The Nuclear Pore Complex: Methods and Protocols (Methods in Molecular Biology, 2502) 1071623362, 9781071623367

Citation preview

Methods in Molecular Biology 2502

Martin W. Goldberg Editor

The Nuclear Pore Complex Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

The Nuclear Pore Complex Methods and Protocols

Edited by

Martin W. Goldberg Department of Biosciences, Durham University, Durham, UK

Editor Martin W. Goldberg Department of Biosciences Durham University Durham, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2336-7 ISBN 978-1-0716-2337-4 (eBook) https://doi.org/10.1007/978-1-0716-2337-4 © Springer Science+Business Media, LLC, part of Springer Nature 2022 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface The Nuclear Pore Complex (NPC) stands at the interface between nuclear and cytoplasmic compartments. It facilitates and controls the exchange of almost all molecules between these two cellular compartments and is therefore a critical component for the control of transcription in the nucleus, and translation in the cytoplasm. It is also physically anchored to the endo-membrane system, as well as linked to the cytoskeleton, nucleoskeleton, and to specific domains of chromatin. It therefore has known, or potential, roles in controlling, or being affected by, all these systems. The “nuclear pore” was discovered in the 1950s by the newly developing methods of electron microscopy, and it soon became evident that a highly elaborate structure resided within the pore. This meant two things: not only was there a gateway to the nucleus, but the gateway had a gate and there had to be keys to that gate as well as mechanisms to carry cargoes through the gate. This led to two parallel, and sometimes overlapping, fields of study: (1) the structure of the NPC and mechanisms of movement through it, and (2) the signals and other factors that govern whether a molecule can access the gateway from either direction. This is now a large field of study, where we have a good, but incomplete, understanding of the structural framework of the NPC, as well as the structural and functional roles of individual nucleoporins. We have a less solid understanding however of the mechanisms of translocation through the NPC channel. We also have a good understanding, in exquisite atomic detail, of the proteins and other molecules responsible for carrying cargoes to and through the NPC. However, due to the myriad of diverse cargoes, the large number of transporter proteins and range of control mechanisms, there is still much to do. In addition, although the fundamentals of nucleocytoplasmic transport appear to be reasonably conserved between the simplest and most complex eukaryotes, as well as being similar in most tissues, it is becoming evident that there is considerable diversity in the details within different cells, and possibly even between different NPCs in the same cell. For this reason, and the fact that nucleocytoplasmic transport is such a central eukaryotic process, understanding the structure, function, interactions, posttranslational modifications, dynamics, and biophysical properties of the NPC, and its associated factors, is crucial to understanding many important biological questions beyond the basic biology of the NPC. The NPC has links to many disease and disorder processes, including cancer, viral infection, degenerative disorders, and aging. It is also important during development, tissue-specific cell functioning as well as genome organization and function. The NPC, therefore, is no longer just of interest to specialist laboratories studying its fundamental biology, but, in addition, to many laboratories studying disease, immunity, and aging as well as development, gene expression, and tissue function in all eukaryotic systems of interest, including humans and other mammals, model organisms such as fungi, worms and flies, as well as plants. The study of the NPC may therefore be crucial in diverse fields from the advancement of medicine to the attainment of food security. This therefore is a large field that requires a huge range of methodologies to answer all questions related to the NPC and its roles in many different processes. Many questions can be answered by standard cell and molecular biology methods, but due to the large size and complexity of the NPC, and its stable integration into membranes, cyto/nucleo-skeletal systems and chromatin, as well as the myriad of “substrates” passing or transporting through

v

vi

Preface

it, and its heterogeneity, such methods often have to be specifically adapted and new approaches developed. We present here a wide range of methods that cover both these cases, including those that have broad applications for isolating and analyzing NPCs and nucleoporins, their interactions and post-translational modifications, as well as protocols that address functional aspects of nucleoporins. Structural studies, imaging and biophysical methods have always played a central role in understanding many aspects of the nuclear transport system, so a number of protocols are also included showing how both new and traditional methods can be applied to NPC research. Apart from mammalian systems, the protocols also cover a wide range of model experimental organisms as well as plants. It is hoped that these will be useful to both specialist and nonspecialist laboratories for both comparative studies and to answer questions that can only easily be answered in specific systems. A single volume cannot cover all essential methods for a large and complex field such as this, but it is hoped that the broad range of protocols provided here will find utility in many laboratories interested in the mechanisms and functional roles of the NPC. Durham, UK

Martin W. Goldberg

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

THE NUCLEAR PORE COMPLEX AND NUCLEOPORINS

1 Affinity Isolation of Endogenous Saccharomyces Cerevisiae Nuclear Pore Complexes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ilona Nudelman, Javier Fernandez-Martinez, and Michael P. Rout 2 Transformation of Chaetomium thermophilum and Affinity Purification of Native Thermostable Protein Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nikola Kellner and Ed Hurt 3 Nuclear Pore Complex Assembly Using Xenopus Egg Extract . . . . . . . . . . . . . . . . Guillaume Holzer and Wolfram Antonin

PART II

v xi

3

35 51

NUCLEO-CYTOPLASMIC PASSAGE

4 Analysis of Nuclear Pore Complex Permeability in Mammalian Cells and Isolated Nuclei Using Fluorescent Dextrans . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Marcela Raices and Maximiliano A. D’Angelo 5 Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects in Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81 Saskia Hutten and Dorothee Dormann 6 Subcellular Fractionation Suitable for Studies of RNA and Protein Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Biljana Culjkovic-Kraljacic and Katherine L. B. Borden 7 Localizing Total mRNA in Plant Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Geraint Parry 8 Using Single Molecule RNA FISH to Determine Nuclear Export and Transcription Phenotypes in Drosophila Tissues . . . . . . . . . . . . . . . . . . . . . . . . . 113 Jennifer R. Aleman, Shawn C. Little, and Maya Capelson

PART III

FUNCTIONAL ANALYSIS OF NUCLEOPORINS

9 Analysis of Nucleoporin Function Using Inducible Degron Techniques . . . . . . . 129 Vasilisa Aksenova, Alexei Arnaoutov, and Mary Dasso 10 Monitoring of Chromatin Organization at the Nuclear Pore Complex, Inner Nuclear Membrane, and Nuclear Interior in Live Cells by Fluorescence Ratiometric Imaging of Chromatin (FRIC).. . . . . . . . . . . . . . . . . 151 Frida Niss, Cecilia Bergqvist, Anna-Lena Stro¨m, and Einar Hallberg

vii

viii

11

12

Contents

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans by Live Imaging and Functional Genomics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Patricia de la Cruz Ruiz, Raquel Romero-Bueno, and Peter Askjaer Protein Retargeting in Aspergillus nidulans to Study the Function of Nuclear Pore Complex Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 Subbulakshmi Suresh and Stephen A. Osmani

PART IV 13

14

15

16

Split-GFP Complementation to Study the Nuclear Membrane Proteome Using Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shary N. Shelton, Sarah E. Smith, and Sue L. Jaspersen Bimolecular Fluorescence Complementation: Quantitative Analysis of In Cell Interaction of Nuclear Transporter Importin α with Cargo Proteins. . . . . . . . . . . Alexander Lee, Marie A. Bogoyevitch, and David A. Jans Validation of Nuclear Pore Complex Protein–Protein Interactions by Transient Expression in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fumika Ikeda and Kentaro Tamura Binding Affinity Measurement of Nuclear Export Signal Peptides to Their Exporter CRM1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ho Yee Joyce Fung and Yuh Min Chook

PART V 17

18

20

21

22

205

215

235

245

POST TRANSLATIONAL MODIFICATIONS

Analysis of Ubiquitylation and SUMOylation of Yeast Nuclear Pore Complex Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259 Catherine Dargemont Purification of Cdk-CyclinB-Kinase–Targeted Phosphopeptides from Nuclear Envelope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Justin D. Blethrow, Amanda L. DiGuilio, and Joseph S. Glavy

PART VI 19

PROTEIN–PROTEIN INTERACTIONS

BIOPHYSICAL METHODS

Crystallization of Nuclear Export Signals or Small-Molecule Inhibitors Bound to Nuclear Exporter CRM1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ho Yee Joyce Fung and Yuh Min Chook Atomic Force Microscopy for Structural and Biophysical Investigations on Nuclear Pore Complexes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ivan Liashkovich, Gonzalo Rosso, and Victor Shahin Multivalent Interactions with Intrinsically Disordered Proteins Probed by Surface Plasmon Resonance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Larisa E. Kapinos and Roderick Y. H. Lim Assembly and Use of a Microfluidic Device to Study Nuclear Mechanobiology During Confined Migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Richa Agrawal, Aaron Windsor, and Jan Lammerding

285

299

311

329

Contents

PART VII 23

24

25 26

27

28

29

30

31

ix

IMAGING NPCS AND TRANSPORT

Speed Microscopy: High-Speed Single Molecule Tracking and Mapping of Nucleocytoplasmic Transport. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steven J. Schnell, Mark Tingey, and Weidong Yang Imaging Fluorescent Nuclear Pore Complex Proteins in C. elegans. . . . . . . . . . . . Courtney Lancaster, Giulia Zavagno, James Groombridge, Adelaide Raimundo, David Weinkove, Tim Hawkins, Joanne Robson, and Martin W. Goldberg Visualizing Nuclear Pore Complexes in Xenopus Egg Extracts . . . . . . . . . . . . . . . . Sampada Mishra and Daniel L. Levy TEM Imaging of Membrane Choreography During Mitosis of Drosophila Tissue Culture Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anton Strunov, Lidiya V. Boldyreva, Alexey V. Pindyurin, Maurizio Gatti, and Elena Kiseleva Scanning Electron Microscopy (SEM) and Immuno-SEM of Nuclear Pore Complexes from Amphibian Oocytes, Mammalian Cell Cultures, Yeast, and Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martin W. Goldberg and Jindrˇisˇka Fisˇerova´ NPC Structure in Model Organisms: Transmission Electron Microscopy and Immunogold Labeling Using High-Pressure Freezing/Freeze Substitution of Yeast, Worms, and Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Christine Richardson, Jindrˇisˇka Fisˇerova´, and Martin W. Goldberg High-Resolution Imaging and Analysis of Individual Nuclear Pore Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Boris Fichtman, Saroj G. Regmi, Mary Dasso, and Amnon Harel Live CLEM Imaging of Tetrahymena to Analyze the Dynamic Behavior of the Nuclear Pore Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tokuko Haraguchi, Hiroko Osakada, and Masaaki Iwamoto Visualizing Nuclear Pore Complex Assembly In Situ in Human Cells at Nanometer Resolution by Correlating Live Imaging with Electron Microscopy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Helena Bragulat-Teixidor, M. Julius Hossain, and Shotaro Otsuka

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

353 373

395

407

417

439

461

473

493

513

Contributors RICHA AGRAWAL • Weill Institute for Cellular and Molecular Biology, Cornell University, Ithaca, NY, USA VASILISA AKSENOVA • Division of Molecular and Cellular Biology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA JENNIFER R. ALEMAN • Department of Cell and Developmental Biology, University of Pennsylvania, Philadelphia, PA, USA WOLFRAM ANTONIN • Institute of Biochemistry and Molecular Cell Biology, Medical School, RWTH Aachen University, Aachen, Germany ALEXEI ARNAOUTOV • Division of Molecular and Cellular Biology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA PETER ASKJAER • Andalusian Center for Developmental Biology (CABD), CSIC/JA/ Universidad Pablo de Olavide, Seville, Spain CECILIA BERGQVIST • Department of Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden JUSTIN D. BLETHROW • Pacific Biosciences, Menlo Park, CA, USA MARIE A. BOGOYEVITCH • Department of Biochemistry and Molecular Biology, University of Melbourne, Parkville, VIC, Australia LIDIYA V. BOLDYREVA • Institute of Molecular and Cellular Biology (IMCB), SB RAS, Novosibirsk, Russia KATHERINE L. B. BORDEN • Department of Pathology and Cell Biology, Institute of Research in Immunology and Cancer, Universite´ de Montre´al, Montre´al, QC, Canada HELENA BRAGULAT-TEIXIDOR • Max Perutz Labs, a joint venture of the University of Vienna and the Medical University of Vienna, Vienna Biocenter (VBC), Vienna, Austria; Vienna BioCenter PhD Program, Doctoral School of the University of Vienna and Medical University of Vienna, Vienna, Austria MAYA CAPELSON • Department of Cell and Developmental Biology, University of Pennsylvania, Philadelphia, PA, USA YUH MIN CHOOK • Department of Pharmacology, UT Southwestern Medical Center, Dallas, TX, USA BILJANA CULJKOVIC-KRALJACIC • Department of Pathology and Cell Biology, Institute of Research in Immunology and Cancer, Universite´ de Montre´al, Montre´al, QC, Canada MAXIMILIANO A. D’ANGELO • Cellular and Molecular Biology of Cancer Program, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, CA, USA; NCI-Designated Cancer Center, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, CA, USA CATHERINE DARGEMONT • Institut de Ge´ne´tique Humaine, Universite´ de Montpellier, Laboratoire de Virologie Mole´culaire CNRS-UMR9002, Montpellier, France MARY DASSO • Division of Molecular and Cellular Biology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA PATRICIA DE LA CRUZ RUIZ • Andalusian Center for Developmental Biology (CABD), CSIC/ JA/Universidad Pablo de Olavide, Seville, Spain AMANDA L. DIGUILIO • Department of Biochemistry and Molecular Biophysics, University of Chicago, Chicago, IL, USA

xi

xii

Contributors

DOROTHEE DORMANN • Johannes Gutenberg Universit€ a t Mainz, Institute of Molecular Physiology, Mainz, Germany; Institute of Molecular Biology (IMB) Mainz, Mainz, Germany JAVIER FERNANDEZ-MARTINEZ • Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, NY, USA; Ikerbasque, Basque Foundation for Science, Bilbao, Spain; Instituto Biofisika (UPV/EHU, CSIC), University of the Basque Country, Leioa, Bizkaia, Spain BORIS FICHTMAN • Azrieli Faculty of Medicine, Bar-Ilan University, Safed, Israel JINDRˇISˇKA FISˇEROVA´ • Department of Biology of the Cell Nucleus, Institute of Molecular Genetics AS CR, Prague, Czech Republic HO YEE JOYCE FUNG • Department of Pharmacology, UT Southwestern Medical Center, Dallas, TX, USA MAURIZIO GATTI • IBPM CNR and Department of Biology and Biotechnology, Sapienza University of Rome, Rome, Italy JOSEPH S. GLAVY • Department of Pharmaceutical Sciences, Fisch College of Pharmacy, University of Texas at Tyler, Tyler, TX, USA MARTIN W. GOLDBERG • Department of Biosciences, Durham University, Durham, UK JAMES GROOMBRIDGE • Department of Biosciences, Durham University, Durham, UK EINAR HALLBERG • Department of Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden TOKUKO HARAGUCHI • Graduate School of Frontier Biosciences, Osaka University, Suita, Japan AMNON HAREL • Azrieli Faculty of Medicine, Bar-Ilan University, Safed, Israel TIM HAWKINS • Department of Biosciences, Durham University, Durham, UK GUILLAUME HOLZER • Institute of Biochemistry and Molecular Cell Biology, Medical School, RWTH Aachen University, Aachen, Germany M. JULIUS HOSSAIN • Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany ED HURT • Biochemistry Center, University of Heidelberg, Heidelberg, Germany SASKIA HUTTEN • Johannes Gutenberg Universit€ at Mainz, Institute of Molecular Physiology, Mainz, Germany FUMIKA IKEDA • Department of Environmental and Life Sciences, School of Food and Nutritional Sciences, University of Shizuoka, Shizuoka, Japan MASAAKI IWAMOTO • Department of Biosciences, College of Humanities and Sciences, Nihon University, Tokyo, Japan DAVID A. JANS • Nuclear Signalling Laboratory, Monash Biomedicine Discovery Institute, Monash University, Clayton, VIC, Australia; Department of Biochemistry and Molecular Biology, University of Melbourne, Parkville, VIC, Australia SUE L. JASPERSEN • Stowers Institute for Medical Research, Kansas City, MO, USA; Department of Molecular and Integrative Physiology, University of Kansas Medical Center, Kansas City, KS, USA LARISA E. KAPINOS • Biozentrum and the Swiss Nanoscience Institute, University of Basel Switzerland, Basel, Switzerland NIKOLA KELLNER • Biochemistry Center, University of Heidelberg, Heidelberg, Germany ELENA KISELEVA • Institute of Cytology and Genetics (ICG), Siberian Branch of Russian Academy of Sciences (SB RAS), Novosibirsk, Russia

Contributors

xiii

JAN LAMMERDING • Weill Institute for Cellular and Molecular Biology, Cornell University, Ithaca, NY, USA; Nancy E. and Peter C. Meinig School of Engineering, Cornell University, Ithaca, NY, USA COURTNEY LANCASTER • MRC Laboratory for Molecular Cell Biology, University College London, London, UK ALEXANDER LEE • Nuclear Signalling Laboratory, Monash Biomedicine Discovery Institute, Monash University, Clayton, VIC, Australia; Department of Biochemistry and Molecular Biology, University of Melbourne, Parkville, VIC, Australia DANIEL L. LEVY • Department of Molecular Biology, University of Wyoming, Laramie, WY, USA IVAN LIASHKOVICH • Institute of Physiology II, University of Mu¨nster, Mu¨nster, Germany RODERICK Y. H. LIM • Biozentrum and the Swiss Nanoscience Institute, University of Basel Switzerland, Basel, Switzerland SHAWN C. LITTLE • Department of Cell and Developmental Biology, University of Pennsylvania, Philadelphia, PA, USA SAMPADA MISHRA • Department of Molecular Biology, University of Wyoming, Laramie, WY, USA FRIDA NISS • Department of Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden ILONA NUDELMAN • Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, NY, USA; Fisher Drug Discovery Resource Center (DDRC), The Rockefeller University, New York, NY, USA HIROKO OSAKADA • Laboratory of Molecular Cell Biology, Research Institute for Diseases of Old Age, Juntendo University Graduate School of Medicine, Tokyo, Japan STEPHEN A. OSMANI • The Department of Molecular Genetics, The Ohio State University, Columbus, OH, USA SHOTARO OTSUKA • Max Perutz Labs, a joint venture of the University of Vienna and the Medical University of Vienna, Vienna Biocenter (VBC), Vienna, Austria GERAINT PARRY • GARNet, School of Biosciences, Cardiff University, Cardiff, UK ALEXEY V. PINDYURIN • Institute of Molecular and Cellular Biology (IMCB), SB RAS, Novosibirsk, Russia MARCELA RAICES • Cellular and Molecular Biology of Cancer Program, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, CA, USA; NCI-Designated Cancer Center, Sanford Burnham Prebys Medical Discovery Institute, La Jolla, CA, USA ADELAIDE RAIMUNDO • Department of Biosciences, Durham University, Durham, UK SAROJ G. REGMI • Division of Molecular and Cellular Biology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA A. CHRISTINE RICHARDSON • Department of Biosciences, Durham University, Durham, UK JOANNE ROBSON • Department of Biosciences, Durham University, Durham, UK RAQUEL ROMERO-BUENO • Andalusian Center for Developmental Biology (CABD), CSIC/ JA/Universidad Pablo de Olavide, Seville, Spain GONZALO ROSSO • Institute of Physiology II, University of Mu¨nster, Mu¨nster, Germany MICHAEL P. ROUT • Laboratory of Cellular and Structural Biology, The Rockefeller University, New York, NY, USA STEVEN J. SCHNELL • Department of Biology, Temple University, Philadelphia, PA, USA VICTOR SHAHIN • Institute of Physiology II, University of Mu¨nster, Mu¨nster, Germany SHARY N. SHELTON • Stowers Institute for Medical Research, Kansas City, MO, USA SARAH E. SMITH • Stowers Institute for Medical Research, Kansas City, MO, USA

xiv

Contributors

ANNA-LENA STRO¨M • Department of Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden ANTON STRUNOV • Institute of Cytology and Genetics (ICG), Siberian Branch of Russian Academy of Sciences (SB RAS), Novosibirsk, Russia; Center for Anatomy and Cell Biology, Medical University of Vienna, Vienna, Austria SUBBULAKSHMI SURESH • The Department of Molecular Genetics, The Ohio State University, Columbus, OH, USA KENTARO TAMURA • Department of Environmental and Life Sciences, School of Food and Nutritional Sciences, University of Shizuoka, Shizuoka, Japan MARK TINGEY • Department of Biology, Temple University, Philadelphia, PA, USA DAVID WEINKOVE • Department of Biosciences, Durham University, Durham, UK AARON WINDSOR • Cornell NanoScale Science and Technology Facility, Cornell University, Ithaca, NY, USA WEIDONG YANG • Department of Biology, Temple University, Philadelphia, PA, USA GIULIA ZAVAGNO • Department of Biosciences, Durham University, Durham, UK

Part I The Nuclear Pore Complex and Nucleoporins

Chapter 1 Affinity Isolation of Endogenous Saccharomyces Cerevisiae Nuclear Pore Complexes Ilona Nudelman, Javier Fernandez-Martinez, and Michael P. Rout Abstract Studying protein complexes in vitro requires the production of a relatively pure sample that maintains the full complement, native organization, and function of that complex. This can be particularly challenging to achieve for large, multi-component, membrane embedded complexes using the traditional recombinant expression and reconstitution methodologies. However, using affinity capture from native cells, suitable whole endogenous protein complexes can be isolated. Here we present a protocol for the affinity isolation of baker’s yeast (S. cerevisiae) nuclear pore complexes, which are ~50 MDa assemblies made up of 552 distinct proteins and embedded in a double-membraned nuclear envelope. Producing this sample allowed us for the first time to perform analyses to characterize the mass, stoichiometry, morphology, and connectivity of this complex and to obtain its integrative structure with ~9 Å precision. We believe this methodology can be applied to other challenging protein complexes to produce similar results. Key words Affinity capture, Native Isolation, Nuclear pore complex, Endogenous macromolecular assembly, Structural and functional analyses, Electron microscopy, Baker’s yeast, S. cerevisiae

1

Introduction Protein macromolecular assemblies are the workhorses of living organism; thus, deciphering their function and how they perform is crucial to understanding cellular processes. A relatively pure sample of a protein complex must be obtained to perform any in vitro biochemical, biophysical, structural, or functional studies. Two main methodologies exist to produce such a sample. One is a “bottom up” approach, namely, to overexpress each of the protein complex components in a host organism, purify them, and reconstitute the assembly. The other “top down” approach is to isolate an endogenous protein or protein complex from its native organism. Both methodologies have advantages and disadvantages, depending on the specific protein complex to be produced. However, in the case of large and multi-component, or membrane embedded assemblies, overexpression in a host organism and subsequent

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2022

3

4

Ilona Nudelman et al.

purification and reconstitution can be extremely challenging, whereas affinity isolation from the native environment offers many advantages, both in terms of feasibility and functional quality of the sample. The concept behind affinity isolation is quite simple—to isolate a protein complex from a cell, one of the proteins in the complex is tagged with an affinity tag (this protein is termed a “handle”), which can tightly bind to an antibody (or another binding partner). Tagging might not be necessary if there is a good (tightly binding and specific) antibody to the protein of interest itself. Cells are grown, harvested, and lysed, and the clarified lysate is incubated with an insoluble matrix (e.g., agarose beads, magnetic beads) coated with the antibody. The “handle” protein binds to the antibody via the affinity tag and if the associated complex is sufficiently stable, it remains attached to the “handle” protein and thereby immobilized on the matrix. The remaining lysate is discarded, and the protein complex is eluted (natively or by denaturation). Now the protein complex can be analyzed in any desired fashion—denatured eluates are usually characterized by SDS-PAGE followed by mass spectrometry (MS), while native assemblies can be used for structural (e.g., cryo-electron microscopy (cryo-EM), cross-linking with mass spectrometric readout (CX-MS)), proteomic (e.g., quantitative MS, native MS), and functional studies [1–7]. Despite the simplicity of the overall concept behind affinity isolation, its successful application requires careful optimization of all the parameters for the protein complex in question. These parameters include performing an efficient lysis, choosing the best antibody/tag pair, choosing the correct “handle,” finding the best buffer composition and optimizing the elution. Our group has pioneered the development of methodologies for optimizing the outcomes of affinity isolations for a large variety of protein complexes in many different organisms [3, 8–13]. Our approach is based on several principles (Fig. 1). First, cells are lysed by flashfreezing in liquid nitrogen followed by cryo-milling to generate micron-scale cell powder. This approach, essentially keeping the lysis stage in an extremely preservative solid phase, enables excellent preservation of native protein complexes and highly efficient cell breakage leading to full access to these complexes, while minimizing proteolytic degradation [11, 14, 15]. In addition, cell powder can be stored almost indefinitely at 80  C and portions of the milled powder can be weighed from a larger stock depending on the scale of the experiment, allowing flexibility and multiplexing of the experimental design—including making exploration of multiple subsequent buffers straightforward (below) (Fig. 1a). Next, the antibody/tag pair used for affinity isolation must be optimized for the specific complex. Important parameters include binding specificity, efficiency, and speed, as well as optimal tag placement on the “handle” in terms of steric hindrance and accessibility to the

Affinity Isolation of Endogenous Whole Yeast NPCs

5

Fig. 1 Affinity isolation methodology. An outline of a general affinity isolation protocol, which can be optimized for various protein complexes from many organisms, as described in this work. (a) Cell lysis using flashfreezing and cryo-milling to obtain cell powder which can be stored at 80  C and used as needed for any scale experiment. (b) Affinity capture by resuspending cell powder in an optimized affinity capture buffer and using the clarified (centrifuged and/or filtered) lysate for incubation with antibody-coupled magnetic beads to capture the complex on the beads followed by discarding the remaining supernatant and washing steps. Complex elution is accomplished by denaturation or by native elution (proteolytic removal or competitive binding) followed by separation from the beads. Eluted complex can be analyzed by SDS-PAGE and other downstream analyses mentioned in the text

antibody [13, 14]. The affinity isolation begins by resuspending frozen cell powder in a buffer (Fig. 1b). Once the powder is in contact with the buffer and so transitioned to the liquid phase in a (by definition) non-native environment, all protein complexes can in principle begin to destabilize and dissociate at different rates, depending on temperature, the strength of their interaction, their environmental conditions, etc. The goal is to preserve the non-covalent interactions of the “handle” with the other proteins in the complex for as long as possible by fast and efficient resuspension into buffer conditions that have been optimized to be the most stabilizing. The physical characteristics of the resuspension process, such as buffer-to-powder ratio, temperature, and resuspension mode (vortex, shaking, brief sonication, etc.) must be empirically optimized for the specific complex. Buffer composition is crucial

6

Ilona Nudelman et al.

for maintaining complex stability throughout the affinity isolation; therefore, many conditions need to be tested to find the optimal pH and buffer type, overall ionic strength, salt type(s) and concentration, detergent type(s) and concentration, and other additives as needed. We developed a method to perform high throughput screening to find the best buffer formulations for a specific affinity purification [10]. The resuspended lysate is clarified by centrifugation and/or filtration to eliminate insoluble cell debris such as cell wall fragments, followed by incubation with antibody conjugated paramagnetic beads to achieve immobilization of the complex on the beads via binding between the tag on the “handle” and the antibody on the matrix (Fig. 1b). This process should be quick and efficient, without introducing substantial non-specific binding to the complex or to the beads, emphasizing the need to use a high affinity and specificity antibody/tag pair, beads with extremely low non-specific binding, and appropriate buffer composition. After immobilization is complete, the lysate is discarded, and the beads are washed a few times to ensure all non-specifically bound components are removed (Fig. 1b). The washing buffer composition as well as number and mode of washing steps should also be empirically optimized to maximize removal of non-specific binders while minimizing complex destabilization. Subsequently, the complex can be eluted from the beads, either by native elution in which the complex structure and activity are preserved or by denaturation (Fig. 1b). If native elution is required, the elution method also needs to be optimized to be quick and efficient, to preserve the complex intact, and to not introduce non-specific binding. Two options can be explored: (1) proteolytic removal via a specific protease (where a protease cleavage target sequence should be inserted between the tag and the “handle” protein) or (2) elution by competitive binding (e.g., anti-FLAG antibody and a FLAG tag to be eluted with FLAG peptide [12, 14]). The eluted complex can now be analyzed by SDS-PAGE and characterized further using any number of techniques mentioned above. Here, we present a protocol for affinity isolation of endogenous whole nuclear pore complexes (NPCs) from S. cerevisiae. The NPC is the sole mediator of all nucleocytoplasmic transport in eukaryotic cells and is involved in many crucial nuclear processes [16]. It is a large macromolecular assembly (~50 MDa in yeast) composed of 552 distinct proteins (called nucleoporins or Nups) and embedded in the nuclear envelope which is a double-membraned barrier that surrounds the nucleus [1, 17]. The NPC is a cylindrical assembly made up of 8 spokes arranged around a C-8 symmetry axis perpendicular to the nuclear envelope. The spokes form concentric rings around the same C-8 axis—an inner ring and two outer rings, nuclear, and cytoplasmic, in turn connected to the nuclear basket and the cytoplasmic mRNA export complex, respectively (see Fig. 4 in [1]). Recently, our group was able to determine a structure for

Affinity Isolation of Endogenous Whole Yeast NPCs

7

the baker’s yeast NPC. We analyzed samples of endogenous whole NPCs prepared using the protocol presented here to determine a cryo-electron tomography (cryo-ET) map of the isolated NPC at ~28 Å average resolution, use mass spectrometric methods to determine its mass and stoichiometry, and subject the isolated NPCs to CX-MS, which provided the distance restraints within and between Nups [1]. All these restraints were used to integratively calculate a structure of the yeast NPC at unprecedented detail, allowing us to draw several paradigm-shifting conclusions as to how the NPC might accomplish its functions and why it is built the way it is. The case of the NPC is a prime example of how the ability to produce an optimized affinity-captured endogenous sample enabled significant progress in the field. Prior research of the NPC focused primarily on in vivo or in situ studies [18–21] or studies of isolated or endogenously overexpressed, reconstituted subcomplexes [22, 23]. While these are extremely valuable, a detailed picture of the complete assembly was needed to put all the data in the correct context [18, 24]; thus, although isolation of any macromolecular complex likely alters it to some degree, the high-resolution data obtained is nonetheless invaluable for understanding its structure and functionalities in situ. This was enabled by producing a sufficiently pure sample of whole NPCs and analyzing it using the methods described above and in [1]. Using this approach on other large macromolecular complexes may lead to similar advances. The specific parameters of the protocol described here were optimized using the methodologies described above [3, 8–10, 14, 15] following extensive trial and error efforts to produce the best possible sample. As a tag we used Protein A (PrA), which is a 42 kDa surface protein originally found in the cell wall of the bacteria Staphylococcus aureus. The IgG-binding domains of PrA are widely used as affinity tags since they bind to IgG via the constant (Fc) region [25, 26] and therefore do not require an antigen-specific antibody for affinity capture. We used rabbit polyclonal IgG for affinity isolation of proteins and protein complexes tagged with the PrA affinity tag. Our “handle” protein is predominantly Mlp1, a nuclear basket component [1], peripherally situated at the nuclear side of the NPC which allows good steric access of the antibody to the tag for efficient binding. We explored other peripherally situated Nups as “handles” and found Nup82 (cytoplasmic mRNA export platform component [1]) and Nup84 (a component of the nuclear and cytoplasmic outer rings [1]) to be equally good. We needed intact NPCs for the downstream analyses, therefore we chose to use protease cleavage as our native elution method. PreScission Protease (PPX), a commercially available version of human rhinovirus (HRV) 3C protease was selected and its cleavage target sequence (LEVLFQ/GP) was inserted

8

Ilona Nudelman et al.

between our “handle” protein (Mlp1, Nup84, or Nup82) and the PrA tag. We routinely use Dynabeads M-270 Epoxy magnetic beads from Thermo Fisher Scientific conjugated to the antibody of choice (in our case, rabbit polyclonal IgG) for our affinity isolations (bead conjugation protocol is included here). After significant empirical exploration, we found that an optimal buffer formulation for the stabilization of the NPCs was 20 mM HEPES pH 7.4, 50 mM potassium acetate (KOAc), 20 mM NaCl, 2 mM MgCl2, 0.5% Triton X-100, 0.1% Tween-20, 1 mM DTT, 10% glycerol. The main buffer species is HEPES at pH 7.4 which was chosen after unsuccessfully trying various other options, such as Bis-Tris at pH 6.5 and sodium phosphate at pH 8. Additives KOAc, NaCl, MgCl2, and DTT were found to contribute to the stability of the intact complex (increasing the ratio of intact NPCs to dissociated subcomplexes). Many different permutations of various detergents were explored but we found that a combination of 0.5% Triton X-100 and 0.1% Tween-20 enabled efficient extraction of NPCs from the surrounding nuclear envelope [1]. Even with this treatment, there are substantial remnants of the native pore membrane present in the isolated NPCs, as shown by cryo-ET analyses [1]. Once the NPCs are extracted from the membrane, we continue the affinity isolation without Triton X-100, since its prolonged presence is not necessary. The addition of 10% glycerol significantly improved the yield of the affinity isolation, indicated by the ratio between the amount of the complex successfully cleaved by the protease and found in the eluate and amount of the complex successfully cleaved by the protease, but remained stuck to the beads probably via non-specific interactions with the bead surface (see Note 1). The NPC is large, flexible, and relatively fragile without the support of the nuclear envelope, therefore, once extracted it is vulnerable to harsh mechanical stresses. Consequently, our protocol employs gentle handling such as low speed centrifugation followed by filtration to eliminate insoluble cell debris from the lysate as well as a single wash step after the immobilization on the beads to remove the non-specifically bound components. We found that the optimal resuspension parameters to preserve intact NPCs are working with 1.0–1.5 g cell powder aliquots weighed out in 50 mL tubes and resuspended by vortexing in 9.0–13.5 mL of buffer (buffer-to-powder ratio 1/10 v/v). Using these conditions allows for faster and more even resuspension with less vortexing, which reduces the mechanical shear forces on the NPCs. It is very important to perform the affinity isolation as quickly as possible from the time of resuspension until the elution (in our hands this takes a few hours) and to keep everything at 4  C for the duration of the protocol to prevent dissociation of the NPCs. This protocol was used to produce samples for the various downstream analysis described in [1] such as mass spectrometry (MS) quantitation and cross-linking and cryo-ET with

Affinity Isolation of Endogenous Whole Yeast NPCs

9

sub-tomogram averaging. Therefore, it constitutes a large-scale preparation of ~175 μL of sample with a final concentration of 0.3–0.4 mg/mL, requiring 15 g of cell powder and a few adjustments in the protocol due to using such large amounts. However, downscaling is possible and encouraged for optimization tests and requires simple ratio adjustments to the protocol. We use 1–1.5 g of cell powder for testing affinity isolation performance. In general, it is not recommended to use this protocol for less than one 1–1.5 g portion of powder, due to resuspension related issues, as mentioned above.

2

Materials

2.1 Growing, Harvesting, and FlashFreezing Yeast Cells

The yeast strain used for this protocol is MLP1-PPX-PrA (see [1], Supplementary Table 5). It is a S. cerevisiae W303 strain in which the endogenous gene of interest (encoding for the “handle” protein used for the affinity isolation), in our case nucleoporin MLP1, is tagged with a PrA affinity tag (see Note 2) and a PPX cleavage target sequence inserted between MLP1 and PrA, to enable native elution. The same protocol can be used for S. cerevisiae strains in which Nup84- or Nup82-encoding genes are similarly tagged (see Nup82-PPX-PrA or Nup84-PPX-PrA in [1], Supplementary Table 5). All strains can be obtained from the Rout lab upon request (http://ncdir.org/cd/). 1. Frozen glycerol stock of Mlp1-PPX-PrA strain (W303: MATα ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 MLP1PPX-ProteinA::HIS5). 2. YPD agar plates. 3. YPD media: 1% yeast extract, 2% peptone, 2% glucose. 4. Small (100 mL) and medium (250 mL) culture flasks. 5. Six large culture flasks (such as 2.8 L Nalgene™ Polycarbonate Fernbach Culture Flasks from Thermo Fisher Scientific). 6. 1 M Ampicillin. 7. Hemocytometer. 8. Refrigerated centrifuge, rotor, and flasks for pelleting large cell volumes (such as Beckman Coulter Avanti J-26XP centrifuge, with JLA-8.1000 rotor and 1 L bottles). 9. Ice cold ddH2O. 10. 50 mL tubes. 11. Refrigerated centrifuge and rotor for 50 mL tubes (such as Beckman Coulter Allegra X-14R centrifuge, with SX4750A ARIES™ Swinging-Bucket Rotor and 50 mL tube adapters).

10

Ilona Nudelman et al.

12. 12% w/v polyvinylpyrrolidone (PVP), an extracellular cryoprotectant. Dissolve 1.2 g PVP in 10 mL ddH2O. 13. 200 mM HEPES, pH 7.4: For 10 mL, dissolve 476.6 mg HEPES in 9 mL ddH2O. Adjust pH by adding 1 M KOH. Add ddH2O to 10 mL. 14. Resuspension buffer: 20 mM HEPES pH 7.5, 1.2% PVP. For 10 mL, combine 1 mL 12% w/v PVP, 1 mL 200 mM HEPES pH 7.4 and 8 mL ddH2O. 15. Protease Inhibitor Cocktail (Sigma Aldrich P8340). 16. Solution P: dissolve 2 mg Pepstatin A and 90 mg PMSF in 5 mL absolute ethanol. Store at 4 ̊C and discard after 3 weeks. Note that PMSF is highly poisonous, use appropriate caution (wear gloves, work in a fume hood, etc.). 17. 1 M Dithiothreitol (DTT) solution: weigh 1.54 g of DTT, dissolve completely in a final volume of 10 mL ddH2O. Filter sterilize, aliquot (1–1.5 mL) and store at 20  C. 18. Vacuum aspirator. 19. Liquid nitrogen. 20. Cryoprotective gloves. 21. Styrofoam box. 22. Syringe needle. 23. 25 mL or 50 mL syringe. 2.2 Cryo-Milling of Yeast Cells

1. Liquid nitrogen. 2. Cryoprotective gloves. 3. Styrofoam box. 4. Retsch Planetary Ball Mill PM100. 5. Retsch cryo-milling jar assembly: stainless steel jar (50 mL or 125 mL), lid, and 20 mm balls. 6. Tongs. 7. Spatula. 8. 50 mL tubes.

2.3 Antibody Conjugation to Magnetic Beads

1. Dynabeads M-270 Epoxy (Thermo Fisher Scientific 14302D). 2. Rabbit polyclonal IgG - purified from normal Rabbit Serum using Protein A affinity chromatography (Innovative Research)—this product comes as a solution with a 10–20 mg/mL concentration. Equivalent products from other sources can also be used. 3. 15 mL tubes. 4. Magnetic separator for 15 mL tubes (e.g., DynaMag™-15 from Thermo Fisher Scientific).

Affinity Isolation of Endogenous Whole Yeast NPCs

11

5. Vacuum aspirator. 6. Vortex mixer. 7. Parafilm. 8. 2 mL round-bottom tubes. 9. Tube shaker or rocker. 10. 0.1 M sodium phosphate, pH 7.4: For 1 L, dissolve 2.6 g NaH2PO4·H2O and 21.7 g Na2HPO4·7H2O in 1 L. Check the pH. Filter and store at RT. 11. 3 M ammonium sulfate: For 100 mL, dissolve 39.6 g (NH4)2SO4 in 50 mL of 0.1 M sodium phosphate buffer, pH 7.4. Filter and store at RT. 12. Rotating wheel or tube shaker/rocker in an incubator (temperature set to 30  C). 13. 0.1 M glycine–HCl pH 2.5. 14. 10 mM Tris–HCl pH 8.8. 15. 0.1 M trimethylamine (TEA)—make fresh. For 5 mL, mix 70 μL of TEA with 4.93 mL ddH2O. 16. Phosphate-buffered saline (PBS). 17. PBS with 0.5% (w/v) Triton X-100. 18. PBS with 0.02% sodium azide (NaN3). 2.4 Affinity Isolation of Yeast NPCs

1. 15 g cryo-milled cell powder (from Subheading 3.1 of this protocol) (see Note 3). 2. 750 μL conjugated IgG Dynabeads slurry (from Subheading 3.2 of this protocol) (see Note 4). 3. 0.22 μm filters. 4. 50 mL tubes. 5. 15 mL tubes. 6. 1.5 mL tubes. 7. 200 mL freshly made affinity capture (AC) buffer: 20 mM HEPES pH 7.4, 50 mM potassium acetate, 20 mM NaCl, 2 mM MgCl2, 0.5% Triton X-100, 0.1% Tween-20, 1 mM DTT, 10% glycerol. To prepare, combine in a 250 mL cylinder: 4 mL 1 M HEPES pH 7.4., 2 mL 5 M potassium acetate, 0.8 mL 5 M NaCl, 0.4 mL 1 M MgCl2, 10 mL 10% Triton X-100, 1 mL 20% Tween-20, 0.2 mL 1 M DTT, 20 mL 100% glycerol, 161.6 mL ddH2O. Mix, filter sterilize, and store on ice. 8. 50 mL freshly made native elution (NE) buffer: 20 mM HEPES pH 7.4, 50 mM potassium acetate, 20 mM NaCl, 2 mM MgCl2, 0.1% Tween-20, 1 mM DTT, 10% glycerol. To prepare, combine in a 50 mL tube: 1 mL 1 M HEPES pH 7.4,

12

Ilona Nudelman et al.

0.5 mL 5 M potassium acetate, 0.2 mL 5 M NaCl, 0.1 mL 1 M MgCl2, 0.25 mL 20% Tween-20, 0.05 mL 1 M DTT, 5 mL 100% glycerol, 42.9 mL ddH2O. Mix, filter sterilize, and store on ice. 9. Protease Inhibitor Cocktail (Sigma Aldrich P8340). 10. Solution P: dissolve 2 mg Pepstatin A and 90 mg PMSF in 5 mL absolute ethanol. Store at 4 ̊C and discard after 3 weeks. Note that PMSF is highly poisonous, use appropriate caution (wear gloves, work in a fume hood, etc.). 11. 1.6 μm Whatman glass microfiber syringe filters (Cytiva Life Sciences 6882-2516). 12. Laboratory syringes (various sizes). 13. Magnetic separator for 15 and 1.5 mL tubes (e.g., DynaMag™-15, DynaMag™-2 from Thermo Fisher Scientific). 14. Vacuum aspirator. 15. Vortex mixer. 16. Laboratory balance. 17. Liquid nitrogen. 18. Refrigerated centrifuge and rotor for 50 mL tubes (such as Beckman Coulter Allegra X-14R centrifuge, with SX4750A ARIES™ Swinging-Bucket Rotor and 50 mL tube adapters). 19. Refrigerated micro-centrifuge and rotor (such as Eppendorf 5417R tabletop centrifuge with an F-45-30-11 rotor). 20. Rotating wheel mixer (or shaker) set at 4  C. 21. Styrofoam box. 22. Spatula. 23. PreScission protease (Cytiva Life Sciences 27084301). 24. Benchtop mini centrifuge (such as mySPIN™ 6 Mini Centrifuge from ThermoFisher Scientific). 2.5 SDS-PAGE Analysis

1. XCell SureLock Mini-Cell Electrophoresis System (ThermoFisher Scientific) or equivalent. 2. Electrophoresis power supply (such as PowerPac™ Basic Power Supply from Bio-Rad). 3. NuPAGE™ 4–12%, Bis-Tris, 1.0 mm, Mini Protein Gel, 10-well (ThermoFisher Scientific) (see Note 5). 4. MES running buffer (such as NuPAGE™ MES SDS Running Buffer (20) from ThermoFisher Scientific). 5. NuPAGE™ Scientific).

LDS

Sample

Buffer

(4)

(ThermoFisher

6. 1 M DTT solution (see Subheading 2.1, step 10).

Affinity Isolation of Endogenous Whole Yeast NPCs

13

7. 1.5 mL tubes. 8. NuPAGE™ Antioxidant. 9. Protein marker (such as Color Prestained Protein Standard, Broad Range (10–250 kDa) from New England Biolabs). 10. 10 mg/mL Bovine Serum Albumin (BSA) stock solution: add 100 mg of BSA (Sigma Aldrich or similar) to a 15 mL tube containing 9.5 mL of ddH2O. Gently rock the capped tube until the BSA is completely dissolved. Do not vortex. Adjust the final volume to 10 mL with ddH2O. Divide into 1 mL aliquots and store at 20  C. 11. 0.25 mg/mL BSA stock solution: add 25 μL of the 10 mg/mL BSA stock solution to 975 μL ddH2O. Gently mix, then divide into 100 μL aliquots and store at 20  C. 12. Sturdy spatula or “gel knife.” 13. Gel staining box or container. 14. Orbital shaker at room temperature. 15. Imperial™ Protein Stain (ThermoFisher Scientific). 16. Standard image scanner or gel scanner (such as ImageQuant LAS 4000 from Cytiva Life Sciences). 2.6 Negative Stain Electron Microscopy (EM) Analysis

1. TEM grids for negative stain EM—200 mesh continuous carbon Cu TEM grids (such as CF200-Cu Electron Microscopy Sciences). 300 or 400 mesh continuous carbon Cu TEM grids are also suitable. 2. A set of fine-tipped tweezers: straight, self-closing, and curved. Dumont biology grade tweezers with style five tips are recommended (such as 72701-01, 0202-N5-PO, and 0208-7-PO, respectively, from Electron Microscopy Sciences). However, any tweezers suited for extra-fine handling are acceptable. 3. Transmission electron microscope—80–120 keV or 80–200 keV side entry microscope (such as JEOL 1400 Plus or FEI TECNAI G2 Spirit BioTwin), equipped with a digital camera. 4. 2% (w/v) Uranyl Acetate stain solution (UA stain)—combine 100 mg Uranyl Acetate powder (such as 22,400 from Electron Microscopy Sciences) with 5 mL ddH2O. Place the solution on a rocker and incubate for 2 h, then sonicate for 5 min. Filter the solution with a 0.22 μm syringe filter (or centrifuge and retain the supernatant). Store the UA stain wrapped in aluminum foil in a dark place (it is light sensitive) and at RT (it aggregates at lower temperatures) up to 1 year. 5. Glow discharge system or plasma cleaner (such as PELCO easiGlow™ Glow Discharge Cleaning System). 6. Parafilm.

14

Ilona Nudelman et al.

7. 1 mL syringes. 8. 0.2 μM filters for 1 mL syringes (such as 4 mm Cellulose Acetate Nalgene™ Syringe Filters, 171-0020, from Thermo Fisher Scientific). 9. Blot paper (e.g., Whatman grade 1 qualitative filter paper, standard grade, 90 mm circle from Cytiva Life Sciences). 10. EM grid storage box (e.g., 71,150 from Electron Microscopy Sciences or use empty boxes from purchased grids).

3

Methods

3.1 Growing, Harvesting, and FlashFreezing Yeast Cells

This section involves growing, harvesting, and flash-freezing of yeast cells. The affinity isolation protocol described in this manuscript requires 15 g of cryo-milled cell powder (see Subheading 3.4). The typical yield of S. cerevisiae cell cultures harvested at 3.0  107 cells/mL is 3–4 g per liter, but it may vary depending on the strain and the final cell density. We usually grow 6–12 L of S. cerevisiae cell culture at a time, producing roughly 20–50 g of cryo-milled cell powder (the protocol described here is for 12 L S. cerevisiae cell culture). Cell culture volume can be adjusted according to need (see Note 6). The protocols for harvesting and flash-freezing of cells are extensively described on the National Center for Dynamic Interactome Research website (https://www. ncdir.org/protocols/), including a video tutorial (titled “Harvesting Cells and Making Noodles”) [27]. Please check the link for updated versions before you perform these procedures. It is crucial to follow the prescribed safety procedures and take extreme care when handling liquid nitrogen. 1. Streak the strain from the frozen glycerol stock onto a YPD agar plate (for this strain, a regular YPD plate is used; for other strains use an appropriate selection plate, if needed). Incubate the plate at 30  C until single yeast colonies reach ~1–2 mm in diameter (this usually takes 2–3 days). Seal plates with Parafilm and store at 4  C for up to 1 month. Day One 2. Starter culture—prepare a small culture flask with 15 mL pre-sterilized YPD media. Add 1:1000 (v/v) 1 M Ampicillin to prevent bacterial contamination. Inoculate with a single colony from the agar plate. Grow overnight at 30  C with shaking at ~200 rpm. 3. Prepare 12 L of YPD media divided equally between six large flasks (2 L per flask). Cover flasks with aluminum foil, autoclave to sterilize, and allow to cool overnight.

Affinity Isolation of Endogenous Whole Yeast NPCs

15

Day Two 4. Estimate cell density of the starter culture using a hemocytometer (following manufacturer’s instructions). At this point the culture should be very dense (in stationary phase). 5. Pre-culture—prepare a flask with 50 mL pre-sterilized YPD media. Add 1:1000 (v/v) 1 M Ampicillin. Inoculate with 0.3 mL of the starter culture (1/50 v/v). Grow at 30  C with shaking at ~200 rpm until mid-log phase (~3  107 cells/mL) is reached (estimate cell density using a hemocytometer). This usually takes ~5 h assuming a doubling time of ~1.5 h. 6. Large-scale cultures—add 1:1000 (v/v) 1 M Ampicillin to the 6 autoclaved flasks with YPD media. Calculate inoculation volume to yield a final 2 L culture at mid-log phase (~3  107 cells/mL) after an overnight incubation (for example: ~18 h incubation requires ~0.58 mL inoculation volume from a 2.5  107 cells/mL pre-culture assuming a doubling time of 1.5 h). Inoculate the large flasks with the calculated inoculation volume from the actively growing pre-culture. 7. Grow the large cultures overnight (for the calculated incubation time) at 30  C with shaking at ~200 rpm. Day Three 8. Estimate cell density of the large cultures using a hemocytometer to make sure cells are at mid-log phase (~3  107 cells/ mL). 9. Harvest cells by spinning down at 5000  g for 5 min at 4  C (for centrifuge and rotor options see Subheading 2.1, step 8). Discard the supernatant. 10. Gently resuspend the cell pellet in 25 mL of ice cold ddH2O.Pool the mixtures into 50 mL tubes and spin down at 2000  g for 5 min at 4  C (for centrifuge and rotor options see Subheading 2.1, step 11). Repeat once more. 11. Add 1/100 Protease Inhibitor Cocktail (v/v), 1/100 Solution P and 1/1000 1 M DTT (v/v) to the resuspension buffer (see Subheading 2.1, step 14). 12. Note the volume of the cell pellet in the tubes and resuspend in an equal volume of resuspension buffer with the Protease Inhibitor Cocktail, Solution P, and DTT. Spin down at 2500  g for 20 min. Aspirate the supernatant, leave the pellet as dry as possible. 13. Fill a clean Styrofoam box with liquid nitrogen. Submerge a conical tube rack for 50 mL tubes into the bath. Take a 50 mL tube and poke its cap with a syringe needle to form tiny holes. Unscrew the cap and place the tube into the tube rack inside the liquid nitrogen. Fill the tube with liquid nitrogen.

