The Unfolded Protein Response and Cellular Stress, Part A

VOLUME FOUR HUNDRED AND EIGHT Y-NINE METHODS IN ENZYMOLOGY The Unfolded Protein Response and Cellular Stress, Part A...

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VOLUME FOUR HUNDRED AND EIGHT Y-NINE

METHODS

IN

ENZYMOLOGY The Unfolded Protein Response and Cellular Stress, Part A

METHODS IN ENZYMOLOGY Editors-in-Chief

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

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

VOLUME FOUR HUNDRED AND EIGHT Y-NINE

METHODS

IN

ENZYMOLOGY The Unfolded Protein Response and Cellular Stress, Part A EDITED BY

P. MICHAEL CONN Divisions of Reproductive Sciences and Neuroscience (ONPRC) Departments of Pharmacology and Physiology Cell and Developmental Biology, and Obstetrics and Gynecology (OHSU) Beaverton, OR, USA

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

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32 Jamestown Road, London NW1 7BY, UK First edition 2011 Copyright # 2011, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@ elsevier.com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made For information on all Academic Press publications visit our website at elsevierdirect.com ISBN: 978-0-12-385116-1 ISSN: 0076-6879 Printed and bound in United States of America 11 12 13 10 9 8 7 6 5 4 3 2 1

CONTENTS

Contributors Preface Volumes in Series

Section I. UPR and Cell Stress in Normal and Diseased Cells 1. The Unfolded Protein Response in Mouse Cerebral Cortex

xi xvii xix

1 3

Nirinjini Naidoo 1. Introduction: The UPR 2. Transduction of the UPR Signal by PERK, IRE1, and ATF6 3. UPR and Apoptosis 4. The UPR in Health and Disease 5. Induction of the UPR by Sleep Deprivation 6. Experimental Protocols 7. Concluding Remarks Acknowledgments References

2. Immunohistochemical Detection of the Unfolded Protein Response in Atherosclerotic Plaques

3 4 5 6 6 7 18 19 19

23

Sˇa´rka Lhota´k, Ji Zhou, and Richard C. Austin 1. Endoplasmic Reticulum and the Unfolded Protein Response 2. Atherosclerosis 3. UPR in the Atherosclerotic Lesion 4. Mouse Models of Atherosclerosis 5. Methods Acknowledgments References

3. The Role of PDI as a Survival Factor in Cardiomyocyte Ischemia

24 24 25 27 29 44 44

47

Stefano Toldo, Anna Severino, Antonio Abbate, and Alfonso Baldi 1. Introduction 2. Identification of PDI as Cardio-Protective Factor

48 50 v

vi

Contents

3. Conclusions References

4. Measurement of the Increase in Endoplasmic Reticulum Stress-Related Proteins and Genes in Adipose Tissue of Obese, Insulin-Resistant Individuals

62 62

67

Guenther Boden and Salim Merali 1. Introduction 2. Study Subjects and Fat Biopsies 3. Proteomic Analysis 4. Western Blot Analysis 5. RT-PCR Analysis Acknowledgments References

5. Measurement of the Unfolded Protein Response (UPR) in Monocytes

68 69 70 79 80 81 82

83

Toma´s P. Carroll, Catherine M. Greene, and Noel G. McElvaney 1. Introduction 2. Investigating UPR Activation in the Monocyte Acknowledgments References

6. Assessment of Endurance Exercise Training in Hepatic Endoplasmic Reticulum Stress

84 86 93 93

97

Natalie Ann Chapados 1. Introduction 2. Assessment of an Endurance-Training Program 3. “Dietary-Inducible” Hepatic ER Stress 4. Assessment of Hepatic ER Stress Molecular Markers 5. Required Materials Acknowledgments References

7. Discovery Approaches to UPR in Athero-Susceptible Endothelium In Vivo

98 99 104 105 105 106 106

109

Mete Civelek, Elisabetta Manduchi, Gregory R. Grant, Christian J. Stoeckert Jr., and Peter F. Davies 1. Introduction 2. Procedures for the Isolation of Endothelial Cells and Preparation of RNA for Microarray Hybridization 3. Microarray Hybridization and Feature Extraction

110 112 116

Contents

4. Bioinformatics Analysis 5. Another Approach to Gene Connectivity: Weighted Gene Coexpression Network Analysis 6. Validation and Follow-up Acknowledgments References

8. Assessment of Endoplasmic Reticulum Stress and the Unfolded Protein Response in Endothelial Cells

vii 117 122 124 125 125

127

Ines Witte and Sven Horke 1. Introduction 2. The Acute Phase of ER Stress 3. The Adaptive Reaction Phase: Unfolded Protein Response in Endothelial Cells 4. The Final Phase: Cell Death Signaling Acknowledgments References

9. Measuring the Impact of Cigarette Smoke on the UPR

128 129 131 140 145 145

147

Hong Zhao, Jin Yang, Lin Shan, and Ellen D. Jorgensen 1. Introduction 2. Preparing Samples for Analysis 3. Analysis of Activation of the PERK Pathway 4. ATF6 Pathway 5. IRE1 Pathway 6. Using LSC to Measure UPR Activation Acknowledgments References

10. Induction and Measurement of UPR and Osmotic Stress in the Yeast Pichia pastoris

148 149 155 158 159 159 161 162

165

Martin Dragosits, Diethard Mattanovich, and Brigitte Gasser 1. Introduction 2. Measuring UPR Induction in P. pastoris 3. Inducing and Measuring Osmotic Stress in P. pastoris Acknowledgments References

11. Induction of ER Stress Response Leading to Programmed Cell Death in Trypanosoma brucei

166 169 179 186 186

189

Hanoch Goldshmidt and Shulamit Michaeli 1. Introduction 2. Induction of ER Stress in Trypanosomes

190 193

viii

Contents

3. Measurements of ER Stress in Trypanosomes Cells 4. Measurements of ER Stress-Induced Cell Death Acknowledgments References

12. Measurement of Activation of the Endoplasmic Reticulum Stress Response in Autoimmune Myositis

194 201 203 203

207

Heather M. Alger, Sree Rayavarapu, and Kanneboyina Nagaraju 1. Introduction 2. Measuring Endoplasmic Reticulum Stress Response in Skeletal Muscle Tissues Using Western Blotting 3. Immunohistochemical Detection of ER Stress Markers in Muscle Tissues 4. Determination of mRNA Levels of ER Stress Targets 5. Concluding Remarks Acknowledgments References

13. Activation of the Unfolded Protein Response in Human Acute Myeloid Leukemia

208 209 216 221 223 223 223

227

Julian A. Schardt, Beatrice U. Mueller, and Thomas Pabst 1. Introduction 2. Technical Aspects 3. Discussion References

14. Modulation of the Unfolded Protein Response by GRP78 in Prostate Cancer

228 232 239 241

245

Gustaaf de Ridder, Rupa Ray, Uma K. Misra, and Salvatore V. Pizzo 1. Introduction 2. Methods for Monitoring UPR Modulation 3. Conclusions References

15. Endoplasmic Reticulum Stress in Brain Damage

246 248 256 256

259

Ram Raghubir, Venkata Prasuja Nakka, and Suresh L. Mehta 1. Introduction 2. ER Stress and Unfolded Protein Response 3. Cross Talk Between ER and Mitochondria 4. Experimental Approaches for the Detection of ER Stress Acknowledgment References

260 261 267 269 271 272

Contents

16. Methods and Models for Monitoring UPR-Associated Macrophage Death During Advanced Atherosclerosis

ix

277

Edward B. Thorp 1. Introduction to ER Stress in Atherosclerosis 2. Significance of Macrophage ER Stress and Apoptosis in Advanced Atherosclerosis 3. Modeling Atherosclerosis-Relevant ER Stress-Induced Apoptosis 4. The UPR and Macrophage Death at the Murine Aortic Root 5. Conclusion and Future Methodological Advances Acknowledgments References

278 279 281 285 293 293 293

17. Detecting Autophagy in Response to ER Stress Signals in Cancer 297 Marıá Salazar, Sonia Hernańdez-Tiedra, Sofıá Torres, Mar Lorente, Manuel Guzmań, and Guillermo Velasco 1. Introduction 2. Monitoring Autophagy Upon ER Stress 3. Monitoring Autophagy Upon ER Stress In Vivo References

18. The Unfolded Protein Response Induced by Salt Stress in Arabidopsis

298 299 309 317

319

Miaoying Wang, Qiangyi Xu, and Ming Yuan 1. Introduction 2. The Treatment of Arabidopsis Seedlings 3. The Analysis of Unfolded Protein Response Acknowledgments References

19. Measurement of ER Stress Response and Inflammation in the Mouse Model of Nonalcoholic Fatty Liver Disease

319 320 322 327 327

329

Ze Zheng, Chunbin Zhang, and Kezhong Zhang 1. Introduction 2. Methods to Measure ER Stress Response and Hepatic Inflammation in the Mouse Model of NAFLD 3. Visualizing ER Stress Response Associated with Liver Pathology in the Mouse Model of NAFLD Acknowledgments References Author Index Subject Index

330 332 341 345 345 349 365

CONTRIBUTORS

Antonio Abbate Victoria Johnson Research Center and VCU Pauley Heart Center, Virginia Commonwealth University, Richmond, Virginia, USA Heather M. Alger Research Center for Genetic Medicine, Children’s National Medical Center, Washington, DC, USA Richard C. Austin Department of Medicine and Division of Nephrology, St. Joseph’s Hospital and McMaster University, Hamilton, Ontario, Canada Alfonso Baldi Department of Biochemistry and Biophysics F. Cedrangolo, Second University of Naples, Italy Guenther Boden Department of Medicine and the Clinical Research Center, Temple University School of Medicine, Philadelphia, PA, USA Toma´s P. Carroll Department of Medicine, Royal College of Surgeons in Ireland, Education and Research Centre, Beaumont Hospital, Dublin, Ireland Natalie Ann Chapados Institut de Recherche de l’Hoˆpital Montfort/Montfort Hospital, Research Institute, Ottawa, Ontario, Canada Mete Civelek Institute for Medicine and Engineering, and Department of Bioengineering, University of Pennsylvania, Philadelphia, Pennsylvania; Division of Cardiology, Department of Medicine, University of California, Los Angeles, California, USA Peter F. Davies Institute for Medicine and Engineering, and Department of Bioengineering; Department of Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA Gustaaf de Ridder Department of Pathology, Duke University Medical Center, Durham, North Carolina, USA xi

xii

Contributors

Martin Dragosits Department of Biotechnology, University of Natural Resources and Life Sciences, Vienna, Muthgasse 18, Vienna, Austria, and UC Davis Genome Center, University of California Davis, One Shields Avenue, Davis, USA Brigitte Gasser Department of Biotechnology, University of Natural Resources and Life Sciences, Vienna, and Austrian Centre of Industrial Biotechnology (ACIB GmbH), Muthgasse 18, Vienna, Austria Hanoch Goldshmidt The Mina and Everard Goodman Faculty of Life Sciences, and Advanced Materials and Nanotechnology Institute, Bar-Ilan University, Ramat-Gan, Israel Gregory R. Grant Center for Bioinformatics, and Department of Genetics, University of Pennsylvania, Philadelphia, Pennsylvania, USA Catherine M. Greene Department of Medicine, Royal College of Surgeons in Ireland, Education and Research Centre, Beaumont Hospital, Dublin, Ireland Manuel Guzmań Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, and Centro de Investigacioń Biome´dica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Madrid, Spain Sonia Hernańdez-Tiedra Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, Madrid, Spain Sven Horke Institute of Pharmacology, University Medical Center of the Johannes Gutenberg University Mainz, Germany Ellen D. Jorgensen Department of Pathology, New York Medical College, Valhalla, New York, USA Sˇa´rka Lhota´k Department of Medicine and Division of Nephrology, St. Joseph’s Hospital and McMaster University, Hamilton, Ontario, Canada Mar Lorente Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, Madrid, Spain Elisabetta Manduchi Center for Bioinformatics, and Department of Genetics, University of Pennsylvania, Philadelphia, Pennsylvania, USA

Contributors

xiii

Diethard Mattanovich Department of Biotechnology, University of Natural Resources and Life Sciences, Vienna, and Austrian Centre of Industrial Biotechnology (ACIB GmbH), Muthgasse 18, Vienna, Austria Noel G. McElvaney Department of Medicine, Royal College of Surgeons in Ireland, Education and Research Centre, Beaumont Hospital, Dublin, Ireland Suresh L. Mehta Department of Pharmaceutical Sciences/BRITE, North Carolina Central University, Durham, North Carolina, USA Salim Merali Department of Biochemistry, Temple University School of Medicine, Philadelphia, PA, USA Shulamit Michaeli The Mina and Everard Goodman Faculty of Life Sciences, and Advanced Materials and Nanotechnology Institute, Bar-Ilan University, Ramat-Gan, Israel Uma K. Misra Department of Pathology, Duke University Medical Center, Durham, North Carolina, USA Beatrice U. Mueller Department of Internal Medicine, University Hospital, Bern, Switzerland Kanneboyina Nagaraju Research Center for Genetic Medicine, Children’s National Medical Center, and Department of Pediatrics, The George Washington University Medical Center, Washington, DC, USA Venkata Prasuja Nakka Department of Neurological Surgery, School of Medicine & Public Health, University of Wisconsin, Madison, Wisconsin, USA Nirinjini Naidoo Center for Sleep and Circadian Neurobiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania, USA Thomas Pabst Department of Medical Oncology, University Hospital, Bern, Switzerland Salvatore V. Pizzo Department of Pathology, Duke University Medical Center, Durham, North Carolina, USA

xiv

Contributors

Ram Raghubir Division of Pharmacology, Central Drug Research Institute, (CSIR), Chatter Manzil Palace, Lucknow, India Rupa Ray Department of Pathology, Duke University Medical Center, Durham, North Carolina, USA Sree Rayavarapu Research Center for Genetic Medicine, Children’s National Medical Center, and Institute of Biomedical Sciences, The George Washington University, Washington, DC, USA Marıá Salazar Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, and Centro de Investigacioń Biome´dica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Madrid, Spain Julian A. Schardt Department of Medical Oncology, University Hospital, Bern, Switzerland Anna Severino Institute of Cardiology, Catholic University of the Sacred Heart, Rome, Italy Lin Shan Pathology Department, Duke University Medical Center, Durham, North Carolina, USA Christian J. Stoeckert Jr. Center for Bioinformatics, and Department of Genetics, University of Pennsylvania, Philadelphia, Pennsylvania, USA Edward B. Thorp Department of Medicine, Division of Molecular Medicine, Columbia University, New York, USA Stefano Toldo Victoria Johnson Research Center and VCU Pauley Heart Center, Virginia Commonwealth University, Richmond, Virginia, USA Sofıá Torres Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, Madrid, Spain Guillermo Velasco Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, and Centro de Investigacioń Biome´dica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Madrid, Spain

Contributors

xv

Miaoying Wang State Key Laboratory of Plant Physiology and Biochemistry, College of Biological Sciences, China Agricultural University, Beijing, China Ines Witte Institute of Pharmacology, University Medical Center of the Johannes Gutenberg University Mainz, Germany Qiangyi Xu State Key Laboratory of Plant Physiology and Biochemistry, College of Biological Sciences, China Agricultural University, Beijing, China Jin Yang Education and Training Systems International, Inc., Chapel Hill, North Carolina, USA Ming Yuan State Key Laboratory of Plant Physiology and Biochemistry, College of Biological Sciences, China Agricultural University, Beijing, China Chunbin Zhang The Center for Molecular Medicine and Genetics, The Wayne State University School of Medicine, Detroit, Michigan, USA Kezhong Zhang The Center for Molecular Medicine and Genetics, and The Department of Immunology and Microbiology, The Wayne State University School of Medicine, Detroit, Michigan, USA Hong Zhao Department of Pathology, New York Medical College, Valhalla, New York, USA Ze Zheng The Center for Molecular Medicine and Genetics, The Wayne State University School of Medicine, Detroit, Michigan, USA, and Currently a graduate student in the program of Biological Science at the Wayne State University Ji Zhou Department of Medicine, Thrombosis and Atherosclerosis Research Institute (TaARI) and McMaster University, Hamilton, Ontario, Canada

PREFACE

The observation that the living cell contains a mechanism to sense and correct the accumulation of unfolded (or incorrectly folded) proteins in the endoplasmic reticulum was formidable in organizing thoughts about cellular integration. This mechanism both halts further protein synthesis and promotes the production of chaperone proteins that act to relieve this problem. If this problem cannot be corrected, the mechanism can initiate programmed cell death. Aspects of this unfolded protein response (UPR) are conserved from yeast to man, an observation that suggests a key role in the process of maintaining a living cell. The UPR presents a way of understanding cellular regulation, a mechanism for disease, and a therapeutic opportunity. The present volume provides descriptions of the occurrence of the UPR, the methods used to assess it, pharmacological tools, and other methodological approaches to analyze its impact on cellular regulation. The authors explain how these methods are able to provide important biological insights. Authors were selected based on research contributions in the area about which they have written and based on their ability to describe their methodological contributions in a clear and reproducible way. They have been encouraged to make use of graphics, comparisons to other methods, and to provide tricks and approaches not revealed in prior publications that make it possible to adapt methods to other systems. The editor expresses appreciation to the contributors for providing their contributions in a timely fashion, to the senior editors for guidance, and to the staff at Academic Press for their helpful inputs. P. MICHAEL CONN

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METHODS IN ENZYMOLOGY

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

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Methods in Enzymology

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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VOLUME 328. Applications of Chimeric Genes and Hybrid Proteins (Part C: Protein–Protein Interactions and Genomics) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 329. Regulators and Effectors of Small GTPases (Part E: GTPases Involved in Vesicular Traffic) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 330. Hyperthermophilic Enzymes (Part A) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 331. Hyperthermophilic Enzymes (Part B) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 332. Regulators and Effectors of Small GTPases (Part F: Ras Family I) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 333. Regulators and Effectors of Small GTPases (Part G: Ras Family II) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 334. Hyperthermophilic Enzymes (Part C) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 335. Flavonoids and Other Polyphenols Edited by LESTER PACKER VOLUME 336. Microbial Growth in Biofilms (Part A: Developmental and Molecular Biological Aspects) Edited by RON J. DOYLE VOLUME 337. Microbial Growth in Biofilms (Part B: Special Environments and Physicochemical Aspects) Edited by RON J. DOYLE VOLUME 338. Nuclear Magnetic Resonance of Biological Macromolecules (Part A) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 339. Nuclear Magnetic Resonance of Biological Macromolecules (Part B) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 340. Drug–Nucleic Acid Interactions Edited by JONATHAN B. CHAIRES AND MICHAEL J. WARING VOLUME 341. Ribonucleases (Part A) Edited by ALLEN W. NICHOLSON VOLUME 342. Ribonucleases (Part B) Edited by ALLEN W. NICHOLSON VOLUME 343. G Protein Pathways (Part A: Receptors) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 344. G Protein Pathways (Part B: G Proteins and Their Regulators) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT


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VOLUME 345. G Protein Pathways (Part C: Effector Mechanisms) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 346. Gene Therapy Methods Edited by M. IAN PHILLIPS VOLUME 347. Protein Sensors and Reactive Oxygen Species (Part A: Selenoproteins and Thioredoxin) Edited by HELMUT SIES AND LESTER PACKER VOLUME 348. Protein Sensors and Reactive Oxygen Species (Part B: Thiol Enzymes and Proteins) Edited by HELMUT SIES AND LESTER PACKER VOLUME 349. Superoxide Dismutase Edited by LESTER PACKER VOLUME 350. Guide to Yeast Genetics and Molecular and Cell Biology (Part B) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 351. Guide to Yeast Genetics and Molecular and Cell Biology (Part C) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 352. Redox Cell Biology and Genetics (Part A) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 353. Redox Cell Biology and Genetics (Part B) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 354. Enzyme Kinetics and Mechanisms (Part F: Detection and Characterization of Enzyme Reaction Intermediates) Edited by DANIEL L. PURICH VOLUME 355. Cumulative Subject Index Volumes 321–354 VOLUME 356. Laser Capture Microscopy and Microdissection Edited by P. MICHAEL CONN VOLUME 357. Cytochrome P450, Part C Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 358. Bacterial Pathogenesis (Part C: Identification, Regulation, and Function of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 359. Nitric Oxide (Part D) Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 360. Biophotonics (Part A) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 361. Biophotonics (Part B) Edited by GERARD MARRIOTT AND IAN PARKER

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VOLUME 362. Recognition of Carbohydrates in Biological Systems (Part A) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 363. Recognition of Carbohydrates in Biological Systems (Part B) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 364. Nuclear Receptors Edited by DAVID W. RUSSELL AND DAVID J. MANGELSDORF VOLUME 365. Differentiation of Embryonic Stem Cells Edited by PAUL M. WASSAUMAN AND GORDON M. KELLER VOLUME 366. Protein Phosphatases Edited by SUSANNE KLUMPP AND JOSEF KRIEGLSTEIN VOLUME 367. Liposomes (Part A) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 368. Macromolecular Crystallography (Part C) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 369. Combinational Chemistry (Part B) Edited by GUILLERMO A. MORALES AND BARRY A. BUNIN VOLUME 370. RNA Polymerases and Associated Factors (Part C) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 371. RNA Polymerases and Associated Factors (Part D) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 372. Liposomes (Part B) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 373. Liposomes (Part C) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 374. Macromolecular Crystallography (Part D) Edited by CHARLES W. CARTER, JR., AND ROBERT W. SWEET VOLUME 375. Chromatin and Chromatin Remodeling Enzymes (Part A) Edited by C. DAVID ALLIS AND CARL WU VOLUME 376. Chromatin and Chromatin Remodeling Enzymes (Part B) Edited by C. DAVID ALLIS AND CARL WU VOLUME 377. Chromatin and Chromatin Remodeling Enzymes (Part C) Edited by C. DAVID ALLIS AND CARL WU VOLUME 378. Quinones and Quinone Enzymes (Part A) Edited by HELMUT SIES AND LESTER PACKER VOLUME 379. Energetics of Biological Macromolecules (Part D) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 380. Energetics of Biological Macromolecules (Part E) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS


VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES

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VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA


VOLUME 417. Functional Glycomics Edited by MINORU FUKUDA VOLUME 418. Embryonic Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 419. Adult Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 420. Stem Cell Tools and Other Experimental Protocols Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 421. Advanced Bacterial Genetics: Use of Transposons and Phage for Genomic Engineering Edited by KELLY T. HUGHES VOLUME 422. Two-Component Signaling Systems, Part A Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 423. Two-Component Signaling Systems, Part B Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 424. RNA Editing Edited by JONATHA M. GOTT VOLUME 425. RNA Modification Edited by JONATHA M. GOTT VOLUME 426. Integrins Edited by DAVID CHERESH VOLUME 427. MicroRNA Methods Edited by JOHN J. ROSSI VOLUME 428. Osmosensing and Osmosignaling Edited by HELMUT SIES AND DIETER HAUSSINGER VOLUME 429. Translation Initiation: Extract Systems and Molecular Genetics Edited by JON LORSCH VOLUME 430. Translation Initiation: Reconstituted Systems and Biophysical Methods Edited by JON LORSCH VOLUME 431. Translation Initiation: Cell Biology, High-Throughput and Chemical-Based Approaches Edited by JON LORSCH VOLUME 432. Lipidomics and Bioactive Lipids: Mass-Spectrometry–Based Lipid Analysis Edited by H. ALEX BROWN

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VOLUME 433. Lipidomics and Bioactive Lipids: Specialized Analytical Methods and Lipids in Disease Edited by H. ALEX BROWN VOLUME 434. Lipidomics and Bioactive Lipids: Lipids and Cell Signaling Edited by H. ALEX BROWN VOLUME 435. Oxygen Biology and Hypoxia Edited by HELMUT SIES AND BERNHARD BRU¨NE VOLUME 436. Globins and Other Nitric Oxide-Reactive Protiens (Part A) Edited by ROBERT K. POOLE VOLUME 437. Globins and Other Nitric Oxide-Reactive Protiens (Part B) Edited by ROBERT K. POOLE VOLUME 438. Small GTPases in Disease (Part A) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 439. Small GTPases in Disease (Part B) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 440. Nitric Oxide, Part F Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 441. Nitric Oxide, Part G Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 442. Programmed Cell Death, General Principles for Studying Cell Death (Part A) Edited by ROYA KHOSRAVI-FAR, ZAHRA ZAKERI, RICHARD A. LOCKSHIN, AND MAURO PIACENTINI VOLUME 443. Angiogenesis: In Vitro Systems Edited by DAVID A. CHERESH VOLUME 444. Angiogenesis: In Vivo Systems (Part A) Edited by DAVID A. CHERESH VOLUME 445. Angiogenesis: In Vivo Systems (Part B) Edited by DAVID A. CHERESH VOLUME 446. Programmed Cell Death, The Biology and Therapeutic Implications of Cell Death (Part B) Edited by ROYA KHOSRAVI-FAR, ZAHRA ZAKERI, RICHARD A. LOCKSHIN, AND MAURO PIACENTINI VOLUME 447. RNA Turnover in Bacteria, Archaea and Organelles Edited by LYNNE E. MAQUAT AND CECILIA M. ARRAIANO


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VOLUME 448. RNA Turnover in Eukaryotes: Nucleases, Pathways and Analysis of mRNA Decay Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 449. RNA Turnover in Eukaryotes: Analysis of Specialized and Quality Control RNA Decay Pathways Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 450. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 451. Autophagy: Lower Eukaryotes and Non-Mammalian Systems (Part A) Edited by DANIEL J. KLIONSKY VOLUME 452. Autophagy in Mammalian Systems (Part B) Edited by DANIEL J. KLIONSKY VOLUME 453. Autophagy in Disease and Clinical Applications (Part C) Edited by DANIEL J. KLIONSKY VOLUME 454. Computer Methods (Part A) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 455. Biothermodynamics (Part A) Edited by MICHAEL L. JOHNSON, JO M. HOLT, AND GARY K. ACKERS (RETIRED) VOLUME 456. Mitochondrial Function, Part A: Mitochondrial Electron Transport Complexes and Reactive Oxygen Species Edited by WILLIAM S. ALLISON AND IMMO E. SCHEFFLER VOLUME 457. Mitochondrial Function, Part B: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases Edited by WILLIAM S. ALLISON AND ANNE N. MURPHY VOLUME 458. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides Edited by DAVID A. HOPWOOD VOLUME 459. Complex Enzymes in Microbial Natural Product Biosynthesis, Part B: Polyketides, Aminocoumarins and Carbohydrates Edited by DAVID A. HOPWOOD VOLUME 460. Chemokines, Part A Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 461. Chemokines, Part B Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 462. Non-Natural Amino Acids Edited by TOM W. MUIR AND JOHN N. ABELSON VOLUME 463. Guide to Protein Purification, 2nd Edition Edited by RICHARD R. BURGESS AND MURRAY P. DEUTSCHER

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VOLUME 464. Liposomes, Part F Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 465. Liposomes, Part G Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 466. Biothermodynamics, Part B Edited by MICHAEL L. JOHNSON, GARY K. ACKERS, AND JO M. HOLT VOLUME 467. Computer Methods Part B Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 468. Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding: Part A Edited by DANIEL HERSCHLAG VOLUME 469. Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding: Part B Edited by DANIEL HERSCHLAG VOLUME 470. Guide to Yeast Genetics: Functional Genomics, Proteomics, and Other Systems Analysis, 2nd Edition Edited by GERALD FINK, JONATHAN WEISSMAN, AND CHRISTINE GUTHRIE VOLUME 471. Two-Component Signaling Systems, Part C Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 472. Single Molecule Tools, Part A: Fluorescence Based Approaches Edited by NILS G. WALTER VOLUME 473. Thiol Redox Transitions in Cell Signaling, Part A Chemistry and Biochemistry of Low Molecular Weight and Protein Thiols Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 474. Thiol Redox Transitions in Cell Signaling, Part B Cellular Localization and Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 475. Single Molecule Tools, Part B: Super-Resolution, Particle Tracking, Multiparameter, and Force Based Methods Edited by NILS G. WALTER VOLUME 476. Guide to Techniques in Mouse Development, Part A Mice, Embryos, and Cells, 2nd Edition Edited by PAUL M. WASSARMAN AND PHILIPPE M. SORIANO VOLUME 477. Guide to Techniques in Mouse Development, Part B Mouse Molecular Genetics, 2nd Edition Edited by PAUL M. WASSARMAN AND PHILIPPE M. SORIANO VOLUME 478. Glycomics Edited by MINORU FUKUDA


VOLUME 479. Functional Glycomics Edited by MINORU FUKUDA VOLUME 480. Glycobiology Edited by MINORU FUKUDA VOLUME 481. Cryo-EM, Part A: Sample Preparation and Data Collection Edited by GRANT J. JENSEN VOLUME 482. Cryo-EM, Part B: 3-D Reconstruction Edited by GRANT J. JENSEN VOLUME 483. Cryo-EM, Part C: Analyses, Interpretation, and Case Studies Edited by GRANT J. JENSEN VOLUME 484. Constitutive Activity in Receptors and Other Proteins, Part A Edited by P. MICHAEL CONN VOLUME 485. Constitutive Activity in Receptors and Other Proteins, Part B Edited by P. MICHAEL CONN VOLUME 486. Research on Nitrification and Related Processes, Part A Edited by MARTIN G. KLOTZ VOLUME 487. Computer Methods, Part C Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 488. Biothermodynamics, Part C Edited by MICHAEL L. JOHNSON, JO M. HOLT AND GARY K. ACKERS VOLUME 489. The Unfolded Protein Response and Cellular Stress, Part A Edited by P. MICHAEL CONN

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S E C T I O N

O N E

UPR AND CELL STRESS IN NORMAL AND DISEASED CELLS


C H A P T E R

O N E

The Unfolded Protein Response in Mouse Cerebral Cortex Nirinjini Naidoo Contents 3 4 5 6 6 7 7 18 19 19

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

Introduction: The UPR Transduction of the UPR Signal by PERK, IRE1, and ATF6 UPR and Apoptosis The UPR in Health and Disease Induction of the UPR by Sleep Deprivation Experimental Protocols 6.1. Tissue procurement 7. Concluding Remarks Acknowledgments References

Abstract The unfolded protein response (UPR) is important for normal cellular homeostasis, development, and physiology. Endoplasmic reticulum stress and the UPR have also been implicated in a variety of diseases and pathologies. Thus the UPR is being studied in many model systems. We describe in this chapter methods for studying the UPR in a mouse model, specifically the cerebral cortex of mouse brain.

1. Introduction: The UPR The endoplasmic reticulum (ER) is a reticular membranous network that extends throughout the cytoplasm. In eukaryotic cells, it is the compartment within which all secretory and transmembrane proteins are folded in an energy-dependent manner. To ensure the proper folding of proteins, the ER utilizes a dedicated machinery of molecular chaperones and folding enzymes that associate directly with nascent polypeptides. These include the Center for Sleep and Circadian Neurobiology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00001-7

#

2011 Elsevier Inc. All rights reserved.

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molecular chaperone BiP/GRP78 (immunoglobulin-binding protein/glucose-regulated protein 78) and PDI (protein disulfide isomerase). Perturbations that alter ER homeostasis, therefore, disrupt folding and lead to the accumulation of unfolded proteins and protein aggregates, which are detrimental to cell survival. These perturbations include disturbances in calcium homeostasis, redox status, elevated secretory protein synthesis, and glucose or energy deprivation. As a consequence, the cell has evolved an adaptive coordinated response to limit accumulation of unfolded proteins in the ER. This signaling pathway is termed the ER stress response or the unfolded protein response (UPR) (for reviews, see Harding et al., 2002; Schroder and Kaufman, 2005a,b, 2006; Zhang and Kaufman, 2004). On a cellular level, the UPR triggers three kinds of protective cellular responses: (a) upregulation of ER-resident chaperones to assist in the refolding of misfolded proteins; (b) attenuation of protein translation which is mediated by the serine–threonine kinase PERK that serves to reduce protein load in the ER; and (c) degradation of misfolded proteins by the proteasome by a process called ER-associated degradation (ERAD). These three responses are protective measures to alleviate ER stress; however, prolonged stress leads to the activation of proapoptotic pathways (Szegezdi et al., 2006).

2. Transduction of the UPR Signal by PERK, IRE1, and ATF6 ER stress signals are transduced across the ER membrane by three proximal sensors of the UPR, inositol-requiring element-1 (IRE-1), PKRlike endoplasmic reticulum kinase (PERK), and activating transcription factor 6 (ATF6). All three of these sensors are maintained in an inactive state at the ER membrane by binding to BiP. Upon accumulation of unfolded proteins, bound BiP dissociates from ATF6, IRE1, and PERK to chaperone the misfolded proteins thereby permitting the activation of one or more of these transducers (Zhang and Kaufman, 2006). BiP, therefore, serves as the master regulator of the UPR. PERK signaling attenuates protein translation. PERK is a type I transmembrane serine–threonine kinase that appears to be present in most cells. Binding to BiP holds it in an inactive monomeric state. When this binding is disrupted, PERK is activated through homodimerization and autophosphorylation. Activated PERK phosphorylates translation initiation factor 2a (eIF2a), thereby reducing the rate of translation through the formation of a stalled 43S ternary complex. This results in a general decrease of most proteins. However, some selected proteins with internal ribosomal entry sites (IRES), such as ATF4 (activating transcription factor 4), GRP94, and

The Unfolded Protein Response in Mouse Cerebral Cortex

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BiP, are translated more efficiently and hence their protein levels actually increase. PERK also activates two different regulators of redox homeostasis: Nrf2 (nuclear respiratory factor -2) and ATF4 (Cullinan et al., 2003). Signaling by both Nrf2 and ATF4 can modulate common targets, including the proapoptotic factor CHOP/GADD153 (C/EBP homologous protein/ growth arrest and DNA damage-inducible gene 153) (Cullinan and Diehl, 2004). In cells lacking Nrf2, CHOP expression levels are elevated compared to their wild-type controls, while Nrf2 overexpression ameliorates CHOP accumulation during the UPR (Cullinan and Diehl, 2004). In contrast, ATF4, a major downstream target of eIF2a phosphorylation, promotes CHOP expression (Harding et al., 2000). When ER stress is not mitigated, CHOP levels increase (Zinszner et al., 1998), which leads to activation of cell cycle arrest and apoptosis. IRE1 has both kinase and endoribonuclease activities. Once activated, the cytoplasmic domain of IRE1a gains endoribonuclease activity and cleaves 26 nucleotides from the mRNA encoding X-box-binding protein (XBP) 1, generating a spliced variant (XBP1s) that functions as a potent transcriptional transactivator of genes involved in ER expansion, protein maturation, folding and export from the ER, as well as export and degradation of misfolded proteins (Yoshida et al., 2001). Recent data indicate that the trigger for IRE1’s endoribonuclease activity is not phosphorylation per se, but rather a conformational change in the kinase domain induced by nucleotide binding, which is promoted by trans-autophosphorylation of the kinase activation loop (Ron and Walter, 2007). IRE1 may also degrade ER-targeted mRNAs, thus decreasing the production of new proteins in the organelle (Hollien and Weissman, 2006). ATF6 is a 90-kDa-bZIP protein that is activated by posttranslational modifications. ATF6 activation as part of the UPR leads to its translocation to the Golgi and cleavage by site-1 and -2 proteases (S1P and S2P). The 50-kDa cleaved ATF6a translocates to the cell nucleus, where it binds to the ER stress response element CCAAT(N)9CCACG (Yoshida et al., 1998) in genes encoding ER chaperone proteins such as BiP and GRP94. This results in increases in the level of these proteins and hence improved protein-folding activity in the ER (Yoshida et al., 1998).

3. UPR and Apoptosis Excessive and prolonged stress leads to apoptosis (Szegezdi et al., 2006). Apoptosis in response to ER stress is specific to metazoan cells (Schroder and Kaufman, 2005a). When cell protective changes mediated by the UPR fail to restore folding capacity, a combination of both the intrinsic and extrinsic apoptotic pathways are activated (Schroder and

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Kaufman, 2005b). CHOP, together with Bcl-2 family members (Bak/Bax), caspase-12, and c-jun NH2 terminal kinase ( JNK) are components of the ER stress-mediated apoptotic pathway (Wu and Kaufman, 2006). Targets of CHOP include Bcl-2, GADD34, ERO1a, and TRB3 (Szegezdi et al., 2003). CHOP is known to repress Bcl-2 which is anti-apoptotic.

4. The UPR in Health and Disease The UPR pathway is important for normal cellular homeostasis, development, and physiology. ER stress and the UPR have also been implicated in a variety of diseases and pathologies including diabetes (Eizirik et al., 2008; Harding and Ron, 2002), cerebral ischemia (DeGracia and Montie, 2004), viral infections (He, 2006), cardiovascular disease (Vasa-Nicotera, 2004), cancer (Ma and Hendershot, 2004), inclusion body myositis (Vattemi et al., 2004), and neurodegenerative diseases (Lindholm et al., 2006; Rao et al., 2004).

5. Induction of the UPR by Sleep Deprivation We have shown that acute sleep loss or sleep deprivation leads to the induction of the UPR in mouse cerebral cortex (Naidoo et al., 2005) and Drosophila brain (Naidoo et al., 2007). In fact, several microarray studies indicate that BiP transcript is increased with sleep deprivation in all species examined, in Drosophila (Shaw et al., 2000), rat cerebral cortex (Cirelli et al., 2004), mouse cerebral cortex (Mackiewicz et al., 2007; Maret et al., 2007), and the telencephalon of the white crowned sparrow ( Jones et al., 2008). In mice, transcript and protein levels of BiP increase with 6, 9, and 12 h of sleep deprivation (Mackiewicz et al., 2007; Naidoo et al., 2005). Other UPR-specific transcripts that change with sleep deprivation in mouse cerebral cortex include DNA-J which is a cochaperone of BiP, XBP-1, calreticulin, caspase-9, ATF4, and ATF6. Immunoprecipitation studies indicate that BiP is dissociated from the kinase PERK with 6 h of sleep deprivation. There is phosphorylation of PERK and eIF2a indicative of inhibition of protein translation. Ribosome profiles from mice that have been subjected to 6 h of sleep deprivation indicate that there is a disaggregation of polysomes into monosomes in the sleep-deprived animals suggesting an attenuation of protein translation (Naidoo et al., 2005). Immunohistochemical studies also indicate that there is an increase in protein levels of ER chaperones GRP94, BiP/GRP78, and ERp72 during


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sleep deprivation in both the dorsal and lateral cortex of mouse brain (Terao et al., 2003) again compatible with the induction of the UPR. The adaptive UPR response to sleep deprivation is attenuated in aged mice (22–24 months of age). Following 3, 6, 9, and 12 h of sleep deprivation, BiP levels did not increase in the cerebral cortex of aged mice in contrast to the robust increase in young animals (Naidoo et al., 2008). Instead, animals display upregulation of proapoptotic markers (Naidoo et al., 2008). GADD34 (growth arrest and DNA damage-inducible protein), a protein phosphatase 1 (PP1)-interacting protein, which causes PP1 to dephosphorylate eIF2a and relieves the translational block imposed by eIF2a phosphorylation (Brush et al., 2003; Szegezdi et al., 2006), was increased with both age and sleep deprivation. There was also a dramatic increase in the proapoptotic factor CHOP with age that was further increased in older animals with sleep loss. A role for CHOP in mediating apoptosis in response to ER stress is well established (Wang et al., 1996; Zinszner et al., 1998).

6. Experimental Protocols 6.1. Tissue procurement We examined molecular changes occurring with sleep deprivation in mouse cerebral cortex. We used C57BL/6J mice in all our studies. The protocol describing procurement and homogenization of cortical tissue follows: 1. Euthanize/sacrifice mice by cervical dislocation. 2. Remove the brain and dissect out the cerebral cortex of each mouse rapidly on ice using mouse brain atlas of Franklin and Paxinos (1996) as a guide. 3. Briefly, the first coronal cut will be performed approximately 1 mm rostral to the point where the optic nerves merge to form the optic chiasm and the second cut approximately 1 mm caudally to the optic chiasm (or just “behind” it). From the resulting slice, the left and right part of the cerebral cortex containing cingulate cortex (Cg1 and Cg2) primary and secondary motor areas (M1 and M2), as well as the somatosensory cortex (S1 and S2), will be separated from the subcortical structures along the corpus callosum. 4. Flash freeze the tissue in liquid nitrogen and store at –70 C until use. All animal protocols should be performed in accordance with National Institutes of Health guidelines and be approved by the Institutional Animal Care and Use Committee.

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6.1.1. Tissue homogenization 1. Prepare lysis buffer for tissue homogenization. Lysis buffer 20 mM Tris–HCl pH 7.5 1 mM EGTA 1 mM EDTA 1% Triton X-100 10% glycerol Protease inhibitors (1 mM PMSF, 2 mg/ml pepstatin, and 4 mg/ml aprotinin). 2. Homogenize the cortical tissue in approximately 300 ml of cold lysis buffer on ice using a hand held or benchtop-motorized homogenizer equipped with a teflon pestle. 3. Centrifuge the lysate at 14,000 rpm for 10 min; save the supernatant on ice or at 4 C until the protein assay is done. 4. Determine protein concentration of the supernatant by the Pierce micro-BCA assay. 5. Store tissue homogenate in small aliquots (50 ml) at 70 C. 6.1.2. SDS-PAGE and westerns We use SDS-PAGE and westerns to determine the protein levels of BiP, activation of ATF6 by cleavage, activation of PERK by phosphorylation, phosphorylation of eIF2a, CHOP, and GADD34—the dilutions and source of antibodies used are listed in Table 1.1. 6.1.3. Determination of BiP/GRP78 BiP is the sentinel marker of the UPR. An increase in BiP expression usually indicates induction of the UPR. There are several commercial antibodies available to measure BiP protein levels. We have tried many of them and found that most work well. We prefer using the Stressgen (now Assay Designs) polyclonal BiP antibody for westerns. While we usually obtain just a single band at 78 kDa on our Western blots when testing a new batch of antibody or a new experiment, we usually run a sample of commercial hamster BiP/GRP78 (100–200 ng; Stressgen). All BiP antibodies tested are listed in Table 1.1. In our experiments, we examine the effect of different lengths of sleep deprivation on UPR induction in the mouse cerebral cortex. Individual sleep-deprived and matching control mouse cortex homogenates are run on SDS-PAGE gels in triplicate. We run several gels so that we can have both technical and biological replicates.

Table 1.1 UPR Antibodies, source, and description Antibody

Company

Immunogen

Host

Application Dilutions

GRP 78 (BiP) Santa Cruz aa 525–653 at the C-terminus of GRP 78 (H-129) Biotechnology GRP78 (BiP) Stressgen C-terminus of rat GRP78, sequence identical to that in mouse GADD 34 Santa Cruz C-terminus peptide of mouse GADD 34 (C-19) Biotechnology GADD 153 Santa Cruz Amino acids 1–168 representing full length mouse (CHOP) Biotechnology GADD153

Rabbit IHC

1:100

Rabbit Western

1:1000

Rabbit Western, IHC Rabbit Western, IHC

Caspase 12

Stressgen

Rabbit Western

1:200, 1:2000 1:500– 1:1000, 1:500 1:2000

P-eIF2a (Ser51) P-PERK

Rabbit Western

1:500

GADD 153 (CHOP) ATF6

Cell Signaling Technology Cell Signaling Technology Thermo Scientific Imgenex

Rabbit Western, IHC Mouse IHC

1:1000, 1:500 1:500

Mouse Western

1:500

eIF2a

Cell signaling

Rabbit Western

1:500

16 residue synthetic peptide corresponding to amino acids 2–17 of murine caspase-12 Synthetic phosphopeptide corresponding to residues surrounding Ser51 of eIF2a Phosphopeptide residues surrounding Thr980 of mouse PERK Corresponds to bacterially expressed mouse fusion CHOP Partial protein containing amino acids 1–273 of human ATF6 C-terminus sequence

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6.1.4. PERK activation and phosphorylation of eIF2a PERK is maintained in an inactive state by binding to the lumenal domain of BiP/GRP78. Under conditions of ER stress, BiP/GRP78 dissociates to service the increasing load of ER client proteins; loss of BiP/GRP78 binding correlates with oligomerization, trans-autophosphorylation, and activation of downstream signaling by PERK (Bertolotti et al., 2000). Activated PERK phosphorylates eIF2a on serine 51, a modification that increases the affinity of eIF2 for eIF2B, a GDP–GTP exchange factor that charges the eIF2–GTP–tRNA-met ternary complex (Kimball et al., 2001). Functioning as a competitive inhibitor of eIF2B, phospho-eIF2a reduces the concentration of the active ternary complex, prevents assembly of the 43S preinitiation complex, and attenuates protein translation. Use of specific antibodies to the phosphorylated state of both PERK and eIF2a are useful to determine activation of PERK. 6.1.5. Activation of ATF6 The ER lumenal domain of ATF6 has two Golgi localization signals: GLS1 and GLS2. Binding to BiP/GRP78 masks the localization signals in the lumenal region of ATF6; dissociation of BiP allows ATF6 to be transported to the Golgi body. Further, the ER lumenal domain of ATF6 has disulfide bonds and ER stress-induced reduction plays an important role in both translocation of ATF6 to Golgi body and the subsequent recognition by the S1P and S2P (Nadanaka et al., 2007; Shen et al., 2002). Once cleaved by S1P and S2P, the 50-kDa ATF6a translocates to the cell nucleus. 6.1.6. Procedure 1. Prepare samples (20 mg protein) representing individual mice by adding 2 Laemmli buffer and boiling for 2–5 min. 2. Run on SDS-PAGE gels (Bio-Rad, 10% Tris–HCl) according to Laemmli (1970). 3. Transfer gels to nitrocellulose membranes (Bio-Rad) using either semidry or wet transfer. 4. Following transfer, place membranes in blocking solution, 5% milk in phosphate buffered saline, 0.05% Tween 20 (PBST), for 1 h at room temperature. 5. Incubate membrane with rabbit polyclonal antibody against BiP/GRP78 (Stressgen, 1:1000 in 5% milk and PBST) either at 4 C overnight or for 90 min at room temperature. 6. Wash the blots three times with PBST for 5–10 min each time. 7. Incubate with horseradish peroxidase-conjugated secondary antibody (anti-rabbit 1:3000, Sigma) made in 5%milk in PBST for 1 h at room temperature. 8. Wash the blots three times with PBST for 5–10 min each time.

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9. Detect protein by enhanced chemiluminescence (ECL, Pierce Supersignal) using the AlphaInnotech Fluorochem 8900 ChemiImager (see Figs. 1.1 and 1.2). Alternatively, the membranes can be exposed to X-ray film until clear bands are observed after developing the film. 10. Quantify protein bands on the AlphaInnotech ChemiImagerTM using the Alphaease software. If X-ray film is being used, quantify bands using densitometry. We now also use the LI-COR Odyssey infrared imaging system (LI-COR, Inc., Lincoln, NE) for quantitative Western blotting.

CON

SD

BiP

P-Perk

P-eIF2a

eIF2a

Figure 1.1 Composite image showing representative Western blots for BiP, P-PERK, P-eIF2a, and eIF2a in the cerebral cortex of young mice that were sleep deprived (SD) for 6 h. Control (Con) mice were undisturbed during sleep.

Young

Aged

Control

SD

Figure 1.2 Western showing CHOP expression in young and aged control and sleep-deprived samples. CHOP levels are markedly higher in aged animals compared to young.

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The Odyssey uses near-infrared fluorescence detection that allows the quantification of proteins over a much wider linear dynamic range than chemiluminescence can. There are several advantages of using the Odyssey over chemiluminescence. The signal is stable. The Odyssey has a two color detection system with two channels at 700 and 800 nm, respectively. Two antibodies can be simultaneously detected on the same membrane. Strong and weak bands are accurately detected on the same blot without the uncertainty and inconvenience of multiple exposures or time in the darkroom. The disadvantage of using the Odyssey includes having to purchase their patented “unknown” buffers which gets to be very expensive. The Odyssey buffer is used for blocking and diluting theTMprimary and secondary TM antibodies. The secondary antibodies are IRDye 800 and IRDye 680 made against the various host species. 6.1.7. Dissociation of BiP from PERK During the UPR, parallel with the increase of chaperone function is an attenuation of translation to decrease the protein-folding load. During ER stress, decrease in protein biosynthesis occurs when the eukaryotic translation initiation factor 2a is phosphorylated on serine 51 by PERK. PERK is an ER-resident type I transmembrane protein, whose N-terminal lumen domain is sensitive to the upstream ER stress signal and whose C-terminal cytoplasmic domain directly phosphorylates eIF2a (Harding et al., 1999). PERK is maintained in an inactive state by the binding of the ER chaperone BiP/GRP78 to its lumenal domain. Under conditions of ER stress, BiP/ GRP78 dissociates to service the increasing load of ER client proteins; loss of BiP/GRP78 binding correlates with oligomerization, trans-autophosphorylation, and activation of downstream signaling by PERK (Bertolotti et al., 2000). The association of BiP/GRP78 with PERK therefore indicates the state of the cell. We examined the effect of sleep deprivation on the degree to which BiP/GRP78 is associated with PERK. We used an antibody to PERK to co-immunoprecipitate BiP/GRP78 from cortical lysates of sleep deprived and control undisturbed animals using the protocol described below. The immunoprecipitated protein was resolved by 7.5% SDS-PAGE under reducing conditions and transferred to nitrocellulose membranes. Blots were incubated with GRP78 antibody (1:1000); signals were observed using horseradish peroxidase labeled secondary antibody. Protein bands were detected and analyzed by enhanced chemiluminescence (Pierce Supersignal) and quantitative imaging (AlphaInnotech Fluorochem 8900). Densitometry was performed using the Alphaease FC software. Alternatively, IR-conjugated secondary antibodies were used and protein bands were detected and quantified directly by infra red imaging on the Odyssey (LI-COR).


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6.1.8. Immunoprecipitation protocol 1. Use about 100 mg of protein/experiment. 2. Preclear sample with about 10 ml of washed Protein-G Sepharose beads/1 h at 4 C; bead washing is described in step 3. 3. Before use, wash beads with lysis buffer containing protease inhibitor. Wash by adding 300 ml lysis buffer, suspend beads, spin down at 10,000 rpm for 1 min. Remove supernatant but do not allow beads to dry out. Repeat three times. 4. Add 25 ml lysis buffer to homogenates. 5. Add 2 ml of PERK antibody to each homogenate sample—incubate 4 C overnight, with end over end rotation. 6. Incubate 30 ml of washed beads (from step 3) with homogenate– antibody mixture for 1.5 h with end over end rotation. 7. Spin down beads at 10,000 rpm for 1 min. 8. Wash beads three times with 200 ml buffer. 9. Add 2 Laemmli buffer (50 ml), then boil for 2–5 min. 10. Spin down beads, 10,000 rpm for 1 min. 11. Remove supernatant for use in SDS gel electrophoresis. 12. Load 20 ml/well; run gel 13. Use appropriate antibody in western to detect interacting proteins

6.1.9. Immunohistochemistry While detection of the proteolytic processing of ATF6 by westerns works well it can sometimes be tricky to detect the cleavage products. It is advisable to carry out immunohistochemistry to look for nuclear localization of ATF6 as a marker of activation. We also use immunohistochemistry to detect localization of the UPR markers to specific neuronal populations within the mouse cerebral cortex and other brain areas. There is an image of ATF6 localization to the nucleus using immunostaining in Fig. 1.3. Using immunofluorescence and immunostaining with DAB (3,30 -diaminobenzidine), we have found that BiP localizes to the pyramidal neurons of layers I–VI of the medial prefrontal cortex, cingulate cortex, and piriform cortex (Fig. 1.4). We also found increased CHOP staining in these areas only in aged mice (Fig. 1.5). In contrast, BiP is reduced in these same cells in the aged animals. Aged mice also display more GADD34 staining in cortical neurons than do young mice; this is shown in Fig. 1.6. GADD34 is a downstream target of CHOP and is a good marker of a maladaptive response. The protocol below describes both immunofluorescence and immunostaining of free-floating brain sections.

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25 mm

ATF6

Figure 1.3 Immunostaining of ATF6 in piriform cortical neurons. DAB-nickel was used to localize ATF6 in the nucleus of the neurons. 400 magnification. Scale bar, 25 mm.

Young

M1

Old

50 mm

Cg2

Pir

Figure 1.4 Decreased immunohistochemical labeling of BiP in with age. DAB labeled BiP in young and aged cingulate, M1 and piriform cortical neurons. 160 magnification. Scale bar, 50 mm.

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Young

Cg1/M2

Old

100 mm

M1

Pir

Figure 1.5 Increased expression of CHOP with age. DAB labeled CHOP in young and aged cingulate, M1 and piriform neurons. 80 magnification. Scale bar, 100 mm (for all images).

6.1.10. General procedure 1. Procure brain tissue after pentobarbital anesthesia and transcardial perfusion with saline and 4% paraformaldehyde. 2. Incubate whole brains in 4% paraformaldehyde, then cryopreserve in 30% sucrose in PBS containing 0.05% sodium azide. 3. Slice the brain along the coronal axis in 20 or 40 mm sections from the medulla through the basal forebrain approximating murine coordinates 1.54–1.94 mm bregma (Franklin and Paxinos, 1996). 4. Store sections as 1:6 or 1:3 series in 0.1% sodium azide in PBS at 4 C before use. 5. Incubate sections with normal serum for 60–90 min to block nonspecific binding of immunoglobulin. Use the same species’ serum as the secondary antibody is made to.

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2M

24 M

GADD34

GADD34

100 mm

Figure 1.6 GADD34 immunofluorescence in piriform cortex. The left panel shows GADD34 in young cerebral cortex and the right panel shows increased GADD34 immunofluorescence in aged mouse cortex. 200 magnification. Scale bar, 100 mm.

6. Incubate sections with primary antibody at appropriate dilution in blocking buffer for 1 h at room temperature or overnight at 4 C. We use MAP2 antibody (Abcam, 1:1000) as a neuronal marker. For BiP, we use GRP78 (H-129; Santa Cruz) antibody at 1:500, and CHOP is detected using the GADD153/CHOP antibody at 1:500 (sc575; Santa Cruz). 7. Rinse sections 3 10 min in PBS. 8. Incubate sections in secondary antibody diluted in PBS with 2% normal serum for 90 min at room temperature. For fluorescence use Alexa Fluor secondary antibodies diluted at 1:500. For DAB staining, use the appropriate biotinylated secondary antibody (see below). 9. Wash sections in PBS for 30 min in six 5 min intervals. 10. Mount sections on the slide and dry for 30 min in the dark. 11. Coverslip with slow-fade gold antifade reagent with DAPI. 6.1.11. DAB and DAB-nickel staining protocol DAB and DAB-Nickel (DAB-Ni) can be used for double staining of tissue sections. The DAB stain is very stable and immunostained sections can be kept for a long time without fading and loss of signal. Use of DAB results in a brown stain that is enhanced with nickel to produce a dark gray stain. Day 1 1. 2. 3. 4.

Wash sections in PBS for 3 5 min. Treat sections with 3% H2O2 in PBS for 30 min. Wash in PBS for 3 5 min. Incubate in blocking buffer (4% normal serum, 1% BSA, 0.4% Triton X-100 in PBS with 0.1% sodium azide at room temperature for 60–90 min).


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5. Incubate sections in primary antibody diluted in blocking buffer at room temperature for overnight or 4 C for 48 h. Day 2 6. Wash sections in PBS for 6 5 min. 7. Incubate sections in biotinylated appropriate secondary antibody with 2% normal serum,0.5% BSA, 0.4% Triton X-100 in PBS at room temperature for 60 min. (Prepare the ABC reagent buffer. Let it sit at room temperature for at least 30 min before use. 1:1000 in PBS with 0.4% Triton X-100.) 8. Wash in PBS for 6 5 min. 9. Incubate in ABC reagent at room temperature for 60 min. 10. Wash in PBS for 3 5 min. 11. Wash in 0.175 M sodium acetate for 3 5 min. 12. Use DAB-Ni1 to visualize the reaction (from 5 to 20 min). 13. Wash in PBS for 6 5 min. For single labeling, go to step 14; for double labeling, go to step 15. 14. Dehydrate with progressive alcohol at 5–10 min each (70%, 95%, 100%, and 100%) and two times xylene for 10 min each. Mount with permount and coverslip. 15. For double labeling, repeat step 5 with the second primary antibody. Day 3 16. Repeat steps 6–14. (You should perform DAB-Ni staining first and it should be an antibody that localizes to the nucleus and then carry out the DAB staining for the cytosolic marker.)

6.1.12. Polysome analyses Actively translating ribosomes exist as polysomes; during a shutdown or attenuation of translation, a large fraction of the ribosomes is disassembled into monosomes (Doutheil et al., 1997; Gulow et al., 2002; Harding et al., 2000). We carried out polysome analyses to further assess whether protein synthesis was being inhibited after 6 h of sleep deprivation. The microsomal fraction of sleep deprived and control cortical lysates were separated by sucrose gradient centrifugation to yield the ribosomal profiles. The ribosomal profile of samples subjected to 6 h of sleep deprivation show a moderate increase in the monosome peak with a concomitant decrease in the polysomes peak when compared to controls, suggesting an attenuation of translation. For a more detailed description, see the chapter on Polysome Analysis in Methods in Molecular Biology by Sagliocco et al. (2006). 1

DAB-Ni: 2 mg DAB, 1 ml 5% Nickel in 0.175 M sodium acetate to 10 ml, add 8.3 ml 3% H2O2.

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6.1.13. Polysome analysis protocol Cortex tissue from individual mice sleep deprived for 6 h and their matching controls was homogenized in 1 ml of polysome extraction buffer (15 mM Tris–HCl pH 7.4, 15 mM MgCl2, 0.3 M NaCl, 1% Triton X-100, 0.1 mg/ml cyclohexamide, 1 mM PMSF, 2 mg/ml pepstatin, and 4 mg/ml aprotinin). The lysates were cleared by centrifugation at 12,000g for 10 min, and then layered on top of a 4 mL 10–50% sucrose gradient prepared in extraction buffer. After a spin of 2 h at 40,000 rpm in a Sorvall AH650 rotor, 200 ml fractions were collected and the absorbance at 254 nm was read. 6.1.14. Assessment of changes in mRNA for BiP/Q-PCR of BiP In addition to assessing changes in protein levels and activation though phosphorylation, one can also determine transcript levels of the various UPR markers. We examined the alteration of BiP transcript levels using quantitative PCR. I outline below the protocol we used for BiP and provide information about the primer and probe sets used and that can be used for other UPR markers. The expression of the BiP gene in the cerebral cortex C57BL/6J mice during sleep and in sleep deprivation was established by quantitative RT-PCR using the TaqManÒ Gene Expression Assays and 7500 realtime PCR system (Applied Biosystems). Total RNA was isolated with Trizol (Invitrogen) and further cleaned using RNeasy purification kit (Qiagen). cDNA was synthesized using 0.5 mg total RNA and TaqManÒ Reverse Transcription Reagents (Applied Biosystems). The following sets of primers and probes for the relevant genes were used: Mm00517691_m1 (Hspa5/BiP) and Mm00545877_m1 (Apc). Each PCR reaction was repeated in triplicate. The relative transcript level was established by the “DD method” as described previously (Livak and Schmittgen, 2001). The Apc (adenomatosis poliposis coli) gene was used as an internal standard; this gene was identified as unchanged in its expression either during sleep or sleep deprivation in the cerebral cortex and hypothalamus of C57BL/6J mice (Mackiewicz et al., 2007).

7. Concluding Remarks This chapter described techniques and protocols for examining the UPR in whole mouse cerebral cortex. Currently, we know from immunohistochemistry that not all neuronal populations in the mouse brain respond to ER stress in a similar manner (Naidoo et al., 2009; unpublished observation). There are temporal and spatial differences. Use of laser capture microscopy and specific neuronal tags should soon allow one to examine


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the UPR in very specific neuronal populations and discern why certain neurons respond the way they do. This would allow the development of neuron-specific therapeutics for the various diseases ER stress and the UPR are implicated in.

ACKNOWLEDGMENTS I am grateful to Dr. Marishka Brown for critical reading and discussions during the writing of this chapter; Drs. Jingxu Zhu and Yan Zhu for the images and Ms. Jennifer Montoya for assistance with the figures.

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Vasa-Nicotera, M. (2004). The new kid on the block: The unfolded protein response in the pathogenesis of atherosclerosis. Cell Death Differ. 11(Suppl. 1), S10–S11. Vattemi, G., et al. (2004). Endoplasmic reticulum stress and unfolded protein response in inclusion body myositis muscle. Am. J. Pathol. 164, 1–7. Wang, X. Z., et al. (1996). Signals from the stressed endoplasmic reticulum induce C/EBPhomologous protein (CHOP/GADD153). Mol. Cell. Biol. 16, 4273–4280. Wu, J., and Kaufman, R. J. (2006). From acute ER stress to physiological roles of the unfolded protein response. Cell Death Differ. 13, 374–384. Yoshida, H., et al. (1998). Identification of the cis-acting endoplasmic reticulum stress response element responsible for transcriptional induction of mammalian glucose-regulated proteins. Involvement of basic leucine zipper transcription factors. J. Biol. Chem. 273, 33741–33749, [erratum appears in J. Biol. Chem. 1999 Jan 22;274(4):2592]. Yoshida, H., et al. (2001). XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor. Cell 107, 881–891. Zhang, K., and Kaufman, R. J. (2004). Signaling the unfolded protein response from the endoplasmic reticulum. J. Biol. Chem. 279, 25935–25938. Zhang, K., and Kaufman, R. J. (2006). The unfolded protein response: A stress signaling pathway critical for health and disease. Neurology 66, S102–S109. Zinszner, H., et al. (1998). CHOP is implicated in programmed cell death in response to impaired function of the endoplasmic reticulum. Genes Dev. 12, 982–995.


C H A P T E R

T W O

Immunohistochemical Detection of the Unfolded Protein Response in Atherosclerotic Plaques Sˇa´rka Lhota´k,* Ji Zhou,† and Richard C. Austin* Contents 24 24 25 27 27 29 29 30 33 33 40 42 44 44

1. 2. 3. 4.

Endoplasmic Reticulum and the Unfolded Protein Response Atherosclerosis UPR in the Atherosclerotic Lesion Mouse Models of Atherosclerosis 4.1. Lesion morphology 4.2. Lesion size 5. Methods 5.1. Sectioning the aortic root 5.2. Measuring lesion size in serial sections 5.3. Immunohistochemistry 5.4. Double immunofluorescence 5.5. Detection of apoptosis Acknowledgments References

Abstract It is well established that activation of the unfolded protein response (UPR) occurs during all stages of plaque development and progression. Mouse models of atherosclerosis have allowed the study of the UPR in the complex environment of lesions which are composed of different cell types, abundant extracellular matrix, as well as necrotic and apoptotic areas. For proper evaluation of the UPR, the processing, sectioning, and staining of lesions have to be performed in a consistent and reproducible manner. In this chapter, we describe in detail how to embed aortic roots in paraffin, cut serial sections through the aortic root, measure lesion size, and stain paraffin sections by immunohistochemistry, * Department of Medicine and Division of Nephrology, St. Joseph’s Hospital and McMaster University, Hamilton, Ontario, Canada { Department of Medicine, Thrombosis and Atherosclerosis Research Institute (TaARI) and McMaster University, Hamilton, Ontario, Canada Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00002-9

#


23

24

Sˇa´rka Lhota´k et al.

immunofluorescence, and the terminal dUTP nick-end-labeling (TUNEL) technique, with special attention to the proteins involved in the UPR.

1. Endoplasmic Reticulum and the Unfolded Protein Response The endoplasmic reticulum (ER) is the site of synthesis, folding, and modification of proteins destined for secretion or the plasma membrane. ER lumenal-resident chaperones such as the 78-kDa glucose-regulated protein (GRP78), 94-kDa glucose-regulated protein (GRP94), protein disulfide isomerase (PDI), calnexin, and calreticulin assist in the folding process. The ER has a sophisticated mechanism of quality control to prevent misfolding of proteins termed the unfolded protein response (UPR). The UPR consists of three integrative pathways whose sensors (PKR-like ER kinase (PERK), inositol requiring kinase 1 (IRE1), and activating transcription factor 6 (ATF6)) reside in the ER membrane. Their activation leads to (i) the attenuation of general protein translation (through the PERK pathway) and (ii) the increased expression of specific ER chaperones, such as GRP78 and PDI (through the ATF6 and IRE1 pathways). In addition, components of the ERassociated degradation pathway are also activated so as to eliminate the accumulation of misfolded proteins. ER stress refers to the disruption of ER homeostasis (accumulation of misfolded proteins, release of ER calcium) leading to the activation of the UPR. Numerous agents and conditions have recently been identified that induce the UPR during the process of atherogenesis, including free cholesterol, peroxynitrite, homocysteine, and hypoxia (Austin, 2009). Under these conditions, the UPR is designed to alleviate ER stress, thereby restoring correct ER homeostasis. However, if the stress cannot be resolved, the cell is destined for ER stress-induced apoptosis via activation of caspase-12 and downstream caspases. For detailed review on the UPR and its role in apoptotic cell death, see Malhotra and Kaufman (2007).

2. Atherosclerosis Atherosclerosis refers to a disease of large and middle size arteries which is characterized by the formation of complex lesions within arterial walls (reviewed in Lusis, 2000). The first stage during atherogenesis consists of the “fatty streak” which is composed of foam cells. The formation of foam cells is initiated by the presence of modified LDL particles in the subendothelial space which leads to the recruitment of blood-borne monocytes to this site and their differentiation into macrophages. Macrophages ingest LDL particles, esterify the cholesterol, and store the lipid in vacuoles

Immunohistochemistry in Atherosclerotic Lesions

25

giving them their “foamy” appearance. Lesions progressively evolve by recruitment of smooth muscle cells from the media, deposition of extracellular matrix, and apoptosis of foam cells, thereby creating defined regions of cellular necrosis and calcification. Such plaques may become unstable and prone to rupture. Plaque rupture is the most devastating clinical complication of atherosclerosis leading to the formation of a thrombus and resulting in myocardial infarction or stroke.

3. UPR in the Atherosclerotic Lesion In the 1990s, increased plasma homocysteine was identified as a risk factor for cardiovascular disease (Boushey et al., 1995). To explain the mechanism behind this association, we as well as others identified a number of genes upregulated by homocysteine. Using mRNA differential display and cDNA analysis, we discovered in cultured endothelial cells that homocysteine induced the expression of GRP78 and the growth arrest and DNA damageinducible gene 153 (GADD153) (Outinen et al., 1998). Around that time, the unfolded protein pathways were being elucidated and these proteins were known to be induced by stress to the ER (Malhotra and Kaufman, 2007). To translate our findings to an in vivo model of atherosclerosis, we utilized a mouse model of hyperhomocysteinemia to assess the induction of the UPR in lesion resident cells (Zhou et al., 2001). In collaboration with Dr. Erling Falk and colleagues, we showed that increased plasma homocysteine levels in apoE/ mice supplemented with methionine led to increased lesion size (Zhou et al., 2001). Using immunohistochemistry, we found that the staining intensities for markers of ER stress, namely the ER chaperones and phosphoPERK, were increased in the lesions of hyperhomocysteinemic mice compared with controls (Zhou et al., 2004). Surprisingly, positive immunostaining for UPR markers was also observed in the aortic root lesions at all stages of lesion progression in control apoE/ mice (Zhou et al., 2005). The association between the UPR and atherosclerosis has now been clearly established (for review, see Austin, 2009; Marciniak and Ron, 2006). Currently, UPR-related research is focused in two main areas: ER stress and activation of the UPR in (i) atheroprone regions of the blood vessel (Civelek et al., 2009; Dickhout et al., 2005; Feaver et al., 2008; Sanson et al., 2009), as well as (ii) apoptotic cell death and plaque stability, which are crucial to the prevention of devastating clinical events. ER stress markers were found to be increased in unstable plaques in human carotid lesions (Dickhout et al., 2007; Myoishi et al., 2007). In both human and mouse lesions, the UPR is observed predominantly in macrophages and macrophage foam cells (Fig. 2.1). This is consistent with in vitro findings showing that cholesterol loading of macrophages causes UPR activation and

26


GRP78

p-PERK

A

I M

D

GADD153

B

C

E

F

A

I

A

M

Figure. 2.1 Endoplasmic reticulum stress in early and advanced lesions from apoE/ mice. Immunohistochemical staining for GRP78, phospho-PERK, and GADD153 is shown in early (A–C) and advanced (D–F) lesions. GRP78 was found in foam cells in the lesions and some cells in adventitia in both stages of the lesion progression (A, D). Phospho-PERK staining was also seen in the foam cells; however, only some of the cells were strongly expressing the protein (B, D), whereas others were devoid of the signal, most likely reflecting the temporary character of the protein phosphorylation. Few of the smooth muscle cells in the media were also positive (B). GADD153 strongly labeled nuclei in the foam cells in early lesions (C, arrows) and in the lumenal portion of the advanced lesion (F, arrows). I, intima; M, media; A, adventitia. Bar ¼ 50 mm.

subsequent apoptosis (Feng et al., 2003). Currently, macrophage apoptosis is under intense study given its direct relevance to plaque growth and stability (Han et al., 2006; Tabas et al., 2009, 2010; Tsukano et al., 2010). The complexity of atherosclerotic lesions and the interplay between its pathways and components dictate that research should be carried out using in vivo models. Immunohistochemistry is an excellent tool for studying the expression of various proteins in specific cells within the lesions. An example of the UPR-related immunostaining in early and advanced lesions from apoE/ mice is presented in Fig. 2.1. The ER chaperone GRP78 is detected in all macrophage foam cells in the early lesion (Fig. 2.1A) and on the lumenal side of the neointima in the advanced lesion (Fig. 2.1D). Only a few cells in the necrotic region of the neointima displayed any staining. Phospho-PERK staining was observed in the macrophage foam cells in both early and advanced lesions (Fig. 2.1B, E); however, the pattern of staining was different than that for GRP78. Some of the cells had intense staining while others were devoid of staining. We believe that this represents the transient activation of PERK in association with an increase in GRP78 expression. GADD153, one of the components of ER


27

stress-induced apoptosis (Oyadomari and Mori, 2004), is strongly expressed in the nuclei (and less so in the cytoplasm) of foam cells in both early and advanced lesions (Fig. 2.1C, F).

4. Mouse Models of Atherosclerosis None of the in-bred strains of mice spontaneously develop atherosclerosis. Only small lesions consisting of foam cells can be induced by modified diets over long periods of time in some strains (Whitman, 2004). These are of limited use because the lesions do not progress to advanced and complicated lesions analogous to those seen in humans. A major advance in using mouse models to study atherosclerosis was triggered by the generation of apoE-deficient (apoE/) and LDL receptor-deficient (ldlr/) mice in 1992 and 1994, respectively (Ishibashi et al., 1994; Piedrahita et al., 1992; Plump et al., 1992). ApoE/ mice develop lesions spontaneously on chow diet, while ldlr/ mice require high cholesterol diet. In both models, lesions progress from simple fatty streaks composed of lipid laden macrophages to complex lesions with necrotic cores, extracellular lipid deposition, and fibrous cap formation, thus resembling the progression in human lesions. These mice have been crossed with other knockout or transgenic mice to study the role of various genes in lesion progression and stability. For designing an experiment using atheroprone mice, there are a number of helpful reviews summarizing the type of lesions, age, gender, diet, and genetic background (Daugherty, 2002; Daugherty and Rateri, 2005; Daugherty et al., 2009; Whitman, 2004).

4.1. Lesion morphology In all mouse models, lesions start in the atheroprone region of the aortic root. With time and increased hyperlipidemia, these lesions increase in size and complexity with new lesions forming in the aortic arch and the thoracic and abdominal aorta, predominantly around the ostia of large arteries. Early lesions are composed of macrophage foam cells. Morphologically, these are large cells filled with lipid droplets (Fig. 2.2A) and can be identified by immunostaining with a macrophage marker (Fig. 2.3). The morphology of advanced lesions is complex. The lumenal side is typically comprised of macrophage foam cells interspersed with smooth muscle cells forming the cap next to the endothelium. Below this layer, there are areas of apoptosis (dense fragmented nuclei) as well as acellular areas characterized by the lack of nuclei and extracellular matrix with cholesterol clefts (Fig. 2.2B). In early lesions, neutral lipids (cholesterol esters and triglycerides, stained with Oil Red O or Sudan IV) and free cholesterol (stained with filipin) can be

28


L

B

A

FC

L A M M

Figure. 2.2 Morphology of early and advanced lesions. Only a few foam cells are seen close to the valve leaflet in an early lesion (arrows, outline in green, A). Advanced lesion (outlined in green, B) consists of a cellular area on the lumenal side formed predominantly by foam cells (FC) and an acellular area devoid of nuclei (A). Cholesterol clefts (arrows) and apoptotic bodies (arrowheads) are embedded in extracellular matrix. L, lumen; M, media; FC, foam cells; A, acellular area; arrows, cholesterol clefts; arrowheads, apoptotic bodies. Bar ¼ 50 mm.

Mac-3 A

SMA B

vWF C

M

D

E

F

M

Figure. 2.3 Cellular composition of atherosclerotic lesions. Lesions are composed mostly of macrophage cells (stained here with the Mac-3 antibody, A, D) overlaid by a thin layer of endothelium (stained with von Wilebrandt factor (vWF), C, F). Smooth muscle cells are rarely seen in early lesions; staining is seen only in the media (M, B). In advanced lesions smooth muscle cells are present in the cap of the lesion (arrows, E), in addition to the media. SMA, smooth muscle actin. Bar ¼ 50 mm.


29

identified in the macrophage foam cells. In advanced lesions, the lumenal side of the lesion stains for both neutral lipids and free cholesterol (not shown). However, the acellular necrotic core does not stain for neutral lipids. Thus, using neutral lipid staining on cryosections to highlight lesions for measurement (as is sometimes seen in literature) will underestimate lesion size.

4.2. Lesion size Several reviews describe how to assess and measure lesions (Daugherty and Rateri, 2005; Daugherty et al., 2009). In the early stages, lesions are only present in the aortic root. However, in more advanced cases of atherosclerosis where lesions are present throughout the aortic tree, lesions can be measured either on sections through the aortic root (paraffin or cryosections), as in early lesions, or in en face whole mount preparations of the aortic arch or the whole aortic tree. It was reported that lesion sizes in the aortic root, aortic arch, and the aortic tree correlate well in several different genetic models (Tangirala et al., 1995); therefore, one method can substitute for another. Due to a large variance between animals, it is recommended to use two methods that complement each other, such as serial sections through the aortic root and the en face preparation of the aortic tree. Both can be assessed from a single mouse. In en face preparations, lesions are stained with a neutral lipid stain (Oil Red O or Sudan IV) and the stained area is measured by an image analysis software and expressed as percentage of the total intimal area (Tangirala et al., 1995). For sectioning, the aortic root may be embedded in OCT and frozen for cryosectioning (detailed protocol in Baglione and Smith, 2006) or processed into paraffin for microtomy. Cryosections are preferred if lipid analysis is of importance, as in paraffin blocks, the lipid is extracted during tissue processing. Paraffin sectioning, however, allows for a more precise control of the angle of sectioning and easier collection of serial sections, in addition to easier storage of blocks and sections. Lipid analysis can still be performed in the corresponding en face whole mount preparation.

5. Methods We describe methods for embedding hearts with aortic roots in paraffin and serial sectioning the aortic root. Some of the serial sections are stained with H&E and used to measure lesion size (see Section 5.2). The rest may be used to study various components of the lesion such as extracellular matrix components or apoptosis, or for immunodetection of antigens of interest, such as cell type markers, markers of UPR activation, and others (see Section 5.3).

30


5.1. Sectioning the aortic root 5.1.1. Embedding the heart To obtain reproducible and comparable data, the lesions have to be sectioned perpendicular to the aortic root. If the aortic root is sectioned obliquely, the lesion size will be overestimated. Second, a starting point for measurements has to be anatomically defined. To aid in this, the following procedure is recommended. The heart is dissected from the mouse with 1–2 mm of the ascending aorta attached and fixed in formalin for 24 h. It is then cut transversally as described in Paigen et al. (1987), along the line A that connects the tips of the atria (Fig. 2.4). (It is helpful to do this under a dissecting microscope.) Line A is parallel to the aortic root (line B, Fig. 2.4B) and the correct angle ensures that the sections through the aortic root will be perpendicular to the aorta. The heart is then processed and embedded in paraffin, the cut face down, using deep molds (Ted Pella, Inc. #27195). Despite great care taken to cut the heart along the line A, this is not always achieved perfectly. However, it can be rectified during sectioning. We use a method that allows finding the perpendicular cut through the aortic root based on the landmarks observed in unstained sections during the sectioning process and correcting the angle by tilting the block. To allow this, the tissue must be embedded in deep molds, and for tilting at more extreme angles, a specimen holder like the one in Fig. 2.5 (e.g., Leica #7998) is useful.

A

B RA AA

LA

B A

A L R

Figure. 2.4 Preparing mouse heart for paraffin embedding. Formalin-fixed heart is positioned with the ascending aorta (AA) curving to the left. The heart should be cut along the line A connecting the tips of the atria (A). In B, the dorsal part of the heart was removed to expose the aortic valve (arrow) which lies in the plane of line B. AA, ascending aorta; L, left ventricle; R, right ventricle; LA, left atrium; RA, right atrium.


A

31

B

Figure. 2.5 (A) Paraffin block and microtome clamp. Heart is embedded with the cut surface down in a deep mold creating a high paraffin block. A special microtome clamp is used that allows the block to be positioned at an angle inside the clamp. The whole assembly is cooled on ice during serial sectioning (B).

5.1.2. Orienting the block Sectioning is commenced at ambient temperature, moving quickly from inside toward the top of the heart with section thickness set at 8 mm. Occasionally an unstained section is collected for microscopic inspection (Fig. 2.6). When the mitral valves are observed (Fig. 2.6A), it is time to slow down and proceed carefully. From this point, every 10–12th unstained section is inspected under the microscope (Fig. 2.6B and C). At the first sign of the aortic valve leaflets (Fig. 2.6C, arrows), a decision has to be made regarding the orientation of the block. In Fig. 2.6C, two valve leaflets are visible (arrows), the third one, however, is missing (asterisk). Note also that the aortic wall in the sinus has a different morphology than cardiac muscle; layers of smooth muscle are seen in the aorta. The top side of the image (Fig. 2.6C) is still formed by cardiac muscle. Therefore, this side has to be brought closer to the knife and sectioned through until the third leaflet comes into view. (Note that in the microscope the image is inverted. We will be actually bringing the lower side of the block toward the knife. It is important to know which side in the image corresponds to which side of the section and the block.) The clamp assembly with the block may be tilted using knobs on the microtome. For larger angles, the block itself may be oriented in the clamp at an angle. After tilting, the block is sectioned through empty paraffin until the tissue is reached. Every other section (8 mm thick) is now captured and inspected under a microscope. The tissue will now be at the edge of the section (Fig. 2.6D, arrow). In Fig. 2.6, after tilting full angle with the microtome adjusting knobs, the third leaflet came into view (Fig. 2.6D). A little further, all three leaflets were visible, spaced evenly around the aorta (Fig. 2.6E). At this point, the serial sectioning begins.

32


A

B

C

D

E

R

L

Figure. 2.6 Finding the perpendicular cut through the aortic root. Upper panel: unstained sections as they present during the sectioning process. The area in the rectangle is shown in the lower panel, stained and enlarged, to better demonstrate landmark features: mitral valve, arrow (A); aortic valve leaflets, arrows, (C) a missing leaflet, asterisk (C). At this stage the block has been tilted and sectioned, new edge of the block, arrow (D). The third leaflet is coming into view, asterisk (D). All three leaflets are visible, spaced equally, double headed arrows (E). R, right ventricle; L, left ventricle.

