Autophagosome and Phagosome

Autophagosome and Phagosome M E T H O D S I N M O L E C U L A R B I O L O G YTM John M. Walker, SERIES EDITOR 457...

Author: Vojo Deretic

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Autophagosome and Phagosome

M E T H O D S

I N

M O L E C U L A R

B I O L O G YTM

John M. Walker, SERIES EDITOR 457. Membrane Trafficking, edited by Ales Vancura, 2008 456. Adipose Tissue Protocols, Second Edition, edited by Kaiping Yang, 2008 455. Osteoporosis, edited by Jennifer J. Westendorf, 2008 454. SARS- and Other Coronaviruses: Laboratory Protocols, edited by Dave Cavanagh, 2008 453. Bioinformatics, Volume 2: Structure, Function, and Applications, edited by Jonathan M. Keith, 2008 452. Bioinformatics, Volume 1: Data, Sequence Analysis, and Evolution, edited by Jonathan M. Keith, 2008 451. Plant Virology Protocols: From Viral Sequence to Protein Function, edited by Gary Foster, Elisabeth Johansen, Yiguo Hong, and Peter Nagy, 2008. 450. Germline Stem Cells, edited by Steven X. Hou and Shree Ram Singh, 2008. 449. Mesenchymal Stem Cells: Methods and Protocols, edited by Darwin J. Prockop, Douglas G. Phinney, and Bruce A. Brunnell, 2008. 448. Pharmacogenomics in Drug Discovery and Development, edited by Qing Yan, 2008. 447. Alcohol: Methods and Protocols, edited by Laura E. Nagy, 2008. 446. Post-translational Modification of Proteins: Tools for Functional Proteomics, Second Edition, edited by Christoph Kannicht, 2008. 445. Autophagosome and Phagosome, edited by Vojo Deretic, 2008. 444. Prenatal Diagnosis, edited by Sinhue Hahn and Laird G. Jackson, 2008. 443. Molecular Modeling of Proteins, edited by Andreas Kukol, 2008. 442. RNAi: Design and Application, edited by Sailen Barik, 2008. 441. Tissue Proteomics: Pathways, Biomarkers, and Drug Discovery, edited by Brian C.-S. Liu, 2008. 440. Exocytosis and Endocytosis, edited by Andrei I. Ivanov, 2008. 439. Genomics Protocols, Second Edition, edited by Mike Starkey and Ramnanth Elaswarapu, 2008 438. Neural Stem Cells: Methods and Protocols, Second Edition, edited by Leslie P. Weiner, 2008 437. Drug Delivery Systems, edited by Kewal K. Jain, 2008 436. Avian Influenza Virus, edited by Erica Spackman, 2008 435. Chromosomal Mutagenesis, edited by Greg Davis and Kevin J. Kayser, 2008 434. Gene Therapy Protocols: Volume 2: Design and Characterization of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 433. Gene Therapy Protocols: Volume 1: Production and In Vivo Applications of Gene Transfer Vectors, edited by Joseph M. LeDoux, 2008 432. Organelle Proteomics, edited by Delphine Pflieger and Jean Rossier, 2008

431. Bacterial Pathogenesis: Methods and Protocols, edited by Frank DeLeo and Michael Otto, 2008 430. Hematopoietic Stem Cell Protocols, edited by Kevin D. Bunting, 2008 429. Molecular Beacons: Signalling Nucleic Acid Probes, Methods and Protocols, edited by Andreas Marx and Oliver Seitz, 2008 428. Clinical Proteomics: Methods and Protocols, edited by Antonia Vlahou, 2008 427. Plant Embryogenesis, edited by Maria Fernanda Suarez and Peter Bozhkov, 2008 426. Structural Proteomics: High-Throughput Methods, edited by Bostjan Kobe, Mitchell Guss, and Huber Thomas, 2008 425. 2D PAGE: Sample Preparation and Fractionation, Volume 2, edited by Anton Posch, 2008 424. 2D PAGE: Sample Preparation and Fractionation, Volume 1, edited by Anton Posch, 2008 423. Electroporation Protocols: Preclinical and Clinical Gene Medicine, edited by Shulin Li, 2008 422. Phylogenomics, edited by William J. Murphy, 2008 421. Affinity Chromatography: Methods and Protocols, Second Edition, edited by Michael Zachariou, 2008 420. Drosophila: Methods and Protocols, edited by Christian Dahmann, 2008 419. Post-Transcriptional Gene Regulation, edited by Jeffrey Wilusz, 2008 418. Avidin–Biotin Interactions: Methods and Applications, edited by Robert J. McMahon, 2008 417. Tissue Engineering, Second Edition, edited by Hannsjörg Hauser and Martin Fussenegger, 2007 416. Gene Essentiality: Protocols and Bioinformatics, edited by Svetlana Gerdes and Andrei L. Osterman, 2008 415. Innate Immunity, edited by Jonathan Ewbank and Eric Vivier, 2007 414. Apoptosis in Cancer: Methods and Protocols, edited by Gil Mor and Ayesha Alvero, 2008 413. Protein Structure Prediction, Second Edition, edited by Mohammed Zaki and Chris Bystroff, 2008 412. Neutrophil Methods and Protocols, edited by Mark T. Quinn, Frank R. DeLeo, and Gary M. Bokoch, 2007 411. Reporter Genes: A Practical Guide, edited by Don Anson, 2007 410. Environmental Genomics, edited by Cristofre C. Martin, 2007 409. Immunoinformatics: Predicting Immunogenicity In Silico, edited by Darren R. Flower, 2007 408. Gene Function Analysis, edited by Michael Ochs, 2007 407. Stem Cell Assays, edited by Vemuri C. Mohan, 2007 406. Plant Bioinformatics: Methods and Protocols, edited by David Edwards, 2007 405. Telomerase Inhibition: Strategies and Protocols, edited by Lucy Andrews and Trygve O. Tollefsbol, 2007

M E T H O D S I N M O L E C U L A R B I O L O G YT M


Edited by

Vojo Deretic Department of Molecular Genetics and Microbiology University of New Mexico Health Sciences Center Albuquerque, NM, USA

Vojo Deretic Department of Molecular Genetics and Microbiology University of New Mexico Health Sciences Center, Albuquerque, NM, USA [email protected]

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

ISBN: 978-1-58829-853-9

e-ISBN: 978-1-59745-157-4

Library of Congress Control Number: 2007940755 ©2008 Humana Press, a part of Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Background: An electron micrograph of a nascent autophagosome in a normal rat kidney cell. The autophagosome limiting membrane is visible as a dark line. Endoplasmic reticulum cisternae with attached ribosomes surround the autophagosome on two sides. Endoplasmic reticulum is also among the contents captured by the autophagosome. Golgi ribbon is seen on the bottom right. Prepared by Eeva-Liisa Eskelinen, University of Helsinki. Inset: A novel autophagy reporter, tandem monomeric RFP-GFP-tagged LC3 (tfLC3) can be used to distinguish between autophagosomes and autolysosomes. The GFP signal is sensitive to the acidic and/or proteolytic conditions of the autolysosomal lumen, and thus GFP fluorescence is lost in autolysosomes, whereas mRFP is more stable and its fluorescence persists. Therefore, colocalization of GFP (green) and RFP (magenta) fluorescence indicates a compartment, such as the phagophore or an autophagosome, that has not yet fused with a lysosome. In contrast, an mRFP signal without GFP fluorescence corresponds to an autolysosome, as evidenced by colocalization with Lamp1 (cyan). Prepared by Shunsuke Kimura, Takeshi Noda, and Tamotsu Yoshimori, Osaka University. Printed on acid-free paper 987654321 springer.com

Preface

The intent of this volume is to provide a comprehensive resource with detailed methods for study of two distinct but partially morphologically similar processes of autophagy and phagocytosis. Autophagy is a rapidly growing field, and there is a need for standards of assessment in identification of autophagosomal organelles and for monitoring various aspects of autophagic functions. Phagocytosis is a relatively mature field that has established methods but can benefit from an update on the current trends. Finally, cross-pollination between the two fields is of interest. Although cross-cutting studies between phagocytosis (which could be viewed as a special case of autophagy of a cell’s exterior) are presently few and far between, it is possible that a merger of methods in both fields will prompt further explorations of similarities and differences. The collection of methods described in this book should allow the reader to find appropriate techniques to identify, monitor, and quantify autophagic processes in cellular and animal models of autophagy. Since the basic autophagic machinery is highly conserved, these methods can be applied nearly universally—of course with appropriate and judicious modifications. Among the core battery of assays are: (1) GFP-LC3 (Atg8) puncta formation, monitored by fluorescence microscopy; (2) lipidation of LC3 and associated electrophoretic mobility shift, monitored by immunoblotting; (3) ultrastructural analysis by electron microscopy; and (4) proteolysis of stable proteins by monitoring radioactive amino acid release during autophagic turnover. These techniques can be complemented by less specific but relatively quick methods of staining with acidotropic dyes (lysotracker and monodansylcadaverine) and more importantly mechanistic studies using pharmacological agonists and antagonists and, very importantly, siRNA knockdowns of key autophagic proteins (e.g., Beclin 1, Atg5, Atg7). Somewhat less accessible, but very important, are Atg knockout cell lines and transgenic animals, including murine and fly models. The core methodologies and approaches are applicable whether the objective is to study cell survival, cell death, cancer, neurodegeneration (Huntington’s, Alzheimer’s, Parkinson’s diseases), development, aging, intrinsic (cell-autonomous) resistance to infection, innate and adaptive immunity, antigen processing, T- or B-cell homeostasis, and numerous other v

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health and disease states. The chapters in this volume, from a number of authorities in the field of autophagy, should facilitate work in laboratories with or without prior experience in autophagy research. There are many important questions to be answered regarding fundamental and applied aspects of phagosomal biology, apart from the partial overlaps with the autophagic pathway highlighted here. The methods described in this volume should allow researchers to find in one place several modern techniques for in vitro and in vivo studies of phagosomal organelles. While this book was in its production stages, several autophagy methods and biological relationships of high relevance have been published, attesting to the impressive speed at which this field is moving. These are: (i) A work in Nature (1), directly linking autophagy and phagocytosis, along the lines anticipated in this book and touched upon in Chapter 1. (ii) A multi-author comprehensive discussion on the use and limitations of various autophagy assays (2). The interested reader is advised to consult this text. (iii) An important methodological refinement on how to monitor and quantify LC3-I-to-II conversion by immunoblotting (3). This method calls for comparison of samples from cells treated with a putative inducer of autophagy in the presence and absence of Bafilomycin A (an inhibitor of acidification and maturation of autophagosomes into degradative organelles). The intensity of the LC3-II band is compared to the intensity of actin (unlike comparisons to LC3-I, as often done in the past). A suspected inducer of autophagy under examination is expected to increase the intensity of the LC3-II band (relative to actin) in the sample treated with both the putative inducer and Bafilomycin A when compared to the intensity seen in a parallel control (without the putative inducer) treated only with Bafilomycin A. (iii) Another significant advance is the use of a tandem RFP-GFP-LC3 fusion (instead of single GFP-LC3 fusion) to monitor LC3 puncta by imaging (see book cover) (4). This assay is based on differential sensitivity of GFP and RFP to lumenal pH in autophagic organelles: GFP is pH-sensitive but RFP is not. Here, doubly positive puncta (green+ red+ ; or yellow when green and red images are merged) represent newly induced autophagosomes, while singly positive (green− red+ puncta represent autophagic organelles that have acidified and matured into degradative organelles. By consulting these new methodological developments and using the detailed, step-by-step protocols in this volume, the researchers entering or already working in this field will have a full panel of methods at their fingertips.

References 1. Sanjuan, M.A. et al. (2007) Toll-like receptor signalling in macrophages links the autophagy pathway to phagocytosis. Nature 450:1253–1257.

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2. Klionsky, D.J. et al. (2008) Guidelines for the use and interpretation of assays for monitoring autophagy in higher eukaryotes. Autophagy 4:151–175. 3. Mizushima, N. & Yoshimori, T. (2007) How to interpret LC3 immunoblotting. Autophagy 3:542–545. 4. Kimura, S., Noda, T. & Yoshimori, T. (2007) Dissection of the autophagosome maturation process by a novel reporter protein, tandem fluorescent-tagged LC3. Autophagy 3:452–460.

January 2008, Placitas, New Mexico, USA

Vojo Deretic

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xiii

1. Autophagosome and Phagosome Vojo Deretic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2. Fine Structure of the Autophagosome Eeva-Liisa Eskelinen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

3. Methods for Assessing Autophagy and Autophagic Cell Death Ezgi Tasdemir, Lorenzo Galluzzi, M. Chiara Maiuri, Alfredo Criollo, Ilio Vitale, Emilie Hangen, Nazanine Modjtahedi, and Guido Kroemer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

4. LC3 and Autophagy Isei Tanida, Takashi Ueno, and Eiki Kominami . . . . . . . . . . . . . . . . . . . . . . . . . 77

5. Amino Acid Regulation of Autophagosome Formation Alfred J. Meijer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

6. Autophagic Proteolysis of Long-Lived Proteins in Nonliver Cells Esteban A. Roberts and Vojo Deretic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .111

7. Autophagosomes in GFP-LC3 Transgenic Mice Noboru Mizushima and Akiko Kuma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119

8. Experimental Control and Characterization of Autophagy in Drosophila Gabor Juhasz and Thomas P. Neufeld . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .125

9. Analysis of Autophagosome Membrane Cycling by Fluorescence Microscopy Julie E. Legakis and Daniel J. Klionsky . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .135

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10. Protein Trafficking into Autophagosomes Andrew Young and Sharon Tooze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .147

11. Sphingolipids in Macroautophagy Grégory Lavieu, Francesca Scarlatti, Giusy Sala, Stéphane Carpentier, Thierry Levade, Riccardo Ghidoni, Joëlle Botti, and Patrice Codogno . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .159

12. Molecular Links Between Autophagy and Apoptosis Iwona A. Ciechomska, Christoph G. Goemans, and Aviva M. Tolkovsky. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .175

13. Clearance of Mutant Aggregate-Prone Proteins by Autophagy Brinda Ravikumar, Sovan Sarkar, and David C. Rubinsztein . . . . . . . . . . . .195

14. Localization and MHC Class II Presentation of Antigens Targeted for Macroautophagy Dorothee Schmid and Christian Münz . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .213

15. Chaperone-Mediated Autophagy S. Kaushik and A. M. Cuervo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .227

16. Microautophagy in the Yeast Saccharomyces cerevisiae Andreas Uttenweiler and Andreas Mayer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .245

17. EM Analysis of Phagosomes Chantal de Chastellier. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .261

18. Analysis of Phosphoinositide Dynamics During Phagocytosis Using Genetically Encoded Fluorescent Biosensors Gabriela Cosío and Sergio Grinstein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .287

19. In Vitro Phagosome–Endosome Fusion Isabelle Vergne and Vojo Deretic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301

20. Real-Time Spectrofluorometric Assays for the Lumenal Environment of the Maturing Phagosome Robin M. Yates and David G. Russell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .311

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21. Monitoring Time-Dependent Maturation Changes in Purified Phagosomes from Dictyostelium discoideum Régis Dieckmann, Navin Gopaldass, Caroline Escalera, and Thierry Soldati . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .327

22. Large-Scale Purification of Latex Bead Phagosomes from Mouse Macrophage Cell Lines and Subsequent Preparation for High-Throughput Quantitative Proteomics Adam Rupper and James Cardelli. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .339

23. Class II MHC Antigen Processing in Phagosomes Lakshmi Ramachandra, W. Henry Boom, and Clifford V. Harding . . . . . .353

24. Analyzing Association of the Endoplasmic Reticulum with the Legionella pneumophila–Containing Vacuoles by Fluorescence Microscopy Alyssa Ingmundson and Craig R. Roy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .379

25. Fractionation of the Coxiella burnetii Parasitophorous Vacuole Dale Howe and Robert A. Heinzen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .389

26. Bacterial Phagosome Acidification Within IFN--Activated Macrophages: Role of Host p47 Immunity-Related GTPases (IRGs) Sangeeta Tiwari and John D. MacMicking . . . . . . . . . . . . . . . . . . . . . . . . . . . . .407

27. SopE-Mediated Recruitment of Host Rab5 on Phagosomes Inhibits Salmonella Transport to Lysosomes Richa Madan, Ganga Krishnamurthy, and Amitabha Mukhopadhyay . . .417

28. The Mycobacterium tuberculosis Phagosome Esteban A. Roberts and Vojo Deretic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .439

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451

Contributors

W. Henry Boom • Division of Infectious Diseases, Case Western Reserve University, Cleveland, OH Joëlle Botti • INSERM U756, Faculté de Pharmacie, Université Paris-Sud, Châtenay-Malabry, France James A. Cardelli • Department of Microbiology and Immunology, Louisiana State University Health Sciences Center, Shreveport, LA Stéphane Carpentier • INSERM U466, Institut Louis Bugnard, Centre Hospitalier Universitaire de Rangueil, Toulouse, France Iwona A. Ciechomska • Department of Biochemistry, University of Cambridge, Cambridge, UK Patrice Codogno • INSERM U756, Faculté de Pharmacie, Université Paris-Sud, Châtenay-Malabry, France Gabriela Cosío • Program in Cell Biology, Hospital for Sick Children, Toronto, Ontario, Canada Alfredo Criollo • INSERM, Unit “Apoptosis, Cancer and Immunity”, Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine – Université Paris-Sud; Villejuif, France Ana Maria Cuervo • Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, NY Chantal de Chastellier • Centre d’Immunologie de Marseille-Luminy, Marseille, France Vojo Deretic • Health Sciences Center, Department Molecular Genetics and Microbiology, University of New Mexico Health Sciences Center, Albuquerque, NM, USA Régis Dieckmann • Department of Biochemistry, University of Geneva, Geneva, Switzerland Caroline Escalera • Department of Biochemistry, University of Geneva, Geneva, Switzerland Eeva-Liisa Eskelinen • University of Helsinki, Department of Biological and Environmental Sciences, Division of Biochemistry, University of Helsinki, Helsinki, Finland xiii

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Lorenzo Galluzzi • INSERM, Unit “Apoptosis, Cancer and Immunity,” Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine–Université Paris-Sud, Villejuif, France Riccardo Ghidoni • Laboratory of Biochemistry and Molecular Biology, San Paolo Medical School, Milan, Italy Christoph G. Goemans • Department of Biochemistry, University of Cambridge, Cambridge, UK Navin Gopaldass • Department of Biochemistry, University of Geneva, Geneva, Switzerland Sergio Grinstein • Program in Cell Biology, Hospital for Sick Children, Toronto, Ontario, Canada Emilie Hangen • INSERM, Unit “Apoptosis, Cancer and Immunity”, Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine – Université Paris-Sud; Villejuif, France Clifford V. Harding • Department of Pathology, Case Western Reserve University, Cleveland, OH Robert A. Heinzen • Coxiella Pathogenesis Section, Laboratory of Intracellular Parasites, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT Dale Howe • Coxiella Pathogenesis Section, Laboratory of Intracellular Parasites, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT Alyssa Ingmundson • Section of Microbial Pathogenesis, Yale University School of Medicine, Boyer Center for Molecular Medicine, New Haven, CT Gabor Juhasz • Department of General Zoology, Eötvös Loránd University, Budapest, Hungary S. Kaushik • Department of Anatomy and Structural Biology, Marion Bessin Liver Research Center, Albert Einstein College of Medicine, New York, NY Daniel J. Klionsky • Life Sciences Institute and Departments of Molecular, Cellular, and Developmental Biology and Biological Chemistry, University of Michigan, Ann Arbor, MI Eiki Kominami • Department of Biochemistry, Juntendo University School of Medicine, Bunkyo-ku, Tokyo, Japan Ganga Krishnamurthy • National Institute of Immunology, Aruna Asaf Ali Marg, New Delhi, India Guido Kroemer • INSERM, Unit “Apoptosis, Cancer and Immunity,” Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine–Université Paris-Sud, Villejuif, France Akiko Kuma • Department of Physiology and Cell Biology, Tokyo Medical and Dental University, Tokyo, Japan and SORST, Japan Science and Technology Agency, Japan

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Grégory Lavieu • Department of Physiology and Cellular Biophysics, Columbia University, New York, NY Julie E. Legakis • Life Sciences Institute and Departments of Molecular, Cellular, and Developmental Biology and Biological Chemistry, University of Michigan, Ann Arbor, MI Thierry Levade • INSERM U466, Institut Louis Bugnard, Centre Hospitalier Universitaire de Rangueil, Toulouse, France John D. MacMicking • Section of Microbial Pathogenesis, Yale University School of Medicine, New Haven, CT Richa Madan • National Institute of Immunology, Aruna Asaf Ali Marg, New Delhi, India Andreas Mayer • Département de Biochimie, Université de Lausanne, Epalinges, Switzerland M. Chiara Maiuri • INSERM, Unit “Apoptosis, Cancer and Immunity”, Villejuif, France; Institut Gustave Roussy, Villejuif, France; Dipartimento di Farmacologia Sperimentale, Facoltà di Scienze Biotecnologiche – Università degli Studi di Napoli “Federico II”; Napoli, Italy Alfred J. Meijer • Department of Medical Biochemistry, Academic Medical Center, University of Amsterdam, Meibergdreef, Amsterdam, The Netherlands Noboru Mizushima • Department of Physiology and Cell Biology, Tokyo Medical and Dental University, Tokyo, Japan and SORST, Japan Science and Technology Agency, Kawaguchi, Japan Nazanine Modjtahedi • INSERM, Unit “Apoptosis, Cancer and Immunity”, Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine – Université Paris-Sud; Villejuif, France Amitabha Mukhopadhyay • National Institute of Immunology, Aruna Asaf Ali Marg, New Delhi, India Christian Münz • Laboratory of Viral Immunobiology and Christopher H. Browne Center for Immunology and Immune Diseases, The Rockefeller University, New York, NY Thomas P. Neufeld • Department of Genetics, Cell Biology and Development, University of Minnesota, Minneapolis MN Lakshmi Ramachandra • Department of Pediatrics, Department of Pathology, and Division of Infectious Diseases, Case Western Reserve University and Rainbow Babies and Children’s Hospital, Cleveland, OH Brinda Ravikumar • Department of Medical Genetics, University of Cambridge, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge, United Kingdom Esteban A. Roberts • Department of Molecular Genetics and Microbiology, University of New Mexico School of Medicine, Albuquerque, NM

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Craig R. Roy • Section of Microbial Pathogenesis, Yale University School of Medicine, Boyer Center for Molecular Medicine, New Haven, CT David C. Rubinsztein • Department of Medical Genetics, University of Cambridge, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge, United Kingdom Adam Rupper • Department of Microbiology and Immunology, Louisiana State University Health Sciences Center, Shreveport, LA David G. Russell • Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY Giusy Sala • Laboratory of Biochemistry and Molecular Biology, San Paolo Medical School, Milan, Italy Sovan Sarkar • Department of Medical Genetics, University of Cambridge, Cambridge Institute for Medical Research, Addenbrooke’s Hospital, Cambridge, United Kingdom Francesca Scarlatti • Laboratory of Biochemistry and Molecular Biology, San Paolo Medical School, Milan, Italy Dorothee Schmid • Laboratory of Viral Immunobiology and Christopher H. Browne Center for Immunology and Immune Diseases, The Rockefeller University, New York, NY Thierry Soldati • Department of Biochemistry, University of Geneva, Geneva, Switzerland Isei Tanida • Department of Biochemistry, Juntendo University School of Medicine, Bunkyo-ku, Tokyo, Japan Ezgi Tasdemir • INSERM, Unit “Apoptosis, Cancer and Immunity,” Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine–Université Paris-Sud, Villejuif, France Sangeeta Tiwari • Section of Microbial Pathogenesis, Yale University School of Medicine, Boyer Center for Molecular Medicine, New Haven, CT Aviva M. Tolkovsky • Department of Biochemistry, University of Cambridge, Cambridge, UK Sharon Tooze • Cancer Research UK, London Research Institute, London, United Kingdom Takashi Ueno • Department of Biochemistry, Juntendo University School of Medicine, Tokyo, Japan Andreas Uttenweiler • Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Isabelle Vergne • Department of Molecular Genetics and Microbiology, University of New Mexico School of Medicine, Albuquerque, NM Ilio Vitale • INSERM, Unit “Apoptosis, Cancer and Immunity”, Villejuif, France; Institut Gustave Roussy, Villejuif, France; Faculté de Médecine – Université Paris-Sud; Villejuif, France

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Robin M. Yates • Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY Andrew Young • Cancer Research UK, London Research Institute London, United Kingdom

1 Autophagosome and Phagosome Vojo Deretic Summary Autophagy and phagocytosis are evolutionarily ancient processes functioning in capture and digestion of material found in the cellular interior and exterior, respectively. In their most primordial form, both processes are involved in cellular metabolism and feeding, supplying cells with externally obtained particulate nutrients or using portions of cell’s own cytoplasm to generate essential nutrients and energy at times of starvation. Although autophagy and phagocytosis are commonly treated as completely separate biological phenomena, they are topologically similar and can be, at least morphologically, viewed as different manifestations of a spectrum of related processes. Autophagy is the process of sequestering portions of cellular interior (cytosol and intracellular organelles) into a membranous organelle (autophagosome), whereas phagocystosis is its topological equivalent engaged in sequestering cellular exterior. Both autophagosomes and phagosomes mature into acidified, degradative organelles, termed autolysosomes and phagolysosomes, respectively. The basic role of autophagy as a nutritional process, and that of phagocytosis where applicable, has survived in present-day organisms ranging from yeast to man. It has in addition evolved into a variety of specialized processes in metazoans, with a major role in cellular/cytoplasmic homeostasis. In humans, autophagy has been implicated in many health and disease states, including cancer, neurodegeneration, aging and immunity, while phagocytosis plays a role in immunity and tissue homeostasis. Autophagy and phagocytosis cooperate in the latter two processes. In this chapter, we briefly review the regulatory and execution stages of both autophagy and phagocytosis.

Key Words: Autophagy; Phagocytosis; Atg; Vps34; Tor. 1. Autophagosome Autophagy is an evolutionarily highly conserved cellular homeostatic process whereby cells control their cytoplasmic biomass, organellar abundance, and distribution, remove potentially harmful protein aggregates, eliminate intracellular pathogens such as bacteria, protozoans, and viruses, and process self or From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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foreign proteins for antigen presentation (1–10). Autophagy comes in several forms: macroautophagy (bulk degradation of cytoplasmic components including proteins and whole organelles), microautophagy (a morphologically distinct form of autophagy often seen in yeast), and chaperone-mediated autophagy (a more subtle form of protein degradation whereby individual proteins are imported directly into lysosomes for degradation). There is also a growing number of function- or target-based classifications of autophagy with terms such as mitophagy (autophagy of mitochondria), pexophagy (autophagy of peroxisomes), ER-phagy (autophagy of the endoplasmic reticulum), xenophagy (autophagy of invading microorganisms), immunophagy (autophagy in innate and adaptive immunity), and virophagy (autophagy of cytoplasmic viruses). Unless otherwise specified, we refer in this volume to macroautophagy simply as autophagy. All cells in our bodies are capable of undergoing autophagy. Its induction is controlled by a specialized regulatory cascade, with the Ser/Thr kinase Tor at its center. When Tor is active, cells increase their biomass and proliferate, while when Tor is inhibited, cells reduce their biomass by autophagy. A short list of the inputs funneled through Tor into the regulation of autophagy is shown in Fig. 1. Once autophagy is induced, morphologically detectable execution processes (Fig. 2) begin to unfold with the following visually discernible stages: (1) initiation, whereby membranous structures (isolation membrane or phagophore) form in the cytoplasm, giving appearance of crescents; (2) elongation, during which membranes increase in size, wrap themselves around sections of the cytosol or a targeted organelle, culminating in phagophore closure and formation of a typical double membrane autophagosome characterized by two lipid bilayers (11); and (3) maturation, whereby a newly formed autophagosome fuses with endosomal organelles, forming a hybrid organelle referred to as amphisomes (11), and lysosomes, forming a terminal, fully acidified, and degradative organelle termed the autolysosome. There are also classification systems in which the autophagic pathway has been separated in additional substages, with nucleation being considered as a well-defined step in yeast. In this organism, a specialized organelle, termed the preautophagosomal structure (PAS), is identifiable as a distinct sorting station supplying components to newly initiated phagophores. A PAS equivalent has been elusive in metazoan cells, although membrane trafficking steps, theoretically leading to and from a putative PAS equivalent, can be recapitulated in mammalian cells (12). The autophagically captured material is eventually degraded in autolysosomes. Depending on the targeted cellular component, this results in (1) removal of damaged organelles (e.g., spuriously compromised mitochondria lest cells undergo an unscheduled apoptosis); (2) trimming of an organelle (e.g., ER) to its appropriate size; (3) generation of free amino acids and energy sources to maintain essential cellular anabolic needs at times of nutrient or energy


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Fig. 1. Core signaling pathways controlling autophagy. Activators of Tor (black letters, white boxes) suppress autophagy, while inhibitors of Tor promote autophagy (white letters, gray boxes). In yeast, Tor negatively regulates Atg1, a factor that sets off the autophagy execution cascade. Hence, when Tor is inhibited by rapamycin, autophagy is induced. Tor is normally regulated by a small GTPase, Rheb, which binds to the N-terminal portion of the Tor kinase domain. As with other small signaling GTPases, which act as molecular switches, the GDP-bound form of Rheb is in the OFF position, while the GTP bound form is in the ON position. Rheb activity is regulated by the GTPase activating protein (GAP) TSC1/2, which receives and integrates various upstream inputs (28) (Fig. 3) that come from (1) growth factor receptor signaling via the Akt/PKB pathway, inhibiting autophagy; (2) energy status via AMPK, a kinase that responds to the AMP/ATP ratio, with active AMPKstimulating autophagy (29), and the recently recognized contributor REDD1, which acts independently of AMPK (30); and (3) Ca2+ effects on CaMKK- and phosphorylation and activation of AMPK and thus induction of autophagy (31). The best understood pathway controlling TSC1/2 is the one stimulated by growth factors. Binding of growth factors to receptors activates type I PI3K p110/p85, which generates phosphatidylinositol-(3,5)-P3 (PIP3). PIP3 recruits PDK1 and Akt/PKB to the plasma membrane, where PDK1 phosphorylates and activates Akt/PKB. Active PKB phosphorylates and inactivates TSC1/2. By inactivating the TSC1/2 GAP, this cascade enhances Rheb-GTP–dependent activation of Tor and phosphorylation of Tor targets. This in turn inhibits autophagy. If growth factors or amino acids are withheld,

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deprivation; (4) removal of intracellular pathogens including bacteria, protozoans, and viruses; (5) capture and processing of self antigens for endogenous antigen presentation in an MHC II–restricted fashion; etc. Many aspects of the molecular machinery controlling autophagy have been delineated, and specific Atg factors have been characterized. In yeast, they number in excess of 30 genes, but in mammalian cells their orthologs, identifiable through bioinformatics, are limited to just over a dozen (not counting multiple isoforms), with many more clearly remaining to be identified as the studies progress. Some of these factors are shown in Fig. 2. 2. Phagosome In contrast to autophagy (a process of phagocytosis of objects/targets already in the cellular interior), conventional phagocytosis (13) represents uptake and internalization of objects initially located outside of the cell (Fig. 3). The two processes (autophagy and phagocytosis) are initiated and driven by different mechanisms. Nevertheless, there is an overlap during the terminal stages in both pathways (Figs. 2 and 3), when autophagosomes become autolysosomes or when conventional phagosomes mature into phagolysosomes. While all cells in our body are capable of undergoing autophagy, most cells, apart from the specialized phagocytic cells, are normally not particularly active in phagocytosis. Even so, they can be “coerced” into doing so in some cases: (1) often, in tissues during organ development, neighboring cells may phagocytose bystander dying cells in the process of organ cavity formation (14); (2) in vitro manipulations can make a nonphagocytic cell actively phagocytic, often requiring expression of a receptor or additional molecules (15); (3) bacteria and other pathogens can stimulate processes in host cells to make them engulf and take up microbes (16). Nevertheless, a strong, prominent phagocytic function is reserved for the cells of the reticuloendothelial system, and macrophages are usually considered to be the prototypic phagocytic cells. Fig. 1. (Continued) autophagy is augmented. Recent work (32) has identified a long missing nucleotide exchange factor for Rheb, which loads Rheb with GTP, thus providing another arm of Rheb regulation, and hence Tor activation (which should result in autophagy inhibition), which yet remains to be explored. Recognized only recently, but probably representing an ancient signaling mechanism, hVPS34 appears to transduce the amino acid–replete conditions to Tor and in this capacity plays a negative role in signaling upstream of autophagy initiation (33,34). However, once autophagy is initiated, hVPS34 complexed with Beclin 1 and PI3P play an essential role in the execution of autophagy. Autophagy can be induced by rapamycin or amino acid starvation and inhibited by 3-methyl adenine, an inhibitor of PI3K.


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Fig. 2. Execution stages of autophagy. Initiation: A nascent autophagosomal organelle, termed a phagophore (isolation membrane), corrals an organelle or a section of the cytoplasm. Elongation: The phagophore elongates and bends, wrapping around the cytoplasmic target to be sequestered. At this stage, Atg factors form two distinct protein–protein or protein–lipid conjugates: (1) Atg5 is covalently linked to Atg12, and the resulting Atg5-Atg12 conjugate associates with Atg16; (2) Atg8 (LC3) is converted from its cytosolic LC3-I form into a C-terminally phosphatidylethanolamine (PE) conjugated form, LC3-II. This stage culminates with phagophore closure and formation of double-membrane (two lipid bilayers) autophagosome. Maturation (flux): During this stage, the autophagosomal pathway merges with the endosomal/lysosomal pathway. Autophagosomes fuse with endosomes (generating amphisomes) and lysosomes, finally being converted into fully lytic, acidified organelles termed autolysosomes. The inner

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Fig. 3. Conventional phagosomes. Depicted are stages in the formation and maturation of a conventional phagosome in a macrophage (25). Several small GTPases from the family of Rab proteins have been implicated at different stages of phagolysosome biogenesis. The phagolysosome represents the default terminal stage of phagosomal maturation pathway, unless it is blocked by factors produced by intracellular pathogens. Phosphoinositides implicated at different stages are shown. PA, phosphatidic acid; DAG, diacylglycerol. EEA1 and Hrs are PI3P-binding proteins acting at different stages of phagosomal maturation.

Phagocytosis studies can be subdivided into investigations of (1) particle recognition by receptors and particle uptake and (2) phagolysosome biogenesis, which involves maturation of the initially formed, nondegradative phagosome into a degradative organelle. It is, however, hard not to draw a parallel with autophagy, if one simply substitutes external space with particles (the principal target of phagocytosis) for an internal space with an organelle or cytosol (the conventional targets of autophagy). There are also physiological overlaps, because phagocytosis and autophagy have common functions when it comes to eliminating intracellular pathogens (17–19) and antigen Fig. 2. (Continued) membrane has been disrupted, and the autolysosome has only the outer, delimiting membrane. The density of the material in autophagosomes resembles that in the surrounding cytosol, but autolysosomal lumen appears denser. Often, internal membranes can be seen, representing remnants of organelles captured by autophagy. MVB, multivesicular bodies; LE, late endosome; Lys, lysosome.


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presentation (20,21). The two pathways start, based on our current understanding, using different machinery, but end in a similar way with a degradative organelle and engagement of similar regulators such as the phosphatidylinositol-3-kinase hVPS34. It is of note that phosphoinositides (22), and in particular phosphatidylinositol-3-phosphate (PI3P), play an essential role in phagolysosome biogenesis (23–25). Furthermore, PI3P is a key element in both initiation and maturation of autophagosomes. The lipid kinase hVPS34 has specific interacting partners (e.g., Beclin1 (26)) that modulate its function: in yeast there are two hVPS34 complexes, one involved in autophagy, and the other controlling conventional endosomal pathway. The hVPS34 complexes in mammalian cells may be similarly specialized or perhaps exist in more than two forms, regulating various processes including autophagy, the endosomal pathway, and maturation of conventional phagosomes. The mammalian factors participating in hVPS34 complexes are beginning to be identified, and presently include UVRAG (27) and at least three additional elements with or without obvious equivalents in the yeast.

Acknowledgments The editor wishes to thank immensely Michal Mudd. She has tirelessly played a pivotal role at all stages of bringing this project to a conclusion. This work was supported by NIH grants AI069345, AI45148, and AI42999.

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9. Yoshimori, T. (2007) Autophagy: paying Charon’s toll. Cell 128:833–836. 10. Kamada, Y., Sekito, T., and Ohsumi, Y. 2004) Autophagy in yeast: a TOR-mediated response to nutrient starvation. Curr. Top. Microbiol. Immunol. 279:73–84. 11. Fengsrud, M., Erichsen, E. S., Berg, T. O., Raiborg, D., and Seglen, P. O. (2000) Ultrastructural characterization of the delimiting membranes of isolated autophagosomes and amphisomes by freeze-fracture electron microscopy. Eur. J. Cell Biol. 79:871–882. 12. Young, A. R., Chan, E. Y., Hu, X. W., et al. (2006) Starvation and ULK1dependent cycling of mammalian Atg9 between the TGN and endosomes. J. Cell Sci. 119:3888–3900. 13. Aderem, A., and Underhill, D. M. (1999) Mechanisms of phagocytosis in macrophages. Annu. Rev. Immunol. 17:593–623. 14. Qu, X., Zou, Z., Sun, Q., Luby-Phelps, K., et al. (2007) Autophagy gene-dependent clearance of apoptotic cells during embryonic development. Cell 128:931–946. 15. Joiner, K. A., Fuhrman, S. A., Miettinen, H. M., Kasper, L. H., and Mellman, I (1990) Toxoplasma gondii: fusion competence of parasitophorous vacuoles in Fc receptor-transfected fibroblasts. Science 249:641–646. 16. Cossart, P., and Sansonetti, P. J. (2004) Bacterial invasion: the paradigms of enteroinvasive pathogens. Science 304:242–248. 17. Gutierrez, M. G., Master, S. S., Singh, S. B., Taylor, G. A., Colombo, M. I., and Deretic, V. (2004) Autophagy is a defense mechanism inhibiting BCG and Mycobacterium tuberculosis survival in infected macrophages. Cell 119:753–766. 18. Singh, S. B., Davis, A. S., Taylor, G. A., and Deretic, V. (2006) Human IRGM induces autophagy to eliminate intracellular mycobacteria. Science 313:1438–1441. 19. Alonso, S., Pethe, K., Russell, D. G., and Purdy, G. E. (2007) Lysosomal killing of Mycobacterium mediated by ubiquitin-derived peptides is enhanced by autophagy. Proc. Natl. Acad. Sci. USA 104:6031–6036. 20. Schmid, D., Pypaert, M., and Munz, C. (2007) Antigen-loading compartments for major histocompatibility complex class II molecules continuously receive input from autophagosomes. Immunity 26:79–92. 21. Ramachandra, L., Noss, E., Boom, W. H., and Harding, C. V. (2001) Processing of Mycobacterium tuberculosis antigen 85B involves intraphagosomal formation of peptide-major histocompatibility complex II complexes and is inhibited by live bacilli that decrease phagosome maturation. J. Exp. Med. 194:1421–1432. 22. Pizarro-Cerda, J., and Cossart, P. (2004) Subversion of phosphoinositide metabolism by intracellular bacterial pathogens. Nat. Cell Biol. 6:1026–1033. 23. Fratti, R. A., Backer, J. M., Gruenberg, J., Corvera, S., and Deretic, V. (2001) Role of phosphatidylinositol 3-kinase and Rab5 effectors in phagosomal biogenesis and mycobacterial phagosome maturation arrest. J. Cell Biol. 154:631–644. 24. Vieira, O. V., Botelho, R. J., Rameh, L., et al. (2001) Distinct roles of class I and class III phosphatidylinositol 3-kinase in phagosome formation and maturation. J. Cell Biol. 155:19–25. 25. Vieira, O. V., Botelho, R. J., and Grinstein, S. (2002) Phagosome maturation: aging gracefully. Biochem. J. 366:689–704.


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26. Pattingre, S., Tassa, A., Qu, X., et al. (2005) Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122:927–939. 27. Liang, C., Feng, P., Ku, B., et al. (2006) Autophagic and tumour suppressor activity of a novel Beclin1-binding protein UVRAG. Nat. Cell Biol. 8:688–699. 28. Wullschleger, S., Loewith, R., and Hall, M. N. (2006) TOR signaling in growth and metabolism. Cell 124:471–484. 29. Meley, D., Bauvy, C., Houben-Weerts, J. H., et al. (2006) AMP-activated protein kinase and the regulation of autophagic proteolysis. J. Biol. Chem. 281:34870–34879. 30. Sofer, A., Lei, K., Johannessen, C. M., and Ellisen, L. W. (2005) Regulation of mTOR and cell growth in response to energy stress by REDD1. Mol. Cell Biol. 25:5834–5845. 31. Hoyer-Hansen, M., Bastholm, L., Szyniarowski, P., et al. (2007) Control of macroautophagy by calcium, calmodulin-dependent kinase kinase-beta, and Bcl-2. Mol. Cell 25:193–205. 32. Hsu, Y. C., Chern, J. J., Cai, Y., Liu, M., and Choi, K. W. (2007) Drosophila TCTP is essential for growth and proliferation through regulation of dRheb GTPase. Nature 445:785–788. 33. Byfield, M. P., Murray, J. T., and Backer, J. M. (2005) hVps34 is a nutrientregulated lipid kinase required for activation of p70 S6 kinase. J. Biol. Chem. 280:33076–33082. 34. Nobukuni, T., Joaquin, M., Roccio, M., et al. (2005) Amino acids mediate mTOR/raptor signaling through activation of class 3 phosphatidylinositol 3OHkinase. Proc. Natl. Acad. Sci. USA 102:14238–14243.

2 Fine Structure of the Autophagosome Eeva-Liisa Eskelinen

Summary This chapter describes the electron microscopic fine structure of early and late autophagic vacuoles in mammalian cells. Detailed instructions are given for the preparation of cells for conventional electron microscopy and for the identification of autophagic vacuoles by morphology. Electron microscopy remains one of the most accurate methods for quantitation of autophagic vacuole accumulation. Therefore, quantitation of autophagic vacuoles by electron microscopy and point counting is also described. Finally, a short description is given for preparation of ultra thin cryosections for immunogold labeling of autophagic vacuoles.

Key Words: Autophagy; electron microscopy; point counting; volume fraction; cryosectioning, immunoelectron microscopy. 1. Introduction 1.1. The Significance of Autophagy Autophagy is a lysosomal degradation pathway for cytoplasmic material (1,2) that is important for survival during short-term starvation. By degrading some nonessential components, cells get nutrients for energy production and vital biosynthetic reactions. Autophagy is essential for energy metabolism during starvation (3,4) and immediately after birth (5) and for cell homeostasis in muscle, liver, and pancreas (6,7). Autophagy contributes to growth regulation—impaired autophagy can lead to cancer (8,9)—and to longevity (10). Further, autophagy has been shown to reduce the toxicity of the protein aggregates in Huntington’s disease (11). In the central nervous system, inhibition of autophagy causes neurodegeneration (12,13). In addition, autophagy plays a role in innate immunity in defense against viral infection (14) and intracellular bacteria (15). From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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1.2. The Autophagic Pathway After an induction signal such as starvation, autophagy starts when a flat membrane cistern wraps around a portion of cytoplasm and forms a closed double-membraned vacuole containing cytosol and/or organelles (16). The membrane cistern has been called the phagophore or the isolation membrane. The sealed vacuole is called the autophagosome and is devoid of any lysosomal proteins. Autophagosomes mature by fusing with endosomal and lysosomal vesicles, which also deliver lysosomal membrane proteins and enzymes (17,18). The segregated cytoplasm is then degraded by lysosomal hydrolases, and the degradation products are transported back to cytoplasm. A schematic drawing of autophagy is presented in Fig. 1. The term amphisome refers to an autophagosome that has fused with an endosome, and autolysosome refers to an autophagosome or amphisome that has fused with a lysosome. The general term autophagic vacuole refers to an autophagosome, amphisome, or autolysosome. In electron microscopy, autophagic vacuoles can be identified as membrane-bound vesicles containing cytoplasmic material or organelles (1). Morphologically, autophagic vacuoles can be further classified into early or initial autophagic vacuoles (AVi),

Fig. 1. Schematic presentation of the formation and maturation of autophagosomes in mammalian cells. The nomenclature of the different autophagosome stages is given on the left side of the drawing. Further details are given in the text.

Fine Structure of the Autophagosome

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containing morphologically intact cytosol or organelles, and to late or degradative autophagic vacuoles (AVd), containing partially degraded cytoplasmic material (17,19,20). (The morphology of early autophagic vacuoles is demonstrated in Figs. 4A,B, 5B, and 6A,B. Late autophagic vacuoles are shown in Figs. 5B and 6A,B.) 1.3. Identification of Autophagic Vacuoles by Light and Electron Microscopy Several novel yeast genes essential for autophagy (Atg genes) were recently characterized (21). Many mammalian homologs have been identified for these yeast genes. The microtubule-associated protein 1 light chain 3, or LC3, was shown to be the mammalian homologue of Atg8 (22). LC3 is the only known marker protein for autophagic vacuoles. Anti-LC3 has been used as a marker for autophagosomes, especially in light microscopy, but also in electron microscopy (23). LC3 is present mainly in autophagosomes. Because it is degraded by the incoming lysosomal hydrolases, less LC3 is present in amphisomes and autolysosomes (23). In electron microscopy, autophagic vacuoles can also be identified by morphology, particularly when using samples embedded in conventional plastic resins. This chapter deals with the fine structure of autophagic vacuoles in plastic embedded samples, with a short description of the quantitation of autophagy. Finally, reference is made to the fine structure of autophagic vacuoles in thin cryosections which are used for immunogold labeling. 2. Material 1. Fixation for Plastic Embedding: 2% glutaraldehyde (electron microscopy grade) in 0.2 M hydroxyethyl piperazine ethane sulfonate (HEPES), pH 7.4. 2. Fixation for Cryosectioning and Immunolabeling: 4% paraformaldehyde (PFA, use the powder form) in 0.2 M HEPES, pH 7.4. This solution may contain 0.05–0.2% glutaraldehyde (electron microscopy grade). 3. Resin for Flat Embedding: Agar 100 (Agar Scientific, Stansted, UK) and LX-112 (Ladd Research, Williston, VT, USA) have been tested with the flat embedding protocol. Other Epon equivalent resins may work as well, but may cause problems with cells seeded on plastic. Many resins partially dissolve plastic, and thus it will become very difficult to detach the block from the dish. Therefore, glass cover slips are recommended when other Epon equivalent resins are used for flat embedding. 4. Resins for Embedding of Cell Pellets: a) Durcupan mixture (Fluka): Component A: 10 g B: 10 g C: 0.3 g D: 0.3 g

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Eskelinen Mixture can be stored at −20°C for some days. Do not open the container before the mixture has reached room temperature. b) Agar 100 mixture (Agar Scientific): see manufacturer’s instructions. Use the medium formula. Other Epon equivalent embedding media will probably work as well.

5. 1.84 M Sucrose–20% PVP for Cryoprotection: Prepare 2.3 M sucrose in phosphate-buffered saline (PBS) by dissolving 39.365 g sucrose in 50 mL PBS. This takes some hours with constant mixing. You may gently warm the solution (37°C) to enhance dissolving. Put 40 mL of 2.3 M sucrose in a 100-mL Erlenmeyer flask. Add 10 g polyvinylpyrrolidone (PVP) (Sigma, Mw 10,000). Mix with a magnetic stirrer overnight to dissolve. Check the pH with a pH paper—if necessary, adjust the pH to 7.4 using 1.1 M Na2 CO3 (usually less than 1 mL is necessary). For 1.7 M sucrose–15% PVP, use 2.1 M sucrose and add 7.5 g PVP.

3. Methods 3.1. Resin Flat Embedding of Aldehyde-Fixed Animal Cell Cultures 1. Grow cells on plastic dishes or glass cover slips. Incubate cells in serum and amino acid free medium for 1–2 h to induce autophagy. From step 2 onward, everything is done in a fume hood, with gloves. Do not let the cells dry out at any point during the procedure. 2. Fix the cells by changing the culture medium to 2% glutaraldehyde in 0.2 M HEPES, pH 7.4. Incubate at room temperature (RT) for 2 h. 3. Wash the cells in 0.2 M HEPES (and storage at 4°C for up to some weeks). 4. Wash the cells in PBS three times. 5. Postfix in 1% Osmium tetroxide in water at RT for 1 h. Osmium is very toxic and volatile; use a good fume cupboard and gloves. 6. Wash the cells in water twice. 7. Stain the cells in 2% uranyl acetate in water at RT, in the dark, for 1 h. This step gives contrast to autophagosome membranes. 8. Dehydration (RT): 70% ethanol in water 3 times 3 min 95% ethanol 3 times 3 min 100% ethanol 3 times 3 min 9. Optional: Dip cover slips in propylene oxide. Proceed quickly to step 10, as propylene oxide evaporates very quickly. Do not use propylene oxide on plastic dishes—it dissolves plastic. 10. Infiltration of resin: Aspirate almost all ethanol from the cells. Add a few drops of 100% resin and swirl the dish to spread the resin to the cells. The resin layer should cover the cells, but it should not be thick. Optimally add less than 1 mm layer of resin. Incubate for 1–2 h at RT, until all ethanol has evaporated. 11. Fill gelatin or Beam capsules with resin. Place the capsules upside down to the cells. Make sure no air bubbles stay between the cells and the resin. It is


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possible to check the cells with an inverted phase contrast microscope and select the desired area for the capsules. 12. Polymerize the resin at 45°C overnight and then at 60°C for one day. 13. The capsules can be released from the plastic dishes by gentle heating on a hot plate, then gently swirling the capsule. Glass cover slips can be removed by dipping the glass to liquid nitrogen for 5–10 s, and then knocking on the glass to remove it from the block. Make sure all glass is removed before using a diamond knife to cut sections. 14. Cut 70- to 80-nm sections with a diamond knife, and stain the sections with uranyl acetate and lead citrate. See Notes 1–3 on the fine structure of autophagosomes. Option 1. Use reduced osmium tetroxide instead of osmium tetroxide in water (step 5). This enhances the contrast of autophagosome-limiting membranes, but reduces the contrast of ribosomes. Thus, the later maturation stages of autophagic vacuoles are more difficult to identify. Protocol: Mix 2% osmium tetroxide in water and 0.2 M sodium cacodylate, pH 7,4, to give the final concentrations of 1% osmium tetroxide and 0.1 M cacodylate, and add 15 mg/mL K4 Fe(CN)6 . Incubate at RT for 1 h. Option 2. Use sucrose in the uranyl acetate en block staining solution (step 7): 1 % uranyl acetate, 0.3 M sucrose in water, 1 h, 4°C. This may reduce extraction of some sample components.

3.2. Resin Embedding of Aldehyde-Fixed Animal Cell Pellets for Quantitation of Autophagy 1. Use 6-cm culture dishes. The cells should be semi-confluent. Fix cells in 2% glutaraldehyde in 0.2 M HEPES, pH 7.4 at RT for 2 h. Scrape the cells from culture plate using a razor blade after 30 min fixation. Pellet the cells at full speed in an Eppendorf centrifuge for 5 min. Continue fixation as a pellet. Do not resuspend the pellet during the rest of embedding, but use gentle mixing during incubations. 2. Wash the pellets in 0.2 M HEPES (and store at 4°C). 3. Wash in PBS three times. 4. Postfix the pellets in 1% osmium tetroxide in water at RT for 1 h. 5. Wash in water twice. 6. Stain the pellets in 2% uranyl acetate in water at RT, in the dark, for 1 h. 7. Dehydration (RT) (mixing is important during dehydration): 70% ethanol in water 15 min 95% ethanol 15 min 100% ethanol twice for 15 min propylene oxide 20 min (toxic and volatile) 8. Infiltration of resin: resin + propylene oxide (1 + 1) at RT for 2 h 100% resin overnight, RT

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Fig. 2. Principles of autophagic vacuole quantitation and point counting. (A) Grid squares can be used as sampling units. It is recommendable to count at least three squares from each sample. The selected squares should be uniformly distributed over the section. When a small magnification (400–600×) is used for this selection, selection bias is easier to avoid because autophagic vacuoles are not visible at low magnification.


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9. Transfer the pellets to fresh resin in embedding molds and incubate for 4–6 h, RT. 10. Polymerize the blocks at 60°C for 2 days. 11. Cut 80-nm sections with a diamond knife and stain with uranyl acetate and lead citrate. See Notes 1–3 on the fine structure of autophagosomes.

3.3. Quantitation of Autophagic Vacuoles in Thin Sections For quantitative estimation of the autophagic accumulation, the orientation of the cells in the sections should be random. This is best achieved when using cell pellets. If flat embedded cells are used, the counting should be done using sections cut from more than one plane, which should locate a few micrometers from each other. This is because it is possible that the structures of interest are more concentrated in a certain region of the cytoplasm (for instance, close to the culture surface or above the nucleus). This problem is most easily avoided when using randomly oriented sections, which can be done using cell pellets. There are two alternatives for the electron microscopic quantitation. First, the volume fraction, e.g., fraction of the cellular volume occupied by autophagic vacuoles, can be estimated. This can be accurately and efficiently estimated by point counting (24) (Fig. 2). Second, it is possible to estimate the number of autophagic vacuole profiles per cell area on sections. The advantage of the latter is that is saves a considerable amount of work, since only the cell volume needs to be estimated by point counting. Square or hexagonal grid openings can be used as sampling units in the quantitation (Fig. 2). The whole grid square is systematically scanned under the microscope for the presence of early and late autophagic vacuoles (Fig. 2B). When estimating the number of vacuoles per cell area, only the number of vacuoles in the grid square is recorded. When estimating volume fraction, each autophagic vacuole is photographed at 12.000× magnification for point counting of the area Fig. 2. (Continued) (B) Screening of the selected grid squares for the presence of autophagic vacuole profiles at higher magnification 12,0000×. (C) Point counting. A square lattice is placed on top of a microscopic photograph or photographic negative. The intersections of the lattice lines are used as test points. Points hitting the structure of interest (autophagic vacuole, or cell in a cell pellet) are counted. The distance between points (d) is calibrated with the final magnification of the photograph. These two parameters are used to calculate the area of the structure, as indicated below the drawing. This procedure can be done on the computer screen, if the photographs are digital. The lattice is created using the computer program. If the microscope is using a film camera, point counting can be performed using a light box. The lattice is printed on an overhead foil which is then placed on top of the film sheet. A magnifying loop can be used to help counting.

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of the vacuoles (point counting 1; Fig. 2C). This can be done directly from photographic negatives if a digital camera is not available in the microscope. The cell area can be measured using about 400× magnification, which allows the whole grid square to fit in one photograph. The area of the cell profiles is then estimated using point counting with a different lattice (point counting 2; Fig. 2C). Finally, the number of vacuoles per cell area is simply calculated by dividing the number of vacuoles in the grid square with the cell area in the same grid square. Similarly, the volume fraction is calculated by dividing the area occupied by autophagic vacuoles (from point counting 1) with the cell area in that grid square (from point counting 2). Thus, the ratio of areas directly gives the ratio of volumes (25). It is recommendable to count at least three grid squares for each sample to get a representative estimate (Fig. 2A). These squares should locate evenly over the whole thin section and should be selected using systematic random sampling (25). In most cases, the number of autophagic vacuole profiles per cell area gives a very similar result compared to the stereologically more accurate volume fraction (Fig. 3A,B). Since the former saves work, it is better suited for experiments containing several samples. It should be kept in mind, however, that the number of profiles per cell area is not equal to the number of vacuoles per cell area. This is because large vacuoles have a greater probability than small ones to hit the thin section. For more exact discussion, please refer to stereology literature (25). For a general discussion of quantitation of autophagic vacuoles, see Notes 4 and 5. 3.4. Preparation of Cultured Cells for Cryosectioning and Immunogold Labeling Use confluent/subconfluent cultures. There are basically two options for fixation: paraformaldehyde mixed with a low concentration of glutaraldehyde or paraformaldehyde only. Glutaraldehyde gives better ultrastructural preservation, but not all antigens or antibodies tolerate glutaraldehyde treatment. Use 4% paraformaldehyde (PFA) in 0.2 M HEPES, pH 7.4, and fix for 1–2 h at room temperature. This solution may contain 0.05–0.2% glutaraldehyde. If you use 4% PFA without glutaraldehyde, incubate the cells in this solution for 2 h, followed by 2% PFA in 0.2 M HEPES at 4°C overnight. In both options, transfer the cells to a fresh buffer after fixation. For long-distance shipping, leave the cells in 2% PFA in 0.2 M HEPES. Gelatin embedding helps to preserve the fine structure of cells during sectioning. Gelatin embedding is not recommended if you wish to label lipid antigens. For gelatin embedding, first wash the cells in PBS. Then scrape the cells from the culture dish in PBS and transfer to an Eppendorf tube. Pellet the cells at full speed in an Eppendorf centrifuge. Add 200 μL 10% gelatin in


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Fig. 3. Estimation of autophagic vacuole accumulation in mouse embryonic fibroblasts. The cells were fixed without treatment (FSC), or incubated in serum and aminoacid–free medium for 30 min (EBS 0.5 h) or 2 h (EBS 2 h), or in serum-free culture medium for 2 h (MEM 2 h). Cells were fixed and prepared for electron microscopy according the protocol given in Subheading 3.2. Accumulation of autophagic vacuoles was estimated both as volume fraction (A) and as number of profiles per cell area (B). The number of profiles is given per 100 μm2 . AVi, initial autophagic vacuoles containing morphologically intact cytoplasm; AVd, late autophagic vacuoles containing partially degraded, but still identifiable, cytoplasmic material.

PBS at 37°C and mix gently with the pipet tip. Incubate at 37°C for 5 min. Pellet the cells again at room temperature (suitable speed needs to be tested for each cell type). Remove the gelatin from the tube, and add a fresh 200 μL of 10% gelatin, and mix with the tip. Pellet again at room temperature to get a firm pellet. Keep the tube in ice for 30 min to set the gelatin. Take a small spatula and dip it to sucrose-PVP (see below). Then use the wet spatula to transfer the gelatin and cells to a Petri dish on ice, and cut the cell pellet to small cubes/sticks (maximum side length about 2 mm).

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Before freezing, the cells need to be incubated in a cryoprotectant. 2.1–2.3 M sucrose in PBS or a mixture of sucrose and PVP in PBS (1.84 M sucrose–20% PVP, or 1.7 M sucrose–15% PVP) are mostly used. PVP gives plasticity to the block and thus helps cryosectioning. Incubate the cell cubes in sucrose-PVP at 4°C, with occasional mixing, for several hours to overnight, until the cubes sink to the bottom. Trim the cubes under a stereo microscope and mount on cryoultramicrotome holders at 4°C (in a cold room, or using an ice bath). For optimal freezing, the maximum size of the cubes should be about 1 mm. Freeze the mounted cubes by punching into liquid nitrogen. Store the samples in liquid nitrogen until sectioning. Cut 70- to 80-nm sections at −100°C using a cryoultramicrotome and a cryodiamond knife. Use a mixture of sucrose and 1.5% methyl cellulose (1 + 1–4 + 1) to pick up the sections. Compared to picking up sections with sucrose only, this greatly improves the ultrastructure of the cells (26). Transfer sections on carboncoated Formwar grids and leave the extra sucrose–methyl cellulose on the grids. The grids can be stored section side up, at 4°C, for several months. To remove the sucrose–methyl cellulose, incubate the grids on PBS twice for 5 min. Immunogold label the sections using standard procedures for Tokuyasu sections (24). Finally embed the sections in methyl cellulose–uranyl acetate. Notes 6 and 7 deal with the use of cryosections to study autophagy. 4. Notes 4.1. Fine Structure of Autophagosomes and Autophagic Vacuoles in Plastic Sections 1. By definition, autophagosomes are double-membrane limited vacuoles that contain undegraded cytoplasm and no lysosomal proteins. In plastic sections, the two limiting membranes are often so close to each other that it is not possible to see them as separate membranes (Fig. 4A, large arrowheads). Sometimes the limiting membrane may appear to contain multiple layers (Fig. 4B, arrow). It is possible that this is an artifact caused by the chemical fixation. On the other

Fig. 4. Fine structure of autophagosomes (early/initial autophagic vacuoles) in plastic embedded hepatocytes isolated from mouse liver. The cells were incubated in serum and amino acid free medium to induce autophagy. (A) The autophagosome on top of the panel contains a mitochondrion, endoplasmic reticulum, and ribosomes. The limiting membrane (large arrowheads) is visible only partially as a double membrane (arrows). Below the autophagosome, a U-shaped membrane cistern, a putative phagophore (small arrowhead), seems to be in the process of wrapping around a peroxisome (p). (B) Rough endoplasmic reticulum


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Fig. 4. (Continued) tightly follows part of the limiting membrane of the autophagosome (small arrowheads). Another rough cistern is located inside the autophagosome, on the other side of the limiting membrane (large arrowheads). The arrow indicates a region where the autophagosome limiting membrane seems to consist of multiple layers. This may be an artifact caused by aldehyde fixation. (From ref. 1.)

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hand, it is also possible to see the two limiting membranes in plastic sections. In these cases, there is a narrow empty (electron lucent) space between the two membranes (Figs. 5B, 6A,B). Sometimes the autophagosome limiting membrane does not have any contrast in thin sections. This is probably caused by extraction of lipids during sample preparation. Occasionally, it is possible to observe putative phagophores, U-shaped membrane cisterns which seem to be engulfing portions of cytoplasm (Fig. 4A, small arrowhead, and Fig. 5A, asterisks). In these cisterns the two separate membranes are usually clearly visible. Typically, a cistern of rough endoplasmic reticulum (ER) is located close to the autophagosome or phagophore. Another rough cistern is often located on the other side of the autophagosome or phagophore limiting membrane (Figs. 4B and 5A, small and large arrowheads). Thus, the limiting membrane runs in the space between these two rough ER cisterns. The role of this autophagosome-associated ER, if any, is currently unknown. 2. The cytoplasmic contents of autophagosomes include organelles, such as ER membranes and mitochondria (Fig. 4A,B). Ribosomes are frequently seen inside autophagosomes (Figs. 4A,B, 5B, and 6A,B) and serve as a good marker structure for the cytoplasmic contents. The diameter of autophagosome profiles varies between 300 and 400 nm and several micrometers. In many cultured cells the average diameter is about 600 nm. Autophagosomes are frequently observed in fusion profiles with endosomal or lysosomal vesicles (18). Autophagosomes can also fuse with each other, or several autophagosomes can fuse with a single endo/lysosomal vesicle or late autophagic vacuole (Fig. 5B). In the fusion event, the outer limiting membrane fuses with the endo/lysosome limiting membrane. The contents, still surrounded by the inner limiting membrane, are delivered to the endo/lysosome lumen. This membrane must then be degraded, or at least permeabilized, to allow degradation of the cytoplasmic contents. In late autophagic vacuoles, partially degraded electron-dense ribosomes serve as a good criterion for identification of the cytoplasmic contents (Fig. 5B, asterisks, and Fig. 6A,B,

Fig. 5. Fine structure of putative phagophores and autophagic vacuoles in plastic embedded mouse embryonic fibroblasts. The cells were incubated in serum and amino acid free medium for 2 h to induce autophagy. (A) Two putative phagophores. Rough endoplasmic reticulum surrounds both phagophores (large arrowheads). Another rough cistern is located on the other side of the phagophore membrane sack in both phagophores (small arrowheads). The asterisks (*) indicate the empty space in the lumen of the phagophore, between the two limiting membranes of the coming autophagosome. (B) Three autophagosomes (arrows) in the process of fusion with a late autophagic vacuole. Asterisks indicate so-called autophagic bodies, portions of partially degraded cytoplasm still partially surrounded by the autophagosome inner limiting membrane. These can be identified by their contents of electron-dense ribosomes. Ribosomes become more electron dense as a consequence of degradation.


Fig. 5. (Continued )

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Fig. 6. Fine structure of early and late autophagic vacuoles in plastic embedded mouse embryonic fibroblasts. The cells were incubated in serum and amino-acid– free medium for 2 h to induce autophagy. Early autophagic vacuoles (AVi) contain morphologically intact cytoplasm, which looks identical to the cytoplasm surrounding the vacuoles. Late autophagic vacuoles (AVd) contain material which can still be recognized as cytoplasmic, in this case ribosomes, and looks partially degraded, i.e., more electron dense than the cytoplasm surrounding the vacuole.

AVd). The cytoplasmic contents often appear partly degraded, even when the inner limiting membrane still seems to be intact (Fig. 6A, AVd). 3. It should be noted that special attention must be paid when identifying autophagic vacuoles in transmission electron microscopy. There are several examples in the current literature of different organelles, including multilamellar or multi-


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vesicular endosomes, even mitochondria, being claimed as autophagic vacuoles. Only vacuoles containing cytoplasmic material, in most cases ribosomes, can be claimed as autophagic.

4.2. Quantitation of Autophagy by Electron Microscopy 4. Before discovery of the Atg proteins, autophagy could only be detected using electron microscopy or biochemical methods (1). Even today, quantitative electron microscopy is one of the most sensitive methods to detect the accumulation of autophagic vacuoles. In addition, it is possible to detect the nature of the accumulating vacuoles, e.g., whether early autophagic vacuoles (AVi) or more matured late autophagic vacuoles (AVd) accumulate (1). The ratio of AVi/AVd gives clues as to the cause of the accumulation. If AVi predominate, there is probably a defect in the maturation of autophagosomes into degradative autophagic vacuoles. On the other hand, large accumulation of AVd suggests that there may be a defect in autolysosome formation. It should be noted that most of the AVd are probably amphisomes, since autolysosomes are likely short lived and thus less frequently observed by microscopy. 5. In cultured animal cells, the accumulation of autophagic vacuoles becomes detectable 15–30 min after the cells have been switched to starvation medium. The accumulation increases until 2 h starvation, but longer starvation times do not seem to increase the net accumulation, unless proteinase inhibitors or microtubule drugs are added to the incubation medium to prevent degradation in or formation of autolysosomes (18).

4.3. Autophagic Vacuoles in Thin Cryosections 6. Postembedding immunogold labeling with thin cryosections is one of the most sensitive methods to immunolabel antigens for electron microscopy (24). Concerning autophagy detection, the drawback of cryosections is that ribosomes have no contrast in these sections. This makes identification of autophagosomes more demanding, because without ribosomes it is difficult to identify the cytoplasmic contents. Also, classification of AVi and AVd is difficult in cryosections. Immunolabeling of the autophagosome marker LC3 can be used to help identification (23), but good antibodies are not always available. The level of endogenous LC3 is also relatively low for clear-cut immunogold labeling. Identification of autophagic vacuoles can still be achieved by morphology, at least in cases where clearly identifiable organelles such as mitochondria are seen among the contents (Fig. 7). Immunolabeling of lysosomal membrane proteins and enzymes can be used to detect fusion with endosomal or lysosomal vesicles (Fig. 7). 7. Some cell types are more suitable for cryosectioning than others. Isolated hepatocytes, for instance, give a good morphology in cryosections even after fixation in 4% paraformaldehyde only (Fig. 7). Cells and tissues with high glycogen content are not suitable for this kind of approach because glycogen is washed away during the sectioning and labeling procedure, leaving behind large empty areas

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Fig. 7. Fine structure of autophagic vacuoles in a thin cryosection. Mouse hepatocytes were isolated and cultured overnight. Autophagy was induced in serum- and amino-acid–free medium. Thin cryosections were prepared as described in Subheading 3.4. and immunogold labeled with anti-cathepsin D (5 nm gold, small arrowheads) and anti-LAMP-2 (10 nm gold, large arrowheads). Autophagic vacuoles were identified by their morphology, using cytoplasmic contents as a criterion. Two autophagic vacuoles are shown. One of them contains a mitochondrion (m), the other partially condensed ER membranes (er). The arrow indicates a small vesicle, possibly about to fuse with the latter autophagic vacuole. MVB, multivesicular body/endosome. in the sections. Therefore, the animals should be fasted before tissue preparation if glycogen-containing tissues such as liver or muscle are to be cryosectioned for electron microscopy.

Acknowledgments The author would like to thank Pirkko Hirsimäki (Turku, Finland) and Hilkka Reunanen (Jyväskylä, Finland) for the initial introduction to the secrets of electron microscopy and autophagy. John Lucocq (Dundee, UK) is thanked for sharing his knowledge on the quantitative aspects of microscopy, and Paul Saftig (Kiel, Germany) is acknowledged for his generous support during the years we have used to explore the functions of lysosomal membrane proteins. Work described in this chapter was supported by The Royal Society (Dundee), Helsel Stiftung (Kiel), University of Helsinki Research Funds, and Biocentrum Helsinki.


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References 1. Eskelinen, E. L. (2005) Autophagy in mammalian cells, in Lysosomes (Saftig, P., ed.), Landes Bioscience/Eurekah.com, Georgetown, pp. 166–180 2. Klionsky, D. J., and Emr, S. D. (2000) Autophagy as a regulated pathway of cellular degradation. Science 290, 1717–1721. 3. Boya, P., Gonzalez-Polo, R. A., Casares, N., et al. (2005) Inhibition of macroautophagy triggers apoptosis. Mol. Cell Biol. 25, 1025–1040. 4. Lum, J. J., Bauer, D. E., Kong, M., et al. (2005) Growth factor regulation of autophagy and cell survival in the absence of apoptosis. Cell 120, 237–248. 5. Kuma, A., Hatano, M., Matsui, M., et al. (2004) The role of autophagy during the early neonatal starvation period. Nature 432, 1032–1036. 6. Eskelinen, E. L., Tanaka, Y., and Saftig, P. (2003) At the acidic edge: emerging functions for lysosomal membrane proteins. Trends Cell Biol. 13, 137–145. 7. Tanaka, Y., Guhde, G., Suter, A., et al. (2000) Accumulation of autophagic vacuoles and cardiomyopathy in LAMP-2 -deficient mice. Nature 406, 902–906. 8. Liang, X. H., Jackson, S., Seaman, M., et al. (1999) Induction of autophagy and inhibition of tumorigenesis by beclin 1. Nature 402, 672–676. 9. Qu, X., Yu, J., Bhagat, G., et al. (2003) Promotion of tumorigenesis by heterozygous disruption of the beclin 1 autophagy gene. J. Clin. Invest. 112, 1809–1820. 10. Melendez, A., Talloczy, Z., Seaman, M., Eskelinen, E. L., Hall, D. H., and Levine, B. (2003) Autophagy genes are essential for dauer development and lifespan extension in C. elegans. Science 301, 1387–1391. 11. Ravikumar, B., Vacher, C., Berger, Z., et al. (2004) Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat. Genet. 36, 585–595. 12. Hara, T., Nakamura, K., Matsui, M., et al. (2006) Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885–889. 13. Komatsu, M., Waguri, S., Chiba, T., et al. (2006) Loss of autophagy in the central nervous system causes neurodegeneration in mice. Nature 441, 880–884. 14. Talloczy, Z., Jiang, W., Virgin IV, H. W., et al. (2002) Regulation of starvation and virus-induced autophagy by the eIF2alpha kinase signaling pathway. Proc. Natl. Acad. Sci. USA 99, 190–195. 15. Nakagawa, I., Amano, A., Mizushima, N., et al. (2004) Autophagy defends cells against invading group A Streptococcus. Science 306, 1037–1040. 16. Arstila, A. U., and Trump, B. F. (1968) Studies on cellular autophagocytosis. The formation of autophagic vacuoles in the liver after glucagon administration. Am. J. Pathol. 53, 687–733. 17. Dunn, W. A. (1994) Autophagy and related mechanisms of lysosomal-mediated protein degradation. Trends Cell Biol. 4, 139–143. 18. Eskelinen, E. L. (2005) Maturation of autophagic vacuoles in mammalian cells. Autophagy 1, 1–10. 19. Dunn, W. A. (1990) Studies on the mechanisms of autophagy: formation of the autophagic vacuole. J. Cell Biol. 110, 1923–1933.

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20. Dunn, W. A. (1990) Studies on the mechanisms of autophagy: maturation of the autophagic vacuole. J. Cell Biol. 110, 1935–1945. 21. Klionsky, D. J., Cregg, J. M., Dunn, W. A. J., et al. (2003) A unified nomenclature for yeast autophagy-related genes. Dev. Cell 5, 539–545. 22. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728. 23. Jäger, S., Bucci, C., Tanida, I., et al. (2004) Role for Rab7 in maturation of late autophagic vacuoles. J. Cell Sci. 117, 4837–4848. 24. Griffiths, G. (1993) Fine Structure Immunocytochemistry, Springer-Verlag, Berlin Heidelberg. 25. Howard, C. V., and Reed, M. G. (1998) Unbiased Stereology. Three-Dimensional Measurement in Microscopy, Springer-Verlag, New York. 26. Liou, W., Geuze, H. J., and Slot, J. W. (1996) Improving structural integrity of cryosections for immunogold labeling. Histochem. Cell Biol. 106, 41–58.

3 Methods for Assessing Autophagy and Autophagic Cell Death Ezgi Tasdemir, Lorenzo Galluzzi, M. Chiara Maiuri, Alfredo Criollo, Ilio Vitale, Emilie Hangen, Nazanine Modjtahedi, and Guido Kroemer

Summary Autophagic (or type 2) cell death is characterized by the massive accumulation of autophagic vacuoles (autophagosomes) in the cytoplasm of cells that lack signs of apoptosis (type 1 cell death). Here we detail and critically assess a series of methods to promote and inhibit autophagy via pharmacological and genetic manipulations. We also review the techniques currently available to detect autophagy, including transmission electron microscopy, half-life assessments of long-lived proteins, detection of LC3 maturation/aggregation, fluorescence microscopy, and colocalization of mitochondrionor endoplasmic reticulum–specific markers with lysosomal proteins. Massive autophagic vacuolization may cause cellular stress and represent a frustrated attempt of adaptation. In this case, cell death occurs with (or in spite of) autophagy. When cell death occurs through autophagy, on the contrary, the inhibition of the autophagic process should prevent cellular demise. Accordingly, we describe a strategy for discriminating cell death with autophagy from cell death through autophagy.

Key Words: Apoptosis; autophagosomes; fluorescence microscopy; endoplasmic reticulum; LC3-GFP; lysosomes; mitochondria; starvation. 1. Introduction Autophagy is an evolutionarily conserved, homeostatic process that allows for the bulk degradation of long-lived proteins and organelles (1). In eukaryotic cells, the autophagic pathway is initiated when organelles and/or portions of the cytosol destined to degradation are enclosed within double-membraned vacuoles, namely autophagosomes (also known as AV, i.e., autophagic vacuoles). To promote the From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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degradation of their luminal content, autophagosomes fuse with lysosomes, thus forming the so-called autophagolysosomes (2). It is important to note that the mere presence of AV in cells is not a proof of elevated autophagy, because a reduced turnover of AV (e.g., due to a decreased fusion of AV with lysosomes) suffices to increase their number severalfold (3). Accordingly, ultrastructural studies have to be accompanied by evidence for increased protein and/or organelle turnover to ascertain increased autophagic activity. The execution and regulation of the autophagic program rely on several autophagy-specific genes (atg), which are characterized by a high degree of conservation among species as distant as humans and yeast (4). Some Atg proteins are directly implicated in the formation of the autophagosome. For instance, the ubiquitination of Atg5 and Atg12 by the E1-like enzymes Atg7 and Atg10 is required to form AV (5). Microtubule-associated protein light chain 3 (LC3) is the mammalian equivalent of yeast Atg8 and exists in two forms, LC3-I and -II. LC3-I is an 18-kDa polypeptide normally found in the cytosol, whereas the product of its proteolytic maturation (LC3-II, 16 kDa) resides in the autophagosomal membranes (6). Beclin-1 is the mammalian orthologue of yeast Atg6 and localizes to the trans-Golgi network, where it participates in autophagosome formation by interacting with the class III phosphatidylinositol 3-kinase (PI3K) human vacuolar protein sorting factor protein 34 (hVps34) (7). Importantly, the pro-autophagic interaction between hVps34 and Beclin-1 can be inhibited by the binding of the latter to the anti-apoptotic proteins Bcl-2 or Bcl-Xl (8). To ensure the turnover of old and damaged organelles, autophagy occurs constitutively at low, basal levels. However, it is also a tightly regulated adaptive mechanism that enhances cell survival under various environmental and cellular stresses, including nutrient deprivation, oxidative stress, accumulation of misfolded proteins, bacterial and viral infection, toxic stimuli, and irradiation (9,10). One of the most efficient triggers of autophagy is starvation. In response to culture in nutrient-free media, cells degrade nonessential components to meet the cell’s energetic demand as well as to provide metabolites for vital biosynthetic reactions. In this scenario, the suppression of autophagy by chemical inhibitors or by the downregulation of essential genes (e.g., Atg5, Atg6/Beclin-1, Atg10, or Atg12) can sensitize cells to starvation-induced death (3). Autophagy inhibition sensitizes cells also to the depletion of obligatory growth (or survival) factors, which result in decreased nutrient import through the plasma membrane. This cell death occurs without massive autophagic vacuolization and is usually accompanied by the hallmarks of apoptosis, including mitochondrial membrane permeabilization (MMP) and activation of caspases (3). In some instances, the inhibition of autophagy tends to shift the cell death subroutine to more necrotic phenotypes (11).

Methods for Assessing Autophagy

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A very different picture emerges when the late stages of AV maturation (e.g., fusion between AV and lysosomes) are inhibited. This may be accomplished by the administration of lysosomotropic bases (hydroxychloroquine or chloroquine) (3), by inhibiting the vacuolar proton pump responsible for the luminal acidification of lysosomes (with bafilomycin A1) (3), or by the knock-down of the gene encoding lysosome-associated membrane protein 2 (LAMP-2) (12). In such conditions, nutrient-starved cells massively accumulate AV in the cytoplasm, as determined by electron microscopy or by following the redistribution of the AV marker LC3 fused with a green fluorescent protein (GFP) moiety (LC3-GFP). In spite of AV accumulation, the autophagic protein turnover is inhibited (12,13). This example illustrates the importance of using a correct combination of methods to assess AV formation, AV turnover, and cell death. In some cases, cells manifesting massive autophagic vacuolization can be rescued from cell death by the removal of lysosomal inhibitors and/or the readdition of nutrients, despite their sometimes disastrous morphological aspect evoking textbook images of autophagic cell death. This is especially true when the still nucleus exhibits a normal aspect and mitochondria remain energized (14). However, upon prolonged nutrient starvation and inhibition of the formation of autophagolysosomes, vacuolated cells manifest the hallmarks of apoptosis (e.g., MMP, release of toxic proteins from the mitochondrial intermembrane space, and caspase activation), indicating that the point of no return that separates cellular life and death has been trespassed (15). Autophagy has been suggested to constitute an effector mechanism of cell death, meaning that self-eating would be the first step of self-killing. Some pharmacological studies based on the use of 3-methyladenine (3-MA, a lowaffinity inhibitor of hVps34) tend to support this hypothesis. For instance, it has been proposed that autophagy is required for the death of sympathetic neurons cultured in the absence of the essential nerve cell growth factor (NGF) and in the presence of caspase inhibitors (16). Similarly, TRAIL-induced autophagy accounts for the formation of hollow acini-like structures in differentiating mammary epithelial cells (17). In this model, 3-MA can provoke luminal filling when caspase-3 is inhibited by overexpression of a Bcl-Xl transgene. However, the fact that 3-MA effectively inhibits hVps34 only at concentrations ≥10 mM sheds major doubts on its specificity. Reportedly, inhibition of Atg genes by small interfering RNAs (siRNAs) can prevent autophagic cell death, at least in some models. For instance, siRNA-mediated downregulation of Atg6/Beclin-1 or Atg7 has been shown to reduce type 2 cell death of human U937 cells dying in response to caspase-8 inhibition (18). siRNAs targeting Atg6/Beclin-1 and Atg5 are able to prevent the autophagic cell death of Bax−/− Bak−/− mouse embryonic fibroblasts succumbing to the inhibition of topoisomerase 2 by

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etoposide or to inhibition of tyrosine kinases by staurosporine (19). Finally, knock-down of Atg5 (or overexpression of a dominant negative variant of Atg5) can inhibit interferon- (IFN-)–induced autophagic vacuolization and subsequent cell death (20). The definition of the features that may be employed to identify dead cells has been the subject of a long debate. Permeabilization of the plasma membrane, disintegration of the cell, and/or phagocytosis of the corpse have been recognized as the characters that delimit the irreversibility of the process (21,22). Several subroutines of cell death can be distinguished according to ultrastructural criteria. The best-studied modality of cell death, apoptosis (type 1 cell death), is characterized by morphological changes that include nuclear pyknosis (chromatin condensation) and karyorhexis (nuclear fragmentation). As the process advances, cells form small round bodies surrounded by membranes that contain intact cytoplasmic organelles and/or nuclear fragments. In vivo, these “apoptotic bodies” are usually engulfed by resident phagocytic cells. As mentioned above, massive autophagic vacuolization is observed in some instances of cell death, which has been named “autophagic cell death” (type 2 cell death) (22). It is an ongoing conundrum, however, in which case “autophagic cell death” is truly mediated through autophagy (meaning that its inhibition would prevent cell death) and in which case it simply occurs together with autophagy (meaning that inhibition of autophagy would affect only the morphology of the process, but not the fate of cells) (12,23). Necrosis (type 3 cell death) is a subroutine of cell death that does not manifest the hallmarks of apoptosis or massive autophagic vacuolization. The principal feature of necrosis is a gain in cell volume (oncosis) that finally culminates in the sudden rupture of the plasma membrane, accompanied by the unorganized dismantling of swollen organelles (11). In spite of the large amount of published data, the methods that are currently available for the detection of autophagy are affected by numerous intrinsic pitfalls. The most reliable and conventional technique to visualize autophagic vacuolization is transmission electron microscopy. Biochemical methods and techniques designed to measure the aggregation of autophagosome markers allow for the monitoring of autophagy, yet are afflicted by relevant problems. Here, we will provide an overview of routine methods for the assessment of autophagy and autophagic cell death. 2. Materials 2.1. Common Materials 2.1.1. Disposables 1. 1.5-mL microcentrifuge tubes (Eppendorf, Hamburg, Germany).100 × 20 culture dishes (Corning Inc. Life Sciences, Acton, MA).


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2. 12-mm Ø cover slips (Menzer-Gläser GmbH, Braunschweig, Germany)—sterilized by incubation for 15–30 min in 100% ethanol (Carlo Erba Reagents, Milan, Italy). 3. 15- and 50-mL conical centrifuge tubes (BD Falcon, San Jose, CA). 175-cm2 flasks for cell culture (BD Falcon). 4. 5-mL, 12 × 75 mm FACS tubes (BD Falcon). 5. 6-, 12-, 24-well plates for cell culture (Corning Inc. Life Sciences). 6. 76 × 26 mm slides for fluorescence microscopy (Carl Roth GmbH, Karlsruhe, Germany).

2.1.2. Solutions 1. Growth medium for HeLa cells: Dulbecco’s modified Eagle’s medium (DMEM) containing 4.5 g/L glucose, 4 mM l-glutamine and 110 mg/L sodium pyruvate (Gibco-Invitrogen, Carlsbad, CA) supplemented with 100 mM HEPES buffer (Gibco-Invitrogen) and 10% fetal bovine serum (FBS, from PAA Laboratories GmbH, Pasching, Austria). 2. PBS (1X): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2 HPO4 , 1.4 mM KH2 PO4 in deionized water (dH2 O), adjust pH to 7.4 with 2 N NaOH. 3. Trypsin/ethylenediaminetetraacetic acid (EDTA): 0.25% trypsin, 0.38 g/L (1mM) EDTA•4Na in Hank’s balanced salt solution (HBSS) (Gibco-Invitrogen).

2.2. Experimental Modulation of Autophagy 1. Autophagy inducers are listed in Table 1. 2. Autophagy inhibitors are listed in Table 2. 3. siRNAs used for the inhibition of the autophagic pathway in human cell lines are reported in Table 3. 4. Oligofectamine™ transfection reagent (Oligofectamine™, from GibcoInvitrogen). 5. Opti-MEM® reduced serum medium (Opti-MEM® ), with Glutamax™ and phenol red (Gibco-Invitrogen).

2.3. Measuring Autophagy 1. 3 MM® Whatmann filter paper. 2. 40% acrylamide:N-N’-methylenediacrylamide (29/1) solution (Bio-Rad, Hercules, CA). 3. 6-mm-thick, 10.5 × 11 cm foam sponges (GE Healthcare Life Sciences Sciences, Uppsala, Sweden). 4. Ammonium persulfate (APS), stock solution at 10% (w/v) in dH2 O (stored at 4°C) (see also Note 37). 5. Antibodies employed in immunoblotting techniques are listed in Table 4. 6. Blocking buffer: 0.1% Tween-20 (Sigma-Aldrich) and 5% (w/v) nonfat powdered milk (commonly found in food stores) in PBS. Storage at 4°C should not exceed one week, unless 0.02% sodium azide is added as preservative.

a

Tocris Biosciences Calbiochem Sigma-Aldrich Not available for purchase Calbiochem

Tocris Biosciences Sigma-Aldrich Sigma-Aldrich

Sigma-Aldrich Sigma-Aldrich Sigma-Aldrich

Companya ER stressing agent IMPase inhibitor Starvation inducer (NF medium) IMPase inhibitor IMPase inhibitor Class I PI3K pathway inhibitor mTOR inhibitor ER stressing agent ER stressing agent IP3 R blocker IP3 R blocker

Activity

mM mM mM mM mM

in in in in in

DMSO −20°C DMSO −20°C DMSO −20°C ethanol −20°C ethanol −20°C

1 μM 3 μM 2.5 μM 2 μM 10 μM

100 μM 10 mM 75–100 μM

10 mM in dH2 O −20°C 1 M in dH2 O4°C 10 mM in DMSO −20°C

1 1 1 2 2

20 μM 50 μM 100%

Final concentration

10 mM in ethanol −20°C 5 mM in ethanol −20°C 4°C

Stock solution storage

Calbiochem, San Diego, CA; Sigma-Aldrich, St. Louis, MO; Tocris Biosciences, Ellisville, MO.

Rapamycin Thapsigargin Tunicamycin Xestospongin B Xestospongin C

Brefeldin A Carbamazepine Earle’s Balanced Salt Solution L-690,330 Lithium Chloride N-Acetyl-d-sphingosine (C2-ceramide)

Reagent

Table 1 Autophagy Inducers

34 Tasdemir et al.


35

Table 2 Autophagy inhibitors Reagent

Companya

Activity

Stock solution Storage

3-Methyladenine Sigma-Aldrich (3-MA) Bafilomycin A1 Sigma-Aldrich

10 mM

Hydroxychloroquine

30 μg/mL

a

hVps34 inhibitor Powder RT H+ -ATPase 0.2 mM in inhibitor DMSO −20°C Sanofi-Aventis Lysosomal 30 mg/mL in lumen alkalizer dH2 O −20°C

Final concentration

0.1 μM

Sanofi- Aventis, Bridgewater, NJ; Sigma-Aldrich, St. Louis, MO.

7. Bovine serum albumin (BSA, from Sigma-Aldrich), stock solution 10 mg/mL in dH2 O, stored at 4°C). 8. Chemiluminescent substrate: SuperSignal West Femto Maximum Sensitivity Substrate (Pierce Biotechnology, Rockford, IL). 9. Chemiluminescent substrate: SuperSignal West Pico Chemiluminescent Substrate (Pierce Biotechnology). 10. DC protein assay (Bio-Rad). 11. Dehybridizing buffer: 570 μL of 1 N acetic acid solution (Sigma-Aldrich) + 100 mL dH2 O. 12. Electrophoresis apparatus: Mini-PROTEAN 3 electrophoresis cell system (BioRad), powered by a PowerPac HC power supply (Bio-Rad). 13. Epon® epoxy resin (Miller-Stephenson, Danbury, CT). 14. Film processor: Curix 60 table-top film processor (Agfa, Mortsel, Belgium). 15. Fixative solution: 4% paraformaldehyde (PFA, from Sigma-Aldrich) + 0.19% picric acid (Sigma-Aldrich) in PBS. For 10 mL: dissolve 0.4 g of PFA in 500 μL of dH2 O, add one drop of 2 N NaOH, heat to 65–70°C until solution clears, add 9.5 mL of PBS, let cool to room temperature (RT), add 38 μL of saturated picric acid (Sigma-Aldrich), and move the fixative solution to ice bath (see also Notes 13–15). 16. Fluorescence microscope: IRE2 microscope equipped with a DC300F camera (Leica Microsystems GmbH, Wetzlar, Germany). 17. Fluorescent dyes used for the detection of autophagy are listed in Table 5. 18. Fluoromount-G™ mounting medium (Southern Biotech, Birmingham, AL). 19. Grade I glutaraldehyde solution, 25% in dH2 O, specially purified for use as an electron microscopy fixative (Sigma-Aldrich). 20. Isopropyl alcohol (isopropanol, CAS No. 67-63-0, from Carlo Erba Reagents). 21. l-[U-14 C]-Valine, aqueous solution containing 2% ethanol, sterilized (GE Healthcare Life).

Symbol

atg5

atg10

atg12

atg6

emd

Target gene

Autophagy related gene 5 homolog



Beclin-1

Emerin

NC_000023.9

AF077301

NM_004707

NM_8031482

BC002699

Accession number sense 5’-GCAGAACCAUACUAUUUGCdTdT-3’ antisense 5’-GCAAAUAGUAUGGUUCUGCdTdT-3’ sense 5’-GCAACUCUGGAUGGGAUUGdTdT-3’ antisense 5’-CAAUCCCAUCCAGAGUUGCdTdT-3’ sense 5’-GGAGUUCAUGAGUGCUAUAdTdT-3’ antisense 5’-GUAGCCAUCAGAACAGUCCdTdT-3’ sense 5’-GGACUGUUCUGAUGGCUACdTdT-3’ antisense 5’-GUAGCCAUCAGAACAGUCCdTdT-3’ sense 5’-CAGAGGAACCUGCUGGCGAdTdT-3’ antisense 5’-UCGCCAGCAGGUUCCUCUGdTdT-3’ sense 5’-GAAGUUGGAACUCUCUAUGdTdT-3’ antisense 5’-CAUAGAGAGUUCCAACUUCdTdT-3’ sense 5’-CUCAGGAGAGGAGCCAUUUdTdT-3’ antisense 5’-AAAUGGCUCCUCUCCUGAGdTdT-3’ sense 5’-GAUUGAAGACACAGGAGGCdTdT-3’ antisense 5’-GCCUCCUGUGUCUUCAAUCdTdT-3’ sense 5’-CCGUGCUCCUGGGGCUGGGdTdT-3’ antisense 5’-CCCAGCCCCAGGAGCACGGdTdT-3’

Sequence

Table 3 siRNAs Used to Knock Down Genes Involved in utophagy in Human Cell Lines

(81)

(3)

(3)

(3)

(3)

Ref.

