Apoptosis and Cancer: Methods and Protocols [1 ed.] 1493916602, 9781493916603

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Methods in Molecular Biology 1219

Gil Mor Ayesha B. Alvero Editors

Apoptosis and Cancer Methods and Protocols Second Edition

METHODS

IN

M O L E C U L A R B I O LO G Y

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

For further volumes: http://www.springer.com/series/7651

Apoptosis and Cancer Methods and Protocols Second Edition

Edited by

Gil Mor and Ayesha B. Alvero Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University, New Haven, CT, USA

Editors Gil Mor Department of Obstetrics, Gynecology and Reproductive Sciences Yale University New Haven, CT, USA

Ayesha B. Alvero Department of Obstetrics, Gynecology and Reproductive Sciences Yale University New Haven, CT, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-4939-1660-3 ISBN 978-1-4939-1661-0 (eBook) DOI 10.1007/978-1-4939-1661-0 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2014949776 © Springer Science+Business Media New York 2015 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is a brand of Springer Springer is part of Springer Science+Business Media (www.springer.com)

Preface Since the release of the first edition of this book in 2008, Pubmed has banked more than 100,000 scientific publications on apoptosis and more than 50,000 articles on apoptosis and cancer—and these numbers are continuing to increase. In the past few decades, the focus has been to delineate the cascade of steps eventually leading to cell death with the objective of identifying the key regulators. As these pathways are catalogued and specific key proteins are identified, most recent studies have been involved in the identification of either inhibitors of apoptosis (i.e., for prevention of cardiac cell death or neuronal cell death following hypoxic stress) or inducers of apoptosis (i.e., for cancer therapy). Whichever clinical application is intended, a mainstay of the studies has been the demonstration of apoptosis induction. This can be achieved, however, only if sensitive and specific assays are available. Thus, the aim of this book has not changed. It is still a detailed reference for the performance of molecular and cellular biology techniques for studying and/or detecting the activation of the apoptotic pathway, especially in the study of cancer. Resistance to apoptosis has been defined as one of the hallmarks of cancer and has been demonstrated to play a major role in chemoresistance. More recently, with the demonstration of hierarchy within the cancer cell subtypes that make up the tumor, focus has turned to understanding the mechanism/s of apoptosis evasion in the tumor-initiating cells or cancer stem cells. The demonstration that this cell population represents the more chemoresistant subtype is a first step in understanding the different mechanisms that are in place to evade apoptosis within the heterogeneous tumor. This book remains a combination of chapters authored by scientists from both industry and academia. In addition, as the field of apoptosis has grown in the past years, new assays have been developed to detect its activation not only in vitro but also in vivo. Assays have also been developed, which are optimized for multiplex analysis and medium- to highthroughput screens. Finally, since cell death can be a culmination of various pathways, this book also includes chapters that focus on cellular processes that can contribute to programmed cell death, such as autophagy and proteasome activation. We wish to thank all the authors for their contributions and we are confident that each chapter will be an excellent reference for any laboratory. We also wish to thank Ms. JoAnn Bilyard for her help in sending out invitations to prospective authors and in the organization of this book. Finally, special thanks to our Series Editor, Dr. John Walker. We hope that each chapter provides investigators with a stand-alone resource for the execution and analysis of the described protocols and that the book will provide an excellent reference for the study and detection of apoptosis within and even outside the area of cancer research. New Haven, CT, USA

Gil Mor, M.D., Ph.D. Ayesha B. Alvero, M.D.

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Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Caspase-3 Activation Is a Critical Determinant of Genotoxic Stress-Induced Apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gaurav S. Choudhary, Sayer Al-harbi, and Alexandru Almasan 2 Flow Cytometry Enumeration of Apoptotic Cancer Cells by Apoptotic Rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . David Diaz, Alfredo Prieto, Eduardo Reyes, Hugo Barcenilla, Jorge Monserrat, and Melchor Alvarez-Mon 3 “Multiplexed Viability, Cytotoxicity, and Caspase Activity Assays” . . . . . . . . . . Andrew L. Niles and Terry L. Riss 4 A Multiplexed Method for Kinetic Measurements of Apoptosis and Proliferation Using Live-Content Imaging . . . . . . . . . . . . . . . . . . . . . . . . Katherine Artymovich and Daniel M. Appledorn 5 Detection of End-Stage Apoptosis by ApopTag® TUNEL Technique . . . . . . . Chandra Mohan, Kevin Long, Manpreet Mutneja, and Jun Ma 6 Detection and Quantification of Apoptosis in Primary Cells Using Taqman® Protein Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christina Pfister, Heike Pfrommer, Marcos S. Tatagiba, and Florian Roser 7 Detection of p53 Protein Aggregation in Cancer Cell Lines and Tumor Samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yang Yang-Hartwich, Jamie Bingham, Francesca Garofalo, Ayesha B. Alvero, and Gil Mor 8 Detection of p53 Protein Transcriptional Activity by Chromatin Immunoprecipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yang Yang-Hartwich, Emily Romanoff, Jamie Bingham, Ayesha B. Alvero, and Gil Mor 9 Homogeneous, Bioluminescent Proteasome Assays . . . . . . . . . . . . . . . . . . . . . Martha A. O’Brien, Richard A. Moravec, Terry L. Riss, and Robert F. Bulleit 10 Laser Capture Microdissection for Gene Expression Analysis . . . . . . . . . . . . . . Mallikarjun Bidarimath, Andrew K. Edwards, and Chandrakant Tayade 11 Using the Peggy Simple Western System for Fine Needle Aspirate Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Erik T. Gentalen and John M. Proctor

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12 Analysis of Autophagosome Formation Using Lentiviral Biosensors for Live Fluorescent Cellular Imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kevin Long, Chandra Mohan, Janet Anderl, Karyn Huryn-Selvar, Haizhen Liu, Kevin Su, Mark Santos, Matthew Hsu, Lucas Armstrong, and Jun Ma 13 Optical Imaging of Ovarian Cancer Using HER-2 Affibody Conjugated Nanoparticles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Minati Satpathy, Rafal Zielinski, Ilya Lyakhov, and Lily Yang 14 Measuring Cardiac Autophagic Flux In Vitro and In Vivo . . . . . . . . . . . . . . . . Michael A. Gurney, Chengqun Huang, Jennifer M. Ramil, Nandini Ravindran, Allen M. Andres, Jon Sin, Phyllis-Jean Linton, and Roberta A. Gottlieb 15 PET Imaging for Tyrosine Kinase Inhibitor (TKI) Biodistribution in Mice . . . Hiroshi Fushiki, Yoshihiro Murakami, Sosuke Miyoshi, and Shintaro Nishimura Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors SAYER AL-HARBI • Department of Cancer Biology, Lerner Research Institute, Cleveland, OH, USA; Department of Human Cancer Genomic Research, King Faisal Specialist Hospital and Research Cancer, Saudi Arabia ALEXANDRU ALMASAN • Department of Cancer Biology, Lerner Research Institute, Cleveland, OH, USA; Department of Radiation Oncology, The Cleveland Clinic, Cleveland, OH, USA MELCHOR ALVAREZ-MON • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Alcalá de Henares, Madrid, Spain; Immune System Diseases and Oncology Service, University Hospital “Príncipe de Asturias”, Madrid, Spain AYESHA B. ALVERO • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA JANET ANDERL • EMD Millipore, Temecula, CA, USA ALLEN M. ANDRES • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA; Cedars Sinai Heart Institute, Los Angeles, CA, USA DANIEL M. APPLEDORN • Essen Bioscience, Ann Arbor, MI, USA LUCAS ARMSTRONG • EMD Millipore, Temecula, CA, USA KATHERINE ARTYMOVICH • Essen Bioscience, Ann Arbor, MI, USA HUGO BARCENILLA • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain MALLIKARJUN BIDARIMATH • Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, ON, Canada JAMIE BINGHAM • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA ROBERT F. BULLEIT • Promega Corporation, Madison, WI, USA GAURAV S. CHOUDHARY • Department of Cancer Biology, Lerner Research Institute, Cleveland, OH, USA; Department of Pathology, Case Western Reserve University, Cleveland, OH, USA DAVID DIAZ • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain ANDREW K. EDWARDS • Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, ON, Canada HIROSHI FUSHIKI • Bioimaging Research Laboratories, Astellas Pharma Inc., Tsukuba, Ibaraki, Japan FRANCESCA GAROFALO • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA ERIK T. GENTALEN • Protein Simple, Santa Clara, CA, USA ROBERTA A. GOTTLIEB • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA; Cedars Sinai Heart Institute, Los Angeles, CA, USA MICHAEL A. GURNEY • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA MATTHEW HSU • EMD Millipore, Temecula, CA, USA

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CHENGQUN HUANG • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA KARYN HURYN-SELVAR • EMD Millipore, Temecula, CA, USA PHYLLIS-JEAN LINTON • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA HAIZHEN LIU • EMD Millipore, Temecula, CA, USA KEVIN LONG • EMD Millipore, Temecula, CA, USA ILYA LYAKHOV • VARNISS, L.L.C., Frederick, MD, USA JUN MA • EMD Millipore, Temecula, CA, USA SOSUKE MIYOSHI • Bioimaging Research Laboratories, Astellas Pharma Inc., Tsukuba, Ibaraki, Japan CHANDRA MOHAN • EMD Millipore, Temecula, CA, USA JORGE MONSERRAT • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain GIL MOR • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA RICHARD A. MORAVEC • Promega Corporation, Madison, WI, USA YOSHIHIRO MURAKAMI • Bioimaging Research Laboratories, Astellas Pharma Inc., Tsukuba, Ibaraki, Japan MANPREET MUTNEJA • EMD Millipore, Temecula, CA, USA ANDREW L. NILES • Promega Corporation, Madison, WI, USA SHINTARO NISHIMURA • Bioimaging Research Laboratories, Astellas Pharma Inc., Tsukuba, Ibaraki, Japan MARTHA A. O’BRIEN • Promega Corporation, Madison, WI, USA CHRISTINA PFISTER • Department of Neurosurgery, University of Tuebingen, Tuebingen, Germany HEIKE PFROMMER • Department of Neurosurgery, University of Tuebingen, Tuebingen, Germany ALFREDO PRIETO • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain JOHN M. PROCTOR • Protein Simple, Santa Clara, CA, USA JENNIFER M. RAMIL • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA; Cedars Sinai Heart Institute, Los Angeles, CA, USA NANDINI RAVINDRAN • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA EDUARDO REYES • CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain; Research Unit, Industrial Farmacéutica Cantabria, Madrid, Spain TERRY L. RISS • Promega Corporation, Madison, WI, USA EMILY ROMANOFF • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA FLORIAN ROSER • Department of Neurosurgery, University of Tuebingen, Tuebingen, Germany MARK SANTOS • EMD Millipore, Temecula, CA, USA MINATI SATPATHY • Department of Surgery, Emory University School of Medicine, Atlanta, GA, USA; Cedars Sinai Heart Institute, Los Angeles, CA, USA JON SIN • Donald P. Shiley BioScience Center, San Diego State University, San Diego, CA, USA KEVIN SU • EMD Millipore, Temecula, CA, USA

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MARCOS S. TATAGIBA • Department of Neurosurgery, University of Tuebingen, Tuebingen, Germany CHANDRAKANT TAYADE • Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, ON, Canada LILY YANG • Department of Surgery, Emory University School of Medicine, Atlanta, GA, USA YANG YANG-HARTWICH • Department of Obstetrics, Gynecology, and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, USA RAFAL ZIELINSKI • MD. Anderson Cancer Center, Houston, TX, USA

Chapter 1 Caspase-3 Activation Is a Critical Determinant of Genotoxic Stress-Induced Apoptosis Gaurav S. Choudhary, Sayer Al-harbi, and Alexandru Almasan Abstract Apoptosis can be measured by number of methods by taking advantage of the morphological, biochemical, and molecular changes undergoing in a cell during this process. The best recognized biochemical hallmark of both early and late stages of apoptosis is the activation of cysteine proteases (caspases). Detection of active caspase-3 in cells and tissues is an important method for apoptosis induced by a wide variety of apoptotic signals. Most common assays for examining caspase-3 activation include immunostaining, immunoblotting for active caspase-3, colorimetric assays using fluorochrome substrates, as well as employing the fluorescein-labeled CaspaTag pan-caspase in situ detection kit. Key words Caspase-3, Apoptosis, Irradiation, PARP-1, Flow cytometry, Immunohistochemistry

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Introduction Apoptosis, or programmed cell death, is a cellular mechanism used to eliminate cells that are injured, infected, or have reached the end of their life-span [1]. This process is tightly regulated by a family of proteases called the caspases that are normally found in healthy cells as inactive precursors but become activated during apoptosis [2]. Apoptotic cell death is characterized by a series of morphological and biochemical features, such as plasma membrane blabbing, chromatin condensation, DNA cleavage, and exposure of phosphatidylserine on the extracellular side of the plasma membrane [3]. A critical process in execution of apoptosis is activation of a cascade of ICE/CED-3 family of cysteine proteases, termed caspases [4, 5]. Caspases are ubiquitously expressed intracellular cysteine proteases that mediate cell death and inflammation. Caspases have been originally identified in C. elegans. Later, mammalian homologs of these caspases have been discovered. Mammalian caspases, 14 members discovered to date, play distinct roles in apoptosis and inflammation. Specifically, caspase-3 is a major mediator

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_1, © Springer Science+Business Media New York 2015

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of both apoptotic and necrotic cell death. Caspases are synthesized as inactive precursors or zymogens, which are activated by proteolytic cleavage to generate active enzymes, which then may further proteolytically cleave other caspases or their cellular target proteins [6, 7]. An active caspase consists of two large (p17) and two small (p12) subunits that form two heterodimers that associate in a tetramer [8–10]. They recognize a 4–5 amino acid sequence on the substrate, which has an aspartic acid residue at P4 position as a critical requirement. This residue, which is the target for specific cleavage, occurs at the carbonyl end of the aspartic acid residue [11]. Active caspase-9 cleaves procaspase-3, which then is required for many of the characteristic apoptotic nuclear changes. Downstream effector caspases, such as caspase-3, cleave and inactivate proteins crucial for the maintenance of cellular cytoskeleton, DNA repair, signal transduction, and cell cycle control [6]. There are over 300 in vivo caspase substrates; amongst them are poly (ADP-ribose) polymerase (PARP-1) and ICAD/DFF45, the cleavage of which results in liberation of a caspase-activated deoxyribonuclease (CAD) that is responsible for the oligonucleosome-size DNA fragmentation that is characteristic of most apoptotic cells [12]. The activation cascade of ICE/CED-3 family of caspases is a common and critical step in the execution phase of apoptosis, triggered by different factors, including genotoxic agents (e.g., γ-irradiation or treatment with anticancer agents [4, 13–19]). Pharmacological inhibitors of caspase-3 can prevent the cell death following irradiation significantly indicating that caspase-3 activation is critical for genotoxic stress-induced apoptosis. Moreover, activation of caspase-3 can be an effective marker for the positive outcome of the different radio- and chemotherapeutic treatments of various cancers. Other assays for genotoxic stress-induced apoptosis have been presented elsewhere [20]. Importantly, various modes of cell death can be induced by DNA damage that can trigger caspase-3 activation [21]. Caspase-3 can be detected via immunofluorescence and immunoblotting using anti-active caspase-3 antibodies, by colorimetric assays employing fluorochrome substrates, and by flow cytometric methods, such as that using fluorochrome inhibitors of caspases (FLICA). Activated caspases cleave many cellular proteins and the resulting “signature” proteolytic fragments may also serve as useful markers. This chapter provides standard protocols that we have successfully used in our laboratory for a number of experimental systems, including cells grown in culture and as xenografts.

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Materials

2.1 Immunocytochemistry

1. Glass cover slips (22 × 22 mm) and slides. 2. Formaldehyde: 4 % in 1× PBS (dilute 37 % formaldehyde stock in 1× PBS), make fresh each time.

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3. Blocking buffer: 2 % goat serum, 0.3 % Triton X-100 in 1× PBS, sterile filtered. 4. Anti-active caspase-3 primary antibody. 5. Appropriate fluorochrome-conjugated secondary antibody. 6. Mounting medium for fluorescence (i.e., Vectashield, Vector Laboratories, Inc., Burlingame, CA), with or without 4′,6′-diamidino-2-phenylindole hydrochloride (DAPI). 7. Nail polish. 2.2 Caspase-3 Activity Determination: Colorimetric Assay

1. Lysis buffer: 1 % NP 40, 20 mM Hepes (pH 7.5), and 4 mM EDTA. Just before use, add the following protease inhibitors: aprotinin (10 μg/ml), leupeptin (10 μg/ml), pepstatin (10 μg/ ml), and 1 mM phenyl methyl sulfonyl fluoride (PMSF). 2. Reaction buffer: 100 mM Hepes (pH 7.5), 20 % v/v glycerol, 5 mM dithiothreitol (DTT), and 0.5 mM EDTA. 3. Caspase-3 substrate (Ac-DEVD-pN), 20 mM stock in DMSO (stable for >1 year at −20 °C), at 100 μM final concentration. Additional colorimetric as well as fluorometric substrates are available. The fluorometric substrates 7-amino-4-methylcoumarin (AMC) and 7-amino-4-trifluoromethylcoumarin (AFC) are more sensitive but require a fluorometer capable of detecting the 380/460 and 405/500 nm excitation/emission spectra, respectively. AFC can be also detected colorimetrically at 380 nm. 4. Microtiter plate reader, spectrophotometer, or fluorometer.

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Immunoblotting

1. 1× PBS. 2. Lysis buffer: 20 mM HEPES, pH 7.5, 1 mM EDTA, 150 mM NaCl, 1 % NP-40, and 1 mM DTT with protease inhibitors 1 mM PMSF, 1 μg/ml leupeptin. 3. Kit/reagents to determine protein concentration. 4. BSA (bovine serum albumin) to use as standard. 5. SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis). 6. Nitrocellulose or PVDF membrane. 7. 1× PBST: 1× PBS with 1 % Tween 20. 8. Milk (nonfat dry milk). 9. Active caspase-3 and PARP-1 primary antibodies. 10. Appropriate peroxidase-conjugated secondary antibodies. 11. Chemiluminescent reagents. 12. Disuccinimidyl suberate (DSS), final concentration 2 mM. 13. Conjugation buffer: 20 mM Sodium phosphate (pH 7.5) containing 0.15 M NaCl, 20 mM Hepes (pH 7.0), and 100 mM carbonate/bicarbonate (pH 9.0). 14. Quenching buffer: 1 M Tris–HCl (pH 7.5).

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2.4 FluoresceinLabeled CaspaTag Pan-Caspase In Situ Assay Kit

1. Cultured cells with media. 2. 15 ml polystyrene centrifuge tubes. 3. Microscope slides. 4. Hemocytometer. 5. Centrifuge. 6. Vortexer. 7. PBS, pH 7. 8. Dimethyl sulfoxide (DMSO). 9. Kit (Millipore) components: (a) FLICA Reagent (FAM-VAD-FMK): For lyophilized vials, reconstitute one vial of lyophilized reagent with 50 μl of DMSO and mix by swirling until completely dissolved (150× stock solution). Working solution: 30×, dilute 1:5 in PBS, pH 7.4. (b) 10× Wash buffer: 60 ml. Working solution: Dilute 1:10 in deionized water. (c) Fixative: 6 ml (d) Propidium iodide (PI): 1 ml at 250 μg/ml. (e) Hoechst 33342 Stain: 1 ml at 200 μg/ml.

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Methods To induce apoptosis, cells (2 × 105/ml) are irradiated at 4–20 Gy (137Cs source; fixed dose rate of 2.8 Gy/min [13, 22, 23]), or treated with any of the DNA-damaging chemotherapeutic agents, such as the topoisomerase inhibitor etoposide (VP16; 10 μM [5, 15]).

3.1 Immunocytochemistry to Detect Active Caspase-3

Caspase-3, the major effector caspase, is one of the key executioners of apoptosis. In response to an apoptotic signal, cleavage of inactive caspase-3 occurs mainly at the Asp175 residue, and thereby, being activated. A specific antibody against active caspase-3 can be used to detect the apoptotic cells by immunocytochemistry. 1. For plating cells for this experiment, we use glass cover slips (sterilized by dipping in ethanol and passing through flame) that are placed into 6-well plates. Cells (1 × 105 cells/well) are seeded and grown overnight. 2. Remove media and rinse cells with 1× PBS warmed to 37 °C. 3. To fix the cells, add 1–2 ml of 4 % formaldehyde to each well. Incubate cells to fix for 20 min at room temperature. 4. Wash each well with 1× PBS, three times for 5 min.

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5. Incubate cells in blocking buffer for 5–10 min at room temperature. 6. Dilute the primary antibody (two primary antibodies could be added at the same time, but they need to be of different origin, e.g., one rabbit and the other one mouse) in 100–200 μl blocking buffer, according to the recommended dilution. 7. Use a different 6-well dish to incubate the cells with the antibodies. Soak filter paper (3-cm diameter circles) in 1× PBS and place them into the wells; this is necessary to maintain the humidity in the chamber. Place the cover slips on top of the soaked filter paper. Add the primary antibody carefully to cover the entire cover slip. Incubate at room temperature for 1–2 h. 8. Wash with 1× PBS for 5 min, three times, each well. 9. Add the secondary antibody (fluorochrome conjugated) in blocking buffer and incubate for 30–45 min at room temperature in the dark. For dual staining the fluorophores need to have different emissions spectra for each individual antibody (e.g., FITC at 525 and phycoerythrin at 578 nm). There are a set of very sensitive and stable Alexa dyes (Molecular Probes, now part of Invitrogen); consult The Handbook—A Guide to Fluorescent Probes and Labeling Technologies for a comprehensive resource for fluorescence technology and its applications (http://probes.invitrogen.com/handbook/). 10. Wash with 1× PBS for 5 min, three times, each well. 11. Pick up cover slips with a forceps and drain away excess 1× PBS. 12. For mounting, add a drop of Vectashield to a clean microscope slide and gently lay the cover slip on top. 13. Remove excess Vectashield by blotting with tissue and seal with nail polish. 14. After adding the secondary antibody, it is important to keep slides in the dark at all times. A similar protocol can be used for tissue sections (see Note 1). 15. Store slides in a −20 °C freezer. 3.2 Caspase-3 Activity Determination: Colorimetric Assay

A simple colorimetric assay can measure the release of the chromogenic group from the synthetic substrate, most commonly p-nitroanilide (pNA) by activated caspases. Ac-DEVD-pNA is most frequently used, with the cleaved pNA being monitored colorimetrically through its absorbance at 405–410 nm. Although DEVD-based substrates are called caspase-3 specific they are in fact cleaved by most caspases, with caspase-3 being the most efficient. In vitro titration experiments and/or use of specific inhibitors may be required to distinguish the activity of various caspases. Other DNA substrates are available for several other caspases.

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1. Wash cells (1 × 106) with cold 1× PBS and resuspend them in 50 μl of cold lysis buffer, vortex, and keep on ice for 30 min. 2. Centrifuge the cell lysates at 12,000 × g for 10 min at 4 °C, collect the supernatant in fresh tubes, and assay the protein concentration for each sample. Keep on ice. 3. To a 96-well plate add reaction buffer, caspase substrate (100 μM final concentration), and 20–50 μl cell lysates for a final 200 μl reaction volume. 4. Incubate samples at 37 °C for 1–2 h and monitor the enzymecatalyzed release of p-NA at 405 nm using a microtiter plate reader. 3.3 Immunoblot Detection of Active Caspase-3

In most cases, the 32 kDA procaspase (inactive caspase)-3 protein is converted to active caspase-3 (p17, p12) that can be detected by Western blot analysis. One of its many cellular substrates is PARP-1. In the apoptotic cells, PARP-1 (110 kDa) is cleaved to form two truncated fragments, with the 86-kDa fragment being most frequently detected by the available commercial antibodies. 1. Collect the treated and untreated cells (1 × 106) by centrifugation (500 × g for 5 min). Decant the medium, resuspend the cell pellet in cold 1× PBS very gently, and spin it down (500 × g for 5 min). Decant the supernatant and repeat the process one more time. Remove 1× PBS carefully without disturbing the cell pellet. 2. Lyse the cells in a lysis buffer, with the cells incubated for 30 min on ice with occasional vortexing. 3. Centrifuge the cells for 15 min at 15,000 × g and collect the supernatants. 4. Determine protein concentration in these samples using a spectrophotometric method and appropriate protein assay reagents. 5. Load 50–100 μg protein, as well as the protein standard marker, on an 8 % SDS-PAGE gel to separate the proteins under denaturing conditions. 6. Transfer the proteins to a nitrocellulose membrane either by the wet or semidry transfer method. 7. Block the membrane with 5 % milk for 1 h at room temperature or overnight at 4 °C. 8. Incubate the membrane with primary antibodies (PARP-1, active caspase-3) for 2 h at room temperature or overnight at 4 °C (following the company’s recommended dilution) (see Note 2). 9. Wash the blot three times with 1× PBST at room temperature at 10-min intervals.

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10. Add the appropriate secondary antibody (anti-mouse or anti-rabbit depending on the primary antibody) with a 1:2,000 dilution to the blot and incubate for 1–1.5 h at room temperature. 11. Wash the blot five times with 1× PBST at room temperature at 10-min intervals. 12. Wash the blot with double-distilled water for very short time to get rid of Tween 20 and develop it using chemiluminescent reagents. 3.4 FluoresceinLabeled CaspaTag Pan-Caspase In Situ Assay Kit

This is a useful approach to detect the active caspase in individual cells. The methodology is based on the fluorochrome inhibitors of caspases (FLICA). The inhibitors are cell permeable and noncytotoxic. This kit contains a carboxyfluorescein-labeled fluoromethyl ketone peptide inhibitor of caspase (FAM-VAD-FMK), which generates a green fluorescence. This probe covalently binds to a reactive cysteine residue on the large subunit of the active caspase heterodimer, thereby inhibiting further enzymatic activity after being taken up into cells. The green fluorescence provides the direct measure of the active caspase present in the cell and it can be quantitated by flow cytometry or fluorescence plate reader, although it can be also analyzed by immunofluorescence to provide information on single cells that can be visualized by microscopy. 1. Transfer ~300 μl of each cell suspension (~106 cells) to sterile tubes. 2. Add 10 μl of freshly prepared 30× FLICA reagent and mix cells by flicking the tubes. 3. Incubate tubes for 1 h at 37 °C under 5 % CO2, protecting tubes from light. Swirl tubes once or twice during this time to gently resuspend the settled cells. 4. Add 2 ml of 1× wash buffer to each tube and mix gently. 5. Centrifuge the cells at 400 × g for 5 min at room temperature. 6. Remove the solution carefully and discard the supernatant. Gently vortex the cell pellet to disrupt any cell-to-cell clumping. 7. Wash the cells with 1 ml of 1× wash buffer. 8. Resuspend the cell pellet in 400 μl of 1× wash buffer. 9. For bicolor analysis, add 2 μl of PI solution to each cell suspension sample. Set aside a second suspension sample without PI. 10. For single-color analysis, the samples can be kept on ice and analyzed on the FL1 channel. Otherwise, 40 μl fixative can be added and cells can be stored at 2–8 °C, protected from light. However, for bicolor analysis, cells to be analyzed with PI cannot be fixed. Instead, they have to be analyzed immediately on the FL1 channel of FACS for fluorescein and FL2 channel for red fluorescence (see Note 3).

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Notes 1. Formalin-fixed and paraffin-embedded mouse [24] or patientderived human [25] tissue sections can be also examined. The slides are deparaffinized with xylene and graded alcohol and treated with citrate buffer (pH 6) for 20 min for antigen retrieval before incubation with primary antibodies. Sections are counterstained with hematoxylin before being examined under the microscope. Immunohistochemistry for caspase-3 can be combined with the in situ detection of apoptotic cells by terminal deoxynucleotide transferase-mediated dUTP nickend labeling (TUNEL) [25]. TUNEL or Comet assays are methods that detect DNA strand breaks that are associated with apoptosis. The samples are first immunostained for caspase-3 and, after washing in PBS, with a horseradish peroxidase (HRP)-linked secondary antibody. Immunoreactivity is visualized by a 10-min incubation with the HRP substrate diaminobenzidine. After staining for caspase-3, the same slides are then processed for in situ detection and localization of apoptosis at the level of single cells. Sections are then stained with anti-fluorescein antibodies linked with alkaline phosphatase, developed with Fast Red substrate and counterstained with hematoxylin. 2. The molecular weight of native PARP-1 is 110 kDa and that of cleaved PARP-1 is 86 kDa. The molecular weight of procaspase-3 is 32 kDa, while active caspase-3 migrates at 17 as well as 12 kDa. Some antibodies recognize only the pro-form of caspase-3, some recognize only the active form, and some can recognize both (information available from the respective data sheets). The primary antibodies can be reused for a couple of times if they are stored at 4 °C in the presence of sodium azide (0.01 %, w/v). 3. Flow cytometry analyses can be done with single-color (FLICA alone) or dual-color staining (FLICA and PI). It is recommended that induced and non-induced samples be run for each labeling condition (unlabeled, FLICA labeled, PI labeled, and FLICA/PI labeled).

Acknowledgments This work was supported by a research grant from the National Institutes of Health to A.A. (CA127264).

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References 1. Fuchs Y, Steller H (2011) Programmed cell death in animal development and disease. Cell 147:742–758 2. Autret A, Martin SJ (2009) Emerging role for members of the Bcl-2 family in mitochondrial morphogenesis. Mol Cell 36:355–363 3. Youle RJ, Strasser A (2008) The BCL-2 protein family: opposing activities that mediate cell death. Nat Rev Mol Cell Biol 9:47–59 4. Chen Q, Gong B, Almasan A (2000) Distinct stages of cytochrome c release from mitochondria: evidence for a feedback amplification loop linking caspase activation to mitochondrial dysfunction in genotoxic stress induced apoptosis. Cell Death Differ 7: 227–233 5. Gong B, Almasan A (2000) Apo2 ligand/ TNF-related apoptosis-inducing ligand and death receptor 5 mediate the apoptotic signaling induced by ionizing radiation in leukemic cells. Cancer Res 60:5754–5760 6. Hengartner MO (2000) The biochemistry of apoptosis. Nature 407:770–776 7. Kumar S (1999) Mechanisms mediating caspase activation in cell death. Cell Death Differ 6:1060–1066 8. Walker NP, Talanian RV, Brady KD, Dang LC, Bump NJ, Ferenz CR et al (1994) Crystal structure of the cysteine protease interleukin-1 beta-converting enzyme: a (p20/p10)2 homodimer. Cell 78:343–352 9. Rotonda J, Nicholson DW, Fazil KM, Gallant M, Gareau Y, Labelle M et al (1996) The three-dimensional structure of apopain/ CPP32, a key mediator of apoptosis. Nat Struct Biol 3:619–625 10. Wilson KP, Black JA, Thomson JA, Kim EE, Griffith JP, Navia MA et al (1994) Structure and mechanism of interleukin-1 beta converting enzyme. Nature 370:270–275 11. Thornberry NA, Rano TA, Peterson EP, Rasper DM, Timkey T, Garcia-Calvo M et al (1997) A combinatorial approach defines specificities of members of the caspase family and granzyme B. Functional relationships established for key mediators of apoptosis. J Biol Chem 272: 17907–17911 12. Fischer U, Janicke RU, Schulze-Osthoff K (2003) Many cuts to ruin: a comprehensive update of caspase substrates. Cell Death Differ 10:76–100 13. Gong B, Chen Q, Endlich B, Mazumder S, Almasan A (1999) Ionizing radiation-induced, Bax-mediated cell death is dependent on activation of cysteine and serine proteases. Cell Growth Differ 10:491–502

14. Mazumder S, Chen Q, Gong B, Drazba JA, Buchsbaum JC, Almasan A (2002) Proteolytic cleavage of cyclin E leads to inactivation of associated kinase activity and amplification of apoptosis in hematopoietic cells. Mol Cell Biol 22:2398–2409 15. Mazumder S, Gong B, Almasan A (2000) Cyclin E induction by genotoxic stress leads to apoptosis of hematopoietic cells. Oncogene 19:2828–2835 16. Mazumder S, Plesca D, Kinter M, Almasan A (2007) Interaction of a cyclin E fragment with Ku70 regulates Bax-mediated apoptosis. Mol Cell Biol 27:3511–3520 17. Plesca D, Mazumder S, Gama V, Matsuyama S, Almasan A (2008) A C-terminal fragment of Cyclin E, generated by caspase-mediated cleavage, is degraded in the absence of a recognizable phosphodegron. J Biol Chem 283:30796–30803 18. Mazumder S, Choudhary GS, Al-Harbi S, Almasan A (2012) Mcl-1 Phosphorylation defines ABT-737 resistance that can be overcome by increased NOXA expression in leukemic B cells. Cancer Res 72:3069–3079 19. Sharma A, Singh K, Mazumder S, Hill BT, Kalaycio M, Almasan A (2013) BECN1 and BIM interactions with MCL-1 determine fludarabine resistance in leukemic B cells. Cell Death & Disease 4:e628 20. Plesca D, Mazumder S, Almasan A (2008) DNA damage response and apoptosis. Methods Enzymol 446:107–122 21. Surova O, Zhivotovsky B (2013) Various modes of cell death induced by DNA damage. Oncogene 32:3789–3797 22. Gong B, Chen Q, Endlich B, Mazumder S, Almasan A (1999) Ionizing radiation-induced, Bax-mediated cell death is dependent on activation of serine and cysteine proteases. Cell Growth Diff 10:491–502 23. Chatterjee P, Choudhary GS, Sharma A, Singh K, Heston WD, Ciezki J et al (2013) PARP inhibition sensitizes to low dose-rate radiation TMPRSS2-ERG fusion gene-expressing and PTEN-deficient prostate cancer cells. PLoS One 8:e60408 24. Ray S, Almasan A (2003) Apoptosis induction in prostate cancer cells and xenografts by combined treatment with Apo2 ligand/tumor necrosis factor-related apoptosis-inducing ligand and CPT-11. Cancer Res 63:4713–4723 25. Masri SC, Yamani MH, Russell MA, Ratliff NB, Yang J, Almasan A et al (2003) Sustained apoptosis in human cardiac allografts despite histologic resolution of rejection. Transplantation 76:859–864

Chapter 2 Flow Cytometry Enumeration of Apoptotic Cancer Cells by Apoptotic Rate David Diaz, Alfredo Prieto, Eduardo Reyes, Hugo Barcenilla, Jorge Monserrat, and Melchor Alvarez-Mon Abstract Most authors currently quantify the frequency of apoptotic cells in a given phenotypically defined population after calculating the apoptotic index (AI), i.e., the percentage of apoptotic cells displaying a specific linage antigen (LAg) within a population of cells that remain unfragmented and retain the expression of the LAg. However, this approach has two major limitations. Firstly, apoptotic cells fragment into apoptotic bodies that later disintegrate. Secondly, apoptotic cells frequently lose, partially or even completely, the cell surface expression of the LAg used for the identification of specific cell subsets. This chapter describes a flow cytometry method to calculate the apoptotic rate (AR) that takes into account both cell fragmentation and loss of lineage antigen expression on measurement of apoptosis using flow cytometry ratiometric cell enumeration that emerges as a more accurate method of measurement of the occurrence of apoptosis in normal and tumoral cell cultures. Key words Apoptosis, Apoptotic rate, Apoptotic index, Cell enumeration, Accurate apoptosis measurement, Microbeads, Annexin V, Antigen loss, Cell fragmentation

1

Introduction

1.1 Apoptosis Measurement

The initial methods developed for the in vitro quantification of apoptosis measured phenomena associated with apoptosis in cultures at the population level, such as the assessment of nucleosomal DNA fragmentation after gel electrophoresis [1–3]. However, it soon became clear that individual cells undergo apoptosis in a heterogeneous and asynchronous manner [4]. It was therefore realized that the accurate measurement of apoptosis required methods that could identify apoptosis events at the single-cell level [5–11]. These methods revealed the heterogeneity of the apoptotic process to be correlated with cell phenotype—at least to a certain extent [12]. The ongoing development of flow cytometric techniques eventually made it possible to simultaneously identify and quantify apoptotic

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_2, © Springer Science+Business Media New York 2015

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cells phenotypically defined by the expression of their surface lineage antigens (LAg). The relevance of apoptosis has promoted active research into new methods of detecting these subcellular lesions at the singlecell level in complex cell mixtures both ex vivo and in cultured cells [5, 7, 13]. A good example is the use of annexin V for the detection of early apoptotic cells. In such cells, PS translocates from the inner side of the plasma membrane to the outer membrane leaflet, where it becomes exposed [14–16]. It can then be bound by annexin V, a phagocyte membrane protein [17]. The availability of fluorochrome-labeled recombinant soluble annexin V provides a useful tool for detecting and quantifying early apoptotic cells by flow cytometry [7, 8, 14]. The annexin V labeling method can be improved by the staining with the vital dye 7-amino-actinomycin D (7AAD) [11] to identify early and late apoptotic cells and necrotic ones. Cell washing, and choice of resuspension buffer, can affect the accuracy of measurements of apoptosis. It has been shown that wash cycles not only cause cell loss, but also affect the viability of cells, as well as the precision of repeat measurements. Therefore, wash cycles should be reduced to a minimum, which also reduces the time required for sample preparation. 1.2 Discrimination Between Whole Cells and Cell Fragments by Flow Cytometry

7AAD labeling can be used to discriminate between either viable and apoptotic whole cells, or cell fragments. Apoptosis led to the fragmentation of apoptotic cells into apoptotic bodies under different culture conditions (Fig. 1). Compared to whole cells, apoptotic bodies are smaller, consistent with the notion that one cell generates several apoptotic bodies. The inclusion of the latter in the cell analysis gate and their subsequent consideration as apoptotic cells result in an overestimation of the frequency of apoptosis and, therefore, discrimination between cells and apoptotic bodies is critical for accurate measurement of apoptosis. As shown in Fig. 1, the discrimination between either viable and apoptotic whole cells or cell fragments was achieved by the analyses of both their bivariate profiles of size (FSC)/DNA staining with 7AAD (left panels), and their FSC/granularity (SSC) distribution (right panels). Contour plots in the top panels show that freshly purified CD19+ lymphocytes formed a homogeneously sized population of viable cells that uniformly excluded 7AAD. After 24 h of culture (bottom panels) both apoptotic cells and apoptotic bodies emerged, but the remaining subset of viable B-cells maintain the characteristics from the original fresh B-lymphocytes, since they shared their FSC/SSC features (panel d), and did not take up 7AAD (contour levels under viable cell arrows in the panels c and d). In contrast, apoptotic cells showed a reduced FSC and a slightly increased SSC (panel d) that was coincident with variable 7AAD staining related to progression into late apoptosis (panel c). Finally, apoptotic bodies showed markedly smaller FCS and SSC signals than

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Fig. 1 Flow cytometry approach used to discriminate whole cells from apoptotic bodies by gating in 7AAD/FSC bivariate dot plots. Freshly purified CD19+ lymphocytes were labeled with CD19-APC, annexin V-FITC, and 7AAD. Flow cytometry analysis was performed prior to and after 24 h of culture. The experiment was repeated six times. Panels (a) and (b) show SSC/FSC and 7AAD/FSC bivariate contour plots of freshly purified B-cells. Panels (c) and (d) show how whole cells (R1) were differentiated from apoptotic bodies (R2, 7AAD−, and lower FSC signal than the lower limit of the 7AAD+ apoptotic cells) through combined analysis of the FSC/SSC/7AAD characteristic of the events measured

did live or apoptotic whole cells independently of their occasionally weak 7AAD staining (panels c and d). Thereafter, an event was considered to correspond to a whole cell when it provided an FSC signal greater than the lower limit of 7AAD+ apoptotic cells (insert of continuous line boxes in bottom panels). Using these criteria, whole apoptotic cells were clearly distinguishable from apoptotic bodies (inserts of discontinuous line boxes). 1.3 Apoptosis Quantification

Most authors currently quantify the frequency of phenotypically defined apoptotic cells after calculating the apoptotic index (AI), i.e., the percentage of apoptotic cells displaying a specific LAg within a population of cells that remain unfragmented and retain the expression of the LAg [18–20]. However, this approach has two major limitations. Firstly, apoptotic cells fragment into apoptotic bodies that later disintegrate. This leads to an underestimation

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of the percentage of apoptotic cells if the debris is excluded from the gates for cell analyses, or, alternatively, to the overestimation of apoptosis if several apoptotic bodies derived from a single cell are misinterpreted as individual apoptotic cells [21]. Secondly, apoptotic cells frequently lose, partially or even completely, the cell surface expression of the LAg used for the identification of specific cell subsets [22–24]; this means that the apoptotic cells from one phenotypically defined cell subset that lose the expression of their characteristic LAg can no longer be identified as targets in the apoptosis quantification, which leads to miscalculations [25]. The limitations of current flow cytometric approaches for evaluating apoptosis warrant the development of a new multiparameter method that (1) identifies and quantifies cells suffering apoptotic lesions in earlier stages of apoptosis; (2) discriminates live, necrotic, and apoptotic cells in a time frame within the death program that is well ahead of LAg loss and the generation of cell debris; and (3) extends AI to provide an estimate of the number of cells that have undergone apoptosis and its relation to the number of seeded cells: the apoptotic rate (AR). The AR overcomes the limitations of current flow cytometric techniques which do not use internal standards to determine absolute numbers. In previously described methods [6, 7, 9–11], AI has been used to measure the proportion of apoptotic cells in relation to the total number of detectable cells in the test tube at the end point of the cell culture assay. The enumeration of apoptotic cells by the AR reflects the proportion of cells that have undergone apoptosis in relation to the total number of cells seeded at the start point of the cell culture assay. This makes the estimation of the incidence of apoptosis more valid, since current methods ignore late apoptotic cells which have suffered LAg loss or fragmentation into apoptotic bodies. Therefore, the AR is a more sensitive indicator of apoptosis than the widely used AI. The ability to accurately and sensitively determine the number and population of cells undergoing apoptosis will allow great advances in evaluating new therapies targeted at inducing or inhibiting apoptosis. In addition, it could provide an early marker of therapeutic outcome, enabling clinicians to quickly determine if, for example, a new chemotherapeutic agent is successfully targeting neoplastic cells or if these are resistant to the therapy. The ease of use of flow cytometric techniques allows apoptosis to be used as a clinical parameter. Well-defined interpretations of results such as AR will help develop the use of apoptosis as a marker in making clinical decisions. A limitation of the proposed method is that AR can only be properly applied in time frames in which the in vitro cell proliferation does not alter significantly the number of cells in the culture. This time frame depends on the rate of proliferation of the studied cells. If the cells do not proliferate (i.e., B-chronic lymphocytic

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leukemia cells) then apoptosis can be measured by AR at 24 h or even 48 h of culture. When cells proliferate vigorously (i.e., certain tumor cell lines) it is necessary to perform the apoptosis assays after shorter periods of culture (3–6 h) to avoid interference of proliferative processes on the quantification of cell loss by apoptosis. In any case, even in conditions in which apoptosis and growth simultaneously occur, methods that enumerate apoptotic cells provide more information than those which only provide relative proportions of apoptotic cells. In summary, apoptosis cannot be accurately quantified by simply taking into account the percentage of cells that show apoptotic lesions. Single-cell approaches must therefore be used with care if occurrence of apoptosis is to be accurately evaluated, and should take into account absolute cell enumeration via the use of an internal microbead standard and the calculation of the AR.

2 2.1

Materials Equipment

1. Sterile 50 ml conical tubes. 2. 5 ml polystyrene round-bottom tubes 12 × 75 mm. 3. 96-Well flat-bottom culture plates. 4. Neubauer chamber. 5. FACSCalibur flow cytometer (Becton & Dickinson Biosciences).

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1. Complete medium: RPMI 1640 supplemented with 10 % heat-inactivated fetal calf serum, 25 mM Hepes, and 1 % penicillin-streptomycin. 2. 7-Aminoactinomycin D (7-AAD)—Highly toxic. 3. Ca2+-binding buffer: Hepes 10 mM, NaCl 150 nM, MgCl2 1 mM, CaCl2 1.8 mM, and KCl 5 mM; pH adjusted to 7.4. 4. Annexin V-binding buffer containing Ca2+: Hepes 10 mM, NaCl 150 nM, MgCl2 1 mM, CaCl2 1.8 mM, and KCl 5 mM; pH adjusted to 7.4. 5. Annexin V-FITC. 6. 6 μm CALIBRITE microbeads (Becton & Dickinson Biosciences). 7. Gelatin. 8. Trypan blue. 9. 0.5 × 10−6 M Staurosporine—Highly toxic. 10. 10−3 M Cycloheximide—Highly toxic. 11. 2 μg/ml Phytohemagglutinin—Highly toxic. 12. T Cell Expander (Dynal, Oslo, Norway).

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Methods The methods described below outline (1) preparation of the microbeads, (2) preparation of the cell suspension, (3) preparation of the culture, (4) preparation of the basal condition, (5) acquisition of cells after culture, and (6) calculation of the apoptotic rate.

3.1 Preparation of the Microbeads

One of the major problems of the use of microbeads in flow cytometric enumeration of cells is the possibility of adherence. Due to this, we need to block the adherence of microbeads to both the tube and the own microbeads by using gelatin in the solution used to dilute the microbeads. Another factor is the sedimentation of microbeads in the tube. Just before adding the microbeads to the cell sample, we need to vortex vigorously the microbead solution. 1. In a 50 ml conical tube, prepare a volume (μl) of Ca2+-binding buffer equal to 100 × number of sample tubes you will use (see Note 1). 2. Add gelatin 0.05 % (w/v). 3. Heat the tube in a thermal bath to 37 ºC for 30 min. 4. Keep at room temperature for 30 min. 5. Add CALIBRITE microbeads to the 50 ml tube to prepare a 1/100 (v/v) dilution. 6. Vortex the 50 ml tube during 1 min. 7. Store at 4 ºC in a refrigerator until use.

3.2 Preparation of the Cell Suspension

A cell suspension of tumor cells in complete medium must be obtained. This suspension could be homogeneous (i.e., tumoral cell line) or heterogeneous (i.e., peripheral blood mononuclear cells from a patient suffering from leukemia or tumor cells obtained from a tumor biopsy). All the protocol must be performed using sterile material in a laminar flow chamber. 1. Take 20 μl of the cell suspension and dilute it with 20 μl of trypan blue (see Note 2). 2. Mix gently and count the viable cells (cells without blue staining) in a Neubauer chamber. 3. Adjust the cells to a cell concentration of 0.5 × 106 viable cells/ml.

3.3 Preparation of the Culture

1. Add 100 μl of complete medium into three wells (triplicate) in 96 flat-bottom culture plates (see Note 3) and into three 5 ml polystyrene round-bottom tubes. 2. Add 100 μl of the diluted cells to the wells with complete medium (see Note 4) and into three 5 ml polystyrene roundbottom tubes to make the basal condition. 3. Culture the plate at 37 ºC in 5 % CO2 (see Note 5).

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1. Add to the tubes a combination of monoclonal antibodies labeled in FL-2 (i.e., phycoerythrin) and FL-4 (i.e., allophycocyanin) (see Note 6). 2. Incubate cells at 4 ºC in the dark for 20 min. 3. Centrifuge cells at 300 × g and 4 ºC for 5 min and decant the supernatant (see Note 7). 4. Resuspend cells and add 100 μl of Ca2+-binding buffer. 5. Add 6 μl of annexin V-FITC diluted 1/5 in Ca2+-binding buffer at 4 ºC in the dark for 10 min. 6. Add 100 μl of the prepared microbeads (remember to make a vigorous vortexing of the microbead solution before adding it to the cell suspension). 7. Add 100 ml of 7AAD diluted in Ca2+-binding buffer to a final concentration of 2.5 μg/ml and wait for 3–5 min (see Note 6). 8. Acquire the cell tubes in a four-color flow cytometer (see Note 8). You must make a cell gate around microbeads and adjust the number of acquired microbeads (i.e., 2,000 microbeads) to simplify the calculations to obtain the apoptotic rate.

3.5 Acquisition of Cells After Culture

1. Take out the volume of each well with a micropipette (see Note 9) and add it into 5 ml polystyrene round-bottom tubes. 2. Prepare the 24-h condition like Subheading 3.4 (see Note 10).

3.6 Calculation of the Apoptotic Rate

The calculation of apoptotic rate (AR) consists of two sequential steps. First, the number of events corresponding to cells which have finished the apoptotic process and have undergone fragmentation into apoptotic bodies or have completely lost the expression of surface markers is calculated from the difference between the number of events corresponding to seeded cells and that of cells which remain in culture and are LAg+ after challenge. Second, we sum to this number the number of events corresponding to annexin V+ cells and calculate the apoptosis occurrence with respect to the total number of seed cells: 1. NFC = NSC − NRC where NFC = events corresponding to fragmented cells or that completely lost the expression of their LAg; NSC = events corresponding to seeded cells; and NRC = events corresponding to the remaining cells, which include both annexin V+ and annexin V− cells. 2. The AR is then calculated by the following equation (see Notes 11 and 12): AR   NAV  C  NFC  / NSC

where NAV+C = events corresponding to the number of annexin V-positive cells.