5.1.3. Serial sectioning Serial sections are cut at 4 mm thickness, and for that, the block has to be cooled. To preserve the angle, the whole clamp assembly with the block is removed and placed with the cut face down on an ice tray (Fig. 2.5B). After reinserting in the microtome, serial sections are cut at 4 mm and floated in a warm water bath. After 10–15 sections, the block has to be cooled again. Sections are collected on numbered Plus slides (#12-55015, Fisher Scientific). The first 20 sections are collected and placed in the top portion of slides 1–20. For the section #21, we go back and place it in the middle portion of slide 1, section #22 on the slide 2, etc. Section #41 will be placed in the lower part of slide 1, and so on until section #60. Sections #61, 81, and 101 are placed on a new slide. Thus the three sections on each slide are 80 mm apart along the aortic root (4 mm section thickness times 20). Lesions can usually be seen in unstained slides. Sectioning and collecting sections continues until there are no more lesions present, often 100 sections or more for larger lesions. An example of serial sections through aortic root with early or advanced lesions is shown in Fig. 2.7.

33


Section #: Microns

1 0

21 80

41 160

61 240

81 320

6.5

2.4

0

0

387.2

261.4

200.6

37.6

9 weeks

Area (×103 mm2) 27.7

23 weeks

Area (×103 mm2) 355.7

Figure. 2.7 Serial sections from early and advanced apoE/ mice on chow diet. Five sections spaced by 80 mm along the ascending aorta are presented for early lesions (upper panel) and advanced lesions (lower panel). The point of origin for measurement was defined at section #1. The lesion size for each image is shown (the measurement was performed at a higher magnification of the same section).

5.2. Measuring lesion size in serial sections The starting point for lesion measurements needs to be consistent and is usually set at the leaflet confluence (such as sections #1 in Fig. 2.7). Lesions should be measured in several sections along the aortic root and ascending aorta to account for the lateral length of the lesion as well as its thickness. It is suggested to serial section the aortic root and ascending aorta as far as lesions are observed and measure the lesion area on sections at 80 mm intervals. The mean lesion area in a given number of sections can then be used, or by plotting the lesion area versus distance from the leaflet origin and determining the area under the curve, the actual volume of the lesion is obtained (Fig. 2.8) (Daugherty and Rateri, 2005; Daugherty et al., 2009; Paigen et al., 1987; Purcell-Huynh et al., 1995). Various image analysis packages may be used to measure the areas of manually outlined lesions (Fig. 2.2). An example of sections selected for the measurement is presented in Fig. 2.7. Five sections each from two apoE/ mice on chow diet, one at 9 weeks of age (early lesions), the other at 23 weeks of age (advanced lesions), are shown. The sections are 80 mm apart along the aortic root. Lesion areas in each section are shown. In Fig. 2.8, these lesion areas were plotted against the distance from the origin.

5.3. Immunohistochemistry Immunohistochemistry on paraffin sections has proven to be a reliable tool for studying atherosclerotic lesions in mouse models. A general protocol for horseradish peroxidase substrate staining will be described. Particulars for

34


Lesion area in the aortic root 450,000 400,000 Lesion area (mm2)

350,000 300,000 250,000 200,000 150,000 100,000 50,000 0 0

100 200 300 Distance from the origin (mm) 23 weeks

400

9 weeks

Figure. 2.8 Lesion areas along the aortic root and ascending aorta. Lesion areas from Fig. 2.7 were plotted against the distance along the aortic root and ascending aorta. Area under the curve represents lesion volume.

some antibodies of interest are shown in Table 2.1, and suggestions for troubleshooting and optimizing new antibodies are included in Section 5.3.5. 5.3.1. Controls Every immunostaining experiment should be designed and performed with positive and negative controls in place. A positive control would include a tissue where the protein expression is expected. For UPR activation, a good positive control is the pancreas where ER chaperone expression is seen in the islets of Langerhans under normal physiological conditions (Harding et al., 2001; Iwawaki et al., 2004) (Fig. 2.9A–C). For a negative control, omitting the primary antibody is the simplest. For a more rigorous negative control, normal immunoglobulins of the species in which the primary antibody was raised, diluted to the same concentration as the primary antibody, are used. The most convincing negative control is using mouse tissue in which the protein of interest is genetically ablated (Fig. 2.10). 5.3.2. Materials and solutions

Various readymade immunohistochemistry kits may be used. We prefer using individual reagents which reduces cost and gives us flexibility to work with a range of antibodies from different species. Humidity chamber for staining

Table 2.1 Antibodies used for immunohistochemical evaluation of the UPR and apoptosis Blocking

Primary antibody Used at Dilution Antigen retrieval

Serum

Biotinylated secondary

Goat 5 Mouse 2

1:40 1:500

– –

Rabbit Goat

Rabbit anti-goat Goat anti-mouse*

Rabbit 5

1:40

Goat

Goat anti-rabbit

Stressgen #SPA-891 Stressgen #SPA-600 Cell signaling #3191þ Pharmingen #55322

Mouse Rabbit Rabbit Rat

2 Unknown Unknown 0.5

1:500 1:500 1:250 1:1000

HIER-Retrieve-all 2, Triton X – – – HIER-citric

Goat Goat Goat Rabbit

Mac-3

Pharmingen #550292

Rat

0.34

1:90

HIER-citric

Rabbit

Von Willebrand factor Smooth muscle actin Cleaved caspase-3 CD3

DakoCytomation A 0082

Rabbit 6

1:500

Protease

Goat

Goat anti-mouse* Goat anti-rabbit Goat anti-rabbit Rabbit anti-rat, mouse adsorbed Rabbit anti-rat, mouse adsorbed Goat anti-rabbit

Sigma A2547 Cell signaling #9661 DakoCytomation A 0452

Mouse Unknown 1:200 Rabbit Unknown 1:100 Rabbit 6 1:100

– HIER-citric HIER-citric

Goat Goat Goat

Goat anti-mouse* Goat anti-rabbit Goat anti-rabbit

Antigen

Source

Host

GRP78 KDEL GADD153

Santa Cruz sc-1050 KDEL Stressgen #SPA827 Santa Cruz sc-575

PDI Calreticulin Phospho-PERK Mac-3

mg/ml

* Alternatively, anti-mouse Dako EnVision #K4000 HRP conjugate may be used.

KDEL

PDI

A

B

Calreticulin C

I

I

I

p-PERK D

GRP78 E

Combined F

Figure. 2.9 Pancreas as a positive control for endoplasmic reticulum (ER) chaperone immunostaining. Various levels of staining for ER chaperones (GRP78 plus GRP94, KDEL, A; PDI, B; and calreticulin, C) were observed in endocrine cells within the islets of Langerhans (I). Occasionally, strong staining was also seen in an exocrine cell (arrowheads, A and C). Double immunofluorescence revealed that phospho-PERK and GRP78 staining were mutually exclusive in the islets (D–F). Cells with high levels of GRP78 exhibited a weak p-PERK staining (arrow) and vice versa (arrowhead). I, islet of Langerhans. Bar ¼ 50 mm. SR-B1 −/−ApoE −/−

SR-B1 +/+ ApoE −/− A

B

C

D

Figure. 2.10 Tissue from a gene-ablated mouse used as a negative control for immunohistochemistry. In tissues from the apoE/ mouse, scavenger receptor B1 (SR-B1) expression was observed in plasma membranes of hepatocytes (A) as well as cells in the adrenal gland (C). The staining was absent in the liver (B) and the adrenal gland (D) of the double knockout SR-B1//apoE/ mouse. Bar ¼ 50 mm.

37


Biotinylated secondary antibodies: Anti-goat BA-5000, anti-mouse BA9200, anti-rabbit BA-1000, anti-rat, mouse adsorbed, BA-4001, Vector Laboratories. HRP-streptavidin, 20 concentrate, cat. # 50-242Z, Zymed, Invitrogen Nova Red (HRP substrate) cat. # SK-4800, Vector Laboratories Gill’s hematoxylin (Sigma–Aldrich) Permount (Fisher Scientific) Superfrost Plus slides (#12-550-15 Fisher Scientific) Rice steamer Tris buffer 20 Tris (hydroxymethyl) methylamine NaCl

121.1 g 170.0 g

Add distilled water Adjust pH to 7.6 with approximately 50–60 ml concentrated HCl Adjust final volume to 1000 ml with distilled water Tris buffer, working solution Dilute Tris buffer 20 with distilled water 5.3.3. General protocol Particulars for each antibody, such as antigen retrieval, blocking serum, and primary and secondary antibody information are listed in Table 2.1. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Sections are dried at room temperature overnight. Sections are deparaffinized in three changes of xylene, 10 min each. Three changes of 100% ethanol, 1 min each Endogenous peroxidase block: 60 ml methanol þ 1 ml 30% H2O2 þ 4 drops of concentrated HCl, 10 min Wash in 70% ethanol, then distilled water three times Perform an antigen retrieval if required (Table 2.1 and Section 5.3.3.1). Rinse with Tris buffer. Move slides to the humidity chamber. Slides are now in horizontal position; avoid drying. Block with 5% normal serum (see Table 2.1). Flick off excess blocking serum; wipe a rectangular area around the tissue with Kleenex. This will keep the primary antibody from spreading over the slide and drying. Incubate with primary antibody diluted in the 5% normal serum, for 1–2 h, or overnight at 4 C. Wash with Tris buffer 2.

38


13. Incubate with biotinylated secondary antibody for 30 min.1 14. Wash with Tris buffer 2. 15. Incubate with streptavidin-peroxidase, diluted 1 drop in 2 ml of Tris buffer, for 10 min. 16. Wash with Tris buffer. 17. Wash with distilled water. 18. Prepare Nova Red solution according to manufacturer’s directions. 19. Incubate for 2–15 min, observing the reaction through microscope on the positive control or a section that is expected to be positive. 20. Wash with distilled water. 21. Counterstain with Gills hematoxylin No. 3, 30 s. 22. Wash with tap water. 23. Dehydrate in two changes of 100% ethanol and two changes of xylene. Use different xylenes and ethanols than for deparaffinizing. Avoid lower concentration of ethanol, go directly to 100% ethanol, and move quickly through ethanols and xylenes. Nova Red staining sometimes tends to disappear in ethanols. 24. Mount with Permount.

5.3.3.1. Antigen retrieval techniques Heat-induced epitope retrieval (HIER) Preheat appropriate buffer in a plastic slide container in rice steamer for 25 min Place slides in the preheated buffer, let steam for 30 min Remove the container from rice steamer, let cool down in the same solution for additional 30 min

Citric buffer, pH 6.0 Dissolve 2.3 g of citric acid in 1000 ml of distilled water. Adjust pH to 6.0 with 2 N NaOH ( 13 ml)

Retrieve-all Antigen Unmasking System 2: Basic pH 10, 1 (SIGNET) #SIG-31922, Covance Protease digestion Dissolve 0.025 g of protease (Streptomyces griseus, #P6911, Sigma) in 50 ml of PBS Treat slides for 5 min at room temperature Wash with PBS, then Tris buffer

1

Alternatively, for mouse primary antibodies, anti-mouse Dako EnVision #K4000 may be used diluted 1:3 in Tris buffer. Incubate for 30 min. It is an HRP conjugate, therefore, go directly to step 16.


39

Triton X treatment 0.1% Triton X in PBS, incubate for 10 min 5.3.4. Helpful hints and pointers Helpful hints and pointers leading to convincing and visually appealing immunostaining

Never let slides dry during the procedure. Change xylenes often. We believe that the remains of undissolved paraffin inhibit the peroxidase reaction. After six racks of slides the last xylene should be replaced with a clean one. If a staining that used to give reliable results in the past stops working, a good rule of thumb is to replace xylenes. Use different set of alcohols and xylenes for deparaffinizing and for dehydrating before mounting with Permount. After each incubation, wash extensively with squeeze bottle pointed at the tissue and sweeping around it with considerable pressure. Expect to use 4 l of Tris buffer for staining 20 slides. Mouse monoclonal antibodies may produce mouse-on-mouse (MOM) background on mouse tissue conspicuous over blood vessels, calcified areas and elsewhere caused by the secondary antibody binding to mouse epitopes in the tissue. An alternative is to use anti-mouse Dako EnVision þ System-HRP Labelled Polymer #K4000 which reduces the problem. Another alternative is MOM kits supplied by various companies. 5.3.5. Designing an immunostaining protocol for a new antibody Before purchasing a new antibody, it is useful to conduct a literature search for immunohistochemical staining because not every antibody will work on formalin-fixed paraffin-embedded tissues. Manufacturers’ datasheets sometimes indicate applications tested though this information and pictures presented are not always convincing. A publication showing clear immunostaining will also provide information on antigen retrieval, concentration, type of tissue. Pilot experiments should be performed on a positive tissue (see Section 5.3.1) to test several primary antibody concentrations in the range of 1–10 mg/ml. If the antibody concentration is not known, a good rule of thumb is to use it 10 times more concentrated than recommended for Western blotting. The first run should include three sets of slides: without antigen retrieval, with protease digestion, and with HIER. In each set, a negative control and one or two primary antibody concentrations in the 1–10 mg/ml range are included. After assessing results from this run parameters may be fine-tuned. If the slide with no antigen retrieval shows staining where it was expected, antigen retrievals are not necessary and should be avoided. Though they may be necessary for some antibodies, with others they can induce

40


nonspecific staining. HIER is particularly problematic: nonspecific nuclear staining is often observed. After the appropriate antigen retrieval has been determined the staining can be further optimized. Background (haze over all cell types) may be reduced by diluting the primary antibody. Several dilutions should be tested.

5.4. Double immunofluorescence On paraffin-embedded tissues, peroxidase staining is preferable to immunofluorescence because it allows better orientation in the tissue. However, when colocalization of two antigens is needed, double immunofluorescence is the method of choice and can be performed on paraffin sections. For example, in Fig. 2.11, cytoplasmic staining of GRP78 and nuclear staining of GADD153 is shown in foam cells in an advanced lesion. 5.4.1. Controls The antibodies should first be optimized by the peroxidase protocol (see Section 5.3.3) to determine appropriate antigen retrievals and primary antibody concentrations. Both antibodies should work well on the same tissue using the same antigen retrieval. In addition, they should be raised in two different species.

GRP78

GADD153 B

A

Combined C

A FC

L

Figure. 2.11 Double immunofluorescence for GRP78 and GADD153 in advanced lesions. Nuclei in the foam cell-rich region were positive for GADD153 (green, A). Red cytoplasmic staining for GRP78 was observed in these cells (B), merged image (C). L, lumen; FC, foam cells; A, acellular area. Bar ¼ 50 mm.


41

5.4.2. Materials Alexa fluorophores are stable dyes with high intensity of fluorescence available in various colors and conjugated to secondary antibodies (Invitrogen). The colors have to be selected based on the microscope available. Alexa Fluor 488 is excited by blue light and fluoresces green; excitation and emission are in the same ranges as fluorescein (FITC). All fluorescent microscopes are equipped with this filter. For orange or red color, Alexa Fluor 546 or 594 (corresponding to rhodamine or Texas Red) can be used, based on the filter available in the microscope.

Alexa Fluor secondaries raised in donkey (e.g., donkey anti-rabbit Alexa Fluor 488, donkey anti-goat Alexa 594, Invitrogen) Normal donkey serum Permafluor mountant (aqueous, Fisher) DAPI (40 ,6-diamidino-2-phenylindole), 10 mg/ml stock solution (D-9564, Sigma) 5.4.3. General protocol 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

Paraffin sections 4 mm thick are dried at room temperature overnight. Sections are deparaffinized in three changes of xylene, 10 min each. Three changes of 100% ethanol, 1 min each 70% ethanol, then distilled water, 3 Perform antigen retrieval if necessary (see Section 5.3.3.1 and Table 2.1). Wash with Tris buffer, 2. Block with 5% normal donkey serum in Tris, 10 min. In a humidity chamber, incubate with the cocktail of the two antibodies, diluted in 5% normal donkey serum, for 1 h at RT. Wash with Tris buffer, 2. Incubate with a mixture of secondary Alexa-conjugated antibodies, diluted 1/200 in Tris. Wash with Tris buffer. Wash with distilled water. Counterstain with DAPI (stock solution diluted 5000 in PBS), 5 min. Wash with distilled water. Mount with Permafluor.

5.4.4. Hints and pointers We do not find it necessary to protect slides from light during staining with Alexa Fluor conjugates. However, slides should be imaged and photographed the same day or within 24 h of staining because, with time, the signal diminishes and the background and autofluorescence increase. When presenting results, each color should be shown separately in addition to the merged image.

42


5.5. Detection of apoptosis Apoptosis is observed in more advanced lesions in mouse models. It is visible on H&E sections as dark condensed nuclei and apoptotic bodies (Fig. 2.2B). It can be detected by the terminal dUTP nick-end-labeling (TUNEL) nuclear staining or by immunostaining for cleaved caspase-3 (see Section 5.3 and Fig. 2.12D). TUNEL assay detects DNA fragmentation. It uses an enzyme, the terminal deoxynucleotidyl transferase (TdT), to insert labeled dUTP at the nicks in DNA. The thymus serves as a good positive control tissue due to abundance of apoptotic thymocytes (Fig. 2.12B–D). For a negative control, the TdT enzyme is omitted from the labeling mixture. In lesions, a false-positive TUNEL staining is sometimes detected in calcified areas. This can be avoided by pretreatment of sections with 3% citric acid for 1 h (Kockx et al., 1996). 5.5.1. TUNEL assay We use the TACSÒ 2 TdT In Situ Apoptosis detection kit (Trevigen). The individual components of the kit can be purchased separately. The dUTP is biotin labeled; therefore it can be used either for an HRP type of staining,

A

B

D

C

Figure. 2.12 Detection of apoptosis. TUNEL staining was detected over apoptotic bodies in advanced lesion from 23-week-old apoE/ mouse on chow diet (arrow, A). The thymus served as a positive control tissue. A number of thymocytes were stained (B), whereas no staining was observed in the control section where the TdT enzyme was omitted from the labeling mixture (C). Fluorescence was used to colocalize TUNEL staining with cleaved caspase-3 immunofluorescence in the thymus. Some of the thymocytes displayed both the red nuclear TUNEL staining and the green cytoplasmic cleaved caspase-3 staining (arrows, D) identifying them as apoptotic cells. Bar ¼ 50 mm.

43


using streptavidin–HRP as in the immunostaining protocol (see Section 5.3.3), or for fluorescence, using a streptavidin-conjugated Alexa dye. For combined immunofluorescence and TUNEL, the TUNEL protocol is followed by blocking with a blocking serum and incubation with the antibody of interest (Fig. 2.12D). If antigen retrieval is required, it should be performed first. 5.5.1.1. Materials Cat# 4810-30-CK TACS 2 Core kit (includes 10 TdT labeling buffer, 10TdT Stop buffer, TdT dNTP Mix, TdT Enzyme) Individual components of the kit may be purchased separately Proteinase K Cat# 4810-30-01 5.5.1.2. TUNEL protocol 1. Sections are deparaffinized in three changes of xylene, 10 min each. 2. Three changes of 100% ethanol, 1 min each. 3. 70% ethanol, 3 distilled water. 4. PBS wash, 2 5 min 5. Move slides to the humidity chamber. Slides are now in horizontal position; avoid drying. 6. Incubate with proteinase K (1 ml pK þ 50 ml deionized water/slide), 30 min, 37 C. 7. Wash 2 with deionized water 2 min each. 8. Quench endogenous peroxidase: 45 ml methanol þ 5 ml 30% H2O2, in Koplin jar, 5 min 9. PBS wash, 1 min 10. Prepare 1 labeling buffer: 45 ml water þ 5 ml 10 labeling buffer 11. Prepare labeling mixture (/slide):

Labeling buffer TdTdNTP mix 50 Co2þ TdT enzyme

50 ml 1 ml 1 ml 1 ml

For negative control, omit the TdT enzyme. 12. 13. 14. 15. 16.

Incubate 1 h at 37 C. Nucleotides are biotin labeled. Prepare 1 stop buffer: 45 ml water þ 5 ml 10 stop buffer. Stop the reaction by incubating in stop buffer for 5 min. Wash 2 in PBS, 2 min each. Perform steps 14–24 of the immunostaining protocol (Section 5.3.3) or steps 10–15 of the immunofluorescence protocol (Section 5.4.3) using a streptavidin–Alexa conjugate, diluted 1/200 in PBS buffer.

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ACKNOWLEDGMENTS We thank Dr. Bernardo Trigatti for providing us with tissues from the SR-B1 knockout mice. This work was supported in part by research grants to Richard Austin from the Heart and Stroke Foundation of Ontario (T-6146, NA-6393) and the Canadian Institutes of Health Research (MOP-74477). Financial support from St. Joseph’s Healthcare Hamilton is also acknowledged. Richard C. Austin is a Career Investigator of the Heart and Stroke Foundation of Ontario and holds the Amgen Canada Research Chair in the Division of Nephrology at St. Joseph’s Healthcare and McMaster University.

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Ishibashi, S., Goldstein, J. L., Brown, M. S., Herz, J., and Burns, D. K. (1994). Massive xanthomatosis and atherosclerosis in cholesterol-fed low density lipoprotein receptornegative mice. J. Clin. Invest. 93, 1885–1893. Iwawaki, T., Akai, R., Kohno, K., and Miura, M. (2004). A transgenic mouse model for monitoring endoplasmic reticulum stress. Nat. Med. 10, 98–102. Kockx, M. M., Muhring, J., Bortier, H., De Meyer, G. R., and Jacob, W. (1996). Biotin- or digoxigenin-conjugated nucleotides bind to matrix vesicles in atherosclerotic plaques. Am. J. Pathol. 148, 1771–1777. Lusis, A. J. (2000). Atherosclerosis. Nature 407, 233–241. Malhotra, J. D., and Kaufman, R. J. (2007). The endoplasmic reticulum and the unfolded protein response. Semin. Cell Dev. Biol. 18, 716–731. Marciniak, S. J., and Ron, D. (2006). Endoplasmic reticulum stress signaling in disease. Physiol. Rev. 86, 1133–1149. Myoishi, M., Hao, H., Minamino, T., Watanabe, K., Nishihira, K., Hatakeyama, K., Asada, Y., Okada, K., Ishibashi-Ueda, H., Gabbiani, G., Bochaton-Piallat, M. L., Mochizuki, N., et al. (2007). Increased endoplasmic reticulum stress in atherosclerotic plaques associated with acute coronary syndrome. Circulation 116, 1226–1233. Outinen, P. A., Sood, S. K., Liaw, P. C., Sarge, K. D., Maeda, N., Hirsh, J., Ribau, J., Podor, T. J., Weitz, J. I., and Austin, R. C. (1998). Characterization of the stressinducing effects of homocysteine. Biochem. J. 332(Pt. 1), 213–221. Oyadomari, S., and Mori, M. (2004). Roles of CHOP/GADD153 in endoplasmic reticulum stress. Cell Death Differ. 11, 381–389. Paigen, B., Morrow, A., Holmes, P. A., Mitchell, D., and Williams, R. A. (1987). Quantitative assessment of atherosclerotic lesions in mice. Atherosclerosis 68, 231–240. Piedrahita, J. A., Zhang, S. H., Hagaman, J. R., Oliver, P. M., and Maeda, N. (1992). Generation of mice carrying a mutant apolipoprotein E gene inactivated by gene targeting in embryonic stem cells. Proc. Natl. Acad. Sci. USA 89, 4471–4475. Plump, A. S., Smith, J. D., Hayek, T., Aalto-Setala, K., Walsh, A., Verstuyft, J. G., Rubin, E. M., and Breslow, J. L. (1992). Severe hypercholesterolemia and atherosclerosis in apolipoprotein E-deficient mice created by homologous recombination in ES cells. Cell 71, 343–353. Purcell-Huynh, D. A., Farese, R. V., Jr., Johnson, D. F., Flynn, L. M., Pierotti, V., Newland, D. L., Linton, M. F., Sanan, D. A., and Young, S. G. (1995). Transgenic mice expressing high levels of human apolipoprotein B develop severe atherosclerotic lesions in response to a high-fat diet. J. Clin. Invest. 95, 2246–2257. Sanson, M., Auge, N., Vindis, C., Muller, C., Bando, Y., Thiers, J. C., Marachet, M. A., Zarkovic, K., Sawa, Y., Salvayre, R., and Negre-Salvayre, A. (2009). Oxidized lowdensity lipoproteins trigger endoplasmic reticulum stress in vascular cells: Prevention by oxygen-regulated protein 150 expression. Circ. Res. 104, 328–336. Tabas, I., Seimon, T., Timmins, J., Li, G., and Lim, W. (2009). Macrophage apoptosis in advanced atherosclerosis. Ann. NY Acad. Sci. 1173(Suppl. 1), E40–E45. Tabas, I., Tall, A., and Accili, D. (2010). The impact of macrophage insulin resistance on advanced atherosclerotic plaque progression. Circ. Res. 106, 58–67. Tangirala, R. K., Rubin, E. M., and Palinski, W. (1995). Quantitation of atherosclerosis in murine models: Correlation between lesions in the aortic origin and in the entire aorta, and differences in the extent of lesions between sexes in LDL receptor-deficient and apolipoprotein E-deficient mice. J. Lipid Res. 36, 2320–2328. Tsukano, H., Gotoh, T., Endo, M., Miyata, K., Tazume, H., Kadomatsu, T., Yano, M., Iwawaki, T., Kohno, K., Araki, K., Mizuta, H., and Oike, Y. (2010). The endoplasmic reticulum stress/EBP homologous protein pathway-mediated apoptosis in macrophages contributes to the instability of atherosclerotic plaques. Arterioscler. Thromb. Vasc. Biol. 30, 1925–1932.

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Whitman, S. C. (2004). A practical approach to using mice in atherosclerosis research. Clin. Biochem. Rev. 25, 81–93. Zhou, J., Moller, J., Danielsen, C. C., Bentzon, J., Ravn, H. B., Austin, R. C., and Falk, E. (2001). Dietary supplementation with methionine and homocysteine promotes early atherosclerosis but not plaque rupture in ApoE-deficient mice. Arterioscler. Thromb. Vasc. Biol. 21, 1470–1476. Zhou, J., Werstuck, G. H., Lhotak, S., de Koning, A. B., Sood, S. K., Hossain, G. S., Moller, J., Ritskes-Hoitinga, M., Falk, E., Dayal, S., Lentz, S. R., and Austin, R. C. (2004). Association of multiple cellular stress pathways with accelerated atherosclerosis in hyperhomocysteinemic apolipoprotein E-deficient mice. Circulation 110, 207–213. Zhou, J., Lhotak, S., Hilditch, B. A., and Austin, R. C. (2005). Activation of the unfolded protein response occurs at all stages of atherosclerotic lesion development in apolipoprotein E-deficient mice. Circulation 111, 1814–1821.

C H A P T E R

T H R E E

The Role of PDI as a Survival Factor in Cardiomyocyte Ischemia Stefano Toldo,* Anna Severino,† Antonio Abbate,* and Alfonso Baldi‡ Contents 1. Introduction 2. Identification of PDI as Cardio-Protective Factor 2.1. Apoptosis detection in autoptic human heart samples after AMI 2.2. PDI is upregulated in ischemic cardiomyocytes 2.3. UPR regulation in hypoxic cardiomyocytes 2.4. Superoxide dismutase-1 is a target of the anti-apoptotic action of PDI 2.5. Protective effects of PDI overexpression during acute MI in the mouse model 2.6. UPR activation and PDI-altered redox state in diabetes 2.7. Detection of PDI-altered redox state in the diabetic heart 3. Conclusions References

48 50 51 52 53 54 55 57 58 62 62

Abstract Acute myocardial infarction (AMI) leads to activation of unfolded protein response (UPR) following endoplasmic reticulum (ER) stress. Failing in the restoration of the proper folding activity in the ER can lead to apoptosis and cell death. While it can be easy to detect transcripts and proteins expression alterations during a pathological state, it can be difficult to address the importance of changes in protein expression in the physiopathological context. We found protein disulfide isomerase (PDI) increased expression in human autoptic heart samples correlating with cell survival following AMI. PDI enzymatic activity resulted to be important to achieve cardiomyocyte protection from hypoxic stress, dependent on its ability to relieve ER stress preventing accumulation of * Victoria Johnson Research Center and VCU Pauley Heart Center, Virginia Commonwealth University, Richmond, Virginia, USA { Institute of Cardiology, Catholic University of the Sacred Heart, Rome, Italy { Department of Biochemistry and Biophysics F. Cedrangolo, Second University of Naples, Italy Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00003-0

#


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48

Stefano Toldo et al.

nonfolded proteins in the ER, and to enhance superoxide dismutase 1 (SOD-1) activity. Furthermore, adenoviral-mediated PDI overexpression in an in vivo mouse model of AMI prevented adverse cardiac remodeling reducing cardiomyocyte apoptosis. Finally, we suggest a method to detect alterations in normal redox state in PDI (and eventually in the PDI family’s proteins) during pathologies in which ER stress is induced. Diabetes pathology correlates with increased risk of AMI and worse cardiac remodeling. We found an alteration in PDI redox state in the diabetic heart and suggest using this system for the detection of the redox state alteration to screen for therapies able to restore the proper redox state.

1. Introduction Ischemic cardiomyopathy (IC) refers to severe myocardial dysfunction resulting from occlusive coronary artery disease (CAD) (Burch and McDonald, 1971; Burch et al., 1970) and it represents the most frequent cause of cardiovascular mortality in Western countries. It is triggered by complete or partial occlusion of a major epicardial coronary artery, and it is associated with an imbalance between blood supply and demand, resulting in acute myocardial infarction (AMI). IC induces a significant impairment of left ventricular (LV) function. Despite treatment advances, complete reperfusion is not always achievable. This results in higher risk for pathologic cardiac enlargement, heart failure (HF), and death. Persistence of ischemia leads to irreversible cell death. Ischemic necrosis and myocardial apoptosis are both implicated in determining myocardiocyte loss in AMI (Cohn et al., 2000; Greenberg et al., 1984). Postinfarction LV remodeling consists of progressive chamber dilatation, wall thinning, and systolic/diastolic dysfunction (Fig. 3.1). This process involves cellular and molecular mechanisms, beginning days after AMI and persisting for weeks and months after the initial insult both at site of infarction (even after complete infarct healing) and in the surviving unaffected areas. Environmental changes and developmental processes can drastically change the workload of the endoplasmic reticulum (ER). To maintain ER homeostasis, a coordinated program known as the unfolded protein response (UPR) adjusts the folding capacity of the ER to meet demand and engages the destructive pathway if necessary (Hirsch et al., 2009). Hypoxic stress plays a pivotal role in the pathogenesis of heart disease, stroke, diabetes, and cancer (Okada et al., 2004; Thuerauf et al., 2006). Perturbations of ER homeostasis, such as prolonged hypoxic condition or metabolic stress, affect protein folding and cause ER stress (ERS). Diabetes is a group of heterogeneous disorders with the common elements of hyperglycemia and glucose intolerance, associated with more than

49

The Role of PDI as a Survival Factor in Cardiomyocyte Ischemia

Adverse cardiac remodeling following AMI Acute MI

Hours Heart failure

Years Months

Days Weeks

Compensatory hypertrophy

Progressive dilatation

Figure 3.1 AMI is the most common cause of acute and chronic HF. The initial damage to the myocardium during AMI activates reparative processes that are initially beneficial but produce pathologic changes over time. In the first hours and days, cardiac cell death can determine the extent of remodeling. This process is activated as a result of the initial loss of functional myocardium and subsequent secondary changes in the surviving portions of the heart and is inevitably associated with an unfavorable clinical course. The greater is the damage to heart, the greater is the remodeling process, and the worse is the expected outcome for the patient.

doubled risk of heart disease and stroke (Haffner et al., 1998; Hayat et al., 2004). Diabetes may be associated with LV dysfunction independent of CAD, hypertension, or any other cause of heart disease (Hayat et al., 2004). Diabetic patients have an increased likelihood of developing HF following myocardial infarction (MI), and once established, outcome is worse than in nondiabetics (Hayat et al., 2004). Recent studies describe activation of the UPR in both diabetes mellitus and IC. Diabetes affects the ability of the UPR to restore the physiologic state by altering the function of its key components (Cai and Kang, 2001; Xu et al., 2009). Protein disulfide isomerase (PDI) is a member of the UPR necessary for the appropriate protein folding and prevention of misfolding during stress, as seen in myocardial ischemic injury (Schroder and Kaufman, 2005; Severino et al., 2007). In the liver of diabetic rodents, PDI has been reported to be in an altered redox state affecting PDI’s ability to function, leading to the accumulation of misfolded protein in the ER lumen (Nardai et al., 2005). PDI catalyzes the formation, reduction, and isomerization of disulfide bonds in the ER lumen, and as a polypeptide-binding protein, it can function as a molecular chaperone which assists the folding of polypeptides. Recently,

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PDI has been described as a hypoxia-induced protein in some in vitro and in vivo models being enhanced at the mRNA level and protein level in human aortic and pulmonary artery endothelial cells (Graven et al., 2002; Myoishi et al., 2007; Tian et al., 2009). Upregulation of PDI specifically in endothelial cells contributes to their ability to tolerate hypoxia (Graven et al., 2002). Upregulation of PDI has been demonstrated also in neuroblastoma cells in response to hypoxia, where it is hypothesized to serve as protective role. Overexpression of PDI in these cells resulted in an increase of cell viability in response to hypoxia and protected hippocampal cells from apoptotic cell death in response to brain ischemia (Tanaka et al., 2000). All these results evidence a PDI protective role in degenerative diseases, brain ischemia, and in general in hypoxic systems, characterized by accumulation of unfolded or misfolded proteins and ER stress. In this chapter, we review (1) how apoptosis inversely correlates with PDI expression following ischemia in the heart of human patients and mouse heart; (2) how PDI expression modulates the extent of tissue injury; and (3) how changes in PDI redox state in vivo in the heart may affect its function in diabetes. Despite an increasing number of ways to detect markers of UPR activation and ER stress, the identification of key factors playing a role in human diseases remains difficult to be assessed. In this chapter, we describe our approach to determine protein expression, its redox state, and its functional consequences in a model of ischemic heart disease.

2. Identification of PDI as Cardio-Protective Factor In this section, we describe a direct approach to evaluate changes in gene expression in the ischemic heart and how to move on to identify and characterize the role and the importance of a specific protein in cell protection following AMI. Autoptic myocardial samples were collected from the proximal ischemic area and the remote zone to perform apoptotic cell death analysis and protein expression analysis. Previously, a gene expression analysis approach of the UPR proteins showed that in the ischemic heart, a cluster of ER stress-related genes was transcriptionally activated during the development of the disease (Azfer et al., 2006). This screening demonstrated elevated transcript levels of genes involved in UPR such as ER and cytoplasmic chaperones, oxidoreductases, PDI family, and ER-associated degradation system such as ubiquitin. PDI gene, as a member of the UPR regulon, was found to be upregulated in the viable myocardium. Using immunohistochemistry and the array technology we identified a correlation between PDI expression and apoptotic cell death following AMI.


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2.1. Apoptosis detection in autoptic human heart samples after AMI Autopsy, in the examined subjects, was performed within 24 h after death (median 16 h). Gross examination of the hearts was performed to measure LV parameters and to define the infarct-related artery and the infarcted area. LV longitudinal and transverse diameters were measured at the atrioventricular section. The LV free wall thickness was measured at the median third of the posterior wall. Specimens were obtained at sites of MI and in regions of the left ventricle remote from the infarcted area by a patent coronary artery and were either snap frozen for RNA extraction or fixed in 10% paraformaldehyde for immunohistochemical analysis. The morphologic analysis of tissue structure, cellular and nuclear integrity, and inflammatory infiltrates was performed by light microscopy. In situ end-labeling of DNA fragmentation (TUNEL) have enabled an easy qualitative and quantitative evaluation of cell death in tissues (Haunstetter and Izumo, 1998; Narula et al., 1996; Olivetti et al., 1996, 1997). Several series of TUNEL-stained sections were subsequently used for the staining for caspase-3 or muscle-actin to better characterize which of the TUNEL positive cells was an apoptotic cardiomyocyte. Cardiomyocytes were defined as apoptotic if colocalization of TUNEL caspase-3 staining was evident. In the infarcted hearts, detection of apoptotic cardiomyocytes showed a significantly higher apoptotic rate in the peri-infarct versus remote areas. Interestingly, the apoptotic rate was higher both in the peri-infarct and remote areas and it significantly correlated with increasing cardiac weight and cardiac transverse diameter (Severino et al., 2007). Myocardial apoptosis peaks at 4–12 h in AMI and is persistently demonstrable up to 10 days (Abbate et al., 2002). In the context of MI, apoptosis represents the major form of death, being several times more common than necrosis (peak value of 43% of apoptotic cardiomyocytes at 4.5 h vs. peak value of 8% of necrotic cells at 24 h) (Kajstura et al., 1996). Several other studies, both experimental and observational, have confirmed such results (Abbate et al., 2002). Apoptosis is a fundamental physiologic and pathologic mechanism that allows elimination of normal but no-longer-useful cells during embryogenesis or of aged or damaged cells during life. A delicate balance between survival and death exists in cells undergoing physiologic and pathologic stress (such as hypoxia) and initiation of apoptosis during initial phases may not always be followed by its completion (Buja and Entman, 1998; Majno and Joris, 1995). Furthermore, balance between apoptosis and primary necrosis depends on available energetic levels, as completion of apoptosis needs adequate ATP cellular concentrations. Necrosis of cardiomyocytes is usually completed within 24–48 h following abrupt coronary occlusion and phagocytosis and removal of cellular debris by inflammatory cells starts at 24 h. Cardiomyocytes surviving through the acute phases undergo major

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metabolic rearrangements in order to favor survival in the delicate balance between hibernation and apoptosis (Abbate et al., 2002). The term “hibernation” refers to a chronic condition of severe energy deprivation of the myocardium due to chronic or repetitive hypoperfusion associated to reversible contractile dysfunction (Braunwald and Kloner, 1982). Cardiomyocytes in the border zone have been described to have multiple overlapping phenotypes of oncosis, apoptosis, and autophagy (Arbustini et al., 2008; Dorn and Diwan, 2008, Kostin et al., 2003). Independent of the markers used to detect cardiomyocyte degeneration (TUNEL, annexin V, caspase-3 activation for apoptosis, or ubiquitin and LC3 staining for autophagy), the degree of this degeneration correlates with clinical signs of adverse cardiac remodeling and predicts outcome. Experimental induction of apoptosis is sufficient to cause dilated cardiomyopathy, HF, and death in the mouse (Wencker et al., 2003). However, therapies able to reduce cardiomyocyte apoptosis ameliorate postinfarction myocardial remodeling and cardiac function (Abbate et al., 2002, 2006; Dorn and Diwan, 2008; Nishida et al., 2008). The data collected from the autoptic heart samples confirmed that the apoptotic rate both in the peri-infarct and remote areas significantly correlated with increasing cardiac weight and cardiac transverse diameter (Severino et al., 2007).

2.2. PDI is upregulated in ischemic cardiomyocytes Impaired oxygen supplementation, inflammation, necrotic cell death, neurohormonal signaling, etc., contribute to altered gene and protein expression in the peri-infarct area (Abbate et al., 2009; Searles, 2002; Temsah et al., 1999, 2000). To evaluate the change in the transcriptome in the bordering zone, we performed a cardiovascular microarray on RNA extracted from the autoptic samples. The array for human cardiovascular specific genes was used to compare the RNA transcripts from two different pools: one from samples obtained in the peri-infarct region and one from the remote area. Five micrograms of pooled RNAs (from border and remote zones) were retrotranscribed using a-P32 dATP. The choice of pooling remote myocardium as comparison for the bordering zone was considered essential to have an internal (rather than external) control and to minimize the possibility of postmortem autolysis-related changes. Two independent RNA preparations were used to perform the experiment in duplicate. The PDI gene was upregulated threefold in the RNA extracted from the peri-infarct areas. Gene upregulation in the border zone was confirmed by semiquantitative reverse transcriptase polymerase chain reaction (qPCR) assay on the same pooled RNAs. Immunohistochemical analysis of PDI expression and the costaining for TUNEL was performed to confirm the upregulation of PDI at the protein level in the peri-infarct areas and to show a correlation between PDI expression and cell death. This analysis showed that most of the


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cardiomyocytes overexpressing PDI were TUNEL negative. In the cases with intense PDI expression, the apoptotic rate was significantly lower compared with the samples having mild to moderate expression. Notably, subjects with intense PDI expression were less likely to be suffering from stage-C New York Heart Association functional class IV congestive HF and to have biventricular enlargement at postmortem examination.

2.3. UPR regulation in hypoxic cardiomyocytes BiP/GRP78, the major molecular chaperone of the ER, works as a sensor of unfolded proteins in the ER and regulates the activation of ER stress transducers. ER stress response leads to a transcriptional upregulation program, which is conducted by the three transducers PERK, ATF6, and IRE1. To test the effects of hypoxia on cardiomyocytes, we used the mouse HL-1 cell line. HL-1 cells are differentiated and proliferating atrial cardiomyocytes derived from mouse AT-1 cells (Yusuf et al., 2000). Hypoxia was induced with a gas mixture of 5% carbon dioxide and 95% gaseous nitrogen for 24 and 48 h (hypoxic groups). We used total RNA extracted from hypoxia-treated HL1 cells to analyze the expression levels of PDI as well as of PERK, ATF6, Bip, and IRE1 and the chaperone GRP94 by qRT-PCR. b-Actin, used as housekeeping gene to normalize the results, is not affected by hypoxic condition, as tested in many experiments conducted in our lab comparing the results with other housekeeping genes (data not shown). PDI, GRP94, and ATF6-a were significantly upregulated (p < 0.05) after exposure to hypoxia. ATF6 expression increased with duration of hypoxia while the expression of IRE1-a increased significantly (p < 0.05) only after 48-h exposure to hypoxia. The mRNA level of PERK did not significantly change, whereas the level of Bip-mRNA decreased under influence of hypoxia. PERK, a type-I ER transmembrane protein with a luminal domain, in normal conditions is bound by Bip/GRP78. Upon ER stress activation, Bip releases the luminal domain of PERK, allowing PERK to dimerize and become active, leading to general shut-off of the translation and upregulation of the genes involved in the recovery from ER stress (Raven and Koromilas, 2008). Our findings are consistent with the activity of these two proteins. The expression analysis of the PDI adjuvant enzyme Ero1 on the same mRNA samples showed that this protein is enhanced with the duration of hypoxia. These results provided independent confirmation for the observed upregulation of PDI during hypoxic condition. We have also repeated the qPCR experiments with cells transfected 24 h prior to hypoxia and overexpressing PDI. These data indicated that overexpression of PDI did not significantly change the expression of the tested genes, suggesting that the detected effects of PDI overexpression are direct. The co-upregulation of Ero1 supported the notion that the enzymatic activity of PDI protects cardiomyocytes after hypoxic stress. To test this hypothesis, we described as HL-1 cells

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overexpressing PDI were protected by hypoxic stress. The forced expression of PDI carrying point mutations in one of the PDI’s active sites (PDIa C40S and PDIa’ C387S) abolished the protective role seen with the wild-type enzyme and actually increased the cell death compared to the controls. A similar set of experiments was performed infecting the cells with an adenoviral vector expressing the GFP and coding either for PDI–GFP/PDI– or for the PDI’s mutants (PDIa C40S–GFP/PDIa–, PDIa0 C387S–GFP/PDIa0 –, and the double mutant PDIaa0 C40S/C387S–GFP/PDIaa0 –) using the GFP vector (GFP/null) as control. The cells were infected with the adenoviruses 24 h before the hypoxic treatment (24 and 48 h). Cells were marked with propidium iodine and were analyzed by flow cytofluorimetry. Cytofluorimetric analysis showed a marked reduction of propidium iodine as the PDI– GFP positive cells compared to the GFP alone showed that the actually the population of cells overexpressing PDI had a less apoptotic rate than the others. Furthermore, all the PDI mutants increased the effects of hypoxia on cell survival (Fig. 3.2). These data indicates that PDI enzymatic activity is necessary to put into effect its anti-apoptotic action.

2.4. Superoxide dismutase-1 is a target of the anti-apoptotic action of PDI Impaired protein folding in the ER can be produced either by the reduction of the oxygen supply associated with the coronary occlusion or with the following oxidative stress associated with reperfusion. In fact, disulfide bond

Cell death %

80 70 60

* **

50

* **

40 30

* **

**

20 10

48 h

24 h

GFP/null GFP/aa⬘

48 h

24 h

Normoxia

Hypoxia GFP/PDI

GFP/PDIa

GFP/PDIa⬘

* p < 0.05 vs. GFP/null ** p < 0.05 vs. GFP/PDIa, – a⬘, – aa⬘

Figure 3.2 PDI overexpression limits cell death induced by hypoxia. Adenoviralmediated GFP-associated overexpression of PDI coupled with cytofluorimetry were used to assess the ratio of green fluorescent protein expressing cells with DNA damage. The infection with the wild-type gene reduces DNA damage, detected with propidium iodide, and cell death after 24 and 48 h of hypoxia. These results show that while PDI reduces propidium staining by 45% at 48 h and 60% at 24 h, the PDI mutants, a, a0 , and the double mutant, aa0 , increase the cell death also in normoxic condition.


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formation requires oxygen and is compromised by the production of reactive oxygen species. Indeed, significant superoxide generation occurs in cardiomyocytes during both ischemia and reperfusion (King et al., 1998). Copper– zinc superoxide dismutase, also known as superoxide dismutase 1 or SOD-1 is the principal superoxide scavenger system in the cytoplasm (Muller et al., 2007). Overexpression of SOD-1, which efficiently dismutases superoxide and prevents postischemic injury in the heart, during ischemia attenuates apoptosis and the inflammatory response (Wang et al., 1998). Recently, in amyotrophic lateral sclerosis (ALS), a neurodegenerative disease in which mutations of SOD-1 (mSOD-1) are associated with familial type of the disease, it has been found an association between SOD-1 with PDI (Atkin et al., 2006). Although SOD-1 is very thermally stable, mSOD-1 aggregates are present in spinal cords of ALS patients and in transgenic mice overexpressing this mutant protein. The mechanism of mSOD-1-mediated toxicity is unclear, but it involves apoptotic signaling (Atkin et al., 2006; Pasinelli et al., 2004). Furthermore, neuronal cells transfected with wtSOD1 or mSOD1 and treated with the PDI inhibitor bacitracin showed an increased formation of aggregates suggesting that PDI prevents the formation of SOD1 inclusions in motor neurons (Atkin et al., 2006). Interestingly, it has been demonstrated that PDI is overexpressed in familial ALS, acting as a survival factor through physical interaction with SOD-1 (Buja and Entman, 1998). This interaction prevents SOD-1 aggregation and consequently neuronal degeneration. To obtain initial insights into protective mechanisms supported by PDI in our model, we investigated whether its anti-apoptotic effects were mediated by a direct action on the activity or the expression of SOD-1, determined by measuring the release of hydrogen peroxide (H2O2) and by Western blot analysis. The HL1 cells overexpressing PDI showed a significantly higher H2O2 release than HL1 cells transfected with the vector alone (Fig. 3.3A). However, this overexpression did not affect SOD-1 expression (Fig. 3.3B). Furthermore, hypoxia had no direct effects SOD-1 expression (Fig. 3.3C). Therefore, PDI in HL-1 cells was able to increase SOD-1 activity. Our finding of increased SOD activity is in line with the previous reports and suggests that increased expression of PDI might protect myocardial tissue from apoptosis mediated by superoxide, through increased SOD activity.

2.5. Protective effects of PDI overexpression during acute MI in the mouse model We tested the effects of PDI gene delivery in an in vivo mouse model of MI, by injecting the adenoviral vector expressing GFP/PDI into the hearts of adult male CD-1 mice before the infarction. The protocol for in vivo adenoviral-mediated gene transfer was set as follows: 48 h before the induction of the ischemic injury, four injections of adenoviral vectors

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A

B

*

1400 H2O2 pM/mg of proteins

GFP null

GFP PDI

PDI b -Actin

1200

SOD-1

1000

C Hypoxia

800 600

–

+ 24 h

+ 48 h

PDI 400 b -Actin 200 SOD-1 Controls

PDI

*p < 0.05 vs. control Figure 3.3 PDI does not regulate SOD-1 expression but increases its activity. In HL-1 cell extracts driven by PDI overexpression, SOD-1 activity increases by 80% (A). This enhancement is not detected with a concomitant increase of SOD-1 expression but only with a boost in the dismutase activity (B). Hypoxic stress, while able to positively regulate PDI expression, does not affect SOD-1 expression (C).

were performed through a 32-gauge needle into the ventricular wall of adult CD-1 mice, two for each side flanking the left coronary artery. The volume of each injection was 2.5 ml and the approximate titer was 1010 PFU/ml. Hearts injected with transgene-null adenoviral vectors or hearts from sham-operated animal were used as a control. To induce MI, the left main coronary artery was ligated. Echocardiography was performed to assess cardiac function 7 days after MI, prior to sacrifice. Efficiency of the viral infection was assessed by Western blots and immunohistochemistry, by increased myocardial PDI expression and GFP detection in PDI/ GFP-injected mice. In vivo assessment of the remodeling pattern with transthoracic echocardiography confirmed a more favorable pattern in GFP/PDI-treated mice. Compared with sham-operated mice, GFP/nulltreated infarcted mice had significantly greater LV end diastolic and systolic diameter (LVEDD and LVESD). Compared with GFP/null animals, GFP/ PDI mice had significantly smaller LVEDD similar to that in sham-operated animals and a trend toward lower LVESD values. Infarct size assessment in the mid-ventricular section with Masson’s trichrome staining showed a significantly smaller infarct size in GFP/PDI-treated mice, determined as reduced circumferential extension and as transmural extension. The apoptotic rate in the peri-infarct region was also significantly reduced. PDI gene

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AMI Myocardial damage

Oxidative stress

↑ SOD-1 activity

↑ Protein-folding activity

UPR activation

ER stress

UPR activation PDI upregulation

Figure 3.4 Role of PDI upregulation following acute myocardial infarction. Following AMI, cardiomyocyte’s fate in the border zone depends on their ability to counteract oxidative stress and endoplasmic reticulum overload-induced stress. Both this pathologic conditions induce activation of the UPR to relieve unfolded and misfolded protein accumulation in the ER lumen. PDI family members, including PDI, are upregulated to shrink the ER stress in response to UPR, acting directly as molecular chaperones to stabilize the proteins’ structure. PDI can also act on SOD-1 activity. This step is dependent on PDI catalytic activity and its enhanced expression. Both these actions on protein folding and SOD-1 activity can relieve ER stress and oxidative damage, reducing apoptosis and minimizing the effects of tissue injury on myocardial function.

transfer therapy was able to prevent both adverse cardiac remodeling and cardiomyocyte apoptosis. We hypothesize that this protective action is realized directly by PDI’s ability to relieve unfolded protein accumulation and indirectly by activation of SOD activity (Fig. 3.4).

2.6. UPR activation and PDI-altered redox state in diabetes We have, so far, described how PDI was identified as survival factor in ischemic cardiomyocytes. We now explore the concept that PDI-altered redox state may affect its function. A recent study in a mouse model of diabetes showed pathological and functional changes in the cardiomyocyte structure as well as increase cardiomyocytes apoptosis accompanied by ER stress activation (Cai and Kang, 2001). In these mice, the analysis of ER chaperones showed an increased expression of eIF2, GRP78, and GRP94 proteins and the activation of ATF6. Diabetes is a disease in which our knowledge of ER stress and UPR derives mainly from studies performed on pancreas, liver, vasculature, and specialized epithelial cells (i.e., ocular

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epithelial cells). Nardai et al. (2005) showed that PDI redox state was altered during diabetes. We believe that understanding of PDI in AMI and diabetes is relevant to understand ER stress, UPR, and cell death responses. It is possible that PDI function is enhanced in diabetes considering the central role of the ER stress. However, it is possible that diabetes impairs PDI expression or function, in which case PDI dysfunction would represent an additional mechanism by which diabetes affects cardiovascular pathophysiology. As diabetes has been described to induce a pathologic state called diabetic cardiomyopathy (Hayat et al., 2004; Kannel et al., 1974; Rubler et al., 1972) in which ER stress plays a pivotal role, in this paragraph, we focus on the evaluation of PDI expression and redox state in the diabetic heart. Several pieces of evidence showed that the intracellular level of small molecules as FAD is lower in both diabetic animal models and diabetic patients (Banhegyi et al., 1998, Reddi, 1986) and that the activity of some FAD-containing enzymes, as Ero1, is significantly lower in experimental diabetes (Nardai et al., 2003). In streptozotocin-induced diabetic rats, despite of the oxidative changes of the extracellular space, the redox environment of liver microsomal vesicles was shifted to a more reduced state with an increase of total thiol content and an increase of protein-thiol/ disulfide ratio (Nardai et al., 2005). Interestingly, during diabetes, PDI has an almost complete shift to its most reduced form. Nardai et al. showed that in the liver of diabetic mice, a large fraction of PDI (and Erp57) is in a reduced form compared to healthy animals. The effect was selective on PDI because the redox state of other proteins, such as calnexin, Grp78, and Grp94 as well as the inactive PDI-homolog ERp28, was unchanged (Nardai et al., 2005). In an opposite way, Ero1 showed a more oxidized state. Further analysis assessed a significant decrease of the PDI enzymatic activity in diabetic samples compared to controls not accompanied by a decrease of the protein level.

2.7. Detection of PDI-altered redox state in the diabetic heart To define the redox state of PDI in the heart of diabetic mice, we used a biochemical assay to determine the differences in the redox status of PDI in heart samples of healthy and diabetic mice, to compare the quote of oxidized protein able to catalyze the disulfide bond formation. This assay can be used to study the effects of drug therapy on the restoration of the oxidized state of PDI. The assay takes advantage of the ability of methoxy-polyethylene glycol-maleimide (mPEG-mal), a polymer with a molecular mass of 5 kDa, to bind to the free thiol groups of a protein to induce a negative electrophoretic shift that is responsible for an apparent increase of molecular weight of almost 25 kDa for each mPEG-mal molecule linked. The pegylation reaction is performed using the method published by Appenzeller-Herzog


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and Ellgaard (2008) with some modifications to be adapted for tissue protein analysis using an immunological-based analysis. Hearts of diabetic mice were harvested after 4 weeks from the injection of streptozotocin (130 mg/kg). In this model, glucose levels were permanently elevated from day 3 to the end of the chosen time point. The snap-frozen hearts were powdered using a mortar and a pestle. The samples were prepared as follow for biochemical modifications to discriminate between different redox states of the sulfhydryl groups of PDI. A control tissue sample was prepared to disclose any nonspecific signal in the Western blot analysis. Ten milligrams of powdered tissue was homogenized in 200 ml of extraction buffer (80 mM Tris–HCl, pH 7.3, 200 mM PMSF) containing DTT (10 mM) for 5 min at 37 C. This sample was prepared to assess the maximum reduction state of the proteins and was used to avoid false-positive signals. The DTT-treated control tissue sample did not undergo the pegylation reaction to maintain the highest electrophoretic mobility. However, all the other samples were pegylated to produce samples with lower electrophoretic mobility. Ten milligrams of the test samples from diabetic and control mice underwent chemical alkylation with N-ethylmaleimide (NEM) to block the sulfhydryl groups of the free-reduced cysteines. NEM is a cell membrane permeable molecule. Briefly, the tissue powdered in liquid nitrogen was incubated in 100 ml of ice-cold extraction buffer supplemented with 20 mM NEM and 30% DMSO (v/v). Samples were vortexed for 5 s and then centrifuged 1 min at 20,000g at 4 C followed by resuspension in the same buffer and 10 min incubation on ice. These steps and the use of high concentration of DMSO help the rapid diffusion of NEM in the powdered tissue. After 1 min centrifugation at 20,000g at 4 C, the tissue was then placed in 200 ml of fresh extraction buffer and homogenized passing the sample 10 times through a 25-gauge needle syringe. After addition of SDS (final concentration 0.1%), the samples were denatured for 10 min at 97 C, centrifuged 1 min at 25 C, and the supernatant was collected for the following biochemical treatments. One hundred microliters of NEM-alkylated samples were reduced for 15 min by addition of Tris(2-carboxyethyl) phosphine hydrochloride (TCEP, final concentration 10 mM) to reduce the oxidize cysteines and were incubated for 1 h with mPEG-mal (15 mM). This sequence of reactions allowed the alkylation of the sulfhydryl groups gained in covalent bounds, producing a molecule with higher molecular weight compared to the native protein. Excess mPEG-mal was removed by using methanol/chloroform protein precipitation. One sample volume of methanol was added to the lysate, followed by vortexing. After the addition of 0.25 sample volumes of chloroform and vortexing the mixture was centrifuged for 5 min at 20,000g at room temperature for phase separation. The lower phase was discarded, and the proteins in the interphase were resuspended adding 0.75

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sample volumes of methanol and vortexing followed by centrifugation for 2 min at 20,000g (all sample volumes compared with the original sample). The supernatant was carefully aspirated, and the pellet dried for 15 min at room temperature. The samples were resuspended in nonreducing Laemmli buffer 1 for Western blot analysis. Samples underwent SDS-PAGE analysis in 8% acrylamide gels to allow better separation of the bands. We recommend analyzing 1/10 of the sample resuspended in Laemmli buffer. The proteins were transferred onto nitrocellulose membranes, incubated with a rabbit anti-mouse polyclonal antibody raised against PDI (DL-11, Sigma-Aldrich) and a mouse anti-mouse b-actin (clone C-2, Sigma-Aldrich), following enhanced-chemiluminescence (ECL) analysis and autoradiography (Fig. 3.5). For the detection of the most oxidized form (with the lower electrophoretic mobility), we recommend to use a more sensitive ECL, because of the low ratio of PDI

Pegylation of cysteines H

H

S

S

S

PDI

PDI Alkilation of reduced + groups/reduction

R

R

S

S

Alkilation of reduced + groups/reduction

PDI

H

H

S

S

PDI + mPEG-mal

mPEG-mal +

R

R

mPEG-mal

S

S

S

PDI

S

WB

WB

mal-PEGm S

PDI

Figure 3.5 Relationship between the sequence of reactions in the cysteines pegylation protocol and Western blot hybridization based on the thiols redox state. In the figure are reported the two sets of reactions developed to induce pegylation of the cysteines gained in disulphide bonds in the two active sites of PDI and the expected result after Western blot analysis. On the left and the right sides are represented the alkylation reactions in the active sites in the case of a reduced (left) or oxidized (right) catalytic site. In reduced catalytic site (left side), the first alkylation produces sulfur-bound N-ethylmaleimide, while the following reactions are ineffective. The product is a PDI bound to a low molecular weight molecule, without any effect on its electrophoretic properties. When the active sites are oxidized (right side), the cysteines are protected by the N-ethylmaleimide alkylation. The reduction with TCEP and the following reaction with mPEG-mal produce a high molecular weight PDI, with decreased electrophoretic mobility.

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completely pegylated. The protein bands were compared by densitometric analysis (Scion Image, Scion Image Corporation) and the results were adjusted to the b-actin quantity in the samples. In Fig. 3.6 are reported the resulting bands after the Western blot analysis. In the diabetic hearts, after 4 weeks from the streptozotocin treatment, PDI expression is unaffected compared to the healthy hearts, but the quantity of oxidized PDI is lower. Compared to the previous published techniques (AppenzellerHerzog and Ellgaard, 2008, Nardai et al., 2005), this method offers both vantages and limitations. We are able to detect easily the quote of the most reduced and most oxidized PDI in the tissue with a good resolution between the bands. However, it is hard to analyze the semioxidized variants of this protein due to the presence of many bands close to each other. This is

A

Control

STZ

Oxidized PDI

Total PDI

b-Actin

Total PDI

C Change compared to the control (%)

Change compared to the control (%)

B 400 350 300 250 200 150 100 50

Controls

STZ

Oxidized PDI p < 0.05 160 140 120 100 80 60 40 20 Controls

STZ

Figure 3.6 Diabetes induces reduction of PDI active sites. PDI expression analysis was performed on total protein extracted from the heart tissue of healthy and diabetic mice to detect differences in the amount of oxidized form of the protein. The detection was performed with Western blot analysis (A). The densitometric analysis revealed that while total PDI expression is unaffected during diabetes (B), the portion of the oxidized protein is markedly reduced by 60% (C).

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an approach to detect how in vivo a pathological condition affecting the ER is able to modify the redox state of PDI’s catalytic sites as well as of other PDI family members. Furthermore, it is a powerful tool to analyze the ability of therapeutic drugs to restore the catalytic activity of PDI in vivo.

3. Conclusions PDI upregulation following hypoxia is part of the survival response due to the oxygen deprivation to maintain the correct protein folding in the cells. Following AMI, overexpression of PDI correlates with less cell death and its forced overexpression induces a better cardiac remodeling due to reduced cardiomyocytes apoptosis. The mechanism of myocardial protection can be determined in part by the PDI ability to reduce the protein overload and the ER stress induced by hypoxia, and in part to the ability of PDI to induce SOD-1 activity in cardiomyocytes. Both these actions are able to reduce the apoptotic cell death preventing ER-dependent apoptosis and the oxidative damage enhanced by reactive oxygen species. The worse outcome seen in both diabetic patients with AMI and animal model of AMI can be due to a preexisting condition disabling the protective pathway activated following ischemic damage. PDI ability to create new disulphide bonds is dependent on the oxidation of its active sites. To oxidize two reduced cysteines, PDI has to be oxidized. The finding that during diabetes, PDI is found mostly reduced in different organs (Nardai et al., 2005) and in the heart can partially explain the induction of UPR and ER stress correlated with reduced protein secretion, protein accumulation in the ER, and increased ER-dependent apoptosis. This shift to a more reduced form of PDI could be responsible for reduced protection of cardiomyocyte following AMI. The possibility to detect PDI-altered redox state can help to characterize pathological condition associated with many diseases, including diabetes, diabetic cardiomyopathy, and ischemic heart disease, and can help to find therapeutic approach able to restore the correct redox state.

REFERENCES Abbate, A., Biondi-Zoccai, G. G., and Baldi, A. (2002). Pathophysiologic role of myocardial apoptosis in post-infarction left ventricular remodeling. J. Cell. Physiol. 193, 145–153. Abbate, A., Bussani, R., Amin, M. S., Vetrovec, G. W., and Baldi, A. (2006). Acute myocardial infarction and heart failure: Role of apoptosis. Int. J. Biochem. Cell Biol. 38, 1834–1840.


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Searles, C. D. (2002). The nitric oxide pathway and oxidative stress in heart failure. Congest. Heart Fail. 8(142–147), 155. Severino, A., Campioni, M., Straino, S., Salloum, F. N., Schmidt, N., Herbrand, U., Frede, S., Toietta, G., Di Rocco, G., Bussani, R., Silvestri, F., Piro, M., et al. (2007). Identification of protein disulfide isomerase as a cardiomyocyte survival factor in ischemic cardiomyopathy. J. Am. Coll. Cardiol. 50, 1029–1037. Tanaka, S., Uehara, T., and Nomura, Y. (2000). Up-regulation of protein disulfide isomerase in response to hypoxia/brain ischemia and its protective effect against apoptotic cell death. J. Biol. Chem. 275, 10388–10393. Temsah, R. M., Netticadan, T., Chapman, D., Takeda, S., Mochizuki, S., and Dhalla, N. S. (1999). Alterations in sarcoplasmic reticulum function and gene expression in ischemic– reperfused rat heart. Am. J. Physiol. 277, H584–H594. Temsah, R. M., Dyck, C., Netticadan, T., Chapman, D., Elimban, V., and Dhalla, N. S. (2000). Effect of beta-adrenoceptor blockers on sarcoplasmic reticular function and gene expression in the ischemic–reperfused heart. J. Pharmacol. Exp. Ther. 293, 15–23. Thuerauf, D. J., Marcinko, M., Gude, N., Rubio, M., Sussman, M. A., and Glembotski, C. C. (2006). Activation of the unfolded protein response in infarcted mouse heart and hypoxic cultured cardiac myocytes. Circ. Res. 99, 275–282. Tian, F., Zhou, X., Wikstro¨m, J., Karlsson, H., Sjo¨land, H., Gan, L. M., Boreń, J., and Akyu¨rek, L. M. (2009). Protein disulfide isomerase increases in myocardial endothelial cells in mice exposed to chronic hypoxia: A stimulatory role in angiogenesis. Am. J. Physiol. Heart Circ. Physiol. 297, 1078–1086. Wang, P., Chen, H., Qin, H., Sankarapandi, S., Becher, M. W., Wong, P. C., and Zweier, J. L. (1998). Overexpression of human copper, zinc-superoxide dismutase (SOD1) prevents postischemic injury. Proc. Natl. Acad. Sci. USA 95, 4556–4560. Wencker, D., Chandra, M., Nguyen, K., Miao, W., Garantziotis, S., Factor, S. M., Shirani, J., Armstrong, R. C., and Kitsis, R. N. (2003). A mechanistic role for cardiac myocyte apoptosis in heart failure. J. Clin. Invest. 111, 1497–1504. Xu, J., Wang, G., Wang, Y., Liu, Q., Xu, W., Tan, Y., and Cai, L. (2009). Diabetes- and angiotensin II-induced cardiac endoplasmic reticulum stress and cell death: Metallothionein protection. J. Cell. Mol. Med. 13, 1499–1512. Yusuf, S., Sleight, P., Pogue, J., Bosch, J., Davies, R., and Dagenais, G. (2000). Effects of an angiotensin-converting-enzyme inhibitor, ramipril, on cardiovascular events in high-risk patients. The Heart Outcomes Prevention Evaluation Study Investigators. N. Engl. J. Med. 20(342), 145–153.


C H A P T E R

F O U R

Measurement of the Increase in Endoplasmic Reticulum Stress-Related Proteins and Genes in Adipose Tissue of Obese, Insulin-Resistant Individuals Guenther Boden* and Salim Merali† Contents 1. Introduction 2. Study Subjects and Fat Biopsies 2.1. Subjects 2.2. Fat biopsies 3. Proteomic Analysis 3.1. Required materials 3.2. Protein extraction protocol for two-dimensional electrophoresis 3.3. First dimension of 2DE 3.4. Second dimension of 2DE 3.5. Fluorescent staining with SYPROÒ Ruby 3.6. 2DE image analysis 3.7. In-gel digestion 3.8. Extraction of peptides 3.9. Preparation of MALDI matrix 3.10. Sample plate preparation and MALDI-TOF/TOF analysis 3.11. Results of proteomic analysis 4. Western Blot Analysis 4.1. Required materials 4.2. Expression analysis 4.3. Results of Western blot analysis

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* Department of Medicine and the Clinical Research Center, Temple University School of Medicine, Philadelphia, PA, USA { Department of Biochemistry, Temple University School of Medicine, Philadelphia, PA, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00004-2

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Guenther Boden and Salim Merali

5. RT-PCR Analysis 5.1. Results of RT-PCR analysis Acknowledgments References

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Abstract Here, we provide a detailed description of proteomic, Western blot and RT-PCR analyses performed to examine fat biopsy samples from lean insulin-sensitive and obese insulin-resistant nondiabetic individuals for evidence of endoplasmic reticulum (ER) stress. Subcutaneous fat biopsies were obtained from the upper thighs of six lean and six obese nondiabetic subjects. Fat homogenates were used for proteomic (two-dimensional gel (2DE) and MALDI-TOF/TOF), Western blot, and RT-PCR analysis. Proteomic analysis revealed 19 differentially upregulated proteins in fat of obese subjects. Three of these proteins were the ER stress-related unfolded protein response (UPR) proteins calreticulin, protein disulfide-isomerase A3, and glutathione-S-transferase P; Western blotting revealed upregulation of several other UPR stress-related proteins, including calnexin, a membranebound chaperone, and phospho c-jun NH2-terminal kinase ( JNK)-1, a downstream effector protein of ER stress; RT-PCR analysis revealed upregulation of the spliced form of X-box-binding protein-1s, a potent transcription factor and part of the proximal ER stress sensor inositol-requiring enzyme-1 pathway. These findings demonstrate of UPR activation in subcutaneous adipose tissue of obese human subjects. As JNK can inhibit insulin action and activate proinflammatory pathways, ER stress activation of JNK may be a link between obesity, insulin resistance, and inflammation.

1. Introduction Obesity is closely associated with insulin resistance and with a state of low-grade inflammation characterized by elevation of inflammatory cytokines in blood and tissues (Bray, 2004). Insulin resistance and inflammation contribute to the development of several disorders including type 2 diabetes, hypertension, atherogenic dyslipidemia, and abnormalities in blood coagulation and fibrinolysis, all of which are independent risk factors for atherosclerotic vascular disease such as heart attacks, strokes, and peripheral arterial disease (Bray, 2004). Therefore, it has become important to understand why and how obesity is so tightly linked with insulin resistance and inflammation. On one hand, free fatty acids (FFA) have been established as important links between obesity, insulin resistance, and inflammation. Most obese people have elevated plasma FFA levels (Reaven et al., 1988) and elevated plasma FFA levels have been shown to cause insulin resistance and

ER Stress Proteins and Genes in Human Fat

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inflammation (Boden et al., 1994, 2001, 2005; Itani et al., 2002; Santomauro et al., 1999; Yu et al., 2002). On the other hand, not all of these insulin resistant subjects have elevated plasma FFA levels which means that there must be other causes for obesity-related insulin resistance and inflammation. One of these appears to be endoplasmic reticulum (ER) stress (Ozcan et al., 2004). This raises the question as to why there is ER stress in obesity and how obesity-associated ER stress can cause insulin resistance and inflammation. Excessive macronutrient intake is the main cause for obesity and several recent studies in rodents have implicated ER stress as an early sign of nutrient excess and a cause for the development of insulin resistance and inflammation. For instance, in mice fed high fat diets for 3 months, the chronic excessive macronutrient intake caused obesity and ER stress in adipose tissue and liver, whereas overexpression or administration of ER stress reducing chaperone proteins reduced ER stress, insulin resistance, and inflammation (Schroder and Kaufman, 2005). ER stress can lead to the development of insulin resistance and inflammation. Proposed mechanisms include ER stress-induced phosphorylation and activation of C-jun Nterminal kinase ( JNK), and activation and nuclear translocation of nuclear factor kB (NFkB), which is a key promoter of inflammation (Zhang and Kaufman, 2008). In addition, ER stress is a major source for the production of reactive oxygen species (ROS) which can produce insulin resistance and inflammation (Schroder and Kaufman, 2005; Zhang and Kaufman, 2008). Thus, it has been proposed that the ER may be a proximal site that senses nutritional excess and translates that into signals producing insulin resistance and inflammatory responses (Zhang and Kaufman, 2008). Until very recently, however, all the evidence linking ER stress with obesity, insulin resistance, and inflammation were based on in vitro and animal data. We have recently shown, however, that there is ER stress in fat of obese people (Boden et al., 2008). In this chapter, details of this study are described.

2. Study Subjects and Fat Biopsies 2.1. Subjects Six lean and six obese healthy volunteers were studied. None of the participants had a family history of diabetes or other endocrine disorders or were taking medications. Their body weights were stable for at least 2 months before the biopsies. Compared with the nonobese volunteers, the obese volunteers were heavier (93.4 vs. 77.4 kg; P < 0.03) and had more body fat 40.7 vs. 19.9 kg; P < 0.004) but had the same fat-free mass (57.6 vs. 57.6 kg) and were insulin resistant (1/homeostasis model assessment 0.44 vs. 0.29; P < 0.05). Informed written consent was obtained from all subjects after explanation of the nature, purpose, and potential risks of these

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studies. The study protocol was approved by the Institutional Review Board of Temple University Hospital.

2.2. Fat biopsies The subjects were admitted to the Temple University Hospital Clinical Research Center on the day before the studies. At 8 a.m. on the day after admission, a venous blood sample was obtained and an open fat biopsy was performed by a surgeon. Fat biopsies were obtained from the lateral aspect of the upper thigh (15 cm above the patella) under local anesthesia, as described (Boden et al.,1994). The excised fat was dropped immediately into isopentane and kept at its freezing point (160 C) by liquid nitrogen. The frozen fat was stored at 80 C until analyzed.

3. Proteomic Analysis 3.1. Required materials

SYPROÒ Ruby (Invitrogen, Carlsbad, California) Milli-Q System (Millipore, Billerica, MA)

The following is from GE Healthcare, Piscataway, NJ:

IPGphor horizontal electrophoresis apparatus IPGphor strip holders, IEF sample applicator strip Immobiline DryPlate 4–7 and/or 3–10 IPG buffers (pH range 4–7 and/or 3–10) IPG cover fluid DeStreak reagent

The following is from Sigma–Aldrich, St. Louis, MO:

CHAPS Urea Thiourea DTT Iodoacetamide

3.2. Protein extraction protocol for two-dimensional electrophoresis 1. Place the amount of tissue sample to be processed in a precooled (–20 C), clean mortar.


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2. Freeze the tissue sample thoroughly by adding a small volume (2–5 mL) of liquid nitrogen. 3. Allow most of the liquid nitrogen to evaporate, leaving 1–2 mL. 4. Grind the tissue with a precooled (–20 C) pestle until a fine powder is obtained. 5. Transfer the powder into a 1.5-mL tube and measure the wet weight. Add 0.5 mL of extraction buffer: 7 M urea,1 2 M thiourea, 4% CHAPS, 43 mM DTT,2 60 mM Tris/HCl (pH 8.8), and 0.1% SDS; additives: 1 protease inhibitor cocktail2 and 0.2 mM EDTA. 6. Sonicate sample with five pulses of 10 s each. Add 2 mM MgCl2 and Benzonase (50 U per 100 ml of sample solution) mix and incubate at room temperature (RT) for 15 min with vortexing every 5 min.3 7. Spin solution at max speed (21,000g) for 30 min at RT. Discard the pellet. 8. Mix protein extract (supernatant) with 20 C cold acetone/0.1% DTT at 1:3 (v/v) ratio and store the mixture at 20 C overnight. 9. Spin at 21,000g for 30 min at 4 C, discard the supernatant and resuspend the pellet with 1.2 mL 80% (20 C) cold acetone/0.2% DTT2 and spin (21,000g/4 C) for 5 min and discard the supernatant. Repeat wash.4 10. Air dry the pellet till no acetone is remaining and redissolved in DeStreak rehydration solution or two-dimensional electrophoresis (2DE) sample buffer: 7 M urea, 2 M thiourea, 4% CHAPS, 1.2% DeStreak, and 0.01% bromophenol blue. Shaking for 3 h at RT to solubilize. 11. Determine the protein concentration using 2D Quant Kit as per manufacturer’s instructions (Amersham).

3.3. First dimension of 2DE 12. Mix sample to the final volume with DeStreak rehydration buffer supplemented with the corresponding immobilized pH gradient (IPG) buffer. 13. Rehydrate the IPG strips overnight at 22 C in rehydration buffer: 7 M urea, 2 M thiourea, 4% CHAPS, 5% glycerol, 15% 2-propanol, 1.2% DeStreak reagent2, 0.5% IPG buffer 7–112, and 0.2% methylcellulose mixed with an appropriate sample volume.5 1 2 3 4 5

Deionize urea stirring it during 10 min with w/v 0.6% mixed-ion exchanger (e.g., Amberlite MB-1). These reagents should be added fresh. In order to prevent carbamylation modifications, avoid incubation at 37 C. Since benzonase is a protein it may appear on the 2D map. To increase protein recovery, sample can be incubated at 20 C/1 h during the first wash prior spin. Rehydration and sample volumes depend on pH range, separation distance, and detection method.

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14. Use Ethan IPGphor system (GE Health system) to perform isoelectric focusing (IEF). Program the settings to temperature of 20 C and use the following settings. Step T ( C) Start volt (V) IEF protocol (7 cm) 1 20 0 2 20 250 3 20 1500 4 20 3000

End volt (V)

Duration (h) Total (Vh)

250 1500 3000 8000

0.5 1 1 3

11,000

3.4. Second dimension of 2DE 1. Dissolve 100 mg of DTT in 10 mL of equilibration buffer 6 M urea, 30% (w/v) glycerol, and 2% (w/v) SDS in 0.05 M Tris–HCl buffer (pH 8.8). Take out the focused IPG gel strips from the freezer and place them into individual test tubes. Add 10 mL of equilibration buffer I and 50 mL of the bromophenol blue (0.25%, w/v) solution. Seal the test tubes with Parafilm, rock them 15 min on a shaker, and then pour off equilibration buffer I. 2. Dissolve 400 mg of iodoacetamide in 10 mL of equilibration buffer. Add equilibration buffer II and 50 mL of bromophenol blue solution to the test tube as above and equilibrate for another 15 min on a rocker. 3. After the second equilibration, rinse the IPG gel strip with deionized water for a second and place it on a piece of filter paper at one edge for a few minutes to drain off excess equilibration buffer. 4. Place IPG gel strip on top of the vertical SDS gel. 5. Fill the electrophoresis chamber with electrode buffer and turn on the SDS gel cassettes in a vertical position to facilitate the application of the first dimension IPG strips. 6. Equilibrate the IPG gel strips as described above for first dimension and immerse them in electrode buffer for a few seconds. 7. Place the IPG gel strip on top of an SDS gel. Carefully press the IPG strip with a spatula onto the surface of the SDS gel to achieve complete contact. Repeat this procedure for the remaining IPG strips. 8. Insert the gel cassettes in the electrophoresis apparatus and start electrophoresis.6 9. Run the SDS–PAGE gels in Bio-Rad Mini-PROTEANÒ 3 System (Hercules, CA) at 200 V for 45 min. 6

Remove the IPG gel strips from the surface of the vertical SDS gel once the proteins have migrated out of the IPG gel strip.


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10. Terminate the run when the bromophenol blue tracking dye has migrated off the lower end of the gel. 11. Open the cassettes carefully with a spatula. Peel the gel off the glass plate carefully, lifting it by the lower edge, and place it in a tray containing fixing solution or transfer buffer, respectively. Then continue with fixing, protein staining, or blotting.

3.5. Fluorescent staining with SYPROÒ Ruby 1. After electrophoresis, place the gel into a clean container with 100 mL of fixing solution (50% methanol, 7% acetic acid) and agitate on an orbital shaker for 30 min. Repeat once more with fresh fixing solution. Pour off the used fixing solution.7 After fixing, perform three washes in ultrapure water for 10 min each, before proceeding to the staining step. 2. Add 60 mL of SYPROÒ Ruby gel stain. Agitate on an orbital shaker overnight. 3. Transfer the gel to a clean container and wash in 100 mL of wash solution (10% methanol, 7% acetic acid) for 30 min. The transfer step helps minimize background staining irregularities and stain speckles on the gel. Before imaging rinse the gel in ultrapure water a minimum of two times for 5 min to prevent possible corrosive damage to the imager. 4. SYPROÒ Ruby protein gel stain has two excitation maxima, one at 280 nm and one at 450 nm, and has an emission maximum near 610 nm. Proteins stained with the dye can be visualized using a 300 nm UV transilluminator, a blue-light transilluminator, or a laser scanner. The stain has exceptional photostability allowing long exposure times for maximum sensitivity.

3.6. 2DE image analysis 1. Perform 2DE analysis by PDQuest software (Bio-Rad), version 8.0. 2. Enumerate and analyze each gel from obese and lean fat samples for spot detection, background subtraction, and protein spot volume quantification. 3. Make manual corrections to validate the matches automatically generated by the software. 4. Normalize spot volume values in each gel by dividing the raw quantity of each spot by the total volume of all spots included in the same gel. 5. Determine SD for each protein spot and the average spot volume values. 7

For IEF gels, place the gel into a clean container with 100 mL of IEF fix solution and agitate on an orbital shaker for 3 h.

