed the full details of the process (Yamada et al., 2006). Observations are performed using a Zeiss Axiovert 100 inverted microscope equipped with an LSM 510 module for confocal microscopy (Carl Zeiss, Jena, Germany). To observe liposome membranes by fluorescence microscopy, fluorescence-labeled phospholipids are mixed with other phospholipids (1:100–1:1000, mol/mol). 2.3.5. Visualization of F-actin Actin prepared as described above is polymerized in buffer C (2 mM Tris– HCl, pH 8.0, 30 mM KCl, and 0.2 mM ATP) and then used for the experiments. To visualize F-actin entrapped within liposomes, a mixture of rhodamine-phalloidin in methanol and buffer C, or alternatively rhodamine-phalloidin redissolved in buffer C after volatilizing the methanol, is added to the actin solution before emulsification in the lipid-containing oil or mixing with HMM. The molecular ratio of rhodamine-phalloidin to the actin monomer was approximately 1:40, which depends on the amount and thickness of the actin bundles formed.

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3. Morphogenesis of Giant Liposomes Encapsulating Actin and Its Cross-linking Proteins 3.1. G-Actin and its polymerization in liposomes (natural swelling method) Liposomes made of neutral and acidic phospholipids assumed spherical or tubular shapes in buffer A, and their shapes always fluctuated in solution (Fig. 3.3A) until the encapsulated actin polymerized. Liposomes containing only actin-cross-linking protein or G-actin also assumed spherical or tubular shapes in buffer A and always fluctuated in shape. The line-like images of liposomes obtained by dark-field microscopy have a uniform brightness which is almost identical to that of single-layered membranes from A

B

C

Figure 3.3 Morphological changes of actin-containing liposomes caused by actin assembly without an actin-cross-linking protein. (A) Dark-field microscopic images of liposomes made of neutral (PC) and acidic (PG) phospholipids (1:1, mol/mol). (B) Transformed liposomes obtained by the polymerization of encapsulated G-actin into F-actin. Four disk-shaped and one flat spoon-shaped liposomes are shown (viewed from the top). The lipid composition was PE and PG (1:1, mol/mol), and the concentration of encapsulated actin was 100 mM. (C) A fluorescent micrograph that shows the F-actin localization within a disk-shaped liposome obtained by actin assembly (top view). The assembled F-actin was labeled with rhodamine-phalloidin. The lipid composition was PC and PG (1:1, mol/mol), and the concentration of the encapsulated G-actin was 50 mM. All scale bars represent 5 mm. Note that here we can distinguish a disk-shaped liposome from a spherical one by shifting the focus plane of the microscope, or by continuous observation of each free-tumbling liposome in a microscope specimen. The figure is adopted from Honda et al. (1999).

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erythrocyte ghosts whose associated proteins have been removed by protease treatment. Polymerization of the encapsulated G-actin into F-actin was achieved by raising the temperature, and the subsequent morphological changes of liposomes were monitored. When the actin polymerized, the liposomes transformed into flat disk or spoon shapes (Fig. 3.3B) (Miyata and Hotani, 1992). The F-actin that had polymerized in liposomes spontaneously aligned along the periphery of the flattened liposomes to form bundles (Fig. 3.3C). These liposomes are fairly rigid, as judged by the low fluctuation in their shapes. Their membranes were quiescent, and bending motions were restrained.

3.2. Actin and fascin (natural swelling method) When G-actin was encapsulated into liposomes together with fascin and then polymerized, the F-actin bundled tightly. Nearly 60% of liposomes transformed into lemon shapes and straight rigid projections subsequently developed from the tip(s), and eventually they became elongated straight tube(s) (Fig. 3.4A). Fluorescence labeled F-actin was always detected in the long projecting membrane regions (Fig. 3.4A, left), indicating that the fascin-mediated actin bundles are responsible for the formation of the membrane projections. The frequency of transformed liposomes and the lengths of their projections increased at higher concentrations of encapsulated actin or fascin.

3.3. Actin and a-actinin (natural swelling method) Unlike fascin, a-actinin had only a weak effect on morphological changes of actin-containing liposomes. When actin polymerized, most liposomes coencapsulating a-actinin transformed into disk shapes similar to those generated by encapsulating and polymerizing G-actin in the absence of a linking protein. The polymerized actin bundled and then aligned along the periphery. Liposomes transformed into disk shapes possessing long projections only in rare cases (<5%), and some of those projections were not straight (Fig. 3.4B). F-actin bundles always existed in those projected regions (Fig. 3.4B, left), indicating that the a-actinin-mediated actin bundles are also responsible for the formation of membrane projections and occasionally acquired enough strength and rigidity to push out the lipid bilayer membrane.

3.4. Actin and filamin (natural swelling method) When actin and filamin were encapsulated together into liposomes, they became very rigid and swollen as the encapsulated actin assembled. More than 50% of liposomes transformed into ellipsoid- or nut-like shapes

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A

B

C

D

Figure 3.4 Morphological changes of liposomes caused by actin assembly with actincross-linking proteins (left: fluorescence, right: dark-field). (A) Transformed liposomes with fascin. 50 mM G-actin and 3 mM fascin were encapsulated and the lipid composition was PC and PG (1:1, mol/mol). Since the fascin-mediated actin bundles were too bright in dark-field microscope, a lower actin concentration was used in the case of fascin. There was, however, no difference in the liposome transformation between 50 and 100 mM actin. (B) Transformed liposomes with a-actinin. 100 mM G-actin and 2 mM aactinin were encapsulated and the lipid composition was PE and PG (1:1, mol/mol). (C) Transformed liposomes with filamin. 100 mM G-actin and 1 mM filamin were encapsulated and the lipid composition was PE and PG (1:1, mol/mol). (D) 10 (upper) or 25 mM (bottom) G-actin and 5 mM BBMI were encapsulated and the lipid composition was PC and PG (1:1, mol/mol). Since actin and BBMI were too difficult to coencapsulate into liposomes by the natural swelling method because they immediately formed large rigor complexes outside the liposomes, lower concentrations of actin were examined. The shape of each transformed liposome was stable and did not change. Fluorescence images show the localization of rhodamine-labeled F-actin bundles. In each case, before the polymerization of actin, the liposomes took spherical or tubular shapes and always fluctuated in shape as did the empty ones. In addition, each transformed shape started out as either spherical or tubular. All scale bars represent 5 mm. The figure is adopted from Honda et al. (1999).

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(Fig. 3.4C) and their inner spaces were filled with three-dimensional networks of F-actin of uniform distribution (Fig. 3.4C, left). Although the F-actin gels mediated by filamin had high viscosity, they could not be observed by dark-field microscopy. Unlike liposomes containing fascin or a-actinin, liposomes with filamin never formed long membrane projections (Fig. 3.4C).

3.5. Actin and BBMI (natural swelling method) BBMI can cross-link between F-actins, as well as between F-actin and a liposomal membrane. We found that because of this property, when BBMI was coencapsulated into liposomes, it effectively transformed them into protruded shapes at a lower concentration of actin compared to the other actin-cross-linking proteins (Fig. 3.4D). In the presence of BBMI, the actin bundles formed are promptly recruited and bound to the inner surface of the liposome membrane resulting in efficient liposome transformation. Consistently, the actin bundles were observed at the membranes of the transformed liposomes (Fig. 3.4D, left).

3.6. F-Actin (spontaneous transfer method) Although we prepared liposomes encapsulating only F-actin (at 200 mM), which had already polymerized before the encapsulation, using the spontaneous transfer method, they showed no transformation (Fig. 3.5). On the other hand, Miyata et al. (1999) showed that liposomes encapsulating the same concentration of G-actin prepared by the natural swelling method grew protrusive structures as the actin polymerized. These results are consistent with the pioneering reports: the assembly of G-actins at the end of the membrane-pushing F-actin, that is, the mechanical force generated during actin polymerization and elongation, is required for the liposomal transformation (Hotani et al., 2003; Maemichi et al., 2008; Miyata et al., 1999).

3.7. ActoHMM (spontaneous transfer method) Figure 3.6 shows liposomes that have entrapped F-actin together with HMM. In contrast to the images with F-actin only (Fig. 3.6A), in the presence of HMM, F-actin assemblies, such as bundles and networks, are generated (Fig. 3.6B–D). We also confirmed the appearance of similar assemblies of F-actin in aqueous solution in control experiments. As exemplified in Fig. 3.6C and D, with increased HMM concentration, deformed liposomes tended to appear reproducibly, although a large fraction of liposomes remained spherical.

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Figure 3.5 Confocal microscopic images of giant PC liposomes encapsulating F-actin obtained by the spontaneous transfer method (upper and bottom left: fluorescence, bottom right: transmission). The concentration of F-actin inside the liposomes is 200 mM. The fluorescence images show the distribution of rhodamine-phalloidinlabeled F-actin. The bottom panels show enlarged images of an individual F-actinencapsulating liposome. The small spherical objects situated on the surfaces of liposomes in the transmission images are attributed to oil droplets in the water phase (Yamada et al., 2006). Scale bars represent 50 mm (upper) or 10 mm (bottom). The figure is adopted from Takiguchi et al. (2008).

3.8. Actin and S-1 (spontaneous transfer method) When S-1 rather than HMM was coencapsulated with F-actin into giant liposomes, the liposomes showed a uniform distribution of F-actins and maintained their spherical shapes, even under conditions where excess S-1 was coencapsulated (Fig. 3.6E).

3.9. Mechanism determining liposome morphology The transformed morphology of liposomes closely correlated with the structure of actin bundles or networks (Fig. 3.1). For instance, when fascin was coencapsulated, actin bundles and liposome projections were as straight

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A

50 mM F-actin alone

B 50 mM F-actin + 3.8 mM HMM

C 50 mM F-actin + 5.0 mM HMM

D 50 mM F-actin + 7.5 mM HMM

E 50 mM F-actin + 60 mM S-1

Figure 3.6 Confocal microscopic images of actoHMM-encapsulating giant liposomes obtained by the spontaneous transfer method (left: fluorescence, right: transmission). The concentrations of F-actin inside the liposomes are 50 mM (A–E). The fluorescence images show the distribution of F-actin. (A) Images of liposomes encapsulating F-actin alone are shown as a control. (B–D) The concentrations of HMM coencapsulated are 3.8, 5.0, and 7.5 mM, respectively (indicated at the top of each panel). In C and D, liposomes possessing nonspherical irregular shapes are shown. (E) An actoS-1encapsulating giant liposome obtained by the spontaneous transfer method. The concentration of coencapsulated S-1 is 60 mM. Transmission images show the existence of small oil droplets around the liposomes, which were squeezed out from the oil phase. Bars represent 10 mm. The figure is adopted from Takiguchi et al. (2008).

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and rigid as needles, and showed no bending (Fig. 3.4A). This might arise from the higher rigidity of actin bundles mediated by fascin. On the other hand, bundles and projections were flexible and usually curved when a-actinin was coencapsulated. F-actin bundles always existed along the periphery of the central discoid portion as well as in the projecting regions (Fig. 3.4B). These results indicate that, in most cases, the elasticity of actin bundles mediated by a-actinin was not sufficiently strong to push out the lipid bilayer. Liposomes encapsulating filamin never formed projections (Fig. 3.4C), which probably results from the fact that filamin makes a three-dimensional gel rather than bundles of F-actin. Liposomes encapsulating BBMI transformed into protruded shapes even when the concentration of encapsulated actin was comparably lower, probably because the actin bundles formed with BBMI were bound to the liposome membrane efficiently by BBMI (Fig. 3.4D). These results indicate that the mechanical force generated by actin polymerization itself is strong enough to overcome the tension of liposomal membranes and can push them out if the actin filaments are tightly cross-linked since individual ones are very flexible. Fascin has a lower affinity and bundling activity to F-actin compared with a-actinin, but liposomes possessing projection(s) were formed much more frequently and their projections had higher rigidity when actin was assembled with fascin rather than with a-actinin (Fig. 3.4A and B). This might result from the fact that fascin forced the F-actin bundles to cross-link more tightly and rigidly than did a-actinin (Fig. 3.1). Moreover, liposomes encapsulating filamin never formed projections (Fig. 3.4C), even though filamin had the highest affinity to F-actin among these three proteins and had a similar stoichiometry to F-actin with a-actinin. This probably results from the fact that filamin makes a three-dimensional gel rather than bundles of F-actin (Fig. 3.1). In liposome-free conditions, it has been shown that the affinity of an actin-cross-linking protein for actin determines the mechanical properties of actin filament gels (Wachsstock et al., 1993, 1994; Xu et al., 1998). However, while the transformation frequency depends on the concentrations of actin and cross-linking proteins, the morphology does not. The results obtained from our study indicate that the manner (Fig. 3.1) rather than the quantity (such as the binding affinity or stoichiometry) of cross-linking is essential for the morphogenesis of actin-containing liposomes (Honda et al., 1999). Myosins can be classified into two groups according to their headstructure, that is, double- or single-headed myosins. HMM, which belongs to the double-headed type and has two actin-binding motor domains, is able to cross-link F-actins into actin bundles or actin gels, and moreover can transform the bundles or gels (Tanaka-Takiguchi et al., 2004). On the other hand, S-1, which belongs to the single-headed type and has only one actin-binding motor domain, is unable to cross-link F-actins. Liposomes coencapsulating HMM but not S-1 show morphological changes. From these results, one can conclude that simply bundling or redistributing

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F-actin, that is, the inter-filament physical interactions, can be a motive force for organizing the actin networks and the liposomal morphogenesis, as well as the polymerization of G-actin and the subsequent bundling of F-actin (Fig. 3.6C and D).

4. Concluding Remarks In this chapter, we describe the experimental procedures (natural swelling and spontaneous transfer method) used to form cell-sized liposomes containing actin and actin-cross-linking proteins. Furthermore, by studying the morphogenesis of the liposomes thus obtained, we successfully demonstrated that the mechanical force generated by the assembly of actin network structures itself is strong enough to transform cell-sized giant liposomes, and that the manner, rather than the quantity, of cross-linking is essential for the morphogenesis of liposomes encapsulating actin and actin-cross-linking proteins. Especially, by the spontaneous transfer method, we successfully constructed giant liposomes encapsulating both F-actin and HMM at high concentrations in the presence of 5 mM MgCl2 and 50 mM KCl. As a result, it could be shown that the encapsulated actoHMM formed self-organized actin network-like structures, and nonspherical liposomes were obtained in a reproducible manner. Divalent cations are indispensable for life, for example, the fuel of most of the machinery of living cells is Mg-ATP, and Ca2þ is a key factor in a number of biological regulatory systems. By the natural swelling method, however, the efficiency of construction of giant liposomes significantly decreased in the presence of divalent cations. In addition, the natural swelling method cannot control the concentrations of encapsulated species. On the other hand, the spontaneous transfer method enables us to efficiently make giant liposomes encapsulating the desired amounts of F-actin up to 200 mM, even in the presence of 5 mM MgCl2. Note that 200 mM is comparable to the actin concentration expressed in living cells ( Janson et al., 1991; Pollard et al., 2000), and is the upper limit of F-actin concentration that allows handling, including pipetting, due to its very high viscosity. By evaluation of the fluorescence intensity, the encapsulated concentration of F-actin inside the liposomes is almost constant and F-actin exists in a homogeneous manner. In previous studies using the natural swelling method, the highest concentration of actin that could be encapsulated in liposomes was 200 mM, when gel-filtrated G-actin was encapsulated and then polymerized into F-actin with KCl introduced by electroporation after the formation of liposomes (Miyata et al., 1999). Although such studies are interesting, it has been impossible to efficiently encapsulate desired amounts of F-actin inside liposomes using the natural swelling method.

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At this time, hundreds of actin-associating proteins and ten classes of different types of myosin motors have been identified. Usual analytical studies, however, might be unable to answer how they share roles and cooperate with each other in cells because living cells are typically a very complex system. This reconstitutive and very efficient system utilizing cell-sized giant liposomes containing actoHMM prepared by the spontaneous transfer method might represent the first critical step for developing a motile artificial cell model, which will generate spontaneous motion in a system similar to but much simpler than living cells. The important problems that remain are how to supply sufficient amounts of ATP to actoHMM existing inside the liposomes, and how to control the reactions between F-actin, HMM, and ATP. Thus, the next step may be to perform experiments by merging different liposomes, where F-actin, HMM, or ATP are encapsulated separately. Recently, a-hemolysin, a membrane-pore forming toxin from bacteria, has found application in molecular transport into vesicles (Gibrat et al., 2008; Noireaux and Libchaber, 2004; Pontani et al., 2009), and that novel methodology may prove useful to supply ATP to actoHMM-containing giant liposomes.

ACKNOWLEDGMENTS The studies described were supported by the Japan Society for the Promotion of Science ( JSPS) under a Grant-in-Aid for Creative Scientific Research (Project No. 18GS0421) and by the MEXT of Japan under a Grant-in-Aid (Scientific Research of Priority Areas, System Cell Engineering by Multi-Scale Manipulation, Project No. 20034024 and No.17076007).

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Otto, J. J., Kane, R. E., and Bryan, J. (1980). Redistribution of actin and fascin in sea urchin eggs after fertilization. Cell Motil. 1, 31–40. Pautot, S., Frisken, B. J., and Weitz, D. A. (2003a). Engineering asymmetric vesicles. Proc. Natl. Acad. Sci. USA 100, 10718–10721. Pautot, S., Frisken, B. J., and Weitz, D. A. (2003b). Production of unilamellar vesicles using an inverted emulsion. Langmuir 19, 2870–2879. Pollard, T. D., Blanchoin, L., and Mullins, R. D. (2000). Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu. Rev. Biophys. Biomol. Struct. 29, 545–576. Pontani, L.-L., van der Gucht, J., Salbreux, G., Heuvingh, J., Joanny, J.-F., and Sykes, C. (2009). Reconstitution of an actin cortex inside a liposome. Biophys. J. 96, 192–198. Rodriguez, O. C., Schaefer, A. W., Mandato, C. A., Forscher, P., Bement, W. M., and Waterman-Storer, C. M. (2003). Conserved microtubule-actin interactions in cell movement and morphogenesis. Nat. Cell Biol. 5, 599–609. Schroeder, T. E. (1973). Actin in dividing cells: Contractile ring filaments bind heavy meromyosin. Proc. Natl. Acad. Sci. USA 70, 1688–1692. Shum, H. C., Lee, D., Yoon, I., Kodger, T., and Weitz, D. A. (2008). Double emulsion templated monodisperse phospholipid vesicles. Langmuir 24, 7651–7653. Stachowiak, J. C., Richmond, D. L., Li, T. H., Liu, A. P., Parekh, S. H., and Fletcher, D. A. (2008). Unilamellar vesicle formation and encapsulation by microfluidic jetting. Proc. Natl. Acad. Sci. USA 105, 4697–4702. Takiguchi, K. (1991). Heavy meromyosin induces sliding movements between antiparallel actin filaments. J. Biochem. 109, 520–527. Takiguchi, K., Yamada, A., Negishi, M., Tanaka-Takiguchi, Y., and Yoshikawa, K. (2008). Entrapping desired amounts of actin filaments and molecular motor proteins in giant liposomes. Langmuir 24, 11323–11326. Tanaka-Takiguchi, Y., Kakei, T., Tanimura, A., Takagi, A., Honda, M., Hotani, H., and Takiguchi, K. (2004). The elongation and contraction of actin bundles are induced by double-headed myosins in a motor concentration-dependent manner. J. Mol. Biol. 341, 467–476. Taylor, D. L., and Condeelis, J. S. (1979). Cytoplasmic structure and contractility in amoeboid cells. International Review of Cytology Vol. 56, pp. 57–144. Academic Press, New York. Wachsstock, D. H., Schwartz, W. H., and Pollard, T. D. (1993). Affinity of a-actinin for actin determines the structure and mechanical properties of actin filament gels. Biophys. J. 65, 205–214. Wachsstock, D. H., Schwartz, W. H., and Pollard, T. D. (1994). Cross-linker dynamics determine the mechanical properties of actin gels. Biophys. J. 66, 801–809. Xu, J., Wirtz, D., and Pollard, T. D. (1998). Dynamic cross-linking by a-actinin determines the mechanical properties of actin filament networks. J. Biol. Chem. 273, 9570–9576. Yamada, A., Yamanaka, Y., Hamada, T., Hase, M., Yoshikawa, K., and Baigl, D. (2006). Spontaneous transfer of phospholipid-coated oil-in-oil and water-in-oil micro-droplets through an oil/water interface. Langmuir 22, 9824–9828. Yamashiro, S., Yamakita, Y., Ono, S., and Matsumura, F. (1998). Fascin, an actin-bundling protein, induces membrane protrusions and increases cell motility of epithelial cells. Mol. Biol. Cell 9, 993–1006. Yamashiro-Matsumura, S., and Matsumura, F. (1985). Purification and characterization of an F-actin-bundling 55-kilodalton protein from HeLa cells. J. Biol. Chem. 260, 5087–5097.

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C H A P T E R

F O U R

Reconstitution of Membrane Budding with Unilamellar Vesicles Anna V. Shnyrova and Joshua Zimmerberg Contents 1. Introduction 2. M Protein Purification 3. Evaluation of the Membrane Activity of M Protein Through its Interaction with Intermediate-Sized Unilamellar Liposomes 3.1. Remarks about IUV formation and characterization 3.2. IUV preparation 3.3. Analysis of M protein binding to IUVs of different lipid compositions 3.4. M protein interaction with IUVs: Leakage of aqueous dyes 4. Reconstitution of M-Protein-Driven Membrane Budding on GUVs 4.1. GUV preparation 4.2. Interaction of M protein with GUVs 5. Concluding Remarks References

56 57 58 58 59 61 65 66 66 69 73 73

Abstract Enveloped virus particles select their lipid–protein components and egress by budding from the host cell membranes. The matrix protein of many enveloped viruses has been proposed as a crucial element for viral budding; however, molecular mechanisms behind membrane remodeling by the matrix protein are yet to be unraveled. Here, we describe a set of in vitro functional reconstitution assays that allow quantitative evaluation of both, membrane binding and creation of membrane curvature by the matrix protein isolated from Newcastle Disease Virus. Individual budding events orchestrated by the matrix protein can be resolved in real time. The assays may be applied for direct reconstitution of the on-membrane action of cellular proteins involved in membrane curvature induction upon binding in vivo. Laboratory of Cellular and Molecular Biology, Program in Physical Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, Maryland, USA Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64004-0

55

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1. Introduction Cellular life and death depends critically on the membrane curvature creation by cellular proteins. Since the works by Palade (1953, 1975), the mechanisms of cellular trafficking and membrane deformation have been the target of extensive studies and many protein candidates for curvature induction have been proposed. However, it is not easy to extract the exact molecular mechanism behind the cellular curvature creation from in vivo observations, as cellular membranes are very crowded environments. The bottom-up approach, where isolated membrane systems are used to reconstitute cellular events with a minimum set of proteins, simplifies greatly this task. Recent reconstitution studies have proved this approach be a handy tool for a better understanding of curvature creation in cells. One illustrative example of these kinds of studies is the reconstitution of budding of enveloped viruses on giant unilamellar vesicles (GUVs) (Shnyrova et al., 2007; Solon et al., 2005). Despite the generally accepted notion that the host cell participates in viral budding, final virus particles include only small traces of cellular proteins (Briggs et al., 2003). This suggests that enveloped viruses ultimately rely on their own protein machinery for vesicle egress ion. Budding of most retroviruses (except Foamy Virus) and many paramixoviruses have been proposed to rely critically on the viral matrix protein, although other viral proteins are often enlisted as a requirement for a minimal budding machinery (reviewed by Welsch et al., 2007). The matrix (M) protein is the major component of most enveloped viruses, where it forms a tightly packed shell beneath the lipid envelope of the virus, thus resembling the caveolin coat of the endocytic vesicles, though with the opposite topology (Welsch et al., 2007). For many paramixoviruses, including Newcastle Disease Virus (NDV), expression of M protein in the absence of other viral proteins in the cell leads to formation of virus-like particles (VLPs) ( Jayakar et al., 2004; Pantua et al., 2006; Takimoto and Portner, 2004). These results indirectly suggested that NDV M protein possesses the energy and functionality necessary to create membrane shape of desired geometry. The mechanism of negative membrane curvature induction, however, still remained unresolved and the question remained open whether M alone, with just a lipid bilayer, is the minimum requirement for this process of membrane remodeling. Here, we describe an in vitro assay that allows to evaluate, to monitor in real time, and to quantify the ability of a membrane bound protein (the NDV M protein) to induce negative curvature upon its interaction with the membrane of giant liposomes. This assay offers unique insights into the molecular mechanisms driving this type of membrane remodeling in vivo.

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2. M Protein Purification An in vitro reconstitution of membrane activity of a protein requires highly purified protein in its native conformation, as misfolded proteins or sample contaminants can bind to the membrane and alter its behavior. These requirements frequently preclude and limit the use of recombinant membrane-active proteins from bacteria, as they often fold incorrectly in the absence of other components of their native environment and tend to aggregate in physiological conditions. Instead, we purify the M protein directly from its native environment: the NDV. This method yields high amounts of sufficiently purified protein. NDV LaSota strain (Charles Rivers Laboratories Inc., MA) is grown in the allantoic cavity of 11-day-old pathogen-free chick embryos (CBT Farm, MD). Upon infection, embryos are allowed to grow for 48 h at 37 ºC and 55% humidity. The allantoic fluid is then collected and stored at
4 ºC. Virus is purified the next day as described previously (Garcia-Sastre et al., 1989). Scheid and Choppin’s (1973) purification protocol is used as the reference for the M protein purification from NDV. To separate the lipid-anchored proteins from the viral nucleocapsid, the virus pellet is resuspended in 10 mL of 1 M KCl, 10 mM Tris–HCl and 5 mM CaCl2 at pH 7.4 (buffer A) with 2% the of detergent Triton X-100 and then incubated 30 min at room temperature with constant agitation. After incubation, the sample is centrifuged at 200,000g for 2 h at 4 ºC. The supernatant, containing the main components of the viral envelope is recovered and dialyzed against 2 L of buffer A, lacking KCl (five changes for 36 h). The low ionic strength of the dialysis buffer induces M protein precipitation and separation from the rest of the viral membrane proteins. The contents of the dialysis bag are centrifuged at 9000g for 30 min at 4 ºC. The pellet containing M protein is then dissolved in 1 mL of buffer B, which contains 1 M KCl, 20 mM HEPES, and 0.2 EDTA, pH 7.4. Final protein concentration is determined using the BCA protein assay kit (Pierce, IL) and is adjusted to lower than 75 mM in order to avoid protein precipitation during storage. Various viral M proteins are known for their ability to self-assemble into ordered structures at low salt conditions. The speed and efficiency of such aggregations are usually determined by the ionic strength and protein concentration (McCreedy et al., 1990). This sensitivity has been routinely used to separate matrix proteins from the rest of the viral proteins: M protein appears mostly as a monomer at high ionic strength, but aggregates slowly and irreversibly at physiological ionic strength (Sagrera et al., 1998). Accordingly, the use of high ionic strength (>1 M) solution during

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purification is critical for the disassembly of the interior of NDV from its envelope. However, experimental observations indicate that M protein interaction with NP protein is not totally disrupted during the purification procedure based only on salt concentration changes, and that the aggregation of M protein may be in part due to the formation of this M–NP complexes. Thus, addition of calcium in millimolar concentration during M protein purification allows for a better separation of M protein from the NP complex, presumably through disruption of some electrostatic interactions, therefore leading to highly purified M protein. The protein purity is estimated from the reducing SDS–PAGE by analysis of the areas under the intensity peaks correspondent to protein bands, as was described previously (Coorssen et al., 2002).

3. Evaluation of the Membrane Activity of M Protein Through its Interaction with Intermediate-Sized Unilamellar Liposomes GUVs provide an excellent tool for studying the membrane activity of a particular protein, as they allows for real time observation of shape transformations by bright-field or fluorescence microscopy. However, experiments on GUVs may end up frustrating if the binding of the protein to a particular lipid composition is not previously analyzed by quantitative methods. In this aspect, large (200–500 nm in diameter) and intermediatesized (100–200 nm in diameter) unilamellar liposomes (LUVs and IUVs, respectively) are suitable for the initial checkup of the protein–membrane interactions.

3.1. Remarks about IUV formation and characterization There are various methods for IUV preparation described elsewhere (a great reference source is New, 1990b). The choice of a particular method mainly depends on the desired lipid composition of the vesicles and the buffer used. For the compositions and conditions described below, a combination of gentle hydration method followed by extrusion through appropriate polycarbonate membrane gives reproducible results. However, we encourage the reader to explore a variety of methods for IUV preparation in order to find an appropriate one or a combination of several that suits the particular experimental case. The success in the IUV preparation relies on achieving both the desirable size and the unilamellarity of the vesicles. The first parameter is crucial in terms of the curvature as well as the lateral membrane tension of the vesicles. For example, the membrane binding of many proteins depends on

Modeling Viral Budding with Unilamellar Vesicles

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membrane curvature (e.g., Peter et al., 2004; Vaccaro et al., 1993). Thus, if the vesicle population exhibits high size variability, results of the binding assays may be truly confusing. Hence, the size distribution of the IUV sample is to be measured routinely, for example, by dynamic light-scattering. The lamellarity of the vesicles is extremely important when quantitative results are obtained from a binding assay. Even if the phospholipid concentration is determined for a particular IUV sample, the amount of the lipid accessible to the proteins, that is, lipids in the outer monolayer of a vesicle, may be up to orders of magnitude lower than for a unilamellar vesicle. Multilamellarity also affects the amount of encapsulated material into the vesicle and the effective membrane rigidity. This may lead to potential artifacts in experiments, for example, in measurements of the proteininduced leakage of encapsulated material by fluorescence spectroscopy. Thus, the routine of checking the unilamellarity of the IUVs should be adopted as well, for example, by TNBS assay (New, 1990a). A direct measurement of the IUVs lamellarity is offered by the electron microscopy imaging of ammonium molibdate negatively stained samples, as described previously (Bugelski et al., 1990). The use of uranyl acetate (UA) stain with this kind of samples is highly discouraged, as UA interferes with lipid membranes, especially with the charged ones (Caffrey et al., 1987; Parsegian et al., 1981).