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Notes 1. Prepare an extra 10 % more volume that you will need to be sure that you will have enough volume for all the tests if any problem arises. The minimum volume you must prepare is 10 ml because less volume cannot be shaken properly. Do not use microbeads prepared 10 or more days ago. 2. If you have an excessive number of cells per count chamber field you can dilute the cells in trypan blue until obtaining the proper dilution. A cell count per field between 30 and 120 cells allows accurate counting. 3. This is to measure spontaneous apoptosis. You can induce apoptosis by several kinds of apoptogens like etoposide, staurosporine, or cycloheximide or even study the activationinduced cell death induced by phytohemagglutinin or microbeads coated with anti-CD3 and anti-CD28 antibodies. 4. Critical step: It is very important to seed cells carefully because it affect so much to cell enumeration. 5. The time of culture should be adjusted depending on the apoptosis and cell growth properties of the tumor cells in culture. If the cells grow quickly then the time frame to measure apoptosis should be sorter. 6. If you want to make several different labeling of the cells with different combinations of antibodies you must prepare three culture wells and three 5 ml polystyrene round-bottom tubes for each combination to assess the precision of apoptosis measurement. If you do not want to label with 7AAD then you can add an additional monoclonal antibody labeled in FL-3 channel (i.e., peridinin chlorophyll protein conjugate). 7. Critical step: It is very important to decant cells carefully because it affect so much to cell enumeration due to cell loss. 8. You can label the cells with more or less fluorochrome-labeled antibodies depending on the technical characteristics of your flow cytometer. 9. Critical step: It is very important to take out cells carefully because it affects so much to cell enumeration due to cell loss. You must take out all the volume of the well. 10. It is critical to use for all the tests for a given experiment the same microbead solution for the reference of the ratiometric enumeration. 11. It should be noted that you can calculate the AR of a cell subpopulation defined by the expression of a cell marker and not only the total AR.

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12. It is possible to make a more immediate calculation of AR using the next equation: AR   NSC – NVC  / NSC

where NSC = events corresponding to seeded cells. NVC = events corresponding to viable cells after culture (number of annexin V-negative cells). References 1. Wyllie AH (1980) Glucocorticoid-induced thymocyte apoptosis is associated with endogenous endonuclease activation. Nature 284: 555–556 2. Russell JH, Dobos CB (1980) Mechanisms of immune lysis. II. CTL-induced nuclear disintegration of the target begins within minutes of cell contact. J Immunol 125:1256–1261 3. Cohen JJ, Duke RC (1984) Glucocorticoid activation of a calcium-dependent endonuclease in thymocyte nuclei leads to cell death. J Immunol 132:38–42 4. Khong HT, Restifo NP (2002) Natural selection of tumor variants in the generation of “tumor escape” phenotypes. Nat Immunol 3: 999–1005 5. Darzynkiewicz Z, Juan G, Li X, Gorczyca W, Murakami T, Traganos F (1997) Cytometry in cell necrobiology: analysis of apoptosis and accidental cell death (necrosis). Cytometry 27:1–20 6. Herault O, Colombat P, Domenech J, Degenne M, Bremond JL, Sensebe L, Bernard MC, Binet C (1999) A rapid single-laser flow cytometric method for discrimination of early apoptotic cells in a heterogeneous cell population. Br J Haematol 104:530–537 7. van Engeland M, Nieland LJW, Ramaekers FCS, Schutte B, Reutelingsperger CPM (1998) Annexin V-affinity assay: a review on an apoptosis detection system based on phosphatidylserine exposure. Cytometry 31:1–9 8. Vermes I, Haanen C, Steffens-Nakken H, Reutelingsperger CPM (1995) A novel assay for apoptosis. Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labelled Annexin V. J Immunol Methods 184:39–51 9. Gorczyca W, Gong J, Darzynkiewicz Z (1993) Detection of DNA strand breaks in individual apoptotic cells by the in situ terminal deoxynucleotidyl transferase and nick translation assays. Cancer Res 52:1945–1951

10. Gong J, Traganos F, Darzynkiewicz Z (1994) A selective procedure for DNA extraction from apoptotic cells applicable for gel electrophoresis and flow cytometry. Anal Biochem 218:314–319 11. Schmid I, Krall WJ, Uittenbogaart CH, Braun J, Giorgi JV (1992) Dead cell discrimination with 7-aminoactinomycin D in combination with dual color immunofluorescence in single laser flow cytometry. Cytometry 13:204–208 12. Pantaleo G, Graziosi C, Demarest JF, Butini L, Montroni M, Fox CH, Orenstein JM, Kotler DP, Fauci AS (1993) HIV infection is active and progressive in lymphoid tissue during the clinically latent stage of disease. Nature 362: 355–358 13. Ashkenazi A, Dixit VM (1998) Death receptors: signaling and modulation. Science 281:1305–1308 14. Koopman G, Reutelingsperger CPM, Kuijten GA, Keehnen RM, Pals ST, van Oers MH (1994) Annexin V for flow cytometric detection of phosphatidylserine expression on B cells undergoing apoptosis. Blood 84:1415–1420 15. Devaux PF (1991) Static and dynamic lipid asymmetry in cell membranes. Biochemistry 30:1163–1173 16. Zachowski A (1993) Phospholipids in animal eukaryotic membranes: transverse asymmetry and movement. Biochem J 294:1–14 17. Fadok VA, Voelker DR, Campbell PA, Bratton DL, Cohen JJ, Noble PW, Riches DW, Henson PM (1993) The ability to recognize phosphatidylserine on apoptotic cells is an inducible function in murine bone marrow-derived macrophages. Chest 103:102 18. Potten CS (1996) What is an apoptotic index measuring? A commentary. Br J Cancer 74: 1743–1748 19. Darzynkiewicz Z, Traganos F (1998) Measurement of apoptosis. Adv Biochem Eng Biotechnol 62:33–73

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20. Darzynkiewicz Z, Bedner E, Traganos F, Murakami T (1998) Critical aspects in the analysis of apoptosis and necrosis. Hum Cell 11:3–12 21. Prieto A, Díaz D, Barcenilla H, García Suárez J, Reyes E, Monserrat J, San Antonio E, Melero D, de la Hera A, Orfao A, Álvarez Mon-Soto M (2002) Apoptotic rate: a new indicator for the quantification of the occurrence of apoptosis in cell culture. Cytometry 48:185–193 22. Potter A, Kim C, Golladon KA, Rabinovith PS (1999) Apoptotic human lymphocytes have diminished CD4 and CD8 receptor expression. Cell Immunol 193:36–47 23. Philippé J, Louagie H, Thierens H, Vral A, Cornelissen M, De Ridder L (1997) Quantification of apoptosis in lymphocyte

subsets and effect of apoptosis on apparent expression of membrane antigens. Cytometry 29:242–249 24. Diaz D, Prieto A, Barcenilla H, Monserrat J, Prieto P, Sanchez MA, Reyes E, HernandezFuentes MP, de la Hera A, Orfao A, AlvarezMon M (2004) Loss of lineage antigens is a common feature of apoptotic lymphocytes. J Leukoc Biol 76:609–615 25. Prieto A, Reyes E, Diaz D, Hernandez-Fuentes MP, Monserrat J, Perucha E, Munoz L, Vangioni R, de la Hera A, Orfao A, AlvarezMon M (2000) A new method for the simultaneous analysis of growth and death of immunophenotypically defined cells in culture. Cytometry 39:56–66

Chapter 3 “Multiplexed Viability, Cytotoxicity, and Caspase Activity Assays” Andrew L. Niles and Terry L. Riss Abstract Multiplexed assay chemistries provide for multiple measurements of cellular parameters within a single assay well. This experimental practice is not only more cost efficient, but also provides more information about a compound or treatment. The ability to combine the activity profiles within the same sample provides a level of normalization not possible with parallel assays. Furthermore, multiplexing caspase activity assays with viability and/or cytotoxicity assays can support conclusions regarding cytotoxic mechanism and provide normalization, which may help correct for differences in cell number. Key words Multiplex, Fluorescence, Luminescence, Cell-based, Cytotoxicity, Caspase

1

Introduction The term “multiplex” is used extensively in biology to describe techniques or assays that capture more than one set of data from the same sample by measuring different parameters. The motivation for combining assays within the same well is twofold: reduction in cost of reagents, consumables, and operator time versus parallel assays, and the intrinsic power of intra-well response normalization. Regardless of which specific experimental application is being described, all multiplex assays have an obvious requirement for the combined assay chemistries to be compatible, distinct, and measurable. Assay signal separation is achieved by various means including using fluorophores with divergent excitation and emission spectra [1], using chemiluminescence and bioluminescence [2], using fluorescence and bioluminescence [3], or using bioluminescence in a sequential manner with the aid of a quenching agent [4]. For the purposes of this chapter, we describe only those chemistries germane to cytotoxicity, viability, and caspase activation assays.

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_3, © Springer Science+Business Media New York 2015

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1.1 A Real-Time Cytotoxicity Assay Multiplexed with an Endpoint Caspase Activity Assay

Caspase activation in cell culture is an early and definitive hallmark of apoptosis. Caspase activity is transient however, and greatly dependent upon the strength and mechanism of action of the stimulus [5]. Because of the variance in the kinetics of any individual caspase response, it is therefore experimentally possible to either underestimate or completely fail to detect caspase activation events using endpoint caspase activity assays employed at inappropriate time points. For this reason, caspase induction responses are typically addressed through time course experimentation using serial dilutions of the test article in parallel assay plates (Fig. 1). Therefore, any same-well multiplexed assay that improves experimental efficiency by indicating the proper “assay window” for optimal caspase measurement, while normalizing the data for individual well-towell variation, would be indicated and useful for profiling a compound or treatment [6]. It is often useful to correlate caspase activity (or lack of caspase activity) with global changes in cellular health [7, 8]. Cytotoxicity assays are one such barometer of cell health and measure the emergence of biomarkers resulting from changes in membrane integrity. Because maintenance of membrane integrity is integral

Fig. 1 The transience of a typical caspase induction response. Terfenadine was twofold serially diluted in RPMI 1640 + 10 % FBS in a sterile, opaque, 96-well plate. RPMI 1640 + 10 % FBS + 0.1 % DMSO served as the untreated/vehicle control. K562 cells were added in an equal volume to all wells of the plate and the compound exposure initiated at 37 °C at 5 % CO2. Replicate wells containing K562, terfenadine, and vehicle control were incubated for 6, 24, 48, and 72 h. Caspase-Glo® 3/7 Reagent (Promega) was prepared and added in 100 μl to specific time point replicates of the assay plate. Luminescence was measured using the GloMax Multi+™ (Promega). Notice the robust caspase activity response at 6 h which degrades to below control values by 48 and 72 h. Also note a nontypical, dose-dependent deviance in caspase activation at the highest concentrations of terfenadine which is indicative of poor compound solubility/availability during the exposure

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for viability, linking these biomarkers to caspase activation may indicate mechanism of action. For instance, caspase activation with a commensurate increase in cytotoxicity relative to the control treatment would strongly support the conclusion of programmed cell death by apoptosis [9]. However, cytotoxicity in the absence of caspase activation may indicate a primary necrotic event. The ability to distinguish between these forms of cytotoxicity in a plate-based format by a homogeneous multiplexed assay is often relevant from a therapeutic perspective, because apoptosis can often be modulated whereas the necrotic process is less understood and typically detrimental [10]. Although classical cytotoxicity chemistries can be combined with caspase activity assays with great success, these methods are relegated to endpoint determinations, which require parallel plates assayed at varying time points. A novel cytotoxicity probe has been recently developed that obviates these limitations [6]. This probe can be delivered to assay plates at the time of cell seeding or at dosing with test substances. Introduction of the probe in this manner is possible because it is both chemically stable and physiologically inert. The probe is a pro-fluorescent, asymmetric cyanine dye, which has an enhanced affinity for genomic DNA. The dye is normally excluded by healthy cell membranes until they lose integrity. Upon binding of the probe with DNA, the molecule becomes intensely fluorescent. Fluorescence associated with cell death can be evaluated either in real time (using environmentally controlled plate readers or optical imagers) or at selected time points using standard fluorometry [11]. A principle application for this probe is to define the first emergence of cytotoxicity, when endpoint caspase activity assays can be employed to determine the presence or absence of the apoptotic biomarkers. Together, the multiplexed biomarkers greatly reduce erroneous interpretations of whether caspase activation has occurred. 1.2 A Real-Time Cytotoxicity Assay Multiplexed with an Endpoint Viability and Caspase Activity Assay

Experimental manipulation of cells in culture leads to four general outcomes after a defined exposure: no effect relative to untreated or vehicle-treated control (with regard to viability or cytotoxicity), antiproliferative effects, enhanced proliferative rates, or cytotoxicity. The importance of establishing whether a compound or treatment produces nil or cytotoxic effects is obvious and well appreciated. Antiproliferative effects, however, are equally important for oncology efforts directed at characterizing existing or new antineoplastic agents [12]. Skillful use of multiplexed viability and cytotoxicity assays can be particularly useful in revealing both cytostatic and early cytotoxic effects and hence the general tolerability or cytotoxic potential of a test compound [13]. For instance, lower viable cell number relative to control without commensurate increases in cytotoxicity is consistent with cytostatic effects. The discordance in these multiplexed biomarkers indicates that

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there is a difference in the proliferative rate in treated versus untreated cells (and therefore viable cell number) that cannot be attributed to cell death. More commonly, cytostatic effects are accompanied by dose-dependent cytotoxic effects, albeit separated by effective potencies early in the progression of the cytotoxic response. Although proliferative effects can also be monitored with multiplexed viability and cytotoxicity assays, these responses are not directly relevant to studies involving caspase activation. Procedurally, multiplexed viability/and or cytotoxicity assays are easily configured by the sequential addition of viability and/or cytotoxicity reagents to the assay well prior to homogeneous caspase reagent addition. This sequence-dependent addition is necessary because homogeneous caspase activity assay buffers contain lytic components, which would affect cellular viability and interfere with meaningful viability or cytotoxicity measurements.

2 2.1

Materials Equipment

1. 15 ml conical tubes. 2. 1.5 ml microfuge tubes. 3. 75 cm2 culture flasks. 4. Reagent reservoirs. 5. Sterile pipette tips. 6. Single- and multichannel micro-pipettors. 7. Hemocytometer and cover slip. 8. Bright-field microscope. 9. Tabletop centrifuge. 10. Opaque multi-well plate. 11. Monochronometer-based fluorometer equipped with the following filter pairs:

or

fluorometer

400ex 505em 485–500ex 520–535em 12. Multi-well luminometer. 13. CO2 incubator. 2.2

Reagents

1. Preferred cell line, for this chapter we used K562. 2. Appropriate cell culture medium. 3. Fetal bovine serum. 4. Trypan blue solution. 5. Cytotoxic compound, for this chapter we used terfenadine and bortezomib. 6. Dimethyl sulfoxide or appropriate solvent.

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7. Caspase-Glo®-3/7 Assay (Promega). 8. CellTox™ Green Assay (Promega). 9. CellTiter-Fluor™ Assay (Promega).

3

Methods The methods described below outline experimental examples of (a) a real-time fluorogenic cytotoxicity assay (Fig. 2) multiplexed with a luminogenic caspase 3/7 activity assay (Fig. 3a, b), and (b) a real-time fluorogenic cytotoxicity assay multiplexed with a spectrally distinct viability assay and a luminogenic caspase 3/7 activity assay (Fig. 4a, b).

3.1 A Real-Time Cytotoxicity Induction Time Course: Establishing Mechanism of Action

The following protocol produces dose-dependent induction of caspases 3 and 7 which leads to time-dependent changes in membrane integrity by apoptosis. The proteasome inhibitor, bortezomib, is used in this model with the myelogenous leukemia cell line, K562. This protocol is written for cells cultured in suspension but can be amended to explore the kinetics of caspase activation using nearly any cell line, treatment, or test article. Attachmentdependent cells should be added to plates and allowed to recover/ equilibrate at least 2–16 h prior to dosing.

Fig. 2 A real-time cytotoxicity induction time course. Bortezomib was twofold serially diluted in RPMI 1640 + 10 % FBS and added to replicate wells of a sterile opaque white plate in 50 μl. A series of wells received medium only to serve as untreated control. K562 cells were adjusted to 100,000 cells/ml in RPMI 1640 + 10 % FBS and made 1:500 with CellTox™ Green, and then seeded at 5,000 cells/well in 50 μl. Fluorescence was measured using a GloMax Multi+™ at 4 and 24 h until a significant cytotoxicity response was observed. The time course was extended in parallel wells to include a 48 h cytotoxicity time point

Andrew L. Niles and Terry L. Riss

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Fig. 3 A real-time cytotoxicity induction time course multiplexed with a caspase activity assay. Bortezomib was twofold serially diluted and added to K562 cells in the presence of CellTox™ Green as described in Fig. 2. (a) After 24 h of exposure, Caspase-Glo™ 3/7 Reagent was added in an additional 100 μl per well and mixed briefly to ensure cell/reagent homogeneity. After 30 min of incubation, luminescence was measured using a GloMax Multi+™. (b) After 48 h of exposure Caspase-Glo™ 3/7 Reagent was added to a second set of replicate wells and luminescence measured. Note the difference in caspase activation magnitude between the two time points and the difference in caspase induction potencies versus cytotoxicity, suggesting appropriate timing to measure the full magnitude of the caspase induction response

To avoid culture contamination by microorganisms, conduct cell-based assay experiments in a laminar flow hood or clean-room environment using aseptic technique with sterile reagents and consumables. 1. Solubilize the caspase induction agent and test compound. For example, dilute bortezomib to a 10 mM stock solution in sterile DMSO.

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Fig. 4 A real-time cytotoxicity induction time course multiplexed with an endpoint viability and caspase activity assay. Bortezomib was twofold serially diluted and added to K562 cells in the presence of CellTox™ Green as described in Fig. 2. After 24 h (a) and 48 h (b) of exposure, 20 μl CellTiter-Fluor was added to the wells, mixed briefly, and then incubated at 37 °C for 30 min. Fluorescence was measured at 400ex 505em using the GloMax Multi+. Next, Caspase-Glo™ 3/7 Reagent was added in an additional 100 μl per well and mixed briefly to ensure cell/reagent homogeneity. After 30 min of incubation, luminescence was measured using a GloMax Multi+™. Notice that bortezomib potency against cell number (viability) is substantially more potent than cytotoxicity, indicating that proliferation was mechanistically halted prior to progression through apoptosis

2. Next, prepare an intermediate 20 μM dilution of compound in complete medium. 3. Using a multichannel pipettor, add 50 μl volumes of complete medium to all wells of columns 2–12 of a sterile, opaque (white) 96-well plate. Twofold serially dilute apoptosis inducer and test compound by adding 50 μl of 20 μM agents

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to columns 1 and 2, rows A–D and E–H, respectively. Gently mix compound/dilution medium in column 2 by aspirating and dispelling 50 μl volumes at least five times. After mixing, remove 50 μl from each well in column 2 and proceed to column 3. Repeat through column 10, but remove 50 μl to waste. Column 11 will serve as untreated control and column 12 will serve as a reagent background control. 4. Harvest cells from culture flasks. Remove a representative volume (1,000 μl or less) for analysis. Gently pellet the remaining pool by centrifugation at 200 × g for 8–10 min at RT. 5. Examine a dilution of the cell sample by trypan blue exclusion to determine the population viability and cell count. If viability is greater than 90 %, add complete medium (with serum and nutrient/cofactor adjuncts) so that the cells are at a density of 100,000 viable cells/ml. 6. Add CellTox™ Green to cells at a dilution of 1:500 (e.g., 20 μl of CellTox™ Green/10 ml of cells). Mix by inversion. Prepare a CellTox™ Green background control by adding 4 μl of the probe to 2 ml of complete medium. 7. Plate the cell/dye mix in 50 μl (5,000 cells/well) to columns 1–11 of the microtiter plate containing diluted compounds. Add 50 μl of dye background control to replicate wells of column 12 to serve as a cell-free reagent background control. 8. Mix the plate by orbital shaking at 500–700 RPM after each addition to insure homogeneity and compound dispersion. 9. Incubate the plate at 37 ºC in a standard humidified CO2 incubator during the exposure period. Alternatively, the plate can be placed in an environmentally controlled fluorometer (such as Tecan Infinite 200 with Gas Control Module™) and fluorescence collected in kinetic mode at 485–500ex 520–535em. 10. If using standard fluorometry, remove the plate from the incubator at predetermined intervals to assess cytotoxicity. The time points of 4, 12, 24, 36, 48, and 72 h are typically suitable for addressing a kinetic response. Measure fluorescence at 485–500ex 520–535em. Analyze data by comparing treated wells to untreated wells. Background subtraction of cell-free wells can be helpful in this analysis. Commercial software (GraphPad by Prism, Sigma Plot, etc.) utilize well-validated regressions and fits that may also aid analysis. 11. At the first emergence of statistically significant increases in fluorescence in test or control inducer wells relative to untreated control, abort the time course and proceed to the luminescent caspase assay. 12. Equilibrate the frozen Caspase-Glo®-3/7 Buffer and lyophilized Caspase-Glo®-3/7 Substrate to room temperature prior

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to use. The buffer may be thawed in a 37 °C water bath. Remove the buffer from the water bath immediately after thawing. 13. Transfer the Caspase-Glo®-3/7 Buffer to the lyophilized Caspase-Glo®-3/7 Substrate to create the reagent. Mix by swirling or by inverting the contents until the substrate is completely dissolved. 14. Add 100 μl of Caspase-Glo®-3/7 Reagent to each well of columns 1–12. Mix briefly using an orbital shaker. After 20–30 min of incubation at RT, measure the luminescent signal associated with caspase-3/7 activity. 15. Plot the background-subtracted fluorescence and luminescence data as a percentage of untreated control versus compound concentration. Determine whether caspase activity correlates positively with cytotoxicity. 3.2 A Real-Time Cytotoxicity Induction Time Course Multiplexed with a Viability and Caspase Activity Assay

As established in Subheading 3.1, the inclusion of a real-time cytotoxicity probe (like CellTox™ Green) can be indispensable for establishing the kinetics and progression of an apoptotic response. However, orthogonal measures of cell health and viable cell number can also provide valuable insight into a compound or treatment’s mechanism of action. The CellTiter-Fluor™ assay multiplexed in this protocol measures a constitutive and conserved proteolytic activity resident only in viable mammalian cells that is notably unaffected by metabolic rate or status of a cell [14]. This “live cell protease activity” is measured using a fluorogenic, cellpermeable, peptide substrate. The substrate enters intact cells where it is cleaved to generate a fluorescent signal proportional to the number of living cells. This live cell protease activity marker becomes inactive upon loss of membrane integrity and leakage into the surrounding culture medium. This biomarker defines the relative number of viable cells in a well after treatment and may, or may not, inversely correlate with the emergence of cytotoxicity biomarkers. 1. Solubilize the caspase induction agent and test compound. For example, dilute bortezomib to a 10 mM stock solution in sterile DMSO. 2. Next, prepare an intermediate 20 μM dilution of compound in complete medium. 3. Using a multichannel pipettor, add 50 μl of complete medium to all wells of columns 2–12 of a sterile, opaque (white) 96-well plate. Twofold serially dilute apoptosis inducer and test compound by adding 50 μl of 20 μM agents to columns 1 and 2, rows A–D and E–H, respectively. Gently mix compound/dilution medium in column 2 by aspirating and dispelling 50 μl at least five times. After mixing, remove 50 μl from each well in column 2 and proceed to column 3. Repeat

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through column 10, but remove 50 μl to waste. Column 11 will serve as untreated control and column 12 will serve as a reagent background control. 4. Harvest cells from culture flasks. Remove a representative volume (1,000 μl or less) for analysis. Gently pellet the remaining pool by centrifugation at 200 × g for 8–10 min at RT. 5. Examine a dilution of the cell sample by trypan blue exclusion to determine the population viability and cell count. If viability is greater than 90 %, add complete medium (with serum and nutrient/cofactor adjuncts) so that the cells are at a density of 100,000 viable cells/ml. 6. Add CellTox™ Green to cells at a dilution of 1:500 (e.g., 20 μl of CellTox™ Green/10 ml of cells). Mix by inversion. Prepare a CellTox™ Green background control by adding 4 μl of the probe to 2 ml of complete medium. 7. Plate the cell/dye mix in 50 μl (5,000 cells/well) to columns 1–11 of the microtiter plate containing diluted compounds. Add 50 μl of dye background control to replicate wells of column 12 to serve as a cell-free reagent background control. 8. Mix the plate by orbital shaking at 500–700 RPM after each addition to insure homogeneity and compound dispersion. 9. Incubate the plate at 37 ºC in a standard humidified CO2 incubator during the exposure period. Alternatively, the plate can be placed in an environmentally controlled fluorometer (such as Tecan Infinite 200 with Gas Control Module™) and fluorescence collected in kinetic mode at 485–500ex 520–535em. 10. If using standard fluorometry, remove the plate from the incubator at predetermined intervals to assess cytotoxicity. The time points of 4, 12, 24, 36, 48, and 72 h are typically suitable for addressing a kinetic response. Measure fluorescence at 485–500ex 520–535em. Analyze data by comparing treated wells to untreated wells. Background subtraction of cell-free wells can be helpful in this analysis. Commercial software (GraphPad by Prism, Sigma Plot, etc.) utilize well-validated regressions and fits that may also aid analysis. 11. At the first emergence of statistically significant increases in fluorescence in test or control inducer wells relative to untreated control, abort the time course and proceed to the CellTiterFluor™ portion of the multiplex. 12. Equilibrate the frozen CellTiter-Fluor™ substrate and buffers to 37 °C by immersion in a water bath. Vortex both substrate and buffer after thawing to insure homogeneity. 13. Create a 5× CellTiter-Fluor™ Reagent by adding 10 μl of the GF-AFC substrate to 2.0 ml of CellTiter-Fluor™ Buffer. The solution may appear “milky” upon addition of the substrate, but will quickly become clear with vortexing.

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14. Add 20 μl of the 5× CellTiter-Fluor™ Reagent to all wells of the 96-well plate. Mix by orbital shaking at 500–700 RPM for 1 min and then return to a 37 °C incubator for at least 30 min. Although 30 min of incubation is typically sufficient to achieve suitable assay sensitivity, additional incubation (up to 3 h) may improve the dynamic range of the assay. 15. Measure fluorescence at 400ex 505em.. The fluorescent product of this assay is reasonably stable, allowing for multiple reads to optimize photomultiplier gain settings, etc. After a satisfactory signal is obtained, proceed to the luminescent caspase portion of the multiplex. 16. Equilibrate the frozen Caspase-Glo®-3/7 Buffer and lyophilized Caspase-Glo®-3/7 Substrate to room temperature prior to use. The buffer may be thawed in a 37 °C water bath. Remove the buffer from the water bath immediately after thawing. 17. Transfer the Caspase-Glo®-3/7 Buffer to the lyophilized Caspase-Glo®-3/7 Substrate to create the reagent. Mix by swirling or by inverting the contents until the substrate is completely dissolved. 18. Add 100 μl of Caspase-Glo®-3/7 Reagent to each well of columns 1–12. Mix briefly using an orbital shaker. After 20–30 min of incubation at RT, measure the luminescent signal associated with caspase-3/7 activity. 19. Plot the background-subtracted fluorescence and luminescence data as a percentage of untreated control versus compound concentration. Determine whether caspase activity correlates positively with cytotoxicity and negatively with viability.

4

Notes 1. The kinetics and potency of apoptosis inducers may vary greatly with respect to cell type tested and molecular target/pathway of caspase induction [15]. Efforts directed at optimizing and standardizing cell culture conditions prior to dosing will improve inter-assay response variation. 2. Although statistically rare, all viability, cytotoxicity, and caspase activity assay chemistries are subject to both specific and nonspecific interferences. Compounds with fluorescent properties or intense absorption in the visible spectrum may complicate assay measures or cause optical quenching. Unusually rich medium (20 % or more animal serum) or complex support matrices (collagen or Matrigel) may also limit the availability of assay reactants and therefore sensitivity. All nonconforming data sets should be further examined for possible artifacts.

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3. CellTox™ Green stains the DNA of membrane-compromised cells. With attachment-dependent cells, dead cell fluorescence may be unequally represented within the geometry of the monolayer of the well. Furthermore, cells that have become membrane compromised or have lifted off the plate surface may no longer retain their original DNA content. Accurate, multi-well measures can be achieved in these situations by either orbital shaking at 700–900 RPM prior to measuring fluorescence or employing an “optical averaging” feature available on most commercial fluorometers. 4. When multiplexing a viability or cytotoxicity assay with a caspase activity measure, always follow a sequential order of assay chemistry addition. This is required because homogeneous caspase assay formulations contain agents that lyse cells. Therefore, addition of caspase reagents prior to viability or cytotoxicity reagents would automatically kill all cells in the culture. 5. Normalization of caspase activity responses to cell number by viability or cytotoxicity assays is particularly useful in replicate wells of multi-well formats. However, care should be taken in such normalization because excessive clumping may lead to nonlinear viability and cytotoxicity responses. Furthermore, bacterially contaminated wells may contribute to viability and cytotoxicity measures. References 1. Grant S, Sklar J, Cummings R (2002) Development of novel assays for proteolytic enzymes using rhodamine-based substrates. J Biomol Screen 7:531–540 2. Bronstein I, Fortin J, Stanley PE, Stewart GSAB, Kricka LJ (1994) Chemiluminescent and bioluminescent reporter gene assays. Anal Biochem 219:169–181 3. Wesierska-Gadek J, Gueorguieva M, Ranftler C, Zerza-Schnitzhofer G (2005) A new multiplex assay allowing simultaneous detection of the inhibition of cell proliferation and induction of cell death. J Cell Biochem 96:1–7 4. Nieuwenhuijsen B, Huang Y, Wang Y, Ramerez F, Kalgaonkar G, Young K (2003) A dual luciferase multiplex high-throughput screening platform for protein-protein interactions. J Biomol Screen 8:676–684 5. Hook B, Schagat T (2012) Profiling compound effects on cell health in a time course using a multiplexed, same-well assay. PubHub Online 6. Niles A, Worzella T, Zhou M, McDougall M, Lazar D (2012) Measuring cytotoxicity in real time with a highly stable green dye. [Internet] December 2012

7. Niles A, Moravec R, Riss T (2004) Characterizing responses to treatments using homogeneous caspase activity and cell viability assays. Cell Notes 9:11–14 8. Niles A, Worzela T, Scurria M, Daily W, Bernad L, Guthmiller P, McNamara B, Rashka K, Lange D, Riss T (2006) Multiplexed viability, cytotoxicity and apoptosis assays for cell-based screening. Cell Notes 16:12–15 9. Niles A, Moravec R, Riss T (2008) Update on in vitro cytotoxicity assays for drug development. Expert Opin Drug Discov 3:655–669 10. Leist M, Jaattela M (2001) Four deaths and a funeral: from caspases to alternative mechanisms. Nat Rev Mol Biol 2:1–10 11. Worzella T et al (2013) Real-time cytotoxicity analysis using CellTox™ green cytotoxicity assay and the Tecan Infinite® 200 PRO with gas control module. [Internet] May 2013 12. Caraglia M, Santini D, Marra M, Vincenzi B, Tonini G, Budillon A (2006) Emerging anticancer molecular mechanisms of aminobisphosphonates. Endocr Relat Cancer 13:7–26 13. Hook B, Bratz M, Schagat T (2013) Gain more informative data by multiplexing a fluorescent

Multiplexed Cell Death Assays real-time cytotoxicity assay with luminescent, fluorescent or colorimetric viability assays. [Internet] June 2013 14. Niles A, Moravec R, Hesselberth P, Scurria M, Daily W, Riss T (2007) A homogeneous assay to measure live and dead cells in the same

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sample by detecting different protease markers. Anal Biochem 366:197–206 15. Riss T, Moravec R (2004) Use of multiple assay endpoints to investigate the effects of incubation time, dose of toxin, and plating density in cellbased assays. Assay Drug Dev Technol 2:51–62

Chapter 4 A Multiplexed Method for Kinetic Measurements of Apoptosis and Proliferation Using Live-Content Imaging Katherine Artymovich and Daniel M. Appledorn Abstract In vitro cell proliferation and apoptosis assays are widely used to study cancer cell biology. Commonly used methodologies are however performed at a single, user-defined endpoint. We describe a kinetic multiplex assay incorporating the CellPlayerTM NucLight Red reagent to measure proliferation and the CellPlayerTM Caspase-3/7 reagent to measure apoptosis using the two-color, live-content imaging platform, IncuCyteTM ZOOM. High-definition phase-contrast images provide an additional qualitative validation of cell death based on morphological characteristics. The kinetic data generated using this strategy can be used to derive informed pharmacology measurements to screen potential cancer therapeutics. Key words Kinetic multiplex cell death assay, Real-time imaging, Caspase-3, Fluorescent imaging

1

Introduction Careful regulation of cell proliferation and cell death is critical to proper tissue development and maintenance. Programmed or regulated cell death—apoptosis—is executed by intrinsic or extrinsic factors which, in most cases, activate a cascade of cysteinyl aspartate proteinases, known as caspases. Abnormalities in the proliferative and/or apoptotic signaling cascades can lead to uncontrolled cell growth and tumor formation [1]. Therefore, in vitro cell proliferation and apoptosis assays have been widely used to study cell biology and screen potential cancer therapeutics. Current in vitro proliferation assays measure metabolic activity and DNA synthesis of viable cells. Metabolic assays utilize substrates that emit a colorimetric/fluorometric signal when reduced by the metabolically active cellular environment. These signals can be quantitatively measured using a spectrophotometer [2]. The disadvantage of these assays is that an exact cell count and morphological changes are often not assessed. Alternatively, antibody and chemical probes can be used to stain nuclei in order to measure cell count using a high-content imaging device; yet, due to

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_4, © Springer Science+Business Media New York 2015

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the toxic nature of these dyes, they cannot be used for the long-term evaluation of proliferation. Caspase-3 is the primary executioner in the apoptotic signaling cascade, which makes it a popular marker for in vitro apoptosis assays. Many of these assays utilize the DEVD (Asp-Glu-Val-Asp) motif, which is recognized by caspase-3 [3]. Fluorescent substrates, such as luciferase, incorporate the DEVD motif and release a fluorescent signal when bound to caspase-3. Similar to proliferation assays, these methodologies are performed at a single, user-defined endpoint and often require multiple wash steps resulting in the loss of cells or critical data when cells undergo apoptosis at different rates. We present a kinetic multiplex assay incorporating the CellPlayer NucLight Red reagent to measure proliferation and the CellPlayer Caspase-3/7 reagent to measure apoptosis using the two-color, live-content imaging platform, IncuCyte ZOOM (Essen BioScience). The NucLight Red reagent is a lentivirusbased reagent which stably integrates into the host cell line and labels the nuclei in a non-perturbing way (Fig. 1a). The caspase3/7 reagent is an inert, non-fluorescent substrate that freely crosses the cell membrane and is cleaved by activated caspase-3/7 resulting in the release of a green DNA-binding dye. In combination, the NucLight Red label and the caspase-3/7 reagent provide a multiplexed way to differentiate inhibition of cell growth and induction of cell death, respectively (Fig. 1b). High-definition phase-contrast images provide an additional qualitative validation of cell death based on morphological characteristics. Finally, using any number of strategies (e.g., area under the curve, max counts, single time points), the kinetic data generated using this strategy can be used to derive informed pharmacology measurements.

Fig. 1 HD images of multiplexed proliferation/apoptosis assay. (a) Untreated HeLa NucLight Red cells. (b) HeLa NucLight Red cells treated with 300 nM staurosporine. Blended phase-contrast and red/green images taken with a 20× objective illustrate homogenous expression of the red nuclear signal and activation of caspase-3/7 (green)

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Materials

2.1 Equipment and Consumables

1. IncuCyte ZOOM™ (Essen BioScience, Ann Arbor, MI, USA): 4× or 10× magnification, Dual Color Model 4459. 2. Two 96-well plates: One plate for setting up dilutions, one tissue culture-treated plate for cells and assay (see Note 1). 3. Multichannel pipette: 12-channel, 30–300 μL. 4. Consumables: Sterile pipette tips, 50 mL reservoirs, 15 mL conical tubes, 1.5 mL microfuge tubes.

2.2 Cell Line Components

1. CellPlayer™ NucLight Red Cell Line (see Note 2). 2. Low riboflavin media: F-12 or MEM (see Note 3). 3. 10 % FBS. 4. Puromycin (see Note 4).

2.3 Reagents and Compounds

1. Caspase-3/7 Reagent (Essen BioScience, Ann Arbor, MI, USA, Cat # 4440). 2. Compounds (see Note 5): (a) Staurosporine: Molecular weight, 466.53 g/mol. Make working stock of 500 μM in 100 % DMSO. (b) Camptothecin: Molecular weight, 348.35 g/mol. Make working stock of 1 mM in 100 % DMSO. (c) Cycloheximide: Molecular weight, 281.35 g/mol. Make working stock of 1 mM in 100 % DMSO.

3

Methods

3.1 Cell Culture and Seeding Assay Plate

1. Prior to beginning the assay, grow cells to 80 % confluence in a 25 or 75 cm2 tissue culture-treated flask. 2. Wash, trypsinize, and count cells. Dilute cells to a final density of 5 × 104 cells/mL in 12 mL complete media without antibiotics (see Note 6). 3. Mix cell solution well and carefully pour into a 50 mL reservoir. 4. Immediately, use a multichannel pipette to dispense 100 μL per well (5,000 cells/well) in the 96-well assay plate. 5. Allow plate to incubate at ambient temperature for 15 min before placing in the 5 % CO2, 37 °C, humidified incubator overnight (see Note 7).

3.2 Preparation of Reagent Plate

1. Dilute caspase-3/7 reagent (5 mM stock) to 5 μM in 15 mL complete media without antibiotics (see Note 8).

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2. Aliquot 1 mL of caspase-3/7 media into three Eppendorf tubes (see Notes 5 and 9). (a) Dilute staurosporine to 1 μM in one tube (2 μL of 500 μM staurosporine) and dispense 200 μL to wells A1–A4 of the reagent plate. (b) Dilute camptothecin to 4 μM in one tube (4 μL of 1 mM camptothecin) and dispense 200 μL to wells A5–A8 of the reagent plate. (c) Dilute cycloheximide to 3 μM in one tube (3 μL of 1 mM cycloheximide) and dispense 200 μL to wells A9–A12 of the reagent plate. 3. Pour the remaining caspase-3/7 media solution into a 50 mL reservoir and using the multichannel pipette, dispense 120 μL of caspase-3/7 media to rows B–H of the reagent plate. 4. Use a multichannel pipette to dilute compounds from rows A to G in a threefold serial dilution by adding 60µl into 120µl changing pipette tips between each row (row H is a negative control). 3.3 Transfer Compounds from Reagent Plate to Assay Plate

3.4 IncuCyte ZOOM™ Setup and Image Processing

1. Remove the 96-well assay plate containing cells from the incubator. Using a multichannel pipette, carefully aspirate the media from the cells. 2. Using a multichannel pipette, immediately transfer 100 μL from rows A to H of the reagent plate to the assay plate containing cells (Fig. 2 for plate setup). Remove any air bubbles if necessary (see Note 10). 1. Immediately place assay plate into the IncuCyte ZOOM™ and allow it to equilibrate for 15–30 min before starting the first scan (see Note 11). 2. Set the scan schedule: (a) Scan type: Standard. (b) Channel selection: Phase, green, red. (c) Spectral unmixing: 8 % of Red removed from green. (d) Set interval every 2 h. (e) Job type and processing definition (see Note 12). 3. Scan every 2–3 h for a total of 48 h (see Note 13). 4. Automated image processing on the fly is accomplished by applying an appropriate processing definition. Data can be graphed as soon as image analysis is completed (see Note 12).

3.5 Exporting and Analyzing Data

1. Kinetic data can be visualized within the IncuCyte ZOOM software. If additional pharmacological calculations are desired, complete the following steps.

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Fig. 2 Plate map. An example plate map showing that NucLight Red cells can be treated with several dilutions of multiple compounds in the presence of caspase-3/7 reagent

2. Export object count/mm2 of green caspase-3/7 objects and red nuclear objects from each well at each time point into excel and copy the data into a graphing software (we recommend GraphPad Prism). 3. Plot the object counts/mm2 versus time and calculate the area under the curve (AUC) for each well (Fig. 3a, b). 4. For each drug, use the replicate AUC values for each concentration to calculate EC50 and IC50 values (see Note 14). (a) Transform the concentrations for each compound using X = Log(X), where X = concentration in molar. (b) Use the nonlinear regression (log(agonist or inhibitor) versus response—variable slope) to calculate EC50 or IC50 values, respectively (Fig. 3c).

4

Notes 1. Any tissue culture-treated 96-well plate supported by IncuCyte ZOOM will work for this assay (we recommend Corning® 96 Well Clear Flat Bottom TC-Treated Microplate, Cat # 3595).

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Fig. 3 Kinetic and pharmacological analysis of nuclear counts and caspase-3/7 activation. HeLa NucLight Red cells were treated with varying concentrations of staurosporine in the presence of the caspase-3/7 reagent. (a) Nuclear objects and (b) caspase-3/7-positive objects were measured over time in response to increasing concentrations of staurosporine. (c) The area under the curve (AUC) from graphs (a) and (b) were used to calculate IC50 and EC50 values, respectively

2. This assay may be performed with a nonnuclear labeled cell line to measure only apoptosis, but the nuclear label is necessary to kinetically quantify proliferation in addition to apoptosis. We suggest using one of Essen BioScience’s pre-made NucLight Red cell lines, HT1080 (fibrosarcoma, Cat # 4485), MDA-MB-231 (breast adenocarcinoma, Cat # 4487), HeLa (cervical carcinoma, Cat # 4489), or A549 (lung epithelial carcinoma, Cat # 4491). Alternatively, any primary or immortal cell line may be infected with Essen BioScience’s nonperturbing lentivirus reagent, NucLight Red (Cat #s 4476 and 4478), to stably express a nuclear restricted red fluorescent protein. 3. Riboflavin, an essential vitamin in many media formulations, contributes to high background fluorescence in the green channel of images. Media with low riboflavin, such as F-12 or MEM, should be used to produce the best signal:noise ratio for the caspase-3/7 reagent. We suggest Ham’s F-12 + GlutaMAX™-1 (Cat # 31765, Gibco/Life Technologies). 4. If using one of Essen BioScience’s pre-made NucLight Red cell lines, maintain expression of the nuclear restricted red fluorescent protein by growing cells in 0.5 μg/mL puromycin. If using a different NucLight Red cell line, it is the user’s

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responsibility to optimize the effective puromycin or bleomycin concentration for that cell type according to the Essen BioScience’s lentivirus reagent instructions. Puromycin or bleomycin should be removed during the assay. 5. Any drug compound can be tested in this assay. Dilute all compounds to a working stock concentration in 100 % DMSO so that the final concentration of DMSO in the experiment does not exceed 0.4 %. The protocol outlined above tests three common apoptotic and cytostatic compounds. If testing new drug compounds, we recommend including one known apoptotic compound (staurosporine) and one known cytostatic compound (cycloheximide). 6. The cell seeding density will vary depending on cell type. Several cell seeding densities should be tested so that cells are 25–35 % confluent at the start of the assay. 7. Cell seeding techniques are a critical part of setting up cellbased assays. It is important to mix cells well by agitating the reservoir and pipetting the solution up and down before dispensing to each row. Incubating the plate at ambient temperature for 15 min prior to placing in the incubator is also important to reduce edge effects in the assay. These techniques ensure an even distribution of cells throughout the wells. 8. All test agents will be diluted in medium containing 5 μM caspase-3/7 reagent, so make up a volume that will accommodate all treatment conditions and dilutions. 9. Making up the starting concentration of compounds in 1 mL media + caspase-3/7 reagent is ideal for replicates of four. We recommend using replicates of at least three for each compound at each concentration in the 96-well assay. 10. Air bubbles can cause anomalies in images ruining the collection and analysis of data. Be sure to avoid making bubbles in pipetting technique. If bubbles are present, remove the inner straw of a plastic ethanol squeeze bottle filled with 70 % ethanol and gently squeeze air from the bottle over wells containing bubbles. 11. Condensation will form on the plate when transferred from ambient temperature to the 37 °C, humidified environment. Scanning right away will cause the images to appear out of focus. Allow the plate to equilibrate in the incubator for 15–30 min to avoid condensation. The effects of very toxic compounds, such as staurosporine, can be seen immediately, so it is imperative to begin imaging less than 30 min–1 h after addition to cells. 12. The first time the apoptosis experiment is performed, a new image collection and processing definition need to be created.

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For future experiments, the user can (and should) apply the processing definition to an experiment at the time of vessel scheduling. For more information on fluorescence processing in the IncuCyte ZOOM, consult IncuCyte ZOOM processing technical notes. 13. The endpoint for this experiment will vary depending on the cell type and compound being tested. At the end of the experiment the user may choose to normalize the number of caspase-3/ 7-expressing cells to total cell number with a membranepermeable dye. We suggest using a final concentration of 1 μM Vybrant(R) DyeCycle(TM) Green (Life Technologies) per well. Make up a 3 μM Vybrant DyeCycle Green solution in PBS or complete media and add 50 μL directly to the 100 μL media in each well. 14. Maximum values or data-informed endpoints can also be used to calculate pharmacology. References 1. Hanahan D, Weinberg RA (2011) Hallmarks of cancer: the next generation. Cell 144:646–674 2. Vega-Avila E, Pugsley MK (2011) An overview of colorimetric assay methods used to assess survival or proliferation of mammalian cells. Proc West Pharmacol Soc 54:10–14

3. Wu X, Simone J, Hewgill D, Siegel R, Lipsky PE, He L (2006) Measurement of two caspase activities simultaneously in living cells by a novel dual FRET fluorescent indicator probe. Cytometry A 69:477–486

Chapter 5 Detection of End-Stage Apoptosis by ApopTag® TUNEL Technique Chandra Mohan, Kevin Long, Manpreet Mutneja, and Jun Ma Abstract DNA fragmentation, the end stage of apoptosis, is the measure of ultimate demise of the cell. A convenient method for examining apoptosis via DNA fragmentation is by the Terminal deoxynucleotidyl transferase (Tdt) dUTP Nick-End Labeling (TUNEL) assay where the DNA strand breaks are detected by enzymatically labeling the free 3′-OH termini with modified nucleotides. ApopTag® kits detect single-stranded and double-stranded breaks associated with apoptosis. This technique is also helpful to distinguish between apoptotic and necrotic cell death where the latter is associated with random DNA fragment lengths producing a DNA smear. Apoptotic cells stained positive with ApopTag® kits are easier to detect and their identification is more certain, as compared to the examination of simply histochemically stained tissues. In addition, quantitative results can be obtained using flow cytometry and apoptotic cells can be differentiated from necrotic cells with greater than tenfold sensitivity. Key words Apoptosis, DNA fragmentation, TUNEL assay, ApopTag®, Anti-digoxigenin conjugate

1

Introduction Apoptosis is a highly regulated normal process in development and morphogenesis; however, failure to regulate apoptosis can lead to several diseases, including autoimmune disorders, neurodegenerative diseases, and cancer [1]. Apoptosis occurs when cells have sufficient time to organize and participate in their own demise. Apoptosis is controlled by multiple signaling events and the cell cycle checkpoint controls are intricately linked to apoptotic cascades [2]. Cells undergoing apoptosis display a number of distinctive biochemical and morphological changes, including shrinkage, membrane blebbing, chromatin condensation, and extensive nuclear fragmentation followed by the formation of apoptotic bodies that undergo phagocytosis [3–5]. This carefully controlled process does not lead to any inflammation, unlike necrosis that leads to edema and disruption of the plasma membrane.