36 Tasdemir et al.

lamp-1

lamp-2

Lysosome-associated membrane protein 1

Lysosome-associated membrane protein 2

NM_002294

NM_005561

NM_002647

sense 5’-GGCUGAAACUACCAGUAAAdTdT-3’ antisense 5’-UUUACUGGUAGUUUCAGCCdTdT-3’ sense 5’-GGAGGCAAAUAUCCAGUUAdTdT-3’ antisense 5’-UAACUGGAUAUUUGCCUCCdTdT-3’ sense 5’-CGAGAAAUGCAACACGUUAdTdT-3’ antisense 5’-UAACGUGUUGCAUUUCUCGdTdT-3’ sense 5’-GGAAUCCAGUUGAAUACAAdTdT-3’ antisense 5’-UUGUAUUCAACUGGAUUCCdTdT-3’ sense 5’-GCUGUGCGGUCUUAUGCAUdTdT-3’ antisense 5’-AUGCAUAAGACCGCACAGCdTdT-3’ sense 5’-GCGGUCUUAUGCAUUGGAAdTdT-3’ antisense 5’-UUCCAAUGCAUAAGACCGCdTdT-3’

(12)

(12)

(82)

Notes: siRNAs are provided either as 100 mM stock solutions (to be stored as such at −20°C), or in dehydrated form. In the latter case, stock solutions should be prepared by adding an appropriate volume of dH2 O or of the specific basic buffer (usually delivered with the siRNA) to obtain the final siRNA concentration of 20–100 mM, and by incubating the tube first at 90°C for 1 min then at 37°C for 1 h. These steps are required to resolve secondary structures that may have formed during lyophilization and that may interfere with the siRNA silencing efficacy.

hVps34

Human vacuolar protein sorting factor protein 34

Methods for Assessing Autophagy 37

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Tasdemir et al.

Table 4 Antibodies for Immunoblotting Primary antibody Anti-GAPDH (6C5) Anti-LC3

Specificity

Source organism

Monoclonal IgG1

Mouse

Polyclonal IgG

Rabbit

Secondary antibody

Label

Source organism

Anti-mouse

Horseradish peroxidase

Goat

Anti-rabbit

Horseradish peroxidase

Goat

Companya Chemicon International Santa Cruz Biotechnology Companya Southern Biotech Southern Biotech

Ref. mAB374 sc-28266

Ref. 1010-05 4010-05

a

Chemicon International, Temecula, CA; Santa Cruz Biotechnology, Santa Cruz, CA; Southern Biotech, Birmingham, AL.

22. Lead citrate: dissolve 0.1 g of lead(II) citrate tribasic trihydrate (Sigma-Aldrich) in 100 mL dH2 O; add dropwise 10 N NaOH (Sigma-Aldrich) until solution becomes clear; store at RT. 23. Lipofectamine™ 2000 transfection reagent (Lipofectamine™, from GibcoInvitrogen). 24. Loading buffer (5X): 313 mM Tris-HCl (pH 6.8), 10 mM EDTA, 500 mM dl-dithiothreitol (DTT), 10% (w/v) sodium dodecylsulfate (SDS), 50% (v/v) glycerol, 0.05% (w/v) bromophenol blue in dH2 O (or commercially available from Fermentas, Ontario, Canada). 25. l-Valine (Sigma-Aldrich), stock solution in dH2 O, at the concentration of 100 mM (stored at 4°C). 26. Lysis buffer: CelLytic™ M (Sigma-Aldrich). 27. Migration buffer (1X): 100 mL 10X Tris/Glycine/SDS buffer (Bio-Rad) + 900 mL dH2 O. 28. Mini-PROTEAN 3 combs (5-, 9- 10- or 15- wells, from Bio-Rad) (see also Note 46). 29. N,N,N’,N’-Tetra-methyl-ethylenediamine (TEMED, from Bio-Rad). 30. Nitrocellulose membrane roll (Bio-Rad). 31. Osmium tetroxide solution for electron microscopy, 2% in dH2 O. 32. Photographic films: 18 × 24 Amersham Hyperfilm™ ECL™ (GE Healthcare Life Sciences).

520 nm

500 nm

535 nm

PI

617 nm

599 nm

579 nm

Red

Green/Red

Red

Green

498 nme

335 nm

5

Red

590 nm

577 nm

Blue

461 nm

Blued

Red

Green

Green

Color

352 nm

MitoTracker® Red (CMXRos) NAO

Hoechst 33342 (bound to DNA) LysoTracker® Red MDC (in AV)

420 nmd

4

518 nm

603 nm

300 nm

501 nm

484 nm

EthBr (bound to DNA) HE

517 nm

492 nm

CellTracker™ Green (CMFDA) DiOC3 (6)

Emission peak

Absorption peak

Fluorescent probes

FACS (FL1/2 channel) FACS (FL3 channel)

IF

IF

IF

not applicabled IF

FACS (FL1 channel) FACS (FL3 channel)

IF

Application

Sigma-Aldrich

Molecular Probes

Molecular Probes

Sigma-Aldrich

Molecular Probes

Molecular Probes

Molecular Probes

Not applicablec

Molecular Probes

Molecular Probes

Companya

1 mg/mL in dH2 O

1 mM in DMSO −20°C 5 mM in 1:1 DMSO/ethanol Freshly made 1 mM in DMSO −20°C 500 μM in ethanol −20°C

5 mM in DMSO –20°C 10 mg/mL in dH2 O 4°C

40 μM in ethanol −20°C Not applicablec

10 mM in DMSO –20°C

Stock solution Storageb

Table 5 Fluorescent Probes, Labels, and Proteins for the Detection of Autophagy and Autophagic Cell Death

1 μg/mL

100 nM

150 nM

50 μM

50–75 nM

2 μM

25 μM

Not applicablec

40 nM

1 μM

Final concentration

Methods for Assessing Autophagy 39

507 nm

583 nm Green

Red

Color

Green

Red

Green

Color

Red

IF

IF

Application

FACS (FL1 channel)

IF

IF

Application

FACS (FL2 channel)

Clontech Laboratories Clontech Laboratories

Companya

Miltenyi Biotec

Molecular Probes

Molecular Probes

Companya

Molecular Probes

150 nM

LC3

mito

Coupling

Secondary antibodies (goat anti-mouse) Secondary antibodies (goat anti-mouse and anti-rabbit) Annexin V

Coupling

15 mM in ethanol –20°C

a Clontech Laboratories, Palo Alto, CA; Miltenyi Biotec, Bergisch Gladbach, Germany; Molecular Probes-Invitrogen, Carlsbad, CA; SigmaAldrich, St. Louis, MO. b To avoid photobleaching, all stock solutions should be stored under protection from light. c EthBr results from the oxidation of HE. d These properties are not exploited for the HE-mediated detection of ROS (see also Note 82). e MDC in solution emits weakly at 425 nm, but its peak is shifted to 498 nm in lipid-rich microenvironments (see also Note 23). Abbreviations: CMFDA, 5-chloromethylfluorescein diacetate; CMXRos, chloromethyl-x-rosamine; dH2 O, deionized water; DiOC6 (3), 3,3 dihexyloxacarbocyanine iodide; DMSO, dimethylsulfoxide; DsRed2, Discosoma sp. red fluorescent protein; EthBr, ethidium bromide; FACS, fluorescence-activated cell sorter; FITC, fluorescein isothiocyanate; GFP, green fluorescent protein; HE, hydroethidine; IF, immunofluorescence; LC3, microtubule-associated protein light chain 3; MDC, monodansylcadaverine (5-dimethylaminonaphthalene-1-(N-(5aminopentyl))sulfonamide); mito, mitochondrial targeting sequence from subunit VIII of human cytochrome c oxidase; NAO, nonyl acridine orange; PI, propidium iodide; TMRM, tetramethylrhodamine methyl ester.

488 nm

GFP

Absorption peak

Fluorescent proteins

558 nm

519 nm

494 nm

DsRed2

603 nm

578 nm

Emission peak

519 nm

495 nm

Alexa Fluor® 488 Alexa Fluor® 568 FITC

Emission peak

573 nm

Absorption peak

543 nm

Fluorescent labels

TMRM

40 Tasdemir et al.


41

33. Ponceau S staining solution: 0.1% (w/v) Ponceau S and 5.0% (w/v) acetic acid (Sigma-Aldrich) in dH2 O (or commercially available from Sigma-Aldrich). 34. Protein molecular weight (MW) markers: Precision Plus Protein™ Standard, Dual Color (Bio-Rad). 35. Rinsing buffer: 0.1% Tween-20 in PBS. Storage at RT. 36. Running gel buffer: 1.5 M Tris buffer (pH 8.8, from Bio-Rad). 37. Sörensen phosphate buffer (1X, 0.1 M, ph 7.3): 23 mL of 0.2 M NaH2 PO4 + 77 mL of 0.2 M Na2 HPO4 + 100 mL dH2 O. 38. Stacking gel buffer: 0.5 M Tris buffer (pH 6.8). 39. Transfer buffer (1X): 100 mL 10X Tris/glycine buffer (Bio-Rad) + 200 mL ethanol + 900 mL dH2 O. 40. Transfer cassette: Mini Trans-Blot Cell (Bio-Rad). 41. Transmission electron microscope: Tecnai G2 Spirit (FEI, Eindhoven, The Netherlands). 42. Transparent plastic film (Saran, as used for food preservation). 43. Trichloroacetic acid (TCA), stock solution 10% (w/v) in dH2 O (stored at RT). 44. Uranyl acetate: dissolve 10 g of uranyl acetate dihydrate in 100 mL of methanol (Carlo Erba Reagents); shake or vortex until complete dissolution; store at −20°C.

2.4. Measurement of the Colocalization of Cytoplasmic Organelles with Autophagic Vacuoles 1. 0.1% (w/v) SDS in PBS. 2. 10% FBS (PAA Laboratories GmbH) in PBS. 3. Antibodies employed for immunofluorescence microscopy assessments are listed in Table 6. 4. BSA buffer: 3 mg/mL BSA (Sigma-Aldrich) in PBS. 5. Confocal microscope: Zeiss LSM 510 (Carl Zeiss AG, Oberkochen, Germany). 6. Fluorochromes used to measure the colocalization of cytoplasmic organelles with autophagic vacuoles are listed in Table 5.

2.5. Measuring Cell Death–Related Parameters 1. Annexin V-FITC kit for the detection of phosphatidylserine externalization (Miltenyi Biotec, Bergisch Gladbach, Germany). 2. Carbonyl cyanide m-chlorophenylhydrazone (CCCP, from Sigma-Aldrich), stock solution in ethanol, 10 mM, stored at −20°C. 3. Cytofluorometer: FACScan (Becton Dickinson, San Jose, CA) equipped with an argon ion laser emitting at 488 nm. 4. Fluorescent probes used to measure cell death–related parameters are listed in Table 5.

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Tasdemir et al.

Table 6 Antibodies for Immunofluorescence Microscopy Primary antibody

Specificity

Source organism

Companya

Ref.

Anti-AIF internal domain Anti-Calreticulin

Polyclonal IgG

Rabbit

Polyclonal IgG

Rabbit

Anti-Caspase-3a (Asp175) Anti-cytochrome c (clone 6H2.B4) Anti-Hsp60 (clone LK1) Anti-LAMP2

Polyclonal IgG

Rabbit

Monoclonal IgG1

Mouse

Chemicon International Stressgen Bioreagents Cell Signalling Technology BD Pharmingen

Monoclonal IgG1

Mouse

Sigma-Aldrich

H4149

Monoclonal IgG1

Mouse

Santa Cruz Biotechnology

sc-18822

Secondary antibody

Label

Anti-mouse Anti-mouse Anti-rabbit

Alexa Fluor® 488 Alexa Fluor® 568 Alexa Fluor® 568

Source organism Goat Goat Goat

AB 16501 SPA-600D 9661 556432

Company

Ref.

Molecular Probes Molecular Probes Molecular Probes

A-11001 A-11031 A-11036

a BD Pharmingen, San Diego, CA; Cell Signalling Technology, Danvers, MA; Chemicon International, Temecula, CA; Molecular Probes-Invitrogen, Carlsbad, CA; Santa Cruz Biotechnology, Santa Cruz, CA; Sigma-Aldrich, St. Louis, MO; Stressgen Bioreagents, Ann Arbor, MI.

3. Methods Cells that activate the autophagic program, manifest in the accumulation of double-membraned autophagic vesicles in the cytoplasm. These vacuoles can be classified according to their electron density, as assessed by transmission electron microscopy, into early and late AV. Early AV (AV1) contain cytoplasmic portions and/or organelles and exhibit an electron density equivalent to the cytoplasm (AV1). Late or degradative AV (AV2) show evidence of partially digested luminal content and are characterized by increased electron density. The mere presence of AV in the cytoplasm does not necessarily indicate an increased level of autophagy, since a reduction of the fusion between autophagosomes and lysosomes suffices to promote AV accumulation. Thus, quantitative


43

methods for the detection of cytoplasmic protein turnover should be employed in addition to AV monitoring, in order to verify augmented levels of autophagy. 3.1. Experimental Modulation of Autophagy Given its importance in cellular homeostasis and response to stress, it is not surprising that autophagy is a precisely controlled process that is regulated by several (cross-talking) signaling pathways (24,25). In this context, a major role is played by the mammalian target of rapamycin (mTOR) protein, a serine/threonine kinase implicated in the control of several cellular functions (26,27). mTOR integrates numerous inputs (including signals from growth factors, insulin, cellular stressors as well as those reporting the availability of nutrients) to stimulate protein synthesis via the phosphorylation of key translation regulators like ribosomal protein S6 kinase (p70SK6) and eukaryotic initiation factor 4E binding protein 1 (eIF4EBP1) (28). Accordingly, mTOR is a major gatekeeper and inhibitor of autophagy, and may exert its antiautophagic effects either by promoting protein synthesis or by a direct inhibition of crucial atg proteins. Other important regulators of autophagy include class I and class III PI3Ks. The increase in class I PI3K products (like phosphatidylinositol 3,4-bisphospate, i.e., PIP2, and phosphatidylinositol 3,4,5-trisphospate, i.e., PIP3), caused by feeding the cells with synthetic lipids or via by stimulating the interleukin-13 receptor, has been reported to inhibit macroautophagy (29). Conversely, class III PI3K (hVps34) activates autophagy by interacting with Beclin-1 and plays a crucial role at early stages of autophagosome formation (30). 3.1.1. Induction of Autophagy In vitro, several strategies may be employed to induce autophagy (Fig. 1). These include: the inhibition of mTOR (e.g., by starvation, by inhibition of the upstream class I PI3K-mediated signaling, or by administration of the mTOR specific inhibitor rapamycin) (27,31); the inhibition of inositol monophosphatase (IMPase), resulting in a reduction of free inositol and inositol-1,4,5triphosphate (IP3) levels (e.g. with lithium, L-690,330 or carbamazepine) (32); the blockade of the IP3 receptor (IP3R) of the endoplasmic reticulum (ER) (e.g., with xestospongin B and C) (33); the induction of ER stress via the activation of the unfolded protein response (e.g., with tunicamycin, thapsigargin or brefeldin A) (34); or the induction of oxidative stress (e.g. with H2 O2 ) (35). 1. Wild-type HeLa cells are cultured in appropriate growth medium and passaged when approaching to confluence with trypsin/EDTA to provide new maintenance cultures in 175-cm2 flasks and experimental cultures. The latter are performed

44

Tasdemir et al. signal

Inducers

Endogenous modulators

Plasma membrane

signal

Phagophore

Cytoplasmic organelles

Atg5 Atg6/Beclin-1 Atg10 Atg12 LAMP-2 hVps34

Inhibitors

Endogenous modulators Bcl-2 Class I PI3K mTOR

Chemical inhibitors Chemical inducers Brefeldin-A Carbamazepine Ceramide L-690,330 Lithium chloride Rapamycin Thapsigargin Tunicamycin Xestospongin B Xestospongin C

Lysosome

3-methyladenine Bafilomycin A1 Hydroxychloroquine

AV1

fusion

Adaptive response to

AV2

siRNAs Atg5 Atg6/Beclin-1 Atg10 Atg12 LAMP-2 hVps34

Bacterial infection Irradiation Oxidative stress Protein aggregates Starvation Viral infection

degradation

Fig. 1. Overview of the autophagic pathway. The formation of autophagic vacuoles begins with the segregation of portions of cytoplasm and/or cytoplasmic organelles by lipid-rich apparatuses also known as phagophores. This generates immature autophagosomes (AV1), which are doubled-membraned vacuoles in which the luminal content has


45

either in 12-well plates (100 × 103 cells in 1 mL of growth medium) or in 24-well plates in which sterile cover slides have been previously deposited (25 × 103 cells in 0.5 mL of growth medium). 2. Twelve to 24 h after plating, induction of autophagy is performed. To this aim, growth medium is removed by aspiration and cells are washed twice with 1 or 2 mL (in 24- or 12-well plates, respectively) sterile PBS. Then, autophagy inducers (see Table 1) are administered in growth medium at the appropriate concentration (see Note 1). Alternatively, growth medium is substituted with 0.5–1 mL of Earle’s balanced salt solution (EBSS, nutrient-free [NF] medium). Finally, cells are incubated at 37ºC in 5% CO2 atmosphere for a variable time frame, depending on the experimental setting (see Note 2). 3. Finally, autophagic activity and cell death can be monitored as described below (see Subheading 3.2.).

3.1.2. Inhibition of Autophagy The cascade of events leading to autophagy can be suppressed at various steps (36) (Fig. 1), which include: (1) the formation of autophagosomes; (2) the fusion between autophagosomes and lysosomes (3,13); and (3) the final, lysosomal degradation phase (15,37). The inhibition of the autophagic process can be achieved by different means, notably (1) chemical inhibitors (see Table 2) (3,13,15); (2) transfection-enforced overexpression of inhibitory proteins (8); and (3) siRNAs targeting essential atg gene products (see Table 3) (3,12,30). 3-Methyladenine (3-MA) prevents autophagy at the sequestration step by inhibiting the class III PI3K hVps34 (29). However, the results of experiments in which 3-MA is employed as an autophagy inhibitor should be evaluated cautiously due to the fact that this compound affects many other signaling pathways (by interacting with kinases such as class I PI3K; c-Jun N-terminal Fig. 1. (Continued) not yet been degraded. It is only after the fusion with lysosomes and the acidification of the luminal content that autophagosomes progress to mature, degradation-competent organelles (AV2). Autophagy represents an adaptive response to different stressful conditions (e.g., the depletion of nutrients, bacterial and viral infections, accumulation of misfolded proteins) and can be triggered by pharmacological inducers such as rapamycin, tunicamycin, or xestospongin B. The inhibition of autophagy (which is regulated by several endogenous signalling pathways) can be achieved by molecules like bafilomycin A1 or hydroxychloroquine as well as by genetic manipulations. In the latter case, the siRNA-mediated depletion of proteins essential for autophagy provides a specific tool to suppress the autophagic pathway. Atg, autophagy-specific gene; hVps34, human vacuole protein sorting factor protein 34; LAMP-2, lysosome-associated membrane protein-2; PI3K, phosphatidylinositol-3 kinase; siRNA, small-interfering RNA.

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Tasdemir et al.

kinase, i.e., JNK; and mitogen-activated protein kinases, i.e., MAPK) (29,38). Moreover, at the concentration of 10 mM (representing the minimal concentration required to block autophagy), 3-MA also inhibits MMP and other catabolic pathways (39). Hydroxychloroquine is a lysosomotropic amine, which is clinically used for the treatment of malaria, rheumatoid arthritis, and systemic lupus erythematosus. As do other immunosuppressant drugs, hydroxychloroquine exhibits cytotoxic properties. This agent induces the alkalinization of the lysosomal lumen, thereby inhibiting the fusion between lysosomes with autophagosomes and interfering with the action of lysosomal hydrolases, which function at an acidic pH. At higher concentrations, hydroxychloroquine induces lysosomal membrane permeabilization (LMP), resulting in the potentially lethal leakage of catabolic hydrolases into the cytosol (15). Bafilomycin A1 is a specific inhibitor of the vacuolar H+ -ATPase, the proton pump that acidifies the lysosomal lumen (37,40). Similarly to hydroxychloroquine, bafilomycin A1 prevents the activation of lysosomal hydrolases and interferes with the fusion between autophagosomes and lysosomes (13), presumably because lysosomal acidification is required for this step. The intrinsic disadvantage of chemical inhibitors is that they can exert several unwanted side effects due to the fact that none of the currently available molecules is truly specific for autophagy. The siRNA-mediated depletion of proteins essential for the autophagic pathway is characterized by a high degree of specificity. siRNAs targeting hVps34, Beclin-1, Atg5, Atg10, Atg12, and LAMP-2 are now widely employed to suppress autophagy (see Table 3) (3,12,30). 3.1.2.1. Chemical Inhibition of Autophagy 1. HeLa cells are maintained in culture and plated as described above (see Subheading 3.1.1.). 2. Chemical inhibition of autophagy is performed 12–24 h after plating. As for the induction of autophagy, growth medium is removed, cells are washed twice with sterile PBS, then autophagy inhibitors (see Table 2) are administered at the appropriate concentration (see Note 1), either in growth medium (negative control condition) or in NF medium (providing a positive control for autophagy induction). Thereafter, cells are cultured at 37ºC in a 5% CO2 incubator. 3. After a time frame varying according to the specific experimental aim (see Note 2), autophagic activity can be measured as described below (see Subheading 3.2.).

3.1.2.2. siRNA-Mediated Inhibition of Autophagy 1. HeLa cells are seeded in 6-well plates (approximately, 200 103 cells in 3 mL of growth medium), in order to obtain confluence levels around 60–70% on the next day.


47

2. If cells have reached appropriate confluence, transfection of siRNA is performed 12–24 h after plating (see Note 3). For the delivery of siRNAs to adherent cells, liposome-based transfection is employed. To this aim, 200 nmol of siRNA (final concentration in wells = 100 nM) are diluted in 180 μL of Opti-MEM® and 4 μL of Oligofectamine™ transfection reagent (Oligofectamine™) are gently mixed with 16 μL of Opti-MEM® . After a first incubation of 5–10 min, the diluted siRNAs and the diluted Oligofectamine™ solution are combined, gently mixed, and incubated for additional 20 min to allow for the formation of Oligofectamine™:siRNA transfection complexes (see Notes 4 and 5). 3. Before the addition of the complexes to the cells, growth medium is replaced by 1.8 mL of fresh medium without serum (see Notes 6 and 7). Then, 200 μL of the solution containing Oligofectamine™:siRNA complexes are added to each well, and plates are incubated at 37ºC in 5% CO2 atmosphere for 4 h. 4. Four hours after transfection, 220 μL of FBS should be added to each well to restore the final concentration of 10% (as in complete growth medium). 5. Twelve to 24 h after transfection, cells can be trypsinized (500 μL of Trypsin/EDTA per well) and seeded in 12- or 24-well plates, according to the specific experimental settings (see Notes 8–10). 6. After the time required for the siRNA-mediated downregulation of the target protein (see Note 8), the desired stimuli can be administered to the cells in which the autophagic pathway has been suppressed by genetic techniques. 7. Finally, following another incubation at 37ºC in 5% CO2 atmosphere (the duration depends again on the specific experimental conditions), autophagy can be assessed as described below (see Subheading 3.2.).

3.2. Measuring Autophagy At present, multiple techniques are available for the quantification of autophagy and the identification of AV. Table 7 summarizes these methods, by providing also the main advantages and drawbacks of each. 3.2.1. Electron Microscopy 1. 2 × 106 cells are cultured in 100 × 20 mm culture dishes. 2. After the treatment with the desired stimuli, cells are fixed for 1 h at 4°C in 1.6% glutaraldehyde in 0.1 M Sörensen phosphate buffer (pH 7.3) and washed once with PBS. 3. Cells are then re-fixed in aqueous 2% osmium tetroxide and finally embedded in Epon® epoxy resin, until imaging. 4. Examination is performed at 80 kV under a transmission electron microscope, on ultrathin sections (80 nm) stained with 0.1% lead citrate and 10% uranyl acetate.

Once the images are obtained, the number of type I and type II AV can be quantified on a per-cell basis (see Fig. 2). In addition, AV volume can be evaluated by morphometric methods.

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Tasdemir et al.

Table 7 Methods for the Detection of Autophagy Method Electron microscopy

LC3-GFP aggregation

Advantages Direct observation of the AV Quantitative (n° of AV per cell, ratio between their surface/volume and that of the cell) Differentiates between AV1 and AV2 Immunoelectron microscopy approaches enable the specific labeling of AV Rather specific Allows for the quantification of AV per cell Can be performed on histological sections Can be coupled to other stainings for co-localization studies

Bulk degradation of long-lived proteins

Specifically measures turnover of long-lived proteins

MDC staining

Stains AV due to ion trapping in acidic compartments and to increased relative fluorescence in hydrophobic milieus Can be used in vivo without construction of transgenic animals

TCA-precipitated radioactivity can be used to estimate the intracellular amount of radiolabeled proteins

Drawbacks Skill requiring, laborious Prone to subjective interpretations (e.g., swollen organelles may be mistaken for AV) Scarcely representative of the overall sample (few cells examined)

Weaker signal from AV2 than from AV1 Possibility of false-negative and false-positive results

Low amounts of aggregates are prone to misinterpretations, due to basal levels of ongoing autophagy Not specific for autophagy nor for lysosomal proteolysis Excess unlabeled amino acids administered during the chase step may inhibit autophagy Not applicable if fusion of AV and lysosomes is interrupted Rather unspecific: stains only acidic vacuoles of the autophagic pathway (AV2), but several other acidic compartments of cells Cannot be used alone for the detection/quantification of autophagy

Methods for Assessing Autophagy CellTracker™ Green staining

LC3-I to LC3-II conversion

Useful in double staining techniques to distinguish AV from other vacuolar compartments

Specific (autophagy never observed in the absence of LC3-I to LC3-II conversion and LC3-II incorporation in AV) LC3-II/LC3-I ratio correlates with the extent of autophagic activity

49 Totally unspecific (it detects all vacuolar compartments)

Cannot be used alone to assess autophagy Provides an estimate of the instantaneous levels of autophagy rather than of the overall autophagic flux (due to relatively short lifetime of AV) Antibodies against LC3 display higher affinity for LC3-II, resulting in overestimation of the corresponding band Possibility of false-positive results

Abbreviations: AV, autophagic vacuoles; GFP, green fluorescent protein; LC3, microtubuleassociated protein light chain 3; MDC, monodansylcadaverine.

3.2.2. LC3-GFP Aggregation 1. HeLa cells (25 × 103 in 0.5 mL growth medium) are seeded in 24-well plates in which sterile cover slips have been previously deposited. 2. Twelve to 24 h later, cells are transfected with a plasmid coding for the autophagosome marker LC3 fused with green fluorescence protein (GFP) (6), according to the following protocol. 4 μg of plasmid are diluted in 200 μL of Opti-MEM® , at the same time as 5 μL of Lipofectamine™ are gently mixed with 200 μL Opti-MEM® . After a first incubation of 5–10 min, the diluted plasmid solution and diluted Lipofectamine™ solution are gently mixed and incubated for another 20 min to promote the formation of Lipofectamine™:plasmid complexes (see Notes 4 and 5). 3. Thereafter, 30 μL of solution containing the Lipofectamine™:plasmid complexes are added to each well, in which medium had been previously replaced with 500 μL of serum-free growth medium. Plates are then incubated at 37ºC in 5% CO2 atmosphere for 4 h before 60 μL of FBS are added to restore the final FBS concentration of 10% (as in complete growth medium) (see Note 6). 4. Cells are cultured for 24 h, or until they start to express the LC3-GFP fusion protein, prior to treatment with the desired stimuli (see Notes 11 and 12). 5. At the end of stimulation, growth medium is removed, cells are washed twice with PBS and fixed in 400 μL of fixative solution for 30 min at RT (see Notes 13–16).

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1 µm

1 µm 1 µm

Fig. 2. Ultrastructural features of autophagy. HeLa cells were treated with 2.5 μM tunicamycin for 6 h and processed for electron microscopy, as described in Subheading 3.2.1. The induction of autophagy is witnessed by massive vacuolization of the cytoplasm. In the same cells, immature autophagic vacuoles characterized by an electron density equivalent to the cytoplasm coexist with late vesicles, in which catabolic processes have been already started (characterized by an increased electron density). Black bars report picture scale. 6. Fixative solution is removed, cells are washed three times with PBS, and nuclear counterstaining is performed by the addition of 2 μM Hoechst 33342 in PBS (200 μL per well) for 10 min (see Note 17). 7. The cover slips are mounted onto slides using Fluoromount-G™ mounting medium prior to examination in a fluorescence microscope. For the visualization of LC3GFP, cells should be examined using an oil immersion objective (50–65× magnification). Suitable excitation and emission filters should be selected according to the following absorption and emission peaks: 488/507 nm for LC3-GFP and 352/461 nm for Hoechst 33342 (see Note 18). The LC3-GFP fusion protein redistributes from a diffuse (cytoplasmic and nuclear) to a vacuolar, punctuate (exclusively cytoplasmic) pattern when AV are formed (see Fig. 3 and Note 19)

3.2.3. Bulk Degradation of Long-Lived Proteins 1. HeLa cells (400 × 103 in 2.5 mL growth medium) are plated in 6-well plates.


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2. Forty-eight hours later, growth medium is replaced by 0.2 μCi/mL l-[U-14 C]valine (see Note 20) in complete medium. Plates are then incubated at 37°C in 5% CO2 atmosphere for 24 h. 3. At the end of the radiolabeling period, cells are washed three times with PBS (to remove unincorporated radioisotopes), and incubated with 10 mM unlabeled l-valine in complete growth medium (2–2.5 mL) for 1 h (prechase period) (see Note 21). 4. After the prechase step, the culture medium is again replaced by fresh medium containing 10 mM unlabeled valine in the presence or absence of an autophagy inhibitor (e.g., 3-MA) (see Note 22). Cells are then incubated with the desired stimuli for 4–12 h (chase period). 5. After the chase step, supernatant is collected and proteins from the medium and from cells are precipitated separately with 10% (w/v) TCA (overnight, 4°C). 6. Thereafter, TCA-precipitated fractions are centrifuged (600g, 20 min, RT) and redissolved in 1 mL of 0.2 N NaOH. Finally, radioactivity is determined by liquid scintillation counting. 7. The rate of degradation of long-lived proteins can be calculated by determining the ratio of TCA-precipitated radioactivity recovered from the medium to the sum of TCA-precipitated radioactivity from the medium and cells: %Proteolysish−1 = Rm /Rc + Rm Hoechst 33342

LC3-GFP

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Fig. 3. Detection of autophagy by the LC3-GFP aggregation technique. HeLa cells transfected with a plasmid encoding the LC3-GFP fusion protein (as described in Subheading 3.2.2.) were left untreated (Control) or treated with 2.5 μM tunicamycin for 6 h (Autophagy). Thereafter, cells were processed for fluorescence microscopy examination, as detailed in Subheading 3.2.2. In untreated cells LC3-GFP exhibits a diffuse cytoplasmic signal. When autophagy is induced, LC3-GFP chimeric proteins aggregate in autophagic vacuoles, leading to a punctuate cytoplasmic staining. As confirmed by Hoechst 33342 counterstaining, LC3-GFP is not found in the nucleus.

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where Rm is TCA-precipitated radioactivity from medium and Rc is TCAprecipitated radioactivity from cells. 3.2.4. Monodansylcadaverine (MDC) Staining of Autophagic Vacuoles 1. HeLa cells (25 × 103 in 0.5 mL of growth medium) are seeded in 24-well plates where sterile cover slips have been previously dropped. 2. Following the desired stimuli for the induction or inhibition of autophagy, AV are labeled with 50 μM MDC in growth medium (0.5 mL) for 30 min at 37°C. 3. Then, cells are washed three times with PBS and fixed in 400 μL of 4% PFA (see Notes 13 and 14) for 30 min at RT. After fixation, PFA is removed and cells are washed once with PBS. Finally, cover slips are mounted as described above (see Subheading 3.2.2., step 7). 4. As for LC3-GFP staining, the assessment of AV is performed by fluorescence microscopy, at magnifications of 50–65×. Usually, MDC is observed with a 335(380)/525 nm (excitation/emission) filter set (see Note 23). The percentage of cells with punctuate structures should be assessed among a sample of statistical relevance (see Notes 19 and 24).

3.2.5. CellTracker™ Green (CMFDA) Staining 1. 25 × 103 HeLa cells are cultured in 24-well plates with sterile coverslips (0.5 mL growth medium). 2. After the stimulation or inhibition of autophagy, cells are stained with 1 μM CMFDA at 37ºC in a 5% CO2 incubator for 30 min. 3. Then, cells can be fixed and counterstained with Hoechst 33342 as described above (see Subheading 3.2.2., steps 5–7, and Notes 13–17). 4. CMFDA staining is routinely examined in fluorescence microscopy with a 488/525 (excitation/emission) filter set (see Note 25). 5. With this technique, AV vacuoles appear as “holes” within a diffuse green cytoplasmic fluorescence. The percentage of cells bearing at least one discernable cytoplasmic vacuole should be determined (see Note 19).

3.2.6. LC3-I–to–LC3-II Conversion by Immunoblotting 3.2.6.1. Preparation of Samples 1. HeLa cells (5 × 105 in 3 mL of growth medium) are cultured in 6-well plates. 2. Following the desired treatments to induce or suppress autophagy, cells are collected by trypsinization, washed once in cold (4°C) PBS and pelleted (300–500g, 5 min, 4°C) (see Note 26). 3. Supernatant is discarded, and pellets are lysed in 30–50 μL of lysis buffer (see Notes 27 and 28). 4. The protein concentration of lysates is determined by means of the Bio-Rad DC Protein Assay, following the manufacturer’s instructions (see Note 29).


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5. Then, 4–6 μL of 5X loading buffer are added to 20–80 μg of proteins from each lysate, and the final volume is adjusted to 20–30 μL with dH2 O (see Notes 30 and 31). 6. Finally, samples are incubated for 5 min at 100°C. After cooling to RT, samples are ready for separation by sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS-PAGE) (see Note 32).

3.2.6.2. SDS-PAGE 1. These instructions assume the use of a BioRad mini-PROTEAN 3 system for gel electrophoresis. The protocol is easily adaptable to other formats. 2. A 1.0-mm-thick, 15% running gel (see Note 33) is prepared by mixing 1.8 mL of 40% acrylamide:N-N’-methylenediacrylamide (29/1) solution (see Note 34), 1.25 mL of running gel buffer (pH 8.8), 1.8 mL of dH2 O, 50 μL of 10% SDS, 10 μL of TEMED (see Notes 35 and 36), and 26 μL of 10% APS (see Notes 37 and 38). Immediately after the addition of APS (to avoid polymerization in tube; see Notes 39 and 40), gel should be poured into preassembled glasses up to approximately two thirds of their height (see Notes 41 and 42). 3. Quickly following, overlay gel with isopropanol (see Note 43) to ensure a flat surface and exclude air. 4. After the polymerization (see Note 44), isopropanol is poured off and the top of the gel is washed with dH2 O. 5. The stacking gel (see Note 45) is prepared by mixing 250 μL of 40% acrylamide/bisacrylamide (29/1) solution (see Note 34), 625 μL of stacking gel buffer (pH 6.8), 1.6 mL of dH2 O, 50 μL of 10% SDS, 9 μL of TEMED (see Notes 35 and 36), and 15 μL of APS (see Notes 37 and 38). Rapidly following the addition of APS, the stacking gel is poured on top of the polymerized running gel and the comb for the creation of wells is inserted (see Notes 38 and 46). 6. Once the stacking gel is polymerized, the comb is gently removed and the gel unit is moved to the cell for separation. Upper and lower chambers of the gel unit are filled with migration buffer (1X) (see Note 47), which is used also to gently wash sample wells. 7. At this stage, each well is loaded with a sample prepared as described above (see Subheading 3.2.6.1.). One well is reserved for protein MW markers (see Note 48). 8. Finally, the gel unit is connected to power supply and the proteins are separated in constant-field mode (see Note 49). The run should be arrested when MW markers are separated throughout entire height of the gel (see Note 50). 3.2.6.3. Immunoblotting (Western Blotting) 1. Two sponges and four sheets of filter paper of approximately the same size as the running gel are incubated in transfer buffer (1X) for 1–2 min at RT. 2. One nitrocellulose membrane of approximately the same size of the running gel is activated by incubation in dH2 O for 5–10 min at RT, then equilibrated in transfer buffer.

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3. The gel unit is disconnected from the power supply and disassembled. The stacking gel is discarded. 4. The transfer “sandwich” is prepared by overlaying the following components (see Note 51): one sponge, two sheets of Whatmann paper, the nitrocellulose membrane, the running gel, two sheets of Whatmann paper, one sponge. 5. The “sandwich” is placed in a transfer cassette (see Note 52), which is subsequently inserted into the transfer apparatus. Special attention should be paid in the orientation of the cassette to ensure that the nitrocellulose membrane is placed between the gel and the anode (see Note 53). 6. To control temperature, an ice block is placed next to the transfer cassette (see Note 54). The cell is then completely filled with transfer buffer (1X). A magnetic stir-bar is placed in the cell to help in maintaining a homogeneous solution and temperature. 7. The power supply is activated, and the transfer is accomplished at the constant voltage of 90 V for 2 h (see Notes 55 and 56). 8. Once the transfer is complete, the cassette is disassembled. Sponges and sheets are removed (see Note 57). The colored MW markers should be visible on the nitrocellulose membrane. 9. The bound proteins can be visualized by a rapid incubation with Ponceau S staining solution (see Note 58). Prior to the blocking, Ponceau S should be removed by rinsing the membrane with PBS or dH2 O. 10. To block unspecific binding sites, the nitrocellulose membrane is incubated on a rocking platform in 10 mL of blocking buffer (45–40 min, RT). 11. Blocking buffer is discarded, membrane is quickly rinsed in rinsing buffer and a 1/200 dilution of rabbit anti-LC3 polyclonal antibody (in 5–10 mL of blocking buffer) is added. Incubation with the primary antibody can be performed either at RT for 2 h or at 4°C overnight, in both cases on the rocking platform. 12. Once the primary antibody is removed, the membrane is washed three times with rinsing buffer (5 min, RT). 13. Upon washing, membrane is incubated for 1 h at RT (on the rocking platform) with a freshly prepared 1/5000 dilution in blocking buffer of goat anti-rabbit horseradish peroxidase (HRP)–labeled secondary antibody. 14. Secondary antibody solution is removed and the membrane is again washed three times (5 min, RT) with rinsing buffer. 15. The membrane is overlaid with 2 mL of chemiluminescent substrate for 1 min (see Note 59). Then, chemiluminescent substrate is removed and the membrane is moved to an x-ray film cassette. 16. In the dark room, a photographic film (see Note 60) is exposed to the membrane for a suitable time, typically 3 or 4 min (see Notes 61 and 62). 17. Finally, exposed films are developed in a common table-top film processor. 18. To ensure equal loading of the samples onto the gel (thus comparability of the bands), membrane can be dehybridized from bound antibodies by incubation in dehybridizing buffer for 15 min at RT, followed by one wash in rinsing buffer and by repetition of the steps 10–17 with the following antibodies:


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Primary antibody: mouse anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) monoclonal antibody (see Note 63), 1/10000 in 10 mL of blocking buffer, 1 h, RT. Secondary antibody: goat anti-mouse HRP-labeled secondary antibody, 1/5000 in 10 mL of blocking buffer, 1 h, RT.

LC3-II is the posttranslationally processed form of LC3, which localizes to autophagosomes and autophagolysosomes during autophagy (6). Under nonautophagic conditions, the AV marker LC3 appears as a single band of approximately 18 kDa, corresponding to LC3-I, the uncleaved isoform. Following the induction of autophagy, LC3-II (the cleaved isoform of LC3) can be visualized as an additional band with a slightly higher electrophoretic mobility than LC3-I (approximately 16 kDa). 3.3. Methods to Measure the Colocalization of Cytoplasmic Organelles with Autophagic Vacuoles 3.3.1. Lysosomes Lysosomes are acidic vesicles that participate in the degradation of extracellular macromolecules that are taken up via endocytosis, micropinocytosis, or phagocytosis. Lysosome are also implicated in the catabolism of cytoplasmic components through autophagy (36). The most abundant proteins found in lysosomal membranes are the lysosome-associated membrane proteins 1 and 2 (LAMP-1 and -2), which together constitute almost 50% of all lysosomal membrane proteins. To study the fusion between AV and lysosomes, colocalization experiments can be performed according to different staining techniques. 3.3.1.1. LAMP-2 Immunofluorescence Staining 1. HeLa cells are cultured in 24-well plates, transfected with the LC3-GFP plasmid and treated as described above (see Subheading 3.2.2., steps 1–4, and Notes 4–6, 11, and 12). 2. After the desired stimulation, growth medium is discarded, cells are washed twice with PBS, and incubated with fixative solution for 30 min at RT. Thereafter, cells are rinsed an additional three times with PBS (see Notes 13–16). 3. Incubation for 10 min (RT) in 0.1% SDS is used to permeabilize cells. Then, cells are rinsed three times with PBS and nonspecific binding sites are blocked by incubating the samples for 20 min at RT with 10% FBS (in PBS). Subsequently, cells are again washed once with PBS. 4. For staining, cells are incubated with the mouse anti-LAMP-2 monoclonal primary antibody (1/100 dilution in 500 μL of BSA buffer). Incubation can be performed either at RT for 1 h or at 4°C overnight.

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5. Primary antibody solution is removed, cells are washed three times with PBS and incubated for 1 h at RT with the Alexa Fluor® 568-conjugated secondary antibody (1/300 dilution in 500 μL of BSA buffer) (see Note 64). 6. Finally, BSA buffer is discarded, stained cells are rinsed three times with PBS and cover slips are mounted as described above (see Subheading 3.2.2., step 7). 7. To determine the colocalization between AV and lysosomes, confocal fluorescence microscopy should be performed at 63X magnification (see Note 65) on a device equipped with a dual FITC-Texas Red filter combination (see Notes 66 and 67). 8. Percentage of colocalization is quantified by software-assisted analysis of green, red, and green-red merged images (see Note 68).

3.3.1.2. LysoTracker® Probes

LysoTracker® probes are fluorescent acidotropic, lysosomotropic, readily cell-permeant fluorochromes for labeling and tracking acidic compartments in living cells. These molecules accumulate in lysosomes and autophagosomes (as well as in other acidic subcellular compartments) due to their chemical character of weak basic amines. Since such probes are not specific for autophagosomes, they cannot be used alone to assess and/or quantify autophagy (41). LysoTracker® probes exist in several variants, which exhibit distinct excitation and emission spectra, to facilitate double or triple stainings. For instance, when AV are labeled with LC3-GFP (emitting in green), colocalization of AV and lysosomes can be visualized with LysoTracker® Blue or Red. 1. 25 × 103 HeLa cells are cultured in 24-well plates, transfected with the LC3-GFP plasmid, and treated as described above (see Subheading 3.2.2., steps 1–4 and Notes 4– 6, 11, and 12). 2. At the end of the desired treatments, growth medium is removed, cells are washed twice with PBS, then prewarmed medium containing the selected LysoTracker® probe (e.g., LysoTracker® Red) at the final concentration of 50–75 nM is added. Staining is performed at 37°C, in 5% CO2 atmosphere, for a minimal time of 30 min (see Note 69). 3. The staining solution is removed, cells are washed once with PBS, and cover slips are mounted as described above (see Subheading 3.2.2., step 7). 4. The cells should be observed using a confocal microscope equipped with a 63X objective (see Notes 65 and 66). A dual FITC-Texas Red filter combination is appropriate to visualize both the LC3-GFP fusion protein and LysoTracker® Red (see Note 67). 5. To determine the percentage of colocalization, software-assisted analysis of green, red, and red-green merged images is performed (see Note 68).

3.3.2. Mitochondria Mitochondria are vital organelles for cellular metabolism and bioenergetics, but they are also the major regulators of cell death. Indeed, in many (if not all)


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paradigms of apoptosis, MMP represents the point of no return in the cascade of events that ultimately leads to the cell’s demise (42). MMP results in the leakage of potentially toxic proteins from the mitochondrial intermembrane space (IMS) into the cytosol. For instance, while cytochrome c (Cyt c) in the IMS has vital functions (by acting as an electron shuttle in oxidative phosphorylation), it participates in the activation of caspases when it is released from mitochondria (43). In addition, mitochondria play a role in stress responses and can produce reactive oxygen species (ROS) when damaged (44,45). Selective degradation of mitochondria by autophagy is also known as “mitophagy” and is thought to be promoted by their functional impairment and/or by MMP. Mitophagy may ensure the removal of damaged and potentially dangerous mitochondria, thus acting as a quality control mechanism (46). 3.3.2.1. Transfection with pDsRed2-mito

pDsRed2-mito is a mammalian expression vector that encodes a fusion of Discosoma sp. red fluorescent protein (DsRed2) (47) and the mitochondrial targeting sequence from subunit VIII of human cytochrome c oxidase (mito) (48,49). Upon expression, this chimeric protein readily localizes to the mitochondrial matrix, thus allowing for the visualization and tracking of mitochondria. The simultaneous detection of AV (with LC3-GFP) and mitochondria (with DsRed2-mito) provides a means to estimate the degree of mitophagy in cells. 1. HeLa cells are seeded in 24-well plates, as described above (see Subheading 3.2.2., step 1). 2. Twelve to 24 h after seeding, cells are co-transfected with LC3-GFP and pDsRed2mito plasmids. For the transient transfection of cells with two plasmids, the same protocol reported above (see Subheading 3.2.2., steps 2–4, and Notes 4–6, 11, and 12) is applied. Importantly, the ratio between the total amount of plasmid DNA and Lipofectamine™ has to be maintained, so 2 μg of LC3-GFP plasmid and 2 μg of pDsRed2-mito should be employed. 3. Following the desired treatments, cells are washed once with PBS and cover slips are mounted as detailed above (see Subheading 3.2.2., step 7). 4. Confocal fluorescence microscopy is then performed at 63X magnification (see Notes 65 and 66). The LC3-GFP and DsRed2-mito fusion proteins can be appropriately visualized by employing a dual FITC-Texas Red filter combination (see Note 67). 5. The degree of colocalization between green and red fluorescence is determined as described above (see Subheading 3.3.1.1. and Note 68).

3.3.2.2. MitoTracker® Probes

MitoTracker® probes are cell permeant, mitochondrion-selective, fluorescent stains that are concentrated by active mitochondria and well retained during

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fixation (50). This latter feature (which is not shared by conventional mitochondrial probes such as rhodamine 123 or tetramethylrosamine) seems to depend on the presence of a mildly thiol-reactive chloromethyl moiety. As for LysoTracker® probes, several variants of MitoTracker® probes exist for use in double or triple staining protocols (e.g., LC3-GFP and MitoTracker® Red to determine the colocalization of AV and mitochondria). 1. For colocalization studies of AV marked with LC3-GFP fusion protein and mitochondria stained with MitoTracker® Red CMXRos, the same protocol described above for LysoTracker® Red (see Subheading 3.3.1.2., steps 1–3, and Notes 4–6, 11, 12, and 69) is followed. 2. In contrast to LysoTracker® probes, MitoTracker® probes should be employed at the final concentration of 150 nM. 3. Confocal microscopy assessments are performed as detailed above (see Subheading 3.3.1.2.,– steps 4 and 5, and Notes 65–68).

3.3.2.3. Heat Shock 60 kDa Protein (Hsp60) Immunofluorescence Staining

In eukaryotes, Hsp60 is mainly localized to the mitochondrial matrix and is not released during cells death. Thus, it has been widely employed as a specific marker of mitochondria (51,52). 1. HeLa cells are cultured, transfected, treated, permeabilized, and fixed as described above for LAMP-2 staining (see Subheading 3.3.1.1., steps 1–3, and Notes 4–6, 11–16). 2. For staining, cells are incubated with the mouse anti-Hsp60 monoclonal primary antibody (1/100 dilution in 500 μL of BSA buffer). Incubation can be performed either at RT for 1 h or at 4°C overnight. 3. Subsequent washing, incubation with the Alexa Fluor® 568-conjugated secondary antibody, mounting and confocal microscopy determinations are carried out as described above (see Subheading 3.3.1.1., steps 5–8 and Notes 64–68).

3.3.3. Endoplasmic Reticulum The ER is a network of or interconnected tubules, vesicles, and sacs involved in multiple processes, including the regulation of cytosolic calcium concentrations. Several studies suggest that the membranes of AV originate from ER. Calreticulin is the most prominent calcium-binding protein found in the ER lumen (53) and can be used as a marker in AV-ER colocalization experiments. 1. HeLa cells are grown, transfected, treated, permeabilized, and fixed as described above for LAMP-2 staining (see Subheading 3.3.1.1., steps 1–3 and Notes 4–6, 11–16).


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2. For staining, cells are incubated with the rabbit anti-calreticulin polyclonal primary antibody (1/100 dilution in 500 μL of BSA buffer). Incubation can be performed either at RT for 1 h or at 4°C overnight. 3. Subsequent washing, staining with the Alexa Fluor® 568-conjugated secondary antibody, mounting, and confocal microscopy determinations are carried out as described above (see Subheading 3.3.1.1., steps 5–8 and Notes 64–68).

3.4. Methods for Measuring Cell Death–Related Parameters Most cell death in vertebrates proceeds via the intrinsic, mitochondrial pathway of apoptosis (54). In several models of cell death, MMP is considered as the point of no return in the pro-apoptotic cascade of events (42). MMP may result from the activity of pro-apoptotic members of the Bcl-2 family (such as Bax, Bak, and BH3-only proteins like Bid) (55,56) or from the opening of a multiprotein complex, the permeability transition pore complex (PTPC) (57,58). The sudden increase in permeability of the mitochondrial inner membrane (IM) that derives from PTPC opening (a process known as mitochondrial permeability transition, MPT) leads to dissipation of the mitochondrial transmembrane potentialm and eventually to the rupture of the mitochondrial outer membrane (OM) (42). Irrespective of its initiation (be it mediated by Bcl-2 pro-apoptotic proteins or by opening of the PTPC), MMP leads to the functional impairment of mitochondria and to the release of IMS proteins into the cytosol. These include caspase activators like Cyt c (43,59), Omi/HtrA2 (Omi stressregulated endoprotease/high temperature requirement protein A 2) (60) and Smac/DIABLO (second mitochondria-derived activator of caspase/direct IAP binding protein with a low pI) (61,62), as well as caspase-independent cell death effectors such as apoptosis-inducing factor (AIF) (63,64) and endonuclease G (EndoG) (65). The assessment of early mitochondrial alterations allows for to identification of cells that are committed to death but have not yet displayed an apoptotic phenotype. The m (providing direct indications on the IM permeability status) can be assessed by means of cationic fluorescent probes (66). Such lipophilic dyes (e.g., DiOC6 (3), TMRM) accumulate in mitochondria driven by m , according to the Nernst equation, and can be used in situ for microscopic assessments of m as well as in cytofluorometric-based procedures for m quantification (52). OM permeabilization can be assessed by the detection of IMS proteins (e.g., Cyt c, AIF) in the cytosol. This can be achieved by immunoblotting subcellular fractions with antibodies against Cyt c or AIF, or by in situ immunofluorescence microscopy on fixed and permeabilized cells (51,67,68). As an alternative, cells can be transfected with cDNA constructs encoding IMS proteins fused to a GFP moiety (69,70). Such chimeric proteins are targeted to IMS as their

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normal counterparts. Upon apoptosis stimulation, videomicroscopy permits to follow the redistribution of GFP-tagged proteins from IMS to other subcellular compartments in real time. Among these techniques, immunofluorescence microscopy seems the most appropriate to detect the release of IMS protein, since it provides more accurate information on their redistribution than subcellular fractionation followed by immunoblotting, and it is not associated with the technical requirements of videomicroscopy (52). One of the classical hallmarks of apoptosis is the externalization of phosphatidylserine (PS) and phosphatidylethanolamine (PE) (71,72). In normal conditions, these phospholipids are sequestered on the cytoplasmic leaflet of the plasma membrane, but they translocate to the outer leaflet upon apoptosis induction. Fluorescent Annexin V that specifically binds to PS exposed on the extracytoplasmic side of the plasma membrane can be used to identify apoptotic cells. Following MMP, mitochondrial uncoupling results in enhanced generation of ROS. This alteration may be quantified (in association with other mitochondrial parameters like the m ) by means of cytofluorometric techniques. For instance, the nonfluorescent probe hydroethidine (HE) is oxidized by ROS to ethidium bromide (EthBr), which emits red fluorescence (73). In particular, HE is more sensitive to superoxide anion than 2 ,7 -dichlorofluorescein diacetate, which preferentially measures H2 O2 formation (45,73,74). Alternatively, ROSinduced damage of mitochondria can be determined indirectly, by assessing the oxidation state of cardiolipin, a lipid restricted to the IM. This may be achieved with nonyl acridine orange (NAO), a fluorochrome that specifically interacts with nonoxidized, intact cardiolipin (75). Consequently, a reduction in NAO fluorescence is an indicator of ROS-mediated cardiolipin oxidation. Downstream MMP, the activation of the caspase cascade is frequently associated with the onset of apoptotic cell death (42,54). However, caspaseindependent routes to death have been reported in several models of apoptosis (76). Moreover, caspase activation occurs in several nonapoptotic scenarios (77). As it stands, the detection of activated caspases cannot be used alone to assess apoptosis, but it may provide a useful complementary assay to other techniques. Among the various caspases that execute the apoptotic program, a prominent one is caspase-3 (78). Hence, detection of the large fragment (17/19 kDa) of active caspase-3 (caspase-3a), resulting from the caspase-9– mediated cleavage adjacent to Asp175, is commonly considered as an indicator of apoptosis induction. The integrity of plasma membrane represents a central difference between apoptosis and necrosis. During primary necrosis, the generalized swelling of the cell and of cytoplasmic organelles results in the early breakdown of plasma membrane. On the contrary, plasma membrane integrity is maintained until


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the late stages of apoptosis, before secondary necrosis occurs (22). Thus, the vital dye propidium iodide (PI, emitting in red), which cannot enter living cells nor cells undergoing apoptosis, can be used to differentiate between apoptotic and necrotic cells by means of flow cytometry. Also, other dyes that bind stoichiometrically to double-stranded DNA, such as 4 ,6-diamidino2-phenylindole dihydrochloride (DAPI) and Hoechst 33342, can be used to measure cell viability. Moreover, these molecules can be employed to label the nuclei of permeabilized cells for fluorescence-based assessments. When examined by fluorescence microscopy, nuclei from normal cells glow brightly and homogeneously and can be easily differentiated from apoptotic nuclei, which display chromatin condensation (nuclear pyknosis, at early stages) or a total segmented morphology (karyorhexis, later during the process (22). 3.4.1. Analysis of m Dissipation 1. 25 × 103 HeLa cells are seeded in 24-well plates (0.5 mL of growth medium per well). After 12–24 h, cells are treated with the desired stimuli. 2. At the end of treatments, cells are collected by trypsinization (300 μL of trypsin/EDTA per well), and spun at 300g for 5 min (RT) (see Note 70). 3. Supernatant is discarded and cells are labeled at 37°C either with 150 nM TMRM (emitting in red) or with 40 nM DiOC6 (3) (emitting in green) in 200 μL of growth medium (see Notes 71 and 72). 4. Cytofluorometric acquisitions are performed by means of a conventional cytofluorometer equipped with a single laser for excitation (see Notes 73 and 74). 5. Analysis should be performed on viable, normal-sized cells by gating a single population with normal forward and side scatter parameters (see Note 75). Each experiment should be supported by an appropriate set of controls (see Note 76).

3.4.2. Analysis of Phosphatidylserine Externalization Apoptosis-related PS translocation to the outer leaflet of plasma membrane is detected by staining the cells with fluorescein isothiocyanate (FITC)-conjugated Annexin V (see Note 77). 1. HeLa cells are cultured, treated, and collected as described above (see Subheading 3.4.1., steps 1 and 2, and Note 70). 2. Supernatant is discarded and cells are washed in 1 mL of binding buffer and centrifuged at 300g for 10 min (RT). Supernatant is removed completely and the washing step is repeated. 3. Cells are resuspended in 100 μL of binding buffer and 10 μL of Annexin V-FITC are added. 4. Samples are carefully mixed and incubated at RT under protection from light for 15 min (see Notes 78 and 79).

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5. Cells are washed by adding 1 mL binding buffer and spun at 300 g for 10 min (RT). Supernatant is discarded and cells are resuspended in 500 μL of binding buffer (see Note 80). 6. Immediately following, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 4 and 5, and Notes 72–73, 75, 80, and 81).

3.4.3. Determination of Mitochondrial ROS Generation and Local ROS Effects 1. 100 × 103 HeLa cells are cultured in 12-well plates (1 mL of growth medium). 2. Twelve to 24 h later, the desired treatments are administered to the cultures. 3. At the end of stimulation, cells are trypsinized (500 μL trypsin/EDTA per well), and spun at 300g for 5 min (RT) (see Note 70). 4. Upon removal of the supernatant, cells are labeled either with 25 μM HE or with 100 nM NAO in 500 μL of culture medium. 5. Labeling is performed at 37°C for 10 min. 6. Then, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 4 and 5, and Notes 72, 73, 75, 82, and 83).

3.4.4. Determination of the Mitochondrial Release of IMS Proteins by Immunofluorescence Microscopy 1. 25 × 103 HeLa cells are seeded in 24-well plates as described above (see Subheading 3.1.1., step 1). After 12–24 h, cells are treated with the desired stimuli. 2. Following treatments, cells are fixed and permeabilized as described before (see Subheading 3.3.1.1., steps 2 and 3 and Notes 13–16). 3. For double staining, cells are then incubated with the rabbit anti-AIF and with the mouse anti-cytochrome c primary antibodies (both at 1/100 dilution in 500 μl of BSA buffer). Staining can be performed indifferently at RT for 1 h or at 4°C overnight (see Note 84). 4. Subsequent washing, staining with anti-mouse and anti-rabbit Alexa Fluor® conjugated secondary antibodies, and mounting are performed as described above (see Subheading 3.3.1.1., steps 5 and 6 and Note 64). 5. Examination is then carried out at the fluorescence microscope by using an oil immersion objective (50–65× magnification). Excitation and emission filters should be selected according to the following absorption/emission peaks: 495/519 nm for Alexa Fluor® 488 (green emission); 578/603 nm for Alexa Fluor® 568 (red emission); 352/461 nm for Hoechst 33342 (blue emission).

A distinct, punctuate cytoplasmic pattern of fluorescence indicates the mitochondrial localization of AIF (red fluorescence) and Cyt c (green fluorescence). This can be verified by counterstaining mitochondria with antibodies directed against sessile mitochondrial markers, including OM integral (e.g.,


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voltage-dependent anion channel) or matrix proteins (e.g., Hsp60). As an alternative, mitochondria can be prestained with MitoTracker® probes, as detailed above (see Subheading 3.3.2.2.). On the contrary, a diffuse staining pattern can be observed upon the mitochondrial release of IMS proteins. Following the release from mitochondria, Cyt c is found mainly in the cytosol (43), whereas AIF accumulates in the nucleus (63,79). The percentage of cells exhibiting a redistribution of AIF and Cyt c should be determined in a statistically meaningful population (see Note 19). 3.4.5. Caspase 3a Staining 1. HeLa cells are seeded in 24-well plates, treated, permeabilized, and fixed as detailed above (see Subheading 3.1.1., step 1, Subheading 3.3.1.1., steps 2 and 3, and Notes 13–16). 2. For staining, cells are then incubated with the rabbit anti-caspase-3a (1/100 in 500 μL of BSA buffer). Staining can be performed indifferently at RT for 1 h or at 4°C overnight (see Note 84). 3. Subsequent washing, staining with anti-rabbit Alexa Fluor® -conjugated secondary antibodies, mounting, and examination are performed as previously detailed (see Subheading 3.3.1.1., steps 5 and 6 and Note 64, Subheading 3.4.4., step 5). 4. Caspase-3 activation results in a bright, diffuse cytoplasmic red fluorescence, markedly in contrast with the background signal of negative cells. For each sample, the percentage of positivity for caspase-3a should be quantified in a cell population of statistical relevance (see Note 19).

3.4.6. Detection of Plasma Membrane Integrity Dead cells incorporate the so-called vital dyes, that are normally excluded by the barrier given by an intact plasma membrane. One of the most widely used vital dyes is propidium iodide (PI). 1. HeLa cells are cultured, treated, and collected as described above (see Subheading 3.4.1., steps 1 and 2 and Note 70). 2. Supernatant is discarded and cells are labeled at 37°C with 1 μg/mL PI in 200 μL of growth medium (see Note 71). 3. Then, cytofluorometric acquisitions are performed as described above (see Subheading 3.4.1., steps 3–5 and Notes 72, 73, 75, 80, and 85).

3.5. A General Strategy for the Resolution of the Central Silemma of Autophagic Cell Death As outlined in the introduction to this chapter, one of the major problems raised by the observation of autophagic cell death (type 2 cell death) concerns

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causality. Does cell death simply occur with autophagy, which would constitute a late and desperate attempt of the cell to adapt to stress? Or is cell death mediated through autophagy, meaning that self-eating is one of steps that leads to self-killing? In the case of cells presenting massive autophagic vacuolization followed by plasma membrane barrier and disintegration, cell death should be considered as occurring through autophagy sensu stricto if all the following requirements are met: 1. Cell death (loss of membrane integrity with positive staining for PI or similar vital dyes) should be preceded by all signs of autophagy (double-membraned vesicles, LC3-GFP aggregation, LC3 proteolytic maturation, increase of the acidic subcellular compartment, enhanced turnover of long-lived proteins, co-localization of lysosomal and mitochondrial or ER markers), and these signs should be inhibitable by siRNAs targeting essential proteins of the process (atg5, atg6/Beclin-1, atg10, atg12, hVps34). 2. Cell death should occur without unequivocal morphological manifestations of apoptosis (e.g., pyknosis, karyorhexis, shrinkage, and formation of the apoptotic bodies). However, cell death may be linked at late stages to mitochondrial permeabilization and caspase activation. These alterations would indicate that apoptotic effector mechanisms have been activated. 3. Most importantly, cell death in all its possible manifestations (autophagic, apoptotic, or necrotic) should be inhibited by genetic manipulations designed to reduce or suppress the expression of atg genes or hVps34. Thus, the inhibition of autophagy by specific methods (as opposed to the unspecific pharmacological modulators that are currently available) should prevent cell death and maintain the cells healthy and viable.

If it is possible to substantiate by means of these methods that cells can effectively die through autophagy, then it will become a challenging endeavor for future investigation to determine the functional importance of autophagic cell death in tissue homeostasis, remodeling, and pathology.

4. Notes 1. Working dilutions of the inducers and inhibitors of autophagy should be prepared from the stock solutions shortly before use. Intermediate dilutions, when necessary, can be made either in sterile PBS or in growth medium. 2. The minimal time necessary to observe a stimulation of the autophagic process varies according to the inducer from 1 h (e.g., upon stimulation with xestospongin C) up to 12 h (e.g., after depletion of serum and amino acids). Thus, to avoid excessive toxicity, it is recommendable NOT to incubate cells for longer than 36–48 h prior to the quantification of autophagic vacuoles. 3. Contrarily to the manufacturer’s instructions (recommending 30–50% of confluence), we found that optimal transfection rates are attained when cells


4.

5.

6.

7.

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are slightly more confluent (50–70%). However, excessive confluence (at levels coinciding with a reduction of proliferation) should be carefully avoided, since it results in a significant drop of the efficacy of transfection. When the diluted Oligofectamine™ and the siRNA solution are mixed, the solution may appear cloudy. The same applies to Lipofectamine™ and plasmid solutions. The liposome-mediated siRNA transfection protocol is carried out entirely at RT under a common safety cabinet. However, it is recommendable to keep the tubes containing the stock solutions of siRNAs and Oligofectamine™ in an ice bath (and to return them to storage conditions immediately after use), to avoid the degradation of reagents, and to minimize solvent evaporation (both of which may eventually affect the concentration of the stocks). The same applies to the liposome-mediated transfection of plasmids. The absence of serum is especially important for optimal formation of the Oligofectamine™:siRNA complexes. Nevertheless, contrary to common beliefs, we found that complexes (once properly assembled in serum-free conditions) can be added to cells cultured in complete growth medium without relevant reductions of the transfection efficiency. This practice removes the need for serum addition 4 h after transfection. The same applies to the liposome-mediated transfection of plasmids. Given the very high affinity of transfection complexes for the plasma membrane, they should be added to cells dropwise, by trying to cover the whole surface of the growth medium, in order to avoid intrawell variations of the transfection efficiency. Due to multiple variables, including the efficacy of different siRNAs (also targeting the same transcript) as well as the half-life of target proteins, the time required for optimal knock-down may range from a few hours to several days. As a guideline, we recorded satisfactory levels of downregulation of several proteins involved in autophagy (e.g., Beclin-1, LAMP-2, Atg5, Atg10, Atg12, hVps34) 48 h after transfection. However, it is recommendable to check the kinetics of protein downregulation for each siRNA and specific experimental setting. This should be accomplished both at the mRNA level (by RT-PCR, using the appropriate primers) and at the protein level (by immunoblotting with the specific antibodies). As a negative control, cells transfected with scrambled siRNAs or with siRNAs targeting irrelevant sequences (not found in the cellular genome) should be carried along the experimental procedure and subjected to the same treatments administered to cells transfected with siRNA targeting proteins essential for autophagy. At this stage, the seeding concentration heavily depends on the duration and strength of the subsequent treatments. As a guideline, for treatments of 24 h or less (administered 24 h after plating) we use seed 100 × 103 cells in 1 mL of growth medium (12-well plates) or 25 × 103 cells in 0.5 mL of growth medium (24-well plates).

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11. We recorded the expression of the LC3-GFP fusion protein already 12 h after transfection. Faster expression is not excluded, but should be controlled in the specific experimental setting. 12. As for siRNAs, an adequate control has to be carried along the entire experimental procedure and treated as samples (cells transfected with the plasmid encoding the LC3-GFP chimeric protein). To this aim, untransfected cells may be used. However, a more stringent control is provided by cells transfected with the empty cloning vector. 13. PFA is toxic by inhalation and should be handled under an appropriate fume hood. 14. Stability of the PFA solution is limited. Accordingly, it should be prepared shortly before use. Once dissolved, it is recommendable to keep PFA on ice bath throughout the experiment. 15. Picric acid (chemical name 2,4,6-trinitrophenol) is a close derivative of trinitrotoluene (TNT) and is toxic and explosive if allowed to dry out. Thus, maximal care should be used when handling picric acid. 16. At washing steps, gentle pipetting is recommended to avoid the detachment of cells from the coverslips. 17. Hoechst 33342 is carcinogenic and mutagenic. 18. While wild-type GFP has two excitation peaks, a major one at 395 nm (in the long UV range) and a smaller one at 478 nm (blue), the red-shifted mutant variant encoded by the plasmid pEGFP-C1 (cloning vector from Clontech Laboratories, Palo Alto, CA) is characterized by a single absorption peak at 488 nm and higher expression in mammalian cells. 19. For quantification purposes, the frequency of cells exhibiting cytoplasmic LC3GFP aggregation should be determined among (at least) 200 cells. Moreover, the number of AV (green dots) per cell should be counted in a representative sample of (at least) 50 cells. The same considerations about the population size apply to other techniques for AV quantification (e.g., MDC or CMFDA staining), as well as to the assessment of translocation of IMS proteins and caspase-3 activation. 20. l-[U-14 C]-Valine is a radiochemical. Its handling and storage should conform to the current safety rules. 21. The prechase period is crucial to get rid of short-lived proteins, which are indeed degraded during this step. 22. At this stage, the use of an autophagy inhibitor provides the required negative control. 23. MDC is an autofluorescent molecule characterized by a relatively weak emission, peaking at 525 nm. However, it has been demonstrated that it behaves as a solvent polarity probe (80) and that its interaction with membrane lipids (as occurring in AV) enhances by severalfold its emission, while shifting its emission to 498 nm. 24. To distinguish the characteristic vesicular distribution of MDC in autophagosomes, each experiment should be performed and scored with the appropriate controls. To this aim, NF medium combined or not with an autophagy inhibitor of choice may provide positive and negative control conditions, respectively.


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25. CMFDA is characterized by an absorption and fluorescence emission maxima of 492 and 517 nm, respectively, as determined in aqueous buffer or methanol. Values may exhibit slight variations in cellular environments. 26. To minimize proteolytic degradation, after washing samples should be kept in ice bath until the addition of loading buffer and boiling or storage. 27. Lysis should be performed in the minimal volume of lysis buffer that allows for total resuspension of the pellets (usually 30 μL will suffice). Lysis may be facilitated by thorough pipetting or quick vortexing. Keeping the lysis buffer volume to a minimum reduces the dilution of the protein content of samples, thus resulting in easier gel loading (see also Note 31). 28. Lysates may be stored at –20°C for prolonged periods (several months) without significant degradation of protein content. 29. Protein quantification is performed by interpolating a calibration curve that is built from a serial dilution of BSA. BSA standards are rarely (if ever) included in the kits for quantification and should be prepared shortly before use from the BSA stock solution. 30. The amount of proteins that should be loaded onto the polyacrylamide gel for separation depends on the size and expected expression levels of protein of interest. For large or highly expressed proteins, 20 μg may suffice to visualize the corresponding band upon immunoblotting. For small or rare proteins larger amounts (up to 60–80 μg) are required. 31. When the quantity of proteins for loading has been determined, samples that will be separated on the same gel are prepared in the minimal volume, which allows one to load the same amount of proteins for each lysate and the addition of 5X loading buffer (to a final concentration of 1X). Keeping the loading volume to a minimum facilitates the loading procedure. In case of diluted samples, 6–7X loading buffer may be employed. 32. Boiling is performed to complement the denaturating activity of lysis and loading buffers (containing SDS, reducing agents, and calcium chelators), with the aim to break the inter- and intramolecular bonds responsible of the higher-level (tertiary and quaternary) protein structures. This step ensures that the subsequent separation is truly based on the size of protein subunits, with little influence of protein–protein interaction and native structural features. 33. The percentage of acrylamide directly determines the sieving properties of the gel, thus influencing its separation range and resolution. As a guideline, proteins with a molecular weight higher than 100 kDa should be resolved on 5–10% total acrylamide gels, whereas smaller proteins can be separated on gels containing 10–15% acrylamide. To facilitate pouring and subsequent handling, concentrations of acrylamide higher than 15% should be avoided. 34. Monomeric acrylamide is a neurotoxin and suspected carcinogen. Avoid skin contact and inhalation. Always wear gloves and protective clothing and handle nonpolymerized acrylamide solutions under a fume hood. 35. In polyacrylamide gels, TEMED acts as crosslinking agent. Also its concentration influences the final properties of the polymerized gel.

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36. TEMED is toxic by inhalation. It should be always handled under a fume hood. 37. APS stock solution (10% in dH2 O) is unstable. Storage at 4°C is possible, but never exceed one week. The use of old APS solutions may result in incomplete polymerization of the acrylamide gel. 38. APS generates the free radicals required for the polymerization of acrylamide, thus acting as the catalyst of the reaction. Accordingly, APS should be added as the last component of the gel solution. 39. This protocol reports the amounts of reagents required for a single gel. These quantities may be readily scaled up to pour more gels from the same starting solution. However, no more than four to six gels should be cast from the same solution, to avoid premature polymerization. 40. Hint: Upon pouring, the gel solution remaining in the tube should not be immediately discarded, but left in contact with air. It will help in determining the polymerization state of the gels. 41. Pouring should be performed quickly, while avoiding the formation of bubbles that would interfere with protein migration. 42. It is recommended to leave at least the upper third of the glass height for the stacking gel. 43. Isopropanol may be substituted with pure water. 44. Polymerization time depends on acrylamide concentration, but usually 10–15 min are sufficient. Prolonged polymerization time or incomplete polymerization may result from an old APS stock (see also Note 38). Polymerization can be verified by examining the remnant solution in the tube in which the gel solution has been prepared (see also Note 40). 45. The use of a stacking gel improves the resolution of electrophoresis because it causes proteins to accumulate and stack as a very thin zone at the stacking gel/running gel boundary and because it arranges the proteins according to their order of mobility before their entry into the running gel. 46. The comb (which determines the size of wells) should be selected according to the number and final volume of the samples. 5-, 9-, 10-, and 15-well combs are available, allowing to load approximately 60, 50, 40, and 20 μL of samples, respectively. 47. Importantly, the migration buffer should completely cover the wells but not create a communication between the upper and lower chambers that would compromise the correct electrical flow in the cell. 48. Prestained markers covering different ranges of MW are commercially available and allow for visual follow-up of migration. 49. Before entry in the running gel, voltage should be set at 50–70 mV. Thereafter, voltage can be augmented to 140 mV. It should be kept in mind that higher voltages result in reduced migration time but also in lower resolution. 50. Migration fronts, as indicated by the fast migrating bromophenol blue component of the loading buffer, may be safely run off the gel, if desired. 51. During assembly, special care should be taken to avoid the formation of bubbles (which will interfere with the transfer of proteins) between the various layers of


52.

53.

54.

55. 56. 57. 58.

59. 60. 61.

62.

63.

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the “sandwich.” To remove bubbles, a common 5-mL pipet can be gently rolled on top of each component, before the addition of the next one. Each transfer cassette allows the insertion of two “sandwiches.” To avoid the loss of proteins during the transfer, they should both be oriented in the same fashion (see also Note 53). If the cassette is inserted with the reverse orientation (membrane between the gel and the cathode), negatively charged proteins will be attracted by the anode across the filter paper sheets and the sponge and eventually be lost in the transfer buffer. Due to the intense current, the transfer apparatus temperature tends to increase excessively, if not controlled. As an alternative to the ice block, the entire transfer procedure can be performed in a cold room (4°C). As an alternative, the transfer can be accomplished overnight at a constant current of 45 mA per membrane. To keep the buffer temperature as homogeneous as possible, the transfer should be performed on top of a magnetic stirrer (gentle stirring). Upon careful rinsing in PBS or dH2 O, sponges can be re-used. Filter papers are discarded. At this stage, problems during the transfer (such as bubbles) can be easily identified. Also, membranes can be wrapped in transparent plastic film and photocopied (or digitalized by scanning) to keep track of the correct transfer. At this step, attention should be paid to ensure the even coverage of the membrane with the chemiluminescent substrate. Photographic films are light sensitive and should be handled only in the dark room until the end of the development processing. Exposure time varies greatly according to the intensity of signals from the HRP-catalyzed chemiluminescent reaction. In turn, this depends on several factors, including the amount of protein in the band, the affinity of primary and secondary antibodies, and the time passed from the incubation of the chemiluminescent substrate. For most proteins, exposure times between 1 and 5 min provide good results. When proteins are small or scarcely expressed (or when the antibodies display reduced affinity), longer exposures (up to 10–12 hours) should be attempted. On the contrary, when proteins are large or well represented in the sample (or when the antibodies exhibit high affinity), a few seconds may be sufficient. Some trials may be required to obtain the correct exposition time. Given the time-dependent decay of the luminescent signal, short exposures should be performed before longer ones. If the band of interest cannot be detected upon overnight exposure, the reagent Supersignal West Femto Maximum Sensitivity Substrate may provide enhanced sensitivity. Loading control should be performed with an antibody directed against a highly represented protein, whose amount is stable in most experimental settings. Usually, GAPDH, -tubulin, or -actin are detected as loading control.

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64. To avoid photobleaching of the fluorescent antibodies, samples should be protected from light from this step onward. 65. As an alternative, 40 or 100X objectives may also be employed. 66. We detected the colocalization of two fluorescent signals by means of a Zeiss LSM 510 confocal microscope. 67. The dual FITC-Texas Red filter combination allows for the visualization of LC3GFP (absorption/emission peaks at 488/507 nm) and DsRed2-mito (558/583 nm) fusion proteins, as well as of LysoTracker® Red (577/590 nm), MitoTracker® Red CMXRos (579/599 nm), and the Alexa Fluor® 488 (495/517) and 568 (578/603 nm) fluorochromes. 68. To this aim, we routinely use the free, Java® -based, open-source software ImageJ. 69. Staining times may be extended up to 2 h, according to cell type and/or general conditions of the culture. 70. At this stage, the supernatant of treated cells should not be discarded, to avoid the loss of part of the population detached from the plate upon the treatment. Rather, supernatants should be collected in FACS tubes, to which the corresponding cell suspension will be added after trypsinization. 71. To avoid probe-dependent toxicity to the cells, cytofluorometric acquisitions should be performed within 30 min. 72. When large series of samples are to be analyzed (>12 tubes), the interval between labeling and cytofluorometric analysis should be kept constant. 73. To this aim, we routinely use a Becton Dickinson FACScan cytofluorometer, equipped with an argon ion laser emitting at 488 nm. The following channels are employed for the detection of fluorescent emissions: FL1 for DiOC6 (3), FITC and NAO (also in FL2); FL2 for NAO (also in FL1) and TMRM; FL3 for PI and EthBr (as resulting from HE superoxide anion-mediated oxidation) (see also Notes 74, 80, and 81). 74. DiOC6 (3) is characterized by absorption/emission peaks at 484/501 nm, TMRM by maxima at 543/573 nm. 75. As a reminder, the forward light scatter gives an indication on cell size whereas the side scatter is related to several factors, including granularity, refractive index, and shape of the cells. 76. Since the incorporation of DiOC6 (3) and TMRM can be influenced by parameters not related to mitochondria (e.g., cell size, plasma membrane permeability, activity of multidrug resistance pumps, etc.) (45,66), results can only be interpreted when the difference in staining profiles between different experimental conditions (e.g., control vs. apoptosis) are NOT affected by the incubation with a ionophore causing the complete disruption of m . Hence, an adequate set of controls can be prepared by splitting in 2 aliquots each sample, and by preincubating one of the two series with 50–100 μM CCCP for 10–15 min before staining (37°C, 5% CO2 ). 77. To this aim, we routinely employ the Annexin V-FITC kit from Miltenyi Biotec, according to the manufacturer’s instructions.


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78. Protection from light prevents photobleaching of FITC fluorochrome. 79. Higher temperatures or longer incubation times lead to nonspecific labeling. 80. If desired, immediately before cytofluorometric acquisition, 5 μL of a 100 μg/mL solution of PI can be added for the simultaneous detection of cells with ruptured plasma membranes. PI is characterized by absorption/emission peaks at 535/617 nm (see also Note 85). 81. FITC exhibits an absorption maximum at 494 nm and an emission one at 519 nm. 82. EthBr resulting from the ROS-mediated conversion of HE readily binds to DNA, and this complex is characterized by absorption/emission peaks of 300/603 nm (as opposite to reduced HE, emitting at 420 nm). 83. NAO displays absorption/emission peaks at 500/520 nm. 84. At the same time, nuclear counterstaining can be performed by the addition of 2 μM Hoechst 33342, together with primary antibodies. 85. PI staining may be associated with either DiOC6 (3) or Annexin V to distinguish normal cells (PI− , DiOC6 (3)high , or Annexin V− ), dying cells (PI− , DiOC6 (3)low , or Annexin V+ ), and dead cells (PI+ ) within the same population.

Acknowledgments This work has been supported by a special grant from Ligue National contre le cancer, as well as by grants from Agence Nationale de Recherche, Agence Nationale pour la Recherche sur le Sida, Fondation pour la Recherche Médicale, Institut National du Cancer, and European Commission (RIGHT, Active p53, Trans-Death, Death-Train, ChemoRes). Xestospongin B was a kind gift of Dr. Jordi Molgó. We are grateful to Dr. Gérard Pierron (Laboratoire “Réplication de l’ADN et Ultrastructure du Noyau”–Villejuif, France) for his contributions to the section on electronic microscopy. The authors would like also to thank Anne-Laure Pauleau, Yael Zermati, and Shahul Mouhamad (INSERM Unit “Apoptosis, Cancer and Immunity”–Villejuif, France) for invaluable help and suggestions. References 1. Klionsky, D. J., and Emr, S. D. (2000) Autophagy as a regulated pathway of cellular degradation. Science 290, 1717–1721. 2. Mizushima, N., Ohsumi, Y., and Yoshimori, T. (2002) Autophagosome formation in mammalian cells. Cell Struct. Funct. 27, 421–429. 3. Boya, P., Gonzalez-Polo, R. A., Casares, N., et al. (2005) Inhibition of macroautophagy triggers apoptosis. Mol. Cell Biol. 25, 1025–1040. 4. Klionsky, D. J. (2005) The molecular machinery of autophagy: unanswered questions. J. Cell Sci. 118, 7–18. 5. Ohsumi, Y. (2001) Molecular dissection of autophagy: two ubiquitin-like systems. Nat Rev Mol. Cell Biol. 2, 211–216.

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4 LC3 and Autophagy Isei Tanida, Takashi Ueno, and Eiki Kominami

Summary Microtubule-associated protein 1A/1B-light chain 3 (LC3) is a soluble protein with a molecular mass of ∼17kDa that is distributed ubiquitously in mammalian tissues and cultured cells. During autophagy, autophagosomes engulf cytoplasmic components, including cytosolic proteins and organelles. Concomitantly, a cytosolic form of LC3 (LC3-I) is conjugated to phosphatidylethanolamine to form LC3-phosphatidylethanolamine conjugate (LC3-II), which is recruited to autophagosomal membranes. Autophagosomes fuse with lysosomes to form autolysosomes, and intra-autophagosomal components are degraded by lysosomal hydrolases. At the same time, LC3-II in autolysosomal lumen is degraded. Thus, lysosomal turnover of the autophagosomal marker LC3-II reflects starvationinduced autophagic activity, and detecting LC3 by immunoblotting or immunofluorescence has become a reliable method for monitoring autophagy and autophagy-related processes, including autophagic cell death. Here we describe basic protocols to assay for endogenous LC3-II by immunoblotting, immunoprecipitation, and immunofluorescence.

Key Words: LC3; lipidation; ubiquitylation-like autolysosome; autophagy; ATG conjugation system.

reaction;

autophagosome;

1. Introduction Autophagy is the major protein degradation system responsible for the turnover of bulky cellular constituents (1). As a pivotal cellular housekeeping system, autophagy contributes to maintain intercellular homeostasis (2). In addition, autophagy has been found to play significant roles in antigen presentation, bacterial and viral infection, and cell death (3–6). Dysfunction of autophagy has been proposed as an underlying mechanism for neurodegenerative diseases, muscle diseases, cancer, and hepatic inflammation. From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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During autophagy, unique double-membraned autophagosomes are formed to engulf intracellular components, including organelles such as mitochondria. Autophagosomes fuse with lysosomes to form autolysosomes, and sequestered intra-autophagosomal components are degraded by lysosomal hydrolases. LC3 was originally identified as one of three light chains (LC1, LC2, and LC3) associated with purified MAP1A and MAP1B (7). Until its autophagyspecific role as a mammalian Atg8 homologue became established, LC3 had been long thought to be involved in the regulation of assembly and disassembly of microtubules (8,9). During the formation of autophagosomal membranes, cytosolic LC3 (LC3-I) is conjugated to phosphatidylethanolamine (PE) (10,11) through two consecutive ubiquitylation-like reactions catalyzed by the E1-like enzyme Atg7 and the E2-like enzyme Atg3 (12–14) to LC3-II (8). During the fusion of autophagosomes with lysosomes, intra-autophagosomal LC3-II is also degraded by lysosomal proteases (Fig. 1). Therefore, as a marker of autophagosomal membranes, changes in cellular LC3-II level are connected to the dynamic turnover of LC3-II via the lysosome, i.e., autophagic activity (9). Monitoring LC3-

Fig. 1. Lysosomal turnover of endogenous LC3-II during autophagy. For nutrientrich conditions, HEK293 cells were cultured in DMEM medium containing 10% FCS. Where indicated, cells were treated with the protease inhibitors, E64d (10 μg/mL) and pepstatin A (10 μg/mL) (Inhibitors +) for 2 h, or, as a negative control (Inhibitors -), with the solvent dimethylsulfoxide. For starvation conditions, cells were incubated for 4 h in Krebs Ringer bicarbonate buffer (KRB) in the presence (+) or absence (-) of protease inhibitors. The cells were lysed, total proteins (10 μg per lane) were separated by SDS-PAGE, and endogenous LC3 in the lysates was recognized by immunoblotting with an anti-LC3 antibody (WB: anti-LC3). LC3-I, soluble form of LC3; LC3-II, membrane-bound form of LC3. During the formation of autophagosomes, LC3 is lipidated, and LC3-II is localized on autophagosomes and autolysosomes. While intra-autophagosomal LC3-II is degraded by lysosomal hydrolases, a transient cellular amount of LC3-II does not simply reflect starvation-induced autophagic activity. Since endogenous LC3-II is considerably degraded by lysosomal hydrolases after formation of autolysosomes, lysosomal turnover of LC3-II should be investigated to estimate an autophagic activity using E64d and pepstatin A.

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I and LC3-II by Western blotting, immunoprecipitation, and immunofluorescence is essential for investigating the mechanism of mammalian autophagy. In assaying endogenous LC3-II by immunoblotting, it is important to consider the hydrophobicity of LC3-II. Since LC3-II is a protein–PE conjugate, only part of it can be extracted in the presence of 1% Triton X100 (11). Insufficient extraction, however, will lead to misleading results (Fig. 2). Similar considerations apply to immunofluorescent analyses of endogenous LC3. In this chapter, we describe a protocol for effective extraction of LC3-II. In the latter section of this chapter, we mention several treatment precautions. 2. Materials 2.1. Cell Culture and Lysis 1. Dulbecco‘s modified Eagle‘s medium (DMEM) (Invitrogen, Carlsbad, CA) supplemented with 10% (v/v) fetal calf serum (Invitrogen). 2. Krebs Ringer bicarbonate buffer (KRB) for nutrient starvation: 118.5 mM NaCl, 4.74 mM KCl, 1.18 mM KH2 PO4 , 23.4 mM NaHCO3 , 6 mM glucose, 2.5 mM

Fig. 2. Immunoprecipitation of LC3-I and LC3-II. Mouse liver postnuclear supernatant (2 mg protein in 100 μL) or total lysate of cultured B16 melanoma cells (0.5 mg in 100 μL) was mixed with an equal volume (100 μL) of SDS-lysis solution and subjected to immunoprecipitation as described in the text. The resulting immunoprecipitate was treated with modified Laemmli buffer, separated by SDS-PAGE, and analyzed by immunoblotting. For comparison, the same postnuclear supernatant or the lysate was mixed with an equal volume of PBS containing 1% TX-100. The mixtures were cleared by centrifugation at 15,000g for 10 min. LC3 was immunoprecipitated from the resultant supernatant and the precipitated antigens were analyzed by immonoblotting. SDS-lysis solution; LC3-I and LC3-II immunoprecipitated from the samples treated with SDS-lysis solution, PBS-TX-100; LC3-I and LC3-II immunoprecipitated from the samples treated with PBS containing 1% TX-100.

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3.

4. 5. 6. 7.

8. 9. 10.

CaCl2 , 1.18 mM MgSO4 , and 6 mg/L phenol red (15) (see Note 1), adjusted to pH 7.6 by titration with 1 N NaOH. E64d (Peptide Institute, Inc., Osaka, Japan) (16,17) and pepstatin A (Peptide Institute, Inc.) (16,18) dissolved at 10 mg/mL in dimethyl sulfoxide (special grade for biochemical analysis) (Wako Pure Chemical Industries, Ltd., Tokyo, Japan). These 1000-fold concentrated stock solutions are stored at -20ºC, at which temperature they are stable for at least 3 months (see Note 2). Phosphate-buffered saline (PBS): 20 mM sodium phosphate, pH 7.2, and 150 mM NaCl. CompleteTM protease inhibitor cocktail (Roche Diagnostics). An ultrasonic generator with a micro-probe (model USP-300, Shimadzu, Tokyo, Japan). Modified Laemmli buffer: 0.1 M Tris-HCl, pH 6.8, 2% (w/v) sodium dodecyl sulfate (SDS), 10% (w/v) glycerol, 6% (v/v) ß-mercaptoethanol, and 0.03% (w/v) bromophenol blue. Teflon cell scrapers (Sumitomo Bakelite, Tokyo, Japan). BCA protein assay kit (Pierce Biotechnology, Inc., Rockford, IL). TX100-lysis solution: 2% polyethylene glycol mono-p-isooctylphenyl ether (Triton X100, Nacalai Tesque, Inc., Kyoto, Japan) in PBS with the complete protease inhibitor cocktail (Roche Diagnostics) (for option 2b).

2.2. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Stock solutions for preparing 12.5% SDS-gels: 1.5 M Tris-HCl, pH 8.8 (at 25ºC), 0.5 M Tris-HCl, pH 6.8 (at 25ºC), 30% acrylamide solution (29:1 (w/w) acrylamide: methylene-bisacrylamide) and N,N,N´,N´-tetramethylethylenediamide (TEMED, Wako). 2. Ammonium persulfate (APS): 10% (w/v) in water, prepared immediately before use. 3. Water-saturated isobutanol: Equal volumes of water and isobutanol are shaken in a glass bottle and allowed to separate; the top layer is stored at room temperature. 4. Running buffer: 25 mM Tris, 192 mM glycine, and 0.1% (w/v) SDS. Do not adjust pH. Store at room temperature. 5. Prestained molecular weight markers, broad range (6–175 kDa) (New England Biolabs, Beverly, MA): The solution contains 0.01% (w/v) phenol red and 0.01% (w/v) bromophenol blue. 6. Apparatuses for SDS-gel electrophoresis: NA-1030 (Nihon-Eido, Tokyo, Japan) for a mini-wide gel (1.0 mm thick, 200 mm wide, 105 mm high, suitable for 16–20 samples per gel).

2.3. Western Blotting for LC3-I and LC3-II 1. Bjerrum and Schafer-Nielsen‘s transfer buffer: 48 mM Tris, 39 mM glycine, 20% methanol (analytical grade), pH 9.0–9.4, depending on reagent purity. Do not adjust pH; if lower than pH 9.0, prepare again.

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2. Polyvinylidene difluoride (PVDF) membrane (Durapore membrane GV, pore size 0.22 μm; Millipore, Bedford, MA) and chromatography paper (cat. no. 590; Advantec Japan, Tokyo, Japan). 3. Transblot SD Semi-Dry Transfer Cell (BioRad, Hercules, CA) for transferring proteins from polyacrylamide gels to PVDF membranes. 4. Tris-buffered saline containing polyixyehlene sorbitan monolaurate/Tween 20 (TTBS): 20 mM Tris-HCl, pH 7.5 at 25ºC, 150 mM NaCl, and 0.05% (v/v) polyoxyethylene sorbitan monolaurate (Tween 20). 5. Blocking buffer: 5% (w/v) nonfat dry milk (Snow Brand Milk Products Co. Ltd., Tokyo, Japan) in TTBS. 6. Primary antibody solution: 1 μg/mL anti-LC3 IgG, 1% (w/v) fraction V bovine serum albumin (BSA), 20 mM Tris-HCl, pH 7.5 at 25ºC, 0.1% NaN3 and 150 mM NaCl. When stored at 4ºC, it can be reused at least 10 times within a year. 7. Secondary antibody solution: 0.005% (v/v) goat anti-rabbit IgG antibody (minimum cross-reactivity with other species) conjugated to horse radish peroxidase and 1.5% (v/v) normal goat serum (Invitrogen) in TTBS. 8. Enhanced chemiluminescent reagents: SuperSignal West Dura Extended Duration Substrate or SuperSignal West Pico Chemiluminescent Substrate (Pierce Biotechnology Inc., Rockford, IL).