3.2. IUV preparation For the M protein experiments, IUVs are prepared by a combination of the gentle hydration method (Reeves and Dowben, 1969) with the extrusion procedure (Mayer et al., 1986). IUV are formed from mixtures of the lipid stocks in chloroform of dioleoyl phosphatidylcholine (DOPC), palmitoyl oleoyl phosphatidylcholine (POPC), dioleoyl phosphatidylethanolamine (DOPE), dioleoyl phophatidylglycerol (DOPG) and cholesterol (chol), doped with 0.2 molar percent of lipid fluorescence marker Lissamine Rhodamine B DOPE (Rh-DOPE). All lipids are from Avanti Polar Lipids (AL). IUVs used for the binding assay with M protein are prepared from the following lipid mixtures (in molar ratios): DOPC:DOPE:Chol:Rh-PE 60:29.8:10:0.2 (PE-chol neutral mixture); POPC:Chol:Rh-DOPE 69.8:30:0.2 (chol neutral mixture); POPC:Rh-PE 99.8:0.2 (neutral mixture); POPC:DOPG:Rh-PE 84.8:15:0.2 (charged mixture). The total lipid concentration in the final IUV sample for these mixtures is 1 g/L. For the assay of the M-protein-induced IUV leakage, the PE–chol neutral mixture is employed, with total lipid concentration of 0.5 g/L in the IUV sample. Each lipid mixture (initially in chloroform) is added to a round bottom of 5 or 10 mL flask, suitable for a rotary–evaporator system (such as VV Micro Evaporator (Heidolph Brinkmann LLC, IL) or equivalent). Chloroform and methanol (9:1, v/v) are added to the flask until a final volume of

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0.5 mL is achieved. The flask if rapidly transferred to the rotary– evaporator, where the evaporation of the organic solvents is carried out with gentle warming of the flask (30 ºC or above the phase transition temperature(s) of the lipids in the mixture) and an intermediate flask rotation speed, in order to obtain a thin and uniform lipid film. Once the visible traces of the organic solvent have disappeared, the evaporation continues for another 15–30 min. Then, the flask is transferred to a highpressure lyophilizer at room temperature for 1 h to ensure the removal of residual solvents. The desired aqueous buffer is added to the flask and the lipid film is left to hydrate overnight at 37 ºC. Special care is taken to avoid shaking the flask during the hydration process. In the case of the IUV preparation for the binding assay, the aqueous buffer is 100 mM KCl, 20 mM HEPES, and 0.2 mM EDTA, pH 7.4 (buffer C). For leakage measurements, an aqueous fluorescent dye should be entrapped inside the IUVs. Thus, buffer D (12.5 mM ANTS, 45 mM DPX, 20 mM KCl, 0.2 mM EDTA, 20 mM HEPES, pH 7.4) contains 8-aminonaphthalene-1,3,6-trisulfonate (ANTS) quenched with p-xylenebis(piridinium bromide) (DPX) (Molecular Probes, Invitrogen Corp., CA) (Du¨zgu¨nes¸, 2003a). The quenching efficiency is proportional to the concentration of ANTS/DPX. The dyes are loaded in to IUV at high concentration ensuring effective quenching of ANTS fluorescence. The leakage measurements are based on the increase of the ANTS fluorescence upon dilution. A small (<1 nm) hole in the IUV membrane is sufficient to let ANTS/DPX release to the bulk solution, thus causing the dilution. Encapsulation of dyes different in size allows estimating the size of such holes. For example, 70 kDa fluorescein isothiocyanate conjugated dextran, FITC-dextran (Sigma-Aldrich Corp., MO), contained in buffer E (75 mg/mL 70 kDa FITC-dextran dissolved in buffer C), requires a much larger (5 nm) hole in the membrane to escape. Iso-osmolarity of buffer D, buffer E, and buffer C is controlled using a Wescor 5500 (UT) vapor pressure osmometer or equivalent. The lipid cloud formed after hydration (mainly containing MLVs and LUVs) is dispersed in the buffer by gentle vortexing of the flask. Using a glass Pasteur pipette, the vesicle dispersion is loaded into an extruder (Lipex Biomembrane Inc., Canada) preassembled with a drain disc and one or two (usually 0.1 mm, if not indicated otherwise) polycarbonate filter (GE Osmonic Labstore, MN). The samples are extruded 15 times. Nonencapsulated ANTS/DPX is separated from IUVs using prepacked Sephadex PD-10 columns (GE Healthcare Biosciences Corp., NJ). Non-trapped 70 kDa FITC-dextran is removed using a column packed with Superose 6 prep grade (GE Healthcare Biosciences Corp.) at a 1 mL/min flow rate. In both cases, buffer C is used as the elution buffer. A crude estimation of the amount of lipids in the preparation is made by measuring the fluorescence signal from Rh-DOPE. For that, 96-well plate apt for

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fluorescence measurements with a plate reader instrument (Synergy 4 microplate reader (Biotek, VT) or similar) is used. Ten microliters aliquots from the IUV suspension before and after extrusion are diluted to 100 mL of buffer C. The Rhodamine (Rh) fluorescence is measured at 550 nm excitation and 590 nm emission wavelengths. The fluorescence of each sample is measured three times and the arithmetic mean of the fluorescence signals is used to estimate the amount of lipid lost during extrusion process by the following formula: lipid loss 

Fbefore
Fafter ; Fbefore

ð4:1Þ

where Fbefore stands for the mean Rh fluorescence signal before extrusion and Fafter, for mean Rh fluorescence signal after the extrusion for each sample prepared. The total lipid concentration of the IUV preparation is then estimated by applying the following formula: CIUV  Ci ð1
lipid lossÞ;

ð4:2Þ

where CIUV stands for the total lipid concentration in the IUV preparation and Ci stands for the total lipid concentration of the initial lipid mixture. We note that this method provides a rough estimation of the amount of total lipid in the final IUV preparation and cannot discriminate between lipid species in nonhomogeneous lipid mixtures. Alternatively the phosphate analysis (Bartlett, 1959; Du¨zgu¨nes¸, 2003b) or liquid chromatography combined with mass spectrometry (Byrdwell, 2005) should be employed if the amount of a particular lipid in the sample is a critical parameter. The size distribution and the lamellarity level of the IUV samples are routinely controlled before using them in M protein experiments. Only IUV preparations that show unimodal size distributions with mean diameters of 120  30 nm by dynamic light-scattering measurements with a dynamic light-scattering instrument (N4 Plus Submicron Particle Sizer (Beckman Coulter Inc., FL) or equivalent) are used in the experiments. Furthermore, the unilamellarity of these samples is also confirmed periodically by negative-staining EM as described in (Bugelski et al., 1990).

3.3. Analysis of M protein binding to IUVs of different lipid compositions Stable binding of M protein to PE–chol neutral, chol neutral, neutral and charged mixtures is assayed by ficoll gradient flotation or sedimentation methods (Fraley et al., 1980). Ficoll solution of 50% (w/v) is prepared by dissolving 25 g of Ficoll PM400 (GE Healthcare Life Sciences, NJ) in buffer C (the amount of this stock increases or decreases proportionally to the amount of samples in the binding assay). This concentration is near the

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solubility limit of ficoll, so the solution preparation requires slight heating and extensively mixing. Solution containing smaller amount of ficoll are prepared by simple dissolving of the appropriate amount of Ficoll PM400 in buffer C. In simple two-step flotation experiments, 111 mg of M protein (2.8 nmol) is incubated with IUVs at 1:200 molar ratio of M protein to lipid for 5 min at room temperature. In the control experiments, the same amount of the protein or IUVs are diluted to the same volume as the IUV/protein mixture and similarly incubated in buffer C. After incubation, each sample is mixed with 50% (w/v) ficoll so that the ficoll is diluted to 20% (w/v). The final volume of this mixture should not exceed 1.2 mL. Then, two-step ficoll gradients are prepared in a 13  51 mm (4 mL total volume) polycarbonate centrifuge tube (Beckman Coulter Inc., CA). The 20% (w/v) ficoll solution mixed with the protein-IUV samples is carefully transferred to the bottom of the centrifuge tubes. Next, the 10% ficoll solution is layered on top (the volume of this layer is usually greater than 2.5 mL). The last gradient step is composed of 200 mL of buffer C. The gradients are then centrifuged using a swinging bucket rotor SW 55Ti (Beckmann Coulter Inc., CA) at 4 ºC and 150,000g for 30 min. IUV, seen as the Rh-labeled band, settle at the boundary between buffer C and 10% ficoll layers. Similar Rh-labeled band is seen in IUV–M protein mixture. These bands can be better observed and documented by illuminating the centrifuge tubes from the top with a flash light and acquiring a photograph (Fig. 4.1). Multistep gradients are required to resolve whether several subpopulations of IUVs are formed upon the protein adsorption. For example, a six-step gradient is prepared by layering, from the bottom to the top of the centrifuge tube, 500 mL layers of 20%, 15%, 12%, 9%, 6%, and 3% ficoll. The samples are deposited very carefully on top of the last layer. The gradients are centrifuged at 150,000g in a SW55Ti rotor (Beckmann Coulter Inc.) at 4 ºC for 90 min. The location of the protein-IUV band/s depends on the amount of protein and membrane deformation upon its binding to the IUVs. Again, the bands can be observed and documented by illuminating the centrifuge tubes from the top with a light source. These images may be very illustrative, as they give qualitative macroscopic description of the behavior of each sample as compared to the IUV control (Fig. 4.1). Independently of the method used, the gradients are fractionated into aliquots (smaller than the volume of a gradient layer, e.g., 100 mL) by using a peristaltic pump connected through tygon type tubing to 1.5 mm capillary borosilicate glass (#PG10150-4, WPI Inc., FL) carefully brought to the bottom of the centrifuge tube. A fraction collector (Model 2110 (BioRad Labs., CA) or equivalent) is attached to the output of the pump. After being collected, the 10 mL of each fraction is diluted to 100 mL with buffer C in a 96-well plate suitable for fluorescence measurements (Fig. 4.1). The Rh

550 nm em 590 nm ex

Rh fluorescence measurement of all the fractions collected IUV control IUV + protein

fmax Peak

Analysis

FLuorescence, a.u.

Fractionation

Baseline 0 10 20 30 40 50 Gradient fraction #

1.0

0.5

0.0

Normalized fluorescence

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Modeling Viral Budding with Unilamellar Vesicles

Figure 4.1 Steps of the assay for protein binding to IUV by sedimentation (or flotation). The left panel shows the photographs of the control (only IUV) and experimental sample (IUV plus protein sample) bands after centrifugation in ficoll gradients. The samples are then fractionated and each fraction is analyzed by measuring the Rh fluorescence at 550/590 excitation/emission wavelengths using a microplate reader. The plot on the right represents the fluorescence intensity of the small aliquots from each fraction versus the fraction number. For the case shown, the IUV in the control (black squares) and in the assayed (gray circles) samples distribute predominantly in two similar populations (two fluorescence peaks). SDS–PAGE analysis of the fraction corresponding to the maximum of the smaller peak shows no significant presence of protein. Thus, the quantification of the protein binding is performed only for the fractions in the peak marked with a gray frame. The plot is normalized by subtracting the baseline (black dashed line) and dividing by the maximum value of the fluorescence peak ( fmax). The normalized scale is shown on the right side of the plot. The detailed analysis of the protein binding is further described in the text.

fluorescence, indicating the amount of lipid in each fraction, is then measured at 550 nm excitation wavelength and 590 nm emission wavelength in a plate reader instrument (Synergy 4 microplate reader (Biotek) or similar). This allows the construction of a plot of fluorescence intensity of each fraction versus the fraction number by using a suitable software (Origin 7.0 (OriginLab Corp., MA) or similar) (Fig. 4.1). The peaks in this plot show the distribution of IUV between fractions. The fractions corresponding to the peaks of the distribution of Rh fluorescence (‘‘Rh-pick fractions’’) are selected to explore for the M protein contents by SDS–PAGE. The same fractions from the control samples are also selected and analyzed by SDS–PAGE. It is recommendable to load the maximum sample volume possible into the gel (the 10-well Pierce Precise Protein Gels (Pierce) are a good option to consider, as they allow to load as much as 50 mL of a sample per line). As it is very difficult to guess a priori the percentage of protein bound to the IUV in each particular case, it is recommendable to initially use sensitive stains, such as SyproRuby Red Gel Stain (Molecular Probes, Invitrogen Corp.) or similar in the initial binding experiments. Once the binding has been characterized, less-sensitive stains, such as SimplyBlue SafeStain (Invitrogen Corp.) or equivalent may be used.

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Equal volumes of the selected fractions are subjected to the reducing treatment and then loaded into the gel together with a line marker. The gel should also include a calibration line with a known amount of protein loaded. Stained gel images are acquired and analyzed using a gel imaging system (FluorChem FC (Alpha Innotech Corp., CA) or equivalent). M protein exhibits stronger binding to chol containing liposomes, with certain preference for the PE–chol neutral composition, likely because of less dense lipid packing in such mixtures related to high spontaneous curvature of PE (Fuller and Rand, 2001). M protein shows lower affinity for the compositions containing charged lipids. No protein signal is detected in the equivalent protein control fraction (Shnyrova et al., 2007). The areas under the intensity peaks of the protein bands are further analyzed using an appropriate software (Origin 7.0 (OriginLab Corp.) or similar). This analysis allows establishing the percentage of the protein bound to each IUV type. First, the amounts of protein in each line of the gel are found by normalizing to the area of the protein intensity peak in the calibration line: gL ¼

AL gCL ; ACL

ð4:3Þ

where gL are the grams of protein loaded in the sample line, AL is the area under the intensity peak of the protein band, ACL is the area under the intensity peak of the protein band in the calibration line, and gCL are the grams of protein loaded into the calibration line. The amount of protein calculated this way is per volume of sample loaded into the gel. Thus, the amount of protein for the fraction with the maximum Rh signal will be: gf max ¼

Vf gL ; VL

ð4:4Þ

where gfmax stands for the grams of protein in the studied fraction, Vf is the total volume of the fraction (100 mL), VL is the volume of the fraction loaded into the gel line, and gL are the grams of protein loaded in the gel line [see Eq. (3)]. However, this procedure underestimates the amount of protein bound to IUVs, as the IUV-protein band usually extends for more than one fraction collected from the centrifuge tube (plot in Fig. 4.1). To correct for the smearing of IUV over the gradient, the plot of Rh fluorescence versus the fraction collected is used. It is assumed that the distribution of M protein follows the distribution of IUV in the fractions, that is, the fractions which signals contribute to the Rh-fluorescence peak on the plot. Each point of the fluorescence represents a homogeneous fraction of IUV with bound M proteins. The maximum point of the peak corresponds to the fraction analyzed by SDS–PAGE and which protein amount is a known

65

Modeling Viral Budding with Unilamellar Vesicles

parameter (gfmax). To calculate the amount of protein in the entire band (i.e., for all the fractions that form the fluorescence peak), first, the plot showing the distribution of Rh fluorescence is baseline-corrected leading to elimination of the fractions with no detectable peaks of Rh fluorescence (plot in Fig. 4.1). Next, the fluorescence values for each fraction of the fluorescence peak are normalized by dividing for the fluorescence value of the peak maximum. By doing so, the amount of protein bound to the single population of IUVs is calculated as follows: gB ¼ gf max

n X Ff n ; F n¼1 f max

ð4:5Þ

where gB denotes the grams of protein bound to the IUVs, gfmax is calculated using Eq. (4), Ffn is the fluorescence value minus the baseline for the fraction n of the fluorescence peak and Ffmax is the fluorescence value minus the baseline for the fraction with the maximum fluorescence intensity of the peak. If peaks of the Rh fluorescence corresponding to different populations of IUV (or gradient bands) overlap, simple deconvolution analysis is needed to estimate the amount of the protein in each IUV population. Finally, the percent of the total protein bound to a single population of IUVs is estimated by: % bound protein ¼

gB  100; gi

ð4:6Þ

where gB is calculated from Eq. (4) and gi denotes the grams of protein initially in the binding assay (111 mg in the case considered). The fractions can be further analyzed for IUV morphological deformations caused by adsorbed proteins via EM imaging. Cryo-EM techniques are favorable as IUV deformations caused by the adhesion and flattering to the supporting film of the grids used for negative-staining EM are avoided altogether.

3.4. M protein interaction with IUVs: Leakage of aqueous dyes The IUV stability upon M protein adsorption is explored using IUVs loaded with aqueous fluorescent markers, either small (ANTS/DPX) or large (70 kDa FITC-dextran). Leakage of these aqueous dyes from IUVs upon the M protein additions is detected at ambient temperature and under constant stirring by spectrofluorimetric measurements using a spectrofluorometer (Aminco-Bowman SLM-2 (Spectronic Instruments Inc., NY) or equivalent). In a cell suitable for fluorescence measurements and containing a stirrer bar IUV sample is diluted with buffer C. Protein is added in an amount such as the protein/lipid molar ratio increases 0.01 units after each

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protein addition. The increase in integral fluorescence due to the release of the dyes is detected. The normalized fluorescence intensity (FN) is recalculated from the integral IUV fluorescence intensity (F) as: FN ¼

F
Fi ; FT
F i

ð4:7Þ

where Fi corresponds to the fluorescence intensity before the protein addition and FT to the fluorescence intensity after complete disruption of IUV (infinite dilution of the fluorophores) by detergent (0.1% of Triton X-100). The excitation/emission wavelengths used are 380/520 nm for ANTS/DPX detection and 550/590 nm for Rh-DOPE detection. The experiments are carried out in triplicate. Data analysis is performed with Origin 7.0 software or similar. Starting at 0.01 protein/lipid ratio, release of both markers is observed with slower kinetics for the larger marker (Shnyrova et al., 2007). The possibility that the observed effects are caused by some artifact present in the protein solution (such as traces of detergent triton that is used during the protein purification) is ruled out by a-chymotrypsin proteolytic (a-chymotrypsin from bovine pancreas; Sigma-Aldrich Corp.). 5 mM protein solution is incubated 5-min at room temperature with a-chymotrypsin in 1:5 molar ratio. The digestion of M protein upon this treatment is confirmed by an SDS–PAGE (Shnyrova et al., 2007). After this treatment, the sample containing pre-digested M protein is added to the IUVs in the same conditions as the M protein without digestion. The efficiency of content release from IUV after protein digestion is greatly impaired (Shnyrova et al., 2007). Release efficiencies are comparable for both high- and low-molecularweight markers for the same amount of the protein added, thus most probably indicating vesicle bursting, rather than membrane pore formation. Also, inward invaginations of the IUV membrane expected upon the M protein adsorption should elevate membrane tension (Shnyrova et al., 2007), thus increasing the possibilities for membrane bursting, versus small pore formation. Thus, ‘‘massive’’ leakage with no size selectivity, indicating bursting, might be used as a rough preliminary criterion for the curvature activity of a protein.

4. Reconstitution of M-Protein-Driven Membrane Budding on GUVs 4.1. GUV preparation GUVs are prepared using a slight modification of the electroformation protocol (Angelova and Dimitrov, 1988). The design of a custom made electroformation chamber used is depicted in Fig. 4.2A. The chamber

B

10

m

m

Function generator ~ V

Pipette holder

m

m

Platinum electrodes

Pressure valve

y

35 mm plastic dish

ΔP 10

m

G

U

5 mm

V f zo orm ne at

m

Lipid application device

io

n

20

xyz micromanipulator

od Eb

Flexible tubing

Pt electrodes

PTF

PT F bi E ng

tu

Connection to function generator

A

Electroformation chamber

Microscope stage Objective

Figure 4.2 Electroformation of GUV. (A) Schematic representation of the lipid film application device and the electroformation chamber with Pt electrodes. (B) Schematic representation of the setup for observation of the GUV formation and for further manipulations of the electrode-bound GUVs.

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Anna V. Shnyrova and Joshua Zimmerberg

consists of a polytetrafluoroethylene (PTFE) body and two 0.8 mm platinum (Pt) electrodes (Goodfellow Corp., PA) inserted into the PTFE block in such a way that on one side both electrodes are parallel to each other (this is the place for the GUVs to form), while on the other side the electrodes go in opposite directions. This is the place where the connector cables to the voltage generator are attached. Lipid mixtures used for GUV preparation are the following (in molar ratios): DOPC:DOPE:Chol:Rh-DOPE (48:43:4:5); POPC:Chol:RhDOPE (90:5:5); POPC:Rh-PE (95:5); POPC:DOPG:Rh-PE (80:15:5), similar to those used in IUV experiments. In some cases, the difference in composition between IUV and GUV is dictated by the electroformation protocol where each particular lipid mixture, upon drying, has to form a film covering platinum electrodes. When lipid mixture includes charged species, like DOPG, the mixture is dissolved in chloroform:methanol:ethyl ether (4:1:5 volume ratio), to a final concentration of 0.1 g/L of total lipid. In all other cases, the mixtures are dissolved in chloroform:methanol (9:1 volume ratio), to a final concentration of 0.1 g/L. A custom made application device is used for application of the lipid mixture to the Pt electrodes (Fig. 4.2A). This device consists of a 0.56 mm ID  1.07 mm OD PTFE microbore tubing (Cole Palmer, IL) of 2 cm length inserted into flexible tubing of 1.02 mm ID (such as Masterflex Tygon Tubing, Saint-Gobain Corp., PA) with 1 cm length that is hermetically closed on the opposite side to the PTFE tubing by a small screw at the end. This device offers the advantage of being disposable after use (alternatively, a 10 mL Hamilton syringe (Hamilton Company, NV) can be used for lipid application, but care should be taken to clean the syringe thoroughly after each use with appropriate lipid solvent combination, such as ethanol, methanol, chloroform. Sonication is also a good option for syringe cleaning). Before using the lipid solution, the device is washed with pure chloroform by repetitive cycles of squeeze/release of the flexible tubing part, while a small part of the PTFE tubing is immersed into the chloroform solvent. After washing, the PTFE end of the device is immersed into the lipid solution. By squeezing-releasing of the flexible part of the device a small amount of the solution is withdrawn. Two small drops of it are deposited on each one of the two platinum electrodes of the electroformation chamber by gently squeezing the flexible part of the application device. After the application, the electrodes are dried under vacuum for at least 1 h or overnight. Then, the electroformation chamber is assembled on a plastic disposable dish with a 0.17 mm thick cover glass bottom (Delta TPG Dish (Bioptech, PA) or similar; Fig. 4.2B) and the dish is mounted on a stage of an inverted microscope (Axiovert 200 (Carl Zeiss Inc., Germany) with a fluorescence excitation source, a filter cube combination suitable for Rh observation and a 40/0.65 and a 100/1.25 objectives (ACHROPLAN; Carl Zeiss Inc.) or equivalent microscope configuration). The electrodes are

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connected to a function generator (GFG-3015 model function generator (Instek America Corp., CA) or equivalent) preset to apply a 10 Hz and 0.2 V sinewave. Right after the initial voltage application, a buffer containing 20 mM HEPES and approximately 200 mM sucrose, pH 7.4 is gently added to the dish in such a way that it is slightly above the electrodes. This buffer should be scrupulously equilibrated osmotically with buffer C by using the procedure described above. During the following 15 min, the sinewave amplitude is gradually increased from 0.2 to 1.0 V. Electroformation times varied from 15 min to 1 h for different lipid compositions of the GUV. After GUVs become visible on the electrode by bright-field or fluorescence microscopy observation with a 40 objective, the voltage is lowered to 0.2 V and then turned off completely. If the GUVs are to be detached from the electrodes, the frequency is lowered to 0.1 Hz before lowering of the voltage. Then the GUVs are collected with 200 mL pipette and stored in a microcentrifuge tube until usage. If the GUVs are to be left on the electrodes, the plastic dish is perfused with buffer C for 15 min by using an appropriate microscope perfusion system (such as the micro-perfusion system (Bioscience Tools, CA) or equivalent). Visualization of GUVs in the attached configuration is performed directly on the inverted microscope stage immediately after perfusion of the chamber with buffer C.

4.2. Interaction of M protein with GUVs Visualization of GUVs in the attached to the electrode configuration (Fig. 4.3A) is performed directly on the inverted microscope stage immediately after perfusion of the chamber with buffer C. The biggest advantage of this configuration is the relative immobilization of the GUVs on the electrodes, which facilitates greatly the application of the protein in the proximity of the GUV, as well as allows to easily patching the GUV membrane. However, the fluorescence background from the electrode and other GUVs in the vicinity sometimes interferes with the imaging of a single GUV. Also, the proximity of the electrodes to the bottom of the dish becomes a critical factor when high-magnification objectives (such as 100 or bigger) are used. These objectives have small working distances, thus there is a requirement for the electrodes to be close to the bottom of the dish, something not easily achievable with 0.8 mm thick electrodes. To observe individual vesicles at high magnifications, the detached GUVs are routinely used. In this case, a 0.17-mm-thin glass 35 mm dish (Delta TPG Dish (Bioptech) or similar) is pretreated with 1 g/L BSA for 1 min and thoroughly washed with buffer C in order to avoid GUV spreading and bursting upon contacting the glass bottom of the dish. The dish is setup on the microscope stage and 1 mL of buffer C is added. GUV suspension of 100–200 mL collected after electroformation is added to the

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A M protein Pipette

GUV Platinum electrode

B M protein Pipette

GUV BSA pretreated cover slip

Figure 4.3 Main configurations used in GUV experiments. (A) GUV remains bound to the electrode and the M protein is applied by patching the GUV membrane with a patch-pipette filled with the protein solution of choice. (B) A GUV detached from the electrode is situated on the glass bottom of the dish (pretreated with BSA) and the M protein is applied to the GUV through the patch-pipette brought in proximity to the GUV. Pulses of positive hydrostatic pressure cause protein efflux.

1 mL of buffer C in the dish. As GUVs are formed in the sucrose buffer, osmotically equilibrated with buffer C, after 5–10 min all the GUVs are settled in the glass coverslip of the bottom of the dish due to a difference in density between the two buffers (Fig. 4.3B). In this case, GUVs are easily visualized by high-resolution fluorescence microscopy, however, the fact that GUVs are only slightly attached to the bottom, makes the experiments with protein delivery more involved. For both configurations, prior to the protein addition, the intrinsic behavior of a target vesicle is recorded for a period of time (1–3 min). The recording is made using a digital camera (iXon (Andor, CT) or similar) attached to the camera port of the microscope. The camera is controlled through a PC with MicroManager open source microscope software (University of California, San Francisco, CA) or equivalent. The unilamellarity of the GUV may be established by fluorescence analysis of the GUV contour, as was described previously (Akashi et al., 1996). Upon establishing the stability (and unilamellarity) of the target GUV, the M protein is applied through a pipette brought to the close proximity of the GUV (Fig. 4.3B). The pipettes for M protein application are obtained by pulling a 1.5 mm capillary borosilicate glass (#PG10150-4, WPI Inc.) with a micropipette puller (Flaming Brown Micropipette puller, Model P80/PC; Sutter Instruments Co., CA). The pipette is back-filled with

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M protein solution using a 1 mL syringe equipped with MicroFilTM (WPI Inc.). Three motorized actuators (12.5 mm Travel Open-loop DC CMA Actuator; Newport Corp., CA) attached to a crossed-roller bearing linear stage, 0.5 in., with XYZ travel (461-XYZ-M, Newport Corp.) are used to control the pipette position. The pipette filled with M protein solution is inserted into a microelectrode holder with pressure port (WPI Inc.) attached to the linear stage (Fig. 4.2B). A 1-mL syringe is attached to one port in a three-way luer valve (WPI Inc.). Another port of the valve is connected through long PTFE tubing to the pressure port of the microelectrode holder. This allows controlling the pressure difference applied to the pipette (i.e., for protein application or liposome patch isolation). The luer valve should be closed (i.e., turned toward the unoccupied port) during the pipette search in the objective focus area. The delivery pipette containing a concentrated solution of the protein (4 mM) is brought carefully to the proximity of the studied GUV, the pipette motion being observed by bright-field microscopy. Once the pipette is located in the proximity of the vesicle, it is directed toward the GUV and the luer lock in the pressure loop is opened. A pulse of positive hydrostatic pressure is applied to the pipette by gently pressing the pressure syringe. Immediately, a small flow of protein can be seen exiting the pipette. Prior to the protein application, the microscope light pass is switched to the fluorescence mode and the morphological changes of the GUV membrane are monitored and documented by acquiring time lapse series. The protein concentration in the vicinity of GUV increases transiently. Shortly after protein application, invagination of the GUV membrane is detected. After each protein application the apparent size of the GUV decreases and the steady-state fluorescence in the GUV projection area increased (Shnyrova et al., 2007). Internalized membrane vesicles are clearly detected, thus justifying the fluorescence changes. Special care should be taken about the pressure applied to the delivery pipette when GUVs slightly attached to the dish bottom are used. If the pressure is too strong, the GUV can be detached from the bottom of the dish and flow away. In this case, the pressure valve of the pressure syringe should be closed, the pipette should be lift immediately and a new GUV, far enough from the initial observation point, should be found. Several experiments can be performed in the same dish. However, the pipette should be exchanged after each one of them to avoid protein dilution effects. GUVs of different compositions are used in order to establish the preferences for M-protein-induced budding. Surprisingly, and against the prediction made upon the binding preferences of M protein detected with IUVs, vesicle formation is more effective when cholesterol or charged lipids are present in the GUV composition. Thus, the budding activity of M protein is not directly related to the efficiency of its membrane binding (Shnyrova et al., 2007). Another important observation is formation of transient, membrane

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domains upon application of M proteins to GUV. The appearance of these domains indicates that M proteins self-organize into fluid-like formations which then bud away from the membrane (Shnyrova et al., 2007). Overall, these experiments directly demonstrate that M protein implements the genetically encoded information required to create the closed spherical membrane particles through its interactions with lipid bilayer. The validity of such observations should be routinely checked by doing buffers and negative protein controls (i.e., by applying the M protein buffer containing no proteins or a reference protein, such as albumin, in similar concentration as M protein). Those are very important controls, as osmotic pressure, as well as bulk protein binding, can greatly affect the behavior of the GUVs. For example, differences in osmotic pressure between the exterior and the interior of the GUV can cause spontaneous budding and formation of vesicles in the GUV lumen without the need of a protein, as in the case described by Guedeau-Boudeville et al. (2000). Thus, iso-osmolarity of every solution used with GUVs should be particularly monitored. When experiments are performed on electrode-attached GUVs, the procedure is almost the same as with detached GUVs. However, the manipulations are eased by the fact that GUV is immobilized. Patch-clamp type experiment, hard on the slightly attached GUV, can be performed using single GUVs on the electrode (Fig. 4.3A). For these experiments, the luer pressure valve on the pressure syringe is opened before the pipette approaches a GUV closely. After touching the GUV, a very small negative pressure is applied. The GUV membrane is sucked in and a stable contact, seen as a small membrane patch inside the pipette, forms. Protein adsorption on this isolated patch inside the pipette starts instantaneously, and the morphological changes of the patch membrane, as well as of the whole liposome are observed and documented by time lapse series as described before. The membrane rearrangements quickly lead to the budding directed toward the interior of the GUV. Round vesicles of different diameters become visible near the patch and inside the GUV. The membrane outside the patched area provided a lipid reservoir to support this small vesicle formation. Thus, the GUV shrinks while more vesicles are produced. Finally, the GUV membrane detaches from the pipette and multiple vesicles are seen moving inside the GUV (Shnyrova et al., 2007). Again, buffer and negative protein controls should be performed in this configuration. The patch-clamp experiments emphasize the localized character of the M protein action. It is an important observation, as GUVs are notorious for their extreme sensitivity to the global changes in GUV membrane, such as introduction of area asymmetry (difference in area) between monolayers of GUV membrane. These experiments also indicate the power of M protein assembly: constitutive formation of membrane vesicles lead to building up the lateral membrane tension powerful enough to induce the GUV detachment from the pipette. Analyzing these observations provide valuable mechanistic insights into curvature-driven activity of M protein (Shnyrova et al. 2007).

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5. Concluding Remarks The assays described in this chapter represent a relatively easy approach for studying the activity of cellular protein involved in controlling membrane curvature. The approach is composed of two parts. The initial analysis is based upon quantification of membrane binding and leakage for membranes of different lipid compositions. Although less spectacular than the GUV-based experiments, these assays offer a significant amount of information about the mechanism of protein–lipid interaction. They can also provide ultrastructural information upon EM analysis of membrane shape transformations by bound proteins. Nevertheless, these measurements are lacking the dynamic aspects of the budding process. Microscopic observations of the GUV membrane provide a versatile tool to visualize the dynamics of membrane curvature creation. These observations reveal dynamic, transient intermediates of the budding process, such as membrane domains, which would not be resolved by structural analysis. However, these observations are limited by the optical resolution of the microscope itself. To resolve smaller structures (i.e., <200 nm), as well as to confirm fission of the vesicle neck, more sensitive techniques should be used in combination with the methods described here (such as measurements of membrane area and permeability by electrical admittance measurements on planar bilayer lipid membranes; see Shnyrova et al., 2007). Ultimately, the combination of ultrastructural, electrophysiological, and microscopy techniques form an experimental environment critical for obtaining real mechanistic insights into complex processes of membrane morphogenesis.

REFERENCES Akashi, K., Miyata, H., Itoh, H., and Kinosita, K. Jr. (1996). Preparation of giant liposomes in physiological conditions and their characterization under an optical microscope. Biophys. J. 71, 3242–3250. Angelova, M. I., and Dimitrov, D. S. (1988). A mechanism of liposome electroformation. Prog. Colloid. Polym. Sci. 76, 59–67. Bartlett, G. R. (1959). Phosphorus assay in column chromatography. J. Biol. Chem. 234, 466–468. Briggs, J. A., Wilk, T., and Fuller, S. D. (2003). Do lipid rafts mediate virus assembly and pseudotyping? J. Gen. Virol. 84, 757–768. Bugelski, P. J., Sowinski, J. M., and Kirsh, R. L. (1990). Negative stain electron microscopy. In ‘‘Liposomes: A Practical Approach,’’ (R. R. C. New, ed.), pp. 140–161. Oxford University Press, New York. Byrdwell, W. C. (2005). Methods for Lipid Analysis by Liquid Chromatography/Mass Spectrometry and Related Techniques. AOCS Press. Caffrey, M., Morris, S. J., and Feigenson, G. W. (1987). Uranyl acetate induces gel phase formation in model lipid and biological membranes. Biophys. J. 52, 501–505.