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_5, © Springer Science+Business Media New York 2015

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DNA fragmentation, the end stage of apoptosis, is the measure of ultimate demise of the cell [6, 7]. In a number of well-researched model systems, during late-stage apoptosis, large fragments of 50–300 kb are first produced by the action of endonucleases. These large DNA fragments are visible on pulsed-field electrophoresis gels [8–10]. The activation of Ca2+- and Mg2+-dependent endonucleases further shortens these fragments by cleaving the DNA at linker sites between nucleosomes [11]. The ultimate DNA fragments are multimers of about 180 bp nucleosomal units that appear as “DNA ladder” in standard agarose gel electrophoresis [10–14]. 1.1 ApopTag® Technology

A convenient method for examining apoptosis via DNA fragmentation is by the Terminal deoxynucleotidyl Transferase (Tdt) dUTP Nick-End Labeling (TUNEL) assay [13]. EMD Millipore’s ApopTag® technology is based on the TUNEL principle. Here the DNA strand breaks are detected by enzymatically labeling the free 3′-OH termini with modified nucleotides. ApopTag® kits detect single-stranded [15] and double-stranded breaks associated with apoptosis. Drug-induced DNA damage is not identified by the TUNEL assay unless it is coupled to the apoptotic response [16]. The ApopTag® technique is helpful to distinguish between apoptotic and necrotic cell death where the latter is associated with random DNA fragment lengths producing a DNA smear. However, it is important to evaluate TUNEL staining results in conjunction with morphological criteria. Although DNA fragments can be identified on agarose gels, the single-cell sensitivity of ApopTag® histochemistry is considered to be a far superior technology. Apoptotic cells stained positive with ApopTag® kits are easier to detect and their identification is more certain, as compared to the examination of tissues that are histochemically stained. Another feature of ApopTag® is that quantitative results can be obtained using flow cytometry, since end-labeling methodology detects apoptotic cells with a >10-fold higher sensitivity than necrotic cells [17, 18]. The protocol described here is for the ApopTag® Fluorescein in situ Apoptosis Detection Kit (Cat. No. S7110). However, the following alternative fluorogenic and chromogenic kits are also available from EMD Millipore: ApopTag® Peroxidase in situ Apoptosis Detection Kit (Cat. No. S7100). ApopTag® Plus Peroxidase in situ Apoptosis Detection Kit (Cat. No. S7101). ApopTag® Plus Fluorescein in situ Apoptosis Detection Kit (Cat. No. S7111). ApopTag® Fluorescein Direct in situ Apoptosis Detection Kit (Cat. No. S7160). ApopTag® Red in situ Apoptosis Detection Kit (Cat. No. S7165).

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ApopTag® Peroxidase in situ Oligo Ligation (ISOL) Apoptosis Detection Kit (Cat. No. S7200). ApopTag® Kits are designed for labeling the free 3′-OH DNA termini in situ with chemically labeled and unlabeled nucleotides. The nucleotides are added enzymatically by terminal deoxynucleotidyl transferase (TdT) [13, 19] that catalyzes a template-independent addition of nucleotide triphosphates (NTPs) to the 3′-OH ends of double-stranded or single-stranded DNA. The incorporated nucleotides form an oligomer composed of digoxigenin nucleotide and unlabeled nucleotide in a random sequence. The ratio of labeled to unlabeled nucleotide in ApopTag® kits is optimized to promote anti-digoxigenin antibody binding, or to minimize fluorescein self-quenching. DNA fragments labeled with the digoxigenin-nucleotide are allowed to bind a fluorescein-conjugated antibody that is specific for digoxigenin (Fig. 1). This provides a sensitive detection in immunohistochemistry or immunocytochemistry and is not affected by experimental variations due to the substrate or the development step.

1.2 Principle of the Procedure

In Direct

Direct

End result of apoptosis: Nucleosome sized DNA fragments

End result of apostosis: Nucleosome sized DNA fragments

ApopTag® Step 1: Tail with digoxigenin-dNTP

ApopTag® Step 1: Tail with fluorescein-nucleotide

ApopTag® Step 2: Blind antibody conjugate

ApopTag® Step 2: Analyze by flow cytometry ApopTag® Step 3: Stain with substrate and view by microscopy (perox.) as depicted. Alternatively, analyze by microscopy or flow cytometry (fluor.)

Fig. 1 ApopTag® Technology

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This mixed molecular biological-immunohistochemical system allows for sensitive and specific staining of very high concentrations of 3′-OH ends that are localized in apoptotic bodies. The ApopTag® system differs from other in situ labeling techniques [13, 20, 21] in which avidin binding to cellular biotin can be a source of error. The digoxigenin/anti-digoxigenin system is equally sensitive, but immunochemically similar ligands for binding of the anti-digoxigenin antibody are generally insignificant in animal tissues, ensuring low background staining. An affinitypurified sheep polyclonal antibody is the specific anti-digoxigenin antibody used in ApopTag® kits and exhibits less than 1 % crossreactivity with the major vertebrate steroids. In addition, the Fc portion of this anti-digoxigenin antibody has been removed by proteolytic digestion to eliminate any nonspecific adsorption to cellular Fc receptors.

2 2.1

Materials Equipment

1. Silanized glass slides (to avoid detachment of tissue sections during processing). 2. Glass cover slips (for oil immersion objective, use 22 × 50 mm). 3. Adjustable micropipettors. 4. Glass or plastic Coplin jars. 5. Microcentrifuge tubes. 6. 15 mL screw-cap polypropylene centrifuge tubes. 7. Forceps for handling plastic cover slips (optional). 8. Humidified chamber. 9. Covered water bath, or incubator at 37 °C. 10. Light microscope equipped with bright-field optics (40× and 10× objectives) and also equipped for fluorescence. 11. Flow cytometer equipped with a 15 mW, 488 nm argon excitation laser, with appropriate filters.

2.2

Reagents

1. Deionized water. 2. Xylene (keep xylene used for de-waxing separate from that used for last dehydration step). 3. Ethanol—absolute, 95 %, 70 %, diluted in dH2O. 4. Ethanol:acetic acid, 2:1 (v:v) (for tissue cryosection or cell protocols). 5. Slide mounting medium (i.e., antifade). 6. 0.5–1.0 μg/mL Propidium iodide in slide mounting medium. 7. 0.5–1.0 μg/mL DAPI (4′-6′ diamino-2-phenylindole) in slide mounting medium.

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8. 10 % Neutral buffered formalin: 10 mL of formalin + 90 mL of phosphate-buffered saline (PBS), pH 7.4. This is the preferred fixative before paraffin embedding (see Note 1). 9. 1 % Paraformaldehyde: 1 mL of methanol-free paraformaldehyde + 15 mL of PBS, pH 7.4. 10. 10× PBS, pH 7.4: Dissolve 55 g of Na2HPO4, 13.5 g of NaH2PO4, and 117 g of NaCl in 800 mL of deionized water and adjust the pH to 7.4 using dilute NaOH or HCl. Adjust the volume to 1 L with deionized water. 11. 1 % Bovine serum albumin prepared in PBS, pH 7.4. 12. 10 mM Citrate buffer, pH 6.0. 13. DN buffer: 30 mM Tris base, pH 7.2, 4 mM MgCl2, 0.1 mM DTT. Adjust pH to 7.2 using concentrated HCl. 14. TRITON® X-100: 10 % (w:v) stock solution. 15. TRITON® X-100 (0.1 % (w:v)) in PBS (for bicolor or triplelabeling protocols): This can be prepared in advance and stored at 4 °C for up to 1 month. 16. Proteinase K (for paraffin-embedded tissue protocol): Dilute the 200 μg/mL stock solution to 20 μg/mL in PBS, pH 7.4 just prior to use. 17. Working strength TdT enzyme: Dilute in reaction buffer just prior to use as follows: 33 μL of TdT + 77 μL of reaction buffer (provided in the kit from EMD Millipore). Mix well by vortexing. This amount is sufficient to treat two 5 cm2 tissue specimens. 18. Working strength stop/wash buffer: Provided in the kit from EMD Millipore. Prepare working solution by diluting 1 mL of buffer with 34 mL water. It can be stored at 4 °C for up to 1 year. 19. Working strength anti-digoxigenin-fluorescein antibody solution: Dilute 62 μL of anti-digoxigenin conjugate with 68 μL of blocking solution (provided in the kit by EMD Millipore) just prior to use. This amount is sufficient to treat two 5 cm2 tissue specimens. 20. Propidium iodide staining solution: Dissolve 50 μg propidium iodide and 10 mg RNase (700 Kunitz units) in 10 mL of PBS, pH 7.4. Prepare fresh each time and keep on ice until use. Precautions 1. The following kit components contain potassium cacodylate (dimethylarsinic acid) as a buffer: Equilibration buffer, reaction buffer, and TdT enzyme. These components are harmful if swallowed; avoid contact with skin and eyes and wash areas of contact immediately. Store kit components as indicated in Table 1.

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Table 1 ApopTag® kit components Component

Volume

Recommended storage

Equilibration buffer

3 mL

−15 to −25 °C

Reaction buffer

2 mL

−15 to −25 °C

TdT enzyme

0.64 mL

−15 to −25 °C

Stop/wash buffer

20 mL

−15 to −25 °C

Blocking solution

2.6 mL

−15 to −25 °C

Anti-digoxigenin-fluorescein

2.1 mL

2 to 8 °C

Plastic cover slips

100

Room temperature

2. Antibody conjugates and blocking solution contain 0.08 % sodium azide as a preservative. 3. TdT enzyme contains glycerol and will not freeze at −20 °C. For maximum shelf life, do not warm this reagent to room temperature before dispensing.

3

Methods The methods described below are immunohistochemistry and flow cytometric techniques to study end-stage apoptosis in tissue sections and cells, respectively. The TUNEL technique is extensively used for the detection and quantification of apoptosis in histological tissue sections. However, for best interpretation of data it is recommended to employ at least two different techniques to assess apoptotic death, for example, caspase-3 assay and DNA fragmentation assay.

3.1 Immunohistochemistry Method

While using immunohistochemistry techniques it is best to use a positive control and a negative control to avoid any misinterpretation of data. One of the following can be used as suitable positive control. In the normal female rodent mammary glands, extensive apoptosis occurs 3–5 days after weaning of rat pups. Sections of this tissue mounted on slides are commercially available. Typically, 1–2 % of the total number of cells on the slide is apoptotic. For biological positive controls, programmed cell death can be induced in young adult rat thymic lymphocytes by dexamethasone. In normal rodent testis, apoptotic spermatogonia spontaneously occur in the seminiferous tubules.

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A positive control sample can be prepared from any tissue sample by treating with DNase I as follows. 1. Pretreat section with DN buffer at room temperature for 5 min. 2. Dissolve DNase I in DN buffer to a final concentration of 1.0– 0.1 μg/mL (specific activity 10,000–1,000 U/mL). 3. Apply DNase I solution and incubate for 10 min at room temperature. 4. Rinse with five changes of dH2O for 3 min each change. As the consistency and prior processing of tissues will differ, testing a range of conditions including Proteinase K digestion is highly recommended. A negative control or sham staining can be performed without active TdT, but including Proteinase K digestion to control for nonspecific incorporation of nucleotides or for nonspecific binding of enzyme conjugate. Water or equilibration buffer may be substituted for the volume of TdT enzyme. Inactive working strength TdT can be prepared by adding to the regular TdT mixture, a 5 % (v:v) dilution from the bottle of stop/wash buffer concentrate, to chelate the divalent cationic enzyme cofactor. 3.2

Sample Fixation

Using a cross-linking fixative is advantageous because it helps to tether the small chromatin fragments to the tissue, preventing their extraction during the processing steps. The preferred fixative for embedding tissue in paraffin for ApopTag® analysis is standard 10 % (v:v) neutral buffered formalin. Formalin-fixed tissue can be embedded in paraffin or in plastic resin [22]. Tissue processing in paraffin increases the number of apparent apoptotic events and reduces the background, in comparison to cryosections of the same tissue. Pretreatment of paraffin-embedded tissue sections is required, after rehydration, to improve the exposure of DNA by digesting DNA-binding proteins. The tissue type and the fixation time used can affect the strength of protease pretreatment needed. 1. Place the slide in 10 mM citrate buffer, pH 3.0–6.0, in a Coplin jar, and gently boil for three to five cycles of 3 min each in a microwave oven [23, 24]. 2. Refill with fresh buffer between cycles, but do not let the sample dry out. A pressure cooker or an autoclave may also be used instead of a microwave. 3. If a detergent pretreatment method is preferred then treat with 0.5 % TRITON® X-100 for 10 min [25]. Do not allow the specimen to dry by evaporation when changing solutions. 4. Remove the slides from the final wash, tap off excess water, then blot or aspirate around the section, and promptly apply

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the next reagent. If there are multiple samples to be processed, slides can be treated at fixed time intervals and immediately placed in a humid chamber. Incubations can then be terminated at similar intervals to maintain a constant incubation time. 3.3 Fluorescent Staining of ParaffinEmbedded Tissue

1. Deparaffinize tissue section (in a Coplin jar): Wash the specimen in three times in xylene for 5 min each followed by two washes in absolute ethanol for 5 min each, one wash in 95 % ethanol, and one in 70 % ethanol for 3 min each. Give a final wash in PBS for 5 min. 2. Pretreat tissue: Treat the specimen with freshly diluted Proteinase K (20 μg/mL) for 15 min at room temperature in a Coplin jar or directly on the slide. Wash the specimen in two changes of PBS in a Coplin jar for 2 min each wash. 3. Apply equilibration buffer: Gently tap off any excess liquid and carefully blot or aspirate around the section. Immediately apply 75 μL/5 cm2 equilibration buffer directly on the specimen. Incubate for at least 10 s at room temperature. 4. Apply working strength TdT enzyme: Gently tap off excess liquid and carefully blot or aspirate around the section. Immediately pipette onto the section 55 μL/5 cm2 of working strength TdT enzyme. Incubate in a humidified chamber at 37 °C for 1 h (see Note 2). 5. Apply stop/wash buffer: Place the specimen in a Coplin jar containing working strength stop/wash buffer. Agitate for 15 s and incubate for 10 min at room temperature. During this incubation, bring the vial of anti-digoxigenin conjugate to room temperature while avoiding any exposure to light. Dilute as suggested in item 19 of Subheading 2.2 for the desired number of specimens. 6. Apply working strength anti-digoxigenin conjugate: Wash the specimen three times in PBS, pH 7.4 for 1 min each. Gently tap off excess liquid and carefully blot or aspirate around section. Apply the working strength anti-digoxigenin conjugate to the slide; use about 65 μL/5 cm2 of surface covered. Incubate in a humidified chamber for 30 min at room temperature while avoiding exposure to light. 7. Wash in PBS, pH 7.4: Wash the specimen in four changes of PBS, pH 7.4 in a Coplin jar for 2 min each wash at room temperature. 8. Counterstain and mount after fluorescein staining: Apply a mounting medium containing 0.5–1.0 μg/mL of propidium iodide or DAPI. Use 15 μL for a 22 × 50 mm cover slip with an oil immersion objective. Mount under a glass cover slip. If storage is required, apply rubber cement to edges of the cover slip (see Notes 3 and 4). Store at −20 °C in the dark.

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9. View fluorescein and counterstain: View by fluorescence microscopy using appropriate excitation and emission filters (see Notes 5 and 6). 3.4 Fluorescent Staining of Tissue Cryosections or Cells

Apoptosis in adherent cell cultures can involve detachment from the substrate, and supernatants should be tested, if possible, by using cytospin processing. 1. Fix specimen according to type: Tissue cryosections or adherent cultured cells: Fix in 1 % paraformaldehyde in PBS, pH 7.4 in a Coplin jar (or cell culture vessel) preferably for 10 min at room temperature or for up to 15 h at 4 °C. Drain off excess liquid. Wash in two changes of PBS for 5 min each wash. Postfix in precooled ethanol:acetic acid (2:1) for 5 min at −20 °C in a Coplin jar. Drain, but do not allow to dry (this solvent permeabilizes cells). Wash in two changes of PBS for 5 min each wash. Skip to step 2. Cell suspensions for microscopy: Fix cells at a density of approximately 5 × 106 cells/mL in freshly diluted 1 % paraformaldehyde in PBS, pH 7.4 for 10 min at room temperature. Dry 50–100 μL of cell suspension on a microscope slide (optionally, cytospin cells). As primary cell isolates may be less easily permeabilized than cultured cells, the use of an ethanol:acetic acid postfix step is recommended. Wash in two changes of PBS for 5 min each wash. Proceed to step 2. 2. Apply equilibration buffer: Gently tap off excess liquid and carefully blot or aspirate around the section. Immediately apply 75 μL/5 cm2 of equilibration buffer directly on the specimen. Incubate for at least 10 s at room temperature. 3. Apply working strength TdT enzyme: Gently tap off excess liquid and carefully blot or aspirate around the section. Immediately pipette onto the section 55 μL/5 cm2 of working strength TdT enzyme. Incubate in a humidified chamber at 37 °C for 1 h. 4. Apply stop/wash buffer: Put the specimen in a Coplin jar containing working strength stop/wash buffer and agitate for 15 s. Incubate for 10 min at room temperature. Remove an aliquot of anti-digoxigenin conjugate from the stock vial sufficient to process the desired number of specimens. Warm the aliquot to room temperature while avoiding exposure to light. 5. Apply working strength anti-digoxigenin conjugate: Wash the specimen in three changes of PBS, pH 7.4 for 1 min each wash. Gently tap off excess liquid and carefully blot or aspirate around the section. Apply working strength anti-digoxigenin conjugate to the slide; use 65 μL/5 cm2 of specimen surface area. Incubate in a humidified chamber for 30 min at room temperature Avoid exposure to light.

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Fig. 2 Detection of apoptotic cells with ApopTag® Fluorescein Direct kit in mouse embryo forelimb bud fixed with 10 % neutral-buffered formalin and paraffin embedded

6. Wash in PBS: Wash the specimen in four changes of PBS in a Coplin jar for 2 min per wash at room temperature. 7. Counterstain and mount after fluorescein staining: Apply a mounting medium containing 0.5–1.0 μg/mL of propidium iodide or DAPI. Use 15 μL for a 22 × 50 mm cover slip with an oil immersion objective. Mount under a glass cover slip. If storage is required, apply rubber cement to edges of the cover slip. Store at −20 °C in the dark. 8. View fluorescein or rhodamine and counterstain: View by fluorescence microscopy using standard fluorescein or rhodamine excitation and emission filters. A typical result is shown in Fig. 2. 3.5 Flow Cytometry Method

Prior to flow cytometric assay prepare the following solutions: 1. Working strength TdT enzyme: Dilute 33 μL of stock enzyme with 77 μL of reaction buffer (provided in the kit from EMD Millipore) just prior to use. Mix well by vortexing. 2. Working strength stop/wash buffer: Dilute 1 mL of stock buffer with 34 mL of deionized water. It can be stored at 4 °C for up to 1 year. 3. Working strength anti-digoxigenin-fluorescein antibody solution: Dilute 49 μL of stock anti-digoxigenin conjugate with 56 μL of blocking solution (provided in the kit from EMD Millipore). Avoid exposure to light and keep on ice.

3.6 Controls for Flow Cytometry

Three types of control samples are recommended for flow cytometry to aid in setting up electronic compensation and quadrant statistics. For bicolor experiments, these are (1) cells stained with ApopTag® Fluorescein only, (2) cells stained with propidium iodide only, and (3) unstained cells.

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Preparation

Positive control cells can be prepared by fixing a cell suspension in 1 % paraformaldehyde in PBS, pH 7.4 for 30 min on ice. Positive control samples can be prepared by inducing cells in suspension cultures, which should contain a mixture of viable, apoptotic, and necrotic cells. Some examples are (a) Jurkat cells treated with 300 ng/mL of anti-Fas monoclonal antibody CH-11 for 5 h, (b) U-937 cells cultured with 2–4 ng/mL of TNFα for 2–3 h or with 4 μg/mL of camptothecin for 4 h, and (c) murine thymocytes left in culture media for 1–3 h.

3.8 Fluorescent Staining of Cell Suspension

In this protocol determination of cell number is important. The signal may be decreased if greater than 4 × 106 cells are used. Avoid moving cells between tubes: use of a single 15 mL screw-cap tube for all steps is recommended to prevent cell loss. Also, centrifugation steps should be performed at 400 × g for 5 min. Sperm cells should be centrifuged at 1,200 × g for 10 min.

3.7

1. Induce cells and sample at time points according to protocol. Count cells. 2. Fix cells: Resuspend 1–2 × 106 cells in 0.5 mL of PBS and with a Pasteur pipette, add the suspension into 5 mL of 1 % paraformaldehyde in PBS, pH 7.4, on ice. Fix for 15 min on ice and spin down the cells. Resuspend in 10 mL of ice-cold PBS and respin the cells. Resuspend in 70 % ice-cold ethanol. Keep at −20 °C for at least 1–2 h (see Note 7). 3. Assay setup: Prepare an ice bath for holding working strength TdT enzyme. Pre-warm an incubator to 37 °C. Prepare sufficient amount of working strength TdT enzyme. Prepare working strength stop/wash buffer. Prepare 0.1 % (w:v) TRITON® X-100 in PBS. Prepare propidium iodide counterstain solution. Prepare working strength anti-digoxigenin-fluorescein conjugate. Pretreat assay tubes with 5 % BSA in PBS for 1 min, and then drain well. 4. Wash fixed cells: Spin down 1–2 × 106 fixed cells per sample. Add 1 mL PBS and vortex gently. Spin down the cells and discard the supernatant. Add 1 mL of PBS, vortex, and spin. Discard the supernatant. 5. Apply equilibration buffer: Resuspend cells in 75 μL/5 cm2 of equilibration buffer. 6. Apply working strength TdT enzyme: Spin down the cells. Remove the supernatant. Resuspend the cells in 50 μL of TdT enzyme and incubate in a water bath for 30 min at 37 °C. At 15 min of incubation, resuspend cells that have settled to the bottom of the tube. 7. Stop/wash: Add 1.0 mL of working strength stop/wash buffer directly to the cell suspension. Spin down the cells and remove the supernatant. Resuspend cells in 1 mL of working strength stop/wash buffer.

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8. Apply working strength anti-digoxigenin-fluorescein: Spin down the cells and remove the supernatant. Resuspend in 100 μL of working strength anti-digoxigenin-fluorescein conjugate. Incubate for 30 min at room temperature. Avoid exposure to light. At 15 min of incubation, resuspend cells that have settled to the bottom of the tube. 9. PBS wash: Add 1.0 mL of 0.1 % TRITON® X-100 in PBS directly to the cell suspension. Spin down the cells and remove the supernatant. Repeat PBS wash and spin down the cells. 10. DNA staining: Add 1.0 mL of propidium iodide staining solution. Incubate for 15 min at room temperature; avoid exposure to light (see Notes 6 and 8). 11. Collect data (for example: using a Becton Dickinson FACScan flow cytometer equipped with a 15 mW argon ion laser). Measure green fluorescence of fluorescein and red fluorescence of propidium iodide. Generate a log FL1 vs. linear FL2 plot.

4

Notes 1. Commercial formalin solution contains about 37 % (w:v) formaldehyde with 10–15 % methanol added as a stabilizer. A standard 1:10 (v:v) dilution of formalin in buffered solution, conventionally known as “10 % neutral buffered formalin,” actually contains about 3.7 % formaldehyde (w:v). 2. Optional stopping points: There are several optional stopping points for temporary storage during sample processing. In the microscopy protocols: Slides may be left in equilibrium buffer or water for up to 60 min at 4 °C to room temperature. After incubating in working strength TDT enzyme, slides can be washed for 5 min in stop/wash buffer, then immersed in 70 % ethanol in a Coplin jar, and stored at −20 °C for at least 3 days. Before continuing with the protocols, samples should be washed with three changes of PBS for 2 min per change. 3. Use of plastic cover slips: Plastic cover slips can be used to assure that a constant volume of solution is applied per unit of specimen area. However, their handling time slows down the protocol. Plastic cover slips may be trimmed to any desired size and shape. The kit’s yield of specimens will be reduced if cover slips are larger than standard. 4. Apply plastic cover slips to microscope slides so as to minimize trapped air bubbles, which may cause variable enzyme reaction or detection. Place the slide across the pipettes, faceup and level, inside the humidified chamber. The slide edges should not touch anything so as to prevent drainage of the reagent.

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5. For (immuno) fluorescence microscopy: Both the FITC signal and the propidium iodide (PI) counterstain can be viewed with a “long-pass” filter for FITC (ex. 490 nm and em. 520 nm). This filter allows sufficient PI signal to “bleed through.” A “dualpass” filter, designed for viewing both FITC and PI, would allow more red light through, possibly competing with and decreasing the FITC signal. As PI binds reversibly to DNA, a PI signal can be modulated up or down by washing the sample and reapplying PI at another concentration. Photobleaching will cause the signal to fade in proportion to the time and intensity of exposure to excitation light. 6. Propidium iodide or DAPI staining intensity, as visualized by microscopy, is affected by variations in the tissue type, the fixation method (type, concentration, freshness, and time), tissue pre-treatments (Proteinase K or other), stain concentration, the light filter used, and photobleaching during imaging. The optimal counterstain concentration will result in fluorescence intensity nearly equal to that of the primary stain. In addition, the fluorescence signal per cell may be less intense when more concentrated samples are tested by flow cytometry. 7. In the flow cytometry protocols: After placing the cells in 70 % ethanol, they can be stored at −20 °C for at least 3 months. After PI is added, the tube containing the cells can be wrapped in foil and stored at 4 °C for 2–3 days. 8. For flow cytometry: In the bicolor protocol, measure red fluorescence of PI at >620 nm using linear amplification. In both flow cytometry protocols, measure FITC fluorescence as a green signal (530 nm peak fluorescence) by the FL1 detector through a band-pass filter (530 ± 15 nm) using logarithmic amplification. References 1. Elmore S (2007) Apoptosis: a review of programmed cell death. Toxicol Pathol 35: 495–516 2. Mohan C (2010) Apoptosis: receptor and mitochondrial gateways to cell death. In: Signal transduction—a short overview of its role in health and disease, 2nd edn. EMD, San Diego, CA, pp 67–71 3. Darzynkiewicz Z, Juan G, Li X, Gorczyca W, Murakami T, Traganos F (1997) Cytometry in cell necrobiology: analysis of apoptosis and accidental cell death (necrosis). Cytometry 27:1–20 4. Kerr JFR, Harmon BV (1991) Definition and incidence of apoptosis: an historical perspective, chapter 1. In: Tomei LD, Cope FO (eds) Apoptosis: molecular basis of cell death. Cold

5.

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Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp 5–29 Majno G, Joris I (1995) Apoptosis, oncosis and necrosis, an overview of cell death. Am J Pathol 146:3–15 Migheli AM, Attanasio A, Schiffer D (1995) Ultrastructural detection of DNA strand breaks in apoptotic neural cells by in situ end labeling techniques. J Pathol 176:27–35 Matassov D, Kagan T, Leblanc J, Sikorska M, Zakeri Z (2004) Measurement of apoptosis by DNA fragmentation. Methods Mol Biol 282: 1–17 Brown DG, Sun XM, Cohen GM (1993) Dexamethasone-induced apoptosis involves cleavage of DNA to large fragments prior to

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Chandra Mohan et al. internucleosomal fragmentation. J Biol Chem 268:3037–3039 Walker PR, Kokileva L, LeBlanc J, Sikorska M (1993) Detection of the initial stages of DNA fragmentation in apoptosis. Biotechniques 15: 1032–1047 Walker PR, Weaver VM, Lach B, LeBlanc J, Sikorska M (1994) Endonuclease activities associated with high molecular weight and internucleosomal DNA fragmentation in apoptosis. Exp Cell Res 213:100–106 Arends MJ, Morris RG, Wyllie AH (1990) Apoptosis: the role of the endonuclease. Am J Pathol 136:593–608 Bursch W, Paffe S, Putz B, Barthel G, SchulteHermann R (1990) Biochemistry of cell death by apoptosis. Biochem Cell Biol 68:1071–1074 Gavrieli Y, Sherman Y, Ben-Sasson SA (1992) Identification of programmed cell death in situ via specific labeling of nuclear DNA fragmentation. J Cell Biol 119:493–501 Staley K, Blaschke AH, Chun J (1997) Apoptotic DNA fragmentation is detected by a semi-quantitative ligation-mediated PCR of blunt DNA ends. Cell Death Differ 4:66–75 McGahon A, Bissonnette R, Schmitt M, Cotter KM, Green DR, Cotter TG (1994) Bcr-Abl maintains resistance of chronic myelogenous leukemia cells to apoptotic cell death. Blood 83:1179–1187 Chapman RS, Chresta CM, Herberg AA, Beere HM, Heer S, Whetton AD, Hickman JA, Dive C (1995) Further characterization of the in situ terminal deoxynucleotidyl transferase (TdT) assay for the flow cytometric analysis of apoptosis in drug resistant and drug sensitive leukemia cells. Cytometry 20:245–256 Gold R (1994) Differentiation between cellular apoptosis and necrosis by the combined use of in situ tailing and nick translation techniques. Lab Invest 71:219–225 Gorczyca W, Gong J, Darzynkiewicz Z (1993) Detection of DNA strand breaks in individual

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apoptotic cells by the in situ terminal deoxynucleotidyl transferase and nick translation assays. Cancer Res 53:1–7 Schmitz GG, Walter T, Seibl R, Kessler C (1991) Non-radioactive labeling of oligonucleotides in vitro with the hapten digoxigenin by tailing with terminal transferase. Anal Biochem 192:222–231 Gorczyca W, Bruno S, Darzynkiewicz RJ, Gong J, Darzynkiewicz Z (1992) DNA strand breaks occurring during apoptosis: their early in situ detection by terminal deoxynucleotidyl transferase and nick translation assays and prevention by serine protease inhibitors. Int J Oncol 1:639–648 Wijsman JH, Jonker RR, Keijzer R, Van De Velde CJH, Cornelisse CJ, Van Dierendonck JH (1993) A new method to detect apoptosis in paraffin sections: in situ end-labeling of fragmented DNA. J Histochem Cytochem 41: 7–12 Mundle S, Iftikhar A, Shetty V, Dameron S, Wright Quinones V, Marcus B, Loew J, Gregory S, Raza A (1994) Novel in situ double labeling for simultaneous detection of proliferation and apoptosis. J Histochem Cytochem 42:1533–1537 Labat Moleur F, Guillermet C, Lorimier P, Robert C, Lantuejoul S, Brambilla E, Negoescu A (1998) TUNEL apoptotic cell detection in tissue sections: critical evaluation and improvement. J Histochem Cytochem 46:327–334 Strater J, Gunthert AR, Bruderlein S, Moller P (1995) Microwave irradiation of paraffinembedded tissue sensitizes the TUNEL method for in situ detection of apoptotic cells. Histochem Cell Biol 103:157–160 Tornusciolo DR, Schmidt RE, Roth KA (1995) Simultaneous detection of TdT mediated dUTP-biotin nick end labelling (TUNEL)— positive cells and multiple immunohistochemical markers in single tissue sections. Biotechniques 19:800–805

Chapter 6 Detection and Quantification of Apoptosis in Primary Cells Using Taqman® Protein Assay Christina Pfister, Heike Pfrommer, Marcos S. Tatagiba, and Florian Roser Abstract There are several methods to detect apoptosis using cleaved caspase-3 and each harbors its own advantages and disadvantages. When primary cell cultures are used, the disadvantages of the standard methods can make apoptosis detection difficult due to their slow growth rate and replicative senescence, thereby limiting the available cell number and experiment time span. In this chapter, we describe apoptosis detection and quantification using an innovative method named TaqMan® protein assay. TaqMan® protein assay uses antibodies and proximity ligation for quantitative real-time PCR. Biotinylated antibodies are labeled with oligonucleotides. When the labeled antibodies bind in close proximity, the oligonucleotides are connected using DNA ligase. The ligation product is amplified and detected using Taqman® based Real-Time PCR. Using this technique, we can not only detect apoptosis with a 1,000-fold higher sensitivity than western blot, but we can also exactly quantify cleaved caspase-3 expression. Thereby apoptosis can be determined and quantified in a fast reliable manner. Key words Apoptosis, Caspase-3, Taqman protein assays, Primary cells

1

Introduction Apoptosis can be reliably detected by assaying for cleaved (active) caspase-3, for which active caspase-3 antibodies are used in methods such as immunocytochemistry, enzyme-linked immunosorbent assay (ELISA), and western blot. These methods, however, have limitations especially when detecting apoptosis in primary cells. Immunocytochemistry and western blot are only semiquantitative as are cell-based ELISAs. In addition, western blot requires a high input quantity and has low sensitivity. Because primary cell cultures often display slow growth rate and undergo replicative senescence, the input quantity required for western blot is often not achieved. Although ELISAs require less sample input and show higher sensitivity than western blots, a matching pair of monoclonal antibodies is however needed for this assay.

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_6, © Springer Science+Business Media New York 2015

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Fig. 1 Schematic overview of active caspase-3 TPA. The workflow consists of four steps: (1) Assay preparation: 3′-Oligonucleotide is bound to polyclonal caspase-3 antibody using a biotin-streptavidin interaction → probe A. 5′-Oligonucleotide is bound to monoclonal active caspase-3 antibody the same way → probe B; (2) Binding step: Probe A and probe B bind to their specific epitopes in a cell lysate. (3) Ligation step: The assay probe oligonucleotides in close proximity hybridize to a short connecting oligonucleotide using DNA ligase. (4) Amplification step: The ligation product is amplified and detected using Taqman-based real-time PCR

Moreover the antibody consumption for ELISAs is high. Thus using these traditional methods to detect apoptosis in primary cell cultures is challenging. There are several benefits of using primary cell cultures. First, primary cells are morphologically and genetically similar to cells in vivo. Second, primary cells display a variety of cell subtypes in cultures. Thus, primary cell cultures remain the closest representation of in vivo conditions. Recently, a new method for protein detection and quantitation was developed. TaqMan® protein assay (TPA) combines antibody detection with quantitative real-time PCR using proximity ligation [1, 2] (see Fig. 1). A polyclonal biotinylated antibody is labeled with 3′- and 5′-oligonucleotides generating a probe pair (probe A + probe B). When this probe pair is mixed with a cell lysate the two probes bind on two different epitopes. When the two probes bind in close proximity the 3′- and 5′-oligonucleotides are connected using a connecting oligonucleotide and DNA ligase. This ligation product is amplified and detected using Taqman® based real-time PCR. To detect active caspase-3 using TPA a polyclonal caspase-3 antibody and a monoclonal active caspase-3 antibody are used. The combination of inactive caspase-3 antibody as probe A and active caspase-3 antibody as probe B forms a TPA specific only for active caspase-3. TPA displays a very high sensitivity with a detection limit of 400 fg/well for caspase-3 (probe A + B polyclonal caspase-3 antibody) and 2 pg/well for active caspase-3 (probe A (polyclonal caspase-3 antibody) + probe B (monoclonal active caspase-3 antibody)), respectively [3]. Thus TPA has a 1,000-fold higher sensitivity than western blot. The following are advantages of using TPA: (1) only a small sample input is needed (typically about 50 ng), which is an advantage when slow-growing primary cell cultures are used. In this protocol the initial amount is rather high for TPA, which is 5 μg. (2) TPA has very small antibody usage. Only 300 ng of antibody is needed

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for about eight 96-well plates. (3) TPA is a highly sensitive and quantitative protein detection method. (4) TPA allows a direct correlation between protein and mRNA expression due to the use of real-time PCR. Our group focuses on meningioma, which is the most common intracranial tumor. Meningiomas originate from the arachnoidal cap cells of the meningeal cover of the spinal cord and brain and constitute approximately 13–26 % of all intracranial pathologies. There are a few stable meningioma cell lines, which are mostly malignant [4]. Thus, most meningioma cell culture research is conducted with primary meningioma cells. Due to their low proliferation rate, cell numbers in primary meningioma cultures are limited. Moreover, these cultures undergo senescence in early passages and cells alter their phenotype with ongoing culture [5]. Thus, the use of ELISA or western blot to detect and quantify cleaved caspase-3 in these cultures is restricted. The method outlined in this chapter describes the detection and quantification of active caspase-3 protein in primary meningioma cells. This method, however, can be used for the detection of other proteins, as long as there is a suitable antibody available. TPA is suitable to detect not only active protein forms, but also posttranslational modifications such as phosphorylation. The method outlined below is utilized for: 1. Lysis of pretreated primary cells. 2. Preparation and storage of the sample to be analyzed. 3. Biotinylation of antibodies. 4. Validation of biotinylated antibodies. 5. Labeling of biotinylated antibodies with oligonucleotides. 6. Apoptosis detection using TPA. 7. TPA analysis.

2 2.1

Materials Equipment

1. Real-time PCR system and analysis software. 2. Spectrometer. 3. Microcentrifuge. 4. 96-Well PCR plates. 5. PCR plate-sealing mats. 6. Filter tips. 7. Sterile 0.5, 1, and 2 ml tubes. 8. Sterile 0.5 ml microcentrifuge tubes (we use Protein LoBind tubes from Eppendorf, Hamburg, Germany). 9. Sterile 15 ml tubes.

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Reagents

2.2.1 Reagents for Cell Preparation and Cell Lysis

1. 12-, 24-, or 48-well plates. 2. D-PBS sterile. 3. Protein Quant Sample Lysis Kit (i.e., 4448536, Life Technologies, Grand Island, NY). 4. Protease inhibitor cocktail. 5. Phosphatase inhibitor cocktail.

2.2.2 Antibodies and Reagents for Antibody Labeling

1. Caspase-3 antibody Minneapolis, MN).

(i.e.,

AF-605-NA,

R&D

Systems,

2. Cleaved Caspase-3 antibody (i.e., 700182, Life Technologies, Grand Island, NY). 3. Biotin-XX microscale Protein Labeling Kit (i.e., B30010, Life Technologies, Grand Island, NY). 4. TaqMan® Protein Assays Open Kit (i.e., 4453745, Life Technologies, Grand Island, NY).

2.2.3 Reagents for Taqman Protein Assay

1. TaqMan® Protein Assays Core Reagents Kit with Master Mix (i.e., 4448591, Life Technologies, Grand Island, NY). 2. Molecular biology-grade nuclease-free water.

3

Methods

3.1 Sample Preparation

See Note 1 for details before you begin: 1. Seed primary cells (~105 cells/ml) in appropriate tissue culture plate in complete medium for 24–48 h. Cell number depends on primary cells and plate used (see Note 2). 2. Treat cells with preferred apoptosis inducer in complete medium for applicable period of time. 3. Discard the medium and carefully wash treated cells twice with ice-cold D-PBS. 4. Place the plate on ice. 5. Lyse cells with ice-cold Sample Lysis Buffer (Protein Quant Sample Lysis Kit). Sample Lysis Buffer should contain each 10 μl/ml protease and phosphatase inhibitor (see Note 3). 100 μl Sample Lysis Buffer is sufficient to lyse 250,000 cells. As cells are not counted with this method the volume of sample lysis buffer should be estimated based on seeded cell number and growth rate. 6. After addition of Sample Lysis Buffer carefully move the plate (on ice) to lyse the cells. After cells are detached carefully pipette cells and buffer up and down to lyse the cells completely (see Fig. 2). Avoid air bubbles and foaming.

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Fig. 2 Preparation of cell lysates. (a) Carefully pipette cells up and down to completely lyse cells. (b) Transfer lysed cells in a 1 ml tube

7. Pipette the lysed cells in a 1 ml tube and place the tube on ice for 10–15 min. 8. Clarify the lysates by centrifugation at 12,000 × g for 10 min at 4 °C. 9. Quantify protein concentration spectrometrically. 10. Divide cell lysates in 50 μl aliquots. 11. Store at −80 °C. 12. All samples need to have the same protein concentration for TPA experiments. The initial concentration in our lab is 12.5 mg/ml. The concentration is optimized for primary meningioma cells. Fresh cell lysates are diluted to 12.5 mg/ml using Lysate Dilution Buffer (TaqMan® Protein Assays Open Kit) and stored at −20 °C. 3.2 Biotinylation of Antibody Labeling

See Note 4 for details before you begin: 1. Dilute lyophilized caspase-3 antibody with 200 μl PBS. Antibody concentration is 0.5 mg/ml. 2. A 50 μl aliquot of the caspase-3 antibody is biotinylated using the Biotin-XX Microscale Protein Labeling Kit as per the manufacturer’s instructions. For 50 μl caspase-3 antibody, 5 μl sodium bicarbonate and 1.05 μl reactive biotin-XX solution are needed. 3. Also 50 μl cleaved caspase-3 antibody is biotinylated accordingly. The appropriate volume of reactive Biotin-XX solution for the cleaved caspase-3 antibody is 2 μl. 4. Biotinylated antibodies are stored in 0.5 ml Protein LoBind tubes at −20 °C. 5. Dilute 3 μl biotinylated antibodies with 47 μl Antibody Dilution Buffer (TaqMan® Protein Assays Open Kit) to 200 nM (30 μg/ml). 6. Store diluted antibodies at −20 °C.

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3.3 Forced Proximity Test

Validation of biotinylation of the antibodies is performed using the “forced proximity test.” 1. Mix 5 μl 200 nM 3′ Prox-Oligo (red cap, TaqMan® Protein Assays Open Kit) with 5 μl 200 nM 5′ Prox-Oligo (purple cap, TaqMan® Protein Assays Open Kit) in a 0.5 ml tube on ice. 2. Centrifuge tube briefly to spin the liquid to the bottom. 3. Label three 0.5 ml tubes: Tube 1: CASP3 Tube 2: active CASP3 Tube 3: NPC (negative protein control) 4. Tube 1: Mix on ice 2 μl Prox-Oligo with 2 μl 200 nM biotinylated caspase-3 antibody. 5. Tube 2: Mix on ice 2 μl Prox-Oligo with 2 μl 200 nM biotinylated active caspase-3 antibody. 6. Tube 3: Mix on ice 2 μl Prox-Oligo with 2 μl Antibody Dilution Buffer. 7. Centrifuge tubes briefly to collect the liquid at the bottom. 8. Incubate the tubes for 60 min at room temperature (RT). 9. Add 396 μl of Assay Probe Dilution Buffer (TaqMan® Protein Assays Open Kit) to each tube. 10. Incubate the three tubes for 30 min at RT. 11. Place the three tubes on ice. 12. Dilute 2 μl DNA Ligase (TaqMan® Protein Assays Core Reagents Kit with Master Mix) in 198 μl Ligase Dilution Buffer (TaqMan® Protein Assays Core Reagents Kit with Master Mix) in a 0.5 ml tube on ice. 13. Invert the tube twice and place the tube back on ice. 14. Mix 100 μl Ligation Reaction Buffer (TaqMan® Protein Assays Core Reagents Kit with Master Mix) with 1,815 μl nucleasefree water and 2 μl diluted DNA ligase (step 12) in a 2 ml tube. 15. Invert the tube twice and place the tube back on ice. 16. Place a 96-well reaction plate (or two eight-well PCR strips) on ice. 17. A total of 16 wells are needed. Choose eight wells in row 1 for caspase-3 and eight wells in row 2 for active caspase-3. 18. Add 96 μl of the ligation solution (step 14) to each well. 19. Add 4 μl of tube 1 (CASP3) to wells 1–4 and 4 μl of tube 3 (NPC) to wells 5–8 in row 1. 20. Add 4 μl of tube 2 (active CASP3) to wells 1–4 and 4 μl of tube 3 (NPC) to wells 5–8 in row 2. 21. Seal the reaction plate with a PCR plate-sealing mat.

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22. Centrifuge PCR plate briefly to spin the liquid to the bottom. 23. Incubate the sealed reaction plate using the following thermalcycling conditions: Ligation: 1 Cycle, 37 °C, 10 min Cooling: 1 Cycle, 4 °C, up to 10 min 24. Continue immediately. 25. Mix 20 μl thawed Universal PCR Assay (TaqMan® Protein Assays Core Reagents Kit with Master Mix) with 200 μl Fast Master Mix (TaqMan® Protein Assays Core Reagents Kit with Master Mix) in a 0.5 ml tube on ice. 26. Centrifuge tube briefly to spin the liquid to the bottom. Place the tube on ice. 27. Place a 96-well reaction plate (or two eight-well PCR strips) on ice. 28. Add 11 μl PCR mix (step 25) to each of the 16 wells. 29. Transfer 9 μl of the ligation product (step 23) to each well. 30. Seal the reaction plate with PCR plate-sealing mats. 31. Centrifuge tube briefly to spin the liquid to the bottom. 32. Incubate the sealed reaction plate using the following real-time PCR conditions (see Note 5): Holding stage: 1 Cycle, 95 °C, 20 s Cycling stage: 40 Cycles, 95 °C, 1 s, and 60 °C, 20 s 33. Save the results of the real-time PCR system software. For analysis see Subheading 3.6.1. 3.4 Antibody Labeling

3.4.1 Assay Probe A

When both biotinylated antibodies passed the “forced proximity test” they are labeled with Prox-Oligos. Antibodies labeled with 3′ Prox-Oligo are designated Assay Probe A. Antibodies labeled with 5′ Prox-Oligo are designated Assay Probe B. Every TPA consists of an Assay Probe A and an Assay Probe B. 1. Mix 5 μl 200 nM biotinylated caspase-3 antibody with 5 μl 3′ Prox-Oligo in a 0.5 ml Protein LoBind tube on ice. 2. Centrifuge tube briefly to collect the liquid at the bottom. 3. Incubate the tube for 60 min at RT. 4. Add 90 μl of Assay Probe Dilution Buffer (TaqMan® Protein Assays Open Kit) to the tube. 5. Mix gently. 6. Centrifuge the tube briefly to collect the liquid at the bottom. 7. Incubate the tube for 20 min at RT. 8. Store Assay Probe A at −20 °C.

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3.4.2 Assay Probe B

1. Mix 5 μl 200 nM biotinylated cleaved caspase-3 antibody with 5 μl 5′ Prox-Oligo in a 0.5 ml Protein LoBind tube on ice. 2. Centrifuge tube briefly to spin the liquid to the bottom. 3. Incubate the tube for 60 min at RT. 4. Add 90 μl of Assay Probe Dilution Buffer (TaqMan® Protein Assays Open Kit) to the tube. 5. Mix gently. 6. Centrifuge tube briefly to spin the liquid to the bottom. 7. Incubate the tube for 20 min at RT. 8. Store Assay Probe B at −20 °C.

3.5 Taqman Protein Assay for Apoptosis Detection

A control lysate (not treated with apoptotic agent) should always be used as control. This TPA is for relative quantification, not absolute quantification. Also every plate has to contain a “noprotein control” (NPC) to calculate ΔCT. Figure 3 shows the standard configuration of a 96-well plate used in our lab. This plate contains one control lysate with a five-point serial dilution in triplicates, five sample lysates also with a five-point serial dilution in triplicates, and an NPC with a double triplicate. See Note 6 for details before you begin: 1. Place a 96-well plate (plate 1) on ice to prepare the serial dilution. 2. Dilution sample 1: Dilute 4 μl 12.5 mg/ml cell lysate in 16 μl Lysate Dilution Buffer (TaqMan® Protein Assays Open Kit) in

Fig. 3 Example of a 96-well plate for TPA with input amount per well in μg

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Fig. 4 Pipetting scheme for TPA with input amount per well in μg. Plate 1 shows the dilution series for a control lysate and five samples. Plate 2 shows the pipetting scheme for TPA with a triplicate reaction for each sample. Therefore 2 μl diluted cell lysate from well A1 (left plate ) is transferred to wells A1–A3 (right plate ) and so on

well A1 (see Fig. 4 for 96-well plate 1). Initial input amount is 5 μg (see Note 7). 3. Place 12 μl Lysate Dilution Buffer in wells B1–E1. 4. Transfer 12 μl cell lysate from well A1 to well B1. 5. Pipette up and down several times to mix the sample. 6. Transfer 12 μl cell lysate from well B1 to well C1. 7. Pipette up and down several times to mix the sample. 8. Transfer 12 μl cell lysate from well C1 to well D1. 9. Pipette up and down several times to mix the sample. 10. Transfer 12 μl cell lysate from well D1 to well E1. 11. Pipette up and down several times to mix the sample. 12. Repeat steps 2–11 for every sample. 13. Seal plate 1 with a PCR plate-sealing mat. 14. Centrifuge plate 1 briefly to spin the liquid to the bottom. 15. Place plate 1 on ice. 16. Place a second 96-well plate (plate 2) on ice (see Note 8). 17. Place thawed Assay Probe Dilution Buffer (TaqMan® Protein Assays Open Kit), Assay Probe A, and Assay Probe B on ice. Do not vortex Assay Probes. 18. Mix gently 216 μl Assay Probe Dilution Buffer with 12 μl Assay Probe A and 12 μl Assay Probe B in a 0.5 ml tube. 19. Add 2 μl Assay Probe solution (step 17) to each well of plate 2. 20. Transfer 2 μl of the serial dilution of plate 1 to the well indicated in Fig. 4 in plate 2.