2.4. Immunoprecipitation of Endogenous LC3 1. SDS-lysis solution: 2% (w/v) SDS in PBS with Complete protease inhibitor cocktail (Roche Diagnostics, Indianapolis, IN). 2. TX100-solution: 1% polyethylene glycol mono-p-isooctylphenyl ether (Triton X100, Nacalai tesque) in PBS with CompleteTM protease inhibitor cocktail (Roche Diagnostics). 3. TX100-wash solution: 2% polyethylene glycol mono-p-isooctylphenyl ether (Triton X100, Nacalai tesque) in PBS with Complete protease inhibitor cocktail (Roche Diagnostics). 4. Protein A-agarose (Santa Cruz Biotechnology, Santa Cruz, CA). 5. Modified Laemmli buffer: 0.1 M Tris-HCl, pH 6.8, 2% (w/v) SDS, 10% (w/v) glycerol, 6% (v/v) ß-mercaptoethanol, and 0.03% (w/v) bromophenol blue. 6. A 5-mL syringe and 25G × 1” needle (Terumo, Tokyo, Japan).

2.5. Immunofluorescence Analysis of Endogenous LC3 1. Cell-fix solution: 4% paraformaldehyde in PBS, pH 7.2, prepared just before use and warmed to 37ºC. 2. Digitonin solution: 50 μg/mL digitonin (Wako) in PBS, pH 7.2. 3. Quenching solution: 50 mM NH4 Cl in PBS. 4. IF-blocking solution: 2% (w/v) BSA, 5% (v/v) normal goat serum (Invitrogen), 20 mM Tris-HCl, pH 7.5, 150 mM NaCl. 5. IF primary antibody solution: 5 μg/ml of affinity purified anti-LC3 IgG in IF-blocking solution.

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6. IF secondary antibody solution: 0.05% (v/v) goat anti-rabbit IgG conjugated to Alexa488, 594 or Q-dot 605 (Invitrogen) in IF-blocking solution. 7. Tris-buffered saline (TBS): 20 mM Tris-HCl, pH 7.5, 150 mM NaCl. 8. Mounting medium: SlowFade light (Invitrogen). 9. Fluorescence microscope (e.g., a Zeiss Axioplan2 fluorescence microscope; Carl Zeiss, Thornwood, NY) and an ORCA-ER CCD camera and Aqua C-imaging system (Hamamatsu Photonics, Tokyo, Japan).

2.6. Antibodies Polyclonal rabbit antibodies and mouse monoclonal antibodies to LC3 were raised to synthetic peptides corresponding to amino-terminal sequences (8) and to LC3 fused with GST expressed in Escherichia coli (19) (see Note 3).

3. Methods It is important to consider the hydrophobicity of LC3-II when preparing cell lysates and during immunoprecipitation. Since LC3-II consists of LC3 conjugated to PE, autophagosomal and autolysosomal LC3-II is only partially solubilized in PBS containing 1% Triton X100, 1% Nonidet P40, or 1% Tween 20. To completely solubilize LC3-II from cells and tissue homogenates, we have boiled samples in PBS containing 1% SDS or used PBS containing 2% Triton X100. Under less stringent conditions, LC3-II is insufficiently extracted (Fig. 2). Rather than immunostaining of endogenous LC3-II, a recombinant fluorescent protein, GFP-LC3, overexpressed in cells has often been employed to visualize autophagosomes. Even after autolysosomal degradation, however, a fraction of the free GFP cleaved from GFP-LC3 remains resistant to digestion in the lumen of autolysosomes. Therefore, when using GFP-LC3 as a reporter, care must be taken to avoid measuring the pseudo-fluorescence of free GFP. To avoid this potential artifact, immunofluorescence analysis of endogenous LC3-II is preferred. Due to the hydrophobicity of LC3-II, use of methanol and ethanol to permeabilize cells should be avoided. Since structures of autophagosomes and autolysosomes are sensitive to some detergents, we normally use digitonin to permeabilize cells. 3.1. Cell Culture and Preparation of Cell Lysates for Recognizing LC3-I and LC3-II 1. To investigate autophagic response under starvation conditions, prewarmed KRB buffer is prepared. When lysosomal degradation of LC3-II is suspected, one thousandth of the stock solutions of E64d and pepstatin A are added to the medium to yield final concentrations of 10 μg/mL. As a control, the solvent DMSO is

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added to the medium. The cells, in a 60-mm dish, are incubated at 37ºC for 3–6 h to accumulate LC3-II; in general, LC3-II accumulates in cultured cells 3 h after adding the inhibitors. As a control of nutrient-rich conditions, HEK293 cells are incubated in DMEM containing 10% FCS. The cells are washed twice in PBS, harvested with Teflon cell scrapers, resuspended in ice-cold 200 μL of PBS with CompleteTM protease inhibitor cocktail (Roche Diagnostics), and lysed with an ultrasonic generator for 10 s (0.5-s pulse with 50% duty) on ice. [Option 2b for preparation of cell-lysate without sonication: TX100-lysis solution (final concentration: 2% Triton X-100) is added to washed cells and incubated with mild shaking.] The protein concentration of the lysate is measured using a BCA protein assay kit. Modified Laemmli buffer is added to the samples, and the samples are boiled for 5 min. (For recognizing endogenous LC3, 10–20 μg per lane should be loaded onto SDS-polyacrylamide gels.) The samples are chilled on ice and centrifuged for 10 min at 15,000g.

3.2. SDS-PAGE for Recognizing Endogenous LC3-I and LC3-II 1. Glass plates are prepared for SDS-PAGE (for mini gels, spacer thickness is 1 mm). The inner plates should be cleaned with 70% ethanol. 2. A running gel solution (3.2 mL of distilled water, 4.2 mL of 30% acrylamide solution, 2.5 mL of 1.5 M Tris-HCl, pH 8.8 (at 25ºC), 0.1 mL of 10% SDS, 33 μL of 10% APS, and 5 μL of TEMED) is prepared and poured into the space between the glass plates. 3. The solution is gently overlaid with 150 μL of water-saturated isobutanol, and the gel is allowed to polymerize for 30–60 min at room temperature. 4. A stacking gel solution (2.89 mL of distilled water, 0.79 mL of 30% acrylamide solution, 1.25 mL of 1.5 M Tris-HCl, pH 6.8 [at 25ºC], 0.05 mL of 10% SDS, 17 μL of 10% APS, and 5 μL of TEMED) is prepared. 5. The isobutanol is removed from the polymerized running gel by washing the top with deionized water. Residual water is gently removed with a paper towel. The stacking gel solution is poured into the space between the glass plates, and a comb is inserted. 6. After the stacking gel polymerizes, the comb is removed, and the wells are washed. 7. The gel is mounted into the SDS-PAGE electrophoresis chamber, and running buffer is added. 8. Samples (10–20 μg per lane) are loaded onto the gel. A 5-μL aliquot of prestained molecular weight markers is employed as a standard for molecular weights. 9. The gel is electrophoresed at 1–1.5 mA current constant per cm of gel width, until the pink color of the phenol red in the markers reaches the bottom of the gel. (Good separation between LC3-I and LC3-II occurs when a fraction of the phenol red runs through the gel.)

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3.3. Western Blotting for LC3-I and LC3-II 1. A PVDF membrane is soaked in methanol for 3 min, transferred to Bjerrum and Schafer-Nielsen‘s transfer buffer, and incubated for 5 min. 2. Four sheets of chromatography paper are soaked in Bjerrum and SchaferNielsen‘s transfer buffer. 3. After removing the glass plates and stacking gel, the running gel is rinsed in Bjerrum and Schafer-Nielsen‘s transfer buffer. Two sheets of chromatography paper are placed on the anode plate and overlaid with a PVDF membrane, the running gel, and two additional sheets of chromatography paper. 4. The stack is set onto a Transblot SD Semi-Dry Transfer Cell. 5. The stack is electrophoresed at 16 V for one hour. 6. The gel and papers are discarded, and the PVDF membrane is allowed to dry on a paper towel. 7. Blocking: The PVDF membrane is incubated in Blocking buffer at room temperature for 30 min. 8. The membrane is washed three times in TTBS at room temperature for 2 min each. 9. Primary antibody reaction: The PVDF membrane is incubated in primary antibody solution for 60 min at room temperature. 10. The membrane is washed at least five times, for 5 min each, in TTBS at room temperature. 11. Secondary antibody reaction: The PVDF membrane is incubated in secondary antibody solution for 60 min at room temperature. 12. The membrane is washed at least three times, for 10 min each, in TTBS at room temperature. 13. The reaction of horse radish peroxidase with chemiluminescent reagent is started on the membrane, and the signal is detected on x-ray film or CCD camera (Fig. 1).

3.4. Immunoprecipitation of Endogenous LC3 1. Cells on a 60-mm dish are washed twice in PBS, harvested with Teflon cell scrapers, and resuspended in 100 μL of PBS. (When immunoprecipitating endogenous LC3 from rat (or mouse) tissues, postnuclear supernatant should be prepared.) 2. An equal volume of SDS-lysis solution is added to the cell suspension or postnuclear supernatant. After mixing gently by repeated inverting, the samples are boiled at 100ºC for 5 min and chilled on ice for 3 min. 3. Seven volumes of TX100-solution are added to each sample and mixed thoroughly. Each sample is centrifuged at 10,000g for 15 min at 4ºC, and the pellets are discarded. 4. One mL of TX100-solution is added to 50 μL of protein A–Agarose beads. The suspensions are centrifuged at 500g for 5 min at 4ºC to collect the beads. 5. Each cell suspension supernatant obtained at step 3 is added to 50 μL of equilibrated protein A–Agarose beads.

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6. The suspensions are incubated at 4ºC for one hour with rotating to remove proteins that bind nonspecifically to the beads. The suspensions are centrifuged at 500g for 5 min at 4ºC, and the supernatants are collected. 7. Protein A–Agarose beads are equilibrated as described in step 4. 8. The precleared supernatants are added to the equilibrated protein A–Agarose beads, along with 1 μg of affinity-purified anti-LC3 IgG or, as a negative control, 1 μg of normal rabbit IgG. 9. The mixtures are incubated at 4ºC for 16 h with rotation. 10. The mixtures are centrifuged at 500g for 5 min at 4ºC, and the supernatants are discarded using a 5-mL syringe with 25G × 1” needle (0.5 mm × 25 mm) (Terumo, Tokyo, Japan). 11. The beads are washed at least five times by vortexing in TX100-wash solution. 12. The beads are collected by centrifugation at 500g for 5 min.

Fig. 3. Accumulation of endogenous LC3-positive puncta under starvation conditions in the presence of E64d and pepstatin A in HeLa cells. HeLa cells were transferred to KRB medium for 4 h (starvation). “E64d & pepstatin A” (B and D) indicates that cells were incubated for 4 h with these protease inhibitors, whereas “nontreated” (A and C) indicates addition of a solvent instead of these inhibitors. The cells were fixed and permeabilized with digitonin, and endogenous LC3 was recognized using rabbit antihuman LC3 antibody and goat anti-rabbit IgG conjugated to Q-dot 605. Intracellular fluorescence of endogenous LC3 in HeLa cells was observed. A series of Z-scanned fluorescent images was deconvoluted by a program for two-dimensional blind deconvolution with Aqua C-imaging software (Hamamatsu Photonics). Images C and D are about fourfold magnifications of the images shown in boxes with dotted lines in A and B, respectively. An arrowhead indicates a cup-shaped preautophagosome, and an arrow indicates an autophagosome/autolysosome.

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13. Fifty μl of modified Laemmli buffer is added to the beads, boiled for 5 min, and chilled on ice for one minute. 14. The immunoprecipitates are analyzed as described in Subheadings 3.2. and 3.3. (Fig. 2).

3.5. Immunofluorescence Analysis of Endogenous LC3 1. After washing in TBS, cells are incubated in cell-fix solution at 37ºC for 5 min. 2. The cells are rinsed twice in TBS. 3. Cells are permeabilized in digitonin solution at 37ºC for 5 min. The solution is discarded by aspirating. 4. Excess digitonin is quenched by incubation in quenching solution at 37ºC for 5 min. The solution is discarded by aspirating. 5. The cells are rinsed twice in TBS. 6. Blocking: Cells are incubated in IF-blocking solution at 37ºC for 30 min. 7. The cells are rinsed three times in TBS. 8. Primary antibody reaction: Cells are incubated in IF-primary antibody solution at 37ºC for 60 min. 9. The cells are washed three times in TBS for 5 min. 10. Secondary antibody solution: Cells are incubated in IF secondary antibody solution at 37ºC for 60 min. 11. The cells are washed five times in TBS for 5 min. 12. The cells are mounted on glass slides with mounting medium. 13. Fluorescent images are obtained using a fluorescence microscope (Fig. 3).

4. Notes 1. Earle‘s balanced salt solution (EBSS) can be substituted for KRB for starvation conditions. If using Hanks’ balanced salt solution (HBSS) instead of KRB as a starvation medium, more care should be taken regarding buffering and incubation. HBSS should be buffered by adding 10 mM hydroxyethyl piperazine ethane sulfonate (HEPES). When cells are incubated in HBSS, it may be better to incubate them at 37ºC without control of CO2 . 2. Storage of pepstatin A for over 3 months should be avoided. 3. Polyclonal and monoclonal anti-LC3 antibodies are now available from the Medical & Biological Laboratories, Co. Ltd (http://www.mbl.co.jp/) and Cell Signaling Technology (http://www.cellsignal.com/).

References 1. Eskelinen, E. L. (2005) Maturation of autophagic vacuoles in mammalian cells. Autophagy 1, 1–10. 2. Kadowaki, M., Karim, M. R., Carpi, A., and Miotto, G. (2006) Nutrient control of macroautophagy in mammalian cells. Mol. Aspects Med. 27, 426–443.

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3. Wileman, T. (2006) Aggresomes and autophagy generate sites for virus replication. Science 312, 875–878. 4. Rubinsztein, D. C., Difiglia, M., Heintz, N., et al. (2005) Autophagy and its possible roles in nervous system diseases, damage and repair. Autophagy 1, 11–22. 5. Munz, C. (2006) Autophagy and antigen presentation. Cell Microbiol. 8, 891–898. 6. Deretic, V., Singh, S., Master, S., et al. (2006) Mycobacterium tuberculosis inhibition of phagolysosome biogenesis and autophagy as a host defence mechanism. Cell Microbiol. 8, 719–727. 7. Mann, S. S., and Hammarback, J. A. (1994) Molecular characterization of light chain 3. A microtubule binding subunit of MAP1A and MAP1B. J. Biol. Chem. 269, 11492–11497. 8. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728. 9. Tanida, I., Minematsu-Ikeguchi, N., Ueno, T., and Kominami, E. (2005) Lysosomal turnover, but not a cellular level, of endogenous LC3 is a marker for autophagy. Autophagy 1, 84–91. 10. Kabeya, Y., Mizushima, N., Yamamoto, A., Oshitani-Okamoto, S., Ohsumi, Y., and Yoshimori, T. (2004) LC3, GABARAP and GATE16 localize to autophagosomal membrane depending on form-II formation. J. Cell Sci. 117, 2805–2812. 11. Sou, Y. S., Tanida, I., Komatsu, M., Ueno, T., and Kominami, E. (2006) Phosphatidylserine in addition to phosphatidylethanolamine is an in vitro target of the mammalian Atg8 modifiers, LC3, GABARAP, and GATE-16. J. Biol. Chem. 281, 3017–3024. 12. Tanida, I., Tanida-Miyake, E., Ueno, T., and Kominami, E. (2001) The human homolog of Saccharomyces cerevisiae Apg7p is a protein-activating enzyme for multiple substrates including human Apg12p, GATE-16, GABARAP, and MAPLC3. J. Biol. Chem. 276, 1701–1706. 13. Tanida, I., Tanida-Miyake, E., Komatsu, M., Ueno, T., and Kominami, E. (2002) Human Apg3p/Aut1p homologue is an authentic E2 enzyme for multiple substrates, GATE-16, GABARAP, and MAP-LC3, and facilitates the conjugation of hApg12p to hApg5p. J. Biol. Chem. 277, 13739–13744. 14. Tanida, I., Ueno, T., and Kominami, E. (2004) LC3 conjugation system in mammalian autophagy. Int. J. Biochem. Cell Biol. 36, 2503–2518. 15. Ueno, T., Muno, D., and Kominami, E. (1991) Membrane markers of endoplasmic reticulum preserved in autophagic vacuolar membranes isolated from leupeptinadministered rat liver. J. Biol. Chem. 266, 18995–18999. 16. Ueno, T., Ishidoh, K., Mineki, R., et al. (1999) Autolysosomal membraneassociated betaine homocysteine methyltransferase. Limited degradation fragment of a sequestered cytosolic enzyme monitoring autophagy. J. Biol. Chem. 274, 15222–15229. 17. Tamai, M., Matsumoto, K., Omura, S., Koyama, I., Ozawa, Y., and Hanada, K. (1986) In vitro and in vivo inhibition of cysteine proteinases by EST, a new analog of E-64. J. Pharmacobiodyn. 9, 672–677.

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18. Umezawa, H., Takeuchi, T., Linuma, H., Suzuki, K., and Ito, M. (1970) A new microbial product, oudenone, inhibiting tyrosine hydroxylase. J. Antibiot. (Tokyo) 23, 514–518. 19. Jager, S., Bucci, C., Tanida, I., et al. (2004) Role for Rab7 in maturation of late autophagic vacuoles. J. Cell Sci. 117, 4837–4848.

5 Amino Acid Regulation of Autophagosome Formation Alfred J. Meijer

Summary Amino acids are not only substrates for various metabolic pathways, but can also serve as signaling molecules controlling signal transduction pathways. One of these signaling pathways is mTOR-dependent and is activated by amino acids (leucine in particular) in synergy with insulin. Activation of this pathway inhibits autophagy. Because activation of mTOR-mediated signaling also stimulates protein synthesis, it appears that protein synthesis and autophagic protein degradation are reciprocally controlled by the same signaling pathway. Recent developments indicate that amino acid–stimulated mTOR-dependent signaling is subject to complex regulation. The mechanism by which amino acids stimulate mTORdependent signaling (and other signaling pathways), and its molecular connection with the autophagic machinery, is still unknown.

Key Words: Amino acid signaling; insulin; phosphatidylinositol 3-kinase (PI3K); mammalian target of rapamycin (mTOR); ribosomal protein S6; AMP-activated protein kinase (AMPK); AICAriboside (AICAR); glutamate dehydrogenase; beclin 1; reactive oxygen species.

1. Introduction Cells adapt to changes in their environment by adjusting anabolic and catabolic pathways. In times of nutrient shortage, for example, macromolecules are degraded to produce substrates for energy production. Macroautophagy (“autophagy”) is the major, lysosomal mechanism by which cells degrade protein, in addition to the multienzyme proteasome system. Unlike the proteasome, however, autophagy can also eliminate damaged or redundant organelles. From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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During autophagy, part of the cytoplasm is sequestered by a double membrane, the origin of which is still not firmly established but which is presumably derived from specialized regions of the endoplasmic reticulum (1). Several Atg proteins, first discovered in yeast, are involved in the formation and maturation of autophagosomes (2). Mammalian counterparts of most of the ATG genes have also been found (2). After their formation, the initial autophagosomes acquire their lytic enzymes by fusion with existing or newly formed lysosomes to form autophagolysosomes upon which degradation of the sequestered material can occur. When the cellular nutrient supply becomes insufficient, autophagy is activated. A classical example is that of the mammalian liver, which, in starvation, degrades proteins by autophagy in order to produce amino acids for gluconeogenesis, glucose being required for the brain and the erythrocytes. However, even under nutrient-rich conditions, some ongoing autophagy is still required in order to allow cells to remove defective cell structures. Thus, mice with specific knock-outs of Atg5 or Atg7 in neuronal cells develop neurodegeneration because of defective autophagy (3,4).

2. Control of Autophagy For decades, amino acids have been known as (product) inhibitors of autophagy and they do so by inhibition of autophagosome formation (5), although an effect of some amino acids on autophagosome fusion with lysosomes and on the intralysosomal pH (leucine) (see ref. 6 for literature) cannot be ruled out. In addition, autophagy has long been known to be inhibited by insulin and promoted by glucagon (5,7). The mechanism by which amino acids inhibit autophagosome formation has been obscure for a long time. However, research carried out in our own and other laboratories over the last 10–15 years has indicated that amino acids can control autophagy via changes in the activity of signal transduction pathways. Among these, the mTOR-dependent signal transduction pathway, also used by insulin and other growth factors, appears to play a prominent role (8). However, other signaling pathways, such as the ras/raf/Erk1/2 pathway (9) and the integrin/p38MAPK pathway, may also be affected (10). 2.1. Insulin, Amino Acids, mTOR-Mediated Signaling, and Autophagy Before discussing the stimulation of mTOR-dependent signaling by amino acids, and its relationship with autophagy, a short description of the insulin signaling pathway is given (Fig. 1) (see refs. 11 and 12 for a detailed discussion of the various components in this pathway).

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Fig. 1. Amino acid–activated signaling and the regulation of autophagy. Abbreviations: IRS, insulin receptor substrate; PI3K, phosphatidylinositol 3-kinase class; PDK1, phosphoinositide-dependent kinase-1; PKB, protein kinase B; AMPK, AMP-activated protein kinase; TSC, tuberous sclerosis complex; Rheb, Ras homolog enriched in brain; raptor, regulatory associated protein of mTOR; mTOR, mammalian target of rapamycin; S6K, 70 kD S6 kinase; S6, ribosomal protein S6; MAPK, mitogen-activated protein kinase; Erk, extracellular regulated protein kinase.

The first part of the insulin-dependent signal transduction pathway is located upstream of the protein kinase mTOR and includes the insulin receptor, IRS1 and 2, PI3K class I (producing PtdIns(3,4,5)P3 and PtdIns(3,4)P2 ) and protein kinase B. This part is involved in the the regulation of carbohydrate metabolism (13). The second part of the insulin-signaling pathway, located downstream of mTOR, includes components such as S6K, 4E-BP1, ribosomal protein S6, eIF2-kinase, and eEF-2 kinase, proteins that are involved in the regulation of protein synthesis (12). mTOR activity is controlled by the heterodimer TSC1/TSC2, which acts as a brake on mTOR-dependent signaling. TSC1/TSC2 acts as a GTPase-activating protein complex for the small G-protein Rheb, which, in its GTP form binds to and activates mTOR (12,14). When TSC2 is phosphorylated by protein kinase B, the TSC1/TSC2 complex becomes inactive and mTOR signaling activated.

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mTOR is present in a complex with raptor, a protein that functions as a scaffold for mTOR-mediated phosphorylation of mTOR substrates; the protein GL is also part of this complex (15). In this form, mTOR activity is inhibited by rapamycin. When, however, mTOR is complexed with another protein, rictor, its kinase activity is rapamycin-insensitive and serves to control protein kinase B phosphorylation (15). Early experiments with protein phosphatase inhibitors in isolated hepatocytes had already indicated that protein phosphorylation was involved in the control of autophagy, but the link with amino acids was not made in these studies (16). A breakthrough in the search for mechanisms by which amino acids inhibit autophagy (and stimulate protein synthesis) was obtained in studies with [32 Pi]phosphate-labeled hepatocytes carried out in our laboratory. In these studies, we discovered that the same amino acids that inhibited autophagy, rapidly (within 20 min) and greatly (up to fivefold) stimulated the phosphorylation of a 31 kDa ribosomal protein that we identified as S6 (17,18). Under a variety of experimental conditions, with various amino acid mixtures, in the absence and presence of insulin and/or glucagon, there was a linear relationship between the percentage inhibition of autophagic proteolysis (measured in the presence of low concentrations of cycloheximide to inhibit simultaneous protein synthesis) and the degree of S6 phosphorylation. Insulin and low concentrations of amino acids acted in synergy: insulin inhibited autophagy and stimulated S6 phosphorylation, but only in the presence of low amino acid concentrations, not in the absence of amino acids (when autophagy was maximal) or at high amino acid concentrations (when autophagy was already minimal). Glucagon had the opposite effect: it stimulated autophagy but only in the presence of low amino acid concentrations, not in the absence of amino acids or in the presence of high amino acid concentrations. Among the various amino acids that inhibit autophagy, leucine (but not valine) was particularly effective in inhibiting autophagy and stimulating S6 phosphorylation. Amino acid–induced S6 phosphorylation was completely inhibited by rapamycin, indicating that mTOR and S6K were components of the signaling pathway (Fig. 1). Of great significance was the fact that rapamycin could partly, albeit not completely, reverse the inhibition of autophagy by amino acids. Because the mTOR pathway was known to be involved in the regulation of protein synthesis, we concluded that, apparently, protein synthesis and (autophagic) protein degradation are controlled by the same signaling pathway, which is efficient from the point of view of metabolic regulation (18). The observation that rapamycin increases autophagy has been confirmed for many other cell types (19), including yeast (20,21). Our studies were important for two reasons. First, they were the first to show that amino acids can act as signaling molecules that stimulate a signal


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transduction pathway sharing components with the insulin signaling pathway, with wide ramifications for the regulation of metabolism, not only for protein synthesis and autophagy, but, as we will see later, also for carbohydrate metabolism. Second, they suggested a possible mechanism for the inhibition of autophagy by amino acids. Since our initial observations, the ability of amino acids to stimulate mTORdependent signaling and the synergy between insulin and amino acids have been confirmed (or rediscovered) for many other insulin-sensitive cell types, with leucine being the most effective amino acid (see refs. 8 and 15 for reviews). Likewise, the inhibition of mTOR signaling by glucagon has been confirmed (22,23). As indicated above, an important feature of amino acid signaling in hepatocytes is that high concentrations of amino acids alone are sufficient to stimulate mTOR-dependent signaling even when insulin is not present (8,18). Conversely, the effect of insulin on mTOR-dependent signaling is potentiated by the presence of low concentrations of amino acids and is absent when amino acids are severely depleted. In contrast to insulin, amino acids do not activate protein kinase B and probably also not PI3K class I (see ref. 8 for review). Because amino acid–stimulated mTOR-dependent signaling is prevented by inhibitors of PI3K, as was first shown by our laboratory (24) and later by others (8,15), it is likely that mTOR receives two parallel inputs, one via PI3K class I / PDK1 / PKB / TSC1,2 / Rheb and one via amino acids (see Fig. 1). Thus, both PI3K class I (by insulin) and mTOR (by amino acids) must be activated to ensure full activation of mTOR downstream targets. Because high concentrations of amino acids are sufficient to stimulate mTOR signaling maximally, one has to assume that basal activity of PI3K class I is sufficient for mTOR-dependent signaling under these conditions (25). Recent information suggests that amino acids may not stimulate PI3K class I but rather PI3K class III, which produces PtdIns(3)P that directly stimulates mTOR without the requirement for protein kinase B (15,26,27) (see also below). Another development which was important for a better understanding of the regulation of autophagy was the finding that interruption of signaling by inhibitors of PI3K class I (wortmannin, LY294002), in contrast to that caused by rapamycin, did not stimulate but rather inhibited autophagy (24). In order to account for this unexpected result we hypothesized that PtdIns(3)P, the product of PI3K class III, is essential for autophagosome formation whereas PtdIns(3,4)P2 and PtdIns(3,4,5)P3 , the products of PI3K class I, are inhibitory and that the PI3K inhibitors are not entirely specific and inhibit both PI3K class I and III (6) (see Fig. 1); the basis of this hypothesis was the situation in yeast cells which are very active in autophagy under nutrientdeprived conditions but only contain a PI3K class III analog, Vps34, but

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not a PI3K class I. The hypothesis was tested in HT-29 cells and appeared to be correct: PtdIns(3)P appeared to be essential for autophagy, whereas PtdIns(3,4)P2 and PtdIns(3,4,5)P3 were inhibitory (28). In addition, overexpression of PTEN, which hydrolyzes PtdIns(3,4,5)P3 to PtdIns(4,5)P2 and PtdIns(3,4)P2 to PtdIns(4)P, stimulated autophagy in human colon cancer HT29 cells (29). 3-Methyladenine, the classical inhibitor of autophagy (30), turned out to be a PI3K inhibitor; inhibition of PtdIns(3)P formation provided a satisfatory explanation for its mechanism of action (24,28). Interestingly, Beclin1, the mammalian homolog of Atg6, which is involved in autophagosome formation (31), is found in a complex with PI3K class III (32,33). In order to be able to bind to PI3K class III and to stimulate autophagy, Beclin 1, which is found in association with the anti-apoptotic protein Bcl-2, must first dissociate from the inhibitory Beclin 1-Bcl-2 complex (34). In addition to their ability to inhibit autophagy via mTOR activation, amino acids also inhibit autophagy by decreasing Beclin1-associated PI3K class III activity (35), because of increased binding of Beclin 1 by Bcl-2 (34). Presumably, this accounts for the observation that inhibition of autophagy by amino acids is rapamycin-insensitive under some conditions (36–38). At variance with a requirement of autophagy for the Beclin 1-PI3K class III complex are recent studies showing that amino acids stimulate mTOR signaling via activation of PI3K class III and that amino acid depletion results in a decrease in the activity, but not the amount, of Beclin 1-associated class III PI3K (15,26,27). In these studies, autophagy was not investigated, however. A possible explanation is that there are different pools of PI3K class III, differentially regulating autophagy and mTOR signaling (15,26,27). Perhaps an excess of free PI3K class III, not bound to Beclin 1 (or bound to Bcl-2, see previous paragraph), signals to mTOR and inhibits the stimulation of autophagy by the Beclin 1-PI3K class III complex (see Fig. 1). According to Hinault et al. (39), amino acids are able to stimulate the insulinsignaling pathway in yet another manner. These authors propose that amino acids can stimulate protein kinase B in a PI3K-independent manner because they stimulated protein kinase B phosphorylation in the presence of insulin and wortmannin (39). It must be pointed out, however, that amino acids can react chemically with wortmannin in a nonenzymic fashion and thus relieve the inhibition of PI3K (40). Although the molecular connection between mTOR and the machinery required for autophagy is not yet firmly established, experiments with yeast have indicated that the Atg1–Atg13 complex, which is required for autophagosome formation, is one of the targets. Starvation of yeast cells, or rapamycin treatment, dephosphorylates Atg1 and enhances its protein kinase activity; Atg13, which binds to and activates Atg1, is hyperphosphorylated under


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nutrient-rich conditions in a Tor-dependent manner, reducing its affinity for Atg1 (41). Recent evidence obtained with yeast indicates that the protein kinase activity of Atg1 is essential for autophagosome formation (42). 2.2. mTOR, AMP-Activated Protein Kinase, and Autophagy In addition to its ability to sense amino acids, mTOR may also act as a sensor of the cellular energy state. Originally it was thought that mTOR senses the intracellular ATP concentration because of its relatively low affinity for ATP (43). However, later simultaneous observations in several laboratories, including our own, have indicated that small decreases in ATP result in activation of AMPK which, in turn, inhibits mTOR (44) and inhibits protein synthesis. This is in agreement with the function of AMPK to shut off ATP-dependent processes (45). AMPK possibly inhibits mTOR signaling by phosphorylating TSC2 (46). In myotubes, amino acids stimulate phosphorylation of mTOR at Ser2448 . In the absence of amino acids, however, or by activation of AMPK, Thr2446 is phoshorylated. The close proximity of these two phosphorylation sites makes them mutually exclusive in that phosphorylation of one site inhibits that of the other and vice versa; they are viewed as switches integrating the opposing signals of growth factors and nutrient deprivation (47). Inhibition of mTOR by AMPK, like that caused by addition of rapamycin may be expected to increase autophagy (Fig. 1). However, there is controversy in the literature on this issue. In yeast, activation of AMPK, indeed, stimulates autophagy (48). By contrast, activation of AMPK by addition of the cellpermeable nucleotide analogue AICAriboside (AICAR) in hepatocytes strongly inhibits autophagy (49). Recent evidence, however, indicates that also in mammalian cells, AMPK is required for autophagy. Thus, apoptotic stimuli, which result in increased mitochondrial permeability and decreased mitochondral membrane potential, target these mitochondria for autophagic degradation (50,51); it is likely that AMPK under these conditions is activated. Inhibition of mitochondrial ATP synthesis with oligomycin in insect cells has been shown to promote massive autophagy, of mitochondria in particular (52). In mouse embryonic fibroblasts, activation of the tumor suppressor p53 inhibits mTOR activity through activation of AMPK, a phenomenon that is accompanied by increased autophagy (53,54). Eukaryotic elongation factor 2-kinase (eEF-2 kinase) is essential for autophagy (55) and is known to be activated by AMPK (56). In a study recently carried out with different mammalian cell types, we obtained direct evidence that AMPK activation, as in yeast, promotes autophagy; we also concluded that the inhibition of autophagy by AICAR is unrelated to its ability to activate AMPK (57).

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2.3. Mechanism of Amino Acid Sensing in mTOR-Mediated Signaling Although amino acids can activate mTOR, the mechanism by which this occurs is unknown. Possible mechanisms are discussed below. 2.3.1. Amino Acid Receptor: Intra- or Extracellular? Most evidence indicates that the amino acid receptor is intracellular rather than extracellular (58,59). It has been proposed that the plasma membrane contains a leucine-specific receptor protein that controls autophagy independent of mTOR (37); this receptor protein cannot be the leucine transporter as proposed (15), because both valine and isoleucine, which are transported via the same transporter, neither affect autophagy (6) nor mTOR-mediated signaling (60,61). It cannot be excluded that both intra- and extracellular sensing mechanisms operate in parallel or that their relative activity is cell-type–dependent. One mechanism for intracellular amino acid sensing is that amino acids inhibit TSC1/2. Recent evidence indicates that at least TSC2 is not involved (62). Because AMPK activation inhibits mTOR signaling by phosphorylating TSC2 (see Subheading 2.2.), the possibility that amino acids stimulate mTOR by inhibition of AMPK can also be ruled out. Another recent study has indicated that amino acids promote the association of Rheb with mTOR, an effect that is probably not caused by increased charging of Rheb with GTP (63) (contrast ref. 64, where an increase in Rheb-GDP upon nutrient depletion was found) but rather by direct action on mTOR (65). Although it is tempting to conclude that mTOR itself is the amino acid sensor, it is still possible that the effect of amino acids is indirect and that they decrease the concentration of an inhibitor that interferes with the Rheb-mTOR association (65). As indicated in Subheading 2.1., the possibility that amino acids are able to stimulate mTOR signaling via activation of PI3K class III, at least under some conditions, cannot be ruled out. But how amino acids exert this effect remains to be elucidated. 2.3.2. Amino Acid Sensing via Glutamate Dehydrogenase? In most cell types, leucine (but not the other branched-chain amino acids) is one of the most potent amino acids in stimulating mTOR-dependent signaling (8); metabolism of leucine is not required (66). In addition, nonmetabolizable analogues of leucine can mimic its effect (60,61). Leucine is also very potent in promoting the association between mTOR and Rheb (65). Interestingly, in pancreatic -cells the specificity of leucine (e.g., valine and isoleucine are not effective) and its analogs in stimulating mTOR signaling is strikingly similar


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to the ability of these amino acids to stimulate glutamate dehydrogenase, the enzyme which is thought to play an important role in insulin production in -cells (67). A possible connection between glutamate dydrogenase and mTOR signaling may be the production of GTP via oxidation of -oxoglutarate, formed by glutamate dehydrogenase, for GTP charging of Rheb. It must be pointed out, however, that this GTP is produced within the mitochondria and somehow must be transported to the cytosol, a function that cannot be carried out by the mitochondrial adenine nucleotide transporter (68,69). Because nucleosidediphosphate kinase is found both within the mitochondria (70) and at the mitochondrial outer membrane (71), the exit of GTP from the mitochondria may be indirect via transfer of its terminal phosphate to ADP within the mitochondria and channeled back to GDP at the mitochondrial outer membrane. As indicated above, there is some controversy on whether (64) or not (63) the charging of Rheb with GTP is increased by amino acids. In principle, this would be an attractive hypothesis, especially because in cancer cells glutamine is an important substrate for energy production (72) so that flux through glutamate dehydrogenase may be high, mTOR signaling is very active and autophagy suppressed in cancer cells (19). Another hypothesis is that NADPH produced by glutamate dehydrogenase in the direction of deamination, via the glutathione redox system, is engaged in the scavenging of reactive oxygen species which may be involved in the initiation of autophagy, at least under some conditions (73,74). 2.3.3. Diadenosine Polyphosphates An intriguing possibility, which has not been mentioned in the literature so far, is that diadenosine polyphosphates (ApnA), byproducts of the aminoacyltRNA synthetase reaction, are involved in mTOR stimulation. Indirect evidence comes from two independent studies: in one study, leucine stimulated Ap4A production by the mitochondria in pancreatic ß-cells (75), while in another study leucine stimulated mTOR activity in ß-cells through increased mitochondrial oxidative metabolism (67), as discussed in the previous section. Because mTOR may be associated, at least in part, with mitochondria (76,77), it is tempting to speculate that Ap4A is another possible signal that connects mitochondrial metabolism to mTOR activity. In this context, it is important to note that Ap4A is a strong inhibitor of AMP-activated protein kinase (78). 2.3.4. eIF2 Another mechanism of amino acid sensing is one in which cells respond to changes in the charging of tRNAs. This mechanism is based on data in yeast showing that, upon amino acid starvation, free uncharged tRNA binds

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to the protein kinase Gcn2 because its active center strongly resembles that of aminoacyl-tRNA synthetases (79). Gcn2 activation results in the phosphorylation of eIF2, which then derepresses GCN4 mRNA translation. Gcn4 is a transcriptional activator that promotes the transcription of many genes involved in nitrogen metabolism; these include not only genes involved in the biosynthesis of amino acids but also genes required for autophagy (80). The activity of Gcn2 itself becomes inhibited by TOR-dependent phosphorylation of Ser577 of Gcn2 (81). Because rapamycin is still able to stimulate autophagy in GCN2disrupted yeast (82), it is possible that Gcn2 may not be downstream of Tor, or that Gcn2 controls autophagy by a mechanism that is independent of Tor. In mammalian cells, the eIF2 kinase PKR (double-stranded RNA-dependent protein kinase), which is the equivalent of Gcn2, contributes to the control of autophagy, and PKR can rescue starvation-induced autophagy in GCN2disrupted yeast (82). Unexpectedly, experiments with aminoalcohols, inhibitors of aminoacyltRNA synthetases, have yielded contradictory results in mammalian cells with regard to their effects on both amino acid signaling and autophagy, presumably because of lack of specificity of these compounds (8,15,59). Furthermore, amino acid deprivation did not affect free-tRNA levels, at least in HEK-293 cells (43). 2.3.5. Phosphatidic Acid The mitogenic messenger phosphatidic acid can also activate mTORdependent signaling provided sufficient amino acids are also present, indicating that phosphatidic acid promotes signaling in parallel to amino acids (83,84). This is similar to the effect of insulin, as discussed earlier (Subheading 2.1.). Whether insulin can affect phosphatidic acid concentrations or whether phosphatidic acid affects autophagy is not known. 2.4. Feedback Interaction in the mTOR Signaling Pathway and Autophagy Overactivation of amino acid–dependent mTOR-mediated signaling can lead to the inhibition of the proximal part of the insulin signaling pathway (85–87) (Fig. 1). This is because of phosphorylation of IRS1 by S6K, which results in decreased binding of the p85 regulatory subunit of PI3K class I to IRS1. Because class I PI3K is required for mTOR downstream signaling, this feedback system may be part of a homeostatic mechanism that is required to prevent the overactivation of mTOR by amino acids. It has been proposed that the overactivation of mTOR contributes to insulin resistance in obesitylinked diabetics (86). Downregulation of proximal insulin signaling can be expected to have consequences for autophagy, because a decline in inhibitory


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PtdIns(3,4,5)P3 will accelerate autophagy. Surprisingly, very little is known about protein turnover in type 2 diabetes. It is likely, however, that it is, indeed, increased in type 2 diabetes (88,89). Whether autophagy contributes to the increased protein turnover in type 2 diabetes is entirely unknown. The possible consequences of feedback interaction of S6K on mTOR upstream signaling for autophagy were discussed by us (90) in connection with a study by Scott et al. (91) concerning the regulation of autophagy in the fat body of Drosophila melanogaster. In this study, the authors convincingly show that S6K is not inhibitory, as previously suggested (18) (see Subheading 2.1.) but is, in fact, essential for autophagy. Thus, in this insect, the same protein kinase (S6K) that is required for protein synthesis, an anabolic process, is also essential for autophagy, a catabolic process (91). A possible explanation for this paradox may be found in the negative-feedback effect of S6K on signaling upstream of Tor, which would then increase autophagy as a result of the fall in class I PI3K activity (90). This may be important, because even under nutrientrich conditions cells must be able to carry out some autophagic activity, not in order to produce nutrients, but in order to eliminate damaged cell structures or structures that are no longer needed by the cell. Conversely, when nutrients become scarce, e.g., during starvation, the inactivation of Tor by a fall in amino acid concentration accelerates autophagy, provided sufficient active S6K is still present. During long-term starvation, S6K activity may fall so low that class I PI3K is activated again and this would then restrain excessive autophagy in order to prevent cell death, as suggested (91). Autophagy has been implicated as a protective mechanism in various neurodegenerative diseases because the process contributes to the removal of defective proteins (2). In Huntington’s disease, for example, expanded polyglutamine proteins accumulate abnormally in intracellular aggregates which in cell models can be prevented by induction of autophagy with rapamycin (92). Unexpectedly, however, in a functional genetic screen of Huntington’s disease it was found that activation of IRS2, which mediates insulin signaling (Fig. 1), results in autophagy-mediated clearance of the polyglutamine protein aggregates (93). The activation of autophagy occurred in spite of activation of protein kinase B, mTOR, and S6K, but was still PI3K class III- and Beclin1– dependent. Perhaps the negative feedback in proximal insulin signaling by S6K, as discussed in the previous paragraph, accounts for these surprising results. 2.5. Other Amino Acid–Sensitive Signaling Mechanisms Controlling Autophagy (see Fig. 1) In human colon cancer HT-29 cells, another amino acid–dependent signaling pathway can control autophagy, in addition to the PI3K/mTOR pathway.

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Activation of Erk1/2 stimulates the GTPase-activating protein GAIP and abolishes the inhibitory effect of trimeric Gi3 protein on autophagy (94). Amino acids, by stimulating the phosphorylation of Ser259 , inactivate the Erk1/2 MAPK kinase Raf-1 and downregulate autophagy (9). By contrast, in C2C12 myotubes the inhibition of autophagy by amino acids is not accompanied by any changes in Erk1/2 phosphorylation (35). Differences in amino acid signaling mechanisms and in the control of autophagy may exist, apparently depending on the cell type and perhaps also on the degree of differentiation (35). Ceramide has been shown to activate autophagy by upregulation of Beclin 1 and inhibition of protein kinase B (95). Ceramide also decreases intracellular amino acid concentrations by inhibition of amino acid transport, resulting in decreased mTOR-dependent signaling (96). Although the link with autophagy was not studied in these studies (96), inhibition of amino acid transport may also contribute to the activation of autophagy by ceramide. In cultured rat hepatocytes and in the flow-through perfused rat liver, amino acid–induced cell swelling, caused by Na+ -dependent concentrative transport of certain amino acids, inhibits autophagy independent of mTOR by activation of p38MAPK (97); likewise, in yeast the p38MAPK orthologue Hog1 also plays an important role in the osmosensitivity of autophagy (21). Integrins and microtubules are part of the osmo-sensing mechanism (10,98). Insulin inhibits autophagy because it also increases cell volume and activates p38MAPK by an integrin-dependent, mTOR-independent mechanism (38,99). By contrast, in C2C12 myotubes, p38MAPK is not involved in the inhibition of autophagy by amino acids (35), whereas in skeletal muscle and in adipocytes insulin also does not affect p38MAPK (100). In contrast to the flow-through perfused rat liver, in the isolated circulatory perfused rat liver, neither amino acids that increase cell volume (glutamine, alanine) nor insulin affect autophagy on their own, unless they are added in combination with other amino acids (e.g., leucine) (101). In isolated hepatocytes, an increase in cell volume alone is also not sufficient to inhibit autophagy, although it does potentiate the inhibitory effect of low concentrations of leucine and other amino acids on autophagy by a mechanism that is mTOR-dependent (18,102). Again, depending on the experimental conditions and on the cell type, different mechanisms appear to operate in the control of autophagy. 3. Concluding Remarks Although it is now generally accepted that amino acids inhibit autophagy by activation of mTOR-dependent signaling in concert with insulin, very little is known about the mechanisms involved and much work needs to be done


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in this exciting area. Among the most compelling questions are the nature of the primary amino acid receptor and the nature of the mechanisms coupling the amino acid signaling pathway to autophagosome formation. If the amino acid receptor turns out to be intracellular rather than extracellular, amino acid metabolism, in addition to plasma membrane amino acid transport, will directly affect intracellular amino acid concentrations, thus signaling and autophagy. Because amino acid metabolism is strongly cell type–dependent, this may also be true for the regulation of amino acid signaling and of autophagy. Recent reviews on autophagy have stressed the role of autophagy in processes such as cancer and aging (103–108). An intriguing question here is whether changes in amino acid signaling, in addition to alterations in the expression of Atg proteins, contribute to the variations in autophagy. As discussed earlier (see Subheading 2.4.), we predict that in type 2 diabetes, which is characterized by insulin resistance, autophagy is activated, possibly as a protective mechanism to eliminate damaged organelles, e.g., mitochondria (103). In this context it is important to note that both caloric restriction (109) and defective insulin signaling (110,111) extend life span; this may be ascribed, at least in part, to increased autophagy (109,112). It is worthwile mentioning that the Klotho protein, which has strong antiaging properties in mammals, confers insulin resistance (113). In relation to this, SIRT4, a mammalian homologue of Sir2, which is an NAD-dependent deacetylase that promotes longevity in yeast, flies, and worms, is a mitochondrial enzyme that ADP ribosylates and downregulates glutamate dehydrogenase, inhibits insulin production in ß-cells (114) (see Subheading 2.3.2.). Increased mitochondrial production of reactive oxygen species, with loss of mitochondrial function, has been implicated not only in the aging process (109), but also in type 2 diabetes (115); elimination of damaged mitochondria will be of vital importance to prolong life. It is tempting to speculate that diminished amino acid signaling contributes to the autophagic protection under these conditions.

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6 Autophagic Proteolysis of Long-Lived Proteins in Nonliver Cells Esteban A. Roberts and Vojo Deretic

Summary Autophagy is a cellular homeostasis pathway used to sustain cellular anabolic needs during times of nutrient or energy deprivation. Autophagosomes sequester cytoplasmic constituents, including macromolecules such as long-lived proteins. Upon fusion of autophagosomes with lysosomes, the engulfed cargo is degraded. The proteolysis of longlived proteins by macroautophagy is a standard, specific measure of autophagic degradation and represents an end-point assay for the pathway. The assay is based on a pulse-chase approach, whereby cellular proteins are radiolabeled by an isotopically marked amino acid, the short-lived, rapidly turned over, proteins are allowed to be degraded during a long chase period, and then the remaining, stable radiolabeled proteins are subjected to autophagic degradation. The classical application of this method has been in hepatocytes, but the recent growth of interest in autophagy has necessitated adaptation of this method in nonliver cells. Here we describe a protocol to quantify autophagic degradation of longlived proteins in macrophages. This chapter details the method of analyzing autophagic proteolysis in RAW264.7 mouse macrophages.

Key Words: Autophagy; proteolysis; long-lived proteins; macrophage. 1. Introduction Autophagy is an intracellular bulk degradation mechanism beginning with the engulfment of cytoplasmic constituents including proteins into doublemembrane autophagosomes and ending with fusion of autophagosomes with lysosomes for final degradation of the sequestered macromolecules (1–3). Fusion of autophagosomes with lysosomes to form autolysosomes is important for delivery of lysosomal hydrolases and completion of the autophagic From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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pathway via degradation of cytoplasmic components. Among the cytoplasmic constituents targeted for autophagolysosome degradation are stable, long-lived proteins within the cell. A simple radiolabeled amino acid–based pulse chase assay can be used to assess the function of the autophagic pathway in cells by quantitatively monitoring the turnover of long-lived proteins during induction of autophagy or during its inhibition (2–4). This assay was traditionally used to examine autophagy in hepatocytes, since liver cells are robust in their autophagic activity (3,4). Since autophagy is now studied in many different cell types, it is necessary to adapt and optimize original methods developed for liver cells (2–4) to other individual cell types. This chapter describes the methodologies used to examine the autophagic proteolysis of long-lived proteins in nonhepatic cell types and details the quantification of autophagic proteolysis in macrophages. The majority of analyses of proteolysis under stimulation of autophagy have utilized a variation of the method developed by Codogno and colleagues in 1996 (5) when they examined the role of Gi3 protein in autophagy in HT29 cells (a human colon carcinoma cell line). This method uses l-(14 C)valine for the radiolabeled incorporation of amino acid into de novo synthesized proteins (pulse). The radioisotope is subsequently chased by incubating cells in media containing cold valine to allow the degradation of short-lived proteins (chase). It is only after this period of short-lived protein degradation that the autophagic induction of long-lived protein degradation can be specifically examined. Upon a period of autophagic induction, the analysis becomes a simple calculation of the ratio of soluble radioactivity present in the culture supernatant to total cellular radioactivity that reflects the rate of stable protein degradation (5,6). Several derivatives of this method have been used depending on the experimental situation and have included using higher concentrations of radioisotope in mouse embryonic fibroblasts (MEFs) and HeLa cells (7,8), different methods of cellular lysis (MEFs and HeLa) (7,8), and various degradation measurements such as percent degradation over a time course rather than single time point measurement as performed in MEFs and human fibroblasts (9,10). One deviation from the method of Ogier-Denis (5) has been the use of tritiated leucine as the amino acid of choice for examining the autophagic proteolysis of long-lived proteins. This is probably a simple issue of preference and availability rather than advantage since it has been shown that, at least in human fibroblasts, l-(3 H)leucine- or l-(3 H)valine-labeled cells lose protein radioactivity at a similar rate, making these amino acids equally useful in evaluating the degradation of stable proteins (9). Recently, autophagy has been linked to innate immune defense against several pathogens ranging from RNA viruses, like poliovirus (11), grampositive bacteria, like Streptococcus (12), gram-negative bacteria, like Shigella

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(13), and even M. tuberculosis in infected macrophages (14,15). Autophagy has been implicated in antigen presentation, which can occur in various cell types, but is best achieved by macrophages and dendritic cells (16). Because of this, quantification of autophagic processes is of interest in antigenpresenting cells in general and specifically in macrophages. Macrophages represent a challenging experimental system while analyzing autophagy because they are robust in phagolysosome biogenesis, free-radical production, and cytokine secretion and signaling. Therefore, establishing reproducible techniques to evaluate autophagic processes in macrophages is of paramount importance to enable researchers to probe the role of autophagy in innate and adaptive immunity. Since the autophagic proteolysis of long-lived proteins is a gold standard method to examine autophagy, we have optimized this assay for autophagic proteolysis in macrophages (Fig. 1). The following methods describe a protocol for examining the autophagic proteolysis of long-lived proteins in the easily transfectable mouse macrophage cell line RAW264.7.

% 3[H] Leucine Release

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Fig. 1. Proteolysis of long-lived proteins in macrophages. RAW264.7 macrophages were examined for autophagic degradation of long-lived proteins using Earle’s balanced salt solution (Starv). Starvation enhances proteolysis twofold (*p < 0.05) over that in complete DMEM (Control). Classical inhibitors of autophagy were examined for their efficacy in preventing proteolysis (Wm = wortmannin [100 nM]; 3MA = 3methyladenine [10 mM]; Noc = nocodazole [20 μM]; NH4 Cl = ammonium chloride [10 mM]; Vin = vinblastine [50 μM]). All inhibitors exerted a significant reduction in levels of autophagic proteolysis when compared to starvation. Individual statistics showed that not all inhibitors reduced proteolysis to control levels (*p < 0.05; † p > 0.05).

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2. Materials 2.1. Cell Culture, Media, and Reagents 1. RAW264.7 cells (ATCC TIB-71) are grown in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco/BRL, Bethesda, MD) supplemented with 10% fetal bovine serum (FBS; Hyclone, Ogden, UT) and 4 mM l-glutamine (BioWhittaker, Walkersville, MD) (complete DMEM). 2. Leucine-free DMEM (Chemicon International, Temecula, CA) created using the formulation from Gibco/BRL (Bethesda, MD). 3. 100% trichloroacetic acid. 4. 10% bovine serum albumen (BSA) in 1X phosphate-buffered saline (PBS) pH 7.2. 5. lysis buffer: 10 mM Tris HCl (pH 8.0), 150 mM NaCl, , 2 mM EDTA, 0.5% deoxycholate, 2% NP-40 in dH2 O (Note 1). 6. “cold” l-leucine (Sigma, St. Louis, MO). 7. Autophagy reagents: Earle’s Balanced Salt Solution (EBSS), wortmannin (Sigma), 3-methyladenine (Sigma), Nocodazole (Sigma), vinblastine sulfate salt (Sigma). 8. 1.7-mL Ultraclear graduated microcentrifuge tubes (GENEMate, Kaysville, UT) (Note 2).

2.2. Radioactive Reagents and Supplies 1. Radioactivity: 1 μCi/mL l-(4,5-3 H)leucine (73.0 Ci/mmol) (Amersham, Waltham, MA). 2. Scintillation fluid: Optiphase HiSafe 3 Scintillation cocktail (PerkinElmer, Waltham, MA). 3. Scintillation vials: Borosilicate glass vial with urea cap (22mm) with foil liner (Wheaton, Millville, NJ). 4. Packard Tri-Carb 2100TR liquid scintillation analyzer, (PerkinElmer, Waltham, MA).

3. Methods 3.1. Pharmacological Inhibitor Preparation 1. Wortmannin (FW = 484.4 g/mol): use at 100 nM final. a. Dilute 1mg (supplied in a single vial) in 2.064 mL 100% DMSO to yield 1 mM stock. b. Perform a 1:10 dilution in 1X PBS to yield 100 μM stock (working solution). c. Dilute 1:1000 (1 μL in 1 mL) for final use = 100 nM final. 2. 3-Methyladenine (FW = 149.16 g/mol): use at 10 mM final. a. Prepare 100 mM stock by dissolving 0.014916 g in 1 mL of dH2 O. Heat between 60 and 80°C/10 min in a heating block with cap-lock. b. Add 100 μL of 100 mM stock per 1 mL of media to achieve 10 mM final.


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3. Nocodazole (FW = 301.32 g/mol): use at 20 μM final. a. Dissolve 2 mg (supplied in a single vial) in 0.332 mL 100% DMSO to yield 20 mM stock. b. Dilute 1:1000 (1 μL in 1 mL) for final use = 20 μM final. 4. Vinblastine sulfate salt (FW = 909.05 g/mol): use at 50 μM final. a. Dissolve 1 mg (supplied in a single vial) in 0.044 mL 100% DMSO to yield 25 mM stock. b. Dilute 1:500 (50 μL in 1 mL) for final use = 50 μM. 5. Ammonium chloride (FW = 53.49 g/mol): use 10 mM final. a. Dissolve 0.5349 g in 10 mL dH2 O to achieve a 1 M stock. b. Dilute 1:100 (10 μL in 1 mL) for final use = 10 mM.

3.2. Proteolysis Assay 1. Scrape a 2-day-old subculture of RAW macrophages (70–80% confluency) into 12 mL complete DMEM and count cells (10 μL) using trypan blue staining (5 μL) in a hemocytometer. 2. Dilute cells to 7 × 104 /mL in complete DMEM and dispense 1 mL per well into 12-well plate. Incubate at 37°C, 5% CO2 overnight. 3. Remove media and place 1 mL DMEM with 1 μCi/mL 3 H-leucine adjusted to 62.4 μM final onto cells and incubate overnight at 37°C, 5% CO2 . 4. Remove tritiated media into radioactive liquid waste container and wash once with 0.5 mL/well cold DMEM containing 62.4 μM l-leucine (adjusted DMEM). Dispose wash in radioactive liquid waste container. 5. Incubate macrophages in 1 mL/well adjusted DMEM overnight (typically 16 h) at 37°C, 5% CO2 to chase out short-lived proteins. 6. Remove media from wells and dispose in radioactive liquid waste container. Wash wells destined for starvation once with 1 mL 37°C EBSS and add 0.5 mL EBSS/well. Incubate at 37°C/10 min. Repeat two times and add 0.5 mL EBSS (or EBSS + inhibitor)/well (Note 3). Wash other wells with 1 mL adjusted DMEM and add 0.5 mL DMEM for control. Incubate samples at 37°C, 5% CO2 for 4 h. 7. Remove sample media to a microcentrifuge tube (see Note 2). Add 50 μL of fresh 10% BSA in 1X PBS followed by 100 μL of room temperature 100% trichloroacetic acid to each tube and incubate on ice at least 10 min. 8. Spin samples at 5,500× gh/10 min at room temperature. The supernatant represents the soluble radioactivity from samples, which contain released tritiated leucine. 9. Add 0.5 mL of lysis buffer to cells remaining in the wells and place on shaker 10 min at room temperature to aid in lysis. Pipet several times to ensure total cell lysis. These samples represent the total cellular radioactivity.

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10. Aliquot soluble radioactivity samples and total cellular radioactivity samples into scintillation vials containing 3–5 mL of scintillation fluid and read in a scintillation counter using 1 min read times. The percent leucine release is calculated by dividing the soluble radioactivity cpm by the total cellular radioactivity sample cpm.

4. Notes 1. Lysis buffer: Lysis buffer can be substituted by 0.1M NaOH or cold, sterile dH2 O (hypotonic lysis). 2. Microcentrifuge tubes: GENEMate cat. no. C-3218-1 Ultraclear microcentrifuge tubes resist leakage, preventing radioactive contamination. 3. Repeated washes for starvation: Repeated 10-min washes with EBSS are necessary to activate the starvation response in RAW264.7 macrophages. This may be the result of, besides the simple carryover of amino acids, a sufficient efflux of free amino acids that will then repress autophagic signal induction. A similar technique was used to enhance amino acid starvation in MCF7 and HeLa cells using hourly replacement of starvation media to discard secreted amino acids (17).

Acknowledgments This work was supported by the New York Community Trust’s Heiser Program for Research in Tuberculosis and Leprosy Postdoctoral Fellowship to E.A.R. and by grants AI45148 and AI069345 from the National Institutes of Health.

References 1. Mizushima, N. (2004) Methods for monitoring autophagy. Int. J. Biochem. Cell Biol. 36(12), 2491–2502. 2. Seglen, P. O. (1983) Inhibitors of lysosomal function. Methods Enzymol. 96, 737–764. 3. Seglen, P. O. and Gordon, P. B. (1982) 3-Methyladenine: specific inhibitor of autophagic/lysosomal protein degradation in isolated rat hepatocytes. Proc. Natl. Acad. Sci. USA 79(6), 1889–1892. 4. Blommaart, E. F., et al. (1997) The phosphatidylinositol 3-kinase inhibitors wortmannin and LY294002 inhibit autophagy in isolated rat hepatocytes. Eur. J. Biochem. 243(1–2), 240–246. 5. Ogier-Denis, E., et al. (1996) Guanine nucleotide exchange on heterotrimeric Gi3 protein controls autophagic sequestration in HT-29 cells. J. Biol. Chem. 271(45), 28593–28600. 6. Gronostajski, R. M. and Pardee, A. B. (1984) Protein degradation in 3T3 cells and tumorigenic transformed 3T3 cells. J. Cell Physiol. 119(1), 127–132.


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7. Mizushima, N., et al. (2001) Dissection of autophagosome formation using Apg5deficient mouse embryonic stem cells. J. Cell Biol. 152(4), 657–668. 8. Nara, A., et al. (2002) SKD1 AAA ATPase-dependent endosomal transport is involved in autolysosome formation. Cell Struct. Funct. 2002. 27(1), 29–37. 9. Fuertes, G., et al. (2003) Changes in the proteolytic activities of proteasomes and lysosomes in human fibroblasts produced by serum withdrawal, amino-acid deprivation and confluent conditions. Biochem. J. 375(Pt 1), 75–86. 10. Talloczy, Z., et al. (2002) Regulation of starvation- and virus-induced autophagy by the eIF2alpha kinase signaling pathway. Proc. Natl. Acad. Sci. USA 99(1), 190–195. 11. Suhy, D.A., Giddings, T. H., Jr., and Kirkegaard, K. (2000) Remodeling the endoplasmic reticulum by poliovirus infection and by individual viral proteins: an autophagy-like origin for virus-induced vesicles. J. Virol. 74(19), 8953–8965. 12. Nakagawa, I., et al. (2004) Autophagy defends cells against invading group A Streptococcus. Science 306(5698), 1037–1040. 13. Ogawa, M., et al. (2005)Escape of intracellular Shigella from autophagy. Science 307(5710), 727–731. 14. Gutierrez, M.G., et al. (2004) Autophagy is a defense mechanism inhibiting BCG and Mycobacterium tuberculosis survival in infected macrophages. Cell 119(6), 753–766. 15. Singh, S.B., et al. (2006) Human IRGM induces autophagy to eliminate intracellular mycobacteria. Science 313(5792), 1438–1441. 16. Schmid, D., Pypaert, M., and Munz, C. (2007) Antigen-loading compartments for major histocompatibility complex class II molecules continuously receive input from autophagosomes. Immunity 26(1), 79–92. 17. Inbal, B., et al. (2002) DAP kinase and DRP-1 mediate membrane blebbing and the formation of autophagic vesicles during programmed cell death. J. Cell Biol. 157(3), 455–468.

7 Autophagosomes in GFP-LC3 Transgenic Mice Noboru Mizushima and Akiko Kuma

Summary Recent studies of the molecular mechanism of autophagy have made available several marker proteins for autophagosomes. These marker proteins allow us to identify autophagic structures easily and accurately by fluorescent microscopy. The most widely used marker for autophagosome is LC3, a mammalian homolog of Atg8. To analyze autophagy in whole animals, we generated GFP-LC3 transgenic mice and describe here how we determine the occurrence of autophagy in vivo using this mouse model.

Key Words: Autophagosome; GFP; green fluorescent protein; LC3; Atg8. 1. Introduction Although the autophagic vacuole was identified almost 50 years ago, specific markers for autophagosomes were not discovered until recently. In the 1990s, the autophagic pathway was dissected at the molecular level in the yeast Saccharomyces cerevisiae. To date, at least 16 genes have been found to be required for autophagosome formation. The nomenclature of these genes was unified under the term ATG genes, which also includes other autophagy-related genes (1). Seven of these ATG gene products function in two ubiquitination-like conjugation systems (2): one system mediating the conjugation of Atg12 to Atg5 (3) and the other mediating covalent linkage between Atg8 and phosphatidylethanolamine (PE) (4). These two systems are conserved in mammals. While the Atg12–Atg5 conjugate is present only on the autophagic isolation membrane or phagophore before complete enclosing, LC3 (a mammalian homolog of Atg8)–PE conjugate is present on both the phagophore and autophagosomes (but less on autolysosomes) (5–7). The localization of LC3 is usually determined with green fluorescent protein From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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(GFP)-conjugated LC3 (GFP-LC3) (5,8). Autophagosomes are easily detected as fluorescent dots or sometimes ring-shaped structures if they are larger than 1 μm. In Atg5−/− cells, punctuate LC3 structures are not detected (6,9). Examination of GFP-LC3 localization is a very simple and highly specific method, requiring only a high-resolution fluorescence microscope. In addition, real-time observation in living cells is feasible. Thus, this method has been applied to animal studies by generating GFP-LC3 transgenic mice in which GFP-LC3 is overexpressed in almost all tissues under the control of the constitutive CAG promoter (10). Use of this transgenic mouse model enables the occurrence of autophagy in mouse tissues to be directly monitored by fluorescence microscopic analysis of cryosections.

2. Materials 2.1. Mice 1. GFP-LC3#53 mice: This mouse line is now distributed through the RIKEN BioResource Center in Japan (http://www.brc.riken.jp/lab/animal/en/dist.shtml). 2. Wild-type C57BL/6 mice for colony maintenance.

2.2. Genotyping 1. Tail digestion solution: 10 mM Tris-HCl, pH 8.4, 50 mM KCl, 2.5 mM MgCl2 , 0.45% NP-40, 0.45% Tween-20. Autoclave and store at room temperature. Add 1/100 vol of 20 mg/mL Proteinase K (final 0.2 mg/mL) prior to use. 2. PCR primers. Primer 1: 5’-ATAACTTGCTGGCCTTTCCACT-3’; Primer 2: 5’-CGGGCCATTTACCGTAAGTTAT-3’; Primer 3: 5’-GCAGCTC ATTGCTGTTCCTCAA -3’. Primer 1 and Primer 2 for amplification of the GFP-LC3 transgenic allele (about 250 bp). Primer 1 and Primer 3 for amplification of the wild-type allele (about 350 bp) (11) (Fig. 1). 3. (optional) GFP macroscopy: Model GFsP-5 from Biological Laboratory Equipment, Maintenance and Service Ltd (http://www.bls-ltd.com/).

2.3. Sample Preparation 1. 2. 3. 4. 5.

Perista pump (ATTO AC-2110). Tissue-tek: OCT compound (Sakura Finetechnical Co., Ltd., Tokyo, Japan). Cryostat: LEICA CM3050S. SlowFade Light Antifade Kit (Molecular Probes). Microscope: Fluorescence microscope (Olympus IX81, Tokyo, Japan) equipped with a 60X oil-immersion objective lens (Plan Apo, 1.40 NA) and a cooled CCD camera (Hamamatsu Photonics, ORCA-ER (1360x1024)) (see Note 4). 6. Software: MetaMorph Series Version 6 (Molecular Device, Sunnyvale, CA).

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Fig. 1. PCR-based genotyping of GFP-LC3 mice. Genomic DNA was extracted from wild-type and hemizygous GFP-LC3 transgenic mouse tails and analyzed by PCR. Bands representing wild-type and GFP-LC3 allele are indicated.

3. Methods 3.1. Mouse Maintenance 1. It is recommended that GFP-LC3 mice be maintained as heterozygous, because it is very important to compare the transgenic mice with wild-type siblings to distinguish the GFP-LC3 signal from background autofluorescent signals. 2. If littermate control is not necessary, GFP-LC3 mice can be maintained as homozygous. No apparent abnormal phenotype has been observed in the homozygous mice.

3.2. Genotyping by Polymerase Chain Reaction (PCR) 1. 2. 3. 4. 6. 7.

Cut 0.2–0.5 cm of tail and place in a 1.5-mL tube. Add 100–400 μL of tail digestion solution (with Proteinase K). Incubate at 55°C for 8 h (or overnight). Mix occasionally. Centrifuge briefly, then boil for 5 min. Centrifuge at 13,000 g for 10 min. Take 1 μL supernatant for PCR reaction. Mix 1 μL tail sample, 0.2 μL primer 1, 0.2 μL primer 2, 0.2 μL primer 3, 2 μL PCR buffer, 1.6 μL dNTP mix (2.5 mM each), 0.2 μL rTaq (Takara), 14.6 μL DW. 8. PCR reaction: Step 1: 94°C 4 min; step 2: 94°C 0.5 min; step 3: 60°C 0.5 min; step 4: 72°C 1 min; 30 cycles to step 2; step 5: 72°C 7 min; step 6: 4°C.

3.3. Genotyping by GFP Macroscopy (Optional) 1. GFP-LC3 transgenic neonates can be easily distinguished from wild-type neonates even in mouse cages using GFP macroscopy. 2. Adult mice can be genotyped by checking the GFP fluorescent signal of mouse palms. 3. A portable UV illuminator does not work for our GFP-LC3 mice.

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3.4. Sample Preparation 1. Perfuse mice transcardially with about three times the volume (body weight) of 4% paraformaldehyde dissolved in 0.1 M Na-phosphate buffer (pH 7.4). Dipping tissues in PFA may be sufficient but quick fixation is important to avoid artificial induction of autophagy during sample preparation. 2. After perfusion, remove tissues and further fix them in the same fixative for an additional 4 h or overnight (depending on antibodies for double staining). 3. Immerse the fixed tissues in 15% sucrose/PBS for at least 4 h, then in 30% sucrose/PBS for at least an additional 4 h (or overnight). 4. Embed the tissue samples using OCT compound (Tissue-Tek) and store at –70°C. 5. Section the tissues at 5–7 μm thickness with a cryostat. Air-dry the sections at room temperature for 30 min. The sections can be stored at –70°C (or –20°C) until use. 6. Wash the well-dried cryosections in PBS and mount on glass slides using SlowFade Light Antifade Kit.

3.5. Fluorescence Microscopy 1. Select a 60X oil-immersion objective lens and put a small drop of immersion oil on the objective lens. 2. Place a glass slide and focus on cells by transmitted light imaging (usually differential interference contrast (DIC)). 3. Select an appropriate dichroic filter set (FITC or GFP). 4. Observe the sample by eye. This step should be as short as possible if the fluorescent signal is weak. 5. Capture images (Fig. 2). It is recommended also to take images using unrelated filter sets such as RFP or Cy5 as controls (see Note 2).

3.6. Quantitative Analysis of GFP-LC3 Dots The number or total area of GFP-LC3 structures can be quantified using software. However, uneven cytosolic background signals make conventional thresholding difficult or weak dot signals nonextractable. To better extract the dot signals, the “Top Hat” algorithm of the the MetaMorph Series Version 6 (Molecular Device) is useful. Small dot peaks can be extracted from the surrounding relatively lower background signals irrespective of absolute signal intensity (see Notes 1 and 3).

4. Notes 1. Although this transgenic mouse model is very useful for in vivo studies, there are a few possible limitations. First, GFP-LC3 localization represents only autophagosome formation. Since autolysosomes have less membrane-bound LC3

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Fig. 2. Example of GFP-LC3 transgenic mouse analysis. Gastrocnemius muscle samples were prepared from GFP-LC3 transgenic mice before (left) or after 24-h starvation (right) and fixed with 4% paraformaldehyde. Cryosections were analyzed by fluorescence microscopy. GFP-LC3 dots represent autophagosomes. (Bar 10 μm.) than autophagosomes, the appearance of GFP-LC3 dots does not definitely constitute autophagic “degradation.” Too fast fusion of autophagosomes with lysosomes may result in a lower number of GFP-LC3 dots, which would underestimate autophagic activity. 2. It is also very important to distinguish true GFP-LC3 dot signals from autofluorescent signals. Certain cells such as neurons show autofluorescent dot structures like lipofuscin. Such artifacts may be avoided by the following two methods. First, it is particularly important to compare samples expressing GFP-LC3 with nontransgenic control samples. Second, specific GFP-LC3 signals should not be detected using other fluorescence filter sets such as rhodamine, Cy5, or UV. True GFP-LC3 signals should be detected specifically by the GFP or FITC filter set. We usually use the U-MGFPHQ unit for GFP observation and the U-MWIG2 unit to check autofluorescence. 3. GFP-LC3 can be incorporated into protein aggregates independent of autophagy. This phenomenon is particularly apparent in hepatocytes and neurons of autophagy-deficient mice such as Atg5−/− (12) and Atg7−/− mice (13,14), or cells having unrelated inclusion bodies such as those induced by polyglutamine expression (15). Additionally, transient transfection of GFP-LC3 often causes aggregation. In these conditions, GFP-LC3 dots should be carefully interpreted. 4. We usually use wide field microscopy rather than confocal laser scanning microscopy. Since the number of autophagosomes is not so large, the highest Z-axial resolution is generally not required to eliminate “out-of-focus” fluorescence.

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References 1. Klionsky, D. J., Cregg, J. M., Dunn, W. A., Jr., et al. (2003) A unified nomenclature for yeast autophagy-related genes. Dev. Cell 5, 539–545. 2. Ohsumi, Y. (2001) Molecular dissection of autophagy: two ubiquitin-like systems. Nat. Rev. Mol. Cell Biol. 2, 211–216. 3. Mizushima, N., Noda, T., Yoshimori, T., et al. (1998) A protein conjugation system essential for autophagy. Nature 395, 395–398. 4. Ichimura, Y., Kirisako, T., Takao, T., et al. (2000) A ubiquitin-like system mediates protein lipidation. Nature 408, 488–492. 5. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728. 6. Mizushima, N., Yamamoto, A., Hatano, M., et al. (2001) Dissection of autophagosome formation using Apg5-deficient mouse embryonic stem cells. J. Cell Biol. 152, 657–667. 7. Kabeya, Y., Mizushima, N., Yamamoto, A., Oshitani-Okamoto, S., Ohsumi, Y., and Yoshimori, T. (2004) LC3, GABARAP and GATE16 localize to autophagosomal membrane depending on form-II formation. J. Cell Sci. 117, 2805–2812. 8. Mizushima, N. (2004) Methods for monitoring autophagy. Int. J. Biochem. Cell Biol. 36, 2491–2502. 9. Kuma, A., Hatano, M., Matsui, M., et al. (2004) The role of autophagy during the early neonatal starvation period. Nature 432, 1032–1036. 10. Mizushima, N., Yamamoto, A., Matsui, M., Yoshimori, T., and Ohsumi, Y. (2004) In vivo analysis of autophagy in response to nutrient starvation using transgenic mice expressing a fluorescent autophagosome marker. Mol. Biol. Cell 15, 1101–1111. 11. Kuma, A., and Mizushima, N. (2007) Chromosomal mapping of the GFP-LC3 transgene in GFP-LC3 mice. Autophagy 4, 61–62. 12. Hara, T., Nakamura, K., Matsui, M., et al. (2006) Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885–889. 13. Komatsu, M., Waguri, S., Ueno, T., et al. (2005) Impairment of starvation-induced and constitutive autophagy in Atg7-deficient mice. J. Cell Biol. 169, 425–434. 14. Komatsu, M., Waguri, S., Chiba, T., et al. (2006) Loss of autophagy in the central nervous system causes neurodegeneration in mice. Nature 441, 880–884. 15. Kuma, A., Matsui, M., and Mizushima, N. (2007) LC3, an autophago some marker, can be incorporated into protein aggregates independent of autophagy; caution in the interpretation of LC3 localization. Autophagy 3, 323–328.

8 Experimental Control and Characterization of Autophagy in Drosophila Gabor Juhasz and Thomas P. Neufeld

Summary Insects such as the fruit fly Drosophila melanogaster, which fundamentally reorganize their body plan during metamorphosis, make extensive use of autophagy for their normal development and physiology. In the fruit fly, the hepatic/adipose organ known as the fat body accumulates nutrient stores during the larval feeding stage. Upon entering metamorphosis, as well as in response to starvation, these nutrients are mobilized through a massive induction of autophagy, providing support to other tissues and organs during periods of nutrient deprivation. High levels of autophagy are also observed in larval tissues destined for elimination, such as the salivary glands and larval gut. Drosophila is emerging as an important system for studying the functions and regulation of autophagy in an in vivo setting. In this chapter we describe reagents and methods for monitoring autophagy in Drosophila, focusing on the larval fat body. We also describe methods for experimentally activating and inhibiting autophagy in this system and discuss the potential for genetic analysis in Drosophila to identify novel genes involved in autophagy.

Key Words: Atg8; autophagy; Drosophila; GFP; TEM; LysoTracker Red; somatic clones.

1. Introduction Autophagic structures in insect tissues were first described nearly 40 years ago in larval cells preparing for metamorphosis (reviewed in ref. 1). More recent analyses in Drosophila have focused on the interactions of autophagic and apoptotic processes (2–9), the regulation of autophagy by nutrient and hormonal signaling pathways (9–12), and its role in neurodegeneration (13). As Drosophila has emerged as a genetically amenable system to analyze From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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autophagy in vivo, a description of currently used methods for manipulating and monitoring autophagy in this system may be of use. Here we describe a number of approaches applicable to the study of autophagy in Drosophila.

2. Materials 2.1. Drosophila Stocks For clonal expression of transgenes, we cross UAS lines of interest carrying X-linked hsp70-FLP to a Act>CD2>GAL4 UAS-GFP strain. For loss of function mosaic studies, mutations of interest are recombined with FRT inserts at the base of the proper chromosome arm, then crossed with fb-GAL4 FRT UAS-GFP lines. Alternatively, clones can be marked using UAS-dsRed or UAS-RFP, which allows GFP-Atg8 to be used as an autophagy indicator in this system. 2.2. Larval Cultures and Starvation 1. Standard cornmeal-molasses-agar fly food. 2. 20% sucrose in water.

2.3. Dissection and Staining of Fat Bodies with LysoTracker Red 1. PBS (phosphate-buffered saline), pH = 7.4. 2. DAPI (4 ,6-diamidino-2-phenylindole dihydrochloride; Sigma, St. Louis, MO) dissolved in water to obtain a 1 mM (1000x) stock solution, which can be stored in a dark vial at 4 C. 3. LysoTracker Red DND-99 (Molecular Probes, Eugene, OR) diluted in PBS to obtain a 20 mM (200x) stock solution, which is stable in a dark vial at 4 C for a few weeks.

2.4. Fixation for Confocal Microscopy 1. Formaldehyde (Fluka, Milwaukee, WI). 2. Vectashield (Vectorlabs, Burlingame, CA).

2.5. Fixation and Embedding for Transmission Electron Microscopy (TEM) 1. Paraformaldehyde, Warrington, PA).

glutaraldehyde,

OsO4 ,

Polybed

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(Polysciences,

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3. Methods 3.1. Experimental Regulation of Autophagy in the Larval Fat Body This section discusses approaches for experimentally inducing or inhibiting autophagy in the Drosophila larval fat body, using manipulations of culture conditions and genetic expression. 3.1.1. Nutrient Control The larval fat body is exquisitely sensitive to nutrient conditions, and thus autophagy is readily induced by a simple starvation regimen. Induction of autophagy can first be detected within 1–2 h of starvation, and by 4 h after food withdrawal a robust response is consistently observed. 3.1.1.1. Pretreatment

Due to the high sensitivity of the fat body to nutrient levels, preincubation of larvae in fresh media under uncrowded conditions is recommended to prevent chronic induction of autophagy, which can diminish the acute response to starvation. We generally transfer 10–20 late second-instar larvae to vials containing fresh fly food supplemented with live yeast paste, 16–24 h prior to starvation. 3.1.1.2. Induction by Starvation

Larvae can be conveniently floated out of fly food media by pouring several mL of a 20% sucrose solution onto the culture; the high density of this solution causes the larvae to float to the surface. Larvae are then transferred by forceps to vials or small dishes containing fresh 20% sucrose and incubated at room temperature for the duration of the starvation period, generally 4 h. Alternatively, larvae can be transferred to vials containing a strip of filter paper saturated with PBS. Following the starvation period, larvae are dissected and immediately processed as described in section 3.2.2. 3.1.1.3. Induction by Rapamycin

The autophagic response to starvation in the larval fat body appears to result largely from inhibition of the target of rapamycin (TOR) signaling pathway (14). As an alternative to starvation, direct inhibition of TOR by the drug rapamycin results in a robust autophagic response. Following pretreatment as above, larvae are transferred to fresh fly food containing rapamycin at a concentration of 1 μM. Weak induction is observed by 4 h, with a much stronger response occurring by 24 h.

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3.1.2. Developmental Control During the final 24 h of the larval period, rising titers of the steroid hormone ecdysone result in a developmentally programmed induction of autophagy in the larval fat body (12,15). This developmentally regulated autophagy can be readily observed by comparing autophagy markers in early vs. late third-instar larvae under fed conditions. Premature induction can be achieved by exposure of dissected fat body to ecdysone in culture. Such methods have been described previously (15) and are not detailed here. 3.1.3. Genetic Control A major advantage of Drosophila is the ease with which genetic function can be manipulated in this system. Gene expression can be readily induced or inhibited in vivo, either singly or in multiple combinations, and at precise times and locations during development. An important component of this control is the ability to alter gene expression using a mosaic approach, in which marked clones of cells differ in their genetic makeup from their wild-type neighbors (Fig. 1A,B). The principal advantage of this approach is that experimental and control cells are subject to the same environmental and developmental conditions, and thus subtle phenotypic differences can be more readily detected than in animal-to-animal comparisons. This technique is especially applicable to studies of autophagy, which vary with developmental and culture conditions. The method of choice for both loss and gain of function mosaic experiments in Drosophila is the Flippase (FLP)-Flippase Recombinase Target (FRT) system of mitotic recombination. In this system, FLP-induced exchange between heterozygous sister chromatids causes loss of heterozygosity for a given chromosomal arm. Alternatively, FLP-induced expression of the GAL4 transcription factor can be used to clonally activate UAS-controlled transgenes. As these methods have been extensively reviewed elsewhere (16,17), we will focus our comments on their application in the larval fat body. 3.1.3.1. Loss of Function Clones

The induction of FLP-mediated, green fluorescent protein (GFP)–marked loss of function clones in the fat body requires two modifications of the general protocols developed for imaginal tissues. First, because this procedure works only in mitotically active cells, clones must be induced during the first 8 h of embryonic development, after which time the fat body enters an endoreduplicative (nonmitotic) cell cycle. We typically perform 6- to 8-h embryo collections, followed immediately by a 1- to 2-hour heat shock at 37°C to activate expression of the hsp70-driven FLP gene. This typically gives rise to several


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Fig. 1. Autophagy in the larval fat body. Examples of Lysotracker Red–stained samples are shown. (A, A ) In fed animals, clonal overexpression of Tsc1 and Tsc2, negative regulators of the TOR signaling pathway, causes a punctate pattern of LysoTracker Red staining (A ) exclusively within the transgene-expressing cells, which are marked by cytoplasmic GFP expression in (A) and indicated by arrowheads. Nuclei are stained with DAPI. (B, B ) In response to a 4-h starvation treatment, wild-type control cells display strong punctate LysoTracker Red staining (B ). In contrast, LysoTracker Red staining is blocked in cells expressing high levels of Rheb, an upstream activator of TOR signaling. Cells overexpressing Rheb are marked by cytoplasmic GFP expression in (B) and indicated by arrowheads. Nuclei are stained with DAPI. (C, C ) Fat body dissected from 4-h starved larvae expressing GFP-Atg8 (C) and stained with LysoTracker Red (C ). Large punctate structures positive for both GFP-Atg8 and LysoTracker Red represent autolysosomes (indicated by arrowheads), whereas smaller structures marked solely by GFP-Atg8 represent autophagosomes.

one- to two-cell clones per fat body lobe. Second, we find that the commonly used GFP marker lines driven by the ubiquitin promoter display nonuniform expression levels in the fat body, which can confound efforts to identify clones by GFP expression. As a remedy, we use Upstream Activating Sequence (UAS)-

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driven GFP under the control of strong fat body GAL4 lines such as fb- or Cg-GAL4 (9,18). These transgenes are recombined onto chromosomes bearing centromere-proximal FRT sites prior to use. A final note is that, unlike in imaginal tissues, fat body cells mix extensively during development, which often results in considerable separation of clones from their sister twinspots. 3.1.3.2. Clonal Expression of Transgenes

A unique feature of the fat body makes it especially amenable to FLPmediated clonal induction of transgene expression. Britton et al. (19) described heat shock–independent spontaneous induction of “flip-out” clones in the fat body and midguts of hsp70-FLP Act>CD2>GAL4-bearing larvae. Presumably these tissues are more sensitive than others to low levels of leaky expression of FLP early in their development. Induction of FLP later in development is not practical in the fat body, as it tends to result in ubiquitous transgene expression throughout the fat body, likely due to the polyploid nature of this tissue. Figure 1 displays examples of spontaneously induced clones of GFPmarked cells overexpressing Tsc1 and Tsc2 (A), which induce autophagy by inhibiting TOR, or Rheb (B), which suppresses starvation-induced autophagy by maintaining TOR activation. 3.1.3.3. Forward Genetic Approaches to Identifying Autophagy Regulators

It is perhaps obvious to point out that the genetic approaches discussed above can easily be adapted for use as tools in gene discovery. The development of comprehensive collections of UAS-regulated lines allowing induction of overexpression or RNA interference will undoubtedly lead to identification of novel regulators and effectors of autophagy. In addition, newly available collections of FRT-linked lethal P insertions provide a valuable resource for loss of function mosaic studies. 3.2. Monitoring Autophagy in the Larval Fat Body Here we describe three independent methods for assaying autophagy in the fat body. LysoTracker Red stains acidic organelles in fixed and living cells and is usually confined to the late endosomal and lysosomal compartments. Under fed conditions, the larval fat body displays virtually no punctate LysoTracker Red staining. Starvation causes a robust induction of staining, which is severely inhibited in autophagy mutants. Thus, LysoTracker Red can be considered a reliable indicator of starvation-induced autophagy in this tissue. Examples of LysoTracker Red staining are shown in Fig. 1. Greater specificity can be achieved by following the localization of GFP-Atg8, which becomes conjugated


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to autophagosomal membranes. Lines expressing GFP fused to both human and fly homologs of Atg8 have been described (10,12) and are available. The combination of LysoTracker Red staining and GFP-Atg8 fluorescence allows distinction between autophagosomes (green signal only) and autolysosomes (both green and red signal) (Fig. 1C). Finally, we include a brief protocol for analysis by TEM. 3.2.1. Larval Cultures and Starvation 1. Set up fly crosses with 5–10 females and a similar number of males. Do a 24-h egg collection. 2. Forty-eight hours after finishing egg collection, transfer approximately 20 L2stage larvae to a fresh vial to reduce crowding. Add yeast paste or dry yeast to make sure all larvae have plenty of food to eat. 3. On the next day, collect larvae to examine (fed), or transfer 8–10 larvae into 20% sucrose for 3 h (starved).

3.2.2. Dissection and Staining of Fat Bodies with LysoTracker Red 1. Wash larvae thoroughly in water or PBS. 2. Transfer three to five larvae into a drop of PBS. Cut them in half using a sharp blade, and turn the carcasses inside out using forceps. 3. Separate fat body lobes and transfer them to staining solution (1× LysoTracker Red + DAPI, prepared fresh daily) for 2 min. 4. Transfer fat body pieces into a drop of PBS or 50% glycerol-PBS on a slide. Cover with a cover slip and evaluate immediately using a fluorescent microscope.

3.2.3. Fixation for Confocal Microscopy 1. Follow the above steps, but dissect in PBS and fix in freshly prepared 3.7% formaldehyde in PBS for 20 min to overnight at 4 C. 2. Wash two times in PBS. 3. Mount in Vectashield mounting medium. 4. Seal preparations by applying nail polish around the edges of cover slips and store at 4 C.

3.2.4. Fixation and Embedding for TEM 1. Follow the steps described earlier, but dissect in PBS and fix overnight at 4 C in 2% paraformaldehyde, 4% glutaraldehyde, 2% sucrose, 100 mM phosphate buffer. 2. Rinse in 3% sucrose in 100 mM PBS. 3. Postfix in 2% OsO4 in 100 mM phosphate buffer overnight. 4. Dehydrate in ethanol and embed into Polybed 812 following the manufacturer‘s recommendations.

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4. Notes 1. All experiments are performed at 25 C except as otherwise noted. 2. With sufficient numbers of females, egg-collection times can be shortened to as little as an hour, resulting in a more synchronous larval culture. 3. Turning carcasses inside-out requires skill and practice. An alternative is to simply tear larvae open and dissect out the fat body instead. 4. To speed up the LysoTracker Red staining procedure, it is acceptable to dissect larvae directly in staining solution. 5. It is important that the appropriate amount of liquid be used when mounting fat body lobes. Too little liquid will result in crushing cells when the cover slip is applied, whereas in too much liquid, the fat body will not be flat enough for good photographs. 6. LysoTracker Red bleaches very rapidly, so it is essential to minimize exposure time when using this dye. It can be useful to first find the proper focal plane using the UV/DAPI channel. 7. LysoTracker Red fixes poorly in larval tissues, and in most cases its signal is gone by 5 min of 3.7% formaldehyde fixation. If you need to work with a fixed tissue (e.g., more time is needed for imaging, or you have a GFP-Atg8 signal also), reduce fixation time to a few seconds to 2 min. LysoTracker signal will be very weak, but you can use an antibleaching agent that way. Another option is to analyze LysoTracker Red staining and GFP-Atg8 signal in different animals using live and fixed tissue, respectively. 8. For TEM, alternate methods of fixation have also been used successfully. Dissected tissues can be fixed in 3.2% paraformaldehyde, 0.5% glutaraldehyde, 1% sucrose, 40 mM CaCl2 in 0.1 N pH 7.4 sodium-cacodylate overnight, rinsed twice in 0.1 N sodium-cacodylate, postfixed in 0.5% OsO4 in 0.1 N sodiumcacodylate for 1 h at room temperature, dehydrated in ethanol and embedded in Araldyte (Fluka) following the manufacturer‘s recommendations.

References 1. Neufeld, T. P. (2004) Role of autophagy in developmental cell growth and death: insights from Drosophila, in Autophagy (Klionsky, D. J., ed.), Landes Bioscience, Georgetown, pp. 224–232. 2. Velentzas, A. D., Nezis, I. P., Stravopodis, D. J., Papassideri, I. S., and Margaritis, L. H. (2007) Mechanisms of programmed cell death during oogenesis in Drosophila virilis. Cell Tissue Res. 327, 399–414. 3. Akdemir, F., Farkas, R., Chen, P., et al. (2006) Autophagy occurs upstream or parallel to the apoptosome during histolytic cell death. Development 133, 1457–1465. 4. Juhasz, G., and Sass, M. (2005) Hid can induce, but is not required for autophagy in polyploid larval Drosophila tissues. Eur. J Cell Biol. 84, 491–502. 5. Martin, D. N., and Baehrecke, E. H. (2004) Caspases function in autophagic programmed cell death in Drosophila. Development 131, 275–284.


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6. Gorski, S. M., Chittaranjan, S., Pleasance, E. D., et al. (2003) A SAGE approach to discovery of genes involved in autophagic cell death. Curr. Biol.13, 358–363. 7. Lee, C. Y., and Baehrecke, E. H. (2001) Steroid regulation of autophagic programmed cell death during development. Development 128, 1443–1455. 8. Lee, C. Y., Clough, E. A., Yellon, P., Teslovich, T. M., Stephan, D. A., and Baehrecke, E. H. (2003) Genome-wide analyses of steroid- and radiation-triggered programmed cell death in Drosophila. Curr. Biol. 13, 350–357. 9. Scott, R. C., Juhasz, G., and Neufeld, T. P. (2007) Direct induction of autophagy by Atg1 inhibits cell growth and induces apoptotic cell death. Curr. Biol .17, 1–11. 10. Scott, R. C., Schuldiner, O., and Neufeld, T. P. (2004) Role and regulation of starvation-induced autophagy in the Drosophila fat body. Dev. Cell 7, 167–178. 11. Lindmo, K., Simonsen, A., Brech, A., Finley, K., Rusten, T. E., and Stenmark, H. (2006) A dual function for Deep orange in programmed autophagy in the Drosophila melanogaster fat body. Exp. Cell Res. 312, 2018–2027. 12. Rusten, T. E., Lindmo, K., Juhasz, G., et al. (2004) Programmed autophagy in the Drosophila fat body is induced by ecdysone through regulation of the PI3K pathway. Dev. Cell 7, 179–192. 13. Ravikumar, B., Vacher, C., Berger, Z., et al. (2004) Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat. Genet. 36, 585–595. 14. Scott, P. H., Brunn, G. J., Kohn, A. D., Roth, R. A., and Lawrence, J. C., Jr. (1998) Evidence of insulin-stimulated phosphorylation and activation of the mammalian target of rapamycin mediated by a protein kinase B signaling pathway. Proc. Natl. Acad. Sci. USA 95, 7772–7777. 15. Sass, M., and Kovacs, J. (1977) The effect of ecdysone on the fat body cells of the penultimate larvae of Mamestra brassicae. Cell Tissue Res. 180, 403–409. 16. McGuire, S. E., Roman, G., and Davis, R. L. (2004) Gene expression systems in Drosophila: a synthesis of time and space. Trends Genet. 20, 384–391. 17. Theodosiou, N. A., and Xu, T. (1998) Use of FLP/FRT system to study Drosophila development. Methods 14, 355–365. 18. Hennig, K. M., Colombani, J., and Neufeld, T. P. (2006) TOR coordinates bulk and targeted endocytosis in the Drosophila melanogaster fat body to regulate cell growth. J. Cell. Biol. 173, 963–974. 19. Britton, J. S., Lockwood, W. K., Li, L., Cohen, S. M., and Edgar, B. A. (2002) Drosophila‘s insulin/PI3-kinase pathway coordinates cellular metabolism with nutritional conditions. Dev. Cell 2, 239–429.