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Coorssen, J. R., Blank, P. S., Albertorio, F., Bezrukov, L., Kolosova, I., Backlund, P. S. Jr., and Zimmerberg, J. (2002). Quantitative femto- to attomole immunodetection of regulated secretory vesicle proteins critical to exocytosis. Anal. Biochem. 307, 54–62. Du¨zgu¨nes¸, N. (2003a). Fluorescence assays for liposome fusion. Methods Enzymol. 372, 260–274. Du¨zgu¨nes¸, N. (2003b). Preparation and quantitation of small unilamellar liposomes and large unilamellar reverse-phase evaporation liposomes. Methods Enzymol. 367, 23–27. Fraley, R., Subramani, S., Berg, P., and Papahadjopoulos, D. (1980). Introduction of liposome-encapsulated SV40 DNA into cells. J. Biol. Chem. 255, 10431–10435. Fuller, N., and Rand, R. P. (2001). The influence of lysolipids on the spontaneous curvature and bending elasticity of phospholipid membranes. Biophys. J. 81, 243–254. Garcia-Sastre, A., Cabezas, J. A., and Villar, E. (1989). Proteins of Newcastle disease virus envelope: Interaction between the outer hemagglutinin-neuraminidase glycoprotein and the inner non-glycosylated matrix protein. Biochim. Biophys. Acta 999, 171–175. Guedeau-Boudeville, M. B., Bradley, J. C., Singh, A., and Jullien, L. (2000). Changes in the morphology of giant vesicles under various physico-chemical stresses. In ‘‘Giant Vesicles,’’ (P.a.W. Luigi Luigi, ed.), pp. 341–351. John Wiley & Sons, LTD, West Sussex, England. Jayakar, H. R., Jeetendra, E., and Whitt, M. A. (2004). Rhabdovirus assembly and budding. Virus Res. 106, 117–132. Mayer, L. D., Hope, M. J., and Cullis, P. R. (1986). Vesicles of variable sizes produced by a rapid extrusion procedure. Biochim. Biophys. Acta 858, 161–168. McCreedy, B. J. Jr., McKinnon, K. P., and Lyles, D. S. (1990). Solubility of vesicular stomatitis virus M protein in the cytosol of infected cells or isolated from virions. J. Virol. 64, 902–906. New, R. R.C (1990a). Characterization of liposomes. In ‘‘Liposomes: A Practical Approach,’’ (R. R. C. New, ed.), pp. 138–139. Oxford University Press, New York. New, R. R. C. (1990b). Liposomes: A Practical Approach. Oxford University Press, New York. Palade, G. E. (1953). An electron microscope study of the mitochondrial structure. J. Histochem. Cytochem. 1, 188–211. Palade, G. (1975). Intracellular aspects of the process of protein synthesis. Science 189, 867. Pantua, H. D., McGinnes, L. W., Peeples, M. E., and Morrison, T. G. (2006). Requirements for the assembly and release of Newcastle disease virus-like particles. J. Virol. 80, 11062–11073. Parsegian, V. A., Rand, R. P., and Stamatoff, J. (1981). Perturbation of membrane structure by uranyl acetate labeling. Biophys. J. 33, 475–477. Peter, B. J., Kent, H. M., Mills, I. G., Vallis, Y., Butler, P. J., Evans, P. R., and McMahon, H. T. (2004). BAR domains as sensors of membrane curvature: The amphiphysin BAR structure. Science 303, 495–499. Reeves, J. P., and Dowben, R. M. (1969). Formation and properties of thin-walled phospholipid vesicles. J. Cell Physiol. 73, 49–60. Sagrera, A., Cobaleda, C., Berger, S., Marcos, M. J., Shnyrov, V., and Villar, E. (1998). Study of the influence of salt concentration on Newcastle disease virus matrix protein aggregation. Biochem. Mol. Biol. Int. 46, 429–435. Scheid, A., and Choppin, P. W. (1973). Isolation and purification of the envelope proteins of Newcastle disease virus. J. Virol. 11, 263–271. Shnyrova, A. V., Ayllon, J., Mikhalyov, I. I., Villar, E., Zimmerberg, J., and Frolov, V. A. (2007). Vesicle formation by self-assembly of membrane-bound matrix proteins into a fluidlike budding domain. J. Cell Biol. 179, 627–633. Solon, J., Gareil, O., Bassereau, P., and Gaudin, Y. (2005). Membrane deformations induced by the matrix protein of vesicular stomatitis virus in a minimal system. J. Gen. Virol. 86, 3357–3363.

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Takimoto, T., and Portner, A. (2004). Molecular mechanism of paramyxovirus budding. Virus Res. 106, 133–145. Vaccaro, A. M., Tatti, M., Ciaffoni, F., Salvioli, R., Barca, A., and Roncaioli, P. (1993). Studies on glucosylceramidase binding to phosphatidylserine liposomes: The role of bilayer curvature. Biochim. Biophys. Acta 1149, 55–62. Welsch, S., Muller, B., and Krausslich, H. G. (2007). More than one door—Budding of enveloped viruses through cellular membranes. FEBS Lett. 581, 2089–2097.

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S E C T I O N

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Detection of Antimycolic Acid Antibodies by Liposomal Biosensors Y. Lemmer,*,‡ S. T. Thanyani,* P. J. Vrey,* C. H. S. Driver,† L. Venter,* S. van Wyngaardt,* A. M. C. ten Bokum,* K. I. Ozoemena,† L. A. Pilcher,† D. G. Fernig,} A. C. Stoltz,*,§ H. S. Swai,‡ and J. A. Verschoor* Contents 1. Introduction 2. Experimental 2.1. Purification of mycobacterial MA 2.2. Fluorescent labeling of MA 2.3. Determination of MA carrying capacity of liposomes 2.4. MA liposome size is influenced by cholesterol and MA content, but not by pH 2.5. MA liposome immobilization on IAsys biosensor cuvettes 2.6. Technology transfer from the IAsys waveguide to the ESPRIT SPR biosensor 2.7. The cholesteroid nature of MA demonstrated on the ESPRIT biosensor 3. Conclusion Acknowledgments References

80 81 81 82 82 84 86 89 100 102 102 102

Abstract Antibodies to mycolic acid (MA) antigens can be detected as surrogate markers of active tuberculosis (TB) with evanescent field biosensors where the lipid antigens are encapsulated in liposomes. Standard immunoassay such as ELISA, where the lipid antigen is not encapsulated, but directly adsorbed to the wellbottoms of microtiter plates, does not yield the required sensitivity and

* { { } }

Department of Biochemistry, University of Pretoria, Pretoria, South Africa Department of Chemistry, University of Pretoria, Pretoria, South Africa Materials Science and Manufacturing, CSIR, Pretoria, South Africa Department of Infectious Diseases, University of Pretoria, Pretoria, South Africa School of Biological Sciences, University of Liverpool, United Kingdom

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64005-2

#

2009 Elsevier Inc. All rights reserved.

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specificity for accurate diagnosis of TB. One reason for this is the cross-reactivity of natural anticholesterol antibodies with MAs. MAs are the major cell wall lipids of mycobacteria. Mycobacterial MA has immunomodulatory properties and elicits specific antibodies in TB patients. Liposomes were optimized for their use as carriers both for the presentation of immobilized purified mycobacterial MA on sensor surfaces, and as a soluble inhibitor of antibody binding in inhibition assays. By using an inhibition assay in the biosensor, the interference by anticholesterol antibodies is reduced. Here, we describe the MA carrying capacity of liposomes with and without cholesterol as a stabilizing agent, optimized concentration and size of liposomes for use in the biosensor assay, comparison of the methods for wave-guide and surface plasmon resonance biosensors and how the cholesteroid nature of MA can be demonstrated by the biosensor when Amphotericin B is allowed to bind to MA in liposomes.

1. Introduction As a result of the development of drug-resistant strains of Mycobacterium tuberculosis and the breakdown of the immune system of its host by HIV, tuberculosis (TB) is no longer a ‘‘controlled’’ disease and has become a major health problem in both developed and developing countries (Houghton et al., 2002). Diagnosis of TB is no longer 100% reliable due to AIDS, and was never adequate for determining extrapulmonary and child TB. HIV coinfection and drug resistance appears to shorten the lifetime of a TB patient considerably, such that it becomes a matter of life and death to be able to diagnose TB within 24 h of sampling. The decision toward treatment cannot be taken lightly, as the treatment regime has to be maintained for at least 6 months to clear all the latent TB from the body. The current drive toward new tools for TB diagnosis arose from these facts, and that the mycobacterial pathogen isolated from sputum samples is slow growing, thereby requiring several weeks to become visible during in vitro growth (Reischl, 1996; Samanich et al., 2000). Although DNA-amplifying technology has reduced the period from sampling to TB diagnosis to within days, it still uses mainly samples obtained from the lungs. Therefore, fast, affordable, and reliable diagnosis of TB has become a high priority in public health (Chan et al., 2000), and is currently actively pursued in several laboratories. Mycolic acids (MAs) appeared to be promising antigens for the design of fast immunodiagnostics for TB (Verschoor and Onyebujoh, 1999). The major cell wall lipids of mycobacteria are the MAs, which are long chain (C60–C90) a-branched, b-hydroxy fatty acids. Mycobacterial MAs are immunogenic. MA was the first nonprotein antigen shown to stimulate the proliferation of human CD4
, CD8
, and CD4þ T-lymphocytes upon CD1 presentation (Beckman et al., 1994; Goodrum et al., 2001). In addition, anti-MA antibodies could be detected in the serum of patients with active

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pulmonary TB (Pan et al., 1999) using the standard ELISA procedure, indicating that free MAs are present in the circulation during active TB disease (Beatty et al., 2000). The prevalence of anti-MA antibodies appeared to be independent of the degree to which a person was suffering from AIDS (Schleicher et al., 2002). Whereas ELISA did not detect anti-MA antibodies well enough to be considered as a basis for a diagnostic test of active TB, biosensor analysis did (Thanyani et al., 2008). It could achieve the required specificity and sensitivity to be regarded seriously as a solution to the current dilemma of standard TB diagnosis taking several weeks after sampling to produce a result. Even the highly sensitive PCR detection of mycobacterial DNA in patient sputum samples still takes a few days to deliver a diagnostic outcome. MAs are soluble in extremely nonpolar organic solvents such as chloroform, dichloromethane, and hexane. Alternatively, they can be ‘‘solubilized’’ in boiling water or aqueous buffers. For the detection of anti-MA antibodies by means of ELISA, MA was either coated from hexane solutions (Pan et al., 1999) or from boiling phosphate-buffered saline (PBS) (Schleicher et al., 2002). This presentation of MA is clearly not physiological. Presentation of MA in liposomal environments more closely resembles the way in which MA is encountered in the body of the patient. Indeed, when encapsulated into liposomes and injected into mice, MA was shown to behave as a typical pathogen-associated molecular pattern (Korf et al., 2005). It could also act as an immunomodulatory compound that suppresses experimental asthma through Treg cell intervention (Korf et al., 2006). It was only logical to derive that diagnosis of TB by detection of surrogate marker antibodies against MA should perform better when MA antigen was presented in liposomes. This was demonstrated by Thanyani et al. (2008) by making use of biosensors. Here, we describe how liposomes are optimized for their use as carriers of MA for immobilization as antigens on sensor surfaces, and as soluble inhibitors of antibody binding in inhibition assays in a TB diagnostic assay. It is dubbed the MARTI-test, as an acronym of Mycolic acid Antibody Real-Time Inhibition. Whereas the original MARTI-test is described for a wave-guide biosensor (Thanyani et al., 2008), we show here how the method is adjusted to also suit the more popularly used surface plasmon resonance (SPR) biosensors. Finally, we demonstrate the cholesteroid nature of MA with the biosensor by measuring the binding of Amphotericin B to MA in liposomes.

2. Experimental 2.1. Purification of mycobacterial MA MAs are isolated and purified from M. tuberculosis H37Rv ATTC 27294 purchased from the American Type Culture Collection (ATTC, Baltimore, MD, USA) as described by Goodrum et al. (2001). The MA is dissolved in

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chloroform (HPLC grade; Merck; Darmstadt, Germany) and aliquotted into glass vials. The chloroform is evaporated under nitrogen gas and the dried MA is stored at 4  C until use. The purified MAs are checked for LPS contamination using the kinetic-QCL Limulus amoebocyte test kit (Sigma, St. Louis, MO). In the experiments shown in this chapter, no LPS is detected.

2.2. Fluorescent labeling of MA MA is fluorescently labeled by derivatization with 5-bromomethyl fluorescein (5-BMF; Molecular Probes, Leiden, The Netherlands), as described by Korf et al. (2005). Quality control is performed by TLC on a silica gel GHL thin layer plate. Chromatography is performed in two dimensions, with chloroform:methanol:water as the mobile phase in the first dimension, and 100% methanol (Merck) as the mobile phase in the second dimension. Iodine vapor is used to visualize the MA. The absence of free 5-BMF, not associated with the MA spot, indicates that the label is covalently bound. Fluorescently labeled 5-BMF–MA is incorporated into liposomes for assessment by the biosensor or flow cytometry.

2.3. Determination of MA carrying capacity of liposomes MAs are dissolved in chloroform and 100 mg quantities are aliquoted into amber vials and stored at 4  C. Liposomes are prepared according to the method described by Bangham (1983). This involves the deposition of a thin lipid film from an organic solvent medium on the wall of a container, followed by agitation with an aqueous medium. In short, phosphatidylcholine (PC from egg yolk, 99% pure; Sigma), cholesterol (Sigma), and dried MA are all dissolved separately in chloroform. PC (9 mg) with or without 4.5 mg cholesterol and varying amounts of MA are mixed together in a glass vial. The chloroform is evaporated under nitrogen gas in a chemical hood. PBS (1 ml) is added and the mixture is heated to 80  C until the lipid film dissociates from the wall of the vial. The hot mixture is then vortexed and sonicated for 5 min using a Branson sonifier for 50 pulses at a 20% output level. MA-carrier liposomes are extracted using a triphase partition method, modified from Dennison and Lovrein (1997). Briefly, a sample (50–200 ml) of MA-carrier liposome solution is diluted to 600 ml with H2O and extracted three times with equal volumes of tert-butanol and chloroform. The chloroform phases are collected and the chloroform is evaporated under nitrogen gas. MA remaining in the vials due to saturation of the liposomes is collected by chloroform rinses. HPLC of the extracted MA is carried out as described (Butler and Kilburn, 1990), after derivatization with p-bromophenacyl bromide. A high molecular weight internal standard

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(Ribi ImmunoChem Research Company, Hamilton, MT, USA) is added to each sample to allow quantification of the extracted MA. Liposomes are made with egg PC, with or without the addition of cholesterol. The maximum amount of MA that can be incorporated reproducibly is about 2 mg/ml for liposomes consisting of phosphatidyl choline and cholesterol, but only about 500 mg MA/ml into liposomes consisting of phosphatidyl choline only (Fig. 5.1). MA not incorporated into the liposomes can in most cases be quantitatively recovered from the wall of the vessel in which the liposomes are prepared (shaded part of the bars). At very high concentrations the recovery is not quantitative, as the MA tends to form clumps which float in the liposome suspension. The MA liposomes

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Figure 5.1 Titration of the maximum amount of MA that can be incorporated in liposomes consisting of egg phosphatidylcholine, with or without cholesterol: (A) liposomes consisting of egg phosphatidylcholine (99% pure) only; (B) liposomes consisting of egg phosphatidylcholine (99% pure) and cholesterol. The shaded part of the bars represent the percentage of MA recovered from the wall of the vessel in which the liposomes were prepared. Each bar represents the mean of three samples and the S.D.

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can be lyophilized and reconstituted without significant loss of MA content. Heating to 80  C in the reconstitution process is an important step to ensure full recovery of MA content.

2.4. MA liposome size is influenced by cholesterol and MA content, but not by pH MA-containing liposomes are prepared with equal amounts of 5-BMF labeled and unlabeled MAs and varying amounts of cholesterol according to Table 5.1. A liposome stock suspension is obtained by adding PBS (1 ml) to the dried lipid components, dissolving it on a heat block for 60 min after vortexing and sonicating the heated (80  C) mixture with a Branson Sonifier B30 Cell Disrupter at 20% duty cycle and output control of 2 for 5 min. Both the size distribution (forward scatter, FS) and the fluorescence (FL-1) of the liposomes are measured on a flow cytometer (a Beckman Coulter Epics Ultra instrument is used to obtain the results in Fig. 5.2). For the analysis, 10 times dilutions of the stock liposomes are prepared using PBS in plastic analyzing tubes covered with foil to prevent the photobleaching of the fluorescent marker. The tubes are kept in a water bath at 37  C during the experiments. One gate is set on the analyzer to exclude background signals and debris events that have FS values below 10. A constant number of 50,000 events are counted per measurement. Flowset (3.6 mm) fluorospheres measured with each experiment is used to correlate the relative size of the liposomes to that of the beads when varying amounts of cholesterol are added to a constant amount of MA and PC. The size distribution of the liposomes changes in accordance with a change in the cholesterol concentration (Fig. 5.2A). Two different liposome states can be identified. At the two highest concentrations of cholesterol, the liposomes are 2 log units bigger than the liposomes that contain none or the lowest concentrations of cholesterol. The liposomes obtained with the addition of 11.25 ml of cholesterol stock give an intermittent size of liposomes in between that of the two states. With an increase in the cholesterol concentration and liposome size, the liposomes also show an analogous increase of 2 log units in the amount of fluorescence per liposome that is Table 5.1 Liposome compositions with various concentrations of cholesterol

Cholesterol (ml from 100 mg/ml stock) PC (ml from 100 mg/ml stock) 5-BMF–MA (mg) Unlabeled MA (mg)

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Figure 5.2 Size (A) and fluorescence (B) of MA-containing liposomes containing various amounts of cholesterol according to Table 5.1 as determined with flow fluorometry. Flowset beads (3.6 mm) were used as a marker for the size distribution of the liposomes.

emitted (Fig. 5.2B), indicating that the change from smaller to larger liposomes is not due to swelling with water/PBS, but to an accumulation of more MA-containing material into the bigger liposomes. The results imply that cholesterol content determines two states of liposomes, where higher concentrations of cholesterol induce the disordered state of the bilayer in liposomes that increase their size 100-fold. All cholesterol-containing liposomes used in the biosensor analysis were in the disordered state at the highest concentration of cholesterol shown here.

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In Fig. 5.2, the biggest sized liposomes measured around 30 mm and were composed of PC (9 parts), cholesterol (4.5 parts), and MA (0.25 parts). When the phosphatidyl choline is combined with either the MA or the cholesterol, significant changes in the sizes of the liposomes occur. With cholesterol alone the size sharply increases, while with MA alone the size sharply decreases (Fig. 5.3). This effect is not significantly influenced in the pH range between 4 and 10 of the PBS to which the liposomes are exposed, probably due to the unchanged surface charge of phosphatidyl choline liposomes over these pH values. The anionic phosphatidyl residues (pKa  3.5) only become neutralized below pH 4, while the cationic state of choline (pKa ¼ 13.9) is maintained over the full pH range measured (Tatulian, 1993). It is therefore expected that the stability of the liposomes described here for use in the biosensor will be quite tolerant to a broad pH window extending to both sides of the typical pH of 7.4 of PBS.

2.5. MA liposome immobilization on IAsys biosensor cuvettes A number of application notes are available for the adsorption of liposomes to hydrophobic sensor surfaces. These do not stand up as rigorous and reliable for the immobilization of MA antigen-containing liposomes aiming at antibody detection. This motivated the use of nonderivatized sensor cuvettes where the glass-like hafnium oxide surface can be made hydrophobic by treatment with the cationic detergent, cetylpyridinium chloride (CPC). CPC is well known for its use as an antiseptic agent. Its property

900 800 700

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Figure 5.3 Relative sizes of PC/cholesterol and PC/mycolic acid liposomes and a mixture of the two liposomes. All liposome types contained 9 parts of phosphatidyl choline and either 4.5 parts of cholesterol, or 0.25 parts of MA, or both.

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of binding to glass when applied at aqueous dilutions below the critical micellar concentration (i.e., <0.1 mg/ml) is long established (Westwell and Anacker, 1959), leaving a ‘‘greasy’’ surface (Hartley and Runnicles, 1938). PBS/AE buffer consists of 8.0 g NaCl, 0.2 g KCl, 0.2 g KH2PO4, and 1.05 g Na2HPO4 per liter of double distilled, deionized water containing 1 mM EDTA and 0.025% (m/v) sodium azide and is adjusted to pH 7.4. CPC (0.02 mg/ml) and saponin (1 mg/ml) are prepared in PBS/AE. The IAsys resonant mirror biosensor system and twin-cell nonderivatized cuvettes are from Affinity Sensors (Farfield Scientific, Crewe, UK). The sensor is set for a data sampling interval of 0.4 s, temperature of 25  C, and stirring rate of 75% for all experiments. The cells are rinsed three times prior to use with 96% ethanol (Saarchem, South Africa), followed by extensive washing with PBS/AE. A 60 ml volume of PBS/AE is pipetted into each cell of the cuvette to obtain a stable baseline for 1 min. Cuvettes are washed with PBS/AE until a stable baseline could be maintained for at least 5 min. The cells are aspirated and 50 ml of a 20 mg/ml CPC in PBS/AE solution is added. After 10 min the cells are washed five times with 60 ml PBS/AE. PBS/AE (25 ml) is added until a stable baseline is achieved. Twenty-five microliters of the desired liposome solution are added and the binding response is monitored for 10 min. The MA-containing liposome concentration consisting of 9:3:1 (m/m) phosphatidyl choline:cholesterol:MA is titrated between 0.02 and 10 mg/ml of total lipid concentration. The MA-containing liposome concentration with the optimum binding capacity is found to be 500 mg total lipid/ml (Fig. 5.4). This concentration is used in all subsequent experiments.

MA-liposome binding (arc. s)

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Figure 5.4 Titration of the optimal concentration of MA/cholesterol-containing liposomes for immobilization on a nonderivatized IAsys biosensor cuvette.

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After liposome binding, the cells are washed five times with 60 ml PBS/AE and immediately after that, treated five times with 60 ml 1 mg/ml saponin in PBS/AE. Due to response differences among saponin batches, it is necessary to titrate the amount of saponin required each time a new batch of saponin is used. The cells are incubated with saponin for at least 10 min and until a stable baseline is achieved before a final five times wash with 60 ml PBS/AE. Antibody interaction analysis could be continued from this point as described by Thanyani et al. (2008). The cells are washed five times with PBS/AE, the content of each cell substituted with 25 ml of PBS/AE and left for about 5–10 min to achieve a stable baseline. Inhibition studies are performed using patient’s serum that is first placed at room temperature to thaw completely. After obtaining a stable baseline, a 1/1000 dilution of serum antibodies (10 ml) in PBS/AE is added in each cell, to compare the responses of the two cells over 10 min. A preincubation of 1/500 dilutions of serum with solutions of liposomes-containing MAs and empty liposomes (PC alone) is allowed for 20 min. These are then added (10 ml) for binding inhibition studies in the two cuvette cells, one with MAs liposomes and the other with empty liposomes as a control, and allowed to bind for 10 min. Finally, dissociation of antibodies is effected with three times PBS/AE washing and measurement of the response for 5 min. Regeneration of the cuvette is effected by three washes with 96% ethanol for 1 min, followed by seven washes with 70 ml PBS/AE for 1 min. The surface is then finally treated with 50 ml potassium hydroxide (12.5 M) for 2 min followed by seven washes with 70 ml PBS/AE for 1 min. A typical profile of a positive and negative TB test using this method, but applied in the ESPRIT biosensor, is shown in Fig. 5.11. The outcome of the test in the IAsys biosensor scored an overall specificity of 48.4% (15/31) and sensitivity of 86.7% (26/30) with 61 patient sera analyzed (Thanyani et al., 2008). When adjusted for the inadequate performance of the reference culture test in HIV-infected patients, the MARTI-test scored a specificity of 76.9%. A disadvantage of the IAsys biosensor in this application is the difficulty of aligning the relative basic signal strength of the two cells in a cuvette, which must be identical during the first serum exposure before a patient diagnosis can be derived from the difference of the binding signal during the second liposome-preincubated serum exposure. We experienced an average success rate of around 30% to obtain comparable signals during the first serum exposure. For this reason and due to the fact that the IAsys cuvettes were relatively expensive, we transferred the technology from the wave-guide IAsys biosensor to the more generally used SPR biosensors.

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2.6. Technology transfer from the IAsys waveguide to the ESPRIT SPR biosensor An ESPRIT SPR biosensor (Eco Chemie B.V., Utrecht, The Netherlands) is used in this study to detect antibodies to MA in human patient sera. The principle of the SPR biosensor is based on the change in the refractive index on a thin gold film surface modified with various materials (Lee et al., 2005) to indicate the binding of ligands, in this case anti-MA antibodies. Both IAsys and ESPRIT biosensors use a cuvette system. The light is totally internally reflected from the sensing surface by means of a prism in both biosensors. SPR signals are related to the refractive index close to the sensor surface, and therefore report the amount of macromolecules bound to the sensor surface. An SPR immunosensor is composed of several important components such as a light source, detector, prism with transducer surface and flow system (Shankaran et al., 2007). The transducer surface is usually a gold film (50–100 nm, on which biomolecules, such as antibody or antigen, are immobilized) on a glass slide optically coupled to the glass prism through refractive index matching oil. The resonance conditions are influenced by the biomolecules interacting with their immobilized ligands on the gold layer. When the molecules interact, the change in the interfacial refractive index can be detected as a shift in the resonance angle. These changes are monitored over time and converted into a sensorgram, from which the kinetics and affinity constants of the interaction can be determined. There has been considerable progress in the development of new methods of immobilizing biological recognition elements onto transducer sensor surfaces (Zhang et al., 2000), a key step in the development of biosensors. The use of self-assembled mono- and multilayers (SAMs) is increasing rapidly in various fields of research, and this applies especially to the construction of biosensors (Zhang et al., 2000, 2008). The uncomplicated procedure for SAM formation and compatibility with metal substrates such as gold for electrochemical measurements enable special benefits for biosensor applications. The term self-assembly involves the spontaneous arrangement of atoms and molecules into an ordered stable form or even aggregate of functional entities (Tecilla et al., 1990). For example, the highly ordered and dense nature of the long-chain alkanethiols of SAMs mimic the cellular microenvironment of lipid bilayer structures, thereby providing novel substrates for immobilized biomolecules (Arya et al., 2006). The molecular self-assembly of long-chain alkanethiol on gold has drawn considerable attention during the past decade, since self-assembled monolayers (SAMs) have strong adhesion to a substrate, high degree of thermal and chemical stability, and mechanical strength (Kim et al., 2001). The stability of the SAMs of the alkanethiol molecules formed on the gold depends on the strength of Au–S bond and the Van der Waals force between a thiol molecule and its surrounding molecules (Han et al., 2004).

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SAMs can be used as interface layers upon which almost all types of biological components, including proteins, enzymes, antibodies, and their receptors can be loaded (Zhang et al., 2000). Here, the preparation of octadecanethiol (ODT) in absolute ethanol to form a SAM is described. It is characterized by cyclic voltammetry (CV) and applied for the measurement of binding, or inhibition of binding of patient serum antibodies to MAs that are immobilized in liposomes onto the alkanethiol coated ESPRIT biosensor surface. The low solubility of ODT in ethanol is preferred to form the SAMs. Kim et al. (2001) demonstrated that the adsorption rate of alkanethiol onto clean gold when using a quartz crystal microbalance (QCM) biosensor depends on the thiol concentration, temperature, and solvent used. Here, a full coverage of the underivatized Au surface is observed when 10 mM of ODT is used. This is confirmed by a strongly hindered redox reaction when the surface is characterized with a CV instrument. CV experiments are carried out using an Autolab potentiostat PGSTAT 30 from Eco Chemie (Utrecht, The Netherlands) driven by the General Purpose Electrochemical Systems (GPES) data processing software version 4.9. Sodium dodecylsulphate (SDS) and absolute ethanol (analytical grade) were obtained from Merck (Gauteng, SA). ODT, ferricyanide [K3Fe (CN)6], ferrocyanide [K4Fe(CN)6], potassium chloride (KCl), and urea, all analytical grade, were obtained from Sigma-Aldrich (St. Louis, MO). Acetic acid (analytical grade), sodium hydrogen carbonate (NaHCO3), isopropanol (chemically pure), and sodium hydroxide (NaOH) were obtained from Saarchem (Gauteng, SA). ODT (10 mM) is dissolved in absolute ethanol using a water bath sonifier (Ultrasonic Cleaner, Optima Scientific CC, Model: DC150H) for 30 min. Sodium hydrogen carbonate (0.2 M), SDS (0.5%, w/v), sodium hydroxide (50 mM), 1 mM ferrocyanide/ferricyanide, 1 M KCl, and urea (6 M ) are prepared with sterile double distilled water. The coating of the gold disk cannot be monitored in real time, since the ODT is dissolved in absolute ethanol that generates too large refractive index jumps in the sensor signals when alternated with PBS/AE. The underivatized gold disk is incubated for 16 h at room temperature in 10 mM ODT. The ODT-coated gold disk is immersed in a solution of 1 mM ferrocyanide/ferricyanide (used as a redox probe) containing 1 M KCl and scanned at a rate of 25 and 50 mV/s at a potential window of
0.1 to 0.5 V (vs. Ag/AgCl, saturated KCl). The results in Fig. 5.5 show that there is a significant drop in the current response of the ODT-coated disk toward the redox probe in comparison to the uncoated gold disk, indicating the formation of a stable SAM of ODT by S–Au bonds on the surface of the gold disk. The SAM can be maintained after the exposure of the coated surface to several regeneration cycles of absolute ethanol and a mixture of 50 mM NaOH with isopropanol (2:3, v/v).

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Underivatised Au ODT coated Fourth regeneration

4.00E-04 3.00E-04 2.00E-04

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Figure 5.5 Testing of the octadecanethiol-coated ESPRIT biosensor gold surface against sequential times of regeneration with a mixture of isopropanol and 50 mM NaOH (2:3, v/v) using cyclic voltammetry. The voltammograms for the different surfaces are shaded to better identify the one from the other.

Kim et al. (2001) reported that partial ODT multilayers on the gold surface could be formed via the formation of disulfides, since thiols are oxidized to disulfides in the presence of oxygen and the solubility of disulfides in ethanol is much less than that of thiols. If a solution of ODT in ethanol is exposed to oxygen and oxidized to disulfide, the oxidized disulfide can be precipitated onto the monolayer. Here, the solution of ODT in absolute ethanol is covered with parafilm to avoid oxygen exposure. The ODT-coated disk is subsequently inserted into the biosensor on a droplet of special refractive index oil, after wiping the glass bottom surface with lens tissue. The PBS/AE is filtered through a 0.2 mm particle retention membrane and degassed with helium for 30 min before use. Degassing is required whenever a hydrophobic surface is created that is exposed to air before liposome addition to prevent the formation of a ‘‘dissolved’’ air layer onto the hydrophobic surface that prevents SPR to occur. The cuvette is flushed with 500 ml ethanol (96%) using the automatic dispenser with simultaneous draining, followed by brief (60 s) flow-washing with PBS/AE. An automated software program sequence can be created to control the addition of all the samples and liquids into the cuvette. Quality of the surfaces are monitored by determining the SPR dips after cleaning the Au ODT-coated surface with 96% ethanol and a mixture of isopropanol and 50 mM sodium hydroxide (2:3, v/v). The samples are transferred from a

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384 multiwell plate (Bibby Sterilin Ltd, Stone, UK) to the cuvette surface by an autopipettor. First, the baseline of the ESPRIT biosensor is set with 10 ml PBS/AE, followed by addition of 50 ml MA liposomes on the disk for 20 min. The immobilized liposomes are then finally washed five times with 100 ml PBS/AE, to prepare for blocking the surface with saponin. 2.6.1. Use of degassed buffers after liposome coating of the sensor surface A recent study (Eastoe and Ellis, 2007) showed that exposure of lipids to degassed buffers resulted in a detergent effect that destabilized the lipids. The removal of hydrophobic gas by pumping created a limited ability of the degassed water or aqueous buffer to dissolve lipids. To determine whether this will also be the case here where buffers are degassed by bubbling through helium gas, we exposed a liposome-coated surface repeatedly to either degassed or nondegassed buffer. The liposomes-containing MAs are immobilized on ODT-coated gold sensor disks for 20 min. The liposomes are washed five times with degassed or nondegassed PBS/AE, and left for 5 min with mixing to achieve a baseline. This procedure is repeated three times. Figure 5.6A demonstrates how the baseline is affected during movement of degassed PBS/AE over the liposome coat; compared to when buffer is used that is not degassed (Fig. 5.6B). A stable baseline is obtained only when a nondegassed PBS/AE is used. The rest of the procedure in the MARTI-assay that follows after liposome coating is subsequently done with buffer that is not degassed, taking special care that air bubbles do not develop in the fluid lines that could affect the working of the pumps. In several attempts where we test samples with continued use of degassed buffer after liposome coating, we obtain nonreliable results, with the liposome layer often coming apart at the final step of antibody incubation. 2.6.2. Optimization of saponin concentration Different concentrations of saponin (%, m/v) prepared in PBS/AE (0.1%, 0.05%, 0.025%, 0.0125%, and 0.00625%) are tested to block the hydrophobic sites of the MA liposome layer. The stock saponin concentration is 0.1% and the subsequent dilutions are prepared from this stock solution. From the results obtained (Fig. 5.7A), there is a tendency of an increase in saponin accumulation onto MA liposomes immobilized on an ODT-coated gold surface, as the saponin concentration is increased from 0.00625% to 0.05%. At a saponin concentration of 0.05% there is an amount of net saponin accumulation after PBS/AE buffer wash (Fig. 5.7B). An unstable baseline is also obtained when 0.05% saponin is used. A saponin concentration of 0.0125% is chosen as optimal, because it gives a stable baseline and acceptable variation after PBS/AE wash (Fig. 5.7B) as compared to 0.00625% and 0.025%. The differences in optimal saponin concentration used on the IAsys (0.03%) and current ESPRIT biosensors

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Figure 5.6 Effect of degassed (A) and nondegassed (B) buffer on immobilized mycolic acids liposomes in the ESPRIT biosensor. The arrows indicate where washing cycles with PBS/AE were introduced before allowing a baseline to be reached with mixing before substitution of cell content.

(0.0125%) could be due to different batches of saponin, or that the CPC and ODT activation before immobilization of the MA produces different surface properties. 2.6.3. Optimization of first serum exposure dilution in PBS/AE After optimization of saponin concentration, the next step is to determine which concentration of serum is optimal for the MARTI-assay in the first exposure to antigen. Chung et al. (2005) indicated that serum should be diluted to minimize the nonspecific binding to the biosensor surface. Serum is a complicated protein mixture for direct application to a biosensor surface. The introduction of a first serum exposure at high dilution was previously done on the IAsys biosensor to provide a practical working dilution that did not fully saturate the antigen coat, but was still concentrated enough to give

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Saponin concentration (m/v, %) Figure 5.7 Optimization of saponin concentration to avoid nonspecific binding on immobilized mycolic acids on the Au surface coated with octadecanethiol. Accumulation of saponin is performed for 5 min (A) and washed with PBS/AE (B). The error bars indicate the standard error of the mean (S.E.M.) and n ¼ 3.

a measurable signal to probe the comparability of the binding signals from the cuvette. This simultaneously blocked off the major nonspecific binding areas and hugely increased the accuracy of the MARTI-test.