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21. Seal plate 2 with a PCR plate-sealing mat. 22. Centrifuge plate 2 briefly to spin the liquid to the bottom. 23. Incubate the sealed plate 2 using the following thermal-cycling conditions: Binding: 1 Cycle, 37 °C, 60 min Cooling: 1 Cycle, 4 °C, up to 10 min 24. Place plate 2 on ice and continue immediately. 25. Place thawed Ligase Dilution Buffer, Ligation Reaction Buffer, and DNA-Ligase (TaqMan® Protein Assays Core Reagents Kit with Master Mix) on ice. 26. Dilute 2 μl DNA ligase in 998 μl Ligase Dilution Buffer in a 1 ml tube on ice. 27. Invert the tube twice and place the tube back on ice. 28. Mix 600 μl Ligation Reaction Buffer with 10,908 μl nucleasefree water and 12 μl diluted DNA ligase (step 25) in a 15 ml tube. 29. Invert the tube twice and place the tube back on ice. 30. Place plate 2 (step 22) on ice. 31. Add 96 μl of the ligation solution (step 27) to each well. 32. Seal the reaction plate with a PCR plate-sealing mat. 33. Centrifuge plate 2 briefly to spin the liquid to the bottom. 34. Incubate the sealed plate 2 using the following thermal-cycling conditions: Ligation: 1 Cycle, 37 °C, 10 min Cooling: 1 Cycle, 4 °C, up to 10 min 35. Place plate 2 on ice and either continue immediately with the real-time PCR step (go to step 43) or perform a protease reaction (steps 36–43), if the real-time PCR step should be delayed (up to 3 days at 4 °C or up to 2 weeks at −20 °C). 36. Place thawed PBS and protease (TaqMan® Protein Assays Core Reagents Kit with Master Mix) on ice. 37. Dilute 4 μl protease in 396 μl PBS in a 0.5 ml tube on ice. 38. Pipette up and down two times to dilute protease. 39. Add 2 μl diluted protease to each well. 40. Seal plate 2 with a PCR plate-sealing mat. 41. Centrifuge plate 2 briefly to spin the liquid to the bottom. 42. Incubate the sealed plate 2 using the following thermal-cycling conditions: Terminate ligation: 1 Cycle, 37 °C, 10 min Inactivate protease: 1 Cycle, 95 °C, 5 min Cooling: 1 Cycle, 4 °C, hold

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43. Place plate 2 on ice. 44. Mix 100 μl thawed Universal PCR Assay (TaqMan® Protein Assays Core Reagents Kit with Master Mix) with 1,000 μl Fast Master Mix (TaqMan® Protein Assays Core Reagents Kit with Master Mix) in a 1 ml tube on ice. 45. Centrifuge tube briefly to spin the liquid to the bottom. Place the tube on ice. 46. Place a third 96-well reaction plate (plate 3) (step 34 or 42) on ice. 47. Add 11 μl PCR mix (step 43) to each well. 48. Transfer 9 μl of the ligation product (step 34) or the proteasetreated ligation product (step 42) to each well. 49. Seal plate 3 with PCR plate-sealing mats. 50. Centrifuge plate 3 briefly to spin the liquid to the bottom. 51. Incubate the sealed plate 3 using the following real-time PCR conditions (see Note 5): Holding stage: 1 Cycle, 95 °C, 20 s Cycling stage: 40 Cycles, 95 °C, 1 s, and 60 °C, 20 s 52. Save the results of the real-time PCR system software. For analysis see Subheading 3.6.2. 3.6 Software and Analysis 3.6.1 Analysis Forced Proximity Test

Use a real-time PCR system software and a spreadsheet program. 1. Set the threshold cycle (CT) to 0.2 and chose automatic baseline. 2. Export the results from the instrument software to a spreadsheet program. 3. Calculate the average CT values for each biotinylated antibody and each NPC. Each biotinylated antibody and the NPC should have four different values. Average CT value = sum of these four values divided by 4. 4. Calculate the ΔCT values for each biotinylated antibody: ΔCT value = average CT value (NPC) minus average CT value (biotinylated antibody). 5. A biotinylated antibody passed the “forced proximity test” if the ΔCT value is equal or higher than 8.5. If the ΔCT value is lower than 8.5 see Note 9.

3.6.2 Analysis Taqman Protein Assay

Data are analyzed using ProteinAssist™ 1.0 (Applied Biosystems, Foster City, CA), which uses the ΔCT2 method to calculate relative protein expression between untreated controls and treated sample. 1. Use the real-time PCR system software to set the threshold cycle (CT) to 0.2 and choose automatic baseline (see Fig. 5).

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Amplification Plot Control Lysate + Sample 1

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2. Open the ProteinAssist™ software and click “Create Study.” 3. Choose μg/well for “Input Quantity Unit.” 4. Click “Per Study” for reference use. 5. Click “Experiment Files.” 6. Click “Import” and choose the real-time PCR file (step 52). ProteinAssist™ software supports *.csv, *.eds, and *.txt files. 7. Import the file. 8. Select the imported file. 9. Right click a well and select “Edit Well(s).” 10. Assign every well of the control lysate to the task “Reference.” 11. Select for every NPC well the task “NPC.” 12. Select for every sample well the task “Unknown.” 13. Right click a well and select “Edit Well(s).” 14. Assign the correct input amount to every well, for example well A1 “Input Quantity” 5. 15. Click “Analyze” (green button at the top) (see Note 10).

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16. Click “Analysis.” 17. The automatic threshold is 2.0. The threshold depends on the expression level of the evaluated protein in the used cell lysates. The optimized threshold for active caspase-3 in primary meningioma cells is 1.0. 18. Choose the appropriate threshold. 19. Click “Analyze” (after every change in the graph the data has to be analyzed again). 20. The analysis of the PCR plate is shown below the graph. 21. Often the automated linear range has to be adjusted manually (see Fig. 6). 22. Select the control lysate (click box before the sample name). 23. The graph only displays the control lysate. 24. Change the linear range if needed (see Note 11). 25. Outliers are marked as an unfilled triangle. Omit outliers by right clicking the data point in the graph. Select “Omit.” 26. Repeat steps 22–25 for every sample. 27. Click “Fold Change.” 28. The graph displays the results as a bar diagram (see Fig. 7). Below the graph the results are detailed (see Note 12).

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Notes 1. There are two different methods to prepare the samples for TPA. The more conventional way is to detach the treated cells with Accutase®, pellet, and count. Then cells are lysed according to their cell number to a concentration of 2,500 cells/μl. As primary cells are often difficult to count exactly spectrometric determination of protein concentration can be used. When protein concentration is used cell lysis can be performed faster by directly adding the Sample Lysis Buffer to each well. The protocol only describes the direct cell lysis as a fast cell lysis is favorable to determine apoptosis. 2. The well number of the plates depends on the available cell number, experiment size, and experiment setting. One advantage of this method to determine apoptosis is the availability of the sample for further protein analysis. 3. Complete Sample Lysis Buffer should be prepared before washing the cells. Also the volume for all wells should be estimated before beginning cell lysis to avoid interruptions. 4. Subheadings 3.2 and 3.3 are only performed in the beginning as the amount of biotinylated antibody is sufficient for a large number of experiments. The antibody volume for biotinylated

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Fig. 6 Linear range graphs before manual adjustment (above ) and after manual adjustment of the linear range (below )

antibody can be increased when needed to avoid a new forced proximity test. Adjust reagent volumes accordingly. 5. These real-time PCR conditions are optimized for a StepOnePLUS system. For other systems refer to the manufacturer.

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Fig. 7 Fold change graph based on results of linear range graphs (Fig. 5)

6. The initial input amount of cell lysate depends on the cells used. The input amount (5 μg) used in Subheading 3.5 is optimized for primary meningioma cells. It is much higher than recommended by Life Technologies. The recommended highest initial amount is 50 ng. To optimize the TPA for the other cells, first use a single control cell lysate, which is certainly positive for active caspase-3. To determine the linear range use a twofold serial dilution with ten dilution steps (as described in Subheading 3.5, step 2). When the amount of active caspase-3 is too high in the cell lysate the ΔCT values are lower than in lower cell inputs. The best initial input amount is the highest cell input in the linear range (see Fig. 8). 7. As Lysate Dilution Buffer is provided in a 25 ml flask, aliquot in 1 or 2 ml tubes to make thawing faster. 8. To make correct pipetting easier mark the wells for each sample as indicated in Fig. 2. 9. When using the same reagents and amounts as stated above the “forced proximity test” should not fail. There are several possible reasons if the biotinylated antibody fails the “forced proximity test.” There could be excess free biotin in the solution, the antibody is not biotinylated, or the antibody concentration is not correct. The most likely reason for failure is incorrect antibody concentration. Determine the antibody concentration (most likely lower than 0.5 mg/ml) of the biotinylated antibody and adjust the concentration accordingly when preparing the 200 nM dilution.

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Fig. 8 Influence of input quantity on results by the example of the caspase-3 TPA using recombinant protein. Too high input amounts result in lower ΔCT values for higher input quantities (minimized linear range). Also the results are falsified. Black line: Input quantity 2,000 pg rec. protein. Grey line: Input quantity 200 pg rec. protein

10. Click “Analysis Settings.” The sensitivity for outlier detection and linear range detection can be adjusted. But mostly the default setting is sufficient. 11. The linear range of the regression lines should contain at least three data points from different input quantities. If there are no outliers the three data points should contain nine ΔCT values. The results are more reliable if all data points are included. To manually change the linear range move the mouse cursor close to the edge of the range, click the left button, and drag the edge to your desired location. The analysis below changes immediately. The ideal regression line has a R2 value of 1.0. Therefore the most reliable results are a combination of most or all data points and a R2 value near 1. 12. The fold change is calculated using the threshold. The fold change between samples is calculated between the crossover point of the linear trend line of the sample and the crossover point of the linear trend line of the control lysate at the ΔCT threshold. Therefore the value of the control lysate is always 1. Be careful. The default scale is “log2.” So are the detailed results below. To change the numbers to linear click the arrow at the right side of “Type” and choose linear. On the far right

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side is the button “Scale” to change the scale of the diagram. To export the data click “Export” at the left side. Data can be exported as *.txt or *.csv file. References 1. Fredriksson S et al (2007) Multiplexed protein detection by proximity ligation for cancer biomarker validation. Nat Methods 4(4):327–329 2. Swartzman E et al (2010) Expanding applications of protein analysis using proximity ligation and qPCR. Methods 50(4):23–26 3. Pfister C et al (2013) Detection and quantification of farnesol-induced apoptosis in difficult

primary cell cultures by TaqMan protein assay. Apoptosis 18(4):452–466 4. Ragel B et al (2008) A comparison of the cell lines used in meningioma research. Surg Neurol 70(3):295–307 5. Pallini R et al (2000) Phenotypic change of human cultured meningioma cells. J Neurooncol 49(1):9–17

Chapter 7 Detection of p53 Protein Aggregation in Cancer Cell Lines and Tumor Samples Yang Yang-Hartwich, Jamie Bingham, Francesca Garofalo, Ayesha B. Alvero, and Gil Mor Abstract The p53 protein plays a central role in regulating apoptosis. The loss of functional p53 is common in many cancers. In cancer cells, the dysfunctional p53 protein often maintains a misfolded, inactive conformation due to genetic mutations or posttranslational deregulation. The misfolded p53 protein can aggregate and form amyloid-like oligomers and fibrils, which abrogate the pro-apoptotic functions of p53. Therefore, the aggregation of p53 may be a crucial factor in carcinogenesis, tumor progression, and the response of cancer cells to apoptotic signals. In this chapter, we provide details on various methods for detecting p53 aggregation in cancer cell lines and tumor samples. Key words p53, Protein aggregation, Amyloid fibril, Thioflavin T, Non-denaturing polyacrylamide gel, Immunofluorescence staining

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Introduction

1.1 p53 Protein in Apoptosis and Cancer

The p53 protein plays a central role in cellular response to stress signals. In response to various stimuli, p53 is key in the execution of DNA repair, cell cycle arrest, senescence, and apoptosis [1]. About 50 % of all cancers have a p53 mutation and most of these mutations occur in the DNA-binding domain. Mutated p53 protein loses its ability to bind to DNA and therefore mutations can abrogate its various functions that depend on its transcriptional activity including apoptosis. In cancers wherein p53 remains wild, p53 is often inactivated by the posttranslational regulation. For example, dis-regulated phosphorylation, acetylation, or ubiquitination can inhibit p53 activation. Overexpression of MDM2/ MDM4 can increase the ubiquitination and proteosome-dependent degradation of p53 [2]. It has been reported that overexpression of the dominant-negative mutant of p53 or p53 isoforms can cause the suppression of p53 activities [3, 4]. Because the p53 protein

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acts as a tumor suppressor that maintains cellular homeostasis, all factors that destabilize p53 protein or affect its proper conformation can be oncogenic. 1.2 Protein Aggregation

Protein aggregation is the pathogenic hallmark of a growing number of diseases such as amyotrophic lateral sclerosis, Alzheimer’s, Parkinson’s, and prion disease [5]. Recent research about p53 aggregation in different types of cancers has indicated that cancer is also a protein aggregation disease [6]. Amyloid-forming proteins are an unusual subset of proteins that have the potential to aggregate. Although the amino-acid sequence of this class of proteins is diverse, when aggregated they may all adopt a similar, insoluble, highly organized structure, known as the cross-β spine consisting of an ordered arrangement of β-sheets. The formation of these aggregates depends on protein concentration, complex interactions with other proteins, and the specific cellular environment [7]. For example, inefficient protein degradation would fail to clear misfolded proteins and therefore induces protein aggregation.

1.3 Cancer as a p53 Protein Aggregation Disease

The p53 protein has recently been suggested to be a potential amyloid-forming protein. Different groups have shown that the p53 transactivation domain, as well as the DNA-binding and tetramerization domains, can all misfold and form fibrillar aggregates in vitro [8–10]. The aggregation of p53 into amyloid fibrils has been shown in several cancer cell lines and different types of tumors [6, 11]. Protein aggregates are often associated with the perturbation of essential cellular functions and various human disorders [5]. The aggregation of p53 can sequestrate the native p53 protein into an inactive conformation that loses its pro-apoptotic functions. Considering the crucial role of p53 in apoptosis and cancer, the aggregation of p53 may be a key mediator in the tumorigenicity and the progression of cancer, as well as in the chemoresistance of tumors. Since both mutant and wild-type p53 protein can aggregate, the aggregation of wild-type p53 may be the direct cause of the loss of functional p53 and the resistance to apoptosis in many tumors with wild-type TP53 gene (the gene encoding p53 protein). Little is known about the cause of p53 aggregation. One possible cause is the imbalance of p53 protein turnover, which fails to clear the misfolded p53 protein. The reactive oxygen species (ROS) that accumulate during metabolic stress may as well play a role in inducing the aggregation of p53.

1.4 Detection of p53 Aggregation in Cancer Cells

Various assays have been developed to monitor protein aggregation. The fluorescent dye Thioflavin T has been widely used for the identification and quantification of amyloid fibrils [12]. When Thioflavin T is added to samples containing β-sheet-rich deposits, such as amyloid fibrils, it fluoresces strongly with excitation at

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Fig. 1 Thioflavin T and p53 co-staining in cancer cell lines. The p53 aggregation-positive cell line A showed co-staining of p53 (red) and Thioflavin T (green). The p53 aggregation-negative cell line B showed intensive p53 staining but no Thioflavin T staining (Color figure online)

440 nm and emission at 490 nm. By combining Thioflavin T staining with p53 immunofluorescence staining, protein aggregates can be detected in cancer cell lines and tumor samples. The correlation between p53 protein expression and protein aggregation can be evaluated in terms of quantity and localization (see Fig. 1). Immunostaining, western blot, dot blot, or ELISA using antibodies that recognize the conformation-dependent generic epitopes of amyloid fibrils can also detect protein aggregates. The polyclonal rabbit antibody (OC), which was generated by immunizing rabbits with Aβ42 fibrils, can detect amyloid fibrils and fibrillar oligomers [13]. The immunofluorescence co-staining with anti-p53 antibody and antibody OC enables the visualization of protein aggregates and p53 protein in cancer cell lines and tumor samples. This is another co-staining assay for evaluating the correlation of p53 protein and aggregates in terms of their quantity and localization (see Fig. 2). Non-denaturing gel electrophoresis and western blot of p53 can more specifically detect p53 protein aggregation in cancer cells and tissues. Non-denaturing gel electrophoresis runs in the absence of sodium dodecyl sulfate (SDS) and therefore proteins retain their folded conformation in the gel. The nature of a protein’s conformation will alter its hydrodynamic size and mobility through the gel. For example, more compact conformations have higher mobility. And larger structures like oligomers and fibrils have lower mobility. Therefore, non-denaturing gel electrophoresis is an excellent tool for detecting oligomers and aggregates. After non-denaturing gel electrophoresis, proteins can be transferred to polyvinylidene difluoride (PVDF) membranes followed by immunoblotting with anti-p53 antibody.

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Fig. 2 Co-staining with anti-p53 antibody and anti-amyloid antibody OC. The p53 aggregation-positive cell line A showed strong co-staining of p53 (red) and Thioflavin T (green). The p53 aggregation-negative cell line B is positive to p53 staining but negative to OC staining (Color figure online)

Fig. 3 Non-denaturing gel and western blot of p53. Aggregated p53 protein is visualized as high molecular (MW) p53 bands

Thus the aggregated p53 protein can be visualized on the PVDF membranes as high-molecular-weight bands (see Fig. 3). Finally, a dot blot-based high-throughput screening method can also be used for large-scale tests of protein aggregation. The protein lysates are prepared from cells or tissues and loaded as dots on PVDF membranes (see Fig. 4a). The protein aggregates are detected by blotting with the antibodies targeting amyloid fibrils and fibrillar oligomers such as antibody OC (see Fig. 4b). Dot blot differs from western blot because proteins are not separated electrophoretically. The complex procedures for running the gels and transferring to membranes are not required. Therefore, dot blot is a fast, simple assay for detecting protein aggregation. It is especially suitable for testing large numbers of samples. The selected samples that are positive in dot blot can be applied to nondenaturing gel and p53 western blot in order to further confirm that p53 is involved in the formation of protein aggregation.

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Fig. 4 Dot blot for detecting protein aggregation. (a) A schematic side view of a 96-well dot blot loading system; (b) an image of dot blot detecting protein aggregation in tumor samples with antibody OC. Each dot was loaded with the protein lysate of one tumor sample. PBS was used as a negative control (arrow)

Below, we describe in detail the methods for detection of p53 aggregates using immunofluorescence, non-denaturing polyacrylamide gel electrophoresis, and dot blot.

2 2.1

Materials Equipment

1. Fluorescence microscope. 2. Western blot system (i.e., Mini Trans-Blot® Electrophoretic Transfer Cell, BioRad, #170-3930). 3. Dot blot system (i.e., Whatman Dot Blot 96 system, Biometra, #053-401). 4. Vacuum pump.

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2.2.1 Immunofluorescence: Thioflavin T and p53 Co-staining

1. 100 mM Thioflavin T: Dissolve Thioflavin T in sterile water as stock solution. Keep stock Thioflavin T solution in the dark at 4 °C. Dilute the stock solution to 5 mM with PBS before use. 2. Anti-p53 antibody (we use clone DO-1). 3. Hoechst 33342. 4. 10 % normal goat serum blocking solution. 5. Appropriate fluorescent-labeled detection antibody (here we use Alexa Fluor 594 goat anti-mouse IgG (H + L) antibody). 6. Fixation buffer: 3.7 % paraformaldehyde. 7. Permeabilization buffer: 0.2 % Triton X-100.

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8. Wash buffer 10× (i.e., Dako, #S3006). 9. Mounting medium. 10. Four- or eight-well cell culture chamber slides. 11. Cover slips. 2.2.2 Immunofluorescence: Amyloid and p53 Double Staining

1. Anti-amyloid fibril antibody (OC). 2. Anti-p53 antibody. 3. Hoechst solution: 1 μg/ml Hoechst 33342 diluted in PBS. 4. 10 % normal goat serum blocking solution. 5. Appropriate fluorescent-labeled detection antibody (here we use Alexa Fluor 594 goat anti-mouse IgG (H + L) antibody and Alexa Fluor 488 goat anti-rabbit IgG (H + L) antibody). 6. Wash buffer 10× (i.e., Dako, #S3006). 7. Mounting medium. 8. Cell culture chamber slide (e.g., BD #354632 for eight wells or #354577 for four wells). 9. Cover slips.

2.2.3 Non-denaturing Gel

1. 30 %/0.8 % (w/v) acrylamide/Bis-acrylamide. 2. 1.5 M Tris–HCl, pH 8.8. 3. 0.5 M Tris–HCl, pH 6.8. 4. Ammonium persulfate. 5. TEMED. 6. 1 M Tris–HCl, pH 6.8. 7. 1 % bromophenol blue. 8. Glycerol. 9. Prestained broad-range protein molecular weight marker. 10. Running buffer: 25 mM Tris, 192 mM glycine. 11. Transfer buffer: 25 mM Tris, 192 mM glycine, 20 % methanol. 12. Sample buffer: 15.5 ml 1 M Tris–HCl (pH 6.8), 2.5 ml 1 % bromophenol blue, 25 ml glycerol, 7 ml H2O. 13. PVDF membranes. 14. Methanol. 15. Nonfat dry milk. 16. Anti-p53 antibody (DO-1). 17. Appropriate peroxidase-labeled IgG antibody. 18. PBST buffer: Phosphate-buffered saline (pH 7.5) with 0.05 % Tween-20. 19. Chemiluminescent reagents.

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1. PVDF membranes. 2. Methanol. 3. Nonfat dry milk. 4. Anti-amyloid fibril antibody (OC). 5. Appropriate peroxidase-labeled IgG antibody. 6. PBST buffer: Phosphate-buffered saline (pH 7.5) with 0.05 % Tween-20. 7. Chemiluminescence reagents.

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3.1 Immunofluorescence Staining of Thioflavin T and p53 in Adherent Cells

1. Culture cells in chamber slide until cells reach about 70 % confluence. Aspirate medium and add fixation buffer for 10 min at room temperature (RT). 2. Aspirate fixation buffer. Add permeabilization buffer for 5 min at RT. 3. Gently wash cells twice with PBS to remove fixation and permeabilization buffer. 4. Add 10 % normal goat serum blocking solution and incubate for 30 min at RT. 5. Dilute anti-p53 antibody with 1× wash buffer (we use at dilution 1:500). Aspirate blocking solution and add p53 antibody solution. Incubate for 1–3 h at RT or overnight at 4 °C. 6. Wash chambers three times with 1× wash buffer, every time for 5–10 min. 7. Dilute Alexa Fluor 594 goat anti-mouse IgG secondary antibody with 1× wash buffer (1:1,000). Incubate cells with secondary antibody solution for 30 min at RT. From this point, the slide should always be protected from light. 8. Aspirate secondary antibody solution and wash cells three times with 1× wash buffer. 9. Aspirate the washing buffer in the chambers. Add Thioflavin T solution and incubate for 10 min at RT. 10. Aspirate the solution. Add 70 % ethanol and incubate for 5 min at RT. 11. Repeat the incubation of 70 % ethanol. 12. Wash the chambers three times with water. 13. Incubate in Hoechst 33342 solution for 5 min at RT. 14. Wash chambers three times with PBS. 15. Remove the chamber and gasket. Mount the slide. 16. Visualize using a fluorescence microscope and filter sets appropriate for each label (see Note 1).

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3.2 Immunofluorescence Staining of Amyloid Fibril and p53 in Adherent Cells

1. Follow steps 1–6 above in Subheading 3.1 for p53 staining. Then proceed to staining with anti-amyloid fibrils. 2. Dilute anti-amyloid fibril rabbit polyclonal antibody (OC) with 1× wash buffer (1:1,000). Aspirate wash buffer. Add antibody solution and incubate overnight at 4 °C. 3. Wash cells three times with 1× wash buffer. 4. Dilute Alexa Fluor 488 goat anti-rabbit IgG secondary antibody with 1× wash buffer (1:1,000). Incubate cells with secondary antibody solution for 30 min at RT. 5. Aspirate secondary antibody solution and wash cells three times with 1× wash buffer. 6. Follow steps 14–16 above in Subheading 3.2 for nuclear staining with Hoechst. 7. Visualize using a fluorescence microscope and filter sets appropriate for each label (see Note 2).

3.3 Detection of p53 Aggregation by Non-denaturing Gel Electrophoresis and Western Blot

1. Prepare the separating gel by mixing the following: (a) 6 ml 30 %/0.8 % (w/v) acrylamide/Bis-acrylamide. (b) 9.5 ml 1.5 M Tris–HCl, pH 8.8. (c) 240 μl 10 % ammonium persulfate. (d) 12 μl TEMED. (e) 8.3 ml H2O. Load into the gel cassette and allow to polymerize for 30 min. 2. Prepare the stacking gel by mixing the following: (a) 1 ml 30 %/0.8 % (w/v) acrylamide/Bis-acrylamide. (b) 2.5 ml 0.5 M Tris–HCl, pH 6.8. (c) 45 μl 10 % ammonium persulfate. (d) 22.5 μl TEMED. (e) 6.5 ml H2O. Load into the gel cassette, attach the combs, and allow to polymerize for 30 min. 3. Prepare each protein sample by mixing 10–50 μg protein with sample buffer. 4. Set up non-denaturing gel and electrophoresis apparatus. Load protein samples and protein molecular weight marker to the gel. 5. Run the gel at a fixed voltage of 100 V for 120 min. Heat may cause protein degradation, so avoid using high voltage and keep the gel system in cold room or on ice. 6. Disassemble the gel cassettes.

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7. Assemble the transfer cassette by layering the following: (a) Sponge. (b) Filter paper. (c) Gel. (d) PVDF membrane. (e) Filter paper. (f) Sponge. 8. Start the electrophoretic transfer at 32 V overnight or 100 V for 1 h. 9. After the transfer, block the membrane with 5 % nonfat milk in PBST buffer for 1 h at RT. 10. Wash the membrane three times with PBST buffer. 11. Incubate the membrane with anti-p53 antibody (we use the DO-1 clone at 1:2,000 dilution in 2 % nonfat milk in PBST) at 4 °C overnight. 12. Wash the membrane three times with PBST buffer. 13. Incubate the membrane with appropriate peroxidase-labeled secondary IgG (usually at 1:2,000 dilution in 2 % nonfat milk in PBST) for 1 h at RT. 14. Wash the membrane three times with PBST buffer. 15. Develop the blot by chemiluminescence. 3.4 Detection of p53 Protein Aggregation by Dot Blot

1. Dilute 10–50 μg protein with PBS to total volume of 100– 200 μl per sample. 2. Pretreat membranes with methanol for 2 min and then equilibrate in PBS. 3. Assemble the dot blot sandwich (see Fig. 4a) and tighten the screws, one pair of opposite screws at a time. 4. Apply vacuum to the blot system (≤150 mbar less than atmospheric pressure) and tighten the blot system again. 5. Pipette the samples into the individual wells with the vacuum off. 6. Filter the samples through the membrane for 30 min with a weak vacuum. 7. After the liquid is drawn off, add 200 μl PBS to each well with vacuum on to wash the membrane three times. 8. Disassemble the system. Block the membrane with 5 % nonfat milk in PBST buffer for 1 h at RT. 9. Incubate the membrane with anti-amyloid fibril antibody (1:1,000 dilution in 2 % nonfat milk in PBST) at 4 °C overnight.

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10. Wash the membrane three times with PBST buffer. 11. Incubate the membrane with peroxidase-labeled IgG secondary antibody (1:2,000 dilution in 2 % nonfat milk in PBST) for 1 h at RT. 12. Wash the membrane three times with PBST buffer. 13. Develop the blot by chemiluminescence.

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Notes 1. Since Vassar and Culling first described the use of Thioflavin T as a fluorescent marker of amyloid in 1959 [14], Thioflavin T has become one of the most widely used compounds for staining amyloid fibrils. The substantial enhancement of its fluorescence emission upon binding to fibrils makes Thioflavin T a particularly powerful and convenient tool. The Thioflavin T stain is very easy to perform and the results are easy to interpret. The Thioflavin stain also has the advantage of detecting very small amounts of amyloid. It can be performed on cultured cells and paraffin-embedded and frozen tissues. However, Thioflavin stains are not permanent. The fluorescence fades rapidly during exposure. Therefore, the samples should be protected from direct light as much as possible to prevent the fading of fluorescent signal. Also, Thioflavin T is not entirely specific to amyloid in some conditions. The structures, like fibrin, keratin, intestinal muciphages, paneth cells, zymogen granules, and juxtaglomerular apparatus, may show positive stains of Thioflavin T [15]. The staining Thioflavin T results should be confirmed by either electron microscopy or other staining methods, such as Congo red staining [16]. 2. The anti-amyloid fibril antibody OC that is used in the immunofluorescence staining and dot blot of protein aggregates recognizes the generic epitope of amyloid fibrils and soluble fibrillar oligomers. Many conformation-dependent monoclonal or polyclonal antibodies have been generated to detect amyloid fibrils and soluble fibrillar oligomers [17–19]. They may have different features. For instance, another antibody A11 is often used to detect oligomers in the study of degenerative diseases [13]. A11 stains small focal or punctuate deposits in Alzheimer’s disease tissues; it does not stain diffuse plaques or other plaque types. A11 specifically recognizes the generic epitope common to prefibrillar oligomers that represent the toxic or pathological species of aggregates and does not detect fibrils, monomers, or natively folded precursor proteins. In our experiments, the samples (cancer cell lines and tumor tissues) that are positive for OC are not always positive for A11. OC can detect a wider range of samples with protein aggregation.

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A11 is more suitable for the detection of oligomers formed by p53 protein when it is used for the co-staining with anti-p53 antibody. Therefore, it is critical to select the proper antibody for protein aggregation study based on the purpose of the experiment. It is also critical to include proper controls in the staining to insure the specificity of the antibody and eliminate the background signals. For instance, negative controls in which the primary antibody is omitted or replaced with normal serum of the same species as primary antibody can test the specificity of the anti-amyloid antibodies. It will be ideal if a positive control that is known to contain protein aggregates is tested to confirm that the experiment procedure works. References 1. Zilfou JT, Lowe SW (2009) Tumor suppressive functions of p53. Cold Spring Harb Perspect Biol 1(5):a001883. doi:10.1101/cshperspect. a001883 2. Michael D, Oren M (2002) The p53 and Mdm2 families in cancer. Curr Opin Genet Dev 12(1):53–59, S0959437X01002647 [pii] 3. Chen J, Ng SM, Chang C, Zhang Z, Bourdon JC, Lane DP, Peng J (2009) p53 isoform delta113p53 is a p53 target gene that antagonizes p53 apoptotic activity via BclxL activation in zebrafish. Genes Dev 23(3):278–290. doi:10.1101/gad.1761609, 23/3/278 [pii] 4. Chan WM, Poon RY (2007) The p53 isoform Deltap53 lacks intrinsic transcriptional activity and reveals the critical role of nuclear import in dominant-negative activity. Cancer Res 67(5):1959–1969. doi:10.1158/0008-5472. CAN-06-3602, 67/5/1959 [pii] 5. Stefani M, Dobson CM (2003) Protein aggregation and aggregate toxicity: new insights into protein folding, misfolding diseases and biological evolution. J Mol Med (Berl) 81(11): 678–699. doi:10.1007/s00109-003-0464-5 6. Silva JL, Rangel LP, Costa DC, Cordeiro Y, De Moura Gallo CV (2013) Expanding the prion concept to cancer biology: dominant-negative effect of aggregates of mutant p53 tumour suppressor. Biosci Rep 33(4):e00054. doi:10.1042/ BSR20130065, BSR20130065 [pii] 7. Aguzzi A, O’Connor T (2010) Protein aggregation diseases: pathogenicity and therapeutic perspectives. Nat Rev Drug Discov 9(3):237– 248. doi:10.1038/nrd3050, nrd3050 [pii] 8. Rigacci S, Bucciantini M, Relini A, Pesce A, Gliozzi A, Berti A, Stefani M (2008) The (1–63) region of the p53 transactivation domain aggregates in vitro into cytotoxic amyloid assemblies. Biophys J 94(9):3635–3646.

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doi:10.1529/biophysj.107.122283, S0006-3495 (08)70440-5 [pii] Ishimaru D, Andrade LR, Teixeira LS, Quesado PA, Maiolino LM, Lopez PM, Cordeiro Y, Costa LT, Heckl WM, Weissmuller G, Foguel D, Silva JL (2003) Fibrillar aggregates of the tumor suppressor p53 core domain. Biochemistry 42(30):9022–9027. doi:10.1021/bi034218k Higashimoto Y, Asanomi Y, Takakusagi S, Lewis MS, Uosaki K, Durell SR, Anderson CW, Appella E, Sakaguchi K (2006) Unfolding, aggregation, and amyloid formation by the tetramerization domain from mutant p53 associated with lung cancer. Biochemistry 45(6):1608–1619. doi:10.1021/bi051192j Xu J, Reumers J, Couceiro JR, De Smet F, Gallardo R, Rudyak S, Cornelis A, Rozenski J, Zwolinska A, Marine JC, Lambrechts D, Suh YA, Rousseau F, Schymkowitz J (2011) Gain of function of mutant p53 by coaggregation with multiple tumor suppressors. Nat Chem Biol 7(5):285–295. doi:10.1038/nchembio.546, nchembio.546 [pii] Khurana R, Coleman C, Ionescu-Zanetti C, Carter SA, Krishna V, Grover RK, Roy R, Singh S (2005) Mechanism of thioflavin T binding to amyloid fibrils. J Struct Biol 151(3):229–238. S1047doi:10.1016/j.jsb.2005.06.006, 8477(05)00130-9 [pii] Kayed R, Head E, Sarsoza F, Saing T, Cotman CW, Necula M, Margol L, Wu J, Breydo L, Thompson JL, Rasool S, Gurlo T, Butler P, Glabe CG (2007) Fibril specific, conformation dependent antibodies recognize a generic epitope common to amyloid fibrils and fibrillar oligomers that is absent in prefibrillar oligomers. Mol Neurodegener 2:18. doi:10.1186/17501326-2-18, 1750-1326-2-18 [pii]

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14. Vassar PS, Culling CF (1959) Fluorescent stains, with special reference to amyloid and connective tissues. Arch Pathol 68:487–498 15. Brancroft J, Gamlble M (2002) Theory and practice of histological techniques, 5th edn. Churchill Livingstone, London 16. Linke R (2006) Congo red staining of amyloid: improvements and practical guide for a more precise diagnosis of amyloid and the different amyloidoses. In: Uversky VN, Fink AL (eds) Protein misfolding, aggregation, and conformational diseases, vol 4. Springer, New York, pp 239–376 17. Iwatsubo T, Odaka A, Suzuki N, Mizusawa H, Nukina N, Ihara Y (1994) Visualization of A beta 42(43) and A beta 40 in senile plaques

with end-specific A beta monoclonals: evidence that an initially deposited species is A beta 42(43). Neuron 13(1):45–53, 0896-6273(94) 90458-8 [pii] 18. Fukumoto H, Asami-Odaka A, Suzuki N, Iwatsubo T (1996) Association of A beta 40-positive senile plaques with microglial cells in the brains of patients with Alzheimer’s disease and in non-demented aged individuals. Neurodegeneration 5(1):13–17 19. Hrncic R, Wall J, Wolfenbarger DA, Murphy CL, Schell M, Weiss DT, Solomon A (2000) Antibodymediated resolution of light chain-associated amyloid deposits. Am J Pathol 157(4):1239– 1246. doi:10.1016/S0002-9440(10)64639-1, S0002-9440(10)64639-1 [pii]

Chapter 8 Detection of p53 Protein Transcriptional Activity by Chromatin Immunoprecipitation Yang Yang-Hartwich, Emily Romanoff, Jamie Bingham, Ayesha B. Alvero, and Gil Mor Abstract p53 is a key transcriptional mediator that controls the expression of hundreds of target genes necessary to maintain cellular homeostasis and genome integrity. An important cellular function that is dependent on p53 transcriptional activity is apoptosis or programmed cell death. Indeed, inhibition of p53 transcriptional activity is often observed in cancers as a result of mutations within its DNA-binding domain. In this chapter, we describe the use of chromatin immunoprecipitation and real-time quantitative polymerase chain reaction to detect p53 transcriptional activity in cancer cells and tumor tissues. This technique enables the determination of the ability of p53 to bind to the promoter region of apoptotic genes and to evaluate the transcription-dependent activity of p53-induced apoptosis. Key words p53, Transcriptional activation, Chromatin immunoprecipitation (ChIP), Real-time quantitative PCR

1 1.1

Introduction Cancer and p53

p53 has been described as the “the guardian of the genome,” as it is a master transcriptional mediator that controls cellular homeostasis [1]. The mutation of TP53 gene (encoding p53 protein) is the most frequent mutation in human cancers. Even in cases of cancer that express wild-type TP53, perturbations in pathways signaling to p53 often exist. Inactivation of p53 is a common feature of cancers [2]. It causes the inappropriate regulation of p53 target genes and the failure to induce p53-dependent responses to stress signals. The majority of p53 mutations located in the DNA-binding domain display loss of function regarding their transcriptional activity [3], resulting in cellular inability to trigger apoptosis upon DNA damage. The inhibition of p53 transcriptional activity is therefore an important factor for tumor initiation and progression in many types of cancers.

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1.2 The p53 Target Genes

The identification of p53 target genes is very important for understanding the functions of p53 and how p53 regulates cell growth and apoptosis. As a transcription factor, p53 specifically recognizes the DNA sequences containing two adjacent copies of the consensus sequence 5′-RRRCWWGYYY-3′ (R = G/A, W = T/A, T = C/T) separated by 0–13 base pairs [4]. Thousands of genes that contain at least one p53 consensus binding sequence in their promoters have been reported although only a small portion of these genes have been confirmed. p21, GADD45, and 14-3-3σ are three p53 target genes that play critical roles in inducing cell cycle arrest, while Bax, CD95(Fas), Noxa, PUMA, Killer/DR5, Ei24/PIG8, PERP, Pidd, and p53AIP1 are target genes that induce apoptosis [5–12].

1.3 Assays for Detecting p53 Transcriptional Activity

Different assays have been developed for the detection of p53 transcriptional activity. For instance, transcriptional activity of p53 can be determined by the analysis of mRNA expression levels of p53 target genes or reporter assays. A reporter assay system requires a plasmid construct that encodes either the firefly luciferase reporter gene or a gene encoding a fluorescent protein under the control of a minimal promoter and tandem repeats of the p53 response element. When this plasmid is transfected into cells, the p53 transcriptional activity in the cells can be evaluated. Chromatin immunoprecipitation (ChIP) is a technique used to study the interaction between protein and DNA. ChIP of p53 protein is an invaluable method for analyzing the binding of p53 to the DNA. In a p53 ChIP experiment, cells or tissues are first treated by formaldehyde to cross-link DNA and the bond p53 protein. Chromatin is then isolated and sheered into small fragments by sonication or enzyme digestion prior to immunoprecipitation (IP). IP specifically isolates the complex of p53 and the binding DNA fragments. Reverse cross-linking releases the DNA and digests the protein and the purified DNA can be used for PCR particularly with primers designed for p53-binding sequences in the promoter regions of different target genes. By real-time quantitative PCR, the amount of DNA fragments bound to p53 can be quantified. Thus the DNA-binding ability of p53, which reflects the level of p53 transcriptional activation, can be evaluated.

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1. Microcentrifuge. 2. Sonicator. 3. Eppendorf tube rotator. 4. Real-time PCR system.

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1. 37 % formaldehyde. 2. 1.25 M glycine. 3. Protease inhibitor cocktail. 4. Cell lysis buffer: 20 mM Tris–HCl (pH 8.0), 85 mM KCl, 0.5 % NP-40. 5. Nuclei lysis buffer: 50 mM Tris–HCl (pH 8.0), 10 mM EDTA, 1 % SDS. 6. Dilution buffer: 0.01 % SDS, 1.1 % Triton X-100, 1.1 mM EDTA, 20 mM Tris–HCl (pH 8.0), 167 mM NaCl. 7. Protein A agarose/salmon sperm DNA beads. 8. Anti-p53 antibody (clone DO-1). 9. Isotype-matched control IgG. 10. Low-salt wash buffer: 0.1 % SDS, 1 % Triton X-100, 2 mM EDTA, 20 mM Tris–HCl (pH 8.0), 150 mM NaCl. 11. High-salt wash buffer: 0.1 % SDS, 1 % Triton X-100, 2 mM EDTA, 20 mM Tris–HCl (pH 8.0), 500 mM NaCl. 12. LiCl wash buffer: 0.25 M LiCl, 1 % NP-40, 1 % deoxycholate, 1 mM EDTA, 20 mM Tris–HCl. 13. Elution buffer: 1 % SDS, 50 mM NaHCO3. 14. TE buffer: 10 mM Tris–HCl (pH 8.0), 1 mM EDTA. 15. RNase. 16. 5 M NaCl. 17. Proteinase K. 18. PCR Purification Kit. 19. SYBR green Real-time PCR mix. 20. Primers for real-time PCR (see Note 1).

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Methods 1. For cross-linking in adherent cells: Add formaldehyde to a final concentration of 1 % (e.g., 0.27 ml of 37 % formaldehyde to 10 ml media) directly to the media and shake for 10 min at room temperature (RT). 2. For cross-linking in non-adherent cells: Add formaldehyde to a final concentration of 1 % directly to the floating cells in the media and shake for 10 min at RT. 3. For cross-linking in tissues: Frozen tissue should be thawed first on ice. Chop thawed frozen tissue or fresh tissue into small pieces using two razor blades. Add 10 ml PBS per gram of tissue. Add formaldehyde to a final concentration of 1 % directly to the tissue in PBS and shake for 15 min at RT.

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4. Stop the cross-linking by adding glycine to a final concentration of 125 mM. Shake for 5 min at RT. 5. Wash the adherent cells with ice-cold PBS three times and then scrape the cells into 1 ml PBS. For non-adherent cells and tissues, centrifuge at 2,000 rpm (300 × g) for 5 min and then wash the pellets with ice-cold PBS three times. 6. Resuspend the pellets in cell lysis buffer containing protease inhibitors and incubate on ice for 10 min. 7. Centrifuge at 5,000 rpm for 5 min. 8. Resuspend the pellets (nuclei fragment) in nuclear lysis buffer containing protease inhibitors. Incubate on ice for 10 min. 9. Sonicate the chromatin to an average length of about 100– 500 bp. Keep samples on ice (see Note 2). 10. Centrifuge at 13,000 rpm for 10 min at 4 °C. The supernatant contains the chromatin. It can be stored at −80 °C. 11. Dilute the chromatin with dilution buffer (1:5–1:20). Add 50 μl of Protein A agarose/salmon sperm DNA bead slurry to 500 μl of diluted chromatin and rotate the tube at 4 °C for 30 min to pre-clear the sample. 12. Quickly centrifuge the beads at 13,000 rpm × 15 s and transfer the supernatant to a new tube. Save 50 μl of the supernatant as input control. 13. Add 1 μg antibody to 500 μl chromatin in each tube. Anti-p53 antibody is added to the test groups. IgG is added to the IP control groups. 14. Incubate with agitation at 4 °C for 3 h to overnight. 15. Add 50 μl of Protein A agarose/salmon sperm DNA beads to each tube and rotate the tube at 4 °C for 1 h. 16. Quickly centrifuge the beads at 13,000 rpm × 15 s and aspirate the supernatant. 17. Wash the beads with the following buffers. For every washing step, add 1 ml of the wash buffer and rotate the tube at 4 °C for 5 min. Then spin down the beads: (a) Low-salt wash buffer. (b) High-salt wash buffer. (c) LiCl wash buffer. (d) TE buffer, twice. 18. Prepare fresh elution buffer. Add 250 μl elution buffer to the beads. Incubate at RT for 15 min. Vortex the beads to mix several times during the incubation. 19. Spin down the beads at 13,000 rpm for 5 min. Transfer the supernatant to a new tube. Repeat the elution step (step 18) once. Combine the supernatant.

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20. Reverse formaldehyde cross-linking by adding 1 μl RNase (from 10 mg/ml stock) and 20 μl of 5 M NaCl to the elution samples and the input controls. Incubate at 65 °C for 4 h. 21. Add 10 μl 0.5 M EDTA (pH 8.0), 12 μl 1.0 M Tris-HCL (pH 6.5), and 5 μl 50 mg/ml proteinase K. Incubate at 45 °C for 1 h. 22. Extract the DNA using a PCR Purification Kit. Alternatively, ethanol extraction or phenol-chloroform extraction can be used, see Notes 3 and 4. 23. The purified DNA can be used for real-time PCR. 24. Analyze the PCR data (see Note 5).

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Notes 1. The PCR primers for p53 ChIP assays are commercially available or can be designed based on the sequence of the p53binding site in the target gene of interest. PUMA taken as an example, there are two p53-binding sites in the promoter region of PUMA, both of which can be directly bound and transactivated by p53 [10]. Primers can be designed by targeting the proximal region of either of the p53-binding sites. A negative control for the PCR should be included to test whether the observed DNA enrichment is specific since some antibodies may cause nonspecific enrichment. The negative control primers are usually designed for a negative locus where the protein of interest (p53) is absent. For example, a region in the exon4 of GAPDH can be used to design the negative control primers [13]. 2. The conditions for sonication must be optimized for each cell line or tissue type and the instrument. For example, Fisher’s Sonicator Model 500 was used here. A good starting point for optimizing the condition is 5, 10, and 15 min with 30 s “on” and 30 s “off” cycle at 50–70 % of maximum power. After sonication, the result can be checked by running a gel. First, use 10 μl sample, add 40 μl H2O, reverse cross-link by adding 2 μl of 5 M NaCl, and then boil for 15 min. After returning to room temperature, add 1 μl of 10 mg/ml RNase A at 37 °C for 10 min. Clean and purify DNA with PCR Purification Kit. Load 5 μl of the purified DNA on gel and determine the size of smear. The sonication condition that gives a smear of DNA sizes from 200 bp to 1 kb with a peak around 500 bp should be used for ChIP reactions (see Fig. 1). 3. Phenol/chloroform extraction of DNA: (a) Add an equal volume of phenol:chloroform (1:1) to the DNA sample.

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Fig. 1 DNA gel for optimizing the conditions of sonication. An ovarian cancer cell line was used in this test. (A) 100 bp DNA marker. (B) Sonication for 5 min. (C) Sonication for 10 min

(b) Mix the contents of the tube vigorously by vortexing. (c) Centrifuge the mixture at 13,000 rpm for 10 min at room temperature. (d) Transfer the aqueous phase (the upper phase) to a fresh tube. (e) Repeat steps (a) to (d) until no protein is visible at the interface of the organic and aqueous phases. (f) Add an equal volume of chloroform and repeat step through. (g) Recover the nucleic acid by ethanol precipitation. 4. The PCR data need to be properly normalized to minimize the effect of different sources of variability, such as the amount of chromatin, the efficiency of immunoprecipitation, and the DNA recovery. Data can be normalized using either of the two methods—the percent input method and the fold enrichment method. In the percent input method, the signals of ChIP samples are divided by the signals of input sample. Both the IgG control and the IP p53 samples are normalized to the input control. In the fold enrichment method, the signals of ChIP samples are divided by the signals of IgG control sample. The result is represented as the fold increase in IP p53 signal relative to the background signal (IP with IgG). The percent input method is preferred since it includes the normalization of both background and the amount of input chromatin. 5. Ethanol precipitation: (a) Add 1/10 volume sodium acetate (3 M, pH 5.5). Mix by inverting tube. (b) Add 2–3 volumes of 100 % ethanol. Mix by inverting tube.