16

Ilona Nudelman et al.

14. With a spatula, transfer the cell paste into a 20 mL or 50 mL syringe depending on the cell pellet volume. 15. Gently press the plunger and extrude the cell paste directly into the cryo-cooled 50-mL tube forming frozen strings of cell paste or “noodles.” 16. When the tube is full, screw on the cap with holes and pour off the excess liquid nitrogen. Store the frozen noodles at 80  C until cryo-milling. Typically, for 12 L of yeast cell culture, the yield is roughly two 50 mL tubes of noodles. 3.2 Cryo-Milling of Yeast Cells

This protocol is extensively described on the National Center for Dynamic Interactome Research website (https://www.ncdir.org/ protocols/), including a video tutorial (Cryogenic Disruption Using PM100 Ball Mill). Please check the link for updated versions before you perform this procedure. It is crucial to follow the prescribed safety procedures and take extreme care when handling liquid nitrogen. Protective lab clothing and cryoprotective gloves must be used at all times during cryo-milling. 1. For the cryo-milling process, every component that will be in touch with the frozen cells must be kept in liquid nitrogen temperatures. Pre-cool the components for cryo-milling (stainless steel jar, lid, and balls) by immersion in a liquid nitrogen bath (e.g., Styrofoam box). Once the vigorous boiling in the bath ceases, take out the pre-cooled milling jar and add the frozen yeast noodles (100 kDa) diffusing into the nucleus as well as much smaller molecules (90% of the cells are lysed but the nuclei are still intact, centrifuge the nuclei suspension at 1000  g for 10 min at 4  C. 15. Resuspend in 1.5 mL of Nuclei Wash Buffer. 16. Centrifuge the nuclei suspension at 800  g for 10 min at 4  C. 17. Resuspend in the nuclei pellet 1.5 mL of Nuclei Wash Buffer. 18. Centrifuge the nuclei suspension at 800  g for 10 min at 4  C. 19. Resuspend nuclei in 50–150 μL of Diffusion buffer. Nuclei tend to aggregate so mix well by pipetting up and down. 20. Place 2 μL of nuclei suspension onto a glass slide, cover with a coverslip and inspect them on a light microscope (see Note 11). 21. If nuclei are clean move forward with the permeability analysis. If the nuclei solution has large amount of cellular debris or partially broken cells that could affect the assay read out, perform the additional purification steps described in Note 12. 3.5.1 Diffusion Analysis on Isolated Nuclei

1. Mix well by gently pipetting up and down. 2. Place 2 μL of the assay mixture onto a glass slide. 3. Place a 12 mm round coverslip on top. 4. Seal coverslip with clear nail polish. 5. Incubate for 5–10 min (see Note 5). 6. Image and quantify as described for permeabilized cells.

78

4

Marcela Raices and Maximiliano A. D’Angelo

Notes 1. We have successfully used Tris–HCl instead of HEPES buffers for the protocols described here. 2. Because digitonin permeabilization is strongly affected by the density of cells on the coverslips it is critical that all conditions have a comparable cell number/confluency. 3. Every new batch of digitonin should be tested for best permeabilization conditions as small variations in concentration and purity can strongly impact the permeabilization efficiency. Cells with higher cholesterol levels in their plasma membrane might need lower digitonin concentrations of shorter times of permeabilization. 4. Trypan blue can be used instead of fluorescent dextrans when setting up the permeabilization conditions. The Trypan blue dye cannot enter the cell cytoplasm or nucleus unless membranes are compromised. Digitonin-permeabilized cells can be incubated with a 0.2% solution of Trypan blue for 2–3 min and inspected under the light microscope. The correct permeabilization conditions should allow the dye to enter the cytoplasm of most cells (>80–90%) while still being excluded from the nucleus. 5. When analyzing nuclear permeability, only a small number of coverslips should be subjected to the diffusion assay and imaged at the same time to ensure the smallest possible time between imaging different samples. Because longer incubation times will affect the diffusion of some molecules into the nucleus, we recommend to not analyze more than 3–4 coverslips at the time and to spend no more than 5 min per coverslip during the imaging process. 6. The time that it takes dextrans to diffuse into the nucleus can vary considerably between cell types and nuclei. Thus, the dextran incubation time for the diffusion assay should be optimized for each cell type and condition. 7. As more fluorescent dextrans become available, a larger combination of molecular weights can be analyzed. Depending on the fluorophores conjugated to each dextran the laser configurations might vary. 8. We recommend imaging and quantifying 50–100 cells per condition. However, it is important to note that the time that it takes to image all conditions should be minimized to reduce the incubation time difference between the first and the last coverslip to be imaged.

In Vitro Nuclear Permeability Assays

79

9. A larger volume of cells (>300 μL) makes the syringe-mediated lysis of cells in hypotonic buffer more efficient which translates to more nuclei and a cleaner nuclear preparation. 10. It is important not to over-lyse the cells because too many syringe passages will result in the breakage of nuclear membranes and shearing of DNA. 11. A small aliquot of purified nuclei can be stained with DAPI or Hoechst 33242 and their integrity can be analyzed by confocal microscopy. 12. For additional nuclei purification, resuspend nuclei in 3 mL of a 2:1 mixture of Buffers B:A. Carefully place mixture on top of 1.5 mL of Buffer B. Centrifuge for 30 min at 15,000  g. Resuspend in 1 mL of buffer A. Centrifuge at 800  g for 5 min. Resuspend in 50–100 μL of diffusion buffer. References 1. Beck M, Hurt E (2017) The nuclear pore complex: understanding its function through structural insight. Nat Rev Mol Cell Biol 18:73–89 2. Wente SR, Rout MP (2010) The nuclear pore complex and nuclear transport. Cold Spring Harb Perspect Biol 2:1–19 3. Paine PL, Moore LC, Horowitz SB (1975) Nuclear envelope permeability. Nature 254: 109–114 4. Allen TD, Cronshaw JM, Bagley S, Kiseleva E, Goldberg MW (2000) The nuclear pore complex: mediator of translocation between nucleus and cytoplasm. J Cell Sci 113:1651– 1659 5. Timney BL, Raveh B, Mironska R, Trivedi JM, Kim SJ, Russel D, Wente SR, Sali A, Rout MP (2016) Simple rules for passive diffusion through the nuclear pore complex. J Cell Biol 215:7–76 6. Dultz E, Huet S, Ellenberg J (2009) Formation of the nuclear envelope permeability barrier studied by sequential photoswitching and flux analysis. Biophys J 97:1891–1897 7. D’Angelo MA, Raices M, Panowski SH, Hetzer MW (2009) Age-dependent deterioration of nuclear pore complexes causes a loss of nuclear integrity in postmitotic cells. Cell 136: 284–295 8. Shahin V, Ludwig Y, Schafer C, Nikova D, Oberleithner H (2005) Glucocorticoids remodel nuclear envelope structure and permeability. J Cell Sci 118:2881–2889 9. Roehrig S, Tabbert A, Ferrando-May E (2003) In vitro measurement of nuclear permeability

changes in apoptosis. Anal Biochem 318:244– 253 10. Feldherr CM, Akin D (1990) The permeability of the nuclear envelope in dividing and nondividing cell cultures. J Cell Biol 111:1–8 11. Belov GA, Lidsky PV, Mikitas OV, Egger D, Lukyanov KA, Bienz K, Agol VI (2004) Bidirectional increase in permeability of nuclear envelope upon poliovirus infection and accompanying alterations of nuclear pores. J Virol 78: 10166–10177 12. Eftekharzadeh B, Daigle JG, Kapinos LE, Coyne A, Schiantarelli J, Carlomagno Y, Cook C, Miller SJ, Dujardin S, Amaral AS, Grima JC, Bennett RE, Tepper K, DeTure M, Vanderburg CR, Corjuc BT, DeVos SL, Gonzalez JA, Chew J, Vidensky S, Gage FH, Mertens J, Troncoso J, Mandelkow E, Salvatella X, Lim RYH, Petrucelli L, Wegmann S, Rothstein JD, Hyman BT (2018) Tau protein disrupts nucleocytoplasmic transport in Alzheimer’s disease. Neuron 99: 925–940 13. Ferrando-May E, Cordes V, Biller-Ckovric I, Mirkovic J, Gorlich D, Nicotera P (2001) Caspases mediate nucleoporin cleavage, but not early redistribution of nuclear transport factors and modulation of nuclear permeability in apoptosis. Cell Death Differ 8:495–505 14. Terasaki M, Campagnola P, Rolls MM, Stein PA, Ellenberg J, Hinkle B, Slepchenko B (2001) A new model for nuclear envelope breakdown. Mol Biol Cell 12:503–510

80

Marcela Raices and Maximiliano A. D’Angelo

15. Lenart P, Ellenberg J (2006) Monitoring the permeability of the nuclear envelope during the cell cycle. Methods 38:17–24 16. Zhu Y, Liu TW, Madden Z, Yuzwa SA, Murray K, Cecioni S, Zachara N, Vocadlo DJ (2016) Post-translational O-GlcNAcylation is essential for nuclear pore integrity and maintenance of the pore selectivity filter J. Mol Cell Biol 8:2–16 17. Samudram A, Mangalassery BM, Kowshik M, Patincharath N, Varier GK (2016) Passive permeability and effective pore size of HeLa cell nuclear membranes. Cell Biol Int 40:991–998

18. Mohr D, Frey S, Fischer T, Guttler T, Gorlich D (2009) Characterisation of the passive permeability barrier of nuclear pore complexes. EMBO J 28:2541–2553 19. Adam SA, Marr RS, Gerace L (1990) Nuclear protein import in permeabilized mammalian cells requires soluble cytoplasmic factors. J Cell Biol 111:807–816 20. Nishikawa M, Nojima S, Akiyama T, Sankawa U, Inoue K (1984) Interaction of digitonin and its analogs with membrane cholesterol. J Biochem 96:1231–1239

Chapter 5 Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects in Neurodegenerative Diseases Saskia Hutten and Dorothee Dormann Abstract In the recent years, defective nuclear import has emerged as an important pathomechanism of neurodegenerative diseases, particularly in amyotrophic lateral sclerosis (ALS). Here, specific nuclear RNA binding proteins (RBPs) mislocalize and aggregate in the cytoplasm of neurons and glial cells in degenerating brain regions. Bona fide transport assays that measure nuclear import in a quantitative manner allow one to distinguish whether disease-linked RBP mutations that cause cytosolic RBP mislocalization directly result in reduced nuclear import or cause increased cytoplasmic localization of the RBP through other mechanisms. Here we describe the quantitative analysis of nuclear import rates of RBPs using a hormone-inducible system by live cell imaging. Key words Nuclear import, RNA binding protein, Neurodegeneration, Nuclear localization sequence, Hormone-inducible, Live cell imaging, Quantitative transport assay

1

Introduction Import of proteins >30–40 kDa into the nucleus is mediated by members of the importin β-family of nuclear transport receptors (NTRs), also known as importins or karyopherins [1, 2]. Importins bind specific sequences, the so-called nuclear localization sequences (NLSs), in their respective cargo protein and mediate their nuclear import by interacting with components of the nuclear pore complex [2, 3]. Defects in nuclear import have been suggested to contribute to several neurodegenerative diseases, where a number of RNA binding proteins (RBPs) and transcription factors have been found to be mislocalized from the nucleus to cytoplasmic inclusions [4]. It is important to note that this mislocalization can be caused either directly by defects in nuclear import of the respective RBP or indirectly, e.g. by enhanced nuclear export and/or aberrant interactions or aggregation of the RBP in the cytoplasm. Bona fide import assays that allow quantitative analysis of nuclear import rates are well suited to distinguish between these two

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_5, © Springer Science+Business Media, LLC, part of Springer Nature 2022

81

82

Saskia Hutten and Dorothee Dormann

Fig. 1 Workflow for the hormone-inducible import assay

possibilities. Here we describe such a nuclear import assay adapted from [5]. It allows for the analysis of import rates in living cells and can be applied to RBPs as well as other proteins that normally localize to the nucleus and carry a NLS. Fusion of the protein-ofinterest to a fluorescent protein (e.g. GFP) allows for direct visualization by live cell fluorescence microscopy. In addition, the reporter protein is fused to one or two hormone binding domains of the glucocorticoid receptor (GCR), which retains the reporter protein in the cytoplasm of transfected cells. Upon addition of a steroid hormone (dexamethasone), the reporter protein is released from the cytoplasm and imported into the nucleus, by virtue of the NLS in the protein-of-interest (Fig. 1). Differences in nuclear import rates can then be determined by measuring fluorescence intensities in nuclear and cytoplasmic compartments over time. Here, we describe this assay using the ALS-linked RBP Fused in Sarcoma (FUS) as a paradigm. We measure nuclear import rates of reporter proteins containing either FUS wildtype (wt) or an ALS-linked FUS NLS mutant (FUS P525L) known to cause cytosolic

Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects. . .

83

Fig. 2 Typical result of an import assay using the hormone-inducible import assay. (a) Selected time points upon dexamethasone induction in an import assay for GCR2-GFP2-FUS wild-type (FUS WT) versus FUS carrying an ALS-associated point mutation in the NLS (FUS P525L). Bar, 20 μm. (b) Import efficiency for GCR2-GFP2-FUS WT and -FUS P525L, derived from the ratios of the fluorescence intensity of Nucleus and Cytoplasm (N/C) plotted over time, shown as mean of ca. 30 cells
SEM

mislocalization of FUS and early onset ALS [6–8] (Fig. 2). Beyond the analysis of disease-associated mutations, the assay can also be used to assess the involvement of putative import receptors or accessory factors in the import reaction, e.g. by silencing candidate receptors/factors [9, 10], co-expressing specific competitor constructs [11, 12], or adding small molecular inhibitors of importins (Soderholm, 2011). In addition to nuclear import, the GCR-reporter can also be used to study nuclear export (or egress by passive diffusion) out of the nucleus upon washout of dexamethasone in real-time [5, 13].

84

2

Saskia Hutten and Dorothee Dormann

Materials

2.1

Cell Lines

1. HeLa cells or similar adherent cell line.

2.2

Reagents

1. Dulbecco’s Modified Eagle’ Medium (DMEM) or other medium suitable for preferred cell line. 2. Dialyzed Fetal bovine serum (FBS) (see Note 1). 3. Trypsin. 4. PBS. 5. Live cell imaging dishes (see Note 2). 6. Serum- reduced medium (e.g. OptiMEM). 7. Transfection reagent. 8. Protein-of-interest in GCR(2)-GFP(2)-reporter. 9. Live cell imaging medium (e.g. Fluorobrite or CO2 independent medium lacking phenol red, see Note 3). 10. Optional: re-usable, flexible adhesive, e.g. Blu-Tac (see Note 4). 11. Dexamethasone. 12. Ethanol p.a.

2.3 Media for Growth, Transfection and Live Cell Imaging

1. HeLa growth and transfection medium: DMEM supplemented with 10% dialyzed FBS.

2.4 Preparation of Dexamethasone

1. Dexamethasone stock solution: Dissolve 25 mg of dexamethasone in 25.4 mL of Ethanol p.a. to obtain a 2.5 mM stock solution. Aliquot and store at 20  C.

2. HeLa imaging medium: Live cell imaging medium supplemented with 10% dialyzed FBS for imaging >3 h (see Note 5).

2. Dexamethasone working solution: To induce import for live cell imaging, prepare a 2–6 concentrated solution in imaging medium (e.g. if import is to be induced with 5 μM dexamethasone in a final volume of 600 μL, add 1.2 μL of 2.5 mM dexamethasone stock solution in 100 μL imaging medium, to be added to 500 μL imaging medium in the dish) (see Note 6). 2.5

Microscope

2.6

Software

Confocal microscope suitable for live cell imaging, e.g. a Zeiss spinning disk microscope, equipped with environmental chamber (and optional CO2 supply). 1. Image acquisition software (e.g. Zeiss ZEN) with build-in plug-in to acquire a time-lapse for a list of individual fields of view (“point-list”).

Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects. . .

85

2. Fiji/Image J: Download and install “Time Series Analyzer” Tool from Fiji website following provided instructions (https://imagej.nih.gov/ij/plugins/time-series.html). 3. Microsoft Excel. 4. Optional: Graph Pad Prism.

3

Methods

3.1 Cell Culture and Transfection

1. Prepare the cell culture medium (e.g. DMEM for HeLa cells) with 10% dialyzed FBS. 2. Maintain cells (e.g. HeLa cells) for at least two passages in DMEM/ 10% dialyzed FBS (see Note 1). 3. The day before transfection, seed cells into a live cell imaging dish so that they reach a confluency suitable for transfection. 4. Transfect GCR-GFP reporter (we prefer GCR2-GFP2-tagged RBPs, see Notes 7 and 8) with transfection reagent of choice. Omitting antibiotics during transfection will increase transfection efficiency and reduce cell stress. Change medium after 3–5 h to reduce formation of stress granules due to transfection stress. 5. Incubate the cells for 16–24 h to allow for reporter expression (see Note 9).

3.2

Live Cell Imaging

1. Heat up the environmental chamber to 37  C (with optional 5% CO2) 4–5 h before start of experiment (see Note 10). 2. Replace the cell culture medium with imaging medium and fix the dish on the stage either by clamps integrated in the stage (if available) or by using Blue-Tac to avoid moving the dish accidentally during the experiment (see Note 4). 3. Allow the cells to settle on the stage for at least 1 h before the start of time-lapse acquisition (see Note 11). 4. Establish the imaging setup by choosing objective, laser intensity, and exposure time as well as temporal settings for the timelapse experiment (e.g. duration of 40 min with 2.5 min intervals) (see Note 12). 5. Using a built-in plug-in of the microscopy software, establish a list of different fields of view of cells with cytoplasmic localization of the reporter (see Note 13). 6. Start time-lapse acquisition. 7. After the image of the first time point has been acquired, add dexamethasone working solution in small drops to cell medium to induce import of the GCR-reporter (see Note 6).

86

Saskia Hutten and Dorothee Dormann

8. Replace the lid of the cell culture dish with large coverslip to provide easy access to cells without the risk of medium evaporation (see Note 14). 3.3

Analysis

1. Open the time-lapse file as a hyperstack in Fiji. 2. Open the ROI Manager (“Analyze” ! “Tools” ! “ROI Manager”). 3. For each transfected cell, draw a representative region of interest (ROI) for (a) the nucleus (“ROI N”) and (b) the cytoplasm (“ROI C”) using built-in ROI tools (see Note 15). Add ROIs sequentially to the ROI manager by pressing [t] or clicking “Add [t]” in the ROI manager. Rename ROIs N1, C1, N2, C2, etc. in the ROI manager. 4. Draw a ROI outside the area of any cell to determine background fluorescence. Rename this ROI “background”. ROIs can be saved for documentation (“File” ! “Save”). 5. Open “Time Series Analyzer” Tool (“Plugins” ! “Time Series Analyzer”). 6. Mark all ROIs in the ROI Manager and click “Get Average” in the Time Series Analyzer Window. 7. Save generated measurements as an excel file (the simultaneously generated graph can be ignored). 8. Open the Excel file. For each image, subtract the background from the measured value for each ROI (N1, C1, etc.). 9. Use background corrected values to calculate N/C ratio for each cell (N1/C1, N2/C2, etc.) (see Note 16). 10. Obtained N/C ratios can be plotted as mean
SEM using either Excel or Graph Pad Prism. We recommend performing at least three independent experimental replicates with at least 15–20 cells for each sample to determine the statistical significance by calculating the “area under the curve”

4

Notes 1. Standard FBS often contains traces of steroid hormones that can interfere with cytosolic retention of the GCR-reporters. Particularly for RBPs that are characterized by a slow nuclear egress/export rate, this will lead to a higher percentage of cells already displaying nuclear localization of the reporter in the absence of dexamethasone. We find that two passages (i.e. six cell doublings) in growth medium containing commercially available dialyzed FBS is sufficient to reduce this background significantly.

Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects. . .

87

2. Any commercially available live cell imaging dishes that are compatible with the microscope of choice are suitable. If transfection efficiency is high and multiple constructs are supposed to be compared within one experiment, the use of multichambered live cell imaging dishes can be useful to reduce the time that cells need to settle on the stage (see Note 10). 3. Phenol red can increase background fluorescence, hence we recommend the use of phenol red-free imaging medium. If the environmental chamber at the microscope cannot be supplied with CO2, the use of CO2-independent medium is recommended to avoid changes in the pH of the imaging medium over time, as this could result in cellular stress and influence nuclear import rates. 4. Accessing the dish to add dexamethasone bears the risk of accidentally moving the dish, resulting in a loss of focus and/or field of view. We recommend using either stage clamps provided by the microscope manufacturer or, alternatively, the use of a re-usable and formable adhesive such as Blu-Tac to “fix” the dish on stage. 3-point fixation for round dishes or 4-corner fixation of square dishes by small amounts of Ble-Tac are sufficient and can be removed without leaving any residues, even at 37  C, and can be re-used. 5. For imaging beyond 3–4 h, we recommend to add dialyzed FBS to the imaging medium to maintain cell health. 6. We recommend adding dexamethasone as 2–6 concentrated stock corresponding to at least 1/5th of the medium in the dish, in order to ensure even distribution. Prepare dexamethasone working solution freshly every time and keep it warm (ideally inside the environmental chamber). 7. While a single GCR-tag is sufficient for cytosolic retention and a single GFP-moiety for detection of reporter proteins, we found the addition of a second GCR domain and use of a double GFP-moiety helpful to reduce passive diffusion across the nuclear pore complex. As each GCR domain and each GFP-moiety adds ~25 kD to the molecular size of the reporter construct, passive nuclear influx of the reporter is greatly diminished in the absence of a NLS-containing protein-ofinterest. 8. It cannot be excluded that some positively charged amino acids at the N-terminus of the GCR domain might act as a very weak NLS. Comparing the nuclear import of the RBP-reporter to a reporter harboring a stop-codon after the second GFP-moiety (GCR2-GFP2-Stop) helps to distinguish passive diffusion/ background import of the GCR2-GFP2-reporter from active import mediated by the NLS in the RBP or protein-of-interest.

88

Saskia Hutten and Dorothee Dormann

9. Limiting the expression time after transfection can help to reduce the percentage of cells displaying already partially nuclear fluorescence due to RBP reporters interacting with nucleic acids in the nucleus upon disassembly of the nuclear membrane during mitosis. 10. Due to its high metal content, the microscope acts as a considerable “heat sink.” To avoid temperature fluctuations and focus drift during the imaging process, we recommend to heat up the environmental chamber for at least 4–5 h before the start of the experiment. 11. Allowing cells to adjust to the environmental chamber for ~1 h helps to avoid focus drift in time-lapse experiments. 12. When setting up exposure time and laser intensity, one needs to consider that accumulation of the fluorescent reporter in the nucleus (or possibly in nuclear foci) may result in higher fluorescence intensity/pixel than initially observed for the diffuse reporter in the (usually) larger cytoplasm. It is essential to avoid pixel saturation for quantification of fluorescence intensities. Duration and interval of the time-lapse acquisition needs to be chosen such that monitoring of nuclear import with sufficient temporal resolution can be ensured, but bleaching of the reporter is avoided. 13. Choose cells with similar levels of cytoplasmic localization of the reporter in order to compare import dynamics. The number of possible fields of view is limited by the distance between individual points to be visited during the time-lapse experiment and the dynamics of nuclear import reaction. Faster import rates require shorter intervals to allow for sufficient temporal resolution of the import reaction. Allow for enough “gap time” to be able to add dexamethasone in the first interval after starting the experiment. 14. To avoid evaporation of medium during image acquisition the dish should be covered. We recommend using a sufficiently large coverslip/cover glass to cover the dish as it can be easily and quickly retrieved. Alternatively, inverting the lid can also be sufficient. If the size of the environmental chamber allows, a beaker with distilled water can also help to reduce evaporation by maintaining constant moisture level during the imaging process. 15. If cells remain stationary during the imaging process, we recommend using the whole nuclear area for quantification by using the polygon selection tool. As a representative area for the cytoplasm, a band of 1–2 pixel around the nuclear ROI can be applied (“Edit” ! “Selection” ! “Make Band”). If cells move in xy-direction, we have found that a representative

Hormone-Inducible Transport Reporter Assay to Study Nuclear Import Defects. . .

89

circular ROI of 4–5 μm in diameter is sufficient to quantify nuclear import rates if placed centrally, such that it does not leave the nuclear or cytoplasmic compartment at any time during the time-lapse experiment and avoids non-representative fluorescent foci. 16. If the protein-of-interest accumulates in bright nuclear foci and this is affected upon a distinct treatment or mutation in the protein-of-interest, the nuclear import rate might be skewed by measuring the nuclear fluorescence of only a single confocal plane (either in its entity or as representative circular area). In this case, an alternative way to compare import rates is measurement and plotting of the loss in the cytoplasmic fluorescence over time.

Acknowledgments We thank Erin Sternburg and Lara Silva for critical comments on the manuscript. We thank Peter Becker for support and acknowledge Michael Kiebler for access to spinning disc confocal microscope (DFG, INST 86/1581-1 FUGG). This work was supported by the Fritz Thyssen Foundation (Az. 10.19.1.001MN) (to D.D.), the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) within grants DO 1804/1-1, DO 1804/1-2, DO 1804/3-1 (to D.D.), the Munich Cluster for Systems Neurology (EXC2145 SyNergy – ID 390857198 to D.D.) and the Junior Researcher Fund of Ludwig-Maximilians-Universit€at Mu¨nchen (to D.D.). References 1. Cautain B et al (2015) Components and regulation of nuclear transport processes. FEBS J 282(3):445–462 2. Fried H, Kutay U (2003) Nucleocytoplasmic transport: taking an inventory. Cell Mol Life Sci 60(8):1659–1688 3. Soniat M, Chook YM (2015) Nuclear localization signals for four distinct karyopherin-beta nuclear import systems. Biochem J 468(3): 353–362 4. Hutten S, Dormann D (2020) Nucleocytoplasmic transport defects in neurodegeneration - cause or consequence? Semin Cell Dev Biol 99:151–162 5. Love DC, Sweitzer TD, Hanover JA (1998) Reconstitution of HIV-1 rev nuclear export: independent requirements for nuclear import and export. Proc Natl Acad Sci U S A 95(18): 10608–10613

6. Chio A et al (2009) Two Italian kindreds with familial amyotrophic lateral sclerosis due to FUS mutation. Neurobiol Aging 30(8): 1272–1275 7. Dormann D et al (2010) ALS-associated fused in sarcoma (FUS) mutations disrupt Transportin-mediated nuclear import. EMBO J 29(16):2841–2857 8. Kwiatkowski TJ Jr et al (2009) Mutations in the FUS/TLS gene on chromosome 16 cause familial amyotrophic lateral sclerosis. Science 323(5918):1205–1208 9. Hutten S et al (2008) The Nup358-RanGAP complex is required for efficient importin alpha/beta-dependent nuclear import. Mol Biol Cell 19(5):2300–2310 10. Hutten S et al (2009) The nuclear pore component Nup358 promotes transportin-

90

Saskia Hutten and Dorothee Dormann

dependent nuclear import. J Cell Sci 122(Pt 8): 1100–1110 11. Cansizoglu AE et al (2007) Structure-based design of a pathway-specific nuclear import inhibitor. Nat Struct Mol Biol 14(5):452–454 12. Kosugi S et al (2008) Design of peptide inhibitors for the importin alpha/beta nuclear

import pathway by activity-based profiling. Chem Biol 15(9):940–949 13. Ederle H et al (2018) Nuclear egress of TDP-43 and FUS occurs independently of Exportin-1/CRM1. Sci Rep 8(1):7084

Chapter 6 Subcellular Fractionation Suitable for Studies of RNA and Protein Trafficking Biljana Culjkovic-Kraljacic and Katherine L. B. Borden Abstract The nuclear pore complex is the major conduit for trafficking between the nucleus and cytoplasm. Nuclear import and export of both proteins and RNAs represent important functional steps for many biological processes. One of the major means to study NPC activity and the nuclear and cytoplasmic distribution of proteins and RNAs is through biochemical fractionation. Here, we describe detailed methods to generate high quality nuclear and cytoplasmic fractions simultaneously capturing RNA and proteins which can be used subsequently for a wide array of biochemical characterizations including proteomics and next generation sequencings. Key words Nuclear- Cytoplasmic Fractionation, RNA, Protein, Localization

1

Introduction Many RNAs and proteins can translocate or shuttle between the nucleus and cytoplasm depending on conditions. The cellular structure that permits the majority of trafficking is the nuclear pore complex (NPC). There are a wide variety of functional impacts based on a protein or RNA’s subcellular location and studying such trafficking also provides a means to study NPC activity under different conditions. There are two main methods to quantify the relative distribution of proteins between these two compartments: microscopy and biochemical fractionation. Here, we focus on this latter method. Isolation of nuclear and cytoplasmic fractions enables one to not only quantify the distribution of proteins or RNAs under different conditions but also physically isolate specific factors or complexes from each fraction for further study. Here we describe a fast and straightforward protocol for obtaining nuclear and cytoplasmic fractions that can be used for simultaneous isolation of RNAs and proteins for various biochemical studies (including next generation sequencing and proteomics). This procedure maintains integrity of protein–protein as well as protein–RNA

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_6, © Springer Science+Business Media, LLC, part of Springer Nature 2022

91

92

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

complexes and can be used for interaction studies such as protein co-immunoprecipitation studies (co-IPs) as well as RNA-immunoprecipitation studies (RIP). This protocol has been successfully used for cellular fractionation of a wide variety of normal and cancer cell lines including mouse and human fibroblasts, osteosarcoma, breast cancer, various human myeloid and lymphoid cell lines [1–10] and thus the protocol described here should also be applicable to other mammalian cell types. Commonly used methods of subcellular fractionation are based on density gradient centrifugation that are lengthy and can lead to varying degree of nuclear leakage and degradation of RNAs and proteins. The protocol described here is a modified version of one described in the 1980’s [1, 11]. This protocol is based on utilizing different mild detergents to obtain high quality nuclear and cytoplasmic fractions. First, cells are lysed in a buffer that contains Nonidet P-40 (IGEPAL CA-630) which solubilizes the plasma membrane while maintaining the integrity of the nuclear membrane. After a brief centrifugation, the cytoplasmic fraction is contained in the supernatant, and two other nonionic detergents (Sodium deoxycholate and Tween 40) are employed to strip membranous organelles such as the endoplasmic reticulum (ER) and mitochondria from the nuclei, while keeping the nuclear membrane intact. Resulting nuclei and cytosolic fractions can be used for RNA isolation, preparing protein lysates for Western blot (WB) analysis, proteomic studies, co-IP, RIP studies, etc. [1–10, 12–16] or extended to further fractionation of nuclear components (i.e. nucleoplasm, nucleolar fractions, and chromatin bound components, etc.). The quality of the obtained nuclear and cytoplasmic fractions should always be confirmed by WB analysis employing antibodies against different subcellular marker proteins (Table 1) and/or by semiQ-RT-PCR (or RT-qPCR) using appropriate primers for small nuclear RNA (snRNA) U6 (nuclear localization) and transfer RNAs (tRNAs) such as tRNAMet or tRNALys (predominantly cytoplasmic localization). We describe fractionation and validation strategies here.

2

Materials

2.1 Cellular Fractionation

1. Prepare Diethyl pyrocarbonate (DEPC)-treated water by diluting DEPC to 0.1% (v/v) solution in double distilled (dd) H2O and incubate at room temperature overnight. Autoclave to destroy the unreacted DEPC. 2. Prepare Protease Inhibitors Cocktail: 1 mg/mL Pepstatin, 1 mg/mL Leupeptin, 2 mg/mL Aprotinin, 100 mM PMSF, 1.6 mg/mL Benzamidine, 1 mg/mL Phenanthroline. This

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

93

Table 1 Protein markers for monitoring subcellular fractionation Localization

Marker

Mw

Antibody source

Nucleus

PolR2A Histone 2B Nopp140 Lamin A

220 kDa 14 kDa 140 kDa 70 kDa

Santa Cruz Abcam Santa Cruz SIGMA

Cytoplasm

α-Tubulin GAPDH MEK-1 Cytochrome C Calreticulin

50 kDa 36 kDa 43 kDa 12 kDa 55 kDa

SIGMA Santa Cruz Santa Cruz BD Abgent

provides protection from aspartyl, cysteine, serine and threonine peptidases, and metalloproteases. Use it at 1:1000 dilution in all buffers. Commercially available protease inhibitor cocktails such Complete™ EDTA-free Protease Inhibitor Cocktail from Roche Biochemicals can also be used instead. 3. Prepare Lysis buffer B (LBB): 10 mM Tris (pH ¼ 8.4), 140 mM NaCl, 1.5 mM MgCl2, 0.5% Nonidet P-40/IGEPAL CA-630 (see Note 1), and store at 4  C. Add fresh (just prior to use), 1 mM DTT and 100 U/mL RNaseOUT (or similar RNase inhibitor), and the protease inhibitor cocktail above. For studies where phosphorylation status is examined, phosphatase inhibitors should also be included such as Sodium Fluoride (inhibits Ser/Thr and acidic phosphatases), Sodium Orthovanadate (inhibits Tyr and alkaline phosphatases), and β-Glycerophosphate (inhibits Ser/Thr phosphatases). Phosphatase inhibitor cocktails are also available from various manufacturers. 4. Prepare Detergent stock solution: 3.3% (w/v) Sodium Deoxycholate, 6.6% (v/v) Tween 40 in DEPC treated water (see Note 2). Do not autoclave. It will dissolve quickly at 37  C. 5. Sonicator such as Sonic Dismembrator Model 500, Fisher, Max Output 400 W. 2.2

RNA Isolation

1. Mix Chloroform and Isoamyl alcohol (IAA) in a 24:1 ratio. A ready to use mix is commercially available from various manufacturers. 2. 50 TAE Buffer: 2 M Tris, 50 mM EDTA disodium salt, 1 M glacial acetic acid. 3. To prepare 1% agarose gel, mix 1 g of agarose with 100 mL 1TAE Buffer in a microwavable flask, cover the top with saranwrap and make a few perforations with 10 μL tip (this

94

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

prevents the solution to boil over and allows excess of steam to escape). Microwave the solution for 3–4 min, until the agarose is completely dissolved (check by swirling the flask). Let solution cool to ~50  C (5–10 min) and pour the agarose gel into the tray with comb in place. 4. To prepare 2RNA Formaldehyde loading dye mix: 7.2 mL Deionized Formamide, 2.6 mL 37% Formaldehyde, 1 mL Glycerol, 1.6 mL 10 MOPS buffer, 0.8 mL saturated Bromphenol Blue solution, 1.8 mL DEPC water, and 75 μL 10 mg/ mL Ethidium bromide solution. Aliquot and store at 20  C. 10MOPS Buffer: 200 mM MOPS pH 7.0, 10 mM EDTA, 50 mM Na Acetate. 5. M-MLV Reverse Transcriptase kit. 6. Random Hexamers and Oligo(dT). 7. RNaseOUT (40 U/μL) or similar RNase inhibitor. 8. 10 mM dNTPs mix (deoxyribonucleotide triphosphates). 9. Taq DNA Polymerase with Buffer. 10. Primers for U6 snRNA and tRNAMet in human cells: U6Fw: CGCTTCGGCAGCACATATAC, U6Rv: AAAATATGGAACGCTTCACGA, tRNAMetFw: AGC AGA GTG GCG CAG CGG, tRNAMetRv: GAT CCA TCG ACC TCT GGG TTA. 11. PCR Thermocycler. 12. UV-Vis spectrophotometer. 13. TRIzol™ and TRIzol™ LS Reagent (TRIzol™ LS Reagent is the formulation of TRIzol™ reagent more suitable for RNA isolation from liquid samples). TRIzol™ is acid-guanidinium thiocyanate-phenol based reagent designed for one-step RNA isolation [17–19], and many other similar products are commercially available from different companies (such as TRI Reagent, QIAzol, RNAzol, RiboZol, etc.). 14. Isopropyl alcohol. 15. Prepare 70% Ethanol by mixing 70 mL of 100% ethanol with 30 mL of DEPC treated water. 16. UV Transilluminator. 2.3 Protein Lysates and WB Analysis

1. Prepare 12% SDS Polyacrylamide gel solution by mixing 3.3 mL ddH2O, 2.5 mL 1.5 M Tris Buffer pH 8.8, 4 mL 30% Acrylamide: Bis-acrylamide solution (37.5:1), and 100 μL of 10% SDS solution. Add 100 μL of 10% Ammonium persulfate and 5 μL of TEMED (N,N,N0 ,N0 -tetramethylethylene diamine) immediately before pouring the gel between glass plates. Let gel to polymerize for 1 h.

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

95

2. Prepare 4% SDS Stacking gel solution by mixing 6 mL ddH2O, 2.5 mL 0.5 M Tris Buffer pH 6.8, 4 mL 30% Acrylamide: Bis-acrylamide solution (37.5:1), and 100 μL of 10% SDS solution. Add 100 μL of 10% Ammonium persulfate and 5 μL of TEMED immediately before pouring the gel. 3. 10 SDS Running Buffer: 0.25 M Tris, 1.92 M Glycine, 1.0% SDS pH 8.3 4. 6 Laemmli loading buffer 0.3 M Tris–HCl, 0.6 M DTT, 10% SDS, 0.06% Bromophenol blue, 30% Glycerol. Add 50 μL β-Mercaptoethanol per 1 mL before use. 5. 10 TBS: 0.2 M Tris, 0.15 M NaCl, pH 7.4. 6. Polyvinylidene difluoride (PVDF) membrane. 7. Vertical mini electrophoresis system or similar system. 8. Electrotransfer unit.

3

Methods

3.1 Cellular Fractionation

All steps should be carried out at 4  C. 1. Collect 3–5  107 mammalian cells and wash twice in ice cold PBS and by spinning at 1200 rpm (300–400  g) for 5 min (see Note 3). 2. Resuspend cell pellets by gently pipetting up and down in 1 mL of LBB until no clumps are visible and transfer lysates to 1.5 mL microfuge tubes. It usually takes around 30–50 up-down strokes using 1 mL pipette for U2Os cells (see Note 4). 3. Spin lysates for 3 min at 1000  g at 4  C in benchtop microcentrifuge. 4. Carefully transfer supernatant (Sn, this is the cytoplasmic fraction) to a fresh 2 mL or 15 mL tube. 5. Resuspend pellet in 1 mL of LBB by gentle pipetting up and down 4–5 times, and transfer to a round bottom polypropylene tube. Add slowly (drop by drop) one-tenth of the volume (100 μL) of Detergent stock solution, under slow vortexing (this will prevent nuclei from clumping) and incubate on ice for 5 min (see Note 5). 6. Transfer to a clean 1.5 mL microfuge tube and spin for 3 min 1000  g at 4  C in benchtop microcentrifuge. 7. Transfer supernatant (this is post-nuclear fraction containing the endoplasmic reticulum, mitochondria, and fragments of other cytoplasmic organelles) to add it to the cytoplasmic fraction or retain it separately for analysis of the post-nuclear fraction.

96

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

8. Resuspend pellet in 1 mL of LBB and spin for 3 min 1000  g at 4  C in benchtop microcentrifuge (see Note 6). 9. Discard Sn. Pellet represents intact nuclei that can be used directly for RNA isolation (see next section) and/or preparation of nuclear lysates. 3.2 RNA Isolation, cDNA Synthesis, and semiQ-PCR

1. Add equal volume of TRIzol™ LS Reagent to the cytoplasmic fraction (combined supernatants from step 4 and  from previous section), and proceed to step 3.

3.2.1 RNA Isolation

2. Resuspend nuclei (pellet from step 9 previous section) in 1 mL of TRIzol™ Reagent. This might take a few minutes since nuclei are quite sticky and can be difficult to resuspend (see Note 7). 3. Add 200 μL of Chloroform: IAA per 1 mL of TRIzol™ or TRIzol™ LS Reagent, mix well by inverting tubes (25 times) and centrifuge 15 min at 12,000  g. 4. Transfer aqueous phase (avoiding interphase which appears as a white ring between upper aqueous phase and red organic phase on the bottom) to a fresh tube. RNAs from this step can be isolated by alcohol precipitation or using commercially available kits. (a) To precipitate RNAs, add equal volume of Isopropyl alcohol to the aqueous phase, mix well and precipitate RNAs by incubating for a few hours or overnight at 20  C. Centrifuge 30 min 16,000  g, the pellet should be visible on the bottom of the tube. Discard supernatant, and wash the pellet by adding 1 mL 70% Ethanol, vortex briefly (5–10 s, maximum speed, to detach pellet from the bottom) and spin for 5 min 16,000  g. Discard Ethanol, leave pellets to air dry, and resuspend in DEPC or any other RNase free water (see Note 8). (b) We also successfully used Direct-Zol RNA Purification kits from Zymo Research. To isolate RNAs from Trizol™ or TRIzol™ LS Reagent directly or aqueous phase follow instructions manual. Other similar kits should work as well. 5. Measure absorbance A260 and A280 to estimate concentration and purity of isolated RNAs using a NanoDrop (or any other UV-Vis spectrophotometer) (see Note 9). 6. Estimate RNA integrity by running RNA samples on 1% agarose gel (or use an Agilent Bioanalyzer or similar equipment). Many RNAs contain substantial secondary structures and thus should be denatured prior to loading on a gel (see Note 10). Mix 300–500 ng of RNA samples with formaldehyde loading dye, incubate 15 min at 65  C, chill on ice for 5 min, spin

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

97

Fig. 1 (a) Nuclear (N) and cytoplasmic (C) RNAs isolated from U2Os cells using this method analyzed on 1% agarose gel. (b) SemiQ-PCR for U6 snRNA and tRNAMet for nuclear and cytoplasmic RNAs

quickly, and load onto gel. Run gel for 30 min at 100 V and visualize on UV transilluminator. Typical results for RNAs isolated from nuclear and cytoplasmic RNAs are shown on Fig. 1a. 3.2.2 cDNA Synthesis and semiQ-PCR

Protocol for cDNA synthesis described here is for using M-MLV (Moloney Murine Leukemia Virus) reverse transcriptase (Invitrogen; see Note 11). 1. Combine 300 ng of DNAse treated RNA (see Note 8) samples with 1 μL of 10 mM dNTPs solution, 0.5 μL of 50 μM Random Hexamers, 1 μL of 50 μM Oligo(dT), and RNase free water to a final volume of 12 μL. Incubate at 65  C for 5 min and immediately transfer to chill on ice. 2. Combine (per sample) 4 μL of 5 First Strand Buffer (from M-MLV reverse transcriptase kit) with 2 μL of 0.1 M DTT 1 μL of RNaseOUT (40 U/μL) and 1 μL of M-MLV reverse transcriptase (200 U), mix well and add to RNA mix. 3. Incubate for 10 min at 25  C, 50 min at 37  C, and 15 min at 70  C (to inactivate the reverse transcriptase). 4. Dilute cDNAs 20 with Glycogen solution (40 ng/mL in ultrapure water). 5. Prepare PCR reactions using U6 or tRNAMet primers by mixing Taq Polymerase Buffer to a final concentration 1 (usually supplied by manufacturer as 10 Buffer), 200 μM dNTPs

98

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

(each), 2 μM forward and reverse primers, 1 U Taq Polymerase, and 5 μL of diluted cDNA in 20 μL PCR reaction. Cycling conditions: 3 min 95  C initial denaturation, 30 cycles (for snRNA U6) and 28 cycles (for tRNAMet) of 30 s 95  C, 30 s 58  C and 30 s 68  C, followed by final extension of 5 min at 68  C (see Note 12). 6. Analyze PCR products on 2% Agarose gel. Typical results are shown in Fig. 1b. 3.3 Preparation of Protein Lysates and Western Blot Analysis

1. Add 300–500 μL LBB buffer to nuclear pellet (see Note 13) and resuspend with microtip and sonicate three cycles of 6 s bursts (with 30 s pause between each burst) using microtip at 20–25% power (Max Output 400 W), where power output is the key variable (see Note 14). 2. To clarify protein lysates (cytoplasmic, nuclear, and total cell lysate) centrifuge them for 10 min at 10,000  g. Transfer the supernatants to clean tubes. 3. Measure concentration of lysates using BCA (or any other) method. 4. For Western blot analysis, boil 10–20 μg of each sample in Laemmli buffer 5–8 min at 95  C, resolve on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and transfer to polyvinylidene difluoride (PVDF) membrane (see Note 15). 5. Incubate membranes in 5% milk in TBS–Tween 20 to block non-specific binding. Dilute primary antibodies (Table 1) in 5% milk (in TBS–Tween 20), and incubate membranes overnight at 4  C with gentle rocking. 6. Wash membranes three times 5–10 min in TBS–Tween 20 and incubate with appropriate HRP-conjugated secondary antibodies diluted in 5% milk (in TBS–Tween 20). 7. Wash membranes three times 5–10 min in TBS–Tween 20, once with TBS and visualize blots using HRP substrate. Typical results are shown in Fig. 2 (see Note 16).

4

Notes 1. Note that NP-40 is not the abbreviation for Nonidet P-40. Unfortunately, many biochemical protocols published from 1960s refer to Nonidet P-40 as NP-40. Nonidet P-40 (originally manufactured by the Shell Chemical Company) was discontinued in the early 2000s, and the most chemically similar substitute commercially available today is IGEPAL CA-630. Nonidet P-40 and IGEPAL CA-630 are milder detergents, and are used to dissolve cytoplasmic membranes only, while

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

99

Fig. 2 Western blot analysis of total (T), cytoplasmic (C), and nuclear (N) lysates isolated from U2Os cells using nuclear and cytoplasmic markers demonstrating fraction quality

NP-40 is often used to disrupt all cellular membranes, including the nuclear membrane. Store buffers containing IGEPAL CA-630 light protected. 2. RNA molecules are very susceptible to degradation either by RNases or non-specific cleavage in the presence of divalent cations. The most common sources of RNAse are microbial growth in solutions and human skin. Thus, gloves should always be worn while handling RNA. To reduce RNase contamination, buffers and solutions should be treated with DEPC or prepared in DEPC treated ddH2O (i.e. Tris and HEPES buffers cannot be treated by DEPC). All buffers used for preparation of RNA samples and lysates for RIPs should be supplemented with RNase inhibitors such as RNaseOUT (Invitrogen, or other commercially available RNase inhibitor). Glassware and equipment (such as sonicator probes or homogenizers) should be treated twice with RNAZap (available from different suppliers) or 0.2% SDS, and rinsed thoroughly in DEPC treated water. 3. Grow adherent cells to 70–90% confluency, and suspension cells to the density as recommended in specification sheet for the particular cell line. Some cell types are more sensitive to crowding than others, which can change the physiological state of cells, and may affect experimental outcomes. We find that trypsinization of adherent cells gives better results when it comes to fractionation. Scraping of cells can lead to damage of nuclei which leads to clumping (as a result of leaked DNA) and contamination of fractions (nuclear components into cytoplasmic fractions and vice versa).

100

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

Typical cellular fractionations require 10–50  106. For RNA or protein isolation 10–20  106 cells are usually enough (in case of U2Os cells or mouse fibroblasts this equals about two 15-cm plates). For preparation of nuclear extracts for immunoprecipitations (including RIPs), we recommend starting with at least 4–5 of 15-cm plates (at least 50  106 cells). Collect cells in 50 mL conical tubes and centrifuge them at 300–400  g at 4  C for 5 min. Decant the supernatant and gently suspend the cells in ice cold 1 PBS to wash them (40–50 mL). Repeat this step. At the point of second PBS wash, take an aliquot (one-tenth to one-fifth) of resuspended cells for isolation of total RNAs and another aliquot for isolation of total protein lysates. Keep the cell pellets on ice at all times to slow metabolic processes and to inhibit RNA and protein degradation. 4. Gentle pipetting is key to successful fractionation, which helps to maintain intact nuclei specially during the initial cell lysis step. If starting with lower number of cells (>1  107) up to 30 up-down pipette strokes will be sufficient for complete lyses of the cells. Number of up-down pipette strokes should also be determined for each cell type. Suspension cell lines such as DLBCL cell lines (e.g. OCI-LY1, SUDHL6, DOHH2) and AML cell lines (such as MonoMac6, THP-1, K562, etc.) are more fragile and require less pipette strokes (10–15 up-down pipetting). On the other hand, if nuclear fractions show cytoplasmic contamination, it is recommended to decrease the number of cells per fractionation or increase the number of pipette strokes. Thus, this step may need some optimization for different cell types. 5. Set your vortex to a minimal speed and touch mode. 6. This step serves to wash detergent solution from a previous step, thus total resuspension of nuclei is not necessary. Take a small aliquot of this suspension (5–10 μL) and spot it on a glass slide to check under the phase contrast microscope (for comparison the suspension of whole cells should be used). Nuclei should appear as oval and rounded without cytoplasmic attachments. Shrunk or irregular shaped nuclei might indicate damaging of the nuclear membranes. In this case, consider a more gentle resuspension of nuclei and decrease the incubation time in Detergent solution. 7. If nuclei are used to make lysates for WB analysis or RIP, aliquots of these lysates can be used to isolate RNAs using TRIzol™ LS Reagent. If lysates obtained from crosslinked cells or nuclei are used for RNA isolation, heating TRIzol™ LS Reagent mixture for 15 min at 65  C is needed to efficiently disrupt crosslinked RNA-protein complexes.

