Nuclear Pore Complexes and Nucleocytoplasmic Transport - Methods [1st Edition] 9780124171787, 9780124171602

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages xiii-xvii
PrefacePages xix-xxValérie Doye
Chapter 1 - Fifty Years of Nuclear Pores and Nucleocytoplasmic Transport Studies: Multiple Tools Revealing Complex RulesPages 1-40Aurélie G. Floch, Benoit Palancade, Valérie Doye
Chapter 2 - Imaging Metazoan Nuclear Pore Complexes by Field Emission Scanning Electron MicroscopyPages 41-58Boris Fichtman, Lihi Shaulov, Amnon Harel
Chapter 3 - Imaging Yeast NPCs: From Classical Electron Microscopy to Immuno-SEMPages 59-79Elena Kiseleva, A. Christine Richardson, Jindriska Fiserova, Anton A. Strunov, Matthew C. Spink, Simeon R. Johnson, Martin W. Goldberg
Chapter 4 - Exploring Nuclear Pore Complex Molecular Architecture by Immuno-Electron Microscopy Using Xenopus OocytesPages 81-98Nelly Panté, Birthe Fahrenkrog
Chapter 5 - Utilizing the Dyn2 Dimerization-Zipper as a Tool to Probe NPC Structure and FunctionPages 99-115Dirk Flemming, Philipp Stelter, Ed Hurt
Chapter 6 - The Use of Targeted Proteomics to Determine the Stoichiometry of Large Macromolecular AssembliesPages 117-146Alessandro Ori, Amparo Andrés-Pons, Martin Beck
Chapter 7 - A Pulse–Chase Epitope Labeling to Study Cellular Dynamics of Newly Synthesized Proteins: A Novel Strategy to Characterize NPC Biogenesis and Ribosome Maturation/ExportPages 147-163Philipp Stelter, Ed Hurt
Chapter 8 - Analysis of Nuclear Reconstitution, Nuclear Envelope Assembly, and Nuclear Pore Assembly Using Xenopus In Vitro AssaysPages 165-191Cyril Bernis, Douglass J. Forbes
Chapter 9 - Xenopus In Vitro Assays to Analyze the Function of Transmembrane Nucleoporins and Targeting of Inner Nuclear Membrane ProteinsPages 193-218Nathalie Eisenhardt, Allana Schooley, Wolfram Antonin
Chapter 10 - Imaging the Assembly, Structure, and Function of the Nuclear Pore Inside CellsPages 219-238Shotaro Otsuka, Anna Szymborska, Jan Ellenberg
Chapter 11 - Cell-Fusion Method to Visualize Interphase Nuclear Pore FormationPages 239-254Kazuhiro Maeshima, Tomoko Funakoshi, Naoko Imamoto
Chapter 12 - An In Vitro System to Study Nuclear Envelope BreakdownPages 255-276Joseph Marino, Lysie Champion, Cornelia Wandke, Peter Horvath, Monika I. Mayr, Ulrike Kutay
Chapter 13 - Modern Tools to Study Nuclear Pore Complexes and Nucleocytoplasmic Transport in Caenorhabditis elegansPages 277-310Peter Askjaer, Vincent Galy, Peter Meister
Chapter 14 - Assessing Regulated Nuclear Transport in Saccharomyces cerevisiaePages 311-330Christopher Ptak, Richard W. Wozniak
Chapter 15 - Analysis of Nucleocytoplasmic Transport in Digitonin-Permeabilized Cells Under Different Cellular ConditionsPages 331-352Maiko Furuta, Shingo Kose, Ralph H. Kehlenbach, Naoko Imamoto
Chapter 16 - Novel Approaches for the Identification of Nuclear Transport Receptor SubstratesPages 353-378Makoto Kimura, Ketan Thakar, Samir Karaca, Naoko Imamoto, Ralph H. Kehlenbach
Chapter 17 - NPC Mimics: Probing the Mechanism of Nucleocytoplasmic TransportPages 379-393Tijana Jovanovic-Talisman, Brian T. Chait, Michael P. Rout
Chapter 18 - Analysis of RNA Transport in Xenopus Oocytes and Mammalian CellsPages 395-413Ichiro Taniguchi, Asako McCloskey, Mutsuhito Ohno
Chapter 19 - Strategies for Investigating Nuclear–Cytoplasmic tRNA Dynamics in Yeast and Mammalian CellsPages 415-436Jacqueline B. Pierce, Shawn C. Chafe, Manoja, B.K. Eswara, George van der Merwe, Dev Mangroo
Chapter 20 - Dissecting Ribosome Assembly and Transport in Budding YeastPages 437-461Martin Altvater, Sabina Schütz, Yiming Chang, Vikram Govind Panse
Chapter 21 - Approaches to Studying Subnuclear Organization and Gene–Nuclear Pore InteractionsPages 463-485Defne Emel Egecioglu, Agustina D’Urso, Donna Garvey Brickner, William H. Light, Jason H. Brickner
IndexPages 487-495
Volumes in SeriesPages 497-508

Citation preview

Series Editors Leslie Wilson Department of Molecular, Cellular and Developmental Biology University of California Santa Barbara, California

Phong Tran Department of Cell and Developmental Biology University of Pennsylvania Philadelphia, Pennsylvania

Academic Press is an imprint of Elsevier 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA 225 Wyman Street, Waltham, MA 02451, USA The Boulevard, Langford Lane, Kidlington, Oxford, OX5 1GB, UK 32 Jamestown Road, London NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands First edition 2014 Copyright # 2014 Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN: 978-0-12-417160-2 ISSN: 0091-679X For information on all Academic Press publications visit our website at store.elsevier.com

Printed and bound in USA 14 15 16 11 10 9 8 7 6

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Contributors Martin Altvater Institute of Biochemistry (IBC), ETH Zu¨rich, Otto-Stern-Weg 3, and MLS Program, Life Science Zurich Graduate School, Winterthurerstrasse 190, Zurich, Switzerland Amparo Andre´s-Pons European Molecular Biology Laboratory, Structural and Computational Biology Unit, Meyerhofstr. 1, 69117, Heidelberg, Germany Wolfram Antonin Friedrich Miescher Laboratory of the Max Planck Society, Tu¨bingen, Germany Peter Askjaer Andalusian Center for Developmental Biology (CABD), CSIC/Junta de Andalucı´a/ Universidad Pablo de Olavide, Carretera de Utrera, Seville, Spain Martin Beck European Molecular Biology Laboratory, Structural and Computational Biology Unit, Meyerhofstr. 1, 69117, Heidelberg, Germany Cyril Bernis Cell and Developmental Biology, University of California, San Diego, California, USA Donna Garvey Brickner Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Jason H. Brickner Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Shawn C. Chafe Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada Brian T. Chait Laboratory of Mass Spectrometry and Gaseous Ion Chemistry, The Rockefeller University, New York, USA Lysie Champion Institute of Biochemistry, ETH Zurich, Zurich, Switzerland Yiming Chang Institute of Biochemistry (IBC), ETH Zu¨rich, Otto-Stern-Weg 3, Zurich, Switzerland Vale´rie Doye Institut Jacques Monod, CNRS, UMR 7592, Univ. Paris Diderot, Sorbonne Paris Cite´, F-75205 Paris, France

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Contributors

Agustina D’Urso Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Defne Emel Egecioglu Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Nathalie Eisenhardt Friedrich Miescher Laboratory of the Max Planck Society, Tu¨bingen, Germany Jan Ellenberg Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany Manoja, B.K. Eswara Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada Birthe Fahrenkrog Institute for Molecular Biology and Medicine, Universite Libre´ de Bruxelles, Charleroi, Belgium Boris Fichtman Faculty of Medicine in the Galilee, Bar-Ilan University, Safed, Israel Jindriska Fiserova Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom Dirk Flemming Biochemie-Zentrum der Universita¨t Heidelberg (BZH), Im Neuenheimer Feld 328, Heidelberg, Germany Aure´lie G. Floch Institut Jacques Monod, CNRS, UMR 7592, Univ. Paris Diderot, Sorbonne Paris Cite´, F-75205 Paris, and Ecole Doctorale Ge`nes Ge´nomes Cellules, Universite´ Paris Sud-11, Orsay, France Douglass J. Forbes Cell and Developmental Biology, University of California, San Diego, California, USA Tomoko Funakoshi Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, and Department of Biochemistry, Faculty of Pharmaceutical Sciences, Toho University, Funabashi, Chiba, Japan Maiko Furuta Department of Molecular Genetics, National Institute of Genetics, The Graduate University for Advanced Studies Sokendai, Mishima, Shizuoka, Japan

Contributors

Vincent Galy Sorbonne Universite´s, UPMC, Univ Paris 06, and CNRS, UMR7622, IBPS, F-75005 Paris, France Martin W. Goldberg Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom Amnon Harel Faculty of Medicine in the Galilee, Bar-Ilan University, Safed, Israel Peter Horvath Institute of Biochemistry, ETH Zurich, Zurich, Switzerland Ed Hurt Biochemie-Zentrum der Universita¨t Heidelberg (BZH), Im Neuenheimer Feld, Heidelberg, Germany Naoko Imamoto Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan Simeon R. Johnson Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom Tijana Jovanovic-Talisman Department of Molecular Medicine, Beckman Research Institute of the City of Hope Comprehensive Cancer Center, Duarte, California, USA Samir Karaca Bioanalytical Mass Spectrometry Group, Max Planck Institute for Biophysical Chemistry, Go¨ttingen, Germany Ralph H. Kehlenbach Department of Molecular Biology, Faculty of Medicine, Georg-August-University of Go¨ttingen, Go¨ttingen, Germany Ralph H. Kehlenbach Department of Molecular Biology, Faculty of Medicine, Georg-August-University of Go¨ttingen, Go¨ttingen, Germany Makoto Kimura Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan Elena Kiseleva Laboratory of Morphology and Function of Cell Structure, Institute of Cytology and Genetics, Russian Academy of Science, Novosibirsk, Russia Shingo Kose Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan Ulrike Kutay Institute of Biochemistry, ETH Zurich, Zurich, Switzerland

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William H. Light Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Kazuhiro Maeshima Cellular Dynamics Laboratory, RIKEN, Wako, Saitama; Biological Macromolecules Laboratory, Structural Biology Center, National Institute of Genetics, and Department of Genetics, School of Life Science, Graduate University for Advanced Studies (Sokendai), Mishima, Shizuoka, Japan Dev Mangroo Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada Joseph Marino Institute of Biochemistry, ETH Zurich, Zurich, Switzerland Monika I. Mayr Institute of Biochemistry, ETH Zurich, Zurich, Switzerland Asako McCloskey Institute for Virus Research, Kyoto University, Kyoto, Japan Peter Meister Cell fate and Nuclear Organization, Institute of Cell Biology, University of Bern, Baltzerstrasse 4, Bern, Switzerland Mutsuhito Ohno Institute for Virus Research, Kyoto University, Kyoto, Japan Alessandro Ori European Molecular Biology Laboratory, Structural and Computational Biology Unit, Meyerhofstr. 1, 69117, Heidelberg, Germany Shotaro Otsuka Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany Benoit Palancade Institut Jacques Monod, CNRS, UMR 7592, Univ. Paris Diderot, Sorbonne Paris Cite´, F-75205 Paris, France Vikram Govind Panse Institute of Biochemistry (IBC), ETH Zu¨rich, Otto-Stern-Weg 3, Zurich, Switzerland Nelly Pante´ Department of Zoology, Life Sciences Centre, University of British Columbia, Vancouver, Canada Jacqueline B. Pierce Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada

Contributors

Christopher Ptak Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada A. Christine Richardson Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom Michael P. Rout Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, USA Allana Schooley Friedrich Miescher Laboratory of the Max Planck Society, Tu¨bingen, Germany Sabina Schu¨tz Institute of Biochemistry (IBC), ETH Zu¨rich, Otto-Stern-Weg 3, and MLS Program, Life Science Zurich Graduate School, Winterthurerstrasse 190, Zurich, Switzerland Lihi Shaulov Faculty of Medicine in the Galilee, Bar-Ilan University, Safed, Israel Matthew C. Spink Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom Philipp Stelter Biochemie-Zentrum der Universita¨t Heidelberg (BZH), Im Neuenheimer Feld, Heidelberg, Germany Anton A. Strunov Laboratory of Morphology and Function of Cell Structure, Institute of Cytology and Genetics, Russian Academy of Science, Novosibirsk, Russia Anna Szymborska Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany Ichiro Taniguchi Institute for Virus Research, Kyoto University, Kyoto, Japan Ketan Thakar Department of Molecular Biology, Faculty of Medicine, Georg-August-University of Go¨ttingen, Go¨ttingen, Germany George van der Merwe Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada Cornelia Wandke Institute of Biochemistry, ETH Zurich, Zurich, Switzerland Richard W. Wozniak Department of Cell Biology, University of Alberta, Edmonton, Alberta, Canada

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Preface Bidirectional transport between the cytoplasm and the nucleoplasm is a finely tuned process, whose regulation is critical for the proper function of numerous biological pathways. Exchanges between these two compartments take place through nuclear pore complexes (NPCs), huge macromolecular assemblies embedded within the nuclear envelope (NE). Since the unearthing of this research field, more than 50 years ago, the combination of a large variety of approaches, performed in a broad range of model organisms, has permitted the deciphering of the main features of NPCs, their assembly and dynamics, as well as the precise rules governing and regulating nucleocytoplasmic exchanges. Although this field of research is extremely dynamic and has both contributed to and benefited from a wealth of innovating methods over recent years, no dedicated volume had been devoted to this topic since Elsevier published the Methods volume “Nuclear cytoplasmic transport and nuclear pore complexes” in 2006 (Volume 39(4): 275–380, ed. B.M.A. Fontoura). As this topic impinges on multiple fields, scattered protocols could however be found in volumes devoted to nuclear architecture, gene expression, small GTPases, or specific model systems. It was thus timely to set up a dedicated volume, and I am grateful to Phong Tran, senior editor of this series, for stimulating me to assemble this book, and to the many leaders in the field who have devoted some of their precious time to contribute to specific chapters. Following the introduction, that presents an overview of the history and of the main tools and rules governing nuclear transport, the 21 chapters in this volume provide protocols developed in the most widespread model systems in the field, namely mammalian cells in culture (Chapters 2, 6, 10–12, 15, 16, 18, 19, and 21), yeast Saccharomyces cerevisiae (Chapters 3, 5, 7, 14, and 19–21), Xenopus laevis oocytes and eggs extracts (Chapters 2, 4, 8, 9, and 18), and Caenorhabditis elegans (Chapter 13). The first chapters of this volume mainly focus on methods that have enabled impressive progress in our understanding of NPC structure and dynamics. These include the visualization of the NPC architecture and the localization of its constituents (the nucleoporins, Nups) that have largely benefited from transmission and scanning electron microscopy approaches (Chapters 2–4 and 13) and more recently from super-resolution imaging (Chapter 10), and recent protocols designed to study large Nups or NPC subcomplexes by negative staining (Chapter 5), to allow the precise determination of Nup stoichiometry (based on NE purifications and targeted proteomics, Chapter 6), or to follow the in vivo assembly of newly synthesized proteins within complexes (Chapter 7). While such approaches have contributed to refine our view of NPC organization, they no doubt could be usefully applied to improve our understanding of any other complex macromolecular assemblies. NPCs are dynamic structures whose disassembly and reassembly upon open mitosis (Chapters 10 and 13) or de novo assembly during interphase (Chapter 11) can be monitored in a quantitative manner using live cell imaging. In addition, in vitro assays based on nuclei assembled using cell-free extracts of Xenopus eggs (Chapters 2, 8, and 9) or on

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semi-permeabilized mammalian cells (Chapters 12 and 15) have been adapted and refined to decipher the mechanisms that underlie these dynamic changes. To assess the critical function of nuclear pores in bidirectional transport of macromolecules, specific reporters and methods have been designed in each model organism to measure nuclear permeability (Chapters 10, 12, and 13), protein import or export (Chapters 8, 9, 13, and 14), or the fate of the various classes of RNAs that are assembled into ribonucleoparticles (Chapters 18–20). With respect to nuclear protein transport, this volume includes recent adaptations to the well-established in vitro assay that relies on digitonin-permeabilized cells (Chapter 15), quantitative mass-spectrometry approaches to identify specific import or export substrates (Chapter 16), and an example of nanodevices that mimic functional NPCs (Chapter 17). Dedicated tools to follow and characterize RNA transport mechanisms are provided in Chapter 18 (that details microinjections in Xenopus oocytes and FISH studies in vertebrates), Chapter 19 (dedicated to tRNA dynamics), and Chapter 20 (ribosome assembly and transport). While not covered by specific chapters, all these tools can be adapted to study nucleocytoplasmic trafficking of a wealth of other macromolecular assemblies, notably viral particles. Finally, the nuclear transport machinery further plays a critical role in multiple cellular processes, including cell cycle regulations (see for instance Chapters 14 and 15), maintenance of genetic integrity, and gene expression. For the latter topic, protocols developed to analyze gene positioning and to evaluate the association of Nups with transcribed genes are detailed in Chapter 21 (and referred to in Chapter 13). Twenty one chapters cannot entirely cover the multiplicity of model systems and approaches so far used in this ever-expanding field, and I apologize to those whose protocols could not be included in this volume. However, all authors have clearly made considerable efforts to provide, in addition to step-by-step protocols that should enable scientists to reproduce their preferred methods, critical references to related or alternative techniques that have been developed to tackle similar questions. Other currently arising model organisms in the field (such as flies, plants, fungi, ciliates, or protozoans, to cite but a few) are unfortunately not included in this volume, and will clearly deserve specific chapters in the future. I hope that all readers, junior scientists joining the nuclear transport field or already acquainted with it as well as outsiders of other fields, will appreciate, as much as I did, going through these carefully detailed methods chapters. I am particularly grateful to all authors for their commitment to this collective project. I would also like to thank the editors at Elsevier, Sarah Lay and Zoe Kruze, for their efficient help during this process, and all my collaborators at the Institut Jacques Monod for their constant support and understanding over these last busy months. Vale´rie Doye

CHAPTER

Fifty Years of Nuclear Pores and Nucleocytoplasmic Transport Studies: Multiple Tools Revealing Complex Rules

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Aure´lie G. Floch*,{, Benoit Palancade*, and Vale´rie Doye* *

Institut Jacques Monod, CNRS, UMR 7592, Univ. Paris Diderot, Sorbonne Paris Cite´, F-75205 Paris, France { Ecole Doctorale Ge`nes Ge´nomes Cellules, Universite´ Paris Sud-11, Orsay, France

CHAPTER OUTLINE Introduction ................................................................................................................ 2 1.1 The NPCs: A Modular Macromolecular Assembly ................................................... 3 1.1.1 Toward a Refined View of the NPC Structure......................................... 3 1.1.2 Nucleoporins: The Building Blocks of NPCs .......................................... 6 1.1.3 Integrating the Nups Into the 3D Architecture of the NPC ...................... 8 1.1.3.1 Nup Localization...........................................................................8 1.1.3.2 Interactions Among Nups .............................................................9 1.1.3.3 Nup Stoichiometry ......................................................................10 1.1.3.4 Toward a Detailed Map of NPCs..................................................10 1.1.4 Nups are Composed of a Limited Set of Structural Domains ................. 10 1.2 Nucleocytoplasmic Trafficking: The Rules of the Road ......................................... 12 1.2.1 Investigating Nucleocytoplasmic Transport ......................................... 12 1.2.1.1 Historical Overview .....................................................................12 1.2.1.2 Expanding the Toolbox................................................................14 1.2.2 The Signals for Nucleocytoplasmic Exchanges .................................... 15 1.2.3 A Family of Protein Transport Receptors: The Karyopherins .................. 16 1.2.4 The Ran GTPase: A Key to Transport Directionality.............................. 17 1.2.5 The Case of INM Targeting ................................................................ 19 1.2.6 Distinct Pathways Contribute to RNA Export ....................................... 20 1.2.7 Translocation Across the NPCs: A Dual Function for FG-Nups as Barrier and Gate ............................................................. 21 1.2.8 Noncanonical Transport Pathways through the NE ............................... 23

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00001-1

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1.3 The Nuclear Transport Machinery: A Dynamic and Versatile Device ...................... 23 1.3.1 NPC Biogenesis Throughout the Cell Cycle ......................................... 24 1.3.1.1 NPC Disassembly .......................................................................24 1.3.1.2 Post-mitotic NPC Assembly.........................................................24 1.3.1.3 De novo NPC Assembly ..............................................................26 1.3.2 Multiple Functions of the Nuclear Transport Machinery During the Cell Cycle ........................................................................ 27 1.3.3 NPCs, Nuclear Organization, and Gene Expression .............................. 29 1.3.4 NPCs and Genetic Stability ............................................................... 30 Concluding Remarks ................................................................................................. 31 Acknowledgments ..................................................................................................... 31 References ............................................................................................................... 32

Abstract Nuclear pore complexes (NPCs) are multiprotein assemblies embedded within the nuclear envelope and involved in the control of the bidirectional transport of proteins and ribonucleoparticles between the nucleus and the cytoplasm. Since their discovery more than 50 years ago, NPCs and nucleocytoplasmic transport have been the focus of intense research. Here, we review how the use of a multiplicity of structural, biochemical, genetic, and cell biology approaches have permitted the deciphering of the main features of this macromolecular complex, its mode of assembly as well as the rules governing nucleocytoplasmic exchanges. We first present the current knowledge of the ultrastructure of NPCs, which reveals that they are modular and repetitive assemblies of subunits referred to as nucleoporins, associated into stable subcomplexes and composed of a limited set of protein domains, including phenylalanine-glycine (FG) repeats and membrane-interacting domains. The outcome of investigations on nucleocytoplasmic trafficking will then be detailed, showing how it involves a limited number of molecular factors and common mechanisms, namely (i) indirect association of cargos with nuclear pores through receptors in the donor compartment, (ii) progression within the channel through dynamic hydrophobic interactions with FG-Nups, and (iii) NTPase-driven remodeling of transport complexes in the target compartment. Finally, we also discuss the outcome of more recent studies, which indicate that NPCs and the transport machinery are dynamic and versatile devices, whose biogenesis is tightly coordinated with the cell cycle, and which carry nonconventional duties, in particular, in mitosis, gene expression, and genetic stability.

INTRODUCTION One of the major evolutionary steps that occurred at the cellular level was the acquisition of internal compartments, enclosed by lipid membranes. Membrane internalization and organelle formation provided a major evolutionary advantage: by

1.1 The NPCs: A Modular Macromolecular Assembly

simultaneously carrying out different functions within these distinct compartments, cells increased their robustness and complexity. The nucleus, observed for the first time by Antonie van Leeuwenhoek in the seventeenth century, is the defining organelle that distinguishes eukaryotic from prokaryotic cells. Its boundary, the nuclear envelope (NE), sequesters the genetic material from the cytoplasm. This separation notably enables eukaryotic cells to spatially and temporally regulate distinct stages of genome expression (mRNA transcription, maturation, translation, and decay). However, the nuclear content is not totally isolated from the cytoplasm, thanks to a gating system called nuclear pore complex (or NPC). These structures, localized at the points of fusion between the outer and inner nuclear membrane of the NE, connect the nucleoplasm to the cytoplasm and allow the bidirectional trafficking of a flow of cellular components. While small molecules (water, sugars, and ions) can freely translocate though the NPCs, large macromolecules (proteins and nucleic acids) and even megadalton-sized macromolecular complexes such as ribosomal subunits or viral particles undergo highly selective nuclear import and/or export processes (Fig. 1.1). Regulating this transport is a crucial issue for the cell, both for the maintenance of nuclear identity and for the control of gene expression. It requires the assembly and maintenance of stable NPCs as well as finely tuned cargo/ transport receptor systems that organize bidirectional transport and provide selectivity. Over the past 50 years, the combination of a huge variety of approaches, performed in a broad range of model organisms, has provided the main rules governing nuclear pore assembly and nucleocytoplasmic transport. In this chapter, we give an overview of the main approaches that have been used in the field to elucidate the fundamental principles that govern NPC organization and nucleocytoplasmic traffic, and to demonstrate the dynamic and versatile nature of the nuclear transport apparatus that also exerts duties beyond transport.

1.1 THE NPCS: A MODULAR MACROMOLECULAR ASSEMBLY 1.1.1 Toward a refined view of the NPC structure In 1950, Callan and Tomlin, who observed the giant nuclei of amphibian oocytes by electron microscopy (EM), were the first to report that pores, organized in annular structures, pierced the NE. Franke (1966) and Gall (1967) subsequently described in various species a macromolecular structure, embedded at fusion points of the NE bilayer, with a eightfold rotational symmetry (Fig. 1.2A). Since then, NPCs have been imaged in multiple organisms (see introduction to Chapter 2 and references therein; Wente & Rout, 2010). The density of NPCs on the NE is most frequently in the range of 3–15 per mm2 of NE, leading to 1000–5000 NPCs per mammalian nucleus, and 100–200 in the much smaller yeast nuclei. In contrast, the large nuclei from amphibian oocytes (0.4 mm diameter) are characterized by a very high NPC density (60 NPCs/mm2) and contain about 50 million pores. Xenopus laevis has thus been an extensively studied model system to uncover NPC structure

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FIGURE 1.1 An Overview of Nucleocytoplasmic Exchanges The main nuclear processes involving nucleocytoplasmic transport of proteins and RNA– protein complexes are represented: chromatin assembly; DNA metabolism; RNA synthesis/ processing; and ribosome biogenesis. Note that viral genomes can enter the nucleus through NPCs either as intact viral particles or upon disassembly of the viral capsids on the cytoplasmic side. ONM, outer nuclear membrane; INM, inner nuclear membrane; and ER, endoplasmic reticulum.

(see Chapters 2 and 4 and references therein). Although their overall architecture appears to be conserved with little change during evolution, NPC diameter varies between 100 and 150 nm, and thickness by 50–70 nm. Reflecting this variation, the total mass of NPCs was initially estimated to be 125 MDa in vertebrates, but only 60 MDa in yeast (reviewed in Stoffler, Fahrenkrog, & Aebi, 1999). The overall NPC architecture is composed of three rings: the nuclear and cytoplasmic rings that sandwich a central spoke ring delineating a central channel of 40 nm. Anchored to this membrane-embedded central framework, peripheral NPC components extend into the cytoplasm and the nucleoplasm. These filamentous structures, although detectable by thin-section EM (see Chapter 4), are best visualized by

1.1 The NPCs: A Modular Macromolecular Assembly

FIGURE 1.2 The Ultrastructure of Nuclear Pore Complexes (A) a. Pioneer EM image of a nuclear pore from a negatively stained preparation of the oocyte envelope from a newt, Triturus, revealing its eightfold rotational symmetry. b. Original schematic representations including the sizes and a three-dimensional view of a nuclear pore in the double-layered nuclear envelope. (B) Scanning EM micrographs of nuclear pores from diverse eukaryotes revealing cytoplasmic (top panels) and nucleoplasmic (lower panels) extensions. Scale bar, 100 nm. (C) Top and lateral views of the cryo-EM structure of human NPCs at 3.2 nm resolution. The cytoplasmic (CR), spoke (SR), and nuclear rings (NR) are indicated and the pore membrane appears in yellow. An additional inner density likely corresponding to molecules with high structural plasticity is colored in purple. Dotted lines highlight shapes similar to the structure of some members of the Nup93 complex. The diameters and thickness of NPCs are indicated. (A) Reproduced from Gall (1967) # (1967), with permission from the author and The Rockefeller University Press. (B) Images provided by Elena Kiseleva and reproduced from Brohawn, Partridge, Whittle, and Schwartz (2009), with permission from Elsevier. (C) Reproduced from Bui et al. (2013) # (2013), with permission from Elsevier.

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CHAPTER 1 Introduction to NPC and Nuclear Transport

scanning electron microscopy (SEM) (Fig. 1.2B; see also Chapter 2). The eight cytoplasmic filaments appear as 50-nm-long disordered structures, whereas the so-called “nuclear basket” is composed of eight filaments (95 nm long in yeast and 120 nm in vertebrates) that join to form a nuclear ring (reviewed in Stoffler et al., 1999). The complexity and size of NPCs have so far precluded the determination of their whole structure at atomic resolution. However, introduction of cryo-electron tomography (cryo-ET) combined with in silico subtomogram averaging procedures has led to substantial progress by refining the 3D architecture of the NPCs to 6 nm resolution (Beck, Lucic, Forster, Baumeister, & Medalia, 2007; Frenkiel-Krispin, Maco, Aebi, & Medalia, 2010; Maimon, Elad, Dahan, & Medalia, 2012) and more recently to 3.2 nm for NPCs imaged from purified human NEs (see Chapter 6; Bui et al., 2013; Fig. 1.2C). These studies validated the dimensions of the NPCs and their overall architecture. They also confirmed that, in addition to the 40-nm-wide central channel, the cytoplasmic and nuclear rings are not entirely apposed to the outer nuclear membrane (ONM) and inner nuclear membrane (INM), but leave peripheral openings of 10 nm which traverse the spoke ring complex and could allow passage of globular particles up to a size of 5 nm.

1.1.2 Nucleoporins: The building blocks of NPCs The composition of NPCs has been elucidated by the combination of multiple biochemicals and genetic approaches. The molecular characterizations of the pore constituents (called nucleoporins or Nups) started in the 1980s by the production of antibodies, notably monoclonal antibodies that bind in a polyspecific manner to multiple Nups. For instance, Gu¨nter Blobel’s lab developed the widely used mouse mAb414 antibody that first led to the characterization of mammalian Nup62 (Davis & Blobel, 1986). However, this antibody in fact recognizes a subset of Nups called “FG-Nups” (see below) and presents a wide interspecies cross-reactivity in eukaryotes. In budding yeast, genetic screens and total genome sequencing also led to the description of new sets of Nups (for review, see Doye & Hurt, 1997). Another important step was the development of approaches based on biochemical purification of Nups subcomplexes, coupled with mass spectrometry analysis. These techniques led to the characterization of multiple vertebrates and yeast Nups. This culminated with the purifications of yeast and mammalian NPCs and their analysis by mass spectrometry (Cronshaw, Krutchinsky, Zhang, Chait, & Matunis, 2002; Rout et al., 2000 and references therein). In these studies, bona fide Nups were validated through observation, upon tagging with GFP, of a typical punctate NE labeling in live cells. Since then, only a few novel Nups have been identified (Fig. 1.3A). Together, these studies have revealed that, despite their huge size, NPCs are composed of only 30 Nups (Fig. 1.3A). However, due to the eightfold symmetry of the assembly, and the additional twofold symmetry of the main scaffold relative to the axis of the NE, each Nup is present in multiple copies leading to an assembly of 500–1000 proteins.

1.1 The NPCs: A Modular Macromolecular Assembly

FIGURE 1.3 The Molecular Composition of NPCs (A) Model of the organization of nucleoporins within the NPC framework in vertebrates (left) and budding yeast (right). FG repeats containing nucleoporins are circled in red. Proteins carrying enzymatic activities appear in bold. NPC-associated proteins other than bona fide nucleoporins are circled with dotted lines. The Y-complex appears in blue. The hNup214/ yNup159–Nup82 subcomplexes appear in yellow. (B) a. Structural model of the human Y-complex. Available crystal structures were fitted within the EM structure of the Y-complex vertex. The arrows and dashed ellipses indicate the position of flexible hinges and static connectors. b, c. Inner and outer Y-complexes segmented from the cryo-EM structure of human NPCs. (C) Positioning of the Y-complex (in blue) and Nup82 (in yellow) within the computational model of yeast NPC. (D) Positioning of two Y-complex rings (in blue and green) and of the Nup214 complex (in yellow) within the cryo-EM structure of the human NPC. (A) Modified from Wozniak, Burke, and Doye (2010) # (2010), with permission from Springer. (B) Reproduced from Bui et al. (2013) # (2013), with permission from Elsevier. (C) Modified from Alber et al. (2007b) # (2007), with permission from Nature Publishing Group. (D) Reproduced from Bui et al. (2013) # (2013), with permission from Elsevier.

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1.1.3 Integrating the Nups into the 3D architecture of the NPC Having a list of constituents in hand, a prerequisite for a detailed understanding of the transport mechanism is to precisely integrate these Nups into the complex NPC architecture.

1.1.3.1 Nup localization A first requirement is to determine the localization of each Nup within the assembly. With the exception of integral membrane Nups, which are evidently positioned within the pore membrane, the localization of most Nups cannot be inferred from their primary sequence. Thin-section EM in combination with immunogold labeling has been the main approach used to assess their localization within the NPCs. These studies were performed on isolated Xenopus nuclei, but also in mammalian or yeast cells using specific antibodies, or tagged Nups (see Chapters 3 and 4 and references therein). In yeast, nearly all Nups were systematically localized by immuno-EM using strains carrying a C-terminal protein A tag (Alber et al., 2007a; Rout et al., 2000). For specific Nups, immunogold labeling performed on SEM samples can further provide a combined view of their localization with surface topology (see Chapter 2). Unlike EM, the resolution of fluorescence microscopy has long impaired the refined localization of Nups within the NPCs. However, the use of low concentrations of digitonin, that permeabilizes the plasma membrane but not the NE, can provide clues on the restricted localization of specific Nups on the nuclear side of the NPCs (see for instance, Bastos, Lin, Enarson, & Burke, 1996). In addition, discrimination of nuclear versus cytoplasmic localizations could be achieved for a few peripheral Nups using confocal microscopy (see for instance, Zhang, Saitoh, & Matunis, 2002). Together, these studies have revealed that although most Nups present a symmetrical localization, some are only localized on the cytoplasmic or nucleoplasmic face, or biased toward one side, conferring a global asymmetry to NPCs (Fig. 1.3A). Recently, super-resolution imaging, combined with single-particle averaging, has permitted the mapping of the average radial positions of individual fluorescent labels on Nups with nanometer precision, thus providing another approach for the precise mapping of Nups within the NPC framework (see Chapter 10). Importantly, this huge molecular assembly is not static, and a systematic analysis of multiple Nups using photobleaching experiments has revealed that in interphase mammalian cells, NPC components exhibit a wide range of residence times from seconds to days. Despite some exceptions, the central parts of the NPC appear to be very stable, consistent with a function as a structural scaffold, whereas peripheral components exhibit more dynamic behavior, suggesting adaptor as well as regulatory functions (Rabut, Doye, & Ellenberg, 2004). Of note, these studies were facilitated in vertebrate cells by the fact that NPCs are stably anchored within the nuclear lamina (Daigle et al., 2001). In contrast, the whole NPCs can diffuse within the plane of the NE in yeasts that lack a bona fide lamina (Belgareh & Doye, 1997; Bucci & Wente, 1997).

1.1 The NPCs: A Modular Macromolecular Assembly

1.1.3.2 Interactions among Nups Information on the interactions between Nups is also crucial to build an accurate picture of the NPC. Biochemical studies, frequently combined with functional assays, have revealed the assembly of subsets of Nups into stable heterooligomeric subcomplexes amenable to affinity purification experiments. These subcomplexes serve in turn as modular building blocks to form larger NPC structures. Despite considerable differences between the primary sequences of orthologous Nups, most yeast and mammalian modules have been well conserved throughout evolution. The best currently characterized modules are the so-called metazoan Nup107–160 (yeast Nup84) and Nup53–Nup93 (yeast Nup53–Nic96) subcomplexes which are essential architectural elements of the NPC scaffold, and the Nup62 (yeast Nsp1) subcomplex that constitutes a major transporter module (reviewed in Brohawn et al., 2009; Wente & Rout, 2010; see Fig. 1.3A for the positioning of the various NPC subcomplexes in vertebrates and budding yeast). The Y-shaped yeast Nup84/metazoan Nup107–160 complex is the best characterized NPC building block, as reflected by the extensive set of genetic, biochemical, structural, and functional data accumulated over the years (reviewed in GonzalezAguilera & Askjaer, 2012). Pioneering single-particle negative stain EM studies from the Hurt lab, using both isolated and in vitro-reconstituted complexes, revealed the characteristic Y-shaped assembly formed by the seven Nups that constitute the yeast Nup84 complex (Nup133, Nup120, Nup145C, Nup85, Nup84, Seh1, and Sec13). The computational integration of biochemical and structural data subsequently enabled the determination of its structure to a precision of 1.5 nm (Fernandez-Martinez et al., 2012 and references therein). In many eukaryotes, notably excluding Saccharomyces cerevisiae, the Y-complex contains two additional proteins, Nup37 and Nup43 (Loiodice et al., 2004). In addition, because of its stable interaction with the Nup107–160 complex, Elys/MEL-28, although solely localized to the nuclear side of the NPCs (see Fig. 1.3A; Bui et al., 2013 and references therein), is sometimes considered as a tenth member of this complex. Recently, a structural model of the hNup107–160 subcomplex was obtained by combining electron tomograms of affinity-purified subcomplex particles, spatial constraints obtained by cross-linking mass spectrometry, and previously available crystal structures or homology models (Bui et al., 2013, see Fig. 1.3B). The metazoan Nup93 complex (that comprises Nup93, Nup188, Nup205, Nup155, and Nup53—also termed Nup35 in vertebrates) forms the central spoke ring of the NPCs. Its constituents are also conserved and display similar interactions in budding yeast (see Fig. 1.3A). Two-hybrid screens and genetic analyses in yeast, combined with in vitro assembly of the orthologous Nups from the fungus Chaetomium thermophilum, enabled reconstitution of this NPC module in vitro (Amlacher et al., 2011). In this study, the structural DID-Dyn2 label, which is easily visible in the electron microscope, was used to precisely position specific Nups within this NPC subcomplex (see Chapter 5).

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1.1.3.3 Nup stoichiometry In addition to their approximate position and specific partners, the copy number of each Nup per NPC is another important parameter required to understand the molecular architecture of NPCs. The first insights toward Nups respective stoichiometry were provided by semiquantitative investigations of their relative abundance in budding yeast (based on immunoblot signals of Nups tagged with the same protein A moiety; Rout et al., 2000) and vertebrates (using quantifications of zinc or Coomassie-stained gels loaded with NPC-enriched fractions; Cronshaw et al., 2002). More recently, the combination of integrated targeted proteomics (see Chapter 6) and super-resolution microscopy approaches have enabled the determination of the absolute stoichiometry of the human NPC. This analysis revealed a stable stoichiometry of most NPC scaffold Nups in distinct human cell types whereas significant variations could be observed for peripheral Nups (Ori et al., 2013). This confirms earlier observations that NPC composition could vary between different tissues or during development (D’Angelo, Gomez-Cavazos, Mei, Lackner, & Hetzer, 2012 and references therein).

1.1.3.4 Toward a detailed map of NPCs With such parameters in hand, integrative approaches have been developed to resolve the molecular architecture of NPCs. A first approach involved the translation of multiple datasets into spatial restraints that were then used to generate an ensemble of structures consistent with the data. Analysis of this ensemble produced the first approximation of the molecular architecture of the S. cerevisiae NPC (Alber et al., 2007a, 2007b; Fig. 1.3C). More recently, by combining single-particle EM and cross-linking mass spectrometry, the Y-complex, as well as a few additional NPC constituents, could be precisely positioned within the refined cryo-ET structure of the human NPC scaffold (Bui et al., 2013). This revealed that each human NPC comprises 32 copies of the Y-shaped Nup107 complex that assemble into two reticulated rings (Fig. 1.3D). Of note, this organization differs from the yeast NPC model, in which only 16 copies of the Nup84 complex were previously integrated (Alber et al., 2007a,2007b). In the future, extending such a refined approach to additional NPC subcomplexes will be required to bridge length scales from overall molecular architecture down to atomic resolution. In addition, comparison of NPC architecture between distant species or distinct tissues/cell types should shed light on its adaptability and specialization.

1.1.4 Nups are composed of a limited set of structural domains Protein structure prediction analyses of Nups obtained in multiple species, including the divergent eukaryote Trypanosoma brucei, as well as atomic resolution of the structure of an increasing number of Nups, revealed that they are composed of a few repetitive structural domains that likely evolved from the duplication of a small set of precursors genes (reviewed in Aitchison & Rout, 2012; Brohawn et al., 2009; Hoelz, Debler, & Blobel, 2011).

1.1 The NPCs: A Modular Macromolecular Assembly

Although NPCs are anchored to the pore membrane, only a few Nups exhibit transmembrane domains. There are three pore membrane proteins (Poms) in mammals (Pom121, gp210, and Ndc1) and four in S. cerevisiae (Pom34, Pom152, Ndc1, and Pom33, of which the three first ones are associated to form the transmembrane ring; Onischenko, Stanton, Madrid, Kieselbach, & Weis, 2009) (Fig. 1.3A). There is apparently no strong selective pressure for their conservation, as among them, only Ndc1 and Pom33/TMEM33 are conserved throughout evolution (note that the NPC localization of vertebrate TMEM33 remains to be demonstrated). In addition, the deletion of the three known integral membrane Nups in Aspergillus nidulans is not lethal (Liu, De Souza, Osmani, & Osmani, 2009). While this may reflect the existence of yet unidentified Poms, there are also alternative modes of interaction between the NPC and the pore membrane. Indeed, non-integral membrane-binding modules have also been identified in several Nups. In particular, two distinct domains with membrane-binding properties were functionally characterized in mammalian Nup53, a constituent of the Nup93 complex that also interacts with Ndc1. The C-terminal domain of Nup53, predicted to form an amphipathic helix, was further shown to have membrane-deforming capabilities (Vollmer et al., 2012). In addition, ALPS motifs (amphipathic lipid-packing sensor, first described in the COPI-coat assembly protein ArfGAP1) were found by in silico analysis in several human and yeast Nups. The ALPS motif within human Nup133 (that belongs to the Y-shaped Nup107 complex) was demonstrated to effectively bind to curved membranes in vitro (Drin et al., 2007). These integral membrane proteins and membrane-binding domains anchor the NPCs within the NE, thanks to their interaction with additional core scaffold proteins. Noteworthy, most Nups of the core scaffold are built of a-solenoid folds (antiparallel pairs of a-helices organized to form a coil), b-propellers folds (6–7 bladeshaped b-sheet subunits arranged around a central axis), or a combination of a N-terminal b-propeller followed by a carboxy-terminal a-solenoid fold (reviewed in Brohawn et al., 2009). Despite primary sequence divergences, these domains are well conserved throughout evolution. As the combination of these two protein fold types is also found in coated vesicle components (COPI, COPII, and clathrin), it was proposed that they derive from an ancestral “protocoatomer,” that would have had the capacity to interact with and stabilize highly curved membrane surfaces (see DeGrasse et al., 2009 and references therein). Another protein fold present in a few Nups corresponds to a-helices with heptad repeat patterns (hxxhcxc, with h for hydrophobic and c for charged residues) that allow homologous or heterologous “coiled-coil” interactions. Extended coiled-coil domains, found in the vertebrate Nup Tpr and in its S. cerevisiae orthologs, Mlp1 and Mlp2, constitute the central element of the nuclear basket (Krull, Thyberg, Bjorkroth, Rackwitz, & Cordes, 2004). In addition, more restrained coiled-coil domains contribute to complex formation and NPC anchoring of “FG-Nups” (see Brohawn et al., 2009; Devos et al., 2006). Besides a domain contributing to their anchoring to NPCs, “FG-Nups” bear one of the most prevalent structural motifs found within Nups. The so-called “FG

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domains” correspond to 5–50 repeats of phenylalanine-glycine (FG), Phe-X-Phe-Gly (FXFG), or Gly-Leu-Phe-Gly (GLFG) residues, separated by variable hydrophilic or charged spacer sequences (for a review on FG-Nups, see Terry & Wente, 2009). Such repeats are found in 11 vertebrate “FG-Nups” and 13 S. cerevisiae Nups that localize within the NPC central channel, the cytoplasmic filaments, or the nuclear basket (Fig. 1.3A). FG domains are largely natively unfolded and form flexible filaments able to take a large scale of dynamic conformations as notably described for Nup153 (see Chapter 4 and references therein). This flexibility, along with the hydrophobic properties of these repeats, allows rapid association and dissociation of FG-Nups with a large range of partners giving to these Nups a key function in nucleocytoplasmic transport (detailed below). In metazoans, several of the FG-containing domains are modified by O-linked N-acetylglucosamine (O-GlcNAc) addition on serine or threonine residues (Holt et al., 1987). This modification leads to the recognition of several FG-Nups by wheat germ agglutinin (WGA), a lectin that was demonstrated in early studies to inhibit nuclear transport, possibly by generating steric hindrance (Finlay, Newmeyer, Price, & Forbes, 1987; see also Chapter 8). Because of the enrichment for this modification within Nups, WGA also provides a convenient tool to label NPCs in fixed or live cells (Hanover, Cohen, Willingham, & Park, 1987; Onischenko, Gubanova, Kiseleva, & Hallberg, 2005). While the biological significance of nuclear pore glycosylation has remained largely unknown, recent studies indicate that GlcNAc may modulate associations between specific Nups, Nup stability, and possibly NPC permeability (reviewed in Li & Kohler, 2014). Other post-translational modifications described in multiple FG and non-FGNups include phosphorylations (notably mediated by cell cycle-dependent kinases, see Section 1.3), sumoylation (Palancade & Doye, 2008), or ubiquitination (Hayakawa, Babour, Sengmanivong, & Dargemont, 2012).

1.2 NUCLEOCYTOPLASMIC TRAFFICKING: THE RULES OF THE ROAD 1.2.1 Investigating nucleocytoplasmic transport 1.2.1.1 Historical overview The first EM observations of NPCs were accompanied by the evidence of material in the process of translocating through the nuclear pores (Anderson & Beams, 1956 and references therein). EM studies in Chironomus salivary glands subsequently enabled the visualization of the nuclear export of Balbiani ring granules that correspond to huge ribonucleoprotein particles (RNPs) (Stevens & Swift, 1966). Independently, pulse chase experiments combined with subcellular fractionation, as well as nuclear transplantation experiments, have revealed the transfer of proteins from the cytoplasm to the nucleus in HeLa cells (Byers, Platt, & Goldstein, 1963; Speer & Zimmerman, 1968). Binucleated cells mainly obtained

1.2 Nucleocytoplasmic Trafficking: The Rules of the Road

by cell-fusion/heterokaryon assays further enabled the characterization of the shuttling of proteins between the cytoplasm and the nucleus (see methods and references in Chapter 11 for mammalian cells and in Chapter 20 for budding yeast). Other early transport studies were based on thin-section EM observations or localization of radiolabeled molecules by autoradiography following microinjections, notably into Xenopus oocytes (Feldherr, 1965, 1969). Thanks to their large size, the nucleus and cytoplasm of amphibian oocytes can be manually separated at various times after injection to determine the fate of the microinjected molecules (see Bonner, 1975 and references therein). In the past, cytoplasmic or nuclear microinjections of colloidal gold particles, amorphous dextrans of various sizes, recombinant proteins, or various RNA species have been successfully employed (for reviews and references related to these techniques, see Chapters 4 and 18). Together, these early studies have revealed the main features of nucleocytoplasmic transport, namely the behavior of NPCs as sieve-like barriers, freely permeable for small molecules, but able to selectively import or export molecules larger than the passive diffusion limit (30–50 kDa, or 5 nm in size) in an energy-dependent manner. In vitro nuclear protein import assays were first performed on isolated or in vitroassembled nuclei incubated with Xenopus extracts and a fluorescently labeled nuclear protein (Newmeyer, Finlay, & Forbes, 1986). However, one of the major advances in the field has been the development of an in vitro transport assay based on digitoninpermeabilized vertebrate cells (Adam, Marr, & Gerace, 1990). In this assay, the plasma membrane of tissue culture cells is specifically permeabilized with digitonin and the system is then supplied with cell extracts or purified transport factors to reconstitute nucleocytoplasmic transport in a semi cell-free environment. Coupled with biochemical fractionations and analyses, these experiments have led to the identification of soluble transport factors required for NPC targeting and nuclear translocation of distinct types of proteins, namely karyopherins/importins and the small GTPase Ran (detailed below) (see Chapters 15 and 16 and references therein). Genetic screens of yeast mutant collections (mainly S. cerevisiae) have also enabled the identification of multiple factors contributing to nucleocytoplasmic transport. Among them, a screen for mutants that missorted a chimeric NLS-cytochrome c1 protein to the mitochondria, allowing growth on glycerol, led to the characterization of several npl (nuclear protein localization) mutants (Sadler et al., 1989). Temperaturesensitive mtr (mRNA transport defective, Kadowaki, Zhao, & Tartakoff, 1992) mutants were initially identified based on their survival at restrictive temperature in the presence of toxic amino acid. Their contribution to mRNA export was then validated using fluorescence in situ hybridization (FISH) using oligo-dT probes. In addition, FISH-based screens identified multiple rat (ribonucleic acid trafficking, Amberg, Goldstein, & Cole, 1992) and brr (bad response to refrigeration, de Bruyn Kops & Guthrie, 2001) mutants impaired for mRNA export. Besides export factors, these screens also uncovered multiple Nups (notably members of the yeast Nup84 complex), as well as factors involved at various stages of mRNA metabolism. Visualization of GFP-tagged ribosomal protein localization (detailed in Chapter 20) and an in vivo

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nuclear tRNA export assay combined with overexpression screens (see Chapter 19) also contributed to identify additional export factors. In addition, synthetic lethal or high-copy suppressor screens based on Nup or nuclear transport mutant yeast strains uncovered multiple players including Gle1 and Gle2 (GLFG lethal), and key RNA export factors such as Mex67 and Los1 (reviewed in Doye & Hurt, 1997). Biochemical approaches, as well as yeast two-hybrid and three-hybrid interaction screens, have also uncovered multiple RNA-binding proteins of which some were subsequently demonstrated to contribute to the export of specific RNA species (Anderson, Wilson, Datar, & Swanson, 1993; see Chapters 19 and 20). Finally, because viruses often exploit nucleocytoplasmic transport pathways to facilitate viral replication or escape the host antiviral response, the study of viral interactions with the host cellular machinery also provided key tools that notably contributed to the discovery of nuclear localization sequences and several mRNA export pathways in vertebrates (reviewed in Yarbrough, Mata, Sakthivel, & Fontoura, 2013).

1.2.1.2 Expanding the toolbox Once identified, the contribution of specific factors in defined yeast nuclear transport pathways can be further assessed using adequate functional studies. These notably include the use of fluorescently tagged cargos or reporters (see Leslie, Timney, Rout, & Aitchison, 2006; Chapters 14 and 20 and references therein) or FISH approaches (see Chapters 19 and 20; and see also Rahman & Zenklusen, 2013 for single-molecule resolution FISH). Likewise, multiple tools are nowadays available to study bidirectional transport of macromolecules in metazoans cells (for specific tools available in Caenorhabditis elegans, Drosophila melanogaqster, Arabidopsis thaliana, and Aspergillus nidulans, see Chapter 13; Mason & Goldfarb, 2009; Meier & Brkljacic, 2010; and Markina-Inarrairaegui et al., 2011, respectively). Besides the widely used GFP, the photoswitchable fluorescent protein Dronpa can be used to assess NE permeability, and, when fused to reporter proteins, to study the kinetics of active import/export through the NPC (Ando, Mizuno, & Miyawaki, 2004; see also Chapter 10, Section 10.2). Various export assays for RNAs have been developed (see Chapter 18), and thanks to improved light microscopy approaches, single messenger ribonucleoprotein (mRNP) imaging and tracking in Chironomus tentans salivary gland cells or mammalian cells can now be used to monitor their export kinetics (Kalo, Kafri, & Shav-Tal, 2013; Kaminski, Spille, Nietzel, Siebrasse, & Kubitscheck, 2013). Structural studies of transport factors have also provided considerable insight into the elaborate protein–protein interactions that orchestrate nucleocytoplasmic transport (for reviews, see Chook & Suel, 2011; Conti, Muller, & Stewart, 2006; Lott & Cingolani, 2011; Stewart, 2010). To better characterize the selectivity and permeability properties of the NPC and the contribution of FG-Nups in this process, biophysical tools have been developed, including FG repeat hydrogels that display permeability properties very similar to authentic NPCs (Labokha et al.,

1.2 Nucleocytoplasmic Trafficking: The Rules of the Road

2013 and references therein) and NPC mimics (see Chapter 17 by and references therein). This vast diversity of experimental model organisms and approaches has now facilitated the uncovering of the main molecular players and mechanisms contributing to the fine-tuning of multiple nucleocytoplasmic transport pathways. As detailed below, active nuclear transport of macromolecules requires specific signals, shuttling nuclear transport receptors recognizing these signals, transport-associated NTPases, which account for the energy requirement of the process, and specific Nups.

1.2.2 The signals for nucleocytoplasmic exchanges As a letter needs an address to reach its destination, cargos need specific signaling sequences to either enter or exit the nucleus. Such amino acid sequences, named NLS (nuclear localization signal) and NES (nuclear export signal), are defined as being both necessary and sufficient to target proteins into and out of the nucleus, respectively. The first “classical” NLSs (cNLSs), initially identified in simian virus (SV40) large-T antigen (Kalderon, Roberts, Richardson, & Smith, 1984), are short stretches of positively charged residues (lysine and arginine). While these monopartite cNLSs are found in 20–30% of the nuclear proteins, another 12–30% of them contain a more complex “bipartite” cNLS, first characterized in nucleoplasmin (Dingwall, Robbins, Dilworth, Roberts, & Richardson, 1988) and composed of two clusters of charged residues separated by a spacer of 10 amino acids (for review, see Marfori et al., 2011). However, multiple non-classical NLSs have also been identified (for review, see Chook & Suel, 2011; Chapters 14 and 16). Note that unlike N-terminal sorting signals, NLS sequences are not cleaved and do not display any specific position within the protein sequence. Such potential NLS can frequently be predicted (see Marfori et al., 2011 and references therein). However, a substantial number of proteins that are imported into the nucleus feature signals that are not detected by NLS-based models. In addition, functional identification of a NLS is sometimes further complicated by the coexistence of multiple distinct NLSs within a given protein. Characterization of NESs has lagged considerably behind the analysis of NLSs. Typical NES sequences, also termed “leucine-rich NESs,” were first described in the protein kinase inhibitor and human immunodeficiency virus type 1 (HIV-l) Rev protein (Fischer, Huber, Boelens, Mattaj, & Luhrmann, 1995; Wen, Meinkoth, Tsien, & Taylor, 1995). These are short stretch of hydrophobic amino acids with variable spacing (typically Fx2Fx3Fx2–3FxF or FPxFx2FxF, where F is an hydrophobic residue, most frequently a leucine; for reviews, see Guttler & Gorlich, 2011; Kutay & Guttinger, 2005). Although algorithms can be used to predict such NESs, the fact that they share sequence similarity to regions that form the hydrophobic cores of many proteins impairs their identification (see Chapter 16 and references therein). In addition, as for NLSs, non-canonical export sequences have also been characterized (reviewed in Hutten & Kehlenbach, 2007).

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1.2.3 A family of protein transport receptors: The karyopherins As mentioned previously, in vitro transport assays combined with biochemical analyses have led to the characterization of soluble factors necessary for efficient transport, notably the karyopherins (or Kaps, from the greek karyon, nucleus and pherein, to bring). Karyopherins-b, also termed importins, exportins, or transportins, are all involved in nuclear import or export and share a similar architecture: multiple HEAT repeats (helix-loop-helix), that contribute to substrate recognition and binding to FG repeats present in Nups, and a N-terminal Ran
GTP-binding domain (see below). Despite their low sequence identity (20%), their organization and similar molecular weights (95–145 kDa) suggest that the Kaps-b probably evolved from a common ancestor. So far, 14 Kaps-b have been identified in yeast (including 3 exportins and 1 bidirectional karyopherin) and 19 have been characterized in mammalian cells (including 6 exportins and 2 bidirectional karyopherins) (for a comprehensive table, see Tran, Bolger, & Wente, 2007; see also Chook & Suel, 2011; Guttler & Gorlich, 2011; Chapters 14 and 16). There are two ways for karyopherins to recognize their cargos: a direct binding or an indirect binding via an adaptor protein. The latter mechanism was the one initially described. Indeed, both monopartite and bipartite cNLSs are recognized by adaptors of the importin-a family (Kap-a). While there is only one importin-a in budding yeast (Kap60), multiple subtypes exist in most metazoans, thus providing an additional means to finely regulate nuclear import of distinct substrates (reviewed in Goldfarb, Corbett, Mason, Harreman, & Adam, 2004; Yasuhara, Oka, & Yoneda, 2009). Importins-a bind cNLSs via three a-helix repeats called armadillo (ARM) domains. They also contain an autoinhibitory N-terminal domain, named IBB (importin-b binding), whose binding to the first described karyopherinb protein (importin-b/Kapb1; Kap95 in budding yeast) releases the ARM domains for cargo binding. A similar IBB domain is also found in a few other adaptor proteins including snurportin (that contributes to the reimport of U snRNAs, see Chapter 18). The IBB thus functions as a specialized NLS that evolved to transport cargos together with importin-b (reviewed in Lott & Cingolani, 2011). However, importin-b, as most other importins, can also bind directly to numerous cargos. The identification of distinct binding sites and conformations further indicates that Kapb1 may possibly interact simultaneously with distinct cargos (reviewed in Chook & Suel, 2011). Conversely, there is a strong redundancy between the importins, especially for cargos with essential functions (Tran et al., 2007; for recent strategies to identify importin-specific cargos, see Chapter 16). This leads to a complex system, where a discrete number of transporters are able to import a huge number of nuclear proteins, thereby possibly providing an additional level of physiological regulation. A major breakthrough in the field of protein export was the demonstration that export of “leucine-rich” NES-bearing proteins is sensitive to a cytotoxic drug, leptomycin B (LMB) (Wolff, Sanglier, & Wang, 1997). Indeed, a mutant allele of Schizosaccharomyces pombe crm1 (Chromosome Region Maintenance/Exportin-1,

1.2 Nucleocytoplasmic Trafficking: The Rules of the Road

Xpo1 in budding yeast) had been independently identified in a genetic screen for LMB resistance (Nishi et al., 1994). Subsequently, several studies demonstrated that Crm1, which belongs to the Kap-b family, is the export receptor for “leucine-rich” NESs. LMB covalent attachment to a cysteine residue within Crm1 interferes with NES binding in most cells (but not in S. cerevisiae, see Chapter 14; reviewed in Guttler & Gorlich, 2011; Kutay & Guttinger, 2005; see also Chapter 16 and references therein). Subsequently, other exportins have been identified, that unlike Crm1, export a narrow range of cargos (reviewed in Guttler & Gorlich, 2011). Among them CAS (also called Exportin-2/Xpo2, Cse1 in S. cerevisiae) plays a key function in nucleocytoplasmic transport as it recycles importin-a, its only known export cargo, back to the cytoplasm. As detailed below, some exportins contribute to RNA export. In addition, a few karyopherins (one in budding yeast and two in vertebrates) can translocate distinct sets of cargos in opposite directions.

1.2.4 The Ran GTPase: A key to transport directionality Following the observation that translocation through NPCs requires energy, the in vitro import assay enabled the identification of the small GTPase Ran (Ras-related nuclear protein/TC4). Along with the observation that non-hydrolyzable analogues of GTP inhibit the rate of in vitro nuclear import, this suggested that the metabolic energy supplied by the RanGTPase system could provide the driving force for directional transport (Melchior, Paschal, Evans, & Gerace, 1993). As other small GTPases, Ran (termed Gsp1/2 in budding yeast; Belhumeur et al., 1993) requires specific cofactors to switch between its GTP and its GDP-bound forms. The RanGEF (guanine nucleotide exchange factor, RCC1 in metazoans and its homologue Prp20 in yeast) is localized to the nucleus by virtue of its association with chromatin. Conversely, the GTPase-activating protein1 (RanGAP1, Rna1 in budding yeast) is localized in the cytoplasm, and even anchored on the cytoplasmic filaments of the NPC in metazoans. The asymmetrical localization of these two enzymes, along with the existence of a conserved transport receptor specific for Ran
GDP (Ntf2), leads to the nuclear accumulation of Ran
GTP. This Ran gradient is the key to establish the transport directionality (Fig. 1.4C). Indeed, all karyopherins preferentially interact with Ran
GTP (as first demonstrated by Rexach & Blobel, 1995), but this interaction has opposite consequences for importins versus exportins/cargos interactions. Indeed, Ran
GTP binding in the nucleus provokes the dissociation of the importin/ cargo complexes, thus leading to the nuclear release of the import substrates. The importin/Ran
GTP complexes then translocate back to the cytoplasm where RanGAP-mediated hydrolysis of GTP regenerates free importins, able to bind new substrates (Fig. 1.4A). Conversely, Ran
GTP binding enhances the affinity of exportins to their cargos inside the nucleus. Following translocation through the NPCs, hydrolysis of Ran
GTP in the cytoplasm leads to the dissociation of the Ran/exportin/cargo(s) complexes, allowing the recycling of exportins (Fig. 1.4B; for reviews, see Conti et al., 2006; Guttler & Gorlich, 2011). These unique properties

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FIGURE 1.4 The Molecular Mechanisms of Nucleocytoplasmic Transport (A, B) Schematic representation of the different components of the protein nuclear import and export machineries including karyopherins (importins and exportins) and their cargos (import or export substrates containing NLS or NES sequences). (C) Factors required for the establishment of the Ran
GTP/GDP nucleocytoplasmic gradient. (D) Schematic representation of the mRNA export process. Note that other cellular RNAs are exported out of the nucleus by a Ran/exportin-dependent mechanism as in (B) (see text for details). Modified from Alves, Palancade, & Doye (2005). The nuclear pore: A control station at the nucleus cytoplasm frontier. Biofutur, 254, 41–45 # (2005).

1.2 Nucleocytoplasmic Trafficking: The Rules of the Road

of Ran and Kaps thus contribute to the directionality, but also to the irreversibility of the transport process.

1.2.5 The case of INM targeting As INM proteins are synthesized in the endoplasmic reticulum (ER), which is continuous with the ONM (Fig. 1.1), it has initially been assumed that INM proteins could diffuse within the pore membrane and freely reach the INM, where they would be retained. But this diffusion–retention model was challenged by the observation that targeting of INM proteins in HeLa cells is energy and temperature dependent (Ohba, Schirmer, Nishimoto, & Gerace, 2004). Subsequently, an INM-sorting motif (a region containing positively charged amino acids adjacent to the hydrophobic transmembrane sequence) was identified. This motif is recognized by a membrane-associated truncated form of importin-a (importin-a16 in insect cells, Kpna-4–16 in vertebrates) that lacks the importin-b-binding domain and is required for the concentration of some INM proteins in the vicinity of the pore membrane (Braunagel, Williamson, Ding, Wu, & Summers, 2007 and references therein). In both diffusion–retention and sorting motif-dependent models, translocation is anticipated to take place through the 10-nm-wide channels located at the periphery of the NPCs, that could accommodate the diffusion or transport of INM proteins, assuming that they have a small extralumenal domain. This restriction is consistent with early observations revealing that there is a 60-kDa extraluminal domain size limitation for INM translocation (Soullam & Worman, 1995). More recently, however, NLS sequences that bind classical importin-a and mediate their translocation through the NPC in an importin-b- and Ran
GTP-dependant manner have been identified in several INM proteins (reviewed in Katta, Smoyer, & Jaspersen, 2013; Lusk, Blobel, & King, 2007). In this case, the peripheral NPC channels would not accommodate the size of the INM protein–importin-a/b complexes that instead would pass through the central channel of the NPC by using long unstructured linkers. These unfolded linkers would slice through the NPC scaffold to enable binding between the transport factors and the FG domains in the center of the NPC (Meinema et al., 2011). Finally, the observation that the lumenal domain of SUN2 supports NE localization highlights the existence of INM targeting pathways other than diffusion–retention and transport factor-mediated trafficking (Turgay et al., 2010). Genetic studies in yeast and RNAi experiments in mammals have further revealed the involvement of transmembrane core scaffold Nups in INM targeting. As these NPC alterations did not affect the kinetics of nuclear import of soluble cargos, this suggests that membrane bound and soluble cargos likely follow different pathways through the NPC (reviewed in Antonin, Ungricht, & Kutay, 2011). In the future, additional studies, of which some might take advantage of a recently developed in vitro assay based on Xenopus nuclei (detailed in Chapter 9), will be required to further clarify the various and redundant paths taken by INM proteins to reach their final destination.

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1.2.6 Distinct pathways contribute to RNA export With the exception of most mRNAs (detailed below), nuclear export of the various classes of RNAs also involve Ran- and exportin-dependent pathways (for a general review see Kohler & Hurt, 2007). Some RNAs directly interact with exportins, thanks to specific secondary structures. This is notably the case for tRNAs whose export, following their maturation and aminoacylation, involve Exportin-t (Xpo-t, Los1 in S. cerevisiae) and Exportin-5 (Xpo-5, Msn5p in S. cerevisiae) (as well as additional, factors, as detailed in Chapter 19). Likewise, highly structured small RNAs, including microRNA precursors (pre-miRNAs), adenovirus VA1 RNA, and human Y1 RNA, but also Dicer mRNA are exported by Exportin-5 (see Bennasser et al., 2011 and references therein). In addition, the Exportin1/Crm1-dependent export pathway indirectly contributes (e.g., via NES-containing adaptor proteins) to the nuclear export of several RNAs. For instance, the export of unspliced HIV mRNAs involves the binding of Rev, a NES-containing HIV-1-encoded protein, to the Rev-responsive element present within these viral RNAs (Fischer et al., 1995). Likewise, export of spliceosomal uridine-rich small nuclear RNAs (U snRNAs) involves the recruitment of a NEScontaining adaptor, PHAX (PHosphorylated Adaptor for RNA eXport) that binds to the U snRNA and its associated cap-binding complex (Ohno, Segref, Bachi, Wilm, & Mattaj, 2000 and references therein; see also Chapter 18). Finally, ribosomal RNAs (rRNA) are exported within two distinct export competent preribosomal particles: the pre-60S subunit that contains the mature 25S, 5.8S, and 5S rRNAs, and the pre-40s subunit that contains the 20S/18S rRNA (reviewed in Zemp & Kutay, 2007; see also Chapters 7 and 20). Export of both subunits was shown to be Crm1/ Xpo1 dependent, and a NES-dependent adaptor, Nmd3, was demonstrated to be essential for the export of pre-60S subunits. In addition, pre-60S subunit export requires additional export receptors, including the Mex67–Mtr2 heterodimer (a key mRNA export factor, see below) in yeast, Exportin-5 in vertebrates, as well as several auxiliary factors (references cited in Chapter 20; see also Wild et al., 2010 and references therein). While all the above-listed factors interact with the FG-rich domains of Nups, a recent study has revealed the contribution in 60S export of yeast Gle2 that interact with the non-FG domain of Nup116 (Occhipinti et al., 2013). Ribosomal subunits, likely because of their huge size, thus need to mobilize several export pathways for an efficient passage through the NPC. A common feature of these various RNA export processes is their tight link with RNA maturation, which provides a quality-control mechanism ensuring that only properly processed RNAs or assembled RNPs are exported. Likewise, mRNAs associate with multiple adaptor proteins and are exported as mRNPs, whose assembly is closely linked with many aspects of mRNA biogenesis, including transcription, processing, and quality control (for reviews, see Natalizio & Wente, 2013; Oeffinger & Zenklusen, 2012; Stewart, 2010; see also Chapter 18). However, unlike other RNA species, export of most mRNAs does not rely on exportinand Ran-dependent pathways (see, however, Natalizio & Wente, 2013 for

1.2 Nucleocytoplasmic Trafficking: The Rules of the Road

Crm1-dependent export of specific mRNPs). Instead, a conserved export dimer (termed Mex67/Mtr2 in yeast and TAP/p15 or NXF1/NXT1 in vertebrates) is in charge of mRNP export. NXF1 was initially demonstrated to directly interact with a structured RNA sequence, termed constitutive transport element (CTE), required for the export of specific viral mRNAs (Kang & Cullen, 1999 and references therein). However, NXF1/yMex67 does not directly bind to cellular mRNAs. Instead, multiple trans-acting adaptor proteins that directly bind the mRNA and couple the recruitment of the export receptor with mRNA transcription, processing, and genome organization, have been identified (Fig. 1.4D; reviewed in Natalizio & Wente, 2013; Nino, Herissant, Babour, & Dargemont, 2013; Oeffinger & Zenklusen, 2012). Like karyopherin-mediated transport processes, this mRNA export pathway also relies on the interaction between the export dimer and FG repeats. However, the main difference is that directionality of mRNA export does not depend on the Ran system. Instead, this process is driven by a DEAD box helicase (yDbp5 in budding yeast, Ddx18 in vertebrates). This protein is positioned at the cytoplasmic filaments, thanks to its binding to yNup159 (vertebrate Nup214). yDbp5, along with additional players localized on the cytoplasmic side of the NPCs, notably yNup159, the mRNA export factor yGle1, and its cofactor inositol hexakisphosphate (IP6), remodels the mRNP, leading to mRNA release and allowing the recycling of the export factors (for review, see Ledoux & Guthrie, 2011). Finally, it is noteworthy that the core scaffold Y-complex (yeast Nup84/ metazoan Nup107–160) is also implicated in mRNA export (Doye & Hurt, 1997 and references therein; Vasu et al., 2001). While these Nups could act by virtue of their function in maintaining NPC integrity, their reported physical interactions with components of the mRNA export machinery (Resendes, Rasala, & Forbes, 2008; Yao, Lutzmann, & Hurt, 2008) suggest that they may provide additional binding sites contributing to mRNP translocation.

1.2.7 Translocation across the NPCs: A dual function for FG-Nups as barrier and gate Although the translocation across NPCs is at the heart of the transport process, its precise mechanism is currently still a rather controversial issue. One thing for sure is the key role played by the unstructured and hydrophobic FG repeated sequences present in multiple Nups for both the establishment of the diffusion barrier and the active transport through the pore channel. Several models have been proposed to explain the molecular mechanisms underlying the contribution of the FG-Nups in the transport process. A common feature of all these models relies on the fact that all nuclear transport factors carry multiple low-affinity binding sites that can interact with the thousands of FG sequences that are provided by the 200 FG-Nups present in each NPC (for detailed reviews, see Aitchison & Rout, 2012; Terry & Wente, 2009). A first set of models might be grouped as “virtual gating” models. The “Brownian affinity gating” argues for a random (Brownian) movement of molecules within the

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pore channel. In this model, the FG domains set an entropic barrier that would be overcome by the affinity of transported cargo-carrier complexes for FG-Nups (Rout, Aitchison, Magnasco, & Chait, 2003). It was later proposed that FG-Nups are organized in a layer anchored and closely apposed to the walls of the transport channel. In this case, transporters are not free to move in 3-dimensions (as in the Brownian movement) but only in 2-dimensions all along this surface, hence the name of reduction of dimensionality, “ROD” model (Peters, 2005). In this model, passive diffusion would be permitted through a 8–10 nm FG-free central channel. Alternatively, the unfolded FG domain might act as a “polymer brush” that would sweep away macromolecules from their vicinity. In this context, it was proposed that transport factor binding would cause the participating FG domain(s) to collapse locally toward their anchoring sites in the NPC (as notably observed using atomic force microscopy), thereby reversibly releasing the entropic barrier (Lim et al., 2007). An alternative model is based on the observation that the FG domains can interact with each other by virtue of hydrophobic (via the phenylalanines residues) as well as hydrophilic interactions, leading to the formation of a sieve-like meshwork. This “hydrogel barrier” would prevent the passage of large molecules (the size of the FG-mesh fixing the upper limit for diffusion) whereas selective translocation of transporters would be achieved by a local dissolving of the FG–FG network. A recent study refined this “selective phase” model by demonstrating that hydrogels formed by distinct FG domains display distinct sieving effects, suggesting that NPCs could contain several gel layers of distinct mesh sizes and capacities for selective transport (Labokha et al., 2013 and references therein). The “virtual gating” and “selective phase” models are not mutually exclusive, and another hybrid model has been proposed, that is based on the existence of distinct types of FG domains: FG domains (mainly FXFG) with a dynamic extended coil conformation and FG domains (mainly GLFG types) with a globular collapsed conformation. In this so-called “forest” model, the bimodal distribution of these “trees” and “bushes” types of FG domains would create a central transporter structure and two distinct zones of traffic along the NPC conduit: a peripheral zone, close to the core scaffold, for small cargos, and a central zone, able to expand to facilitate translocation of large cargos (Adams & Wente, 2013; Yamada et al., 2010 and references therein). In all these models, the FG-network does not provide any directionality to the transport path. In line with this concept, high-resolution single-molecule studies have revealed that cargos explore the pore channel, until they reach the exit site (Grunwald & Singer, 2010; Yang & Musser, 2006; see discussion in Adams & Wente, 2013). In contrast, the asymmetric distribution of some FG-Nups along the axis of the NPC, along with the observation that specific transport receptors display a higher affinity for FG-Nups positioned at the end of their transport path through the NPC, had initially led to the hypothesis that an “affinity gradient” (Ben-Efraim & Gerace, 2001; Pyhtila & Rexach, 2003) may contribute to the polarized transport through the NPC. However, genetic studies in yeast, based on the combined deletion of various FG domains, have indicated that this asymmetric

1.3 The Nuclear Transport Machinery: A Dynamic and Versatile Device

distribution of FG-Nups is dispensable for basal in vivo transport (Strawn, Shen, Shulga, Goldfarb, & Wente, 2004). Currently, none of these models may fully account for the complex and multiple translocation mechanisms through NPCs. Thanks to the multiple in vitro and in vivo approaches that have already been developed, future studies integrating both the diversity of FG repeats and their positioning within the NPC framework will likely help to better understand the fundamental role played by the FG-Nups in the bidirectional transport process.

1.2.8 Noncanonical transport pathways through the NE In addition to these well-established protein or RNA transport mechanisms, there is an ever-expanding repertoire of alternative nuclear transport pathways that may be critical for particular cargos or under specific physiological or developmental conditions (see Wagstaff & Jans, 2009 and references therein). Transport molecules distinct from karyopherins include calmodulin (reviewed in Wagstaff & Jans, 2009), or Hikeshi, which mediates the nuclear import of Hsp70s in a FG-dependent but Ranindependent manner (detailed in Chapter 15). In addition, some proteins harboring karyopherin-like HEAT repeats, such as b-catenin, cytoskeletal proteins containing spectrin repeats, or nuclear shuttling/signaling molecules containing ARM repeats do not require soluble receptors and translocate independently of a carrier (reviewed in Wagstaff & Jans, 2009; see also Kumeta, Yamaguchi, Yoshimura, & Takeyasu, 2012 and references therein). For such proteins, it is anticipated that the driving force for their accumulation in the nucleus is their affinity for binding targets in the nucleoplasm, a mechanism that also contributes to the nuclear accumulation of small proteins that can otherwise diffuse through the NPCs. Finally, although most transport through the NE envelope takes place through the NPC, studies of herpes virus egress have uncovered an unexpected vesicle-mediated pathway (reviewed in Yarbrough et al., 2013). Subsequently, nuclear export of mRNP granules harboring specific synaptic transcripts in Drosophila larvae was shown to occur through a similar pathway (Speese et al., 2012). This suggests that herpes virus might have subverted a genuine cellular transport mechanism that could provide an alternative way to travel across the NE, as also recently hypothesized for INM proteins (Katta et al., 2013).

1.3 THE NUCLEAR TRANSPORT MACHINERY: A DYNAMIC AND VERSATILE DEVICE Beyond its role as selective gates between cytoplasm and nucleus, the nuclear transport machinery is also remodeled to adapt to specific stages of cell and organism life, and further plays a critical role in multiple cellular processes, including cell cycle regulation, gene expression, and maintenance of genetic integrity. Although any of these topics would deserve a full chapter, this part aims to highlight the main

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features of these multiple and complex mechanisms, while referring to recently published reviews in which these aspects have been exquisitely detailed.

1.3.1 NPC biogenesis throughout the cell cycle The complex NPC architecture is fundamental to maintain the proper functioning of pores, and consistently, the NPC scaffold is maintained with little to no turnover throughout the life span of post-mitotic differentiated cells. In fact, the deterioration of NPCs and the concomitant loss of the nuclear permeability barrier in aging cells highlight the requirement for cells to maintain the integrity of this assembly (reviewed in Toyama & Hetzer, 2013). In contrast, NPCs are dynamic structure in dividing cells that need to assemble new pores to prevent dilution of preexisting NPCs upon the successive rounds of cell division.

1.3.1.1 NPC disassembly The most impressive NPC rearrangements are observed in cells characterized by an open mitosis in which the NE and NPCs disassemble at mitotic onset to allow the formation of the mitotic spindle. Following nuclear envelope breakdown (NEBD), NPCs are largely dismantled, yet most Nups remain assembled into stable subcomplexes in the mitotic cytoplasm. By combining GFP-tagged Nups and specific transport reporters, the disassembly kinetics of Nups and transport competence of the NPCs could be simultaneously monitored by quantitative time-lapse fluorescence microscopy in single living cells (see Chapter 10 and references therein). In addition, in vitro assays that recapitulate NPC disassembly at mitotic onset have also been developed, using in vitro-reconstituted oocyte nuclei, or more recently, semi-permeabilized HeLa cells (see Chapter 12 and references therein). Together, these approaches have confirmed the implication of distinct classes of mitotic kinases, including Cdk1, that phosphorylate a broad range of Nups, most frequently on multiple sites (reviewed in FernandezMartinez & Rout, 2009). In particular, hyperphosphorylation of Nup98 was demonstrated to affect the NPC permeability barrier at an early stage of NEBD (see Chapter 12).

1.3.1.2 Post-mitotic NPC assembly Starting from anaphase onset, NE and Nups reassemble around the newly formed nuclei in the daughter cells, a process required to reestablish nuclear compartmentalization. How the simultaneous and rapid reassembly of thousands of NPCs, each composed of 500–1000 Nups, is spatially and temporally coordinated with the other cellular rearrangements that take place upon mitotic exit is a fascinating question. First hints into the sequential stages of post-mitotic NPC reassembly were obtained by combining distinct antibodies in immunofluorescence studies in mammalian cells (Bodoor et al., 1999). The advent of live cell imaging has provided a refinement of the kinetics of this highly dynamic process. As with disassembly, the reassembly kinetics of Nups and the acquisition of transport competence of the NPCs can be monitored by quantitative time-lapse fluorescence microscopy in

1.3 The Nuclear Transport Machinery: A Dynamic and Versatile Device

single living cells (see Chapter 10 and references therein). This assay revealed that Nups are successively recruited to reassemble new import competent NPCs within 10 min of anaphase onset. Live cell imaging of C. elegans embryos (see Chapter 13) or synticial Drosophila embryos (see Katsani, Karess, Dostatni, & Doye, 2008; Onischenko et al., 2005 and references therein) provides alternative models system that allow the simultaneous observation of multiple and successive disassembly and reassembly reactions. All these assays, when combined with molecular perturbations, such as RNAi depletion, provided insights into the sequential roles of various Nups in the NPC assembly process. However, in vivo depletion studies bear several limitations due to the extent of RNAi-induced depletion, loss of viability caused by altered transport capacities, and other secondary effects that may occur at other stages of the cell cycle. In addition, nuclei assembled in vitro using cell-free extracts of Xenopus eggs and exogenous DNA or chromatin have been used extensively to study NE and NPCs assembly (see Chapters 2, 8, and 9). This approach, combined with EM, made possible the visualization of potential NPC assembly intermediates (Goldberg, Wiese, Allen, & Wilson, 1997). Moreover, the Xenopus system provides the possibility to perform a “biochemical knockout” strategy to study the assembly of NPCs that occurs during in vitro nuclear formation. Following a highly efficient depletion of a protein of interest, it is possible to evaluate the recruitment of distinct Nups by immunofluorescence microscopy, to visualize NE integrity and NPC architecture using EM or SEM, and to assess transport properties of the assembled nuclei, thanks to specific reporters (Chapters 2, 8, and 9). In these assays (as also in the case of RNAi-mediated depletion), add-back experiments further enable the characterization of specific functional domains within a given protein, thereby refining the molecular mechanisms contributing to pore biogenesis. Together, these approaches have helped elucidate the order and interdependence of the main Nup recruitment steps (see Schooley, Vollmer, & Antonin, 2012 and references therein). Post-mitotic assembly is initiated by the recruitment of the Nup107–160 complex on chromatin, which is achieved thanks to its interaction with Elys (also called MEL-28 in C. elegans) that binds directly DNA with its C-terminal AT-Hook domain. This is followed by the recruitment of the transmembrane Nups Pom121 (thought to be mediated by its interaction with the Nup107–160 complex), and Ndc1. The components of the Nup93 complex, that also interact with these two transmembrane Nups, are then incorporated in a stepwise manner in the assembling NPCs. Nup93 subsequently recruits the Nup62 complex that, together with another FG-Nup, Nup98, contributes to the establishment of the NPC selectivity barrier. NPC assembly is then completed by the formation of the peripheral NPC structures (cytoplasmic filaments and nuclear basket). Besides their key role in bidirectional traffic in interphase, the Ran GTPase and karyopherins also play a critical role in post-mitotic NPC reassembly. Indeed, interaction of several Nups, including Elys, the Nup107–160 complex, Nup53, and FG-Nups, with importin-b and transportin was shown to inhibit NPC reassembly (Lau et al., 2009 and references therein). These negative regulatory events are

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counteracted by Ran
GTP, whose high concentration is maintained around chromosomes during mitosis (Kalab, Weis, & Heald, 2002; see also Chapter 15). Ran
GTP thus locally releases these Nups to allow their incorporation into the assembling NPCs. This spatial information must, however, be coordinated to the temporal regulation of the process, which is likely achieved by the reversal of mitosis-specific phosphorylation events (Walther et al., 2003; for review, see Schooley et al., 2012). While these studies have revealed that post-mitotic assembly is initiated by the recruitment of specific Nups on chromatin, how the ER membranes are reorganized to enclose the chromatin and form the NE, and how this process is coordinated with NPC reassembly remain debated (discussed in Schooley et al., 2012). Two distinct models have been proposed: (i) NPC assembly intermediates (seeds or “prepores”) may become subsequently enclosed by the outgrowing membranes of the reforming NE (enclosure model). Alternatively, (ii) the ER network on the chromatin surface may first form a closed NE into which NPC assembly subsequently proceeds (insertion model). Assembly of new NPCs by insertion into the intact nuclear membranes also occurs in all species during interphase, when the number of NPCs doubles in preparation for reentry into next mitosis, and is the unique mode of pore biogenesis in organisms, such as yeast, that employ closed mitosis for cell division (reviewed in Doucet & Hetzer, 2010; Jaspersen & Ghosh, 2012; Rothballer & Kutay, 2013). The post-mitotic NPC insertion model is thus appealing, as it would represent a unifying mechanism for NPC assembly in all stages of the cell cycle and across species. However, in contrast to the simultaneous post-mitotic assembly of thousands of NPCs in metazoan cells, interphase NPC assembly occurs as a collection of sporadic events. Although the NPCs assembled are identical, these two processes may thus accommodate distinct mechanisms.

1.3.1.3 De novo NPC assembly First insights into the molecular mechanisms involved in the biogenesis of new NPCs in the intact NE (a process also termed “de novo” biogenesis) have been provided by genetic studies in budding yeast (reviewed in Fernandez-Martinez & Rout, 2009). In particular, members of the Nup84 complex, Pom33, and reticulons (proteins involved in the maintenance of reticular ER membranes) are all required for proper NPC distribution, suggesting a role in NPC biogenesis or stability. A genetic screen based on the altered localization of GFP-tagged Nups (nuclear pore assembly mutants, npa, Ryan & Wente, 2002) revealed the contribution of additional factors in this process. In vertebrates, approaches to examine interphase NPC assembly have been developed using in vitro nuclear reconstitution assays based on Xenopus egg extracts (D’Angelo, Anderson, Richard, & Hetzer, 2006). Visualization of interphase NPC formation in mammalian cells has long been hampered by the interference of preexisting NPCs with the observation of nascent NPCs. While total fluorescence intensity of Nups at the NE in G1 versus G2 cells can provide clues into NPC numbers, high-resolution cell imaging now enables the visualization of individual NPCs and the detection of new assembly events at previously pore-free sites. In addition, a photobleaching

1.3 The Nuclear Transport Machinery: A Dynamic and Versatile Device

approach (FRAP) and an assay based on the cell-fusion technique have also been developed to tackle this problem (see Chapter 11 and references therein). Together, these approaches have provided the first clues on the de novo NPC assembly pathway (see reviews by Doucet & Hetzer, 2010; Jaspersen & Ghosh, 2012; Rothballer & Kutay, 2013 and references therein). A prerequisite for the assembly of new NPCs during interphase is the formation of aqueous pores in the NE, a step that requires remodeling of the nuclear membranes. Although this process is far from being understood, interactions between the luminal domains of integral membrane Nups, such as POM121, and other transmembrane proteins, such as SUN1 in vertebrates or Heh1/2 in yeast, may act in an early step to decrease the distance between the INM and ONM. In parallel, membrane-binding scaffold Nups, such as vertebrate Nup53, and membrane-shaping proteins like reticulons (Rtn1/Yop1 in budding yeast, Rtn4a in vertebrates) may assist pore formation by bending the nuclear membranes toward each other and facilitating their tight approximation. Other membrane remodeling events, such changes in the lipid composition of the NE, are also likely involved in the formation of the pore membrane (reviewed in Rothballer & Kutay, 2013). Recruitment of membrane coat-related scaffold Nups, notably the Nup107–160 complex (yeast Nup84 complex), as well as the previously mentioned ALPS domain-containing Nups, could then help to stabilize membrane curvature. As for post-mitotic assembly, this initial step is followed by the recruitment of other Nups required for the construction of the mature NPC and the establishment of its barrier and transport functions (reviewed in Rothballer & Kutay, 2013). Of note, as NPC assembly was shown to proceed from both sides of the NE, preexisting NPCs are likely required to import Nups, possibly preassembled as subcomplexes, into the nucleus. Interestingly, both in vivo studies in yeast (Ryan & Wente, 2002) and Xenopus-based cell-free assays (D’Angelo et al., 2006) have revealed the contribution of Ran and Kaps in de novo NPC biogenesis. However, the specific (possibly multiple) steps to which they contribute remain uncertain (discussed in Fernandez-Martinez & Rout, 2009).