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Serum samples collected for another study (Schleicher et al., 2002) were used that were obtained from 61 adult patients (aged between 18 and 65 years), who were admitted to the general medical wards of the Helen Joseph Hospital, Johannesburg, South Africa, including a number with active pulmonary TB. The TB-negative patients that were used as controls had medical conditions other than TB and were recruited from the general medical wards. The liposomes are immobilized as described earlier, the surface blocked with 0.0125% saponin, after which 50 ml of PBS/AE is added and left for 5 min to affect a stable baseline. This is followed by addition of 35 ml of either 1/500, 1/1000, 1/2000, or 1/4000 dilutions of serum in PBS/AE. For the assessment of the optimal dilution of the first serum exposure, a second exposure of serum preincubated in MAs-containing or empty liposomes is kept constant at 1/250 in all the experiments. The results show that the chosen serum dilution range of 1/4000–1/500 responded in an almost linear positive correlation between antibody-binding signal and serum concentration with a slight running out at 1/4000 that indicates that the lower limit of the serum concentration is reached. The results obtained in Fig. 5.8 gave a positive linear correlation with a coefficient (r2) of 0.9749 between the serum concentrations and their signal binding response over the range measured, which is a requirement for a successful MARTI-assay.

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Figure 5.8 Optimization of the dilution of serum (P135) for the first exposure to antigen in the MARTI-assay, after 0.0125% saponin blocking of the mycolic acid liposome coat of the ESPRIT biosensor. The error bars indicate the standard deviation. Correlation coefficient (r2) ¼ 0.9749, n  3.

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The 1/4000 and 1/2000 dilutions are adequate for the first serum exposure, leaving enough room for a positive binding event at the second serum exposure. 2.6.4. Second serum exposure with liposome preincubation P129 (TB-positive) serum was used to optimize the dilution of the second exposure to preincubated serum in MA-containing, or empty liposomes for inhibition studies, following on a first serum exposure to immobilized antigen at a dilution of either 1/4000 or 1/2000. The first exposure should avoid the saturation of antigen with antibody before the addition of preincubated serum. Different dilutions (1/250, 1/500, 1/1000, and 1/2000) of preincubated serum in MA-containing and empty liposomes are applied by 35 ml addition to either 1/4000 or 1/2000 of first serum exposure in PBS/AE, after 10 min of incubation. This is followed by washing away of the unbound antibody with five times 100 ml PBS/AE. The TB-positive patient P129 serum showed a significant decrease of signal when the serum was preincubated in MA-containing liposomes, compared to empty liposomes over a range of 1/250, 1/500, or 1/1000 dilution (Fig. 5.9) after a first serum exposure at 1/4000. There was no inhibition of antibody by MA preincubation observed when 1/2000 dilution of serum was used and binding response signals were also too low.

Binding inhibition response (m⬚)

140 PC

MA

120 100 80 60 40 20 0 1/250

1/500 1/1000 Serum dilution in liposomes

1/2000

Figure 5.9 MARTI antibody-binding inhibition response of preincubated TB-positive P129 serum dilutions inhibited with mycolic acids (MAs)-containing and empty (PC) liposomes after first serum exposure of 1/4000 on immobilized MAs. The error bars indicate the standard deviation. P129 showed significant MA inhibition signals at 1/250, 1/500, and 1/1000 serum dilutions, with P-values of 0.00014, 0.01411, and 0.0393, respectively, but no significant inhibition at 1/2000 serum dilution (P-value of 0.7857). A 95% (0.05) confidence limit was used, n ¼ 3.

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Binding inhibition response (m⬚)

120 PC

MA

100 80 60 40 20 0 1/250

1/500 1/1000 Serum dilution in liposomes

Figure 5.10 MARTI-binding inhibition response of various dilutions of preincubated TB-positive patient serum (P129) with mycolic acids (MA)-containing and empty (PC) liposomes after first exposure serum dilution of 1/2000 to surface immobilized MAs. The error bars indicate the standard deviation. No statistical difference (at 95% confidence limit) is obtained at 1/250 and 1/500 dilutions between MA- and PC-inhibited serum, with P-values of 0.116 and 0.356, respectively, while a significant inhibition was observed at 1/1000 (P-value of 0.0086) n ¼ 3.

This shows that the lower limit of serum concentration was reached at 1/2000 dilution to measure the inhibition of anti-MA antibody binding. The results in Fig. 5.10 indicate that inhibition values of 17%, 19%, and 41% were obtained at 1/250, 1/500, and 1/1000 dilutions of serum in liposome solution, respectively, after first serum exposure at 1/2000 dilution. At first sight, it appeared that a better value was obtained by using a first serum exposure of 1/2000 dilution, followed by a second, antigen preincubated serum dilution at 1/1000 dilution (numerical difference: 12.50 millidegrees). However, when looking at the numerical signal difference between MA- and empty liposome-inhibited serum, then the 1/4000 dilution of first serum exposure followed by second serum exposure at 1/500 still gave the best value (numerical difference: 21.53 millidegrees). In addition, the significance of the difference between antibody-binding inhibition with MA liposomes and empty liposomes was significant at 1/250, 1/500, and 1/1000 dilution of serum after first serum exposure at 1/4000 dilution, while only the 1/1000 dilution of inhibited serum produced a significant difference after a first serum exposure of 1/2000 dilution (P-value limit of 0.05). The 1/2000 dilution of first serum exposure appears, therefore, to restrict the workable range of serum dilutions at the second critical serum exposure that provides the inhibition end result. This was confirmed when another TB-positive—HIV-negative serum (P96) was tested and for which a better inhibition response was obtained at the

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preferred serum dilutions of 1/4000 and 1/500 for first and second serum exposures, respectively, compared to the result obtained with first exposure at 1/2000. At the preferred serum dilutions of exposure, the TB-negative— HIV-negative serum P94 gave the expected zero inhibition value, with a P-value of 0.9863. 2.6.5. Regeneration of the ODT-coated gold disks After dissociation of the unbound serum antibodies to MAs, the surface is regenerated with 100 ml mixture of isopropanol and 50 mM NaOH (2:3, v/v) for 2 min and finally washed with 100 ml of 99% (absolute) ethanol. The surface is washed five times with 100 ml of PBS/AE after each regeneration step to prepare it for a next round of liposome coating on the stable ODT layer. 2.6.6. Cleaning of cuvette and needles A flow wash sequence is used to clean the needles after analyzing approximately 30 sample runs. Sequential washes with 0.5% (w/v) SDS, 6 M urea, 1% (v/v) acetic acid, 0.2 M sodium hydrogen carbonate (NaHCO2), and ddd H2O are done in order to maintain the quality of the SPR signals during repeated measurements. 2.6.7. The optimized MARTI-assay With the lesson learnt of avoiding degassed buffers after liposome coating and the conditions optimized for the blocking of the liposome layer with saponin, titrations of the optimal dilutions for first exposures to serum and second exposure to antigen-inhibited serum dilutions were done. It is concluded that best results were obtained with 1:4000 dilution of serum at first exposure and 1:500 dilution of serum at second exposure. In the second exposure, the serum is preincubated with antigen in order to effect an inhibition of binding signal, as graphically demonstrated in Fig. 5.11. The SPR dips (Fig. 5.11 inserts) between 0% and 10% reflectivity that are associated with the binding profiles proved that the sensor surfaces remained intact and fully activated during the run of the experiments. Using this optimized protocol, four serum samples were selected from the Schleicher et al. (2002) collection and assessed for the presence of anti-MA antibodies. In Table 5.2, the MARTI-assay results are presented and compared with that obtained on ELISA by Schleicher et al. (2002). From Table 5.2, P129 and P96 tested false negative on ELISA and true positive on ESPRIT biosensor, while P94 tested equivocally on ELISA and true negative on ESPRIT biosensor, as it was previously shown on the IAsys biosensor (Thanyani et al., 2008). The MARTI-assay on IAsys biosensor was successfully validated to an accuracy of 82% for the serodiagnosis of active pulmonary TB. The IAsys biosensor system applied to the

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A 400

Angle (m⬚)

300

d b

c

Reflectivity [%] 100

200

PC MA

80 60 40

100

30 -2000

a

0

0

1000

2000 Time (s)

0 2000 Angle [m⬚]

3000

4000

B 400

Angle (m⬚)

300 d

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b

Reflectivity [%] 100

c

PC MA

80 60 40

100

30 -4000 -2000 0 2000 4000 Angle [m⬚]

a

0

0

1000

2000 Time (s)

3000

4000

Figure 5.11 Typical sensorgrams summarizing the process of measuring serum antibody (A ¼ TB-positive P129 and B ¼ TB-negative P94) binding, or inhibition of binding by mycolic acid-containing and empty liposomes, on an ESPRIT biosensor with ODT-coated gold surface and immobilized mycolic acid liposomes. Mycolic acids liposomes were immobilized on the ESPRIT biosensor surface (a), blocked with saponin (b), calibrated with a 1/4000 first exposure of serum (c), and applied to measure the binding and dissociation of 1/500 diluted sera inhibited with empty (thick line) or mycolic acid-containing (thin line) liposomes at lesser dilution (d). The arrows indicate washing with PBS/AE. Table 5.2 MARTI (ESPRIT biosensor) and ELISA analysis compared for their ability to detect antibody to MA in three selected human sera

a b

Patient no.

TB/HIV status

ELISA assaya

P129 P96 P94

TBþHIV
TBþHIV
TBþHIV


1.59 1.05 1.69

MARTI-assayb

25.09 29.53
0.23

Signal to background value of absorbance at 450 nm. Values higher than 2 are taken as positive. Percentage inhibition of antibody binding to MA liposomes. Values higher than 20% are taken as positive.

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MARTI-test has a weakness in that the channels often do not give matching results, while the cuvettes are 10 times more expensive than the gold disks provided for the ESPRIT biosensor. The ESPRIT biosensor is provided with an adjustable laser setting to compensate for differences in the channel readings as well as an automated pipettor system that reduces variance from one sample to the next. The MARTI-assay as applied in the ESPRIT biosensor has now reached the stage where a result of sample analysis can be guaranteed within 4 h of receipt of the serum. This is the first time that such reliability has been achieved. However, more sera need to be analyzed to confirm the reproducibility of the assay among the HIV-positive population, to prove the value of the MARTI-test against the many studies reported of low sensitivity and specificity with HIV-positive samples using standard techniques of TB diagnosis (Antunes et al., 2002; Hendrickson et al., 2000; Schleicher et al., 2002).

2.7. The cholesteroid nature of MA demonstrated on the ESPRIT biosensor In our previous studies, we have provided evidence for a structural relationship and attraction between free MAs and cholesterol (Benadie et al., 2008). This was supported by demonstrating the interaction between MA and Amphotericin B—an antifungal macrolide agent known to bind to cholesterol (Baginski et al., 2002)—on the IAsys biosensor system. The same principle was confirmed with the ESPRIT biosensor as demonstrated below, in an attempt to determine what the effect of labeling of MA would be on its manifestation of cholesteroid nature. For the preparation of the different liposomes, phosphatidyl choline stock solution (90 ml, 100 mg/ml chloroform) is added to an amber glass vial containing either MA (1 mg) or an equimolar amount of 5-BMF labeled MA (1.35 mg). For the preparation of cholesterol-containing liposomes, phosphatidyl choline stock solution (60 ml, 100 mg/ml chloroform) is added to a cholesterol solution (30 ml, 100 mg/ml chloroform). The samples are mixed well until dissolved, then dried under a stream of N2 gas at 85  C. Saline (2 ml) is then added and the sample is heated on a heat block for 20 min at 85  C. The sample is then vortexed for 1 min, sonicated with a Virsonic probe sonicator until a clear solution forms to indicate vesicle formation, aliquoted at 0.2 ml per vial, lyophilized and stored at
70  C until use. Before use, lyophilized liposomes are reconstituted with buffer (2 ml). The liposomes are placed on a heat block for 30 min at 85  C, vortexed for 2 min and sonified. The final liposome concentration comes to 500 mg lipid/ml.

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The binding interaction between Amphotericin B and either cholesterol-, MA-, or 5-BMF labeled MA-containing immobilized liposomes are tested here. An ODT-coated gold disk is mounted in the ESPRIT instrument and the individual liposomes immobilized as described before. The instrument is operated and reagents used at RT. The liposomes are washed five times with nondegassed PBS/AE, and left for 5 min with mixing to obtain a baseline. Amphotericin B (1  10
4 M ) is added to the liposome layer and the direct interaction is recorded for 10 min after which the disk is washed five times with nondegassed PBS/AE, and left for 5 min. The results (Fig. 5.12) confirm the ability of the ESPRIT biosensor to demonstrate that Amphotericin B recognizes both cholesterol and MA, as was previously shown with the IAsys biosensor. In addition, it shows for the first time the intolerance of the system for fettering with the structure of MA by adding a bulky label on its carboxylic acid group.

Relative binding

120

80

40

70 60 50 40 30 20 10 0

Cholesterol

MA

Chol MA

Angle (m⬚)

Angle (m⬚)

0

5BMF-MA 70 60 50 40 30 20

Chol 5BMF-MA

10 0

5

10 15 Time (s)

20

25

0

0

5

10 15 Time (s)

20

25

Figure 5.12 Normalized AmB-binding capacity on immobilized lipid antigens cholesterol, MA, and 5BMF–MA and binding curves of MA versus cholesterol and 5BMF–MA versus cholesterol. The error bars indicate the standard deviation, n ¼ 3 for each set.

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3. Conclusion Antibodies to MA in serum as surrogate markers of active TB can be detected by making use of liposomes as MA antigen carriers in a wave-guide evanescent field biosensor (Thanyani et al., 2008). However, SPR biosensors are more generally in use, more amenable to high-throughput screening, have lower running costs, and provide more comparable binding signals in twin-cell cuvettes. The more preferred technique of SPR biosensor technology, using standard methods to form a hydrophobic surface onto the gold disks on which the MA-containing liposomes can be immobilized, is reported here. This demonstrates the use of liposomes in biosensors to detect large biomolecules such as antibodies. By using Amphotericin B as a small ligand to bind cholesterol and MA in a wave-guide biosensor the cross-reactivity of antibodies to MAs in ELISA can be explained (Benadie et al., 2008). This can also be demonstrated using an SPR biosensor. The biosensor antibody detection approach (MARTI—TB serodiagnostic test) remains unaffected by the antibody cross-reactivity between MA and cholesterol.

ACKNOWLEDGMENTS This research was supported by a Marie-Curie Early Stage Training Fellowship (Molfun) to Y. Lemmer, the Cancer and Polio Research Fund and the North West Cancer Research Fund, Adcock Ingram Limited (South Africa), the National Research Foundation of South Africa (Technology and Human Resources for Industry Programme), the European and Developing Countries Clinical Trial Partnership Grant No. 2004.1.R.d1, and the Medical Research Council of South Africa. Dr. ten Bokum held a Claude Leon Harris Foundation postdoctoral fellowship.

REFERENCES Antunes, A., Nina, J., and David, S. (2002). Serological screening for tuberculosis in the community: An evaluation of the Mycodot procedure in an African population with high HIV-2 prevalence (Republic of Guinea-Bissau). Res. Microbiol. 153, 301–305. Arya, S. K., Solanki, P. R., Singh, R. P., Pandey, M. K., Datta, M., and Malhotra, B. D. (2006). Application of octadecanethiol self-assembled monolayer to cholesterol biosensor based on surface plasmon resonance technique. Talanta 69, 918–926. Baginski, M., Resat, H., and Borowski, E. (2002). Comparative molecular dynamics simulations of amphotericin B-cholesterol/ergosterol membrane channels. Biochim. Biophys. Acta 1567, 63–78. Bangham, A. D. (1983). Liposomes: An Historical Perspective. Marcel Dekker, New York. Beatty, W. L., Rhoades, E. R., Ullrich, H. J., Chatterjee, D., Heuser, J. E., and Russell, D. G. (2000). Trafficking and release of mycobacterial lipids from infected macrophages. Traffic 1, 235–247.

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Beckman, E. V., Porcelli, S. A., Morita, C. T., Behar, S. M., Furlong, S. T., and Brenner, M. B. (1994). Recognition of a lipid antigen by CD1-restricted ab þ T cells. Nature 372, 691–694. Benadie, Y., Deysel, M., Siko, D. G., Roberts, V. V., Van Wyngaardt, S., Thanyani, S. T., Sekanka, G., Ten Bokum, A. M., Collett, L. A., Grooten, J., Baird, M. S., and Verschoor, J. A. (2008). Cholesteroid nature of free mycolic acids from M. tuberculosis. Chem. Phys. Lipids 152, 95–103. Butler, W. R., and Kilburn, J. O. (1990). High-performance liquid chromatography patterns of mycolic acids as criteria for identification of Mycobacterium chelonae, Mycobacterium fortuitum, and Mycobacterium smegmatis. J. Clin. Microbiol. 28, 2094–2098. Chan, E. D., Reves, R., Belisie, J. T., Brennan, P. J., and Hahn, W. E. (2000). Diagnosis of tuberculosis by a visually detectable immunoassay for lipoarabinomannan. Respir. Crit. Care Med. 161, 1713–1719. Chung, J. W., Kim, D. S., Bernhardt, R., and Pyun, J. C. (2005). Application of SPR biosensor for medical diagnostics of human hepatitis B virus (hHBV). Sens. Actuator B 111, 416–422. Dennison, C., and Lovrein, R. (1997). Three-phase partitioning: Concentration and purification of proteins. Protein Exp. Purif. 11, 149–161. Eastoe, J., and Ellis, C. (2007). De-gassed water and surfactant-free emulsions: History, controversy, and possible applications. Adv. Colliod Interface Sci. 134, 89–95. Goodrum, M. A., Siko, D. G. R., Niehues, T., Eichelbauer, D., and Verschoor, J. A. (2001). Mycolic acids from Mycobacterium tuberculosis: Purification by countercurrent distribution and T cell stimulation. Microbios 106, 55–67. Han, J., Wang, X., and Kwok, D. L. (2004). Structure and stability of self-assembled monolayer for octadecanethiol adsorbed on flame annealing gold substrate and its potential application to microfluidics. ICMENS 4, 1–4. Hartley, G. S., and Runnicles, D. F. (1938). The determination of the size of paraffin-chain salt micelles from diffusion measurements. Proc. Roy. Soc. London: Math. Phys. Sci. 168, 420–440. Hendrickson, R. C., Douglass, J. F., Reynold, L. D., McNeill, P. D., Carter, D., Reed, S. G., and Houghton, R. (2000). Mass spectrometric identification of Mtb81, a novel serological marker for tuberculosis. J. Clin. Micribiol. 38, 2354–2361. Houghton, R. L., Lodes, M. J., Dillon, D. C., Reynolds, L. D., Day, C. H., McNeill, P. D., Hendrickson, R. C., Skeiky, Y. A. W., Sampaio, D. P., Badaro, R., Lyashchenko, K. P., and Reed, S. G. (2002). Use of multiepitope polyproteins in serodiagnosis of active tuberculosis. Clin. Diagn. Lab. Immunol. 9, 883–891. Kim, H. D., Noh, J., Hara, M., and Lee, H. (2001). An adsorption process study on the selfassembled monolayer formation of octadecanethiol chemisorbed on gold surface. Bull. Korean Chem. Soc. 22, 276–280. Korf, J., Stoltz, A., Verschoor, J. A., De Baetselier, P., and Grooten, J. (2005). The Mycobacterium tuberculosis cell wall component mycolic acid elicits pathogen-associated host innate immune responses. Eur. J. Immunol. 35, 890–900. Korf, J. E., Pynaert, G., Tournoy, K., Boonefaes, T., van Oosterhout, A., Ginneberge, D., Haegeman, A., Verschoor, J. A., De Baetseller, P., and Grooten, J. (2006). Macrophage reprogramming by mycolic acids promotes a tolerogenic response in experimental asthma. Am. J. Respir. Crit. Care Med. 174, 152–160. Lee, W. J., Sim, J. S., Cho, M. S., and Lee, J. (2005). Characterization of self-assembled monolayer of thiol on a gold surface and the fabrication of a biosensor chip based on surface plasmon resonance for detecting anti-GAD antibody. Biosens. Bioelectron. 20, 1422–1427. Pan, J., Fujiwara, N., Oka, S., Maekura, R., Ogura, T., and Yano, L. (1999). Anti-cord factor (trehalose 6, 60 -dimycolate) IgG antibody in tuberculosis patients recognizes mycolic acid subclasses. Microbiol. Immunol. 43, 863–869.

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Reischl, U. (1996). Application of molecular biology-based methods to the diagnosis of infectious diseases. Front. Biosci. 1, 72–77. Samanich, K. M., Keen, M. A., Vissa, V. D., Harder, J. D., Spencer, J. S., Belisle, J. T., Zolla-Pazner, S., and Laal, S. (2000). Serodiagnostic potential of culture filtrate antigens of Mycobacterium tuberculosis. Clin. Diagn. Lab. Immunol. 7, 662–668. Schleicher, G. K., Feldman, C., Vermaak, Y., and Verschoor, J. A. (2002). Prevalence of anti-mycolic acid antibodies in patients with pulmonary tuberculosis co-infected with HIV. Clin. Chem. Lab. Med. 40, 882–887. Shankaran, D. R., Gobi, K. V., and Miura, N. (2007). Recent advancements in surface plasmon resonance immunosensors for detection of small molecules of biomedical, food and environmental interest. Sens. Actuators B 121, 158–177. Tatulian, S. (1993). Phospholipids Handbook. Marcel Dekker Inc., New York. Tecilla, P., Dixon, P. D., Slobodkin, G., Alavi, D. S., Waldeck, D. H., and Hamilton, A. H. (1990). Hydrogen-bonding self-assembly of multichromophore structures. J. Am. Chem. Soc. 112, 9408–9410. Thanyani, S. T., Roberts, V., Siko, D. G., Vrey, P., and Verschoor, J. A. (2008). A novel application of affinity biosensor technology to detect antibodies to mycolic acid in tuberculosis patients. J. Immunol. Methods 332, 61–72. Verschoor, J. A., and Onyebujoh, P. (1999). The menace of the AIDS-tuberculosis combo: Any solutions? Bioessays 21, 365–366. Westwell, A. E., and Anacker, E. W. (1959). Adsorption of cetylpyridinium chloride on glass. J. Phys. Chem. 63, 1022–1024. Zhang, S., Wright, G., and Yang, Y. (2000). Materials and techniques for electrochemical biosensor design and construction. Biosens. Bioelectron. 15, 273–282. Zhang, Y., Wang, H., Yan, B., Zhang, Y., Li, J., Shen, G., and Yu, R. (2008). A reusable piezoelectric immunosensor using antibody-adsorbed magnetic nanocomposite. J. Immunol. Methods 332, 103–111.

C H A P T E R

S I X

Solid Lipid Nanoparticle Formulations: Pharmacokinetic and Biopharmaceutical Aspects in Drug Delivery Eliana B. Souto*,† and Slavomira Doktorovova´* Contents 106 107 107 112 113

1. 2. 3. 4.

Introduction Production of SLN Pharmacokinetics and Pharmacodynamics Modified Release Profile 4.1. Short or relatively long half-life drugs 4.2. Targeting to a specific tissue by implantation for long-term release 5. Biopharmaceutical Aspects of Administration Routes 6. Clinical Pharmacology 6.1. Local effects and systemic distribution 6.2. Dose size 6.3. Concentration fluctuations and therapeutic range 6.4. Toxicological concentration 7. Concluding Remarks References

113 114 117 117 120 121 121 121 122

Abstract Solid lipid nanoparticles (SLNs) have emerged as important tools to modify the release profile for a large number of drugs including protein and peptide molecules. SLNs are produced from biocompatible and biodegradable lipid materials, making them a promising therapeutic strategy for drug targeting and delivery, and surmounting the inherent limitations of regulation acceptance. Due to their versatility in loading both lipophilic and hydrophilic molecules in the solid lipid matrix, SLNs depict the ability to prolong, extend or * {

Department of Pharmaceutical Technology, Faculty of Health Sciences, Fernando Pessoa University, Porto, Portugal Institute of Biotechnology and Bioengineering, Centre of Genetics and Biotechnology, University of Tra´s-os-Montes and Alto Douro (CGB-UTAD/IBB), Vila Real, Portugal

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64006-4

#

2009 Elsevier Inc. All rights reserved.

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sustain the release profile of the loaded molecules, therefore reducing the repeated administration, and increasing the therapeutic value of a certain treatment. Additional advantages include reduction of drug toxicity and increase of drug bioavailability. To develop SLN formulations for drug targeting and delivery, a basic pharmacokinetic understanding of drug distribution is of major relevance, as well as the biopharmaceutical aspects of the administration route. This chapter provides a fundamental understanding of the pharmacokinetic properties of SLNs, which influence both biopharmaceutical and clinical profiles of the loaded molecules.

1. Introduction Lipid-based drug delivery systems (LDDS) are an interwoven field of science requiring a holistic approach of several research areas, including concepts and applications of pharmaceutical technology, pharmacology, pharmacokinetics, and biopharmaceutics. Within the LDDS, solid lipid nanoparticles (SLN) have emerged as novel modified release systems for several drugs, including protein and peptide molecules (Martins et al., 2007, 2009; Mu¨ller et al., 2006, 2008). SLN are composed of a solid matrix based on very chemically different lipid molecules (e.g., waxes, mixtures of mono-, di-, and triacylglycerols, fatty acids) generally recognized as being safe. This provides the advantage of using physiological components and/or excipients accepted for oral and topical administration, reducing the risk of acute and chronic toxicity. For the production of thermodynamically stable SLN, these particles need to be stabilized by a film of surfactant molecules surrounding the lipid matrix in aqueous dispersion. Surfactants such as polysorbates, poloxamers, and phospholipids are very common in SLN formulations for several administration routes. One can have a neat delivery system, but a major obstacle for its introduction into clinical practice is the need to undertake toxicity studies, in particular for parenteral drug delivery. The status of the raw materials used for SLN production is therefore discussed as a function of the administration route. Topical and oral administration routes for SLN are usually nonproblematic. As a result of the biodegradation studies, one can expect that the degradation velocity of SLN composed of different lipids but stabilized with the same surfactant or surfactant mixture will depend on the chemical nature of the lipid matrix. On the other hand, for SLN composed of the same lipid but stabilized by different surfactants or surfactant mixtures, the degradation velocity will be mainly determined by the chemical nature of the surfactants. As a result, the degradation velocity of the lipid matrix can

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be adjusted (accelerated or delayed) by modifying the surfactant composition. Based on the differences in degradation velocity, SLN are expected to be a system with great flexibility to modify the release profiles for different drugs.

2. Production of SLN SLN are easily produced by a melt emulsification followed by a hot or cold high-pressure homogenization (HPH) process. Briefly, for the hot HPH the lipid phase is previously melted 5–10
C above its melting point, followed by drug dissolution or dispersion in the melted phase. Stirring this melted phase in a hot surfactant solution, a pre-emulsion is produced. The obtained pre-emulsion is homogenized under high pressure, producing a hot nanoemulsion, which is further cooled down, recrystallizing the lipid, and forming SLN. The cold HPH technique requires a previous step of melting the solid lipid so that the drug can be dissolved and/or admixed to this phase. Applying liquid nitrogen or dry ice, the lipid phase cools down rapidly, and solidifies and then, by means of mortar milling, it is ground, resulting in microparticles. These microparticles are further dispersed in a cold aqueous surfactant solution, producing a presuspension that is homogenized at or below room temperature using the HPH.

3. Pharmacokinetics and Pharmacodynamics For the majority of the modified release LDDS, a basic pharmacokinetic understanding of a given drug’s disposition in the body is of paramount importance, since LDDS are not designed only to release the drug at a controlled, delayed, or prolonged rate, but are expected to reach the target and maintain a certain drug concentration. Drug disposition is a sequentially and simultaneously occurring process, depending on the absorption, distribution, metabolism, and elimination of the drug. For modified release SLN, the occurrence of general absorbability of the drug when released from the lipid matrix must be established. The intrinsic rate of absorption will be dictated by the lipid matrix (lipid nature, surfactant composition, particle size, surface properties), which means that the rate-limiting step is the release of the drug from the SLN. From a technological point of view, modified release LDDS should be multiple-dose systems designed to depict a steady-state concentration on the target tissue. The magnitude of this concentration is dependent on the amount of drug per unit of time (dose) and on the loss of drug from the volume of distribution per unit of time (clearance).

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In SLN, the release profile is also governed by the polymorphism of the lipid matrix. Polymorphism is the ability of a compound to crystallize in more than one distinct crystalline species with different internal lattices (Souto et al., 2006). Lipid crystallization and polymorphic transformations during storage time are important parameters for the performance of SLN as delivery system (Bunjes and Unruh, 2007; Kuntsche et al., 2005). The internal structure of the lipid matrix can vary depending on the threedimensional structure of triacylglycerols (TAGs) that can adopt the following structures upon crystallization: the hexagonal (a, H) phase, with a subcell lattice spacing of 0.415–0.42 nm, the orthorhombic perpendicular (b0 , O?) phase, with a subcell strong lattice spacing of 0.42–0.43 and 0.37–0.40 nm, or as the triclinic parallel (b, T||) phase with a subcell strong lattice spacing of 0.46 nm. Other polymorphic forms may also be found with complex acylglycerols including mixed acid TAGs or partial acylglycerols; for example, multiple b0 and b, sub-a, or intermediate forms, usually mentioned as bi. However, the nomenclature and properties of monoacid TAGs can also be used for these complex acylglycerols that have similar crystal packing. Once a stabilized SLN formulation has been produced, the release is faster from a, than from b0 and b phases. Crystallization of bulk TAGs from the melt after rapid cooling usually occurs in the less stable a-form, which transforms via the b0 -form, into the most stable b-form upon heating or during storage. In colloidal dispersions such as SLN, these transformations of TAGs are faster than in the bulk material, which leads to a change in the relative fraction of the polymorphic forms. Depending on the chemical nature of the lipid and on the production conditions, different fractions of a and b0 modifications may occur. This phenomenon can lead to a reduction of the melting point, or more precisely, changes in form and shift of the melting peak. The long-term stability of these polymorphic forms is less likely to occur, leading to a gradual transformation into more stable polymorphic forms, that is, increasing the content of b0 /bi and finally b. The occurrence of polymorphic transformations is not desired because the change in lipid structure is responsible for drug expulsion during storage and changes in the release profile of the incorporated drug (Bunjes and Unruh, 2007; Johnsson et al., 2005). When a drug is administered into the systemic circulation either directly introduced by intravascular injection or reaching the blood by absorption after extravascular administration, it is distributed not only throughout the bloodstream in the organs of immediate equilibrium, but also to other organs and tissues. It may also be eliminated by metabolism and/or by excretion. Since the molecules entering first will have been distributed and perhaps eliminated before the last molecules are absorbed, the entire process is dynamic, that is, made up of simultaneously occurring individual kinetic processes.

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Drug accumulative release wt (%)

120 Free solution

100

SLN formulation

80 60 40 20 0 0

20

40 Time (h)

60

80

Figure 6.1 Temozolomide release profiles from free solution (♦) and from SLN (e) in pH 6.8 PBS medium (modified after Huang et al., 2008a).

In modified release LDDS, the absorption of the drug after released from their matrix does not occur instantaneously, obviously delaying the distribution and elimination of the drug from the bloodstream. This can play an important role in reducing the adverse side effects of drugs with a short therapeutic window, since the drug is effectively released within the safety range (Huang et al., 2008a; Mu¨ller et al., 2006). SLN were reported to reduce the cardio toxicity and nephrotoxicity of temozolomide in comparison to a solution of free drug (Huang et al., 2008a). After i.v. administration of the drug dissolved in a liquid vehicle a high peak of drug concentration was depicted early after administration, followed by a relatively rapid decrease (Fig. 6.1). The peak concentration of temozolomide administered in SLN was lower, resulting from the fact that the drug is not immediately available for redistribution nor elimination until it is released from the nanoparticle structure. The mechanism of drug release form SLN may involve diffusion or degradation of the lipid matrix and subsequent diffusion (Manjunath and Venkateswarlu, 2006; Mu¨ller et al., 1996; Olbrich et al., 2002). Only after occurrence of one of these phenomena, the drug molecule is released into systemic circulation. As the drug is released subsequently, the maximal achieved concentration is not as high as the Cmax of drug administered in solution or emulsion. This may be particularly important in case of highly potent drugs, where initial concentration peak may reach toxic concentration and side effects may occur. Studies performed by Manjunath and coworkers using clozapine (Manjunath and Venkateswarlu, 2005) or nitrendipine (Manjunath and Venkateswarlu, 2006) as model drugs encapsulated in SLN revealed that SLN could increase the bioavailability of both drugs in comparison to free drug suspensions at all-time points. Although the concentration versus time

110 Drug plasma concentration (ng/m)

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400 350 300 250 200 150 100 50 0

SLN formulation Free solution 0

10

20

30 Time (h)

40

50

60

Figure 6.2 Plasma concentration versus time profiles of after administration of 10 mg/kg temozolomide as free solution () and as SLN (▲) in mice (modified after Huang et al., 2008a).

profiles were similar, the free drug suspensions appear to be less effective in delaying the distribution and elimination of the drug. Assuming first-order elimination, which is the case with most drugs, and plotting the concentration versus time profiles, the terminal slope is the elimination phase, characterized by a straight line. Figure 6.2 gives an example of such a profile obtained after i.v. administration of temozolomide-loaded SLN composed of stearic acid and lecithin (Huang et al., 2008a). The reasons behind the profile obtained as free solution may be the following: (i) the drug is rapidly absorbed and rapidly distributed between the systemic circulation (including organs of instant equilibrium) and the tissues to which the drug eventually goes; (ii) the drug is absorbed, but slowly (peripheral compartment); and (iii) the drug is slowly absorbed but rapidly distributed between the central and peripheral compartments. In SLN, the drug release usually takes longer than the distribution phase. As such, a monocompartmental model is usually obtained. In case of the peroral administration route, the uptake of particles from the gastrointestinal (GI) mucosa also needs to be considered. Due to their small size, SLN are directly taken up in the GI tract. The typical adhesiveness of SLN to the mucosal surface of the intestine walls provides longer contact of the nanoparticles with the gut, and, thus, enhanced time for drug absorption. The encapsulation of drugs into the lipid matrix also protects the drug from exposure to the environment with different pH or degradating enzymes and bacteria. Furthermore, the presence of the surfactant film used to stabilize SLN may also contribute to the higher permeability of the intestinal wall cell membranes (Manjunath and Venkateswarlu, 2005). The resulting concentration versus time profiles may therefore show higher concentrations of the drug in plasma for a longer time when compared to a solution or suspension of the same drug (Fig. 6.2). The exception may be the first few minutes after administration, when the concentration of the drug administered as a solution or suspension will rise faster (Zara et al., 2002a).