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

101

Samples in TRIzol™ or TRIzol™ LS Reagent can be stored at 80  C for at least several weeks. 8. For precipitation of small amounts of RNA, polyacryl carrier (2–8 μL, MRC) or glycogen (0.01–1 mg/mL) should be added to the RNA solution to facilitate precipitation. Do not over-dry RNA pellets, it will make them difficult to resuspend. For downstream applications such as RT-PCR, RNAs should be DNase treated. For DNase treatment several commercial kits which include inactivation and cleaning reagents are available (i.e. Turbo DNase Free from Invitrogen). Kits for RNA isolation on columns usually include DNase treatment as well. 9. Ratio of A260/280 for pure RNAs should be ~2.0 for pure RNAs. Abnormal ratios indicate contamination with proteins or TRIzol™ components [20, 21]. Contaminants in RNA samples can differentially inhibit the reverse transcription reaction lead to misleading results from RT-qPCR. The absorbance A260 and A280 of RNA samples does not give information about RNA quality (integrity), which can be determined only by analyzing samples on an agarose gel or using Agilent Bioanalyzer. 10. Homemade or any commercially available Formaldehyde loading dye can be used at a final concentration of 1. To monitor integrity of RNAs, 1% agarose gel in 1TAE buffer can be used. If separation of RNAs on agarose gel is used for Northern blot, Agarose gels containing Formaldehyde in MOPS running buffer should be used [21]. 11. Other Reverse Transcriptases or commercially available kits that utilize random primers can also be used for cDNA synthesis. Efficiency of reverse transcription can vary, and thus optimization of PCR steps might be needed (such as increase or decrease of number of PCR cycles). To control for genomic DNA contamination, mock reactions that contain RNA samples and lack M-MLV reverse transcriptase should be included. 12. If PCR products for U6 snRNA and tRNAMet appear saturated after analysis on agarose gel electrophoresis (i.e. very strong signals and no/very slight differences between fractions) consider lowering the number of PCR cycles (4–5 cycles less), or start with less cDNA. If PCR products are faint, increase the number of cycles. After confirming purity and integrity of RNA samples, cDNAs can be obtained to measure levels of specific mRNAs in nuclear and cytoplasmic fractions by quantitative Real-Time PCR. We successfully used this protocol to measure Cytoplasmic/Nuclear ratio of specific RNAs resulting from enhanced

102

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

RNA export upon eIF4E overexpression or repression with drugs or RNAi strategies [2, 4–10, 12, 13, 15, 16]. Several articles extensively cover quantitative PCR strategies and the details of how to design primers and calculate expression of specific RNAs [22–26]. 13. Volume of buffer should be about 5 times volume of nuclear pellet. Nuclei can be lysed in any other lysing buffer depending on downstream application. For co-IPs and RIPs homogenize nuclei using Dounce homogenizer with tight pestle (2–3 rounds of 50 strokes) in buffer of choice (i.e. RIPA or NT-2 buffer). Nuclei can be also crosslinked for co-IPs and RIPs before lysis using paraformaldehyde. To crosslink nuclei, first wash nuclear pellets two times in 1PBS (nuclei are very sticky and can get stuck in the pipette tip, thus gentle and slow pipetting is recommended for this step). To crosslink resuspend nuclei in 1% paraformaldehyde (made in PBS), incubate with rotation 10 min at room temperature and quench with 0.15 M Glycine for 5 min. After washing three times in 1 PBS, crosslinked nuclei can be lysed in buffer of choice by sonication as described in Subheading 3.3. 14. Sonication time for the nuclei can vary for different cell types and the type of sonicator probe. Volume of buffer used should not be too high in order to obtain efficient disruption of nuclei. On the other hand, small volume of buffer and too high power can cause foaming during sonication which in turn can lead to protein degradation. Viscous lysates indicate inefficient lysis, in this case, increase sonication time (i.e. number of rounds), and consider increasing the volume of lysis buffer (dilute and aliquot lysates in multiple microfuge tubes with 0.5 mL of lysate per tube). Extracts should be kept on ice at all time during experiments and can be flash-frozen and stored at 80  C. 15. Nitrocellulose membranes can also be used. We typically use 12% SDS-PAGE to cover various sizes of all markers for cellular fractionation (see Table 1), and cut membranes horizontally in 3–4 strips to simultaneously detect proteins with different molecular weight from the same blots. 16. This protocol typically produce high quality fractions. For instance, nuclear fractions are considered clean if there is little or no evidence of cytoplasmic markers such as α-Tubulin, GAPDH or MEK kinase, ER markers such as calreticulin or mitochondrial markers such as Cytochrome C using WB analysis (Fig. 1) while these are features of the cytoplasmic fraction. Conversely, the nuclear fraction should reveal the presence of nuclear-specific proteins such as PolR2A, Lamin A, Nopp140,

Nucleo-Cytoplasmic Fractionation of Mammalian Cells

103

or histone H2B, while these should be absent from the cytoplasmic fraction. Note that Actin and GAPDH are considered to be cytoplasmic proteins, but they can also be detected in the nucleus at least in some cell types [27–30]. References 1. Daar IO, Maquat LE (1988) Premature translation termination mediates triosephosphate isomerase mRNA degradation. Mol Cell Biol 8(2):802–813 2. Topisirovic I et al (2005) Eukaryotic translation initiation factor 4E activity is modulated by HOXA9 at multiple levels. Mol Cell Biol 25(3):1100–1112 3. Topisirovic I et al (2003) The proline-rich homeodomain protein, PRH, is a tissuespecific inhibitor of eIF4E-dependent cyclin D1 mRNA transport and growth. EMBO J 22(3):689–703 4. Topisirovic I et al (2003) Aberrant eukaryotic translation initiation factor 4E-dependent mRNA transport impedes hematopoietic differentiation and contributes to leukemogenesis. Mol Cell Biol 23(24):8992–9002 5. Culjkovic B, Topisirovic I, Skrabanek L, RuizGutierrez M, Borden KL (2005) eIF4E promotes nuclear export of cyclin D1 mRNAs via an element in the 3’UTR. J Cell Biol 169(2): 245–256 6. Culjkovic-Kraljacic B, Baguet A, Volpon L, Amri A, Borden KLB (2012) The oncogene eIF4E reprograms the nuclear pore complext to promote mRNA export and oncogenic transformation. Cell Rep 2(2):207–215 7. Culjkovic-Kraljacic B et al (2016) Combinatorial targeting of nuclear export and translation of RNA inhibits aggressive B-cell lymphomas. Blood 127(7):858–868 8. Urtishak KA et al (2019) Targeting EIF4E signaling with ribavirin in infant acute lymphoblastic leukemia. Oncogene 38(13): 2241–2262 9. Pettersson F et al (2011) Ribavirin treatment effects on breast cancers overexpressing eIF4E, a biomarker with prognostic specificity for luminal B-type breast cancer. Clin Cancer Res 17(9):2874–2884 10. Assouline S et al (2009) Molecular targeting of the oncogene eIF4E in acute myeloid leukemia (AML): a proof-of-principle clinical trial with ribavirin. Blood 114(2):257–260 11. Nevins JR (1980) Definition and mapping of adenovirus 2 nuclear transcription. Methods Enzymol 65(1):768–785

12. Davis MR, Delaleau M, Borden KLB (2019) Nuclear eIF4E stimulates 30 -end cleavage of target RNAs. Cell Rep 27(5):1397–1408.e4 13. Topisirovic I et al (2009) Molecular dissection of the eukaryotic initiation factor 4E (eIF4E) export-competent RNP. EMBO J 28(8): 1087–1098 14. Topisirovic I et al (2009) Stability of eukaryotic translation initiation factor 4E mRNA is regulated by HuR, and this activity is dysregulated in cancer. Mol Cell Biol 29(5):1152–1162 15. Volpon L et al (2016) Importin 8 mediates m7G cap-sensitive nuclear import of the eukaryotic translation initiation factor eIF4E. Proc Natl Acad Sci U S A 113(19):5263–5268 16. Culjkovic B, Topisirovic I, Skrabanek L, RuizGutierrez M, Borden KL (2006) eIF4E is a central node of an RNA regulon that governs cellular proliferation. J Cell Biol 175(3): 415–426 17. Chomczynski P, Sacchi N (1987) Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162(1):156–159 18. Weber K, Bolander ME, Sarkar G (1998) PIG-B: a homemade monophasic cocktail for the extraction of RNA. Mol Biotechnol 9(1): 73–77 19. Chomczynski P (1993) A reagent for the single-step simultaneous isolation of RNA, DNA and proteins from cell and tissue samples. BioTechniques 15(3):532–534, 536–537 20. Glasel JA (1995) Validity of nucleic acid purities monitored by 260nm/280nm absorbance ratios. BioTechniques 18(1):62–63 21. Sambrook JF, Russell D (2001) Molecular cloning: a laboratory manual (3-volume set). Cold Spring Harbor Laboratory Press, Cold Spring Harbor 22. Wang Y, Zhu W, Levy DE (2006) Nuclear and cytoplasmic mRNA quantification by SYBR green based real-time RT-PCR. Methods 39(4):356–362 23. Wilhelm J, Pingoud A (2003) Real-time polymerase chain reaction. Chembiochem 4(11): 1120–1128

104

Biljana Culjkovic-Kraljacic and Katherine L. B. Borden

24. Larionov A, Krause A, Miller W (2005) A standard curve based method for relative real time PCR data processing. BMC Bioinformatics 6: 62 25. Deprez RHL, Fijnvandraat AC, Ruijter JM, Moorman AFM (2002) Sensitivity and accuracy of quantitative real-time polymerase chain reaction using SYBR green I depends on cDNA synthesis conditions. Anal Biochem 307(1): 63–69 26. Derveaux S, Vandesompele J, Hellemans J (2010) How to do successful gene expression analysis using real-time PCR. Methods 50(4): 227–230 27. Butera G et al (2019) Regulation of autophagy by nuclear GAPDH and its aggregates in cancer

and neurodegenerative disorders. Int J Mol Sci 20(9):2062 28. Virtanen JA, Vartiainen MK (2017) Diverse functions for different forms of nuclear actin. Curr Opin Cell Biol 46:33–38 29. Carlile GW, Tatton WG, Borden KLB (1998) Demonstration of a RNA-dependent nuclear interaction between the promyelocytic leukaemia protein and glyceraldehyde-3-phosphate dehydrogenase. Biochem J 335:691–696 30. de Lanerolle P, Cole AB (2002) Cytoskeletal proteins and gene regulation: form, function, and signal transduction in the nucleus. Sci STKE 2002(139):pe30

Chapter 7 Localizing Total mRNA in Plant Cells Geraint Parry Abstract Visualizing the location of the total cellular mRNA pool can be important to understand how different genes affect cellular physiology. Over the past decade researchers investigating RNA processing, nuclear transport and the function of the nuclear pore complex have used in situ hybridization protocol to visualize and quantify the accumulation of the total mRNA pool within the plant cell nucleus. Key words In situ hybridization, Nucleus, mRNA, Nuclear pore complex, Nuclear transport

1

Introduction Messenger RNA (mRNA) is the critical intermediate that transfers the genetic information encoded in DNA through to the generation of functional proteins. Transcription within the cell nucleus generates thousands of mRNAs that then travel through the nuclear pore complex to the cytosol where they are recognized by ribosomes, by whose action strings of amino acids are assembled into higher-order proteins. Obtaining an understanding of where mRNA is expressed and localised within a tissue and cell provide significant insights into the function of the underlining genes. In situ hybridization (in situ) is the technique that has been historically used to reveal the positioning of specific mRNAs either through radioactive or non-radioactive labeling of short anti-sense DNA probes. This technique can be used with both tissue sections or in whole mount samples [1, 2]. In situs allow the identification of an expression domain, usually at the resolution of a small number of cells or to a single cell. Over recent years a refined in situ technique has been developed that uses multiple fluorescently labeled probes (up to 40 probes) targeted to a single mRNA. This allows the localization of single RNAs with subcellular resolution [3]. Each of these in situ techniques aim to visualize the expression of a single gene. However, certain experimental circumstances

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_7, © Springer Science+Business Media, LLC, part of Springer Nature 2022

105

106

Geraint Parry

require visualization of the total cellular mRNA pool. Over the past decade these have included experiments that are investigating mechanisms of RNA processing and transport or the factors that control nuclear export, including the function of the nuclear pore complex [4–11]. Generally, this technique uses a fluorescently labeled anti-sense poly(dT) probe to evaluate the localization of all the mRNA within a plant cell. Engler et al [12] first published on this technique but more recently Gong et al (1995), who were investigating the function of the DEAD box RNA helicase LOS4, have modified this protocol. In this study, poly(A) RNA export is blocked in los4–2 cells at warm temperatures. Wildtype and los4–2 mutant plants were grown at 4  C for 2 months or 22  C for 2 weeks. In situ hybridization with fluorescein-labeled oligo(dT) probe was performed with seedlings growing at 4  C (a) or 22  C (b) as shown in Fig. 8 panel A in Gong et al. (2005) [4]. In comparison to wildtype cells, mutant cells show that mRNA accumulates within the nucleus. Parry (2014) showed that different NPC mutants, nup160 and nup62, which have similar whole plant phenotypes, show different accumulation of total mRNA. In nup160 mutants the total mRNA accumulates to the nucleus while in nup62 mutant this accumulation is normal [8]. Similarly when looking at the function of factors involved in nuclear export, Sørensen et al. (2017) showed that tex1 and mos11 single mutants show wildtype levels of nuclear mRNA accumulation but that tex1mos11 double mutants accumulate mRNA in their nuclei [10]. Therefore, this protocol is able to define different cell biological phenotypes between seemingly phenotypically similar mutants. This protocol has only been used in Arabidopsis so might need to be adapted in order to correctly fix tissues in other organisms. The protocol has been used equally well to assess total mRNA accumulation in both root or leaf tissue.

2

Materials 1. Glass vials and tissue culture dishes (see Note 1). 2. Forceps and razor blade for tissue manipulation. 3. Buffer 1: 120 mM NaCl, 7 mM Na2HPO4, 3 mM Na2PO4, 2.7 mM KCl. 80 mM EGTA, 0.1% Tween, 10% dimethylsulfoxide (DMSO) (see Note 2). 4. Paraformaldehyde (PFA). 5. Xylene. 6. 100% Ethanol. 7. 100% Methanol.

Localizing Total mRNA in Plant Cells

107

8. 12-well plastic tissue culture dish. Other sizes of culture dishes may be used but will require a larger volume of reagent. 9. Perfect Hyb Plus Buffer (Sigma-Aldrich). 10. Oligo(dT)25 DNA primer labeled with Fluorescein (see Note 3). 11. 20 SSC: 20 SSC is 3 M NaCl and 300 mM sodium citrate (see Note 4). 12. 10% sodium dodecyl sulfate (SDS) (see Note 4). 13. Glass microscope slides. 14. Appropriate cover slips. 15. Antifade mounting material (e.g. VectaShield, Vectorlabs). 16. Antifade mounting material with DAPI (e.g. VectaShield, Vectorlabs). 17. Propidium iodide .

3

Methods Carry out all procedures at room temperature unless otherwise specified. 1. Grow Arabidopsis seedlings using your standard growth conditions. 2. Prepare required tissue samples from leaf and/or root tissue (see Note 5). 3. Place samples into glass container with Buffer 1 + 5% PFA (freshly made with PFA). Depending on the container use enough liquid to ensure tissue was fully submerged. Incubate at room temperature with slow shaking for >30 min (see Note 6). 4. Remove liquid and wash with slow shaking for 5 min with 100% methanol (2) and 100% ethanol (2) for a total of four washes. Completely cover tissue samples with wash solutions. 5. Remove liquid and wash for 30 min with slow shaking with 1:1 ethanol:xylene (see Note 7). Always use a glass vial for this wash. 6. Remove liquid and wash for 5 min with 100% ethanol. 7. Remove liquid and wash for 5 min with 1:1 methanol:Buffer 1 (without PFA) mix. 8. Remove liquid and postfix for Buffer 1 with 5% PFA for >30 min.

108

Geraint Parry

9. Remove liquid and wash for 5 min with Buffer 1 (without PFA) (2) and for 5 min with Perfect Hyb Plus Buffer (1) (see Note 8). 10. Tissue samples are pre-hydridized at 50  C in fresh Perfect Hyb Plus Buffer for >1 h (see Note 9). 11. Add 1 pmol μL 1 oligo(dT) probe labeled with fluorescein (see Note 10) to Perfect Hyb Plus Buffer and incubate with tissue sample overnight at 50  C in the dark (see Note 11). 12. Remove liquid and incubate at 50  C with 2  SSC, 0.1% SDS wash solution for 1 h and then 0.2  SSC, 0.1% SDS wash solution for 20 min. 13. Samples are transferred to fresh 0.2xSSC (no SDS) and can be held at 4  C for weeks prior to imaging (see Note 12). 14. Samples are processed for size in order to fit under available cover slips (see Note 13) and mounted in VectaShield Antifade mounting material +/ DAPI depending on the visualization requirements (see Note 14). 15. Samples should be evaluated using appropriate filters to visualize position of nucleus (with DAPI or propidium iodide) alongside the expression of the fluorescein-labeled probe. Both confocal and epifluorescence microscopes have been successfully used for visualizing fluorescein expression. 16. If necessary, quantification of mRNA accumulation can be achieved by selecting a single point or set area in the DAPI image, one within the nucleus and another in the cytosol adjacent to the nucleus (Fig. 1). The pixel intensity is then measured at the equivalent positions in the fluorescein image. This will give a value that represents the amount of fluorescein expression in the nucleus compared to cytoplasm. It is important to select the location for measurement using the DAPI image to rule-out the chance of any bias that might occur if the position was selected in the fluorescein image. 17. Within a single experiment multiple nuclei should be visualized across independent tissue samples. Experiments should be repeated on at least three occasions to verify any findings.

4

Notes 1. Plastic vials will be melted by xylene. Use of a shallow glass vial or dish allows for easier manipulation of tissue samples. 2. Buffer 1 without DMSO and PFA can be prepared in advance. DMSO and PFA are added prior to the experiment. 3. This type of labeled oligo(dT) can be ordered as from most oligonucleotide suppliers.

Localizing Total mRNA in Plant Cells

DAPI

Wildtype

a

FLO b

#1

c

nup85

#2

#4 #3

109

#2a #1a

d

#4a #3a

Fig. 1 Epifluorescence microscope images (63) of cells in the root elongation zone from 7do seedlings treated with an oligo(dT)-Fluorescein (FLO) probe (b, d) and post-stained with DAPI (a, c). Nuclei in nup85 mutant roots show an increase in FLO accumulation when compared to wildtype. FLO accumulation is quantified by setting single-point measurement positions in images of DAPIstained nuclei (a, #1, #2; c, #3, #4) and then measuring pixel intensity in an identically positioned FLO-image (b, #1a, #2a; d, #3a, #4a). The pixel intensity is compared between #1a and #2a (or #3a and #4a) to give a value for FLO accumulation in the nucleus. Scale bar- 10 μm. (Figure adapted from Parry (2014) [8])

4. Dilute to working concentrations from 20XSSC and 10% SDS stocks. 5. Most studies either use small leaves ( 0.05. P-values obtained with paired t-tests

Using Single Molecule RNA FISH to Determine Nuclear Export and. . .

119

2. Preheat diluted probe solution to 37  C in a thermomixer. 3. Remove as much methanol as possible without disturbing heads at the bottom of the tube and then add 1 mL of Wash Buffer A (see Note 7). Incubate for 5 min on an orbital shaker. 4. Remove Wash Buffer A and then add 100 μL of the preheated diluted probe to the sample and incubate in the dark at 37  C overnight (see Note 8) in an incubator with gentle rocking or in a thermomixer with gentle mixing (around 300 rpm). Day 3

1. Remove diluted probe and add 1 mL of Wash Buffer A. Incubate for 30 min at 37  C in the dark (dark incubator with gentle rocking or thermomixer again). 2. Remove Wash Buffer A and add Hoechst DNA stain at (1: 1000) diluted in Wash Buffer A. Incubate for 30 min at 37  C in the dark with gentle rocking. 3. Remove diluted Hoechst and add 1 mL of Wash Buffer B. Incubate for 5 min at room temperature on an orbital shaker in the dark (covered with aluminum foil). 3.4

Mounting

1. To mount or transfer samples, cut the thinnest part of the tip off of a 1 mL pipet tip (so that it is wide enough to transfer heads without damaging them). Rinse the tip several times in PBS and then using the wide tip, carefully transfer larval heads to a glass dissecting dish filled with PBS for salivary gland dissection, with minimal light under a dissecting scope. Carefully dissect off salivary glands, keeping the pair attached at the hook if possible (glands are very fragile here so it can be harder to dissect any fat off at this stage). 2. Once glands are separated, carefully place them onto a cleaned slide in a drop of Vectashield (about 10–12 μL) and using forceps, carefully arrange them so they are not overlapping (see Note 9). 3. Carefully place a coverslip onto the mounted salivary glands. This works best when you hold the coverslip with clean forceps, and first allow one edge of the coverslip to come in contact with Vectashield, then slowly lower the rest of the coverslip down, letting the Vectashield slowly come in contact with the glass without allowing bubbles to form. 4. Apply a dot of clear nail polish carefully to each corner of the coverslip. Allow slides to dry for ~30 min in the dark before completely sealing with nail polish. 5. Slides can be imaged immediately or stored at 4  C for imaging the next day (see Note 10).

120

Jennifer R. Aleman et al.

3.5 Confocal Microscopy

1. Acquire 3D stacks by confocal microscopy using a high numerical aperture, high magnification objective. We have conducted our confocal imaging at room temperature on a Leica TCS SP8 Confocal using a PL APO 63
/1.40 Oil objective at 1.6
Zoom, using Type F Immersion Oil Leica 11513859, and Leica Software LAS-X 3.3. 2. Imaging settings used were as described in Little and Gregor, 2018 [22]. Briefly, the most essential settings are described below in steps 3–7. 3. Voxel size: Images should be obtained with pixels that are 76
76 nm with a z increment between 250 nm and 420 nm for the most efficient detection of diffraction-limited objects (mRNAs) by our custom MATLAB software. 4. Section thickness: Image stacks range from 10 μM to 20 μM in the Z-plane. Wild-type salivary gland nuclei are typically around 8–10 μM thick but stacks can be up to 15 μM in Z depending on the mutant background. 5. Scanner settings and image acquisition: smFISH signal from salivary glands covers an extremely wide range of fluorescence intensities. Whereas single mRNAs are dim, endoreplication generates hundreds of copies of individual loci on polytene chromosomes, and thus actively transcribed genes can contain thousands of nascent transcripts. The wide range in signal requires low-noise photon detectors capable of spanning a dynamic range of 1000-fold or more. The Leica SP8 is equipped with low-noise hybrid (HyD) detectors capable of acting as either traditional photomultiplier tubes or as singlephoton detectors. Placing these detectors in photon-counting mode provides optimal results. The limited number of fluorophores per single mRNA requires multiple scan accumulations at slow scanning speeds with long pixel dwell times (roughly 10 μs per pixel). For multi-channel imaging, it is preferable to alternate channels with every scanned line. Additionally, to accommodate the wide dynamic range, data must be collected as 12- or 16-bit images. 6. Laser power: this must be determined on a case-by-case basis per experiment. It is important to use as little laser power as possible to avoid photobleaching but to provide sufficient excitation so that signal can be visualized above background or autofluorescence signal. Furthermore, care must be taken to avoid saturation of the detectors from the extremely bright fluorescence found in the nascent transcription of salivary glands, which can contain thousands of nascent transcripts for highly active genes. A good practice is to collect a test image at the brightest focal plane and ensure that the brightest pixel is less than 50% of the maximum attainable value.

Using Single Molecule RNA FISH to Determine Nuclear Export and. . .

3.6

Analysis

121

Image analysis described here was carried out using a custom MATLAB code. Code is available upon request from Dr. Shawn Little ([email protected]). Image Segmentation of Cytoplasm and Nucleoplasm in Three Dimensions 1. 3D stacks of Hoechst staining are segmented into volumes representing nucleoplasm and cytoplasm of individual cells. A balanced histogram thresholding approach is applied to find a threshold separating Hoechst-labeled from non-labeled voxels after taking the log of voxel intensity across all voxels in the 3D stack.

2. From the resulting binary masks, each slice is filtered in 2D with a circular averaging filter with a radius of 10 pixels, then morphologically closed with a disk-shaped structuring element to create contiguous nuclei in 3D composed of voxels unequivocally assigned to the nucleus. 3. The resulting 3D nuclear masks are morphologically closed to create contiguous objects comprised of pixels assigned to Hoechst-labeled nucleoplasm. 4. To ensure that only the nucleus interior is used for density calculations, 3D nuclear volumes are eroded by 3% of the mean nuclear diameter. The maximum distance between edges of a bounding box describing each nucleus in three dimensions is found, and the average across nuclei used as the mean nuclear diameter. 5. Cytoplasmic volumes corresponding to each cell are determined by dynamic dilation of nuclear masks using increments of 3% of nuclear diameter, extending each individual nuclear mask using a disk-shaped structuring element until dilated objects overlap. A new 3D mask is created by dilation of the original nuclear mask using a disk with whose radius is half of the first distance that results in object overlap. The interior of these objects, corresponding to a nuclear volume padded (by dilation) with an additional 6% of nuclear diameter, is removed from the interior of each nuclear mask. This generates the mask corresponding to the cytoplasmic volume. Removal of the additional 6% of nuclear diameter is necessary due to uncertainty in the exact boundary between nucleoplasm and cytoplasm. The 6% shell therefore corresponds to a volume of uncertainty, which was disregarded when calculating mRNA densities. Masks are visually inspected to ensure nucleus and cytoplasmic assignments are to the correct cells. All subsequent calculations are performed on the density of mRNAs found in the cytoplasmic and nucleoplasmic volumes.

122

Jennifer R. Aleman et al.

mRNA Quantification

1. Individual mRNAs appear as diffraction-limited puncta, the detection of which is performed as previously described using difference-of-Gaussian filtering [24]. Detection generates a list of centroid coordinates for each mRNA object. 2. The mean intensity of single mRNAs is estimated by fitting a 2D circular Gaussian with the radius of the point spread function to the mean image of all cytoplasmic mRNAs, as described [25]. 3. Densities of mRNA puncta in the cytoplasm or nucleoplasm are calculated as the count of objects in corresponding 3D masks divided by the volume of the mask. The ratios of derived nucleoplasmic to cytoplasmic mRNA densities are used to represent mRNA export functionality and compared among genotypes. An example of obtained ratios for control vs. Mtordepleted cells is shown in Fig. 1c, d. 4. Quantifying transcriptional output of nascent loci requires two approaches, the choice of which depends on whether transcribing loci were easily identifiable. For most polytenized cells, the region of the nucleus containing transcribing endoreplicated loci is readily apparent in the FISH images (as in Fig. 1a). This volume is easily segmented from the surrounding image using histogram thresholding. For cells in which volumes containing the transcribing loci could not be identified by histogram analysis (as in Fig. 1b), the brightest objects with fit intensities greater than four times the mean and present within 500 microns of each other are selected as actively transcribing loci. The selection of putative transcribing sites by both methods is visually inspected for accuracy. 5. After selection, the total fluorescence intensity within volumes containing transcribing signal is determined by summation. 6. Fluorescence offset (background) is determined by averaging the signal in voxels surrounding the selected volumes within a distance of 250 nm. 7. The fluorescence corresponding to signal is determined by subtracting the background signal per voxel multiplied by the volume of the transcribing volumes. 8. The resulting values are then divided by the mean intensity of individual mRNAs determined in step 2, yielding an estimate of instantaneous transcriptional activity. For example and further details, see also [21].

Using Single Molecule RNA FISH to Determine Nuclear Export and. . .

4

123

Notes 1. Probe designer settings are as detailed in [22]—Make sure to indicate Drosophila as the organism. Leave the masking number set to 5 and the max number of probes set to 48 (this is the optimal number of probes needed, but we have had experience visualizing good signal with as little as 35 probes). Oligo length should be set to 20 with 2 nt spacing which is the preset on the webpage. Once the software generates the oligos for the sequence you provided, go through this list and discard and oligos with GC content below 40% (want GC content to be between 40% and 60%). 2. You can read more about Stellaris Dyes and Modifications here: https://www.biosearchtech.com/suppor t/education/ stellaris-rna-fish/dyes-and-modifications-for-stellaris . Probes are shipped as dry pellets and should be stored at 4  C until ready to be resuspended. 3. It is essential to pick larva that are at the wandering third instar stage from minimally crowded vials– not too early or too late third instar (if developmental timing during the wandering period is not being used as a variable). Too early would be larvae that are moving around the vial quickly and that are likely too young and will have smaller salivary glands and polytene nuclei. Too late would be larvae that are higher up the vial, moving minimally with anterior sphericals protruded and will have salivary glands that are starting to fill with too much salivary gland “glue” protein (these glands have a glassy appearance), which will lead to a lot of cytoplasmic background when imaging making single RNA molecules in the cytoplasm too hard to detect. The perfect third instar larvae would be moving minimally (mostly mouth movements), but not yet at pre-pupa stage. Females tend to have larger salivary glands, if possible to pick them for your experimental set up. 4. The dissection procedure can take some practice to precisely cut larvae open at the right position. If you cut too posteriorly, you will have too much of the gut attached and it will be hard to invert the cuticle inside out because you will have too far of a distance to flip the cuticle. 5. The number of animals needed to proceed with FISH is up to you and depends on the sample size needed for your particular experiment. In the beginning, we suggest 5–6 animals per condition as in the washing and hybridization steps, you tend to lose some tissue along the way, depending on how rough you are. 5–6 animals ensure that you will have about three usable salivary glands to mount in the end. If you have trouble

124

Jennifer R. Aleman et al.

with tissue retrieval at the mounting steps, we suggest adding more dissected animals to your tubes to start with and also being gentler with your pipetting during wash steps. 6. Methanol vs. Ethanol permeabilization—Biosearch recommends permeabilization in 70% ethanol. We usually permeabilize solely in 100% methanol in order to quench endogenous GFP signal in the salivary glands (a product of the genotype of larvae we often use). We tested lower percentages of methanol, methanol for up to 30 min and then switching into 70% ethanol for storage, but storing the sample overnight in 100% methanol was necessary to quench GFP signal and did not interfere with the subsequent hybridization of FISH probes. Permeabilization in 70% ethanol should be sufficient if your larvae do not contain GFP, but some optimization may be required. 7. Be careful with washes—do not quickly shoot wash liquid directly and forcefully on heads, add wash liquid to Eppendorf tubes gently and at an angle in order to avoid directly shooting liquid on heads. Wash steps are the points at which you may lose the most tissue from heads—rough washing leads to detachment of tissue (especially salivary glands). It is hard to salvage tissue once it is detached and floating in wash buffer in the tube. 8. Probe incubation should be at least 4 h; however, we mainly conduct experiments with an overnight probe incubation (16 h). Optimization for each new probe set may be required if you want to incubate for less time. 9. It is best to use hooked forceps (forceps that have been bent so the two fine tips don’t come to a close anymore so that you can pick up the glands in a small bubble of liquid and then transfer this bubble to the Vectashield drop on the slide). If attempting to move the salivary glands without hooked forceps, try grabbing at the top of the gland near the place where the two glands are attached. The glands are very fragile at this step and can break into pieces if you try to grab them at any other position. 10. Slide storage after mounting—it is best to store slides coverslip down and image 1 day after mounting. This will allow the salivary glands to settle to the bottom of the cover slip and allow for less slight tissue movement when acquiring Z-stacks on an inverted confocal microscope. Slides may be stored for up to a month at 4  C for later imaging, but the sooner you image, the better results you will obtain (within 1–2 weeks is the optimal time period).

Using Single Molecule RNA FISH to Determine Nuclear Export and. . .

125

References 1. Kuhn TM, Capelson M (2019) Nuclear pore proteins in regulation of chromatin state. Cell 8:1414 2. Raices M, D’Angelo MA (2017) Nuclear pore complexes and regulation of gene expression. Curr Opin Cell Biol 46:26–32 3. Ibarra A, Hetzer MW (2015) Nuclear pore proteins and the control of genome functions. Genes Dev 29:337–349 4. Pascual-Garcia P, Debo B, Aleman JR, Talamas JA, Lan Y, Nguyen NH, Won KJ, Capelson M (2017) Metazoan nuclear pores provide a scaffold for poised genes and mediate induced enhancer-promoter contacts. Mol Cell 66(63–76):e66 5. Ibarra A, Benner C, Tyagi S, Cool J, Hetzer MW (2016) Nucleoporin-mediated regulation of cell identity genes. Genes Dev 30:2253– 2258 6. Pascual-Garcia P, Jeong J, Capelson M (2014) Nucleoporin Nup98 associates with Trx/MLL and NSL histone-modifying complexes and regulates Hox gene expression. Cell Rep 9: 433–442 7. Light WH, Freaney J, Sood V, Thompson A, D’Urso A, Horvath CM, Brickner JH (2013) A conserved role for human Nup98 in altering chromatin structure and promoting epigenetic transcriptional memory. PLoS Biol 11: e1001524 8. Liang Y, Franks TM, Marchetto MC, Gage FH, Hetzer MW (2013) Dynamic association of NUP98 with the human genome. PLoS Genet 9:e1003308 9. Light WH, Brickner DG, Brand VR, Brickner JH (2010) Interaction of a DNA zip code with the nuclear pore complex promotes H2A.Z incorporation and INO1 transcriptional memory. Mol Cell 40:112–125 10. Kalverda B, Pickersgill H, Shloma VV, Fornerod M (2010) Nucleoporins directly stimulate expression of developmental and cell-cycle genes inside the nucleoplasm. Cell 140:360– 371 11. Capelson M, Liang Y, Schulte R, Mair W, Wagner U, Hetzer MW (2010) Chromatinbound nuclear pore components regulate gene expression in higher eukaryotes. Cell 140:372–383 12. Wente SR, Rout MP (2010) The nuclear pore complex and nuclear transport. Cold Spring Harb Perspect Biol 2:a000562 13. Terry LJ, Wente SR (2007) Nuclear mRNA export requires specific FG nucleoporins for

translocation through the nuclear pore complex. J Cell Biol 178:1121–1132 14. Galy V, Gadal O, Fromont-Racine M, Romano A, Jacquier A, Nehrbass U (2004) Nuclear retention of unspliced mRNAs in yeast is mediated by perinuclear Mlp1. Cell 116:63–73 15. Guglielmi A, Sakuma S, D’Angelo MA (2020) Nuclear pore complexes in development and tissue homeostasis. Development 147: dev183442 16. Cho UH, Hetzer MW (2020) Nuclear periphery takes center stage: the role of nuclear pore complexes in cell identity and aging. Neuron 106:899–911 17. Aksenova V, Smith A, Lee H, Bhat P, Esnault C, Chen S, Iben J, Kaufhold R, Yau KC, Echeverria C et al (2020) Nucleoporin TPR is an integral component of the TREX2 mRNA export pathway. Nat Commun 11: 4577 18. Vinciguerra P, Iglesias N, Camblong J, Zenklusen D, Stutz F (2005) Perinuclear Mlp proteins downregulate gene expression in response to a defect in mRNA export. EMBO J 24:813–823 19. Iovine MK, Watkins JL, Wente SR (1995) The GLFG repetitive region of the nucleoporin Nup116p interacts with Kap95p, an essential yeast nuclear import factor. J Cell Biol 131: 1699–1713 20. Hieronymus H, Yu MC, Silver PA (2004) Genome-wide mRNA surveillance is coupled to mRNA export. Genes Dev 18:2652–2662 21. Aleman JR, Kuhn TM, Pascual-Garcia P, Gospocic J, Lan Y, Bonasio R, Little SC, Capelson M (2021) Correct dosage of X chromosome transcription is controlled by a nuclear pore component. Cell Rep 35(11):109236 22. Little SC, Gregor T (2018) Single mRNA molecule detection in drosophila. Methods Mol Biol 1649:127–142 23. Kennison JA (2008) Dissection of larval salivary glands and polytene chromosome preparation. CSH Protoc 2008:pdb.prot4708 24. Little SC, Tikhonov M, Gregor T (2013) Precise developmental gene expression arises from globally stochastic transcriptional activity. Cell 154:789–800 25. Little SC, Sinsimer KS, Lee JJ, Wieschaus EF, Gavis ER (2015) Independent and coordinate trafficking of single Drosophila germ plasm mRNAs. Nat Cell Biol 17:558–568

Part III Functional Analysis of Nucleoporins

Chapter 9 Analysis of Nucleoporin Function Using Inducible Degron Techniques Vasilisa Aksenova, Alexei Arnaoutov, and Mary Dasso Abstract Over the last decade, the use of auxin-inducible degrons (AID) to control the stability of target proteins has revolutionized the field of cell biology. AID-mediated degradation helps to overcome multiple hurdles that have been encountered in studying multisubunit protein complexes, like the nuclear pore complex (NPC), using classical biochemical and genetic methods. We have used the AID system for acute depletion of individual members of the NPC, called nucleoporins, in order to distinguish their roles both within established NPCs and during NPC assembly. Here, we describe a protocol for CRISPR/Cas9-mediated gene targeting of genes with the AID tag. As an example, we describe a step-by-step protocol for targeting of the NUP153 gene. We also provide recommendations for screening strategies and integration of the sequence encoding the Transport Inhibitor Response 1 (TIR1) protein, a E3-Ubiquitin ligase subunit necessary for AID-dependent protein degradation. In addition, we discuss applications of the NUP-AID system and functional assays for analysis of NUP-AID tagged cell lines. Key words CRISPR/Cas9, Nuclear Pore Complex, Auxin-Inducible Degradation

Abbreviations AID GOI gRNA HDR NE NPC NUP NUP-AID POI TIR1

Auxin-Induced Degron Gene of Interest guide RNA Homology Directed Repair Nuclear Envelope Nuclear Pore Complex Nucleoporin/NPC constituent protein Nucleoporin-AID fusion protein Protein of Interest Transport Inhibitor Response 1 protein/ubiquitin ligase subunit

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_9, © Springer Science+Business Media, LLC, part of Springer Nature 2022

129

130

1

Vasilisa Aksenova et al.

Introduction Nucleoporins (NUPs) are components of the Nuclear Pore Complex (NPC), a large multisubunit protein complex which perforates the nuclear envelope (NE) to form mammoth channels that permit selective exchange of macromolecules between the cytosol and the nucleus. The NPC has a highly complex modular structure and is composed of multiple copies of ~30–35 distinct NUPs. Most nucleoporins are essential for cell viability [1], and mutations in NUP genes have been well documented to cause a plethora of human diseases [2]. However, studying NUPs has been challenging for several reasons. First, NUPs are very long-lived proteins that, once synthesized, are maintained in some tissues throughout an organism’s lifespan [3]. Therefore, conventional RNAi- or CRISPR-mediated silencing of expression for a NUP-of-interest may not immediately result in the desired loss of function. Second, as cells in multicellular organisms undergo mitosis, their NPCs disassemble into NUP subcomplexes and disperse in the cytoplasm, while the NE is dissolved to allow spindle formation. At the end of mitosis, the NE and NPCs are rebuilt in newly formed daughter cells. Hence, prolonged removal of a particular NUP may disrupt both the NPC’s interphase function and the mitotic NPC disassembly/reassembly cycle, so it may be difficult to distinguish how either of these aspects contribute toward observed pleiotropic phenotypes. Third, silencing the expression of a particular NUP in rapidly dividing culture cells may lead to destabilization of either the whole NPC or particular groups of nucleoporins. For instance, removal of NUP133 by siRNA causes a simultaneous loss of multiple nucleoporins and reduced density of NPCs on the nuclear envelope [4], again making it difficult to distinguish how individual nucleoporins contribute toward observed phenotypes. Here, we discuss an approach for studying nucleoporins and other long-lived proteins that can decisively overcome these barriers: CRISPR/Cas9-mediated targeting of endogenous NUP genes with Auxin-Inducible Degron (AID) tags and subsequent analysis of NUP function by their Auxin-controlled acute depletion. Nishimura and colleagues developed AID tags as an elegant degradation system to control protein stability conditionally [5]. The system was applied for the rapid depletion of CENP-H in chicken DT40 cells [5]. This system was subsequently used to deplete other proteins [6], and has become a popular tool to study protein functions in many model systems, including yeast [7, 8], worms [9], flies [10], and mammalian cell lines [11–14]. One of the most frequently used approaches in mammalian cell lines is gene knock-out followed by overexpression of AID-tagged fusions of the targeted protein [15]. However, overexpression of NUPs has been commonly observed in cancer cells [2, 16] raising

Analysis of Nucleoporin Function Using Inducible Degron Techniques

131

questions about the physiological impact of NUP overexpression. Depletion of NUPs in nondividing primary or terminally differentiated human cells can also be efficiently achieved through the Trim Away method [17]. However, antibody-based methods are limited by antibody specificity to target proteins and this strategy is not facile for large scale experiments such as mass-spectrometry or nextgeneration sequencing. Therefore, AID-mediated degradation of endogenously tagged proteins is a highly advantageous alternative for studying various NUP functions. We provide a detailed protocol for endogenous targeting of nucleoporins with the AID tag and for Auxin-mediated rapid degradation of AID-tagged NUPs. In particular, we provide guidance for AID gene targeting, screening strategies, E3-Ubiquitin ligase subunit TIR1 integration, functional assays, and applications of the AID system to studies of NPC organization. In this protocol, we use NUP153 as an example, but this approach could be applied to virtually any protein of interest (POI).

2 2.1

Materials Plasmids

1. pX330-U6-Chimeric_BB-CBh-hSpCas9 (Addgene, 42230), codon-optimized SpCas9 and chimeric guide RNA expression plasmid. 2. pcDNA5-EGFP-AID-BubR1 (Addgene, 47330), codonoptimized full-length Auxin-Inducible Degron. 3. pBABE TIR1-9Myc (Addgene, 47328), codon-optimized OsTIR1. 4. pmiRFP670-N1 (Addgene, 79987), codon-optimized monomeric near-infrared fluorescent protein miRFP670. 5. pQCXIB (Clontech), blasticidin-resistance gene.

2.2

Cells

1. Dukes’ type C, colorectal adenocarcinoma, DLD-1 (ATCC CCL-221). 2. Colorectal carcinoma, HCT 116 (ATCC CCL-247).

2.3 Software and Online Tools

1. SnapGene. 2. NEBuilder Assembly tool https://nebuilderv1.neb.com 3. CRISPR Design Tool https://crispr.zhaopage.com 4. UniProt data resource, https://www.uniprot.org 5. NCBI BLAST, https://blast.ncbi.nlm.nih.gov/Blast.cgi

2.4 Reagents and Consum-ables

1. DMEM. 2. McCoy’s 5a Medium Modified. 3. Opti-MEM I Reduced Serum Medium.

132

Vasilisa Aksenova et al.

4. Fetal Bovine Serum (e.g. Atlanta Biologicals, S11550). 5. Trypsin-EDTA. 6. PBS, pH 7.4. 7. Water, Ultra-Pure. 8. Penicillin-Streptomycin. 9. GlutaMAX Supplement. 10. ViaFect (Promega, E4982). 11. Hygromycin. 12. Blasticidin. 13. DMSO. 14. Auxin (e.g. Indole-3-acetic acid sodium salt, Sigma, I514810G). 15. Counting Slides (e.g. Bio-Rad, 145-0011). 16. Cell Culture Plates; 12-, 24-, 6-well. 17. Lumox multiwell, 24-well (e.g. Sarstedt, 94.6110.024). 18. BbsI-HF, supplied with CutSmart® Buffer. 19. Nde I, supplied with CutSmart® Buffer. 20. NotI-HF, supplied with CutSmart® Buffer. 21. Alkaline Phosphatase, Calf Intestinal. 22. Wizard® Genomic DNA Purification Kit. 23. Gibson assembly master mix. 24. T4 Polynucleotide Kinase (PNK), supplied with 10
T4 Polynucleotide Kinase Reaction Buffer. 25. T4 DNA Ligase, supplied with 10
T4 DNA Ligase Reaction Buffer. 26. High Efficiency NEB® 5-alpha Competent E. coli. 27. SOC outgrowth medium. 28. Platinum™ SuperFi™ DNA Polymerase.

3

Method The CRISPR/Cas9-NUP-AID system consists of the plant hormone Auxin and a plant-specific ubiquitin E3 ligase subunit (TIR1), as well as the endogenous SCF ligase complex and proteasome machinery [5]. Nishimura and colleagues applied the Auxinmediated degradation system to mammalian cells, and the AID system has further been successfully adapted to address many important issues. One of the major caveats of this method is that AID tag addition has been observed to destabilize target proteins even in the absence of Auxin in some cases, presumably through the

Analysis of Nucleoporin Function Using Inducible Degron Techniques

133

mass action of co-expressed TIR1. To circumvent this problem, the system has been modified to control TIR1 expression by either using the doxycycline-inducible system [18], by expressing repressive ARF-PB1 domain [19], or replacing OsTIR1(WT) with the mutant form of OsTIR1(F74G) and auxin with a ligand, 5-Ph-IAA [20]. Here, we describe an alternative strategy to lower TIR1 levels by linking TIR1 expression with the product of the essential but low-expressed RCC1 gene. This approach allows incorporation of the AID degron at an endogenous gene-of-interest (GOI), fostering expression of POI-AID fusion protein at levels that are similar to endogenous POI levels before auxin. We applied this approach for targeting multiple nucleoporins [21], including major members of the Y-complex, cytoplasmic filaments, channel and basket nucleoporins, as well as other non-nucleoporin genes. In the present method, we describe CRISPR/Cas9-AID targeting of NUP153, an NPC basket component protein, at its C-terminus, as well as essential functional assays required after the generation of NUP-AID cell lines. An overview of the CRISPR/Cas9-NUP-AID degradation scheme and the targeting protocol are shown in Fig. 1 and Fig. 2, respectively. 3.1

gRNA Cloning

We used the pX330 vector (Addgene, 42230) to drive simultaneous expression of S. pyogenes Cas9 (SpCas9) protein and guide RNA (gRNA) sequence in the same cell. To increase the efficiency of DSB (Double-Strand Breaks) formation, we use two pX330 SpCas9-gRNA plasmids that recognize different 3’ NGG PAM sequences around the targeting site (Fig. 3a). NUP153 gRNA design and cloning incorporated the following steps: 1. Import accession number with the position of a NUP153 gene location (NC_000006.12:17615035.0.17706925) from the NCBI database to SnapGene software. For targeting NUP153 gene use gene ID: 9972 and GRCh38.p13 assembly: https://www.ncbi.nlm.nih.gov/gene/9972. 2. To choose the guide RNA (gRNA) sequence, we use the CRISPR Design Tool. Select a region of ~300 nt around the STOP codon, and indicate classic NGG PAM type, target gRNA size 18 bp, and human hg38 assembly to analyze the sequence. https://crispr.zhaopage.com/task.php?id¼5611e87d591 dba87928f79db58521a8e High- and low-quality gRNAs are marked in green and red colors, respectively. Select a gRNA with a high gRNA score, a low number of off-targets and a position close to the STOP codon. Avoid gRNAs with off-targets at Transcription Start Sites (TSS) and inside the exon sequences of off-target genes.

134

a

Vasilisa Aksenova et al.

b

untreated NUP153-NG-AID

RCC1-iRFP670

t=0h

auxin NUP153-NG-AID

RCC1-iRFP670

t=2h

Fig. 1 AID targeting and rapid degradation of NUP153. (a) A scheme of auxin mediated degradation. NUP153, endogenously tagged with both fluorescent marker and Auxin-inducible degron (NeonGreen-AID) degrades in the presence of plant specific ubiquitin ligase TIR1 and plant hormone auxin. TIR1 is an auxin receptor; it binds the mammalian SKP1-CUL1-F-Box (SCF) complex and directs AID-tagged proteins for ubiquitination and proteasome-mediated degradation. (b) Live imaging of NG-AID-tagged NUP153 in untreated and auxin-treated DLD-1 cells. We incorporated iRFP670 at the end of the RCC1 open reading frame, to use it as a chromosomal marker. E3 ubiquitin ligase TIR1 was integrated into RCC1 locus downstream of sequence encoding iRFP670 but separated from RCC1-iRFP670 gene fusion by sequence encoding P2A self-cleavage peptide. Scale bar: 10 μm. NPC – Nuclear Pore Complex, NG – NeonGreen, AID – Auxin Inducible Degradation, Aux – Auxin, RCC1 - Regulator of Chromosome Condensation 1, iRFP – infra-Red Fluorescent Protein

We recommend using gRNA sequences within the last exon, upstream of the STOP codon for the GOI. If the GOI needs to be tagged at the N-terminus, use gRNA sequences that recognize the first exon, downstream of the Start codon. If the analyzed sequence lacks NGG PAM signatures or shows only low-quality gRNAs, alternative Cas9-derived variants can be chosen that target distinct PAM sequences, such as NG, GAA, or GAT [22]. As shown in Fig. 3a, these criteria caused us to select the following gRNAs for targeting of the NUP153 gene at C-terminus of the coding sequence (PAM sequence is underlined): gRNA-1 AAGACTGCTGTTAGACGCAGG gRNA-9 GTTAGACGCAGGAAATAAAGG 3. Exclude the PAM sequence and add CACC and CAAA overhangs to the desired oligonucleotides, as shown on the scheme below. Add “g” right after the CACC overhang (highlighted in bold) if the gRNA sequence does not start with a “g” to drive

Analysis of Nucleoporin Function Using Inducible Degron Techniques

135

Fig. 2 A timeline workflow of AID (left) and TIR1 (right) integration into endogenous locus of GOI and RCC1 genes, respectively. GOI – Gene-of-Interest

expression from human U6 promoter [23]. Synthesize salt-free oligonucleotides: 50 – caccgNNNNNNNNNNNNNNNNNN – 30 30 – cNNNNNNNNNNNNNNNNNNcaaa – 50 50 and 30 overhangs are required for cloning gRNA sequences into the BbsI-digested pX330 vector [24]. Use the following primers for C-terminus targeting of NUP153 gene (Fig. 3b, see Note 1): C-term_gRNA-1F caccgAAGACTGCTGTTAGACGC C-term_gRNA-1R aaacGCGTCTAACAGCAGTCTTc C-term_gRNA-9F caccGTTAGACGCAGGAAATAA C-term_gRNA-9R aaacTTATTTCCTGCGTCTAAC 4. Digest 1 μg of pX330 with BbsI-HF for 3 h at 37  C: 1 μg of pX330 1 μL BbsI-HF 5 μL 10
CutSmart Buffer X μL H2O up to 50 μL total Add 1 μL of Calf Intestinal alkaline phosphatase, vortex, and incubate the reaction at 37  C for 1 h to dephosphorylate the plasmid. Perform gel extraction of the digested plasmid and elute it in ultra-pure water. Measure DNA concentration. If using

136

Vasilisa Aksenova et al.

a Cas9 gRNA-1

Cas9 gRNA-9

NUP153 gene. Last exon TAA

Cas9 gRNA-1

Cas9 gRNA-9

PAM

PAM

AGATAAAGACTGCTGTTAGACGCAGGAAATAAAGGTCACA TCTAT

TCCTTTATTTCCAGTG AAGACUGCUGUUAGACGC gRNA

AGATAAAGACTGCTGTTAGACGCAGGAAATAAAGGTCACATTGGT TCTATTTCTGACGA TCCAGTGTAACCA GUUAGTCGCAGGAAAUAA gRNA

b gRNA-1

BbsI U6 promoter

GAAA

BbsI

5’ – CACCGAAGACTGCTGTTAGACGC – 3’

CTTTGTGG

GTTTT

3’ – CTTCTGACGACAATCTGCGCAAA – 5’

A

gRNA scaffold

Pol III terminator

Bbs1digested

Cas9

pX330

Fig. 3 Design of gRNAs for targeting the NUP153 gene. (a) Position and sequences of gRNAs targeting last exon of NUP153 gene. SpCas9 cut sites on genomic DNA are indicated with blue arrows, gRNA sequences are highlighted in orange, PAM sequences are shown in yellow. Note, PAM sequence should be present only in genomic DNA. (b) Cloning of gRNA sequences with added CACC and CAAA overhangs into pX330 vector. gRNA-1 requires additional G nucleotide in the beginning of gRNA sequence to initiate expression of gRNA from U6 promoter. Note that the gRNA-9 starts from a G nucleotide and does not require additional G nucleotide

Quick CIP, incubate at 37  C for 10 min and inactivate by heating at 80  C for 2 min. 5. Phosphorylation and annealing of oligonucleotides: Resuspend dry oligonucleotides in ultra-pure water to a concentration of 1 mM, then assemble the reaction: 1 μL gRNA forward 1 μL gRNA reverse 1 μL 10
T4 DNA Ligase Buffer (contains 1 mM ATP) 6.5 μL H2O 0.5 μL T4 PNK

Analysis of Nucleoporin Function Using Inducible Degron Techniques

137

Anneal in a thermocycler using the following parameters: 37  C 30 min, 95  C 5 min and then ramp down to 25  C at 5  C/min. 6. Ligation reaction: We recommend setting up phosphorylation with the following annealing of oligonucleotides right before ligation step to maximize the cloning efficiency. Dilute phosphorylated and annealed oligonucleotides duplex mix 1:200 in ultra-pure water. Assemble the reaction: X μL dephosphorylated BbsI-digested plasmid (50 ng) 1 μL phosphorylated and annealed diluted oligo duplex 1 μL 10
T4 DNA Ligase Buffer 1 μL T4 DNA Ligase X μL H2O, to a total volume of 10 μL Incubate the reaction overnight at 16  C. Omit annealed oligos in a separate reaction as a negative control. 7. Bacterial transformation: 1. Thaw an aliquot of frozen competent cells (High Efficiency NEB® 5-alpha Competent E. coli) on ice. 2. Add 5 μL of ligation mixture to the competent cells and incubate 35 min on ice. 3. Heat shock bacteria for 45 s at 42  C and put back on the ice for 2 min. 4. Add 500 μL of SOC outgrowth medium and incubate bacteria for 1 h at 37  C on a shaker. 5. Plate the bacteria on prewarmed ampicillin agar plates. To make agar plates with ampicillin follow Cold Spring Harbor Protocols: http://cshprotocols.cshlp.org/con tent/2011/2/pdb.rec12396.full 6. Incubate plates overnight at 37  C. The Negative control plate should have only 0–5 colonies. 7. Choose two-three colonies from the plate and inoculate them into 5 mL LB media containing ampicillin. Incubate bacteria overnight (see Note 2). 8. Perform standard miniprep plasmid purification protocol to isolate plasmid DNA. Purified plasmid DNA can be stored at 20  C. 9. Check insertion of gRNA sequence using universal U6 primer: GACTATCATATGCTTACCGT.