1.3.2 Multiple functions of the nuclear transport machinery during the cell cycle Besides their roles in nucleocytoplasmic transport and NPC biogenesis, multiple factors of the nuclear transport machinery have also been implicated in diverse mitotic processes. In particular, the role of Ran and karyopherins in the spatial coordination of multiple cell cycle events has been extensively reviewed (Clarke & Zhang, 2008; see also Lau et al., 2009; Chapter 15 and references therein). Among the contributing mechanisms, the enrichment of Ran
GTP around the mitotic chromosomes provides spatial information to locally release spindle assembly factors from Kap-containing inhibitory complexes. In addition, the nuclear export factor Crm1 also plays a role in mitosis through its RCC1-dependent localization to kinetochores, where it was shown to recruit RanBP2/Nup358 and RanGAP1. The cytoplasmic nucleoporin RanBP2 is an E3

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SUMO ligase that forms a complex with two other proteins, RanGAP1, stably conjugated to SUMO (small ubiquitin-like modifier) and the E2 SUMO-conjugating enzyme Ubc9 (RRSU complex, RanBP2-RanGAP1:SUMO-Ubc9; see Fig. 1.3A). At kinetochores, this complex may locally regulate the Ran
GTP gradient and was also shown to sumoylate both topoisomerase II, a key player in sister chromatid separation, and the chromosome passenger complex (CPC), a mitotic regulator which functions in spindle and kinetochore assembly (reviewed in Bukata, Parker, & D’Angelo, 2013; Wozniak et al., 2010). This dual location at NPCs in interphase and kinetochores in mitosis is also an intriguing feature of the mammalian and C. elegans Nup107–160 complex and Elys/MEL-28 (a property, however, not shared by the Drosophila or yeast complexes). In mammalian cells, efficient depletion of this Y-shaped complex from kinetochores impairs mitotic progression. This mitotic phenotype was correlated with altered kinetochore recruitment of specific complexes including the abovementioned RRSU and CPC complexes, and the microtubule nucleation complex (g-tubulin ring complex, g-TuRC). In addition, the Nup107–160 complex is localized to spindle poles and proximal spindle fibers in prometaphase mammalian cells and was demonstrated to contribute to the in vitro assembly of bipolar spindles in Xenopus egg extracts (reviewed in Wozniak et al., 2010). Active mitotic functions have also been assigned for Rae1, a nucleoporin sharing homology with the mitotic checkpoint protein Bub3, that is localized at kinetochores, spindle, and spindle poles in human and Xenopus mitotic cells. Rae1, through interactions with several partners, seems to regulate spindle assembly and chromosome segregation through multiple mechanisms (reviewed in Bukata et al., 2013; Chatel & Fahrenkrog, 2011). Mitotic localizations and functions have now been reported for an increasing number of Nups including Nup98, Nup188, and Nup62 (reviewed in Bukata et al., 2013; Chatel & Fahrenkrog, 2011; see also Hashizume et al., 2013; Itoh et al., 2013). The association of Nups with distinct parts of the mitotic apparatus (mitotic chromosomes, spindle, centrosomes, and kinetochores) may contribute to the precise choreography of NE disassembly/reassembly with other mitotic events. Conversely, NPCs behave as an anchor for the localization of key cell cycle regulators. Indeed, besides their localization to kinetochores, the spindle assemble checkpoint (SAC) proteins Mad1 and Mad2 are also constitutively recruited to the nuclear pore basket by Tpr (yeast Mlp1/2) (see Fig. 1.3A). In yeast, the shuttling of Mad1 between unattached kinetochores and nuclear pores upon SAC activation was demonstrated to inhibit Kap121p-mediated protein import, thus indirectly changing spindle dynamics (Cairo, Ptak, & Wozniak, 2013). In mammalian cells, Tpr functions as a scaffold for regulating the stability of Mad1–Mad2 before their targeting to kinetochores, a mechanism likely underlying Tpr requirement for a robust SAC response (see Schweizer et al., 2013 and references therein). Because Tpr is also required to recruit the SUMO-isopeptidase SENP1 to NPCs (Fig. 1.3A), these authors further speculated that the function of Tpr in SAC might involve spatial

1.3 The Nuclear Transport Machinery: A Dynamic and Versatile Device

control of SUMO-dependent proteostasis at the NPCs. SENP proteins, and possibly other factors associated with the nuclear basket, may also account for the implication of Nup153 in the coordination between post-mitotic NPC basket assembly and abscission (discussed in Mackay & Ullman, 2011). In addition, two distinct NPC-anchored pathways were found to independently recruit the microtubule motor dynein to the NE in mammalian G2 cells, a process contributing to the proper localization of centrosomes close to the NE in mitotic prophase. These two G2-specific mechanisms, that depend upon associations with either the cytoplasmic filament Nup, RanBP2, or the Nup107–160 scaffold constituent, Nup133, independently recruit dynein through distinct interaction networks (Bolhy et al., 2011; Splinter et al., 2010). Of particular note, these two modes of G2-specific dynein recruitment to nuclear pores were recently demonstrated to contribute to another cell cycle-dependent mechanism, namely, the apical migration of nuclei from radial glial progenitor cells that takes place in G2 (Hu et al., 2013). These functions in mitosis and/or nuclear migration may contribute to the developmental defects reported in a functionally null allele of mouse Nup133 (Lupu, Alves, Anderson, Doye, & Lacy, 2008). These nonexhaustive examples thus highlight the multiple means of cross talk between NPC functions and cell cycle progression, and their possible contribution to developmental processes.

1.3.3 NPCs, nuclear organization, and gene expression Microscopists originally noticed that chromatin is not randomly distributed at the periphery of the nucleus and that NPCs are surrounded by heterochromatin-free areas possibly enriched in transcriptionally active genes (reviewed in Raices & D’Angelo, 2012). This observation raised the possibility that NPCs could act as genome organizers, leading in the 1980s to the so-called “gene gating” hypothesis which stipulated that gene expression could be regulated by physical association with nuclear pores (Blobel, 1985). Over the past 10 years, NPCs were confirmed to be important players in the definition of transcriptionally active regions using a panel of experimental approaches, in particular, in budding yeast (detailed in Chapter 21). On one hand, genome-wide chromatin immunoprecipitation studies demonstrated the association of several Nups and Kaps with actively transcribed genes and their importance in the definition of chromatin boundaries (Casolari et al., 2004; Ishii, Arib, Lin, Van Houwe, & Laemmli, 2002). On the other hand, GFP tagging of specific chromosomal loci with the LacO/LacI system indicated that a number of inducible genes relocate to NPCs upon transcriptional activation (reviewed in Van de Vosse, Wan, Wozniak, & Aitchison, 2011). NPC anchoring of activated genes requires transient interactions between multiple players at different stages of the gene expression process. These include Nups, mRNA export factors, chromatin-associated proteins, specific DNA sequences, and/or the mRNA itself (reviewed in Dieppois & Stutz, 2010). While

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relocalization of activated loci to NPCs has been shown to contribute to optimal mRNA production, their retention at NPCs following repression is likely to represent a form of epigenetic transcriptional memory and to prime them for faster reactivation (reviewed in Egecioglu & Brickner, 2011). Possible mechanisms of NPC-associated transcriptional memory include the maintenance of gene loops, as probed by chromosome conformation capture (3C), and/or the integration of specific epigenetic marks at the anchored locus (reviewed in Egecioglu & Brickner, 2011; Raices & D’Angelo, 2012). It has to be noted that consistent with their strategic location at the interface between the NPC and the nuclear interior, nuclear basket-associated proteins are especially important for these processes. While the aforementioned experimental data were mainly collected in yeast, accumulating evidence suggests that NPCs are also involved in the control of gene expression in metazoans (reviewed in Capelson, Doucet, & Hetzer, 2010; Light & Brickner, 2013; see also Chapters 13 and 21). A particular feature of the NPC-gene connection in higher eukaryotes is illustrated by the recruitment of a subset of Nups to actively transcribed genes in the nucleoplasm. Understanding in detail how the association of Nups with genes, either at NPCs or within the nucleus, regulates their expression, will be an important challenge for the future.

1.3.4 NPCs and genetic stability Beside their functions in chromosome segregation and gene expression, Nups have crucial roles in the maintenance of genetic integrity by directly influencing the metabolism of DNA damage. The first evidence for a connection between NPCs and the DNA damage response (DDR) came from genome-wide screens performed in yeast (reviewed in Bukata et al., 2013). These approaches revealed that Nups mutants are hypersensitive to clastogen or ionizing radiation, and identified a strong genetic interaction between the NPC and the homologous recombination pathway. In particular, the Y-complex as well as Nups of the nuclear basket appear to be critical for preventing the accumulation of unrepaired double-strand breaks (DSBs) (reviewed in Bukata et al., 2013). The combination of imaging approaches with genetics and ChIP studies in yeast further helped to dissect the molecular function of NPCs in DSB repair. On one hand, the NPC seems to act as a platform capable of recruiting several types of damaged DNA, including persistent DSBs, collapsed replication forks, and eroded telomeres, possibly channeling them into alternative repair pathways (reviewed in Nagai, Davoodi, & Gasser, 2011). On the other hand, the reported NPC association of enzymes of the SUMO pathway appears to be critical in preventing the accumulation of DNA damage (reviewed in Palancade & Doye, 2008). The SUMO-deconjugating enzyme Ulp1/SENP2 and the SUMO-dependent ubiquitin ligases Slx5/8, which are both anchored to NPCs likely through interactions with the Y-complex, are expected to target proteins of the DNA repair machinery for desumoylation and/or ubiquitination, thereby regulating their activity in the DDR. Accordingly, NPCs could position damaged chromosomes and their associated factors in the vicinity of these enzymes to finely tune the outcome of the DDR. While

Acknowledgments

dedicated studies as well as proteomic analyses have identified a number of DNA repair and replication factors whose sumoylation may account for this regulation (discussed in Nagai et al., 2011; Palancade & Doye, 2008), it is likely that similar mechanisms could regulate multiple other cellular processes. In this respect, it is noticeable that Ulp1 was recently shown to target for desumoylation two factors at the chromatin/NPC interface (Bretes et al., 2014; Texari et al., 2013). The connection between NPCs and the DDR, albeit studied largely in yeast, appears to be conserved during evolution, as revealed by the genetic instability caused by the loss-of-function of Y-complex components in several distant species (see Paulsen et al., 2009 and references therein). However, while the association of Ulp1/SENP2 with NPCs is conserved in mammals, its role in the DDR remains to be investigated. Of note, the mammalian-specific NPC localization of another enzyme of the SUMO pathway, namely, the SUMO ligase RanBP2 (see Werner, Flotho, & Melchior, 2012 and references therein), could also contribute to regulate SUMO modification of targets in the vicinity of nuclear pores.

CONCLUDING REMARKS Over the past 50 years, a combination of approaches in several model organisms has revealed the main rules of nuclear pore organization and nucleocytoplasmic transport. Despite its repetitive and modular structure, this fascinating assembly appears more complex and flexible than initially estimated with, for instance, its cell type or tissue specificities. The reported importance of nuclear pore and nucleocytoplasmic transport for several cellular processes, as well as its intimate connection with gene expression, probably explains why several Nups were found to be mutated in cancer (Chow, Factor, & Ullman, 2012; Kohler & Hurt, 2010) or other diseases (Jamali, Jamali, Mehrbod, & Mofrad, 2011). Future challenges in the field will include, among other aims, (i) a refined determination of NPC structure and biogenesis pathways, (ii) an improved understanding of the non-conventional functions of Nups and nucleocytoplasmic transport components, in and out of the pore, for instance, at gene loci, and (iii) a deeper comprehension of NPC composition and nucleocytoplasmic transport pathways and their (mis)regulations occurring during development, differentiation, and disease. For this purpose, some of the traditional methods used in the field, as well as innovative techniques and tools, are described in this volume. Importantly, certain of these approaches that have revolutionized our view of NPCs could be usefully applied to improve our understanding of other equally complex macromolecular assemblies.

Acknowledgments We thank Brian Burke (Institute of Medical Biology, Singapore) for critical reading of the manuscript, and colleagues for authorizing the reproduction of previously published images.

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We apologize to colleagues whose work could not be cited directly due to space constraints. Work in this laboratory is supported by CNRS, University Paris Diderot and “Who am I?” laboratory of excellence (ANR-11-LABX-0071/ANR-11-IDEX-0005-01), the French National Research Agency (ANR-12-BSV2-0008-01 to V. D.), Fondation ARC pour la Recherche sur le Cancer (to V. D. and B. P.), and Ligue Nationale contre le Cancer (to B. P.).

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Nino, C. A., Herissant, L., Babour, A., & Dargemont, C. (2013). mRNA nuclear export in yeast. Chemical Reviews, 113(11), 8523–8545. Nishi, K., Yoshida, M., Fujiwara, D., Nishikawa, M., Horinouchi, S., & Beppu, T. (1994). Leptomycin B targets a regulatory cascade of crm1, a fission yeast nuclear protein, involved in control of higher order chromosome structure and gene expression. Journal of Biological Chemistry, 269(9), 6320–6324. Occhipinti, L., Chang, Y., Altvater, M., Menet, A. M., Kemmler, S., & Panse, V. G. (2013). Non-FG mediated transport of the large pre-ribosomal subunit through the nuclear pore complex by the mRNA export factor Gle2. Nucleic Acids Research, 41(17), 8266–8279. Oeffinger, M., & Zenklusen, D. (2012). To the pore and through the pore: A story of mRNA export kinetics. Biochimica et Biophysica Acta, 1819(6), 494–506. Ohba, T., Schirmer, E. C., Nishimoto, T., & Gerace, L. (2004). Energy- and temperaturedependent transport of integral proteins to the inner nuclear membrane via the nuclear pore. Journal of Cell Biology, 167(6), 1051–1062. Ohno, M., Segref, A., Bachi, A., Wilm, M., & Mattaj, I. W. (2000). PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell, 101(2), 187–198. Onischenko, E. A., Gubanova, N. V., Kiseleva, E. V., & Hallberg, E. (2005). Cdk1 and okadaic acid-sensitive phosphatases control assembly of nuclear pore complexes in Drosophila embryos. Molecular Biology of the Cell, 16(11), 5152–5162. Onischenko, E., Stanton, L. H., Madrid, A. S., Kieselbach, T., & Weis, K. (2009). Role of the Ndc1 interaction network in yeast nuclear pore complex assembly and maintenance. Journal of Cell Biology, 185(3), 475–491. Ori, A., Banterle, N., Iskar, M., Andres-Pons, A., Escher, C., Khanh Bui, H., et al. (2013). Cell type-specific nuclear pores: A case in point for context-dependent stoichiometry of molecular machines. Molecular Systems Biology, 9, 648. Palancade, B., & Doye, V. (2008). Sumoylating and desumoylating enzymes at nuclear pores: Underpinning their unexpected duties? Trends in Cell Biology, 18(4), 174–183. Paulsen, R. D., Soni, D. V., Wollman, R., Hahn, A. T., Yee, M. C., Guan, A., et al. (2009). A genome-wide siRNA screen reveals diverse cellular processes and pathways that mediate genome stability. Molecular Cell, 35(2), 228–239. Peters, R. (2005). Translocation through the nuclear pore complex: Selectivity and speed by reduction-of-dimensionality. Traffic, 6(5), 421–427. Pyhtila, B., & Rexach, M. (2003). A gradient of affinity for the karyopherin Kap95p along the yeast nuclear pore complex. Journal of Biological Chemistry, 278(43), 42699–42709. Rabut, G., Doye, V., & Ellenberg, J. (2004). Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nature Cell Biology, 6(11), 1114–1121. Rahman, S., & Zenklusen, D. (2013). Single-molecule resolution fluorescent in situ hybridization (smFISH) in the yeast S. cerevisiae. Methods in Molecular Biology, 1042, 33–46. Raices, M., & D’Angelo, M. A. (2012). Nuclear pore complex composition: A new regulator of tissue-specific and developmental functions. Nature Reviews. Molecular Cell Biology, 13(11), 687–699. Resendes, K. K., Rasala, B. A., & Forbes, D. J. (2008). Centrin 2 localizes to the vertebrate nuclear pore and plays a role in mRNA and protein export. Molecular and Cellular Biology, 28(5), 1755–1769. Rexach, M., & Blobel, G. (1995). Protein import into nuclei: Association and dissociation reactions involving transport substrate, transport factors, and nucleoporins. Cell, 83(5), 683–692.

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CHAPTER

Imaging Metazoan Nuclear Pore Complexes by Field Emission Scanning Electron Microscopy

2

Boris Fichtman, Lihi Shaulov1, and Amnon Harel Faculty of Medicine in the Galilee, Bar-Ilan University, Safed, Israel

CHAPTER OUTLINE Introduction .............................................................................................................. 42 Prehistoric Landmarks (before EM): From Cells to the Nuclear Envelope ................... 42 The EM Era: From the Nuclear Envelope to Nuclear Pore Complexes ........................ 43 High Resolution SEM: Direct Surface Imaging of NPCs.............................................. 43 2.1 Rationale........................................................................................................... 44 2.1.1 Purpose ........................................................................................... 44 2.1.2 An Overview of Sample Preparation.................................................... 45 2.1.2.1 Fixation ......................................................................................45 2.1.2.2 Dehydration and Critical Point Drying ..........................................47 2.1.2.3 Sputter Coating...........................................................................47 2.2 Materials........................................................................................................... 48 2.2.1 Equipment....................................................................................... 48 2.2.2 Materials ......................................................................................... 48 2.2.3 Reagents ......................................................................................... 48 2.2.4 Buffers ............................................................................................ 48 2.3 Anchored Nuclei ................................................................................................ 49 2.3.1 Coating Silicon Chips........................................................................ 49 2.3.2 Chromatin Decondensation and Attachment........................................ 50 2.3.3 In Vitro Assembly Reaction................................................................ 50 2.4 Mammalian Cell Nuclei ...................................................................................... 51 2.5 Immunogold Labeling ......................................................................................... 52 2.6 Sample Preparation for FESEM............................................................................ 54

1

Current address: Department of Microbiology, Faculty of Health Sciences, Ben-Gurion University of the Negev, Beer Sheva, 84105, Israel.

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00002-3

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2.6.1 Fixation, Dehydration, and CPD ......................................................... 55 2.6.2 Sputter Coating and FESEM Analysis ................................................. 55 Acknowledgments ..................................................................................................... 56 References ............................................................................................................... 56

Abstract High resolution three-dimensional surface images of nuclear pore complexes (NPCs) can be obtained by field emission scanning electron microscopy. We present a short retrospective view starting from the early roots of microscopy, through the discovery of the cell nucleus and the development of some modern techniques for sample preparation and imaging. Detailed protocols are presented for assembling anchored nuclei in a Xenopus cell-free reconstitution system and for the exposure of the nuclear surface in mammalian cell nuclei. Immunogold labeling of metazoan NPCs and a promising new technique for delicate coating with iridium are also discussed.

INTRODUCTION Prehistoric landmarks (before EM): From cells to the nuclear envelope Our story begins in 1661 when King Charles II of England commissioned a series of microscopical studies from the Royal Society, an assignment which was entrusted to a young self-educated scientist, Robert Hooke. Considerable technical efforts led to a microscope with magnifications of up to 50
, providing insight into uncharted territories: “. . .by the help of Microscopes, there is nothing so small, as to escape our inquiry; hence there is a new visible World discovered to the understanding.” In a thin cutting of cork, Hooke observed empty spaces contained by walls, which reminded him of the living quarters of monks. He termed them cells and discovered one of the fundamental features of all life. Hooke was also able to appreciate the size of these cells: “in a Cubick Inch, above twelve hundred Millions. . . a thing almost incredible, did not our Microscope assure us of it” (Hooke, 1665). In the following decades, Antonie van Leeuwenhoek perfected optical lenses and microscopes, reaching magnifications of up to 275
and discovering microorganisms. Among his many discoveries, he described an internal “lumen,” the nucleus, in the red blood cells of salmon. Walther Flemming discovered chromatin in the late nineteenth century and presented detailed drawings of chromatin enclosed in a distinct compartment, surrounded by a clearly visible border (Flemming, 1878). In 1913, Kite used dyes and crystalloids to study the permeability of internal cellular structures and provided early experimental evidence for the existence of the nuclear envelope (Kite, 1913). However, the matter was far from resolved and even in the late 1930s some scientists still denied the existence of an organized membrane surrounding the nucleus.

Introduction

The EM era: From the nuclear envelope to nuclear pore complexes The first commercial transmission electron microscope (TEM) was constructed in 1939 and in the years that followed, fixation and staining techniques were developed for viewing biological samples, opening a new era in the study of the cell nucleus. Porter and colleagues used TEM imaging at magnifications up to 4100
to study cultured cells and chick embryos, fixed and stained with OsO4. The nucleus, demarcated by the nuclear envelope, was clearly seen in their micrographs, but the samples were too thick to distinguish additional details (Porter, Claude, & Fullam, 1945). This was soon improved with the use of high-speed microtomes and in the late 1940s much of the debate revolved around the lipid content of this nuclear border and the first clues indicating that it consists of two distinct layers (Baud, 1948; Callan, Randall, & Tomlin, 1949; Gessler & Fullam, 1946). Interestingly, Callan and Tomlin used angled metal shadowing on nuclear envelopes from amphibian oocytes to obtain topographical information on an array of pores of regular size, although they did not yet recognize that these structures were embedded in two concentric layers of membranes (Callan & Tomlin, 1950). By the end of that decade, Watson proposed the term “pore complex” for the large, approximately cylindrical formations seen to penetrate the two separate membranes of the nuclear envelope (Watson, 1959). These intricate molecular gateways or channels, now known to us as nuclear pore complexes (NPCs), became a new focus of attention for many subsequent TEM studies. The octagonal, rather than circular, nature of NPC structure was revealed in 1967 (Gall, 1967) and the first 3D reconstruction of its central framework was obtained in 1982 from negatively stained nuclear envelopes of Xenopus laevis oocytes (Unwin & Milligan, 1982). Many refinements and higher resolution models followed, including the use of cryoelectron tomography (reviewed in: Bui et al., 2013; Lim, Aebi, & Fahrenkrog, 2008; Maimon & Medalia, 2010). However, our overall knowledge of NPC structure and many of the advances brought about by TEM would not be complete without the additional input and surface views obtained by scanning electron microscopy (SEM).

High resolution SEM: Direct surface imaging of NPCs The NPC is an ideal target for structural studies by high resolution SEM (Schatten & Pawley, 2008). It is small enough to be beyond the capabilities of regular SEM, but large and complex enough to present a formidable challenge for averaging and 3D reconstruction techniques based on TEM imaging. Many of the architectural features of the NPC are still evident after coating with a 1- to 2-nm conductive metal layer and the distinctive, asymmetrically oriented filamentous extensions on both sides of the central scaffold are particularly suited for SEM imaging. Moreover, this large supramolecular structure is composed of many distinct nucleoporins, which can be probed by specific antibodies, using immunogold labeling techniques. Hans Ris was the first to successfully apply high resolution SEM to the nuclear envelope and discovered the basket structure on the nucleoplasmic side of the

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NPC (Ris, 1989). This was followed by a series of seminal microscopic studies by Martin Goldberg and Terry Allen, using SEM with a field emission electron source, in-lens detectors and specimens coated with chromium or tantalum (Goldberg & Allen, 1992, 1993, 1996). Most of these studies were performed on the giant nuclei (germinal vesicles) of amphibian oocytes, from which, the nuclear envelope can be manually isolated, providing direct access to both its cytoplasmic and nucleoplasmic sides. This approach yielded a wealth of structural information on NPCs, the nuclear envelope and adjacent structures. However, despite the clear advantages of oocytes, they represent an arrested, preembryonic state, and cannot be used to study dynamic processes like NPC assembly. Instead, an in vitro nuclear reconstitution system utilizing extracts made from unfertilized Xenopus laevis eggs has taken center stage in the study of NPC assembly, including analysis by SEM (Goldberg, Wiese, Allen, & Wilson, 1997; Harel et al., 2003; Wiese, Goldberg, Allen, & Wilson, 1997). Mammalian cell culture is an additional area of interest for studying metazoan NPCs, because SEM imaging can be combined with systems for genetic manipulation and even with the study of human disease. A major obstacle for this approach is the need to expose the surface of the nucleus from intact cells, which has mainly been achieved by physical fracturing techniques or the use of detergents (Allen et al., 2007; Drummond, Rutherford, Sanderson, & Allen, 2006; Walther et al., 2003).

2.1 RATIONALE In this chapter, we present detailed protocols for the visualization of metazoan nuclear envelopes and NPCs by high resolution field emission scanning electron microscopy (FESEM). For FESEM and other EM approaches in budding yeast, see Chapter 3. We describe the use of anchored chromatin templates (Section 2.3) in the Xenopus cell-free nuclear reconstitution system. This method is particularly suited for the visualization of fragile assembly intermediates, inhibited at early stages of nuclear envelope and NPC formation (Rotem et al., 2009; Shaulov, Gruber, Cohen, & Harel, 2011; Shaulov & Harel, 2012). A second method (Section 2.4) is our recently developed protocol for the exposure of mammalian cell nuclei without the use of detergents (Shaulov & Harel, 2012). In both cases, it is possible to proceed directly to the fixation of the sample and subsequent preparation steps for ultrastructural analysis by FESEM. Alternatively, we describe a detour into an immunogold labeling protocol (Section 2.5) leading to a combined view of surface topology and the localization of specific nucleoporins by antibodies. We also discuss a promising new coating technique with iridium, which provides a virtually grain-less coating layer and clearer images of NPC substructures.

2.1.1 Purpose Anchored chromatin templates consist of partially decondensed chromatin attached directly to the surface of poly-lysine (poly-L)-coated silicon chips. The use of these

2.1 Rationale

anchored templates in the Xenopus reconstitution system has several advantages. First, unlike the situation in intact cells or manually removed oocyte nuclei, there is no need for any treatment in order to remove the plasma membrane or other intracellular obstacles and gain direct access of the microscope’s electron beam to the nuclear surface. Second, nuclear assembly can be terminated at any time point (Fichtman, Ramos, Rasala, Harel, & Forbes, 2010) or by the use of chemical inhibitors (Macaulay & Forbes, 1996). This can also be extended to the well-known use of immunodepleted extracts and intervention by the addition of recombinant proteins (Finlay & Forbes, 1990; Harel et al., 2003; Shaulov et al., 2011). Third, when assembly is carried out directly on this solid surface, there is no need for centrifugation, which can damage fragile assembly intermediates like BAPTA (1,2-bis-[oaminophenoxy]-ethane-N,N,N0 ,N0 -tetra acetic acid)-inhibited nuclei (Shaulov & Harel, 2012). Fourth, the anchored assembly intermediates can easily be subjected to immunogold labeling since their attachment to the silicon chips simplifies repeated incubations and washes. Our new method for exposing the nuclear surface in mammalian tissue culture cells is based on repeated hypotonic treatments without the addition of any detergents. This helps to preserve the nuclear envelope and NPCs as close as possible to their native morphology (Shaulov, Fichtman, & Harel, 2014; Shaulov & Harel, 2012). Immunogold labeling protocols have been adapted for FESEM (Goldberg, 2008; Shaulov & Harel, 2012) and allow the localization of certain proteins to surface structures with relatively high accuracy ( 25 nm for gold-conjugated secondary antibodies and 15 nm for protein A-gold). This approach can be used for generating combined images depicting surface topology via an in-lens detector for secondary electrons, as well as gold particle positions determined by a separate EsB detector for backscattered electrons (Goldberg, 2008; Rotem et al., 2009).

2.1.2 An overview of sample preparation A flowchart outlining the main steps in each protocol and different options for the workflow is shown in Fig. 2.1. As mentioned above, both of the procedures for in vitro assembly and for mammalian cells can be combined with immunogold labeling. The bottom section of the chart is common to all protocols and includes several preparation steps, which are required to make biological specimens suitable for the high vacuum of the microscope chamber (Schatten & Pawley, 2008). These steps, with some theoretical background are outlined below.

2.1.2.1 Fixation Conventional (non-cryo) sample preparation for SEM relies on chemical fixation to preserve the fine structure in biological samples. This fixation is often divided into a primary and a secondary phase. The most commonly used agents for primary fixation are aldehydes that cross-link free amino groups in proteins and amino-lipids (Johnson, 1985). Glutaraldehyde quickly and irreversibly cross-links its substrates, but penetrates samples more slowly than paraformaldehyde (PFA), therefore they are

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FIGURE 2.1 An Overview of the Experimental Procedures This flowchart describes the main steps in the protocols for imaging in vitro assembled and mammalian cell nuclei. Both of these protocols can be extended to include immunogold labeling. The bottom part of the chart outlines the main steps in sample preparation for FESEM imaging.

2.1 Rationale

often combined in the primary fixation phase. The secondary fixative is osmium tetroxide (OsO4) which cross-links unsaturated phospholipids within biological membranes. This heavy metal oxide also increases the electron density of specimens, improving the ability to scatter the primary beam and produce more backscattered and secondary electron signals. However, OsO4 does not improve specimen conductivity since metal oxides are insulators.

2.1.2.2 Dehydration and critical point drying Since specimens placed in the SEM chamber are examined under vacuum, they must first be thoroughly dried. When aqueous biological samples are directly air-dried, the surface tension of the evaporating water causes distortion and damage to the specimens. The principle of critical point drying (CPD) was proposed in 1951 and is based on a specific combination of temperature and pressure (the critical point), at which the physical characteristics of the liquid and gaseous phases of a solvent become indistinguishable (Anderson, 1951). The critical point for water represents extremely harsh conditions (374  C, 229 bar) and it is therefore replaced with liquid CO2 (31  C, 74 bar), which is more compatible with fixed biological samples. Since water and CO2 are not miscible, the specimens are first dehydrated through a graded series of an intermediate solvent, ethanol. Ethanol is replaced with liquid CO2 in a sealed chamber of a CPD apparatus and the CO2 is brought to its critical point and converted to the gaseous phase without the damaging effects of surface tension.

2.1.2.3 Sputter coating Once the samples are dry, they can in principle, be examined by FESEM. However, dried biological specimens are exceptionally troublesome, as they are good insulators and have low material density even after OsO4 fixation. These properties make them predisposed to accumulate negative charge when irradiated by the probing electron beam and inefficient at scattering the electron beam to produce secondary and backscattered electrons, thus reducing the brightness and degrading the resolution of the image. To overcome these difficulties, biological specimens are usually coated with a thin conductive layer of metal. However, the coating may obscure the fine structural details of the specimen, so its thickness must be carefully optimized. The sputter deposition technique was invented over 150 years ago and was later adapted for biological samples and electron microscopy (Grove, 1852; Williams & Wyckoff, 1944). Some of the popular coating materials for biology are platinum, gold, gold/palladium alloy, and chromium. Chromium has been the metal of choice for high resolution imaging of NPCs because it is suitable for depositing very thin (down to 1 nm) films of fine granulation. The relatively low atomic mass of chromium (52, compared to 197 of gold) makes it compatible with immunogold labeling applications. Unfortunately, chromium is also a highly reactive metal and is susceptible to rapid oxidation upon exposure to air. Chromium oxide has a lower secondary electron emission coefficient and a lower density compared to chromium (Joy, 1991). In practical terms, this means that the actual thickness of chromium

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coating layers changes with the inevitable exposure of specimens to oxygen and this also affects their conductivity and the brightness of the signal. We have recently attempted to use iridium instead of chromium for the coating and imaging of NPCs by FESEM. Iridium is a stable metal and provides very thin, virtually grain-less coating layers, which are not prone to oxidation. Although we only have limited experience with this type of coating material, it seems to be a promising alternative for chromium (see Section 2.6.2).

2.2 MATERIALS 2.2.1 Equipment Critical point dryer (e.g., EM CPD300, Leica) attached to a high purity liquid CO2 cylinder. Sputter-coating apparatus (e.g., K575X, Emitech, or Q150R S, Quorum), preferably with exchangeable chromium and iridium targets. High resolution field emission scanning electron microscope (e.g., Merlin, Zeiss).

2.2.2 Materials Silicon chip support, 5
5 mm2 (#16008, Ted Pella). Aluminum SEM specimen mounting stubs (#75150, Electron Microscopy Sciences). Carbon adhesive tabs (#77825-12, Electron Microscopy Sciences).

2.2.3 Reagents Poly-L-lysine hydrobromide (P1524, Sigma). Dissolve at 0.2 mg/ml in sterile ultrapure water, filter through a 0.22-mm filter, and divide into 120 ml aliquots. Store at 20  C. Glutaraldehyde, 8% EM grade (#16019, Electron Microscopy Sciences). Store at 4  C. PFA, 16% EM grade (#15710, Electron Microscopy Sciences). Sodium cacodylate buffer, 0.2 M, pH 7.4 (#11652, Electron Microscopy Sciences). Store at 4  C. Osmium tetroxide, 2% (#19152, Electron Microscopy Sciences). PELCO conductive silver 187 (#16045, Ted Pella). ˚ beads, 8–12 mesh (#208582, Sigma) Dehydrating molecular sieves, 3 A (Table 2.1).

2.2.4 Buffers ELB (egg lysis buffer): 10 mM HEPES, pH 7.8, 50 mM KCl, 2.5 mM MgCl2. ELBS: ELB þ 250 mM sucrose. ELBS-K: ELBS þ 50 mM KCl.

2.3 Anchored Nuclei

Table 2.1 Antibodies used for immunogold labeling of NPCs by FESEM Antibodies

Source, comments

References

mAb414 (anti-FG Nups) anti-Nup62 (pc)a

Covance (#MMS-120R); dilute 1:300 from ascites fluid Douglass Forbes (La Jolla, USA)b

Unpublished

anti-ELYS (pc)a

Amnon Harel (Safed, Israel)b

anti-Nup107 (pc)a

Ulrike Kutay (Zurich, Switzerland)b

anti-Nup133 (pc)a 12 or 18 nm colloidal gold-goat anti-mouse IgG

Douglass Forbes (La Jolla, USA)b Jackson (#115-205-146); dilute 1:10–1:40, note variability between commercial batches of colloidal gold reagents Jackson (#111-205-144); dilute 1:10–1:40, as above

12 or 18 nm colloidal gold-goat anti-rabbit IgG

Shaulov and Harel (2012) Rotem et al. (2009) and Shaulov et al. (2011) Rotem et al. (2009) and Shaulov et al. (2011) Rotem et al. (2009) All of the above

All of the above

a

pc—affinity purified rabbit polyclonal antibodies generated by researchers in the field. Test individual batches of affinity-purified anti-Nup polyclonals in the 1–10 mg/ml range.

b

5
Buffer F: 400 mM PIPES KOH, pH 6.8, 5 mM MgCl2, 750 mM sucrose. PBS: 2.67 mM KCl, 1.47 mM KH2PO4, 138 mM NaCl, 8.1 mM Na2HPO4, pH 7.4. Hypotonic buffer: 15 mM Tris–HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2.

2.3 ANCHORED NUCLEI This section focuses on the assembly of nuclear envelopes on anchored chromatin templates using Xenopus egg extracts and a cell-free reconstitution system. For a detailed description of the assembly system, including the preparation of Xenopus egg cytosol and membranes and of demembranated sperm chromatin, see Chapters 8 and 9.

2.3.1 Coating silicon chips To increase the adhesion of biological material to the solid support and enable subsequent fixation to this surface, silicon chips are coated with poly-L. 1. Mark individual silicon chips by engraving numbers or letters in one corner with a diamond pen.

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2. Pipette 25 ml of 0.2 mg/ml poly-L and spread the solution over the entire surface of each chip. Incubate for 15 min at room temperature (RT). 3. Remove the poly-L solution and wash 3
in ultrapure water; air dry.

2.3.2 Chromatin decondensation and attachment We use demembranated sperm chromatin prepared as detailed in Chapter 8. The chromatin needs to be partially decondensed before it can be attached to poly-Lcoated chips. For this purpose, it is preincubated in a crude preparation of nucleoplasmin. Nucleoplasmin is a heat-stable protein which is able to displace protamines from sperm chromatin. It is easy to obtain an enriched preparation of nucleoplasmin, as it is the major protein remaining soluble after heat denaturation of total egg cytosol (Macaulay & Forbes, 1996). 1. Prepare crude nucleoplasmin by incubating Xenopus egg cytosol (prepared as detailed in Chapter 8) at 90  C, for 5 min, followed by a 10-min centrifugation at 20,000
g. Collect the supernatant and store in aliquots at 80  C. 2. Gently pipet 1 ml of chromatin stock (150,000 sperm units) into 10 ml of crude nucleoplasmin. Incubate 10 min at RT. Note: When pipetting chromatin always use wide gorge orifice tips or cut off the end of regular yellow tips. 3. Dilute the decondensed chromatin with 140 ml of ELB. Mix by very gentle pipetting. 4. Apply 20 ml of the suspension to the surface of each poly-L-coated chip. Allow the chromatin to settle onto the chip in a humidified chamber for 1 h at RT. Note: a humidified chamber is essential for preventing evaporation from the limited volume of solution covering the chips. Use pipet-box lids or petri dishes on parafilm and wet tissue paper for a humid environment. Avoid flooding. 5. Remove the solution by gentle pipetting (no vacuum aspiration) and replace with 20 ml ELB. Handle each chip separately and avoid drying. 6. Block the anchored chromatin templates with 20 ml of 2 mg/ml BSA in ELB. Incubate 15 min at RT in a humidified chamber.

2.3.3 In vitro assembly reaction While incubating the chromatin templates in blocking buffer prepare the assembly reaction mixture on ice. For each chip use: 25 ml of Xenopus egg cytosol, 1 ml energy mix (a 1:2:1 mixture of: 0.2 M ATP, 1 M phosphocreatine, 5 mg/ml phosphocreatine kinase), 0.3 ml Nocodazole from a 0.6-mg/ml stock solution, and 1 ml of diluted membranes. For each new preparation of fractionated egg extracts, the optimal ratio between the membrane and cytosolic fractions needs to be determined empirically. This is judged by assessing the amount of membranes needed for forming continuous, fully sealed nuclear envelopes. As a general rule, 20–25% of the optimal amount of membranes used for forming spherical nuclei in a standard assembly reaction is sufficient for the assembly of anchored nuclei (see Shaulov & Harel, 2012).

2.4 Mammalian Cell Nuclei

We recommend overlaying membrane aliquots with argon, before storage at 80  C, to prevent the oxidation of unsaturated lipids. 1. Wash each chip 3
with up to 50 ml of ELB, followed by one wash with ELBS. Use gentle pipetting and handle each chip separately to avoid detaching the chromatin templates. 2. Remove the buffer and apply 20 ml of the reaction mixture to the surface of each chip. Incubate 45 min at RT in a humidified chamber to assemble nuclei. Note: After assembly is completed, it is possible to move the chips into a 24-well cell culture plate and perform subsequent washes and incubations in wells. Take care, however, to avoid large bursts of liquid from pipetmans which might overturn the chips or detach the anchored nuclei. 3. Wash 3
with up to 50 ml of ELBS-K and proceed to the first fixation step in the immunogold labeling protocol (Section 2.5) or in sample preparation for FESEM (Section 2.6). The results of a typical anchored assembly experiment as visualized by FESEM at different magnifications are shown in Fig. 2.2.

2.4 MAMMALIAN CELL NUCLEI A brief, generalized protocol is described here for mammalian tissue culture cells. This protocol has been successfully applied to different types of adherent cells, including immortalized cell lines and primary cultures. The experimental design takes into account progressive material loss and duplicate samples. For additional technical details and notes on optimizing the protocol for individual cell types see Shaulov et al. (2014). 1. Prepare poly-L-coated silicon chips as in Section 2.3.1. Use four 5
5 mm2 chips for 1 million cells. 2. Grow cells under standard conditions, detach by trypsinization, and pellet by centrifugation for 3 min at 400
g, RT. Note: at this stage, detached cells can be frozen in medium supplemented with fetal calf serum and DMSO, stored under liquid nitrogen, and subsequently thawed for treatment. 3. Wash twice in PBS. 4. Resuspend cells in cold hypotonic buffer. Use 200 ml of buffer for the 250,000 cells destined for each chip. Incubate on ice for 5 min. 5. Pass the cells twice through a syringe with a 21-gauge needle, centrifuge for 3 min at 1000
g, 4  C in a fixed-angle rotor. 6. Optional: resuspend pellet in 200 ml of hypotonic buffer þ 10% glycerol and repeat steps 4–5. This repeated cycle might be needed for some cell lines that prove to be more resistant to the hypotonic treatment. 7. Resuspend each pellet in 1 ml of PBS þ 10% glycerol and centrifuge directly onto the poly-L-coated silicon chip for 10 min at 800
g, 4  C in a swing-bucket rotor.

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FIGURE 2.2 FESEM Imaging of Anchored Nuclei Assembled In Vitro (A–D) Increasing magnifications of a chromium-coated nuclear envelope assembled on a single chromatin template tethered to the support. Chromatin is completely enclosed by the nuclear envelope and NPCs exhibit the characteristic radial symmetry and are embedded in intact membranes. The arrow in D points to one NPC with a large particle symmetrically oriented within its central channel. Such particles are thought to be large cargoes caught in transit through the NPC.

Note: to improve the yield and spread the material over the entire surface of the chips, improvised low-speed centrifugation holders can be used as shown in Fig. 2.4B. 8. Carefully remove the chips into a 24-well cell culture plate filled with PBS and proceed to the first fixation step in the immunogold labeling protocol (Section 2.5) or in sample preparation for FESEM (Section 2.6). The results of a typical experiment as visualized by FESEM are shown in Fig. 2.3.

2.5 IMMUNOGOLD LABELING This protocol is designed to be carried out in a 24-well cell culture plate. One to three silicon chips can be handled in a single well to conserve antibody stocks. It is important to keep track of chip numbering and not to overturn the chips. The most

2.5 Immunogold Labeling

FIGURE 2.3 Direct Surface Imaging of Mammalian Cell Nuclei by FESEM (A) Cells from a primary culture of mouse lung fibroblasts were subjected to hypotonic treatment, spun down onto silicon chips, and coated with chromium. A field of cells is shown at low magnification. Only some of the smaller spherical bodies prove to be detached nuclei after examination at higher magnifications. Other bodies cannot be unequivocally identified, and in some cases, partial expanses of the nuclear surface are exposed from within the cell body. (B) An intact nucleus, completely detached from the cell body. (C) A higher magnification of a selected surface area of the nucleus shown in B. Mature NPCs are embedded in the undamaged membrane surface of the nuclear envelope. Smaller particles protruding from the surface are thought to be ribosomes.

important caution is to prevent the chips from drying, even for a few seconds. As shown in Fig. 2.4A, this can be accomplished by working simultaneously with two pipetmans: one for removing the liquid covering the chips in the well and the other for introducing a new solution. This practice should be continued all the way up to the CPD stage in Section 2.6.1. The approximate dilution range for each primary antibody can be determined in preliminary trials by indirect immunofluorescence on anchored nuclei or culture cells (Rotem et al., 2009). The background staining levels of the secondary gold-conjugated antibodies should be determined by omitting the primary antibodies in one of the experimental controls. Dilute primary and secondary antibodies in PBS þ 0.5% BSA. PBS can be replaced with

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FIGURE 2.4 Practical Tips for Handling Silicon Chips (A) The “in-out” double pipetman method prevents sample drying. Silicon chips are processed in 24-well plates. Pipet tips are pressed against the bottom and the edges of the well to prevent damage from direct currents. Two pipetmans are used simultaneously: one for removing the liquid covering the chips in the well and the other for introducing a new solution. (B) An improvised low-speed centrifugation holder for spinning mammalian cell nuclei onto a poly-L-coated silicon chip. Cut off the bottom of a 1.5-ml Eppendorf tube and fit the tube into a lid removed from another tube, to create a flat platform for the chip. Wrap parafilm around the tube to prevent leakage. Place inside a 14-ml Falcon tube stuffed with paper up to the 10.5-ml mark and spin in a swing-bucket rotor. See also Shaulov et al. (2014).

the ELB-based buffer system in all steps. However, we have obtained good results with both buffers, as long as phosphate ions were avoided in the last few steps preceding the secondary fixation with OsO4 in Section 2.6.1. 1. Carefully transfer the silicon chips, using high precision tweezers, into a 24-well cell culture plate filled with PBS. 2. Prefix with 3.7% PFA, 0.2% glutaraldehyde (GA) in 1
Buffer F for 5 min at RT. 3. Wash 2
5 min with PBS þ 0.2% glycine. 4. Block with PBS þ 0.5% BSA for 30 min. 5. Incubate with primary antibody for 45 min–1 h at RT. 6. Wash 3
with PBS þ 0.5% BSA. 7. Incubate with secondary gold-conjugated antibody for 30–45 min at RT. 8. Wash twice in PBS.

2.6 SAMPLE PREPARATION FOR FESEM This part of the procedure is common to all the options of the workflow shown in Fig. 2.1. The silicon chips carrying the samples are placed in a 24-well cell culture

2.6 Sample Preparation for FESEM

plate, if this has not already been done previously. It is essential to prevent the chips from drying in all the steps up to the CPD stage (see Section 2.5 and Fig. 2.4). Purchase the fixing agents PFA, GA, and OsO4 in single-use, sealed ampoules and prepare the working solutions for fixatives with 0.22 mm filtered buffers, immediately before use. Special attention should be paid to the purity of the 100% ethanol used for dehydration. Store this solution with dehydrating molecular sieves and filter through a 0.22-mm filter before use.

2.6.1 Fixation, dehydration, and CPD 1. Wash chips 3
in 20 mM HEPES pH 7.4. 2. Incubate in primary fixative containing 2% PFA and 2% GA in 1
Buffer F for 30 min at RT. 3. Wash 3
2 min in 0.1 M cacodylate buffer, pH 7.4. 4. Incubate in secondary fixative containing 1% OsO4 in 0.1 M cacodylate buffer for 10 min at RT. OsO4 must be protected from light. 5. Wash 3
2 min in ultrapure water. 6. Dehydrate through a graded ethanol series: 30%, 50%, 70%, 90%, 95%, and 100% ethanol; 2
2 min for each step. 7. Transfer the silicon chips to a CPD apparatus filled with ethanol as an intermediate fluid. Exchange into high purity liquid CO2 and subsequently from liquid to gas, according to the manufacturer’s instructions. If an automated apparatus (e.g., Leica EM CPD300) is available, perform 10 cycles to ensure the complete exchange of fluids. The dried samples should be stored under vacuum, in a desiccator, from this point on. For best results, coat the samples and perform FESEM imaging on the same day, minimizing their exposure to air. Alternatively, the procedure can be stopped at the end of the dehydration process and samples can be stored overnight in 100% ethanol.

2.6.2 Sputter coating and FESEM analysis Mount chips on aluminum stabs using carbon adhesive tabs or conductive silver paint and transfer them to a sputter-coating apparatus. The thickness of the coating layer can be adjusted for different applications. One- to two-nanometre chromium layers are most suitable for immunogold labeling and the detection of gold particles by an EsB detector. Some samples of mammalian cell nuclei are particularly susceptible to specimen charging and require a thicker (3–5 nm) coating layer. As mentioned above, we have also attempted to use a 2-nm thick iridium coat as an alternative for chromium coating on samples of in vitro assembled anchored nuclei. Figure 2.5 shows the encouraging results of this trial: iridium-coated NPCs reveal additional architectural details of the cytoplasmic filaments as well as views of the internal nuclear basket structure. This appears to be a significant improvement in the imaging of metazoan NPCs and needs to be tested on additional types of samples.

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FIGURE 2.5 Improved Ultrastructural Analysis of Iridium-coated NPCs (A) Anchored nuclei were assembled in vitro and processed for FESEM imaging as in Fig. 2.2, except for coating with a 2-nm thick layer of iridium. (B–D) Magnified views of the same image. Note the additional details of internal and external substructures observed with this delicate coating procedure, in comparison to the chromium-coated NPCs in Figs. 2.2 and 2.3.

Acknowledgments The authors thank Eugenia Klein and Michael Elbaum for advice on FESEM analysis, Ralph Neujahr and Xiong Liu (Carl Zeiss group, Germany) for their assistance in obtaining the images in Fig. 2.5 and Nitzan Shabtay for assistance with Fig. 2.4. This work was supported by a research grant from the Israel Science Foundation (1072/10) to A.H.

References Allen, T. D., Rutherford, S. A., Murray, S., Gardiner, F., Kiseleva, E., Goldberg, M. W., et al. (2007). Visualization of the nucleus and nuclear envelope in situ by SEM in tissue culture cells. Nature Protocols, 2(5), 1180–1184. Anderson, T. F. (1951). Techniques for the preservation of three-dimensional structure in preparing specimens for the electron microscope. Transactions of the New York Academy of Sciences, 13(4 Series II), 130–134. Baud, C. A. (1948). A cytochemical study of the perinuclear lipidic layer in the liver cell. Nature, 161(4093), 559. Bui, K. H., von Appen, A., Diguilio, A. L., Ori, A., Sparks, L., Mackmull, M. T., et al. (2013). Integrated structural analysis of the human nuclear pore complex scaffold. Cell, 155(6), 1233–1243.

References

Callan, H. G., Randall, J. T., & Tomlin, S. G. (1949). An electron microscope study of the nuclear membrane. Nature, 163(4138), 280. Callan, H., & Tomlin, S. (1950). Experimental studies on amphibian oocyte nuclei. I. Investigation of the structure of the nuclear membrane by means of the electron microscope. Proceedings of the Royal Society of London, Series B: Biological Sciences, 137(888), 367–378. Drummond, S. P., Rutherford, S. A., Sanderson, H. S., & Allen, T. D. (2006). High resolution analysis of mammalian nuclear structure throughout the cell cycle: Implications for nuclear pore complex assembly during interphase and mitosis. Canadian Journal of Physiology and Pharmacology, 84(3–4), 423–430. Fichtman, B., Ramos, C., Rasala, B., Harel, A., & Forbes, D. J. (2010). Inner/Outer nuclear membrane fusion in nuclear pore assembly: Biochemical demonstration and molecular analysis. Molecular Biology of the Cell, 21(23), 4197–4211. Finlay, D. R., & Forbes, D. J. (1990). Reconstitution of biochemically altered nuclear pores: Transport can be eliminated and restored. Cell, 60(1), 17–29. Flemming, W. (1878). Zur Kenntniss der Zelle und ihrer Theilungs-Erscheinungen. Schriften des Naturwissenschaftlichen Vereins fu¨r Schleswig-Holstein, 3, 23–27. Gall, J. G. (1967). Octagonal nuclear pores. The Journal of Cell Biology, 32(2), 391–399. Gessler, A. E., & Fullam, E. F. (1946). Sectioning for the electron microscope accomplished by the high speed microtome. The American Journal of Anatomy, 78, 245–283. Goldberg, M. W. (2008). Immunolabeling for scanning electron microscopy (SEM) and field emission SEM. Methods in Cell Biology, 88, 109–130. Goldberg, M. W., & Allen, T. D. (1992). High resolution scanning electron microscopy of the nuclear envelope: Demonstration of a new, regular, fibrous lattice attached to the baskets of the nucleoplasmic face of the nuclear pores. The Journal of Cell Biology, 119(6), 1429–1440. Goldberg, M. W., & Allen, T. D. (1993). The nuclear pore complex: Three-dimensional surface structure revealed by field emission, in-lens scanning electron microscopy, with underlying structure uncovered by proteolysis. Journal of Cell Science, 106(Pt 1), 261–274. Goldberg, M. W., & Allen, T. D. (1996). The nuclear pore complex and lamina: Threedimensional structures and interactions determined by field emission in-lens scanning electron microscopy. Journal of Molecular Biology, 257(4), 848–865. Goldberg, M. W., Wiese, C., Allen, T. D., & Wilson, K. L. (1997). Dimples, pores, star-rings, and thin rings on growing nuclear envelopes: Evidence for structural intermediates in nuclear pore complex assembly. Journal of Cell Science, 110(Pt 4), 409–420. Grove, W. R. (1852). On the electro-chemical polarity of gases. Philosophical Transactions of the Royal Society of London, 142, 87–101. Harel, A., Orjalo, A. V., Vincent, T., Lachish-Zalait, A., Vasu, S., Shah, S., et al. (2003). Removal of a single pore subcomplex results in vertebrate nuclei devoid of nuclear pores. Molecular Cell, 11(4), 853–864. Hooke, R. C. (1665). Micrographia: Or some physiological descriptions of miniature bodies made by magnifying glasses. London, England: Jo. Martyn and Ja. Allestry. Johnson, T. J. (1985). Aldehyde fixatives: Quantification of acid-producing reactions. Journal of Electron Microscopy Technique, 2(2), 129–138. Joy, D. C. (1991). Contrast in high-resolution scanning electron microscope images. Journal of Microscopy, 161(2), 343–355. Kite, G. (1913). The relative permeability of the surface and interior portions of the cytoplasm of animal and plant cells. Biological Bulletin, 25(1), 1–7 (A Preliminary Paper).

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Lim, R. Y., Aebi, U., & Fahrenkrog, B. (2008). Towards reconciling structure and function in the nuclear pore complex. Histochemistry and Cell Biology, 129(2), 105–116. Macaulay, C., & Forbes, D. J. (1996). Assembly of the nuclear pore: Biochemically distinct steps revealed with NEM, GTP gamma S, and BAPTA. The Journal of Cell Biology, 132(1–2), 5–20. Maimon, T., & Medalia, O. (2010). Perspective on the metazoan nuclear pore complex. Nucleus, 1(5), 383–386. Porter, K. R., Claude, A., & Fullam, E. F. (1945). A study of tissue culture cells by electron microscopy: Methods and preliminary observations. The Journal of Experimental Medicine, 81(3), 233–246. Ris, H. (1989). Three-dimensional imaging of cell ultrastructure with high resolution lowvoltage SEM. Institute of Physics Conference Series, 98, 657–662. Rotem, A., Gruber, R., Shorer, H., Shaulov, L., Klein, E., & Harel, A. (2009). Importin beta regulates the seeding of chromatin with initiation sites for nuclear pore assembly. Molecular Biology of the Cell, 20(18), 4031–4042. Schatten, H., & Pawley, J. B. (2008). Biological low voltage field emission scanning electron microscopy. New York, USA: Springer. Shaulov, L., Fichtman, B., & Harel, A. (2014). High resolution scanning electron microscopy for the imaging of nuclear pore complexes and Ran-mediated transport. Methods in Molecular Biology, 1120, 253–261. Shaulov, L., Gruber, R., Cohen, I., & Harel, A. (2011). A dominant-negative form of POM121 binds chromatin and disrupts the two separate modes of nuclear pore assembly. Journal of Cell Science, 124(Pt 22), 3822–3834. Shaulov, L., & Harel, A. (2012). Improved visualization of vertebrate nuclear pore complexes by field emission scanning electron microscopy. Structure, 20(3), 407–413. Unwin, P. N., & Milligan, R. A. (1982). A large particle associated with the perimeter of the nuclear pore complex. The Journal of Cell Biology, 93(1), 63–75. Walther, T. C., Alves, A., Pickersgill, H., Loiodice, I., Hetzer, M., Galy, V., et al. (2003). The conserved Nup107-160 complex is critical for nuclear pore complex assembly. Cell, 113(2), 195–206. Watson, M. L. (1959). Further observations on the nuclear envelope of the animal cell. The Journal of Biophysical and Biochemical Cytology, 6, 147–156. Wiese, C., Goldberg, M. W., Allen, T. D., & Wilson, K. L. (1997). Nuclear envelope assembly in Xenopus extracts visualized by scanning EM reveals a transport-dependent ‘envelope smoothing’ event. Journal of Cell Science, 110(Pt 13), 1489–1502. Williams, R. C., & Wyckoff, R. W. (1944). The thickness of electron microscopic objects. Journal of Applied Physics, 15(10), 712–716.

CHAPTER

Imaging Yeast NPCs: From Classical Electron Microscopy to Immuno-SEM

3

Elena Kiseleva*, A. Christine Richardson{, Jindriska Fiserova{, Anton A. Strunov*, Matthew C. Spink{, Simeon R. Johnson{, and Martin W. Goldberg{ *

Laboratory of Morphology and Function of Cell Structure, Institute of Cytology and Genetics, Russian Academy of Science, Novosibirsk, Russia { Department of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom

CHAPTER OUTLINE Introduction .............................................................................................................. 60 3.1 Conventional TEM .............................................................................................. 61 3.1.1 Materials ......................................................................................... 62 3.1.1.1 Equipment..................................................................................63 3.1.1.2 Spheroplast Preparation..............................................................63 3.1.1.3 Fixation, Dehydration, Contrasting, and Embedding .....................63 3.1.1.4 Sectioning and Poststaining ........................................................63 3.1.2 Procedure ........................................................................................ 64 3.1.2.1 Spheroplast Preparation..............................................................64 3.1.2.2 Fixation ......................................................................................64 3.1.2.3 Dehydration, Contrasting, and Embedding...................................65 3.1.2.4 Sectioning and Staining of Semithin and Ultrathin Sections..........65 3.2 SEM and Immuno-SEM of Yeast Nuclei................................................................ 65 3.2.1 Materials ......................................................................................... 66 3.2.1.1 Equipment..................................................................................66 3.2.1.2 Isolation of Nuclei from Spheroplasts ..........................................66 3.2.1.3 Immunolabeling..........................................................................66 3.2.1.4 Fixation ......................................................................................67 3.2.2 Procedure ........................................................................................ 67 3.2.2.1 Nuclear Isolation.........................................................................67 3.2.2.2 Ribosome Removal.....................................................................68 3.2.2.3 Prefixation and Attachment to Chip .............................................68 3.2.2.4 Immunolabeling..........................................................................68 3.2.2.5 Preparation of Samples for FESEM ..............................................69

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00003-5

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3.3 Immunogold Labeling of Yeast Ultrathin Cryosections........................................... 69 3.3.1 Materials ......................................................................................... 71 3.3.1.1 Fixation, Embedding in Gelatin, and Sucrose Infiltration...............71 3.3.1.2 Cryosectioning ............................................................................72 3.3.1.3 Immunogold Labeling .................................................................73 3.3.2 Procedure ........................................................................................ 73 3.3.2.1 Yeast Culture..............................................................................73 3.3.2.2 Fixation, Embedding in Gelatin, and Sucrose Infiltration...............73 3.3.2.3 Cryosectioning ............................................................................74 3.3.2.4 Immunogold Labeling .................................................................75 Conclusions and Perspectives ................................................................................... 76 Acknowledgments ..................................................................................................... 77 References ............................................................................................................... 77

Abstract Electron microscopy (EM) has been used extensively for the study of nuclear transport as well as the structure of the nuclear pore complex (NPC) and nuclear envelope. However, there are specific challenges faced when carrying out EM in one of the main model organisms used: the yeast, Saccharomyces cerevisiae. These are due to the presence of a cell wall, vacuoles, and a densely packed cytoplasm which, for transmission EM (TEM), make fixation, embedding, and imaging difficult. These also present problems for scanning EM (SEM) because cell wall removal and isolation of nuclei can easily damage the relatively fragile NPCs. We present some of the protocols we use to prepare samples for TEM and SEM to provide information about yeast NPC ultrastructure and the location of nucleoporins and transport factors by immunogold labeling within that ultrastructure.

INTRODUCTION Electron microscopy (EM) has many applications in the study of nuclear transport and has been used extensively. Early work in amphibians showed details of nuclear pore complex (NPC) structure (Gall, 1967), later built upon by a variety of EM techniques including scanning EM (SEM) (Goldberg & Allen, 1992; Ris, 1989; see also Chapter 2), cryo-EM (Akey & Radermacher, 1993), and electron tomography (Beck et al., 2004). The transport process was visualized using transmission EM (TEM) of insect salivary glands exporting large mRNP particles (Stevens & Swift, 1966) and gold particle tagged import cargoes (Feldher, Kallenbach, & Schultz, 1984). Despite the importance of yeast, particularly, Saccharomyces cerevisiae, as a model organism in the study of nuclear transport, the use of EM has been limited. S. cerevisiae has a cell wall, large vacuoles, and a dense cytoplasm, all of which conspire to make EM challenging, as fixation is impaired, structures are difficult to preserve, and details difficult

3.1 Conventional TEM

to perceive. For these reason, when studying yeast nuclear transport by EM, it is often necessary to use a range of techniques in order to extract different types of information. Techniques detailed in this chapter include (1) conventional TEM methods which can be carried out in any well-setup EM lab, but which require some harsh procedures that must be considered when interpreting the images; (2) methods for imaging yeast NPC structure by field emission SEM (feSEM) (Kiseleva et al., 2007; Murray & Kiseleva, 2008), to give detailed information on the structure, which can also be labeled with colloidal gold-tagged antibodies (Goldberg & Fiserova, 2010), but which is limited to surface details; and (3) immunogold labeling for TEM which is best achieved using thawed cryosections (Tokuyasu, 1973), because in the absence of any embedding material there is excellent access for antibodies to epitopes within the section, giving the most sensitive labeling. Yeasts present some particular problems with this method concerning fixation, epitope accessibility, and membrane visibility, but a protocol was developed by Griffith, Mari, Maziere, and Reggiori (2008) to alleviate these problems, for which we present the details as used in our laboratory. The ultimate preservation of yeast ultrastructure is achieved by high pressure freezing (HPF) and low temperature fixation and low temperature embedding/freeze substitution (FS), followed by room temperature sectioning (McDonald, 2007). The drawback of HPF/FS is that it is time consuming, not always consistent, and therefore requiring much optimization, as well as expensive, specialized equipment. Moreover, although it can sometimes be extended to immunogold labeling using certain suitable antibodies, this is not always possible and the use of the Tokuyasu method is often necessary. HPF/FS for NPC and nuclear transport studies will not be covered here (for details and examples see Fiserova & Goldberg, 2010; Fiserova, Richards, Wente, & Goldberg, 2010; Fiserova, Spink, Richards, Saunter, & Goldberg, 2014).

3.1 CONVENTIONAL TEM Conventional TEM (room temperature fixation, embedding, and sectioning) has been used, for instance, to study changes in cell/nuclear morphology and NPC distribution in mutant yeast strains (e.g., Doye, Wepf, & Hurt, 1994; Titus, Dawson, Rexer, Ryan, & Wente, 2010; Wente & Blobel, 1993). In order to obtain good morphology, it is necessary to remove the cell wall after aldehyde fixation, in the presence of sorbitol to protect the cells from osmotic shock, followed by membrane fixation with osmium tetroxide, and enhancement of membrane contrast with K3[Fe(CN)6]. Cell pellets are then embedded in agarose to prevent dispersion during subsequent processing and embedded in standard epoxy resins, then sectioned, stained, and viewed by standard methods. The resulting images (Fig. 3.1B–D) show good contrast of membranes as well as protein structures, although many details are difficult to observe due to the high cytoplasmic density. Some shrinkage/distortion of membranes, particularly, of the nuclear envelope, is observed when compared to HPF/FS samples (Fig. 3.1E), which should be taken into account when interpreting images and considering comparative controls. However, some of the details at high

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FIGURE 3.1 Visualization of budding yeast cells and spheroplasts by phase contrast microscopy. (A) Yeast cells at stationary phase (left image), at logarithmic phase (cells contain small buds) (middle image), and spheroplasts obtained by lyticase treatment of cells (right image): round shape spheroplast shows no signs of damage. (B) Irregular shaped nucleus with smooth nuclear envelope which is a sign of good structural preservation. (C0 and D0 ) High magnification of nuclear envelope from “B” (C and D insets, respectively). Outer and inner nuclear membranes are distinguished and NPCs (arrows) are visible. For comparison, TEM thin sections of HPF/FS yeast are shown at low (E) and high (E0 ) magnification.

magnification, of NPCs for instance, are comparable, although with different contrast to HPF/FS (Fig. 3.1C0 , D0 compared to E0 ).

3.1.1 Materials Many of the chemicals used here are highly toxic and gloves should be used at all times. All fixation and embedding steps should additionally be carried out in a fume hood.

3.1 Conventional TEM

3.1.1.1 Equipment 1. 2. 3. 4. 5. 6.

Centrifuge with swing out rotor. Horizontal shaker. 60  C oven. Ultramicrotome (Leica Microsystems GmbH, Wetzlar, Germany). Phase contrast light microscope. Transmission electron microscope.

3.1.1.2 Spheroplast preparation 1. Liquid YPD medium: 1% (w/v) yeast extract (DIFCO Laboratories, Detroit, MI), 2% Bacto peptone (w/v) (DIFCO Laboratories), 2% dextrose (w/v) (Fisher Scientific, Loughborough, UK). 2. Fixative 1: 4% Paraformaldehyde (PFA) (Panreac, Barcelona, Spain) in 0.1 M sodium cacodylate (Fluka, St. Gallen, Switzerland). To dissolve PFA, warm buffer on a magnetic stirrer with heating then add drops of 20% sodium hydroxide until solution clears. Check pH is 7.4 and adjust using drops of concentrated HCl. 3. Buffer 1: 0.1 M Tris–HCl pH 7.4 (Promega, Madison, WI), 10 mM Dithiothrietol (Sigma–Aldrich, Saint Louis, MO). 4. Buffer 2: 20 mM K3PO pH 7.4, 0.5 mM MgCl2 with (a) 1.2 M sorbitol (Sigma– Aldrich), (b) 1.0 M sorbitol, or (c) 0.5 M sorbitol. 5. Buffer 3a: 10,000 U Lyticase (Sigma–Aldrich) in 800 ml of Buffer 2.

3.1.1.3 Fixation, dehydration, contrasting, and embedding 1. Fixative 2: 1% OsO4 (Sigma–Aldrich), 1% K3[Fe(CN)6] (potassium ferricyanide, Sigma–Aldrich improves membrane staining) in 0.1 M sodium citrate pH 7.4 2. Low Melting temperature Agarose (Sigma–Aldrich). 3. Ethanol (30%, 50%, 70%, 95%). 4. Stain 1: 2% uranyl acetate (Serva, Heidelberg, Germany) in 70% ethanol. 5. 100% ethanol. 6. 100% acetone. 7. Agar 100 epoxy resin kit (Epon-812, MNA, DDSA, DMP-30) (Agar Scientific, Essex, UK). 8. Plastic sample embedding capsules (Agar Scientific).

3.1.1.4 Sectioning and poststaining 1. Formvar-coated 200-mm mesh copper and nickel grids (Agar Scientific). 2. 1% Toluidine Blue Stain. Dissolve 0.8 g of sodium tetraborate in 100 ml of distilled water, add 0.8 g of toluidine blue, and 0.2 g of pyronin Y stir well and filter before use. 3. Filter paper (Filtrak Brandt GmbH, Wiesenbad, Germany). 4. Stain 2: 1% uranyl acetate in ethanol filtered through 0.45 mm filter (PharmAssure, Hayward, CA). 5. Stain 3: Reynold’s lead citrate: 0.28 g of lead nitrate (Fluka), 0.325 g of sodium citrate, 6 ml of boiled distilled water, and 1 M sodium hydroxide (Panreac).

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To prepare, dissolve lead nitrate in water, add sodium citrate, and mix for 30 min. Add sodium hydroxide drop by drop until solution turns clear. Make up to 10 ml. Filter and store in a fridge, centrifuge the necessary volume of lead citrate at 15000  g in a sealed tube for 5min.

3.1.2 Procedure 3.1.2.1 Spheroplast preparation

1. Grow yeast for 1–2 days at 30  C in 50 ml YPD medium in 100 ml conical flask with shaking (140 rpm). 2. Add 5 ml of culture to 50 ml fresh medium and incubate for 3–4 h with shaking (170 rpm) at 30  C to mid-log phase (when most cells have buds). Check after 3–4 h by phase-contrast light microscopy and proceed to the next step when approximately 70% of cells have buds (Fig. 3.1A). It is important that cells are in mid-log phase. Different strains will achieve this at different times. 3. Add Fixative 1 at a 1:1 (v/v) ratio to culture and shake for 30 min. 4. Transfer to 50 ml tubes and centrifuge at 3000  g in a swing out rotor for 5 min at room temperature. 5. Resuspend the pellet in 20 ml dH2O for 1–2 min. 6. Pellet cells by centrifugation (3000  g, 3 min). 7. Resuspend pellet in Buffer 1 and shake for 25 min at 30  C with shaking (140 rpm). 8. Pellet (3000  g, 3 min). 9. Resuspend pellet in 50 ml Buffer 2a. 10. Pellet (3000  g, 3 min). 11. Resuspend in 20 ml Buffer 3a and incubate for 20–40 min, 30  C, shaking. 12. Every 10 min, take 1 ml of sample onto a glass slide, place on a cover slip, and observe with a phase contrast microscope with 60  or 100  objective. Proceed when  70% of the cells are spheroplasts. Spheroplasts are round and less bright than whole cells (Fig. 3.1A). Spheroplasts are fragile and must be handled carefully. 13. Pellet spheroplasts (3000  g, 3 min). 14. Resuspend in 10 ml Buffer 2a.

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

Pellet (3000  g, 3 min). Resuspend in 10 ml Buffer 2b (with 1.0 M sorbitol) and pellet (3000  g, 3 min). Resuspend in 10 ml Buffer 2c (0.5 M sorbitol). Pellet (3000  g, 3 min). Resuspend in 10 ml Fixative 2 and incubate for 1 h with shaking (140 rpm). Pellet (3000  g, 3 min). Resuspend in 10 ml 0.1 M sodium citrate, pellet (3000  g, 3 min), and repeat 2 . Check spheroplasts at each step using phase contrast microscope

3.2 SEM and Immuno-SEM of Yeast Nuclei

3.1.2.3 Dehydration, contrasting, and embedding 1. 2. 3. 4. 5. 6. 7. 8. 9.

Place a small quantity of wet pellet in a drop of molten agarose. Incubate for 10 min at 30  C. Place on ice until agarose sets. Transfer agarose to a Petri dish containing water and cut into small pieces (1 mm3). Dehydrate in an ethanol series (30%, 50%, 70% for 10 min each) on a specimen rotator. Place in Stain 1 for 1 h, shaking, in the dark. Dehydrate 2  10 min in 100% ethanol then 2  10 min in 100% acetone. Incubate in a 3:1 mixture of acetone:Agar 100 resin, 2 h, room temperature, then 1:1 acetone:Agar 100 for 2 h and then 1:3 acetone:Agar 100 overnight. Transfer to fresh Agar 100, 2 h, then embed in freshly prepared Agar 100 in plastic capsules. Polymerize in 60  C oven 2–3 days.

3.1.2.4 Sectioning and staining of semithin and ultrathin sections 1. Trim block and prepare semithin sections (0.5–1.0 mm). 2. Stain sections with 1% Toluidine Blue Stain and choose region containing many spheroplasts. 3. Cut ultrathin sections (60 nm thick) with ultramicrotome. 4. Place copper grids in acetone for 5 min and dry on filter paper. 5. Pickup sections on grids. 6. Float grid on 30–40 ml droplet Stain 2 for 5 min. 7. Rinse with distilled water for 5 min. 8. Float grid on 30–40 ml droplet of Stain 3 for 5 min. Put NaOH tablets around the drop. 9. Wash with distilled water 5 min. 10. Withdraw water from grid using filter paper and allow to dry. 11. Observe with TEM at 100 kV accelerating voltage.

3.2 SEM AND IMMUNO-SEM OF YEAST NUCLEI High resolution SEMs using field emission electron guns (feSEM) have been used in the nuclear transport field to study nuclear envelope structure of amphibian oocytes leading to the discovery of the NPC basket and details of its structure (Goldberg & Allen, 1992; Ris, 1989) as well as details of surrounding structures such as the nuclear lamina (Goldberg, Huttenlauch, Hutchison, & Stick, 2008). Using immunolabeling protocols similar to those designed for immunofluorescence, but with colloidal gold-tagged secondary antibodies, the location of individual proteins or other epitopes can be determined, assuming the labeling is at the surface being imaged (e.g., Kiseleva, Morozova, Voeltz, Allen, & Goldberg, 2007). Imaging yeast nuclear envelope and NPCs is particularly challenging because the surface of interest (the nuclear envelope), which has to be exposed, is deep within the cell, surrounded

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by a dense cytoplasm and enclosed in a strong cell wall. To make matters worse yeast NPC structure appears to be particularly fragile. The protocol below is based on previously published protocols (Kiseleva et al., 2004; Kiseleva, Allen, et al., 2007; Murray & Kiseleva, 2008), which have been modified to more consistently maintain NPC integrity. We have found it advantageous to include protease inhibitors in the spheroplasting buffer, because unlike the TEM preparation, fixation (which halts any proteolysis) is done after removal of the cell wall and isolation of the nuclei. We also get more consistent results and more extensive NPC structures if we leave ribosomes in place and do not include a salt wash step (Fiserova et al., 2014). However, in the presence of ribosomes NPCs can be difficult to recognize.

3.2.1 Materials 3.2.1.1 Equipment 1. 1 ml screw cap glass vials (Biotrace International) 2. High vacuum coating unit with chromium target using high purity Argon (>99.999% Air Liquide) (e.g., Cressington 328HR) 3. Critical Point dryer (e.g., Leica Microsystems) 4. Dumont tweezers 3 Dumostar (Agar Scientific) 5. High Resolution Scanning Electron Microscope with Back Scattered Electron detector (e.g., Hitachi S5200) 6. Phase-contrast microscope with 60  or 100  lens 7. Silicon chips 5 mm  5 mm (Agar Scientific)

3.2.1.2 Isolation of nuclei from spheroplasts (for spheroplast preparation see Section 3.1) 1. 1 mg/ml Poly-L-lysine in dH2O (Sigma–Aldrich) 2. Buffer 3b: 10,000 U Lyticase (Sigma–Aldrich) in 800 ml of Buffer 2 þ 0.1% Pepstatin A (Sigma–Aldrich), 0.1% Phenylmethanesulfonyl fluoride (Sigma– Aldrich) (Fiserova et al., 2014) 3. Buffer 4: 10 mM potassium phosphate, 0.5 mM MgCl2, pH 6.5. 4. Buffer 5: 0.5 mM MgCl2, pH 6.5. 5. Buffer 6: 20 mM Tris–HCl, 0.5 mM MgCl2, 0.2 M Sucrose, pH 6.5. 6. Buffer 7: 20 mM Tris–HCl, pH 6.5, 0.5 mM MgCl2.

3.2.1.3 Immunolabeling 1. 1% Bovine serum albumin (BSA) in Buffer 7. or 1% Fish skin gelatin (FSG) in Buffer 7. 2. 0.1 M Glycine in PBS Buffer 7. 3. Primary antibody. 4. Secondary antibody 10 nm gold conjugate EM grade (GE Healthcare or British Biocell International).

3.2 SEM and Immuno-SEM of Yeast Nuclei

3.2.1.4 Fixation 1. Fixative 3: 4% PFA (Agar Scientific), 20 mM potassium phosphate pH 6.5, 0.5 mM MgCl2, 0.2 M sucrose filtered using 0.4 mm syringe filter. 2. Fixative 4: 2% glutaraldehyde (Fluka), 0.2% tannic acid (TAAB laboratories) in 20 mM potassium phosphate, 0.5 mM MgCl2, pH 7.4 filtered using 0.4 mm syringe filter. 3. Fixative 5: 1% Osmium tetroxide (Agar Scientific) in dH2O. 4. Fixative 6: 1% Uranyl acetate (Agar Scientific) in dH2O. 5. Ethanol series: 30%, 50%, 70%, 95%, 100%.

3.2.2 Procedure 3.2.2.1 Nuclear isolation 1. Grow 50 ml yeast cell cultures to mid-log phase. 2. Pellet 50 ml yeast culture (3000  g, 3 min, room temperature). 3. Quickly pour off YPD and gently resuspend pellet by pipetting in 20 ml distilled water. 4. Pellet (3000  g, 3 min, room temperature). 5. Poor off supernatant and gently resuspend pellet in 30 ml of Buffer 1 (Section 3.1.1.2) and shake at 140 rpm for 35 min. 6. Pellet (3000  g, 3 min, room temperature). 7. Discard supernatant and gently resuspend in 30 ml Buffer 2a. 8. Pellet (3000  g, 3 min). 9. Discard supernatant and gently resuspend in 5 ml Buffer 3b. 10. Incubate for 30–40 min at room temperature Check every 10 min for degree of spheroplasting as describe in Section 3.1.2.1. 11. Pellet (3000  g, 3 min). 12. Resuspend pellet in 5 ml Buffer 2a. 13. Pellet (3000  g, 3 min). 14. Resuspend pellet in 5 ml Buffer 2a. 15. Subsequent steps must be kept at 4  C. 16. Prechill 200, 400, and 600 ml of Buffer 5 in 3 microfuge tubes. 17. Add 200 ml of spheroplasts (from step 14) to each tube. Causes gentle osmotic shock. 18. Flick the tubes to mix and incubate 5 min. 19. Remove 1–2 ml of each sample and check under phase contrast microscope (60–100  objective). Released nuclei are small round and phase dark and may be varied in size. Vacuoles are similar but phase bright. 20. Choose a sample dilution which gives a reasonable number of released nuclei without causing excessive lysis (usually 1:1 dilution) and prepare a fresh sample containing 200 ml spheroplasts. 21. To prepare samples for feSEM without ribosome removal, continue to Section 3.2.2.3.

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3.2.2.2 Ribosome removal To reveal certain components of the NPCs it may be necessary to remove ribosomes. We find that some features of the NPC, particularly cytoplasmic filaments, are less well preserved after salt extraction (Fiserova et al., 2014) but NPCs may be difficult to recognize when densely surrounded by ribosomes. 1. Pellet nuclei for 3 min at 3000  g. 2. Resuspend in 0.3–0.5 M KCl in 20 mM potassium phosphate pH 6.5. Try a range of KCl concentrations to optimize extraction for specific strains and preparations. 3. Incubate for 10 min at room temperature. 4. Proceed to Section 3.2.2.3.

3.2.2.3 Prefixation and attachment to chip 1. Pipette 40 ml of 1 mg/ml poly-L-lysine onto acetone cleaned silicon chips for 30 min. 2. Rinse poly-L-lysine chips in dH2O and transfer to Buffer 4. 3. Place chip face up in a modified Eppendorf tube (“microtube chamber” see Chapter 2; Kiseleva, Allen, et al., 2007). Cut off lid, place chip into the inside of the lid, then cut off bottom of the tube so that it fits tightly into the lid. 4. Pipette 0.5 ml Fixative 3 into a microtube chamber with a silicon chip at the bottom. 5. Layer 4–8 ml nuclei (with or without ribosomes) on top. You may have to optimize the amount. 6. Place microtube chamber inside 14 ml centrifuge tube with a tissue or agarose plug at the bottom to make a flat cushion. 7. Centrifuge in a swing out rotor for 10 min at 3000  g at 4  C. 8. If images of the nucleoplasmic face of the NPC are required, centrifuge at 5000  g, 10 min at 4  C. 9. Remove microtube chamber from the centrifuge tube, pipette off most of the liquid and separate chamber from lid. Remove chip and place in Fixative 3 (with sucrose omitted) for 10 min. 10. Proceed to Section 3.2.2.4 for immunogold labeling or Section 3.2.2.5 if labeling is not required.

3.2.2.4 Immunolabeling 1. Wash chips 2  in Buffer 6, 5 min in a Petri dish. 2. Transfer to 0.1 M glycine in Buffer 7 for 10 min 3. Transfer to 1% BSA in Buffer 7 for 20 min. Other blocking agents, such as fish skin gelatin (Sigma–Aldrich), may be tried if problems with high background are encountered. 4. Place wet filter paper in a 9-cm Petri dish and place Parafilm on top to make a moist chamber.

3.3 Immunogold Labeling of Yeast Ultrathin Cryosections

5. Dilute the primary antibody to an appropriate concentration in Buffer 7. You will need 10 ml of diluted antibody/chip. 6. Place chip on dry filter paper to dry the back. Ensure the front of the chip with the sample attached remains wet at all times, however, the back must be dry to prevent antibody solution wicking off. 7. Place chip on the Parafilm in the moist chamber and pipette on 10 ml antibody, close lid and incubate for 1 h. 8. Wash 2  Buffer 7, 10 min in a small Petri dish. 9. Dilute secondary antibody–gold conjugate 1:10 in Buffer 7. Higher dilutions may be tried. 10. Place chip on dry filter paper to dry the back. 11. Place chip on Parafilm in moist chamber and pipette on 10 ml secondary antibody, close lid and incubate for 1 h. 12. Wash 3  10 min Buffer 7.

3.2.2.5 Preparation of samples for FESEM 1. Transfer chips to Petri dish with Fixative 4 for 10 min. Samples may be stored in Fixative 4 at 4  C for 48 h. 2. Wash chips briefly in dH2O. 3. Fix in Fixative 5 for 10 min. 4. Wash briefly in dH2O. 5. Stain/fix 10 min with Fixative 6. 6. Dehydrate samples through ethanol series: 2 min each 30%, 50%, 70%, 95% (twice), 100% (3 ). 7. Transfer chips to critical point dryer containing 100% dry ethanol. 8. Critical point dry using high purity (< 5 ppm water) liquid CO2. Samples may be stored under vacuum for up to 1 week. 9. Coat samples with 1–3 nm chromium. Samples may be stored under vacuum to prevent rehydration and oxidation of the chromium coat, however, chromium coat will gradually deteriorate over time. 10. Visualize by feSEM (see Fiserova et al., 2014; Kiseleva et al., 2004 for examples).

3.3 IMMUNOGOLD LABELING OF YEAST ULTRATHIN CRYOSECTIONS The power of thin section TEM is the wealth of information contained in a section, with a large proportion of all the macromolecular components of the cell visible. Many of these cell components can be identified by their morphology, but many more cannot. On the other hand individual components can be readily identified by immunofluorescence, or by fluorescent protein tagging. Antibodies can be tagged with small (5–10 nm) gold nanoparticles which can be unequivocally identified in the TEM. Excellent secondary antibody–gold conjugates are commercially available so that immunofluorescence protocols can be adapted for immunogold labeling by

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substituting fluorescent- for gold-tagged antibodies. Ideally, all the information available in a TEM thin section should be maintained, but standard fixation and embedding methods are not usually compatible with maintaining antigenicity and antigen accessibility. The Tokuyasu (1973) method was developed to preserve antigenicity, accessibility, and ultrastructure. It involves light fixation, followed by embedding in saturated sucrose, freezing in liquid nitrogen, then cryosectioning. Sections are thawed and the sucrose washed out, leaving a resin-less section where the ultrastructure is preserved, but antigens are accessible. Immunogold labeling can then be carried out much like immunofluorescence, although generally antibody reagents are used at higher concentrations and/or for longer. Unlike many immunofluorescence protocols no detergents are used and membranes are well preserved. In fact the position of membranes is the main ultrastructural information present. As with the other methods presented here, the main difficulty with yeast is the presence of the cell wall because it inhibits the entry of the viscous saturated sucrose into the cell which is essential to prevent ice crystal formation and damage during freezing. Treatment of fixed cells with periodic acid extracts cell wall components and allows excellent penetration of sucrose (Griffith et al., 2008), allowing the implementation of the Tokuyasu technique in yeast. Figure 3.2 shows the labeling of the N-terminal domain of Nup116p in wild-type budding yeast using this technique.

FIGURE 3.2 Immunogold labeling of thawed cryosection of a yeast cell using an antibody raised against Nup116p (WU956, Bucci & Wente, 1998), then a 10-nm gold-tagged secondary antibody. Arrows indicate NPCs. Black dots are the colloidal gold particles.

3.3 Immunogold Labeling of Yeast Ultrathin Cryosections

3.3.1 Materials 3.3.1.1 Fixation, embedding in gelatin, and sucrose infiltration 1. 1 N NaOH (VWR International, Lutterworth, UK). 2. 8% PFA stock solution: To make 100 ml solution, mix 8 g of PFA (Agar Scientific) with 90 ml of distilled water and heat to 60  C. While stirring add 1 N NaOH drop wise until the solution clears. Cool and make it up to 100 ml with distilled water. 3. 0.4 M PHEM buffer: Dissolve three pellets of NaOH in 600 ml of distilled water. Add 72.5 g PIPES (240 mM final) and stir until dissolved. Gradually add 15.2 g EGTA (40 mM final) while stirring. Dissolve 23.8 g HEPES (100 mM final) in 200 ml of distilled water and add this to the above mixture. Add 1.63 g MgCl2 (8 mM final). Adjust pH to 6.9 and adjust volume to 1l with distilled water. 4. Double strength fixative: 4% PFA, 0.4% glutaraldehyde (Agar Scientific) in 0.1 M PHEM buffer pH 6.9: To make double strength fixative mix together 25 ml of 8% PFA, 800 ml of 25% glutaraldehyde and 12.5 ml of 0.4 M PHEM buffer. Adjust pH to 6.9 and make up to 50 ml with distilled water. 5. Single strength fixative: dilute double strength fixative 1:1 with water. 6. 1% Periodic acid in 0.1 M PHEM: Dissolve 100 mg of periodic acid (Sigma– Aldrich, UK) in 2.5 ml of 0.4 M PHEM and make up to 10 ml. Periodic acid helps permeabilize cell wall and aids sucrose infiltration for improved freezing and ultrastructure (Griffith et al., 2008; van Tuinen & Riezman, 1987). 7. 0.2 M phosphate buffer pH 7.4. Make stock solution (A) by dissolving 35.6 g Na2HPO4 2H2O in 500 ml of distilled water and adjust the volume to 1l. Stock solution (B): dissolve 27.6 g NaH2PO4H2O in 500 ml of distilled water and adjust the volume to 1l. Mix 81 ml of stock A and 19 ml of stock B to produce 100 ml of 0.2 M phosphate buffer pH 7.4. 8. 10% Sodium azide. 9. 10  PBS. Dissolve in 750 ml of water: 80 g NaCl, 2.0 g KCl, 14.4 g Na2HPO4, and 2.4 g KH2PO4. Adjust pH to 7.4 with NaOH and bring the volume to 1l with distilled H2O and sterilize. Store at 4  C. Make 1  fresh as required. 10. 10% Gelatin (Dr. Oetker GmbH, Villach, Austria) in PBS. Add 20 g of gelatin to 200 ml PBS. Stir at room temp for a few minutes then heat to 60  C for 4–6 h. Allow to cool to 37  C then add 400 ml of 10% Sodium azide. Pour gelatin into 5 ml vials and refrigerate. 11. 2.3 M sucrose in 0.1 M phosphate buffer: Dissolve 78.7 g of sucrose in 0.1 M phosphate buffer pH 7.4 to a final volume of 100 ml. Aliquot into glass vials and store at 4  C. 12. Liquid nitrogen. 13. 1.5 ml microfuge tubes. Use clear walled microfuge tubes. Since the pellets are pale in color, they will be easier to see in clear walled tubes.