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Improved bioavailability of poorly soluble drugs may also be achieved by formulating them into LDDS. Formulating those drugs in SLN may provide concentration versus time profiles with comparable low variations and elimination rates, but the initial burst of the drug is avoid (Mu¨ller et al., 2006). The effect of lipid matrix composition on pharmacokinetic parameters has been examined by Manjunath and Venkateswarlu (2005). Higher plasma concentrations of clozapine were achieved from SLN composed of long acyl chain length TAGs because of the extended time required for their degradation. Likewise, higher drug concentrations were achieved with positively charged SLN compared to that with SLN of lower or negative charge, even if composed of the same lipid. Elimination or terminal half-life is the time required to reduce the concentration in blood, plasma, or serum to one-half after equilibrium has been reached. This parameter can be determined from the slope of the terminal line of a semilogarithmic plot by regression analysis. This parameter is important for the selection of the type (matrix) of SLN. The shorter the terminal half-life the greater will be the amount of the drug that will be incorporated into the system. Only drugs whose terminal half-life can be correlated with the pharmacological response are candidates for modified release LDDS. The reticuloendothelial system (RES) is responsible for removal of small xenobiotics from systemic circulation. Kuppfer cells take up all xenobiotics smaller than 7 mm. The elimination of small xenobiotics from the bloodstream is facilitated by adherence of serum opsonines onto the surface of the particles, making their surface more suitable to be recognized and eliminated by phagocytic cells (Wang and Wu, 2006). Opsonization may be avoided by coating the nanoparticles with hydrophilic polymers, such as poloxamers or polyethylenglycols (PEG), producing the so-called ‘‘stealth SLN’’ (Fundaro et al., 2000; Sanjula et al., 2009; Wang and Wu, 2006; Zara et al., 2002b; Zhang et al., 2008a). Poloxamer 188 also serves as a surfactant in SLN formulations, in addition to providing a hydrophilic coating that enhances the mean residence time (MRT) of the encapsulated drug (Huang et al., 2008a). The MRT is the mean time a drug molecule resides in the body, being defined as the time corresponding to 63.2% elimination from the body. As such, modified release LDDS should have a MRT significantly longer than that obtained with conventional dosage forms. Formulation of drugs in SLN generally leads to enhanced MRT in the systemic circulation as well as in various tissues. Yang et al. (1999) reported that MRT of camptothecin in systemic circulation increased 18 times when administered as SLN, and a fourfold increase has been observed for liver, spleen, kidneys, lungs, and heart. The reason behind this enhanced MRT may be the delayed elimination of camptothecin, since it takes place only after the drug is released from the lipid

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matrix. Another factor influencing the elimination, and thus the MRT, is the stealthiness of the SLN provided by poloxamer. Presystemic loss of drug can occur regardless of whether the drug is administered systemically or locally, except when given intravascularly. It is that fraction of drug that is degraded, inactivated, or metabolized after its release from the dosage form. Tissue degradation should not be underestimated. A decrease of presystemic loss when formulating drugs in SLN may also be a reason for the enhanced bioavailability that is usually observed. Most prominent is the presystemic loss of drug upon peroral administration, namely degradation by intestinal content and enzymes, biotransformation by intestinal microbial flora, the first-pass effect, the metabolism in the gut wall, mesenteric veins, portal vein, and liver. Many drugs are given perorally or deep rectally, undergoing an extensive first-pass effect, which is the biotransformation of the drug in the gut lumen prior to absorption and in the intestinal epithelium and/or liver after permeation of the intestinal mucosa (presystemic), but before entering systemic circulation. For such compounds, the GI tract is a poor choice for administration, and other routes (transdermal, buccal, nasal) need to be used. In case of topical administration the drug may be metabolized in the skin, or in the case of ocular administration the drug may be lost in the tear fluid, which is drained into the nasal cavity.

4. Modified Release Profile One of the most important features of SLN is the modified release profile usually depicted by the loaded drugs. The localization of the drug within the SLN matrix will influence its release. Three theoretical models have been proposed to describe the SLN structure, each of them being responsible for a particular release profile (Mu¨ller et al., 2002; Souto and Mu¨ller, 2007). The drug-enriched shell model describes the situation when the drug is present at such a concentration that it is fully solubilized by the lipid. In the course of SLN production, the solidification of lipid starts first, followed by solidification of drug-enriched lipid melt around the lipid cores. The drug is therefore located mostly near the surface of the particle, thus creating a drug-enriched shell. The release of the drug from particles with this structure will be fast, and burst release of the drug shortly after the administration may occur. This is the case of glucocorticoids when formulated into SLN, where the molecules are attached to the particle surface rather than being incorporated into the particle core (Lombardi Borgia et al., 2005), thereby causing their fast release after administration (Hu et al., 2006). Extended or prolonged release of the drug occurs when the drug is located in the core of the SLN, as described by the drug-enriched core

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model. This structure may be created when the drug is present at concentrations near or higher than its solubility limit in the lipid phase. During the production, the drug precipitates, creating the core of the particle, which is subsequently wrapped by a lipid shell as the lipid solidifies around the crystallized drug–lipid core. The drug is therefore covered by a practically drug-free lipid shell, which hinders its diffusion from the matrix. The release profiles may be prolonged for several hours, to several days (Esposito et al., 2008; Huang et al., 2008a; Kumar et al., 2007). The matrix structure is dependent on the composition and on the production parameters (Souto and Mu¨ller, 2007). The in vitro release will correlate with the concentration versus time profile in vivo. The prolonged release of the drug will delay its redistribution and elimination, thus providing greater MRT both in systemic circulation and tissues, at the same time avoiding high peak concentrations that may occur after i.v. administration of free drug in a solution or in a suspension.

4.1. Short or relatively long half-life drugs Short-half-life drugs, such as butyrate or peptides (calcitonin, insulin), were shown to be protected from immediate clearance by encapsulation into SLN. For example, butyric acid, that is normally eliminated from the bloodstream within minutes and does not reach a therapeutic level even upon large dose administration, could achieve a concentration sufficient to exhibit its anti-inflammatory and antineoplastic effects (Brioschi et al., 2008; Dianzani et al., 2006). Moreover, the enhanced protection against drug degradation and longer circulation enables reached the brain (Brioschi et al., 2007). In oral administration, cholesteryl butyrate-based SLN proved to be more efficient than the prodrug or salt of the active drug form. A relatively long half-life drug concentration could also be improved. Bromocriptine loaded in SLN could maintain the therapeutic concentration in rats for over 5 h and improve the mobility, while the effect of free drug was nearly absent within this period of time (Esposito et al., 2008).

4.2. Targeting to a specific tissue by implantation for long-term release Enhanced circulation time of SLN may favor their localization in specific tissues. Different tissue distribution was found in mice after administration of temozolomide (Huang et al., 2008a). Of particular relevance was the increased concentration in the brain and decreased concentration in heart and kidneys, where the drug may exhibit toxic effects. In general, the nanoparticles tend to accumulate in macrophages of the RES organs that remove them from the circulation. Passive targeting to RES organs is achieved solely by increasing the circulation time (Xu et al., 2009;

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Ye et al., 2008). Targeting to these organs may be even enhanced by surfacing the nanoparticles with galactosides (Lian et al., 2008). SLN may also be passively targeted to solid tumors due to the leaky vasculature of these latter. If present in the circulation during a sufficient time, SLN tend to accumulate in the tumor. This enhanced retention and permeation effects may be used for passive targeting provided that the surface characteristics of the nanoparticle enable them to escape the RES for a sufficient time (Reddy et al., 2006; Ruckmani et al., 2006b; Tausch et al., 2008).

5. Biopharmaceutical Aspects of Administration Routes Several routes have recently gained increased attention for LDDS. Based on the documentation and state-of-art of topical administration of such systems, the first lipid nanoparticle formulations have entered the market (Mu¨ller et al., 2007; Pardeike et al., 2008). These are in general designed for cosmetic purposes; however, their suitability for application on skin, documented by these products, is of high importance when considering the application of actives for skin diseases treatment. Several drugs used for the treatment of skin disorders may cause side effects either in the skin or, if they happen to enter the systemic circulation, even systemic side effects. This is of paramount importance in terms of application for long periods of time, which is the case of most skin diseases. The highly lipophilic drugs partition into the skin lipids of its horny layer and permeate toward the epidermis, and eventually to the dermis which is rich in vasculature. Although the dermis is less lipophilic than the upper parts of the skin, if the drug reaches this layer, it may enter into the bloodstream, a condition not acceptable for drugs meant for local action. Due to insolubility in water, the drugs are formulated in systems containing solubilizing agents that may subsequently increase their penetration into deeper layers of the skin. The extent of drug permeation through the skin is dependent on more factors. The penetration of the drug into the deeper layers of the skin is influenced by the skin/vehicle partition coefficient. Encapsulation of a lipophilic drug into SLN may decrease this coefficient, maintaining a higher portion of the drug in the upper parts of the skin and preventing the drug to permeate through the skin (Fang et al., 2008; Iscan et al., 2006). The simple fact that the drug is soluble in the lipid matrix of SLN explains this effect. As the horny layer and epidermis are highly lipophilic, SLN may penetrate them easily and accumulate preferentially in these upper parts of the skin. As shown for cyproterone acetate-loaded SLN, the nanoparticles accumulated mostly within the first 100 mm of the skin (Stecova et al., 2007).

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Similar results were observed for a fluorescent dye loaded in a lipid nanoparticle formulation composed of cetyl palmitate and medium-chain TAGs (Teeranachaideekul et al., 2008a). The accumulation of drug administered in SLN in the epidermis was reported for several drugs for which systemic absorption is unwanted, for example, podophyllotoxin (Chen et al., 2006), psoralens (Fang et al., 2008), isotretinoin (Liu et al., 2007b), and several NSAIDS ( Joshi and Patravale, 2006, 2008; Puglia et al., 2008). Enhancement of skin bioavailability of a hydrophilic drug (penciclovir) was also achieved by loading it in SLN composed of glyceryl monostearate and lecithin (Lv et al., 2009). The skin penetration effects depicted by SLN are also dependent on the occlusive effect that these particles show upon topical administration, in addition to the permeation enhancing effects of the lipid materials employed for SLN production. The occlusive properties of SLN are documented by several studies (Souto and Mu¨ller, 2008). These properties can be controlled by the degree of crystallinity of the lipid matrix. The presence of small amounts of liquid lipids inside the solid lipid matrix of SLN destabilize the crystal order, leading to lower occlusion (Wissing and Mu¨ller, 2002). The surfactants employed in SLN formulations will also influence the extent to which the drug permeates through the skin, due to their own permeation enhancement effects. Lecithin-stabilized SLN showed significantly higher epidermis localization than polysorbate stabilized SLN (Chen et al., 2006). The SLN may penetrate the skin both along the stratum corneum and the hair the follicle route (Chen et al., 2006). The residence time on the skin surface of drugs delivered by means of SLN formulations was also found to be significantly longer than when administered by classical topical dosage forms (Puglia et al., 2008). With improved penetration into the upper layer of the skin, together with release of the drug prolonged over a period of time, lower doses of the drug can be administered. As the permeation through the skin is usually low, side effects of the administered drug may be avoided. Due to their small size, SLN adhere onto the mucosal surfaces, which can also be used for improvement of bioavailability, not only in skin but also in ocular and pulmonary applications. The increased residence time is of particular relevance in ophthalmic applications, where the drug solution is removed within a short time. Increased corneal permeability and precorneal retention of ibuprofen formulated in lipid nanoparticles has been reported recently (Li et al., 2008). Particle size is of utmost importance in effective pulmonary application. SLN even seem promising for application of peptides and proteins for systemic delivery through the lung, as shown for insulin-loaded SLN (Liu et al., 2008a), which increased significantly the bioavailability of insulin, in comparison to control. Uptake of SLN from alveoli to the lymph system was shown by radiolabeled SLN (Videira et al., 2002),

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providing a possibility for targeting the lymph nodes, and to reach the systemic circulation. Soybean phosphatidylcholine is used as an absorption enhancer in pulmonary delivery of proteins and peptides, and was also reported to facilitate nebulization of a nanoparticle formulation (Liu et al., 2008a). At the same time it can be employed as the surfactant stabilizing the SLN. Another promising administration route for SLN is the oral route. SLN undergo enzymatic degradation by lipases/co-lipases in the gut, the structure is disrupted and micellar structures with bile salts are assembled. These are subsequently taken up actively by intestinal epithelial cells or M-cells of Peyer’s patches (des Rieux et al., 2006). The drug incorporated into SLN will therefore be incorporated into the micelles and be absorbed together with the lipids. This effect has been suggested as the mechanism for increased absorption of drugs from the gut, as observed in the case of SLN carrying cyclosporine A (Mu¨ller et al., 2006) or salmon calcitonin (Martins et al., 2009). The absorption of SLN from the gut into systemic circulation was confirmed by Yuan et al. (2007), who monitored SLN loaded with a fluorescent dye, which was not absorbed on its own in the lymph. As much as 78% of the absorbed fluorescent SLN were detected in the thoracic duct, suggesting the lymphatic uptake as the major absorption mechanism for drugs encapsulated in SLN. Increased oral bioavailability has been reported for several drugs encapsulated in SLN. The size of the particles improves the adhesion to mucosal surfaces, providing closer and longer contact with the intestine wall. Moreover, the use of surfactants to stabilize the SLN may improve the absorption once the nanoparticles reach the gut. The great advantage is that the lipid structures are passed into the lymphatic circulation, avoiding the passage through the portal vein and the first-pass metabolism. This provides the opportunity of bioavailability enhancement upon oral administration of drugs with high first-pass effect. Lovastatin administered by means of SLN was shown to reach the systemic circulation in higher concentrations, as the first-pass loss is decreased (Suresh et al., 2007). The enhancement of oral bioavailability of drugs incorporated into SLN was proven for both lipophilic and hydrophilic drugs. Absorption rate of fenofibrate was higher and subsequent Area Under the Curve (AUC) in plasma was greater compared to micronized drug (Hanafy et al., 2007). Moreover, the interindividual variability of studied pharmacokinetic parameters decreased upon administration of this drug either in SLN or as a nanosuspension. Vinpocetine-loaded SLN were suggested as a potential formulation for this drug, as it can be protected from enzymatic degradation prior to absorption, thereby enhancing enhance its subsequent plasma concentration (Luo et al., 2006). Recently, a fivefold increase in oral bioavailability was reported for quercetin when incorporated into SLN (Li et al., 2009), and SLN proved a suitable solution for the administration of peptides and proteins via the oral route.

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SLN are of particular interest for increasing the penetration and permeation of skin to several drugs, but may also be employed for peroral, buccal, nasal, rectal, intramuscular, and subcutaneous routes of administration. SLN may increase the amount absorbed by several orders of magnitude and they are intended to increase the absolute bioavailability of a drug for a given route of administration. Since SLN are pharmacologically inert, the mechanisms behind their role in increasing drug bioavailability may include increasing drug solubility (used as cosolvent), wettability by reducing surface tension, spreadability (increased contact surface area), miscibility with the epithelialium (mucus), and the skin (skin lipids), permeability of membranes by interacting with phospholipids (surfactant molecules), permeability of membranes by binding surface calcium and magnesium ions, as well as by changing the hydrostatic pressure to act on water-filled pores (sugars, glycols). These phenomena may be observed for both hydrophobic and hydrophilic drugs if the mechanism is based on a change a of membrane permeability.

6. Clinical Pharmacology The clinical pharmacological studies may supply pertinent information on the desired interaction between the drug molecules and receptors, whether a local effect, a general systemic concentration, or a specific target (tissue) are expected. Furthermore, they also contribute to the understanding of the desired or required drug concentration at the site of action, and its acceptable range and toxic limitation, for the development of a feasible LDDS.

6.1. Local effects and systemic distribution Classical examples of local applications are the topical, ocular, and other mucosal routes. The application of LDDS onto the skin is very well explored. Provided that the drug within the lipid nanoparticle matrix accumulates preferentially in the epidermis, the action of the drug will be limited to the local skin effects. Several examples of drugs for topical application on skin are listed in Table 6.1. Local effects are also expected for SLN for application in the eye, for example, NSAIDs applied in eye to prevent the inflammation after surgery on eye and to suppress the pain. Lipid nanoparticles containing the sodium salt of diclofenac (Attama et al., 2008) and ibuprofen (Li et al., 2008) were proposed as nonirritant and effective carriers. Cyclosporine A-loaded SLN were shown to penetrate corneal cells (Gokce et al., 2008).

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Table 6.1 Examples of drugs administered by LDDS for local distribution Drug

References

Alpha lipoic acid Ascorbyl palmitate Beta-carotene

Ruktanonchai et al. (2009) Teeranachaideekul et al. (2007a, 2008c) Hentschel et al. (2008), Triplett and Rathman (2009) Pojarova et al. (2006), Polovyanenko et al. (2008), Shahgaldian et al. (2008) Hu et al. (2006)—NLC Stecova et al. (2007) Xiang et al. (2007) Lu et al. (2008)

Calixarenes Clobetasol proprionate Cyproterone acetate Dexamethasone acetate Dexamethasone palmitate Insect repelent Ketoconazole Ketoprofen Minoxidil Naproxen Nile red Octadecylaminefluorescein isothiocynate Podophylotoxin Psoralen Retinoids

Iscan et al. (2006) Souto and Mu¨ller (2006), Souto et al. (2006) Puglia et al. (2008)—NLC Silva et al. (2009)—NLC Puglia et al. (2008)—NLC Kuchler et al. (2009), Teeranachaideekul et al. (2008b) Yuan et al. (2007)

Chen et al. (2006), Zhu et al. (2009) Fang et al. (2008) Castro et al. (2007), Jee et al. (2006), Liu et al. (2007b), Mandawgade and Patravale (2008), Pople and Singh (2006), Shah et al. (2007) Rhodamine B Wang and Wu (2006) Rhodamine 123 Chattopadhyay et al. (2008) Sunscreens Cengiz et al. (2006), Lee et al. (2007a), Scalia and Mezzena (2009), Xia et al. (2007) Tilmicisine Han et al. (2009) Triamcinolone acetonide Liu et al. (2008b) Ubidecarenone Teeranachaideekul et al. (2007b) Valdecoxib Joshi and Patravale (2006)

With respect to systemic distribution, the bloodstream must be reached usually by oral or parenteral administration. Miscellaneous drugs incorporated into LDDS reported recently are listed in Table 6.2. Systemic effects of encapsulated drugs within SLN may also be achieved by transdermal administration, upon creating a supersaturated system

SLN Formulations: Pharmacokinetics and Biopharmaceutics

Table 6.2 Examples of drugs administered by LDDS for systemic distribution Drug

References

Actarit Arthemeter Baclofen Bromocriptine Buprenorphine Calcitonin Camptothecin 10-Hydroxycamptothecin Carvedilol Chlorambucil Cholesteryl butyrate Cisplatin Ciprofloxacine Cyclosporine A Diazepam pDNA Docetaxel Doxorubicin Ferrulic acid Flavonoids Flurbiprofen Floxuridinyl diacetate Ibuprofen Insulin

Ye et al. (2008) Joshi et al. (2008) El Assawy et al. (2008) Esposito et al. (2008) Wang et al. (2009) Martins et al. (2009) Huang et al. (2008b) Liu et al. (2008c), Zhang et al. (2008b) Faisal et al. (2008), Sanjula et al. (2009) Sharma et al. (2009) Brioschi et al. (2007, 2008) Tian et al. (2008) Jain and Banerjee (2008) Mu¨ller et al. (2008) Abdelbary and Fahmy (2009) Asasutjarit et al. (2007) Xu et al. (2009) Serpe et al. (2006), Ying et al. (2008) Bondi et al. (2009b) Li et al. (2009), Wang et al. (2007) Bhaskar et al. (2009a), Han et al. (2008) Lian et al. (2008) Casadei et al. (2006), Paolicelli et al. (2008) Battaglia et al. (2007), Bi et al. (2009), Gallarate et al. (2008), Liu et al. (2008a,d), Sarmento et al. (2007) Hsu and Su (2008) Priano et al. (2007), Rezzani et al. (2009) Paliwal et al. (2008), Ruckmani et al. (2006b) Lu et al. (2006b) Bondi et al. (2009a) Bhaskar et al. (2009b), Kumar et al. (2007), Manjunath and Venkateswarlu (2006) Vivek et al. (2007) Dong et al. (2008), Lee et al. (2007b), Yuan et al. (2008) Yang et al. (2009) de Ven et al. (2009) Kuo and Chen (2009), Kuo and Su (2007) Zhang et al. (2007) Kuo and Chen (2009), Kuo and Su (2007) Zhang et al. (2007)

Magnetite Melatonin Methotrexate Mitoxantrone Nimesulide Nitrendipine Olanzapine Paclitaxel Praziquantel Saponins Saquinavir Silibinin Saquinavir Silibinin

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consisting of drug-loaded SLN dispersed in a drug-loaded medium, and applied under occlusion (Souto et al., 2007). The achieved bioavailability, expressed as the AUC, was reported similar to the AUC obtained after oral administration of nitrendipine (Bhaskar et al., 2009b), with the advantage of sustained and more prolonged residence of the drug in the blood. SLN were also shown to be suitable for transdermal delivery of flurbiprofen (Han et al., 2008) and melatonin (Priano et al., 2007), which showed systemic effects after encapsulation in SLN. Similar to the local effect, a desired drug concentration is often required only in a specific organ or tissue when designing a targeting approach. The LDDS may be administered intra- or extravascularly. However, the drug is either in the form of a prodrug which is activated at the target organ only, or somehow entrapped and released at the target organ. The rationale for organ targeting is to reach a therapeutic drug concentration at the receptor site with minimal systemic body burden. Examples of targeting SLN formulations include ferritin-surfaced SLN to carry 5-fluorouracil to breast cancer cells ( Jain et al., 2008), and monostearin-based SLN containing folic acid-conjugated stearic acid for paclitaxel delivery (Yuan et al., 2008). Other targeting applications include the delivery of gene therapeutic agents and diagnostics for brain tumors (Brioschi et al., 2007).

6.2. Dose size Considerations must be given to usual (noncontrolled release) dose size and dose rate. Drugs that act systemically and are therapeutically effective only in relatively large doses do not qualify a priori for modified release LDDS. A typical modified release system is essentially a multiple-dose unit. Even though the dose is not exactly the amount of a single dose multiplied by the number of dosing intervals, for usual doses equivalent to the duration of a modified release system, the total amount per unit is much higher than that of a single dose. Modified release LDDS imply a decrease of the usual dose used to achieve therapeutic concentrations. It was reported that mitoxantrone administered by means of SLN yielded the same tumor cell growth inhibition in half the dose of the free drug (Lu et al., 2006a). In another study, to assure a concentration within the therapeutic range, the lovastatin dose was lower if administered as SLN, because in this formulation it has higher bioavailability and is released over a prolonged period of time (Suresh et al., 2007).

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6.3. Concentration fluctuations and therapeutic range Theoretically and desirably, a modified release LDDS, unless programed for a triggered mechanism, should release the drug by a zero-order process, which would result in a concentration versus time profile similar to that after i.v. constant rate infusion. However, zero-order release is rarely achieved, and if so, the resulting steady-state concentration will probably not be a straight line parallel to the abscissa, but more likely will be a soft wave due to many factors such as circadian rhythm, body activity, stress, and food intake. Nevertheless, in the design of a modified release SLN, it is necessary to select a desired (or required) target concentration or concentration range. The target concentration should usually be the same as for conventional dosage forms. The difference will be that the fluctuations of drug concentration in blood will be much smaller for the duration of the device as observed within each dosing interval of a conventional drug product. Ideally, candidates for modified release LDDS will be drugs of such wide therapeutic range that neither batch-to-batch variations in release rate nor individual physiological variations exceed the therapeutic range.

6.4. Toxicological concentration Toxicological concentration of a candidate for a modified release LDDS should be much higher than the therapeutic range. This is of utmost importance in case of accidental disruption or degradation of the LDDS, rupture or malfunction. Another consideration is the question of whether the drug is metabolized pharmacologically into highly potent structures or to toxic metabolites. Since SLN control only the release of the parent compound, but not its biotransformation, and since the rate of metabolism may vary widely among lipid materials and in the same material obtained by the same supplier, long-range predictions are difficult to make.

7. Concluding Remarks SLN play a prominent role as novel LDDS in pharmaceutical technology. The present chapter encompasses a thorough report on the use of SLN as promising LDDS to increase the bioavailability of several drugs. This chapter shortly outlined the pharmacokinetic and biopharmaceutical properties governing the modified release profile of SLN in drug delivery and summarized their applications for different routes of administration. The basics of pharmacokinetic parameters were explained in detail and examples of their use in the evaluation of SLN performance in controlling the release profile of the loaded drugs were given.

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cancer: Findings from the Austrian Sentinel Node Study Group. Ann. Surg. Oncol. 15, 3378–3383. Teeranachaideekul, V., Mu¨ller, R. H., and Junyaprasert, V. B. (2007a). Encapsulation of ascorbyl palmitate in nanostructured lipid carriers (NLC)—Effects of formulation parameters on physicochemical stability. Int. J. Pharm. 340, 198–206. Teeranachaideekul, V., Souto, E. B., Junyaprasert, V. B., and Mu¨ller, R. H. (2007b). Cetyl palmitate-based NLC for topical delivery of Coenzyme Q(10)—Development, physicochemical characterization and in vitro release studies. Eur. J. Pharm. Biopharm. 67, 141–148. Teeranachaideekul, V., Boonme, P., Souto, E. B., Mu¨ller, R. H., and Junyaprasert, V. B. (2008a). Influence of oil content on physicochemical properties and skin distribution of Nile red-loaded NLC. J. Control. Release 128, 134–141. Teeranachaideekul, V., Souto, E. B., Mu¨ller, R. H., and Junyaprasert, V. B. (2008b). Physicochemical characterization and in vitro release studies of ascorbyl palmitate-loaded semi-solid nanostructured lipid carriers (NLC gels). J. Microencapsul. 25, 111–120. Teeranachaideekul, V., Boonme, P., Souto, E. B., Mu¨ller, R. H., and Junyaprasert, V. B. (2008c). Influence of oil content on physicochemical properties and skin distribution of Nile red-loaded NLC. J. Control. Release 128, 134–141. Tian, J., Pang, X., Yu, K., Liu, L., and Zhou, J. (2008). Preparation, characterization and in vivo distribution of solid lipid nanoparticles loaded with cisplatin. Pharmazie 63, 593–597. Triplett, M. D., and Rathman, J. F. (2009). Optimization of beta-carotene loaded solid lipid nanoparticles preparation using a high shear homogenization technique. J. Nanopart. Res. 11, 601–614. Videira, M. A., Botelho, M. F., Santos, A. C., Gouveia, L. F., Pedroso De Lima, J. J., and Almeida, A. J. (2002). Lymphatic uptake of pulmonary delivered radiolabelled solid lipid nanoparticles. J. Drug Target. 10, 607–613. Vivek, K., Reddy, H., and Murthy, R. S. (2007). Investigations of the effect of the lipid matrix on drug entrapment, in vitro release, and physical stability of olanzapine-loaded solid lipid nanoparticles. AAPS PharmSciTech 8, E83. Wang, Y., and Wu, W. (2006). In situ evading of phagocytic uptake of stealth solid lipid nanoparticles by mouse peritoneal macrophages. Drug Deliv. 13, 189–192. Wang, D., Zhao, P., Cuia, F., and Li, X. (2007). Preparation and characterization of solid lipid nanoparticles loaded with total flavones of Hippophae rhamnoides (TFH). PDA J. Pharm. Sci. Technol. 61, 110–120. Wang, J. J., Liu, K. S., Sung, K. C., Tsai, C. Y., and Fang, J. Y. (2009). Skin permeation of buprenorphine and its ester prodrugs from lipid nanoparticles: Lipid emulsion, nanostructured lipid carriers and solid lipid nanoparticles. J. Microencapsul. 1–14. Wissing, S., and Mu¨ller, R. (2002). The influence of the crystallinity of lipid nanoparticles on their occlusive properties. Int. J. Pharm. 242, 377–379. Xia, Q., Saupe, A., Mu¨ller, R. H., and Souto, E. B. (2007). Nanostructured lipid carriers as novel carrier for sunscreen formulations. Int. J. Cosmet. Sci. 29, 473–482. Xiang, Q. Y., Wang, M. T., Chen, F., Gong, T., Jian, Y. L., Zhang, Z. R., and Huang, Y. (2007). Lung-targeting delivery of dexamethasone acetate loaded solid lipid nanoparticles. Arch. Pharm. Res. 30, 519–525. Xu, Z., Chen, L., Gu, W., Gao, Y., Lin, L., Zhang, Z., Xi, Y., and Li, Y. (2009). The performance of docetaxel-loaded solid lipid nanoparticles targeted to hepatocellular carcinoma. Biomaterials 30, 226–232. Yang, S. C., Lu, L. F., Cai, Y., Zhu, J. B., Liang, B. W., and Yang, C. Z. (1999). Body distribution in mice of intravenously injected camptothecin solid lipid nanoparticles and targeting effect on brain. J. Control. Release 59, 299–307.

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C H A P T E R

S E V E N

Preparation of Complexes of Liposomes with Gold Nanoparticles Chie Kojima,* Yusuke Hirano,† and Kenji Kono† Contents 132 134 134 136 137 137

1. 2. 3. 4. 5. 6. 7.

Introduction Preparation of Complexes of EYPC Liposomes with Au NPs Time-Dependent SPR of the Complexes TEM Analysis of the Complexes DLS Analysis of the Complexes Calcein Release from the Complexes Estimation of Numbers of the Au NP and the Liposome in the Complexes 8. Optimization of Lipid Components of the Complexes 9. Concluding Remarks Acknowledgment References

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Abstract Liposomes have been widely used as drug carriers. Visible liposomes have recently become more attractive as drug carriers in personalized medicine. Gold nanoparticles (Au NPs) have unique size- and shape-dependent properties based on their surface plasmon resonance. They can be visualized by computed tomography (CT) and laser optoacoustic imaging. In addition, their photothermogenic properties are useful for photothermal therapy and photoresponsive drug release from liposomes. Therefore, complexation of liposomes with Au NPs is of considerable interest. There are three types of complex: Liposomes containing Au NPs in the inner phase, liposomes with Au NPs at the lipid membrane, and liposomes modified with Au NPs on the surface. This chapter focuses on the preparation and characterization of the third type of complex that is prepared by direct mixing of a Au NP dispersion with a liposome suspension.

* {

Nanoscience and Nanotechnology Research Center, Research Institutes for the Twenty First Century, Osaka Prefecture University, Osaka, Japan Graduate School of Engineering, Osaka Prefecture University, Osaka, Japan

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64007-6

#

2009 Elsevier Inc. All rights reserved.

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1. Introduction Drug delivery systems (DDS) are attractive for chemotherapy, because they reduce severe side effects and facilitate effective drug action. Drug carriers are essential for the success of DDS. There are many types of drug carriers available, such as micelles, polymers, virus particles, proteins, and liposomes. Liposomes have classically been used as drug carriers and some, such as Doxil, have already gained Food and Drug Administration (FDA) approval (Immordino et al., 2006). Liposomes are vesicle structures composed of phospholipids with a hydrophobic tail and a hydrophilic head. Due to their amphiphatic character, liposomes can encapsulate water-soluble drug molecules in the inner phase and lipid-soluble ones in the hydrophobic membrane. Improvements to the next generation of liposome drug carriers include controllable release and visibility of the liposome, obtained by addition of functional molecules. Functional molecules that have been added to liposomes include thermosensitive, pH-sensitive, and visible molecules (Al-Jamal and Kostarelos, 2007; Chilkoti et al., 2002; Immordino et al., 2006; Kono, 2001; Kono and Arshady, 2006). Gold nanoparticles (Au NPs) could also be of use as functional molecules because of their interesting shape- and size-dependent physical and chemical properties (Burda et al., 2005; Daniel and Astruc, 2004). Due to their surface plasmon resonance (SPR), Au NPs strongly absorb visible light (Link and El-Sayed, 1999). In addition, they convert this light energy to heat energy (Link and El-Sayed, 2000). Consequently, Au NPs have been considered for photothermal therapy, imaging, and photosensitive drug release (Govorov and Richardson, 2007; Jain et al., 2007; Kim et al., 2007; Pissuwan et al., 2006). Complexes of liposomes with Au NPs are attractive because they can act as both stimuli-responsive and visible drug carriers (Hong et al., 1983; Kojima et al., 2008; Li et al., 2004; Paasonen et al., 2007; Volodkin et al., 2009; Wu et al., 2008). These photochemical properties are only expressed by Au NPs ranging from approximately 2 to 100 nm in diameter (Burda et al., 2005; Daniel and Astruc, 2004). When Au NPs aggregate, they lose their unique photochemical properties. Therefore, the preparation of the complexes also has to be performed without the Au NPs aggregating. Three types of Au NP–liposome complexes exist (Fig. 7.1). The first of these contains Au NPs in the inner phase of the liposome. This type of complex can be prepared by reducing Au ions in the presence of a reductant (Wu et al., 2008), and has been used to investigate in vivo liposome distribution (Hong et al., 1983). In DDS applications, this reduction may be detrimental to drug activity. In the second type of complex, Au NPs are present in the lipid membrane (Paasonen et al., 2007; Park et al., 2006).