138

Vasilisa Aksenova et al.

3.2 Repair Donor Template Design

Here, we describe a vector design that allows insertion of a ~2600 bp cassette that encodes NeonGreen fluorescent protein, full-length AID degron tag, and the Hygromycin B-resistance gene. To create a repair donor template, we recommend either using plasmid from any commercial or home-made vector that contains multiple unique cloning sites (MCS) or using the backbone of any commercial vector for subsequent Gibson assembly. This protocol describes the designs of the repair template based on the pEGFPN1 vector (Clontech) using the Gibson assembly. Take a vector of your choice, preferably less than 5 kb. Here, we use pEGFP-N1 vector. 1. Perform restriction digestion to remove unnecessary sequences or sequences which could interfere with HDR (mammalian promoters, enhancers, internal ribosomal sites sequences encoding fluorescent proteins or antibiotics for stable cell lines selection). In this case, we used NdeI and NotI restriction sites to remove the CMV-EGFP cassette. 2. Purify genomic DNA from your target cell line. We isolated DNA from DLD-1 or HCT116 cell line using Wizard® Genomic DNA Purification Kit according to manufacturer’s recommendation (see Note 3). 3. Choose a ~1–2 kb region around the STOP codon (C-term targeting). The length of the homology arm should be 500–1000 nt long, starting less than 10 nt up- or downstream of the Cas9-induced double-strand break (DSB). The recommended position of homologous arms for NUP153 gene is shown in Fig. 4a. When designing primers for homology arms, it is important to mutate the target PAM sequence(s) or/and the gRNA binding sites (8–10 bases from the 30 end) [24] to prevent unwanted cleavage of the HDR construct by CRISPR/Cas9. Change the third position in the coding nucleotide triplets, according to the degeneracy rule, and ensure that the frequency of codon usage of the resultant codons is similar to that of the original ones. Synonymous mutations do not affect the encoded amino acid; however, codon bias might affect translation efficacy, folding, and stability of tagged protein [25]. We arranged codons based on frequency usage and demonstrate silent nucleotide changes performed in NUP153 gene (Fig. 4a, b). 4. Go to NEBuilder: change preferences according to assembly kit, number of fragments, and polymerase settings. We recommend keeping a minimum 20 nt overlap length between fragments. Hit “Build Construct”–> Add fragment –> Add cloning vector either from the database or add sequence

Analysis of Nucleoporin Function Using Inducible Degron Techniques

a

139

STOP I

K

T

A

V

R

R

R

K

ATA•AAG•ACT•GCT•GTT•AGA•CGC•AGG•AAA•TAAAGGTCACATTGGTGTTGTACTCAA LHA I

K

T

A

PAM V

R

R

R

PAM K

D

I

RHA S

G

G

G

ATA•AAG•ACT•GCT•GTT•AGA•CGG•AGA•AAG•GAT•ATC•TCC•GGA•GGT•GGT LHA NUP153 SGGG3-NeonGreen-AID

••••••••••••

AGGTCACATTGGTGTTGTACTCAA RHA

b Triplet

Fraction

Frequency/ Thousand

AA

Triplet

M

ATG

1.00

22.3

P

CCC

0.33

*

TGA

0.52

1.3

P

CCT

0.28

*

TAA

0.28

0.7

P

CCA

*

TAG

0.20

0.5

P

R

CGG

0.21

11.9

R

AGA

0.20

11.5

AA

Fraction Frequency/ Thousand

AA

Triplet

Fraction

Frequency/ Thousand

AA

Triplet

Fraction

Frequency/ Thousand

20.0

T

ACC

0.36

19.2

V

GTG

0.47

28.9

17.3

T

ACA

0.28

14.8

V

GTC

0.24

14.6

0.27

16.7

T

ACT

0.24

12.8

V

GTT

0.18

10.9

CCG

0.11

7.0

T

ACG

0.12

6.2

V

GTA

0.11

7.0

L

CTG

0.41

40.3

S

AGC

0.24

19.4

G

GGC

0.34

22.8

L

CTC

0.20

19.4

S

TCC

0.22

17.4

G

GGA

0.25

16.3

R

AGG

0.20

11.4

L

CTT

0.13

12.8

S

TCT

0.18

14.6

G

GGG

0.25

16.4

R

CGC

0.19

10.9

L

TTG

0.13

12.6

S

AGT

0.15

11.9

G

GGT

0.16

10.8

R

CGA

0.11

6.3

L

CTA

0.07

6.9

S

TCA

0.15

11.7

R

CGT

0.08

4.7

L

TTA

0.07

7.2

S

TCG

0.06

4.5

A

GCC

0.40

28.5

A

GCT

0.26

18.6

H

CAC

0.59

14.9

E

GAG

0.58

40.8

C

TGC

0.55

12.2

A

GCA

0.23

16.0

H

CAT

0.41

10.4

E

GAA

0.42

29.0

C

TGT

0.45

9.9

A

GCG

0.11

7.6

D

GAC

0.54

26.0

Q

CAG

0.75

34.6

F

TTC

0.55

20.4

I

ATC

0.48

21.4

D

GAT

0.46

22.3

Q

CAA

0.25

11.8

F

TTT

0.45

16.9

I

ATT

0.36

15.7

I

ATA

0.16

7.1

W

TGG

1.00

12.8

K

AAG

0.58

32.9

Y

TAC

0.57

15.6

N

AAC

0.54

19.5

K

AAA

0.42

24.0

Y

TAT

0.43

12.0

N

AAT

0.46

16.7

Fig. 4 Integration of silent mutations in repair template. (a) Integration of silent mutations in the gRNA “seed” sequences of the repair template to prevent unwanted cleavage by Cas9-gRNA complexes. PAM sequences are shown in yellow, silent mutations are highlighted in blue. (b) A table of synonymous codons in H.sapiens. Codons are arranged based on their frequency usage within each amino acid group. Table color coding was used for visualization purposes

manually in FASTA format –> Choose restriction digestion as a method of backbone linearization and upstream and downstream restriction sites –> Add fragments encoding full-length 229-amino acid AID degron (Addgene #47330) or codonoptimized minimal functional AID tag (PKDPAKPPAKAQVVGWPPVRSYRKNVMVSCQKSSGGPEAAAFVK), Hygromycin B gene (Addgene #47330), and NeonGreen fluorescent protein (see Note 4). We recommend adding a flexible linker - (SGGG)3 between the tag and GOI [26]. We advise adding unique restriction sites between components of the tag if they will need to be replaced in the future. It is also critical to insert a sequence encoding P2A self-cleavage peptide between the selection marker and AID degron to separate tagged GOI and the selection marker (Fig. 5a).

140

Vasilisa Aksenova et al.

Fig. 5 Analysis of selected AID-NUP153 homozygous clone. (a) A diagram of the repair template, demonstrating integration site of NG-AID-P2A-Hygromycin cassette into NUP153 endogenous locus, and oligonucleotides used for genotyping. (b) Genomic PCR from NUP153-NG-AID cells and parental DLD-1 cell line. (c) Western blot analysis of NUP153-NG-AID homozygous clone before and after 2 h of auxin treatment (lane 3 and 4). Black and green arrows indicate the molecular weight of unmodified and AID-NG-tagged NUP153. LHA – Left Homology Arm, RHA – Right Homology Arm

5. Synthesize required salt-free oligonucleotides suggested by NEBuilder. 6. Amplify all required fragments using High Fidelity polymerase (for example, Platinum™ SuperFi™ DNA Polymerase). Extract fragments from agarose gel and set up Gibson assembly reaction according to the kit assembly protocol recommendations: https://international.neb.com/applications/cloning-andsynthetic-biology/dna-assembly-and-cloning/gibsonassembly. Use a total of 0.2 pmols of DNA fragments and 3:1 insert: vector molar ratio for donor template assembly. To calculate the mass of insert and vector use NEB ligation calculator: http://nebiocalculator.neb.com/#!/ligation. 5 μL Gibson Assembly Master Mix (2
) X μL dephosphorylated NdeI- and NotI -digested plasmid (40 ng) X μL DNA fragment corresponding LHA X μL DNA fragment corresponding RHA X μL DNA fragment corresponding AID degron X μL DNA fragment corresponding NeonGreen X μL DNA fragment corresponding P2A-Hygromycin X μL H2O up to 10 μL total 7. Incubate reaction in a thermocycler at 50  C for 1 h.

Analysis of Nucleoporin Function Using Inducible Degron Techniques

141

8. Proceed to bacteria transformation using 5–10 μL of reaction mix. Incubate bacterial plates overnight and select 4–6 clones next day. 9. Inoculate clones into 5 mL LB media with selection antibiotics. Incubate bacteria overnight and perform standard miniprep plasmid purification protocol. Purified plasmid DNA can be stored at 20  C. 10. Check donor template assembly using restriction digestion with unique restriction enzymes and sequencing with the following primers: F_pBR322 AACGCCAGCAACGCGGCCT R_EVB GATGAGTTTGGACAAACCAC F_Hygro AGCGAGAGCCTGACCTATTGCA R_Hygro CGCAGGACATATCCACGC F_NG AGACGGAATTAAAACACTCGA F_NUP153 TGCTCAGAGCATCATGTCACATTA The last exon of the NUP153 gene needs to be in frame with the sequences encoding NeonGreen fluorescent proteins, the AID degron, P2A, and Hygromycin resistance marker. The STOP codon is required at the end of Hygromycin gene before RHA sequence. 3.3 Transfection and Clone Isolation

DLD-1 and HCT116 cell lines are both stably pseudodiploid adherent cell lines with the modal chromosome number of 46, occurring in 86% and 62% of cells, respectively. Both cell lines are sensitive to Hygromycin B (200 μg/mL), Blasticidin (10 μg/ mL), and Puromycin (3 μg/mL) (see Note 5). To culture DLD-1 and HCT116 cell lines we use either DMEM or McCoy’s Medium supplemented with heat-inactivated 10% FBS, antibiotics (100 IU/mL penicillin and 100 μg/mL streptomycin) and 2 mM GlutaMAX, in 5% CO2 atmosphere at 37  C. We recommend propagating cells at least two passages before the transfection, maintaining cells confluency at 70–80%, and testing cells for Mycoplasma contamination with any commercially available kit. 1. Seed cells for transfection at ~1.2
105 cells/well in 2 mL of complete media on 12-well plates. We recommend transfecting cells at 50–60% of confluency. ViaFect transfection reagent can be used with antibiotics-free media containing regular 10% serum. We advise changing regular complete media to OptiMEM antibiotics-free media containing 10% serum 2 h before the transfection to maximize the transfection efficiency. We use ratio 4:1 (ViaFect:DNA) and 2:1:1 for repair template:gRNA-1:gRNA-2 plasmids; 1 μg of total DNA per well of 12-well plate.

142

Vasilisa Aksenova et al.

2. Prepare 1.5 mL sterile tube. 3. Add 100 μL of Opti-MEM media into 1.5 mL tube. 4. Add 500 ng of repair template, 250 ng of gRNA-1, and 250 ng of gRNA-2. Vortex for 5 s. 5. Add 4 μL of ViaFect. Mix DNA-Viafect complexes by tapping or vortexing for 30 s using medium speed. 6. Incubate DNA-Viafect complexes at room temperature for 15 min. 7. Add 100 μL of DNA-Viafect complexes to a well of 12-well plate containing 2 mL of Opti-MEM media with serum. Add mixture in a dropwise manner to ensure equal distribution of complexes among cells. 8. Incubate DNA-Viafect complexes with cells for 24 h. 9. Change media to regular complete media 24 h posttransfection and incubate for another 48 h. 10. Trypsinize cells in 300 μL of Trypsin-EDTA solution, add 700 μL of complete DMEM, resuspend trypsinized cells to achieve single-cell density and plate them (100 μL and 900 μL) into two 10 cm plates containing 12 mL of complete media and the selection antibiotic. We use Hygromycin B in final concentration 200 μg/mL. 11. Return plates in incubator and wait for 1–2 weeks until singlecell colonies are formed. Evaluate plates every 4 days to prevent overgrowth of colonies. 12. Replace cell culture media to 1
PBS. 13. Use a sterile pipette tip to pick up an entire colony. Press the plunger of the 10 μL pipette to remove the air from the tip. Place pipette tip into 10 cm dish nearby the colony, scrape colony on the side and suck in detached clone into the tip with the PBS solution. Transfer the colony into a new 24-well plate. Change the pipet tip after each colony (see Note 6). We usually pick 12–18 clones. 14. Add 65 μL of Trypsin-EDTA solution into each well. Incubate colonies for 20 min at 37  C in the incubator. 15. Vigorously shake the cell culture plate. 16. Add 2 mL of fresh regular complete media and incubate for several days until cells reach 50–70% confluency. If the AID tag is properly integrated, you should see a bright green signal at the nuclear rim of the cells as shown on Fig. 1b. Transparent Lumox 24-well plates can be used both to visualize NeonGreen signal and to allow selection of brightest colonies with the proper localization of NUP153.

Analysis of Nucleoporin Function Using Inducible Degron Techniques

3.4 Screening of Homozygous Clones

143

Complete protein degradation is achieved if all alleles of the gene are tagged with the AID degron. To identify homozygous clones, we use PCR analysis of genomic DNA extracted from the colonies and immunoblot analysis using protein-specific antibodies. We recommend performing genotyping and immunoblot analysis at early steps of clones’ propagation to minimize number of passages of the final cell line. 1. Add 250 μL of transfected well.

Trypsin-EDTA

solution

into

each

2. Incubate for 20 min. 3. Add 1 mL of complete media into each well of plate. Mix gently by pipetting with 1 mL pipette tip. 4. Prepare three separate plates. 5. Divide cells of each trypsinized clone (total volume of each well 1250 μL) among three separate 24-well plates: first plate 500 μL of cells per well, second plate - 500 μL of cells per well, third plate - the remaining 250 μL of cells per well. The first and the second plate will be used for genotyping and immunoblot analysis, the third plate to propagate clones for storage. 6. Add 2 mL of fresh regular complete media to all wells and incubate for several days until they form a monolayer. 3.4.1 Genomic DNA extraction

1. Add 200 μL of Nuclei Lysis buffer from Wizard® Genomic DNA Purification Kit into each well. Scrape cells with 1 mL pipette tip and collect lysate into 1 mL Eppendorf tubes. 2. Add 6 μL of 20 mg/mL Proteinase K, incubate for 3 h or overnight at 55  C. Optional: add 1 μL of RNase Solution provided with the kit and incubate for 30 min at 37  C. 3. Allow the sample to cool down at 4  C for 2 min before proceeding to the next step. 4. Add 70 μL of Protein Precipitation Solution and invert tubes multiple times. Leave samples at 4  C for 5 min. 5. Centrifuge for 5 min at 13,000–16,000
g at 4  C to form a tight white pellet. 6. Collect supernatant into new 1 mL Eppendorf tubes with 600 μL of isopropanol. Mix samples multiple times and leave at RT for 3 min. At this step, you can move samples at 20  C and store there until further analysis. 7. Centrifuge samples for 10 min at 13,000–16,000
g at 4  C. 8. Decant supernatant. 9. Add 1 mL of 70% ethanol. Invert tubes multiple times.

144

Vasilisa Aksenova et al.

10. Centrifuge samples for 7 min at 13,000–16,000
g at 4  C. Decant supernatant. 11. Repeat steps 15 and 16. 12. Carefully aspirate remaining ethanol and let DNA pellet to air-dry for 10–20 min. 13. Resuspend DNA pellet in 50 μL of ultra-pure water. Rehydrate DNA overnight at 4  C. DNA samples are ready for PCR or can be stored at 4  C until further analysis. To confirm a proper integration of the degron in all alleles use genomic PCR amplification with primers annealing outside of homology arms. A position of primers for genotyping of NUP153-AID clones is shown on Fig. 5a. We recommend using NCBI Primer-BLAST tool to design and verify the annealing temperature of your specific primers (see Note 7). 14. Design primers annealing outside of homology arms. We recommend the following set of primers to screen for homozygous integration of the degron at the C-terminal part of NUP153 gene. Annealing temperature - 60.5  C. Length of PCR product from: parental DLD-1 cell line or untagged allele – 1934 bp tagged allele – 4453 bp. F_NUP153_Cterm_out AACCTATCATGTTGCGGCCG R_NUP153_Cterm_out ATTATCCAAAGTCACCACAGTCCAT Genotyping results of homozygously tagged NUP153 clones are shown in Fig. 5b. To further verify the tag integration into NUP153 locus, we recommend purifying PCR fragments from the gel and sequencing the region of insertion, including homology arms. 3.4.2 Western Blot Analysis

Western blotting is required to confirm both integration of the tag into locus of the interest and to ensure that the tag does not interfere with the stability of the tagged protein. Use SDS-PAGE and Western blotting protocol commonly used in your laboratory or method described here [27] to separate and transfer proteins into PVDF membrane. To detect degron integration into the locus of GOI we recommend using either antibodies recognizing a specific epitope at the opposite terminus of the AID-tagged protein (here: NUP153 N terminus) or antibodies raised against the whole protein that recognizes multiple epitopes. We used rabbit polyclonal antibody A301788A from Bethyl Laboratories that recognize a region between residue 50 and 100 of human NUP153 (see Note 8). Western blot results of homozygously tagged NUP153 clone is shown in Fig. 5c.

Analysis of Nucleoporin Function Using Inducible Degron Techniques

145

3.5 Integration of OsTIR1

Ubiquitin ligase TIR1 is required for Auxin-mediated degradation of AID-tagged proteins. We recommend CRISPR/Cas9-mediated integration of TIR1 into the RCC1 (Regulator of Chromosome Condensation 1) locus. We inserted cDNA encoding TIR1 at the C-terminus of RCC1 gene product. We additionally added iRFP670 sequence to the C-terminus of RCC1 to simultaneously visualize cell nuclei and confirm a proper TIR1 integration. A sequence encoding a P2A self-cleavage peptide separates RCC1iRFP670 and TIR1 proteins. Transfect NUP153-homozygous clone with RCC1:iRFP670P2A-TIR1 donor plasmids and gRNAs targeting RCC1 locus. See Subheading 3.3 for transfection and clone isolation details. The sequence of gRNAs, oligonucleotides for homology arms amplification, and donor design are provided in Aksenova et al. [28] and Yau et al. [29]. Select cells on blasticidin (10 μg/mL). TIR1positive clones will express chromatin-bound RCC1-iRF670 which can be visualized in the infra-red channel. Collect 6–10 clones. We recommend splitting each TIR1-positive clone into two wells on Lumox, 24-well plates as shown in Fig. 2 (right panel). Allow cells to adhere for 2–3 days. Add Auxin (1 mM final) to one of the wells and analyze the disappearance of NeonGreen signal over time. Degradation times can vary among different cell lines and clones with hetero- or homozygous integration of TIR1 (see Note 9). Select clone with the brightest level of NeonGreen expression and shortest time of NUP153 degradation. We isolated the clone with degradation time of NUP153 ~ 40 min [28]. Confirm NUP153 degradation using Western blot analysis as shown in Fig. 5c (see Note 10).

3.6 Tag Selection and Functional Assays

NUP153 can be successfully tagged both at the N- or C-terminus. However, N-terminally tagged NUP153 cells display a dimmer NeonGreen signal and require a longer time for complete NUP153 degradation. This may reflect the fact that the N-terminus of NUP153 encodes its NPC-targeting domain, potentially influencing the fluorescence level of the NeonGreen signal and accessibility of the degron to TIR1 ligase. We tagged NUP153 gene with the sequences encoding NeonGreen protein and full-length AID degron. If a smaller tag is required, we recommend using a codon-optimized minimal functional AID tag (microAID, 71-114 aa) [30]. The presence of the tag can potentially affect the interaction of AID-tagged protein with other binding partners. We recommend analyzing NUP-AID cell line in several functional assays. We developed a pipeline of functional assays performed for each NUP-AID tagged cell line:

146

Vasilisa Aksenova et al.

1. Analysis of the cell growth of AID-tagged cells and parental untagged cell line [31]. The growth rate of AID-tagged cells should be similar to the parental cell line. Auxin treated NUP153-AID tagged cells should display cell growth abnormalities within 24 h. 2. Analysis of the protein NUP153 degradation using either mass spectrometry (MS) of whole-cell lysates or/and Western blot analysis (Fig. 5c). 3. Nucleoporins are primary gatekeepers regulating nuclearcytoplasmic transport. We recommend performing the import–export assay [32] to analyze transport function of AID-tagged nucleoporins. In the absence of Auxin, AID-tagged nucleoporins’ cell line should display transport rates similar to those in the parental cell line. 4. The addition of tag can interfere with both protein’s function and with its ability to interact with corresponding partners. We recommend performing immunostaining and Western blot analysis of key interacting partners. We analyzed localization of NUP50, TPR, RanBP2, RanGAP1, NUP133, and NUP98 in both AID-tagged NUP153 and parental cell lines in the absence of Auxin. 3.7 NUP-AID System Applications

The rapid degradation of AID-tagged proteins facilitates investigation of the NPC’s architectural composition, including investigations of questions which were previously unattainable. For instance, AID-tagged nucleoporins can be specifically degraded within a particular phase of the cell cycle, allowing their roles during interphase and post-mitotic NPC assembly to be experimentally distinguished. Furthermore, the CRISPR/Cas9-NUP-AID system can be combined with endogenous targeting of other NPC-components using other fluorescent proteins. Multi-color CRISPR/Cas9-NUP-AID system allows simultaneous visualization of up to three nuclear pore components. This degron-targeting strategy can also be used to distinguish redundancy of nucleoporin phenotypes. For example, the loss of NUP153 results in mislocalization of NUP50 from the nuclear envelope into the nucleoplasm [28]. It remains unclear if such aberrant NUP50 nuclear localization is partially responsible for the NUP153 phenotype. The double degron cell line with the simultaneous AID-targeting of NUP153 and NUP50 can help to address this question. DLD-1 and HCT116 cell lines maintain a pseudodiploid number of chromosomes, where each gene is encoded by two alleles which can be tagged with the fluorescent protein using CRISPR/ Cas9. Homozygously tagged nucleoporins preserve a physiologically relevant protein level in comparison to ectopically expressed

Analysis of Nucleoporin Function Using Inducible Degron Techniques

147

nucleoporins. CRISPR/Cas9 homozygously tagged nucleoporins’ cell lines can be used to determine the number of nucleoporins per pore and estimate nucleoporin exchange at the pore using microscopy-based methods as super-resolution microscopy (SRM) and fluorescence recovery after photobleaching (FRAP). Removal of Auxin results in restoration of NUP-AID protein levels, depending on the translation efficiency from the corresponding mRNA. This approach may be used to analyze the rates of recovery of “damaged pores” after the loss of critical NUPs.

4

Notes 1. The native form of the guide RNA sequence targeting Cas9 to its binding site within the genome is 20 nt long. The guide RNA could be truncated to 18–19 bp by removing the 5 ‘nucleotides after the first G nucleotide. This truncation reduces off-target activity without the loss of on-target genome-editing efficiency [33]. 2. Use ampicillin and kanamycin at a final concentration of 100 μg/mL and 50 μg/mL, respectively. To make LB media use the following recipe http://cshprotocols.cshlp.org/con tent/2006/1/pdb.rec8141.full?sid=7f7ce2e0-966b-40bd-93 dc-2e7818003b97. 3. We recommend using ultra-pure water rather than TE buffer to dissolve genomic DNA, plasmid DNA, and oligonucleotides. 4. If you need to perform codon optimization of the degron or TIR1 sequences, use IDT Codon Optimization Tool: https:// www.idtdna.com/pages/tools/codon-optimization-tool? returnurl¼%2FCodonOpt. OsTIR1 and full-length AID degron encoded by pBABE TIR1-9Myc (Addgene, 47328) and pcDNA5-EGFP-AID-BubR1 (Addgene, 47330) plasmids are codon-optimized to maximize expression of these genes in H. sapiens. The degron sequence of the minimal functional AID tag (microAID) 71-114 amino acids has been optimized in our laboratory. 5. If it is not possible to use these cell lines as a model, check a copy number of the GOI and stability of the karyotype of the desired cell line. If you fail to obtain targeting of all alleles within the genome, you may perform a second round of selection. To achieve this, introduce sequences that correspond to a different antibiotic-resistant gene into your original HDR construct and perform transfection and HDR-based selection of the cell clone that contains heterozygous insert. However, to follow that approach care must be taken to verify that the

148

Vasilisa Aksenova et al.

untargeted allele(s) are not cleaved by the CRISPR/Cas9 and repaired by the non-homologous end joining pathway, as the gRNA target sequence in those allele(s) will be lost. 6. Colonies formed after selection with the selection marker should be round and preserve the same morphology as original cell line. Avoid colonies with an uneven shape because they can be formed by neighboring colonies fused together. 7. To design primers for amplification of homology arms use the NCBI Primer-BLAST tool: https://www.ncbi.nlm.nih.gov/ tools/primer-blast/index.cgi?LINK_LOC¼BlastHome. We recommend designing primers outside of regions with repetitive sequences. Choose two-three pairs of primers for genotyping clones with similar annealing temperature. 8. If protein-specific antibodies are not available, we suggest performing a mass-spectrometry analysis of whole-cell lysate to confirm full protein degradation. 9. Depletion of nucleoporins can be achieved by the cultivation of cells in complete media in the presence of 1 mM Auxin (Sigma Aldrich). Auxin stock solution (500 mM, in water) can be stored at 20  C up to several months. Auxin is light-, temperature-, and pH-sensitive. 10. The AID-TIR1 system can decrease level of AID-tagged protein [12], possibly due to affinity of AID degron to TIR1 ligase [34]. We advise to first integrate the AID degron and keep frozen aliquots of TIR1 negative homozygous clones, then integrate TIR1. TIR1 positive clones should be stored in liquid nitrogen and passaged from fresh aliquots no more than 12 times after TIR1 integration. Note that the AID-tagged protein level can drop at late passages or during improper or prolonged cell storage. Because the DLD-1 cell line is not entirely homogeneous, GOI expression levels may differ between clones. We recommend analyzing level of AID-tagged proteins in several separate clones before and after TIR1 integration. AID-tagged proteins should show expression levels similar to the untagged endogenous protein.

Acknowledgments This work was supported by Intramural Research Program of the Eunice Kennedy Shriver National Institute of Child Health and Human Development at the National Institutes of Health, USA (Intramural Project #Z01 HD008954). We thank Ashley Person, NIH postbaccalaureate fellow, for comments on this manuscript.

Analysis of Nucleoporin Function Using Inducible Degron Techniques

149

References 1. Hampoelz B, Andres-Pons A, Kastritis P, Beck M (2019) Structure and assembly of the nuclear pore complex. Annu Rev Biophys 48: 515–536. https://doi.org/10.1146/annurevbiophys-052118-115308 2. Sakuma S, D’Angelo MA (2017) The roles of the nuclear pore complex in cellular dysfunction, aging and disease. Semin Cell Dev Biol 68:72–84. https://doi.org/10.1016/j. semcdb.2017.05.006 3. Toyama BH, Savas JN, Park SK, Harris MS, Ingolia NT, Yates JR, Hetzer MW (2013) Identification of long-lived proteins reveals exceptional stability of essential cellular structures. Cell 154(5):971–982. https://doi.org/ 10.1016/j.cell.2013.07.037 4. Walther TC, Alves A, Pickersgill H, Loiodice I, Hetzer M, Galy V, Hulsmann BB, Kocher T, Wilm M, Allen T, Mattaj IW, Doye V (2003) The conserved Nup107-160 complex is critical for nuclear pore complex assembly. Cell 113(2):195–206. https://doi.org/10.1016/ s0092-8674(03)00235-6 5. Nishimura K, Fukagawa T, Takisawa H, Kakimoto T, Kanemaki M (2009) An auxinbased degron system for the rapid depletion of proteins in nonplant cells. Nat Methods 6(12):917–922. https://doi.org/10.1038/ nmeth.1401 6. Hoffmann S, Fachinetti D (2018) Real-time De novo deposition of centromeric histoneassociated proteins using the auxin-inducible degradation system. Methods Mol Biol 1832: 223–241. https://doi.org/10.1007/978-14939-8663-7_12 7. Shetty A, Reim NI, Winston F (2019) Auxininducible Degron system for depletion of proteins in Saccharomyces cerevisiae. Curr Protoc Mol Biol 128(1):e104. https://doi.org/10. 1002/cpmb.104 8. Nishimura K, Kanemaki MT (2014) Rapid depletion of budding yeast proteins via the fusion of an auxin-inducible Degron (AID). Curr Protoc Cell Biol 64:20.9.1–20.916. https://doi.org/10.1002/0471143030. cb2009s64 9. Zhang L, Ward JD, Cheng Z, Dernburg AF (2015) The auxin-inducible degradation (AID) system enables versatile conditional protein depletion in C. elegans. Development 142(24):4374–4384. https://doi.org/10. 1242/dev.129635 10. Bence M, Jankovics F, Lukacsovich T, Erdelyi M (2017) Combining the auxin-inducible degradation system with CRISPR/Cas9-based

genome editing for the conditional depletion of endogenous Drosophila melanogaster proteins. FEBS J 284(7):1056–1069. https://doi. org/10.1111/febs.14042 11. Nielsen CF, Zhang T, Barisic M, Kalitsis P, Hudson DF (2020) Topoisomerase IIalpha is essential for maintenance of mitotic chromosome structure. Proc Natl Acad Sci U S A 117(22):12131–12142. https://doi.org/10. 1073/pnas.2001760117 12. Sathyan KM, McKenna BD, Anderson WD, Duarte FM, Core L, Guertin MJ (2019) An improved auxin-inducible degron system preserves native protein levels and enables rapid and specific protein depletion. Genes Dev 33(19–20):1441–1455. https://doi.org/10. 1101/gad.328237.119 13. Nora EP, Goloborodko A, Valton AL, Gibcus JH, Uebersohn A, Abdennur N, Dekker J, Mirny LA, Bruneau BG (2017) Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169(5):930–944.e22. https://doi.org/10.1016/j.cell.2017.05.004 14. Lambrus BG, Moyer TC, Holland AJ (2018) Applying the auxin-inducible degradation system for rapid protein depletion in mammalian cells. Methods Cell Biol 144:107–135. https://doi.org/10.1016/bs.mcb.2018. 03.004 15. Holland AJ, Fachinetti D, Han JS, Cleveland DW (2012) Inducible, reversible system for the rapid and complete degradation of proteins in mammalian cells. Proc Natl Acad Sci U S A 109(49):E3350–E3357. https://doi.org/10. 1073/pnas.1216880109 16. Boer J, Bonten-Surtel J, Grosveld G (1998) Overexpression of the nucleoporin CAN/NUP214 induces growth arrest, nucleocytoplasmic transport defects, and apoptosis. Mol Cell Biol 18(3):1236–1247. https://doi. org/10.1128/mcb.18.3.1236 17. Clift D, So C, McEwan WA, James LC, Schuh M (2018) Acute and rapid degradation of endogenous proteins by trim-away. Nat Protoc 13(10):2149–2175. https://doi.org/10. 1038/s41596-018-0028-3 18. Natsume T, Kiyomitsu T, Saga Y, Kanemaki MT (2016) Rapid protein depletion in human cells by auxin-inducible degron tagging with short homology donors. Cell Rep 15(1): 210–218. https://doi.org/10.1016/j.celrep. 2016.03.001 19. Sathyan KM, Scott TG, Guertin MJ (2020) ARF-AID: a rapidly inducible protein

150

Vasilisa Aksenova et al.

degradation system that preserves basal endogenous protein levels. Curr Protoc Mol Biol 132(1):e124. https://doi.org/10.1002/ cpmb.124 20. Yesbolatova A, Saito Y, Kitamoto N, MakinoItou H, Ajima R, Nakano R, Nakaoka H, Fukui K, Gamo K, Tominari Y, Takeuchi H, Saga Y, Hayashi KI, Kanemaki MT (2020) The auxin-inducible degron 2 technology provides sharp degradation control in yeast, mammalian cells, and mice. Nat Commun 11(1):5701. https://doi.org/10.1038/s41467-02019532-z 21. Lin DH, Hoelz A (2019) The structure of the nuclear pore complex (an update). Annu Rev Biochem 88:725–783. https://doi.org/10. 1146/annurev-biochem-062917-011901 22. Hu JH, Miller SM, Geurts MH, Tang W, Chen L, Sun N, Zeina CM, Gao X, Rees HA, Lin Z, Liu DR (2018) Evolved Cas9 variants with broad PAM compatibility and high DNA specificity. Nature 556(7699):57–63. https:// doi.org/10.1038/nature26155 23. Kim HK, Lee S, Kim Y, Park J, Min S, Choi JW, Huang TP, Yoon S, Liu DR, Kim HH (2020) High-throughput analysis of the activities of xCas9, SpCas9-NG and SpCas9 at matched and mismatched target sequences in human cells. Nat Biomed Eng 4(1):111–124. https://doi.org/10.1038/s41551-0190505-1 24. Cong L, Ran FA, Cox D, Lin S, Barretto R, Habib N, Hsu PD, Wu X, Jiang W, Marraffini LA, Zhang F (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339(6121):819–823. https://doi.org/10. 1126/science.1231143 25. Quax TE, Claassens NJ, Soll D, van der Oost J (2015) Codon bias as a means to fine-tune gene expression. Mol Cell 59(2):149–161. https://doi.org/10.1016/j.molcel.2015. 05.035 26. Chen X, Zaro JL, Shen WC (2013) Fusion protein linkers: property, design and functionality. Adv Drug Deliv Rev 65(10):1357–1369.

https://doi.org/10.1016/j.addr.2012. 09.039 27. Blancher C, Jones A (2001) SDS -PAGE and Western blotting techniques. Methods Mol Med 57:145–162. https://doi.org/10.1385/ 1-59259-136-1:145 28. Aksenova V, Smith A, Lee H, Bhat P, Esnault C, Chen S, Iben J, Kaufhold R, Yau KC, Echeverria C, Fontoura B, Arnaoutov A, Dasso M (2020) Nucleoporin TPR is an integral component of the TREX-2 mRNA export pathway. Nat Commun 11(1):4577. https:// doi.org/10.1038/s41467-020-18266-2 29. Yau KC, Arnaoutov A, Aksenova V, Kaufhold R, Chen S, Dasso M (2020) RanBP1 controls the Ran pathway in mammalian cells through regulation of mitotic RCC1 dynamics. Cell Cycle 19(15):1899–1916. https://doi.org/10.1080/15384101.2020. 1782036 30. Morawska M, Ulrich HD (2013) An expanded tool kit for the auxin-inducible degron system in budding yeast. Yeast 30(9):341–351. https://doi.org/10.1002/yea.2967 31. Feoktistova M, Geserick P, Leverkus M (2016) Crystal violet assay for determining viability of cultured cells. Cold Spring Harb Protoc 2016(4):pdb.prot087379. https://doi.org/ 10.1101/pdb.prot087379 32. Niopek D, Wehler P, Roensch J, Eils R, Di Ventura B (2016) Optogenetic control of nuclear protein export. Nat Commun 7: 1 0 6 2 4 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / ncomms10624 33. Fu Y, Sander JD, Reyon D, Cascio VM, Joung JK (2014) Improving CRISPR-Cas nuclease specificity using truncated guide RNAs. Nat Biotechnol 32(3):279–284. https://doi.org/ 10.1038/nbt.2808 34. Tan X, Calderon-Villalobos LI, Sharon M, Zheng C, Robinson CV, Estelle M, Zheng N (2007) Mechanism of auxin perception by the TIR1 ubiquitin ligase. Nature 446(7136): 6 4 0 – 6 4 5 . h t t p s : // d o i . o r g / 1 0 . 1 0 3 8 / nature05731

Chapter 10 Monitoring of Chromatin Organization at the Nuclear Pore Complex, Inner Nuclear Membrane, and Nuclear Interior in Live Cells by Fluorescence Ratiometric Imaging of Chromatin (FRIC). Frida Niss, Cecilia Bergqvist, Anna-Lena Stro¨m, and Einar Hallberg Abstract The image analysis tool FRIC (Fluorescence Ratiometric Imaging of Chromatin) quantitatively monitors dynamic spatiotemporal distribution of euchromatin and total chromatin in live cells. A vector (pTandemH) assures stoichiometrically constant expression of the histone variants Histone 3.3 and Histone 2B, fused to EGFP and mCherry, respectively. Quantitative ratiometric (H3.3/H2B) imaging displayed a concentrated distribution of heterochromatin in the periphery of U2OS cell nuclei. As a proof of concept, peripheral heterochromatin responded to experimental manipulation of histone acetylation as well as expression of the mutant lamin A protein “progerin,” which causes Hutchinson-Gilford Progeria Syndrome. In summary FRIC is versatile, unbiased, robust, requires a minimum of experimental steps and is suitable for screening purposes. Key words Chromatin, Nuclear pore complex, Nuclear membrane, Live imaging, Fluorescence ratiometric

1

Introduction Nuclear envelope (NE) proteins organize chromatin. Silent areas of the genome are organized in a condensed form called heterochromatin, whereas active genes distribute in less condensed areas called euchromatin. Generally, euchromatin is common in the nuclear interior whereas heterochromatin is more frequent in the nuclear periphery. The organization of heterochromatin in the nuclear periphery is believed to involve proteins of the nuclear lamina and the inner nuclear membrane [1]. There is a strong statistical correlation between the activity of genes and their radial position within the nucleus [2], pointing out the nuclear periphery

Frida Niss and Cecilia Bergqvist contributed equally to this work. Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2022

151

152

Frida Niss et al.

as a “silencing” region. Both A- and B-type lamins [3], and many inner nuclear membrane (INM) proteins, including LBR, Samp1 and the LEM-domain proteins [3–5] have been shown to be essential for maintenance of peripheral heterochromatin. On the other hand, proteins of the nuclear pore complex (NPC) were reported to bind active genes and insulator elements, most likely to promote euchromatin locally to avoid clogging nucleocytoplasmic transport through the nuclear pores [6, 7]. Maintenance of heterochromatin is recognized as a crucial protective factor in aging. Loss of heterochromatin with age increases the incidence of de-repression of genes and transposons causing cancer and neurological disorders [8–11], This points to an important role for the NE in harnessing peripheral heterochromatin, which is compromised in some laminopathies, e.g., Hutchinson-Gilford progeria syndrome [5, 11, 12]. Chromatin organization has been extensively studied on a cell population level, but there is a need to understand dynamic chromatin reorganization in single live cells. To meet this need, we have developed a novel quantitative imaging tool called FRIC (Fluorescence Ratio Imaging of Chromatin) [4]. We have exploited the fact that a specific histone variant, H3.3, is found in euchromatin whereas H2B is present in all chromatin. H2B fused to the monomeric fluorescent protein “mCherry” (587/610 nm) and H3.3 fused to EGFP (488/507 nm) expressed at stoichiometrically constant levels from the pTandemH vector (Fig. 1A) can easily be monitored in two separate channels for several days after transient transfection. FRIC is especially well suited for monitoring spatial redistributions of heterochromatin and euchromatin in the nuclear periphery. The localization of heterochromatin designated by low ratios in the periphery of the nucleus (Fig. 1C, c–e) is much more clearly seen using FRIC compared to viewing only H3.3 (Fig. 1B) or only H2B (Fig. 1B). As proof of concept, a clear and expected increase in peripheral heterochromatin was observed after treatment with the histone acetyltransferase inhibitor Anacardic acid (AA) (Fig. 2A– C). In contrast, decreased peripheral heterochromatin was observed after treatment with the histone deacetylase inhibitor, Trichostatin A (TSA) (Fig. 2D, E) or expression of progerin (Fig. 2G, H) [4], a lamin A mutant causing Hutchinson-Gilford Progeria Syndrome [12]. In order to illustrate dynamic reorganization of chromatin as a result of TSA treatment, cells were followed by time-lapse FRIC, showing that significant decreases in peripheral heterochromatin was evident within an hour of exposure to TSA (Fig. 3). In comparison to contemporary immunostaining methods, FRIC is unbiased in terms of accessibility in dense heterochromatic areas. In addition, FRIC is relatively unlaborious, cheap, and versatile, which makes it ideal for screening purposes.

Monitoring Chromatin Organisation by FRIC

153

a 6000

herry mC

500

0

00 10

2B er ot m

CM CMV enhance V pro H r

2

ES IR

200 0

pTandem H2b-mcherry & H3.3-EGFP 6100 bp

40 .

3

00 E G FP

H3

3000

b H3.3-GFP

H2B-mCherry

c Masked H3.3-GFP Masked H2B-mCherry a b

Ratio

Radial Profile

Distance Map c

d

1,2 1,1 1 0,9 0,8 0,7 0,6 0,5

e

0

10

20 Zone

30

40

Fig. 1 Monitoring epigenetic changes in spatiotemporal distribution of chromatin in live cells using FRIC. (A) Schematic illustration of the bicistronic tandem vector (pTandemH) for stoichiometrically constant expression of H2B-mCherry and H3.3-EGFP. (B) Confocal fluorescence microscopy images of H3.3-EGFP (left) and H2B-mCherry (right) in live U2OS cells transfected with pTandemH. (C) The segmented images from B, of H2B-mCherry (a) and H3.3-EGFP (b). The normalized intensity ratio (H3.3/H2B) (c, bright ¼ high ratio, dark ¼ low ratio) plotted on the nuclear distance map as mean ratios (d). Radial profile of zones from the nuclear periphery to the interior (P ! I) with incision showing the division of zones in relation to the profile above (e). P-values >0.05 were considered not significant, P < 0.01 considered *, P < 0.001 **, P < 0.0001 ***, and P < 0.0001 ****. (Reproduced from [4])

154

Frida Niss et al.

Fig. 2 Effects of histone acetylation and Progerin on chromatin organization. U2OS cells transfected with pTandemH were treated with Anacardic Acid (AA) (A–C), Trichostatin A (TSA) (D-F) or transfected with a plasmid encoding progerin (G-I). (A) Ratiometric images (H3.3/H2B) of an AA treated U2OS cell. Heterochromatin (darker) and euchromatin (brighter) regions. (B) Radial profile (P ! I; only the first 10 zones are shown in the graph; see supplementary [4] for all zones). Cells treated with AA had significantly ( p < 0.0001) lower ratios in the nuclear periphery (zones 1–5) in comparison to control cells (n ¼ 144 AA treated, n ¼ 107 control). (C) Mean relative ratio (P/I, zone 1–3/zone 4–40) of AA treated and control cells ( p < 0.0001). (D) Ratiometric images of TSA treated U2OS cells. (E) Radial profile of cells treated with TSA and control cells (only the first 10 peripheral zones (P ! I) are shown in the graph, see supplementary [4] for all zones). Cells treated with TSA had significantly ( p < 0.00052) higher ratio in the nuclear periphery (zone 1–2) in comparison to

Monitoring Chromatin Organisation by FRIC

2

155

Materials 1. Materials for culturing cells. (a) Human U2OS cells. (b) Dulbecco’s modified Eagle medium. (c) Fetal bovine serum. (d) Penicillin-Streptomycin. (e) PBS: NaCl, KCl, Na2HPO4, KH2PO4. (f) Trypsin-EDTA. 2. Materials for FRIC. (a) Glass bottom dishes for microscopy. (b) XtremeGENE™ HP. (c) Trichostatin A. (d) Anacardic acid. (e) pTandemH [4].

3

Methods FRIC can be implemented for either live or fixed cells. This protocol will mainly describe how to implement the method for live cells. Further tips on how to modify the method for fixed cells can be found in the Notes section. 1. Culture U2OS cells in DMEM medium supplemented with 10% FBS and 1% Penicillin-Streptomycin. Seed 200,000 cells on day 0 in 35 mm glass bottom dishes adapted for microscopy imaging (see Notes 1 and 2). Throughout the experiment, change the medium every 2–3 days. 2. Prepare a transfection mixture in a 200 μL tube by adding serum-free medium, 0.5 μg of pTandemH and 1.5 μL of XtremeGENE ™ HP transfection reagent, in that order, to a total volume of 100 μL (see Note 3). For experiments using a second

ä Fig. 2 (continued) control cells (n ¼ 45 TSA treated, n ¼ 52 control cells). (F) Mean relative ratio (P/I, zone 1–3/zone 4–40) of TSA treated and control cells ( p ¼ 0.002). (G) Ratiometric images of progerin expressing U2OS cells. (H) Radial profile of cells treated with progerin transfected and control cells (only the first 10 peripheral zones (P ! I) are shown in the graph, see supplementary [4] for all zones). Cells transfected with progerin had significantly ( p < 0.0103) higher ratio in the nuclear periphery (zone 1–2) in comparison to control cells (n ¼ 55 progerin, 52 control cells). (I) Mean relative ratio (P/I, zone 1–3/zone 4–40) of progerin transfected and control cells ( p ¼ 0.0057). Experiments were performed three times. P-values >0.05 were considered not significant, P < 0.01 considered *, P < 0.001 **, P < 0.0001 *** and P < 0.0001 ****. (Reproduced from [4])

156

Frida Niss et al.

Fig. 3 Monitoring of dynamic changes in chromatin organization in live cells treated with TSA. U2OS cells transfected with pTandemH were treated with 200 nM TSA and monitored every 15 min. (A) Ratiometric images of control cells and cells treated with TSA after 0, 15, 30, 45, and 60 min. (B) Time lapse study displaying change in ratio of the nuclear periphery (zone 1) of control and TSA treated cells. After 60 min, cells treated with TSA had a significantly ( p ¼ 0.0115) higher ratio in the nuclear periphery (zone 1) in comparison to control cells (n ¼ 56 control, n ¼ 45 TSA cells). Experiments were performed three times. P-values >0.05 were considered not significant, P < 0.01 considered *, P < 0.001 **, P < 0.0001 ***, and P < 0.0001 ****. (Reproduced from [4])

plasmid to, e.g. induce a phenotype, add 0.5 μg of this plasmid into the mixture as well and increase the amount of XtremeGENE to 3 μL (see Note 4). Before starting, allow all reagents to equilibrate to room temperature and mix well (see Note 5). After mixing the reagents, incubate the tube at room temperature for at least 30 min.