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6. Perform Student’s t-test on the match spots in order to determine the spots that were differentially expressed. 7. Choose spots that show a statistically significant difference with a confidence level of 0.05. 8. To test for inter gel reproducibility, perform 2DE analysis in triplicate using one representative sample.

3.7. In-gel digestion 1. Wearing gloves and sleeve protectors, wipe down ALL surfaces in the hood with methanol/water-moistened lint-free cloth, including the outside of all the tubes (make sure to not wipe off the labeling), the outside and inside of the Speed Vac and centrifuge, tube racks, bottles, etc. Wipe razor blades with methanol-soaked lint-free cloth. 2. Prepare the following solutions: 25 mM NH4HCO3 (100 mg/50 mL) 25 mM NH4HCO3 in 50% ACN 50% ACN/5% formic acid (may substitute TFA or acetic acid) 12.5 ng/mL trypsin (Promega) in 25 mM NH4HCO3 (freshly diluted) 3. Dice each gel slice into small pieces (1 mm2) and place into 0.65-mL siliconized tubes (PGC Scientific). 4. Add 100 mL (or enough to cover) of 25 mM NH4HCO3/50% ACN and vortex for 10 min. 5. Using gel loading pipet tip, extract the supernatant and discard. 6. Repeat steps 3 and 4 once or twice. 7. Speed Vac the gel pieces to complete dryness ( 20 min). 8. Prepare fresh solutions: 10 mM DTT in 25 mM NH4HCO3 (1.5 mg/mL) 55 mM iodoacetamide in 25 mM NH4HCO3 (10 mg/mL) 9. Add 25 mL (or enough to cover) 10 mM DTT in 25 mM NH4HCO3 to dried gels. Vortex and spin briefly. Allow reaction to proceed at 56 C for 1 h. 10. Remove supernatant, add 25 mL 55 mM iodoacetamide to the gel pieces. Vortex and spin briefly. Allow reaction to proceed in the dark for 45 min at RT. 11. Remove supernatant (discard). Wash gels with 100 mL NH4HCO3, vortex 10 min, spin. 12. Remove supernatant (discard). Dehydrate gels with 100 mL (or enough to cover) of 25 mM NH4HCO3 in 50% ACN, vortex 5 min, spin. Repeat one time. 13. Speed Vac the gel pieces to complete dryness (20 min). Proceed with trypsin digest.


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14. Add trypsin solution (step 2) to just barely cover the gel pieces. Estimate the gel volume and add about 3 volume of trypsin solution. This volume will vary from sample to sample, but on average 5–25 mL is sufficient. 15. Rehydrate the gel pieces on ice or at 4 C for 10 min Spin. Add 25 mM NH4HCO3 as needed to cover the gel pieces. 16. Spin briefly and incubate at 37 C for 4 h—overnight.

3.8. Extraction of peptides 1. Transfer the digest solution (aqueous extraction) into a clean 0.65-mL siliconized tube. 2. To the gel pieces, add 30 mL (enough to cover) of 50% ACN/5% formic acid, vortex 20–30 min, spin, sonicate 5 min. Repeat. 3. Vortex the extracted digests, spin and Speed Vac to reduce volume to 10 mL. 4. Either proceed with C18 ZipTip (Millipore) cleanup or analyze with LC–MS. Add 2–5 mL of 5% formic acid. When analyzing low levels of protein, concentrate the peptides by eluting from ZipTips using 3 mL of elution solution, into a clean 0.65-mL siliconized tube. 5. Use 1 mL of the unseparated digests for analysis by Matrix Assisted Laser Desorption Ionization/Time of Flight (MALDI/TOF).

3.9. Preparation of MALDI matrix 1. Dissolve the contents of a 10-mg tube of a-cyano-4-hydroxycinammic acid in 1 mL of the 50% acetonitrile in 0.05% triflouroacetic acid solution. For best performance, once in solution, the matrix should be stored in the dark and used within 1 week. 2. Some residual crystals may be visible in the matrix solution. The acetonitrile concentration can be adjusted to suit individual preferences. A mixture of 70% ACN and 30% of the 0.1% TFA solution can also be used.

3.10. Sample plate preparation and MALDI-TOF/TOF analysis 1. Transfer 10 mL of the matrix solution to a small tube. 2. Add 1 mL of the standard/sample to the tube containing the matrix and vortex. 3. Apply 1.0 mL from the mixture to wells of an AnchorChipTM sample target plate used for the Bruker Auto-flex MALDI-TOF/TOF. 4. Once the liquid has evaporated (2 h), the target is ready for analysis.

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5. Obtain peptide mass fingerprints using the reflective and positive ion mode of Autoflex MALDI TOF/TOF mass spectrometer (Bruker, Daltonics Inc., Billerica, MA) 6. Use FlexAnalysisTM software with signal-to-noise ratio of 2:1 to collect mass spectra from 100 to 400 laser shots. 7. Generate mono-isotopic peaks by mass peak value calculation. 8. Use two trypsin auto-digestion peptides with M þ H values 842.509 and 2211.104 as internal standards. 9. Indentify protein by matching the calibrated peptide mass values within NCBInr protein databases using an in-house version of Mascot Server 2.2 imbedded in Bruker’s Biotool software. 10. Select Homo sapiens species for taxonomy. 11. Use the following match variances: mass tolerance of 40 ppm, one missed trypsin cleavage, fixed modification of carbamidomethyl cysteine, and variable modification of methionine oxidation.

3.11. Results of proteomic analysis Gels with isoelectric focusing ranges of PI 4–7 and 6–10 produced a total of 900 protein spots in each gel (Fig. 4.1). The comparison of all spots visualized yielded 24 spots that were significantly different in lean versus obese volunteers. Three of these spots represented multiple isoforms of vimentin. Three vimentin isoforms were considered as one protein (24 2 ¼ 22). Two other spots could not be identified (22 2 ¼ 20). Of 20 remaining differentially expressed proteins, 10 were upregulated and 1 (a-enolase) was downregulated in obese versus lean volunteers (Table 4.1). The differentially expressed proteins were grouped into the following categories: (1) UPR and stress (seven proteins), (2) energy and FFA metabolism (five proteins), (3) structural proteins (four proteins), and (4) protein transport and signaling (four proteins). UPR and stress. Levels of expression of the following UPR proteins were overexpressed in 2DEs in adipose tissue from obese volunteers: CRT, a protein chaperone, increased from undetectable to 1019 236 arbitrary units; PDI, a protein foldase, increased approximately threefold; and glutathione-S-transferase P, an antioxidant protein belonging to a UPR-upregulated pathway increased 1.8-fold. Several cytosolic small heat-shock proteins (HSPs) (20 and 27 kDa HSP) and one mitochondrial HSP (60 kDa HSP) were also overexpressed in the adipose tissue of obese volunteers, suggesting the presence of cytosolic and mitochondrial stress in addition to ER stress (Table 4.1).

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Lean

Obese PDI PDI Calreticulin

Lean

Obese

14 9 12 7 8

7

19

19

8 9 12

3 2

3

4

2

18 5

18

4 11

5

11

15

16

13

16 13

15 17 1

6

1

20

14

20

17

6 10

10

pI 4

7 pI 4

7

Lean

Obese

21

21

23

23 24

24

22

pI 6

22

10 pI 6

10

Figure 4.1 Expression of the UPR proteins CRT and PDI in adipose tissue of one lean and one obese subject (upper panels). Differentially expressed proteins in fat homogenates from one lean and one obese subject. Proteins were separated by isoelectric focusing and molecular weight (2DE) as described in Section 3. The subproteom from each sample was assessed using PI ranges of 4–7 (middle panels) and 6–10 (lower panels). The proteins were stained with SPYROÒ Ruby and images compared by PD Quest software. The numbers correspond to the spot numbers in Table 4.1. The arrows indicate upregulated proteins in the fat of obese subjects (from Boden et al., 2008 with permission).

Table 4.1 Proteins differentially expressed in 2DGs

Spot no.

Protein identification

UPR and stress 3 CRT 23 PDI 20 20 kDa HSPb-6 21 27 kDa HSPb-1 25 HSPb-5 16 60 kDa HSP 19 Glutathione-S-transferase P Energy and FFA metabolism 12 ATP synthase subunit-b 15 Perilipin 22 Aldehyde dehydrogenase 24 a-Enolase 26 Carbonic anhydrase-1 Structural proteins 1 Myosin light-chain polypeptide-6 2 Tropomyosin b-chain 4 Tropomyosin a4-chain 7–9 Vimentin Protein transport and signaling 6 g-Synuclein 11 pGDP dissociation inhibitor-1 5 14-3-3 protein-g 10 Galectin-1 Data are means S.D.

Swiss protein accession no.

Mouse score

Peptides matched

Normalized spot volumes SD (lean; n ¼ 6)

Normalized spot volumes SD (lean; n ¼ 6)

P27797 P30101 014558 P04792 P02511 P10809 P09211

66 61 76 78 74 69 107

7 6 5 8 9 6 8

0 421 328 2534 1157 4242 1438 4162 558 600 558 2231 698

1019 1170 7530 8140 8967 1868 3936

P06576 060240 P05091 P06733 P00915

138 57 87 65 198

11 6 9 7 11

2413 801 1602 381 1341 161 803 214 2463 774

5499 1251 3960 1019 4088 799 293 74 4846 523

0.01 0.007 0.001 0.018 0.012

P60660

60

6

1792 528

3837 284

0.002

P07951 P67936 P08670

58 94 204

6 8 19

1141 443 770 355 3136 1418

4072 531 3429 779 11,005 3353

0.03 0.002 0.007

076070 P52565

66 75

4 5

488 465 2163 612

1714 670 3168 286

0.034 0.049

P61981 P09382

78 113

6 10

875 525 9030 1387

3619 1005 25,049 4175

0.005 0.001

236 63 248 1643 2918 56 390

P

< 0.001 < 0.01 0.001 0.001 0.049 < 0.001 0.013


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4. Western Blot Analysis 4.1. Required materials

Blotting chamber (Bio-Rad) Protein disulfide isomerase A3 (Santa Cruz, CA, SC-20132) Calreticulin (Santa Cruz, CA, SC-11398) Calnexin (Santa Cruz, CA, SC-11397) Mouse anti-rabbit horseradish peroxidase-conjugated secondary antibody (Biomeda, Foster City, CA)

4.2. Expression analysis 1. Separate the proteins (30–80 mg) from the same adipose tissue lysates as used for the 2DEs by 10–14% gradient SDS-PAGE. 2. Transfer the separated proteins to a nitrocellulose membrane in a semidry blotting chamber according to the manufacturer’s protocol. 3. Block the blots with 5% milk in Tris-buffered saline solution (pH 7.6) containing 0.05% Tween-20. 4. Probe with the following rabbit anti-human antibodies from Santa Cruz Biotechnology (Santa Cruz, CA) at a concentration of 0.4 mg/mL: protein disulfide isomerase A3 (PDI), calreticulin (CRT), and calnexin (CNX). 5. Use a rabbit antiserum that detects phosphor c-jun NH2-terminal kinase ( JNK)-1, -2, and -3 (Cell Signaling Technology, Danvers, MA) and a rabbit antiserum that detects total JNK. 6. Incubate the blots with primary antibody overnight at 4 C at with gentle shaking and then incubate with a mouse anti-rabbit horseradish peroxidase-conjugated secondary antibody (1:10,000) (Biomeda) for 1 h at RT. 7. Expose the blots using a chemiluminescent detection method (Enhanced ECL Detection System; GE Healthcare BioSciences).

4.3. Results of Western blot analysis Western blotting confirmed upregulation of CRT and PDI and revealed upregulation of CNX ( 1.8-fold), a membrane bound chaperone and phospho-JNK-1 2.0-fold, a downstream effector protein of the UPR. Phospho-JNK 2/3, however, was unchanged (Fig. 4.2).

80

p-JNK-1

cnx b -actin

0.8

JNK-1

0.8

P < 0.05 P = 0.024

0.6

0.4

0.4 0.2

Calreticulin/b -actin

0.0 1.5

0.0 p-JNK-2,3

crt b -actin

P < 0.005

ns

0.8

1.0

0.4

0.5 0.0 2.0

PDI/b -actin

1.2

JNK-2,3

1.5

Lean

PDI b -actin

Obese

0.0

Phospho-JNK-2,3/JNK-2,3

Calnexin/b -actin

1.0

Phospho-JNK-1/JNK-1


P = 0.05

1.0 0.5 0.0

Lean

Obese

Figure 4.2 Protein abundance (by Western blots) of CNX/b-actin, CRT/b-actin, PDI, phospho-JNK-1/JNK-1, and phospho-JNK-2,3/JNK-2,3 in subcutaneous adipose tissue from four lean (insufficient fat was available for Western analysis from two of six lean subjects) and six obese nondiabetic subjects. Inserts show representative Western blots (from Boden et al., 2008 with permission).

5. RT-PCR Analysis Total RNAs were isolated from frozen adipose tissues, and real-time RT-PCR was performed with a SYBR Green One-Step qRT-PCR kit (Invitrogen) and a Light-Cycler (Roche, Indianapolis, IN). Primers for X-box-binding protein (XBP)-1s (NM-005080) were sense: TTGAGAACCAGGAGTTAAG and antisense CCTGCACCTGCTGCGGACT.

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XBP-1

0.8

12 mRNA/18S rRNA arbitrary units

TNF-a P = 0.03 0.6

8 P = 0.024

0.4

4

0.2

0

0.0 IL-6

IL-1b 0.8

0.8

0.6

0.6

0.4

0.4

02

02

0.0

Lean

Obese

Lean

Obese

0.0

Figure 4.3 Messenger RNA (mRNA) corrected for 18S ribosomal RNA (18S rRNA) of sXBP-1, tumor necrosis factor-a, interleukin-1b, and interleukin-6 in six lean and six obese nondiabetic subjects (from Boden et al., 2008 with permission).

5.1. Results of RT-PCR analysis Upregulation of the spliced form of XPP-1s, a part of the IRE-1/XBP-1 proximal ER stress sensor, and of tumor necrosis factor-a, a proinflammatory cytokine, were documented with RT-PCR, whereas there were no differences in interleukin-1b and -6 mRNA in adipose tissue of lean and obese subjects (Fig. 4.3).

ACKNOWLEDGMENTS This work was supported by National Institutes of Health grants R01-DK58895, a grant from the American Diabetes Association (1-10-CT-06), a grant from the Department of Health, Commonwealth of Pennsylvania, and a grant from the Groff Foundation (all to G. B.) and R01-A1064017 (to S. M.).

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REFERENCES Boden, G., Chen, X., Ruiz, J., and Rossetti, L. (1994). Mechanisms of fatty acid induced inhibition of glucose uptake. J. Clin. Invest. 93, 2438–2446. Boden, G., Lebed, B., Schatz, M., Homko, C., and Lemieux, S. (2001). Effects of acute changes on plasma FFA on intramyocellular fat content and insulin resistance in healthy subjects. Diabetes 50, 1612–1617. Boden, G., She, P., Mozzoli, M., Gumireddy, K., Reddy, P., Xiang, X., Luo, Z., and Ruderman, N. (2005). Free fatty acids produce insulin resistance and activate the proinflammatory NFkB pathway in rat liver. Diabetes 54, 3458–3465. Boden, G., Duan, X., Homko, C., Molina, E. J., Song, W., Perez, O., Cheung, P., and Merali, S. (2008). Increase in endoplasmic reticulum stress-related proteins and genes in adipose tissue of obese insulin-resistant individuals. Diabetes 57, 2438–2444. Bray, G. A. (2004). Medical consequences of obesity. J. Clin. Endocrinol. Metab. 89, 2583–2589. Itani, S. I., Ruderman, N. B., Schmieder, F., and Boden, G. (2002). Lipid-induced insulin resistance in human muscle is associated with changes in diacylglycerol, protein kinase C, and IkB-a. Diabetes 51, 2005–2011. Ozcan, U., Cao, Q., Yilmaz, E., et al. (2004). Endoplasmic reticulum stress links obesity, insulin action, and type 2 diabetes. Science 306, 457–461. Reaven, G. M., Hollenbeck, C., Jeng, C.-Y., Wu, M. S., and Chen, Y. D. (1988). Measurement of plasma glucose, free fatty acid, lactate and insulin for 24 h in patients with NIDDM. Diabetes 37, 1020–1024. Santomauro, A. T., Boden, G., Silva, M., Rocha, D. M., Santos, R. F., Ursich, M. J., Strassmann, P. G., and Wajchenberg, B. L. (1999). Overnight lowering of free fatty acids with acipimox improves insulin resistance and glucose tolerance in obese diabetic and nondiabetic subjects. Diabetes 48, 1836–1841. Schroder, M., and Kaufman, R. J. (2005). The mammalian unfolded protein response. Annu. Rev. Biochem. 74, 739–789. Yu, C., Chen, Y., Cline, G. W., et al. (2002). Mechanism by which fatty acids inhibit activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3-kinase activity in muscle. J. Biol. Chem. 277, 50230–50236. Zhang, K., and Kaufman, R. J. (2008). From endoplasmic-reticulum stress to the inflammatory response. Nature 454, 455–462.

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Measurement of the Unfolded Protein Response (UPR) in Monocytes Toma´s P. Carroll, Catherine M. Greene, and Noel G. McElvaney Contents 84 84 85 86

1. Introduction 1.1. ER stress and the unfolded protein response 1.2. The monocyte 2. Investigating UPR Activation in the Monocyte 2.1. Isolation, culture, and treatment of peripheral blood monocytes 2.2. Enzyme-linked immunosorbent assay 2.3. RNA isolation, cDNA synthesis, and RT-PCR 2.4. Detection of UPR markers by immunoblotting 2.5. Immunofluorescence Acknowledgments References

86 88 89 91 92 93 93

Abstract In mammalian cells, the primary function of the endoplasmic reticulum (ER) is to synthesize and assemble membrane and secreted proteins. As the main site of protein folding and posttranslational modification in the cell, the ER operates a highly conserved quality control system to ensure only correctly assembled proteins exit the ER and misfolded and unfolded proteins are retained for disposal. Any disruption in the equilibrium of the ER engages a multifaceted intracellular signaling pathway termed the unfolded protein response (UPR) to restore normal conditions in the cell. A variety of pathological conditions can induce activation of the UPR, including neurodegenerative disorders such as Parkinson’s disease, metabolic disorders such as atherosclerosis, and conformational disorders such as cystic fibrosis. Conformational disorders are characterized by mutations that modify the final structure of a protein and any cells that express abnormal protein risk functional impairment. The monocyte is an important and long-lived immune cell and acts as a key immunological orchestrator, dictating the intensity and duration of the host immune response. Department of Medicine, Royal College of Surgeons in Ireland, Education and Research Centre, Beaumont Hospital, Dublin, Ireland Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00005-4

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Monocytes expressing misfolded or unfolded protein may exhibit UPR activation and this can compromise the host immune system. Here, we describe in detail methods and protocols for the examination of UPR activation in peripheral blood monocytes. This guide should provide new investigators to the field with a broad understanding of the tools required to investigate the UPR in the monocyte.

1. Introduction 1.1. ER stress and the unfolded protein response Newly synthesized secreted and transmembrane proteins are transported into the lumen of the endoplasmic reticulum (ER) where they are folded and correctly assembled. The homeostasis of this organelle is vital to continued normal cell function and survival. Perturbations that disrupt ER homeostasis, such as fluctuations in calcium storage or increased demand for protein folding arising from elevated production of secretory proteins, compromise ER protein folding capacity resulting in the accumulation of unfolded or misfolded protein within the lumen of the ER. An imbalance between the load of unfolded proteins entering the ER and the ability of the ER to process this load is termed ER stress, and can be detrimental to cell survival. ER stress can also be induced by a range of pathophysiological conditions, including stroke, ischemia, diabetes, obesity, viral infection, and mutations that impair protein folding (Kaufman, 1999). To maintain homeostasis, the ER has evolved highly specific signaling pathways collectively referred to as the unfolded protein response (UPR), which strives to adapt for survival or induce apoptosis. In the event of ER stress, this protective pathway causes the translational attenuation of general protein synthesis, induces a transcriptional program of ER-resident chaperones, and coordinates the degradation of misfolded proteins through ERassociated degradation (ERAD) (Mori, 2000). Three proximal ER-resident transmembrane sensors, protein kinase RNA (PKR)-like ER kinase (PERK), inositol-requiring kinase 1 (IRE1), and activating transcription factor 6 (ATF6) form a tripartite management system that orchestrates the mammalian UPR (Ron and Walter, 2007). The luminal domain of these three molecules is sequestered by the ER chaperone glucose-regulated protein 78 (GRP78), however, upon misfolded protein accumulation, GRP78 dissociates from PERK, IRE1, and ATF6, leading to their activation (Malhotra and Kaufman, 2007). The most immediate step in the UPR is translational attenuation to prevent further accumulation of unfolded proteins in stressed cells. This is mediated by PERK, through phosphorylation of eukaryotic initiation factor 2 (eIF2) at Serine 51 on its alpha subunit (eIF2a) (Harding et al., 1999).

Measuring UPR Activation in the Monocyte

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While reducing the rate of protein synthesis and the protein load in the ER, eIF2a phosphorylation paradoxically promotes the translation of ATF4 mRNA, a basic leucine zipper (bZIP) transcription factor. ATF4 induces a pro-survival program of genes essential for amino acid import, glutathione biosynthesis, and resistance to oxidative stress (Harding et al., 2003). The next step in the UPR employs IRE1 and ATF6, which regulate chaperone induction, expansion of the ER in response to increased client protein load, and ERAD (Mori, 2000). Upon UPR engagement, IRE1 is activated and its endoribonuclease activity causes removal of a 26-nucleotide intron from X-box-binding protein 1 (XBP-1) mRNA (Tirasophon et al., 1998). The spliced XBP-1 mRNA is translated into a potent bZIP transcription factor which translocates to the nucleus and acts as a key regulator of ER folding capacity. Concurrently, another bZIP transcription factor ATF6 is released from GRP78 and transported to the Golgi where it undergoes sequential cleavage by site-1 protease (S1P) and site-2 protease (S2P). Cleaved ATF6 cooperates with spliced XBP-1 to induce the expression of ER chaperones, ER quality control genes, folding enzymes, and ERAD (Yoshida et al., 1998, 2001). ERAD ensures the removal of terminally misfolded proteins from the ER lumen to the cytoplasm for ultimate degradation by the ubiquitin–proteasome system (Travers et al., 2000).

1.2. The monocyte The human immune system has been historically divided into innate immunity and acquired or adaptive immunity. Monocytes and their multitude of differentiated offspring fulfill important regulatory and effector roles in both arms of the immune system (Medzhitov and Janeway, 2000). Peripheral blood monocytes in the circulation migrate through blood vessel walls into various organs and differentiate into more specialized macrophages. Monocytes and their macrophage progeny act as immunological orchestrators and serve three major functions, antigen presentation, phagocytosis, and immunomodulation (Dale et al., 2008). Upon activation, monocytes and macrophages produce large amounts of cytokines and chemokines that recruit other immune cells such as neutrophils to sites of infection and inflammation. The first reports of ER stress and UPR activation in immune cells were in professional antibody-secreting plasma cells (Calfon et al., 2002). This developmental ER stress is a natural consequence of the transition from B cell to plasma cell, which produces huge amounts of antibodies and requires a massive expansion in ER folding capacity (Iwakoshi et al., 2003). However, there is emerging evidence that UPR activation in monocytes and macrophages may play a role in the pathogenesis of a number of diseases. In mouse models of atherosclerosis, macrophages overloaded with free cholesterol were shown to exhibit UPR activation (Li et al., 2005), and

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monocytes from Type II diabetes patients showed elevated expression of GRP78 (Komura et al., 2010). Monocytes from cystic fibrosis (CF) patients were shown to be intrinsically abnormal in their cytokine responses (Zaman et al., 2004), but this can be reversed by inhibitors designed to prevent DF508 CFTR (CF transmembrane conductance regulator) degradation by ERAD and increase the secretion of CFTR (Vij et al., 2006). In addition, work from this group has demonstrated intracellular accumulation of Z alpha-1 antitrypsin (AAT) in the ER of monocytes, and this causes sustained activation of the UPR with subsequent effects on immune function (Carroll et al., 2010). As the tissue macrophage can present a challenge in terms of isolation with unreliable sampling techniques or lengthy differentiation protocols from monocytes that risk cell activation, the monocyte is a useful tool for investigating immune system derangement in a variety of disorders.

2. Investigating UPR Activation in the Monocyte 2.1. Isolation, culture, and treatment of peripheral blood monocytes In this section, we will provide details for isolating monocytes from freshly drawn whole blood from human donors. It must be noted that studying the behavior and function of human monocytes is a difficult undertaking. Monocytes constantly observe their surroundings and swiftly react to changes, as is their nature. The isolation, purification, and culture of these cells in vitro can affect their phenotype, and care must be taken not to stimulate the cells. The faster the isolation process the less chance of cell activation, and all buffers to be used should be prewarmed to 37 C. We use Lymphoprep (Axis-Shield) to perform density gradient separation but Ficoll-Paque can also be used (Yeo et al., 2009). 2.1.1. Required materials

Lymphoprep (Axis-Shield) 1 saline (0.9% NaCl) Hanks balanced salt solution (Biosciences) EasySep Human CD14 positive selection kit (Stemcell Technologies) and EasySep magnet (Stemcell Technologies). The kit is designed to isolate CD14 positive cells from fresh or previously frozen peripheral blood mononuclear cells by positive selection. 5 mL polystyrene round bottom Falcon tube (BD Biosciences, product)


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RPMI 1640 (Biosciences) supplemented with 10% FCS, 2% glutamine and penicillin/streptomycin (necessary for culturing monocytes) Thapsigargin (Molecular Probes) and tunicamycin (Sigma) for chemically inducing UPR activation. Thapsigargin depletes calcium stores in the ER by inhibiting Ca2þ ATPase, while tunicamycin inhibits N-linked glycosylation of newly synthesized proteins. 2.1.2. Monocyte isolation Mononuclear cells are isolated from venous peripheral blood obtained from donors by density gradient separation. The blood must be collected in tubes containing an anticoagulant (EDTA or heparin). Once drawn, blood is immediately placed in 50-mL tube and mixed with an equal volume of 0.9% NaCl (1 saline). The diluted blood is then layered over Lymphoprep in a fresh 50-mL tube. For example, if 15 mL blood is obtained, mix with 15 mL 1 saline and then slowly and carefully layer over 15 mL Lymphoprep. Alternatively, Lymphoprep can be under-layered below the diluted blood using a long pipette. It is essential to avoid mixing of blood and Lymphoprep before centrifugation. Centrifuge the Lymphoprep and blood at 800g for 10 min at room temperature, with the centrifuge brake switched off. After centrifugation, the mononuclear cells (which include monocytes and lymphocytes) form a distinct band at the blood/Lymphoprep interface. Carefully remove this band using a Pasteur pipette and place in a fresh tube. Add an equal volume of 1 saline or HBSS to reduce the density of the solution and pellet these cells by gentle centrifugation (300 g, 10 min). Resuspend cells in 1 mL EasySep recommended medium (1 mM EDTA, 2% fetal calf serum (FCS) in PBS Dulbecco) and transfer this 1 mL cell suspension to a 5-mL Falcon tube. It is useful to count the number of mononuclear cells isolated at this step. Monocytes should typically represent 10% of this total mononuclear fraction. From this point, we follow the EasySep CD14 kit protocol exactly as described in order to isolate monocytes from the mononuclear cell population. In our experience, the most critical step in the Lymphoprep protocol is to ensure the centrifuge brake is switched off as this will ensure a tight band of mononuclear cells forms at the interface between serum and Lymphoprep. Another critical step is the careful layering of diluted blood over Lymphoprep to ensure no mixing occurs between the layers. Leaving the centrifuge brake on or inefficient layering technique will result in a diffuse mononuclear band at the interface, and a poor monocyte yield. Once monocytes are successfully isolated and counted they can be used in a variety of applications. For example, if performing stimulation experiments, monocytes can be cultured in RPMI containing 10% (v/v) FCS and 1% penicillin/streptomycin at 37 C in a 5% CO2 atmosphere for up to 24 h. However, it is essential that monocytes are cultured in polypropylene tubes,

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and not polystyrene tubes, as polystyrene promotes significant adherence and probable activation of monocytes.

2.2. Enzyme-linked immunosorbent assay In addition to its role in maintaining ER homeostasis, it has become apparent that ER stress-induced UPR activation can influence the expression of a subset of inflammatory genes. The spliced form of XBP-1 has been shown to control the production of IL-6 in the mouse B cell (Iwakoshi et al., 2003). XBP-1 was also shown to regulate the production of IFN-b in mouse macrophages subjected to ER stress (Smith et al., 2008). Another study in a mouse model of atherosclerosis demonstrated that accumulation of free cholesterol in macrophages caused UPR activation and the secretion of significant amounts of TNF-a and IL-6 (Li et al., 2005). Thus, the underestimated technique of enzyme-linked immunosorbent assay (ELISA) is a reliable method of investigating an aberrant immune response in monocytes, often a consequence of UPR activation. This quantitative method is highly reproducible, relatively cheap, and easy to perform. However, due to the large quantities of chemokines and cytokines secreted by monocytes, dilution of supernatants is often required and as little as 2 mL of supernatant can be sufficient in some cases, for example, when assaying IL-8. Another application for ELISA is in the investigation of impaired secretion of specific proteins. For example, we have used this technique to show impaired secretion of AAT from monocytes isolated from AATdeficient individuals (Carroll et al., 2010). 2.2.1. Required materials for IL-8 ELISA

Monocyte supernatants Immulon 2HB 96 well high-binding microtitre plates (Thermo Electron Corporation) Voller’s coating buffer (100 mM bicarbonate/carbonate buffer, pH 9.6) Wash buffer (0.05% Tween in PBS, pH 7.4) Blocking buffer (1% BSA, 0.05% Tween in PBS, pH 7.4) Recombinant human IL-8 (R&D Systems) Monoclonal antibody to human IL-8 (MAB208, R&D Systems) Biotinylated antibody to human IL-8 (BAF208, R&D Systems) Streptavidin–horseradish peroxidase (HRP) (Biolegend) ABTS substrate (Invitrogen)

2.2.2. ELISA to measure IL-8 Supernatants are recovered and protein concentrations determined by ELISA with specific antibodies to the secreted protein of interest, in this example, the chemokine IL-8. The day before, a high-binding 96-well


89

plate is coated with 100 mL per well of capture antibody (MAB208, 1/500 dilution) diluted in Voller’s buffer—this is a key point. If PBS is used as a diluent for the coating antibody, as stated in the manufacturer’s datasheet, the ELISA will not work. The plate is sealed and incubated overnight at 4 C. Next day, each well is aspirated and filled with wash buffer and repeated for a total of three washes using a multichannel pipette. Complete removal of the liquid at each step is achieved by inverting the plate and blotting on clean tissue paper. Plate is blocked by the addition of 200 mL of blocking buffer and incubated for 1 h at room temperature. Repeat aspiration and wash steps as before for three washes. Plates are filled with 100 mL per well of monocyte supernatant (diluted). Serial dilutions of recombinant human IL-8 in serum-free RPMI medium are performed to construct a standard curve (31.25–2000 pg/mL) and the plate is incubated for 2 h at room temperature. Plate is washed three times as before, 100 mL of the detection antibody (BAF208, 1/2500 dilution) is added to each well, and plate is incubated for a further 2 h. Next, plate is washed three times, 100 mL streptavidin–HRP is added (1/2500 dilution) and incubated for 30 min at room temperature. Another three washes are performed, and the plate is incubated with 100 mL ABTS for 5–30 min in a dark box or cupboard protected from direct light. It is useful to check the plate periodically for the development of a green color, and once a significant color is observed, measure the absorbance of each well at 405 nm on a Victor2 microplate reader (Wallac). Prepare a standard curve from the data produced from the serial dilutions with concentration on the X axis versus absorbance on the Y axis (linear). Interpolate the concentration of the sample from this standard curve using Prism 4.0 statistical analysis software (GraphPad). As an ancillary step to ELISA, cytokine arrays can be employed to measure multiple proteins in the monocyte supernatants (RayBiotech, Inc.). This technique, although expensive, allows the semiquantitative measurement of over 100 cytokines, chemokines, proteases, antiproteases, and growth factors from a single sample. Any potential leads can then be confirmed by ELISA or quantitative RT-PCR.

2.3. RNA isolation, cDNA synthesis, and RT-PCR There are inherent difficulties involved in measuring the activation of the major UPR orchestrators. The phosphorylation of PERK and IRE1 and the proteolytic cleavage of ATF6 are hallmarks of their activation. However, these proteins are expressed at very low levels and detection is hindered by a lack of high-quality commercial antibodies. As surrogate markers of activation, a number of UPR genes downstream of these proximal sensors have been identified, and can be easily measured by realtime PCR. For example, ATF4 is selectively activated by PERK-induced phosphorylation of eIF2a, and while IRE1 activation is difficult to

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reproducibly and consistently measure, the processing of unspliced XBP-1 mRNA into mature spliced XBP-1 (sXBP-1) mRNA is a convenient, widely used indicator of IRE1 activation (Marciniak et al., 2004). The expression of several other UPR-responsive genes can be easily measured in this way, including calreticulin, calnexin, CHOP, ERdj5, p97/VCP, GRP78, and GRP94. 2.3.1. Required materials

TRI reagent (Sigma) QuantiTect reverse-transcription cDNA synthesis kit (Qiagen) SYBR Green I Master mix (Roche) for real-time quantitative RT-PCR GoTaq Green Master mix (Promega) for conventional RT-PCR Oligonucleotide primers (Eurofins MWG Operon)

2.3.2. Real-time PCR For analysis of UPR markers, 500 mL TRI reagent is added to the isolated monocytes (in a fume hood) and RNA is recovered according to the manufacturer’s instructions. We prefer to resuspend RNA in 0.1% diethylpyrocarbonate (DEPC)-treated water, as DEPC is an efficient, nonspecific inhibitor of RNases. Store the isolated RNA at 80 C and avoid repeated freeze–thaw cycles. Quantify RNA concentration on a spectrophotometer prior to each separate cDNA synthesis. Equal quantities of RNA are then reverse transcribed into cDNA using the Quantitect Reverse Transcription kit. The resulting cDNA is used as the template for quantitative real-time PCR. Oligonucleotide primers are synthesized specific to each target UPR gene and quantitative PCRs performed containing 2 mL template cDNA, 10 mL 2 SYBR Green I master mix, 10 pmol of forward and reverse primers, and nuclease-free water to give a final volume of 20 mL. Amplification is performed on the Roche LightCycler 480 PCR system with the expression of target genes relative to the housekeeping gene b-actin determined using the 2 DDCT method (Livak and Schmittgen, 2001) (Fig. 5.1). 2.3.3. Analysis of XBP-1 mRNA cleavage XBP-1 mRNA splicing is analyzed in our laboratory using a conventional semiquantitative RT-PCR assay developed by Harding et al. (Calfon et al., 2002). RNA is isolated using TRI reagent and cDNA synthesized as described. The resulting cDNA is template in a reaction with GoTaq master mix and specific XBP-1 forward and reverse primers. The primers used to amplify XBP-1 cDNA: forward 50 -AAACAGAGTAGCAGCTCAGACTGC-30 ; reverse 50 -TCCTTCTGGGTAGACCTCTGGGA-30 . PCR products are resolved on a 2.5% agarose gel with unspliced XBP-1 yielding a product of 480 bp, while spliced XBP-1 is 454 bp. Agarose gel

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40 Relative mRNA

CRT

30

sXBP-1 GRP78

20 10 0 0

100 Thapsigargin (nM)

300

Figure 5.1 Quantitative RT-PCR analysis of calreticulin (CRT), sXBP-1, and GRP78 mRNA induction in peripheral blood monocytes after treatment with thapsigargin for 4 h.

electrophoresis may need to be performed for up to 2 h to detect an appreciable difference between spliced and unspliced XBP-1.

2.4. Detection of UPR markers by immunoblotting As previously described, the phosphorylation of PERK and IRE1 and the proteolytic cleavage of ATF6 are difficult to detect. However, the activation of a number of other downstream UPR proteins such as ATF4, CHOP, GRP78, and p97/VCP can be detected by Western blotting. 2.4.1. Immunoblotting method Monocytes are isolated as described, pelleted by gentle centrifugation (400g, 5 min) and cells resuspended in 1 mL hypotonic buffer (10 mM Hepes (pH 7.9), 1.5 mM MgCl2, 10 mM KCl, 0.5 mM PMSF, and 0.5 mM DTT) (Sigma). Cells are lysed for 30 min on ice before centrifugation at 14,000g for 10 min at 4 C. The supernatant is then recovered for immunoblotting and stored at 80 C until required. Protein concentration in the monocyte lysate is determined by the method of Bradford (1976). Whole cell lysates are separated by electrophoresis on SDS-polyacrylamide gel and transferred to a PVDF membrane (Sigma). The percent polyacrylamide used will depend on the size of the UPR protein being investigated. PVDF membranes can be cut to investigate the levels of multiple proteins if there is sufficient difference in the size of the proteins being examined, and this can eliminate the need to strip and reprobe membranes. Nonspecific binding is blocked with 5% bovine serum albumin (Sigma) in PBS containing 0.1% Tween-20 (Sigma). Immunoreactive proteins are detected by incubating the membrane with specific antibodies to UPR proteins of interest, for example, the excellent anti-KDEL antibody

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which detects GRP78 and GRP94 (Stressgen), and comparing with appropriate loading controls such as GAPDH or b-actin.

2.5. Immunofluorescence Immunofluorescence is a powerful tool that can be used to demonstrate the accumulation of misfolded protein within the ER. For example, we have used this technique to show intracellular accumulation of AAT within the ER of monocytes from AAT-deficient individuals, as well as increased GRP78 and GRP94 expression (Carroll et al., 2010). 2.5.1. Required materials

Poly-L-lysine coated slides (Sigma) 4% paraformaldehyde (Sigma) diluted in PBS 0.2% Triton X-100 diluted in PBS (permeabilization buffer) 4% BSA/1% gelatin in PBS (blocking buffer) Vectashield (Vector Laboratories)

2.5.2. Colocalization using immunofluorescence This method is used to investigate AAT localization to the ER by doublestaining monocytes with an antibody to AAT and an antibody to ERresident chaperones containing the tetrapeptide KDEL motif. Monocytes are isolated as described and 100 mL of monocyte suspension pipetted onto a poly-L-lysine glass slide (“P” side up). Score the slide with a diamond pen to indicate the position of cells. Poly-L-lysine coated slides should be used as they provide higher adhesion, reducing the chances of tissue or cell loss during processing. The cells are allowed to adhere to slide for 10 min and then fixed in 4% paraformaldehyde. The cell membranes are disrupted in 0.2% Triton X-100. Nonspecific binding of antibodies and fluorescent conjugates are blocked by preincubation in 4% BSA/1% gelatin. Colocalization of AAT and ER-specific chaperones is detected by immunofluorescence using goat polyclonal anti-AAT-FITC (Abcam) and mouse monoclonal anti-KDEL (Stressgen), with an anti-mouse tetramethylrhodamine isothiocyanate (TRITC) secondary conjugate (Abcam) for visualization of the anti-KDEL antibody. PBS washes (3) are performed after each incubation, and ensure that the addition and removal of solutions are performed carefully in the corner of the slides so as not to disturb or dislodge the monocytes. Cells are then mounted in Vectashield, containing 40 ,6diamidino-2-phenylindole di-lactate (DAPI), and examined using a LSM510 Meta laser scanning confocal microscope (Zeiss). Images are captured at 63 magnification and 4 zoom with excitation wavelengths for FITC, TRITC, and DAPI of 488, 543, and 364 nm, respectively (Fig. 5.2).

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AAT

KDEL

Merge

MM

DAPI

5 mm

5 mm

5 mm

5 mm

5 mm

5 mm

5 mm

ZZ

5 mm

Figure 5.2 Intracellular accumulation of AAT and increased KDEL expression in the ER of ZZ monocytes. The subcellular distribution of AAT in monocytes isolated from MM (normal) and ZZ (AAT-deficient) individuals was determined by immunofluorescence using antibodies for AAT (green) and the ER marker KDEL (red), with colocalization indicated by yellow staining. Nuclei were stained with DAPI (blue). Imaging was acquired using a Zeiss LSM510 Meta confocal microscope and the images presented are single focal plane scans of 1 mm depth at the midsection of the fixed cells.

ACKNOWLEDGMENTS We are thankful to Dr. Warren Thomas and Dr. David Bergin for assistance with confocal imaging and analysis, and all the members of the Department of Medicine, RCSI Beaumont Hospital, both past and present. This work was supported by financial assistance from the Alpha One Foundation (Ireland), the Alpha One Foundation (US), the Health Research Board (HRB), the Medical Research Charities Group, the Children’s Research Centre Crumlin Hospital, the Program for Research in Third Level Institutes (PRTLI), Talecris Biotherapeutics, and the Royal College of Surgeons in Ireland. Confocal imaging experiments were performed on equipment supported by the Higher Education Authority (HEA) of Ireland through the National Biophotonics and Imaging Platform Ireland (NBIPI).

REFERENCES 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. Calfon, M., Zeng, H., Urano, F., Till, J. H., Hubbard, S. R., Harding, H. P., Clark, S. G., and Ron, D. (2002). IRE1 couples endoplasmic reticulum load to secretory capacity by processing the XBP-1 mRNA. Nature 415, 92–96.

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Carroll, T. P., Greene, C. M., O’Connor, C. A., Nolan, A. M., O’Neill, S. J., and McElvaney, N. G. (2010). Evidence for unfolded protein response activation in monocytes from individuals with alpha-1 antitrypsin deficiency. J. Immunol. 184, 4538–4546. Dale, D. C., Boxer, L., and Liles, W. C. (2008). The phagocytes: Neutrophils and monocytes. Blood 112, 935–945. Harding, H. P., Zhang, Y., and Ron, D. (1999). Protein translation and folding are coupled by an endoplasmic-reticulum-resident kinase. Nature 397, 271–274. Harding, H. P., Zhang, Y., Zeng, H., Novoa, I., Lu, P. D., Calfon, M., Sadri, N., Yun, C., Popko, B., Paules, R., Stojdl, D. F., Bell, J. C., et al. (2003). An integrated stress response regulates amino acid metabolism and resistance to oxidative stress. Mol. Cell 11, 619–633. Iwakoshi, N. N., Lee, A. H., Vallabhajosyula, P., Otipoby, K. L., Rajewsky, K., and Glimcher, L. H. (2003). Plasma cell differentiation and the unfolded protein response intersect at the transcription factor XBP-1. Nat. Immunol. 4, 321–329. Kaufman, R. J. (1999). Stress signaling from the lumen of the endoplasmic reticulum: Coordination of gene transcriptional and translational controls. Genes Dev. 13, 1211–1233. Komura, T., Sakai, Y., Honda, M., Takamura, T., Matsushima, K., and Kaneko, S. (2010). CD14þ monocytes are vulnerable and functionally impaired under endoplasmic reticulum stress in patients with type 2 diabetes. Diabetes 59, 634–643. Li, Y., Schwabe, R. F., DeVries-Seimon, T., Yao, P. M., Gerbod-Giannone, M. C., Tall, A. R., Davis, R. J., Flavell, R., Brenner, D. A., and Tabas, I. (2005). Free cholesterol-loaded macrophages are an abundant source of tumor necrosis factor-alpha and interleukin-6: Model of NF-kappaB- and map kinase-dependent inflammation in advanced atherosclerosis. J. Biol. Chem. 280, 21763–21772. Livak, K. J., and Schmittgen, T. D. (2001). Analysis of relative gene expression data using real-time quantitative PCR and the 2(Delta Delta C(T)) Method. Methods 25, 402–408. Malhotra, J. D., and Kaufman, R. J. (2007). The endoplasmic reticulum and the unfolded protein response. Semin. Cell Dev. Biol. 18, 716–731. Marciniak, S. J., Yun, C. Y., Oyadomari, S., Novoa, I., Zhang, Y., Jungreis, R., Nagata, K., Harding, H. P., and Ron, D. (2004). CHOP induces death by promoting protein synthesis and oxidation in the stressed endoplasmic reticulum. Genes Dev. 18, 3066–3077. Medzhitov, R., and Janeway, C., Jr. (2000). Innate immunity. N. Engl. J. Med. 343, 338–344. Mori, K. (2000). Tripartite management of unfolded proteins in the endoplasmic reticulum. Cell 101, 451–454. Ron, D., and Walter, P. (2007). Signal integration in the endoplasmic reticulum unfolded protein response. Nat. Rev. Mol. Cell Biol. 8, 519–529. Smith, J. A., Turner, M. J., Delay, M. L., Klenk, E. I., Sowders, D. P., and Colbert, R. A. (2008). Endoplasmic reticulum stress and the unfolded protein response are linked to synergistic IFN-beta induction via X-box binding protein 1. Eur. J. Immunol. 38, 1194–1203. Tirasophon, W., Welihinda, A. A., and Kaufman, R. J. (1998). A stress response pathway from the endoplasmic reticulum to the nucleus requires a novel bifunctional protein kinase/endoribonuclease (Ire1p) in mammalian cells. Genes Dev. 12, 1812–1824. Travers, K. J., Patil, C. K., Wodicka, L., Lockhart, D. J., Weissman, J. S., and Walter, P. (2000). Functional and genomic analyses reveal an essential coordination between the unfolded protein response and ER-associated degradation. Cell 101, 249–258. Vij, N., Fang, S., and Zeitlin, P. L. (2006). Selective inhibition of endoplasmic reticulumassociated degradation rescues DeltaF508-cystic fibrosis transmembrane regulator and


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suppresses interleukin-8 levels: Therapeutic implications. J. Biol. Chem. 281, 17369–17378. Yeo, C., Saunders, N., Locca, D., Flett, A., Preston, M., Brookman, P., Davy, B., Mathur, A., and Agrawal, S. (2009). Ficoll-Paque versus Lymphoprep: A comparative study of two density gradient media for therapeutic bone marrow mononuclear cell preparations. Regen. Med. 4, 689–696. Yoshida, H., Haze, K., Yanagi, H., Yura, T., and Mori, K. (1998). Identification of the cisacting endoplasmic reticulum stress response element responsible for transcriptional induction of mammalian glucose-regulated proteins. Involvement of basic leucine zipper transcription factors. J. Biol. Chem. 273, 33741–33749. Yoshida, H., Matsui, T., Yamamoto, A., Okada, T., and Mori, K. (2001). XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor. Cell 107, 881–891. Zaman, M. M., Gelrud, A., Junaidi, O., Regan, M. M., Warny, M., Shea, J. C., Kelly, C., O’Sullivan, B. P., and Freedman, S. D. (2004). Interleukin 8 secretion from monocytes of subjects heterozygous for the deltaF508 cystic fibrosis transmembrane conductance regulator gene mutation is altered. Clin. Diagn. Lab. Immunol. 11, 819–824.


C H A P T E R

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Assessment of Endurance Exercise Training in Hepatic Endoplasmic Reticulum Stress Natalie Ann Chapados Contents 98 99 99 103 104 105 105 105 106 106

1. Introduction 2. Assessment of an Endurance-Training Program 2.1. Procedures 2.2. Inculcate good running behaviors 3. “Dietary-Inducible” Hepatic ER Stress 4. Assessment of Hepatic ER Stress Molecular Markers 4.1. Procedures: Handling of the liver 5. Required Materials Acknowledgments References

Abstract Hepatic endoplasmic reticulum (ER) stress has recently been associated with several health complications such as obesity, type 2 diabetes, and hepatic steatosis. Exercise training has been recognized for many years to have important beneficial effects in these metabolic complications, however much remains unknown regarding the effects of exercise training on ER stress in the liver. A better understanding of the effects of exercise training on hepatic ER stress response requires studies in which the exercise training protocol is well assessed. Therefore, the purpose of this chapter is to provide detailed description of an endurance-training program and suggest a dietary approach in order to help induce and study changes in ER stress molecular markers expression levels in the liver of rats in response to exercise training.

Institut de Recherche de l’Hoˆpital Montfort/Montfort Hospital, Research Institute, Ottawa, Ontario, Canada Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00006-6

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Natalie Ann Chapados

1. Introduction The endoplasmic reticulum (ER) plays an important role in the folding process of newly synthesized protein in a living organism. Alteration of this process is associated with ER stress appearance and activation of the unfolded protein response (UPR). Briefly, the UPR pathway implicates the activation of three transmembrane proteins (PERK, ATF6, and XBP1) by a cytosolic chaperone protein (BiP/GRP78) in order to establish ER homeostasis (Flamment et al., 2010). Researchers have detected the presence of ER stress in various tissues such as the neuron (Um et al., 2008), the adipose tissue (Xu et al., 2010), and the liver (Ozcan et al., 2004; Wang et al., 2006), providing new insights on ER stress implication in different health complications such as Alzheimer’s disease (Um et al., 2008), diabetes (Ozcan et al., 2004), and hepatic steatosis (Wang et al., 2006). The physiological effects of an exercise training program have been well documented over the years. In fact, exercise training is known to have beneficial metabolic effects in various health complications such as obesity, type 2 diabetes, and hepatic steatosis in humans (Eriksson et al., 2010; Kantartzis et al., 2009) and animals (Marques et al., 2010). These findings imply that the systemic effects of exercise training in the development of these health problems could possibly translate at the cellular level, that is, ER stress. Exercise training programs have been showed to have potential effects on ER stress in tissues such as the skeletal muscle and the brain. For instance, studies have reported two- to threefold increase of GRP78 protein expression in the soleus and extensor digitorum longus in rats following a 3-month treadmill program compared to sedentary rats (Gonzalez et al., 2000). Furthermore, 16 weeks of treadmill training in mice resulted in a significant increase in GRP78 protein expression in the brain of Alzheimer patients (Um et al., 2008). As for the liver, the only evidence is demonstrated by the study of the acute effects of exercise. Gonzalez et al. (Gonzalez and Manso, 2004) reported that rats exposed to a single exercise bout increased GRP78 protein expression by twofold compared to sedentary rats. When investigating the impact of exercise in animal models, just as in humans, the importance of using a well-designed training program must not be overlooked. Only a carefully crafted training protocol is likely to provide relevant and interpretable data. While a plethora of information is available for establishing standard human training regimen, the design of a training program for rodents could be seen as rather challenging. Therefore, the purpose of this chapter is to provide a detailed assessment of an endurance exercise protocol to study molecular changes in hepatic ER stress in rats. Hopefully, this technique will help minimize the difficulties regarding the

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Endurance Training on Liver ER Stress in Rats

assessment of an adequate training program and induce cellular changes in hepatic ER stress molecular makers in rats.

2. Assessment of an Endurance-Training Program Adopting a fit exercise program to induce changes.

2.1. Procedures The next section consists of three types of exercise training protocol on treadmill: (1) increasing speed, constant slope; (2) increasing slope, constant speed; and (3) increasing speed and slope. 2.1.1. Increasing speed, constant slope The exercise training protocol presented in this section is based on established values of VO2max levels conducted in rats. Shepherd et al. (Shepherd and Gollnick, 1976) assessed VO2max in Sprague–Dawley rats using a motor-driven work wheel equipped with a metabolic chamber. Oxygen uptake measurements were performed during exercise at seven different speeds (16–67 m min 1) when the slope was null (0 of inclination). Values ranged from 6.44 ml (100 g min) 1 when rats ran at 16 m min 1 and increased 30% at 67 m min 1. As shown in Table 6.1, VO2 ml (100 g min) 1 increased linearly with running speed intensity (m min 1) up to a maximal value of 9.51 0.14 ml (100 g min) 1 at 49.5 m min 1. Thereafter, the average VO2 ml (100 g min 1) decreased slightly to 9.39 0.15 ml (100 g min) 1 at a running speed of 67.0 m min 1. In addition, I have included in Table 6.1 Table 6.1 Oxygen uptake in rats during exercise at different speeds (m min 1) Speed (m min 1) VO2 ml (100 g min 1) VO2 (ml kg 1 min 1) % VO2max

0 16.0 22.5 28.5 39.0 49.5 53.5

2.42 6.44 7.26 7.75 8.85 9.51 9.49

0.1 0.16 0.15 0.14 0.14 0.14 0.14

24.2 64.4 72.6 77.5 88.5 95.1 94.9

1.0 1.6 1.5 1.4 1.4 1.4 1.4

Baseline 67.7 76.3 81.5 93.0 100 99.8

Adapted from Shepherd et al. (Shepherd and Gollnick, 1976). Values are expressed by means S.E. Pooled data from three groups of rats, 16 rats per group.

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120

VO2 (ml kg−1 min−1)

100 80 60 40 20 0

0

10

20

30

40

50

60

m min-1

Figure 6.1 Linear relationship between increasing speed (16–49.5 m min 1) and VO2 (ml kg 1 min 1) in Sprague–Dawley rats. Adapted from Shepherd et al. (Shepherd and Gollnick, 1976). Values are expressed by means S.E. of pooled data from three groups of rats, 16 rats per group.

converted values of VO2 expressed in ml kg 1 min 1 derived from initial values expressed in ml (100 g min) 1 obtained from Shepherd et al. (Shepherd and Gollnick, 1976) as well as % VO2max. Thereafter, VO2 (ml kg 1 min 1) and speed (m min 1) values were plotted and presented in Fig 6.1 in order to obtain Eq. (6.1) derived from the linear relationship between increasing speed (16–49.5) and VO2. y ¼ 0:9169x þ 51:106ðr ¼ 0:98Þ

ð6:1Þ

Equation (6.1) will therefore help to predict VO2 at a given speed (between 16 and 49.5 m min 1) when the slope is constant at 0 . 2.1.2. Increasing slope, constant speed As mentioned earlier, values provided from Shepherd et al. (Shepherd and Gollnick, 1976) presented in Table 6.1 and the Eq. (6.1) derived from the linear relationship between VO2 (ml kg 1 min 1) and speed (m min 1) (Fig. 6.1) were obtained when the slope is constant at 0 . Therefore, it has been reported that an increase of one slope incline increases 0.44 mlO2 kg 1 min 1 in oxygen consumption, representing about 0.5% of VO2max for each slope incline (Norton et al., 1990). Figure 6.2 represents an example of the relationship between the increasing slope ( ) and the VO2 (mlO2 kg 1 min 1) when the speed is constant (e.g., here the

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74.5 74

VO2 (ml kg -1 min-1)

73.5 73 72.5 72 71.5 71 70.5 70 69.5 0

2

4

6

8

10

Incline (⬚)

Figure 6.2 Relationship between speed (m min 1) and VO2 (ml kg 1 min 1) in Sprague–Dawley rats when speed is constant at 20 m min 1. An increase of each inclination of 1 increases 0.5% of VO2max. Here, a speed of 20 m min 1 corresponds to an oxygen uptake of 69.44 ml kg 1 min 1 at 0 incline.

speed is constant at 20 m min 1). Equation (6.2) derived from the relationship presented in Fig. 6.2 is y ¼ 0:4403x þ VO2 mlO2 kg1 min1 ð6:2Þ at 0 slope value (e.g., here VO2 corresponds to 69.44 mlO2 kg 1 min 1 at speed 20 m min 1). The exercise training protocol that I proposed in the following section is an enforced training protocol on a treadmill that is generally used in our laboratory. This type of program ensures a precise control of the training stimulus and therefore could serve as a rigorous model to study the physiological effects of exercise training in rats. 2.1.3. Increasing speed and slope (proposed endurance-training program) The endurance-training program (Table 6.2) consists of a 6-week endurance running program in Sprague–Dawley rats starting when rats are 6 weeks of age. Rats were trained to run 5 days week 1 for 6 weeks on a motor-driven treadmill with 10 individual lanes (Quinton Instruments, Seattle, WA, USA). We kept rats in a 12 h light–dark cycle (6 a.m.–6 p.m. light; 6 p.m.–6 a.m. dark). Since rodents are not normally active during the

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Table 6.2 Proposed endurance-training program for rats

Period

Time (min)

Acclimation Day 1 15 Day 2 15 Day 3 20 Day 4 20 Day 5 25 2-day rest Training program Week 1 30 30 35 35 35 Week 2 40 40 45 45 45 Week 3 45 50 50 55 55 Week 4– 60 5–6

Speed (m min 1)

Slope ( )

VO2 (ml kg 1 min 1)

% VO2max

15 20 20 20 20

0 0 0 2 2

64.9 69.4 69.4 70.3 70.3

68.2 73 73 74 74

21 21 21 23 23 23 23 23 23 23 26 26 26 26 26 26

2 4 4 6 6 8 8 8 10 10 10 10 10 10 10 10

71.2 72.1 72.1 74.8 74.8 75.7 75.7 75.7 76.6 76.6 79.3 79.3 79.3 79.3 79.3 79.3

75 76 76 78.7 78.7 79.6 79.6 79.6 80.5 80.5 83.4 83.4 83.4 83.4 83.4 83.4

light period, the optimal training period should be during the dark cycle. Therefore, by personal experience and convenience, we usually trained our rats in the early morning beginning the light cycle (7 a.m.). As shown in Table 6.2, exercise training time (min day 1), speed (m min 1), and slope is initially set at 30 min day 1, 21 m min 1, and 2% slope following the 5-day acclimation period. During the first 3 weeks, speed, duration, and slope of the exercise progressively increased until the rats could run from 30 min day 1 at 21 m min 1 and 2% slope to 60 min day 1 at 26 m min 1 and 10% slope by the beginning of week 4. Thereafter, speed, duration, and slope of the exercise were maintained for the last 3 weeks. Progression of VO2max throughout the endurance-training protocol, including the acclimation and the rest periods, is summarized in Fig. 6.3. In this protocol, the target VO2max is reached by week 4;

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90

Acclimation

Rest

5

2

Training

% VO2 max

80

70

60

1

Days

2 Weeks

3

4+

Figure 6.3 Summarized progression of VO2max throughout the endurance-training program in Sprague–Dawley rats including the acclimation and rest periods.

therefore, it should be noted that this endurance-training protocol can be prolonged for longer training periods (e.g., 8, 16, or 24 weeks). This exercise training program is considered as moderate (83.4% VO2max) for rats as more intense exercise training programs are currently used in the literature (Laughlin and Armstrong, 1982; Luginbuhl et al., 1984). As for the duration and the intensity, our previous laboratory experience indicates that sedentary rats can run continuously for an hour between 70% and 80% VO2max immediately after the acclimation period (Chapados and Lavoie, 2010; Chapados et al., 2008, 2009). Accordingly, the proposed endurance-training program is adequate to induce physiological changes without leading to exhaustion.

2.2. Inculcate good running behaviors Although most all rats will run on the treadmill, some problems with this type of exercise protocol should be noted. First, it is necessary to expose rats to a habituation period of five exercise bouts in order to familiarize the animals to the exercise training protocol and the motor-driven treadmill. This adaptation period will also allow exercise responders to be distinguished from nonresponders. The nonresponders correspond to rats that refuse to run or demonstrate sporadic (stop and go) running behavior. By this, following the 5-day acclimation to exercise and 2-day rest periods, experimental groups will be established by performance and motivation in response to exercise, that is, runners or nonrunners (control or sedentary).

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Second, negative reinforcement performed during the acclimation is usually needed in order to encourage the rats to run on the treadmill. Without this kind of reinforcement, most animals avoid running. Air jets and mild electrical shocks are two types of negative reinforcement commonly employed in treadmill exercise training protocols in order to implement running behaviors and do not induce any harm to animals. Air jets combined with light brushing underneath the paws when animals are at the end of the corridor can be repeatedly used until the adequate running behaviors are obtained. However, if this technique proves to be ineffective in motivating the rats to run, mild electrical stimulation combined with tapping twice with a stick on the plexiglass can be used thereafter. Basically, this behavioral reinforcement method includes tapping twice on the plexiglass immediately followed by an electrical discharge when the rats stop running and find themselves on the electrical grid at the end of the corridor. The electrical intensity should not exceed 0.5 W (300 V and 1.6 mA). This electrical intensity is not harmful and proves to be sufficient to displace the rats that stopped running and are at the end of the corridor on an electrical grid installed on the motor-driven treadmill. Our experience in the laboratory has shown that rats quickly avoid electrical grid contact and respond to the tapping stimulus alone within five exercise bouts resulting rapidly in the avoidance of the use of the electrical stimulation. Last, it is generally assumed that the residual effects of an exercise bout may remain for some period posttraining (Armstrong et al., 1983; Kesaniemi et al., 2001). In order to discriminate the effects of the acute exercise bout from training (chronic), all experiments should be performed 36–48 h after the last exercise bout. This enforced training protocol on treadmills has been widely used as a model for studying the physiological effects of exercise training in rats. This type of training program permits a precise control of the training stimulus.

3. “Dietary-Inducible” Hepatic ER Stress Here, I present a dietary approach that was used in our laboratory to induce hepatic ER stress in order to study the effects of training in animals. Recently, the relationship between ER stress and lipid accumulation in the liver has been studied in models of hepatic lesion in animals. For example, dietary-induced nonalcoholic hepatic steatosis has been recently associated with the presence of ER stress in animals (Ozcan et al., 2004; Wang et al., 2006). Hepatic BiP/GRP78 mRNA levels are upregulated in mice fed a high-fat diet for 16 weeks (Ozcan et al., 2004). Furthermore, high-saturated fat seems to induce more important hepatic lesions. Wang et al. (2006) showed even though liver triacylglycerol (TAG) levels were equal, a twofold


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increase in GRP78 protein content and XBP1 gene expression were noted in the liver of rats fed a high-saturated-fat diet compared to rats fed highpolyunsaturated-fat diet. In the same line of idea, our group of researchers has demonstrated that MTP inhibition in high-saturated-fat-fed rats for 6 weeks leads to a dramatic increase in liver TAG content and results in higher gene expression of ER stress molecular markers (i.e., BiP/GRP78, PERK, ATF6, and XBP-1) (Chapados and Lavoie, 2010). Our laboratory highsaturated-fat diet consisted of 43% lipid (80% lard, 20% corn oil), 38% carbohydrate, and 19% protein (kcal) and was provided from Harlan, Teklad (WI, USA) (Chapados and Lavoie, 2010; Chapados et al., 2008, 2009) and given concomitantly with the training program.

4. Assessment of Hepatic ER Stress Molecular Markers 4.1. Procedures: Handling of the liver On the day of the sacrifice, immediately following the liver excision one piece of the liver is extracted (our laboratory always used the median lobe as we find it more accessible). The sample is immediately snap frozen using precooled clamps (5 min) to the temperature of liquid nitrogen prior to use. The clamps containing the liver sample are placed in liquid nitrogen for 2 min. Then, the liver sample is wrapped in aluminum foil and put in liquid nitrogen (if the experiment is not completed) or stored at 78 C until further analyses. As for liver RNA extraction and measurements, follow the manufacturer’s manual.

5. Required Materials Training protocol

Smooth brush (nylon) Air jets system Motor-driven treadmill adapted for animals and equipped with an electrical grid (see Chapados and Lavoie, 2010; Chapados et al., 2008, 2009). Stick Handling of the liver

Clamps (precooled in liquid nitrogen)

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ACKNOWLEDGMENTS The research relevant to this chapter was supported by Natural Sciences and Engineering Research Council of Canada (NSERC).

REFERENCES Armstrong, R. B., Ogilvie, R. W., and Schwane, J. A. (1983). Eccentric exercise-induced injury to rat skeletal muscle. J. Appl. Physiol. 54(1), 80–93. Chapados, N. A., and Lavoie, J. M. (2010). Exercise training increases hepatic endoplasmic reticulum (er) stress protein expression in MTP-inhibited high-fat fed rats. Cell Biochem. Funct. 28(3), 202–210. Chapados, N., Collin, P., Imbeault, P., Corriveau, P., and Lavoie, J. M. (2008). Exercise training decreases in vitro stimulated lipolysis in a visceral (mesenteric) but not in the retroperitoneal fat depot of high-fat-fed rats. Br. J. Nutr. 100(3), 518–525. Chapados, N. A., Seelaender, M., Levy, E., and Lavoie, J. M. (2009). Effects of exercise training on hepatic microsomal triglyceride transfer protein content in rats. Horm. Metab. Res. 41(4), 287–293. Eriksson, M., Uddeń, J., Hemmingsson, E., and Agewall, S. (2010). Impact of physical activity and body composition on heart function and morphology in middle-aged, abdominally obese women. Clin. Physiol. Funct. Imaging 30(5), 354–359. Flamment, M., Kammoun, H. L., Hainault, I., Ferre, P., and Foufelle, F. (2010). Endoplasmic reticulum stress: A new actor in the development of hepatic steatosis. Curr. Opin. Lipidol. 21(3), 239–246. Gonzalez, B., and Manso, R. (2004). Induction, modification and accumulation of HSP70s in the rat liver after acute exercise: Early and late responses. J. Physiol. 556(Pt. 2), 369–385. Gonzalez, B., Hernando, R., and Manso, R. (2000). Stress proteins of 70 kDa in chronically exercised skeletal muscle. Pflugers Arch. 440(1), 42–49. Kantartzis, K., Thamer, C., Peter, A., Machann, J., Schick, F., Schraml, C., et al. (2009). High cardiorespiratory fitness is an independent predictor of the reduction in liver fat during a lifestyle intervention in non-alcoholic fatty liver disease. Gut 58(9), 1281–1288. Kesaniemi, Y. K., Danforth, E., Jr., Jensen, M. D., Kopelman, P. G., Lefebvre, P., and Reeder, B. A. (2001). Dose-response issues concerning physical activity and health: An evidence-based symposium. Med. Sci. Sports Exerc. 33(6 Suppl.), S351–S358. Laughlin, M. H., and Armstrong, R. B. (1982). Muscular blood flow distribution patterns as a function of running speed in rats. Am. J. Physiol. 243(2), H296–H306. Luginbuhl, A. J., Dudley, G. A., and Staron, R. S. (1984). Fiber type changes in rat skeletal muscle after intense interval training. Histochemistry 81(1), 55–58. Marques, C. M., Motta, V. F., Torres, T. S., Aguila, M. B., and Mandarim-deLacerda, C. A. (2010). Beneficial effects of exercise training (treadmill) on insulin resistance and nonalcoholic fatty liver disease in high-fat fed C57BL/6 mice. Braz. J. Med. Biol. Res. 43(5), 467–475. Norton, K. I., Delp, M. D., Jones, M. T., Duan, C., Dengel, D. R., and Armstrong, R. B. (1990). Distribution of blood flow during exercise after blood volume expansion in swine. J. Appl. Physiol. 69(5), 1578–1586. Ozcan, U., Cao, Q., Yilmaz, E., Lee, A. H., Iwakoshi, N. N., Ozdelen, E., et al. (2004). Endoplasmic reticulum stress links obesity, insulin action, and type 2 diabetes. Science 306 (5695), 457–461.


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Shepherd, R. E., and Gollnick, P. D. (1976). Oxygen uptake of rats at different work intensities. Pflugers Arch. 362(3), 219–222. Um, H. S., Kang, E. B., Leem, Y. H., Cho, I. H., Yang, C. H., Chae, K. R., et al. (2008). Exercise training acts as a therapeutic strategy for reduction of the pathogenic phenotypes for Alzheimer’s disease in an NSE/APPsw-transgenic model. Int. J. Mol. Med. 22(4), 529–539. Wang, D., Wei, Y., and Pagliassotti, M. J. (2006). Saturated fatty acids promote endoplasmic reticulum stress and liver injury in rats with hepatic steatosis. Endocrinology 147(2), 943–951. Xu, L., Spinas, G. A., and Niessen, M. (2010). ER stress in adipocytes inhibits insulin signaling, represses lipolysis, and alters the secretion of adipokines without inhibiting glucose transport. Horm. Metab. Res. 42(9), 643–651.


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Discovery Approaches to UPR in Athero-Susceptible Endothelium In Vivo Mete Civelek,*,†,k Elisabetta Manduchi,‡,§ Gregory R. Grant,‡,§ Christian J. Stoeckert Jr.‡,§ and Peter F. Davies*,†,} Contents 1. Introduction 1.1. Site-specific adaptive ER stress and UPR phenotype in athero-susceptible endothelium in vivo 1.2. Genomics approach to arterial endothelial phenotype 2. Procedures for the Isolation of Endothelial Cells and Preparation of RNA for Microarray Hybridization 2.1. Arterial tissue preparation 2.2. Endothelial cell harvest 2.3. Assessing endothelial cell purity 2.4. Endothelial RNA extraction and quality control 2.5. Integrity of isolated endothelial RNA 2.6. Messenger RNA amplification and evaluation 2.7. Amplified RNA fluorescent dye conjugation 3. Microarray Hybridization and Feature Extraction 4. Bioinformatics Analysis 4.1. Annotation of the porcine microarray 4.2. Microarray data preprocessing 4.3. Analysis of differential gene expression 4.4. Identification of enriched biological themes 4.5. Gene set enrichment analysis 4.6. Ingenuity pathway analysis

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* Institute for Medicine and Engineering, University of Pennsylvania, Philadelphia, Pennsylvania, USA { Department of Bioengineering, University of Pennsylvania, Philadelphia, Pennsylvania, USA { Center for Bioinformatics, University of Pennsylvania, Philadelphia, Pennsylvania, USA } Department of Genetics, University of Pennsylvania, Philadelphia, Pennsylvania, USA } Department of Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, Pennsylvania, USA k Current address: Division of Cardiology, Department of Medicine, University of California, Los Angeles, California, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00007-8

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5. Another Approach to Gene Connectivity: Weighted Gene Coexpression Network Analysis 5.1. Network visualization 6. Validation and Follow-up Acknowledgments References

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Abstract The endothelium is a monolayer of cells that lines the entire inner surface of the cardiovascular and lymphatic circulations where it controls normal physiological functions through both systemic and local regulation. Endothelial phenotypes are heterogeneous, dynamic and malleable, properties that in large- and medium-sized arteries lead to a central role in the development of focal and regional atherosclerosis. The endothelial phenotype in athero-susceptible sites is different from that in nearby athero-resistant regions. Understanding the in vivo gene, protein, and metabolic expression profiles of susceptible endothelium is, therefore, an important spatiotemporal challenge in atherosclerosis research. Recent studies have demonstrated that endoplasmic reticulum (ER) stress and the UPR are characteristics of susceptible endothelium. Here, we outline global genomic profiling, pathway analyses, and gene connectivity approaches to the identification of UPR and associated pathways as discrete markers of athero-susceptibility in arterial endothelium.

1. Introduction Atherosclerosis is not a diffuse disease; it has been noted for over a century that lesion development is associated with arterial curvatures, asymmetries, and branches where the nonuniform arterial geometry generates patterns of blood flow that are considerably more complex than elsewhere. Since it is well established that endothelial cells are highly sensitive to flow/ shear stress, a hemodynamics contribution to localized susceptibility is likely. Athero-susceptible endothelium in vivo expresses a different repertoire of cell phenotypes than that in nearby protected locations (Davies, 2009). Identification of important differences in gene and protein expression and the mechanisms responsible requires both global profiling and classic cell and molecular approaches. Recently, systems biology and discovery science methodologies identified the unfolded protein response (UPR) as a prominent differential component of endothelial phenotype in regions in vivo that are susceptible to atherosclerosis. Here, we first summarize the findings that led to UPR identification. This is followed by a description of the transcriptomics approaches and bioinformatics analyses employed. Mention is then made of the conventional cell and molecular biochemistry used to validate and extend the results of the genomics predictions.

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1.1. Site-specific adaptive ER stress and UPR phenotype in athero-susceptible endothelium in vivo ER stress is an adaptive protective mechanism that arises because of excessive protein biosynthesis or interference with normal protein-folding mechanisms in the ER lumen in response to multiple kinds of cellular stress. In athero-susceptible regions of the arterial circulation, such stresses are likely generated by locally complex hemodynamics that create adverse biomechanical forces and, through the formation of flow separation zones, promote the retention of pro-pathological biochemicals such as free radicals. These and other stresses result in excessive newly synthesized and/or misfolded polypeptides in the ER lumen that exceed its protein-folding capacity. The resulting activation of UPR is an ubiquitous adaptive cell response that activates a set of compensatory intracellular signaling pathways. The UPR elicits a coordinated transcriptional upregulation of ER chaperones and folding enzymes to promote the correct assembly of unfolded polypeptides and prevent incompletely folded proteins from aggregating, thereby assisting cell survival in an adverse environment. In the unstressed state, the ER chaperone Binding Protein (BiP; also known as heat shock protein A5, HSPA5, and glucose-related protein 78, GRP78) binds to each of three ER stress transducers. These are ER transmembrane proteins each having an ER-luminal domain for the sensing of unfolded proteins and a cytosolic domain for signaling. Bound BiP maintains the inactive state of the transducers. In ER stress, BiP dissociates from the chaperones to bind unfolded/misfolded polypeptides in the ER lumen, causing chaperone phosphorylation. Activation of ATF6a (activating transcription factor 6a), IRE1a (inositol requiring kinase 1a), and PERK (protein kinase-like ER kinase) together with the downstream consequences of their activation constitute the UPR. The products of the activated UPR transducers converge as transcriptional regulators in the nucleus to upregulate ER chaperones and UPR transducer synthesis and to ubiquitinate unfolded proteins for degradation through the proteosome; both processes relieve ER stress accumulation of unfolded proteins and restore ER protein equilibrium to a normal range. Failure leads to apoptosis through transcriptional induction of the transcription factor CHOP (C/ ERB homologous protein), inflammation through activation of NFkB, and generation of reactive oxygen species (ROS) through excessive protein oxidation in the ER (Malhotra and Kaufman, 2007).

1.2. Genomics approach to arterial endothelial phenotype In a multisite study (Civelek et al., 2009) on 45 normal adult swine, endothelium in susceptible regions of the aortic arch (AA), proximal brachiocephalic artery, aorto-renal branch region, and abdominal aorta were

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analyzed relative to protected sites of the common carotid artery, descending thoracic aorta (DT), and the distal renal artery. All athero-susceptible regions are associated with complex disturbed blood flow. From this multisite study, the most abundant common feature of the endothelium of all athero-susceptible regions was the upregulation of genes associated with ER processing of proteins, ER stress, and the UPR. Differential gene expression analysis identified a highly connected and coordinated network of genes upregulated in the susceptible regions. Three independent pathway mining approaches—Gene Ontology (GO) terms overrepresentation (using DAVID), gene set enrichment analysis (GSEA), and Ingenuity Pathway Analysis (IPA)—identified ER stress and the UPR to be overrepresented functional categories in athero-susceptible endothelium including genes that function in protein folding, synthesis, and posttranslational protein modification. To validate the transcriptome analyses, endothelial cell proteins were isolated from AA and DT and also from the athero-susceptible aorto-renal branch and the protected distal renal artery. At each athero-susceptible disturbed flow site, BiP transcript and/or protein expression was significantly upregulated. Western blot demonstrated significantly elevated phospho ATF6a, phospho IRE1a and its target, spliced XBP-1. However, the third transducer pathway PERK was not activated. Overall, this study, approached without preconceived expectations of differential expression of genes and proteins associated with ER stress/UPR, suggests that stresses associated with flow disturbance in vivo elicit activation of the UPR, an ER response common to other forms of stress, and that chronic UPR is a signature for athero-susceptible endothelial phenotype in vivo. A schematic of the overall experimental approach is outlined in Fig. 7.1. The reader is also referred to Civelek et al. (2009) for specific UPR output data associated with the procedures outlined in this chapter in addition to the public domain bioinformatics data sets referred to in the text.

2. Procedures for the Isolation of Endothelial Cells and Preparation of RNA for Microarray Hybridization In this section, the endothelial isolation procedure from swine arterial tissue is outlined. Contamination by smooth muscle cells and leukocytes should be avoided. In order to preserve the integrity of RNA for downstream applications, RNase-free conditions must be strictly followed. We recommend preparing buffers in DEPC-treated water, cleaning the surgical instruments with RNAaseZap (Ambion, Inc.), and general care for avoiding contamination by changing gloves frequently.

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B

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mRNA Nucleic acid amplification

Sites of athero-susceptibility; flow disturbance Sites resistant to atherosclerosis; flow unidirectional Validation (qRTPCR; Western blot; IHC)

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Figure 7.1 Schematic outline of discovery approach to site-specific endothelial phenotype in swine arteries. Regions of athero-susceptibility and resistance are indicated in the coronary arteries, aorta, and renal arteries. (A) Heart and great vessels, from left to right: distal right coronary (RC) artery, proximal RC, aortic arch, proximal left coronary artery, proximal left anterior descending (LAD) coronary artery, distal LAD, distal circumflex artery. (B and C) (opened) Aortic arch and descending thoracic aorta. (D and E) (opened) Proximal and distal regions of renal arteries. Transcriptome analyses of endothelium isolated from discrete regions identified protein biosynthesis, ER stress, and UPR as dominant pathway phenotypes differentially expressed in regions of athero-susceptibility. Transcript validation by quantitative real-time PCR (qRTPCR) is followed by isolation of endothelial proteins for Western blotting, to determine regional protein and phospho-protein expression differences (indicative of UPR activation), and immunohistochemistry (IHC) of proteins in situ.

2.1. Arterial tissue preparation Ascending, descending, and abdominal aortas with their branches and carotid artery are harvested within 30–45 min after animal death. Immediately following excision from the animal, they should be cooled on ice to inhibit metabolism. The vessel lumen is rinsed with ice-cold RNAase-free PBS. Surrounding tissue is dissected away and the vessels are cut open longitudinally with blunt artery scissors (Fine Scientific Tools) to minimize damage to endothelial cells then pinned onto waxed trays and rinsed once again with cold RNase-free PBS.

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2.2. Endothelial cell harvest Endothelial cells are gently scraped (fine scalpel blade angled in the direction of the “stroke”) from discrete arterial regions identified to correspond to sites susceptible to atherosclerosis (or any other variable of interest to the investigator). For our work, this information is known from the geometry of the arteries, various in vivo imaging modalities of blood flow (where flow disturbances predict sites of athero-susceptibility), and by direct observations of the nuclear shape of the endothelial cells. The last is conducted by immersing the artery for 1 min in a solution of 10 mg/ml DAPI (40 ,6diamidino-2-phenylindole) dye (Sigma) in PBS, a fluorescent stain that binds strongly to DNA; flow disturbance is indicated by the absence of alignment between endothelial cells in contrast to most arterial locations where the nuclear alignment reflects directional undisturbed laminar flow (Passerini et al., 2004). For most of our studies, sample size ranges between 0.1 and 1 cm2 (several hundred to 10,000 cells). Since small regions yield insufficient endothelial RNA for direct labeling and hybridization to microarrays, mRNA amplification is necessary (below). We have retained good fidelity of linear amplification from as little as 1 ng total RNA which corresponds to about 100 cells. To avoid sample collection and processing bias as well as other confounding factors, it is advisable if possible to harvest randomly from comparative groups of animals. Although this approach introduces increased variance in gene expression, the emerging results are likely to represent that of a general population. In our studies, no more than three samples came from the same animal; in most cases, each sample was from a single animal (45 animals in total). Scrapes from various regions from multiple animals were pooled to obtain a reference sample for microarray hybridizations.