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14. Light microscope. 15. Dissecting microscope. 16. Single edged razor blades, acetone washed (Agar Scientific). 17. Fine forceps (Agar Scientific). 18. End over end rotator (Agar Scientific). 19. Fixed angle specimen rotator (Agar Scientific). 20. Cryovials and racks or canes. 21. Liquid nitrogen storage facilities. 22. Bench centrifuge. A swing out rotor is preferred as the pellet will form nicely in the bottom of the microfuge tube. If a fixed angle rotor is used the pellet may smear up one side.

3.3.1.2 Cryosectioning 1. 2. 3. 4. 5.

Leica EM UC6 Ultramicrotome and Leica EM FC6 cryochamber. Liquid nitrogen. Ultramicrotome Specimen pins (Leica Microsystems GmbH). 2.3 M sucrose solution in 0.1 M phosphate buffer 1% Toluidine Blue Stain. see Section 3.1.1.4. 6. 1.1 M Na2CO3 7. Polyvinylpyrrolidone (PVP), mean mol wt: 10,000 (Sigma–Aldrich). 8. PVP/Sucrose mixture. Mix 4 g of PVP and 0.6 ml of 1.1 M Na2CO3 to a paste and then add 17 ml of 2 M sucrose solution. The liquid will clear if you allow it stand overnight at room temperature. Store in aliquots at 20  C. 9. Haug static line Antistatic device (Haug GmbH, Germany). 10. Diamond knife (e.g., Diatome dry cryo 45  C). 11. Leica EM KMR2 knife making machine (Leica Microsystems). 12. Glass knives. 13. Wire loops of 2.5 and 4 mm diameter. 14. Eyelash probes. Made by attaching a cleaned eyelash or eyebrow hair to a wooden stick approximately 15 cm long. 15. Glass microscope slide. 16. 60  C Hot plate. 17. Freshly carbon coated Formvar 200-mm mesh nickel grids (Agar Scientific) on a stickered glass slide. Avoid using copper grids for immunogold labeling as some buffers (e.g., Tris, PBS) will react with the copper and produce a fine precipitate over the specimen. Cu2þions may also have an inhibitory effect on antibody function. A stickered microscope slide makes it easier to remove the grids with cryosections after storage and also allows for easy labeling. We use Avery No. L7162 Quick peel addressing labels. To render it hydrophilic the Formvar film on the grids is coated with a thin layer carbon just before commencing sectioning.

3.3 Immunogold Labeling of Yeast Ultrathin Cryosections

3.3.1.3 Immunogold labeling 1. 10% gelatin in PBS. 2. 2% gelatin plates. Warm 20 ml of 10% gelatin solution to 37  C and make it up to 100 ml with warm PBS. Pour a thin layer into 2.5 cm Petri dishes and cool. Store at 4  C. 3. 2% Methyl cellulose. Heat 196 ml of distilled water to 90  C then, while stirring, add 4 g methyl cellulose (25 centipoise) (Sigma–Aldrich). Cool the solution quickly to 10  C on ice, while stirring. Place in a 4  C room overnight and maintain stirring. The following day discontinue stirring and incubate for a further for 3 days at 4  C. Centrifuge solution at 4  C and 60,000 rpm for 60 min. Store 25 ml aliquots of the supernatant in tubes wrapped in foil at 4  C. The solution can store like this for 3 months. 4. 0.1% BSAc in PBS (Aurion, Netherlands). 5. 4% Uranyl acetate pH 4. Dissolve 0.4 g of uranyl acetate in 10 ml of distilled water and the pH 4. Add 90 ml 6. 1.8% Methyl cellulose, 0.4% uranyl acetate pH 4 (MC/UA). methyl cellulose solution to 10 ml 4% uranyl acetate pH 4 and mix gently. Can be stored at 4  C in the dark for 3 months.

3.3.2 Procedure 3.3.2.1 Yeast culture 1. Plate frozen yeast stock on YPD or SD selective plates and grow colonies. 2. On the morning of the day before fixation, inoculate 10 ml of liquid YPD or SD media with a single colony and grow on a horizontal shaker at 180 rpm for about 8–10 h at 30  C. Check the OD600 of the culture. 3. For wild-type cells, assuming a culture doubling time is 2 h; calculate the culture dilution required to inoculate 50 ml media so that after overnight growth, the culture will be mid-log phase with an OD600 value of 0.5–1, equivalent to 1–5  107 cells/ml. The dilution usually corresponds 1/100–1/200 of the final media volume (50 ml). 4. Inoculate 50 ml medium with the required dilution and grow overnight on a horizontal shaker at 180 rpm at 30  C.

3.3.2.2 Fixation, embedding in gelatin, and sucrose infiltration 1. Grow culture to 1 OD600 2. Add an equal volume of double strength fixative to the culture and fix for 15–20 min at room temperature in an end over end tumbler. 3. Pellet the yeast by centrifugation at 200  g for 2–3 min. Resuspend in fresh single strength fixative and continue fixation for a further 3 h at room temperature. 4. Pellet and resuspend in 1 ml of 0.1 M PHEM buffer in a 1.5-ml microfuge tube. 5. Pellet and resuspend three more times in 1 ml of 0.1 M PHEM buffer.

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6. Pellet and resuspend in 1 ml of freshly prepared 1% periodic acid in 0.1 M PHEM buffer and incubate for 1 h at room temperature on an end over end tumbler. 7. Pellet and resuspend the clumps of cells 3  in 1 ml of 0.1 M PHEM buffer. 8. Pellet and resuspend in 10% gelatin in PBS buffer at 37  C for 10 min. 9. Repellet and then place tubes on ice until gelatin sets. 10. Carefully cut the tip, containing the pellet, off the microfuge tube, and then under a dissecting microscope prepare small blocks 1 mm3 or smaller Must be done on ice to keep from the pellet using a clean razor blade. gelatin solid. 11. Infiltrate the pieces in 2.3 M sucrose 0.1 M phosphate buffer at 4  C overnight on a fixed angle specimen rotator. Care must be taken to ensure that the tissue stays submerged at all times and is not allowed to stick to the sides of the vial where the sucrose could crystallize. 12. Transfer infiltrated specimen blocks onto clean metal pins, remove excess sucrose with a damp filter paper, and immerse the specimen pin and specimen in liquid nitrogen until freezing is complete (it stops bubbling) and then store liquid nitrogen until required for sectioning. Do this step quickly over ice or in a cold room. Any changes to the concentration of the sucrose will alter the cutting properties of the frozen block. Water evaporating from the specimen will increase the sucrose concentration producing softer blocks and possibly crystalline sucrose. When plunging the pin into the LN2 move it back and forth rapidly a few times until the bubbling stops to minimize the Leidenfrost effect which can slow down the freezing rate.

3.3.2.3 Cryosectioning 1. Setup the cryoultramicrotome according to the manufacturer’s instructions. 2. Set the cryochamber, knife, and specimen temperature to 100  C. Face and trim the block with a glass knife using the “mesa trimming technique” and then, with a fresh part of the knife, cut semithin cryosections at a thickness of 200 nm, and a cutting speed of 2.5–3 mm/s. Mesa trimming produces a raised flat topped rectangular platform with smooth parallel sides which is ideal for producing ribbons of sections. Briefly a glass knife is used to flatten the surface of the block and then moved to the right of the area of interest where a 50- to 100-mm step is cut into the surface of the block. Now the whole block is turned through 90  and another step is cut into block, avoiding the area of interest. This is repeated twice more until the small raised mesa containing the area of interest is produced (Bozzola & Russell, 1999). 3. Retrieve sections using a 2.5-mm wire loop filled with a drop of PVP/Sucrose mixture. Touch the thawed droplet to the surface of a clean microscope slide the sections will stay on the slide. Cover the area where the sections are with 1%

3.3 Immunogold Labeling of Yeast Ultrathin Cryosections

4. 5.

6.

7.

Toluidine Blue Stain in and place on a hotplate for 2 min, wash off the stain with distilled water and view with a brightfield microscope. Retrim the block face using the “mesa trimming technique” with a glass knife to produce a block face 0.6–0.4  0.3–0.4 mm. Swing the cryo diamond knife into the cutting position and reset the cryochamber, knife, and specimen temperature to 120  C. Cut ultrathin sections at a thickness of 70–80 nm and a cutting speed of 0.4 and 1.00 mm/s. If static causes the sections to stick and pile up on the knife, use the anti static device. Using the Anti static device: (a) Start sectioning with the ionizer at a high position and slow cutting speed, 0.4–0.6 mm/s. (b) Reduce the voltage after a few sections, try increasing the cutting speed 0.8–1.00 mm/s. (c) If sections stick at the cutting edge: increase the voltage. (d) If sections lift up from the knife surface: reduce the voltage. (e) Switch ionizer to position 1 or 2 or even off while moving sections with a hair. (f ) Always switch the ionizer off during pickup of the sections. Carefully guide the sections away from the knife edge down the knife with the eyelash and move them to the side to make pickup easier and safer. Turn off the anti static device and retrieve the ribbon of sections with the loop containing PVP/sucrose solution. Wait about 10 s for the droplet to thaw then slowly touch it to the surface of a freshly carbon coated Formvar 200-mm mesh nickel grid that is held on a stickered glass slide. The sections will adhere to the grid. The grids can be stored on the glass at 4  C in a microscope slide storage box for up to 6 months with no adverse affects.

3.3.2.4 Immunogold labeling For steps 1–6: Put the grids onto cold gelatin plates before melting then warmed to 37  C for 30 min to remove the dried mixture of sucrose and methyl cellulose. It is important from now on to always keep the section side of the grid wet and the back surface dry. Non specific antibody reactions are dealt with by the following strategies: residual aldehyde groups are quenched in 0.1% glycine, positively charged specimen compounds are blocked with a BSA and 0.1% BSAc. Acetylated 0.1%, BSAc blocking is used during both primary and secondary antibody incubation as it has increased negative charge. 1. Place the grids, section side down onto cold 2% gelatin plates and melt in 37  C oven for 30 min. 2. Rinse with 0.1% glycine in PBS 5  for 1 min. 3. Block in 1% BSA in PBS 3  for 1 min. 4. Incubate in 0.1% BSAc in PBS 3  for 5 min. 5. Incubate with primary antibody diluted in 0.1% BSAc in a wet chamber using 10 ml droplets per grid for 1 h at room temperature or overnight at 4  C.

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Test serial dilutions to find the best working concentration of your antibody. To confirm that any labeling is specific a number of controls should also be performed in parallel with the antibody in use, for example, omit the primary antibody and replace with incubation buffer only, replace the primary antibody with nonimmune serum from the same species, use another antibody from the same species that is specific for an antigen known to be absent in the tissue. 6. Rinse in 0.1% BSAc in PBS 5  for 2 min. 7. Incubate with colloidal gold-conjugated secondary antibody in a wet chamber using a 5- to 10-ml antibody droplet per grid diluted 1:20–1:100 in 0.1% BSAc in PBS. Dilution of the secondary antibody for EM is usually between 1:20 and 1:100 depending on the secondary antibody. It is recommended to test serial dilutions to find the appropriate working concentration. 8. Incubate for 1 h at room temperature. 9. Wash in 0.1% BSAc in PBS 3  for 5 s each. 10. Rinse in 0.1% BSAc PBS 4  for 5 min each. 11. Wash in PBS times for 3  for 5 min each. 12. Stabilize with 1% glutaraldehyde in PBS for 5 min each. Optional step but recommended to stabilize antigen–antibody–gold complex by fixation. This may prevent loss of labeling during subsequent steps. 13. Wash in distilled water 10  for 1 min each. Uranyl acetate precipitates in the presence of phosphate ions so the PBS must be thoroughly washed from the sections before final staining and embedding in MC/UA. 14. Rinse quickly over two drops of cold MC/UA pH 4. MC/UA embedding is done on ice because the solution is less viscous in the cold. 15. Incubate in fresh drop of cold MC/UA pH 4 for 6–8 min each. 16. Loop out the grids with the 4-mm wire loops and reduce the MC/UA to an even thin film and let air dry for 30 min. Drain excess MC/UA from the loop by placing the loop at a 45–60 angle onto a clean filter paper. Once the methyl cellulose starts to be absorbed by the paper, gently drag the loop away from the wet patch until no more liquid is removed. When the film is dry, the grids can be carefully removed from the wire loop using pointed forceps. 17. View in TEM (see Fig. 3.2 for example of immunogold labeling of GLFG repeats of Nup116p).

CONCLUSIONS AND PERSPECTIVES We present here three protocols used to analyze ultrastructural aspects of yeast NPCs, nuclear envelope, and nuclear transport. Conventional TEM is straightforward, reliable, and readily available to most cell biologist. It can, for instance, be used to obtain information concerning cell and nuclear envelope morphology in mutant strains, but careful comparison to wild-type strains is essential to take into

References

account fixation artifacts. Information about NPC dimensions is also obtainable, but this must be quantified in order to take account of the plane of section (Fiserova et al., 2010). If serial sections are collected, information concerning NPC numbers in different strains/conditions can be accurately estimated using stereological techniques (Mayhew, Muhlfeld, Vanhecke, & Ochs, 2009). If fine morphological details are required, however, HPF/FS methods are preferred (Fiserova & Goldberg, 2010), but these are time consuming, expensive, and can be difficult to troubleshoot. FeSEM can be used to image surfaces at relatively high resolution and can give information about NPC and NE structure that is difficult to obtain any other way. It is therefore ideal for studying certain peripheral NPC structures (cytoplasmic and nucleoplasmic filaments, rings, central structures, etc.). Because the surface of interest has to be exposed and yeast NPCs appear to be fragile and are often lost during nuclear isolation, particular care must be taken to prevent proteolysis and physical damage. An alternative, that avoids nuclear isolation, is freeze fracture EM, which because it involves freezing cell components in situ, mitigates for any reorganization that may occur during sample preparation, and the physical and biochemical fragility of NPCs is not a problem. However, because of the nature of freeze fracturing, structural details are not easily visible or interpretable, and this is a very technically demanding technique, requiring specialist equipment, and expertise. On the other hand it is an excellent method to study NPC distribution (Doye et al., 1994), although current “super resolution” light microscopy methods may soon be preferred (Lo¨schberger et al., 2012; see also Chapter 10). A further advantage of feSEM is that existing immunofluorescence protocols can usually be adapted for feSEM. Immunogold labeling for TEM is not usually possible with conventional resin embedded sections. It is sometimes possible with HPF/FS sections, but the most reliable and simplest method for immuno-EM is the use of resin-less sections for labeling. It may be possible to combine HPF with resin-less section labeling (Griffith et al., 2008, unpublished data), but at the cost of a more demanding, time consuming method with uncertain benefits.

Acknowledgments Thanks to Helen Grindley for technical assistance. Work was supported by Biotechnology and Biological Sciences Research Council, UK (grant numbers BB/E015735/1 and BB/ G011818/1).

References Akey, C. W., & Radermacher, M. (1993). Architecture of the Xenopus nuclear pore complex revealed by three-dimensional cryo-electron microscopy. The Journal of Cell Biology, 122, 1–19. Beck, M., Fo¨rster, F., Ecke, M., Plitzko, J. M., Melchior, F., Gerisch, G., et al. (2004). Nuclear pore complex structure and dynamics revealed by cryoelectron tomography. Science, 306, 1387–1390.

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Bozzola, J. J., & Russell, L. D. (1999). Electron microscopy: Principles and techniques for biologists (2nd ed.). Sudbury, MA: Jones and Bartlett Publishers. Bucci, M., & Wente, S. R. (1998). A novel fluorescence-based genetic strategy identifies mutants of Saccharomyces cerevisiae defective for nuclear pore complex assembly. Molecular Biology of the Cell, 9, 2439–2461. Doye, V., Wepf, R., & Hurt, E. C. (1994). A novel nuclear pore protein Nup133p with distinct roles in poly(A)þ RNA transport and nuclear pore distribution. The EMBO Journal, 13, 6062–6075. Feldher, C. M., Kallenbach, E., & Schultz, N. (1984). Movement of a karyophilic protein through the nuclear pores of oocytes. The Journal of Cell Biology, 99, 2216–2222. Fiserova, J., & Goldberg, M. W. (2010). Immunoelectron microscopy of cryofixed freeze substituted Saccharomyces cerevisiae. Methods in Molecular Biology, 657, 191–204. Fiserova, J., Richards, S. A., Wente, S. R., & Goldberg, M. W. (2010). Facilitated transport and diffusion take distinct spatial routes through the nuclear pore complex. Journal of Cell Science, 123, 2773–2780. Fiserova, J., Spink, M., Richards, S. A., Saunter, C., & Goldberg, M. W. (2014). Entry into the nuclear pore complex is controlled by a cytoplasmic exclusion zone containing dynamic GLFG-repeat nucleoporin domains. Journal of Cell Science, 127, 124–136. Gall, J. G. (1967). Octagonal nuclear pores. The Journal of Cell Biology, 32, 391–399. Goldberg, M. W., & Allen, T. D. (1992). High resolution scanning electron microscopy of the nuclear envelope: Demonstration of a new, regular, fibrous lattice attached to the baskets of the nucleoplasmic face of the nuclear pores. The Journal of Cell Biology, 119, 1429–1440. Goldberg, M. W., & Fiserova, J. (2010). Immuno-gold labelling for scanning electron microscopy. Methods in Molecular Biology, 657, 297–313. Goldberg, M. W., Huttenlauch, I., Hutchison, C. J., & Stick, R. (2008). Filaments made from A- and B-type lamins differ in structure and organization. Journal of Cell Science, 121, 215–225. Griffith, J., Mari, M., Maziere, A. D., & Reggiori, F. (2008). A cryosectioning procedure for ultrastructural analysis and the immunogold labelling of yeast Saccharomyces cerevisiae. Traffic, 9, 1060–1072. Kiseleva, E., Allen, T. D., Rutherford, S., Bucci, M., Wente, S. R., & Goldberg, M. W. (2004). Yeast nuclear pore complexes have a cytoplasmic ring and internal filaments. Journal of Structural Biology, 145, 272–288. Kiseleva, E., Allen, T. D., Rutherford, S. A., Murray, S., Morozova, K., Gardiner, F., et al. (2007). A protocol for isolation and visualization of yeast. Nature Protocols, 2, 1943–1953. Kiseleva, E., Morozova, K. N., Voeltz, G. K., Allen, T. D., & Goldberg, M. W. (2007). Reticulon 4a/NogoA locates to regions of high membrane curvature and may have a role in nuclear envelope growth. Journal of Structural Biology, 160, 224–235. Lo¨schberger, A., van de Linde, S., Dabauvalle, M. C., Rieger, B., Heilemann, M., Krohne, G., et al. (2012). Super-resolution imaging visualizes the eightfold symmetry of gp210 proteins around the nuclear pore complex and resolves the central channel with nanometer resolution. Journal of Cell Science, 125, 570–575. Mayhew, T. M., Muhlfeld, C., Vanhecke, D., & Ochs, M. (2009). A review of recent methods for efficiently quantifying immunogold and other nanoparticles using TEM sections through cells, tissues and organs. Annals of Anatomy, 191, 153–170.

References

McDonald, K. (2007). Cryopreparation methods for electron microscopy of selected model systems. Methods in Cell Biology, 79, 23–56. Murray, S., & Kiseleva, E. (2008). A protocol for isolation and visualization of yeast nuclei by scanning electron microscopy. Methods in Cell Biology, 88, 367–387. Ris, H. (1989). Three-dimensional imaging of cell ultrastructure with high resolution low voltage SEM. Institute of Physics Conference Series, 98, 657–662. Stevens, B. J., & Swift, H. (1966). RNA transport from nucleus to cytoplasm in Chironomus salivary glands. The Journal of Cell Biology, 31, 55–77. Titus, L. C., Dawson, T. R., Rexer, D. J., Ryan, K. J., & Wente, S. R. (2010). Members of the RSC chromatin-remodeling complex are required for maintaining proper nuclear envelope structure and pore complex localization. Molecular Biology of the Cell, 21, 1072–1087. Tokuyasu, K. T. (1973). A technique for ultracryotomy of cell suspensions and tissues. The Journal of Cell Biology, 8, 377–383. van Tuinen, E., & Riezman, H. (1987). Immunolocalization of glyceraldehyde-3-phosphate dehydrogenase, hexokinase, and carboxypeptidase Y in yeast cells at the ultrastructural level. The Journal of Histochemistry and Cytochemistry, 35, 327–333. Wente, S. R., & Blobel, G. (1993). A temperature-sensitive NUPll6 null mutant forms a nuclear envelope seal over the yeast nuclear pore complex thereby blocking nucleocytoplasmic traffic. The Journal of Cell Biology, 123, 275–284.

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CHAPTER

Exploring Nuclear Pore Complex Molecular Architecture by Immuno-Electron Microscopy Using Xenopus Oocytes

4

Nelly Pante´*, and Birthe Fahrenkrog{ *

Department of Zoology, Life Sciences Centre, University of British Columbia, Vancouver, Canada Institute for Molecular Biology and Medicine, Universite Libre´ de Bruxelles, Charleroi, Belgium

{

CHAPTER OUTLINE Introduction .............................................................................................................. 82 4.1 Materials........................................................................................................... 86 4.1.1 Equipment...................................................................................... 86 4.1.2 Materials ........................................................................................ 86 4.1.3 Xenopus ......................................................................................... 86 4.1.4 Reagents ........................................................................................ 86 4.1.5 Buffers ........................................................................................... 87 4.1.6 Plasmids ........................................................................................ 87 4.2 Experimental Strategies ..................................................................................... 87 4.3 Preparation of Antibodies Conjugated with Colloidal Gold Particles....................... 89 4.4 Immunogold Labeling of Nucleoporins Using Anti-Nucleoporin Antibodies ............. 90 4.5 Immunogold Labeling of Nucleoporins Using Epitope-Tagged Nucleoporins ........... 93 4.5.1 Design of the Constructs .................................................................. 95 4.5.2 Protocol.......................................................................................... 95 Concluding Remarks ................................................................................................. 96 Acknowledgments ..................................................................................................... 96 References ............................................................................................................... 96

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00004-7

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CHAPTER 4 Imaging Xenopus Oocytes NPCs by Immuno-EM

Abstract Xenopus oocytes are large in size and perfectly suited for microinjection experiments. Their nuclei, which can be readily isolated manually, are characterized by an extremely high density of nuclear pore complexes (NPCs). Therefore, Xenopus oocytes are an excellent system to study NPC structure and molecular architecture, as well as nucleocytoplasmic transport on an ultrastructural level. A wide range of electron microscopy (EM) techniques can be employed to do so and thin-sectioning immuno-EM has been proven to be a powerful tool in this context. NPCs are composed of multiple copies of a set of about 30 different nucleoporins, which are often large, multidomain proteins. Their complex organization within NPCs can be unraveled by using domain-specific antibodies to individual nucleoporins in combination with microinjection and expression of epitope-tagged nucleoporins. Here, we describe the immuno-EM methods using Xenopus oocyte that allow for precise ultrastructural localization of nucleoporins within the structure of the NPC.

INTRODUCTION The large size of Xenopus oocytes, typically 2–3 mm in diameter, and their nuclei (about 0.4 mm in diameter) has made them an excellent system to study the function, ultrastructure, and molecular architecture of the nuclear pore complex (NPC). The NPC is an elaborate macromolecular complex of about 120 MDa embedded in the double membrane of the nuclear envelope (NE) that mediates the bidirectional trafficking of macromolecules between the nucleus and the cytoplasm. The large size of both oocytes and their nuclei makes their manipulation relatively easy and allows the work in an intact system without the need of, for example, detergents to access the nucleus. Consequently, not only does the nuclear transport system remain intact, but also the integrity of NPCs is retained. Microinjection of Xenopus laevis oocytes was first used to study nuclear transport (Dworetzky & Feldherr, 1988; Feldherr, 1969; Feldherr, Kallenbach, & Schultz, 1984), and this system has provided insights into nuclear import and export mechanisms. For this procedure, the transport substrate (e.g., proteins or RNAs) is first conjugated with colloidal gold particles and then microinjected into the oocytes. After preparation of the oocytes by embedding and thin-section electron microscopy (EM), the fates of the gold particles are then determined by transmission EM (TEM). Comparison of the location of gold particles at different times with reference to landmarks on NPC structure visible by high-resolution EM allows to deduce the path taken by a particular transport substrate. In addition, when combined with experimental conditions that inhibit nuclear transport, the gold particles (coated with nuclear transport substrates) can be arrested at distinct sites of the NPC, revealing transport arrested intermediates and distinct molecular steps in the mechanism of nuclear transport (Gorlich, Pante, Kutay, Aebi, & Bischoff, 1996; Pante & Aebi, 1996;

Introduction

Rollenhagen, Muhlhausser, Kutay, & Pante, 2003; Rollenhagen & Pante, 2006). This approach can be used to study nuclear import or nuclear export by injecting the oocytes with the transport substrate in the cytoplasm or the nucleus respectively. This system has also been adapted for the study of nuclear import of several viruses (Au, Cohen, & Pante, 2010; Au & Pante, 2012; Cohen & Pante, 2005; Pante´ & Kann, 2002; Rabe, Vlachou, Pante, Helenius, & Kann, 2003). The detailed protocols for microinjection of Xenopus oocytes with transport substrates and their preparation for TEM have been previously described (Au et al., 2010; Pante, 2006; see also Chapter 18). In addition, to the ease for microinjection and well-established EM protocols to yield very well preserved NPCs, the Xenopus oocyte system also has the advantage of having a high number of NPC, 5  107 NPCs (60 NPCs/mm2; Grossman, Medalia, & Zwerger, 2012). This is extremely large compared for example with nuclei from human cells, which contain only 3–5 NPC/mm2. Thus, a wide spectrum of EM techniques that range from negative staining to tomographic studies have been used to study the ultrastructure of the Xenopus oocyte NPC (Akey & Radermacher, 1993; Grossman et al., 2012; Hinshaw, Carragher, & Milligan, 1992; Maco, Fahrenkrog, Huang, & Aebi, 2006; Stoffler et al., 2003). A frequently used method is embedding and thin-sectioning EM, which is more accessible to researches than, for example, cryoelectron tomography. For this technique, specimen is fixed, dehydrated, and embedded in an epoxy-based resin, such as Epon 812, which assures a good structural preservation of the sample. This good structural preservation is of high importance when looking at the ultrastructure of NPCs. The epoxy-embedded specimen is then cut into extremely thin sections (50–70 nm thick), and the sections are stained and visualized with a TEM. When intact Xenopus oocytes or isolated Xenopus nuclei are prepared by this methodology, long stretches along the NE containing a large number of NPCs are revealed in cross sections or side views (Fig. 4.1A and B) and in top views from the cytoplasmic side (Fig. 4.1C). In intact oocytes, the cytoplasm and nuclear structure adjacent to the NE are revealed (Fig. 4.1A), whereas in isolated nuclei the cytoplasm is absent and the nuclear structure is often lost because the nuclear content leaks during the sample preparation. Therefore, Xenopus nuclei appear in thin-section electron micrographs as isolated NEs, which allows to distinguish not only individual NPCs but also their fine structural subunits (Fig. 4.1B). In these micrographs, the NPC yields a tripartite structure with (1) an electron dense membrane-embedded central framework, which includes a cytoplasmic and a nuclear ring; (2) cytoplasmic filaments in the NPC cytoplasmic periphery; and (3) an assembly of filaments at the nuclear periphery, known as the nuclear basket (Fig. 4.1D and E). At the molecular level, the NPC is composed of multiple copies of roughly 30 different proteins, known as nucleoporins (Cronshaw, Krutchinsky, Zhang, Chait, & Matunis, 2002; Ori et al., 2013; Rout et al., 2000). Embedding thin-sectioning EM of isolated Xenopus nuclei in combination with immunogold labeling has been used to determine the detailed localization of some of these nucleoporins within the structure of the NPC (Chatel, Desai, Mattheyses, Powers, & Fahrenkrog, 2012;

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CHAPTER 4 Imaging Xenopus Oocytes NPCs by Immuno-EM

A

C

B

D

c

c

n

n

E Cytoplasmic filaments Nuclear envelope

Central framework

Nuclear basket

FIGURE 4.1 EM visualization of nuclear pore complexes. Cross section along an NE from (A) an intact Xenopus oocyte with the adjacent cytoplasm (c) and nucleoplasm (n) and (B) an isolated Xenopus nucleus largely freed from cytoplasmic and nuclear contents. (C) Radial section of a small patch of an NE revealing the cytoplasmic face of NPCs. (D) Selected example of a well-preserved NPC revealing its roughly tripartite structural organization. (E) Schematic representation of the main structural components of the NPC. These include the central framework embedded in the NE, the cytoplasmic filaments, and the nuclear basket. Scale bars, 100 nm.

Fahrenkrog & Aebi, 2003; Fahrenkrog et al., 2002; Mansfeld et al., 2006; Pante, Bastos, McMorrow, Burke, & Aebi, 1994; Pante, Thomas, Aebi, Burke, & Bastos, 2000; Paulillo et al., 2005; Schwarz-Herion, Maco, Sauder, & Fahrenkrog, 2007). For this method, nuclei are manually isolated from the Xenopus oocytes, and then incubated with antibodies against the nucleoporin of interest, which have been directly conjugated to colloidal gold. After immunogold labeling, the Xenopus oocyte nuclei are most commonly prepared for embedding–thinsectioning TEM. However, immunogold-labeled nuclei could also be prepared for

Introduction

TEM by quick-freezing/freeze-drying/rotary metal shadowing (Fahrenkrog et al., 2002; Pante´ & Aebi, 1994), or their surface can be visualized by high-resolution field emission scanning EM (Allen et al., 2008). Nucleoporins are often rather large proteins with multiple domains so that antibodies against the distinct domains are required to precisely determine the position and organization of a nucleoporin within the structure of the NPC (Fahrenkrog et al., 2002; Frosst, Guan, Subauste, Hahn, & Gerace, 2002). To overcome the limitation of unavailable anti-nucleoporin antibodies, microinjection of mRNA or plasmid DNA into the oocytes can be used to express epitope-tagged versions of nucleoporins in the oocytes (Fahrenkrog & Aebi, 2003; Fahrenkrog et al., 2002; Pante et al., 2000). The tagged nucleoporins can then be detected by immunogold labeling using antibodies against the respective tag. This approach can be useful (1) to confirm the results of the standard immunogold-labeling experiment using antibodies against the nucleoporins, (2) to overcome a lack of specificity of antibodies resulting from the high cross-reactivity of many anti-nucleoporin antibodies, and (3) to determine the exact location of the N- and C-terminus of a nucleoporin within a single NPC by expressing a double-tagged nucleoporin (Fahrenkrog & Aebi, 2003). For both approaches, preembedding labeling is used (i.e., labeling with the antibody is performed prior to embedding of the sample), which brings about the big advantage of using epoxy resins to preserve the detailed structure of the NPC. The localization of the nucleoporin Nup153 nicely illustrates the power of combining these two methodologies. Nup153 is organized in three domains: an N-terminal domain, a central zinc finger, and a C-terminal domain (Ball & Ullman, 2005). Several immuno-EM studies consistently revealed that Nup153 resides at the nuclear face of the NPC, but the exact location had remained controversial: two studies found Nup153 close to the NE at the nuclear ring (Pante et al., 2000; Walther et al., 2001) and one study found it at the distal ring of the nuclear basket (Pante et al., 1994). The use of antibodies against the three domains of Nup153 in Xenopus oocyte nuclei unraveled the discrepancy and revealed that the N-terminal domain of Nup153 resides at the nuclear ring, the zinc-finger domain is found at the distal ring of the nuclear basket, and the C-terminal domain, which is highly flexible, was found all over the nuclear basket and even at the cytoplasmic face of the NPC (Fahrenkrog et al., 2002). Similarly, expression of epitope-tagged versions of Nup153 in Xenopus oocyte nuclei was useful to confirm (1) the location of the N-terminal domain of Nup153 at the nuclear ring (Fahrenkrog et al., 2002; Pante et al., 2000) and (2) the multiple locations of the C-terminal domain of Nup153 within the NPC (Fahrenkrog & Aebi, 2003; Fahrenkrog et al., 2002). In this chapter, we describe in detail the protocols for performing preembedding immunogold labeling used to localize nucleoporins within the ultrastructure of the NPC. This includes the protocol using anti-nucleoporin antibodies, as well as the protocol to localize epitope-tagged nucleoporins.

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CHAPTER 4 Imaging Xenopus Oocytes NPCs by Immuno-EM

4.1 MATERIALS 4.1.1 Equipment Beckman tabletop ultracentrifuge with rotor TLA 120.2 Dissecting microscope EM specimen trimmer device (Leica, Reichert Ultratrim) Leica ultramicrotome EM (Leica Microsystems, Vienna, Austria) Microinjector and manipulator for Xenopus oocyte injection Oven Transmission EM

4.1.2 Materials 200 ml and 20-ml pipettors and pipette tips Anesthetic and surgical equipment to isolate a small piece of ovary from a frog Copper EM grids coated with Parlodion (200 mesh) Double depression glass slides (Menzel Gla¨ser) Fine forceps (Dumont 7, curved) Razor blades Terasaki microwell plates (60 wells, 10 ml volume; Greiner, Nunc) Tissue culture dishes (35 mm; Falcon, BD) Ultra 45  C room temperature diamond knife (Diatome, Biel, Switzerland)

4.1.3 Xenopus Female X. laevis frogs, larger than 9 cm (Nasco, Fort Atkinson, WI, USA).

4.1.4 Reagents Acetone Antibodies (Table 4.1) BSA Collagenase (C0130, Sigma-Aldrich) Epon 812 resin (Fluka, Buchs, Switzerland) Ethanol Glutaraldehyde, EM grade (G7776, Sigma-Aldrich) Lead nitrate (11520, Sigma-Aldrich) Low melting point agarose (A2576, Sigma-Aldrich) Osmium tetroxide (4% solution, 251755, Sigma-Aldrich) Sodium citrate (W302600, Sigma-Aldrich) Tannic acid (403040, Sigma-Aldrich) Tetrachloroauric acid (G4022, Sigma-Aldrich) Uranyl acetate (Fluka 73943, Sigma-Aldrich)

4.2 Experimental Strategies

Table 4.1 Antibodies that had been used successfully for detecting epitope tags by immuno-electron microscopy Epitope tag

Antibody

Source

EGFP EGFP Histidine (6 )

Mouse monoclonal anti-GFP Rabbit poyclonal anti-GFP Mouse monoclonal anti-His (C-term) (clone 3D5) Mouse monoclonal anti-His (clone 13/45/31-2) Mouse monoclonal antic-myc (clone 9E10)

Dianova (MA1-26343) Alexis (ALX-210-199/1) Invitrogen (R930-25)

Histidine (6 ) myc

Dianova (dia 900) Santa Cruz (sc-40)

4.1.5 Buffers Modified Barth’s saline (MBS): 88 mM NaCl, 1 mM KCl, 0.82 mM MgSO4, 0.33 mM Ca(NO3)2, 0.41 mM CaCl2, 10 mM HEPES, pH 7.5 Calcium-free MBS: MBS without Ca(NO3)2 and CaCl2 Low-salt buffer (LSB): 1 mM KCl, 0.5 mM MgCl2, 10 mM HEPES, pH 7.5

4.1.6 Plasmids pcDNA3.1/myc-His (Invitrogen) pEGFP-C (Clontech)

4.2 EXPERIMENTAL STRATEGIES (FIG. 4.2) The protocol to isolate Xenopus oocytes is well documented in the literature (Maco et al., 2006; Pante, 2006; Sive, Grainger, & Harland, 2010) and includes the removal of a small piece of the ovary from a narcotized female X. laevis. Oocytes are kept in MBS for a maximum of 5 days at 4  C. Oocytes differ in their developmental stage, and for all immuno-EM experiments, we use stage VI oocytes, which are large with a clear separation of the black animal pole and the off-white vegetal pole. For microinjection experiments, oocytes are defolliculated by collagenase treatment (Au et al., 2010; Pante, 2006). For direct immunolabeling of nucleoporins, oocytes are transferred into LSB, and nuclei are isolated manually from the animal pole by popping a small hole with a fine forceps. For labeling of epitope-tagged nucleoporins, the nuclei are isolated after the oocyte has expressed the desired protein, which is achieved by microinjecting the oocyte in their cytoplasm with the appropriate mRNA or plasmid DNA into their nuclei. Nuclei (can then be permeabilized if required) are then incubated with appropriate antibodies (directly conjugated with colloidal gold particles) and processed for fixation, embedding, sectioning, and EM.

87

Surgical removal of a small piece of ovary

Defolliculation of oocytes with collagenase

Manual separation of forceps

Microinjection of plasmid DNA into nuclei of stage VI oocytes Manual isolation of nuclei from animal hemisphere

Incubation at room temperature Manual isolation of nuclei from animal hemisphere

Incubation with antibodies directly conjugated to colloidal gold

Fixation and preparation for EM

c n

FIGURE 4.2 Diagram illustrating the steps for preparing Xenopus oocytes for immuno-labeling of nucleoporins followed by thin-sectioning EM. For details, see Sections 4.4 and 4.5. The electron micrograph shows the typical result of a NE cross section of an isolated Xenopus nucleus labeled with an antibody against the Zn-finger of Nup153 coated with 8-nm colloidal gold particles. The cytoplasm and nucleoplasm are indicated by c and n, respectively. Scale bar, 100 nm.

4.3 Preparation of Antibodies Conjugated with Colloidal Gold Particles

REMARKS Oocytes can be easily handled under a binocular microscope using a 200-ml pipettor with a pipette tip that has been cut at the end to allow undisrupted suction of the oocytes; similarly, isolated nuclei are handled with a 20-ml pipettor with a pipette tip that has been cut at the end (Au et al., 2010; Maco et al., 2006). For all ultrastructural analysis, isolated nuclei and preservation of NPC structure are best at the day of oocyte removal from the Xenopus and we typically carry out the experiments directly at the day of surgery or the next day at the latest.

4.3 PREPARATION OF ANTIBODIES CONJUGATED WITH COLLOIDAL GOLD PARTICLES Typical immunogold-labeling protocols use primary antibodies against the epitope to be localized and commercially available gold-conjugated antibodies. This procedure has the disadvantage that the localization of the gold particles are several nanometers away from the epitope, and in the case of labeling Xenopus oocyte nuclei, it introduces several manipulations of the nuclei, which result in very long protocols and distortion of the NE and NPC structure. Thus, in our protocol, we use primary antibodies directly conjugated with gold particles. Colloidal gold particles are electron dense and readily detectable in the EM. They can be easily coated with antibodies to map nucleoporins within the NPC. The standard sizes for colloidal gold particles we use for immuno-EM are 8–14 nm. For some experiments, two or more different sizes of colloidal gold particles can be used to do multiple labeling in the same oocyte, for example, to localized different domains of a nucleoporin (Fahrenkrog & Aebi, 2003). Colloidal gold particles are commercially available but can also be easily produced in the laboratory through the reduction of tetrachloroauric acid (HAuCl4) with tannic acid (TA) and sodium citrate in the presence of potassium carbonate (KCO3) (Slot & Geuze, 1985) (see detailed protocol in Pante, 2006). Colloidal gold particles have an atomic nucleus, which is surrounded by negatively charged gold chloride anions (AuCl4 ) (Horisberger, 1981). Proteins, such as antibodies, are typically positively charged and can bind to the colloidal gold particles by electrochemical interactions. No cross-linking is necessary so that bound macromolecules typically maintain their biological activity. It is important to add the correct amount of protein molecules that neutralize the negative charge of the gold particles (see below). If not enough molecules are available, the gold particles are electronically unstable and would either precipitate or interact unspecific with other positively charged cellular molecules upon microinjection into the oocyte. On the contrary, an excess of antibodies or transport substrates would compete with the gold-labeled molecules for antigen binding or nuclear transport in the oocyte (Pante, 2006). Colloidal gold particles stabilized by antibodies have a red to dark red color depending on the size and the density of the gold particles. They can be

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stored at 4  C for months. Unstable particles typically precipitated at the bottom of the test tube, they become black, whereas the supernatant is clear. To determine the correct amount of a given antibody that is needed to stabilize the gold particles, one has to carry out a so-called flocculation test (for details see Pante, 2006). The amount of antibody needed is not predictable and not related to the antibody concentration. When antibodies are used repeatedly, the flocculation test has to be carried out for each new batch. Independent of the concentration, we therefore start with a 1:1 dilution of the antibody (in PBS or LSB) and carry out serial dilutions (typically 1:1, 1:2, 1:4, 1:8, 1:16, 1:32, and so on), until the required dilution has been reached. For most antibodies, this is in the range of 1:50 to 1:100, but higher dilutions can be necessary as well the use of undiluted antibody. Dilutions required for colloidal gold conjugation are not related to dilutions of the same antibody used for immunofluorescence. Also antibodies that work well for immunofluorescence do not necessarily work well for EM (in most cases in fact they do not). For our studies on nucleoporin localization, we have typically used custom made antibodies (polyclonal and monoclonal) and tested their suitability for EM directly in the oocytes.

4.4 IMMUNOGOLD LABELING OF NUCLEOPORINS USING ANTI-NUCLEOPORIN ANTIBODIES Nucleoporins are often large, multidomain proteins and are characterized by a complex organization. This complexity necessitates the use of multiple, domain-specific antibodies against an individual nucleoporin to precisely map its localization in the NPC. The use of domain-specific antibodies in Xenopus oocyte nuclei has notably revealed the precise topology of several multidomain nucleoporins, such as Nup153, Nup214, Nup98, and the Nup62 complex (Chatel et al., 2012; Paulillo et al., 2005; Schwarz-Herion et al., 2007). A typical experimental strategy for immunogold labeling of Xenopus oocyte nuclei using antibodies against nucleoporins is outlined in Fig. 4.2 (right part), and a representative result (using anti-Nup153 antibodies) is also shown in this figure. The protocol includes the following steps: ISOLATION OF XENOPUS OOCYTE NUCLEI 1. A piece of ovary is surgically removed from a female Xenopus, and placed in a small beaker (50–100 ml) containing MBS. Individual oocytes are carefully transferred with a 200-ml pipettor (adjusted to 50–100 ml) with a cut-off pipette tip into the lit of a 35-mm tissue culture dish filled with LSB. The tip of the pipette is cut at the end so that suction through the pipettor does not disrupt the oocytes. 2. All subsequent manipulations are then carried out using a dissecting microscope. Oocytes are oriented with fine forceps so that their black animal pole, where the nucleus is located, is facing the eyepieces of the microscope.

4.4 Labeling of Nucleoporins Using Anti-Nucleoporin Antibodies

Next, the nucleus is isolated by popping a small hole with a curved Dumont forceps in the animal pole of the oocyte. In LSB, nuclei remain intact and are partially swelling so that they easily pop out of the opened oocyte. Note: The isolation of nuclei can be carried out in LSB at room temperature or in LSB kept on ice and slightly warmed up during isolation. The optimal conditions and the ease of isolation vary from batch to batch of oocytes. 3. Isolated nuclei are transferred with a cut-off pipette tip of a 20-ml pipettor (adjusted to 5–10 ml) into a 35-mm tissue culture dish containing LSB. 10–15 nuclei are collected for each labeling experiment. Notes: – Typically, the released nuclei are cleaned from yolk rapidly in LSB without further manipulation. However, if too much yolk remains attached to the nuclei, they can be cleaned by gently aspirating them up and down into the cut pipette tip of a 20-ml pipettor in LSB. – Isolated nuclei are best at day 1 after oocyte removal from the Xenopus and are typically not isolated later than at day 3 after surgery.

ANTIBODY LABELING 4. (optional) To label nucleoporins located in the luminal domain of the NPC, the oocytes are briefly treated with Triton X-100 (0.1% for 1 min) before the incubation with the antibodies (Mansfeld et al., 2006). 5. Antibody labeling is performed in a double depression glass slide. One depression is filled with the appropriate antibody directly conjugated to colloidal gold diluted in LSB (80 ml of antibody solution is used per depression), and the second depression is filled with LSB containing 0.1% BSA. The nuclei are transferred into the depression containing the antibody and are incubated for 2 h at room temperature. The nuclei are then transferred to the next depression and washed for 5 min in LSB containing 0.1% BSA to block unspecific binding of the antibodies. Note: We place the glass slide in a normal petri dish. A filter paper is positioned in the bottom of the petri dish, and an area of the filter paper is cut out so that the slide depressions are excluded from the filter paper.

FIXATION, EMBEDDING, AND THIN SECTIONING 6. The labeled nuclei are then prefixed in LSB containing 2% glutaraldehyde and 0.3% tannic acid for 1 h at room temperature. For this step, the nuclei are transferred into a depression of a slide, which contains the fixative solution. Note: Tannic acid is always made fresh from a 2% stock solution. 7. After fixation, the nuclei are washed twice for 5 min each in LSB and embedded in a drop of 2% low-melting agarose. The washing steps are performed by transferring the fixed nuclei into depressions of a slide, which contain the washing solution. After the last washing step the nuclei are not transferred

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8.

9.

10.

11.

12. 13.

anymore. To embed them in agarose, the buffer is aspirated from the depression slide as much as possible with a pipettor, and any excess of LSB is carefully removed with a small piece of filter paper. To embed them in agarose, 50–80 ml of agarose is carefully added to the depression slide. After the agarose solidifies, small blocks containing each a nucleus are cut out with a razor blade under the dissecting microscope. The blocks are transferred with a toothpick into a small (4-ml) glass vial containing 750 ml of LSB. Nuclei are postfixed for 1 h in 1% OsO4 by adding 250 ml of a 4% stock solution (1% final concentration) and subsequently washed three times in LSB for 5 min. If necessary, the samples are stored at 4  C overnight and the protocol is continued the next day. Note: After postfixation with OsO4, the nuclei are brown to black and can be easily seen by eye so that manipulations under the dissecting microscope are no longer necessary. A Pasteur pipette is used to remove the OsO4 and the buffer during the washing. The fixed samples are then sequentially dehydrated by placing them in a series of increasing concentrations of ethanol: 50%, 70%, and 90% ethanol for 10 min each, then three times 100% ethanol for 10 min each, and finally 10 min in 100% acetone. Note: The contrast of the samples can be increased if necessary by an additional step of bloc staining. In this case, the 70% ethanol incubation is followed by an incubation step of 70% ethanol containing 2% uranyl acetate for 1 h. After that the protocol is continued with the 90% ethanol incubation. Epon is not soluble in ethanol, and therefore requires the use of a transition solvent, which is typically acetone. The dehydrated samples are next infiltrated with a 1:1 mixture of Epon resin and acetone for 1 h, followed by infiltration with a 2:1 mixture of Epon resin and acetone for 1 h, and finally in fresh pure Epon resin for 3–4 h. Nuclei are next placed in flat molds filled with fresh pure Epon and polymerized in an oven overnight at 60  C. Notes: – If no flat molds are available, samples can also be polymerized in gelatin capsules. – Complete polymerization of the Epon might occasionally take longer than overnight incubation and may be extended as necessary. After polymerization, the specimen block is trimmed using an EM specimen trimmer device so that section through the embedded nucleus can be made. The trimmed block is transferred to an ultramicrotome and ultrathin (50–70 nm thick) sections are cut using a diamond knife. Sections are collected with a small loop and place on parlodion-coated EM copper grids, dried, and stained with 6% uranyl acetate for 1 h, followed by 2% lead citrate for 2 min. The grids are then carefully washed with double distilled water in a jet of water from a small, 50 ml plastic flask, and air-dried by sucking off residual water on the grid with filter paper. The samples are finally examined under a TEM. Note: If a diamond knife is not available, one might use glass knifes instead.

4.5 Labeling of Nucleoporins Using Epitope-Tagged Nucleoporins

IMAGING AND STATISTICAL ANALYSIS To determine the position of a nucleoporin within the NPC and for a statistical analysis of the gold particle distribution, digitally recorded EM micrographs are analyzed using Image J. Alternatively, film recorded EM micrographs are either digitalized or printed, and analyzed using Image J. The position of gold particles associated with NPCs is measured from images of cross sections along an NE and selected NPCs (Fig. 4.3A). The NPCs should have a good structural preservation so that the major architectural elements are clearly visible. First, the overall linear dimensions of the Xenopus NPC in Epon-embedded nuclei are determined, as they may differ from the dimensions seen in tomographic reconstructions due to the different sample preparation used. Next, we determine the horizontal position of each gold particle with respect to the midplane of the NPC, that is, the NPC central plane, and the vertical position along the central axis through the NPC, that is, the NPC eightfold symmetry axis (Fig. 4.3B). The central plane of the NPC is defined as position 0, so positive numbers represent the location of gold particles on the cytoplasmic face of the NPC, and negative numbers represent the location of gold particles on the NPC nuclear side. For each antibody, the position of at least 100 (preferentially 200) gold particles is scored. From these data, histograms are generated to show the overall distribution of gold particles with respect to the two planes of the NPC (Fig. 4.3B) and the average distances and their standard deviations are calculated. For further visualization of the gold particle distribution, diagrams of the NPC are generated were we illustrate the so-called location clouds (Fig. 4.3C). The center of a cloud represents the mean position of the gold particles in the NPC. The radii of the clouds are defined by the standard deviations of the horizontal and vertical distances. In doing so, it is possible to determine the relative position of the epitope recognized by the used antibody within the NPC and with this the location of the nucleoporin domain. This analysis further allows us to determine the position of a given protein domain with respect to other domains of the same nucleoporin or with respect to other nucleoporins.

4.5 IMMUNOGOLD LABELING OF NUCLEOPORINS USING EPITOPE-TAGGED NUCLEOPORINS Xenopus oocytes are an excellent system for ectopically expression of proteins (Markovich, 2008). The general strategy to express proteins of interest in Xenopus oocytes is to microinject either mRNA into the cytoplasm or plasmid DNA into the nucleus of the oocyte. To overcome the limitation of the standard immunogoldlabeling protocol resulting from the high cross-reactivity of many anti-nucleoporin antibodies, caused by the presence of highly antigenic repetitive sequence motifs within many nucleoporins, and the lack of available antibodies for some nucleoporins, we have exploited the power of Xenopus oocyte as an efficient translation system to express epitope-tagged nucleoporins and localize these by immunogold EM. The protocol to localize epitope-tagged nucleoporin has also the advantage that

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Xenopus NPCs N-terminal domain Zinc-finger domain C-terminal domain

FIGURE 4.3 Immunolocalization of the N-terminal domain of Nup153 within the nuclear pore complex. (A) Intact nuclei isolated from Xenopus oocytes were preimmunolabeled with antibodies against the N-terminal domain of Nup153 coupled with 8-nm colloidal gold. Shown are a cross section along a nuclear envelope after Epon embedding and thin sectioning as well as selected examples of gold-labeled NPCs. c, cytoplasm; n, nucleus. Scale bars, 100 nm. (B) Histogram representation of the quantitative analysis of the gold particle distribution associated with NPCs. (C) Schematic representation of the epitope distribution of the three domains of Nup153 by elliptic “location clouds” along with the linear dimensions of Xenopus NPCs in thin sections. Reprinted from Fahrenkrog et al. (2002), Copyright (2002), with permission from Elsevier.

experiments can be designed using double-tagged nucleoporins in the same oocyte to determine the exact location of the N- and C-terminus of a nucleoporin even within a single NPC (Fahrenkrog & Aebi, 2003). A typical experimental strategy to perform immunogold EM of epitope-tagged nucleoporin is outlined in Fig. 4.2 (left part). The protocol includes the following steps:

4.5 Labeling of Nucleoporins Using Epitope-Tagged Nucleoporins

4.5.1 Design of the constructs For epitope tagging of nucleoporins it is possible to either inject mRNA (Pante et al., 2000) or plasmid DNA (Fahrenkrog et al., 2002) into the oocytes. We prefer to use plasmids, as it is easier and more convenient to handle them compared to mRNA. Plasmids should have a cytomegalovirus (CMV) promoter for mammalian expression, such as the pcDNA3.1/myc-His and pEGFP-C plasmids. We typically cloned the nucleoporins with an N-terminal GFP-tag and a C-terminal myc-His-tag, respectively. For double tagging, we subcloned the nucleoporin including the GFP-tag from the pEGFP into the pcDNA3.1/myc-His. Antibodies to detect the epitope tags, which work well for immuno-EM, are listed in Table 4.1. Note: Other epitope tags, such as HA or FLAG, may also be used. However, we found that mouse monolclonal anti-HA antibodies deriving from the clones HA-7 or 12CA5, respectively, were not suitable for immuno-EM detection.

4.5.2 Protocol 1. A piece of ovary from female X. laevis is removed and collected in MBS as described in step 1 of Section 4.4. 2. The oocytes are washed in calcium-free MBS and defolliculated by incubation for 2 h at room temperature while gently rocking with collagenase. For this, oocytes are transferred into a 50-ml Falcon tube containing 10 ml of 5 mg/ml collagenase in calcium-free MBS. 3. Once the oocytes are sufficiently defolliculated, they are washed three times in MBS and transferred to a 100-mm diameter Petri dish containing MBS. Oocytes are kept at 4  C and used within the next 2 days for microinjection of plasmid DNA. 4. Using a dissecting microscope, mature stage VI oocytes are selected for microinjection. These oocytes are large, with good contrast between the black animal hemisphere and the creamy-colored vegetal hemisphere. 5. A Terasaki microwell plate is filled with MBS, and air bubbles in the wells are removed manually with fine forceps. Next individual stage VI oocytes are placed into each well and oriented for nuclear injection, that is, with the animal pole facing the eyepieces of the microscope. Note: To increase the accuracy of nuclear injection, it is possible to centrifuge the oocytes prior to injection. To do so, the microwell plate is covered with a lid, placed into a low-speed centrifuge, and spun for 15 min at 500  g. After centrifugation, the nucleus will be visible through the animal pole, which facilitates the injection. 6. About 10 nl of plasmid DNA (5 ng/ml) is injected into the nucleus of each oocyte (about 20 nuclei/oocytes per sample). One microliter of 1% bromophenol blue is added to 10 ml of the solution to be injected. The bromophenol blue dye aids the visualization of the microinjection. 7. The injected oocytes are then transferred into small glass (4-ml) vials containing MBS and are incubated for 20 h at room temperature.

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8. Next the nuclei (which should be light blue) of the injected oocytes are isolated, immunogold labeling using an antibody against the tag that is directly conjugated to colloidal gold, and processed for embedding EM as described in steps 2-12 of Section 4.4. Note: If the epitope tag used is GFP, isolated nuclei can be visualized under a fluorescence microscope before performing the immunogold labeling.

CONCLUDING REMARKS Xenopus oocytes in combination with well-established immuno-EM protocols provide a powerful system for studying the precise ultrastructural localization of nucleoporins at the level of NPC components. Experiments can be designed with specific antibodies against defined domains in the nucleoporin of interest, or by expressing epitope-tagged nucleoporins and using antibodies against the tag. We highly recommend the use of both methodologies (i.e., immunogold with anti-nucleoporin antibodies, and localization of epitope-tag nucleoporins) to be confident about the topology of any given nucleoporin within the NPC. When combined, these two experimental strategies allow the mapping of the entire nucleoporin within the structure of the NPC. In doing so, EM studies using Xenopus oocytes have significantly contributed to define the molecular architecture of the NPC and to advance the field of nuclear transport. We hope that detailed immuno-EM methods described in this chapter will further contribute to the utility of this system to refine the molecular architecture and function of the NPC.

Acknowledgments This work was supported by grants from the Fonds de la Recherche Scientifique (FNRS) Belgium and the Fonds Jean Brachet to B. F., and the Natural Sciences and Engineering Research Council of Canada, the Canada Foundation for Innovation, and the Michael Smith Foundation for Health Research to N. P.

References Akey, C. W., & Radermacher, M. (1993). Architecture of the Xenopus nuclear pore complex revealed by three-dimensional cryo-electron microscopy. The Journal of Cell Biology, 122(1), 1–19. Allen, T. D., Rutherford, S. A., Murray, S., Drummond, S. P., Goldberg, M. W., & Kiseleva, E. (2008). Scanning electron microscopy of nuclear structure. Methods in Cell Biology, 88, 389–409. http://dx.doi.org/10.1016/S0091-679X(08)00420-2. Au, S., Cohen, S., & Pante, N. (2010). Microinjection of Xenopus laevis oocytes as a system for studying nuclear transport of viruses. Methods, 51(1), 114–120. http://dx.doi.org/ 10.1016/j.ymeth.2010.02.001.

References

Au, S., & Pante, N. (2012). Nuclear transport of baculovirus: Revealing the nuclear pore complex passage. Journal of Structural Biology, 177(1), 90–98. http://dx.doi.org/10.1016/j. jsb.2011.11.006. Ball, J. R., & Ullman, K. S. (2005). Versatility at the nuclear pore complex: Lessons learned from the nucleoporin Nup153. Chromosoma, 114(5), 319–330. Chatel, G., Desai, S. H., Mattheyses, A. L., Powers, M. A., & Fahrenkrog, B. (2012). Domain topology of nucleoporin Nup98 within the nuclear pore complex. Journal of Structural Biology, 177, 81–89. http://dx.doi.org/10.1016/j.jsb.2011.11.004, PII: S1047-8477(11) 00319-4. Cohen, S., & Pante, N. (2005). Pushing the envelope: Microinjection of Minute virus of mice into Xenopus oocytes causes damage to the nuclear envelope. The Journal of General Virology, 86(Pt 12), 3243–3252. http://dx.doi.org/10.1099/vir.0.80967-0. Cronshaw, J. M., Krutchinsky, A. N., Zhang, W., Chait, B. T., & Matunis, M. J. (2002). Proteomic analysis of the mammalian nuclear pore complex. The Journal of Cell Biology, 158(5), 915–927. Dworetzky, S. I., & Feldherr, C. M. (1988). Translocation of RNA-coated gold particles through the nuclear pores of oocytes. The Journal of Cell Biology, 106(3), 575–584. Fahrenkrog, B., & Aebi, U. (2003). The nuclear pore complex: Nucleocytoplasmic transport and beyond. Nature Reviews. Molecular Cell Biology, 4(10), 757–766. Fahrenkrog, B., Maco, B., Fager, A. M., Koser, J., Sauder, U., Ullman, K. S., et al. (2002). Domain-specific antibodies reveal multiple-site topology of Nup153 within the nuclear pore complex. Journal of Structural Biology, 140(1–3), 254–267. Feldherr, C. M. (1969). A comparative study of nucleocytoplasmic interactions. The Journal of Cell Biology, 42(3), 841–845. Feldherr, C. M., Kallenbach, E., & Schultz, N. (1984). Movement of a karyophilic protein through the nuclear pores of oocytes. The Journal of Cell Biology, 99(6), 2216–2222. Frosst, P., Guan, T., Subauste, C., Hahn, K., & Gerace, L. (2002). Tpr is localized within the nuclear basket of the pore complex and has a role in nuclear protein export. The Journal of Cell Biology, 156(4), 617–630. Gorlich, D., Pante, N., Kutay, U., Aebi, U., & Bischoff, F. R. (1996). Identification of different roles for RanGDP and RanGTP in nuclear protein import. The EMBO Journal, 15(20), 5584–5594. Grossman, E., Medalia, O., & Zwerger, M. (2012). Functional architecture of the nuclear pore complex. Annual Review of Biophysics, 41, 557–584. http://dx.doi.org/10.1146/annurevbiophys-050511-102328. Hinshaw, J. E., Carragher, B. O., & Milligan, R. A. (1992). Architecture and design of the nuclear pore complex. Cell, 69(7), 1133–1141. Horisberger, M. (1981). Colloidal gold: A cytochemical marker for light and fluorescent microscopy and for transmission and scanning electron microscopy. Scanning Electron Microscopy, 2, 9–31. Maco, B., Fahrenkrog, B., Huang, N. P., & Aebi, U. (2006). Nuclear pore complex structure and plasticity revealed by electron and atomic force microscopy. Methods in Molecular Biology, 322, 273–288. http://dx.doi.org/10.1007/978-1-59745-000-3_19. Mansfeld, J., Guttinger, S., Hawryluk-Gara, L. A., Pante, N., Mall, M., Galy, V., et al. (2006). The conserved transmembrane nucleoporin NDC1 is required for nuclear pore complex assembly in vertebrate cells. Molecular Cell, 22(1), 93–103. Markovich, D. (2008). Expression cloning and radiotracer uptakes in Xenopus laevis oocytes. Nature Protocols, 3(12), 1975–1980. http://dx.doi.org/10.1038/nprot.2008.151.

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Ori, A., Banterle, N., Iskar, M., Andres-Pons, A., Escher, C., Khanh Bui, H., et al. (2013). Cell type-specific nuclear pores: A case in point for context-dependent stoichiometry of molecular machines. Molecular Systems Biology, 9, 648. http://dx.doi.org/10.1038/ msb.2013.4. Pante, N. (2006). Use of intact Xenopus oocytes in nucleocytoplasmic transport studies. Methods in Molecular Biology, 322, 301–314. http://dx.doi.org/10.1007/978-1-59745000-3_21. Pante, N., & Aebi, U. (1996). Molecular dissection of the nuclear pore complex. Critical Reviews in Biochemistry and Molecular Biology, 31(2), 153–199. Pante´, N., & Aebi, U. (1994). Toward the molecular details of the nuclear pore complex. Journal of Structural Biology, 113(3), 179–189. Pante, N., Bastos, R., McMorrow, I., Burke, B., & Aebi, U. (1994). Interactions and threedimensional localization of a group of nuclear pore complex proteins. The Journal of Cell Biology, 126(3), 603–617. Pante´, N., & Kann, M. (2002). Nuclear pore complex is able to transport macromolecules with diameters of about 39 nm. Molecular Biology of the Cell, 13(2), 425–434. Pante, N., Thomas, F., Aebi, U., Burke, B., & Bastos, R. (2000). Recombinant Nup153 incorporates in vivo into Xenopus oocyte nuclear pore complexes. Journal of Structural Biology, 129(2–3), 306–312. Paulillo, S. M., Phillips, E. M., Koser, J., Sauder, U., Ullman, K. S., Powers, M. A., et al. (2005). Nucleoporin domain topology is linked to the transport status of the nuclear pore complex. Journal of Molecular Biology, 351(4), 784–798. Rabe, B., Vlachou, A., Pante, N., Helenius, A., & Kann, M. (2003). Nuclear import of hepatitis B virus capsids and release of the viral genome. Proceedings of the National Academy of Sciences of the United States of America, 100(17), 9849–9854. http://dx.doi.org/10.1073/ pnas.1730940100. Rollenhagen, C., Muhlhausser, P., Kutay, U., & Pante, N. (2003). Importin beta-depending nuclear import pathways: Role of the adapter proteins in the docking and releasing steps. Molecular Biology of the Cell, 14(5), 2104–2115. http://dx.doi.org/10.1091/mbc.E02-060372. Rollenhagen, C., & Pante, N. (2006). Nuclear import of spliceosomal snRNPs. Canadian Journal of Physiology and Pharmacology, 84(3–4), 367–376. http://dx.doi.org/10.1139/y05-101. Rout, M. P., Aitchison, J. D., Suprapto, A., Hjertaas, K., Zhao, Y., & Chait, B. T. (2000). The yeast nuclear pore complex: Composition, architecture, and transport mechanism. The Journal of Cell Biology, 148(4), 635–651. Schwarz-Herion, K., Maco, B., Sauder, U., & Fahrenkrog, B. (2007). Domain topology of the p62 complex within the 3-D architecture of the nuclear pore complex. Journal of Molecular Biology, 370(4), 796–806. Sive, H. L., Grainger, R. M., & Harland, R. M. (2010). Isolation of Xenopus oocytes. Cold Spring Harbor Protocols, 2010(12), pdb.prot5534. http://dx.doi.org/10.1101/pdb.prot5534. Slot, J. W., & Geuze, H. J. (1985). A new method of preparing gold probes for multiplelabeling cytochemistry. European Journal of Cell Biology, 38(1), 87–93. Stoffler, D., Feja, B., Fahrenkrog, B., Walz, J., Typke, D., & Aebi, U. (2003). Cryo-electron tomography provides novel insights into nuclear pore architecture: Implications for nucleocytoplasmic transport. Journal of Molecular Biology, 328(1), 119–130. Walther, T. C., Fornerod, M., Pickersgill, H., Goldberg, M., Allen, T. D., & Mattaj, I. W. (2001). The nucleoporin Nup153 is required for nuclear pore basket formation, nuclear pore complex anchoring and import of a subset of nuclear proteins. The EMBO Journal, 20(20), 5703–5714.

CHAPTER

Utilizing the Dyn2 Dimerization-Zipper as a Tool to Probe NPC Structure and Function

5

Dirk Flemming, Philipp Stelter, and Ed Hurt Biochemie-Zentrum der Universita¨t Heidelberg (BZH), Im Neuenheimer Feld, Heidelberg, Germany

CHAPTER OUTLINE Introduction ............................................................................................................ 100 5.1 Common Preparatory Steps............................................................................... 101 5.1.1 Fusion of eDID to a Target Protein ................................................... 101 5.1.1.1 Design of eDID ........................................................................ 101 5.1.1.2 Cloning of eDID to the N- or C-terminal End of a Target Protein................................................................................ 102 5.1.1.3 Integrating the eDID Sequence Within a Yeast ORF .................. 103 5.1.1.4 Expression and Purification of Proteins Modified with eDID ..........................................................................................104 5.1.2 Purification and Handling of Recombinant (e)DID ............................. 105 5.1.3 Purification of Recombinant Dyn2 ................................................... 106 5.2 Dyn2 as an EM Label to Map Subcomplexes and Single Nups ............................. 107 5.2.1 Purification Protocol for Labeled Particles ........................................ 108 5.2.2 Negative Stain EM Analysis............................................................. 109 5.2.2.1 Grid Preparation and Negative Staining .................................... 109 5.2.2.2 Basic Image Processing........................................................... 111 5.3 Probing the FG Network by Selective Insertion of the eDID–Dyn2 Complex into FG Repeat Domains ...................................................................... 111 5.4 Materials and Reagents.................................................................................... 113 5.4.1 Plasmids ....................................................................................... 113 5.4.2 Strains .......................................................................................... 113 5.4.3 Materials and Reagents................................................................... 113 Conclusion ............................................................................................................. 114 Acknowledgment..................................................................................................... 114 References ............................................................................................................. 114

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00005-9

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CHAPTER 5 Using the DID-Dyn2 System to Study NPC

Abstract The discovery of dynein light chain 2 (Dyn2) as a member of the nucleoporins in yeast led to a series of applications to study NPC structure and function. Its intriguing ability to act as a hub for the parallel dimerization of two short amino acid sequence motifs (DID) prompted us to utilize it as a tool for probing nucleocytoplasmic transport in vivo. Further, the distinct structure of the Dyn2–DID rod, which is easily visible in the electron microscope, allowed us to develop a precise structural label on proteins or protein complexes. This label was used to identify the position of subunits in NPC subcomplexes or to derive at pseudo-atomic models of single large Nups. The versatility for various applications of the DID–Dyn2 system makes it an attractive molecular tool beyond the nuclear pore and transport field.

INTRODUCTION The size of the nuclear pore complex (NPC) with a molecular mass of
120 MDa in metazoan and
50 MDa in yeast makes it a formidable task to study its structure and function. Although the inventory of the components building up an NPC is rather complete, new nucleoporins (Nups) are occasionally still discovered (Chadrin et al., 2010; Stelter et al., 2007). Such a case was the identification of the dynein light chain (Dyn2) as a member of the Nup family in Saccharomyces cerevisiae. Dyn2 might have been previously unnoticed as a nucleoporin because of its small size of
10 kDa till Stelter et al. discovered it co-eluting with an oligomeric form of the Nup82 complex. Dyn2 is known from its role in the dynein motor complex where it interacts with the dynein heavy chain (Dyn1) and the intermediate chain (Pac11) to perform various functions within the cell, including spindle orientation and nuclear migration. Nevertheless, a huge amount (25%) of the Dyn2 cytoplasmic pool of yeast was found associated with the NPCs (Romes, Tripathy, & Slep, 2012). However, when yeast cell are depleted of Dyn2, growth is not significantly affected. Like its human ortholog DLC1 (identity 47%), Dyn2 forms predominantly a stable homodimer with two opposed binding grooves for binding motifs. Hereby a platform is created, which can induce and stabilize the parallel dimerization of two target sites. In higher eukaryotes, these protein targets typically contain a consensus motif that is a sequence of 12 residues in length (Navarro-Lerida et al., 2004). Nup159 harbors in its primary sequence six of the recognition motifs for a Dyn2 interaction, separated by short linkers, between its FG (phenylalanine–glycine) repeats and the C-terminal coiled-coil domain (see Fig. 5.1). Under physiological conditions, Nup159 recruits Dyn2 and helps to dimerize the Nup82–Nsp1– Nup159 complex located at the cytoplasmic pore filaments. The DID motif of Nup159 contains variations of the most common consensus K/RXTQT motif (Lo, Naisbitt, Fan, Sheng, & Zhang, 2001). NMR experiments showed that this sequence (called DID, for dynein light-chaininteraction domain) is intrinsically disordered and that binding of one Dyn2 dimer

5.1 Common Preparatory Steps

FIGURE 5.1 Domain organization of Nup159 with the primary sequence of the DID (dynein light-chaininteracting domain). Two DIDs dimerize upon addition of Dyn2. The resulting rodlike structure can be visualized by negative stain electron microscopy.

promotes the binding of the subsequent (Nyarko, Song, & Barbar, 2012). In the DID of Nup159, only five of the six predicted motifs bind Dyn2. In vitro reconstitution of the Dyn2–DID complex and negative stain electron microscopy (EM) analysis revealed a readily visible, prominent 20-nm rod-like structure in which five Dyn2 homodimers were aligned between two extended DID strands (Fig. 5.1). The unique properties and direct visibility also made it a precise and versatile tool for mapping subunits of the Nup84 complex (Flemming, Thierbach, Stelter, Bottcher, & Hurt, 2010) as well as for deciphering a pseudo-atomic model of Nup188 and Nup192 by EM (Amlacher et al., 2011). The ability of the nucleoporin Dyn2 to bind to varying recognition motifs (here used: DID and Pac11) serving as a molecular glue prompted us to use it as a tool to specifically block parts of the FG transport channel and thereby probing the transport mechanisms (Stelter, Kunze, Fischer, & Hurt, 2011).

5.1 COMMON PREPARATORY STEPS 5.1.1 Fusion of eDID to a target protein 5.1.1.1 Design of eDID Several binding partners have different recognition motifs in the region that binds Dyn2. The binding sites display a highly conserved QT sequence motif. For example, Pac11 contains two tandem, cooperatively working Dyn2-binding sites (Rao et al., 2013). As a longer, alternative binding platform for Dyn2, an artificially engineered DID was created from the binding motif of Pac11 (dynein intermediate chain) via PCR modification techniques. By joining three tandem QT 11-mer peptides (residues 46–57 and 76–87), a binding motif with six consecutive Dyn2 was created. This DID was termed DID1 or eDID (here further called eDID, e stands for engineered; Fig. 5.2) and showed by EM to bind six Dyn2 dimers.

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FIGURE 5.2 Nucleotide and amino acid sequence of the artificially designed eDID with six Dyn2 binding motifs. Annealing sequences for primers to amplify are shown in italic.

REMARKS Shortening the eDID motif to three binding domains was performed and resulted in a rod exhibiting three Dyn2 dimers as expected (unpublished data). A further elongation to more than eight domains caused a precipitation of the protein upon binding of Dyn2 and is therefore not applicable.

5.1.1.2 Cloning of eDID to the N- or C-terminal end of a target protein The eDID construct will next be fused to the protein of interest, that is, the protein to be labeled or dimerized. For EM-labeling experiments, it is recommended to create constructs bearing the DID at either the N- or the C-terminal end. The position of the affinity tag does not interfere with the function of the Dyn2–DID interaction. In multisubunit complexes, different subunits can bear different tags for tandem affinity purifications (see Fig. 5.3). For labeling approaches, no linker should be between the protein and the DID sequence to avoid unnecessary flexibility. Plasmids containing the sequence of the eDID (e.g., pFa6a-eDID-Flag-PLox) for further subcloning are available upon request in the lab of Ed Hurt, University of Heidelberg. For the PCR amplification of the eDID sequence, primers with the appropriate overhang should be designed. The overhang should contain recognition sites for restriction enzymes for a subsequent cloning to the target protein: forward primer: 50 -overhang-ACATACGACAAAGGCATTCAAACAGATC AAATTG-30 reverse primer: 50 -overhang-CGTTCCTTCTTCCATATCGGTTTGCACC-30 Since the eDID is rather short (237 nucleotides), a short elongation time for the PCR reaction can be chosen (98), 3 mg/ml L-a-phosphatidylserine (sodium salt, brain, porcine), 3 mg/ml L-a-phosphatidylinositol (sodium salt, liver, bovine), 6 mg/ml L-a-phosphatidylethanolamine (egg, chicken), 15 mg/ml L-aphosphatidylcholine (egg, chicken; all from Avanti Polar Lipids, USA) in 10% n-octyl-b-D-glucopyranoside DiIC18: 1 mg/ml 1,10 -dioctadecyl-3,3,30 ,30 -tetramethylindocarbocyanine perchlorate in DMSO (“DiI,” DiIC18(3), crystalline; Life Technologies GmbH, Germany) Sucrose buffer as in Section 9.1.1 and PBS as in Section 9.3.1

9.3.2.3 Method 1. Swell G50 beads in sucrose buffer for 10 min at room temperature. 2. Remove bottom and top lids from the chromatography column and fill column with swollen G50 beads. Let beads settle by gravity. Add more beads carefully until the glass cylinder of the column is nearly filled (leave about 0.5 cm space at the top). The upper edge of the bead layer should remain visible. Note: Avoid air bubbles in the column. Be careful that columns do not run dry. 3. Fill column with sucrose buffer, put top lid on, and connect the lid to a buffer reservoir via tubing. 4. Wash column with sucrose buffer from the reservoir by letting the buffer pass by gravity flow. 5. To block the column, take top lid off, carefully remove excess buffer and add 100 ml of 1 mg/ml BSA in sucrose buffer directly onto the bead layer. Let the solution enter the beads, then fill the column immediately but carefully with buffer, put top lid back on, and wash the column as in step 4. 6. Block column again with 20 ml lipid mix diluted 1:5 with PBS in 100 ml total volume as in step 5. Note: Lipid mix can be altered as well as the detergent used to dissolve the lipids. A relatively high-CMC (critical micelle concentration) value of the detergent and small aggregation number is important to ensure its removal by gel filtration. 7. Wash column with sucrose buffer for 30 min as in step 4. 8. Close bottom lid of the column. To store your columns for longer term, add 0.1% NaN3 to the sucrose buffer. Note: For long-term storage of the columns, let some buffer pass from time to time.

9.3 Biochemical Procedures

9.3.3 Depletion of transmembrane proteins from Xenopus membranes To specifically deplete transmembrane proteins, Xenopus membranes are solubilized by detergent and the protein of interest is immunodepleted by passage of the solubilized membrane fraction over an antibody column (Fig. 9.2A). Detergent removal from the solubilized and depleted membrane fraction reconstitutes the membranes. An efficient way for detergent removal is passage of a gel filtration column (Allen, Romans, Kercret, & Segrest, 1980) under the precondition that the detergent has a relative high-CMC value, which defines the concentration of the free detergent in solution in contrast to its micellar form, and a not small aggregation number. For add-back experiments, the purified integral membrane protein (see Section 9.2.1) is added to the solubilized and depleted membrane fraction and coreconstituted by passage of the gel filtration column.

9.3.3.1 Materials and equipment • • • • •

n-Octyl-b-D-glucopyranoside or equivalent detergent as in Section 9.3.2 Beckman Optima TLX Ultracentrifuge, TLA100 rotor and tubes (or equivalent system) Mobicol columns (Mobicol “classic” with 1 closed screw cap and plug and 35 mm pore size filters, MoBiTec GmbH, Germany) Cooled tabletop microcentrifuge Crude membranes, antibody beads, and G50-chromatography columns described in Sections 9.1.2, 9.3.1, and 9.3.2

9.3.3.2 Buffers and solutions •

Sucrose buffer, lipid mix, and DiIC18 as in Sections 9.1.1 and 9.3.2.

9.3.3.3 Method

1. Solubilize 20 ml of crude membranes (for preparation, see Section 9.1.2) in sucrose buffer with 1% n-octyl-b-D-glucopyranoside for 10 min at 4  C. Notes: Frozen membrane aliquots can be used. Ensure proper membrane solubilization with marker proteins by western blotting. If necessary other detergents (with relative high-CMC values and small aggregation numbers to allow for their removal afterwards) might be used such as CHAPS. Use high-quality detergent, some batches of n-octyl-b-D-glucopyranoside need to be purified before use by passage over a mixed bead resin (e.g., AG501X8 from Bio-Rad, Germany). 2. Clear by centrifugation for 10 min at 200,000
g and 4  C in a TLA100 rotor and take supernatant. 3. Equilibrate 40 ml of antibody beads (50% slurry, prepared as described in Section 9.3.1) in a Mobicol column with sucrose buffer immediately before use, spin dry by centrifugation for 30 s at 5000
g and 4  C in a cooled tabletop

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A

B k oc

M

ck ck ba ba ck d d o Ad Ad M

Membrane

C k oc

M

ac

k

b dd

A

Membrane

FIGURE 9.2 Depletion and Functional Add-Back of Membrane Proteins. (A) Schematic illustration of immunodepletion of transmembrane proteins from Xenopus membranes. Membranes are solubilized with a detergent and the target membrane protein is depleted from the solubilized membrane fraction by specific antibody beads. Finally, the depleted membranes are reconstituted by detergent removal. (B) Western blot analysis of Xenopus membranes, which were mock-depleted, immunodepleted of endogenous NDC1 or NDC1-depleted and substituted with recombinant EGFP–NDC1 expressed in E. coli (addback). Endogenous and recombinant NDC1 are detected by NDC1-antibodies, the recombinant protein also by EGFP-antibodies. The NDC1 depletion does not affect levels of other integral membrane proteins including the transmembrane nucleoporins GP210 and POM121. (C) Immunofluorescence of nuclear assembly reactions with Xenopus cytosol and membranes from (B). Recombinant EGFP–NDC1 is faithfully integrated into the nuclear membranes as detected by NDC1- and EGFP-antibodies. NDC1 depletion blocks NPC formation (DNDC1). This phenotype is rescued by the addition of recombinant EGFP–NDC1 (addback) as indicated by the presence of Nup58, a nucleoporin located in the inner part of the NPC. DNA is stained with DAPI. Bar: 10 mm.

9.3 Biochemical Procedures

4. 5.

6.

7. 8.

microcentrifuge. Apply supernatant of step 2 to dried beads and incubate for 30 min at 4  C on a rotating wheel. Note: Optimal bead to solubilized membranes ratio needs to be determined. However, the given conditions work for most of the proteins we tested. Elute unbound supernatant (spin as in step 3) and incubate a second time as in step 3 with fresh antibody beads. Elute unbound supernatant and add 20 ml lipid mix and 0.2 ml of 1 mg/ml DiIC18 in DMSO. For add-back experiments, add your protein of interest in approximately endogenous concentration together with the lipid mix to the eluate. Notes: The optimal amount of readded protein needs to be determined. In our hands, for most add-back attempts achieving endogenous protein levels works fine (Fig. 9.2B). If necessary one can avoid the use of a fluorescent dye and reconstituted membranes are collected blindly. For this, a test run in which one can follow the reconstituted fraction with a marker (e.g., a fluorescent dye) is performed and the number of drops counted until the reconstituted membrane fraction runs out of the column. Reconstitute membranes by detergent removal by passing the sample over a G50-chromatography column (prepared as described in Section 9.3.2) equilibrated to sucrose buffer. Remove excess of buffer and load the samples directly on the bead layer. Fill the column with buffer as soon as the sample has entered the beads. Collect the membrane-containing fraction (400 ml or 8–9 drops), which appears pink due to addition of DiIC18. Pellet reconstituted membranes by centrifugation for 30 min at 200,000
g in a TLA100 rotor. Resuspend membrane pellet in 20 ml sucrose buffer. Reconstituted membranes are now ready to use in the nuclear assembly reaction (Section 9.4.1).

9.3.4 Reconstitution of recombinant integral membrane proteins in proteoliposomes For purification, integral membrane proteins are detergent solubilized from membranes. At the end of the purification process, the detergent is removed to maintain the proper functionality of the integral membrane protein by reconstitution in proteoliposomes, which can be used as a tool for functional studies as in Section 9.4.

9.3.4.1 Materials and equipment • •

Beckman Optima TLX Ultracentrifuge, TLA120.2 rotor and tubes (or equivalent system) G50-chromatography columns described in Section 9.3.2

9.3.4.2 Buffers and solutions •

PBS, lipid mix, DiIC18 and sucrose buffer as in Sections 9.1.1, 9.3.1, and 9.3.2

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9.3.4.3 Method 1. Prepare preblocked G50-columns as described in Section 9.3.2 but use PBS instead of sucrose buffer. 2. Apply a mix consisting of 20 ml lipid mix, 5 ml purified protein (conc. 40 h (Rabut et al., 2004), cells need to be incubated at least for 2 days after transfection to allow efficient incorporation of the tagged Nups into the NPC. For the dynamic Nups such as Nup153 and Nup50, 1 day is sufficient for their expression and incorporation into the NPC. Making stable cell lines is recommended especially when the cell line you use is hard to transfect and the fraction of cells with an adequate expression level is low. To boost signal further and control for the functionality of the tagged Nups, the untagged

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endogenous protein can be depleted simultaneously by RNAi, which typically increases the fraction of fluorescent Nups and therefore the specific signal in the NPC (the detailed protocol is described in Section 10.3) (Szymborska et al., 2013). Alternatively, endogenous genes can be replaced with tagged ones by using targeted genome editing techniques that have been developed recently (reviewed in van der Oost, 2013). On the day of imaging, the medium is replaced with prewarmed CO2-independent medium without phenol red supplemented with fetal calf serum (FCS), glutamine, penicillin and streptomycin, and 0.2 mg/ml Hoechst 33342, and imaging is performed at 37  C in a microscope body-enclosing incubator.

10.1.2 Live imaging of NPC assembly Since postmitotic assembly of a given Nup occurs typically within
4 min, the process has to be sampled with a temporal resolution of 1 min or better. We typically start recording cells in late metaphase, to reliably catch anaphase onset and then sample 3–5 optical sections every 30 s or for detailed kinetic analysis single-optical sections every 6–12 s (Fig. 10.1A). Mitotic cells, especially in the relatively longlived metaphase stage, are easily found in a log phase growing cell population and thus cell cycle synchronization is often dispensable. The population of mitotic cells can be increased by G1/S arrest and release using Thymidine for HeLa cells or Aphidicolin for normal rat kidney (NRK) cells if necessary (Dultz et al., 2008; Mall et al., 2012). You should avoid arresting cells in mitosis using Nocodazole or MG132 because these treatments interfere with the normal cell division process close to the observation time and are therefore prone to cause artifacts. Figure 10.1A shows a typical result of triple color time-lapse imaging of dividing cells. Nup assembly around chromosomes (Hoechst signal) and the accumulation of import substrate (IBB-DiHcRed; importin b-binding domain of importin a; Gorlich, Henklein, Laskey, & Hartmann, 1996) fused with a tandem repeat of HcRed in the nuclei are monitored. The different spectral channels should be recorded simultaneously since cell division is a highly dynamic process and a time delay between the acquisition of each channel causes spatial and temporal offsets. When simultaneous recording is not possible due to cross-talk of fluorescence between channels, a confocal microscope system which sequentially switches the excitation wavelength during laser scanning line-by-line to allow pseudo-simultaneous interleaved recording can be advantageous and makes it, for example, possible to separate the GFP channel from the CFP channel (Fig. 10.1C). We performed our imaging on Zeiss LSM 780, equipped with 405, 458, 488, 514, and 561 nm lasers and Zeiss 63  1.4 NA Plan Apochromat objective. For four color imaging (Fig. 10.1C), dichroic beamsplitters MBS 405 and T90/R10 were used and bandpass filters were adjusted to 415–447, 464–490, 499–551, and 616–673 nm. For triple color imaging (Fig. 10.1A), MBS 405 and 488/561 were used and bandpass filters were adjusted to typically 415–447, 499–551, and 603–673 nm.

FIGURE 10.1 Kinetic Analysis of the Postmitotic NPC Assembly in Living HeLa Cells (A and C) The mitosis of HeLa cells expressing IBB-DiHcRed and EGFPx3–Nup107 (A) or Pom121–EGFPx3 and Srprb-ECFP (C) was monitored by confocal microscopy. Time-lapse sequences of single 3-mm confocal slices were recorded every 12 s with a pixel size of 0.13 mm. Time after anaphase onset is indicated. Images were filtered with a median filter (kernel size: 2 pixels) for presentation purposes. (B and D) Mean intensities of IBB in the nucleus and Nups at the nuclear rim were quantified. The total intensity of Nup107 (B) and the mean intensity of ER-subtracted Pom121 (D) at the nuclear rim are also shown. The quantification results of four (B) and five (D) different cells were normalized to the lower and upper plateau, aligned to the time point of anaphase onset, and averaged. Error bars show the standard deviation. (E and F) Summary of NPC assembly kinetics in HeLa cells (E), and in NRK cells (F). EGFPx3–Nup133, EGFP–Nup98, EGFPx3–Nup153, and EGFPx3–Nup214 were expressed and their assembly kinetics were analyzed as in (A and B) (E). The number of analyzed cells is indicated. For HeLa cells, the data of ER-subtracted Pom121 are shown (Pom121*). For NRK cells, t1/2 of IBB is used to temporally align the assembly time series. Modified from Dultz et al. (2008).