Monitoring Chromatin Organisation by FRIC

157

3. Add 100 μL of the transfection mixture dropwise to each newly seeded dish, and gently swirl the dish. 4. On day 2–4, depending on confluency, add Trichostatin A at a concentration of 50 nM to the appropriate dishes for a 24 h incubation (see Note 6). 5. On day 3–5 the cells should be confluent. 1 h before imaging add Anacardic acid to a concentration of 10 μM to the appropriate dishes (see Notes 7–9). 6. Use a confocal laser scanning microscope with a stage incubator that keeps the temperature at 37  C and the CO2 at 5% for live imaging. Locate nuclei that are expressing both H3.3EGFP and H2B-mCherry (see Notes 10 and 11). Use a 63/1.4 oil immersion objective (see Note 12) and collect two-channel images of around 50 cells per treatment and replicate (see Note 13). 7. Every nucleus should be imaged to capture the section of the nucleus with the largest area (see Note 14). 8. Optional: Before analysis, an image quality module can be added that excludes outliers (see Note 15). 9. Use the free software CellProfiler [13] to analyze the imaged nuclei. (a) Measure the minimum pixel diameter of the imaged nuclei using the MeasureObjectSizeShape module in Cellprofiler, or manually in a small sample of images using any image handling program (see Note 16). (b) Use the module IdentifyPrimaryObjects in CellProfiler version 3.1.9 to find the nuclei, using the H2B-mCherry channel. In this module, use the automatic thresholding strategy and discard objects smaller than the minimum pixel diameter or touching the image border. (c) Segment the nuclei from the background, using the module MaskImage and the mask created in the previous step. (d) Calculate the intensity of both channels in each segmented nucleus, using the module MeasureImageIntensity. (e) Normalize the H2B-mCherry and H3.3-EGFP images using the module ImageMath to divide each channel by the measurement of their mean intensities. In a second ImageMath module, divide them by their variation. This ensures normalization intra- and intercellularly. (f) Create a ratiometric image by dividing the normalized H3.3-EGFP channel by the normalized H2B-mCherry channel using a second ImageMath module.

158

Frida Niss et al.

(g) Use the MeasureObjectIntensityDistribution module in CellProfiler to divide the nuclei into an appropriate number of concentric zones (bins) of equal width by dividing the radius into equal parts (see Note 17). Subsequently use this module to calculate the mean ratio intensity per zone. (h) Save and export the data for further analysis in Excel. 10. Plot the mean intensity of each zone from the nuclear periphery to nuclear interior as a radial profile (see Note 18).

4

Notes 1. The human osteosarcoma cell line U2OS has cells with large nuclei that grow in a perfect monolayer without overlap. Furthermore, they are contact inhibited, meaning that they stop proliferating when 100% confluence is reached. This provides excellent conditions for image acquisition. 2. To image fixed samples, either fix the cells in the same glass bottom dish as used for live imaging or seed the same number of cells into the well of a 6-well plate containing coverslips of no. 1.5. Proceed to fix and mount samples as appropriate. Depending on the cell line, a coating that promotes cell adhesion to the glass coverslips might be needed, such as, e.g. polylysine. This is not necessary for U2OS cells. 3. There might be need for optimization of the transfection protocol depending on which cell line is used. XtremeGENE™ HP normally works best in a 1:1, 3:1, or 6:1 ratio (μL XtremeGENE:μg plasmid). 4. A plasmid expressing progerin (Lamin A-L647R prelamin A) can be used as a control phenotype that should result in a decreased amount of heterochromatin in the nuclear periphery. 5. It is important to vortex the XtremeGENE transfection reagent immediately before adding it to the reaction mixture. It is equally important to vortex the transfection mixture before the room temperature incubation. 6. Trichostatin A is a Histone Deacetylase inhibitor and can be used as a control that should result in a lower amount of total heterochromatin. 7. Anacardic acid is a Histone Acetyltransferase inhibitor that can be used as a control that should result in a higher amount of total heterochromatin. 8. As both of these treatments (see Notes 6–7) are time sensitive, and imaging with a confocal laser scanning microscope can take time, make sure to give yourself enough time to image both at the end of their respective incubation times.

Monitoring Chromatin Organisation by FRIC

159

9. For imaging of fixed samples, use a fixative such as 3.7% paraformaldehyde to fix the cells at this point. 10. Since the transient transfection creates a heterogeneous population of nuclei with differing expression levels, the intensity of the fluorescent proteins will also vary. To be able to use the same image acquisition settings for all nuclei across treatments and replicates, determine what the average intensity of both channels are during the first experiment. Adjust your settings for optimal image acquisition of the average nucleus intensity. Exclude nuclei that are over- or underexposed according to these settings in each treatment and replicate in the following experiments. 11. Due to restrictions of the CellProfiler pipeline, the imaged nuclei must be round or oval shaped, without large invaginations. If the studied treatment/mutation causes many morphological abnormalities in the nuclei, FRIC can be used to study the milder/earlier effects of the treatment/mutant, when nuclear shape has not yet been compromised. 12. For a faster image acquisition, it is possible to use a 40x objective instead. However, the lower magnification might mean the nuclei will have to be divided into fewer concentric zones by the CellProfiler pipeline, leading to a lower resolution of the chromatin organization. 13. As live cell imaging and confocal laser scanning can take time, it would be prudent to image both channels simultaneously in case the cell moves and the overlap of the two channels changes. 14. To make sure to capture the widest section of the nucleus, capture a z-stack of three or more images around the visually determined largest section. In the image analysis, the section with the largest area can be chosen from these images. 15. To ensure that the intensity levels lie within appropriate limits, an ImageQuality module can be added to the CellProfiler pipeline. If 0.2% of the pixels in the nucleus are saturated or underexposed, or if the Otsu-threshold value is above 0.15, the image should be removed from the data set. 16. The minimum pixel diameter is usually larger than 80 pixels when imaging using a 40 objective for most cell lines, including U2OS cells. 17. An appropriate maximal number of concentric zones can be determined by dividing the minimum nuclear diameter by 2, to ensure that each zone has a width of at least two pixels. 18. In the CellProfiler export file, the zone numbered 1 is localized in the nuclear center.

160

Frida Niss et al.

Acknowledgments We would like to thank the Imaging Facility at Stockholm University (IFSU). This work was made possible by grants from the Swedish Research Council #621-2010-448, Cancerfonden #110590 and the foundation Olle Engkvists minne. References 1. Stewart CL, Roux KJ, Burke B (2007) Blurring the boundary: the nuclear envelope extends its reach. Science 318:1408–1412 2. Guelen L, Pagie L, Brasset E, Meuleman W, Faza MB, Talhout W, Eussen BH, de Klein A, Wessels L, de Laat W et al (2008) Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453:948–951 3. Solovei I, Wang AS, Thanisch K, Schmidt CS, Krebs S, Zwerger M, Cohen TV, Devys D, Foisner R, Peichl L et al (2013) LBR and lamin A/C sequentially tether peripheral heterochromatin and inversely regulate differentiation. Cell 152:584–598 4. Bergqvist C, Niss F, Figueroa RA, Beckman M, Maksel D, Jafferali MH, Kulyte A, Strom AL, Hallberg E (2019) Monitoring of chromatin organization in live cells by FRIC. Effects of the inner nuclear membrane protein Samp1. Nucleic Acids Res 47:e49 5. Berk JM, Tifft KE, Wilson KL (2013) The nuclear envelope LEM-domain protein emerin. Nucleus 4:298–314 6. Kalverda B, Fornerod M (2010) Characterization of genome-nucleoporin interactions in Drosophila links chromatin insulators to the nuclear pore complex. Cell Cycle 9:4812–4817 7. Kind J, Pagie L, Ortabozkoyun H, Boyle S, de Vries SS, Janssen H, Amendola M, Nolen LD, Bickmore WA, van Steensel B (2013)

Single-cell dynamics of genome-nuclear lamina interactions. Cell 153:178–192 8. Lee SK, Wang W (2019) Roles of topoisomerases in heterochromatin, aging, and diseases. Genes (Basel) 10:884 9. Villeponteau B (1997) The heterochromatin loss model of aging. Exp Gerontol 32:383–394 10. Larson K, Yan SJ, Tsurumi A, Liu J, Zhou J, Gaur K, Guo D, Eickbush TH, Li WX (2012) Heterochromatin formation promotes longevity and represses ribosomal RNA synthesis. PLoS Genet 8:e1002473 11. Andrenacci D, Cavaliere V, Lattanzi G (2020) The role of transposable elements activity in aging and their possible involvement in laminopathic diseases. Ageing Res Rev 57:100995 12. Goldman RD, Shumaker DK, Erdos MR, Eriksson M, Goldman AE, Gordon LB, Gruenbaum Y, Khuon S, Mendez M, Varga R et al (2004) Accumulation of mutant lamin A causes progressive changes in nuclear architecture in Hutchinson-Gilford progeria syndrome. Proc Natl Acad Sci U S A 101:8963– 8968 13. Carpenter AE, Jones TR, Lamprecht MR, Clarke C, Kang IH, Friman O, Guertin DA, Chang JH, Lindquist RA, Moffat J et al (2006) CellProfiler: image analysis software for identifying and quantifying cell phenotypes. Genome Biol 7:R100

Chapter 11 Analysis of Nuclear Pore Complexes in Caenorhabditis elegans by Live Imaging and Functional Genomics Patricia de la Cruz Ruiz, Raquel Romero-Bueno, and Peter Askjaer Abstract Nuclear pore complexes (NPCs) are essential to communication of macromolecules between the cell nucleus and the surrounding cytoplasm. RNA synthesized in the nucleus is exported through NPCs to function in the cytoplasm, whereas transcription factors and other proteins are selectively and actively imported. In addition, many NPC constituents, known as nuclear pore proteins (nucleoporins or nups), also play critical roles in other processes, such as genome organization, gene expression, and kinetochore function. Thanks to its genetic amenability and transparent body, the nematode Caenorhabditis elegans is an attractive model to study NPC dynamics. We provide here an overview of available genome engineered strains and FLP/Frt-based tools to study tissue-specific functions of individual nucleoporins. We also present protocols for live imaging of fluorescently tagged nucleoporins in intact tissues of embryos, larvae, and adult and for analysis of interactions between nucleoporins and chromatin by DamID. Key words Caenorhabditis elegans, CRISPR-Cas9, DamID, FLP, Live imaging, NPC, npp, Nuclear pore complex, Nucleocytoplasmic transport, Nucleoporin

1

Introduction The nematode Caenorhabditis elegans has been a popular and powerful model organism for more than half a century thanks to pioneering work by Sydney Brenner [1]. The transparency allows live observation of complex processes, such as organ development and neuronal activity in the intact organism with non-invasive techniques. Cell divisions are stereotypic in space and time, producing an invariant cell lineage [2, 3]. The development of C. elegans from egg through four larval stages (L1-L4) to fertile adult takes approximately 3 days. The adult hermaphrodite produces ~250 progeny over a time course of 3–5 days, and has a lifespan of 2–3 weeks. C. elegans embryos are surrounded by a resistant eggshell, which facilitates mounting for long time lapse observation

Patricia de la Cruz Ruiz and Raquel Romero-Bueno contributed equally to this work. Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_11, © Springer Science+Business Media, LLC, part of Springer Nature 2022

161

162

Patricia de la Cruz Ruiz et al.

from the zygote to the fast-moving three-fold embryo ready to hatch. Larvae and adults are small (up to ~1 mm) which enables high-resolution live imaging of individuals that can be either anaesthetized or immobilized by microfluidics devices [4] (Fig. 1). Despite their short lifespan, changes in nuclear envelope morphology are observed in old individuals, making C. elegans an attractive model to study aging [5]. C. elegans can easily be manipulated in large numbers and is thus very suitable for high-throughput forward and reverse genetic studies. This has revealed roles of C. elegans nucleoporins (nups; in C. elegans nomenclature known as Nuclear Pore Proteins or NPPs) in a variety of processes, such as germ granule distribution [6], transposon silencing [7], sensitivity to ionizing radiation [8], transposon silencing [7], and cell polarity [9]. Most nups are conserved in C. elegans (Table 1) and it is reasonable to assume that the three-dimensional structure of the NPC and its function in nucleocytoplasmic transport are also maintained [10, 11]. Similarly to the situation in other organisms, C. elegans nups have been shown by chromatin immunoprecipitation (ChIP) and DNA adenine methylation identification (DamID) to interact with the genome, suggesting that certain nups might be directly involved in regulation of gene expression [12–14]. Interestingly, recent single cell analyses suggest that the ratio between individual C. elegans nups varies significantly between cell types although this remains to be confirmed at the protein level [11, 15, 16]. Moreover, it has been proposed that certain nups are expressed only during embryogenesis and larval development and remain stably integrated in NPCs during the entire lifespan of the animal [17]. Transgenes can be inserted efficiently into specific sites in the genome of C. elegans and CRISPR technologies allow precise point mutations, gene knockouts, and insertion of short tags or genes encoding fluorescent proteins (Table 1) [18–20]. We have recently developed a versatile toolkit for FLP/Frt-mediated recombination in C. elegans, which enables spatiotemporal control of gene expression, cell ablation, and conditional knockout (Fig. 2 and Table 2) [21–23]. This has opened the possibility to perform tissue-specific DamID experiments to determine protein-DNA contacts in intestine and muscles [24, 25]. Efficient systems for inducible and rapid protein degradation are also available [26, 27] and the various tools can be further combined to increase their utility (Fig. 2). We provide here a protocol for live imaging of embryos, larvae, and adults, using strains with endogenously GFP-tagged nups as examples. Protocols for analysis of fixed embryos by immunofluorescence and electron microscopy are available elsewhere [28]. We also present details on how to identify chromatin domains bound by proteins of interest by DamID. Finally, we encourage the reader to consult excellent chapters on C. elegans biology and methods at WormBook (http://www.wormbook.org).

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans. . .

163

Fig. 1 Live imaging of endogenously tagged nucleoporins. (a) Schematic representation of the NPC. Nucleoporins are categorized based on their localization and form in several cases biochemically stable subcomplexes. Modified from [10]. (b) Expression levels of GFP-tagged MEL-28/ELYS, NPP-21/TPR, and NPP-24/NUP88 in hypodermal cells of L1 larvae were compared by confocal live microscopy. NPP-21/TPR is the most abundant of the three, followed by NPP-24/NUP88. Scale bar 10 μm. (b) During aging, the accumulation of MEL-28/ELYS at NPCs is diminished compared to the nuclear interior. (c) C. elegans is suitable for whole-animal imaging by live SPIM microscopy. In this example, NPP-21/TPR::GFP was observed on a MuVi Luxendo Light-Sheet Microscope. The magenta signal in (a) and (c) represents mCherry-tagged histone H2B (HIS-58)

164

Patricia de la Cruz Ruiz et al.

Table 1 C. elegans nucleoporins

C. elegans Human

Mutant allelesa

FP knock-inb

Other Other strainsc reagentsd Reference

MEL-28

ELYS/ AHCTF1

tm2434; t1578; t1684

NPP-1

NUP54

syb207

NPP-2

NUP85

tm2199

NPP-3

NUP205

ok1999

NPP-4

NUPL1

ok617

XA3548 (GFP)

Aff; Y2H

[43, 44, 46]

NPP-5

NUP107

ok1966; tm3039

BN69 (GFP)

Abs

[47]

NPP-6

NUP160

ok2821; tm4329

NPP-7

NUP153

ok601

WLP799 (GFP)

JH2686 (GFP)

Abs

[48, 49]; Pintard lab

NPP-8

NUP155

tm2513

WLP800 (mCh)

XA3546e (GFP)

Abs

[46]; Pintard lab

NPP-9

NUP358

gk3059

YY1566 (RFP) JH2184 (GFP)

Abs

[49–51]

JH2458 (GFP)

Abs

[47–49]

Abs

[47, 48]

Aff; Y2H

[43, 44, 52] [53, 54]; Askjaer lab

NPP-10Nf NUP98

BN426 (GFP) BN208 (Dam) Abs

LW1089 (GFP); OCF22 (mCh);

Aff; Y2H

BN1044 (GFP)

[13, 34, 41]

[42–44]

Askjaer lab Abs; Y2H [43, 45]

ok467

NPP-10Cf NUP96 NPP-11

NUP62

ok1599; gk5507

NPP-12

NUP210

ok2424; tm2320

BN1136 (GFP), DG4460 (GFP)

Abs

NPP-13

NUP93

ok1534

WLP805 (GFP)

Abs; Y2H [43, 45]; Pintard lab

NPP-14

NUP214

ok1389; sm160

dsRed

NPP-15

NUP133

ok1954

BN75 (GFP)

NPP-16

NUP50

ok1839; tm1596

GFP

[55] Abs

[17, 47]

Abs

[12] (continued)

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans. . .

165

Table 1 (continued)

C. elegans Human

Mutant allelesa

NPP-17/ RAE-1

RAE1

ok1720; tm2784; tm2796

NPP-18

SEH1

ok3278

NPP-19

NUP35

tm2886

NPP-20

SEC13R

NPP-21

TPR

NPP-22/ NDC1/ NDC-1 TMEM48

FP knock-inb

BN1015 (GFP)

Other Other strainsc reagentsd Reference mCh

Aff

BN46 (GFP), BN510 (Dam)

Abs; Y2H [47, 57]; Askjaer lab

EGD496 (GFP) tm1541; tm2952

BN1062 (GFP)

tm1845

DG4557 (GFP), WLP801 (mCh)

[56]

[58] Askjaer lab

EU1485 (GFP), BN375 (Dam) BN150 (GFP)

Abs

[54, 59, 60]; Pintard lab; Askjaer lab

NPP-23

NUP43

[47]

NPP-24

NUP88

BN739 (GFP)

Askjaer lab

NPP-25

TMEM33

BN1089 (GFP)

Askjaer lab

NPP-26

GLE1

NPP-27

ZC3HC1

BN1369 (GFP)

Askjaer lab

No clear C. elegans homologues were found for the mammalian nups AAAS/ALADIN, NUP37, NUP188, NUPL2/ hCG1, POM121 a Only selected alleles are listed. Most of these and other alleles are available from the Caenorhabditis Genetics Center (CGC; University of Minnesota; https://cgc.umn.edu) 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) b-c Strains expressing nucleoporin fusion proteins, either endogenously taggedb or as transgenec; Dam, DNA adenine methyltransferase; GFP, green fluorescent protein; mCh, mCherry; RFP, red fluorescent protein. Strain names are indicated when known; strains available from CGC are written in bold d Abs, antibodies; Aff, expression of affinity-tagged protein; Y2H, plasmids to study yeast two hybrid interactions e Prone to germ-line silencing f 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

166

Patricia de la Cruz Ruiz et al.

A

GFP Prom

ex1

ex3

ex2 Frt

gene of interest ex4

ex1

exn

B

GFP Prom

3’UTR

ex1

ex3

ex2

gene of interest ex4

ex1

exn

Frt

Frt

3’UTR Frt

+ FLP

+ FLP

Tissue-specific frame shift and nonsense-mediated mRNA decay

C

Tissue-specific complete gene knockout

D

hsp-16.41p

mCherry

his-58 3’UTR vhhGFP4::zif-1

Frt

3’UTR

hsp-16.41p

Frt

mCherry

his-58 3’UTR

Frt

+ FLP

Dam

gene of interest 3’UTR

Frt

+ FLP vhhGFP4::ZIF-1

Tissue-specific, heat-inducible degradation of GFP-tagged proteins

Dam fusion protein

Tissue-specific expression of Dam fusion protein in non-induced conditions

Fig. 2 Strategies for analysis of C. elegans nups by FLP/Frt-mediated genome recombination. (a) Insertion of a GFP cassette containing Frt sites in introns 1 and 2 of GFP immediately upstream of a gene of interest serves two purposes [21]. Firstly, the gene of interest will be tagged endogenously with GFP, enabling the study of its expression by live imaging. Secondly, co-expression of FLP in a specific tissue or at a particular moment during development will excise the second exon of GFP, thereby introducing a reading frame shift and a premature termination codon that together will abolish expression of the gene of interest. Several of the GFP knock-in strains in Table 1 contains this GFP variant. (b) For FLP-controlled complete removal of the gene of interest, an upstream GFP cassette with one or two Frt sites can be combined with insertion of a Frt site in the 3’ UTR of the gene of interest. (c) Inducible degradation of GFP-tagged protein using a vhhGFP4::zif-1 transgene [27] downstream of a Frt-flanked mCherry::HIS-58 stop cassette. VHHGFP4::ZIF-1 triggers ubiquitination of GFP-tagged proteins, thus enabling rapid and specific degradation in a spatiotemporal controlled manner [62]. (d) DamID experiments for mapping of protein-DNA contacts rely on low expression of Dam fusion proteins [14, 29]. The FLP/Frt system allows tissue-specific basal expression of Dam fusion proteins from an uninduced heat shock promoter [24, 25]

2 2.1

Materials Live Imaging

1. C. elegans strains expressing a protein(-s) of interest fused to a fluorescent protein(-s) (see Table 1). 2. Pipette and tips (10–1000 μL). 3. Glass slides (25 mm
75 mm
1 mm). 4. Label tape. 5. Cover slips (12 mm
12 mm and 22 mm
22 mm). 6. Worm pick with platinum wire. 7. Eyelash mounted on toothpick or Pasteur pipette. 8. 2% agarose solution in distilled water, melted in microwave oven and kept molten in a heat block at 65  C.

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans. . .

167

Table 2 Strains stably expressing codon-optimized FLP Promoter

Cell type

Strain

Referencea

ckb-3

Somatic gonad

BN854

[62]

ceh-60

Multiple neurons and intestine

BN813

[61]

dat-1

Dopaminergic neurons

BN617

[21]

dpy-7

Hypodermal

BN551

[21]

eat-4

Glutamatergic neurons

BN993

Unpublished

gpa-14

Neurons

BN816

[62]

hlh-8

M lineage

BN502

[21]

hlh-12

Distal tip cell

BN1204

[62]

hsp-16.41

Ubiquitous; heat inducible

BN646

[21]

lag-2

Multiple

BN558

[21]

lin-31

P lineage

BN1023

[62]

mec-7

Mechanosensory neurons

BN498

[21]

mex-5

Germ line

BN711

[22]

myo-2

Pharyngeal muscle

BN543

[21]

myo-3

Body wall muscle

BN503

[21]

nhr-82

Seam cell lineage

BN455

[21]

nhx-2

Intestine

BN999

[62]

rgef-1

Pan-neuronal

BN507

[21]

tph-1

Serotonin-producing neurons

BN499

[21]

UAS

Gal4-compatible

BN908

[23]

unc-17

Cholinergic neurons

BN1123

Unpublished

unc-47

GABAergic motor neurons

BN544

[21]

unc-122

Coelomocytes

BN1029

[62]

a

[62] strains can be requested from the corresponding author

9. M9 buffer: 22 mM KH2PO4, 34 mM Na2HPO4, 86 mM NaCl, 1 mM MgSO4. 10. Meiosis buffer: 25 mM HEPES pH 7.4, inulin 0.5 mg/mL, Leibovitz L-15 medium 60%, fetal bovine serum 20%. 11. Levamisole stock solution: 100 mM tetramisole hydrochloride (e.g. Sigma-Aldrich L9756). Dilute 1:10 in M9 buffer to obtain 10 mM working solution. 12. VALAP: 1:1:1 mixture of Vaseline or petroleum jelly, lanolin, and paraffin. Melts at 60  C.

168

Patricia de la Cruz Ruiz et al.

13. Potassium phosphate 1 M pH 6: 132 mM KH2PO4, 868 mM KH2PO4. 14. NGM plates (25 mM potassium phosphate pH 6, 51 mM NaCl, 1 mM CaCl2, 1 mM MgSO4, 17 g/L agar, 2.5 g/L peptone, 5 mg/L cholesterol) seeded with OP50 Escherichia coli bacteria (see Note 1). 2.2

DamID

1. C. elegans strains expressing E. coli Dam fused to either a protein of interest (test protein) or GFP (normalization control) (see Table 1 and Note 2). 2. NGM plates seeded with E. coli that does not express Dam, for instance GM119. 3. M9 buffer: 22 mM KH2PO4, 34 mM Na2HPO4, 86 mM NaCl, 1 mM MgSO4. 4. Hypochlorite solution: 1 N NaOH, 30% household bleach solution. 5. DNeasy Blood and Tissue Kit (QIAGEN #69504). 6. RNAse A (Qiagen #19101). 7. DpnI enzyme with 10
CutSmart buffer (NEB R0176S, 20 U/μL). 8. T4 DNA ligase (Roche #10799009001, 5 U/μL). 9. DpnII enzyme with 10xDpnII reaction buffer (NEB R0543S, 10 U/μL). 10. Taq DNA Polymerase with ThermoPol Buffer (NEB M0267S, 5 U/μL). 11. dNTP mix 2.5 mM (Takara SD0696). 12. Primer AdRt 50 CTAATACGACTCACTATAGGG CAGCGTGGTCGCGGCCGAGGA (100 μM). 13. Primer AdRb 50 -TCCTCGGCCG (100 μM). 14. AdR primers: combine equal volumes of: – 4 N: 50 -NNNNGTCCTCGCGGCCGAGGATC (50 μM) – 5 N: 50 -NNNNNGTCCTCGCGGCCGAGGATC (50 μM) – 6 N: (50 μM).

50 -NNNNNNGTCCTCGCGGCCGAGGATC

15. QIAquick® PCR Purification Kit (QIAGEN #28104). 16. NEBNext® Ultra DNA Library Prep Kit for Illumina® (NEB E7370). 17. NEBNext® Multiplex Oligos for Illumina® (Index Primers Set 1 E7335S; for up to 12 pooled samples. Combine with Index Primers Set 2 E7500S for up to 24 pooled samples). 18. SpeedBead Magnetic Carboxylate (GE Healthcare #65152105050250).

Modified

Particles

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans. . .

169

19. Ethanol 100% as well as freshly prepared 70% and 80% (store at 20  C). 20. Sodium acetate 3 M (pH 5.2). 21. QUBIT® fluorometer and dsDNA HS Assay Kit (Invitrogen Q32854). 22. Tubes suitable for QUBIT (e.g. DNA LoBind Tube 0.5 mL; Eppendorf #0030108035). 23. Thermocycler. 24. Magnetic particle concentrator (e.g. DynaMag-PCR; Invitrogen #492025). 25. Glycogen 20 mg/mL (Roche #10901393001).

3

Methods

3.1 Live Imaging of Whole Animals

Live imaging requires both restriction of movement during image acquisition and preservation of cell and tissue integrity. If a developmental process is to be studied for tens of minutes or even hours, particular care has to be devoted to establishing mounting conditions that do not interfere with the process, nor cause photo toxicity or bleaching by sample illumination (see Note 3). 1. Place two glass slides labeled with tape in parallel separated by a third clean microscope slide on a flat surface. 2. Using a pipette, place approximately a 50 μL drop of 2% melted agarose onto the center of the clean slide. 3. Instantly, cover the agarose drop with another glass slide in perpendicular way to make agarose flat before it solidifies. Avoid bubbles in the agarose to ensure a flat, intact pad (see Note 4). 4. Once agarose is solidified, move away the labeled slides and detach the two remaining slides carefully so that the agarose pad stays on one of them. Place the slide on the bench with the agarose facing upwards. 5. Place a 3 μL drop of 10 mM levamisole onto the center of the agarose pad (see Note 5). 6. Transfer 2–3 animals using a worm pick into the drop where they will float off. To avoid transfer of excess bacteria, pick worms from a bacteria-free zone of the plate or transfer worms to a plate without bacteria before making the agarose pad. Relocate the animals in the desired positions with the eyelash while the drop progressively dries up around them. Immediately place the coverslip before the drop disappears and refill the space under the coverslip with M9 buffer to avoid desiccation of the animals (see Note 6).

170

Patricia de la Cruz Ruiz et al.

7. Seal the coverslip with melted VALAP. This is most easily done with a fine paintbrush. 8. Acquire microscopy images at 20–24  C within 30 min after mounting the sample (see Note 7). 9. Recover the animals from the agarose pad if they are to be analyzed again later. Gently remove the coverslip while observing the animals in a stereoscope. Add 5 μL of M9 buffer and transfer the animals to a 10 μL drop of M9 on an NGM plate seeded with OP50 bacteria (see Note 1). 3.2 Live Imaging of Embryos

The C. elegans embryo is protected by a robust eggshell and can easily be prepared for live imaging. However, it requires some practice to obtain early 1-cell stage embryos (see Note 8). 1. Prepare a slide with a 2% agarose pad as described in steps 1–4 above (Subheading 2.1). 2. Transfer two young adult hermaphrodites to a 3 μL drop of M9 buffer on a 12-mm square glass coverslip (see Notes 9 and 10). Selection of L4 stage animals the day before imaging ensures having semi-synchronized young adults. 3. Use two syringe needles (e.g. 25–26 GA) to cut the animals in the middle to release the embryos from the uterus. 4. Lower gently the agarose pad (upside down) until it contacts the drop containing the embryos. The coverslip will adhere to the agarose pad. 5. Optionally, seal the coverslip using melted VALAP if the recording is planned to last for long time (more than 40 min). 6. Using a low-magnification objective, quickly select a suitable embryo and switch to a 60
or 100
oil objective for recording with the best possible image resolution using fluorescent microscopy combined with DIC imaging (see Notes 11–14).

3.3

DamID

The protocol presented here is optimized for using ~4000 nematodes per genotype and replica (see Note 15). 1. Collect embryos from asynchronous cultures by standard hypochlorite treatment (see Notes 16 and 17). 2. Leave embryos to hatch overnight at 16–20  C in M9 with gentle agitation (~120 rpm). 3. Determine the number of hatched L1 larvae and unhatched embryos in 2–10 μL aliquots. The hatching rate should be higher than 70%. 4. Aliquot ~1000 L1s onto 100 mm NGM plates containing Dam-negative E. coli as food source (see Note 18).

Analysis of Nuclear Pore Complexes in Caenorhabditis elegans. . .

171

5. Incubate nematodes at a suitable temperature for your experiment until they reach the desired life stage and collect them with M9 in a conical 15 mL tube. 6. Wash the nematodes 10 times with 15 mL of M9. Pellet them by centrifugation at 800
g for 1 min in each wash. Extensive washes are essential to avoid E. coli sequences in the final library. 7. Perform five rounds of freezing/thawing in liquid nitrogen and a 37  C water bath or thermoblock to break the animals’ cuticle. 8. Purify genomic DNA using the DNeasy Blood and Tissue Kit (including the optional RNase treatment step). Handle the DNA with care to avoid shearing, which can lead to amplification of unspecific sequences at later steps in the protocol. Elute the DNA in two steps: first in 100 μL and next in 200 μL. Measure DNA concentration using QUBIT. Mix both eluates if needed and precipitate the DNA if the concentration is 2 weeks, cleistothecia (fruiting bodies) will start to appear on the plate. Under a dissecting microscope, carefully move the cleistothecia onto water agar plates using a glass needle and gently roll them over to remove any attached mycelia. Be very gentle in this step to prevent breakage of cleistothecia and premature release of ascospores. Once cleaned, transfer the cleistothecia to a 1.5 mL tube filled with 1 mL 0.2% tween80 and break it with a toothpick to release ascospores. The ascospores are now ready for plating to analyze the meiotic progeny and can be stored at 4  C. 7. Plate some of the ascospores on MM to test if they have crossed. If the strains have crossed, growth will be observed. If there was no crossing, the parent strains will not be able to grow on MM due to auxotrophy for the different nutritional markers. 8. If they have crossed, perform genotyping by plating on media with different nutrients to identify the progeny of interest. In this case, for example, the test would be to look for pyrG+ and pyroA+ prototrophs. 9. The progeny of interest are streaked 2–3 times on selective media to isolate clonal colonies. We now have cells of interest that have GFP and GBP fusion proteins. 3.5 Microscopy of Strains Expressing GBP Fused Anchor and GFP Fused Protein of Interest

1. The final strains with the GBP fused anchor protein and GFP fused protein of interest can now be examined by microscopy. For quick screening, an epifluorescence microscope can be used. For more extensive imaging to test protein–protein linking through the GFP-GBP system, we recommend live cell imaging using a spinning disc confocal microscope to minimize photodamage during imaging. 2. For quick screening, the cells of interest can be cultured on a coverslip. To do this, spread parafilm on a sterile 10 cm plate and place sterile coverslips on it. Up to 4–5 coverslips can be placed in a single petri dish. Mix a small amount of spores with MM + 5 V (no riboflavin), add 300 μL of the suspension on the coverslip and incubate for 12–14 h at room temperature. The surface tension will keep a dome of media on the coverslips if 41) for a single typical experiment. **p < 0.01

GraphPad and reveal significantly ( p < 0.01) higher specific nuclear fluorescence for the c-Fos-c-Jun pair, compared to the FosΔzip-c-Jun samples. 3.5.2 Quantitative In-Cell Assessment of IMPα-Cargo Interaction Using the BiFC System

IMPα-NLS-cargo interaction can also be assessed in the BiFC system. 1. After transfection of the appropriate BiFC constructs, CLSM imaging is used to visualize reconstituted Venus fluorescence in the cell samples. As Fig. 5a shows, nuclear fluorescence is evident in cells cotransfected to express IMPα and NS5 (top panels), confirming interaction of the two in a cellular context. 2. The specificity of interaction is indicated by the fact that an NLS-binding site mutant of IMPα (IMPα2K192R396) shows markedly reduced nuclear fluorescence (Fig. 5a bottom panels). 3. As indicated above, IMPα only binds to cargoes with high affinity when heterodimerized with IMPβ1 through binding of the IBB of IMPα by IMPβ1; if IMPβ1 is not bound, the IBB prevents high affinity IMPα-NLS interaction. The IMPα2ΔIBB construct is deleted for the IBB domain, and so can bind NLS-containing cargoes with high affinity. Figure 5b

226

Alexander Lee et al.

Fig. 5 IMPα interaction with the NLS-containing protein DENV NS5 in the BiFC system; demonstration of specificity. HeLa cells transfected to express the

In-Cell Interaction of Importin α with Cargo Proteins

227

highlights this in a cellular context, where, although IMPα interaction with NS5 results in robust nuclear fluorescence (top panels), stronger nuclear fluorescence is evident for cells expressing IMPα2ΔIBB coexpressed with NS5 (bottom panels). 4. Image analyses (Fig. 5a, b lower panels) supports these results; IMPα recognition of NS5 is significantly ( p < 0.05) reduced by mutations in the NLS-binding site of IMPα (IMPα2K192R396 construct—left), and significantly enhanced ( p < 0.01) in the absence of the IBB domain in IMPα (IMPα2ΔIBB construct— right). 5. Interactions can be validated using BiFC constructs where DV2 NS5 and IMPα2 are expressed in the opposite BiFC vector (NS5 into VN and IMPα constructs into VC). As shown in Fig. 5c, d, results for these construct combinations are essentially identical to those in the other construct orientation, with NS5-IMPα interaction reduced by the K192R396 mutation (Fig. 5c), and IMPα2 lacking IBB (IMPα2ΔIBB) shows increased interaction (Fig. 5d). 6. Quantitative analysis (Fig. 5c, d lower panels) confirms statistically significant ( p < 0.01) differences for both comparisons. 7. Interaction of IMPα2 with other cargoes can also be tested. The NLS of T-ag (see Subheading 1) is the best understood IMPα-recognized NLS, and shown to be functional as a nuclear targeting module out of context of T-ag in a whole range of expression systems, including in transfected cells [22, 23]. Figure 6 shows in-cell analysis for interaction in the BiFC system of IMPα2 with a construct containing four tandem, in-frame copies of the T-ag NLS. Nuclear fluorescence due to T-ag-Interaction with IMPα2 is clearly evident, with the IMPα2ΔIBB construct showing increased nuclear fluorescence (Fig. 6a). Quantitative analysis confirms statistically significant ( p < 0.001) difference in the extent of interaction of T-ag with IMPα2ΔIBB compared to IMPα2 (Fig. 6b).

ä Fig. 5 (continued) indicated BiFC constructs were CLSM imaged 18 h later. Representative images and results for quantitation of specific nuclear fluorescence, in this case relative to VN-IMPα2 complementing with VC-NS5 for a, b; and VC-IMPα2 complementing with VN-NS5 for c, d. Results represent the mean  SEM for a single typical experiment. Results were for (a) n ¼ 57, 52 (b) n ¼ 68, 92 (c) n ¼ 60, 67 (d) n ¼ 60, 73 nuclei per sample. *p < 0.05; **p < 0.01

228

Alexander Lee et al.

Fig. 6 IMPα interaction with the T-ag-NLS-containing protein in the BiFC system; demonstration of specificity. Mutations affect the interaction and nuclear accumulation of IMPα2 with 4NLS. HeLa cells transfected to express the indicated BiFC constructs were CLSM imaged 18 h later. (a) Representative images are shown are for BiFC fluorescence (yellow, left), and BiFC and Hoechst nuclear stain (blue) merge image (right). (b) Quantitation of specific nuclear fluorescence relative to VN-IMPα2 complementing with VC-4xNLS. Results represent the mean  SEM for a single typical experiment (n ¼ 43, 58 nuclei per sample). ***p < 0.001 3.5.3 Quantitative In-Cell Assessment of Inhibition of IMPα-Cargo Interaction Using the BiFC System

The BiFC assay can be used to test the effects of inhibitors on in-cell interactions between two proteins. 1. In the case of IMPα2-NLS interactions, HeLa cells can be treated without or with an inhibitor, such as the IMPα targeting agent ivermectin (see Sect. 1). Ivermectin is known to bind IMPα directly and inhibit binding to NLSs as well as IMPβ1 [34]; this is the basis of ivermectin inhibiting infection by a number of different viruses [34–38]. 2. Figure 7 shows results for analysis in the BiFC system, demonstrating that ivermectin reduces the specific nuclear fluorescence of IMPα2 with 4xNLS and DENV NS5 (Fig. 7a, b top panels). 3. Quantitative analysis shows a significant ( p < 0.05) 60% reduction in binding (Fig. 7a, b bottom panels) [34], consistent with the idea that the BiFC system can be used to test inhibitors.

3.6

Summary

BiFC is a powerful tool to study protein–protein interactions, and can be used to examine interactions of IMPα with NLS-containing cargoes in a cellular context. Importantly, it is possible to analyze interactions quantitatively. Interaction specificity can be assessed by examining interaction of derivatives of IMPα either with mutations that impair NLS recognition, or with the autoinhibitory IBB

In-Cell Interaction of Importin α with Cargo Proteins

229

Fig. 7 IMPα interaction with NLS-containing proteins can be inhibited by ivermectin in the BiFC system. HeLa cells transfected to express the indicated BiFC constructs were CLSM imaged 18 h later. Cells were treated with either DMSO or 25 μM Ivermectin overnight post-transfection. Specific nuclear fluorescence was determined by dividing the average fluorescence of the sample relative to the control. Representative images for BiFC fluorescence (yellow, left), and BiFC and Hoechst nuclear stain (blue) merge image (right). Results for quantitation of specific nuclear fluorescence relative to VC-4xNLS complementing with VN-IMPα2 treated with DMSO (a, b) are graphed. Results represent the mean  SEM for a single typical experiment; results were for (a) n ¼ 73, 25 (b) n ¼ 88, 31 nuclei per sample. *p < 0.05; ***p < 0.001

domain deleted. Clearly, the system can be used to compare recognition of different NLSs, in different cell-type contexts of interest, and potentially in the absence and presence of the activation of signaling pathways/phosphorylation and so on. Excitingly, it is possible to use inhibitors of IMPα such as ivermectin in the BiFC system. This can help confirm the specificity of IMPα-cargo binding, but perhaps more interestingly, be used to assess the efficacy of various different inhibitors of the IMPα-NLS interaction. In principle, it may be possible to design a high throughput screening protocol around the assay, in the search for new inhibitors of interest, that in turn may have antiviral, anticancer, or other activities [27].

230

Alexander Lee et al.

Finally, it seems reasonable to speculate that the BiFC system can be used to assess other protein–protein interactions relevant to the nuclear transport process in a cell. These could include the recognition by other IMPs (e.g., IMPβ1) of their cargoes, potentially IMP-nucleoporin interaction, and inhibitors thereof. The power of the BiFC system in this context is that these interactions can be analyzed quantitatively in a living cell, in the context of the many competing interactions.

4

Notes 1. For Gibson cloning method, primers include sequences from both insert and vector. A 15–20 nucleotide with a melting temperature ~55  C is recommended for the insert and vector sequences in the primer. Ensure that the insert is in frame with both the identification tag (N terminal) as well as the Venus protein (C terminal). Forward and reverse primers should have as close to equivalent as possible Tm in the sequences that bind the insert to ensure efficient PCR reaction. Primers should end with either C or G on both ends to promote binding. The GC content should be between 40% and 60%. As here, restriction enzyme sites should be chosen from the multiple cloning sites of the vectors. 2. Gibson cloning kit was used to simplify and speed up the cloning process for multiple constructs of interest. Traditional ligation method can be used for cloning however primer design would be slightly different. 3. Lipid transfection was used to introduce the plasmids into HeLa cells. Although other methods (e.g., electroporation) may be used, we found that this method resulted in high transfection rate and most cells were cotransfected with both plasmids of interest. 4. Gibson recommends total 0.02–0.2 pmols of fragments with 10 μL of Gibson Assembly Master Mix (2) for a total volume of 20 μL. Optimal cloning efficiency of 50–100 ng of vector with two- to threefold of excess inserts. If purified fragments were of low yield, this may not be practically feasible. Therefore two- to threefold volume of insert relative to vector for a total of 10 μL can be used. 5. Electroporation is performed using 0.1 cm electrode gap cuvette in a Gene Pulser. 2 μL of Gibson reaction mix is added to 40 μL DH10B competent cells. Cells are pulsed at 1.6 kV, 200 Ω and 25 μF. 1 mL of LB media is immediately added and incubated at 37  C for 1 h. Centrifuge at 16,000  g for 1 min. Remove 800 μL of media. Resuspend cells and plate on agar plate containing appropriate antibiotic.

In-Cell Interaction of Importin α with Cargo Proteins

231

6. Forward sequencing primer binding to CMV promoter and reverse sequencing primer binding to either VC155 or VN155 (I152L) are used. 7. For overnight drug treatment, drugs are added at the same time as transfection. For timed treatments, samples can be treated at specified times prior to imaging, but imaging would usually be 18–24 h post-infection. Dimethyl sulfoxide (DMSO) is used as the solvent of ivermectin. Equal total volume is added to control and drug treatment. 8. Minimum size is estimated based on the usual size of the nucleus of a HeLa cells. Under our conditions 550 pixel units was suitable, ignoring small cells or cells undergoing apoptosis. 9. Background fluorescence is determined from the nucleus of a healthy cell stained with Hoechst 33342, but does not show Venus fluorescence. References 1. Nagai T, Ibata K, Park ES, Kubota M, Mikoshiba K, Miyawaki A (2002) A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat Biotechnol 20(1):87–90. https:// doi.org/10.1038/nbt0102-87 2. Kodama Y, Hu C-D (2010) An improved bimolecular fluorescence complementation assay with a high signal-to-noise ratio. BioTechniques 49(5):793–805. https://doi.org/ 10.2144/000113519 3. Shyu YJ, Hu C-D (2008) Fluorescence complementation: an emerging tool for biological research. Trends Biotechnol 26(11):622–630. https://doi.org/10.1016/j.tibtech.2008. 07.006 4. Hemerka JN, Wang D, Weng Y, Lu W, Kaushik RS, Jin J, Harmon AF, Li F (2009) Detection and characterization of influenza A virus PA-PB2 interaction through a bimolecular fluorescence complementation assay. J Virol 83(8):3944–3955. https://doi.org/10.1128/ jvi.02300-08 5. Hu C-D, Chinenov Y, Kerppola TK (2002) Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Mol Cell 9(4):789–798. https://doi.org/10. 1016/s1097-2765(02)00496-3 6. Bischof J, Duffraisse M, Furger E, Ajuria L, Giraud G, Vanderperre S, Paul R, Bjo¨rklund M, Ahr D, Ahmed AW, Spinelli L, Brun C, Basler K, Merabet S (2018) Generation of a versatile BiFC ORFeome library for analyzing protein–protein interactions in live

Drosophila. eLife 7:e38853. https://doi.org/ 10.7554/eLife.38853 7. Kerppola TK (2008) Bimolecular fluorescence complementation (BiFC) analysis as a probe of protein interactions in living cells. Annu Rev Biophys 37:465–487. https://doi.org/10. 1146/annurev.biophys.37.032807.125842 8. Patel LR, Curran T, Kerppola TK (1994) Energy transfer analysis of Fos-Jun dimerization and DNA binding. Proc Natl Acad Sci U S A 91(15):7360–7364. https://doi.org/10. 1073/pnas.91.15.7360 9. Sjo¨hamn J, Ba˚th P, Neutze R, Hedfalk K (2016) Applying bimolecular fluorescence complementation to screen and purify aquaporin protein:protein complexes. Protein Sci 25(12):2196–2208. https://doi.org/10. 1002/pro.3046 10. Kodama Y, Hu C-D (2012) Bimolecular fluorescence complementation (BiFC): a 5-year update and future perspectives. BioTechniques 53(5):285–298. https://doi.org/10.2144/ 000113943 11. Kerppola TK (2006) Design and implementation of bimolecular fluorescence complementation (BiFC) assays for the visualization of protein interactions in living cells. Nat Protoc 1(3):1278–1286. https://doi.org/10.1038/ nprot.2006.201 12. Kerppola TK (2013) Multicolor bimolecular fluorescence complementation (BiFC) analysis of protein interactions with alternative partners. Cold Spring Harbor Protocols 2013(9): pdb.top077164. https://doi.org/10.1101/ pdb.top077164

232

Alexander Lee et al.

