Cell Cycle Oscillators: Methods and Protocols (Methods in Molecular Biology, 2329) 1071615378, 9781071615379

Table of contents :
Preface
Contents
Contributors
Chapter 1: Cell Cycle Control: A System of Interlinking Oscillators
1 Introduction
1.1 Anatomy of a Cell Cycle Oscillator
1.2 Checkpoints: Putting a Break on Cell Cycle Oscillators
2 Entering the Cell Cycle and G1-S
2.1 Cyclin D as a Mitogenic Sensor for the Cell Cycle
2.2 Phosphorylation of pRb Underlies the Restriction Point
2.3 The G1 DNA Damage Checkpoint
3 Control of S Phase
3.1 Initiation of S Phase
3.2 Prevention of Rereplication
3.3 Replication Checkpoint
4 G2-Mitosis
4.1 Cyclin B-CDK1 as the Engine of Mitosis
4.2 Feedback Control Ensures Biphasic Activation of CDK1
4.3 Kickstarting the CDK1 Activating Loops
4.4 Greatwall (MASTL) Helps Maintain the Mitotic State
4.5 The G2 DNA Damage Checkpoint
5 Mitosis-G1
5.1 APC/C Drives Mitotic Exit
5.2 The Spindle-Assembly Checkpoint
References
Chapter 2: Substrate Phosphorylation Rates as an In Vivo Measurement of Kinase Activity
1 Introduction
1.1 Kinases and Their Activity
1.2 Caveats to Current Approaches to Measuring Kinase Activity
1.3 Direct Measurement of Initial Substrate Phosphorylation Rates In Vivo Can Be Used to Estimate Kinase Activity
2 Materials
2.1 Strains and Media
2.2 Kinase Inhibitor
2.3 Filtration Units
2.4 Yeast Protein Extraction
3 Method
3.1 Growing Cells
3.2 Inhibition, Release, and Time Course
3.2.1 Inhibition
3.2.2 Release
3.2.3 Time Course
3.3 Protein Extraction
3.4 Data Analysis
4 Notes
References
Chapter 3: Real-Time Monitoring of APC/C-Mediated Substrate Degradation Using Xenopus laevis Egg Extracts
1 Introduction
2 Materials
2.1 Xenopus laevis Egg Extract Preparation and Manipulation
2.2 Immunodepletion
2.3 Preparation of Fluorescently Labeled Substrate by In Vitro Translation
2.4 Measuring APC/C Activity Using a Fluorescence Plate Reader
3 Methods
3.1 Egg Extract Preparation
3.1.1 Preparing Interphase-Arrested Extract from CSF Extract
3.1.2 Preparing Mitotic Extract from Interphase Extract
3.2 Immunodepletion
3.3 Preparation of Fluorescently Labeled Substrate by In Vitro Transcription/Translation
3.4 Measuring APC/C Activity Using a Fluorescence Plate Reader
3.5 Data Analysis
4 Notes
References
Chapter 4: Fluorescent Peptide Biosensors for Probing CDK Kinase Activity in Cell Extracts
1 Introduction
2 Materials
2.1 Peptide Biosensor Preparation and Labeling
2.2 Source of Kinases: Recombinant Protein or Cell Extracts
2.3 Activity Assay in Black 96-Well Plates
3 Methods
3.1 Fluorescent Biosensor Preparation
3.1.1 Prelabeled CDKACT
3.1.2 Unlabeled CDKACT
3.2 Cell Extract Sample Preparation
3.3 Performing Kinase Assays with CDKACT Biosensors
3.4 Determination of Kinase Curves
3.5 Histogram Representation of Relative Maximal Activity
4 Notes
References
Chapter 5: Phosphatase and Kinase Substrate Specificity Profiling with Pooled Synthetic Peptides and Mass Spectrometry
1 Introduction
2 Materials
3 Methods
3.1 Defining Instrument Linear Response Range
3.2 Calculate Ionization Correction Factors for Substrate and Product Peptide Pairs
3.3 Enzyme Assay
3.4 LC-MS Analysis
3.5 Data Analysis
3.5.1 Substrate and Product Peak Identification
3.5.2 MS1 Peak Integration
3.5.3 kcat/KM Calculation
4 Notes
References
Chapter 6: Whole-Mount Immunostaining for the Identification of Histone Modifications in the S-Phase Nuclei of Arabidopsis Roo...