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10.1.3 Computational image analysis and quantitation of assembly kinetics Nups are distributed throughout the cell in the cytoplasm or endomembranes at metaphase and accumulate at the nuclear rim after anaphase onset (Fig. 10.1A). High-resolution microscopy has demonstrated that Nups appear as discrete patches around the dividing chromosomes, suggesting that the binding of Nups to the chromosome represents the formation of NPCs rather than a general coating of the chromosome (Dultz et al., 2008; Lu, Ladinsky, & Kirchhausen, 2011). In order to quantify the assembly kinetics of Nups as well as the functional state of the assembling NPCs by nuclear import of IBB, the signal intensities of Nups in the NE and IBB in the nucleus are measured. The regions of the NE and the nucleus can be segmented using the chromatin (Hoechst/H2B-FP) signal as a reference in Image J (http://rsb.info.nih.gov/ij/). Briefly, a mask of the nucleus is generated by filtering the chromatin images with a median filter followed by binarization. The mask of the nuclear periphery is obtained by eroding and dilating the binary images and then subtracting the eroded from the dilated image. You can measure either the mean or the total intensity, which gives you information about the NPC density and number, respectively. It is worth noting, that after the characteristic intensity increase due to NPC assembly, the mean intensity of many Nups, for example, Nup107 (Fig. 10.1B, green line), decreases again at later times after anaphase onset. This is due to chromatin decondensation and nuclear surface growth during telophase that causes a decrease in the NPC density (Mora-Bermudez, Gerlich, & Ellenberg, 2007). Meanwhile, the total intensity of Nup107 stays constant, reflecting the fact that most postmitotic NPCs have been formed and have incorporated the Nup (Fig. 10.1B, light green line). Another interesting feature to note is the behavior of integral membrane Nups that in mitosis reside in the endoplasmic reticulum (ER), such as Pom121 (Fig. 10.1C, 0 min, shown in green). The intensity of Pom121 at the nuclear rim increases 2–5 min after anaphase onset although no enrichment of the protein around nuclei is observed visually (Fig. 10.1C and D, green line). This increase of signal at the nuclear periphery is caused by the movement of the segregating chromosomes, which brings them so close to the ER membranes that their signal is picked up by the dilated nuclear periphery measurement region, although an NPC-specific Pom121 enrichment has not yet started (Fig. 10.1C). In order to correct for the presence of the ER population of Pom121 and only quantify the Pom121 which newly assembles into the NPC, we coexpress an ER-localizing protein, for example, Srprb (signal recognition particle receptor, beta subunit) fused to an FP in a third color (e.g., ECFP) and perform four color time-lapse imaging (Fig. 10.1C). The ER population of Pom121 can then be masked during analysis in Image J as follows: (i) background is subtracted from each channel, (ii) the total intensities of Pom121 and Srprb at metaphase are normalized, and (iii) the images of Srprb are subtracted from the Pom121 images before quantitation of the Pom121 channel (Fig. 10.1C and D, gold line).

10.2 Monitoring NE Permeability

After such corrections, the assembly kinetics of different Nups can be compared by aligning their time series to the time of anaphase onset or the time of the half maximal IBB intensity increase (t1/2) in the nucleus (Fig. 10.1E and F). Either way of temporal alignment gives a similar overall result but the alignment to IBB t1/2 typically yields lower errors, as this temporal reference is closer to the time of assembly of most Nups. The general order of the Nup assembly in HeLa cells is very similar to the one we previously reported in NRK cells (Dultz et al., 2008) except for Pom121 (Fig. 10.1E and F). This is because the ER population of Pom121 was not subtracted at the time the NRK cell data was generated. Although strong overexpression of Nups has been reported to alter the timing of import competence of assembling NPCs (Anderson, Vargas, Hsiao, & Hetzer, 2009), the moderate amount of Nups expression used here does not significantly affect the import kinetics of IBB.

10.2 MONITORING NE PERMEABILITY IN LIVING CELLS BY SEQUENTIAL PHOTOSWITCHING Dronpa is a photoswitchable FP cloned from Pectiniidae corals (Ando, Mizuno, & Miyawaki, 2004) and can be reversibly photobleached and photoactivated over 50 times by
490 and
400 nm light, respectively, without significant loss of fluorescence (Habuchi et al., 2005). Like most monomeric FPs with GFP fold, Dronpa has a molecular weight of 28.8 kDa and is small enough to pass through the NPC by passive diffusion. By sequential photoswitching of Dronpa and monitoring its flux from cytoplasm to the nucleus, the permeability of the NE can be monitored in single-living cells at different cell cycle stages, allowing for repeated measurements independent of cell-to-cell variation.

10.2.1 Sample preparation A plasmid encoding Dronpa is introduced into HeLa cells with the transfection reagent Fugene6 24–48 h before imaging. We use cells expressing IBB-DiHcRed because the IBB signal allows us to segment the region of the nucleus and quantify the nuclear translocation of Dronpa. Other nuclear proteins such as histones fused to FPs spectrally distinguishable from Dronpa can also be used as a nuclear marker.

10.2.2 Measuring nuclear permeability by sequential photoswitching Mitotic cells expressing Dronpa are imaged and the timing of anaphase onset is recorded. Around 15 min after anaphase onset, Dronpa fluorescence in a whole cell is switched off by illuminating entire field of view with
490 nm light. Next, Dronpa

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is photoactivated by a short pulse of
400 nm light in a small region of the cytoplasm (Fig. 10.2A). Photobleaching can be achieved by illumination with a 514 nm or 488 nm argon-ion laser or a mercury lamp via a yellow FP filter (BP 500/20 FT 515 BP535/50) for few seconds. Independent of the precise wavelength, illumination should be stopped immediately after sufficient photobleaching is achieved to avoid undesired phototoxic effects and potential cell cycle delay/arrest of the observed cells. For photoactivation, a 405 nm blue diode laser is used. Again, the light dose should be as low as possible while still providing sufficient contrast immediately after photoactivation. Following photoactivation of a pool of Dronpa in the cytoplasm, the nuclear accumulation of activated Dronpa is monitored by time-lapse imaging with low dose of 488 nm laser light. This photoswitching experiment is then repeated several times in the same cell as the cell cycle progresses, to monitor changes in permeability. As shown in Fig. 10.2A and C, Dronpa translocation to the nucleoplasm reaches a steady state within 6, 25, 75, and 150 s in a cell 17, 25, 35, and 45 min after anaphase onset, respectively. The frame rates of the time-lapse sequence should be higher for cells early after anaphase onset since Dronpa rapidly enters the nucleoplasm at this time. Typically, images are recorded every
0.75,
1.0, and
1.5 s for cells
15,
25, and
35 min after anaphase onset, respectively, and
2.0 s for the later time points. It should be noted that with properly adjusted light dose, no deleterious effects of the repetitive photoswitching on cell cycle progression are observed (Dultz et al., 2009).

10.2.3 Computational image analysis and extraction of equilibrium rate constants The fluorescence of Dronpa decreases to some extent during time-lapse image acquisition since the 488 nm light used to excite Dronpa also photobleaches it. In our experimental conditions, we adjust the imaging intensity to a sufficiently low level, so that the fluorescence is bleached by no more than 15–30%. Since this loss is still significant and could distort the kinetic analysis of the translocation, photobleaching should be corrected as follows: (i) subtract background intensity from mean intensities of Dronpa in the nucleus and whole cell, respectively, and (ii) divide the nuclear intensity by the whole cell intensity. The IBB signal is used for the segmentation of the nucleus. The first image just after photoactivation (Fig. 10.2A, 0 s) is excluded from the analysis since it takes 1–2 s for Dronpa to equilibrate throughout the cytoplasm, as is the case for photoactivatable GFP (Beaudouin, Mora-Bermudez, Klee, Daigle, & Ellenberg, 2006). By fitting the data with a model for the NE permeability (described below), the equilibration rate constant of Dronpa between the cytoplasm and the nucleoplasm can be measured (Fig. 10.2B–D). Assuming that the translocation of Dronpa into the nucleoplasm is governed by first-order kinetics, the translocation can be described as d½Nuc=dt ¼ kout ½Nuc þ kin ½Cyto,

ð10:1Þ

FIGURE 10.2 Kinetic Analysis of the Nuclear Envelope Permeability in Living HeLa Cells (A) Nuclear envelope permeability was repeatedly measured in the same HeLa cell at different time points (17, 25, 35, and 45 min) after anaphase onset. Dronpa molecules in the regions indicated by white circles were photoactivated by a 405-nm laser and their translocation into the nucleus was recorded by confocal microscopy with a confocal slice of 3 mm. Time after photoactivation is indicated above images. Transmitted light images combined with IBB-DiHcRed channels are also shown. Images were filtered with a median filter (kernel size: 2 pixels) for presentation purposes. (B) Schematic representation of the rate constants used in the model for NE permeability. (C) Mean intensities of Dronpa in the nucleus were quantified, corrected for photobleaching, normalized to an upper plateau, and fitted with single exponential functions. (D and E) The measured equilibration rates are plotted on a log scale with base of 10 along the time points of anaphase onset. Data from HeLa cells (D) and NRK cells (E). The data points of the single cell shown in (A and B) are highlighted in red (D). The number of analyzed cells is indicated. AO, anaphase onset. (E) Modified from Dultz et al. (2009), shown with permission from Elsevier# is shown.

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where [Nuc] and [Cyto] are the concentrations of Dronpa in the nucleoplasm and the cytoplasm, respectively, and kin and kout are the inward and outward translocation rate constants (Fig. 10.2B). Since the number of total Dronpa molecules is constant (ND) in the cell, ½NucVNuc þ ½CytoVCyto ¼ ND ,

ð10:2Þ

where VNuc and VCyto are the volume of the nucleoplasm and the cytoplasm, respectively. Substitution of Eq. (10.2) into Eq. (10.1) yields
d½Nuc=dt ¼ ND kin =VCyto  kout þ kin VNuc =VCyto ½Nuc: ð10:3Þ From this differential equation, we obtain

½Nuc ¼ ½Nuc1  ½Nuc1  ½Nuc0 exp  kout þ kin VNuc =VCyto t ,

ð10:4Þ

where [Nuc]1 and [Nuc]0 are the final and initial concentrations of Dronpa in the nucleoplasm, respectively. Eq. (10.4) can be converted as


½Nuc  ½Nuc0 = ½Nuc1  ½Nuc0 ¼ 1  exp  kout þ kin VNuc =VCyto t : ð10:5Þ kout þ kinVNuc/VCyto is defined as the equilibration rate constant. As shown in Fig. 10.2C, the accumulation curves of Dronpa into the nucleoplasm are well fitted to Eq. (10.5). The data fitting is done in MATLAB (The MathWorks, Inc) and the equilibration rates are determined in single cells at different time points after anaphase onset (Fig. 10.2D). The result shows that in HeLa cells, very similar to our previous measurements in NRK cells (Dultz et al., 2009), the NE barrier is established about 1 h after the assembly of import competent NPCs has been completed (Fig. 10.2D and E).

10.3 STRUCTURAL ANALYSIS OF THE NPC BY SUPER-RESOLUTION MICROSCOPY The advances in the field of SR light microscopy brought the resolution achievable in biological samples down to approximately 20 nm, allowing for observation of cellular structures at unprecedented detail. One approach to SR imaging, comprising a number of techniques such as stochastic optical reconstruction microscopy (STORM), photoactivated localization microscopy (PALM), and ground-state depletion followed by individual molecule return (GSDIM), relies on precise localization of single-fluorophore molecules cycling between fluorescent and nonfluorescent states (reviewed in Bates, Jones, & Zhuang, 2013; Furstenberg & Heilemann, 2013). The SR image of biological structures is reconstructed using the positions of a large number of localized fluorophores which are conjugated to specific cellular macromolecules. Recently, localization microscopy allowed

10.3 Structural Analysis of the NPC by Super-resolution Microscopy

for direct visualization of several different scaffold Nups (Ori et al., 2013; Szymborska et al., 2013) and the luminal part of the pore membrane protein gp210 (Loschberger et al., 2012). All these Nups were observed arranged in a ring around the transport axis of the pore. We combined STORM imaging with a single-particle averaging approach to study the organization of the scaffold of the NPC (Szymborska et al., 2013). With this method we have mapped the average distances of several GFP-tagged components of the Nup107–160 complex from the center of the pore (radial position) with subnanometer precision. Below, we outline the key steps in the sample preparation and imaging procedure and describe the image analysis method used for determination of radial positions of the fluorescent labels. Our protocol was optimized for imaging of the NPC scaffold components in U2-OS cells, but it can easily be adapted for a different set of Nups and a different cell line.

10.3.1 Sample preparation and imaging The most critical aspect to consider during the sample preparation is the size of the reagent carrying the fluorescent label and the density or completeness of labeling of the Nup of interest. Although specific labeling of the endogenous Nups can be achieved using antibodies, the large size of an IgG antibody can potentially offset the fluorescent molecules from the targeted epitope by 7–15 nm, depending on whether a fluorophore-conjugated primary or a primary and secondary antibody pair are used. An alternative approach relies on expression of GFP-tagged Nups and immunostaining with fluorophore-conjugated anti-GFP cameloid antibodies (nanobodies) which are approximately 10 times smaller than an antibody (Ries, Kaplan, Platonova, Eghlidi, & Ewers, 2012). Unless the GFP-tag is introduced directly by genome engineering (van der Oost, 2013) of all alleles encoding the Nup of interest, the density of the NPC labeling with the anti-GFP nanobody is limited by the presence of the untagged protein in the pore, making parts of the structure unrecognizable to the nanobody. To circumvent the laborious procedure of establishing homozygous knock-in cell lines for every Nup of interest, in our experiments we combined transient expression of an siRNA-resistant GFP-tagged Nup with RNAi-mediated depletion of the endogenous protein. We found that high rate of replacement for many Nups can be achieved with a single round of transient transfection of both the GFP construct and the siRNA. However, since many Nups are long lived, it can be beneficial to generate stable Nup–GFP lines and transfect them with siRNAs multiple times over the course of several days. Alternatively, both the GFP-tagged Nup and the siRNA-coding gene can be stably expressed under a bidirectional promoter (Ori et al., 2013). The Nup of interest can be tagged at either N- or C-terminus with a green FP such as mEGFP. Although most GFP-tagged Nups are functional (Rabut et al., 2004), it is important to verify that a new fusion protein localizes correctly and complements an RNAi phenotype of the endogenous protein, if present.

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Pre-designed or custom-made siRNAs against the Nups’ coding regions are available from several manufactures. The cDNA is made siRNA-resistant by introducing 3 or 4 silent mutations towards the 30 -end of the siRNA-target sequence. The potency of the siRNA and the resistance of the tagged Nup to knock-down can be verified by Western blotting (if an appropriate antibody is available) or quantitative PCR. In our experiments, we introduce the siRNA into the cells by solidphase transfection on siRNA-coated plates, because it causes less cell death and is more convenient than liquid phase transfection. A detailed protocol for siRNA coating is available elsewhere (Erfle et al., 2008). The siRNA-coated plates can be prepared ahead of time and stored for months in sealed boxes with drying pearls. The second factor influencing the quality of the NPC labeling is the choice of the fluorescent dye and the degree of conjugation of the nanobody to the fluorophore. Although many commercial fluorophores exhibit switching behavior, the Alexa Fluor 647 and Cy5 are the best performing STORM dyes to date (Dempsey, Vaughan, Chen, Bates, & Zhuang, 2011). The conjugation is usually achieved using the NHS-ester chemistry (Ries et al., 2012) and ideally the ratio of dye to nanobody is close to 1. At higher ratios, the target binding of the nanobody may become compromised, resulting in high background. At lower ratios, some of the nanobodies are not fluorescent which reduces the achievable labeling density. For STORM imaging, the samples are embedded in a buffer containing a primary thiol, which drives the fluorophore switching, an oxygen scavenging system which reduces irreversible photobleaching, and a buffer which maintains the pH during the measurement time on the microscope. The measurements are performed on a widefield or TIRF microscope equipped with a sensitive EMCCD camera. Commercial setups for localization microscopy are now available from several manufacturers (e.g., Leica, Zeiss, and Nikon). For details on the hardware, which are beyond the scope of this chapter, the reader is referred to Bates et al. (2013) and Dempsey et al. (2011). We performed our imaging on Leica SR GSD, equipped with 500 mW 642 nm continuous wave excitation laser, a 30 mW 405 nm diode activation laser; DBP 405/10 642/10 excitation filter; BP 710/100 emission filter and Leica HCX PL APO 100, NA 1.47 Oil CORR TIRF PIFOC objective; and an additional 1.6  magnification lens. 1. U2-OS cells are cultured in DMEM supplemented with 10% FCS according to standard protocols. For transfection, plate 100,000 cells in a two-well LabTek. Next day, transfect the cells using Fugene6, according to the manufacture’s protocol. The amount of DNA and transfection reagent should be optimized for any new cell line or a construct. A good starting point is 0.5–1 mg of DNA and 1:3 DNA to Fugene6 ratio. 2. Twenty-four hours after Nup–GFP transfection, transfer 50,000 cells into a well of a 24-well plate coated with 1.6 pmol of Silencer Select siRNA (Life Technologies) complexed with Lipofectamine 2000 as described in Erfle et al. (2008). For different brands of siRNA the amount should be optimized.

10.3 Structural Analysis of the NPC by Super-resolution Microscopy

3. Forty-eighty hours later, transfer 40,000 cells onto an eight-well LabTek and allow them to attach for 12–16 h. Alternatively, the cells can be plated on acid washed and poly-L-lysine-coated #1.5 cover slips. 4. Rinse the cells with phosphate-buffered saline (PBS) and then incubate for 20 s with fixing solution (2.4% PFA/PBS). This prevents the detaching of the cells during the subsequent steps. Permeabilize and extract the cells by incubating for 3 min in 0.4% Triton X-100/PBS to remove soluble cytoplasmic protein and then fix for 30 min. Quench for 5 min with 50 mM NH4Cl/PBS and wash with PBS. Block the cells for 30 min with a few drops of Image-iT FX Signal Enhancer solution and then with 2% BSA in PBS for 1 h. Stain the cells with Alexa Fluor 647-conjugated nanobody diluted in 2% BSA/PBS for 90 min and wash with PBS. The concentration of the nanobody should be optimized for each batch of the conjugate, but 100–300 ng/ml is a good starting point. All steps are performed at room temperature. The samples can be stored in PBS at 4  C for a few days, but should imaged as soon as possible after preparation. 5. Embed the samples in 0.5 ml of freshly prepared imaging buffer containing: 50 mM Tris pH 8.5, 10 mM NaCl, 10% (w/v) glucose, 0.5 mg/ml glucose oxidase, 40 mg/ml catalase, and 10 mM b-mercaptoethylamine (MEA, store diluted in water as 100 mM stock at 20  C). If cover slips are used, they can be mounted with a drop of buffer on a depression slide and sealed with two-component silicone glue such as Twinsil. The imaging buffer undergoes acidification over time, which adversely influences the switching behavior of the cyanine dyes (Dempsey et al., 2009); therefore, the buffer should be exchanged after about 1–1.5 h. 6. Locate a cell with high and specific signal of anti-Nup–GFP nanobody using low intensities of laser light to prevent photoswitching and irreversible bleaching. The number of cells with high incorporation of the tagged Nup into the pore will vary depending on the construct and the transfection efficiencies, and typically is quite low. Avoid imaging cells with abnormal nuclear shape, exhibiting pore clustering or other aberrant phenotypes. Focus the microscope on the bottom of the cell nucleus, where the NE is flat and several hundreds of NPC are visible in a similar orientation. 7. To drive the fluorophores into dark state, illuminate the sample in epifluorescence mode with high power of 642-nm laser light until singlemolecule switching is observed. For Alexa Fluor 647, this typically takes 0.5–1 min. Begin the data collection. The signal-to-noise ratio should be maximized by proper setting of the EM gain and the frame rate should be adjusted to the switching speed of the fluorophore. For Alexa Fluor 647, we typically acquire 40,000–100,000 frames with 10–30 ms exposure times. The images can be acquired in epifluorescence mode or using highly inclined light (Tokunaga, Imamoto, & Sakata-Sogawa, 2008). Optionally, illumination with 405 nm activation light can be used to increase the rate of fluorophore return from the dark states.

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FIGURE 10.3 Super-resolution Imaging of NPCs and Determination of Radial Position of a Fluorescent Marker (A) Representative STORM image of a 9 mm2 area of the bottom surface of the nucleus of a U2-OS cell transiently expressing mEGFP–Nup160–siRNA-resistant cDNA, stained with an anti-GFP nanobody conjugated to Alexa Fluor 647, 72 h after transfection with siRNA against endogenous Nup160. Scale bar (SB): 0.5 mm. (B) Four exemplary pores suitable for averaging. SB: 0.1 mm. (C) An average image generated by summing of 783 translationally aligned quality-controlled pores. (D) Average radial profile of the image in (C) (black points) fitted with Eq. (10.6) (red line). The mean radial position of the fluorescent nanobody on mEGFP–Nup160 determined in this experiment is 52.8 nm.

After acquisition, the STORM image is reconstructed using the localized positions of the individual molecules (Fig. 10.3A). The details on single-molecule detection, correction of the data for lateral drift, and image reconstruction are reviewed extensively in the literature (e.g., Bates et al., 2013) and commercial STORM systems offer software solutions to perform these tasks; we will therefore not discuss them here. We typically render our images with 10-nm pixels and assign a single gray value of a 16-bit image per each molecule localized within the pixel.

10.3 Structural Analysis of the NPC by Super-resolution Microscopy

10.3.2 Determination of the average radial position of the fluorescent marker To determine the average radial position of the fluorescent marker, single pores are interactively cropped from the SR images in 30  30 pixel boxes with Boxer (EMAN1 image processing package, Baylor College of Medicine) (Fig. 10.3B). The particles are then centered using an iterative alignment method. Next, an average image is generated. The radial position of the marker is determined from the radial intensity profile of the average image. The quality of the SR images is important for the accuracy of the radius measurement. The SR image should have little to no background, be devoid of drift and contain only high-precision localizations (Fig. 10.3A). The average localization precision in an image, measured as the full width at half maximum of signal from a single nanobody, should be about 20–30 nm. The most important factor influencing the precision of the alignment is the labeling density of the structures (see Fig. 10.3B for examples of pores suitable for averaging). We found that images of pores formed by less than fiveresolvable clusters of localizations organized around the pore’s center, cause alignment errors, and lead to an underestimation of the radial position of the marker. We therefore exclude such particles from the analysis. We evaluate the number of localization clusters per image by decomposing the signal at the circumference of the pore to a minimal number of Gaussian peaks (Szymborska et al., 2013). For highest precision alignment, at least 500 quality-controlled images of single pores are required. We perform the translational alignment with IMAGIC (Image Science Software, GmBH), a software for processing of single-particle EM data. First, the images of single pores are normalized to a constant variance of pixel intensities. Next, the images are summed and rotationally averaged to generate the first reference. The normalized images are filtered with a low pass filter to 28 nm. The filtered images are aligned to the reference with a phase correlation method (Kuglin & Hines, 1975). Next, the calculated subpixel shifts are applied to the normalized images. The translated images are then summed and rotationally averaged to generate the next reference and the procedure is iterated. We found that a stable alignment is obtained after 5–7 iterations. The average image is generated by summing all translated normalized images after the last iteration (Fig. 10.3C). To determine the average radial profile, the average image is transformed into a polar coordinate system and the mean intensity along the angular direction is calculated. The average profile is then normalized between 0 and 1 (Fig. 10.3D). Finally, the average radial position of the fluorophore is determined by nonlinear least squares fitting of the profile with the equation (Fig. 10.3D):   2 2 2 R ð10:6Þ f ðr Þ ¼ 2pAeððr þR Þ=2s Þ I0 2 r s where R is the average radial position of the fluorescent marker, s and A are spread and amplitude of the 2D Gaussian peaks forming the pore image, I0 is the modified

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Bessel function of the first kind, r is the position along the radial profile, and f(r) is the intensity along the radial profile. Unless otherwise stated, we performed the analysis in MATLAB (The MathWorks, Inc).

10.4 FUTURE PERSPECTIVE Live-cell imaging enables the detailed kinetic analysis of postmitotic NPC assembly. The protocols used to study NPC assembly (Section 10.1) and NE permeability (Section 10.2) can also be applied to the study of NPC disassembly by observing cells followed from prophase to prometaphase (Dultz et al., 2008). The cells ready to enter prometaphase can be identified by their chromosome condensation state. Combined with molecular perturbations such as RNAi depletion, these assays can be very useful to provide new insight into the molecular mechanism of the process. Automatic imaging systems such as Micropilot, which detects cells of interest in a low-resolution prescan mode and carries out high-resolution imaging experiments once it finds them (Conrad et al., 2011; Terjung et al., 2010), allow to deal with a large number of samples and thus make it possible to perform even large-scale screens for genes involved in the assembly process. The fact that the assembly kinetics is similar between HeLa and NRK cells suggests that the assembly process is conserved in mammals. In a similar manner, the kinetic analysis of NE permeability can be used as an assay to identify the molecular factors required for establishing the postmitotic diffusion barrier. Although slightly more complex to automate, due to the repetitive time series after photoswitching, in principle also this approach is amenable to RNAi screening. Moreover, Dronpa, when fused to additional reporter proteins, may also be used to assess the diffusion of larger molecules and the kinetics of active import/export through the NPC (e.g., Ando et al., 2004). SR imaging is a relatively simple method for probing the structural organization of specific components of the NPC in whole cells and, combined with single-particle averaging, allows to map the average radial positions of individual fluorescent labels on Nups with subnanometer precision. The method will improve with new developments in the labeling technologies, which place the fluorophore closer to the targeted epitope (e.g., Plass, Milles, Koehler, Schultz, & Lemke, 2011) and advancements in the field of SR microscopy, including multicolor, live-cell, and 3D techniques. We expect that this methodology will be useful for addressing questions pertaining to the structure of the mature NPC, as well as NPCs in partially assembled states, in different cell types or for structure-function analysis of the NPC after molecularly defined perturbations.

10.5 MATERIALS AND INSTRUMENTS (A) Microscopes Zeiss LSM780 Leica SR GSD

Acknowledgments

(B) Plasmids and cell lines pIBB-DiHcRed (Dultz et al., 2008) pEGFP3–Nup133 (Belgareh et al., 2001) pEGFP3–Nup107 (Belgareh et al., 2001) pPom121–EGFP3 (Daigle et al., 2001) pEGFP–Nup98 (Rabut et al., 2004) pEGFP3–Nup153 (Daigle et al., 2001) pEGFP3–Nup214–cDNA of Nup214 (von Lindern et al., 1992) was subcloned into the expression vector with appropriate restriction enzymes as described in Daigle et al. (2001). pSrprb–ECFP (Daigle et al., 2001) pDronpa (Ando et al., 2004) pmEGFP–Nup160–siRNA-resistant (Szymborska et al., 2013) HeLa cells stably expressing IBB-DiHcRed (unpublished) For published plasmids used in our studies, please visit EUROSCARF website at the following address to obtain aliquots: http://web.uni-frankfurt.de/fb15/mikro/ euroscarf/data/ellenberg.html (C) Reagents Alexa Fluor 647 NHS-ester (Life Technologies) Anti-GFP nanobody (Chromotek #gt-250) b-Mercaptoethylamine (MEA) (Sigma #30070) Bovine serum albumin (BSA) (Sigma #A7906) Catalase (Sigma #C3155) Fugene6 (Promega) Glucose Oxidase (Sigma #G0543) Hoechst 33342 (Sigma #B2261) Image-iT FX Signal Enhancer (Life Technologies) Lipofectamine 2000 (Life Technologies) Paraformaldehyde (PFA) 16% (Electron Microscopy Sciences) PBS (2.7 mM KCl, 1.5 mM KH2PO4, 8.1 mM Na2HPO4, 137 mM NaCl, pH 7.4) Triton X-100 (Sigma #T8787) Trizma base, NaCl (Sigma #T1503) (D) Other materials 18-mm #1.5 cover slip (Gerhard Menzel GmbH) Glass depression slides (neoLab) Nunc LabTek-chambered cover glass, two- and eight-well (ThermoScientific) Twinsil silicone glue (Picodent, Wipperfu¨rth, Germany)

Acknowledgments We thank Elisa Dultz and Antonio Politi for helpful discussion. This work was supported by funding from the German Research Council to J. E. (DFG EL 246/3-2 within the priority program SPP1175). S. O. is supported by a JSPS fellowship (postdoctoral fellowship for research abroad) and an EMBL interdisciplinary postdoctoral fellowship.

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Szymborska, A., de Marco, A., Daigle, N., Cordes, V. C., Briggs, J. A., & Ellenberg, J. (2013). Nuclear pore scaffold structure analyzed by super-resolution microscopy and particle averaging. Science, 341, 655–658. Terjung, S., Walter, T., Seitz, A., Neumann, B., Pepperkok, R., & Ellenberg, J. (2010). Highthroughput microscopy using live mammalian cells. Cold Spring Harbor Protocols, 2010, 297–314. Tokunaga, M., Imamoto, N., & Sakata-Sogawa, K. (2008). Highly inclined thin illumination enables clear single-molecule imaging in cells. Nature Methods, 5, 159–161. van der Oost, J. (2013). Molecular biology. New tool for genome surgery. Science, 339, 768–770. von Lindern, M., van Baal, S., Wiegant, J., Raap, A., Hagemeijer, A., & Grosveld, G. (1992). Can, a putative oncogene associated with myeloid leukemogenesis, may be activated by fusion of its 3’ half to different genes: Characterization of the set gene. Molecular and Cellular Biology, 12, 3346–3355.

CHAPTER

Cell-Fusion Method to Visualize Interphase Nuclear Pore Formation

11

Kazuhiro Maeshima*,{,{, Tomoko Funakoshi*,}, and Naoko Imamoto* *

Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan Biological Macromolecules Laboratory, Structural Biology Center, National Institute of Genetics, Mishima, Shizuoka, Japan { Department of Genetics, School of Life Science, Graduate University for Advanced Studies (Sokendai), Mishima, Shizuoka, Japan } Department of Biochemistry, Faculty of Pharmaceutical Sciences, Toho University, Funabashi, Chiba, Japan

{

CHAPTER OUTLINE Introduction ............................................................................................................ 240 Interphase NPC Assembly ...................................................................................... 241 Cell-fusion Assays in Mammalian Cells (heterokaryons Assays)................................ 243 11.1 Materials and Equipment .................................................................................246 11.1.1 Cell Lines .................................................................................. 246 11.1.2 Reagents ................................................................................... 246 11.1.3 Equipments ...............................................................................246 11.2 Quantitative Analysis of Interphase NPC Formation using Cell-Fusion Method .....247 11.3 Combining the Cell-Fusion Method with Drug and siRNA Treatments .....................249 11.3.1 Combination of Cell-fusion Method with Drug Treatment................ 249 11.3.2 Combination of Cell-fusion Method with Protein Knockdown Using siRNAs............................................................................. 249 11.4 Visualization of Interphase NPC Formation Using Photobleaching ......................250 Conclusions............................................................................................................ 252 Acknowledgments ................................................................................................... 252 References ............................................................................................................. 252

Abstract In eukaryotic cells, the nucleus is a complex and sophisticated organelle that organizes genomic DNA to support essential cellular functions. The nuclear surface

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00011-4

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contains many nuclear pore complexes (NPCs), channels for macromolecular transport between the cytoplasm and nucleus. It is well known that the number of NPCs almost doubles during interphase in cycling cells. However, the mechanism of NPC formation is poorly understood, presumably because a practical system for analysis does not exist. The most difficult obstacle in the visualization of interphase NPC formation is that NPCs already exist after nuclear envelope formation, and these existing NPCs interfere with the observation of nascent NPCs. To overcome this obstacle, we developed a novel system using the cell-fusion technique (heterokaryon method), previously also used to analyze the shuttling of macromolecules between the cytoplasm and the nucleus, to visualize the newly synthesized interphase NPCs. In addition, we used a photobleaching approach that validated the cell-fusion method. We recently used these methods to demonstrate the role of cyclin-dependent protein kinases and of Pom121 in interphase NPC formation in cycling human cells. Here, we describe the details of the cell-fusion approach and compare the system with other NPC formation visualization methods.

INTRODUCTION In eukaryotic cells, the genomic DNA and cytoplasm are physically separated by the nuclear envelope (NE). The nuclear pore complexes (NPCs) provide aqueous gates for nucleocytoplasmic transport during interphase. They are very large protein complexes (estimated mass, 60–125 MDa) assembled with eightfold symmetry from multiple copies of approximately 30 different nucleoporins (Nups). In vertebrates, the NPC contains three integral membrane pore proteins (Ndc1, gp210, and Pom121), which anchor the NPC scaffold to the nuclear membrane (reviewed in Antonin, Ellenberg, & Dultz, 2008). Some Nups that form the NPC structural scaffold, such as members of the Nup107–Nup160 complex, Nup93–Nup205 complex, and Pom121, are stably embedded in the NPC during interphase with a residence time of several tens of hours (Rabut, Doye, & Ellenberg, 2004; Rabut, Lenart, & Ellenberg, 2004). In contrast, the more peripheral Nups, such as Nup153 and Nup50, are highly dynamic, with residence times of seconds or minutes. While the NPC scaffold is maintained with little to no turnover throughout the life span of postmitotic differentiated cells (D’Angelo, Raices, Panowski, & Hetzer, 2009), NPCs are dynamic structure in dividing cells (reviewed in FernandezMartinez & Rout, 2009; Kutay & Hetzer, 2008). NPCs are assembled twice in the cell cycle of dividing metazoan cells. In mitosis, NPCs are disassembled (see Chapter 12 from Marino et al. in this volume) and must be reassembled at the end of mitosis (postmitotic NPC assembly, see Chapters 8, Bernis and Forbes, and 9, Eisenhardt et al.). During interphase, assembly also occurs to increase the number of NPCs for eventual distribution of their components into two daughter cells (interphase NPC formation; Maul et al., 1972). Postmitotic NPC assembly occurs concomitantly with the formation of the NE around chromatin (Maul, 1977).

Introduction

Assembly begins in early anaphase, with the recruitment of the essential Nup107– Nup160 scaffold complex to the chromatin. This recruitment is mediated by ELYS/Mel28, a Nup that contains an AT-hook DNA-binding motif. This is followed by the recruitment of the other scaffold Nups, Pom121, and Nup93, and finally, of more peripheral Nups such as Nup153 and Nup50 (reviewed in Antonin et al., 2008; Franz et al., 2007; Imamoto & Funakoshi, 2012; see also Otsuka et al., Chapter 10 in this volume).

Interphase NPC assembly During interphase, the number of NPCs on the nuclear surface doubles in preparation for reentry into next mitosis (Imamoto & Funakoshi, 2012; Maul et al., 1972; Fig. 11.1A). This process, that involves the assembly of NPCs into an intact NE (also termed de novo NPC assembly), also occurs in some nondividing vertebrate cells, such as in oogenesis, and is the unique mode of pore assembly in organisms with closed mitosis such as budding and fission yeasts (reviewed in Doucet & Hetzer, 2010). However, as its study required the development of specific assays, less is so far known about the mechanism of interphase NPC formation than about postmitotic processes. To study interphase NPC formation, one must quantify the newly assembled (nascent) NPCs on the nuclear surface. While approaches to examine interphase NPC assembly have been developed using in vitro nuclear reconstitution assays based on Xenopus egg extracts (reviewed in Doucet & Hetzer, 2010), studies of this process in mammalian cells are still limited: the current approaches notably involve (i) comparisons of NPC levels, based on total fluorescence intensity at the NE in G1 versus G2 in synchronized cells (Doucet, Talamas, & Hetzer, 2010), (ii) highresolution live-cell imaging in cell lines stably expressing GFP–Nups that enabled the observation of new NPC assembly events at previously pore-free sites (Dultz & Ellenberg, 2010), or (iii) simultaneous localization of two distinct Nups at the NE surface in fixed or live cells that notably revealed the earlier recruitment of Pom121 as compared to Nup107 in interphase NPC assembly (Doucet et al., 2010; Dultz & Ellenberg, 2010). However, the counting of NPCs, even when semiautomatized (Dultz & Ellenberg, 2010), proved laborious and the resolution of light microscopy is insufficient to distinguish between adjacent NPCs. In addition, increase in the nuclear surface area as the cell cycle progressed also affects the NPC density, thus complicating the analysis (Dultz & Ellenberg, 2010). The presence of pore-free islands within the NE in early G1 cells of human dividing cells may also potentially introduce bias in these analyses (Maeshima et al., 2006). As an alternative approach to directly visualize nascent NPCs on the nuclear surfaces during interphase, we have used a photobleaching approach (FRAP, Figs. 11.1C and 11.4) (Iino et al., 2010; Maeshima et al., 2010) and also developed a new method based on the cell-fusion technique (Figs. 11.1B, 11.2, and 11.3; Maeshima et al., 2010, 2011; Funakoshi et al., 2011).

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A

B Control cells

C Cell-fusion method Donor

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G1 cells Cell fusion by PEG

16-18 h

FRAP

16 h

G2 cells

Nascent Venus–Nup-labeled NPC Preexisting Venus–Nup-labeled NPC Preexisting and nascent nonfluorescent NPC Bleached Venus–Nup-labeled NPC

FIGURE 11.1 Cell-fusion and Photobleaching Methods for the Visualization of Interphase NPC Assembly (A) The number of nuclear pore complexes (NPCs) almost double during interphase in dividing cells. (B) Schematic representation of the cell-fusion method. HeLa cells expressing Venus–Nup are used as donor cells, and HeLa cells expressing CFP–H2B are used as acceptor cells (top). G1-synchronized donor and acceptor cells are treated with polyethylene glycol (PEG) to make heterokaryons (middle). Sixteen to eighteen hours later, new Venus– Nup-labeled NPCs should appear on the acceptor nucleus (bottom right), which has the brighter CFP–H2B signal. (C) The photobleaching method. Nuclear surface regions in G1-phase cells expressing Venus–Nup (top) are photobleached by a 488 nm laser (middle). At 16 h after bleaching, new Venus–Nup-labeled NPCs appear in the bleached areas, representing newly synthesized NPCs (bottom).

Introduction

FIGURE 11.2 Visualization of New NPC Formation by Cell-fusion Method (A) A simplified scheme of cell-fusion methods. (B) Immediately after fusion (0 h), donor nuclei from Nup133–Venus-expressing cells display many bright fluorescent dots representing NPCs, whereas no detectable dots are observed on the acceptor nuclei from CFP–H2B-expressing cells. (C) The Venus fluorescent signals in acceptor nuclei increase in time-dependent manner (4, 8, 12, and 18 h). In the middle view (second row), the relative mean intensity (acceptor rim/donor rim) is shown in the brackets on the image. In the surface view (third row), the number in the brackets is dot density on the nuclear surface. Scale bar, 10 mm. The images were reproduced from Maeshima et al. (2010), with permission.

Cell-fusion assays in mammalian cells (heterokaryons assays) For heterokaryons-based assays in budding yeast, see Altvater et al., Chapter 20 in this volume. It is now well established that the fusion event of heterotypic cells is important for development, repair of tissues and the pathogenesis of disease (for review Ogle, Cascalho, & Platt, 2005). Induction of cell fusion by the Sendai virus (HVJ,

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Fusion

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+ aphidicolin

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(% of Pom121 KD(–), time 0: indicated by*)

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FIGURE 11.3 Combining siRNA Treatment and Cell-fusion Method Reveals the Implication of Pom121 in Interphase NPC Assembly (A) Schematic representation of the interphase NPC assembly method combined with siRNA treatment. (B) Effect of Pom121 depletion (Pom121KD) on interphase NPC assembly. Arrowheads indicate acceptor nuclei in heterokaryons. At 16-h postfusion (16 h, panels

Introduction

Hemagglutinating Virus of Japan) was first introduced by Okada (1962a, 1962b) and Okada and Tadokoro (1962). Harris and colleagues used this technique to demonstrate that both differentiated and undifferentiated cells from different animal species could be successfully fused together to form viable heterokaryons (cells formed by the fusion of two or more cells of different types and having two or more distinct nuclei; Harris, 1965). An alternative approach to induce cell fusions was developed using polyethylene glycol (PEG) that allows cell membrane fusion of attached cells within seconds (Hales, 1977; Harel et al., 2003). This approach has notably been used to follow the shuttling of proteins between the cytoplasm and the nucleus (Nakrieko, Ivanova, & Dagnino, 2010; Pin˜ol-Roma & Dreyfuss, 1992; Yokoya, Imamoto, Tachibana, & Yoneda, 1999) or to visualize the dynamics of NPC-associated proteins (Katahira et al., 1999). Cell-fusion methods require distinguishing donor from recipient cell. Therefore, either cells from distinct species (distinguished by speciesspecific antibodies or typical chromatin features), or identical cell types transiently or stably expressing distinct markers have been used. To follow cell cycle-specific events, cells that are to be fused must be synchronized at the same cell cycle stage. In this chapter, we detail a modified cell-fusion method that allows monitoring interphase NPC assembly in mammalian cells. This method fuses early G1 synchronized cells stably expressing a yellow-fluorescently tagged scaffold Nup (e.g., Venus–Nup107 or Venus–Nup133) (donor cells) and CFP–H2B (acceptor cells), respectively. We describe how this method can be combined with drug treatments or protein knockdown using RNAi to further analyze the molecular mechanisms contributing to interphase NPC assembly. Finally, we also detail an alternative protocol, based on photobleaching of G1 cells and discuss the respective advantages of these two methods.

7–12), a clear nuclear rim signal is observed with Venus–Nup107 on the acceptor nuclei of control heterokaryons (panel 7) that are also strongly labeled with the anti-Pom121 antibody (panel 9). In contrast, a very low Venus–Nup107 signal is observed on the acceptor Pom121 KD nuclei (panels 10–12). Scale bar, 10 mm. (C) Quantification results of fluorescence intensities of Pom121 and Venus–Nup107 at the nuclear rim of nuclei in heterokaryons. Fluorescent intensities of Venus–Nup107 (graphs b, d), or Pom121 detected by immunofluorescence staining (a, c) at the rim of nuclei derived from acceptor nuclei (a, b) and donor nuclei (c, d) in cells with or without Pom121 KD were measured. Mean values were plotted and standard deviations are shown as relative percentages to the level of the Pom121 KD(
) at time ¼ 0 (marked with asterisks in graphs). Pom121 KD(
): n ¼ 4 (0 h), n ¼ 22 (16 h). Pom121 KD(þ): n ¼ 9 (0 h), n ¼ 32 (16 h). The schematic and images were reproduced from Funakoshi, Clever, Watanabe, and Imamoto (2011), with permission.

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11.1 MATERIALS AND EQUIPMENT 11.1.1 Cell lines – HeLa cells expressing Nup133–Venus, Venus–Nup107, or CFP–H2B can be supplied upon request (Funakoshi et al., 2011; Maeshima et al., 2010). – Cells are grown in DMEM medium (Sigma C11965500BT) containing 10% fetal bovine serum (Gibco-BRL) at 37  C under 5% CO2.

11.1.2 Reagents – Aphidicolin (1 mg/ml in ethanol stored at
20  C, Sigma A0781) – Cell culture dish (10-cm diameter and 6-cm diameter, e.g., Corning 430167 and 430166) – Formaldehyde (16% solution stored at room temperature, Polyscience 50-00-0) – Glass-base 3.5-cm dish (Iwaki 3910–035) – Lipofectamine 2000 (stored at 4  C, Life Technologies 11668–027) – Nail polish for sealing coverslips – Nocodazole (5 mg/ml in DMSO stored at
20  C, Sigma M1404) – OptiMEM (stored at 4  C, Life Technologies 31985062) – Phosphate-buffered saline (PBS) – Polyethylene glycol 1500 (50% PEG 1500 (w/v) stored at 4  C, Roche 10783641001) – Poly-L-lysine-coated coverslips (12- or 18-mm diameter; Iwaki): the coverslips are coated with poly-L-lysine solution [0.1% (w/v) in H2O, Sigma P8920] for 5 min, then washed with dH2O twice, and air-dried. – Pom121-specific siRNA duplex (50 CCACCACAGUCACCACCUUCAGCCA-30 , 50 -UGGCUGAAGGUGGUGACUGUGGUGGUU-30 ) (20 mM in RNase-free dH2O stored at
20  C, Integrated DNA Technologies). The siRNA duplex used was previously shown to be effective (oligo-2 in Funakoshi et al., 2007). – PPDI mounting medium: 20 mM HEPES, pH 7.4, 1 mM MgCl2, 100 mM KCl, 78% glycerol, 1 mg/ml paraphenylene diamine (Sigma P6001). The medium should be kept at
80  C with light protection. – Purvalanol A (10 mM in DMSO stored at
20  C, Sigma P4484) – Roscovitine (10 mM in DMSO stored at
20  C, Sigma R7772) – Thymidine (100 mM in PBS stored at
20  C, Sigma T9250)

11.1.3 Equipments – Cell culture chamber for live-cell imaging (Olympus MI-IBC) – DeltaVision RT microscope (Applied Precision) includes an inverted Olympus IX70 with a cooled CCD camera (Photometrics CoolSnap HQ) and standard CFP and YFP color channels.

11.2 Quantitative Analysis of Interphase NPC Formation

– DeltaVision RT quantifiable laser module (50 mW, 488-nm solid-stable laser) (Applied Precision) (for photobleach method) – DeltaVision Softworx softwear (Applied Precision) – PlanApo 60 /1.40 oil-immersion objective (Olympus) (for cell-fusion and photobleach methods)

11.2 QUANTITATIVE ANALYSIS OF INTERPHASE NPC FORMATION USING CELL-FUSION METHOD For our cell-fusion method, we established two HeLa cell lines stably expressing fusions of a bright yellow fluorescent protein Venus (Nagai et al., 2002) with Nup133 or Nup107, scaffold proteins that are essentially immobile in NPCs. Many fluorescent dots of NPCs, visualized by Nup133–Venus or Venus–Nup107, were observed on the nuclear surface (Figs. 11.2 and 11.3). In these cell lines, endogenous Nup133 or Nup107 is dramatically downregulated, perhaps to maintain a constant level of these scaffold Nups, suggesting that their ectopic expression was unlikely to be problematic (Maeshima et al., 2010). The cells expressing Nup133–Venus or Venus–Nup107 were used as donor cells, and cells stably expressing a cyan fluorescence protein (CFP: Rekas, Alattia, Nagai, Miyawaki, & Ikura, 2002) histone H2B (H2B) were used as acceptor cells (Fig. 11.1B). Synchronized G1-donor and -acceptor cells were fused with PEG to make heterokaryons (Fig. 11.1B; protocol adapted from Katahira et al., 1999; Pin˜ol-Roma & Dreyfuss, 1992). To prevent mitotic entry of the cells, we used the DNA polymerase a/d-specific inhibitor aphidicolin (Ikegami et al., 1978), which arrests synchronized control cells at the G1/S transition (Pedrali-Noy et al., 1980). Aphidicolin did not inhibit NPC formation during interphase (Fig. 11.2C; Maul, Hsu, Borun, & Maul, 1973). Immediately after cell fusion (defined as time 0), many bright Nup–Venus dots are observed on the surface of the donor nuclei (that has no CFP–H2B signal), whereas no signal is detectable on acceptor nuclei derived from CFP–H2B-expressing cells (Fig. 11.2B). 18 h after fusion, the acceptor nuclear surfaces (characterized by brighter CFP–H2B signal than the donor nuclei) are covered with many bright Nup–Venus fluorescent dots (Fig. 11.2C). In addition, the number of fluorescent dots on the acceptor nuclear surfaces increases in a time-dependent manner (Fig. 11.2C). Because Nup133 and Nup107 are immobile scaffold proteins of NPCs (Rabut, Doye, et al., 2004), these bright Nup–Venus dots correspond to newly formed NPCs. PROTOCOL: CELL SYNCHRONIZATION To monitor interphase NPC formation, use of synchronized early G1 cells is essential. 1. One day before the experiment, seed the both donor and acceptor HeLa cell lines on 10-cm cell culture dishes (five dishes for each cell line, at 2  106 cells/dish).

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2. Block cells in metaphase: add 0.1 mg/ml nocodazole and incubate for 4 h (approximately 15–20% of the cells should be arrested in mitosis). Note: Do not use longer nocodazole treatment. While it may increase the percentage of mitotic cells, this may interfere with release from the nocodazole block. 3. Collect mitotic cells by shaking the culture dishes; wash the cells three times with PBS and count the number of cells. Releasing the nocodazole block will allow the cells to exit mitosis and enter the G1 phase in a synchronous manner.

CELL FUSION 4. Mix equal numbers of mitotically released Nup133–Venus (or Venus–Nup107) donor cells and CFP–H2B acceptor cells (for each, 2.5  105 cells) and seed them on 6-cm dish containing 5 poly-L-lysine-coated coverslips. Note: For successful cell-fusion, an appropriate density, where the donor and acceptor cells are only partially in contact, is essential. High densities of cells and too much contact cause the cells to become multinucleated. The appropriate density for the cells used depends on their cell size and should be determined before experiment. 5. After completion of cytokinesis (approximately 3 h after releasing the mitotic cells), wash the coverslips in the 6-cm dish twice with prewarmed PBS. Apply 200 ml of 50% PEG solution onto each coverslip and incubate them for 2 min at room temperature to allow for cell fusion. 6. To remove the PEG, wash the cells on coverslips twice with 10 ml of prewarmed PBS. 7. Transfer the cells with coverslips into new 6-cm dish containing prewarmed medium (defined as t ¼ 0), and incubate for 1 h at 37  C under 5% CO2 for completion of fusion and settling the fused cell. For one coverslip, move to step 9 (t ¼ 0 time point) prior to the 1 h incubation. 8. At that stage (t ¼ 0), add aphidicolin (5 mg/ml) to the medium to prevent mitotic entry of the cells (aphidicolin can be added just before or just after starting the 1 h incubation in step 7). FLUORESCENCE IMAGING AND QUANTIFICATION 9. At t ¼ 0 and after 16–18 h (or at various time points; see Fig. 11.2C), briefly wash the cells with coverslips with PBS and fix them in 2% formaldehyde for 15 min at room temperature; mount the coverslips using PPDI mounting medium, and seal with nail polish. 10. Record images of heterokaryons with donor and acceptor nuclei using the DeltaVision RT microscope. Note: We usually record image stacks with the DeltaVision microscope using an Olympus PlanApo 60/1.40 objective and a step size of 0.2 mm. Images are acquired without deconvolution. In the stacks obtained, optical sections including the nuclear surfaces close to the coverslips allow to visualize distinct NPCs.

11.3 Combining the Cell-Fusion Method with Drug and siRNA Treatments

11. Quantify the Venus signals on the nuclear surfaces or rim regions (Figs. 11.2 and 11.3). Note: We use the Softworx software. For the nuclear surface signals, fluorescent dot density on each nuclear surface (dots/mm2) was determined. For the rim signals, the nuclear edge is manually traced based on the H2B–CFP signal or Nup133–Venus signal. Regions with a 13-pixel width, with the nuclear edges as the center, are defined as the nuclear rim regions. A mean value of fluorescence intensity in the rim regions is measured using cytoplasmic signals as a background.

11.3 COMBINING THE CELL-FUSION METHOD WITH DRUG AND SIRNA TREATMENTS 11.3.1 Combination of cell-fusion method with drug treatment Use of specific inhibitors is a powerful way to examine specific cellular functions. To understand how interphase NPC formation is regulated during the cell cycle, we used roscovitine, a selective inhibitor of cyclin-dependent protein kinases (Cdks; Bach et al., 2005; Whittaker et al., 2007), as Cdks are the master regulators of the eukaryotic cell cycle (Morgan, 2007). We added aphidicolin or roscovitine (40 mM) 1 h after cell fusion. Quantitative measurements also showed that the acceptor rims in roscovitine-treated heterokaryons had significantly weaker Nup133 signals (0.151  0.049) than those in the aphidicolin-treated cells (0.504  0.068; p-value < 0.0001). These data indicated that roscovitine inhibits NPC formation during interphase (Maeshima et al., 2010, 2011), and Cdk activity is involved in interphase NPC formation.

11.3.2 Combination of cell-fusion method with protein knockdown using siRNAs In vertebrates, Pom121 is one of transmembrane Nups. Because Pom121 is stably associated with NPC (Rabut, Doye, et al., 2004), it was possible to observe the effect of Pom121-depletion on interphase NPC assembly by performing Pom121 knockdown only in acceptors cell prior to cell fusion (see scheme in Fig. 11.3A). Signals of Venus–Nup107 on the acceptor nuclei of heterokaryons after cell fusion indicate nascent NPC formation on the acceptor nuclei. Unlike in control experiments, when acceptor cells were treated with siRNA against Pom121 prior to cell fusion, fewer Venus–Nup107 fluorescent dots were detected on acceptor nuclei after 16 h (35% decrease in Venus–Nup107 intensity compared to that of untreated nuclei— Fig. 11.3B and C; Funakoshi et al., 2011). These data demonstrated that Pom121 is indispensable for interphase NPC formation in HeLa cells.

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PROTOCOL (SEE ALSO FIG. 11.3A): 1. Seed donor (Venus–Nup107) cells and acceptor (CFP–H2B) cells (for each cells, 2  105 cell/3.5-cm dish). 2. Six hours later, treat both cell lines with 2 mM thymidine for 16 h. 3. Release the thymidine block by rinsing cells twice with prewarmed PBS and once with culture medium, and then incubate the cells for 4 h. 4. Using Lipofectamine 2000, transfect the acceptor CFP–H2B cells with Pom121-specific siRNA duplex as detailed below: – Add 6.25 ml of 20 mM siRNA to 250 ml of OptiMEM and mix. As control, siRNA buffer (30 mM Hepes, pH 7.5, 100 mM potassium acetate, in RNasefree dH2O) is used. – Add 2.5 ml Lipofectamine 2000 solution to 250 ml of OptiMEM, mix, and incubate for 15 min at room temperature. – Mix diluted siRNA and Lipofectamine solutions, incubate for 15 min at room temperature, and then add 500 ml of the mixture to the acceptor CFP–H2B cells in a 3.5 cm dish. 5. Four hours after siRNA transfection (about 8 h after thymidine release) replace the culture media of both donor and dsRNA-transfected acceptor cell lines by fresh medium containing 2 mM thymidine. Incubate for further 16 h and then release thymidine block (e.g., repeat steps 2 and 3). 6. Eight hours after release treat both cell lines with 0.1 mg/ml nocodazole and incubate for 3–4 h. 7. Collect the mitotic cells by mitotic shake-off and wash three times with prewarmed PBS (see steps 1–3 of the general protocol in Section 11.2.1). 8. Mix equal numbers of mitotically released Venus–Nup107 donor cells and CFP–H2B acceptor cells (2  104 cells for each) and seed them on 24-well plate-containing 12-mm poly-L-lysine-coated coverslips. 9. Perform cell fusion as described in the previous protocol (Section 11.2.1, starting from step 5). Note: To evaluate the knock down efficiency of the siRNA, immunofluorescence can be performed using specific antibodies (Pom121 antibodies in our case). 10. Record images of heterokaryons with donor and acceptor nuclei using the DeltaVision RT microscope and quantify the Venus signals on the nuclear surfaces or rim regions (see Section 11.2.1, steps 10 and 11).

11.4 VISUALIZATION OF INTERPHASE NPC FORMATION USING PHOTOBLEACHING As an alternative approach to visualize interphase NPC formation, we photobleached certain nuclear surface areas in Nup133–Venus or Venus–Nup107-expressing G1 cells with a 488-nm laser (Figs. 11.1C and 11.4; Iino et al., 2010; Maeshima et al., 2010, 2011). Sixteen hours after bleaching, many bright dots appeared in

11.4 Visualization of Interphase NPC Formation Using Photobleaching

FIGURE 11.4 Visualization of new NPC formation by photobleaching method (A) A simplified scheme of the photobleaching method. (B) Typical outcome of a FRAP method on two distinct Nup133–Venus-expressing cells. Sixteen hours after bleaching, many bright Nup133–Venus dots appear in the bleached areas. Relative mean intensity (bleached area/unbleached area) is shown in the brackets on the image. Scale bar, 10 mm. The schematic was reproduced from Maeshima, Iino, Hihara, and Imamoto (2011).

the bleached areas, as they did in the acceptor nuclei in heterokaryons. These bright dots did not appear when G1 cells were treated for 16 h with Cdk inhibitors roscovitine (40 mM) or purvalanol A (30 mM), confirming that the contribution of Cdk in interphase NPC formation (for additional studies using this photobleaching method; see Daigle et al., 2001; D’Angelo, Anderson, Richard, & Hetzer, 2006). PROTOCOL: 1. Prepare synchronized mitotic cells according to the previous protocol (see Section 11.2.1, steps 1–3). Following nocodazole release seed cells onto a poly-L-lysine-coated glass-base 3.5-cm dish (1  105 cells). 2. Three hours after releasing the mitotic cells, add aphidicolin (5 mg/ml) or Cdk inhibitors roscovitine (40 mM) [or purvalanol A (30 mM)]. 3. Move the dish to the MI-IBC live-cell chamber on the DeltaVision RT microscopy and record the prebleach images. 4. Photobleach half of the nuclear surface area using the quantifiable laser module (50 mW, 488-nm solid-state laser) of the DeltaVision RT microscope, and acquire postbleach images immediately after photobleaching.

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Note: The nuclear surface in a region of interest is generally completely bleached using a 1-s stationary pulse at 50% laser power. If required, photobleaching can be repeated twice or more. 5. Sixteen hours later, acquire the postbleach images. Note: To quantify the fluorescent signals in the bleached area, mean values of fluorescent intensity in the bleached, and unbleached areas are measured using the cytoplasmic signals as a background. The relative values of the mean intensity (bleached area/unbleached area) are then measured for each cell and plotted (Fig. 11.4B).

CONCLUSIONS We describe two complementary approaches for direct visualization of newly formed interphase NPCs on nuclear surfaces: a cell-fusion method and a photobleaching method. The cell-fusion method, which reveals NPC formation events on whole acceptor nuclei, has a low background signal and avoids risk of laser-induced cell damage. This method is also suitable to quantitative analysis on a large number of nuclei. Alternatively, the photobleaching methods offers the advantage of selective monitoring particular cells of interest. It is especially useful for combination with knockdown experiments using selective siRNAs (Maeshima et al., 2010, 2011). These imaging systems should provide useful tools to further our understanding of the mechanism of interphase NPC assembly.

Acknowledgments We are grateful to all the collaborators, especially Mr. Iino, who contributed to the development of the cell-fusion methods. We also thank Dr. Tachibana for helpful discussion. This work was supported by a MEXT grant-in-aid and RIKEN special project funding for Basic Science in Cellular System.

References Antonin, W., Ellenberg, J., & Dultz, E. (2008). Nuclear pore complex assembly through the cell cycle: Regulation and membrane organization. FEBS Letters, 582(14), 2004–2016. Bach, S., Knockaert, M., Reinhardt, J., Lozach, O., Schmitt, S., Baratte, B., et al. (2005). Roscovitine targets, protein kinases and pyridoxal kinase. The Journal of Biological Chemistry, 280(35), 31208–31219. Daigle, N., Beaudouin, J., Hartnell, L., Imreh, G., Hallberg, E., Lippincott-Schwartz, J., et al. (2001). Nuclear pore complexes form immobile networks and have a very low turnover in live mammalian cells. The Journal of Cell Biology, 154(1), 71–84. D’Angelo, M. A., Anderson, D. J., Richard, E., & Hetzer, M. W. (2006). Nuclear pores form de novo from both sides of the nuclear envelope. Science, 312(5772), 440–443.

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Maul, G. G. (1977). Nuclear pore complexes. Elimination and reconstruction during mitosis. The Journal of Cell Biology, 74(2), 492–500. Maul, H. M., Hsu, B. Y., Borun, T. M., & Maul, G. G. (1973). Effect of metabolic inhibitors on nuclear pore formation during the HeLa S3 cell cycle. The Journal of Cell Biology, 59(3), 669–676. Maul, G. G., Maul, H. M., Scogna, J. E., Lieberman, M. W., Stein, G. S., Hsu, B. Y., et al. (1972). Time sequence of nuclear pore formation in phytohemagglutinin-stimulated lymphocytes and in HeLa cells during the cell cycle. The Journal of Cell Biology, 55(2), 433–447. Morgan, D. O. (2007). The cell cycle-principles of control. London: New Science Press Ltd. Nagai, T., Ibata, K., Park, E. S., Kubota, M., Mikoshiba, K., & Miyawaki, A. (2002). A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nature Biotechnology, 20(1), 87–90. Nakrieko, K. A., Ivanova, I. A., & Dagnino, L. (2010). Analysis of nuclear export using photoactivatable GFP fusion proteins and interspecies heterokaryons. Methods in Molecular Biology, 647, 161–170. http://dx.doi.org/10.1007/978-1-60761-738-9_9. Ogle, B. M., Cascalho, M., & Platt, J. L. (2005). Biological implications of cell fusion. Nature Reviews. Molecular Cell Biology, 6(7), 567–575. Okada, Y. (1962a). Analysis of giant polynuclear cell formation caused by HVJ virus from Ehrlich’s ascites tumor cells. I. Microscopic observation of giant polynuclear cell formation. Experimental Cell Research, 26, 98–107. Okada, Y. (1962b). Analysis of giant polynuclear cell formation caused by HVJ virus from Ehrlich’s ascites tumor cells. III. Relationship between cell condition and fusion reaction or cell degeneration reaction. Experimental Cell Research, 26, 119–128. Okada, Y., & Tadokoro, J. (1962). Analysis of giant polynuclear cell formation caused by HVJ virus from Ehrlich’s ascites tumor cells. II. Quantitative analysis of giant polynuclear cell formation. Experimental Cell Research, 26, 108–118. Pedrali-Noy, G., Spadari, S., Miller-Faures, A., Miller, A. O., Kruppa, J., & Koch, G. (1980). Synchronization of HeLa cell cultures by inhibition of DNA polymerase alpha with aphidicolin. Nucleic Acids Research, 8(2), 377–387. Pin˜ol-Roma, S., & Dreyfuss, G. (1992). Shuttling of pre-mRNA binding proteins between nucleus and cytoplasm. Nature, 355(6362), 730–732. Rabut, G., Doye, V., & Ellenberg, J. (2004). Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nature Cell Biology, 6(11), 1114–1121. Rabut, G., Lenart, P., & Ellenberg, J. (2004). Dynamics of nuclear pore complex organization through the cell cycle. Current Opinion in Cell Biology, 16(3), 314–321. Rekas, A., Alattia, J. R., Nagai, T., Miyawaki, A., & Ikura, M. (2002). Crystal structure of venus, a yellow fluorescent protein with improved maturation and reduced environmental sensitivity. The Journal of Biological Chemistry, 277(52), 50573–50578. Whittaker, S. R., Te Poele, R. H., Chan, F., Linardopoulos, S., Walton, M. I., Garrett, M. D., et al. (2007). The cyclin-dependent kinase inhibitor seliciclib (R-roscovitine; CYC202) decreases the expression of mitotic control genes and prevents entry into mitosis. Cell Cycle, 6(24), 3114–3131. Yokoya, F., Imamoto, N., Tachibana, T., & Yoneda, Y. (1999). beta-Catenin can be transported into the nucleus in a Ran-unassisted manner. Molecular Biology of the Cell, 10(4), 1119–1131.

CHAPTER

An In Vitro System to Study Nuclear Envelope Breakdown

12

Joseph Marino, Lysie Champion, Cornelia Wandke, Peter Horvath, Monika I. Mayr, and Ulrike Kutay Institute of Biochemistry, ETH Zurich, Zurich, Switzerland

CHAPTER OUTLINE Introduction ............................................................................................................ 256 12.1 Preparative Steps............................................................................................258 12.1.1 Generation of Cell Lines Expressing a Fluorescent NE Marker......... 258 12.1.2 Mitotic Extract ........................................................................... 260 12.1.2.1 HeLa S3 Cell Culture.......................................................... 260 12.1.2.2 Preparation of Mitotic Extract ............................................. 261 12.1.3 Fluorescently Labeled Dextrans ................................................... 262 12.1.3.1 Preparation of 155 kDa TRITC-dextran............................... 263 12.1.3.2 Labeling of a 70 kDa Aminodextran with Alexa Fluor 405 Carboxylic Acid Succinimidyl Ester.................................................... 263 12.1.4 Energy Mix................................................................................. 263 12.2 NEBD Assay ....................................................................................................264 12.2.1 Cell Seeding .............................................................................. 264 12.2.2 Semi-permeabilization of Cells .................................................... 265 12.2.3 NEBD Reaction Mix .................................................................... 265 12.2.4 Image Acquisition ...................................................................... 266 12.2.4.1 Microscopic Setup and Acquisition Settings........................ 266 12.2.4.2 Image Acquisition .............................................................. 267 12.3 Special Treatments .........................................................................................267 12.3.1 Modification of Reporter Cells ..................................................... 267 12.3.1.1 Overexpression of Nucleoporin Mutants ............................. 267 12.3.1.2 Knockdown of Endogenous Protein .................................... 267 12.3.2 Modification of Cell Extract ......................................................... 269 12.3.2.1 Chemical Inhibitors............................................................ 269 12.3.2.2 Protein Depletion and Add-back......................................... 269 12.4 Data Analysis..................................................................................................270 12.4.1 Computational Steps .................................................................. 271 12.4.2 Drift Correction of the Image Sequences....................................... 271 12.4.3 Detection of Cell Nuclei .............................................................. 271 Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00012-6

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12.4.4 Statistical Measurements ............................................................ 271 12.4.5 Reporting................................................................................... 273 12.5 Future Directions ............................................................................................273 12.6 Material and Reagents ....................................................................................273 12.6.1 Material (listed in Alphabetical Order) .......................................... 273 12.6.2 Reagents (listed in Alphabetical Order) ........................................ 274 Acknowledgments ................................................................................................... 275 References ............................................................................................................. 275

Abstract During mitosis in vertebrate cells, the nuclear compartment is completely disintegrated in the process of nuclear envelope breakdown (NEBD). NEBD comprises the disassembly of nuclear pore complexes, disintegration of the nuclear lamina, and the retraction of nuclear membranes into the endoplasmic reticulum. Deciphering of the mechanisms that underlie these dynamic changes requires the identification of the involved molecular components and appropriate experimental tools to define their mode of action. Here, we describe an in vitro, imaging-based experimental system, which recapitulates NEBD. In our assay, we induce NEBD on nuclei of semi-permeabilized HeLa cells expressing fluorescently tagged nuclear envelope (NE) marker proteins by addition of mitotic cell extract that is supplemented with fluorescently labeled dextran. Time-lapse confocal microscopy is used to monitor the fate of the selected NE marker protein, and loss of the NE permeability barrier is deduced by influx of the fluorescent dextran into the nucleus. This in vitro system provides a powerful tool to follow NEBD and to characterize factors required for the reorganization of the NE during mitosis.

INTRODUCTION The genetic material of eukaryotic cells is enclosed by the nuclear envelope (NE) (Hetzer, Walther, & Mattaj, 2005; Schirmer, Florens, Guan, Yates, & Gerace, 2003), a double lipid bilayer perforated by large nuclear pore complexes (NPCs) that mediate nucleocytoplasmic transport of macromolecules. NPCs are each composed of about 30 nucleoporins (Nups) that occur in multiples of eight (Rothballer & Kutay, 2012a, 2012b; Suntharalingam & Wente, 2003). At the onset of mitosis in higher eukaryotic cells, the NE is dismantled in a process referred to as nuclear envelope breakdown (NEBD) (De Souza & Osmani, 2009). NEBD starts with the gradual dispersal of Nups into the cytoplasm leading to an increased permeability of the NE (Dultz et al., 2008). Further, the nuclear lamina is depolymerized and nuclear membrane components are redistributed into the mitotic endoplasmic reticulum (ER). Retraction of NE and ER membranes from chromatin is supported by microtubule-dependent forces (reviewed in Gu¨ttinger, Laurell, & Kutay, 2009).

Introduction

NPC disassembly and more generally NEBD have been originally investigated using in vivo models. By injecting large dextran dyes into the cytoplasm of starfish oocytes, Terasaki and colleagues assessed loss of the NE permeability barrier during meiotic maturation (Terasaki et al., 2001). In mammalian cells, NEBD has been mainly investigated in vivo by fluorescence microscopy of fluorophore-tagged NE proteins (Beaudouin, Gerlich, Daigle, Eils, & Ellenberg, 2002; Salina et al., 2002) (see also Chapter 10). Although these in vivo studies have extended our understanding of nuclear disassembly, in vitro systems provide powerful complementing tools to decipher mechanistic aspects of cellular processes. A few studies describing in vitro assays for NEBD have been previously reported. In 1998, Collas has used in vitro reconstituted nuclei incubated in mitotic sea urchin extract to investigate NE disassembly (Collas, 1998). Later, Ullman and colleagues employed in vitro assembled Xenopus laevis nuclei and egg extract (Liu, Prunuske, Fager, & Ullman, 2003). More recently, we have established a visual, in vitro nuclear disassembly assay using mitotic X. laevis egg extracts and somatic HeLa cell nuclei bearing a GFP-tagged NE protein (Mu¨hlhausser & Kutay, 2007), also described in Shankaran, Mackay, and Ullman (2013). Our in vitro NEBD assay relies on digitonin-permeabilized cells and exogenous cytosol—a system originally used to study nuclear transport in vitro (Adam, Sterne-Marr, & Gerace, 1992). Here, we describe the methodological details of an in vitro NEBD assay based on HeLa cell components, which recapitulates NPC disassembly at mitotic onset in human cells. In brief, HeLa cells stably expressing a GFP-tagged NE marker (here an NPC protein) are first semi-permeabilized with digitonin. Whereas the NE membrane stays intact upon treatment with a low concentration of digitonin, the plasma membrane gets perforated. Cytosolic components of these leaky cells are lost during subsequent washing steps. To induce NEBD, the semi-permeabilized cells are then incubated with mitotic HeLa cell extract, which is supplemented with fluorescent dextran serving as permeability marker for the NE. Loss of NE permeability and NPC disassembly are visualized by fluorescence microscopy, monitoring the gradual diffusion of dextran into the nuclei as well as the loss of fluorescent signal from the NE. Quantification of both processes is performed using an automated MatLab-based algorithm. This in vitro assay provides a powerful tool to study synchronized NEBD events in mammalian somatic cells, and offers the possibility to biochemically manipulate the system at will. Using this in vitro system, we have previously shown that phosphorylation of Nup98 is rate limiting for NPC disassembly at the onset of mitosis (Laurell et al., 2011). In this chapter, we describe our protocol to perform the in vitro NEBD assay using the example of HeLa cells expressing 2xGFP-Nup58 (Rabut, Doye, & Ellenberg, 2004) as NE marker and TRITC-labeled 155 kDa dextran to monitor loss of the permeability barrier (see Section 12.2). Additionally, we explain how to prepare crucial reagents, such as cell lines bearing fluorescent NE markers and mitotic HeLa S3 cell extract (see Sections 12.1.1 and 12.1.2). Further, we provide information for automated image analysis and data quantification (see Section 12.4). Finally, we specify how this system can be altered to assess the contribution of a specific factor to NEBD (see Section 12.3). Figure 12.1 summarizes the experimental workflow of the in vitro NEBD assay.

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FIGURE 12.1 Flowchart of the in vitro NEBD assay including preparatory steps. Dark gray boxes indicate key steps of the process. Optional steps are shown in white boxes.

12.1 PREPARATIVE STEPS 12.1.1 Generation of cell lines expressing a fluorescent NE marker Visualization of the NE during the NEBD assay requires a fluorescently tagged NE marker protein, which is homogeneously expressed in a stable cell line. The generation of such a cell line requires the integration of a DNA fragment encoding a fluorescently labeled NE protein into the multiple cloning site of a suitable vector, such as a bicistronic pIRES vector. pIRES plasmids possess an internal ribosome

12.1 Preparative Steps

entry site (IRES) that allows for the simultaneous expression of a protein of interest, for example, a fluorescently tagged NE protein, and an antibiotic resistance marker from a single mRNA. To generate a cell line, the respective plasmid is first transiently transfected into cells, and marker-positive cells are selected by the addition of an appropriate antibiotic. Generation of double- or triple-stable cell lines expressing various fluorescently labeled NE proteins may be accomplished by a combination of pIRES vectors containing different antibiotic resistances. The following example describes the generation of HeLa cells stably expressing 2xGFP-Nup58 selected by puromycin resistance. 1. Grow HeLa cells in DMEM supplemented with 10% FCS, 100 U/ml penicillin, and 100 mg/ml streptomycin (further referred to as DMEM) in a 100-mm tissue culture dish to 80–90% confluency in a humidified incubator at 37
C and 5% CO2. 2. Transiently transfect cells with 6 mg of plasmid DNA using X-tremeGENE transfection reagent according to the manufacturer’s instructions. 3. Split cells into five 100-mm dishes 24 h post transfection. 4. 24 h later, start the selection for positive clones by exchanging the medium with DMEM containing puromycin at a final concentration of 1 mg/ml (selection medium). 5. Replace selection medium every 3–4 days until single colonies are visible. 6. Pick and spread single colonies as follows: • Wash colonies with PBS. • Carefully add 2 ml of 0.5 mM EDTA (in PBS) to each dish and incubate 2 min at 37
C in the humidified CO2 incubator. • Carefully suck single colonies into the tip of a 200-ml pipette. • Resuspend each colony into a well of a 24-well plate containing 500 ml selection medium. 7. Expand cells into 6-well plates containing fresh selection medium as soon as they reach confluency. Additionally, transfer 5 104 cells/ml into a 24-well plate and assay your gene of interest by using an appropriate analysis method (e.g. fluorescence microscopy, Western blotting). 8. Choose positive colonies and expand the corresponding clones into 100-mm dishes. 9. Store selected cell lines by cryopreservation: • Detach cells by addition of 0.5 mM EDTA (in PBS). • Pellet the cell suspension at 1000 rpm for 5 min at 4
C. • Resuspend the cell pellet with 1 ml ice-cold FCS and 1 ml ice-cold DMEM (without 10% FCS, 100 U/ml penicillin, and 100 mg/ml streptomycin) containing 20% DMSO. • Transfer into cryovials and incubate on ice for 10 min. • Slowly freeze the cells at 80
C in a polystyrene box for 1–3 days. • Place the cryovials in liquid nitrogen for long-term storage.

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REMARKS! I. To generate a stable cell line, you may first need to determine the minimum concentration of antibiotic required to kill untransfected cells of the recipient cell line. We recommend testing a range of concentrations of the respective antibiotic beforehand. Untransfected cells should die within 4–5 days of antibiotics treatment. II. The expression level of NE marker may vary between positive colonies. Therefore, freeze a selection of positive clones with different expression levels. III. Depletion of the endogenous protein by RNAi may facilitate the integration of the overexpressed protein at the NE.

12.1.2 Mitotic extract 12.1.2.1 HeLa S3 cell culture For the preparation of mitotic HeLa cell extract (ME), we use HeLa S3 cells—a HeLa cell clone, which can be grown either adherently or in suspension. Suspension cell culturing offers the advantage of high cell yield from relatively low culture volumes. Note that this HeLa ME is not suitable to study microtubule (MT)-dependent processes, since the extract is prepared in the presence of the microtubuledepolymerizing agent nocodazole. For analyses of MT-dependent processes during NEBD, it is recommended to use CSF-arrested Xenopus egg extract (Mu¨hlhausser & Kutay, 2007). BUFFERS AND SOLUTIONS • Fixative: 3% paraformaldehyde, 0.1% TX-100—store at room temperature; add 1 mg/ml Hoechst (from a 10 mg/ml stock). 1. Culture HeLa S3 cells in RPMI medium supplemented with 10% fetal calf serum, 100 U/ml penicillin and 100 mg/ml streptomycin, and 1  “MEM Non-Essential Amino Acids” in a humid atmosphere at 37
C and 5% CO2. Grow cells in adherent culture and expand the cells every 2 days according to the scheme shown in Table 12.1.

Table 12.1 Expansion protocol for HeLa S3 cells Day

Procedure

Passage

1 3 5 7 9 10

Thaw S3 cells to 1  100-mm dish Transfer cells to 1  150-mm dish Transfer cells to 3  150-mm dish Transfer cells to 12  150-mm dish Transfer cells to 1 spinning flask (500 ml) for growth in suspension Add nocodazole to 100 ng/ml

p0 p1 p2 p3 p4

12.1 Preparative Steps

2. Harvest HeLa S3 cells from twelve 150-mm dishes. Transfer cells into a 1-l spinner flask containing 500 ml RPMI medium containing all supplements. Grow cells in a humid atmosphere at 37
C and 5% CO2 with constant stirring at 75 rpm (cellspin control unit). After 24 h, determine the cell number, for example, using a Neubauer counting chamber. 3. At a density of 0.8–1.2  106 cells/ml, add nocodazole (10 mg/ml stock, in DMSO) to a final concentration of 100 ng/ml and grow cells for another 24 h. When the cell density is higher than 1.2  106 cells/ml, dilute cells accordingly with medium. 4. To determine the mitotic index by Hoechst staining, transfer 1 ml of cell suspension into an Eppendorf tube, pellet cells (13,000 rpm, 5 min, tabletop centrifuge) and resuspend the cell pellet in 100 ml fixative supplemented with Hoechst. Transfer 5 ml of fixed cells onto a cover slip and inspect DNA condensation status based on Hoechst fluorescence by microscopy. Calculate mitotic index as the percentage of cells with condensed chromosome content. Cells should only be harvested for preparation of ME if the mitotic index is 95% or above.

12.1.2.2 Preparation of mitotic extract BUFFERS AND SOLUTIONS • ATP: 200 mM in H2O; adjust pH to 7.5 with NaOH; store at 80
C. • “Modified EBS”: 40 mM Na-b-glycerophosphate (from a 1 M stock in H2O, stored at 20
C), 15 mM MgCl2, 20 mM EGTA, 1 mM glutathione, 1 mM PMSF (from a 250 mM stock in ethanol), 10 mg/ml pepstatin A (from a 10 mg/ml stock in DMSO), 10 mg/ml aprotinin (from a 10 mg/ml stock in H2O), 10 mg/ml leupeptin. Prepare 100 ml prior to use. Keep at 4
C. • “Modified EBS (þATP)”: add ATP to a final concentration of 2 mM in 50 ml of modified EBS. To obtain ME of high quality, the following steps need to be performed as fast as possible. Cells and lysate have to be kept on ice throughout the whole procedure. Rotors and centrifuges need to be cooled in advance. 1. Transfer HeLa S3 cells into 500-ml conical centrifugation tubes, place tubes in a swinging bucket rotor, and centrifuge at 1000 rpm for 15 min at 4
C (swing-out rotor, Heraeus Multifuge 3S-R). 2. Resuspend cell pellet in 30 ml ice-cold modified EBS (w/o ATP) and centrifuge at 1000 rpm for 15 min (swinging bucket rotor, Heraeus Labofuge 400R; for this and the following centrifugation steps). 3. Estimate volume of the cell pellet (e.g., 3.2 ml). Resuspend cells in 50 ml of 33% ice-cold modified EBS (þATP). Incubate cell suspension on ice for 5 min and centrifuge for 15 min at 1000 rpm and 4
C. 4. Wash cells in 20 ml modified EBS (þATP) and centrifuge at 1000 rpm for 15 min.

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5. Resuspend cell pellet in 0.7 volumes (compared to the pellet volume estimated in step 3, e.g., 2.2 ml) of modified EBS (þATP) and incubate on ice for 5 min. Snap freeze cell suspension in liquid nitrogen. At this point, frozen cells can be stored at 80
C for several hours up to overnight. 6. For cell lysis, first slowly thaw frozen cells in lukewarm water and mix occasionally. Then, press cell suspension 3–5 times through a 27G needle using a 5-ml syringe. Avoid air bubbles. Transfer crude cell lysate into TLA-100.3 centrifuge tubes (3.5 ml, 13  51 mm) and spin for 5 min at 100,000 rpm (Beckman Optima TLX ultracentrifuge with TLA100.3 rotor). 7. Carefully remove the upper layer (membrane fraction) using a 27G needle attached to a syringe. Transfer the middle layer (cytosolic fraction) into a fresh TLA-100 tube. Discard the pellet (cell debris). 8. Centrifuge cytosolic fraction for 30 min at 100,000 rpm (Beckman Optima TLX ultracentrifuge with TLA100.3 rotor). 9. After this second centrifugation step, collect cytosolic supernatant into a 15-ml tube and estimate the volume. Add sucrose to a final concentration of 250 mM and aliquot the extract into PCR tube strips (we prepare 20, 60, and 100 ml aliquots). Snap freeze aliquots in liquid nitrogen and store at 80
C. 10. Store remaining modified EBS at 20
C. It may be used as washing buffer during protein depletion experiments as described in Section 12.3.2.2. REMARKS! For preparation of interphase cell extract, incubate cells in presence of 3 mM thymidine for 24 h (instead of using nocodazole). Again, the mitotic index is determined by visual inspection under the microscope. Cells should only be used for preparation of interphase extract if the mitotic index is close to zero.

12.1.3 Fluorescently labeled dextrans Dextrans are hydrophilic polysaccharides that are commercially available (e.g., from Sigma or Invitrogen) in a variety of sizes ranging from 3 to 2000 kDa. Dextran conjugates have relatively high stability, good water solubility, and low toxicity. They can be purchased either in a fluorescently labeled form or as amino derivates that can be labeled with one of the numerous fluorescent derivatives of succinimidyl esters. Fluorescent dextrans are ideal markers to visualize the gradual loss of the nuclear permeability barrier during NEBD. Since molecules smaller than 40 kDa can freely diffuse through NPCs (Suntharalingam & Wente, 2003), 70 kDa or larger dextrans should be used. The combination of differently labeled dextrans of varying size can be exploited to monitor different steps of NPC disintegration in the same experiment (Lenart et al., 2003). First, we describe the preparation of a solution of TRITC-labeled 155 kDa dextran (commercially available as powder from Sigma). Second, we provide information on dextran labeling, using the example of labeling of a 70 kDa aminodextran with Alexa Fluor 405 carboxylic acid succinimidyl ester (both from Invitrogen).

12.1 Preparative Steps

12.1.3.1 Preparation of 155 kDa TRITC-dextran 1. Dissolve the lyophilized dextran in permeabilization buffer (see Section 12.2.2 for buffer composition) to a final concentration of 25 mg/ml. Sonication and heating to 45
C for 5 min may increase solubility of the dextran conjugate. Vortex solution briefly. 2. Remove insoluble dextran particles by centrifugation at 13,000 rpm (tabletop centrifuge, Eppendorf ) for 5 min at 4
C. 3. Store aliquots at 80
C.

12.1.3.2 Labeling of a 70 kDa aminodextran with Alexa Fluor 405 carboxylic acid succinimidyl ester BUFFERS AND SOLUTIONS • Amine-free reaction buffer (50 mM HEPES pH 7.5, 150 mM KOAc). • 1.5 M hydroxylamine chloride pH 8.5 (prepare freshly). 1. Dissolve lyophilized dextran powder in amine-free reaction buffer to a final concentration of 10 mg/ml (143 mM). To increase solubility, you may briefly sonicate the solution or incubate it at 45
C for 5 min. 2. Pellet insoluble particles by centrifugation in a tabletop centrifuge at 13,000 rpm for 5 min at 4
C. Store supernatant at 80
C or proceed to labeling reaction. 3. Dissolve 1 mg Alexa Fluor 405 succinimidyl ester (Invitrogen) in 100 ml DMSO to a final concentration of 10 mM. The dye can be stored at 80
C. 4. For labeling, supplement 0.5 mg dextran solution with 4–5 equivalents (3.5 ml) of Alexa Fluor 405 active ester and incubate the reaction for 1 h at room temperature by constant stirring and protected from light. 5. Stop labeling reaction by adding 25 mM hydroxylamine chloride to the dextrandye solution and incubate it for further 15 min. 6. Remove free dye by gel filtration, for example, by using a NAP-5 column: • Equilibrate the column with permeabilization buffer. • Load dextran conjugate. • Elute with permeabilization buffer and collect fractions of 4 drops each. • Pool the fractions from the first fluorescent peak. 7. Store fluorescently labeled dextran at 80
C. REMARKS! Dextran conjugates can be checked for size homogeneity and, if required, further purified by gel filtration using a Superdex 200 16/60 column (GE Healthcare).

12.1.4 Energy mix As NEBD requires the activity of enzymes, including NTPases, the ME needs to be supplemented with an energy-regenerating system (energy mix). A 20  energy mix is prepared as follows:

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1. Prepare a solution of 200 mM creatine phosphate, 10 mM GTP, and 10 mM ATP in 50 mM HEPES pH 7.0. 2. Add 1 mg/ml creatine kinase. 3. Add 250 mM sucrose. 4. Snap freeze and store 20 energy mix in small aliquots at  80
C. Avoid multiple freeze–thaw cycles.

12.2 NEBD ASSAY In vitro NEBD assays are performed using semi-permeabilized HeLa cells stably expressing 2xGFP-Nup58. Nuclear disassembly is triggered by the addition of ME supplemented with energy mix and fluorescently-labeled dextran. Progression through NEBD is monitored by time-lapse confocal microscopy and quantified using a MatLab-based application (Fig. 12.2). In the next sections, we describe how to perform and analyze in vitro NEBD experiments.

12.2.1 Cell seeding Cells stably expressing 2xGFP-Nup58 are seeded onto an Ibidi imaging chamber 12–24 h before the NEBD assay. Ibidi chambers contain 18 wells with a volume capacity 50 ml. Seeding cells at an appropriate density is crucial for the following permeabilization step. While a high cell density might reduce the permeabilization efficiency, too low cell density results in overpermeabilization or detachment of the cells. 1. Place an Ibidi chamber in a 100-mm sterile dish, remove the lid of the chamber, and keep it in the 100-mm dish. 2. Detach 2xGFP-Nup58 cells and determine the cell concentration.