13. Miller KE, Kim Y, Huh W-K, Park H-O (2015) Bimolecular fluorescence complementation (BiFC) analysis: advances and recent applications for genome-wide interaction studies. J Mol Biol 427(11):2039–2055. https://doi. org/10.1016/j.jmb.2015.03.005 14. Ohashi K, Kiuchi T, Shoji K, Sampei K, Mizuno K (2012) Visualization of cofilinactin and Ras-Raf interactions by bimolecular fluorescence complementation assays using a new pair of split Venus fragments. BioTechniques 52(1):45–50. https://doi.org/10.2144/ 000113777 15. Lin J, Wang N, Li Y, Liu Z, Tian S, Zhao L, Zheng Y, Liu S, Li S, Jin C, Xia B (2011) LEC– BiFC: a new method for rapid assay of protein interaction. Biotech Histochem 86(4): 2 7 2 – 2 7 9 . h t t p s : // d o i . o r g / 1 0 . 3 1 0 9 / 10520295.2010.483068 16. Nakagawa C, Inahata K, Nishimura S, Sugimoto K (2011) Improvement of a Venusbased bimolecular fluorescence complementation assay to visualize bFos-bJun interaction in living cells. Biosci Biotechnol Biochem 75(7): 1399–1401 17. Isogai M, Kawamoto Y, Inahata K, Fukada H, Sugimoto K, Tada T (2011) Structure and characteristics of reassembled fluorescent protein, a new insight into the reassembly mechanisms. Bioorg Med Chem Lett 21(10): 3021–3024. https://doi.org/10.1016/j. bmcl.2011.03.039 18. Poon IKH, Jans DA (2005) Regulation of nuclear transport: central role in development and transformation? Traffic 6(3):173–186. https://doi.org/10.1111/j.1600-0854.2005. 00268.x 19. Wagstaff KM, Jans DA (2009) Importins and beyond: non-conventional nuclear transport mechanisms. Traffic 10(9):1188–1198. https://doi.org/10.1111/j.1600-0854.2009. 00937.x 20. Lonhienne TG, Forwood JK, Marfori M, Robin G, Kobe B, Carroll BJ (2009) Importin-beta is a GDP-to-GTP exchange factor of Ran: implications for the mechanism of nuclear import. J Biol Chem 284(34): 22549–22558. https://doi.org/10.1074/jbc. M109.019935 21. Pryor MJ, Rawlinson SM, Butcher RE, Barton CL, Waterhouse TA, Vasudevan SG, Bardin PG, Wright PJ, Jans DA, Davidson AD (2007) Nuclear localization of dengue virus nonstructural protein 5 through its importin α/β–recognized nuclear localization sequences is integral to viral infection. Traffic 8(7): 795–807. https://doi.org/10.1111/j. 1600-0854.2007.00579.x

22. Hu¨bner S, Xiao C-Y, Jans DA (1997) The protein kinase CK2 site (Ser111/112) enhances recognition of the Simian virus 40 large T-antigen nuclear localization sequence by importin. J Biol Chem 272(27): 17191–17195. https://doi.org/10.1074/jbc. 272.27.17191 23. Xiao C-Y, Hu¨bner S, Jans DA (1997) SV40 large tumor antigen nuclear import is regulated by the double-stranded DNA-dependent protein kinase site (Serine 120) flanking the nuclear localization sequence. J Biol Chem 272(35):22191–22198. https://doi.org/10. 1074/jbc.272.35.22191 24. Yasuda Y, Miyamoto Y, Yamashiro T, Asally M, Masui A, Wong C, Loveland KL, Yoneda Y (2012) Nuclear retention of importin α coordinates cell fate through changes in gene expression. EMBO J 31(1):83–94. https:// doi.org/10.1038/emboj.2011.360 25. Bello´n-Echeverrı´a I, Carralot J-P, Del Rosario AA, Kueng S, Mauser H, Schmid G, Thoma R, Berger I (2018) MultiBacMam bimolecular fluorescence complementation (BiFC) tool-kit identifies new small-molecule inhibitors of the CDK5-p25 protein-protein interaction (PPI). Sci Rep 8(1):5083. https://doi.org/10.1038/ s41598-018-23516-x 26. Fulcher AJ, Jans DA (2011) Regulation of nucleocytoplasmic trafficking of viral proteins: an integral role in pathogenesis? Biochim Biophys Acta 1813(12):2176–2190. https://doi. org/10.1016/j.bbamcr.2011.03.019 27. Jans DA, Martin AJ, Wagstaff KM (2019) Inhibitors of nuclear transport. Curr Opin Cell Biol 58:50–60. https://doi.org/10.1016/j.ceb. 2019.01.001 28. Gregory WM, David AJ (2012) Editorial [Hot Topic: Subcellular trafficking of pathogens: targeting for therapeutics (Guest Editors: Gregory W. Moseley and David A. Jans)]. Infect Disord Drug Targets 12(1):2–3. https://doi. org/10.2174/187152612798994975 29. Caly L, Ghildyal R, Jans DA (2015) Respiratory virus modulation of host nucleocytoplasmic transport; target for therapeutic intervention? Front Microbiol 6:848. https:// doi.org/10.3389/fmicb.2015.00848 30. Fontoura BM, Faria PA, Nussenzveig DR (2005) Viral interactions with the nuclear transport machinery: discovering and disrupting pathways. IUBMB Life 57(2):65–72. h t t p s : // d o i . o r g / 1 0 . 1 0 8 0 / 15216540500078608 31. Ghildyal R, Ho A, Dias M, Soegiyono L, Bardin PG, Tran KC, Teng MN, Jans DA (2009) The respiratory syncytial virus matrix protein possesses a Crm1-mediated nuclear export

In-Cell Interaction of Importin α with Cargo Proteins mechanism. J Virol 83(11):5353–5362. https://doi.org/10.1128/jvi.02374-08 32. Ito N, Moseley GW, Blondel D, Shimizu K, Rowe CL, Ito Y, Masatani T, Nakagawa K, Jans DA, Sugiyama M (2010) Role of interferon antagonist activity of rabies virus phosphoprotein in viral pathogenicity. J Virol 84(13):6699. https://doi.org/10.1128/jvi. 00011-10 33. Yang SNY, Atkinson SC, Fraser JE, Wang C, Maher B, Roman N, Forwood JK, Wagstaff KM, Borg NA, Jans DA (2019) Novel flavivirus antiviral that targets the host nuclear transport importin α/β1 heterodimer. Cell 8(3):281. https://doi.org/10.3390/cells8030281 34. Yang S, Atkinson S, Wang C, Lee A, Bogoyevitch M, Borg N, Jans D (2020) The broad spectrum antiviral ivermectin targets the host nuclear transport importin α/β1 heterodimer. Antivir Res 177:104760. https://doi. org/10.1016/j.antiviral.2020.104760 35. Tay MYF, Fraser JE, Chan WKK, Moreland NJ, Rathore AP, Wang C, Vasudevan SG, Jans DA (2013) Nuclear localization of dengue virus (DENV) 1–4 non-structural protein 5; protection against all 4 DENV serotypes by the inhibitor Ivermectin. Antivir Res 99(3): 301–306. https://doi.org/10.1016/j. antiviral.2013.06.002

233

36. Fraser JE, Watanabe S, Wang C, Chan WKK, Maher B, Lopez-Denman A, Hick C, Wagstaff KM, Mackenzie JM, Sexton PM, Vasudevan SG, Jans DA (2014) A nuclear transport inhibitor that modulates the unfolded protein response and provides in vivo protection against lethal dengue virus infection. J Infect Dis 210(11):1780–1791. https://doi.org/10. 1093/infdis/jiu319 37. Wang C, Yang SNY, Smith K, Forwood JK, Jans DA (2017) Nuclear import inhibitor N-(4-hydroxyphenyl) retinamide targets Zika virus (ZIKV) nonstructural protein 5 to inhibit ZIKV infection. Biochem Biophys Res Commun 493(4):1555–1559. https://doi.org/10. 1016/j.bbrc.2017.10.016 38. Caly L, Druce JD, Catton MG, Jans DA, Wagstaff KM (2020) The FDA-approved drug ivermectin inhibits the replication of SARS-CoV2 in vitro. Antiviral Res 178:104787. https:// doi.org/10.1016/j.antiviral.2020.104787 39. Shyu YJ, Liu H, Deng X, Hu C-D (2006) Identification of new fluorescent protein fragments for bimolecular fluorescence complementation analysis under physiological conditions. BioTechniques 40(1):61–66. https://doi.org/10.2144/000112036

Chapter 15 Validation of Nuclear Pore Complex Protein–Protein Interactions by Transient Expression in Plants Fumika Ikeda and Kentaro Tamura Abstract The nuclear pore complex (NPC) is the largest protein complex, consisting of multiple copies of over 30 different nucleoporins. The interactions between the nucleoporins are critical elements for the NPC functions of the nuclear envelope in plant cells. In recent years, transient expression-based validations of protein-protein interactions have been widely used in plants. Bimolecular fluorescence complementation assay and coimmunoprecipitation assays are powerful tools to identify the molecules that interact with specific proteins. Here, as an example, we describe these techniques using nucleoporin protein interactions in plants. Key words Arabidopsis thaliana, Nuclear pore complex, Bimolecular fluorescence complementation, Coimmunoprecipitation, Agroinfiltration, Nicotiana tabacum, Nicotiana benthamiana

1

Introduction The nuclear envelope (NE) is composed of the outer nuclear membrane, the inner nuclear membrane, and the nuclear pore complex (NPC). The NPC is a large multiprotein complex that opens up an ffi50 nm-wide channel. It consists of multiple copies of over 30 different nucleoporins that share a similar protein domain structure. It has been estimated that 38% of nucleoporins contain an α-solenoid fold composed of 2–3 α-helix units, 29% contain phenylalanine– glycine (FG) repeats that provide interaction domains to form an elastic and hydrogel-like structure, and 16% contain β-propeller folds that function as a scaffold for the NPC [1]. The nucleoporins assemble into several different subcomplexes, including a transmembrane ring, core scaffold (inner ring, outer ring, and linker), cytoplasmic filaments, nuclear basket, and central barrier [2]. Multiple copies of these subcomplexes oligomerize into a unique higher-order structure. The NPC mediates nucleocytoplasmic exchange and trafficking that is highly regulated and involved in a considerably broader range of cellular activities in all eukaryotes

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_15, © Springer Science+Business Media, LLC, part of Springer Nature 2022

235

236

Fumika Ikeda and Kentaro Tamura

[3]. In plants, each nucleoporin has been shown to have diverse and specific functions, including regulation of development and responses to environmental stimuli [4–6]. Therefore, the structure of NPCs in plants has been subjected to intensive research. Due to the limited sequence similarity of nucleoporins in plants, interactome approaches have been the most successful strategies in understanding the NPC structure [7]. Bimolecular fluorescence complementation (BiFC) assay and coimmunoprecipitation assays are powerful tools to identify molecules that interact with specific proteins. The BiFC assay is based on protein-fragment complementation methods in living cells. The bait and prey proteins to be tested for interaction are fused to two fragments of a reporter fluorescent protein, neither of which by itself exhibits fluorescence activity. During the interaction, the bait and prey proteins bring the two fragments together and reconstitute the fluorescent protein. This reconstitution is normally irreversible; thus, transient and weak interactions can be detected. In the coimmunoprecipitation assay, the bait protein is precipitated with a specific antibody, and the interactors are identified by Western blotting. This assay can purify the stable protein complex, allowing efficient identification of whole protein components. Here, we describe methods to validate the interaction between nucleoporins on the NE in plants. Using transient expression in tobacco epidermal cells that have a higher transformation frequency than other systems [8], it is possible to perform high throughput validation for protein-protein interactions. These approaches are also a convenient and powerful method for revealing proteome-wide interactome maps that provide significant insights into the functions of unknown proteins.

2

Materials

2.1 Plant Expression Vectors for the BiFC Assay and Coimmunoprecipitation

Suitable Gateway technology-compatible binary vectors are available [9, 10] and can be used to insert the nucleoporin sequences of interest by standard methods. 1. BiFC Binary Vectors: YFP N Fragment Fused to Either N-Terminal (pB4NY0) or C-terminal (pB4NY2) of the nucleoporin of interest (e.g., Arabidopsis Nup136, and YFP C fragment fused to either N-terminal (pB4CY0) or C-terminal (pB4CY2) of the nucleoporin of interest (e.g., Arabidopsis Nup82) [9] (Table 1). 2. Internal control vector for BiFC: A nuclear-localized RFP (e.g., Histone-tagRFP) is used to determine BiFC efficiency and subcellular localization.

NPC Protein-Protein Interactions in Plants

237

Table 1 BiFC and Co-IP binary vectors used to verify the interactions between nucleoporins in planta Name

Fusion protein

BiFC binary vectors

pB4NY0 pB4NY2 pB4CY0 pB4CY2

nYFP-nucleoporin nucleoporin-nYFP cYFP-nucleoporin nucleoporin-cYFP

Co-IP binary vectors

pGWB405 pGWB406 pGWB454 pGWB455

nucleoporin-GFP GFP-nucleoporin nucleoporin-RFP RFP-nucleoporin

3. Coimmunoprecipitation (Co-IP) binary vectors for: GFP fused to either N-terminal (pGWB 405) or C-terminal (pGWB 406) of the nucleoporin of interest, and RFP fused to either N-terminal (pGWB 454) or C-terminal (pGWB 455) of the nucleoporin of interest [10] (Table 1). 2.2

Agroinfiltration

1. Agrobacteria (GV3101::pMP90) were transformed using a binary vector containing a BiFC construct of the nucleoporins of interest. 2. Infiltration medium: acetosyringone.

1

mg/mL

D-glucose,

0.1

mM

3. 1 mL syringes without a needle. 4. 1- to 2-month-old Nicotiana tabacum or Nicotiana benthamiana plants (see Note 1). 2.3 BiFC Assay of Nucleoporins

1. Agrobacteria transformed Subheading 2.1).

with

BiFC

constructs

(see

2. A confocal microscope with lasers and filters to image the BiFC signal. 2.4 Coimmunoprecipitation of Nucleoporins

1. Transient expression of fusion proteins in tobacco leaves.

2.4.1 Immunoprecipitation

1. Lysis buffer: 50 mM Hepes-KOH (pH 7.5), 150 mM NaCl, 0.5% (v/v) Triton X-100, 0.1% (v/v) Tween 20. Store at 4  C.

2. Agrobacteria transformed with either GFP or RFP fusion protein constructs (see Subheading 2.1).

2. Wash buffer: 50 mM Hepes-KOH (pH 7.5). Store at 4  C. 3. Elution buffer: 100 mM Tris-HCl (pH 6.8), 2% (w/v) sodium n-dodecyl sulfate, 20% (w/v) glycerol, 5% (v/v) 2-mercaptoethanol, 0.005% (w/v) bromophenol blue.

238

Fumika Ikeda and Kentaro Tamura

4. Magnetic beads conjugated to an anti-GFP antibody (e.g., μMACS Anti-GFP MicroBeads, Miltenyi Biotec, Germany) (see Note 2). 5. Columns and magnetic separator (e.g., Miltenyi μColumns and μMACS Separator). 2.4.2 Reagents for Sodium Dodecyl SulfatePolyacrylamide Gel Electrophoresis (SDSPAGE) and Immunoblotting

1. Running gel buffer: 1.5 M Tris-HCl (pH 8.8). 2. Stacking gel buffer: 1 M Tris-HCl (pH 6.8). 3. 30% acrylamide/bis-acrylamide mixed solution (29:1). 4. Ammonium persulfate: 10% (w/v) solution in water. Make aliquots and store at 20  C (see Note 3). 5. N,N,N,N0 -tetramethyl-ethylenediamine (TEMED). Store at 4  C. 6. SDS-PAGE running buffer: Dissolve 3.04 g Tris, 14.42 g glycine, and 1 g SDS in 1 L water. 7. Blotting buffer: Dissolve 3 g Tris, 14.4 g glycine, 1 g SDS, and 200 mL methanol in 800 mL water (see Note 4). 8. Tris-buffered saline with Tween 20 (TBST): 50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 0.05% (v/v) Tween 20. 9. Blocking buffer: 5% (w/v) skimmed milk in TBST. 10. Antibodies: Anti-GFP monoclonal antibody (e.g., clone JL-8, Clontech, USA), anti-RFP monoclonal antibody (e.g., clone RF5R, ThermoFisher, USA), HRP-linked anti-mouse IgG sheep antibody (e.g., GE, USA). Store at 4  C.

3

Methods

3.1 Preparation of Vectors and Agrobacterium Transformation

1. Generate constructs. Each nucleoporin gene of interest is cloned into entry vectors (e.g., pENTR1A). After verification of the gene sequences in the entry vector, the gene of interest is cloned into the binary vectors by LR recombination reaction. 2. Culture Agrobacterium cells in 50 mL of LB media at 28  C until OD600 ¼ 0.5–0.8. 3. Wash the cells twice with 20 mL of water. 4. Resuspend the cells in 1 mL of 10% glycerol. 5. Transform the cells with binary vector (10 mg/mL using centrifugal concentrators with 10 kDa MWCO. Flash-freeze using liquid nitrogen and store in
80  C until use.

3.3 Purification of MBP-NES

1. Grow 1 L of BL21 E. coli cells expressing MBP-NES in LB with 100 μg/mL ampicillin (shaking at 230 rpm) to OD 0.4–0.6 at 37  C. Induce expression with 0.4 mM IPTG for 10 h at 25  C or 3 h at 37  C with 200 rpm shaking. Chill to 4  C until collection. 2. Centrifuge to pellet the E. coli cells, resuspend the pellets in 20 mL/L growth of Lysis C buffer, and freeze at
20  C until use. 3. Perform steps 3 and 4 from Subheading 3.1. 4. Equilibrate ~10 mL of amylose resin beads with Lysis C buffer. 5. Add supernatant of clarified lysate to amylose resin and collect the flow-through. Return the flow-through to the amylose resin column for the second time, and collect a second flowthrough.

Measurement of CRM1-NES Binding Affinities

251

6. Wash the column with 50 mL of Q A buffer (see Note 11). 7. Elute MBP-NES with 50 mL of Q A buffer that contains 10 mM maltose. 8. Purify the eluted MBP-NES by ion exchange chromatography (HiTrap Q) using Q A and Q B buffer with a gradient of 0–100% Q B over 20–30 column volume. 9. Use SDS-PAGE to visualize proteins in the HiTrap Q chromatogram peaks and concentrate the fractions that contain MBP-NES using 30 kDa MWCO centrifugal concentrators. 10. Use a Superdex 75 column that is equilibrated with TB buffer to exchange the buffer of MBP-NES into TB buffer. 11. Use SDS-PAGE to visualize proteins in the Superdex 75 chromatogram peaks and concentrate the fractions containing clean MBP-NES to >10 mg/mL using 30 kDa MWCO centrifugal concentrators, flash-freeze and store at
80  C until use. 3.4 Fluorescence Polarization Assay

1. Dialyze CRM1, RanGTP, and MBP-NES overnight into TB buffer. 2. Collect the proteins and some TB buffer from the dialysis device. Measure the concentration of the proteins using the collected TB buffer as the blank sample. 3. For direct titration of CRM1 to the FITC-PKINES probe, dilute CRM1 to 20 μM in 50 μL volume in a PCR tube at room temperature, supplementing the mixture with 0.05% Tween 20 (see Note 12) (Fig. 1, step I). 4. Pipet 25 μL of TB buffer with 0.05% Tween 20 into 15 more PCR tubes at room temperature (Fig. 1, step II). 5. Take 25 μL from the master-mix in step 3 and serially dilute the CRM1 15 times using the tubes in step 4 at room temperature (Fig. 1, step III). 6. Distribute 7.5 μL from each of the 16 PCR tubes containing serially-diluted CRM1 into three columns of wells in the 384-well plate in room temperature (Fig. 1, step IV). 7. Make a 2 master-mix containing 40 nM FITC-PKINES and 120 μM RanGTP in TB buffer with 0.05% Tween 20 in a total volume of 400 μL, which is sufficient for one set of titrations done in triplicate (see Note 13) (Fig. 1, step V). 8. Mix 7.5 μL of the 2 master mix from step 7 to the wells of 2 mix distributed in step 6 to yield final 1 reaction mixes, 15 μL each, at room temperature (Fig. 1, step VI). 9. Incubate the reaction mixes for ~1 h at room temperature in the dark, before performing measurements in the plate reader. 10. For competition titrations of MBP-NES to CRM1-FITCPKINES, perform procedures described in steps 3–6 to

252

Ho Yee Joyce Fung and Yuh Min Chook

Fig. 1 Schematic for setting up 384-well plate FP assay. Step I: Make 2 master mix for 20 μM CRM1 (direct titration) or 100 μM MBP-NES (competition titration) in a PCR tube (50 μL volume). Step II: Pipet 25 μL of TB in each of 15 more PCR tubes. Step III: Serially dilute the 2 master mix from step I using the 15 tubes made in step II. Step IV: Distribute the titration series from step III into three columns of the plate (16  3 wells). Step V: Make another 2 master mix with 40 nM FITC-PKINES and 120 μM Ran (direct) or 40 nM FITC-PKINES, 120 μM Ran and 300 nM CRM1 (competition) in an microcentrifuge tube (400 μL volume). Step VI: Distribute the mixes from step V across the three columns for their respective titrations (16  3 wells)

generate serially-diluted MBP-NES in TB buffer with 0.05% Tween 20, starting with 100 μM in 50 μL (see Notes 12 and 14) (Fig. 1, steps I to IV). 11. Make a 2 master mix with 40 nM FITC-PKINES, 120 μM RanGTP, and 300 nM CRM1 in TB buffer with 0.05% Tween 20 in total volume of 400 μL, which is sufficient for one triplicate titration. Increase the volume according to the number of MBP-NES titrations you want to perform (see Note 15) (Fig. 1, step V). 12. Mix 7.5 μL of the 2 master mix from step 11 to the triplicate set of the serially diluted MBP-NES from step 10 to yield final 1 reaction mixes of 15 μL each (Fig. 1, step VI). 13. Incubate reaction mixes for at least 1.5 h at room temperature in the dark before performing measurements. 14. Perform fluorescence polarization measurements using plate reader (see Note 16).

Measurement of CRM1-NES Binding Affinities

253

15. Using PALMIST (see Note 17) or another analysis program of your choice, calculate the polarization (P) of each reaction using the measured parallelly (Ik) and perpendicularly I
I (I⊥) polarized intensities with the equation P ¼ I kk þI ⊥⊥ , averaging your data across triplicates. Fit the direct and competition binding data points with the appropriate binding model (1:1 A + B* for direct binding and B* and C competes for A in competition binding) to obtain the affinities for the FITCPKINES (B*) and the MBP-NES (C) to CRM1 (A) and their associated confidence intervals/errors.

4

Notes 1. This protocol describes the purification of full length wild-type human CRM1. Changes in the expression or purification protocol may be needed if applied to other CRM1 constructs. 2. This ScRan construct stabilizes the GTP-bound state of Ran, purifies easily and is easy to handle biochemically. 3. We usually add two to three residues before and after the consensus match (from Φ0 to Φ4), for example: XXΦ0XXΦ1XXXΦ2XXΦ3XΦ4XX for a class 1a NES. Residues that flank the NES likely affect secondary structural propensity of the peptide and thus could affect binding affinity for CRM1 [7]. It is better to be consistent and include the same number of residues before and after the NES consensus. 4. Yields can range from 0.4 to 2 mg/L growth depending on mutations included. Since the yield is low, we purify protein from at least 4 L (if not 6–12 L) of E. coli at a time. 5. During the first pass through the cell disruptor, pressure applied can be lower to prevent blockage, but peak pressure for the last past should reach ~10,000–15,000 psi. Sample should be chilled during homogenization. 6. If the TEV protease is stored in more than 10% (v/v) glycerol, it is important to ensure that the final glycerol percentage in the TEV cleavage reaction does not exceed 10%. Cleaving the fusion protein with TEV overnight at 4  C will decrease the final yield. 7. Elution can be continued if there is protein remaining in the elution after two elutions of 15 mL each. However, large volumes of eluted protein will increase the time needed to concentrate CRM1, which in turn will lead to significantly lower yields of the protein. 8. There is often a peak of aggregated CRM1 and a second peak of monomeric CRM1. It is acceptable to inject more sample into the size-exclusion column as long as the two peaks remain

254

Ho Yee Joyce Fung and Yuh Min Chook

separated. Completing the purification within 24 h from lysis of bacteria is important for optimal yield and activity of the protein. 9. Stop concentrating the protein if the concentrate starts to turn yellow. Raise the salt level in the buffer to ~150 mM sodium chloride to clarify a cloudy concentrate. 10. Since the construct is engineered to stabilize the GTP-bound state, you may omit the EDTA stripping/GTP reloading step and instead add GTP to the protein at every step of the purification. 11. Protease inhibitors can be added to buffer used in steps 6–8 if proteolytic degradation of the MBP-NES is observed. 12. Using this method, pipetting error in the titration series will not be captured and the error captured will be mostly instrumental. In order to capture pipetting error, the triplicate serial dilution should be performed separately, but this is very labor intensive if many titrations need to be performed. To do so, you can make a CRM1 or MBP-NES solution with 2 the maximum concentration in the first well of a 384-well plate in 15 μL instead of a PCR tube in step 3, then pipet 7.5 μL TB buffer into the 15 other wells in the same column, perform the serial dilution (remember to discard 7.5 μL from the last well of the serial dilution), and then repeat the same process two more times to generate a triplicate set. 13. Direct titration is performed on each day of the experiment to ensure that the CRM1 and RanGTP proteins used are of similar quality. Data from the direct titration is used to fit data for the competition mode experiments performed that same day. The fitted dissociation constant for the probe should be ~30 nM. The RanGTP concentration in the assay is kept at 60 μM to be consistent with the previous set-up where the maximum CRM1 concentration used was 20 μM and thus there is 3 molar excess Ran [7]. The current assay has a maximum CRM1 concentration of 10 μM, and thus Ran concentration can be lowered to 30 μM. 14. For NESs that bind CRM1 with lower affinities, if possible, do start the serial dilution at a higher concentration so that MBP-NES can displace all of the FITC-PKINES probe bound to CRM1 and capture a complete sigmoidal curve when the data is fitted. 15. 150 nM of wild-type CRM1 is appropriate to establish sufficient dynamic range and good baseline signal from FITCPKINES-bound CRM1. This concentration should be adjusted for CRM1 variants.

Measurement of CRM1-NES Binding Affinities

255

16. Using our CLARIOstar plus instrument and Corning 384-well plates, we measure FP using top optic, 50 flashes per well at focal height 5.1 mm with a 0.1 s settling time. Gain is adjusted using the first well with target mP of 200 and kept constant for the rest of the measurements. 17. We use PALMIST as it is the original software for analysis of our MST data, which has a competition mode to fit each MBP-NES titration in conjunction with the direct titration performed on the same day with F-statistics and error-surface projection method for error estimation [12–14]. This program has been configured to analyze 384-well plates FP data by importing a excel file with the raw and calculated polarization data exported in plate format from the CLARIOstar data analysis software.

Acknowledgments We thank Jordan Baumhardt and Bini Shakya for their assistance. This work is funded by the Cancer Prevention Research Institute of Texas (CPRIT) Grants RP180410 and RP170170 (Y.M.C.), Welch Foundation Grant I-1532 (Y.M.C), NIGMS of NIH under Award R01GM069909 (Y.M.C.), the Mabel and Alfred Gilman Chair for Molecular Pharmacology (Y.M.C.), and the Eugene McDermott Scholar in Biomedical Research (Y.M.C.). References 1. Fu SC, Fung HYJ, Cagatay T, Baumhardt J, Chook YM (2018) Correlation of CRM1-NES affinity with nuclear export activity. Mol Biol Cell 29(17):2037–2044. https://doi.org/10. 1091/mbc.E18-02-0096 2. Engelsma D, Bernad R, Calafat J, Fornerod M (2004) Supraphysiological nuclear export signals bind CRM1 independently of RanGTP and arrest at Nup358. EMBO J 23(18): 3643–3652. https://doi.org/10.1038/sj. emboj.7600370 3. Paraskeva E, Izaurralde E, Bischoff FR, Huber J, Kutay U, Hartmann E, Luhrmann R, Gorlich D (1999) CRM1mediated recycling of snurportin 1 to the cytoplasm. J Cell Biol 145(2):255–264. https:// doi.org/10.1083/jcb.145.2.255 4. Askjaer P, Bachi A, Wilm M, Bischoff FR, Weeks DL, Ogniewski V, Ohno M, Niehrs C, Kjems J, Mattaj IW, Fornerod M (1999) RanGTP-regulated interactions of CRM1 with nucleoporins and a shuttling DEAD-box

helicase. Mol Cell Biol 19(9):6276–6285. https://doi.org/10.1128/mcb.19.9.6276 5. Guttler T, Madl T, Neumann P, Deichsel D, Corsini L, Monecke T, Ficner R, Sattler M, Gorlich D (2010) NES consensus redefined by structures of PKI-type and Rev-type nuclear export signals bound to CRM1. Nat Struct Mol Biol 17(11):1367–1376. https://doi. org/10.1038/nsmb.1931 6. Koyama M, Matsuura Y (2010) An allosteric mechanism to displace nuclear export cargo from CRM1 and RanGTP by RanBP1. EMBO J 29(12):2002–2013. https://doi. org/10.1038/emboj.2010.89 7. Fung HY, Fu SC, Brautigam CA, Chook YM (2015) Structural determinants of nuclear export signal orientation in binding to exportin CRM1. elife 4:e10034. https://doi.org/10. 7554/eLife.10034 8. Fung HY, Fu SC, Chook YM (2017) Nuclear export receptor CRM1 recognizes diverse conformations in nuclear export signals. elife 6:

256

Ho Yee Joyce Fung and Yuh Min Chook

e23961. https://doi.org/10.7554/eLife. 23961 9. Monecke T, Haselbach D, Voss B, Russek A, Neumann P, Thomson E, Hurt E, Zachariae U, Stark H, Grubmuller H, Dickmanns A, Ficner R (2013) Structural basis for cooperativity of CRM1 export complex formation. Proc Natl Acad Sci U S A 110(3):960–965. https://doi.org/10.1073/ pnas.1215214110 10. Petosa C, Schoehn G, Askjaer P, Bauer U, Moulin M, Steuerwald U, Soler-Lopez M, Baudin F, Mattaj IW, Muller CW (2004) Architecture of CRM1/Exportin1 suggests how cooperativity is achieved during formation of a nuclear export complex. Mol Cell 16(5): 761–775. https://doi.org/10.1016/j.molcel. 2004.11.018 11. Askjaer P, Jensen TH, Nilsson J, Englmeier L, Kjems J (1998) The specificity of the CRM1Rev nuclear export signal interaction is

mediated by RanGTP. J Biol Chem 273(50): 33414–33422. https://doi.org/10.1074/jbc. 273.50.33414 12. Baumhardt JM, Walker JS, Lee Y, Shakya B, Brautigam CA, Lapalombella R, Grishin N, Chook YM (2020) Recognition of nuclear export signals by CRM1 carrying the oncogenic E571K mutation. Mol Biol Cell 31(17): 1879–1891. https://doi.org/10.1091/mbc. E20-04-0233 13. Tso SC, Chen Q, Vishnivetskiy SA, Gurevich VV, Iverson TM, Brautigam CA (2018) Using two-site binding models to analyze microscale thermophoresis data. Anal Biochem 540-541: 64–75. https://doi.org/10.1016/j.ab.2017. 10.013 14. Scheuermann TH, Padrick SB, Gardner KH, Brautigam CA (2016) On the acquisition and analysis of microscale thermophoresis data. Anal Biochem 496:79–93. https://doi.org/ 10.1016/j.ab.2015.12.013

Part V Post Translational Modifications

Chapter 17 Analysis of Ubiquitylation and SUMOylation of Yeast Nuclear Pore Complex Proteins Catherine Dargemont Abstract Posttranslational modifications and in particular ubiquitylation and SUMOylation of the nuclear pore complex (NPC), have been shown to regulate some of its functions, particularly in response to diverse stress signals. Although proteomic approaches are extremely powerful to identify substrates and modification sites, dissecting specific mechanisms and regulation functions of ubiquitylation and SUMOylation of the diverse NPC proteins, in different genetic backgrounds or cell environmental conditions, requires specific biochemical assays based on purification and precise analysis of 6His-tagged ubiquitylated or SUMOylated protein of interest. Here we describe an approach that can be easily employed without specific equipment. It allowed to successfully analyze yeast NPC proteins but can easily be adapted to the study of the mammalian NPC. Key words Nuclear pore complex, Ubiquitin, SUMO, Yeast, 6His-Tag purification

1

Introduction Transport of macromolecules between the nucleus and the cytoplasm of eukaryotic cells occurs through gigantic dedicated structures, called nuclear pore complexes (NPCs). Their overall molecular architecture presenting an eightfold rotational symmetry is highly conserved across species and within cell types. NPCs are organized by biochemically defined submodules- namely the coat nucleoporin complex, the inner ring complex, the central channel nucleoporin complex, the nuclear basket, the cytoplasmic domain and the transmembrane proteins that anchors the NPC within the nuclear envelope [1–4]. However, NPCs also display a high degree of heterogeneity and plasticity. In particular, NPC composition but also NPC density within the nuclear envelop can vary as a function of cell type, cell differentiation or aging [5–7]. Although mechanisms responsible for such a diversity have not yet been extensively dissected, studies

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_17, © Springer Science+Business Media, LLC, part of Springer Nature 2022

259

260

Catherine Dargemont

mostly performed over the past decade highlighted the role of posttranslational modifications in the dynamic changes of the NPC architecture and functions. Glycosylation by O-linked β-N-acetyl glucosamine (OGlcNAc) has been the first type of posttranslational modification described for nucleoporins. In metazoans, 16 of ~30 nucleoporins are potential substrates for this glycosylation process that is not only involved in the structural integrity of the NPC but also in its function as conduits in nucleocytoplasmic transport [8]. Phosphorylation of Nups by mitotic kinases participates to the NPC disassembly at the onset of prophase and also contributes to the proper expression and localization of nucleoporins involved in postmitotic NPC assembly [9–15]. In this respect, it has been recently shown that Tpr, a nuclear basket nucleoporin, recruits the extracellular signal-regulated kinase (ERK) that, in turn, controls the number of NPC by phosphorylating Nup153, a nuclear basket nucleoporin essential for early stages of NPC biogenesis [16, 17]. Phosphorylation of central channel nucleoporins (FG-Nups) can modulate the selective permeability of the NPCs [18–20] indicating that phosphorylation events not only affect the architecture but also alter the functions of the NPC. Of note, O-GlcNAcylation and phosphorylation can cross-talk as both modifications can occur on the same or closely located serine and threonine residues [8]. More recently, both yeast and human nucleoporins emerged as targets of acetylation [21, 22]. Interestingly, daughter cell-specific NPC deacetylation in asymmetrically dividing budding yeast, determines the time of commitment to a new division cycle [23]. Nucleoporins are also extensively subjected to dynamic conjugation to ubiquitin and ubiquitin-like proteins, and in particular SUMO. Covalent attachment of monoubiquitin, polyubiquitin chains or ubiquitin-like molecules is mediated through a thiolester cascade of reactions catalyzed by a ubiquitin/SUMO-activating enzyme (E1), a ubiquitin/SUMO-conjugating enzyme (E2), and a ubiquitin/SUMO-protein ligase (E3) [24, 25]. Notably, Nup358, a cytoplasmic mammalian nucleoporin, displays itself an E3 SUMO ligase activity, with RanGAP being its major substrate [26] and, in yeast, the SUMO-dependent ubiquitin ligase Slx8Slx5, associates with the NPC [27]. Both ubiquitin and SUMO moieties can me cleaved off by specific hydrolases. Interestingly, some mammalian (SENP1, SENP2) and yeast (Ulp1) SUMO proteases are specifically associated with the NPC [28–32]. The NPC cannot be considered as a single entity toward the ubiquitin/ SUMO conjugation system, but is rather the target of multiple ubiquitin/SUMO-modifying enzymes [33, 34]. A systematic analysis of the ubiquitylation of the nucleoporins recently conducted in S. cerevisiae revealed that more than 50% of the nucleoporins displayed a modification by ubiquitin [35]. Prevalence of

Ubiquitylation and SUMOylation of the Yeast NPC

261

Fig. 1 SUMOylation of yeast nuclear basket proteins is highly sensitive to cellular conditions. In absence of stress, Nup60, Nup1, and Nup2 are SUMoylated but the level of modification is rather low, particularly for Nup60. Upon genotoxic stress, SUMOylation of Nup60 increases whereas modification of Nup2 or Nup1 is under limit of detection. In contrast, upon osmotic stress, SUMOylation of Nup60, Nup2, and Nup1 strongly increases [38]

monoubiquitylation suggests broad functions of this posttranslational modification rather than a degradative role. Unlike ubiquitylation, SUMOylation does not trigger protein degradation. As shown by the role of monoubiquitylation of Nup159 in nuclear segregation at the onset of mitosis, one can predict that posttranslational modifications might be associated with regulation of described or yet unexplored functions of the NPC [35]. Over the past years, increasing number of studies indeed enlighten transport-independent functions of the NPC, including gene regulation, chromatin organization, DNA repair, RNA processing, RNA quality control, and cell cycle regulation [36, 37]. Posttranslational modifications of the yeast nuclear basket nucleoporins have been specifically analyzed upon cellular stresses including genotoxic and osmotic stresses (Fig. 1). Although major nuclear transport routes are not regulated by Nup60 nor Nup2 ubiquitin and SUMO modifications, monoubiquitylation of Nup60 controls the plasticity of the NPC nuclear basket and regulates the DNA damage response and telomere repair [34]. More generally, the posttranslational modification landscape of nuclear basket proteins is extremely sensitive to both genotoxic and osmotic stresses suggesting that the NPC acts as a stress sensor serving to transmit extracellular stress signals into the nucleus [38]. The various major posttranslational modifications including ubiquitylation and SUMOylation have been extensively studied through proteomic approaches, with thousands of substrates and modification sites being revealed. However, addressing questions mechanisms responsible for nucleoporin modifications, impact of the cell environment, the different layers of cross talk between

262

Catherine Dargemont

S.cerevisiae

Nup-HA

Nup-HA 6His-Ub 6His-SUMO

6His-Ub-Nup-HA 6His-SUMO-Nup-HA

Genomic tagging

Expression of 6His-Ub or 6His-SUMO

Ni-NTA column purificaon Western blong analysis

Fig. 2 Strategy of the approach. The Saccharomyces cerevisiae strains are HA-tagged at the nucleoporin genomic locus of interest. Cells are then transformed with a plasmid encoding 6His–ubiquitin or 6His–SUMO. 6His-modified cellular proteins are purified on Ni-NTA column and protein of interest is analyzed by western blotting using anti-HA antibody

nucleoporins and their modifications relies on specific approaches that allows to precisely dissect ubiquitylation or SUMOylation processes. Although fastidious, methods described here (summarized in Fig. 2) have been proven to be particularly well adapted to the systematic analysis of the yeast NPC, and in particular the low stoichiometry of modified proteins.

2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water) and analytical grade reagents. Prepare and store all reagents at room temperature. Buffers and reagents that need to be freshly prepared are indicated.

2.1

Buffers

1. Minimal synthetic media with drop-out medium supplements is used to create selective media for growing auxotrophic yeast cultures. Minimal synthetic medium contains yeast nitrogen base (1.7 g/L), ammonium sulfate (5 g/L), and glucose (2%). 2. Loading buffer (LB): 100 mM K2HPO4, 20 mM Tris-HCl pH 8.0, 100 mM NaCl, 6 M Guanidine Hydrochloride. This buffer can be stored at 4  C. Before use, add 10 mM imidazole, 10 mM β-mercaptoethanol, and 0.1% Triton X-100. 3. Wash Buffer: 10 mM Tris-HCl pH 6.4, 100 mM Na-phosphate buffer pH 6,4, 8 M Urea, 10 mM imidazole, 10 mM

Ubiquitylation and SUMOylation of the Yeast NPC

263

β-mercaptoethanol, and 0.1% Triton X-100. This buffer has to be freshly prepared. The 1 M stock sodium phosphate buffer pH 6.4 is obtained by mixing 735 mL of 1 M NaH2PO4 and 265 mL of 1 M Na2HPO4. 4. 2 SDS-Blue loading buffer: 125 mM Tris-HCl pH 6.8, 4% SDS, 20% glycerol, 10% β-mercaptoethanol, 0.004% Bromophenol Blue. 5. Trichloroacetic acid (TCA). 2.2

Equipment

1. Cell disrupter (e.g., MagNA Lyser, Roche Diagnostics). 2. Glass beads (0.5 mm diameter).

2.3

Antibodies

1. Anti-HA tag antibody (e.g., BioLegend HA-11, 0.75 μg/mL). 2. Anti-6His tag antibody (e.g. Millipore, 0.2 μg/mL). 3. Monoclonal anti-ubiquitin antibody (e.g., Santa Cruz Biotechnology). 4. Anti-SUMO antibody (polyclonal rabbit anti-Smt3; [39]).

3

Methods

3.1 Culture and 6HisUb or 6His-SUMO Induction

1. The Saccharomyces cerevisiae strains are HA-tagged at the nucleoporin genomic locus of interest using PCR-based homologous recombination ([40]; see Note 1). Cells transformed with a plasmid encoding 6His–ubiquitin or 6His– SUMO under the CUP1 promoter (such as YEp352-pCUP16His-Ub or YEp352-pCUP1-6His-SMT3, [34]; see Notes 2– 4) are grown at 30  C (unless strain of interest is temperaturesensitive) in synthetic media with the appropriate supplements. At the end of day 1, inoculate freshly transformed cells in 150 mL of selective media supplemented with 100 μM CuSO4 at 30  C ON (for temperature-sensitive cells, see Note 5). Don’t forget to inoculate cells transformed with empty vector as a negative control. 2. At day 2, collect 100 OD600 units (OD600 0.7–1.2) of each cell culture by centrifugation at 3000  g for 3 min at room temperature. Resuspend the pellet into 20 mL of H2O for each tube. Centrifuge the yeast at 3000  g for 3 min at room temperature. Resuspend cells in 1 mL H2O, transfer to 2 mL cap tubes, recover the cell pellets by centrifugation. Cell pellets can be stored at 80  C (see Note 6).

3.2 Lysate Preparation

1. Resuspend cell pellets corresponding to 100 OD600 units in 0.5 mL of 20% TCA. 2. Transfer the yeast suspension to a 2 mL cap tube (tube 1) containing 800 μL of glass beads (see Note 7).

264

Catherine Dargemont

3. Lyse the yeast cell suspensions with the cell disrupter for 60 s at 6000 rpm twice at 4  C (in the cold room). 4. Poke two holes in the bottom of the tubes with a heated 26-guage needle (one hole is not enough to pass through the tube). 5. Immediately place the tubes in another 2 mL tube (tube 2). 6. Centrifuge for 2 min at 1000  g with a swing-out rotor (regular cell culture centrifuge) at room temperature (the cell lysate is thus recovered in the recipient tube). 7. Add 1 mL of 7.5% TCA to the 2 mL cap tubes containing glass beads (tube 1) and centrifuge the tubes (tubes 1 in tubes 2) for 2 min at 1000  g at room temperature. 8. Add 0.2 mL of 10% TCA to the 2 mL of cap tubes containing glass beads (tube 1) and centrifuge the tubes (tubes 1 in tubes 2) for 2 min at 1000  g at room temperature. 9. Incubate the recipient tubes containing the complete supernatant (tube 2; approximately 1.7 mL) on ice for 45 min. Discard tubes 1. 10. Centrifuge the tubes for 10 min at 20,000  g at 4  C and remove the supernatant. 11. Gently wash pellets with 750 μL of 0.1% TCA (see Note 8). 12. Centrifuge the tubes for 10 min at 20,000  g at 4  C and remove the supernatant. At this stage, the pellets can be frozen (see Note 6). 13. Add 30 μL of 1 M Tris (unbuffered) to each pellet and 450 μL of Loading Buffer (LB) and resuspend the pellets. It is crucial to be careful and patient. Centrifuge the tubes for 10 min at 20,000  g at 4  C and collect the supernatants. 14. Add 450 μL LB to the remaining pellets and resuspend them. Centrifuge the tubes for 10 min at 20,000  g at 4  C and collect the supernatants. 15. Mix supernatants from steps 13 and 14. The final volume of cell lysates should be approximately 900 μL. Check the pH with pH indicator paper. The pH should be 7–8. If not, add 20 μL maximum of 1 M Tris (unbuffered). 16. Collect a 30 μL sample of each cell lysate and proceed to TCA precipitation (see Subheading 3.3). These samples will correspond to the input. 3.3 TCA Precipitation for the Input Samples

1. Take 30 μL of lysate corresponding to 3 OD yeast cell culture (step 15 of Subheading 3.2) to a 1.5 mL tube, add 240 μL of H2O, and 30 μL of 50% TCA, vortex. 2. Incubate the samples on ice for 30 min.

Ubiquitylation and SUMOylation of the Yeast NPC

265

3. Centrifuge the samples at 20,000  g for 15–30 min at room temperature. Gently discard the supernatant. 4. Add 150 μL of 1 Blue SDS-loading buffer (to 0.02 OD/μL) and boil at 95  C for 10 min under agitation. 3.4 6His-Ub/SUMO Purification

1. Transfer 80 μL of 50% slurry of Ni2+-nitrilotriacetic acid–agarose (Ni-NTA) beads (Qiagen) in a 1.5 mL tube and wash with 1 mL LB, centrifuge the tubes for 1 min at 1000  g at 4  C. Carefully discard supernatant with a micropipette and wash twice in LB. 2. Resuspend the beads in 80 μL LB to obtain 50% of Ni-NTA preequilibrated beads in buffer. Steps 1 and 2 should not be performed in advance. 3. Transfer the complete supernatants (step 15 of Subheading 3.2) to the 1.5 mL tubes containing 80 μL of Ni-NTA beads preequilibrated suspension and rotate for 1 h at room temperature. 4. Centrifuge the tubes for 2 min at 1000  g. Discard supernatant carefully with a micropipette and wash with 1 mL of the LB for 5 min at room temperature. 5. Centrifuge the tubes for 2 min at 1000  g. Discard supernatant carefully with a micropipette and wash with 1 mL of the Wash Buffer for 5 min at room temperature. Repeat this step twice. Spin down the tubes and remove the excess supernatant. 6. Elute with 50 μL of 2 SDS-Blue loading buffer for 5 min at 95  C under agitation. Centrifuge the samples at max speed for 1 min and transfer the supernatant to fresh tubes.

3.5 Analysis of 6HisUb/SUMO Modified Proteins

4

Cell lysates and Ni-purified 6His-Ubiquitin/SUMO–conjugated forms of nucleoporin-HA from cells transformed or not transformed with a plasmid encoding 6His-Ubiquitin/SUMO are separated on appropriate SDS–polyacrylamide gel electrophoresis gels and transferred to nitrocellulose or polyvinylidene difluoride membranes (see Note 9). Western blots are processed with an anti-HA antibody. Ubiquitin or SUMO expression and efficiency of purification was controlled using an anti-6His tag or an anti-Ubiquitin or anti-SUMO antibody (see Note 10) respectively (for an example, see Fig. 3).

Notes 1. Genomic tagging of yeast nucleoporin genes by homologous recombination allows to analyze the diverse nucleoporins using the same anti-tag antibody. To this respect, it is essential to use a tag that cannot be modified by Ubiquitin or SUMO which

266

Catherine Dargemont

Fig. 3 SUMOylation of Nup2. Ni-purified 6His-SUMO–conjugated forms of Nup2HA were purified from cells transformed (+) or not transformed ( ) with an expression plasmid of 6His-SUMO, treated or not with 200 mM hydroxy-urea (genotoxic/replicative stress). Cell lysates (top) and Ni-purified material (middle) were examined by Western blotting with an anti-HA antibody. SUMO expression and efficiency of purification were controlled using an anti-SUMO antibody (bottom)

essentially target lysine residues. For this reason, we generally use 3HA-tag that does not contain any lysine residue. 2. Ubiquitin or SUMO modifications are highly labile upon cell lysis. To preserve these modifications, we use 6His-tagged version of ubiquitin or SUMO in order to lyse cells in guanidium-containing buffer without affecting purification of modified proteins. 3. Plasmids with different auxotrophy or antibiotic-resistance genes are chosen depending on the yeast strain of interest. 4. To distinguish different kinds of ubiquitin linkage, polySUMOylation from multisite SUMOylation, plasmids encoding for 6His-Ubiquitin/SUMO mutants can be used in which specific or all lysines have been mutated into arginines (see for example [41]). 5. Strains containing a thermosensitive allele are grown at a permissive temperature (25  C) and subsequently shifted to the restrictive temperature for 3 h before harvesting for biochemical analysis.

Ubiquitylation and SUMOylation of the Yeast NPC

267

6. We observed that processing more than 8 samples for purification did not allow an optimal preservation of modifications and eventually led to irreproducible results. We thus advise to process samples in different batches in case more than 8 are required for the full experiment. 7. Glass beads should be sterile. 8. Be careful to not resuspend the pellets. 9. For a proper analysis, 5–10 μL of cell lysate (input) are sufficient whereas 15–20 of purification eluate are necessary. However, this has to be tested as it depends on the stoichiometry of both abundance of the protein of interest and its modification level. 10. From our experience, anti-6His Tag and anti-ubiquitin antibodies are similar in terms of sensitivity. In contrast, anti-Smt3 antibody is much more sensitive and appropriate to SUMOylation analysis than the anti-6His Tag. References 1. Knockenhauer KE, Schwartz TU (2016) The nuclear pore complex as a flexible and dynamic gate. Cell 164(6):1162–1171. https://doi. org/10.1016/j.cell.2016.01.034 2. Hoelz A, Glavy JS, Beck M (2016) Toward the atomic structure of the nuclear pore complex: when top down meets bottom up. Nat Struct Mol Biol 23(7):624–630. https://doi.org/10. 1038/nsmb.3244 3. Beck M, Hurt E (2017) The nuclear pore complex: understanding its function through structural insight. Nat Rev Mol Cell Biol 18(2): 73–89. https://doi.org/10.1038/nrm. 2016.147 4. Kim SJ, Fernandez-Martinez J, Nudelman I, Shi Y, Zhang W, Raveh B, Herricks T, Slaughter BD, Hogan JA, Upla P, Chemmama IE, Pellarin R, Echeverria I, Shivaraju M, Chaudhury AS, Wang J, Williams R, Unruh JR, Greenberg CH, Jacobs EY, Yu Z, de la Cruz MJ, Mironska R, Stokes DL, Aitchison JD, Jarrold MF, Gerton JL, Ludtke SJ, Akey CW, Chait BT, Sali A, Rout MP (2018) Integrative structure and functional anatomy of a nuclear pore complex. Nature 555(7697):475–482. https://doi.org/10.1038/nature26003 5. D’Angelo MA, Gomez-Cavazos JS, Mei A, Lackner DH, Hetzer MW (2012) A change in nuclear pore complex composition regulates cell differentiation. Dev Cell 22(2):446–458. https://doi.org/10.1016/j.devcel.2011. 11.021

6. Gomez-Cavazos JS, Hetzer MW (2015) The nucleoporin gp210/Nup210 controls muscle differentiation by regulating nuclear envelope/ ER homeostasis. J Cell Biol 208(6):671–681. https://doi.org/10.1083/jcb.201410047 7. Lupu F, Alves A, Anderson K, Doye V, Lacy E (2008) Nuclear pore composition regulates neural stem/progenitor cell differentiation in the mouse embryo. Dev Cell 14(6):831–842. https://doi.org/10.1016/j.devcel.2008. 03.011 8. Eustice M, Bond MR, Hanover JA (2017) O-GlcNAc cycling and the regulation of nucleocytoplasmic dynamics. Biochem Soc Trans 45(2):427–436. https://doi.org/10. 1042/BST20160171 9. Macaulay C, Meier E, Forbes DJ (1995) Differential mitotic phosphorylation of proteins of the nuclear pore complex. J Biol Chem 270(1): 254–262. https://doi.org/10.1074/jbc.270. 1.254 10. Glavy JS, Horwitz SB, Orr GA (1997) Identification of the in vivo phosphorylation sites for acidic-directed kinases in murine mdr1b P-glycoprotein. J Biol Chem 272(9): 5909–5914. https://doi.org/10.1074/jbc. 272.9.5909 11. Glavy JS, Krutchinsky AN, Cristea IM, Berke IC, Boehmer T, Blobel G, Chait BT (2007) Cell-cycle-dependent phosphorylation of the nuclear pore Nup107-160 subcomplex. Proc

268

Catherine Dargemont

Natl Acad Sci U S A 104(10):3811–3816. https://doi.org/10.1073/pnas.0700058104 12. Onischenko EA, Gubanova NV, Kiseleva EV, Hallberg E (2005) Cdk1 and okadaic acidsensitive phosphatases control assembly of nuclear pore complexes in Drosophila embryos. Mol Biol Cell 16(11):5152–5162. https://doi.org/10.1091/mbc.e05-07-0642 13. Lusk CP, Blobel G, King MC (2007) Highway to the inner nuclear membrane: rules for the road. Nat Rev Mol Cell Biol 8(5):414–420. https://doi.org/10.1038/nrm2165 14. Laurell E, Beck K, Krupina K, Theerthagiri G, Bodenmiller B, Horvath P, Aebersold R, Antonin W, Kutay U (2011) Phosphorylation of Nup98 by multiple kinases is crucial for NPC disassembly during mitotic entry. Cell 144(4):539–550. https://doi.org/10.1016/j. cell.2011.01.012 15. Maeshima K, Iino H, Hihara S, Funakoshi T, Watanabe A, Nishimura M, Nakatomi R, Yahata K, Imamoto F, Hashikawa T, Yokota H, Imamoto N (2010) Nuclear pore formation but not nuclear growth is governed by cyclin-dependent kinases (Cdks) during interphase. Nat Struct Mol Biol 17(9): 1065–1071. https://doi.org/10.1038/nsmb. 1878 16. McCloskey A, Ibarra A, Hetzer MW (2018) Tpr regulates the total number of nuclear pore complexes per cell nucleus. Genes Dev 32(19–20):1321–1331. https://doi.org/10. 1101/gad.315523.118 17. Vollmer B, Lorenz M, Moreno-Andres D, Bodenhofer M, De Magistris P, Astrinidis SA, Schooley A, Flotenmeyer M, Leptihn S, Antonin W (2015) Nup153 recruits the Nup107160 complex to the inner nuclear membrane for interphasic nuclear pore complex assembly. Dev Cell 33(6):717–728. https://doi.org/10. 1016/j.devcel.2015.04.027 18. Kehlenbach RH, Gerace L (2000) Phosphorylation of the nuclear transport machinery down-regulates nuclear protein import in vitro. J Biol Chem 275(23):17848–17856. https://doi.org/10.1074/jbc.M001455200 19. Kosako H, Imamoto N (2010) Phosphorylation of nucleoporins: signal transductionmediated regulation of their interaction with nuclear transport receptors. Nucleus 1(4): 309–313. https://doi.org/10.4161/nucl.1.4. 11744 20. Mishra A, Sipma W, Veenhoff LM, Van der Giessen E, Onck PR (2019) The effect of FG-Nup phosphorylation on NPC selectivity: a one-bead-per-amino-acid molecular dynamics study. Int J Mol Sci 20(3):596. https://doi. org/10.3390/ijms20030596

21. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M (2009) Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science 325(5942):834–840. https:// doi.org/10.1126/science.1175371 22. Henriksen P, Wagner SA, Weinert BT, Sharma S, Bacinskaja G, Rehman M, Juffer AH, Walther TC, Lisby M, Choudhary C (2012) Proteome-wide analysis of lysine acetylation suggests its broad regulatory scope in Saccharomyces cerevisiae. Mol Cell Proteomics 11(11):1510–1522. https://doi.org/10. 1074/mcp.M112.017251 23. Kumar A, Sharma P, Gomar-Alba M, Shcheprova Z, Daulny A, Sanmartin T, Matucci I, Funaya C, Beato M, Mendoza M (2018) Daughter-cell-specific modulation of nuclear pore complexes controls cell cycle entry during asymmetric division. Nat Cell Biol 20(4):432–442. https://doi.org/10. 1038/s41556-018-0056-9 24. Cappadocia L, Lima CD (2018) Ubiquitin-like protein conjugation: structures, chemistry, and mechanism. Chem Rev 118(3):889–918. https://doi.org/10.1021/acs.chemrev. 6b00737 25. Ovaa H, Vertegaal ACO (2018) Probing ubiquitin and SUMO conjugation and deconjugation. Biochem Soc Trans 46(2):423–436. https://doi.org/10.1042/BST20170086 26. Pichler A, Gast A, Seeler JS, Dejean A, Melchior F (2002) The nucleoporin RanBP2 has SUMO1 E3 ligase activity. Cell 108(1): 109–120. https://doi.org/10.1016/s00928674(01)00633-x 27. Nagai S, Dubrana K, Tsai-Pflugfelder M, Davidson MB, Roberts TM, Brown GW, Varela E, Hediger F, Gasser SM, Krogan NJ (2008) Functional targeting of DNA damage to a nuclear pore-associated SUMO-dependent ubiquitin ligase. Science 322(5901):597–602. https://doi.org/10.1126/science.1162790 28. Hang J, Dasso M (2002) Association of the human SUMO-1 protease SENP2 with the nuclear pore. J Biol Chem 277(22): 19961–19966. https://doi.org/10.1074/jbc. M201799200 29. Zhao X, Wu CY, Blobel G (2004) Mlp-dependent anchorage and stabilization of a desumoylating enzyme is required to prevent clonal lethality. J Cell Biol 167(4):605–611. https://doi.org/10.1083/jcb.200405168 30. Goeres J, Chan PK, Mukhopadhyay D, Zhang H, Raught B, Matunis MJ (2011) The SUMO-specific isopeptidase SENP2 associates dynamically with nuclear pore complexes through interactions with karyopherins and

Ubiquitylation and SUMOylation of the Yeast NPC the Nup107-160 nucleoporin subcomplex. Mol Biol Cell 22(24):4868–4882. https:// doi.org/10.1091/mbc.E10-12-0953 31. Chow KH, Elgort S, Dasso M, Ullman KS (2012) Two distinct sites in Nup153 mediate interaction with the SUMO proteases SENP1 and SENP2. Nucleus 3(4):349–358. https:// doi.org/10.4161/nucl.20822 32. Takahashi Y, Mizoi J, Toh EA, Kikuchi Y (2000) Yeast Ulp1, an Smt3-specific protease, associates with nucleoporins. J Biochem 128(5):723–725. https://doi.org/10.1093/ oxfordjournals.jbchem.a022807 33. Nino CA, Hayakawa A, Dargemont C, Babour A (2012) Mapping ubiquitin modifications reveals new functions for the yeast nuclear pore complex. Cell Logist 2(1):43–45. https://doi.org/10.4161/cl.19720 34. Nino CA, Guet D, Gay A, Brutus S, Jourquin F, Mendiratta S, Salamero J, Geli V, Dargemont C (2016) Posttranslational marks control architectural and functional plasticity of the nuclear pore complex basket. J Cell Biol 212(2):167–180. https://doi.org/10.1083/ jcb.201506130 35. Hayakawa A, Babour A, Sengmanivong L, Dargemont C (2012) Ubiquitylation of the nuclear pore complex controls nuclear migration during mitosis in S. cerevisiae. J Cell Biol 196(1):19–27. https://doi.org/10.1083/jcb. 201108124 36. Raices M, D’Angelo MA (2017) Nuclear pore complexes and regulation of gene expression.