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A

B

C

Figure 7.1 A schematic image of the complexes formed between liposomes and Au NPs. (A) A liposome encapsulating Au NPs, (B) a liposome loaded with Au NPs in the membrane, and (C) a liposome modified with Au NPs at the surface (from Kojima et al., 2008).

However, as the thickness of the lipid bilayer is only about 4 nm, a limited number of Au NPs can be incorporated using this strategy. The third complex type is a liposome modified with Au NPs on its surface. This type of complex is simply prepared by mixing an Au NP dispersion with a liposome suspension (Kojima et al., 2008; Volodkin et al., 2009). In this chapter, the third preparation method is described in detail (Kojima et al., 2008), and the influence of lipid components on the liposome are described. The lipids include: egg yolk phosphatidylcholine (EYPC), distearyldimethylammonium bromide (DDAB), distearoyl-sn-glycero-3phosphoethanolamine-N-[methyl(poly(ethylene glycol))] (PEG-PE), and stearylmercaptan (C18-SH). The complexes are characterized by a variety of techniques. UV–Vis spectrometry is used to investigate SPR and its influence on the stability of Au NPs. The size and morphology of the complexes are analyzed by dynamic light scattering (DLS) and transmission electron microscopy (TEM). The release of encapsulated molecules from the liposome is examined by using the fluorescent dye calcein.

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2. Preparation of Complexes of EYPC Liposomes with Au NPs Au NPs with a diameter of 13 nm are prepared by reducing Au ions with citric acid, according to a previous report (Grabar et al., 1995). A 5-min reflux is carried out with 50 ml of 1 mM HAuCl4 (Wako Pure Chemical Industries Ltd.). Five milliliters of 38.8 mM sodium citrate (Kishida Chemical) are added with vigorous stirring. The solution changes color from yellow to dark wine red. After cooling, an Au ion dispersion with a concentration of 0.91 mM is obtained by filtration. EYPC can be obtained from a variety of sources (Avanti Polar Lipids, Sigma, etc.). For our studies, it was kindly provided by NOF Corp. (Tokyo, Japan). A chloroform solution of EYPC (10 mg/ml, 500 ml) is evaporated to remove the solvent. The obtained thin lipid membrane (5.0 mg, 6.25 mmol) is further dried under vacuum for at least 2 h, and then dispersed in 0.5 ml of phosphate-buffered saline (PBS; 20 mM Na2HPO3–NaH2PO3, 150 mM NaCl, pH 7) with sonication in a bath type sonicator for 3 min. The liposome suspension is freeze-thawed four times. The obtained liposome suspension is extruded through a polycarbonate membrane with a pore diameter of 200 nm (Kojima et al., 2008; Kono et al., 1999). Lipid concentrations are estimated by Phospholipids C-Test Wako (Wako Pure Chemical Industries Ltd.) according to the manufacturer’s instructions. Sample, blank, and standard solutions are each mixed with the color reagent, which contains phospholipase D, choline oxidase, peroxidase, 4-aminoantipyrine, 3,5-dimethoxy-N-ethyl-N-(2-hydroxy-3-sulfopropyl)aniline sodium, and ascorbic oxidase. After reaction at 37
C for 5 min, the absorbance at 600 nm is measured to estimate the EYPC concentration. The concentration-estimated liposome suspension in PBS is diluted up to 950 ml by addition of distilled water and 10 times concentrated PBS (10 PBS) in an appropriate ratio. Fifty microliters of Au NP dispersion ([Au] ¼ 0.91 mM ) is added to the liposome suspension and vortexed. The final solution is 46 mM of gold ions in 1 PBS. For use in TEM analysis, complexes with an EYPC/Au ratio of 10/1 are prepared by vortex mixing 500 ml of the Au NP dispersion ([Au] ¼ 0.91 mM) with 500 ml of the liposome suspension, diluted with distilled water and 10 PBS. The final solution is 4.6  102 mM of gold ions in 1 PBS.

3. Time-Dependent SPR of the Complexes One of characteristic properties of Au NPs is surface plasmon absorption around 523 nm. To investigate this in each of the complex dispersions, the UV–Vis absorption spectra ranging from 400 to 800 nm are measured

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using a Jasco Model V-560 spectrophotometer ( Jasco Inc., Japan) at 25
C (Haba et al., 2007). As a control, the spectrum of a dispersion solution without liposomes is also measured. The SPR absorption of Au NPs in the absence of liposomes almost disappeared under physiological conditions (Fig. 7.2A). This is due to aggregation resulting from shielding of electrostatic repulsion (Burda et al., 2005; Daniel and Astruc, 2004). In contrast, the SPR absorption in spectra of Au NPs in the presence of EYPC liposomes was retained; this was dependent on the EYPC/Au ratio (Fig. 7.2). At ratios of 1/10 and 1/1, the SPR signal decreased (rapidly and slowly, respectively), while at a ratio of 10/1, the signal remained after 12 h. This indicated that EYPC liposomes contributed to stable dispersion of Au NPs under isotonic conditions, preventing their aggregation.

A

B

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4h 8h 12 h

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0.2 0.1

0 400

Liposome only

Liposome only 500 600 700 Wavelength (nm)

800

0 400

500 600 700 Wavelength (nm)

800

Figure 7.2 Time-dependent UV–Vis spectra of the complex of EYPC liposomes (200 nm in diameter) with Au NPs at different ratios in PBS. The time-dependent UV–Vis spectra of Au NPs are only shown as a control (A). The mole ratio of EYPC to Au is 1/10 (B), 1/1 (C), and 10/1 (D). [Au] ¼ 46 mM (from Kojima et al., 2008).

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4. TEM Analysis of the Complexes TEM analysis is performed as follows (Hayashi et al., 1998; Kojima et al., 2008). Collodion-coated grids are coated with a carbon thin film, using an ion sputtering device (E-1030, Hitachi High-Technologies Corp., Japan). After incubation for 1 day in a desiccator, a small drop of sample is placed on the grid for 5 min and then the excess drawn off with filter paper. The Au concentration of the complex at an EYPC/Au ratio of 0/10 and 1/10 was 46 mM. At a ratio of 10/1, the Au concentration of the complex is increased up to 4.6  102 mM, because it is difficult to identify the liposome under this diluted condition. A drop of 2% (w/v) phosphotungstic acid (pH 7) is applied to the grid, drawn off with filter paper, and the stained sample is allowed to dry. The grid is viewed under an electron microscope at 200 kV ( JEOL Ltd., JEM-2000FEX II). Figure 7.3 shows the TEM images of Au NPs in the absence and presence of EYPC liposomes. Large aggregates were observed in TEM

Figure 7.3 TEM images of the complexes of EYPC liposomes with Au NPs at the EYPC/Au ratio of 0/10 (A), 1/10 (B), and 10/1 (C) in PBS after the 6 h incubation. Arrows indicate the Au NPs. Bar ¼ 100 nm (from Kojima et al., 2008).

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images of both the sample comprising only Au NPs and the complex at an EYPC/Au ratio of 1/10. In contrast, no Au NPs aggregates were observed for the complex at an EYPC/Au ratio of 10/1. These findings are consistent with the SPR analysis results. In the TEM image of the complex at an EYPC/Au ratio of 10/1, many Au NPs were observed at the boundary surface within the liposomal assembly. This indicates that the Au NPs complexed with the liposomes.

5. DLS Analysis of the Complexes DLS analysis is performed in the vesicle mode of Nicomp ZLS380 (Nicomp) at room temperature using EYPC suspension (2.5 ml, approximately 0.01 mM) (Kojima et al., 2008). This is performed in both the absence and presence of the Au NP at an EYPC/Au ratio of 10/1. The size of liposome was unchanged before and after the addition of Au NPs (Fig. 7.4), suggesting that Au NPs can complex with liposomes without the aggregation. This finding is not consistent with our TEM results, in which liposomal assembly was observed. It is possible that the liposomal aggregation was an artifact of TEM sample preparation.

6. Calcein Release from the Complexes The collapse behavior of liposomes is analyzed by adding Au NPs to calcein-loaded liposomes. Although the fluorescence of the calcein encapsulated in the liposomes is essentially quenched, an intense florescence is observed after its release from the liposome. The loaded and unloaded percent of calcein is determined from the fluorescence at the initial step and the fluorescence after adding detergent to collapse the liposome, respectively. Calcein release measurements are performed according to the method previously reported (Kono et al., 1994), with some modification (Kojima et al., 2008). Fluorescence intensity is largely influenced by the photochemical properties of the Au NP, so the concentration of Au NPs was decreased to the minimum detection level of calcein. A chloroform solution of EYPC (10 mg/ml, 500 ml) is evaporated to remove the solvent. The obtained thin lipid membrane (5.0 mg, 6.25 mmol) is further dried under vacuum for at least 2 h, and then dispersed in 0.5 ml of 63 mM calcein aqueous solution (pH 7.4) with sonication for 3 min. The liposome suspensions are freeze-thawed four times. The obtained liposome suspension is extruded through a polycarbonate membrane with a pore diameter of 100 nm. Free calcein is removed by gel permeation chromatography using a

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A

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7305

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Figure 7.4 Intensity-weighted distributions of the intact liposomes (A) and the complexes of liposomes with Au NPs at the EYPC/Au ratio of 10/1 (B) in PBS after the 6 h-incubation by DLS (from Kojima et al., 2008).

Sepharose 4B column and PBS (10 mM Na2HPO3–NaH2PO3, 150 mM NaCl, pH 7.4). The lipid concentrations are estimated by Phospholipids C-Test Wako (Wako Pure Chemical Industries Ltd.) according to the manufacturer’s instructions, as described earlier. An aliquot of the calceinloaded liposome dispersion is added to 3 ml of PBS containing 0.5 mM ethylenediaminetetraacetic acid (EDTA, Kishida Chemical); the final lipid concentration is 0.2 mM. The fluorescence intensity of the solution is monitored using a spectrofluorometer ( Jasco Inc., FP-6500) at excitation and monitoring wavelengths of 480 and 515 nm, respectively. Measurements are taken before and after the addition of Au NPs to the liposome

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suspension at an EYPC/Au ratio of 1/10. The amount of calcein remaining in the liposome is estimated from Eq. (7.1). 0

Ft  Ft  100 Fluorescence intensity ð%Þ ¼ 0 F0  F0

ð7:1Þ

where Ft and F0 refer to the fluorescence at t and 0 h after the addition of the Au 0 0 NP dispersion or PBS, and Ft and F0 refer to the fluorescence of these samples with the addition of 10% Triton X-100 solution (Kishida Chemical, final concentration 0.03%) at t and 0 h. The fluorescence intensity (%) of calcein for the intact liposome decreased after the 24-h incubation (Fig. 7.5). This suggests that the calcein absorbed on the liposome membrane might be released. The fluorescence intensity of calcein after the addition of Au NPs at an EYPC/Au ratio of 1/10 was the same as that in the intact liposome, suggesting that adding Au NPs did not promote calcein release. This implies that the liposomal membrane remained intact after interaction with Au NPs.

7. Estimation of Numbers of the Au NP and the Liposome in the Complexes

Fluorescence intensity (%)

As described earlier, complexed Au NPs at an EYPC/Au ratio of 10/1 were stably dispersed. However, complexes at ratios of 1/1 and 1/10 were not. The particle numbers for Au NPs and liposomes in the complexes can be calculated. A liposome of 200 nm in diameter was determined to contain 3.5  105 EYPC molecules, from Eq. (7.2).

100 80 60 40 20 0

0

5

10 15 20 Incubation time (h)

25

Figure 7.5 Time-dependent fluorescence intensity (%) of calcein in the liposomes with (open symbols) and without (closed symbols) of Au NPs is shown (from Kojima et al., 2008).

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4pr 2  2 ð7:2Þ A where N, r, and A refer to the number of lipids, the radius of the liposomes, and the section area of the EYPC head group (0.717 nm2), respectively (Lasic, 1993). Whereas an Au NP of 13 nm in a diameter is composed of 6.8  104 gold atoms, as determined from Eq. (7.3).    3 2 D ð7:3Þ U¼ p 3 a N¼

where U, D, and a refer to the number of Au atoms, the diameter of the Au NPs, and the edge length of a unit cell (0.40786 nm), respectively (Chithrani et al., 2006). From these calculations, the respective numbers of Au NPs and EYPC liposome in the complex at an EYPC/Au ratio of 10/1 are estimated to be 7.8  1011 and 4.0  1011, under our conditions with 45.5 nmol Au ion and 455 nmol EYPC lipid. Therefore, an EYPC liposome might interact with approximately one Au NP. Overall, these results show that the complexes at an EYPC/Au of 10/1 had a particle ratio of approximately one-to-one and were stably dispersed without disturbing the liposome structure. The enhancement of affinity between the liposome and the Au NPs is of significance for the application.

8. Optimization of Lipid Components of the Complexes As the affinity of liposome to Au NPs should be affected by the lipid component of the liposome, complexes are prepared using both cationic and PEG-modified liposomes. It is expected that cationic lipids and PEG will interact with anionic Au NPs more efficiently. These liposomes are prepared using DDAB (Tokyo Chemical Industry Co. Ltd., Tokyo, Japan) and PEG-PE (Avanti Polar Lipids Inc., Alabaster, AL), in addition to EYPC (Fig. 7.6B and C). DDAB-containing liposomes (10 mol% DDAB (0.625 mmol) and 90 mol% EYPC (5.625 mmol)) are prepared according to the same preparation method of the EYPC liposome before the sonication. The liposomes are incubated at 4
C for 1 day, followed by sonication for 10 min. The size distribution of this liposome preparation is determined to be approximately 200 nm by DLS. A PEG-PE-containing liposome with a diameter of 200 nm and composition of 5 mol% PEG-PE (0.31 mmol) and 95 mol% EYPC (5.94 mmol) is also prepared, according to the same procedure as the EYPC liposomes. The concentration of these lipids is

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O-

+

O

CH−O−C−R −

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−

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−−

CH3

−

Br−

−

B

CH−O−C−(CH2)16CH3 =

CH2−O−C−(CH2)16CH3 O

Figure 7.6 Structures of EYPC (A), DDAB (B), and PEG-PE (C).

estimated from the measured EYPC concentration, considering the in feed ratios. The DDAB- and PEG-bearing liposomes also inhibited the decrease of the SPR signal of the Au NPs, with a dependence on the ratio of lipid to Au (Fig. 7.7). The stability of the SPR signal for these liposomes was similar to the EYPC liposomes. This suggests that complex formation was not improved by DDAB and PEG-PE. Complex formation using liposomes with alkanethiol and different molecular weight preparations of PEG-PE may also be investigated. Liposomes are prepared using C18-SH (Tokyo Chemical Industry Co. Ltd.) and PEG2000-PE or PEG5000-PE (NOF Corp.). PEG-PE- and C18-SHcontaining liposomes with diameters of 100 nm and composition of 5 mol% PEG2000-PE or PEG5000-PE (0.31 mmol), 15 mol% of C18-SH (0.93 mmol), and 80 mol% of EYPC (5.00 mmol) is prepared, according to the same procedure as the EYPC liposome minus the reduction. Before the addition of Au NPs, the liposomes are reduced by reacting with dithiothreitol (DTT; Wako Pure Chemical Industries Ltd.) for 1 h, followed by dialysis using degassed PBS (10 mM Na2HPO3–NaH2PO3, 150 mM NaCl, pH 7.4). The PEG2000- and PEG5000-bearing liposome complexes containing C18-SH inhibited efficiently the decrease of the Au NP SPR signal in the time-dependent SPR absorption spectra, even at a lipid/Au ratio of 1/10 (Fig. 7.8). The PEG5000-PE-bearing liposome was the most stably dispersed complex, suggesting that both the alkanethiol bound to the surface of the Au NPs and PEG play a role in good dispersal of the complex.

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0 400

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Figure 7.7 Time-dependent UV–Vis spectra of the complex of (A, C, E) DDAB- or (B, D, F) PEG-bearing liposomes with Au NPs at different ratios. The mole ratio of lipid to Au is (A, B) 10/1, (C, D) 1/1, and (E, F) 1/10. [Au] ¼ 46 mM.

9. Concluding Remarks We have described the preparation of various complexes of dispersions of liposomes and Au NPs. Liposomes improve the stability of Au NP dispersions under isotonic conditions. The complexes are formed without disturbing the liposome structure. In addition, complex formation is enhanced by using PEG- and thiol-modified liposomes. It is known that

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800

B 0.2 PEG-5000

Abs

0.15

1h 4h 8h 12 h

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Figure 7.8 Time-dependent UV–Vis spectra of the complex of (A) PEG-2000- or (B) PEG-5000-bearing liposomes containing alkanethiol with Au NPs at the lipid/Au of 1/10. [Au] ¼ 46 mM.

PEG-bearing liposomes are used widely as a drug carrier due to the biocompatibility and the prolonged blood circulation of the liposomes (Greenwald et al., 2000). These types of complexes have potential diagnostic and therapeutic applications in nanomedicine.

ACKNOWLEDGMENT This work was supported by Grants-in-Aid from the Ministry of Education, Culture, Sports, Science and Technology of Japan (MEXT).

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REFERENCES Al-Jamal, W. T., and Kostarelos, K. (2007). Liposome-nanoparticle hybrids for multimodal diagnostic and therapeutic application. Nanomedicine 2, 85–98. Burda, C., Chen, X., Narayanan, R., and El-Sayed, M. A. (2005). Chemistry and properties of nanocrystals of different shapes. Chem. Rev. 105, 1025–1102. Chilkoti, A., Dreher, M. R., Meyer, D. E., and Raucher, D. (2002). Targeted drug delivery by thermally responsive polymers. Adv. Drug Deliv. Rev. 54, 613–630. Chithrani, B. D., Ghazani, A. A., and Chan, W. C. W. (2006). Determining the size and shape dependence of gold nanoparticle uptake into mammalian cells. Nano Lett. 6, 662–668. Daniel, M. C., and Astruc, D. (2004). Gold nanoparticles: Assembly, supramolecular chemistry, quantum-size-related properties, and applications toward biology, catalysis, and nanotechnology. Chem. Rev. 104, 293–346. Govorov, A. O., and Richardson, H. H. (2007). Generating heat with metal nanoparticles. Nanotoday 2, 30–38. Grabar, K. C., Freeman, R. G., Hommer, M. B., and Natan, M. J. (1995). Preparation and characterization of Au colloid monolayers. Anal. Chem. 67, 735–743. Greenwald, R. B., Conover, C. D., and Choe, Y. H. (2000). Poly(ethylene glycol) conjugated drugs and prodrugs: A comprehensive review. Crit. Rev. Ther. Drug Carrier Syst. 17, 101–161. Haba, Y., Kojima, C., Harada, A., Ura, T., Horinaka, H., and Kono, K. (2007). Preparation of poly(ethylene glycol)-modified poly(amidoamine) dendrimers encapsulating gold nanoparticles and their heat-generating ability. Langmuir 23, 5243–5246. Hayashi, H., Kono, K., and Takagishi, T. (1998). Temperature-dependent associating property of liposomes modified with a thermosensitive polymer. Bioconjug. Chem. 9, 382–389. Hong, K., Friend, D. S., Glabe, C. G., and Papahadjopoulos, D. (1983). Liposomes containing colloidal gold are a useful probe of liposome-cell interactions. Biochim. Biophys. Acta 732, 320–323. Immordino, M. L., Dosio, F., and Cattel, L. (2006). Stealth liposomes: Review of the basic science, rationale, and clinical applications, existing and potential. Int. J. Nanomed. 1, 297–315. Jain, P. K., El-Sayed, I. H., and El-Sayed, M. A. (2007). Au nanoparticles target cancer. Nanotoday 2, 18–29. Kim, D., Park, S., Lee, J. H., Jeong, Y. Y., and Jon, S. (2007). Antibiofouling polymercoated gold nanoparticles as a contrast agent for in vivo X-ray computed tomography imaging. J. Am. Chem. Soc. 129, 7661–7665. Kojima, C., Hirano, Y., Yuba, E., Harada, A., and Kono, K. (2008). Preparation and characterization of complexes of liposomes with gold nanoparticles. Colloids Surf. B 66, 246–252. Kono, K. (2001). Thermosensitive polymer-modified liposomes. Adv. Drug Deliv. Rev. 53, 307–319. Kono, K., and Arshady, R. (2006). Smart Nano and Microparticles. Kentus Books, London. Kono, K., Hayashi, H., and Takagishi, T. (1994). Temperature-sensitive liposomes: Liposomes bearing poly(N-isopropylacrylamide). J. Control. Release 30, 69–75. Kono, K., Nakai, R., Morimoto, K., and Takagishi, T. (1999). Thermosensitive polymermodified liposomes that release contents around physiological temperature. Biochim. Biophys. Acta 1416, 239–250. Lasic, D. D. (1993). Liposomes, from physics to applications. Elsevier, Amsterdam. Li, X., Li, Y., Yang, C., and Li, Y. (2004). Liposome induced self-assembly of gold nanoparticles into hollow spheres. Langmuir 20, 3734–3739.

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Link, S., and El-Sayed, M. A. (1999). Spectral properties and relaxation dynamics of surface plasmon electronic oscillations in gold and silver nanodots and nanorods. J. Phys. Chem. B 103, 8410–8426. Link, S., and El-Sayed, M. A. (2000). Shape and size dependence of radiative, nonradiative, and photothermal properties of gold nanocrystals. Int. Rev. Phys. Chem. 19, 409–453. Paasonen, L., Laaksonen, T., Johans, C., Yliperttula, M., Konttun, K., and Urtti, A. (2007). Gold nanoparticles enable selective light-induced contents release from liposomes. J. Control. Release 122, 86–93. Park, S.-H., Oh, S.-G., Mun, J.-Y., and Han, S.-S. (2006). Loading of gold nanoparticles inside the DPPC bilayers of liposome and their effects on membrane fluidities. Colloids Surf. B 48, 112–118. Pissuwan, D., Valenzuela, S. M., and Cortie, M. B. (2006). Gold nanoparticles as therapeutics. Trends Biotechnol. 24, 62–67. Volodkin, D. V., Skirtach, A. G., and Mohwald, H. (2009). Near-IR remote release from assemblies of liposomes and nanoparticles. Angew. Chem. Int. Ed. Engl. 48, 1807–1809. Wu, G., Mikhailovsky, A., Khant, H. A., Fu, C., Chiu, W., and Zasadzinski, J. A. (2008). Remotely triggered liposome release by near-infrared light absorption via hollow gold nanoshells. J. Am. Chem. Soc. 130, 8175–8177.

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C H A P T E R

E I G H T

Bio-Nanocapsule–Liposome Conjugates for In Vivo Pinpoint Drug and Gene Delivery Takeshi Kasuya,* Joohee Jung,*,1 Rie Kinoshita,† Yasumasa Goh,† Takashi Matsuzaki,* Masumi Iijima,*,‡ Nobuo Yoshimoto,*,‡ Katsuyuki Tanizawa,* and Shun’ichi Kuroda*,†,‡ Contents 1. 2. 3. 4. 5.

Introduction First-Generation Bio-Nanocapsules Second-Generation BNCs Retargeting of BNC–LP Conjugates Overexpression of BNCs in S. cerevisiae 5.1. Purification of BNCs by ultracentrifugation (
10 mg protein per lot) 5.2. Purification of BNCs using column chromatography 5.3. Purification of ZZ–BNC using column chromatography 6. Conjugation of BNCs with LPs 6.1. Example 1.1: Preparation of BNC–LP conjugates containing DNA (BNC–lipoplex conjugates) 6.2. Example 1.2: In vitro transfection with BNC–lipoplex conjugates 6.3. Example 1.3: In vivo transfection with BNC–lipoplex conjugates 6.4. Example 2.1: Preparation of BNC–LP conjugates containing DOX 6.5. Example 2.2: In vitro cytotoxic effects of BNC–LP conjugates containing DOX 6.6. Example 2.3: In vivo therapeutic effects of BNC–LP conjugates containing DOX

* { { 1

148 149 150 152 153 154 154 155 155 156 156 158 158 159 161

Institute of Scientific and Industrial Research, Osaka University, Ibaraki, Osaka, Japan Beacle Inc., ORIC, Haga, Okayama, Japan Graduate School of Bioagricultural Sciences, Nagoya University, Chikusa, Nagoya, Japan Present address: Institute for Innovative Cancer Research, ASAN Medical Center, Pungnap-2, Songpa, Seoul, Korea

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64008-8

#

2009 Elsevier Inc. All rights reserved.

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7. Preparation of Antibody-Displaying BNC–LP Conjugates 8. Preparation of Biotin-Displaying BNC–LP Conjugates 9. Concluding Remarks Acknowledgments References

161 163 163 164 164

Abstract A bio-nanocapsule (BNC) is an  50-nm hepatitis B virus (HBV) subviral particle comprising HBV envelope L proteins and a lipid bilayer, and is synthesized in recombinant Saccharomyces cerevisiae. When BNCs are administered intravenously in a mouse xenograft model, they can accumulate specifically in human liver-derived tissues and enter cells efficiently by the HBV-derived human liverspecific infection machinery, localized at the outer-membrane pre-S region of the L protein. BNC specificity for the human liver can be altered to other tissues by substituting the pre-S region using targeting molecules (e.g., antibodies, lectins, cytokines). BNCs can spontaneously form complexes with liposomes (LPs) by the membrane fusogenic activity of the pre-S region. LPs containing various therapeutic materials (e.g., chemicals, proteins, DNA, RNA) can therefore be covered with BNCs to form an  150-nm BNC–LP conjugate. BNC–LP conjugates injected intravenously can deliver incorporated materials to target tissues specifically and efficiently by utilizing the HBV-derived infection machinery. The stability of BNC–LP conjugates in the blood circulation is similar to that of PEGylated LPs. In this chapter, we describe the preparation and in vivo application of BNC–LP conjugates, and the potential of BNC–LP conjugates as in vivo pinpoint drug delivery systems.

1. Introduction Liposomes (LPs) are one of the most promising drug delivery system (DDS) carriers for genes and drugs, and several LP-based medicines have entered clinical use (Lorusso et al., 2007; Markman, 2006; Richardson, 2006; Rosenthal et al., 2002). Most LP-based medicines are nanoscaled. They are used for the delivery of anticancer drugs to carcinomatous lesions by taking advantage of the enhanced permeation and retention effect of solid tumors (Maeda et al., 2000). The blood vasculature in tumors is leaky, because it possesses larger sized pores than the blood vasculature in healthy tissues. A nanomedicine of diameter 100 nm can therefore readily seep out from the blood circulation and accumulate in tumors (‘‘passive targeting’’). The major obstacle for passive targeting is the immune system in the liver and spleen, that is, the reticuloendothelial system (RES) in which macrophages (including Kupffer cells) capture opsonized micro- and nanoscaled compounds (Moghimi et al., 2001). To escape the RES, recent LPs have been modified with polyethylene glycol (PEG) (i.e., PEGylated LPs),

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thereby enhancing their stability in the blood circulation by avoiding LPs from binding to serum proteins (‘‘opsonization’’) (Lasic and Martin, 1995; Moghimi and Szebeni, 2003). PEGylated LPs containing anticancer drugs could dramatically reduce adverse side effects, but several reports have noted that the LP encapsulation of anticancer drugs contribute only to the reduction of side effects, not to the enhancement of therapeutic effects (Hong et al., 1999; Parr et al., 1997). This indicates that modifying LPs for the delivery of drugs to actively target foci (‘‘active targeting’’) is required. After attachment onto the cell surface, LPs are usually incorporated within target cells through an endocytotic cascade. Most of the drugs in LPs are degraded in late endosome and lysosome without exhibiting sufficient therapeutic effects. In contrast to LPs, several viral vectors (e.g., adenovirus, adeno-associated virus, lentivirus, retrovirus) are widely used in gene delivery. This is because recombinant viral vectors can efficiently pass across the plasma membrane and deliver their genomic information to the cell nucleus as well as escaping from the endocytotic cascade. These viral vectors may have unexpected severe side effects originating from the viral genome (Marshall, 2002; Savulescu, 2001), but the efficiency of gene delivery is much higher than that of lipoplexes (LP–DNA complexes) (Hama et al., 2006). Taken together, the next generation of LP-based nanomedicines must have the following features: (1) viral genome-free structure; (2) active targeting machinery; (3) escape machinery from the RES; (4) escape machinery from the endosomal degradation pathway; and (5) intracellular targeting machinery.

2. First-Generation Bio-Nanocapsules In the process of developing a hepatitis B virus (HBV) vaccine, we succeeded in overexpressing HBV subviral L particles in Saccharomyces cerevisiae (Kuroda et al., 1992). The L particle is an 50-nm hollow capsule comprising HBV envelope L proteins and a lipid bilayer (Yamada et al., 2001). In 2003, we unexpectedly found that the L particles could incorporate genes and drugs by electroporation, and deliver the incorporated materials specifically and efficiently to cell lines (in vitro) and tumors (in vivo) derived from the human liver (Yamada et al., 2003). We therefore designated the HBV L particle as a ‘‘bio-nanocapsule’’ (BNC) (Yu et al., 2005). The hepatophilic and highly efficient delivery properties of BNCs are considered to be based on the infection mechanism of HBV. BNCs and HBV display the N-terminal half of the outer-membrane pre-S region of L protein (Fig. 8.1), which is postulated to have a pivotal role in the human liver-specific attachment of HBV (Kasuya et al., 2008a,b). We succeeded in the continuous expression (>1 month) of the human blood clotting factor

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Bio-nanocapsule (BNC)

Hepatitis B virus (HBV) Core protein Viral genome

DNA polymerase

Hollow (genome free) pre-S

Cellular internalization

Human hepatocytespecific binding site

S

HBV envelope L protein

Figure 8.1 HBV and BNC (schematic).

IX gene in a mouse xenograft model by a single intravenous injection of an electroporated BNC (Yamada et al., 2003). We also succeeded in obtaining a significant reduction in the size of human liver-derived tumors in a mouse xenograft model by a single intravenous injection of an electroporated BNC containing the herpes simplex virus thymidine kinase (HSV-tk) gene and sequential treatment with gancyclovir (an HSV-tk-dependent inhibitor of DNA polymerases) (Iwasaki et al., 2007). Engineered BNCs, of which the human liver-specific attachment site of the pre-S region is changed to other targeting molecules (epidermal growth factor (EGF), anti-EGF receptor (EGFR) monoclonal antibody), have been shown to accumulate in nonhuman liver tissues in vitro and in vivo (Tsutsui et al., 2007; Yamada et al., 2003). This strongly suggests that BNCs are promising in vivo pinpoint DDS carriers targeting not only human liver tissue but also other tissues.

3. Second-Generation BNCs Electroporation was used to transiently induce micropores across the lipid bilayer of BNCs. It was found that BNCs stored for long periods tended to show lower incorporation efficiency than freshly prepared BNCs. The intra- and intermolecular disulfide bonds of L proteins (14 Cys residues in three transmembrane segments of the L protein) were postulated to gradually form in a time-dependent manner, which would make BNCs

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resistant to electroporation. We substituted each Cys residue with Ala or Ser by genetic modification, and identified at least eight Cys residues of the L protein unnecessary for BNC formation (Nagaoka et al., 2007). The BNC harboring the eight Cys ! Ala or Cys ! Ser mutations showed good incorporation efficiency in an electroporation method. This method was not suitable for introducing large materials into a BNC (e.g., >20-kbp plasmid for gene therapy; >10-nm fluorescence-labeled polystyrene beads for bioimaging) and adopting the electroporation method for the good manufacturing practice (GMP)-based production of BNC-based nanomedicines would be very difficult. We recently found that the N-terminal half of the L protein possesses membrane fusogenic activity (Matsuzaki et al., unpublished data). BNCs spontaneously form an 150-nm rigid complex with LPs (BNC–LP conjugate) in which multiple BNCs are embedded on the surface of LPs (Fig. 8.2) ( Jung et al., 2008). This property of BNC allows incorporation of various therapeutic materials into BNC–LP conjugates as follows. First, various materials (even 40-kbp plasmid, 100-nm fluorescent polystyrene beads) are constantly incorporated into LPs by conventional methods. Second, LPs are covered with BNCs harboring tissue specificity and high infectivity by the fusogenic activity of BNC. Third, BNC–LP conjugates can deliver various materials incorporated specifically and efficiently into human liver-derived tumors in a mouse xenograft model

Liposome

Incorporation

DNA

siRNA BNC

Conjugation Protein

Chemical compounds

BNC-liposome conjugate

Figure 8.2

Preparation of BNC–LP conjugates (bars: 50 nm).