FIGURE 12.2 Scheme illustrating the in vitro assay for NEBD. HeLa cells expressing 2xGFP-Nup58 are semi-permeabilized with digitonin. Addition of mitotic extract (supplemented with energy mix and TRITC-labeled dextran) leads to NPC disassembly, allowing for dextran influx. Disassembly of the NE is monitored by confocal laser-scanning microscopy.

12.2 NEBD Assay

3. Seed 3000–5000 cells per well in a total volume of 40–50 ml. The final cell number should be tested empirically. The optimal cell number depends on the growth rate of the stable cell line. 4. Allow cells to attach. Grow cells for 12–24 h in a humidified incubator at 37
C with 5% CO2.

12.2.2 Semi-permeabilization of cells Before proceeding with the in vitro NEBD assay, 2xGFP-Nup58 cells need to be semi-permeabilized. During this process, the cell membrane is selectively perforated by treatment with a low concentration of the nonionic detergent digitonin, while the nuclear membrane remains intact. Soluble cytoplasmic proteins are removed from the cells during subsequent washing steps and are then replaced by untreated or modified ME (see Section 12.3). BUFFERS AND SOLUTIONS (PREPARE FRESHLY) • Ice-cold permeabilization buffer: 20 mM KOH–HEPES (pH 7.5), 110 mM KOAc, 5 mM Mg(OAc)2, 250 mM sucrose, and 0.5 mM EGTA; prepare 10 ml. • Ice-cold permeabilization buffer supplemented with digitonin: prepare 5 ml of permeabilization buffer with 0.002–0.003% digitonin (according to cell density). Work on ice during the entire procedure. Transfer the 100-mm dish containing the Ibidi chamber on a metal block, which is placed on ice. To prevent any cell detachment during the permeabilization and washing steps, pipette solutions carefully at the edge of the wells. 1. Remove medium and wash cells with 35 ml of ice-cold PBS. 2. Remove PBS, add 30 ml permeabilization buffer supplemented with digitonin, and incubate for 10 min. 3. Wash cells with 30 ml permeabilization buffer (without digitonin) successively for 0, 2, 5, and 10 min.

12.2.3 NEBD reaction mix In parallel to semi-permeabilization of cells, prepare the NEBD reaction mix (shown in Table 12.2) as follows: Table 12.2 Pipetting scheme for NEBD reaction mix Component

Volume (ml)

ME Permeabilization buffer TRITC-labeled 155 kDa dextran 20  Energy mix

20.00 2.75 1.00 1.25

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1. Supplement the ME with dextran and 1  energy mix. Prepare a total volume of 25 ml for each well. The final concentration of dextran should be tested empirically; 0.5 mg/ml dextran is a good starting concentration. 2. Add 18 ml of supplemented ME to semi-permeabilized cells, place the lid back onto the Ibidi imaging chamber, and start image acquisition (see Section 12.2.4). This time point is defined as “time 0.” 3. Snap freeze remaining samples of ME for further analysis (e.g., kinase assay, Western blot) and store these aliquots at 80
C.

12.2.4 Image acquisition NEBD is monitored by influx of fluorescent dextran into nuclei, which reflects the gradual loss of the NE permeability barrier. Simultaneously, the release of 2xGFPNup58 from the NE can be followed. This section describes how NEBD is microscopically recorded over time.

12.2.4.1 Microscopic setup and acquisition settings Laser-scanning microscopy of NEBD assays is performed using a 40 water immersion objective mounted on an inverted confocal microscope (Zeiss LSM710 FCS). The system is equipped with an Argon laser, a DPSS 561 laser, and a temperaturecontrolled incubator box. NEBD assays are performed at 25
C to slow down the kinetics of the reactions. For stable imaging conditions, it is crucial to preequilibrate the incubation chamber of the microscope to 25
C for at least 30 min prior to imaging. Unstable temperature will result in drifts of the focal plane. Acquisition parameters are selected using the ZEN 2010 software (Carl Zeiss MicroImaging, Inc.): • • • •

•

• •

Time lapse: 4 min; 15–25 cycles, depending on the stable cell line, the quality of the ME, and the treatment applied (e.g., protein depletion). Multi-positioning: a minimum of 3 positions per well is acquired. Z-stack: for each position, 3 Z-slices are acquired with an interval distance of 1.3 mm. The Z-stack is centered according to the selected focal plane. Excitation and detection windows: – GFP: excitation at 488 nm; detection window between 500 and 560 nm – TRITC: excitation at 561 nm, detection window between 580 and 680 nm Laser intensity and photomultiplier sensitivity are optimized to obtain a good image quality without inducing photobleaching damages, which could possibly affect NEBD kinetics. Additional settings: GFP and TRITC channels are acquired sequentially (channel mode, switch track every line) at a speed of 7 (pixel dwell time: 3.15 ms). Image parameters: pixel size: 0.26 mm; image size: 134.7  134.7 mm; bit depth: 8 bit; frame size: 512  512.

12.3 Special Treatments

12.2.4.2 Image acquisition After addition of ME to semi-permeabilized cells, proceed directly to image acquisition: 1. Add water immersion medium to the objective and place Ibidi chamber on the slide holder. Moisten the wells to be imaged by moving the objective along the Ibidi chamber for several rounds. Water immersion medium evaporates more rapidly than the usual immersion oil does. Apply a sufficient amount of immersion medium. 2. Adjust the focal plane of the first well. 3. Using the 488-nm laser (green channel), select 3–4 positions per well according to the following criteria: • Appropriate cell density • Normal nuclear shape • Low number of cells without GFP signal • Low number of cells closely juxtaposed to each other 4. Select appropriate positions for the remaining wells. 5. Fine-tune the focal plane of each position. The NE should appear as a sharp ring. 6. Center the Z-stack at this Z-plane and start image acquisition. Since NEBD usually occurs between 30 and 50 min after addition of ME to the cells, image acquisition should start at the latest 20 min after adding the ME. An example of a NEBD assay using 2xGFP-Nup58 HeLa cells treated with mitotic or interphase cell extract is shown in Fig. 12.3.

12.3 SPECIAL TREATMENTS In general, protein function during NEBD can be addressed by modification of the reporter cell line and/or by modifying the ME.

12.3.1 Modification of reporter cells 12.3.1.1 Overexpression of nucleoporin mutants To directly study the function of a NE protein, fluorescently tagged NE markers are stably expressed in a cell line of choice. NEBD kinetics of cells expressing a mutant NE protein (e.g., phospho-mimetic or phospho-deficient mutant) can be compared to those expressing the wild-type protein.

12.3.1.2 Knockdown of endogenous protein Cell lines bearing a NE marker can be subjected to RNAi for two purposes: 1. To enhance the incorporation of recombinant NE marker into the NE after siRNA-mediated depletion of the endogenous protein.

267

FIGURE 12.3 An in vitro NEBD experiment. (A) In vitro NEBD reactions were performed by addition of interphase or mitotic HeLa cell extracts to semi-permeabilized HeLa cells stably expressing 2xGFP-Nup58. Loss of the NE permeability barrier (influx of TRITC-labeled 155 kDa dextran) was monitored by time-lapse confocal laser-scanning microscopy. Representative images of different time points are shown. (B) Quantification of dextran-positive nuclei over the time course of the experiments. (C) Quantification of the intensity of 2xGFP-Nup58 at the NE over the time course of the experiment. (D) In vitro kinase assay using histone H1 phosphorylation as readout for CDK1 kinase activity.

12.3 Special Treatments

2. To investigate the function of a protein that is a component of the nuclei of the reporter cell line. To deplete such factor, cells should be treated with siRNAs prior to the NEBD assay. Cells are usually subjected to RNAi for 48 or 72 h using 10 or 20 nM siRNA, optimized for the protein of interest. SiRNA transfection is performed in 6-well plates, for example, using INTERFERin® as a transfection reagent. One day prior to the in vitro NEBD assay, RNAi-treated cells are harvested and seeded into Ibidi imaging chambers.

12.3.2 Modification of cell extract ME can be modified in two different ways: chemical inhibition of a protein’s function (e.g., for enzymes such as kinases) or antibody-mediated depletion of a protein from the extract. Specificity of effects observed after protein depletion is verified by add-back of the respective recombinant protein, which should restore NEBD to control kinetics.

12.3.2.1 Chemical inhibitors For chemical inhibition of mitotic kinases, small-molecule inhibitors such as the PLK1 inhibitor BI2536 (100 mM stock in DMSO) or the CDK1 inhibitor RO3306 (10 mM stock in DMSO) are added directly to the ME (e.g., 500 nM final concentration for BI2536 and 1–10 mM final concentration for RO3306) and incubated at room temperature for 10–15 min. To keep the DMSO concentration in the extract low, inhibitors should preferably be prediluted in permeabilization buffer and not in DMSO. A solvent control reaction is strongly recommended. The incubation of ME with inhibitors is performed in parallel to the permeabilization of cells. The extract–inhibitor mixture is supplemented with fluorescently labeled dextran and 1  energy mix prior to its addition to the semi-permeabilized cells.

12.3.2.2 Protein depletion and add-back Efficient protein depletion requires the availability of several milligrams of a highly specific antibody. This antibody is then covalently coupled to protein A/G beads (9:1 mixture; GE Healthcare) using 2 mg of antibody per 1 ml A/G beads. Antibodycoupled A/G beads are stored at 4
C. Protein depletion from ME has to be performed directly prior to the NEBD assay. In general, we use a 1:6 ratio of antibody-coupled A/G beads to extract and perform two subsequent rounds of depletion. Depending on the abundance of protein and the quality of the antibody, this protocol has to be adapted for each case (e.g., additional rounds of depletion, extended depletion time, different “beads-to-extract” ratio). 1. For depletion of 60 ml of ME, wash 20 ml of antibody-coupled beads (or DMPcrosslinked A/G beads as control) in 1 ml of modified EBS. Centrifuge 1 min at 1000 rpm (tabletop centrifuge, Eppendorf ) and discard supernatant. Repeat once. Resuspend antibody-coupled beads in 1 ml EBS and centrifuge 500 ml of beads suspension through a Mobicol spin column. Insert plug to the bottom of the column.

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Table 12.3 Pipetting scheme for NEBD after protein depletion. Component

Volume (ml)

Depleted ME Recombinant protein (or permeabilization buffer) TRITC-labeled 155 kDa dextran 20 Energy mix

20.00 2.75 1.00 1.25

2. Thaw 60 ml of ME and add it to the spin column containing antibody-coupled or A/G (control) beads. 3. Add lid on top of the column, place it into an 1.5-ml Eppendorf tube, and incubate for 15 min at 4
C with gentle agitation. Ensure that beads and extract are well mixed. (We perform this incubation step at 900 rpm in a thermomixer placed in the cold room.) 4. Centrifuge “1-depleted” extract into a fresh tube (1000 rpm, 1 min). Prepare a fresh spin column by centrifuging another 500 ml of a preequilibrated antibody-coupled or A/G beads suspension through a Mobicol spin column. Again, insert plug to bottom of the column. 5. Add “1-depleted” extract to the freshly prepared spin column, screw cap, and incubate another 15 min for a second round of depletion. 6. Centrifuge the “2-depleted” extract into a fresh tube and immediately apply in the NEBD assay (see Table 12.3 for a typical NEBD reaction mixture after depletion). 7. For rescue experiments, add the recombinant protein in one or several concentrations (in permeabilization buffer) to the depleted extract. REMARKS! Depletion efficiency for each experiment should be controlled by Western blotting. Take a 2 ml aliquot of ME before and after depletion and mix it with 10 ml of Laemmli buffer. In addition, elute bound proteins from EBS-washed beads by incubation with Laemmli buffer for 5 min. For depletion of kinases, residual enzymatic activity can be determined in a kinase assay. Use 1 ml of the respective NEBD reaction mix (see Section 12.2.3) for a 10 ml total sample volume, containing 100 mM ATP, 1 mCi 32P-g-ATP, and 2–5 mg of kinase-specific substrate (e.g., H1 for CDK1/B1, casein for NEK kinases) in permeabilization buffer. Incubate reaction for 10–15 min at 30
C, add Laemmli buffer, and subject samples to gel electrophoresis. Dry the Coomassie-stained gel and expose it to a phospho-imaging screen.

12.4 DATA ANALYSIS In this section we describe a software to quantitatively analyze the progression of NEBD in the in vitro assay (http://www.highcontentanalysis.org/). The implementation of algorithms and graphical interfaces is in MatLab (The MathWorks, Inc., Natick, MA, USA).

12.4 Data Analysis

12.4.1 Computational steps The analysis consists of the following computational steps: 1. 2. 3. 4.

Drift correction of the image sequences Detection of cell nuclei Statistical measurements Reporting

Here, we describe the details of the computational steps.

12.4.2 Drift correction of the image sequences Due to multi-position acquisition, the image sequences sometimes have a stage drift of a few pixels. This drift is due to the mispositioning of the stage motors, which only causes misalignment parallel to the x and y image axes. Without loss of generality, we can assume only translational errors, and ignore rotation, shearing, and any other nonlinear deformations. We also assume that cell migration is uniform and significantly smaller than the stage drift between consecutive images. For image sequences that satisfy these two criteria, we calculate the phase correlation (Kuglin & Hines, 1975; Riess et al., 2012) between consecutive image pairs and determine the drift shift between them with subpixel resolution (see Fig. 12.4A).

12.4.3 Detection of cell nuclei Detection of the exact location and morphology of nuclei is the most crucial step of the analysis. There are two inherent difficulties. First, nuclei cannot be detected after NEBD because they are flooded by fluorescent dextran. Second, detection of cell nuclei only on the first time frame (and the use of the corresponding analysis masks on subsequent images) creates noisy segmentation and imprecise object boundaries, due to cell migration and confocality. To overcome this problem, we use the mean projection of the first several images (in practice 4, but it is an adjustable parameter in the software) and detect the nuclei on this projection. The first step of the detection is an automated Otsu thresholding. This is followed by filling holes and removing small objects (in practice we remove objects smaller than 51.50 mm2 (300 pixels)). The location of the NE is then approximated by a few pixels wide ring. To separate touching cells, Voronoi tessellation is used (Gonzalez & Woods, 2002). Finally, cells and the corresponding rings are uniquely labeled and a graphical interface is displayed to the user to validate and manually remove objects (see Fig. 12.4).

12.4.4 Statistical measurements For every nucleus, the mean fluorescence intensities in the nuclear interior and at the nuclear rim (NE) are monitored over time. The nuclear fluorescence intensity can either be measured in the entire segmented area (Fig. 12.4B, green regions), or in the center of mass of the nuclei in a user-defined square or circular area. The intensities are normalized using the background image, which is determined in the inverse of the thresholded masks. Loss of the NE permeability barrier, that is NEBD, is

271

FIGURE 12.4 Depiction of the computational steps performed by the data analysis software. The image processing algorithm consists of three steps (A–C). (A) Alignment of the image sequence to account for the paraxial movement of the microscopic object. This registration is based on phase correlation (left). An example for a significant positioning error between consecutive images is indicated by a red arrow (middle). Intensity difference maps before and after correction (right). (B) Segmentation of cell nuclei in the first image of the sequence using the Otsu adaptive thresholding method (left). Nuclei that cannot be separated are excluded by filtering out segments that are too large (not shown). Small objects are excluded based on a size criterion (inset, middle). Nuclear intensity is measured in the entire segmented area (green regions). (C) Time course of in vitro nuclear disassembly. Images of selected time points are shown. The influx of TRITC-labeled 155 kDa dextran into segmented nuclei is visualized. Intact nuclei are shown in green, and nuclei classified as dextran-positive are colored in red (mean fluorescence intensity ratio between inside and outside of nucleus is larger than 0.3).

12.6 Material and Reagents

defined as the time point when the fluorescent intensity in the nuclear region reaches 30% of the fluorescence intensity of the labeled dextran outside of nuclei (Fig. 12.4C). This value can be defined by the user in the algorithm. The nuclear envelope intensities are measured on the GFP image (see Fig. 12.4B). Since the size of the NE is subpixel size, it is very critical to determine its exact position. In practice, using thresholding and other classical edge-based methods proved unreliable. Therefore, we do not measure the intensity directly on the image, but rather on its grayscale dilated version, which extends bright intensities around a defined kernel size and shape, in a sort of “maximum filter.” By using this and assuming that the NE is the brightest object within the template’s extension, the algorithm is robust against slight segmentation mistakes. The intensities are then measured and normalized to the first frame.

12.4.5 Reporting As a final step, different measurement reports and visual outputs are created. The single cell-based mean intensity measurements and their standard deviations, as well as the NEBD event statistics, are saved to comma-separated text files. A visual overview file is created for every experiment, which shows the original image series, the segmentation, and the NE detection results. In addition, graphs displaying the fluorescence intensity ratio over time are generated for the analyzed nuclei.

12.5 FUTURE DIRECTIONS We have developed a robust, imaging-based in vitro system to study NEBD using human somatic cell extracts and semi-permeabilized cells expressing fluorescent NE marker proteins. Our in vitro assay bears several advantages that could prove as an essential asset to the analysis of NEBD in vivo. The addition of cell extract at a given time point (fixed reference point) and the combination of different fluorescent NE marker proteins enable to follow the timing and kinetics of NPC disassembly. This is of particular interest when studying a very rapid and synchronous process like the dispersal of nucleoporins from the NE. Moreover, the use of a fully activated mitotic extract allows to dissect protein functions independently of their role, for example, in regulating mitotic entry. The system can be easily altered biochemically by depleting or adding factors of interest. Thus, it could also prove as a versatile tool to identify new factors involved in NEBD, and thereby pave the way for reconstitution of the process.

12.6 MATERIAL AND REAGENTS 12.6.1 Material (listed in alphabetical order) • • •

40 Water immersion objective (C-Apochromat 40 /1.2 W Korr M27) Argon laser (Lasos Lasertechnik GmbH, RMC 7812 Z1) Cellspin control unit (IBS Integra Biosciences, 183013)

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• • • • • • • • • • • • • • • • • • • •

Cover slips (12 mm) and microscopic slides (Thermo Scientific) Ibidi chamber, u-Slide 18 well flat (Ibidi, 81826) Illustra NAP-5 column (GE Healthcare, 17-0853-01) Incubation chamber (incubator controller, EMBL GP168) Inverted confocal microscope (Zeiss LSM710 FCS) Labofuge 400R (Heraeus, 75008375) Mobicol spin column (MoBiTec) Multifuge centrifuge 3S-R (Heraeus, 75004375) Needle 27G (B. Braun, 9186174) PCR strips (Thermo Scientific, AB-114) Screw cap conical bottom centrifuge tubes (Corning, 431123) Small filter—35 mm pore size (MoBiTec, M513515) Spinning flask (IBS Cellspin1000, Integra Biosciences) Swing-out rotor (Heraeus, 75006445) Swinging bucket rotor (Heraeus, 75008172) Syringe 5 ml (Codan, 62.5607) Tabletop centrifuge 5415D (Eppendorf, 022621408) Tissue culture dishes (100 mm), 6-well plates, 24-well plates (Greiner Bio-One) TLA-100.3 centrifuge tubes (open-top, polycarbonate, thickwall, 3.5 ml, 13  51 mm; Beckman, 349622) Ultracentrifuge, optima TLX (Beckmann, A95761)

12.6.2 Reagents (listed in alphabetical order) • • • • • • • •

• • •

Amino-dextrans of different sizes (Invitrogen or Sigma-Aldrich), storage at 80
C Aprotinin (10 mg/ml stock, Applichem, A2132) Dextran (Sigma, T1287) Digitonin (5% stock solution, storage at 20
C) Dimethyl sulfoxide (DMSO, Sigma, 41640) DMEM (Sigma, D6429) EDTA (Sigma, E9884) Energy mix (20 ), storage at  80
C • creatine phosphate, 400 mM dissolved in H2O, storage at 80
C (Sigma, P7936) • creatine kinase, 50 mg/ml dissolved in 50 mM HEPES pH 7.0, 250 mM sucrose (Roche, 10127566001) • GTP, 100 mM in H2O, storage at 80
C (Sigma, G8877) • ATP, 200 mM in H2O, storage at 80
C (Sigma, A2383) FCS (PAA Laboratories GmbH, A15-101) Fluorescent succinimidyl ester dyes (e.g., Alexa Fluor 405 succinimidyl ester, Invitrogen, A30000) Glutathione (Sigma, 64251)

References

• • • • • • • • • • • • • • • • • •

Hoechst (10 mg/ml stock solution, Sigma, 861405) Immersion medium Immersol W 2010, oiler 20 ml (Carl Zeiss, 444969-0000-000) INTERFERin (Polyplus Transfection, 409-10) Leupeptin (Applichem, A2183) MEM Non-Essential Amino Acids (NEAA 100 , Gibco, 11140-035) Mitotic cell extract (ME), storage at  80
C Nail polish Opti-MEM I reduced serum medium (Gibco, 31985047) Paraformaldehyde (PFA, Sigma, P6148) PBS (137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 1.4 mM KH2PO4, pH 7.4) Penicillin/streptomycin (100 , PAA Laboratories GmbH, P11-010) Puromycin (Sigma, P9620) RPMI 1640 Medium þ GlutaMax 1  (Gibco,61870-010) Sucrose (stock solution 2.5 M, Sigma, S9378) Thymidine (Sigma, T9250) Triton X-100 (Sigma, 93426) Vectashield mounting medium (Vector Laboratories, H-1000) X-tremeGENE 9 transfection reagent (Roche, 06 365 787 001)

Acknowledgments We thank Petra Mu¨hlha¨usser and Eva Laurell, two former PhD students in the laboratory, who have invested tremendously into the development of the in vitro NEBD assay. Our work on NEBD is supported by a grant of the ERC to U. K.

References Adam, S. A., Sterne-Marr, R., & Gerace, L. (1992). Nuclear protein import using digitoninpermeabilized cells. Methods in Enzymology, 219, 97–110. Beaudouin, J., Gerlich, D., Daigle, N., Eils, R., & Ellenberg, J. (2002). Nuclear envelope breakdown proceeds by microtubule-induced tearing of the lamina. Cell, 108(1), 83–96. Collas, P. (1998). Nuclear envelope disassembly in mitotic extract requires functional nuclear pores and a nuclear lamina. Journal of Cell Science, 111(Pt. 9), 1293–1303. De Souza, C. P., & Osmani, S. A. (2009). Double duty for nuclear proteins—The price of more open forms of mitosis. Trends in Genetics, 25(12), 545–554. Dultz, E., Zanin, E., Wurzenberger, C., Braun, M., Rabut, G., Sironi, L., et al. (2008). Systematic kinetic analysis of mitotic dis- and reassembly of the nuclear pore in living cells. The Journal of Cell Biology, 180(5), 857–865. Gonzalez, R., & Woods, R. (2002). Digital image processing (2nd ed.). Upper Saddle River, NJ: Prentice Hall. Gu¨ttinger, S., Laurell, E., & Kutay, U. (2009). Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nature Reviews. Molecular Cell Biology, 10(3), 178–191.

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Hetzer, M. W., Walther, T. C., & Mattaj, I. W. (2005). Pushing the envelope: Structure, function, and dynamics of the nuclear periphery. Annual Review of Cell and Developmental Biology, 21, 347–380. Kuglin, C. D., & Hines, D. C. (1975). The phase correlation image alignment method. In Conference on cybernetics and society. Laurell, E., Beck, K., Krupina, K., Theerthagiri, G., Bodenmiller, B., Horvath, P., et al. (2011). Phosphorylation of Nup98 by multiple kinases is crucial for NPC disassembly during mitotic entry. Cell, 144(4), 539–550. Lenart, P., Rabut, G., Daigle, N., Hand, A. R., Terasaki, M., & Ellenberg, J. (2003). Nuclear envelope breakdown in starfish oocytes proceeds by partial NPC disassembly followed by a rapidly spreading fenestration of nuclear membranes. The Journal of Cell Biology, 160(7), 1055–1068. Liu, J., Prunuske, A. J., Fager, A. M., & Ullman, K. S. (2003). The COPI complex functions in nuclear envelope breakdown and is recruited by the nucleoporin Nup153. Developmental Cell, 5(3), 487–498. Mu¨hlhausser, P., & Kutay, U. (2007). An in vitro nuclear disassembly system reveals a role for the RanGTPase system and microtubule-dependent steps in nuclear envelope breakdown. The Journal of Cell Biology, 178(4), 595–610. Rabut, G., Doye, V., & Ellenberg, J. (2004). Mapping the dynamic organization of the nuclear pore complex inside single living cells. Nature Cell Biology, 6(11), 1114–1121. Riess, T., Marino, J., Wandke, C., Merhof, D., Deussen, O., Csucs, G., et al. (2012). Image analysis of nuclear envelope breakdown events using KNIME. In: Proceedings of the 9th international workshop on computational systems biology (WCSB’12), Ulm, Germany. Rothballer, A., & Kutay, U. (2012a). SnapShot: The nuclear envelope I. Cell, 150(4), 868, e861. Rothballer, A., & Kutay, U. (2012b). SnapShot: The nuclear envelope II. Cell, 150(5), 1084, e1081. Salina, D., Bodoor, K., Eckley, D. M., Schroer, T. A., Rattner, J. B., & Burke, B. (2002). Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell, 108(1), 97–107. Schirmer, E. C., Florens, L., Guan, T., Yates, J. R., 3rd., & Gerace, L. (2003). Nuclear membrane proteins with potential disease links found by subtractive proteomics. Science, 301(5638), 1380–1382. Shankaran, S. S., Mackay, D. R., & Ullman, K. S. (2013). A time-lapse imaging assay to study nuclear envelope breakdown. Methods in Molecular Biology, 931, 111–122. Suntharalingam, M., & Wente, S. R. (2003). Peering through the pore: Nuclear pore complex structure, assembly, and function. Developmental Cell, 4(6), 775–789. Terasaki, M., Campagnola, P., Rolls, M. M., Stein, P. A., Ellenberg, J., Hinkle, B., et al. (2001). A new model for nuclear envelope breakdown. Molecular Biology of the Cell, 12(2), 503–510.

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13

Modern Tools to Study Nuclear Pore Complexes and Nucleocytoplasmic Transport in Caenorhabditis elegans

Peter Askjaer*, Vincent Galy{,}, and Peter Meister} *

Andalusian Center for Developmental Biology (CABD), CSIC/Junta de Andalucı´a/Universidad Pablo de Olavide, Carretera de Utrera, Seville, Spain { Sorbonne Universite´s, UPMC, Univ Paris 06, UMR7622, IBPS, F-75005 Paris, France } CNRS, UMR7622, IBPS, F-75005 Paris, France } Cell fate and Nuclear Organization, Institute of Cell Biology, University of Bern, Baltzerstrasse 4, Bern, Switzerland

CHAPTER OUTLINE Introduction ............................................................................................................ 278 13.1 Forward and Reverse Genetics .........................................................................279 13.1.1 RNAi ......................................................................................... 285 13.1.2 Construction of RNAi Clone ......................................................... 286 13.1.3 Preparation of RNAi Plates.......................................................... 286 13.1.4 C. elegans RNAi Feeding............................................................. 286 13.1.5 Materials ................................................................................... 287 13.2 Transgenesis ..................................................................................................287 13.3 Live Imaging of Embryos..................................................................................290 13.3.1 Sample Preparation .................................................................... 295 13.3.2 How to Limit Phototoxicity .......................................................... 295 13.3.3 Materials ................................................................................... 296 13.4 In Vivo Methods to Evaluate Structural and Functional Integrity of the NE ........................................................................................................296 13.4.1 Preparation of Fluorescently Labeled Dextran for Germline Injection ................................................................................. 298 13.5 Immunofluorescence and Electron Microscopy .................................................298 13.5.1 Immunofluorescence .................................................................. 299 13.5.2 Transmission Electron Microscopy ............................................... 302 13.6 Interaction of Nups with Chromatin ..................................................................303

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00013-8

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Summary and Future Perspectives............................................................................ 304 Acknowledgments ................................................................................................... 304 References ............................................................................................................. 305

Abstract The nematode Caenorhabditis elegans is characterized by many features that make it highly attractive to study nuclear pore complexes (NPCs) and nucleocytoplasmic transport. NPC composition and structure are highly conserved in nematodes and being amenable to a variety of genetic manipulations, key aspects of nuclear envelope dynamics can be observed in great details during breakdown, reassembly, and interphase. In this chapter, we provide an overview of some of the most relevant modern techniques that allow researchers unfamiliar with C. elegans to embark on studies of nucleoporins in an intact organism through its development from zygote to aging adult. We focus on methods relevant to generate loss-of-function phenotypes and their analysis by advanced microscopy. Extensive references to available reagents, such as mutants, transgenic strains, and antibodies are equally useful to scientists with or without prior C. elegans or nucleoporin experience.

INTRODUCTION About half a century ago, Sydney Brenner decided to use Caenorhabditis elegans, a little (
1 mm) free-living soil nematode, to identify genes responsible for animal behavior and morphology (Brenner, 1974). C. elegans became immediately a popular model organism as transparency allowed observing organ development and muscle activity in the intact organism with noninvasive techniques. Careful observations by Sulston and colleagues uncovered that the cell lineage of C. elegans is invariant, implying that all individuals contain a fixed number of cells (959 somatic cells in the adult hermaphrodite) stereotypically dividing and positioned within the body (Sulston & Horvitz, 1977; Sulston, Schierenberg, White, & Thomson, 1983). The life cycle of C. elegans proceeds quickly: development from egg through four larval stages (L1–L4) to fertile adult takes approximately 3 days. Moreover, C. elegans reproduces mainly as a self-fertilizing hermaphrodite and under optimal conditions a single adult will produce
250 progeny over a time course of 3–5 days. For genetic crosses, male animals can be induced by temperature shifts or genetic tricks, allowing classical genetic interaction studies (e.g., complementation or double mutants). C. elegans embryos are particularly well suited for analysis of mitotic processes, cell differentiation, and morphogenesis with high temporal and spatial resolution. Surrounded by a resistant eggshell, they can easily be mounted for long time-lapse observation from the zygote to the fast-moving threefold embryo ready to hatch. The first mitotic division is asymmetric, producing two daughter cells of unequal size, composition, and fate. Genetic screens based on this feature have

13.1 Forward and Reverse Genetics

identified numerous conserved factors involved in polarity establishment through control of mitotic spindle positioning (reviewed in Gonczy, 2008; McNally, 2013). C. elegans can be grown on solid media or in liquid culture, which makes it ideal for high-throughput forward or reverse genetic studies. Genome-wide RNAi screens have for instance uncovered roles of C. elegans nucleoporins (nups; in C. elegans nomenclature also known as Nuclear Pore Proteins or NPPs) in diverse processes such as transposon silencing (Vastenhouw et al., 2003), germ granule distribution (Updike & Strome, 2009) sensitivity to ionizing radiation (van Haaften et al., 2006). Moreover, the rapid life cycle of C. elegans facilitated the striking observation that certain nups are expressed only during embryogenesis and larval development and remain stably integrated in NPCs during the entire lifespan of the animal (D’Angelo, Raices, Panowski, & Hetzer, 2009). The above characteristics combined with the fact that most nups (Table 13.1) and transport factors (Table 13.2) are conserved in C. elegans and that the nuclear envelope disassemble and reassemble at each round of cell division make this organism attractive to study nuclear pore complexes and nucleocytoplasmic transport in a physiological, multicellular but simple in vivo context. Tables 13.1 and 13.2 list both general and specific phenotypes ascribed to individual proteins and provide an overview of mutant alleles and available reagents. Scientists who are not familiar with C. elegans are encouraged to also consult two recent volumes of Methods in Cell Biology dedicated to this model system (vol. 106–107) as well as WormBook available at http://www.wormbook.org.

13.1 FORWARD AND REVERSE GENETICS C. elegans has been extensively used to conduct genetic experiments. Most screens used forward genetics strategies in which animals are mutagenized using chemical or physical DNA-damaging agents. Mutagenized progeny is then screened for the phenotype of interest. Traditionally, positional mapping through genetic crosses, a process that can last several years, was used to identify mutated genes. Forward genetics has recently regained interest with the possibility to rapidly characterize mutations using highthroughput sequencing (Doitsidou, Poole, Sarin, Bigelow, & Hobert, 2010; Zuryn, Le Gras, Jamet, & Jarriault, 2010). An alternative method to forward genetics is ‘reverse’ screening, in which genes are knocked down individually by RNA interference (RNAi). The creation of genome-wide RNAi libraries made reverse genetic screens easy and cost-efficient (Kamath & Ahringer, 2003; Rual et al., 2004). Reverse approaches also allow fast and independent confirmation of mutations obtained using forward genetic strategies. While reverse genetic screens are usually faster, forward genetic screens can uncover temperature-sensitive mutations or reduction of function mutants. Strains carrying deletions for individual genes are available for approximately one third of all protein coding genes. These deletions have mainly been obtained using mutagenesis and PCR-based deletion screening of large strain libraries (http:// www.shigen.nig.ac.jp/c.elegans/, http://celeganskoconsortium.omrf.org/). Such a

279

Table 13.1 C. elegans nucleoporins Frequent phenotypesa

Specific phenotypes

Emb; Lva; Lvl; Nmo; Pgl; Stp Clr; Emb; Lva; Nmo; Pgl; Pvl; Stp Clr; Emb; Lva; Nmo; Pgl; Ste

Spindle orientation; RNAi efficiency NPC assembly; synthetic lethal with NPP-5, -14, -15, -17 NPC exclusion limit; spindle orientation; timing of mitosis Spindle orientation; transposon silencing

Worm

Human

NPP-1

NUP54

NPP-2

NUP85

NPP-3

NUP205

NPP-4

NUPL1

Emb; Stp

NPP-5

NUP107

Emb; Pgl

NPP-6

NUP160

NPP-7

NUP153

NPP-8

NUP155

NPP-9

NUP358

Emb; Lva; Lvl; Nmo; Pgl Emb; Nmo; Lva; Pgl; Ste Emb; Lva; Lvl; Nmo; Pgl; Pvl Emb; Nmo; Pgl; Pvl

Interaction with spindle assembly checkpoint; kinetochore assembly NPC assembly

NPC assembly RNAi efficiency; spindle assembly;

Mutant allelesb

Reagentsc

Reference

FP; Y2H

Kim et al. (2005) and Schetter, Askjaer, Piano, Mattaj, and Kemphues (2006) Galy, Mattaj, and Askjaer (2003) and Rodenas, Gonzalez-Aguilera, Ayuso, and Askjaer (2012) Galy et al. (2003), Hachet et al. (2012), and Schetter et al. (2006)

tm2199

FP

ok1999

Abs; Y2H

ok617

FP; Y2H

ok1966; tm3039

Abs; FP

ok2821; tm4329 ok601

FP

tm2513

Abs; FP

Abs; FP

Abs; FP

Franz et al. (2005), Schetter et al. (2006), Updike, Hachey, Kreher, and Strome (2011), and Vastenhouw et al. (2003) Franz et al. (2005) and Rodenas et al. (2012)

D’Angelo et al. (2009) and Rodenas et al. (2012) D’Angelo et al. (2009), Galy et al. (2003), and Voronina and Seydoux (2010) Franz et al. (2005) Askjaer, Galy, Hannak, and Mattaj (2002), Kim et al. (2005), Sheth, Pitt, Dennis,

NPP10Nd NPP10Cd NPP-11

NUP98

NPP-12

NUP210

NPP-13

NUP93

Emb; Nmo; Pgl

NPP-14

NUP214

wt

NPP-15

NUP133

Lvl

NPP-16

NUP50

wt

NPP-17/ RAE-1

RAE1

Emb; Hya; Pvl; Ste; Stp

NPP-18

SEH1

wt

NUP96 NUP62

Emb; Lva; Lvl; Nmo; Pgl; Ste Emb; Lva; Lvl; Nmo; Ste Emb; Lva; Lvl; Nmo Emb; Lva

nuclear envelope formation NPC assembly; P granule integrity NPC assembly

Abs

and Priess (2010), and Voronina and Seydoux (2010) Galy et al. (2003), Rodenas et al. (2012), and Voronina and Seydoux (2010) Galy et al. (2003) and Rodenas et al. (2012)

ok467

Spindle orientation

ok1599

FP; Y2H

Schetter et al. (2006)

Nuclear envelope breakdown

ok2424; tm2320

Abs

NPC exclusion limit; spindle orientation; timing of mitosis Synthetic lethal with NPP-2 Sensitivity to ionizing radiation RNAi efficiency; anoxia-induced prophase arrest Axon termination and synapse formation

ok1534

Abs; Y2H

Audhya, Desai, and Oegema (2007), Cohen, Feinstein, Wilson, and Gruenbaum (2003), and Galy et al. (2008) Galy et al. (2003), Hachet et al. (2012), and Schetter et al. (2006)

Abs; FP

ok1389 ok1954

Galy et al. (2003) Abs; FP

D’Angelo et al. (2009), Rodenas et al. (2012), and van Haaften et al. (2006) Hajeri, Little, Ladage, and Padilla (2010) and Kim et al. (2005)

Aff; FP

Grill et al. (2012)

ok1839; tm1596 ok1720; tm2784; tm2796

Continued

Table 13.1 C. elegans nucleoporins—cont’d Worm

Human

NPP-19

NUP35

NPP-20

SEC13R

NPP-21

TPR

NPP-22 / NDC-1

NDC1/ TMEM48

NPP-23 MEL-28

NUP43 ELYS/ AHCTF1

Frequent phenotypesa

Specific phenotypes

Mutant allelesb

Emb; Nmo; Pgl; Stp Emb; Lva; Lvl; Nmo; Pgl; Stp Clr; Emb; Lva; Ste Clr; Emb; Lva; Lvl; Nmo; Ste

NPC assembly

tm2886

Regulation of tumor growth and apoptosis NPC assembly; modification of dynein activity

tm1541; tm2952 tm1845

NPC assembly; spindle assembly

tm2434; t1578; t1684

wt Emb; Lva

Reagentsc

Reference

Abs; FP; Y2H

Rodenas et al. (2012) and Rodenas, Klerkx, Ayuso, Audhya, and Askjaer (2009)

Pinkston-Gosse and Kenyon (2007) Abs; FP

FP Abs; FP

O’Rourke, Dorfman, Carter, and Bowerman (2007) and Stavru et al. (2006) Rodenas et al. (2012) Fernandez and Piano (2006) and Galy, Askjaer, Franz, Lopez-Iglesias, and Mattaj (2006)

No clear C. elegans homologues were found for the mammalian nups AAAS/ALADIN, NUP37, NUP88, NUP188, NUPL2/hCG1, and POM121. a Gross phenotypes, which for most genes were reported in large-scale RNAi studies. See Galy et al. (2003) and WormBase (http://www.wormbase.org) for details and references. Clr, clear/transparent body; Emb, embryonic lethal; Hya, hyper active; Lva, larval arrest; Lvl, larval lethal; Nmo, (pro-)nuclear morphology alteration in early embryo; Pgl, P-granule abnormality; Pvl, protruding vulva; Ste, sterile; Stp, sterile progeny; wt, wild type. Abnormal P granule distribution (Pgl) was observed for many npp genes (Updike & Strome, 2009; Voronina & Seydoux, 2010). b Only selected alleles are listed. These and other alleles are available from the Caenorhabditis Genetics Center (CGC; University of Minnesota; http://www.cbs.umn.edu/CGC/) and the National Bioresource Project for the Experimental Animal “Nematode C. elegans” (Tokyo Women’s Medical University School of Medicine; http://www.shigen.nig.ac.jp/c.elegans/index.jsp). c Abs, antibodies; Aff, expression of affinity-tagged protein; FP, expression of fluorescently tagged protein; Y2H, plasmids to study yeast two hybrid interactions. d Because NPP-10N and NPP-10C are produced from a single protein precursor, a given RNAi phenotype will generally reflect the combined effect of depleting both proteins. P granule phenotypes are, however, specific to NPP-10N depletion.

Table 13.2 C. elegans transport receptors and Ran GTPase-associated proteins Worm

Human a

Frequent phenotypesa

Specific phenotypes

Mutant alleles

Reagents

Reference

wt

Silencing of repetitive DNA

gk200

Abs; Aff

Nuclear envelope formation; spindle assembly NPC assembly; RNAi efficiency; silencing of repetitive DNA Nuclear envelope formation; spindle assembly Redox-dependent nuclear import; RNAi efficiency

ok256

Abs; Aff

Geles and Adam (2001) and Robert, Sijen, van Wolfswinkel, and Plasterk (2005) Askjaer et al. (2002) and Geles, Johnson, Jong, and Adam (2002)

ok715

Abs; Aff

Geles and Adam (2001), Kim et al. (2005), and Robert et al. (2005)

Abs

Askjaer et al. (2002), Fernandez and Piano (2006), and Ikegami and Lieb (2013) Kim et al. (2005) and Putker et al. (2013)

IMA-1

KPNA

IMA-2

KPNA

Emb; Lvl; Nmo

IMA-3

KPNA

Emb; Lvl; Nmo; Pgl

IMB-1

KPNB1/ IMB1

Emb; Nmo

IMB-2 IMB-3 IMB-4/ XPO-1

TNPO1/ TRN RANBP6 XPO1/ CRM1

Emb; Lva; Lvl; Stp Emb; Lva; Ste Emb; Lva; Lvl; Pgl; Ste

IMB-5/ XPO-2 IMB-6/ XPO-3

CSE1L/ CAS XPOT/ XPO3

Emb; Lva; Lvl; Nmo; Pgl; Pvl wt

tm6328

Abs

ok1795 b-Catenin nucl. export; miRNA biogenesis; mRNA export

Chromosome segregation; RNAi efficiency Sensitivity to ionizing radiation

tm1437; tm1889

Bussing, Yang, Lai, and Grosshans (2010), Kuersten, Segal, Verheyden, LaMartina, and Goodwin (2004), Nakamura et al. (2005), and Updike and Strome (2009) Kim et al. (2005) Updike and Strome (2009), Walther et al. (2003) van Haaften et al. (2006) Continued

Table 13.2 C. elegans transport receptors and Ran GTPase-associated proteins—cont’d Frequent phenotypesa

Worm

Human

RAN-1

RAN

Emb; Lva; Lvl; Nmo; Pgl; Ste

RAN-2

RANGAP1

Emb; Nmo; Ste

RAN-3

RCC1

Emb; Lvl; Nmo; Pvl; Ste

RAN-4

NUTF2/ NTF2 RANBP3

Emb; Lva; Nmo; Pgl; Ste Nmo

RAN-5

Specific phenotypes b-Catenin nucl. export; Eph receptor trafficking; nuclear envelope formation; spindle assembly Nuclear envelope formation; SMN complex component: spindle assembly; b-Catenin nucl. export; nuclear envelope formation; RNAi efficiency

Mutant alleles

Reagents

Reference

tm5197

Abs; Aff

Askjaer et al. (2002), Bamba, Bobinnec, Fukuda, and Nishida (2002), Cheng, Govindan, and Greenstein (2008), Nakamura et al. (2005)

ok1939; tm3590

Y2H

Askjaer et al. (2002), Bamba et al. (2002), and Burt, Towers, and Sattelle (2006)

ok3709

FP

Askjaer et al. (2002), Bamba et al. (2002), Kim and Yu (2011), and Nakamura et al. (2005) Updike and Strome (2009)

tm1439 b-Catenin nucl. export

Nakamura et al. (2005)

Annotated C. elegans importins, exportins, and Ran GTPase-related proteins. Other transport factors, such as RXF-1, -2, ALY-1, -2, -3, etc., are omitted due to space constraints. See Table 13.1 for column legends. a The homology between C. elegans IMA proteins and human importin alpha (KPNA) proteins is insufficient to make pair-wise assignments (Geles et al., 2002).

13.1 Forward and Reverse Genetics

strategy has the intrinsic drawback that the rest of the genome is mutagenized too. Mutants therefore have to be outcrossed extensively to wild-type strains to clean the genetic background. This remains very inefficient for linked mutations and a few background ‘equilibrating’ mutations can persist over a number of backcrosses. To overcome this and create targeted short deletions, the newly developed CRISPR-Cas9 system has been adapted for use in C. elegans (Chiu, Schwartz, Antoshechkin, & Sternberg, 2013; Friedland et al., 2013; Lo et al., 2013; Waaijers et al., 2013). Cas9 is an RNA-guided nuclease, which induces a sequence-specific double strand break that when repaired by nonhomologous end joining creates small insertions and deletions. The efficiency appears high enough that the technique is likely to become routine in the next years. Moreover, when oligonucleotides spanning over the break site are coinjected with CRISPR-Cas9, the repair machinery is able to use these as a template for repair, enabling the creation of targeted point mutations in the genome.

13.1.1 RNAi The discovery of RNAi as means to efficiently knockdown expression of individual genes (Fire et al., 1998) increased dramatically the popularity of C. elegans. Despite that RNAi-based tools are now available in many systems, C. elegans is still an attractive organism to induce and study loss-of-function phenotypes (Simpson, Davis, & Boag, 2012). RNAi efficiency is remarkably high and persists through generations due to an endogenous amplification step mediated by RNA-dependent RNA polymerases found in nematodes, fungi, and plants. Moreover, RNAi in C. elegans spreads throughout the body (although with a lower effect in the nervous system) causing a systemic knockdown. Finally, C. elegans can easily be cultivated in liquid in 96-well plates, thus facilitating genome-wide scale studies. Three methods have been developed to introduce the triggering double-stranded RNA (dsRNA) in C. elegans: (1) injection of dsRNA into the nematodes, (2) soaking of the nematodes in a solution containing dsRNA, and (3) feeding bacteria expressing the dsRNA to the nematodes. The injection and soaking methods both require in vitro dsRNA synthesis and purification and the former also necessitates dedicated microinjection equipment and training. The feeding method relies on simple cloning and microbiology techniques and is easily scaled up in terms of quantity of nematodes and/or number of genes to be analyzed. Moreover, bacteria clones can be amplified and distributed inexpensively. We therefore focus here on the feeding method, but the reader should keep in mind that the efficiency in some cases have been found to be slightly inferior to injection. Hence, if by feeding an expected phenotype is not observed or if quantification shows an incomplete depletion, injection, or soaking should be considered. The protocol below is designed to analyze a few genes. An in-depth discussion of methods and protocols for large-scale RNAi screening is available (Cipriani & Piano, 2011). Bacteria expressing dsRNA corresponding to the gene(s) to be analyzed can be generated by standard molecular biology techniques, obtained from a C. elegans laboratory that possesses the relevant clone(s) or, for most annotated genes, purchased

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CHAPTER 13 Analysis of C. elegans Nuclear Pore Complexes

(e.g., Source BioScience [http://www.lifesciences.sourcebioscience.com] or Thermo Scientific [http://www.thermoscientificbio.com]).

13.1.2 Construction of RNAi clone 1. Design PCR primers to amplify a 300- to 1000-bp fragment of your gene. If you use genomic DNA as template place the primers to get maximum exon content. Alternatively, use cDNA as template. BLAST your fragment against the C. elegans genome (e.g., at http://www.wormbase.org) to ensure that the fragment does not target other genes (avoid matching sequences >20 nt). Add restriction sites to the primers to facilitate insertion of the fragment between the two convergent T7 promoters of the vector pPD129.36 L4440 (available from http://www.addgene.org or any C. elegans lab). 2. Restriction digest, purify, and ligate PCR fragment(s) and vector. 3. Transform the ligation reaction into standard bacteria and plate on LB-Amp plates. 4. Miniprep DNA from several colonies and check for correct insert. 5. Transform HT115(DE3) bacteria with miniprep DNA. Include also a reaction with pPD129.36 to use as negative RNAi control. HT115(DE3) is RNase IIITet þ and can be obtained from the Caenorhabditis Genetics Center (http://www. cgc.cbs.umn.edu). Plate on LB-Amp-Tet (optional: using carbenicillin instead of ampicillin may improve RNAi efficiency due to higher stability of this antibiotics).

13.1.3 Preparation of RNAi plates 1. Inoculate 3 ml LB-Amp with a single colony from a fresh LB-Amp-Tet plate and incubate at 37  C, 200 rpm until OD600 ¼ 0.6–0.8 (
8 h). Alternatively, grow bacteria overnight. 2. Add 1 mM IPTG to the bacteria culture just before seeding the plates. 3. Seed NGM plates containing 1 mM IPTG and 100 mg/ml ampicillin or 20 mg/ml carbenicillin with 100 ml of bacteria culture. 4. Incubate at room temperature overnight and use immediately or store at 16  C for a few days.

13.1.4 C. elegans RNAi feeding 1. In the days prior to the experiment check that worms are growing well without suffering starvation or contamination. 2. Transfer worms to an empty NGM plate and leave for 30–60 min remove bacteria sticking to the cuticle. Select L4 larvae for analysis of RNAi-depleted embryos. 3. Transfer 10–15 L4s to RNAi plates (optional: after the worms have moved away from the spot on the RNAi plate where they were placed, put 5 ml of 50% household bleach/1 N NaOH solution to kill any bacteria transferred with the worms).

13.2 Transgenesis

4. Incubate plates for 16–48 h at 16–25  C. For several genes RNAi efficiency has been observed to be temperature-dependent so it is advisable to test several conditions. Likewise, kinetics of protein synthesis and turnover will influence the rate of RNAi-mediated depletion. 5. Dissect worms to obtain embryos for live imaging as described below. To study postembryonic phenotypes adult worms should be transferred to fresh RNAi plates every 12–24 h to generate semisynchronous populations of offspring.

13.1.5 Materials C. elegans genomic DNA as PCR template For 1-few reactions, sufficient DNA can be obtained by disrupting 5–10 worms in a PCR tube containing 2.5 ml lysis buffer. Overlay with a drop of mineral oil to prevent evaporation. Incubate at 80  C for 15 min; 60  C for 1 h; 95  C for 15 min. Add 22.5 ml PCR mixture to PCR tube with lysed worms (for amplification of a single target sequence) or dilute and split lysed worms in 2–5 PCR tubes (for several target sequences). Worm lysis buffer Tris/HCl 10 mM pH 8.3 KCl 50 mM MgCl2 2.5 mM NP-40 0.45% Tween-20 0.45% Gelatin 0.01% Add 1 ml proteinase K (10 mg/ml; Sigma P6556) to 99 ml of worm lysis buffer immediately before use NGM plates NaCl 50 mM Agar 1.7% Peptone 0.25% Autoclave and add sterile Cholesterol (5 mg/ml in EtOH) 1 ml/L CaCl2 1 mM MgSO4 1 mM Potassium phosphate pH 6.0 25 mM For RNAi add 1 mM IPTG and 100 mg/ml ampicillin or 20 mg/ml carbenicillin

13.2 TRANSGENESIS Transgenesis in C. elegans has greatly improved in the last few years, offering a variety of methods with different advantages and limitations (Table 13.3). The simplest and most frequently used transgenes are extrachromosomal arrays.

287

Table 13.3 Comparison of transgene types Transgene type Size Insertion site Chromatin state Mitotic and meiotic stability Expression level Germline/early embryo expression Nature of exogenous DNA Mode of transgenesis Genome mutagenic load Price Difficulty/workload

Extrachromosomal arrays

Integrated extrachromosomal array

Bombarded transgenes

Homologous recombinationa

Very large (>100 copies) Extrachromosomal Heterochromatic Variable

Very large (>100 copies) Random insertion Heterochromatic 100% stable

From 1 to 100 copies Random insertion Variable 100% stable

Single insertion Targeted insertion Variable 100% stable

Usually high to very high expression levels Usually silenced

Usually high to very high expression levels Usually silenced

Mixture of plasmids

Uses an extrachromosomal array Irradiation (UV/X-ray)

Low/middle expression levels Variable (promoter dependent) Mixture of plasmids Bombardment

Low (endogenous) expression levels Usually expressed (insertion site dependent) Mixture of plasmids/homologous recombination template Injection

Low (backcrossing recommended)

Low (backcrossing recommended)

Expensive Labor intensive (worm amplification)

Inexpensive Labor intensive (large number of injections)

Injection None

Inexpensive Easy and fast

High (extensive backcrossing recommended) Inexpensive Labor intensive (selection and backcrossing)

a Homologous recombination through repair of a double-stranded DNA break can be initiated either by Mos1 transposase or RNA-guided Cas9 nuclease activity. Insertion site is based on the presence of Mos1 transposons in the genome or design of small guide RNA molecules. See text for details.

13.2 Transgenesis

Extrachromosomal arrays are created by injection of DNA inside the worm gonad. Injected DNA is concatenated into an array which stability varies both mitotically and meiotically, depending on its sequence composition and complexity. Transgenic worms are identified either by a fluorescent or phenotypic marker or by the rescue of a mutation. Due to their high copy number, expression from arrays is usually very high (overexpressed) and in the case of fluorescent proteins readily observed with a dissecting scope. However, arrays are usually poorly expressed in the germline and early embryos, most likely because of their highly repetitive nature and their heterochromatic structure. A number of strategies can be employed to increase complexity, improve stability, and germline expression of the arrays, for instance by coinjecting heterologous DNA. Arrays can also be integrated into the genome by irradiation and screening for stable transmission. The method for germline injection is extensively described (Berkowitz, Knight, Caldwell, & Caldwell, 2008). An alternative to injected arrays is microparticle bombardment (Hochbaum, Ferguson, & Fisher, 2010; Praitis, 2006). Gold beads coated with DNA are shot at worms and occasionally the bombarded DNA gets randomly integrated into the genome. Germline integration is recognized by 100% transmission to the progeny and/or rescue of a phenotypically screenable mutation. Transgene arrays obtained by bombardment are smaller in size, ranging from 1 to 100 copies and less subject to germline silencing. The drawbacks of the bombardment method is the cost of the device and consumables as well as the difficulty to check for homozygozity when crossing strains as the insertion site is most often unknown. To avoid insertion-site artifacts and better control copy number, site-specific integration of transgenes using homologous recombination was recently developed. These methods are based on induction of a site-specific DNA double strand break through the action of either Mos1 transposase or Cas9 nuclease. In Mos1-mediated single-copy insertion (MosSCI) and Mos1 excision induced transgene-instructed gene conversion (MosTIC), the transposase is expressed in a strain harboring a characterized insertion of the Drosophila transposon Mos1 (Frokjaer-Jensen, Davis, Ailion, & Jorgensen, 2012; Frokjaer-Jensen et al., 2008; Robert & Bessereau, 2011). More than 13,000 individual Mos1 insertion sites have been isolated, which in principle can all be used for genetic engineering (Vallin et al., 2012). The CRISPR-Cas9 method allows even more flexibility because the system can be designed to cleave the genome specifically at any site that contains a G/A(N)19NGG sequence motif (Chen, Fenk, & de Bono, 2013; Dickinson, Ward, Reiner, & Goldstein, 2013; Katic & Grosshans, 2013; Tzur et al., 2013). In both methods, a transgene surrounded by sequences homologous to the flanking sequences of the break is injected into the germline and serves as template for recombination repair. Depending on the design of the repair template, the methods can be used to introduce transgenes into intergenic regions of the genome or to tag endogenous loci. To discriminate between homologous recombination and the creation of an array, additional markers are injected, which can later be counter-selected. Detailed protocols for both methods, including tools to identify suitable CRISPR-Cas9 target sequences, are available online

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(e.g., https://sites.google.com/site/jorgensenmossci/; http://wormcas9hr.weebly.com; http://crispr.mit.edu). Although Mos1- and CRISPR-Cas9-mediated homologous recombination offer the advantage of mimicking endogenous expression levels, detection of weakly expressed fluorescent constructs may be challenging. When transgenes should be expressed only in some cells, a number of characterized promoters are available for expression in differentiated somatic tissues. Expression in these cells is usually not an issue, although using heterologous, cell-type specific promoters might lead to overexpression. The germline and the early embryo remain the most difficult tissues to express transgenes, due to mechanisms defending the genome against exogenous DNA. As stated above, arrays are usually not expressed at that stage, while bombarded and single-copy transgenes have variable expression success. Promoters that have been shown proficient for expression in the germline include pie-1 and mex-5 (Merritt, Gallo, Rasoloson, & Seydoux, 2010; Zeiser, Frokjaer-Jensen, Jorgensen, & Ahringer, 2011). Other promoters (e.g., housekeeping genes, such as tbb-1, baf-1, his-72) can also efficiently drive expression in the germline, but get silenced more often.

13.3 LIVE IMAGING OF EMBRYOS Live imaging of early C. elegans embryos is a powerful way to examine the consequences of an RNAi depletion or mutation of a given nuclear pore complex (NPC) component by DIC microscopy (Galy et al., 2003; Sonnichsen et al., 2005) or fluorescence microscopy (Galy et al., 2003). Many different read-outs allow detecting defects in protein nuclear import, nuclear envelope (NE), and NPC integrity (Askjaer et al., 2002; D’Angelo et al., 2009; Galy et al., 2003), and NPC distribution (Lee, Gruenbaum, Spann, Liu, & Wilson, 2000) (Tables 13.1, 13.2, 13.4, and 13.5). Furthermore, imaging of embryos expressing NPC or NE components fused to green fluorescent protein (GFP) provide information about the timing of their insertion during postmitotic NE formation (Franz et al., 2005) as well as their turnover at the NE (Galy et al., 2006). Importantly, validation of these fluorescent reporters as reliable reporters for their endogenous counterparts is facilitated by the availability of knockout alleles in public repositories (see Section 13.1.1 and Tables 13.1 and 13.2). Fertilization naturally occurs inside the worm body every
20 min in each of the two gonad arms. This is followed within half an hour by the completion of meiosis, pronuclear formation, meeting, and centration and finally the first mitotic cell division (Fig. 13.1). Completion of meiosis is coordinated with the production of the chitin-containing eggshell that protects and isolates the embryo from the external environment and physical constrains (Olson, Greenan, Desai, Muller-Reichert, & Oegema, 2012). This timing is important to consider since drugs and lipophilic fluorescent dyes are able to enter the embryo only prior to egg-shell synthesis simply by soaking early meiotic embryos. Alternatively, drugs and dyes can be injected into the gonads of the hermaphrodite a few hours before collecting the embryos for imaging (Galy et al., 2003). For exposure of embryos at a given time-point, the egg-shell may

13.3 Live Imaging of Embryos

Table 13.4 Useful fluorescent markers Strain

Reporter

Tissue

Comments

Reference

BN46

GFP:: NPP-19

Embryos and germ line

Rodenas et al. (2009)

BN69

GFP:: NPP-5

Embryos and germ line

GZ264

GFP:: PCN-1

JH1327

PIE-1:: GFP

Embryos and oocytes Embryos and oocytes

GFP::NPP-19 (NUP35) accumulates in germ line and embryonic NEs; endogenous npp-19 is mutated but rescued by GFP::NPP-19 expression GFP::NPP-5 (NUP107) accumulates in germ line and embryonic NEs; endogenous npp-5 is mutated but rescued by GFP::NPP-5 expression; co-expression of mCherry::HIS58 GFP::PCN-1 (PCNA) accumulates in all embryonic nuclei PIE-1::GFP accumulates in the germ line blastomer where it is imported into the nucleus

MR164

NLS:: GFP GFP:: LEM-2

Intestine

OD139

YFP:: LMN-1

Embryos and germ line

PS3808

NLS:: GFP:: LacZ GFP:: TBB-2

Uterus, vulva and neurons Embryos and germ line

OD83

WH204

Embryos and germ line

Bright signal; strains carries also lin-35 mutation GFP::LEM-2 accumulates in germ line and embryonic NEs; co-expression of mCherry::HIS58 YFP::LMN-1 (lamin) accumulates in germ line and embryonic NEs; co-expression of mCherry::HIS-58 Bright signal; expression restricted to specific cell types within these tissues Bright expression of GFP::TBB-2 (b-tubulin); useful to analyze nuclear exclusion of soluble tubulin

Rodenas et al. (2012)

Brauchle, Baumer, and Gonczy (2003) Reese, Dunn, Waddle, and Seydoux (2000) Kostic and Roy (2002) Audhya et al. (2007)

Audhya et al. (2007)

Gupta and Sternberg (2002) Strome et al. (2001)

Nonexhaustive list of strains expressing NE or nuclear import markers. All strains are available from the Caenorhabditis Genetics Center (CGC; University of Minnesota; http://www.cbs.umn.edu/CGC/).

physically be disrupted by applying a gentle pressure on the embryo (Gonczy et al., 2001) or by perm-1 RNAi, which when combined with immobilized embryos allows precisely timed drug inhibitions (Carvalho et al., 2011). In the latter study, a custombuild microdevice consisting of an array of wells (300 mm  300 mm and 150 mm deep) was used to exchange a neutral medium for a drug-containing medium (or vice

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Table 13.5 Subset of relevant antibodies Antigen

Codea

Typeb

IFc

WBc

Actin

C4

Mm

þ

EMR-1

3272

Mp

þ

3598

Rp

þ

FG nups

MAb414

Mm

þ

þ

GFP

D153-3

Rm

þ

þ

IMB-1

SDQ4154; 4155 Q3891; Q4051

Lp*

þ

3597

Rp

þ

SDQ2349

Lp*

3932

Lp*

þ

þ

LMN1

Mm

þ

þ

BUD3

Lp*

þ

þ

MEL28

Lp*

þ

NPP-3

SY1539; 1540

Lp*

þ

NPP-5

SG4839

Lp*

þ

þ

NPP-9

P5A6

Mm

þ

þ

SDQ3854

Lp*

þ

MEL-28

Abcam, Covance MBL þ

þ

LMN-1

Source

Reference

MP Biomedicals

Lp*

LEM-2

IPc

þ

þ

þ

Novus Biologicals Novus Biologicals

Novus Biologicals þ The Developmental Studies Hybridoma Bank

Gruenbaum, Lee, Liu, Cohen, and Wilson (2002) Gruenbaum et al. (2002) Galy et al. (2003) Rohner et al. (2013) Ikegami and Lieb (2013) Ikegami, Egelhofer, Strome, and Lieb (2010) Gruenbaum et al. (2002) Ikegami and Lieb (2013) Gruenbaum et al. (2002) Hadwiger, Dour, Arur, Fox, and Nonet (2010) Galy et al. (2006) Fernandez and Piano (2006) Hachet et al. (2012) and Ikegami and Lieb (2013) Rodenas et al. (2012) Sheth et al. (2010)

þ

Novus Biologicals

13.3 Live Imaging of Embryos

Ab#1; Ab#2

Cp

þ

þ

GBJQ

Lp

þ

þ

NPP-10C

GBLC

Lp

þ

þ

NPP-12

NPP12

Lp

þ

þ

NPP-13

SDQ3897; 4094 JL00007

Lp*

NPP13

Lp*

þ

Lp*

þ

þ

NPP-19

SDQ3896; 4093 OWYL

Lp*

þ

þ

SUN-1

41970002

Lp*

þ

þ

a-Tubulin

DM1a

Mm

þ

þ

12G10

Mm

þ

AB18251 KT23

Lp* Mm

þ þ

NPP-10N

NPP-16

Unknown NE antigen

þ

Novus Biologicals

þ

Lp*

þ

Voronina and Seydoux (2010) Galy et al. (2003) Galy et al. (2003) Galy et al. (2008) Ikegami and Lieb (2013) Ikegami and Lieb (2013) Hachet et al. (2012)

Novus Biologicals Rodenas et al. (2009) Novus Biologicals Sigma The Developmental Studies Hybridoma Bank Abcam The Developmental Studies Hybridoma Bank

Askjaer et al. (2002) Fernandez and Piano (2006)

Takeda, Watanabe, Qadota, Hanazawa, and Sugimoto (2008)

Nonexhaustive list of antibodies that efficiently recognize C. elegans antigens. Please refer to Tables 13.1 and 13.2 for a complete listing of NE-related proteins for which antibodies are available. a Code of serum, antibody batch, or monoclonal description. b Cp, guinea pig polyclonal; Lp, rabbit polyclonal; Lp*, affinity purified rabbit polyclonal; Mm, mouse monoclonal; Mp, mouse polyclonal; Rm, rat monoclonal; Rp, rat polyclonal. c IF, immunofluorescence; WB, western blot; IP, immunoprecipitation.

versa) in
1 min with constant observation of the samples (see Carvalho et al., 2011 for details). Drugs that have been successfully utilized in C. elegans through any of these delivery methods include nocodazole to destabilize microtubules (10–100 mg/ml, Sigma M1404; Gonczy et al., 2001), latrunculin A to inhibit actin (10 mM, Sigma

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FIGURE 13.1 Time-lapse observation of nucleoporin dynamics. Selected still images from confocal timelapse recording of C. elegans npp-5 (NUP107) mutant embryo expressing a rescuing GFP:: NPP-5 fusion protein (green) and mCherry::HIS-58 (hisH2B; magenta) (Rodenas et al., 2012). Boxed regions in the merged panels are shown at higher magnification to the left. Embryo is mounted with anterior to the left. (A) GFP::NPP-5 localize to separating chromosomes in meiosis I (anterior) and to sperm-derived chromatin (posterior,
5 mm from cortex); (B) GFP::NPP-5 accumulates on kinetochores during meiosis II; (C) NE assembly around sperm-derived chromatin; (D) NE assembly around oocyte-derived chromatin; (E) juxtapositioning of fully grown pronuclei; (F) NE breakdown and accumulation of GFP::NPP-5 at kinetochores of condensing chromosomes in prometaphase; (G) GFP::NPP-5 localizes to kinetochores of holocentric chromosomes, which are arranged as two straight lines facing the mitotic spindle poles in metaphase; (H) signal of GFP::NPP-5 spreads to cover chromosomes in anaphase; (I) enrichment of GFP::NPP-5 at the nuclear periphery during NE reassembly in telophase; and (J) nuclear growth in interphase. Bars, 5 mm.

13.3 Live Imaging of Embryos

L5163; Carvalho et al., 2011), the proteasome inhibitor c-lactocystin-ß-lactone (20 mM, Calbiochem 426102; Carvalho et al., 2011), and the XPO-1 (CRM1) inhibitor leptomycin B (1 ng/ml, Sigma L2913; Pushpa, Kumar, & Subramaniam, 2013).

13.3.1 Sample preparation Embryos are extracted by dissection of gravid hermaphrodites in a drop of appropriate physiological buffer. Embryos that have completed meiosis (>30 min post fertilization) at the time of dissection will survive in M9 buffer, while earlier embryos devoid of a mature egg-shell are sensitive to physical and osmotic pressure and require meiosis buffer or egg salts to develop. 1. Using a worm pick, transfer one to five young adult gravid hermaphrodites to a 25-ml drop of buffer to remove bacteria. 2. Transfer the worms to a 5-ml drop of buffer on a 22-mm square glass coverslip. 3. Cut the worms in the middle using thin forceps (Dumont N 5) and a syringe needle (26 GA) to release the embryos from the uterus. 4. Collect gently the embryos by approaching and contacting the drop of buffer with a freshly made wet agarose pad (2% agarose in water or meiosis buffer lacking FBS) on a slide. 5. Seal the coverslip using melted VALAP. 6. For optimum temperature control during the live imaging, glue the slide onto a fast response temperature controller using a thin layer of vacuum grease. Recordings are typically performed at 20  C to minimize the risk of local overheating but the temperature controller may be set up to temperatures ranging from 16 to 25  C and allows rapid temperature shifts to analyze temperaturesensitive mutants (Gorjanacz et al., 2007). However, when DIC imaging is required we recommend to not use the temperature controller. 7. Place a drop of immersion oil on the coverslip and put the slide onto the microscope. Using a low magnification objective, quickly select a suitable onecell stage embryo and switch to a 63  or 100 oil objective for recording with the best possible image resolution using fluorescent microscopy.

13.3.2 How to limit phototoxicity As a good practice, effects of illumination on embryogenesis should be assayed. Under appropriate live imaging conditions, the monitored embryos develop normally (no defects in asymmetric cell divisions, timing of cell division nor chromatin segregation) and hatch under the coverslip after 12–14 h. Minimizing the light dose applied to the specimen is critical to reduce photobleaching and associated phototoxicity. Laser power should be kept at a minimum value and the number of slides in Z stacks as well as frequency of the stacks should be limited to what is strictly required. For a given dose of light it causes less damage when delivered slowly (increased exposure time with reduced laser power or light intensity) (Tinevez et al., 2012). Typically, a 488-nm laser power 25 amino acid residues in length that contain regions enriched in basic amino acid residues. Although recognition of a nuclear transport signal is generally linked to a specific Kap, for some cargos multiple Kaps may bind. Thus, there is always the potential for redundancy with respect to the number of Kaps capable of transporting a specific cargo (reviewed in Chook & Su¨el, 2011; Xu et al., 2010).

Introduction

Table 14.1 Nuclear transport signals Yeast karyopherin

Nuclear transport signal

Kap60p/Kap95p Kap60p/Kap95p Kap121p Kap121p

Monopartite classic NLS Bipartite classic NLS Basic NLS Basic NLS

Kap123p

Basic NLS

Kap104p Kap114p

Basic PY-NLS RS domain

Xpo1p

Leucine-rich NES

Amino acid sequence K-(K/R)-X-(K/R)1 (K/R)-(K/R)-X10–12-(K/R)13/5 K-(V/I)-X-K-X1–2-(K/H/R)2 Basic enriched peptide, >25 amino acids1 Basic enriched peptide, >25 amino acids1 (Basic enriched)4–20-(R/K/H)-X2–5-PY1 >40% arginine–serine dipeptide content1 f-X2–3-f-X2–3-f-X-f where f ¼ L, V, I, F, or M3

Amino acid sequences obtained from 1Chook & Su¨el, 2011; 2Kobayashi & Matsuura, 2013; 3 Xu, Framer, & Chook, 2010. X ¼ any amino acid 3/5 refers to 3 out of 5 consecutive amino acids are either a K or an R.

Although Kap/cargo complexes appear to have no intrinsic directionality with respect to how they move through pores, their role in directional transport, as importers or exporters, depends upon the small GTPase Ran (yeast Gsp1p). During import, importin/cargo complexes move through pores by transiently associating/dissociating with multiple Nups, rich in phenylalanine–glycine repeats, that line the central NPC channel. Once through the pore, the importin/cargo complex interacts with nuclear RanGTP resulting in cargo release and the export of the remaining importin/RanGTP complex back to the cytoplasm. Hydrolysis of RanGTP to RanGDP, catalyzed by a cytoplasmic Ran GTPase-activating protein (yeast Rna1), releases the importin and RanGDP is then imported back into nucleus. Once in the nucleus, RanGDP is converted to RanGTP by a Ran guanine nucleotide exchange factor (yeast Prp20p). During export, the exportin/cargo complex forms in the nucleoplasm and is stabilized through its association with RanGTP. This trimeric complex traverses the pore to the cytoplasm where the Ran GTPase-activating protein stimulates Ran hydrolysis of GTP and conversion of RanGTP to RanGDP. This destabilizes the trimeric complex, releasing the cargo and allowing both the exportin and RanGDP to independently reenter the nucleoplasm (reviewed in Cook, Bono, Jinek, & Conti, 2007). Often, cargos will possess both NLSs and NESs, thus their steady-state localization is dependent upon a competition between these two transport events. Which process dominates is often dependent upon extracellular cues and a cell’s physiological state including, for example, nutrient availability, stress conditions, and cellcycle stage. Often these triggers induce posttranslational modifications of the cargo

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that alter the balance between import and export. For example, in yeast cells, grown under conditions of phosphate starvation, the importin Kap121p actively imports the transcription factor Pho4p into the nucleus where it upregulates the expression of phosphate-responsive genes. Upon phosphate addition, however, Pho4p becomes phosphorylated, reducing its affinity for Kap121p, but augmenting its interaction with the exportin Msn5p. Together these events favor export and cytoplasmic accumulation of phosphorylated Pho4p (Kaffman, Rank, O’Neill, Huang, & O’Shea, 1998; Kaffman, Rank, & O’Shea, 1998). In this chapter, we will describe methods used in the analysis of regulated nuclear transport in the yeast Saccharomyces cerevisiae. These experiments center on the use of fluorescence microscopy to probe the in vivo localization of cargos or nuclear transport reporters fused to a fluorescent protein tag. We also detail specific genetic backgrounds and growth conditions used to study how pathways that control progression through the cell cycle regulate nuclear transport.

14.1 OBSERVING STEADY-STATE LOCALIZATION OF KAP CARGO PROTEINS 14.1.1 GFP-tagged nuclear transport cargos and reporters Assaying nuclear transport in vivo has been facilitated by the use of fusion proteins consisting of a nuclear transport signal-containing cargo protein and a fluorescent protein tag. In this chapter, we will focus our discussion on GFP, but other fluorescent proteins such as mCherry offer alternatives. These fusion proteins generally consist of the cargo protein fused at either their carboxy- or amino-terminus to GFP. Strains producing carboxy-terminal GFP fusions of most yeast proteins may be found as part of a library (Huh et al., 2003) that is commercially available (Invitrogen). Alternatively, a targeted cargo may be tagged in a specific genetic background using transformation and homologous recombination. For this purpose, DNA cassettes, constructed to integrate the coding sequence of a fluorescent protein at the carboxy- or amino-terminal ends of a specific ORF, can be synthesized by PCR using plasmids that have been previously described (e.g., Gauss, Trautwein, Sommer, & Spang, 2005; Longtine et al., 1998) and are available through various agencies (e.g., EUROSCARF). Alternatively, transport signals in specific cargoes can be used in isolation to examine transport events that regulate their nuclear import. This approach has the advantage of separating the analysis of a given transport pathway from other transport signals and interactions that may retain or anchor the cargo in the cytoplasm or the nucleus. These experiments employ nuclear transport reporter proteins consisting of either, an NLS–GFP fusion used for nuclear import studies, or an NLS–NES–GFP fusion used for nuclear export studies. The export reporter includes an NLS, along with the NES, to ensure that the fusion enters the nucleus prior to its export (Stade, Ford, Guthrie, & Weis, 1997). Most nuclear transport signals are fused to two or three tandem GFP moieties in order to raise the reporter size to >50 kDa, which is above

14.1 Observing Steady-State Localization of Kap Cargo Proteins

the NPC diffusion limit of
40 kDa. This reduces the contribution of passive diffusion, thus ensuring that reporter localization is predominantly dependent upon facilitated nuclear transport. In addition, these fusions typically employ GFP variants containing the S65T amino acid substitution, (GFPS65T, eGFP, GFPþ) as, relative to GFP, they exhibit stronger fluorescent signals that are not reduced at elevated temperatures, such as 37  C, and are less susceptible to photobleaching (see Ha, Schwarz, Turco, & Beverly, 1996; Sample, Newman, & Zhang, 2009). A caveat of using transport reporters is that the nuclear transport signal of the target is not always straightforward as only some signals conform to a consensus sequence or exhibit enrichment for a specific amino acid(s) (Table 14.1). As a result, signals are often identified through the generation of cargo truncations, each fused to GFP, that narrow down the region and ultimately the amino acid sequence required to produce an expected localization. A second caveat is that while an extracted sequence can function as a transport signal, this may not be the case within the context of the full-length protein from which it was extracted. Thus, complementary analysis may be required, such as the introduction of amino acid substitutions within a cargo’s putative nuclear transport signal and observing its effect on cargo localization. For an example, see Scott, Cairo, Van de Vosse, and Wozniak (2009) where the NES of Mad1p was defined. Beyond their use in defining a cargo’s nuclear transport signal, reporters may also be used to probe Kap function within specific biological pathways. For example, we employed the NLSPho4-GFP3 reporter (Kaffman, Rank, & O’Shea, 1998) to show that general Kap121p-mediated import is inhibited under conditions of spindle assembly checkpoint activation in yeast (Cairo, Ptak, & Wozniak, 2013).

14.1.2 Observing nuclear transport cargos and reporters by fluorescence microscopy Using fluorescence microscopy, the in vivo localization of cargo–GFP, NLS–GFP, and NLS–NES–GFP fusions can be observed. The steady-state localization of these fusions may vary depending upon the cellular levels of the fusion protein, the composition of the signal, and the numbers of nuclear transport signals in the cargo molecule. Those containing solely an import or export signal are predicted to exhibit a predominantly nuclear or cytoplasmic localization. Those cargos containing both signals can present with various ratios of nuclear to cytoplasm signal dictated by the relative strength of the NLS versus the NES (see Fig. 14.1A, for examples of these localizations). For example, a predominantly nuclear signal may reflect either the en masse import of a cargo or the dynamic import and export of the cargo where the import process is favored over export. In addition, changes in the physiological state of a cell may impact the localization of the cargo either by favoring import or export, or through the inhibition of nuclear transport (Fig. 14.1A and C). It is important to keep in mind that steady-state localization of a fusion protein is also influenced by the presence of cargo-binding partners in the cytoplasm or nucleus that could anchor the fusion protein in a compartment. Thus, the localization of a cargo is not necessarily a reflection of the relative strength of import versus export signal.

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CHAPTER 14 Nuclear Transport in Saccharomyces cerevisiae

FIGURE 14.1 (A) Various cargo protein localization phenotypes, in this case for the reporter NLSPho4-GFP3, are shown. The NLSPho4-GFP3 gene has been integrated into the strain shown. These include predominantly nucleoplasmic (cells grown in YPD) or, under conditions of nuclear import inhibition, predominantly cytoplasmic (cells grown in YPD þ 20 mg/ml nocodazole), or an intermediate localization between both nucleoplasmic and cytoplasmic (cells grown in YPD þ 20 mg/ml nocodazole). The position of a vacuole is indicated by the asterisk. (B) Shown are cells expressing NLSPho4-GFP3 from either a low-copy CEN/ARS plasmid or from a genomic locus (integrated). Note that the cell-to-cell variation of the fluorescent signal, observed when the reporter is expressed from a plasmid, is lost when the reporter gene is expressed from a genomic locus. (C) In the left-hand panels, cells producing either the NLSPho4-GFP3 or NLSSV40-GFP2 reporter are shown. Genes encoding each reporter have been integrated within their respective strains. These cells were treated with nocodazole and imaged by fluorescence microscopy at the times indicated. These images were used to quantify the average nuclear/cytoplasmic fluorescence intensity ratio (nuclear/cytoplasmic ratio) from 50 cells and these values graphed for each time point. Error bars express standard error. Panel C has been reprinted from Cairo et al. (2013) with permission from Elsevier.

14.1.3 Using plasmid-based or genomically integrated transport reporters Coding sequences for cargo–GFP and transport reporters (e.g., NLS–GFP) are often cloned into yeast CEN/ARS-based expression plasmids (e.g., pRS31X and pRS41X family of plasmids; Sikorski & Hieter, 1989). For example, Kaffman, Rank, and O’Shea (1998) cloned the PHO4 promoter and the NLSPho4-GFP3 coding sequence into the pRS316 plasmid. We have used this construct for the generation of other

14.1 Observing Steady-State Localization of Kap Cargo Proteins

plasmid-based reporters (Scott et al., 2009; Scott, Lusk, Dilworth, Aitchison, & Wozniak, 2005) as: the NLSPho4 sequence may be readily replaced with a coding sequence of interest; the GFP3 provides a strong signal and generates fusions above the NPC diffusion limit; the PHO4 promoter leads to sufficient reporter production and ultimately fluorescent signal. However, cells transformed with these “low-copy” plasmids contain one to several copies of the plasmid per cell. Thus, there is often significant cell-to-cell variability in the expression of the fusion gene and levels of the GFP signal strength (Fig. 14.1B). This variability is further exacerbated when high-copy number plasmids are used. This phenomenon can make it difficult to assess fusion protein localization across the cell population. Where necessary, this shortcoming can be overcome by integrating the ORF for GFP at the 30 -end of the cargo gene ORF allowing examination of the cargo–GFP encoded by the gene and regulated by its promoter. Strains producing carboxy-terminal GFP fusions of most yeast proteins may be found as part of a library (Huh et al., 2003) that is commercially available (Invitrogen). Variability in the expression levels of plasmid-based NLS- or NLS–NES–GFP transport reporter genes can also be reduced by their integration. To integrate these reporter genes, we use the plasmids as templates in a PCR to generate a DNA cassette that includes the reporter-coding sequence, relevant regulatory elements (promoter, terminator), and a marker gene. The oligonucleotides used include sequences that base pair with plasmid sequence adjacent to the region that encodes all of these elements. In addition, the oligonucleotides include 60 nucleotides of flanking sequence used for integration of the DNA cassette at a genomic locus. The loci we have used include auxotrophic markers (e.g., ura3 or leu2) within the yeast strain of choice (Cairo et al., 2013). We observe that strains producing these reporters from a genomic locus produce a reporter-GFP signal that is equivalent from cell to cell (Fig. 14.1B).

14.1.4 Preparing yeast cells for fluorescence microscopy The following describes a generalized approach to assess the localization of GFPlabeled fusion proteins from which the underlying nuclear transport process is inferred. Variations of this protocol form the basis of those described in subsequent sections. Equipment and reagents required are listed in Section 14.3. Growth media used will depend upon whether the GFP fusion gene under study is expressed from a plasmid (Synthetic complete (SC) media designed for plasmid selection) or integrated and expressed from a genomic locus (Yeast extract Peptone Dextrose (YPD) media). Note: Adenine supplementation. When working with any ADE3þ ade2-1 strain of S. cerevisiae, (e.g., W303- or YPH-based strains), adenine is added from a 50  stock solution (6 mg/ml adenine in 0.1 N NaOH) to a final concentration of 120 mg/ml. This helps prevent background arising from the accumulation of a fluorescing metabolic intermediate produced as a consequence of the ade2-1 mutation.

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CHAPTER 14 Nuclear Transport in Saccharomyces cerevisiae

PROTOCOL 1. Inoculate 5 ml of appropriate liquid culture media (YPD or SC) with desired yeast strain and incubate the culture overnight at room temperature (RT) with agitation, for example, on a roller-drum. For larger cultures ( 10 ml), use an appropriately sized erlenmeyer flask and grow the cultures on an orbital shaker. This is required for methods described in subsequent sections where cultures are split or used in a time course. Incubation at RT helps prevent overgrowth, producing an overnight culture at mid to late log phase or OD600 between
0.5 and 2.0. An OD600 of 1.0 is generally equivalent to
2  107 cells/ml. 2. The following day, the culture is diluted to an OD600 of
0.1–0.3, and incubated at desired temperature for two or more generations to an OD600 of
0.5–1.5. We avoid using cultures with too low (2.0) an OD600. 3. Transfer 1 ml of culture to a microcentrifuge tube and collect the cells by centrifugation at
13,000  g for 30 s in a microcentrifuge. Cells grown in SC media tend not to pellet completely. To circumvent this, 100 ml of YPD can be added to the 1 ml of culture prior to centrifugation. This improves the efficiency of cells pelleting. 4. Aspirate off the media and wash the cells with 1 ml of fully supplemented SC media, that is, all amino acids present. As components of YPD will fluoresce, and hence obscure sample observation, this step is particularly useful when YPD is used. 5. Collect the cells by microcentrifugation. Aspirate off the media and resuspend the cells in
20–50 ml of fully supplemented SC media. This should provide a cell density giving between 20 and 100 cells per field when viewed by microscopy at 100  magnification. 6. Spot 1.5 ml of the cell suspension onto a slide. From just above the spot, drop a coverslip onto the cells to promote cell spreading. This should provide an even, single layer of cells that do not move and are observable in the same focal plane. Slides made in this manner should be useable for
15 min before they begin to dry out. 7. We typically visualize cells by epifluorescence microscopy at 100  magnification. Acquisition times will be dependent upon the specific GFP fusion, but should be limited so as not to produce a saturated signal. This is critical for signal quantification as described in the next section.

14.1.5 Quantifying nucleoplasmic/cytoplasmic fluorescence intensity ratios Alterations in transport mediated by specific Kaps, or of targeted cargos, in response to changes in cell physiology often represent an area of keen interest. Changes in the steady-state localization of the cargo–GFP can often be subtle and assessing potential alterations often requires quantifying the relative nuclear/cytoplasmic ratio of cargo– GFP fusions. To do this, we determine the relative amount of GFP fusion

14.2 Perturbing Nuclear Transport

fluorescence observed in the nucleus and cytoplasm of cells. Specifically, Image J software is used to determine the mean integrated fluorescence intensity per unit area in each compartment, which is then expressed as a nuclear/cytoplasmic fluorescence intensity ratio. This analysis is performed as follows. PROTOCOL 1. Acquire images by fluorescence microscopy as described in Section 14.1.4, using various acquisition times. Various exposures yielding subsaturated signal intensity will be used in the following steps to quantify nuclear/cytoplasmic fluorescence intensity ratios. 2. Open a micrograph in Image J and, using the “rectangular” tool, make a 5 pixel  5 pixel box. 3. Place the box within the nucleus of one cell and determine the nuclear mean integrated fluorescence intensity/unit area for the boxed area by clicking the “Measure” tool found under the “Analyze” heading. For GFP fusions possessing an NLS, the nucleus is usually observed as a spot enriched in GFP fluorescence. When the nucleus is not apparent, NPC or NE markers, such as Sec63-mCherry, may be used as a marker for the nuclear periphery (e.g., see Van de Vosse et al., 2013). 4. Repeat step 2 for a cytoplasmic region, as well as a region outside the cell that will define the background signal. When choosing a cytoplasmic region avoid vacuoles as they generally lack fluorescence and will give a false low value for cytoplasmic fluorescence intensity (Fig. 14.1A, asterisk). 5. Subtract the background from both the nuclear and cytoplasmic intensities. 6. Calculate the intensity ratio by dividing the nuclear intensity by the cytoplasmic intensity. 7. Repeat the process for 50 or more cells. Use these values to calculate an average nuclear/cytoplasmic fluorescence intensity ratio. Calculated ratios for a given exposure time should be similar to those determined for other subsaturated exposures. See Fig. 14.1C, for an example.

14.2 PERTURBING NUCLEAR TRANSPORT Cargo localization, as observed using the techniques described above, reflects the steady-state distribution of the cargo as determined by relative rates of import and export under a given growth condition. Understanding the regulation of these transport events requires identifying the factors and physiological conditions that mediate and modulate the cargos localization, that is, what Kaps control the cargo’s localization and what cellular events alter its transport. To probe these mechanistic details, changes in cargo localization within specific mutant backgrounds and/or under specific growth conditions are used. Below, we describe some conditions we have used to assess mechanisms of cargo localization, focusing on the use of conditional yeast mutants and cell-cycle analysis.