269

Curr Opin Cell Biol 46:26–32. https://doi. org/10.1016/j.ceb.2016.12.006 37. Simon MN, Kalousi A, Soutoglou E, Ge´li V, Dargemont C (2018) NPC in DNA repair and telomere maintenance. In: Nuclear pore complexes in genome organization, function and maintenance. Springer. https://doi.org/10. 1007/978-3-319-71614-5_9 38. Folz H, Nino CA, Taranum S, Caesar S, Latta L, Waharte F, Salamero J, Schlenstedt G, Dargemont C (2019) SUMOylation of the nuclear pore complex basket is involved in sensing cellular stresses. J Cell Sci 132(7):jcs224279. https://doi.org/10.1242/ jcs.224279 39. Bonnet A, Bretes H, Palancade B (2015) Nuclear pore components affect distinct stages of intron-containing gene expression. Nucleic Acids Res 43(8):4249–4261. https://doi.org/ 10.1093/nar/gkv280 40. Longtine MS, McKenzie A 3rd, Demarini DJ, Shah NG, Wach A, Brachat A, Philippsen P, Pringle JR (1998) Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14(10):953–961. https://doi.org/10. 1002/(SICI)1097-0061(199807)14: 103.0.CO;2-U 41. Psakhye I, Jentsch S (2012) Protein group modification and synergy in the SUMO pathway as exemplified in DNA repair. Cell 151(4): 807–820. https://doi.org/10.1016/j.cell. 2012.10.021

Chapter 18 Purification of Cdk-CyclinB-Kinase–Targeted Phosphopeptides from Nuclear Envelope Justin D. Blethrow, Amanda L. DiGuilio, and Joseph S. Glavy Abstract We describe a method for rapid identification of protein kinase substrates within the nuclear envelope. Open mitosis in higher eukaryotes is characterized by nuclear envelope breakdown (NEBD) concerted with disassembly of the nuclear lamina and dissociation of nuclear pore complexes (NPCs) into individual subcomplexes. Evidence indicates that reversible phosphorylation events largely drive this mitotic NEBD. These posttranslational modifications likely disrupt structurally significant interactions among nucleoporins (Nups), lamina and membrane proteins of the nuclear envelope (NE). It is therefore critical to determine when and where these substrates are phosphorylated. One likely regulator is the mitotic kinase: Cdk1Cyclin B. We employed an “analog-sensitive” Cdk1 to bio-orthogonally and uniquely label its substrates in the NE with a phosphate analog tag. Subsequently, peptides covalently modified with the phosphate analogs are rapidly purified by a tag-specific covalent capture and release methodology. In this manner, we were able to confirm the identity of known Cdk1 targets in the NE and discover additional candidates for regulation by mitotic phosphorylation. Key words Cdk1-Cyclin B kinase (cdk1), Nuclear Envelope (NE), Nuclear Pore Complex (NPC), nucleoporins (Nups), Nuclear Lamina, Nuclear Envelope Breakdown (NEBD), Mass Spectrometry (MS, MS/MS), Analog-sensitive kinase (as-kinase), ATP analog N6-(benzyl)ATP-γ-phosphorothioate

1

Introduction In eukaryotes, cellular membranes compartmentalize the cell into specialized organelles. These membrane bound compartments provide unique local environments required by biological molecules and allow biochemical reactions to proceed without interference from one another. The separation of DNA from surrounding cellular organelles and materials is achieved by the nuclear envelope (NE) [1, 2]. The passage of medium- and large-sized biomolecules through this barrier is mediated in both directions by nuclear pore complexes (NPCs). They provide a critical point for the regulation of communication between cytoplasmic and nucleoplasmic components [3]. The double lipid membrane bilayer of the NE, consists

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_18, © Springer Science+Business Media, LLC, part of Springer Nature 2022

271

272

Justin D. Blethrow et al.

of inner and outer nuclear membrane (INM and ONM, respectively) networks with circular openings that are occupied by NPCs. These NPCs span the NE and are the gateways for macromolecular traffic into and out of the nucleus [4]. These are highly regulated transport pathways that control nuclear entry and exit of molecules such as transcription factors, RNAs, kinases and viral particles [4, 5]. NPCs are the cell’s largest discrete protein assembly, at roughly 110 MDa in humans [6]. Both in yeast and mammals, NPCs are assembled from about 30 distinct proteins, collectively termed nucleoporins (Nups) and in a transport competent NPC, they occur in multiples of 8 [7, 8]. These Nups are organized into modular subcomplex units which confer an eightfold rotational symmetry about the axis of transport [9]. The structural core of the NPC also displays twofold symmetry about the plane of the NE. In human cells this structural core is composed predominately of the Nup107 and Nup93 complexes [10–12]. Throughout interphase, NPCs are anchored to transmembrane proteins of the NE such as POM121, gp210, and NDC1 which are positioned at the curvature of the NE where INM and ONM meet [13]. The ONM is continuous with the ER, to which it recedes during NEBD. The INM contains a unique array of integral membrane proteins including lamina-associated polypeptides 1 and 2 (LAP1 and LAP2) and Lamin B receptor (LBR) [13]. The overall shape of the NE is thought to be maintained by the meshwork of intermediate filaments and membrane proteins known as the nuclear lamina [14, 15]. The NE is a complex, yet dynamic, barrier and must undergo enormous physical changes in order to facilitate cell division [16]. NEBD occurs at the very onset of mitosis, the G2/M phase transition [17]. Temporally regulated kinase activity results in cellcycle dependent phosphorylation events [18]. Phosphorylation of Nup98 and structural scaffold is essential to NPC disassembly [19, 20]. It has been shown by Onischenko et al., that Cdk1 activity is required to keep NPCs dissociated during mitosis, while reassembly of NPCs in daughter cells is phosphatase dependent [21]. Therefore, it is crucial to identify Cdk1 substrates in the NE and to map their sites of phosphorylation in order to understand their roles in NEBD and regulation of the assembly and disassembly of NPCs during cell division. Through the use of an “analog sensitive kinase” (or as-kinase) approach, Cdk1 substrates can be specifically labeled at their sites of phosphorylation with a chemical tag that facilitates their subsequent purification [22–24]. The kinase is engineered to accept a bulky ATP analog that is a very poor substrate for wildtype kinases. The ATP analog also includes a thiophosphate moiety in the gamma position, which is transferred to substrates. A covalent “capture-and release” approach is then used to purify the

Capturing Phosphopeptides from the Nuclear Envelope

273

peptides of substrates that harbor the (thio)phosphate groups. Mass-spectrometric analysis of these fragments reveals both the identity of the parent proteins, and the location of the phosphorylation site(s).

2 2.1

Materials NE Isolation

1. Complete Protease Inhibitors (Roche, 1873580001 or similar). 2. Nucleases: RNase and DNase I. 3. Low molecular weight heparin (LMWH) sodium salt (Aprox. 3000 da, Millipore Sigma 1304118 formerly H-3400). 4. Phenylmethylsulfonyl fluoride (PMSF). 5. 1,4-Dithiothreitol (DTT). Solutions

1. Lysis Buffer: 0.1 mM MgCl2, 1 mM DTT, 5 μg/mL DNase l, 5 μg/mL RNase plus complete protease inhibitor and 0.5 mM PMSF. 2. Extraction Buffer I: 10% sucrose (w/v), 20 mM triethanolamine (pH 8.5) 0.1 mM MgCl2, 1 mM DTT plus complete protease inhibitor and 0.5 mM PMSF. 3. Extraction Buffer II: 10% sucrose (w/v), 20 mM triethanolamine (pH 7.5) 0.1 mM MgCl2, 1 mM DTT plus complete protease inhibitor and 0.5 mM PMSF. 4. Sucrose cushion: 30% sucrose (w/v), 20 mM triethanolamine (pH 7.5) 0.1 mM MgCl2, 1 mM DTT plus complete protease inhibitor and 0.5 mM PMSF. 2.2 Kinase Substrate Labeling

1. 10 mM N6-(benzyl)ATP-γ-phosphorothioate [6-Bn-ATP-γ-S] (BIOLOG Life Science Institute, Bremen, Germany) in 20 mM HEPES, pH 7.4. Alternative: 10 mM ATP-γ-S in 20 mM HEPES, pH 7.4. 2. as-Cdk1-Cyclin B kinase in 20 mM HEPES pH 7.4, 150 mM NaCl, at 0.15 mg/mL (see ref. 21 for production of as-Cdk1Cyclin B). Alternative: wild-type Cdk1-Cyclin B (Millipore 14-450). Stock Solutions and Buffers

1. Kinase buffer: 20 mM HEPES, pH 7.4. 2. 50 mM MgCl2 in deionized water.

274

Justin D. Blethrow et al.

3. 5% Triton X-100 in deionized water. 4. 40 mM EDTA, pH 8. 2.3 Tryptic Digest of NE Proteins

1. Sonicating water bath. 2. Trypsin Solution: 0.5 μg/μL - mass spectrometry grade modified trypsin. Stock Solutions and Buffers

1. 1 M Tris, pH 8.5. 2. 1.5% Empigen BB in deionized water. 3. Digestion Buffer (5
): 500 mM Tris pH 8.5, 1% SDS, 1.5% Empigen. 2.4 Thiophosphopeptide Purification

1. Oxone. 2. Sulfolink iodoacetyl-agarose beads (Pierce Chemical 20401). 3. 1-mL disposable micro-spin columns. Stock solutions and buffers

1. Oxidation solution: Dissolve 10 mg of Oxone in 10 mL of water for 1 mg/mL. 2. 1% formic acid in deionized water. 3. 50% acetonitrile in deionized water. 4. 5 M NaCl in deionized water.

3

Methods Purified NE is prepared from rat liver nuclei [7, 25–28], essentially as previously described with minor modifications. The livers are harvested from male Sprague-Dawely rats and nuclei are isolated following the described methods [25]. The ATP analog is designed to enable specific labeling of as-Cdk1 substrates in the presence of competing kinases [24]. This is accomplished by virtue of the benzyl group appended to the N6 amine, which confers specificity for the F80G mutant version of Cdk1. A kinase-transferable tag is provided by a thiophosphate group in the gamma position. Synthesis of this reagent is described in Allen et al., [20]. Design, expression and purification of as-Cdk1 is presented in Polson et al., [21]. If the target material containing candidate substrates is sufficiently simple, such that it may contain very few or no other kinases, the benefit conferred by this system may be insufficient to justify the effort required to obtain or produce these reagents. In that case, it may be appropriate to use wild-type kinase with commercially available ATP-γ-S. Doing so will preserve the ability of the kinase to tag substrates with

Capturing Phosphopeptides from the Nuclear Envelope

275

thiophosphate, but will abrogate the specificity of the kinasenucleotide interaction. In this case, it is particularly appropriate to conduct a control labeling experiment lacking added kinase to appraise background labeling by endogenous kinases. 3.1 Isolation of NE and Heparin Treatment

Starting material for the subsequent protocol can be obtained by the detailed procedure described in the classic work of Blobel and Potter [25]. Here, we refer specifically to the isolation and heparin treatment of NE from rat liver. However, we note that it can easily be adapted for NE from a variety of cell lines including the widely used HeLa [19] and Hek293 cell lines (see Note 1) [29]. 1. Rapidly thaw frozen rat liver nuclei (100 units)(see Note 2) in 30  C degree bath for a few seconds. Gently resuspend frozen aliquots back into mononuclei mixture by gentle vortexing and/or pipetting then transfer to a 15 mL glass round-bottom tube and centrifuge for 1 min at 1000
g using a swinging bucket rotor at 4  C. Carefully discard the supernatant leaving the pellet. 2. Dropwise add 1 mL of lysis buffer containing the DNase l and RNase (room temperature) while continuously vortexing at the midrange setting just enough for the liquid to reach the threequarter height of the tube. Add 4 mL of extraction buffer I (room temperature) in the same fashion. Adjust vortexing speed if needed to avoid spilling the resuspenison. 3. Incubate this digestion for 15 min at room temperature then underlay with 4 mL of ice-cold sucrose cushion solution. 4. Pellet by centrifugation at 4000
g for 15 min at 4  C using a swinging bucket rotor. Aspirate off the supernatant. 5. Dropwise resuspend the pellet with 1 mL of ice-cold extraction buffer II while continuously vortexing then carefully add 0.5 mL of ice-cold extraction buffer II containing 0.3 mg/ mL of heparin. 6. Underlay with 4 mL of the sucrose cushion solution and centrifuge for 15 min at 4000
g at 4  C. 7. Resuspend the heparin-treated NE in ice-cold extraction buffer ll and centrifuge at 4000
g for 15 min at 4  C. Remove the supernatant and repeat this step twice to fully remove excess heparin (see Note 3). 8. Enrichment of the NE can be visualized by gel staining and immunoblot (Fig. 1). Negative stain electron microscopy shows the NE and embedded NPCs (Fig. 2) (see Note 4). 9. Store the material at 80  C in 100-U aliquots.

276

Justin D. Blethrow et al.

Fig. 1 Nuclear envelope preparation and heparin treatment in human and rat cell lines. (a) Four cell fractions: cell extract (CE), cytoplasm (CY), nucleoplasm (NP) and nuclear envelope (NE), were separated with 4–20% SDS/PAGE, transferred to nitrocellulose and probed with MAb414 (against FG-Nups) (b) NE from different sources were separated with 4–20% SDS-PAGE and Coomassie stained. HeLa NE before and after heparin treatment (Lanes 1 and 2). Rat liver NE before and after heparin treatment (Lanes 3 and 4). (c) Heparin-treated HeLa NE (0 mg/mL to 0.3 mg/mL) were separated with 4–20% SDS/PAGE, transferred to nitrocellulose and probed with MAb414 (Against FG-Nups). Heparin-treated HeLa NE. HeLa NE was treated with increasing levels of heparin (0 mg/mL to 0.3 mg/mL) (left to right) then separated into pellet (bound) and supernatant (unbound) under low speed centrifugation conditions. S ¼ Supernatant, P ¼ Pellet and T ¼ Total, an aliquot removed before centrifugation

3.2 Protocol for Labeling Phosphorylation Sites of Cdk1 Substrates at the NE

The described substrate-tagging procedure is scaled for 20 units of NE. This scale is readily adjusted as necessary. The logic of the purification chemistry is described in Blethrow et al., [22] (Fig. 3). Synthesis of the thiophosphate donor nucleotide analog N6(benzyl)ATP-γ-phosphorothioate is described in Allen et al., [30] and also available commercially (Subheading 2.2). 1. Add 9.5 μL of Kinase Buffer to 20 units of pelleted NE material, and gently resuspend the pellet by pipette. 2. Add 1.5 μL of 5% Triton X-100 with gentle mixing. 3. Add 150 ng (1 μL) of activated as-Cdk1-Cyclin B kinase with gentle mixing.

Capturing Phosphopeptides from the Nuclear Envelope

277

Fig. 2 Negative stain EM image of HeLa NE treated with low levels of heparin (0.3 mg/mL). Visible are intact segments of NE with embedded NPCs with distinct features like the cytoplasmic filaments and nuclear basket and both hetero- and euchromatin regions on the nuclear side of the NE. Scale bar, 100 nm

Fig. 3 Strategy for the purification of thiophosphopeptides by as-kinase capture of Cdk1-Cyclin B substrates in the NE. Cdk1-CyclinB complexes harboring the analog-sensitive mutant are added to segments of purified and heparin-treated NE are depicted with intact NPCs (green crescents) along with the modified thiophosphate. Following modification of NE substrates by the modified kinase, proteins are tryptically digested and incubated with iodoacetyl-agarose beads to specifically isolate thio-tagged substrates digested to peptides. Thiolcontaining groups covalently attach to beads and unbound peptides are washed away. Thiophosphopeptides are specifically liberated by oxidation-promoted hydrolysis of the sulfur-phosphorus bond, and are converted to phosphopeptides in the process. Inset: Chemical structure of N6-(benzyl)ATP-γ-phosphorothioate responsible for specifically tagging NE substrates of the analog-sensitive kinase Cdk1

278

Justin D. Blethrow et al.

4. Add 1.5 μL of 50 mM MgCl2 with gentle mixing. 5. Add 1.5 μL of 10 mM N6-(benzyl)ATP-γ-phosphorothioate with gentle mixing. 6. Allow the kinase reaction to sit for 30 min at room temperature. 7. Stop the kinase reaction by adding 5 μL of 40 mM EDTA, then place the reaction on ice or store at 80  C for later use. 3.3 Tryptic Digest of NE Proteins

1. Add 5 μL of 5
Digestion Buffer to the tube containing the kinase reaction and mix. Pipette carefully to avoid frothing. 2. Add 5 μL of Trypsin Solution and mix. Pipette carefully to avoid frothing. 3. Incubate the reaction tube in a sonicating water bath at 37  C for 1 h. 4. Centrifuge the reaction at 18,000
g (room temperature) for 10 min to pellet any insoluble material and carefully transfer the supernatant to a new tube. The supernatant contains the desired tryptic peptides.

3.4 Purification of Thio-Labeled Phosphopeptides

1. Add 25 μL of acetonitrile to the peptide solution. 2. Add 20 μL of iodoacetyl-agarose beads to a separate tube. The beads are provided as a 1:1 slurry in the commercial product when resuspended, so it is necessary to use 40 μL of slurry. It may be necessary to trim the pipette tip with a blade to facilitate pipetting the beads. 3. To wash the bead slurry, add 500 μL of deionized water to the beads. Mix by pelleting and centrifuging for 30 s at 3000
g. Remove the supernatant and repeat. 4. Add the peptide solution to the washed iodoacetyl-agarose beads and wrap the tube in aluminum foil to protect it from light. 5. Allow the peptides to react with the beads for 1 h with continuous rotational mixing. 6. Pellet the beads by centrifuging for 30 s at 3000
g. Remove the supernatant and transfer to a separate tube. This contains unbound peptides, buffer, and detergents. Store this fraction for later troubleshooting and analysis if desired. 7. Suspend the beads in 500 μL of 50% acetonitrile/water and transfer them to a 20-mL column body. Allow the liquid to drain to waste (see Note 5). 8. Slowly add 10 mL of 50% acetonitrile/water to the beads and allow the liquid to drain to waste. 9. Slowly add 10 mL of deionized water to the beads and allow the liquid to drain to waste.

Capturing Phosphopeptides from the Nuclear Envelope

279

10. Slowly add 10 mL of 5 M NaCl to the beads and allow the liquid to drain to waste. 11. Slowly add 10 mL of deionized water to the beads and allow the liquid to drain to waste. 12. Slowly add 10 mL of 1% formic acid to the beads and allow the liquid to drain to waste (see Note 6). 13. Suspend the beads in 1 mL or less of deionized water and transfer them to a 1 mL disposable micro-spin column. Centrifuge the column for 30 s at 3000
g. Discard eluate. 14. Add 100 μL of Oxidation Solution to the beads. Mix briefly and allow the reaction to stand for 30–60 s. Centrifuge the column for 30 s at 3000
g and retain the eluate. The eluate contains recovered substrate-derived phosphopeptides. 15. Add 100 μL of 50% acetonitrile in water to the beads. Mix, and centrifuge the column for 30 s at 3000
g. Combine the eluate with that obtained in step 14. 16. Add 1 μL of 1 M DTT to the combined eluates and mix. Allow the mixture to stand for a few minutes. 17. Freeze the eluate by incubation at 80  C or in liquid N2, then lyophilize and store, or proceed to analysis (see Note 7). 18. Perform mass spectrometric analysis of the recovered peptides. As the technical details of mass spectrometric analysis depend substantially on the analytical instruments to be used, these details are beyond the scope of this work (see Note 8).

4

Notes 1. A number of steps can be adjusted to adapt protocols for the isolation of nuclei and NE from different cell types, such as HeLa. They are discussed in detail in the literature [6, 19, 29]. In short, they include composition of hypotonic buffer, duration of cell swelling, and number of Dounce strokes for cell lysis. Molarity of sucrose cushions and centrifugation parameters may depend on cell/nuclear size. Finally the amounts of nucleases should be individually optimized. Success of each step can be monitored by observation under a simple light microscope. 2. One unit equals 1
108 nuclei. Gentle resuspension of frozen aliquots back into mononuclei mixtures is required. 3. Preparation of the NE was modified from the previous protocol with some additional steps to prevent any interference with later covalent capture kinase applications [19, 26, 28, 29].

280

Justin D. Blethrow et al.

4. In Fig. 1, we show immunoblots probed with MAb414 (against FG-Nups: 358, 214, 153 and 62) across the cellular fractions: cell extract, cytoplasm, nucleoplasm and NE (Fig. 1a). A load of one unit per lane in 4–20% SDS-PAGE is stained with Coomassie blue to visualize bands of NE proteins in untreated and heparin-treated samples (Fig. 1b). Optional: A series of NE preparations treated with increasing levels of heparin were evaluated. Up to 0.3 mg/mL heparin yields NE amenable to subsequent kinase labeling while retaining membrane associations with the majority of nucleoporin proteins (Fig. 1c). 5. It is important to wash the beads thoroughly to remove detergents, which will otherwise complicate subsequent mass spectrometric analysis. If detergents are observed in the recovered material (as indicated by polymer ladders, often exhibited at 44 Da intervals), increase the water and water/organic wash volumes. 6. In addition to phosphopeptides, the recovered material contains by-products such as potassium sulfate salts and oxidized DTT. These species must be removed prior to analysis, for example by reverse-phase cleanup using C18 microcolumns, or by online desalting using a trap column in LC-MS. 7. If the wash steps are performed adequately, all peptides in the sample will be phosphopeptides (though it is common to see one or two peptides from very high abundance proteins persisting as contaminants). As such, no further enrichment of phosphopeptides should be necessary prior to analysis. In our experience, nanoscale LC-MS/MS is the preferred method for analysis with incorporation of an online desalting step. 8. We recovered 20 phosphorylation sites on 11 proteins [22]. Confirming identifications in the larger screen, we identified known Cdk1 phosphorylation sites in the nuclear lamina constituents lamin A/C and B. We also identified sites on three transmembrane proteins that associate directly with the lamin network: Lap1, Lap2B, and MAN1 [22]. We also identified phosphorylation sites within constituents of the NPC (Nup53, Nup133, and Nup358) and in the nucleoplasmic domains of three transmembrane proteins (gp210, POM121, and NDC1) that assemble and/or anchor the NPC into the NE [22] (Fig. 4).

Capturing Phosphopeptides from the Nuclear Envelope

281

Fig. 4 A Cdk1-Cyclin B phosphorylation site found in Man1 (S145). A tandem mass spectrum is shown for an extract-recovered phosphopeptide spanning S145 in Man1. The peptide sequence is shown in the inset, and observed b-series and y-series ions are indicated above select peaks. b0 and y0 indicate loss of water, and b* and y* indicate loss of ammonia from the corresponding b and y ions. The fragmentation pattern indicates phosphorylation at S145: y2 is observed unmodified while the addition of 80 Da is seen starting at y3, and modified fragments readily lose phosphate (loss of 98 Da). The b series shows a corresponding shift, confirming the presence of a phosphate group at S145

Acknowledgments The corresponding author would first like to thank the late Gu¨nter Blobel for his mentorship and encouragement. Also, we would like to Andrew Krutchinsky and Kevan Shokat for their support. We thank Thomas Cattabiani for critical reading of the manuscript. This work was supported in part by the Howard Hughes Medical Institute. J.S.G. was supported by the National Institutes of Health Individual National Research Service Award 5F32GN20520 and NIGMS-7R15GM119118. References 1. Gall JG (1954) Observations on the nuclear membrane with the electron microscope. Exp Cell Res 7(1):197–200 2. Gall JG (1967) Octagonal nuclear pores. J Cell Biol 32(2):391–399 3. Stewart M (2007) Molecular mechanism of the nuclear protein import cycle. Nat Rev Mol Cell Biol 8(3):195–208

4. Lin DH, Hoelz A (2019) The structure of the nuclear pore complex (an update). Annu Rev Biochem 88:725–783 5. Mackmull MT, Klaus B, Heinze I, Chokkalingam M, Beyer A, Russell RB et al (2017) Landscape of nuclear transport receptor cargo specificity. Mol Syst Biol 13(12):962

282

Justin D. Blethrow et al.

6. 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. Mol Syst Biol 9:648 7. Cronshaw JM, Krutchinsky AN, Zhang W, Chait BT, Matunis MJ (2002) Proteomic analysis of the mammalian nuclear pore complex. J Cell Biol 158(5):915–927 8. Rout MP, Aitchison JD, Suprapto A, Hjertaas K, Zhao Y, Chait BT (2000) The yeast nuclear pore complex: composition, architecture, and transport mechanism. J Cell Biol 148(4):635–651 9. Beck M, Lucic V, Forster F, Baumeister W, Medalia O (2007) Snapshots of nuclear pore complexes in action captured by cryo-electron tomography. Nature 449(7162):611–615 10. Bui KH, von Appen A, Diguilio AL, Ori A, Sparks L, Mackmull MT et al (2013) Integrated structural analysis of the human nuclear pore complex scaffold. Cell 155(6): 1233–1243 11. Kosinski J, Mosalaganti S, von Appen A, Teimer R, DiGuilio AL, Wan W et al (2016) Molecular architecture of the inner ring scaffold of the human nuclear pore complex. Science 352(6283):363–365 12. von Appen A, Kosinski J, Sparks L, Ori A, DiGuilio AL, Vollmer B et al (2015) In situ structural analysis of the human nuclear pore complex. Nature 526(7571):140–143 13. Lusk CP, Blobel G, King MC (2007) Highway to the inner nuclear membrane: rules for the road. Nat Rev Mol Cell Biol 8(5):414–420 14. Moir RD, Spann TP, Lopez-Soler RI, Yoon M, Goldman AE, Khuon S et al (2000) Review: the dynamics of the nuclear lamins during the cell cycle-- relationship between structure and function. J Struct Biol 129(2–3):324–334 15. Moir RD, Yoon M, Khuon S, Goldman RD (2000) Nuclear lamins A and B1: different pathways of assembly during nuclear envelope formation in living cells. J Cell Biol 151(6): 1155–1168 16. Hampoelz B, Andres-Pons A, Kastritis P, Beck M (2019) Structure and assembly of the nuclear pore complex. Annu Rev Biophys 48: 515–536 17. 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

18. Olsen JV, Vermeulen M, Santamaria A, Kumar C, Miller ML, Jensen LJ et al (2010) Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis. Sci Signal 3(104):ra3 19. Diguilio AL, Glavy JS (2013) Depletion of nucleoporins from HeLa nuclear pore complexes to facilitate the production of ghost pores for in vitro reconstitution. Cytotechnology 65(4):469–479 20. Glavy JS, Krutchinsky AN, Cristea IM, Berke IC, Boehmer T, Blobel G et al (2007) Cellcycle-dependent phosphorylation of the nuclear pore Nup107-160 subcomplex. Proc Natl Acad Sci U S A 104(10):3811–3816 21. Onischenko EA, Gubanova NV, Kiseleva EV, Hallberg E (2005) Cdk1 and okadaic acidsensitive phosphatases control assembly of nuclear pore complexes in drosophila embryos. Mol Biol Cell 16(11):5152–5162 22. Blethrow JD, Glavy JS, Morgan DO, Shokat KM (2008) Covalent capture of kinase-specific phosphopeptides reveals Cdk1-cyclin B substrates. Proc Natl Acad Sci U S A 105(5): 1442–1447 23. Polson AG, Huang L, Lukac DM, Blethrow JD, Morgan DO, Burlingame AL et al (2001) Kaposi’s sarcoma-associated herpesvirus K-bZIP protein is phosphorylated by cyclindependent kinases. J Virol 75(7):3175–3184 24. Blethrow J, Zhang C, Shokat KM, Weiss EL (2004) Design and use of analog-sensitive protein kinases. Curr Protoc Mol Biol Chapter 18: Unit 18.1 25. Blobel G, Potter VR (1966) Nuclei from rat liver: isolation method that combines purity with high yield. Science 154(757):1662–1665 26. Bornens M, Courvalin JC (1978) Isolation of nuclear envelopes with polyanions. J Cell Biol 76(1):191–206 27. Dwyer N, Blobel G (1976) A modified procedure for the isolation of a pore complex-lamina fraction from rat liver nuclei. J Cell Biol 70(3): 581–591 28. Matunis MJ (2006) Isolation and fractionation of rat liver nuclear envelopes and nuclear pore complexes. Methods 39(4):277–283 29. Ori A, Andres-Pons A, Beck M (2014) The use of targeted proteomics to determine the stoichiometry of large macromolecular assemblies. Methods Cell Biol 122:117–146 30. Allen JJ, Lazerwith SE, Shokat KM (2005) Bio-orthogonal affinity purification of direct kinase substrates. J Am Chem Soc 127(15): 5288–5289

Part VI Biophysical Methods

Chapter 19 Crystallization of Nuclear Export Signals or Small-Molecule Inhibitors Bound to Nuclear Exporter CRM1 Ho Yee Joyce Fung and Yuh Min Chook Abstract The Karyopherin protein CRM1 or XPO1 is the major nuclear export receptor that regulates nuclear exit of thousands of macromolecules in the cell. CRM1 recognizes protein cargoes by binding to their 8–15 residue-long nuclear export signals (NESs). A ternary CRM1–Ran–RanBP1 complex engineered to be suitable for crystallization has enabled structure determination by X-ray crystallography of CRM1 bound to many NES peptides and small-molecule inhibitors. Here, we present a protocol for the purification of the individual proteins, formation of the ternary CRM1–Ran–RanBP1 complex and crystallization of this complex for X-ray crystallography. Key words CRM1, XPO1, Nuclear export signals, NES, Leptomycin B, LMB, SINE, KPT, X-ray crystallography

1

Introduction The Chromosome Region of maintenance 1 protein (CRM1), also known as Exportin-1 (XPO1), is the most general nuclear exporter in the cell, which is responsible for the localization of possibly thousands of macromolecules [1]. Crystallization of CRM1 in complex with protein cargoes and other nuclear export cofactors was previously challenging due to conformational flexibility of CRM1. However, the ternary CRM1–Ran–RanBP1 ternary complex, which is an intermediate state formed in nuclear export, is easily crystallized and produced high resolution structures [2]. Further manipulation and engineering of the complex has allowed for efficient crystallization of small-molecule inhibitors or nuclear export signal (NES) peptides bound to CRM1. CRM1 in this engineered CRM1–Ran–RanBP1 complex adopts a very stable conformation, which allows it to be easily crystallized in the same condition reliably. However, for the same reason, the complex is appropriate only for studying inhibitor or NES interactions with the NES binding groove of the CRM1 as the rest of the CRM1

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_19, © Springer Science+Business Media, LLC, part of Springer Nature 2022

285

286

Ho Yee Joyce Fung and Yuh Min Chook

protein is significantly constrained by interactions with RanGTP and RanBP1 to one stable conformation. We have so far crystallized a total of 14 CRM1–inhibitor and 25 CRM1–NES complexes, which have allowed us to understand the mechanisms of CRM1 inhibition [3–7] and the recognition of NES peptides with diverse sequences [8–10].

2

Materials All water used is Milli-Q purified deionized water (sensitivity of 18 MΩ-cm at room temperature).

2.1 Plasmids and Strains

1. CRM1 crystallization construct for inhibitor complexes: pGexTEV-ScCRM1 residues 1-1058, Δ377-413, 537DLTVK541 to GLCEQ (to resemble the NES groove of human CRM1) (see Note 1). 2. CRM1 crystallization construct for NES complexes: pGexTEV-ScCRM1 residues 1-1058, Δ377-413, 537DLTVK541 to GLCEQ (to resemble the NES groove of human CRM1) and V441D mutation (see Note 1). 3. Ran expression construct: pET15b-HsRan (full-length) with N-terminal HisX6 tag. 4. RanBP1 expression construct: pGex-TEV-ScRanBP1 (Yrb1 residues 61-201). 5. NES expression construct: pMal-TEV-NES with desired NES peptide sequence inserted after TEV cleavage site (see Note 2). 6. E. Coli. BL21 (DE3) cells: BL21 cells are used for all protein expressions.

2.2 Chemicals and Reagents

1. LB Media: Miller LB broth is used for growth. Autoclaved before use. 2. Ampicillin: 100 mg/mL stocks in water, filtered and stored in
20  C. 3. Isopropyl β-D-1-thiogalactopyranoside (IPTG): 1 M stocks in water, filtered and stored in
20  C. 4. TEV protease: We purify our own His-tagged TEV protease, stored in 1 mg/mL in
20  C with 30% (v/v) glycerol or
80  C with 10% (v/v) glycerol. 5. Guanosine-50 -triphosphate (GTP): 100 mM stock in 50 mM Tris 8.0, stored in
20  C. 6. 50 -Guanylyl imidodiphosphate (GMPPNP): 100 mM stock in 50 mM Tris pH 8.0, stored in
20  C.

Crystallization of CRM1-NES or CRM1-Inhibitor Complexes

287

7. Alkaline phosphatase agarose (Sigma-Aldrich): ammonium sulfate suspension, stored in 4  C. 8. Maltose. 2.3

Buffers

All reducing agents, protease inhibitors, and nucleotides are added fresh, before use. 1. Lysis A: 40 mM HEPES pH 7.5, 2 mM magnesium acetate, 200 mM sodium chloride, 10 mM DTT, 1 mM benzamidine, 10 μg/mL leupeptin, 50 μg/mL AEBSF. 2. Wash A: 40 mM HEPES pH 7.5, 5 mM magnesium acetate, 100 mM sodium chloride, 2 mM DTT. 3. Wash B: 40 mM HEPES pH 7.5, 5 mM magnesium acetate, 300 mM sodium chloride, 2 mM DTT. 4. Wash C: 40 mM HEPES pH 7.5, 5 mM magnesium acetate, 100 mM sodium chloride, 1 mM DTT. 5. GF: 20 mM HEPES pH 7.5, 5 mM magnesium acetate, 100 mM sodium chloride, 2 mM DTT. 6. Lysis B: 50 mM Tris pH 8.0, 5 mM magnesium acetate, 400 mM sodium chloride, 10% (v/v) glycerol, 20 mM imidazole pH 7.8, 2 mM 2-mercaptoethanol, 1 mM Benzamidine, 10 μg/mL Leupeptin, 50 μg/mL AEBSF, 0.2 mM GTP. 7. Ni A: 50 mM Tris pH 8.0, 5 mM magnesium acetate, 200 mM sodium chloride, 10% (v/v) glycerol, 2 mM 2-mercaptoethanol. 8. Ni B: 50 mM Tris pH 8.0, 5 mM magnesium acetate, 200 mM sodium chloride, 10% (v/v) glycerol, 500 mM imidazole pH 7.8, 2 mM 2-mercaptoethanol. 9. TB: 20 mM Tris pH 8.0, 2 mM magnesium acetate, 110 mM potassium acetate, 10% (v/v) glycerol, 2 mM DTT. 10. BT: 20 mM Bis-Tris pH 6.5, 5 mM magnesium acetate, 100 mM sodium chloride. 11. Lysis C: 50 mM HEPES pH 7.5, 200 mM sodium chloride, 10% (v/v) glycerol, 2 mM DTT, 1 mM benzamidine, 10 μg/ mL leupeptin, 50 μg/mL AEBSF. 12. Q A: 20 mM HEPES pH 7.5, 30 mM sodium chloride, 10% (v/v) glycerol, 2 mM DTT. 13. Q B: 20 mM HEPES pH 7.5, 1 M sodium chloride, 10% (v/v) glycerol, 2 mM DTT. 14. Crystallization condition: 16 to 18% (w/v) PEG3350, 100 mM Bis-Tris pH 6.4 or 6.6, 200 mM ammonium nitrate (see Note 3).

288

2.4

Ho Yee Joyce Fung and Yuh Min Chook

Equipment

1. High-pressure homogenizer. 2. Affinity chromatography resins: Glutathione Sepharose (GSH), HisTrap HP (Cytiva), Amylose. 3. Glass chromatography columns. 4. Fast protein liquid chromatography (FPLC) instrument. 5. Size-exclusion chromatography columns: Superdex 200 and Superdex 75 (Cytiva). 6. Ion exchange chromatography column: HiTrap Q (Cytiva). 7. Centrifugal concentrators with 2, 10, 30, and 50 kDa molecular weight cutoff (MWCO). 8. Centrifugal filters. 9. 24-well plate for hanging drop crystallization. 10. 22 mm siliconized square cover slides.

3

Methods All protein purification and centrifugation steps are performed on ice or in a cold room or cold box at 4  C, unless specifically stated.

3.1 Purification of CRM1

1. Grow 6 L of BL21 cells expressing the desired CRM1 construct (see Note 1) in LB with 100 μg/mL Ampicillin, at 37  C (shaking at 230 rpm) to OD 0.8 to 1.5. Lower the temperature of the shaker and induce protein expression with 0.5 mM IPTG for 10–12 h, at 25  C (shaking at 200 rpm). Chill to 4  C until collection (see Note 4). 2. Centrifuge to pellet the cells, resuspend the pellets in 20 ml/L growth of Lysis A buffer and freeze in
20  C until use. 3. Thaw frozen bacteria cell suspensions and lyse the cells by passing the thawed lysate through the high-pressure homogenizer/cell disruptor three times (see Note 5). 4. Centrifuge the lysate at 20,000 rpm or 48,400  g for 40 min in a fixed-angle centrifuge. 5. While the lysate is in the centrifuge, wash 5–10 mL bead volume of GSH beads, placed in a glass chromatography column, with water followed by Lysis A buffer. 6. Add supernatant of the centrifuged lysate to the washed GSH beads (Fig. 1a, lane 1) and collect the flow-through (Fig. 1a, lane 2). 7. Wash the GST-CRM1 GSH beads with 20–50 mL each of Wash A followed by Wash B (Fig. 1a, lane 3 and 4). 8. Wash/equilibrate the column with 20 mL Wash C buffer.

Crystallization of CRM1-NES or CRM1-Inhibitor Complexes

289

Fig. 1 Generation of the components of the CRM1–Ran–RanBP1 complex. (a) Coomassie-stained SDS-PAGE gel of a CRM1 purification. Lanes are samples from: supernatant after centrifugation (1), flow-through over GSH beads (2), washes A (3) and B (4), pooled flow-through containing CRM1 after TEV cut (5) and the remaining TEV-cleaved GST on glutathione Sepharose beads (6). (b) Typical chromatogram profile of a CRM1 purification in a HiLoad 26/600 Superdex S200 column (320 mL column volume). First peak is aggregation and second peak is CRM1. (c) Example Ran–RanBP1 purification on a Superdex 75 10/300 (24 mL column volume). Coomassie-stained SDS-PAGE gel (left) of a typical scouting run (chromatogram on the right). Samples are from fractions indicated by the bracket on the chromatogram. First peak is the Ran–RanBP1 heterodimer, and second peak contain Ran and RanBP1 monomers which comes off after the complexed heterodimer

9. Add ~1 mg of TEV to Wash C buffer to a total volume of 10 mL, add the TEV-containing buffer to the GST-CRM1 bound GSH beads and let sit for 2 h at room temperature. Use a spatula to gently stir the beads to resuspend the mixture every 15 min (see Note 6). 10. Elute and wash the beads with 10–15 mL of Wash C buffer twice. Pool the eluate and washes to obtain a total of 30–40 mL CRM1 at room temperature (Fig. 1a, lane 5 and 6) (see Note 7). 11. Concentrate the CRM1 pool using centrifugal concentrators with 50 kDa MWCO at max speed permitted for the concentrator, for a maximum of 15 min total (see Note 8). 12. Use a Superdex 200 column equilibrated in GF buffer to further purify the CRM1 (see Note 8).

290

Ho Yee Joyce Fung and Yuh Min Chook

13. Use SDS-PAGE to visualize proteins in the Superdex 200 chromatogram peaks. Concentrate the fractions that contain monomeric CRM1 to >5 mg/mL, flash-freeze the purified protein in aliquots in liquid nitrogen and store in
80  C (Fig. 1b). 3.2 Purification of the Ran–RanBP1 Heterodimer

1. Grow BL21 E. coli cells (transformed with Ran and RanBP1 expression vector respectively, 4 L each) in LB with 100 μg/mL Ampicillin at 37  C, shaking at 230 rpm, to OD 0.4 to 0.6. Induce expression of the proteins with 0.5 mM IPTG for 10 h at 25  C (200 rpm shaking). Chill to 4  C until collection (see Note 9). 2. Centrifuge to pellet the cells, resuspend the pellets in 20 ml/L growth of Lysis B buffer (for Ran) or Lysis A buffer (for RanBP1). Freeze and store the cell suspensions at
20  C until ready to begin protein purification. 3. To purify Ran, start with steps 3 and 4 from Subheading 3.1. 4. Equilibrate a 5 mL HisTrap HP column with Ni A buffer (see Note 10). 5. Load the supernatant of the clarified Ran lysate onto the HisTrap column using a peristaltic pump. 6. Wash off unbound proteins using 98% Ni A and 2% Ni B buffer on a FPLC. 7. Elute bound proteins from the HisTrap using a gradient of 2 to 100% Ni B buffer. 8. Use SDS-PAGE to visualize proteins in the peaks of the chromatogram. Concentrate fractions that contain Ran using centrifugal concentrators with 10 kDa MWCO to exportforkinT.ipf. Follow the instructions to prepare and export curves for fitting in MatLab. The following files will be created: – dateFCx.txt (sensogram), – datekinconsFCx.txt (contains an array of the analyte concentrations), – datetaFCx.txt (contains an array of the dissociation times for each injection), – datetdFCx.txt (contains an array of the association times for each injection), – datestFCx.txt (contains information regarding the starting point of the first injection). 7. Place all exported files into the same folder with the MatLab analysis codes/programs (see https://git.scicore.unibas.ch/ lim-group/spr-analysis-code-public). 8. Open in MatLab “kinparc.m” code (see Subheading 2.6) and insert names of the exported files above, which should be executed as indicated and run script. Details of the kinetic analysis and used model for the multivalent interaction of NTRs with the FG layer is provided in Note 9 and was published previously [6]. 9. At the end of the processing the following result files would be created. – dateFCx_kon_matrix.txt, – dateFCx_koff_matrix.txt, – dateFCx_t.txt, – dateFCx_fit.txt, – dateFCx_data.txt, – dateFCx_res.txt. 10. Use “resplotSchoch.m” to plot data together with the fit and residual (see https://git.scicore.unibas.ch/lim-group/spr-anal ysis-code-public). 11. Use “matrixsum.m” to plot the kinetic maps (see link for codes above and Note 10). The files “matrix1” and “matrix2” created by executing “matrixsum.m” could be also used to replot kinetic maps in Python using script “MakePlotKinAnalysisUNO.py” or the overlap of two kinetic maps using “MakePlotKinAnalysisDUE.py”. The starting parameters for the histograms fits in these plots could be selected by previewing the maps using “MakePlotKinAnalysisUNO_maponly.py” or “MakePlotKinAnalysisDUE_maponly.py” (see https://git. scicore.unibas.ch/lim-group/spr-analysis-code-public).

Study of the Multivalent Interactions at Biointerface by SPR 3.5.5 Evaluation of FG Nup Layer Height

325

1. To measure an initial FG layer height (d2(initial)), execute three consecutive BSA injections with a duration of 30 s each after the FG Nup immobilization on the sensor surface with an interval of 60 s (Figs. 3 and 4). 2. Each triple injection is repeated after an analyte (NTR) injection to detect conformational changes (Δdj¼ d2,j - d2(initial)) following NTR binding (see Note 11). 3. The layer height is calculated, using the below eq. (2):   R1,j ∙ m2 ld d 2,j ¼ ln þ d1, 2 R2,j ∙ m1

ð3Þ

where d1 is the thickness of the reference layer in flow cell 1 (see Note 12), d2,j is the FG Nup layer thickness in flow cell 2 at the j-th NTR injection, R1,j and R2,j are the SPR responses to the BSA injection in flow cells 1 and 2 at each j-step, ld is decay length of the evanescent field (it was estimated to be 320 nm for our setup, see Schoch et al., 2013). See Fig. 4 for measurements of R1 and R2. m1/m2 is a calibration factor that is approximately 1, given the SPR sensitivity is the same in all flow cells (see Note 13).

4

Notes 1. Use high quality PUT3 with at least 95% purity. Prepare 100 mM stock solutions of this compound in pure ethanol (pro analysis) and then dilute it with ethanol to 10 mM. This stock solution can be used to prepare 1 mM PUT3 in the running buffer without risking that it precipitates. Always spin down the final solution before being used for passivating the reference channel. 2. FG Nup solubility is lower in the cleavage buffer compared with the buffer containing 8 M urea. Thus, FG Nup containing fractions should be diluted upon collection and prior to dialysis into the cleavage buffer. 3. Prepare PUT3 solution immediately before the experiment. 4. Secure the gold chip with a Teflon holder (or equivalent) during the sonication procedure to avoid physical contact with other surfaces to minimize potential contamination or scratching. 5. Immobilize PUT3 in the reference channel before introducing the FG Nups in the sample channels so as to avoid any crosscontamination of FG Nups in the reference channel. 6. Both SLMs in either the reference and sample channels are expected to exhibit a similar thickness. Hence, their height difference should be zero.

326

Larisa E. Kapinos and Roderick Y. H. Lim

7. Measure ΔRU as a difference between the sensogram baseline immediately preceding FG Nup injection and the new baseline after the injection, ensuring that signal drift is no longer detected (see Fig. 4). 8. Minimize errors in calculating the mean grafting distance by washing the flow channel until the baseline drift has ceased. This ensures that noncovalently bound molecules are removed from the sensor surface. Likewise, a short injection of 0.1–0.2 mM NaOH may be used to check if noncovalently bound molecules are completely removed from the sensor surface. When a stable baseline is observed, the error in the calculation of mean grafting distance (g) depends only on the signal-to-noise ratio of the instrument. 9. The regularization method by Svitel et al. [21] is used to extract the association and dissociation kinetic constants associated with the multivalent binding of NTRs to the FG-Nups. Please see Kapinos et al. for details [6]. 10. The issue of MTL can be accounted for by knowing the surface density of the FG Nups, dimensions of the flow channels and the analyte diffusion constant. The surface ligand density (i.e., FG Nups) is calculated as follows. 5

2 ∙ 10 σ ¼ ΔRU 1000MW [mol/dm ].

The mass transfer coefficient is calculated as:  2=3  1=3 f D km ¼ 0:98   , h 0:3  w  l where h, w and l are height, width, length [m] (the following values are available in the BiacoreT200 manual for this instrument: v ¼ h  w  l ¼ 0.6 μL ¼ 6  1010 m3; h ¼ 0.04 mm ¼ 4 h10i5 m); 3 11 f is the flow rate ms (e.g., 10 μL/min orh 1.710 m3/s); i m2 D is the analyte diffusion coefficient s : D ¼ 342:3  1  1011 , MW 1=3 f η rel

where ηrel ¼ 0.89 at 25  C (for 100 kDa proteins could be taken D~6  1011 [m2/s] at 25  C) (see also BiacoreT200 manual). In the current context, km ¼ 2  105dm/s for a 100 kDa protein (Kapβ1) at 10 μL/min flow rate. Accordingly, the mass transport limitation is not an issue if km[Ab]  kon[B0][Ab] or (kon[B0])/km  1, where [B0] is an initial surface density (σ) of the bound ligand (FG Nups) and [Ab] is the analyte (NTR) concentration.