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through intravenous injection ( Jung et al., 2008). Additionally, this method can be expanded to GMP-based mass production more readily than electroporation. We therefore designated the BNC–LP conjugate as a ‘‘secondgeneration BNC’’ (Kasuya and Kuroda, 2009). With respect to the attachment of BNC–LP conjugates to human liver cells, the N-terminal half of the pre-S region has as a specific receptor for the human liver, which was demonstrated in HBV and BNCs. The intracellular drug release from BNC–LP conjugates may be mediated by the cellular internalization activity (another fusogenic activity) of the C-terminal half of the pre-S region and S region (see Fig. 8.1), as demonstrated in HBV (Glebe and Urban, 2007). In fact, an engineered BNC lacking the N-terminal half of the pre-S region can form a BNC–LP conjugate and deliver incorporated materials to target cells (Kasuya et al., 2008c). BNC–LP conjugates therefore possess the advantages of LPs and viral vectors, namely the use of versatile materials (as with LP) and the HBV-derived infection and active targeting machinery. These results indicate that BNC–LP conjugates are more promising than BNC per se as in vivo pinpoint DDS carriers.

4. Retargeting of BNC–LP Conjugates Due to the narrow tropism of HBV (because of the function of the N-terminal half of the pre-S region), systemically injected BNCs accumulate specifically in human liver-derived tissues in vivo. For expanding the indications of BNC-based nanomedicines, it is important to establish the methodology for retargeting BNC from human liver to nonliver tissues by substitution of the pre-S region by other targeting molecules (e.g., cytokines, growth factors, receptors, antibodies, glycans, lectins, aptamers). First, the pre-S (3–77) region is replaced with EGF by genetic engineering. The EGF-displaying BNC lost the specificity to human liver cells and obtained new specificity to EGFR-overexpressing A431 cells in vitro (Yamada et al., 2003). This approach needs a time-consuming step for constructing the expression system and sometimes fails to achieve the high productivity of BNCs in yeast cells. Next, a large part (50–159) of the pre-S region is replaced with the Staphylococcus aureus protein A-derived IgG Fc-binding ZZ domain. Beyond expectation, the ZZ domain-displaying BNC (ZZ–BNC) is efficiently synthesized in yeast cells. After ZZ–BNC is mixed with anti-EGFR IgG, the mixture is injected intracranially to a glioma-transplanted mouse orthograft model. The anti-EGFR IgG-displaying BNC accumulated in the EGFR-overexpressing transplanted glioma in the mouse corpus striatum (Kurata et al., 2008; Tsutsui et al., 2007), indicating that the antibody-displaying ZZ–BNC is applicable for

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in vivo use. In 2008, because the ZZ domain contains many Lys residues, ZZ–BNC was modified with N-hydroxysuccinimide (NHS)-biotin to display biotin molecules onto the BNC surface. The biotinylated ZZ– BNC can be used for displaying various biotinylated targeting molecules (see above) by using an avidin (e.g., streptavidin, neutravidin) as an adaptor. For instance, the Phaseolus vulgaris agglutinin-L4 isolectin (L4-PHA)-displaying BNC (PHA–BNC) has been shown to accumulate in vivo in highly metastatic malignant tumors which overexpress b1-6-branching N-acetylglucosamine (GlcNAc), a specific ligand of L4-PHA (Kasuya et al., 2008c). PHA–BNC–LP conjugate can deliver luciferase-expressing plasmid to the b1-6GlcNAc-overexpressing cells, showing that the lack of a large part of the pre-S region does not affect the formation of BNC–LP conjugates. We recently revealed that the fusogenic activity of BNC required for LP conjugation is delineated in the short sequence at the N-terminal of pre-S region, which remains in ZZ–BNC (Matsuzaki et al., unpublished data). These data strongly suggested that the engineered BNCs (ZZ–BNC, biotinylated ZZ–BNC) and their LP conjugates can be used for lesion-specific pinpoint DDS carriers. In this chapter, we describe methods to prepare BNC–LP conjugates containing DNA, and BNC–LP conjugates containing an anticancer drug (doxorubicin (DOX)), and the effects of these BNC–LP conjugates in in vitro and in vivo systems.

5. Overexpression of BNCs in S. cerevisiae BNC is produced in S. cerevisiae AH22R (a leu2 his4 can1 cirþ pho80) strain (Kobayashi et al., 1988). The yeast cells are transformed by the spheroplast method (Hinnen et al., 1978) with the YEp plasmid pGLDLIIP39-RcT (Kuroda et al., 1992), which encodes the N-terminal chicken lysozyme signal sequence-fused HBV envelope L protein (subtype adr) under the glyceraldehyde-3-phosphate dehydrogenase (GLD; also called TDH3) gene promoter. For production of ZZ–BNC, the DNA segment encoding the pre-S region (50–159) in the plasmid pGLDLIIP39-RcT is replaced with the DNA segment encoding the IgG Fc-binding ZZ domain derived from S. aureus protein A (Tsutsui et al., 2007). The LEU2þ yeast transformants are cultured in a synthetic selection medium 8S5N-P400 (Yamada et al., 2001) at 30  C for 7 days, harvested by centrifugation, and stored at 80  C. The amount of BNC in yeast cells is estimated to be 40% of total soluble proteins (Kuroda et al., 1992).

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5.1. Purification of BNCs by ultracentrifugation (
10 mg protein per lot) BNC and ZZ–BNC can be readily purified by ultracentrifugation owing to the overexpression of BNCs in yeast cells. The yeast cells (about 20 g wet weight) are disrupted with glass beads (0.5 mm in diameter, 175 ml) using a BEAD-BEATER (BioSpec Products, Bartlesville, OK, USA) in 160 ml of 0.1 M sodium phosphate buffer (pH 7.2) containing 7.5 M urea, 15 mM ethylenediaminetetraacetic acid (EDTA), 4 mM phenylmethylsulfonyl fluoride (PMSF), 0.1 mM 4-amidinophenyl-methanesulfonyl fluoride (APMSF), and 0.1% (v/v) Tween 80. The crude extract obtained by centrifugation at 34,780g at 4  C for 30 min (about 6 g of protein) is mixed with PEG 6000 solution (15%, w/v at final concentration) to precipitate BNCs. The precipitants (about 3 g of protein) are subjected to CsCl isopycnic ultracentrifugation (10–40%, w/v) using an SW28 rotor (Beckman Coulter, Inc., Fullerton, CA, USA) at 24,000 rpm at room temperature for 15 h. Fractions containing BNCs are determined by sandwich enzyme-linked immunosorbent assay (ELISA) for BNC using an IMx HBsAg assay system (Abbott Laboratories, Abbott Park, IL, USA). Positive fractions are subjected twice to sucrose density gradient ultracentrifugation (10–50%, w/v) using an SW28 rotor at 24,000 rpm at room temperature for 15 h. Fractions containing BNCs are concentrated by ultrafiltration using an Amicon Ultra 100,000 NMWL (Millipore, Billerica, MA, USA) and subjected to a Sephacryl S-500 HR (GE Healthcare, Waukesha, WI, USA) gel-filtration column equilibrated with phosphate-buffered saline (PBS) containing 1 mM EDTA. The protein concentration of BNCs is measured by a bicinchoninic acid (BCA) assay kit (Sigma-Aldrich, St Louis, MO, USA) using bovine serum albumin (BSA) as a calibration standard. Approximately 10 mg (as a protein) of BNC is obtained from recombinant yeast cells grown in 2 l of culture medium (Yamada et al., 2001).

5.2. Purification of BNCs using column chromatography The ultracentrifugation step is rate-limiting, time-consuming, and produces a low yield, so obtaining >10 mg of BNC per lot in 1 week is difficult. Based on the heat stability of BNCs (Yamada et al., 2001), a large amount of yeast crude extract can be processed immediately by heat treatment ( Jung et al., unpublished data). The crude extract of yeast cells is obtained by the method described earlier, and then dialyzed three times against PBS containing 1 mM EDTA at 4  C for 2 h to remove urea. The crude extract (about 6 g of protein) is dispensed to 40-ml plastic tubes, incubated at 70  C for 20 min, and then centrifuged at 34,780g at 4  C for 30 min to remove yeast-derived proteins. The supernatant (about 800 mg of protein) is subjected to a sulfate-cellulofine column (1.6  20 cm; Chisso Corp., Tokyo,

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Japan) equilibrated with PBS containing 150 mM NaCl. After the stepwise elution of PBS containing 1 M NaCl, the fraction containing a BNC is concentrated by ultrafiltration using an Amicon Ultra 100,000 NMWL (about 120 mg of protein) and then subjected to a Sephacryl S-500 HR gel-filtration column (1.6  60 cm) equilibrated with PBS containing 1 mM EDTA. About 60 mg of the highly purified BNC is obtained from the yeast cells grown in 2 l of culture medium.

5.3. Purification of ZZ–BNC using column chromatography Porcine IgG is precipitated from porcine serum (Sigma-Aldrich) with (NH4)2SO4 (40% saturation), dissolved in PBS, and dialyzed against PBS at 4  C for 48 h. The dialyzed solution is mixed with two-times volume of 60 mM acetate buffer (pH 4.8) and then mixed with n-caprylic acid to achieve a final concentration of 6.8% (v/v). After incubation at room temperature for 30 min with gentle stirring, the supernatant containing purified IgG is obtained by brief centrifugation. Purified porcine IgG is conjugated to an NHS-activated Sepharose 4B Fast Flow (GE Healthcare) column according to manufacturer’s instructions. The IgG-conjugated Sepharose column is equilibrated with PBS. Preparation of the crude extract of yeast and heat treatment are the same as the BNC protocol. The supernatant is subjected to the porcine IgGconjugated Sepharose column (affinity chromatography) to capture ZZ– BNC specifically using the IgG–ZZ domain interaction. The column is washed extensively with 75 mM Tris–HCl (pH 7.2) containing 10 mM NaCl, and then ZZ–BNC is obtained by the stepwise elution of 10 mM Tris–HCl (pH 7.2) containing 3.5 M NaSCN, 500 mM NaCl, and 10 mM EDTA. Fractions containing ZZ–BNC are subjected to a Sephacryl S-500 HR gel-filtration column (1.6  60 cm) equilibrated with PBS containing 1 mM EDTA. About 30 mg of highly purified ZZ–BNC can be obtained from yeast cells grown in 2 l of culture medium.

6. Conjugation of BNCs with LPs The LPs used for BNC conjugation (Fig. 8.2) can be prepared by conventional methods such as solvent evaporation (Bangham and Horne, 1964), ethanol injection (Batzri and Korn, 1973), and reverse-phase evaporation (Szoka and Papahadjopoulos, 1978). The lipid composition of LPs should be optimized for the materials to be incorporated, for example, cationic lipids for the preparation of DNA- or siRNA-containing LPs (i.e., lipoplexes; Chapter 14 of volume 465, Du¨zgu¨nes¸ et al., 2002; Li and Huang, 2006), and anionic/neutral lipids for chemical compounds

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containing LPs. To our knowledge, all LPs that have been employed have resulted in the formation of BNC–LP conjugates.

6.1. Example 1.1: Preparation of BNC–LP conjugates containing DNA (BNC–lipoplex conjugates) For preparation of a DNA–LP complex (lipoplex), we routinely use commercially available lyophilized cationic LPs containing O,O0 -ditetradecanoyl-N-(a-trimethylammonioacetyl)diethanolamine chloride (DC-6-14; Kikuchi et al., 1999) as the major cationic lipid, an essential component for the cellular uptake of DNA, for example, Coatsoame-EL-01-D (NOF, Tokyo, Japan) and LipoTrust series (Hokkaido System Science, Sapporo, Japan). In the case of Coatsome-EL-01-D, 1.5 mg of lyophilized LP (as lipids) is dissolved in 1 ml of 250 mg/ml luciferase (LUC) expression vector pGL3 (Promega, Madison, WI, USA) and incubated for 15 min to allow lipoplex formation. Aliquots of lipoplex (100 mg LP, 16.7 mg DNA) are mixed with freeze-dried BNC (100 mg as protein) and incubated at room temperature for 15 min. By changing the amount of LP used for 16.7 mg DNA, a series of lipoplex and BNC–lipoplex conjugates are prepared with an N/P ratio (molar ratio of nitrogen-atom content in cationic lipids to phosphorous-atom content in plasmid DNA) from 0.3 to 2.4. The z-averaged sizes and z-potentials of lipoplex and BNC–lipoplex conjugate are measured at 25  C using a Zetasizer Nano-ZS (Malvern Instruments Ltd., Worcestershire, UK). When the content of cationic lipids in lipoplex is increased, z-averaged sizes and z-potentials are apt to increase to >1 mm and >0 mV, respectively, in accordance with the increase in N/P ratio (Fig. 8.3A). BNCs are found to keep the sizes of BNC–lipoplex conjugates to <200 nm and z-potentials negative.

6.2. Example 1.2: In vitro transfection with BNC–lipoplex conjugates About 1  105 of human cells (hepatocarcinoma HepG2, colon carcinoma WiDr, cervical carcinoma HeLa) are seeded on a 24-well cell culture plate (Iwaki, Tokyo, Japan) and incubated for 24–48 h in a humidified atmosphere at 37  C in 5% (v/v) CO2. Aliquots of BNC–lipoplex conjugates containing pGL3 (N/P ratio ¼ 1.4; 100 mg LP, 16.7 mg DNA, and 100 mg BNC (as protein)) are diluted with complete cell culture medium containing 10% (v/v) fetal bovine serum and antibiotics, and applied to each well with 1 mg/ml (as protein of BNC, 0.5 mg/well) at final concentration. The medium is exchanged after incubation for 3–6 h to avoid nonspecific cellular uptake, and then incubated for 48 h to allow LUC expression.

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RLU (%)

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1 1.5 N/P ratio

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HepG2

HeLa

WiDr

Figure 8.3 Effect of BNC conjugation on lipoplex (DNA-containing LPs). (A) z-averaged sizes (upper panel) and z-potentials (lower panel) of BNC–lipoplex conjugates (circles with solid line) and lipoplex (squares with dashed line) at the indicated N/P ratio. (B) In vitro transfection with LUC expression plasmid by BNC–lipoplex (closed bars) and lipoplex (open bars). LUC activities in cell lysates are measured 48 h after transfection. Mean  S.D. (n ¼ 9). *p < 0.05.

After lysis with passive lysis buffer (Luciferase Assay Kit, Promega), intracellular activities of LUC can be measured by a GloMax 20/20n Luminometer (Promega) using the LUC substrate luciferin. Human liverderived HepG2 cells transfected with BNC–lipoplex conjugates containing pGL3 showed approximately five times higher LUC expression than those transfected with lipoplex containing pGL3, whereas human nonliverderived HeLa cells and WiDr cells showed no significant differences in LUC expression between BNC–lipoplex conjugates and lipoplexes (Fig. 8.3B). These data indicated that the BNC–lipoplex conjugates deliver the LUC plasmid to human liver cells specifically in vitro. This prompted us to examine if BNC–lipoplex conjugates are more suitable than conventional lipoplexes for an in vivo pinpoint gene delivery system.

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6.3. Example 1.3: In vivo transfection with BNC–lipoplex conjugates Xenograft mouse models bearing tumors on their backs are generated using nude mice (5 weeks, male, BALB/c-nu/nu; CLEA Japan, Tokyo, Japan) under guidelines of the Institute of Scientific and Industrial Research, Osaka University, Japan. The cancer cell lines used are NuE and Huh-7 (human hepatocarcinoma), WiDr (human colon carcinoma), A431 (human epithelial carcinoma), PC12 (rat adrenal pheochromocytoma), and MDA-MB435 (human breast carcinoma). About 1  106 cells are mixed with 50 ml of BD Matrigel Matrix HC (BD Biosciences, Bedford, MA, USA) and subcutaneously injected into the back of mice. After 2–4 weeks, mice possessing a tumor of appropriate diameter 1 cm can be obtained. For the delivery of green fluorescent protein (GFP)-expression plasmid pEGFP-C1 (Clontech, Mountain View, CA, USA), BNC–lipoplex (N/P ratio ¼ 1.4; 100 mg LP, 16.7 mg pEGFP-C1, and 100 mg BNC (as protein)) are intravenously injected to each tumor-bearing mouse. At 1 week after injection, mice are killed, and tumors and tissues (liver, lung, spleen, brain, heart, kidney) are isolated. Tissues and tumors are embedded in plastic resin or cryomold, cut into sections of 5 mm thickness, and observed using fluorescent microscopy or confocal laser scanning microscopy. GFP expression can be observed in only human liver-derived tumors, not in other tumors and tissues ( Jung et al., 2008).

6.4. Example 2.1: Preparation of BNC–LP conjugates containing DOX Stock solutions of dipalmitoylphosphatidylcholine (DPPC, NOF), dipalmitoylphosphatidylethanolamine (DPPE, NOF), dipalmitoylphosphatidylglycerol sodium (DPPG-Na, NOF) are prepared in a chloroform/methanol (2:1, v/v) mixture at a concentration of 50 mM. A stock solution of cholesterol (Chol, NOF) is prepared in the same chloroform/methanol mixture at a concentration of 200 mM. All stock solutions are stored at 80  C and prewarmed to 60  C before use. Phospholipids and cholesterol (DPPC:DPPE:DPPG-Na:Chol ¼ 15:15:30:40 molar ratio) are dissolved in the chloroform/methanol mixture in a round-bottomed flask, and then evaporated at 60  C by utilizing a rotary evaporator to produce a thin hemispherical lipid film. The film is hydrated in buffer containing 10 mM HEPES (pH 4.0) and 120 mM (NH4)2SO4 at 60  C. The freeze–thaw cycle is repeated five times. The crude solution of LP is subjected to a Lipex extruder (Northern Lipids, Vancouver, BC, Canada) equipped with a polycarbonate filter (pore size, 200 nm) at 60  C five times, followed by seven times with a filter (pore size, 50 nm). The z-average size of LP is

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measured by dynamic light scattering at 25  C using a Zetasizer Nano-ZS. The LPs are subjected to a Sephadex G-25 (GE Healthcare) gel-filtration column equilibrated with 10 mM HEPES buffer (pH 7.4) containing 100 mM NaCl and 3.4% (w/v) sucrose. Total lipid concentration is calculated from that of DPPC, which is determined by a Laboassay Phospholipid Kit (Wako, Osaka, Japan) utilizing choline oxidase and [N-ethyl-N-(2hydroxy-3-sulfopropyl)-3,5-dimethoxyaniline] (DAOS) (Takayama et al., 1977). To introduce DOX (also called adriamycin; Wako) into LPs by a remote-loading method, 200 ml of 10 mg/ml DOX–HCl solution is added to 15 mg of LP solution (as total lipids, prewarmed at 60  C), and then incubated for 20 min at 60  C with gentle stirring. After free DOX is removed by a Sephadex G-25 gel-filtration column, LPs containing DOX (about 2 mg DOX–HCl per 15 mg total lipid) can be obtained. Aliquots of LPs containing DOX (2 mg LP, 0.27 mg DOX–HCl) are gradually added to freeze-dried BNC (100 mg as protein) and incubated at room temperature for 15 min to form a BNC–LP conjugate containing DOX. The diameter of the conjugate is estimated to be 140 nm by the dynamic laser scattering method.

6.5. Example 2.2: In vitro cytotoxic effects of BNC–LP conjugates containing DOX About 5  103 cells (HepG2, Huh-7, MDA-MB-435) cultured on a 96well plate for 24–48 h are incubated with BNC–LP conjugates containing DOX at appropriate concentration (1–100 mg/ml as DOX) for 6–24 h and then incubated for 48–72 h in fresh complete medium. Cellular viability can be measured by the formation of formazan from a 3-(4,5-dimethylthiazol2-yl)-2,5-diphenyltetrazolium bromide (MTT), a tetrazole probe using a CellTiter 96 Aqueous Non-Radioactive Cell Proliferation Assay (Promega). The strong cytotoxicity of BNC–LP conjugates containing DOX, comparable to that of DOX itself, can be observed in only human liverderived HepG2 and Huh-7 cells, not in human nonliver-derived MDAMB-435 cells (Fig. 8.4A). When comparing the cytotoxicity of BNC–LP conjugates with that of LP, BNC–LP conjugates showed strong cytotoxicity only toward HepG2 cells and Huh-7 cells, whereas LP did not show severe cytotoxicity to any cells. The half maximal inhibitory concentration (IC50) of BNC–LP containing DOX for Huh-7 cells is 12 mg/ml, whereas that of LP containing DOX is 100 mg/ml. Doxil, a commercial PEGylated LP containing DOX (Markman, 2006), also showed a cytotoxicity curve similar to that of LP containing DOX. These data indicated that BNCs confer a human liver-specific active targeting function to LPs containing DOX.

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Figure 8.4 Effect of BNC conjugation on LP containing DOX. (A) In vitro cytotoxicity curves of BNC–LP conjugates. BNC–LP conjugates containing DOX (triangles with gray lines), LP containing DOX (squares with solid lines), DOX alone (diamonds with solid lines), and doxil (crosses with dashed lines). HepG2 cells (left panel), Huh-7 cells (center panel), and MDA-MB-435 cells (right panel). Mean  SD (n ¼ 9). (B) In vivo tumor suppression by BNC–LP conjugates. BNC–LP conjugates containing DOX (triangles with gray lines), DOX alone (diamonds with solid lines), and untreated control (crosses with dashed lines). Tumor volume (left panel) and body weight (right panel). Mean  S.D. (n ¼ 9).

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6.6. Example 2.3: In vivo therapeutic effects of BNC–LP conjugates containing DOX To the mouse xenograft model bearing Huh-7-derived tumors (about 1 cm in diameter), about 6 mg/kg (as DOX–HCl) of BNC–LP conjugates containing DOX is intravenously injected every 4 days under 10 ml/kg conditions. Largest (a) and smallest (b) superficial diameters of the tumor are measured every day, and tumor volume (V ) is calculated as V ¼ ab2/2. BNC–LP conjugates containing DOX efficiently suppressed tumor growth, but identical amounts of DOX injected by LP only and injected alone failed to suppress it (Fig. 8.4B). As for toxicity, mice given about 8 mg/kg (as DOX–HCl) every 4 days by a BNC–LP conjugate showed less loss of body weight than those by DOX alone. A significant change in weight was not observed in tissues (liver, spleen, kidney, heart) 12 days after the first injection (data not shown). These data indicated that BNC–LP conjugates containing DOX possess not only active targeting machinery but also less toxicity. When a mouse xenograft model bearing Huh-7-derived tumors received a single intravenous injection of BNC–LP conjugate containing DOX, LP containing DOX, doxil (PEGylated LP containing DOX), and DOX alone (8 mg/kg as DOX), blood is collected from the tail vein at 0, 30, 60, 120, and 180 min after the first injection. Plasma is collected by centrifugation at 3000 rpm at 4  C, and DOX is extracted with a five volumes of the chloroform/methanol (4:1, v/v) mixture. After evaporation of solvent with nitrogen gas, the dried DOX fraction is subjected to high-performance liquid chromatography (HPLC) equipped with an octylsilanized gel column (Nucleosil 100 5C-18, Chemco Scientific Co. Ltd., Osaka, Japan) equilibrated with a 20 mM PBS/methanol mixture (25:75, v/v) containing 40 mM sodium heptane sulfonate. Flow rate is 1 ml/min, and DOX-derived fluorescence (excitation, 475 nm; emission, 554 nm) is determined using purified DOX as the internal standard. The initial concentration of DOX (0 min) in plasma is about 100 mg/ml in all mice (Fig. 8.5). The plasma concentration of DOX after 60 min in mice injected with a BNC–LP conjugate containing DOX is much higher than those with LP containing DOX and DOX alone, which is comparable to those with doxil. BNC conjugation, like PEGylation, enhances the robustness of LP against the RES.

7. Preparation of Antibody-Displaying BNC–LP Conjugates ZZ–BNC–LP conjugates are prepared using ZZ–BNC instead of BNC by the method described earlier for in vitro and in vivo retargeting of BNC–LP conjugates according to antibody affinity. LP conjugation must

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be done before displaying antibodies on the surface of ZZ–BNC because antibodies displayed onto ZZ–BNC significantly reduce the fusogenic activity of ZZ–BNC (presumably by increased steric hindrance). Briefly, an aliquot of LPs (2 mg LP) containing materials of interest is gradually added to freeze-dried ZZ–BNC (100 mg as protein). We routinely add the smaller amount of antibody than ZZ–BNC (as protein) ranging from 1/5 to 1/100 (as protein) to the ZZ–BNC–LP solution. Antibodies may be spontaneously aligned on the surface of ZZ–BNC–LP conjugates by interaction of the IgG Fc domain with the ZZ domain. In the case of displaying IgG harboring weak affinity to the ZZ domain (Bjo¨rck and Kronvall, 1984), a membrane-impermeable amine-reactive cross-linking agent such as bis(sulfosuccinimidyl)suberate (BS3; Thermo Fisher Scientific, Waltham, MA, USA) was used to cross-link the binding between the IgG Fc domain and ZZ–BNC. Five millimolar BS3 dissolved in dimethylsulfoxide (DMSO) is added to the ZZ–BNC–LP displaying IgG to a concentration of 50 mM, allowed to react for 1 h, and free BS3 removed by the reaction with 0.1 mM glycine. The antibody-displaying ZZ–BNC–LP conjugate is ready for in vitro and in vivo use. To our knowledge, it is not necessary to display the antibodies as much as possible onto ZZ-BNC for maximizing the delivering ability of ZZ-BNC-LP conjugate, strongly suggesting that the amounts of antibody on ZZ-BNC should be optimized in in vitro system.

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8. Preparation of Biotin-Displaying BNC–LP Conjugates For in vitro and in vivo retargeting of BNC–LP conjugates by various biorecognition molecules (e.g., cytokines, peptides, lectins, glycans), the surface of BNCs should be modified with these molecules without affecting fusogenic activity. The ZZ domain displayed on ZZ–BNC possesses many Lys residues, so the e-amino residues are modified with biotin (the smallest high-affinity tag) using sulfo-biotin-NHS-ester (Thermo Fischer Scientific) according to the manufacturer’s protocol. The number of biotins displayed on the surface of ZZ–BNC can be measured utilizing a 40 -hydroxyazobenzene-2-carboxylic acid (HABA) assay kit (Thermo Fisher Scientific). Usually, 1 mol of ZZ–BNC is modified with about 35 mol of biotin. Next, biotinylated ZZ–BNC–LP conjugate is mixed with an equimolar amount of biotinylated biorecognition molecules and avidins (e.g., avidin, streptavidin, neutravidin) to display the biorecognition molecules onto ZZ–BNC. As described in the antibody-displaying ZZ–BNC, the display of biorecognition molecules should be done after formation of the BNC–LP conjugate. The steric hindrance of biotinylated biorecognition molecules and avidins is too large for ZZ–BNC to exhibit fusogenic activity. The biorecognition molecules displaying the ZZ–BNC–LP conjugate is ready for in vitro and in vivo use. To our knowledge, it is not necessary to maximize the biorecognition molecules on ZZ–BNC to maximize the delivery capacity of the ZZ–BNC–LP conjugate, strongly suggesting that the amounts of biorecognition molecules on ZZ–BNC should be optimized in in vitro systems.

9. Concluding Remarks The studies and methods described in this chapter demonstrate that the BNC–LP conjugate, as a hybrid of a viral vector and LPs, is a promising and rational approach to achieving a pinpoint delivery system for drugs and genes in vivo. Two major problems associated with BNC–LP conjugates must be resolved before they are in clinical use. The first problem is the immunogenicity of BNC–LP. BNC was initially developed as an immunogen of a recombinant hepatitis B vaccine, so BNC per se sometimes elicits an unexpected level of immune response. Recently, we succeeded in reducing the immunogenicity by incorporating HBV escape mutants, which can proliferate in humans vaccinated with hepatitis B vaccine. The second problem is the nonspecific incorporation by the RES. Like other nanoparticle-based medicines, BNC–LP conjugates cannot fully escape from the

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RES, whereas HBV can accomplish the escape. We must analyze the endogenous escape mechanism of HBV at the molecular level, and introduce it to BNC–LP conjugates.

ACKNOWLEDGMENTS The authors thank Professor M. Seno (Okayama University), Professor A. Kondo (Kobe University), and Professor M. Ueda (Keio University) for their helpful advice. We are grateful to Ms. Y. Matsushita, Ms. N. Shikaku, and Ms. Y. Matsui for their technical support. This work was supported in part by the Regional Research and Development Resources Utilization Program from the Japan Science and Technology Agency.

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Nanoliposomal Dry Powder Formulations Gaurang Patel,* Mahavir Chougule,† Mandip Singh,† and Ambikanandan Misra* Contents 1. Introduction 2. Preparation of Nanoliposomal DPFs 2.1. Ternary mixture 2.2. Spray-drying 2.3. Freeze-drying 2.4. Spray freeze-drying 3. Physicochemical Characterization of NLDPFs 3.1. Assay 3.2. Flow behavior 3.3. Moisture content determination 3.4. Reconstitution time and volume 3.5. Particle size of NLDPFs 3.6. Drug retention and stability studies 3.7. Scanning electron microscopy (photomicrographs and image analysis) 3.8. Differential scanning calorimetry 3.9. In vitro lung deposition studies 3.10. In vitro drug release study 3.11. In vivo studies 4. Concluding Remarks References

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Abstract Liposomal dry powder formulations (DPFs) have proven their superiority over conventional DPFs due to favorably improved pharmacokinetics and pharmacodynamics of entrapped drugs, and thus, reduced local and systemic toxicities. * {

TIFAC-CORE in NDDS, Pharmacy Department, Faculty of Technology and Engineering, The Maharaja Sayajirao University of Baroda, Kalabhavan, Vadodara, Gujarat, India Pharmaceutics Department, College of Pharmacy and Pharmaceutical Science, Florida A&M University, Tallahassee, Florida, USA

Methods in Enzymology, Volume 464 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)64009-X

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Nanoliposomal DPFs (NLDPFs) provide stable, high aerosolization efficiency to deep lung, prolonged drug release, slow systemic dilution, and avoid macrophage uptake of encapsulated drug by carrier-based delivery of nano-range liposomes. This chapter describes methods of preparation of nanoliposomes (NLs) and NLDPFs, using various techniques, and their characterization with respect to size distribution, flow behavior, in vitro drug release profile, lung deposition, cellular uptake and cytotoxicity, and in vivo pharmacokinetics and pharmacodynamics. Some examples have been detailed for better understanding of the methods of preparation and evaluation of NLDPFs by investigators.

1. Introduction To conquer limitations of pressurized meter dose inhalers (pMDIs) and nebulizers, dry powder inhalers (DPIs) have been developed. DPIs are devices for delivering a dry powder formulation (DPF) of an active drug, including macromolecules and biotechnologicals, for local or systemic therapies (Martin and Anthony, 2005). DPFs are stable, one-phase solid blends, easy to process, very portable, patient-friendly, easy to use, do not require spacers, and have no requirement for coordination of inhalation and actuation (Ashurst et al., 2000; Geller, 2005; Hickey et al., 1994). Other benefits include the capacity to carry a high drug dose, high lung deposition (from 50% to 70%), minimal extra-pulmonary loss of drug due to low oropharyngeal deposition, low device retention, and low exhaled loss. Pulmonary disposition of drugs is associated with lung clearance by alveolar macrophages of the reticuloendothelial system (RES), irrespective of the method used for delivery. This drawback may lead to high drug dosing frequency, causing patient noncompliance (four to six times in some diseases) and increase in dose with time. Liposomal drug delivery to the respiratory tract is particularly attractive, because drugs accumulate locally in lung as the target organ. It is biocompatible (Kimelberg and Mayhew, 1978) as more than 85% of lung surfactant is composed of phospholipids. It provides controlled release of drugs for local or systemic action (Taylor and Newton, 1992), has reduced local/ systemic side effects, and thus improved therapeutic index of the drug and reduced dosing frequency, total dose, and probably cost of therapy (Courrier et al., 2002; Kimelberg and Mayhew, 1978; Konduri et al., 2005; Newman, 2004). The delivery of drugs encapsulated in liposomes in suspension form to the lung using a nebulizer or pMDIs are already under clinical investigation. In suspension form, liposomes have problems such as lipid degradation, aggregation, and fusion of liposomes resulting, in drug leakage during storage and aerosolization to lung (Hu and Rhodes, 1999). If a drug is delivered at the right site in the lung, at the required dose, at

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optimum frequency, and it stays for the required period of time, we can expect optimum therapeutic response of the drug with minimum side effects (Oku and Namba, 2005). Conventional drug DPFs fulfill the first three requisites, but fail in the fourth one. Liposomal drugs enhance the drug residence time in the lungs, prevents enzymatic degradation of the drug, and the nano-size prevents its rapid removal through the clearance mechanism, alleviating the limitations of plain drug or conventional liposomal drug. Drug encapsulated nanoliposomes (NLs) can be processed into DPF form using freeze-drying, spray-drying, and spray freeze-drying to achieve long-term stability, and overcome problems associated with the suspension form of liposomes (Courrier et al., 2002; Niven, 1995).