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It should be noted that the methods described below are not useful in studying nuclear import processes where either, there is little or no exchange of the cargo between nucleoplasm and cytoplasm after import, or when treating the cells causes a weak transport defect with no observable change in the nuclear signal of an NLS– GFP reporter. The first limitation may be overcome through the use of an inducible system where production of the cargo–GFP is activated only after the cells have been subjected to a specific perturbation, and its import, or lack thereof, observed (see Leslie, Timney, Rout, & Aitchison, 2006). To observe weak import defects, yeast cells, producing a reporter composed of an NLS fused to a single GFP, are treated with metabolic poisons that inhibit facilitated nuclear transport. This allows the NLS–GFP to passively diffuse and equilibrate between cytoplasm and nucleus. Facilitated transport is then restored upon removal of the poisons and the import rate of the reporter determined. By comparing the import rate in treated and untreated cells, the affect of a specific perturbation on nuclear import can be assessed (see Leslie, Timney, Rout, & Aitchison, 2006; Roberts & Goldfarb, 1998).

14.2.1 Conditional yeast mutant strains The use of conditional mutants has proved to be a powerful tool for investigating nuclear transport processes. In yeast, strains carrying a null mutation for nonessential genes are readily made (Tong et al., 2001) and are also available commercially (Invitrogen). Transport in mutants lacking nonessential genes can be directly examined in parallel with a WT counterpart. However, many genes involved in transport and/or its regulation are essential and can only be studied using a conditional mutation. Most of these are temperature-sensitive mutations and include those that are either directly engaged in transport (e.g., kapts, gsp1ts, prp20ts, etc.) or in other biological pathways that directly or indirectly regulate transport, for example, cell-cycle and/or signal transduction pathways. Alternatively, genes of interest may be placed under control of a conditional promoter element, such that they are only expressed under specific growth conditions. An example is use of the MET3 promoter (PMET3) that activates gene expression only when cells are grown in media lacking methionine (Mao, Hu, Liang, & Lu, 2002). A third case is use of a mutant that sensitizes the S. cerevisiae exportin Xpo1p to the antifungal drug Leptomycin B (LMB). While most eukaryotic homologs of Xpo1p are LMB sensitive, the S. cerevisiae homolog lacks a conserved cysteine residue that when alkylated by LMB results in export inhibition. Inserting this cysteine residue (xpo1T539C) renders S. cerevisiae Xpo1p sensitive to LMB (Neville & Rosbash, 1999). Use of these conditional strains is described below.

14.2.1.1 Preparing temperature-sensitive (ts) yeast strains for fluorescence microscopy Note: When using ts yeast strains, fusions should include GFP derivatives carrying the S65T amino acid substitution as GFP itself produces a significantly weaker fluorescent signal at higher temperatures such as 37  C (Sample et al., 2009).

14.2 Perturbing Nuclear Transport

PROTOCOL 1. Follow the protocol described in Section 14.1.4 to establish actively growing cultures of the targeted ts mutant and its WT counterpart. The WT culture is used as a control to ensure that any defects observed in the ts mutant are attributable to a loss of function rather than temperature changes. Cultures are initially grown at a permissive temperature, preferably one that minimizes any defects associated with the mutation. As many mutants isolated based on restricted function at 37  C also show partial defects at 30  C, we tend to grown all cultures at RT prior to shifting to the nonpermissive temperature. 2. Once cultures grown at the permissive temperature have reached the desired cell density, they are split each into two separate cultures. If necessary, dilute the culture to an OD600 of
0.1–0.3 with media at the same temperature to prevent saturation over the course of the temperature shift. Keep one culture of each strain at the permissive temperature and transfer the other culture directly to an air shaker-incubator set at the nonpermissive temperature (37  C for most strains). An air shaker allows for a gradual increase in the temperature of the culture and prevents a heat-shock response that might occur if the cells were transferred directly to a preheated water bath shaker at the nonpermissive temperature. 3. The time of incubation at the nonpermissive temperature, prior to the analysis of transport, is dependent on the mutant. Generally, we examine various time points after temperature shift (between 0 and 3 h) to assess transport changes and, if applicable, their relationship to known phenotypes of the mutant. 4. At the various time points, cells in each culture are processed for microscopy as outlined in Section 14.1.4. Processing and image acquisition are usually done at RT, thus, the time spent to carry out these steps should be quick (
10–15 min maximum) to minimize potential recovery of cells grown at the nonpermissive temperature. If recovery of mutant protein function is rapid, solutions used to prepare the cells can be maintained at the nonpermissive temperature and a heated microscope stage can be used during image acquisition. Note: When many cultures are being analyzed during the same experiment, it is useful to stagger their initial transfer to the nonpermissive temperature, for example, every 15 min and then process in a similar staggered fashion to ensure that each culture is treated in an identical manner. Alternative approaches: Cells sampled at various time points can also be fixed by the direct addition of formaldehyde (Sigma) to a culture to a final concentration of 4%. Cultures are then incubated an additional 5–30 min prior to preparation for microscopy as described in Section 14.1.4. As fixation may inhibit the fluorescence of some GFP fusions, the affect of formaldehyde treatment should be tested prior to its experimental use. For in vivo studies, microfluidic chambers (available from CellASIC Corporation, San Leandro, CA, USA), designed to allow for rapid changes in media and environment (e.g., temperature) using a flow control system, can also be used.

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14.2.1.2 Using the MET3 promoter to shutoff gene expression This method requires replacing the promoter driving a gene of interest with PMET3. We have employed a PCR-based integrative approach described elsewhere, using, as a template, a plasmid that has been modified to include PMET3 (Makio et al., 2009). Methionine auxotrophs may not be used for this technique, as strains are grown in media lacking methionine. The DNA cassette used to introduce PMET3 at a given locus also introduces a selectable marker, which can be used to identify putative positive integrants. Of note, when essential genes are placed under control of the PMET3 promoter, the strain must be grown in media lacking methionine to allow expression of the essential gene. In addition, we generally use a PMET3 cassette containing the coding sequence for an HA3 tag engineered to insert at the amino-terminal end of the target gene open reading frame (ORF). PROTOCOL 1. Inoculate 5 ml of SC media lacking methionine with a colony of the desired yeast strain and incubate the culture overnight with agitation to mid-log phase (OD600
0.5). For most strains, incubate at 30  C. 2. Pellet the cells by centrifugation at
1700  g for 2–3 min using a tabletop swinging bucket centrifuge and then discard the media. 3. Resuspend the pellet in 5 ml of YPD and supplement the culture with 20 mg/ml methionine (5 ml of 20 mg/ml methionine in ddH20). Alternative: If the cultures used must remain in SC media, for example, to select for a plasmid, methionine can be added directly to the culture in step 1 to a final concentration of 200 mg/ml. 4. Incubation time required to deplete the protein encoded by the gene of interest must be determined for each gene product as it is dependent upon the turnover rate of the protein. Preliminary experiments should include a time course that follows depletion of the HA3-tagged protein using Western blot analysis. 5. Process cells in each culture for microscopy as outlined in Section 14.1.4.

14.2.1.3 Inhibition of xpo1T539C using LMB To analyze the function of yeast Xpo1 in the localization of a targeted cargo, the endogenous XPO1 gene can be replaced with the xpo1T539C allele in a genetic background of interest (Neville & Rosbash, 1999). For example, we have defined a putative NES present in the yeast spindle assembly checkpoint protein Mad1p, by engineering the Mad1p NES sequence into a plasmid encoding a NLSSV40-NESMad1pGFP3 reporter and transforming this plasmid into an xpo1T539C-containing strain (Scott et al., 2009). As first demonstrated using an NLSSV40-NESPKI-GFP3 reporter (Stade et al., 1997), an active NES will prevent nuclear accumulation of the reporter. Thus, this reporter can be used to assess the functionality of a putative NES by examining its nuclear exclusion. Moreover, the role of Xpo1 for mediating the export of the NES-containing reporter or cargo can be assessed by treatment of the xpo1T539C strain with LMB and monitoring its nuclear accumulation.

14.2 Perturbing Nuclear Transport

PROTOCOL 1. Cultures of each strain are grown following the protocol outlined in Section 14.1.4 using the appropriate SC media to select for the plasmids if necessary. 2. Once the cells reach an OD600
0.5, split each culture into two. Treat one set with vehicle (20 ml methanol/ml of culture) and the other with LMB to a final concentration of 100 ng/ml (20 ml of 5 mg/ml LMB in methanol per ml of culture). 3. Incubate the cultures at 30  C (for most strains) and examine reporter localization using fluorescence microscopy at 15 min intervals. Generally, for a reporter such as NLSSV40-NESPKI-GFP3, nuclear accumulation is visible in 0.5–1 h.

14.2.2 Assessing cell-cycle dependence of nuclear transport Many nuclear transport processes are regulated in a cell cycle-dependent manner. These may include regulatory events that occur as the cell passes through a particular cell-cycle stage, or may result from cell-cycle arrest in response to activation of a specific checkpoint. Examples include a transport switch for the Sumo E3 ligase Siz1p from principally import (i.e., nuclear accumulation) during G1 and S-phase to export during mitosis (i.e., cytoplasmic localization and septin ring association) ( Johnson & Gupta, 2001; see Fig. 14.2A). In another case, we find that Kap121pmediated nuclear import is inhibited in response to kinetochore/microtubule detachment leading to the loss of nuclear accumulation (Cairo et al., 2013).

14.2.2.1 Asynchronous cultures Possible cell cycle-dependent changes in the localization of a cargo may be assessed in asynchronous cultures. A rough gauge of which cell-cycle stage a cell is currently in employs a visual comparison of mother cell size with the presence and size of the emerging daughter bud, where, unbudded and small budded cells are in G1 phase, small-medium budded cells are in S–G2, and medium–large budded cells are in various stages of mitosis. A finer indicator of cell-cycle stage combines bud size with an additional cell-cycle indicator such as nuclear position and morphology (Hartwell, Culotti, Pringle, & Reid, 1974). The nucleus may be visualized using the cargo if the GFP fusion is imported. Alternatively NPC or NE markers, such as Sec63-mCherry, may be used as a marker for the nuclear periphery. Visualization of Siz1-GFP provides an example of using these cell-cycle characteristics to observe its reduced nuclear localization and appearance in the cytoplasm and at the budneck as cells enter mitosis (Fig. 14.2A).

14.2.2.2 Stage-specific cell-cycle arrest In some cases, it is desirable to arrest cells at a particular cell-cycle stage prior to visualization of the cargo by fluorescence microscopy. There are numerous methods that arrest yeast cells in particular stages of the cell cycle (see Amon, 2002). Here, the

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FIGURE 14.2 (A) Changes in Siz1p localization at various cell-cycle stages are shown. Cells are shown as a progression through the cell cycle and include unbudded (G1), small budded (G1–S), medium budded (S–G2), and large budded (mitosis) cells. The nucleus is apparent as the region showing accumulation of Siz1-GFP signal. Of note, the location and morphology of nuclei in these various cells are indicative of their position in the cell cycle. Cells in G1, S, and G2 phases possess a single nucleus in the mother cell that is positioned away from the budneck. In metaphase, a single nucleus is found at the budneck. During early anaphase, chromosome segregation begins and the nucleus enters the daughter cell. As anaphase progresses, two distinct nuclei form and eventually migrate to the distal ends of the mother and daughter cells by late anaphase/telophase. (B) Shown are FACS profiles for yeast cells from a single culture incubated under various conditions. Initially, the culture was grown asynchronously (AS) to mid-log phase in pH 5.0 YPD. a-Factor was then added and the culture incubated for 3 h prior to sampling for FACS. The cells were then pelleted, washed, and resuspended in YPD containing 20 mg/ml nocodazole and 1% DMSO. Samples for FACS analysis were then taken from this culture every 20 min. FACS profiles shown measure the number of cells (cell count) with a specific DNA content (fluorescence). DNA content is characteristic of specific cell-cycle stages where 1n represents cells with unreplicated DNA in G1, while cells with a DNA content of 2n have replicated DNA and are in G2 or M. a-Factor arrest causes accumulation of cells in START (G1) with a 1n DNA content, while nocodazole treatment arrests cells in M-phase with a 2n DNA content. Cells in S-phase have a DNA content intermediate to these, for example, the small peak at the 40 min time point after release into nocodazole represents cells undergoing replication.

14.2 Perturbing Nuclear Transport

methods we favor are described, including protocols to arrest cells at START, S-phase, or metaphase. Observations made under these various arrest conditions can reveal changes in the localization of cargo molecules arising from modifications of the cargo itself or the nuclear transport machinery. These data can provide important insight into the functions of the nuclear transport machinery in regulating cellular pathways.

14.2.2.2.1 START arrest using a-factor

The mating pheromone a-factor induces a signal transduction pathway within haploid MATa cells that leads to their arrest, prior to cell-cycle commitment, at START. This arrest is characterized by the accumulation of unbudded cells that form a mating projection or “shmoo” (reviewed in Bardwell, 2005). 1. a-factor is solubilized in ddH20 to 1 mg/ml and stored at 20  C as a 200  stock. Strains to be treated with a-factor must have a MATa mating type. 2. Follow the steps in Section 14.1.4 but use YPD or SC media whose pH has been adjusted to 5.0. The efficacy of a-factor in mediating arrest is much higher at this pH as the Bar1p protease, that degrades a-factor, is inhibited under this condition (Futcher, 1999). 3. Once cells have reached an OD600 no > 0.5, add a-factor to a final concentration of 5 mg/ml and incubate
2.5–3 h. The extent of arrest can be visually determined by microscopy using a brightfield microscope. Efficient arrest generally leads to >90% of cells appearing unbudded and exhibiting a schmooing phenotype (Amon, 2002). 4. Alternative: To reduce the amount of a-factor required for cell arrest, the BAR1 gene can be deleted in the strain being used. Loss of Bar1p hypersensitizes cells to a-factor such that only 10–50 ng/ml are required to induce arrest (MacKay et al., 1988). This also eliminates the need for using pH 5.0 media. 5. Process cells in each culture for microscopy as outlined in Section 14.1.4. For these and subsequent cell-cycle arrest protocols, it is important to confirm the efficiency of arrest using FACS analysis (e.g., see Fig. 14.2B). Cells to be analyzed by FACS are pelleted from 1 ml of mid-log phase culture (
1  107 cells) then fixed by resuspension in 1 ml of 70% ethanol. Prior to further preparation, cells are incubated at RT for 1 h or can be left at 4  C overnight to several days prior to further treatment. Cells are then treated sequentially with RNase, protease, and a fluorescing DNA dye (propidium iodide or SYTOX green) prior to determination of the cells DNA content by flow cytometry (for details, see Hasse & Reed, 2002). 6. Release from arrest: After step 3, cells can be released from arrest, to reenter the cell cycle, by washing the cells twice with media lacking a-factor. After the second wash, the cells are resuspended in the same media and incubated at 30  C. By observing a sample of the culture every 10–20 min by fluorescence microscopy, progression of the cell population, and localization of the transport cargo, through the cell cycle can be followed. We generally do not sample beyond
3 h postrelease as the cultures become asynchronous.

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14.2.2.2.2 S-phase arrest using hydroxyurea 1. Follow the steps in Section 14.1.4; however, on day 2, grow the cultures to an OD600 no greater than 0.5. 2. Add hyroxyurea powder directly to the culture for a final concentration of 250 mM. 3. Incubate for 3 h (approximately two generations). Cells should appear as large budded. These cells generally have a rounder appearance then large budded cells observed in asynchronous populations and also have a single nucleus. 4. Process cells in each culture for microscopy as outlined in Section 14.1.4.

14.2.2.2.3 Metaphase arrest A metaphase arrest can be induced by activation of the spindle assembly checkpoint. Activation of this checkpoint occurs in response to defects in kinetochore– microtubule attachments. Drugs that cause microtubule depolymerization, such as nocodazole, can be used to disrupt these attachments (protocol detailed below). Alternatively, ts mutations in kinetochore components, for example, ndc80ts or ask1ts alleles, sufficiently destabilize the kinetochore–microtubule interface to activate the spindle assembly checkpoint (Pinsky, Kung, Shokat, & Biggins, 2006). The protocol outlined in Section 14.2.1.1 to evaluate cargos in a ts mutant can be employed for kinetochore mutants producing a GFP–cargo. Alternatively, metaphase arrest can be achieved without disrupting kinetochore– microtubule interactions by altering levels of key mitotic regulates such as Cdc20. During mitosis, increasing levels of Cdc20 bind to and activate the anaphase promoting complex. By downregulating the expression of CDC20 using the regulatable MET3 promoter, cells can be arrested in metaphase (Cairo et al., 2013). Using a PMET3-CDC20 strain producing a GFP–cargo, the protocol outlined in Section 14.2.1.2 can be followed to deplete cells of Cdc20p and examine transport in metaphase arrested cells. Moreover, we have used PMET3-CDC20-mediated metaphase arrest to distinguish between transport regulatory events controlled by metaphase arrest (detected in Cdc20 depleted cells) from those activated by disruption of kinetochore–microtubules interactions (detected in nocodazole arrested cells or Cdc20 depleted cells treated with nocodazole). PROTOCOL: NOCODAZOLE TREATMENT 1. Follow the steps in Section 14.1.4; however, on day 2, grow the cultures to an OD600 no greater then 0.3, then split each culture into two. 2. To one of the split cultures, add DMSO to a final concentration of 1% v/v (vehicle only). The other half of the culture is treated with nocodazole as follows. To 1 ml of culture media, add 10 ml of a 200 mg/ml solution of nocodazole in DMSO. The latter solution is made fresh by diluting a nocodazole stock (2 mg/ml in DMSO) 10-fold with DMSO. Nocodazole can also be used at 12.5 and 15 mg/ml concentrations; however, 20 mg/ml works best in our hands.

14.3 Materials and Reagents

In addition, we have found that the efficacy of nocodazole is not equivalent from all vendors. We have had the best success with that obtained from EMD Millipore. 3. Incubate the cells for 2–3 h. We generally find that the majority of cells are arrested and appear large budded by 2–2.5 h. By 3 h and longer, a significant proportion of the population begins to rebud, indicative of slippage out of the metaphase arrest. 4. Process cells in each culture for microscopy as outlined in Section 14.1.4. We generally image at 0.5 h time points starting 1.5 h after nocodazole addition.

14.3 MATERIALS AND REAGENTS 14.3.1 Plasmids pRS31X and pRS41X family of plasmids; (Sikorski & Hieter, 1989) pNLSPho4-GFP3 (pRS316 based; Kaffman, Rank, & O’Shea, 1998) pTM1046 (contains PMET3 that replaces PGAL1 of pFA6a-kanMX6-PGAL1-3HA) (Longtine et al., 1998; Makio et al., 2009)

14.3.2 Strains Yeast deletion library strains (available from Invitrogen) ask1ts strain (W303; Pinsky et al., 2006) bar1△ (BY4741; our lab) ndc80ts strain (W303; Pinsky et al., 2006) PMET3-CDC20 strain (W303; our lab) SEC63-mCherry-NAT (BY4741; our lab) xpo1T539C (W303; Neville & Rosbash, 1999)

14.3.3 Material/equipment Erlenmeyer flasks, orbital shaker, test tubes (16  150-mm tubes for cultures of 5 ml or less), roller-drum, incubator, 1.5 ml polypropylene microcentrifuge tubes, microcentrifuge, glass microscope slides, glass coverslips, fluorescence microscope, flow cytometer (we used a FACScan, Becton Dickinson).

14.3.4 Reagents, buffers, and media Yeast media – 20% Dextrose (per 500 ml): Solubilize 100 g dextrose in double deionized water (ddH2O) to a final volume of 500 ml, and then autoclave. – SC liquid media (per l): 1.7 g yeast nitrogen base, 5 g ammonium sulphate, and 0.8 g amino acid dropout powder (specific for plasmid auxotrophic marker used, ref if commercial, or briefly mention prepared by mixing equivalent amounts of all

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aa þ ade þ ura at concentration XX each?– company is Sunrise Science Products) are solubilized in ddH2O to a final volume of 900 ml and then autoclaved. Once autoclaved, 100 ml of sterilized 20% (w/v) dextrose is added to a final concentration of 2% (w/v). – YPD liquid media (per l): 10 g yeast extract and 20 g peptone are solubilized in ddH2O to a final volume of 900 ml and then autoclaved. Once autoclaved, 100 ml of sterilized 20% (w/v) of dextrose is added to a final concentration of 2% (w/v). a-factor (acetate salt; Sigma): Stock solution: 1 mg/ml in ddH2O. Store aliquots at 20  C Formaldehyde (36.5–38% solution; Sigma) LMB (Sigma): Stock solution: 5 mg/ml in 100% methanol. Store aliquots at 20  C. Hydroxyurea (Sigma): Store powder at 4  C. Nocodazole (EMD Millipore): Stock solution: 2 mg/ml in DMSO.

Acknowledgments Funding sources for this work were provided by the Howard Hughes Medical Institute, Alberta Innovates Health Solutions, and the Canadian Institutes of Health Research (MOP 106502).

References Aitchison, J. D., & Rout, M. P. (2012). The yeast nuclear pore complex and transport through it. Genetics, 190(3), 855–883. Amon, A. (2002). Synchronization procedures. Methods in Enzymology, 351, 457–467. Bardwell, L. (2005). A walk-through of the yeast mating pheromone response pathway. Peptides, 26(2), 339–350. Cairo, L. V., Ptak, C., & Wozniak, R. W. (2013). Mitosis-specific regulation of nuclear transport by the spindle assembly checkpoint protein Mad1p. Molecular Cell, 49(1), 109–120. Chook, Y. M., & Su¨el, K. E. (2011). Nuclear import by karyopherin-bs: Recognition and inhibition. Biochimica et Biophysica Acta, 1813(9), 1593–1606. Cook, A., Bono, F., Jinek, M., & Conti, E. (2007). Structural biology of nucleocytoplasmic transport. Annual Review of Biochemistry, 76, 647–671. Futcher, B. (1999). Cell cycle synchronization. Methods in Cell Science, 21(2–3), 79–86. Gauss, R., Trautwein, M., Sommer, T., & Spang, A. (2005). New modules for the repeated internal and N-terminal epitope tagging of genes in Saccharomyces cerevisiae. Yeast, 22(1), 1–12. Ha, D. S., Schwarz, J. K., Turco, S. J., & Beverly, S. M. (1996). Use of the green fluorescent protein as a marker in transfected Leishmania. Molecular and Biochemical Parasitology, 77(1), 57–64. Hartwell, L. H., Culotti, J., Pringle, J. R., & Reid, J. T. (1974). Genetic control of the cell division cycle in yeast. Science, 183(4120), 46–51.

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Hasse, S. B., & Reed, S. I. (2002). Improved flow cytometric analysis of the budding yeast cell cycle. Cell Cycle, 1(2), 132–136. Hoelz, A., Debler, E. W., & Blobel, G. (2011). The structure of the nuclear pore complex. Annual Review of Biochemistry, 80, 613–643. Huh, W.-K., Falvo, J. V., Gerke, L. C., Carroll, A. S., Howson, R. W., Weissman, J. S., et al. (2003). Global analysis of protein localization in budding yeast. Nature, 425(6959), 686–691. Johnson, E. S., & Gupta, A. A. (2001). An E3-like factor that promotes SUMO conjugation to the yeast septins. Cell, 106(6), 735–744. Kaffman, A., Rank, N. M., O’Neill, E. M., Huang, L. S., & O’Shea, E. K. (1998). The receptor Msn5 exports the phosphorylated transcription factor Pho4 out of the nucleus. Nature, 396(6710), 482–486. Kaffman, A., Rank, N. M., & O’Shea, E. K. (1998). Phosphorylation regulates association of the transcription factor Pho4 with its import receptor Pse1/Kap121. Genes and Development, 12(17), 2673–2683. Kobayashi, J., & Matsuura, Y. (2013). Structural basis for cell-cycle-dependent nuclear import mediated by the karyopherin Kap121p. Journal of Molecular Biology, 425(11), 1852–1868. Leslie, D. M., Timney, B., Rout, M. P., & Aitchison, J. D. (2006). Studying nuclear protein import in yeast. Methods, 39(4), 291–308. Longtine, M. S., McKenzie, A., 3rd., Demarini, D. J., Shah, N. G., Wach, A., Brachat, A., et al. (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast, 14(10), 953–961. MacKay, V. L., Welch, S. K., Insley, M. Y., Manney, T. R., Holly, J., Saari, G. C., et al. (1988). The Saccharomyces cerevisiae BAR1 gene encodes an exported protein with homology to pepsin. Proceedings of the National Academy of Sciences of the United States of America, 85(1), 55–59. Makio, T., Stanton, L. H., Lin, C.-C., Goldfarb, D. S., Weis, K., & Wozniak, R. W. (2009). The nucleoporins Nup170p and Nup157p are essential for nuclear pore complex assembly. Journal of Cell Biology, 185(3), 459–473. Mao, X., Hu, Y., Liang, C., & Lu, C. (2002). Met3 promoter: A tightly regulated promoter and its application in construction of conditional lethal strain. Current Microbiology, 45(1), 37–40. Neville, M., & Rosbash, M. (1999). The NES-Crm1 export pathway is not a major mRNA export route in Saccharomyces cerevisiae. The EMBO Journal, 18(13), 3746–3756. Pinsky, B. A., Kung, C., Shokat, K. M., & Biggins, S. (2006). The Ilp1-Aurora protein kinase activates the spindle checkpoint by creating unattached kinetochores. Nature Cell Biology, 8(1), 78–83. Roberts, P. M., & Goldfarb, D. S. (1998). In vivo nuclear transport kinetics in Saccharomyces cerevisiae. Methods in Cell Biology, 53, 545–557. Sample, V., Newman, R. H., & Zhang, J. (2009). The structure and function of fluorescent proteins. Chemical Society Reviews, 38(10), 2852–2864. Scott, R. J., Cairo, L. V., Van de Vosse, D. W., & Wozniak, R. W. (2009). The nuclear export factor Xpo1p targets Mad1p to kinetochores in yeast. Journal of Cell Biology, 184(1), 21–29. Scott, R. J., Lusk, C. P., Dilworth, D. J., Aitchison, J. D., & Wozniak, R. W. (2005). Interactions between Mad1p and the nuclear transport machinery in the yeast Saccharomyces cerevisiae. Molecular Biology of the Cell, 16(9), 4362–4374.

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CHAPTER

15

Analysis of Nucleocytoplasmic Transport in DigitoninPermeabilized Cells Under Different Cellular Conditions

Maiko Furuta*, Shingo Kose{, Ralph H. Kehlenbach{, and Naoko Imamoto{ *

Department of Molecular Genetics, National Institute of Genetics, The Graduate University for Advanced Studies Sokendai, Mishima, Shizuoka, Japan { Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan { Department of Molecular Biology, Faculty of Medicine, Georg-August-University of Go¨ttingen, Go¨ttingen, Germany

CHAPTER OUTLINE Introduction ............................................................................................................ 332 15.1 Equipment, Material, Reagents, and Buffers......................................................335 15.1.1 Equipments ............................................................................... 335 15.1.2 Material and Reagents ................................................................ 336 15.1.3 Strain and Cell Line.................................................................... 338 15.1.4 Buffers ...................................................................................... 338 15.2 Purification of Recombinant Transport Factors ..................................................339 15.3 Use of Digitonin-Permeabilized Cells to Study Nuclear Transport Under Normal and Heat-Shock Conditions ..................................................................342 15.3.1 Preparation of Cytosol ................................................................. 343 15.3.2 Preparation of Fractionated Cytosol (Importin-depleted/Containing Fractions) .................................................................................. 344 15.3.3 Preparation of Fluorescent His-Tagged GFP-Hsc70 Substrate......... 344 15.3.4 Preparation of Permeabilized Cell ................................................ 345 15.3.5 Transport Assay.......................................................................... 346 15.4 Interphase Nucleocytoplasmic Transport and Mitotic Chromosome Loading of Chromokinesin hKid ........................................................................................346 15.4.1 Preparation of GST-FLAG-hKid .................................................... 347 15.4.2 Preparation of Permeabilized Mitotic Cells.................................... 348 15.4.3 In Vitro Chromosome-Loading Reaction ........................................ 348

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00015-1

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Conclusions and Perspectives ................................................................................. 349 Acknowledgments ................................................................................................... 349 References ............................................................................................................. 349

Abstract The regulation of nucleocytoplasmic transport is crucial not only for basic cellular activities but also for physiological adaptation to specific situation during the cell cycle, development, or stress. Although a wide variety of transport pathways have been identified in eukaryotic cells, the functional significance of their multiplicity remains unclear. The best-characterized nuclear transport receptors (NTRs) are the members of the importin b family (karyopherin, transportin) whose association with specific cargoes is regulated by the GTPase Ran. In this chapter, we first provide an overview of the various expression vectors used to purify recombinant NTRs. We then describe two sets of recent examples of using well-established digitonin-permeabilized cellfree transport systems in mammalian cells to mimic different cellular conditions in living cells: normal/heat-shock conditions and interphase/mitosis. In the former case, physiological regulation impacts different transport pathways in opposite ways. In the latter case, the importin b–Ran system is used at different cell-cycle stages but with the same biochemical principle to specify the nuclear localization and chromatin loading of a specific protein, respectively. This in vitro transport assay, when adapted to specific cellular conditions or particular substrates, should help to uncover specific transport pathways or transport factors function under different cellular conditions.

INTRODUCTION The nucleus is the control center in the eukaryotic cell, crucial for various cellular activities. Since the nucleus is isolated from the cytoplasm by the nuclear envelope (NE), molecules that enter and exit the nucleus must translocate through nuclear pore complexes (NPCs), which are large protein assemblies that span the NE. NPCs provide a size selectivity that allows the passive diffusion of small molecules such as ions and proteins smaller than
30 kDa. The larger molecules such as proteins and RNAs must be actively transported by nucleocytoplasmic transport receptors (NTRs). Research over the past two decades has greatly advanced the understanding of the mechanisms of nucleocytoplasmic transport. The best-characterized NTRs are the members of the importin b (also called karyopherin b, transportin) family. The importin b family-mediated transport pathways are coupled with the GTPase cycle of Ran, which is driven by the chromatin-bound guanine nucleotide exchange factor RCC1 (RanGEF) and the cytoplasmic GTPase activating protein1 (RanGAP1). The asymmetric localization of Ran’s regulators and function of the Ran-GDP import factor p10/NTF2 lead to the nuclear accumulation of Ran-GTP in cells. Ran-GTP regulates the association between the NTRs and their cargoes. Cargo proteins contain

Introduction

specific signals for nuclear transport: nuclear localization signals (NLSs) direct the nuclear import of proteins, whereas nuclear export signals (NESs) specify protein export from the nucleus. Import receptors, importins, bind to their NLS-bearing cargo in the cytoplasm where the concentration of Ran-GTP is low. After translocation into the nucleus, importins release their cargo when bound to Ran-GTP (Stewart, 2007). On the other hand, export receptors, exportins, form ternary complexes within the nucleus that contain both NES-bearing cargo molecules and Ran-GTP. Exportins release their cargo within the cytoplasm upon Ran-GTP hydrolysis by RanGAP1 (Gu¨ttler & Go¨rlich, 2011). In cells that undergo open mitosis, there is obviously no NE to allow for a sharp nucleocytoplasmic Ran-GTP gradient during mitosis. However, RCC1 continuously binds to chromatin; therefore, increased concentration of Ran-GTP is present at the vicinity of mitotic chromosomes. Based on the same principles as in interphase, importins bind to cargoes, which are required for spindle assembly, NPC assembly, and NE assembly in mitotic cytosol. At the vicinity of mitotic chromosomes, importins release their cargoes by binding to Ran-GTP (Kalab & Heald, 2008; Lau et al., 2009). Conversely, the formation of ternary complexes between exportin 1, its specific cargoes, and Ran-GTP at the vicinity of mitotic chromosomes is important for kinetochore fiber formation and chromosome segregation (Arnaoutov et al., 2005). Therefore, the activity and/or localization of cargoes (mitotic regulators) can be regulated by Ran-GTP through binding with NTRs. Importin/exportin-Ran systems thus control multiple cellular processes throughout the cell cycle; while they are essential for nucleocytoplasmic transport, they also play important roles in mitotic progression in the absence of NE (see Fig. 15.3). In vitro transport reconstitution systems using digitonin-permeabilized cells (Adam, Marr, & Gerace, 1990) have been widely used to study nuclear transport, as they are simple to use and enable the analysis of transport in both directions (nuclear import and export). These experimental systems are based on the use of digitonin, a nonionic detergent that, when used at low concentration, permeabilizes the plasma membrane but not the NE. Following permeabilization and subsequent washing steps, the permeabilized cells, which are depleted of shuttling transport factors, are incubated with either cytosol or purified NTR and transport factors such as Ran, an energy-regenerating system, and fluorescently tagged cargoes. Transport of the cargo can then be monitored by either fluorescent microscope or flow cytometry. This system allowed the identification of particular factors involved in specific transport pathways (Adam & Gerace, 1991; Go¨rlich, Prehn, Laskey, & Hartmann, 1994; Imamoto et al., 1995; Pollard et al., 1996). Details of these assays have been previously described (Adam, Sterne-Marr, & Gerace, 1992; Cassany & Gerace, 2009; Kehlenbach & Gerace, 2002; Kehlenbach & Paschal, 2005; Paschal & Gerace, 1995). In this chapter, we first provide an overview of various expression vectors that allow the purification of recombinant NTRs. Such information should be useful for researcher performing not only the in vitro transport assay, but also pull-down experiments to assess interactions between importins and their specific cargoes.

333

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We next detail two recent examples of the use of the in vitro system that closely mimic different cellular conditions: – Nucleocytoplasmic transport of heat-shock protein Hsp70 under normal condition and heat-shock condition (see Figs. 15.1 and 15.2). – The interphase nucleocytoplasmic transport and mitotic chromosome loading of chromokinesin hKid (see Figs. 15.3 and 15.4). These two examples address completely different phenomena. In the first case, physiological regulation impacts different transport pathways in opposite ways: the downregulation of conventional nuclear import pathways and the upregulation of heat-stress-induced transport pathways (see Fig. 15.1). In the second example, the importin b-Ran system is used at different cell-cycle stages (in the presence or

Nucleocytoplasmic transport during heat shock Normal conditions Conventional Importin β-dependent pathway

Heat shock Heat-shock (HS)

NLSs

Importins

Hikeshi-dependent pathway Hsp70s ATP

NLSs

Hsp70s

Importins

Hikeshi

Down regulation

Up regulation

Cytoplasm Nucleus Hsp70s

Recovery

Attenuation HS-phenotype Reversion “Nuclear damage”

Cell death

FIGURE 15.1 Models of Nuclear Transport Under Normal Versus Heat-Shock Condition During heat-shock stress, the conventional nuclear import pathways mediated by importin b-family members are downregulated, whereas the nuclear import pathway of Hsp70s molecular chaperones, which is mediated by Hikeshi, is upregulated. This figure was reproduced from Kose, Furuta, and Imamoto (2012), #2012 with permission from Elsevier.

15.1 Equipment, Material, Reagents, and Buffers

FIGURE 15.2 In vitro Reconstitution of Heat-Shock-Induced Nuclear Import The nuclear transport of recombinant His6-GFP-Hsc70 was examined with the combination of permeabilized cells and cell extracts prepared from either normal or heat-shocked HeLa cells. The efficient nuclear accumulation of Hsc70 is supported only by cytosol extracted from heat-shocked cells. In the same setting, the conventional M9 NLS (GST-M9-GFP) nuclear import mediated by transportin was downregulated. Cytosol prepared from heatshocked cells was separated into importin-containing and importin-depleted fractions using a phenyl-Sepharose column. The nuclear import of Hsc70 was not supported by the importindepleted cytosol but was reconstituted when the importin-containing fraction was supplemented to the depleted cytosol. The nuclear import of Hsc70 was also recovered when bacterially expressed purified recombinant Hikeshi was added to the importin-depleted cytosol. Modified from Kose et al. (2012), #2012 with permission from Elsevier.

absence of NE) but with the same biochemical reaction: the association between importin b and its cargo is regulated by Ran-GTP (see Fig. 15.3).

15.1 EQUIPMENT, MATERIAL, REAGENTS, AND BUFFERS 15.1.1 Equipments 37  C Orbital shaker for bacteria cultures Digital Sonifier 450D (Branson) Rotating wheel Centrifuges

335

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CHAPTER 15 In Vitro Analysis of Nucleocytoplasmic Transport

A

(Nucleotide hydrolysis) Kid α

p10

GDP

Ran

GTP

GTP

Ran

Ran β

RanGAP1 (RanGAP)

GDP

Ran

CAS

β

α _

Cytoplasm

Interphase nucleus GTP

Kid α

GTP

Ran

Cargo release

β

Ran

β

CAS α

(Nucleotide exchange) GDP

p10

GTP

RCC1 (RanGEF)

Ran

Ran

Kid

B _ α

_ α

Spindle binding inhibition

Kid β

β

GTP

Ran

Chromosome targeting

β

Kid α _

Ran

RCC1 (RanGEF)

CAS α _

Kid β

GDP

Mitotic spindle

Ran

GTP

Kid Cargo release Kid and chromosome deposition ? GTP

(Nucleotide exchange)

Kid

Mitotic chromosome

Ran CAS

FIGURE 15.3 Models of Importin a/b–Ran System in Interphase and Mitosis hKid possesses two conventional NLSs and consequently localizes in the nucleus during interphase using the heterodimeric importin a/b nuclear import receptor. The chromosome loading of hKid in mitosis is mediated by importin a/b and Ran systems with the same principle utilized in its interphase nuclear transport (see text for details). Panel B was adapted from Tahara et al. (2008), #2008.

Spinner (Bellco Glass, Inc.) Hand-operated steel homogenizer (Dura-Grind, Wheaton)

15.1.2 Material and reagents Amicon Ultra-4 (10 k MWCO, 50 k MWCO; Millipore) Ampicillin (Nacalai tesque, Inc.)

15.1 Equipment, Material, Reagents, and Buffers

A

B

Interphase +Imp α/β +Ran-GDP +ATP

Mitosis +Imp α/β +Apy

+Imp α/β +Imp α/β +Ran-GDP +ATP +Ran-GDP +ATP +hCAS

DNA

DNA

Imp β

Imp α

FLAG-hKid

hKid

+Imp α/β +Ran-GDP

FIGURE 15.4 Nuclear Import (Interphase) and Chromosome Loading (Mitosis) of hKid (A and B) Nuclear transport and chromosome loading of flag-tagged hKid was examined with purified recombinant transport factors followed by immunofluorescence with an anti-FLAG antibody. (A) The importin a/b-mediated nuclear import of hKid is coupled to the GTPase cycle of Ran during interphase. Nuclear transport of flag-tagged hKid was examined with purified recombinant importin a, importin b, and Ran-GDP, with or without an energy regeneration system (þATP). (B) The importin a/b-mediated chromosome loading of hKid in mitosis is also facilitated by the addition of Ran together with energy sources. Nuclear transport of flag-tagged hKid was examined with purified recombinant CFP-importin a, YFPimportin b, in the presence of either Apyrase (þApy) or Ran-GDP þ an energy regenerating system (Ran-GDP þ ATP), in the presence or absence of recombinant hCAS. Panels A and B are adapted from Tahara et al. (2008), #2008.

Anti-FLAG monoclonal antibody (Sigma) Apyrase (Sigma) ATP-agarose (linked through C-8; Sigma) ATP-REGENERATING SYSTEM – ATP (Sigma): 33 mM ATP in 20 mM HEPES (adjust pH to 7.4 with NaOH), store at 20  C – Creatine phosphate (Calbiochem): 150 mM creatine phosphate in TB without EGTA, store at 20  C – Creatine phosphokinase (Calbiochem): 600 U/ml creatine phosphokinase in TB without EGTA, store at 20  C Dissolve and mix 1:1:1 (ATP:creatine phosphate:creatine phosphokinase) prior to use (this will correspond to a 10 stock solution)

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CHAPTER 15 In Vitro Analysis of Nucleocytoplasmic Transport

CHT-II column (1.5 ml, column prepacked with CHT ceramic hydroxyapatite; #732-4756, Bio-Rad) Dialysis tubes (10 and 50 kDa cutoff ) Digitonin (Calbiochem): prepare stock 20 mg/ml in DMSO, store at 20  C. Formaldehyde (methanol-free) (16% UltraPure EM grade) (Polysciences) Glutathione Sepharose 4 Fast Flow (GE Healthcare) Glutathione (GE Healthcare) Imidazole Isopropyl-b-D-thiogalactopyranoside (IPTG) Microcon YM-30 columns (30 k MWCO; Millipore) MonoQ column (GE Healthcare) MonoS column (GE Healthcare) Multitest 8-wells slides (MP Biomedicals LLC) Ni-NTA agarose (Qiagen) PD10 column (GE Healthcare) Phenyl-Sepharose Fast Flow (low sub; GE Healthcare) PreScission Protease (GE Healthcare) Protease inhibitors: aprotinin, leupeptin, and pepstatin Thymidine (Sigma) Thrombin (Sigma Chemical Co., St. Louis, MO)

15.1.3 Strain and cell line Escherichia coli strain BL21(DE3) Following transformation with plasmids, E. coli are grown in LB þ 50 mg/ml ampicillin. HeLa-S3 (RIKEN Cell Bank) HeLa cells are grown in RPMI-1640 medium supplemented with 5% FBS at 37  C with a 5% CO2 atmosphere. For heat-shock treatments, the medium is further complemented by 20 mM HEPES (pH 7.3).

15.1.4 Buffers 0.9% NaCl Transport Buffer (TB): 20 mM HEPES (pH 7.3), 110 mM potassium acetate, 2 mM magnesium acetate, 5 mM sodium acetate, 0.5 mM EGTA, 2 mM DTT, and 1 mg/ml each of aprotinin, pepstatin A, and leupeptin FOR PURIFICATION OF GST-TAGGED IMPORTINS Bacteria lysis buffer: 50 mM Tris–HCl (pH 8.3), 500 mM NaCl, 1 mM DTT, and 2 mM PMSF Elution buffer: 100 mM Tris–HCl (pH 7.3), 100 mM NaCl, 20 mM glutathione, 1 mM DTT, and 1 mg/ml each of aprotinin, pepstatin, and leupeptin

15.2 Purification of Recombinant Transport Factors

FOR PREPARATION OF CYTOSOL Wash buffer: 50 mM Tris–HCl (pH 7.3), 50 mM NaCl, 1 mM phenylmethylsulfonyl fluoride (PMSF), 2 mM dithiothreitol (DTT), and 1 mg/ml each of aprotinin, pepstatin A, and leupeptin Lysis buffer: wash buffer containing 5 mM MgOAc and 20 mM cytochalasin B Elution buffer: 50% ethylene glycol in 50 mM Tris–HCl (pH 7.3), 2 mM DTT, and 1 mg/ml each of aprotinin, pepstatin A, and leupeptin FOR PURIFICATION OF HIS-TAGGED GFP-HSC70 Bacteria lysis buffer: 50 mM Tris–HCl (pH 8.0), 300 mM NaCl, 1 mM 2-mercaptoethanol, and 2 mM PMSF Purification buffer I: 20 mM HEPES (pH 7.3), 100 mM NaCl, 1 mM 2-mercaptoethanol, 1 mg/ml each of aprotinin, pepstatin, and leupeptin Purification buffer II: buffer I containing 5 mM MgCl2 FOR PURIFICATION OF GST-FLAG-HKID Buffer A: 50 mM Tris–HCl (pH 8.3), 500 mM NaCl, 2 mM DTT, and 200 mM PMSF Buffer P: buffer A containing 0.1% Tween 20 and 1 mg/ml each of aprotinin, leupeptin, and pepstatin A Buffer B: 50 mM Tris–HCl (pH 8.8), 500 mM NaCl, 2 mM DTT, 0.1% Tween 20, and 1 mg/ml each of aprotinin, leupeptin, and pepstatin A Buffer C: 20 mM HEPES (pH 7.3), 400 mM NaCl, 2 mM DTT, and 1 mg/ml each of aprotinin, leupeptin, and pepstatin A

15.2 PURIFICATION OF RECOMBINANT TRANSPORT FACTORS Ran-GDP was expressed in E. coli and purified as previously described (Dasso, Seki, Azuma, Ohba, & Nishimoto, 1994; Hieda et al., 1999; Melchior, Sweet, & Gerace, 1995). See also Chapter 16 for purification of GST-Ran. Bacterial expression vectors for several importin b NTRs and transport factors (p10/NTF2, importin a) fused to glutathione-S-transferase (GST) or other tags have been previously described (see Cassany & Gerace, 2009; Kose, Imamoto, Tachibana, Shimamoto, & Yoneda, 1997). Those listed in Table 15.1 are available from our lab. Various purification procedures have been previously reported (see original references indicated in Table 15.1 and Cassany & Gerace, 2009). Below is a typical protocol that can be adapted to all importin b recombinant NTRs listed in Table 15.1. Note: The expression and purification conditions are same for all importin b NTRs (including the importin a export receptor CAS). The amount that can be purified from 1 l culture differs between proteins. For importin b, transportin, RanBP5, and CAS, we generally obtain
5–10 mg/l of E. coli culture, while for some other NTRs the yield is
1–2 mg/l.

339

Table 15.1 Plasmid information for the purification of recombinant NTRs Transport factor

Gene name

Human ID

Other designations

MW (kDa)

Plasmid backbone

Tag

Importin ß

KPNB1

3837

Karyopherin ß1; NTF97; PTAC97

97

pGEX2T/ HA pGEX6P1/EYFP pGEX6P3/FLAG

GSTHA-N GSTEYFP -N GSTFLAG-N

Remove GST-tag using

Original publication(s)

Thrombin

Kose et al. (1997)

PreScission Protease PreScission Protease

Tahara et al. (2008)

Importin 4

IPO4

79711

RanBP4

119

Importin 5

IPO5

3843

124

pGEX6P1-GW

GST-N

PreScission Protease

Importin 7

IPO7

10527

RanBP5; Karyopherin ß3 RanBP7

120

Importin 8

IPO8

10526

RanBP8

120

Importin 9

IPO9

55705

RanBP9

116

pGEX6P3/FLAG pGEX6P3/FLAG pGEX6P-2

GSTFLAG-N GSTFLAG-N GST-N

Importin 11

IPO11

51194

RanBP11

113

Importin 13

IPO13

9670

108

GSTFLAG-N GST-N

Transportin 1

TNPO1

3842

RanBP13; Karyopherin 13 Karyopherin ß2

pGEX6P1/FLAG pGEX6P-3

PreScission Protease PreScission Protease PreScission Protease PreScission Protease PreScission Protease

pGEX6P3/FLAG

GSTFLAG-N

101

PreScission Protease

Rout, Blobel, and Aitchison (1997), Schlenstedt et al. (1997) Deane et al. (1997)

Ja¨kel et al. (1999) Go¨rlich et al. (1997) Mu¨hlha¨usser, Mu¨ller, Otto, and Kutay (2001) Plafker and Macara (2000) Mingot, Kostka, Kraft, Hartmann, and Go¨rlich (2001) Pollard et al. (1996)

Transportin 2

TNPO2

30000

Karyopherin ß2b

100

pGEX6P-3

GST-N

PreScission Protease

Transportin 3

TNPO3

23534

TransportinSR

104

RanBP6

RANBP6

26953

125

RanBP10

RANBP10

57610

67

CAS

CSE1L

1434

pGEX6P3/FLAG pGEX6P1/FLAG pGEX6P1/FLAG pGEX6P-1

GSTFLAG-N GSTFLAG-N GSTFLAG-N GST-N

Importin a1

KPNA2

3838

pGEX6P1/FLAG pGEX6P1/ECFP pET-3a

GSTFLAG-N GSTECFP-N No tag

PreScission Protease PreScission Protease PreScission Protease PreScission Protease PreScission Protease PreScission Protease –

pET-16b/ pQE80L

No tag/ 6xHis-N

Exportin 2 (XPO2), CSE1 Rch1; hSRP1; Karyopharin a2; PTAC58

110 58

NTF2

NUTF2

10204

Nuclear transport factor 2; p10

14

Ran

RAN

5901

TC4; Rasrelated nuclear protein

24

–

Gu¨ttinger, Mu¨hlha¨usser, Koller-Eichhorn, Brennecke, and Kutay (2004) Kataoka, Bachorik, and Dreyfuss (1999)

Schulze et al. (2008) Kutay, Bischoff, Kostka, Kraft, and Go¨rlich (1997) Kose et al. (1997) Tahara et al. (2008) Paschal and Gerace (1995), Tachibana, Hieda, Sekimoto, and Yoneda (1996) Dasso et al. (1994), Melchior et al. (1995), Hieda et al. (1999)

Original publication(s) refer to either initial publication of the listed factor or initial publication of purification methods described in this chapter.

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CHAPTER 15 In Vitro Analysis of Nucleocytoplasmic Transport

PROTOCOL: 1. Transform E. coli strain BL21(DE3) with the NTR expression vectors (see Table 15.1). Grow E. coli cells in 4 l of LB medium containing 50 mg/ml ampicillin at 37  C to a density of 0.6 (OD550). Bacteria are chilled in the cold room before induction to decrease the temperature to 20  C. 2. Induce expression by addition of 0.1 mM IPTG and incubate for 12–18 h at 20  C. 3. Harvest the cells by centrifugation at 1500  g for 15 min at 4  C. 4. Wash the cells with 0.9% NaCl. 5. Harvest the cells and resuspend them in 40-ml lysis buffer. 6. After cell disruption by freeze–thaw (freeze in liquid N2 and thaw, twice) and sonication (on ice, 30–40% amplitude, three times for 1 min each), clarify the extract by centrifugation at 200,000  g for 15 min. 7. Incubate the extract with 1 ml bed volume of Glutathione Sepharose previously equilibrated with lysis buffer for 1 h at 4  C on a rotating wheel. 8. Wash the beads with 10 ml of lysis buffer, and elute the recombinant proteins with 1 ml of elution buffer. 9. Cleave off the GST portion of chimeras by 2 h incubation at room temperature with 1 NIH unit of thrombin per 100 mg of chimeras. In case of GST proteins derived from PGEX6p vectors, incubate overnight at 4  C with 1 unit of PreScission Protease per 100 mg of chimeras. 10. Separate GST and thrombin or PreScission Protease from recombinant proteins on a MonoQ column at flow rate of 0.5 ml/min with a linear gradient from 0.1 to 1.0 M NaCl in 20 mM HEPES (pH 7.3), 2 mM DTT, 1 mg/ml each of aprotinin, leupeptin, and pepstatin. 11. Dialyze the recombinant proteins against transport buffer containing 2 mM DTT and concentrate using an AmiconUltra-4 (50 k MWCO; Millipore) when necessary (protein concentration of above 1 mg/ml is convenient for use). Snapfreeze aliquots in liquid nitrogen and store at 80  C.

15.3 USE OF DIGITONIN-PERMEABILIZED CELLS TO STUDY NUCLEAR TRANSPORT UNDER NORMAL AND HEAT-SHOCK CONDITIONS This approach has been initially published in Kose et al. (2012). Cellular stresses significantly affect nuclear transport systems. During heatshock stress, the conventional nuclear import pathway is downregulated (Furuta et al., 2004; Kodiha, Chu, Matusiewicz, & Stochaj, 2004; Miyamoto et al., 2004), whereas the nuclear import of molecular chaperone Hsp70s is upregulated (Pelham, 1984; Velazquez & Lindquist, 1984; Welch & Feramisco, 1984). The combination of the digitonin-permeabilized cells and cytosol prepared from either normal or heat-shock-treated HeLa cells mimics the nuclear import as observed in living

15.3 Use of Digitonin-Permeabilized Cells

control cells or heat-shocked cells, respectively (Fig. 15.2). With this approach, we found that efficient nuclear accumulation of the constitutive member of the Hsp70 (Hsc70) was supported only by cytosol extracted from heat-shocked cells and not by heat-shock-treated permeabilized cells. In the same setting, the conventional nuclear import mediated by importin b family members, importin b and transportin, was suppressed (Fig. 15.2). Downregulation of the importin b family member mediated import and upregulation of Hsp70s nuclear import thus mimic reactions observed in heat-shocked living cells. Nuclear transport carriers are thought to have hydrophobic properties necessary to translocate through NPCs and are enriched by phenyl-Sepharose beads under stringent binding conditions (Ribbeck & Go¨rlich, 2002). This principle could be applied to fractionate transport factor(s) required for the nuclear import of Hsc70. Cytosol prepared from heat-shocked cells was separated into importin-containing and importin-depleted fractions using a phenyl-Sepharose column (Fig. 15.2). The nuclear import of Hsc70 was not supported by the importin-depleted fraction but was reconstituted when the importin-containing fraction was supplemented to the depleted fraction (Fig. 15.2). By further biochemical purification from the importin-containing fraction, we identified a novel transport pathway that was mediated by a novel factor called Hikeshi. The nuclear import of Hsc70 was recovered when bacterially expressed Hikeshi was added to the importin-depleted fraction. Unlike importin b family-mediated nuclear transport, the Hikeshi-mediated nuclear import of Hsp70s is not coupled to the GTPase cycle of Ran but instead to the ATPase cycle of Hsc70.

15.3.1 Preparation of cytosol Note: A slightly different procedure also compatible with the nuclear transport assay has also been previously described (Cassany & Gerace, 2009; Kehlenbach & Gerace, 2002; Kehlenbach & Paschal, 2005). 1. Culture HeLa-S3 cells in a spinner bottle at 37  C with a 5% CO2 atmosphere. Expand the cell culture every 24 h with RPMI-1640 medium supplemented with 5% FBS, keeping the cell density at 2–5  105 cells/ml. Start the extract preparation when the cells reach 4–5  105cells/ml in 3–6 l medium. For preparation of the heat-shocked cytosol, incubate cells at 43  C for 1 h with prewarmed RPMI-1640 medium/5% FBS/20 mM HEPES (pH 7.3). 2. Collect cells by centrifugation at 800  g for 5 min at 4  C. All of the following steps are performed on ice or at 4 C. 3. Wash twice with PBS by suspending and centrifuging at 800  g for 5 min. Cells from a 6 l culture (
10 g) can be spun in a 50-ml conical tube in the second washing step. Weigh the cell pellet. 4. Wash once with 50 ml of wash buffer. 5. Resuspend in 1 volume (1 ml/g of cell pellet) of lysis buffer and swell the cells for 10 min on ice.

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CHAPTER 15 In Vitro Analysis of Nucleocytoplasmic Transport

6. Lyse the cells with 10–15 strokes of a hand-operated steel homogenizer. 7. Centrifuge the homogenate at 800  g for 10 min. The supernatant is used for cytosolic extract preparation, whereas the pellet is used for the nuclear extract (see Chapter 16). Clear the supernatant by centrifugation at 200,000  g for 15 min. 8. Dialyze overnight against 4 l transport buffer (10 kDa cutoff ). The next day, remove aggregates by centrifugation at 200,000  g for 15 min, aliquot the resulting cytosol in 100 ml aliquots, freeze in liquid nitrogen and store at 80  C.

15.3.2 Preparation of fractionated cytosol (importin-depleted/ containing fractions) The retention of importin-b family NTRs on phenyl-Sepharose (Ribbeck & Go¨rlich, 2002) is used to prepare importin-depleted and importin-containing fractions. 1. Incubate the cytosolic extracts with 20% bed volume of phenyl-Sepharose equilibrated with lysis buffer and rotate the mixture gently for 1 h. Recover the extract after centrifugation at 800  g for 5 min. Keep the phenyl-Sepharose beads for step 3. Repeat this depletion procedure. 2. After centrifugation at 800  g for 5 min, take flow through as the importindepleted fraction. Protein concentrations of this depleted cytosolic extracts should be
10 mg/ml. 3. Combine the phenyl-Sepharose beads and wash them five times with 5 volumes of lysis buffer. 4. Elute the bound materials (importin-containing fraction) with 1 bed volume of elution buffer. Repeat this elution procedure five times. 5. Dialyze both importin-depleted cytosol and importin-containing fractions against transport buffer. Concentrate the importin-containing fraction using an Amicon Ultra-4 (10 k MWCO) approximately to 1/10 volume. Aliquot the importindepleted cytosol in 50 ml aliquots, the importin-containing fraction in 5 ml aliquots, freeze in liquid nitrogen, and store at 80  C.

15.3.3 Preparation of fluorescent His-tagged GFP-Hsc70 substrate Detailed purification of several other import substrates (e.g., SV40 T antigen basic NLS conjugated to fluorescently labeled BSA) can be found in Cassany and Gerace (2009). However, in general, we use genetically manipulated model cargo substrates, such as NLSs sequences cloned in between GST and GFP. These recombinant import substrates are purified from E. coli on Glutathione Sepharose essentially as described for GST-tagged NTFs in Section 15.2, except that bacteria are chilled in the cold room before harvest for 2 h. Indeed, the chilling often enhances the fluorescent intensity of GFP-fusion proteins.

15.3 Use of Digitonin-Permeabilized Cells

Note: When NLSs are fused to the C-terminus of GST or GFP, they can be easily degraded. In Fig. 15.2 for example, we used the M9-NLS fused between GST and GFP, which is a typical cargo for transportin 1 (Yokoya, Imamoto, Tachibana, & Yoneda, 1999). PURIFICATION OF HIS6-GFP-HSC70 Note: Some common purification steps are further detailed in Section 15.2. 1. Transform E. coli strain BL21 (DE3) with His-tagged GFP-Hsc70 expression pQE80L vector. Grow E. coli cells in 4 l of LB medium containing 50 mg/ml ampicillin at 37  C to a density of 0.6 (OD550). Bacteria are chilled in the cold room before induction to decrease the temperature to 20  C. 2. Induce expression with the addition of 0.1 mM IPTG and incubate for 14 h at 20  C. 3. Bacteria are chilled in the cold room before harvest for 2 h. Harvest the cells by centrifugation at 1500  g for 15 min at 4  C. 4. Wash the cells with 0.9% NaCl. 5. Harvest the cells and resuspend them in 20 ml lysis buffer. 6. After cell disruption by freeze–thaw and sonication as described above for importins, clarify the extract by centrifugation at 200,000  g for 15 min. 7. Incubate the extract with 1 ml bed volume of Ni-NTA agarose, equilibrated with bacteria lysis buffer, for 1 h at 4  C on a rotating wheel. 8. Wash the beads with 10 ml of lysis buffer containing 10 mM imidazole, and elute the recombinant proteins with 1 ml buffer I containing 250 mM imidazole. 9. Incubate the eluate for 2 h at 4  C on a rotating wheel with 1 ml of ATP-agarose (linked through C-8; Sigma). 10. Wash the beads with 10 ml buffer II, and elute the bound proteins with 1 ml buffer II containing 5 mM ATP. 11. Load onto a CHT-II column (Bio-Rad), and elute the recombinant proteins with the linear gradient of 10–250 mM potassium phosphate (pH 7.3) in 100 mM NaCl and 1 mM DTT at flow rate of 0.5 ml/min. 12. Dialyze against transport buffer (containing 2 mM DTT), measure protein concentration and concentrate to 1 mg/mL using an AmiconUltra-4 (50 k MWCO; Millipore). Snap-freeze aliquots in liquid nitrogen and store at 80  C.

15.3.4 Preparation of permeabilized cell All of the following steps are performed in a 10-cm plastic dish containing a multitest 8-well slide. 1. 24–36 h prior to the import reaction, seed 2  106 HeLa-S3 cells in 10-cm plastic dishes containing an 8-well slide (diameter, 6 mm). For the preparation of heat-shocked permeabilized cell, incubate cells at 43  C for 1 h with prewarmed RPMI-1640 medium/5% FBS/20 mM HEPES (pH 7.3).

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2. Wash cells twice with 15 ml of cold transport buffer in the 10-cm plastic dish. Permeabilize cells with 15 ml of digitonin-containing transport buffer (final concentration of 40 mg/ml) for 5 min on ice. Note: The permeabilization temperature is very important in our hands, but different temperatures are used in other protocols (see Cassany & Gerace, 2009). 3. Wash cells twice with cold transport buffer, and preincubate cells with 15 ml transport buffer for 5 min on ice. Subject cells to transport reaction.

15.3.5 Transport assay The transport reaction mix contains – Cytosol (5 ml at
10 mg/mL) prepared from either normal or heat-shock-treated HeLa cells) or fractionated cytosols (5 ml of the importin-depleted cytosol either alone, or combined with either 0.5 ml of the importin-containing fraction or purified Hikeshi (2 mM final concentration of recombinant protein)). – Fluorescent substrates (0.8 mM His6-GFP-Hsc70 or GST-M9-GFP). – ATP-regenerating system (final concentration of 1 mM ATP, 5 mM creatine phosphate, 20 U/ml creatine phosphokinase). 1. Use 10 ml transport reaction per well in an 8-well slide and incubate for 20 min at 30  C in a moisturized plastic dish. (Use the permeabilized cells prepared from either normal or heat-shock-treated HeLa cells.) 2. Wash twice with cold transport buffer. 3. Fix cells with 3.7% formaldehyde in transport buffer for 10 min at room temperature. 4. Analyze the cells by fluorescence microscopy, using appropriate filters. Examples of nuclear import reactions using recombinant Hsc70 and M9-NLS are shown in Fig. 15.2.

15.4 INTERPHASE NUCLEOCYTOPLASMIC TRANSPORT AND MITOTIC CHROMOSOME LOADING OF CHROMOKINESIN hKid This approach was initially described in Tahara et al. (2008). The cellular function of the chromokinesin human kinesin-like DNA-binding protein (hKid) is better characterized in mitotic cells than in the interphase nucleus. In metaphase, hKid promotes polar ejection force through binding to both mitotic spindle and mitotic chromosomes, and regulates anaphase onset. For this activity, a large population of hKid is localized on mitotic chromosomes (Funabiki & Murray, 2000; Tokai et al., 1996). hKid possesses two conventional NLSs and consequently localizes in the nucleus during interphase using the heterodimeric importin a/b nuclear import receptor (Tokai et al., 1996; see also Fig. 15.3A). Using the in vitro system, we could

15.4 Interphase Nucleocytoplasmic Transport and Mitotic Chromosome

reconstitute the interphase nuclear import of hKid using either cytosol or purified recombinant transport factors that are present in the cytosol required for the reaction: importin a/b, Ran-GDP, energy regeneration system, and flag-tagged hKid (Tahara et al., 2008; Figs. 15.3A and 15.4A). We also examined the mitotic behavior of hKid in digitonin-permeabilized mitotic cells using purified recombinant NTRs, importin a, and Ran (Tahara et al., 2008). In the presence of importin a/b, hKid efficiently targeted mitotic chromosomes (Fig. 15.4B, left panel). This effect of importin a/b on hKid did not depend on the mitotic spindle, because treatment of nocodazole, which disrupts the mitotic spindle, did not affect the importin a/b-dependent chromosome targeting of hKid. Our data further showed that (1) the addition of Ran, together with energy sources (ATP and its regenerating system), causes further accumulation of hKid, while cotargeted importin b dissociates from chromosomes (Fig. 15.4B central panel). (2) Addition of CAS, an importin family NTR mediating nuclear export of importin a, tends to release cotargeted importin a from chromosomes (Fig. 15.4B right panel). The use of digitonin-permeabilized cells revealed that the chromosome loading of hKid in mitosis is mediated by importin a/b and the Ran system, with the same principle utilized in its interphase nuclear transport (Fig. 15.3B).

15.4.1 Preparation of GST-FLAG-hKid Note: Some common purification steps are further detailed in Section 15.2 1. Transform E. coli strain BL21(DE3) with the GST- and FLAG-tagged hKid expression pGEX vectors. Grow E. coli cells in 4 l of LB medium containing 50 mg/ml ampicillin at 37  C to a density of 0.6 (OD550). Bacteria are chilled in the cold room before induction to decrease the temperature to 20  C. 2. Induce expression by addition of 0.1 mM IPTG and incubate for 18 h at 20  C. 3. Harvest the cells by centrifugation at 1500  g for 15 min at 4  C. 4. Wash the cells with 0.9% NaCl. 5. Harvest the cells and resuspend them in buffer A. 6. After cell disruption by freeze–thaw and sonication, clarify the extract by centrifugation at 200,000  g for 15 min. 7. Incubate the extract with Glutathione-Sepharose 4B (GE Healthcare) for 1 h at 4  C on a rotating wheel. 8. Wash the beads with buffer P and elute the recombinant proteins with buffer B containing 20 mM glutathione. 9. Cleave off GST with PreScission Protease. 10. Separate GST and PreScission Protease from recombinant proteins on a MonoS column and elute the recombinant proteins with a linear gradient of 0.25–1.0 M NaCl in 20 mM HEPES (pH 7.3) and 2 mM DTT. 11. Adjust the NaCl concentration to 1 M. Concentrate the proteins by ultrafiltration on Microcon YM-30 columns and desalt using a PD10 column equilibrated with buffer C.

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15.4.2 Preparation of permeabilized mitotic cells 1. 36–48 h prior to the reaction, seed 2  105 cells/ml HeLa-S3 cells in 10-cm plastic dishes containing an 8-well slide. 2. To enrich for mitotic cells, cells are treated with 2 mM thymidine for 16 h, rinsed twice with prewarmed PBS and once with culture medium, and released into fresh medium for 8 h. After a second 16 h thymidine block, cells are released into fresh medium 8–9 h prior to the assay (about 80% of cells are mitotic). Note: To determine the contribution of spindles on the chromosome loading of recombinant proteins, cells are treated with 2 mM nocodazole for 2 h prior to digitonin-permeabilization. 3. Wash cells twice with 15 ml of cold transport buffer in the 10-cm plastic dish. Then permeabilize cells with 15 ml of digitonin-containing transport buffer (final concentration of 40 mg/ml) for 5 min on ice. 4. Wash cells twice with cold transport buffer and preincubate with 15 ml transport buffer for 5 min on ice. Subject cells to transport reactions.

15.4.3 In vitro chromosome-loading reaction The transport reaction mix contains: 0.4 mM importin a, 0.4 mM importin b, 4 mM Ran-GDP, 0.4 mM FLAG-tagged substrate (hKid), an ATP-regenerating system (final concentration of 1 mM ATP, 5 mM creatine phosphate, 20 U/ml creatine phosphokinase). In some case, we also included 1.7 mM recombinant CAS. Note: Expression and purification of recombinant importin a, importin b, and CAS was performed as described above in Section 15.2 for the other importin NTRs. Ran-GDP was expressed and purified as described (see references in Section 15.2 and Table 15.1). For experiments performed in the absence of ATP, add 0.1 unit/ml apyrase to completely deplete residual ATP retained in the permeabilized cell. Note: Addition of apyrase is also applicable to the interphase transport assay. However, note that apyrase sometimes leads to the degradation of some purified recombinant substrate proteins. 1. Use 10 ml transport reaction per well of an 8-well slide and incubate for 20 min at 30  C in moisturized plastic dishes (use the permeabilized cell prepared from either normal or mitotic HeLa cells in the same transport reaction setting). 2. Wash twice with cold transport buffer. 3. Fix cells with 3.7% formaldehyde in transport buffer for 10 min at room temperature and treat with 0.2% Triton X-100 for 5 min. 4. Subject cells to indirect immunofluorescence. For FLAG-hKids, we used antiFLAG monoclonal antibody combined with Cy3-conjugated secondary antibodies. Counterstain DNA with DAPI. 5. Analyze the cells by fluorescence microscopy, using appropriate filters. Examples of nuclear import and chromatin-binding reactions of hKid are shown in Fig. 15.4.

References

CONCLUSIONS AND PERSPECTIVES Digitonin-permeabilized cell-free assays, initially developed by Adam et al. (1990) for nuclear import assay, can be used in a wider variety of experiments than initially anticipated. Historically, this system allowed the identification of the first nuclear import receptors, importin a and importin b. The digitonin-permeabilized cell assay can be used to mimic interphase nuclear transport under different cellular conditions with many different substrates. In Chapter 16, Kimura et al. describe the application of the digitonin-permeabilized cell system for substrate determination using MS/MS. In addition, we also detail in this chapter how digitonin-permeabilized cells can be used to investigate events in mitotic cells. In Chapter 12, Marino et al. further describe how this assay can be adapted to study NE breakdown and NPC disassembly at mitotic entry, using semipermeabilized cells expressing fluorescent NE marker proteins. However, care must be taken when these assays are performed. Different digitonin concentrations or different permeabilization conditions (temperature, permeabilization time) are used in different labs, as can be seen in previous protocols (Cassany & Gerace, 2009). This may depend on cell-type used or on the cellular conditions to be examined. Digitonin concentrations as well as permeabilization conditions must be carefully titrated before performing experiments. Furthermore, the activity of cytosol or transport factors at the beginning of experiments must be checked. The reproducibility of the results is also a key aspect that has to be validated.

Acknowledgments We are grateful to members of Cellular Dynamics Laboratory. Our work was supported by the Japan Society for the Promotion of Science (JSPS) through the “Funding Program for Next Generation World-Leading Researchers (NEXT Program),” initiated by the Council for Science and Technology Policy (CSTP), and MEXT grant-in aids.

References Adam, S. A., & Gerace, L. (1991). Cytosolic proteins that specifically bind nuclear location signals are receptors for nuclear import. Cell, 5, 837–847. Adam, S. A., Marr, R. S., & Gerace, L. (1990). Nuclear protein import in permeabilized mammalian cells requires soluble cytoplasmic factors. The Journal of Cell Biology, 111, 807–816. Adam, S. A., Sterne-Marr, R., & Gerace, L. (1992). Nuclear protein import using digitoninpermeabilized cells. Methods in Enzymology, 219, 97–110. Arnaoutov, A., Azuma, Y., Ribbeck, K., Joseph, J., Boyarchuk, Y., Karpova, T., et al. (2005). Crm1 is a mitotic effector of Ran-GTP in somatic cells. Nature Cell Biology, 6, 626–632. Cassany, A., & Gerace, L. (2009). Reconstitution of nuclear import in permeabilized cells. Methods in Molecular Biology, 464, 181–205.

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Dasso, M., Seki, T., Azuma, Y., Ohba, T., & Nishimoto, T. (1994). A mutant form of the Ran/ TC4 protein disrupts nuclear function in Xenopus laevis egg extracts by inhibiting the RCC1 protein, a regulator of chromosome condensation. The EMBO Journal, 13, 5732–5744. Deane, R., Scha¨fer, W., Zimmermann, H. P., Mueller, L., Go¨rlich, D., Prehn, S., et al. (1997). Ran-binding protein 5 (RanBP5) is related to the nuclear transport factor importin-beta but interacts differently with RanBP1. Molecular and Cellular Biology, 17, 5087–5096. Funabiki, H., & Murray, A. W. (2000). The Xenopus chromokinesin Xkid is essential for metaphase chromosome alignment and must be degraded to allow anaphase chromosome movement. Cell, 102, 411–424. Furuta, M., Kose, S., Koike, M., Shimi, T., Hiraoka, Y., Yoneda, Y., et al. (2004). Heat-shock induced nuclear retention and recycling inhibition of importin alpha. Genes to Cells, 9, 429–441. Go¨rlich, D., Dabrowski, M., Bischoff, F. R., Kutay, U., Bork, P., Hartmann, E., et al. (1997). A novel class of RanGTP binding proteins. The Journal of Cell Biology, 138, 65–80. Go¨rlich, D., Prehn, S., Laskey, R. A., & Hartmann, E. (1994). Isolation of a protein that is essential for the first step of nuclear protein import. Cell, 5, 767–778. Gu¨ttinger, S., Mu¨hlha¨usser, P., Koller-Eichhorn, R., Brennecke, J., & Kutay, U. (2004). Transportin2 functions as importin and mediates nuclear import of HuR. Proceedings of the National Academy of Sciences of the United States of America, 101, 2918–2923. Gu¨ttler, T., & Go¨rlich, D. (2011). Ran-dependent nuclear export mediators: A structural perspective. The EMBO Journal, 17, 3457–3474. Hieda, M., Tachibana, T., Yokoya, F., Kose, S., Imamoto, N., & Yoneda, Y. (1999). A monoclonal antibody to the COOH-terminal acidic portion of Ran inhibits both the recycling of Ran and nuclear protein import in living cells. The Journal of Cell Biology, 144, 645–655. Imamoto, N., Shimamoto, T., Takao, T., Tachibana, T., Kose, S., Matsubae, M., et al. (1995). In vivo evidence for involvement of a 58 kDa component of nuclear pore-targeting complex in nuclear protein import. The EMBO Journal, 15, 3617–3626. Ja¨kel, S., Albig, W., Kutay, U., Bischoff, F. R., Schwamborn, K., Doenecke, D., et al. (1999). The importin beta/importin 7 heterodimer is a functional nuclear import receptor for histone H1. The EMBO Journal, 18, 2411–2423. Kalab, P., & Heald, R. (2008). The RanGTP gradient—A GPS for the mitotic spindle. Journal of Cell Science, 121, 1577–1586. Kataoka, N., Bachorik, J. L., & Dreyfuss, G. (1999). Transportin-SR, a nuclear import receptor for SR proteins. The Journal of Cell Biology, 145, 1145–1152. Kehlenbach, R. H., & Gerace, L. (2002). Analysis of nuclear protein import and export in vitro using fluorescent cargoes. In E. Manser & T. Leung (Eds.), Methods in molecular biology, Vol. 189. (pp. 231–245). New York, NY: Springer. Kehlenbach, R. H., & Paschal, B. M. (2005). Analysis of nuclear protein transport in vitro. In J. Celis (Ed.), Cell biology: a laboratory handbook, Vol. 2. (pp. 267–276). Amsterdam, Netherlands: Elsevier Science Publishers. Kodiha, M., Chu, A., Matusiewicz, N., & Stochaj, U. (2004). Multiple mechanisms promote the inhibition of classical nuclear import upon exposure to severe oxidative stress. Cell Death and Differentiation, 11, 862–874. Kose, S., Furuta, M., & Imamoto, N. (2012). Hikeshi, a nuclear import carrier for Hsp70s, protects cells from heat shock-induced nuclear damage. Cell, 149, 578–589.

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Kose, S., Imamoto, N., Tachibana, T., Shimamoto, T., & Yoneda, Y. (1997). Ran-unassisted nuclear migration of a 97-kD component of nuclear pore-targeting complex. The Journal of Cell Biology, 139, 841–849. Kutay, U., Bischoff, F. R., Kostka, S., Kraft, R., & Go¨rlich, D. (1997). Export of importin alpha from the nucleus is mediated by a specific nuclear transport factor. Cell, 90, 1061–1071. Lau, C. K., Delmar, V. A., Chan, R. C., Phung, Q., Bernis, C., Fichtman, B., et al. (2009). Transportin regulates major mitotic assembly events: From spindle to nuclear pore assembly. Molecular Biology of the Cell, 20, 4043–4058. Melchior, F., Sweet, D. J., & Gerace, L. (1995). Analysis of Ran/TC4 function in nuclear protein import. Methods in Enzymology, 257, 279–291. Mingot, J. M., Kostka, S., Kraft, R., Hartmann, E., & Go¨rlich, D. (2001). Importin 13: A novel mediator of nuclear import and export. The EMBO Journal, 20, 3685–3694. Miyamoto, Y., Saiwaki, T., Yamashita, J., Yasuda, Y., Kotera, I., Shibata, S., et al. (2004). Cellular stresses induce the nuclear accumulation of importin alpha and cause a conventional nuclear import block. The Journal of Cell Biology, 165, 617–623. Mu¨hlha¨usser, P., Mu¨ller, E. C., Otto, A., & Kutay, U. (2001). Multiple pathways contribute to nuclear import of core histones. EMBO Reports, 2, 690–696. Paschal, B. M., & Gerace, L. (1995). Identification of NTF2, a cytosolic factor for nuclear import that interacts with nuclear pore complex protein p62. The Journal of Cell Biology, 129, 925–937. Pelham, H. R. (1984). Hsp70 accelerates the recovery of nucleolar morphology after heat shock. The EMBO Journal, 3, 3095–3100. Plafker, S. M., & Macara, I. G. (2000). Importin-11, a nuclear import receptor for the ubiquitin-conjugating enzyme, UbcM2. The EMBO Journal, 19, 5502–5513. Pollard, V. W., Michael, W. M., Nakielny, S., Siomi, M. C., Wang, F., & Dreyfuss, G. (1996). A novel receptor-mediated nuclear protein import pathway. Cell, 6, 985–994. Ribbeck, K., & Go¨rlich, D. (2002). The permeability barrier of nuclear pore complexes appears to operate via hydrophobic exclusion. The EMBO Journal, 21, 2664–2671. Rout, M. P., Blobel, G., & Aitchison, J. D. (1997). A distinct nuclear import pathway used by ribosomal proteins. Cell, 89, 715–725. Schlenstedt, G., Smirnova, E., Deane, R., Solsbacher, J., Kutay, U., Go¨rlich, D., et al. (1997). Yrb4p, a yeast ran-GTP-binding protein involved in import of ribosomal protein L25 into the nucleus. The EMBO Journal, 16, 6237–6249. Schulze, H., Dose, M., Korpal, M., Meyer, I., Italiano, J. E., & Shivdasani, R. A. (2008). RanBP10 is a cytoplasmic guanine nucleotide exchange factor that modulates noncentrosomal microtubules. The Journal of Biological Chemistry, 283, 14109–14119. Stewart, M. (2007). Molecular mechanism of the nuclear protein import cycle. Nature Reviews Molecular Cell Biology, 3, 195–208. Tachibana, T., Hieda, M., Sekimoto, T., & Yoneda, Y. (1996). Exogenously injected nuclear import factor p10/NTF2 inhibits signal-mediated nuclear import and export of proteins in living cells. FEBS Letters, 397, 177–182. Tahara, K., Takagi, M., Ohsugi, M., Sone, T., Nishiumi, F., Maeshima, K., et al. (2008). Importin-beta and the small guanosine triphosphatase Ran mediate chromosome loading of the human chromokinesin Kid. The Journal of Cell Biology, 180, 493–506. Tokai, N., Fujimoto-Nishiyama, A., Toyoshima, Y., Yonemura, S., Tsukita, S., Inoue, J., et al. (1996). Kid, a novel kinesin-like DNA binding protein, is localized to chromosomes and the mitotic spindle. The EMBO Journal, 15, 457–467.

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Velazquez, J. M., & Lindquist, S. (1984). hsp70: Nuclear concentration during environmental stress and cytoplasmic storage during recovery. Cell, 36, 655–662. Welch, W. J., & Feramisco, J. R. (1984). Nuclear and nucleolar localization of the 72,000dalton heat shock protein in heat-shocked mammalian cells. The Journal of Biological Chemistry, 259, 4501–4513. Yokoya, F., Imamoto, N., Tachibana, T., & Yoneda, Y. (1999). beta-catenin can be transported into the nucleus in a Ran-unassisted manner. Molecular Biology of the Cell, 4, 1119–1131.

CHAPTER

Novel Approaches for the Identification of Nuclear Transport Receptor Substrates

16

Makoto Kimura*, Ketan Thakar{, Samir Karaca{, Naoko Imamoto*, and Ralph H. Kehlenbach{ *

Cellular Dynamics Laboratory, RIKEN, Wako, Saitama, Japan Department of Molecular Biology, Faculty of Medicine, Georg-August-University of Go¨ttingen, Go¨ttingen, Germany { Bioanalytical Mass Spectrometry Group, Max Planck Institute for Biophysical Chemistry, Go¨ttingen, Germany

{

CHAPTER OUTLINE Introduction ............................................................................................................ 354 Bioinformatic Approaches for the Identification of Transport Receptor Substrates .... 355 Mass-spectrometry-based Approaches for the Identification of Transport Receptor Substrates ............................................................................................... 356 16.1 Identification of CRM1-Dependent Export Cargos...............................................357 16.1.1 Equipment and Reagents ............................................................ 358 16.1.2 Protocol..................................................................................... 360 16.1.2.1 Adaptation of Cells From Normal DMEM to SILAC Medium ................................................................................. 360 16.1.2.2 Transfection ...................................................................... 360 16.1.2.3 Subcellular Fractionation.................................................... 361 16.1.2.4 Gel Electrophoresis and in-gel Digestion ............................. 362 16.1.2.5 LC–MS/MS ........................................................................ 362 16.1.2.6 Data Analysis..................................................................... 362 16.2 Identification of Nuclear Import Cargos ............................................................364 16.2.1 Equipment, Reagents, and Buffers............................................... 366 16.2.2 Protocol..................................................................................... 367 16.2.2.1 Preparation of Recombinant Proteins ................................. 367 16.2.2.2 Preparation of Importin-Depleted Cytosol............................ 368 16.2.2.3 Preparation of Depleted Nuclear Extract ............................. 369 16.2.2.4 SILAC................................................................................ 370 16.2.2.5 In Vitro Import Reaction ..................................................... 370

Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00016-3

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16.2.2.6 Sample Preparation for LC–MS/MS..................................... 372 16.2.2.7 Data Analysis..................................................................... 373 Conclusions............................................................................................................ 375 Acknowledgments ................................................................................................... 375 References ............................................................................................................. 376

Abstract Nucleocytoplasmic transport affects the subcellular localization of a large proportion of cellular proteins. Transported proteins interact with a set of
20 transport receptors, importins and exportins, which mediate translocation through the nuclear pore complex. Here we describe two novel methods based on quantitative proteome analysis for the identification of cargo proteins that are transported by a specific importin or exportin. The first approach is based on SILAC (stable isotope labeling of amino acids in cells) using cells that have been treated or not with specific reagents, followed by subcellular fractionation. Applying this approach to cells treated with or without the selective CRM1 inhibitor leptomycin B, we identified substrates of CRM1, the major nuclear export receptor. In the second SILAC approach, digitonin-permeabilized cells are incubated with nuclear and cytosolic extracts in the absence or presence of particular import receptors of interest. Proteomic analysis of the permeabilized cells then yields proteins whose nuclear import depends specifically on the added import receptor. Using this system, we identified substrates of two representative import receptors, transportin and importin-a/b.

INTRODUCTION It has been estimated that up to 35% of all cellular proteins enter the nucleus at some point during the cell cycle (Kumar et al., 2002). Many of these may be imported or exported either constantly or upon a certain stimulus. About 30 years ago, the first regions and sequences were identified that direct proteins for transport into the nucleus (NLSs, nuclear localization signals). More than a decade later, sequences (NESs, nuclear export sequences) that mediate export of proteins from the nucleus back into the cytoplasm were found. Today, we have a detailed knowledge about the transport machinery that controls the localization of nucleocytoplasmic shuttling proteins. A group of
20 proteins belonging to the superfamily of importin-b-related nuclear transport receptors (NTRs) interacts with transport cargos via characteristic NLSs (importins) or NESs (exportins). All members of this family bind RanGTP, a small G-protein that controls the loading of transport receptors with cargo molecules: after transport of an import complex into the nucleus, binding of RanGTP to the importin initiates the dissociation of the import complex. In nuclear export, by contrast, RanGTP is an integral component of a trimeric transport complex containing the exportin, the cargo and RanGTP. Dissociation of the export complex in the

Introduction

cytoplasm is then initiated by GTP hydrolysis on Ran, which requires the GTPaseactivating protein RanGAP as a cofactor. Resulting from the activities of p10/NTF2, a dedicated nuclear import factor for RanGDP and RCC1, a chromatin-bound nucleotide exchange factor for Ran, RanGTP itself is highly concentrated in the nucleus and is a defining factor for the directionality of nuclear transport. For general references on the players and the mechanisms of nuclear transport, see Fried and Kutay (2003), Wa¨lde and Kehlenbach (2010), and Wente and Rout (2010). Transport across the nuclear envelope occurs through the nuclear pore complexes, very large protein assemblies that contain
30 nucleoporins (Nups) occurring in copy numbers of eight or multiples of eight (Cronshaw, Krutchinsky, Zhang, Chait, & Matunis, 2002; Ori et al., 2013). A subset of these Nups interacts with transport receptors, mediating the translocation of transport complexes. In all models that try to explain the biophysical underpinnings of nuclear transport, these Nups play a very general role, as they affect all transport pathways as defined by the individual transport receptors (Wente & Rout, 2010). Furthermore, some Nups have been described to affect nucleocytoplasmic transport of individual substrates in a manner that does not depend on their specific receptor (Wa¨lde & Kehlenbach, 2010). After more than three decades of intensive research to elucidate the role of the nuclear pore complex, of soluble NTRs and of accessory proteins like Ran and RanGAP, it will be important (i) to identify the cellular substrates of the
20 NTRs, (ii) to characterize proteins whose subcellular localization is affected by specific Nups, and (iii) to describe the effects of certain stimuli (e.g., heat shock, nutrient deprivation, and viral infection) on the subcellular localization of the proteome. We now focus on novel approaches toward the first goal, some of which might be applicable for the other goals as well.

Bioinformatic approaches for the identification of transport receptor substrates The “classical” NLS (cNLS) with a dimer of importin-a and importin-b as transport receptor is characterized by either one (monopartite) or two (bipartite) stretches of amino acids that are enriched in basic residues (Lange, McLane, Mills, Devine, & Corbett, 2010). Examples are the monopartite NLS of the SV40 large T antigen and the bipartite NLS of nucleoplasmin (126PKKKRRV132) (155KRPAATKKAGQAKKKK170). In yeast, up to 45% of all proteins contain stretches corresponding to these consensus sequences (Lange et al., 2007) and several algorithms/programs are available to predict such NLSs in protein sequences. Importin-a is the adapter protein that recognizes the cNLS. Importin-b then binds to the IBB (importin-b-binding) domain of importin-a and mediates interaction of the import complex with the nuclear pore. Alternative NLSs were described for the importin-b-related transport receptor transportin. Again, two classes can be distinguished, namely, NLSs containing either a basic or a hydrophobic stretch of amino acids close to characteristic PY-residues and programs are available to predict such PY-NLSs (Lee et al., 2006). For nuclear export, only one class of NESs has been

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described so far that is recognized by the nuclear export receptor CRM1. This NES, which was originally termed “leucine-rich NES,” is characterized by a stretch of hydrophobic amino acids with variable spacing (for review, see Hutten & Kehlenbach, 2007). As for nuclear import, search algorithms can be used to predict CRM1dependent NESs (Fu, Imai, & Horton, 2011; la Cour et al., 2004). Several problems, however, arise with all prediction programs: first, they cannot take into account the three-dimensional structure of the protein. Hence, they may identify sequences that are buried in the structure of the protein and are not exposed for binding to a transport receptor. This is a particular issue for the hydrophobic NESs that mediate interaction with the export receptor CRM1. Furthermore, as the programs analyze linear sequences, they miss NLSs or NESs that only arise upon proper folding of the protein (Ayers, Nedrow, Gillilan, & Noy, 2007). Second, there are no proper consensus sequences known for the vast majority of NTRs. In addition, basic amino acid sequences resembling basic cNLSs could also interact with import receptors other than the importin-a/b dimer, but such receptors have different substrate specificities. For the majority of import cargos that interact with importin-a/b in vitro in a cNLSdependent manner, it has not unequivocally been demonstrated that importin-a/b is also the responsible import receptor in vivo. In the future, improved prediction programs might be available that take into account more validated NLSs and NESs and also the three-dimensional structure of potential cargo proteins.