1 Introduction
2 Materials
2.1 Plant Material
2.2 Seed Sterilization
2.3 Media
2.4 Seed Sowing
2.5 Reagents
2.6 Immunostaining
2.7 Moisture Chamber
2.8 Observation
3 Methods
3.1 Seed Sterilization
3.2 Seed Sowing
3.3 Cultivation
3.4 EdU Staining
3.5 Immunostaining
3.6 Microscopic Observation
4 Notes
References
Chapter 7: Construction of a Full-Length 3′UTR Reporter System for Identification of Cell-Cycle Regulating MicroRNAs
1 Introduction
2 Materials
2.1 Cloning
2.2 DNA Transfection and Selection of Stable Cell Lines
2.3 miRNA Transfection and Assaying of Luciferase Expression
3 Methods
3.1 Cloning of 3′UTR from Human Genomic DNA into pmirGLO Vector
3.2 Establishment of 3′UTR Dual Luciferase Reporter Cell Lines
3.3 miRNA Library Screening of 3′UTR Reporter Targeting
4 Notes
References
Chapter 8: Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis
1 Introduction
1.1 Cyclin-Cdk Complexes in Cell Cycle Control
1.2 Purification of Cyclin-Cdk Complexes for In Vitro Analysis
1.3 Use of Suitable Linker Between Cyclin and Cdk
1.4 Advantages of the Cyclin-Cdk1 Fusion Strategy in Purification
1.5 Cyclin-Cdk Fusion Complex Purification and Activity Measurement
2 Materials
2.1 Buffers
2.1.1 Lysis Buffer for Controlling Cyclin- L-Cdk1 Expression
2.1.2 Buffers for the Purification of Cyclin- L-Cdk1 Complex (See Note 1)
2.1.3 Kinase Assay Buffers
2.2 Reagents
2.3 Media
2.4 Expression Vectors
2.5 Yeast Strains
2.6 Equipment
2.7 Software
3 Methods
3.1 Construction of Yeast Expression Plasmid
3.2 Yeast Transformation and Characterization of Expression
3.3 One Potential Strategy for Inducing Large-Scale Cultures
3.4 Purification Procedure of Cyclin-Dependent Kinase Complex from Budding Yeast
3.5 In Vitro Kinase Assay to Measure the Activity of Purified Cyclin- L-Cdk1 Complexes
4 Notes
References
Chapter 9: Optimizing Cell Synchronization Using Nocodazole or Double Thymidine Block
1 Introduction
2 Materials
2.1 Cell Cycle Drugs
2.2 DNA Staining for Viability or FACS Analysis
2.3 Cell Culture
3 Methods
3.1 Preliminary Assessment of Nocodazole Concentrations
3.2 Selection of Nocodazole and Thymidine Timings
3.3 Collection of Cells After Release
4 Notes
References
Chapter 10: Highly Synchronous Mitotic Progression in Schizosaccharomyces pombe Upon Relief of Transient Cdc2-asM17 Inhibition
1 Introduction
2 Materials
2.1 Cell Culture and Filtration
2.2 Calcofluor Staining to Score the Septation Index
2.3 Generation of p13Suc1 Beads
2.4 Preparing for the Histone H1 Kinase Assay
3 Methods
3.1 cdc25-22 Induction Synchronization
3.2 cdc2-asM17 Induction Synchronization (See Note 4)
3.3 Calcofluor Staining
3.4 Histone H1 Kinase Assay
3.4.1 Preparation of Magnetic p13Suc1 Beads
3.4.2 Histone H1 Kinase Assay: Sample Collection (See Note 16)
3.4.3 Histone H1 Kinase Assay: Reaction
3.4.4 Histone H1 Kinase Assay: Detection
4 Notes
References
Chapter 11: Elucidating Human Mitosis Using an Anaphase-Like Cell-Free System
1 Introduction
2 Materials
2.1 Buffers and Reagents
2.1.1 Cell Culture Maintenance
2.1.2 Generation of NDB Cell System
2.1.3 Validating Mitotic Arrest in NDB Cells Expressing Nondegradable Cyclin B1
2.1.4 Preparation of NDB Mitotic Protein Lysate
2.1.5 Preparation of NDB Mitotic Extracts
2.1.6 Degradation and Mobility-Shift Assays in NDB Mitotic Extracts
2.1.7 G1-Like NDB Extracts
2.1.8 Immunoprecipitation of APC/C from NDB Extracts and Immunoblot of Cdc20/Cdh1
2.2 Equipment
2.2.1 General Use
2.2.2 Cell Culture Maintenance
2.2.3 Microscopy
2.2.4 Flow Cytometry
2.2.5 Extract Preparation
2.2.6 Degradation/Phosphorylation Assays
2.2.7 Immunoblot and Immunoprecipitation
3 Methods
3.1 Generation of NDB Cell Line
3.2 PI Staining Protocol for Quantifying DNA Content by Flow Cytometry
3.3 Chromosome Spreads
3.4 Harvesting Mitotic NDB Cells for Whole-Cell Protein Lysate Preparation and Immunoblot Assays
3.5 Preparation of NDB Mitotic Extracts
3.6 Degradation and Mobility Shift Assays in NDB Extracts
3.7 Recapitulating a G1-Like State in NDB Extracts
3.7.1 Mitotic Exit in Tet-Induced NDB Cells
3.7.2 Mitotic Exit in NDB Extracts
3.7.3 Immunoprecipitation of APC/C in Mitotic vs. G1-Like NDB Extracts
4 Notes
References
Chapter 12: EDU (5-Ethynyl-2′-Deoxyuridine)-Coupled Fluorescence-Intensity Analysis: Determining Absolute Parameters of the Ce...