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(c) Store the solution on ice for 15–30 min (or at −80 °C for 5–10 min) to allow the precipitation to form. (d) Spin down at 13,000 rpm at 4 °C for 10–15 min. (e) Aspirate supernatant carefully not to disturb pellet. (f) Add 500 μl 70 % ethanol (stored at −20 °C). (g) Spin in microcentrifuge at 13,000 rpm at 4 °C for 10 min. (h) Aspirate supernatant. Dry pellet at room temperature for 10 min. (i) Resuspend the pellet in water or TE buffer. TE buffer is preferred, because the DNA would be more stable. References 1. Beckerman R, Prives C (2010) Transcriptional regulation by p53. Cold Spring Harb Perspect Biol 2(8):a000935. doi:10.1101/ cshperspect.a000935, cshperspect.a000935 [pii] 2. Olivier M, Hollstein M, Hainaut P (2010) TP53 mutations in human cancers: origins, consequences, and clinical use. Cold Spring Harb Perspect Biol 2(1):a001008. doi:10.1101/ cshperspect.a001008 3. Smeenk L, van Heeringen SJ, Koeppel M, van Driel MA, Bartels SJ, Akkers RC, Denissov S, Stunnenberg HG, Lohrum M (2008) Characterization of genome-wide p53-binding sites upon stress response. Nucleic Acids Res 36(11):3639–3654. doi:10.1093/nar/gkn232, gkn232 [pii] 4. el-Deiry WS, Kern SE, Pietenpol JA, Kinzler KW, Vogelstein B (1992) Definition of a consensus binding site for p53. Nat Genet 1(1): 45–49. doi:10.1038/ng0492-45 5. Miyashita T, Reed JC (1995) Tumor suppressor p53 is a direct transcriptional activator of the human bax gene. Cell 80(2):293–299, 0092-8674(95)90412-3 [pii] 6. Gu Z, Flemington C, Chittenden T, Zambetti GP (2000) ei24, a p53 response gene involved in growth suppression and apoptosis. Mol Cell Biol 20(1):233–241 7. Oda E, Ohki R, Murasawa H, Nemoto J, Shibue T, Yamashita T, Tokino T, Taniguchi T, Tanaka N (2000) Noxa, a BH3-only member of the Bcl-2 family and candidate mediator of p53-induced apoptosis. Science 288(5468): 1053–1058, 8508 [pii]

8. Wu GS, Burns TF, McDonald ER 3rd, Jiang W, Meng R, Krantz ID, Kao G, Gan DD, Zhou JY, Muschel R, Hamilton SR, Spinner NB, Markowitz S, Wu G, el-Deiry WS (1997) KILLER/DR5 is a DNA damage-inducible p53-regulated death receptor gene. Nat Genet 17(2):141–143. doi:10.1038/ng1097-141 9. Attardi LD, Reczek EE, Cosmas C, Demicco EG, McCurrach ME, Lowe SW, Jacks T (2000) PERP, an apoptosis-associated target of p53, is a novel member of the PMP-22/gas3 family. Genes Dev 14(6):704–718 10. Yu J, Zhang L, Hwang PM, Kinzler KW, Vogelstein B (2001) PUMA induces the rapid apoptosis of colorectal cancer cells. Mol Cell 7(3):673–682, S1097-2765(01)00213-1 [pii] 11. Oda K, Arakawa H, Tanaka T, Matsuda K, Tanikawa C, Mori T, Nishimori H, Tamai K, Tokino T, Nakamura Y, Taya Y (2000) p53AIP1, a potential mediator of p53dependent apoptosis, and its regulation by Ser46-phosphorylated p53. Cell 102(6):849–862, S0092-8674(00)00073-8 [pii] 12. Owen-Schaub LB, Zhang W, Cusack JC, Angelo LS, Santee SM, Fujiwara T, Roth JA, Deisseroth AB, Zhang WW, Kruzel E et al (1995) Wildtype human p53 and a temperature-sensitive mutant induce Fas/APO-1 expression. Mol Cell Biol 15(6):3032–3040 13. Kaeser MD, Iggo RD (2002) Chromatin immunoprecipitation analysis fails to support the latency model for regulation of p53 DNA binding activity in vivo. Proc Natl Acad Sci U S A 99(1):95–100. doi:10.1073/pnas.012283399, 012283399 [pii]

Chapter 9 Homogeneous, Bioluminescent Proteasome Assays Martha A. O’Brien, Richard A. Moravec, Terry L. Riss, and Robert F. Bulleit Abstract Protein degradation is mediated predominantly through the ubiquitin–proteasome pathway. The importance of the proteasome in regulating degradation of proteins involved in cell-cycle control, apoptosis, and angiogenesis led to the recognition of the proteasome as a therapeutic target for cancer [1–6]. The proteasome is also essential for degrading misfolded and aberrant proteins, and impaired proteasome function has been implicated in neurodegerative and cardiovascular diseases [7, 8]. Robust, sensitive assays are essential for monitoring proteasome activity and for developing inhibitors of the proteasome. Peptideconjugated fluorophores are widely used as substrates for monitoring proteasome activity, but fluorogenic substrates can exhibit significant background and can be problematic for screening because of cellular autofluorescence or interference from fluorescent library compounds. Furthermore, fluorescent proteasome assays require column-purified 20S or 26S proteasome (typically obtained from erythrocytes), or proteasome extracts from whole cells, as their samples. To provide assays more amenable to highthroughput screening, we developed a homogeneous, bioluminescent method that combines peptideconjugated aminoluciferin substrates and a stabilized luciferase. Using substrates for the chymotrypsin-like, trypsin-like, and caspase-like proteasome activities in combination with a selective membrane permeabilization step, we developed single-step, cell-based assays to measure each of the proteasome catalytic activities. The homogeneous method eliminates the need to prepare individual cell extracts as samples and has adequate sensitivity for 96- and 384-well plates. The simple “add and read” format enables sensitive and rapid proteasome assays ideal for inhibitor screening. Key words Proteasome assay, 20S Proteasome, Bioluminescence, Luciferase, Aminoluciferin, Bioluminescent proteasome assay, Cell-based proteasome assay

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Introduction

1.1 The UbiquitinProteasome Pathway

In eukaryotic cells, the turnover of intracellular proteins is mediated mainly by the ubiquitin–proteasome pathway, a nonlysosomal proteolytic pathway. The 26S proteasome is a 2.5-MDa multiprotein complex found in both the nucleus and the cytosol of all eukaryotic cells and is comprised of a single 20S core particle and 19S regulatory particles at one or both ends [9, 10]. Three major proteolytic

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activities are contained within the 20S core. Together these three activities are responsible for much of the protein degradation required to maintain cellular homeostasis including degradation of critical cell cycle proteins, tumor suppressors, transcription factors, inhibitory proteins, and damaged cellular proteins [11, 12]. Proteins destined to be degraded by the proteasome are first selectively targeted by the addition of a series of covalently attached ubiquitin molecules. The 26S proteasome degrades polyubiquitinated proteins in an ATP-dependent manner. The 19S regulatory unit binds and removes the ubiquitin chains from tagged proteins, and ATPases within the regulatory complex appear to unfold protein substrates and translocate the unfolded polypeptides into the 20S core [11, 12]. There the polypeptides are degraded to yield peptides ranging from 3 to 25 amino acids in length [13]. The 20S catalytic core and the 19S regulatory complex are highly conserved from yeast to mammals [14]. The catalytic core of the complex, the 20S proteasome, is a barrel-shaped assembly of 28 protein subunits that possesses three different proteolytic activities designated as chymotrypsin-like, trypsin-like, and caspase-like (also termed post-glutamyl peptide hydrolase) [14, 15]. The catalytic sites are located on the inner surface of the central β-rings of the cylindrical particle, and access to them is controlled by narrow and gated channels in the outer α-rings of the complex. The association of the 20S particle with a 19S regulatory complex at one or both ends of the barrel forms the 26S proteasome and confers an open channel conformation, resulting in much higher rates of peptide hydrolysis in the 26S proteasome [16, 17]. Robust, sensitive assays for the catalytic activities of the proteasome will aid in the discovery and validation of new inhibitors. The proteasome has been validated as a therapeutic target for cancer treatment. Proteasome inhibitors induce apoptosis, and transformed cells, especially multiple myeloma cells, display greater susceptibility to proteasome inhibition than nonmalignant cells [3, 6]. The enhanced proliferative rate of malignant cells may cause accumulation of damaged proteins at a higher rate that in turn would increase dependency on proteasomal degradation [18]. The first-generation proteasome inhibitor, bortezomib (PS-341, Velcade®), is now an FDA-approved drug for the treatment of multiple myeloma and mantle cell lymphoma, and several secondgeneration inhibitors are being developed [19–22]. The clinical testing of bortezomib, as well as other new proteasome inhibitors, for efficacy on an array of cancers is currently in progress [23–27]. Useful high-throughput assays can increase the efficiency of screening for new inhibitors. We describe here the bioluminescent proteasome assays, compare them with fluorescent assays, and demonstrate their utility in several cell lines.

Bioluminescent Proteasome Assays

1.2 Bioluminescent Proteasome Assay Concept

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Peptide-conjugated fluorophores are widely used as substrates for monitoring proteasome activity, but sensitivity of fluorescent assays can be limited for a variety of reasons. Peptide-conjugated fluorophores can have residual fluorescence or spectral overlap with their cleaved fluorescent products, thus increasing background and reducing sensitivity [28, 29]. Cells can exhibit autofluorescence and compounds in natural product, and synthetic chemical libraries frequently exhibit fluorescence that can cause assay interference [30]. To provide an alternative to fluorescence, we synthesized luminogenic substrates using standard Fmoc chemistry as previously described [31]. We synthesized luminogenic versions of the commonly used fluorogenic coumarin-based substrates, SucLLVY-aminoluciferin, Z-LRR-aminoluciferin, and Z-nLPnLDaminoluciferin, to monitor the chymotrypsin-like, trypsin-like, and caspase-like activities of the proteasome, respectively [32] (see Fig. 1). The bioluminescent assays are homogeneous assays, such that the proteasome and luciferase function simultaneously. As a result of this coupled-enzyme format, the proteasome and luciferase rapidly reach a steady-state, where the rate of proteasome cleavage of the substrate is equal to the rate of luciferase utilization of the released aminoluciferin, and stable light output is achieved. Steadystate is typically reached in 10–20 min, and stable light output persists for several hours (see Fig. 2). At steady-state, the light output is proportional to the rate of proteasome cleavage and thus the H Suc-LLVY- N or Z-LRRor Z-nLPnLD-

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Fig. 1 The luminogenic, aminoluciferin substrates containing the Suc-LLVY, Z-LRR, or Z-nLPnLD sequence are recognized by the 20S proteasome. Following 20S proteasome cleavage, the substrate for luciferase (aminoluciferin) is released, allowing the luciferase reaction to produce light

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Fig. 2 The proteasome assays give stable signal. The proteasome assays (Proteasome-Glo™ 3-Substrate System) were tested with human purified 20S proteasome (1 μg/ml) (closed symbols) or without 20S proteasome as a control (open symbols) in 96-well plates in 100 μl total volume. Luminescence was monitored at various times for 3 h on a GloMax™ 96 Microplate luminometer. The signals peak rapidly and then are very stable for all three assays as shown on a log scale

amount of proteasome activity (see Fig. 3). Eventually, the light output decreases when the proteasome and luciferase become inactivated, but the half-life for each of the proteasome assays is greater than 3 h (see Fig. 2). Another feature of the homogeneous, bioluminescent format is that any free aminoluciferin that is a by-product of the peptideconjugating synthesis is removed from the reagent prior to exposing the proteasome substrate to the test samples. Consequently, the background is very low, and the linear dynamic range is very large. The broad dynamic range and stable signal results in increased sensitivity and flexibility for the bioluminescent proteasome assays. A comparison of bioluminescent and fluorescent proteasome assays demonstrates that the bioluminescent assays are significantly more sensitive and have a much lower limit of detection for proteasome activity (see Fig. 3). 1.3 Cellular Bioluminescent, Proteasome Assay Concept

Historically, proteasome activity was measured in cells by making cell lysates, using various methods to enrich for proteasome, and then testing for activity using fluorogenic substrates [33, 34]. Being able to monitor proteasome activity for all three catalytic sites directly in cells in multiwell culture dishes has obvious advantages for high-throughput screening applications [35]. The sensitivity of the bioluminescent, homogeneous format enabled the

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Suc-LLVY-Glo Z-LRR-Glo

10,000

Z-nLPnLD-Glo Suc-LLVY-AMC

Signal/Noise

Suc-LLVY-AMC+SDS 1,000

Boc-LRR-AMC Ac-nLPnLD-AMC

100

10

1 0.00001

0.0001

0.001

0.01

0.1

1

10

20S (ug/ml)

Fig. 3 Luminescent proteasome assays are more sensitive than fluorescent proteasome assays. Human 20S proteasome was serially diluted in 10 mM HEPES (pH 7.6) in 96-well plates. For each catalytic activity, half the plate received the appropriate Proteasome-Glo™ Reagent and half the plate received the comparable fluorogenic substrate, diluted in 100 mM HEPES, pH 7.5, 1 mM EDTA, to the same concentration as the luminescent substrates. The fluorescent assay for chymotrypsin-like proteasome activity was run with and without 0.02 % SDS (see Note 1). Thirty minutes after addition of the Proteasome-Glo™ Reagent, luminescence was recorded as relative light units (RLU) on a GloMax™ 96 Microplate luminometer. Fluorescence was measured 30 min after adding the appropriate substrate on a LabSystems Fluoroskan Ascent fluorometer and recorded as relative fluorescence units (RFU). To normalize between RLU and RFU, the results were plotted as signal to noise ratios. Each point represents the average of four wells. The Proteasome-Glo™ Assays were linear over four logs of 20S proteasome concentration for all three assays. The limit of detection is defined as a signal to noise ratio = 3 (dotted line). The luminescent proteasome assays give higher signal to noise ratios and lower limits of detection than the fluorescent assays

development of direct cellular assays for proteasome activity. In addition to sensitivity, specificity is critical when developing a cellular assay. The proteasome catalytic activities are described as chymotrypsin-like, trypsin-like, and caspase-like, indicating the relationship to catalytic properties of other proteases. To minimize nonspecific protease activity on the proteasome substrates, we developed a permeabilizing agent that enhances access to the proteasome while leaving lysosomal vesicles intact [32]. Specificity of the Suc-LLVY-aminoluciferin substrate for the chymotrypsin-like activity of the proteasome was demonstrated with lactacystin and epoxomicin, highly specific and irreversible inhibitors of the proteasome (see Fig. 4). However, when using

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Fig. 4 Comparison of two proteasome inhibitors tested in cultured cells. U266 multiple myeloma cells were grown in RPMI-1640 containing 10 % FBS and 1 mM sodium pyruvate. Cells were added to 96-well plates at 10,000 cells/90 μl/ well. Cells were then equilibrated at 37 °C, 5 % CO2 for 2 h. Serial dilutions of lactacystin or epoxomicin were prepared in culture medium, and 10 μl of each dilution was added to wells. The cells were incubated with the drugs for 105 min at 37 °C, 5 % CO2. The plate was allowed to equilibrate to 22 °C before 100 μl/ well of Proteasome-Glo™ Chymotrypsin-like Cell-Based Reagent was added. Luminescence was measured with a DYNEX MLX® luminometer 15 min after adding reagent. The relative potency for the two inhibitors is consistent with published information [39]

the Z-LRR-aminoluciferin to measure trypsin-like activity directly in cells, we initially observed significant nonspecific background luminescence originating from fetal bovine serum-supplemented culture medium, as well as from the cells. Protease inhibitors were screened for their specific ability to decrease both of these background activities in this assay format, while having minimal impact on the proteasome. A mixture of three inhibitors was found to reduce background luminescence [32] (see Fig. 5). These protease inhibitors decrease serum background by approximately 95 % when added to the trypsin-like luminescent proteasome reagent. Cellular background in U266 cells, defined as activity which was not inhibited by a 2 h epoxomicin pretreatment, was reduced from 45 to 4.4 %. Inclusion of the protease inhibitors into the luminescent reagent was an effective way to significantly increase specificity of trypsin-like proteasome measurements and still retain a no-wash homogeneous format (see Fig. 5). The specificity of the assays can be confirmed by inhibiting a majority of the activity with the selective proteasome inhibitors, lactacystin, epoxomicin [36–40], or bortezomib (see Fig. 7). Numerous cell lines have been tested with the assays, including Jurkat, U937, U266, H929, RPMI-8226, HL-60, H226, PA-1, DU 145, SW 620, MOLT-4, and MCF-7, and the specificities have been confirmed for all three assays (see Fig. 8).

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800,000 Medium

700,000

Luminescence (RLU)

Cells (-) epoxomicin

600,000 Cells (+) epoxomicin

500,000 400,000 300,000 200,000 100,000 0 No inhibitors

With inhibitors

Fig. 5 Use of inhibitors to reduce background (nonproteasomal) trypsin-like activity from serum-supplemented culture medium and cells. Samples containing culture medium without cells (gray bars), U266 cells (25,000 cells/well in 96-well plate) treated with DMSO vehicle (black bars), or U266 cells treated 2.5 h with 5 μM epoxomicin (hatched bars) to inhibit proteasome activity were assayed using Z-LRR-aminoluciferin to measure the trypsin-like activity of the proteasome either in the absence (no inhibitors) or in the presence of inhibitors to reduce background trypsin-like activity. Luminescence was recorded 15 min after reagent additions using a Promega GloMax 96 luminometer and plotted without any subtractions. (Reprinted from [32])

2 2.1

Materials Equipment

1. White-walled multiwell plates. Solid bottom plates are optimal for enzyme assays, and clear bottom plates are optimal for cellular assays. 2. Multichannel pipette or automated pipetting station for delivery of Proteasome-Glo™ Reagent. 3. Plate shaker for mixing multiwell plates. 4. Luminometer capable of reading multiwell plates (e.g. GloMax™ Microplate Luminometer). 5. Fluorimeter capable of reading multiwell plates (e.g. LabSystems Fluoroskan Ascent). 6. 37 °C incubator with 5 % CO2.

2.2

Reagents

1. Proteasome-Glo™ Chymotrypsin-Like Assay, ProteasomeGlo™ Trypsin-Like Assay, and Proteasome-Glo™ CaspaseLike Assay (Promega, Madison, WI) for in vitro assays with purified proteasome.

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2. 20S proteasome enzyme (i.e. ENZO, Plymouth Meeting, PA, or Boston Biochem, Cambridge, MA) or 26S proteasome (ENZO). 3. 10 mM HEPES buffer, pH 7.6 (for proteasome dilution). 4. Proteasome-Glo™ Chymotrypsin-Like Cell-Based Assay, Proteasome-Glo™ Trypsin-Like Cell-Based Assay, and Proteasome-Glo™ Caspase-Like Cell-Based Assay (Promega, Madison, WI) for assays with cultured cells. 5. The proteasome inhibitors: lactacystin, clasto-lactacystin-βlactone, and epoxomicin. 6. Dimethylsulfoxide (DMSO). 7. Suc-LLVY-AMC (i.e. Calbiochem). 8. Boc-LRR-AMC (i.e. ENZO). 9. Ac-nLPnLD-AMC (i.e. ENZO). 10. Apo-ONE® Homogeneous Caspase 3/7 Assay (Promega).

3

Methods The methods described below outline (1) assays for monitoring all three catalytic activities of the proteasome using purified 20S proteasome, (2) assays for monitoring all three catalytic activities of the proteasome using cultured cells, (3) inhibition studies using both formats, and (4) a protocol for multiplexing the bioluminescent, chymotrypsin-like, cell-based proteasome assay, and a fluorescent assay for caspase activity.

3.1 Detection of Proteasome Activity Using Purified 20S Proteasome

Directions are given for performing the Proteasome-Glo™ Assays in a total volume of 100 μl using 96-well plates and a luminometer. However, the assays can be easily adapted to different volumes providing the 1:1 ratio of Proteasome-Glo™ Reagent volume to sample volume is preserved (e.g., 25 μl sample + 25 μl ProteasomeGlo™ Reagent in a 384-well format).

3.1.1 Proteasome-Glo™ Reagent Preparation

1. Thaw the Proteasome-Glo™ Buffer and equilibrate both buffer and the lyophilized Luciferin Detection Reagent to room temperature prior to use. 2. Reconstitute the Luciferin Detection Reagent in the amber bottle by adding the appropriate volume of Proteasome-Glo™ Buffer (10 ml each for cat. no. G8621, G8631, G8641; 50 ml each for cat. no. G8622, G8632, or G8642). The Luciferin Detection Reagent should go into solution easily in less than 1 min. 3. Thaw the appropriate substrate and equilibrate to room temperature prior to use. For the Chymotrypsin-like Assay, use the

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Suc-LLVY-Glo™ Substrate; for the Trypsin-like Assay, use the Z-LRR-Glo™ Substrate; and for the Caspase-like Assay, use the Z-nLPnLD-Glo™ Substrate. A slight precipitate may be observed. Mix well by vortexing briefly. 4. Prepare the Proteasome-Glo™ Reagent by adding the Proteasome-Glo™ Substrate to the resuspended Luciferin Detection Reagent as per Table 1. Label the reagent bottle to identify the substrate used. 5. Allow the Proteasome-Glo™ Reagent to sit at room temperature for 30 min prior to use. This allows for the removal of any contaminating free aminoluciferin. Although free aminoluciferin is not detected by HPLC, it is present in trace amounts. 3.1.2 Proteasome-Glo™ Assay Conditions

Prepare the following reactions to detect proteasome activity (or inhibition of activity) in purified enzyme preparations: ●

Blank: Proteasome-Glo™ Reagent + vehicle control for test compound or inhibitor, if used.

Table 1 Instructions for making the Proteasome-Glo™ Reagent and final substrate concentrations Proteasome-Glo™ assay

Volume substrate added (μl)

Substrate concentration in reagent (μM)

Cat. No.

Substrate

Chymotrypsin-like assay

G8621

Suc-LLVY-Glo™

50

40

Chymotrypsin-like assay

G8622

Suc-LLVY-Glo™

250

40

Trypsin-like assay

G8631

Z-LRR-Glo™

100

30

Trypsin-like assay

G8632

Z-LRR-Glo™

500

30

Caspase-like assay

G8641

Z-nLPnLD-Glo™

50

40

Caspase-like assay

G8642

Z-nLPnLD-Glo™ 250

40

Chymotrypsin-like cell-based assay

G8660, G8661

Suc-LLVY-Glo™

50

40

Chymotrypsin-like cell-based assay

G8662

Suc-LLVY-Glo™

250

40

Trypsin-like cell-based assay

G8760, G8761

Z-LRR-Glo™

100

30

Trypsin-like cell-based assay

G8762

Z-LRR-Glo™

500

30

Caspase-like cellbased assay

G8860, G8861

Z-nLPnLD-Glo™

50

40

Caspase-like cellbased assay

G8862

Z-nLPnLD-Glo™ 250

40

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●

Positive Control: Proteasome-Glo™ Reagent + vehicle control + purified proteasome enzyme (20S or 26S). Assay: Proteasome-Glo™ Reagent + test compound + purified proteasome enzyme (20S or 26S) (see Notes 1 and 2).

The blank is used as a measure of any background luminescence associated with the test compound vehicle and the Proteasome-Glo™ Reagent and should be subtracted from experimental values. The positive control is used to determine the maximum luminescence obtainable with the purified enzyme system. Vehicle refers to the solvent used to dissolve the inhibitor or test compound used in the study (see Note 3). 3.1.3 Proteasome-Glo™ Standard Assay (96-Well, 100 μl Final Reaction Volume)

1. Add 50 μl Proteasome-Glo™ Reagent to each well of a white 96-well plate containing 50 μl blank, control, or test sample. If reusing tips, be careful not to touch pipette tips to the wells containing samples to avoid cross-contamination (see Note 4). 2. Gently mix contents of wells using a plate shaker at 500– 700 rpm for 30 s. Incubate at room temperature for 10 min–3 h depending on convenience of reading time. Maximal signal is reached typically within 10–30 min using purified 20S proteasome (see Fig. 2). At this time, sensitivity is optimal. Temperature fluctuations will impact the luminescent readings; if the room temperature fluctuates too much, a constant-temperature incubator may be desired (see Note 5). 3. Record luminescence with a plate-reading luminometer as directed by the manufacturer.

3.1.4 Determining Inhibition Curves for the Three Proteasome Catalytic Activities

1. When generating IC50 curves for a proteasome inhibitor, titrate the inhibitor in HEPES (10 mM, pH 7.6) and add 25 μl per well in a 96-well plate. Dilute 20S or 26S proteasome in the same buffer to 2 μg/ml and add 25 μl per well for a final proteasome concentration of 1 μg/ml. 2. Incubate at room temperature for 1 h to allow irreversible inhibitors such as lactacystin and epoxomicin to bind completely (see Note 6). 3. Add 50 μl Proteasome-Glo™ Reagent, containing Suc-LLVYGlo™, Z-LRR-Glo™, or Z-nLPnLD-Glo™, to each well of a white 96-well plate. 4. Gently mix contents of wells using a plate shaker at 500– 700 rpm for 30 s. Incubate at room temperature for at least 10 min. 5. Record luminescence with a plate-reading luminometer. Luminescence can be read at various times (see Fig. 6 and Table 2).

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Fig. 6 Inhibition of proteasome with clasto-lactacystin-β-lactone. Inhibitor titrations and 26S proteasome were combined in 96-well plates as described above. All three proteasome activities were tested with clasto-lactacystin-β-lactone, the active analog of lactacystin [38]. Inhibition curves using the Proteasome-Glo™ Chymotrypsin-like Assay demonstrate consistent IC50 values when readings are taken between 10 min and 18 h after addition of the Proteasome-Glo™ Reagent. Within 10 min, the dynamic range is maximal

Table 2 IC50 values for clasto-lactacystin-B-lactone on all three proteasome activities Inhibitor IC50 (μM) Substrate

Clasto-lactacystin β-lactone

Suc-LLVY-Glo™

0.02

Z-LRR-Glo™

0.76

Z-nLPnLD-Glo™

2.6

Values were calculated from readings taken at 60 min. The relative potencies are consistent with previous reports [33, 36]

3.2 Detection of Proteasome Activity from Cultured Cells

This protocol provides instructions for performing the ProteasomeGlo™ Cell-Based Assays in a total volume of 200 μl using 96-well plates and a luminometer. However, the assays can be easily adapted to different volumes if the 1:1 ratio of Proteasome-Glo™ CellBased Reagent volume to sample volume is preserved (e.g., 25 μl sample + 25 μl Proteasome-Glo™ Cell-Based Reagent in a 384-well format). If you are using the Proteasome-Glo™ Trypsin-Like or Chymotrypsin-Like Cell-Based Assays and are preparing assay plates using trypsinized cells, follow the protocol for cell trypsinization and preparation to avoid contamination with trypsin or chymotrypsin in the plated cells (see Note 7).

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3.2.1 Proteasome-Glo™ Cell-Based Reagent Preparation

1. Thaw the Proteasome-Glo™ Cell-Based Buffer, and equilibrate both the buffer and the lyophilized Luciferin Detection Reagent to room temperature before use. 2. Reconstitute the Luciferin Detection Reagent in the amber bottle by adding the appropriate volume of Proteasome-Glo™ Cell-Based Assay Buffer (10 ml each for cat. no. G8660, G8661, G8760, G8761, G8860, or G8861; 50 ml for cat. no. G8662, G8762, or G8862). The Luciferin Detection Reagent should go into solution easily in less than 1 min. 3. Thaw the appropriate substrate and equilibrate to room temperature prior to use. For the Chymotrypsin-like Cell-Based Assay, use the Suc-LLVY-Glo™ Substrate; for the Trypsin-like Cell-Based Assay, use the Z-LRR-Glo™ Substrate; and for the Caspase-like Cell-Based Assay, use the Z-nLPnLD-Glo™ Substrate. A slight precipitate may be observed. Mix well by vortexing briefly. 4. Prepare the Proteasome-Glo™ Cell-Based Reagent by adding the Proteasome-Glo™ Substrate to the resuspended Luciferin Detection Reagent as per Table 1. For the Proteasome-Glo™ Trypsin-Like Assay only, add Inhibitor 1 and Inhibitor Mix 2 as detailed in Table 3. After adding Inhibitor 1 to the resuspended Luciferin Detection Reagent the inhibitor will appear white and cloudy; however, upon mixing the resulting solution will clarify. After adding each inhibitor, rinse pipette tip several times. Mix to homogeneity by swirling the contents or inverting the bottle. Label the reagent bottle to identify the substrate used. 5. Allow the Proteasome-Glo™ Cell-Based Reagent to stand at room temperature for 30 min before use (see Subheading 3.1.1).

3.2.2 Trypsinization and Cell Preparation

1. From the parent T-75 cm2 flask of cells destined to be used, remove medium to waste, and rinse flask with D-PBS (without calcium and magnesium). 2. Add minimal (0.5–1.0 ml) amount of prewarmed trypsin:EDTA solution to flask surface and incubate until cells detach. 3. Add 9 ml of complete medium (containing serum) to cell suspension, mix, and pellet cells by gentle centrifugation.

Table 3 Instructions for making the Proteasome-Glo™ Reagent for the trypsin-like cell-based assay Proteasome-Glo™ assay

Cat. No.

Substrate

Volume substrate added (μl)

Inhibitor 1 (μl)

Inhibitor mix 2 (μl)

Trypsin-like cellbased assay

G8760, G8761

Z-LRR-Glo™

100

15

100

Trypsin-like cellbased assay

G8762

Z-LRR-Glo™

500

75

500

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107

4. Remove medium to waste, and wash the cell pellet with 12–15 ml of complete medium. Pellet cells by gentle centrifugation. 5. Remove medium to waste, suspend cell pellet in medium, count, and adjust to desired density. Cells are now ready to be plated. 3.2.3 Controls and Assay Conditions

Prepare the following reactions to detect proteasome activity (or inhibition of activity) using cells in culture: ●

●

●

●

Blank: Proteasome-Glo™ Cell-Based Reagent + culture medium (without cells) and vehicle control used. No-Treatment Control: Proteasome-Glo™ Cell-Based Reagent + culture medium containing cells and vehicle control (without test compound). Inhibitor Control: Proteasome-Glo™ Cell-Based Reagent + culture medium containing cells with a specific proteasome inhibitor such as lactacystin or epoxomicin (see Notes 6 and 8). Test: Proteasome-Glo™ Cell-Based Reagent + culture medium containing cells with test compound.

The blank is used as a measure of background luminescence contributed by the cell-culture medium, the vehicle used to deliver test compounds, and the Proteasome-Glo™ Cell-Based Reagent and should be subtracted from all control and assay values. Vehicle refers to the solvent used to dissolve the inhibitor or test sample used in the study. The no-treatment control is used to determine the maximum luminescence obtained from untreated cells. The inhibitor control is used to determine the maximum inhibition of proteasome activity and helps identify nonspecific protease activity not related to the proteasome. Test samples represent the cells with their respective treatments. 3.2.4 Proteasome-Glo™ Cell-Based Standard Assay (96-Well Plates)

1. Prepare the appropriate Proteasome-Glo™ Cell-Based Reagent for the desired catalytic activity as described above and mix thoroughly before starting the assay. 2. You may need to optimize cell number and treatment duration for each cell line. For a 96-well plate format, we recommend working with approximately 10,000–20,000 suspension cells per well or 5,000–10,000 adherent cells per well (see Note 9). 3. For consistent results, equilibrate assay plates to a constant temperature before performing the assay (see Note 5). 4. Use identical cell numbers and volumes for the assay and control reactions. 5. If preparing multiple plates, replicate controls on each plate. 6. Add 100 μl Proteasome-Glo™ Cell-Based Reagent to each 100 μl sample and appropriate controls as needed. Cover the plate with a plate sealer or lid.

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7. Mix the contents of the wells at 500–700 rpm using a plate shaker for 2 min. Incubate at room temperature for a minimum of 10 min. 8. Measure the luminescence of each sample in a plate-reading luminometer as directed by the manufacturer. 3.2.5 Determining Inhibition of the Proteasome in Cultured Cells

1. Add cells to 96-well plates in a 90 μl volume at 5,000–20,000 cells per well (see Note 9) and allow the cells to equilibrate for 2 h (suspension cells) or overnight (attached cells) at 37 °C, 5 % CO2. 2. Serial dilute the inhibitor in culture medium and add 10 μl per well, including a no inhibitor control. 3. Incubate the inhibitor with the cells for 1–2 h. 4. Add the Proteasome-Glo™ Cell-Based Reagent, shake the plate, and measure the luminescence as above (see Figs. 7 and 8).

3.2.6 Multiplexing the Proteasome-Glo™ Chymotrypsin-Like, Cell-Based Assay with a Caspase Activity Assay

1. Add cells in culture medium to 96-well plates in 90 μl per well and allow to equilibrate overnight at 37 °C, 5 % CO2. 2. Add inhibitor or test drug to cells in 10 μl per well and incubate for various times at 37 °C, 5 % CO2. 3. Remove plates from the incubator and allow them to equilibrate to room temperature. 4. Prepare the Proteasome-Glo™ Chymotrypsin-like, Cell-Based Reagent as above. 5. Prepare a modified Apo-ONE® Reagent by adding the ApoONE® substrate 1:20 into the Apo-ONE® Buffer. This gives a 10× Apo-ONE® Homogeneous Caspase 3/7 Reagent. 6. Add the Proteasome-Glo™ Chymotrypsin-like, Cell-Based Reagent to the cells at 100 μl per well. 7. Incubate for 15 min and measure luminescence on a platereading luminometer. 8. Add 20 μl per well of the 10× Apo-ONE® Reagent. 9. Shake the plates for 2 min at 700 rpm and incubate at room temperature for 30 min. 10. Measure fluorescence at 485/527 nm (see Fig. 9).

4

Notes 1. The final concentration of 20S proteasome should be within the linear range of the assay (see Fig. 3). With the enhanced sensitivity of the bioluminescent assays, less 20S proteasome is typically needed for the assays. We recommend defining the linear range for the particular proteasome preparation.

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Fig. 7 Inhibition profiles for epoxomicin and bortezomib for all three protease activities of the proteasome. U266 cells (20,000 cells/well in 96-well plates) were treated with epoxomicin or bortezomib for 1 h at 37 °C. Plates were then equilibrated to 22 °C before addition of luminescent proteasome reagent for each protease activity. (a) Chymotrypsin-like activity measured using Suc-LLVY-aminoluciferin. (b) Trypsin-like activity measured using Z-LRR-aminoluciferin. (c) Caspase-like activity measured using Z-nLPnLDaminoluciferin. Luminescence was recorded using a Promega GloMax 96 luminometer 10 min after reagent addition, with data shown as a percentage of the untreated control samples. Results shown are the average ± SD from three independent experiments. Data fitted using GraphPad Prism software (GraphPad, San Diego, CA) [32]. Bortezomib (PS-341) was obtained from Millennium Takeda Oncology (Cambridge, MA). (Reprinted from [32])

2. SDS cannot be used as an activating agent for the bioluminescent assays. Although SDS is frequently used to enhance the chymotrypsin-like activity of the proteasome, it is detrimental to luciferase and is not necessary for these bioluminescent assays. Superior sensitivity is achieved even in the absence of SDS (see Fig. 3). 3. The chemical environment of the luciferase reaction will affect the enzymatic rate and thus luminescence intensity. Solvents used for various chemical compounds may interfere with the luciferase reaction and thus the light output from the assay. DMSO, commonly used as a vehicle to solubilize organic chemicals, has been tested at final concentrations up to 1 % in

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Fig. 8 Bortezomib inhibition of proteasome activities with multiple cancer cell lines. Various cancer lines were plated to contain 5,000–20,000 cells/well in 96-well plates and equilibrated before treating with bortezomib for 1 h at 37 °C. Plates were then equilibrated to 22 °C before addition of luminescent proteasome reagent for each site. (a) Chymotrypsin-like activity measured using Suc-LLVY-aminoluciferin. (b) Trypsin-like activity measured using Z-LRR-aminoluciferin. (c) Caspase-like activity measured using Z-nLPnLD-aminoluciferin. Luminescence was recorded using a Promega GloMax 96 luminometer 10 min after reagent addition. Data are shown as a percentage of untreated samples. For each cell line, results shown are the average ± SD from replicates wells (n = 4). Data were fitted using GraphPad Prism software [32]. Bortezomib (PS-341) was obtained from Millennium Takeda Oncology (Cambridge, MA). (Reprinted from [32])

the assay and found to have a minimal effect on light output. Libraries stored in DMSO are compatible with the bioluminescent proteasome assays. 4. Owing to the sensitivity of the Proteasome-Glo™ Assays, contamination with other luciferin-containing reagents can result in high background luminescence. Be sure that shared luminometers are cleaned thoroughly before performing this assay. Avoid workspaces and pipettes that are used with luciferincontaining solutions, including luminescence-based cell viability, apoptosis, or gene reporter assays.

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RLU (proteasome) or RFU (caspase)

8000 7000 6000 1.5hr proteasome 5000

4.5hr proteasome

4000

1.5 hr caspase 4.5 hr caspase

3000 2000 1000 0 0.0001

0.001

0.01

0.1

1

epoxomicin (µM)

Fig. 9 Sequential multiplex to determine proteasome and caspase-3/7 activity. H929 multiple myeloma cells were grown in RPMI-1640 containing 10 % FBS and 1 mM sodium pyruvate; they were added to 96-well plates (10,000 cells/ well) at 90 μl/well and incubated overnight at 37 °C, 5 % CO2. Epoxomicin was titrated in culture medium and added at 10 μl/well. Cells were incubated with the inhibitor for 1.5 or 4.5 h at 37 °C, 5 % CO2. The plate was removed from the incubator and equilibrated to room temperature before adding ProteasomeGlo™ Cell-Based Reagent. The plate was mixed and luminescence was recorded after 15 min. A 10× Apo-ONE® Homogeneous Caspase 3/7 Reagent containing a (Z-DEVD)2-Rhodamine 110 was then added at 20 μl/well and fluorescence was recorded after 30 min. Epoxomicin treatment for 1.5 h did not induce caspase 3/7 activity, but caspase 3/7 activity was induced after 4.5 h treatment with ≥0.04 μM epoxomicin

5. Environmental factors that affect the rate of the luciferase reaction will also affect the intensity of the light output and the stability of the luminescent signal. Temperature can affect the rate of this enzymatic assay and thus the light output. For consistent results, equilibrate assay plates to a constant temperature before performing the assay. For batch-mode processing of multiple plates, positive and negative controls should be included for each plate. Additionally, precautions should be taken to ensure complete temperature equilibration. 6. The Proteasome-Glo™ Assays are optimized for use with purified 20S or 26S proteasome and the Proteasome-Glo™ CellBased Assays are optimized for use with cultured cells. It may be possible to assay for proteasome activity in crude cell lysates using the luminogenic substrates, but controls for specificity should be included [41]. The Proteasome-Glo Trypsin-like

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Cell-Based Assay with the inhibitor mixture should be used to monitor trypsin-like proteasome activity in cell lysates. None of the substrates are uniquely cleaved by the proteasome; therefore, depending on the extraction method and level of purity of the proteasome, confirming specificity with specific inhibitors may be critical. Lactacystin and epoxomicin are natural inhibitors that are very selective for the proteasome [39, 40], and are most potent against the chymotrypsin-like activity, followed by the trypsin-like and caspase-like activities (see Table 2 and Fig. 6) [40]. Epoxomicin and lactacystin are the most selective commercially available inhibitors although lactacystin has been reported to inhibit cathepsin A and tripeptidyl peptidase II under certain circumstances [39]. 7. Following trypsinization, minute quantities of trypsin or chymotrypsin present in the resulting cell suspension used for plating and preparing assay plates, will seriously affect the assay results. It is important to follow the protocol to remove any contaminating trypsin or chymotrypsin. 8. Some cell lines may contain protease activity that cannot be inhibited using either lactacystin or epoxomicin. We recommend performing a proteasome-inhibitor control as well as an untreated cell control in each assay plate to help define the window of activity attributable to the proteasome. Uninhibitable activity is typically low and can be subtracted (see Fig. 7). 9. An appropriate number of cells should be used to stay within the linear range for the Proteasome-Glo™ Cell-Based Assay. For a 96-well plate format, we recommend preparing bioassays to contain approximately 10,000–20,000 suspension cells per well or 5,000–10,000 adherent cells per well. Cell number can be scaled accordingly when using smaller formats. Empirical determination of the optimal cell number and treatment duration for each cell line and plate format may allow the use of even fewer cells; proteasome activity may vary significantly depending on cell type.

Acknowledgements The authors wish to thank our colleagues at Promega Biosciences, Michael Scurria, Laurent Bernad, Bill Dailey, and James Unch, for synthesizing the bioluminescent proteasome substrates. We are indebted to Keith Wood and Dieter Klaubert for the homogeneous, bioluminescent assay concept. We also thank Kay Rashka, Sandra Hagen, Jeri Culp, Debra Lange, Brian McNamara, Anissa Moraes, and Pam Guthmiller for translating the concepts into products.

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Chapter 10 Laser Capture Microdissection for Gene Expression Analysis Mallikarjun Bidarimath, Andrew K. Edwards, and Chandrakant Tayade Abstract Laser capture microdissection (LCM) is an excellent and perhaps the only platform to isolate homogeneous cell populations from specific microscopic regions of heterogeneous tissue section, under direct microscopic visualization. The basic operations of the LCM system are based on (a) microscopic visualization of phenotypically identified cells of interest, (b) selective adherence of cells to a melting thermolabile film/membrane using a low-energy infrared laser (IR system) or photovolatization of cells within a selected region (UV system), (c) capturing or catapulting of structurally intact cells from a stained tissue section. RNA/DNA or protein can be extracted from the cell or tissue fragments for downstream applications to quantitatively study gene expression. This method can be applied to many downstream analyses including but not limited to quantitative real-time polymerase chain reaction (PCR), microarray, DNA genotyping, RNA transcript profiling, generation of cDNA library, mass spectrometry analysis, and proteomic discovery. The application of LCM is described here to specifically and reliably obtain a homogeneous cell population in order to extract RNA to study microRNA expression by quantitative real-time PCR. Key words Endometrial lymphocytes, Microdissection, PALM microbeam, RNA extraction, microRNA

1

Introduction Laser capture microdissection (LCM) is a versatile technology to procure cytologically and or phenotypically identified cell populations from a heterogeneous tissue [1]. Both the pathological and healthy tissues are comprised of various cell types or populations within the heterogeneous tissues in a multicellular organism [1, 2]. The molecular analysis of DNA, RNA, and protein extracted using LCM provides for an enhanced genomic or proteomic profiling of specifically targeted cell populations from complex tissue sections [3]. However, the molecular analysis of cell or tissue extract becomes less reliable if the relative abundance of the cell population of interest is low. LCM enables researchers to isolate pure

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_10, © Springer Science+Business Media New York 2015

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cellular subpopulations from cytological preparation and or live cell culture under microscope without contamination from surrounding cells [1]. Several methods have been reported over the decades in order to conduct an accurate molecular analysis of pure cell populations from complex tissue. In the 1970s, Lowry and Passonneau developed a biochemical microanalysis protocol, which used “freehand” microdisssection under direct microscopy of a tissue section [4, 5]. At the same time, others reported using manual tools, such as razor blades, needles, and fine glass pipettes to isolate cells of interest under the microscope [5]. However, these initial protocols were time consuming, tedious, did not allow for the precise control over area of interest identified, and required a high degree of manual dexterity [6]. Shibata proposed a microdissection procedure in 1993 which utilized an ultraviolet laser beam to destroy the DNA of cellular components of all undesired tissue by negative selection. A specific dye was used to protect the cell or area of interest from ultraviolet laser beam. However, this technique was only useful for DNA and not for other analytes that are susceptible to UV-light degradation [7]. In the mid-1990s, Emmert-Buck and colleagues developed the LCM system at the National institutes of Health (Bethesda, MD, USA). This system was initially designed for accurate and efficient dissection of cells from histological tissue sections of solid tumors [1]. Later in 1997, LCM technology was commercialized through a collaborative research and development agreement partnership with Arcturus Engineering Inc. (Sunnyvale, CA, USA) as the PixCell system [8]. PixCell series was the most widely used laser-based microdissection system. PixCell series is being heavily used in the “Cancer Genome Anatomy Project” (CGAP) sponsored by the National Cancer Institute [8]. Currently, Arcturus engineering Inc. has multiple generations of LCM system including PixCell IIe on the market. Recently they have commercialized Veritas microdissection system which has a combination of infrared laser and UV laser microdissecting possibilities. LCM remains the standard terminology for any laser based microdissection system, irrespective of the type of laser used [9]. Based on the type of laser and procurement of dissected tissues, LCM systems are generally divided into three classes: infrared (IR LCM, [1], ultraviolet (UV LCM, [10]), and combined IR-UV system [10, 11]. Arcturus PixCell II and PixCell IIe instruments are classified under IR system whereas PALM microbeam (Carl Zeiss), LMD6000 (Leica Microsystems), and mmi CellCut (Molecular Machines and Industries) are classified under UV LCM systems. UV systems often include a combination of UV laser microdissection and catapulting systems. Recent version of Arcturus Veritas microdissection instrument utilizes both IR and UV laser sources to cut and catapult the cells or tissues [12].

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Fig. 1 Principal components of laser capture microdissection consists of (a) Monitor connected to a personal computer installed with PALM ROBOSoftware, (b) Supplementary fluorescence unit, (c) Digital microscope interface, (d) Joystick, (e) LPC cut, (f) Laser unit, (g) inverted microscope, and (h) Control Unit

The principal of the LCM system is based on (a) visualization of stained cells under direct microscopy, (b) selective attachment of cells, using a low-energy infrared laser (IR system), to a melting thermolabile film made of an ethylene vinyl acetate (EVA) membrane or a UV based photovolatization of cells within a selected area, (c) laser-guided catapulting of microdissected cell population [1]. The EVA membrane has its absorption maximum near the wavelength of the laser. This results in membrane melting only in the vicinity of laser path and later expanding into the section filling small gaps in the tissue section [1]. The typical microdissected areas have a dark outer border and a clear center, indicating that the thermolabile polymer has melted and clear center containing cell or area of interest is intact [8]. Generally membrane slides are incorporated with dye which absorbs laser energy thus preventing damage to enclosed cell or tissue fragment as well as helps in accurately identifying the areas within melted polymer [8]. The standard LCM instrument consists of an inverted microscope, control box to monitor all operations, a low energy infrared diode and laser control unit, a mouse or joystick-controlled microscope stage with a chuck for slide immobilizaiton, UV source for microscopic observation, camera and a color monitor [1, 13, 14]. A personal computer is generally connected to the LCM microscope for laser control and other associated settings (Fig. 1). The cap or microcentrifuge tube, for collection of transferred isolated cells, is manufactured with a thermoplastic membrane on the bottom surface of cap. It has a diameter of 6 mm and tightly fits on microcentrifuge tube to enable successive cell lysate preparation [13, 14]. UV based LCM systems works on a totally different

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principle compared to infrared LCM systems. The UV laser is directed around the cells or area of interest on a tissue section mounted on membrane slide. The undesired tissue around the cell or area of interest ablates away by narrow beam of UV laser while preserving the cells of interest intact. Later these cells are catapulted into an adhesive cap suspended on a robotic transport arm [13, 14]. LCM can be easily adapted for downstream applications such as microRNA expression studies. microRNAs are small noncoding RNAs, of an average 22 nucleotides, function in transcriptional and posttranslational regulation of gene expression [3]. The significance of this group of small RNAs arises from the ever growing evidence that they regulate many biological processes, such as cellular differentiation, proliferation, angiogenesis, and apoptosis. Dysregulation of these miRNAs often results in abnormalities in many biological processes. The expression levels of microRNAs are fine-tuned and can reach more than 50,000 copies of one miRNA within single cell [3]. A small number of miRNAs are differentially expressed when normal and diseased tissues are compared for their miRNA signatures. These miRNAs are of high diagnostic value and have implications in biomarker studies. Many miRNAs share high sequence similarities between pre- and mature miRNAs or between mature miRNAs within families. This similarity among the miRNAs might produce false positive results as biologically inactive pre-miRNAs as well as contaminating bystander cells may falsify the signal [3]. Accurate measurement of miRNAs is technically challenging due to sequence similarity, and spatial and temporal variation in expression. A well-defined pure cell population which are at the same developmental stage may provide precise miRNA quantification. LCM provides a technology to isolate rare cell types and subsequent accurate measurement of tissue-specific miRNAs using Q-PCR. LCM followed by Q-PCR can be applied not only to frozen tissue sections but also on formalin-fixed paraffin embedded (FFPE) sections for miRNA profiling. In our laboratory, LCM followed by Q-PCR protocol has been optimized to study mRNA as well as miRNA expression in porcine endometrial lymphocytes, endometrial endothelium, trophoblasts [15, 16], and mRNA expression in immunostained dendritic cell-specific intercellular adhesion molecule-grabbing nonintegrin (DC-SIGN+) cells [17] and mouse uterine natural killer cells at various stages of gestation [18, 19]. 1.1 Advantages of LCM

1. Reliable technology to study gene expression in small number of precise population of target cell or area of interest from their in situ environment. 2. The original cellular architecture and morphology of transferred tissue fragment or cell on a membrane polymer is preserved. 3. Combined with its precision and versatility, LCM can be performed as quickly as regular imaging of a histological section.

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4. Does not require manual dexterity as it is a “no touch” technique which reduces contamination. 5. Accurate microdissection is possible without destroying the adjacent tissues which allows for sequential sampling of several tissue components from same slide. 6. Other microdissection techniques require a microcapillary or a needle tip to remove the isolated cells. 7. LCM can be documented with the help of built-in camera to record images of isolated cells in addition to the selected area of interest as well as residual tissue after microdissection. 8. LCM is easily applied for FFPE tissue material, which is one of the most widely practiced clinical sample preservation and archiving. 9. Downstream analysis of archived FFPE material after LCM has tremendously improved retrospective molecular studies and our understanding of disease pathogenesis. 10. Laser beam does not affect nucleic acid integrity or protein composition isolated from captured cells which can be successfully utilized for downstream applications such as Q-PCR and proteomic analysis. 1.2 Disadvantages of LCM

1. Complete dehydration of tissue section and absence of cover slip on membrane slide can affect the cellular integrity and morphology. 2. Cell or tissue of interest identification requires expertise and can only be performed with an experienced histologist. 3. Standard histo-chemical staining protocols require several hours, which can result in further degradation of RNA or proteins by ubiquitous RNases and proteinases. 4. The extracted RNA, DNA or protein is always in small quantity. 5. Failure to remove selected cells due to lack of adherence of cells to the thermolabile membrane or inappropriate laser setting can be a technically challenging variable to overcome. 6. Precision of LCM and purity of isolated cells is affected due to minimum laser spot size of 7.5 μm which is very common with older machines; however most recent generations of LCM instrument had overcome this limitation.