Mass-spectrometry-based approaches for the identification of transport receptor substrates One way to identify novel transport cargos is to analyze proteins that interact with import or export receptors (Fukumoto, Sekimoto, & Yoneda, 2011; Guttinger, Muhlhausser, Koller-Eichhorn, Brennecke, & Kutay, 2004; Ja¨kel and Go¨rlich, 1998; Mingot, Bohnsack, Ja¨kle, & Go¨rlich, 2004; Miyamoto et al., 2013). As an alternative to biochemical-interaction-based approaches, a quantitative proteomic approach was recently applied to identify importin-a–substrates whose subcellular localization was affected by a specific gene knockdown (Wang et al., 2012). Quantitative proteomics has previously been used to analyze the spatial distribution between the nucleus and the cytoplasm (Boisvert et al., 2012). We now describe two proteomic approaches that are based on the differential subcellular localization of potential transport cargos under certain conditions. Both methods involve SILAC (stable isotope labeling of amino acids in cells), followed by quantitative mass spectrometry (Mann, 2006). For the comparison of the proteome of cells (or of subcellular fractions) under different conditions, cells are grown in medium containing either “light” (normal) or “heavy” amino acids. Heavy amino acids contain, for example, stable isotopes 2H instead of 1H, 13C instead of 12 C, or 15N instead of 14N. Complete labeling (>95% is recommended) with heavy or light amino acids is achieved after approximately five cell doublings. Essential amino acids like arginine and lysine are chosen for labeling. In a typical SILAC experiment, population A is labeled with light amino acids and population B is labeled with the

16.1 Identification of CRM1-Dependent Export Cargos

corresponding heavy amino acids. Incorporation of heavy amino acids into proteins causes a defined mass shift compared to proteins containing light amino acids, but no other chemical alterations. Hence, by all biological criteria, protein function should not be affected by the labeling. After appropriate treatment of cells as per the requirements of the experiment, the cells are mixed at a 1:1 ratio. Proteins are extracted, digested with trypsin or other proteases, and analyzed by mass spectrometry. In the mass spectra, each specific peptide appears as a pair with a defined difference in mass. The first peptide with a lower mass contains the light amino acids that originate from population A. The second peptide with a higher mass contains the heavy amino acids from population B. A 1:1 ratio of heavy and light peptides suggests an equal abundance of the corresponding protein in the two populations. An higher peak intensity for the heavy peptide indicates that the corresponding protein was more abundant in population B and vice versa. The ratio of peak intensities of multiple peptides derived from one protein directly yields the ratio of the proteins in the original samples. When whole cell extracts or subcellular fractions are analyzed, the complexity and the dynamic range of the samples are very high. Therefore, prefractionation, for example, by SDS-PAGE or isoelectric focusing should be performed. Using this mass-spectrometry-based approach, thousands of individual proteins can be analyzed in parallel in a quantitative manner. To increase the accuracy of the analysis, we recommend at least two biological and/or technical replicates (consider costs and instrumentation time). Furthermore, the labeling can be switched. Applying stringent criteria, cutoff ratios of 1.3–2.0-fold are usually considered significant (for review, see Mann, 2006). Two SILAC-based approaches are presented here: first, we compare the subcellular localization of proteins in cells that have been treated with or without the selective CRM1 inhibitor leptomycin B (LMB). The method involves a subcellular fractionation of treated and untreated cells, followed by proteome analysis of the fractions (Thakar, Karaca, Port, Urlaub, & Kehlenbach, 2013). The second approach takes advantage of an established nuclear import assay in digitonin-permeabilized cells (Adam, Marr, & Gerace, 1990; see also Chapter 15). Nuclei of permeabilized cells are incubated with nuclear and cytosolic extracts, which are depleted of endogenous NTRs, in the absence or presence of particular import receptors of interest. After the reaction, proteomic analysis of the permeabilized cells yields proteins whose nuclear import depends specifically on the added import receptor. Using this system, we systematically identified substrates of two representative import receptors, transportin (Kimura, Kose, et al., 2013) and importin-a/b (Kimura, Okumura, Kose, Takao, & Imamoto, 2013).

16.1 IDENTIFICATION OF CRM1-DEPENDENT EXPORT CARGOS This approach was initially published in Thakar et al. (2013). The fungal metabolite LMB from Streptomyces (Hamamoto, Gunji, Tsuji, & Beppu, 1983) is a selective inhibitor of the CRM1-mediated nuclear export pathway

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(Wolff, Sanglier, & Wang, 1997). LMB acts by covalently binding to cysteine 528 in the NES-binding region of human CRM1 (Kudo et al., 1999), thus preventing the formation of export complexes (for review, see Hutten & Kehlenbach, 2007). The workflow for analyzing the effect of LMB on the subcellular distribution of proteins is shown in Fig. 16.1. LMB leads to an accumulation of CRM1-substrates in the nucleus and/or their depletion from the cytoplasm. Changes in the abundance of substrates in subcellular fractions can be detected by SILAC, followed by quantitative mass spectrometry.

16.1.1 Equipment and reagents ANTIBODIES Mouse anti-HA antibody (16B12, Covance, Emeryville, CA, USA) dilution 1:1000 Rabbit anti-SP1 antibody (Thermo Scientific, Rockford, IL, USA) dilution 1:1000 Rabbit antitubulin (Proteintech, Chicago, IL, USA) dilution 1:1000 Rabbit anti-GAPDH antibodies (Proteintech, Chicago, IL, USA) dilution 1:1000 Mouse antilamin A/C (BD Biosciences, San Jose, CA, USA) dilution 1:1000 Benzonase: Sigma-Aldrich Chemie GmbH, Taufkirchen, Germany C18 capillary column, analytical: Dr. Maisch GmbH, Germany (15 cm, 360-m ˚ , 5 mm, C18-AQ) m outer diameter, 75-mm inner diameter; Reprosil-Pur 120 A C18 trap column: Dr. Maisch GmbH, Germany (1.5 cm, 360-mm outer diameter, ˚ , 5 mm, C18-AQ) 150-mm inner diameter; Reprosil-Pur 120 A Fetal calf serum, dialyzed: PAA Laboratories GmbH, Pasching, Austria Digitonin: Merck KGaA, Darmstadt, Germany, 1% stock solution in H2O, store at 20  C HeLa P4 cells (Charneau et al., 1994) grown at 37  C in a humidified incubator with a 5% CO2 atmosphere. These cells are readily transfectable using the calcium phosphate method (see below) Leptomycin B (LMB): Alexis Biochemicals, Switzerland. 10 mM stock solution in ethanol, store at 20  C LiChrosolv water: Merck KGaA, Darmstadt, Germany Plasmid: pRev(48–116)-GFP2-M9 (Thakar et al., 2013) Penicillin–streptomycin: PAA Laboratories GmbH, Pasching, Austria; at concentrations of 100 IU/ml and 100 mg/ml, respectively Protease inhibitors: complete EDTA free protease inhibitors cocktail tablets (Roche Diagnostics GmbH, Mannheim, Germany) SDS-PAGE: Life Technologies GmbH, Darmstadt, Germany, 4–12% NuPAGE Bis–Tris Gels SILAC DMEM media: Supplement Dulbecco’s Modified Eagle Medium (DMEM) containing high glucose (4.5 g/l) deficient in arginine and lysine (PAA Laboratories GmbH, Pasching, Austria) with 10% dialyzed (to minimize the

FIGURE 16.1 Experimental Workflow for SILAC Identification of CRM1-Dependent Export Cargos HeLa cells are grown in DMEM containing either light or heavy amino acids. Equal number of cells are mixed prior to subcellular fractionation. Soluble proteins from cytosolic and nuclear fractions are separated by SDS-PAGE. After in-gel digestion, extracted peptides are analyzed by LC–MS/MS and raw data are processed with MaxQuant. A similar figure was originally published by Thakar et al. (2013).

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concentration of amino acids) fetal calf serum (FCS), penicillin–streptomycin (100 IU/ml and 100 mg/ml, respectively), and either unlabeled L-arginineHCl and L-lysineHCl (SILAC: R0K0, “light”; Cambridge Isotope Laboratories, Andover, MA, USA) or L-arginine-U-13C6HCl and L-lysine-U-13C6-15N22HCl (SILAC: R6K8, “heavy”; nomenclature derives from the mass shifts of 6- and 8-Da, respectively, for “heavy” amino acids compared to light amino acids) at concentrations of 50 mg/ml each FOR MASS-SPECTROMETRY ANALYSES Agilent 1100 Nanoflow LC System (Agilent Technologies) LTQ-Orbitrap Xl or LTQ Orbitrap Velos hybrid mass spectrometer (Thermo Electron, Bremen, Germany) MaxQuant software

16.1.2 Protocol 16.1.2.1 Adaptation of cells from normal DMEM to SILAC medium 1. Culture HeLa P4 cells (Charneau et al., 1994) in normal DMEM to 80–90% confluency and further divide into two culture dishes containing light or heavy SILAC medium. 2. Change medium every 2–3 days. Subculture cells at least twice in SILAC medium and allow at least five cell doublings. Remove a small number of cells (
0.5–1  106 cells) to control the incorporation rate of light or heavy amino acids at different time points. Here, a small-scale MS analysis is sufficient. Almost complete incorporation (99%) of heavy amino acids can be achieved after five passages of the cells. Mass spectrometric quantification accuracy can be determined by checking logarithmic fold change of mixed untreated heavy and light cells. Samples can be used for further analysis if the level of quantified proteins showing a fold change close to zero is higher than 95%.

16.1.2.2 Transfection Transfect
2  106 cells with a plasmid coding for a control protein that is expected to change its subcellular localization upon the addition of LMB (here: Rev(48–116)GFP2-M9) using the Ca-phosphate method. This transfection method yields high transfection efficiencies in HeLa P4 cells. It avoids use of expensive kits and all required solutions can be prepared in the lab. 1. Briefly, transfection is performed by mixing 1.5 mg/ml of plasmid with 500 ml of sterile 250 mM CaCl2 solution in a 1.5-ml eppendorf cup. Vortex the tube thoroughly. Add 500 ml of sterile 2 HBS buffer solution (50 mM HEPES, 250 mM NaCl, 1.5 mM Na2HPO4, pH 6.98) and vortex thoroughly. Incubate mixture at room temperature for 20 min and then add drop by drop to cells in SILAC media. Gently rock the plate for homogenous distribution of transfection mix.

16.1 Identification of CRM1-Dependent Export Cargos

2. After 24 h, add 10 nM LMB and further incubate for 3 h at 37  C to block CRM1dependent nuclear export. 3. To generate results with a high confidence interval, perform two biological replicates and analyze each biological replicate twice. Additionally, to avoid false positives perform label-swap experiments. Proteins with conflicting results are eventually removed from the list.

16.1.2.3 Subcellular fractionation 1. Trypsinize and mix 4  106 untreated “light” cells with an equal number of LMB-treated “heavy” cells (“forward experiment”) or LMB-treated “light” cells with untreated “heavy” cells (“reverse experiment”) in a ratio of 1:1. 2. Wash cells in cold medium and centrifuge at 4  C for 5 min at 100  g. Resuspend pellet in PBS and centrifuge again at 4  C for 5 min at 100  g. Remove 10% of the cells and boil directly in SDS sample buffer, centrifuge at 14,000  g for 15 min and collect supernatant: this supernatant is referred to as total lysate (T) and can be analyzed to detect global changes in protein composition. 3. Subcellular fractionation: resuspend remaining cells in 400 ml of ice-cold buffer 1 (150 mM NaCl, 50 mM HEPES pH 7.4, 0.02% digitonin and protease inhibitors), incubate at 4  C for 10 min, and then centrifuge at 2000  g for 5 min. Digitonin is used to permeabilized the plasma membrane, leaving the nuclear envelope intact. The supernatant is referred to as a “cytosol-enriched fraction” (C). 4. Resuspend pellet in 400 ml of ice-cold buffer 2 (150 mM NaCl, 50 mM HEPES pH 7.4, 1% NP-40 and protease inhibitors), incubate for 30 min on ice, and centrifuge at 7000  g for 10 min. The supernatant mainly contains membranebound organelles such as endoplasmic reticulum, Golgi, mitochondrial, and some nuclear luminal proteins and was not used in our analysis. 5. Resuspend pellet in 400 ml of ice-cold buffer 3 (150 mM NaCl, 50 mM HEPES pH 7.4, 0.5% sodium deoxycholate, 0.1% SDS, benzonase (1 U/ml), and protease inhibitors), incubate at 4  C for 1 h, and centrifuge at 7000  g for 10 min. The supernatant comprises the extracted nuclear membranes and soluble nuclear proteins (N). 6. The quality of the fractionation can be tested by western blot by probing for marker proteins like a-tubulin and GAPDH which are present predominantly in the cytosolic fraction and absent from the nuclear fraction. As nuclear marker proteins, lamin A/C and the transcription factor SP1 can be used. Note: This protocol is optimized for HeLa P4 cells. For subcellular fractionation of other cell types, certain adjustments, like incorporation rate of labeled amino acids, optimizing transfection efficiency, testing quality of subcellular fractions, may be required. For alternative nuclear purification/subcellular fractionation procedures, see also Chapter 6 (swelling of cells in a hypotonic buffer followed by mechanical rupture) or Chapter 19 (using commercially available isolation kit).

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16.1.2.4 Gel electrophoresis and in-gel digestion 1. Load equal protein amounts of nuclear and cytosolic extracts (
50–75 mg) and separate by SDS-PAGE, followed by Coomassie Brilliant Blue staining. Leave empty lanes between each sample. 2. Wash the gel several times with LiChrosolv water. The most common contaminant in MS analysis is keratin, so it is important to work under laminar flow and to wear gloves. 3. After scanning the gel, cut each lane into the desired number of slices (e.g.,
20; depending on the required proteome coverage). 4. Chop slices into small pieces (
1 mm2) and collect them in 0.5 ml tubes. Rinse scalpel after each slice. 5. Dehydrate gel pieces by incubation in acetonitrile for 10 min. 6. To reduce disulfide bonds, incubate gel pieces first in 100 ml dithiothreitol (DTT) (10 mM) at 56  C for 50 min and then in 100 ml iodoacetamide (55 mM) for 20 min at room temperature in the dark to prevent reformation of disulfide bonds. 7. Subject proteins to in-gel digestion with trypsin (Roche) overnight using a trypsin:protein ratio of
1:50 and assuming similar protein amounts per gel piece (i.e.,
50–75 ng trypsin per gel piece). 8. To extract peptides from the gel pieces, cover them with acetonitrile. Shake for 10 min at room temperature. Spin down the gel pieces and transfer the liquid to a fresh tube. Incubate gel pieces with 50 ml 1% formic acid for 20 min, add 50 ml acetonitrile, and incubate for 10 min. Spin down gel pieces, recover supernatants, and combine them with those from the previous step. 9. Dry samples in speed-vac and store dried samples at 20  C.

16.1.2.5 LC–MS/MS 1. Dissolve peptides in 1% formic acid and load extracted peptides on a C18 trap column (1.5 cm) at a flow rate of 10 ml/min. 2. Elute and separate retained peptides on an analytical C18 capillary column (15 cm) at a flow rate of 300 nl/min with a gradient from 5% to 38% acetonitrile in 0.1% formic acid for 50 min using an Agilent 1100 Nanoflow LC System (Agilent Technologies) coupled to an LTQ-Orbitrap Velos hybrid mass spectrometer (Thermo Electron, Bremen, Germany) operated in datadependent mode. 3. Acquire survey scans in the Orbitrap (m/z 350–1600) with a resolution of 30,000 at m/z 400 with a target value of 1  106. 4. Sequentially isolate up to 15 most intense ions with charge þ2 for collisioninduced dissociation with normalized collision energy of 35. Set dynamic exclusion to 60 s in order to avoid repeating sequencing of peptides.

16.1.2.6 Data analysis 1. Analyze raw MS files from LTQ-Orbitrap Velos by MaxQuant software (version 1.0.13.13) using Mascot search engine or later versions with built in Andromeda search engine. Use the human UniProt FASTA database to match peak lists.

16.1 Identification of CRM1-Dependent Export Cargos

2. Use the following search parameters: set carbamidomethylation of cysteine as a fixed modification, oxidation of methionine, and N-terminal protein acetylation as variable modifications; tryptic specificity with no proline restriction and use up to two missed cleavages. 3. Set MS survey scan mass tolerance to 7 ppm and for MS/MS 0.5 Da. Set false discovery rate to 1%, both at the peptide and the protein level. Consider peptides with a posterior error probability of less than 0.05 for identification and quantification. Enable “Re-quantification” and disable “keep low scoring versions of identified peptides.” 4. Perform quantification of SILAC pairs with a minimum ratio count of two by considering unique and razor peptides. 5. Set significance B as a main criterion for data analysis. This avoids setting any empirical cutoff values, but shows a better statistical significance of outlier proteins, as these are identified from the bulk of the distribution by calculating the variances of all proteins. Furthermore, it takes into account the fact that highly abundant proteins are more accurately quantified than rare ones. 6. Proteins with p-value  0.01 are considered significant. The data can be represented as shown in Fig. 16.2. A total of
3300 proteins (cytosolic fraction, Fig. 16.2A) or
3200 proteins (nuclear fraction, Fig. 16.2B) was identified by LC–MS/MS after SILAC labeling, with an overlap of
1900 proteins (Thakar et al., 2013). Applying a high level of stringency, we detected changes for 138 proteins that upon LMB treatment were either depleted from the cytosolic fraction (84 proteins) or accumulated in the nuclear fraction (59 proteins) or both (5 proteins). Among the identified proteins were our positive A

B

Cytosol Depleted

Enriched

Depleted

Enriched

9

Log10(intensity)

9

Log10(intensity)

Nucleus 10

10

RanBP1

8 7

Rev-GFP2-M9

6

8 7 NMD3

6 5

5

4

4 -2

-1

0

-1

Fold change (log2ratio)

-2

-2

-1

0

-1

-2

Fold change (log2ratio)

FIGURE 16.2 Scatter Plots of Quantified Proteins in Cytosolic (A) or Nuclear Fractions (B) After LMB-treatment of HeLa Cells Proteins are colored according to significance B, where gray triangles signify p-values >0.01, blue 300 evolutionarily conserved nonribosomal trans-acting factors, which transiently associate with preribosomal subunits at distinct assembly stages. A subset of trans-acting and transport factors passage assembled preribosomal subunits in a functionally inactive state through the nuclear pore complexes (NPC) into the cytoplasm, where they undergo final maturation before initiating translation. Here, we summarize the repertoire of tools developed in the model organism budding yeast that are spearheading the functional analyses of trans-acting factors involved in the assembly and intracellular transport of preribosomal subunits. We elaborate on different GFP-tagged ribosomal protein reporters and a pre-rRNA reporter that reliably monitors the movement of preribosomal particles from the nucleolus to cytoplasm. We discuss the powerful yeast heterokaryon assay, which can be employed to uncover shuttling trans-acting factors that need to accompany preribosomal subunits to the cytoplasm to be released prior to initiating translation. Moreover, we present two biochemical approaches, namely sucrose gradient analyses and tandem affinity purification, that are rapidly facilitating the uncovering of regulatory processes that control the compositional dynamics of trans-acting factors on maturing preribosomal particles. Altogether, these approaches when combined with traditional analytical biochemistry, targeted proteomics and structural methodologies, will contribute to the dissection of the assembly and intracellular transport of preribosomal subunits, as well as other macromolecular assemblies that influence diverse biological pathways.

INTRODUCTION To construct eukaryotic ribosomal subunits (40S and 60S), a cell must assemble >70 ribosomal proteins (r-proteins) with four different rRNA species. A concerted effort of all three transcriptional machineries (RNA Polymerases (Pol) I, II, and III) is required in order to ensure the high efficiency and accuracy of ribosome production. Pol I produces 35S rRNA that is cleaved and processed into

Introduction

18S, 5.8S, and 25S rRNAs, Pol III synthesizes 5S rRNA, whereas Pol II transcribes the mRNAs encoding the ribosomal proteins. Eukaryotic ribosome assembly is aided by >300 nonribosomal trans-acting factors that transiently interact with evolving precursor particles to perform specific maturation steps (Kressler, Hurt, & Baßler, 2010; Strunk & Karbstein, 2009). While the structure and function of the mature ribosome is better characterized at the molecular level, our knowledge regarding the ribosome assembly pathway is only emerging. Pioneering work performed in the early 1970s by the Planta and Warner laboratories identified the earliest preribosomal particle termed 90S, which is subsequently processed to the precursors of the mature 60S and 40S subunits, respectively (Kruiswijk & Planta, 1974; Trapman, Rete`l, & Planta, 1975; Udem & Warner, 1973). Today, we know that these particles contain the pre-rRNAs, ribosomal proteins, and trans-acting factors that are removed as the preribosomes migrate from the nucleolus to the cytoplasm. Early biochemical work indicated that the process of ribosome synthesis in budding yeast is similar to that in higher eukaryotes. This validated the use of yeast as a model system to which powerful genetic, cell biological, and biochemical tools could be applied. In the late 1990s, visual approaches using large and small subunit reporters (Rpl25–GFP, Rpl11–GFP, and Rps2–GFP) identified factors involved in the intranuclear transport and nuclear export of preribosomal subunits (Hurt et al., 1999; Milkereit et al., 2003; Stage-Zimmermann, Schmidt, & Silver, 2000). Despite these advances, the composition of preribosomal particles remained largely unknown until the advent of tandem affinity purification (TAP) protocols in budding yeast and sensitive mass spectrometry (Puig et al., 2001; Rigaut et al., 1999). These technologies allowed the isolation and compositional analysis of maturing pre-60S and pre-40S particles, and have advanced the field by expanding the inventory of factors that are involved in ribosome biogenesis/export (Grandi et al., 2002; Nissan, Baßler, Petfalski, Tollervey, & Hurt, 2002; Scha¨fer, Strauß, Petfalski, Tollervey, & Hurt, 2003). These analyses have provided “biochemical snapshots” of the highly dynamic assembly process and thus aided the sequential ordering of preribosomal particles along the 60S and 40S pathways. The precursor 35S pre-rRNA is generated by the RNA Pol I driven transcription of rDNA repeats in the nucleolus. The emerging pre-rRNA is cotranscriptionally modified and loaded with a subset of small subunit r-proteins and trans-acting factors to form the earliest precursor of the 40S subunit: the 90S particle (Fig. 20.1). Cleavage of the 35S pre-rRNA releases the pre-40S particle and permits the remaining pre-rRNA to assemble with large subunit r-proteins and biogenesis factors to form pre-60S particles. After separation of the 90S particle into pre-40S and pre-60S subunits, the precursors follow independent assembly pathways. Pre-40S particles undergo few compositional changes as they travel through the nucleoplasm and are rapidly exported to the cytoplasm. In contrast, pre-60S particles associate with
200 trans-acting factors along their assembly pathway and therefore undergo dynamic compositional changes as they travel towards the NPC. At distinct stages of biogenesis, trans-acting factors are released from preribosomal particles and

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FIGURE 20.1 Current model for assembly and transport of 40S and 60S ribosomal subunits. The earliest ribosomal precursor, the 90S particle is composed of 35S pre-rRNA, the large U3 snoRNP-containing processome, r-proteins, and assembly factors of the 40S subunit. Following cleavage at site A2, the 90S precursor separates into 40S and 60S pre-ribosomes. The processome is released from the pre-40S particles, and a few assembly factors join the pre-40S subunits before nuclear exit. The majority of pre-60S assembly factors join after cleavage of pre-rRNA at A2 site. Many pre-60S assembly factors are released in subsequent steps, whereas a few proteins associate at later stages and passage the pre-60S subunit through the nuclear pore. Final maturation of preribosomes before initiating translation occurs in the cytoplasm. TAP purifications of the depicted assembly factors purify preribosomal particles at that maturation stage.

recycled back to participate in new rounds of maturation. These events are triggered by diverse energy-consuming enzymes (ATP-dependent RNA helicases, AAAATPases, ABC-ATPases, GTPases), which associate with maturing preribosomal particles at distinct maturation stages. The stripping and remodeling action of diverse energy-consuming enzymes is thought to sequentially reduce the complexity of preribosomal particles leading to acquisition of export competence (Tschochner & Hurt, 2003). Export competent preribosomal particles are transported from the nucleus to the cytoplasm separately upon interaction with the general nuclear export factor Xpo1 that directly recognizes a nuclear export sequence. Nmd3 is the only known adaptor between the pre-60S particle and Xpo1 (Gadal et al., 2001; Ho, Kallstrom, & Johnson, 2000). An essential adaptor protein for Xpo1 on the surface of the pre40S particle has not been yet identified. Additionally, a pre-60S particle employs

20.1 Localization of Preribosomal Subunits by Fluorescence Microscopy

multiple trans-acting factors that shield the highly negative charge of the rRNA and position the subunit for entry into the disordered FG-repeats of the NPC transport channel (Altvater et al., 2012; Baßler et al., 2012; Bradatsch et al., 2007; Hackmann, Gross, Baierlein, & Krebber, 2011; Hung, Lo, Patel, Helmke, & Johnson, 2008; Oeffinger, Dlakic´, & Tollervey, 2004; Yao et al., 2010). The majority of trans-acting factors that associate with preribosomal particles during early biogenesis are released and recycled back to the nucleolus prior to nuclear export. However, a few factors remain associated with preribosomal particles and facilitate their nuclear export into the cytoplasm. The release of shuttling transacting factors from preribosomal particles constitutes “cytoplasmic maturation steps” in the ribosome biogenesis pathway (Panse, 2011; Panse & Johnson, 2010). These final steps are crucial not only for completing ribosomal subunit maturation, but also because a failure to recycle a factor back to the nucleus leads to its depletion from its nucleolar/nuclear sites of action, inducing pre-rRNA processing delays, assembly defects, and impaired nuclear export. While large-scale proteomic approaches have greatly advanced our understanding of ribosome assembly and provided a general framework to investigate this conserved process, the precise functions of the >300 trans-acting factors remain largely unclear. Here, we focus on the biochemical and cell biological tools developed in the model organism yeast that are facilitating the functional analysis of the uncharacterized trans-acting factors.

20.1 LOCALIZATION OF PRERIBOSOMAL SUBUNITS BY FLUORESCENCE MICROSCOPY The transport of preribosomal particles from the nucleus to the cytoplasm can be monitored by different large (L25-GFP, L11-GFP, and L5-GFP) and small (S2GFP) subunit reporter constructs (Altvater et al., 2012; Hurt et al., 1999; Milkereit et al., 2003; Stage-Zimmermann et al., 2000). An important premise for these visual in vivo assays is the stable incorporation of the reporter construct into the preribosomal subunits during early assembly steps, and that the assembled preribosomal particles are rapidly exported to the cytoplasm. Thus the steady-state localization of the reporters in wild-type (WT) cells is cytoplasmic, with nuclear exclusion (Fig. 20.2, left). Impairment in the assembly and/or export of preribosomal particles to the cytoplasm should therefore lead to the nuclear accumulation of the reporters as seen for bud20△ (L5-GFP) and yrb2△ mutants (S2-GFP) (Fig. 20.2, left). PROTOCOL 1. Yeast strains are transformed with the reporter plasmids. Cells expressing reporters are grown in appropriate liquid media (for mutants at permissive temperature) until saturation.

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FIGURE 20.2 (Left) Localization of preribosomal subunits by fluorescence microscopy. Localization of L5-GFP or S2-GFP reporters in the indicated strains was monitored by fluorescence microscopy. Wild-type (WT) cells show cytoplasmic L5-GFP and S2-GFP localization whereas bud20△ and yrb2△ mutant cells display L5-GFP or S2-GFP mislocalization to the nucleus, respectively. Scale bar ¼ 5 mm. (Right) Polysome analyses after subjecting cell lysates to sucrose density gradient centrifugation. The strains were grown to mid-log phase and treated with 100 mg/ml cycloheximide to preserve polysomes. Four O.D.260 units of the clarified lysates were fractionated by 7–50% sucrose density ultracentrifugation and analyzed at 254 nm using a density gradient fractionator. The peaks for 40S and 60S subunits, 80S ribosomes, polysomes, and halfmers (polysomes containing an additional 40S subunit, asterisks) are indicated. In comparison to WT, the bud20△ mutant shows an increased 40S/ 60S subunit ratio, a decreased number in 80S ribosomes and appearance of halfmers (asterisks). The yrb2△ mutant that is impaired in the nuclear export of pre-40S subunits shows a decreased ratio of 40S/60S subunits.

2. Cells are diluted into 10 ml fresh media and grown until mid-log phase for at least 2–3 divisions. (In the case of cold or temperature sensitive mutants, the cultures are shifted to the appropriate restrictive temperature after 2–3 divisions of growth at permissive temperature.) 3. Cells are pelleted by centrifugation in a 15-ml Falcon tube (2300 rpm, 3 min) and washed once with 10 ml dH2O.

20.2 Fractionation of Cell Extracts by Sucrose Gradient Sedimentation

4. Supernatant is poured off and cell pellet is resuspended in remaining liquid. 5. 2 ml of cell suspension are placed on a slide, a cover slip is pressed over before inspection under a fluorescence microscope. REMARKS 1. The assay can be performed with strains containing the reporters growing on solid selective media. In this case, strains typically in stationary phase are spread on fresh YPD plates to induce ribosome production. After growing the cells for 4–8 h at the appropriate temperature, cells can be scraped from plate using an inoculation loop, mixed with 2 ml H2O placed on a glass slide and analyzed under a fluorescence microscope. 2. The L25-GFP and S2-GFP reporter plasmids with different selection markers (LEU2, URA3, TRP1, and HIS3) originate from the Hurt laboratory (Hurt et al., 1999; Milkereit et al., 2003), the L11-GFP reporter (LEU2) originates from the Silver laboratory (Stage-Zimmermann et al., 2000), and the L5-GFP reporter plasmids (LEU2, URA3, TRP1, and HIS3) can be requested from the Panse laboratory (Altvater et al., 2012). All the above reporters are expressed under the endogenous promoters. In principle, the ribosomal protein L5 can be C-terminally tagged with GFP at the genomic locus for monitoring the nuclear export of the large preribosomal subunit.

20.2 FRACTIONATION OF CELL EXTRACTS BY SUCROSE GRADIENT SEDIMENTATION Polysome profiles obtained by ultracentrifugation of cell extracts on sucrose gradients are a powerful approach to analyze deficits in free ribosomal subunits due to impairment in assembly and/or transport (Fig. 20.2, right; Warner & Knopf, 2002). Furthermore, association of trans-acting factors with preribosomal particles and the mechanisms that drive their dissociation after fulfilling their task can be reliably investigated by combination of sucrose gradient sedimentation, fractionation, and Western analysis.

20.2.1 Preparation of 7–50% sucrose gradients 20.2.1.1 Buffers and solutions Sucrose solutions Tris–HCl pH 7.5 NH4Cl MgCl2 Dithiothreitol (DTT) Sucrose

50 mM 50 mM 12 mM 1 mM (add before use) 7% or 50% (w/v)

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20.2.1.2 Setup for preparing gradients Sucrose density gradients are poured as well as fractionated using an ISCO Teledyne system equipped with a Model 160 TrisTM pump and a Foxy Jr. Fraction Collector. Outlined below is a step-by-step protocol and program used to pour and collect 7–50% sucrose gradients. – Preparing the TrisTM pump: 1. Turn pump speed turning wheel to 100%. 2. Switch pump direction to counter clock-wise operation. 3. Move locking levers of the tubing to vertical position. – Preparing Gradient mixer: 4. Put tubing inlets into the corresponding buffers (buffer A: low sucrose, buffer B: high sucrose). – Prepare Foxy Jr. for sucrose gradient collection: 5. Press “Standby/Operate.” 6. Check whether the correct rack is inserted. 7. Put waste bottle to drain outlet.

20.2.1.3 Priming – Fill 8. 9. 10. 11. 12. 13. 14. 15. 16. – Fill 17. 18. 19. 20. 21. 22.

tubing with buffer A to the first valve: Press “ADVANCE” (leads to a new menu). Choose “Acc,” then “Spd”: set pump speed to 100% (confirm with “Enter”). Return one step with “Back.” Choose “Grad”: set gradient to 0% B (confirm with “Enter”). Return two steps by pressing “Back” twice. Press “Pump”-this starts pumping the buffer into the system. When the solution reaches the first valve, stop the pump (use the direction switch on the TrisTM pump, switch it to “STOP”). Turn off pump in the Foxy Jr.: press “ADVANCE,” then “Pump.” Put pump direction switch back to counter clock-wise operation. tubing with buffer B to mixing chamber: Press “ADVANCE,” choose “Acc,” then “Grad”: set gradient to 100% B (confirm with “Enter”). Return two steps by pressing “Back” twice. Press “Pump”-this starts pumping the buffer into the system. When the solution reaches the mixing chamber, stop the pump (use the direction switch on the TrisTM pump, switch it to “STOP”). Turn off pump in the Foxy Jr.: press “ADVANCE,” then “Pump.” Put pump direction switch back to counter clock-wise operation.

20.2.1.4 Programming for preparing the gradient 23. Choose program number 24. Press “Edit” on the control panel

20.2 Fractionation of Cell Extracts by Sucrose Gradient Sedimentation

25. 26. 27. 28. 29. 30.

31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50.

Type ¼ Time windows Rack type ¼ 25/28 mm vials (make sure the correct rack is installed) Last tube ¼ x (enter number of tubes you want to prepare) Fraction by Time Fraction time ¼ 12:00 (fraction time has to be higher than the time needed to pour a gradient þ delay time) Flow delay ¼ 1:00 (delay time corresponds to the time needed for the solution to flow from the mixing chamber to the drop former; 1:00 for our system if pump speed is 40% in the program) Window 1 start ¼ 0:00 Window 1 end ¼ 4:05 (time needed to fill a SW41 Polyallomer tube) Window 2 start ¼ 0:00 ¼ Off (no second time window is used) Nonpeak/window ¼ Drain (for times when no window is active, the solution goes down the drain) Restart ¼ Time, Next Tube Restart time ¼ 5:07 (take time needed for system to fill tube þ delay time þ 2 s ! 4:05 þ 1:00 þ 0.02 ¼ 5.07) Event: 1 ¼ Pump Speed change 1: Speed % ¼ 40, 0:00 (at time ¼ 0:00, pump speed changes to 40%) Event: 2 ¼ Gradient change 2: Grad. %B ¼ 100, 0:00 (buffer composition at 0:00 is 100% B) Event: 3 ¼ Gradient change 3: Grad. %B ¼ 0, 4:05 (at 4:05, 0% B is pumped on the gradient) Event: 4 ¼ Gradient change 4: Grad. %B ¼0, 4:25 (for further 20 s, 0% B is pumped) Event: 5 ¼ Gradient change 5: Grad. %B ¼100, 4:25 (still at 4:25, switch to 100% B occurs) Event: 6 ¼ Pump Speed change 6: Speed % ¼ 0, 5:06 (pump is turned off after time needed for system to fill tube þ delay time þ 1 s ! 4:05 þ 1:00 þ 0.01 ¼ 5.06) Event: 7 ¼ None The program is saved and the display automatically returns to the starting window.

20.2.1.5 Running the program for pouring sucrose gradients 51. 52. 53. 54. 55.

Add empty 13.2 ml SW41 Polyallomer tubes to the rack. Start program: press “Run.” Tubes will now be filled according to the program. At the end, press “End” to return to the starting window. Remove tubes from rack, store at 20  C or load samples and perform ultracentrifugation (see below).

20.2.1.6 Cleaning 56. Thoroughly wash the setup with warm ddH2O.

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20.2.2 Preparation of cell extracts under polysome preserving conditions 20.2.2.1 Lysis Buffers Lysis buffer Tris–HCl pH 7.5 NaCl MgCl2 Cycloheximide

10 mM 100 mM 30 mM 100 mg/ml (add before use)

20.2.2.2 Protocol 1. Grow cells in 200 ml of appropriate medium to O.D.600 ¼ 0.8. 2. To prepare extracts under polysome preserving conditions, add cycloheximide to a final concentration of 100 mg/ml to the cell culture, mix well and leave on ice for 5 min. 3. Cells are harvested in a 50-ml Falcon tube by centrifugation (4000 rpm for 5 min at 4  C) and washed with 20 ml lysis buffer. 4. Cells are resuspended in 1 ml lysis buffer and transferred to a 2-ml Eppendorf tube. 5. Cells are quickly pelleted (14,000 rpm for 5 sec at 4  C) and the supernatant is discarded. 6. Cells are resuspended in 1 ml lysis buffer with 0.2 g glass beads (400–600 mm) and lysed by vortexing (Disruptor Genie, Scientific Industries) at maximum speed for 6 min at 4  C. (Critical step!) 7. To recover the lysate, punch a hole through the bottom of the tube with a needle (Microlance 3, 0,9 mm  40 mm). Collect the lysate in a new 1.5 ml Eppendorf tube by pushing the lysate through the hole using a 2.5-ml syringe plug (Terumo). The glass beads should remain in the old tube. Wash glass beads once with 200 ml lysis buffer. 8. The collected lysate is further clarified by centrifugation (14,000 rpm for 10 min at 4  C). 9. Transfer supernatant to a 1.5-ml Eppendorf tube with a pipette avoiding the lipid layer on the surface. 10. Measure the RNA concentration at 260 nm. 11. Add glycerol to a final concentration of 5% and make aliquots of 4 units of A260. (At least 10 aliquots can be made out of a 200-ml starting culture.) 12. Snap freeze the aliquots in liquid N2 and store at 80  C.

20.2.3 Sucrose density gradient sedimentation 1. Sucrose gradients are made in 13.2 ml SW41 Polyallomer tubes containing a 7–50% (w/v) sucrose gradient (see above).

20.2 Fractionation of Cell Extracts by Sucrose Gradient Sedimentation

2. An aliquot of the cell lysate (4 units of A260) is thawed on ice and carefully layered on top of the sucrose density gradient using a pipette. 3. For a typical polysome profile, the sucrose gradient is subjected to ultracentrifugation either at 39,000 rpm for 3 h, or 27,000 rpm for 15 h at 4  C (low deceleration) using a Beckman SW41 swing bucket rotor. 4. Polysome profiles are recorded at 254 nm and gradients are fractionated (12 drops (ca. 500 ml) per fraction resulting in about 15 fractions in total) using a TELEDYNE Isco density gradient recording system (see below).

20.2.4 Sucrose gradient fractionation 20.2.4.1 Buffers and solutions Chase solution: Tris–HCl pH 7.5 NH4Cl MgCl2 Dithiothreitol (DTT) Sucrose

50 mM 50 mM 12 mM 1 mM (add before use) 60% (w/v)

20.2.4.2 Setup preparation – Prepare UV detection: 1. Turn on UV detection system (>15 min prior to use, red light switches to green if UV lamp is ready). 2. Adjust sensitivity (e.g., 0.2–0.5). – Prepare TrisTM pump: 3. Turn pump speed turning wheel to 100%. 4. Switch pump direction to counter clock-wise operation. 5. Move locking levers of the tubing to vertical position. 6. Put tubing inlet into 60% sucrose solution (chase solution). – Prepare Foxy Jr.: 7. Press “standby/operate.” 8. Check whether the correct rack is inserted. 9. Put waste bottle to drain outlet. – Prepare Tube piercer: 10. Assemble collar, ring, and retaining nut (choose correct size for collar/ring, depending on tube size you are using). 11. Assemble piercer: connect bottom cap to tubing, add center column, spring, and septum; lower needle below septum level. 12. Connect outlet tubing to drop former on Foxy Jr.: Press “ADVANCE,” choose “Tube” (Foxy Jr. arm will move to tube position). 13. Remove drop former from arm.

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14. Switch tubing to connect tube piercer outlet. 15. Put drop former back to arm. 16. Press “ADVANCE,” choose “Park” (arm will move back to drain).

20.2.4.3 Priming – Pump 60% sucrose into needle: 17. Press “ADVANCE,” then “Pump.” 18. Pump speed turning wheel can be used to adjust pumping speed. 19. When the solution reaches the needle, stop the pump (use the direction switch on the TrisTM pump, switch it to “STOP”). – Pump off water/buffer from a tube to rinse system and adjust baseline: 20. Fill tube with ddH2O (or low sucrose buffer). 21. Tighten nut retainer on Tube piercer. 22. Insert full tube and tighten once more. 23. Raise the piercing system to bottom of the tube (septum should make tight contact). 24. Turn bottom cap to raise needle: pierce tube and make sure the needle is inserted far enough into the tube. 25. Turn pump speed turning wheel to 55%. 26. Turn pump back on with switch ! 60% sucrose is pumped into the tube, water/buffer is raised and pumped out through the UV cell to Foxy Jr. fraction collector (pump speed can be increased). 27. When water/buffer is in the UV cell, adjust baseline with the recorder offset. 28. Stop pumping once the tubing is rinsed and the baseline is adjusted. – Pump water back in the reverse direction to save some of the 60% sucrose solution and to rinse the UV cell: 29. Press “ADVANCE,” then “Tube.” 30. Remove drop former from Foxy Jr. arm and place in ddH2O. 31. Press “ADVANCE,” then “Park.” 32. Switch pump direction to clock-wise operation. 33. Press “ADVANCE,” then “Pump.” 34. Pump until upper level of 60% sucrose in the tube approaches needle level, then stop pump (switch to “STOP”). 35. Press “ADVANCE,” then “Tube.” 36. Dry drop former and put back into Foxy Jr. arm. 37. Press “ADVANCE,” then “Park.”

20.2.4.4 Programming 38. Choose program number. 39. Press “Edit.”

20.2 Fractionation of Cell Extracts by Sucrose Gradient Sedimentation

40. 41. 42. 43. 44. 45.

Type ¼ Simple Rack type ¼ Micro tubes (to fractionate into Eppendorf tubes) Last tube ¼ x (enter number of fractions to collect) Fraction by Drops Fraction drops ¼ 12 (12 drops correspond to ca. 500 ml) The display automatically returns to the starting window.

20.2.4.5 Fractionation 46. Add empty Eppendorf tubes to the rack. – Remove tube in the piercer: 47. Loosen retaining nut. 48. Lower piercer. 49. Remove tube. 50. Do not dry needle and septum. 51. Lower needle to a level just below the septum (ready for the next tube). – Insert tube to fractionate: 52. Tighten nut retainer on Tube piercer. 53. Insert tube and tighten once more. 54. Raise the piercing system to bottom of the tube (septum should make tight contact). 55. Turn bottom cap to raise needle: pierce tube and make sure the needle is inserted far enough into the tube. – Start fractionation: 56. Turn pump speed turning wheel to 55%. 57. Turn pump back on: counter clock-wise operation. 58. Start detection: remove cap of pen, lower pen to paper, set paper speed (e.g., to 300 cm/h). 59. Start program before solution reaches the drop former (press “Run”). 60. During the run, check whether system is leaky, whether fractions have equal (and correct) volume. 61. At the end, press “End” to return to the starting window. – End fractionation: 62. Pump stops automatically at the end of the last fraction. 63. Elevate pen. 64. Keep fractions on ice/freeze in liquid nitrogen/etc., as desired. 65. Press “ADVANCE,” then “Tube.” 66. Remove drop former from Foxy Jr. arm and place in ddH2O (use warm ddH2O if you are fractionating the last tube). 67. Press “ADVANCE,” then “Park.” 68. Switch pump direction to clock-wise operation. 69. Check paper of UV detection: switch off paper movement after the profiles are separated enough.

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70. Pump speed can be increased but has to be back at 55% for next run. 71. Pump until upper level of 60% sucrose in the tube approaches needle level, then stop pump (switch to “STOP”). 72. Press “ADVANCE,” then “Tube.” 73. Dry drop former and put back into Foxy Jr. arm. 74. Press “ADVANCE,” then “Park.” – Repeat fractionation steps for all other tubes. REMARKS 1. For run-off gradients that do not require preservation of polysomes, extracts are prepared in lysis buffer without cycloheximide treatment (Kemmler, Occhipinti, Veisu, & Panse, 2009). 2. To assess the salt stability of ribosomal subunits, cell extracts are prepared in lysis buffer containing 50 mM Tris–HCl pH 7.5, 50 mM KCl, 1 mM DTT (no MgCl2 and cycloheximide) and separated by ultracentrifugation (40,000 rpm, 4  C, 6 h) in 11.2 ml 15–30% (w/v) sucrose gradients made in the same lysis buffer (Parnell & Bass, 2009). 3. For determining the subunit stoichiometry under high salt conditions, lysates are prepared in lysis buffer containing 50 mM Tris–HCl pH 7.5, 800 mM KCl, 10 mM MgCl2, 1 mM DTT, and separated by ultracentrifugation (40,000 rpm, 4  C, 4 h) in 11.2 ml of 7–50% (w/v) sucrose gradients made in the same lysis buffer (Parnell & Bass, 2009).

20.3 ISOLATION OF PRERIBOSOMAL PARTICLES BY TAP The eukaryotic ribosome assembly pathway remained largely refractory to biochemical interrogation for nearly 30 years. The breakthrough occurred in 2001 with the development of efficient affinity purification protocols combined with sensitive mass spectrometry (Puig et al., 2001; Rigaut et al., 1999). By employing the powerful “TAP” several groups have unraveled the compositions of the 90S, 60S, and 40S preribosomal particles which accelerated the analysis of the ribosome assembly pathway (Grandi et al., 2002; Nissan et al., 2002; Scha¨fer et al., 2003). These studies culminated in the sequential ordering of preribosomal particles along the 40S and 60S biogenesis pathways (Figs. 20.1 and 20.3). Yeast strains expressing TAP-tagged versions of trans-acting factors are commercially available as collections from EUROSCARF (http://web.unifrankfurt.de/fb15/mikro/euroscarf/ord_tap.html) and Thermo-Scientific (http:// www.thermoscientificbio.com/gene-expression-cdnas-orfs/non-mammalian-cdnasand-orfs/yeast). Alternatively, trans-acting factors of interest can be endogenously tagged at the C-terminus by homologous recombination using either pFA6a- or pYM-based TAP cassette plasmids ( Janke et al., 2004; Longtine et al., 1998).

20.3 Isolation of Preribosomal Particles by TAP

FIGURE 20.3 Tandem affinity purification of pre-60S and pre-40S subunits. TAP was performed via the indicated TAP-tagged bait proteins of Ssf1, Rix1, Arx1, Kre35 for pre-60S subunit and Noc4, Enp1, Rio2, Asc1 for pre-40S subunit purification. The Calmodulin Sepharose eluates were analyzed on NuPAGE 4–12% gradient gels followed by Silver staining.

20.3.1 Buffers Lysis buffer Tris–HCl pH 7.5 NaCl MgCl2 Igepal CA-630

50 mM 50–200 mM 1.5 mM 0.15% (v/v)

Elution buffer Tris–HCl pH 8 NaCl Ethylene glycol tetraacetic acid (EGTA)

10 mM 50 mM 5 mM

20.3.2 Protocol 1. Grow 2–4 l of yeast culture in appropriate medium up to O.D.600 ¼ 3–3.5 and harvest by centrifugation (5000 rpm, 15 min, 4  C).

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2. Wash and combine cells in 50 ml dH2O and transfer cells to a 50-ml Falcon tube. 3. Pellet cells (5000 rpm, 5 min, 4  C) and discard supernatant. At this step, the cell pellet can be snap frozen in liquid N2, and stored at 80  C. 4. Thaw the cell pellet and adjust the volume to 25 ml with lysis buffer containing 1/2 tablet of complete protease inhibitors (Roche), 1 mM PMSF and 1 mM DTT. Transfer the cell suspension to the grinding bowl and wash the tube with additional 5 ml lysis buffer. Add 25 ml glass beads (400–600 mm) and lyse the cells at 500 rpm for 20 min in a Pulverisette 6 planetary mill (Fritsch) at 4  C. 5. Transfer the cell lysate into a 50-ml Falcon tube by pushing the lysate through a 50-ml syringe, such that glass beads are retained in the syringe. Wash the glass beads once with 5 ml lysis buffer and combine the wash with the cell lysate. 6. The lysate is clarified by centrifugation first at 5000 rpm for 10 min at 4  C, and subsequently after transferring to a new centrifuge tube at 18,000 rpm for 30 min at 4  C. 7. Add 150 ml equilibrated IgG Sepharose (in lysis buffer, GE Healthcare) to the cleared cell lysate and incubate for 1.5 h at 4  C on a rotating wheel. 8. The IgG Sepharose beads are collected in a disposable 10 ml column (Bio-Rad) and washed twice with 5 ml lysis buffer containing 0.5 mM DTT. Let liquids flow through. 9. Close the bottom of the column and seal with parafilm. Tobacco etch virus (TEV) protease cleavage is performed in 5 ml lysis buffer containing 0.5 mM DTT and 10 ml TEV protease (1 mg/ml) to release bound preribosomal particles from IgG Sepharose. Seal the column and rotate for 2 h at 16  C (or overnight at 4  C). 10. The 5 ml TEV-eluate is drained by gravity flow into a new disposable 10 ml column (closed and sealed at the bottom). Wash IgG Sepharose with 4 ml lysis buffer (no DTT) and collect the flow through in the same column. 11. Add CaCl2 and DTT to the TEV-eluate to final concentrations of 2 and 1 mM, respectively. 12. Add equilibrated 150 ml Calmodulin Sepharose (in lysis buffer, GE Healthcare) to the column containing the TEV-eluate. Close and seal the column and incubate for 1.5 h at 4  C on a rotating wheel. 13. Collect the Calmodulin Sepharose beads by opening the column, the flow through is drained by gravity. Wash beads with 10 ml lysis buffer containing 2 mM CaCl2 and 1 mM DTT. 14. Elution of bound TAP particles is performed by incubating the Calmodulin beads with 300 ml elution buffer at 35  C for 10 min. Repeat this step 3. 15. The proteins in the Calmodulin eluates are precipitated by adding trichloroacetic acid (TCA) to final concentration of 10% (v/v), and incubated on ice for 15 min. 16. The contents are pelleted (14,000 rpm, 4  C, 10 min), washed once with 1 ml cold acetone and pelleted again.

20.4 Monitoring Localization of the 40S Preribosome

17. The protein pellet is air dried and resuspended in 20–50 ml 1  LDS sample buffer (Invitrogen). 18. Samples are heated at 70  C for 10 min, separated on NuPAGE 4–12% Bis–Tris gradient gels and subjected to silver staining and Western analyses. REMARK To analyze the isolated preribosomal particles by different mass spectrometry approaches (shotgun, selecting reaction monitoring, data independent acquisition SWATH-MS), proteins in the TEV or Calmodulin eluates are precipitated using TCA and resuspended in denaturing buffer (8 M urea, 50 mM ammonium bicarbonate, and 5 mM EDTA). Next, proteins are reduced with 12 mM dithiothreitol for 30 min at 32  C and alkylated with 40 mM iodoacetamide for 45 min at 25  C. The samples are diluted 1:5 with 0.1 M ammonium bicarbonate and digested overnight with sequencing-grade porcine trypsin (Promega) at an enzyme/substrate ratio of 1:100. The digestion is stopped with formic acid to a final concentration of 2%. The peptide mixtures are desalted on Sep-Pak C18 cartridges (Waters), eluted with 80% acetonitrile, dried by vacuum centrifugation, and resuspended in 0.15% formic acid (Altvater et al., 2012).

20.4 MONITORING LOCALIZATION OF THE 40S PRERIBOSOME BY FLUORESCENCE IN SITU HYBRIDIZATION Nucleoplasmic localization of the small subunit reporter S2-GFP indicates impairment in early nucleolar assembly or late maturation/nuclear export of pre-40S subunits. In vivo localization of the 50 portion of the internal transcribed spacer 1 (ITS1), present within 20S pre-rRNA, monitored by fluorescence in situ hybridization (FISH) is a rapid way to distinguish between the two possibilities. In WT cells, due to rapid nuclear export of pre-40S subunits, Cy3-ITS1 is seen in the nucleolus (arrow), but not in the nucleoplasm (DAPI, arrowhead, Fig. 20.4) (Grosshans, Hurt, & Simos, 2000; Moy & Silver, 1999). Upon nuclear exit of pre-40S subunits, cytoplasmic cleavage of the 20S pre-rRNA into mature 18S rRNA is followed by the efficient degradation of the 50 portion of ITS1 by the nuclease Xrn1. An increase in nucleoplasmic signal of Cy3-ITS1 fluorescence as compared to WT cells indicates the accumulation of pre-40S subunits in the nucleoplasm as seen for yrb2△ cells (Fig. 20.4). In contrast, mutants that are impaired in early nucleolar assembly steps (bud22△) show only nucleolar Cy3-ITS1 fluorescence that is very similar to that in WT cells (Fig. 20.4). •

The protocol for FISH we use is adapted from Grosshans et al. (2000) and nearly identical to the one described in Chapter 19 in the case of tRNA probes, except that we include 10 mM Ribonucleoside Vanadyl Complex along with zymolyase during spheroplasting. In addition, we use RNase-free formaldehyde.

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FIGURE 20.4 Localization of 20S pre-rRNA by ITS1 fluorescence in situ hybridization. Indicated strains were grown to mid-log phase and the in vivo localization of the 50 portion of ITS1, present within 20S pre-rRNA, was monitored by FISH using a Cy3-labeled oligonucleotide complementary to the 50 portion of ITS1. Nuclear and mitochondrial DNA was stained with DAPI. The yrb2△ mutant, impaired in nuclear export of pre-40S subunits, exhibits a nuclear accumulation of Cy3-ITS1. In the bud22△ mutant, impaired in early nucleolar assembly steps, ITS1 localization is restricted to the nucleolus that is very similar to WT cells. Arrow, nucleolus; arrowhead, nucleoplasm (DAPI). Scale bar ¼ 5 mm.

The Cy3-labeled ITS1 oligonucleotide probe (50 -Cy3-ATG CTC TTG CCA AAA CAA AAA AAT CCA TTT TCA AAA TTA TTA AAT TTC TT-30 , ThermoScientific) is complementary to the 50 end of ITS1 (Faza, Chang, Occhipinti, Kemmler, & Panse, 2012; Pertschy et al., 2009).

20.5 ANALYSIS OF SHUTTLING TRANS-ACTING FACTORS BY HETEROKARYON ASSAYS Maturation of preribosomal particles occurs in the nucleolus, nucleoplasm, and in the cytoplasm before translation initiation. Cytoplasmic maturation involves sequential removal of transport receptors and nucleolar/nuclear trans-acting factors that travel with preribosomal particles to the cytoplasm to be released. Shuttling trans-acting factors and transport factors very often display nucleolar/nuclear localization (Bud20-GFP, Fig. 20.5). This steady-state localization is often misleading since it

20.5 Analysis of Shuttling Trans-Acting Factors by Heterokaryon Assays

FIGURE 20.5 Characterization of shuttling trans-acting factors by the heterokaryon assay. (A) Principle of the heterokaryon shuttling assay. Upon mating, the kar1-1 mutation prevents the fusion of the two nuclei, resulting in a heterokaryon that shares the same cytoplasm. A factor that shuttles between the nucleus and cytoplasm enters the nucleus of the kar1-1 mutant (labeled with Nup82-mCherry, left). In contrast, a nonshuttling factor does not appear in the Nup82-mCherry-labeled nucleus (right). (B) Cells expressing GFP-tagged shuttling transacting factor Bud20 or nonshuttling protein Gar1 were mated with a strain expressing kar1-1 and Nup82-mCherry. Heterokaryons were analyzed by fluorescence microscopy. Scale bar ¼ 5 mm.

does not reveal their need to travel to the cytoplasm for their release from preribosomal particles. The heterokaryon assay is a simple and reliable assay that can confirm the shuttling behavior of nucleolar/nuclear localized trans-acting factors and transport factors (Belaya, Tollervey, & Kosˇ, 2006; Conde & Fink, 1976; Vallen, Hiller, Scherson, & Rose, 1992). A strain expressing trans-acting factor fusion with GFP is mated with a kar1-1 mutant, in which mating and cell conjugation is not followed by nuclear fusion, leading to heterokaryon formation. To distinguish the two nuclei in the resulting heterokaryon, the nucleoporin Nup82 of the kar1-1 mutant is fused to mCherry. Typical controls for these analyses must include a shuttling (Bud20-GFP) and a nonshuttling (Gar1-GFP) factor. Gar1-GFP is never seen in the nucleus of the kar1-1 mutant (red signal), whereas Bud20-GFP localized to both nuclei (Fig. 20.5).

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20.5.1 Strains a. MATa strain containing GFP fusion of a trans-acting factor of interest b. MATa kar1-1 mutant expressing Nup82-mCherry

20.5.2 Protocol 1. Strains are grown to O.D.600  1. 2. In a 2-ml Eppendorf tube, 1 ml of both strains a and b are mixed and incubated at 30  C for 15 min. 3. The mixture is concentrated onto a 0.45-mm nitrocellulose filter by using a vacuum filtration system (Millipore) and the filter is subsequently placed on a YPD plate and incubated at 30  C for
1 h (until heterokaryons begin to form). 4. The filter is transferred to a YPD plate containing cycloheximide (50 mg/ml) to block de novo translation of the GFP-tagged protein and incubated for 1–2 h. 5. Use an inoculation loop to gently scrape off cells from the plate and resuspend in 2 ml dH2O on a microscopy glass slide. Press over a cover slip and inspect the GFP and mCherry signal under a fluorescence microscope. REMARK Others (Yao et al., 2010) and we have observed that the heterokaryon assay can be performed reliably without treatment with cycloheximide (in this case step 4 can be skipped). We advice to perform the heterokaryon assay both with and without cycloheximide treatment.

20.6 MATERIAL, REAGENTS AND YEAST MEDIA 20.6.1 Material (listed in alphabetical order) • • • • • • • • • • • •

50/60 ml syringe (Sanitex, 1060) Centrifuge 5804 R (Eppendorf, 5805 000.327) Centrifuge Sorvall RC3BP with H-6000A Rotor (Thermo-Scientific, 75007530, 11250) Cover glasses (VWR, 631-0137) Density gradient fractionator system and gradient former: TrisTM pump, Foxy Jr., Gradient Former Model 160, UA-6 detector (TELEDYNE Isco) Disruptor Genie (Scientific Industries, SI-D278) Fluorescence microscope Leica DM6000 B (Leica) Glass beads 400–600 mm (SiLibeads, 45015) Incubator IFE 600 (Memmert) Incubator shaker ISF1-X (Kuhner) Inoculation loop (Greiner, 731101) Membrane filters 0.45 mm (Millipore, HAWP04700)

20.6 Material, Reagents and Yeast Media

• • • • • • • • • • •

Microcentrifuge 5415R (Eppendorf, 022621408) Microscope slides (VWR, 631-1550) Needle Microlance 3, 0,9 mm  40 mm (BD, 301300) Planetary mill Pulverisette 6 with 80 ml grinding bowl (Fritsch, 06.2000.00 50.4100.00) Poly-Prep Chromatography Columns (Bio-Rad, 731-1553) Polyallomer tubes SW41 (Beckman Coulter, 331372) Spectrophotometer Ultrospec 2100 pro (GE Healthcare, 80-2112-21) Sterifil Aseptic System (Millipore, XX1104700) Syringe plug 2.5 ml (Terumo, SS02S) Thermomixer comfort (Eppendorf ) Ultracentrifuge Optima L-90K with Swing bucket rotor SW 41 Ti (Beckman Coulter, 365672, 331362)

20.6.2 Reagents (listed in alphabetical order) • • • • • • • • • • • • • • • • • • • • • • • •

• •

Acetone (Merck, 1.00014.1000) CaCl2 (Sigma–Aldrich, 22,350-6) cOmplete protease inhibitor cocktail tablets, EDTA free (Roche, 11873580001) Cy3-labeled ITS1 oligonucleotide probe ( Jakovljevic et al., 2004) (Thermo-Scientific) Cycloheximide (AppliChem, A0879) DAPI (Sigma–Aldrich, D9542) Denhardt’s solution (Sigma–Aldrich, D2532) Difco Agar (BD, 214 530) Dithiothreitol (DTT) (AppliChem, A1101) Ethylene glycol tetraacetic acid (EGTA) (Sigma–Aldrich, E8145) Formaldehyde (Sigma–Aldrich, 47608) Glucose (Sigma–Aldrich, G8270) Glycerol (AppliChem, A1123) Igepal CA-630 (Sigma–Aldrich, I88969) IgG Sepharose 6 Fast Flow (GE Healthcare, 17-0969-01) LDS sample buffer (Invitrogen, NP0008) MgCl2 (Fluka, 63072) NaCl (Merck, 1064045000) NH4Cl (Sigma–Aldrich, A9434) NuPAGE 4–12% Bis–Tris (Novex, NP0321BOX) Phenylmethanesulfonyl fluoride (PMSF) (AppliChem, A0999) Ribonucleoside Vanadyl Complex (Sigma–Aldrich, R3380) Sucrose (Sigma–Aldrich, 84100) TEV protease (expressed and purified from plasmid pTH24-TEVSH that originates from the Berglund laboratory) (van den Berg, Lo¨fdahl, Ha¨rd, & Berglund, 2006) TCA (Sigma–Aldrich, T6399) Tris–HCl (Sigma–Aldrich, T3253)

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20.6.3 Yeast media •

• •

•

Synthetic complete (SC) media: 0.69% yeast nitrogen base without amino acids (ForMedium, CYN0410) 2% glucose (Sigma–Aldrich, G8270) 0.6–0.8% appropriate drop-out supplements complete mixtures (ForMedium) SC plates (SC, 2% agar) Yeast-extract peptone dextrose (YPD) media: 1% yeast extract, (ForMedium, YEM03) 2% peptone (ForMedium, PEP03) 2% glucose (Sigma–Aldrich, G8270) YPD plates (YPD, 2% agar)

CONCLUSIONS The development of diverse approaches in budding yeast have created an exciting time for the community involved in studying ribosome assembly and transport. Genetic perturbation of trans-acting factors, in particular >50 energy-consuming enzymes can now be combined with TAP and sensitive targeted proteomic approaches, together with cell biological approaches. These analyses will give us mechanistic insights into the highly dynamic nature of preribosomal particles as they travel from the nucleolus to potentially specific sites in the cytoplasm. Moreover, the combination of TAP and Cryo-electron microscopy (Bradatsch et al., 2012; Nissan et al., 2004; Scha¨fer et al., 2006) should give us high-resolution snapshots of the ribosome assembly line.

Acknowledgments V. G. P. is supported by grants from the Swiss National Science Foundation and the ETH Zurich. V. G. P. is the recipient of a Starting Grant Award (EURIBIO260676) from the European Research Council.

References Altvater, M., Chang, Y., Melnik, A., Occhipinti, L., Schu¨tz, S., Rothenbusch, U., et al. (2012). Targeted proteomics reveals compositional dynamics of 60S pre-ribosomes after nuclear export. Molecular Systems Biology, 8. Baßler, J., Klein, I., Schmidt, C., Kallas, M., Thomson, E., Wagner, M. A., et al. (2012). The conserved Bud20 zinc finger protein is a new component of the ribosomal 60S subunit export machinery. Molecular and Cellular Biology, 32(24), 4898–4912. Belaya, K., Tollervey, D., & Kosˇ, M. (2006). FLIPing heterokaryons to analyze nucleocytoplasmic shuttling of yeast proteins. RNA, 12(5), 921–930.

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Milkereit, P., Strauss, D., Bassler, J., Gadal, O., Ku¨hn, H., Schu¨tz, S., et al. (2003). A Noc complex specifically involved in the formation and nuclear export of ribosomal 40 S subunits. Journal of Biological Chemistry, 278(6), 4072–4081. Moy, T. I., & Silver, P. A. (1999). Nuclear export of the small ribosomal subunit requires the Ran-GTPase cycle and certain nucleoporins. Genes & Development, 13(16), 2118–2133. Nissan, T. A., Baßler, J., Petfalski, E., Tollervey, D., & Hurt, E. (2002). 60S pre-ribosome formation viewed from assembly in the nucleolus until export to the cytoplasm. EMBO Journal, 21(20), 5539–5547. Nissan, T. A., Galani, K., Maco, B., Tollervey, D., Aebi, U., & Hurt, E. (2004). A pre-ribosome with a tadpole-like structure functions in ATP-dependent maturation of 60S subunits. Molecular Cell, 15(2), 295–301. Oeffinger, M., Dlakic´, M., & Tollervey, D. (2004). A pre-ribosome-associated HEAT-repeat protein is required for export of both ribosomal subunits. Genes & Development, 18(2), 196–209. Panse, V. G. (2011). Getting ready to translate: Cytoplasmic maturation of eukaryotic ribosomes. CHIMIA International Journal for Chemistry, 65(10), 765–769. Panse, V. G., & Johnson, A. W. (2010). Maturation of eukaryotic ribosomes: Acquisition of functionality. Trends in Biochemical Sciences, 35(5), 260–266. Parnell, K. M., & Bass, B. L. (2009). Functional redundancy of yeast proteins Reh1 and Rei1 in cytoplasmic 60S subunit maturation. Molecular and Cellular Biology, 29(14), 4014–4023. Pertschy, B., Schneider, C., Gna¨dig, M., Scha¨fer, T., Tollervey, D., & Hurt, E. (2009). RNA helicase Prp43 and its co-factor Pfa1 promote 20 to 18 S rRNA processing catalyzed by the endonuclease Nob1. Journal of Biological Chemistry, 284(50), 35079–35091. Puig, O., Caspary, F., Rigaut, G., Rutz, B., Bouveret, E., Bragado-Nilsson, E., et al. (2001). The tandem affinity purification (TAP) method: A general procedure of protein complex purification. Methods, 24(3), 218–229. Rigaut, G., Shevchenko, A., Rutz, B., Wilm, M., Mann, M., & Seraphin, B. (1999). A generic protein purification method for protein complex characterization and proteome exploration. Nature Biotechnology, 17(10), 1030–1032. Scha¨fer, T., Maco, B., Petfalski, E., Tollervey, D., Bo¨ttcher, B., Aebi, U., et al. (2006). Hrr25dependent phosphorylation state regulates organization of the pre-40S subunit. Nature, 441(7093), 651–655. Scha¨fer, T., Strauß, D., Petfalski, E., Tollervey, D., & Hurt, E. (2003). The path from nucleolar 90S to cytoplasmic 40S pre-ribosomes. EMBO Journal, 22(6), 1370–1380. Stage-Zimmermann, T., Schmidt, U., & Silver, P. A. (2000). Factors affecting nuclear export of the 60S ribosomal subunit in vivo. Molecular Biology of the Cell, 11(11), 3777–3789. Strunk, B. S., & Karbstein, K. (2009). Powering through ribosome assembly. RNA, 15(12), 2083–2104. Trapman, J., Rete`l, J., & Planta, R. J. (1975). Ribosomal precursor particles from yeast. Experimental Cell Research, 90(1), 95–104. Tschochner, H., & Hurt, E. (2003). Pre-ribosomes on the road from the nucleolus to the cytoplasm. Trends in Cell Biology, 13(5), 255–263. Udem, S. A., & Warner, J. R. (1973). The cytoplasmic maturation of a ribosomal precursor ribonucleic acid in yeast. Journal of Biological Chemistry, 248(4), 1412–1416.

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Vallen, E., Hiller, M., Scherson, T., & Rose, M. (1992). Separate domains of KAR1 mediate distinct functions in mitosis and nuclear fusion. Journal of Cell Biology, 117(6), 1277–1287. ˚ ., Ha¨rd, T., & Berglund, H. (2006). Improved solubility of TEV van den Berg, S., Lo¨fdahl, P.-A protease by directed evolution. Journal of Biotechnology, 121(3), 291–298. Warner, J. R., & Knopf, P. M. (2002). The discovery of polyribosomes. Trends in Biochemical Sciences, 27(7), 376–380. Yao, Y., Demoinet, E., Saveanu, C., Lenormand, P., Jacquier, A., & Fromont-Racine, M. (2010). Ecm1 is a new pre-ribosomal factor involved in pre-60S particle export. RNA, 16(5), 1007–1017.

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Approaches to Studying Subnuclear Organization and Gene–Nuclear Pore Interactions

21

Defne Emel Egecioglu, Agustina D’Urso, Donna Garvey Brickner, William H. Light, and Jason H. Brickner Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA

CHAPTER OUTLINE Introduction ............................................................................................................ 464 21.1 A Quantitative Assay for Gene Localization to the Nuclear Pore Complex in Yeast ........................................................................................... 465 21.1.1 Strain Construction.....................................................................465 21.1.1.1 Inserting the LacO Array Into the Yeast Genome.................465 21.1.1.2 Inserting DNA Zip Code Variants ........................................467 21.1.1.3 Introducing GFP-LacI and a Fluorescent ER Marker ...........468 21.1.2 Microscopy Experiments .............................................................469 21.1.2.1 Materials and Reagents Required for Microscopy Experiments ..................................................................................... 469 21.1.2.2 Microscope Settings........................................................... 469 21.1.2.3 Data Acquisition through Confocal Microscopy ...................469 21.1.2.4 Data Analysis..................................................................... 470 21.1.2.5 Conclusion: Analysis of Gene Localization in Live Cells through Confocal Microscopy............................................................ 471 21.2 Monitoring Interchromosomal Clustering of Genes at the NPC ........................... 471 21.2.1 Strain Construction.....................................................................472 21.2.1.1 RFP-LacI and GFP-TetR Two-dot Assay .............................472 21.2.1.2 GFP-LacI Two-dots Assay .................................................. 473 21.2.2 Microscopy Settings and Methods for Clustering Experiments.........473 21.2.3 Conclusion: Monitoring Interchromosomal Clustering of Genes at the NPC ...................................................................474 21.3 Using Chromatin Immunoprecipitation to Probe Nuclear Organization, Transcription, and Chromatin Structure in Yeast and Human Cells .................... 474 21.3.1 Equipment, Buffers, Solutions, and Reagents for ChIP ..................475 21.3.2 Yeast ChIP.................................................................................477 21.3.3 ChIP in HeLa Cells .....................................................................479 Methods in Cell Biology, Volume 122 Copyright © 2014 Elsevier Inc. All rights reserved.

ISSN 0091-679X http://dx.doi.org/10.1016/B978-0-12-417160-2.00021-7

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21.3.4 qPCR Reaction and Analysis........................................................480 21.3.4.1 qPCR Set Reaction ............................................................ 480 21.3.4.2 Data Analysis..................................................................... 480 21.4 List of Plasmids and Strains ........................................................................... 481 Concluding Remarks, Possible Caveats, and Troubleshooting .................................... 482 Acknowledgments ................................................................................................... 483 References ............................................................................................................. 483

Abstract Many genes in budding yeast Saccharomyces cerevisiae associate with the nuclear pore complex (NPC), which impacts their location within the nucleus and their transcriptional regulation. To understand how eukaryotic genomes are spatially organized, we have used multiple approaches for analyzing the localization and transcription of genes. We have used these approaches to study the role of DNA elements in targeting genomic loci to the NPC and how these interactions regulate transcription, chromatin structure and the spatial organization of the yeast genome. These studies combine yeast molecular genetics with live-cell microscopy and biochemistry. Here, we present detailed protocols for these cytological and molecular approaches.

INTRODUCTION Eukaryotic genomes are spatially organized within the nucleus. Chromosomes fold back on themselves and are positioned in distinct “territories.” The localization of genes with respect to each other and with respect to nuclear landmarks can be coupled to their expression (Egecioglu & Brickner, 2011). One model for this type of regulation is the movement of genes from the nucleoplasm to the nuclear periphery through interaction with the nuclear pore complex (NPC) upon activation. This phenomenon was discovered in the brewer’s yeast Saccharomyces cerevisiae (Brickner & Walter, 2004; Casolari et al., 2004) and has since been observed in flies, worms, and human cells (Liang & Hetzer, 2011). Genome-wide molecular approaches suggest that hundreds of yeast genes physically associate with the NPC (Casolari, Brown, Drubin, Rando, & Silver, 2005; Casolari et al., 2004). Therefore, the interaction of nuclear pore proteins with genes is both widespread and conserved. We have found that interaction of yeast genes with the NPC is controlled by cis-acting promoter elements (Ahmed et al., 2010; Brickner et al., 2012; Light, Brickner, Brand, & Brickner, 2010). These DNA elements are both necessary and sufficient to confer interaction with the NPC and to target genomic loci to the nuclear periphery. For this reason, we have called them DNA zip codes. These DNA elements may control a more global spatial organization of the yeast nucleus; the targeting of genes to the NPC also results in the interchromosomal clustering of genes that share common zip codes (Brickner et al., 2012). To study these phenomena, we have employed cytological and molecular approaches described in this chapter.

21.1 Quantitative Assay for Gene Localization to the NPC

21.1 A QUANTITATIVE ASSAY FOR GENE LOCALIZATION TO THE NUCLEAR PORE COMPLEX IN YEAST We have used a simple quantitative assay to monitor the targeting of genes to the NPC in yeast. As a proxy for interaction of genomic loci with the NPC, we tag the locus of interest with a protein (or fluorescent protein) and then localize this protein with respect to the nuclear envelope through immunofluorescence and confocal microscopy (Brickner, Light, & Brickner, 2010) (Fig. 21.2A). We then quantify the fraction of the cells in which the locus of interest colocalizes with the nuclear envelope. Here, we describe current methods utilized by our lab to visualize fluorescently tagged genomic loci through confocal microscopy.

21.1.1 Strain construction We use a strategy similar to that described previously (Brickner et al., 2010) to create yeast strains that allow the determination of the subnuclear localization of genomic loci. However, whereas the strains described previously are only useful for immunofluorescence, the strains described here can be visualized live. Classical and general molecular biology methods used for this section include lithium acetate yeast transformation (Amberg, Burke, & Strathern, 2006a, 2006b) for both plasmids and fragments and standard bacterial plasmid transformation. All plasmids and strains are listed in the summary table at the end of this chapter. These plasmids, plasmid maps, and strains are available to academic scientists from the Brickner Lab upon request. To visualize the location of a gene of interest with respect to the nuclear envelope, three elements are required: an array of Lac Operators (LacO array) inserted at the genomic locus of interest, the GFP-tagged Lac repressor (GFP-LacI), and an mCherry-tagged marker for the nuclear envelope/endoplasmic reticulum.

21.1.1.1 Inserting the LacO array into the yeast genome The LacO array we use consists of 128 repeats of the Lac Operator cloned into the yeast-integrating plasmid pRS306 (Sikorski & Hieter, 1989), resulting in the plasmid p6LacO128 (Brickner et al., 2010; Brickner & Walter, 2004) (Fig. 21.1A). This yeast shuttle vector carries the URA3 and Ampr (bla, for b-lactamase in Fig. 21.1) markers for selection in yeast and E. coli, respectively. We have used two different approaches to insert the LacO array into the yeast genome: (1) digesting p6LacO128 with a restriction enzyme that cleaves within URA3 to target integration to the endogenous URA3 locus (Fig. 21.1A) or (2) cloning sequences downstream of a gene of interest into the multiple cloning site in p6LacO128 and digesting the resulting plasmid with a restriction enzyme that cleaves within these sequences to direct integration of the LacO array and URA3 at that locus (Fig. 21.1C). The URA3 locus localizes primarily in the nucleoplasm and colocalizes with the nuclear envelope in only 25–30% of the cells (Brickner & Walter, 2004; Taddei et al., 2006) (e.g., Fig. 21.2B). This represents the fraction of the yeast nuclear volume that cannot be resolved from the nuclear envelope by light microscopy and is expected for an

465

FIGURE 21.1 Methodology Used in Strain Construction for Microscopy Inserting the LacO array and testing sequences for zip code activity in the yeast genome. (A) The p6LacO128 plasmid is integrated at URA3 by digestion with StuI. Putative DNA zip codes can be cloned bedside the LacO array using the unique restriction sites on either side of the array. (B) Integration of putative zip codes (yellow Zip segments) next to the LacO array already incorporated within yeast genome through Kanr/Ampr exchange. Left: Putative DNA zip codes are either cloned into the PacI site in the pZipKan plasmid and then introduced into yeast by digestion with KpnI þ EcoRV or incorporated into the cassette by including the putative zip code in the primer and amplifying the cassette by PCR. Right: The KmR cassette with a zip code can be transformed into a yeast strain having the LacO array integrated at URA3. Transformants are selected on G418 medium and screened by PCR for proper integration. (C) General strategy to incorporate the LacO array next to a gene of interest (GOI1). The 30 -end and 30 -UTR of the gene of interest is cloned into p6LacO128. Digestion within this sequence at a unique site promotes integration of the LacO plasmid downstream of the gene of interest. Uraþ transformants are selected on medium lacking uracil.

21.1 Quantitative Assay for Gene Localization to the NPC

FIGURE 21.2 Quantifying Gene Localization at the Nuclear Periphery in Live Cells (A) Individual z-slices from confocal microscopy of yeast cells having the LacO array integrated at URA3 and expressing both GFP-LacI and mCherry on the endoplasmic reticulum/nuclear envelope. The slice corresponding to the brightest, most focused dot was scored (indicated with yellow arrows). Cells in which the center of the green dot does not colocalize with the nuclear envelope were scored as “OFF” (open arrowheads). Cells in which the center of the green dot colocalizes with the nuclear periphery were scored as “ON” (closed arrowheads). (B) Histogram depicting the localization of the URA3 locus with either the LacO array and KmR cassette inserted in place of the Ampr gene (as in Fig. 21.1B) or with the LacO array and the KmR gene with the GRSI zip code inserted in place of the Ampr gene. Three biological replicates of 30–50 cells each were scored for each strain. For reference, the mean peripheral localization of the URA3 gene from Brickner and Walter (2004) is shown with the dashed line.

unbiased distribution (Brickner & Walter, 2004). Therefore, URA3 serves as a negative control for targeting to the NPC. For genes that interact with the NPC, we observe between 50% and 75% colocalization with the nuclear envelope (Fig. 21.2B). The fact that this number is lower than 100% reflects the dynamic nature of the association of genes with the NPC; these genes continuously move and occasionally dissociate from the nuclear periphery (Cabal et al., 2006). Furthermore, most experiments represent a snapshot(s) of an asynchronous culture of cells and targeting of active genes to the NPC is regulated through the cell cycle; for 20–30 min after the initiation of S-phase, localization to the nuclear periphery is lost (Brickner & Brickner, 2010). Cells in G1 or G2/M show higher percent colocalization with the nuclear periphery (Brickner & Brickner, 2010).

21.1.1.2 Inserting DNA zip code variants Much of our work has focused on deciphering the molecular mechanism(s) by which genes are targeted to the NPC. Many genes are targeted to the NPC by cis-acting promoter elements that function as DNA zip codes. To test the ability of DNA

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sequences to function as DNA zip codes, we insert them at URA3, along with a LacO array. We define DNA zip codes as DNA elements that are sufficient to cause URA3 to localize at the nuclear periphery. To test elements for zip code activity, DNA sequences can be cloned adjacent to the LacO array in p6LacO128 and the resulting LacO plasmid can be inserted at URA3 (Ahmed et al., 2010). For small DNA elements, we integrate them directly into the backbone of the p6LacO128 plasmid that has already been integrated at URA3 in yeast (Ahmed et al., 2010; Light et al., 2010, 2013) (Fig. 21.1A). Candidate sequences can be either cloned into the PacI site in an integration cassette within the plasmid pZipKan or included in the primers used for PCR amplification of the Kanr marker from this plasmid (KmR in Fig. 21.1B). Yeast transformants that have replaced a portion of the Ampr gene in the p6LacO128 plasmid at URA3 with the putative zip code and the Kanr gene are selected by plating on G418 medium. The resulting yeast colonies are confirmed through PCR from genomic DNA. The restriction sites available for cloning a desired fragment of DNA or annealed oligonucleotides encoding zip code variants into p6LacO128 are as follows: Between the LacO array and ori (Fig. 21.1A): SciI, XhoI, AbsI, PspXI, KpnI, and Acc65I. Between the URA3 gene and the LacO array (Fig. 21.1A): DrdIV, SacII, BstXI, SacI, AleI, EagI, NotI, SpeI, BamHI, XmaI, SmaI, HindIII, SphI, SalI, and Eco53kI. The primers used to amplify the KmR cassette from the pZipKan plasmid for integration of putative zip codes (Fig. 21.1B) are as listed below. This method is recommended for relatively short inserts (15–20mers). To test longer sequences, cloning annealed oligonucleotides into the PacI site in the pZipKan plasmid followed by digestion of pZipKan with KpnI and EcoRV (Fig. 21.1B) is recommended. Forward primer (without zip code): 50 -AAAAAGGCCGCGTTGCTGGCGTTTTTCCATAGGCTCCGCCCCC CTGCGGATCCCCGGGTTAATTAACATCTTTTACCC-30 Forward primer (with zip code): 50 -AAAAAGGCCGCGTTGCTGGCGTTTTTCCATAGGCTCCGCCCCC CTGCGGATCCCCGGG-[Zip Code]-TTAATTAACATCTTTTACCC-30 Reverse primer: 50 -CGAAATCAAAAAAAAGAATAAAAAAAAAATGATGAATTGAA TTGAGAATTCGAGCTCGTTTAAAC-30

21.1.1.3 Introducing GFP-LacI and a fluorescent ER marker Before introducing the LacO array, we introduce the GFP-LacI and the fluorescent ER marker. This allows screening for transformants with the best LacO arrays by microscopy. The plasmid pAFS144 expressing GFP-LacI is digested with NheI to target for integration at the HIS3 locus. To mark the endoplasmic reticulum and nuclear envelope, we use mCherry fused to an endoplasmic reticulum membrane protein under the control of the GPD promoter. This plasmid (pmCh-ER04) is digested with either BstXI or AflII to target for integration at the TRP1 locus. This plasmid is

21.1 Quantitative Assay for Gene Localization to the NPC

derived from pAC08-mCh-L-TM from the Veenhoff lab (Meinema et al., 2011). The GPD promoter from p416-GPD (Mumberg, Muller, & Funk, 1995) was cloned as a SacI–SpeI fragment in place of the GAL1-10 promoter (using SacI–AvrII sites) in the parent plasmid pAC08-mCh-L-TM to create pGPD-mCh-ER16. The entire promoter-fusion protein-30 -UTR was removed from this plasmid as a KpnI–SacI fragment into the integrating plasmid pRS304 (Sikorski & Hieter, 1989).

21.1.2 Microscopy experiments 21.1.2.1 Materials and reagents required for microscopy Experiments • • • • •

Confocal laser scanning microscope with 488 and 561 nm lasers. In our case, we use the Leica SP5 II LSCM. Immersion oil for microscopes. Microscope slides, 25  75  1 mm: Fisher CAT#12-544-4. Premium cover glass for microscope slides, 24  50 mm: Fisher CAT#12-544-14. LAS AF or LAS AF Lite software, available from Leica.

21.1.2.2 Microscope settings We have used several confocal microscopes with 100 1.44NA objectives with success (Fig. 21.2A). Our lab currently uses a Leica SP5 line scanning confocal microscope with the following settings: the Argon 488 nm and Diode Pumped Solid State 561 nm lasers are used at 10–15% power and images are acquired for a 150 mm  150 mm field at 2048  2048 pixel resolution (without zoom). We collect a z-stack of 20  0.73 mm slices, with a step size of 0.29 mm. This redundancy provides a smoother transition between each slice and enables better visualization of the green dot generated by the GFP-LacI:LacO array interaction (Fig. 21.2A). For each yeast culture, at least two samples are analyzed per slide, and care is taken to ensure that the cells are immobile.

21.1.2.3 Data acquisition through confocal microscopy Even without deconvolution, laser scanning confocal microscopy produces images that are well resolved in the z dimension and have minimal out-of-focus light. Below is the procedure we use to quantify the localization of genes at the nuclear periphery in the strains described in the previous section: 1. Because rich medium (YPD) produces significant fluorescence, cells are grown in Synthetic Dextrose Complete (SDC) medium for several generations (usually overnight) at either room temperature (23  C) or 30  C. Data collection is performed on cultures that are in early to mid-log phase of growth (OD600  0.5). 2. 10–50 ml of cells are harvested by centrifugation at 4000 rpm (3041  g) for 1 min, resuspended in fresh SDC media and concentrated 10–20  (final OD600 ¼ 5–10), and stored at room temperature until visualized. Note: Chilling

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cells on ice or chemically inhibiting metabolism to arrest them has effects on gene targeting to the NPC and is not recommended. 3. The cells should be promptly visualized under the microscope, although we have not observed any significant difference between cells that were scored immediately after harvesting and cells that were scored 2 h after harvesting. 4. Immediately before imaging, 1 ml of the concentrated cells is spotted onto a microscope slide and covered with a cover glass. The droplet of cells should spread in the thin space between the slide and the cover glass in an even fashion, without reaching the edges. Using a small volume reduces the movement of cells during microscopy.