2. Preparation of Nanoliposomal DPFs Conventional DPFs composed of saccharide carriers containing surface adsorbed micronized drug and having particle diameter in between 3 and 5 mm (Hersey, 1975). The particle size is restricted to 3–5 mm as below 3 mm particles are engulfed by macrophages, and above 5 mm particles show extensive oropharyngeal deposition, which is often associated with widespread systemic side effects (Bisgaard, 1996; Byron and Patton, 1994; Hickey, 1992; Newman and Clarke, 1983). Desirable characteristics of ideal DPFs include enhanced powder performance, higher lung deposition, good flow properties, and efficient delivery of a wide variety of therapeutic agents. Because of including controlled drug release, biocompatibility, biodegradability, and nontoxic nature, liposomes have been extensively researched in pulmonary drug delivery (Shek and Barber, 1986). Formulation flexibility makes liposomal drug delivery more versatile than other lipid-based delivery systems. Methods such as the Bangham method, the organic solvent injection method, and the reverse-phase evaporation method (Betagiri et al., 1993a; New, 1990a) have been commonly used for the preparation of liposomes. Liposomes having a nano-size range can be obtained by passing the liposomal suspension through a high-pressure homogenizer (10,000–25,000 psi) or extrusion through polycarbonate membranes having a pore size of less than about 100 nm at 300–400 bars pressure, and 2–10 cycles above the glass transition temperature of respective lipids of liposomal membrane. Liposomes, lipid- and polymer-based nanoparticles and nanosuspensions can also be effectively formulated using the supercritical fluid technique (SCF) (Kompella and Koushik, 2001; Sunkara and Kompella, 1999). SCF technology offers advantages over the other techniques, such as narrow size distribution, uniform morphology, and assurance of complete removal of organic solvent (Kadimi et al., 2007; Misra et al., 2008). Fixed amounts of drug, phospholipid, and cholesterol are dissolved

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in an organic solvent; the solution thus formed is loaded into a high-pressure reactor maintained below the glass transition temperature and optimized pressure (2200 psi) of supercritical fluid. The sample solution is atomized in the reaction vessel, allowing the supercritical fluid to extract the organic solvent and leave behind nano-sized liposomes (Otake et al., 2001). According to the physicochemical properties of the drug to be encapsulated into liposomes, the method of preparing liposomes is selected and the formulation and process variables are optimized to maximize the percentage of drug entrapment, and to achieve the desired vesicle size and size distribution, lamellarity, and stability. Prepared NLs are separated from unentrapped drug by centrifugation or dialysis. Unentrapped hydrophobic drug is separated at low speed (4000 rpm), and for hydrophilic drug, liposomes are separated by centrifugation at high speed (35,000 rpm). NL pellets are redispersed in aqueous medium. Required quantities of carriers (lactose, sucrose, mannitol, etc.), cryoprotectants (trehalose, mannitol, etc.), and antiadherents (glycine, L-leucine, serine, etc.) are added into drug incorporated nanoliposomal suspension and water is removed by either spray-drying, freeze-drying, or spray freeze-drying to obtain nanoliposomal DPFs (NLDPFs) (Fig. 9.1). Details of the methods for converting NL suspensions in DPF suitable for inhalation are elaborated with examples in this chapter.

2.1. Ternary mixture Freeze-dried NLDPFs are either subjected to jet milling or sieved successively through sieves # 100, 200, 350, and 500. The micronized powder is mixed with small quantity of coarse carrier such as lactose, antiadherent, and lubricant as a ternary component to get NLDPFs having improved flow behavior and enhanced lung deposition. Amphotericin B liposomes are prepared from a mixture of hydrogenated soyphosphatidylcholine; cholesterol and either saturated soyphosphatidylglycerol (7:3:0.5) for negative charge or stearylamine (1:1:0.1) using reverse-phase evaporation technique. The soyphosphatidylglycerol and stearylamine are used to prepare negatively and positively charge liposomes, respectively. Liposomes are extruded from 2 mm polycarbonate membrane to reduce size, separated from unentrapped drug and are lyophilized using sucrose as cryoprotectant. Coarse lactose (Sorbolac 400) in varying mass ratio, with or without addition of fines (500# sieved Pharmatose 325M), is used in different mixing sequence to formulate amphotericin B NLDPFs. We observed that the 10% fine with coarse lactose is found to be optimum blend at 1:6 mass ratio of liposome:lactose. The mixing sequence of the three components, that is, liposome powder, coarse carrier, and fines are also found to influence the fine particle fraction. The maximum fine particle fraction in the range of 17–28% is observed using a Rotahaler (Cipla, India)

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A

B Formulation optimization of nanoliposomes

Formulation optimization of NLDPFs

Method of preparation Thin film hydration Reverse phase evaporation Spray drying Supercritical fluid technology

Method of preparation Physical mixing of dry liposomes with carriers Freeze drying Spray drying Spray-freeze drying Supercritical fluid technology

Optimization of process parameters Temperature and time for drying of film Hydration time Speed of rotation Sonication parameters

Optimization of process parameters Total freezing time Total drying time Spray drying parameters

Optimization of formulation parameters Total drug: lipid ratio Phospholipid: cholesterol ration ratio Amount and type of charged lipids Amount and type of hydration medium

Physicochemical characterization Percentage drug entrapment Particle size and size distribution Zeta-potential Morphology Entrapped volume of aqueous phase Oxidative index

Optimization of formulation parameters Type and amount of cryoprotectant Type and amount of carrier Type and amount of antiadherent

Physicochemical characterization Flow behavior Moisture content Reconstitution time and volume Particle size of reconstituted vesicles Drug retention Stability study Morphology In vitro lung deposition study In vivo study

Figure 9.1 (A) Detailed flow chart describing formulation optimization of nanoliposomes and (B) detailed flow chart describing formulation optimization of NLDPFs.

as a delivery device at 30, 60, and 90 l/min. Although amphotericin NLDPFs are prepared from micron size liposomes and same process can be applied for NLs to process into NLDPFs (Shah and Misra, 2004b).

2.2. Spray-drying The spray-drying method to prepare NLDPFs offers many advantages, including better control over particle formation, and hence can be easily translated to large-scale production (Woolfe et al., 2001). During spraydrying, materials come into contact with hot air or gas for a fraction of a second, otherwise actual drying takes place at ambient temperature, and that

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is why the spray-drying technique is used for the preparation of NLDPFs composed of less temperature-sensitive materials (Bosquillona et al., 2004). Respirable properties of spray-dried NLDPFs containing superoxide dismutase (Briscoe et al., 1995), ciprofloxacin (Sweeney et al., 2005), and antiasthmatic drugs (Patel and Misra, 2007; Sunkara and Kompella, 1999) have been evaluated. In our laboratory, we have developed spray-dried liposomal powder formulations of tacrolimus (Chougule et al., 2007), dapsone (Chougule et al., 2008), and amiloride hydrochloride (AH) (Chougule et al., 2006) with good inhalation properties. Physical properties such as particle size and size distribution, surface energy, surface rugosity, particle density, surface area and microviscosity greatly depend on processing and formulation variables (Chougule et al., 2006, 2008; Parmar, 2006). By controlling such parameters we can get DPI formulation having desired properties for different classes of drugs and widen the application window of pulmonary delivery. AH-incorporated multilamellar vesicles are prepared by thin film hydration. Two hundred milligrams AH, hydrogenated soyaphosphatidylcholine and cholesterol (8:2) are dissolved in CHCl3:CH3OH (1:1) and dried as a thin film under 400 mmHg vacuum and at 65
C temperature. The film is then hydrated at 65
C for 1 h with 50 ml of phosphate-buffered saline (PBS), pH 7.4. Hydration is followed by size reduction by passing through a high-pressure homogenizer (Emulsiflex-C5, Avestin Inc., Ottawa, Canada) preheated to 65
C using a thermostat for three cycles at 10,000 psi. Resultant NLs are subjected to centrifugation at 35,000 rpm, 20
C for 20 min using ultracentrifuge (Sigma Laboratory centrifuge, 3 K 30, Osterode, GmbH). Liposomal pellets are separated, resuspended, and characterized for vesicle size and percent drug entrapment. The liposome pellets equivalent to 50 mg of AH are dispersed in 200 ml PBS containing 15 mg/ml additive (lactose/sucrose/mannitol), and 10% of glycine, based on the weight of the powder at room temperature. For spray-drying, a laboratory spray-dryer (LSD-48, JISL, Mumbai, India) is used. Inlet and outlet temperature are kept at 120  5 and 65–70
C, respectively. The yield of the product is found to be 60  5%. The spray-dried product is filled in capsules (size 2) individually by weight of powder equivalent to 500 mg of AH. HDPE bottles containing capsules and silica bags are placed in a desiccator kept in a refrigerator, after purging with nitrogen and covering with PVC-coated aluminum foil until evaluation (Chougule et al., 2006).

2.3. Freeze-drying NLDPFs of highly heat-sensitive materials such as proteins, peptides, and enzymes can be prepared using freeze-drying method. NLDPFs with different aerodynamic properties can be prepared using different proportions of carriers,

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cryoprotectants, and antiadherents (Lu and Hickey, 2005; Shah and Misra, 2004a; Shahiwala and Misra, 2005). Freeze-dried LDPFs for pulmonary administration of therapeutic molecules such as budesonide, ketotifen, amphotericin B, leuprolide acetate (LA), and levonorgestral have been formulated ( Joshi and Misra, 2001a, 2003; Shah and Misra, 2004a; Shahiwala and Misra, 2004, 2005). The method is explained using leuprolide DPF as an example. The reverse-phase evaporation method is employed to prepare LA-loaded liposomes. LA, hydrogenated soyaphosphatidylcholine, and cholesterol, at 2:4:1 molar ratio, are dissolved in chlorform:methanol:acetate buffer, pH 5.2 (2:4:1) in a quick fit glass tube. After vortexing the tube for 5 min, it is attached to a rotary evaporator (Superfit Continental Pvt Ltd, Mumbai, India) to dry the contents at 55
C under vacuum (400 mmHg) until a gel formation. Again the tube is agitated on vortex mixture for 5 min to collapse the gel into fluid. Ten minutes rotary drying and 5 min vortexing is again repeated twice. The liposomal dispersion is extruded by passing through polycarbonate membranes (below 100 nm). After addition of sucrose in liposomal dispersion, it is sufficiently diluted to obtain a lipid:sugar (sucrose) ratio of 1:6. The dispersion is frozen at  40
C overnight and dried for 24 h under negative displacement pressure (model DW1 0–60E; Heto Drywinner, Birkerod, Denmark). The obtained porous cake is passed through #200 and #240 sieves. To get the final strength of 250 mg LA/10 mg of powder, equivalent amount of Sorbolac 400 is geometrically added. The DPF is filled in capsules (size 2) individually by weight of powder equivalent to 250 mg of LA. HDPE bottles containing capsules and silica bags are placed in a desiccator kept in a refrigerator after purging with nitrogen and covering with PVC-coated aluminum foil until evaluation (Shahiwala and Misra, 2005).

2.4. Spray freeze-drying Spray freeze-drying involves spraying the drug solution into a freezing medium (cryogen) followed by lyophilization of the suspension. Advantages of spray freeze-drying involves almost 100% yield, production of light and porous particles with enhanced aerosol performance, and minimum chances of aggregation of particles. Besides these advantages, disadvantages of the spray freeze-drying process include the expense and the time required (Sweeney et al., 2005). Dimyristoylphosphatidylglycerol, lactose, and ciprofloxacin in a weight percent ratio 5:17:1, respectively, form a smooth suspension upon vortexing (2  30 s to 4  30 s in 1 h), and the suspension remains stable at 4
C for several days. Spray freeze-drying is performed with a two-fluid nozzle, in which compressed nitrogen and a peristaltic pump are used to drive the suspension in fine atomized form into a flask containing liquid nitrogen. The cryogen is then allowed to evaporate, and the resulting powder is dried for 48 h in a lyophilizer. The collector is held at 52
C while the vacuum is 0.004 mbar. The powder is subsequently collected and stored in a sealed vial at 4
C (Sweeney et al., 2005).

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3. Physicochemical Characterization of NLDPFs Prepared liposomes are characterized for percent drug entrapment, size and size distribution, zeta potential, entrapped volume, morphology, oxidative index, and differential scanning calorimetry (DSC) (Akwete and Hsu, 1993; Betagiri et al., 1993b;Chougule et al., 2006, 2007; Goel, 1988; Hyung et al., 2008; Joshi and Misra, 2001b; Kaszuba, 1999; Kent et al., 2007; Maria et al., 2004; Markus et al., 2004; Mina et al., 2002; New, 1990b; Shah and Misra, 2004a; Vyas et al., 2005), and NLDPFs are evaluated for following parameters.

3.1. Assay The amount of drug present in a fixed quantity of DPF can be obtained by dissolving the required quantity of DPF in methanol or a methanol– chloroform mixture. The resultant solution is filtered to remove the sugar part. The amount of drug in the filtrate is estimated by a suitable analytical method (Chougule et al., 2006).

3.2. Flow behavior The angle of repose is measured to determine the flow properties of a powder. The compressibility index reflects the percent voids in the powder along with the flow and dispersibility properties of formulation. Bulk and tapped density of the powder is determined for the calculation of the compressibility index and the theoretical mean mass aerodynamic diameter (MMAD) of the powder. Dispersibility index is the dispersing ability of agglomerated particles on application of pressure equivalent to inhalation air pressure. A higher dispersibility index is expected to get a higher lung deposition (Carr, 1965; Chougule et al., 2006).

3.3. Moisture content determination Stability and flow properties mainly depend on the amount of moisture present in the NLDPFs. A higher amount of moisture causes powder aggregation, leading to poor lung deposition. The moisture content of LDPF is accurately determined by the Karl Fischer titration method for abrupt idea loss on drying (LOD) moisture balance can be used (Chougule et al., 2006). Solid-state characterization and residual water content data for AH NLDPFs are tabulated in Table 9.1.

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Table 9.1 Solid-state characterization and residual water content of amiloride hydrochloride nanoliposomal dry powder formulations

Formulations

AHL AHNLDPFlactose AHNLDPFmannitol AHNLDPFsucrose

Angle of repose (
)

Carr’s compressibility index

Residual water content (%)

0.9  0.1 0.3  0.05

45.6  2.4 31.1  1.9

25.8  1.4 35.8  2.1

5.9  0.6 4.4  0.5

0.2  0.01

27.3  1.5

39.8  1.5

2.5  0.4

0.36  0.04

30.8  1.5

33.4  1.9

5.7  0.8

Tapped density (g/ml)

Reprinted with permission from Chougule et al. (2006). Copyright # American Scientific Publishers, www.aspbs.com.

3.4. Reconstitution time and volume Time and volume of phosphate buffer (pH 7.4) required to reconstitute a fixed quantity of NLDPF is measured (Chougule et al., 2007).

3.5. Particle size of NLDPFs Instruments based on laser light scattering and the photon correlation principle are mainly used to determine the particle size of NLDPFs (Chougule et al., 2006; Goel, 1988; Kaszuba, 1999). Figure 9.2 shows the particle size distribution graph of AH NLDPFs.

3.6. Drug retention and stability studies As per ICH guidelines, comparative drug retention (PDR) studies are performed on the NLDPFs for long-term stability at 5  3
C for 12 months, and accelerated testing at 25  2
C/60% RH  5% RH for 6 months (ICH Guidelines, 2003; Singh, 1999). The product in its final packing is stored at the above-mentioned storage conditions. Samples are withdrawn at definite time intervals and evaluated for assay, water content, PDR, fine particulate fraction, and emission and physical changes (Chougule et al., 2007; Singh, 1999). Data of drug retention and stability of tacrolimus NLDPFs are recorded in Table 9.2.

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Particle size distribution

Volume (%)

12 10 8 6 4 2 0 0.01

0.1

1

10 Particle size (mm)

100

1000

3000

Figure 9.2 Particle size distribution of amiloride hydrochloride NLDPFs. (Reprinted with permission from Chougule et al. (2006). Copyright # American Scientific Publishers, www.aspbs.com.)

3.7. Scanning electron microscopy (photomicrographs and image analysis) Prepared NLDPFs are evaluated for surface morphology, size, shape, and aggregation behavior by using scanning electron microscopy (SEM) (Chougule et al., 2008). Scanning electron microphotograph and b surface texture analysis of dapsone NLDPF are depicted in Fig. 9.3. Topographical features of dapsone NLDPFs measured by SEM image analysis are tabulated in Table 9.3.

3.8. Differential scanning calorimetry Prepared NLDPFs are evaluated for thermal properties using DSC (Chougule et al., 2006). DSC curves of AH NLDPFs are contained in Fig. 9.4.

3.9. In vitro lung deposition studies Anderson cascade impactor (official in USP) and multiple-stage liquid impinger (official in BP and EP) resembles human respiratory track and thus used to evaluate target region within the lung and deposition of inhaled NLDPFs in each region of lung. From that data the emitted dose, fine particle dose, fine particle fraction, MMAD, and geometric standard deviation (GSD) can be calculated according to the procedure given in USP (USP 30 NF 25, 2007). The other parameters to be estimated include recoverable dose, emitted dose, percent emission, and percent dispersibility.

Table 9.2 Stability data of tacrolimus nanoliposomal dry powder formulations

Stability conditions

Initial

40  1M

2M

3M

6M

30  1M

2M

Description

Assay (%)

101.16  1.86 White free flowing powder 2
C and 75  5% RH 101.08  3.24 White free flowing powder 99.43  2.97 White free flowing powder 98.09  3.95 White free flowing powder 99.72  3.54 White free flowing powder
2 C and 65  5%RH 103.46  2.72 White free flowing powder 102.16  2.91 White free flowing powder

Water content

Percent drug retained

Liposomal size (VMD) (mm)

Emission (%)

Fine particle fraction (FPF)

2.8  0.3

101.38  2.47

0.140  0.02

82  3.0

71.1  2.5

2.7  0.5

95.92  3.51

0.145  0.03

80.6  2.2

70.45  2.7

3.3  0.5

91.45  3.95

0.152  0.03

78.4  3.0

67.13  2.5

3.4  0.6

87.43  3.91

0.158  0.02

77.2  2.9

62.91  1.9

3.0  0.7

85.71  3.98

0.161  0.03

75.4  2.6

60.11  3.0

2.8  0.4

100.26  3.29

0.144  0.03

81.87  3.7

70.93  2.1

2.9  0.5

101.49  3.96

0.138  0.03

81.46  3.4

71.08  2.4

(continued)

Table 9.2 (continued) Stability conditions

Description

Assay (%)

101.13  2.58 White free flowing powder 100.18  3.1 6M White free flowing powder 99.42  2.9 9M White free flowing powder 100.51  3.59 12 M White free flowing powder
25  2 C and 60  5% RH 102.19  2.53 3M White free flowing powder 101.17  3.5 6M White free flowing powder 101.83  3.92 9M White free flowing powder 102.81  4.59 12 M White free flowing powder 3M

Water content

Percent drug retained

Liposomal size (VMD) (mm)

Emission (%)

Fine particle fraction (FPF)

2.9  0.4

100.17  3.21

0.148  0.02

81.4  3.9

70.84  2.3

3.0  0.4

102.41  4.23

0.151  0.03

80.19  3.2

68.42  2.9

2.9  0.5

99.59  3.91

0.153  0.04

80.07  3.0

67.82  2.8

2.9  0.4

99.14  4.1

0.152  0.03

78.76  3.5

65.19  2.8

2.8  0.3

101.80  2.95

0.138  0.03

82.4  2.6

71.20  2.7

2.8  0.4

102.11  3.44

0.143  0.02

82.1  3.1

70.12  2.5

2.9  0.5

101.74  4.28

0.148  0.02

80.74  2.7

68.17  2.9

2.9  0.6

101.62  4.38

0.143  0.02

79.13  3.4

68.28  2.8

Reprinted with permission from Chougule et al. (2008). Copyright # Dove Medical Press.

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A

B

Figure 9.3 Scanning electron micrograph (A) and surface texture analysis (B) of dapsone NLDPF. (With kind permission from Springer Science þ Business Media: Chougule et al. (2008).

NLDPF density and geometric diameter are the key parameter which decides MMAD of NLDPF and thus lung deposition and effectiveness of the therapeutic delivery system (Chougule et al., 2007). Tacrolimus NLDPFs were evaluated for in vitro pulmonary deposition using an eight stage Andersen cascade impactor with a preseparator (Graseby-Andersen, Atlanta, GA, USA) and data obtained are tabulated in Table 9.4.

3.10. In vitro drug release study In vitro drug release study is a tool that provides an estimate of the in vivo pharmacokinetic and pharmacodynamic performance of the formulation (Fielding and Abra, 1992). Therefore an in vitro release technique is proposed, validated, and utilized for drug release studies from optimized NLDPFs. To examine the drug release kinetics and mechanism, the percent drug diffused (Shah et al., 1993), the kinetics of release, the mean steadystate flux (Chien, 1992; Robinson and Lee, 1987), and the diffusion coefficient (Chougule et al., 2006) are measured. The curve of percent drug released versus time for AH NLDPFs is graphically presented in Fig. 9.5.

3.11. In vivo studies In vivo studies are to be performed to evaluate lung absorption and the lung deposition pattern. For this purpose, the following three approaches have been utilized.

Table 9.3 Topographical features of dapsone NLDPFs measured by SEM image analysis Formulations

Roundness

Aspect ratio

Fractal dimension

Heterogeneity

Clumpiness

NLDPF-lactose NLDPF-trehalose NLDPF-sucrose

1.106  0.076 1.015  0.065 1.376  0.092

1.356  0.210 1.163  0.196 1.453  0.209

1.129  0.064 0.856  0.052 1.341  0.064

0.492  0.057 0.351  0.051 0.674  0.153

0.194  0.057 0.109  0.047 0.274  0.095

With kind permission from Springer Science þ Business Media: Chougule et al. (2008).

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Nanoliposomal Dry Powder Formulations

40.00 a b c

DSC mW

20.00

0.00

-20.00 -40.00

200.00

100.00

300.00

Temp (C)

Figure 9.4 Differential scanning calorimetric graphs of developed amiloride hydrochloride NLDPF (a), plain drug (b), and physical mixture (c). (Reprinted with permission from Chougule et al. (2006). Copyright # American Scientific Publishers, www. aspbs.com.) Table 9.4 In vitro aerosol deposition data of tacrolimus nanoliposomal dry powder formulations

Formulations

NLDPFtrehalose NLDPFsucrose NLDPFlactose

Emitted dose (%)

Fine particle fraction (%)

Mean median aerodynamic diameter (mm)

Geometric standard deviation

82  3.0

71.1  2.5

2.2  0.1

1.7  0.2

63  5.5

53.7  3.6

2.8  0.2

2.3  0.1

74  4

62.8  3.1

2.6  0.2

2.2  0.1

Reprinted with permission from Chougule et al. (2008). Copyright # Dove Medical Press.

3.11.1. Animal models (in vivo) Evaluation of pharmacokinetic behavior and distribution in different sections of the lungs are carried out using different animal models, including rats and guinea pigs. Methods such as intratracheal instillation, micro spray, nebulization, and aerosol puff are used for dosing of NLDPFs in animals. Drug present in homogenate of various lobes of the lung and in bronchoalveolar lavage is estimated at various time intervals to get the idea of amount of drug deposition as well as pattern of distribution within the lungs. The amount of drug present in blood and homogenates of other major organs is estimated to assess systemic absorption after inhalation of NLDPFs. Validated analytical methods are developed in our laboratory to

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Gaurang Patel et al.

100 90 80 70 60 50 40 30 20 10 0 0

2

SAHL

4

6 Time (h)

SLDPIM

8

SLDPIL

10

12

SLDPIS

Figure 9.5 In vitro release profile of developed amiloride hydrochloride NLDPFs and plan drug DPFs (SAHL ¼ plain drug formulation, SLDPIM ¼ NLDPF containing mannitol, SLDPIL ¼ NLDPF containing lactose, and SLDPFs ¼ NLDPF containing sucrose). (Reprinted with permission from Chougule et al. (2006). Copyright # American Scientific Publishers, www.aspbs.com.)

estimate the pharmacokinetic profiles of dapsone, rifampicin, amikacin, ketotifen, tacrolimus, AH, isoniazid, and amphotericin B in rats (Chougule et al., 2006, 2007, 2008; Joshi and Misra, 2001a,c, 2003; Shah and Misra, 2004a,c). Serum LH levels are measured to estimate the pharmacodynamic response after intratracheal administration of leuprolide and levonorgestral liposomes (Shahiwala and Misra, 2004, 2005). However, these methods require sacrifice of animals, and hence other techniques, including in vitro cell lines and ex vivo techniques, may also be used (Chougule et al., 2006). AH-loaded NLDPFs are evaluated for in vivo performance following intratracheal instillation. For each time interval, six albino rats (220–240 g) of either sex are housed in individual plastic cages at a constant temperature. Rats are fasted overnight prior to each experiment. After anesthetizing rats by intraperitoneal administration of a urethane solution (1.2 gm/kg), the trachea is exposed by blunt dissection of the sternohyoideus muscle and a small midline incision is made over the trachea between the fifth and sixth tracheal rings using a 20-gauge needle. The trachea is cannulated with PE200 tubing (5–7 cm) with the tip positioned approximately at the tracheal bifurcation. The PE50 (10– 15 cm) tubing connected to a glass Hamilton syringe (Waters, Bangalore, India) is inserted into the cannula and advanced to the bifurcation of the trachea. Prepared formulations equivalent to 100 mg of AH are administered, and animals are sacrificed at 2, 4, 8, 10, and 12 h after administration (Fig. 9.6). Bronchoalveolar lavage is performed on anesthetized and recannulated animals with 12 ml PBS, prewarmed to 37
C. To perform the lavage, the Hamilton syringe connected to the PE50 tubing is replaced with a three-way stopcock attached to two 20 ml syringes. Approximately 12 ml

Nanoliposomal Dry Powder Formulations

Figure 9.6

183

Intratracheal instillations and bronchoalveolar lavage on rat.

sterile (0.22 mm filtered) PBS is injected slowly in fractions to fill the lungs. The fluid is withdrawn by gentle aspiration; this bronchial alveolar lavage yields between 7 and 11 ml liquid, which is centrifuged at 2000 rpm for 5 min. One percent Triton X-100 is added to the supernatant and analyzed to determine unreleased AH. The lungs and portions of the trachea below the instillation site are excised and homogenized in 10 ml PBS containing 1% Triton X-100. Sulphosalicylic acid (10%) is used as a deproteinizing agent. AH released is analyzed in the supernatant after suitable dilution (Chougule et al., 2006). 3.11.2. Lung epithelial cell models (in vitro) In vitro cell line models have become more popular than in vivo models to evaluate prepared formulations for cell uptake, cell toxicity, and cell death kinetics studies (Anabousi et al., 2006; Elbert et al., 1999; Fang et al., 2004; Fuchs et al., 2003; Pang et al., 2005; Sakagami et al., 2002a,b). Intact lung studies have two limitations: (i) the contribution of different distal lung epithelial cells can not be studied separately, and (ii) the surface area for fluid absorption can only be approximated. Lung epithelial cell lines include tracheobronchial, bronchial, and alveolar cell lines and they are mainly of rat and human origin. The examples of cancer-derived cell lines include Calu-1, Calu-3, Calu-6, H441, HBE1, A427, A549, 16HBE140, and BEAS-2B. The examples of cell lines transformed from normal lung include 9HTE16o-, 16HBE14o-1HAEo-, BEAS-2BCF/T43, and AK-D. These cell lines have been tested for the change in transepithelial transport kinetics (i.e., Papp) and for predicting in vivo lung absorption (Tronde et al., 2003).

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An in vitro cell culture model that mimics the respiratory epithelium, in particular cells that form tight monolayers and secrete components of mucus or surfactant, would be a valuable tool for NLDPFs characterization. The air-interfaced culture (AIC) or liquid-covered culture (LCC)-grown Calu3 monolayers can be used as an in vitro model to evaluate therapeutic large porous powder particles (Fiegel et al., 2003). Similar experiments can be designed for evaluation of NLDPFs. Briefly, Calu-3 cells are seeded onto Transwell clear permeable filter inserts at a density of 105 cells/cm2. After seeding, cells are grown in 1500 ml (apical) and 2600 ml (basolateral) of cell culture media at 37
C in a 5% CO2 incubator. For LCC conditions, the media bathing the cells are removed and cells are fed with 1500 ml fresh apical and 2600 ml fresh basolateral media 1 day after seeding, and then once daily. For AIC conditions, the apical medium of cells are removed 1 day after seeding and cells are fed daily with 1400 ml fresh basolateral media only after. When the cells reach sufficient confluency, the apical fluid is removed from the LCC-grown Calu-3 monolayers, and the basolateral media is reduced to 1400 ml. AIC-grown monolayers are used as a control. Filter inserts containing cell monolayers are transferred from the 6-well plate to a cut-out single well containing prewarmed media. One well is placed directly under the second-stage nozzle of an Astra-type liquid impinger (Erweka, Heusenstamm, Germany) and the impinger is sealed, as illustrated in Fig. 9.7. Dry powder particles are aerosolized onto the monolayers for 30 s at 30 l/min from #2 gelatin capsules, using a Spinhaler device. The filter inserts are then returned to the 6-well plate and placed back into an incubator at 37
C. Slightly wetted Transwell inserts without cells are placed in the impinger and treated as described earlier to obtain reproducible data. After particle impinging, 0.8 N NaOH is added to the apical and basolateral compartments of the filter to dissolve the dry powder. Following a 24 h incubation at 37
C, the complete degradation of the particles on the filter inserts are verified by light microscopy. Samples are taken and analyzed for fluorescein-sodium using a fluorescence plate reader (at excitation and emission wavelengths of 485 and 530 nm, respectively; Fiegel et al., 2003). 3.11.3. Lung tissue models (ex vivo) Ex vivo lung tissue models have been preferred when the information on the mechanisms involved in drug transport or lung disposition kinetics are not predictable from in vivo or in vitro models. The isolated perfused lung (IPL) model is most widely used for the purpose of establishing the mechanisms of drug absorption and deposition in the lungs. In this type of model, the lung are isolated from the body and placed in a simulated environment to perform the desired experiment. IPL prepared from small rodents have been used for drug deposition and organ toxicity studies (Pang et al., 2005; Sakagami et al., 2002a,b; Tronde et al., 2003). The limitations of IPL models includes. The only

185

Nanoliposomal Dry Powder Formulations

Inhaler device

Transwell containing cell monolayer

Outlet to vacuum

Figure 9.7 Astra-type liquid cascade impinger used with a Transwell under the second-stage nozzle. (With kind permission from Springer Science þ Business Media: Fiegel et al. (2003)).

drawbacks associated with these models are the short viable periods of (2–3 h), and the absence of blood and mucous circulation. The IPL preparation has been employed mostly with rats and guinea pigs rather than with mice, dogs and monkeys. The preparation consists of (a) peristaltic pumps and a tubing assembly that is used to carry the perfusate to and from the lungs and (b) a double-jacketed artificial thorax to house the IPL at a certain temperature, normally at 37
C. A flow rate of 12–15 ml/ min (rat lung) is used to pump the perfusate from the reservoir to the IPL through a central porthole of the thorax is lid. The opening at the bottom of the thorax is used to return the perfusate to the reservoir or to collection. The air-tight sealed thorax enables negative- or positive-pressure ventilation, or the maintenance of a certain pressure. Under anesthesia, the pulmonary circulation of the lung is cannulated via the pulmonary artery and lung perfusion is initiated and maintained. The pulmonary vein is also cannulated as an exit of the lung perfusion, or as an alternative, the bottom of the left auricle and both sides of the ventricles in the heart are amputated to allow the free flow of perfusate without any unnecessary back pressure in

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95%O2 + 5%CO2 Thermometer pH probe

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B Air Fluid dispenser

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Figure 9.8 Isolated perfused rat lung (IPRL) preparations of (A) Horizontally positioned IPRL with a scheme of forced solution instillation and (B) vertically positioned IPRL with a scheme of nebulization catheter dosing. (Reprinted from Sakagami et al. (2006). Copyright 2006, with permission from Elsevier.)

the IPL. The lung and heart are excised from the body en bloc. The trachea is also cannulated and often exteriorized out of the thorax. This setup supports the IPL suspension in the thorax, and for pulmonary biopharmaceutics research, allows airway lumienal administration of test molecules. The IPL can be suspended in a horizontal or vertical fashion, as shown in Fig. 9.8. Autologous whole blood or a buffered artificial medium, such as Krebs– Ringer or Krebs–Henseleit buffer (pH 7.4) supplemented with 5 mM glucose and/or 4–5% albumin can be used as the perfusate. The perfusate medium is equilibrated with a mixture of oxygen and carbon dioxide

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throughout the experiments. Generally, the perfusion is recirculated, but can also be a single-pass system. Using sufficient surgical technique, it has been shown that the IPL viability can be maintained for a maximum of 2–3 h at 37
C, after which the lung gradually develops translucent areas, indicating edema formation (Sakagami et al., 2006). After the perfusion begins, the perfusate samples are with drawn via the tracheal port at different time intervals, enabling the characterization of