2

Materials

2.1 Preparation of Tissue Sections from Fresh or SnapFrozen Tissues

1. Optimal cutting temperature medium (OCT, I.e. Tissue-Tek® O.C.T.™ Sakura Finetek). 2. Cryomolds. 3. RNase inhibitor (for preparation of staining solutions).

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4. RNase away (for cleaning lab-wares and instruments). 5. Dry ice. 6. Frozen tissue block or Paraffin-embedded formalin-fixed blocks. 7. Disposable blades for cryostat. 8. 1.00 mm PEN Membrane Glass Slide (i.e. Carl Zeiss cat# 415190-9041-000). Slides are available in different thickness and covered with either polyethylene naphthalate or polyethylene terephthalate membrane. We recommended using a biochemically inert 1.0 mm PEN-membrane slide for the maximum recovery of DNA, RNA or protein. During the catapulting process, membrane acts as a backbone that enables the isolation of selected area without affecting the tissue morphology. 9. Desiccator-containing slide storage box. 2.2 Staining Reagents and Fixatives

All staining reagents (ethanol 70, 95 %, hematoxylin, eosin) are prepared using RNase/DNase-free water and RNase inhibitor 1. Harris hematoxylin. 2. Eosin-Y. 3. 100 % Ethanol Molecular Biology Grade. Graded alcohol prepared v/v in RNAse/DNAse-free dH2O (75, 95 %). 4. Xylene. 5. Hydrogen peroxide. 6. Bovine serum albumin. 7. Primary antibody of interest (in this chapter we describe staining with anti-human DC-SIGN). 8. Tris buffer saline: 0.15 M Sodium Chloride, 0.05 M Tris–HCl in deionized water, pH = 7.6 at 25 °C. 9. Appropriate antibody.

horse

radish

peroxidase-tagged

secondary

10. 1× Diaminobenzidine Working Solution. 2.3 Laser Capture Microdissection

1. LCM Platform (i.e. PALM Microbeam). 2. Tissue sections prepared on PEN Membrane Glass Slide. 3. 500 μL capacity RNase-free microcentrifuge tubes.

2.4 Quantifying microRNA Expression

1. Total RNA purification kit (in this chapter we describe the protocol using Total RNA purification kit from Norgen Biotek Corp.). 2. Reverse transcription kit (i.e. miscript II RT kit from Qiagen).

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3. SYBR Green PCR Master Mix (i.e. 2× QuantiTect, Qiagen). 4. 10× miScript Primer Assay (Qiagen, Mississauga, ON, Canada, Cat#MS00044100). 5. Thermal cycler. 6. Filter pipette tips (RNAse-, DNAse-, and pyrogen-free 10, 100, 200 μL). 7. Spectrophotometer to measure RNA concentration. 8. Incubator. 9. Pipettes (10, 100, 200 μL). 10. −80 °C freezer for RNA sample storage.

3

Methods The experimental workflow to quantify the microRNA expression using LCM is represented schematically in Fig. 2. Each step is discussed in detail as follows:

3.1 Tissue Processing for LCM

Various tissue processing methods are being used on a diverse range of samples. However, these methods vary according to downstream applications. Tissue specimens are generally fixed by snap freezing or a formalin fixation method. Snap freezing is recommended for optimal recovery of high-quality RNA or DNA. Formalin fixation (10 % formalin) preserves the morphology by completely penetrating the tissue. However, the quality of RNA, DNA or protein quality will be compromised due to crosslinking of these molecules with formalin. Recently, new protocols have been developed to extract RNA/DNA or protein from

Fig. 2 Workflow of LCM and gene expression analysis

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paraffin embedded archived specimens [5, 6]. The following section discusses only frozen specimen processing. 1. Collect tissue specimens using presterilized surgical instruments such as scissors, scalpel blade and forceps on RNase-free working platform. 2. Prepare a flat working space by placing a metal tray on dry ice in a Styrofoam box. 3. Fill a thin layer (1/3rd) of OCT medium into a suitable size cryomold placed on metal tray. 4. The dissected specimen is manually oriented in the OCT containing cryomold and quickly filled until specimen is completely immersed in OCT. 5. After approximately 2 min, OCT turns white after forming a mold around the specimen. 6. These cryomolds may be used immediately for sectioning or stored at −80 °C until required. 3.2

Sectioning

1. The cryostat microtome is precooled so that chamber and block temperatures attain −17 °C and −20 °C, respectively. 2. Clean the cryostat chamber including blade holder and antiroll plate with RNase away after removing an old blade. 3. The frozen cryomolds are immediately transferred on dry ice to cryostat chamber and allowed to condition for a few minutes. 4. The OCT is layered onto a chuck (specimen holder) and immediately the tissue block is pressed against the chuck with the help of the anvil. 5. Wait for 10 min until tissue block attaches to the chuck. Fix this assembly onto the cutting block. 6. Start cutting a series of sections of 10–12 μm thickness on a parallel cutting surface. 7. Continue cutting until several sections with inappropriate size and shape are sliced away and discard them. 8. Adjust the blade and cut the section of 7–8 μm thickness and carefully transfer this section onto RNase-free room temperature membrane slide. For better adherence of section, membrane covered side of the slide is pressed gently against the tissue section and place thumb onto opposite side of the membrane slide. The higher temperature of thumb, in comparison with the chamber, will provide a better attachment between the section and the membrane. 9. It is recommended to change the blade, if multiple specimens are handled at the same session. 10. Slides with frozen sections are kept in cryostat or on dry ice if staining is performed immediately. If not, they may be stored in slide box at −80 °C until required.

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3.3 Staining and HistoMorphological Review

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The staining protocols should be selected depending on the suitability for downstream applications. Rapid staining procedures involving H&E [15, 16] and Toluidine blue are described in Tables 1 and 2, respectively. These methods have been optimized in our laboratory for use in frozen sections. However, FFPE sections should be deparaffinized (2 min in xylene for two times, 1 min in 100 % ethanol, 1 min in 95 % ethanol, and 1 min in 70 % ethanol) prior to following the steps in Table 1. Staining station with graded alcohols and staining reagents should be arranged well before the procedure. It is advised not to reuse the staining reagents in order to protect the integrity of biomolecules in the cell or tissue fragment isolated from LCM. If downstream application involves the extraction of RNA/DNA or protein, RNase or protease inhibitor should be used to minimize Table 1 Staining protocol using hematoxylin and eosin Step Staining reagent

Time

Comment

1

70 % Ethanol

30 s

Tissue fixation

2

RNase/DNase-free water 15 s

Rehydration and OCT removal

3

Hematoxylin stain

Nuclei staining

4

RNase/DNase-free water 15 s

Excess hematoxylin removal

5

Eosin stain

Cytoplasm staining

6

RNase/DNase-free water 15 s

Excess eosin removal

7

70 % Ethanol

30 s

Dehydration

8

95 % Ethanol

30 s

Dehydration

9

95 % Ethanol

30 s

Dehydration

10

100 % Ethanol

30 s

Dehydration

11

100 % Xylene

5 min Removal of ethanol

15 s

15 s

Table 2 Staining protocol using toluidine blue Step Staining reagent

Time

Comment

1

Toluidine Blue solution

3 min

Permanently stains cells

2

RNase/DNase-free water 15 s

Rehydration and OCT removal

3

RNase/DNase-free water 15 s

Excess Toluidine Blue removal

4

75 % Ethanol

3 min

Dehydration

5

Air-dry or dry on heater

5 min

At 40 °C in incubator

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the degradation. The membrane slides should be blotted using absorbent paper between each step in order to prevent carryover of excess reagents. The stained sections on the slides should be air-dried as quickly as possible (approximately 5 min) and immediately proceed with LCM. Before starting the microdissection procedure, the sections on the slides should be briefly reviewed under microscope for the histology and to plan the LCM session (see Notes 1 and 2). 3.4

Immuno-LCM

3.5 Microdissection Procedure

Immuno-histochemical staining of the frozen sections prior to LCM provides a precise identification of immunophenotypically defined cell populations and is commonly referred as immuno-LCM. ImmunoLCM overcomes the difficulty in identification based on their morphology alone [19]. Using specific antibodies to label the cells, it is possible to identify as well as isolate pure population, even of identical morphology depending on their antigen expression [19–21]. Routine immunohistochemical staining procedures require prolonged incubation period, which causes a significant degradation or loss of RNA due to ubiquitous RNases. Immuno-LCM to identify and obtain DC-SIGN+ cells from porcine endometrial frozen sections is modified in our laboratory from a prolonged 48 h to rapid 45 min protocol using brown diaminobenzidine staining [17]. The detailed steps in the immunostaining procedure are provided in Table 3. At the end of staining protocol, slides should be allowed to air-dry for 3–5 min and proceed with LCM. In our laboratory, microdissection is performed using PALM Microbeam from Carl Zeiss Inc. This instrument features a program, PALM RoboSoftware, which enables the visualization of composite images of the stained tissue section. PALM RoboSoftware controls all operations in the system such as platform movement, objective selection, light intensity, exposure time, condenser control, white balance and contrast, autofocussing, slide and region of interest selection, camera specifications, and laser parameters like speed, thickness, and energy (Fig. 3). Stepwise instructions are provided below. 1. Switch on the computer, microscope and the laser 5 min before performing LCM. 2. Load the slides (maximum three slides, Fig. 4d) carrying frozen sections onto holding frame of the stage and allow it for few seconds to adjust to the room temperature. 3. Load the capped microcentrifuge tubes (Fig. 4a) onto the robotic arm to collect the cell or tissue fragments of interest. 4. Move the robotic arm over top of the tissue sections to allow for proper microscopic visualization.

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Table 3 Immunostaining protocol using diaminobenzidine Step

Staining reagent

Time

Comment

1

75 % Ethanol

30 s

Tissue fixation

2

RNase/DNase-free water

30 s

Rehydration and OCT removal

3

3 % Hydrogen peroxide

5 min

Incubation

4

Nuclease-free tris buffer saline

30 s

Rinse

5

2 % Bovine Serum Albumin

5 min

Blocking at room temperature

6

25 μg/mL Anti-human DC-SIGN antibody or Isotype control antibody

15 min

At 37 °C

7

Nuclease-free tris buffer saline

30 s

Rinse

8

1:500 Anti-mouse horse radish peroxidase-tagged secondary antibody

15 min

At 37 °C

9

Nuclease-free tris buffer saline

30 s

Rinse

10

Diaminobenzidine working solution

5 min

Incubation

11

Nuclease-free tris buffer saline

30 s

Rinse

3

Hematoxylin stain

5s

Nuclei staining

7

75 % Ethanol

30 s

Dehydration

8

95 % Ethanol

30 s

Dehydration

9

95 % Ethanol

30 s

Dehydration

10

100 % Ethanol

2 min

Dehydration

11

100 % Xylene

5 min

Removal of ethanol

5. The laser should be set accurately according to the requirements, typically UV-energy to 50, UV-focus to 60, and the laser speed to 25. Among the options for laser setting, set it for “cut” and for “Laser” set it to “LPC” for optimum cutting. 6. The sections should be quickly reviewed by choosing 10× objective lens to identify the area or cell of interest prior to dissection. 7. The inner side of the lid of microcentrifuge tube has to rest, facing directly, over the section of tissue when the robotic arm is moved onto the slide. 8. Make sure that laser focus and intensity are accurately set (Fig. 5) and change the objective to 40× in order to cut narrow refined sections. The laser focus and intensity should be refocused and recalibrated every time the objective is changed.

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Fig. 3 PALM ROBOSoftware layout. After launching the program, the above window appears showing the menu on top left corner and tool bar below. Frequently used major tools during the LCM include (A ) camera and display settings to autolive or autoconfigure, exposure time and to adjust the white balance. (B ) Microscope tools to set the inverted microscope with different objectives as well as fluorescence. (C ) Light source and condenser. (D ) Laser tools to manually set the energy, speed and focus of the laser beam. (E ) Laser key to start cutting with laser and status display. (F ) Color palette and graphic tool to mark the area of interest with color for laser cutting. Drawing tools are available in different shapes including free hand, line, rectangle, and oval

9. A microscopic region on the slide devoid of tissue section should be located to optimize the focus and intensity of laser beam. 10. Calibrate the alignment and position of the laser marker with respect to stage movement (Fig. 6); this is very important as an inappropriate alignment will not allow for specific isolation of cell populations. 11. Draw the square box, circle or a serpentine line using freehand tool selected from the graphic toolbar. Click the laser function after the initial settings of speed of 25 μm/s with laser energy to 50. Adjust the cut focus and width of laser once laser beam start dissecting the membrane. To increase or decrease the laser width, the laser cut function should be manipulated (Fig. 3). 12. Select “RoboLPC” from the dropdown menu to simultaneously cut and catapult the cell or area of interest. Draw a line around the area of interest using freehand drawing tool. Click start laser to cut and catapult the preselected specific area. The “AutoLPC” option should only be used to cut and

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Fig. 4 Images of (a) silicon coated adhesive cap (indicated by black arrowhead). (b) Cryomold (c) Optimum cutting temperature medium (OCT Cryomatrix) (d) polyethylene naphthalate coated PEN membrane slide

catapult sections on a glass slide. Poorly adhering specimens like brain sections may be cut and catapulted using only superfrost plus charged slides since these require high laser energy to catapult into adhesive cap. 13. The cell or tissue fragment is collected into an overhanging silicon layered adhesive cap (Fig. 4a). This silicon layer is completely dry and does not need any capturing liquid in the cap. 14. After the collection of desired number of cells, presence of catapulted sample should be double checked using “cap check” option (Fig. 7b). Cap check enables the visualization of catapulted cells by moving the underlying stage and leaving only the cap in the microscopy light path. A typical microdissection procedure involving isolation of lymphocytes from porcine endometrium, catapulting them into adhesive cap and Cap check is demonstrated in Fig. 8. 15. Press the “Home” button on the control pad associated with robotic arm to bring it back to the home position (Fig. 7b); remove and close the capped tube and proceed to the incubation prior to RNA extraction.

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Fig. 5 Laser thickness and intensity calibration before microdissecting the specific cells or area of interest during the laser capture microdissection. In the above picture, the thickness of the top laser cut is too broad which might obliterate or inaccurately microdissect the cells or area of interest. Bottom laser thickness is accurately set by manipulating the laser cut energy and focus. The thickness and intensity should be selected manually depending on thickness of section as well as cell or tissue of interest. Magnification: 400×, Scale Bar: 75 μm 3.6 Quality Control of LCM

Successful microdissection of cells or area of interest for downstream applications is associated with best quality control practices. The factors affecting the quality of LCM dissected material and possible solutions are discussed below. 1. During specimen handling and preparation, RNase- and proteinase-free conditions should be maintained. 2. Wear proper gloves and use RNase-free surgical instruments and associated dissecting materials. 3. Avoid thawing of specimen or cryomold by using dry ice during transportation. 4. All reagents associated with LCM procedure should be of molecular biology grade and prepared in RNase- and DNasefree water before processing samples. 5. Membrane slides are highly recommended for optimum and efficient recovery of LCM material (see Notes 3 and 4). 6. Filtered RNase/DNase-free pipette tips are advised for this procedure.

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Fig. 6 Calibration of the laser in relation to stage movement (a) Inaccurate or incorrect position of the laser beam (Blue rectangle) in relation to platform. The gap between blue line and actual cutting line can be noticed: this offset might damage the cells. (b) Accurate positioning of laser marker in relation to platform. Actual cutting line and blue marker line are overlapped. Magnification: 400×, Scale Bar: 75 μm (color figure online)

Fig. 7 PALM Navigator, Cap Mover II, and Element List windows are frequently used functions in laser capture microdissection. (a) PALM Navigator containing an image of slideholder (maximum three slides), provides overview of the sample or section of choice. Within the area of scanned image, a single click of mouse on any spot will take the stage to that spot rapidly and display the image on the monitor. (b) PALM CapMover II enables ROBOSoftware-guided automatic movement of caps or diffuser directly over the tissue section. (c) Element list consists of all elements and their properties in a tabular form. It provides a series of functions including laser, objective, color, number, name, and type of element

7. Tissue fixation is the most critical step in maintaining RNA/ DNA or protein integrity, which depends on penetration time, temperature of fixation and size of sample. These criteria should be considered while fixing the sections (see Note 5). 8. For high-quality yield of RNA and DNA, LCM should be performed as quickly and efficiently as possible. 9. Check each reagent and chemicals supplied with kits for visible precipitation.

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Fig. 8 Laser capture microdissetion of lymphocytes from frozen porcine endometrial sections at gestation day 50, rapidly stained with Toluidine Blue. (a) Using a circular drawing tool from taskbar of PALM RoboSoftware, lymphocytes were selected and circled in blue before microdissecting and catapulting into overhanging adhesive cap. (b) Marked empty circular spaces or areas of interest after microdissection of lymphocytes. (c) Presence of lymphocytes in the adhesive cap (obtained through CapCheck function) catapulted from the tissue sections. Magnification: 400×, Scale Bar: 75 μm

3.7 Incubation and Cell Lysate Preparation for Total RNA Extraction

The following procedure is adapted from manufacturer’s instructions and guides with the cell lysate preparation from frozen sections using Total RNA kit from Norgen BioTek Corp. FFPE material may also be used to prepare cell lysate. However, the RNA integrity of FFPE material is compromised due to RNA degradation. Preheat the incubator to reach 42 °C at least 30 min before this step. 1. The microcentrifuge capped tube containing microdissected cells or tissue fragments is added with 300 μL of “Lysis Solution” (or equivalent RNA lysis solution) and briefly vortexed for 10 s. 2. Incubate the capped tube by keeping it upside down for 30 min at 42 °C; after every 10 min, gently vortex the tube for 15 s and put it back into the incubator. 3. Following incubation, vortex the tube for 15 s. 4. The lysate should be added with 300 μL of 70 % graded ethanol and vortex briefly for 15 s to mix and proceed to total RNA extraction.

3.8 Total RNA Extraction

The following methods are adapted from the manufacturer’s instructions and guides using Total RNA kit from Norgen BioTek Corp. 1. Assemble RNA binding columns with flowthrough collection tubes. 2. Pipette up to 600 μL of the cell lysate prepared with 70 % graded ethanol into the column and spin down at 8,000 × g for 1 min. 3. Visually inspect the column for presence of any residual lysate, if the lysate volume has not passed through, centrifuge this assembly for an additional minute.

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4. Completely discard the cell debris/flow through and reassemble the binding column with collection tube. 5. Repeat the above steps (steps 3 and 4) for any left-over or extra lysate volume if required; this causes binding of all RNA to the membrane of the column (see Note 6). 6. An optional On-column residual genomic DNA that may affect downstream application may be removed using RNaseFree DNase. 7. Pipette 400 μL of “wash solution” (or equivalent wash solution) to the binding column and spin down at 8,000 × g for 1 min (see Note 7). 8. Again visually inspect the column for presence of any residual lysate and centrifuge for an additional minute if the lysate volume has not passed through the column (see Note 8). 9. Completely discard the cell debris/flow through and reassemble the binding column with collection tube. 10. Repeat column washing steps (steps 7, 8, and 9) for three times and discard the flow through lysate. 11. Completely remove the lysate or dry the resin by spinning down the assembly for 2 min and discard the collection tube. 12. Reassemble the binding column with RNase-free elution tube (1.7 mL microcentrifuge tube). 13. Apply 50 μL of “Elution Solution” (or equivalent elution solution) to the binding column and spin down for 2 min at 200 × g followed by 1 min at 14,000 × g. 14. Make sure entire 50 μL “Elution Solution” is passed through into the elution tube, if not; centrifuge the assembly for an additional minute at 14,000 × g. 15. A second elution (Repeat steps 13 and 14) is recommended in fresh elution tube for maximum recovery from traces of RNA bound to the column. 16. Measure the concentration and purity of total RNA (see Notes 9–11). 17. The total RNA should be used immediately or stored at −80 °C until required (see Notes 12–16). 3.9 cDNA Synthesis from microRNAs

The following procedure describes the reverse transcription of miRNA into cDNA using miscript II RT kit (Qiagen) as per the manufacturer’s instructions. For mature microRNA quantification using HiSpec buffer, the recommended starting RNA material ranges from 10 ng up to a maximum of 2 μg depending on the enrichment and number of target microRNAs measured. For simultaneous quantification of mature or precursor microRNAs, other noncoding RNAs and mRNA, it is recommended to use miScript Hiflex buffer and starting material in the range of 0.5 μg

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Table 4 Reverse transcription reaction components Component

Volume/reaction

5× miScript HiSpec Buffer or 5× miScript HiFlex Buffer

4 μL

10× miScript Nucleics Mix

2 μL

RNase-free water

Variable

miScript Reverse Transcriptase Mix

2 μL

Template RNA

Variable

Total volume

20 L

(for precursor microRNA detection) up to a maximum of 1 μg. It is recommended to prepare at least 10 % greater volume of master mix than required to overcome the loss due to residual reaction mix in the tube and pipetting error. 1. Thaw all buffers provided in the kit and template RNA on ice. 2. Briefly centrifuge each solution to collect residual drops of buffer attached to the sides and store on ice. 3. Take out the miScript Reverse Transcriptase Mix from −20 °C just before the preparation of reaction mix and place back into −20 °C freezer immediately. 4. Prepare and gently mix the reverse transcription reaction mixture containing miScript Hispec Buffer, miScript Nucleics Mix, and miScript Reverse Transcriptase Mix according to Table 4. 5. Add master mix of 8 μL to each 0.2 mL PCR tube followed by template RNA and make up the final volume of 20 μL by adding RNase-free water. 6. Gently mix, centrifuge for few seconds and set each tube on ice. 7. Tighly close the tubes and perform the reverse transcription reaction by incubating at 37 °C for 60 min and later at 95 °C for 5 min to inactivate the reverse transcriptase mix and store it on ice. 8. Measure the concentration and purity of cDNA. 9. Dilute the cDNA sample ten times with RNA/DNAase-free water immediately prior to RT-PCR or stop at this step and store the undiluted cDNA at −20 °C until further use. 3.10 microRNA Real Time PCR Reaction

This section describes the real-time quantification of mature microRNA or noncoding RNA using a target-specific forward primer (miScript primer assay), and a SYBR Green PCR master mix as per the manufacturer’s instructions (Qiagen). Although this

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Table 5 Reaction setup for real-time PCR Component

Volume/reaction (96-well)

2× QuantiTect SYBR Green PCR master mix 12.5 μL 10× miScript universal primer

2.5 μL

10× miScript primer assay

2.5 μL

RNase-free water

Variable

Template cDNA (added at step 3)

≤2.5 μL

Total volume

25 μL

procedure is optimized to use with various brands of light cyclers, our laboratory applies LightCycler 480 (Roche Diagnostics, Laval, QC, Canada) to measure microRNA expression. The final concentration of diluted cDNA should be within 50 pg–3 ng per PCR reaction. The accepted normalization controls (RNU6-2, SNORD 72, SNORA and RNU5A) should be selected based on stability of the expression across the tissues. The forward primers for specific target microRNA may be custom designed using sequences available in the miRBase version 19 [22] (Table 5). Stepwise instructions are as follows: 1. Thaw diluted template cDNA on ice and 2× QuantiTect SYBR Green Master Mix, 10× miScript Universal Primer, 10× miScript Primer Assay and RNase-free water to room temperature and gently mix. 2. Prepare and gently mix the reaction mix containing 2× QuantiTect SYBR Green Master Mix, 10× miScript Universal Primer, 10× miScript Primer Assay for each reaction according to Table 5. 3. If multiple reactions are planned, calculate each reaction components to be added and prepare a final reaction mix. 4. Gently mix the reaction mix and pipette accurate volumes into designated wells. 5. 10× miScript Primer Assay for individual-specific target microRNA should be added separately into designated wells of the PCR plate after pipetting the reaction mix. 6. Apply diluted template cDNA into each designated wells and make up the final reaction volume to 25 μL using RNase-free water. 7. Carefully seal the PCR plate with adhesive film and centrifuge at 1,000 × g for 1 min at room temperature; this will remove any existing air bubbles.

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Table 6 Cycling conditions for real-time PCR Step

Time

Temperature

Additional comments

PCR Initial activation step

15 min

95 °C

HotStarTaq DNA Polymerase is activated by this heating step.

Denaturation

15 s

94 °C

Annealing

30 s

55 °C

Extension

30 s

70 °C

Cycle number

45 cycles

3-step cycling

Perform fluorescence data collection

Melting Peaks 1.915 1.765

miR-150

-(d/dT) Fluorescence (465-510)

1.615 1.465 1.315 1.165 1.015 0.865 0.715 0.565 0.415 0.265 0.115 -0.035

54

56

58

60

62

64

66

68

70

72 74 76 78 Temperature [°C]

80

82

84

86

88

90

92

94

96

Fig. 9 Melt curve analysis showing only one melting peak (74.5–76 °C) from a specific amplification product (n = 3) of mature microRNA, miR-150 extracted from laser capture microdissected porcine endometrial lymphocytes. Dissociation curve analysis was performed using LightCycler LC 480 program

8. Place the sealed plate in the Real-Time Cycler and perform the PCR as per the standard cycling conditions provided in Table 6. 9. At the end of reaction, the PCR product may be stored at −20 °C or used for primer specificity confirmation and sequencing. 3.11

Data Analysis

Using LightCycler LC 480 built-in software, determine the specificity and identity of the individual PCR product by doing a dissociation curve analysis (Fig. 9). In order to determine relative

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quantification, define a baseline for amplification plot resulting from PCR reaction. Baseline is noise level in early amplification cycles where no fluorescence is detected in PCR products. Define the threshold by placing a threshold point just above the background signal; however, this point should be within lower half of the amplification plot. Construct a standard curve for target and control by preparing a tenfold dilutions. Plot the CT values (Y-axis) against the log of template amount (X-axis). Using standard curve and CT values, determine the amount of target and control in the test sample. Calculate the normalized amount of target by dividing the amount of target by amount of control. Set the normalized target amount as calibration value and calculate the expression level of each target by dividing the normalized target amount by calibration value. Calculate and report the fold change values for each target gene.

4

Notes 1. Make sure that the frozen sections on LCM stage should not get dry in order to protect RNA integrity of the sample. This might lead to firm attachment of tissue section to the membrane slide. Incomplete dehydration of frozen sections along with low-energy settings for laser beam makes microdissecting and catapulting of the cells difficult. In order to troubleshoot this problem, properly calibrate laser focus and intensity. 2. Always replace old with fresh stains after every 20–25 slides within treatment and group. Use separate staining jars for tissue sections from different experimental groups. 3. The possible reasons for inefficient recovery of RNA from sample are either poor quality RNA in starting material or degradation of RNA during extraction. Another reason may be inefficient detachment of cells from the adhesive cap. 4. The good quality RNA depends on source tissue, proper staining procedure, duration of fixation and time spent on LCM settings. 5. Fixatives such as paraformaldehyde or formalin may diminish the RNA quality, so it is highly recommended to use snap frozen tissues along with alcohol fixation method. 6. Poor quality RNA may not bind properly to the membrane in the RNA extraction assembly which might lead to low RNA yield. 7. Buffer concentrations of the extraction kit should be checked regularly to ensure optimum RNA yield. 8. Visually inspect the binding columns for any clogging which might affect the RNA yield.

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9. Proper and optimum incubation period is recommended as shorter incubation periods affect the RNA yield. 10. Poor quality adhesive capped tubes may not form tight seals with the cap. Leakage may occur due to inappropriate seal and especially when the tube is kept upside down with lysis buffer inside for incubation. 11. It is advised to place tissue sections near the middle third of the membrane slide. The extreme edges of the slide will not be within usable LCM area. 12. Various stains are suitable with LCM including H&E, Toluidine blue, Wright-Giemsa, Methylene blue, Methyl green and Cresyl Violet acetate. However, eosin is not usually required for visualization of cells or cellular cytoplasm during the LCM. 13. Alternatively stains may be prepared in 50 mL centrifuge tubes, which can accommodate two slides with stained sections facing opposite sides. This will prevent the wastage of stain if there are very few slides per LCM session. 14. It is possible to videograph whole LCM session; however, camera is sufficient to document the LCM isolated cells or tissue fragments. Click the camera button in the Inspect tool bar. The image will be saved in one folder specified by the user. Typical images required from each LCM session are: Before microdissection (selected area of interest using freehand tool), after microdissection (the empty spaces after the cell or area of interest has been catapulted into cap), and finally the cap check image (cells or tissue fragment present in the adhesive cap) to confirm that all cells are successfully catapulted into the cap. 15. It is possible to automatically count the number of cells isolated using element list and to determine the efficiency of LCM as well as spot size. 16. If the diameter of the dissected material exceeds the 1,000 μm, it is usually possible to collect using forceps rather than adhesive caps. References 1. Emmert-Buck MR, Bonner RF, Smith PD et al (1996) Laser capture microdissection. Science 274:998–1001 2. Bonner RF, Emmert-Buck M, Cole K (1997) Laser capture microdissection: molecular analysis of tissue. Science 278:1481–1483 3. Hoefig PK, Heissmeyer V (2010) Measuring microRNA expression in size-limited FACSsorted and microdissected samples. In: Monticelli S (ed) MicroRNAs and the immune system: methods and protocols, vol 667,

Methods in molecular biology. Humana Press, New York, NY, pp 47–63 4. Braakman RB, Tilanus-Linthorst MM, Liu NQ et al (2012) Optimized nLC-MS workflow for laser capture microdissected breast cancer tissue. J Proteomics 75:2844–2854 5. Esposito G (2007) Complementary techniques: laser capture microdissectionincreasing specificity of gene expression profiling of cancer specimens. Adv Exp Med Biol 593:54–65

Laser Capture Microdissection 6. Eltoum IA, Siegal GP, Frost AR (2002) Microdissection of histologic sections: past, present, and future. Adv Anat Pathol 9:316–322 7. Shibata D (1993) Selective ultraviolet radiation fractionation and polymerase chain reaction analysis of genetic alterations. Am J Pathol 143:1523–1526 8. Decarlo K, Emley A, Dadzie OE et al (2011) Laser capture microdissection: methods and applications. In: Murray G (ed) Methods in Molecular Biology, 2nd edn. Humana Press, New York, NY, pp 1–15 9. Edgley AJ, Gow RM, Kelly DJ (2010) Lasercapture microdissection and pressure catapulting for the analysis of gene expression in the renal glomerulus. Methods Mol Biol 611:29–40 10. Xiang CC, Mezey E, Chen M et al (2004) Using DSP, a reversible cross-linker, to fix tissue sections for immunostaining, microdissection and expression profiling. Nucleic Acids Res 32:e185 11. Ordway GA, Szebeni A, Duffourc MM et al (2009) Gene expression analyses of neurons, astrocytes, and oligodendrocytes isolated by laser capture microdissection from human brain: detrimental effects of laboratory humidity. J Neurosci 87:2430–2438 12. Tayade C, Edwards A, Bidarimath M (2014) Laser capture microdissection. The guide to investigations of mouse pregnancy, 1st edn. Elsevier, Amsterdam 13. Golubeva Y, Salcedo R, Mueller C (2013) Laser capture microdissection for protein and NanoString RNA analysis. In: Taatjes DJ, Roth J (eds) Cell imaging techniques: methods and protocols, 1st edn. Humana Press, Totowa, NJ, pp 213–257

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14. Espina V, Wulfkuhle JD, Calvert VS et al (2006) Laser-capture microdissection. Nat Protoc 1: 586–603 15. Tayade C, Black GP, Fang Y (2006) Differential gene expression in endometrium, endometrial lymphocytes, and trophoblasts during successful and abortive embryo implantation. J Immunol 176:148–156 16. Tayade C, Fang Y, Hilchie D et al (2007) Lymphocyte contributions to altered endometrial angiogenesis during early and midgestation fetal loss. J Leukoc Biol 82:877–886 17. Linton NF, Wessels JM, Cnossen SA et al (2009) Angiogenic DC-SIGN+ cells are present at the attachment sites of epitheliochorial placentae. Immunol Cell Biol 88:63–71 18. Tayade C, Fang Y, Black GP et al (2005) Differential transcription of Eomes and T-bet during maturation of mouse uterine natural killer cells. J Leukoc Biol 78:1347–1355 19. Tayade C, Hilchie D, He H et al (2007) Genetic deletion of placenta growth factor in mice alters uterine NK cells. J Immunol 178: 4267–4275 20. Fend F, Michael R, Emmert-Buck MR et al (1999) Immuno-LCM: laser capture microdissection of immunostained frozen sections for mRNA analysis. Am J Pathol 154:61–66 21. Nakamura N, Ruebel K, Jin L et al (2007) Laser capture microdissection for analysis of single cells. In: Thornhill A (ed) Single cell diagnostics: methods and protocols, 1st edn. Humana Press, Totowa, NJ, pp 11–18 22. Griffiths-Jones S (2004) The microRNA registry. Nucleic Acids Res 32(Database issue): D109–D111

Chapter 11 Using the Peggy Simple Western System for Fine Needle Aspirate Analysis Erik T. Gentalen and John M. Proctor Abstract Simple Western™ assays are capillary-based electrophoretic immunoassays, similar in scope to SDS-PAGE (molecular weight separation, “size”) and IEF (isoelectric focusing, “charge”) immunoblotting. The enhanced sensitivity and automation of the Simple Western makes it better suited to cancer diagnostics and research than the traditional Western platform. Because of its smaller sample volume requirements, primary cells, such as those obtained from fine needle aspirates (FNAs), and solid tumor slices may be used to generate quantitative comparable data. The Peggy™ instrument is capable of performing either size or charge assays on up to 96 samples in a single unattended run. Key words Fine needle aspirates (FNAs), Western, Immunoassay, Simple Western, Capillary electrophoresis, Isoelectric focusing

1

Introduction Protein analysis is a critical part of characterizing a tumor in both basic research and clinical diagnostics. Understanding altered signaling pathways is necessary to guide the drug development process and can potentially alter the course of treatment for patients [1]. Protein identification is still often done by Western blot, a workhorse of proteomics research which has not changed significantly in the 30 years since it was invented [2]. However, several shortcomings make the Western blot incompatible with clinical applications. The many steps that require hands-on manipulations lead to variation from run to run and make standardization a challenge. Also, the sample and protein concentration requirements are high relative to the small amount of sample available from fine needle aspirates (FNAs) and other primary cell sources. Peggy (Fig. 1a) was developed to simplify protein analysis and can separate proteins by either size or charge in a fully automated assay. The user simply prepares a microtiter plate with samples, antibodies, and a few other reagents, loads the plate into the

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_11, © Springer Science+Business Media New York 2015

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Fig. 1 Peggy. (a) Peggy instrument. (b) View inside the Peggy instrument showing five compartments where capillaries are shuttled to and from as the different steps of the assay are performed

instrument along with the necessary capillaries, and then waits for the run to complete with analyzed data to interpret. All the steps of the protein separation and immunoassay are performed inside a 5 cm long, 100 μm internal diameter capillary. The instrument moves the capillaries between function-specific trays (separation, immobilization, antibody incubation, detection, and washing) as shown in Fig. 1b. The small capillaries require only nanoliter volumes of sample and the automation enables a very high level of precision in the assay.

2

Materials

2.1 FNA Collection/ Lysis

1. Sample. 2. 2 mL syringe, 20 gauge needle. 3. Phosphate buffered saline. 4. RIPA lysis buffer: 25 mM HEPES pH 7.5, 150 mM NaCl, 1 % NP-40, 0.25 % Na deoxycholate, 10 % glycerol, phosphatase inhibitor cocktail, protease inhibitor cocktail.

2.2 Reagents/ Consumables

1. Peggy Size Separation Master Kit (ProteinSimple): Buffers, matrices, microtiter plates, capillaries, diluents, ladders, secondary antibodies, luminol-S, peroxide for size-based assays. 2. Peggy Charge Separation Master Kit (ProteinSimple): buffers, anolyte, catholyte, microtiter plates, capillaries, luminol, peroxide XDR for charge-based assays. 3. Ampholyte premix (ProteinSimple). 4. pI standards (ProteinSimple).

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5. Sample Diluent (ProteinSimple). 6. DMSO inhibitor (ProteinSimple). 7. Secondary antibodies and antibody diluent (ProteinSimple). 8. Primary Antibodies. 9. Water: 0.22 μm filtered, deionized. 2.3

Equipment

1. Peggy (ProteinSimple). 2. Centrifuge with microtiter plate adapter. 3. Heat block. 4. Standard microbiology tools: vortexer, microcentrifuge, pipets.

3

Methods The ProteinSimple Peggy instrument analyzes tumor and other protein samples through either size or charge separation and subsequent immunoassay. All the steps from sample loading to detection are automatically performed by Peggy as described in Fig. 2. Samples, antibodies, luminol/peroxide, and other reagents are loaded onto a 384 well plate, in sets of 12 wells, corresponding to the 12 capillaries run simultaneously on Peggy. It will multitask up to 8 sets of 12 capillaries at a time, for as many as 96 samples per run.

3.1 Sample Preparation

Following the procedure of Fan et al. [3], tumor samples are collected from mouse by fine needle aspirate. 1. Continuous negative pressure is applied to a 2 mL syringe with 20 gauge needle while 10 passes are made through a subcutaneous tumor (see Note 2). 2. Specimen is collected in PBS. 3. Specimen is lysed in RIPA lysis buffer (ProteinSimple) (see Note 1). 4. Concentration of lysates is established by BCA assay. Procedures for preparing lysate from intact tumor, other primary cell sources, and cell culture are available at http://www. proteinsimple.com/resources.html There are several differences in the sample preparation for size and charge assays, which are outlined in Subheadings 3.2 and 3.3 below.

3.2 Size Assay Preparation

For size-based analysis, samples are boiled in an SDS-containing buffer and separated in a polymer matrix, similar to SDS-PAGE. A biotinylated molecular weight ladder, run in the first capillary of the set of 12, is used to calculate apparent molecular weight for the proteins detected in the other capillaries. Every sample is prepared with three fluorescent markers to aid with the alignment of the sample capillary to the ladder capillary.

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Fig. 2 Outline of the Simple Western process. Sets of (12) 5 cm capillaries are automatically processed through each step. For a size-based assay, the capillary is loaded with separation and stacking matrix before the sample is loaded; for a charge-based assay, the capillary is completely filled with sample that has been mixed with ampholytes (~400 nL). Proteins are then resolved by electrophoresis and immobilized using UV light. Primary antibodies targeted to the protein of interest are flushed through the capillaries, followed by HRPlabeled secondary antibodies. Chemiluminescent substrate is loaded and signal is recorded on a CCD camera. The signal is then processed to produce electropherograms which are analyzed for peak identification. Electropherograms can be digitally converted into virtual “blots” for a more traditional view of the data (see Fig. 14)

3.2.1 Fluorescent Standard

Fluorescent standard is run in every capillary to help align to the biotinylated ladder in capillary 1. The following stock solution will be added to each sample as described in Subheading 3.2.4. Prepare the fluorescent standard as follows: 1. Add 22 μL water to lyophilized single-use tube of fluorescent standard (ProteinSimple). 2. Add 20 μL 10× Sample Buffer (ProteinSimple). 3. Add 8 μL freshly prepared 1 M DTT.

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4. Pipet up and down to resuspend the lyophilized material. 5. Store on ice until ready for use. 3.2.2 Biotinylated Ladder

1. In a fresh microcentrifuge tube, add 96 μL Ladder Resuspension Buffer (ProteinSimple) and 4 μL freshly prepared 1 M DTT. Vortex to mix. 2. Add 20 μL of this solution to lyophilized single-use tube of biotinylated ladder (ProteinSimple). Pipet up and down to resuspend the lyophilized material. 3. Transfer solution to 0.5 mL microcentrifuge tube. 4. Denature at 95 °C for 5 min. 5. Store on ice until ready for use.

3.2.3 Sample

1. Dilute lysate to 1.3 mg/mL in 1× Sample Buffer (ProteinSimple) (see Note 3). 2. Combine 1.25 μL fluorescent standard with 3.75 μL diluted sample lysate. Vortex to mix. 3. Denature at 95 °C for 5 min. 4. Vortex. Spin briefly in microcentrifuge. 5. Store on ice until ready for use.

3.2.4 Luminol-S/ peroxide

1. Combine 150 μL of Luminol-S and 150 μL of Peroxide in a 0.5 mL microcentrifuge tube. 2. Vortex. Store on ice until ready for use.

3.2.5 Bulk Reagents for Size-Based Separation Assays

Fill reservoir cups as follows (see Fig. 3): Wash buffer

30 mL

Running buffer

20 mL

Matrix Removal Buffer 20 mL

3.2.6 Other Plate Reagents

Primary antibodies should be diluted in Antibody Diluent Plus (ProteinSimple) to recommended or empirically determined concentration. Recommended concentrations for many antibodies are available in the ProteinSimple Antibody Database at http://www. proteinsimple.com/antibody/antibodies.html (see Note 6). Initial dilutions of 1:50 should be used if the concentration has not been optimized yet. Anti-rabbit and anti-mouse HRP labeled secondary antibodies, and Streptavidin-HRP are available from ProteinSimple in ready to use (1×) format. Separation matrix and Stacking Matrix are also supplied ready to use. Recommended volumes for all plate reagents are shown in Table 1.

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Fig. 3 Peggy resource tray. The resource tray will be loaded with capillaries and the bulk reagents outlined in Subheadings 3.2.5 and 3.3.3

Table 1 Reagent volume guidelines for plate layout in Fig. 5 Reagent

3.2.7 Plate Preparation

Amount (μL)

Biotinylated ladder

5

Prepared samples

5

Antibody diluent plus

20

Primary antibody

15

Streptavidin-HRP

15

Secondary antibody

15

Luminol-S/peroxide (HRP substrate)

15

Separation matrix

15

Stacking matrix

15

Water

30

A plate preparation guide (Fig. 4) is provided to aid with pipetting. There are two temperature zones on the plate for size-based analysis. The top portion of the plate (row A through row J) is kept at 10 °C throughout the run. The lower portion of the plate is at

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Fig. 4 Peggy plate prep guide. This plastic overlay fits on the 384 well plate to assist with pipetting the samples and other plate reagents

room temperature. Load samples, ladder, antibodies, and luminol peroxide mixture in the 10 °C portion of the plate (see Fig. 5 and Table 1) (see Notes 4 and 5). The Stacking Buffer and Separation Matrix should be loaded last in wells N7-N18 and O7-O18, respectively. These viscous reagents need to be kept at room temperature throughout the run. The separation matrix is especially viscous (~300 mPa s). Care should be taken to ensure that 15 μL is loaded into each well. It is recommended to use a repeating pipettor or reverse pipetting technique to ensure delivery of the correct volume. To reduce evaporation, load 30 μL of water into wells M5-M20, N5-6, N19-N21, O5-6, O19-20, and P5-20. Centrifuge plate at 750 × g for 5 min at room temperature to remove any air bubbles from the plate. 3.2.8 Compass Software Setup

In the Simple Western Compass software, select Assay mode (see Fig. 6), then File – New Assay – Peggy Size. This will open a default size-based assay. Modify the sample, antibody, and luminol/ peroxide names and locations to match the desired plate layout. If necessary, adjust the number of cycles and antibody incubation times. Other run parameters will not normally need to be adjusted. To change reagent names, highlight the appropriate wells and click

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Fig. 5 Suggested plate layout for the size-based assay. Samples, antibodies, Antibody Diluent Plus, and luminol peroxide are placed in the 10 °C region of the plate, while separation matrix, stacking matrix, and water are loaded into the room temperature region. Capillary 1 is reserved for Biotinylated Ladder and has its own reagent well designations: L—Biotinylated ladder, A—Antibody diluent plus, S—Streptavidin

Fig. 6 Default layout for new size-based separation assay

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Fig. 7 Well content edit window

Fig. 8 Example of run-specific edits to default assay for size-based separation

“Edit” to bring up well content edit window (Fig. 7). The name and attribute entered will be displayed in the template window. To add additional rows of reagents, highlight the rows in which you want to add reagents, then click “S” to designate it a sample row, “1” for primary antibody, etc. You can also drag and drop the rows in the existing plate layout to rearrange them. If necessary, adjust the number of cycles and antibody incubation times. Other run parameters will not normally need to be adjusted. For an example of a complete run edited as described here, see Fig. 8. Proceed to Subheading 3.4.

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3.3 Charge Assay Preparation

For charge analysis, samples will be mixed with solution-phase carrier ampholytes. The ampholytes will create the pH gradient used to focus the proteins at their isoelectric point (pI). A fluorescently labeled pI ladder is added to each sample for pI calculation of the protein of interest. No ladder capillary is required for charge assays.

3.3.1 Sample Preparation

Depending on the expected isoelectric point of the protein of interest, the user will choose the appropriate pH gradient from the ProteinSimple catalog or can purchase the ampholyte gradient separately to be added to Ampholyte-free Premix (ProteinSimple). A pI standard ladder will be chosen to match the gradient. When analyzing a protein for the first time, it is recommended to use the broad 3–10 gradient (“premix G2”, ProteinSimple) with pI ladder 1 (pI = 4.0, 4.9, 6.4, 7.0, 7.3). Once the ladder and gradient have been selected, the sample is prepared as follows: 1. Prepare Sample Diluent—Mix 46 μL of Sample Diluent (ProteinSimple) with 4 μL DMSO inhibitor (ProteinSimple). 2. Prepare Sample matrix—mix 176 μL premix G2 with 4 μL pI standard ladder, vortex for 30 s. 3. Dilute sample to 0.4 mg/mL in Sample Diluent from step 1. 4. Mix 12 μL premix + ladder solution prepared in step 2 with 4 μL diluted sample for a final lysate concentration of 0.1 mg/mL.

3.3.2 Luminol/ Peroxide-XDR

1. Combine 150 μL of Luminol and 150 μL of Peroxide-XDR in a 0.5 mL microcentrifuge tube. 2. Vortex. Store on ice until ready for use.

3.3.3 Bulk Reagents for Charge-Based Separation Assays

Fill reservoir cups as follows (see Fig. 3): Wash buffer

20 mL

Anolyte

15 mL

Catholyte

15 mL

3.3.4 Other Plate Reagents

Primary antibodies should be diluted in Antibody Diluent (ProteinSimple) to recommended or empirically determined concentration. Recommended concentrations for many antibodies are available in the ProteinSimple Antibody Database at http://www. proteinsimple.com/antibody/antibodies.html (see Note 6). Initial dilutions of 1:50 should be used if the concentration has not been optimized yet. Anti-rabbit and anti-mouse HRP labeled secondaries are available from ProteinSimple and should be diluted 1:100.

3.3.5 Plate Preparation

A plate preparation guide is provided to aid with pipetting. See Fig. 4. Recommended plate layout and volumes are shown in Table 2.

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Table 2 Plate layout and recommended volumes for charge-based assay Plate row Wells Reagent

Amount (μL)/well

A

1–12

Sample preparation containing 12 lysate + Premix G2 + pI standards

B

1–12

Primary antibody

20

C

1–12

Secondary antibody

20

J

1–12

Luminol/peroxide XDR

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Fig. 9 Default layout for new charge-based assays

There are two temperature zones on the plate. The top portion of the plate (row A through row J) will be kept at 4 °C throughout the run. The lower portion of the plate is at room temperature and is not used in a charge-based run. Load samples, antibodies, and luminol peroxide mixture in the 4 °C portion of the plate. Centrifuge plate at 750 × g for 5 min at 4 °C to remove any air bubbles from the plate. 3.3.6 Compass Software Setup

In the Simple Western Compass software, select Assay mode (Fig. 9), then File – New Assay – Peggy Charge. This will open a default charge-based assay. Modify the sample, antibody, luminol/peroxide names and locations to match the desired plate layout. To change

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Fig. 10 Real-time images of separation during a run. In charge (a) and size (b) based assays, fluorescent standards are loaded into each capillary for pI or molecular weight calculation. They can also be monitored in real time during the run as a diagnostic for run performance. In panel (a), there are five pI standards (4.9, 6.0, 6.4, 7.0, and 7.3) focusing in 12 capillaries. In panel (b), the size separation proceeds from right to left and standards of molecular weight 1, 29, and 180 kDa are visible in all 12 capillaries, showing the progress of the electrophoretic separation

reagent names, highlight the appropriate wells and click “Edit” (Fig. 7). The name and attribute entered will be displayed in the template window. To add additional rows of reagents, highlight the rows in which you want to add reagents, then click “S” to designate it a sample row, “1” for primary antibody, etc. You can also drag and drop the rows in the existing plate layout to rearrange them. If necessary, adjust the number of cycles and antibody incubation times. Other run parameters will not normally need to be adjusted. 3.4 Start Run (Size and Charge)

When the desired assay has been defined in Compass, select Start. The Compass software will prompt the user to save the new assay file and open the start run wizard. The wizard will guide the user through preparing the instrument—loading water, emptying waste, and loading the bulk reagents and sample plate. Once the run starts, the instrument will automatically proceed through all the steps in Fig. 2. The movement of the fluorescent standards through the capillaries can be monitored in real time through the Run Summary window, see Figs. 10 and 11. Additionally, the instrument calculates a schedule for all events in the run which can be viewed in the Run Summary—Status window (Fig. 11). This gives the user an estimated completion time for the run.