9 Analysis of Autophagosome Membrane Cycling by Fluorescence Microscopy Julie E. Legakis and Daniel J. Klionsky

Summary Autophagy is a physiological process functionally linked to cellular dynamics during starvation, cardiomyopathies, neurodegeneration, cellular immunity, and certain cancers. Although nearly 30 autophagy-related (ATG) genes have been identified and characterized, the molecular mechanisms of this process are only partially understood. One aspect of the pathway that has been intensely studied is the identity of the membrane source for newly formed autophagosomes. Although it occurs at a basal level, autophagy is an inducible process. The process of autophagosome formation involves recruitment and delivery of membrane and recycling of Atg proteins. Despite continuing attempts to identify the source of the autophagosome membrane, we are only recently beginning to understand the nature of autophagosome formation and the role of membrane protein cycling in this process. There now exists an assay utilizing fluorescence microscopy to monitor the localization, and therefore the movement, of membrane-associated Atg proteins. We describe here a method that allows visualization of Atg membrane proteins in order to observe their potential source membranes and also to determine the temporal order of action of other Atg proteins with regard to their movement.

Key Words: Autophagy; autophagosome formation; membrane cycling; membrane transport; vacuole.

1. Introduction Cell survival requires the ability to adapt to rapidly changing conditions. Autophagy is a cellular recycling mechanism involving either targeted or bulk degradation and reuse of intracellular components, which enables cells to survive periods of nutrient limitation, to combat viral and bacterial infection, From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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and is implicated in preventing some types of neurodegenerative disease and cancer (1–3). Autophagy is a general term for a number of related processes and exists both as a typically nonspecific, inducible process called macroautophagy and as one of a number of pathways responsible for exclusive delivery of specific cargo to the lysosome (or the vacuole in yeast). Here we discuss a method that can be used to observe membrane protein recruitment during macroautophagy and the cytoplasm to vacuole targeting (Cvt) pathway, although the protocol can potentially be used to observe membrane protein cycling for other autophagy-related pathways. In brief, macroautophagy involves the sequestration of bulk cytoplasm, including entire organelles, at the putative site of autophagosome formation, the phagophore assembly site (PAS). Fusion of the enclosing vesicle with the lysosome/vacuole follows the completion of sequestration and ultimately results in the degradation and recycling of the autophagosome cargo. Other autophagyrelated pathways differ from macroautophagy only in the material sequestered and the site of enclosure. Current models have divided these pathways into a series of discrete steps: (1) induction; (2) cargo selection and packaging (for specific types of autophagy); (3) vesicle nucleation; (4) vesicle expansion and completion; (5) retrieval of Atg components; (6) targeting, docking and fusion with the lysosome/vacuole; (7) breakdown of the vesicle contents; and (8) recycling of the resulting macromolecules (4). At present, nearly 30 autophagyrelated (ATG) genes have been identified in yeast (5–7). Current investigations seek to clarify the role of the corresponding gene products in these pathways. The nature of membrane recruitment in autophagosome formation is not well defined. Various intracellular compartments have been proposed as the source of autophagosome membranes. Included in these potential membrane contributors are the endoplasmic reticulum (ER), Golgi complex, plasma membrane, and mitochondria (8–12; J. Legakis, W.-L. Yen, and D.J. Klionsky, unpublished observations). Recent studies have shed some light on the dynamics of this process, suggesting that membrane for the newly forming autophagosomes is recruited in part from mitochondria and the Golgi complex by specific Atg proteins (11–13; J. Legakis, W.-L. Yen, and D.J. Klionsky, unpublished observations). One of these is the transmembrane protein Atg9, which has been shown to cycle between the PAS and the mitochondria (11,13). The transport of Atg9 after knocking out ATG1 (TAKA) assay is a fluorescence microscopy method to monitor movement of Atg9, thus following membrane recruitment to the PAS (13–15). The principle behind this assay relies upon the properties of Atg9 and its movement between the PAS and other subcellular compartments. Atg9 is one of two known integral membrane proteins involved in autophagosome formation (16). In wild-type cells, Atg9 resides in multiple

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punctate structures within the cell, including the PAS, mitochondria, and other unidentified compartments, and cycles between these sites (11). In contrast, in cells with ATG1 deleted, Atg9 is restricted to the PAS, unable to travel to the other subcellular locations (13). Utilizing an epistasis approach, by introducing a second mutation (for example, atgX) in addition to the ATG1 deletion, a temporal function in Atg9 movement can be assigned to the protein of interest. Specifically, the protein can be categorized as either (1) required for anterograde movement of Atg9 to the PAS; (2) needed for retrograde Atg9 movement (from the PAS); or (3) not involved in Atg9 cycling. For example, Atg9-YFP is manifest as a single punctate structure in atg13 cells, and shows a similar phenotype in atg1 atg13 cells. Thus, Atg13 appears to act at the same time or after Atg1 in Atg9 cycling and is required for retrograde Atg9 transport Fig. 1). In contrast, Atg9-YFP is localized at multiple punctate structures in both atg27 and atg1 atg27 cells, suggesting that Atg27 acts before Atg1 in movement of Atg9 and is involved in anterograde Atg9 trafficking Fig. 1). A protein is not involved in Atg9 cycling if Atg9 is localized in multiple punctate structures (one of which must be shown to correspond with the PAS) in the single mutant, and in a single punctate structure in conjunction with atg1. Moreover, real-time cycling of Atg9 can be observed using a temperature-sensitive allele of ATG1 (atg1ts). By

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Fig. 1. Epistasis analysis of Atg9-YFP. Wild-type, atg1, atg13, atg1, atg13 atg27, and atg1 atg27 cells expressing the Atg9-YFP fusion protein were grown to mid-log phase, harvested and analyzed by fluorescence microscopy. Atg9-YFP in the atg27 and atg1 atg27 strains is not located at the PAS (data not shown) indicating that Atg27 is needed for anterograde movement of Atg9. DIC, differential interference contrast.

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repeatedly shifting between the permissive and nonpermissive temperatures, the movement of Atg9-GFP can be followed from its wild-type localization at multiple punctate structures to concentrating at the PAS (anterograde transport) and redistributing back to the multiple punctate sites (retrograde movement). This assay also allows the identification of other factors (e.g., non-Atg proteins) required for Atg9 cycling. In addition, the TAKA assay can be utilized to follow trafficking of two other membrane-associated Atg proteins, Atg23 and Atg27 (12; J. Legakis, W.-L. Yen, and D. J. Klionsky, unpublished observations). Atg23 is a peripheral membrane protein that cycles between the mitochondria and the PAS (13,17; J. Legakis, W.-L. Yen, and D. J. Klionsky, unpublished observations), and Atg27 is a transmembrane protein that has recently been shown to cycle from the mitochondria/Golgi to the PAS (12). Although these proteins, similar to Atg9, cycle between the PAS and other subcellular compartments, their movement is not entirely coincident with that of Atg9 (14; J. Legakis, W.-L. Yen, and D. J. Klionsky, unpublished observations). Continued use of the TAKA assay will likely lead to a more complete understanding of the subcellular compartments in which these three proteins reside, possibly to discovery of other proteins involved in the cycling process, and to a better overall understanding of membrane recruitment for autophagosome formation. 2. Materials 2.1. Cell Culture 1. Rich medium (YPD): 1% yeast extract, 2% peptone, 2% glucose (see Notes 1 and 2). 2. Synthetic minimal medium (SMD): 0.67% yeast nitrogen base, 2% glucose, amino acids, and vitamins, as needed. 3. Synthetic medium lacking nitrogen (SD-N): 0.17% yeast nitrogen base without amino acids, 2% glucose.

2.2. Yeast Stable Transformation 1. 50% PEG-4000: polyethylene glycol, molecular weight 3,350 g/mol, 50% w/v. 2 Lithium acetate solution: 1 M lithium acetate, pH 7.5 (adjusted with acetic acid). Dilute this solution 10-fold for 100 mM lithium acetate. 3. Denatured single-stranded DNA (ssDNA): 2 mg/mL heat-denatured (5 min boiling) salmon sperm DNA, sheared by sonication (approximately 3 min with a probe sonicator). 4. Transformation mix: 240 μL 50% PEG-4000, 36 μL lithium acetate solution, 50 μL ssDNA, 9 μL sterile double-distilled or Millipore filtered H2 O, 25 μL transforming DNA (PCR product).


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2.3. Yeast Plasmid Transformation 1. 10X TE buffer: 100 mM Tris-HCl, pH 7.5. 2. 10X lithium acetate buffer: 1 M lithium acetate, pH 7.5 (adjusted with acetic acid). 3. TE/lithium acetate buffer: 1 mL 10X TE buffer (10 mM final), 1 mL 10X lithium acetate buffer (100 mM final), 8 mL sterile H2 O. This solution should be made fresh for each set of transformations. 4. 50% PEG-4000: polyethylene glycol, molecular weight 3350 g/mol, 50% w/v. This solution should be filter sterilized and made fresh every 3–4 weeks. 5. Denatured single-stranded DNA (ssDNA): 2 mg/mL heat-denatured (5 min boiling) salmon sperm DNA, sheared by sonication (approximately 3 min with a probe sonicator). 6. Transformation mix: 150 μL 50% PEG-4000, 10 μL ssDNA, 1–2 μL plasmid DNA (from a miniprep).

2.4. Fluorescence Microscopy 1. Fixation buffer: 50 mM KH2 PO4 , pH 6.5–8.0 (see Note 3), 1.5% formaldehyde, 1 μM MgCl2 . 2. Wash buffer: 50 mM KH2 PO4 , pH 6.5–8.0 (same pH as the fixation buffer), 1 μM MgCl2 .

3. Methods The proteins monitored for this assay, Atg9, Atg23 and Atg27, are all dynamic proteins residing in multiple punctate structures. It is possible to visualize these proteins in live cells, but because they cycle rapidly between the compartments, for accurate colocalization studies with most microscopes, the cells must be fixed. Fixation also enhances the fluorescent signal, as the proteins are stationary. Furthermore, for accurate evaluation of their behavior, fluorescent chimeras of the proteins should be created by chromosomal tagging, thus allowing visualization of the proteins expressed at physiological levels. Other fluorescent organelle markers, for example, RFP-Atg8 for the PAS, Mito-BFP for the mitochondria, etc., can either be introduced by plasmid transformation, or the tag can be integrated into the chromosome, similar to Atg9GFP. Furthermore, many subcellular structures can be stained using dyes such as MitoTracker Red (Invitrogen) to visualize mitochondria prior to fixation. 3.1. Cell Culture 1. All yeast colonies are maintained on agar plates containing rich or selective media. Colonies on plates can be stored for up to one month at 4°C. Liquid cultures are grown from a single colony.

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2. Cells are first grown in liquid medium overnight, then diluted and re-grown to mid-log phase in rich (YPD), or selective (SMD), medium, as needed (see Note 4). For starvation conditions, the cells are grown in medium lacking nitrogen (SD-N) for one hour prior to viewing.

3.2. Yeast Stable Transformation: Strain Construction and Stable Transformations for Integration into the Chromosome 1. Fluorescent tagging of Atg9 with GFP (or RFP, BFP, etc.) is achieved by PCRbased integrations of the tag at the 3 end of the gene. This allows expression of the fusion protein under the control of the native promoter. The template for integration of Atg9-GFP is pFA6-GFP-HIS3 S.k. (or -KAN; 18). Approximately 10 μg (generally 25 μL of a standard PCR reaction) of the appropriate PCR product is sufficient to generate transformants. 2. Cells are grown overnight at 30°C (see Note 5), diluted in 5 mL culture medium/transformation (referred to as one unit of competent cells) to OD600 ≤ 0.25, regrown to OD600 0.8–1.0, harvested by centrifugation, and washed once with 4 mL of H2 O per unit of competent cells. Resuspend in 100 μL of 100 mM lithium acetate per unit of competent cells and aliquot 100 μL of the competent cells into a separate microcentrifuge tube for each transformation (see Note 6). 3. Pellet the cells by centrifugation at 3000 rpm for 3 min, resuspend by pipetting in transformation mix totaling 360 μL and incubate at 30°C for 30 min to 3 h (see Note 5). A negative control should be included in which the cells are incubated in transformation mix lacking DNA. 4. Transfer the cells to 37–42°C (heat shock) and incubate 30 min to 1 h (see Note 5). 5. Pellet the cells by centrifugation at 4000 rpm for 2 min, aspirate the supernatant, resuspend each sample in 200 μL H2 O, and spread transformed cells on plates selective for the appropriate auxotrophic marker or antibiotic resistance factor (see Note 7). 6. Incubate plates 2–3 days at 30°C (see Note 5). 7. Individual colonies can be screened for Atg9-GFP expression by Western blot using anti-GFP antibodies (commercially available), or by PCR-based screening.

3.3. Yeast Plasmid Transformation 1. Cells are grown overnight at 30°C (see Note 5), diluted in 10 mL culture medium/6 transformations to OD600 0.5, regrown to OD600 0.8–1.0, harvested by centrifugation at 3000 rpm for 3 min and washed once with 1 mL of H2 O, followed by a wash in 1 mL TE/lithium acetate buffer. Cells are then resuspended in 150 μL lithium acetate, and are now referred to as competent cells. 2. Twenty-five μL of competent cells are aliquoted into microcentrifuge tubes, transformation mix is added to each condition, and incubated at 30°C for 30 min (see Note 5). A negative control should be included in which the cells are incubated in transformation mix lacking plasmid DNA.


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3. Tubes are then transferred to 37–42°C for 15 min (see Note 5). 4. Cells are pelleted by centrifugation at 4000 rpm for 2 min, resuspended in 200 μL H2 O and spread onto selective agar plates (see Note 7). 5. Plates are grown at 30°C for 2–3 days (see Note 5), and colonies can be screened by Western blotting for expression of the protein encoded by the plasmid.

3.4. Fluorescence Microscopy with Fixation (Basic Protocol) 1. Cells expressing chromosomally tagged Atg9-GFP (and, if desired, plasmids encoding RFP-Atg8 and/or Mito-BFP) are first grown overnight at the appropriate temperature, followed by dilution of 5 mL of culture to OD600 ≤ 0.5 and regrowth to OD600 0.8–1.0. 2. Cells are harvested by centrifugation at 3000 rpm for 3 min in 15 mL conical tubes, resuspended by pipetting in 2.5 mL fixation buffer, and incubated with gentle mixing for 30 min at room temperature. 3. Wash cells once with 2.5 mL wash buffer, and resuspend in 200 μL wash buffer. 4. Drop 2 μL of cell solution onto a microscope slide and carefully place a cover slip over the top, avoiding the introduction of bubbles under the cover slip. 5. The cells are visualized with a fluorescence microscope with deconvolution software for image analysis (or a confocal microscope), first establishing the focus with differential interference or phase contrast microscopy. If using a deconvolution microscope, software must be used to deconvolve the images in order to evaluate a single focal plane of the cells. Typical localization patterns of Atg9GFP, in both wild-type (colocalized with the PAS marker, RFP-Atg8 in Fig. 2B) and atg1 cells are shown in Fig. 2.

3.5. Fluorescence Microscopy with Fixation (Cycling Protocol) 1. Cells expressing chromosomally tagged Atg9-GFP, harboring an ATG1 deletion and a plasmid-based, reversible temperature-sensitive allele of ATG1 (atg1ts), are first grown overnight at the permissive temperature of 24°C, followed by dilution of 10 mL of cell culture to OD600 ≤0.5 and regrowth to OD600 0.8–1.0. Plasmids encoding organelle markers or other fluorescent proteins can be introduced as necessary. This experiment requires continuing the growth of cells in liquid culture for a number of hours; therefore it is recommended that the culture be periodically diluted to maintain it at mid-log growth phase. In order to accurately evaluate the movement of these proteins within a single population of cells, it is necessary to simultaneously prepare an aliquot of the cells for microscopy while maintaining the remainder of the liquid culture at the appropriate temperature for subsequent analysis. Therefore, the complete steps required for one condition (a) will be outlined, followed by the instructions for the next condition (b), and so on Fig. 3). 2a. One mL of cells is removed from the culture, pelleted in a microcentrifuge tube at 3000 rpm for 3 min, resuspended in 0.5 mL of fixation buffer, and incubated at room temperature, with gentle shaking for 30 min.

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Fig. 2. Atg9-GFP localization in wild-type and atg1 cells. (A) Wild-type and atg1 cells expressing an Atg9-GFP chimera were grown to mid-log phase, harvested and viewed by fluorescence microscopy. (B) Wild-type cells harboring chromosomally tagged Atg9-GFP and transformed with a plasmid encoding RFP-Atg8 were grown to mid-log phase, subjected to mild fixation, and examined by fluorescence microscopy. A white arrow marks the position of RFP-Atg8. DIC, differential interference contrast. 3a. The aliquot of cells is pelleted as above, washed once in 0.5 mL of wash buffer, and resuspended in 50 μL of wash buffer, followed by evaluation by fluorescence microscopy, as outlined in Subheading 3.4., step 5. 2b. At the same time, the remaining cell culture is shifted to the nonpermissive temperature of 37°C, and incubated for 1 h. 3b. One mL of cells is removed from the culture, pelleted in a microcentrifuge tube as above, resuspended in 0.5 mL of fixation buffer, and incubated at room temperature, with gentle shaking for 30 min. 4b. The aliquot of cells is pelleted as above, washed once in 0.5 mL of wash buffer, resuspended in 50 μL of wash buffer, followed by evaluation by fluorescence microscopy, as outlined in Subheading 3.4., step 5. 2c. Meanwhile, the remaining cell culture is shifted to the permissive temperature of 24°C and incubated for 1 h. 3c. One mL of cells is removed from the culture, pelleted in a microcentrifuge tube as above, resuspended in 0.5 mL of fixation buffer, and incubated at room temperature, with gentle shaking for 30 min. 4c. The aliquot of cells is pelleted as above, washed once in 0.5 mL of wash buffer, resuspended in 50 μL of wash buffer, followed by evaluation by fluorescence microscopy, as outlined in Subheading 3.4., step 5. An example of Atg9-GFP cycling evaluated by a similar protocol is shown in Fig. 4.

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Fig. 3. Flow chart for sample preparation during TAKA assay with a temperature sensitive atg1 mutant strain. See text for details.

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Fig. 4. Atg9 can cycle to and from the PAS. The atg1 strain expressing Atg9-YFP was transformed with a plasmid bearing atg1ts. Cells were grown in SMD selective medium at 24°C to OD600 0.8, shifted to 37°C for 60 min., then shifted back to 24°C for another 60 min. Before each temperature shift, cells were imaged by fluorescence microscopy. DIC, differential interference contrast. (Reproduced from ref. 14 with permission from Cell Press.)

4. Notes 1. All solutions should be prepared in water that has a resistance of 18.2 M-cm (referred to throughout as H2 O). 2. All solutions for cell culture and transformations should be sterile. Furthermore, all of these procedures should be carried out using sterile conditions.

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3. The pH of the fixation and wash buffers can be adjusted to optimize staining of other intracellular structures by pH-dependent dyes. 4. Yeast cells in culture can be diluted and regrown indefinitely, as long as they are evaluated at or near mid-log phase (OD600 = 0.8–1.0). 5. Temperature-sensitive mutant strains should be grown at the appropriate permissive temperature (usually 23–26°C) and the heat shock should be carried out at 30–37°C. 6. The cells can be stored overnight at 4°C in the 100 mM lithium acetate solution. 7. If using the kanr marker (kanamycin resistance), pellet cells, resuspend in 1 mL YPD/transformation and incubate 30°C for 1 h (see Note 5) prior to spreading onto plates.

Acknowledgments This work was supported by the National Institutes of Health Public Health Service grant GM53396 (to D.J.K.). References 1. Levine, B. and Klionsky, D. J. (2004) Development by self-digestion: molecular mechanisms and biological functions of autophagy. Dev. Cell 6, 463–477. 2. Shintani, T. and Klionsky, D. J. (2004) Autophagy in health and disease: a doubleedged sword. Science 306, 990–995. 3. Legakis, J. E. and Klionsky, D. J. (2006) Overview of autophagy, in Autophagy in Immunity and Infection: A Novel Immune Effector (Deretic, V., ed.), Wiley-VCH, Weinheim, pp. 3–17. 4. Klionsky, D. J. (2005) The molecular machinery of autophagy: unanswered questions. J. Cell Sci. 118, 7–18. 5. Klionsky, D. J., Cregg, J. M., Dunn, W. A., Jr., et al. (2003) A unified nomenclature for yeast autophagy-related genes. Dev. Cell 5, 539–545. 6. Kawamata, T., Kamada, Y., Suzuki, K., et al. (2005) Characterization of a novel autophagyspecific gene, ATG29. Biochem. Biophys. Res. Commun. 338, 1884–1889. 7. Stasyk, O. V., Stasyk, O. G., Mathewson, R. D., et al. (2006) Atg28, a novel coiled-coil protein involved in autophagic degradation of peroxisomes in the methylotrophic yeast Pichia pastoris. Autophagy 2, 30–38. 8. Legesse-Miller, A., Sagiv, Y., Glozman, R., and Elazar, Z. (2000) Aut7p, a soluble autophagic factor, participates in multiple membrane trafficking processes. J. Biol. Chem. 275, 32966–32973. 9. Fengsrud, M., Lund Sneve, M., Overbye, A., and Seglen, P.O. (2004) Structural aspects of mammalian autophagy, in Autophagy (Klionsky, D.J., ed.) Landes Bioscience, Georgetown, TX, pp. 11–25. 10. Reggiori, F., Wang, C.-W., Nair, U., Shintani, T., Abeliovich, H., and Klionsky, D.J. (2004) Early stages of the secretory pathway, but not endosomes, are


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required for Cvt vesicle and autophagosome assembly in Saccharomyces cerevisiae. Mol. Biol. Cell 15, 2189–2204. Reggiori, F., Shintani, T., Chong, H., Nair, U., and Klionsky, D.J. (2005) Atg9 cycles between mitochondria and the pre-autophagosomal structure in yeasts. Autophagy 1, 101–109. Yen, W.-L., Legakis, J.E., Nair, U., and Klionsky, D.J. (2007) Atg27 is required for autophagy-dependent cycling of Atg9. Mol. Biol. Cell 18, 581–593. Reggiori, F., Tucker, K. A., Stromhaug, P. E., and Klionsky, D. J. (2004) The Atg1- Atg13 complex regulates Atg9 and Atg23 retrieval transport from the preautophagosomal structure. Dev. Cell 6, 79–90. Shintani, T. and Klionsky, D. J. (2004) Cargo proteins facilitate the formation of transport vesicles in the cytoplasm to vacuole targeting pathway. J. Biol. Chem. 279, 29889–29894. Cheong, H., Yorimitsu, T., Reggiori, F., Legakis, J. E., Wang, C.-W., and Klionsky, D.J. (2005) Atg17 regulates the magnitude of the autophagic response. Mol. Biol. Cell 16, 3438–3453. Noda, T., Kim, J., Huang, W.-P., et al. (2000) Apg9p/Cvt7p is an integral membrane protein required for transport vesicle formation in the Cvt and autophagy pathways. J. Cell Biol. 148, 465–480. Tucker, K. A., Reggiori, F., Dunn, W. A., Jr., and Klionsky, D. J. (2003) Atg23 is essential for the cytoplasm to vacuole targeting pathway and efficient autophagy but not pexophagy. J. Biol. Chem. 278, 48445–48452. Longtine, M. S., McKenzie, A., III, Demarini, D. J., et al. (1998) Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953–961.

10 Protein Trafficking into Autophagosomes Andrew Young and Sharon Tooze

Summary The methods described are designed to enable the assignment of an intracellular localization of secretory proteins, either soluble or membrane associated, to later secretory compartments, such as the trans-Golgi network (TGN) or endosome. These two subcellular compartments are closely linked through extensive protein trafficking, in both an anterograde and a retrograde direction. These compartments are likely to be important in the formation of autophagosomes during the process of autophagy. Our current knowledge of how autophagosomes form is scarce, and further investigation into the role that other subcellular compartments have in this process is needed.

Key Words: Autophagy; protein trafficking; subcellular fractionation; Atg; mAtg9;Golgi;endosome

1. Introduction This chapter addresses methods to determine to which intracellular organelle a protein localizes. Determining where in a cell a protein localizes is often the first stage in characterizing a novel protein. The localization of a novel protein may provide clues as to its function or, if the protein is already of known function, then it is important to know where within the cell it performs that function. Here we will focus on techniques concerned with determining whether a protein localizes to pathways leading to the formation of autophagosomes. Protocols to determine whether a protein localizes to Golgi and endosomal membranes are described as these potentially have inputs into the induction of autophagy and formation of autophagosomes. Recent investigations into a transmembrane protein called mammalian Atg9 (mAtg9), originally identified From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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in yeast as a protein required for autophagy, have exploited these techniques to start to gain an understanding of this question (1). The analysis of the glycosylation of secretory proteins or transmembrane proteins (only proteins that have passed through the secretory pathway are glycosylated) can be informative in determining a protein‘s localization. However, not all proteins synthesized in the ER are N-glycosylated; some may be O-glycosylated. N-linked glycosylation is the most common form of glycosylation, which is particularly useful for pinpointing precisely where a protein is in the early secretory pathway. The presence of a NXS/T motif within the primary amino acid sequence of the protein is a good indication that the protein is glycosylated. However, not all these motifs will be glycosylated; for example, for reasons of steric hindrance, or limits in accessibility of the site to the glycosylation enzymes may inhibit glycosylation. Proteins that traffic to and though the Golgi acquire a complex form of N-linked glycosylation, which is detectable using a combination of endoglycosidase H (Endo H) and PNGase F glycosidases. Endo H cleaves only high-mannose forms of glycans present on proteins in the ER and early Golgi. The presence of Endo H–resistant glycosylation can therefore be used to determine whether the protein of interest has passed through the later compartments of the Golgi. PNGase F is able to also cleave off the complex forms of glycosylation that are added in later Golgi compartments and can therefore be used to determine if a protein is N-glycosylated. Immunofluorescence analysis can be used to determine where a protein of interest is localized to at steady state by assessing its overlap with a panel of proteins that have a previously determined localization within the cell. Centrifugation methods can be used to the same end. Differential, velocitycontrolled, or equilibrium centrifugation of cellular material after mechanical disruption of the plasma membrane can be used to separate subcellular organelles, and the distribution of marker proteins of previously determined localization can be compared to that of the protein of interest and its localization ascertained. Examples of the marker proteins mentioned above are: protein disulfide isomerase (PDI), a protein resident in the lumen of the ER; GM130, a protein resident in the cis/medial cisternae of the Golgi apparatus (2); mannosidase II, a medial Golgi glycosidase; TGN38, at steady state displaying a predominantly trans-Golgi localization (3); EEA1, an early endosomally associated tether protein (4); and Rab proteins, specific Rabs that have been shown to localize to specific organelles (5). Antibodies to a majority of these proteins are commercially available. The best characterized and most frequently used marker for autophagosomes is microtubule associated protein (MAP) light chain 3, or LC3 (6,7). Several commercial antibodies are available, but use of these

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should be carefully controlled with co-localization with GFP-LC3, or increased accumulation of LC3-II under conditions expected to induce autophagy, for example, vinblastine (8). GFP-LC3 expressed by transfection of plasmid is frequently used, but caution should be used in the interpretation of the results if GFP-LC3 is overexpressed as this may create artefacts.

2. Materials 2.1. N-Glycosylation 1. Endoglycosidase H (Endo H) and PNGase F (New England Biolabs). 2. Ball-bearing-based cell cracker (EMBL Workshop, Heidelberg, Germany). 3. Phosphate-buffered saline (PBS): 0.137 M NaCl, 2.7 mM KCl, 10 mM Na2 HPO4 , 18 mM KH2 PO4 (Sigma). Adjust to pH 7.4 with HCl if necessary and autoclave before storage at room temperature. 4. Cell scraper can be made from a semi-circular piece of rubber stopper (bung) mounted on the end of a plastic 10 mL pipet or purchased (e.g., from Fischer Scientific). 5. Trypan blue (Sigma). 6. Ultracentrifuge, and ultracentrifuge rotor, capable of achieving >100,000g, for example, Beckman TLA45.

2.2. Immunofluorescence 1. Glass cover slips, 22 mm2 square, 1.5 mm thick, and glass microscopy slides, 76 × 26 mm (e.g. from Fisher Scientific). 2. Paraformaldehyde, 16% (v/v) (Agar Scientific), dilute to 3% using PBS, and add MgCl2 and CaCl2 to give 84 μM and 96 μM final, respectively. 3. PBS: 0.137 M NaCl, 2.7 mM KCl, 10 mM Na2 HPO4 , 18 mM KH2 PO4 . Adjust to pH 7.4 with HCl if necessary and autoclave before storage at room temperature. 4. NH4 Cl, make up as a 1 M stock in water and dilute using PBS to the working concentration, 50 mM solution. Store both at room temperature. 5. Triton X-100 (Sigma). Made up as a 20% (w/v) stock and store at 4ºC. 6. Porcine skin gelatin (Sigma), make up as a 4% (w/v) stock and store in ∼15 mL aliquots at −20ºC. 7. 3MM paper (Whatmann). 8. Parafilm (Pechiney Plastic Packaging). 9. Secondary antibodies conjugated to Alexa 488, 555, and 647 (Molecular Probes). These Alexa dyes have a strong signal and are quite fade resistant. 10. Mowiol 4-88 (Calbiochem). Add 2.4 g Mowiol to 6 g glycerol and 6 mL distilled water and incubate at room temperature for 2 h. Then add 12 mL 0.2 M Tris-HCl pH 8.5 and incubate at 53ºC, stirring occasionally, until the Mowiol dissolves. Make 1 mL aliquots and freeze at –20ºC for storage.

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2.3. Subcellular Fractionation 2.3.1. Golgi Preparation 1. Six female Sprague-Dawley rats, weight 180–200 g. 2. Potassium phosphate buffer (PPB): mix sufficient 0.1 M K2 HPO4 solution and 0.1 M KH2 PO4 solution to achieve pH 6.7. Measure with pH electrode. Add MgCl2 to 5 mM , pepstatin A to 5 μM final and 1 complete EDTA free protease inhibitor tablet (Roche) per 50 mL. 3. Scissors, sharp/surgical quality. 4. Sucrose, ultra-pure. 5. Ultracentrifuge, and ultracentrifuge rotor, capable of achieving >100,000g, for example, Beckman SW28 rotor and SW28 tubes (or equivalent). 6. 150 μm mesh steel laboratory sieve (Endecotts Ltd, UK).

2.3.2. Endosome Preparation 1. Predialyzed Ficoll (Sigma). 2. Delta Refractometer (Bellingham and Stanley Ltd., UK). 3. 0.4 M TES (2-[2-hydrox-1,1-bis(hydroxymethyl)ethyl)amino]ethanesul-fonic acid) pH 7.4, sucrose, 0.5 M EDTA pH 7.4. 4. STE buffer: 250 mM sucrose, 10 mM TES pH 7.4 and 1 mM EDTA pH 7.4. 5. STM buffer: 250 mM sucrose, 10 mM TES pH 7.4, 1 mM MgCl2 . 6. Gradient maker, with reservoirs capable of holding at least 15 mL each. 7. Nycodenz (Axis-Shield, Norway). 8. Ultracentrifuge and ultracentrifuge rotor VTi50 (Beckman). 9. 39 mL Quick-seal centrifuge tubes (Beckman, polyallomer, 25 × 89 mm) and heat sealer for Vti50 rotor. 10. Peristaltic pump. 11. Fraction collector, capable of holding 30–40 tubes. 12. Potter Elvejhem homogeniser (Thomas Scientific). 13. 10-mL syringe and long (∼10 cm) blunt-ended needle. 14. 2 mm diameter stainless steel tube ∼10cm in length.

3. Methods 3.1. N-Glycosylation Analysis 1. Seed one 35-mm dish for each experimental data point to achieve 80–90% confluency (typically 1.6 × 106 cells) on the day of the experiment. 2. Using a cell scraper, scrape the cells in PBS. 3. Homogenize the cells using the ball-bearing-based cell cracker (passing the cells through a narrow gauge needle can be just as effective). Check the cells for breakage using trypan blue (see Note 1). Keep the homogenate on ice. 4. Centrifuge the homogenate at 2200g for 10 min at 4ºC.


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5. Centrifuge the resulting post nuclear supernatant (PNS) at 45,000 g in a Beckman TLA45 ultracentrifuge rotor (or equivalent) for 1 h at 4ºC to pellet the membranes. This allows resuspension of the proteins in a sufficiently small volume to allow glycosidase enzyme treatment. 6. Resuspend the membrane pellet in PBS containing protease inhibitors (see Notes 2 and 3). 7. Treat the resulting solution with the glycosidase enzymes Endo H and PNGase F (New England Biolabs [NEB]) as per the manufacturer’s instructions. 8. Analyze by SDS-PAGE and Western blotting, and look for an increase in the protein‘s electrophoretic mobility upon removal of its N-glycosylation. Shown in Fig. 1 is an example of a glycosidase digest performed on mAtg9.

3.2. Immunofluorescence 1. Seed the cells at least the day before at a sufficient density that they will be reasonably sparse upon fixation, approximately 50–60% confluency (typically 2.6 × 106 cells), in order that individual cells will be clearly visible in the microscope. Seed the cells in a 6-well dish, containing the cover slips. Depending upon the adherence properties of the cell type, pretreatment of the cover slips can help avoid cells washing off the cover slip during the procedure (see Note 4). 2. Fix the cells using 3% (v/v) formaldehyde (Agar Scientific) in PBS containing 84 μM MgCl2 and 96 μM CaCl2 (for subsequent staining with Rab antibodies, see Note 5). Alternatively, cells can be fixed using pure methanol at −20ºC (see Note 6) (if using methanol, leave out the quenching and permeabilization steps, steps 3 and 4). 3. Incubated cells in 50 mM NH4 Cl for 10 min to quench residual formaldehyde, wash three times with PBS. 4. Permeabilize the cells by incubating them in 0.2% (w/v) Triton X-100 in PBS for 5 min. Then wash three times in PBS.

Fig. 1. Mammalian Atg9 is present in a late or post-Golgi fraction. mAtg9 with an HA-tag was transfected into HEK293 cells. After preparation of a lysate, aliquots were subjected to digestion with Lane 1, Endo H (H), Lane 2, untreated (UN), or Lane 3, with PNGase F (F). The samples were subjected to SDS-PAGE, followed by Western blotting with anti-HA antibodies. HA-mAtg9 has Endo H–resistant, N-linked glycans.

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5. Wash three times with 0.2% (w/v) gelatin in PBS and leave in the last gelatin wash for at least 20 min to block. 6. Dilute the primary antibody in the gelatin solution and incubate with the cover slips for 20 min (see Note 7). 7. After washing three times with the gelatin solution, incubate with the secondary antibodies. 8. Wash the cover slips three times with the gelatin solution, three times with PBS, and then finally once with water. 9. Finally, drain cover slips by touching on 3 MM paper, and mount the cover slips on slides using Mowiol 4-88.

3.3. Subcellular Fractionation 3.3.1. Golgi Preparation Rat liver Golgi are prepared as described by Slusarewicz et al. (9). The procedure uses a discontinuous sucrose gradient made up of sucrose solutions of different concentrations (see Note 8) in PPB. Organelles, and in particular Golgi membranes, are here subjected to differential centrifugation, which separates them by density. 1. Starve the rats for 24 h prior to the experiment (see Note 9). 2. Prepare six discontinuous sucrose gradients consisting of 13 mL of 0.86 M sucrose in PPB underlain with 7.5 mL of 1.3 M sucrose in PPB in Beckman Ultraclear SW28 tubes (see Note 10). 3. Sacrifice the rats by asphyxiation with CO2 followed by cervical dislocation. 4. Excise livers using scissors and quickly immersed in 200 mL ice cold 0.5 M sucrose in PPB and swirl and squeeze the livers occasionally to expel any blood and to speed cooling. 5. Place 36 g of liver into fresh 0.5 M sucrose in PPB and cut into pieces with scissors to release as much blood as possible. 6. Decant excess liquid to leave a volume of less than 80 mL. 7. Mince the livers into small pieces, approximately 4–5 mm square, using scissors. 8. Homogenize the liver by gently pressing it through the 150 μm mesh steel laboratory sieve using the bottom of a 250 mL conical flask (see Note 11). Collect the homogenate in a plastic dish. 9. Pour the homogenate (H) into a 100 mL measuring cylinder and make up to a final volume of 80 mL with 0.5 M sucrose in PPB, then thoroughly mix. 10. Overlay 13 mL of the homogenate onto each of the six gradients and top up with 0.25 M sucrose in PPB. 11. Centrifuge in a Beckman ultracentrifuge using a SW28 rotor at 28,000 g for 1 h at 4ºC. Keep an aliquot of the homogenate and snap freeze in liquid nitrogen. 12. After centrifugation, remove the lipid layer on the surface of the gradient by aspiration.


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13. Collect the intermediate (I) Golgi fraction from the interface between the 0.5 M and 0.86 M using a Pasteur pipet and bulb (approximately 2–3 mL from each gradient). 14. Pool the fractions from the six gradients and diluted to 8–9% (w/w) sucrose (0.25–0.28 M) using PPB. Check the concentration using the Delta refractometer. 15. Pour this intermediate fraction into two SW28 tubes (Beckman) and top up with 0.25 M sucrose in PPB. Keep an aliquot of the intermediate fraction and snap freeze in liquid nitrogen. 16. For each tube underlay 0.5 mL of 1.3 M sucrose in PPB (see Note 12). 17. Centrifuge at 7000 g in an SW28 rotor for 30 min at 4ºC. 18. Remove the supernatant by aspiration and collect the final membrane felt, the Golgi membranes (G) with a P200 Gilson pipet (or equivalent). This should yield 1–1.5 mL of Golgi membranes. 19. Divide into 100-μL aliquots and snap-freeze in liquid nitrogen, then store at −80ºC. These membranes can be thawed and refrozen at least twice without significant change of morphology or loss of enzymatic activity. 20. Analyze the fractions collected using organelle markers. Figure 2 shows the result of immunoblotting for Mannosidase II (MannII), TGN38, and mAtg9. Note that Golgi cisternae are enriched by this preparation.

3.3.2. Endosome Preparation The endosome preparation is from rat liver according to the protocol of Ellis et al. (10), which essentially consists of a continuous Ficoll gradient on which organelles are separated by density. 1. Prepare the gradients the day before the experiment. Dissolve predialyzed Ficoll (Sigma) in water, 1 mL water per gram of Ficoll. After the Ficoll has dissolved, dilute it until the refractive index is 1.37, to give a 25% (w/v) solution.

Fig. 2. Mammalian Atg9 fractionates in a similar fashion to TGN38, but not Mannosidase II. Fractions from the Golgi preparation, homogenate (H), intermediate fraction (I), and purified Golgi (G) were analyzed by SDS-PAGE, followed by Western blotting with antibodies to Mannosidase II (MannII), TGN38, and mAtg9.

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Fig. 3. Mammalian Atg9 is found in fractions partly overlapping with TGN38, but not EEA1 or PDI. Rat liver postnuclear supernatant (PNS) was subjected to the endosome gradient analysis as described. Fractions were collected and numbered as shown. The individual fractions were analyzed by SDS-PAGE and immunoblotted with the antibodies shown. mAtg9 sediments more closely to the Cation-independent mannose-6-phosphate receptor (CI-MPR, data not shown; see ref. 1) but can be found in membranes containing TGN38, as expected from Fig. 2. The overlap with EEA1 is only partial, and EEA1 exhibits a different profile, peaking at Fraction. No. 27, whereas mAtg9 has a peak at Fraction. No. 24.

2. Add 400 mM TES pH 7.4, solid sucrose and 0.5 M EDTA pH 7.4 to the 25% Ficoll solution to give 22% Ficoll, 250 mM sucrose, 10 mM TES and 1 mM EDTA. 3. Use buffer STE to dilute the 25% Ficoll to 1% Ficoll solution. 4. Add 15 mL of the 22% Ficoll to the chamber of the gradient maker closest to the exit tube and 15 mL of the 1% Ficoll to the other chamber. Stir the 22% Ficoll chamber by means of a magnetic stir bar. 5. Place 4 mL of a 45% (w/v) Nycodenz (Axis-Shield, Norway) solution into the bottom of a 39 mL Quick-Seal centrifuge tube (Beckman, polyallomer, 25 × 89 mm). 6. Overlay the Ficoll gradient onto the Nycodenz by connecting the gradient maker to a peristaltic pump and placing the tubing into the neck of the centrifuge tube such that the Ficoll solution drips down onto the Nycodenz. 7. To start pouring the gradient, turn on the pump and open the tap separating the chambers simultaneously. Once poured, leave the gradients to sit overnight at 4ºC in order to let them smooth by diffusion. 8. Sacrifice the rat by asphyxiation with CO2 followed by cervical dislocation and excise the liver and transfer it to a beaker containing 3 mL/g liver of ice cold STM buffer. 9. Homogenize the liver in a 4ºC cold room using a Potter Elvejhem homogenizer (Thomas Scientific) fitted with a Teflon pestle rotating at 2400g, with three complete up-and-down strokes. 10. Centrifuge the homogenate at 1500g for 10 min at 4ºC. 11. Load the resulting post nuclear supernatant (PNS) on top of the Ficoll gradients, 5 mL per gradient, using a syringe coupled to a long blunt-ended needle.


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12. Heat seal the tubes closed and centrifuge in a VTi50 rotor (Beckman) at 50,000g (206,000g) for 1 h at 4ºC using slow initial acceleration and no braking for deceleration. 13. After centrifugation, remove the top of each tube using a scalpel. Working in a 4ºC cold room, pass a 2 mm diameter stainless steel tube down to the bottom of the centrifuge tube. 14. Connect the metal tube via the peristaltic pump to a fraction collector. Precalibrate the pump speed to run at ∼1 mL/min with water and program the fraction collector to move to the next tube every 60 s (see Note 13). 15. Snap-freeze the fractions in liquid nitrogen and store at −80ºC. 16. Analyze the fractions by SDS-PAGE and Western blotting. Shown in Fig. 3 is an analysis of EEA1, TGN38, and PDI, compared to mAtg9.

4. Notes 1. Trypan blue is a dye that crosses the plasma membrane but is actively pumped out of live cells. Dead cells or disrupted cells cannot pump out the dye and stain blue. 2. The membrane pellet can be difficult to spot after centrifugation; mark the top of the centrifuge tube on the side facing outwards in order to determine on which side of the tube the pellet should appear after removal from the rotor. Often the pellet can appear as a round opaque spot. 3. The membrane pellet from the high-speed spin (>100,000g) can be difficult to resuspend. Resuspension is achieved through a combination of disruption with a glass bulb made from melting a disposable glass pipet, repeated pipetting in a small volume of the PBS, and vortexing or shaking at 4°C. 4. One method of cover slip pretreatment is to overlay the cover slips with a 0.1 mg/ mL solution of poly-d-lysine and incubating them for ≥15min at room temperature before washing three times with water to remove any that has not stuck to the cover slip. 5. For subsequent staining with Rab antibodies, in order to remove the non– membrane associated cytoplasmic pool of the Rab, cells can be prepermeabilized with 0.05% saponin in 80 mM Pipes pH 6.8, 1 mM MgCl2 , 5 mM EGTA (a physiological buffer) for 5 min before fixation and then washed with PBS before fixation. 6. Different antibodies will work best with different fixation methods, which must be empirically determined. 7. For economy, use only 100 μL of diluted antibody solution. In order to do this, line the bottom of a plastic container with wetted 3MM paper (Whatmann) and place a sheet of Parafilm (Pechiney Plastic Packaging) on top of the paper. The wetted 3MM paper will maintain a humidified atmosphere and the Parafilm will cause the 100 μL of antibody solution to form a droplet. Drain the cover slip on dry 3MM paper by touching the edge against the paper, and invert the cover slips onto to the antibody droplets using tweezers.

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8. To ensure the sucrose solutions are the exact concentrations, check them using a Delta refractometer (Bellingham and Stanley Ltd., UK). Often it is necessary to either dilute or add more sucrose to the solution, despite careful preparation. 9. The rats are starved so as to stop lipoprotein synthesis, which would otherwise dramatically alter the Golgi’s physical characteristics. 10. Underlaying of the heavier sucrose solution is performed using a syringe connected to a long (∼10 cm) metal needle by passing the needle down to the bottom of the tube. 11. This relatively gentle method of homogenization reduces the possibility of cisternal unstacking by mechanical shear (11). 12. The dense sucrose will act as a cushion to stop the membranes impacting on the bottom of the tube and potentially breaking. 13. The varying density of the Ficoll will mean that the volume of the fractions collected will vary. To allow comparison with published markers, it is recommended that the refractive index is measured. This will also allow the quality of the gradient formed after centrifugation to be checked. The gradient should be a linear gradient.

References 1. Young, A. R., Chan, E. Y. W., Hu, X. W., et al. (2006) Starvation and ULK1dependent cycling of mammalian Atg9 between the TGN and endosomes. J. Cell Sci. 119, 3888–3900. 2. Nakamura, N., Rabouille, C., Watson, R., et al. (1995) Characterization of a cisGolgi matrix protein, GM130. J. Cell Biol. 131, 1715–1726. 3. Luzio, J. P., Brake, B., Banting, G., Howell, K. E., Braghetta, P., and Stanley, K. K. (1990) Identification, sequencing and expression of an integral membrane protein of the trans-Golgi network (TGN38).Biochem. J. 270(1), 97–102. 4. Mu, F.-T., Callaghan, J. M., Steele-Mortimer, O., et al. (1995) EEA1, an early endosome-associated protein.J. Biol. Chem. 270(June 2), 13503–13511. 5. Zerial, M., and McBride, H. (2001) Rab proteins as membrane organizers. Nat. Rev. Mol. Cell Biol. 2(2), 107–117. 6. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19(21), 5720–5728. 7. Tanida, I., Ueno, T., and Kominami, E. (2004) LC3 conjugation system in mammalian autophagy. Int. J. Biochem. Cell Biol. 36(12), 2503–2518. 8. Köchl, R., Hu, X., Chan, E., and Tooze, S. A. (2006) Microtubules facilitate autophagosome formation and fusion of autophagosomes with endosomes. Traffic 7(2), 129–145. 9. Slusarewicz, P., Hui, N., and Warren, G. (1994) Purification of rat liver Golgi stacks, in Cell Biology: A Laboratory Handbook (Celis, J. E., ed.), Academic Press Inc., Orlando, FL, pp. 509–517.


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10. Ellis, J., Jackman, M., Perez, J., Mullock, B., and Luzio, J. (1992) Membrane Traffic Pathways in Polarised Epithelial Cells, IRL Press, Oxford, UK. 11. Hino, Y., Asano, A., Sato, R., and Shimizu, S. (2001) Biochemical studies of rat liver Golgi apparatus. J. Biochem. (Tokyo) 83, 909–923.

11 Sphingolipids in Macroautophagy Grégory Lavieu, Francesca Scarlatti, Giusy Sala, Stéphane Carpentier, Thierry Levade, Riccardo Ghidoni, Joëlle Botti, and Patrice Codogno

Summary Sphingolipids are constituents of biological membranes. Ceramide and sphingosine 1-phosphate (S1P) also act as second messengers and are part of a rheostat system, in which ceramide promotes cell death and growth arrest, and S1P induces proliferation and maintains cell survival. As macroautophagy is a lysosomal catabolic mechanism involved in determining the duration of the lifetime of cells, we raised the question of its regulation by sphingolipid messengers. Using chemical and genetic methods, we have shown by GFP-LC3 staining and analysis of the degradation of long-lived proteins that both ceramide and S1P stimulate autophagy.

Key Words: Autophagy; ceramide; proteolysis; sphingosine 1-phosphate; sphingosine kinase; sphingolipids.

1. Introduction The metabolism of sphingolipids is a highly dynamic process generating second messengers that include ceramide and sphingosine 1-phosphate (S1P) (1). The formation of ceramide (an N-acylated sphingoid base) is followed by a deacylation to generate sphingosine, which is phosphorylated by sphingosine kinases (SK) to produce S1P. It is now generally admitted that ceramide and S1P have contrasting roles in the response to cell stress (2). Whereas ceramide is generally associated with cell growth arrest and cell death induction, S1P promotes cell proliferation and maintains cell survival. Macroautophagy (hereafter referred to as “autophagy”) is a lysosomal, catabolic pathway regulating the turnover of macromolecules and organelles (3). From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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Its impact on cell lifetime remains unclear, since autophagy appears to be a cytoprotective mechanism (when nutrient supply is limited), but can also lead to type 2 cell death (also known as autophagic cell death) as distinct from apoptosis or type 1 cell death (4). The similar functions of the sphingolipid rheostat and of autophagy in determining cell fate led us to wonder whether sphingolipids could regulate autophagy. To find out, we investigated autophagic capacities after manipulating the levels of ceramide and S1P in MCF-7 cells and HT-29 cancer cells (5,6). The endogenous ceramide level was increased by feeding cells with the short cell-permeant C2-ceramide, whereas its biosynthesis was prevented by treating cells with the ceramide synthase inhibitor fumonisin B1 (FB1). Overexpression of sphingosine kinase 1 (SK1) was used to increase the level of S1P, and treatment with dimethylsphingosine (DMS) was used to inhibit its biosynthesis. Autophagic parameters were evaluated by measuring long-lived protein degradation as described in the following sections to estimate the autophagic flux into the lysosome and determine the number of autophagosmes formed using GFP-LC3 (7,8). The findings show that increases in both endogenous ceramide and S1P levels can indeed trigger autophagy.

2. Materials 2.1. Cell Culture and Treatment 1. Dulbecco’s modified Eagle’s medium (DMEM) (Invitrogen, Fisher Biosciences, ILLkirch, France) supplemented with 10% fetal bovine serum (FBS) (Invitrogen, Fisher Biosciences) and 1% penicillin-streptomycin (Invitrogen, Fisher Biosciences). 2. C2 -Ceramide (C2 -Cer, Calbiochem 110145), C2 -dihydroceramide (C2 -DHCer, Calbiochem 219537), and dimethylsphingosine (DMS, Calbiochem 310500) dissolved in ethanol at 117 mM, 117 mM, and 75 mM, respectively. 3. 3-Methyladenine (3-MA, Sigma M9281) and fumonisin B1 (Sigma F1147) dissolved in water before use at 10 mM and 5 mM, respectively.

2.2. Degradation of Long-Lived Proteins 1. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2 HPO4 , 1.4 mM KH2 PO4 , pH 7.3. 2. Hanks’ balanced salt solution without sodium bicarbonate (HBSS) or Earle’s balanced salt solution (EBSS) from Invitrogen, Fisher Biosciences. 3. l-[U-14 C] Valine (266 mCi/mmol, Amersham Biosciences). 4. Trichloroacetic acid (TCA) (v/w 100%). 5. 0.2 N NaOH.

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2.3. Lipid Phosphate Evaluation 1. Methanol and chloroform. 2. Sodium phosphate, monobasic, anhydrous (NaH2 PO4 ), dissolved in water. Store at room temperature. 3. Washing buffer: 10 N H2 SO4 -70%HClO4 -H2 O, 9:1:40, (v/v/v) respectively. Store at room temperature. 4. Ammonium molybdate (Sigma), 0.9% (w/v) dissolved in water. 5. L-Ascorbic acid, 9%, (w/v) dissolved in water, prepare freshly just before use.

2.4. Preparation of Octyl--d-Glucopyranoside: Dioleoylphosphatidylglycerol (OG-DOPG) Mixed Micelles 1. 2. 3. 4.

Octyl--d-glucopyranoside (OG) (Calbiochem 494459). Pyrex sintered glass funnel from E. Pasquali Srl (Italy). Diethyl ether (Fluka 32203). l--Dioleoylphosphatidylglycerol (DOPG sodium salt) (Avanti Polar Lipids 840475).

2.5. Diacylglycerol Kinase (DGK) In Vitro Assay 1. Sn-1,2-Diacylglycerol kinase, recombinant, Escherichia coli (≥2 units/mg protein, MW 13,700, Calbiochem 266724). 2. 2x buffer: 0.1 M imidazole (Sigma I0125), pH 6.6, 0.1 M LiCl (Sigma L8895), 25 mM MgCl2 anhydrous (Sigma M8266), 2 mM EGTA (Sigma E4378), pH 6.6 dissolved in water. Store at 4°C up to 6 mo. 3. Dithiothreitol (DTT) (Sigma D5545) dissolved in water at 1 M; aliquots stored at –80°C. 4. Assay mixture: 50 μL of 2x buffer, DTT 3 mM (0.2 μL of a 1 M solution) and 5 μg of diacylglycerol kinase, Recombinant enzyme for each sample added just before use. 5. Dilution buffer: 10 mM imidazole, pH 6.6, 1 mM diethylenetriaminepentaacetic acid (DTPA, Sigma), pH 7.0. Store at 4°C for up to 6 mo. 6. ATP from (Amersham Pharmacia Biotech) dissolved in water at 20 mM and aliquots stored at –80°C. 7. Adenosine 5 -[-32 P]triphosphate, triethylammonium salt (3 Ci/μmol specific activity) from Amersham Biosciences. 8. ATP mixture: ATP, 10 mM and [−32 P]ATP (about 0.13 μCi/μL) for each sample, added just before use. 9. Ceramide from porcine brain used for internal standard purchased from Avanti Polar Lipids Inc. (860052). 10. Diacylglycerol used for internal standard (Avanti Polar Lipids Inc. (800811)). 11. Recrystallized octyl--d-glucopyranoside (OG): a. Add 5 g of OG to 20 mL of acetone. b. Heat mixture at 40°C to completely dissolve the OG.

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Lavieu et al. c. Filter the solution through a sintered Pyrex glass funnel. d. Slowly add 100 mL diethyl ether to the filtered solution and reheat if necessary (if precipitation appears in the solution). e. Place in –20°C freezer overnight (or longer) to form crystals. f. Decant the solution containing recrystallized OG through a sintered glass funnel. g. Wash the crystals with 500 mL ice-cold diethyl ether. h. Completely dry the crystals and scrape out of the funnel. i. Weigh the crystals and store in powder form at –20°C for up to 12 mo.

2.6. Sphingosine Kinase Activity and Sphingosine 1-Phosphate Production 1. d-erythro-Sphingosine (Biomol EI-155) dissolved in ethanol at 50 mM, in a screw-cap glass tube, and stored at –20°C. 2. [-32 P]Adenosine 5´-triphosphate ([-32 P]ATP, 6000 Ci/mmol, PerkinElmer Life Sciences). 3. d-erythro-[3-3 H]Sphingosine (23 Ci/mmol, PerkinElmer Life Sciences) dissolved in ethanol. 4. Sphingosine kinase buffer : 20 mM Tris-HCl pH 7.4, 20% glycerol, 1 mM mercaptoethanol, 1 mM EDTA, 1 mM phenylmethylsulfonylfluoride, 15 mM NaF, 10 μg/mL leupeptin, 10 μg/mL aprotinin, 40 mM -glycerophosphate, 0.5 mM deoxypyridoxine, and 1 mM sodium orthovanadate.

2.7. Separation of Reaction Products (Ceramide 1-Phosphate) by Thin-Layer Chromatography 1. Thin-layer chromatography (TLC) plates from Whatman (4865-821, LK6D Silica Gel 60A, Size 20 × 20, thickness 250 μm). 2. Acetic acid (Fluka 27221) and acetone (Fluka 00570). 3. X-Omat AR from Kodak (1651454, size 20.2 × 25.4). 4. Transfer a mixture of chloroform–acetone–methanol–acetic acid–water (10:4:3: 2:1, by vol.) into a well-sealed TLC chamber (use silicone and apply pressure in order to obtain good adherence of the lid). Saturate the chamber with vapors by using a sheet of filter paper as a wick (a circular strip, approximately the same height as the tank). For a 26 × 20 × 7 cm chamber, use 150 mL of solvent and allow to saturate for 5–12 h.

2.8. Separation of Reaction Products (Sphingosine 1-Phosphate) by TLC 1. Solvent A: 1-butanol/methanol/acetic acid/distilled water (80:20:10:20, by vol.) 2. Whatman LK6D TLC plates (20 cm × 20 cm).


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3. Chromatography tank, containing about 100 mL of solvent A. The tank should be prepared (and sealed) at least 2 h before use. 4. Autoradiography films (Kodak BioMax MR).

3. Methods 3.1. Assay of Long-Lived Protein Degradation The protocol described below was originally validated in human colon cancer HT-29 cells, and then in various different cancer cells, in particular in human breast cancer MCF7 cells (5,6) (Fig. 1), and can be optimized depending on the cell system used. Autophagy has been also assayed after transfection of the autophagy marker (Fig. 1). For a description of this method see refs. 7 and 8. 1. Cells are seeded in 6-well plates at 106 cells/plate and used near confluence. 2. Intracellular proteins are labeled for 18 h at 37°C with 0.2 μCi/mL of l-[U14 C]valine in complete medium. 3. Any unincorporated radioactivity is removed by rinsing three times with PBS. 4. Cells are then incubated with fresh complete medium and 10 mM valine for one hour (see Note 1). 5. Short-lived proteins are degraded after incubating for one hour, when the medium is replaced; the cells are then incubated in HBSS (or EBSS) plus 0.1% of bovine serum albumin (see Note 2) and 10 mM valine to stimulate autophagy or with the appropriate fresh complete medium, and incubated for a further 4 h. Throughout the chase period, 3-MA can be added to inhibit de novo formation of autophagic vacuoles (9; see also Note 3). Under the experimental conditions shown in Fig. 2, the chase medium was complete medium supplemented with 10 mM valine (Fig. 1B). 6. The medium is then precipitated overnight with TCA (100%) added to produce a final concentration of 10%. 7. After centrifuging the culture medium for 10 min at 470g at 4°C, the acid-soluble radioactivity is measured by liquid scintillation counting. 8. The cells are washed twice with cold 10% TCA, dissolved at 37°C in 0.2 N NaOH. Radioactivity is then measured by liquid scintillation counting. The rate of long-lived protein degradation is calculated from the ratio of the acid-soluble radioactivity in the medium to the acid-precipitable cell fraction.

3.2. Quantification of Ceramide The nonpolar properties of ceramide mean that it has to be extracted from cells in organic solvents. This is done using a modification of the Bligh and Dyer method (10), which involves lysis of the cells with an organic solvent followed by dilution with chloroform and water to obtain phase separation. To obtain a single-phase mixture, it is important to maintain the following

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Fig. 1. Effect of sphingolipids on autophagy. (A) GFP-LC3 staining and quantification of cells with GFP-LC3 dots in MCF-7 cells transfected with an empty vector (control), transfected with a cDNA encoding the wild-type sphingosine kinase 1 (SK1wt ). Control cells were treated with C2 -Cer (75 μM) for 2 h. When required, the cells were treated with DMS (1.5 μM). The bar represents 10 μm. (B, left) Degradation of [14 C]valine-labeled, long-lived proteins in MCF-7 cells transfected with an empty vector (control), transfected with a cDNA encoding the wild-type sphingosine kinase 1 (SK1wt ). Control cells were treated with C2 -Cer (75 μM) for 2 h. When required, cells were treated with DMS (1.5 μM). (B, right) Degradation of [14 C]valine-labeled, long-lived proteins in HT-29 cells treated with C2 -Cer (75 μM) or C2 -DHCer (75 μM) for 2 h in the presence or the absence of FB1 (100 μM) or 3MA (10 mM). (Reproduced from refs. 5 and 6 with permission from ASBMB.)

proportions: methanol, chloroform, water—2:1:0.6, (by vol.). A double-phase system is then obtained by adding chloroform and water in order to achieve a final methanol, chloroform, water ratio of 2:2:1.6 (by vol.). The lower phase, containing less polar lipids, is separated from the upper phase, containing more polar lipids plus nonlipidic molecules. The critical parameters of the lipid extraction are the ratios of chloroform, methanol, and buffer/water. Although the absolute volumes may be changed to reflect the amount of mass (tissues) being extracted, it is important that the ratios be maintained at 2:1:0.6 (by vol.) and 2:2:1.6 (by vol.) before and after dilution, respectively (10).


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Fig. 2. Sphingolipid level and sphingosine kinase activity. (A) MCF-7 cells were transfected with the cDNA encoding the wild-type sphingosine kinase 1 (SK1wt ) or an empty vector (control). SK activity and sphingolipid level were analyzed 24 h posttransfection. When required, cells were treated with DMS (1.5 μM), FB1 (100 μM), 3MA (10 mM). Cells were treated for 2 h with C2 -Cer (75 μM). The S1P level was determined after [3 H]sphingosine incorporation and endogenous long-chain ceramides were quantified by the DGK assay. (B) Long-chain ceramide level was quantified in HT-29 cells by the DGK assay after treatment with C2 -Cer (75 μM) or C2 -DHCer (75 μM) for 2 h in the presence or the absence of FB1 (100 μM) or 3MA (10 mM). (Reproduced from refs. 5 and 6 with permission from ASBMB.)

3.2.1. Lipid Extraction 1. After treatment, the cells are washed twice in ice-cold PBS, pH 7.4, scraped straight off the plate, and harvested by centrifuging at 4°C for 5 min. About one to two times 106 HT-29 cells (one 10-cm Petri dish, 80% confluent) can be processed using 2 mL of methanol, 1 mL of chloroform, and 0.6 mL of water. After centrifuging, the solvents are added to dry pellets of cells, in 13 × 100 mm glass screw-cap test tubes, and the capped tubes are then vortexed immediately for 15 s (see Note 4). 2. The samples are kept at room temperature for at least 10 min and are then vortexed again for 15 s. Chloroform (1 mL) and water (1 mL) are then added, and samples are vortexed once more for 15 s. This results in a biphasic mixture. The biphasic mixture is composed of a lower lipid-containing chloroform phase and an upper phase consisting of methanol and water. 3. The mixture is centrifuged at 2000g for 5 min at 4°C and then the two phases are separated: the organic phase (lower one) can be collected in fresh glass tubes, and

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the lipids are evaporated under a stream of nitrogen or under vacuum. To provide values for phospholipids within the range of the standard values, take a 1-mL aliquot of the lower phase for ceramide determination and 0.3-mL duplicates for determining the phospholipid content (see Note 5). 4. Dry the samples for phospholipid quantification under a stream of nitrogen or under vacuum, and store the remaining sample at 20°C for the DG assay.

3.2.2. Lipid Phosphate Evaluation 1. The cellular phosphate is extracted with the lipid and is subjected to colorimetric assay, being quantified by referring to a standard curve, and provides a measurement of the cell mass of the samples. The standard measurements, ranging from 5 to 120 nmol of phosphate, are acquired by taking 5- to 120-μL aliquots of a 1 mM NaH2 PO4 aqueous solution in screw-cap glass test tubes. 2. Add 0.6 mL of washing buffer to the dried lipid samples and standard tubes. 3. Put the open tubes in a heating block at 157°C for 4–16 h. Let the samples reach this temperature slowly, without preheating the block. This procedure evaporates the aqueous phase, leaving approximately 100 μL of free phosphate in the tubes. 4. Once the samples have cooled, add 0.9 mL of water, and vortex the samples for 30 s to resuspend thoroughly (see Note 6). 5. Add 0.5 mL of ammonium molybdate and vortex samples. Molybdate and phosphate will react to produce phosphomolybdic acid. 6. Add 0.2 mL of L-ascorbic acid to the samples, and mix gently. This step converts phosphomolybdic acid into molybdenum blue, which has its highest absorbance at 820 nm. 7. After incubating the samples at 45°C (water bath) for 30 min, cool for 5–10 min and read the absorbance at 820 nm in the spectrophotometer. Standard absorbances are used to plot a standard curve. The phosphate content can be determined by referring to the standard curve, allowing for the appropriate dilution factors resulting from the extraction process (see Note 7).

3.2.3. Preparation of OG-DOPG Mixed Micelles All glassware must be acid washed, rinsed thoroughly with water, and then rinsed with acetone and dried. 1. Take an aliquot of 0.97 mL of 20 mg/mL l--dioleoylphosphatidylglycerol (DOPG) in a Pyrex screw-cap tube (see Note 8). 2. Dry the DOPG as a thin film around the bottom of the glass tube (see Note 9). 3. Take up a 7.5 % solution of OG in water, and add 1 mL to each tube containing dried DOPG. 4. Sonicate the mixture until the DOPG has completely dissolved.

Vortex vigorously and repeat the process. Store at –20°C once the micellar solution is completely clear.


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3.2.4. Diacylglycerol Kinase After lipid extraction by following the Bligh-Dyer protocol the ceramide is labelled in vitro using DG kinase (11,12), which phosphorylates both diacylglycerols and ceramides to form phosphatidic acid and ceramide-1-phosphate, respectively. This in vitro assay is precise, and makes it possible to perform a quantitative analysis of ceramide (Fig. 2). To obtain the quantitative conversion of ceramide to ceramide-1-phosphate, it is crucial to perform the assay with an excess of enzyme so that the substrate is totally converted into the product (13,14). The quantification of ceramide can be determined from the slope of a standard curve using known amounts of naturally occurring ceramide and diacylglycerol after calculating the mass of ceramide-1-phosphate produced by DG kinase. The lipids extracted are phosphorylated by the kinase, using [-32 P]ATP. The amount of enzyme per sample indicated below can be changed, depending on the activity of the protein. 1. A standard curve can be plotted using substrates of DG kinase, such as diacylglycerol and ceramide. After resuspending in chloroform, samples containing 80–2560 pmol of these standard lipids are used to plot a standard curve by taking aliquots in glass test tubes and then evaporating to dryness under a stream of nitrogen or under vacuum. 2. The extracted lipids and standards are resuspended in the mixed micelles containing a nonionic detergent and phospholipids. 20 μl of mixed micelles (see Subheading 2.) is added to the samples. 3. Vortex the lipids for 30 s, leave for 5 min at room temperature, sonicate in a water bath for 30 s and then immediately vortex again for 30 s. 4. Carefully resuspend the samples (do not vortex) in 70 μL of assay mixture containing DG kinase. 5. Adjust the volume of each sample to 70 μL using dilution buffer. 6. Begin the reaction by adding 10 μL/sample of ATP mixture. 7. Vortex the samples carefully and briefly. The reaction is then allowed to proceed for 30 min at room temperature. 8. Stop the reaction by adding 1 mL chloroform, 2 mL methanol, and 0.6 mL water. Vortex vigorously for 30 s. 9. Extract the phosphorylated lipids again according to the Bligh-Dyer procedure by adding 1 mL chloroform and 1 mL water, and vortex vigorously. 10. Separate the two phases by centrifuging at 2000g for 5 min at 4°C. 11. Discard the upper phase that contains about 95% of the radioactivity. Aliquot the lower phase, containing the phosphorylated lipids, into fresh tubes and dry under a stream of nitrogen or under vacuum.

3.2.5. Separation of Reaction Products by TLC Various substrates present in lipid extracts are commonly phosphorylated by DG kinase. This means that liquid scintillation counting of the organic extract

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obtained after the DG kinase reaction does not provide an exact determination of the ceramide 1-[32 P]phosphate content. The products of the DG kinase reaction are therefore subjected to TLC, and the ceramide-1[32 P]phosphate is identified by its co-migration with phosphorylated ceramide standards and on the basis of its Rf value. 1. Resuspend dried lipids in 80 μL of a mixture of chloroform-methanol (1:1, by vol.) and vortex in capped tubes for 30 s. 2. Immediately afterwards, apply 20 μL at the origin of a prescored silica gel 60 plate (see Note 10). 3. Use an appropriate solvent system to separate lipids by migration on the silica. Place the silica plate in the chamber (inside the paper ring), and allow the solvent front to migrate to the top of the plate (see Note 11). 4. After marking the solvent front lane, dry the plate in a chemical hood for 15–30 min before exposing to an x-ray film (16–24 h of exposure at –80°C is usually necessary). 5. Spots of interest are scraped from the plate and quantified by liquid scintillation counting (see Note 12). Diacylglycerols and short-chain ceramides will appear as their phosphorylated derivatives, with Rf values of 0.62 and 0.25–0.3, respectively. 6. The values of the radioactive counts are normalized for the concentrations of total phospholipids in the samples.

3.3. Sphingosine Kinase Activity and Sphingosine 1-Phosphate Production Phosphorylation of sphingosine can be catalyzed by two sphingosine kinases, SPHK1 and SPHK2, the enzymatic properties of which exhibit some differences (15,16). To obtain reliable and reproducible estimation of the kinase activity, the activities of S1P lyase and phosphatases have to be blocked; this is accomplished by adding the required inhibitors to the sphingosine kinase buffer. The assay for sphingosine kinase activity presented here is essentially that developed by Spiegel and coworkers (17,18) (Fig. 2). In this assay, sphingosine is converted by sphingosine kinase into radiolabeled S1P in the presence of [-32 P]ATP. The radiolabeled S1P is then separated from the substrates by solvent extraction and TLC. In this procedure, which uses acidic conditions, most of the S1P is extracted in the organic phase. To determine the amount of S1P produced (Fig. 2), we propose an assay based on the ability of intact living cells to phosphorylate an exogenously administered radioactive sphingoid base ([3-3 H]sphingosine). The radiolabeled S1P formed by the cells is then extracted from the cells or culture medium (S1P has amphiphilic properties, which make it likely that it will cross cell membranes and be secreted into the extracellular medium) using organic


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solvents. Provided that the lipid extraction is carried out under alkaline conditions, S1P can be recovered from the aqueous phase (19). 3.3.1. Preparation of Samples for the Assay of Sphingosine Kinase Activity 1. Preparation of cell lysates: Control MCF7 cells or SK1-overexpressing MCF-7 cells (5 × 105 cells/well in 6-well plates) are incubated and exposed to the appropriate stimulus. At the end of the incubation time, wash the cells twice with PBS. Scrape the cells with a rubber policeman and lyse in sphingosine kinase buffer. Centrifuge cell lysates at 100,000g for 90 min at 4°C (using a Beckman Ti50 rotor). Collect the supernatants. Determine the protein concentration of aliquots using the technique described by Bradford (20), and the Bio-Rad protein assay reagent. Store supernatants at –80°C. 2. Preparation of sphingosine-Triton X-100 micelles: prepare a 1 mM solution of sphingosine containing 5% Triton X-100 by mixing 5 μL of the sphingosine stock solution and 245 μL of 5% Triton X-100 and sonicating for 5 min in a water bath. 3. Preparation of the ATP mixture: just before the enzyme assay, prepare a solution that contains 20-mM unlabeled ATP and 200-mM MgCl2 . Since each assay sample requires 10 μL of the mixture, to 9 μL of this solution add 1 μL of a [-32 P]ATP solution at approximately 10 μCi/μL. Take an aliquot of this mixture and count in a scintillation counter.

3.3.2. Assay of Sphingosine Kinase Activity on Cell Lysates 1. Place glass test tubes on a rack in ice. 2. Add cytosolic extract (50–200 μg of protein) or, for the blank, sphingosine kinase buffer. Then, add sphingosine kinase buffer to obtain a volume of 180 μL. Add 10 μL of the 1 mM sphingosine-Triton X-100 micelles (see Note 13). 3. Start the reaction by adding 10 μL of the [-32 P]ATP solution. 4. Incubate for 30 min at 37°C in a shaking water bath. 5. Transfer the tubes into ice. Stop the reaction by adding 20 μL 1 N HCl and 0.8 mL of chloroform/methanol/12 N HCl (100:200:1, by vol.). Mix thoroughly, and allow to stand at room temperature. Five minutes later, add 0.24 mL chloroform and 0.24 mL 2 N KCl. Vortex again and, 5 min later, centrifuge at 1000g for 10 min. 6. Aspirate 0.4 mL of the lower organic phase, and transfer into a fresh test tube. Add 0.9 mL of the theoretical upper phase, and mix vigorously. Five minutes later, centrifuge at 1000g for 10 min. 7. Aspirate 0.4 mL of the lower organic phase, and dry under a stream of nitrogen. 8. Dissolve in 60 μL of chloroform/methanol (2:1, by vol.), and spot 30 μL onto a TLC plate. Samples should be applied 2 cm from the bottom of the plate. Also spot an S1P standard on a separate lane. Applications can be made by spotting 5-μL aliquots and then drying with a hair dryer.

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9. Place the TLC plate in the TLC tank, put the lid on, and then let the solvent migrate until it reaches about 1–2 cm from the top of the plate. Remove the plate and let dry in a hood. 10. Expose the plate to iodine vapors by putting it in a TLC chamber containing iodine. Once the lipids are visualized, take a picture of the plate or mark the area corresponding to the S1P standard (with a pencil). Then let the iodine evaporate by leaving the plate in a hood (a warm hair dryer can be used to accelerate the evaporation of the iodine). 11. Wrap the plate in plastic (Saran) film, and expose to autoradiography film. After overnight exposure, develop the film. Mark the areas corresponding to the radiolabeled S1P, and then scrape these areas into a piece of paper. Transfer the silica to scintillation vials, add scintillation fluid, shake, and count in a scintillation counter. 12. Calculate the sphingosine kinase activity (expressed as pmol/min/mg) in each sample as follows: first calculate the number of cpm per pmol of total ATP (see Note 14). Assuming that the S1P formed has the same specific radioactivity as ATP, convert the number of cpm scraped off the plate (after subtracting the blank) into pmol of S1P. Correct for the incubation time, the quantity of protein, and the volume of sample spotted onto the TLC plate.

3.3.3. Determination of S1P Production by Intact Cells 1. Control MCF7 cells or SK1-overexpressing MCF-7 cells (105 cells) are plated in 25-cm2 cell culture flasks. 2. Twenty-four hours later, when confluence has almost been reached, the culture medium is replaced by fresh FBS-free medium containing 0.3 μCi/mL of [3-3 H]sphingosine (as an ethanolic solution). Any compound of interest to be tested is then added. 3. After incubating at 37°C for the desired time (from 3 to 24 h), the culture medium is collected and frozen at –20°C. The cells are washed twice with PBS, and scraped off with a rubber policeman. Sedimented cells are then frozen at –20°C. 4. The cell pellets are resuspended in 200 μL distilled water and lysed by freezethawing. An aliquot can be kept to determine the protein content. 5. Add 1.5 mL of chloroform/methanol (1:2, by vol) to cell lysates or aliquots (0.2 mL) of culture medium, mix, and then add 0.7 mL of 0.5 N NaOH followed by 1 mL chloroform to induce phase separation (see Note 15). 6. After vortexing thoroughly and centrifuging (at 1000g for 15 min), a 1-mL aliquot of the upper phase is aspirated for liquid scintillation counting.

4. Notes 1. Amino acids are physiological inhibitors of autophagy (21). The choice of amino acid used during the pulse chase is important, because some of them, such as leucine, are potent inhibitors of autophagy. Valine is frequently used because


2. 3.

4. 5. 6. 7. 8. 9. 10.

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this amino acid does not interfere with autophagy in most cell types. A cocktail of amino acids (with the following final concentrations of amino acids (in μM: asparagine: 60; isoleucine: 100; leucine: 250; lysine: 300; methionine: 40; phenylalanine: 50; proline: 100; threonine: 180; tryptophan: 70; valine: 180; alanine: 400; aspartate: 30; glutamate: 100; glutamine: 350; glycine: 300; cysteine: 60; histidine: 60; serine: 200; tyrosine: 75; ornithine: 100 corresponding to concentrations four times greater than those in the portal vein of starved rats) also very potently inhibits autophagy. HBSS is used when the cells are incubated in a humidified chamber at 37°C in the absence of CO2 ; otherwise EBSS is used. 3-MA blocks autophagy by inhibiting class III phosphatidylinositol 3-kinase (22). However, it should be kept in mind that 3-MA is a phosphatidylinositol 3-kinase inhibitor (23), which interferes with other intracellular trafficking pathways dependent on phosphatidylinositol 3-kinases (24). 3-MA also affects some other intracellular events (25). Cell pellets, resuspended in methanol, can be stored for up to 15 d at –80°C. Dried lipids can be stored for up to 30 d at –80°C. Decanting water around the tube walls will make it easier to collect of the residues of samples spread by fumes during heating. Approximately 1 × 106 HT-29 cells correspond to 30–40 nmol of lipid phosphate. The DOPG stock solution should contain 20 mg/mL or 27 mM of DOPG in chloroform. If the DOPG is dried as a pellet at the bottom of the tube, it is more difficult to dissolve as an aqueous solution. To resuspend all the lipids contained in the tubes, take a sufficient volume of solvent. When taking 20-μL aliquots out of 80 μL, it is crucial not to let the solvents evaporate since this would modify the concentration of lipids. The remaining lipids can be dried and saved for another run, if this is required. It is necessary to preclean the silica plate in acetone for 2 h before use, and let it dry completely before applying the samples. Prepare a well-sealed chamber at least 12 h before use to achieve equilibrium between liquid-vapor phases of the solvent mixture. Since saturating conditions are important during the run, it is essential to leave the tank open as briefly as possible while inserting the plate. Radioactivity associated with ceramide 1-32 phosphate is represented by two spots with Rf values of 0.47–0.41 corresponding to ceramides with different acyl chain length and/or dihydroceramides. Determining sphingosine kinase activity in different concentrations of Triton X-100 may makes it possible to discriminate between the activities of SPHK1 and SPHK2. High concentrations (e.g., 0.5%) of Triton X-100 have been reported to inhibit SPHK2 activity, whereas they stimulate SPHK1 activity (16). The range of protein concentrations giving a linear response for enzyme activity is cell type–dependent.

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15. The extraction of S1P from cell lysates or culture medium and their separation from sphingosine can be performed according to the procedure reported by Vessey et al., which also uses alkaline conditions (26).

Acknowledgments We are grateful to Stuart M. Pitson, Brian Wattenberg, and Tamotsu Yoshimori for providing us with cDNAs encoding SPHK1 and GFP-LC3, respectively.

References 1. Spiegel, S., and Milstien, S. (2003). Sphingosine-1-phosphate: An enigmatic signaling lipid. Nat. Rev. Mol. Cell Biol. 4, 397–407. 2. Ogretmen, B. and Hannun, Y. A. (2004). Biologically active sphingolipids in cancer pathogenesis and treatment. Nat. Rev. Cancer 4, 604–616. 3. Klionsky, D. J. and Emr, S. D. (2000). Autophagy as a regulated pathway of cellular degradation. Science 290, 1717–1721. 4. Levine, B. and Yuan, J. (2005). Autophagy in cell death: an innocent convict? J. Clin. Invest. 115, 2679–2688. 5. Lavieu, G., Scarlatti, F., Sala, G., et al. (2006). Regulation of autophagy by sphingosine kinase 1 and its role in cell survival during nutrient starvation. J. Biol. Chem. 281, 8518–8527. 6. Scarlatti, F., Bauvy, C., Ventruti, A., et al. (2004). Ceramide-mediated macroautophagy involves inhibition of protein kinase B and up-regulation of Beclin 1. J. Biol. Chem. 279, 18384–18391. 7. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000). LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728. 8. Mizushima, N. (2004). Methods for monitoring autophagy. Int. J. Biochem. Cell Biol. 36, 2491–2502. 9. Seglen, P. O. and Gordon, P. B. (1982). 3-Methyladenine: specific inhibitor of autophagic/lysosomal protein degradation in isolated rat hepatocytes. Proc. Natl. Acad. Sci. USA 79, 1889–1892. 10. Bligh, E. G. and Dyer, W. J. (1959). A rapide method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. 11. Hokin, L. E. and Hokin, M. R. (1959). Diglyceride phosphokinase: an enzyme which catalyzes the synthesis of phosphatidic acid. Biochim. Biophys. Acta 31, 285–287. 12. Preiss, J., Loomis, C. R., Bishop, W. R., Stein, R., Niedel, J. E. and Bell, R. M. (1986). Quantitave measurement of sn-1,2-diacylglycerols present in platelets, hepatocytes, and ras-, and sis-transformed normal rat kidney cells. J. Biol. Chem. 261, 8597–8600.


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13. Perry, D. K. and Hannun, Y. A. (1999). The use of diglyceride kinase for quantifying ceramide. Trends Biochem. Sci. 24, 226–227. 14. Van Veldhoven, P. P., Bishop, W. R., Yurivich, D. A. and Bell, R. M. (1995). Ceramide quantitation: evaluation of a mixed micellar assay using E. coli diacylglycerol kinase. Biochem. Mol. Biol. Int. 36, 21–30. 15. Kohama, T., Olivera, A., Edsall, L., Nagiec, M. M., Dickson, R. and Spiegel, S. (1998). Molecular cloning and functional characterization of murine sphingosine kinase. J. Biol. Chem. 273, 23722–23728. 16. Liu, H., Sugiura, M., Nava, V. E., et al. (2000). Molecular cloning and functional characterization of a novel mammalian sphingosine kinase type 2 isoform. J. Biol. Chem. 275, 19513–19520. 17. Olivera, A., Barlow, K. D. and Spiegel, S. (2000). Assaying sphingosine kinase activity. Methods Enzymol. 311, 215–223. 18. Olivera, A. and Spiegel, S. (1998). Sphingosine kinase. Assay and product analysis. Methods Mol. Biol. 105, 233–242. 19. Gijsbers, S., Van der Hoeven, G. and Van Veldhoven, P. P. (2001). Subcellular study of sphingoid base phosphorylation in rat tissues: evidence for multiple sphingosine kinases. Biochim. Biophys. Acta 1532, 37–50. 20. Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. 21. van Sluijters, D. A., Dubbelhuis, P. F., Blommaart, E. F. and Meijer, A. J. (2000). Amino-acid-dependent signal transduction. Biochem. J. 351(Pt 3), 545–550. 22. Petiot, A., Ogier-Denis, E., Blommaart, E. F., Meijer, A. J. and Codogno, P. (2000). Distinct classes of phosphatidylinositol 3’-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells. J. Biol. Chem. 275, 992–998. 23. Blommaart, E. F., Krause, U., Schellens, J. P., Vreeling-Sindelarova, H. and Meijer, A. J. (1997). The phosphatidylinositol 3-kinase inhibitors wortmannin and LY294002 inhibit autophagy in isolated rat hepatocytes. Eur. J. Biochem. 243, 240–246. 24. Punnonen, E. L., Marjomaki, V. S. and Reunanen, H. (1994). 3-Methyladenine inhibits transport from late endosomes to lysosomes in cultured rat and mouse fibroblasts. Eur. J. Cell Biol. 65, 14–25. 25. Tolkovsky, A. M., Xue, L., Fletcher, G. C. and Borutaite, V. (2002). Mitochondrial disappearance from cells: a clue to the role of autophagy in programmed cell death and disease? Biochimie 84, 233–240. 26. Vessey, D. A., Kelley, M. and Karliner, J. S. (2005). A rapid radioassay for sphingosine kinase. Anal. Biochem. 337, 136–142.

12 Molecular Links Between Autophagy and Apoptosis Iwona A. Ciechomska, Christoph G. Goemans, and Aviva M. Tolkovsky

Summary Macroautophagy (herein referred to as autophagy) contributes to the control of life and death throughout the animal and plant kingdoms. Bilateral links have been found between apoptosis and autophagy where inducers of apoptosis also induce autophagy and vice versa. In some cases, autophagy delays the onset of apoptosis and thus prolongs life although it may also promote apoptosis and other forms of cell death. It is thus of great biological and medical interest to understand the molecular connections between these two pathways, and try to utilize—or block—them selectively to aid induction of cell death (e.g., cancer cells) or inhibit death (e.g., in degenerative disorders). This chapter describes methods for studying apoptotic induction of autophagy and its effects on cell function. We also discuss potential pitfalls. Although cell lines are used as model systems, the substances and methods described here can be applied to primary cells and tissues.

Key Words: Apoptosis; autophagy; Bax; caspase; LC3; PARP cleavage.

1. Introduction Although macroautophagy (henceforth referred to as autophagy) is activated primarily as a response to nutritional deprivation (1,2), especially through lack of nitrogen/amino acids, autophagy is also induced by many—if not all— apoptotic stimuli (3–6). The mechanisms by which apoptotic stimuli activate autophagosome formation are not well defined, but our studies on neurons have established that amino acid starvation—a classical method of inducing autophagy especially in liver, heart, and kidney (7,8)—is not necessary (5). In fact, apoptotic stimuli that cause DNA damage (5), ER stress (9), or the ones that originate as extrinsic signals (Fas, tumor necrosis factor [TNF]-) (10) From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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also promote autophagosome formation. Moreover, at least for induction of autophagy by TNF-, no new protein synthesis is required as autophagy is activated in the presence of protein synthesis inhibitors (I. A. Ciechomska et al., unpublished data). Importantly, Beclin1, a key intermediate in the transduction of proautophagic signals to autophagosome formation, binds and alters the properties of some of the antiapoptotic Bcl2 family members and vice versa (11). Moreover, ER stress, which is a key mechanism of apoptosis induction, is also a mechanism for inducing autophagy (12–15). In this chapter, methods of apoptosis induction that produce autophagy and measurement of both in cell culture systems are described. Although the focus is on adherent cells, where detailed microscopic studies can be obtained, induction of apoptosis can be carried out on many types of nonadherent cells, as well as in isolated tissues and whole organisms (plants and animals). It is important to note that some cancer cell lines are highly resistant to apoptosis (e.g., some MCF7 lines lack expression of caspase 3). Hence, it is crucial to verify that apoptosis has been induced by the treatment being used before connections to autophagy can be considered. 2. Materials 1. Cell lines for studying apoptotic induction of autophagy (HeLa, 293, Cos7, MCF7) can be obtained from American Type Culture Collection (ATCC, www.atcc.org) or European Collection of Cell Cultures (ECACC, www.hpa.org.uk/business/ecacc.htm). Recommended growth conditions are provided with each batch as well as the passage number. It is important to restrict experiments to a limited number of passages as the genetic status of cells drifts on prolonged propagation. 2. For the above cells, use Dulbecco’s modified Eagle’s medium (DMEM, 5–10 mM d-glucose) supplemented with 10% fetal bovine serum (FBS) and antibiotics (100 U/mL penicillin and 100 μg/mL streptomycin) (all from SigmaAldrich, www.sigmaaldrich.com; or Invitrogen, www.invitrogen.com). 3. Staurosporine (Sts, www.sigmaaldrich.com) is a potent kinase inhibitor that induces the intrinsic mitochondrial pathway of apoptosis in most cell types. Prepare a 1 mM stock in dimethyl sulfoxide (DMSO); store in aliquots at –20ºC. 4. Etoposide (www.sigmaaldrich.com) is representative of proapoptotic DNAdamaging agents (e.g., cisplatin and cytosine arabinoside). Prepare a 50–100 mM stock in DMSO; store in aliquots at –20ºC. 5. Thapsigargin (Tg, a sarco/endoplasmic reticulum Ca2+ ATPase [SERCA] inhibitor that causes release of Ca2+ from the ER) and tunicamycin (Tm, which disrupts N-glycosylation of newly synthesized proteins in the ER) promote ER stress-induced apoptosis. Prepare stocks of 2 mM Tg and 10 mg/mL Tm in DMSO and store aliquots at –20ºC. 6. TNF- not only activates the extrinsic pathway of apoptosis through the death-inducing signaling complex (DISC) and caspase 8 but also activates a

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protective mechanism via induction of nuclear factor (NF)-B. Therefore, it is commonly applied together with the protein synthesis inhibitor cycloheximide to suppress NF-B expression. Prepare stocks of 100 ng/μL TNF- (Peprotech, www.peprotech.com) as instructed. Store in aliquots at –80ºC. Cycloheximide (www.sigmaaldrich.com) is dissolved at 10 mg/mL in medium or water. Store at –20ºC. UV-C irradiation is obtained using a Stratalinker® UV Crosslinker (www.stratagene.com). Vinblastine activates autophagy so is a useful positive control. Dissolve at 10–50 mM in DMSO; store in aliquots at –20ºC. Earle’s or Hanks basal salt solution (EBSS, HBSS; www.sigmaaldrich.com) is used to activate autophagy by amino acid starvation. Hoechst 33342 and propidium iodide (PI, www.sigmaaldrich.com). Prepare stocks at 100 μg/mL in water and store at 4ºC. More concentrated stocks can be prepared and stored at –20ºC. Boc-aspartyl(O-methyl)fluoromethylketone (BAF, www.mpbio.com) is one of a family of cell-permeable caspase inhibitors that can be used to demonstrate that apoptosis is indeed occurring. Prepare 50 mM stocks in DMSO, aliquote, and store and –20ºC. Use at 50–100 μM, but beware that these inhibitors may target other cysteine proteases (e.g., cathepsins) differentially depending on concentration. Bafilomycin A1 (Baf A1, www.biomol.com) alkalinizes the lysosomal lumen by inhibiting the vacuolar-type (V-type) H+ ATPase. Stock at 1 μM in DMSO; store at –20ºC. Cell lysis buffer (recipe is also good for collecting phosphoproteins): 50 mM Tris-HCl, pH 7.4, 1% NP-40; 120 mM NaCl, 1 mM EDTA, 25 mM sodium fluoride, 40 mM sodium glycerophosphate, 1 mM sodium orthovandate, 1 mM benzamidine, 5 mM sodium pyrophosphate, protease inhibitor cocktail (e.g., Complete™ , Roche, www.roche.com). 4X loading buffer: 50 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 100 mM dithiothreitol (DTT), and 0.01% bromophenol blue. Caspase dilution buffer: 10 mM HEPES, pH 7.4, 42 mM KCl, 5 mM MgCl2 , 1 mM DTT, 1 μg/mL pepstatin A, 1 μg/mL leupeptin, 5 μg/mL aprotinin, and 0.1% Triton X-100. Caspase assay buffer: 20 mM HEPES, pH 7.4, 1 mM EDTA, 10% sucrose, 5 mM DTT, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1propanesulfonate (CHAPS), and 15 μM DEVD-AMC (Asp-Glu-Val-Aspaminomethylcoumarin).