2.3. Assessing endothelial cell purity Periodically throughout the harvest procedure, samples of scraped endothelial cells are spread on glass microscope slides and fixed in ice-cold acetone. In order to assess the purity, cells are double stained using an anti-porcine CD31 and von Willebrand factor antibodies for the detection of endothelium. This double staining is important since certain leukocyte subtypes also express CD31. Anti-porcine alpha-actin antibody for the detection of smooth muscle cells and anti-porcine CD45 antibody for the detection of leukocytes are used. DAPI staining indicates the total number of isolated cells in each optical field on the slide. Counting of the cells with different staining patterns indicates the presence/absence of contaminating tissue and blood cells. The gentle scraping isolation technique yields on average 96.5% endothelium (Fig. 7.2).

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a -SMC actin

CD31 and vWF

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Figure 7.2 Purity of isolated endothelial samples. Scraped samples were fixed onto microscope slides. Purity and contamination was assessed by antibody staining. (Left panel) CD31 (green) and von Willebrand Factor (vWF) (red) double staining for endothelial cells. (Middle panel) a-smooth muscle actin (green) staining for smooth muscle. (Right panel) CD45 (green) staining for leukocytes. Average endothelial purity was 96.5% with 2.78% smooth muscle cell and 0.72% leukocyte contamination. Nuclei were observed with blue Hoechst 33258 staining. Red arrows show a smooth muscle cell and a leukocyte. Bar ¼ 20 mm.

2.4. Endothelial RNA extraction and quality control Freshly isolated cells are transferred directly to a lysis buffer containing the RNase inhibitors guanidine isothiocyanate and b-mercaptoethanol (0.143 M; Absolutely RNA Nanoprep Kit, Stratagene, La Jolla, CA) and stored on dry ice. Total RNA is isolated using the Absolutely RNA Nanoprep or Microprep kit depending on the size of the samples (cell numbers) according to manufacturer’s instructions. Briefly, an equal volume of 70% RNAase-free ethanol is added to thawed cell lysates. They are loaded onto a silica-based fiber matrix, which binds RNA during centrifugation. Contaminating DNA is digested by a 15-min DNase treatment at 37 C. Proteins and DNA are removed by high- and low-salt buffer washes. Total RNA is purified by two successive elutions of 10 ml for Nanoprep or 50 ml for Microprep kits in 65 C elution buffer and RNAase-free water. Our experience suggests that warming the elution buffer to 65 C greatly enhances the amount of recovered RNA from cells.

2.5. Integrity of isolated endothelial RNA Total RNA integrity is evaluated using an Agilent Bioanalyzer 2100 and RNA 6000 Nano Labchips (Agilent Technologies, Palo Alto, CA) according to manufacturer’s instructions. Total RNA is judged to be intact if two ribosomal bands (28S and 18S) are present in approximately a 2:1 ratio and if the RNA integrity number is above 9.5. RNA quantity is measured using a Nanodrop ND-1000 spectrophotometer (Nanodrop Technologies, Inc., Rockland, DE). Total RNA with 260/280 and 260/230 wavelength (nm) ratios, which denote the purity of isolated nucleic acid, between 1.8 and 2.1 and concentrations higher than 10 ng/ml is used in subsequent procedures.

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2.6. Messenger RNA amplification and evaluation It is not possible to obtain enough RNA from small arterial regions used in this study. Therefore, linear RNA amplification is used to increase the RNA amount for subsequent microarray hybridizations while preserving the ratio of transcripts intact. Total RNA (range 1–100 ng) is amplified using the MessageAmp aRNA Kit (Ambion, Austin, TX). This procedure is based upon the antisense RNA (aRNA) linear amplification procedure described by Van Gelder et al. (1990). Poly(A) RNA is reverse-transcribed with an oligo (dT) primer containing a T7 RNA polymerase promoter sequence. RNase H treatment cleaves the mRNA into small fragments that serve as primers during second-strand synthesis, resulting in a double-stranded cDNA template for T7-mediated linear amplification by in vitro transcription. Aminoallyl UTP nucleotides are incorporated for subsequent dye conjugation steps. Typically 2–5 mg aRNA is produced from one round of amplification of 100 ng RNA; a second round of amplification for small samples yields substantial amounts (> 20 mg) of aRNA with little loss of fidelity (Polacek et al., 2003). aRNA is quantified using the Nanodrop ND-1000 and evaluated for size distribution using Agilent RNA 6000 Nano Labchips. In addition to the sample preps, pooled reference RNA is amplified once, collected and frozen in 10 mg aliquots for reference use.

2.7. Amplified RNA fluorescent dye conjugation Amplified RNA is dried using a vacuum dryer at low heat setting. Higher temperatures degrade RNA and, therefore, should be avoided. RNA is reconstituted in 9 ml coupling buffer (from the MessageAmp kit). Monofunctional NHS ester Cy3 or Cy5 dye, reconstituted in 11 ml DMSO, is added (Amersham CyTMDye Post-labeling Reactive Pack, GE Healthcare, UK). Reference aRNA and sample aRNA are labeled with Cy3 and Cy5 fluorescent dyes, respectively. Samples are incubated in the dark for 30 min at room temperature. Hydroxylamine (4.5 ml; 4 molar) is added for 15 min to quench the dye coupling reaction. Dye-coupled RNA is purified into nuclease-free water using the aRNA filter cartridges (MessageAmp kit) in order to remove excess dye. Final volume of dye-conjugated RNA is typically 150 ml.

3. Microarray Hybridization and Feature Extraction A reference design is used in which samples are labeled with Cy5 and the amplified RNA from pig common reference RNA is labeled with Cy3. Pig common reference RNA consists of aRNA amplified from endothelial


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total RNA which is pooled from all arterial sites in the study. Samples are vacuum dried to 27 ml at a low heat setting. In order to facilitate hybridization efficiency, samples are fragmented using fragmentation reagents (Ambion, Cat# 8740, Austin, TX). Three microliter of 10 fragmentation reagent is added and the samples are incubated at 70 C for 15 min. The fragmentation reaction is inhibited by the addition of 3 ml stop reagent. Each Cy5-conjugated sample is combined with an equal volume of Cy3-conjugated reference RNA. Nuclease-free water is added to each sample to a total volume of 70 ml followed by the addition of 1 ml of 10 mg/ml herring sperm DNA. Seventy-one microliter of 2 hybridization buffer (Proteomics Research Services, PRS16003050) is added to the samples which are incubated at 95 C for 5 min followed by centrifugation at 10,000g for 1 min. Samples are loaded onto custom-printed or commercial swine microarrays. Microarrays are hybridized in a Genomics Solution HybStation (Ann Arbor, MI) using a step-down protocol (42, 35, 30 C each for 5 h) which ensures uniform hybridization across all probes. Microarrays are later washed with medium stringency buffer (PRS-16004001, Proteomics Research Services, Ann Arbor, MI) at 30 C for 2 min, followed by a high stringency buffer wash (PRS-16004501, Proteomics Research Services, Ann Arbor, MI) at 25 C for 2 min. Finally, microarrays are washed with postwash buffer (PRS-16003501, Proteomics Research Services, Ann Arbor, MI) at 25 C for 2 min. They are dipped into deionized water for 30 s and dried by centrifugation at 500g for 1 min. In our study, a total of 98 samples were hybridized. Sample hybridizations were performed in batches of 10–12 in random order over several days to avoid experimental bias (day-today variations in hybridization conditions occur remarkably arbitrarily despite best practices). Arrays are scanned with an Agilent DNA Microarray Scanner at 5 mm resolution (single pass) with 100% laser power and 100% photo multiplier tube sensitivity. Images are analyzed with Agilent Feature Extraction Software (version 9.1) with raw fluorescence intensity values determined using the “CookieCutter” method of spot analysis. Each .TIF image file is examined for the quality of hybridization. If artifacts (e.g., substantial uneven hybridization) are present, those microarrays are discarded. GAL file grid, which contains probe annotations, is fitted by hand for each microarray image to ensure correct alignment for each spot.

4. Bioinformatics Analysis 4.1. Annotation of the porcine microarray Since the pig genome is fully sequenced, commercial swine microarrays are now readily available and often cost effective; manufacturer’s procedures should be followed. We custom-print porcine oligonucleotide microarrays

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(University of Pennsylvania Microarray Core Facility) using Qiagen’s Pig Array-Ready Oligo Set on 16.94 52.94 mm Codelink slides. This set includes 70-mer probes for 10,665 genes from The Institute for Genomic Research (TIGR) porcine database. The average melting temperature for the primers is 78 C. They are designed to have minimal hairpin structure and cross-hybridization and are 30 biased (within 1000 base pairs of the 30 end) to ensure adequate signal tolerance for some RNA degradation. Several control Cy3 spots, Stratagene alien controls, and 133 custom 70mer probes including genes known to play key roles in endothelial function and in atherosclerosis are also printed onto the microarrays. Oligos are suspended in 50 mM sodium phosphate buffer at a final concentration of 8.33 mM for printing. The resulting array has 12,288 spots arranged in 32 subgrids, each with 17 rows and 22 columns. Several sequencing projects have contributed to the sequencing of the porcine transcriptome in recent years (Tuggle et al., 2007). Frequent updating of the porcine expressed sequence tags (ESTs) necessitated the most recent annotation of the 70mers used in printing the microarrays for subsequent bioinformatics analyses. Each of the printed 70mers is derived from 64,746 (currently 110,744) Tentative Clusters (TCs) built from 575,730 (currently >1 million) ESTs and 6854 expressed transcripts from a total of 257 cDNA libraries (Current Porcine Gene Index: SsGI Release 14.0; March 11, 2010: Dana Farber Cancer Institute; http://compbio.dfci. harvard.edu/tgi/cgi-bin/tgi/gimain.pl?gudb¼pig). Using the 70mer sequences, a text file in FASTA format is created to store the printed oligomer sequence information. In order to obtain the most recent annotation, these sequences are first compared to the available 1185 porcine Reference Sequence Collection (RefSeq) from the National Center for Biotechnology Information (NCBI) using the Basic Alignment Search Tool (BLAST) with a required match of 64 bases with 94% identity. As a result, 830 printed oligomers were annotated with porcine RefSeq IDs. Second, the same sequences were compared to human RefSeqs with a stringency of blast p-value less than 0.001. This resulted in the annotation of 4180 printed oligomers with human RefSeq IDs. Third, 70mers were translated into peptide sequences using “blastx” in all six possible reading frames and the resulting peptide sequences were compared to UniProt100 database, which contains the translation of coding sequences of multiple genomic databases. Matches with blast p-value of less than 0.001 were retained. Since UniProt100 contains information about multiple species, a word comparison script was used to collapse the matches preferentially to porcine, human, mouse, and rat species for a total of 4111 UniProt IDs. In rare instances, matches for other species are allowed if porcine, human, mouse, or rat were not available. Finally, similar to step three, the TC sequences (instead of the 70mers) are translated into peptide sequences using “blastx” in all six possible reading frames and the resulting peptide sequences were compared


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to UniProt100. Using these approaches, 8962 of the 10,798 printed oligomers are fully annotated. Functional annotation of the microarray is achieved by mapping the Uniprot IDs to GO IDs. The GO describes gene products based on their associated biological processes, cellular components, and molecular functions in a species-independent manner. Microarray spots mapped to 3153 unique GO “biological process,” 1885 GO “molecular function,” and 599 GO “cellular component” IDs. Information about the microarray and its full annotation can be found with accession number A-CBIL-16 at ArrayExpress http://www.ebi.ac.uk/ microarray-as/ae/.

4.2. Microarray data preprocessing One thousand four hundred and ninety spots which correspond to control spots (blanks, Cy3, and Strategene Alien) are first filtered out. For each assay, saturated spots in at least one channel are set to NA. For each channel, the mean signal measure from the Agilent Feature Extraction software is used as input signal intensity. No background subtraction is performed. M and A values are calculated from the raw data using Eqs. (7.1) and (7.2) where R and G are signals of the Cy5 (red) and Cy3 (green) channels, respectively. M ¼ log2 R log2 G

ð7:1Þ

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The M values are normalized with print-tip loess normalization using the Bioconductor marray package (version 1.12.0) for R (version 2.4.0).

4.3. Analysis of differential gene expression Differential expression analysis is performed using Patterns of Gene Expression (PaGE version 5.1.6; http://www.cbil.upen.edu/PaGE; Grant et al., 2005). PaGE is a false discovery rate (FDR)-based method of controlling false positives. It uses a permutation-based algorithm to estimate the FDR. In PaGE, for any specified constant, permutations of the data matrix are used to estimate the rate of false positives in any set of genes having a T-statistic greater than the constant. An appropriate constant is chosen to guarantee the desired FDR, typically ranging between 0.25 and 0.05 (75–95% confidence). Confidences are then assigned to all genes in the set. For example, if 100 genes are discovered as differentially expressed with a confidence of 0.75 (i.e., FDR of 0.25), the expected number of false

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positives is 25. PaGE also produces “levels” of differential expression, based on the confidence parameter. Detailed information about the PaGE algorithm can be found at http://www.cbil.upenn.edu/PaGE/doc/PaGE_documentation_technical_manual.pdf. Differentially expressed genes in various comparisons are obtained using processed M values as input to PaGE. The above procedures usually result in a long list of genes differentially expressed above a significant threshold set by the investigator through PaGE. Further data processing procedures allow the physiological and pathological implications of subsets of these genes to be evaluated through their commonalities, associations, and network connectivities, and the identification of dominant differential pathways. The use of several of these procedures is next outlined in the context of UPR as a prominent differentially expressed characteristic of athero-susceptible endothelium.

4.4. Identification of enriched biological themes Differentially expressed genes are interrogated for overrepresented biological themes using database for annotation visualization and integrated discovery (DAVID) and based on GO terms (Huang da et al., 2009). The DAVID functional annotation clustering tool highlights the most relevant GO terms associated with a differentially expressed gene list. Details of the DAVID algorithm can be found at http://david.abcc.ncifcrf.gov/.

4.5. Gene set enrichment analysis GSEA is an algorithm that performs differential expression analysis at the level of gene sets (Subramanian et al., 2005). The input to GSEA consists of a collection of gene sets and microarray expression data with replicates for two conditions to be compared. GSEA employs a permutation-based test which uses Kolmogorov–Smirnov running sum statistic to determine which of the gene sets from the collection are differentially expressed between the two conditions. GSEA differs from differential gene expression analysis in the sense that it might identify genes which are part of a differentially expressed set but which might not be identified as significantly differentially expressed alone. The details of the GSEA algorithm can be found at http://www.broad.mit.edu/gsea/. For GSEA of various comparisons, for example, athero-susceptible versus athero-protected arteries, gene sets made up of 15–500 genes were created using GO mappings of the microarray. A total of 548 GO biological process (BP), 256 GO molecular function (MF), and 142 GO cellular component (CC) gene sets were used for this analysis. For each comparison, GSEA is performed separately for BP, MF, and CC gene sets. Gene sets are analyzed at an FDR set by the investigator.


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4.6. Ingenuity pathway analysis Differentially expressed genes are analyzed for interactions (from scientific literature) with each other and other molecules using IPA (Ingenuity Systems, www.ingenuity.com; Calvano et al., 2005). A list of differentially expressed genes with multispecies UniProt identifiers (to cover homologies) with their corresponding confidence and expression values is uploaded into the IPA application. A confidence cutoff is set by the investigator to identify genes which are used to interrogate the relationships curated to include literature findings. Details of the IPA algorithm can be found at http://www.ingenuity.com/library/index.html. In our particular endothelial experiments, IPA identified 73% of the upregulated genes in atherosusceptible endothelium to form a tightly connected network of interactions based on known gene–protein and protein–protein direct relations (Fig. 7.3) many of which are linked to ER stress and UPR (Civelek et al., 2009).

Figure 7.3 IPA of endothelial gene expression. A dataset containing gene identifiers with corresponding expression values and FDRs were analyzed using IPA. An FDR value cutoff of 25% was set to identify genes whose expression was significantly differentially regulated. Networks of these focus genes were then algorithmically generated based on their connectivity. Only direct interactions of gene–protein and protein–protein were considered. Seven networks were combined to form one large network. Red color indicates EC gene upregulation in athero-susceptible regions of arteries compared to athero-protected regions. Intensity of red is proportional to expression ratio. Gray color indicates molecules present in the data set that were not significantly differentially expressed (FDR > 25%).

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5. Another Approach to Gene Connectivity: Weighted Gene Coexpression Network Analysis In recent studies (Civelek et al., submitted for publication), we are using an additional approach to global endothelial gene expression across the arterial system. Weighted gene coexpression network analysis (WGCNA) is a systems biology approach to construct a weighted gene coexpression network to identify groups of genes (modules) whose expression is highly correlated using the WGCNA R package (Langfelder and Horvath, 2008). A comprehensive tutorial with several examples for the methodology is available at http://www.genetics.ucla.edu/labs/horvath/ CoexpressionNetwork/Rpackages/WGCNA/. Eighty seven samples were used for a recent network analysis. For each probe, a connection strength measure is determined by the pairwise correlations between expression profiles. Overall connectivity (k) for each probe is calculated by taking the sum of its connection strength with all other probes in the network (see WGCNA R tutorial). About 5579 probes with k > 5 are used for further analysis (Ghazalpour et al., 2006). A similarity matrix is constructed by calculating the biweight midcorrelation, which is robust to outliers, for all pairwise comparison of probe expression across all microarray samples (Wilcox, 2005). This correlation matrix is then transformed into a matrix of connection strengths (adjacency matrix) using a power function (connection strength ¼ (correlation)b), resulting in a weighted network. The parameter b was chosen by using the scale-free topology criterion (Langfelder and Horvath, 2008) to be six such that the resulting network connectivity distribution approximated scale-free topology. The adjacency matrix is used to calculate the topological overlap matrix (TOM) (Yip and Horvath, 2007). Modules are groups of genes with similar patterns of connection strengths with all other genes of the network and “high topological overlap” (Zhang and Horvath, 2005). Modules are identified using the TOM in conjunction with average linkage hierarchical clustering. A dynamic tree-cutting algorithm is used to identify the modules (Langfelder et al., 2008). Each module is summarized by its first eigengene (first principal component of the expression values across samples) and modules with similar gene expression identified by highly correlated eigengenes (correlation coefficient > 0.95) are merged. In our study, samples are grouped according to athero-susceptibility (susceptible vs. protected) or circulatory bed (coronary vs. noncoronary). Significant association of each module with either athero-susceptibility (susceptible ¼ 1 and protected ¼ 0) or circulatory bed (coronary ¼ 1 and noncoronary ¼ 0) is identified by calculating the Pearson correlation of module eigengene and each classification and by calculating the Student asymptotic p-value for the given

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correlations. The significant modules are then assessed for enrichment in GO terms using DAVID based on Fisher’s exact test utilizing the 5579 probe annotations as reference gene list (Huang da et al., 2009). This is then visualized.

5.1. Network visualization Gene interactions are visualized using Cytoscape (Cline et al., 2007). Intramodular connectivity for each gene is calculated taking the sum of its connection strength with all other genes in the same module. The gene connectivity is scaled by the maximum connectivity strength which results in the gene with the most connections having a connectivity value of 1. Genes with connectivity (k) less than 0.3 are filtered, resulting in 3192 genes (nodes) and 48,990 interactions (edges) for network visualization. The layout is obtained using an edge-weighted spring embedded algorithm using connectivity as the edge weight (Fig. 7.4). The distance between two nodes is proportional to the strength of the connection between two nodes, with highly connected genes being the closest to each other. The nodes are colored according to the module colors they are assigned by the WGCNA algorithm. For individual module network visualization, all the interactions between module genes are used without filtering. The node size is proportional to the intramodular connectivity of each gene with

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Figure 7.4 Arterial endothelium gene coexpression network. Highly connected genes were portioned into modules of similar expression profiles using the WGCNA algorithm. An interaction network was constructed using gene connectivity between two genes as the distance of an edge between two genes (nodes). The colors indicate different modules of highly correlated endothelial genes.

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the others in the same module. Largest node represents the gene with the highest connectivity and is denoted as the hub gene of the module. This particular approach revealed that endothelium in coronary and noncoronary arterial beds exhibit markedly different gene expression profiles but that transcript profiles in coronary arteries are consistent with the presence of ER stress/UPR, as previously shown in non-coronary arteries.

6. Validation and Follow-up Identification of differentially expressed pathways using global genomics directs the investigator to pathways, biomarkers, or clusters of molecules of interest for further hypothesis-driven, focused investigations. Typically, the important initial steps in this transition are to validate differential expression of representative genes by quantitative real-time PCR. The development of commercial multiplex-based qRT-PCR instrumentation has replaced gene-by-gene analyses with chip analyses. This can readily be performed on very small numbers of cells (thus retaining spatial relevance). Expression of proteins encoded by genes of interest can be quantified by Western blot if sufficient cells are available (potential loss of spatial resolution). Standard protocols for these procedures are readily available. Our studies of athero-susceptible endothelial phenotyping of UPR were validated and expanded at both the transcript (by qRT-PCR) and protein (Westerns) levels as outlined in the Section 1.1 (Civelek et al., 2009). Besides our study, two additional reports using complementary approaches support our findings and provide further mechanistic insights about site-specific endothelial ER stress, hemodynamics, and athero-susceptibility. Feaver et al. (2008) used an in vitro model to simulate human arterial shear stress waveforms. Atherosusceptible or athero-protective flow was applied to human endothelial cells. BiP (GRP78) was found to be significantly upregulated in a sustained manner under athero-susceptible, but not athero-protective flow up to 24 h. This response was dependent on both sustained activation of p38, as well as integrin a2b1. Increased BiP expression correlated with the activation of the ER stress sensing element promoter by athero-susceptible flow as a marker of the UPR. Shear stress regulation of BiP was through increased protein stability when compared to other flow regulated proteins, such as connexin-43 and vascular cell adhesion molecule (VCAM)-1. Increased endothelial expression of BiP was also observed in athero-susceptible versus athero-protective regions of C57BL6 mice. The study supports a role for the hemodynamic environment in preferentially inducing BiP and the UPR in athero-susceptible regions before lesion development. Zeng et al. (2009) reported that spliced XBP-1 (sXBP-1) encodes the XBP-1 transcription factor that translocates to the nucleus to


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activate selective pro-apoptotic target genes as one of the three transduction arms of the UPR response. Following the observation of endothelial expression of the XBP-1 pathway of UPR in branching regions of apoE/ mice arteries and in atherosclerotic lesions that developed there, their study reported that athero-susceptible flow waveforms induced XBP-1 splicing in cultured endothelial cells. Overexpression of (activated) sXBP-1 induced apoptosis in cultured human endothelial cells. To extend the findings to an in vivo assay for atherogenesis, adenoviral-mediated overexpression of sXBP-1 was induced in an apoE/ murine aortic isograft model. In these animals, enhanced intimal hyperplasia and atherosclerosis developed in normally protected regions of the aorta suggesting that when the XBP-1 UPR pathway is greatly overstimulated, the adaptive protective function of UPR reverts to a pathological imbalance. While overexpression was not entirely limited to the endothelium in the isograft model, the data are supportive for a prominent role for endothelial sXBP-1. These three different but complementary approaches to endothelial ER stress provide compelling evidence for the existence of site-specific chronic adaptive UPR in endothelial cells in vivo; the hemodynamic environment associated with sites of athero-susceptibility likely plays a significant role. The principles and procedures outlined for global genomics in this chapter can be applied to other levels of regulation in the same cells. For example, we have recently reported differential microRNA analyses of athero-susceptible endothelium by microarray to show posttranscriptional regulation of pro-inflammatory phenotypes by these small RNAs without preconceived expectation of their target pathways (Fang et al., 2010).

ACKNOWLEDGMENTS We gratefully acknowledge American Heart Association Fellowship support to MC (0315286U) and National Institute of Health grants HL062250 (PFD) and HG004521 (CS, EM, GG).

REFERENCES Calvano, S. E., Xiao, W., Richards, D. R., Felciano, R. M., Baker, H. V., Cho, R. J., Chen, R. O., Brownstein, B. H., Cobb, J. P., Tschoeke, S. K., Miller-Graziano, C., Moldawer, L. L., et al. (2005). A network-based analysis of systemic inflammation in humans. Nature 437, 1032–1037. Civelek, M., Manduchi, E., Riley, R. J., Stoeckert, C. J., Jr., and Davies, P. F. (2009). Chronic endoplasmic reticulum stress activates unfolded protein response in arterial endothelium in regions of susceptibility to atherosclerosis. Circ. Res. 105, 453–461. Cline, M. S., Smoot, M., Cerami, E., Kuchinsky, A., Landys, N., Workman, C., Christmas, R., Avila-Campilo, I., Creech, M., Gross, B., Hanspers, K., Isserlin, R., et al. (2007). Integration of biological networks and gene expression data using Cytoscape. Nat. Protoc. 2, 2366–2382.

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Davies, P. F. (2009). Hemodynamic shear stress and the endothelium in cardiovascular pathophysiology. Nat. Clin. Pract. Cardiovasc. Med. 6, 16–26. Fang, Y., Shi, C., Manduchi, E., Civelek, M., and Davies, P. F. (2010). MicroRNA-10a regulation of proinflammatory phenotype in athero-susceptible endothelium in vivo and in vitro. Proc. Natl. Acad. Sci. USA 107, 13450–13455. Feaver, R. E., Hastings, N. E., Pryor, A., and Blackman, B. R. (2008). GRP78 upregulation by atheroprone shear stress via p38-, alpha2beta1-dependent mechanism in endothelial cells. Arterioscler. Thromb. Vasc. Biol. 28, 1534–1541. Ghazalpour, A., Doss, S., Zhang, B., Wang, S., Plaisier, C., Castellanos, R., Brozell, A., Schadt, E. E., Drake, T. A., Lusis, A. J., and Horvath, S. (2006). Integrating genetic and network analysis to characterize genes related to mouse weight. PLoS Genet. 2, e130. Grant, G. R., Liu, J., and Stoeckert, C. J., Jr. (2005). A practical false discovery rate approach to identifying patterns of differential expression in microarray data. Bioinformatics 21, 2684–2690. Huang da, W., Sherman, B. T., and Lempicki, R. A. (2009). Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources. Nat. Protoc. 4, 44–57. Langfelder, P., and Horvath, S. (2008). WGCNA: An R package for weighted correlation network analysis. BMC Bioinform. 9, 559. Langfelder, P., Zhang, B., and Horvath, S. (2008). Defining clusters from a hierarchical cluster tree: The Dynamic Tree Cut package for R. Bioinformatics 24, 719–720. Malhotra, J. D., and Kaufman, R. J. (2007). The endoplasmic reticulum and the unfolded protein response. Semin. Cell Dev. Biol. 18, 716–731. Passerini, A. G., Polacek, D. C., Shi, C., Francesco, N. M., Manduchi, E., Grant, G., Pritchard, W. P., Powell, S. J., Chang, G., Stoeckert, C., and Davies, P. F. (2004). Coexisting pro-inflammatory and anti-oxidative endothelial transcription profiles in a disturbed flow region of the adult porcine aorta. Proc. Natl. Acad. Sci. USA 101, 2482–2487. Polacek, D. C., Passerini, A. G., Shi, C., Francesco, N. M., Manduchi, E., Grant, G. R., Powell, S., Bischof, H., Winkler, H., Stoeckert, C. J., Jr., and Davies, P. F. (2003). Fidelity and enhanced sensitivity of differential transcription profiles following linear amplification of nanogram amounts of endothelial mRNA. Physiol. Genomics 13, 147–156. Subramanian, A., Tamayo, P., Mootha, V. K., Mukherjee, S., Ebert, B. L., Gillette, M. A., Paulovich, A., Pomeroy, S. L., Golub, T. R., Lander, E. S., and Mesirov, J. P. (2005). Gene set enrichment analysis: A knowledge-based approach for interpreting genomewide expression profiles. Proc. Natl. Acad. Sci. USA 102, 15545–15550. Tuggle, C. K., Wang, Y., and Couture, O. (2007). Advances in swine transcriptomics. Int. J. Biol. Sci. 3, 132–152. Van Gelder, R. N., von Zastrow, M. E., Yool, A., Dement, W. C., Barchas, J. D., and Eberwine, J. H. (1990). Amplified RNA synthesized from limited quantities of heterogeneous cDNA. Proc. Natl. Acad. Sci. USA 87, 1663–1667. Wilcox, R. R. (2005). Introduction to Robust Estimation and Hypothesis Testing, second edn. Elsevier/Academic Press, Amsterdam, Boston. Yip, A. M., and Horvath, S. (2007). Gene network interconnectedness and the generalized topological overlap measure. BMC Bioinform. 8, 22. Zeng, L., Zampetaki, A., Margariti, A., Pepe, A. E., Alam, S., Martin, D., Xiao, Q., Wang, W., Jin, Z. G., Cockerill, G., Mori, K., Li, Y. S., et al. (2009). Sustained activation of XBP1 splicing leads to endothelial apoptosis and atherosclerosis development in response to disturbed flow. Proc. Natl. Acad. Sci. USA 106, 8326–8331. Zhang, B., and Horvath, S. (2005). A general framework for weighted gene co-expression network analysis. Stat. Appl. Genet. Mol. Biol. 4, 1–45.

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Assessment of Endoplasmic Reticulum Stress and the Unfolded Protein Response in Endothelial Cells Ines Witte and Sven Horke Contents 1. Introduction 2. The Acute Phase of ER Stress 2.1. Calcium influxes analyzed by laser scanning microscopy 3. The Adaptive Reaction Phase: Unfolded Protein Response in Endothelial Cells 3.1. ATF6 pathway 3.2. eIF2a phosphorylation (PERK pathway) 3.3. XBP1 splicing (IRE1 pathway) 3.4. GRP78 3.5. CHOP 4. The Final Phase: Cell Death Signaling 4.1. Measurement of mitochondrial membrane potential DCm by microplate reader or laser scanning microscopy 4.2. Use of flow cytometry (FACS) to detect ER stress-induced oxidative stress 4.3. ER stress-induced caspase activation Acknowledgments References

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Abstract In the vascular wall, the most inner cell layer that separates the blood from organelles is comprised of only a single layer of endothelial cells (ECs). This cell type is fundamental to a large variety of processes, ranging from blood coagulation and interaction with inflammatory cells to cardiovascular diseases such as hypertension, diabetes, and atherosclerosis. Dysfunction of ECs is often causally linked to these processes such that research exploring such events attracted much attention. Damage of ECs and subsequent disruption of the intact endothelial barrier can result not only from oxidative stress, but also from Institute of Pharmacology, University Medical Center of the Johannes Gutenberg University Mainz, Germany Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00008-X

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conditions that stress the endoplasmic reticulum (ER) and induce a signaling pathway termed unfolded protein response (UPR). While its primary goal is to alleviate ER stress, the UPR can also induce cell death. Cultured ECs are often used in in vitro approaches to understand various pathophysiological events, but they behave differently from many other cell types such that cell-typespecific procedures are needed. Here, we describe how ER stress can be induced and assessed in cultured ECs and demonstrate their specific responses to classical ER stress conditions.

1. Introduction Endothelial cells (ECs) form the inner luminal layer of lymphatic and vascular walls, which is also known as the intima. They coat the interior surface as a single layer independent of the type and size of blood vessel and are found in the entire vascular system. Importantly, ECs do not just form a passive barrier between blood and organs. Rather, they have pleiotropic functions and regulate, and therefore impact on, coagulation, fibrinolysis, angiogenesis, adhesion and transmigration of inflammatory cells, homing of macrophages in the subendothelial cells (where they may form foam cells), and also blood pressure (through the production of vasoactive substances such as nitric oxide or endothelin). Moreover, ECs transport and/or secrete a large variety of (macro-) molecules and/or hormones, and they play important roles in the degradation of lipoprotein particles. Therefore, EC dysfunction has serious consequences and contributes to many diseases, such as hypertension, atherosclerosis, diabetes, systemic lupus erythematosus, arthritis, or bleeding disorders, all of which involve an enhanced generation of oxidative stress and endoplasmic reticulum (ER) stress (Feng et al., 2003; Forstermann, 2008; Forstermann and Munzel, 2006; Simionescu, 2007). ER stress activates the unfolded protein response (UPR) pathway, which increases the expression of proteins that promote the capacity of the ER to handle client proteins. This involves activation of ER stress transducers PERK, IRE1, and ATF6 and aims at reducing a cell’s ER stress burden. However, programmed cell death signaling pathways are also induced and the balance of these responses determines cellular survival or death. Given the vital roles of ECs, there is a clear need to analyze their functions in vitro, and many different primary and immortalized cells are available (Bouis et al., 2001). An often used immortalized EC line is EA.hy 926, which is a good and reliable alternative to primary human umbilical vein endothelial cells (HUVECs), from which they have been generated; they still share many, if not all, specific characteristics such as production of NO and vWF. We found no major differences between these two cells, and data presented for EA.hy 926 cells can be easily adapted to primary

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HUVECs. However, ECs clearly differ from other cell types, and when stress pathways are to be experimentally induced in ECs, it is mandatory to explore their specific responses. In the vasculature, ECs produce and secrete many different factors which predispose them to ER stress, a process augmented during atherogenesis when ECs are exposed to accumulating oxidized lipids and adjacent macrophage foam cells. Therefore, the concentration of pharmacological compounds needed to experimentally induce ER stress is much lower and may induce a specific reaction profile. There are many situations when a researcher wants to investigate whether a certain treatment or stimulus induces ER stress and activates the UPR, and whether a compound or protein enhances or attenuates this response. In order to compare to a specific treatment of choice, there are several commercially available toxins that cause ER stress in cultured ECs. Tunicamycin inhibits N-glycosylation, which causes accumulation of unmodified proteins in the ER. The lactone antibiotic brefeldin A blocks ER to Golgi protein transport and induces retrograde protein transfer, likewise leading to the accumulation of cargo proteins in the ER. Thapsigargin inhibits the sarco-/ER Ca2þ ATPase (SERCA) that transports Ca2þ into the ER, causing depletion of ER Ca2þ stores. Dithiothreitol (DTT) is a reducing agent which induces ER stress because it causes the accumulation of unfolded proteins by inhibiting disulfide bond formation of proteins. However, DTT nonspecifically alters the redox status of practically all organelles and thus could affect many pathways. To apply these compounds and analyze a protein’s function, reasonable concentrations and time points should be used to avoid adverse side effects and misinterpretations. Moreover, acute reactions must be discriminated from long-term effects, and it should be considered that up to 40% of ECs can undergo cell death after 2 days of ER stress by tunicamycin (1 mg/ml; measured by 7-AAD/ Annexin-V flow cytometry; not shown). Here, we present a detailed description of how UPR activation can be analyzed and show endothelial-specific responses to various concentrations of tunicamycin, thapsigargin, and/or DTT. To this end, we demonstrate the impact of ER stress on Ca2þ oscillations during the first 20 min, UPR activation within the first 8 h and ROS production, and mitochondrial membrane potential and caspase activation after 1–3 days.

2. The Acute Phase of ER Stress 2.1. Calcium influxes analyzed by laser scanning microscopy Several stimuli involve Ca2þ signaling or cause Ca2þ disturbances, particularly when considering ER stress and signaling to mitochondria, as these organelles are the cells’ main Ca2þ storage compartments. These may be

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very fast Ca2þ fluxes and can be observed as acute effects or long-lasting events. Many methods and dyes exist to detect Ca2þ-fluxes. An alternative to common fura dyes is Fluo-4-AM (Molecular Probes), which is retained in cells and generates bright green fluorescence upon Ca2þ interaction. Actually, detection is very similar to that of DCm, as confocal microscopy, microplate readers, or FACS analysis may be used, depending on the availability and the goal of the experiment. Results from a microplate approach can be seen elsewhere (Horke et al., 2010). For end-point analysis of longer time points, flow cytometry may be an alternative and can be performed similarly to the protocol in Section 4.2. In this section, we wished to analyze linkage between ER stress and acute cytosolic Ca2þ influx with emphasis on the inducing stimulus. To this end, cells were stimulated with low or high doses of thapsigargin and tunicamycin. Thapsigargin involves direct interference with Ca2þ homeostasis (via SERCA inhibition), whereas tunicamycin blocks protein glycosylation. Both induce ER stress, but the underlying mechanism determines Ca2þ fluxes and Ca2þ-responsive functions, for example, the ER stressattenuating role of PON2 (Horke et al., 2008). Here, we used a laser scanning microscopy (LSM) method to reveal acute Ca2þ fluxes. This approach allows simultaneous quantification of many and inspection of individual cells. Also, it reveals Ca2þ oscillations and outperforms microplate readers, which can only collect data from populations, rather than individual cells. Since cells in one population do not necessarily react synchronously, reading individual cells is much more advantageous. Required material

Chambered lab-Tek slides (Nunc or Greiner Bio-One) with thin glass bottoms or opaque cell culture dishes suitable for confocal microscopy; surfaces ranging from that of 96- to 24-well dishes are appropriate, for example, Greiner Bio-one black m-clear 96- or 24-well plates. Sterile cell culture bench, autoclaved pipettes, and plastic tips. ECs seeded the day before the experiment at 8 104 cells/24-well. HBSS buffer with Ca2þ and Mg2þ (PAA) at 37 C. Fluo-4-AM (1–5 mM; Molecular Probes) and nuclear staining dye (depending on the available filter sets, you may use Dapi, Draq5, or ToPro3; e.g., from Molecular Probes or Cell Signaling Technology) appropriately diluted in prewarmed HBSS. A laser scanning microscope equipped with suitable lasers and filter sets, a heating chamber, and the ability to record and quantify time-lapse series. We use a Zeiss LSM-710 equipped with ZEN-2008 software, the 488nm laser for Fluo-4-AM (emission 492–629 nm) and the 633-nm laser for Draq5 (emission 616–757 nm), an EC Plan-Neofluar 10/0.30 M27 or 20/0.50 M27 objective, and a heating chamber.


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LSM procedure: Seed ECs at 8 104 cells/well of a 24-well dish the day before the experiment. Then, growth medium is removed, cells are washed twice with HBSS buffer, loaded with Fluo-4-AM in HBSS buffer (200 ml/24-well) and stored in an incubator for 20–40 min. Meanwhile, arrange the microscope to allow immediate measurement; prepare solutions of HBSS containing twice the concentration of tunicamycin or thapsigargin that needs to be analyzed. Wash cells and add HBSS, 200 ml/well and include nuclear markers; we observed that after only single staining, Draq5 is not very stable and rather diffuse after >1 h, thus we analyzed cells in buffer containing this nuclear dye or we added it freshly to each sample. Embed the culture plate in the heating chamber of the microscope and focus on a field containing well-stained cells. Add 200 ml of HBSS (with 2 concentration of tunicamycin or thapsigargin), quickly ensure that cells are still focused and start time-lapse series immediately. The latter part of the experiment is critical and must be performed as fast as possible, since the fluorescence changes significantly within 20–120 s under stress conditions. We used the following parameters: EC Plan-Neofluar 10/0.30 M27 objective, pixel dwell 1.58 ms, image size 1024 1024, 90 cycles 10 s each, pinhole 2) after recovery of cells or the next day, which may be experimentally tested. After treatment, aspirate the growth medium and wash the cells once with PBS, add 100 ml (per 24-well) of freshly prepared passive lysis buffer contained in the Dual-Glo Luciferase Assay System, and rock at room temperature for 15 min. You may pause at this step and store plates at 20 C. Perform two freeze–thaw steps with liquid nitrogen to break cells, either with the whole plate or after previous transfer into test tubes. Follow the details of the Dual-Glo Luciferase Assay System for subsequent sample handling. Record luciferase chemiluminescence with a Berthold microplate


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luminometer. Normalize firefly to renilla activity; include untransfected samples to estimate signal intensity (transfected cells must have much higher activities). GRP78 Western blotting Follow the above guidelines for detection of p-eIF2a with the following modifications: PhosphoStop may be omitted; extend the time of protein transfer to the membrane to 25 min; we used goat-anti-GRP78 antibody C-20 from Santa Cruz with a 1:500 dilution in 5% milk powder in TBS-T buffer overnight at 4 C. The enhanced GRP78 protein expression in response to ER stress is dose-dependent and clearly visible with low concentrations of tunicamycin (>0.3 mg/ml) or thapsigargin (>1 nM) (Fig. 8.2C).

3.5. CHOP The CHOP (Gadd153) protein is a very potent pro-apoptotic factor induced by the UPR pathway and other signaling events. The UPR is a protective pathway, but if the stress condition remains insurmountable, the pro-apoptotic arm of the pathway dominates the outcome. It is not fully understood how CHOP signals to the cell death machinery and which genes are modulated by CHOP, but several studies demonstrated its central role in ER stress-induced cell death. If ER stress occurs, CHOP expression is significantly enhanced and can be followed by promoter reporter studies, qRT-PCR, and Western blotting. Required material

For mRNA analysis by qRT-PCR, follow the above protocol for detection of XBP1. Taqman primers for CHOP: sense (50 -GACCTGCAAGA GGTCCTGTC-30 ); antisense (50 -CTCCTCCTCAGTCAGCCAAG30 ); probe (50 -CACCACTCTTGACCCTGCTT-30 ). To detect CHOP at the protein level, modifications of the above protocols are needed Tank blot buffer: 25-mM Tris, 192-mM glycine, 20% methanol RIPA cell lysis buffer: 50-mM Tris–HCl (pH 8.0), 150-mM NaCl, 1% NP-40 (or Triton X-100), 0.5% Na-deoxycholate, 0.1% SDS Rabbit-anti-CHOP F168 antibody from Santa Cruz Mini-Protean-III SDS–PAGE/Western blotting apparatus from BioRad. Use 1.5-mm glass plates to load up to 70 ml per lane.

Procedure for CHOP protein detection: Seed ECs the day before the experiment using 8 104/dish of a 24-well dish. After treatment, lyse cells in a 65-ml RIPA/well, denature for 5 min at 95 C, and separate by 12% SDSPAGE in Mini-Protean-III systems (Bio-Rad). Run gels with 100 V for 10 min and 150 V for approx. 1 h. Then, replace the SDS-PAGE inset with the Western blotting accessories and transfer protein by tank blotting, under

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constant cooling for 1.5 h and with 200-mA/minigel. Remove membrane, block with 5% milk powder in TBS-T, and then incubate with anti-CHOP antibody (1:200) at 4 C overnight. Subsequent steps follow the above standard procedures. It is important to note that several commercial antibodies detect other proteins but not CHOP (e.g., see Haataja et al., 2008) or are too unspecific such that many bands appear which complicates evaluation. Therefore, we generally recommend to verify correct mass, repeat with pre-immune serum if available, compare normal and stressed conditions (8 h; tunicamycin > 1 mg/ml), use siRNA (smart-pool; Dharmacon) to knock down CHOP, or add suitable CHOP-peptides to block the antibody (if available). Due to the absence of CHOP under unstressed conditions and the high induction after ER stress, it is advisable to perform time and/or dose–response experiments.

4. The Final Phase: Cell Death Signaling 4.1. Measurement of mitochondrial membrane potential DCm by microplate reader or laser scanning microscopy Mitochondrial membrane potential is a reliable measure of cell stress and apoptosis, as DCm is compromised early during stress response and a good indicator of ongoing cell death. Several dyes can be used to detect DCm, which allow flexibility in wavelengths and combinations with other fluorescence markers. Due to their bright signal and stability, detection can be performed with a (confocal laser scanning) microscope, microplate reader, or by flow cytometry. Microscope and microplate approaches allow following DCm during the course of the experiment or as an end-point analysis, while FACS determines end-points only. Here, we describe end-point analysis using the two former methods: Required material

Chambered lab-Tek slides (Nunc or Greiner Bio-One) with thin glass bottoms or other opaque cell culture dishes suitable for (confocal) microscopy or microplate readers; surfaces ranging from that of 96- to 24-well dishes are appropriate, for example, Greiner Bio-one black m-clear 96- or 24-well plates. Microplate readers can be used provided appropriate fluorescence detection is available; continuous measurements require a heating chamber. We use a FluoStar-Optima from BMG Labtech with heating unit and 485/520-nm filters for excitation/emission, respectively. Sterile cell culture bench, autoclaved pipettes, and plastic tips. ECs seeded the day before the experiment at an appropriate density (see below). HBSS buffer with Ca2þ and Mg2þ (PAA), prewarmed


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DiOC6 (Molecular Probes) and nuclear staining dye (see above) diluted in prewarmed HBSS Though confocal microscopy may not be mandatory, we use a Zeiss LSM-710 equipped with ZEN-2008 software, 488-nm laser for DiOC6 (emission 492–629 nm), and 633-nm laser for Draq5 (emission 616– 757 nm), an EC Plan-Neofluar 10/0.30 M27 or 20/0.50 M27 objective. This allows simultaneous inspection of 150–200 cells/visible field. LSM procedure: ECs are seeded in appropriate culture dishes. The duration of the experiment must be considered and subconfluent cultures are preferred, as confluent cells have lower respiratory activity and are much densely packed, which together complicates analysis of DCm. Here, we describe the procedure for a 4-chambered slide (where each surface resembles that of one 24-well dish) and use 0.5–0.8 105 cells/chamber. One day after seeding or after 8 h at the earliest, cell treatment is started. Subsequently, cells are washed twice with a 500-ml HBSS and stained with 25-nM DiOC6 and Draq5 for 20 min in a cell culture incubator, 200–300 ml/chamber. Draq5 allows simultaneous assessment of nuclear morphology and identification of cells with absent DiOC6 staining. In the meantime, prepare the microscope to record fluorescence at the appropriate wavelengths. Then, cells are washed, transferred to the microscope, and immediately processed. Using the control samples, adjust all relevant parameters, such as resolution (usually 1096 1096), pixel dwell (> 2 ms), laser power, gain, and pinhole (4/group/experiment) and a number of repetitions to reach statistical significance, as the differences in signal intensity may be small and also refer to the sensitivity of your reader. If DCm is to be followed for a longer time, microplate readers may be advantageous, as they do not have the putative drawback of cells moving out of focus; however, you only receive fluorescence intensities and cannot inspect cells visually.

4.2. Use of flow cytometry (FACS) to detect ER stress-induced oxidative stress A large body of evidence showed that ER stress directly or indirectly provokes the generation of reactive oxygen species (ROS) (for an overview, see Gorlach et al., 2006; Haynes et al., 2004; Malhotra and Kaufman, 2007; Santos et al., 2009). This occurs at multiple sites and different time points, for example, during disulfide bond formation in the ER (as a by-product or via glutathione depletion) or in response to activity of the mitochondrial electron transport chain. Superoxide (O2) is converted to hydrogen peroxide (H2O2) by spontaneous or enzymatic reaction; the intracellular iron content in turn determines the formation of highly reactive hydroxyl radical (OH) via the Fenton reaction. But O2 can also generate peroxynitrite (ONOO), hypochlorous acid (HOCL), or singlet oxygen (1O2); thus, many antioxidant defense mechanisms exist to combat overwhelming ROS generation and damage. When analyzing ROS, there may be inconclusiveness regarding techniques and reporter systems, as there exists a plethora of dyes and methods. Generally, it is critical to consider the radical species detected (referring to specificity of the reporter system), intracellular localization, time point, and stimulus (see other chapters for a detailed insight; Gomes et al., 2005). As a starting point and to address the overall occurrence of oxidative stress, it seems advisable to use conditions unspecific for particular radicals or subcellular structures and systems allowing easy detection suitable with many devices and applications. The fluorescent reporter H2DCFDA fulfills many of these criteria, as it fluoresces in the FITC (GFP) A, but employing flow cytometry to analyze ROS formation by assessment of H2DCFDA fluorescence in ER stress-induced cells. (C) EA.hy 926 cells were treated with the indicated concentrations of DTT, tunicamycin, or thapsigargin for 24 h and subsequently analyzed for caspase-3/7 activation. Reproduced with permission from Horke et al. (2008). # The Biochemical Society.

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channel upon ROS, reacts with several radicals, and has low technical demand. Therefore, H2DCFDA-loaded cells can be analyzed by many methods such as microscopy and microplate readers (which allow timelapse analysis; see, e.g., Altenhofer et al., 2010; Horke et al., 2007). An alternative is flow cytometry (FACS) to determine end-point ROS formation: Required material

ECs seeded the day before the experiment at 8 104 cells/well of a 24-well dish HBSS buffer with Ca2þ and Mg2þ (PAA), prewarmed Cm-H2DCFDA (2 mM; Molecular Probes) diluted in prewarmed HBSS Trypsin–EDTA solution (PAA), prewarmed, for cell removal from growth plates Warm growth medium for trypsin neutralization 5-ml Polystyrene Round-Bottom Tubes (BD Falcon) Appropriate flow cytometry equipped with suitable laser and filter sets (FITC). We use FACS-Calibur equipped with CellQuest Pro software (BD Biosciences).

FACS procedure: EA.hy 926 cells are seeded in 24-well plates with a density at 8 104 cells/well. One day after seeding, cell treatment is started for the desired time. Subsequently, cells are washed with a 500-ml HBSS and stained with 2-mM cm-H2DCFDA for 30 min in a cell culture incubator, 500 ml/well. Then, cells are trypsinized, resuspended in growth medium, pelleted (350 g, 5 min, RT), resuspended in a 200-ml HBSS and immediately analyzed by flow cytometry (FITC filter set). Use control samples to adjust all relevant parameters. The comparison between untreated and tunicamycin/thapsigargin-treated samples reveals a pronounced generation of ROS production secondary to ER stress (Fig. 8.3B).

4.3. ER stress-induced caspase activation If the ER stress-inducing stimulus remains insurmountable, cell death is induced. Which precise signaling events are involved is a matter of debate, especially because this not only differs between mice and humans, but also depends on the inducing stimulus and the cell type. We and others found that mitochondrial oxidative signaling and also caspase activation can be involved in ER stress. However, several methodological approaches must be used to discriminate between survival and different cell death pathways, especially since the UPR intersects, for example, with autophagy (Galluzzi et al., 2009; Klionsky et al., 2008). In addition to microscopic inspection, the effects of ER stress on DCm (see above) can be expanded by determination of caspase activation.


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Caspase-Glo-3/7 assay kit from Promega Inc. Opaque, white 96-well plates, cell culture grade, from Greiner-Bio-One Sterile cell culture bench, autoclaved pipettes, and plastic tips; PBS buffer (PAA); ECs seeded the day before the gene reporter experiment with 1.2–2 104 cells/well of a 96-well dish. Centro LB960 plate luminometer (Berthold Technologies, Germany) Procedure: Seed cells as mentioned above include untreated and empty wells (a positive control is helpful, such as treating ECs with 0.5-mM staurosporine for 16 h). The day after seeding, treat cells as indicated and measure caspase activity according to the Caspase-Glo-3/7 assay kit instructions. For your treatment, mind the final volume per well, since an equal volume of Caspase-Glo reagent is added at the end. We found that ECs tolerated volumes as low as 40 ml/96-well. However, depending on the intensity of caspase activation, it must be verified that maximal signal intensity is not exceeded if only low amounts of caspase substrate reagent were added (as the activated caspases could cleave the substrate in an unproportional fashion). Include >3 samples for each group and repeat the experiment accordingly to reach statistical significance. The activation of caspase-3/7 in ECs in response to different concentrations of tunicamycin, thapsigargin, or DTT can be seen in Fig. 8.3C. Assaying caspase activation (cleavage of pro-caspases) by Western blotting may add to this approach.

ACKNOWLEDGMENTS We thank Petra Wilgenbus for excellent technical advice and performance, Ulrich Fo¨rstermann for support and scientific discussions, and David M. Zuckerman for critically reading the manuscript. Financial support came from the University Medicine Mainz and the German Research Foundation (DFG-grant HO3925/4-1). The authors have no conflict of interest to declare.

REFERENCES Altenhofer, S., Witte, I., Teiber, J. F., Wilgenbus, P., Pautz, A., Li, H., Daiber, A., Witan, H., Clement, A. M., Forstermann, U., and Horke, S. (2010). One enzyme, two functions: PON2 prevents mitochondrial superoxide formation and apoptosis independent from its lactonase activity. J. Biol. Chem. 285, 24398–24403. Bouis, D., Hospers, G. A., Meijer, C., Molema, G., and Mulder, N. H. (2001). Endothelium in: A review of human vascular endothelial cell lines for blood vessel-related research. Angiogenesis 4, 91–102. Chen, X., Shen, J., and Prywes, R. (2002). The luminal domain of ATF6 senses endoplasmic reticulum (ER) stress and causes translocation of ATF6 from the ER to the Golgi. J. Biol. Chem. 277, 13045–13052.

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Feng, B., Yao, P. M., Li, Y., Devlin, C. M., Zhang, D., Harding, H. P., Sweeney, M., Rong, J. X., Kuriakose, G., Fisher, E. A., Marks, A. R., Ron, D., et al. (2003). The endoplasmic reticulum is the site of cholesterol-induced cytotoxicity in macrophages. Nat. Cell Biol. 5, 781–792. Forstermann, U. (2008). Oxidative stress in vascular disease: Causes, defense mechanisms and potential therapies. Nat. Clin. Pract. Cardiovasc. Med. 5, 338–349. Forstermann, U., and Munzel, T. (2006). Endothelial nitric oxide synthase in vascular disease: From marvel to menace. Circulation 113, 1708–1714. Galluzzi, L., Aaronson, S. A., Abrams, J., Alnemri, E. S., Andrews, D. W., Baehrecke, E. H., Bazan, N. G., Blagosklonny, M. V., Blomgren, K., Borner, C., Bredesen, D. E., Brenner, C., et al. (2009). Guidelines for the use and interpretation of assays for monitoring cell death in higher eukaryotes. Cell Death Differ. 16, 1093–1107. Gomes, A., Fernandes, E., and Lima, J. L. (2005). Fluorescence probes used for detection of reactive oxygen species. J. Biochem. Biophys. Methods 65, 45–80. Gorlach, A., Klappa, P., and Kietzmann, T. (2006). The endoplasmic reticulum: Folding, calcium homeostasis, signaling, and redox control. Antioxid. Redox Signal. 8, 1391–1418. Haataja, L., Gurlo, T., Huang, C. J., and Butler, P. C. (2008). Many commercially available antibodies for detection of CHOP expression as a marker of endoplasmic reticulum stress fail specificity evaluation. Cell Biochem. Biophys. 51, 105–107. Haynes, C. M., Titus, E. A., and Cooper, A. A. (2004). Degradation of misfolded proteins prevents ER-derived oxidative stress and cell death. Mol. Cell 15, 767–776. Horke, S., Witte, I., Wilgenbus, P., Kruger, M., Strand, D., and Forstermann, U. (2007). Paraoxonase-2 reduces oxidative stress in vascular cells and decreases endoplasmic reticulum stress-induced caspase activation. Circulation 115, 2055–2064. Horke, S., Witte, I., Wilgenbus, P., Altenhofer, S., Kruger, M., Li, H., and Forstermann, U. (2008). Protective effect of paraoxonase-2 against endoplasmic reticulum stress-induced apoptosis is lost upon disturbance of calcium homoeostasis. Biochem. J. 416, 395–405. Horke, S., Witte, I., Altenhofer, S., Wilgenbus, P., Goldeck, M., Forstermann, U., Xiao, J., Kramer, G. L., Haines, D. C., Chowdhary, P. K., Haley, R. W., and Teiber, J. F. (2010). Paraoxonase 2 is down-regulated by the Pseudomonas aeruginosa quorumsensing signal N-(3-oxododecanoyl)-L-homoserine lactone and attenuates oxidative stress induced by pyocyanin. Biochem. J. 426, 73–83. Klionsky, D. J., Abeliovich, H., Agostinis, P., Agrawal, D. K., Aliev, G., Askew, D. S., Baba, M., Baehrecke, E. H., Bahr, B. A., Ballabio, A., Bamber, B. A., Bassham, D. C., et al. (2008). Guidelines for the use and interpretation of assays for monitoring autophagy in higher eukaryotes. Autophagy 4, 151–175. Kyhse-Andersen, J. (1984). Electroblotting of multiple gels: A simple apparatus without buffer tank for rapid transfer of proteins from polyacrylamide to nitrocellulose. J. Biochem. Biophys. Methods 10, 203–209. Malhotra, J. D., and Kaufman, R. J. (2007). Endoplasmic reticulum stress and oxidative stress: A vicious cycle or a double-edged sword? Antioxid. Redox Signal. 9, 2277–2293. Santos, C. X., Tanaka, L. Y., Wosniak, J., and Laurindo, F. R. (2009). Mechanisms and implications of reactive oxygen species generation during the unfolded protein response: Roles of endoplasmic reticulum oxidoreductases, mitochondrial electron transport, and NADPH oxidase. Antioxid. Redox Signal. 11, 2409–2427. Simionescu, M. (2007). Implications of early structural–functional changes in the endothelium for vascular disease. Arterioscler. Thromb. Vasc. Biol. 27, 266–274. Wek, R. C., Jiang, H. Y., and Anthony, T. G. (2006). Coping with stress: eIF2 kinases and translational control. Biochem. Soc. Trans. 34, 7–11. Witte, I., Altenho¨fer, S., Wilgenbus P., Amort, J., Clement, A.M., Pautz, A., Li, H., Fo¨rstermann, U., Horke, S. (2011). Beyond reduction of atherosclerosis: PON2 provides apoptosis resistance and stabilizes tumor cells. Cell Death Dis. (in press).

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Measuring the Impact of Cigarette Smoke on the UPR Hong Zhao,* Jin Yang,† Lin Shan,‡ and Ellen D. Jorgensen* Contents 148 149 150

1. Introduction 2. Preparing Samples for Analysis 2.1. Exposure of cells to CS in vitro 2.2. Collection and preparation of minimally invasive clinical samples 3. Analysis of Activation of the PERK Pathway 3.1. Transfection of A549 human lung cells to silence PERK 3.2. Preparation of nuclear fraction to measure ATF4 nuclear translocation 4. ATF6 Pathway 4.1. Required materials 4.2. Transfection of A549 cells with ATF6 plasmid 5. IRE1 Pathway 5.1. Required materials 5.2. XPB1 splicing assayed by PCR 6. Using LSC to Measure UPR Activation 6.1. Required materials 6.2. Procedure Acknowledgments References

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Abstract The unfolded protein response (UPR) is a set of pathways activated by the accumulation of improperly folded proteins. It can be triggered by a broad range of stressful conditions which disrupt successful maturation of proteins in the endoplasmic reticulum (ER) by interfering with proper folding, assembly, and posttranslational modification. Recent studies have demonstrated the induction of ER stress and activation of elements of the UPR in human lung * Department of Pathology, New York Medical College, Valhalla, New York, USA { Education and Training Systems International, Inc., Chapel Hill, North Carolina, USA { Pathology Department, Duke University Medical Center, Durham, North Carolina, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00009-1

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cells exposed to diesel exhaust particles, airborne particulate matter, and tobacco smoke. ER stress has been found to play a role in a variety of lung maladies, including cancer, infections, idiopathic pulmonary fibrosis, and chronic obstructive pulmonary disease. Lung cancer is one of the few diseases where the etiological agent, cigarette smoke (CS), is well known. It is, therefore, desirable to measure dysregulation of the UPR pathway in samples representing both the earliest events (cells exposed to CS in vitro) and in clinical samples from healthy smokers and individuals with smoking-related lung diseases. We hereby provide a detailed description of methods for assessing the degree and timing of cellular response to CS with respect to the three major UPR pathways.

1. Introduction The unfolded protein response (UPR) is a set of pathways activated by the accumulation of improperly folded proteins in the endoplasmic reticulum (ER). It can be triggered by a broad range of stressful conditions which disrupt successful maturation of proteins in the ER by interfering with proper folding, assembly, and posttranslational modification (for review, see Ron and Walter, 2007). The UPR functions to reduce the accumulation of aberrantly folded proteins and promote cell survival by reducing the ER client protein burden, as well as by increasing the ER folding and degradation capacities. However, if the stress is severe and homeostatic equilibrium in the ER is not restored, a secondary response (triggered, at least in part, by the same molecular sensors) promotes the induction of apoptosis and leads to cell death (Szegezdi et al., 2006). Thus, it is not surprising that ER stress has been found to play a role in a variety of maladies, including cancer, infections, and lung diseases such as idiopathic pulmonary fibrosis and chronic obstructive pulmonary disease (COPD) (Healy et al., 2009; Lawson et al., 2008; Lee and Hendershot, 2006; Lin et al., 2010; Malhotra et al., 2009). Analysis of human lung tissue samples reveals differential expression of UPR components in smokers versus nonsmokers and in lung cancer compared to nonmalignant lung ( Jorgensen et al., 2008; Kelsen et al., 2008). Recent studies have demonstrated the induction of ER stress and activation of elements of the UPR in human lung cells exposed to diesel exhaust particles, airborne particulate matter, and tobacco smoke (Hengstermann and Muller, 2008; Jorgensen et al., 2008; Jung et al., 2007; Kelsen et al., 2008; Laing et al., 2010; Watterson et al., 2009; Yoshida, 2007). The combination of these results suggests that chronic induction of one or more components of the UPR pathway may play an etiological role in lung disease. Upregulation of UPR protein effectors in lung cancers may strengthen cellular resistance to various cytotoxic stresses such as hypoxia,

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chemotherapeutic drugs, and immune attack, thus providing a pro-survival advantage (Fels and Koumenis, 2006; Koong et al., 2006; Li and Lee, 2006; Shu et al., 2007; Wang et al., 2008). This could be particularly true for cells that have one or more pro-apoptotic pathways disabled as is commonly the case in lung cancers (Haura et al., 2004). In mammalian cells, the first step in activation of UPR is the dissociation of the ER-resident master chaperone regulator binding immunoglobulin protein/glucose response protein 78 (BiP) from three UPR transducer proteins in the ER: activation of transcription factor 6 (ATF6), inositol requiring enzyme 1a (IRE1), and protein kinase R-like ER kinase (PERK). Each of these transmembrane proteins initiates a separate signaling branch of the UPR. The PERK pathway controls attenuation of translation via phosphorylation of the alpha subunit of eukaryotic translation initiation factor 2 (eIF2a), while the ATF6 and IRE1 pathways regulate transcriptional induction of various chaperones and components of protein degradation machinery in the ER. The three branches display distinct sensitivities toward different forms of ER stress (DuRose et al., 2006). Thus, it is of interest to measure the degree of involvement of the PERK, ATF6, and IRE1 pathways in response to cigarette smoke (CS) and the timing with which it occurs. In this chapter, we provide methods for characterizing the effect of CS on the three major pathways of the UPR. Many events in the PERK, IRE1, and ATF6 pathways can be assayed by Western blotting, but much depends on the quality of the available antibodies (Haataja et al., 2008). We describe which Western blot assays work well for each pathway, as well as procedures that provide more detailed information such as fractionation of cellular compartments and gene silencing. For the ATF6 pathway, we provide a method for monitoring the cleavage of ATF6, an event which signals the activation of that pathway. Indications that the IRE1 pathway is activated include phosphorylation of IRE1, which is detected by Western blot and cleavage of XBP1 RNA, which is assayed using PCR. We describe the preparation of RNA samples for microarrays and/or quantitative PCR; however, we will not cover these techniques in this chapter. The final method we will discuss is the use of laser scanning cytometry (LSC) to quantitate expression of UPR proteins in cells collected from smokers and nonsmokers.

2. Preparing Samples for Analysis Lung cancer is one of the few diseases where the etiological agent, CS, is well known. It is, therefore, possible to measure dysregulation of the UPR pathway in samples representing both the earliest events (cells exposed

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to CS in vitro) and in clinical samples from healthy smokers and individuals with smoking-related lung diseases.

2.1. Exposure of cells to CS in vitro One of the most important decisions made by the investigator wishing to measure the effect of CS is how to deliver it to cells. No in vitro exposure system can faithfully recreate the complicated environment of the human lung, so there must be an awareness of the inherent shortcomings of any particular setup. Many groups choose to use CS condensates or cigarette extracts dissolved in liquids such as dimethylsulfoxide (DMSO) that can be added to the cell culture medium (e.g., Fields et al., 2005). This approach avoids the need for a smoke delivery machine, as these materials can sometimes be purchased or otherwise obtained and stored frozen until use. However, there is a major caveat to using these materials, as CS contains not only a particulate phase but also a vapor phase which is not adequately represented in such preparations. The vapor phase of CS has been shown to be crucial to tumor formation in animal models (Witschi, 2005) and to strongly induce UPR components in experiments using in vitro systems ( Jorgensen et al., 2008). For these reasons, it is preferable to use whole smoke generated by a smoking machine. Such machines are commercially available and produce smoke in a calibrated manner. Parameters that can be varied include the puff volume, puff duration, and time interval between puffs. A second consideration is how the smoke from the machine will be delivered to the cells. Experiments using monolayer cell cultures have been criticized as less relevant than systems using so-called three-dimensional cultures. The latter rely on the propensity of normal human bronchial epithelial (NHBE) cells to form cultures that are several cells thick and show some differentiation into basal and epithelial layers. The cultures are prepared on a permeable mesh membrane in the form of an insert that fits into tissue culture systems in which the culture medium contacts the bottom of the membrane, feeding the basolateral layer, while the surface epithelium is in direct contact with the air/smoke (Deschl et al., 2010; Maunders, et al., 2007). In our experience, the molecular effects of CS on monolayer versus three-dimensional cultures are not divergent enough to warrant the extra effort of preparing and analyzing the more complicated system. Therefore, we recommend the use of human lung cells in monolayer cultures as described below for most purposes. 2.1.1. Required materials 2.1.1.1. Smoking machine/smoke exposure chamber The effect of direct exposure of cells to smoke in vitro is a comparatively new area of exploration, hence there is no widely accepted standard apparatus designed


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for this purpose. One common method for delivering CS to cells is to bubble the output from a commercial smoking machine through a phosphate buffered saline (PBS) solution and then replace the cell culture medium with this solution to treat the cells. Again, we prefer to use a direct exposure method that delivers the vapor phase gasses in a manner consistent with human smoking. To deliver smoke uniformly, it is necessary to construct an exposure chamber that is coupled to a commercial smoking machine (we use a KC 5 Port Smoker from KC Automation, Richmond, VA, but any machine capable of delivering CS in a calibrated manner is acceptable). In our device, each puff of CS is first drawn into a cylindrical aluminum 250 cc chamber which has a removable lid for cleaning. During cell treatment, 5% CO2 in air flows through the chamber and is mixed with incoming CS from the smoking machine via a stir bar. This dilution is necessary because direct exposure to whole smoke straight from the smoking machine results in delivery of smoke components at a level much higher than observed in human smoking behavior and was so toxic that even a 5 min exposure resulted in nearly 100% cell death. The CS/air mixture then passes through a separate rectangular aluminum exposure chamber containing the cell samples at a constant flow rate of 500 cc/min (maintained by an external pump and regulated by mass flow controllers). The chamber rests on a 1-in. thick aluminum platform which contains channels through which flow 37 ºC water, maintaining the exposure chamber temperature. Our setup also has an additional mass flow controller which allows for further dilution of the CS if desired. The standard smoke dilution used in most of our experiments is 35 cc delivered over 1 min in either a 250 or 500 cc volume, and the intensity of exposure can also be controlled by varying the length of time the cells spend in the exposure chamber (typically between 5 and 20 min). The time and distance that the smoke travels from the end of the cigarette to the exposure chamber is minimized by using the shortest lengths of tubing possible between the parts of the apparatus. Absorbance at 300 nm is used to quantify the “tar” deposition in PBS solutions that have been exposed to CS. By using PBS solutions containing known amounts of “tar” dissolved in DMSO, we have found the absorbance to be linear with respect to “tar” concentration in the range of 2.5–100 mg/mL. These measurements should be used to test the variability of CS exposure in different locations within your chamber, if it is large enough to accommodate several samples at once. 2.1.1.2. Human lung cells and media A549 cells (#CCL-185, American Type Culture Collection, Manassas, VA) cultured in Ham’s F12K medium with 2-mM L-glutamine adjusted to contain 1.5 g/L sodium bicarbonate (Gibco/Invitrogen, Carlsbad, CA) and supplemented with 10% fetal bovine serum (ATCC) NHBE cells (with retinoic acid) from nonsmoking, nondiabetic donors (#CC2540, Lonza, Walkersville, MD) cultured in complete

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bronchial epithelial cell growth medium, prepared by supplementing bronchial epithelial basal medium with retinoic acid, epidermal growth factor, epinephrine, transferrin, T3, insulin, hydrocortisone, antimicrobial agents, and bovine pituitary extract by addition of SingleQuotsTM (Lonza). 2.1.1.3. Other materials

Dulbecco’s PBS with Ca2þ and Mg2þ(Gibco/Invitrogen) 0.25% trypsin–EDTA RIPA cell lysis buffer (#8990, Thermo Scientific, Rockford, IL) HaltTM protease inhibitor cocktail kit (#78410, Thermo Scientific) Phosphatase inhibitor cocktails 1 and 2 (#P2850 and #P5726, respectively, Sigma–Aldrich Corp., St. Louis, MO) Protein concentration assay kit and standards (e.g., kits from Bio-Rad Laboratories, Hercules, CA) RNeasy mini kit for RNA extraction (#74104, Qiagen, Inc., Valencia, CA) QiaShredder columns (#79654, Qiagen, Inc.) Thapsigargin, tunicamycin, and dithiothreitol (DTT) (Sigma–Aldrich, St. Louis, MO) 35-mm tissue culture dishes Two-chambered glass slides (#154461, Thermo Scientific) Cell scrapers (#83-1830, Sarstedt, Inc., Newton, NC) 1.5-mL microcentrifuge tubes Cigarettes

2.1.2. CS treatment Seed A549 or NHBE cells into 35-mm cell culture dishes at a density of 105 cells/35-mm dish or into two-chambered slides at 5 104 cells per chamber to achieve approximately 70% confluency at the time of exposure to CS. Always treat NHBE cell cultures before their fifth passage. Immediately before treatment, replace the cell culture medium with 37 C PBS containing calcium and magnesium, because it is superior to media in terms of maintaining pH. Prepare your mock-treated samples first. Remove the covers from the dishes or chambers and place them in the smoke exposure chamber. We typically generate CS under Federal Trade Commission (FTC, as per Federal Register, 1967) smoking conditions (35 0.3 cc puff, one puff every 60 s, 2-s puff duration with none of the ventilation holes blocked) using 2R4F reference research cigarettes (designed to represent the average “lights” cigarette marketed in the USA with FTC values of 9.7 mg “tar” and 0.85 mg nicotine; University of Kentucky, Louisville, KY). For the mock exposure, do not place a cigarette in the smoking port. Following mock treatment, remove the PBS covering the cells and replace it with 1 mL per chamber of fresh culture medium at 37 C. Place cells in the 37 C, 5% CO2 incubator for the desired time period. Next, prepare


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CS-exposed samples. The procedure is the same as for the mock-treated, except that a cigarette is placed in the smoking port and lit. Smoke cigarettes to within 3 mm of the filter tip, replacing them with a fresh cigarette if longer exposure times are desired. 2.1.3. Positive control treatments Prepare positive control samples. Use 1-mM thapsigargin, 10-mg/mL tunicamycin, or 2-mM DTT added to the cell culture medium in DMSO (thapsigargin and tunicamycin) or water (DTT) and incubate in a 37 C, 5% CO2 incubator. 2.1.4. Sample preparation: Cell lysis for Western blots Prepare lysis buffer by adding protease and phosphatase cocktails to RIPA buffer according to the manufacturer’s instructions. Treat cells in 35-mm cell culture dishes. Following treatment, remove the culture medium and rinse the cell monolayers twice with 2 mL per dish 4 ºC PBS. Add 1 mL (per 106 cells) of RIPA cell lysis buffer containing protease and phosphatase inhibitors. Dislodge cells using a cell scraper and transfer to a 1.5-mL microcentrifuge tube. Pipet each sample up and down vigorously a few times to promote lysis, then let sit on ice for 25 min. Centrifuge tubes at 10,000g for 25 min at 4 C and transfer the supernatants to fresh tubes. Determine protein concentration of the supernatants using protein standards and assay kit. Lysates can be frozen and stored at 80 ºC for future use. 2.1.5. Sample preparation: RNA extraction for PCR and microarray analysis Treat cells in 35-mm cell culture dishes. For microarray experiments, we typically prepare four dishes for each experimental condition and use the RNA from each dish for a separate array. Aspirate the medium from the dish and rinse twice with 1 mL 37 ºC PBS. Add 350 mL of Buffer RLT per dish. Pipet up and down a few times to lyse cells. Homogenize cell lyzates by putting them through a QIAshredder spin column and then follow the manufacturer’s instructions to extract RNA using an RNeasy mini kit. For NHBE and A549 cells, each 35-mm dish should yield about 10 mg RNA. Freeze the eluted RNA and store at 80 C if it will not be used immediately. Using this procedure, we typically produce RNA of extremely high quality for use in either PCR or microarray experiments. The details of both these technologies are outside the scope of this chapter, but we have used them in the past to assay for transcription of UPR-related genes ( Jorgensen et al., 2008).

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2.1.6. Sample preparation: Fixing slides for LSC Seed cells onto chambered slides 2 days before treatment at 5 104 cells per chamber. Cells should be at about 70% confluence at the time of treatment. Following treatment, fix cells with 1% formaldehyde in PBS on ice for 15 min. Rinse with PBS and store in ethanol at 20 C for at least 2 h before proceeding with immunochemistry.

2.2. Collection and preparation of minimally invasive clinical samples One of the difficulties of characterizing the molecular pathology of CS in vivo is that tissue from the target organ, the lung, is accessible only via invasive procedures. In order to investigate early stage events, it would seem necessary to use such techniques as broncoscopy to obtain relevant samples for analysis (Beane et al., 2007; Steiling et al., 2009). Recently, however, it has been reported that there is a correlation between the molecular pattern of lung and the tissues from buccal and nasal regions (Boyle et al., 2010; Sridhar et al., 2008; Zhang et al., 2010), suggesting that these tissues may be an acceptable surrogate for lung tissue in studies aimed at identifying the biomarkers of tobacco-related harm in CS-exposed individuals. 2.2.1. Required materials

Toothbrush (for buccal cells) Cytosoft cytology brush (#CYB-1, Medical Packaging Corporation, Camarillo, CA) (for nasal passage cells) Glass microscope slides Dako delimiting pen (#S2002 Dako North America, Carpinteria, CA) 50-mL polypropylene tubes (#352098 Becton Dickinson Corp.) Ice-cold PBS Coplin jars with screw caps Ice-cold 80% ethanol Ice-cold 1% formaldehyde (methanol-free) in PBS

2.2.2. Collecting buccal mucosal and nasal passage cells Prepare glass slides. Using the Dako delimiting pen, mark the boundaries of the analysis area by drawing a box about 1 in. square on the slides. Place slides on a level surface before collecting the sample. Immediately before the sample is taken, add about 10-mL ice-cold PBS to a 50-mL tube. Collect cheek cell samples by using a toothbrush to scrape the insides of both cheeks gently from the upper to the lower portion of the mouth four times (this process will take approximately 5–10 s). Then, dip the toothbrush into the tube containing ice-cold PBS. Gently agitate the toothbrush for 30 s to


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release the cells. Cap the tube and vortex for 3 s to break up cell clumps and form a cell suspension. Using a plastic transfer pipette, remove a small amount of the cell suspension and carefully place one drop within the box drawn on each slide. Note that it is very important that this step be done immediately after vortexing. The cells will start to settle to the bottom of the tube almost immediately, so it is crucial to put the drops of cell suspension on the slides before this happens. Gently tilt each slide until the drop spreads out over most of the boxed area. Leave on a level bench top to air dry for at least 30 min. When slides show no signs of wetness, carefully submerge the slides without agitation in a Coplin jar that has been filled to the top of the slide holder grooves with ice-cold 1% formaldehyde. Let them fix for 15 min. Fill a second Coplin jar with ice-cold 80% ethanol and transfer the slides into it. Cap the jar and store the slides at 4 C where they should be stable for several months. Nasal cell collection is carried out essentially the same as for buccal mucosal cells, except for the initial collection step where a cytology brush is used to gently scrape cells from the inferior nasal turbinates.

3. Analysis of Activation of the PERK Pathway In response to ER stress, PERK undergoes autophosphorylation and activation of a kinase function that phosphorylates eIF2a, resulting in a transient repression of protein synthesis. The PERK pathway is particularly amenable to Western blot analysis, as there are good commercially available antibodies for many key pathway members, including phosphorylated and unphosphorylated forms of PERK and eIF2a. The general procedure for Western blotting is well known and we have described it previously in terms of measuring UPR components in cells treated with CS ( Jorgensen et al., 2008). In our experience, eIF2a becomes phosphorylated relatively quickly, within 30 min following smoke treatment. Table 9.1 lists the timing of UPR events as assayed by Western blot in A549 cells responding to a 20-min treatment with CS. Useful information regarding the appearance of various UPR proteins after treatment with thapsigargin, tunicamycin, and DTT has already been published (Durose et al., 2006).

3.1. Transfection of A549 human lung cells to silence PERK In mammalian cells there are three other kinases in addition to PERK that are capable of phosphorylating eIF2a (PKR, HRI, and GCN2, reviewed in Wek et al., 2006). Although they appear to be regulated by different stress stimuli (e.g., accumulation of unfolded proteins, amino acid deprivation, or oxidative stress), their catalytic domains are homologous. Phosphorylation of eIF2a results in similar downstream events including translation attenuation and

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Table 9.1 Sequence of UPR events in human lung cells following 20 min exposure to cigarette smoke Event

Pathway

Time posttreatment (h)

Increase in BiP Phosphorylation of eIF2a Appearance of GADD34 Nuclear translocation of ATF4 Appearance of ATF3 Appearance of GADD153/CHOP Cleavage of ATF6 Cleavage of XBP1 mRNA Increase in IRE1

All PERK PERK PERK PERK PERK ATF6 IRE1 IRE1

24 0.5 4–7 4 2–4 2–4 1 None None

activation of transcriptional programs that augment the cell’s ability to cope with stress, regardless of whether it is accomplished via PERK or by one of the other kinases. PERK-induced activation of eIF2a is believed to occur exclusively upon the induction of ER stress (Ron and Walter, 2007). Therefore, it is necessary to demonstrate the dependence of CS-induced phosphorylation of eIF2a on the phosphorylation of PERK in order to show that CS treatment specifically induces ER stress. This can be accomplished by transfecting cells with a plasmid containing siRNA designed to silence PERK, followed by exposure to CS. A similar procedure could be used to silence the other kinases, if desired. 3.1.1. Required materials

100-mm2 tissue culture dishes A549 cells and appropriate growth medium, with and without serum (see Section 2.1.1) 20 mM stock of PERK siRNA oligos (Dharmacon, Lafayette, CO, catalog # L-004883-00) 20 mM stock of nontarget siRNA oligos (Dharmacon, catalog # D001810-10) DharmaFECT1 transfection reagent (Dharmacon, catalog # T-2001-02). 3.1.2. Transfection procedure Seed A549 cells in 100-mm2 culture dishes at 106 cells per dish. Place in 37 ºC/ 5% CO2 incubator. At 20 h postseeding, prepare transfection medium. Add 21 mL of DharmaFECT1 to 679 mL of serum-free medium and incubate at room temperature for 5 min. In a separate tube, add 70 mL of siRNA oligos to 630 mL of serum-free medium and incubate at room temperature for 5 min.


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Then, combine the DharmaFECT1 and siRNA oligo solutions and incubate at room temperature for an additional 20 min. At the end of the incubation period, add 12.6 mL of complete growth medium to complete the transfection medium. Remove the existing medium from the cell culture dishes and replace with a 14-mL transfection medium per dish. Place in 37 ºC/5% CO2 incubator for 6 h, after which the medium is replaced with fresh prewarmed growth medium. Return to the incubator for another 20 h. Trypsinize and reseed into 35-mm dishes at 1 105 cells per dish and return to the incubator for another 48 h prior to CS treatment.