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Fig. 11 Run Summary window. The Run Summary window provides real-time images of electrophoresis and a detailed schedule of all events over the course of a run

Fig. 12 Assigning peak names in the Compass software

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Analysis

After the run is complete, the data will be bundled together in a . cbz file and available for analysis. The analysis steps include entering information about the run—such as which reference standards were used and how Compass should identify and name the detected proteins.

3.5.1 Naming Standards

The standards used, whether pI standards in the charge assay or molecular weight standards in the size assay, must be defined correctly for the analysis to proceed. After opening the cbz file, select the Analysis mode. Then, select Edit – Analysis to bring up the window in Fig. 13. Size assays will generally use “Biotinylated ladder 1” in capillary 1. If the ladder has been loaded into a different capillary, select the capillary number using the drop-down menu next to “Ladder Capillary” (Fig. 13a). The fluorescent standards MW 1, 29, and 180 are prepared in every sample and run in every capillary. These are used to align the “Biotinylated ladder 1” with each capillary’s data and determined the molecular weight of proteins detected in each capillary. For pI standards (Fig. 13b), set the appropriate ladder using the drop-down menu next to “Default”. This is the ladder that will be applied to all capillaries, and used for calculating the pI value for detected proteins in each capillary.

3.5.2 Naming Peaks

Once the standards are properly designated, the Compass software will proceed to identify the corresponding standard peaks within each capillary, and translate the X-axis of the electropherogram to isoelectric point or molecular weight. Detected proteins will be displayed as peaks in the electropherogram and will be assigned pI value or MW. These peaks can be assigned names as designated by the user. To assign peak names, the user opens the Analysis window (select Edit – Analysis and choose the Peak Names option). Create a peak group name under “Analysis Settings”. In the example in Fig. 12, there are two peak group names, “ERK1” and “ERK2”. For each peak group name, create a list of peaks within the rightmost panel (labeled “Analysis Settings: ERK1” in the figure). Designate the peak’s pI or MW and range (+/−) and a unique name for each peak. Continue creating peak groups under “Analysis Settings” and defining peak names and attributes in the rightmost panel. After all the peak groups have been created for an assay, designate when they should be applied to the data under “Apply Settings”. In this example, whenever the antibody is listed as “ERK1/2”, the ERK1 and ERK2 settings will be applied. One can set up peak naming conventions for many different antibodies in a single run. Once the settings are entered, the user can click “Export…” to save the analysis settings to a file as a template for import into future experiments.

Fig. 13 Defining standards in Compass software. (a) For size-based assays. (b) For charge-based assays

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Fig. 14 Lane View representation of size data

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Once the peaks have been named, Compass will automatically process the data and calculate peak area and height and organize data by peak name. As shown in Fig. 15, compass will show the named peaks on the electropherogram and create a table below, where the peak area will be displayed in raw numbers or as relative percent area within each peak group. The user can cut and paste these values from the table into a spreadsheet, or select “Export Tables…” from the File menu to create a text file containing the areas and heights of all named peaks in a run. This file can then be opened in Excel or another program for more detailed analysis.

Notes 1. Lyse the sample in as small a volume as possible. This will help in assay development, and also allows for further dilution of lysis buffers, which can affect the electrophoresis. 2. Chill all reagents and syringe and needle before collecting cells from the tumor. 3. For initial experiments, a lysate concentration of 1 mg/mL (size) or 0.1 mg/mL (charge) is recommended. Optimal protein concentration will depend on the expression level of the protein of interest and antibody performance, and should be determined empirically.

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Fig. 15 Analyzing peaks using the Compass software

4. When pipetting into plates, contact the bottom of the well with the pipet tip and slowly raise the tip as you push the plunger on the pipettor to minimize air bubbles at the bottom of the wells. Large air bubbles may not be removed by centrifugation and can cause failure to load reagent. 5. Always load at least 5 μL into each reagent well on the microtiter plate. This will ensure the capillary tip is below the liquid height when loading reagents. 6. For an up to date list of publications using the Simple Western systems, see http://www.proteinsimple.com/citations.html ProteinSimple, the ProteinSimple logo, Simple Western, Compass, and Peggy are trademarks and/or registered trademarks of ProteinSimple. Excel is a trademark of Microsoft. References 1. Bild AH, Yao G, Chang JT et al (2006) Oncogenic pathway signatures in human cancers as a guide to targeted therapies. Nature 439:353–357 2. Towbin H, Staehelin T, Gordon J (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure

and some applications. Proc Natl Acad Sci U S A 76(9):4350–4354 3. Fan AC, Deb-Basu D, Orban MW et al (2009) Nanofluidic proteomic assay for serial analysis of oncoprotein activation in clinical specimens. Nat Med 5:566–571

Chapter 12 Analysis of Autophagosome Formation Using Lentiviral Biosensors for Live Fluorescent Cellular Imaging Kevin Long, Chandra Mohan, Janet Anderl, Karyn Huryn-Selvar, Haizhen Liu, Kevin Su, Mark Santos, Matthew Hsu, Lucas Armstrong, and Jun Ma Abstract Autophagy, a highly regulated homeostatic degradative process, allows cells to reallocate nutrients from less important to more essential processes under extreme conditions of starvation. Autophagy also prevents the buildup of damaged proteins and organelles that cause chronic tissue damage and disease. Although a topic of great interest with involvement of multiple signaling pathways, there are limitations in real-time detection of the autophagic process. EMD Millipore has developed technologies where prepackaged, ready-to-use, high-titer lentiviral particles, “lentiviral biosensors,” encoding GFP- or RFP-tagged proteins provide a convenient and robust solution for fluorescent imaging of cells undergoing autophagy. Compared to nonviral transfection methods, lentiviral transduction, in many cases, offers higher transfection efficiency and more homogeneous protein expression, particularly for traditionally hard-to-transfect primary cell types. Lentiviral biosensors are ideal for use with fixed and live cell fluorescent microscopy, and are nondisruptive towards cellular function. GFP- or RFP-protein localization matches well with antibodybased immunostaining and demonstrates altered patterns of expression upon treatment with modulators of cell function and phenotype. Lentiviral biosensors provide a broadly effective, convenient method for visualization of cell behavior under a variety of physiological and pathological treatment conditions, in both endpoint and real-time imaging modalities. In this study, we focus on lentiviral biosensors containing GFP-LC3 and RFP-LC3 to study the formation of autophagosomes. Key words Lentivirus, Autophagy, Biosensor, LC3, Live cell imaging

1

Introduction Autophagy is a highly regulated homeostatic degradative process where cells destroy their own components via the lysosomal machinery and recycle them. Under extreme conditions of starvation, cells utilize this process to reallocate nutrients from the less important processes to more essential processes required for survival. In eukaryotes, autophagy functions solely as a degradative and remodeling pathway, whereas in yeasts it also plays a role in

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_12, © Springer Science+Business Media New York 2015

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biosynthesis. Generally, three types of autophagic processes have been recognized. They are chaperone-mediated autophagy, microautophagy, and macroautophagy. Chaperone-mediated autophagy involves the direct translocation of cytosolic proteins across the lysosomal membrane. It requires cytosolic and lysosomal chaperones to unfold substrates. During microautophagy, cytoplasm is sequestered directly at the lysosomal surface by separation and/or invagination of the lysosomal membrane. In macroautophagy the sequestering of membrane is distinct from lysosome and it involves the formation of autophagosome that fuses with lysosome, which provides the hydrolytic enzyme machinery. Correlation between autophagy and apoptotic cell death has become an emerging topic of great scientific interest, especially in the field of tumor biology. On one hand, autophagy may induce cell death by degrading essential components, but on the other, it may facilitate survival of cancer cells under unfavorable metabolic conditions. Hence, cancer cells, with mutated Bcl-2, may survive chemotherapy by employing a protective autophagic process. A better understanding of autophagy will allow us to develop therapeutic agents to either increase or decrease the extent of this process. A number of disease states, including those where mutant proteins cause pathological changes, could become target of autophagy inducing agents. Some of the examples include Parkinson’s disease, Huntington’s disease, and Alzheimer’s disease where undesirable aggregates of proteins are possibly causative factors of disease. 1.1 Lentivirus-Based Biosensors for Subcellular Visualization

Analysis of the dynamics of subcellular structures has been revolutionized in the past 15 years by the development and refinement of genetically encoded fusions between fluorescent proteins and cellular structural proteins. Such fusion proteins, if designed properly, incorporate into the structure of interest without disturbing its function, and permit visualization of the structure in live cells and in real time by fluorescence microscopy [1]. The cDNAs encoding the fusion proteins have traditionally been delivered into cells by chemical transfection or electroporation. However, such transient transfection procedures have drawbacks, including highly variable expression levels and low efficiencies for transfecting primary cells. Selection of clonal cell lines stably expressing the construct of interest allows for optimized expression levels, but the process is time-consuming and is not feasible for primary cells. Fortunately, recently developed viral gene delivery vectors, such as lentiviral and adenoviral vectors, permit transduction of virtually any cell type, at more tightly controlled expression levels. These viral vectors have been utilized successfully for expressing genetically encoded subcellular markers, with the aim of imaging specific cellular states and events for a variety of diseases [2] and are becoming a mainstay in cancer research [3]. Most often, researchers have to

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develop the lentiviral constructs themselves using standard molecular biology protocols that are outside the scope of this paper. See multiple sources online for lentiviral construction and basic transfection issues from sources like Protocol-online.org, etc. [4]. More recently, there are available a growing number of commercially prepackaged lentiviral particles encoding fluorescent fusion proteins with subcellular markers for cell fate, cytoskeletal structures, and adhesion, making cellular analysis considerably easier and routine (see Note 1). Modern lentiviral biosensors are packaged with third generation lentiviral packaging plasmids, which produce pseudoviral particles that have vanishingly low probabilities of pathogenicity [5] (see Note 2). The fluorescent proteins employed are TagGFP2 and TagRFP, which have been demonstrated to be monomeric for minimal interference with the function of the fusion partner proteins and have quantum yields comparable to fluorescent proteins from other species [6, 7]. In this chapter, we highlight lentivirus-based biosensors for visualizing the intriguing process of autophagy. The biosensors used involve a TagGFP2 and TagRFP fused at their C-termini to the autophagosome marker LC3. LC3 precursors, diffusely distributed in the cytosol, are proteolytically processed to form LC3-I. Upon initiation of autophagy, the C-terminal glycine is modified by addition of a phosphatidylethanolamine to form LC3-II, which translocates rapidly to nascent autophagosomes in a punctate distribution [8]. DNA constructs encoding fluorescent proteins fused to LC3 in conjunction with a GFP-LC3 control mutant are now widely employed for intracellular monitoring of autophagosome formation by fluorescence microscopy.

2 2.1

Materials Equipment

1. Eight-well glass chamber slides for fixed cell imaging, or chambered cover glasses for live cell imaging (Millicell® EZ Slide, Millipore). 2. CO2 Incubator. 3. Inverted wide-field fluorescent microscope with a 63× oilimmersion objective lens and illumination/filters appropriate for GFP or RFP visualization. 4. Flow cytometer (guava easyCyte™ 8HT flow cytometer, EMD Millipore). 5. Sterile pipette tips. 6. General cell culture ware for HUVEC cells.

2.2

Reagents

1. LentiBrite™ GFP-LC3, RFP-LC3, and GFP-LC3 Control Mutant Lentiviral Biosensors (EMD Millipore, Temecula, CA, USA). See Subheading 3.1 for construction comments.

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2. Earle’s balanced salt solution (EBSS). 3. Lysosomal inhibitor. 4. 3-Methyladenine (3-MA). 5. 3.7 % Formaldehyde in Dulbecco’s phosphate-buffered saline (DPBS). 6. Fluorescent staining buffer (DPBS with 2 % blocking serum and 0.25 % Triton X-100). 7. The LentiBrite™ GFP-LC3-II Enrichment Kit (EMD Millipore; Cat. No. 17-10230). 8. FlowCellect™ GFP-LC3 Reporter Autophagy Assay Kit (U2OS) (EMD Millipore; Cat. No. FCCH100181). 9. Human Umbilical Vein Endothelial Cells (HUVEC) (EndoGro™, EMD Millipore; Cat. No. SCCE001). 10. Human umbilical vein endothelial cell (HUVEC) growth medium.

3

Methods

3.1 Construction of Lentiviral Vectors Encoding Fluorescent Protein Fusions

LentiBrite™ GFP-LC3, RFP-LC3, and GFP-LC3 Control Mutant Lentiviral Biosensors are constructed as follows: 1. Obtain the cDNAs encoding TagGFP2 and TagRFP (Evrogen) and the cDNA encoding human LC3A residues 1–120, which represents the proteolytically processed, mature form of LC3A, cloned in-frame at the 3′ end of the fluorescent protein cDNA. The resulting fusion proteins, TagGFP2-LC3 and TagRFP-LC3, leave the C-terminal glycine (Gly120) of LC3 available for lipidation upon induction of autophagy. To generate a control mutant that does not translocate upon induction of autophagy, site-directed mutagenesis was employed to mutate LC3 Gly120 to alanine, which renders the protein refractory to lipidation. 2. Transfer constructs to pCDH-EF1-MCS (System Biosciences Inc.), a lentiviral vector containing the constitutive, moderately expressing EF1α promoter. Generate third generation HIV-based VSV-G pseudotyped lentiviral particles using the pPACKH1 Lentivector Packaging System from System Biosciences (see Note 3).

3.2 Cell Seeding and Lentiviral Transduction

1. Seed cells in growth medium onto eight-well glass chamber slides for fixed cell imaging, or chambered cover glasses for live cell imaging. Select seeding densities to provide for 50–70 % confluency after overnight culture (e.g., 20,000–40,000 cells/cm2). 2. The next day after seeding, replace medium with fresh growth medium.

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3. Dilute high-titer lentiviral stock 1:40 with growth medium, and add lentiviral volume to the seeded cells for the desired multiplicity of infection (MOI). MOI refers to the ratio of the number of infectious lentiviral particles to the number of cells being infected. Typical MOI values ranged from 10 to 40 (see Note 4). 4. Incubate infected cells at 37 °C, 5 % CO2 for 24 h. 5. After 24 h of lentiviral transduction, remove lentiviruscontaining medium and replace with fresh growth medium (see Note 5). 6. Disinfect all lentivirus-containing medium and plasticware in direct viral contact with 10 % bleach before disposal. 7. Culture cells for another 24–48 h, changing medium every 24 h. For autophagy experiments, cells are either left in growth medium or incubated in Earle’s balanced salt solution (EBSS) containing a lysosomal inhibitor for 2–4 h. In some cases, 5 mM 3-methyladenine (3-MA) can be included to inhibit autophagosome formation. 3.3

Live Cell Imaging

1. For live cell visualization, place the chambered cover glass in a temperature-controlled microscope stage insert. 2. Initiate imaging as rapidly as possible following addition of modulator. LC3-expressing cells are imaged in EBSS containing a lysosomal inhibitor. 3. Perform live cell imaging on an inverted wide-field fluorescent microscope with a 63× oil-immersion objective lens and illumination/filters appropriate for GFP or RFP visualization (Leica DMI6000B).

3.4 Cell Fixation, Staining, and Imaging

1. Fix cells for 30 min at room temperature with 3.7 % formaldehyde in Dulbecco’s phosphate-buffered saline (DPBS). During fixation and for all subsequent steps, protect cells from light to minimize photobleaching. 2. Rinse samples twice with fluorescent staining buffer (DPBS with 2 % blocking serum and 0.25 % TRITON® X-100). 3. For immunocolocalization studies, primary antibody in fluorescent staining buffer is added to each well for 1 h incubation at room temperature. 4. Samples are then rinsed three times with fluorescent staining buffer. 5. Incubate with fluorescent secondary antibody and DAPI (1 μg/ml) in staining buffer for 1 h at room temperature. 6. Rinse samples twice with fluorescent staining buffer and DPBS. Coverslip slides with mounting media containing anti-fade

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Fig. 1 Plasmid vs. lentivirus transfection in easy- and hard-to-transfect cell types. HeLa cells and HUVECs were transfected with a TagGFP2-tubulin-encoding construct, either utilizing plasmid DNA in conjunction with a lipid-based chemical transfection reagent or using LentiBrite™ lentiviral particles. Images were obtained via wide-field fluorescent imaging with a 20× objective lens (blue = DAPI nuclear counterstain, green = GFPtubulin). Lentiviral transduction resulted in higher transfection efficiency (particularly for HUVEC, for which plasmid transfection was unsuccessful) and GFP-tubulin signal of more uniform fluorescence intensity

reagent and No. 0 cover glasses (Ted Pella). Allow to cure as indicated by manufacturer. 7. Image mounted specimens on inverted wide-field (as above) or Leica DMI4000B confocal fluorescent microscope, utilizing illumination and filters appropriate for GFP, RFP, Cy5 (for immunocolocalization), or DAPI excitation and emission wavelengths. 63× oil-immersion objective lens are recommended (our results shown in this paper are using these optics). Select magnification, exposure settings, etc. as appropriate for desired resolution and signal intensity. DAPI counterstain may be visualized at excitation/emission: 358 nm/461 nm. 3.5 Lentiviral Transfection Validation

Using LentiBrite™ lentiviral biosensors, and Subheadings 3.2–3.4 above, we demonstrated the improved efficiency of gene delivery and homogeneity of gene expression achieved by lentiviral transduction. In Fig. 1, easily transfectable HeLa cells were transfected with GFP-labeled tubulin via plasmid (with a chemical transfection reagent) or via lentivirus. Lentivirally transduced cells demonstrated higher transfection efficiency (percentage of cells positive for signal, compared to total number of cells) compared to chemically transfected cells, as well as more homogeneous expression (compared to the range of high and low expressers in the plasmid-transfected population). For a typically “hard-to-transfect” primary cell type such as human umbilical vein endothelial cells (HUVEC), lentiviral transduction produced homogeneously bright signal in a significant proportion of cells, in contrast to plasmid transfection, which resulted in minimal GFP-tubulin expression.

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3.6 Specificity of Localization of Lentivirally Delivered GFP-LC3

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Genetically encoded biosensors for studying autophagy have become a widely employed technique since the first description of the use of GFP-tagged LC3 to detect autophagosome formation [5]. Although immunofluorescent detection of endogenous LC3 can be performed, genetically encoded biosensors achieve greater sensitivity for detecting changes in autophagosome formation. However, the use of transient transfection of plasmid DNA for expression of GFP-LC3 has been criticized for causing artifactual, autophagy-independent punctae, and the preferred approach is to use cell lines stably expressing the GFP-LC3 construct [9]. To determine whether lentiviral delivery of DNA encoding fluorescent proteins fused to LC3 avoids such artifacts, we produced lentivirus encoding TagGFP2-LC3 for transduction into a broad variety of cell types (Subheading 3.1). We found that lentiviral delivery of fluorescent protein-tagged LC3 allowed for accurate detection of autophagosome formation, as determined by (1) immunofluorescent colocalization of LC3, (2) live cell imaging of autophagosome formation, (3) use of a mutant version of LC3 that is resistant to lipidation and fails to localize to autophagosomes, and (4) use of autophagy inhibitor (Subheadings 3.2–3.4). To compare localization patterns of genetically encoded fluorescent proteins with antibody-based immunofluorescence, HeLa cells were lentivirally transduced with GFP-LC3. GFP-LC3expressing cells were either left untreated or subjected to starvation conditions to induce autophagy by incubation in EBSS. A lysosomal inhibitor was also included to prevent degradation of LC3-containing autophagosomes. Both the fluorescent protein and anti-LC3 antibody displayed diffuse nuclear and cytoplasmic signal under fed conditions and a punctate distribution following starvation (Fig. 2). Next, we analyzed time-dependent LC3 translocation following autophagic induction in live cells. By wide-field microscopy, lentivirally transduced cells were imaged every minute over the course of approximately 2 h following treatment. Full-length video is available at www.millipore.com/autophagyvideo. Autophagy was induced in GFP-LC3-transduced HT-1080 cells via EBSS/ lysosomal inhibitor starvation, resulting in accumulation of punctate LC3 in newly formed autophagosomes. As shown in Fig. 3, significant formation of autophagosomes was visible at 50 min. At 110 min, nearly the entire visible GFP signal was localized to autophagosomes. For an additional assessment of the specificity of the GFP-LC3 biosensor, we employed two controls: a mutant LC3 that is resistant to lipidation, and an autophagy inhibitor. In Fig. 4, cells were lentivirally transduced with TagGFP2-LC3 or a TagGFP2-LC3 non-translocating control mutant. Transduced cells were starved in EBSS with a lysosomal inhibitor, in the presence or absence of

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Fig. 2 GFP-LC3 fluorescent signal (green) colocalizes with signal from fluorescent staining using LC3 antibody (red ). HeLa cells were transduced with TagGFP2-LC3, and 72 h later, either left in growth medium or starved for 4 h in EBSS with a lysosomal inhibitor. Cells were subsequently fixed, immunostained, and imaged by wide-field microscopy. Starved, autophagic cells displayed punctate cytoplasmic LC3 distribution, in contrast to diffuse nuclear and cytoplasmic localization under fed conditions. Fluorescently tagged protein colocalized with staining obtained with anti-LC3

Fig. 3 Live cell time-lapse imaging of lentivirally transduced cells. HT-1080 cells were lentivirally transduced with TagGFP2-LC3, and imaged by oil-immersion wide-field microscopy in real time. The cells were starved in the presence of a lysosome inhibitor, and imaging was immediately initiated, with images obtained every minute for 2 h. Still-frame captures demonstrate formation of GFP-LC3positive discrete cytoplasmic punctae

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Fig. 4 Autophagy inhibitor 3-MA prevents translocation of GFP-LC3. HT-1080 cells were lentivirally transduced with TagGFP2-LC3 wild-type (GFP-LC3 wt) or TagGFP2-LC3G120A (GFP-LC3 mutant) at MOI of 20. Transduced cells were either left in growth medium or starved in EBSS with lysosome inhibitor in the presence or absence of 3-methyladenine (3-MA). Cells were fixed, mounted, and imaged by wide-field fluorescence microscopy. Cells transduced with wild-type GFP-LC3 no longer exhibited cytoplasmic punctae under starvation conditions in the presence of 3-methyladenine. In addition, cells transduced with a negative control mutant GFP-LC3 maintained diffuse nuclear and cytosolic distribution under all conditions

3-methyladenine, an inhibitor of PI3 kinase that blocks autophagosome formation. The mutant LC3 fusion protein did not translocate to a punctate cytoplasmic distribution upon starvation. Also, when starved in the presence of 3-methyladenine, both the wild-type and mutant LC3 fusion proteins displayed a diffuse distribution throughout the nucleus and cytoplasm, as typical of fed cells. 3.7 Analysis of GFP-LC3 Localization by Flow Cytometry

The LentiBrite™ Autophagosome Enrichment Kit can be used to prepare samples for high-sensitivity analysis of autophagosome formation in primary cells. 1. Incubate human umbilical vein endothelial cells (HUVEC) with lentivirus encoding TagGFP2-LC3 or TagGFP2LC3G120A (control mutant) at an MOI of 40 for 24 h. 2. After removal of the lentivirus, culture the cells for an additional 48 h. Cells can be either left in complete growth medium or incubated in EBSS containing a lysosomal inhibitor. 3. Detach cells with Accutase™ and permeabilize. Treat U2OS cells stably expressing TagGFP2-LC3 (FlowCellect™ GFP-LC3 Reporter Autophagy Assay Kit (U2OS)), in parallel as a positive control. 4. Analyze samples immediately on a Guava easyCyte™ 8HT or similar flow cytometer. Data shown here were analyzed with the InCyte™ Software Module for the Guava machine.

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Fig. 5 Lentiviral transduction enables analysis of autophagy in hard-to-transfect primary cell types. HUVEC and HuMSC were lentivirally transduced at an MOI of 40 with TagGFP2-LC3 or TagRFP-LC3, and fed or starved as in Fig. 2. Cells were then fixed and imaged by wide-field microscopy. The transduced fluorescent proteins displayed diffuse distribution in growth media and a punctate distribution following starvation-induced autophagy

3.7.1 Analysis of Autophagy in Difficultto-Transfect Cell Types by Imaging and Flow Cytometry

Primary cell types, including HUVEC and human mesenchymal stem cells (HuMSC), are traditionally considered difficult-totransfect cell types for plasmid DNA-based chemical transfection. LentiBrite™ lentiviral transduction is shown to be capable of inducing fluorescent protein expression in these cell types in Fig. 5. Both HUVEC and HuMSC were successfully transduced at high efficiency with TagGFP2-LC3 and TagRFP-LC3. As seen with the lentivirally transduced HT-1080 cell line in the previous figures, the fluorescent protein fusions in the primary cells expressed diffusely when cultured in growth medium, and adopted a punctate distribution following starvation in the presence of a lysosome inhibitor. To more accurately assess the extent of LC3 reporter redistribution in primary cells, we employed a flow cytometry assay in which the plasma membrane is selectively permeabilized such that free cytosolic fluorescent protein-tagged LC3 is released while autophagosome-bound LC3 fusion protein is retained. HUVECs were lentivirally transduced with TagGFP2-LC3 or TagGFP2LC3G120A (control mutant). The cells were subsequently starved of amino acids in the presence of a lysosome inhibitor or left untreated, then detached and either permeabilized (using the LentiBrite™ Autophagosome Enrichment Kit) or left intact. Upon permeabilization, the GFP-LC3 in starved cells was almost completely retained, but was greatly depleted in fed cells (Fig. 6 ). This result was similar to the pattern observed in U2OS cells stably expressing TagGFP2-LC3 (FlowCellect™ GFP-LC3 Reporter Autophagy Assay Kit (U2OS)). In contrast,

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Fig. 6 Analysis of GFP-LC3 localization in HUVEC by flow cytometry. HUVECs were lentivirally transduced with TagGFP2-LC3 wild-type (GFP-LC3 wt, top row ) or TagGFP2-LC3G120A control mutant (GFP-LC3 mutant, center row). U2OS cells stably expressing TagGFP2-LC3 wild-type were also analyzed (U2OS-GFP-LC3, bottom row ). Transduced cells were detached and either permeabilized to release free, cytosolic LC3 (green peaks) or left intact (gray peaks). After processing, the cells were analyzed by flow cytometry on a guava easyCyte™ 8HT instrument. Upon permeabilization, only TagGFP2-LC3 wild-type-expressing cells under starvation conditions display retention of the fusion protein, indicative of tight association of LC3 with autophagosomes

permeabilization caused a large reduction in TagGFP2-LC3G120A in both starved and fed cells. In U2OS cells transiently transfected with plasmid encoding TagGFP2-LC3, a very broad distribution of fluorescence was observed, and the shift upon permeabilization of fed cells was much less pronounced (data not shown).

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Notes 1. Lentiviral particles provided higher efficiency of gene delivery and more homogeneous expression of introduced proteins compared to nonviral transfection methods. Other available constructs (GFP or RFP) in the LentiBrite™ portfolio target foundational proteins associated with apoptosis, cell structure, and adhesion: calreticulin, α-tubulin, β-actin, vimentin, α-actinin, and paxillin. Ready-to-use structural biosensors offer a solution for researchers seeking to fluorescently visualize the presence/absence or trafficking of a protein, under normal, abnormal, diseased, or induced cellular states. 2. IMPORTANT SAFETY NOTE: Replication-defective lentiviral vectors, such as the third generation vector provided in this product, are not known to cause any diseases in humans or animals. However, lentiviruses can integrate into the host cell genome and thus pose some risk of insertional mutagenesis. Material is a Risk Group 2 and should be handled under BSL2 controls. A detailed discussion of biosafety of lentiviral vectors is provided in ref. [5]. 3. Whether using commercially available constructs or developing your own, lentivirus storage can be a source of error. Lentivirus is stable for at least 4 months from date of receipt when stored at −80 °C. After first thaw, place immediately on ice and freeze in working aliquots at −80 °C. Frozen aliquots may be stored for at least 2 months. Further freeze/thaws may result in decreased virus titer and transduction efficiency by about 30 % each event. In addition, at higher temperatures, cells might methylate some toxic sequences within 10–14 days. 4. Typical MOI values for high transduction efficiency and signal intensity are in the range of 10–40. For this LC3 target, some cell types may require lower MOIs (e.g., HT-1080, HeLa, human mesenchymal stem cells (HuMSC)), while others may require higher MOIs (e.g., human umbilical vein endothelial cells (HUVEC), U2OS). MOI should be titrated and optimized by the end user for each cell type and lentiviral target to achieve desired transduction efficiency and signal intensity. Increasing MOI while ensuring that cells are not overgrown (50–80 % is the range) seems to eliminate many reported issues. 5. An infection enhancer, e.g., Polybrene (EMD Millipore Cat. No. TR-1003-G), may also be utilized, if desired. The efficiency of retroviral infection is enhanced significantly, 100– 1,000-fold in some cells, by including polybrene during the infection. Polybrene with DMSO shock is also used to mediate DNA transfer into a variety of cell types, such as CHO, chicken embryo fibroblasts, NIH3T3 cells, and myeloid cells.

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Protocol: Recombinant retroviral stocks are prepared by adding 5 ml of growth medium with 5 % serum to a near confluent monolayer of transfected retroviral packaging cells in a 100 mm plate. After 24 h the medium is removed and filtered through a 0.45 μm filter. Cells to be infected with this recombinant retroviral stock are plated at 500,000 cells per 100 mm plate in 10 ml of complete medium. 24 h later, remove the growth medium from the cells. Infect cells with 2 ml of the viral supernatant (or a dilution of the virus stock into 2 ml) in the presence of 5 μg to 10 μg of polybrene per ml (final concentration). Incubate cells for 3–6 h at 37 °C. Add 8 ml of complete medium. Three days after infection, split the cells 1:5 into selection medium. References 1. Goldman RD, Swedlow JR, Spector DL (2009) Live cell imaging: a laboratory manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY 2. Coutu DL, Schroeder T (2013) Probing cellular processes by long-term live imaging—historic problems and current solutions. J Cell Sci 126:3805–3815 3. Fein MR, Egeblad M (2013) Caught in the act: revealing the metastatic process by live imaging. Dis Model Mech 6:580–593 4. Protocol Online. Your Lab’s reference book. http://www.protocol-online.org/ 5. Pauwels K, Gijsbers R, Toelen J, Schambach A, Willard-Gallo K, Verheust C, Debyser Z, Herman P et al (2009) State-of-the-art lentiviral vectors for research use: risk assessment and biosafety recommendations. Curr Gene Ther 9:459–474

6. Merzlyak EM, Goedhart J, Shcherbo D, Bulina ME, Shcheglov AS, Fradkov AF, Gaintzeva A, Lukyanov KA, Lukyanov S, Gadella TW, Chudakov DM (2007) Bright monomeric red fluorescent protein with an extended fluorescence lifetime. Nat Methods 4:555–557 7. Subach OM, Gundorov IS, Yoshimura M, Subach FV, Zhang J, Grüenwald D, Souslova EA, Chudakov DM, Verkhusha VV (2008) Conversion of red fluorescent protein into a bright blue probe. Chem Biol 15:1116–1124 8. Kabeya Y, Mizushima N, Ueno T, Yamamoto A, Kirisako T, Noda T, Kominami E, Ohsumi Y, Yoshimori T (2000) LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J 21:5720–5728 9. Klionsky DJ et al (2008) Guidelines for the use and interpretation of assays for monitoring autophagy in higher eukaryotes. Autophagy 4:151–175

Chapter 13 Optical Imaging of Ovarian Cancer Using HER-2 Affibody Conjugated Nanoparticles Minati Satpathy, Rafal Zielinski, Ilya Lyakhov, and Lily Yang Abstract Computed Tomography (CT), Ultrasound (US), and Magnetic Resonance Imaging (MRI) have been the mainstay of clinical imaging regimens for the detection of ovarian cancer. However, without tumor specific contrast enhancement, these imaging modalities lack specificity and sensitivity in the detection of small primary and disseminated tumors in the peritoneal cavity. Herein, we illustrate a fairly new near infrared (NIR) optical imaging approach developed in our laboratory for the noninvasive detection of ovarian tumors using a HER-2 targeted nanoparticle-based imaging agent in an orthotopic mouse model of ovarian cancer. We used multimodal imaging approaches to detect the disease accurately and rapidly by utilizing a single imaging agent, NIR dye-labeled HER-2 affibody conjugated iron oxide nanoparticles. This agent targets HER-2 receptors, which are overexpressed in ovarian tumors. This chapter outlines materials and methods for the: (1) production of HER-2 targeted nanoparticles; (2) establishment of an orthotopic human ovarian cancer xenograft model; (3) monitoring of tumor growth by bioluminescence imaging; (4) administration of targeted nanoparticles followed by NIR optical imaging for the detection of orthotopic ovarian cancers with targeted accumulation of the nanoparticle imaging probes. Key words Near infrared optical imaging, Iron oxide nanoparticles, HER-2 affibody, Orthotopic model of ovarian cancer, Bioluminescence imaging

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Introduction Peritoneal metastases are the major contributing factor for the increased mortality rate among ovarian cancer patients. Early diagnosis and accurate staging are the key determinants for the successful treatment of this disease. Recent advancement in nanotechnology and molecular imaging bring a ray of hope for the patients with advanced ovarian cancer. These technologies are emerging as powerful tools for the precise detection and identification of many cancers by using biomarker targeted contrast agents. Targeted imaging and personalized therapy should have a significant impact on overall management of ovarian cancer patients [1]. One such biomarker, HER-2/neu, is known to be up-regulated in about 25 % of ovarian cancer patients and is correlated with poor survival [2].

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_13, © Springer Science+Business Media New York 2015

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HER-2 positive tumors have aggressive tumor biology and are resistant to chemotherapy and molecular targeted therapy, such as monoclonal antibody against HER-2 or Herceptin [3]. Surgical resection of ovarian tumors is a common treatment for ovarian cancer. Current methods for identification of an ovarian cancer lesion in the peritoneal cavity rely on the gross appearance of the tumor tissues, which lacks specificity and sensitivity. The development of tumor targeted imaging agents and imaging methods will provide a useful means for sensitive detection of ovarian tumors by preoperative noninvasive imaging and intraoperative image-guided surgery. Recently, a novel affinity molecule, HER-2 affibody, has been developed that is small in size, highly stable, and engineered with a unique cysteine residue at the C terminus end for site specific labeling of imaging contrast agents [4]. More importantly, HER-2 affibody (ZHER2:342) specifically binds to HER-2, a different epitope than Herceptin [5], which makes it suitable for patients subjected to Herceptin treatment. Near infrared (NIR) fluorescence imaging is an emerging imaging approach receiving significant attention as a potential means to detect cancer by noninvasive or intraoperative optical imaging [6, 7]. This method is accomplished by labeling an optical contrast agent (e.g., NIR dye) to a targeting ligand that binds to a target molecule highly expressed in tumor cells or tumor microenvironment. Systemic delivery of targeted imaging probes leads to the accumulation of the probes at the tumor site due to the molecular recognition of the tumor biomarker. At present, optical imaging using nontargeted NIR dyes, such as indocyanine green (ICG), has been used in patients for various clinical applications [8]. A recent study revealed the detection of ovarian tumors in human patients by using fluorescein isothiocyanate-labeled folic acid probes that target to folate receptor highly expressed in ovarian cancers [9]. A number of other preclinical studies has used NIR optical imaging probe for in vivo tumor imaging [7, 10]. Currently the Food and Drug Administration (FDA) has approved two NIR fluorophore dyes: ICG and methylene blue (MB). However, a number of research groups are developing novel NIR molecular imaging agents for optical imaging of tumors. Magnetic iron oxide nanoparticles (IONPs) are potential multifunctional clinical tools for targeted imaging and drug delivery [11]. The iron oxide nanoparticle is biocompatible and biodegradable with low toxicity. Clinically nontargeted IONPs have been used as a contrast agent for magnetic resonance imaging in the detection of lymph nodes [12]. We have conjugated a NIR dye-labeled HER-2 affibody to amphiphilic polymer coated IONPs to produce a dual optical and MR imaging probe, NIR-830-ZHER-2:342–IONPs [13]. This multifunctional nanoprobe is capable of targeting HER-2 overexpressing ovarian tumors that is detectable by multimodal imaging methods such as NIR optical imaging, magnetic resonance imaging

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(MRI), photoacoustic tomography (PAT), fluorescence molecular tomography (FMT), and spectroscopic imaging technique in an orthotopic ovarian tumor mouse model [14]. This chapter outlines our methods of (a) production of recombinant HER-2 affibody; (b) preparation of NIR dye-labeled HER-2 affibody conjugated IONPs; (c) generation of an orthotopic mouse model for ovarian cancer; (d) detection of ovarian tumor progression by bioluminescence imaging (BLI); and (e) detection of orthotopic ovarian cancer by NIR optical imaging and related image processing techniques.

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2.1 Production of Recombinant HER-2 Affibody

1. HER-2 affibody (obtained from Affibody AB, Stockholm, Sweden on Collaborative Research and Development agreement); this protocol can also be used with other antibodies but the disulfide bonds in the protein has to be broken by appropriate reducing agent before labeling with the NIR amine ester dye. 2. Super Broth (Quality Biological, Gaithersburg, MD); LB broth can also be used but yield may be lower. 3. Ampicillin. 4. Isopropyl β-D-1-thiogalactopyranoside (IPTG). 5. Imidazole. 6. Protease Inhibitor Cocktail. 7. Dithiothreitol (DTT). 8. Tobacco Etch Virus (TEV) protease. 9. HIS TRAP column. 10. Protein purification and concentration filters (i.e., Amicon Ultra Sample concentration unit, EMD Millipore). 11. Modular liquid chromatography system (i.e., AktaPurifier10, GE Healthcare). 12. Centrifuge. 13. Sonic Dismembrator. 14. Temperature-controlled benchtop orbital shaker.

2.2 Preparation of Near Infrared Labeled HER-2 Affibody Conjugated Iron Oxide Nanoparticles

1. Iron oxide nanoparticles (core size 10 nm) coated with amphiphilic polymers (we use particles from Ocean NanoTech, LLC, Springdale, AR). 2. Maleimide dye (we use NIR-830 maleimide dye synthesized from IR-783 dye (Sigma-Aldrich)) using a synthesizing method as described in [15] but any commercially available can be used following this protocol.

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3. Borate buffer pH 5.5 and 8.6. 4. Tris[2-carboxyethyl] phosphine (TCEP). 5. Ethyl-3-dimethyl amino propyl-carbodiimide (EDAC). 6. Sulfo-N-hydroxysuccinimide (sulfo-NHS). 7. Ultrafiltration system (i.e., Nanosep 100 K column, Pall Corporation, Ann Arbor, MI). 8. Microfuge tubes. 9. Lab shaker. 10. Spectrophotometer. 11. Dynamic light scattering (DLS) instrument (i.e., Zetasizer Nano-ZS S-90, Malvern Instruments Inc., Southborough, MA). 2.3 Establishment of Orthotropic Human Ovarian Cancer Xenograft Model

1. 6–8 week old female athymic nude mice. 2. SKOV3 ovarian cancer cell line stably transfected with luciferase gene (SKOV3-luc) (kindly provided by Dr. Daniela Matei at Indiana University Purdue University at Indianapolis); any other cancer cell line overexpressing the antigen of interest (in our case HER-2) may be used and transplanted orthotopically as long as they express the luciferase reporter. 3. McCoy’s 5A culture medium with 10 % fetal bovine serum or appropriate culture medium. 4. Trypsin. 5. 1× PBS. 6. D-luciferin. 7. Tissue culture incubator maintained at 37 °C with 5 % CO2. 8. Water bath adjusted to 37 °C. 9. 15 ml conical tubes. 10. Hemocytometer. 11. 0.5 ml insulin syringe. 12. Ketamine hydrochloride and xylazine hydrochloride in sterile saline. 13. Autoclaved surgical instruments (scissors, forceps, surgical wound staples, ear punch, polypropylene suture 18″). 14. Bio shield sterile wrap super, sterile surgical sponges, surgical gloves, and mask. 15. Alcohol Prep pad and Povidone-Iodine prep pad. 16. Heating pad. 17. Digital camera. 18. Meloxicam (Metacam stock 5 mg/ml). 19. Mouse restrainer.

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IVIS in vivo imaging system (Xenogen Corporation, Caliper Life Sciences). Kodak in vivo imaging system Fx (Carestream Health Inc., New Haven, CT).

Methods Production of recombinant HER-2 Affibody molecule (ZHER2:342(GS)7-Cys) (see Note 1) involves: (a) overexpression and purification of MBP-TEV-6His-ZHER2:342-(GS)7-Cys construct (see Note 2); (b) proteolytic removal of MBP tag; and (c) purification of the final product. The plasmid pMTZC for expression of HER-2 Affibody molecules (ZHER2:342-(GS)7-Cys or ZHER2:342-GSC) was cloned as described in [16–18]. The yield of the protocol described below is approximately 4 mg/l.

3.1 Overexpression of MBP-TEV-6HisZHER2:342-(GS)7-Cys

1. Inoculate 100 ml of LB broth medium supplemented with ampicillin at 100 μg/ml with frozen stock of E. coli BL21(DE3) transformed with pMTZC plasmid. Grow bacterial culture O/N with intensive shaking at 37 °C. 2. Measure the OD600 of the O/N culture and inoculate 1 l of LB broth medium supplemented with ampicillin at 100 μg/ml. The initial OD600 should be 0.1–0.15 AU. Grow bacteria at 37 °C with intense shaking 180–220 rpm, periodically checking OD600. 3. Once bacteria have reached the density 1.0 AU, induce protein expression by adding IPTG to final concentration 1 mM, continue bacterial culture for 3 h. 4. Centrifuge bacteria 4,000 × g, 10 min, resuspend in 100 ml ice-cold PBS supplemented with complete protease inhibitor cocktail tablets, and sonicate in ice-bath 2 × 5 min using Sonic Dismembrator at 20 W. 5. Remove cell debris by centrifugation 48,000 × g, 20 min and filter the supernatant through 0.22 μm filter.

3.2 Purification and Proteolytic Removal of MBP Protein

1. Equilibrate HisTrap (5 ml) column with at least 20 volumes of ice-cold 50 mM phosphate buffer containing 300 mM NaCl and 10 mM imidazole, pH 8. 2. Load the supernatant onto the column at 0.5 ml/min using loading superloop (GE HealthCare). 3. Washout unbound proteins by running through the column icecold 50 mM phosphate buffer containing 300 mM NaCl and 10 mM imidazole until UV monitor returns to baseline level. 4. Elute resin-bound protein using linear gradient of imidazole from 0 to 300 mM in 50 mM phosphate buffer with 300 mM

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NaCl, pH 8 over 10× column volumes. Collect eluate in 0.5 ml fractions. Analyze fraction by SDS/PAGE. 5. Based on electropherogram, combine fractions containing MBPTEV-6His-ZHER2:342-GSC (expected molecular weight approximately 50 kDa). Exchange imidazole-containing buffer to PBS, using Amicon Ultracentrifugal filter units (30 kDa, cut off). 6. Add DTT to final concentration 1 mM followed by incubation with TEV protease at 0.1 mg/ml overnight at room temperature. Dilute the sample to 1 mg/ml of protein in PBS. 7. Repeat purification on HisTrap columns (steps 1–6), collect fractions based on electrophoretic migration (expect the band of approx. 10 kDa in reducing SDS/PAGE condition). Dilute protein to 1 mg/ml, pass through 0.22 μm filters using sterile PBS, aliquot in 0.5 ml. Store at −80 °C. 3.3 Preparation of NIR-830-HER-2 Affibody-IONPs

1. Prepare fresh 0.5 M TCEP in 1× PBS. 2. Add 1 μl of TCEP (stock 0.5 M) to 150 μg ZHER2:342-GSC (hereafter referred to as ZHER2:342) for a total volume of 100 μl in PBS, pH 7.4. 3. Incubate the mixture at room temperature for 15–30 min. 4. Prepare a 10 mM stock of NIR-830 maleimide (mol wt. 965.327) in DMSO. 5. Add fourfold molar excess of NIR-830 maleimide dye or commercially available NIR maleimide dyes over the concentration of ZHER2:342 directly to the mixture and incubate the mixture in a sealed tube at 4 °C overnight with rotation (see Note 3). 6. Take 400 μl of amphiphilic polymer coated iron oxide nanoparticles (Stock 5 mg/ml), activate with 200 M of EDAC and 100 M of sulfo-NHS in 10 mM Borate buffer at pH 5.5 according to the carbodiimide method, and then allow it to rotate for 15 min at room temperature (see Note 4). 7. Immediately after incubation, neutralize the reaction by adding 10 mM Borate buffer, pH 8.6, and wash the mixture by using a Nanosep 100 K spin column. 8. Add the bi-conjugate NIR-830-ZHER2:342 to the activated IONP (at a molar ratio ZHER2:342-IONP10:1) and allow it to incubate for 4 h at room temperature with rotation (see Note 5). 9. Purify the final product NIR-830-ZHER2:342–IONP (Fig. 1a) with 10 mM Borate buffer, pH 8.6, using Nanosep 100 K spin column to remove the reaction by-products. 10. Resuspend in 10 mM Borate buffer, pH 8.6, and store at 4 °C. 11. NIR-830-ZHER2:342–IONP generated by using this method is stable for at least 6 months at 4 °C. 12. Measure the concentration of iron in the resulting conjugates by taking the absorbance at O.D. 500 nm (see Note 6).

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Fig. 1 (a) Schematic illustration of optical imaging probe conjugated to iron oxide nanoparticles. (b) Hydrodynamic size measurement by dynamic light scattering (DLS) instrument

13. Measure the hydrodynamic size of NIR-830-ZHER2:342–IONP by using the dynamic light scattering (DLS) instrument by preparing a 1 mg/ml solution in water followed by a quick sonication (Fig. 1b). 3.4 Establishment of Orthotropic Human Ovarian Cancer Xenograft Model

1. Culture SKOV3-luc ovarian cancer cells in McCoy 5A medium supplemented with 10 % fetal bovine serum and 1 % penicillin and streptomycin, and then allow them to grow at 37 °C, 5 % CO2.