21.1.2.4 Data analysis Each nucleus in which the GFP-LacI dot colocalizes with the nuclear envelope is scored as peripheral (ON); all other nuclei are scored as nucleoplasmic (OFF) (Fig. 21.2A and B). There are a few key considerations when deciding if a dot is ON or OFF: 1. The z-stack should include the entire nucleus. This ensures that the nucleus is visible and accounted for and that within the total volume of the entire nucleus there is only a single, relatively immobile dot. We do not score dots that move between the nucleoplasm and the nuclear periphery during the scan or nuclei with multiple dots. Dividing nuclei can also add some ambiguity to the scoring of the position of the dot, so it is important to track the entire volume of the nucleus through the z-stacks. 2. The step size (the distance between z-slices) should be less than half the thickness of a z-slice (i.e., 0.34 mm step size for a 0.73-mm slice thickness). This oversampling ensures a smooth transition between each z-stack and a greater chance of finding the z-position at which the dot is brightest and most focused. 3. The position of the LacI dot should be scored within the z-slice where it is brightest and most focused. 4. The nuclear envelope should be clearly visible as a bright, continuous ring with a dark nucleoplasm. Do not score green dots that are at the top or the bottom of the nucleus, where the nuclear envelope appears as a sheet or in nuclei in which the membrane is not a continuous ring (Fig. 21.2A). 5. If the center of the green dot overlaps the nuclear envelope, it is scored as ON (Fig. 21.2A). 6. We do not score nuclei if the position of the dot is ambiguous because it has moved during the scan, the nuclear envelope is not well resolved or there are two dots within the nucleus. 7. It is helpful to view each channel (green and red) separately as well as the merged image. This can resolve any ambiguity as to the extent of overlap of the LacI dot (green) with the nuclear envelope (red). The Leica LAS AF or LAS AF Lite software also enables the drawing of lines and arrows for highlighting dots and marking cells.

21.2 Monitoring Interchromosomal Clustering of Genes at the NPC

8. As cells in a field are scored, we mark them using the basic shape drawing utility to indicate that they have been scored and to identify them as peripheral or nucleoplasmic. This helps avoid scoring the same nucleus more than once and speeds the counting of each class of cells. The .lif file can be saved with the shapes drawn around the cells. For each experiment, we score at least 30 nuclei per biological replicate and the mean percentage peripheral localization and the standard error of the mean is calculated from at least three biological replicates (Fig. 21.2B). An unpaired t test is applied to determine if two strains or conditions are significantly different. An alternate and more laborious method for performing this experiment has been used by a number of research groups (Meister, Gehlen, Varela, Kalck, & Gasser, 2010; Taddei et al., 2006). This method divides the nucleus into three concentric “zones” of equal area and the fraction of the population in which the LacI dot localizes within each of these zones is determined. This method can reveal more subtle changes in localization than the method we have described. Because yeast cells do not have perfectly spherical nuclei, computational aid is required to calculate the zones for each nucleus.

21.1.2.5 Conclusion: Analysis of gene localization in live cells through confocal microscopy The analysis of fluorescently labeled yeast cells through confocal microscopy provides a robust method for studying the localization of a genomic locus. Our studies have focused on quantifying the localization of a gene with respect to the nuclear envelope in a population of cells under specific conditions. In addition to this type of “snapshot” experiment, it is also possible to examine the dynamics of gene positioning within the nucleus over time, preferably using spinning-disc confocal microscopy (Meister et al., 2010).

21.2 MONITORING INTERCHROMOSOMAL CLUSTERING OF GENES AT THE NPC In addition to causing genes to move from the nucleoplasm to the nuclear periphery, the interaction of DNA zip codes with the NPC can also lead to interchromosomal clustering of genes that share the same zip codes (Brickner et al., 2012). To observe such clustering, we have used two different strategies: (1) tagging loci with different repressor proteins fused to different fluorescent proteins (i.e., RFP-LacI/LacO array and GFP-TetR/TetO array) or (2) integrating LacO arrays of different length that give rise to a larger and a smaller dot. The position of these two loci with respect to each other is determined by measuring the distribution of distances between them in the population (Brickner et al., 2012). Here, we describe both approaches and the important considerations for the microscopy that differ from the method described above.

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21.2.1 Strain construction 21.2.1.1 RFP-LacI and GFP-TetR two-dot assay All plasmids and strains are listed in the summary table at the end of this chapter. These plasmids, plasmid maps and strains can be obtained from the Brickner Lab upon request. 1. RFP-LacI þ GFP-TetR two-dot assay We have used this technique to mark the INO1 gene with GFP-Tet repressor and compare its localization to other genes marked with RFP-Lac repressor (Fig. 21.3A). For introduction of the Lac repressor array into the yeast genome, the methods described above can be used. To integrate the Tet repressor array at INO1,

FIGURE 21.3 Two-Dot Experiments in Live Yeast Cells for Studying Interchromosomal Clustering by Confocal Microscopy For both panels, white scale bar is 1 mm and distances between dots increase from left to right. (A) Examples from the GFP-LacI two-dots assay. This is a diploid cell having both copies of the GAL1 gene marked with the LacO array. The dots are indicated with arrows that have closed arrowheads and straight lines. The large arrows indicate the localization of the large dot (LacO256) and the small arrows indicate the localization of the smaller dot (LacO128). (B) Examples of the RFP-LacI þ GFP-TetR two-dot assay. The green GFP-TetR dot marks the TetO array integrated at INO1 and the RFP-LacI red dot marks the LacO array and component(s) of the INO1 gene integrated at URA3. The green dots are indicated with arrows that have closed arrowheads and the red dots are indicated with arrows that have closed arrowheads and dashed lines.

21.2 Monitoring Interchromosomal Clustering of Genes at the NPC

the 30 -end of INO1 was cloned as an XbaI–HindIII fragment into the TetO array plasmid p15816 (Cabal et al., 2006; Grund et al., 2008), producing p15816-INO1. Into a strain expressing GFP-TetR (integrated at LEU2) (Grund et al., 2008) and RFP-LacI under the control of the ADH2 promoter (CEN/ARS plasmid pME08; select for by growth in SDC-TRP; Jiang, Frey, Evans, Friel, & Hopper, 2009), plasmid p15816-INO1 is integrated at INO1 (after digestion with MscI) and LacO array plasmids are integrated at various genomic sites using the strategy schematized in Fig. 21.1C. To derepress the ADH2 promoter and allow expression of RFP-LacI, cells were grown in media containing 2% ethanol as a carbon source for at least 6 h prior to harvesting.

21.2.1.2 GFP-LacI two-dots assay We have also localized two GFP-LacI marked genes with respect to each other (Brickner et al., 2012). For these experiments, we have localized the HSP104 gene with respect to other genes. Plasmid pFS3013, having 256 Lac repressor binding sites beside the HSP104 gene, was integrated at HSP104 by digesting with BsrGI (Dieppois, Iglesias, & Stutz, 2006) in a strain transformed with GFP-LacI plasmid pAFS144 (Robinett et al., 1996) and a plasmid having 128 Lac repressor-binding sites integrated at other sites (i.e., INO1, GAL1, and GAL2). The different sizes of the arrays give rise to dots of distinct size (Brickner et al., 2012). One advantage of this system is that it offers the ability to combine clustering analysis with localization of each gene with respect to the nuclear envelope (Fig. 21.3B) (Brickner et al., 2012). We have also localized the two alleles of the same gene with respect to each other in diploid cells (Fig. 21.3B). In this case, the size of the dots is not distinguishable.

21.2.2 Microscopy settings and methods for clustering experiments These cells can be either fixed and processed for immunofluorescence (Brickner et al., 2012) or imaged as live cells as described above. We have imaged these strains using both a Zeiss 510 line scanning confocal microscope as well as the Leica SP5. Cells are selected for measurement based on the appearance of both dots in the same confocal slice. Thus, we have not attempted to measure distances between spots in the z dimension. We measure the distances between the centers of the dots in at least 100 cells using Zeiss LSM or the Leica LAS AF software. The distribution of distances is then plotted for the population. Also, we have defined the fraction of cells in which the two dots are “clustered” as the fraction of cells in which the center of the dots are 0.55 mm apart. Dots that are 0.5 mm apart are localized within the same 8% of the area of the nucleus (Brickner et al., 2012). These metrics can be used to compare the localization of pairs of loci using either a t-test (for distributions) or a Fisher’s Exact Test (for the fraction of dots that are clustered). Genes that are not clustered show a distribution with a mean distance of  0.85 0.3 mm and 10–15% clustering. Genes that cluster show a very significant shift to shorter distances, with a mean distance of 0.5  0.3 mm and 65% clustering (Brickner et al., 2012).

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21.2.3 Conclusion: Monitoring interchromosomal clustering of genes at the NPC We have described the techniques used in the Brickner Lab to visualize multiple genomic loci and their position in relation to each other on the nuclear envelope in the yeast nuclei. With this method, multiple DNA zip codes can be tested for interchromosomal clustering of different genes or genomic loci.

21.3 USING CHROMATIN IMMUNOPRECIPITATION TO PROBE NUCLEAR ORGANIZATION, TRANSCRIPTION, AND CHROMATIN STRUCTURE IN YEAST AND HUMAN CELLS An orthogonal approach to studying gene localization to the nuclear periphery by microscopy has been to use chromatin immunoprecipitation (ChIP) to monitor the interaction of nuclear pore proteins with chromatin. In fact, this type of experiment led to the initial discovery of this widespread nuclear pore–gene interaction (Casolari et al., 2004). ChIP is a powerful technique that allows the characterization of specific protein/DNA interactions (both direct and indirect) at a given genomic location of interest or genome wide (Hecht, Strahl-Bolsinger, & Grunstein, 1996; Kuo & Allis, 1999). In this procedure, protein and DNA complexes are fixed by crosslinking with formaldehyde, followed by mechanical shearing into small DNA fragments (Orlando, 2000; Orlando, Strutt, & Paro, 1997). Immunoprecipitation of an antigen leads to the coincident recovery of the DNA associated with the target antigen. This DNA can then be recovered, purified and its immunoprecipitation can be quantified by real-time PCR (qPCR). Relative recovery of a locus of interest can then be compared with appropriate control loci (Nelson, Denisenko, & Bomsztyk, 2006). Using ChIP, our lab has studied the interaction of nuclear pore proteins with chromatin, as well as the downstream events associated with transcription or alterations in chromatin structure. In yeast, we have used Tandem Affinity Purification (TAP)tagged forms of several nuclear pore proteins, including components of the core channel (i.e., Nup100), the nuclear basket (i.e., Nup2), and associated factors (i.e., Mlp2, SAGA). We have also examined the binding of transcription factors, the recruitment of RNA polymerase II and the preinitiation complex, incorporation of the histone variant H2A.Z, and the methylation of histone H3 using ChIP (Ahmed et al., 2010; Brickner et al., 2012, 2007; Light et al., 2010, 2013). Finally, we have performed ChIP against nuclear pore proteins, RNA polymerase II, and chromatin marks in HeLa cells (Light et al., 2013). In this section, we will describe our general protocol for ChIP (Fig. 21.4). We include antigen-specific details for ChIP against RNA polymerase II and H3K4me2 or TAP-tagged proteins in yeast and Nup98 in HeLa cells. The immunoprecipitated material is quantified using real-time quantitative PCR and the yield is expressed relative to a sample of the Input.

21.3 Nuclear Organization, Transcription, Chromatin Structure

FIGURE 21.4 Flow chart for the ChIP protocol described in this chapter.

21.3.1 Equipment, buffers, solutions, and reagents for ChIP EQUIPMENT • Branson Sonifier 450 • 8-Tip microtip attachment for sonifier (Cole Parmer catalog# 630-0586) • Magnetic bead harvester (for 1.5–2.0 ml microfuge tubes)

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•

BIO-RAD iCycler iQ5 (Multicolor Real-Time PCR Detection System), Bio-rad Cat# 170-9780

COMMON REAGENTS FOR YEAST AND HUMAN CELL PROTOCOL • TE Buffer • 20% SDS • 36% Formaldehyde stock: Sigma CAT# F8775-500ML • 2.5 M Glycine stock • 100 mM Tris–HCl (pH 7.0) • 2  ChIP Lysis Buffer Stock (no detergent added): 100 mM HEPES–KOH (pH 7.5), 280 mM NaCl, 2 mM EDTA • 1  ChIP-Lysis Buffer: 50 mM HEPES–KOH (pH 7.5), 140 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% NaDOC þ protease inhibitors • Elution Buffer: 50 mM Tris (pH 8), 10 mM EDTA, 1% SDS • Qiagen Buffers PB (CAT# 19066), PE (CAT# 19065) • Qiagen PCR cleanup column (QIAquick® Spin Column LOT# 136244976, MAT#1018215). These might be sold separately or provided with the QIAquick PCR Purification Kit CAT#28104) • Pan-Mouse IgG (Pol ll-antibody): (Dynabeads® Pan Mouse IgG, CAT # 110.41) • Protein-G (H3K4me2) magnetic beads: (Dynabeads® Protein G for Immunoprecipitation CAT# 10004D) • RNase A • Proteinase K • Recombinant Taq purified from E. coli (Engelke, Krikos, Bruck, & Ginsburg, 1990) • 10  SYBR® Green I Nucleic Acid Gel Stain (Life Technologies/Invitrogen CAT# S-7567. This stock is 10,000 ) FOR YEAST PROTOCOL • Acid-washed 0.5-mm glass beads: BioSpec Products, Inc. CAT#11079105 • Nitrocellulose Membrane Filters: Millipore CAT# RAWP09025 • Pierce BCA Assay Kit: Pierce™ CAT# 23225 (500 assays) or CAT#23227 (250 assays) • Protease Inhibitor cocktail tablets “complete” from Roche CAT#11 697 498 001 (with EDTA): Use 1 tablet per 50 ml of lysis buffer. • anti-RNA Polymerase ll (AbCam, ab32356 or Covance 8WG16) • anti-H3K4me2 antibody (AbCam, ab5408) FOR HELA PROTOCOL • TBS Buffer (Tris Buffered Saline, pH 7.5) • MC Lysis Buffer: 3 mM MgCl2, 10 mM Tris (pH 7.5), 10 mM NaCl, and 1% NP-40 • DMEM þ 10% serum þ Pen/Strep • Rabbit Nup98 antibody (AbCam 45584 or Cell Signaling Technologies 2292)

21.3 Nuclear Organization, Transcription, Chromatin Structure

21.3.2 Yeast ChIP Day 1 Label one set of 50-ml conical tubes and one set of microfuge tubes for the day (for each sample). Obtain ice and an aluminum ice block that fits the sonicator tip spacing. Obtain liquid N2. 1. Grow 100 ml of yeast cells in the appropriate medium to an OD600 of 0.8–1 (¼80–100 ODs). Caution: Do not over grow cells as the following crosslinking steps are optimized for the specified OD range. 2. Crosslink cells with fresh 1% formaldehyde. Add 2.7 ml of 36% formaldehyde stock to 100 ml of culture, fix for 15 min to 1 h at room temperature with occasional mixing/swirling. Stop fixation by adding 6 ml of 2.5 M glycine to a final concentration of 150 mM, incubate for 5 min. Crosslinking time must be optimized for each antigen. 3. Harvest cells by vacuum filtration. Wash nitrocellulose membrane filter with 100 ml of 100 mM Tris–HCl (pH 7.0). 4. Scrape cells by washing gently with buffer (do not use spatulae) off of the filter, resuspend in 1 ml of 100 mM Tris (pH 7.0) and transfer to a microfuge tube. Spin 30 s at top speed (13,000 rpm), remove supernatant, and snap freeze cell pellets in liquid N2. Store cell pellets at 80  C. Day 2 Label two new sets of microfuge tubes for all the samples. Label a third set of tubes for the BCA assay. 1. Resuspend cell pellets in 600 ml ice cold ChIP Lysis Buffer þ protease inhibitors. Note: Make fresh from 2  stock buffer. 2. To lyse the cells, add cells to 600 ml of acid-washed glass beads and vortex at 4 C, six times for 45 s each, with 2 min rests on ice. 3. Puncture the bottom of tubes and caps with an 18G needle, place into a second 1.5-ml tube and collect lysate by gentle centrifugation at 4  C, 100 RCF for 5 min (do not use lid in the centrifuge). Discard the dry glass beads. The lysate should all be in the new 1.5-ml tube. Caution: If the lysate did not travel properly to the new tube, check the puncture holes and spin samples again. 4. Spin lysate 10 min at 7000  g at 4  C. 5. Discard supernatant and resuspend pellet in 1 ml fresh Lysis Buffer. Note: Most of the chromatin is found in pellet before sonication. 6. With an 8-tip microtip attachment on a Branson Sonifier 450, sonicate at setting #7, 10  10 s*. All of the tips should be in a tube immersed in either lysate or buffer/water. To prevent foaming, place tip as close to the bottom of tube as possible while sonicating. *Sonication time (i.e., the number of cycles) should be optimized for each fixation condition because longer fixation usually requires more sonication.

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7. 8. 9. 10.

11.

12.

13.

Check the range of DNA sizes after sonication in a 2% agarose gel. They should be centered  500 bp. Mix well. Pellet insoluble material by spinning for 10 min at maximum speed in a microcentrifuge at 4 C. Transfer supernatant to a new 1.5-ml tube and, if necessary, spin again to remove additional debris and collect the supernatant. Determine protein concentration in the supernatant fraction using BCA assay (Pierce). Samples should range between 2 and 5 mg/ml. Dilute all samples to equal protein concentrations using Lysis buffer and use 1 ml for subsequent steps. For 5 mg of protein in each sample, add 5 mg of anti-RNA Polymerase ll or anti-H3K4me2 antibody. Rotate for 2 h at 4  C. For TAP-tagged proteins, skip this step. While step 11 is in progress, equilibrate the magnetic beads. Per 5 ml of antibody, 8 ml of Pan-Mouse IgG (Pol ll-antibody), or Protein-G (H3K4me2) magnetic beads will be required. Harvest beads for 2 min with magnetic bead harvester, remove supernatant (suspension buffer) and resuspend beads in 1-ml Lysis Buffer. Rotate at 4  C for 2 h. For TAP-tagged proteins, use 5–10 ml Pan-Mouse IgG beads per 5 mg of chromatin. Harvest IgG beads for 2 min with magnetic bead harvester. Remove supernatant with a pipette and resuspend beads in the initial volume of Lysis Buffer that was removed from the stock bottle. Add the appropriate volume of beads to each experimental sample and rotate at 4  C overnight.

Day 3 Label one new set of microfuge tubes for the Input and one set of new microfuge tubes for the IP samples. 1. Before harvesting beads, collect 50 ml of “Input” fraction. Add 200 ml TE þ 12.5 ml of 20% SDS to Input fractions and set aside. 2. Pellet beads for 2 min with magnetic bead harvester (Protein G Dynabeads take longer than Pan-mouse IgG Dynabeads), remove supernatant with a pipette and wash 4  1-ml Lysis Buffer, resuspending and harvesting beads for 2 min between each wash. 3. Elute in 100 ml Elution Buffer (50 mM Tris (pH 8), 10 mM EDTA, 1% SDS), incubate 15 min at 65  C. 4. Harvest beads on magnetic rack, transfer eluate to new microfuge tube. Do a second elution by washing beads with 150 ml TE þ 5ml of 20% SDS. Harvest beads again and pool eluates. Label this tube as your IP samples. 5. RNAse treat with 50 mg RNAse A for 10 min at 37  C followed by Proteinase K treatment with 100 mg Proteinase K for 1 h at 42  C. 6. Incubate both IP and Input samples at 65  C for 6–15 h (usually overnight) to reverse crosslinks.

21.3 Nuclear Organization, Transcription, Chromatin Structure

Day 4 Label one set of tubes for each sample (Input or IP) for the Qiagen cleanup step. Remark: In lieu of the cleanup with the Qiagen column, extraction with phenol:chloroform:isoamyl alcohol (25:24:1) can also be performed—see Note (1). 1. Add 1.25-ml Qiagen PB Buffer to each sample, mix and load onto Qiagen PCR cleanup column. 2. Wash column 2  750 ml Qiagen PE. 3. Spin columns dry for 2 min, top speed in the microcentrifuge, let air dry for 2 min and then elute in 30 ml water.

21.3.3 ChIP in HeLa cells 1. Harvest 20 to 50  106 cells by trypsin: Aspirate media and add 6 ml of Trypsin–EDTA to 15-cm plate and incubate at 37  C until most cells have lifted off the plate (about 5 min or so). Add 19 ml of DMEM þ 10% serum and pen/strep to stop reaction and pipet up and down several times to resuspend. Transfer to a 50 ml conical flask. Finally, fix with 1% formaldehyde (from a 36% stock, as for the yeast protocol) at room temperature for 15 min, quenching with 125 mM glycine for 5 min. 2. Harvest cells by spinning for 5 min at 1750 rpm. Wash the pellet twice with TBS. Note: Cell pellet can be frozen in liquid N2 and stored at 80  C. 3. Suspend cells in 10 ml hypotonic cell lysis with MC Lysis Buffer. 4. Spin lysate 5 min at 4  C twice: To do this, pour off supernatant, and add an additional 10 ml of MC Lysis Buffer, spin again at 4  C 1500 rpm for 5 min. Discard the supernatant and then snap freeze the pellet. 5. Freeze the pellet fraction in liquid N2 and store at 80  C. 6. Suspend cell pellets in 1 ml ChIP Lysis Buffer and sonicate using 8-tip microtip attachment on a Branson 450 sonicator. Use cycles of 15 s on 100% duty cycle, 15 s off on ice for approximately 5 min of total sonication time on tip setting #5. 7. Spin 10 min at 4  C at maximum speed in the microcentrifuge. 8. Collect the supernatant fraction and measure the protein concentration using the BCA assay. 9. Equalize the chromatin concentration for all samples and proceed with 1 ml of each. Using a minimum 1 mg/ml chromatin in 1 ml, add 5 mg Rabbit Nup98 antibody (AbCam 45584 or Cell Signaling Technologies 2292) or m414/ H3K4me3/H3K4me2 antibody and Protein-G DynaBeads (use 5–10 mg/1 ml of chromatin). 10. All subsequent steps are the same as listed from step #12 on Day 2 of the yeast ChIP protocol described above.

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Note (1): In lieu of the cleanup with the Qiagen column on Day 4, extraction with phenol:chloroform:isoamyl alcohol (25:24:1) can also be performed: Bring volume of the IP and Input samples to same volume with TE þ 0.67% SDS (usually about 300 ml) and add an equal volume of phenol:chloroform:isoamyl alcohol. Vortex thoroughly, followed by spinning at room temperature for 5 min at maximum speed in a microcentrifuge. Remove top aqueous layer and add to fresh microfuge tube with equal volume chloroform (300 ml). Vortex to mix and spin for 2.5 min at maximum speed at room temperature. Remove the upper aqueous layer and add to 1 ml ice cold 100% ethanol. Add 1/9th volume of 3 M NaOAc (pH 5.2) and 20 mg glycogen or 2 ml of linear acrylamide (50 mg/ml stock) and mix by inversion several times. Incubate at least 1 h at 20  C and pellet by centrifugation at maximum speed at 4  C for 30 min. Remove supernatant and wash pellet with 70% ethanol and spin for 15 min at 4  C. Remove supernatant and let the pellet air dry until pellet becomes translucent. Resuspend in 30 ml TE and heat for 10 min at 42  C to solubilize DNA. Yields using phenol–chloroform extractions are higher than the yields obtained with Qiagen columns. Determining the concentration of DNA is also more straightforward due to the absence of the characteristic background at A260 that comes from Qiagen buffers.

21.3.4 qPCR Reaction and analysis 21.3.4.1 qPCR set Reaction Dilute Input fractions 1:400 and IP samples 1:10 with water. For standard curves, use six fivefold serial dilutions of genomic DNA (the most concentrated is 20 ng/ml for yeast and 100 ng/ml for HeLa). Set up triplicate 25 ml reactions as follows: 5 ml DNA Template 2.5 ml 10  PCR buffer 0.5 ml 50 mM primer mix 0.25 ml dNTP mix (25 mM stock) 0.25 ml 10  SYBR® Green I Nucleic Acid Gel Stain 0.125 ml Taq 16.4 ml water 25 ml total reaction volume We use the BIO-RAD iCycler iQ5 (Multicolor Real-Time PCR Detection system) for amplification. The PCR reaction, along with real-time settings, is listed in Table 21.1.

21.3.4.2 Data analysis To analyze the data, we use the threshold CT qPCR reaction in the linear range of amplification (Brickner et al., 2007). To determine the relationship between DNA concentration and CT value, we use the genomic DNA standard curve. The data

21.4 List of Plasmids and Strains

Table 21.1 PCR cycling settings used for the data analysis of ChIP Set points (temperature) ( C)

Dual time (min or s)

Camera

Cycle 1 Cycle 2

95 95 50 72

   Real-time

Cycle 3 Cycle 4 Cycle 5

95 55 55

3 min 10 s 30 s 45 s Return to Cycle 2, repeat 40 1 min 1 min 10 s

  Melt

The settings used for the qPCR are shown with their corresponding time frames.

are fit to an exponential curve described by: y ¼ ke(rx), where y is the DNA concentration, k and r are constants, and x is the CT value. This standard curve is performed with every qPCR reaction to confirm that the reagents and the machine are working well. We take the average CT of the three technical replicates of the qPCR for each sample, and DNA concentrations are used to calculate enrichment by dividing the IP/Input for the corresponding samples and plotting. For negative controls in yeast ChIP, we have used primers to amplify repressed loci such as PRM1 and GAL1 genes or the intergenic region downstream of the RPA34 gene. For HeLa Nup98 ChIP negative control, we have used primers to the CIITA promoter, GAPDH and b-ACTIN locus.

21.4 LIST OF PLASMIDS AND STRAINS More information on the plasmids and yeast strains described in this chapter is available from the Brickner Lab. Plasmids Plasmid name

Purpose/description

p6LacO128 p6lacO128-GRSIINO1 or p6LacO128-GRSITSA2 pZipKan pZipKan-GRSIINO1 or pZipKan-GRSITSA2 pAFS144 pmCh-ER04 p15816-INO1

Integration of LacO128 at URA3 Integration of LacO128-GRSI at URA3 Replacement of Ampr with Kanr Replacement of Ampr with Kanr-GRSI Integration of GFP-LacI at HIS3 Integration of mCherry-ER marker at TRP1 Integration of TetO array at INO1

Continued

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Plasmids Plasmid name

Purpose/description

p128-tetR-GFP

Integration of GFP-TetR at LEU2 (Michaelis, Ciosk, & Nasmyth, 1997) Nonintegrated RFP-LacI, select for TRP Integration of LacO256 array at HSP104

pME08 pFS3013

Yeast strains Yeast strain

Description

NDY012 DBY434 DEY09 or TKY01 DBY339

Live-microscopy acceptor strain for inserts at URA3 using pZipKan NDY012 transformed with Kanr NDY012 transformed with Kanr-GRSI

DBY546

TetO at INO1 (green) and LacO at INO1 integrated at URA3 (red). Strain is transformed with GFP-TetR and RFP-LacI Diploid cell with both copies of GAL1 with the LacO128 integration. Strain is transformed with GFP-LacI

CONCLUDING REMARKS, POSSIBLE CAVEATS, AND TROUBLESHOOTING We have presented here the methods used in the Brickner Lab to investigate the spatial organization of the yeast genome and its interactions with nuclear envelope and the NPC. In the sections on microscopy, we have discussed our methods for visualizing the nuclear periphery in yeast cells with one (21.1, pp 463-469) or multiple (21.2, pp 469-472) genomic loci. The position of these genomic loci in relation to the nuclear envelope or in relation to each other on the nuclear periphery reveals a spatial component to regulating gene expression that is mediated by DNA zip codes. Data acquired through confocal microscopy via “snapshots” or over time in live cells can provide important information about the molecular basis of this phenomenon. Movies of over a longer time period can provide important information about the dynamics of this process. An important concern when visualizing live cells is their physiological state. We have found that excess centrifugation, chilling on ice, heating, and overgrowth can alter localization results in live cells. As a recent study has pointed out, the localization and expression of certain chromosomal sites can be affected by the utilization of biological repressors that target these loci (Dubarry, Loiodice, Chen, Thermes, & Taddei, 2011). These include tagged-LacI or TetR used to target LacO and TetO arrays, which is a frequently used method in the localization experiments we describe. Tight binding by LacI or TetR, in combination with cryptic silencing elements can lead to transcriptional repression.

References

Thus, it is important to compare expression with and without the arrays and to use orthogonal techniques such as ChIP to confirm that the localization and expression of a gene of interest are not perturbed by the arrays. Also, mutant LacI proteins with lower affinity can also avoid some of these problems (Dubarry et al., 2011). The study of interactions between Nups, PolII, and other markers of gene expression with chromatin as described in detail in Section 21.3 has been indispensable in understanding nuclear and genomic structure and transcriptional regulation. We stress the importance of optimizing the crosslinking and sonication to obtain the best signal-to-noise ratio. Some proteins or complexes are poorly crosslinked, which can lead to discrepancies between proteins, experiments, or laboratories.

Acknowledgments The authors acknowledge support from NIH grant GM 080484 (J. H. B.), a W.M. Keck Young Scholars in Biomedical Research Award (J. H. B.), Cellular and Molecular Basis of Disease Training Program (institutional predoctoral training grant) 2T32GM008061-31 (A. D.), a Rappaport Award for Research Excellence (W. H. L.), and an American Heart Association Postdoctoral Fellowship 13POST14580066 (D. E. E.). The authors also thank members of the Brickner lab for helpful comments on the chapter.

References Ahmed, S., Brickner, D. G., Light, W. H., Cajigas, I., McDonough, M., Froyshteter, A. B., et al. (2010). DNA zip codes control an ancient mechanism for gene targeting to the nuclear periphery. Nature Cell Biology, 12(2), 111–118 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t]. Amberg, D. C., Burke, D. J., & Strathern, J. N. (2006a). High-efficiency transformation of yeast. CSH Protocols, 2006(1). Amberg, D. C., Burke, D. J., & Strathern, J. N. (2006b). “Quick and dirty” plasmid transformation of yeast colonies. CSH Protocols, 2006(1). Brickner, D. G., Ahmed, S., Meldi, L., Thompson, A., Light, W., Young, M., et al. (2012). Transcription factor binding to a DNA zip code controls interchromosomal clustering at the nuclear periphery. Developmental Cell, 22(6), 1234–1246 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, Non-P.H.S.]. Brickner, D. G., & Brickner, J. H. (2010). Cdk phosphorylation of a nucleoporin controls localization of active genes through the cell cycle. Molecular Biology of the Cell, 21(19), 3421–3432 [Research Support, N.I.H., Extramural]. Brickner, D. G., Cajigas, I., Fondufe-Mittendorf, Y., Ahmed, S., Lee, P. C., Widom, J., et al. (2007). H2A.Z-mediated localization of genes at the nuclear periphery confers epigenetic memory of previous transcriptional state. PLoS Biology, 5(4), e81 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t]. Brickner, D. G., Light, W., & Brickner, J. H. (2010). Quantitative localization of chromosomal loci by immunofluorescence. Methods in Enzymology, 470, 569–580 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t].

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Brickner, J. H., & Walter, P. (2004). Gene recruitment of the activated INO1 locus to the nuclear membrane. PLoS Biology, 2(11), e342 [Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, P.H.S.]. Cabal, G. G., Genovesio, A., Rodriguez-Navarro, S., Zimmer, C., Gadal, O., Lesne, A., et al. (2006). SAGA interacting factors confine sub-diffusion of transcribed genes to the nuclear envelope. Nature, 441(7094), 770–773 [Research Support, Non-U.S. Gov’t]. Casolari, J. M., Brown, C. R., Drubin, D. A., Rando, O. J., & Silver, P. A. (2005). Developmentally induced changes in transcriptional program alter spatial organization across chromosomes. Genes & Development, 19(10), 1188–1198 [Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, P.H.S.]. Casolari, J. M., Brown, C. R., Komili, S., West, J., Hieronymus, H., & Silver, P. A. (2004). Genome-wide localization of the nuclear transport machinery couples transcriptional status and nuclear organization. Cell, 117(4), 427–439 [Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, P.H.S.]. Dieppois, G., Iglesias, N., & Stutz, F. (2006). Cotranscriptional recruitment to the mRNA export receptor Mex67p contributes to nuclear pore anchoring of activated genes. Molecular and Cellular Biology, 26(21), 7858–7870. Dubarry, M., Loiodice, I., Chen, C. L., Thermes, C., & Taddei, A. (2011). Tight protein-DNA interactions favor gene silencing. Genes & Development, 25(13), 1365–1370 [Research Support, Non-U.S. Gov’t]. Egecioglu, D., & Brickner, J. H. (2011). Gene positioning and expression. Current Opinion in Cell Biology, 23(3), 338–345 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t Review]. Engelke, D. R., Krikos, A., Bruck, M. E., & Ginsburg, D. (1990). Purification of Thermus aquaticus DNA polymerase expressed in Escherichia coli. Analytical Biochemistry, 191(2), 396–400 [Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, Non-P.H.S. Research Support, U.S. Gov’t, P.H.S.]. Grund, S. E., Fischer, T., Cabal, G. G., Antunez, O., Perez-Ortin, J. E., & Hurt, E. (2008). The inner nuclear membrane protein Src1 associates with subtelomeric genes and alters their regulated gene expression. The Journal of Cell Biology, 182(5), 897–910 [Research Support, Non-U.S. Gov’t]. Hecht, A., Strahl-Bolsinger, S., & Grunstein, M. (1996). Spreading of transcriptional repressor SIR3 from telomeric heterochromatin. Nature, 383(6595), 92–96 [Research Support, NonU.S. Gov’t Research Support, U.S. Gov’t, P.H.S.]. Jiang, F., Frey, B. R., Evans, M. L., Friel, J. C., & Hopper, J. E. (2009). Gene activation by dissociation of an inhibitor from a transcriptional activation domain. Molecular and Cellular Biology, 29(20), 5604–5610. Kuo, M. H., & Allis, C. D. (1999). In vivo cross-linking and immunoprecipitation for studying dynamic Protein:DNA associations in a chromatin environment. Methods, 19(3), 425–433 [Research Support, U.S. Gov’t, P.H.S. Review]. Liang, Y., & Hetzer, M. W. (2011). Functional interactions between nucleoporins and chromatin. Current Opinion in Cell Biology, 23(1), 65–70 [Research Support, Non-U.S. Gov’t Review]. Light, W. H., Brickner, D. G., Brand, V. R., & Brickner, J. H. (2010). Interaction of a DNA zip code with the nuclear pore complex promotes H2A.Z incorporation and INO1 transcriptional memory. Molecular Cell, 40(1), 112–125 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t].

References

Light, W. H., Freaney, J., Sood, V., Thompson, A., D’Urso, A., Horvath, C. M., et al. (2013). A conserved role for human Nup98 in altering chromatin structure and promoting epigenetic transcriptional memory. PLoS Biology, 11(3), e1001524 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t]. Meinema, A. C., Laba, J. K., Hapsari, R. A., Otten, R., Mulder, F. A., Kralt, A., et al. (2011). Long unfolded linkers facilitate membrane protein import through the nuclear pore complex. Science, 333(6038), 90–93 [Research Support, Non-U.S. Gov’t]. Meister, P., Gehlen, L. R., Varela, E., Kalck, V., & Gasser, S. M. (2010). Visualizing yeast chromosomes and nuclear architecture. Methods in Enzymology, 470, 535–567. Michaelis, C., Ciosk, R., & Nasmyth, K. (1997). Cohesins: Chromosomal proteins that prevent premature separation of sister chromatids. Cell, 91(1), 35–45 [Research Support, Non-U.S. Gov’t]. Mumberg, D., Muller, R., & Funk, M. (1995). Yeast vectors for the controlled expression of heterologous proteins in different genetic backgrounds. Gene, 156(1), 119–122 [Research Support, Non-U.S. Gov’t]. Nelson, J. D., Denisenko, O., & Bomsztyk, K. (2006). Protocol for the fast chromatin immunoprecipitation (ChIP) method. Nature Protocols, 1(1), 179–185 [Research Support, N.I.H., Extramural Research Support, Non-U.S. Gov’t]. Orlando, V. (2000). Mapping chromosomal proteins in vivo by formaldehyde-crosslinkedchromatin immunoprecipitation. Trends in Biochemical Sciences, 25(3), 99–104 [Research Support, Non-U.S. Gov’t Review]. Orlando, V., Strutt, H., & Paro, R. (1997). Analysis of chromatin structure by in vivo formaldehyde cross-linking. Methods, 11(2), 205–214. Robinett, C. C., Straight, A., Li, G., Willhelm, C., Sudlow, G., Murray, A., et al. (1996). In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. The Journal of Cell Biology, 135(6 Pt. 2), 1685–1700. Sikorski, R. S., & Hieter, P. (1989). A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics, 122(1), 19–27 [Research Support, Non-U.S. Gov’t Research Support, U.S. Gov’t, P.H.S.]. Taddei, A., Van Houwe, G., Hediger, F., Kalck, V., Cubizolles, F., Schober, H., et al. (2006). Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature, 441(7094), 774–778 [Research Support, Non-U.S. Gov’t].

485

Index Note: Page numbers followed by f indicate figures and t indicate tables.

A Affinity-purification. See also Immuno-depletion; Recombinant protein production and purification split tag purification, 103I tandem affinity purification (TAP) ChIP against TAP-tagged proteins in yeast, 474 TAP in budding yeast, 102, 439, 450–453, 474 TAP on Epitope Pulse labeled proteins (yeast), 149, 154f Aminoacylation (tRNA), yeast, human cells separation, detection, 432–433, 434f Amphibian oocytes. See Xenopus laevis Anchored nuclei (Xenopus, in vitro nuclear reconstitution assay, FESEM imaging), 49–51, 52–54, 52f, 55, 56f, 168 Antibodies. See also Immuno-depletion; Immunofluorescence; Immunoglold labelling antibodies used for anti-Nup116p, 69–70, 70f, 49t anti-FG-Nups, mAb414, 49t, 85, 90–94, 170, 181, 184–186, 292t anti-FLAG antibody, affinity gel, FLAG peptide, 108, 113, 156, 160, 337f anti-GFP cameloid antibodies (nanobody), 229–233, 232f, 235 anti-importins/RANBPs/transportins, 366–367 anti-Lamin A/C, 358 anti-Nup116p, 69–70, 70f anti HA-TAG, 358 colloidal gold-conjugated antibodies, 49, 66, 69, 89–90 nucleoporins and transport factors reagents in C. elegans, 280t relevant antibodies in C. elegans, 292t Antibody-mediated depletion. See Immuno-depletion Artificially engineered Dyn2 interaction domain (Artificially eDID). See Dynein light-chain2 (Dyn2) dimerization-Zipper

B BAPTA, 45, 182, 184–185, 185f, 186

C Caenorhabditis elegans antibodies, 298–299 forward and reverse genetics, 278–287

genome-wide RNAi screens, feeding method, 278–279, 285, 286–287 immunofluorescence, 298–303, 300f live imaging, embryos, cell cycle, 287, 290–296, 291t, 294f, 304 nuclear envelope (NE) integrity, 296–298, 297f nuclear pore proteins (NPPs), nucleoporins, 278–279, 280t, 282t, 291t, 294f transgenesis, 287–290, 288t transmission electron microscopy (TEM), 302–303, 302f transport receptors and Ran-associated proteins, 283t–284t Cap-binding complex (CBC), 20, 396, 397f Cell cycle, 23–29, 167, 171–172, 195, 225, 240–241, 245, 249, 323–327. See also Nuclear envelope breakdown (NEBD); Nuclear pore complexes (NPCs) assembly/disassembly/biogenesis, 24–27 mitotic HeLa extracts, 257, 260–261, 268f, 348 in Xenopus, cytostatic factor (CSF), meiosis, 167, 171, 196 in yeast asynchronous cultures, 323, 324f metaphase arrest, 326–327 S-phase arrest, hydroxyurea, 326 START arrest, a-factor, 325 Cell-fusion. See Heterokaryons assays Chemical inhibitors. See Drugs Chromatin. See also Xenopus laevis demembranted sperm chromatin chromatin immunoprecipitation (ChIP) ChIP in C. elegans, 303–304 ChIP equipment, 473–474 ChIP in yeast and human cells; data analysis, 472–479, 473f Nups-chromatin interaction in C. elegans, 303–304 Chromokinesin hKid, 334, 346–348 Chromosome, mitotic chromosome loading of hKid, 334, 346–348 Conditional yeast mutant strains. See Yeast conditional/mutant strains Confocal microscopy, 8, 381, 469, 471 CRISPR-Cas9, 279–285, 289–290, 304

487

488

Index

CRM1 (Exportin-1, Xpo-1). See Karyopherins (Kaps), importins/exportins, nuclear transport receptors (NTR)Leptomycin B (LMB)

D Data analysis. See Image analysis/processingMass spectrometry (MS) Depleted nuclear extracts GST-RanGDP-coated GSH-Sepharose, 369, 370 NTRs and RCC1 depletion, 366, 368–370 Depletion. See Immuno-depletion; Genetic tools/ screens; RNA intererence (RNAi); Yeast conditional/mutant strains Dextrans. See Fluorescent reporters Digitonin-permeabilized cells (semi-permeabilized cells, mammalian) in vitro system to study NEBD, 24, 256–275, 264f in vitro transport assays, 332–334, 347–348 large scale in vitro import assay and SILAC-Tp, 365, 365f, 366, 367–368, 369–371, 372, 373–375, 374f normal vs. heat shock condition, 334, 334f, 335f, 342–346 for Nups localization, 8 permeabilized mitotic cells, 337f, 348 subcellular fractionation, cytosol-enriched fraction, 361 DNA-binding domain (DBD), 419–420, 421f See also Yeast three-hybrid interaction screen Dronpa (photoswitchable FP), 14, 225–228, 227f, 234 Drugs, chemical inhibitors aphidicolin, 222, 246–251 BAPTA, 45, 182, 183f, 184–185, 185f, 186, 187 CDK1 inhibitor RO3306, 24, 269 cytostatic factor (CSF), 167, 171 latrunculin A, 293–295 leptomycin B (LMB) (see Leptomycin B) nocodazole, 50–51, 222, 248, 251, 260–261, 293–295, 294f, 296, 297f PLK1 inhibitor BI2536, 269 proteasome inhibitor, 293–295 roscovitine, 246, 249, 251 thymidine, 222, 250 Dynein light-chain2 (Dyn2) dimerization-Zipper, 9, 100–114 Dyn2 interaction domain (DID) in Nup159, 100, 101f eDID (artificially engineered Dyn2 interaction domain), 101–106, 102s, 103s, 104s

negative stain (of Nups and complexes, EM), 109–111, 107–111, 109s, 110f probing the FG-network in yeast, 111–113, 112f Dynein light-chain-interaction domain (DID). See Dynein light-chain2 (Dyn2) dimerization-Zipper

E E. coli. See Recombinant protein production and purification Eggs from frogs. See Xenopus egg extracts Electron microscopy (EM). See also Immunogold labeling/immuno-EMTransmission electron microscopy (TEM); Scanning electron microscopy (SEM) cryoelectron tomogram (cryoET) of NPCs and NE, 5f, 6, 7f, 43, 129f Dyn2 as an EM label to map subcomplexes and single Nups, 9, 107–111, 109f, 110f historical overview, 3–6, 5f, 12–13, 43 negative stain (of Nups and complexes), 100–101, 101f, 108–111 See also Immunogold labeling/Immuno-EM, Transmission Electron Microscopy (TEM), Scanning Electron Microscopy (SEM) EM. See Electron microscopy (EM) Endoplasmic reticulum (ER), 4f, 19, 26–27, 119–120, 124, 171, 182, 214, 215f, 223f, 224, 225, 256, 302f, 361, 465, 467f, 468–469 inner nuclear membrane (INM) protein synthesis, 19, 215f signal recognition particle receptor, beta subunit (Srprb) imaging, 224 Epitope pulse-chase labeling (NPC biogenesis, ribosome maturation/export), 24–27, 148–161 description, mRNA organization, 148–149, 150f immunoprecipitation, 154–156 indirect immunofluorescence, 156–157, 158f subcellular fractions, 157–160 tetracycline-dependent RNA aptamer, 149 tracking newly synthesized proteins, western blot analysis, 151–153, 157–160 Epitope-tagged nucleoporins immuno-EM localization of epitope-tagged FG-nucleoporins, 93–96 (see also Fluorescent reporters) integrating the eDID sequence within a yeast ORF, 103–104, 112 pFa6a-eDID-Flag-PLox-HisMX4-PLox construct (yeast), 103, 104f ER. See Endoplasmic reticulum (ER)

Index

Exportins. See Karyopherins (Kaps), importins/ exportins/transportins, nuclear transport receptors (NTR)

F FG repeat domain, FG-Nups antibodies, mAb414, 49t, 170, 181, 184–186, 292t FG repeat production and purification, 381–384 immuno-EM localization of epitope-tagged FG-nucleoporins, 93–96 introduction, 11–12, 21–23 probing the FG-network in yeast, 111–113 use in NPC mimics, 381 Field emission scanning electron microscopy (FESEM). See Scanning electron microscopy (SEM) FISH. See Fluorescence in situ hybridization (FISH) Fluorescence in situ hybridization (FISH) detection of endogenous poly(A)þRNA in mammalian cells, 407–409, 408f protocol for mammalian cells (tRNA), 429–430 protocol in S. cerevisiae, 427–429 40S pre-ribosome localization in S. cerevisiae, 453, 454f reagents for in situ hybridization experiment in mammalian cells, 400–401 tRNA cellular distribution (yeast, human), 426–431, 426f Fluorescence microscopy. See Fluorescent reporters; Immunofluorescence (IF); Live imaging Fluorescently labeled RNAs, 398, 405–407, 411 Fluorescent reporters C. elegans, useful fluorescent markers, 290, 291t Dronpa (photoswitchable FP), 225–226 fluorescently labeled dextran, 13, 262–263, 264, 264f, 268f, 272f, 297f, 298 generation of cell lines expressing a fluorescent NE marker (HeLa), 258–260 genomically integrated transport reporters (yeast), 316–317 GFP-LacI, 470–471, 472f GFP-Nups plasmids and mammalian cell lines, pIBB-DiHcRed, pSrprb-ECFP, 235 GFP-tagged nuclear transport and reporters (yeast), 314–315 His6-GFP-Hsc70, 334, 335f, 346 IbB-GFP, fluorescent cargoes (nanopore studies), 383–384 Nup133-Venus/Venus-Nup107-expressing cells, 242f, 243f, 244f, 245, 247, 248, 249–252 Rev(48–116)-GFP2-M9 (HeLa cells), 360–361

TetR-GFP and RFP-LacI, 465–473, 466s, 467f, 472f, 481–482 2xGFP-Nup58, 257, 259–260, 264–265, 264f, 267, 268f Flux measurements (nanopores), 387–389, 388f Forward and reverse genetics. See Genetic tools/ screens Frogs. See Xenopus laevis

G Gene-nuclear pore interactions. See also Chromatin chromatin IP (ChIP) in yeast and human cells, 474–481 gene localization to the NPC in yeast, live cells, 465–473, 466s, 467f, 472f gene-NPC interactions in C. elegans, 303–304 Genetic tools/screens, 6, 13–14, 16–17, 26–27. See also RNA interference (RNAi); Yeast conditional/mutant strains C. elegans nucleoporins and transport factor mutants alleles, 279, 280t CRISPR-Cas9 system, 279–285, 289–290, 304 genetics screens in yeast, 13–14 transgenesis in C. elegans, 287–289 GFP. See Green fluorescent protein (GFP) GFP mRNA, mammalian cells, hybridization, probe detection, 410–411 Glutathione-S-transferase (GST). See Recombinant Protein production and purification Green fluorescent protein (GFP). See Fluorescent reporters, Live imaging

H HeLa cells, HeLa Kyoto (HeLaK), HeLa-S3. See mammalian cells Heterokaryons assays cell-fusion assays in mammalian cells, history, Sendai virus, PEG, 243–245 kar1-1, shuttling assay in yeast, 454–456, 455f interphase NPC formation, 243–250, 242s, 243f, 244f

I IF. See Immunofluorescence (IF) Image analysis/processing automated MatLab-based algorithm, 257 basic image processing of negative staining particles, 111 computational image analysis and extraction of equilibrium rate constants (NE permeability, Dronpa), 226–228, 227f GFP-LacI localization, data analysis, 469–471

489

490

Index

Image analysis/processing (Continued) immuno-EM in Xenopus, statistical analysis, 93, 94f NEBD, in vitro assay, 270–273, 272f nuclear/cytoplasmic ratio in S. cerevisiae, 316f, 318–319 quantitation of nucleoporins assembly kinetics, 223f, 224–225 single-particle averaging approach, SR microscopy, 229 Imaging NPCs. See Electron microscopy; Image analysis/processing; Immunofluorescence Live imaging; Super-Resolution (SR) imaging/microscopy Immuno-depletion depletion reconstitution in proteoliposomes, transmembrane proteins, (Xenopus), 195–196, 207–210 importin-depletion using phenyl-Sepharose (HeLa S3), 335f, 338, 343, 344, 367, 368–369, 370 protein depletion and add-back (HeLa, NEBD assay), 269–270 Immunofluorescence (IF), 24–25, 65–66, 69–70, 90, 128, 129f, 156–157, 158f, 181, 208f, 211–212, 215f, 216, 244f, 250, 298–303, 337f, 348, 364f, 465, 473. See also Antibodies Super-Resolution (SR) imaging/microscopy confocal microscopy, 8, 381, 469, 471 detection of epitope pulse-chased yeast proteins, 156–157 IF and super resolution imaging in C. elegans, 299–301 individual NPCs detection, super resolution microscopy, 299–301, 300f Immunogold labeling/Immuno EM colloidal gold-conjugated antibodies, 49, 66, 69, 89–90 immuno-SEM on yeast nuclei, 66, 68–69 on in vitro assembled and mammalian cell nuclei, 52–54 on thin-sections of isolated Xenopus oocyte nuclei, 82–96, 89f, 94f on yeast cryosections, 69–76, 70f Immunoprecipitation. See Affinity-purification Importins (nuclear import factor/receptor). See Karyopherins (Kaps), importins/ exportins/transportins, nuclear transport receptors (NTR) Inner nuclear membrane (INM) proteins passage through the NPC, in vitro assay, Xenopus, 196, 214–216, 215f targeting of INM proteins, 19

Internal ribosome entry site (IRES), 258–259 In vitro assays, 19, 24, 257, 264f, 270, 273 In vitro import assay/reaction introduction, 332–342 large scale in vitro import assay for SILAC-Tp, 370–372 normal vs. heat shock condition, 334f, 342–346 In vitro mitotic chromosome-loading reaction, 336f, 346–348 In vitro nuclear reconstitution assays. See Xenopus In vitro system, NE breakdown. See Nuclear envelope breakdown (NEBD) In vivo tRNA export assay (amber suppression), 423–425, 424f IRES. See Internal ribosome entry site (IRES) Iridium coating, 55, 56f

K Karyopherins (Kaps), importins/exportins/ transportins, nuclear transport receptors (NTR)16–17, 18f, 20, 312, 314. See also Digitonin-permeabilized cells (semi-permeabilized cells, mammalian); In vitro import assay/reaction; Leptomycin B (LMB); Nuclear transport signals; Yeast regulated nuclear transport CRM1-dependent export cargoes, 357–364, 364f, 375 Crm1/Xpo1 and ribosomal export, 20, 440 nuclear transport receptor (NTR) substrate identification, 354–375 production and purification, 340–341t, 367–368, 383–384, 390 tRNA transport factors, 416–419, 418f U snRNA transport factors, 396–398, 397f Kinetochores. See also Mitosis, meiosis in C. elegans, 28, 208t, 294f nuclear pores, nuclear transport machinery and kinetochores, 16–17

L Leptomycin B (LMB) CRM1-dependent export cargoes, 357–364, 364f, 375 drugs used in C. elegans, 293–295 introduction, 16–17, 357–358 S. cerevisiae LMB-sensitive mutant, 320, 322–323 Live imaging analysis of shuttling factors by yeast heterokaryon assays, 454–456, 455f de novo NPC assembly, cell-fusion, 243–250, 242s, 243f, 244f

Index

de novo NPC assembly, photobleaching approach, 241, 242f, 250–252, 251f flux measurements in NPC mimics, 385–389, 388f gene localization to the NPC in yeast, 65–473, 466s, 467f, 472f in vitro NEBD assay, 264–267, 268f, 272f kinetics of postmitotic NPC assembly by multicolor 4D imaging, 221–223, 223f live imaging in C. elegans embryos, 290–298, 291t, 294f, 297f localization of Kap cargo proteins in yeast, 314–318, 316f, 324f monitoring NE permeability by sequential photoswitching, 225–228, 227f transport competence of assembling NPCs, 221–223, 223f LMB. See Leptomycin B (LMB)

M mAb414 C. elegans, 292t, 298–299, 300f, 301 Xenopus, 170, 181, 184–185, 185f, 186 Mammalian cells. See also Digitonin-permeabilized cells (semi-permeabilized cells, mammalian) identification of nuclear transport receptor substrates (HeLa, HeLa-S3), 354–375, 359f, 363f, 364f, 365f interphase NPC formation (HeLa), 240–252, 242f, 243f, 244f, 251f in vitro system to study NEBD (HeLa), 256–274, 258f, 264f, 268f, 272f isolation of nuclei and NE (HeLaK, Hek293, K562, RKO), 119–130, 122t, 142f, 125f, 129f NPC assembly, stucture and function (HeLa, NRK, U2OS), 220–235, 223f, 227f, 232f nuclear organization, transcription, chromatin structure, ChIP, 474, 479–481 nucleocytoplasmic transport, digitoninpermeabilized cells (interphase, mitosis, heat-shock; HeLa-S3), 332–349, 337f RNA transport (HeLa), 396–398, 407–411, 408f SEM on isolated nuclei (adherent cells), 51–52, 53f tRNA detection and aminoacylation (HeLa), 426, 426f, 429–430, 431–434, 434f tRNA in vivo export assay (COS7 cells), 423–425 targeted proteomics, 130–142, 141f, 143f Mass spectrometry (MS) MS following affinity purification, 154, 453 selected reaction monitoring (SRM) assays/ targeted proteomics, 130–143, 131f, 135t, 136, 138f

absolute/relative protein quantifications, 10, 118–119, 139–141, 141f, 141–142, 143f protein modification, 139 PTPs (peptides unique to proteins), selection and development, 130–132, 131f, 134, 136, 137, 138f, 139 SRM data processing, mProphet, 133, 137, 139–142 stable isotope labeling of amino acids in cells (SILAC), 148–149 data analysis, 362–363, 363f, 373–375, 374f SILAC-based in vitro transport (SILAC-Tp) assay/identification of nuclear import cargoes, 365f, 365–375, 374f SILAC identification of CRM1-dependent export cargos, 357–363, 359f, 363f Mitosis, meiosis. See also Cell cycle CSF-arrested Xenopus egg extract, 260–261 mitotic HeLa cell extract (HeLA S3), 260–261 permeabilized mitotic cells, 348 Mutant strains. See Yeast conditional/mutant strains

N Nanopores. See NPC mimics NEBD. See Nuclear envelope breakdown (NEBD) Negative stain (of Nups and complexes). See Electron microscopy (EM) NPCs. See Nuclear pore complexes (NPCs) NPPs. See Nuclear pore proteins (NPPs), C. elegans NTF. See Nuclear transport factor (NTF) NTRs. See Karyopherins (Kaps), importins/ exportins/transportins, nuclear transport receptors (NTR) Nuclear/cytoplasmic fractionation. See Subcellular fractionation Nuclear envelope (NE) integrity/permeability. See also nuclear envelope breakdown (NEBD) in C.elegans, 14, 225–228, 279–287, 290–304 dextrans, 182–184, 183f, 257, 266, 268f, 271–273 photoswitching, 14, 225–228, 231 in Xenopus (NE integrity), 25, 100, 168, 182, 183f, 241, 257 Nuclear envelope (NE) isolation/purification, 119–130, 129f Nuclear envelope breakdown (NEBD) fluorescent dextran, 257, 262–263, 266, 271, 297f GFP-tagged NE marker, HeLa cells, 13–14, 24, 26–27, 112, 223f, 227f, 229, 257, 264f, 296–298, 314–315 in vitro system to study NEBD, 24, 257, 258f, 264, 264f, 265, 268f, 269, 273 mitotic HeLa cell extract, 260–261, 264, 267, 268f

491

492

Index

Nuclear envelope breakdown (NEBD) (Continued) nucleoporin mutants, 267, 294f semi-permeabilization, cells, 24, 257, 264, 264f, 265–266, 267, 268f, 269, 349 Nuclear export, 12–13, 15, 20, 23, 27–28, 82–83, 312, 314–315, 332–333, 354–356, 361, 370–371, 380, 396, 397f, 398, 401–405, 402f, 407–411, 418f, 423–425, 424f, 440–441, 442f, 443, 453–454, 454f Nuclear export signals (NESs). See Nuclear transport signals Nuclear import, 2–3, 16, 17–19, 18f, 23, 82–83, 166, 170, 184, 185f, 214, 224, 290, 298, 312, 314–315, 316f, 320, 323, 332–333, 334–335, 334f, 335f, 336f, 337f, 342–343, 346–347, 348, 349, 354–355, 357, 364–375, 380, 383 Nuclear isolation. See Subcellular fractionation Nuclear localization signals (NLSs). See Nuclear transport signals Nuclear pore complexes (NPCs). See also Nucleoporins assembly/disassembly/biogenesis de novo/interphase NPC assembly, 26–27, 240–252, 242f, 243f, 244f, 251f NPC disassembly, 24, 234, 257, 264f, 294f, 349. See also Nuclear Envelope Breakdown (NEBD) post-mitotic NPC assembly, 24–26, 221–225, 223f, 234, 240–241, 290, 294f mimics, 14–15, 380–391, 386f, 388f Nuclear pore-free intermediates, 25, 26, 44–45, 168, 184–185, 185f, 186, 187 Nuclear pore proteins (NPPs), C. elegans nucleoporins, 278–279, 280t, 291t, 294f Nuclear reconstitution assay. See Xenopus laevis, egg extracts; In vitro assays Nuclear transport factor (NTF). See also Karyopherins (Kaps), importins/ exportins/transportins, nuclear transport receptors (NTRs); Ran GTPase, Ran cycle, Ran-associated factors mRNA export receptor TAP/NXF1-p15, U snRNA transport factors, 396–398, 397f NTF2, 17, 18f, 339, 341t, 355, 367, 372, 382–390, 388f nuclear tRNA transport factors, 416–419, 418f trans-acting factors in the ribosome biogenesis pathway, 439–441, 440f Nuclear transport receptors (NTRs). See Karyopherins (Kaps), importins/ exportins/transportins, nuclear transport receptors (NTR)

Nuclear transport signals (NLSs, NESs), 3, 13–14, 15, 16–17, 18f, 19, 215f, 312, 313t, 314–315, 354–355, 375 nuclear transport receptor (NTR) substrate identification, 312, 354–375 Nucleoporins (Nups), 2–12, 23–31 See also Antibodies; FG repeat domain, FG-Nups; Imaging NPCs, Nuclear pore proteins (NPPs); Pore membrane proteins (Poms); Stoichiometry NusA-TEV. See Tobacco etch virus (TEV)

P PEG. See Polyethylene glycol (PEG), cell fusion, heterokaryons Permeability, 12, 14–15, 24, 42, 129, 225–228, 234, 256, 257, 262, 265, 266, 268f, 271–273, 372, 387 Phenylalanine-glycine (FG) repeat. See FG repeat domain, FG-Nups Phenyl-Sepharose column, 335f, 338, 343–344, 367, 368–369, 370 Photobleaching, photoactivation, photoswitching. See Live imaging Polyethylene glycol (PEG), cell fusion, heterokaryons, 242f, 245, 247 Pore membrane proteins (Poms)/integral membrane nucleoporins, 5f, 8, 11, 19, 25, 26–27 functional studies in Xenopus, immunodepletion and add-back, Ndc1, gp210, Pom121, 195–196, 208f interphase NPC formation (Pom121), 240–241, 244f, 246, 249–250 post-mitotic NPC assembly (Pom121), 223f, 224, 225 Posttranslational protein modifications, epitope pulse-chase labeling in yeast, 149, 151–153 Protein complex stoichiometry. See Stoichiometry Protein depletion. See Immuno-depletion; Genetic tools/screens; RNA intererence (RNAi); Yeast conditional/mutant strains Protein purification. See Recombinant protein production and purification; Affinity purification Proteoliposomes, 209–210, 214, 215f, 216 Pulse-chase labeling. See Epitope pulse-chase labeling

R Ran GTPase, Ran cycle, Ran-associated factors C. elegans Ran-associated factors, 284t, 298–299 GST-RanGDP-coated GSH-Sepharose (RCC1 depletion), 369, 370

Index

importin/Ran system in interphase and mitosis, 333, 334–335, 336f, 347 introduction, nucleocytoplasmic transport, 17–19, 18f NTF2, 17, 18f, 339, 341t, 355, 367, 372, 382–390, 388f purification, 333, 339–342 Ran-GTP, 332–333, 336f, 380 a/b-Ran system, 336f unloading of the tRNA cargo, 419 Recombinant protein production and purification Dyn2, DID (Dyn2 interaction domain), eDID, proteins modified with eDID, 104–107 FG-nups, 381–383 GST-FLAG-hKid, 339, 347 GST-RanGDP, 369 His6-GFP-HSC70, 345 IbB-eGFP-His6, 383–384, 390 integral membrane proteins, 201–202 NTF2-GST, Kap95-GST, Kap95, Kap121-GST, 383–384, 390 nuclear transport factors/receptors (vertebrates), 339–342, 340–341t, 367–368 NusA-TEV, 203–204 reconstitution in proteoliposomes, 209–210 Ribosomal subunits biogenesis and export in budding yeast, 438–456 current knowledge and model, 438–441, 440f isolation of preribosomal particle by TAP, 439, 450–453 localization of 40S preribosome by FISH, 453–454 localization of preribosomal subunits by fluorescence microscopy, 441–443, 442f polysome profiles, sucrose gradient sedimentation, 442f, 443–450 shuttling of nucleolar/nuclear trans-acting factors, 455–456 study using epitope pulse–chase labeling, 154f, 154–156, 158f, 159f Ribosome Internal ribosome entry site (IRES), 258–259 Ribosome removal from isolated yeast nuclei (for FESEM), 68 RNA. See also Fluorescence in situ hybridization; Ribosomal subunits biogenesis and export; tRNA cap binding complex (CBC), 396, 397f control of gene expression, transcription, 29–30, 411 digoxigenin-labeled GFP mRNA probe, in situ hybridization, 400–401, 408f, 409–411

electrophoresis (PAGE), 401, 402f, 403–405, 406f, 409 endogenous poly(A)þRNA, mammalian cells, 407–409 export of radiolabeled RNAs in Xenopus oocytes, 401–405 export pathways, 4f, 18f, 20–21, 23, 112f, 396–398, 397f fractionation and isolation, mammalian cells, 432 fractionation and isolation, S. cerevisiae, 431–432 GFP mRNA, mammalian cells, 400, 401, 408f, 409–411 in vitro transcription, 398, 401, 402–403, 405, 406f localization of fluorescently labeled RNAs, Xenopus oocytes, 398, 406f, 405–407, 411 tetracycline-dependent RNA aptamer, (epitope pulse-chase labeling), 149, 150f U snRNAs, nuclear export and re-import, 396–398, 397f, 401, 402f Xenopus oocyte microinjection experiment, 13, 82–83, 398–399, 401, 402f, 403–404, 411 RNA-FISH. See Fluorescence in situ hybridization (FISH) RNAi. See RNA interference (RNAi) RNA interference (RNAi) C. elegans, reverse genetic screens, feeding method, 279–287 C. elegans, RNAi efficiency, 280–282t Nup–GFP transfection combined with siRNA treatment (U2-OS cells), 229–230 protein knockdown in HeLa cells (NEBD assay), 267–269 protein knockdown, siRNA treatment, (cell fusion assays), 241, 244f, 249–250

S Saccharomyces cerevisiae (budding yeast). See Yeast (Sacharomyces cerevisiae, budding yeast) Scanning electron microscopy (SEM), 41–44, 46f, 54–55 combined with immunogold labeling, 52–54, 66, 68–69 iridium coating technique, 55, 56f on anchored in vitro assembled nuclei (Xenopus), 49–51, 52f on mammalian cell nuclei, 51–52, 53f SEM and immuno-SEM on yeast nuclei, 65–69 Selected reaction monitoring (SRM) assays. See Mass spectrometry SEM. See Scanning electron microscopy (SEM)

493

494

Index

Semi-permeabilized cells. See Digitonin-permeabilized cells SILAC-based in vitro transport (SILAC-Tp) system, 365–375, 365f, 374f SR. See Super-resolution (SR) imaging/microscopy Stable isotope labeling of amino acids in cells (SILAC). See Mass spectrometry Stochastic optical reconstruction microscopy (STORM). See Super-resolution (SR) imaging/microscopy Stoichiometry absolute quantification, nucleoporins, 10, 118–119, 139–141, 141f data processing, mProphet, 133, 137, 139–142 native protein complexes, NPCs, 118–119, 141f, 143f relative quantifications across multiple samples, 141–142, 143f targeted proteomics/ selected reaction monitoring (SRM), 130–143, 131f, 135t, 138f STORM. See Stochastic optical reconstruction microscopy (STORM) Subcellular fractionation cytosol-enriched fraction (HeLa), 361 importin-depleted/containing fractions (HeLa S3), 335f, 343, 344, 346 importin-depleted cytosol using phenylSepharose (HeLa S3), 368–369 isolation of nuclei and nuclear envelopes from mammalian cells in culture, 119–130 HeLaK cells, 121–125, 122t, 124f, 125f, 126, 128, 129f nuclear envelope isolation, 127–128 protocols for other cell lines, 122t, 125–127, 125f, 126t quality controls, 128–130, 138f polysome preserving conditions, 446 RCC1-depleted nuclear extract (HeLa S3), 369–370 subcellular fractionation of yeast spheroplasts (crude nuclear fraction), 157–160, 159f sucrose gradient fractionation (ribosomes), 442f, 443–450 Super-Resolution (SR) imaging/microscopy advances in the field of SR light microscopy, 228–229 C.elegans indivudual NPCs visualized by SR-SIM, 299–302, 300f determination of radial position of a fluorescent marker, 229, 232f, 233–234 single-particle averaging approach, 229, 234 stochastic optical reconstruction microscopy (STORM) imaging, 228, 232

super-resolution imaging of human NPCs using STORM, 232f super-resolution structured illumination microscopy (SR-SIM), 299–302, 300f

T Tandem affinity purification (TAP). See Affinity-purification Targeted proteomics. See Mass spectrometry (MS); Stoichiometry TEM. See Transmission electron microscopy (TEM) TEV. See Tobacco etch virus (TEV) Time-lapse observations. See Live imaging Tobacco etch virus (TEV) GST-TEV-Flag, split tag purification, 103f, 106, 108, 109f, 114 NusA-TEV, INM protein localization, 203–204, 214–216, 215f ProtA-TEV-Cbp-Flag, Flag-TEV-ProtA, Pulse–Chase epitope labeling, 150f, 151f, 156, 161 Tandem Affinity purification (TAP), 452–453, 457 Transfection Ca-phosphate method (HeLa P4 cells), 360–361 Nup-GFP transfection (Fugene6) and siRNA treatment (U2-OS cells), 221–222, 225, 229, 230, 232f, 249–250 Transgenesis (in C. elegans), 287–290, 288t Transmembrane proteins/integral membrane proteins. See Inner nuclear membrane (INM) proteins; Pore membrane proteins (Poms) Transmission electron microscopy (TEM) conventional TEM in S. cerevisiae, 60–61, 62f, 112f gold nanoparticles, 69–70 TEM in C. elegans, 302–303, 302f TEM of nuclear assembly reactions, 212–214 TEM on Xenopus oocytes, 82–85, 84f Transport channel. See FG repeat domain, FG-Nups Transport kinetics, 221–225, 223f, 227f, 380–391 tRNA amber suppressor tRNA (pulse chase labeling), 148, 150f amber supression in vivo tRNA export assay, 423–425, 424f FISH detection of tRNA localization (yeast, human), 426–430, 426f identification of tRNA interacting proteins (yeast3-hybrid), 419–422, 421f

Index

introduction, nuclear-cytoplasmic tRNA dynamic, 416–419, 418f tRNA aminoacylation status, 417–418, 418f, 431–435, 434f

U UAA. See Unnatural amino acid (UAA) Unnatural amino acid (UAA), 149, 162 Uridine-rich small nuclear RNAs (U snRNAs), 16, 20, 396–398, 397f, 402f U snRNAs. See Uridine-rich small nuclear RNAs (U snRNAs)

V Vesicle-vesicle fusion, 182, 183f

X Xenopus laevis egg extracts anchored nuclei, assembly reaction, SEM, 50–51, 52f depletion of transmembrane proteins, 207–209, 208f EM studies on reconstituted nuclei, 96, 170, 211–212, 257 INM targeting assays, 196, 214–216, 215f NPC assembly, 24–27 nuclear in vitro nuclear reconstitution assays, 166–168, 180–184, 183f, 187, 210–212 pore-free nuclear intermediates, 184–187, 185f preparation and fractionation, 172–177, 174f, 195–200, 198f use for in vitro transport assays, 13, 16, 333 demembranted sperm chromatin decondensation, 50, 210–211 preparation, 177–180 frog housing conditions, egg laying, 169, 171–172 oocytes dissection, nuclei isolation, 90–91, 402f, 404, 406f EM and immuno-EM studies, 5f, 9, 12–13, 81–96, 84f, 88f, 94f export of radiolabeled RNAs, 401–405, 402f localization of fluorescently labeled RNAs, 398, 405–407, 406f, 411 microinjection experiments, 13, 82–83, 85, 87, 88f, 95, 398–399, 401, 402f, 403–404, 411 RNA transport studies, 13, 23, 396–411

Y Yeast (Sacharomyces cerevisiae, budding yeast)3–6, 7f, 8, 9, 10, 12–14, 16–17, 19, 20, 29–30. See also Cell cycle in yeast, Fluorescence in situ hybridization (FISH) analysis of shuttling factors by yeast heterokaryon assays, 454–456, 455f electron microscopy, 59–79, 70f epitope pulse-chase labeling (NPC biogenesis, ribosome maturation/export), 148–161 gene–nuclear pore interactions, ChIP, 474–481, 475t genetics screens in yeast, 13–14 nuclear–cytoplasmic tRNA Dynamics, 416–435, 424f nuclear transport signals (NLSs, NESs), 312–328 probing the FG-network in yeast, 111–113 regulated nuclear transport, nuclear/cytoplasmic ratio, 311–328, 316f, 324f ribosome assembly and transport, 438–458 RNA fractionation and isolation, 431–432 subnuclear organization, GFP-lacI, 465–474, 466f, 467f, 472f TAP in budding yeast, 102, 149, 154f, 439, 450–453, 474 Yeast conditional/mutant strains epitope pulse-chase vectors (galactose-inducible, doxycycline-inducible), 153, 156–157 MET3 promoter to shutoff gene expression, 322 temperature-sensitive (ts), 320–321 xpo1T539C, LMB sensitive, 322–323 Yeast culture, 73, 327–328, 456–458, 469 Yeast epitope tagging cloning into epitope pulse-chase vector, 150–151, 151f cloning of eDID to the N-/C-terminal end of a target protein, 102–103, 103f genomically integrated transport reporters, 316–317 GFP-tagged nuclear transport and reporters, 314–315 integrating the eDID sequence within a yeast ORF, 103–104, 104f Yeast spheroplast preparation, 63, 64 isolation of nuclei from spheroplasts, 66, 67 subcellular fractionation (crude nuclear fraction), 157–160, 159f sucrose gradient fractionation (ribosomes), 442f, 443–450 Yeast three-hybrid interaction screen, 419–422, 421f

495

Volumes in Series Founding Series Editor DAVID M. PRESCOTT Volume 1 (1964) Methods in Cell Physiology Edited by David M. Prescott Volume 2 (1966) Methods in Cell Physiology Edited by David M. Prescott Volume 3 (1968) Methods in Cell Physiology Edited by David M. Prescott Volume 4 (1970) Methods in Cell Physiology Edited by David M. Prescott Volume 5 (1972) Methods in Cell Physiology Edited by David M. Prescott Volume 6 (1973) Methods in Cell Physiology Edited by David M. Prescott Volume 7 (1973) Methods in Cell Biology Edited by David M. Prescott Volume 8 (1974) Methods in Cell Biology Edited by David M. Prescott Volume 9 (1975) Methods in Cell Biology Edited by David M. Prescott Volume 10 (1975) Methods in Cell Biology Edited by David M. Prescott

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498

Volumes in Series

Volume 11 (1975) Yeast Cells Edited by David M. Prescott Volume 12 (1975) Yeast Cells Edited by David M. Prescott Volume 13 (1976) Methods in Cell Biology Edited by David M. Prescott Volume 14 (1976) Methods in Cell Biology Edited by David M. Prescott Volume 15 (1977) Methods in Cell Biology Edited by David M. Prescott Volume 16 (1977) Chromatin and Chromosomal Protein Research I Edited by Gary Stein, Janet Stein, and Lewis J. Kleinsmith Volume 17 (1978) Chromatin and Chromosomal Protein Research II Edited by Gary Stein, Janet Stein, and Lewis J. Kleinsmith Volume 18 (1978) Chromatin and Chromosomal Protein Research III Edited by Gary Stein, Janet Stein, and Lewis J. Kleinsmith Volume 19 (1978) Chromatin and Chromosomal Protein Research IV Edited by Gary Stein, Janet Stein, and Lewis J. Kleinsmith Volume 20 (1978) Methods in Cell Biology Edited by David M. Prescott

Advisory Board Chairman KEITH R. PORTER Volume 21A (1980) Normal Human Tissue and Cell Culture, Part A: Respiratory, Cardiovascular, and Integumentary Systems Edited by Curtis C. Harris, Benjamin F. Trump, and Gary D. Stoner

Volumes in Series

Volume 21B (1980) Normal Human Tissue and Cell Culture, Part B: Endocrine, Urogenital, and Gastrointestinal Systems Edited by Curtis C. Harris, Benjamin F. Trump, and Gray D. Stoner Volume 22 (1981) Three-Dimensional Ultrastructure in Biology Edited by James N. Turner Volume 23 (1981) Basic Mechanisms of Cellular Secretion Edited by Arthur R. Hand and Constance Oliver Volume 24 (1982) The Cytoskeleton, Part A: Cytoskeletal Proteins, Isolation and Characterization Edited by Leslie Wilson Volume 25 (1982) The Cytoskeleton, Part B: Biological Systems and In Vitro Models Edited by Leslie Wilson Volume 26 (1982) Prenatal Diagnosis: Cell Biological Approaches Edited by Samuel A. Latt and Gretchen J. Darlington

Series Editor LESLIE WILSON Volume 27 (1986) Echinoderm Gametes and Embryos Edited by Thomas E. Schroeder Volume 28 (1987) Dictyostelium discoideum: Molecular Approaches to Cell Biology Edited by James A. Spudich Volume 29 (1989) Fluorescence Microscopy of Living Cells in Culture, Part A: Fluorescent Analogs, Labeling Cells, and Basic Microscopy Edited by Yu-Li Wang and D. Lansing Taylor

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500

Volumes in Series

Volume 30 (1989) Fluorescence Microscopy of Living Cells in Culture, Part B: Quantitative Fluorescence Microscopy—Imaging and Spectroscopy Edited by D. Lansing Taylor and Yu-Li Wang Volume 31 (1989) Vesicular Transport, Part A Edited by Alan M. Tartakoff Volume 32 (1989) Vesicular Transport, Part B Edited by Alan M. Tartakoff Volume 33 (1990) Flow Cytometry Edited by Zbigniew Darzynkiewicz and Harry A. Crissman Volume 34 (1991) Vectorial Transport of Proteins into and across Membranes Edited by Alan M. Tartakoff Selected from Volumes 31, 32, and 34 (1991) Laboratory Methods for Vesicular and Vectorial Transport Edited by Alan M. Tartakoff Volume 35 (1991) Functional Organization of the Nucleus: A Laboratory Guide Edited by Barbara A. Hamkalo and Sarah C. R. Elgin Volume 36 (1991) Xenopus laevis: Practical Uses in Cell and Molecular Biology Edited by Brian K. Kay and H. Benjamin Peng

Series Editors LESLIE WILSON AND PAUL MATSUDAIRA Volume 37 (1993) Antibodies in Cell Biology Edited by David J. Asai Volume 38 (1993) Cell Biological Applications of Confocal Microscopy Edited by Brian Matsumoto

Volumes in Series

Volume 39 (1993) Motility Assays for Motor Proteins Edited by Jonathan M. Scholey Volume 40 (1994) A Practical Guide to the Study of Calcium in Living Cells Edited by Richard Nuccitelli Volume 41 (1994) Flow Cytometry, Second Edition, Part A Edited by Zbigniew Darzynkiewicz, J. Paul Robinson, and Harry A. Crissman Volume 42 (1994) Flow Cytometry, Second Edition, Part B Edited by Zbigniew Darzynkiewicz, J. Paul Robinson, and Harry A. Crissman Volume 43 (1994) Protein Expression in Animal Cells Edited by Michael G. Roth Volume 44 (1994) Drosophila melanogaster: Practical Uses in Cell and Molecular Biology Edited by Lawrence S. B. Goldstein and Eric A. Fyrberg Volume 45 (1994) Microbes as Tools for Cell Biology Edited by David G. Russell Volume 46 (1995) Cell Death Edited by Lawrence M. Schwartz and Barbara A. Osborne Volume 47 (1995) Cilia and Flagella Edited by William Dentler and George Witman Volume 48 (1995) Caenorhabditis elegans: Modern Biological Analysis of an Organism Edited by Henry F. Epstein and Diane C. Shakes Volume 49 (1995) Methods in Plant Cell Biology, Part A Edited by David W. Galbraith, Hans J. Bohnert, and Don P. Bourque

501

502

Volumes in Series

Volume 50 (1995) Methods in Plant Cell Biology, Part B Edited by David W. Galbraith, Don P. Bourque, and Hans J. Bohnert Volume 51 (1996) Methods in Avian Embryology Edited by Marianne Bronner-Fraser Volume 52 (1997) Methods in Muscle Biology Edited by Charles P. Emerson, Jr. and H. Lee Sweeney Volume 53 (1997) Nuclear Structure and Function Edited by Miguel Berrios Volume 54 (1997) Cumulative Index Volume 55 (1997) Laser Tweezers in Cell Biology Edited by Michael P. Sheetz Volume 56 (1998) Video Microscopy Edited by Greenfield Sluder and David E. Wolf Volume 57 (1998) Animal Cell Culture Methods Edited by Jennie P. Mather and David Barnes Volume 58 (1998) Green Fluorescent Protein Edited by Kevin F. Sullivan and Steve A. Kay Volume 59 (1998) The Zebrafish: Biology Edited by H. William Detrich III, Monte Westerfield, and Leonard I. Zon Volume 60 (1998) The Zebrafish: Genetics and Genomics Edited by H. William Detrich III, Monte Westerfield, and Leonard I. Zon Volume 61 (1998) Mitosis and Meiosis Edited by Conly L. Rieder

Volumes in Series

Volume 62 (1999) Tetrahymena thermophila Edited by David J. Asai and James D. Forney Volume 63 (2000) Cytometry, Third Edition, Part A Edited by Zbigniew Darzynkiewicz, J. Paul Robinson, and Harry Crissman Volume 64 (2000) Cytometry, Third Edition, Part B Edited by Zbigniew Darzynkiewicz, J. Paul Robinson, and Harry Crissman Volume 65 (2001) Mitochondria Edited by Liza A. Pon and Eric A. Schon Volume 66 (2001) Apoptosis Edited by Lawrence M. Schwartz and Jonathan D. Ashwell Volume 67 (2001) Centrosomes and Spindle Pole Bodies Edited by Robert E. Palazzo and Trisha N. Davis Volume 68 (2002) Atomic Force Microscopy in Cell Biology Edited by Bhanu P. Jena and J. K. Heinrich Ho¨rber Volume 69 (2002) Methods in Cell–Matrix Adhesion Edited by Josephine C. Adams Volume 70 (2002) Cell Biological Applications of Confocal Microscopy Edited by Brian Matsumoto Volume 71 (2003) Neurons: Methods and Applications for Cell Biologist Edited by Peter J. Hollenbeck and James R. Bamburg Volume 72 (2003) Digital Microscopy: A Second Edition of Video Microscopy Edited by Greenfield Sluder and David E. Wolf Volume 73 (2003) Cumulative Index

503

504

Volumes in Series

Volume 74 (2004) Development of Sea Urchins, Ascidians, and Other Invertebrate Deuterostomes: Experimental Approaches Edited by Charles A. Ettensohn, Gary M. Wessel, and Gregory A. Wray Volume 75 (2004) Cytometry, 4th Edition: New Developments Edited by Zbigniew Darzynkiewicz, Mario Roederer, and Hans Tanke Volume 76 (2004) The Zebrafish: Cellular and Developmental Biology Edited by H. William Detrich, III, Monte Westerfield, and Leonard I. Zon Volume 77 (2004) The Zebrafish: Genetics, Genomics, and Informatics Edited by William H. Detrich, III, Monte Westerfield, and Leonard I. Zon Volume 78 (2004) Intermediate Filament Cytoskeleton Edited by M. Bishr Omary and Pierre A. Coulombe Volume 79 (2007) Cellular Electron Microscopy Edited by J. Richard McIntosh Volume 80 (2007) Mitochondria, 2nd Edition Edited by Liza A. Pon and Eric A. Schon Volume 81 (2007) Digital Microscopy, 3rd Edition Edited by Greenfield Sluder and David E. Wolf Volume 82 (2007) Laser Manipulation of Cells and Tissues Edited by Michael W. Berns and Karl Otto Greulich Volume 83 (2007) Cell Mechanics Edited by Yu-Li Wang and Dennis E. Discher Volume 84 (2007) Biophysical Tools for Biologists, Volume One: In Vitro Techniques Edited by John J. Correia and H. William Detrich, III

Volumes in Series

Volume 85 (2008) Fluorescent Proteins Edited by Kevin F. Sullivan Volume 86 (2008) Stem Cell Culture Edited by Dr. Jennie P. Mather Volume 87 (2008) Avian Embryology, 2nd Edition Edited by Dr. Marianne Bronner-Fraser Volume 88 (2008) Introduction to Electron Microscopy for Biologists Edited by Prof. Terence D. Allen Volume 89 (2008) Biophysical Tools for Biologists, Volume Two: In Vivo Techniques Edited by Dr. John J. Correia and Dr. H. William Detrich, III Volume 90 (2008) Methods in Nano Cell Biology Edited by Bhanu P. Jena Volume 91 (2009) Cilia: Structure and Motility Edited by Stephen M. King and Gregory J. Pazour Volume 92 (2009) Cilia: Motors and Regulation Edited by Stephen M. King and Gregory J. Pazour Volume 93 (2009) Cilia: Model Organisms and Intraflagellar Transport Edited by Stephen M. King and Gregory J. Pazour Volume 94 (2009) Primary Cilia Edited by Roger D. Sloboda Volume 95 (2010) Microtubules, in vitro Edited by Leslie Wilson and John J. Correia Volume 96 (2010) Electron Microscopy of Model Systems Edited by Thomas Mu¨eller-Reichert

505

506

Volumes in Series

Volume 97 (2010) Microtubules: In Vivo Edited by Lynne Cassimeris and Phong Tran Volume 98 (2010) Nuclear Mechanics & Genome Regulation Edited by G.V. Shivashankar Volume 99 (2010) Calcium in Living Cells Edited by Michael Whitaker Volume 100 (2010) The Zebrafish: Cellular and Developmental Biology, Part A Edited by: H. William Detrich III, Monte Westerfield and Leonard I. Zon Volume 101 (2011) The Zebrafish: Cellular and Developmental Biology, Part B Edited by: H. William Detrich III, Monte Westerfield and Leonard I. Zon Volume 102 (2011) Recent Advances in Cytometry, Part A: Instrumentation, Methods Edited by Zbigniew Darzynkiewicz, Elena Holden, Alberto Orfao, William Telford and Donald Wlodkowic Volume 103 (2011) Recent Advances in Cytometry, Part B: Advances in Applications Edited by Zbigniew Darzynkiewicz, Elena Holden, Alberto Orfao, Alberto Orfao and Donald Wlodkowic Volume 104 (2011) The Zebrafish: Genetics, Genomics and Informatics 3rd Edition Edited by H. William Detrich III, Monte Westerfield, and Leonard I. Zon Volume 105 (2011) The Zebrafish: Disease Models and Chemical Screens 3rd Edition Edited by H. William Detrich III, Monte Westerfield, and Leonard I. Zon

Volumes in Series

Volume 106 (2011) Caenorhabditis elegans: Molecular Genetics and Development 2nd Edition Edited by Joel H. Rothman and Andrew Singson Volume 107 (2011) Caenorhabditis elegans: Cell Biology and Physiology 2nd Edition Edited by Joel H. Rothman and Andrew Singson Volume 108 (2012) Lipids Edited by Gilbert Di Paolo and Markus R Wenk Volume 109 (2012) Tetrahymena thermophila Edited by Kathleen Collins Volume 110 (2012) Methods in Cell Biology Edited by Anand R. Asthagiri and Adam P. Arkin Volume 111 (2012) Methods in Cell Biology Edited by Thomas Mu¨ler Reichart and Paul Verkade Volume 112 (2012) Laboratory Methods in Cell Biology Edited by P. Michael Conn Volume 113 (2013) Laboratory Methods in Cell Biology Edited by P. Michael Conn Volume 114 (2013) Digital Microscopy, 4th Edition Edited by Greenfield Sluder and David E. Wolf Volume 115 (2013) Microtubules, in Vitro, 2nd Edition Edited by John J. Correia and Leslie Wilson Volume 116 (2013) Lipid Droplets Edited by H. Robert Yang and Peng Li Volume 117 (2013) Receptor-Receptor Interactions Edited by P. Michael Conn

507

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Volumes in Series

Volume 118 (2013) Methods for Analysis of Golgi Complex Function Edited by Franck Perez and David J. Stephens Volume 119 (2014) Micropatterning in Cell Biology Part A Edited by Matthieu Piel and Manuel The´ry Volume 120 (2014) Micropatterning in Cell Biology Part B Edited by Matthieu Piel and Manuel The´ry Volume 121 (2014) Micropatterning in Cell Biology Part C Edited by Matthieu Piel and Manuel The´ry