3.2. Preparation of nuclear fraction to measure ATF4 nuclear translocation Phosphorylation of eIF2a promotes the induction of activating transcription factor 4 (ATF4) which accumulates in the nucleus and upregulates genes whose function is to adapt the cell to ER stress. These include CCAAT/ enhancer-binding protein-homologous protein (CHOP/GADD153) and activating transcription factor 3 (ATF3), which act in concert to coordinate the upregulation of growth arrest and DNA damage-inducible protein 34 (GADD34) which has a feedback inhibitory effect on UPR (Novoa et al., 2001). While the upregulation of ATF3, CHOP, and GADD34 can be assessed via Western blotting of whole cell lyzates ( Jorgensen et al., 2008), the translocation of ATF4 requires isolation of nuclei prior to analysis. 3.2.1. Required materials

NHBE or A549 cells (see Section 2.1.1) 100-mm2 cell culture plates 0.25% trypsin–EDTA Qproteome Cell Compartment Kit (# 37502, Qiagen, Inc.)

3.2.2. Fractionation procedure Seed NHBE or A549 cells at 6 105 cells per 100-mm2 dish 2 days prior to CS treatment. Note that this kit recommends 5 106 cells as an adequate amount of starting material, so you will have to treat several dishes. After treatment, remove cells from dishes by trypsinization. Using the Qproteome Cell Compartment Kit, carry out the cellular compartment according to the manufacturer’s instructions. Briefly, pellet the cells by centrifugation and then sequentially extract with a series of three buffers, resulting in isolation of proteins from different cell compartments, including the cytosol, organelles, and the nucleus. This material can be used for Western blot analysis. Make sure to verify the purity of nuclear extracts by including an antibody for the specific nuclear protein Lamin A/C ( Jorgensen et al., 2008).

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4. ATF6 Pathway ATF6 is a 90-kDa protein that translocates from the ER to the Golgi in response to stress, where it is proteolytically processed, resulting in a functional 50-kDa transcription factor. In mammalian cells, it has been reported to be responsible for inducing the transcription of ER chaperone genes and, in concert with the IRE1 pathway, the transcription of ER-associated degradation components (Yamamoto et al., 2007). Since the level of ATF6 in human lung cells is low, some sort of enrichment process is needed in order to use Western blotting to monitor the proteolytic process. We approached the problem by transfecting our cells of interest with a plasmid constituitively expressing ATF6 ( Jorgensen et al., 2008). The elevated amounts of ATF6 in the transfected cells allowed us to visualize the disappearance of the 90-kDa band on a Western blot without prior immunoprecipitation.

4.1. Required materials

NHBE or A549 cells (see Section 2.1.1) 35-mm cell culture dishes ATF6 expression plasmid ATF6/pCMV6-XL5 (# SC115551, OriGene Technologies, Inc., Rockville, MD) pCMV6-XL5 control plasmid (#PCMV6XL5 OriGene Technologies, Inc.) Lipofectamine LTX (#15338-100, Invitrogen) PLUS Reagent (#11514-015, Invitrogen)

4.2. Transfection of A549 cells with ATF6 plasmid At 24 h before transfecting, seed A549 cells in 35-mm culture dishes at 1.2 105 cells per dish. When ready to transfect, dilute 2-mg ATF6/ pCMV6-XL5 or control plasmid DNA in a 400-mL serum-free medium and mix with PLUS Reagent at 1:1 ratio (DNA mg: PLUS vol. in mL). Incubate the mixture at room temperature for 5 min. Add Lipofectamine LTX to the DNA/PLUS mixture and incubate at room temperature for an additional 25 min. At the end of the incubation, remove the original growth medium and replace with 2 mL of fresh prewarmed growth medium per dish. Immediately add the DNA/PLUS/Lipofectamine complex to the dishes, mix gently, and return to the incubator. After 6 h, replace the medium with fresh prewarmed complete growth medium and return the cells to the incubator. Treat cells at 24 h posttransfection.


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5. IRE1 Pathway Dissociation from BiP leads to IRE1 oligomerization and autophosphorylation. This activates the ribonuclease function in its cytoplasmic domain, resulting in the catalysis of an unconventional splicing event in the X-box-binding protein 1 (XBP1) mRNA. The alternative carboxyl terminal domain in the translated protein facilitates its role as a transcriptional activator and allows it to modulate the expression of several genes involved in ER homeostasis. The most straightforward method of assessing the involvement of the IRE1 pathway is to measure phosphorylation of IRE1. However, since we have had limited success with the phosphorIRE1 antibodies commercially available at this writing, we recommend that a PCR-based assay be used for the cleavage of XBP1 mRNA.


RNA harvested using the Qiagen RNeasy mini kit according to the manufacturer’s instructions. High-Capacity cDNA Reverse Transcription kit for RT-PCR (#4374966, Applied Biosystems, Foster City, CA). PCR primers 50 -CTGGAAAGCAAGTGGTAG A-30 and 50 -CTGG GTCCTTCTGGGTAG AC-30 AmpliTag Cold DNA polymerase (#N808-0241, Applied Biosystems). Agarose gel DNA electrophoresis apparatus and reagents

5.2. XPB1 splicing assayed by PCR Perform first-strand cDNA synthesis the High-Capacity cDNA Reverse Transcription kit for RT-PCR, following the manufacturer’s instructions. Using the primers, amplify via PCR for 35 cycles (95 C for 30 s; 58 C for 30 s; 72 C for 1 min) with AmpliTag Cold DNA polymerase. Fragments representing spliced and unspliced XBP1 (398 and 424 bp, respectively) can be visualized by analyzing the samples on 2% agarose gels with ethidium bromide staining.

6. Using LSC to Measure UPR Activation LSC analysis combines the advantages of flow and image cytometry (Powzarowsky et al., 2006). The cells are fixed on slides, so there is minimal loss of sample during cell staining and measurement. Therefore,

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multiparametric measurements can be achieved using samples containing only small numbers of cells. The dissociation of BiP from PERK, ATF6, and IRE1 in response to an accumulation of misfolded proteins is an early event in the UPR; however, continued stress leads to an accumulation of BiP ( Jorgensen et al., 2008). Using LSC, we have measured BiP expression in buccal mucosal cells from smokers and nonsmokers (Fig. 9.1). Nuclear staining, secondary antibodies, and instrument parameters were as described in Jorgensen et al. (2010). LSC technology can be adapted to assay any UPR-related protein or phosphorylated protein for which there is a robust, specific antibody that works in formaldehyde-fixed cells. We typically use DAPI to stain for DNA content and to allow the instrument to contour the cell nuclei. In this manner, we can distinguish between nuclear and cytoplasmic fluorescence. Since every cell is imaged, we can also determine on a cell-by-cell basis if a protein is confined to a particular subcellular location. A detailed description of LSC and its parameters may be found in Powzarowsky et al. (2006).


Slides with fixed cells (see Sections 2.1.6 and 2.2.2) Coplin jar PBS 1% Triton in PBS 1% bovine serum albumin (BSA) in PBS Primary antibody (e.g., rabbit anti-BiP) Nonsmoker

Smoker

4 BiP expression

BiP expression

4 3 2 1

3 2 1

0

1 2 DNA content

0

1 2 DNA content

Figure 9.1 Comparison of BiP protein levels in buccal mucosal cells harvested from smokers and nonsmokers. Samples were prepared as described in Section 2.2. Each dot represents data from an individual cell on the slide. DNA content as assayed by DAPI labeling is plotted against BiP expression measured by fluorescent antibody labeling.


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Secondary antibody (e.g., mouse anti-rabbit) conjugated with fluorophore DAPI (40 ,6-diamidino-2-phenylindole) 1 mg/mL in PBS Long cover slips SlowFade Gold antifade reagent (#S36936, Invitrogen, Eugene, OR) Plastic box with lid

6.2. Procedure Remove the slides from ethanol storage. Note that between each step of this procedure, it helps to shake off the excess solution from the slides before proceeding to the next step. Place the slides in a Coplin jar filled with PBS for 5 min. Repeat with fresh PBS. Remove the slides and add several drops of 1% Triton to the area of the slide containing the cells. Let sit at room temperature for 15 min. Rinse the slides with PBS. Add several drops of 1% BSA to the area of the slide containing the cells. Incubate at room temperature for 30 min. Shake off excess and add 100–200 mL primary antibody. The volume will depend on how large an area is covered by the cells on your slide. We typically start with a 1:100 dilution of the antibody in 1% BSA, but if an unknown antibody is being used, then a series of dilutions may be advisable to test compatibility. Incubate either at room temperature for 1.5 h or overnight at 4 ºC (preferred). If incubating overnight, place the slides on top of a paper towel that has been dampened with water inside a plastic box with a lid. This prevents evaporation of the antibody solution. Afterward, place the slides in a Coplin jar filled with PBS at room temperature for 5 min. Following this step, all subsequent steps are at room temperature in the dark. Remove slides and add 100–200 mL secondary antibody and incubate for 45 min. Place the slides in a Coplin jar filled with PBS for 5 min. Remove the slides and place one or two drops of DAPI solution on each slide. Incubate for 15 min, rinse with PBS, and shake off the excess. Put one drop of antifade onto each slide and carefully place a cover slip on top, trying not to retain any bubbles. It helps to set one edge of the coverslip on the slide and, maintaining a little tension on it so it bends slightly, lay it down gently from left to right. The sample is now ready for LSC analysis. We typically use DAPI to stain for DNA content and to allow the instrument to contour the cell nuclei. A detailed description of LSC and its parameters may be found in Powzarowsky et al. (2006).

ACKNOWLEDGMENTS We thank Dr. Zbigniew Darzynkiewicz and Dr. Frank Traganos of the Brander Cancer Research Institute at New York Medical College for the use of their laser scanning cytometry facility.

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REFERENCES Beane, J., Sebastiani, P., Liu, G., Brody, J. S., Lenburg, M. E., and Spira, A. (2007). Reversible and permanent effects of tobacco smoke exposure on airway epithelial gene expression. Genome Biol. 8, R201. Boyle, J. O., Gu¨mu¨s, Z. H., Kacker, A., Choksi, V. L., Bocker, J. M., Zhou, X. K., Yantiss, R. K., Hughes, D. B., Du, B., Judson, B. L., Subbaramaiah, K., and Dannenberg, A. J. (2010). Effects of cigarette smoke on the human oral mucosal transcriptome. Cancer Prev. Res. 3, 266–278. Deschl, U., Vogel, J., and Aufderheide, M. (2010). Development of an in vitro exposure model for investigating the biological effects of therapeutic aerosols on human cells from the respiratory tract. Exp. Toxicol. Pathol. [Epub ahead of print]. DuRose, J. B., Tam, A. B., and Niwa, M. (2006). Intrinsic capacities of molecular sensors of the unfolded protein response to sense alternate forms of endoplasmic reticulum stress. Mol. Biol. Cell 17, 3095–3107. Federal Register (1967). 32, 11–178. Fels, D. R., and Koumenis, C. (2006). The PERK/eIF2alpha/ATF4 module of the UPR in hypoxia resistance and tumor growth. Cancer Biol. Ther. 5, 723–728. Fields, W. R., Leonard, R. M., Odom, P. S., Nordskog, B. K., Ogden, M. W., and Doolittle, D. J. (2005). Gene expression in normal human bronchial epithelial (NHBE) cells following in vitro exposure to cigarette smoke condensate. Toxicol. Sci. 86, 84–91. Haataja, L., Gurlo, T., Huang, C. J., and Butler, P. C. (2008). Many commercially available antibodies for detection of CHOP expression as a marker of endoplasmic reticulum stress fail specificity evaluation. Cell Biochem. Biophys. 51, 105–107. Haura, E. B., Cress, W. D., Chellappan, S., Zheng, Z., and Bepler, G. (2004). Antiapoptotic signaling pathways in non-small-cell lung cancer: Biology and therapeutic strategies. Clin. Lung Cancer 6, 113–122. Healy, S. J., Gorman, A. M., Mousavi-Shafaei, P., Gupta, S., and Samali, A. (2009). Targeting the endoplasmic reticulum-stress response as an anticancer strategy. Eur. J. Pharmacol. 625, 234–246. Hengstermann, A., and Muller, T. (2008). Endoplasmic reticulum stress induced by aqueous extracts of cigarette smoke in 3T3 cells activates the unfolded-protein-response-dependent PERK pathway of cell survival. Free Radic. Biol. Med. 44, 1097–1107. Jorgensen, E., Stinson, A., Shan, L., Yang, J., Gietl, D., and Albino, A. P. (2008). Cigarette smoke induces endoplasmic reticulum stress and the unfolded protein response in normal and malignant human lung cells. BMC Cancer 8, 229–259. Jorgensen, E. D., Zhao, H., Traganos, F., Albino, A. P., and Darzynkiewicz, Z. (2010). DNA damage response induced by exposure of human lung adenocarcinoma cells to smoke from tobacco- and nicotine-free cigarettes. Cell Cycle 9(11), [Epub ahead of print]. Jung, E. J., Avliyakulov, N. K., Boontheung, P., Loo, J. A., and Nel, A. E. (2007). Prooxidative DEP chemicals induce heat shock proteins and an unfolding protein response in a bronchial epithelial cell line as determined by DIGE analysis. Proteomics 7, 3906–3918. Kelsen, S. G., Duan, X., Ji, R., Pere, O., Liu, C., and Merali, S. (2008). Cigarette smoke induces an unfolded protein response in the human lung: A proteomic approach. Am. J. Respir. Cell Mol. Biol. 38, 541–550. Koong, A. C., Chauhan, V., and Romero-Ramirez, L. (2006). Targeting XBP-1 as a novel anti-cancer strategy. Cancer Biol. Ther. 5, 756–759. Laing, S., Wang, G., Briazova, T., Zhang, C., Wang, A., Zheng, Z., Gow, A., Chen, A. F., Rajagopalan, S., Chen, L. C., Sun, Q., and Zhang, K. (2010). Airborne particulate matter selectively activates endoplasmic reticulum stress response in the lung and liver tissues. Am. J. Physiol. Cell Physiol. [Epub ahead of print].


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Lawson, W. E., Crossno, P. F., Polosukhin, V. V., Roldan, J., Cheng, D. S., Lane, K. B., Blackwell, T. R., Xu, C., Markin, C., Ware, L. B., Miller, G. G., Loyd, J. E., et al. (2008). Endoplasmic reticulum stress in alveolar epithelial cells is prominent in IPF: Association with altered surfactant protein processing and herpesvirus infection. Am. J. Physiol. Lung Cell Mol. Physiol. 294, L1119–L1126. Lee, A. S., and Hendershot, L. M. (2006). ER stress and cancer. Cancer Biol. Ther. 5, 721–722. Li, J., and Lee, A. S. (2006). Stress induction of GRP78/BiP and its role in cancer. Curr. Mol. Med. 6, 45–54. Lin, Y., Wang, Z., Liu, L., and Chen, L. (2010). Akt is the downstream target of GRP78 in mediating cisplatin resistance in ER stress-tolerant human lung cancer cells. Lung Cancer [Epub ahead of print]. Malhotra, D., Thimmulappa, R., Vij, N., Navas-Acien, A., Sussan, T., Merali, S., Zhang, L., Kelsen, S. G., Myers, A., Wise, R., Tuder, R., and Biswal, S. (2009). Heightened endoplasmic reticulum stress in the lungs of patients with chronic obstructive pulmonary disease: The role of Nrf2-regulated proteasomal activity. Am. J. Respir. Crit. Care Med. 180, 1196–1207. Maunders, H., Patwardhan, S., Phillips, J., Clack, A., and Richter, A. (2007). Human bronchial epithelial cell transcriptome: Gene expression changes following acute exposure to whole cigarette smoke in vitro. Am. J. Physiol. Lung Cell Mol. Physiol. 292, L1248–L1256. Novoa, I., Zeng, H., Harding, H. P., and Ron, D. (2001). Feedback inhibition of the unfolded protein response by GADD34-mediated dephosphorylation of eIF2alpha. J. Cell Biol. 153, 1011–1022. Powzarowsky, P., Holden, E., and Darzynkiewicz, Z. (2006). Laser scanning cytometry: Principles and applications. Methods Mol. Biol. 319, 165–192. Ron, D., and Walter, P. (2007). Signal integration in the endoplasmic reticulum unfolded protein response. Nat. Rev. Mol. Cell Biol. 8, 519–529. Shu, C. W., Sun, F. C., Cho, J. H., Lin, C. C., Liu, P. F., Chen, P. Y., Chang, M. D., Fu, H. W., and Lai, Y. K. (2007). GRP78 and Raf-1 cooperatively confer resistance to endoplasmic reticulum stress-induced apoptosis. J. Cell. Physiol. 215, 627–635. Sridhar, S., Schembri, F., Zeskind, J., Shah, V., Gustafson, A. M., Steiling, K., Liu, G., Dumas, Y. M., Zhang, X., Brody, J. S., Lenburg, M. E., and Spira, A. (2008). Smokinginduced gene expression changes in the bronchial airway are reflected in nasal and buccal epithelium. BMC Genomics 9, 259–272. Steiling, K., Kadar, A. Y., Bergerat, A., Flanigon, J., Sridhar, S., Shah, V., Ahmad, Q. R., Brody, J. S., Lenburg, M. E., Steffen, M., and Spira, A. (2009). Comparison of proteomic and transcriptomic profiles in the bronchial airway epithelium of current and never smokers. PLoS ONE 4, e5043. Szegezdi, E., Logue, S. E., Gorman, A. M., and Samali, A. (2006). Mediators of endoplasmic reticulum stress-induced apoptosis. EMBO Rep. 7, 880–885. Wang, Y., Wang, W., Wang, S., Wang, J., Shao, S., and Wang, Q. (2008). Downregulation of GRP78 is associated with the sensitivity of chemotherapy to VP-16 in small cell lung cancer NCI-H446 cells. BMC Cancer 8, 372. Watterson, T. L., Hamilton, B., Martin, R., and Coulombe, R. A., Jr. (2009). Urban particulate matter causes ER stress and the unfolded protein response in human lung cells. Toxicol. Sci. 112, 111–122. Wek, R. C., Jiang, H. Y., and Anthony, T. G. (2006). Coping with stress: eIF2 kinases and translational control. Biochem. Soc. Trans. 34, 7–11. Witschi, H. (2005). Carcinogenic activity of cigarette smoke gas phase and its modulation by beta-carotene and N-acetylcysteine. Toxicol. Sci. 84, 81–87.

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C H A P T E R

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Induction and Measurement of UPR and Osmotic Stress in the Yeast Pichia pastoris Martin Dragosits,*,1 Diethard Mattanovich,*,† and Brigitte Gasser*,† Contents 166 169 171 174 179 179 182 183 186 186

1. Introduction 2. Measuring UPR Induction in P. pastoris 2.1. Inducing UPR in P. pastoris 2.2. Measuring UPR 3. Inducing and Measuring Osmotic Stress in P. pastoris 3.1. Induction of osmotic stress 3.2. Measuring intracellular osmolytes 3.3. Measuring UPR targets Acknowledgments References

Abstract Unfolded protein response (UPR) is a major reaction to intrinsic stress of eukaryotic organisms and is also related to environmental stress reactions. Among yeasts, stress regulation has mainly been investigated in Saccharomyces cerevisiae, while other species with biotechnological or medical interest are less well understood. Pichia pastoris as one example has emerged as a favorite production platform for recombinant proteins during the last two decades. UPR and environmental stress are well known to interfere with the production of recombinant proteins as well as other technologically relevant processes, so that the demand for well-documented protocols to measure such stress reactions has strongly increased. Here, we describe protocols for the induction of UPR and osmotic stress, as well as for the quantitative measurement of cellular stress reactions at the levels of transcripts, proteins, and * Department of Biotechnology, University of Natural Resources and Life Sciences, Vienna, Muthgasse 18, Vienna, Austria { Austrian Centre of Industrial Biotechnology (ACIB GmbH), Muthgasse 18, Vienna, Austria 1 Current address: UC Davis Genome Center, University of California Davis, One Shields Avenue, Davis, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00010-8

#


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metabolites. As such protocols need to be adapted for a new species of interest, the guidelines presented here should enable researchers to study P. pastoris directly without the hassle to modify standard protocols designed for the model organism S. cerevisiae first.

1. Introduction Stress reactions are essential for microorganisms to cope with changes of environmental conditions. Whereas stress reactions have been studied in detail in all kingdoms of life, in yeasts, the best analyzed species is Saccharomyces cerevisiae (for a recent review, see Shima and Takagi, 2009). A deeper understanding of microbial stress response mechanisms is not only important for improved understanding of the general physiology of living cells but also for biotechnological processes involving growing microbial cultures (Mattanovich et al., 2004). Additionally, stress-response mechanisms are of great importance regarding food conservation and food safety issues (Smits and Brul, 2005). Microorganisms are capable of responding to environmental stress stimuli by certain sensor and signaling mechanisms that effectively trigger cellular responses and adapt gene expression patterns in order to maintain growth under new conditions. Bacterial as well as eukaryotic cells such as yeast possess various response mechanisms to counterbalance fluctuations in growth parameters such as temperature, pH, solute concentration, oxygenation, metal ion concentration, or high levels of organic compounds. Although specific stresses result in stress-specific responses, it has been shown that overlaps between the different response pathways exist (e.g., Piper, 1995). Early evidence for these overlaps was acquired in the model bacterium Escherichia coli in the late 1980s of the last century. Stress survival experiments showed that exposure to a certain environmental stress can result in a cross-protection effect when cells were subsequently exposed to a second growth-inhibiting stress condition ( Jenkins et al., 1988, 1990). With the rise of DNA microarrays to a powerful tool for whole transcriptome profiling striking evidence soon came also for other species such as yeasts. Profiling of changes in global gene transcription revealed overlapping responses of stress signaling pathways to different stimuli (Causton et al., 2001; Gasch et al., 2000). The unfolded protein response (UPR) is a regulatory circuit of high relevance for heterologous protein production in eukaryotic cells. As Pichia pastoris is one of the most commonly used yeast hosts for recombinant protein production purposes, this chapter aims at the elucidation of interconnected stress in this yeast rather than in the model yeast S. cerevisiae. Nevertheless, many processes have only been studied in this yeast, so that

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literature data usually refers to S. cerevisiae, but most approaches can also be used for other yeasts with only slight modifications. Yeasts respond to endoplasmic reticulum (ER) stress conditions with the activation of the conserved UPR pathway. The UPR can be activated by a variety of conditions that cause ER stress. In S. cerevisiae, there are three main components (reviewed by Patil and Walter, 2001): Ire1p, a transmembrane protein kinase with endonuclease activity, Hac1pi, which is the transcription factor responsible for the initiation of transcription of genes underlying the UPR-regulation, and the tRNA ligase, which is encoded by the RLG1 gene. Briefly, accumulation of unfolded protein within the ER leads to activation Ire1p, which initiates unconventional splicing of the HAC1u mRNA and enables translation of induced form HAC1i. The UPR transcription factor Hac1p then binds to UPRE (UPR elements) within promoters of its target genes and regulates transcription of several hundred genes (see Travers et al., 2000 for S. cerevisiae; Graf et al., 2008 for P. pastoris; Wimalasena et al., 2008 for Candida albicans). Knowledge of genome-wide transcriptional patterns under UPR conditions is a very important prerequisite when choosing target genes to monitor ER stress response. While S. cerevisiae is often used as a model for other yeasts as well, it is known that some yeasts use partially different patterns of stress regulation. P. pastoris, widely used for recombinant protein production, has been characterized for stress regulation during the last few years (Carnicer et al., 2009; Dragosits et al., 2009, 2010; Graf et al., 2008; Guerfal et al., 2010; Hohenblum et al., 2004; Hyka et al., 2010; Sauer et al., 2004; Yano et al., 2009). Osmotic stress is caused by fluctuations in extracellular solute concentrations. Microorganisms encounter adversarial osmotic conditions in both their natural habitats and in biotechnological processes. Generally, hypoosmotic conditions (low solute concentrations) and hyperosmotic (high solute concentrations) can pose a stress condition to a microbial cell as imbalanced extracellular solute concentration may interfere with cellular water availability, ion homeostasis, and turgor pressure regulation. In biotechnological processes, hyperosmotic stress is more relevant, as in high cell density bioreactor cultivations, media initially contain high concentrations of major nutrients (e.g., carbon sources such as glucose) and salts. Additionally, during the production process, microbial cells may release metabolic (by)products which also add to increased osmotic pressure. Osmotically active compounds may either be ionic (dissociated salts or organic acids) or uncharged (e.g., sugars, sugar alcohols, or undissociated organic acids). Although the response to uncharged and charged osmolytes is widely overlapping, ionic charges such as dissociated salts may trigger additional responses when compared with uncharged solutes. This chapter aims at the impact of nutrient salt concentration on cellular stress response,

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including the response to unfolded protein among others. Therefore, KCl as a charged osmolyte is applied here. Regarding the response of yeast to hyperosmotic pressure, S. cerevisiae is the best studied example. A sudden increase of extracellular osmolarity leads to a temporary growth arrest and subsequently, gene transcription is readjusted in order to resume growth under high osmolarity (Norbeck and Blomberg, 1997). Depending on the severity of the osmotic shock, either the environmental stress response (ESR) or the high osmolarity glycerol (HOG) pathway or both are induced (Gasch et al., 2000; O’Rourke and Herskowitz, 2004). The HOG pathway is a very well-studied signaling pathway in yeast (Hohmann, 2002). The HOG MAP kinase can be activated through two distinct pathways, involving the transmembrane osmosensors, Sln1 and Sho1, both activating Hog1 by distinct phosphorelay systems (Posas and Saito, 1997). In S. cerevisiae among the multitude of cellular readjustments, the upregulation of glycerol-3-phosphate dehydrogenase GPD1 (Larsson et al., 1993) transcription and the repression of the plasma membrane glycerol efflux channel FPS1 (Mager and Siderius, 2002) transcription are of particular importance. In response to hyperosmotic shock, yeasts start production of compatible solutes such as glycerol to cope with osmotic stress. However, different yeast species may produce and accumulate or release different compatible solutes upon osmotic stress (Kayingo et al., 2001). Thus, it is important not only to measure glycerol concentrations but also to include other polyols in poorly characterized yeast species. Recently, evidence of the involvement of the HOG pathway in the response to ER stress conditions has been provided, leading to transcriptional induction of GPD1 and glycerol accumulation upon treatment of S. cerevisiae with tunicamycin (Torres-Quiroz et al., 2010). However, a direct interaction between the UPR and the HOG pathway has been ruled out in this study, as HOG1 deletion had no effect on UPR activity. Nevertheless, a protective effect of the compatible solute glycerol occurs. In another study, Hog1 has been reported to regulate gene expression during persistent ER stress conditions after the initial UPR response in S. cerevisiae (Bicknell et al., 2010). While so far there is no evidence that UPR triggers an osmo-response such as production of compatible solutes (see Fig. 10.1) or confers resistance to salt (Fig. 10.2), adaptation to hyperosmotic conditions has been shown to activate an UPR-like response including upregulation of chaperones in P. pastoris. In strains already having UPR activation due to recombinant protein overproduction, the effect of elevated osmolarity is less pronounced (Dragosits et al., 2010). Methods to analyze responses to ER stress and salt stress conditions, as well as potential interaction between these stress responses in P. pastoris, are described below.

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12

0

200 180 160 140 120 100 80 60 40 20 0

7

18

6

15

Arabitol (mg g–1)

Glycerol (mg g–1)

10 8 6 4

Trehalose (mg g–1)

Mannitol (mg g–1)

2

5 4 3 2 1

12 9 6 3 0

0 X-33 low

Fab low

X-33 Fab Medium Medium

X-33 high

Fab high

X-33 low

Fab low

X-33 Fab X-33 Medium Medium high

Fab high

Figure 10.1 Intracellular mannitol, glycerol, arabitol, and trehalose levels during carbon-limited steady-state cultivation of P. pastoris at different osmolarities. Low 140 mOs/kg, medium 860 mOs/kg, and high 1350 mOs/kg; all osmolarities adjusted with KCl. Osmolytes are given in mg/g yeast dry mass. X-33—P. pastoris wild-type strain. Fab—P. pastoris strain expressing extracellular Fab 3H6 and UPR induced. Error bars represent the standard error of the mean of three biological replica.

2. Measuring UPR Induction in P. pastoris To study the UPR in yeasts, researchers have mainly applied chemical treatment to interfere with oxidative protein folding or protein glycosylation in the ER. Treatment with the reducing agent dithiothreitol (DTT) triggers not only strong UPR, but also other stress responses such as those to general chemical stress and response to a disturbed redox homeostasis. Tunicamycin inhibits the initial step in the synthesis of N-glycans, thus preventing protein glycosylation and further protein processing and secretion, so that UPR is induced. Additionally, tunicamycin leads to a cell cycle arrest and activation of the HOG pathway in S. cerevisiae, but a thorough analysis of other responses to tunicamycin is still missing. Therefore, we suggest alternatives to simulate the UPR in P. pastoris, as well as in other yeasts in a more natural way. This may either be the overexpression of recombinant proteins with slow folding kinetics (Gasser et al., 2006; Hohenblum et al., 2004), or the constitutive stimulation of UPR by overexpression of the active form of the HAC1 gene (Graf et al., 2008). Interestingly, both the heterologous Hac1pi of S. cerevisiae and the homologous Hac1pi of P. pastoris (Guerfal et al., 2010) induce UPR in this yeast.

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YPD A B C KCl

NaCl

0.6 M

1.2 M

1.4 M

1.6 M

A

A

B

B

C

C

A

A

B

B

C

C

A

A

B

B

C

C

A

A

B

B

C

C

Figure 10.2 Growth of S. cerevisiae HA232 (A), P. pastoris X-33 (B), and P. pastoris overexpressing HAC1i (C) on YPD agar with different concentrations of NaCl or KCl. YPD agar plates and plates containing 0.6 M NaCl or KCl were incubated at 28 C for 4 days, whereas all other plates were incubated at 28 C for 6 days.

For comparative purposes, chemical induction of UPR by DTT and tunicamycin as well as overexpression of S. cerevisiae HAC1i are described in this chapter, followed by a presentation of methods to measure UPR induction on transcript and protein level, specifically focussing on few known UPR targets.


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2.1. Inducing UPR in P. pastoris Based on our experience, chemical induction of UPR suffers the drawback that other regulatory circuits are activated as well (see regulation patterns in Table 10.1), so that we recommend HAC1i overexpression as the most authentic approach. As most data on UPR in yeast have been accumulated with DTT or tunicamycin treatment, these methods are described here as well for P. pastoris. 2.1.1. Required materials Strains and vectors P. pastoris strains, for example, X-33, SMD1168 (his4 pep4), or GS115 (his4) (all Invitrogen) Vector for overexpression of HAC1, for example, pGAPHis To construct the induced variant of the HAC1 gene of S. cerevisiae, it is necessary to combine the DNA fragment coding for the first 220 amino acids with the fragment coding for the 18-amino acid exon of the induced Hac1p (Mori et al., 2000) in a two-step PCR reaction (using the primers HACfor SfiI and HACback1 in the first step, and HACfor SfiI and

Table 10.1 Specific regulation of GO functional groups upon constitutive induction of the UPR by HAC1 overexpression, or treatment of P. pastoris with DTT

Both UP Protein folding Vesicular transport Conjugation

Hac1 UP Ribosome biogenesis RNA metabolism Translation Organelle biosynthesis DTT UP Chemical stimulus Cellular homeostasis

Both DOWN Carbohydrate metabolism Amino acid biosynthesis Lipid metabolism Vitamin biosynthesis Metabolism of cofactors Metabolism of heterocyclic and aromatic compounds Hac1 DOWN Membrane biogenesis Cell wall biogenesis Cytoskeleton biogenesis Stress response DTT DOWN Electron transport Respiration Nuclear biogenesis

UP and DOWN means up- and downregulation of the respective groups (data adapted from microarray analysis in Graf et al., 2008).

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HACback2 SacII in the second step, primer sequences are given in Table 10.2), and subsequent ligation of the resulting fragment into pGAPHis. Alternatively, HAC1 can be amplified from P. pastoris mRNA as described by Guerfal et al. (2010). Media

YPD medium (per liter: 10 g yeast extract, 20 g peptone, 20 g glucose); main culture medium (per liter: 10 g yeast extract, 20 g peptone, 100 mM potassium phosphate buffer, pH 6.0, 13.4 g yeast nitrogen base with ammonium sulfate, 0.4 mg biotin) Chemicals

Tunicamycin stock (5 mg/mL in AD); DTT stock (1 M in AD)

2.1.2. Induction of UPR by chemical treatment Inoculate a P. pastoris strain (e.g., X-33) in a 5-mL YPD and shake overnight at 25–28 C and 180 rpm. For each experiment, at least two separate cultures are needed (as biological replica); additionally untreated cultures serve as controls. After measuring optical density at 600 nm (OD600), inoculate 10 mL of main culture medium with an OD600 of 1.0 and grow the culture for approximately 5 h to reach an exponential growth phase (OD600 should be around 5–7), before adding either DTT or tunicamycin to induce the UPR. The control cultures remain untreated. Cultivate as before for another 1–2 h. Then, immediately take up the cells in the appropriate buffer or extraction reagent, as described below, and record the final OD600. Common concentrations for UPR induction with DTT in different yeasts range from 2 to 20 mM. For P. pastoris, addition of 2.5 mM DTT for 1 h was found to be sufficient to induce UPR (Graf et al., 2008) without severely inhibiting growth. Tunicamycin is usually added at a final concentration of 0.5–5 mg/mL. For P. pastoris, we recommend 5 mg/mL for at least 1 h. 2.1.3. Induction of UPR by overexpression of HAC1 Inoculate a P. pastoris strain overexpressing HAC1i under control of a constitutive promoter (e.g., PGAP) and a respective control strain in a 5-mL YPD, and shake overnight at 25–28 C and 180 rpm. For each experiment, at least two separate cultures are needed (as biological replica). After measuring OD600, inoculate 10 mL of main culture medium with an OD600 of 1.0 and grow the culture for approximately 6 h to reach an exponential growth phase (conditions as described above). Then, immediately take up the cells in the appropriate buffer or extraction reagent, as described below, and record the final OD600 (should be around OD 5–7).

Table 10.2 Primers for qRT-PCR and HAC1 cloning Target gene

Oligo name

ACT1

ACT1 fw ACT1 bw PDI1 PDI1 fw PDI1 bw HAC1 HAC1 fw HAC1 bw ERO1 ERO1 fw ERO1 bw ScHAC1i Hac for SfiI amplification Hac back2 SacII Hac back1

Sequence (50 –30 )

Amplicon size

GC content

CCTGAGGCTTTGTTCCACCCATCT 148 45.9% GGAACATAGTAGTACCACCGGACATAACGA GGAAAGGCCCACGATGAAGTTGTC 140 46.4% GCATCCTCATCATTGGCGTAAAGAGTAG GCGGCCCATGCTTCCAGAGAG 163 63.6% CGGTACCACCTAAGGCTTCCAACC GTTGGAAAAGCCGCATATAAACAAAACA 141 36.9% CAGCTTGGGCAAAGTCCTGTAAGAGTTC GCAAGGCCCAGCCGGCCACGATGGAAATGACTGATTTTGAAC CCTCCCGCGGTCATGAAGTGATGAAGAAATCATTCAATTCAAATGAATTCA AACCTGACTGCGCTTCTGGATTACGCCAATTGTCAAG TGG TCA TCG TAA TCA CGG C

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Alternatively, the strains can also be cultivated in the main culture medium containing methanol as a carbon source instead of glucose. However, in the case of methanol as carbon source, cells will need to be cultivated for at least 12 h before samples from the exponential growth phase can be taken.

2.2. Measuring UPR Activation of the UPR can be detected on the level of (i) gene expression by either monitoring single genes, (ii) protein abundance using immunofluorescent staining and visualization or immunoblotting methods, or (iii) through the expression of reporter genes fused to UPRE/Hac1-induced promoters. As an alternative to the methods outlined above, genome scale methods measuring the transcriptome or proteome may provide a broader picture of UPR specific and other stress reactions. For this purpose, transcriptomics by DNA microarrays and differential proteomics using 2D gel electrophoresis can be performed as described in Dragosits et al. (2010). 2.2.1. Measuring UPR by analyzing gene expression levels In yeasts, UPR is usually detected by measuring specific gene products that are induced directly by the activated Hac1 protein. These are typically ER-resident folding catalysts like PDI1, ERO1, and chaperones like BiP (encoded by KAR2). Other valid targets identified by DNA microarray analysis are components of the glycosylation machinery such as PMT genes, or genes coding for proteins involved in the secretory transport (e.g., certain SEC genes). As the transcription of P. pastoris HAC1 mRNA is also induced upon UPR activation, this gene should also be included in the analysis. In contrast to S. cerevisiae and other yeasts such as Yarrowia lipolytica (Oh et al., 2010) or C. albicans (Wimalasena et al., 2008), in P. pastoris, HAC1 mRNA always seems to be present in the spliced form (Guerfal et al., 2010). Therefore, monitoring of the HAC1 splicing event, which is commonly performed as a measure for UPR induction in other yeasts, is not possible in P. pastoris. Induction of transcription of these target genes is usually determined by quantitative real-time PCR (qRT-PCR), as described below. Alternative methods like transcriptional profiling with the aid of affinity capture (TRAC; Rautio et al., 2006) or Northern blotting may be employed as well. For a description of TRAC applied for P. pastoris, the reader is referred to Gasser et al. (2007), while Resina et al. (2007) uses a bead-based sandwich hybridization assay to study induction of UPR during recombinant protein production in P. pastoris. qRT-PCR is suitable to quantify relative amounts of specific transcripts. Usually, a constitutive gene like actin is used for standardization. As the transcription rates of all genes, including actin or other constitutive standards depend on cell growth, it is important that cells


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are harvested at comparable growth conditions (mid-exponentional phase, as described in Section 2.1) or constant growth rates (e.g., chemostat cultures, as described below). 2.2.1.1. Required materials

Cultures of P. pastoris as described in Section 2.1 5% (v/v) phenol in ethanol (cooled on ice) Microcentrifuge tubes with sealed caps Acid-washed glass beads (0.50 mm diameter) Fast Prep cell homogenizer (Thermo Savant) RNAse-free microcentrifuge tubes and tips TRI (Trizol) reagent (e.g., from Ambion, Invitrogen, Sigma) Chloroform, Isopropanol, Ethanol, RNAse free water DNAfree (Ambion) Reverse transcriptase (e.g., Superscript III cDNA synthesis kit (Invitrogen)) SensiMix Plus PCR premix (GenXpress) Real-time PCR machine (e.g., iCycler Thermal Cycler (Bio-Rad), Rotorgene 6000 (Corbett Life Sciences)) and respective real-time PCR tubes Primers designed for qRT-PCR for target genes and internal reference gene (here actin)—the amplicon should be at the 30 end of the gene, with a length of around 150 bp and a GC content between 40% and 60%, the melting temperature of the primers has to be approximately 60 C, otherwise follow general guidelines for PCR primer design.

Sampling For subsequent RNA isolation, the metabolic activity of the cultures has to be immediately quenched after sampling. Therefore, 6 mL of cell culture is taken up in 3 mL of ice-cold 5% (v/v) phenol in ethanol (purity > 95%). Mix by inverting the tube and keep on ice. Aliquote at 2 mL each in fresh microcentrifuge tubes with sealed caps and centrifuge for 2 min at 0–4 C and full speed (13,000 rpm). Discard the supernatant and store the pellet at 80 C until further use. RNA extraction For total RNA extraction, cells need to be disrupted. As P. pastoris cells are rather hard to break, mechanical cell disruption is recommended. Take one tube per experiment and resuspend the pellet in 1 mL of TRI reagent. Add 0.5 mL of glass beads and seal the tube tightly. Cells are disrupted in a FastPrep cell homogenizer (Thermo Savant) using 6 m s 1 for 2 30 s and chilling the cells on ice in between. After 10 min incubation at RT, total RNA is isolated according to the protocol supplied with TRI reagent by the manufacturer. Briefly, to extract RNA, 200 mL chloroform is added to the tube and the tube is vigorously shaken and incubated for approximately

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15 min at room temperature. After centrifugation at full speed and 4 C, the sample is separated in three distinct phases: A colorless upper aqueous phase—containing desired RNA, an interphase—containing DNA, and a red organic phase—containing protein. The aqueous phase is transferred to a fresh, RNase-free microcentrifuge tube and 500-mL isopropanol are added to precipitate the RNA. After 10 min of incubation at RT, the sample is centrifuged again at full speed and 4 C. After this step, the RNA precipitate should be visible as a pellet on the side or bottom of the tube. The supernatant is removed and the pellet washed in 1 mL of 75% ethanol. The final step is the solubilization of the air-dried RNA pellet in an appropriate volume (50 mL) of RNase free water and incubation at 55 C for about 10 min. To eliminate remaining DNA, samples are treated with DNAfree (Ambion) following the manufacturer’s instructions. Extracted RNA is quantified by measuring absorbance at 230/260/280 nm (e.g., on a ND-1000 (Nanodrop) or any other suitable spectrophotometer), while RNA integrity is checked using the Agilent Bioanalyzer 2100 and the RNA Nano 6000 Assay Kit (Agilent). Alternatively, conventional agarose gels can be run to verify RNA quality. cDNA synthesis 2.5 mg of total RNA is reverse-transcribed in a 20-mL reaction volume using the Superscript III (Invitrogen, USA) reagent set. After inactivation of the enzyme by heating to 70 C for 15 min, template RNA is removed by treating the reaction with 1-mL RNase H at 37 C for 20 min. The obtained cDNA is then purified with a PCR-Purification Kit and dissolved in 50-mL 10-mM Tris/Cl with pH 8.0. The quantity of cDNA is determined on a ND-1000 (Nanodrop). qRT-PCR analysis Quantitative real-time PCR is carried out using the dsDNA intercalating dye SYBR Green for the detection of amplified DNA on a Rotorgene 6000. Samples are measured in triplicates. In 10-mL reactions, 1 mL of cDNA is applied with SensiMix Plus PCR premix and the respective primers. A nontemplate control should be run in every experiment for each of the primer pairs to avoid detection of unspecific priming. The reactions are incubated at 95 C for 10 min to activate the Taq polymerase and then subjected to a three-step cycling protocol including melting (94 C, 15 s), annealing (60 C, 20 s), and extension (72 C, 15 s) for a total of 45 cycles. The acquisition of the signal is detected during the annealing step in each cycle. After a final extension, a melting curve is generated by data collection during 70 cycles starting at 65 C, with 0.5 C increase/cycle (2 s intervals). A correct melting curve consists only of one distinct peak, additional peaks are an indication for primer-dimers or unspecific PCR products. When using a set of primers for the first time,


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some PCR reactions should be checked by running them on a conventional 1% agarose gel. Computational data analysis and interpretation Data analysis can be performed, for example, via the Rotorgene Software package and Microsoft Excel. ACT1 is suggested here as reference to determine relative mRNA levels of the other genes. 2.2.2. Measuring UPR by analyzing abundance of target proteins BiP and other ER-resident proteins can be detected immunochemically by antiserum against the ER retention peptide HDEL, which also allows the quantification in single cells by immunofluorescence. Also, specific antibodies against UPR target proteins such as Pdi are available for S. cerevisiae. Here, we describe immunoblotting of P. pastoris cell lyzates, as well as monitoring of UPR signaling by flow cytometric analysis of ethanol-fixed cells using anti-HDEL antibody. 2.2.2.1. Immunoblotting of cell lyzates Required materials

Cultures of P. pastoris as described in Section 2.1 Acid-washed glass beads (0.50 mm diameter) Fast Prep cell homogenizer (e.g., MP Fastprep-24) Disintegration buffer (PBS containing protease inhibitor cocktail (Sigma8820) and 1% Triton X-100) Equipment needed for SDS–PAGE and Western blotting Blocking buffer (PBS, pH 7.4, containing 0.1% Tween and 2% skimmed milk powder or BSA) Washing buffer solution (PBS, pH 7.4, containing 0.1% Tween) Antibodies, for example, rabbit-anti-BiP (StressGen) and anti-rabbit AP- or HRP-conjugate as secondary antibody; or mouse monoclonal HDEL (2E7):sc-53472 (Santa Cruz Biotechnology) and suitable secondary antibody such as antimouse AP- or HRP-conjugate Detection reagent for Western blot (e.g., AP substrate Kit for AP-conjugates and colorimetric detection; ECL Kit for HRP-conjugate and chemiluminescent detection)

Mechanical disintegration of P. pastoris cells Two milliliters of P. pastoris cultures are harvested by centrifugation in microcentrifuge tubes with sealed caps, the resulting pellet is resuspended in a 500-mL disintegration buffer, and the same volume of glass beads is added. Cells are disrupted in a Fastprep for 3 20 s at 6 m s 1, with cooling on ice in between. Add 2 % SDS and denature proteins by heating to 99 C for 10 min and then centrifuge for 15 min at 4 C at full speed. Transfer the liquid phase into a fresh tube. Cell lyzates can be stored at 20 C.

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Immunostaining Perform separation of protein by SDS–PAGE and subsequent Western blotting onto a nitrocellulose membrane. Slightly shake the blot in a blocking buffer for at least 2 h and then incubate the blot with an antiHDEL antibody (1:1000 in a blocking buffer) for 1 h at RT. After three washing steps with a washing buffer (5–10 min each), incubate the blot with a secondary antibody (in the blocking buffer, dilution as recommended by the supplier) for 1 h at RT. Wash the blot again to remove the unbound antibody and then stain the blot with the appropriate reagent to visualize the protein bands. The size of BiP is approximately 78 kDa and other possible bands are ER-hsp40 chaperones (ca. 40 kDa), Pdi1 (ca. 50 kDa), or Lhs1 (ca. 100 kDa). 2.2.2.2. Flow cytometry analysis of cells stained by immunofluorescence Required material

70% ethanol Washing buffer (100-mM Tris/HCl buffer or PBS, pH 7.4, containing 0.1% Triton X-100 and 2-mM MgCl2) Blocking buffer (washing buffer with 2% BSA) Fixation of cells in 70% ethanol To make the cell walls permissible for proteins such as antibodies, samples of cells intended for immunofluorescent intracellular staining have to be fixed in 70% ethanol. Approximately 107 cells are taken up in a 500-mL PBS and centrifuged at 6000 rpm for 5 min at room temperature; the supernatant is then discarded. The pellets are resuspended in 1-mL 70% ethanol while vortexing and stored at –20 C immediately. Intracellular staining ER-resident proteins such as BiP Fifty microliters of ethanol-fixed cell solution (corresponding to an OD of approximately 0.2) are added to 1 mL of washing buffer and then the cells are harvested by centrifugation at full speed (13,000 rpm) and RT for 5 min. The pellet is resuspended in a 1-mL blocking buffer and incubated for 30 min at RT with shaking. After centrifugation, the pellet is incubated in 200 mL of the primary antibody solution (rabbit-anti-BiP (StressGen) or HDEL (2E7): sc-53472 (Santa Cruz Biotechnology); 1:100 in blocking buffer) for 1 h at RT with shaking. Afterward, the cells are centrifuged again and then washed in 1 mL of washing buffer. The resulting pellet is then resuspended in a 200-mL secondary antibody solution (either goat-anti-rabbit IgG FITC-conjugate or anti-mouse IgG (Fab specific) FITC-conjugate; 1:100 in blocking buffer) for 1 h at RT with shaking. Then, the cells are pelleted by centrifugation and the redundant antibodies are washed away with 1 mL of washing buffer. Finally, the cells are resuspended in 200 mL of PBS.


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Flow cytometer—analysis The stained cells are analyzed on a FACS Calibur (Becton Dickinson) with a 488-nm argon laser and a 630-nm diode laser. For each analysis, 10,000 cells of a sample are measured through a 530/30 BP filter (FL1) in logarithmic mode. Cell debris has to be excluded from the data by adjusting the thresholds appropriately. The immunofluorescent data are then evaluated by calculation of the geometric mean of each immunofluorescent signal and related to the cell size according to Eq. (10.1), where relFL is the relative fluorescence signal, FL1 is the measured fluorescence signal, and FSC is the forward scatter, a signal proportional to the cross-sectional area of the cells. FL1 relFL ¼ pffiffiffiffiffiffiffiffiffi3 FSC

ð10:1Þ

3. Inducing and Measuring Osmotic Stress in P. pastoris 3.1. Induction of osmotic stress The aim of a study defines the correct choice of an osmolyte to be added to a microbial culture. Different culture types based on suspension culture or surface culture on solidified (e.g., agar) media can be applied for studying osmotic stress response. Liquid suspension cultures are best suited to prepare homogenous cell samples for biochemical and physiological analysis of stress response. Chemostat cultures are typically used to obtain cells that are in steady state, which ensures highest possible homogeneity and reproducibility. Thus, chemostat cultivation at different osmolarities is described in this chapter. One might also be interested in the immediate response to increased osmolarity. Thus, the aim of the study greatly determines the mode of cultivation. In this case, we would suggest performing shake flask or chemostat cultivations where cells are shock-treated with high or low osmolyte concentration and samples are taken in a range of 5 min to 2 h after shock application. The data presented here as examples were obtained in a study where we were interested in the response of adapted cells; therefore, we took samples at time points where cells had already fully adapted their metabolism to high osmotic conditions (five residence times, see Section 3.1.3). In the case the aim of the study or experiment is to analyze differences in osmotolerance between strains or species, growth tests on solid media such as agar plates represent a convenient method. Such comparisons may also be performed in shake flask cultures with OD600 readings made in regular intervals; however, this might be more time consuming

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than tests on solid media. The data that were obtained by such growth tests on solid media and are presented here will show differences in osmotolerance (caused by the ionic osmolytes NaCl and KCl) between P. pastoris and S. cerevisiae. 3.1.1. Required materials Strains, plates, and chemostat media

P. pastoris X-33 and S. cerevisiae HA232 or any other suitable strains YPD medium (2% (w/v) peptone, 1% (w/v) yeast extract, 2% (w/v) glucose) YPG medium (2% (w/v) peptone, 1% (w/v) yeast extract, 1% (w/v) glycerol) YPD agar plates (2% (w/v) peptone, 1% (w/v) yeast extract, 1% (w/v) agar, 2% (w/v) glucose) containing 0, 0.6, 1.2, 1.4, and 1.6 M NaCl or KCl. Batch medium (amounts per liter): 39.9 g glycerol, 1.8 g citric acid, 12.6 g (NH4)2HPO4, 0.022 g CaCl2 2H2O, 0.9 g KCl, 0.5 g MgSO4 7H2O, 2-mL Biotin (0.2 g L 1), and 4.6-mL trace salts stock solution. The pH was set to 5.0 with 25% (w/w) HCl. Chemostat medium (amounts per liter): 50 g glucose 1H2O, 0.9 g citric acid, 4.35 g (NH4)2HPO4, 0.01 g CaCl2 2H2O, 1.7, or 29.9, or 48.5 g KCl, 0.65 g MgSO4 7H2O, 1 mL biotin (0.2 g L 1), and 1.6 mL trace salts stock solution. The pH was set to 5.0 with 25% (w/w) HCl. Trace salts stock solution (amounts per liter): 6.0 g CuSO4 5H2O, 0.08 g NaI, 3.0 g MnSO4 H2O, 0.2 g Na2MoO4 2H2O, 0.02 g H3BO3, 0.5 g CoCl2, 20.0 g ZnCl2, 5.0 g FeSO4 7H2O, and 5.0 mL H2SO4 (95–98%, w/w). 250–1000 mL baffled Erlenmeyer flasks A 2–3 L benchtop bioreactor with equipment for continuous cultivations

3.1.2. Plate growth assay Grow a starter culture of the strains to be tested in YPD medium at 28 C on a shaker at 170 rpm overnight. On the next day, dilute each of these cultures to an OD of 0.1 in sterile PBS and then prepare 4 serial 1:10 dilutions in sterile PBS. Then, spot 3 mL of each dilution onto YPD agar plates with different concentrations of the osmotically active substances that are to be tested. Incubate the plates for several days at 28 C. Whereas growth at low concentrations of solutes may already be observed after 24 h of growth, higher concentrations may result in largely reduced but still relevant growth. Therefore, it is important to monitor growth for at least up to 5 days. To prevent desiccation of agar plates due to prolonged incubation at 28 C, plates might be sealed with parafilm or stored in a plastic bag.


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3.1.3. Cultivation in chemostat In order to maintain cells in steady state, chemostat cultures in a bioreactor are best suited. Working volumes around 1 L allow for precise control of the dilution rate and frequent sample removal while avoiding too high volumes of required media. Here, a 3.5-L bioreactor is used at a working volume of 1.5 L. On the day before inoculating the bioreactor, prepare a starter culture in a 1-L shake flask containing a 150-mL YPG medium with a 1-mL cryostock of the respective P. pastoris clone and grow this culture at 28 C, shaking at 170 rpm for approximately 24 h. Meanwhile, prepare the bioreactor with a 1.5-L sterile batch medium and inoculate it with the starter culture to a calculated optical density (OD600) of 1.0. Control pH at 5.0 with 25% ammonium hydroxide (w/w) and dissolved oxygen at 20% by controlling the stirrer speed. Gas flow rate is kept constant at 1.5 vvm (volume gas per volume medium and minute). The batch phase lasts for approximately 24 h, and its end is indicated by a sudden increase in dissolved oxygen (as no more carbon source is available for consumption). When this point is reached, start the continuous culture by initiating the feed of chemostat medium at a dilution rate of D ¼ 0.1 h 1 (growth medium and waste efflux flow rates of 150 g/h). As a convention, five residence times of continuous cultivation are performed before assuming steady state and taking samples (residence time refers to the time that is necessary to feed the total cultivation volume once). In our case one residence time is 10 h as at a total volume of 1.5 L (we feed 10 150 g (mL) in this time). When changing the chemostat medium (using media with different osmolarities) between sampling points, it is advised to wait for more than five residence times, for example 8, in order to compensate for the continuous dilution of the medium. For each sample, determine dry biomass concentration by washing duplicates of 10-mL chemostat culture in distilled water and drying the cell suspensions to constant weight at 105 C in preweighed glass beakers. Prepare samples for the respective analyses as indicated in the chapters describing the analytical methods. 3.1.4. Determination of culture supernatant osmolarity As the cells consume solutes from the medium and secrete soluble metabolites, the actual osmolarity of the culture supernatants will differ from that of the initial medium and/or the calculated value. Therefore, it is important to measure the actual osmolarity of culture supernatants, for example, with a Semi-Microosmometer K-7400 (Knaur) (freezing-point measurement). Perform at least three independent measurements for each culture supernatant samples.

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3.2. Measuring intracellular osmolytes One typical reaction of cells to hyperosmotic stress is the increase of solute concentration in the cells. These natural osmolytes are usually sugars or sugar alcohols, glycerol and trehalose being the most well known. While S. cerevisiae mainly increases the intracellular concentration of glycerol, other yeasts accumulate different polyols as osmolytes such as arabitol, mannitol, sorbitol, and erythritol in addition to glycerol. P. pastoris, for example, mainly increases the arabitol concentration (Dragosits et al., 2010). This indicates that it is necessary to know the relevant intracellular osmolytes before quantifying them as indicators of osmotic stress. When the main osmolytes of a certain species are not known, it is a good advice to measure, in a first round, major sugar compounds and polyols (e.g., glycerol, arabitol, mannitol, sorbitol, erythritol, and trehalose) before reducing the number of tested compounds. However, once known they can easily be measured and provide at least a semiquantitative or relative measure of osmostress. In this context, it is crucial to remind the reader of extensive interrelations of stress responses. As metabolite production and accumulation may serve several purposes in the cell and may even be interconnected with overflow metabolism, for example, in oxygen-limited conditions (Carnicer et al., 2009), it is important to interpret the data cautiously. 3.2.1. Required materials Cell samples (fresh cell pellets or samples that were frozen at 80 C) 0.5 M Tris–Cl (pH 5.5) Standard compounds (glycerol, trehalose, mannitol, arabitol, erythritol) HPLC equipment including a Aminex HPX-87H column and refractometric detector 4 mM H2SO4

3.2.2. HPLC measurements Centrifuge 2 mL aliquots of chemostat culture at 4 C and 13,000 rpm for 2 min to pellet cells. Remove the supernatant and freeze cells immediately at 80 C until use. To quantify the intracellular levels of glycerol, arabitol, mannitol, erythritol, and trehalose, perform heat extraction as described by Philips and coworkers (Philips and Herskowitz, 1997). Cell pellets are resuspended in 0.5 M Tris–Cl, pH 7.5, heated to 95 C for 10 min, and centrifuged for 10 min to remove cell debris. Alternatively to the original protocol by Philips et al., heat extraction may also be performed in distilled water as the Tris buffer may interfere with HPLC measurements in the case that higher molecular weight substances are to be analyzed (Tris will give a very intense and relatively broad peak at approximately 5.5 min in HPLC measurements as it is described below). Supernatants are filtered to remove


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particles and are then kept for analysis via HPLC (samples can be kept at 4 C for 24–48 h, but it is recommended that samples be frozen). Isocratic conditions, using 4-mM H2SO4 as solvent and a flow rate of 0.6 mL/min on a Aminex HPX-87H column (Biorad) at 40 C and a Biologic DuoFlow (Bio-Rad) combined with a Smartline RI Detector 2300 (Knaur), can be successfully applied to separate and analyze intracellular substances. For the determination of concentrations, external standard solutions of potential osmolytes in a heat extraction buffer (2–0.01 g/L) are measured in the same run as the samples. Solute concentrations are then correlated with biomass used for heat extraction. To extract detectable amounts of osmolytes, the amount of cell material is crucial. Using the chemostat conditions as described here, it is recommended that at least 2 mL of culture sample be used and the osmolytes extracted using 1.5 mL of extraction buffer. For HPLC measurements, use undiluted samples when injecting 100 mL of sample. When samples are taken from shake flasks cultivation (with considerably lower OD600), we recommend that at least the cell pellet from 10 to 20 mL of culture be used.

3.3. Measuring UPR targets Because cellular stress responses are highly interrelated, the protein-folding machinery may be affected by changes of osmotic conditions as indicated by data from halotolerant yeasts (Lahav et al., 2004). When performing realtime PCR, suitable PCR targets might vary depending on the experimental setup (studying the shock response or adapted cells), thus we highly recommend performing an initial screening by large-scale qRT-PCR or microarray analysis. Suitable targets for specific UPR-related genes that may be used to monitor osmotic stress-related UPR response can be found by performing microarray analysis, as described in Dragosits et al. (2010). Differential proteomics Fluorescence 2D difference in-gel electrophoresis (2D-DIGE) proved to be a very useful tool when studying changes on the protein level. Whereas low abundance proteins are hard to quantify and detect when this method is used, it is perfectly fine when the changes affect high abundance proteins such as proteins involved in energy metabolism, amino acid synthesis, translation, transcription, and of course, protein folding. Heat shock proteins of the HSP70 and 90 family and proteins such as Hac1p and BiP, which are of major interest when studying interrelations of UPR and osmotic stress, can be found in a cluster in the pH range of 2–5 and an MW range of 50–90 kDa (Dragosits et al., 2009). 3.3.1. Required materials

Fastprep (Thermo Savant) Glass beads (0.5–1 mm diameter)

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Cell lysis buffer (according to GE Healthcare DeCyder Manual) Cell resuspension buffer (according to GE Healthcare DeCyder Manual) 2D-Clean Up Kit (GE Healthcare) 2D-Quant Kit (GE Healthcare) 2D-DIGE labeling Kit (GE Healthcare) IPG Strips pH 3-11NL 24 cm (GE Healthcare) Electrophoresis chamber IPG Phor Isoelectric focusing unit (GE Healthcare) Typhoon 9400 gel image scanner (GE Healthcare) GE Healthcare DeCyder Software package

3.3.2. 2D fluorescence difference in-gel electrophoresis 2D-DIGE is essentially performed as described previously (Dragosits et al., 2009). It should be noted that the following protocol is very well suited to isolate soluble proteins, whereas integral membrane proteins and cell wall proteins will need specialized extraction protocols for efficient extraction. Cell material from 1 to 2 mL of chemostat culture is resuspended in 1.5 mL cell lysis buffer and combined with 500-mL glass beads. Cells are disrupted in a Fastprep FP120 (2 30 s at speed 6.0) and centrifuged at full speed, 4 C for 10 min to pellet cell debris. The protein-containing supernatant is transferred to a new tube and purified using the 2D-CleanUP kit according to the manufacturer’s protocol (use protocol for samples containing more than 100 mg total protein). Resuspend the pellet that is obtained after the clean-up in a 300 -mL cell resuspension buffer. The protein concentration in the final sample will usually be between 1 and 2 mg/L. Before measuring the protein concentration and performing the labeling reaction, it is essential to check the pH of the samples, which should be between pH 7 and 9. Acidic samples will label inefficiently, resulting in very poor data quality. To check the pH value, drop 5–10 mL of sample on a pH indicator strip. Add 2–5 mL of 5 M NaOH to the sample if necessary. Then, perform protein quantification using the 2D-Quant Kit according to the manufacturer’s manual and label 50 mg of each sample with the respective fluorescent dye. Three samples can be applied per IPG Strip, whereas one sample (usually labeled with Cy2) is an internal standard, composed of equal amounts of all samples of the study. A third real sample may be considered when data analysis software other than DeCyder Software package that does not depend on this internal standard is used. For first-dimension separation on IPG Drystrip pH 3-11NL on the IPGPhor Isoelectric focusing unit, we recommend that the cup-loading procedure be used (preferred over the rehydration loading technique). The following setting should be used for the IPGPhor Isoelectric focusing unit: 20 C, 50 mA per strip: 6 h at 500 V, 3 h at 500–1000 V, 8 h at 1000–8000 V, and 8000 V for a total of 65 kVh.


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If several runs are performed, we strongly recommend that the samples be run until a certain amount of kVh is reached (in our case, 65 kVh total) rather than running them for only a certain amount of time. This will help to yield very reproducible spot patterns on the 2D-gels. After focusing, IPGDryStrips can either be frozen at 80 C or directly moved to second-dimension separation. Before second-dimension separation, the IPG strips are reequilibrated as described in the DeCyder manual (once reequilibrated, the samples have to be used for SDS–PAGE and cannot be stored at 80 C). Second-dimension separation by SDS– PAGE is performed on a 12% polyacrylamide gel. When six gels are run in one chamber, the following setting is applied: 12.5 V for 20 min followed by 100 V for 3–4 h (until the marker front appears approximately 2 cm above the bottom of the gel). The gels can be immediately scanned on a Typhoon 9400 scanner or they can be stored (wrapped in plastic foil) at 4 C overnight before scanning. It is not advisable to store the gels for more than 16 h at 4 C before scanning. 3.3.3. Image analysis and protein identification Fluorescence gel images are taken at a resolution of 100 mm on a Typhoon 9400 Fluorescence scanner (higher resolutions can be used but will not significantly improve data quality but only result in a massive increase of data amounts). Then, the DeCyder Software package v.5 is used to analyze the obtained gel images. Significantly regulated protein spots (fold-change 1.5, 1-way ANOVA 0.05 in at least one comparison of cultivation conditions and present on at least 80% of the spot maps) can be picked from Coomassie stained gels. After the gel-scanning process, 2D gels can be treated as standard SDS–PAGE gels to fix and stain proteins (Coomassie stain or mass spectrometry compatible silver staining protocols). Gel spots may be stored in microcentrifuge tubes and overlaid with water (stored at 4 C) until MS-based protein identification. For protein identification, proteins are subjected to a tryptic digest to reversed phase capillary chromatography (BioBasic C18, 5 mm, 100 0.18 mm, Thermo) and ESI–MS/ MS on a quadrupole time-of-flight (Q-TOF) Ultima Global (Waters Micromass) mass spectrometer. Mass spectra are analyzed either by using the Protein Lynx Global Server 2.1 software (Waters) or X!Tandem (http:// www.thegpm.org/tandem/). Only proteins identified by at least two peptides should be considered to represent confident hits (Dragosits et al., 2009). We also suggest not using data for spots that contain more than one protein, as a quantification procedure in these cases might be inaccurate. Additional gels with different pH gradients for isoelectric focusing might be used to resolve these multiprotein spots.

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ACKNOWLEDGMENTS The experiments described in this chapter are based on projects supported by the European Science Foundation (ESF, program EuroSCOPE), the Austrian Science Fund (FWF), project no. I37-B03, and the Austrian Research Promotion Agency (Program FHplus). Thanks to Martina Chang (Polymun Scientific) and Burghardt Scheibe (GE Healthcare) for their support and advice in 2D-DIGE, Alexandra Graf (University of Applied Sciences FH Campus Vienna) for extensive support in microarray data analysis, Hans Marx (University of Natural Resources and Life Sciences) for his advice in HPLC analytics, and Astrid Mecklenbraüker and Corinna Rebnegger (University of Natural Resources and Life Sciences) for their help concerning real-time PCR.

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Induction of ER Stress Response Leading to Programmed Cell Death in Trypanosoma brucei Hanoch Goldshmidt*,† and Shulamit Michaeli*,† Contents 190 193 193 194 194 195 197 198 201 201 202 203 203

1. Introduction 2. Induction of ER Stress in Trypanosomes 2.1. DTT and 2-deoxy-D-glucose treatment 3. Measurements of ER Stress in Trypanosomes Cells 3.1. BiP upregulation 3.2. mRNA decay 3.3. Measurement of intracellular calcium 3.4. ER expansion 4. Measurements of ER Stress-Induced Cell Death 4.1. Sub-G1 fraction 4.2. DAN ladder Acknowledgments References

Abstract Trypanosomes are parasitic protozoans that include several medically and a variety of economically important parasites, such as Trypanosoma brucei, the causative agent of sleeping sickness. This parasite cycles between the insect host (procyclic form) and mammalian host (bloodstream form). These parasites lack transcription regulation, including factors that govern the unfolded protein response (UPR) in other eukaryotes. Gene expression is controlled posttranscriptionally by unique mechanisms such as trans-splicing and RNA editing and by mRNA stability. In trans-splicing, a common exon, the spliced leader (SL) is donated to all mRNAs from a small RNA, the SL RNA. The SL RNA is transcribed from a defined promoter assisted by the tSNAP complex. Despite the lack of transcriptional regulation, induction of ER stress elicits changes in the transcriptome similar to those induced by conventional UPR found in other * The Mina and Everard Goodman Faculty of Life Sciences, Bar-Ilan University, Ramat-Gan, Israel { Advanced Materials and Nanotechnology Institute, Bar-Ilan University, Ramat-Gan, Israel Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00011-X

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eukaryotes. The mechanism of upregulation under UPR is dependent on differential stabilization of mRNAs. The transcriptome changes result in ER expansion and elevation in the ER chaperone, BiP. Prolonged ER stress induces the spliced leader RNA silencing (SLS) pathway. SLS is the trypanosome-specific stress response mechanism that elicits the shut-off of SL RNA transcription by perturbing the binding of the transcription factor tSNAP42 to its cognate promoter, eliminating trans-splicing of all mRNAs. SLS was discovered in the RNAi silenced cells depleted for functions that mediate translocation of proteins to the ER such as the signal recognition particle receptor SRa, SEC63— a factor that participates in protein translocation across the ER membrane, or SEC61— the translocation channel. Induction of SLS, either by prolonged ER stress or silencing of the genes associated with the ER membrane that function in ER protein translocation led to programmed cell death (PCD), evident by the exposure of phosphatidyl serine, DNA laddering, increase in ROS production, increase in cytoplasmic Ca2þ, and decrease in mitochondrial membrane potential. Here, we describe the protocols to induce ER stress and to observe the resulting morphological changes by transmission electron microscopy (TEM), changes in cytoplasmic Ca2þ, and DNA fragmentation which are the hallmarks of programmed cell death.

1. Introduction Trypanosomatids are known for their nonconventional gene expression mechanisms, including trans-splicing (Agabian, 1990; Liang et al., 2003) and RNA editing (Feagin, 1990; Stuart et al., 2005). Trans-splicing is essential for the maturation of all mRNAs. In trans-splicing, a small exon, the spliced leader (SL), encoded by a small RNA, the SL RNA, is donated to all pre-mRNA by trans-splicing (Agabian, 1990). No promoters upstream to each protein coding were identified, and transcription is initiated only at numerous positions across the chromosomes (Kolev et al., 2010). As a result of this mode of transcription, the protein-coding genes are arrayed in long polycistronic transcription units which are processed by the concerted action of trans-splicing and polyadenylation (Liang et al., 2003). Thus, gene expression in these parasites is regulated primarily posttranscriptionally at the level of mRNA degradation and translation; for most genes, the signals that dictate this regulation are situated at the 30 UTR (Clayton and Shapira, 2007). Recent RNA-seq data also suggest the existence of alternative splicing of gene transcripts during the developmental cycle of the parasite (Nilsson et al., 2010). Trypanosomes do not completely lack transcriptional regulation. The SL RNA has a defined promoter that binds specific transcription factors such as tSNAPc (Campbell et al., 2003; Das et al., 2005; Schimanski et al., 2005). Moreover, the synthesis of SL RNA was found to regulate gene expression

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under a variety of stresses, especially ER stress; this mechanism was named SLS, for SL RNA silencing (Lustig et al., 2007). SLS was first discovered in cells silenced for the signal recognition particle receptor, SRa. Depletion of SRa results in the shut-off of SL RNA transcription. ChIP assay further demonstrated that under SRa depletion, the SL RNA transcription factor tSNAP42 fails to bind to the SL RNA promoter, thus avoiding the formation of the transcription complex (Lustig et al., 2007). SLS can also be induced by the silencing of two other important ER translocation factors; the SEC63, which is essential for translocation of proteins via the posttranslational and the cotranslational translocations pathways, and SEC61, the translocation channel, pointing out the major importance of functional integrity of the ER and its membrane (Goldshmidt et al., 2010). The endoplasmic reticulum (ER) is the site for folding of proteins that traverse the ER, on their way to other cellular compartments. The ER also maintains the cellular calcium homeostasis (Malhotra and Kaufman, 2007). Resident ER chaperones, high levels of calcium, and an oxidating environment are most essential for ER function. Proteins that translocate to the ER undergo proper folding and those can exit the ER. ER function is sensitive to changes in calcium level, inhibition of glycosylation, oxidative stress, and exposure to reducing agents therefore deviation of homeostasis of these physiological aspects induce the ER stress response, which triggers specific signaling pathways, including the unfolded proteins response (UPR). UPR leads to reduction in the load of proteins to be translocated, enhanced degradation of misfolded proteins, and increased folding capacity of the ER (Bernales et al., 2006b; Ron and Walter, 2007; Travers et al., 2000). In mammals and yeast, UPR is dependent on IRE1, which is a bifunctional kinase and endonuclease that cleaves a nonconventional intron from HAC1 mRNA in yeast, or XBP1 in mammals which encodes for a transcription factor (bZIP) (Cox et al., 1997; Kawahara et al., 1997). In mammals, PERK and ATF6 also participate in UPR. PERK phosphorylates eIF2a to mediate translational attenuation. ATF6 is cleaved in the Golgi and translocates to the nucleus to activate the transcription of chaperones such as BiP (Haze et al., 1999; Lee et al., 2002; Ye et al., 2000; Yoshida et al., 2003). When the ER stress is persistent, the cells activate the apoptotic pathway via the activation of Caspase 12 (Morishima et al., 2002; Rao et al., 2002). Based on the peculiarities in trypanosome gene expression, it is not surprising that trypanosomes lack homologues to the UPR-mediating factors. Moreover, studies claim that although transcriptome changes were observed in T. brucei during the developmental cycle, very minor changes take place as response to ER stressors as well as serum starvation (Koumandou et al., 2008). Our recent studies, however, detected a clear response to ER stressors using higher concentration of DTT (Goldshmidt et al., 2010).

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The transcriptome of cells exposed to DTT was compared to analogous transcriptomes of other eukaryotes, including man, yeast, nematodes, and drosophila, and the results demonstrate that in trypanosomes, like in these other eukaryotes, similar changes were observed (Goldshmidt et al., 2010). The largest category of upregulated genes in response to ER stress is the category of genes that function in protein secretion. It was further demonstrated that the upregulation of genes, such as the chaperon DnaJ, protein disulfide isomerase, thioredoxin, and syntaxin, results from selective stabilization of their mRNAs. Thus, the upregulation of genes essential for executing the ER stress that is achieved in other eukaryotes by stressinduced transcriptional activation, is mediated in trypanosomes by selective mRNA stabilization. Further studies should unravel the 30 UTR sequences and the RNA binding proteins that execute this regulation. The induction of the ER stress was also demonstrated by upregulation of the ER chaperone BiP as well as distinct morphological changes, including ER expansion (Goldshmidt et al., 2010). Persistent exposure of T. brucei procylic and bloodstream stages to ER stressor induced SLS that was diagnosed both by the reduction in SL RNA and the accumulation of tSNAP42. Since persistent exposure to UPR induces apoptosis in mammalian cells, it was of interest to explore whether SLS that is induced upon long exposure to ER stressor is a programmed death pathway. Accumulated data suggest that programmed cell death (PCD) exists in trypanosomatids, and plays a major role in maintaining and regulating the parasite population (Duszenko et al., 2006; Welburn et al., 2006). In metazoa, type I PCD is induced by caspases. Trypanosomes lack homologs to metazoan caspases, and their five metacaspases (Gonzalez et al., 2007; Helms et al., 2006; Lee et al., 2007) were shown not to be involved in apoptosis (Gonzalez et al., 2007; Lee et al., 2007). To examine whether SLS is a death pathway or not, different experimental approaches were examined. DNA fragmentation was examined using the TUNEL assay as well by observing DNA laddering which is the hallmark of PCD. SLS was shown to induce membrane flipping assayed by the Annexin V staining. SLS also induces membrane depolarization as well as ROS production. The observation that during SLS, cytoplasmic Ca2þ increases, most probably because of perturbations in the ER, resembles observations in mammalian cells where Ca2þ release from the ER induced by milder insults promotes cell death through apoptosis (Pinton et al., 2008). In this chapter, we describe in detail the activation of ER stress by treatment with DTT and 2-deoxy-D-glucose, and the measurements of BiP induction and electron microscopy analysis to visualize morphological changes accompanying the ER stress and SLS induction. Detailed protocols to evaluate DNA fragmentation, and changes in cytoplasmic Ca2þ, are given as classical hallmarks of PCD in these parasites.


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2. Induction of ER Stress in Trypanosomes 2.1. DTT and 2-deoxy-D-glucose treatment 2.1.1. Required materials Cells of interest Procyclic forms of T. brucei strain 29-13 (obtained from Dr. Paul. T. Englund, Johns Hopkins University, Baltimore, USA). Growth medium Grown in SDM-79 (Brun and Schonenberger, 1979) supplemented with 10% fetal calf serum and antibiotics (penicillin, streptomycin). The medium should be used at least 24 h after preparation. Reagents

Dithiothreitol (DTT) (Sigma) 2-deoxy-D-glucose (2-DG) (Sigma) Disposables

1.8-ml (Eppendorf) tubes and 50-ml or 250-ml tissue culture flasks. Cells culture

One day before induction Subculture trypanosomes from logarithmic growing cells (1 106 cells/ml) by diluting 1:10 into a 10-ml growth medium in 50-ml flasks or into a 100-ml growth medium in 250-ml flasks. Cells are grown at 27 C with moderate agitation (50 rpm). On the day of induction Cells should be in the logarithmic phase. The number of cells should be (5 105–1 106 cells/ml) and should not exceed this number. The reagents to be used should be freshly prepared from stock solutions (1 M DTT or 1 M 2-DG). Add DTT or 2-DG directly into the growing media to a final concentration of 4 mM DTT or 20 mM 2-DG. Tightly seal the flask and incubate at 27 C up to 12 h. Sample collection Collect samples after the desired time points and centrifuge the cells (2200g) at 4 C.

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3. Measurements of ER Stress in Trypanosomes Cells If several time points are required, the culture for each time point should be grown in different flask. However, before inducing the ER stress by adding DTT or 2-DG, the cells grown in the individual flasks should be mixed into a single flask to generate a homogenous population and only then divided to the initial flasks. We recommend that the ER stress for each flask be induced separately in order to harvest the samples treated for the different time points, simultaneously. If three time points are chosen, time zero, half an hour, and an hour, start the 1-h induction and after half an hour, start the 30-min induction, so at 1 h time, the three samples will be handled together.

3.1. BiP upregulation The ability to induce the production of ER chaperones to increase the folding capacity of the ER is a main phenotype of ER stress response (Ellgaard et al., 1999; Malhotra and Kaufman, 2007; Ron and Walter, 2007; Travers et al., 2000). Induction of chaperones is not only mediated by increase in transcript level but also by preferential translation and reduced proteolysis under stress (Rutkowski et al., 2006). Although BiP mRNA is not among the mRNAs whose level is most highly increased under ER stress (Goldshmidt et al., 2010; Lee, 2005; Travers et al., 2000), the elevation of this protein serves as a major hallmark for ER stress response. To be able to detect changes in BiP level as a result of ER stress, it should be noted that the BiP level might be affected by the way the cells are treated prior to protein extraction. It is absolutely essential to keep the cells under unstressed conditions to avoid BiP induction in the control. Therefore, it is essential to keep the following roles: 1. Keep the cells in appropriate physiological conditions (except the treatment itself ). 2. Keep the cells and buffers in a chilled environment when starting to pellet the cells. 3. Shorten the process as possible.

3.1.1. Solutions Protein extraction buffer (50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 5 mM EDTA, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS)


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3.1.2. Disposables

1.8-ml (Eppendorf) tubes 15-ml tubes 50-ml tissue culture flasks

3.1.3. Treatment Induce ER stress by using 4 mM DTT or 20 mM 2-DG as explained above, in a volume of 10 ml in 50-ml tissue culture flasks. 3.1.4. Sample collection In this experiment, collecting the samples (control and treated cells) in the same batch is crucial to achieve reliable results. Transfer the control and treated cells to 15-ml tubes and pellet the cells at 4 C for only 2 min at 3500g. Immediately discard the supernatant by turning down the tube, discard the residual medium with a 200-ml pipette, and put the cell on ice. Add 50 ml of protein extraction buffer (kept on ice) previously supplemented with protease inhibitor cocktail (complete mini EDTA free, Roche Inc.) and resuspend the sample a few times, but avoid foaming. Leave for 10 min on ice and then centrifuge for 10 min at 23,000g at 4 C. Discard the pellet and keep the supernatant. The concentration of protein level in all the samples should be the same, but you can read it and make corrections accordingly. Separate the protein samples by electrophoresis on 10% SDS polyacrylamide gel and perform western blot analysis using anti-BiP antibodies (we used specific antitrypanosomes BiP diluted 1:5000 that were kindly provided by Prof. James Bangs from the University of Wisconsin– Madison, Madison, USA), followed by additional antibody on the same blot to control for the loading. We commonly used antibodies made to trypanosome proteins such as anti-PTB1, -PTB2 (Stern et al., 2009), or -hnRNPDo (Michaeli et al., 1992).