Study of the Multivalent Interactions at Biointerface by SPR

327

11. Be careful to avoid incurring errors in the FG Nup layer height. This includes contamination and damage to the sensor surface, or unspecific absorption due to poor passivation of the reference channel. Protein solutions should also be prepared very carefully to minimize impurities or aggregates. 12. The thickness of PUT3 is approximately 2 nm [23]. 13. These calculations are a part of the macro “SPR Analysis” that was written for IgorPro (see Subheading 2.6).

Acknowledgments We acknowledge R.L. Schoch for formulating the method to measure layer heights in SPR. We thank R.L. Schoch and R.S. Wagner for writing the SPR Analysis codes for MatLab, IgorPro, and Python. References 1. Schasfoort R, Tudos A (2008) Preface. In: Handbook of surface plasmon resonance. pp X - X i . h t t p s : // d o i . o r g / 1 0 . 1 0 3 9 / 9781847558220-Fp010 2. Schoch RL, Kapinos LE, Lim RYH (2012) Nuclear transport receptor binding avidity triggers a self-healing collapse transition in FG-nucleoporin molecular brushes. Proc Natl Acad Sci U S A 109:16911–16916 3. Schoch RL, Lim RYH (2013) Non-interacting molecules as innate structural probes in surface plasmon resonance. Langmuir 29:4068–4076 4. Hayama R et al (2019) Interactions of nuclear transport factors and surface-conjugated FG nucleoporins: insights and limitations. PLoS One 14:e0217897 5. Kapinos LE, Huang B, Rencurel C, Lim RYH (2017) Karyopherins regulate nuclear pore complex barrier and transport function. J Cell Biol 216:3609–3624 6. Kapinos LE, Schoch RL, Wagner RS, Schleicher KD, Lim RYH (2014) Karyopherincentric control of nuclear pores based on molecular occupancy and kinetic analysis of multivalent binding with FG nucleoporins. Biophys J 106:1751–1762 7. Wagner RS, Kapinos LE, Marshall NJ, Stewart M, Lim RYH (2015) Promiscuous binding of Karyopherinbeta1 modulates FG nucleoporin barrier function and expedites NTF2 transport kinetics. Biophys J 108:918– 927

8. Grossman E, Medalia O, Zwerger M (2012) Functional architecture of the nuclear pore complex. Annu Rev Biophys 41:557–584 9. Sakiyama Y, Mazur A, Kapinos LE, Lim RYH (2016) Spatiotemporal dynamics of the nuclear pore complex transport barrier resolved by high-speed atomic force microscopy. Nat Nanotechnol 11:719–723 10. Rout MP, Aitchison JD, Magnasco MO, Chait BT (2003) Virtual gating and nuclear transport: the hole picture. Trends Cell Biol 13: 622–628 11. Lim RYH et al (2007) Nanomechanical basis of selective gating by the nuclear pore complex. Science 318:640–643 12. Rexach M, Blobel G (1995) Protein import into nuclei: association and dissociation reactions involving transport substrate, transport factors, and nucleoporins. Cell 83:683–692 13. Paci G, Zheng T, Caria J, Zilman A, Lemke EA (2020) Molecular determinants of large cargo transport into the nucleus. elife 9:e55963 14. Aramburu IV, Lemke EA (2017) Floppy but not sloppy: interaction mechanism of FG-nucleoporins and nuclear transport receptors. Semin Cell Dev Biol 68:34–41 15. Isgro TA, Schulten K (2005) Binding dynamics of isolated nucleoporin repeat regions to importin-beta. Structure 13:1869–1879 16. Fasting C et al (2012) Multivalency as a chemical organization and action principle. Angew Chem Int Ed Engl 51:10472–10498

328

Larisa E. Kapinos and Roderick Y. H. Lim

17. Cooper MA, Try AC, Carroll J, Ellar DJ, Williams DH (1998) Surface plasmon resonance analysis at a supported lipid monolayer. Biochim Biophys Acta 1373:101–111 18. Eisele NB, Andersson FI, Frey S, Richter RP (2012) Viscoelasticity of thin biomolecular films: a case study on nucleoporin phenylalanine-glycine repeats grafted to a histidine-tag capturing QCM-D sensor. Biomacromolecules 13:2322–2332 19. Emilsson G et al (2018) Polymer brushes in solid-state nanopores form an impenetrable entropic barrier for proteins. Nanoscale 10: 4663–4669 20. Emilsson G et al (2015) Strongly stretched protein resistant poly(ethylene glycol) brushes prepared by grafting-to. ACS Appl Mater Interfaces 7:7505–7515

21. Svitel J, Balbo A, Mariuzza RA, Gonzales NR, Schuck P (2003) Combined affinity and rate constant distributions of ligand populations from experimental surface binding kinetics and equilibria. Biophys J 84:4062–4077 22. Goldstein B, Coombs D, He X, Pineda AR, Wofsy C (1999) The influence of transport on the kinetics of binding to surface receptors: application to cells and BIAcore. J Mol Recognit 12:293–299 23. Palegrosdemange C, Simon ES, Prime KL, Whitesides GM (1991) Formation of selfassembled monolayers by chemisorption of derivatives of oligo(ethylene glycol) of structure Hs(Ch2)11(Och2ch2)meta-oh on gold. J Am Chem Soc 113:12–20

Chapter 22 Assembly and Use of a Microfluidic Device to Study Nuclear Mechanobiology During Confined Migration Richa Agrawal, Aaron Windsor, and Jan Lammerding Abstract Cancer metastasis, that is, the spreading of tumor cells from the primary tumor to distant sites, requires cancer cells to travel through pores substantially smaller than their cross section. This “confined migration” requires substantial deformation by the relatively large and rigid nucleus, which can impact nuclear compartmentalization, trigger cellular mechanotransduction pathways, and increase genomic instability. To improve our understanding of how cells perform and respond to confined migration, we developed polydimethylsiloxane (PDMS) microfluidic devices in which cells migrate through a precisely controlled “field of pillars” that closely mimic the intermittent confinement of tumor microenvironments and interstitial spaces. The devices can be designed with various densities of pillars, ranging from a very low density that does not require nuclear deformation to high densities that present microenvironment conditions with severe confinement. The devices enable assessment of cellular fitness for confined migration based on the distance traveled through the constriction area over several days. In this protocol, we present two complementary techniques to generate silicon master molds for the device fabrication: (1) SU-8 soft lithography for rapid prototyping and for devices with relatively large features; and (2) reactive ion etching (RIE) to achieve finer features and more durable molds. In addition, we describe the production, use, and validation of the devices, along with the analysis pipeline for experiments using the devices with fluorescently labeled cells. Collectively, this protocol enables the study of confined migration and is readily amendable to investigate other aspects of confined migration mechanobiology, such as nuclear pore complex function in response to nuclear deformation. Key words Confined migration, Microfluidics, Cancer, Mechanobiology, Mechanotransduction, Cell nucleus, Metastasis, Invasion, Confinement

1

Introduction During in vivo migration, cells such as immune cells, fibroblasts, or metastatic tumor cells traverse interstitial spaces as small as 1–2 μm in diameter. This “confined migration” requires the deformation not only of the soft cell body but also of the large (5–10 μm diameter) and relatively rigid nucleus. The substantial deformation of the nucleus and the associated mechanical stress can lead to nuclear envelope rupture, DNA damage, and even fragmentation

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_22, © Springer Science+Business Media, LLC, part of Springer Nature 2022

329

330

Richa Agrawal et al.

of the nucleus [1–3]. At the same time, the mechanical deformation of the nucleus may also induce various biological responses that could alter the function of the migrating cell. Recent studies have identified the nucleus as a cellular mechanosensor that contributes to the cellular response to force, including changes in chromatin organization, nuclear stiffness, gene expression, and cell contractility [4–9]. Several non–mutually exclusive mechanisms have been proposed to explain how the nucleus can translate physical forces into biochemical signals (“nuclear mechanotransduction”), including stretch-induced opening of nuclear pores and stretch-sensitive ion channels in the nuclear membrane [4, 5, 8, 9]. Nonetheless, much remains to be understood about nuclear mechanotransduction mechanisms, particularly in the context of confined migration. Here, we present the fabrication and use of a novel microfluidic device to study nuclear mechanobiology during confined migration. Many current systems to study confined migration have significant limitations: porous membranes such as transwell plates and Boyden chambers are not well suited for live-cell imaging and thus allow for characterization only at fixed endpoints. Threedimensional (3D) matrices made of collagen, other biomaterials, or decellularized tissues are more physiological and allow some flexibility in pore size [10], but the pore size is very heterogeneous, and cells often migrate out of the focal plane. Furthermore, control of the cellular microenvironment is often quite limited in these systems [11]. In contrast, microfluidic devices functionalized with extracellular matrix proteins allows the study of cells in precisely controlled 3D environments. To date, however, most microfluidic devices used to study confined migration consist of straight channels with various widths and heights, providing continuous confinement [12, 13], or branching “tree-like” channels of increasing confinement to probe migratory fitness [14]. Here, we describe the design and fabrication of devices that instead mimic the intermittent confinement of interstitial environments using a precisely controlled but heterogeneous “field of pillars” with variable spacing (Fig. 1a–c). Experiments typically include devices with increasing pillar density and a control device with very low pillar density, which does not require substantial nuclear deformation for migration. The devices are well suited for continuous time-lapse microscopy (Fig. 1d), but also allow assessing migratory fitness based on the distance traveled by the cells from the seeding port. This method of endpoint measurements provides a significant advantage over previous versions which required long time-lapse imaging and transit time calculation [15]. The migration devices are fabricated by using a silicon mold to produce polydimethylsiloxane (PDMS) replicates, which are then bonded to coverslips to create confined 3D microenvironments, as described previously [16, 17]. The device also contains larger

Microfluidic Device to Study Nuclear Mechanobiology

331

Fig. 1 Overview of microfluidic device design. (a) Top-down and side view of microfluidic device layout. Cells are injected through seeding ports (biopsy punched areas, white), migrate into constrictions (red), and then migrate into the outer collection ring (blue). “Low” areas contact glass and assist with adhesion (green). (b) Photograph of a migration device with a 1-cent coin shown for scale. The seeding areas have been filled with water containing food coloring (red, blue, and green). The collection port has been filled with water containing yellow food coloring. (c) Schematic of different representative pillar densities with representative swatches of 15 μm pillar densities, featuring a control, low, and high density. Scale bar: 30 μm. (d) Representative timelapse image series of MDA-MB-231 cells expressing mCherry-actin and H2B-mNeonGreen migrating through medium density pillared devices. Scale bar: 20 μm

features to ensure adequate fluid flow and transport to the cells in the confined environments (Fig. 1a). In this protocol, we present two complementary techniques to fabricate the silicon mold for the PDMS replicates: (1) soft lithography using SU-8 photoresist and (2) reactive ion etching (RIE) (Fig. 2a). In soft SU-8 lithography, a layer of SU-8 photoresist remains on the wafer to create the desired topography for the PDMS replicates. By contrast, RIE uses a sacrificial photoresist as a photomask, which is subsequently removed to reveal features etched directly into the silicon wafer. To date, SU-8 soft lithography predominates the field of microfluidics for cell biology [16] and is ideally suited for coarse (>2 μm) and “tall” (>10 μm) features. Its advantages are the ease of use, low cost, and rapid prototyping, but it is often limited in its ability to produce small features with high-aspect ratios. RIE overcomes these limitations, and the resulting devices etched directly into silicon are highly reproducible and durable. We performed most rapid prototyping using SU-8 soft lithography but produce our final migration devices using two-layer fabrication, in which small constrictions are fabricated using RIE, and large features such as

332

Richa Agrawal et al.

Fig. 2 Microfluidic device fabrication. (a) Overview of silicon wafer fabrication workflow. Silicon wafer fabrication process in which the constriction layer was created using (top) RIE or (bottom) SU-8 soft

Microfluidic Device to Study Nuclear Mechanobiology

333

cell seeding ports and liquid handling channels are formed using a thick (>200 μm) SU-8 photoresist layer. In this protocol, we detail our fabrication methods, device design, use and analysis of the microfluidic migration devices. We validated the devices by comparing the confined migration behavior of mouse embryonic fibroblasts (MEFs) lacking the nuclear envelope proteins lamin A/C (Lmna
/
) and wild-type controls (Lmna+/+). We have previously demonstrated that Lmna
/
MEFs have more deformable nuclei and migrate faster through 2  5 μm2 pores than Lmna+/+ controls [18]. In the migration devices here, the Lmna
/
MEFs travel further through the confined conditions than the Lmna+/+ cells, validating distance traveled also as a reliable measurement for confined migration fitness (Fig. 3). The protocol presented here is particularly useful for researchers who want to rapidly design and prototype microfluidic devices, require devices with closely spaced, high depth-to-width ratios, want to create a highly durable master mold, or wish to study the determinants of fitness for and consequences of confined migration.

2

Materials

2.1 General Equipment and Reagents

1. Oven capable of temperatures up to 90  C. 2. Cleanroom swabs. 3. Hot plate capable of up to 200  C, with temperature ramping capabilities. 4. Deionized (DI) water. 5. Semiconductor grade isopropyl alcohol (IPA). 6. Semiconductor grade acetone. 7. Brightfield microscope with 10 to 100 objectives.

2.2 Preparation for Wafer Fabrication

1. AutoCAD software such as L-edit (Tanner EDA) or equivalent. 2. CZ silicon wafer, 4-in. diameter, type N, 525 μm thick, orientation.

ä Fig. 2 (continued) lithography. (b) Schematic of photomask design and layer overlay. The first layer forms the constriction area, and the second layer forms seeding and collection ports. (c) Casting, mounting, functionalization, and seeding of device. (i) The fabricated wafer is (ii) used to cast a PDMS replicate (ii), from which, recommended but optionally, (iii) a plastic mold is created, and (iv) a second PDMS replicate is cast. (v) Similar to as in (i), a second PDMS replicate is cast and then cut into eight “chips”, each containing a single migration device (vi). Each PDMS chip is biopsy punched three times (once with a 2 mm punch and twice with a 1 mm punch) in each of the three sections, (vii) then washed, (viii) plasma cleaned, and (ix) bonded to a glass-bottomed petri dish. Finally, each microfluidic device is functionalized with an extracellular matrix coating and is ready for cell seeding (x)

334

Richa Agrawal et al.

Fig. 3 Monitoring and quantifying cell migration. (a) Cells moving from seeding area into constriction area. Distance traveled through pillars assessed as distance from each nucleus to beginning of seeding port for Lmna
/
(top) and wild-type (Lmna+/+, bottom) MEFs through control condition. Scale bar: 50 μm (b) Distance traveled by Lmna
/
and wild-type cells through low density pillars and control devices. (c) Average distances traveled into the low-density pillar region for Lmna
/
and wild-type MEFs, normalized to distance traveled by each cell line in control conditions and averaged across all days (n ¼ 50–75 cells per condition from three independent experiments; *, p ¼ 0.039 using a two-sample t-test)

3. Acid piranha (3:1 mixture of sulfuric acid and 30% hydrogen peroxide). 4. Chrome photomask on a quartz substrate, 500  500  0.09000 . 5. Heidelberg DWL 2000 mask writer (Heidelberg Instruments, Heidelberg, Germany. 2.3 Fabricating Constriction Using SU-8 Lithography

1. Spin coater capable of speeds up to 4000 rpm. 2. SU-8 2005 Photoresist. 3. Mask aligner system (e.g., Suss MA6, Garching, Germany). 4. SU-8 developer. 5. Long-pass filter for near-pass UV light (e.g., PL-360LP from Omega Optical or equivalent).

2.4 Fabricating Constriction Layer Using RIE

1. Spin coater. 2. Mask aligner system (e.g., Suss MA6, Garching, Germany).

Microfluidic Device to Study Nuclear Mechanobiology

335

3. Escoat® P-20 solvent. 4. AZ® nLOF 2020 resist. 5. Oxford Cobra Abingdon, GB).

ICP

etcher

(Oxford

instruments,

6. Profilometer (e.g., KLA-Tencor, Milpitas, CA). 7. Anatech plasma asher (Anatech USA, Sparks, NV). 2.5 Upper Layer SU-8 and Final Processing

1. SU-8 100 Photoresist. 2. Mask aligner system (e.g., Suss MA6, Garching, Germany). 3. Long-pass filter for near-pass UV light (PL-360LP from Omega Optical or equivalent). 4. >95% (1H,1H,2H,2H-perfluorooctyl)trichlorosilane (FOTS) 5. Molecular vapor deposition system 100 (MVD100, Applied MicroStructures, San Jose, CA). 6. Video contact angle system such as VCA Optima.

2.6 Casting, Mounting, and Seeding of Devices

1. Sylgard 184 silicone elastomer base and curing agent (Dow Corning, Midland, MI). 2. Stirring rod for mixing elastomer base and curing components. 3. Vacuum pump. 4. Vacuum desiccator. 5. Biopsy punches with plungers, 2.0- and 1.0-mm diameter. 6. FluoroDish 35-mm glass-bottomed petri dish. These dishes are better suited for repeated imaging than glass coverslips, as they make it easy to maintain a sterile environment when imaging the devices every day. 7. Forceps. 8. 70% ethanol in water 9. Oxygen plasma cleaner (e.g., Harrick Plasma, Catalog # PDC-001). 10. Type I collagen (50 μg/mL in 0.02 M glacial acetic acid) or fibronectin (5 μg/mL in PBS) solution. Specific ECM coating and buffer may vary based on cell type used and require some optimization. 11. Dulbecco’s phosphate-buffered saline (DPBS), no calcium, no magnesium. 12. Media for cell culture supplemented as necessary. For MDA-MB-231 breast cancer cells and HT1080 fibroblasts, we used Dulbecco’s Modified Eagle Medium supplemented with penicillin/ streptavidin and 10% fetal bovine serum (FBS).

336

Richa Agrawal et al.

2.7 Imaging and Analysis

1. Inverted fluorescence microscope. The microscope should have objectives with at least 10 or 20 magnification and fluorescence excitation/emission filters for GFP and/or other fluorophores of interest. Some specific applications (e.g., analysis of intracellular dynamics) may require higher magnification. 2. Microscope-mounted CCD or CMOS camera for image acquisition. 3. Image acquisition software, such as ZEN BLUE (Zeiss), Micromanager, or others. 4. ImageJ, FIJI, MATLAB, or other software for image analysis. 5. Cell culture media used for imaging. If the microscopy setup does not control CO2 levels, experiments should be conducted using media with 20–25 mM HEPES buffer. We used Fluorobrite™ DMEM supplemented with penicillin–streptavidin and 10% FBS, buffered with 25 mM HEPES.

3

Methods

3.1 Photomask Fabrication and Preparation for Wafer Fabrication

All fabrication steps should be performed in a dedicated clean-room facility under standard clean-room conditions with sufficient ventilation, protective equipment, and other standard safety precautions. The photomasks (Fig. 2b) contain an image of all of the features that will be etched into or deposited on the silicon wafer, such as the pillars and fluid reservoirs. Note that our design uses two photomasks, one for each layer. 1. Generate a design for the photomasks using AutoCAD software (e.g., L-edit or KLayout), and convert to GDSII file format for mask-writing, ensuring the tone of your mask is correct (positive or negative), performing a tone inversion if necessary, and including alignment marks (Fig. 2b). If microfluidic designs are highly complex, we recommend using a GenISys BEAMER to perform Boolean subtraction (see Note 1). Detailed instructions on mask design for microfluidics have been detailed elsewhere [19]. Our device design files are available upon request. 2. Using a Heidelberg DWL 2000 Mask Writer or equivalent, print the desired design onto the photomask. Develop the photomask and perform a chrome etch, then strip the photoresist. This photomask will be used to expose the design features into photolithography using a near-UV light. 3. Clean bare silicon wafer with a 10 min soak in Acid Piranha solution to remove all organic materials. If this is not practical in your clean room organization, instead clean silicon wafer by

Microfluidic Device to Study Nuclear Mechanobiology

337

rinsing thoroughly for 30 s with each water, IPA, and then acetone. After either treatment, dry wafer with N2 gun and bake wafer for 12 h at 90  C to dehydrate surface completely. Proceed to Subheading 3.2 within a week. The dehydration step is most critical for SU-8 features. We had success using SU-8 photolithography (Fig. 2a (top), Section 3.2A) to produce rapid coarse prototypes to resolve adhesion and fluid flow issues, especially using our initial mask in combination with features cut by hand or from rubylith (see Note 2). However, SU-8 was insufficient to resolve some of our closely spaced features with the desired depth-to-width ratio, and so we also describe our use of reactive ion etching (RIE) to overcome these challenges (Fig. 2a (bottom), Section 3.2B), which additionally ensures increased durability for repeated PDMS castings (see Note 9). 3.2 Fabricating Constriction Layer Using SU-8 Lithography

1. Center wafer on spin-coater and dispense 2 mL of SU-8 2005 in the center. Our spin settings were as given in Table 1. See Note 3 for additional details. The spin protocol may need to be adjusted slightly due to inherent variability, using the spin curve in the SU-8 2005 datasheet as a guideline. 2. Perform a preexposure bake on a preheated 95  C hot plate for 2.5 min (see Note 4). Remove wafer from hot plate and allow wafer to gradually cool to room temperature on a cooling plate undisturbed. Remove the resist from the outer 5 mm periphery of the wafer and the entirety of the backside using cleanroom swab soaked in acetone. 3. Expose the resist using hard or vacuum contact and UV light with a long-pass filter (see Notes 5 and 6). 4. Post-exposure bake the wafer on a preheated hot plate for 3.5 min, then remove from hot plate and allow to cool to room temperature on level surface or cooling block. Check for defects such as craters, cracks, or scratches in photolithography under a microscope at this step, and repeat if needed (Fig. 4a).

Table 1 Settings for spin coating of 5-μm thickness SU-8 layer Spin curve for 5 μm of SU-8 2005

RPM

R/S (acceleration/ deceleration)

Seconds

Ramp up

500

100

10

Spin

3000

300

30

Ramp down

100

100

30

338

Richa Agrawal et al.

Fig. 4 Troubleshooting microfluidic device fabrication. (a) Successful photolithography (left), compared to defects in photolithography of SU-8 (middle, right) Scale bars: 30 μm, 20 μm, 4 mm, respectively. See Note 14. (b) Good adhesion (left), as defined by observing fluorescently labeled nuclei deforming around pillars in all areas. Incomplete adhesion during bonding of microfluidic devices, likely caused by issues during washing and bonding, will cause issues with focus (middle) or cells migrating directly under pillars (right). Scale bars: all 30 μm. Revisit Notes 18–24. (c) Uniform dispersion of cells across entrance of constrictions (left). Air bubbles can prevent cells from dispersing evenly (middle), or otherwise cells may be unevenly distributed (right). See Note 27. Scale bars: all 1 mm. (d) top view and side reconstruction of z-stacks taken on a confocal microscope, using fluorescent Texas Red ™ labeled dextran for easy visualization. The visible slight sidewallsloping did not prove to be an issue of concern. Scale bar: 30 μm. (e) A closer view of dashed region of orthogonal projection in (d), used to determine sidewall angle, α, of 97.8 . Scale bar: 4 μm

5. Soak the wafer in SU-8 developer for 5 min to wash away undeveloped resist. Rinse several times with IPA and then water (see Note 7). 3.3 Fabricating Constriction Layer Using RIE

1. Center wafer on spin-coater and dispense 2 mL of P-20 solvent to cover the wafer on spin coater and spin for 1 min to remove any dangling water bonds. The same settings used for photolithography spin should be adequate. 2. Dispense 2 mL of AZ nLOF 2020 resist in the center of the wafer and spin for 45 s at 3000 rpm with ramping of 500 rpm/ s (see Note 3). This should give a resist thickness of 1.7 μm. 3. Perform a preexposure bake on a preheated 110  C hot plate for 60 s (see Note 4). Remove wafer from hot plate and allow wafer to gradually cool to room temperature on a cooling plate

Microfluidic Device to Study Nuclear Mechanobiology

339

undisturbed. Remove edge bead from the outer 5 mm periphery of wafer and backside using cleanroom swab soaked in acetone. 4. Expose the wafer using the Suss MA6 contact aligner with hard vacuum contact (see Note 6). 5. Perform a post-exposure bake on a preheated 110  C hotplate for 60 s to catalyze the photoacid crosslinking. Take wafer off and place on a level surface or cooling block to let the wafer cool to room temperature. Check for defects such as craters, cracks, or scratches in photolithography under a microscope at this step, repeat if needed (see Note 7). 6. Perform a low power oxygen plasma descum at 100 standard cubic centimeters per minute (sccm) oxygen for 30 s in the Anatech Asher to remove any nanometer-thick areas of resist that could affect etching. Addition of 5–10 sccm nitrogen will allow visualization of the plasma as a purple gas to ensure that it has covered the wafer. 7. Etch using HBr reactive ion etching in the Oxford Cobra ICP or equivalent (see Note 9), quantifying etch depth periodically with a profilometer to etch to a total depth of 5.6 μm with resist on (see Note 10). 8. Perform a high-power oxygen plasma in the Anatech Asher or equivalent (900 sccm oxygen for 10 min) to remove all photoresist. Visually inspect the wafer after this step, as photolithography constrictions may not be identically sized when etched into silicon. Characterize with a profilometer to determine final etched height into silicon after removing the photoresist. Ensure that the constriction geometry and sidewall profiles align with that measured during dextran characterization of final devices (see Subheading 3.4, step 20). 3.4 Upper SU-8 Layer and Final Processing

1. Center wafer on spin coater and dispense 2 mL of SU-8 100 in center of wafer. Our spin settings were as given in Table 2. The spin protocol may need to be adjusted slightly due to inherent variability, using the spin curve in the SU-8 100 datasheet as a guideline (see Note 3). 2. Preexposure bake wafer ramping from room temperature to 62.5  C at 1.5  C/min, holding for 8 hours, then increase temperature to 67.5  C for 14 h or overnight (see Notes 2 and 12). Cover with a crystallization dish resting on two microscope slides such that it does not touch the wafer (see Note 13). 3. Use the Suss MA6 aligner with a long-pass filter to align these features to the first constriction layer using the alignment

340

Richa Agrawal et al.

Table 2 Settings for spin coating of 200-μm thickness SU-8 layer Spin curve for 200 μm of SU-8 100

RPM

R/S (acceleration/ deceleration)

Seconds

Ramp up

500

100

10

Spin

1500

100

60

Ramp down

100

100

15

marks and expose. Our successful protocol was 30 s, 6 s of rest, followed by 30 s. Optimization may be required (see Note 6). 4. Postexposure bake again with petri dish lid ramping from room temperature to 95  C at 1.5  C/min and hold for 1 min. Turn the hot plate off and let cool slowly to room temperature. 5. Leave wafer feature-side up in SU-8 developer overnight to remove undeveloped resist, then rinse three times for 10 s each with IPA and deionized water. 6. Apply a fluoro-octyl trichlorosilane (FOTS) antistiction coating using the Molecular Vapor Deposition (MVD) tool to ensure that the PDMS cast detaches easily from the wafer on the constrictions and other small features. Verify a contact angle of 95 on bare silicon (see Note 15). 3.5 Casting and Mounting Devices (Fig. 2c)

1. Remove dust or other particulates from wafer surface using an air gun, and then place silicon wafer with features facing upward in a 150 mm petri dish (or use plastic mold, see Note 16). 2. Mix 50 g of PDMS components at a 10:1 ratio of base–curing agent and stir to mix thoroughly. Bubbles may appear but will be removed in the following step. 3. Place PDMS into a vacuum desiccator at 30 psi for 30 min to eliminate bubbles. 4. Pour PDMS over wafer. If air bubbles are still present, repeat degassing. A gentle stream of pressurized air aimed at the PDMS may be useful to remove all remaining bubbles. 5. Bake PDMS mixture in oven for at least 2 h at 65  C. 6. When baking PDMS directly on the wafer with constrictions made from SU-8 features, it is recommended to turn the oven off and let the wafer cool gradually as this prevents excess stress on the SU-8 features and elongates the life of the wafer. This step is less crucial when the constriction layer has been formed using RIE, as SU-8 features are more susceptible to cracking or baking.

Microfluidic Device to Study Nuclear Mechanobiology

341

7. Use knife or razor blade to gently cut around the wafer and peel the PDMS. Pipetting some isopropyl alcohol into the separation between the wafer and PDMS may be helpful for PDMS removal. However, it is best to never let any solvents dry on the wafer. After removal of the PDMS, rinse the wafer with deionized water and dry with an air gun. 8. Cut holes into the PDMS to load media and cells into the PDMS devices by using a biopsy punch in each of the three sections with one 2-mm biopsy punch (center) and two 1-mm biopsy punches (outer edge), in locations shown in black on Fig. 1a (see Note 17). 9. Hold device with forceps and gloved hands, ensuring not to disturb the area with the constrictions, and rinse with IPA and deionized water for 5 s each. Repeat twice (see Note 18). 10. Use pressurized air to dry the device, ensuring all water is removed from within the biopsy punched holes and from the constrictions. Place constriction-side up in the plasma cleaner. 11. Rinse 35 mm glass-bottomed petri dish with IPA and water for 5 s each, three times and place in the very back of the plasma cleaner, media-side up (see Note 19). Place alongside the PDMS in the plasma cleaner. 12. Turn on plasma power and close air valves in the plasma cleaner. Turn on vacuum for 10 min and then turn plasma to high (see Note 20). 13. Plasma treat the PDMS devices and glass petri dish for 15 min. 14. Turn off plasma and very slowly release vacuum so the devices are not overturned or disturbed. 15. Place PDMS feature-side down onto the glass portion of the petri dish (see Note 21). 16. Gently press down with one gloved finger, starting at one side and moving to the other to gently push out air bubbles at the contact interface (see Note 22). 17. To improve the bonding, place on a preheated 95  C hot plate for 5 s, remove for 2 s and repeat for 60 s total (see Note 23). 18. Let devices cool for at least 10 min, then move to a cell culture hood for sterilization and collagen coating. 19. Inspect the device for any defects in bonding (Fig. 4b, see Note 24). 20. If using devices for the first time, it is highly recommended to verify that device geometry matches the intended design. This can be achieved by filling devices with a fluorescent solution such as dextran in PBS and using a high magnification objective to take confocal z-stacks. Create an orthogonal projection across the z-stacks and examine the feature profile to check

342

Richa Agrawal et al.

for sidewall sloping due to resist undercut (Fig. 4d, e). It is also recommended to verify in the x-y plane that constriction sizes match the desired dimensions. 3.6 Functionalization of the Devices and Cell Seeding (Fig. 2c)

1. Pipette 3 mL of 70% ethanol into glass-bottomed dishes with the devices, thoroughly rinsing by adding and removing some liquid from the seeding ports as well using a pipette. Incubate devices in 70% ethanol for 10 min to sterilize. 2. Remove ethanol from the devices and rinse with deionized water three times, once again ensuring to rinse the seeding ports. 3. At this point, the devices can be functionalized with extracellular matrix proteins or other biologically relevant coating, diluted in the relevant buffer (PBS, media, etc.). The specific selection will depend on the cell line used. For mouse embryonic fibroblasts, these devices were functionalized using 50 μg/ mL rat tail collagen type I diluted in 0.02 M acetic acid, and then incubated at 4  C overnight to ensure adequate adsorption to the device (see Note 25). 4. Rinse the inside of the devices with media three times and fill devices with media while preparing cell suspension. 5. Trypsinize, count, and resuspend cells in cell culture media to ten million cells/mL or desired concentration for seeding into devices (see Note 26). 6. Remove all media from devices. Pipette 50,000 cells resuspended in 5 μL into each of the three large seeding ports. Check for successful cell seeding with a bright-field microscope, as the cells should be uniformly distributed against the entry of the constriction area (Fig. 4c, see Note 27). 7. Gently add several drops of fresh media to each of the large seeding ports, while being careful that the devices do not overflow. Place in cell culture incubator for 4 h to allow cells to adhere. 8. After this time, any cells that have not adhered should be rinsed out from the device by gently adding liquid to the seeding ports and removing several times with a fine pipette. Add approximately 4 mL of media to submerge the devices and allow for sufficient nutrient access.

3.7 Imaging and Analysis of Cell Migration

The devices enable time-lapse microscopy, as described previously [17], but can also be used to assess migratory fitness as a function of the distance traveled by the cells from the seeding port into the constriction area. This avoids time-consuming time-lapse imaging, phototoxicity, and photobleaching due to repeated imaging, and does not require a heated chamber or focus stabilization mechanisms. Here, we briefly outline our image acquisition and analysis

Microfluidic Device to Study Nuclear Mechanobiology

343

protocol to image the cells daily and compare the distance traveled between confined conditions and control conditions. We have validated this method to assess migratory fitness using previously characterized Lmna
/
and wild-type (Lmna+/+) MEFs, showing that the Lmna
/
MEFs travel further than the wild-type MEFs over the course of 3 days (Fig. 3). 1. Seed the cells at least 12–24 h before beginning imaging, as it takes some time for the cells to enter constrictions. Cells with fluorescently labeled nuclei (e.g., expressing H2B-GFP) will be easiest to visualize. Some optimization should be performed for each cell type to determine the most suitable pillar densities and time-points (see Note 28). 2. Change media in devices to imaging media. Place the dish on the appropriate stage to hold the dish securely. 3. Bring the cells into focus. Capture several (5–10) images around the perimeter of the device, containing the interface between the seeding port and the constriction area to determine the distance traveled in each image (Fig. 3a). After the cells have moved sufficiently far through the devices, it may be necessary to use a tile scan to stitch together several images. We used a 20 objective to measure the traveled distance, however, an objective with higher magnification and NA may be required to capture subcellular details and dynamics. 4. Using ZEN, or an equivalent image analysis software, draw a line from the edge of the seeding port to the nucleus in the constriction area (Fig. 3a, see Note 28). 5. Repeat this process for other pillar densities and low-density control, genotype(s), and treatment(s) (e.g., Lamin A/Cdeficient cells, LINC complex disrupted cells). 6. Compare the distance traveled in each confined condition to the respective unconfined distances to account for differences in unconfined migration speeds between cell lines. Larger normalized distances indicate an improved confined migration ability (Fig. 3b, c).

4

Notes 1. GenISys BEAMER is superior to L-edit for machine fracturing of complex curved layouts. Thus, performing Boolean subtraction in BEAMER with complex designs can drastically reduce mask writing time. Our mask design contained a very large number of data points, even after converting most circles to 32-sided polygons, and this could only be performed with LayoutBEAMER.

344

Richa Agrawal et al.

2. If large scale modifications are desired, it is often easier and faster to laser cut coarse features out of light-blocking rubylith art film. The rubylith masks can be overlaid on top of an existing photomask to create additional features in the same layer, or overtop a “blank” mask consisting of transparent glass to create a layer consisting of just the new rubylith features. The major drawback of this approach is that alignment to other layers on the wafer is only possible visually, and thus not very accurate. Nonetheless, we were able to produce several coarse modifications to our designs using this method during our prototyping phase, resulting in substantial time and cost savings. 3. Ramping the speed gradually while accelerating and decelerating is recommended to allow for more even dispersion of photoresist on the wafer and to prevent buildup of excess resist near the edge of the wafer. This step is particularly crucial for thick films of SU-8. If the resulting film is too thin or too thick, the hold spin speed should be adjusted accordingly. However, if the spin speed is below 1000–1500 RPM, this may result in nonuniform thickness of the resist. 4. A preexposure bake before development evaporates some of the solvent to reduce mask adhesion, prevent photolithographic defects or delamination. This soft bake is particularly critical for highly viscous and “sticky” materials such as SU-8, which is very susceptible to craters and defects. 5. To achieve maximal resolution with SU-8 2005, use a longpass filter to remove wavelengths below 365 nm, and use high vacuum contact settings to minimize diffraction effects. 6. A rough estimate of exposure time can be determined by using the manual to determine the dosage for the specific film thickness, and then dividing the dosage by the lamp intensity. However, there is some inherent variability at this step and lamp intensity naturally varies with lamp age. We recommend to perform an exposure array using a manually positioned rubylith film section on top of the mask to try several different exposure times on different parts of the wafer to determine an ideal exposure time. We personally found that the ideal exposure time for 5 μm SU-8 with a long pass filter was 15 s with a lamp intensity of 11 mJ/cm2. After performing the exposure array, post-exposure baking, and developing the wafer, inspect using a microscope to determine the ideal feature size and check for any defects. 7. It is very important to ensure that all residual undeveloped SU-8 has been removed before proceeding to prevent faulty adhesion in the second layer. A longer soak in SU-8 developer or an acetone rinse is usually sufficient.

Microfluidic Device to Study Nuclear Mechanobiology

345

8. If there are defects in the AZ nLOF 2020 photolithography, stripping of the photoresist can be performed in any plasma tool using a high-power oxygen plasma (900 sccm oxygen, for 10 min). Then, proceed from step 3. 9. Reactive ion etching is a type of dry etching where high-energy ions are bombarded at the wafer surface that is most useful for resolving high aspect ratio features. Various RIE methods alter gas composition, methods of energy generation, and other parameters to generate varying profiles and depths. All RIE methods wear away at both the resist and the silicon, with a characteristic selectivity for resist–silicon. The features must therefore be etched into a sufficiently thick resist to allow for etching to the desired depth, but at some point the resist thickness cannot be increased while still resolving fine features. We tried several methods of RIE to achieve the desired feature resolution, depth to 5 μm, and vertical sidewall profiles. While deep reactive ion etching (DRIE) seemed promising, scalloping along the sidewalls of the constrictions caused the PDMS pillars to become permanently lodged in the wafer. In the end, we had the best success with HBr etching in the Oxford Cobra etcher, as this highly anisotropic etch produced a vertical sidewalls and the silicon–resist selectivity of 4.8:1 was sufficient to etch to 5 μm depth with 2 μm of our selected resist. It is likely that Cl2 etching would also work. We also had success with using an etch designed for photonic device fabrication and using the Inductively Coupled Plasma (ICP-RIE) etch in the Unaxis 770, but ultimately switched to HBr etching due to cost and time saving. 10. A rough characterization of etch depth can be done by using a profilometer as described in Note 11, but this measurement is imprecise because the resist will be removed over time due to a selectivity of 5:1 for silicon–resist. It was found that etching in the Oxford Cobra etcher to a total depth of 5.6 μm resulted in 5.2 μm pillars after the resist was removed. Note that as etch depth increases, etch rate decreases. 11. Characterization by profilometer of the etch depth must be performed at the edge of the pillared area to the flat, unetched area, as 5 μm deep features that are 5 μm wide exceed the aspect ratio capabilities of many pointed conical stylus. Because it is not possible to know exactly how much resist has been removed, several rounds of etching may be necessary for obtaining a precise height. 12. This protocol varies significantly from the guidelines in the SU-8 manual, however we found it to more resilient to cracking. A gradual temperature ramping process minimizes film stress and is very helpful for preventing cracks and delamination.

346

Richa Agrawal et al.

13. Cover the wafer with a crystallization dish resting on two microscope slides to create a more humid microenvironment that prevents formation of a “skin” of SU-8 that stalls solvent evaporation. 14. Defects in SU-8 indicate that the exposure time needs further optimization. However, often these cracks can be merged and delamination alleviated using a curing bake in an oven by ramping from room temperature to 200  C at 10  C/minute. Afterward, turn the oven off and let cool gradually to room temperature. 15. Verification of contact angle can be performed using the VCA optima. This ensures that the FOTS coating was applied correctly to the wafer. In our experience, residual SU-8 caused an improper FOTS coating on the wafer and a 20 s rinse with SU-8 remover solvent or acetone followed by water and a repeated dehydration step resolved this issue. 16. To protect the wafer, we recommend that the first PDMS casting be used to produce a plastic mold, and this plastic mold is used to reproduce migration devices. Detailed notes on producing a secondary plastic mold can be found in Desai et al. [20]. 17. Punching through the PDMS while it rests constriction-side up will both prevent pressure from being applied and destroying the features and prevent the ridge caused by biopsy punching from causing issues during glass bonding. 18. Ensure that IPA and water do not run off gloves and onto devices during rinsing, as the runoff from gloves may cause issues during glass bonding. 19. The plasma is often strongest in the back of most plasma cleaners. Ensure that the dishes and devices are as far back as possible within the plasma cleaning chamber. 20. Toggle air flow valves in the plasma cleaner to ensure plasma is bright pink for the duration of the treatment, as many adhesion problems result from improper plasma treatment. 21. Once the PDMS is placed on the glass, it should not be removed or disturbed. If placing multiple devices on a coverslip, it may be helpful to align them in a similar orientation to make imaging and analysis easier. 22. Some practice may be required to ensure proper adhesion. Using too little force will result in some nonadhered areas, whereas too much force will collapse pillared areas. 23. This activates the surface of the PDMS and glass and improves bonding and adhesion; however, the plastic of the petri dishes warps at this temperature. Alternating 5 s on and off heat for a minute increases bonding strength and does not warp the

Microfluidic Device to Study Nuclear Mechanobiology

347

plastic. We tried to treat these dishes for a longer time at a lower temperature, but this led to both deadhesion and collapse. 24. Inspection should be performed under a brightfield microscope to ensure that the device has properly adhered, as any nonadhered areas will appear a different color than the rest. These will become increasingly apparent as cells are seeded and imaged, as cells will appear under the constrictions. 25. Due to the highly hydrophobic nature of PDMS, it is possible that the constriction area may not be initially filled entirely with the coating solution. It is helpful to check for bubbles again after an hour and vigorously pipette through seeding ports and from the outside edge of devices to remove any air bubbles. Repeat this process until the entire constriction area is filled and no air bubbles are observed. Fibronectin can also be used as described previously; other biological coatings should also be suitable for surface functionalization to promote cell adhesion and migration. 26. The exact number of cells will depend on the specific experimental goals and the particular cell line being used. Seeding 50,000 MEF cells into each of the three seeding ports was ideal for imaging distance traveled after 3 days. If conducting longer-term experiments, it may be better to seed fewer cells such that the cells in the seeding port do not become overcrowded. Similarly, if intending to image the cells the same day, it may be better to seed more so that the cells enter the constrictions. The volume of the seeding port is 5 μL, so it is best to adjust the concentration of cell resuspension, rather than the volume. 27. If the distribution of cells is uneven, withdraw some liquid from the side biopsy punches and replace into central punch to draw cells toward the leading edge. Repeat this process until cells are evenly distributed and no air bubbles are present. If necessary, aspirate cells and repeat. 28. Various cell types will move through the devices at varying speeds and may display unequal abilities to perform confined migration, so some initial characterization may be required to determine what the ideal pillar densities and time-point used will be ideal. It is recommended that the “leading edge” cells are allowed to travel through at least 200 μm to obtain a robust measurement. This will likely require the initial characterization of cell migration through several different density of pillars over time to determine the ideal density of pillars. Additionally, at higher densities, we have observed stalling of migration and substantial cell death after cells had passed 30 1-μm

348

Richa Agrawal et al.

tight constrictions or 75–100 of the 2–4-μm wide constrictions, and not in control conditions. It is best to choose an endpoint at least 1–2 days before migration stalls. 29. We tried two methods of analysis: measuring the average distance of all cells in the constriction area, as well as only measuring the furthest cells in every area. We found the most consistent data was produced by only assessing the furthest cells.

Acknowledgments This work was supported by NIH awards R01 GM137605 and U54 CA210184 and the Department of Defense Breast Cancer Research Program Breakthrough Award BC150580 to J.L. This work was performed in part at the Cornell NanoScale Science & Technology Facility (CNF), a member of the National Nanotechnology Coordinated Infrastructure (NNCI), which is supported by the National Science Foundation (Grant NNCI-2025233). References 1. Shah P, Hobson CM, Cheng S et al (2021) Nuclear deformation causes DNA damage by increasing replication stress. Curr Biol 31: 753–765.e6. https://doi.org/10.1016/j.cub. 2020.11.037 2. Denais CM, Gilbert RM, Isermann P et al (2016) Nuclear envelope rupture and repair during cancer cell migration. Science 352: 353–358. https://doi.org/10.1126/science. aad7297 3. Pfeifer CR, Irianto J, Discher DE (2019) Nuclear mechanics and cancer cell migration. In: la Porta CAM, Zapperi S (eds) Cell migrations: causes and functions. Springer, pp 117–130 4. Elosegui-Artola A, Andreu I, Beedle AEM et al (2017) Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171:1397–1410.e14. https://doi.org/10. 1016/j.cell.2017.10.008 5. Golloshi R, Martin RS, Das P et al (2019) Constricted migration contributes to persistent 3D genome structure changes associated with an invasive phenotype in melanoma cells. bioRxiv 856583 6. Kirby TJ, Lammerding J (2018) Emerging views of the nucleus as a cellular mechanosensory. Nat Cell Biol 20:373–381. https://doi. org/10.1038/s41556-018-0038-y

7. Venturini V, Pezzano F, Castro FC et al (2020) The nucleus measures shape changes for cellular proprioception to control dynamic cell behavior. Science 370:eaba2644. https://doi. org/10.1126/science.aba2644 8. Lomakin AJ, Cattin CJ, Cuvelier D et al (2020) The nucleus acts as a ruler tailoring cell responses to spatial constraints. Science 370: aba2894. https://doi.org/10.1126/science. aba2894 9. Nava MM, Miroshnikova YA, Biggs LC et al (2020) Heterochromatin-driven nuclear softening protects the genome against mechanical stress-induced damage. Cell 181:800–817. e22. https://doi.org/10.1016/j.cell.2020. 03.052 10. Fraley SI, Wu PH, He L et al (2015) Threedimensional matrix fiber alignment modulates cell migration and MT1-MMP utility by spatially and temporally directing protrusions. Sci Rep 5:1–13. https://doi.org/10.1038/ srep14580 11. Zuela-Sopilniak N, Lammerding J (2019) Engineering approaches to studying cancer cell migration in three-dimensional environments. Philos Trans R Soc Lond B Biol Sci 374:20180219. https://doi.org/10.1098/ rstb.2018.0219

Microfluidic Device to Study Nuclear Mechanobiology 12. Richards et al (2018) Engineered models of confined cell migration. Physiol Behav 176: 139–148. https://doi.org/10.1146/annurevbioeng-071114-040654.Engineered 13. Paul CD, Mistriotis P, Konstantopoulos K (2017) Cancer cell motility: lessons from migration in confined spaces. Nat Rev Cancer 17:131–140. https://doi.org/10.1038/nrc. 2016.123 14. Zanotelli MR, Rahman-Zaman A, VanderBurgh JA et al (2019) Energetic costs regulated by cell mechanics and confinement are predictive of migration path during decision-making. Nat Commun 10:1–12. https://doi.org/10. 1038/s41467-019-12155-z 15. Elacqua JJ, Mcgregor AL, Lammerding J (2018) Automated analysis of cell migration and nuclear envelope rupture in confined environments. PLoS One 13:e0195664 16. Gale BK, Jafek AR, Lambert CJ et al (2018) A review of current methods in microfluidic

349

device fabrication and future commercialization prospects. Inventions 3:60. https://doi. org/10.3390/inventions3030060 17. Keys J, Windsor A, Lammerding J (2018) Assembly and use of a microfluidic device to study cell migration in confined environments. Methods Mol Biol 1840:101–118 18. Davidson PM, Denais C, Bakshi MC, Lammerding J (2014) Nuclear deformability constitutes a rate-limiting step during cell migration in 3-D environments. Cell Mol Bioeng 7:293–306. https://doi.org/10. 1007/s12195-014-0342-y 19. Perumal V, Hashim U (2013) Chrome mask design for microfluidic fabrication. Adv Mater Res 795:276–280. https://doi.org/10.4028/ www.scientific.net/amr.795.27620 20. Desai SP, Freeman DM, Voldman J (2009) Plastic masters - Rigid templates for soft lithography. Lab Chip 9:1631–1637. https://doi. org/10.1039/b822081f

Part VII Imaging NPCs and Transport

Chapter 23 Speed Microscopy: High-Speed Single Molecule Tracking and Mapping of Nucleocytoplasmic Transport Steven J. Schnell, Mark Tingey, and Weidong Yang Abstract The nuclear pore complex (NPC) functions as a gateway through which molecules translocate into and out of the nucleus. Understanding the transport dynamics of these transiting molecules and how they interact with the NPC has great potentials in the discovery of clinical targets. Single-molecule microscopy techniques are powerful tools to provide sub–diffraction limit information about the dynamic and structural details of nucleocytoplasmic transport. Here we detail single-point edge-excitation subdiffraction (SPEED) microscopy, a high-speed superresolution microscopy technique designed to track and map proteins and RNAs as they cross native NPCs. Key words Single-molecule microscopy, Sub–diffraction limit imaging, Superresolution light microscopy, Single-point edge-excitation subdiffraction microscopy, Nucleocytoplasmic transport, Live cell imaging

1

Introduction The nuclear pore complex (NPC) is a complex assembly of proteins spanning the inner and outer membranes of the nuclear envelope (NE), forming a gateway between the nucleus and the cytoplasm of eukaryotic cells. The number of NPCs in the NE varies from hundreds in yeast to thousands in human cells [1–7]. As the major exchange pathway for proteins and genetic materials, the NPC or its various subunits plays a crucial role in the regulation and control of many cellular processes [8–17]. The structure of the NPC consists of an eightfold rotationally symmetrical core channel with an associated nuclear basket structure and eight cytoplasmic fibrils [18]. The NPC measures approximately 50 nm in diameter at the narrowest point, and about 200 nm axially in total length in vertebrate cells [18–21]. About one-third of the approximately 30 different proteins that make up the NPC (in copies of multiples of eight each) are filamentous nucleoporins (Nups) containing regions of extensive phenylalanine–glycine (FG) repeats

Martin W. Goldberg (ed.), The Nuclear Pore Complex: Methods and Protocols, Methods in Molecular Biology, vol. 2502, https://doi.org/10.1007/978-1-0716-2337-4_23, © Springer Science+Business Media, LLC, part of Springer Nature 2022

353

354

Steven J. Schnell et al.

[19]. Termed FG Nups, these proteins make up a selective barrier [18, 19, 22–26] in the NPC and are the primary players in regulation of the selective transport of macromolecules into and out of the nucleus. Translocation of signal-independent small molecules (