3. Methods Apoptosis can be induced by a variety of compounds or treatments depending on cell type and receptor/metabolic repertoire. For this reason, to demonstrate generalized (universal) mechanisms, it is important to use several

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compounds or treatments and different cell types. Each of the treatments described here initiates apoptosis through a different mechanism although all of these ultimately converge on the mitochondrial pathway that comprises induction/activation of BH3 only proteins; activation of Bax and/or Bak; mitochondrial outer membrane permeabilization; release of cytochrome-c from the intermembrane space; activation of Apaf-1, caspase 9, and downstream caspases; and cleavage of multiple substrates. Key caspase substrates are acinus and caspase-activated DNase (CAD), which cause DNA clumping and fragmentation (16,17), and poly-ADP-ribose polymerase (PARP), which reports on caspase 3/7 activities within the cell (18–20). Evidence that apoptosis is occurring in intact cells is obtained by monitoring translocation of Bax to mitochondria using, for example, conformation-specific antibodies (21) or GFPtagged Bax (not described here), release of cytochrome-c from mitochondria (5,22–24), nuclear morphological changes using DNA-binding dyes (e.g., see ref. 5), and phosphatidylserine (PS) exposure using fluorescent Annexin V staining (25). Biochemical events monitored by immunoblotting include PARP cleavage by immunoblotting and caspase activity using fluorogenic substrates. Concomitant autophagosome formation in cells can be demonstrated by recording the formation of LC3-II-dependent puncta (using fluorescent fusion protein tags or antibodies) and by immunoblotting for LC3-II. A parallel assay using an inhibitor of autolysosomal-mediated degradation of LC3-II (26,27) (e.g., by inhibition of the V-type H+ ATPase with Baf A1) is crucial to demonstrate that productive autophagy, namely, LC3 turnover, is occurring. Otherwise, LC3-II puncta might be accumulating because there is a block in the formation of autolysosomes or in lysosomal proteolytic activity. 3.1. Induction of Apoptosis 1. Seed cells into 24-wells containing glass cover slips (13 mm, #0–1 thickness, or glass-bottom dishes, www.glass-bottom-dishes.com) at a density that will give 70–80% confluence the next day (see Notes 1 and 2). For biochemistry, seed proportionately in six-well plates (see Note 3). 2. Replace growth medium with medium containing the apoptotic inducer. Dose efficacy should be tested for each cell type at different times after addition as described below. a. Sts: efficacy range 0.1–2 μM. Note that high concentrations can cause necrosis and that Sts can alter cell shape independently of apoptosis due to cytoskeletal deformation. b. Etoposide: efficacy range 20–200 μM. c. Tg and Tm: efficacy range 1–4 μM Tg and 5–20 μg/mL Tm. In some cells, these can be used separately to induce apoptosis, but we find them especially efficacious when they are added together.


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d. TNF- and cycloheximide: efficacy range 5–100 ng/mL TNF-, 5–20 μg/mL cycloheximide. e. UV: efficacy range 100–600 J/m2 . Replace medium with sterile PBS. Place plate/dish on bottom center of Stratalinker chamber, remove lid from wells to be irradiated (UV does not penetrate tissue culture plastic), and activate irradiation protocol as instructed after carefully calculating the time required to give the energy required. Replace the lid immediately. We usually activate the protocol once or twice before submitting the cells to irradiation as this also sterilizes the chamber.

3.2. Quantification of Apoptosis 3.2.1. Analysis of Nuclear Morphology Nuclear condensation and fragmentation during apoptosis can be quantified by co-staining with two nuclear dyes: Hoechst 33342 and PI. Because PI is a membrane-impermeable dye, it only stains the nucleus of dead cells that have lost plasma membrane integrity, that is, cells that have undergone necrosis or secondary necrosis (following on from apoptosis). Hoechst 33342 is membrane permeable so it stains live cells. Nuclei of healthy cells are relatively dimly stained by Hoechst and are not stained by PI. Nuclei of apoptotic cells are fragmented or highly condensed and display bright blue fluorescence due to Hoechst staining. Occasionally, nuclei will appear pink, indicating that the plasma membrane is beginning to be permeable to PI. Necrotic cells show a bright red fluorescence staining with PI, with normal or slightly condensed size. If in doubt, use phase microscopy to examine whether the plasma membrane is shattered as these cells are phase dark. Note that this method cannot be used for fixed cells as PI will stain live as well as dead cells. 1. At the indicated time after treatment, add Hoechst 33342 and PI (1–5 μg/mL final concentration of each) directly to the growth medium. 2. Incubate cells for 10 min at 37ºC. 3. Examine cells by fluorescence microscopy using a UV filter (340–380 nm excitation, 425 nm emission) and count the number healthy and apoptotic bluestained cells and PI-positive red dead cells in at least four randomly selected fields, at least 200 cells in total. 4. Calculate the percentage of apoptotic and necrotic cells out of the total (apoptotic + necrotic + healthy) cells counted.

3.2.2. PARP Cleavage PARP, a 113-kDa protein in humans, binds specifically to DNA strand breaks and is a substrate for caspase 3, which is activated during apoptosis in most cell types. However, in MCF7 cells lacking caspase 3, caspase 7 cleaves

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PARP albeit inefficiently. Caspase 3 cleaves PARP into two fragments of approximately 89 and 24 kDa. Thus, detection of the 89-kDa PARP fragment serves as a biochemical marker of apoptosis. To detect PARP in untreated cells and after exposure to drugs: 1. Collect healthy and dead cells from a six-well plate by scraping them off the dish in the culture medium using the plunger of a 1-mL syringe. 2. Centrifuge at 380×g for 5 min; resuspend cells in 1 mL ice-cold PBS (to remove serum remnants), transfer cell suspension to a microfuge tube, and pellet cells again. 3. After removing PBS, add 100–150 μL cell lysis buffer. Keep on ice for 20–30 min with occasional vortexing to swell the cells and facilitate protein extraction. 4. Centrifuge the lysate at 13,000×g at 4ºC for 10 min. Transfer the supernatant to a fresh microcentrifuge tube and discard the pellet (containing nuclei; transcription factors will be extracted but histones remain with the pellet). 5. Estimate the amount of total protein in 2–5 μL of each sample using a bicinchoninic acid assay (BCA). BCA protein assay kit (www.sigmaaldrich.com) is used according to the manufacturer’s instructions. 6. Add the appropriate amount of 4X gel loading buffer to each sample and heat at 98ºC for 5–10 min. 7. Separate proteins (at least 20 μg per lane) on a 10% denaturing polyacrylamide gel (SDS–PAGE). 8. Transfer the separated proteins to a nitrocellulose/PVDF filter by electroblotting. 9. Stain the filter with Ponceau Red solution (1% Ponceau S [www.sigmaaldrich.com] in 5% acetic acid/water) for 5 min to visualize protein bands. 10. Rinse the membrane in water until protein bands are distinct and mark the position of the molecular weight markers with a ballpoint pen or pencil. Scan, photograph, or photocopy the blot. Image may be used to demonstrate equality of protein loading. 11. Cut membrane across into two pieces at the approximately 70 kDa marker. PARP (113 and 89 kDa) is on the upper piece and -tubulin (∼50 kDa) is on the lower piece of membrane. Detection of -tubulin is a useful loading control for immunoblotting. If you are using an anti-PARP monoclonal antibody, ensure that it recognizes the 89-kDa fragment, namely that the epitope is not within the 24-kDa fragment. 12. After blocking with 5% nonfat milk in TBST (Tris-buffered saline/Tween20: 10 mM Tris-HCl, pH 7.4; 150 mM NaCl; 0.1% Tween-20), incubate the membranes overnight at 4ºC with primary antibodies diluted in blocking buffer or TBST. Antibodies can be reused several times if stored in the presence of 0.02% sodium azide. 13. Wash with TBST four times for 5 min each and incubate for 1 h at room temperature with appropriate horseradish peroxidase (HRP)-conjugated anti-speciesspecific IgG. For best results, choose antibodies that have been subtracted against


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all species other than the species in which the first antibody was raised (e.g., see www.jacksonimmuno.com). Note that HRP is potently inhibited by sodium azide. 14. Visualize immunocomplexes using an enhanced chemiluminescence detection system (ECL, www.amershambiosciences.com). 15. After exposure of blots to X-ray film, bands can be digitized using a scanner. To obtain meaningful data, blots must not be overexposed, and band intensities should be in the linear range of the densitometric sensitivity of the film. This can be tested by running increasing amounts of lysates on parallel lanes and scanning the X-ray film. The range of linearity of the scanner also has to be determined. For this purpose, special calibration strips are commercially available (www.kodak.com). Density can be quantified using ImageJ (NIH Image) analysis system (open source software at http://rsb.info.nih.gov/ij/). See Fig. 1 for image of blot probed for PARP cleavage.

3.2.3. Caspase 3 Activity Caspase 3 is one of the key executioners of apoptosis, being responsible for the proteolytic cleavage of many key proteins, including PARP. Caspase 3 activity can be sensitively assessed by measuring the cleavage of the fluorogenic substrate Ac-DEVD-AMC (N-acetyl-Asp-Glu-Val-Aspaminomethylcoumarin); the signal is emitted when AMC is released on cleavage. Specificity can be determined by adding excess DEVD aldehyde (DEVD-CHO), a high-affinity reversible inhibitor, to a parallel cell extract. Indeed, the DEVD amino acid sequence is derived from the caspase 3 cleavage site in PARP. The same protein extracts used for immunoblotting can be used for caspase activity measurement as the protease inhibitor mix Complete in the extraction buffer does not seem to interfere with the caspase activity assay. A key requirement is the presence of DTT as caspases are cysteine proteases that require a reduced thiol group for activity. Measurement of caspase 3 activity with this substrate is best performed in a 96-well microplate format using a fluorometric microplate reader. 1. Add 20 μg protein extract to ice-cold caspase dilution buffer to produce a final volume of 50 μL. Alternatively, cells can be lysed directly in the caspase dilution buffer by sonication on ice. 2. Transfer each sample to a 96-well microplate. 3. Prepare blank without protein extract (only 50 μL hypotonic buffer) and an AMC calibration standard (0.5–10 μM AMC) to determine the linear range of detection. 4. To each well, add 150 μL of ice-cold caspase assay buffer. 5. Cleavage of DEVD-AMC (intensity of fluorescence) is measured at 340 nm (excitation) and 460 nm (emission) in the microplate reader. Record fluorescence at 5- to 10-min intervals for up to 120 min (for low protein concentrations, one

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can measure up to 24 h as long as the readings are linear). See Fig. 2 for caspase assay results. 6. Analyze the initial rate (slope) of fluorescence signal along the linear portion of the curve depicting fluorescence accumulation. 7. To ensure that the fluorescence is due to caspase activity, add 5- to 10-fold excess DEVD-CHO over DEVD-AMC to parallel samples.


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3.2.4. Bax Activation As discussed above, Bax is a key mediator of apoptosis induced through the mitochondrial pathway on which both extrinsic and intrinsic signals converge. On apoptotic stimulation, Bax forms oligomers and translocates from the cytosol to the mitochondrial outer membrane and increases the membrane’s permeability, which leads to the release of apoptogenic factors, such as cytochrome-c. Active Bax can be distinguished from the inactive cytosolic form using conformation-specific antibodies that bind to the N-terminal domain of the protein. The monoclonal antibodies described here were originally raised in Dr. R. J. Youle’s laboratory (28) and are available from commercial sources (e.g., www.labvision.com). Polyclonal antibodies that recognize the exposed N-terminus are also available (www.scbt.com). Note that the same immunostaining protocol described here for Bax can be used to examine cytochrome-c release from mitochondria, where cytochrome-c displays a punctate staining pattern when confined to mitochondria, but a diffuse cytoplasmic staining pattern when it is released into the cytoplasm (5). See Fig. 3 for example of cytochrome-c and active Bax staining. 1. Treat cells with proapoptotic drugs in the presence of 50–75 μM BAF to prevent caspase-dependent execution of cell death as this may cause the cells to dislodge from the plate. BAF does not prevent Bax translocation (but see Note 4 for an important exception with regard to TNF-). Fix cells in 3% paraformaldehyde in PBS for 15 min at room temperature.

Fig. 1. (A) Immunoblot for LC3-II and poly-ADP-ribose polymerase (PARP) from HeLa cells treated with 1 μM staurosporine (Sts) for 15 h in the absence or presence of 20 nM bafilomycin A1 (Baf A1). Postnuclear extracts were separated on a 10% (PARP, tubulin) or a 15% (LC3) gel. Immunoblots were probed for PARP (here we used a monoclonal antibody from Drs. Said Aoufouchi and Sidney Shall but have also successfully used PARP Antibody #9542; www.cellsignaling.com). LC3 was probed with an antibody raised by Dr. E. Kominami, but we have also successfully used the monoclonal antibody 5F10 from www.nanotools.de. -Tubulin was probed with T5168 from www.sigmaaldrich.com. Note that STS induces an increase in LC3-II whose intensity increases further in the presence of Baf A1. The intensity of LC3-I also increases possibly because some LC3-II that cannot be consumed by the lysosomes is regenerated into LC3-I (note that Baf A1 is slightly toxic to the cells when incubated for long times). (B) Images of Cos7 cells transfected with GFP-LC3, treated as indicated in the figure. were captured using a PerkinElmer™ UltraView system (6). (C) Autophagy induced by apoptotic stimuli compared with proautophagic stimuli in HeLa, Cos7, and MCF7 cells after transfection with GFP-LC3.

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Fig. 2. Caspase 3 activity measured in HeLa cells treated with the indicated proapoptotic factors using DEVD-AMC and a fluorescent microplate reader. (A) Raw traces. (B) Rates calculated from the early portion of the slopes in (A). 2. Wash cells twice with PBS and permeabilize the cells in PBS containing 0.1% CHAPS (see Note 5). 3. Dilute the anti-Bax antibody (0.4 μg/mL) in PBS-CHAPS, add to the cells, and incubate for 1 h at room temperature. Note that the monoclonal antibodies described in ref. 28 are species specific. 4. Wash in PBS four times (we dip the cover slips sequentially in four pots of PBS) and incubate with the appropriate secondary antibody conjugated to a fluorescent reporter (e.g., anti-mouse FITC). 5. Wash the cells and stain nuclei with Hoechst 33342 (1–5 μg/mL) added for 5 min at room temperature. 6. Dip cover slips in water and mount face down onto slides using an appropriate mounting medium such as Fluoromount-G (a water-soluble permanent mounting medium that suppresses photobleaching, www.southernbiotech.com) and analyze by fluorescence (or even better, by confocal) microscopy. 7. Count a minimum of 100 cells that display Bax translocation from a few separate visual fields, typically revealed by intense punctate staining (see Note 6 for method that confirms co-localization of Bax and mitochondria). Determine the percentage of cells displaying active Bax.

3.2.5. Detection of a Subdiploid Population by Fluorescence-Activated Cell Sorting Although detection of apoptotic cells by method in Subheading 3.2.1 is fast after some practice, counting cells under the microscope is painstaking, which can be avoided using fluorescence-activated cell sorting (FACS). One


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Fig. 3. Cells stained for active Bax and cytochrome-c release. Top row shows cultured rat sympathetic neurons from which nerve growth factor (NGF) has been withdrawn for 15 h. Cells were stained with rat-specific anti-Bax monoclonal antibody 1D1 followed by anti-mouse Alexa488. DNA was stained with Hoechst 33342. Note that the cell undergoing DNA fragmentation (arrow) is the only Bax-positive cell. Middle row shows HeLa cells treated with etoposide in the presence of BAF for 21 h. Antiactive Bax was probed with monoclonal antibody 6A7 followed by anti-mouse Cy3. Note that the cells indicated express active Bax although the nuclei have not fragmented due to the presence of BAF. Bottom row shows the same set of HeLa cells probed with anti-cytochrome-c (cyt-c). The cell indicated shows diffuse cytoplasmic staining for cytochrome-c, whereas the adjacent cells show cyt-c localized to mitochondria.

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of the characteristics of apoptosis is the degradation of DNA due to the activation of several endonucleases, some of which are activated by caspases. Fixation of cells with precipitating fixatives (such as ethanol) causes the leakage of the cleaved low-molecular-weight DNA fragments that are produced during apoptosis. As a consequence, apoptotic cells can be identified as a hypodiploid peak (sub-G1 peak), whereas healthy cells generate a typical cell cycle histogram (without sub-G1 peak). See ref. (30) for detailed information on using FACS to analyze apoptosis. 1. After induction of apoptosis, harvest cells in the appropriate manner (trypsinize adherent cells) and prepare a single-cell suspension in PBS. 2. Wash cells twice with PBS (centrifuge 380×g for 5 min) and resuspend at 1 × 106 cells in 200 μL. 3. Fix cells in cold 70% ethanol (2 mL) by adding it dropwise to the cell pellet while vortexing. This should ensure fixation of all cells and minimize clumping. 4. Fix cells for at least 30 min at 4°C. Specimens can be left at this stage at –20ºC for several weeks. 5. Wash cells twice with PBS. Spin at 380×g and be careful to avoid cell loss while discarding supernatant especially after spinning out of ethanol (it may be necessary to centrifuge cells at a slightly higher “g” to pellet after ethanol fixation as the cells become flocculent). 6. Resuspend pellet in 1 mL staining solution containing 50 μg/mL PI and 100 μg/mL RNase A. RNase A should be boiled for 10 min to inactivate any contaminating DNase. RNase A is used to eliminate all free RNA, so that PI will only stain cellular DNA (see Note 7 for the often used complementary method of Annexin V staining). 7. Incubate for 30 min at room temperature. 8. If cells are clumped, pass them through a 25-gauge needle using a 1-mL syringe. 9. Store samples at 4ºC until analyzed by flow cytometry (within 24 h). 10. Collect a minimum of 20,000 events per datafile.

3.3. Analysis of Autophagy The term macroautophagy comprises two processes: (1) formation of autophagosomes and the transport of vesicles containing cargo and (2) lysosomal degradation of the cargo after fusion of the autophagosomes with endo/lysosomes (also called amphisomes or autolysosomes; see refs. 31 and (32)). As with apoptosis, which is regulated at several levels (secondmessenger signaling, mitochondrial events, postmitochondrial events, etc.), one must distinguish between the different phases of autophagy-execution while assessing the interaction between apoptosis and autophagy. To this end, it is important to recognize the molecular principles of each assay. Here, we restrict ourselves to the analysis of microtubule-associated protein light-chain-3


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(MAP-LC3) commonly named LC3. During the formation of autophagosomes, LC3 is proteolytically cleaved and lipidated. This LC3–phospholipid conjugate (LC3-II) is localized on autophagosomes and autolysosomes (prior to LC3-II digestion by lysosomal enzymes or removal from the membrane due to cleavage of the conjugated lipid; see ref. (33)), forming numerous small puncta (34). We describe measurement of LC3-II formation by techniques similar to those we have described for apoptosis. It should be noted, however, that despite the advent of molecular markers for autophagy, demonstration of autophagosome formation and maturation by electron microscopy is still an indispensable technique, for example as practiced in ref. (36). There are also important pitfalls in the analysis of autophagy: in a system in which productive macroautophagy is running at a high pace, there may be even lower steady state levels of LC3-II than in unstimulated cells and this might lead to the false conclusion that cells are hardly undergoing autophagy. Hence, as pointed out by Tanida et al. (33), the true measurement of autophagic flux requires a control where lysosomal degradation of LC3-II is prevented. We use Baf A1 but cysteine and asparate protease inhibitors E64d and pepstatin are also efficacious (27). See also Subheading 3.4. for discussion regarding use of inhibitors of autophagy that impact on apoptosis and vice versa. 3.3.1. Quantification of Autophagosomes by Detection of GFP-LC3 Punctation To determine whether apoptotic stimuli activate autophagosome formation, cells can be transfected with GFP-tagged LC3 plasmid and the change in the distribution of GFP-LC3 from a diffuse cytoplasmic pattern to a punctate pattern after induction of apoptosis can be documented. The mammalian expression plasmid for GFP-LC3 is available through the Addgene plasmid collection (www.addgene.org). As a positive control for autophagic stimulation, cells can be treated with vinblastine (31,35), starved of amino acids, or treated with ammonium chloride (6). 1. Transfect cells (grown on glass cover slips) with GFP-LC3 using appropriate reagents. 2. After 24 h, treat the cells with an apoptotic stimulus. Stain the nuclei with Hoechst 33342 and analyze GFP fluorescence by microscopy. To prevent the loss of cells due to execution of cell death, treat cells in the presence of a pan-caspase inhibitor, such as 50 μM BAF. 3. Treat a cohort of cells with 20 nM Baf A1 to examine the net flux of LC3-II through the autophagic pathway. 4. Prepare a positive control for autophagy induction (GFP-LC3 punctation) by treating cells with 50 μM vinblastine or 20 mM ammonium chloride (1–2 h), or

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starve cells by washing them five times with EBSS or HBSS and incubating them in this medium for 1–2 h. 5. Determine the percentage of GFP-LC3-positive cells that contain GFP-LC3 puncta (see Fig. 3), counting at least 100 cells per sample. 6. Alternatively, stain nuclei with Hoechst 33342 and assess the number of autophagosomes per cell, counting GFP-LC3-positive puncta per nucleus. In cells that have high basal autophagy, this technique may be difficult.

3.3.2. Quantification of LC3-II by Immunoblotting Lipidated LC3-II migrate more rapidly (∼16 kDa) than LC3-I (∼18 kDa) when proteins are separated by SDS–PAGE. Hence, the amount of LC3-II can be used to indicate autophagic activity with the proviso that a parallel experiment is conducted in the presence of lysosomal inhibitors that prevent LC3-II degradation to measure the net extent of autophagic induction (27). 1. After treatment with apoptotic inducers, isolate cell proteins and normalize loading as described in Subheading 3.2.2. 2. Resolve proteins on a 15% denaturing gel (SDS–PAGE) and blot. 3. After blocking with 5 % nonfat milk, probe membranes with anti-LC3. A commercially available monoclonal anti-LC3, which we find very clean, is available from www.nanotools.de. 4. Follow the procedures outlined in Subheading 3.2.2., steps 12–16.

3.4. Use of Inhibitors to Examine the Relationship Between Apoptosis and Autophagy A standard way to address whether autophagy contributes to apoptotic regulation or signaling (e.g., refs. (37–42)) is to inhibit autophagy with chemical inhibitors. 3-Methyl adenine (3-MA) is widely used as a specific inhibitor of autophagy (43), although the authors used the word “specific” to indicate only that it did not inhibit other ATP-dependent processes. 3-MA is a low-affinity PI3-kinase inhibitor that inhibits the class III PI3-kinase Vsp34, which partners Beclin1 and initiates autophagy (44,45). In some reports, other PI3-kinase inhibitors such as wortmannin or LY294002 are used for the same reason. However, our laboratory has demonstrated that 3-MA exhibits—at concentrations used to suppress autophagy—profound inhibitory effects on stress-activated MAPK kinases JNK and p38 that are in many cases key regulators of apoptosis (5). Indeed, 3-MA alters many metabolic processes in cells (46). Moreover, LY294002 and wortmannin are also widely used and potent inhibitors of class I PI3-kinases, which activate an antiapoptotic signaling branch mediated by Akt in many cell types but also inhibit autophagy (47). Therefore, in studies addressing the interplay of apoptosis and autophagy,


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one cannot pay enough attention to the molecular targets of chemical inhibitors whose use otherwise could undermine the purpose of the study. For this reason, it is highly advisable to back up experiments by using a variety of independent inhibitors, or, if possible, to knockdown key regulators in the pathways of autophagy or apoptosis using highly specific RNAi constructs. 4. Notes 1. Glass cover slips should be grease free. Using 100–200 cover slips at a time, we soak cover slips in nitric acid for a few hours in the fume cupboard while slowly rotating on a shaker, then wash them in water to remove all traces of the acid, followed by one wash in methanol (so that they do not stick to each other during drying) and oven-bake in a glass Petri dish at 250ºC overnight. 2. Some cells adhere better to poly-lysine-coated cover slips. Prepare stocks of 1 mg/mL poly-d-lysine in water (e.g., P-0899, Sigma-Aldrich). Keep frozen at −20ºC. For coating cover slips, prepare a 10 μg/mL solution in 0.1 M sodium borate, pH 8.4, immerse the cover slips, and coat from 1 h to overnight at room temperature. Wash three times with water. Air-dry in a sterile cabinet and store at 4ºC. 3. For electron microscopy, it is best to prepare a plastic cover slip made of polychlorotrifluoroethylene such as Aclar® which is consistent with embedding chemicals (www.proscitech.com.au/catalogue/g2.asp). Light microscopy on these cover slips is of low quality. 4. When using TNF-, apoptotic signaling begins by activating caspase 8, and thus, BAF will inhibit the apoptotic signaling from reaching the stage of Bax activation. BAF can be omitted as long as cells are fixed early enough before execution of apoptosis, resulting in blebbing and/or detaching of the cells from the plate. As cell types vary remarkably in the rate of progression through the different stages of the apoptotic program, we recommend to include BAF when possible so that the cells keep beyond the mitochondrial stage of apoptosis. 5. CHAPS in used in lieu of Triton X-100 because Triton—but not CHAPS—can cause Bax to display the N-terminus associated with its activation independently of an apoptotic stimulus (29). Although the cells are fixed, it is best to be safe while performing this assay. 6. It is possible to counterstain mitochondria prior to initiation of apoptosis using a mitochondrial dye that resists paraformaldehyde fixation such as MitoTracker Orange (MTO) (CM-H2TMRos [M7511], www.invitrogen.com). Load cells by adding 0.3–1 μM final concentration of MTO into the growth medium. Incubate for 15–30 min in the incubator. Wash the medium and commence apoptosis by using the appropriate treatment. Note that while using MTO, the secondary antibody used for detection of Bax should not be coupled to TRITC or Cy3. Alternatively, cells can be transfected with a plasmid expressing a mitochondrial-targeted fluorescent protein (available through www.clontech.com) to visualize mitochondria.

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7. Another established marker for apoptosis is PS translocation from the inner leaflet of the plasma membrane to the outer leaflet, which can occur in both caspase-dependent and caspase-independent manners. Externalized PS can be visualized with fluorophore-conjugated Annexin V. Apoptotic cells are those that are Annexin V positive and PI negative (25). When Annexin V is applied to cells, cells must not be fixed or else Annexin V will enter the plasma membrane and stain PS inside the cell (PS is abundant in the inner leaflet of the plasma membrane). See ref. 30 for comprehensive description of analysis of apoptosis by FACS.

Acknowledgments The work described here was supported by PG 064232 from the Wellcome Trust. We thank Nigel Miller, Department of Pathology, Cambridge University, for helping with the FACS protocol. References 1. Yorimitsu, T., and Klionsky, D. J. (2005) Autophagy: molecular machinery for self-eating. Cell Death Differ. 12, 1542–1552. 2. Klionsky, D. J., and Emr, S. D. (2000) Autophagy as a regulated pathway of cellular degradation. Science 290, 1717–1721. 3. Crighton, D., Wilkinson, S., O’Prey, J., et al. (2006) DRAM, a p53-induced modulator of autophagy, is critical for apoptosis. Cell 126, 121–134. 4. Kessel, D., and Reiners, J. J., Jr. (2006) Initiation of apoptosis and autophagy by the Bcl-2 antagonist HA14–1. Cancer Lett. 18, 18. 5. Xue, L., Fletcher, G. C., and Tolkovsky, A. M. (1999) Autophagy is activated by apoptotic signalling in sympathetic neurons: an alternative mechanism of death execution. Mol. Cell Neurosci. 14, 180–198. 6. Bampton, E. T. W., Goemans, C. G., Niranjan, D., Mizushima, N., and Tolkovsky, A. M. (2005) The dynamics of autophagy visualised in live cells; from autophagosome formation to fusion with endo/lysosomes. Autophagy 1, 23–36. 7. Ashford, T. P., and Porter, K. R. (1962) Cytoplasmic components in hepatic cell lysosomes. J. Cell Biol. 12, 198–202. 8. Mizushima, N., Yamamoto, A., Matsui, M., Yoshimori, T., and Ohsumi, Y. (2004) In vivo analysis of autophagy in response to nutrient starvation using transgenic mice expressing a fluorescent autophagosome marker. Mol. Biol. Cell. 15, 1101–1111. Epub 2003 December 29. 9. Momoi, T. (2006) Conformational diseases and ER stress-mediated cell death: apoptotic cell death and autophagic cell death. Curr. Mol. Med. 6, 111–118. 10. Thorburn, J., Moore, F., Rao, A., et al. (2005) Selective inactivation of a Fasassociated death domain protein (FADD)-dependent apoptosis and autophagy pathway in immortal epithelial cells. Mol. Biol. Cell. 16, 1189–1199. 11. Pattingre, S., Tassa, A., Qu, X., et al. (2005) Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122, 927–939.


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13 Clearance of Mutant Aggregate-Prone Proteins by Autophagy Brinda Ravikumar, Sovan Sarkar, and David C. Rubinsztein

Summary The accumulation of mutant aggregate-prone proteins is a feature of several human disorders, collectively referred to as protein conformation disorders or proteinopathies. We have shown that autophagy, a cytosolic, non-specific bulk degradation system, is an important clearance route for many cytosolic toxic, aggregate-prone proteins, like mutant huntingtin and mutant -synucleins. Induction of autophagy enhances the clearance of both soluble and aggregated forms of the mutant protein, and protects against toxicity caused by these mutations in cell, fly, and mouse models. Inhibition of autophagy has opposite effects. Thus, the autophagic pathway may represent a possible therapeutic target in the treatment of certain protein conformation disorders.

Key Words: Autophagy;

aggregate-prone

proteins;

Huntington’s

disease;

rapamycin.

1. Introduction Several human diseases, referred to as protein conformation disorders (PCDs) or proteinopathies, are caused by the accumulation of misfolded, mutant proteins. These conditions include Alzheimer’s disease, Parkinson’s disease (PD), Huntington’s disease (HD), and many of the dominant spinocerebellar ataxias (1). In many cases, the mutation confers a toxic gain-of-function on the target protein and there is a strong correlation between the accumulation of the mutant protein and disease severity (2). An attractive pharmacological approach toward treatment of these PCDs would thus be to enhance the clearance of the toxic mutant proteins. From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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Autophagy is a nonspecific bulk degradation system involved in the clearance of cytosolic long-lived proteins and organelles. During autophagy, doublemembraned structures called autophagic vacuoles enclose a portion of the cytosol with the contents to be degraded. These autophagic vacuoles then fuse with the lysosomes where their contents are degraded by lysosomal hydrolases (3). A key regulator of mammalian autophagy is a phosphatidyl inositol (PI) kinase-related kinase, mammalian target of rapamycin (mTOR) (4). Inhibition of mTOR by treatment with rapamycin results in induction of autophagy. Another kinase involved in the regulation of mammalian autophagy is the class III PI-3-kinase, Vps34. Inhibition of Vps34 by 3-methyl adenine (3-MA) leads to an inhibition of autophagy (5). Furthermore, lysosomal acidification is important for the fusion of autophagic vacuoles with the lysosomes. Thus, autophagy can also be inhibited by using compounds like Bafilomycin A1, which affects the acidification of lysosomes by inhibiting the proton pump (6). Autophagy occurs constitutively in mammalian cells. Inhibition of autophagy by conditional knockout of autophagy-specific genes in the brains of mice resulted in an aberrant accumulation of ubiquitinated protein aggregates reminiscent of those seen in the PCDs (7,8). This suggests the importance of autophagy in the clearance of normal soluble cellular proteins. We have shown that autophagy is an important clearance route for mutant aggregate-prone proteins, like mutant huntingtin that causes HD and mutant -synuclein that causes forms of PD (9–12). Our data suggest that autophagy is clearing soluble forms of the proteins. It is likely that the numbers of aggregates also decrease when autophagy is induced, as this process lowers the concentration of the aggregate precursors—the soluble aggregate-prone species. We used different modulators that act at distinct steps of the autophagic pathway to study aggregation/clearance of a mutant huntingtin fragment and clearance of mutant -synuclein (9–12). We also measured changes in huntingtin-induced cell death with autophagy modulators (9).

2. Materials 2.1. Cell Culture 1. Cell lines: African green monkey kidney cells (COS-7), stable inducible rat pheochromocytoma (PC12) cell lines expressing a variety of proteins; enhanced green fluorescent protein (EGFP)-tagged huntingtin exon-1 fragment with 23 or 74 polyglutamine repeats (EGFP-HDQ23 or EGFP-HDQ74) (13), or hemagglutinin (HA)-tagged A53T or A30P mutants of -synuclein (10). 2. Dulbecco’s modified Eagle’s medium (DMEM) (Sigma-Aldrich, Gillingham, Dorset, UK). 3. Fetal bovine serum (FBS) (Sigma-Aldrich, Gillingham, Dorset, UK).

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Horse serum (Sigma-Aldrich, Gillingham, Dorset, UK). l-Glutamine (Sigma-Aldrich, Gillingham, Dorset, UK). Penicillin/Streptomycin solution (Sigma-Aldrich, Gillingham, Dorset, UK). 1X Trypsin–ethylenediaminetetraacetic acid (EDTA) solution (Sigma-Aldrich, Gillingham, Dorset, UK). Geneticin/G418 sulfate (Gibco Invitrogen, Paisley, UK) is dissolved in sterile water to make a 100 mg/mL stock and frozen at –20°C in aliquots. Hygromycin-B (Merck Chemicals Ltd., Nottingham, UK) solution is mixed with DMEM to a 26 mg/mL stock and aliquots are frozen at –20°C. Doxycycline (Sigma-Aldrich, Gillingham, Dorset, UK) is dissolved in sterile water to make a 100 mg/mL stock and stored at –20°C (see Note 1). 6-well plates (NUNC; Invitrogen, Paisley, UK). 22 × 22 mm thickness No. 1 glass cover slips (VWR international, Lutterworth, UK). 75-cm2 flasks (NUNC; Invitrogen). Cell scrapers (Sarstedt Ltd., Leicester, UK).

2.2. Modulators of Autophagy 1. 3-Methyl adenine (3-MA) (Sigma-Aldrich, Gillingham, Dorset, UK), an inhibitor of autophagy, is dissolved in water to make a 50 mM stock and stored at room temperature (see Note 2). 2. Rapamycin (Rap) (Sigma-Aldrich, Gillingham, Dorset, UK), an autophagy inducer, is dissolved in dimethyl sulfoxide (DMSO) (Sigma-Aldrich, Gillingham, Dorset, UK) to make a 0.5 μg/μL stock, and the aliquots are stored at –20°C in dark Eppendorf tubes to keep away from direct light. 3. Bafilomycin A1 (BafA1) (Sigma-Aldrich, Gillingham, Dorset, UK), an autophagy inhibitor, is solubilised in dimethyl sulfoxide (DMSO) to prepare a 100 μM stock and stored at –20°C. The aliquots are kept in dark Eppendorf tubes to keep away from direct light.

2.3. Transient Transfection 1. Lipofectamine transfection reagent (Invitrogen). 2. Plasmid DNA: EGFP-tagged huntingtin exon-1 fragment with 74 polyglutamine repeats (EGFP-HDQ74) (2). 3. Plasmid DNA: EGFP tagged LC3 (EGFP-LC3; kind gift from T. Yoshimori) (14).

2.4. Immunocytochemistry 1. 1X Phosphate-buffered saline (PBS) solution. 2. Paraformaldehyde (PFA) is dissolved in 1X PBS to make a 4% stock (see Note 3). 3. Antifadent, citifluor AF1 (Citifluor Ltd., London, UK).

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4. 4´,6-Diamidino-2-phenylindole (DAPI) (Sigma-Aldrich, Gillingham, Dorset, UK) is dissolved in citifluor to make a 3 μg/mL stock and stored at 4°C. 5. Nikon Eclipse E600 fluorescence microscope (Nikon, Welwyn GC, UK). 6. Zeiss LSM 510 META confocal microscope (Carl Zeiss, Inc. UK Ltd., Kingston upon Thames, UK).

2.5. Lysis of Mammalian Cells 1. Lysis buffer (2X): 20 mM Tris-HCl pH 6.8, 137 nM NaCl, 1 mM ethylene glycol tetraacetic acid (EGTA), 1% Triton X-100, 10% glycerol; store at 4°C. 1X lysis buffer is prepared fresh with protease inhibitor (see below) just before harvesting cells for lysis. 2. Complete protease inhibitor cocktail tablet (Roche Diagnostics Ltd., Burgess Hill, UK) is dissolved in 2 mL sterile water and stored at –20°C. While making 1X lysis buffer, complete protease inhibitor cocktail solution is added at 1:25 dilution.

2.6. Sodium Dodecyl Sulfide–Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Resolving gel (12%): 30% acrylamide:bis-acrylamide solution, 1.5 M Tris-HCl pH 8.8, 10% SDS, 10% ammonium persulfate (APS) (see Note 4), N,N,N´,N´tetramethyl-ethylenediamine (TEMED) (see Note 5) and distilled water. 2. Stacking gel (5%): 30% acrylamide:bis-acrylamide solution, 1 M Tris-HCl pH 6.8, 10% SDS, 10% APS, TEMED, and distilled water. 3. Water-saturated isobutanol: Take equal volumes of distilled water and isobutanol in a glass bottle, mix in a stirrer overnight, allow to separate and store at room temperature. Use the top (butanol) layer. 4. Prestained molecular weight markers: Kaleidoscope markers (Bio-Rad, Hemel Hempsted, UK). 5. 3X sample buffer: 187.5 mM Tris-HCl pH 6.8, 6% w/v SDS, 30% glycerol, 150 mM DTT, 0.03% w/v bromophenol blue. 6. 10X Running buffer: 250 mM Tris, 1.92 M glycine. While making 1X gel running buffer with distilled water, add 10% SDS at 1:100 dilution. 7. SDS-PAGE apparatus (Bio-Rad). 8. Power pack (Bio-Rad).

2.7. Western Blotting 1. 10X Transfer buffer: 250 mM Tris, 1.92 M glycine. Make 1X transfer buffer with distilled water. 2. Hybond nitrocellulose membrane (Amersham Bioscience UK Ltd., Little Chalfont, UK). 3. Trans blot SD semi-dry transfer cell (Bio-Rad). 4. Power pack (Bio-Rad). 5. Extra-thick filter paper (Bio-Rad).

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6. Blocking buffer: 6% (w/v) nonfat dry milk, 0.1% (v/v) Tween 20 in 1X PBS. 7. Platform rocker (Bibby Sterilin Ltd., Pontypridd, UK). 8. Primary antibody: Mouse anti-EGFP (8362-1, Clontech Laboratories Ltd., Mountain View, CA, USA), mouse anti-HA (MMS-101P, Covance Laboratories, Harrogate, UK). Dilutions are made in blocking buffer (see Note 6). 9. Secondary antibody: Enhanced chemiluminescent (ECL) anti-mouse IgG conjugated to horseradish peroxidase (NA931, Amersham Biosciences). Dilution is made in blocking buffer. 10. ECL Western blotting detection reagent (Amersham Biosciences). 11. High-performance chemiluminescence film (Hyperfilm ECL, Amersham Biosciences). 12. Hypercassette (Amersham Biosciences). 13. RP X-OMAT Processor, Model M6B (Developer, Kodak Ltd., Watford, UK).

2.8. Stripping and Reprobing Immunoblots 1. Stripping buffer: 10% SDS, 1 M Tris-HCl pH 6.8, 14 M -mercaptoethanol. 2. Hot block (Bibby Sterilin Ltd.). 3. Primary antibody: Rabbit anti-actin (A 2066, Sigma-Aldrich, Gillingham, Dorset, UK). Dilution is made in blocking buffer. 4. Secondary antibody: Enhanced chemiluminescent (ECL) anti-rabbit IgG conjugated to horseradish peroxidase (NA934, Amersham Biosciences). Dilution is made in blocking buffer.

3. Methods 3.1. Mammalian Cell Culture 3.1.1. Culturing of COS-7 Cells 1. COS-7 cells are maintained at subconfluent densities in DMEM supplemented with 10% FBS, 100 U/mL penicillin/streptomycin, 2 mM l-glutamine at 37°C, 5% carbon dioxide (CO2 ). 2. The cells are grown on coverslips in 6-well plates for immunofluorescence analysis. 3. The stock cultures are maintained in 75-cm2 flasks (NUNC; Invitrogen) and subcultured periodically as described in Subheading 3.1.1.1.

3.1.1.1. COS-7 Subculture 1. The medium from the flask is removed and the cells are rinsed once with DMEM (without serum). 2. Two mL of 1X trypsin-EDTA is added to cover the cell layer and the flask is incubated at 37°C for 5 min or until (no longer than 10 min) the cells are dissociated from the flask.

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3. The enzyme activity is quenched by adding 8 mL of fresh supplemented medium and the cells are dissociated by pipetting the medium up and down a few times. This also mechanically frees cells from individual clumps into suspension. 4. The cells are passaged in 1:10 to 1:20 ratio in fresh supplemented medium and maintained at 37°C, 5% CO2 incubator with humidity.

3.1.2. Culturing of Stable PC12 Cells 1. The PC12 stable cells expressing either the huntingtin exon-1 fragment tagged to EGFP with 23 or 74 polyglutamine repeats (EGFP-HDQ23 or EGFP-HDQ74) (13) or the wild-type or A53T or A30P mutant -synuclein (10) are maintained in DMEM supplemented with 10% heat-inactivated horse serum, 5% Tet-approved FBS, 100 U/mL penicillin/streptomycin, 2 mM l-glutamine, 100 μg/mL G418 (to maintain tet-on element) and 75 μg/mL hygromycin (to maintain the huntingtin exon-1 or -synuclein components) at 37°C, 10% CO2 with humidity. 2. The cells are seeded at 1–2 × 105 per well in 6-well plates either directly (for Western blot analysis) or on glass cover slips (for immunofluorescence analysis). 3. The expression of the transgene is switched on with 1 μg/mL (for huntingtin) or 2 μg/mL (for -synuclein) doxycycline. 4. The expression of the transgene is switched off by removing the medium containing doxycycline followed by a few rinses with fresh supplemented doxycycline-free medium.

3.1.2.1. PC12 Subculture 1. PC12 cells are subcultured using a protocol similar to that used for COS-7 cells (Subheading 3.1.1.1.), except that a much shorter incubation period with trypsin is required. Since PC12 cells tend to form clumps, care is taken to dissociate the clumps to a fine suspension before reseeding them.

3.2. Transient Transfection 3.2.1. Transient Transfection with EGFP-HDQ74 Plasmid DNA 1. COS-7 cells are grown on coverslips in 6-well plates for 24 h prior to transfection. 2. Transfection is performed using LipofectAMINE reagent (Invitrogen) (see Note 7). 4.5–6 μL of the reagent prediluted in 100 μL of serum-free DMEM is mixed with 1.5–2 μg of plasmid DNA (empty pEGFP or EGFP-tagged HDQ74) also diluted in 100 μL of serum-free DMEM. 3. The mixture is allowed to stand at room temperature for 30 min. 4. In the mean time, cells in 6-well plates are rinsed once with serum-free DMEM and a further 800 μL of the serum-free DMEM is added to the cells. 5. The lipofectAMINE-DNA mixture is then added to the cells, making the total transfection volume up to 1 mL. 6. The transfection mixture is replaced with fresh supplemented culture medium after 3–5 h incubation at 37°C, 5% CO2 . The transfected cells are treated with

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Fig. 1. Analysis of mutant huntingtin (EGFP-HDQ74)-induced aggregation and cell death in transiently transfected COS-7 cells in the presence of an autophagy inhibitor. (A) COS-7 cells were transfected with 1.5 μg of EGFP-HDQ74 construct for 4 h and then grown in complete media for 48 h. Cells were either left untreated or treated with 10 mM 3-MA (an autophagy inhibitor) for the last 15 h of the 48 h post-transfection period, and then fixed for immunofluorescence analysis. Fluorescent microscopy images of EGFP-positive COS-7 cells with EGFP-HDQ74 aggregates were

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3.2.2. Treatment of EGFP-HDQ74-Transfected COS-7 Cells with Modulators of Autophagy Pathway 1. COS-7 cells transiently transfected as above are treated with either 0.2 μg/mL rapamycin for 48 h posttransfection or 10 mM 3-methyl adenine or 400 nM bafilomycin A1 for the last 15 h of the 48 h posttransfection period prior to fixation (see Subheading 3.2.5.). Respective vehicles are used as controls.

3.2.3. Transient Transfection with EGFP-LC3 Plasmid DNA 1. COS-7 cells are grown on cover slips in 6-well plates for 24 h, followed by treatment with autophagy modulators for 24 h (see Subheading 3.2.4.) prior to transfection. 2. Transfection is performed using LipofectAMINE reagent (Invitrogen) and EGFPLC3 construct, as described in Subheading 3.2.1. 3. The transfection mixture is replaced with fresh supplemented culture medium after 3–5 h incubation at 37°C, 5% CO2 . The transfected cells are treated with the autophagy modulators for a further 2 h (see Subheading 3.2.4.) and then fixed for immunofluorescence (see Subheading 3.2.5.) (Fig. 2).

3.2.4. Treatment of EGFP-LC3-Transfected COS-7 Cells with Modulators of Autophagy Pathway 1. COS-7 cells are pretreated with 10 mM 3-methyl adenine or 0.2 μg/mL rapamycin for 24 h prior to transfection. EGFP-LC3 transfected cells are treated with these autophagy modulators for a further 2-h posttransfection period before fixing for immunofluorescence (see Subheading 3.2.5.).

Fig. 1. (Continued) taken at high (top panel) and low intensity (middle panel) to denote the number EGFP-positive transfected cells (top panel) and aggregates (indicated by arrows, middle panel), respectively. Nuclei were stained with DAPI (bottom panel) for analysis of cells showing apoptotic morphology (indicated by thick arrowheads). (Bar, 50 μm.). 3-MA treatment increased the proportion of transfected cells with aggregates and cell death, compared to the control. (B and C) Approximately 200 EGFP-positive COS-7 cells were counted for the proportion of cells with EGFPHDQ74 aggregates (B) and cell death (C). The effect of 3-MA treatment on the percentage of EGFP-positive cells with aggregates or apoptotic morphology (cell death) was expressed as odds ratios, and the control (untreated cells) was taken as 1. 3-MA significantly increased the proportion of cells with aggregates (p < 0.0001) and cell death (p = 0.006), compared to the control.

Odds ratio of EGFPpositive COS-7 cells containing >5 EGFP LC3 vesicles

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Fig. 2. Effect of autophagy modulators on EGFP-LC3 vesicles in transiently transfected COS-7 cells. COS-7 cells were pretreated with or without 0.2 μM rapamycin (Rap) or 10 mM 3-MA for 24 h prior to transfection. Cells were then transfected with 1.5 μg of EGFP-LC3 construct for 4 h and treated with or without the autophagy modulators for a further 2 h. Cells were fixed and analyzed by fluorescence microscopy for the proportion of transfected cells with more than five EGFP-LC3 vesicles. The effects of treatment on the percentage of EGFP-positive cells with more than five EGFP-LC3 vesicles were expressed as odds ratios and the control (untreated cells) was taken as 1. Rapamycin (an autophagy inducer) significantly increased the number of cells with more than five EGFP-LC3 vesicles (p < 0.0001), whereas 3-MA (an autophagy inhibitor) significantly reduced the number of cells with more than five EGFP-LC3 vesicles (p < 0.0001).

3.2.5. Fixing Cells for Immunofluorescence Analysis 1. The cells on cover slips are rinsed once with 1X PBS and fixed with 4% paraformaldehyde in 1X PBS for 20 min. 2. The fixed cells are further rinsed three times with 1X PBS, air-dried, and mounted onto glass slides on the antifadent citifluor, supplemented with 3 μg/mL DAPI to allow visualization of nuclear morphology (see Note 8).

3.3. Quantification of Aggregate Formation and Abnormal Cell Nuclei 1. Aggregate formation and nuclear morphology are assessed using a fluorescence microscope (Nikon eclipse E600W fluorescent microscope) (Fig. 1B,C). 2. Two hundred EGFP-positive COS-7 cells are randomly selected and the proportion of cells with EGFP-HDQ74 aggregates (bright fluorescent foci) is assessed. If a cell has no aggregate, a score of zero is given, while a cell having one or more aggregates is given a score of one (9). The observer is blinded to the identity of the slides and the experiments are performed in triplicate and are repeated twice. 3. Cells are considered dead if the DAPI-stained nuclei showed apoptotic morphology (fragmentation or pyknosis). Pyknotic nuclei are typically less than 50% the diameter of normal nuclei and show obvious increased DAPI intensity. These criteria are specific for cell death, as they show a very high correlation with propidium iodide staining of live cells (15). Furthermore, these nuclear abnormalities are reversed with caspase inhibitors (13,15).

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3.4. Quantification of Cells with EGFP-LC3 Vesicles 1. The proportion of cells with EGFP-LC3 vesicles are assessed using a fluorescence microscope (Nikon eclipse E600W fluorescent microscope) (Fig. 2). 2. Two hundred EGFP-positive COS-7 cells are counted for the proportion of cells with more than five EGFP-LC3 vesicles (green fluorescent punctuate structures). If a cell has 0–5 EGFP-LC3 vesicles, a score of zero is given, while a cell having more than 5 EGFP-LC3 vesicles is given a score of one (12). The observer is blinded to the identity of the slides and the experiments are performed in triplicate and are repeated twice.

3.5. Clearance Experiments in PC12 Stable Inducible Cell Lines 1. The transgene expression in the PC12 stable cells is induced by adding doxycycline (see Subheading 3.1.2.) for 8 h (for huntingtin cell lines) or 48 h (for the -synuclein cell lines) (9,10). 2. The expression is then turned off by removing doxycycline from the media (see Subheading 3.1.2.). 3. Concurrent with switching off the expression, the cells are treated with the modulators of autophagy, 3-MA, Rap, or BafA1 (as in Subheading 3.2.1.) for 24, 48, or 72 h (for immunofluorescence analysis), or 120 h (for Western blot analysis). Equal amounts of water or DMSO are added as controls, where relevant. 4. The cells are either fixed for immunofluorescence as explained in Subheading 3.2.2. (Fig. 3) or collected for Western blot analysis as explained in Subheading 3.6. (Fig. 4).

3.6. Western Blot Analysis for the Clearance of Aggregate-Prone Proteins 3.6.1. Preparation of Samples 1. The cells from 6-well plates are scraped into the media and collected in labeled Eppendorf tubes for each sample. 2. The cells are centrifuged at 5000 g at 4o C for 5 min. 3. The supernatant is discarded and the cell pellets are washed once with 1X PBS. 4. The cells are then lysed with appropriate volumes of lysis buffer for 30 min on ice. 5. The lysed cells are centrifuged at 13,000 g for 5 min at 4o C to remove cell debris and any unlysed cells. 6. The supernatant is stored at –80o C until further analysis. 7. Protein assay is performed on the samples with Bio-Rad protein assay kit. 8. 30 μg of protein from each sample is mixed with 3X sample buffer (to a final 1X) and boiled at 100o C for 5 min on a heating block before loading on the gel for SDS-PAGE.


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Fig. 4. Effect of rapamycin on the clearance of mutant aggregate-prone proteins in stable inducible PC12 cell lines expressing EGFP-HDQ74 or A53T -synuclein. (A) Stable inducible PC12 cells expressing EGFP-HDQ74 were induced with doxycycline for 8 h and transgene expression was switched off (by removing doxycycline) for 120 h, with (+) or without (–) 0.2 μM rapamycin (Rap). Clearance of soluble EGFP-HDQ74 was analyzed by immunoblotting with antibody against EGFP. Rapamycin (120 h OFF + Rap) enhanced the clearance of soluble EGFP-HDQ74 compared to the control (120 h OFF). Note that the lanes appearing as separate gel strips were derived from nonadjacent lanes from the same exposure of immunoblot from a single gel where both the samples were loaded; the intervening lanes were excised to simplify presentation. (B) Densitometry analysis for the clearance of soluble EGFP-HDQ74 was done by the ratio of intensity of EGFP relative to actin. Rapamycin treatment greatly enhanced the clearance of soluble EGFP-HDQ74 compared to the control condition (120 h OFF, which is set to 100%). (C) Stable inducible PC12 cells expressing the A53T mutant of -synuclein were induced with doxycycline for 48 h and transgene expression was switched off (by removing doxycycline) for 24 h, with (+) or without (–) 0.2 μM rapamycin (Rap). Clearance of A53T -synuclein was analyzed by immunoblotting with antibody against HA. Rapamycin (24 h OFF + Rap) enhanced the clearance of A53T -synuclein compared to the control (24 h OFF). Note that the lanes appearing as separate gel strips were derived from nonadjacent lanes from the same exposure of immunoblot from a single gel where both the samples were loaded; the intervening lanes were excised to simplify presentation. (D) Densitometry analysis for the clearance of A53T -synuclein was done by the ratio of intensity of HA relative to actin. Rapamycin treatment greatly enhanced the clearance of A53T -synuclein compared to the control (24 h OFF, set to 100%).


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3.6.2. SDS-PAGE 1. SDS-PAGE is performed using Bio-Rad gel apparatus. 2. Resolving gel is first poured in between a gel plate with a 0.75- or 1-mm integrated spacer and a glass plate sandwich that are held together in the gel casting unit. The gel is poured up to 75% of the length of the plates, leaving space for pouring the stacking gel. 3. A small volume (500 μL) of water-saturated isobutanol is added on the surface of the resolving gel, which is later washed with distilled water when the gel has polymerised. 4. The stacking gel is poured over the resolving gel and a 10-well comb is inserted into it. 5. After polymerization of the stacking gel, the combs are removed and the wells thus created are rinsed with distilled water. 6. The casted gels are fitted into the gel apparatus and 1X gel running buffer is poured into the tank. 7. The boiled samples are then loaded into the wells along with a prestained molecular weight marker. 8. The gels are run at a constant current of 15 mA per gel until the marker bands migrate to the desired position.

3.6.3. Western Blotting 1. The proteins in the samples that have been separated by SDS-PAGE are transferred electrophoretically onto a nitrocellulose membrane using Bio-Rad semidry transfer apparatus. 2. Nitrocellulose membranes are cut according to the size of the gel and equilibrated in 1X transfer buffer along with extra-thick filter papers for 5–10 min. The gel is taken out and is also equilibrated in the transfer buffer. 3. The gel transfer unit is then assembled by placing a filter paper on the base plate of the transfer apparatus, then the nitrocellulose membrane, followed by the gel on top of the membrane and finally two more filter papers on the top. While placing the gel/filter papers on the membrane, the surface is rolled gently with a plastic pipet so as to remove any air bubbles. 4. After placing the compression plate and connecting the lid, the transfer is performed at a constant voltage of 15 V for approximately 1 h. 5. The membranes (immunoblots) are then removed and incubated in 25 mL of blocking buffer with gentle shaking on a rocker at room temperature for 1 h. 6. The blocking buffer is discarded and primary antibody diluted in 10 mL of blocking buffer (anti-EGFP at 1:2000 dilution for detecting mutant huntingtin and anti-HA at 1:1000 dilution for detecting mutant -synuclein) is added to the immunoblots and incubated overnight on a rocker in a cold room at 4°C. Blotting can also be carried out by incubating the membrane with anti-EGFP antibody for 1 h at room temperature on a rocker (see Note 9).

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7. The nonspecific binding of the primary antibody is then removed by washing the immunoblots with 0.1% Tween 20 in 1X PBS (PBS-T) solution, three times for 10 min each on a rocker at room temperature. 8. Secondary antibody (anti-mouse IgG conjugated to horseradish peroxidase at 1:2000 dilution) is added to the immunoblots and incubated for a minimum of 1 h (not more than 3 h) on a rocker at room temperature. 9. The nonspecific binding of the secondary antibody is then removed by washing the immunoblots in PBS-T solution, three times for 10 min each on a rocker at room temperature. 10. After the final wash and discarding the PBS-T, the following procedures are performed in a dark room under safe light conditions. 11. 1.5 mL each of the detection reagents 1 and 2 of the ECL Western blotting detection system are mixed and added to the immunoblots for 1 min, while ensuring coverage of the entire surface. 12. The detection solution is discarded and the immunoblots are then wrapped in cling films and placed in hypercassettes along with a hyperfilm with appropriate exposure times (Fig. 4A,C). 13. The films are developed in an automated Kodak developer.

3.6.4. Stripping and Reprobing the Immunoblots 1. The immunoblots are stripped of the signal for EGFP or HA by immersing them in stripping buffer at 65o C on a hot block for 10 min (see Note 10). 2. The immunoblots are then washed with 1X PBS twice for 5 min each on a rocker at room temperature, followed by blocking with blocking buffer for 1 h. 3. After discarding the blocking buffer, primary antibody (rabbit anti-actin at 1:2000 dilution) is added to the immunoblots and incubated either overnight at 4o C or for 1 h at room temperature on a rocker. 4. The immunoblots are then washed three times with PBS-T as before, incubated with secondary antibody (ECL anti-rabbit IgG conjugated to horseradish peroxidase at 1:2000 dilution) for a minimum of 1 h on a rocker at room temperature, followed by three washes with PBS-T. The signal is detected with the ECL Western blotting detection system as before (Fig. 4A,C).

3.7. Statistical Analysis 3.7.1. Odds Ratio for Quantification of Cells with Mutant Huntingtin Aggregation and Cell Death and EGFP-LC3 Vesicles 1. Pooled estimates for the changes in aggregate formation (Fig. 1B), cell death (Fig. 1C), or EGFP-LC3 vesicles (Fig. 2), resulting from perturbations assessed in multiple experiments, are calculated as odds ratios with 95% confidence intervals [e.g., odds ratio of aggregation = (percentage of cells expressing construct with aggregates in perturbation conditions/percentage of cells expressing construct without aggregates in perturbation conditions)/(percentage of cells expressing


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construct with aggregates in control conditions/percentage of cells expressing construct without aggregates in control conditions)]. Odds ratios were considered to be the most appropriate summary statistic for reporting multiple independent replicate experiments of this type, because the percentage of cells with aggregates under specified conditions can vary between experiments on different days, whereas the relative change in the proportion of cells with aggregates induced by an experimental perturbation is expected to be more consistent. Our lab has used this method frequently in the past to allow analysis of data from multiple independent experiments (12,13,15). 2. Odds ratios and p values are determined by unconditional logistical regression analysis, using the general log-linear analysis option of SPSS 9 software (SPSS, Chicago, USA). The control condition is set to one. *** p < 0.001; ** p < 0.01; * p < 0.05; NS, nonsignificant.

3.7.2. Densitometry Analysis on the Immunoblots for Assessing the Clearance of Mutant Aggregate-Prone Proteins 1. Densitometry analysis on immunoblots from three independent experiments is performed by Scion Image Beta 4.02 software (Scion Corporation, Maryland, USA). The control condition is set to 100%. 2. The clearance of the mutant aggregate-prone proteins is determined by the ratio of the intensity of EGFP to actin for mutant huntingtin clearance (Fig. 4B) or the ratio of the intensity of HA to actin for mutant -synuclein clearance (Fig. 4D).

4. Notes 1. Doxycycline is stored as a 100 mg/mL stock solution at –20°C in dark Eppendorf tubes to protect it from direct light. For experiments, it is diluted in sterile water to a 10 mg/mL working stock solution, which can also be stored at –20°C and can be used for a few months. The working stock is then diluted again in the cell culture medium to a final concentration of 1 μg/mL. 2. When 3-MA is stored at room temperature, it crystallizes. When using the stock solution for experiments, 3-MA needs to be warmed in a 37°C water bath until the solution becomes transparent. This should be started at least 30 min prior to use. 3. The PFA solution is warmed to about 50°C with constant stirring in a fume hood to help it dissolve. If the solution is cloudy, a few drops of 1 M NaOH may be added to make the solution clear. It is then cooled at room temperature, aliquoted, and stored at –20°C. 4. APS must be prepared fresh every time prior to pouring a gel. 5. TEMED is best stored at room temperature. As it may decline in quality after opening, it is better to buy small bottles. 6. Mouse anti-EGFP is an excellent antibody for Western blotting and can be used at 1:10,000 dilution. Mouse anti-HA antibody is diluted at 1:1000 in blocking buffer.

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7. For 1 μg of DNA to be transfected in COS-7 cells in 6-well plates, use 3 μL of lipofectamine reagent. 8. The cover slips are mounted carefully onto the glass slides using forceps to prevent formation of air bubbles. Nail varnish is then applied to the edges of the cover slips to hold them firmly on the slides. 9. The primary antibodies can be reused a few times for subsequent experiments if stored in 0.02% sodium azide (done by dilution from a 10% stock solution) at –20°C. The only adjustment that is required is to increase the exposure time of the film at the ECL step. 10. Stripping of the immunoblots must be carried out in glass beakers on a hot plate inside a fume hood, as this generates a foul smell.

Acknowledgments We thank T. Yoshimori (National Institute of Genetics, Japan) for EGFP-LC3 construct. We are grateful to the Wellcome Trust (Senior Clinical Fellowship to DCR), MRC, EU (EUROSCA) and Muscular Dystrophy Campaign for funding.

References 1. Ross, C. A. and Poirier, M. A. (2004) Protein aggregation and neurodegenerative disease. Nat. Med. 10(Suppl), S10–17. 2. Narain, Y., Wyttenbach, A., Rankin, J., Furlong, R. A. and Rubinsztein, D. C. (1999) A molecular investigation of true dominance in Huntington’s disease. J. Med. Genet. 36, 739–746. 3. Klionsky, D. J. and Ohsumi, Y. (1999) Vacuolar import of proteins and organelles from the cytoplasm. Annu. Rev. Cell Dev. Biol., 15, 1–32. 4. Schmelzle, T. and Hall, M. N. (2000) TOR, a central controller of cell growth. Cell 103, 253–262. 5. Kovacs, A. L., Gordon, P. B., Grotterod, E. M. and Seglen, P. O. (1998) Inhibition of hepatocytic autophagy by adenosine, adenosine analogs and AMP. Biol. Chem. 379, 1341–1347. 6. Yamamoto, A., Tagawa, Y., Yoshimori, T., Moriyama, Y., Masaki, R. and Tashiro, Y. (1998) Bafilomycin A1 prevents maturation of autophagic vacuoles by inhibiting fusion between autophagosomes and lysosomes in rat hepatoma cell line, H-4-II-E cells. Cell Struct. Funct. 23, 33–42. 7. Hara, T., Nakamura, K., Matsui, M., et al. (2006) Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885–889. 8. Komatsu, M., Waguri, S., Chiba, T., et al. (2006) Loss of autophagy in the central nervous system causes neurodegeneration in mice. Nature 441, 880–884. 9. Ravikumar, B., Duden, R. and Rubinsztein, D.C. (2002) Aggregate-prone proteins with polyglutamine and polyalanine expansions are degraded by autophagy. Hum. Mol. Genet. 11, 1107–1117.


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10. Webb, J. L., Ravikumar, B., Atkins, J., Skepper, J. N. and Rubinsztein, D. C. (2003) Alpha-Synuclein is degraded by both autophagy and the proteasome. J Biol. Chem. 278, 25009–25013. 11. Ravikumar, B., Vacher, C., Berger, Z., et al. (2004) Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat. Genet. 36, 585–595. 12. Sarkar, S., Floto, R.A., Berger, Z., et al. (2005) Lithium induces autophagy by inhibiting inositol monophosphatase. J. Cell Biol. 170, 1101–1111. 13. Wyttenbach, A., Swartz, J., Kita, H., et al. (2001) Polyglutamine expansions cause decreased CRE-mediated transcription and early gene expression changes prior to cell death in an inducible cell model of Huntington’s disease. Hum. Mol. Genet. 10, 1829–1845. 14. Kabeya, Y., Mizushima, N., Ueno, T., et al. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J. 19, 5720–5728. 15. Wyttenbach, A., Sauvageot, O., Carmichael, J., Diaz-Latoud, C., Arrigo, A. P. and Rubinsztein, D.C. (2002) Heat shock protein 27 prevents cellular polyglutamine toxicity and suppresses the increase of reactive oxygen species caused by huntingtin. Hum. Mol. Genet. 11, 1137–1151.

14 Localization and MHC Class II Presentation of Antigens Targeted for Macroautophagy Dorothee Schmid and Christian Münz

Summary Intracellular antigens can be presented on major histocompatibility complex (MHC) class II molecules after degradation via macroautophagy. To enhance MHC class II presentation of potential vaccine antigens, we have developed a method to target antigens for autophagic degradation via fusion to the Atg8/LC3 protein: Atg8/LC3 is specifically incorporated into autophagosomes via coupling to phosphatidylethanolamine, and subsequently degraded in MHC class II loading compartments (MIICs). Antigens fused to the N-terminus of Atg8/LC3 follow the same pathway and get preferentially presented on MHC class II molecules. The localization of Atg8/LC3 fusion antigens in MIICs can be visualized by confocal microscopy, and MHC class II presentation can be quantified in a presentation assay with antigen-specific CD4+ T-cell clones. These assays are good measures of autophagosome formation and lysosomal degradation of macroautophagy cargo and therefore are useful for studying regulation of the autophagic pathway under various experimental conditions and physiological perturbations.

Key Words: Atg8/LC3; influenza matrix protein 1; MHC class II; MHC class II loading compartment (MIIC); confocal microscopy; CD4+ T-cell clones; IFN- ELISA.

1. Introduction Major histocompatibility complex (MHC) class II molecules present products of lysosomal proteolysis to CD4+ T cells, which orchestrate the adaptive immune response and therefore are key to successful immune surveillance (1–3). From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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One pathway by which intracellular antigens gain access to lysosomal degradation and subsequent MHC class II presentation is macroautophagy (reviewed in refs. 4 and 5). Macroautophagy has been suggested to deliver viral, model, and tumor antigens for MHC class II presentation to CD4+ T cells (6–9). More recently, we have shown that autophagosomes frequently fuse with MHC class II loading compartments (MIICs) and that this pathway can be targeted to improve MHC class II presentation of potential vaccine antigens (10). Our targeting method takes advantage of the properties of the Atg8/LC3 protein: During autophagosome formation, Atg8/LC3 is covalently coupled to a lipid in the nascent autophagosome membrane via residue Gly120 and thus is incorporated into autophagosomes (11,12). After fusion of autophagosomes with lysosomes, Atg8/LC3 is degraded by lysosomal proteases (10) and Atg8/LC3-derived peptides can be found to be presented on MHC class II molecules (13). To target antigens for autophagic degradation via the same mechanism, we fused the Atg8/LC3 sequence to the 3´ end of the influenza A virus matrix proteins 1 (MP1) sequence. We could demonstrate that this fusion antigen was indeed targeted to MIICs and was presented on MHC class II molecules up to 20-fold more efficiently than MP1 itself (10) (Figs. 1 and 2). Furthermore, we could show that the targeting strategy was dependent on covalent coupling to the autophagosomal membrane, since mutation of Gly120 to Ala120 completely abrogated autophagosome targeting and eliminated enhanced MHC class II presentation of the fusion antigen (10) (Fig. 2). In addition, we could show that MHC class I presentation was not affected, since the fusion protein was equally well recognized by MP1-specific CD8+ T-cell clones (10) (Fig. 2). We propose that Atg8/LC3 fusion antigens are useful tools for studying the autophagy pathway under various experimental conditions and perturbations, since their MIIC localization and MHC class II presentation require an intact autophagic pathway, including autophagosome formation, fusion with late endosomes and lysosomal degradation of autophagic cargo. Any treatment that affects macroautophagy should also affect MIIC localization and MHC class II presentation of Atg8/LC3 fusion antigens and therefore these readouts can be used to measure autophagosome formation and lysosomal degradation. Here we describe in detail how LC3 fusion antigens can be expressed by transient transfection in suitable target cell lines. Furthermore, we describe a method to visualize MIIC localization of LC3 fusion antigens by confocal immunofluorescence microscopy. Finally, we describe how to set up MHC class II presentation assays with antigen specific CD4+ T-cell clones and how their recognition of MHC class II presented epitopes, derived from autophagic cargo, can be measured by IFN- ELISA.

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Fig. 1. Localization of MP1-LC3 in MHC class II loading compartments. (A) MDAMC cells transfected with the MP1-LC3 construct were treated with 200 U/mL IFN- for 36 h to upregulate MHC class II expression. To prevent degradation of MP1LC3 by lysosomal proteases, cells were treated with 50 μM chloroquine (CQ) during the last 6 h of the culture, where indicated (+CQ). Cells were fixed, stained with MP1and MHC class II–specific antibodies and DAPI and analyzed by confocal microscopy. MP1-LC3 staining partially overlaps with the MHC class II staining, especially in CQ-treated cells, indicating that MP1-LC3 is degraded in MIICs. (Scale bar: 10 μm.) Representative fields from one experiment out of two are shown. (From ref. 10 with permission from Elsevier.) (B) As in (A), except that cells were transfected with MP1 construct. In contrast to MP1-LC3, MP1 is homogeneously distributed in the cytoplasm and nucleus and even after CQ-treatment does not accumulate in MHC class II loading compartments. (Adapted from ref. 10 with permission from Elsevier.)

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Fig. 2. MHC class II presentation assay with MP1-specific CD4+ T-cell clones. (A) MP1-specific CD4+ T-cell clone 11.46 was cocultured at various effector to target cell (ET) ratios with HaCat target cells transfected with either MP1, MP1-LC3, or MP1-LC3(G120 A). The next day, IFN- in culture supernatants was measured by ELISA to assess presentation of MP1 on MHC class II. MHC class II presentation is strongly enhanced by the Atg8/LC3 fusion, and this is dependent on covalent coupling to the autophagosome membrane via Gly120 . Error bars indicate standard deviations and p-values for paired, one-tailed Student’s t-test statistics across all E:T ratios are shown. One of two experiments is shown. (B) MHC class I presentation assay with MP1-specific CD8+ T-cell clone 9.2. The setup was the same as in (A), except that MDAMC cells transfected with MP1, MP1-LC3 or MP1-LC3(G120 A) were used to assess presentation on MHC class I. This control experiment shows that MHC class I presentation is not affected by the Atg/LC3 fusion. (From ref. 10 with permission from Elsevier.)

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2. Materials 2.1. Cell Culture 1. Cell lines: HaCat keratinocyte cell line, a gift of Rajiv Khanna, Brisbane, Australia. MDAMC human breast carcinoma cell line, a gift of Irene Joab, Paris, France. (For choice of cell lines, see Note 1). 2. Cell culture medium: Dulbecco’s modified Eagle’s medium (DMEM, Gibco, Carlsbad, CA) with 10% fetal bovine serum (FBS; Sigma), 2 mM glutamine, 110 μg/mL sodium pyruvate, and 2 μg/mL gentamicin (Gibco). 3. Sterile Dulbecco’s phosphate-buffered saline (DPBS; Gibco) with 0.5 mM ethylenediamine tetraacetic acid (EDTA; Sigma, St. Louis, MO). To prepare, make 0.5 M EDTA stock, pH 8.0 in H2 O, filter through a 0.2-μm filter to sterilize, and dilute 1:1000 in sterile DPBS. 4. Solution of 0.05% trypsin/0.53 mM EDTA (Gibco). 5. Recombinant human interferon- (IFN-; ProSpec-Tany TechnoGene LTD, Rehovot Israel). Reconstitute in sterile H2 O + 0.1% human serum albumin (HSA, Sigma) to prepare a 2 × 106 U/mL stock and freeze in aliquots at −20ºC.

2.2. Expression of MP1-LC3 Fusion Construct by Transient Transfection 1. Mammalian expression vector encoding for LC3 fusion protein of influenza A matrix protein 1 (MP1, genebank entry X08088), as described in ref. 10. The construct was designed by cloning the human Atg8/LC3 cDNA sequence (genebank entry NM_022818) into the mammalian expression vector pEGFP-C2 (BD Biosciences, San Jose, CA) and subsequently replacing the EGFP coding sequence with the MP1 coding sequence (without a Stop codon at the 3´ end). As controls, mammalian expression vectors encoding for MP1 or MP1-LC3(G120 A) (10) should be used in parallel with MP1-LC3. These constructs were designed the same way, except that the MP1 construct contains a Stop codon at the 3´ end of the MP1 sequence and the MP1-LC3(G120 A) construct contains a point mutation at nucleotide 358 of the Atg8/LC3 sequence (A instead of G). For best transfection results, maxipreps of the different DNA plasmids should be prepared and DNA should be eluted in sterile DPBS. Determine DNA concentration and purity by OD260 /OD280 reading in a spectrophotometer. OD260 /OD280 ratio should be greater than 1.8. Store DNA in aliquots at −20ºC. 2. Cell culture medium (see Subheading 2.3.1.) without gentamicin (see Note 2). 3. Lipofectamine 2000 transfection reagent (Invitrogen, Carlsbad, CA). 4. OptiMEM-I medium (Gibco).

2.3. Immunocytochemistry and Confocal Microscopy to Analyze Localization of MP1-LC3 in MHC Class II Compartments 1. Circular 1.5-mm cover slips (Fisher, Pittsburgh, PA). 2. 70% (v/v) ethanol and sterile DPBS.

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3. Recombinant human IFN- (see Subheading 2.1.5.). 4. Chloroquine (CQ, Sigma). Prepare a 20 mM stock in ddH2 O and sterilize through a 0.2-μm filter. Freeze in aliquots at –20ºC. Use at a 1:400 dilution. 5. Phosphate-buffered saline (PBS): Prepare 10x stock (1.37 MNaCl, 27 mM KCl, 100 mM Na2 HPO4 , 18 mM KH2 PO4 , adjust pH to 7.4 with HCl) and autoclave before storage at room temperature. Prepare 1x PBS by dilution of one part with nine parts ddH2 O. 6. 4% Paraformaldehyde (PFA): Prepare a 4% (w/v) solution in PBS by heating PFA (Sigma) in PBS in a covered jar until PFA dissolves (careful, since PFA is toxic and vapors should not be inhaled). Let cool to room temperature for immediate use or store in aliquots at –20ºC. Thaw a fresh aliquot for each experiment. 7. Permeabilization solution: 0.1% (v/v) Triton-X 100 in PBS. 8. Blocking buffer: PBS + 1% bovine serum albumin (BSA, Sigma) + 5% normal donkey serum (NDS) + 0.1% saponin (Calbiochem, San Diego, CA) (see Note 3). 9. Wash buffer: PBS + 0.1% saponin. 10. Primary antibodies: Rabbit polyclonal MP1-specific antiserum (14) (use at 1:2000) and mouse monoclonal HLA-DR/DP/DQ-specific hybridoma IVA12 (ATCC, Manassas, VA). Grow hybridoma in RPMI-1640 with 10% FBS, 2 mM glutamine, 2 μg/mL gentamicin and harvest supernatant by spinning cells down at 300g for 10 min. Filter supernatant through a 0.2-μm filter and store at 4ºC. Use at a 1:10 dilution. 11. Secondary antibodies: Donkey anti-rabbit IgG-Alexa 488 (Invitrogen-Molecular Probes, Carlsbad, CA, use at 1:500) and donkey anti-mouse IgG-RhodamineRedTM -X (Jackson ImmunoResearch, West Grove, PA, use at 1:300) (see Note 4). 12. DAPI nucleic acid stain: Prepare a 5 mg/mL stock of 4,6-diamidino-2phenylindole (DAPI, Invitrogen-Molecular Probes) in ddH2 O and store in aliquots at −20ºC. Dilute 1:10,000 in PBS to prepare working solution and store at 4ºC wrapped in aluminium foil. 13. Mounting medium: Prolong Gold Antifade Reagent (Invitrogen-Molecular Probes).

2.4. MHC Class II Presentation Assay Using MP1-Specific CD4+ T-Cell Clones 1. MP1-specific CD4+ T-cell clones, generated as described in ref. 15 and cultured in RPMI-1640 with 8% pooled human serum (PHS, Mediatech Inc.), 450 U/mL recombinant human IL-2 (Chiron, Vacaville, CA), 2 mM glutamine, 2 μg/mL gentamicin in round-bottom 96-well plates (see Note 5). 2. Recombinant human IFN- (see Subheading 2.1., step 5.). 3. RPMI-1640. 4. Coculture medium: RPMI-1640 with 5% PHS, 2 mM glutamine, 2 μg/mL gentamicin.


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5. Positive control stimulus: Specific MP1 peptide (1 mM stock in 10% DMSO, for T-cell stimulation dilute 1:1000 in coculture medium) or phytohemagglutinin (PHA-L, Sigma, 1 mg/mL stock, for T-cell stimulation dilute 1:1000 in coculture medium).

2.5. IFN- ELISA to Analyze Secretion of IFN- by MP1-Specific CD4+ T-Cell Clones 1. High-protein-binding 96-well ELISA plate (e.g., Maxisorp from Nunc, Rochester, NY). 2. ELISA kit for human IFN- (Mabtech, Nacka Strand, Sweden). Prepare a 10 μg/mL stock of human recombinant IFN- provided with kit and freeze in aliquots at -20ºC. Use a freshly thawed aliquot for each experiment. 3. PBS (see Subheading 2.3.). 4. Coculture medium (see Subheading 2.4.). 5. Blocking buffer: PBS + 1% BSA (Sigma). 6. Wash buffer: PBS + 0.05% Tween-20. 7. Incubation buffer: PBS + 0.1% BSA + 0.05% Tween-20. 8. TMB peroxidase substrate solution (Sigma) and Stop solution (1 N sulfuric acid).

3. Methods 3.1. Cell Culture 1. HaCat and MDAMC cell lines are maintained in DMEM + 10% FBS medium in 75-cm2 tissue culture flasks or 100-mm plates until they approach confluence. 2. To split cells, wash monolayer once with 5 mL of DPBS/0.5 mM EDTA and incubate with 2 mL of trypin/EDTA solution at 37ºC for 2–3 min (MDAMC) or 10–15 min (HaCat) to detach cells. To set up new maintenance cultures, replate 1/20 of cells and add fresh culture medium. These cultures will approach confluence after 2–3 d. 3. To induce expression of MHC class II machinery, cells have to be cultured with 200 U/mL IFN- for 24 h (for HaCat) or 36 h (for MDAMC). At least 50% of cells then will express MHC class II on their cell surface.

3.2. Expression of MP1-LC3 Fusion Construct by Transient Transfection 1. To set cells up for transfection, detach cells as described in Subheading 3.1.2. and plate onto a 6-well tissue culture plate in antibiotic-free culture medium at a density of 2 × 105 cells/well (see Note 2). 2. The next day, cultures should be about 70–80% confluent. For each well to be transfected, dilute 2.5 μg plasmid DNA in 250 μL OptiMEM-I medium and mix by vortexing briefly. In a separate tube, dilute 7.5 μL lipofectamine 2000 in

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Schmid and Münz 250 μL OptiMEM-I medium and vortex briefly. Incubate both tubes for 5 min at room temperature. Combine both solutions, vortex briefly and incubate 20 min at room temperature to allow formation of DNA–lipofectamine complexes. Add complexes in a dropwise manner to the culture medium of cells in the 6-well plate and incubate at 37ºC for 4–6 h. After 4–6 h, replace the complex-containing medium with fresh culture medium and culture cells for 18–20 h at 37ºC (see Note 6). Twenty-four hours posttransfection, cells are ready to be split onto microscopy cover slips for localization analysis by confocal immunofluorescence microscopy (see Subheading 3.3.) or for use in MHC presentation assays (see Subheading 3.4.).

3.3. Immunocytochemistry and Confocal Microscopy to Analyze Localization of MP1-LC3 in MHC Class II Compartments 1. Place round 1.5-mm microscopy cover slips into 24-well tissue culture plate. Use eight extra cover slips for control stainings (see Note 7). 2. Sterilize cover slips by washing once with 70% ethanol and twice with sterile DPBS. Remove any traces of ethanol by completely aspirating off ethanol and wash solutions with vacuum suction flask. 3. Trypsinize cells transfected with different MP1 constructs (see Subheading 3.2.) and plate onto sterilized cover slips in cell culture medium, at a density of 5 × 104 cells/well. Plate two wells of each sample, so that cells can be analyzed with and without chloroquine treatment (see Note 8). 4. Treat cells with 200 U/mL IFN- for 24 h (HaCat cells) or 36 h (MDAMC cells) to induce expression of MHC class II molecules. 5. During the last 6 h of the culture, treat one set of cells with 50 μM chloroquine (CQ), to prevent degradation of MP1-LC3 by lysosomal proteases. Leave the other set of cells untreated (see Note 8). 6. Wash cells once in PBS (0.5 mL/well) and fix in 4% paraformaldehyde (PFA, 200 μL/well) for 15 min at room temperature (see Note 9). 7. Wash cells once in PBS (0.5 mL/well) and permeabilize in 0.1% Triton X-100 (200 μL/well) for 5 min. 8. Wash cells once in PBS (0.5 mL/well) and add blocking buffer (200 μL/well) for 30 min. 9. Dilute primary antibodies in blocking buffer and add to cells (200 μL/well) for 30 min at room temperature or for longer periods (up to overnight) at 4ºC. 10. Wash cells three times in wash buffer (0.5 mL/well), incubate for 5 min each time. 11. Dilute secondary antibodies in blocking buffer and add to cells (200 μL/well) for 30 min. 12. Aspirate secondary antibody solutions and add DAPI nucleic acid stain for 20–30 sec (200 μL/well). Afterwards, immediately wash cells as in step 9.


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13. Wash cells once in PBS and mount cover slips by inverting them onto a drop of mounting medium on a microscope slide, up to four cover slips per slide. Carefully press down on cover slip, aspirate excess mounting medium and let dry at room temperature (see Note 10). Afterwards, slides can be stored in the dark at 4ºC for several months. 14. Analyze slides with a confocal laser scanning microscope, using a high N.A. oil immersion lens (e.g., 63x/1.4 N.A.). Excitation at 405 nm induces DAPI fluorescence (blue emission), excitation at 488 nm induces Alexa 488 fluorescence (green emission), and excitation at 543 nm induces Rhodamine-RedTM -X fluorescence (red emission). Software can be used to overlay the different fluorescence channels and to quantify colocalization (see Note 11). An experiment with MP1LC3 and MP1-expressing MDAMC cells is shown as an example in Fig. 1.

3.4. MHC Class II Presentation Assay Using MP1-Specific CD4+ T-Cell Clones 1. For MHC class II presentation assay, use HaCat cells transfected with different MP1 constructs in a 6-well format, as described in Subheading 3.2. Treat cells with 200 U/mL IFN- for 24 h to initiate expression of the MHC class II machinery. 2. Remove any traces of IFN- from cells by washing cell monolayers three times in RPMI-1640 medium. Trypsinize cells to prepare a cell suspension, wash once in coculture medium and count with hemacytometer. Prepare cell suspensions in coculture medium at three different cell concentrations (2 × 105 , 105 , and 6.67 × 104 cells/mL) (see Note 12). 3. Collect MP1-specific CD4+ T-cell clones from 96-well culture plates, wash once in coculture medium and count. Adjust cell concentration to 2 × 106 cells/mL. 4. Set up cocultures of T cells and target cells in doublets (2 wells/condition) in a 96-well round-bottom plate. Per well, plate 50 μL of T-cell suspension (105 cells/well) and 100 μL of the different target cell suspensions (either 2 × 104 , 104 , or 6.67 × 103 cells/well). This will result in effector to target (E:T) ratios of 5, 10 and 15. As a positive control, stimulate T-cell clones with specific MP1 peptide (1 μM) or PHA-L (1 μg/mL). As negative control, stimulate T-cell clones with coculture medium only. 5. Culture cells overnight (18–24 h) at 37ºC.

3.5. IFN- ELISA to Analyze Secretion of IFN- by MP1-Specific CD4+ T-Cell Clones 1. One day prior to ELISA, coat high-protein-binding ELISA plate with primary anti-IFN- antibody (1-D1K, included in IFN- ELISA kit), diluted 1:500 in PBS, 100 μL/well. Incubate plate overnight at 4ºC. 2. The next day, wash plate two times with PBS (200 μL/well) and block with blocking buffer (200 μL/well) for 1 h at room temperature.

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3. Thaw an aliquot of IFN- standard (10 μg/mL) and prepare serial dilutions in coculture medium (prepare 2000, 1000, 500, 250, 125, 62.5, 31.25, 0 pg/mL standards, at least 300 μL each). 4. To make sure that IFN- secreted by T cells is homogeneously distributed in culture supernatants, mix supernatants by pipetting up and down with a multichannel pipet and pellet cells by centrifugation of plates at 300g for 5 min. 5. With a multichannel pipet, carefully remove 120 μL of supernatant from each well and transfer to a new 96-well plate. 6. Wash ELISA plate four times with wash buffer. 7. Add supernatants or IFN- standards (100 μL/well) and incubate for 2 h at room temperature. Freeze remaining 20 μL of supernatant at –20ºC in case ELISA has to be repeated on diluted supernatants (see Note 13). 8. Wash as in step 6 and add 100 μL/well of secondary antibody (7-B6-1-biotin, provided in ELISA kit), diluted 1:1000 in incubation buffer. Incubate one hour at room temperature. 9. Wash as in step 6 and add 100 μL/well of streptavidin-HRP (provided in ELISA kit), diluted 1:1000 in incubation buffer. Incubate one hour at room temperature. 10. Wash as in step 6 and add 100 μL/well of TMB peroxidase substrate. Incubate until blue reaction product has sufficiently developed, then stop reaction by adding 100 μL/well of Stop solution. 11. Measure optical density at 450 nm (OD450 ) in an ELISA plate reader and convert OD450 values into IFN-concentration in pg/mL (see Note 13). An example of the results produced is shown in Fig. 2.

4. Notes 1. Cell lines were chosen based on their ability to upregulate MHC class II expression upon IFN- treatment (10) and their HLA haplotype. HaCat is HLA-DRB1*15, -DRB1*04, -DQBI*03 and -DQB1*03 positive, and therefore could be used in MHC class II presentation assays with MP1-specific CD4+ T-cell clones from a healthy lab donor with the following MHC haplotype: HLA-A*0201, -A*6801, -B*4402, -B*0702, -C*0501, -C*0702, -DRB1*1501, -DRB1*0401, -DRB5*01, -DRB4*01, -DQBI*0602 and -DQB1*0301. MDAMC is HLA-A*02-positive and therefore could be used in MHC class I presentation assays with HLA-A2-restricted MP1-specific CD8+ T-cell clones derived from the same donor. If other cell lines have to be used, determine MHC class II expression/upregulation upon IFN- treatment and compare their HLA haplotype with that of T-cell donors. 2. For transfection with lipofectamine 2000, cell culture medium should not contain any antibiotics. Therefore, omit gentamicin. 3. Normal donkey serum is used as blocking reagent because secondary antibodies are derived from donkey. If secondary antibodies come from a different species, use 5% normal serum from that species as blocking reagent.


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4. For choice of fluorophores, consider which lasers and filter sets are available for your confocal microscope. For colocalization analysis it is important that emission spectra of the green and red fluorochromes do not overlap. Also consider brightness and photostability of fluorophores. 5. For generation of MP1-specific CD4+ T-cell clones, CD14− cells isolated from a healthy lab donor were stimulated with autologous mature DCs that were electroporated with 10 μg of in vitro transcribed MP1-RNA. On day 8, the stimulation was repeated and 10 U/mL IL-2 were added to enhance T-cell survival. On day 21, the surviving cells were cloned by limiting dilution at 10, 1, or 0.3 cells/well and expanded in RPMI-1640 + 8% PHS + 150 U/mL rhIL-2 (Chiron) + 1 μg/mL PHA-L (Sigma). 105 irradiated PBMCs/well and 104 irradiated LCLs/well were added as feeder cells. On day 40, expanded cells were tested in split-well IFN- ELISPOT assays for recognition of an MP1 peptide mix. T-cell clones can be frozen and stored for several years in liquid nitrogen (about 106 cells/aliquot). Frozen cultures can be reexpanded as described above. 6. Removal of complex-containing medium 4–6 h after transfection was found to improve viability of MDAMC and HaCat cells lines. 7. For correct interpretation of results, the following control stainings should be included: a) Replace primary and secondary antibodies with blocking buffer. These stainings should be completely negative. b) Replace primary antibody with blocking buffer, but use secondary antibodies. Background from secondary antibodies should be low. If background turns out to be too high, titer down concentration of secondary antibodies. c) Use untransfected cells and cells that were not treated with IFN- Stainings should be negative. d) Do single-labeling of cells and check signal in the “wrong channel” (red channel for Alexa488 labeling and green channel for Rhodamine-Red Xlabeling). There should be no bleed-through into the wrong channel. e) Do single-labeling with “wrong” secondary antibodies. There should be no cross-reactivity with the wrong species. 8. Chloroquine inhibits lysosomal acidification and thus prevents degradation of lysosomal substrates, including LC3 fusion proteins (see ref. 10). Without inhibition of lysosomal proteases, MP1-LC3 is rapidly degraded and its detection in lysosomal compartments is more difficult. However, when cells are treated with chloroquine, the fusion protein accumulates in lysosomal compartments/MIICs and now can be visualized much more readily (see Fig. 1 and ref. 10). Therefore, it is recommended that one set of cells be treated with chloroquine for 6 h prior to the staining. 9. From the fixation step onwards, cells can be handled outside sterile hood on a laboratory bench. Use a vacuum suction flask to change solutions, exchange plastic tip of suction device when handling different solutions (e.g., antibodies). All incubation steps are done at room temperature, unless noted otherwise.

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10. Prolong Gold antifade mounting medium (Invitrogen-Molecular Probes) should be allowed to dry at room temperature overnight. During this time, the mounting medium will gel and its refractive index increases. Sealing of cover slips with nail polish is not necessary for Prolong Gold, but is recommended for other, water-based mounting media. 11. If a confocal microscope is not available, alternatively slides can be analyzed with a conventional wide-field fluorescence microscopes with a motorized z-stage. To remove out-of-focus light and accurately analyze colocalization of fluorochromes, z-stacks subsequently have to be deconvolved using deconvolution software. 12. The optimal number of target cells and T-cell clones may vary, depending on the T-cell clone and the type of target cell. Therefore it is recommended to try a range of different effector and target cell numbers, ranging from 104 to 2 × 105 T cells/well and 103 –105 target cells/well. 13. In case IFN- levels in supernatants exceed the linear range of the ELISA (approximately 20–1000 pg/mL), dilute frozen supernatants 1:10 in coculture medium and repeat ELISA.

Acknowledgments We thank Rajiv Khanna and Irene Joab for the gift of cell lines and Ari Helenius for the gift of the MP1-specific antiserum. This work was supported by the Arnold and Mabel Beckman Foundation, the Alexandrine and Alexander Sinsheimer Foundation, the Burroughs Wellcome Fund, the National Cancer Institute (R01CA108609 and R01CA101741) (to C.M.), and in part by a General Clinical Research Center grant (M01-RR00102) from the National Center for Research Resources at the National Institutes of Health (to the Rockefeller University Hospital). D.S. is a recipient of a predoctoral fellowship from the Schering Foundation.

References 1. Chapman, H. A. (2006) Endosomal proteases in antigen presentation. Curr. Opin. Immunol. 18, 78–84. 2. Bryant, P., and Ploegh H. (2004) Class II MHC peptide loading by the professionals. Curr. Opin. Immunol. 16, 96–102. 3. Bevan, M. J. (2004) Helping the CD8+ T-cell response. Nat. Rev. Immunol. 4, 595–602. 4. Schmid, D., Dengjel, J., Schoor, O., Stevanovic, S., and Münz. C. (2006) Autophagy in innate and adaptive immunity against intracellular pathogens. J. Mol. Med. 84, 194–202. 5. Münz, C. (2006) Autophagy and antigen presentation. Cell Microbiol. 8, 891–898.