3.2. mRNA decay As detailed in Section 1, the response to ER stress in trypanosomes is mediated not by transcriptional regulation but by stress-induced mRNA stabilization. Here, we describe the method to measure the changes in mRNA stability under ER stress. 3.2.1. Reagents

TRI-Reagent (Sigma) Actinomycin D (Sigma) Sinefungin (Sigma)

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3.2.2. Disposables

50-ml tubes 250-ml tissue culture flasks 1000-ml sterile beakers

3.2.3. Procedure Given that the half-life of mRNAs can range between a few minutes to more than an hour, we recommend the use of five time points: 0, 15, 30, 45, and 90 min. For each time point, a 50 ml of 5–10 106 cells/ml should be used. Grow 500 ml in five 250-ml tissue culture flasks (100 ml each) or in 1000-ml beakers. It is important to mix all the grown cells and then separate them again on the day of the experiment into two 1000-ml sterile beakers, to keep a homogenous population. It is most important to start the measurement of mRNA stability in the time frame that ER stress response is induced but before the cell death process starts. For 4 mM DTT treatment, we recommend that the stress for 1.5 h be induced before the measurements of the half-life are started. Add DTT as described in Section 2.1. 1. After 1.5 h of treatment, divide into 50-ml tubes (5 for control and 5 for treatment) and concentrate by centrifugation (5 min,3000g), discarding 45 ml and resuspending the pellet in the remaining 5-ml culture (this is recommended in order to keep the cells in the same physiological conditions and not to change the DTT concentration in the experiment). 2. After 2 h from stress induction, add sinefungin (2 mg/ml) and mix gently. 3. After additional 10 min (130 min from ER stress induction, add Actinomycin D (30 mg/ml), mix gently, and immediately spin down the cells of control and the time zero of the treatment with ER stressors (30 s, 3500g, 4 C), discard the supernatant and quickly extract the RNA by resuspending the cells using a 1-ml pipette in TRI-Reagent. 4. Collect the other samples at the different time points recommended above, and treat the samples as described above for the zero time point. 5. Resuspend the RNA, fractionate it on Northern blot, and hybridize with DNA probes, as described in Goldshmidt et al. (2010). 6. To control for equal loading, use long-lived RNA (7SL RNA or rRNA) as probes. 7. Expose the blot to phosphor-imager and calculate the half-life of the mRNAs. Results can be seen in Goldshmidt et al. (2010).


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3.3. Measurement of intracellular calcium 3.3.1. Reagents

EGTA (ethylene glycol tetraacetic acid) (Sigma) Fluo-4-AM (Invitrogen) CaCl2 (Sigma) Ionomycin (Sigma) (2 mM stock) 10% Triton X-100 (Sigma) Phosphate-buffered saline (PBS), pH 7.4 (Sigma)

3.3.2. Solutions

PBS, pH 7.4

3.3.3. Disposables

1.8-ml (Eppendorf) tubes 15-ml centrifuge tube 50-ml tissue culture flasks Flow cytometry tubes

3.3.4. Cell culture Culture cells (10 ml, 106 cells/ml) in 50-ml tissue culture flasks and induce ER stress as explained above. 3.3.5. Method To measure the intracellular calcium level, we used Fluo-4-AM. This reagent becomes fluorescent only after its entry into the cell. The acetoxymethyl (AM) ester is cleaved in the cell to give the free indicator Fluo-4 that cannot exit the cell, and its fluorescence measures the intracellular calcium concentration. Procedure 1. Count the cells and take 106 cells from each sample (control and experiment/s). 2. Collect the cells (5 min, 3000g, 4 C) and wash once in PBS. 3. Resuspend the cells in 0.5-ml PBS and add 0.5 ml of Fluo-4-AM (1 mM stock) to a final concentration of mM. Incubate at 27 C for 1 h. 4. Wash the cells three times (2 min, 3000g) with PBS and finally resuspend in 1-ml PBS. Separate into 2 flow cytometry tubes—0.5 ml in each. 5. Evaluate the staining by flow cytometry and analyze the data by a software such as CellQuest software.

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3.3.6. Controls 1. To assure that the measurements are performed on live cells, the following protocol was used during the reading in the flow cytometry. Read the level of fluorescence in the sample. 2. Add to the second tube of the same sample CaCl2 to a final concentration of 2 mM and read. No change should be observed, since the Fluo-4 is present only inside the cell and should not interact with the external CaCl2. 3. Add Triton X-100 to 0.1% or use Ionomycin (1:1000 from 2 mM stock solution) to enable the entry of CaCl2 into the cells. This raises the intracellular Ca2þ and the fluorescence accordingly. The intracellular Ca2þ level could increase because of Ca2þ release from intracellular storage organelles (like the ER). To be sure that increase in intracellular calcium level is due to calcium release from intracellular organelles (in the case of ER stress, the ER is generally the source for the Ca2þ), it is important to perform the experiment also in a Ca2þ-free environment. To this goal, the medium was treated with EGTA that chalets Ca2þ with a special high affinity. EGTA (1 mM) should be added 1 h before the treatment.

3.4. ER expansion The expansion of the ER is a well-described phenomenon. In yeast and mammals, it was suggested that ER dilation helps to accommodate newly synthesized ER proteins and inhibit aggregation of unfolded protein by reducing their concentration (Bernales et al., 2006a; Bommiasamy et al., 2009). Such morphological changes are best observed using electron microscopy. Changes in the ER could be observed within 1 h after DTT addition, but are developing during the treatment. The embedding procedure described here gave the best results. 3.4.1. Cells of interest

Procyclic forms of T. brucei strain 29-13

3.4.2. Reagents

Dithiothreitol (Sigma) OsO4 (Strem) SPI-PonTM 812 Epoxy Embedding Kit (SPI Supplies) Uranyl acetate (SPI Supplies) Lead citrate (Acros)


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3.4.3. Solutions

Karnovsky-cacodylate solution (4% parafomaldehyde, 2.5% glutaraldehyde) buffered in 100 mM sodium cacodylate (pH 7.4) 1% osmium solution 100 mM sodium cacodylate buffer

3.4.4. Disposables

1.8-ml (Eppendorf) tubes 50-ml centrifuge tube 250-ml tissue culture flasks Plastic Pasteur pipettes 200-ml tubes suitable for microtome

3.4.5. Cell culture One day before induction A suitable amount of cells for TEM analysis should be around 109 cells. To reach this amount, it is best to prepare a 100 ml of 106 cells in the logarithmic phase and dilute the culture 1:10 to 10 flasks containing 100 ml each. The culture has to be mixed gently before diluting to be sure that the inoculum is a homogenous population. Day of induction On the day of treatment, the culture should not exceed 106 cells/ml. Add the DTT to each flask and pellet the cells after 1 h of treatment or longer. The morphology changes are most prominent after 3 h of DTT treatment.

3.4.6. Sample collection The procedure starts with the preparation of 1 ml of Karnovski solution at room temperature for each sample. Pellet the cells in four 250-ml tubes (1000g) 10 min at 4 C. Discard the supernatant by leaving a small volume of the medium in the tube. Resuspend the pellet gently with a 1-ml pipette, collect the entire cell suspension from all 250-ml centrifuge tubes into a 50ml tube, and centrifuge again 1000g for 5 min at 4 C. Discard the supernatant completely, immediately resuspend in the Karnovski solution, and transfer to a 1.8-ml tube. From here, all the work is done in a chemical hood. Leave the samples at room temperature for 1.5 h and then transfer to 4 C for at least 1 h before proceeding to embedding. The sample can be left at 4 C for up to 1 month.

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3.4.7. Embedding All centrifugation steps are done very gently at 1000g for 2 min unless differently recommended. A swing-out bucket should be used. Be sure to discard the supernatant to the appropriate disposable toxic waste. The first step is to wash the Karnovski fixatives three times with cacodylate buffer. At each step, use a vortex to mix the cells and then wash by rotating for 15 min. We use a circular rotator with a 25 fixed angle that rotates 20 rpm. Osmium fixation is done in the dark. Resuspend the sample in 0.5 ml of the preheated to a room temperature of 1% OsO4 solution and rotate for 1 h at room temperature. Cover the sample to keep it in the dark. After 1 h, the sample color should be changed to black. Discard the supernatant into OsO4 neutralizer and use the neutralizer also to wash the plastic Pasteur pipettes before discarding them to the toxic waste.

3.4.8. Dehydration

Each wash is done using 1 ml of 50% ethanol (twice for 10 min) and then with 70% ethanol (twice for 10 min).

At this stage, it is possible to stop and leave the samples at 4 C for overnight. The following washes should be done:

Twice with 90% ethanol for 10 min, then twice with 100% for 10 min, and finally, twice with propylene oxide for 8 min. The cell pellets do not hold together very tightly, and it is desired to carefully discard the wash during the many steps. Prepare the Epoxy Resin mixture from the SPI-PonTM 812 Epoxy Embedding Kit. Mix with 1 ml of propylene oxide with Epoxy Resin at a ratio of (3:1). Wash the sample twice with this mixture for 10 min; while discarding, just turn down the Eppendorf tube and let the supernatant leak out. Finally, add 0.5 ml of the Epoxy Resin and only partially mix the cells with the resin by gently twisting it with a wooden sliver. Rotate overnight uncovered to enable evaporation of the propylene oxide. Centrifuge (14,000g for 20 min) (or until the cells are concentrated at the bottom of the tube). It can take longer than 20 min depending on the cell weight. Discard the liquid by turning down the Eppendorf tube, allowing the supernatant to leak outside. Now, very gently, without pressing the cells, transfer the sample to 200-ml tubes and fill with the Epoxy Resin. Leave in 60 C for 3 days for polymerization. Now, the samples are ready for sectioning and visualizing by EM.


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4. Measurements of ER Stress-Induced Cell Death 4.1. Sub-G1 fraction The method is based on measuring the content of DNA molecules per cell. When cells undergo apoptosis, the DNA breaks into small pieces; thus, apoptotic cells display a characteristic broad hypodiploid (sub-G1) peak in FACS analysis, which can be easily distinguished from the narrow peak of cells with normal (diploid) DNA content. 4.1.1. Reagents RNase A (Roche) (10 mg/ml stock) (heat-inactivate by heating at 95 C for 15 min) Propidium iodide (Sigma) (2.5 mg/ml stock)

4.1.2. Solutions

PBS, pH 7.4 70% ethanol in PBS (70 ml ethanol absolute and 30-ml PBS1)—store at 20 C PI staining solution (50 mg/ml RNase A, 25 mg/ml propidium iodide in PBS) 4.1.3. Disposables

15-ml centrifuge tubes 1.8-ml (Eppendorf) tubes 50-ml tissue culture flasks Flow cytometry tubes

4.1.4. Cell culture

Induce ER stress as described in Section 2.1 in 50-ml culture flasks.

4.1.5. Procedure Fixation 1. Harvest the treated and control cells (5 min, 3000g, 4 C) in 15-ml tubes and resuspend in 300-ml ice cold PBS. 2. Add 5 ml of the 20 C cold 70% ethanol, dropwise with simultaneous gentle mixing with a vortex. 3. Leave overnight at 4 C (can be kept up to 2 weeks).

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Staining 1. Centrifuge (5 min, 3000g, 4 C) and resuspend in 1-ml ice-cold PBS. Leave on ice for 0.5 h. 2. Centrifuge again (5 min, 3000g, 4 C) and resuspend in 1-ml ice cold PI staining solution. 3. Cover the tubes with a silver wrap and leave for 20 min at room temperature. Measure the samples by FACS. Analyze the FACS data using the cell cycle mode of your software.

4.2. DAN ladder 4.2.1. Reagents

Proteinase K (Roche) RNaseA (Roche) (25 mg/ml stock in ddw) Bio-Phenol Chloroform 100% ethanol 10% sodium dodecyl sulfate (SDS) Sodium acetate (3 M stock, pH 8)

4.2.2. Solutions 50 mg/ml RNaseA in 10 mM Tris (pH 7.5), 15 mM NaCl (heat to 95 C for 15 min before use to destroy DNases) DNA extraction buffer (10 mM Tris, pH 8.0, 0.1 mM EDTA, pH 8.0, 20 mg/ml RNaseA, 0.5% SDS) PBS, pH 7.4 (Sigma) TE (10 mM Tris, pH 8, 1 mM EDTA, pH 8.0)

4.2.3. Disposables

1.8-ml (Eppendorf) tubes

4.2.4. Cell culture Induce ER stress as explained in Section 2.1. However, in this experiment, the timing for harvest is very important. You have to try different time points to be sure that you collect cells during the dying process but not when DNA is completely degraded.


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4.2.5. Procedure 1. Harvest 5 107 cells by centrifugation (5 min, 3000g, 4 C), wash with 1-ml ice cold PBS, and transfer to a 1.8-ml Eppendorf tube. 2. Resuspend in a 450-ml DNA extraction buffer and incubate at 37 C for 1 h. 3. Add proteinase K to a final concentration of 4 mg/ml and incubate at 50 C for 3 h. 4. Add 0.5 ml of phenol:chloroform (1:1, v/v) and mix by inverting the tube several times. 5. Centrifuge (23,000g, 2 min at room temperature) and very carefully, using a 200-ml pipettor, remove the upper (aqueous) phase into a new Eppendorf tube. (Try to avoid the interface denatured proteins, but take as much as possible of the aqueous phase.) The DNA is more concentrated near the interface. 6. Repeat Steps 4 and 5 again. 7. Add 0.5 ml of chloroform, mix by inversion, centrifuge (23,000g, 2 min at room temperature), and decant the upper phase once again. 8. Put the tube in ice for 5 min. Add twice the volume 20 C chilled 100% ethanol and 0.1 of the volume NaOAc 3 M and mix gently by inverting the tube few times. 9. Keep at 80 C for 1 h and spin at 32,000g 4 C for 20 min. 10. Carefully discard the supernatant and wash carefully with 70% ethanol. 11. Resuspend the DNA pellet in 50 ml of TE. 12. Optional: Add 10 mg/ml RNase A and incubate 1 h at 37 C to degrade the residual RNA that can mask the small DNA fragments. Measure the DNA concentration and run 10–15 mg of DNA on 1.8% agarose gel containing 0.5 mg/ml ethidium bromide at 10 V/cm for 2–3 h and visualize under UV light.

ACKNOWLEDGMENTS The research relevant to this work was supported by a grant from the Israel Science Foundation (ISF) and by an International Research Scholar’s Grant from the Howard Hughes Foundation to S.M. S.M. holds the David and Inez Myers Chair in RNA silencing of diseases.

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Measurement of Activation of the Endoplasmic Reticulum Stress Response in Autoimmune Myositis Heather M. Alger,* Sree Rayavarapu,*,† and Kanneboyina Nagaraju*,‡ Contents 1. Introduction 2. Measuring Endoplasmic Reticulum Stress Response in Skeletal Muscle Tissues Using Western Blotting 2.1. Freezing of skeletal muscle tissue for Western blotting 2.2. Preparation of muscle lysates from frozen tissue 2.3. Estimation of protein concentration in muscle lysates 2.4. Western blot for proteins in the muscle lysates 2.5. Interpretation of Western blotting data 2.6. Commonly used positive controls to assess ER stress-activation markers 3. Immunohistochemical Detection of ER Stress Markers in Muscle Tissues 3.1. Freezing and sectioning of skeletal muscle tissue for immunostaining 3.2. Immunohistochemistry for ER stress markers in the skeletal muscle 3.3. Immunofluorescence for ER stress markers in the skeletal muscle 3.4. Interpretation of immunostaining results 4. Determination of mRNA Levels of ER Stress Targets 4.1. Isolation of RNA from frozen muscle tissues 4.2. Semiquantitative reverse transcriptase PCR 4.3. Interpretation of RT-PCR data 5. Concluding Remarks Acknowledgments References

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* Research Center for Genetic Medicine, Children’s National Medical Center, Washington, DC, USA { Institute of Biomedical Sciences, The George Washington University, Washington, DC, USA { Department of Pediatrics, The George Washington University Medical Center, Washington, DC, USA # 2011 Elsevier Inc. Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00012-1 All rights reserved.

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Abstract Evidence suggests that both immune (cell-mediated and humoral) and nonadaptive immune (endoplasmic reticulum (ER) stress and autophagy) mechanisms play a role in muscle fiber damage and dysfunction in idiopathic inflammatory myopathies (IIM). More recently, the ER stress response pathway, the activation of unfolded protein response, and the ER overload response are being studied to understand their contribution in the progression of IIM. A variety of qualitative and quantitative techniques are used to measure the activation of the endoplasmic reticulum stress response in myopathy. Accurately assessing the activation of ER stress response pathway would not only help in the understanding of disease pathogenesis but would also help to assess the response to therapy. Here, we describe common techniques such as western blotting, immunohistochemistry, immunofluorescence, and determination of mRNA levels for the gene of interest to monitor the ER stress in skeletal muscle tissues.

1. Introduction Idiopathic inflammatory myopathies are a heterogeneous group of muscle diseases of unknown etiology, characterized by symmetrical proximal muscle weakness, elevation of serum muscle enzymes, infiltration of mononuclear cells in the skeletal muscle, and autoantibodies. Several recent studies revealed that nonadaptive immune mechanisms such as endoplasmic reticulum (ER) stress response may play a role in disease pathogenesis (Askanas and Engel, 2003; Henriques-Pons and Nagaraju, 2009; Nagaraju et al., 2005; Nogalska et al., 2007; Vattemi et al., 2004). Molecular mechanisms that initiate and perpetuate ER stress in the skeletal muscle have not been fully understood. However, recent evidence suggests that upregulation of major histocompatibility complex class I (MHC-Class I) molecules in skeletal muscle cells, independent of local inflammatory (mononuclear) infiltration, may be responsible for ER stress in the skeletal muscle (Emslie-Smith et al., 1989). MHC-class I antigen processing and presentation and activation of ER stress are highly interlinked processes. Generally, constitutive MHC-class I expression is absent in normal muscle fibers. It has been proposed that ER stress may have a role in the pathogenesis of myositis, because MHC-class I assembly and transport occurs through ER and its expression is widespread on the muscle fibers of polymyositis (PM), inclusion body myositis (IBM), and dermatomyositis (DM) (Nagaraju, 2005; Nagaraju et al., 2005). Further, ER stress is known to be involved in myogenesis and muscle differentiation as well as in other noninflammatory skeletal muscle conditions such as diabetes, because the skeletal muscle is an insulin-sensitive tissue and an important site for glucose utilization in the body. Therefore, it needs to respond to the increased ER load during

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IGF-1-induced muscle hypertrophy or transient hypoglycemic conditions due to impaired ER function (Schroder and Kaufman, 2005). In normal nonstressed cells, improper or unfolded proteins are sequestered by ER chaperones (e.g., calreticulin, calnexin, and GRP78 (Glucose-regulated protein 78)) to facilitate proper protein folding (Khan and Schroder, 2008; Schroder, 2008). However, under severely stressed conditions, these chaperones may be insufficient to prevent the accumulation of unfolded proteins, thus initiating the ER stress pathway as a cellular defense mechanism. Generally, ER stress can also be provoked by: (a) pathological conditions (ischemia, hyperhomocysteinemia, and viral infection) (Kaufman, 1999; Mori, 2000); (b) prevention of protein N-linked glycosylation (UDP glucose deficiency and tunicamycin treatment) (Flores-Diaz et al., 2004; Liu et al., 1997); (c) lipid dysregulation/accumulation (Askanas and Engel, 2003; Devries-Seimon et al., 2005; Hotamisligil and Erbay, 2008; Kedi et al., 2009); and (d) disrupted calcium homeostasis, as can occur with certain drug treatments (e.g., thapsagargin; Devries-Seimon et al., 2005). Default cellular responses to ER stress include (1) a reduction in protein synthesis via activation of sensors localized in the ER (e.g., IRE-1 (Inositol-requiring enzyme 1), PERK (PKR like ER kinase), ATF6 (Activating transcription factor 6)); (2) upregulation of ER chaperone proteins (e.g., BiP/Grp78, Grp94, calmodulin, and calreticulin) to improve protein folding, reduce protein aggregation, and normalize calcium homeostasis; and ultimately, (3) cell death in response to severe, irreparable cell damage via activation of CHOP and Caspase-12 (Matsumoto et al., 1996; Nakagawa et al., 2000). A myositis mouse model closely recapitulating the human cellular disease process was generated by muscle-specific overexpression of MHC class I (Nagaraju et al., 2000). These mice share several salient features of autoimmune myositis, including (1) presentation of muscle inflammation coupled with autoimmune response in some mice, (2) muscle weakness, (3) elevated levels of serum muscle enzymes, and (4) significant defects in skeletal muscle functions. This is a useful model to investigate the role of ER stress in muscle fiber damage and dysfunction in myositis (Li et al., 2009; Nagaraju et al., 2000). In this chapter, we detail some of the common techniques used to monitor ER stress in the skeletal muscle.

2. Measuring Endoplasmic Reticulum Stress Response in Skeletal Muscle Tissues Using Western Blotting 2.1. Freezing of skeletal muscle tissue for Western blotting 2.1.1. Required materials and reagents Isopentane (2-methylbutane) (Sigma, St. Louis, MO), liquid nitrogen, dry ice, aluminum foil, long forceps, glass beaker, and liquid nitrogen-holding container.

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2.1.2. Method 1. Briefly rinse the freshly collected muscle (preferably cut in mid belly) in cold phosphate buffered saline (optional) and blot dry with a paper towel to remove the excess buffer. 2. Snap-freeze the tissue immediately in liquid nitrogen (150 C)-cooled isopentane. It might be helpful to snap-freeze small portions of muscle tissue samples for future use to avoid repeated freeze-thaw cycles of the same sample. 3. Place the muscle piece onto an aluminum foil; immerse the foil with tissue into a beaker containing isopentane chilled in liquid nitrogen for 20 s using long forceps. It is important not to immerse the tissue directly into liquid nitrogen as this may cause nitrogen to boil, and the resulting gas bubbles may insulate the specimen and slow down its quenching. 4. If liquid nitrogen is not available, tissues can be frozen using dry ice and acetone. Note that slow freezing facilitates the formation of ice crystal artifacts. Ice crystal growth occurs relatively slowly below 40 C and does not occur at liquid nitrogen temperatures (150 C). 5. At this stage, frozen specimens can be stored at 80 C in a freezer until thawed for further analysis.

2.2. Preparation of muscle lysates from frozen tissue 2.2.1. Required materials and reagents Cryostat (Model # CM1900, Leica Microsystems), curved scissors, 1.5-ml Eppendorf tubes, Nonidet P-40 (US Biological, Swampscott, MA), 20 mM Tris (pH 7.4), 150 mM NaCl, 1 mM EDTA, and 4 C high speed centrifuge (Model # 5415 R, Eppendorf). Depending on the source (human muscle biopsy or muscle tissue from euthanized animals) and the method (open biopsy vs. needle biopsy) of muscle tissue collection, one can decide either to make lysates directly from tissues or make muscle sections on a cryostat and use them for preparing lysates. The former method is preferred when there is ample tissue sample, and the later method is preferred when the sample is very small (collected from needle biopsy). For the later method, tissue sections should be oriented and embedded into the optimal critical temperature (OCT) compound. This technique is described in detail in Section 3.1. Schematic representation for the preparation of muscle lysates from both methods is shown in Fig. 12.1. 2.2.2. Method 1. Prepare 1% NP40 lysis buffer (LB). (To make 50 ml of 1% NP40 lysis buffer, add 5 ml of 10% NP40, 500 ml of 1 mM EDTA, 1.5 ml of 150 mM NaCl, and 1 ml of 20 mM Tris–Cl, pH 7.4; and bring the total volume to 50 ml using dH2O.)

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Muscle tissue Limited tissue (needle biopsy)

Ample tissue (Open biopsy/muscle from euthanized animal) Store frozen muscle tissue in multiple aliquots (avoid repeated freeze thaw)

Tissue embedded in optimal critical temperature (OCT) medium

Thaw the required amount of tissue on ice

Transfer to cryostat and make 8–10 mM sections

Cut into small pieces and pool them into chilled eppendorf tube

Pool 25–50 tissue sections into chilled eppendorf tubes

Add 75–100 ml of cold 1% NP40 lysis buffer with protease inhibitors Homogenize the tissue at 4 °C (add more lysis buffer if needed) Incubate for 15 min and vortex five times at 4 °C Centrifuge at 4 °C and 13,000 rpm Collect the supernatant (aliquot and store at –80 °C) Dilute the protein lysate and perform protein estimation Prepare the samples appropriately and perform western blot

Figure 12.1 Flow chart for the preparation of muscle lysate for Western blotting.

2. Take 1 ml of the above 1% NP40 buffer and add protease inhibitors (2 ml Leupeptin (500), 2 ml Antipain (500),1 ml Pepstatin (1000), 5 ml PMSF (200), and 2 ml DTT (500 mM)). Mix thoroughly after each of these steps. 3. Label 1.5-ml Eppendorf tubes accordingly and place frozen tissues or the sections in the respective tubes. All steps from this point should be done preferably on ice.

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4. Add about 75–100 ml of 1% NP40 LB with protease inhibitors to each tube (the amount of lysis buffer added might vary based on the size of the tissue in use). 5. Next, using curved dissection scissors or plastic mortar/pestle cut, chop, and grind the muscle tissue. Add another 75–100 ml of 1% NP40 LB with protease inhibitors to each tube and repeat the homogenization procedure, and thoroughly vortex the lysate. 6. Let the tubes sit on ice for 15 min and vortex two to three times during the incubation period. 7. Centrifuge tubes at 13,000 rpm at 4 C for 15 min and then transfer the supernatants into a fresh set of Eppendorf tubes. It is better to make multiple aliquots of lysate and store at 80 C if you are planning to run multiple gels at different times to avoid repeated freeze-thaw of the samples.

2.3. Estimation of protein concentration in muscle lysates 2.3.1. Required materials and reagents Micro titer plates (Costar, Corning, NY), pipettes (20–1000 ml), 1.5-ml Eppendorf tubes, Bio-Rad Dc protein assay kit (Bio-Rad, Hercules, CA), and a microplate reader with appropriate filters (Molecular devices). 2.3.2. Method 1. Label the Eppendorf tubes accordingly for each lysate and decide the dilution to use for each of the samples. 2. Prepare BSA standard stock solution (15 mg/ml). Then, prepare at least four working standards (1.5, 0.75, 0.375, and 0.18 mg/ml) by making serial dilutions using 1% NP40 LB without protease inhibitors. 3. Take a 96-well plate and load the standards and samples (in triplicate), 5 ml per well. 4. Mix 1 ml of Bio-Rad protein reagent A with 20 ml of protein reagent S, mix thoroughly, and add 25 ml of the A þ S mix to each well. 5. Next add 200 ml of protein reagent B to each well. 6. Put the plate in dark for 15 min, and then read the plate at 650 nm using spectrophotometric plate reader. 7. Estimate the amount of protein present in each sample based on BSA standard curve. 8. Determine the amount of protein to load on to a gel and then add loading solution (LDS) (4) (Invitrogen, Carlsbad, CA), reducing reagent (10) (Invitrogen), and dH2O to each sample accordingly. 9. Next heat the samples at 85 C for 5 min, place them on ice for 1 min, and centrifuge at 13,000 rpm for 20 s. Now, the samples are ready to load on to a gel or can be stored at 20 C for later use.


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The procedure described above is a general procedure and requires optimization in different situations.

2.4. Western blot for proteins in the muscle lysates 2.4.1. Required materials and reagents Commercially available gel running (Bio-Rad), and transfer apparatuses (Bio-Rad), laboratory prepared or precast gels (Invitrogen), running buffer (MOPS/MES) (20) (Invitrogen), transfer buffer (20) (Invitrogen), antioxidant (Invitrogen), methanol (Fisher Scientific, Fair Lawn, NJ), suitable pre stained-protein molecular weight standards (Invitrogen), Ponceu stain (Sigma), nitrocellulose membrane (Bio-Rad), nonfat dry milk (Fisher Scientific), Tris-buffered saline (TBS) (10) (Fisher Scientific), tween-20 (Fisher Scientific), Amersham ECL Western Blotting Analysis System (GE Healthcare, Piscataway, NJ), X-ray film (Kodak BioMax film) (Sigma), Hypercasette (Amersham, UK), GS-800 Densitometer (Bio-Rad), and Quantity-one software (Bio-Rad). 2.4.2. Method Electrophoresis of the proteins 1. Determine the percent of the running gel used based on the molecular weight of the protein being studied (e.g., 10–12% gel for proteins 10– 300 kDa and 15% gels for 3–100 kDa) and the amount of protein loaded per well based on the size of the gel. 2. Running buffer is prepared by adding 25 ml of (20) MOPS/MES buffer to 475 ml of deionized water. The MES buffer can be used for lower molecular weight bands (120 kDa). 3. Take out precast gels and assemble them into the gel-running apparatus as per the manufacturer’s instructions. Make sure the gels are packed tightly to avoid leaking. Add 300 ml of running buffer mix to the outside chamber and 200 ml to the inside chamber filling all the way to the top. Next add 500 ml antioxidant (Invitrogen) to the inside chamber. 4. Load 10 ml of suitable marker into first well, followed by samples in other wells. Note that the volume of sample loaded will vary depending on protein concentration. Now, the proteins are electrophoresed by applying a voltage of 200 V for 1 h 30 min. The amount of time might vary depending on the size of the protein. Stop the gel when the dye front reaches the bottom of the gel apparatus. Transfer of electrophoresed proteins onto a membrane 1. Prepare 1 l of transfer buffer by adding 50 ml of 20 transfer buffer to 750 ml of deionized water and 200 ml of methanol.

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2. Also, six pieces (per blot) of white filter paper, padding sponges, and nitrocellulose membrane are made ready for transfer of proteins. 3. Presoak the sponge, filter papers, and membrane in transfer buffer and then arrange “transfer sandwich” by placing the membrane toward the anode and the gel toward the cathode, and these should be appropriately padded with the available filter papers and the sponges. Care should be taken to avoid air bubbles between the gel and the membrane. 4. Fill the transfer apparatus with buffer and transfer the proteins at 300 mA and 4 C for 90 min or 80 mA at 4 C overnight. The transfer time might vary depending on the experimental conditions. Posttransfer development of membranes for protein identification 1. After completion of transfer, remove the membrane from the apparatus and immerse it in 3 ml of Ponceu stain (diluted with water 1:1) for 1 min. Ponceu stain aids to visualize the proteins on the membrane and guides to cut the membranes appropriately. 2. Make sure to label each piece of the membrane appropriately and then block the membranes with 5% milk in Tris-buffered saline (1) with 0.02% tween (TBST) (blotto) or BSA for 1 h. 3. Make up primary antibody dilutions according to the manufacturer’s instructions either in TBST or in blotto buffer. 4. After blocking, rinse the membranes quickly with TBST once, add primary antibody to the respective blots, and incubate the membranes overnight at 4 C. 5. The following day, wash the membranes with TBST five times for 6 min each, using a table top rocker at room temperature (RT). Next, add the appropriate peroxidase conjugated secondary antibody to the blots and incubate for 1 h at RT. Repeat the wash step again. 6. Next, perform ECL reaction (Amersham ECL Western Blotting Analysis System, GE Healthcare) according to the manufacturer’s instructions and expose Kodak Biomax X-ray film (Sigma) using the hypercasette for a length of time that gives best visualization of the blotted bands. 7. Relative tissue expression of ER stress protein of interest can be determined by calculating the ratio of that protein to loading control (vinculin or tubulin). This can be done by scanning the X-ray film and the protein band of interest to be quantified using densitometry (GS-800 Densitometer; Bio-Rad). Several proteins have been identified in the sequential progression of the ER stress pathway. These proteins can be particularly useful in basic determination of the presence and activation of ER stress. Relevant proteins that have been detected by immunoblot were given in Table 12.1.

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Table 12.1 Different ER stress markers identified using immunoblotting protocols (primary antibodies used and their dilutions were listed belowa,b,c ) Proteina

Primary antibodya

Grp78/BIP Grp94 Calreticulin Calnexin Caspase-12 ERP72

Rabbit polyclonal (Stressgen) Rabbit polyclonal (Stressgen) Rabbit polyclonal (Affinity bioreagent) Rabbit polyclonal (Stressgen) Mouse monoclonal (Cell signaling technology) Mouse monoclonal (BD transduction laboratories) Grp75 Mouse monoclonal (SPS-825; Stressgen) SERCA2 Mouse monoclonal (Affinity bioreagents) Phospho-PERK Rabbit monoclonal (Cell signaling) Total PERK Goat polyclonal (Santa Cruz biotechnologies) Phospho-eIF2a Rabbit polyclonal (Cell signaling) CHOP Mouse monoclonal (Affinity bioreagents) Spliced XBP1 Rabbit polyclonal (Santa Cruz biotechnologies) ATF4 Rabbit polyclonal (Aviva systems biology) IRE-1a Rabbit monoclonal (Cell signaling) a b c d

Dilutiona

1:1000 1:500 1:2000 1:2000 NAd 1:200 NAd 1:2500 1:2000 1:1000 1:2500 1:1000 1:2000 1:5000 1:1000

The proteins and the antibody dilutions listed were obtained from Vitadello et al. (2010), Nagaraju et al. (2005), Vattemi et al. (2004). Secondary antibodies used are HRP conjugated anti-mouse or antirabbit depending on the primary antibody used. The blots were developed using an enhanced chemiluminescence system (Amersham). Not available.

2.5. Interpretation of Western blotting data Appropriate interpretation of the results obtained from the above-described methods is very important to understand the unfolded protein response and in turn, the ER stress. For example, higher ratios of Grp78/vinculin (Fig. 12.2) or CAAT homologous protein (CHOP)/vinculin or sXBP-1/ vinculin or the ratio of phospho eIF2alpha to Total-eIF2alpha (or other loading control such as actin or tubulin) indicates activation of the unfolded protein response in compromised tissues. Likewise, the levels of sarco endoplasmic reticulum Caþ 2-ATPase (SERCA2) (110 kDa) increases in the muscle with the progression of UPR and ER stress (Ikezoe et al., 2009). Similarly, Caspase-12 is highly expressed in mouse skeletal muscle tissues, and cleavage of full-length Caspase-12 (45 kDa) to the activated form of Caspase-12 (28 kDa) can be used as a marker of ER stress (Kalai et al., 2003; Nagaraju et al., 2005) in skeletal muscle tissues (Fig. 12.3). ER stress can also be conveniently assessed by probing for the phosphorylated forms of IRE-1, PERK, and ATF6 in cell cultures using Western blotting.

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A

Human myositis

B

Mouse model

GRP 78 – Vinculin –

Ratio of GRP 78/vinculin

C

C2

C3

P1

P2

P3 D

0.35 0.30 0.25 0.20 0.15 0.10 0.05

*

Controls

Patients

Ratio of GRP 78/vinculin

C1

C1

C2

C3

HT-1 HT-2 HT-3

0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1

*

Controls

HT mice

Figure 12.2 GRP78 expression is significantly increased in muscle tissue in human myositis and the mouse model. (A) Skeletal muscle tissue lysates prepared from muscle biopsies obtained from normal controls (n ¼ 4) and myositis patients (n ¼ 8): immunoblotted with anti-GRP78 and antivinculin antibodies. Equal amounts of the protein were electrophoresed in each gel lane. Representative GRP78 and vinculin immunoblots of three controls (C1–C3) and three myositis patients (P1–P3) are shown. (C) The data were quantitated, as described in Section 2.4.2. The differences in GRP 78 levels between normal controls and myositis patients are highly statistically significant (p < 0.0001). (B, D) Skeletal muscle tissue lysates from double transgenic HT mice (n ¼ 4) and control transgenic mice (n ¼ 4) were analyzed as described in (A). A representative blot of GRP78 and vinculin on samples made from three single transgenic control mice (C1–C3) and three double transgenic mice (HT-1–HT-3) is shown. The differences in GRP78 levels between control single transgenic and double transgenic mice are very highly statistically significant (p < 0.0001) (copyright 2010 John Wiley and Sons; Nagaraju et al., 2005).

2.6. Commonly used positive controls to assess ER stress-activation markers Several methods are available to generate positive control samples for monitoring ER stress. Tunicamycin (inhibits N-linked glycosylation),thapsigargin (inhibits ER calcium ATPase pump), and Brefeldin A (inhibits protein transport from ER to Golgi apparatus) are commonly used to induce ER stress in cell culture conditions (Devries-Seimon et al., 2005; Malhotra and Kaufman, 2007).

3. Immunohistochemical Detection of ER Stress Markers in Muscle Tissues At the histological level, muscle biopsies reveal information on the status of muscle fibers, blood vessels, infiltrating immune cells, and neuromuscular junctions. Techniques such as Western blotting are quantitative

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45 kDa

H+

H+

H+T+

H+T+

31 kDa

Figure 12.3 Caspase-12, a critical mediator of ER stress-induced cell death is significantly activated in HT transgenic mice. Skeletal muscle cell lysates from the affected double (HT) and control single (H or T) transgenic mice were prepared in NP-40 lysis buffer with protease inhibitors. Lysates were electrophoresed on SDS-PAGE and transferred onto the nitrocellulose membrane. The blots were incubated with rabbit anti-mouse Caspase-12 antibodies (Cell Signaling Technology) and developed with anti-rabbit HRP. Note a significant increase in the amount of cleaved Caspase-12 ( 28 kDa) in the double (HT) transgenic (lanes 1 and 2) but not control single (H) transgenic (lanes 3 and 4) mice. Vinculin was used as loading control (data not shown) (copyright 2010 John Wiley and Sons; Nagaraju et al., 2005).

and give overall status of the ER stress in an affected tissue, but they fail to reveal qualitative information on the localization of ER stress to a particular cell type in the muscle biopsy. Therefore, techniques such as immunohistochemistry and immunofluorescence can be used for specific cellular localization of ER stress response markers. Qualitative (semiquantitative) immunostaining techniques in corroboration with quantitative Western blot or RT-PCR techniques will provide a comprehensive idea on the status of ER stress in the affected tissues. These techniques are described in detail in later sections of this chapter.

3.1. Freezing and sectioning of skeletal muscle tissue for immunostaining 3.1.1. Required materials and reagents Phosphate-buffered saline (PBS) without Ca2þ, Mg2þ (pH 7.4), isopentane (2-methylbutane) (Sigma), liquid nitrogen, dry ice, aluminium foil, a long forceps, a glass beaker to hold isopentane, Optimal Critical Temperature (OCT) compound (Sakura Finetek, CA), corks (Electron microscopy sciences, Hatfield, PA) Cryostat, microscopic slides (Fisher Scientific), a tissue sectioning blade, and Cryostat (Model # CM1900, Leica Microsystems).

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3.1.2. Method 1. Immediately after biopsy, rinse the tissue in cold PBS, and blot-dry with a paper towel. 2. Add a little amount of OCT compound on to a cork and place the tissue on the cork with OCT compound using forceps. Immediately, snapfreeze the tissue in the liquid nitrogen-chilled isopentane. Take care to orient the tissue appropriately for sectioning and further staining. Tissues can be stored at 80 C or sectioned immediately for staining. 3. For sectioning, cut 8–12-mm thick serial frozen sections by properly adjusting the cork on to the cryostat. Place the sections onto the glass slides and store them at 80 C until further processing.

3.2. Immunohistochemistry for ER stress markers in the skeletal muscle 3.2.1. Required materials and reagents Tris-buffered saline (10) (TBS) (Invitrogen), acetone (Sigma), rack to hold the slides (Humidified chamber), normal goat and horse serum (Vector Laboratories, Burlingame, CA), peroxidase blocking solution (DAKO, Carpinteria, CA), bovine serum albumin (BSA) (Sigma), primary antibodies directed to antigens activated under ER stress (see Table 12.2), secondary antibodies conjugated to HRP (Horse radish peroxidase) or a fluorescent dye (FITC or Texas red), and directed specifically to the species of the primary antibodies, Di-amino-benzoicacid (DAB) chromogen (Vector Laboratories), Hematoxylin (Sigma), light microscope and camera software (Olympus C.A.S.T. stereology system, Olympus America Inc., Center Valley, PA), fluorescent microscope (Nikon Eclipse E800, Japan) fitted with laser and appropriate filters and spot digital color camera with SPOT advanced software (Diagnostic instruments, Sterling heights, MI), cover slips (Fisher Scientific), and Vectashield Hard Set mounting medium for immunofluorescence (Vector Laboratories), and Permount mounting medium for immunohistochemistry (Fisher Scientific).

3.2.2. Method 1. Obtain slides from the freezer and allow them to warm at room temperature. 2. Prepare 1 TBS solution (100 ml Tris (pH 7.4), 60 ml of 5 mM NaCl, and 1840 ml of distilled water) and blocking solution (50% normal goat serum and 50% normal horse serum) before starting the actual staining process.

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Table 12.2 Different ER stress markers identified using immunostaining protocols (primary antibodies used and their dilutions were listed belowa,b,c ) Protein

Primary antibody

Dilutiona

Grp78/BIP Grp94 Calreticulin Calnexin Caspase-12 SERCA2 Phospho-PERK Phospho-eIF2a HERP ERp72

Rabbit polyclonal (Abcam) Rabbit polyclonal (Stressgen) Rabbit polyclonal (Affinity bioreagent) Rabbit polyclonal (Stressgen) Rabbit polyclonal (Chemicon) Mouse monoclonal (Affinity bioreagents) Rabbit monoclonal (Cell signaling) Rabbit polyclonal (Cell signaling technology) Goat polyclonal Mouse monoclonal (BD transduction laboratories) Mouse monoclonal (Stressgen) Rabbit polyclonal (Santa Cruz) Rabbit polyclonal (Santa Cruz biotechnologies)

1:200 1:50 1:1000 1:1000 1:200 1:500 1:100 1:100 1:100 1:20

Grp75 CHOP Spliced XBP1 a b c d

NAd 1:400–1:800 1:100

The proteins and the antibody dilutions listed were obtained from Vitadello et al. (2010), Nagaraju et al. (2005), Vattemi et al. (2004). Secondary antibodies used are HRP/fluorescence tagged anti-mouse or anti-rabbit depending on the primary antibody used. Nonspecific binding of antibody to Fc receptors was blocked by preincubating the sections with normal goat and horse serum (1:10) dilution. Not available.

3. Fix the slides by immersing them in cold acetone solution for 10 min and allow them to air-dry at least 5 min. Then, outline the tissue section with DAKO wax pen and label the slides. 4. Rehydrate the slides by immersing them in TBS buffer for 10 min. Then, gently tap the slide and drain the excess solution with a paper towel and then incubate with peroxidase blocking solution for 10 min. 5. Wash in TBS buffer and apply 100-ml blocking solution to each slide (to cover the entire section) and incubate in a humidified slide chamber for 60 min at RT. 6. Prepare the diluents for primary antibody solutions using blocking buffer as per the manufacturer’s instructions. The antibodies commonly used for immunostaining purposes are listed in Table 12.2. 7. Gently tap the slides to remove most of the blocking solution and apply 100 ml of primary antibody to each slide and incubate overnight at 4 C (make sure the incubation chamber has moisture). It is advisable to stain some slides with appropriate isotype control antibodies to confirm the specificity of the antibody.

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8. The following day, wash the slides in TBS solution (1) three times with gentle agitation. 9. Prepare the secondary antibody (using the same diluents as above) and apply 100 ml of secondary antibody to each slide. Incubate for 1 h at RT. 10. Wash the slides in TBS solution (1) five times with gentle agitation. 11. Prepare the DAB substrate solution as per the manufacturer’s instructions and apply 100 ml of substrate to each slide. Observe for the development of color over the next 2–10 min. Once the color is satisfactorily developed, tap-off the substrate and wash the slides immediately in deionized water. 12. For nuclear staining, dip slides in hematoxylin solution for 30 s and immediately rinse them in running tap water for 5–10 min to “blue.” Leave the slides in water for some more time and then air-dry them at RT. 13. To complete, mount the cover slip using permount medium (take care to avoid formation of air bubbles). 14. Digital pictures of stained sections can be taken using a light microscope connected to a camera and can be used for quantification.

3.3. Immunofluorescence for ER stress markers in the skeletal muscle Similar to immunohistochemical localization of target antigens, immunofluorescence can be used to localize ER stress markers within the tissue. Proteins that are upregulated in ER-stressed homogenates by immunoblot are good targets for immunolocalization within the muscle tissue sections. Immunofluorescence has an added benefit over immunohistochemistry in that multiple proteins can be detected within the same section and colocalization with organelles can be determined. DAPI (40 ,6-diamidino-2-phenylindole) is a commonly used counter stain to identify the nucleus in sections. Additionally, calnexin and other ER chaperones can also be used as ER markers. The protocol for immunofluorescence is very similar to that of immunohistochemistry; however, some differences exist which are described below. 3.3.1. Method 1. Follow the Steps 1–5 described under Section 3.2.2. However, it is advised to keep the slides in a dark humidified chamber for immunofluorescence. 2. For immunofluorescence, the secondary antibodies will be conjugated to a wavelength-specific fluorescent tag. Secondary antibodies should be selected based on the species in which the primary antibody is developed


3. 4.

5. 6.

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and the wavelength of laser available. Primary antibodies commonly used for immunostaining purposes are listed in Table 12.2. Follow the steps for primary and secondary antibody incubation as described in Steps 6, 7 of Section 3.2.2. Especially for nuclear translocation purposes, muscle tissues can be stained with a fluorescent secondary antibody (FITC or Texas Red). Next, the sections are counterstained with DAPI (Molecular Probes, OR) to visualize the nuclei (blue staining). The specificity of the staining can be confirmed by blocking antibody reactivity with specific peptide or isotype controls. Mount slides using Vectashield medium, place the cover slip (avoid air bubbles), and store the slides in a dark place. Digital pictures of stained sections can be taken using an appropriate fluorescent microscope (Nikon Eclipse E800, Japan) fitted with spot digital color camera with SPOT advanced software (Diagnostic instruments). These pictures can be used for further quantification.

3.4. Interpretation of immunostaining results Immunostaining of ER chaperones (e.g., calnexin, calreticulin, ERP72, Bip/Grp78, and GRP94) can be useful in determining ER stress within the individual cell types in a muscle biopsy (Vattemi et al., 2004; Vitadello et al., 2010). Other ER stress markers such as Caspase-12 and SERCA2 have also been shown by immunolocalization in the affected muscle sections (Ikezoe et al., 2009).

4. Determination of mRNA Levels of ER Stress Targets ER stress can be monitored at the mRNA level because expression of genes that orchestrate stress response will be significantly increased. These genes will typically have ER stress response elements (ERSE-I, ERSE-II) and unfolded protein response elements (UPRE) in their promoters. Targets commonly assessed at the mRNA level using reverse transcriptase polymerase chain reaction (RT-PCR) include GRP78, GRP94, calreticulin, Homocysteine-induced ER protein (HERP), ERP72, tryptophanyltRNA synthetase (WARS), and ER degradation enhancer mannosidase alpha-like 1 (EDEM-1). Required materials and reagents Commercially available RNeasy kit (Qiagen, Valencia, CA), liquid nitrogen, TRIzol reagent (Life Technologies, Gaithersberg, MD), SYBRÒ

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Green PCR Master Mix (Applied Biosystems, Carlsbad, CA), Nanodrop (Thermo Scientific, Wilmington, DE), Gel documentation instrument (Syngene, Frederick, MD), Gel analysis software (Genetools) (Syngene).

4.1. Isolation of RNA from frozen muscle tissues 4.1.1. Method 1. Total RNA is extracted from snap-frozen muscle biopsies or tissue samples ( 100 mg) using TRIzol reagent (Life Technologies, Carlsbad, CA) or similar RNA extraction reagent and protocol. 2. RNA can be purified using RNeasy Mini kit (Qiagen) by following the manufacturer’s instructions. The purity of the total RNA should be confirmed spectrophotometrically at 260 nm. Concentration of the RNA can be measured using the instrument Nanodrop (Thermo Scientific).

4.2. Semiquantitative reverse transcriptase PCR 1. cDNA is prepared from a constant amount of total RNA for each individual sample (0.5–5 mg) with random primers and reverse transcriptase system (Promega, Madison, WI). 2. Primers for the selected target genes can be generated using commercial suppliers such as Invitrogen (www.Invitrogen.com). 3. PCR is performed with primers directed to the gene of interest, under cycling conditions optimized to the target gene. 4. PCR products can be resolved by standard agarose (2%) gel electrophoresis and can be observed with appropriate molecular weight standards using gel documentation instrument (Syngene). Further, these bands can be quantified using gel analysis software (Genetools) (Syngene).

4.3. Interpretation of RT-PCR data The relative transcript abundance can be determined by normalizing the expression values obtained for the transcript of interest to GAPDH expression. ER stress activation can also be assessed by measuring the splicing of XBP-1 mRNA. For example, unfolded protein response initiates oligomerization of IRE-1 and trans-autophosphorylation of the kinase domain of IRE1 which activates splicing of the mRNA that encodes XBP-1 transcription factor. The splicing of XBP1 mRNA can be assessed semiquantitatively by RT-PCR using XBP1 specific primers that detect both unspliced and spliced isoforms under standard conditions (Table 12.3).

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Table 12.3 Identification of XBP1 using RT-PCR; primers used and the sizes of the amplified PCR products were listed belowa Sizeb (bp) Transcript

Forward primera

Reverse primera

USc

Sd

Human

50 TTACGAGAGAAA ACTCATGGCC 30 0 5 GAACCAGGAGTT AAGAACACG 30

50 GGGTCCAAGTTG TCCAGAATGC 30 0 5 AGGCAACAGTG TCAGAGTCC 30

289

263

205

179

Mouse a b c d

The primers listed were obtained from Samali et al. (2010). Expected size (bp) of the PCR products. US: unspliced XBP1 transcript. S: spliced XBP1 transcript.

5. Concluding Remarks The information presented in this chapter is designed to establish a basic series of protocols to use when designing studies of ER stress in myositis muscle, including protein expression, histology, and intracellular localization of antigens. Clearly, additional techniques will become available as scientific breakthroughs advance the field of myositis research. Using the techniques described in the current chapter, researchers have been able to begin to dissect the elegant involvement of ER stress in autoimmune myositis. In future, it might be possible to use a transgenic mouse model to obtain insight into the mechanisms involved in ER stress and the pathogenesis of myositis (Iwawaki et al., 2004). These mechanistic studies could lead to exciting information and advancements about myositis and other devastating muscle disorders. However, readers are advised to thoroughly read the background publications that are references in the current chapter and related chapters for a clear understanding of the role of ER stress and UPR in myositis.

ACKNOWLEDGMENTS Dr. Nagaraju is supported by NIH grant R01-AR050478 (K. Nagaraju), NIH grant 1U54HD053177-01A1 (Hoffman). Also, the authors are thankful to Mr. William Coley for his help in editing this chapter.

REFERENCES Askanas, V., and Engel, W. K. (2003). Unfolding story of inclusion-body myositis and myopathies: Role of misfolded proteins, amyloid-beta, cholesterol, and aging. J. Child Neurol. 18, 185–190.

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Devries-Seimon, T., et al. (2005). Cholesterol-induced macrophage apoptosis requires ER stress pathways and engagement of the type A scavenger receptor. J. Cell Biol. 171, 61–73. Emslie-Smith, A. M., et al. (1989). Major histocompatibility complex class I antigen expression, immunolocalization of interferon subtypes, and T cell-mediated cytotoxicity in myopathies. Hum. Pathol. 20, 224–231. Flores-Diaz, M., et al. (2004). A cellular UDP-glucose deficiency causes overexpression of glucose/oxygen-regulated proteins independent of the endoplasmic reticulum stress elements. J. Biol. Chem. 279, 21724–21731. Henriques-Pons, A., and Nagaraju, K. (2009). Nonimmune mechanisms of muscle damage in myositis: Role of the endoplasmic reticulum stress response and autophagy in the disease pathogenesis. Curr. Opin. Rheumatol. 21, 581–587. Hotamisligil, G. S., and Erbay, E. (2008). Nutrient sensing and inflammation in metabolic diseases. Nat. Rev. Immunol. 8, 923–934. Ikezoe, K., et al. (2009). Amyloid-beta accumulation caused by chloroquine injections precedes ER stress and autophagosome formation in rat skeletal muscle. Acta Neuropathol. 117, 575–582. Iwawaki, T., et al. (2004). A transgenic mouse model for monitoring endoplasmic reticulum stress. Nat. Med. 10, 98–102. Kalai, M., et al. (2003). Regulation of the expression and processing of caspase-12. J. Cell Biol. 162, 457–467. Kaufman, R. J. (1999). Stress signaling from the lumen of the endoplasmic reticulum: Coordination of gene transcriptional and translational controls. Genes Dev. 13, 1211–1233. Kedi, X., et al. (2009). Free cholesterol overloading induced smooth muscle cells death and activated both ER- and mitochondrial-dependent death pathway. Atherosclerosis 207, 123–130. Khan, S. U., and Schroder, M. (2008). Engineering of chaperone systems and of the unfolded protein response. Cytotechnology 57, 207–231. Li, C. K., et al. (2009). Overexpression of MHC Class I Heavy Chain Protein in Young Skeletal Muscle Leads to Severe Myositis. Implications for Juvenile Myositis. Am. J. Pathol. Liu, H., et al. (1997). Endoplasmic reticulum chaperones GRP78 and calreticulin prevent oxidative stress, Ca2þ disturbances, and cell death in renal epithelial cells. J. Biol. Chem. 272, 21751–21759. Malhotra, J. D., and Kaufman, R. J. (2007). The endoplasmic reticulum and the unfolded protein response. Semin. Cell Dev. Biol. 18, 716–731. Matsumoto, M., et al. (1996). Ectopic expression of CHOP (GADD153) induces apoptosis in M1 myeloblastic leukemia cells. FEBS Lett. 395, 143–147. Mori, K. (2000). Tripartite management of unfolded proteins in the endoplasmic reticulum. Cell 101, 451–454. Nagaraju, K., et al. (2000). Conditional up-regulation of MHC class I in skeletal muscle leads to self-sustaining autoimmune myositis and myositis-specific autoantibodies. Proc. Natl. Acad. Sci. USA 97, 9209–9214. Nagaraju, K. (2005). Role of major histocompatibility complex class I molecules in autoimmune myositis. Curr. Opin. Rheumatol. 17, 725–730. Nagaraju, K., et al. (2005). Activation of the endoplasmic reticulum stress response in autoimmune myositis: Potential role in muscle fiber damage and dysfunction. Arthritis Rheum. 52, 1824–1835. Nakagawa, T., et al. (2000). Caspase-12 mediates endoplasmic-reticulum-specific apoptosis and cytotoxicity by amyloid-beta. Nature 403, 98–103.


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Nogalska, A., et al. (2007). Endoplasmic reticulum stress induces myostatin precursor protein and NF-kappaB in cultured human muscle fibers: Relevance to inclusion body myositis. Exp. Neurol. 204, 610–618. Samali, A., et al. (2010). Methods for monitoring endoplasmic reticulum stress and the unfolded protein response. Int. J. Cell Biol. 830307, 2010, 1–11. Schroder, M. (2008). Endoplasmic reticulum stress responses. Cell. Mol. Life Sci. 65, 862–894. Schroder, M., and Kaufman, R. J. (2005). ER stress and the unfolded protein response. Mutat. Res. 569, 29–63. Vattemi, G., et al. (2004). Endoplasmic reticulum stress and unfolded protein response in inclusion body myositis muscle. Am. J. Pathol. 164, 1–7. Vitadello, M., et al. (2010). Myofiber stress-response in myositis: Parallel investigations on patients and experimental animal models of muscle regeneration and systemic inflammation. Arthritis Res. Ther. 12, R52.


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Activation of the Unfolded Protein Response in Human Acute Myeloid Leukemia Julian A. Schardt,* Beatrice U. Mueller,† and Thomas Pabst* Contents 1. Introduction 2. Technical Aspects 2.1. Isolation of leukemic cells from whole blood samples and bone marrow aspirates 2.2. Methods for the detection of the activated UPR in AML patient samples 2.3. Induction of ER-stress in human leukemic cell lines in vitro 2.4. Construction of a calreticulin-inducible leukemic cell line 3. Discussion 3.1. General considerations 3.2. Which event(s) trigger the UPR in leukemic cells? 3.3. AML patients with activated UPR represent a distinct subtype of AML with specific clinical characteristics 3.4. Role of the UPR during leukemogenesis 3.5. Future aspects References

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Abstract There is accumulating evidence for the involvement of the unfolded protein response (UPR) in the pathogenesis of many tumor types in humans. This is particularly the case in rapidly growing solid tumors in which the demand for oxygen and nutrients can exceed the supply until new tumor-initiated blood vessels are formed. In contrast, the role of the UPR during leukemogenesis remains largely unknown. Acute myeloid leukemia (AML) is a genetically heterogeneous clonal disorder characterized by the accumulation of somatic mutations in hematopoietic progenitor cells that alter the physiological regulation of self-renewal, survival, proliferation, or differentiation. The CCAAT/ * Department of Medical Oncology, University Hospital, Bern, Switzerland { Department of Internal Medicine, University Hospital, Bern, Switzerland Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00013-3

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enhancer-binding protein alpha (CEBPA) gene is a key myeloid transcription factor and a frequent target for disruption in AML. In particular, translation of CEBPA mRNA can be specifically blocked by binding of the chaperone calreticulin (CALR), a well-established effector of the UPR, to a stem loop structure within the 50 region of the CEBPA mRNA. The relevance of this mechanism was first elucidated in certain AML subtypes carrying the gene rearrangements t(3;21) or inv(16). In our recent work, we could demonstrate the induction of key effectors of the UPR in leukemic cells of AML patients comprising all subtypes (according to the French-American-British (FAB) classification for human AML). The formation of the spliced variant of the X-box binding protein (XBP1s) was detectable in 17.4% (17 of 105) of AML patients. Consistent with an activated UPR, this group had significantly increased expression of the UPR target genes CALR, the 78 kDa glucose-regulated protein (GRP78), and the CCAAT/enhancer-binding protein homologous protein (CHOP). Consistently, in vitro studies confirmed that calreticulin expression was upregulated via activation of the ATF6 pathway in myeloid leukemic cells. As a consequence, CEBPA protein expression was inhibited in vitro as well as in leukemic cells from patients with activated UPR. We therefore propose a model of the UPR being involved in leukemogenesis through induction of calreticulin along the ATF6 pathway, thereby ultimately suppressing CEBPA translation and contributing to the block in myeloid differentiation and cell-cycle deregulation which represent key features of the leukemic phenotype. From a more clinical point of view, the presence of activated UPR in AML patient samples was found to be associated with a favorable disease course.

1. Introduction The unfolded protein response (UPR) is triggered by the accumulation of misfolded proteins within the endoplasmic reticulum (ER). ERstress can also be caused by environmental perturbations, for example, starvation, hypoxia, viral infections, and heat (Malhotra and Kaufman, 2007; Schroder and Kaufman, 2005a). Physiologic conditions such as increased immunoglobulin production in mature B cells also activate the UPR (Iwakoshi et al., 2003). Only correctly folded proteins are exported to the Golgi complex while incompletely folded proteins are retained in the ER in order to complete the folding process or to be targeted to degradation via the ER-associated protein degradation pathway (ERAD) (Hebert and Molinari, 2007). There are at least three classes of stress sensors residing in the ER to initiate UPR signaling cascades: inositol requiring kinase 1 (IRE1), activating transcription factor 6 (ATF6), and the double-stranded RNA-activated protein kinase (PKR)-like endoplasmic reticulum kinase (PERK). By controlling

The Unfolded Protein Response in AML

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the expression of a set of transcription factors, each of the stress sensors is transmitting information about protein folding status from the ER to the nucleus to initiate global changes in gene expression and protein translation in order to restore cellular homeostasis (Malhotra and Kaufman, 2007). Prolonged activation of the UPR results in the activation of apoptotic pathways (Kim et al., 2008). Recently, there is accumulating evidence for ER-stress being involved in the pathogenesis of a wide range of human diseases like neurodegenerative diseases (e.g., Alzheimer’s disease), diabetes type 1 and 2, cardiovascular diseases (e.g., atherosclerosis or heart failure), immune disorders (e.g., rheumatoid arthritis or inflammatory bowel diseases), and cancer (Hosoi and Ozawa, 2009). The involvement of ER-stress and the UPR in malignant transformation and tumor progression is primarily based on studies in solid tumors. Due to rapid growth and inadequate vascularization, the microenvironment challenges tumor cells. These conditions include decreased levels of oxygen, glucose, and nutrients as well as of intracellular calcium, all of which are detrimental to cell growth and survival (Ma and Hendershot, 2004). For example, the expression of the spliced form of X-box binding protein 1 mRNA (XBP1s) and the nuclear form of activating transcription factor 6 (ATF6n), as well as many classical ER-stress-inducible chaperones, such as 94 kDa glucose-regulated protein (grp94) and 78 kDa glucose-regulated protein (GRP78), can be increased in breast, lung, liver, and colon cancers (Moenner et al., 2007). Direct evidence that the UPR is necessary for tumor growth particularly under hypoxic conditions came from studies targeting the IRE1/XBP1 pathway: first, XBP1-deficient mouse embryonic fibroblasts (MEFs) are selectively impaired for survival following exposure to hypoxia. Second, transformed MEFs as well as the human fibrosarcoma cells (HT1080), which are deficient for XBP1, were unable to grow when transplanted as tumor xenografts in SCID mice. Furthermore, hypoxic conditions activate the PERK pathway and its downstream target eIF2alpha (Fels and Koumenis, 2006). Finally, IRE1 is a critical determinant linking hypoxia and hypoglycemiadependent responses to the induction of vascular endothelial growth factor-A (VEGF-A), indicating an important role of the IRE1 pathway for both angiogenesis and tumor development (Drogat et al., 2007). First evidence for the UPR being involved in hematologic malignancies arose from studies in patients with multiple myeloma. Multiple myeloma constitutes a multifocal plasma cell neoplasm characterized by serum monoclonal gammopathy and focal skeletal lesions. Carrasco et al. (2007) could demonstrate a shift toward increased expression of the spliced form of XBP1 in CD138þ primary myeloma cells in the majority of patients compared to normal plasma cells. A direct role of XBP1s in the pathogenesis of multiple myeloma was uncovered in a transgenic mouse model in which XBP1s expression is under the control of the immunoglobulin VH promoter and Em enhancer element (Carrasco et al., 2007). Em regulatory elements drive

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gene expression throughout B-cell development, as well as in some T cells. In this mouse model, enforced expression of XBP1s in the B-cell compartment enhanced B-cell proliferation and activated known myeloma-relevant pathways, leading (after the transition through a premalignant stadium termed MGUS) to the myeloma phenotype. The first study addressing the role of the UPR in leukemogenesis was performed in the human leukemic cell line (NB4) harboring the t(15;17) translocation which represents the cytogenetic hallmark of acute promyelocytic leukemia (APL) and encodes the oncogenic fusion protein promyleocytic leukemia-retinoic acid receptor alpha (PML-RARA). Khan et al. (2004) could demonstrate that PML-RARA induces the accumulation of insoluble N-CoR protein primarily in the ER which leads to decreased levels of soluble N-CoR protein in the nucleus and subsequent activation of the UPR determined by the generation of the nuclear form of ATF6 (ATF6n). It was suggested that tight binding of N-CoR protein to PML-RARA protein generates an aberrant insoluble N-CoR protein and recruits the molecule to the ER for ER-associated degradation (ERAD). Since N-CoR protein in the nucleus is vital for the transcriptional repression mediated by the tumor suppressor max dimerization protein 1 (MXD1), the authors speculated that this reduction of soluble N-CoR protein levels in the nucleus may be responsible for deregulated proliferation and transformation in the pathogenesis of APL. The cornerstone of current APL treatment is the use of all-trans retinoic acid (ATRA) in combination with cytotoxic chemotherapy. ATRA mediates terminal differentiation of promyelocytes to mature neutrophils. Interestingly, no nuclear form of ATF6 could be detected in NB4 cells treated with ATRA and the solubility of the N-CoR protein was increased by ATRA treatment. The general perception is that prolonged ER-stress leads to induction of apoptosis (Ng et al., 2006). In APL cells, the toxic effect of prolonged activation of the UPR is neutralized through selective expression of the glycoprotease O-sialoglycoprotein endopeptidase (OSGEP) which processes misfolded N-CoR protein, thereby regulating the response of APL cells to UPR-induced apoptosis. The question whether OSGEP is a direct target of the UPR remains elusive, and a causative role of the UPR in the pathogenesis of APL remains to be determined. Furthermore, Tanimura et al. (2009) demonstrated a link between the leukemic fusion protein breakpoint cluster region-c-abl oncogene 1 receptor tyrosine kinase (Bcr-Abl) and concomitant activation of the UPR. The leukemogenic fusion protein Bcr-Abl generated by the translocation t(9;22) is crucial for the pathogenesis of chronic myeloid leukemia and Bcr-Abl positive acute leukemia. The investigators demonstrated increased expression of the UPR target genes GRP78 in kursiveletters and of the UPR effector molecule XBP1S on mRNA and protein level in murine myeloid cells stably


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transfected with the leukemic fusion gene Bcr-Abl as well as on mRNA level in Bcr-Abl-positive acute leukemia cells derived from patients. In addition, targeting the IRE1- and ATF6-pathway of the UPR by dominant negative mutants increased drug-induced apoptosis (drugs used in these experiments were etoposide and imatinib, a nonselective inhibitor of the Abl tyrosine kinase), providing evidence that ER-stress sensors might modulate the ability of Bcr-Abl to protect leukemic cells from apoptosis. Acute myeloid leukemia (AML) is a genetically heterogeneous, clonal disorder characterized by the accumulation of somatic mutations in hematopoietic progenitor cells that alter the physiological regulation of selfrenewal, survival, proliferation, and/or differentiation. There is increasing evidence that a rare cell fraction, the so-called leukemia initiating cells (LICs), also referred as leukemia stem cells (LCS), give rise to the bulk of leukemic blasts detected in the blood and bone marrow of affected individuals (Chan and Huntly, 2008). For mature myeloid cells, there is a stepwise requirement of specific transcription factors during myeloid differentiation from stem cells toward the monocytic or the neutrophil lineage. One of these key transcription factors essential for myelopoiesis is the transcription factor CCAAT/enhancer binding factor alpha (CEBPA) (Rosenbauer and Tenen, 2007). CEBPA is a multifunctional protein which regulates the expression of many myeloid-specific genes, but it is also an inhibitor of cell proliferation by coordinating the exit from the cell cycle, thereby regulating two essential features of terminally differentiated cells ( Johnson, 2005). During myelopoiesis, CEBPA is important for the maintenance of the long-term hematopoietic stem cell pool and the transition from common myeloid progenitors GMPs toward granulopoiesis. Underlining its important role in myelopoiesis, CEBPA was found to be a frequent target for disruption in AML. For instance, CEBPA is mutated in 5–14% of AML patients predominantly in the M1 and M2 subtype of AML (according to the FAB classification). Its transcription can be suppressed by promoter methylation and through the RUNX1–MTG8 fusion product which arises from the t(8;21) translocation. On the posttranscriptional/translational level, inhibition of translation by mRNA-binding proteins (like RNA-binding protein poly(rC) binding protein 2 (PCBP2)), various protein modifications (phosphorylation and SUMOylation) as well as direct protein–protein interactions that alter/inhibit proper CEBPA function have been described (Pabst and Mueller, 2009). In a search for RNA-binding proteins that interact with GC-rich RNAs, calreticulin was identified to interact with GCN repeats within CEBPA and CEBPB mRNAs. GCN repeats within the CEBP mRNAs form stable stem loop structures. In the 50 region of CEBPA mRNA, two putative stem loop structures were identified. Binding of calreticulin to the

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more internal stem loop repressed translation of the CEBPA protein in vitro and in vivo (Timchenko et al., 2002). First evidence for a role of this posttranscriptional mechanism during leukemogenesis came from the following observations: Primary leukemic cells from patients carrying the t(3;21) or inv(16) translocation, leading to the expression of the fusion products AML1-MDS1-EVI1 and CBFBSMMHC, respectively, showed induced calreticulin expression and increased binding of calreticulin to the CEBPA mRNA (Helbling et al., 2004; 2005). As a consequence, CEBPA protein was dramatically suppressed in affected individuals. In a leukemic cell line model, conditional expression of the fusion gene CBFB-SMMHC led to induced expression of calreticulin, increased binding of calreticulin to the CEBPA mRNA, and consecutive suppression of CEBPA protein translation. In vitro, conditional expression of the fusion gene CBFB-SMMHC resulted in a block in differentiation when incubating the cells with differentiating agents like ATRA. Blocking CBFB-SMMHC induced calreticulin expression (using siRNA specifically directed against calreticulin) restored the differentiation potential of leukemic cells despite expression of the fusion gene CBFBSMMHC. These data implement calreticulin-mediated suppression of CEBPA as a mechanism leading to a differentiation block in leukemic cells. Calreticulin is a ubiquitously expressed multifunctional protein, located predominantly in the ER lumen of the cell. It belongs to the lectine family of chaperone molecules and is important for the maintenance of intracellular calcium homoeostasis. Calreticulin is a component of the calnexin/calreticulin cycle within the ER lumen which is responsible for the N-linked glycosylation of proteins along the secretory pathway. This calnexin/calreticulin cycle constitutes one arm of the quality control machinery in the ER that monitors protein conformations and dictates whether a molecule is exported to the Golgi complex or targeted for ERAD via the proteasomal pathway (Schroder and Kaufman, 2005b). As a chaperone molecule calreticulin is a well known target gene of the UPR. In the following chapter, we will discuss the role of the UPR in AML, with particular emphasis on the methodological and technical aspects of our work.

2. Technical Aspects 2.1. Isolation of leukemic cells from whole blood samples and bone marrow aspirates Mononuclear cells (lymphocytes, monocytes, and leukemic cells) have a lower buoyant density than erythrocytes and the polymorphonuclear leukocytes (PMN). Most mononuclear cells have densities below 1.077 g/ml. These cells can therefore be isolated by centrifugation on an iso-osmotic

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medium with a density of 1.077 g/ml, which allows the PMNs and erythrocytes to sediment through the medium while retaining the mononuclear cells at the sample–medium interface. The contamination of erythrocytes in the mononuclear cell suspension is usually less than 10% of the total cell number. The centrifugation steps in the following protocol were carried out on a centrifuge 5810 R (Eppendorf, Hamburg, Germany). EDTA blood sample in 10-ml tubes are centrifuged at 1800 rpm for 10 min at room temperature to separate the cellular elements from the plasma (Fig. 13.1). The supernatant is discarded; the cells are diluted in phosphatebuffered saline (PBS) and transferred to a 50-ml falcon tube containing PBS to a maximum of 40 ml. In the next step, 10-ml blood cells/PBS mix are transferred to a 15-ml falcon provided with 5 ml of LymphoprepTM solution prewarmed at room temperature. Caution should be made that the blood cells/PBS suspension does not mix with the LymphoprepTM solution.

+ PBS

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EDTA blood

10 min, 1800 ⫻ g, at RT

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Discard plasma

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20 min, 450 ⫻ g, at RT Mononuclear cells (MC) Erythrocytes + granulocytes

LymphoprepTM displaced upward

Figure 13.1 Isolation of mononuclear cells from whole blood samples and bone marrow aspirates from AML patients. (A) Cellular blood elements are isolated from whole blood samples (supplemented with ethylenediaminetetraacetate) through centrifugation and are resuspended in phosphate-buffered saline (PBS). (B) Mononuclear cells (MC) can be separated from erythrocytes and granulocytes by centrifugation on the iso-osmotic LymphoprepTM which allows the polymorphonuclear cells (PMN) and erythrocytes to sediment through the medium while retaining the mononuclear cells at the sample–medium interface.

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The 15-ml falcons are centrifuged for 20 min at 450 rpm g. The mononucleated cells accumulate at the medium–sample interface (Fig. 13.1) and are harvested with a 5-ml pipette. The collected cells are transferred to a new 50-ml falcon provided with 30-ml PBS. The cells are washed once in 30-ml PBS, centrifuged at 340g for 10 min at room temperature, and resuspended in 5–10-ml PBS and counted in a cell chamber. Cell pellets are stored at 80 C until further analysis. For the isolation of mononucleated cells from bone marrow aspirates, the protocol starts at the step where cells are mixed in PBS and dispersed on two to four 15-ml falcon tubes provided with 5 ml of LymphoprepTM solution prewarmed at room temperature.

2.2. Methods for the detection of the activated UPR in AML patient samples 2.2.1. Measuring activation of the IRE1 pathway In response to the occurrence of unfolded proteins, IRE1 oligomerizes in the plane of the membrane, allowing trans- and autophosphorylation of juxtaposed kinase domains. Oligomerization can be triggered directly by binding of unfolded proteins to the IRE1 lumenal domain, or might involve the release of oligomerization-repressing chaperons. The only known substrate of the IRE1 kinase is IRE1 itself. Trans- and autophosphorylation of the kinase domain of IRE1 causes the precise endonucleolytic cleavage of the mRNA that encodes the X-box binding protein-1 (XBP1). IRE1 cuts the precursor XBP1 mRNA twice, excising a fragment of 21 base pairs (bps). The 50 and 30 mRNA fragments are then ligated, generating a spliced mRNA that encodes a potent transcriptional activator of UPR target genes (Yoshida et al., 2001; Zhou et al., 2006). In our work, total mRNA from primary leukemic cells from AML patients was extracted and transcribed to cDNA. We used 100-ng cDNA as a template in the PCR reaction. Using an oligonucleotide primer pair which spans the region of the XBP1 mRNA (Yoshida et al., 2001) excised by IRE1alpha, two different PCR products (442 bp for the unspliced form and 416 pb for the spliced form of XBP1 mRNA) were generated. The two PCR bands can then be separated in a two-dimensional agarose gel electrophoresis and visualized after ethidium bromide staining on a UV-light screen (Fig. 13.2). The primers for detecting human XBP1U/XBP1S cDNA were Forward primer: 50 -CCTGGTAGTTGAGAACCAGG-30 Reverse primer: 50 -GGGGCTTGGTATATATGTGG-30 The PCR reaction was prepared in a total volume of 30 ml using 100 ng from the generated patient cDNA as a template, with a final primer concentration of 0.4 mM each, dNTPs 1 mM each, glycerol 10%, H2O, and 10 PCR reaction buffer (Roche Molecular Diagnostics, Pleasanton, USA),

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1 Taq DNA Polymerase (Roche Molecular Diagnostics, Pleasanton, USA), 1 U for each reaction, and was run on a MJ Mini Personal thermal cycler (Bio-Rad Laboratories, Munich, Germany) at the following conditions: first step at 95 C for 2 min, second step at 95 C for 15 s, at 58 C for 30 s, and at 72 C for 30 s repeated 37 times, with a final third step at 72 C for 7 min for extension. PCR bands were visualized by two-dimensional agarose gel electrophoresis (Fig. 13.2). In order to obtain a more quantitative determination of the IRE1 splicing activity, we applied an analog scale using differing concentrations of cloned XBP1U and XBP1S as a template to make the determination observer-independent (Fig. 13.3A). We arbitrarily set the ratio of intensities S(–) AML −

#1−

#2−

S(+) AML

#3−

#4−

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#2+

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#4+ 442 bp(U)

XBP1

416 bp(S) ABL

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Figure 13.2 The IRE1-pathway is activated in leukemic cells and leads to the expression of XBP1(S). Representative examples of AML patients are depicted, illustrating the results of the semiquantitative RT-PCR assay detecting only the unspliced (U) form of XBP1 mRNA (S() left panel), or both the spliced (S) and unspliced form in S(þ) AML patients (right panel) (Schardt et al., 2009a,b).

Sensitivity assay A

50 0

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Figure 13.3 Determining the sensitivity and development of an analog scale for the detection and confirmation of IRE1 splicing activity. (A) The sensitivity of the PCR assay was determined using pCDNA3 expression plasmids encoding for the human XBP1U and XBP1S form. Competitive PCR of decreasing amounts of plasmid encoding for the XBP1U and increasing amounts of plasmid encoding for the spliced form XBP1S are depicted. (B) Competitive PCR with constant amounts of plasmid encoding for XBP1U combined with increasing amounts of the spliced form of XBP1S. The ratio observed with 30 ng of XBP1U plasmid and 20 ng of the spliced form of XBP1 was determined to separate AML patients with induced versus uninduced XBP1s mRNA (Schardt et al., 2009a,b).

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of the PCR products generated from 30 ng of cloned XBP1U cDNA, together with 20 ng of cloned XBP1S cDNA after 37 PCR cycles as the minimum ratio of XBP1U/XBP1S by which a patient sample was classified as XBP1S positive, thereby indicating activation of the IRE1 pathway. We could not detect a preferential amplification of one of the cloned cDNAs for XBP1U or XBP1S in the PCR reaction. Instead, we observed that the intensities of the XBP1U and XBP1S PCR products in the agarose gel electrophoresis assay were not independent of each other: Using a fixed amount of cloned XBP1U cDNA (30 ng) and an increasing amount of cloned XBP1S cDNA in the same PCR reaction, the intensity of the XBP1U band was becoming weaker and vice versa, indicating competition (Fig. 13.3B), since we measured both isoforms in the same reaction with consecutive shortage of the oligonucleotide primers and/or desoxynucleotides in the PCR mix. Therefore, sufficient concentrations of these reaction components are recommended. To assess the specificity of our test, we evaluated peripheral blood samples from 10 healthy volunteers, prepared as described above. Although the importance of XBP1S has been well established during B-cell maturation and has been reported to be physiologically expressed in this cell type, we would not expect to detect any UPR activity due to the low number of B-lymphocytes among mononucleated cells in the peripheral blood. Based on our patient results above, we expected at least one out of the 10 samples to be positive for XBP1S expression. However, we found that the spliced form of XBP1 mRNA was not detectable in any samples from healthy volunteers. Therefore, we suggest that the detection of the XBP1 spliced form in AML patients constitutes a characteristic feature of the specific leukemic cells. Recently, Davies et al. (2008) reported a different approach in order to apply a quantitative assay to assess activation of the UPR. They defined non-splice-specific flanking primers as well as splice-specific Taqman probes to identify each isoform separately. Their assay was validated on cloned cDNAs and quantification was performed using standard curves of diluted cloned amplicons. Input RNA was corrected to mean values of the control genes (HPRT and GAPDH). Nonsplice-specific flanking primers were Forward primer: 50 -AAGCCAAGGGGAATGAAGT-30 Reverse primer: 50 -CCAGAATGCCCAACAGGATA-30 Splice-specific Taqman probes detecting each isoform separately were XBP1U: 50 -Yakima Yellow-AGCACTCAGACTACGTGCACCTBlackHoleQuencher130 XBP1S: 50 -6FAM-GCTGAGTCCGCAGCAGGTGCAG-BlackHoleQuencher1-30 Using this approach, the ratio of XBP1U to XBP1S reflecting the IRE1 splicing activity could be determined more accurately.


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2.2.2. Determination of UPR effector gene expression in primary leukemic cells To confirm the presence of an activated UPR in primary leukemic cells, we measured the relative expression levels of known UPR target genes such as GRP78, CALR, and CHOP. Quantitative RT-PCR for the assessment of relative gene expression was carried out on ABI PRISM 7700 Sequence Detection System (Applied Biosystems, Foster City, USA) using Taqman PCR Mix (Roche, Branchburf, USA) and assay-on-demand gene expression probes (Applied Biosystems, Foster City, USA). The relative quantification was indicated by CT values based on duplicate reactions for each target gene. As control genes, we measured the expression levels of GAPDH and ABL1 using the following oligonucleotide primers/ probes: For GAPDH detection, the primers had the following sequence: GAPDH (sense) 50 -GAAGGTGAAGGTCGGAGT 30 , GAPDH (antisense) 50 -GAAGATGGTGATGGGGATTT 30 , GAPDH Probe 50 -FAM-CAAGCTTCCCGTTCTCAGCC-TAMRA-30 (Eurogentec, Seraing, Belgium). For ABL1 detection, the primers had the following sequence: ABL1 (sense) 50 -TGGAGATAACACTCTAAGCATAACTAAAGGT-30 , ABL1 (antisense) 50 -GATGTAGTTGCTGGGACCCA-30 , ABL1 Probe 50 - FAM-CCATTTTTGGTTTGGGCTTCACACCATTAMRA-30 .