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2. When cells reach 80 % confluence, trypsinize and wash cells twice with culture medium at 200 × g for 5 min followed by a final PBS wash. Count cells using a hemocytometer (see Note 7). 3. Resuspend cells in PBS and transfer 6 × 104 cells in 200 μl PBS to each microfuge tube depending on the number of mice to be injected. Keep extra cells in tubes in case of loss or improper injection. 4. Spin down the cell suspension at 200 × g for 5 min and remove the supernatant carefully without dislodging the cells by using a pipette (critical step) leaving 15–20 μl PBS in the tube and immediately place the tube with cells on ice in preparation for orthotopic injection. 5. Each mouse will be injected orthotopically with 15–20 μl PBS containing at least 5 × 104 cells into the ovarian bursa (see Note 8). 6. Anesthetize the mice by injecting a mixture of 95 mg/kg ketamine hydrochloride and 5 mg/kg xylazine of body weight in sterile saline intramuscularly. 7. Place the mouse on sterile pad dorsally and disinfect the operating skin area with alternate povidone-iodine pad swab followed by an alcohol pad (three times). 8. Make an abdominal incision using a sharp scissor and forceps to expose the right ovary. Carefully look for fallopian tube using forceps to locate the ovarian bursa on the top of the fallopian tube. Gently lift the ovarian bursa along with the fat pad and inject 20 μl of the cell suspension using a 0.5 ml insulin syringe attached to a 27G needle into the bursa area (Fig. 2a–c, f) (see Note 9). 9. Close the abdominal incision in two separate layers. First close the peritoneal muscle followed by the outer skin using suture

Fig. 2 Stepwise illustration of orthotopic implantation of SKOV3-luc cells into ovarian bursa: (a) Location of right and left ovaries (blue arrowheads). (b) Surgical site. (c) Exposed implantation site and implantation of SKOV3-luc cells into ovarian bursa. (d) Closing of surgical site in layers using suture. (e) Completely sewed. (f) Fallopian tube with ovarian bursa (upper panel) and fallopian tube with the primary ovarian tumor (lower panel)

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(18″) (Fig. 2d–e). Punch the ear or use a number tag to identify individual mice for each experiment. 10. Move the mouse back to the cage and place the cage on the heating pad until the mouse is completely awake. 11. Administer pain medication, Meloxicam (metacam), 2 mg/kg immediately after surgery and then 24 and 48 h after the operation (as needed), examine the mice daily and monitor the tumor growth noninvasively by bioluminescence imaging (BLI) weekly. 3.5 In Vivo Bioluminescence Imaging (BLI) for Tumor Progression

1. Anesthetize the mice by injecting a mixture of 95 mg/kg ketamine hydrochloride and 5 mg/kg xylazine in sterile saline intraperitoneally. Start the BLI to monitor the tumor growth on week 2 after implanting the SKOV3-luc cells into the ovarian bursa (see Note 10). 2. Inject the luciferin substrate 30 mg/kg body weight intraperitoneally into the nude mice bearing SKOV3-luc tumor 10 min prior to imaging using a 27G, 0.5 ml insulin syringe (see Note 11). 3. Place the anesthetized mice inside the BLI system. Use the field of view at “B” position so that five mice can be imaged at a time. 4. Follow the instruction of the manufacturer from Caliper Life Sciences imaging system to capture the BLI. For luminescent image acquisition, use an integration time of 10 s and reduce or increase the exposure time (Fig. 3a) as per the intensity of the luminescence signal to avoid saturation. For comparative studies, capture multiple images weekly at the same exposure time (see Note 12). 5. For quantification of luminescence signal, select region of interest (ROI) by encircling tumor area as well as body background from displayed images using IVIS software. Calculate integrated flux of photons (total flux p/s) in each region and

Fig. 3 (a) Representative mouse showing longitudinal tumor progression by bioluminescence imaging; (b) Analysis of luminescence signal selecting region of interest using inbuilt software; (c) Tail vein injection of NIR-830-ZHER2:342-IONPs to the SKOV3-luc tumor bearing mouse

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subtract total flux p/s value of body background from tumor intensity (total flux p/s) every week to follow the longitudinal tumor progression (Fig. 3b). 6. At week 6, when tumor is >6 mm in size, inject 400 pm of NIR-830-ZHER2:342–IONP conjugate dissolved in PBS pH 7.4 in a total volume of 120 μl intravenously via the tail vein (Fig. 3c) of the mouse. 3.6 Near Infrared (NIR) Optical Imaging by Kodak In Vivo Imaging System Fx

1. Anesthetize the mice 24 h following the imaging probe administration by injecting a mixture of 95 mg/kg ketamine hydrochloride and 5 mg/kg xylazine body weight per mouse in sterile saline intramuscularly. 2. Inject luciferin (30 mg/kg) intraperitoneally to each mouse for noninvasive whole body bioluminescence imaging to determine tumor locations and sizes as described in Subheading 3.5. 3. Place the animal on a clear tray ventrally for NIR optical imaging. Acquire all the images with a filter set of excitation wavelength at 800 nm and emission wavelength at 830 or 850 nm, exposure time at 180 s with a gamma value of 0.2. Positioning of tumor bearing mouse is very critical for capturing the NIR images by Kodak in vivo imaging system FX (Carestream Health Inc., New Haven, CT) (see Notes 13 and 14). Follow other equipment settings as per the manufacturer’s instructions. 4. After acquiring NIR images in each position, acquire a corresponding X-ray image to provide a clear anatomical location of the tumor (Fig. 4a). 5. ROIs containing equal pixels are drawn for tumors, kidney area, corresponding body background and tray background using accompanied Kodak in vivo imaging system Fx software package (Fig. 4b). Mean fluorescence intensities (MFI) are computed from the accompanied software. Signal-to-noise ratios (SNR) are calculated by dividing the MFI of tumor by MFI of body background. It is expected to visualize some NIR signal in the kidney within 24 h of nanoparticle delivery due to the excretion of cleaved nanoparticle products. In addition, iron oxide nanoparticles are readily taken up by kupffer cells in the liver and macrophages in the spleen which render bright NIR signals in the liver and spleen ex vivo but their signals were very low in noninvasive NIR imaging. When comparing kidney signal with tumor signal is needed, subtract the body

Fig. 4 (continued) a representative mouse. Corresponding bioluminescence and white field images are also included; (d) Noninvasive NIR optical imaging of targeted delivery of nanoparticles (24 h) of a representative mouse whole body showing NIR signal intensities from dorsal, ventral, tumor cells injected site and noninjected site of the mouse along with gamma scale. Bioluminescence imaging of whole body indicating the tumor location of the corresponding mouse

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Fig. 4 (a) Representative mouse showing target specificity of the nanoparticles at the tumor site after 24 h of nanoparticle delivery; (b) Analysis of NIR optical signal about selecting the region of interest using inbuilt Kodak in vivo imaging FX software. Analysis window also showing a table detailed about all the parameters that depend on the goal of each study; (c) Ex vivo NIR imaging of primary (#1) and metastatic tumors (#2 and #3) from

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background from both the tumor site and noninjected site. Finally divide the MFI of tumor by MFI of kidney site to get a fold increase MFI in the tumor region. At least three to five mice are needed to calculate the standard deviation. 6. Delivery of NIR dye-labeled HER-2 targeted nanoparticles to small metastatic ovarian tumors is difficult to be visualized by noninvasive optical imaging. Although NIR signals can be detected on the optical images, they usually lack sufficient resolution to determine size and locations of the tumor lesions. After 24 and 48 h of nanoparticles delivery, capture the whole body imaging for specific accumulation of nanoparticles at the tumor site and subsequently euthanize the mice by cervical dislocation. 7. Open the skin and abdominal muscle, and place the mouse facing down on the tray to capture optical images of the exposed mouse organs. Immediately excise the tumors, normal organs and tissues for ex vivo organ imaging using NIR, BLI, and bright field imaging (see Note 13). 8. Keep blood and muscle samples for background correction. Primary and metastatic tumors from a representative mouse showing the targeted delivery of the nanoparticles (Fig. 4c). 9. Keep tumor and normal organ samples at −80 °C for future histological analysis. 10. Figure 4d illustrates the critical positioning of mice as well as the gamma scale adjustment in the Kodak in vivo imaging system Fx (see Note 14).

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Notes All animal experiments should be performed in guidance with the protocol approved by the appropriate Institutional Animal Care and Use Committee IACUC. Current animal experiment protocol has been approved by Emory University, IACUC, Atlanta, GA, USA. Ketamine can be obtained from the pharmacy with a special order from a licensed physician. 1. (GS)7-flexible linker is added to HER-2 affibody to lower the probability of steric interaction of the label attached to the C-terminal cysteine with the binding site. 2. Maltose binding protein (MBP) is used to increase the stability and yield of ZHER2:342-GSC. 3. HER-2 affibody contains a disulfide bond which needs to be reduced before labeling with a maleimide. TCEP reduces the disulfide bond effectively as compared to DTT. Since it does not contain thiols, it does not have to be removed from the solution before adding maleimide reaction chemistry.

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4. EDAC is used as a catalyst to improve the reaction activity of sulfo-NHS with COOH functional groups on the polymer coating of the nanoparticles. Conjugation is prepared via crosslinking of carboxyl group of the amphiphilic polymer coating to the amino side groups of ZHER2:342 affibody. 5. HER-2 affibody is small in size and removal of the reducing agent after the reduction step may cause lose of some HER-2 affibody resulting in poor conjugation, so avoid washing the bi-conjugate (NIR-830-ZHER2:342) at this step. 6. Dilute the final products NIR830-ZHER2:342–IONP, 50–100 times in water or PBS buffer then take the OD at 500 nm. Multiply the absorbance value with the dilution factor then divide by a factor of 4.3 to get the iron concentration in mg/ml. 7. Calculate the total number of cells to be used for the animal experiments, pipette out calculated volume of cell suspension, then divide by number of animals to be injected in different microfuge tubes. Include extra cells in each tube in case of cell loss during the washing steps. 8. While putting 20 μl cell suspensions using P-200 pipette by removing the plunger of a 0.5 ml insulin syringe, make sure no bubbles are being introduced. 9. Close attention should be paid to avoid any spillage of cell suspension outside the bursa. 10. SKOV3-luc cells injected orthotopically into the ovary and tumors do not project outside the skin until their sizes reach >1.0 cm. So BLI is the most sensitive regimen to monitor kinetics of the bioluminescent signals for quantification and comparison of signal during longitudinal studies for the firefly luciferase expressing cells. When injecting 5 × 104 SKOV3-luc cells into the tumor bursa, the expected size of the tumor by days 7–10 is about 1–2 mm. 11. Typically a dose of 150 mg/kg D-luciferin substrate is recommended. This dose can be reduced if the luciferin is freshly made and good activity of stably expressing luciferase gene in cancer cell line is observed. 12. Since the ovary is located deep inside the abdominal cavity, capture images both dorsally and ventrally so that any spontaneous or time dependent metastases can be viewed. Our purpose is to include BLI to visualize metastases noninvasively and based on the location of metastases we can pay closure attention to those areas more carefully for the delivery of targeted nanoparticles by NIR in vivo imaging. 13. To study the bio-distribution of NIR labeled targeted nanoparticles on organs, first open the abdominal muscle and remove any accumulated ascites which may interfere in NIR signal, using a 1 ml syringe or pipette. Image the ascites to determine

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whether the floating tumor cells can be targeted by the HER-2 targeted imaging agent. Place the mouse face down keeping all the organs intact. Use clear tape to press the mouse so that it is aligned very closely to the tray. When the primary tumor size is >6 mm, it is hard to visualize NIR signals on the metastatic small tumors. In this case, remove all the tumors, which have the highest NIR signal sequentially and adjust the gamma scale to visualize the NIR signal in any smaller tumors present in the peritoneal cavity. 14. Since mouse ovary is located anatomically very close to kidney, when the tumor grows to over 6 mm in diameter, it is almost adhered to the kidney. However un-conjugated and cleaved products of NIR dye-labeled ZHER2:342 conjugated IONPs are eliminated through the kidney, which is a potential problem for NIR imaging in an ovarian orthotopic mouse model. To solve this issue, acquire the images from the dorsal side, ventral side, tumor injection site, and opposite of injection site. In addition, body background, especially in folded skin areas, may generate some artifacts in this preclinical imaging system.

Acknowledgements The authors gratefully acknowledge Dr. Malgorzata Lipowska, Department of Radiology, Emory University School of Medicine, Atlanta, GA for generous supply of NIR-830 dye. This study was supported by NIH R01CA133722 to Dr. Lily Yang. References 1. Bast RC Jr, Hennessy B, Mills GB (2009) The biology of ovarian cancer: new opportunities for translation. Nat Rev Cancer 9:415–428 2. Hellstrom I, Goodman G, Pullman J, Yang Y, Hellstrom KE (2001) Overexpression of HER-2 in ovarian carcinomas. Cancer Res 61: 2420–2423 3. Bookman MA, Darcy KM, Clarke-Pearson D, Boothby RA, Horowitz IR (2003) Evaluation of monoclonal humanized anti-HER2 antibody, trastuzumab, in patients with recurrent or refractory ovarian or primary peritoneal carcinoma with overexpression of HER2: a phase II trial of the Gynecologic Oncology Group. J Clin Oncol 21:283–290 4. Orlova A, Magnusson M, Eriksson TL, Nilsson M, Larsson B, Hoiden-Guthenberg I, Widstrom C, Carlsson J, Tolmachev V, Stahl S, Nilsson FY (2006) Tumor imaging using a picomolar affinity HER2 binding affibody molecule. Cancer Res 66:4339–4348

5. Orlova A, Tolmachev V, Pehrson R, Lindborg M, Tran T, Sandstrom M, Nilsson FY, Wennborg A, Abrahmsen L, Feldwisch J (2007) Synthetic affibody molecules: a novel class of affinity ligands for molecular imaging of HER2-expressing malignant tumors. Cancer Res 67:2178–2186 6. Weissleder R, Ntziachristos V (2003) Shedding light onto live molecular targets. Nat Med 9: 123–128 7. Hilderbrand SA, Weissleder R (2010) Nearinfrared fluorescence: application to in vivo molecular imaging. Curr Opin Chem Biol 14: 71–79 8. Marshall MV, Rasmussen JC, Tan I-V et al (2010) Near-infrared fluorescent imaging in humans with indocyanine green: a review and update. Open Surg Oncol J 2:12–25 9. van Dam GM, Themelis G, Crane LM, Harlaar NJ, Pleijhuis RG, Kelder W, Sarantopoulos A, de Jong JS, Arts HJ, van der Zee AG, Bart J,

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Low PS, Ntziachristos V (2011) Intraoperative tumor-specific fluorescence imaging in ovarian cancer by folate receptor-alpha targeting: first in-human results. Nat Med 17:1315–1319 Lee SB, Hassan M, Fisher R, Chertov O, Chernomordik V, Kramer-Marek G, Gandjbakhche A, Capala J (2008) Affibody molecules for in vivo characterization of HER2-positive tumors by near-infrared imaging. Clin Cancer Res 14:3840–3849 Lee GY, Qian WP, Wang L, Wang YA, Staley CA, Satpathy M, Nie S, Mao H, Yang L (2013) Theranostic nanoparticles with controlled release of gemcitabine for targeted therapy and MRI of pancreatic cancer. ACS Nano 7(3): 2078–2089 Harisinghani MG, Barentsz J, Hahn PF, Deserno WM, Tabatabaei S, van de Kaa CH, de la Rosette J, Weissleder R (2003) Noninvasive detection of clinically occult lymph-node metastases in prostate cancer. N Engl J Med 348:2491–2499 Satpathy M, Wang L et al (2014) Active targeting using HER-2-affibody-conjugated nanoparticles enabled sensitive and specific imaging of orthotopic HER-2 positive ovarian tumors. Small 10(3):544–555 Satpathy M, Qian W, Zielinski R, Wang L, Lei X, Capala J, Wang YA, Lipowska M, Kairdolf

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BA, Jiang H, Nie S, Mao H, Yang L (2013) Multimodality imaging of ovarian cancer using HER2 affibody conjugated nanoparticles. American Association for Cancer Research, Washington, DC Strekowski L, Mason CJ, Lee H, Gupta R, Sowell J, Pantonay G (2003) Synthesis of water soluble near-infrared cyanine dyes functionalized with [(succinimido)oxy] carbonyl group. J Heterocyclic Chem 40:913–916 Smith B, Lyakhov I, Loomis K, Needle D, Baxa U, Yavlovich A, Capala J, Blumenthal R, Puri A (2011) Hyperthermia-triggered intracellular delivery of anticancer agent to HER2(+) cells by HER2-specific affibody (ZHER2-GS-Cys)-conjugated thermosensitive liposomes (HER2(+) affisomes). J Control Release 153:187–194 Zielinski R, Lyakhov I, Jacobs A, Chertov O, Kramer-Marek G, Francella N, Stephen A, Fisher R, Blumenthal R, Capala J (2009) Affitoxin—a novel recombinant, HER2-specific, anticancer agent for targeted therapy of HER2positive tumors. J Immunother 32:817–825 Lyakhov I, Zielinski R, Kuban M, KramerMarek G, Fisher R, Chertov O, Bindu L, Capala J (2010) HER2- and EGFR-specific affiprobes: novel recombinant optical probes for cell imaging. Chembiochem 11:345–350

Chapter 14 Measuring Cardiac Autophagic Flux In Vitro and In Vivo Michael A. Gurney, Chengqun Huang, Jennifer M. Ramil, Nandini Ravindran, Allen M. Andres, Jon Sin, Phyllis-Jean Linton, and Roberta A. Gottlieb Abstract Autophagy is a lysosomal-dependent catabolic pathway that recycles various cytoplasmic-borne components, such as organelles and proteins, through the lysosomes. This process creates energy and biomolecules that are used to maintain homeostasis and to serve as an energy source under conditions of acute stress. Autophagic flux is a measure of efficiency or throughput of the pathway. Here, we describe a method for determining autophagic flux in vitro and in vivo using the autophagosomal/lysosomal fusion inhibitors chloroquine or bafilomycin A1 and then probing for the autophagosomal marker LC3-II via Western Blot. Key words LC3, Autophagy, Cardiac Autophagic Flux, Chloroquine, Bafilomycin A1

1

Introduction Autophagy is a highly conserved constitutive cellular pathway that degrades damaged and/or dysfunctional organelles, protein aggregates, and other cellular components via the lysosomal machinery. Autophagy provides nutrients and energy to maintain cellular homeostasis, and under conditions of acute cellular stress, such as heart attack or nutrient deprivation, autophagy promotes cellular survival. Due to its key role in cellular homeostasis and response to stress, autophagy impacts several processes including inflammation, the cell cycle, and metabolism. Additionally, autophagy has been shown to play a role in the initiation and progression of human diseases such as diabetes, cancer, cardiovascular disease, bacterial and viral infection, Alzheimer’s disease, and Crohn’s disease, making it an important pathway to study. In short, upon induction of the autophagy pathway a doublemembrane structure forms around the cargo to be degraded, forming an autophagosome. The autophagosome fuses with a lysosome, forming an autophagolysosome, followed by degradation of the

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_14, © Springer Science+Business Media New York 2015

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Fig. 1 The mechanism of autophagic flux inhibitors. LC3-I is converted to the LC3-II phosphatidylethanolamine (PE) conjugate and inserted into both leaflets of the autophagosomal membrane forming around the cargo. Eventually, the cargo is enveloped and LC3-II is recycled off the outer leaflet of the autophagosome. (a) Normally, the autophagosome binds and fuses to a lysosome, forming an autophagolysosome. The cargo, along with the LC3-II, is then degraded by the hydrolytic enzymes originating from the lysosome. (b) A lysosomal fusion inhibitor raises the pH of the lysosome, making autophagosomal/lysosomal fusion impossible. As the autophagosomes accumulate, LC3-II levels increase and can be detected by Western Blot. (c) The lysosomal protease inhibitors prevent the hydrolytic enzymes in the lysosomes from functioning. Consequently, autophagosolysosomes can form, but the cargo and LC3-II cannot be degraded. The accumulation of LC3-II can be measured by Western Blot

cargo and export of its constituent biomolecules. One of the key molecules associated with the autophagosome is LC3 (microtubuleassociated protein light chain 3—the mammalian homolog of Atg8). LC3 primarily exists in two forms, an 18 kDa cytosolic form (LC3-I) and a 16 kDa phosphatidylethanolamine-conjugated form (LC3-II) that is found in the autophagosomal membrane (see Fig. 1—for a detailed cardiac autophagy review, see Rotter et al. Pharmacological Research, 2012 [1]). Typically, autophagy is measured by Western Blot and microscopic analysis. Observing autophagy by Western Blot is achieved by measuring protein expression of LC3, specifically LC3-II. Since LC3-II is incorporated into autophagosomes, changes in LC3-II levels are indicative of a change in the numbers of autophagosomes. This can be quantified via densitometric analysis of blots. Microscopic observation of autophagy is usually done using LC3 conjugated to a fluor such as Green Fluorescent Protein (GFP-LC3). As autophagosomes form, the fluorochrome-LC3

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molecules are incorporated into the double membrane, creating puncta which can be counted. Consequently, changes in puncta numbers are indicative of a change in autophagy. The aforementioned techniques are static measurements of autophagy and are well suited to experiments comparing relative autophagy levels between different conditions. However, autophagy is a dynamic process and interpretation of a static measurement can be challenging. For example, if LC3-II or puncta levels increase, then is the pathway upregulated, or is degradation of autophagolysosomes deficient (promoting the buildup of LC3-II) [2]? To address this question, one can measure autophagic flux. Autophagic flux is a measure of the efficiency of the autophagy pathway, i.e., autophagic flux is a measure of the number of autophagosomes that progress (from nucleation to formation to degradation) through the autophagy pathway per unit time. There are two basic methods for determining autophagic flux. The first method utilizes a tandem fluorescent-tagged LC3 protein, monomeric Red Fluorescent Protein (or mCherry) coupled to Green Fluorescent Protein-LC3 (mRFP/mCherry-GFP-LC3), to report simultaneously both autophagy induction and reduction via changing numbers of puncta and autophagic flux [3]. In short, the GFP is quenched under acidic conditions, while the mRFP retains fluorescence; consequently, newly formed autophagosomes fluoresce in the green and red channels (yellow when the two channels are overlaid) while the acidic environment of the autophagolysosome quenches the GFP, leaving only red fluorescence [3, 4]. To determine flux, the numbers of yellow and red puncta are compared over time. Samples may be fixed; however, some fixation reagents may artificially create green fluorescence by neutralizing the low pH in the autophagolysosome, so care must be taken when selecting the fixation reagents [4]. To date, this method has been successfully used to assess flux in vitro. The second method used to measure flux introduces a pharmacological blockade that prevents lysosomal/autophagosomal fusion or prevents lysosomal-mediated enzymatic degradation (Fig. 1). There are a variety of agents that can be used for autophagic flux analysis. Bafilomycin A1, chloroquine, and ammonium chloride raise the intralysosomal pH, preventing autophagosomal fusion with the lysosome [4–9]. Moreover, protease inhibitors such as Pepstatin A, E-64d, and leupeptin act to prevent autophagosomal destruction in the lysosome [4, 10, 11]. Samples subjected to an autophagy blockade as well as untreated control samples are collected at a specified time after treatment for comparison. Treatment with any of the aforementioned reagents results in the accumulation of autophagosomes that would have progressed through the pathway during that period, which can be measured by Western Blot as an accumulation of LC3-II vs. a nontreated control [11]. A significant increase in LC3-II, as compared

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Fig. 2 Example of chloroquine/bafilomycin blots and rendered data. (a) LC3 and Actin Western Blots and graphs of HL-1 cells treated with and without chloroquine (3 μM—2 h) under three experimental conditions. (b) LC3 and Actin Western Blots of HL-1 cells treated with and without bafilomycin A1 (100 nM—2 h) under three experimental conditions. (c) LC3 and Actin blots and graph of the hearts collected from male C57BL/6 mice subjected to a 48 h fast with and without 50 mg/kg chloroquine injected i.p. 4 h prior to sacrifice [Tx is treatment ± chloroquine (panels (a) and (c)) or ± Bafilomycin A1 (panel (b))]

to untreated controls, is indicative of robust flux. This method allows for the measurement of flux in vitro and in vivo (see Fig. 2). Below are the protocols that assess flux by blocking the autophagy pathway. The following methods are used for: 1. In vitro analysis of flux 2. In vivo analysis of flux 3. LC3 SDS-PAGE, Western Blot analysis, and data quantification/ reporting

2 2.1

Materials Equipment

1. Pestle for 1.5 mL microfuge tubes or Polytron Homogenizer Power Gen Series and Accessories. 2. Plate reader—e.g., Spectra Max Plus 384 with Softmax Pro5 Software (Molecular Devices). 3. Centrifuge. 4. Hotplate and stirrer. 5. Gel imaging system. 6. Gel boxes and transfer accessories. 7. Rocker.

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8. 1.7 and 0.6 mL sterile microfuge tubes. 9. Non-sterile forceps. 2.2 Reagents and Materials

1. Chloroquine diphosphate. 2. Bafilomycin A1. 3. 1 mL TB Syringe Slip Tip with Needle. 4. Liquid nitrogen. 5. Lysis buffer: Triton-X 100 buffer: [50 mM Tris Base pH = 7.4, 150 mM NaCl, 1 mM EGTA, 1 mM EDTA, 1 % Triton-X 100]; RIPA buffer: [50 mM Tris Base pH=8.0, 150 mM NaCl, 0.1 % SDS, 0.5 % sodium deoxycholate, 1 % NP-40]. 6. Protease inhibitor cocktail—EDTA Free. 7. 100 × 20 mm style tissue culture dishes. 8. 60 × 15 mm style tissue culture dishes. 9. Disposable cell lifter. 10. Nonfat powdered milk. 11. Lowry Assay reagents—e.g., Bio-Rad; DC Protein Reagent A [500-0113], DC Protein Assay Reagent B [500-0114], DC Protein Assay Reagent [500-0115]. 12. Albumin standards (BSA). 13. Sample buffer: 5× Laemmli buffer [10 % SDS, 0.05 % bromophenol blue, 40 % glycerol, 250 mM Tris Base pH = 6.8]—supplement with 10 % β-mercaptoethanol before use. 14. 4–20 or 10–20 % Tris-Glycine Gradient Gel. 15. Molecular weight marker. 16. 10× SDS Running Buffer: 10 L—300.3 g Tris Base, 1.441 kg Glycine, 100 g SDS, pH = 8.3. 17. Transfer Buffer: 25 mM Tris Base, 192 mM Glycine, and 10 % methanol. 18. PVDF or Nitrocellulose Transfer Membrane. 19. Tris Buffered Saline with Tween-20: 20×—2 M NaCl, 200 mM Tris Base pH = 7.4, 2 % Tween-20. 20. Anti-LC3AB Antibody [Cell Signaling (4108S); 1:1000 dilution]. 21. Anti-β-Actin Antibody for loading control [Sigma (A4700); 1:1000 dilution]. 22. Appropriate Purified Peroxidase Labeled IgG (secondary antibody). 23. Enhanced chemiluminescent substrate.

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Methods

3.1 Determination of Autophagic Flux Using Chloroquine or Bafilomycin A1 In Vitro

The Gottlieb and Linton labs have used the following protocols solely on HL-1 and C2C12 mouse cardiomyocyte and myoblast cell lines, respectively. We highly recommend performing an analysis of in vitro chloroquine or bafilomycin A1 dose/time responses under the desired experimental conditions to optimize autophagic flux detection in the cell line of interest. Chloroquine or Bafilomycin A1 treatment will reveal the magnitude of autophagic flux during the 2 h period prior to the end of the experiment. Consequently, autophagic flux inhibitors can be injected before, after, or concurrent with the onset of the experiment or treatment. 1. Incubate cells until appropriate confluence is reached (1–2 days) (see Notes 1 and 2). 2. Replace media with fresh complete media with 3 μM chloroquine (dissolved in PBS or saline) or 100 nM bafilomycin A1 (dissolved in DMSO) for 2 h (see Note 3). 3. Quickly wash 3× with 1 mL warm 1× PBS (37 °C) and scrape cells in the presence of Lysis Buffer (300–400 μL for a 60 mm dish) with added 1× Protease Inhibitor Cocktail (and phosphatase inhibitors NaF (10 μL/mL), Na3VO4 (5 μL/mL), and Beta-glycerophosphate disodium (35 μL/mL) if the lysate is to be probed with additional antibodies specific for phosphorylated proteins) (see Note 4). 4. Pipet lysate up and down several times to homogenize lysate. 5. Centrifuge at a minimum of 600 × g at 4 °C for 5 min, transfer supernatant to a fresh tube, and freeze at −80 °C until time to run PAGE and Western Blot (Subheading 3.3).

3.2 Determination of Autophagic Flux Using Chloroquine In Vivo

Bafilomycin A1 is expensive and the Gottlieb lab has found it to be unsuitable in vivo [12] because it needs to be administered at low doses for short periods or induces nonspecific effects such as a disruption of proteasomal and vesicular dynamics and other key cellular processes [6, 13]; however, there are reports of it being used successfully in vivo [14]. The Gottlieb and Linton labs have opted to use chloroquine for the determination of autophagic flux in vivo. Additionally, the Gottlieb and Linton labs have found that autophagy responses differ with age, tissue, strain, and possibly gender in mice. Therefore, we highly recommend performing a chloroquine dose/time response with your animals under your experimental conditions to optimize autophagic flux analysis. An additional observation that warrants mentioning is that circadian rhythm can have a dramatic impact on basal autophagy [1, 15], so ensuring that animals are treated and tissues are harvested at the same time of day is very important. Our work focuses exclusively on the hearts of C57BL/6, BALB/c, and FVB/N mice, so

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treatment times and doses stated below were optimized for those animals subjected to a 24–48 h fast, with tissues harvested at 10 AM, in a facility with lights on at 6 AM and off at 6 PM. Chloroquine treatment will reveal the magnitude of autophagic flux during the 2–4 h period prior to the end of the experiment. Consequently, autophagic flux inhibitors can be injected before, after, or concurrent with the onset of the experiment or treatment. 3.2.1 Preparing and Administering the Chloroquine and Freezing the Heart Tissue

1. Weigh mice just prior to preparing the chloroquine solution to determine appropriate chloroquine dosage. 2. Prepare the chloroquine solution in sterile saline or 1× PBS to administer at 10–50 mg/kg (see Note 5). Inject the mice i.p. and wait a minimum of 2–4 h. Injection volumes should be between 50 and 200 μL. 3. Sacrifice the mouse, harvest the heart, snap freeze the tissue in liquid nitrogen, and keep tissues at −80 °C. (Autophagy is different between atria and ventricles, so we trim the atria before snap freezing the ventricles.)

3.2.2 Preparing the Heart Tissue for SDS-PAGE and Western Blot

1. If the tissue is to be further divided, cut the tissue with a razor blade on dry ice to prevent thawing. 2. Prepare an appropriate volume of ice-cold lysis buffer with 1× Protease Inhibitor (and phosphatase inhibitors NaF (10 μL/ mL), Na3VO4 (5 μL/mL), and beta-glycerophosphate disodium (35 μL/mL) if the lysate is to be probed with additional antibodies that recognize phosphorylated proteins) (see Note 4). 3. Homogenize the heart in ice-cold lysis buffer. The Douncing (done on ice), Polytron (3–5 short bursts at high speed on ice), and the Pestle (place pestle in 1.7 mL microfuge tube and homogenize) methods have been successfully used, but regardless of the method utilized, each sample is treated in exactly the same way. To homogenize half a mouse heart, 400–600 μL of lysis buffer is usually sufficient. 4. Centrifuge for 10 min at a minimum of 600 × g at 4 °C in a centrifuge. Collect the supernatant and freeze at −80 °C in aliquots to minimize freeze/thaw cycles (repeated freeze/thaw cycles are detrimental to LC3 analysis [4]). Keep frozen until thawed for PAGE and Western Blot analysis (Subheading 3.3).

3.3 LC3 SDS-PAGE and Western Blot Analysis

The times and materials used in the text below are specific to Life Technologies Novex Tris-Glycine Gradient Gels. Other materials may be substituted, if needed. 1. Thaw sample lysate aliquots on ice and keep on ice to prevent protein degradation. 2. Quantify total protein in each sample by performing a Lowry Assay in accordance with the manufacturer’s instructions (see Note 6). The protein concentration in the lysate often

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exceeds the Albumin Standards, so 1:10–1:50 dilutions are not uncommon. 3. Combine 25–30 μg total protein, 1× Sample Buffer (supplemented with 10 % β-Mercaptoethanol), and water in a test tube (check the manufacturer’s instructions to determine the appropriate volume to load per well for your gels and make your sample accordingly). If the protein concentration in the lysate is too high to pipet 25–30 μg accurately, dilute the lysate with lysis buffer with protease and phosphatase inhibitors before adding it to the Sample Buffer and Water. 4. Boil sample for 5–10 min, centrifuge briefly, and load 30 μL lysate and protein standard onto a 4–20 to 10–20 % TrisGlycine Gel (see Note 7). Run PAGE using 1×SDS Running Buffer. 5. While gel is running, prepare and refrigerate transfer buffer. 6. Stop the electrophoresis when the lowest molecular weight marker (10 kDa if using the Precision Plus Protein Standard) is just at the bottom of the gel. 7. Transfer onto a PVDF membrane at 30 V for 3 h at 4 °C in accordance with the manufacturer’s instructions using TrisGlycine Transfer Buffer (see Note 8). [Remember PVDF requires methanol treatment.] 8. Block 1 h in 5 % Powdered Milk/1× Tris Buffered Saline with Tween-20 (TBST). Incubate at room temperature on a rocker. 9. Briefly wash twice with 1× TBST at room temperature on a rocker. 10. Add 1:1,000 dilution of rabbit-anti-LC3AB antibody diluted in 5 % nonfat powdered milk/1× TBST and incubate overnight at 4 °C on a rocker (see Note 9). 11. Wash 3× for 7–10 min with 1× TBST at room temperature on a rocker (see Note 10). 12. Add 1:2,500 dilution of peroxidase labeled goat-anti-rabbit antibody in 5 % nonfat powdered milk/1× TBST for 60–75 min at room temperature on a rocker (see Note 10). 13. Wash three or four times for 10 min with 1× TBST at room temperature on a rocker. 14. Develop blot with SuperSignal Substrate. Expect LC3-I and LC3-II bands at approximately 18 kDa and 16 kDa, respectively. 15. (Optional) Strip blots with a Stripping Buffer at room temperature on a rocker in accordance with manufacturer’s instructions. (If looking to probe for additional proteins at or around the molecular weight of LC3 or the loading controls (Actin, etc.), then stripping the blot is recommended.)

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16. Reprobe blot with appropriate loading control/housekeeping protein primary antibody such as Rho GDIα and Tubulin, β-Actin. We prefer using a 1:1,000 mouse-anti-actin or the Rho GDIα primary antibody for 1 h followed by 1:2,500 peroxidase labeled goat-anti-mouse secondary antibody as outlined in steps 9–12 (steps 8–12 if blot was stripped). 17. Quantify LC3-I (optional), LC3-II, and housekeeping protein bands (loading controls) using ImageJ (NIH; available for free download at http://rsb.info.nih.gov/ij/) and report data as LC3-II/Housekeeping (see Fig. 2). (There are a myriad of tutorials for using ImageJ available online.) Remember that a significant increase in LC3-II as compared to the untreated control is indicative of robust flux.

4

Notes 1. Flasks/Dishes for HL-1 cell assays are coated with 0.5 % fibronectin/2 % gelatin for a minimum of 1 h at 37 °C. 2. Our labs and others have noted that when working in vitro, cell passage number, media depth (hypoxia induces autophagy through Hif-1α), and cell confluence can have an impact on autophagy. Consequently, an effort to compare samples from similar passage numbers in conjunction with similar cell densities is important. 3. Our labs have noticed that DMSO can be a potent inducer of autophagy, so when we prepare compounds that must be dissolved in DMSO we try to minimize the amount of DMSO added to the cells (ethanol will do similar things). Examples of such compounds are Chloramphenicol, Pepstatin A, and Bafilomycin A1. If using one of the aforementioned compounds, it is important to add a similar amount of DMSO to the control cells. In general, we try to keep the concentrations of DMSO in our experiments to 0.1 % or lower. Additionally, some groups are using up to 125 μM chloroquine for their experiments on their cell lines (at 250 μM chloroquine they demonstrated cell death) [16]. 4. Both RIPA and a standard Triton-X 100-based Lysis Buffer have been successfully used. Try to minimize the amount of lysis buffer used with in vitro samples in order to maximize the protein concentration of the lysate. If the lysate is too dilute, then it may not be possible to load sufficient total protein on the gel. 5. Some groups report using in vivo chloroquine dosages as low as 10 mg/kg [12] and as high as 112 mg/kg [10, 12, 17]. We have determined that the optimal chloroquine dose is 50 mg/ kg for our studies in C57BL/6 and BALB/c mice.

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6. Lowry Assay is the preferred method of determining protein concentration in the presence of the detergents used in the cell lysis buffers. 7. LC3 blots are best done on 4–20 or 10–20 % gradient gels. For running LC3 and Actin blots either will work equally well; however, if probing for other proteins in addition to LC3 and Actin, then use either a 10–20 % for probing for high or 4–20 % for low molecular weight proteins. 8. PVDF membrane is the preferred membrane for detecting low molecular weight proteins. Nitrocellulose can be substituted if needed. 9. Some reports suggest that the LC3B isoform is the only autophagosome-associated protein that changes upon autophagy induction, implying that the LC3B antibody should be used exclusively for flux analysis. However, it is assumed that each of the different LC3 homologs may be expressed differently in different tissues [4]. According to Daniel Klionsky, “it should not be assumed that LC3B is the optimal protein to monitor [4].” Consequently, the Gottlieb and Linton labs have opted to use an LC3A/B antibody, which detects two of the LC3 isoforms, for our studies. Our labs have found that overnight incubation at 4 °C with the LC3A/B antibody gives the best results. Additionally, we have found that if a blot is to be stripped and re-probed, LC3 must be the first probed. Repeated stripping of the blot reduces the LC3 signal substantially and could potentially confound results. Finally, if the primary antibodies are to be reused several times, it is preferable to make primary antibody dilutions in 5 % BSA/1× TBST. 10. In the case of probing for LC3 by Western Blot, we have found that longer 1× TBST washes following antibody incubations improve the appearance of the blots substantially. Hence, we recommend no less than 7 min 1× TBST washes. References 1. Rotter D, Rothermel BA (2012) Targets, trafficking, and timing of cardiac autophagy. Pharmacol Res 66:494–504 2. Rubinsztein DC et al (2009) In search of an “autophagomometer”. Autophagy 5:585–589 3. Kimura S et al (2007) Dissection of the autophagosome maturation process by a novel

reporter protein, tandem fluorescent-tagged LC3. Autophagy 3:452–460 4. Klionsky DJ et al (2012) Guidelines for the use and interpretation of assays for monitoring autophagy. Autophagy 8:445–544 5. Seglen PO, Reith A (1976) Ammonia inhibition of protein degradation in isolated rat

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hepatocytes. Quantitative ultrastructural alterations in the lysosomal system. Exp Cell Res 100:276–280 Klionsky DJ et al (2008) Does bafilomycin A1 block the fusion of autophagosomes with lysosomes? Autophagy 4:849–950 Yamamoto A et al (1998) Bafilomycin A1 prevents maturation of autophagic vacuoles by inhibiting fusion between autophagosomes and lysosomes in rat hepatoma cell line, H-4II-E cells. Cell Struct Funct 23:33–42 Poole B, Ohkuma S (1981) Effect of weak bases on the intralysosomal pH in mouse peritoneal macrophages. J Cell Biol 90: 665–669 Kawai A et al (2007) Autophagosomelysosome fusion depends on the pH in acidic compartments in CHO cells. Autophagy 3: 154–157 Haspel J et al (2011) Characterization of macroautophagic flux in vivo using a leupeptinbased assay. Autophagy 7:629–642

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Chapter 15 PET Imaging for Tyrosine Kinase Inhibitor (TKI) Biodistribution in Mice Hiroshi Fushiki, Yoshihiro Murakami, Sosuke Miyoshi, and Shintaro Nishimura Abstract Receptor tyrosine kinases play a critical role in cell growth, survival, and proliferation, and are considered potential molecular targets for the treatment of cancer. Although several tyrosine kinase inhibitors (TKIs), such as erlotinib and gefitinib, have demonstrated clinical efficacy via the inhibition of the epidermal growth factor receptor (EGFR), most TKIs are only effective in a small proportion of patients. Positron emission tomography (PET) imaging is a methodology of molecular imaging based on nuclear imaging. PET imaging in combination with radiolabeled TKIs improves accuracy of quantitative imaging strategies and the probability of successful drug development, and may facilitate the stratification of patients. Here, we describe a protocol for PET imaging using radiolabeled TKI in preclinical trials. Key words Tyrosine kinase inhibitor (TKI), PET, Biodistribution, Bioluminescent imaging, Orthotopic pulmonary tumor model

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Introduction Molecular imaging technology has accelerated the translation of preclinical research to the clinical stage by providing physiological, anatomic, and metabolic information through interrogating certain targets [1]. Of the current imaging techniques, positron emission tomography (PET) is an established clinical molecular imaging modality with high sensitivity and deep tissue penetration. PET is a nuclear imaging technique used to map biological and physiological processes in vivo following the administration of positron emitting radioligands, and is based on the detection of photons released by annihilation of positrons emitted by radioisotopes. Positron-emitting radionuclides are first produced in a cyclotron by bombarding target material with accelerated protons. In the body, these radionuclides introduced into a radioligand emit positrons that undergo annihilation with nearby electrons, resulting in the release of two annihilation photons. These photons are detected

Gil Mor and Ayesha B. Alvero (eds.), Apoptosis and Cancer: Methods and Protocols, Methods in Molecular Biology, vol. 1219, DOI 10.1007/978-1-4939-1661-0_15, © Springer Science+Business Media New York 2015

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by imaging instruments and the resulting data can be used to reveal the distribution of the radiotracer within the body. Receptor tyrosine kinases (RTKs) are one of a series of cell surface receptors for growth factors, cytokines, and hormones. The activation and signal transduction initiated by the tyrosine kinase activity of RTKs induces many cellular events such as proliferation and anti-apoptotic processes [2]. RTKs can also play a critical role in the development and progression of many types of cancer [3–5]. Tyrosine kinase inhibitors (TKIs) have become a major category of anti-cancer drugs due to their good efficacies and distinct rationale in treating cancer. For some antitumor drugs including TKIs, the drugs themselves or closely related analogs were labeled for PET imaging studies. Penetration or accumulation of labeled drugs into tumor burden could be evaluated by PET imaging, and the observations might be used for the prediction of the anti-cancer effect of TKIs [6–10], monoclonal antibodies [11–13], and other antitumor agents [14–16]. In this chapter, a brief methodology for performing PET imaging of TKIs in a preclinical cancer model is described, demonstrating PET imaging using our original ALK inhibitor ASP3026 as a drug candidate for the echinoderm microtubuleassociated protein-like 4 (EML4)-anaplastic lymphoma kinase (ALK) positive fraction of lung cancer patients [17]. It should be noted, however, that this technique requires modification depending on the purpose or nature of the agent under evaluation. In addition, we also describe the bioluminescent imaging (BLI) technique and genetic engineering required for stable expression of luciferase in tumor cells [18, 19]. This approach enables the tumor growth to be monitored and may help facilitate future in vivo cancer research.

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2.1 For Preparation of Murine Cancer Model and Bioluminescent Imaging

1. Luciferase expressing NCI-H2228 cells: Parent NCI-H2228 cells from the American Type Culture Collection (Manassas, VA, USA), transfected with the luciferase gene (see Note 1). 2. Culture media: RPMI1640 supplemented with 10 % heatinactivated fetal bovine serum. 3. Solution of 0.25 % Trypsin containing 0.1 M EDTA. 4. Phosphate buffered saline (PBS, pH 7.4). 5. Matrigel (i.e., BD Bioscience, Franklin Lakes, NJ, USA). 6. Immune-incompetent mice (i.e., NOD.CB17-Prkdcscid/J mice). 7. Anesthesia controller (i.e., RC2 Rodent Anesthesia System, VetEquip Inc., Pleasanton, CA, USA). 8. Isoflurane.

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9. 29-gauge needle attached to a 0.3 mL syringe. 10. Surgical instruments (knives, scissors, tweezers, clips, sutures, and razors). 11. Ethanol (70 %). 12. In vivo imaging system (i.e., IVIS-spectrum (Perkin Elmer Inc. [formerly Xenogen Inc.]), Waltham, MA, USA). 13. In vivo grade d-luciferin (i.e., VivoGlo Luciferin, Promega corporation, Madison, WI, USA) dissolved with PBS to a concentration of 15 mg/mL. d-luciferin solution should be stored at −20 °C until use. 2.2 For Synthesis of Radiolabeled Anti-cancer Drugs

1. Cyclotron (i.e., Cyclone 18/9, JFE Engineering Corp., Tokyo, Japan). 2.

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N-N2 gas target.

3. Automated chemical synthesizer (i.e., from Dainippon Seiki, Kyoto, Japan). 4. Dose calibrator (i.e., CRC-127R, Capintec, Inc., Ramsey, NJ, USA). 5. Chemicals, including optimal precursor of radioligand. 6. Sterile saline. 7. Sterile 0.2 μm pored hydrophilicity filter. 8. Sterile glass vial with butyl rubber. 2.3

For PET Imaging

1. PET scanner (i.e., Inveon docked system, Siemens AG, Munich, Germany). 2. Anesthesia controller. 3. Indwelling needle (see Note 2). 4. Injection needle and syringe: 26-gauge needle attached to a 1 mL syringe. 5. Dual chamber m2m (i.e., from Imaging Corp., Cleveland, OH, USA [see Fig. 1]). 6. Sterile saline.

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3.1 Preparation of Murine Cancer Model

Current imaging techniques enable tumor progression to be monitored and the efficacy of anti-cancer drugs to be evaluated, even in orthotopic or metastatic tumor models. Of these techniques, bioluminescent imaging (BLI) is widely used and recognized as an effective tool for monitoring tumor burden. X-ray computed tomography (CT) imaging is suitable for monitoring lung cancer in mice as well as in humans. The combination of BLI/CT and TKI-PET is therefore an effective strategy for assessing TKIs.

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Fig. 1 Dual chamber for mice. Anesthesia gas line and compatible slide connector for Inveon system are pertained

Here, we outline the protocol for an intrapulmonary injection method in a murine model of orthotopic lung cancer [20, 21]. 1. Maintain and expand cancer cells with optimal culture medium. 2. Harvest cells with 0.25 % Trypsin containing 0.1 M EDTA solution, wash with PBS, and resuspend at an optimal cell concentration in PBS containing 500 μM/mL Matrigel. 3. Animal experiments and surgery should be conducted in accordance with ethical guidelines following the approval of the relevant Institutional Animal Care and Use Committee (IACUC). Pinch the tail of the mouse to ensure that the nociceptive response is controlled, and anesthetize with 2–3 % isoflurane. 4. Sterilize the left chest wall with 70 % ethanol, and shave the area of implantation. 5. Make a small skin incision on the left chest wall with a surgical knife, or scissors. 6. Inject 20 μL of cell suspension directly into the lung with a 29-gauge needle syringe attached to a 0.3 mL syringe. During this procedure, monitor the respiratory rate of the left lung. 7. Administer appropriate anti-biologics and analgesic agents, such as NSAIDs, and close the incision with a surgical clip or surgical silk suture. 8. Maintain the mice until the imaging experiments.

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BLI is the most suitable and effective methodology for monitoring growth of implanted tumor cells in the lung. A brief protocol of BLI is outlined below. 1. Anesthetize mice with 2–3 % isoflurane. 2. Select the optimal condition of BLI on your in vivo imaging software. Imaging times range from 0.5 s to 5 min (binning = 2–8), depending on the total tumor burden as a function of light emission from tumor cells. Ensure that the camera is not saturated, and acquire BLI with five mice with the field of view set at D. 3. Inject 10 mL/kg of d-luciferin solution into peritoneal cavity in tumor-bearing mice. 4. Keep the mice for 10 min. Prior to the BLI acquisition, place the mice on the stage of the imaging system. For the intrapulmonary tumor model, position the mice with their left side facing the camera (i.e., the side of cancer cell injection). 5. Ten minutes after the administration of d-luciferin, acquire bioluminescent images. 6. By following the manufacturer’s instructions, quantify the photoluminescent signals on the bioluminescent images by region of interest (ROI) analysis.

3.3 Synthesis of Radiolabeled Anti-cancer Drugs

The chemical synthetic procedure for radiolabeling anti-cancer drugs, including TKIs, varies depending on their chemical structure. Specific articles should be referred to for the details of radiochemistry. Here, we describe a brief protocol concerning the synthesis of [11C]ASP3026, which is the parent compound of an ALK inhibitor under development in our clinic. 1. Prepare or purchase optimal precursors of [11C]ASP3026. 2. Perform a 14N(p, α)11C nuclear reaction by proton bombardment of a 14N-N2 target using a cyclotron-target system to obtain [11C]CO2. And then produce [11C]CH3I from [11C]CO2. 3. Introduce 11C-atom into [11C]ASP3026 by reacting the precursor and [11C]CH3I using automated chemical synthesizer. 4. Purify the crude [11C]ASP3026 with HPLC, and the collected [11C]ASP3026 fraction was evaporated under reduced pressure. 5. Add sterile saline to the residue, and transfer the solution into a sterile glass vial through a sterile filter for use in in vivo experiments.

3.4

PET Imaging

PET imaging acquisition is defined into two methodologies: dynamic PET and static PET. Dynamic PET imaging obtains PET data via correction of a series frame of sinogram data, which is sequentially separated from 10 s to 10–20 min. PET images can be

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reconstructed from the data of each time frame. We can obtain kinetic information regarding PET tracers from a series of images from dynamic PET acquisition. In contrast, static PET imaging has the advantage of being high-throughput and therefore does not lack kinetic information regarding PET tracers. In this section, animal handling and PET imaging acquisition are described. 3.4.1 Dynamic PET Imaging Acquisition

1. Following the manufacturer’s instructions set up the Inveon system to demonstrate PET imaging acquisition. Generally, dynamic PET acquisition will be performed for 60–90 min. 2. Insert an indwelling needle in the tail vein of tumor-bearing mice (Subheading 3.1). In our indwelling system, blood is observed in the indicator of the indwelling cassette when an insertion is successful. To confirm a flow pass, infuse saline (