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6. Brazil, M. I., Weiss, S., and Stockinger, B. (1997) Excessive degradation of intracellular protein in macrophages prevents presentation in the context of major histocompatibility complex class II molecules. Eur. J. Immunol. 27, 1506–1514. 7. Dorfel, D., Appel, S., Grunebach, F., Weck, M. M., Muller, M. R., Heine, A., and Brossart, P. (2005) Processing and presentation of HLA class I and II epitopes by dendritic cells after transfection with in vitro transcribed MUC1 RNA. Blood 105, 3199–3205. 8. Nimmerjahn, F., Milosevic, S., Behrends, U., Jaffee, E. M., Pardoll, D. M., Bornkamm, G. W., and Mautner, J. (2003) Major histocompatibility complex class II-restricted presentation of a cytosolic antigen by autophagy. Eur. J. Immunol. 33, 1250–1259. 9. Paludan, C., Schmid, D., Landthaler, M., Vockerodt, M., Kube, D., Tuschl, T., and Münz, C. (2005) Endogenous MHC class II processing of a viral nuclear antigen after autophagy. Science 307, 593–596. 10. Schmid, D., Pypaert, M., and Münz, C. (2007) MHC class II antigen loading compartments continuously receive input from autophagosomes. Immunity 26, 79–92. 11. Kabeya, Y., Mizushima, N., Ueno, T., Yamamoto, A., Kirisako, T., Noda, T., Kominami, E., Ohsumi, Y., and Yoshimori, T. (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. Embo J. 19, 5720–5728. 12. Mizushima, N., Yamamoto, A., Matsui, M., Yoshimori, T., and Ohsumi, Y. (2004) In vivo analysis of autophagy in response to nutrient starvation using transgenic mice expressing a fluorescent autophagosome marker. Mol. Biol. Cell 15, 1101–1111. 13. Dengjel, J., Schoor, O., Fischer, R., Reich, M., Kraus, M., Muller, M., Kreymborg, K., Altenberend, F., Brandenburg, J., Kalbacher, H., Brock, R. R., Driessen, C., Rammensee, H. G., and Stevanovic, S. (2005) Autophagy promotes MHC class II presentation of peptides from intracellular source proteins. Proc. Natl. Acad. Sci. USA 102, 7922–7927. 14. Martin, K., and Helenius, A. (1991) Transport of incoming influenza virus nucleocapsids into the nucleus. J. Virol. 65, 232–244. 15. Fonteneau, J. F., Larsson, M., Somersan, S., Sanders, C., Münz, C., Kwok, W. W., Bhardwaj, N., and Jotereau, F. (2001) Generation of high quantities of viral and tumor-specific human CD4+ and CD8+ T-cell clones using peptide pulsed mature dendritic cells. J. Immunol. Methods 258, 111–126.

15 Chaperone-Mediated Autophagy S. Kaushik and A. M. Cuervo

Summary Chaperone-mediated autophagy (CMA) is the only type of autophagy in mammalian cells able to selectively degrade cytosolic proteins in lysosomes. CMA is maximally activated in response to stressors such as prolonged starvation, exposure to toxic compounds, or oxidative stress. We have found that CMA activity decreases in aging and in some age-related disorders such as Parkinson’s disease. Impaired CMA under these conditions may be responsible for the accumulation of damaged proteins inside cells and for their higher vulnerability to stressors. In contrast to other forms of autophagy, where substrates are engulfed or sequestered along with other cytosolic components, CMA substrates are translocated one-by-one across the lysosomal membrane. Changes in the levels/activity of the lysosomal components required for substrate translocation can be used to stimulate CMA activity. However, the most unequivocal method to measure CMA is by directly tracking the translocation of substrate proteins into isolated lysosomes.

Key Words: Lysosomes; metabolic labeling; protease protection assay; chaperones; lysosomal membrane proteins. 1. Introduction Two properties differentiate chaperone-mediated autophagy (CMA) from the other types of autophagy in mammalian cells: its selectivity towards a particular pool of cytosolic proteins and the mechanism of delivery of the substrate proteins to lysosomes (1). Only proteins bearing a particular targeting motif in their amino acid sequence, biochemically related to the pentapeptide KFERQ (2), are selectively recognized by the heat shock cognate protein of 70 kDa (hsc70), the chaperone that mediates their delivery to lysosomes for their degradation via CMA (3). It is estimated that about 30% of soluble cytosolic From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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proteins contain this CMA-targeting motif. The already identified substrate proteins include, among others, some glycolytic enzymes (glyceraldehyde-3phosphate dehydrogenase, aldolase, phosphoglyceromutase), subunits of the 20S proteasome, transcription factors and inhibitors of transcription factors (c-fos, the inhibitor of NFB [IB]), cytosolic forms of secretory proteins (2 -microglobulin), calcium-binding proteins (Annexins I, II, IV, and VI), and proteins associated to vesicular trafficking (-synuclein) (reviewed in refs. 1 and 4). CMA substrates access the lysosomal lumen directly across the lysosomal membrane. After the CMA targeting motif is recognized by hsc70, the substrate/chaperone complex is targeted to the surface of the lysosomes where it binds to the lysosome-associated membrane protein type 2A (LAMP-2A), a receptor for CMA (5). Once bound, the substrate unfolds (6) and crosses the lysosomal membrane assisted by a lysosomal form of hsc70 (lys-hsc70) present in the lumen (7). Transport is saturable, requires a source of energy (ATP), and is temperature-dependent (binding occurs at temperatures as low as 10°C, but transport is only detected at temperatures above 25°C) (8,9). The selectivity associated to degradation via CMA seems beneficial under particular conditions in which discrimination between different types of proteins for degradation is required. For example, activation of CMA during prolonged starvation provides the amino acids required for protein synthesis under those conditions but favors degradation of unnecessary proteins against that of proteins essential for cell survival (10,11). Similarly, activation of CMA during mild-oxidative stress (12) or after exposure to protein-modifying toxic compounds (13) allows the selective removal of the proteins damaged or altered under these conditions. Malfunctioning of CMA has been described in familial forms of Parkinson’s disease where -synuclein, the protein mutated in this disorder, is delivered to lysosomes for degradation via CMA, but after binding to LAMP-2A it cannot be translocated to the lysosomal lumen resulting in blockage of this pathway (14). CMA activity also decreases with age due to a decrease in the levels of LAMP2A at the lysosomal membrane (15), contributing thus to the accumulation of abnormal proteins and the higher susceptibility to stressors characteristic of old organisms. Measurement of changes in the lysosomal levels of the main CMA components (LAMP-2A and hsc70) and of the intracellular abundance and location of the lysosomes active for this pathway (those enriched in the CMA components) provide a general idea about CMA activity in tissues and cells in culture. However, accurate measurement of CMA is only attained by tracking the translocation of CMA substrates into isolated lysosomes using proteaseprotection assays (8,16–18).

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2. Materials 2.1. Isolation of Rat Liver Lysosomes 1. Wistar rats (200–250 g). To enrich in CMA-active lysosomes, rats should be starved (complete food removal but with water ad libitum) for 24–48 h before lysosomal isolation (minimum 10 h). 2. Tools: dissection instruments (forceps, scissors, clamps), double cloth gauze, funnels, teflon/glass homogenizer (for motorized homogenizer). 3. Centrifugation supplies: polycarbonate tubes (30 mL), ultraclear tubes for SW28 (Beckman, Fullerton, CA), SW28 rotor (Beckman). 4. 0.25 M sucrose (American Bioanalytical, Natick, MA) adjusted at pH 7 (with NaOH). All sucrose-containing solutions can be stored at 4°C but for no more than 3 d (as slow hydrolysis of sucrose occurs when stored for longer periods of time). 5. Metrizamide (Amresco, Solon, OH) prepared as a 85.6% (w/v) stock in water (stock solution adjusted at pH 7.2 with NaOH). Store in aliquots at −20°C in dark (see Note 1).

2.2. Purification of Recombinant GST-hsc70 1. Glutathione transferase (GST)- hsc70 was purified from XLBlue E. coli bacteria transformed with a plasmid (pGX-2T, Pharmacia, Uppsala, Sweden) containing the cDNA for the fusion protein of human hsc70 (19). 2. Isopropyl--d-thiogalactopyranoside (IPTG, American Bioanalytical) is prepared freshly by dissolving it at 0.1 M in sterile water. 3. Glutathione-agarose (Sigma, St. Louis, MO) is reconstituted as a 50% v/v stock in TBS following manufacture’s instructions. 4. Ampicillin (American Bioanalytical) is dissolved at 100 mg/mL in sterile water and stored at 4°C. 5. STE buffer: 10 mM Tris-HCl, pH 8, 150 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA). 6. Tris buffer saline (TBS): 50 mM Tris-HCl, pH 7.4, 150 mM NaCl. 7. Lysozyme (Sigma) is prepared fresh by dissolving it at 100 mg/mL in sterile water. 8. Dithiothreitol (DTT, Sigma) is prepared as 5 M stock in water and stored in aliquots at −20°C. 9. Triton X-100 (Bio-Rad, Hercules, CA) is prepared as 20% (v/v) stock solution in sterile water and stored at room temperature. 10. N-Lauryl sarcosine (Sarkosyl, Sigma) is prepared as 15% (w/v) stock solution in sterile water and stored at room temperature. 11. Elution buffer: 10 mM reduced GSH (Sigma) in 50 mM Tris-HCl, pH 8 is prepared fresh. 12. Wash buffer: 50 mM Tris-HCl, pH 7.5, 150 mM NaCl.

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13. Thrombin (Sigma) diluted as 10x stock in thrombin cleavage buffer (amount should be adjusted according to the activity of the particular batch used) (optional). 14. Thrombin cleavage buffer: 2.5 mM CaCl2 in wash buffer (optional).

2.3. Radiolabeling of Substrate Proteins by Reductive Methylation 1. Purified protein of interest is prepared at 10 mg/mL in the reaction buffer (see Note 2). 2. Reaction buffer: 10 mM MES (Sigma) pH 7. Store at 4°C. 3. Bovine serum albumin (BSA) (American Bioanalytical) dissolved at 20 mg/mL in the reaction buffer. 4. [14 C]Formaldehyde (Amersham, Piscataway, NJ) 1–3% aqueous solution in sealed ampoule (1.85–2.29 GBq/mmol, 50–62 mCi/mmol). Store at 4°C. 5. Sodium cyanoborohydrate (Sigma) (9 mg/mL) in reaction buffer. Prepare fresh. 6. Sephadex G-25 to G-100 fine (Sigma) (depending on the size of the labeled protein), equilibrated in sterile water following manufacture’s recommendation.

2.4. Protein Degradation Assay 1. Proteolysis buffer: 10 mM 3-(N-Morpholino)propanesulfonic acid (MOPS, Sigma) pH 7.3, 250 mM sucrose, 1 mM DTT, 5.4 mM cysteine. Prepare fresh. 2. Energy regenerating system (ERS): prepare a 6x cocktail at these final concentrations 60 mM ATP, 12 mM phosphocreatine (Sigma), 0.3 mg/mL creatine phosphokinase (Sigma), 60 mM MgCl2 in 0.25 M sucrose adjusted at pH 7.3. 3. Trichloroacetic acid (American Bioanalytical) dissolved at 20% (w/v) in water and stored at room temperature. 4. BSA (American Bioanalytical) disolved at 20 mg/mL in water. 5. Millipore multiscreen assay system (Millipore, Bedford, MA), 0.22-μm durapore filter 96-well plates, and polystyrene flat-bottom 96-well plates.

2.5. Protease-Protection Transport Assay 1. Incubation buffer: 10 mM MOPS pH 7.3, 250 mM sucrose. Prepared fresh and stored at 4°C. 2. Chymostatin (Sigma) prepared at 10 mM stock dissolved in incubation buffer and stored at −20°C. 3. ERS (see Subheading 2.4.). 4. Proteinase K (Sigma) dissolved at 5 mg/mL stock in 10 mM Tris-HCl pH 7.5, 1 mM CaCl2 5. 4-(2-Aminoethyl)-benzenesulfonyl fluoride hydrochloride (AEBSF, American Analytical) dissolved in water as 1mM stock and stored at −20°C. 6. Reagents for standard SDS-PAGE and immunoblot.


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2.6. Indirect Immunofluorescence 1. Dulbecco’s modified Eagle’s medium (DMEM) (Sigma) supplemented with 10% newborn calf serum (or any other media that cells require for growth). 2. Microscope cover slips (22 × 22 mm) (Fisher Scientific, Pittsburgh, PA). 3. Phosphate-buffered saline (PBS): 1.37 M NaCl, 0.03 M KCl, 0.07 M Na2 HPO4 , 0.11 M K2 HPO4 pH 7.4. Store at room temperature. 4. Methanol fixing solution: 20% methanol (v/v) (in PBS). Store at −20°C. 5. Blocking solution: 0.2% (w/v) powdered nonfat milk, 2% newborn calf serum, 0.1 M glycine, 1% BSA, and 0.01% Triton X-100 in PBS. Prepare fresh and maintain at 4°C until use. 6. Secondary antibodies: Fluorescein-labeled donkey anti-mouse IgM, μ-chain specific, and cyanine5-labeled goat anti-rabbit IgG (H+L chain specific) (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). 7. Mounting media: SlowFade Light Antifade Kit with DAPI 1 (Molecular Probes, Invitrogen).

3. Methods CMA activity can be measured in confluent cells in culture using pulsechase experiments. After 2-d labeling with a radiolabeled amino acid (to preferentially label long-lived proteins), CMA can be activated in cultured fibroblasts at about 10 h after serum removal. CMA-dependent degradation is defined under those conditions as that activated by serum removal, which is sensitive to ammonium chloride (the same as any other form of autophagy), but insensitive to phosphatydil-inositol-3-kinase inhibitors (which block macroautophagy) (Fig. 1) (14,20–23). Although this method is useful to discriminate from degradation via macroautophagy, two limitations make it inconvenient to rely only on this method: (1) it only measures activation of CMA in response to serum removal—it does not measure changes in basal CMA activity as it is not possible to differentiate from microautophagy by this approach; (2) it requires the use of inhibitors of macroautophagy, and it is well supported now that maintained blockage of a proteolytic pathway leads to the compensatory upregulation of other proteolytic pathways. The most unequivocal method to measure CMA is by directly tracking the translocation of substrate proteins into lysosomes. CMA can be reproduced in vitro using freshly isolated intact lysosomes by incubating the substrate protein and lysosomes at 37°C in an isotonic media supplemented with hsc70 and an ATP-regenerating system (8,16–18). Using standard protease protection assays, as those commonly used for the study of translocation of proteins across other organelles membrane, the three main steps in CMA (binding, uptake, and degradation) can be separately analyzed. Binding of substrate to the lysosomal membrane can be determined as the amount of substrate associated

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Fig. 1. Measurement of CMA-dependent protein degradation in culture cells. (A) Cultured cells are labeled by incubation with a radiolabeled amino acid for more than 24 h. At the end of the incubation, cells are extensively washed, and degradation of protein is followed by calculating the percentage of acid precipitable radioactivity at time 0 (protein), converted to acid soluble radioactivity (amino acids and small peptides) at each time. (B) Protein degradation increases after serum removal. The percentage of ammonium chloride–sensitive degradation inhibited by PI3K inhibitors corresponds to macroautophagy, while that insensitive to the inhibitor corresponds to CMA and microautophagy.

to the lysosomes recovered by centrifugation, because any protein that has been translocated will be rapidly degraded by the lysosomal proteases (the half-life of substrate proteins in the lysosomal lumen is ∼5 min). Uptake of substrates can be measured in lysosomes pretreated with protease inhibitors. In this case, the substrate remains in the lysosomal lumen after crossing the lysosomal membrane. The amount of substrate translocated into the lysosomal


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lumen can be determined by comparing the level of total substrate (bound plus translocated) to the substrate bound to the membrane (in lysosomes untreated with protease inhibitors), or by measuring the amount of substrate resistant to cleavage by an exogenously added protease (in lysosomes treated with protease inhibitors) (Fig. 2). Because lysosomal isolation requires a large number of cells and some level of practice in the handling of isolated organelles, we have recently optimized an indirect method to evaluate CMA activity in those tissues or cells in which lysosomal isolation is not feasible. This method is also discussed in this chapter and it is based on the analysis of changes in the population of lysosomes responsible for CMA, which is defined as those containing lys-hsc70 (the lysosomal chaperone required for substrate translocation) in their lumen (12,24). This population can be detected in tissue sections or fixed cultured cells by immunoelectron microscopy with an antibody directed against hsc70 (CMA+ lysosomes are the ones containing the chaperone in their lumen) or by immunofluorescence in cultured cells, as the number of vesicles containing hsc70 and LAMP-2A. It is also possible to indirectly track changes in CMA activity by measuring changes in the major components of this pathway (hsc70 and LAMP-2A) in lysosomes. It is important to point out that total cellular levels of these components do not necessarily correlate with changes in CMA, because both proteins are also present in other cellular compartments and play other roles in the cell. It is thus necessary to determine that the increase or decrease in these components takes place in the lysosome-associated fraction. Protein levels usually result more accurate measurement than changes in mRNA levels. Hsc70 is constitutively expressed and we have not found changes at the transcriptional level in any condition associated to changes in CMA activity. In the case of LAMP-2A, conditions such as oxidative stress associate to transcriptional upregulation of LAMP-2A (12), but in other conditions, such as prolonged starvation, the increase in levels of LAMP-2A in the lysosomal compartment does not require de novo synthesis of the protein (25). 3.1. Isolation of Rat Liver Lysosomes 1. Livers from two starved rats are washed extensively with cold 0.25 M sucrose (see Note 3), weighed, and then minced. 2. The minced livers are homogenized in three volumes of 0.25 M sucrose/g of liver with 8–10 strokes at maximum speed. The homogenates are filtered through double gauze, and four volumes of cold 0.25 M sucrose are added. 3. After centrifugation of the homogenate at 6800g for 5 min at 4°C, the supernatant is collected into a clean tube (avoid collecting the white floating material close to the pellet, which mainly consists of heavy mitochondria). The pellet is gently resuspended with a dry glass test tube filled with ice (“cold finger”) in starting

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Fig. 2. In vitro assay for the measurement of substrate protein translocation into lysosomes via CMA. Lyosome isolation: Lysosomes are isolated from rat liver by centrifugation in a discontinuous gradient of metrizamide of a mitochondria/lysosome enriched fraction obtained by differential centrifugation. Protein degradation: A radiolabeled substrate protein (glyceraldehyde-3-phosphate dehydrogenase [GAPDH] in this example) is incubated with freshly isolated intact lysosomes, and their proteolytic activity is calculated as the percentage of acid precipitable radioactivity at time 0 (protein) transformed to acid soluble radioactivity (amino acids and small peptides) at the end of the incubation. Protease-protection assay: Freshly isolated intact lysosomes, treated or not with protease inhibitors, are incubated with the substrate protein (GAPDH) and at the end of the incubation lysosomes are collected by centrifugation and subjected to SDS-PAGE and immunoblot to detect the amount of substrate protein associated. Binding: substrate associated to lysosomes untreated with protease inhibitors; Association: substrate associated to lysosomes treated with protease inhibitors (bound + internalized); Uptake: substrate associated to lysosomes treated with protease inhibitors after removal of the membrane bound substrate with an exogenous protease (proteinase K [Prot. K] in this example).


4.

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8. 9.

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volume of 0.25 M sucrose and spun under the same conditions. Both supernatants are then pooled together. The pooled supernatants are centrifuged at 17,000g for 10 min and the pellet resuspended with the “cold finger” in dry to guarantee complete resuspension. After adding 3.5 volumes of 0.25 M sucrose/g liver to the resuspended pellet, it is centrifuged under the same conditions (this step assures that any cytosolic component trapped in the initial pellet is released into the supernatant). The pellet from the previous centrifugation, highly enriched in light mitochondria and lysosomes (mitochondrial-lysosomal fraction), is then resuspended with the “cold finger,” and 1 mL 0.25 M sucrose/3 g liver and two volumes of 85.6% Metrizamide are added and gently mixed with the sample to a final concentration of 57% metrizamide (see Note 4). Each 10 mL of the 57% metrizamide light mitochondrial-lysosomal fraction are loaded at the bottom of an ultraclear tube and a discontinuous metrizamide gradient is generated on top by overlying: 6 mL of 32.8% Metrizamide, 10 mL 26.3% Metrizamide, and 11 mL 19.8% Metrizamide (all diluted in water pH 7.3). The tube is filled up to 3 mm of the border with 0.25 M sucrose, and it is subjected to centrifugation in a SW 28 rotor at 141,000g for 1 h at 4°C (see Note 5). After centrifugation, white to brownish material is clearly visible in each of the interphases. Mitochondria are preferentially located in the first interphase of the gradient (starting from the bottom); the following interphase is enriched in a mixture of mitochondria and lysosomes. Lysosomes are present in the third (I3) and fourth (I4) interphases, which are collected separately (approx 3–4 mL) with a Pasteur pipet in 30-mL polycarbonate tubes (Fig. 2). After dilution in five volumes of 0.25 M sucrose, the fractions are washed by centrifugation at 37,000g for 10 min at 4°C. The pellet of I3 is carefully resuspended with a blunt Pasteur pipet in 1 mL 0.25 M sucrose and spun at 10,000g for 5 min at 4°C. The pellet of this centrifugation is preferentially enriched in secondary lysosomes with low CMA activity (lacking hsc70 in their lumen). The supernatant of the I3 centrifugation is used to resuspend the pellet of the I4 fraction to produce the secondary lysosomes with high CMA activity (enriched in hsc70) (if resuspended in 0.4 mL, the final protein concentration is 10 mg/mL approx.) (see Notes 6 and 7).

3.2. Purification of Recombinant GST-hsc70 1. Bacteria strains carrying the pGx-GST-hsc70 plasmid are grown overnight at 37°C with agitation in Luria Broth (LB) with 100 μg/mL ampicillin. Cultures are refreshed next morning by 1/10 dilution in LB ampicillin and incubated under the same conditions. 2. When the cultures are visibly turbid (OD600 = 0.3), IPTG is added to a final concentration of 0.1 mM and they are incubated for an additional 4–8 h. 3. Bacteria are collected by centrifugation (7000g for 5 min) and the pellet is washed with 1/10 of the starting volume of STE with 100 μg/mL lysozyme.

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4. After incubation for 15 min on ice, the samples are adjusted to 5 mM DTT and are lysed by the addition of 1.5% sarkosyl (N-lauryl sarcosine) and sonication on ice (1 min in a bath sonicator). 5. Lysates are subjected to centrifugation (10,000g 5 min at 4°C) and the supernatant is adjusted to 2% Triton X-100 (to minimize association of bacterial proteins with hsc70) and vortexed gently. 6. A 1/50 starting volume of GSH-agarose beads (50% v/v in TBS) is added to the supernatant and after incubation for 15 min at 4°C on a shaker, the beads are collected by centrifugation and washed 6–8 times with ice-cold TBS (see Note 8). 7. The final beads slurry is resuspended in the same volume of wash buffer supplemented with reduced GSH and incubated for 30 min at room temperature to elute the protein from the beads. After spinning, the eluted protein is recovered in the supernatant. If desired the GSH can be removed by dialysis against washing buffer without GSH.

3.3. Radiolabeling by Reductive Methylation 1. The protein of interest is dissolved in reaction buffer to a final concentration of 3 mg/mL, and [14 C]formaldehyde (250 μCi) and sodium cyanoborohydrate (final concentration of 1.8 mg/mL in reaction buffer) are added. Handling of this amount of isotope requires the use of filtered tips, double closure microfuge tubes, and disposal of all materials as solid radioactive residues waste. 2. After incubation of the reaction in a final volume of 500 μL at 25°C for one hour, the radiolabeled protein is separated from the nonincorporated isotope by gel filtration through the pertinent Sephadex matrix (according to the molecular weight of the protein). We routinely use mini-spin columns packed with the matrix (1 mL approx.) previously blocked with 20 mg/mL BSA (5 vol) (to prevent nonspecific binding) and equilibrated with the reaction buffer (10 vol). Spinning time is adjusted depending on the protein of interest and the characteristics of the mini-spin column. 3. The eluted radiolabeled protein is collected in separate aliquots (one for each spin with the column first loaded with the reaction mixture, and then with similar volumes of reaction buffer—three times). 4. The amount of radiolabeled protein and free isotope in each aliquot (total 4 aliquots) is determined by measurement of the radioactivity associated to the protein precipitated with TCA. Briefly, an aliquot of each fraction is diluted (1:100) in reaction buffer, and 50 μL of this dilution are added to a microfuge tube containing 50 μL of 20% TCA and 30 μL of 20 mg/mL BSA. The samples are incubated at 4°C for 30 min and then spun at 15,000g for 15 min at 4°C. The supernatant of each tube is collected in a scintillation vial, and the pellet is washed with other 50 μL of 20% TCA. After centrifugation in the same conditions, the second supernatant is added to the vial, while the pellet is resuspended in 50 μL of 0.1 M NaOH and then transferred to a different scintillation vial. After adding the pertinent amount of scintillation liquid, samples are counted in a scintillation -counter. The amount of acid precipitable radioactivity (protein) in the sample


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is calculated by the formula: [(dpm pellet/(dpm pellet + dpm supernatant)) *100], and the amount of radioactivity incorporated by microgram of protein (dpm/μg) by measuring the amount of protein in the collected aliquots by the Lowry procedure (26). Reasonable specific activities are in the order of 1 - 100 × 106 dpm/nmol. Radiolabeled proteins are stored in small aliquots (10 μL) in double sealing microfuge tubes at –80°C for months (avoid repeated freezing and thawing).

3.4. Protein Degradation Assay 1. Incubations are carried out in a 0.22-μm filtered durapore 96-well plate previously wet with sterile water for 10 min at room temperature. Freshly isolated rat liver lysosomes (25 μg protein) (10 μL of a 1:4 dilution in proteolysis buffer) are incubated with 10 μL of radiolabeled protein (2000 dpm/μL), 1 μL of the (6x) ERS and 10 μg/mL GST-hsc70 in a final volume of 60 μL (adjusted with proteolysis buffer) for 30 min at 37°C. One separate well should contain the same reagents, except for the lysosomes, to determine the amount of protein spontaneously cleaved (blank). 2. At the end of the incubation 90 μL of 20% TCA and 30 μL of 20 mg/mL BSA are added to each well to stop the reaction. After incubation at 4°C for at least 30 min, the acid-soluble radioactivity is collected in a polystyrene 96-well plate using the Millipore multiscreen vacuum system. The flow throughs (acid-soluble) are collected in separate scintillation vials and counted. 3. Proteolysis is calculated as the amount of acid precipitable radioactivity (protein) transformed in acid-soluble radioactivity (amino acids and small peptides) at the end of the incubation: [((dpm flow through sample – dpm flow through blank) / dpm pellet time 0)*100] (see Note 9).

3.5. Protease-Protection Transport Assay 1. Freshly isolated rat liver lysosomes are incubated with 100 μM chymostatin for 10 min on ice. Depending on the substrate, a protease inhibitor cocktail (x100) can be used instead (10 mM leupeptine, 10 mM AEBSF, 1 mM pepstatin and 100 mM EDTA). 2. Transport assay is carried out in 0.5-mL microfuge tubes by adding freshly isolated rat liver lysosomes treated or not with protease inhibitor (100 μg protein), GAPDH, or another protein of interest (50 μg), 5 μL of (x6) ERS and 10 μg/mL GST-hsc70 in 30 μL final volume of incubation buffer. Samples are incubated for 20 min at 37°C. 3. At the end of the incubation half of the tubes containing lysosomes treated with chymostatin are cooled down on ice (1 min) and proteinase K is added (5 μL). Samples are maintained on ice for 10 min, AEBSF is added (5 μL) and all samples are centrifuged at 25,000g for 5 min at 4°C. 4. Supernatants are aspirated and pellets are washed with 100 μL of incubation buffer by centrifugation under the same conditions. The final pellet is resuspended in 30 μL of Laemmli buffer (27) with protease inhibitors, boiled for 5 min at 95°C

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and subjected to SDS-PAGE and immunoblot for GAPDH. A lane with 1/10 of the amount of GAPDH added should be included in the immunoblot. 5. Densitometric analysis is performed on the immunoblots directly (if developed by the alkaline phosphatase method) or using the film from an exposure that does not contain saturated bands (if developed by the chemiluminiscence method). For each antibody a standard curve with increasing concentrations of antigen is generated to determine the linear range. These values allow the calculation of (1) binding as the percentage of total added GAPDH associated to lysosomes untreated with protease inhibitors, (2) sssociation as the percentage of GAPDH recovered in lysosomes treated with protease inhibitors, and (3) uptake, calculated as either the difference between association and binding or the amount or the percentage of GAPDH associated to lysosomes treated with proteinase inhibitors after proteinase K treatment (see Note 10).

3.6. Indirect Immunofluorescence 1. Cells grown on sterile cover slips at the bottom of 6-well plates until semiconfluent (50–70% confluence) are washed (twice) with serum-free culture medium (to remove all remaining IgGs in the serum), and fixed in −20°C methanol (20% in PBS) for 1 min at room temperature (7). Cells can be stored in PBS containing 0.02% NaN3 at 4°C in dark for weeks. 2. To start the immunofluorescence procedure, PBS is aspirated and 2 mL of blocking solution are added per well. Blockage is completed by incubating the slides for 30 mins at room temperature and washing the slides (three times) with PBS. 3. The primary antibody (rabbit IgG anti-LAMP-2A (Zymed Laboratories, Invitrogen, Carlsbad, CA) is diluted 1:100 in filtered 0.1% BSA in PBS. Slides are placed facing up in a Petri plate layered with Parafilm and 25 μL of diluted primary antibody are spotted on top of each slide. Incubation with the antibody is carried out in a wet chamber for 1 h at room temperature. 4. Slides are then extensively washed with PBS by picking up the slides with flat forceps and dipping them sequentially in four different reservoirs with 200 mL PBS each. 5. After aspirating the excess of PBS in the slides, they are incubated with 25 μL of the secondary antibody (diluted in filtered 0.1% BSA in PBS) for 30 min at room temperature in the wet chamber. Better results are obtained if the dilutions of the secondary antibodies are spun down before adding to the slides to remove any possible aggregated fluorochrome. 6. After extensive washes (as described in step 4), slides are incubated with 25 μL of the other primary antibody (mouse IgM anti-hsc70 (Abcam Inc., Cambridge, MA)) diluted 1:150 in filtered 0.1% BSA in PBS, under the same conditions (one h at room temperature in wet chamber). 7. After extensive washes (as described in step 4), slides are incubated with 25 μL of the secondary antibody (diluted in filtered 0.1% BSA in PBS) for 30 min at room temperature in the wet chamber as described in step 5, and washed (as described in step 4).


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8. Slides are mounted by placing them face down on top of 15 μL of DAPI-containing mounting media spotted on a glass slide, and sealed with nail polish to prevent drying. 9. Slides are visualized in a fluorescence microscope (Axoivert 200, Carl Zeiss Ltd. Thornwood, NY), and captured images are subjected to deconvolution with the manufacture’s software. Colocalization of the two antibodies is quantified in thresholded images with the JACoP plug-in of the Image J software (NIH, MD). The mean distance of the vesicles positive for each antibody to the nucleus is calculated with the “straight lane tool” and the “analyze particles” function of the Image J software, by drawing lanes from the most distant vesicle positive for each antibody to the nucleus and computing the particle distribution (distance and density). An average of six different radial lines per cell and 20 cells per field is usually calculated to determine changes in the intracellular distribution of CMA+ lysosomes (see Note 11).

4. Notes 1. Metrizamide has to be dissolved in the dark (beaker wrapped with foil paper) slowly to avoid the formation of solid clumps of undissolved material. We usually start with half of the final volume of water and keep adding metrizamide in little amounts while stirring. Adjusting the pH of the solution should be done with 0.01 M NaOH once the metrizamide is dissolved (wait until it stabilizes before adding the next drop). Our laboratory has tested different replacements for metrizamide (sucrose, premade solutions of Percoll or Nycodenz), but in all cases the level of enrichment of the lysosomes more active for CMA was considerably low and the stability of the lysosomal membrane (critical for the uptake experiments) was compromised. 2. We have used more than 20 different proteins for transport assays. The method of radiolabeling by reductive methylation is similar for all of them, but the size of the molecular exclusion matrix (Sephadex) for the separation of the labeled protein from the non-incorporated isotope should be chosen based on the molecular weight of the protein. As an example we have used Sephadex G-50 for GAPDH (144,000 kDa) and Sephadex G-25 for Ribonuclease A (16,000 kDa). 3. Liver perfusion is not strictly necessary. Extensive washing of the livers with sucrose removes most of the contaminant blood, and the remaining blood cells are eliminated in the pellet of the first centrifugation. Two critical factors to obtain a fraction of lysosomes with preserved lysosomal membrane stability are the temperature of isolation (all solutions should be kept on ice to prevent proteases released during the homogenization for damaging the cytosolic face of the lysosomal membrane) and the time required for the whole procedure. Eliminating liver perfusion shortens the time without compromising the purity of the lysosomal fraction.

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4. Pipetting the lysosomal fraction through regular plastic 1000-μL micropipet tips reduces the lysosomal membrane stability considerably. We prefer instead to use 10-mL plastic pipets (wide mouth) when possible. Pipetting up and down the sucrose/metrizamide mixture four to five times should be enough to homogeneously mix the two solutions. 5. As pointed out before, it is critical to keep the temperature as low as possible (without freezing) to prevent protease activation. In order to do that when preparing the discontinuous metrizamide gradient, the tube should be placed inside an ice bucket (or a bigger tube filled with ice), leaving visible only the region where the metrizamide gradient is being formed. A gradient maker can be used if preferred, but with a little practice it is possible to overlay the different density metrizamide solutions with a Pasteur pipet and a rubber bulb without mixing of the phases. 6. As in any isolation procedure, it is essential to test the purity, enrichment, and stability of the isolated organelles. We analyze systematically for the activity of two different lysosomal enzymes (hexosaminidase and -Nacetylglycosaminidase) in the homogenate and in the lysosomal fraction (which allows us to calculate the recovery and enrichment in lysosomal enzymes of the fraction). As possible contamination markers we measure lactic dehydrogenase activity (cytosolic marker), succinyl dehydrogenase activity (mitochondrial marker), and catalase activity (peroxisomal marker) using standard fluorimetric and colorimetric procedures as described before (28). Preparations with more than 1% of contamination are discarded. To test the stability of the lysosomal membrane, we centrifuge an aliquot of 5 μL of the lysosomal preparation at 25,000g for 5 min at 4°C, and measure the hexosaminidase activity in the supernatant (outside lysosomes) and in the pellet (inside lysosomes). Preparations with more than 10% of hexosaminidase activity outside lysosomes are discarded and they cannot be used for uptake experiments. 7. Secondary lysosomes for CMA uptake assays can also be isolated from cultured cells as described in detail in ref. 28. Cell lysis is a critical step in that procedure in order to guarantee proper lysosomal membrane stability. Nitrogen cavitation is the most effective method to disrupt the collected cells without altering the organelle’s membranes. Although sonication or mechanical homogenization is appropriate for isolation of other intracellular organelles, they consistently result in higher than 10% lysosomal membrane breakage after isolation. 8. Although we have not found differences between GST-hsc70 and hsc70 in their ability to recognize CMA substrates and target them for degradation to lysosomes, if desired, the GST tag can be cleaved by incubation with thrombin in cleavage buffer. The amount of GST-hsc70 can be estimated running a small aliquot of the bead-bound fraction on SDS-PAGE and staining with Coomassie blue. After incubation of the beads with 0.2 to 1% w/w thrombin/fusion protein for one hour at 25°C, they are washed three to four times with wash buffer where the cleaved protein is collected.


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9. It is essential to determine that degradation is taking place inside the lysosomal lumen and not as result of the lysosomal enzymes released into the media. We systematically incubated lysosomes in parallel filtered plates at 37°C in the absence of radiolabeled substrate and measure (1) the hexosaminidase activity

Low CMA activity

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Fig. 3. Changes in the components and intracellular distribution of lysosomes active for CMA with the activation of this autophagic pathway. (A) Confluent cells with low CMA activity (i.e., growth in the presence of serum) or with high CMA activity (i.e., after prolonged (>10 h) serum removal) are subjected to immunofluorescence for LAMP-2A and hsc70 (the two main CMA components). Under conditions when CMA is maximally activated, the lysosomes active for CMA (those enriched in LAMP-2A and hsc70) relocate to the perinuclear region. (B) Increase in CMA activity usually results in an increase in the amount of CMA-active lysosomes, and this can be quantified as an increase in the colocalization of hsc70 and LAMP-2A in vesicular structures. (C) The mobilization of the CMA-active lysosomes from the cell periphery toward the perinuclear region can be quantified by drawing straight lines and comparing the distribution of the population of LAMP-2A/hsc70-positive vesicles with respect to the nucleus (distance to nucleus measured in μm).

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present at different times in the flow through (our values right after isolation are around 1–2% of the total lysosomal activity and reach 5–7% after a 30 min incubation) and (2) the proteolytic activity of the flow through when incubated with the same amount of radiolabeled protein as the experimental samples in the same conditions (to determine the percentage of degradation due to release of lysosomal enzymes). Experiments in which hexosaminidase activity or proteolysis by extralysosomal content is more than 10% of the one obtained with the total lysosomal fraction are discarded. 10. Proper controls should be added to verify that the treatment with proteinase K is effective but it is not disrupting the lysosomal membrane. We usually include an extra tube in which Triton X-100 (0.5% final concentration) is added to the samples in the presence of proteinase K. Under these conditions, the substrate is accessible to the protease and should be completely degraded. Persistence of undegraded GAPDH indicates that the amount of proteinase K is not enough to completely degrade GAPDH. To determine that the degradation by proteinase K is efficient we routinely monitor cleavage of the cytosolic tail of LAMP-2A (accessible to proteinase K) by immunoblot with a specific antibody. Persistence of the LAMP-2A tail indicates that the amount of proteinase K is not enough to remove all the GAPDH bound to the lysosomal membrane. 11. Methanol fixation is required to eliminate the soluble cytosolic hsc70 and to preserve the membrane-bound form. Routine negative controls in these studies include slides incubated with (1) only one of the secondary antibodies, (2) both secondary antibodies together, and (3) the primary LAMP-2A antibody with the secondary antibody for hsc70 and vice versa. Activation of CMA associates to the mobilization of the hsc70/LAMP-2A-enriched lysosomes from the periphery of the cell to the perinuclear area (Fig. 3). If the treatment of the cells results in changes in their shape/volume, values should be normalized to the total cellular area (calculated with the “freehand selection” and the “measure” tool of ImageJ).

Acknowledgments We would like to gratefully acknowledge the members of our laboratory for their valuable suggestions. Research in our laboratory is supported by National Institutes of Health/National Institute of Aging grants AG021904 and AG19834 and an Ellison Medical Foundation Award.

References 1. Massey, A., Zhang, C. and Cuervo, A. (2006) Chaperone-mediated autophagy in aging and disease. Curr. Top. Dev. Biol. 73, 205–235. 2. Dice, J. (1990) Peptide sequences that target cytosolic proteins for lysosomal proteolysis. Trends Biochem. Sci. 15, 305–309.


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3. Chiang, H., Terlecky, S., Plant, C. and Dice, J. (1989) A role for a 70 kDa heat shock protein in lysosomal degradation of intracellular protein. Science 246, 382–385. 4. Majeski, A. and Dice, J. (2004) Mechanisms of chaperone-mediated autophagy. Int. J. Biochem. Cell Biol. 36, 2435–2444. 5. Cuervo, A. and Dice, J. (1996) A receptor for the selective uptake and degradation of proteins by lysosomes. Science 273, 501–503. 6. Salvador, N., Aguado, C., Horst, M. and Knecht, E. (2000) Import of a cytosolic protein into lysosomes by chaperone-mediated autophagy depends on its folding state. J. Biol. Chem. 275, 27447–27456. 7. Agarraberes, F., Terlecky, S. and Dice, J. (1997) An intralysosomal hsp70 is required for a selective pathway of lysosomal protein degradation. J. Cell. Biol. 137, 825–834. 8. Terlecky, S. and Dice, J. (1993) Polypeptide import and degradation by isolated lysosomes. J. Biol. Chem. 268, 23490–23495. 9. Aniento, F., Papavassiliou, A. G., Knecht, E. and Roche, E. (1996) Selective uptake and degradation of c-fos and v-fos by rat liver lysosomes. FEBS Lett. 390, 47–49. 10. Wing, S., Chiang, H. L., Goldberg, A. L. and Dice, J. F. (1991) Proteins containing peptide sequences related to KFERQ are selectively depleted in liver and heart, but not skeletal muscle, of fasted rats. Biochem. J. 275, 165–169. 11. Cuervo, A., Knecht, E., Terlecky, S. and Dice, J. (1995) Activation of a selective pathway of lysosomal proteolysis in rat liver by prolonged starvation. Am. J. Physiol. 269, C1200–C1208. 12. Kiffin, R., Christian, C., Knecht, E. and Cuervo, A. (2004) Activation of chaperonemediated autophagy during oxidative stress. Mol. Biol. Cell. 15, 4829–4840. 13. Cuervo, A., Hildebrand, H., Bomhard, E. and Dice, J. (1999) Direct lysosomal uptake of alpha2-microglobulin contributes to chemically induced nephropathy. Kidney Int. 55, 529–545. 14. Cuervo, A. M., Stefanis, L., Fredenburg, R., Lansbury, P. T. and Sulzer, D. (2004) Impaired degradation of mutant alpha-synuclein by chaperone-mediated autophagy. Science 305, 1292–1295. 15. Cuervo, A. M. and Dice, J. F. (2000) Age-related decline in chaperone-mediated autophagy. J. Biol. Chem. 275, 31505–31513. 16. Cuervo, A. M., Dice, J. F. and Knecht, E. (1997) A population of rat liver lysosomes responsible for the selective uptake and degradation of cytosolic proteins. J. Biol. Chem. 272, 5606–5615. 17. Cuervo, A., Terlecky, S., Dice, J. and Knecht, E. (1994) Selective binding and uptake of ribonuclease A and glyceraldehyde-3-phosphate dehydrogenase by isolated rat liver lysosomes. J. Biol. Chem. 269, 26374–26380. 18. Aniento, F., Roche, E., Cuervo, A. M. and Knecht, E. (1993) Uptake and degradation of glyceraldehyde-3-phosphate dehydrogenase by rat liver lysosomes. J. Biol. Chem. 268, 10463–10470. 19. Dworniczak, B. and Mirault, M. (1987) Structure and expression of a human gene coding for a 71 kd heat shock ‘cognate’ protein. Nucl. Acids Res. 15, 5181–5197.

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20. Auteri, J., Okada, A., Bochaki, V. and Dice, J. (1983) Regulation of intracellular protein degradation in IMR- 90 human diploid fibroblasts. J. Cell Physiol. 115, 159–166. 21. Finn, P. F. and Dice, J. F. (2005) Ketone bodies stimulate chaperone-mediated autophagy. J. Biol. Chem. 280, 25864–25870. 22. Fuertes, G., Martin De Llano, J.,Villarroya, A., Rivett, A. and Knecht, E. (2003) Changes in the proteolytic activities of proteasomes and lysosomes in human fibroblasts produced by serum withdrawal, amino-acid deprivation and confluent conditions. Biochem. J. 375, 75–86. 23. Massey, A. C., Kaushik, S., Sovak, G., Kiffin, R. and Cuervo, A. M. (2006) Consequences of the selective blockage of chaperone-mediated autophagy. Proc. Natl. Acad. Sci. USA 103, 5905–5910. 24. Cuervo, A. and Dice, J. (2000) Unique properties of lamp2a compared to other lamp2 isoforms. J. Cell Sci. 113, 4441–4450. 25. Cuervo, A. and Dice, J. (2000) Regulation of lamp2a levels in the lysosomal membrane. Traffic 1, 570–583. 26. Lowry, O., Rosebrough, N., Farr, A. and Randall, R. (1951) Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275. 27. Laemmli, U. (1970) Cleavage of structural proteins during the assembly of the head of the bacteriophage T4. Nature 227, 680–685. 28. Storrie, B. and Madden, E. (1990) Isolation of subcellular organelles. Methods Enzymol. 182, 203–225.

16 Microautophagy in the Yeast Saccharomyces cerevisiae Andreas Uttenweiler and Andreas Mayer

Summary Microautophagy involves direct invagination and fission of the vacuolar/lysosomal membrane under nutrient limitation. In Saccharomyces cerevisiae microautophagic uptake of soluble cytosolic proteins occurs via an autophagic tube, a highly specialized vacuolar membrane invagination. At the tip of an autophagic tube vesicles (autophagic bodies) pinch off into the vacuolar lumen for degradation. Formation of autophagic tubes is topologically equivalent to other budding processes directed away from the cytosolic environment, e.g., the invagination of multivesicular endosomes, retroviral budding, piecemeal microautophagy of the nucleus and micropexophagy. This clearly distinguishes microautophagy from other membrane fission events following budding toward the cytosol. Such processes are implicated in transport between organelles like the plasma membrane, the endoplasmic reticulum (ER), and the Golgi. Over many years microautophagy only could be characterized microscopically. Recent studies provided the possibility to study the process in vitro and have identified the first molecules that are involved in microautophagy.

Key Words: Calmodulin; microautophagy; vacuole; VTC (vacuolar transporter chaperone); yeast.

1. Introduction When yeast cells sense nutrient (e.g., nitrogen) limitation, they stop dividing and enter the stationary phase. They adapt to the new environmental conditions by breaking down large quantities of cellular constituents by autophagy. The phenomenon enables cells to survive long periods of starvation. Autophagy is a highly conserved lysosomal transport and degradation pathway of all eukaryotic cells (1). In the yeast Saccharomyces cerevisiae the lysosomal From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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compartment is called the “vacuole.” It is the main compartment for the storage, recycling, and breakdown of cellular constituents. The organelle can easily be stained for in vivo fluorescence microscopy. Yeast vacuoles can be prepared in milligram amounts per day. This is why vacuoles represent an excellent tool for biochemical studies. The highly dynamic vacuolar organelle undergoes morphological changes dependent on the cell cycle and environmental changes such as nutrient availability (Fig. 1). Concomitant with daughter cell emergence, vacuoles begin to fragment into small vesicles that are transported into the daughter cell. There, the vesicles fuse to build up a new vacuolar compartment.

Fig. 1. Morphological changes of the yeast vacuole. When cells divide, vacuoles fragment into small vesicles that are transported into the daughter cell. There they fuse to build up new vacuoles. When cells are starved they stop dividing. Then, the cell compensates the influx of lipid and protein into vacuolar membranes that is caused by macroautophagy. This is achieved by means of microautophagy, a direct membrane invagination, and fission of autophagic bodies into the vacuolar lumen.

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Upon entry into stationary phase, yeast cells develop a large single central vacuole, which then degrades large quantities of cytosol and/or organelles. There are two entry pathways for bulk cytosolic compounds: Macroautophagy is defined as the formation of double-layered vesicles (autophagosomes) that enclose cytosolic compounds. They target these compounds for degradation by fusing their outer membrane with vacuoles (2). Vacuolar fusion and macroautophagy have been studied intensively over the last decade. In contrast, little is known about microautophagy, a process comprising a direct invagination and budding of vesicles (autophagic bodies) into the vacuolar lumen. It has been characterized microscopically (Fig. 2) (3) and could be reconstituted in a cell-free system composed of purified vacuoles and cytosolic extracts. This in vitro system measures the uptake of a luciferase reporter substrate (Fig. 3) (4). Using pharmacological substances, the in vitro uptake reaction can be dissected into different kinetic stages (5). According to their ability to block the reaction at different kinetic stages, these inhibitors have been defined as early-acting class A inhibitors (nystatin, GTP?S, aristolochic acid) and lateacting class B inhibitors (W-7, valinomycin/FCCP, K252a and rapamycin) (5). Recent findings have identified the first molecular players acting during microautophagy. Both apocalmodulin (i.e., the calcium free conformation of calmodulin) and the vacuolar transporter chaperone (VTC) complex act late in the reaction (6,7). Microautophagy of soluble cytosolic components is topologically equivalent to invaginations occurring during (1) multivesicular body (MVB) formation at the endosome (8,9), (2) retroviral budding at the endosome or the plasma membrane (10), (3) piecemeal microautophagy of the nucleus (PMN), which tranfers parts of the nucleus into vacuoles (11), and (4) micropexophagy in methylotrophic yeasts which leads to the degradation of peroxisomes (12– 17). Although microautophagy of soluble components, like macroautophagy, is induced by nitrogen starvation and Rapamycin (a pharmacological agent inhibiting Tor kinase signaling) and although pexophagic vacuole invagination depends on Atg proteins (18–21), there is no evidence to date that Atg proteins are directly involved in either PMN (11) or microautophagic uptake. Macroautophagy seems to be a prerequisite for sustained microautophagy, however (4). Microautophagy is controlled by the TOR and EGO (composed of Ego1p, Gtr2p, and Ego3p) signaling complexes (22). It leads to direct uptake and degradation of the vacuolar boundary membrane. The main functions of microautophagy may lie in the transition from starvation-induced growth arrest to logarithmic growth (22) and in maintenance of organellar size and membrane composition (3). Since microautophagy leads to uptake and degradation of the vacuolar boundary membrane, it could compensate the enormous influx of membrane caused by macroautophagy.

Fig. 2. Microautophagic invaginations in vivo as seen by (A) fluorescence microscopy using a strain expressing GFP-Vtc4p and (B) ultrathin sectioning electron microscopy. Bars correspond to 1 μm in (A) and (B).


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Fig. 3. Schematic presentation of reconstituted in vitro system for microautophagy. Isolated vacuoles are incubated with cytosol, buffer, salts, ATP-regenerating system, and the uptake substrate for 1 h at 4 or 27°C. After the uptake reaction, the vacuoles are washed thoroughly and processed for detection of the uptake substrate (firefly luciferase or FITC-dextran) (4).

Microautophagic vacuole invagination (with the exception of PMN and pexophagy) might hence be responsible for maintenance of organellar size and membrane composition rather than for cell survival under nutrient restriction. A function in organelle homeostasis should render microautophagic membrane invagination dependent on membrane influx via macroautophagy, which has been observed (4). This hypothesis is also consistent with ultrastructural data. In freeze-fracture electron microscope nascent microautophagic vesicles that bud off from the tips

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of autophagic tubes appear as virtually free from intramembranous particles, i.e., there appears to be lateral heterogeneity induced along the autophagic tube that prevents large membrane proteins from entering the microautophagic vesicle membrane. Microautophagic vesicles share this exceptional ultrastructural feature with nascent autophagosomes, which form in the cytosol by macroautophagy and are also are virtually free of intramembraneous particles. This suggests that microautophagy might compensate macroautophagic membrane influx in terms of both quantity and quality. The yeast vacuole is a highly dynamic organelle that undergoes dramatic morphological changes (Fig. 1). Therefore, the vacuole serves as an excellent model system to study membrane dynamic processes. The organelle can easily be stained for in vivo fluorescence microscopy. Besides, yeast vacuoles can be prepared in milligram amounts per day making vacuoles an excellent system for biochemical studies and in vitro assays. These assays allow one to reconstitute authentic reactions with a minimum of factors necessary to drive the process. These factors (e.g., vacuoles, cytosol, purified proteins, salts, chemicals) can be added independent of each other. In addition, they serve to render the process accessible to defined concentrations of pharmaceuticals and compounds (e.g., purified proteins or reporter molecules) which cannot easily penetrate whole cells due to their chemical nature or size. In vitro reactions can also be arrested at different stages by inhibitors to define kinetic phases. Often certain protein conformations or interactions accumulate in these phases and can then be further characterized. This allows one to not only identify new factors implicated in the process but also to explore their mechanisms of action in detail.

2. Materials 2.1. Buffers 1. PS buffer: 10 mM piperazine-N,N -bis(2-ethanesulfonate) [PIPES]/KOH pH 6.8, 200 mM sorbitol. 2. Cytosol buffer: 40 mM PIPES/KOH, pH 6.8, 0.5mM MgCl2 , 150 mM KCl, 200 mM sorbitol, 1 mM dithiothreitol [DTT], 0.2 mM phenylmethylsulfonyl fluoride [PMSF], 0.1 mM pefabloc SC, 0.5 μg/mL pepstatin A, 50 μM o-phenanthroline. 3. Spheroplasting buffer for vacuole preparation: 50 mM potassium phosphate, pH 7.5, 600 mM sorbitol in YPD with 0.2% glucose. 4. Washing buffer for lyticase preparation: 25 mM Tris-HCl, pH 7.4.

2.2. Media 1. YPD: 1% yeast extract, 2% Bacto peptone 2% glucose. 2. YPD containing 200 nM Rapamycin (Alexis, dissolve 100 X in DMSO).


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3. SD(-N): 0.67% Difco yeast nitrogen base without amino acids and without ammonium sulfate, 2% glucose, for starvation. 4. LB medium: 2% tryptone, 1% yeast extract, 1% NaCl, pH 7.

2.3. Cytosol Preparation Cytosol is usually prepared from yeast strain K91-1A. This strain is deficient for soluble phosphatates. This is advantageous when cytosol preparations are used for in vitro microautophagy assays, especially when values are normalized to vacuolar alkaline phosphatase activity. 2.4. Lyticase for Vacuole Preparation -1,3-Glucanase from Oerskovia xanthineolytica (23,24) is expressed in E. coli strain RSB 805. The protein is purified from the periplasmic space (25). 2.5. Fluorescence Microscopy 1. FM4-64 (Molecular Probes) is dissolved as a 100X stock solution (10 mM) in dimethyl sulfoxide (DMSO) or ethanol and stored at –20°C. 2. For immobilization of yeast cells use Seaplaque agarose in 10 mM PS buffer.

3. Methods 3.1. Yeast Cell Culture Yeast cells are precultured in YPD for 6–8 h at 30°C and then diluted for logarithmic overnight growth (14–16 h, 30°C, 225 rpm) in 2-L Erlenmeyer flasks with 1 L of YPD medium. For starving cells, overnight cultures were harvested at an optical density (OD 600) of 2, centrifuged (4 min 3800g), washed with sterile water, resuspended in 1 L of SD(-N) starvation medium, and incubated (3 h, 30°C, 225 rpm) (see Note 2). 3.2. Cytosol Preparation 1. Harvest overnight yeast cultures at OD600 = 4.5, centrifuged (4 min, 3800g), wash with sterile water, harvest again, resuspend in 1 L of SD(-N), and incubate for 3–4 h at 30°C and 255 rpm. For preparation of cytosol from nonstarved cells, replace SD(-N) by YPD in this incubation. 2. Harvested cells and wash as described above, first with water, and then with one pellet volume of chilled cytosol buffer. After centrifugation (5 min, 3,800g 4°C), resuspend the pellet in a small volume of cytosol buffer so that a thick slurry results.

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3. Freeze the suspension as little nuggets in liquid nitrogen and blend (6–8 times for 30 s) in a Waring blender filled with liquid nitrogen. 4. Thaw cells and centrifuge the lysate (10 min, 12,000g, 4°C). Ultracentrifuge the supernatant (20 min, 125,000g, 2°C), discard the fatty top fraction, and recover the clarified cytosol. 5. Adjust the protein concentration of the cytosolic fraction to 25–30 mg/mL with cytosol buffer (do not use preparations below 10 mg/mL for in vitro microautophagy assays). Freeze 50- to 300-μL aliquots in liquid nitrogen and store at –80°C.

3.3. Lyticase Preparation for Vacuole Preparation 1. Grow 10 l RSB 805 in LB medium with 100 μg/mL ampicillin to OD600 = 0.6. 2. Add 0.4 mM isopropyl thriogalactose (IPTG) and let cells grow for another 5 h at 30°C. 3. Harvest cells (10 min, room temperature, 4200g), wash with 50 mL washing buffer and centrifuge (10 min, room temperature, 4200g). 4. Resuspend pellets in 200 mL washing buffer, supplemented with 2 mM EDTA and an equal volume of 40% (w/v) sucrose in washing buffer and gently shake at room temperature for 20 min. 5. Harvest cells (10 min, room temperature, 4200g) and completely remove the supernatant. Chill flasks on ice. 6. Resuspend the cells from 1 L of culture in 20 mL ice cold 0.5 mM MgSO4 , shake gently (4°C, 20 min) and centrifuge (10 min, 4°C, 4200g). 7. The supernatant contains Lyticase and some E. coli periplasmic proteins. Adjust the protein concentration of the supernatant to 1 mg/mL. The lyticase solution can be stored at –20°C in 6-mL aliquots without further purification. It should be thawed directly before use for digestion of yeast cell walls (see Note 3).

3.4. Vacuole Preparation 1. Grow cells in YPD overnight to OD600 = 2.5-3.0, harvest (2 min, 3,800 g, 4°C), and resuspend them in 50 mL of 30 mM Tris-HCl, pH 8.9 with 10 mM DTT. Incubate cells (5 min, 30°C). 2. Centrifuge cells as above, resuspend in 12 mL of spheroplasting buffer, add 3 mL of a lyticase solution (concentration of stock solution: 1 mg/mL) and transfer into 30-mL Corex tubes. Incubate cells (25 min, 30°C) (see Note 2). 3. Reisolate spheroplasts (4°C, 1 min, 2500g) and resuspend in 2.5 mL 15% Ficoll 400 in PS buffer by gentle stirring with a plastic rod and/or gentle vortexing. 4. Add DEAE-dextran (300 μL) from a 0.4-mg/mL stock in 15% Ficoll 400 in PS buffer. Incubate spheroplasts (2 min at 0°C, then 90 s at 30°C), chill again, transfer to an SW41 ultracantrifugation tube, and overlay with 3 mL 8% Ficoll 400, 3 mL 4% Ficoll 400, and 1.5–2 mL 0% Ficoll 400 in PS buffer. After centrifugation (85 min, 154,000g, 2°C,), harvest vacuoles from the 0–4% interphase.


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5. For storage of vacuoles, add glycerol (final concentration 10% w/v from a 50% stock) to a fresh vacuole suspension. Freeze the suspension as little nuggets in liquid nitrogen and store them at –80°C.

3.5. Fluorescence Microscopy 3.5.1. Staining of Vacuoles In Vivo 1. Grow cells logarithmically in 5 mL of YPD overnight. (20-mL tubes, 30°C, 225). Then, supplement 1 mL of the suspension with 10–20 μM FM4-64 and incubate for 1 h. 2. Reisolate the cells (2 min, 1300g), wash twice with deionized water, reisolate again, and resuspend to an OD600 of 0.5-1 in YPD, YPD containing Rapamycin, or SD-(N) medium without stain. 3. Let cells grow in these media for 3–4 h. Centrifuge 400 μL of the cell suspension (1 min, 12,000 g) and resuspend the cells in 40 μL of the identical medium for concentration. 4. Transfer 5–8 μL of the concentrated cell suspension to a slide, cover immediately with a cover slip, and investigate immediately with a confocal microscope or a convential fluorescence microscope (see Note 4).

3.5.2. GFP-Vtc-Proteins 1. Grow cells logarithmically in 5 mL of YPD overnight. (20-mL tubes, 30°C, 225). 2. Reisolate the cells (2 min, 1300g), wash twice with deionized water, reisolate again, and resuspend to an OD600 of 0.5-1 in YPD, YPD containing Rapamycin, or SD-(N) medium. 3. Let cells grow in these media for 3–4 h. Centrifuge 400 μL of the cell suspension (2 min, 1300g) and resuspend the cells in 40 μL of the identical medium for concentration. 4. Transfer 5–8 μL of the concentrated cell suspension to a slide, cover immediately with a cover slip, and investigate immediately with a confocal microscope or a convential fluorescence microscope.

3.6. Electron Microscopy 3.6.1. Thin Sectioning for Ultrastructural Studies To cryoimmobilize yeast cells by high-pressure freezing proceed as described previously (26): 1. Suck living specimen into cellulose microcapillaries of 200 μm diameter and transfer 2-mm-long capillary tube segments to aluminum platelets of 200 μm depth containing 1-hexadecene.

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2. Sandwich the platelets with platelets without any cavity and then freeze with a high-pressure freezer (Bal-Tec HPM 010, Balzers, Liechtenstein). Remove extraneous hexadecene from the frozen capillary tubes under liquid nitrogen and transfer to 2 mL-microtubes with screw caps (Sarstedt #72.694) containing the substitution medium precooled to –90°C. 3. Keep samples for ultrastructural studies in a freeze-substitution unit (Balzers FSU 010, Bal-Tec, Balzers, Liechtenstein) in 2% osmium tetroxide in anhydrous acetone at –90°C for 32 h, warm up to –60°C within 3 h, keep at –60°C for 4 h, warm up to –40°C within 2 h, and keep there for 4 h. 4. Wash the samples with acetone and transfer them into an acetone-Epon mixture at –40°C, infiltrate at room temperature in Epon, and polymerize at 60°C for 48 h. 5. View ultrathin sections, stained with uranyl acetate and, if required, with lead citrate, in a Philips CM10 electron microscope at 60 kV.

3.6.2. Freeze-Fracture Analysis 1. Sandwich living yeast cells or isolated vacuoles between thin copper sheets (BalTec; Balzers AG), mount on tweezers, an rapidly inject into melting propane (–185°C), as described before (27). 2. Insert the sandwiches under liquid nitrogen into a freeze-fracture unit Type BAF 300 (Balzers AG), fracture at –100°C, and replicate by 45° platinum-shadowing. 3. Transfer replicas into 2.5% sodium dodecyl sulfate (SDS) with 30 mM sucrose in 10 mM Tris-HCl buffer, pH 8.3. After vigorous shaking for 30 min, wash replicas several times with distilled water, and mount onto copper grids for routine electron microscopic analysis in an electron microscope (EM) 10 (Zeiss).

3.6.3. Thin Sectioning for Immunogold Labeling 1. Process samples as described in Subheadings 1. and 2. for thin sectioning for ultrastructural studies. 2. Process samples for immuno/affinity labeling in 0.5% acrolein or uranyl acetate (depending) in anhydrous ethanol using the same temperature/time schedule for freeze-substitution. 3. After washing with ethanol, transfer the samples into an ethanol-Lowicryl K11M mixture, infiltrate with the polar methacrylate resin Lowicryl K11M (Polysciences, Eppelheim, Germany) and polymerize by UV irradition at –40°C for 48 h. 4. Immunolabeling can be done as described previously (28,29) using affinity purified antibodies (typically 0.5–3 μg/mL in 0.1% acetylated bovine serum albumen [BSA] or 0.2% gelatine and 0.5% BSA in phosphate=buffered saline [PBS]) to proteins or lipids of interest. Detect antibodies from rabbit or mouse with protein A labeled with 15-nm gold particles (the signal for antibody derived from goat should be enhanced with anti-goat antibody in between). View ultrathin sections, stained with uranyl acetate, in a Philips CM10 EM at 60 kV.


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3.7. Reconstituted Microautophagic Assay In Vitro Microautophagic activity can be reconstituted in a cell-free system composed of purified vacuoles and cytosolic extracts (4). Vacuoles internalize a reporter enzyme (firefly luciferase, or, if assay by microscopy is desired, fluorophores coupled to high MW carriers—e.g., FITC-dextranes) in an ATP-dependent fashion. After the uptake reaction, internalized luciferase reporter can be reisolated with the vacuoles: It is protected against proteinase K digestion by the vacuolar boundary membrane and by the membrane of the microautophagic vesicle. After proteolytically removing luciferase that has not been taken up, the vacuolar membranes can be lysed and reporter enzyme activity can be quantified by a chemoluminescent assay.

3.7.1. Standard Reaction 1. A standard reaction has a volume of 45 μL and contains: vacuoles (0.2 mg/mL, either freshly prepared or thawed from a –80°C stock), 3 g/mL cytosol from starved cells, 105 mM KCl, 7 mMMgCl2 , 2.2 mM ATP, 88 mM disodium creatine phosphate, 175 U/mL creatine kinase, 17 μg/mL luciferase, 100 μM DTT, 0.1 mM pefabloc SC, 0.5 mM o-phenanthrolin, 0.5 μg/mL pepstatin A, 200 mM sorbitol, 10 mM PIPES/KOH pH 6.8 (see Note 5). Incubate this mixture for 1 h at 27°C. 2. For measuring luciferase uptake, chill the samples on ice, dilute with 300 μL 150 mM KCl in PS buffer, centrifuge (6500g, 3 min, 2°C), wash the pellet once more with 300 μL 150 mM KCl in PS buffer by gently pipetting three to four times up and down with a 1-mL pipet. 3. Centrifuge again and resuspended in 55 μL150 mM KCl in PS buffer by shaking on a shaker (1400 rpm, 8 min, 4°C). 4. Add Proteinase K (0.3 mg/mL from 18x stock) and incubate on ice for 23 min. Stop digestion by adding 55 μL 1 mM PMSF/150 mM KCl in PS buffer. 5. Determine luciferase activity using an assay kit according to the manufacturer’s instruction (Berthold Detection Systems, Pforzheim, Germany): mix 25 μL sample with 25 μL lysis buffer and add 50 μL substrate mix (30) directly before counting light emission in a microplate luminometer (LB 96 V, Berthold Technologies, Bad Wildbad, Germany) (see Note 1). 6. If you run uptake reactions in the presence of pharmaceuticals or antibodies, we recommend determining alkaline phosphatase activity in a 25-μL aliquot as described previously (4). This serves as an internal reference for the quantity of pelleted vacuoles. Calculate uptake activity as the quotient of luciferase activity over alkaline phosphatase activity (counts per second/ OD405 per min) and normalize to a standard reaction run without any pharmaceuticals or antibodies (60 min, 27°C), which is set to 100%. When comparing different yeast strains, check the level of mature alkaline phosphatase in vacuolar preparations (e.g., by Western blotting or alkaline phosphatase assay). Protein levels can differ in

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Uttenweiler and Mayer mutants in comparison to wild-type strains. In these cases uptake activity is not referred to alkaline phosphatase activity.

3.7.2. Kinetic Analysis Using a pharmacological approach including low molecular weight inhibitors, the in vitro uptake reaction can be dissected into different kinetic stages (5). According to their ability to block the reaction at different kinetic stages, these inhibitors have been defined as early-acting class A inhibitors (nystatin, GTPS, aristolochic acid) and late-acting class B inhibitors (W-7, valinomycin/FCCP, K252a, and rapamycin). For this kind of analysis, run standard uptake reactions and add inhibitors at different time points. Then, continue the incubation until the end of a standard 60-min reaction period. Inhibitors influencing very early steps of microautophagy (class A inhibitors) lose their activity if added late during the reaction, whereas inhibitors acting on late events of microautophagy (class B inhibitors) remain active throughout the reaction. As a control, transfer samples to ice, which also stops the reaction (Fig. 4). 3.7.3. Rapid Uptake Late-acting inhibitors in microautophagy can be further classified depending on their ability to inhibit rapid uptake (5). Rapid uptake of luciferase occurs after preincubation of vacuoles under standard conditions supporting microautophagic membrane invagination, but in the absence of the reporter enzyme. It is assumed that this allows the vacuoles to complete all preparatory reactions for uptake, e.g., to form an invagination, without producing a luciferase signal. The formation of vesicles can then be scored by adding luciferase for a short period of time. This allows only rapid uptake from preformed invaginations but is too short for the formation of new invaginations. Thus, this criterion can be used to distinguish the preparation for uptake (tube formation) from its completion (vesicle scission). 1. After 60 min of incubation without luciferase, add the reporter enzyme to an uptake reaction. Incubate the samples for another 5 min to permit reporter enzyme uptake. 2. Terminate uptake by chilling, diluting, and centrifuging the reactions and analyze pelleted vacuoles for luciferase uptake as described above.

4. Notes 1. Buffers. Do not add leupeptin to the lysis buffer for cytosol preparation because this inhibits the firefly luciferase reporter enzyme.


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(a)

(b)

Fig. 4. Kinetic analysis of in vitro microautophagy. (A) Schematic example of a time course experiment with inhibitors. In vitro microautophagic reactions are started. At the indicated time points, the samples receive inhibitors, or control buffer, and are incubated further at 27°C until the end of the 60-min reaction period. For the ice curve, an aliquot is set on ice at the indicated time points and kept there until the end of the reaction period. Reactions performed without inhibitor at 27°C are set to 100% (B) Schematic overview of the kinetic steps. The in vitro assay can be inhibited at different stages by addition of inhibitors or by transfer to 0°C. GTPS: guanosine 5O-(3-thiotriphosphate); FCCP: p-trifluoromethoxy carbonyl cyanide phenyl hydrazone; VTC: vacuolar transporter chaperone. (Adapted from refs. 5 and 6.).

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2. Yeast cell culture. For temperature-sensitive yeast strains, all steps are carried out at 25°C as long as no temperature shock is desired. We usually carry out temperature shock at 37°C for the desired time. Note that heating of large volumes (e.g., 1 L) from 25 to 37°C will take a considerable time. 3. Vacuole preparation. The optimal OD600 for harvesting cells, the time for spheroplasting and the amount of DEAE-dextran added can vary for different yeast strains. If necessary, vacuoles can be prepared with up to 1 mM PMSF in the spheroplasting buffer to reduce protein degradation. 4. Fluorescence microscopy. To avoid fast movement of cells when investigated with a fluorescence microscope, cells can be immobilized the following way: 7 μL of the cell suspension is mixed with 7μL of 0.4% low-melting-point agarose in PS buffer (kept liquid at 35°C). Twelve μL of this mixture are transferred to a slide, covered immediately with a cover slip, and chilled at 4°C for 5 min to immobilize the cells. The intensity of illumination has to be minimized to avoid structural damage to the vacuoles that can occur at higher light intensities or after prolonged illumination of the same field. 5. Reconstituted microautophagic assay in vitro. Prepare all samples on ice. Thaw cytosol/vacuole nuggets directly before use and do not freeze again. Direct comparisons of activity should be performed only with vacuoles from the same batch of preparation because there can be variations in absolute activities from batch to batch. These are often related to changes the composition of media ingredients supplied. Our experience is that the quality of, e.g., peptones or yeast extracts varies considerably over the year, even when purchased from the same supplier. Addition of antibodies/pharmaceuticals to the in vitro reaction can induce formation of large clusters of vacuoles, which may be hard to resuspend. In theses cases you should prolong the resuspension steps carefully without destroying the vacuoles. Alternatively, prepare, e.g., Fab fragments from antibodies.

Acknowledgments This work was funded by grants from the DFG (SFB466-A10), BMBF, SNF, and Boehringer Ingelheim Foundation to A.M. and from the Boehringer Ingelheim Fonds to A.U.

References 1. Reggiori, F., and Klionsky, D. J. (2002) Eukaryot. Cell 1, 11–21. 2. Baba, M., Takeshige, K., Baba, N., and Ohsumi, Y. (1994) J. Cell Biol. 124, 903–913. 3. Muller, O., Sattler, T., Flotenmeyer, M., Schwarz, H., Plattner, H., and Mayer, A. (2000) J. Cell Biol. 151, 519–528. 4. Sattler, T., and Mayer, A. (2000) J. Cell Biol. 151, 529–538. 5. Kunz, J. B., Schwarz, H., and Mayer, A. (2004) J. Biol. Chem. 279, 9987–9996.


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17 EM Analysis of Phagosomes Chantal de Chastellier

Summary Electron microscopy (EM) is the only technique that can combine sensitive proteindetection methods with detailed information on the substructure of cellular compartments. The purpose of this chapter is to describe some of the methods at the EM level that can be used to analyze the spatial organization of cell organelles with respect to phagosomes or vacuoles in which pathogens are sequestered, characterize the compartment in which pathogens are harbored, ie immature phagosomes, phagolysosomes, autophagosomes, and ER-derived vacuoles, to cite a few, and decipher the molecular mechanisms involved in survival of pathogens within infected host cells.

Key Words: Electron microscopy; enzyme cytochemistry; phagosomes; autophagosomes; pathogens.

1. Introduction Intracellular pathogens have evolved a wide variety of strategies to manipulate host cell organelles and/or constituents, thus enabling them to find favorable conditions for survival and multiplication (for review see, e.g, refs. 1 and 2). After binding to cell surface receptors, microorganisms and particles are internalized by phagocytosis into membrane-bound compartments called phagosomes. Under normal conditions, the newly formed phagosomes intermingle contents and membrane with the successive compartments of the endocytic pathway (early endosomes, late endosomes, lysosomes) through a complex series of fusion and fission events. As they are processed into phagolysosomes, they undergo gradual modifications by specific addition and removal of membrane constituents. In addition, they become acidified due From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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to the vacuolar proton pump ATPase located in the membrane and acquire toxic constituents, including hydrolases, which will ultimately destroy bacteria. One of the major strategies used by endoparasites, but by no means the only one, is to modulate these interactions. A wide variety of situations have been described. Pathogens can (1) use the acidic and hydrolase-rich phagolysosomal environment to survive and multiply or (2) avoid the cytolytic environment of the phagolysosome by preventing phagosome maturation and fusion with lysosomes at different steps of the endocytic pathway. Mycobacteria, for example, do not inhibit phagosome–lysosome fusions directly but rather affect the preceding step of phagosome maturation without which fusion with lysosomes cannot occur (reviewed in refs. 3 and 4). Other pathogens escape the endocytic pathway. They can either (1) escape from the phagosome, after lysis of the phagosome membrane, and invade the cytoplasm, in which they multiply, (2) exclude, from the phagosome membrane, plasma membrane–derived constituents and/or non-plasma membrane–derived fusion-mediating factors, thereby depriving the phagosome membrane of recognition signals required for fusion with the successive compartments of the endocytic pathway, or even (3) segregate from the endocytic pathway to interact with the endoplasmic reticulum in which they multiply. Whatever the strategy used to modulate or prevent interactions with compartments of the endocytic pathway, it is of the utmost importance since it will profoundly affect drug targeting to the intracellular site of replication of pathogens and also antigen presentation. More recently, it has been shown that under certain conditions, several pathogens can be sequestered within autophagic compartments (5; reviewed in ref. 6). The fate of bacteria within autophagosomes as well as the significance of these observations remains unclear, but autophagy might have implications for the development of vaccines against such pathogens. Our understanding of the molecular mechanisms of pathogen survival depends a great deal on the tools and techniques we use to obtain information about the pathogen and the cellular machinery. Electron microscopy methods, especially when combined with molecular biology tools (mutants, knockouts) or with drugs that modify the cellular machinery, are extremely valuable tools to unravel the cellular and molecular mechanisms that enable pathogens to survive and multiply within the host cells. EM is indeed at the highest-resolution limit of a spectrum of complementary morphological techniques. It is the only technique that can combine sensitive protein-detection methods with detailed information on the substructure of intracellular compartments and on the spatial organization of organelles within the cell, especially when the latter are in intimate contact but not necessarily interacting with one another.

EM Analysis of Phagosomes

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Before going into the more complex and sensitive EM protein-detection approaches, it is important to master the landscape in which pathogenic bacteria survive and replicate within their host cell. I have, therefore, chosen to devote this chapter to basic conventional EM and to a number of methods, including enzyme cytochemistry, that are extremely useful in gaining insight into many relevant questions regarding pathogen survival within its host, such as: 1. What is the morphological state of bacteria, i.e., are they intact, altered, or damaged (intact as well as certain altered bacteria are usually live, and damaged ones are dead)? 2. What is the correlation between CFU counts and replication, in particular when CFU counts remain stationary; is it because bacteria have become dormant or because some bacteria are being degraded while others multiply? 3. Do bacteria replicate in a single, increasingly larger, phagosome, or does bacterial division induce separation of the phagosome? 4. Is there any specific interaction between the pathogen and the phagosomal membrane? For example, is a close apposition between the phagosome membrane and the bacterial surface all around the bacterium required for prevention of normal phagosome processing? 5. Do the phagosomes contain vesicular or tubular structures of either bacterial or cellular origin which might be important for bacterial replication? 6. What is the spatial organization of the phagosome with respect to other cellular compartments such as the endocytic organelles, the endoplasmic reticulum, or mitochondria? 7. Do phagosomes intermingle content and membrane with the successive compartments of the endocytic pathway, or do they block phagosome processing at some stage? 8. Do they escape the endocytic pathway and interact with other organelles? 9. Do bacteria lyse the phagosome membrane to reside in the cytoplasm? 10. Do bacteria induce a reorganization of the cytoskeletal network, thereby modifying the interactions with the organelles of the endocytic pathway?

Given the broad spectrum of EM methods and applications, it is not possible to describe here all the possible methods that one can use. The analysis of phagosome membrane composition by quantitative autoradiography (described in ref. 7) or by a variety of EM immunolocalization approaches (described in refs. 8 and 9), as well as more recently developed approaches such as electron tomography for three-dimensional imaging (see, e.g., ref. 10) will not be discussed here. Although many EM approaches have come within reach of every laboratory that is willing, and able, to invest resources, and although EM equipment has become increasingly user-friendly, consultation and training with experienced researchers who are familiar with the cell and pathogen ultrastructure is invaluable for learning EM methods, especially these more sophisticated ones, and ensuring that data are interpreted accurately.

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2. Materials 2.1. Basic Conventional Transmission Electron Microscopy (TEM) Many EM products are very toxic and even carcinogenic. Handle with care, under fume hood, and follow security measures indicated on product data sheets. 2.1.1. Products 1. Glutaraldehyde, 25% solution in water (EM grade I, Sigma, St. Louis, MO, USA). Store at –20°C. Once thawed, store at 4°C, no longer than 1 mo. Avoid putting in refrigerator containing cell culture products as fumes might affect such products. Very toxic. 2. Osmium tetroxide 4% solution in water (Electron Microscopy Sciences [EMS], Hatfield, PA, USA or Sigma). Store sealed ampullae at 4°C. Once opened, keep remainder in a clean glass vial with a glass stopper. Stable at 4°C for several months. If solution turns brown or black, discard. Fumes will blacken refrigerator: seal vial with parafilm, store in a plastic container also sealed with parafilm. Avoid putting in same refrigerator as cell culture products as fumes will affect such products. Very toxic. 3. Cacodylic acid, sodium salt, trihydrate (Sigma or EMS). Store at room temperature (RT). Toxic: contains arsenate. 4. CaCl2 and MgCl2 (Sigma). Prepare 1M solutions. Store at RT. 5. Uranyl acetate (EMS or Merck, White house Station, NJ, USA). Store powder at RT. Radioactive. 6. 5,5’-Diethylbarbituric acid, sodium salt (Fluka, Sigma-Aldrich, Buch, SG, Switzerland) for preparation of Na-veronal buffer. Store powder at RT. Very toxic. Regulations on purchase and delivery are strict in many countries. If unavailable, buy maleic acid (Sigma) for preparation of Na-H-maleate-NaOH buffer instead of Na-veronal buffer. 7. Agarose low melting point, molecular biology grade (Sigma). 8. Molecular sieve dehydrate with indicator for drying solvents (Fluka). Store at RT. Toxic, carcinogenic by inhalation. 9. Ethyl alcohol 96.2% and acetone (Carlo Erba, Rodano, Italy). 10. Spurr resin. The medium-grade TLV low-viscosity resin premix kit, from TAAB, Reading, Berkshire, UK, gives excellent results. Store at RT (see Note 1). Very toxic. 11. Lead nitrate (Fluka). Store at RT. Very toxic. 12. Other products of common use: sucrose, sodium acetate, trisodic sodium citrate, NaCl, NaOH, HCl 1 N. 13. Rubber policeman. 14. Microvettes CB 300 Z (Sartorius, Goettingen, Germany). 15: Microcentrifuge with fixed vertical axis or with free angle rotor. 16. Glass vials (content: 2–3 mL).


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17. Flat embedding molds, single tapered ends, in clear silicone. Measures: 14 mm(L) × 5 mm (W) 4 mm(D) (from EMS). Gelatin or plastic capsules of different shapes and sizes can also be used (see with local EM facility). 18. 60°C incubator.

2.1.2. Preparation of Buffers, Dehydrating Agents, Resins, Stains for EM Processing 1. Na-cacodylate buffer 0.1 M, pH 7.2 + 5 mM CaCl2 and 5 mM MgCl2 +/– 0.1 M sucrose: Dissolve, e.g., 5.35 g Na-cacodylic acid in 220 mL distilled water, adjust pH to 7.2 with HCl 1 N, complete to 250 mL with distilled water. Add 1.25 mL CaCl2 1 M and 1.25 mL MgCl2 1 M. When necessary, add 8.5 g sucrose to maintain osmolarity. Store at 4°C. Stable for several months but beware of molds. Very toxic. 2. Na-veronal buffer, pH 6.0. (a) Solution A: dissolve 1.94 g of sodium acetate, 2.94 g of 5,5´- diethylbarbituric acid, sodium salt, and 2.8 g NaCl in 100 mL distilled water. Store at 4°C until further use. Stable for several months. (b) Na-veronal buffer: Mix 20 mL of solution A, 28 mL of HCl 0.1 N and 56 mL of distilled water. Adjust pH to 6.0 with a few drops of 10% acetic acid if necessary. Add 2.5 mL of CaCl2 1 M. Stable at 4°C for 2–3 mo. If crystals form, warm up at 37°C before use. 3. 0.05 M Na-H-Maleate-NaOH buffer, pH 6.0. Dissolve 0.58 g maleic acid and 0.2 g NaOH in 20 mL distilled water, adjust pH to 6.0 with NaOH, complete to 100 mL with distilled water. Store at 4°C. 4. Uranyl acetate 1% in Na-veronal buffer or in Na-H-maleate-NaOH buffer. Dissolve uranyl acetate in buffer, e.g., 250 mg in 25 mL, in a brown glass bottle (because light sensitive) and store at 4°C. Stable for 1 mo. 5. Agar 2% solution in Na-cacodylate buffer devoid of sucrose. 6. Graded series of ethanol and acetone, i.e., 25, 50, 75, 90 and 100% solutions. The 100% solution must be completely dried as follows: add the commercial solution (∼96%) to a glass bottle (with a glass stopper) containing molecular sieve, mix vigorously, and let settle for at least 24 h before use. Avoid mixing before use. Stable at RT for several months. 7. Spurr resin: Follow instructions, i.e., add all three components in same vial, mix vigorously, let settle for an hour to remove bubbles, use immediately afterward or store in 5- or 10-mL syringes (use only Norm-ject devoid of black rubber stopper inside the syringe). Add caps to tip to avoid contact with air and store at –20°C. Once thawed, use content of syringe on the same day; do not refreeze. 8. Uranyl acetate 1% in distilled water for staining of thin sections. Should be available in EM facility. Otherwise, dissolve 200 mg uranyl acetate in 20 mL of distilled water. Store in 10-mL Norm-ject syringes with a 0.22-μm filter at the tip of the syringe. Light sensitive, cover syringe with aluminum foil. Stable for several months at 4°C. 9. Lead citrate for staining of thin sections. Should be available in EM facility. Otherwise, prepare the following solutions only in boiled distilled water. Solution A: lead nitrate 1 M, i.e., 3.3 g in 10 mL water. Solution B: trisodic sodium citrate 1 M, i.e., 3.57 g in 10 mL water. Solution C: NaOH 1 N, i.e., 1 g in 25 mL water.

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de Chastellier To 16 mL of boiled distilled water, add 3 mL of solution B and mix gently with hand. Add 2 mL of solution A and mix gently with hand. A milky precipitate will form. Add, dropwise, 4 mL of solution C while mixing gently with hand. The precipitate must disappear completely. If not, discard and start over. Dispatch in 2- or 5-mL syringes with a 0.22-μm filter at the tip of the syringe. Stable for several months at 4°C. Discard if nonspecific fine precipitates are observed on the thin sections under the EM.

2.2. Acquisition of the Newly Endocytosed Content Marker, Horseradish Peroxidase (HRP), by Preexisting Phagosomes In addition to all the EM products listed in Subheading 2.1.: 1. Peroxidase from horseradish type II, RZ ∼2.0 (Sigma). Store at –20°C. 2. 3,3´-Diaminobenzidine tetrahydrochloride (DAB-HCl) from EMS. Store tablets at RT. Carcinogenic. 3. H2 O2 30% solution, reagent ACS (EMS). Store at 4°C. Once opened, stable for 2 mo at 4°C. 4. Tris-HCl 0.1 M, pH 7.6. Store at 4°C, stable for several months. 5. Prepare a 10 mg/mL solution of DAB in 0.1M Tris-HCl, pH 7.6. Filter the solution and store small aliquots at –20°C. Stable for about 6 mo. Once thawed, do not refreeze an aliquot.

2.3. Acquisition of the Content Marker Bovine Serum Albumin Coupled to Gold Particles (BSA-Au) Chased to Lysosomes Prior to Phagocytic Uptake In addition to all the EM products listed in Subheading 2.1.: 1. BSA coupled to gold particles (BSA-G10). Store at 4°C. Stable for several months (see Note 2 for preparation).

2.4. Enzyme Cytochemistry: Staining for Acid Phosphatase In addition to all the EM products listed in Subheading 2.1.: 1. 2. 3. 4. 5. 6. 7.

Ammonium chloride 1 M. Stable at 4°C for several months. Na-acetate, trihydrate, reagent ACS (EMS). Store at RT. Acetic acid 10% solution. Store at RT. Stable for several months. Glycerol 2-phosphate, disodium salt hydrate (Sigma). Store powder at RT. Ammonium sulfide 20% solution in water (Sigma). Store at RT. Na-fluoride (Sigma). Prepare a 10 mM solution in water. Store at 4°C. Water for gradient elution for high-performance liquid chromatography (HPLC) (Fluka).


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2.5. Enzyme Cytochemistry: Staining for Glucose 6-Phosphatase In addition to all the EM products listed in Subheading 2.1.: 1. 2. 3. 4. 5.

Water for gradient elution for HPLC (Fluka). d-Glucose-6-phosphate, disodium salt dehydrate (Sigma). Store at –20°C. Piperazine-N,N´-bis(2-ethanesulfonate) (PIPES) (Sigma). Store powder at RT. Maleic acid (EMS). Store powder at RT. Tris base, purissimo.

2.6. Assessment of Phagosome Acidity: DAMP Treatment and Immunolocalization In addition to all products from Subheading 2.1. (except for osmium tetroxide and Spurr resin): 1. DAMP (3-(2,4-dinitroanilino)-3´-amino-N-methyldipropylamine) from Oxford Biomedical Research (Oxford, MI). 2. Phosphate-buffered saline (PBS). 3. Bovine serum (do not heat inactivate). 4. Rabbit anti-dinitrophenol antibodies. 5. Protein A coupled to gold particles 10 nm in diameter (PAO-G10) (see Note 2 for preparation. 6. Resin: LRWhite. Purchase kit from EMS. Store kit at 4°C. Stable for several months.

3. Methods 3.1. Basic Conventional TEM (see Figs. 1 and 2) Cells must be fixed, dehydrated, and embedded in resin prior to sectioning and observation under the TEM. Important: Do not let cells dry up at any stage during the entire processing of samples. Illustrations of this method can be found in refs. 7, 11, and 12. 1. Fixative 1 (F1): Prepare a 2.5% solution of glutaraldehyde in 0.1M Na-cacodylate buffer, pH 7.2 containing 5 mM CaCl2 , 5 mM MgCl2 , and 0.1 M sucrose. Prepare fresh; can be kept on ice for half a day but warm up to RT before use. 2. Remove medium from cells. If necessary, wash cells with medium without serum. Do not use PBS for washes as this can introduce tiny calcium or magnesium phosphate precipitates during the fixation steps. 3. Add F1 immediately to culture dish. Fix for 1 h at RT under fume hood (see Note 3). 4. Remove fixative, wash twice (2 × 15 min) with above buffer at RT (see Note 4). 5. Fixative 2 (F2): Prepare a 1% solution of osmium tetroxide in sucrose-free 0.1 M Na-cacodylate buffer, pH 7.2, containing 5 mM CaCl2 and 5 mM MgCl2 . Sucrose is not necessary at this or following steps. Prepare fresh and use immediately.

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Fig. 1. Illustration of how one can use basic conventional microscopy (Subheading 3.1.) to determine morphological appearance of bacteria and identify the compartment in which they survive/replicate. (A, B, C): Bone marrow–derived macrophages were infected with Listeria monocytogenes (Lm), (D) Brucella abortus (Ba) or (E) Legionella pneumophila (Lp), fixed and processed for conventional TEM. These thin sections show whether bacteria are morphologically intact or degraded and give indications on the compartment in which they are enclosed. In (A) Lm is morphologically intact and therefore viable and is in a membrane-bound (arrow) phagosome; in (B) all the Lm are enclosed in membrane-bound (arrow) phagosomes and are degraded, as shown by their distorted shape; in (C) Lm is intact.


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6. Remove buffer, add F2. Fix for 1 h at RT under fume hood. 7. Wash cells three times rapidly to remove remaining osmium. Add 1-1.5mL of buffer in the culture dish (see Note 5). 8. Scrape cells off gently from culture dishes with a rubber policeman and put sample in a conical 1.5-mL Eppendorf tube (see Note 6). 9. Spin down samples in a microcentrifuge (10,000 rpm, 3 min). 10. Prepare a water bath at 42°C. Heat 2% agar. Once liquefied, put in water bath. Prepare a microvette containing about 0.3 mL agar. 11. Take samples in turn, remove supernatant, put Eppendorf tube in water bath, add about 0.3 mL agar to Eppendorf, mix cells and agar, and transfer to a microvette. Centrifuge immediately at 10,000 rpm for 2 min. Step 10 must be done rapidly in order to avoid solidification of agar before the cells have concentrated in the cap at the bottom of the microvette. Put microvette in ice, to harden the agar, for about 5 min. 12. Prepare a glass vial (2–3 mL content) with 1 mL of distilled water for each sample. Water will serve to wash samples briefly before putting them in Naveronal buffer. 13. Remove microvette from ice. With a razor blade, cut along one side of the bottom cap, then remove the cap gently. Gently push through the agar with a match over a glass slide. The sample, easily recognizable by its black color due to osmium fixation, will come out first, then the agar. Keep 3–4 mm of agar to protect the sample during further manipulations, and cut off the remainder. Put the sample in the glass vial. Do not leave samples in water for more than 5 min. 14. Remove water. Wash samples for 2 min with 1 mL of Na-veronal buffer (or Na-H-maleate-NaOH buffer). 15. Remove buffer, add 1 mL of 1% uranyl acetate prepared in Na-veronal buffer (or Na-H-maleate-NaOH buffer). Incubate cells for 1 h at RT. This step serves to improve fixation of membrane phospholipids. 16. Dehydrate samples at RT in a graded series of ethanol or acetone (ethanol is preferable when samples are embedded in Spurr resin), i.e., 25, 50, 75, and 90% followed by three successive baths in 100% dried on molecular sieve, 10 min per bath.

Fig. 1. The bacterium has completely lysed the phagosome membrane and resides in the cytoplasm where it is surrounded by a thick actin filament meshwork (arrow); in (D) the Ba-containing vacuole (BCV) has fused with the endoplasmic reticulum (arrows). The latter is easily identifiable by the presence of ribosomes (double arrows); in (E) the Lp-containing vacuole is decorated with ribosomes (double arrows), thereby indicating that the LCV membrane has acquired ER characteristics. Bar = 0.5 μm. Figures A, B, C) reproduced with kind permission from ASM Press; D) with permission from the Rockefeller University Press and E) with kind permission from Blackwell Publishing Ltd, as indicated in Note 17.

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Fig. 2. Illustration of how one can use basic conventional microscopy (Subheading 3.1.) to identify cell components involved in phagosome processing. Bone marrow– derived macrophages were infected with Mycobacterium avium. At day 7 postinfection, cells were treated for 0–6 h with 5 mM methyl--cyclodextrin (CD), which leads to a gradual depletion of cholesterol in cellular membranes, including phagosomes. (A) Thin section of a control cell, no CD. The phagosome membrane is smooth and closely apposed to the surface of M. avium all around (arrow). Such phagosomes remain immature and, therefore, do not fuse with surrounding lysosomes (Ly). (B) After a 30min treatment with CD, the phagosome membrane starts to become loose (arrows) and undulates around the bacterial surface. Notice the close proximity of the endoplasmic reticulum (ER). (C) Cells were loaded for 30 min with bovine serum albumin coupled to 10-nm-diameter gold particles (BSA-Au10) as described in Subheading 3.3., followed by a 2-h chase to label lysosomes and then treated with CD for 2 h. Phagosomes contain BSA-Au, thereby indicating that they have matured and fused with Ly. (D) After a 3-h


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17. Embedding in Spurr resin at RT: Prepare 25, 50 and 75% solutions of Spurr resin (1 mL per sample). The solutions must be prepared with the 100% dried solution of the dehydrating agent used for dehydration, i.e., either ethanol or acetone. Incubate the samples in the successive baths of resin for 30 min each. Then incubate in a first bath of pure resin for 30 min and a second bath for 2 h. 18. Put pure resin in molds. Add sample. (Do not forget to identify it! Write with a pencil on a small piece of paper, with number towards manipulator.) Add resin to fill mold. 19. Polymerize in a 60°C incubator for 24 h. 20. Section samples (70- to 75-nm-thick sections) at EM facility. 21. Staining of thin sections. Usual case: (a) Deposit grids on a drop of 1% uranyl acetate prepared in distilled water, incubate for 6 min, wash rapidly on five successive drops of distilled water, dry on filter paper; (b) then deposit grids on Reynold’s lead citrate for 2.5 min, wash rapidly on five successive drops of distilled water. Dry on filter paper.