2.2.3. Analysis of quantitative RT-PCR data The relative quantization was given by Ct values, determined in duplicate reactions for each target and internal control gene. The mean value of duplicate Ct values was calculated and subtracted to obtain DCt (DCt ¼ Ct (target gene)Ct (control gene)). The relative expression level was determined as 2 DDCt, where DDCt is DCt (target sample)D Ct (reference sample). For the reference sample, DDCt equals zero and 2 0 equals one by definition. For the other samples, evaluation of 2 DDCt indicates the fold change in gene expression relative to the reference sample (Livak and Schmittgen, 2001). The relative expression levels of the UPR effector molecules in patient samples were finally normalized to the mean relative expression level of a given gene in samples from patients suffering from AML of the M0 subtype (according to the French-American-British (FAB) classification). The results were finally presented as a box plot diagram in a log 2 scale.

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2.3. Induction of ER-stress in human leukemic cell lines in vitro We used three different chemical compounds such as thapsigargin, A23187, and tunicamycin to induce ER-stress in our experimental settings. The chemicals were purchased from Sigma-Aldrich (St. Louis, USA). Thapsigargin (final concentration in the medium 300 nM), the calcium ionophore A23187 (final concentration in the medium 7 mM), or tunicamycin (final concentration in the medium 5 mg/ml) were added to the culture medium of K562, U937, and HL60 leukemic cells. We incubated the different cell lines for 4–24 h before harvesting the cells for further analysis. Thapsigargin and the calcium ionophore A23187 mediate ER-stress through depletion of calcium from the ER lumen and consecutive accumulation of unfolded or misfolded proteins in the ER. Thapsigargin specifically inhibits the ER Ca2þ-ATPase, while the calcium ionophore A23187 mediates the efflux of Ca2þ from the endoplasmic lumen. Tunicamycin is an antibiotic which blocks the initial step of glycoprotein synthesis thus inhibiting the synthesis of N-linked glycoproteins. All these compounds induce a robust UPR in leukemic cell lines as measured by induction of the XBP1S form and increased expression levels of the effector molecules GRP78, CALR, and CHOP by quantitative RT-PCR as described above.

2.4. Construction of a calreticulin-inducible leukemic cell line To study the effect of increased calreticulin expression on human leukemic cells in vitro, we established a conditional cell line model using the tet-off system. Thus, calreticulin expression was under the control of a tetracycline-responsive promoter element. The myeloid U937-T cell line was kindly provided by Gerard Grosveld (Memphis, USA). These cells are stably transfected with the tetracycline transactivator (tTA) under the control of a tetracycline (tet)-responsive element (TRE). To allow conditional expression of calreticulin, a 4.0-kb Sca1-BamH1 fragment of the pcDNA3 vector, containing the neomycin resistance gene, was ligated with the 0.95-kb fragment of the tet-off response plasmid pTRE to generate the pTREneo vector. A 1.4-kb EcoR1-Xho1 fragment encoding the hemagglutinine (HA)-tagged human full-length calreticulin cDNA was then introduced into the pTRE-neo plasmid. The plasmid was linearized and stably transfected into U937-T cells by electroporation. Twenty-four single-cell clones were expanded under selection with puromycin (100 mM) and neomycin (2 mM). To test the clones for conditional calreticulin induction, the cells were washed three times in 50-mL PBS and then cultured in the absence of tetracycline. The increase of calreticulin mRNA transcripts and protein was measured by quantitative real-time PCR and Western blot.


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3. Discussion 3.1. General considerations The assessment of activation of the UPR in leukemic cells may provide challenges at multiple levels. When transducer molecules of all the three major pathways of the UPR are detectable and activated in a cell—such as detection of ATF6N, eIF2alpha-P, and XBP1S—activation of the UPR appears obvious. In contrast, a single transducer/effector molecule might be activated by mechanisms unrelated to activation of the UPR. This suggests that analyzing only a single marker of the UPR may be carefully considered to be indicative for activation of the UPR per se. Moreover, it remains unclear whether ER-stress consistently involves activation of all the three major pathways of the UPR or whether it can selectively activate single pathways depending on cell conditions and cell types. We determined the activation of all the three UPR pathways in human AML. First, activation of the IRE1 pathway was detected by assessing the XBP1s mRNA; second, activation of the ATF6 pathway of the UPR was determined by measuring known effector molecules of this pathway such as GRP78 and CALR; and third, activation of the PERK/ATF4-pathway was determined by the induction of its known target gene CHOP. Establishing a cut-off value represents a notorious problem in quantitative RT-PCR assays. In particular, defining a value indicating increased expression of a transducer or effector of UPR and thereby suggesting activation of the UPR can appear arbitrary. Therefore, we suggest that measuring transducer/effector molecules of all the three UPR main pathways may be more accurate when evaluating the role of the UPR in human diseases such as AML.

3.2. Which event(s) trigger the UPR in leukemic cells? Due to the disseminated nature of acute leukemias, environmental factors like hypoxia or nutrient starvation seem to be unlikely to cause ER-stress in leukemic cells. Therefore, cell-intrinsic factors may be more promising candidates such as mutations in genes encoding secreted proteins or in genes that are involved in protein folding and protein quality control. Some chromosomal translocations such as the t(15;17) or t(9;22) leading to the expression of the leukemic fusion proteins PML-RARA or BCRABL, respectively, can mediate activation of the UPR (Khan et al., 2004; Tanimura et al., 2009). In our study population, activated UPR was detected in 16% (17/105) of AML patients involving nearly all AML subclasses, except M5 and M7

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(according to the FAB classification) and, in particular, also including patients carrying the t(15;17) translocation (Schardt et al., 2009a,b). The presence of an activated UPR correlated with the abnormalities (numerical and structural) of chromosome 7. All patients with chromosome 7 abnormalities displayed activation of the UPR. It is tempting to speculate that chromosome 7 abnormalities might therefore be involved in the activation of the UPR in these cells. However, the genes on chromosome 7 mediating this effect still remain to be identified.

3.3. AML patients with activated UPR represent a distinct subtype of AML with specific clinical characteristics The group of AML patients with activation of the UPR comprised a higher proportion of secondary AML patients. Also, a higher white blood cell count and increased values for lactate dehydrogenase were characteristic for these patients. Furthermore, the FLT3-ITD mutation, which represents a rather frequent genetic abnormality in AML patients, (Marcucci et al., 2005) seems to be a rare event in AML patients with activated UPR. Interestingly, the presence of an activated UPR was associated with a prolonged eventfree, disease-free, and overall survival, assessed at 2 years after diagnosis. The reason for this observation remains elusive, but our observation is in line with numerous reports suggesting that the disrupted function of CEBPA, for example, through mutations such as in roughly 10% of all AML patients, represents an independent prognostic factor indicating a favorable course of the disease (Schlenk et al., 2008). In summary, the presence of an activated UPR defines a subpopulation of AML patients which is characterized by distinct clinical features as well as a favorable course of the disease.

3.4. Role of the UPR during leukemogenesis Our work summarized above suggested that activation of the UPR in primary leukemic cells leads to the suppression of CEBPA protein, a key myeloid transcription factor, through induction of the UPR effector molecule calreticulin along the ATF6 pathway of the UPR (Schardt et al., 2009a,b). Activated calreticulin binds to a stem loop structure in the 50 -region of the CEBPA mRNA and inhibits its translation. During myelopoiesis, CEBPA is important for the maintenance of the long-term hematopoietic stem cell pool and the transition from common myeloid progenitors to the bilineage granulocyte-monocyte precursor cells (GMP) and granulopoiesis (Rosenbauer and Tenen, 2007). Furthermore, CEBPA is a negative regulator of cell-cycle progression through its transcriptional control of c-Myc as well as direct binding and consecutive inhibition of CDK2/4 ( Johansen et al., 2001; Porse et al., 2001; Slomiany et al., 2000; Wang et al., 2001).


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Therefore, we propose a model in which the UPR is involved in leukemogenesis through suppression of CEBPA, ultimately leading to both a block in differentiation and cell-cycle dysregulation, both representing hallmarks of the leukemic phenotype. In this context, calreticulin possesses oncogenic potential in hematopoietic cells.

3.5. Future aspects On the molecular level, the identification of ER-stress initiating event(s) in leukemic cells would be of prominent interest. In particular, further characterization of the genes affected by specific aberrations on chromosome 7 might contribute to a better understanding of mechanisms triggering the UPR in AML cells. Also, it was suggested that prolonged or excessive activation of the UPR ultimately leads to apoptosis of affected cells. Therefore it seems quite paradoxical that an activated UPR might provide a selective advantage to cancer cells. Proceeding along the apoptotic pathway might be circumvented in tumor cells by activation of the nuclear factorkappa B (NFkB), a downstream target of the PERK pathway (Ma and Hendershot, 2004). Further studies need to be done to elucidate the mechanisms by which leukemic cells with activated UPR circumvent apoptosis. From a clinical point of view, the presence of an activated UPR might offer a promising target of novel treatment strategies in AML. As evidenced from studies in multiple myeloma, a plasma cell disorder, these cells exhibit activated UPR due to the high demand on their protein secretory pathway. Multiple myeloma cells are particularly sensitive to treatment with the proteasomal inhibitor bortezomib which induces a terminal UPR, ultimately leading to apoptosis. Therefore, our findings may provide a rationale for the use of proteasomal inhibitors in AML patients with activated UPR. In addition, one could think of ER-stress-inducing compounds given together with bortezomib as a novel treatment strategy. Currently, clinical studies are on the way to test this hypothesis.

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C H A P T E R

F O U R T E E N

Modulation of the Unfolded Protein Response by GRP78 in Prostate Cancer Gustaaf de Ridder, Rupa Ray, Uma K. Misra, and Salvatore V. Pizzo Contents 1. Introduction 1.1. The unfolded protein response (UPR) 1.2. UPR, GRP78, and disease 2. Methods for Monitoring UPR Modulation 2.1. Cancer cell lines 2.2. Media and cell culture 2.3. Western blotting for UPR signaling cascade components 2.4. Measurement of apoptosis of cells treated with anti-CTD antibodies to GRP78 2.5. Measurement of [3H]-thymidine uptake in response to antibodies against GRP78 2.6. Measurement of the activation of Aktthr308 and Aktser473 kinase 2.7. Silencing GRP78 gene expression by RNAi 3. Conclusions References

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Abstract The unfolded protein response (UPR) is an adaptive survival mechanism through which cells can weather the stress of misfolded protein accumulation induced by a wide variety of pathophysiologic and pharmacologic insults. The ER chaperone GRP78 is a central modulator of the UPR both through its proteinbinding capacity and its direct regulation of the UPR signaling molecules IRE1a, PERK, and ATF6. Recent reports have revealed the presence of GRP78 on the surface of cancer cells. Biological roles for cell-surface GRP78 include competing NH2-domain and COOH-domain agonist receptor activities that induce opposite effects on proliferation and apoptosis. Modulation of the UPR impacts both of these processes directly and indirectly. Here, we outline methods that we use to investigate UPR modulation via direct ligation of cell-surface GRP78. Specifically, we review methods of cell culture, cell-signaling analysis with Department of Pathology, Duke University Medical Center, Durham, North Carolina, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00014-5

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emphasis on UPR components, and ultimately, the impact that these have on cell proliferation, survival, and apoptosis.

1. Introduction 1.1. The unfolded protein response (UPR) The UPR is an adaptive response by the cell to overcome the accumulation of misfolded proteins in the endoplasmic reticulum (ER). In cells, proteins enter the lumen of the ER where they are modified and properly folded. However, stresses such as hypoxia, Ca2þ dysregulation, and glucose deprivation can lead to the accumulation of misfolded and unfolded proteins. This activates UPR signaling in order to restore protein homeostasis through the suppression of protein translation and subsequent reduction of protein loading into the ER, upregulation of ER chaperones to assist in folding of accumulated proteins, the degradation of misfolded proteins via ER-associated degradation (ERAD) or autophagy, or the induction of apoptosis. Upregulation of the UPR has been shown to provide a growth advantage to tumor cells (Fels and Koumenis, 2006). The ER transmembrane proteins IRE1 (inositol requiring enzyme 1), PERK (double-stranded RNA-activated protein kinase (PKR)-like ER kinase), and ATF6a (activating transcription factor 6) act as the sensors for the accumulation of unfolded proteins in the ER. These three proteins are normally in an inactive state due to their association with the amino terminal of GRP78, an ER chaperone. However, when there is an increase in the burden of misfolded proteins in the ER, GRP78 preferentially binds to the misfolded proteins, and these three sensors dissociate from GRP78 and initiate the UPR signaling cascade (Quinones et al., 2008). ER stress leads to the activation of IRE1 and the generation of the transcription factor XBP1, which is involved in UPR and ERAD regulation (Lass et al., 2008). Specifically, XBP1 upregulates chaperone expression to facilitate protein folding and genes involved with ERAD to target misfolded proteins for degradation (Yoshida et al., 2006). In addition, procaspase-12 is recruited by activated IRE1 and associates with other proapoptotic proteins such as TRAF2, BAK, and BAX (Yoneda et al., 2001). The dissociation of PERK from GRP78 upon ER stress leads to the phosphorylation of eIF2a. This prevents the initiation of translation, thus preventing the entry of nascent polypeptides into the ER. The ATF4 transcription factor is upregulated by phosphorylated eIF2a (Harding et al., 2000) and induces the expression of proapoptotic genes such as CHOP (GADD154) that suppress the expression of BCL-2, an antiapoptotic gene (Misra and Pizzo, 2005).

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ATF6 is cleaved by S1P and S2P proteases upon dissociation from GRP78. ATF6f is the resultant cytosolic domain of the cleaved protein and acts as a UPR transcriptional activator (Namba et al., 2007). It also regulates gene expression of ER chaperones, ERAD-associated proteins, and apoptotic genes (Kaneko and Nomura, 2003). Together, these three pathways work to alleviate ER stress by decreasing the amount of misfolded/unfolded proteins in the ER, or if this cannot be achieved, by initiating cell apoptosis.

1.2. UPR, GRP78, and disease UPR plays a critical role in maintaining protein homeostasis in eukaryotic cells. Dysregulation of this pathway can lead to the development of protein folding/aggregation diseases such as Alzheimer’s Dementia (Salminen et al., 2009). Additionally, studies demonstrate that genes directly involved in UPR or targets of UPR are upregulated in a number of tumor types. One of these proteins is GRP78, which is overexpressed in breast, colon, prostate, and other adenocarcinoma cell lines, as well as in melanoma. Aberrant GRP78 cell surface expression is seen in many of these malignant neoplasms and is associated with a more aggressive phenotype (Quinones et al., 2008). In sera from prostate cancer patients, increased anti-GRP78 antibodies are associated with prostate cancer progression and shorter survival time (Arap et al., 2004). Although GRP78 is primarily located in the ER, under certain conditions, it translocates to the cell surface where it can associate with MTJ1 and the G proteins, Gs and Gaq11 (Misra et al., 2005). On the cell surface, GRP78 acts as a receptor for activated forms of the proteinase inhibitor, alpha 2-macroglobulin (a2M*). a2M* binds GRP78 with extremely high affinity and causes cellular proliferation and survival through the activation of the MAPK, PI3K/Akt, cAMP-dependent, and UPR signaling (Quinones et al., 2008; Gonzalez-Gronow et al., 2009). The auto-antibodies found in the sera of those prostate cancer patients may be directed toward the a2M* binding site in the NH2-terminal domain (NTD) of GRP78, as they are shown to cause proliferation and survival, similar to that seen with a2M* (Gonzalez-Gronow et al., 2006). Additionally, the ligation of cell surface GRP78 with NTD-directed heterologous antibodies leads to the upregulation of cell surface GRP78 through the synthesis of the transcription factor TFII-I (Misra et al., 2009). Interestingly, antibodies directed toward the COOH-terminal domain (anti-CTD) of GRP78 elicit a quite opposite effect. These antibodies initiate apoptosis, inhibit DNA synthesis and cell proliferation, and in the cell lines, 1-LN and DU145, upregulate the tumor suppressor p53. CTD-directed antibodies also lead to the downregulation of GRP78 expression (Misra and Pizzo, 2010a,b; Misra et al., 2010). This may result in apoptosis due to the fact that ER stress is not

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relieved. By analyzing the aforementioned components of the UPR signaling cascade, we can determine the effects of cell-surface GRP78 ligation on the modulation of the UPR. UPR signaling pathways are very important for maintaining normal cellular homeostasis, and perturbation of these pathways can lead to the development of disease. The importance of ER stress in cancer progression is becoming increasingly recognized, and it is therefore important to define methods to monitor the UPR. Here, we use the specific example of GRP78 ligation by antibodies to study the effects on UPR in the context of prostate cancer. The methods outlined here will provide a guide in establishing the function of various cell surface ligands in UPR regulation.

2. Methods for Monitoring UPR Modulation 2.1. Cancer cell lines The 1-LN cell line (a kind gift from Dr. Phillip Walther, Duke University Medical Center) is a highly metastatic cell line derived from the less metastatic PC-3 cell line. The 1-LN cell line expresses high levels of GRP78 on the cell-surface, while PC-3 cells do not express cell-surface GRP78. As such, the PC-3 cell line is a perfect negative control for investigation of cell surface GRP78 ligation on 1-LN cells. PC-3 and DU-145 cells were purchased from the American Type Culture Collection. Our laboratory can supply 1-LN cells to investigators.

2.2. Media and cell culture PC3, 1-LN, and DU145 cells are maintained in log-phase growth in RPMI 1640 containing 10% fetal bovine serum, 2-mM glutamine, 12.5 units/ml penicillin, 6.5-mg/ml streptomycin, and 10-nM human insulin. 1. Grow cells in 6-well plates at 3 105 cells/well in humidified CO2 (5%) incubator. 2. At 90% confluency, aspirate medium and wash with ice-cold Hanks’ balanced salt solution (HBSS) containing 10-mM HEPES (pH 7.4) and 3.5-mM NaHCO3. 3. Add a fresh volume of medium to the cells for the experiments described below.

2.3. Western blotting for UPR signaling cascade components We use Western blotting to determine the protein levels of established UPR target genes that are induced during ER stress.


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1. 1-LN, DU-145, and PC-3 cells are grown in fresh medium as described above. 2. To induce UPR signaling, cells are treated with either buffer (HBSS), a2M* (100 pM for 15 min) or anti-CTD antibody (Stressgen, Victoria, BC, Catalog number SPA-826, 5 mg/ml for 30 min) and incubated at 37 C in a humidified CO2 (5%) incubator. 3. After treatment, medium is aspirated and 1 volume of buffer A (50-mM Tris–HCl, pH 7.4, 120-mM NaCl, 1% Nonidet-P-40, 25-mM NaF, 1-mM sodium pyrophosphate, 0.1-mM sodium orthovanadate, 1-mM benzamidine, and leupeptin (20 mg/ml)) is added to the cells. 4. Cells are lysed for 10 min on ice, scraped into tubes, and centrifuged at 800g for 10 min. 5. Protein concentration is determined by Pierce BCA Assay (Catalog number 23225) according to the manufacturer’s instructions. 6. An equal amount of protein per lane is electrophoretically separated on either 10% or 12.5% SDS-polyacrylamide gels. 7. Proteins are transferred to nitrocellulose membranes and immunoblotted with antibodies specific for UPR signaling components (Table 14.1). All antibodies are used at 1:1000 dilution. 8. To determine antibody specificity for immunoblotting, control experiments were performed with nonimmune serum and processed as above. 9. Membranes were stripped and reprobed for unphosphorylated target protein or actin as the protein loading control. The detection of protein on immunoblots was performed using enhanced chemifluoresence (ECF) (GE Healthcare, Piscataway, NJ) and phosphoimaging on the Storm 840 Optical Scanner (Molecular Dynamics). Densitometry for quantification was performed in the ImageQuant suite of applications (Figs. 14.1 and 14.2).

2.4. Measurement of apoptosis of cells treated with anti-CTD antibodies to GRP78 2.4.1. Western blotting for proapoptotic and antiapoptotic proteins 1. Cell treatment, lysis, electrophoresis, and transfer are identical to those described above. 2. Membranes are immunoblotted with antibodies specific for cleaved caspase-3, procaspase-9, procaspase-12, and caspase-4 (Table 14.1) 2.4.2. DNA fragmentation assay The Cell Death Detection ELISAÒ (Roche, Indianapolis, IN, Catalog number 11544675001) measures cytoplasmic histone-associated DNA fragments (mono- and oligonucleosomes) after induction of cell death (Fig. 14.3).

Table 14.1

List of antibodies

Antibody

Company

Catalog number

Akt Phospho-Aktthr308 Phospho-Aktser472 ASK1 TRAF2 PERK Phospo-PERK PARP-1 Cleaved PARP-1 eIF2a Phospho-eIF2a Caspase-4 Procaspase-12 Cleaved Caspase-3 Procaspase-9 IRE1a XBP1 ATF6 a ATF4 GADD153 GADD34 Actin

Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Cell Signaling Technologies, Inc. (Beverly, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Santa Cruz Biotechnology (Santa Cruz, CA) Sigma (St. Louis, MO)

2670 4056 5048 3762 4724 3192 3191 9542 9541 2103 3597 4450 2202 9661 9502 sc-10510 sc-7160 sc-166659 sc-200 sc-166682 sc-825 A3653

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7 1-LN

∗

Ratio (Test protein/actin)

6 5 4

∗

3 2

∗

∗

1 0

1

2

3

ATF6a

IRE1-a

Actin

Actin

XBP1

p-PERK

Actin

PERK 1

2

3

1

2

3

Figure 14.1 Modulation of UPR transducer expression by anti-CTD antibodies in 1-LN cells. Representative immunoblots of ATF6, IRE1-a, XBP1, and p-PERK with their respective protein loading controls are shown. The lanes are (A) buffer treated, (B) a2M* (15 pM/15 min), and (C) anti-CTD antibody (1:200/30 min). Quantification of antibody-induced changes in the expression of transducer proteins is shown in the bar diagram above the immunoblots. The ratio of test protein/protein loading control is the mean SE from three to four individual experiments. The bars in each panel are ATF6a (open square box), IRE1a (light-shaded gray box), XBP1 (dark-shaded gray box), and p-PERK (filled square box). *, Values significantly different between buffer and anti-CTD antibody-treated cells, p < 0.05 (adapted from Misra and Pizzo, 2010a,b).

1. Plate cells at 1 105 cells/well in a 96-well plate in the above RPMI 1640 medium. Incubate overnight as normal. 2. Treat cells with buffer, a2M* (50 pM for 15 min), anti-CTD antibody (1:200 for 2 h), or staurosporine (5 mM for 2 h) and incubate at 37 C in a humidified CO2 (5%) incubator. 3. DNA fragmentation is measured according to the manufacturer’s instructions. 4. DNA fragmentation is expressed as the specific enrichment of cytosol with mono- and oligonucleosomes.

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5

1-LN

∗∗

∗

Ratio (Test protein/actin)

4

3

∗ 2

∗

1

0

1

2

3

ATF4

p-e1F2a

Actin

e1F2a

GADD34

GADD153 Actin

Actin 1

2

3

1

2

3

Figure 14.2 Anti-CTD antibody-induced changes in expression of UPR effectors in 1-LN cells. Representative immunoblots of ATF4, p-eIF2a, GADD34, and GADD153 with their respective protein loading controls are shown. The lanes are (A) buffer treated, (B) a2M* (50 pM/15 min), and (C) anti-CTD-antibody (1:200/30 min). Antibody-induced changes in the expression of proteins are shown in the bar diagram above the immunoblots as the ratio of test protein/protein loading control and are the mean SE from three to four individual experiments. The bars are ATF4 (open square box), p-eIF2a (light-shaded gray box), GADD34 (dark-shaded gray box), and GADD153 (filled square box). *, Values significantly different from buffer treated and CTD-GRP78 treated cells, p < 0.05. **, Values significantly different from cells treated with buffer and a2M*, p < 0.05 (adapted from Misra and Pizzo, 2010a,b).

2.4.3. Annexin V assay for the measurement of apoptosis The Guava NexinÒ (Annexin V) Assay (Millipore, Billerica, MA, Catalog number 4500-0455) measures the translocation of phosphatidyl serine to the outer surface of the cell membrane. This event is associated with early apoptic events. 1. Cells are plated and treated as described above for DNA fragmentation assay. 2. Apoptosis is measured using the Guava Nexin ReagentÒ on a Guava EasyCyte PlusÒ flow cytometer according to the manufacturer’s instructions.

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∗

1-LN

MBP phosphorylation (% change over basal)

180

∗

130

80

30

1

2

p-Aktthr308

3

p-Aktser473 1

2

3

1

2

3

Figure 14.3 Downregulation of Akt kinase activation by anti-CTD antibody in 1-LN prostate cancer cells. Changes in myelin basic protein phosphorylation by p-AktThr308 (open square box) and p-AktSer473 (filled square box) kinase immunoprecipitates are shown in the bar diagram above autoradiographs. Changes in MBP phosphorylation are expressed as percent change from basal (100%) and are mean SE from two experiments. *, Values significantly different between buffer and anti-CTD-antibody-treated cells, p < 0.05 (adapted from Misra and Pizzo, 2010a,b).

2.5. Measurement of [3H]-thymidine uptake in response to antibodies against GRP78 Binding of a2M* to cell-surface GRP78 results in a two- to threefold increase in [3H]-thymidine uptake by 1-LN and DU145 cells, but not PC-3 cells, as they do not express GRP78 on the cell surface. We have studied the effects of anti-NTD and anti-CTD antibodies to GRP78 on cell proliferation as measured by [3H]thymidine uptake. 1. Grow cells in the above RPMI 1640 medium to 80–90% confluency. 2. Aspirate medium, add fresh RPMI 1640, and use cell scraper to remove cells. 3. Centrifuge cells at 200g for 5 min. 4. Aspirate medium and resuspend the pellet in a volume of medium. 5. Plate cells at 8–10 104 cells/well in a 48-well plate. 6. Allow cells to adhere and replace the medium with fresh RPMI 1640 containing 0.5% fetal bovine serum. 7. After incubation for 10–12 h, [3H]thymidine (2 mCi/ml) is added to each well. This is followed by the addition of either buffer, a2M* (50 pM) or anti-GRP78 antibodies (Anti-CTD antibody, Stressgen,

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Catalog number SPA-826, Anti-CTD antibody, Santa Cruz Biotechnology, Santa Cruz, CA, Catalog number sc-1051, Anti-CTD antibody, Santa Cruz Biotechnology, Santa Cruz, CA, Catalog number sc-13968, or Anti-NTD antibody Santa Cruz Biotechnology, Santa Cruz, CA, Catalog number sc-1050). All the antibodies are used at a concentration of 1:200 and cells are incubated overnight as described above. 8. Following overnight incubation, reactions are terminated by aspirating the medium, and cell monolayers are washed twice with ice-cold 5% trichloroacetic acid, followed by three washes with ice-cold PBS. 9. Cells are lysed with 1N NaOH for 2 h at 40 C. 10. Aliquots of the lyzates are used for determination of radioactivity in a liquid scintillation counter.

2.6. Measurement of the activation of Aktthr308 and Aktser473 kinase Akt promotes cell survival and Akt activation is observed in a number of human cancers. We have shown that anti-CTD GRP78 antibodies inhibit Akt activation and promote apoptotic signaling. 1. Cells are stimulated with either buffer, a2M*, anti-CTD, or anti-NTD antibodies as described above. 2. Aspirate medium and lyse cells in buffer A. 3. Determine protein concentration by Pierce BCA Assay (Catalog number 23225) according to the manufacturer’s instructions. 4. Divide protein equally into Eppendorf tubes. 5. Add antibodies against p-Aktthr308 and p-Aktser473 at 1:50. 6. Add 40 ml of protein A-agarose and incubate overnight at 4 C. 7. Wash immunoprecipitates with buffer A containing 0.5 mM NaCl, followed by buffer A supplemented with 1-mM DTT, 1-mM PMSF, and 1-mM benzamidine. 8. Centrifuge at 2500 rpm for 5 min at 4 C. 9. To each immunoprecipitate, add 40 ml of kinase buffer containing 50-mM Tris–HCl (pH 7.4), 10-mM MgCl2, 1-mM DTT, 1-mM PMSF, and 20-mg/ml leupeptin. 10. Add 5-mg myelin basic protein (MBP) and initiate the kinase reaction with the addition of 50-mM ATP and 5 mCi of [g33P]ATP (Perkin Elmer Life Sciences, Waltham, CA). 11. Incubate tubes for 30 min at 30 C with shaking. 12. Terminate reaction with the addition of 10-ml SDS sample buffer and process as above to quantify phosphorylated Akt by immunoblotting.


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2.7. Silencing GRP78 gene expression by RNAi GRP78 plays a pivotal role in the regulation of UPR signaling during ER stress. By modulating GRP78 expression through RNAi, we can study the effects of GRP78 expression on various components of the UPR signaling cascade, most notably, those components affected by a2M*, anti-CTD, or anti-NTD antibody stimulation. This protocol for GRP78 knockdown by siRNA results in a reduction of GRP78 by 60–70% as measured by GRP78 mRNA and protein levels. Knockdown of GRP78 also results in a loss of cell-surface GRP78 as measured by receptor binding and abrogation of a2M* signaling. 1. The chemical synthesis of dsRNA homologous to GRP78 mRNA (50 ! 30 ) AAAATACAGCAATTAGTAAAG (Swiss-Prot primary accession number P11021) was done by Ambion, Austin, TX. 2. To make the dsRNA, the sense (5 0 ! 30 AAUACAGCAAUUAGUAAAG) and antisense (50 ! 30 CUUUACUAAUUGCUGUAUUTT) oligonucleotides were annealed as per the manufacturer’s instructions. 3. Wash confluent 1-LN cell monolayers (1.5 106 cells/ well in a 6-well plate) twice with Hanks balance salt solution. Add 2-ml DMEM containing 10% fetal bovine serum, 12.5 units/ml penicillin, and 6.5-mg/ml streptomycin. Incubate for 16 h. 4. Just before transfection, dilute 25-mg GRP78 dsRNA to 100 ml with serum and antibiotic-free DMEM in a tube. In a separate tube, dilute 10-ml Lipofectamine 2000 (Invitrogen, Carlsbad, CA) to 100 ml with serum and antibiotic-free DMEM. 5. Mix both solutions together gently and incubate for 45 min at room temperature. 6. Add 800 ml of serum and antibiotic-free medium to each tube. 7. Wash cell monolayers with serum and antibiotic-free DMEM. Layer 1 ml of Lipofectamine 2000-DMEM (control) or Lipofectamine 2000dsRNA mixture onto the cells. Incubate for 5 h at 37 C in a humidified CO2 (5%) incubator. 8. After 5 h, add 1 ml of antibiotic-free DMEM containing 10% fetal bovine serum to each well and incubate for 16 h. 9. Replace the medium with DMEM containing 10% fetal bovine serum, 12.5 units/ml penicillin, and 6.5-mg/ml streptomycin 24 h after the start of transfection. 10. Incubate for an additional 24 h (48 h total). 11. Aspirate the medium and wash monolayers with fresh complete DMEM. Add a volume of the same medium and use cells for the experiments described above.

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12. In order to demonstrate that transfection of dsRNA to GRP78 does not produce nonspecific effects on GRP78 expression, 1-LN cells are transfected with equimolar concentrations of scrambled small interference RNA (SilencerTM negative control, Catalog number 4610, Ambion, Austin, TX).

3. Conclusions The methods outlined here provide techniques to study the effect of cell-surface ligands on UPR signaling and modulation. Although we use the specific example of GRP78, these methods can be extended to any number of cell-surface receptors, the ligation of which leads to ER stress and induction of the UPR signaling cascade. As our knowledge of the UPR pathways in disease is furthered, we will develop new methods to monitor this highly important signaling cascade.

REFERENCES Arap, M. A., Lahdenranta, J., Mintz, P. J., Hajitou, A., Sarkis, A. S., Arap, W., and Pasqualini, R. (2004). Cell surface expression of the stress response chaperone GRP78 enables tumor targeting by circulating ligands. Cancer Cell 6, 275–284. Fels, D. R., and Koumenis, C. (2006). The PERK/eIF2alpha/ATF4 module of the UPR in hypoxia resistance and tumor growth. Cancer Biol. Ther. 5, 723–728. Gonzalez-Gronow, M., Cuchacovich, M., Llanos, C., Urzua, C., Gawdi, G., and Pizzo, S. V. (2006). Prostate cancer cell proliferation in vitro is modulated by antibodies against glucose-regulated protein 78 isolated from patient serum. Cancer Res. 66, 11424–11431. Gonzalez-Gronow, M., Selim, M. A., Papalas, J., and Pizzo, S. V. (2009). GRP78: A multifunctional receptor on the cell surface. Antioxid. Redox Signal. 11, 2299–2306. Harding, H. P., Novoa, I., Zhang, Y., Zeng, H., Wek, R., Schapira, M., and Ron, D. (2000). Regulated translation initiation controls stress-induced gene expression in mammalian cells. Mol. Cell 6, 1099–1108. Kaneko, M., and Nomura, Y. (2003). ER signaling in unfolded protein response. Life Science 74, 199–205. Lass, A., Kujawa, M., McConnell, E., Paton, A. W., Paton, J. C., and Wojcik, C. (2008). Decreased ER-associated degradation of alpha-TCR induced by Grp78 depletion with the SubAB cytotoxin. Int. J. Biochem. Cell Biol. 40, 2865–2879. Misra, U. K., and Pizzo, S. V. (2005). Up-regulation of GRP78 and antiapoptotic signaling in murine peritoneal macrophages exposed to insulin. J. Leukoc. Biol. 78, 187–194. Misra, U. K., and Pizzo, S. V. (2010a). Modulation of the unfolded protein response in prostate cancer cells by antibody-directed against the carboxyl-terminal domain of GRP78. Apoptosis 15, 173–182. Misra, U. K., and Pizzo, S. V. (2010b). Ligation of cell surface GRP78 with antibody directed against the COOH-terminal domain of GRP78 suppresses Ras/MAPK and


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PI 3-kinase/AKT signaling while promoting caspase activation in human prostate cancer cells. Cancer Biol. Ther. 9, 142–152. Misra, U. K., Gonzalez-Gronow, M., Gawdi, G., and Pizzo, S. V. (2005). The role of MTJ-1 in cell surface translocation of GRP78, a receptor for alpha 2-macroglobulindependent signaling. J. Immunol. 174, 2092–2097. Misra, U. K., Mowery, Y., Kaczowka, S., and Pizzo, S. V. (2009). Ligation of cancer cell surface GRP78 with antibodies directed against its COOH-terminal domain up-regulates p53 activity and promotes apoptosis. Mol. Cancer Ther. 8, 1350–1362. Misra, U. K., Kaczowka, S., and Pizzo, S. V. (2010). Inhibition of NF-kappaB1 and NFkappaB2 activation in prostate cancer cells treated with antibody against the carboxyl terminal domain of GRP78: Effect of p53 upregulation. Biochem. Biophys. Res. Commun. 392, 538–542. Namba, T., Ishihara, T., Tanaka, K., Hoshino, T., and Muzushimna, T. (2007). Transcriptional activation of ATF6 by endoplasmic reticulum stressors. Biochem. Biophys. Res. Commun. 355, 543–548. Quinones, Q. J., de Ridder, G. G., and Pizzo, S. V. (2008). GRP78: A chaperone with diverse roles beyond the endoplasmic reticulum. Histol. Histopathol. 23, 1409–1416. Salminen, A., Kauppinen, A., Suuronen, T., Kaarniranta, K., and Ojala, J. (2009). ER stress in Alzheimer’s disease: A novel neuronal trigger for inflammation and Alzheimer’s pathology. J. Neuroinflammation 6, 41. Yoneda, T., Imaizumi, K., Oono, K., Yui, D., Gomi, F., Katayama, T., and Tohyama, M. (2001). Activation of caspase-12, an endoplastic reticulum (ER) resident caspase, through tumor necrosis factor receptor-associated factor 2-dependent mechanism in response to the ER stress. J. Biol. Chem. 276, 13935–13940. Yoshida, H., Nadanaka, S., Sato, R., and Mori, K. (2006). XBP1 is critical to protect cells from endoplasmic reticulum stress: Evidence from Site-2 protease-deficient Chinese hamster ovary cells. Cell Struct. Funct. 31, 117–125.


C H A P T E R

F I F T E E N

Endoplasmic Reticulum Stress in Brain Damage☆ Ram Raghubir,* Venkata Prasuja Nakka,† and Suresh L. Mehta‡ Contents 1. Introduction 2. ER Stress and Unfolded Protein Response 2.1. The PERK pathway 2.2. The IRE1 pathway 2.3. The ATF-6 pathway 2.4. The CHOP pathway 2.5. The JNK pathway 2.6. The caspase pathway 3. Cross Talk Between ER and Mitochondria 4. Experimental Approaches for the Detection of ER Stress 4.1. Detection of IRE1 activation and splicing of XBP-1 mRNA 4.2. Detection of ATF-6 translocation from ER to the nucleus with fluorescent microscopy 4.3. mRNA and protein levels of UPR target genes 4.4. Use of transgenic animal models for monitoring ER stress Acknowledgment References

260 261 262 263 264 264 265 266 267 269 269 269 270 271 271 272

Abstract The efficient functioning of the ER is indispensable for most of the cellular activities and survival. Disturbances in the physiological functions of the ER result in the activation of a complex set of signaling pathways from the ER to the cytosol and nucleus, and these are collectively known as unfolded protein response (UPR), which is aimed to compensate damage and can eventually trigger cell death if ER stress is severe or persists for a longer period. ☆

C.D.R.I. Communication No. 8806

* Division of Pharmacology, Central Drug Research Institute, (CSIR), Chatter Manzil Palace, Lucknow, India { Department of Neurological Surgery, School of Medicine & Public Health, University of Wisconsin, Madison, Wisconsin, USA { Department of Pharmaceutical Sciences/BRITE, North Carolina Central University, Durham, North Carolina, USA # 2011 Elsevier Inc. Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00015-7 All rights reserved.

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The precise molecular mechanisms that facilitate this switch in brain damage have yet to be understood completely with multiple potential participants involved. The ER stress-associated cell death pathways have been recognized in the numerous pathophysiological conditions, such as diabetes, hypoxia, ischemia/reperfusion injury, and neurodegenerative disorders, including Alzheimer’s disease, Parkinson’s disease, and bipolar disorder. Hence, there is an emerging need to study the basic molecular mechanisms of ER stress-mediating multiple cell survival/death signaling pathways. These molecules that regulate the ER stress response would be potential drug targets in brain diseases.

1. Introduction The endoplasmic reticulum (ER) is a large membranous network found in all eukaryotes and is the cellular site for synthesis, folding, and maturation of most secreted and transmembrane proteins (Kaufman, 1999; Paschen and Doutheil, 1999). Under normal physiological conditions, these proteins are correctly folded, modified, and assembled by numerous chaperons and catalysts in the ER. Additionally, nearly 30% of newly synthesized proteins are rapidly degraded, possibly as a result of improper protein folding (Schubert et al., 2000). The ER lumen has a high redox potential to maintain an oxidizing environment, which, together with the high protein and Ca2þ concentration, provides the ideal milieu for folding and the posttranslational modifications of proteins. However, failure of this machinery to fold newly synthesized or misfolded proteins, which start accumulating in the ER during physiological or pathological situations, generates a unique danger to the cell in the form of ER stress. Thus, even an acute rise in the translation of secretory proteins would impose a major problem for the cell due to a potential build up of misfolded proteins. The situation becomes even more serious when ER environment is disturbed due to alterations in redox state, calcium levels, or failure to posttranslationally modify secretory proteins. These conditions compromise the overall ability of protein folding capacity of ER and result in aggregation of misfolded and/or unfolded proteins (Lai et al., 2007). To cope with this condition, cells initiate an adaptive response in the form of group of signal transduction pathways, collectively termed as unfolded protein response (UPR). The UPR initially tries to reestablish the normal ER function and prevent cellular damage but can eventually trigger cell death if the stress is severe and prolonged during the course of pathological states such as cerebral ischemia, hypoxia, and hypoglycemia/hyperglycemia. Furthermore, cytoplasmic or nuclear accumulation of insoluble or misfolded protein aggregates in the affected brain regions in various neurodegenerative disorders can cause neuronal death; otherwise, failure of UPR may implicate the onset of neurodegenerative disorders (Kaufman, 2002).

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Therefore, activation of UPR is necessary to restore the normal functional architecture of the cell. The adaptive UPR mechanisms once activated, may initiate transcriptional induction of genes involved in the protein folding or ER-associated protein degradation to remove misfolded proteins. The translational machinery in such situations only translates some selected mRNA during this period to reduce accumulation of new proteins into the ER, until mRNAs encoding UPR proteins are produced. However, upon failure of this process, ER stress triggers cell suicidal mechanisms usually in the form of apoptosis. Further investigations involving modulation of ER stress signaling might allow the development of therapeutic strategies for some human diseases that are caused by ER dysfunction (Salminen et al., 2009; Sokka et al., 2007).

2. ER Stress and Unfolded Protein Response The accumulated deposition of misfolded proteins leading to ER stress affects various cell signaling systems as well as neuronal connectivity and cell death during various neurodegenerative diseases (Bence et al., 2001; Soto, 2003). In addition, the threshold for complete suppression of protein synthesis during cerebral stroke indicates the association of a more comprehensive cellular response involved in deciding the fate of affected neurons and suggests that the size of an infarct after focal ischemia is not only determined by the breakdown of energy metabolism but also by the suppression of protein synthesis (Xie et al., 1989). Therefore, under such circumstances, UPR in the cells including neurons integrate signaling network in adaptation to secretory load. It is comprised by three main signaling systems initiated by three prototype ER-localized stress sensors viz: pancreatic ER kinase (PERK), inositol-requiring enzyme 1 (IRE1), and activating transcription factor 6 (ATF-6) which mediate regulation at both the transcriptional and translational levels upon ER stress (Harding and Ron, 2002; Kaufman, 1999; Mori et al., 2000). Under the normal physiological state, all the three effectors bind to the ER chaperone glucoseregulated protein (GRP78) on their luminal domains; thus, GRP78 acts to suppress their activity (Bertolotti et al., 2000; Shen et al., 2002). Further, under conditions of ER stress, when misfolded proteins accumulate in the ER lumen, GRP78 dissociates from the PERK, ATF-6, and IRE1, allowing their activation (Fig. 15.1). However, the downregulation of UPR can increase the vulnerability to ER stress as in the case of Alzheimer’s disease (familial and sporadic), where disease-linked presenilin-1 (PS1) mutation and aberrant splicing isoform (PS2V), generated by exon 5 skipping of the presenilin-2 (PS2) gene transcript affect the ER stress response by inhibiting activation of ER stress transducers such as IRE1, PERK, and ATF-6 (Katayama et al., 2004).

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Figure 15.1 The proposed mechanism of unfolded protein response. Under conditions of ER stress, when misfolded proteins accumulate in the ER lumen, GRP78 dissociates from the three ER stress receptors, PERK, ATF-6, and IRE1, allowing their sequential activation. The disassociated GRP78 binds to the unfolded proteins to assist in refolding. Activated PERK blocks general protein synthesis by phosphorylating eIF2a but also leads to increased paradoxical translation of activating transcription factor 4 (ATF-4), which further induces transcription of genes required to restore ER homeostasis and proapoptotic CHOP/GADD153. ATF-6 is activated by limited proteolysis after its translocation to the Golgi apparatus. Active ATF-6 regulates the expression of ER chaperones and X box-binding protein 1 (XBP-1). Activated IRE1a (an ubiquitous isoform) cleaves XBP-1 mRNA that leads to enhanced translation of spliced XBP-1 protein, a transcription factor for ER-resident enzymes and chaperones, the cochaperone and PERK-inhibitor P58IPK, as well as genes involved in protein degradation.

2.1. The PERK pathway The primary response to ER stress in mammalian cells is a transient global translation attenuation. Dissociation of GRP78 from PERK initiates the dimerization and autophosphorylation of the kinase and generates active


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PERK. Once activated, PERK phosphorylates serine-51 residue of eukaryotic translation initiation factor 2 subunit a (eIF2 a), which leads to inhibition of global protein synthesis (Prostko et al., 1993; Ron, 2002). However, activated PERK phosphorylates eIF2a to avoid further accumulation of proteins by suppressing protein synthesis and also leads to the paradoxical increased translation of transcription factors ATF-4 and C/EBP homologous protein/growth arrest and DNA damage-inducible gene 153 (CHOP/ GADD153; DeGracia and Montie, 2004). ATF-4 promotes cell survival by inducing ER stress target genes involved in amino acid metabolism, redox reactions, stress response, and protein secretion (Harding et al., 2003). Translational recovery is mediated by the stress-induced phosphatase growth arrest and DNA damage-inducible gene 34 (GADD34), which is upregulated by ATF-4 and dephosphorylates eIF2a. Attenuation of translational recovery by pharmacologically inhibiting eIF2a dephosphorylation protects some cells from ER stress-induced apoptosis. Indeed, selective inhibition of eIF2a dephosphorylation by a small molecule inhibitor, salubrinal (Sal), protects cells from ER stress in a dose-dependent manner (Boyce et al., 2005). The available evidence suggests that Sal inhibits the formation of the eIF2a/GADD34/protein phosphatase 1 (PP1) complex, sustaining the phosphorylation of eIF2a, thereby suppressing protein translation for a longer period during the ER stress (Boyce et al., 2005). The efficacy of Sal against brain damage has been assessed in vivo, which proved to be neuroprotective (Nakka et al., 2010; Sokka et al., 2007). Therefore, modulation of the phosphorylated state of eIF2a can be used as a strategy to protect cells from cerebral damage.

2.2. The IRE1 pathway IRE1 is a dual-activity enzyme, having a serine–threonine kinase domain and an endoribonuclease domain. On activation, the endonuclease activity of IRE1 specifically cuts out a 26-nucleotide intron from the XBP-1 mRNA, which leads to a shift of the open reading frame of XBP-1 mRNA (Calfon et al., 2002). Processed XBP-1 mRNA is translated into a new protein of 54 kDa that functions as a transcription factor and has diverse targets specific for ER stress genes, including grp78, grp94, and HSP40 family member P58IPK (Lee et al., 2003). P58IPK binds and inhibits PERK, thereby providing a negative feedback loop that relieves the PERKmediated translational block (Yan et al., 2002). The adequate newly synthesized GRP78 protein shut down the UPR and restore ER function by inactivating PERK and IRE1, thereby refolding the accumulated unfolded proteins in the ER lumen. Although the IRE1– XBP-1 pathway seems to be prosurvival through the induction of ER chaperones and P58IPK, overexpression of IRE1 in HEK293T cells resulted in apoptotic cell death (Wang et al, 1998). How IRE1 could initiate cell death is currently one of the most interesting and unresolved questions in

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this field? The answer might lie with the activation of kinase pathways, most notably, the c-Jun N-terminal kinase (JNK) pathway, BCL-2, and caspases. Therefore, understanding the switch between prosurvival and proapoptotic IRE1 signaling can be a useful strategy to protect cells from the degenerative brain injuries.

2.3. The ATF-6 pathway After dissociation, GRP78 facilitates ATF-6 translocation to the Golgi apparatus, where it is cleaved by site-1 and site-2 proteases to yield an active N-terminal 50-kDa domain (N-ATF-6/p50ATF-6) that translocates to the nucleus (Hong et al., 2004; Nadanaka et al., 2006). In the nucleus, it induces genes with an ER stress response element (ERSE) in their promoter (Schroder and Kaufman, 2005). So far, the recognized targets of ATF-6 include ER chaperone proteins such as GRP78, GRP94, protein disulfide isomerase, and the transcription factors CHOP and XBP-1. Although ATF-6 can induce CHOP mRNA expression, but there is no available evidence to link ATF-6 to ER stress-induced apoptosis; therefore, it seems that ATF-6-mediated signals seems to be purely prosurvival and counteract ER stress (Szegezdi et al., 2006). However, the role of ATF-6 pathway in cerebral ischemia is unclear and therefore requires extensive investigation. Signaling through PERK, IRE1, and ATF-6 can also trigger apoptosis, when the cellular protective mechanisms mediated by the UPR fail to restore folding capacity. However, they do not directly cause cell death but rather initiate the activation of downstream molecules such as CHOP, JNK, and caspases, which further execute the cell death machinery. In addition, Bcl-2 family of proteins also plays an important role in the regulation of apoptosis. However, the ER-mediated apoptotic machinery involving key regulatory molecules is less defined in cerebral ischemia, which may be delineated in future.

2.4. The CHOP pathway CHOP, also known as GADD153, is induced by ER stress more than growth arrest or DNA damage (Wang et al., 1996). CHOP is 29 kDa protein with 169 (human) or 168 (rodent) amino-acid residues. The role of CHOP in ER stress-induced apoptosis has been demonstrated in CHOP/ mice. Indeed, mouse embryonic fibroblasts lacking CHOP are partially resistant to ER stress and exhibits reduced ER stress-induced apoptosis (Oyadomari et al., 2002; Zinszner et al., 1998). Conversely, overexpression of CHOP promotes apoptosis in response to ER stress caused by thapsigargin and tunicamycin (McCullough et al., 2001). The UPR activation also leads to transcription and translation of CHOP, and both ATF-4 and XBP-1/ATF-6 can contribute to CHOP transcription (Ma et al., 2002). Induction of CHOP mRNA


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serves as a hallmark of ER stress and a marker of UPR activation, and its expression is usually associated with apoptosis (McCullough et al., 2001; Zinszner et al., 1998). Recent studies on cerebral ischemia have documented the involvement of CHOP in apoptosis. For instance, increased expression of CHOP mRNA was observed in the rat hippocampus subjected to transient cerebral ischemia (Paschen et al., 1998). Furthermore, mice lacking CHOP have smaller infarcts than wild-type animals subjected to bilateral common carotid arteries occlusion (Tajiri et al., 2004). Since acute neuronal cell death in the ischemic core region is mostly due to necrosis, ER stressmediated apoptosis in vulnerable neurons contributes to delayed cell death or to neurodegeneration in the penumbra of focal ischemia. Further studies which are aimed to elucidate the mechanism of CHOPinduced apoptosis, identified many target genes, including Bcl-2, GADD34, endoplasmic reticulum oxidoreductin 1 (ERO1a), and tribbles-related protein 3 (TRB3; Szegezdi et al., 2006). Although CHOP mainly induces gene expression, it downregulates the expression of Bcl2 and increases cellular ROS, which contributes to ER-associated cell death (Harding et al., 2003; McCullough et al., 2001). Further studies on ER stress-mediated apoptosis involving CHOP are expected to provide new insights into the pathogenesis of cerebral ischemia and other neurodegenerative diseases, which may prove CHOP as an important target for therapeutic intervention to prevent secondary progression of brain injuries.

2.5. The JNK pathway The activation of JNK in response to ER stress has been found to be IRE1a- and TRAF2-dependent (Urano et al., 2000). During ER stress, IRE1a recruits the adaptor protein TRAF2 to the ER membrane. This recruitment is regulated by c-Jun NH2-terminal inhibitory kinase ( JIK), which has been reported to interact with both IRE1a and TRAF2 (Urano et al., 2000; Yoneda et al., 2001). The complex of IRE1a/TRAF2 then recruits apoptosis signal regulating kinase 1 (ASK1) thereby leading to the activation of ASK1 and the downstream JNK pathway (Nishitoh et al., 1998, 2002). Overexpression of JIK promotes interaction between IRE1a and TRAF2 and JNK activation in response to tunicamycin, whereas overexpression of an inactive JIK mutant inhibits JNK activation (Yoneda et al., 2001). The importance of ASK1 in mediating ER stress-induced apoptosis has been demonstrated in ASK/ primary neurons and MEFs, which are resistant to ER stress inducers and are defective in JNK activation and apoptosis (Nishitoh et al., 2002). However, the downstream mechanism by which ASK1 and JNK lead to apoptosis is not completely understood but may involve the regulation of Bcl-2 family of proteins.

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Activation of JNK is a common response to many forms of stress and is known to influence the cell-death machinery through the regulation of Bcl-2 family proteins (Davis, 2000). It is known that phosphorylation of Bcl-2 by JNK, which occurs primarily at the ER, suppresses the antiapoptotic activity of Bcl-2. Besides Bcl-2, JNK also phosphorylates BH3 (Bcl-2 homology domain 3)-only members of the Bcl-2 family such as Bim, which enhances their proapoptotic potential (Szegezdi et al., 2006). Systemic administration of SP600125, a small molecule JNK-specific inhibitor, results in diminished JNK activity and reduced infarct volume after ischemia in a dose-dependent manner. Moreover, inhibition of JNK prevents mitochondrial translocation of Bax and Bim, release of cytochrome c and Smac, and activation of caspase-9 and caspase-3 (Gao et al., 2005), whereas activation of JNK causes serine phosphorylation of 14-3-3, thus leading to disassociation of Bax from 14-3-3 and subsequent translocation to mitochondria (Gao et al., 2005). Thus, there is a growing evidence to substantiate the role of JNK as a critical cell death mediator involving one of the mechanisms as above in the degenerative brain injuries.

2.6. The caspase pathway ER stress-induced apoptosis is initiated by caspase-12 (Van de Craen et al., 1997), which is mainly located on the cytoplasmic side of the ER and regarded as a representative lead molecule implicated in cell death-executing mechanisms related to ER stress (Nakagawa et al., 2000). Caspase-12 is activated by ER stress stimuli, such as tunicamycin, brefeldin A, and thapsigargin, but not by death receptor or mitochondria-mediated apoptotic signals (Nakagawa and Yuan, 2000; Nakagawa et al., 2000). A preliminary report on caspase-12/ mice and MEFs showed resistance to ER stress-mediated apoptosis, suggesting involvement of the caspase-12 (Nakagawa et al., 2000). So far, several possible mechanisms have been proposed for the activation of caspase-12. It has been suggested that calpains, a family of Ca2þ-dependent cystein proteases, have been shown to play a key role to generate an active caspase-12 (Nakagawa and Yuan, 2000). Another study also reported that caspase-7 in response to ER stress translocates from the cytosol to the ER membrane to interact with caspase12, leading to its activation, as the dominant negative catalytic mutant of caspase-7 inhibited caspase-12 activation and cell death (Rao et al., 2001). In addition to the role of calpains and caspase-7 in the activation of caspase-12, TRAF2 also has been shown to play a role in the activation of caspase-12 and to transduce signals to the nucleus via IRE1a (Yoneda et al., 2001). In unstressed conditions, TRAF2 recruits procaspase-12 to promote the clustering of procaspase-12 at the ER membrane (Yoneda et al., 2001). The interaction between TRAF2 and procaspase-12 is inhibited during ER


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stress or by overexpression of IRE1. Therefore, it has been proposed that during ER stress, procaspase-12 is released from TRAF2 complex and gets activated by proximity-induced dimerization. The activated caspase-12 may subsequently be recruited to IRE1a (Yoneda et al., 2001). While the activation of caspase-12 has been documented in rodent models of cerebral ischemia, the question of whether or not a human isoform of caspase-12 exists remains controversial (Szegezdi et al., 2003). However, despite the controversial role of caspase-12 in humans, a recent study demonstrated that ER stress and caspase-12 activation has been implicated in neurodegeneration in Creutzfeldt–Jakobdisease-affected individuals, and the activation of caspase-12 correlates with the higher levels of ER molecular chaperones GRP58, GRP78, and GRP94 expression (Hetz et al., 2003). Moreover, human caspase-4, which plays a key role in ER stress-induced apoptosis, might functionally substitute for mouse caspase-12 in the human system (Hitomi et al., 2004). Although there is no known caspase-12 substrate and even the downstream pathway also remains to be elucidated, several possible models of caspase-12 processing have been suggested, which includes Apaf-1 and cytochrome c independent processing of caspase-9 by caspase-12 (Morishima et al., 2002; Rao et al., 2002). In contrast, recently, it has been suggested that caspase-12 is cleaved downstream of the mitochondria in an Apaf-1-dependent manner and that it cotranslocates with AIF to the nucleus (Sanges and Marigo, 2006; Shiraishi et al., 2006). Nevertheless, extensive in vivo studies are needed essentially to elucidate mechanism(s) responsible for caspase-12 activation and its downstream signaling, and also to understand the basic molecular mechanisms involved in brain injury-induced ER stress. This would further help in establishing possible ER-targeted therapeutic interventions in various neurological disorders.

3. Cross Talk Between ER and Mitochondria The ER is the main intracellular store organelle for Ca2þ, which is an important secondary messenger and is essential for numerous cellular functions. Apoptosis is a consequence of the perturbation of cellular Ca2þ homeostasis such as intracellular Ca2þ overload, ER Ca2þ depletion, and mitochondrial Ca2þ increase. The exposure of cells to apoptotic agents that disturb ER functions reveals a novel cross talk between ER and mitochondria. Furthermore, close physical contacts of ER and mitochondria provide the environment for a local and privileged communication between the two organelles. Pretreatment with tunicamycin, a drug that blocks the ER-resident protein glycosylation, also affects cytochrome c release from the mitochondria,

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followed by caspase-3 activation and DNA fragmentation. Moreover, both cytochrome c released from the mitochondria and caspase-3 activation blockade occurs, when cells are transfected with Bcl-2 specifically targeted to the ER compartment (Putney and Ribeiro, 2000). Recently, it has been demonstrated that a caspase cleavage product B-cell-associated protein 31 (BAP31), an integral membrane protein of ER, induces mitochondrial fission through ER Ca2þ signals, enhancing cytochrome c release to the cytosol (Rutter and Rizzuto, 2000). BAP31 contains three predicted transmembrane domains, followed by a leucine zipper and a death effector domain-like (DED-L) region that associates with certain isoforms of procaspase-8 in the cytosol. The caspases cleave the cytosolic tail of BAP31 that exhibits apoptotic features, whereas overexpression of full-length BAP31 blocks the Fas-mediated apoptosis. In addition, BAP31 also binds to Bcl-2 and Bcl-xL (Xu et al., 2005). Although the role of BAP31 in cerebral ischemia remains unexplored, these observations suggest that apoptotic cross talk between the ER and the mitochondria might be associated with the pathological states of the brain. The temporal profile of ER and mitochondrial dysfunction induced by transient cerebral ischemia suggests that ER dysfunction may be a process upstream of mitochondrial dysfunction. Phosphorylation of PERK and eIF2a during the early reperfusion after transient cerebral ischemia indicates ER dysfunction to be an early pathological process. Moreover, mitochondrial cytochrome c release has not been observed before 2 h of reperfusion, implying that ER dysfunction does precede impairment of mitochondrial function (Hacki et al., 2000). However, a recent study on HeLa cell line has shown that, at early stages of apoptosis, cytochrome c translocates to the ER, where it selectively binds IP3R, resulting in a sustained increase of cytosolic Ca2þ (Boehning et al., 2003). In agreement with the above observation, inositol 1, 4, 5-trisphospate receptor 1 (IP3R1) and ryanodine receptors get activated after global cerebral ischemia by cytochrome c, resulting in Ca2þ efflux from ER (Beresewicz et al., 2006). Thus, it is possible that ER stress triggers a set of reactions leading to leakage of cytochrome c from the mitochondria that further stimulates ER receptors to release more Ca2þ, resulting in a positive feedback loop. It has been reported that ER stress inducers cause coactivation of AIF and caspase12 and their subsequent redistribution to the nucleus (Sanges and Marigo, 2006). Furthermore, reduction in the AIF or caspase-12 expression by RNA interference revealed that AIF primarily controls apoptosis caused by changes in Ca2þ homeostasis but not necessary for protein misfolding apoptosis, whereas caspase-12 seems to regulate both AIF activation and apoptosis (Sanges and Marigo, 2006). Thus, it suggests a novel cross talk between the ER and the mitochondria that might be linked to the pathogenesis of cerebral ischemia.


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4. Experimental Approaches for the Detection of ER Stress The proper use of various experimental approaches of UPR targets is of immense value to understand the basic molecular mechanisms of ER stress-mediated brain damage. These approaches have been depicted in a schematic manner (Fig. 15.2).

4.1. Detection of IRE1 activation and splicing of XBP-1 mRNA The UPR adaptive response up regulates the transcription of genes encoding ER-resident chaperones. The XBP1, which is one of the important factors in UPR signaling during ER stress, is processed at mRNA level by IRE1 dependant unconventional splicing. IRE1 is an ER-located type I transmembrane protein with a kinase domain and an RNase domain in the cytoplasmic region. This gets activated when IRE1 is oligomerized, allowing the transautophosphorylation of IRE1 kinase domain. The active IRE1 then removes 26-nucleotide intron from XBP-1 transcript, leading to a shift in the codon reading frame. The spliced XBP-1 mRNA is translated into a functional transcription factor for UPR, which is a more potent and stable transcription factor. The spliced product of XBP-1 mRNA can be detected by semiquantitative RT-PCR using primer such as human XBP-1 forward primer (FP)-TTACGAGAGAAAACTCATGGCC and reverse primer (RP): GGGTCCAAGTTGTCCAGAATGC; rat: FP-CTGAGTCCGCACAG and RP-GGATCTCTAAAACTAGAGGCT, and mouse: FP-GAACCAGGAGTTAAGAACACG and RP-GGCAACAGTGTCAGAGTCC that can detect both the spliced and unspliced isoforms (Nakka et al., 2010; Samali et al., 2010).

4.2. Detection of ATF-6 translocation from ER to the nucleus with fluorescent microscopy ATF-6, which is another ER transmembrane protein, is activated in response to ER stress by sequential proteolysis, resulting in the 50-kDa p50ATF-6 that translocates to the nucleus. During activation, ATF-6 is transported from the ER to Golgi body, where it is processed by site-1 and site-2 proteases. In this process, the cytoplasmic fragment of ATF-6 after releasing from the membrane is translocated to the nucleus to activate its transcriptional targets. Translocation of ATF-6 can be monitored and detected using ATF-6-specific antibodies by confocal microscopy to ascertain the role of UPR and ER stress in brain damage. However in vitro, ATF-6 activation and translocation can be monitored by using cells

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ER stress and brain (Cerebral stroke, Alzheimer’s etc.) UPR

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Figure 15.2 Illustrates various approaches that may be used to detect and evaluate the activation of UPR and UPR-associated ER stress signaling. The use of multiple assays is important to ascertain and verify UPR activation. WB, Western blotting; IF, Immunofluorescence; t-mice, transgenic mice (/).

transfected with FLAG-tagged ATF-6 or GFP–ATF-6 fusion protein. The FLAG-tagged ATF-6 can be detected using anti-FLAG fluoisothiocynatelabeled antibodies; whereas the N-terminal-cleaved ATF-6 can be detected using anti-FLAG antibody.

4.3. mRNA and protein levels of UPR target genes ER stress has been well documented in various brain pathologies, including cerebral stroke, when stress leads to mitochondrial dysfunction, depletion of ER luminal Ca2þ, or inhibition of ER protein glycosylation. The ER stress response in such a condition autoregulates by temporarily slow accumulation of new proteins in the ER lumen and simultaneously upregulates transcription of genes for ER-resident chaperones and enzymes that abate the effects of ER stress. The target ER stress-responsive genes, such as GRP78, GRP79, and calreticulin, with ERSE consensus sequence can be


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detected with semiquantitative or real-time RT-PCR (Samali et al., 2010). Similarly, protein levels of UPR target genes can be detected using specific antibodies by Western blotting (WB) and immunohistochemistry. The increased phosphorylation of PERK eIF2a can be detected by WB using phospho-specific antibodies.

4.4. Use of transgenic animal models for monitoring ER stress ER stress caused by accumulation of unfolded proteins in the ER is associated with neurodegenerative diseases such as Parkinson’s disease, Alzheimer’s disease, and bipolar disorder, as well as cerebral stroke. However, in vitro approach sometimes initiates cellular damage mechanism in response to insufficient ER stress. Therefore, studies that involve monitoring the ER stress during pathology and development will precisely help to elucidate the association of ER stress to various other cellular signaling. To facilitate the monitoring and analysis of ER stress in vivo, two different transgenic mouse models have been described to exploit various issues regarding ER stress in human diseases and drug development. The first model which was developed by Miura and colleagues is referred as “ER stress-activated indicator” (ERAI; Iwawaki et al., 2004). This was constructed by fusing gene encoding venus variant of green fluorescent protein, as a reporter downstream of a partial sequence of human XBP-1, including the 26-nt ER stress-specific intron. Therefore, ER stress in the cells can be examined by monitoring the fluorescence activity of venus when fusion protein of XBP-1 and venus is produced in the cells (Iwawaki et al., 2004). This is a good model to monitor in vivo the specific effects of ER stress with high sensitivity during development, pathophysiological states, as well as for analysis of drug effects on ER function. However, this model can be only used to detect activation of IRE1 and does not reveal any information about ATF-6 and PERK activation. The second model is known as ERSE-LacZ model (Mao et al., 2006). This was constructed by using a LacZ reporter gene driven by 3 kb of the rat GRP78 promoter. The ERSE-LacZ model helps to determine the expression profile and specificity of ERSE-mediated ER stress in vivo. However, this model system does not reveal any information about the other components of UPR.

ACKNOWLEDGMENT We acknowledge the research support of Central Drug Research Institute (CSIR), Lucknow, 226001, India.

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Methods and Models for Monitoring UPR-Associated Macrophage Death During Advanced Atherosclerosis Edward B. Thorp Contents 1. Introduction to ER Stress in Atherosclerosis 2. Significance of Macrophage ER Stress and Apoptosis in Advanced Atherosclerosis 3. Modeling Atherosclerosis-Relevant ER Stress-Induced Apoptosis 3.1. Method: Athero-relevant ER stress-induced apoptosis in primary macrophages 4. The UPR and Macrophage Death at the Murine Aortic Root 4.1. Method: Laser-capture microdissection of the UPR in atherosclerosis 4.2. Method: Detection of apoptotic macrophages in ER-stressed plaque 5. Conclusion and Future Methodological Advances Acknowledgments References

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Abstract During advanced atherosclerosis, chronic activation of the endoplasmic reticulum (ER) stress pathway, otherwise known as the unfolded protein response (UPR), is strongly associated with atherosclerotic plaque destabilization, the precursor to acute myocardial infarction and sudden death. Destabilized or vulnerable plaques are characterized by features that include thinning of a protective collagenous cap at the interface between the plaque and the vascular lumen and expansion of the necrotic core, a lipid-rich graveyard of dead macrophages. The cell biology of advanced plaque progression is complex and includes multiple cellular stressors that combine to promote chronic inflammation and progressive plaque deterioration. Several of these stressors converge at the ER, leading to activation of the UPR in multiple cell types. In macrophages, prolonged UPR activation triggers apoptosis, which when Department of Medicine, Division of Molecular Medicine, Columbia University, New York, USA Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00016-9

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coupled to defective phagocytic clearance of these dying cells, leads to secondary necrosis and expansion of the plaque necrotic core. Though much insight has been gained recently on the role of the UPR in atherosclerosis, future studies are warranted to determine the cell-type specific contributions of ER stress to athero-progression and the therapeutic potential of UPR modulation. For such objectives to be met, reliable and standardized methodology must be utilized and developed. This chapter summarizes our current understanding of ER stress-induced macrophage apoptosis in atheromata and outlines both in vitro and in vivo methodologies to quantify the UPR in the context of experimental murine-advanced atherosclerosis.

1. Introduction to ER Stress in Atherosclerosis The adaptive cellular response known as the unfolded protein response (UPR) occurs under conditions that perturb endoplasmic reticulum (ER) homeostasis and induce ER stress (Ron and Walter, 2007). Three ER resident proteins sense and differentially regulate the UPR. These include inositol-requiring enzyme-1 (IRE-1; Cox et al., 1993), activating transcription factor-6 (ATF6; Yoshida et al., 1998), and PKR-like eukaryotic initiation factor 2 kinase (PERK; Harding et al., 1999). The function of the UPR or integrated stress response (ISR) is to protect the ER from normal and pathophysiological perturbations that include increased protein synthesis, disruption of ER calcium homeostasis, changes in redox potential, and alterations in the physical properties of the ER membrane bilayer (Hossain et al., 2003; Ma and Hendershot, 2001). In response to interference with ER homeostasis, the UPR initiates transcription of ER chaperones, oxidoreductases, and ER-associated degradation (ERAD) proteins (Travers et al., 2000). The UPR also exerts translational control by phosphorylating eif2a, which is activated by PERK (Harding et al., 2000). This leads to generalized inhibition of protein translation, except in the case of ATF4, which is selectively upregulated. ATF4 induces the proapoptotic transcription factor CHOP (GADD153; Zinszner et al., 1998). In the case of ATF6, its activation by intramembrane proteolysis leads to the production of protein-folding chaperones. This is partially accomplished through ATF6-mediated transcription of XBP-1 (Yoshida et al., 2001). XBP-1 is also activated by IRE-1. IRE-1, through its endoribonuclease domain, promotes XBP-1 mRNA splicing. Spliced XBP-1 targets genes that enhance ER protein folding capacity (Lee, 2003). IRE-1 is also implicated in nonadaptive proapoptotic signaling that includes ribonuclease-mediated mRNA decay and proapoptotic c-Jun kinase ( JNK) signaling (Urano, 2000). If the UPR is successful in alleviating ER stress, negative feedback

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signaling leads to UPR suppression (Merksamer et al., 2008). If on the other hand, these adaptive responses are not sufficient to restore ER equilibrium, the UPR triggers programmed cell death (Lin et al., 2007). Over the last 10 years, ER-stress signaling has emerged to be a key factor in events that promote atherosclerosis. Recent evidence in experimental mice and humans support a scenario whereby chronic ER stress in the vascular wall promotes atherosclerotic progression and events that directly contribute to plaque vulnerability, the precursor to acute myocardial infarction and sudden death. During atherosclerosis, ER stress is activated in multiple cell types of the vascular wall. Though little is known about the role of ATF6 signaling during atherosclerotic development, the other arms of the UPR, that is, IRE-1 and PERK, have been shown to play a significant role in endothelial cells and macrophages of the vascular wall. In endothelial cells, the potentially pro-atherogenic molecule homocysteine causes activation of ER stress-induced growth arrest (Outinen et al., 1999). Also in endothelial cells, IRE-1-mediated XBP-1 splicing leads to endothelial apoptosis and atherosclerosis development in response to disturbed arterial flow (Zeng et al., 2009). The PERK pathway, through its induction of ATF4 and CHOP, appears to be especially important during atherosclerotic progression. Several studies have documented increased mRNA and protein of CHOP in both experimental animals and in human coronary artery plaques. Myoishi et al. reported a strong correlation between expression of CHOP and plaque necrosis and rupture in human coronary lesions (Myoishi et al., 2007). Furthermore, deficiency of CHOP in both the Ldlr/ and Apoe/ models of atherosclerosis reduces advanced lesional macrophages apoptosis and plaque necrosis (Thorp et al., 2009). Thus, the activation of multiple arms of ER stress in distinct cell types combines to affect disparate events that advance atherosclerotic progression. The aforementioned reports shed light on the physiological relevance of ER stress during atherosclerosis; however, these few studies have only scratched the surface. Future studies are needed to test the significance of UPR modulation in atherosclerosis and its amenability to therapeutic intervention. This chapter discusses experimental methods for the interrogation and characterization of one of the best characterized UPR pathways in atheromata, that which occurs in the macrophage during ER stress-mediated apoptosis.

2. Significance of Macrophage ER Stress and Apoptosis in Advanced Atherosclerosis There is sufficient evidence that the UPR is chronically activated in cells of the atherosclerotic vascular wall, including macrophages (Tabas, 2009). Macrophages are a central protagonist of both early and advanced

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atherosclerotic development. Early atherogenesis is characterized by an inflammatory response to retained apolipoproteins in the subendothelium (Tabas et al., 2007; Williams and Tabas, 1995). This involves infiltration of monocytes that later differentiate into macrophages and ingest the retained lipids. During atherosclerotic progression, lesions exhibit evidence of significant accelerated macrophage death (Ball et al., 1995). Increased macrophage apoptosis, in tandem with defective phagocytic clearance of apoptotic macrophages, contributes to expansion of the necrotic core (Tabas, 2005). This destabilizes plaques and in turn promotes plaque disruption and acute thrombotic events, including myocardial infarction and stroke (Kolodgie et al., 2003; Tabas, 2005; Virmani et al., 2006). Work by Feng et al. and later Austin and colleagues showed that ER stress was significantly elevated in lesional macrophages (Feng et al., 2003a; Zhou et al., 2005). These studies also provided evidence for increased expression of the UPR effector CHOP in advanced lesions. These data in experimental mice were later confirmed in studies of human carotid endarterectomy specimens. In these studies, robust CHOP expression and lesional apoptosis was only identified in advanced vulnerable plaques (Myoishi et al., 2007). Together, the experimental evidence supports a working model whereby ER stress-induced macrophage apoptosis is a key event in plaque destabilizing necrotic core generation. The mechanisms of ER-stressed macrophage apoptosis have been explored in detail using cultured macrophage models. These studies indicate that robust apoptotic induction requires the CHOP pathway and that this is upstream of canonical Fas and mitochondrial death pathways (Timmins et al., 2009). CHOP promotes apoptosis by activating the transcription of ER oxidase, which in turn promotes ER calcium release through the IP3 receptor (Harding et al., 2003; Li et al., 2009). Increased cytosolic calcium activates calcium/calmodulin-dependent protein kinase II, which then triggers proapoptotic pathways that include Fas, mitochondrial release of apoptogenic factors, signal transducer and activator of transcription-1 (STAT1) and NADPH oxidase-mediated reactive oxygen species (ROS; Lim et al., 2008). In vitro, 70% of macrophages from Chop/ mice are protected from ER stress-induced apoptosis (Feng et al., 2003a). This is consistent with the report described above in which Chop/ mice are protected from macrophage apoptosis and necrosis in plaque. Thus, the combination of mechanistic data from primary cultured cells with in vivo genetic tests of causality have uncovered key roles for the UPR in advanced lesional macrophage death and plaque necrosis (Feng et al., 2003a,b). Below we describe (1) in vitro methods to measure ER stress-induced apoptosis in macrophages under conditions that model the in vivo milieu and later (2) in vivo methodologies for monitoring ER stress and macrophage apoptosis in atherosclerotic lesions.


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3. Modeling Atherosclerosis-Relevant ER Stress-Induced Apoptosis Mechanistic insight from in vitro studies has proven essential to explain the biology of atherosclerotic progression in vivo. Such studies have implicated multiple ER-stress pathways that may contribute to macrophage apoptosis during atherosclerosis. Athero-relevant inducers are identified from molecules and processes within athermoata that can lead to UPR activation. These include intracellular unesterified cholesterol, oxysterols, oxidant stress, hypoxia, and peroxynitrite (Seimon and Tabas, 2009). For example, in early lesions, aggregated lipoprotein-derived cholesterol is trafficked to the macrophage ER after phagocytic internalization, where acyl-coenzyme A-cholesterol acyltransferase (ACAT) esterifies cholesterol to its cholesteryl acyl ester (Brown et al., 1980). However, in advanced atherosclerosis, macrophages accumulate large amounts of unesterified “free” cholesterol (FC; Katz et al., 1976). In vitro, FC accumulation in macrophages activates the UPR and promotes apoptosis (Feng et al., 2003a). Robust ER stress can also be achieved with 7-ketocholesterol (7KC). 7-KC is an ER stressor that is the most abundant oxysterol in human atherosclerotic lesions (Myoishi et al., 2007). Another scenario of atherorelevant ER stress-induced apoptosis is triggered by the combination of a low-dose ER stressor and a “second hit,” each of which are unable to induce apoptosis by themselves (Seimon et al., 2006). For the second hit, activation of combinatorial pattern recognition receptors (PRR) triggers apoptosis through NADPH oxidase and an ROS pathway in combination with suppression of cell survival pathways. Below, we outline the methodology for measuring athero-relevent ER stress-induced apoptosis in primary macrophages, specifically FC-induced macrophage apoptosis.

3.1. Method: Athero-relevant ER stress-induced apoptosis in primary macrophages 3.1.1. Materials 3.1.1.1. Reagents for eliciting peritoneal macrophages by methyl-BSA or concanavalin A Concanavalin A can be obtained from Sigma (L-7647). Make a 2-mg/ml stock in sterile PBS and store aliquots at 70 C. If eliciting by mBSA injection: Stock methyl-BSA solution: Dissolve 20-mg methyl-BSA (Sigma A1009) in 10-ml sterile 0.9% NaCl. Methyl-BSA in Complete Freund’s adjuvant (CFA DIFCO 231131): Place 2 ml of CFA that has been shaken well before use in a syringe. Fill 2 ml of methyl-BSA solution in another syringe. Connect the 2 syringes to a double-hub microemulsifying needle (20 2–7/8 from Popper & Sons, Inc. #7972)

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and push back and forth 30 times. The mixture becomes very viscous. Methyl-BSA in Incomplete Freund’s adjuvant (IFA: DIFCO 263910). Prepare methyl-BSA-IFA solution same as CFA solution above. MethylBSA in PBS: mix 1-ml methyl-BSA solution above with a 9-ml sterile PBS. 3.1.1.2. Media for primary macrophage culture Tissue culture media, cell culture reagents, and heat-inactivated fetal bovine serum/FBS (GIBCO BRL) can be purchased from invitrogen. Macrophage medium is Dulbecco’s modified eagle’s medium (DMEM) supplemented with 10% FBS and 20% L-cell conditioned medium. L-cells (L929) are from ATCC. L-cell conditioned medium is collected 24 h after adding fresh DMEM 10% FBS to L-cells at 80% confluency. 3.1.1.3. Preparation of LDL by density ultracentrifugation Isolate lowdensity lipoprotein/LDL (d, 1.020–1.063 g/ml) from freshly pooled human plasma (never frozen) by ultracentrifugation. The density of the plasma is assumed to be 1.006; however, the actual density can be measured by weighing 1 ml of solution on a precision scale. Adjust the density to 1.019 with NaBr. The amount to be added is calculated by the following equation: Grams of NaBr is equal to Vi(Df Di)/1 (V0 Df) where Vi ¼ initial volume in ml, Df ¼ final density, and Di ¼ initial density, and V0 ¼ 0.24 when Df ¼ 1.019, V0 ¼ 0.244 when Df ¼ 1.063, and V0 ¼ 0.246 when Df ¼ 1.09. Spin 45 k in 50.2 Beckman centrifuge rotor or equivalent for 24 h at 10 C. The VLDL is now in the top fraction and can be discarded. Harvest the orange layer of each tube and then pool together. Adjust the density to 1.063 with NaBr. To the remaining plasma, adjust the density to 1.063. Spin 45 k in a 50.2 Beckman centrifuge rotor for 24 h at 10 C. After spin, harvest the top orange LDL layer of each tube and pool together. Adjust the density to 1.09 with NaBr. Spin 35 k in a 50.3 Beckman centrifuge rotor for 16 h. Harvest the top layers and dialyze against LP buffer (150-mM NaCl, 1-mM EDTA pH 7.4). Sterilize with a 0.45 millipore syringe filter and store under argon gas. 3.1.1.4. Acetylation of LDL Acetyl-LDL is prepared from a reaction with acetic anhydride and used at a concentration of 50–100 mg/ml in all experiments. In a clean 3-ml stirring vial (Vial—Wheaton, Cat. no. 986257; stirring bar, VWR, Cat. no. 58949-272), place 20–22 mg of LDL. Add an equal volume of saturated sodium acetate. Add LP buffer (formulation below) to a final concentration of 18 mg/ml LDL. Stir briefly at room temperature to mix and take to a 4 C cold room. Determine the amount of acetic anhydride (Sigma A6404) to add. Volume of acetic anhydride required ¼ 2 amount in mg of LDL (e.g., 20-mg LDL required 40 ml of acetic anhydride). Aliquot more than the desired total amount of acetic anhydride and keep on ice. Every 10 min add 1/6 of the


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acetic anhydride in a dropwise manner to the diluted LDL (rinse the pipette tip in the protein solution). After the second or third addition, the solution should become cloudy and a precipitate will form. If the solution becomes too viscous, then add more LP buffer. Stir for an additional hour in the cold room. Place in a dialysis sac (12,000–14,000 MW cutoff) and dialyze against approximately 2 l of LP buffer. Change three times. Collect acetylated LDL and spin in a 1.5-ml microfuge tube for 3 min. Filter the supernatant through a 0.45-mm filter. Determine the protein concentration by a Lowry assay. LP buffer: dissolve 175 g of NaCl (Fisher S271-3) and 2.5 g of EDTA (BP121–500) in 20-l DI water. Adjust pH to 7.4 and chill to 4 C before use. It is important to extensively dialyze after acetylation using multiple bath changes. Keep stored under argon gas and at 4 C. We typically use AcLDL that is less than 0.01 units of endotoxin as measured by Lonza LAL endotoxin assay. 3.1.1.5. Additional Reagents The ACAT inhibitor is 58035 (3-[decyldimethylsilyl]-N-[2-(4-methylphenyl)-1-phenylethyl] propanamide (Ross et al., 1984). Prepare a 10-mg/ml stock in DMSO and use at a concentration of 10 mg/ml in all experiments. CHOP antibody: GADD153 (B-3), Mouse monoclonal IgG1, 200 ml/mg from Santa Cruz Biotechnology, Cat. no. sc-7351 (we have been using Lot # B2603). The Vybrant Annexin V/Propidium Iodide Apoptosis Assay kit can be purchased from molecular probes.