3.2. Acquisition of HRP by Preexisting Phagosomes (see Fig. 3A, B, D, E, G) HRP has been widely used by endocytologists as a content marker because it is very sensitive to staining by cytochemical methods even in standard fixation conditions, and the reaction product forms a dense insoluble deposit that is easily visualized under the EM. The great advantage of this marker is that early endosomes can be distinguished from prelysosomes and lysosomes in terms of two parameters (7). First, they differ in their cytochemical staining pattern after HRP uptake. In early endosomes, the HRP reaction product only lines the inner face of the membrane (Fig. 3A; see also refs. 7 and 11). In prelysosomes and lysosomes, the entire lumen is filled with HRP reaction product (Fig. 3B; see also refs. 7 and 11). Second, early endosomes acquire newly internalized HRP immediately, whereas lysosomes display the marker only after a characteristic lag of 5–10 mins in macrophages and up to 15–30 min in other cells. The first parameter is used to observe whether phagosomes fuse Fig. 2. treatment with CD, M. avium–containing phagolysosomes (PhLy) become deformed. (E) After a 4- to 6-h treatment, cytoplasmic organelles, including other phagosomes, have been engulfed during the formation of an autophagolysosome (APhLy), which also contains gold particles accumulated in Ly prior to cholesterol depletion. In this autophagosome one can still see remnants of the inner membrane (arrow). Notice that bacteria are morphologically intact and surrounded by their electron translucent zone which is part of the cell wall. Bar = 0.5 μm. Reproduced with kind permission from Blackwell Publishing Ltd as indicated in Note 17.

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with early endosomes or with lysosomes (see Fig. 3D,E,G), and the second is used to classify phagosomes as resembling either early endosomes or lysosomes (see, e.g., refs. 7 and 11 for illustration of method; see also Note 7 for other possible markers). 1. Prepare a concentrated solution of HRP in culture medium (RPMI or DMEM). 2. Dilute HRP in complete medium. Prepare the solution fresh, warm up to 37°C, and use immediately (see Note 8 for appropriate concentration).

Fig. 3. (Continued)

EM Analysis of Phagosomes (e)

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(g)

Fig. 3. (Continued)

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3. At selected time points after exposure to HRP, between 0 and 60 min (or more), proceed to fixation with 2.5% glutaraldehyde, followed by washes with cacodylate buffer, as described in steps 1–4 in Subheading 3.1. 4. Wash cells 3 × 1 min with Na-cacodylate buffer devoid of sucrose and then 3 × 1 min with Tris-HCl 0.1M. Warm buffers to RT before use. 5. During the washes, prepare the cytochemical reaction medium: for 10 mL of medium, add 1 mL of DAB 10 mg/mL to 9 mL of 0.1 M Tris-HCl buffer, pH 7.6. Then add 100 μL of a 1% solution of H2 O2 (final concentration 0.01%). The 1% solution of H2 O2 must be prepared immediately prior to use. This medium is light sensitive, proceed swiftly to next step. 6. Remove last washing medium, add reaction medium to cells, and incubate for 15 min at RT in the dark. Caution: DAB is carcinogenic and must be handled with care. After incubation, inactivate DAB with H2 O2 before discarding. 7. Wash three times (3 × 1 min) with Tris-HCl buffer, then three times (3 × 1 min) with Na-cacodylate buffer devoid of sucrose. 8. Process samples for EM from Subheading 3.1., steps 5–21.

Fig. 3. Illustration of how one can use Subheadings 3.2. (A, B, D, E, G) and 3.3. (C, F) to analyze phagosome processing. (A, B) Morphological appearance of early endosomes (eEN) and lysosomes (Ly) stained with HRP. Bone marrow–derived mouse macrophages were given HRP (25 μg/mL) for 30 min, fixed, and stained for the endocytic content marker, HRP. In eEN (A), the reaction product only lines the inner face of the membrane, whereas in Ly (B), it entirely fills the lumen. (D, E, G) Fusion of phagosomes with eEN or Ly stained with HRP. Macrophages were given different types of latex beads. At 2 h after phagocytic uptake, cells were given HRP for 60 min as an endocytic content marker, then fixed and stained for HRP. The 1-μm-diameter hydrophobic bead–containing phagosomes (Ph(La)) fuse with eEN (arrows) (D) but not with Ly (G). The 1-μm-diameter hydrophilic bead–containing phagosomes (Ph(La)) have matured and fuse with Ly (arrow) (E). (C, F) Acquisition of lysosomal marker by phagosomes. Macrophages were incubated for 30 min with BSA tagged with 10-nmdiameter gold particles (BSA-Au10), washed and incubated for 2 h in medium devoid of BSA-AU10 to chase the marker to lysosomes (Ly). Cells were then infected with M. avium for 4 h. Phagosomes containing a single bacterium and for which the phagosome membrane is closely apposed to the bacterial surface do not fuse with Ly (C) , whereas those containing several bacteria and for which the phagosome membrane is, therefore, not closely apposed to the bacterial surface all around mature and fuse with Ly (F). Bar = 0.5 μm. A, B, E, G) reproduced with kind permission of ASM Press, C) with kind permission from Blackwell Science Ltd, D) with kind permission from Elsevier and F) with kind permission from Springer Science and Business media as indicated in Note 17.


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3.3. Acquisition of BSA-Au Chased to Lysosomes Prior to Phagocytic Uptake (see Figs. 2C and 3C,F) A widely used method to determine whether phagosomes have been processed into phagolysosomes or not involves chasing an endocytic content marker to lysosomes prior to phagocytosis and then analyzing acquisition of the marker, by phagosomes, at selected intervals during or after phagocytic uptake. The most frequently used marker, at present, is BSA-Au. This method is illustrated in Figs. 2C and 3 C,F to illustrate no fusion or fusion with lysosomes. (See also refs. 11–13 for illustration of method; see Note 9 for other possible markers.) 1. Sixteen hours prior to labeling, dialyze BSA-Au against serum-free culture medium (DMEM or RPMI) to remove azide. 2. Dilute BSA-Au in serum-free medium in order to have an OD of 3–5. Make fresh and use immediately. 3. Remove medium from cells, wash twice with serum-free medium and add BSAAu solution. Incubate cells for 30 min (60 min if uptake is low) at 37°C. 4. Wash cells three times rapidly with serum-free medium. 5. Remove last wash, add complete medium, and incubate for a further 2 h to chase marker to lysosomes (see Note 10). 6. Process samples as in Subheading 3.1., steps 1–21.

3.4. Enzyme Cytochemistry: Staining for Acid Phosphatase (see Fig. 4A) To determine whether phagosomes have been processed into phagolysosomes, it is possible to stain cells for the presence of lysosomal enzymes. Enzymes are not electron dense and are, therefore, not visible in electron microscopy unless enzyme cytochemistry methods are applied. Such methods do not visualize the enzyme itself but the product of the enzymatic reaction. The fact that the latter must be electron dense to be visualized has limited the number of enzymes that can be localized. Acid phosphatase (AcPase), aryl sulfatase, and trimetaphosphatase are enzymes of choice, because the phosphate liberated during the enzymatic reaction in the presence of substrate will react with the lead citrate used as a capture agent to form insoluble and electrondense lead phosphate precipitates. These precipitates remain in the organelles containing the enzyme provided that the ultrastructure is well preserved (see ref. 14 for illustration of method). 1. All products for the reaction medium must be prepared with boiled distilled water (boiled water for gradient elution for HPLC [Fluka] is even better) (see also Note 11). 2. Prepare Na-acetate buffer 0.05 M, pH 5.0. Stable at 4°C for 5–7 d.

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Fig. 4. Illustration of how one can use acid phosphatase (AcPase) cytochemistry (Subheading 3.4.) to characterize phagosomes and Subheading 3.6. to assess phagosome acidification. (A) Bone marrow–derived mouse macrophages were infected with M. avium. Seven days later, cells were fixed, stained for AcPase, and processed for EM. Lysosomes (Ly) were strongly labeled as shown by the dark deposits (arrow) in the entire lumen of the Ly, but most M. avium–containing phagosomes (Ph (Mav)) were not stained. Only a few of them displayed small deposits (arrow). (B, C) Macrophages were infected either for 4 h with M. avium as in (B) or for 45 min with Listeria monocytogenes (Lm) as in (C), washed and reincubated in fresh medium. One hour later, cells were exposed to DAMP (60 mM, 30 min), fixed and embedded in LRWhite. The probe was localized by the postembedding method described in Subheading 3.6. (B) Many gold particles were observed in Ly, but not in Ph(M av), thereby indicating that the latter were poorly acidic. (C) A high abundance of gold particles was observed in both Ly and Ph(Lm), thereby indicating that the latter are acidic. Bar = 0.5 μm. A, C) Reprinted with kind permission from ASM Press, (B) with kind permission from Elsevier as indicated in Note 17. 3. Prepare a solution of lead nitrate 6% (60 mg in 1 mL of freshly boiled water) immediately before preparing the reaction medium. Toxic. 4. Cytochemical reaction medium: add 30 mg of glycerol 2-phosphate, disodium salt hydrate to 11 mL of 0.05 M Na-acetate buffer. Mix, adjust pH to 5.0


5.

6.

7. 8.

9.

10. 11. 12.

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with 10% acetic acid (one or two drops usually sufficient). Then add dropwise, under vigourous stirring, 200 μL of the lead nitrate solution. If mixture becomes cloudy, discard and start over. Incubate reaction medium at 37°C for at least 2 h before adding to cells (overnight is often better). If the medium is still slightly cloudy, one can try to filter it on a 0.22-μm filter. Prepare a 1.25% solution of glutaraldehyde in 0.1 M Na-cacodylate buffer, pH 7.2, containing 5 mM CaCl2 , 5 mM MgCl2 , and 0.1 M sucrose. Prepare fresh, keep on ice, stable for half a day. At time points of interest, remove medium from cells. If necessary, wash cells with serum-free medium. Do not use PBS for washes as this can introduce tiny calcium or magnesium phosphate precipitates during the fixation and incubation steps. Add cold fixative immediately to culture dish. Fix for 1 h at 4°C (see Note 12). Remove fixative, wash twice (2 × 15 min) at RT with above buffer containing 50 mM NH4 Cl, twice (2 × 15 min) at RT with same buffer devoid of NH4 Cl, and twice (2 × 5 min) with Na-acetate buffer prewarmed to 37°C. Remove buffer, add prewarmed reaction medium and incubate cells at 37°C for 30 min. Controls: (a) incubate cells in substrate-free reaction medium or (b) in complete reaction medium containing 10 mM sodium fluoride. Wash cells twice (2 × 2 min) with Na-acetate buffer and twice (2 × 2 min) with sucrose-free Na-cacodylate buffer. Remove last wash and process cells for conventional electron microscopy as indicated in Subheading 3.1., steps 5–21 (see Note 13). Interpretation: see Note 14.

3.5. Enzyme Cytochemistry: Staining for G6Pase (see Fig. 5) G6Pase is a specific marker of the endoplasmic reticulum (ER) compartment. In addition to basic conventional EM, staining for G6Pase can, therefore, be used to determine whether the ER interacts with and transfers its contents to phagosomes, and whether phagosomes acquire ER characteristics (see refs. 15–17). The method described here was adapted from Griffiths et al. (18). 1. All buffers as well as the reaction medium must be prepared with boiled bidistilled water Water for gradient elution for HPLC (Fluka) is even better. Boil water immediately before preparation of products (see Note 11). 2. Prepare PIPES 0.1 M, pH 7.0: Dissolve 3.02 g of PIPES in 75 mL of boiled water. Solution is milky (do not worry!). Adjust the pH to 7.0 with NaOH 1 N (about 15 mL). The solution clears up. Complete to 100 mL with boiled water. Split into two 50-mL aliquots, add either 2.5 or 5 g of sucrose to obtain PIPES with 5 or 10% sucrose, respectively. Store at 4°C. Use within a week (but better to prepare the day before). 3. Prepare Tris-maleate 0.08 M, pH 6.5: Add 0.485 g Tris base and 0.465 g maleic acid to 40 mL of boiled water, Adjust pH to 6.5 with NaOH 1 N (about 1–2 mL)

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Fig. 5. Illustration of glucose-6-phosphastase (G6Pase) cytochemistry (Subheading 3.5.). Bone marrow–derived mouse macrophages were infected with Brucella abortus wild-type strain 2308 for various times or given 1-μm-diameter hydrophilic latex beads for 30 min. Samples were processed for detection of G6Pase according to Subheading 3.5. Dense deposits in the endoplasmic reticulum (ER) correspond to the reaction product. (A) Representative G6Pase-negative Brucella-containing vacuole (BCV) at 4 h postinfection. (C) Representative G6Pase-negative latex bead-containing phagosome (Ph(La)) processed at 30 min after phagocytic uptake. Notice in both cases that phagosomes are negative, although they are in intimate contact with several G6Pase-positive ER compartments. (B, D) Representative G6Pase-positive BCVs at 24 h postinfection as shown by the presence of dense deposits (arrow) within the BCV. (D) shows a BCV fusing with the ER (double arrows). Bar = 0.5μm. (A, B, D) Reproduced with kind permission from The Rockefeller University Press, as indicated in Note 17.


4. 5.

6.

7. 8.

9.

10.

11. 12. 13.

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and complete to 50 mL with water. Store at 4°C. Use within a week (but better to prepare the day before). Prepare a 6% solution of lead nitrate: Dissolve 60 mg lead nitrate in 1 mL of freshly boiled water. Prepare fresh and use immediately in reaction medium. Prepare cytochemical reaction medium: Dissolve 95 mg of glucose-6-phosphate, disodium salt in 10 mL of Tris-maleate 0.08 M, pH 6.5, adjust pH to 6.5 with a few drops of HCl 0.1 N. Add, dropwise, 160 μL of the 6% lead nitrate solution under vigourous stirring. The incubation medium is often slightly milky. It is, therefore, preferable to filter it through a 0.22-μm filter. Stable at RT for 2 h. Prepare fixatives: (a) F1: 1.25% glutaraldehyde in 0.1 M PIPES, pH 7.0, containing 5% sucrose; (b) F2: 1.25% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, containing 0.1 M sucrose, 5 mM CaCl2 , and 5 mM MgCl2 . Keep on ice. Remove medium, add F1, and fix cells for 30 min on ice. Remove F1 and wash cells 3 × 3 min at RT with 0.1 M PIPES, pH 7.0, containing 10% sucrose, followed by 1 × 30 s at RT with 0.08 M Tris-maleate buffer, pH 6.5. Remove last wash, add cytochemical medium, incubate cells for 2 h at 37°C in a CO2 -free incubator. Possible controls: incubate cells (a) in substrate-free reaction medium or (b) in complete reaction medium containing 10 mM sodium fluoride. Remove cytochemical medium, wash cells 3 × 2 min at RT with 0.08 M Trismaleate buffer and 3 × 2 min at RT with 0.1 M cacodylate buffer, pH 7.2, containing 0.1 M sucrose, 5 mM CaCl2 , and 5 mM MgCl2 . Remove last wash, add F2, and fix cells for 1 h at 4°C. Remove F2 and process cells for conventional electron microscopy as indicated in Subheading 3.1., steps 4–21. Interpretation: see Note 15.

3.6. Assessment of Phagosome Acidity: DAMP Treatment and Immunolocalization (see Fig. 4B,C) DAMP (3-(2,4-dinitroanilino)-3‘amino-N-methyldipropylamine) is a probe that has been widely used to study phagosome acidification at the TEM level (19). This weak base accumulates by diffusion within acidic compartments. Once protonated, it can no longer diffuse out. During chemical fixation with aldehydes, it becomes covalently linked to proteins, which allows it to be retained in acidic organelles during processing of samples for EM. DAMP is then localized with an appropriate postembedding immunolabeling method on thin sections of cells embedded in LRWhite resin as described below (see Note 16). 1. Prepare fixatives: F1: 1.25% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, containing 0.1 M sucrose, 5 mM CaCl2 , and 5 mM MgCl2 ; F2: paraformaldehyde 1% in same buffer. Keep on ice.

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2. At selected intervals postinfection, remove cell culture medium and incubate cells for 30 min at 37°C with 100 μM DAMP diluted in prewarmed complete medium. 3. Wash cells three times (rapidly) with medium (DMEM or RPMI) containing 10% serum (same serum as for cell culturing). 4. Remove last wash, add F1 (warmed to RT), and fix cells for 1 h at RT. 5. Remove F1, add F2, and store sample at 4°C. Cells can be kept at 4°C in F2 for 1 wk. 6. Wash fixed cells twice (2 × 15 min) at RT with complete Na-cacodylate buffer containing 50 mM NH4 Cl and three times (3 × 5 min) at RT with the same buffer devoid of NH4 Cl. 7. Process samples as indicated in Subheading 3.1., steps 8–14. 8. Put samples on ice. Remove buffer, add 1 mL of 0.5% uranyl acetate prepared in Na-veronal buffer (or Na-H-maleate-NaOH buffer). Incubate for 1 h on ice. 9. Dehydrate samples at RT in 50% ethanol (2 × 15 min) followed by 70% ethanol (3 × 10 min). 10. Embed in LRWhite: three successive baths of 20 min in pure resin. 11. Put pure resin in gelatin capsules. Add sample (do not forget to identify it! Write with a pencil on a small piece of transparent paper, with number towards manipulator). Add resin to fill capsule. Put cap on the capsule to avoid contact with air. 12. Polymerize in a 37°C incubator for 3 d. 13. Sectioning (in EM facility): Pick up sections on formvar and carbon-coated nickel grids and put grids on PBS containing 0.5% bovine serum. 14. For immunolabelling, prepare PBS containing 0.5, 5, and 10% fetal bovine serum (FBS). Do not heat inactivate the serum prior to use. 15. Incubate the grids (face with thin sections against the drops of reagents) on the following antibodies or reagents: preincubate sections for 10 min on PBS containing 10% FBS to block nonspecific sites, and then sequentially incubate for 60 min at RT with rabbit antidinitrophenol (anti-DNP) antibodies and for 30 min with protein A coupled to 10-nm-diameter gold particles (PA-Au10). Antibodies and conjugate are diluted in PBS containing 5% FBS. Sections are washed five times rapidly with PBS containing 0.5% FBS between the incubations and after treatment with the conjugate. After three washes with PBS and distilled water, dry the grids of filter paper. As a control, incubate grids on non specific antibody followed by PA-Au10 or on PA-Au10 only (see Note 16 for estimating intraphagosomal pH). 16. Staining of thin sections: Deposit grids on a drop of 1% uranyl acetate prepared in distilled water, incubate for 1 min, wash rapidly on five successive drops of distilled water, dry on filter paper; then deposit grids on Reynold’s lead citrate for 30 s, wash rapidly on five successive drops of distilled water. Dry on filter paper.


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4. Notes 1. I suggest Spurr resin because of its low viscosity, which allows for embedding of samples containing difficult to embed bacteria, such as mycobacteria. It is easy to prepare and it polymerizes within 24 h at 60°C. Other resins, such as Epon, are widely used and available as kits from EMS. 2. Preparation of BSA-G10 or PAO-G10. The method has been described at length by J.W. Slot and H. J. Geuze (20), who introduced the immunogold labeling procedure. 3. Cells can also be fixed overnight but at 4°C instead of RT. Proceed to step 4 the next morning. 4. Cells can also be washed overnight at 4°C instead of 2 × 15 min at RT. Proceed to step 5 the next morning. 5. At this stage, one can add buffer to cells and store at 4°C for up to 2 wk. This is practical if one is doing a kinetic study and wishes to process all EM samples at the same time. 6. Cell scraping: This is a critical step. It is important to scrape cells only after they have been fully fixed with both glutaraldehyde and osmium. Otherwise, cells will be damaged during the scraping process. 7. Endocytic content markers added after phagocytic uptake: Other markers can be used, provided that they can be tagged with electron dense probes. One possibility is to use biotinylated ligands (dextran or albumin) as content markers and then exploit the biotin–streptavidin interaction, with streptavidin coupled to HRP or to gold particles, to localize the ligands intracellularly. One can also tag ligands with gold particles (see also Subheading 3.3.). However, with all these ligands, it is not possible to morphologically distinguish early endosomes from late compartments, as with HRP. 8. Certain primary macrophage cultures, such as mouse bone marrow–derived, are rich in mannose receptors at their surface. In this case, prepare a 1 mg/mL solution and use it at 25 μg/mL (40-fold dilution). HRP will then be essentially internalized via the mannose receptor (21). If cells have no or very few mannose receptors, one must add more HRP, which will be endocytosed by fluid phase endocytosis. Usually one adds 1 mg/mL—up to 6 mg /mL in certain cell types. It is advisable to try different concentrations of marker and different incubation times in the presence of marker to define the optimal conditions and to use control particles for which it is known whether they fuse with early endosomes only or early endosomes and then lysosomes (7,11). 9. Other markers: Ferritin, thorotrast, or ligands tagged with electron-dense probes as indicated in Note 7. 10. Incubation and chase conditions with BSA-Au: As indicated in Note 8, it is advisable to define the optimal working conditions. To obtain the most reliable data, the marker must be internalized by the cell in sufficient amounts so that it will label the entire lysosomal compartment when chased after uptake. It might, therefore, be necessary to expose cells to the marker for 60–120 instead

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13.

14.

15.

de Chastellier of 30 min. However, the marker must not be too densely packed within the lysosome, or it might form a sort of rigid gel or network. In such conditions contents would not be transferred to phagosomes upon fusion with lysosomes, and this would be misinterpreted as a “no-fusion” event. Finally, it is important to keep the chase period short because lysosomal contents are recycled out of the lysosomal compartment via small recycling vesicles, and this will eventually lead to excretion of the marker. In macrophages, about 50% of the label is secreted within a 20-h chase period. For enzyme cytochemistry, use only disposable plastic, no reusable glassware or pipets. Enzyme susceptibility to fixatives: The cells must be fixed before the enzymatic reaction occurs. A compromise must, therefore, be reached between preservation of ultrastructure and of enzymatic activity. This is why I use a lower concentration of glutaraldehyde and fix cells in the cold for detection of AcPase (and other phosphatases). Most enzymes are inactivated by chemical fixatives, and below a threshold concentration they will no longer be detected. When studying the acquisition of other lysosomal enzymes by phagosomes, it is advisable to try different fixation conditions and include controls such as phagocytosis of particles that do not prevent phagosome maturation and fusion with lysosomes, and, therefore, acquisition of hydrolases by phagosomes (see, e.g., refs. 13 and 14). One can check under the light microscope whether one was successful with this method. Grow cells on cover slips; proceed through steps 1–9. Wash cells once with distilled water, incubate the cover slips for 1 min over a drop of 1% ammonium sulfide (under fume hood), wash cover slips in two successive drops of distilled water. Mount on glass slide. Under the light microscope, the reaction product appears as dark brown deposits in lysosomes (and eventually phagosomes). Material treated with ammonium sulfide cannot be processed for EM. Interpretation: First, one must keep in mind that hydrolases such as AcPase, which are concentrated in lysosomes, are also present in small amounts in early compartments and can, therefore, be acquired by phagosomes via multiple fusion events with early endosomes. The amount of enzyme transferred to phagosomes in such conditions, is, however, usually below the threshold level of detection. Second, the hydrolysis of the substrate can also be catalyzed by other phosphatases, including glucose-6-phosphatase, an ER- and nuclear membrane-specific marker, or alkaline phosphatase, which is enriched in the plasma membrane. Their contribution should, however, be minor if the reaction medium is of good quality. Small or large nonspecific deposits can also occur if the reaction medium is not of good quality. It is advisable to consult a specialist to avoid misleading and wrong conclusions! The hydrolysis of the substrate can also be catalyzed by other phosphatases, including acid phosphatase, which is enriched in lysosomes and phagolysosomes, or alkaline phosphatase, which is enriched in the plasma membrane. If the


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cytochemical reaction is done properly, their contribution should, however, be minor. Small or large nonspecific deposits can also occur if the reaction medium is not of good quality. It is advisable to consult a specialist to avoid misleading and wrong conclusions! 16. An important advantage of this method is that the number of gold particles per phagosome can be converted to values for intraphagosomal acidification by the method described by Orci et al. (22) provided that the phagosome contains sufficient amounts of protein for all the protonated DAMP molecules to be retained in the phagosome after fixation. (For examples of assessment of intraphagosomal pH, see refs. 7, 12, and 13.) 17. Permissions to reprint figures: Fig 3F: Reprinted, with kind permission of Springer Science and Business Media, from the book “Intracellular pathogens in membrane interactions and vacuole biogenesis”, JP Gorvel (ed), 2004, chapter title: “Phagosome biogenesis in relation to intracellular survival mechanisms of mycobacteria”, pp 153–165, authors: Lutz Thilo and Chantal de Chastellier, Fig 2A, copyright 2004 Kluwer Academic/Plenum Publishers, New York, NY, USA. Figs 3D and 4B: Reprinted, with kind permission, from the European Journal of Cell Biology, 2003, 68 (Oct): 167–182, authors: Chantal de Chastellier, Thierry Lang and Lutz Thilo, Title of article: Phagocytic processing of the macrophage endoparasite, Mycobacterium avium, in comparison to phagosomes which contain Bacillus subtilis or latex beads, Figs 3A and 10C, copyright 1995 Wissenschaftliche Verlagsgesellschaft, Stuttgart, Germany. Fig 1D and 5A, B, D: Reproduced, with kind permission, from the Journal of Experimental Medicine, 2003, 198: 545–556, copyright 2003 The Rockefeller University Press, New York, NY, USA. Figs 1A, B, C, 3A, B, E, G and 4A, C: Reprinted, with kind permission, from “Cellular Microbiology, 2nd edition”, Au: Chantal de Chastellier, editors: P. Cossart, P. Boquet, S. Normark and R. Rappuoli, copyright 2004 ASM Press, Washington, DC, USA. Fig 1E: Reprinted, with kind permission, from Cellular Microbiology, 2006, online early, doi: 10.1111/j. 1462-5822.00839.x, Au: Anne Fortier, Chantal de Chastellier, Stéphanie Balor and Philippe Gros, title of article: “Birc1e/Naip5 rapidly antagonizes modulation of phagosome maturation by Legionella pneumophila”, Fig 6A, copyright 2006 Blackwell Science Ltd, Oxford, United Kingdom. Figs 2A, B, C, D, E: Reprinted, with kind permission, from Cellular Microbiology, 2006, 8(2): 242–256, Au: Chantal de Chastellier and Lutz Thilo, title of article: “Cholesterol depletion of Mycobacterium avium-infected macrophages overcomes the block in phagosome maturation and leads to the reversible sequestration of mycobacteria in phagolysosome-derived autophagic vacuoles”, Figs 2A, 2D, 6A, 6C, 8A copyright 2005 Blackwell Science Ltd, Oxford, United Kingdom.

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de Chastellier Fig 3C: Reprinted, with kind permission, from Cellular Microbiology, 2002, 4(8): 541–556, Au: Claude Fréhel, François Cannone-Hergaux, Philippe Gros and Chantal de Chastellier, title of article: “Effect of Nramp1 on bacterial replication and on maturation of Mycobacterium avium-containing phagosomes in bone marrow-derived mouse macrophages”, Fig 7B, copyright 2002 Blackwell Science Ltd, Oxford, United Kingdom.

References 1. Haas, A. (1998) Reprogramming the phagocytic pathway—intracellular pathogens and their vacuoles. Mol. Membr. Biol. 15, 103–121. 2. Knodler, L. A., Celli, J. and Finlay, B. B. (2001) Pathogenic trickery: deception of host cell processes. Nat. Rev. Mol. Cell Biol. 2, 578–588. 3. Russell, D. G. (2001) Mycobacterium tuberculosis: here today, and here tomorrow. Nat. Rev. Mol. Cell Biol. 2, 569–577. 4. Thilo, L. and de Chastellier, C. (2004) Phagosome biogenesis in relation to intracellular survival mechanisms of mycobacteria, in Intracellular Pathogens in Membrane Interactions and Vacuole Biogenesis (Gorvel, J. P., ed.), Kluwer Academic/Plenum Publishers, New York, pp 153–169. 5. Gutierrez, M. G., Master, S. S., Singh, S. B., Taylor, G. A., Colombo, M. I. and Deretic V. (2004) Autophagy is a defense mechanism inhibiting BCG and Mycobacterium tuberculosis survival in infected macrophages. Cell 119, 753–766. 6. Gorvel, J. P. and de Chastellier, C. (2005) Bacteria spurned by self-absorbed cells. Nat. Medicine 11, 18–19. 7. de Chastellier, C., Lang, T. and Thilo, L. (1995) Phagocytic processing of the macrophage endoparasite Mycobacterium avium, in comparison to phagosomes which contain Bacillus subtilis or latex beads. Eur. J. Cell Biol. 68, 167–182. 8. Raposo, G., Kleijmeer, M. J., Posthuma, G., Slot, J. W. and Geuze, H. G. (1997) Immunogold labeling of ultrathin cryosections: application in immunology, in Weir’s Handbook of Experimental Immunology (Herzenberg, L. A. and. Weir, D. M., eds.), Blackwell Science Inc., Oxford, United Kingdom, vol. 4, pp. 208.1–208.11 9. de Chastellier, C. (2004) Electron microscopy, in Cellular Microbiology, 2nd ed. (Cossart, P., Boquet, P., Normark, S. and Rappuoli, R., eds.), ASM Press, Washington, DC, pp 451–471. 10. Koster, A. J. and Klumpermann, J. (2003) Electron microscopy in cell biology: integrating structure and function. Nat. Rev. Mol. Cell Biol. 4, SS6–SS10. 11. de Chastellier, C. and Thilo, L. (1997) Phagosome maturation and fusion with lysosomes in relation to surface property and size of the phagocytic particle. Eur. J. Cell Biol. 74, 49–62. 12. de Chastellier, C. and Thilo, L. (2006) Cholesterol depletion of Mycobacterium avium-infected macrophages overcomes the block in phagosome maturation and leads to the reversible sequestration of mycobacteria in phagolysosome-derived autophagic vacuoles. Cell. Microbiol. 8, 242–256.


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13. Fréhel, C., Canonne-Hergaux, F., Gros, P. and de Chastellier, C. (2002) Effect of Nramp1 on bacterial replication and on maturation of Mycobacterium aviumcontaining phagosomes in bone marrow-derived mouse macrophages. Cell. Microbiol. 4, 541–556. 14. de Chastellier, C., Fréhel, C., Offredo, C. and Skamene, E. (1993) Implication of phagosome-lysosome fusion in restriction of Mycobacterium avium growth in bone marrow macrophages from genetically resistant mice. Infect. Immun. 61, 3775–3784. 15. Celli, J., de Chastellier, C., Franchini, D.-M., Pizarro-Cerda, J., Moreno, E. and Gorvel, J. P. (2003) Brucella evasion of macrophage killing through VirBdependent sustained interactions with the endoplasmic reticulum. J. Exp. Med. 198, 545–556. 16. Touret, N., Paroutis, P., Terebiznik, M., et al. (2005) Quantitative and dynamic assessment of the contribution of the endoplasmic reticulum to phagosome formation. Cell 123, 157–170. 17. Fortier, A., de Chastellier, C., Balor, S. and Gros, P. (2007) Birc1e/Naip5 rapidly antagonizes modulation of phagosome maturation by Legionella pneumophila. Cell Microbiol. 9, 910–923. 18. Griffiths, G., Quinn, P. and Warren, G. (1983) Dissection of the Golgi complex. I Monensin inhibits the transport of viral membrane proteins from medial to trans Golgi cisternae in baby hamster kidney cells infected with Semliki forest virus. J. Cell Biol. 96, 835–850. 19. Anderson, R. G. W., Falck, J. R., Goldstein, J. L. and Brown, M. S. (1984) Visualization of acidic organelles in intact cells by electron microscopy. Proc. Natl. Acad. Sci. USA 81, 4838–4842. 20. Slot, J. W. and Geuze, H. J. (1985) A new method of preparing gold probes for multiple-labelling cytochemistry. Eur. J. Cell Biol. 38, 87–93. 21. Lang, T. and de Chastellier, C. (1985) Fluid and mannose receptor-mediated uptake of horseradish peroxidase in mouse bone marrow-derived macrophages. Biochemical and ultrastructural study. Biol. Cell 53, 149–154. 22. Orci, L., Halban, P., Perrelet, A., Amherdt, M., Ravazzola, M. and Anderson, R. G. W. (1994) PH-independent and-dependent cleavage of proinsulin in the same secretory vesicle. J. Cell Biol. 126, 1149–1156.

18 Analysis of Phosphoinositide Dynamics During Phagocytosis Using Genetically Encoded Fluorescent Biosensors Gabriela Cosío and Sergio Grinstein

Summary Phosphoinositide signaling is essential for successful phagocytosis. Phosphoinositides regulate processes such as actin assembly and the recruitment of molecular motors required for ingestion, as well as fusion events required for the maturation of the phagosome. Phosphoinositides not only serve as substrates for the generation of second messengers, but also function to anchor to the membrane cytosolic proteins that contain phosphoinositidebinding motifs. Conventional methods for the detection of phosphoinositides involve their extraction from the cells and separation by chromatographic procedures. These approaches are laborious and expensive and fail to provide spatio-temporal information, which is critical when analyzing localized and transient phenomena like phagocytosis. In this chapter we describe a method to monitor phosphoinositides dynamically by transfection of fluorescently tagged probes (biosensors) into cultured macrophages. These biosensors are based on the fusion of phosphoinositide-binding protein domains with fluorescent proteins. Some specifications for live cell imaging of such phosphoinositide-specific probes are also provided.

Key Words: Phosphoinositide; domain; confocal microscopy.

phagocytosis;

phagosome;

macrophage;

PH

1. Introduction Phagocytosis is defined as the engulfment of large particles, greater than 0.5 μm in diameter, by specialized cells such as neutrophils and macrophages. Phagocytosis is essential for the destruction of pathogens, the clearance of apoptotic cells, and tissue remodeling (1). The phagocytic process comprises From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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a variety of events that are initiated by the engagement of surface receptors and involve multiple signaling pathways. The number and nature of the signals elicited vary depending on the type of receptor engaged. A wide variety of phagocytic receptors have been described, among which the FcR and the CR3 receptors are the most thoroughly characterized. Receptor engagement and the ensuing signals ultimately lead to internalization of the particle into a membrane-bound vacuole or phagosome. The signaling cascade triggered by the activated receptors usually involves activation of protein kinases, alterations in phospholipid metabolism, remodeling of the actin cytoskeleton, and localized acceleration of membrane traffic (2). Upon phagocytosis, the invading organism or alien particle is confined in the phagosome, which then undergoes a process of maturation. During the course of maturation the phagosome interacts with components of the endocytic pathway by a series of fusion and fission events that ultimately yield a hybrid organelle, the phagolysosome (3). Throughout this remodeling process the phagosome acquires degradative properties such as an acidic luminal pH, the delivery and activation of hydrolytic enzymes, and the production of reactive oxygen species that are essential to its microbicidal function (3). Recent studies indicate that phosphoinositides play various, important roles in phagosome formation and maturation. Phosphoinositides are derivatives of the lipid phosphatidylinositol and are thought to be found in virtually all cellular membranes (4). Polyphosphoinositides can be reversibly phosphorylated on positions 3, 4, and/or 5 of the inositol head group, resulting in the generation of seven different species (5–8). In addition to their role as substrates for the phospholipases that generate second messengers, phosphoinositides also act themselves as membrane anchors for cytosolic proteins that possess a cognate phosphoinositide-binding domain (9–11). As a result, changes in phosphoinositide composition provide signals for the selective recruitment and activation of signaling proteins to specific membranes in the cell (4). Phosphoinositides have been shown to be involved in signaling the early stages of phagocytosis, leading to actin assembly and the recruitment of molecular motors to the site of ingestion (1). They also regulate multiple budding, fission and fusion events required for maturation (1). Phosphatidylinositol-4,5-bisphosphate (PI(4,5)P2 ), which is constitutively present in the plasma membrane, becomes transiently and modestly accumulated in pseudopods of forming phagosomes, but disappears rapidly thereafter (1). At least two reactions contribute to the disappearance of PI(4,5)P2 : its degradation by PLC, with the concomitant production of diacylglycerol (DAG) and inositol trisphosphate (IP3) (11), and its conversion to phosphatidylinositol3,4,5-trisphosphate (PI(3,4,5)P3 ) by class I phosphatidylinositol 3-kinase (PI3K) (2). PI(4,5)P2 itself can promote actin polymerization, and its disappearance is

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therefore important for the termination of actin assembly (12). On the other hand, PI(3,4,5)P3 undergoes a significant accumulation at sites of phagosome formation and remains briefly associated with nascent phagosomes (2,13). Newly formed PI(3,4,5)P3 in turn stimulates the hydrolysis of PI(4,5)P2 by promoting the recruitment and activation of PLC (14). PI(3,4,5)P3 also contributes directly to the mechanical events that lead to particle intake by recruiting WAVE and myosin X to the phagosome (15). In addition, 3phosphoinositides seem to be involved in the fusion of endomembranes at sites of phagocytosis, which promotes the conservation of the cell surface area (16). 3-Phosphoinositides, particularly phosphatidylinositol 3-phosphate (PI(3)P), appear to be crucial for phagosome maturation. PI(3)P accumulates on the phagosomal membrane within 1–2 min of sealing and persists for about 10 min (17). PI(3)P seems to be important for the recruitment of ligands such as EEA1 and Hrs, which are important for the fusion on membranes that bear Rab5 and Rab4 and for the inward membrane budding that generates multivesicular bodies (18). To date, the majority of approaches to detect and measure the cellular content of phosphoinositides have relied on the use of radioisotope labeling, in which cellular lipids are metabolically tagged, followed by separation using laborious chromatographic procedures such as high-performance liquid chromatography (HPLC) and/or thin-layer chromatography (TLC) (19,20). Recently, mass spectrometry has also been utilized to measure phosphoinositides, but while this approach bypasses the need for isotopic labeling, its sensitivity is insufficient to detect several minor, yet physiologically very important inositide species (21). These methods are all extremely time-consuming and require the use of large numbers of cells and expensive reagents. Most importantly, because they are all end-point determinations, these approaches fail to reveal the spatiotemporal dynamics of phosphoinositides in the cell. These limitations were recently overcome by the introduction of fluorescently tagged phosphoinositidespecific probes that can be monitored continuously by noninvasive means. In essence, these novel probes consist of selected protein domains (modules) with well-defined phosphoinositide-binding properties, which are tagged with a fluorescent moiety. For convenience, fluorescent properties are conferred to these domains by fusion with a humanized version of the jellyfish green fluorescent protein (GFP), or any of its multiple variants (e.g., CFP, YFP), or with one of the red fluorescent proteins from other invertebrate species (Fig. 1). In this manner, the distribution of the fluorescently labeled phosphoinositidebinding domains can be monitored by light microscopy in living cells. One of the major advantages of this approach is that the probe is genetically encoded and can be delivered into the cells by transfection, viral infection, or microinjection of the corresponding cDNA.

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Fig. 1. Biosensor strategy used to detect phosphoinositides. (A) A chimeric construct is generated by fusing a well-defined phosphoinositide-binding motif, such as the PH domain illustrated here, and a fluorescent protein like GFP. Expression of the cDNA encoding for the chimera will generate a specific fluorescent probe detectable in live cells by noninvasive means. (B) Distribution of PI(3)P accumulation during phagosome maturation. PI(3)P was monitored by using the PX domain from p40phox (35) coupled to GFP. The present picture was taken 2 min after phagosome sealing. Notice that endosomes are also labeled by this chimeric construct due to their PI(3)P content.

The vast repertoire of known phosphoinositide-interacting proteins has led to the identification and characterization of a variety of phosphoinositidebinding modules with distinct specificities and affinities (Table 1). The best characterized module is the PH (pleckstrin-homology) domain (22), which is present in proteins such as Akt/PKB, PDK1 and GRP1. More recently, a variety of other phosphoinositide-binding domains have been identified and characterized, including PX (Phox) (23), ENTH (epsin N-terminal homology) (24), FYVE (Fab1, YOTB, Vac1p, and EEA1) (25), FERM (Ezrin, Radixin, and Moesin) (26), and GRAM (glucosyltransferases, Rab-like GTPase activators, and myotubularins) (27) domains that are present in a variety of proteins. The suitability of phosphoinositide-binding probes for live cell imaging has been the subject of recent debate. It is important to consider that, even though in vitro binding assays may indicate that a particular lipid-binding domain binds preferentially to one of several purified lipids, the probe may not show the same selectivity inside cells. Often cellular domains are coincidence detectors that sense two or more binding determinants simultaneously. While only lipids are present in the in vitro assays, other factors, such as proteins binding to other moieties of the probe are likely to exist in vivo, contributing to their recruitment (52). In this regard, it is desirable to validate the observations made with any one probe by other techniques or utilizing different lipid-binding


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Table 1 Phosphoinositide-Binding Domains Phosphoinositide PtdIns4P

PtdIns3P PtdIns(4,5)P2 PtdIns(3,4)P2 PtdIns(3,5)P2 PtdIns(3,4)P2 / PtdIns(3,4,5)P3 PtdIns(3,4,5)P3

Protein domain FAPP1-PH OSH2-PH OSBP-PH EEA1-FYVE, Hrs-FYVE p40phox-PX PLC1-PH Tubby TAPP1-PH Svp1p Centaurin2-PH AKT-PH PDK1-PH CRAC-PH GRP1-PH ARNO-PH Btk-PH Cytohesin-1-PH

Ref. (28–30) (31,32) (28,29,33) (34) (35) (36,37) (38) (39) (40) (41) (42,43) (44) (45) (46,47) (48) (49) (50,51)

Source: Adapted from ref. (53).

domains that may have distinct binding determinants yet similar lipid selectivity in vitro. Furthermore, it is important to exercise care in the interpretation of results when using probes that can bind to more than one phosphoinositide species. In this case, as before, it is particularly helpful to confirm the findings using alternate probes. Finally, the level of expression of a given probe can be critical. Excessive expression of certain probes may result in interference with the cellular process they are supposed to monitor. This complication can be minimized by analyzing exclusively those cells with the lowest expression level compatible with adequate imaging and by comparing the phenotype of cells with varying levels of expression. Introduction of the appropriate cDNA into phagocytes is essential for the expression of the biosensors and the subsequent imaging of phosphoinositide dynamics during phagosome formation and maturation. Primary professional phagocytes like macrophages and neutrophils are notoriously refractory to transfection or viral infection, so that heroic efforts are often required to introduce the plasmids. On the other hand, some cultured phagocytic cells, such as the mouse monocytic RAW 264.7 cell line, are transfectable at reasonable rates by either lipofection or electroporation. The use of RAW 264.7 cells for the dynamic study of phosphoinositide metabolism during phagocytosis is described below.

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2. Materials 2.1. Cell Culture 1. Dulbecco’s modified Eagle’s medium 1X (DMEM) (cat. no. 319-005-CL. WISENT, Inc., Mississauga, ON) supplemented with 5% fetal bovine serum (FBS PREMIUM, cat. no. 080450 WISENT, Inc.). 2. Hydroxyethyl piperazine ethane sulfonate (HEPES)-buffered solution RPMI 1640 1X (HPMI) (Cat. No. 350-025-CL WISENT, Inc.). 3. Phosphate-buffered saline (PBS) (cat. no. 311-010-CL WISENT, Inc.). 4. Solution of Trypsin (0.05%) and ethylenediamine tetraacetic acid (EDTA) (0.53 mM) (cat. no. 325-042-EL WISENT, Inc.). 5. Round glass cover slips (25 mm diameter) (Fisher, Pittsburgh, PA).

2.2. Transfection Reagents 1. Cell line Nucleofector Kit V (cat. no. VCA-1003. Amaxa BioSystems. Cologne, Germany).

2.3. cDNA Preparation 1. HiSpeed Plasmid Maxi Kit (cat. no. 12663 QIAGEN Inc. Mississauga, ON).

Fig. 2. Spinning disk confocal microscope. The system, such as the one illustrated, consists of a microscope with arc lamp, illumination lasers, spinning disk assembly, and detector cooled CCD camera, all driven by suitable software (e.g., Volocity) from a desktop computer. A system assembled and marketed by Quorum is shown here.


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2.4. Particle Opsonization 1. Uniform latex microspheres (3.87 μm) (cat. no. PS05N, Bangs Laboratories, Inc., Fishers, IN). 2. Human IgG (Sigma-Aldrich, St. Louis, MO).

2.5. Confocal Microscopy 1. Quorum Spinning Disk Confocal Microscopy System (Quorum, ON, CA) (Fig. 2). Consists of a Leica DMIRE2 inverted fluorescence microscope equipped with a Hamamatsu back-thinned EM-CCD camera and spinning disk confocal scan head. The unit is equipped with four separate diode-pumped solid-state laser lines (Spectral Applied Research, Richmond Hill, Ontario, Canada: 405, 491, 561, 652 nm), an ASI motorized XY stage, an Improvision Piezo Focus Drive and a 1.5X magnification lens (Spectral Applied Research). The equipment is driven by Volocity acquisition software and powered by an Apple Power Mac G5 computer. 2. Attofluor chamber (cat. no. A7816. Molecular Probes, Eugene, Oregon, Invitrogen). 3. Digital temperature regulator with a P-insert (PECON, Germany).

3. Methods 3.1. Cell Culture RAW 264.7 macrophages from the American Tissue Culture Collection (ATCC) grown in DMEM plus 5% FBS are split at 70–80% confluency, diluted approximately 1:2 to 1:8 and seeded at a density of 2–4 × 104 viable cells/cm2 in the same medium (see Note 1). Cells are removed by addition of trypsin and incubation at 37°C for 5–10 min. 3.2. Transfection RAW 264.7 macrophages are transfected by electroporation using the Amaxa system (Cologne, Germany) (see Note 2) by following the manufacturer’s instructions with some modifications: 3.2.1. Day 1 1. Cells are subcultured the day before transfection so that they are in a growth phase by the next day (approximately 70% confluence).

3.2.2. Day 2 1. Prewarm the Nucleofector solution (previously prepared by the addition of the “supplement solution”) to room temperature. Prewarm an aliquot of culture medium (DMEM/5% FBS) to 37ºC.

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2. Prepare a 6-well plate (Becton, Dickinson, Falcon cat. no. 353046) by placing one 25-mm sterile coverslip per well and adding 2 mL of DMEM/5% FBS. Place the plate in the tissue culture incubator. 3. Retrieve cells cultured from Day 1, rinse them once with PBS, and add 2 mL of the trypsin solution. Incubate cells with trypsin for about 5–10 min at 37°C (see Note 3). Stop trypsinization by adding 5 mL of DMEM/5% FBS. Take an aliquot of the cells and determine the concentration of viable cells by adding Trypan blue and counting the cells on a hemocytometer. 4. Centrifuge the required number of cells (2 × 106 cells per sample) at 100g for 10 min. Discard the supernatant. 5. Resuspend the pellet in Nucleofector solution pre-equilibrated to room temperature to a final concentration of 2 × 106 cells/100 μL. Add 2 μg of cDNA (final concentration 0.5–1 μg/μL), mix by pipetting and transfer the suspension to the electroporation cuvette. It is important to avoid maintaining the cell suspension longer than 15–20 min, since prolonged exposure to the Nucleofector solution reduces cell viability and gene transfer efficiency. 6. Insert the cuvette into the electroporator and apply program D-32 (see Note 4). 7. Remove the cuvette from the electroporator, add 500 μL of warm culture medium (DMEM/5% FBS) as soon as possible, remove the cells with a transfer pipet provided by the supplier, and seed them onto the prepared 25-mm coverslip, previously placed on the 6-well plate. 8. Place the plate in the tissue culture incubator. Depending on the construct utilized, expression is detectable after 6–24 h.

Generally, 40% of the surviving cells are transfected, although these results may vary with the particular gene expressed. 3.3. Opsonization of Latex Beads 1. Aliquot 200 μL of latex beads directly from the stock suspension (10% solids). Spin the beads in a microcentrifuge at maximum speed for 1 min, remove the supernatant, and wash the beads three times by resuspending them in 1 mL of PBS. 2. After the last wash, resuspend the beads in 300 μL of PBS and add 10 μL of human IgG (stock 2 mg/mL). Incubate for 1 h at room temperature with continuous rotation. 3. Wash 3 times in 1 mL of PBS. Resuspend beads in 1 mL of PBS after the last wash.

3.4. Phagocytosis Assay 1. Some time after transfection (6–24 h), the coverslip with adherent cells is removed from the culture plate and transferred to a clean “Attofluor” chamber and 1–1.5 mL of cold solution HPMI is added (see Note 5). 2. Add 50 μL of the opsonized latex bead suspension.


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3. Place the chamber on ice and bring it to the spinning disk confocal microscope (see Note 6). 4. Place the chamber on the temperature regulator. 5. Locate the cells and identify the focal plane under bright field illumination by using 63X or 100X oil immersion objectives. Detect the transfected cells by epifluorescence using the binoculars and the appropriate fluorescence cube and switch immediately to confocal mode using the appropriate laser line and filter set of the spinning disk. Adjust the conditions for image acquisition, particularly the detector gain and amplifier offset, as well as the frequency and total time of acquisition (see Note 7). 6. Replace the medium for prewarmed serum-free HPMI and set the temperature regulator at 37°C. Initiate recording of fluorescence during the course of particle ingestion and/or maturation.

3.5. Fluorescence Measurements To quantify the recruitment of the probes to a specific cell location, fluorescence intensity can be measured precisely by acquiring digital images, which are then analyzed with suitable software (see Note 8). Quantitation is performed by selecting the region of interest within the cell and comparing its intensity to the cytosolic fluorescence. To allow comparison among cells and between experiments, the background-corrected value is normalized for expression level by dividing it by the cytosolic fluorescence of the same cell. 4. Notes 1. RAW 264.7 cells should not be allowed to reach confluence, as this can lead to cell rounding and overcrowding, which reduces the phagocytic index and complicates the analysis of individual cells. Each batch of cells should be passaged a maximum of 20 times to avoid senescence. 2. In our hands, electroporation of RAW 264.7 macrophages using the Amaxa system is the method that yields the highest transfection efficiency. However, only a fraction of the cells subjected to electroporation survive the procedure, so a comparatively large number of cells need to be transfected in order to obtain a reasonable number of transfectants. Another complication is that the levels of expression of the transfected construct can be very high, potentially interfering or causing loss of cell viability. Other methods for delivery of cDNA that may be considered include the infection of cells with retroviral or lentiviral vectors. This approach is often efficient and yields high cell survival. The level of expression is in general more moderate, less likely to interfere with cell function. On the other hand, construction and packaging of viral vectors can be very laborious and expensive. 3. If the cells haven’t detached completely from the plate after 10 min of incubation with trypsin, scrape them very gently after neutralizing the protease.

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4. We have noticed that the application of program U-14 yields higher transfection efficiency than program D-32. However, the survival of the cells is reduced and 3 million cells should be used per transfection, instead of 2 million. 5. The incubation in cold medium slows down phagocytosis, which facilitates synchronization of the process. 6. The most common type of confocal microscope, the laser scanning microscope, uses a single focused laser beam that progressively rasters an entire region of interest (54). In contrast to the scanning microscope, the spinning disk confocal microscope uses a parallelized approach of multibeam scanning (54) by means of a rapidly spinning Nipkow disk with multiple pinholes in the light path, which results in the near-simultaneous excitation of the entire field of view. This enables the spinning disk system to acquire entire images at a very high speed, unlike the scanning laser microscope. In addition, the quantum efficiency of the spinning disk confocal detector (a CCD, or charge coupled device) is significantly higher than that of the photomultiplier tube used by the scanning microscope. Furthermore, since in the spinning disk confocal the excitation light is split into many mini-beams of correspondingly lower intensity, the overall photobleaching and phototoxicity is minimized, making it ideal for live cell, timelapse imaging. However, it is important to mention that spinning disk microscopes are not equipped for regional bleaching that is required for photobleaching and photoactivation measurements. 7. For phagocytosis assays we usually acquire images every 5–10 s over a 5- to 7-min period. 8. Programs that can be used for this purpose include Image J, Inc. (freely accessible to the general public at http://rsb.info.nih.gov/ij/) or MetaMorph (Universal Imaging Corp.).

Acknowledgments Original work in the authors’ laboratory is supported by the Heart and Stroke Foundation of Ontario, the Canadian Cystic Fibrosis Foundation, and the Canadian Institutes for Health Research. G.C. is a fellow of the McLaughlin Centre for Molecular Medicine, and S.G. is the current holder of the Pitblado Chair in Cell Biology.

References 1. Yeung, T., Ozdamar, B., Paroutis, P. and Grinstein, S. (2006) Lipid metabolism and dynamics during phagocytosis. Curr. Opin. Cell Biol. 18, 429–437. 2. Stuart, L. M. and Ezekowitz, R. A. (2005) Phagocytosis: elegant complexity. Immunity 22, 539–550. 3. Vieira, O. V. et al. (2001) Distinct roles of class I and class III phosphatidylinositol 3-kinases in phagosome formation and maturation. J. Cell Biol. 155, 19–25.


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19 In Vitro Phagosome–Endosome Fusion Isabelle Vergne and Vojo Deretic

Summary Phagolysosome biogenesis plays a pivotal role in elimination of foreign particles and pathogens by leukocytes. This process is achieved by multiple cycles of membrane fusion between the phagosome and the endosomal system. In vitro reconstitution of phagosome fusion with endosomes is instrumental in studying this intricate process. Such an assay is also invaluable for the identification of effectors produced by intracellular pathogens that may hamper the pathway. In this chapter we describe a highly sensitive and relatively rapid method to measure fusion between phagosomes and early, as well as late, endosomal compartments. The assay is based on the formation of a stable biotin–streptavidin complex upon fusion between biotinylated–peroxidase loaded endosomes and magnetic streptavidin conjugated bead-containing phagosomes. Fusion is quantified using a fluorescence-based detection method that measures the peroxidase activity associated with the beads.

Key Words: Phagosome; endosomes; macrophages; fusion; cell-free system; magnetic beads.

1. Introduction Phagolysosome biogenesis, also referred as phagosome maturation, is a highly regulated membrane trafficking process essential for killing and degradation of pathogens, as well as for antigen presentation by professional phagocytes (1,2). Once engulfed by phagocytosis, microorganisms or inert particles reside in a specialized organelle, the phagosome. The biogenesis of the phagolysosome results from a sequential series of fusion events between the phagosome and the compartments of the endocytic pathway. Typically, the newly formed phagosome undergoes maturation by fusing initially with early endosomes, then with the late endosomes, and finally with lysosomes. The From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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use of cell-free assay that reconstitutes fusion of phagosomes with endocytic compartments is a very powerful tool to dissect this extremely complex and dynamic process and to uncover new pathogenic effectors interfering with normal biogenesis. The first biochemical assay that monitored fusion between phagosomes and early endosomes was developed by Stahl and colleagues (3). This pioneering work found that the system required cytosol, adenosine triphosphate (ATP), and NEM-sensitive fusion protein (NSF) (3). However, this procedure did not allow the reconstitution of phagosome fusion with later components of the endocytic pathway. A few years later, Jahraus et al. published a new assay and showed the involvement of Rab5 in fusion of latex bead–containing phagosomes with early endosomes (4). Although this protocol had the advantage to permit measurement of fusion between phagosomes and late endosomes, the time-consuming step of phagosome isolation and reisolation had prevented the utilization of this assay for extensive studies. The procedure described in this chapter is a modified and optimized version of the one developed by Jahraus et al. (4), with the yield of a rapid and sensitive biochemical assay in mind. The method is based on the same principle as the one used in the earliest studies (3,4), which is the content mixing of phagosomes and endosomes and the formation of a stable complex upon fusion (Fig. 1). The endosomes are loaded with biotinylated peroxidase, and the specific loading of early endosomes and late endosomes is achieved by employing different pulse-chase incubation times. Separately, macrophages are allowed to uptake magnetic streptavidin-conjugated beads and to form phagosomes. The introduction of magnetic beads permits rapid and simple isolation of phagosomes and, therefore, significantly reduces the experimental time. It also allows an increase in the number of conditions tested per set of experiments and facilitates washing steps. The endosome and phagosome populations are mixed in the presence of cytosol and ATP. Fusion between the bead-containing phagosome and endosomes results in formation of a stable biotin–streptavidin complex that can be quantified after membrane permeabilization by measuring peroxidase activity associated with the beads. Fusion is directly proportional to the amount of complexes formed and, thereby, to peroxidase activity. High sensitivity is obtained by using a fluorogenic peroxidase substrate. Another advantage of this procedure is that it does not require prelabeling of the beads or the endosome probe. This assay can be used to directly test the effect of pathogenic factors on different steps of phagolysosome biogenesis and to examine the involvement of host proteins and of signaling lipids in phagosome maturation. In combination with in vivo studies, this assay was key in demonstrating the role of Rab14 in the fusion of phagosomes with early endosomes, as well as examining the

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Fig. 1. Outline of the method measuring in vitro fusion between phagosomes and endosomes. Isolated phagosomes containing magnetic streptavidin beads are incubated with endosomes loaded with biotinylated peroxidase in presence of ATP-regenerating system, salts, and cytosol for 1 h at 37°C. Then, organellar membranes are lysed and magnetic beads are washed several times. Peroxidase activity associated with the beads is measured by using a fluorogenic substrate.

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action of SapM, an M. tuberculosis PI3P phosphatase, and phosphatidylinositol mannoside on phagolysosome biogenesis (5–7). 2. Materials 2.1. Cell Culture, Transfection 1. Macrophage culture: J774A.1 (ATCC TIB-67) and RAW264.7 (ATCC TIB-71) cells are grown in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco/BRL, Bethesda, MD) supplemented with 10% fetal bovine serum (FBS, Hyclone, Ogden, UT) and 4 mM l-glutamine (BioWhittaker). 2. 293T cell medium: DMEM supplemented with 10% FBS, 4 mM l-glutamine, penicillin 100 units/mL and streptomycin 100 μg/mL (Invitrogen, Carlsbad, CA). 3. 293T cell transfection: Calcium phosphate transfection kit (#K2780-01, Invitrogen).

2.2. Phagosome, Endosomes, and Cytosol Preparation 1. Streptavidin magnetic beads: BioMag® binding sreptavidin (Polysciences, Warrington, PA). 2. Peroxidase–biotinamidocaproyl conjugate (Biot-HRP, Sigma, St. Louis, MO). 3. Internalization medium: 150 mM NaCl, 20 mM hydroxyethyl piperazine ethane sulfonate (HEPES) pH 7.4, 6.5 mM glucose, 1 mg/mL bovine serum albumin (BSA). 4. BioMag Multi-32 Microcentrifuge Tube Separator (Polysciences). 5. Protease inhibitors. Prepare the following stock solutions: 10 mg/mL Na -Tosyl-llysine chloromethyl ketone (TLCK, Sigma), 1.79 mg/mL E64 (in DMSO, Sigma), 1 mM pepstatin (in DMSO, Sigma), 10 mg/mL leupeptin (Sigma). 6. Homogenization buffer. For cytosol: 250 mM sucrose, 3 mM imidazole pH 7.2 (HB1). For endosomes: 250 mM sucrose, 20 mM HEPES/KOH pH 7.2, 0.5 mM ethylene glycol-bis(-aminoethylether)-N,N,N´,N´-tetraacetic acid (EGTA) (HB2). For phagosomes: 250 mM sucrose, 3 mM imidazole pH 7.2, 0.5 mM EGTA (HB3). Add to each homogenization buffer 1/1000 of protease inhibitors. 7. Phosphate-buffered saline (PBS) and PBS with 5 mg/mL BSA. 8. Syringe apparatus for homogenization: two 22-gauge needles, two plastic syringes (3 mL), one tubing (5 inches, Pharmed AYX42605). Fit 22-gauge needle to each syringe and connect syringes with 5 inches tubing (Fig. 2A).

2.3. Fusion Reaction 1. Biotinylated-insulin: Insulin-biotinamidocaproyl labeled (Sigma) is dissolved at 5 mg/mL in distilled water, aliquoted, and stored at –20°C. 2. ATP-regenerating system: Prepare MgATP solution (42 mM, adjust to pH 7 with KOH, Sigma), creatine phosphate solution (336 mM, Sigma), and creatine


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Fig. 2. (A) Syringe apparatus for cell homogenization. Two 3-mL syringes fitted with 22-gauge needle are connected with 5 in. tubing. (B) Isolation of phagosomes containing magnetic beads. After phagocytosis of magnetic beads, macrophages are homogenized and phagosomes are isolated by placing the cell extract in a multi32 microcentrifuge tube separator for 15 min. Before separation (right tube); after separation (left tube). phosphokinase solution in 50% glycerol (1180 units/mL, Sigma). Each solution is aliquoted and stored at –20°C. Just before starting the fusion reaction, prepare ATP-regenerating system by mixing 1/1/1 ATP, creatine phosphate, and phosphocreatine kinase (in that order). 3. ATP-depleting system: Prepare apyrase solution in distilled water (278 units/mL), aliquot and store at –20°C. Prepare glucose solution (278 mM) and store at 4°C. Just before starting the fusion reaction, prepare ATP-depleting system by mixing 1/1 apyrase and glucose solutions. 4. Fusion buffer: 190 mM HEPES/KOH pH 7.2, 10 mM dithiothreitol (DTT), 15 mMMgCl2 , 500 mM KCl, 3 mM EGTA, 750 mM sucrose.

2.4. Permeabilization and Washing Buffers 1. Permeabilization buffer: 1% Triton X100, 0.1% (w/v) sodium dodecyl sulfate (SDS), 50 mM NaCl, 10 mM Tris-HCl, pH 7.5, 1 mg/mL heparin, 100 μg/mL biotinylated insulin. 2. Washing buffer: 1% (v/v) TritonX100, 0.1% (w/v) SDS, 50 mM NaCl, 10 mM Tris-HCl, pH 7.5, 1 mM ethylenediamine tetraacetic acid (EDTA), 0.5% fish skin gelatin (preheated 15 min, 100°C) (Sigma).

2.5. Peroxidase Activity 1. Fluorescent substrate: QuantaBlu fluorogenic peroxidase substrate (Pierce, Rockford, IL). 2. 96-well black plates (Nunc). 3. Fluorescence plate reader (Gemini, Molecular Devices, Sunnyvale, CA).

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3. Methods 3.1. 293T Transfection 1. 293T cells are seeded at 1 × 106 per 100-mm tissue culture dish. Prepare 20 dishes. 2. 293T cells are transfected with calcium phosphate tranfection kit, according to the manufacturer’s protocol (http://www.invitrogen.com). 3. Transfections are incubated for 48 h. 4. Wash cells with PBS and prepare cytosol as described in Subheading 3.2. (resuspend cells in 1 mL of HB1 prior to homogenization) (see Note 1).

3.2. Cytosol Preparation 1. 2. 3. 4. 5. 6.

7. 8. 9. 10. 11.

Grow macrophages in T175 flasks (24 flasks) to 80% confluency (see Note 2). Wash cells twice with cold PBS. Scrape cells in 12.5 mL cold PBS. Pool cells, wash them once with cold PBS and once with cold HB1. Resuspend cells in 2 mL HB1. Homogenize cells by passing them, 10–20 times, through a syringe apparatus (see Note 3). The homogenization is carried out until about 90% of cells are broken without major breakage of the nucleus, as monitored by light microscopy. Centrifuge the homogenate at 3000g for 20 min at 4°C. Centrifuge the supernatant at 100,000g for 1 h at 4°C in 1.5 mL polyallomer microcentrifuge tube (Beckman, Palo Alto, CA). Cytosol is obtained by collecting the supernatant. Make 50-μl aliquots, freeze rapidly in ethanol–dry ice and store at –80°C, up to 1 yr. The protein concentration of the cytosol should be 6 mg/mL minimum. The day of the fusion reaction: thaw aliquots in 37°C water bath, adjust the protein concentration to 6 mg/mL with HB1, and keep them on ice.

3.3. Endosome Preparation The preparation of endosome-enriched fractions has been adapted from Mayorga et al. (3). 1. 2. 3. 4. 5.

Grow macrophages in T175 flasks (24 flasks) to 80% confluency. Wash cells twice with cold PBS. Scrape cells in 12.5 mL cold PBS. Pool cells, wash them once with cold PBS and once with internalization medium. Cells are resuspended in 3 mL of internalization medium containing 1.7 mg/mL biot-HRP. 6. For loading of early endosomes: incubate cells for 5 min at 37°C, 5% CO2 . Wash cells twice with cold PBS with 5 mg/mL BSA and twice with cold PBS. For loading of late endosomes: incubate cells for 20 min at 37°C, 5% CO2 .


7. 8. 9. 10. 11.

12. 13. 14. 15.

16. 17.

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Wash cells twice with cold PBS with 5 mg/mL BSA and once with cold PBS. Resuspend cells with 12.5 mL warm media and incubate for 40 min at 37°C, 5% CO2 . Wash cells once with cold PBS. Cells are washed once with HB2. Cells are resuspended in 7.5 mL of HB2. Homogenize cells by passing them, 5–10 times, through a syringe apparatus (2.5 mL of cell suspension per apparatus). Pool the homogenates. Pellet unbroken cells and nuclei at 200g for 6 min. Take the supernatant and repeat this step until no more pellet is observed. The last supernatant is the postnuclear supernatant (PNS). Make aliquots (approximately 400 μL), freeze in liquid nitrogen, and store at –80°C up to 1 yr. The day of the fusion reaction: thaw 3 aliquots of PNS in a 37°C water bath. Pool the aliquots and dilute up to 5.5 mL with HB2. Transfer the suspension to Swti55 ultra-clear centrifuge tube (Beckman, Palo Alto, CA). For early endosomes: centrifuge 1 min, 20,000g at 4°C. Then, centrifuge the supernatant 5 min, 23,000g at 4°C. Resuspend the pellet with 90 μL of cold HB2 (enough for 15 fusion reactions). For late endosomes: centrifuge 1 min, 16,000g at 4°C. Then, centrifuge the supernatant 5 min, 23,000g at 4°C. Resuspend the pellet with 90 μL of cold HB2. Keep the endosomes on ice.

3.4. Phagosome Preparation 1. Grow macrophages in T175 flasks (8 flasks) for 2 d to 80% confluency. 2. Remove media, add in each flask 12.5 mL of internalization medium containing 2.3% (v/v) of magnetic beads (37°C), and incubate cells for 30 min at 37°C, 5% CO2 . 3. For late phagosomes only: remove medium, wash three times with warm DMEM, 10% FBS, and incubate cells with DMEM, 10% FBS for 10 min at 37°C, 5% CO2 . 4. Wash cells three times with cold PBS. 5. Scrape cells in 12.5 mL cold PBS. 6. Pellet cells (centrifugation 10 min at 200g) and wash once with cold HB3. 7. Resuspend cells with 5 mL cold HB3. 8. Homogenize cells and prepare PNS as described in Subheading 3.3. 9. Isolate magnetic bead containing phagosomes using the microcentrifuge tube separator, 15 min at 4°C (Fig. 2B). 10. Remove the supernatant and resuspend beads with 900 μL of cold HB3. 11. Keep the phagosomes on ice.

This amount of phagosomes allows 15 fusion reactions.

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3.5. Fusion Reaction 1. Place 15 Eppendorf tubes (five conditions in triplicate) on ice. 2. Add to each Eppendorf tubes the following in order: 50 μL of phagosome suspension, 4.2 μL of biotinylated-insulin (see Note 4), 50 μL of cytosol, 5 μL of endosome suspension, 15 μL of fusion buffer. 3. Add for one condition 10.8 μL of ATP-depleting system and the other conditions 10.8 μL of ATP-regenerating system. 4. Add 15 μL of compound to be tested (inhibitors, effectors) or buffer used for solubilization (control). 5. The fusion reaction is initiated by placing the eppendorf tubes at 37°C for 1 h on a rocking table.

3.6. Permeabilization and Washes 1. 2. 3. 4. 5. 6. 7. 8.

Stop the reaction by placing the tubes on ice. Add 650 μL of HB2 in each tube. Isolate the magnetic beads with separator for 15 min at 4°C. Resuspend the beads in permeabilization buffer (900 μL). Incubate 30 min on ice. Isolate beads. Wash beads two times with washing buffer and three times with PBS. Resuspend beads in 50 μL PBS.

3.7. Peroxidase Activity Associated with Beads 1. Mix 9/1 (v/v) QuantaBlu substrate and QuantaBlu stable peroxide solution, keep on ice. 2. Add 150 μL of QuantaBlu mix to each 50 μL of bead suspension, vortex, and incubate for 30 min at 37°C. 3. Stop reaction by putting tube on ice and by adding 150 μL of Stop solution. 4. Vortex and spin down beads for 5 min at 12,000g (microcentrifuge). 5. Transfer 250 μL of supernatant to 96-well plate. 6. Read relative fluorescence unit (RFU) at 405 nm by exciting at 320 nm using a fluorescence plate reader.

3.8. Data Analysis Fusion efficiency is determined by subtracting RFU of fusion reaction with ATP-depleting system to RFU of reaction with ATP-regenerating system. 4. Notes 1. Because they are highly transfectable, 293T cells are used instead of macrophages to prepare cytosol containing large amount of ectopically expressed protein. To


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allow fusion 293T cytosol is mixed with cytosol prepared from macrophages (1/5, v/v). 2. Cytosol is depleted of specific protein by transfecting RAW264.7 cells for 24 h with siRNA (see Chapter 26 for protocol). Use scrambled siRNA as a control. 3. To avoid the formation of bubbles during homogenization, the needles and the tubing are preloaded with homogenization buffer. 4. Biotinylated insulin is added to the fusion reaction to prevent formation of complexes that could result from damaged organelles.

Acknowledgments The author would like to thank Alex S. Davis for helping with the pictures and Esteban Roberts for critical reading of the manuscript. This work was supported by Grant AI45148 from the National Institutes of Health. References 1. Vergne, I., Chua J., Singh S. B., and Deretic, V. (2004) Cell biology of mycobacterium tuberculosis phagosome. Annu. Rev. Cell Dev. Biol. 20, 367–394. 2. Jutras, I., and Desjardins, M. (2005) Phagocytosis: at the crossroads of innate and adaptive immunity. Annu. Rev. Cell Dev. Biol. 21, 511–527. 3. Mayorga, L. S., Bertini, F., and Stahl, P. D. (1991) Fusion of newly formed phagosomes with endosomes in intact cells and in a cell-free system. J. Biol. Chem. 266, 6511–6517. 4. Jahraus, A., Tjelle, T. E., Berg T., et al. (1998) In vitro fusion of phagosomes with different endocytic organelles from J774 macrophages. J. Biol. Chem. 273, 30379–30390. 5. Kyei, G. B., Vergne, I., Chua, J. et al. (2006) Rab14 is critical for maintenance of Mycobacterium tuberculosis phagosome maturation arrest. EMBO J. 25, 5250–5259. 6. Vergne, I., Chua, J., Lee, H. H., Lucas, M., Belisle, J., and Deretic, V. (2005) Mechanism of phagolysosome biogenesis block by viable Mycobacterium tuberculosis. Proc. Natl. Acad. Sci. USA 102, 4033–4038. 7. Vergne, I., Fratti, R. A., Hill, P. J., Chua, J., Belisle, J., and Deretic, V. (2004) Mycobacterium tuberculosis phagosome maturation arrest: mycobacterial phosphatidylinositol analog phosphatidylinositol mannoside stimulates early endosomal fusion. Mol. Biol. Cell 15, 751–760.

20 Real-Time Spectrofluorometric Assays for the Lumenal Environment of the Maturing Phagosome Robin M. Yates and David G. Russell

Summary The ultimate goal of phagosomal maturation is the delivery of internalized, particulate cargo to acidic, hydrolytically competent compartments capable of mediating its degradation. Here we outline in detail three fluorometric techniques that allow the study of phagosomal maturation in macrophages by quantifying functionally important features of the lumenal environment of the developing phagosome in real time. The first assay utilizes a particle-restricted, pH-sensitive fluorochrome to measure the acidification of the phagosome. The second reports on the development of the proteolytic capacity of the phagosome by following the hydrolysis of a fluorogenic, generic proteinase substrate. The third quantifies the accumulation of lysosomal constituents within the phagosome by measuring the fluorescence resonance energy transfer (FRET) efficiency between a particle-restricted, donor fluor and a fluid phase acceptor fluor that had been chased previously into lysosomes. The assays are described as population-based methodologies utilizing a spectrofluorometer but, alternatively, can be adapted readily to confocal-based technologies for single phagosomal measurements.

Key Words: Phagosome; lysosome; phagocytosis; macrophage; pH; proteolysis; lumen.

1. Introduction The phagosome is an exquisitely dynamic organelle. While traditional, static techniques have led to much of today’s knowledge of the phagosome, real-time fluorescent approaches are necessary for the kinetic analysis of phagosomal development. The majority of the more recent techniques focus on changes to the cytoplasmic aspect of the phagosome, concentrating on regulatory or From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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signaling components. In contrast, techniques that report on the dynamics of the lumenal environment remain scarce. This chapter focuses on three real-time, fluorometric techniques that quantify the development of functionally relevant aspects of phagosomal lumenal biology, namely pH, proteolytic capacity, and relative acquisition of lysosomal cargo. These assays give the investigator three distinct yet interdependent parameters that proceed with different kinetics and collectively document the maturation of the lumenal aspect of the phagosome. Lumenal acidification occurs within seconds of phagosomal formation principally via recruitment and activity of vacuolar-ATPases (V-ATPases) (1,2). Typically, within 15–30 min, the phagosome reaches and maintains a lumenal pH between 4.5 and 5 (3). This rapid and complete acidification is an important early feature of the phagosome and directly influences its enzymatic activites and physical properties (4). The pH-sensitive fluorochromes carboxyfluorescein and Oregon green have both been utilized by a number of investigators to measure pH of various subcellular compartments (1,2,5,6). The fluorescent emission from carboxyfluorescein and Oregon green are differentially affected by changes in pH, varying with the wavelength of excitation. In particular, fluorescent emission of carboxyfluorescein at 520 nm is more sensitive to decreasing pH when excited at 490 nm than it is at 450 nm. This allows the excitation ratio to be used as an internally controlled value that reports on the pH of the environment immediately surrounding the fluorophore. This value can be converted to pH with polynomial regression to a standard curve. Here we outline the measurement of phagosomal pH in macrophages using carboxyfluorescein covalently coupled to experimental particles. The phagosome of the macrophage is adept at degrading most biological macromolecules, this process being central to its function. In particular, a large proportion of the hydrolytic machinery is dedicated to the hydrolysis of protein (7). The development of the proteolytic capacity of the phagosome is multifactorial, requiring delivery and activation of proteases such as the cathepsins and an acidic pH (4). In this chapter we describe an assay that measures the bulk proteolytic activity of the maturing phagosome, utilizing particle-associated albumin labeled heavily with a self-quenching fluor. Following phagocytosis, proteolysis results in the dequenching of the substrate’s fluor relative to a calibration fluorochrome on the particle. The rate of change of the ratio of the two fluorescent intensities is indicative of the bulk proteolytic capacity of the phagosomal lumen. Finally, it is accepted generally that the maturation of a phagosome is more intricate than the single fusion event with one lysosome. Rather, the phagosome progressively accumulates membranous and lumenal components of the late endosomes and lysosomes through multiple fusion events (8). Yet many investigators use phagosomal-lysosomal (P-L) fusion as an indicator of

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phagosomal maturation by counting phagosomes that are deemed “fused” with a lysosome, implying a single, absolute, fusion event. In essence, they are counting the number of phagosomes that have accrued sufficient lysosomal reporter exceeding a threshold imposed by the sensitivity of the apparatus or, most frequently, the human eye. The assay outlined in this chapter quantifies the accumulation of preformed lysosomal constituents within the maturing phagosome in real-time by exploiting fluorescence resonance energy transfer (FRET) between an acceptor fluor that has been chased into lysosomes and a particle-restricted donor fluor. The profiles generated report on the kinetics and extent of lysosomal contribution to the phagosomal content. The particular methodologies outlined in this chapter are described as population-based assays with measurements taken from an area of a monolayer containing up to 4 × 104 macrophages. Population-based methodologies offer a number of advantages over single-cell techniques: they require comparatively basic instrumentation and technical expertise; there is a reduced need for qualitative assessment, hence reducing user bias; and as phagosomal populations are relatively heterogeneous (9,10), measurements taken over a large population are often statistically sound, thus offering the investigator the opportunity to more confidently test numerous variables in a relatively short period. Populationbased assays, however, do not necessarily obviate the need for single-cell measurements. As an alternative, the techniques outlined in this chapter can be readily applied to a single cell format, although the confocal instrument required has to be capable of performing spectral separation for the FRET assays. 2. Materials 2.1. Cells, Reagents, and Buffers 1. Macrophages: Both bone-marrow–derived murine macrophages (BMMØ) and macrophage-like cell lines have been used (see Note 1). BMMØ s are derived from the bone marrow extracted from the femurs, tibias, and iliums of euthanized mice and maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS), 5% horse serum, 2 mM l-glutamine, 1 mM sodium pyruvate, and 20% L-cell conditioned media (BMMØ media). RAW 264.7 cells (available from the American Type Culture Collection, Rockville, MD) are maintained in DMEM supplemented with 10% FBS and 1.5 g/mL sodium bicarbonate. 2. Cover slips: Clean 0.13 × 12.5 × 25 mm cover glass (see Note 2). Sterilize by autoclave. 3. Binding buffer: Tissue culture tested PBS pH 7.2 (Gibco) adjusted to contain 1 mM CaCl2 , 2.7 mM KCl, 0.5 mM MgCl2 , 5 mM dextrose, 10 mM hydroxyethyl piperazine ethane sulfonate (HEPES), and 5 % FBS.

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4. Cuvette buffer: Tissue culture tested phosphate-buffered saline (PBS) pH 7.2 (Gibco) adjusted to contain 1 mM CaCl2 , 2.7 mM KCl, 0.5 mM MgCl2 , 5 mM dextrose, 10 mM HEPES, and 0.1 % calf skin gelatin (see Note 3). 5. Binding dish: A microbiological Petri dish containing a square piece of parafilm adhered to the lower plate surrounded by a damp Kimwipe® tissue. 6. 0.4 % Trypan blue (Gibo). 7. Experimental particles: 3.0 μm carboxylate-modified silica particles (Si-COOH) 5% suspension (Kisker Biotech, Steinfurt, Germany) (see Note 4). 8. Amine reactive fluorescent reagents: 5-(and-6)-carboxyfluorescein, succinimidyl ester (mixed isomers) (CF-SE); Alexa Fluor 488® carboxylic acid, succinimidyl ester (mixed isomers) (Alexa488-SE); Alexa Fluor 594® carboxylic acid, succinimidyl ester (mixed isomers) (Alexa594-SE) (Molecular Probes, Eugene, OR). Dissolve in high quality anhydrous dimethylsulfoxide (DMSO) (Sigma Sigma, St. Louis, MO) at 5 mg/mL before use. Stock solutions can be aliquoted and stored at −20ºC. Protect reagents from light and moisture. 9. DQ green bodipy bovine serum albumin (DQ-BSA) (Molecular Probes). Store as desiccate at −20ºC until use. Protect from light. 10. Alexa Fluor® 594 hydrazide, sodium salt (Alexa594-HA) (Molecular Probes). Dissolve in tissue-culture grade PBS pH 7.2 at 1 mg/mL. Store at −20ºC. Protect from light. 11. Cyanamide (Sigma). Store as desiccate at 4ºC. 12. Coupling buffer: 0.1 M sodium borate in ddH2 O Adjust pH to 8.0 with 10 M NaOH. Filter sterilize through 0.22-μm filter. 13. Quenching buffer: 250 mM glycine in PBS pH 7.2. Filter sterilize through 0.22-μm filter. 14. Defatted BSA (Sigma). 15. -d-Mannosylated-PITC-albumin (Sigma). 16. Sodium azide 2 % aqueous solution. Very toxic. 17. Reference pH buffers (pH 4–5.5): 0.15 M potassium acetate. Adjust pH to 4.0, 4.5, 5.0, and 5.5 with 10 M NaOH. 18. Reference pH buffers (pH 6–7.5): 0.1 M piperazine-N,N´–bis(2-ethanesulfonate (PIPES), 0.1 M KCl. Adjust pH to 6.0, 6.5, 7.0, 7.5, and 8.0 with 5 M HCl.

2.2. Instruments 1. Temperature controlled-spectrofluorometer with variable excitation and emission monochromators. The authors use the QMSE4 model spectrofluorometer from Photon Technologies International (Lawrenceville, NJ) equipped with a thermostat-controlled four-chambered turret for simultaneous measurement of four experimental variables. The QMSE4 is interfaced with a PCcompatible computer and is managed by Felix32 software (Photon Technologies International). 2. Quartz 10 × 10 × 45 mm cuvettes (Fisher Scientific, Pittsburgh, PA). 3. Fluorescent microscope with standard FITC and Texas Red filter sets. The authors use the Zeiss Axioskop 2 plus (Carl Zeiss MicroImaging Inc., Thornwood, NY).


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3. Methods 3.1. Macrophage Monolayer Preparation and Handling 1. Fully differentiated BMMØ monolayers are grown to confluency in untreated Petri dishes. Growth media is removed and replaced with cold PBS pH7.2 (without Ca2+ and Mg2+ ) and incubated at 4ºC for 10 min to facilitate BMMØ detachment from the plastic. BMMØ s are then gently dislodged with a rubber policeman and centrifuged at 230g at 4 ºC for 10 min. 2. Sterile, clean 12.5 × 25 mm cover slips are placed in a sterile Petri dish using fine-point forceps that have been dipped in 70% ethanol and flamed (see Note 5). 3. BMMØ s are gently resuspended in 1 mL BMMØ media, counted using a hemocytometer, and diluted appropriately in BMMØ media to achieve ∼1.25 × 106 macrophages/mL. 4. 10 mL of BMMØ suspension is added to the Petri dish and incubated at 37ºC for 24 h to allow a monolayer to establish on the cover slips. Care should be exercised to prevent excessive movement of the cover slips in the Petri dish. The BMMØ monolayer–covered cover slips (subsequently referred to as monolayers) are then ready for fluorometric analysis.

3.2. Spectrofluorometer Setup and Operation 1. The spectrofluorometer should be set up according to manufacturer’s directions such that optimal measurements can be taken using the desired wavelengths (see Note 6). 2. Clean quartz cuvettes containing cuvette buffer are inserted into a thermostatcontrolled sample holder and warmed to 37ºC prior to the loading of the monolayers (see Note 7). 3. Using fine-point forceps, a monolayer-covered cover slip is grasped at one end (see Note 8) and dipped 10 times into a sterile 50-mL tube containing binding buffer to remove BMMØ growth media and loosely adhered cells. The cover slip is then placed in the cuvette with a vertical orientation (length of the cover slip parallel to the long axis of the cuvette), on the diagonal (width of the cover slip at 45º to the short axes of the cuvette) and with the cellular side facing the emission slit, as shown in Fig. 1. 4. At this point, sufficient background measurements of each monolayer are recorded for the wavelengths required. 5. At the conclusion of the background determination, the cover slips are carefully removed from the cuvettes using forceps and placed with the cellular side up on the parafilm within the binding dish. 6. Eighty-five μL of a suspension of the appropriate experimental particles is carefully laid over each monolayer. The meniscus should be maintained over the cover slip at room temperature for 3 min (see Note 9). 7. The cover slips are once again dipped 10 times in binding buffer to remove unbound particles and are placed in the cuvettes with the same orientation.

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Fig. 1. Orientation of the BMMØ monolayer in the spectrofluorometer. Cover glass is oriented to achieve a 45º incidence with the excitation beam (excitation ) and with the cells facing the emission slit.

8. The appropriate successive fluorescent measurements are recorded, alternating between each sample. 9. At the conclusion of the assay, the cover slips are removed by forceps and placed cellular side down onto 30 μL of 0.4 % Trypan blue on a glass slide and examined with bright-field and fluorescence microscopy. Careful attention should be paid to macrophage viability, optimal bead to BMMØ ratio, and the presence of extracellular beads. This is an extremely important control and must be completed with every sample. Data should be disregarded should there be an decrease in macrophage viability, overloaded macrophages, or the presence of any extracellular particles.

3.3. Kinetic Analysis of Phagosomal pH 3.3.1. Preparation of pH-Sensitive Particles 1. Fifty mg of carboxylate-modified silica particles are washed three times in 1 mL of PBS by brief vortexing and centrifugation in a tabletop microfuge at 2000g for 60 s. 2. Particles are resuspended in PBS (pH 7.2) with 25 mg/mL of the heterobifunctional crosslinker cyanamide (freshly made) and incubated at room temperature with


3.

4. 5.

6.

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agitation for 15 min. Excess cyanamide is removed by washing the particles twice with coupling buffer. Particles are resuspended in 1 mL of coupling buffer with 5 mg of defatted BSA and 1 mg of -d-mannosylated-PITC-albumin, and incubated with agitation for 6 h. This step covalently attaches the albumin to the particles through the cyanamide crosslinker. The BSA generally serves as a coupler itself for further derivation with carboxyfluorescein-SE while also serving to coat the electrostatic surface of the particle with protein. The -d-mannosylated-PITC-albumin serves as an opsonic ligand for the mannose receptor. We have previously shown that the phagocytosis of particles coated in this way is primarily directed through the mannose receptor (3) (see Note 10). Particles are washed twice with quenching buffer to quench unreacted cyanamide and twice with coupling buffer to remove soluble amine groups. The particles are resuspended in 1 mL of coupling buffer, and 10 μL of the 5 mg/mL stock of carboxyfluorescein-SE in DMSO is added (see Note 11). Incubation at room temperature with agitation for 1 h allows the particle-bound albumin to be sufficiently labeled with the amine-reactive fluor. The now fluorescent particles are washed three times with quenching buffer and finally resuspended in 1 mL PBS (see Note 12). Ten μL of 2% solution of the preservative sodium azide can be added for storage at 4ºC.

3.3.2. Measurement of Phagosomal Excitation Ratios 1. Thirty μL of stock carboxyfluorescein, mannosylated particles (CF beads) are washed twice with 1 mL sterile PBS to remove traces of sodium azide. 2. The CF beads are resuspended in an appropriate volume of binding buffer to achieve ∼1 × 107 beads/mL (see Note 13). 3. After 450|520 nm and 490|520 nm (excitation |emission ) background measurements are recorded for BMMØ monolayers alone, 85 μL of the diluted CF beads is laid onto the monolayers in the binding dish and incubated at room temperature for 3 min (as per Subheading 3.2.). 4. The monolayers are dipped in binding buffer to remove unbound CF beads and returned to the spectrofluorometer. 5. Emission at 520 nm is recorded with excitation alternating between 450 and 490 nm. Typically an integration time of 1 s per data point is optimal. Data are collected for at least 30 min to follow the complete acidification of the phagosomal lumenal space. 6. At the conclusion of the assay, the monolayers are examined via microscopy with Trypan blue (as per Subheading 3.2.).

3.3.3. Conversion of Excitation Ratio to pH 1. Fifty μL of CF-bead stock is washed twice in ddH2 O and resuspended in 1 mL ddH2 O.

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2. Three mL of each pH standard buffer from pH 4 to 7.5 is added to clean cuvettes, and background values at 450|520 nm and 490|520 nm are determined on the buffers alone. 3. One hundred μL of the CF-bead suspension is added to each cuvette, and emission at 520 nm is measured with excitation alternating between 450 and 490 nm (see Note 14). 4. The average CF-bead 490/450 excitation ratio for each pH standard (minus background if not already deducted by acquisition software) is calculated and plotted against pH. Confirm that the pKa of the bound carboxyfluorescein is close to the pH that is of most interest and that the curve generated is reasonably flat over the pH range required (see Note 15). 5. The polynomial equation that best describes the curve is calculated (Fig. 2A). Usually a third or fourth order polynomial is sufficient. This can be done with standard mathematical software such as Microsoft Excel® or MATLAB® . 6. This equation is used to convert the real time phagosomal 490/450 excitation ratios into pH units and these values are plotted against time (Fig. 2B) (see Note 16).

3.4. Kinetic Analysis of Phagosomal Proteolytic Capacity 3.4.1. Preparation of Proteolytic Reporter Particles 1. Fifty mg of carboxylate-modified silica particles is washed three times in 1 mL of PBS by brief vortexing and centrifugation in a tabletop microfuge at 2000g for 60 s. 2. Particles are resuspended in PBS (pH 7.2) with 25 mg/mL of the heterobifunctional crosslinker cyanamide (freshly made) and incubated at room temperature with agitation for 15 min. Excess cyanamide is removed by washing the particles twice with coupling buffer. 3. Particles are resuspended in 500 μL of coupling buffer with 1mg of DQ green BSA® and 250 μg of -d-mannosylated-PITC-albumin, and incubated with agitation for 12 h. This step attaches the proteolytic reporter DQ green BSA covalently to the particles through the cyanamide crosslinker. The -D-mannosylated-PITC-albumin serves as an opsonic ligand for the mannose receptor. We have shown previously that the phagocytosis of particles coated in this way is primarily directed through the mannose receptor (3) (see Note 10). 4. Particles are washed twice with quenching buffer to quench unreacted cyanamide and twice with coupling buffer to remove soluble amine groups. 5. The particles are resuspended in 1 mL of coupling buffer, and 10 μl of the 5 mg/mL stock of the calibration fluor (Alexa Fluor 594® -SE) in DMSO is added (see Note 11). Incubation at room temperature with agitation for 1 h allows the particle-bound albumin to be sufficiently labeled with the amine-reactive fluor. 6. The now fluorescent particles are washed three times with quenching buffer and finally resuspended in 1 mL PBS (see Note 12). Ten μL of 2% solution of the preservative sodium azide can be added for storage at 4ºC.


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(a)

(b)

Fig. 2. Determination of phagosomal pH. (A) Excitation ratio vs. pH standard curve generated by CF beads in buffers on known pH. The curve of best-fit can be described by the equation y = 0.4233x3 - 2.8693x2 + 7.3391x - 1.5901 (where x = 490/450 excitation ratio and y = pH). (B) Phagosomal acidification profiles in BMMØ s. Phagosomal acidification can be abolished by the V-ATPase inhibitor concanamycin A (100 nm).

3.4.2. Fluorometric Measurement of Phagosomal Proteolysis 1. Thirty μL of proteolytic reporter particles (PR-beads) are washed twice with 1 mL sterile PBS to remove traces of sodium azide.

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2. The PR beads are resuspended in an appropriate volume of binding buffer to achieve ∼1 × 107 beads/mL (see Note 13). 3. After 594|620 nm and 490|515 nm (excitation |emission ) background measurements are recorded for BMMØ monolayers alone, 85 μL of the diluted PR beads is laid onto the monolayers in the binding dish and incubated at room temperature for 3 min (as per Subheading 3.2.). 4. Unbound PR beads are washed from the monolayers by dipping in binding buffer and returned to the spectrofluorometer. 5. Both calibration and reporter fluorescent intensities are measured. The fluorescence of the calibration fluor Alexa Fluor 594 (emission at 620 nm with excitation at 594 nm) should remain constant throughout the assay (see Note 17). The fluorescence of the reporter fluor FL bodipy® (emission at 515 nm with excitation at 490 nm) will increase with progressive proteolysis of the DQ bodipy BSA substrate. Typically an integration time of 1 s per data point is optimal. Data are collected for at least 90 min or until exhaustion of the substrate. At the conclusion of the assay, monolayers are examined via microscopy with trypan blue (as per Subheading 3.2.). 6. Data is exported into a standard spreadsheet application such as Microsoft Excel® . 7. The appropriate background values are deducted (if not already deducted by the acquisition software) and the ratio SRT /C (where SRT = substrate fluorescence in real time and C = calibration fluorescence) is plotted against time (Fig. 3).