3.1.2. Method 3.1.2.1. Elicitation of primary macrophages Macrophages are obtained from 8 to 10-week-old female C57BL/6J mice. Harvest macrophages either 3 days after intraperitoneal (i.p.) injection of concanavalin A or 4 days after immunization with methyl-BSA (Cook et al., 2003). For the methyl-BSA elicitation, 2 mg/ml methyl-BSA in 0.9% saline is emulsified in an equal volume of CFA (DIFCO). Mice are immunized intradermally with 100 ml of the emulsion. After 14 days, the immunization protocol is repeated, except use IFA instead of CFA. Seven days later, the mice are injected i.p. with 0.5-ml PBS containing 100-mg methyl-BSA and then the macrophages are harvested 4 days after that by peritoneal lavage. For conA: Add 20 ml of stock (40 mg) to 480-ml sterile PBS and inject i.p. 3.1.2.2. Harvesting and culturing peritoneal macrophages Sacrifice mice by CO2 asphyxiation and pin mice down, abdomen side-up. Do not stretch mice too much when pinning as tightness may result in peritoneal leakiness during harvest. Sterilize abdomen by spraying with 70% EtOH. Lift skin with forceps and with scissors expose peritoneum without puncture. Inject peritoneum with 5 ml of ice-cold PBS and agitate stomach to mix. Hold skin up with forceps and push the pasteur pipette into the

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abdomen to aspirate and harvest macrophages. Rinse peritoneum to remove residual cells and filter cells with a 40-mm nylon cell strainer. Spin down cells and plate in medium containing 10% FBS and 20% L-cell conditioned medium. Discard any harvests that have contamination from the intestinal lumen or the blood. Wash away nonadherent cells (nonmacrophages) after 60 min of culture. On the day of plating, the wells should be 70–80% confluent with adherent cells and the cells can be used the next day. If the cells are more sparse, they can be grown in a medium for extra days and then used when the monolayers are more confluent. If the cells are plated too sparsely (100% confluent (i.e., overcrowded), biological artifacts can occur, but the experiment can be salvaged by washing the cells soon after harvest to dislodge the overcrowded cells. 3.1.2.3. Inducing ER stress by FC- loading for analysis of CHOP expression and apoptosis When cells are 80–90% confluent, rinse cells with warm PBS and incubate in a macrophage medium containing 50–100 mg/ml acetyl-LDL plus 10 mg/ml compound 58-035. As a control, incubate cells only with acetyl-LDL to generate foam cells which will be readily visible under the microscope. At desired timepoints (see Fig. 16.1), aspirate media and rinse cells in ice-cold PBS, and again aspirate and add sample lysis buffer (Laemmli buffer) to each well. Scrape cells (the lysate will become viscous), collect the lysate in the Eppendorf tube, and boil samples for 5 min. Freeze at 20 C or load on a 4–20% gel for immunoblotting of CHOP. For apoptosis, macrophages are assayed for early to mid-stage apoptosis by staining with Alexa 488-conjugated Annexin V. Late-stage apoptosis is determined by costaining with propidium iodide. View cells immediately at room temperature on an inverted fluorescent microscope with appropriate imaging software. Enumerate representative fields (five fields containing 1000 cells) per condition. Count the number of Annexin V- and PI-positive cells and express as a percent of the total number of cells in at least four separate fields from duplicate wells. An example of FC-induced CHOP expression and annexin positivity is shown in Fig. 16.1.

3.1.3. Notes/troubleshooting Because apoptosis is a rapid process, perform analysis immediately after staining. If fixation is required, incubate cells with annexin V prior to fixation and then add 2% paraformaldehyde in PBS. Elevated Annexin V and/or PI staining may reflect that apoptosis continues even after removing the plates from temperature-controlled incubators. Therefore, cells stained with Annexin V should not be kept to linger for prolonged times before measurement. Cells that maintain membrane integrity for longer incubation times may become positive for PI, as this dye can slowly permeate and enter

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A

0

5

10 h

CHOP

ACTIN

AcLDL + 58035

* 15 10 5 0 AcLDL + 58035

AcLDL

20

AcLDL

% Annexin positive MFs

B

Figure 16.1 CHOP expression and annexin staining of FC-loaded primary macrophages. Peritoneal macrophages were elicited and incubated in macrophage medium. At 90% confluence, cells were either loaded with AcLDL or alternatively FC-loaded (AcLDL þ 58-035) and (A) immunoblotted for CHOP at indicated times. Blots were stripped and reprobed for actin as a protein loading control. (B) Monolayers were in parallel stained with annexin V (green) and propidium iodide (red) at 16 h to detect mid-stage and late-stage apoptosis. The graph is a quantitation of annexin V-positive, propidium iodide-negative (apoptotic) macrophages in AcLDL cholesteryl ester (CE)loaded, or FC-loaded macrophages. *p < 0.05.

intact cells. Thus, add PI solution just before the analysis. Different batches of AcLDL will vary with respect to their potency of UPR induction and apoptosis. This may be due to variability in the donor that may include a drug or serum component that promotes cell survival.

4. The UPR and Macrophage Death at the Murine Aortic Root Advanced atherosclerotic lesions consist of a heterogeneous population of cell types, each of which is differentially susceptible to ER stress. Precise analysis of UPR gene expression in plaque therefore requires methodology to selectively capture cell-type specific mRNA. Laser-capture

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microdissection (LCM) is a method to harvest subpopulations of tissue cells by direct microscopic and immunohistochemical identification (Espina et al., 2006). IHC-navigated cell-type identification is complicated by the fact that immunoidentification requires prolonged antibody incubations that expose tissue sections to the activity of exogenous and endogenous RNases. Fisher and colleagues have cleverly utilized rapid immunostaining protocols with high-affinity antibodies. This approach permits detection and capture from the same section with limited loss in RNA integrity and yield (Trogan and Fisher, 2005). However, in some cases, immunodetection of antigen may not be amenable to rapid staining. This may be due to low antibody affinity or low expression levels of antigen. Herein, we describe an alternative approach that couples immunodetection of macrophage markers with laser-capture of UPR mRNA on separate sections. This method allows users to incorporate prestandardized immunohistochemical staining protocols with LCM. IHC is performed in tandem with LCM on separate but closely cut (6 mm) adjacent sections. As UPR stress signaling is closely linked to cellular and macrophage apoptosis, we subsequently outline a protocol for monitoring macrophage apoptosis in ER-stressed lesions by double immunofluorescence staining.

4.1. Method: Laser-capture microdissection of the UPR in atherosclerosis 4.1.1. Materials and Equipment 4.1.1.1. Materials required for harvesting and preparation of lesional sections from the aortic root Isoflurane and mouse container for anesthesia. Dissecting platform and dissecting microscope for aortic root harvest. Small animal surgical tools, including fine scissors and forceps. A 1-ml syringe for drawing blood from the heart (prefill with 15 ml 0.5 M EDTA pH 8.0). PBS in a 10-mL syringe with a 24-gauge needle for perfusion of organs. Optimal cutting temperature (OCT) embedding medium. Cryostat with disposable microtome blades (we recommend Leica). Fisher Plus coated microscope slides (Fisher, Cat. no. 12-550-15). All experiments should be performed in compliance with NIH guidelines for the care and use of laboratory animals. 4.1.1.2. Reagents for staining and immunohistochemistry of frozen sections For rapid staining of slides prior to LCM: The AM1935 LCM Staining Kit from Ambion includes reagents for processing 80 slides. This kit avoids exposure of tissue sections to pure water and potential reactivation of endogenous nucleases. The active ingredient during this staining protocol is Cresyl Violet, which permits nuclear staining and visualization of cell morphology from dehydrated tissue. For IHC: Normal goat serum can be used for blocking and can be purchased from Vector Laboratories, Cat. no.


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S-1000. For detection of F4/80þ phagocytes: Anti-F480 can be purchased from Serotec (biotin-conjugated). Biotin-Alexa 488 is sold by the Molecular Probes division of Invitrogen or alternatively, biotin-FITC may be used. 4.1.1.3. Equipment and reagents during laser-capture microdissection LCM work station: This protocol was optimized using the Zeiss PALM (Positioning and Ablation with Laser Microbeams) laser microdissection system equipped with fluorescence imaging. Bring gloves and change frequently. Bring RNA lysis buffer, pipette and tips, and dry ice to store the harvested sample. The LCM work station should be situated in a temperature-controlled and low-humidity work space. 4.1.1.4. Reagents for RNA purification and quantitative RT-PCR From Ambion, AM1931 RNAqueous-Micro Kit for RNA purification downstream of LCM. RNA integrity can be monitored using the RNA 6000 Nano Lab Chip kit from Agilent per manufacturer’s instructions. For reverse transcription, you will need standard oligo-dT (from Invitrogen Cat. no. 18418-012) and dNTP mix (Invitrogen, 10 mM dNTP MIXPCR grade, Cat. no. 18427-013) and lamda superscript II (RNaseH reverse transcriptase # 18064-014). For qPCR: The forward primer for CHOP is CCA CCA CAC CTG AAA GCA GAA. The reverse primer is AGG TGA AAG GCA GGG ACT CA. If using probe: CTG GTC CAC GTG CAG TCA TGG. For control CypA, the forward primer is GGC CGA TGA CGA GCC C and reverse primer: TGT CTT TGG AAC TTT GTC TGC AA. The probe for CypA is TGT CTT TGG AAC TTT GTC TGC AA. The PCR conditions for Chop cDNA detection are 95 C for 1 min, 58 C for 30 s, and 72 C for 30 s, for 40 cycles.

4.1.2. Method 4.1.2.1. Aortic root processing Feed B6 Ldlr/ mice (Jackson, stock # 002207) high-fat, high-cholesterol diet from Harlan Teklad for 12 weeks (diet #88137) starting at 8–10 weeks of age. Anesthetize mice under isoflurane and open thorax. It is important to be relatively expedient during the remaining steps to limit RNA degradation. Remove the blood from the left ventricle and perfuse at physiological pressure with PBS. Remove heart and aorta and cut heart in half, parallel to the aortic valve. Place the upper half on the heart in an embedding mold filled with OCT cryoembedding medium. Position the heart with the cut surface on the bottom of the embedding mold. Fill the embedding mold with OCT and freeze on dry ice. Store specimens at –80 C. Set cryostat temperature at 25 and 30 C for knife. Clean the sectioning platform with 70% EtOH. Adjust the tissue block surface parallel to the plane that the blade cuts. Cut the thick sections until you see the valves. Section tissue blocks at 6 mm thickness and mount on positively charged slides. We generally cut 50 sections up the

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aortic root, of which 6 are used for H&E staining and morphometric and area analysis. The remaining 38 sections are used for LCM (half for IHC staining and the other half for capture). 4.1.2.2. Immunodetection of F4/80þ phagocytes Allow frozen sections to air-dry for 2 min and then fix in cold acetone for 5 min. Block in 3% goat blocking serum (made in PBS) and then incubate slides with antimouse F4/80 IgG (1:100 dilution) in 3% goat serum. Rinse slides in PBS 3 times for 5 min each. Incubate slides with streptavidin FITC antibody in blocking serum for 1.5 h and subsequently rinse slides in PBS 3 times for 5 min each. Image under a fluorescence microscope to identify the phagocyte regions of interest. Capture images from immunostained sections and identify immunopositive regions of interest for later alignment with the immediately adjacent 6 mm serial section. In parallel analyses, we have confirmed that the majority of directly adjacent sections exhibit equivalent immunostaining for F4/80þ regions of interest when cut at 6 mm. A similar approach has been applied for LCM combined with IHC in brain (Kase et al., 2007). For parallel LCM slides, perform rapid staining as described by Ambion (AM1935 LCM Staining Kit) to provide contrast and for alignment with the identified regions of interest. After staining as per the manufacturer’s instructions, the slides are dehydrated in increasing concentrations of EtOH and finally, xylene. We typically process 4 slides at a time by incubating 1 min each in 70% EtOH, followed by 95% EtOH, followed by 100% EtOH, and then two 1-min incubations in fresh xylene. Slides are then air-dried for 10 min and immediately taken to the LCM work station. Slides are stored in a slide box with a desiccant. 4.1.2.3. LCM on PALM To isolate RNA from frozen sections by LCM, it is important to minimize RNA degradation during sample preparation. Capture is performed on a PALM series LCM instrument with fluorescence imaging capability. After marking the region of interest by immunostaining as described above, quickly identify the parallel ROI in the rapid-stained and dehydrated immediate serial section for capture. For laser-capture, this is best done at 40 magnification. Catapult the specimen into a 500-ml tube that is mounted above the slide. The tube should be filled with a RNA catapult buffer (RNA lysis buffer) and later stored on dry ice prior to buffer crystallization. Do not let the tube sit at room temperature longer than 30 min. Alternatively, one can use the PALM zeiss membrane cap and solubilize RNA at a later time. Select the region of interest by demarcating the area as described in the PALM software program. Settings on the PALM must be optimized. We have used the following settings: UV energy -63, Focus, -35. Perform catapult of tissue into the tube and store the specimen in dry ice. After 12 weeks of western diet and capture of intimal F4/80þ regions from the 20 rapid-stained sections described above, more than

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enough RNA should be obtained for an analysis of UPR gene expression. For a thorough analysis and gene expression profiling, we recommend that at least 1 106 square microns of surface area be collected. RNA yield and integrity will vary as described below. 4.1.2.4. RNA processing RNA integrity can be examined on a Nano Lab Chip kit from Agilent. Total RNA is isolated using the RNAqueous-Micro Kit from Ambion and reverse-transcribed into cDNA using standard procedures and materials (Invitrogen). Perform quantitative PCR for Chop and control mRNA cyclophilin A (CypA). In some cases, two rounds of linear amplification may be needed prior to qPCR. When designing future primers for PCR of LCM-captured material, you want to limit the size of the amplicon to less than 300 base pairs if possible due to increased RNA degradation. An example of qPCR of CHOP expression from LCM of atherosclerotic lesions, normalized to CypA, is shown in Fig. 16.2.

A

B

C

D

8

Relative F4/80 + CHOP mRNA

lumen

6

*

4 2 0

10 week 15 week

Figure 16.2 Laser-capture microdissection of CHOP in F4/80þ phagocytes of atherosclerotic plaque. (A) Aortic root lesions from LDLR/ mice were stained with F4/80 and (B) a parallel section captured by LCM. Sections were cut at 6-mm thickness. (C) An image of the lesion after capture is shown. (D) CHOP qRT-PCR of targeted F4/80þ areas reveals increased CHOP expression after increased diet duration. *p < 0.05.

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4.1.3. Notes/troubleshooting Perform the procedure under RNase-free conditions and with RNase decontamination solutions. Change disposable gloves frequently. Note that high humidity or moisture promotes activation of endogenous RNases. Therefore, minimize slide exposure to high room temperature. Keep cryosections at 80 C to prevent RNA degradation and microdissect cells immediately after staining. Also, change cryostat blade between processing of separate murine samples to avoid sample contamination. Also add RNase inhibitors to staining reagents and minimize the staining time. Expected RNA yield can vary depending on sample integrity. Decreased RNA yield may occur by degradation of RNA before processing of sample for LCM. Therefore, minimize the time required for tissue harvesting and for microdissection at PALM station. A single section should not be exposed to room temperature for longer than 20 min. To test the quality of the RNA before LCM, simply scrape the entire section into a RNA extraction buffer and analyze for 18S/28S RNA. LCM can be performed on paraffin-embedded PFA-fixed sections; however, you will need special extraction techniques for the captured material. This will lower RNA yield and integrity and likely entail a protease treatment to break cross-links formed during fixation.

4.2. Method: Detection of apoptotic macrophages in ER-stressed plaque 4.2.1. Materials 4.2.1.1. Reagents for TUNEL staining Bovine serum albumin, doubledistilled water, DNase solution (300 U/ml in 50-mM Tris–HCl, pH 7.4, 1 mg/ml BSA) ethanol, GVA (glycerol-vinyl-alcohol) mounting solution, Histopen, Hoechst 33342 (2 mg/ml in PBS), phosphate-buffered saline, protease K (PCR-grade, nuclease-free at 20 mg/ml in 10 mM (Tris–HCl pH 7.4), 0.1% TX-100, Tunel Assay Kit from Roche 12-156-792-910, and xylene. 4.2.1.2. Reagents for macrophage immunohistochemistry after TUNEL staining Mac-3 is from BD Biosciences, San Diego, California, USA. Rabbit pan antimacrophage antibody (AIA31240) is from Accurate Chemical and Scientific Corporation.

4.2.2. Method 4.2.2.1. Section preparation This protocol is for paraformaldehydefixed, paraffin-embedded lesions, which in our hands have yielded the best results in terms of lesional morphology and preservation of apoptotic DNA. If frozen sections are utilized from slides prepared for LCM above, then we advise first fixing sections with PFA, followed by TUNEL staining


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below. For paraffin-embedded and PFA-fixed sections: Under chemical hood, dewax samples in xylene 3 times, for 10 min each (alternatively perform 12 sequential dips). Rehydrate samples by dipping in 100% EtOH 2, followed by 2 dips in 95% EtOH, and finally, 10 dips in clean H2O. Hold the samples in clean H2O. Circle the samples on slides with histopen for downstream TUNEL and IHC. 4.2.2.2. TUNEL staining of atherosclerotic lesions Add 100 ml of protease K solution to each slide and incubate for 30 min at 37 C. Then, rinse 2 in PBS and hold in PBS solution. First process positive control: Add a 50-mL DNase solution and leave for 10 min at room temperature. Wash 2 in PBS. Next, dry the area around all samples, including positive and negative control. Prepare Roche TUNEL reaction mix as per the manufacturer’s instructions. This is done by mixing 50 mL of vial 1 (TDT enzyme solution) þ 450 mL vial 2 (TMR-red dUTP label solution). This is for nine samples and a positive control. TUNEL staining is performed by adding 50 ml of enzyme mixture to each sample and to positive control. Do not add TUNEL reaction mixture to negative control. Incubate all samples plus controls 60 min at 37 C in a dark, humidified incubator. Rinse the slides in PBS 3 (or 12 dips). Add Hoechst solution to each slide for 5 min. Wash in PBS two times and coverslip using GVA mounting solution. Check for apoptosis per microscopy (excitation 520–560, detection 570–620; may store slides at 4 C in dark prior). 4.2.2.3. Stain with macrophage-specific antibody After performing TUNEL labeling, sections are blocked in 1% BSA and 1% normal goat serum and stained with rat antimouse Mac-3 mAb at a dilution of 1:50 dilution overnight at 4 C h, followed by an incubation of a secondary goat antirat IgG-FITC. Alternatively, macrophages are detected using a rabbit pan antimacrophage antibody (AIA31240). To determine antibody specificity, a preimmune or isotype control should be included in parallel sections. 4.2.2.4. Quantification of TUNEL-positive phagocytes TUNEL-positive lesions are enumerated as TUNEL-positive nuclei. Apoptotic images are merged using Photoshop CS analysis software (version 8.0; Adobe), and the number of TUNEL- and HOECST-positive cells is counted from each lesion. Apoptotic phagocytes are analyzed by fluorescence microscopy by identification and colocalization of fluorescein and TUNEL-red staining after image overlay in Adobe Photoshop. The percentage of TUNEL-positive phagocytes is determined by counting the number of double-stained phagocytes and total phagocytes per aortic root cross-section. An example of TUNEL-positive macrophage enumeration is shown in Fig. 16.3.

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A

DNA

TUNEL

Mf

% TUNEL + MFs

B 3

* 2

1

0 4

12

Figure 16.3 Staining TUNEL-positive macrophages from PFA-fixed paraffin-embedded sections. (A) Representative micrographs show TUNEL-positive signal (red) in nuclei (blue) of aortic root lesions from lesions. Blue is Hoechst nuclear dye. Green is F4/80. (B) Enumeration of TUNEL-positive F4/80þ macrophages from early (4 week Western Diet fed) and advanced (12-week Western Diet fed) lesions. *p < 0.05.

4.2.3. Notes/troubleshooting It is important to avoid nonspecific TUNEL reactivity by optimization of the enzyme concentration or duration of enzyme incubation. Under optimal conditions, TUNEL-positive nuclei in atherosclerotic plaque are typically less than 2% (Kockx and Knaapen, 2000). Optimization of protease K treatment is not only important for accessibility of the TDT enzyme but also for preservation of macrophage epitopes, which can be destroyed with high levels of protease. Finally, apoptosis is a function of lesion maturity. Increased levels of TUNEL and caspase positive cells are found in advanced lesions. Given the heterogeneity of plaque size on the vascular wall, it is important that analyses between two experimental groups are done in lesions at similar stages of development (e.g., equal lesion size). This is critical to avoid measuring differences that manifest simply due to changes in lesion maturation stage.


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5. Conclusion and Future Methodological Advances Future methods will exploit endogenous in vivo ER stress reporters that eliminate processing artifacts that are often the result of sample degradation or nonspecific epitope recognition. For example, a transgenic fluorescent reporter that is induced during ER stress in vivo would eliminate extra secondary detection steps that are required for tissue analysis. In this context, Iwawaki, Miura, and colleagues have created a mouse that is transgenic for a UPR reporter. This consists of the UPR X-box binding protein 1 (XBP-1) fused to venus fluorescent protein and driven by the universal chicken bactin promoter (Iwawaki et al., 2004). Murine cells that are transgenic for this marker specifically fluoresce after activation of the UPR and therefore can be used to monitor physiological and pathological ER stress in vivo. Thus, information about the level of ER stress during atherosclerosis could be obtained by crossing the ERAI-transgenic mouse with hyperlipidemic mouse models. The combination of in vitro mechanistic studies of ER stress with in vivo models of physiological significance are together key to understanding how ER stress modulates atherosclerotic progression.

ACKNOWLEDGMENTS These methods were developed in the laboratory of Dr Ira Tabas at Columbia University Medical Center. I thank and credit this manuscript to my training with Dr Tabas. Thanks also to Dr Alain Borczuk for LCM consultation and Manikandan Subramanian for critical reading of the manuscript. Grant Support from NIH NHLBI 1K99HL097021-01.

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Detecting Autophagy in Response to ER Stress Signals in Cancer Marıá Salazar,*,† Sonia Hernańdez-Tiedra,* Sofıá Torres,* Mar Lorente,* Manuel Guzmań,*,† and Guillermo Velasco*,† Contents 1. Introduction 2. Monitoring Autophagy Upon ER Stress 2.1. Electron microscopy 2.2. Analyzing the autophagy protein LC3 to monitor autophagy 2.3. Note: Analyzing autophagy through changes in the phosphorylation of mTORC1 substrates? 2.4. Monitoring autophagy by knockout or knockdown of essential autophagic proteins 3. Monitoring Autophagy Upon ER Stress In Vivo 3.1. Generation of tumor xenografts 3.2. Analysis of autophagy in samples obtained from tumor xenografts 3.3. In vivo silencing of autophagy or ER stress genes 3.4. Generation of tumor xenografts with transformed embryonic fibroblasts derived from genetically modified mice References

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Abstract Different physiological and pathological situations that produce alterations in the endoplasmic reticulum, lead to a condition known as ER stress. ER stress activates a complex intracellular signal transduction pathway, called unfolded protein response (UPR). UPR is tailored essentially to reestablish ER homeostasis. However, when persistent, ER stress can switch the cytoprotective functions of UPR into cell death promoting mechanisms. One of the cellular mechanisms that are regulated by ER stress is autophagy. Autophagy is a cellular process by which different cytoplasmic components including organelles are targeted for * Department of Biochemistry and Molecular Biology I, School of Biology, Complutense University, Madrid, Spain { Centro de Investigacioń Biome´dica en Red sobre Enfermedades Neurodegenerativas (CIBERNED), Madrid, Spain Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00017-0

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degradation to the autophagosomes. Interestingly, like ER stress, autophagy can be a protective or a cell death promoting mechanism. Recently, a variety of anticancer therapies (including those that stimulate ER stress) have been shown to activate autophagy in tumor cells, which has been proposed to either enhance cancer cell death or act as a mechanism of resistance to chemotherapy. In this chapter, we will describe some of the procedures that are currently used to analyze autophagy as well as some of the experimental approaches that can be undertaken to investigate the connection between ER stress and autophagy in cancer.

1. Introduction Autophagy is a cellular process by which entire cytoplasmic portions—including organelles and other cytoplasmic components—are engulfed within double-membrane vesicles designated autophagosomes (He and Klionsky, 2009). The maturation of these vesicles involves their fusion with lysosomes, which leads in turn to the degradation of the autophagosomal components by the lysosomal degradative enzymes (Levine and Kroemer, 2008; Mizushima et al., 2008). The molecular mechanisms responsible for the regulation of autophagy have not been completely elucidated yet, although genetic and biochemical analyses performed during the last few years have identified several autophagy genes (Atg) that participate in the regulation of this cellular process (He and Klionsky, 2009). The final outcome of the activation of the autophagy program is highly dependent on the cellular context and the strength and duration of the stress-inducing signals. Thus, besides its role in cellular homeostasis, autophagy can be a form of programmed cell death or play a cytoprotective role, for example, in situations of nutrient starvation (Eisenberg-Lerner et al., 2009; Maiuri et al., 2007). Of relevance, different situations that induce ER stress lead in turn to autophagy stimulation (He and Klionsky, 2009). As discussed along this issue of Methods in Enzymology, the ER stress response is initially a protective mechanism. However, when the intensity or duration of the ER damage cannot be restored by this response, ER stress can also lead to cell death (He and Klionsky, 2009). Likewise, autophagy can help to cope with ER stress (e.g., contributing to the elimination of unfolded or aggregated proteins) or participate in the mechanism of ER stress-induced cell death (Matus et al., 2008; Verfaillie et al., 2010; Winslow and Rubinsztein, 2008). In cancer, autophagy has been proposed to play a tumor suppressor function by providing the minimal supply of ATP required for DNA repair, preventing oxidative stress and reducing intratumoral necrosis and local

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inflammation (Maiuri et al., 2009; Mathew et al., 2007; Morselli et al., 2009). In addition, different anticancer treatments (including those that stimulate ER stress) activate autophagy in tumor cells, which has been proposed to either enhance cancer cell death or act as a mechanism of resistance to chemotherapy (Maiuri et al., 2009; Mathew et al., 2007; Morselli et al., 2009; Salazar et al., 2009; Verfaillie et al., 2010).

2. Monitoring Autophagy Upon ER Stress Several experimental approaches can be used to analyze whether a certain stimulus (and in particular those situations that activate ER stress) can also induce autophagy in cultured cells. In most cases, it will be necessary to combine two or more of these approaches to confirm that this cellular process becomes activated. In this section, we will describe some of the most frequently used methodologies to analyze autophagy.

2.1. Electron microscopy Autophagosomes (the characteristic double-membrane vesicles containing cytoplasmic components) have been observed by electron microscopy in mammalian cells since the 1950s cells (Klionsky, 2007) and this methodology still remains as one of the most conclusive ways of confirming the presence of autophagosomes in the cell. In Fig. 17.1A we show microphotographs of cells subjected to an autophagic stimulus in which the morphological features of early and late autophagosomes can be appreciated. Nevertheless, a detailed description of electron microscopy protocols is out of the scope of this chapter and will not be included here.

2.2. Analyzing the autophagy protein LC3 to monitor autophagy The microtubule-associated protein 1 (LC3 or Atg8) is an autophagy protein initially synthesized in its unprocessed form (proLC3). Upon autophagy stimulation, the C terminus of LC3 is cleaved by the Atg4 proteases to form LC3-I, which is then modified (with the sequential participation of at least two additional autophagy proteins) to form the phosphatidylethanolamide (PE)-conjugated form of LC3, named LC3-II (He and Klionsky, 2009). LC3-II remains associated to the autophagosomal membrane until this vesicle fuses with the lysosome to form the autophagolysosome where LC3-II is digested together with the rest of the autophagosomal content. Conversion of LC3-I into LC3-II and changes in the subcellular

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Figure 17.1 Techniques used to analyze autophagy in cultured cells. (A) Electron microscopy. Representative electron microscopy photomicrographs of U87MG cells after 6 h treatment with 4 mM D9-tetrahyrocannabinol (THC, a potent autophagy inducer; bars: 500 nm). Note the presence of early (open arrows) and late (close arrows) autophagosomes in THC but not vehicle (veh)-treated cells. Low panels: Details of early (left) and late (middle and right) autophagosomes. (B) LC3 immunostaining. Effect of THC (18 h) treatment on LC3 immunostaining (green) of U87MG cells (bar: 20 mm). Note the presence of LC3 puncta in cells treated with the autophagic stimulus but not in vehicle-treated cells. (C) LC3 lipidation. Effect of E64d (10 mM) and pepstatin A (PA; 10 mg/ml) on THC-induced LC3 lipidation of U87MG cells. Note the presence of the LC3-II band (higher electrophoretic mobility). In agreement with the stimulation of dynamic autophagy in this experimental setting, incubation with the inhibitors of the lysosomal proteases enhances LC3-II levels.


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distribution of LC3 are thus two hallmarks of autophagy that can be monitored to determine whether this cellular process becomes activated. 2.2.1. Analysis of LC3 subcellular location by immunofluorescence As described above, upon induction of autophagy, LC3-II becomes associated to the autophagosomal membrane. This change in the subcellular distribution of LC3 can be followed by immunoflourecence microscopy (a characteristic pattern of LC3 puncta can be observed in autophagic cells stained with anti-LC3 antibodies; Fig. 17.1B). Alternatively, cells can also be manipulated to express a recombinant form of LC3 fused with GFP (or other fluorescent protein). This recombinant form of LC3 is processed in the same way than endogenous LC3, and therefore when these cells are subjected to an autophagic stimulus exhibit a characteristic pattern of GFP–LC3 punta. 2.2.1.1. Analysis of endogenous LC3

Required material Cells of interest: Autophagy can be detected by this procedure in many different cell types (we will describe the protocol used for detecting LC3 in the human glioma cell line U87MG). Nevertheless, we recommend selecting for this type of experiment cells with large cytoplasm (in which the presence of LC3 dots can be more easily appreciated). Cell culture medium: Certain cell types rapidly undergo autophagy when serum is removed from the medium. If the experimental conditions require incubation in a defined (low-serum content) medium, we recommend including an additional control with cells incubated in complete medium. Anti-LC3: We recommend using the rabbit polyclonal anti-LC3 antibody from MBL (Naka-Ku Nagoya, Japan, ref. PM036). Secondary antibody: Alexa 488 goat anti-rabbit IgG (Invitrogen). Other reagents: PBS, 4% paraformaldehyde, 10% Triton X-100 solution, goat serum, Mowiol (mounting medium), Hoechst 33342 (Stock 5 mg/ ml; Invitrogen). Procedure Grow cells in sterilized cover glasses. After incubation with the desired stimuli, carefully remove the medium, rinse with PBS and fix the cells with 4% paraformaldehyde in PBS for 10 min. Wash twice with PBS and permeabilize the cells with 0.03% Triton X-100 in PBS containing 3% BSA for 30 min. Wash again with PBS and incubate with the primary antibody (polyclonal anti-LC3, MBL; ref. PM036; 1:300 in PBS; we recommend using 30 ml of diluted antibody for each cover glass). After incubation for 1 h at room temperature (or overnight at 4 C), wash with

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PBS for 30 min and incubate with the secondary antibody (anti-rabbit 1:500 in PBS) and Hoechst 33342 (1/4000) for 1 h, wash with PBS for 30 min and mount with Mowiol. Analysis in the confocal microscope We recommend using a confocal microscope rather than a microscope of fluorescence to analyze LC3 immunostaining. The size and number of the LC3 puncta present in the autophagic cells can vary depending on the autophagic stimulus and the cell type. A strong autophagic stimulus (e.g., treatment with D9-tetrahydrocannabinol) is expected to produce a large number of LC3 puncta in the cells (see Fig. 17.1B). However the presence of LC3 dots could be less evident in response to other stimuli. We recommend quantifying the percentage of cells exhibiting at least five to six countable LC3 dots per cell. If controls have some basal autophagy, the number of LC3 puncta per cell rather than the total number of cells displaying puncta should be determined (in this case a threshold based on the average number of puncta found in control cells can be used to determine the rate of basal autophagy in a particular experimental situation). 2.2.1.2. Analysis of recombinant GFP–LC3 expressed in the cell of interest In some cell types, or under some experimental conditions, endogenous LC3 puncta are difficult to detect through immunofluorescence. In those cases, an alternative possibility to monitor autophagy is analyzing the cellular distribution of recombinant GFP–LC3 (or of other fluorescent proteins fused with LC3). This experimental approach makes easier to detect autophagy in some cells types. However, this procedure has some drawbacks. Firstly, transfection with a GFP–LC3 plasmid (or viral transduction with an adenoviral or retroviral vector carrying this construct) may activate stress pathways in the cell, which could stimulate autophagy. Thus, most cells— even in nutrient-rich conditions—display some basal levels of GFP–LC3 puncta. This makes it more tedious to quantify the number of GFP–LC3 positive cells. In this experimental setting it is particularly important to use appropriate controls. In addition, we recommend determining the number of GFP–LC3 puncta per cell (and set for quantification a threshold based on the average number of puncta found in control cells) rather than simply quantifying the total number of cells displaying puncta.

Required material Cells of interest: Autophagy can be detected by this procedure in many different cell types. Nevertheless, we recommend using for this type of experiment cells with large cytoplasm. Packaging cells: In the case of transduction with an adenoviral vector carrying the GFP–LC3 construct, HEK293A packaging cells are required for viral particle amplification.


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Cell culture medium: Same considerations than for the analysis of endogenous LC3 should be taken into account for the analysis of GFP–LC3 distribution. Please note that cells will be plated in medium without antibiotics prior to initiate the transfection or viral transduction procedures. DNA of interest: DNA of high quality (ratio Abs 260/280 > 1.9) is recommended for transfecting mammalian cells. Transfection reagents: LipofectamineTM 2000 (Invitrogen)

Transfection Each cell type may require a different transfection method. Please use the one that works better in your cell type. We use LipofectamineTM 2000 (Invitrogen) for transfecting U87MG cells. Adenoviral particle amplification Adenoviral vectors are frequently used to transiently overexpress proteins in mammalian cells and therefore the use of these vectors results particularly interesting in cells that are difficult to transfect. The disadvantage of this methodology is that the purification and titration of adenoviral vectors is tedious and time-consuming. In addition, adenoviruses are quite labile and their title rapidly decreases even under the appropriate storage conditions. These drawbacks can be circumvented by using fresh viral particleenriched crude supernatants. We include here a brief description of the procedure used to infect U87MG cells with this type of viral supernatant. Packaging cells Plate HEK293A (packaging cells) in 60 cm2 dishes at approximately 90% confluence. Infect the cells the next day with purified adenoviral particles (or with a supernatant obtained from a previous infection). Two days after the infection numerous cells will appear rounded indicating that have been efficiently infected and are producing adenoviral particles. Collect the viral particle-enriched medium, filter it using a 45-mm diameter filter, and add directly to the cells to be transduced with the GFP–LC3 recombinant protein (the adenoviral vector-enriched supernatant can also be frozen and stored at 80 C). Infection of target cells Plate cells to be infected with the adenoviral vector carrying the GFP–LC3 construct at 80% confluence. At the next day, aspirate the medium and add 7 ml of the viral supernatant (obtained from one confluent- HEK293A 60 cm2 dish) to one 60 cm2 dish, incubate at 37 C/5% CO2 in a cell incubator for 4–6 h and replace the medium in which cells were infected by

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complete medium. 48 h after transfection/transduction, cells can be subjected to an autophagic stimulus. GFP–LC3 puncta can be analyzed in a fluorecence (or confocal) microscope. 2.2.1.3. Analyzing ER stress and autophagy markers by immunofluorescence ER stress as induced by certain stimuli is accompanied by morphological changes (normally dilation) in this organelle. This dilation can be monitored by immunofluorecence (e.g., using an antibody raised against protein disulphide isomerase, PDI, a protein located in the lumen of the ER). As mentioned above, ER stress has been shown to activate autophagy. By double staining with anti-PDI and anti-LC3 antibodies, ER stress and autophagy can be monitored in parallel at different timepoints. This experimental approach permits determining whether one of the two cellular events precedes the other in response to a certain stimulus (see Fig. 17.2A). A 0h

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Figure 17.2 Investigating the connection between ER stress and autophagy. (A) Timecourse analysis of PDI and LC3 immunostaining. Effect of THC on PDI (red) and LC3 (green) immunostaining of U87MG cells at different time-points (bar: 20 mm). Note that the increase in PDI dots (a characteristic of ER stressed cells with enlarged ER) occurs at earlier time-points than the increase in LC3 puncta, suggesting that ER stress precede autophagy in the mechanisms of action of this anticancer agent. (B) Silencing of ER stress genes. Effect of THC (18 h) on LC3 immunostaining (green) of U87MG cells transfected with control (siC) or p8-selective (sip8) siRNA. [p8 is an ER stress-related transcription factor that mediates the proautophagic actions of THC]. To identify cells that had been transfected, a red-fluorescent stable control siRNA was included in the transfection mix together with siC or sip8 in 1:4 proportion (red dots within the cells correspond to red-fluorescent control siRNA; bar: 20 mm). Note that THC induces autophagy in siC- but not in sip8-transfected cells.


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2.2.2. LC3 Western blot Western blot analysis of LC3 lipidation is perhaps the most widely used procedure to monitor autophagy. Required material Cells of interest: Autophagy can be detected by this procedure in many different cell types (we will describe the protocol used for detecting LC3 in the human glioma cell line U87MG). Lysis buffer: 50 mM Tris–HCl, pH 7.5, 1 mM phenylmethylsulfonyl fluoride, 50, mM NaF, 5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 0.1% Triton X-100, 1 mg/ml leupeptin, 1 mM EDTA, 1 mM EGTA and 10 mM sodium b-glicerophosphate, 0.1% (v/v) b-mercaptoethanol (add fresh) and 200 mM PMSF (add fresh). Anti-LC3: We recommend using the anti-LC3 rabbit polyclonal antibody (MBL, Naka-Ku Nagoya, Japan, ref. PM036). To note, certain anti-LC3 antibodies have lower sensitivity to detect LC3-I than others. Loading control: Tubulin can be used as loading control (We use anti-alpha tubulin mouse monoclonal antibody from Sigma). Other reagents: Running buffer (Gly 200 mM; Tris–base 25 mM; SDS 0.1%) Transfer buffer (Gly 200 mM; Tris–base 25 mM; Methanol 20%) TBS 10 (60.6 g/l Tris–base; 58.4 g/l NaCl) and Tween 20 (1:1000) PVDF membranes for protein transfers (Amersham Hybond-P) Cell stimulation Culture conditions and treatment with the different stimuli should be adjusted for each type of treatment. Lysates Place dishes in ice (4 C), wash twice with PBS, and remove the medium. Add lysis buffer (normally 60 ml per 60 cm2 plate) and detach the cells using a cell scraper. Collect lysates in Eppendorf tubes, vortex for a couple of minutes and centrifuge for 15 min (15,000g, 4 C) in order to discard the cell debris and quantify protein concentration. Prepare samples with 25–30 mg of protein and the corresponding volume of loading buffer containing SDS and b-mercaptoethanol. Boil samples at 90 C for 5 min before loading in a polyacrylamide gels. Note: LC3-I is more labile than LC3-II and gets more easily degraded in PAGE-SDS sample buffer. We recommend preparing electrophoresis samples (including heating at 90 C) rapidly after cell lysis. Avoid freezing and thawing the sample more than once.

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Electrophoresis and Western blot For detecting LC3 (18–16 kDa) we recommend using 15% acrilamide electrophoresis gels and transferring the proteins (2 h at constant amperage—250 mA per container) to PVDF membranes. After washing block membranes with 5% fat-free milk in TBS-Tween 0.1% for at least half hour, incubate overnight at 4 C with the primary anti-LC3 antibody (1:1000 in TBS-Tween 0.1%), wash the membranes with TBS-Tween 0.1%, incubate with the secondary antibody and follow the standard ECL procedure to reveal the bands corresponding to LC3. Interpretation of LC3 Western blots LC3-II has higher electrophoresis mobility than LC3-I. Thus, a band of apparent MW lower than LC3-I (corresponding to the lipidated form of LC3) is detected when autophagy is induced (see Fig. 17.1C). As mentioned above, some anti-LC3 antibodies have different affinity for LC3-I than for LC3-II. Moreover, total levels of LC3 do not necessarily change, as there might be an increase in the conversion of LC3-I to LC3-II or a decrease in the level of LC3-II relative to LC3-I due to the degradation of this latter form of LC3 in the lysosomes (see below). Thus, we recommend evaluating the changes in the amount of LC3-II rather than in the ratio LC3-II/LC3-I. Dynamic autophagy It is important to bear in mind that autophagy is a dynamic process (i.e., autophagosome formation is followed by its degradation upon fusion of these vesicles with the lysosomes). Accordingly LC3-II—that is present in the autophagosomal membrane—becomes degraded in the autophagolysosome. Thus, one way of confirming that dynamic autophagy—rather than accumulation of autophagosomes—is occurring is to verify that the amount of LC3-II present in the cell increases when the lysosomal proteases are inhibited. To this aim, cells can be treated with inhibitors of the lysosomal proteases [e.g., with pepstatin A (10 mg/ml) and E64d (5 mg/ ml) see Fig. 17.1C] or with pharmacological inhibitors of the fusion of autophagosomes and lysosomes [e.g., Bafilomycin A1 (0.1 mM)]. 2.2.2.1. Note: Analyzing ER stress and autophagy markers by electrophoresis Activation of ER stress is associated with changes in the levels and/or activity of several ER stress-related genes. For example, in many experimental settings eIF2a becomes phosphorylated upon exposure to ER stress (see other chapters of this issue of Methods in Enzymology for further details). By analyzing some of these ER stress markers (e.g., eIF2a phosphorylation) and LC3-II lipidation, ER stress and autophagy can be


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monitor in parallel at different time-points. This experimental approach permits determining whether one of the two cellular events precedes the other in response to a certain stimulus.

2.3. Note: Analyzing autophagy through changes in the phosphorylation of mTORC1 substrates? The mammalian target of rapamycin (mTOR) complex 1 (mTORC1) is a large protein complex involved in the regulation of many cellular processes including protein synthesis and autophagy (Guertin and Sabatini, 2007; He and Klionsky, 2009). In many cellular settings, inhibition of mTORC1 leads to autophagy via derepression of the autophagy protein ULK-1/Atg1 that plays a crucial role in the early stages of autophagosome formation (He and Klionsky, 2009). To note, mTORC1 activity is regulated by several upstream kinases including the prosurvival and anti-apoptotic protein Akt, AMP-activated protein kinase (AMPK) and extracellular signal regulated kinase (ERK) that have also been shown to participate in the regulation of autophagy in response to ER stress (Guertin and Sabatini, 2007; Verfaillie et al., 2010). Autophagy is frequently monitored by indirectly analyzing the phosphorylation status the mTORC1 substrates eukaryotic translation initiation factor 4E binding protein 1 (4EBP-1) and p70 S6 kinase (p70S6K) as well as the downstream target of this latter kinase (the ribosomal protein S6) for which excellent antibodies are available. Nevertheless, the mechanisms that determine the selectivity of mTORC1 for its substrates have not been completely clarified yet. For example, rapamycin (a well-established inhibitor of mTORC1 activity that is widely used as an autophagy inducer) strongly reduces S6 phosphorylation whereas does not affect 4EBP-1 phosphorylation (Guertin and Sabatini, 2007). Thus, a decrease in the phosphorylation of one of the substrates of mTORC1 does not necessarily correlate with changes in the phosphorylation of the others and in particular of ULK-1. Until reliable antiphospho-ULK-1 antibodies are available, we recommend using additional experimental approaches—as the genetic manipulation of the mTORC1/ULK-1 axis—to verify the participation of these signaling complexes in the triggering of autophagy.

2.4. Monitoring autophagy by knockout or knockdown of essential autophagic proteins As indicated in other chapters of this issue of Methods in Enzymology, different ER stress-related genes have been genetically inactivated in mice (Schroder and Kaufman, 2005). In addition, during the last few years mice deficient in different Atg have also been generated (Cecconi and Levine, 2008; He and Klionsky, 2009; some of them, being lethal at different stages of the embryonic development or early after birth). Mouse embryonic

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fibroblast (MEFs) can be obtained from most of these mice. These cells can be immortalized or transformed by using different oncogenes (see Section 3.4.2 for a description of the transformation protocol) and constitute an excellent tool to investigate the participation of a gene in the regulation of autophagy. For example, unlike their wild-type counterparts, embryonic fibroblasts derived from eIF2a Ser51Ala knock-in mice (in which this protein cannot be phosphorylated in response to ER stress) are resistant to autophagy as induced by different stimuli (He and Klionsky, 2009; Verfaillie et al., 2010). An alternative possibility is to transiently or stably reduce the expression of the gene of interest by using short interference RNA (siRNA). These approaches have been very useful to investigate the participation of ER stress-related genes in the regulation of autophagy (He and Klionsky, 2009; Verfaillie et al., 2010). 2.4.1. Knockdown of autophagic proteins Required material Cells of interest: Silencing with siRNA duplexes can be performed in many different cell types. We will describe the protocol used in the human glioma cell line U87MG. Cell culture medium: Please note that cells will be plated in medium without antibiotics prior to initiate the transfection with siRNA duplexes. Transfection reagent: Each different cell line may require a different reactive. For U87MG cells, we use DharmaFECT 1 (Dharmacon, Lafayette, CO) Interference RNA: Sequences for many different genes related with ER stress and autophagy can be found in different data bases and/or are commercially available. For monitoring transfection efficiency by immunofluorescence, cells can be cotransfected with a control fluorescent siRNA (siGLO RISC-Free siRNA, Dharmacon).

Procedure The day before transfection plate cells in fresh complete medium without antibiotics (cells should reach a 75–80% confluence on the day of transfection). Transfect cells using standard procedures for siRNA duplexes. Twenty-four hours after transfection, trypsinize cells and seed them in complete medium. Silencing efficacy should be evaluated by analyzing mRNA (e.g., by using RT-PCR or real time quantitative PCR) or protein (by using Western blot or immunofluorescence) levels of the targeted gene. Note: If the efficacy of transfection is not 100%, and the experiment requires analyzing the effect of knocking down the selected gene in each individual cell (e.g., in experiments of immunofluorescence), we recommend transfecting with a 1:4 mix of a control fluorescent siRNA (e.g.,


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siGLO RISC-Free siRNA, Dharmacon) and the siRNA designed to target the gene of interest. Those cells that have been effectively transfected will show a few characteristic bright spots on its cytoplasm (see Fig. 17.2B).

3. Monitoring Autophagy Upon ER Stress In Vivo The analysis of the biological activity of an anticancer agent frequently requires in vivo confirmation of the results obtained in vitro. ER stress and autophagy can be analyzed in tumor samples (see Section 3.1 for a protocol to generate tumor xenograft with the human glioma cell line U87MG and Section 3.2 for the protocol to analyze autophagy in tumor samples). Likewise, one of the strategies that can be used to confirm the participation of ER stress or autophagy in the mechanism of action of an anticancer agent is to generate tumors that are deficient (or have a decreased expression) of some the genes involved in the regulation of these cellular processes (see Sections 3.3 and 3.4).

3.1. Generation of tumor xenografts Required materials Mice: Male nude mice (Hsd:Athymic Nude-Foxn1; nu/nu) 4–5 weeks of age can be purchased from several companies. We normally order from Harlan, (Indianapolis, IN). Nude mice should be maintained under pathogen-free conditions (use autoclaved water and food and filtered cages). Let mice acclimatize for 5–7 days before injecting tumor cells. Cell culture medium: For U87MG cells we use DMEM supplemented with 10% FBS and antibiotics (penicillin-streptomycin) Trypsin- EDTA solution (Sigma) Other materials

PBS; PBS filtered containing 0.1% glucose

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1.5-ml (Eppendorf) tubes, 1 ml-syringes, needles (25 gauge) Growth of cancer cells for inoculation

Generation of tumor xenografts requires the inoculation of approximately 5–10 106 cells per mice. To facilitate engraftment, cells should be dividing actively (approximately 80% confluence) on the day of injection. As a significant fraction of cells can be lost during trypsinization and inoculation, we recommend growing cells in excess.

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Preparation and injection of cancer cells Add 2–3 ml of prewarmed trypsin-EDTA solution to each 150 cm2 disc with growing cells, incubate 3–5 min at 37 C and add 10% FBS-containing medium to the plates to neutralize trypsin digestion. Collect the detached cells by centrifugation and determine the total number of cells. Finally, after recentrifugation, carefully aspirate the medium and resuspend the cells in PBS with glucose (0.1%, w/v) at a density of 10 106 cells/100 ml. After shaking gently, inject subcutaneously 100 ml of the cell suspension in each mouse with a 1-ml syringe using a 25-gauge needle. We recommend collecting in one sterile tube enough cells as for injecting five to six mice (50–60 106 cells in 500 ml PBS-glucose). Note: In order to optimize the injection procedure, we suggest to take into account the following tips: (i) Keep the cells in ice and warm them (holding with the hand for 30 seconds) before injection (ii) once the mice is immobilized dispose it in a position in which it is inclined 135º with respect your view to facilitate the injection; (iii) introduce the needle subcutaneously (make sure that you can see the needle under the skin) and; (iii) subcutaneously introduce 1–1.5 cm of the needle within the mice before releasing the cell suspension to avoid losing cells during the inoculation. Measuring tumor growth Depending on the cell type and the number of cells inoculated there could be large differences in the time required for engraftment and growth of the tumors up to a size in which they can be visualized. In the case of U87MG cells, 7–10 days after cell inoculation, tumors can be observed in the flank of the mice nearby the zone of injection. Nevertheless, we recommend monitoring tumor establishment daily after inoculation by touching the zone of injection with the fingerprints (growing tumors feel like a small piece of hard tissue). Allow the tumors to growth until they reach an average volume of 200– 250 mm3 before starting the corresponding treatments. [The size of the tumors can be measured with an external caliper (the volume is calculated using the following formula: (4p/3) (width/2)2 (length/2)].

3.2. Analysis of autophagy in samples obtained from tumor xenografts Required material

Tumor sections Anti-LC3: For in vivo detection of LC3, we recommend using the anti-LC3 monoclonal antibody (Nanotools 0231-100/LC3-5F10).


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Secondary antibody: Alexa 488 goat anti-mouse IgG (Invitrogen) Nuclei staining: Hoechst 33342 (Invitrogen; stock solution 5 mg/ml) Blocking solution: 10% goat serum þ 0,5% Triton X-100 in PBS Other reagents: Mowiol (mounting medium) Disposables

Pap pen Procedure

Dissect tumor xenografts, embed the samples obtained in Tissue-Tek (Sakura, Torrance, California) and rapidly freeze them (store at 80 C). (We recommend using a flat bottom recipient to embed and freeze the dissected samples so the frozen tumor block can be more easily store and handled). Slice tumour sections in the cryostat (8 mM) placing the sliced sections in TESPA-coated microscope slides (three to four slices per slide), fix the slides in cold acetone for 10 min and wash twice with PBS. Individually remove slides and carefully absorb the liquid surrounding each section without drying the tissue. Draw a circle around the tissue with a Pap pen, add a drop of PBS, and place the slide in a humid chamber. After this last wash, incubate with blocking solution for 30 min at room temperature. Remove blocking solution, add the required volume of anti-LC3 antibody (nanotools 0231-100/LC3-5F10; 1/100 in PBS) to completely cover the sample and incubate overnight at 4 C in a humid chamber. At the next day, wash three times with PBS for 10 min, add the secondary antibody (Alexa 488 goat anti-mouse) and Hoechst 33342 1/ 1000 in blocking solution and incubate 1 h at room temperature. Wash three times with PBS for 10 min, mount with Mowiol and dry overnight at room temperature. Analysis in the microscope Identification of individual puncta within cells is easier with a confocal microscope and we therefore recommend using it for acquiring definitive images. To note, LC3 puncta in tumor tissue are frequently more difficult to detect than in cultured cells. In these cases, we recommend evaluating not only the formation of LC3 dots but also changes in LC3 intensity. See Fig. 17.3 for examples of LC3 immunostaining in tumor tissue.

3.3. In vivo silencing of autophagy or ER stress genes As described in Section 2.4, one strategy to analyze the participation of ER stress and autophagy in the regulation of different cellular processes is knocking down the expression of crucial ER stress or autophagy-related

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Figure 17.3 Analysis of LC3 immunostaining in samples obtained from tumor xenografts. (A) Identification of cells with LC3 dots. Effect of THC administration on LC3 immunostaining of glioma cell-derived tumor xenografts. Arrows point cells with LC3 dots (bar: 20 mm). (B) Evaluation of the intensity of LC3 immunostaining. Effect of THC treatment on LC3 immunostaining of glioma cell-derived tumor xenografts. Note that—in addition to the presence individual cells exhibiting LC3 dots—THC-treated tumors exhibit a higher intensity of LC3 immunofluorescence than vehicle-treated tumors.

genes in cultured cells. The same approach can be undertaken in vivo. To increase the efficiency of siRNA delivery in vivo (the main limitation of this procedure), we recommend using atelocollagen. The combination of atelocollagen and HPLC-purified siRNA allows an efficient silencing of gene expression in vivo. In the case of subcutaneous tumor xenografts, the mixture of atelocollagen and siRNA can be injected around the engrafted tumor facilitating thus the absorption and incorporation of the siRNA within the tumor cells. In experiments run in our lab, we regularly observe that the targeted gene remains silenced at least for 1 week after the injection of the atelocollagen–siRNA mixture. Nevertheless each gene and siRNA could behave differently and therefore, when setting the conditions for the silencing of new gene in vivo, we recommend analyzing the expression of the targeted gene by RT-PCR or quantitative-real time PCR (mRNA), Western blot (protein), and immunoflourescence (protein and distribution) 2 and 7 days after the administration of atelocollagen and siRNA to verify


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that the gene has been efficiently knocked down. Atelocollagen (AteloGeneTM) is produced and distributed by Koken (Tokyo, Japan) and a detailed protocol for its utilization can be found in the product data sheet.

3.4. Generation of tumor xenografts with transformed embryonic fibroblasts derived from genetically modified mice One possible strategy to investigate the participation of ER stress or autophagy-related genes in tumor progression and response to therapy is to generate tumor xenografts with transformed MEFs derived from mice deficient in the gene of interest. Here we describe the procedure to obtain and transform MEFs. 3.4.1. Isolation of embryonic fibroblasts Required material

Male and female mice (heterozygous mice for the gene of interest) Cell culture medium: DMEM with 10% FBS and antibiotics (penicillin and streptomycin) PBS Dissection material Tissue culture material Procedure Cross mice (normally heterozygous for the gene of interest) and sacrifice pregnant female on gestational day 13.5–14.5. Disinfect with alcohol, practice and incision in the abdominal region, extract the uterus (containing the embryos) and place it in a 15–50-ml tube containing PBS with antibiotics (use the same concentration as in the culture medium). Place the uterus (containing the embryos) in a sterile 10-cm diameter dish; remove the amniotic sac (the membrane around the embryos), add sterile PBS, and transfer each embryo to a new dry plate. Dissect the embryos by removing (and discarding) head, limbs, and liver (a red spot located in the abdominal zone of the embryo) and finely mince the rest of the body with scissors for 30 s. (DNA for genotyping can be isolated from the excised head of each embryo). Add 3 ml of trypsin, incubate at 37 C for 5 min, pipette up, and down until all clumps disaggregate and incubate for five additional minutes at 37 C. Transfer the cell suspension (if there are still large chunks of tissue at this stage, first transfer the suspension to a new tube and let the chunks settle for a few seconds) to a 15-cm diameter plate (or to a 75-cm2 flask)

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with 25 ml of culture medium (DMEM, 10% FBS, and antibiotics) and replace with fresh complete medium 8–16 h after plating. On average cells from a single E13.5 embryo reach confluence 2–3 days after plating. Cells should be trypsinized and frozen just before this happens. Note 1: All the procedure should be perform in sterile conditions Note 2: Prepare one primary culture of MEFs from each embryo 3.4.2. Transformation of primary MEFs In this protocol we use supernatants enriched in retroviral particles obtained from Phoenix Ecotropic cells to transduce MEFs. By using this procedure, we avoid the time-consuming titration procedure. Nevertheless, it is important to bear in mind that viral title is very sensitive to freezing– thawing. Thus, we recommend using fresh supernatants of Phoenix Ecotropic cells. In addition, perform the whole experiment throughout the same week to ensure an efficient infection of primary MEFs. Required material

Cells: Primary cultures of mouse embrionary fibroblasts (MEFs) [these cells enter in senescence after a few (three to four) passages and therefore should be transformed (or frozen) rapidly after isolation]. Packagin cells: PhoenixTM Ecotropic cells will be used to generate retroviral particles carrying the transforming oncogene. Cell culture medium: Please note that cells will be plated in medium without antibiotics prior to viral transduction. DNA of interest: DNA of high quality is recommended for transfections. For infection of MEFs, we use a plasmid encoding a retroviral vector carrying the viral protein E1A and the constitutively active form of Ras (RasV12) as well as a hygromycin resistance gene for selection. Transfection reagents: LipofectamineTM 2000 Polybrene Hygromycin Day 1. Preparing Phoenix Ecotropic cells for transfection

Plate the Phoenix Ecotropic cells (packaging cells) in 10 cm diameter dishes with fresh medium (DMEM with 10% FBS). Cells should reach 90% confluence on day of transfection. Day 2. Transfection of Phoenix cells Transfect Phoenix Ecotropic cells with the plasmid encoding the retroviral vector with the gene(s) of interest using Lipofectamine 2000TM (follow


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the standard protocol of transfection with this reagent). After transfection, incubate the cells at 37 C/5% CO2 for 4–6 h remove the medium and replace by fresh complete medium. Day 3. Preparing MEFs for transduction with retroviral particles Plate the cells in 10 cm diameter dishes with fresh complete medium (DMEM with 10% FBS). MEFs should reach 80–85% confluence on the day of infection. MEFs should be transduced when they are at a low pass (2 or 3 at maximum) to enhance transduction efficiency. Note: We recommend transforming in parallel wild-type and KO MEFs derived from littermate mice to minimize as much as possible differences between wild-type and KO cells other than the expression of the gene of interest. Day 4. Transduction of MEFs Forty-eight hours after transfection, collect the medium from Phoenix Ecotropic cells (8 ml per plate), centrifuge to eliminate cell debris (110 g for 5 min), filter (45 mm), and add polybrene (5mg/ml) to enhance the efficacy of retroviral infection. Note: Add fresh medium to the dishes with Phoenix Ecotropic cells once the first supernatant has been removed to obtain additional viral particles for the next day. Remove medium from MEFs, add retroviral-enriched supernatant (7 ml per 10 cm diameter plate) and incubate cells at 37 C/5% CO2. Day 5. Retransduction of MEFs To enhance the efficacy of infection, we recommend reinfecting MEFs with fresh retroviral supernatant obtained from Phoenix Ecotropic cells using the procedure described above. Day 6. Change the media of MEFs to DMEM with 10% FBS. Selection of transformed cells Start the selection with hygromycin 3 or 4 days after transducing the cells with the retroviral particles. Increase gradually the concentration of antibiotics during the first few days until reaching 1 mg/ml. Individual clones of hygromycin-resistant cells are evident after several weeks of

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incubation in the selection medium. Collect and amplify individual clones and verify that they have the expected genotype. We recommend mixing at least 20 different clones derived from the same embryo to obtain a homogeneous culture of transformed MEFs. Note: Unlike viral particles obtained from Phoenix Amphotropic cells, retrovirus produced by Phoenix Ecotropic cells cannot infect human cells. Nevertheless standard procedures for safe handling of materials that have been in contact with viral particles should be followed during the whole experiment.

3.4.3. Generation of tumor xenografts with transformed MEFs Transformed MEFs can be used to generate tumor xenografts in immunodeficient mice using the procedure described in Section 3.1. Tumor xenografts can be generated in parallel with cells deficient in the protein of interest and their wild-type counterparts to analyze the relevance of that gene (and the process in which is involved) in the action of the antitumoral agent of interest. See Fig. 17.4 for an example of the different response to treatment with THC of Atg5 KO and wild-type tumors (ATG5 is an essential gene for autophagosome formation (He and Klionsky, 2009) and therefore ATG5-deficient tumors do not undergo autophagy in response to this treatment.

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REFERENCES Cecconi, F., and Levine, B. (2008). The role of autophagy in mammalian development: Cell makeover rather than cell death. Dev. Cell 15, 344–357. Eisenberg-Lerner, A., et al. (2009). Life and death partners: Apoptosis, autophagy and the cross-talk between them. Cell Death Differ. 16, 966–975. Guertin, D. A., and Sabatini, D. M. (2007). Defining the role of mTOR in cancer. Cancer Cell 12, 9–22. He, C., and Klionsky, D. J. (2009). Regulation mechanisms and signaling pathways of autophagy. Annu. Rev. Genet. 43, 67–93. Klionsky, D. J. (2007). Autophagy: From phenomenology to molecular understanding in less than a decade. Nat. Rev. Mol. Cell Biol. 8, 931–937. Levine, B., and Kroemer, G. (2008). Autophagy in the pathogenesis of disease. Cell 132, 27–42. Maiuri, M. C., et al. (2007). Self-eating and self-killing: Crosstalk between autophagy and apoptosis. Nat. Rev. Mol. Cell Biol. 8, 741–752. Maiuri, M. C., et al. (2009). Control of autophagy by oncogenes and tumor suppressor genes. Cell Death Differ. 16, 87–93. Mathew, R., et al. (2007). Role of autophagy in cancer. Nat. Rev. Cancer 7, 961–967. Matus, S., et al. (2008). The stress rheostat: An interplay between the unfolded protein response (UPR) and autophagy in neurodegeneration. Curr. Mol. Med. 8, 157–172. Mizushima, N., et al. (2008). Autophagy fights disease through cellular self-digestion. Nature 451, 1069–1075. Morselli, E., et al. (2009). Anti- and pro-tumor functions of autophagy. Biochim. Biophys. Acta 1793, 1524–1532. Salazar, M., et al. (2009). TRB3 links ER stress to autophagy in cannabinoid anti-tumoral action. Autophagy 5, 1048–1049. Schroder, M., and Kaufman, R. J. (2005). The mammalian unfolded protein response. Annu. Rev. Biochem. 74, 739–789. Verfaillie, T., et al. (2010). Linking ER stress to autophagy: Potential implications for cancer therapy. Int. J. Cell Biol. 2010, 930509. Winslow, A. R., and Rubinsztein, D. C. (2008). Autophagy in neurodegeneration and development. Biochim. Biophys. Acta 1782, 723–729.


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The Unfolded Protein Response Induced by Salt Stress in Arabidopsis Miaoying Wang, Qiangyi Xu, and Ming Yuan Contents 319 320 320 321 322 322 323 327 327

1. Introduction 2. The Treatment of Arabidopsis Seedlings 2.1. Salt treatment 2.2. Additional treatments 3. The Analysis of Unfolded Protein Response 3.1. RNA isolation 3.2. The determination of unfolded protein response Acknowledgments References

Abstract Salt stress has a major impact on plant growth and crop production, pointing to the importance of understanding the mechanism of salt tolerance in plants. Disruption of the protein-folding capacity in the endoplasmic reticulum (ER) induces the accumulation of unfolded protein and ER stress, which activates an “unfolded protein response” (UPR). Although reports show that salt stress leads to UPR in various organisms, including plants, it remains to be determined how salt stress induces UPR. Zinc deficiency also induces UPR in a wide range of organisms. Here we provide a detailed description of the role of zinc in initiating UPR in the plant response to salt stress along with details of the methodology required for its investigation.

1. Introduction High salt content in soil causes a wide range of stresses affecting plant growth, which include ion toxicity, osmotic stress, and low-oxygen stress (Chinusamy and Zhu, 2003). Salt stress and drought induce various physiological State Key Laboratory of Plant Physiology and Biochemistry, College of Biological Sciences, China Agricultural University, Beijing, China Methods in Enzymology, Volume 489 ISSN 0076-6879, DOI: 10.1016/B978-0-12-385116-1.00018-2

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and gene regulation responses in plants (Affenzeller et al., 2009; Zhang et al., 2008; Zhu, 2002), including “the unfolded protein response” (UPR) in the endoplasmic reticulum (ER) (Liu et al., 2007b). bZIP60 and bZIP28 are two ER stress sensors/transducers (Liu et al., 2007a) that are involved in UPR which mitigate the accumulation of unfolded or misfolded proteins in plants (Liu et al., 2007a). Under salt-stress conditions, bZIP17 (Liu et al., 2007b), which localizes at the ER in normal conditions, is cleaved. The N-terminal part, containing a bZIP domain, moves into the nucleus and induces genes involved in salt-stress responses, including BiP2 (Martinez and Chrispeels, 2003). Zinc involvement in ER functioning is evolutionarily conserved and zinc deficiency induces UPR in a wide range of organisms (Ellis et al., 2004; Palmiter and Huang, 2004). Zinc deficiency upregulates the ER stress response in yeast and mammalian cells, indicating a conserved requirement for zinc in ER function among eukaryotes (Ellis et al., 2004; Harding et al., 2002). In a recent study, we reported that ZTP29, a putative zinc transporter located at the ER membrane, is involved in the response to salt stress through regulation of the zinc level in the ER and which is required to induce the UPR pathway (Wang et al., 2010). Salt stress induces upregulation of ZTP29 expression and ztp29 mutant plants exhibit various phenotypes of salt hypersensitivity, and the salt stress-induced upregulation of the UPR pathway genes BiP2 and bZIP60 is inhibited. ZTP29 functions in the transportation of zinc from the ER to the cytoplasm and this leads to a reduced ER zinc level and induction of UPR. Activation of UPR thus leads to the tolerance of plants to salt stress. In this chapter, we describe the procedures for examination of how zinc is involved in the UPR in Arabidopsis seedlings under salt stress.

2. The Treatment of Arabidopsis Seedlings Salt stress induces UPR in the ER in Arabidopsis (Liu et al., 2007b). Plants undergoing UPR mitigate the accumulation of unfolded or misfolded proteins through upregulation of the expression of genes that code for ER-folding proteins (Kaufman et al., 2002; Rutkowski and Kaufman, 2004). UPR results in an increase in the activity of chaperone proteins, which play crucial roles in facilitating protein folding during ER stress (Mori, 2000; Wilson, 2002).

2.1. Salt treatment 2.1.1. Required plant materials Seeds of Arabidopsis thaliana ecotype Columbia are germinated on agar plates containing half-strength Murashige and Skoog (MS) salts (Sigma), 1% sucrose, and 0.7% agar, pH 5.7. After stratification at 4 C for at least

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2 days, seedlings are grown in an illuminated growth chamber with 14 h light/10 h dark cycles at 21 C prior to treatment. The salk-144194 Arabidopsis line, which has a T-DNA insert in the eighth exon of ZTP29, can be obtained from the Arabidopsis Biological Resource Center (ABRC, OH, U.S.A.). Homozygous ztp29 mutants are selected before experimentation. 2.1.2. Treatment with salt In order to induce UPR under salt stress, seedlings of 7-day-old wild-type and ztp29 mutant are treated with 100 mM NaCl. The seedlings are transferred into 4 mL centrifuge tubes containing a solution of half-strength MS and 1% sucrose, plus 100 mM NaCl, each tube containing about 0.5 g seedlings. The centrifuge tubes are capped and placed in an illuminated growth chamber at 21 C for 6 h prior to use. UPR can be induced in a few hours after the treatment. In our laboratory, we treat the seedlings for 6 h before the examination of UPR. Although agar medium containing NaCl is widely used in the observations of Arabidopsis phenotypes under salt stress, liquid medium containing halfstrength MS and 1% sucrose, plus 100 mM NaCl (Sinopharm Chemical Reagent Co. Ltd, China) for the treatments has proved efficacious in our hands. Treatment with liquid medium may be more efficient because the salt-stress conditions might affect the cells more directly and rapidly. However, agar plates containing NaCl are better for the observation of seed germination and root growth under salt stress. In addition, half-strength MS should be employed for seedling growth since seedlings grown on halfstrength MS are more sensitive to salt treatment than those on full-strength MS. Overall consistency is important for the experiments, for example, use of plant materials and reagents from the same source and batch number allow for more reproducibility in experiments. Higher purity NaCl should be used in the experiments to avoid contaminant by minor ions.

2.2. Additional treatments 2.2.1. Treatments that interfere with cellular zinc To investigate the involvement of zinc in UPR under salt stress, the seedlings are first prepared as described above. Seven-day-old wild-type and ztp29 mutants are transferred into 4-mL centrifuge tubes containing a solution of (1) half-strength MS plus100 mM NaCl; (2) half-strength MS, 100 mM NaCl, and 400 mM ZnCl2; or (3) half-strength MS, 100 mM NaCl, and 60 mM N,N,N0 ,N0 -Tetrakis-(2-pyridylmethyl)ethylenediamine (TPEN). The tubes are capped and placed in an illuminated growth chamber at 21 C for 6 h prior to experimentation. TPEN is a cell membrane permeable reagent chelating heavy ions, and is used for reducing the zinc level in cells (Hyae et al., 2001). TPEN is dissolved with 100% ethanol to

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make a 100 mM stock solution which is then diluted with distilled water to make a working solution. 2.2.2. The treatment of UPR-inducing reagents ER stress occurs when proteins in the ER are unfolded or misfolded (Urade, 2007). Turicamycin (TM) and dithiothreitol (DTT) are two traditional UPR-inducing reagents which can cause an ER stress response (Iwata and Koizumi, 2005; Kamauchi et al., 2005; Martinez and Chrispeels, 2003). DTT is a dithiol-reducing agent that breaks disulfide bonds and also inhibits their formation. TM inhibits protein glycosylation and therefore protein folding. In Arabidopsis, DTT and TM induce the expression of genes that code for protein-folding enzymes and reduce the expression of genes that code for secretory proteins (Kamauchi et al., 2005; Martinez and Chrispeels, 2003). To examine UPR under salt stress, the seedlings are prepared as described above. Seven-day-old wild-type and ztp29 mutants are transferred into 4-mL centrifuge tubes containing a solution of (1) half-strength MS plus 70 mM NaCl, (2) half-strength MS plus 2 mM DTT, or (3) halfstrength MS plus 5-mg/mL TM (DTT and TM are obtained from SigmaAldrich (U.S.A.). The centrifuge tubes are capped and placed in an illuminated growth chamber at 21 C for 6 h prior to further experimentation.

3. The Analysis of Unfolded Protein Response After the treatments described above, UPR is examined by measurement of the expression levels of the UPR pathway genes BiP2 and bZIP60. BiP2 and bZIP60 have been previously selected for the detection of UPR in Arabidopsis (Iwata et al., 2008; Liu et al., 2007a; Zhang et al., 2008).

3.1. RNA isolation The protocol used for the isolation of RNA used by the authors is essentially the same as that described in the instructions with TRIZOL reagent (Roche, Germany), but with modifications. To prevent RNase contamination, all the experimental labware used in the RNA isolation procedure should be soaked in 0.1% diethypyrocarbonate (DEPC) in water overnight, sterilized at 121 C for 30 min, and dried at 80 C for 6 h before use. 0.1% DEPC is prepared by diluting 1 mL DEPC stock solution (purity >98.5%, purchased from Sangon Biotech Co., Ltd, Shanghai, China) into a 1-L working solution with sterile distilled water, which is kept in the dark for 12 h to allow the DEPC to be fully dissolved. Electrophoresis tanks should be treated with detergent, washed with distilled water, followed by soaking

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in 3% H2O2 overnight in the dark. Before use, the treated electrophoresis tanks should be also washed with 0.1% DEPC. 100–200 mg Arabidopsis seedlings, prepared as described above, are ground to a fine powder in liquid nitrogen. The powder is placed in a 1.5-mL centrifuge tube to which is added 1-mL TRIZOL reagent with immediate mixing by vortexing for 30 s. The homogenized samples are then incubated to permit the complete dissociation of nucleoprotein complexes by placing the tubes on ice for 5 min. After the incubation, 0.2 mL chloroform is added to the centrifuge tube. The centrifuge tube is then capped securely, shaken vigorously by hand for 15 s, and is incubated at 15 C for a further 2 min. The sample is then centrifuged at 12,000 g for 15 min at 4 C. After centrifugation, the sample separates into a lower red, phenol–chloroform phase, an interface and a colorless upper aqueous phase. RNA should remain exclusively in the aqueous phase. The RNA is precipitated from the aqueous phase by mixing with 0.5 mL precooled isopropyl alcohol and incubation at 20 C for 30 min. The solution is then centrifuged at 12,000 g for 15 min at 4 C, and the RNA pellet obtained is washed once with 75% ethanol. The centrifuge tube with the RNA pellet is then placed in a fume-hood to allow the water to evaporate from the RNA pellet. The RNA sample is then redissolved in 40 mL 0.1% DEPC and stored at 20 C for later use. RNA samples should have an A260/280 ratio of 25%).

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50 mm

19.67 min

50 mm

14.83 min

50 mm

12.00 min

50 mm

Tunicamycin 10 mg/ml

0.00 min

50 mm

50 mm

2.00 min

Tunicamycin 1 mg/ml

0.00 min

12.33 min

11.00 min

Thapsigargin 10 nM

0.00 min

7.33 min

Thapsigargin 1 nM

0.00 min

16.00 min Control

50 mm

50 mm

50 mm

50 mm

50 mm

Ines Witte and Sven Horke, Figure 8.1 Depending on the intensity of the stimulus, ER stress induces acute Ca2þ oscillations, a transient or a sustained cytosolic Ca2þ influx. EA.hy 926 cells were seeded in microscopy-suited dishes, loaded with Ca2þ indicator Fluo-4-AM (green) and nuclear marker Draq5 (blue), and treated with the indicated concentrations of tunicamycin or thapsigargin. In parallel, Fluo-4-AM and Draq5 fluorescence were recorded in time-lapse experiments by confocal laser scanning microscopy. Fluorescence intensity reports Ca2þ influx. Scale bar ¼ 50 mm; time after stress induction given in minutes (upper left corner).

A

0

5

10 h

CHOP

ACTIN

% Annexin positive MFs

B

* 15 10 5 0 AcLDL

AcLDL + 58035

AcLDL + 58035

AcLDL

20

Edward B. Thorp, Figure 16.1 CHOP expression and annexin staining of FC-loaded primary macrophages. Peritoneal macrophages were elicited and incubated in macrophage medium. At 90% confluence, cells were either loaded with AcLDL or alternatively FC-loaded (AcLDL þ 58-035) and (A) immunoblotted for CHOP at indicated times. Blots were stripped and reprobed for actin as a protein loading control. (B) Monolayers were in parallel stained with annexin V (green) and propidium iodide (red) at 16 h to detect mid-stage and late-stage apoptosis. The graph is a quantitation of annexin V-positive, propidium iodide-negative (apoptotic) macrophages in AcLDL cholesteryl ester (CE)-loaded, or FC-loaded macrophages. *p < 0.05. A

DNA

TUNEL

Mf

% TUNEL + MFs

B 3

* 2

1

0 4

12

Edward B. Thorp, Figure 16.3 Staining TUNEL-positive macrophages from PFA-fixed paraffin-embedded sections. (A) Representative micrographs show TUNEL-positive signal (red) in nuclei (blue) of aortic root lesions from lesions. Blue is Hoechst nuclear dye. Green is F4/80. (B) Enumeration of TUNEL-positive F4/80þ macrophages from early (4 week Western Diet fed) and advanced (12-week Western Diet fed) lesions. *p < 0.05.

A

B

C

veh

THC

veh

THC

THC E64d + PA

− −

+ −

+ +

− +

LC3-I LC3-II a-tubulin

Marıá Salazar et al., Figure 17.1 Techniques used to analyze autophagy in cultured cells. (A) Electron microscopy. Representative electron microscopy photomicrographs of U87MG cells after 6 h treatment with 4 mM D9-tetrahyrocannabinol (THC, a potent autophagy inducer; bars: 500 nm). Note the presence of early (open arrows) and late (close arrows) autophagosomes in THC but not vehicle (veh)-treated cells. Low panels: Details of early (left) and late (middle and right) autophagosomes. (B) LC3 immunostaining. Effect of THC (18 h) treatment on LC3 immunostaining (green) of U87MG cells (bar: 20 mm). Note the presence of LC3 puncta in cells treated with the autophagic stimulus but not in vehicle-treated cells. (C) LC3 lipidation. Effect of E64d (10 mM) and pepstatin A (PA; 10 mg/ml) on THC-induced LC3 lipidation of U87MG cells. Note the presence of the LC3-II band (higher electrophoretic mobility). In agreement with the stimulation of dynamic autophagy in this experimental setting, incubation with the inhibitors of the lysosomal proteases enhances LC3-II levels.

A 0h

PDI

2h

LC3 PDI

B

4h

LC3 PDI

LC3 PDI

siC veh

18 h

8h

LC3 PDI

LC3

sip8 THC

veh

THC

Marıá Salazar et al., Figure 17.2 Investigating the connection between ER stress and autophagy. (A) Time-course analysis of PDI and LC3 immunostaining. Effect of THC on PDI (red) and LC3 (green) immunostaining of U87MG cells at different time-points (bar: 20 mm). Note that the increase in PDI dots (a characteristic of ER stressed cells with enlarged ER) occurs at earlier time-points than the increase in LC3 puncta, suggesting that ER stress precede autophagy in the mechanisms of action of this anticancer agent. (B) Silencing of ER stress genes. Effect of THC (18 h) on LC3 immunostaining (green) of U87MG cells transfected with control (siC) or p8-selective (sip8) siRNA. [p8 is an ER stress-related transcription factor that mediates the proautophagic actions of THC]. To identify cells that had been transfected, a red-fluorescent stable control siRNA was included in the transfection mix together with siC or sip8 in 1:4 proportion (red dots within the cells correspond to red-fluorescent control siRNA; bar: 20 mm). Note that THC induces autophagy in siC- but not in sip8-transfected cells.

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