Fig. 3. Phagosomal generic proteolytic profiles in BMMØ s. Phagosomal proteolysis can be diminished with the serine-cysteine protease inhibitor leupeptin (100 μg/mL) and the V-ATPase inhibitor concanamycin A (100 nm).


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8. Gradients of portions of the profiles will give relative values of the generic proteolytic capacity of the phagosomal population over that period.

3.5. Kinetic Analysis of Lysosomal Contribution to the Phagosome 3.5.1. Preparation of Donor Fluorescent Particles 1. Fifty mg of carboxylate-modified silica particles is washed three times in 1 mL of PBS by brief vortexing and centrifugation in a tabletop microfuge at 2000g for 60 s. 2. Particles are resuspended in PBS (pH 7.2) with 25 mg/mL of the heterobifunctional crosslinker cyanamide (freshly made) and incubated at room temperature with agitation for 15 min. Excess cyanamide is removed by washing the particles twice with coupling buffer. 3. The particles are resuspended in 1 mL of coupling buffer with 5 mg of defatted BSA and 1 mg of -d-mannosylated-PITC-albumin, and incubated with agitation for 6 h. This step covalently attaches the albumin to the particles through the cyanamide crosslinker. The BSA generally serves as a coupler itself for further derivation with Alex Fluor 488-SE while also serving to coat the electrostatic surface of the particle with protein. The -d-mannosylated-PITC-albumin serves as an opsonic ligand for the mannose receptor. We have shown previously that the phagocytosis of particles coated in this way is primarily directed through the mannose receptor (3) (see Note 10). 4. Particles are washed twice with quenching buffer to quench unreacted cyanamide and twice with coupling buffer to remove soluble amine groups. 5. The particles are resuspended in 1 mL of coupling buffer, and 10 μL of the 5 mg/mL stock of Alex Fluor 488-SE in DMSO is added (see Note 11). Incubation at room temperature with agitation for 1 h allows the particle-bound albumin to be sufficiently labeled with the amine-reactive fluor. 6. The now fluorescent particles are washed three times with quenching buffer and finally resuspended in 1 mL PBS (see Note 12). Ten μL of 2% solution of the preservative sodium azide can be added for storage at 4ºC.

3.5.2. Lysosomal Loading of Acceptor Fluor in Macrophages 1. BMMØ monolayers on glass cover slips (prepared as per Subheading 3.1.) are transferred, using sterile forceps (see Note 8), to a Petri dish containing 100 μg/mL of the acceptor fluor Alexa Fluor 594 hydrazide (Alexa 594-HA) in 6 mL BMMØ growth media (see Note 18). 2. The monolayers are incubated at 37ºC for 3–5 h to allow sufficient pinocytic uptake of the membrane impermeable acceptor fluor. 3. The pulsed BMMØ monolayers are removed by sterile forceps, dipped in BMMØ growth media to remove extracellular acceptor fluor, and placed in a new Petri dish containing fresh, warm BMMØ growth media. The monolayers are then incubated at 37ºC for a further 4–24 h to ensure the lysosomal location of the acceptor fluor (see Note 19).

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3.5.3. Measurement of Phagosomal Acceptor Fluor Accumulation Using FRET 1. Thirty μL of donor fluorescent particles (donor beads) is washed twice with 1 mL sterile PBS to remove traces of sodium azide. 2. The donor beads are resuspended in an appropriate volume of binding buffer to achieve ∼1 × 107 beads/mL (see Note 13). 3. Three fluorescent measurements from BMMØ monolayers are recorded before binding of the donor beads. First, the total acceptor fluor emission at 620 nm is measured with excitation at 594 nm. While this measurement is not used in any further calculation, it is a necessary control to ensure equivalent acceptor fluor loadings and BMMØ monolayer densities of the samples (see Note 20). Second, time should be taken to obtain accurate background values for both the donor and FRET wavelengths at 488|520 nm and 488|620 nm (excitation |emission ), respectively. 4. BMMØ monolayers are carefully removed from the spectrofluorometer, and 85 μL of the diluted donor beads is laid onto the monolayers in the binding dish and incubated at room temperature for 3 min (as per Subheading 3.2.). 5. Unbound donor beads are washed from the monolayers by dipping the cover slips in binding buffer and returning them to the spectrofluorometer (as per Subheading 3.2.).

Fig. 4. Profile of the lysosomal contribution to the phagosome in BMMØ s. Rates and extents of lysosomal contribution to the phagosome can be manipulated by the calmodulin inhibitor W7 (15 μM) and the V-ATPase inhibitor concanamycin A (100 nm).


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6. Both donor and FRET-generated emissions at 520 and 620 nm, respectively, are recorded during excitation at 488 nm. Typically an integration time of 1 s per data point is optimal. Data is collected for at least 3 h to allow the phagosomal population to reach equilibrium with respect to accumulation of lysosomal constituents. At the conclusion of the assay, the monolayers are examined via microscopy with trypan blue (as per Subheading 3.2.). 7. The proportion of the “FRET” signal due to a bleed-though artifact contributed by the beads themselves is then determined. This is a constant used in later calculations. Background measurements at 488|520 nm and 488|620 nm are made with a clean cuvette containing PBS. Five μL of stock donor beads is added to the cuvette and stirred (see Note 14). Emissions at 520 and 620 nm are recorded during excitation at 488 nm. 8. All data are exported into a standard spreadsheet application such as Microsoft Excel. 9. The appropriate background values are deducted (if not already deducted by the acquisition software) and relative FRET units (RFU) defined by the equation: RFU = FRT /DRT – FB /DB (where FRT = FRET generated fluorescent emission in real time, DRT = donor emission in real time, FB = “FRET” signal contribution of the beads alone, DB = donor emission of the beads alone) are calculated and plotted against time (Fig. 4). The RFU at time zero should be close to zero.

4. Notes 1. Primary BMMØ s are generally preferred for their enhanced phagocytic proficiency and adhesion. 2. 0.13 × 12.5 × 25 mm cover glass is not commercially available. Cover glass can be custom ordered from ProSciTech (Thuringowa, Qld, Australia). Alternatively, 25 × 25 mm cover glass is available from Fisher Scientific (Pittsburgh, PA) and can be cut in half by diamond pencil in house. 3. FBS is substituted for gelatin for spectrofluometric assays as it has low autofluorescence with excitation wavelengths above 450 nm. If assays are expected to take greater than 6 h, FBS is preferred for sustained macrophage viability. 4. The authors prefer the 3.0-μm -COOH-modified silica particles due to their ease of handling and low autofluorescence, but polystyrene particles and particles of smaller size can be used successfully. Generally, macrophage-like cell lines do not efficiently phagocytose particles with diameters over 2.0 μm. Therefore particles larger than 2.0 μm are not advised for those cell types. 5. Arrange cover slips so as not to overlap, taking care not to overcrowd them in the Petri dish as cover slips can move after monolayers have been established and damage to BMMØ s can occur. Alternatively, cover slips can be separated from each other using partitioned Petri dishes or 6-well plates. 6. Some general considerations are: the focusing of illumination on sample, the addition of long-pass and short-pass filters, and the adjustment of excitation and

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14.

15.

16.

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Yates and Russell emission slit width to maximize signal-to-noise ratio and to minimize photobleaching. Cuvette buffer should be of a similar temperature to the cuvettes at addition. This prevents bubble formation that can create unwanted scatter of light. Cover slips should only be grasped by forceps at the uppermost edge to avoid damaging the area of the monolayer that is illuminated in the spectrofluorometer. Binding time may need to be increased if smaller particles or polystyrene beads are used. Particles can be directed though Fc receptors by substitution of the -dmannosylated-PITC-albumin with the equivalent amount of human IgG. Ensure that the fluor/DMSO aliquot is fully thawed and at room temperature before opening to avoid wetting the DMSO. Labeled particles should be microscopically examined to ensure adequate fluorescence and absence of clumping. A suitable dilution of beads should be determined for each batch. This should be done by applying the bead suspension to a BMMØ monolayer on cover glass for the desired binding time, washing off unbound beads, and assessing degree of bead adhesion under magnification. For most applications a target of one to two beads/macrophage is desired. Due to their low density, significantly higher concentrations of polystyrene beads are needed to achieve optimal binding to monolayers. Ensure that beads are homogeneously suspended in the cuvette. If the spectofluorometer is not equipped with magnetic stirrers, use a pipet to disperse CF beads immediately before measurement. The pKa of free carboxyfluorescein-SE is 6.4. We have found that, when bound to BSA on our experimental particles as described in Subheading 3.3.1., the pKa is shifted to ∼5.5, making it particularly useful in the generation of phagosomal pH profiles. However, should the pKa of the CF beads be inappropriately high, thus rendering measurement of lower pH values inaccurate, then Oregon greenSE should be used in addition to carboxyfluorescein-SE. Standard curves can also be generated using phagocytosed beads on BMMØ monolayers using the ionophore nigericin (10 μM) in K+ -containing buffers of known pH. This eliminates the possibility of modification of the fluor’s pKa by the intracellular environment. For convenience we routinely generate standard curves using a suspension of beads as described in Subheading 3.3.3. In each experimental setup, however, it should be determined that these curves are equivalent to those generated with bead-containing monolayers treated with nigericin. Standard pH curves need to be generated at the conclusion of every phagosomal pH experiment. Subtle changes to components of the experiment, such as degree of bead labeling, slit width, and PMT voltage, can have profound effects on the relationship between excitation ratio and actual pH. It is preferred, but not usually necessary, to record the calibration fluorescence throughout the entire assay. Recording a calibration fluorescence at the beginning


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and/or conclusion of the assay will increase time resolution of reporter fluor profiles and may allow the use of filter sets that improve signal-to-noise ratios. 18. Pulse media should be filter sterilized through a 0.22-μm filter after the Alexa 594-HA is added. If sterility is maintained, the pulse media can be reused up to three times. 19. A gradual loss of acceptor fluor will occur over time. This loss, however, is minimal for chase periods up to 12 h and can be compensated by increasing pulse periods. 20. A large discrepancy in total acceptor fluorescence values indicates that there are inequities in either loading of acceptor fluor or BMMØ monolayer density. In either case, a monolayer with altered acceptor fluorescence should be replaced with one with acceptor fluorescence commensurate with the other samples.

References 1. Lukacs, G. L., Rotstein, O. D., and Grinstein, S. (1990) Phagosomal acidification is mediated by a vacuolar-type H(+)-ATPase in murine macrophages. J. Biol. Chem. 265, 21099–21107. 2. Hackam, D. J., Rotstein, O. D., Zhang, W. J., et al. (1997) Regulation of phagosomal acidification. Differential targeting of Na+/H+ exchangers, Na+ /K+ ATPases, and vacuolar-type H+ -atpases. J. Biol. Chem. 272, 29810–29820. 3. Yates, R. M., and Russell, D. G. (2005) Phagosome maturation proceeds independently of stimulation of toll-like receptors 2 and 4. Immunity 23, 409–417. 4. Yates, R. M., Hermetter, A., and Russell, D. G. (2005) The kinetics of phagosome maturation as a function of phagosome/lysosome fusion and acquisition of hydrolytic activity. Traffic 6, 413–420. 5. Schlesinger, P. H. (1994) Measuring the pH of pathogen-containing phagosomes. Methods Cell Biol. 45, 289–311. 6. Dunn, K. W., Mayor, S., Myers, J. N., and Maxfield, F. R. (1994) Applications of ratio fluorescence microscopy in the study of cell physiology. FASEB J. 8, 573–582. 7. Garin, J., Diez, R., Kieffer, S., et al. (2001) The phagosome proteome: insight into phagosome functions. J. Cell. Biol. 152, 165–180. 8. Desjardins, M. (1995) Biogenesis of phagolysosomes: the ‘kiss and run’ hypothesis. Trends Cell. Biol. 5, 183–186. 9. Griffiths, G. (2004) On phagosome individuality and membrane signalling networks. Trends Cell. Biol. 14, 343–351. 10. Henry, R. M., Hoppe, A. D., Joshi, N., and Swanson, J. A. (2004) The uniformity of phagosome maturation in macrophages. J. Cell Biol. 164, 185–194.

21 Monitoring Time-Dependent Maturation Changes in Purified Phagosomes from Dictyostelium discoideum Régis Dieckmann, Navin Gopaldass, Caroline Escalera, and Thierry Soldati

Summary The amoeba Dictyostelium discoideum is an established model to study phagocytosis. The sequence of events leading to the internalization and degradation of a particle is conserved in D. discoideum compared to metazoan cells. As its small haploid genome has been sequenced, it is now amenable to genome-wide analysis including organelle proteomics. Therefore, we adapted to Dictyostelium the classical protocol to purify phagosomes formed by ingestion of latex beads particles. The pulse-chase protocol detailed here gives easy access to pure, intact, and synchronized phagosomes from representative stages of the entire process of phagosome maturation. Recently, this protocol was used to generate individual temporal profiles of proteins and lipids during phagosome maturation generating a proteomic fingerprint of six maturation stages (1). In addition, immunolabeling of phagosomes on a coverslip was developed to visualize and quantitate antigen distribution at the level of individual phagosomes.

Key Words: Phagosomes, organelle purification, phagocytosis, Dictyostelium.

1. Introduction The social amoeba Dictyostelium discoideum is a recognized eukaryotic model for professional immune phagocytes (2). The range of ingested particles includes latex beads, live and dead cells of the same species, yeast, and bacteria. It is used to study host–pathogen interactions with bacteria such as Legionella pneumophilia (3), Pseudomonas aeruginosa (4), and Mycobacterium marinum (5,6). The sequence of events leading to the internalization From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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and degradation of a particle is conserved in D. discoideum compared to metazoan. This includes particle recognition and signaling, recruitment of the actin cytoskeleton leading to the formation of pseudopods, and the intense vesicular trafficking involving the delivery of digestive enzymes and membrane retrieval (2,7). Two peculiarities of D. discoideum are the difficulty to identify phagocytic receptors (8–10) and constitutive exocytosis of undigested remnants. However, phagosome/endosome exocytosis is also used by dendritic cells to present antigens on their surface. Additionally, exocytosis of latex beads can be triggered by appropriate secretagogues in macrophages (11). Compared to macrophage-like cell lines, the use of D. discoideum as a phagocytic model has several advantages. The rate of phagocytosis is up to 20-fold higher. The time scale of phagosome maturation from uptake to exocytosis does not exceed 3 h. Its small haploid genome (33 Mbp, 12,000 predicted genes) is now sequenced and thoroughly annotated, allowing rapid targeted mutagenesis and large-scale analysis by random insertion of plasmid sequences, microarrays, and proteomics (1,8,12). The protocol presented here has been used notably to generate individual temporal profiles of proteins and lipids during phagosome maturation (1, Brügger et al., in preparation). The method is based on the purification of low-density latex bead–containing phagosomes via their flotation in sucrose step gradients. The method was originally introduced by Wetzel and Korn for Acanthamoeba and adapted successfully to macrophages by Desjardins and collaborators (13,14). The latter protocol has been further refined and adapted into a pulse-chase protocol that covers the entire maturation process. A large number of pure, intact, and synchronized phagosomes are isolated at six different times spanning the entire maturation process of 3 h. Synchronicity is ensured by a preincubation in the cold of cells and beads at high concentration. Cells are broken with a ball homogenizer for optimal breakage while keeping the phagosomal membrane intact. Phagosome purity is significantly improved by a brief incubation of the cell lysate with a physiological concentration of ATP, which releases the rigor mortis actin–myosin interactions, thereby avoiding co-purification of enmeshed organelles (15). This protocol has been fine-tuned to yield equal amounts of phagosomes at each time point. As it has been shown that phagocytosis was sensitive to particle size (16), shape (17), and nature (18), small adjustments were introduced to isolate phagosome formed with different bead sizes (0.5, 0.8, 3.0 μm). The quantitation of the number of purified phagosomes is precisely performed by measuring light scattering in the fraction collected from the sucrose gradient. An average of 4 × 1010 phagosomes per time point can be isolated. Phagosome concentration is normalized according to absorbance at 600 nm (15). Protein quantitation indicates that the total amount of protein

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per phagosome does not vary significantly throughout maturation (less than 15%) and is approximately 12 μg/109 phagosomes (≈450 μg total) (1). This is further demonstrated by the equal loading of total proteins on 1D gels after normalization (1,7,15). 2. Materials 2.1. Buffers and Equipment 1. Soerensen buffer (SB): 15 mM KH2 PO4 , 2 mM Na2 HPO4 , pH 6.0. 2. Soerensen/Sorbitol buffer (SSB): Soerensen buffer containing 120 mM sorbitol. 3. HESES: 20 mM hydroxyethyl piperazine ethane sulfonate (HEPES)-KOH, pH 7.2, 0.25 M sucrose. 4. Homogenization buffer (HB): HESES, 2X Complete ethylenediamine tetraacetic acid (EDTA)-free (protease inhibitor cocktail; Roche, Hertfordshire, UK). 5. Membrane buffer: 20 mM HEPES-KOH, pH 7.2, 20 mM KCl, 2.5 mM MgCl2 , 1 mM dithiothreitol (DTT), 20 mM NaCl. 6. Storage buffer: 25 mM HEPES-KOH, pH 7.2, 1.5 mM Mg-acetate, 1 mM NaHCO3 , 1 μM CaCl2 , 25 mM KCl, 1 mM ATP, 1 mM DTT, 1X Complete EDTA-free, 100 mM sucrose. 7. Ball homogenizer (Isobiotec, Heidelberg, Germany, barrel 8.000 mm, ball diameter 7.990 mm, resulting in an annular void clearance of 5 μm). 8. Piperazine-N,N´–bis(2-ethanesulfonate) (PIPES) buffer: 20 mM PIPES, pH 6.0. 9. Saturated picric acid solution. Dissolve 3 g of solid picric acid in 1 L of doubledistilled water, warm up to 80°C overnight. Cool down to room temperature and adjust the pH to 6.0. Store at 4°C. 10. Picric acid/PFA fixative. Mix 0.4 g of paraformaldehyde, 7 mL of double distilled water, 10 mL of PIPES buffer. Microwave in brief pulses until it dissolves, then cool immediately to room temperature (on ice). Add 3 mL of a saturated picric acid solution. 11. Phosphate-buffered saline (PBS) buffer: 140 mM NaCl, 2.7 mM KCl, 10 mM Na2 HPO4 , 1.8 mM KH2 PO4 , pH 7.4.

2.2. Cell Culture Dictyostelium discoideum cells of wild-type strain Ax2 are grown axenically in HL5c medium (ForMedium Ltd, Norwich, UK) in shaking culture (at 180 rpm) at 22°C to a density of 5 × 106 cells/mL. 2.3. Preparation of Latex Beads Always prepare fresh. 2 × 2 mL of 0.8-μm latex beads suspension (Sigma, St. Louis, MO) are spun down in an Eppendorf tube (10,000g, 5 min). The beads are then washed twice in SSB, pH 8, to remove the detergent and sodium azide

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contained as preservatives in the suspension supplied by the manufacturer, and finally resuspended in 2 mL of SSB, pH 8, and kept on ice. This appears critical, as insufficient washing blocks phagocytosis. Before use, the bead suspension is sonicated for 5 min in a bath sonicator. For use of other bead sizes see Note 1. 2.4. Preparation of Sucrose Step Gradients Prepare sucrose step gradients in disposable centrifuge tubes (polyallomer tubes 25 × 89 mm, Beckman) by layering the following sucrose solutions on top of each other: 4 mL of 60%, 12 mL of 35%, 12 mL of 25% sucrose in 20 mM HEPES-KOH pH 7.2. The gradients can be stored up to overnight avoiding vibrations. The last layer composed of 4 mL of 10% sucrose in 20 mM HEPES-KOH pH 7.2 is added after the phagosome samples have been loaded to obtain an undisturbed 25%/10% interface and to balance the tubes exactly. 2.5. Preparation of Poly-l-Lysine–Coated Cover Slips Coverslips (12 mm diameter, No. 1, Assistant, Germany) are coated with a 1 mg/mL poly-l-lysine hydrobromide (MW (vis) 93,800, Sigma) solution for 1 h. The coverslips are then dipped once in double-distilled water and left to dry. 3. Methods 3.1. Phagosome Isolation After Pulse-Chase Feeding of Latex Beads 1. Cells from an overnight culture are counted with a hemocytometer, and 8 × 109 cells are centrifuged at 500g for 5 min. Subsequent steps are performed at 4°C or on ice when possible, except indicated otherwise. 2. Resuspend the cells in 50 mL SSB pH 8 and pool into a 50-mL Falcon tube. 3. Wash cells by spinning them down at 500g for 5 min. 4. Gently resuspend cell pellet in 20 mL of ice cold SSB pH 8.0. If desired, biotinylate the cell surface (for steps 4-6, see Note 2). 5. Add the bead suspension to the cells. Mix by inverting the tube. 6. Incubate for 15 min on ice. 7. Our typical setup is based on six time points (see Note 3): • • • • • •

P1: P2: P3: P4: P5: P6:

5 min pulse 15 min pulse 15 min pulse/15 min chase 15 min pulse/45 min chase 15 min pulse/1 h 45 min chase 15 min pulse/2 h 45 min chase


8. 9.

10. 11. 12.

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Pour the bead/cell mixture into 100 mL of HL5c kept at 22°C in a 250-mL flask. Starting with the latest time point P6, pipet 5 mL of the cell/beads suspension for P6 to P3, 6 mL for P2, and 7 mL for P1 (see Fig. 1 and Note 4). Incubate at 22°C while shaking at 120 rpm for the appropriate period of time. At the chosen time point, in order to stop phagocytosis, pour the 100-mL sample into 330 mL of ice-cold SSB prealiquoted in centrifugation bottles (500 mL, 69 × 160 mm) kept on ice (see Note 5). Centrifuge cells for 8 min at 2000 g (Beckman rotor JLA-10.500). Resuspend the cell pellet in 50 mL of ice-cold HESES and centrifuge again for 5 min at 500g. For time point P3 to P6, resuspend the cells in 5 mL of ice cold HL5c and pour into 100 mL of HL5c kept at 22°C. Incubate at 22°C while shaking at 120 g for the appropriate chase period. Proceed then from step 9 to step 11 to stop phagocytosis. Following step 11, repeat the HESES washing step twice for all time points to thoroughly wash away uningested latex beads. Repeat the HESES wash once more if 3-μm beads are used.

Fig. 1. Effect of beads adsorption on phagocytic uptake as measured by FACS. Cells were either preincubated (black curve) or not (grey curve) at 4°C with fluorescent beads (carboxylated Fluoresbrite YG microspheres, 1.0 μm diameter, Polysciences Inc.) for 15 min before starting the uptake at 22°C. Preadsorption of the latex beads onto cells in the cold enables an efficient and synchronous uptake during the pulse period, resulting in a more homogeneous population of isolated phagosomes. In comparison, the initial uptake without preincubation is reduced and more than 20 min are needed to get the same number of beads inside the cells as with preincubation in the cold. Uptake was stopped by plunging the bead/cell suspension in an azide-containing buffer. As cells round up and contract due to azide treatment, subsequent centrifugation detaches uningested beads from the cell surface (1). Therefore, the only beads monitored by FACS have been internalized.

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14. Resuspend the cell pellet (approximately 1 mL) in 2 mL of HB-containing a protease-inhibitor cocktail (Complete EDTA-free, Roche) at 2X concentration, resulting in a 1X final concentration. 15. Homogenize cells by eight passages through a ball homogenizer. (see Fig. 2 and Note 6). 16. Adjust the final concentrations of ATP to 10 mM, of MgCl2 to 10 mM and of sucrose to about 45–50% from freshly made stocks (0.1 M ATP in 40% sucrose, 20 mM HEPES-KOH buffered to pH 7.2; 1 M MgCl2 ; 71.4% sucrose in 20 mM HEPES-KOH pH 7.2) (see Note 7). 17. Mix gently for 15 min using an overhead tumbler or a wheel. 18. Load the density-adjusted homogenate between the 60% and 35% layer of the sucrose step gradients using a syringe and a needle (100 mm × 1.5 mm), and overlay with 4 mL of 10% sucrose solution (for steps 18 and 19, see Note 8). 19. Centrifuge gradients at 28,000 g (Beckman rotor SW 28, 100,000g avg), 4°C for 2 h 30 min. 20. Collect the interphase between 10% and 25% with a Pasteur pipet and dilute with membrane buffer to a final volume of 15 mL, mix by inverting the tube (for steps 20–23, see Note 9). 21. Take a 50-μL aliquot to measure scattering at 600 nm to calculate the number of phagosomes in the sample. A standard curve can be made with serial dilutions

Fig. 2. Optimal cell breakage and effect of ATP incubation. Phase contrast pictures of cells before (A) and after (B, C, D upper panel) six to eight passages through a ball homogenizer. The cell suspension and homogenate were dropped on a glass slide and covered with a cover slip. Pictures were taken with an Axiophot 2 and a 20x objective. Optimal cell breakage is obtained when 95% of the cells have been lysed and have lost their refractile appearance (compare A to B/C). The contrast-inverted picture reveals the presence of beads inside the cells (A, small panel). ATP treatment in the cold releases phagosomes from enmeshed organelles. Aggregates present in B are absent from C (arrows). The intactness of the nuclei (arrowheads) is also essential to avoid aggregation of the organelles around free chromatin. As illustrated here by DAPI staining of a homogenate (D), ball homogenization preserves the nuclei from breakage.


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of the beads, generating a linear correlation between scattering and bead concentration in a range between 2 × 107 and 2 × 108 beads/mL (15). 22. The 14.95 mL of phagosome suspension are further diluted with membrane buffer up to 37 mL and pelleted in the SW 28 again for 50 min at 28,000 g (100,000g avg). 23. Resuspend the pellet in storage buffer. Adjust the amount of buffer according to the scattering measurements so as to reach the same concentration of phagosomes in all samples. 24. Snap-freeze aliquots in liquid nitrogen and store at –80°C until further use. Samples are routinely analysed by SDS-PAGE (see Fig. 3).

3.2. Immunofluorescence on Purified Phagosomes Purified phagosomes are taken from step 21. The amount of phagosomes to be used per coverslip is determined as follow: 300/OD600 = n, where n is the volume in μL of phagosome suspension per cover slip (see step 21 for OD600 measurements). Adjust the volume up to 0.5 mL per cover slip with HESES buffer. Place the coverslips in a 24-well plate and add 0.5 mL of the phagosome suspension per well. Centrifuge 7 min at 1500 g in a clinical centrifuge with multiwell plate swinging buckets. Wash the poly-L-lysine–coated coverslips

Fig. 3. Characterization of the phagosomal fractions. The fractions obtained after a pulse-chase feeding experiment with latex beads of, respectively, 0.5, 0.8, and 3.0 μm in diameter have been separated on a 10% SDS-PAGE. The gels containing the 0.5- and 3.0-μm samples are Sypro Ruby stained (BioRad, according to the manufacturer). The gel containing the 0.8-μm samples is silver stained (GE Healthcare, according to the manufacturer). Total membrane extracts (ME) and/or total cell extract at different dilutions (1 [0.5 × 106 cells/lane], ½, ¼) have been loaded beside the phagososme fractions. The maturation process can be monitored through the appearance/disappearance (open/closed arrows) of bands. Comparison with total cell lysate, total membrane extract shows the unique band pattern of the phagosome fractions. Differences in composition and temporal profiles (asterisks) are visible between phagosome samples obtained by ingestion of beads of different sizes.

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by dipping them once into SSB and place them back in a 24-well plate. The phagosomes are fixed for 30 min at room temperature with a PFA/picric acid solution, washed once with PBS and placed for 30 min with 100 mM glycine in PBS in order to quench the free aldehyde groups, and blocked for 1 h in 2% FCS in PBS. The coverslips are then processed as for standard immunofluorescence (see Fig. 4). Alternatively, a rapid freezing fixation procedure in methanol at –85°C can be used (19).

Fig. 4. Visualization of membrane trafficking at the level of single phagosomes. Plasma membrane proteins were biotinylated prior to latex bead uptake and phagosome purification (see Note 1). Phagosomes isolated at the indicated time points of a pulse/chase feeding experiment were centrifuged on poly-l-lysine–coated coverslips and processed for immunofluorescence. Biotinylated membrane proteins were revealed by streptavidinTRITC (upper row) and PM4C4 recognized by a monoclonal antibody (20) followed by a goat–anti-mouse Alexa488-coupled IgG. Shortly after bead ingestion (5’ and 15’), the vast majority of phagosomes contain biotinylated plasma membrane proteins, but during maturation (from 15’/15’ to 15’/165’), while this marker is retrieved to the surface, it is replaced by nonbiotinylated PM4C4 from later endosomes. Interestingly, the transition from one to the other marker appears not to be gradual but abrupt, reflecting the very intense trafficking to and from maturing phagosomes (1). Arrowhead: a patch of PM4C4 on a biotin-positive phagosome; circle: a biotin-positive, PM4C4-negative phagosome; triangle: an unstained bead; square: a PM4C4-positive, biotin-negative phagosome. Images were taken with a Leica SP2 confocal microscope. The fluorescence and phase signals were superimposed to allow easier visualization of all beads.


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4. Notes 1. For phagocytosis of different bead sizes, we used approximately the same bead surface–to–cell surface ratio (360 μm2 of bead/cell). But to compensate for the low uptake of 3.0-μm beads, we decided to double the offered bead surface. Therefore, the calculated volumes of the supplied bead suspensions are 2.5 mL of 0.5-μm beads (14.62 × 1011 beads/mL, LB-5, Sigma), 4 mL of 0.8-μm beads (3.57 × 1011 beads/mL, LB-8, Sigma), 10 mL of 3.0-μm beads (0.20 × 1011 beads/mL, LB-30, Sigma). As bead size increases, the offered bead surface does not increase linearly. Therefore, the used bead to cell ratios are, respectively, 460 beads/cell (0.5 μm), 180 beads/cell (0.8 μm), and 25 beads/cell (3.0 μm). 2. The preincubation step is performed to maximize the number of latex beads adsorbed onto the cells. If desired, it is also possible to concomitantly biotinylate plasma membrane proteins (7), using 30 mg of Imuno-Pure NHS-LC-Biotin from Pierce (Rockford, IL). The high pH is thus necessary for efficient biotinylation, but the cells should not be kept for extended periods of time at this pH. 3. This pulse-chase time course has been shown to cover the entire phagosome maturation process (1,7). The time points of 5 and 15 min are extremely dynamic and cover the first signaling, cytoskeleton, and membrane trafficking phases. The 5-min sample is processed and homogenized before the 15-min chase is stopped, whereas the other four pellets are processed for the chase period as described in step 12. SDS-PAGE and Western blot analysis show the enrichment of the fractions in phagosomal markers and highlight the progression of the maturation process and the unique identity of phagosomes at each stage (see Fig. 3). 4. To start phagocytosis, a 20-fold excess (100 mL) of HL5c at 22°C is added to the sample, instantaneously raising the temperature and thus generating a sharp synchronous wave of uptake (see Fig. 1). The uneven distribution of the cell/beads suspension (5, 6, or 7 mL) ensures an equal yield for all the time points. 5. To stop the phagocytic process, the 100-mL cell suspension is poured into 3.3 volumes of ice-cold SSB and immediately centrifuged. The use of 120 mM sorbitol is crucial to increase bead flotation and nevertheless not perturb cell pelleting. These cycles of resuspension/centrifugation are sufficient to eliminate the vast majority of uningested beads. 6. Homogenization using a ball homogenizer results in homogeneous cell breakage together with preservation of organelle integrity. The osmolarity of the buffer, the void clearance, and the number of passages is optimized to yield about 95% cell breakage (see Fig. 2). Higher ratios of cell breakage only result in increased nuclear lysis and contamination of cytoplasm with chromatin. 7. Use of a physiological concentration of ATP for a few minutes in the cold avoids artefactual formation of a rigor mortis meshwork of actin and myosins that was shown to entrap contaminating organelles (7). Omission of ATP has visible consequences as it prevents fast and clean flotation of the phagosomes and promotes co-fractionation of an actin–myosin II meshwork (as judged by Coomassie staining of the resulting fractions after SDS-PAGE). This can only partially be compensated by longer centrifugation.

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8. If the centrifugation time is reduced to below 2 h 30 min, banding is not as sharp, and phagosome yield is about 20–30% lower. The rotor has to stop without brake to avoid vibrations. Alternatively, centrifugation can be performed overnight. 9. This dilution is necessary to decrease the sucrose concentration and thus decrease the buoyancy of the latex beads phagosomes. We determined that scattering was caused only by the latex beads and not contaminating particles and/or organelles by treating the latex beads phagosome sample with SDS and pelleting. The resulting “clean” latex beads yielded the same scattering values as when residing inside phagosomes.

Acknowledgments This work was mainly carried out at the University of Geneva and was supported by a grant from the Swiss National Science Foundation. The TS research group participates in the “Non-mammalian Experimental MOdels for the study of bacterial infections (NEMO)” network, supported by the 3R Foundation. A big “thank you” goes to all the lab members who have participated in the establishment of the protocols and have suggested amendments and improvements.

References 1. Gotthardt, D., Blancheteau, V., Bosserhoff, A., Ruppert, T., Delorenzi, M., and Soldati, T. (2006) Proteomic fingerprinting of phagosome maturation and evidence for the role of a Galpha during uptake. Mol. Cell Proteomics, 5, 2228–2243. 2. Maniak, M. (2002) Conserved features of endocytosis in Dictyostelium. Int. Rev. Cytol. 221, 257–287. 3. Hagele, S., Kohler, R., Merkert, H., Schleicher, M., Hacker, J. and Steinert, M. (2000) Dictyostelium discoideum: a new host model system for intracellular pathogens of the genus Legionella. Cell. Microbiol. 2, 165–171. 4. Pukatzki, S., Kessin, R. H., and Mekalanos, J. J. (2002) The human pathogen Pseudomonas aeruginosa utilizes conserved virulence pathways to infect the social amoeba Dictyostelium discoideum. Proc. Natl. Acad. Sci. USA 99, 3159–3164. 5. Solomon, J. M., Leung, G. S., and Isberg, R. R. (2003) Intracellular replication of Mycobacterium marinum within Dictyostelium discoideum: efficient replication in the absence of host coronin. Infect. Immun., 71, 3578–3586. 6. Hagedorn, M., and Soldati, T. (2007) Flotillin and the RacH GTPase modulate intracellular immunity of Dictyostelium to Mycobacterium marinum infection. Cell Microbiol. 9, 2716–2733. 7. Gotthardt, D., Warnatz, H. J., Henschel, O., Bruckert, F., Schleicher, M., and Soldati, T. (2002) High-resolution dissection of phagosome maturation reveals distinct membrane trafficking phases. Mol. Biol. Cell 13, 3508–3520.


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8. Cornillon, S., Pech, E., Benghezal, M., et al. (2000) Phg1p is a nine-transmembrane protein superfamily member involved in Dictyostelium adhesion and phagocytosis. J. Biol. Chem. 275, 34287–34292. 9. Cornillon, S., Gebbie, L., Benghezal, M., et al. (2006) An adhesion molecule in free-living Dictyostelium amoebae with integrin beta features. EMBO Rep. 7, 617–621. 10. Fey, P., Stephens, S., Titus, M. A., and Chisholm, R. L. (2002) SadA, a novel adhesion receptor in Dictyostelium. J. Cell Biol. 159, 1109–1119. 11. Di, A., Krupa, B., Bindokas, et al. (2002) Quantal release of free radicals during exocytosis of phagosomes. Nat. Cell Biol. 4, 279–285. 12. Van Driessche, N., Demsar, J., Booth, E.O., et al. (2005) Epistasis analysis with global transcriptional phenotypes. Nat. Genet. 37, 471–477. 13. Wetzel, M. G., and Korn, E. D. (1969) Phagocytosis of latex beads by Acanthamoeba castellanii (Neff). 3. Isolation of the phagocytic vesicles and their membranes. J. Cell Biol. 43, 90–104. 14. Desjardins, M., Huber, L. A., Parton, R. G., and Griffiths, G. (1994) Biogenesis of phagolysosomes proceeds through a sequential series of interactions with the endocytic apparatus. J. Cell Biol. 124, 677–688. 15. Gotthardt, D., Dieckmann, R., Blancheteau, V., Kistler, C., Reichardt, F., Soldati, T. (2006) Preparation of intact, highly purified phagosomes from Dictyostelium. Methods Mol. Biol. 346, 439–448. 16. Lefkir, Y., Malbouyres, M., Gotthardt, D., et al. (2004) Involvement of the AP-1 adaptor complex in early steps of phagocytosis and macropinocytosis. Mol. Biol. Cell 15, 861–869. 17. Champion, J. A. and Mitragotri, S. (2006) Role of target geometry in phagocytosis. PNAS 103, 4930–4934. 18. Bozzaro, S. (1985) Cell surface carbohydrates and cell recognition in Dictyostelium. Cell Differ. 17, 67–82. 19. Hagedorn, M., Neuhaus, E. M., and Soldati, T. (2006) Optimized fixation and immunofluorescence staining methods for Dictyostelium cells. Methods Mol. Biol. 346, 327–338. 20. Neuhaus, E. M., and Soldati, T. (2000) A myosin I is involved in membrane recycling from early endosomes. J. Cell Biol. 150, 1013–1026.

22 Large-Scale Purification of Latex Bead Phagosomes from Mouse Macrophage Cell Lines and Subsequent Preparation for High-Throughput Quantitative Proteomics Adam Rupper and James Cardelli

Summary Phagocytosis involves the engagement of a diverse array of cell surface receptors whose signals must be integrated on the membrane of the forming phagosomal cup. This method enables the quantitative proteomic analysis of phagosome fractions derived from phagocytes stimulated under two different conditions, thus allowing the complexity of phagosomal signaling to be analyzed in terms of the quantitative changes in phagosomal fraction protein content.

Key Words: Phagosome purification; macrophage; isotope coded affinity tag; liquid chromatography tandem mass spectrometry; sucrose gradient; RAW 264.7. 1. Introduction The process of phagocytosis enables immune cells to interrogate the environment around them. This interrogation involves a plethora of cell surface pattern recognition receptors, opsonin receptors, and receptors that recognize self ligands such as those presented by apoptotic cells (1). The integration of these signals occurs on the phagosomal membrane and leads to molecular decisions that determine whether inflammatory or anti-inflamatory mediators will be released, whether antigen presentation will be performed, and what path phagosomal maturation will take (2,3). The protocol described here began as a method to purify latex bead– containing phagosomes on sucrose step gradients using the density of latex as a From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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way to float phagosomal membranes away from other cellular membranes (4). Proteomic analysis has been used to identify the protein content of latex bead– isolated phagosome fractions and provides the standard from which to measure new techniques (5). With the advent of quantitative proteomic techniques, such as the use of cleavable ICAT (isotope coded affinity tag) reagent, the relative abundance of two sister peptides labeled differentially with heavy and light markers (identified by the collision dissociated peptide fingerprint) can be determined from the original ions’ peak intensities which differ by 9 mass units (6,7). In this way the relative abundance of many peptides from a complex mixture of two samples can be determined. The ability to determine how different activation states of the macrophage affect the protein makeup of the phagosome has become a reality. The protocol to purify latex bead–containing phagosomes from mouse macrophage cell lines was adapted in order to maximize the yield of phagosomal protein from one purification experiment (though multiple repetitions are required to yield one mg of total protein) and enable high-throughput proteomic techniques (which are best performed with mg quantities of protein) to be used as a method to ask questions about how different opsonins, pathogenassociated patterns, or inflammatory mediators might change the protein content of the phagosome. It consists of four major steps: the presentation of IgG opsonized latex beads to macrophages in an efficient and large-scale procedure, the purification of the latex bead phagosomes by isopycnic centrifugation with sucrose step gradients, labeling of the proteins found in the purified fraction with cleavable ICAT reagent, and separation of labeled peptides from this complex mixture by strong cation exchange chromatography and avidin affinity chromatography in order to reduce the complexity of each fraction. Due to space limitations and the complexity of this process, the method for determination of the relative abundance and identity of sample peptides by LC-MS/MS and the use of software tools designed to enable high-throughput analysis of the data with statistical confidence will not be covered in this protocol. The end result will be purified peptide fractions labeled with heavy and light ICAT reagent that can be analyzed by a variety of mass spectrometry platforms. References will be given for those who are interested in the specific details of the platform used by the authors. 2. Materials 2.1. Latex Bead Preparation 1. Nonpyrogenic Eppendorf tubes. 2. Latex beads (0.82 μm, 5081A), (Duke Scientific Corporation, Palo Alto, CA). 3. Mouse monoclonal anti-Ova (A 6075 Sigma Aldrich, St. Louis, MO).

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4. 5X crystalline chicken ovalbumin (32467 Calbiochem) other highly purified sources acceptable. 5. Bead wash buffer (BWB), 25 mM 2-(N-morpholino) ethanesulfonic acid (MES) pH 6.1, 0.05% Triton X 100 (TX100). 6. Ova binding buffer (OBB), 25 mM MES pH 6.1, 0.1 % w/v ovalbumin, 0.05% TX100, 0.05% NaN3 ; filter sterilize with a 0.2-μm low-protein-binding filter; can be stored at 4°C for multiple months. 7. Final resuspension buffer, 25 mM MES pH 6.1. 8. Limulus amoebocyte assay (50-647U, Cambrex). 9. High-speed desktop centrifuge for 1.5-mL Eppendorf tubes. 10. Rotator or shaker in a refrigerator.

2.2. RAW 264.7 Cell Preparation and Bead Presentation 1. 125- or 250-mL Erlenmeyer flasks heat treated at 180°C for 4 h to destroy endotoxin. 2. RPMI 1640 with l-glutamine (Mediatech, Inc., Herndon, VA) supplemented with 10% fetal bovine serum (FBS, Hyclone, Ogden, UT) and penicillin/ streptomycin (full medium). 3. Full medium with 25 mM hydroxyethyl piperazine ethane sulfonate (HEPES) (from 1 M HEPES buffer stock). 4. RAW 264.7 cells (ATCC). 5. 15-cm Petri dishes, non-tissue culture treated, sterile, nonpyrogenic. 7. 15-, 50-, and 250-mL conical cell culture centrifuge tubes. 6. Dulbecco’s phosphate-buffered saline without calcium and magnesium (DPBS), Cellgro.com. 7. DPBS, 1.5 mM ethylenediaminetetraacetic acid (EDTA), filter sterilize. 8. DPBS, 120 mM sorbitol, filter sterilize. 9. Refrigerated benchtop centrifuge for cell culture applications. 10. 37°C shaking incubator.

2.3. Cell Disruption 1. Nitrogen decompression bomb (4639, Parr Instrument Company). 2. N2 gas tank. 3. Homogenization buffer (HB) 20 mM HEPES pH 7.4 (use KOH to titrate pH), 0.25 M sucrose, filter sterilize, and store at 4°C. 3. Homogenization buffer with protease inhibitors and Ca (HB + PI + Ca) 20 mM HEPES pH 7.4, 0.25 M sucrose, 1x protease inhibitors cocktail (Complete Mini EDTA free, 11 836 170 001, Roche); dissolve one tablet in 10 mL of buffer, 0.2 mM CaCl2 , add protease inhibitors on day of use, and store on ice or at 4°C. 4. 15% sucrose w/v in 20 mM HEPES pH 7.4; see Subheading 2.4. for instructions on sucrose solutions.

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2.4. Phagosome Purification by Sucrose Step Gradient 1. 2. 3. 4. 5.

6. 7. 8.

Beckman Ultra-Clear 1 × 3.5 in centrifuge tubes (344058). Sucrose, molecular biology grade. 20 mM HEPES pH 7.4 (use KOH to titrate pH), filter sterilize and store at 4°C. 62% sucrose w/v, 20 mM HEPES pH 7.4 (use KOH to titrate pH), filter sterilize and store at 4°C. Using the 62% sucrose and 20 mM HEPES solutions above, make enough 40, 35, 25, 15, and 10% sucrose solutions to perform the experiment and equilibrate to 4°C. Beckman L8-55M ultracentrifuge. Beckman SW-28 swinging bucket rotor. ICAT labeling buffer: 200 mM Tris-HCl pH 8.3, 6 M urea, 5 mM EDTA, 0.1% SDS; make fresh on day of use.

2.5. Assessment of Phagosome Purity 1. Antibodies: anti-Lamp1 (1D4B rat monoclonal antibody (mAB), University of Iowa) anti-rab5 (BD, mouse mAB, 610282), p62 nuclear porin (mAB414, mouse mAB, Covance), GM130 (BD, 610822, mouse mAB), anti-mouse IgG horseradish peroxidase (HRP, Amersham), anti-rat IgG HRP (Pierce). 2. Mini gel sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) apparatus and electroblotting apparatus. 3. 4x SDS-PAGE loading buffer: 0.25M Tris-HCl pH 6.8, 8% SDS, 30% glycerol, 0.02% bromophenol blue; add 10% -mercaptoethanol before use. 4. Acrylamide: Bis 37.5:1. 5. Ammonium persulfate, make a fresh10% w/v solution in ddH2 O. 6. Upper-Tris buffer: 0.5 M Tris-HCl pH 6.8, 0.3% SDS. 7. Lower-Tris buffer: 1.5 M Tris-HCl pH 8.8, 0.4% SDS. 8. Towbin buffer: 25 mM Tris, 192 mM glycine, 20% v/v methanol. 9. Methanol. 10. Polyvinylidene fluoride transfer membrane (PVDF). 11. 3mm filter paper. 12. Tris-buffered saline with Tween 20 (TBS-T): 10 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.05% Tween 20 w/v. 13. TBS-T with 5% nonfat dry milk w/v. 14. TBS-T with 5% bovine serum albumin (BSA) w/v. 15. ECL plus detection reagent (Amersham). 16. Kodak Biomax XAR film. 17. Silver Stain Plus kit (BioRad).

2.6. Protein Labeling with Isotope Coded Affinity Tag 1. BCA protein assay (Pierce). 2. Tris(2-carboxyethyl)phosphine (TCEP).

Phagosome Purification 3. 4. 5. 6. 7.

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Cleavable ICAT reagent (Applied Biosystems). Dithiothreitol (DTT). 50 mM Tris-HCl, pH 8.3. Sequencing Grade Modified Trypsin (V5113, Promega). 0.2-μm low-protein-binding syringe filter.

2.7. Cation Exchange Chromatography and Avidin Affinity Chromatography 1. High-performance liquid chromatography (HPLC) system with ability to make linear gradients and track UV absorbance at 214 nm. 2. Two-mL sample loop for HPLC system. 3. Polysulfoethyl A column, 2.1 mm × 200 mm, 5-μm particles, 300 Å pore size (PolyLC, Inc.). 4. Strong cation exchange buffer A (SCX buffer A), 25% acetonitrile (volatile, flammable, hazardous), 5 mM KH2 PO4 , pH 3.0. 5. SCX buffer B, 25% acetonitrile, 5 mM KH2 PO4 , pH 3.0, 350 mM KCl. 6. 5% Phosphoric acid. 7. 100 mM Phosphate buffer pH 10.0. 8. 2x PBS pH 7.2. 9. ICAT avidin affinity buffer pack (4326740, Applied Biosystems) good for 50 purifications. 10. One-mL glass Hamilton syringe with a blunt needle.

3. Methods 3.1. Latex Bead Preparation 1. Prepare enough latex beads for a 1:50 dilution with the resuspended RAW 264.7 cells. (see Subheading 3.2. to have RAW 264.7 cells prepared for the day of the experiment). 2. Add 750 μL of beads/1.5 mL Eppendorf tube. Centrifuge at 10,000g for 2 min, remove supernatant and resuspend with 1 mL of BWB, repeat centrifugation step, remove supernatant, and resuspend with OBB and rock or rotate overnight at 4°C (shorter incubation times are not sufficient). Alhough inclusion of surfactant in the buffers helps to reduce bead compaction upon centrifugation, rapid resuspension of the beads is best performed by digging with the pipet tip while pipetting up and down. 3. Wash two times with BWB with centrifuge at maximum g force for 5 min. 4. Resuspend the beads in BWB with a 1:1000 dilution of anti-ova antibody and rock or rotate at room temperature for 30 min (see Notes 1 and 2). 5. Wash once with BWB and resuspend with 25 mM MES pH 6.1. Perform final wash after the cells are prepared for incubation with beads and equilibrated to 37°C in a shaking incubator.

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3.2. RAW 264.7 Cell Preparation and Bead Presentation 1. Expand RAW 264.7 cells to yield the number of cells required for the experiment. Use 5-6 × 108 cells/sucrose gradient. To obtain this number of cells requires 10 15-cm Petri dishes grown to high density (see Note 3). An SW-28 rotor has six buckets, and thus six gradients can be run per rotor, this requires 60 confluent 15-cm dishes be ready on the day of the experiment. We seed the cells from confluent, healthy (not a lot of floating cells) plates at 1:5 into 12 plates and allow them to grow for 3 d, then expand 1:5 into 60 plates and allow the cells to grow for 2 d prior to the experiment, thus optimizing the density and health of the cells. 2. Remove growth medium and wash the cells gently with 10 mL of DPBS. Remove DPBS and add 10 mL DPBS with 1.5 mM EDTA. Allow the cells to incubate at room temperature while all plates are washed and filled with DPBS, 1.5 mM EDTA. Remove cells by vigorously pipetting up and down with an electric pipet aid. 3. Collect the cells by centrifugation at 250g (average) for 10 min and resuspend once in DPBS. Collect the cells by centrifugation again and resuspend in prewarmed (37°C) full growth medium with 25 mM HEPES at 1 × 107 cells/mL. Place cells in a sterile, heat treated 250-mL Erlenmeyer flask, shake at 250 rpm in a 37° C incubator. 4. Present prepared beads as a 1:50 dilution (1mL of prepared beads for 50 mL of prepared cells) into the suspended cells and replace flask into the 37°C shaking incubator. Incubate with shaking for the desired period of time (see Note 4).

3.3. Cell Disruption 1. Collect cells by centrifugation at 250g and wash with cold (4°C) DPBS with 120 mM sorbitol. The sorbitol increases the density of the solution and helps to remove uninternalized beads, but is not necessary. Repeat until supernatant appears clear, usually two to three washes. 2. Wash one time with HB. Resuspend half of the cells in 15 mL HB + PI + Ca and keep on ice. Repeat for the other half. We often performed the experiment where 30 plates were treated with interferon (IFN)- for 18 h and the others were unstimulated. If all 60 plates are used as a control or experimental, then break the cells in two separate 15-mL aliquots. Conditions can be worked out for different volumes and numbers of cells. 3. Load resuspended cells into a prechilled cell disruption bomb. Follow the manufacturer’s procedures and pressurize the bomb with N2 gas to 400 psi (see Note 5). Allow the bomb to incubate on ice for 10 min. Slowly release the pressurized contents of the bomb by turning the release valve counterclockwise until the cell lysate begins to run through the release tube into a 50-mL conical tube. Carefully, watch the released volume as pressurized gas will follow the last contents of the bomb and can blow the cell lysate out of the 50-mL conical tube. This can be avoided by releasing the lysate slowly and holding the angle of the


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release tube such that released gas will not directly blow down into the tube but against a side wall. The pressure and time of incubation in the bomb should be determined empirically by visual inspection of the cell lysate under a tissue culture microscope. Stain the homogenate with Trypan blue and note the number of unbroken cells and the state of the nuclei in a counting chamber. The optimal pressure will disrupt more than 98%of the cells, but all the nuclei will appear intact. 4. Layer the cell homogenate over 15% sucrose (10 mL) in a 50-mL conical tube and centrifuge at 525g (avg) for 30 min at 4°C without the brake. The nuclei can also be spun down without layering over sucrose, but the pellet is often difficult to see. 5. Remove the supernatant, which contains the latex bead phagosomes. Save an aliquot (100 μL) of the supernatant for Western blot and/or silver stain analysis. This is the postnuclear supernatant (PNS).

3.4. Phagosome Purification by Sucrose Step Gradient 1. Preequilibrate all solutions on ice and perform the rest of the procedure on ice. 2. Mix the phagosome sample with 62% sucrose to give a final (c) of 40% sucrose. 3. Layer sequentially in a Beckman Ultra Clear SW-28 rotor tube: 1 mL 62% sucrose, 8–10 mL phagosome sample at 40% sucrose, 8 mL 35% sucrose, 12 mL 25% sucrose, 7 mL 10% sucrose. The final volume is 38 mL; the tube will hold 40 mL. 4. Assure that the tubes are balanced and centrifuge in a swinging bucket rotor (Beckman SW-28) at 100,000g (rmax) for 1 h. 5. Carefully remove the tubes from the rotor and collect the phagosome fraction from the 10–25% sucrose interface using a transfer pipet. The bead layer will be clearly visible. 6. Mix the collected fraction with 20 mM HEPES pH 7.4 to 38 mL and centrifuge again in an SW-28 rotor at 100,000g (rmax) for 30 min. Fractions may be pooled, but the final concentration of sucrose must be 10% or less or the beads will not pellet. Much of the 25% layer often gets aspirated with the beads. 7. Carefully remove the tubes from the rotor and aspirate the supernatant completely. 8. Resuspend the pellet with an appropriate volume of ICAT labeling buffer, or 2x SDS-PAGE buffer, if the phagosome fraction will only be used for Western blot analysis.

3.5. Assessment of Phagosome Purity The easiest method of assessing phagosome fraction purity is by Western blot analysis. The phagosome fractions should be highly enriched with lamp-1, cathepsin D, and rab proteins such as Rab7 (8). Determinations of which antibodies to use must be made based on the nature of the fraction (a very short 5-min pulse with latex beads will not result in a phagosome fraction

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enriched with lamp-1 or cathepsin D, but should be enriched with rab5). In our project, we were interested in IFN-–induced proteins that might be localized to phagosomal membranes. One candidate was Lrg-47 (9), so we used an antibody to Lrg-47 to demonstrate enrichment in our phagosome fraction from INF- treated cells (see Fig. 1). Other data had suggested that Lrg-47 was localized to Golgi membranes in macrophages, so we demonstrated the absence of contamination with Golgi membranes by using antibodies against Golgi membrane markers such as GM130 (10). The nature of the questions being asked will dictate which proteins one should be concerned about when assessing purity. Western blot analysis is also a great way to confirm that protein hits by mass spectrometry are truly enriched in the phagosome fraction. 1. Pour a 10% polyacrylamide mini gel with a stacking gel using the discontinuous buffer system of Laemmli (11). 2. Prepare the samples that have not been resuspended in SDS-PAGE buffer by mixing the samples with 4x SDS-PAGE buffer (including 10% mercaptoethanol) to yield 1x SDS-PAGE buffer in the final sample. Boil the samples for 5 min before loading on to the gel. 3. Load 5–10 μg of protein from the PNS fraction and the phagosome fraction and run the gel until the desired molecular weight markers are near the bottom of the gel. 4. Generate Western blots by electoblotting the gel to PVDF membrane (must be wet with methanol) using Towbin buffer in a mini-gel electroblotter for 1 h at 100 V constant voltage (12). 5. Block the membrane with 5% milk in TBS-T for 1 h.

Fig. 1. Assessment of purified phagosome fraction purity. 5 μg of protein (pns) from cells stimulated or not with 30 units/mL IFN- purified latex bead phagosome fractions after a 1-h pulse, and nuclear fractions from those cells were separated by SDS-PAGE and Western blotted. Lanes: pns (p), phagosome fraction (ph), nuclear fraction (n); fractions from IFN-–treated cells are denoted by a symbol. Western blots were decorated with abs as described in Subheading 2.5.


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6. Wash one time with TBS-T. 7. Add primary antibody in TBS-T with 5% BSA and incubate overnight with agitation at 4°C. 8. Wash the blot with TBS-T three times for 5 min. 9. Add the secondary antibody in TBS-T with 5% milk and incubate at room temperature with shaking for 1 h. 10. Wash the blot 3 × 5 min and leave in TBS-T to keep wet. 11. Incubate the blot for 5 min with ECL plus detection reagent according to the manufacturer’s protocol. 12. Expose the blot to Kodak Biomax XAR film for various times and develop the film with an automated film processor. 13. Alternatively, silver stains can be performed with the gel after SDS-PAGE using the BioRad Silver Stain Plus kit according to the manufacturer’s protocol. When performing silver stains, it is imperative to use clean glass plates in order to reduce background staining.

3.6. Protein Labeling with Isotope Coded Affinity Tag Cleavable ICAT reagent is a thiol reactive molecule that is manufactured in heavy (9 × C13 ) and light (9 × C12 ) forms with an acid-cleavable biotin group (7). The ICAT buffer denatures the proteins in the sample, which are then reduced, giving the ICAT reagent access to reduced cysteine residues. This enables two samples—in our case, one collected from IFN-–stimulated cells and one collected from unstimulated cells—to be labeled in separate reactions with heavy and light reagents and then be combined and processed for mass spectrometry analysis. 1. Determine the concentration of protein in your samples with a BCA or Bradford assay. Follow the manufacturer’s instructions and dilute the BSA standard with ICAT buffer to make a standard curve and assure there are no inhibitory components in the buffer. This protocol works very well with 1 mg of total protein, 0.5 mg from each sample. Less protein will not yield as good a result, and much less will not yield any reasonable result by this protocol. More protein may be labeled, but the column referred to for cation exchange in this protocol can only bind up to 5 mg of protein and runs optimally with 1 mg of total protein. 2. Reduce the samples by adding TCEP to a final concentraton of 5 mM. TCEP in H2 O is very acidic, but the amount of Tris in the ICAT buffer used in this protocol is high in order to account for this step. 3. Check the sample pH; it must remain between 8.0 and 8.5 for ICAT labeling to proceed. Adjust the pH with ammonium bicarbonate if necessary. The sample pH can be checked by spotting a small amount (1 μL) on pH paper with an appropriate range. 4. Incubate the samples for 30 min at 37°C.

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5. Remove 4 μg of protein from each sample to use as a control for labeling efficiency and digestion efficacy by silver stain. 6. Estimate the moles of cysteine residues in each sample. Assumptions: average protein 50 kDa, average of six cysteines per protein. Formula: x g protein/ 50,000 g/mol = mol protein × 6 = mol of cyteine residues. 7. Determine the amount of ICAT reagent in nmol that will be necessary to achieve a 1.2 mM concentration in the samples. Determine how many nmol of ICAT reagent are necessary to achieve a twofold molar excess of ICAT reagent to cysteine residues. Use the greater amount. Each tube contains 175 nmol of ICAT reagent. Try to label both samples under very similar conditions. Use the same amount of protein; try to use the same volume for the reaction. 8. Determine the number of tubes required to label the samples, round up to the nearest full tube. First, spin the tubes in a desktop centrifuge to collect the reagent powder in the bottom. Add the reduced sample to the tube, vortex, spin, and add the sample to any additional tubes necessary to achieve the predetermined molar ratio. Label one fraction (control) with light ICAT reagent and the other fraction (stimulus) with heavy ICAT reagent. 9. Incubate the tubes for 2 h at 37°C with agitation in the dark. 10. Stop the reaction by adding a 10-fold molar excess of DTT to the reaction tube, vortex, and incubate at room temperature for 5 min. 11. Remove 4 μg of protein from each sample to use as a control for labeling efficiency and digestion efficacy by silver stain. 12. Combine the light and heavy samples and dilute the combined sample sixfold with 50 mM Tris-HCl pH 8.3; check the final pH to ensure it is at 8.3. 13. Add trypsin 1:50 w/w with respect to μg of protein and let the sample digest overnight at 37°C with agitation. 14. Remove 4 μg of protein from the combined sample to use as a control for labeling efficiency and digestion efficacy by silver stain. 15. Confirm by SDS-PAGE and silver stain (see Subheading 3.5.) that the sample has been labeled and digested, if the digestion is incomplete, return the sample to 37°C incubation. When the digestion is complete, continue with the next step. 16. Centrifuge the sample in a desktop refrigerated centrifuge at 2500g (rmax) for 1.5 h, carefully remove the supernatant, and filter out the remaining latex beads with a 0.2-μm low-protein-binding syringe filter.

3.7. Cation Exchange Chromatography and Avidin Affinity Chromatography 1. Prepare the cation exchange column as per the manufacturer’s instructions. 2. Dilute the sample 1:1 with SCX buffer A and verify that the sample pH is less than or equal to 3.0. If the pH is not less than or equal to 3.0, titrate with 5% phosphoric acid to achieve the appropriate pH.


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3. Wash the column with 5 mL of SCX buffer A at 0.2 mL/min. 4. Using a 2-mL sample loop, load 2 mL of sample sequentially and wash with SCX buffer A until the UV baseline returns to zero, approximately 3 mL. Continue to load the entire sample. 5. Run a linear gradient to 25% SCX buffer B for 25 min and collect 1-min fractions. 6. Run a linear gradient to 100% SCX buffer B for 50 min and collect 1-min fractions. 7. Wash the column with 8 mL of SCX buffer B. 8. Wash the column with 8 mL of SCX buffer A. 9. Prepare the column for storage as per the manufacturer’s recommendations. 10. Neutralize the fractions which contain peptides as determined by the 214-nm trace with 0.4 volumes of 100 mM NaPO4 pH 10.0. Add 1 volume of 2x PBS pH 7.2. 11. Perform Avidin affinity chromatography and acid cleavage of the ICAT reagent according to the manufacturer’s instructions for each neutralized fraction. 12. The fractions are now ready to be analyzed by a mass spectrometry platform. We further fractionate the avidin purified fractions by reverse phase chromatography in line with MS/MS using an LCQ-Deca ion-trap mass spectrometer (Thermofinnigan) (13). The recorded data are searched against a database of mouse proteins maintained in house by SEQUEST software (14). The results are then submitted to a suite of analysis programs that provide a probability that the search results are correct at the peptide level (PeptideProphet) and determine a combined probability score and a best matched list of proteins found in the sample (ProteinProphet) (15–17). Information about the relative abundance of heavy- and light-labeled peptides is determined by XPRESS software (18), and averages are generated for all peptides determined to belong to a protein by ProteinProphet. The final combined results from two separate labeling experiments of one latex bead phagosome purified sample obtained using IFN-as a differential stimulus are shown in Table 1.

Table 1 Combined Results from LC-MS/MS Analysis of Two Labeling Experiments (each 1 mg protein total)

Number p-Value/range

Total peptides

Total proteins

Differential proteins

7932 0.05a

582 0.5b

11 (1.7–16.7)c

Peptide p-values ≥ 0.05, determined by PeptideProphet. Protein p-values ≥ 0.5, determined by ProteinProphet. c Range for average ratio of heavy to light peptides identified in 11 proteins, determined by XPRESS. a b

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4. Notes 1. RAW 264.7 cells are very sensitive to lipopolysaccharide (LPS) and other bacterial contaminants. All reagents should be screened for LPS by use of the Limulus Amoebocyte Lysate (LAL) assay. All glassware must be baked in order to destroy LPS. Antibodies are often highly contaminated with LPS. The antibody used in this step showed no detectable LPS by LAL, but a rabbit anti-ovalbumin antibody was highly contaminated and required clean-up with LPS-removing chromatography beads (Acticlean Etox, Sterogene Bioseparations, Inc.). 2. Confirmation that the beads are IgG opsonized can be performed by immunofluorescence with a fluorescent secondary to the opsonizing antibody. IgG opsonization increases the efficiency of bead uptake by RAW264.7 cells and also elicits an oxidative burst when the cells are prestimulated with IFN-as measured by a luminol-based assay (19). 3. RAW 264.7 cells stick tenaciously to many tissue culture–treated flasks. The use of nontreated Petri dishes allows for rapid cell removal and reduced space requirements in an incubator, but one must be careful not to lose cells during the DPBS wash step. 4. We often load the beads for 1 h to maximize the number of internalized beads, but any pulse-chase strategy can be performed. The RAW 264.7 cells are resilient to culture in shaking suspension, but we have not performed chase periods beyond 4 h in shaking suspension. During the chase period, we dilute the cells to 5 × 106 /mL in full medium with 25 mM HEPES. 5. Other methods of cell disruption can be used, such as Dounce homogenization, but the composition of the HB would have to be changed. We have found the nitrogen decompression method to be extremely reproducible.

Acknowledgments This work was supported by an NIH grant CA104242 to J.C. We would like to acknowledge the prior work of Alan Aderem, Ruedi Aebersold, and David Goodlet, which made the development of this protocol possible. Both Derek Einhaus and Sam Donahoe provided technical assistance with the development of this protocol. Finally, many others at the Institute for Systems Biology in Seattle provided help to answer many questions. References 1. Underhill, D. M. and Ozinsky, A. (2002) Phagocytosis of microbes: complexity in action. Annu. Rev. Immunol. 20, 825–852. 2. Blander, J. M. (2006) Coupling Toll-like receptor signaling with phagocytosis: potentiation of antigen presentation. Trends. Immunol. 28, 19–25. 3. Blander, J. M. and Medzhitov, R. (2006) On regulation of phagosome maturation and antigen presentation. Nat. Immunol. 7(10), 1029–1035.


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4. Wetzel, M. G. and Korn, E. D. (1969) Phagocytosis of latex beads by Acahamoeba castellanii (Neff). 3. Isolation of the phagocytic vesicles and their membranes. J. Cell Biol. 43(1), 90–104. 5. Garin, J., et al. (2001) The phagosome proteome: insight into phagosome functions. J. Cell Biol. 152(1), 165–180. 6. Gygi, S. P., et al. (1999) Quantitative analysis of complex protein mixtures using isotope-coded affinity tags. Nat. Biotechnol. 17(10), 994–999. 7. Yi, E. C., et al. (2005) Increased quantitative proteome coverage with (13)C/(12) C-based, acid-cleavable isotope-coded affinity tag reagent and modified data acquisition scheme. Proteomics 5(2), 380–387. 8. Yeung, T., et al. (2006) Lipid metabolism and dynamics during phagocytosis. Curr. Opin. Cell Biol. 18(4), 429–437. 9. Feng, C. G., et al. (2004) Mice deficient in LRG-47 displayincreased susceptibility to mycobacterial infection associated with the induction of lymphopenia. J. Immunol. 172(2), 1163–1168. 10. Martens, S., et al. (2004) Mechanisms regulating the positioning of mouse p47 resistance GTPases LRG-47 and IIGP1 on cellular membranes: retargeting to plasma membrane induced by phagocytosis. J. Immunol. 173(4), 2594–2606. 11. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227(5259), 680–685. 12. Towbin, H., Staehelin, T. and Gordon, J. (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl. Acad. Sci. USA 76(9), 4350–4354. 13. Yi, E. C., et al., Approaching complete peroxisome characterization by gas-phase fractionation. Electrophoresis 23(18), 3205–3216. 14. Yates, J. R., 3rd, et al. (1995) Method to correlate tandem mass spectra of modified peptides to amino acid sequences in the protein database. Anal. Chem. 67(8), 1426–1436. 15. Keller, A., et al. (2002) Empirical statistical model to estimate the accuracy of peptide identifications made by MS/MS and database search. Anal. Chem. 74(20), 5383–5392. 16. Nesvizhskii, A .I., et al. (2003) A statistical model for identifying proteins by tandem mass spectrometry. Anal. Chem. 75(17), 4646–4658. 17. von Haller, P. D., et al. (2003) The application of new software tools to quantitative protein profiling via isotope-coded affinity tag (ICAT) and tandem mass spectrometry: II. Evaluation of tandem mass spectrometry methodologies for largescale protein analysis, and the application of statistical tools for data analysis and interpretation. Mol. Cell Proteomics 2(7), 428–442. 18. Han, D. K., et al. (2001) Quantitative profiling of differentiation-induced microsomal proteins using isotope-coded affinity tags and mass spectrometry. Nat. Biotechnol. 19(10), 946–951. 19. Gantner, B. N., et al. (2003) Collaborative induction of inflammatory responses by dectin-1 and Toll-like receptor 2. J. Exp. Med. 197(9), 1107–1117.

23 Class II MHC Antigen Processing in Phagosomes Lakshmi Ramachandra, W. Henry Boom, and Clifford V. Harding

Summary Phagocytic antigen-presenting cells (APCs) are involved in innate and adaptive immune responses to bacteria. Adaptive responses to bacteria involve processing of bacterial antigens for presentation by class II major histocompatibility complex (MHC II) molecules and class I MHC (MHC I) molecules to stimulate CD4+ and CD8+ T cells, respectively. To examine the role of phagosomes in processing of antigens for presentation by MHC II molecules to CD4+ T cells, phagosomes have been biochemically and functionally analyzed by a variety of techniques that include flow analysis (flow organellometry), SDS-PAGE/Western blotting, and an antigen-presenting organelle assay. Using these techniques, we have demonstrated that phagosomes containing latex beads or Mycobacterium tuberculosis (MTB) contain components of the MHC II processing pathway and support the formation of peptide–MHC II complexes.

Key Words: Phagosome; antigen processing; class II MHC; Mycobacterium tuberculosis; macrophage.

1. Introduction Macrophages and immature dendritic cells are phagocytic cells that internalize particulate antigens (Ags, such as microbes or latex beads conjugated to Ag) and present peptides derived from these Ags on MHC I and MHC II molecules for T-cell recognition (1–3). During the processing of particulate Ags, peptide–MHC II complexes may be formed within phagosomes or within endocytic compartments (e.g., MHC II compartment [MIIC]) that receive Ag fragments from phagosomes. From: Methods in Molecular Biology, vol. 445: Autophagosome and Phagosome Edited by: V. Deretic © Humana Press, Totowa, NJ

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To assess the role of phagosomes in the formation of peptide–MHC II complexes, we have developed and modified a variety of techniques to biochemically and functionally evaluate phagosomes. Phagosomes were analyzed for the presence of components of the MHC II processing pathway by flow organellometry (4–7) and sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE)/Western analysis of purified phagosomes (5–7). Degradation of phagosome-associated Ag was analyzed by flow organellometry (4–6). Finally, presence of peptide–MHC II complexes in phagosomes was functionally evaluated by the Ag-presenting organelle assay (5–8). Analysis of phagosomes by these techniques has to be done with the exclusion of other organelles. This can be achieved either by analytic isolation by flow organellometry or by physical isolation. Latex bead phagosomes can be analytically isolated by flow organellometry from other components of the cells without the need for extensive prior physical purification (4–6). Latex bead phagosomes can also be physically isolated on sucrose density gradients or Percoll density gradients, and magnetic latex bead phagosomes can be isolated using a magnet. Phagosomes containing Mycobacterium tuberculosis (MTB) can be purified on Percoll density gradients. By analyzing purified phagosomes by the techniques described, we have demonstrated that latex bead phagosomes rapidly degrade Ag (not evaluated for MTB phagsomes (4–6)) . Phagosomes containing either latex beads or MTB acquire lysosomal-associated membrane protein-1 (LAMP-1) and MHC II as well as some of the other components of the MHC II–processing pathway, e.g., H2-DM and invariant chain (5,7,9). Both latex bead and MTB phagosomes also support the formation of peptide–MHC II complexes, which are subsequently transported to the cell surface and presented to T cells (5,7).

2. Materials 2.1. Cell Culture Macrophages can be obtained directly from the peritoneal cavity of mice or from progenitor cells in the bone marrow. Generation of peritoneal exudate cells (PECs) for experiments such as subcellular fractionation requires a large number of mice (1 million PECs is usually obtained per mouse). Significantly fewer mice (10-fold less) are required to generate the same number of bone marrow–derived macrophages (BMMs). For experiments described in the Methods section, we have found interferon (IFN)--activated BMMs to be functionally equivalent to PECs. For studies using human macrophages/monocytes, activated human monocytic cell lines (e.g., THP-1) or human monocyte-derived macrophages (MDMs) can be used.

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To analyze either phagocytic Ag processing and presentation in intact cells or the presence of specific peptide–MHC II complexes in subcellular fractions (including phagosomes), Ag-specific, MHC II–restricted T cells have been used. Therefore, for these types of studies, it is essential to use macrophages expressing the MHC II alleles recognized by the T cells. The MHC II alleles expressed by murine macrophages can be determined by the strain of mice being used. The MHC II alleles expressed by human macrophages can be determined by HLA typing of DNA extracted from cells. 1. For murine macrophages: Mice expressing known MHC II alleles for generation of PECs or BMMs. Sterile scissors and forceps for dissection. 2. For human macrophages: Human monocytic cell line THP-1 (ATTC, Manassas, VA) and/or human MDMs expressing known MHC II alleles. 3. T cells: Murine T hybridoma cells with appropriate Ag specificity and MHC II restriction. Murine T-cell hybridomas derived from HLA-transgenic mice can be used to detect peptide:MHC complexes presented by human APCs (10). Alternatively, human T cell lines may be used, but they may not respond to fixed APCs or subcellular organelles due to requirements for (fixation-sensitive) accessory molecules. 4. CTLL-2 cells (ATTC) or IL-2 ELISA (eBioscience, San Diego, CA).

2.2. Media 1. Standard medium for murine cells: Dulbecco’s modified Eagle’s medium (DMEM) with glutamine (Hyclone, Logan, UT) supplemented with 10% heatinactivated fetal calf serum (FCS, Hyclone), 50 μM 2-mercaptoethanol (2-ME, Sigma, St. Louis, MO), 1 mM sodium pyruvate, 10 mM hydroxyethyl piperazine ethane sulfonate (HEPES) buffer, 100 U/mL penicillin, and 100 μg/mL streptomycin (Invitrogen Corporation, Carlsbad, CA). 2. Medium for generation of BMMs: Standard medium supplemented with 20% LADMAC cell-conditioned media (11). For generation of LADMAC cellconditioned media, grow LADMAC cells in Alpha medium (Eagle’s minimal essential medium [EMEM, Cambrex, Walkersville, MD], supplemented with 10% FCS [Hyclone] and 4mM l-glutamine [Hyclone]) for 7 d and harvest supernatants. Store supernatants at –80°C for up to 1 yr. 3. Standard medium for human cells: For THP-1 cells use RPMI 1640 (BioWhitaker, Walkersville, MD) supplemented with 10% heat-inactivated FCS (Hyclone), 50 μM 2-mercaptoethanol (2-ME, Sigma), 1 mM sodium pyruvate, 2 mM l-glutamine, 10 mM HEPES buffer, nonessential amino acids, 100 U/mL penicillin, and 100 μg/mL streptomycin (Invitrogen). For human monocytes, use standard medium with 10% pooled human serum (PHS) instead of FBS.

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2.3. Reagents for Activation/Maintenance of Cells 1. For activation of murine BMMs: Murine IFN- (Genzyme, Cambridge, MA). Working concentration is 50–100 U/mL. 2. For activation of THP-1 cells: Phorbol 12-myristate 13-acetate (PMA, Sigma) and human IFN- (Endogen, Woburn, MA). Working concentration is 10 ng/mL for PMA and 50 U/mL for IFN-. 3. For maintanence of CTLL-2 cells: Rat interleukin (IL)-2 (BD Pharmingen, San Diego, CA) (12).

2.4. Preparation of Particulate Ag (Ag-Conjugated Latex Beads and Bacteria) 1. Latex beads: 2-μm latex beads (fluorescent or nonfluorescent; Polysciences, Warrington, PA) or 1- to 2-μm magnetic latex beads (Polysciences). 1-μm latex beads will not readily pellet in a typical tabletop centrifuge and should not be used in experiments described in this chapter. 2. Preparation of antigen for conjugation to latex beads: The antigen to be used for conjugation to latex beads is determined by the antigen specificity of the T-cell hybridomas available for the project. For example, DOBW T hybridoma cells recognize ovalbumin (OVA) peptide 323-339 bound to MHC II molecules I-Ad or I-Ab (5). Therefore, latex beads have to be conjugated to OVA in experiments using DOBW T hybridoma cells. Polysciences provides instructions for conjugation of proteins to beads (covalent or noncovalent). Noncovalent conjugation (passive adsorption) is sufficient for most purposes. The pH of the conjugation buffer may require adjusting for each protein. For example, prepare solution of OVA (Sigma, St. Louis, MO) or hen egg lysosyme (HEL, Sigma) at 5 mg/mL in citrate buffer (pH 4.2) or phosphate-buffered saline (PBS) (pH 7.4), respectively. 3. Bacteria such as MTB, E. coli, Salmonella, or Streptococcus. 4. For declumping MTB: 18- and 22-gauge needles. 5. For fluorescent labeling of MTB or other bacteria: FLUOS (Boehringer), prepared when needed in dimethyl sulfoxide (DMSO) at a concentration of 20 mg/mL, and PBS (pH 9.0).

2.5. Inhibitors 1. Cytochalasin D (Sigma) is dissolved in DMSO at a concentration of 1 mg/mL and stored in aliquots at –20°F. Working concentration is 10 μg/mL. 2. Chloroquine (Sigma) is prepared when needed in water at a concentration of 10 mM. Working concentration is 100 μM. 3. Brefeldin A (BFA, Sigma) is dissolved in ethanol at a concentration of 1 mg/mL and stored in aliquots at –20°F. Working concentration is 1 μg/mL.


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2.6. Sucrose Density Gradients 1. Versene (Gibco-BRL). 2. Homogenization buffer: 0.25 M sucrose (Mallinckrodt, St. Louis, MO) in 10 mM HEPES buffer solution, pH 7.5 (100 mM stock from Gibco BRL). Homogenization buffer should be sterile filtered and stored at 4°C. On the day of the experiment, supplement a small portion of the homogenization buffer with the following protease inhibitors: 1.0 mM phenylmethysulfonyl fluoride (PMSF, Sigma), 1 μg/mL pepstatin (Sigma), 20 μg/mL leupeptin (Sigma). Protease inhibitors: PMSF is dissolved in 100% isopropanol at a concentration of 100 mM and stored in the dark at room temperature. Pepstatin is dissolved in ethanol (with heat up to 60°C) at 1 mg/mL and stored in aliquots at –20°C. Leupeptin is dissolved in Millipore water at 2 mg/mL and stored in aliquots at –20°C (see Note 1). 3. Sucrose solutions of different percentages (62, 32, 26, 21, and 10%) are made by dissolving sucrose in 10 mM HEPES buffer solution (pH 7.5, Gibco-BRL). Sucrose solutions should be sterile filtered through a 0.22-μm filter and stored at 4°C (see Note 2). 4. Cell lifters (Corning). 5. Dounce homogenizer (7 mL capacity, Kontes, Vineland, NJ), autoclaved prior to use.

2.7. Percoll Density Gradients 1. Versene (Gibco-BRL). 2. Homogenization buffer (see Subheading 2.6., step 2). 3. Percoll solutions: Made by diluting Percoll (Amersham) to 20, 23, 27, or 40% in homogenization buffer lacking protease inhibitors. 4. Cell lifters (Corning). 5. Dounce homogenizer (7 mL capacity, Kontes, Vineland, NJ), autoclaved prior to use. 6. Polycarbonate centrifuge tubes (for ultracentrifuge), autoclaved prior to use.

2.8. Biochemical Analysis of Percoll Density Gradient Fractions 1. -Hexosaminidase assay: Assay buffer is 0.1 M 2-(N-morpholino)ethanesulfonic acid (MES, Sigma, pH 6.5) in 0.2% Triton X-100 (Sigma). The substrate is p-nitrophenyl-acetyl--d-glucosaminide and is made just before addition at a concentration of 1.36 mg/mL in Millipore water. 2. For identification of plasma membrane: 125 I-labeled anti-MHC II Abs or sulfoNHS-biotin (Pierce) at 0.5 mg/mL in PBS (pH 8.0). 3. For identification of phagosomes: Fluorescent latex beads (2 μm, Polysciences) or FLUOS-labeled MTB (see Subheadings 2.4., step 5 and 3.2.2.).

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2.9. Flow Analysis of Phagosomes 1. Homogenization buffer (see Subheading 2.6., step 2). 2. 2% paraformaldehyde (Polysciences) is made in PBS (pH 7.4). Solution should be stirred on a heating plate maintained at 56°C until paraformaldehyde is dissolved and then vacuum-filtered through 0.22-μm filter and stored at 4°C (Caution: see Note 3). 3. 0.4 M lysine (dl-lysine, Sigma) is made in Millipore water. Solution should be stirred on a heating plate maintained at 56°C until lysine is dissolved and then sterile filtered through 0.22-μm filter and stored at 4°C. 4. Cell lifters (Corning). 5. 96-Well round bottom plates. 6. FACS buffer is made by adding 1% FCS, 1% bovine serum albumin (BSA), and 0.1% saponin to PBS (pH 7.4). A 1% stock solution of saponin should be made in PBS and added to FACS buffer. 7. Antibodies (Abs) against MHC II, LAMP-1 and against Ags that have been conjugated to latex beads (anti-OVA and anti-HEL).

2.10. Western Blot Analysis of Phagosomes Lysis buffer: PBS (pH 7.4) containing 1% Nonidet P-40 (Sigma) and the protease inhibitors 1 mM PMSF, 20 μg/mL leupeptin, and 2 μg/mL pepstatin (see Subheading 2.6., step 2 on how to make and store protease inhibitors and Note 4). 2.11. Ag Processing and Presentation Assay (T-Cell Assay) 1. 96-Well flat-bottom plates. 2. Murine PECs or BMMs, human monocytes, or monocyte-derived macrophages (see Subheading 2.1.) expressing known MHC II molecules. 3. Ag-specific, MHC II-restricted T hybridoma cells (see Subheading 2.1., step 3). 4. Ag: Latex-beads conjugated to Ag or bacteria (see Subheadings 2.4. and 3.2.). 5. 2% paraformaldehyde and 0.4 M lysine (see Subheading 2.9., step 2 and 2.9., step 3). 6. For detection of IL-2 by the CTLL-2 proliferation assay: CTLL-2 cells (ATCC, Manassas, VA (12)), culture supernatants containing IL-2 and Alamar Blue (Trek Diagnostics, Cleveland, OH). Alternatively, an IL-2 ELISA may be used (eBioscience).

2.12. Ag-Presenting Organelle Assay 1. 96-Well flat-bottom plates. 2. Percoll density gradient fractions or purified phagosomes derived from macrophages expressing known MHC II molecules. 3. MHC II-restricted T hybridoma cells (see Subheading 2.1., step 3). 4. Percoll solution (see Subheading 2.7., step 3). 5. Reagents for the detection of IL-2 by the CTLL-2 proliferation assay or IL-2 ELISA (see Subheading 2.11., step 6).


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3. Methods 3.1. Isolation/Preparation and Maintenance of Cells 3.1.1. Murine Macrophages 3.1.1.1. PECs 1. For generation of PECs, inject mice intraperitonally with 1 mL of 1 × 104 live Listeria monocytogenes (LM) in PBS (pH 7.4; see Note 5) (13). Alternatively, inject mice intraperitonally with 100 μg of concanavalin A (Con A) in 500 μL of PBS (see Note 6). 2. After 4 d (for Con A) or 10–14 d (for LM), harvest cells from the peritoneal cavity of mice (13). 3. Plate cells in 96-well (2 × 105 cells/well) or 6-well plates (3 × 106 cells/well) for 2 h to allow macrophages to adhere to plastic. Rinse off nonadhered cells with DMEM.

3.1.1.2. Murine BMMs 1. For generation of BMMs, isolate both femurs from mice. Cut ends of femurs with sterile scissors and flush bone with DMEM using a 23-gauge needle. 2. Spin and resuspend cells in 12 mL of LADMAC-supplemented media. Plate cells in 6-well plates (2 mL per well). Grow cells at 37°C in 5% CO2 . Replace media after 3 d with 3 mL of LADMAC-supplemented media and subsequently replace after every 2 d. BMMs can be used after 7–14 d. Cells have to be activated with 50–100 U recombinant murine IFN- for 24 h prior to use in Ag-processing assays to upregulate MHC II.

3.1.2. Human Macrophages 3.1.2.1. Activation of THP-1 Cells

To induce differentiation of THP-1 cells into macrophages, activate cells in media supplemented with 10 ng/mL PMA. After 24 h, remove media and incubate cells for an additional 24 h with new medium containing 50 U/mL recombinant human IFN- to upregulate MHC II. 3.1.2.2. Human Monocyte-Derived Macrophages

For generation of human monocyte-derived macrophages, isolate peripheral blood mononuclear cells (PBMCs) and enrich monocytes by allowing them to adhere to tissue culture plates (plastic adherence) for 1 h at 37°C. Alternatively, purify CD14+ monocytes by immunomagnetic cell separation according to the manufacturer’s instructions (Miltenyi Biotec, Auburn, CA). Culture monocytes for 5–7 d to differentiate them into MDMs.

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3.1.2.3. Maintenance of CTLL-2 Cells

Seed CTLL-2 cells at a concentration of 1 × 105 cells/10 mL for a 2-d passage, 5 × 104 cell/10mL for a 3-d passage, or 2.5 × 104 for a 4-d passage. Add 2 U/mL of rat IL-2 (BD Pharmingen). Use only 3- or 4-d passage for IL-2 assay; 2-, 3-, or 4-d passages may be used to set up additional passages (12). 3.2. Preparation of Particulate Ags 3.2.1. Conjugation of Ag to Latex Beads 1. Ag is conjugated to latex beads by passive adsorption. For conjugation of OVA to latex beads, wash 100 μL of latex beads in 1 mL of citrate buffer (pH 4.2). Resuspend beads in 1 mL of 5 mg/mL OVA solution. For conjugation of HEL to latex beads, wash 100 μL of latex beads in 1 mL of PBS (pH 7.4). Resuspend beads in 1 mL of 5 mg/mL HEL solution. 2. Rotate beads with Ag for 2 h at room temperature or at 4°C overnight. Wash beads a minimum of three times in 1 mL of DMEM and resuspend in DMEM corresponding to the original volume of beads. Samples should be stored at 4°C and can used for up to 3 wk. Yield obtained is approximately 1012 beads/mL and 100 μg OVA/mL or 400 μg HEL/mL.

3.2.2. Preparation of MTB 1. MTB has to be declumped prior to use in experiments. For declumping, passage MTB twice through an 18-gauge needle and thrice through a 22-gauge needle. Centrifuge sample at 150g for 5 min to remove clumps. Estimate concentration of viable MTB by analyzing colony forming units (CFUs) on Middlebrook 7H11 plates (Difco, Detroit, MI). 2. For labeling MTB with fluorescein, pellet 109 bacteria in an Eppendorf tube and resuspend in 1 mL of PBS (pH 9.1). Combine with 25 μL of 20 mg/mL FLUOS in DMSO for 5 min at room temperature. Wash labeled MTB twice in DMEM and declump before use.

3.3. Ag-Presenting Assay (T-Cell Assay Using Intact Cells) Prior to addition of Ag to wells, warm up centrifuge to 30–37°C (see Note 7). An example of results from a T-cell assay is shown in Fig. 1. 3.3.1. Standard T-Cell Assay Plate macrophages at a concentration of 2 × 105 cells/well in 96-well flatbottom plates. Leave overnight for BMM. In the case of PECs remove nonadherent cells after 2 h by washing the wells once with 100 μL of medium. Certain cells may have to be activated with IFN- prior to use (see Subheading 3.1.).


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Fig. 1. Requirements for processing of MTB bacilli for presentation of MTB Ag 85B to T cells. Macrophages were pretreated with or without 10 μg/mL cytochalasin D (to inhibit phagocytic uptake) or 100 μM chloroquine (to inhibit acidification of vacuolar compartments) beginning 15 min before addition (pulse) of HK bacteria for 20 min. Samples were washed to remove extracelluar bacteria, chased for 10 min at 37°C and fixed. BB7 T hybridoma cells were used to detect Ag 85B(241-256):I-Ab complexes. MOI = multiplicity of infection. (Adapted from Ramachandra et al., 2001, with permission from the Journal of Experimental Medicine, The Rockefeller University Press.)

1. Add particulate Ag (see Subheadings 2.4. and 3.2.) to the wells in a final volume of 100 μL. Spin the plate at 900g for 5 min (10 min for MTB) to pellet the Ag onto the cells. Incubate the cells at 37°C for an additional 5 min (10 min for MTB) to provide a total pulse time of 10 min (20 min for MTB). 2. Place the plate on ice and wash the wells twice in 100 μL ice-cold DMEM to remove extracellular particulates (observe cells under the microscope and repeat wash if necessary). 3. Add prewarmed media to the cells and incubate cells at 37°C to achieve the desired chase incubation. 4. Fix the cells as follows to prevent any additional processing. Prepare a 1:1 solution of DMEM (no FCS) and 2% paraformaldehyde (fixative solution) as well as a 1:1 solution of 0.4 M lysine and DMEM (Barf solution). Wash wells with 150 μL of DMEM. Add 120 μL of fixative solution to the wells and incubate for 15 min at room temperature. Wash wells with 150 μL of DMEM and then incubate cells in 175 μL of Barf solution for 30 min at room temperature. Wash wells four times with 200 μL of DMEM and then resuspend cells in 100 μL of media.

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5. Add T hybridoma cells (100 μL at 106 cells/mL) to achieve 105 T hybridoma cells/well in a total volume of 200 μL. Incubate cells for 24 h (see Note 8). 6. Harvest supernatants (100 μL) and assess for IL-2 content using the CTLL-2 proliferation assay or IL-2 ELISA (see Note 9). 7. For the CTLL-2 assay, add CTLL-2 cells (50 μL at 5 × 103 /mL) to 100 μL of supernatant. After 24 h add 15 μL of the indicator dye Alamar blue to each well and measure the difference between absorbance at 550 and 595 nm (or 570 and 600 nm) after another 24 h. Blanks for spectrophotometry are provided by wells containing medium alone (added at the initiation of the CTLL-2 assay) and Alamar blue (added at the same time as for the other wells).

3.3.2. Antigen-Presenting Assay with Inhibitors The following inhibitors can be used in T-cell assays to functionally characterize phagocytic Ag processing and should be present throughout the duration of the Ag-processing step (added prior to addition of Ag and present until fixation of APCs, but not present during the T-cell assay) (Fig. 1). 1. Cytochalasin D inhibits phagocytic uptake and should be added at a concentration of 10 μg/mL to cells, 15 min prior to addition of particulate Ag. 2. Chloroquine inhibits acidification of vacuolar compartments and hence formation of peptide–MHC II complexes and should be added at a concentration of 100 μM to cells 15 min prior to addition of particulate Ag. 3. BFA is a fungal metabolite that inhibits anterograde transport through the endoplasmic reticulum (ER) and Golgi complexes. BFA blocks the supply of nascent MHC II molecules to the endocytic/phagocytic pathway and inhibits the formation of most peptide:MHC II complexes in the MIIC compartment as well as in phagosomes (6). Add BFA at a concentration of 1 μg/mL to cells 3 h prior to the addition of particulate Ag. BFA blocks ER-Golgi transport rapidly, but a period of 3 h is required to deplete the post-Golgi reservoir of nascent MHC II.

3.4. Isolation of Latex Bead Phagosomes by Sucrose Density Gradients Phagosomes are isolated by sucrose density gradients essentially as described by Desjardins et al., with minor modifications (14). Use fluorescent latex beads to easily identify phagosomes in the sucrose density gradient. This technique produces highly purified phagosomes as judged by electron microscopy and the low degree of contamination by plasma membrane marker (

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