1 Introduction
2 Materials
2.1 Cell Culture and EDU Supplementation
2.2 Cell Collection and Flow Cytometry
3 Methods
3.1 Cell Culture and Labelling with EDU
3.2 EDU-DNA Detection
3.3 Flow Cytometry and Data Analysis
4 Notes
References
Chapter 13: High-Resolution Analysis of Centrosome Behavior During Mitosis
1 Introduction
2 Materials
2.1 Cell Lines and Culture Conditions
2.2 Transient Plasmid DNA and Small Interfering RNAs (siRNAs) Transfections
2.3 Micropatterning on Glass Coverslip with Deep UV Light
2.4 Cell Confinement
2.5 Live-Cell Imaging
2.6 Analysis and Data Extraction from Live-Cell Imaging Datasets
3 Methods
3.1 Transient Plasmid DNA and Small Interfering RNAs (siRNAs) Transfections
3.2 Micropatterning on Glass Coverslip with Deep UV Light
3.2.1 Coating of Glass Coverslips with PLL-g-PEG
3.2.2 UV Illumination of PLL-g-PEG-coated Glass Coverslips Using a Quartz Photomask
3.2.3 Seeding Cells on FBN-Patterned Glass Coverslips
3.3 Cell Confinement
3.3.1 Cell Seeding for Confinement
3.3.2 Using the Dynamic Cell Confiner
3.4 Live-Cell Imaging
3.5 Analysis and Data Extraction from Live-Cell Imaging Datasets
3.5.1 Single-Cell Analysis
3.5.2 Compile Results
4 Notes
References
Chapter 14: Assaying Cell Cycle Progression via Flow Cytometry in CRISPR/Cas9-Treated Cells
1 Introduction
2 Materials
2.1 Cell Culture and Transfection
2.2 Flow Cytometry Assay
3 Methods
3.1 Transfection and EdU Treatment of Cells
3.2 Click Chemistry Labeling Flow Cytometry-Based Cell Cycle Progression Assay
3.3 Flow Cytometry
4 Notes
References
Chapter 15: Use of Mitotic Protein Kinase Inhibitors and Phospho-Specific Antibodies to Monitor Protein Phosphorylation During...
1 Introduction
2 Materials
2.1 Generation and Purification of Phospho-Specific Antibodies
2.2 Cell Culture Reagents and Drugs
2.3 Cell Lysis, SDS-PAGE and Immunoblotting
3 Methods
3.1 Generation, Purification, and Testing of Phospho-Specific Antibodies
3.1.1 Generation of Phospho-Specific Antibodies
3.1.2 Preparing Phosphopeptide Affinity Column
3.1.3 Purification of Phospho-Specific Antibodies
3.1.4 Validation of Purified Phospho-Specific Antibodies Using Dot Blots
3.2 Cell-Cycle Synchronization, Kinase Inhibition, and Immunoblotting
3.2.1 Maintaining Cell Lines
3.2.2 Mitotic Cell Synchronization
3.2.3 Sample Collection for the Duration of Cell Cycle
3.2.4 Kinase Inhibitor Treatment in Mitotic Cells and Sample Preparation
3.2.5 Cell Lysis and Sample Preparation
3.2.6 SDS-PAGE and Western Blotting
3.2.7 Immunoblotting with Phospho-Specific Antibodies
4 Notes
References
Chapter 16: Visualization of Radiation-Induced Cell Cycle Kinetics with a Fluorescent Ubiquitination-Based Cell Cycle Indicato...
1 Introduction
2 Materials
2.1 Plasmids
2.2 Cell Lines
2.3 Reagents
2.4 Equipment
3 Methods
3.1 Fucci Lentivirus Production
3.2 Establishment of Cells Expressing Fucci (Fig. 2a)
3.3 Time-Lapse Imaging in Monolayer Culture After Irradiation
3.4 Time-Lapse Imaging in Spheroids After Irradiation
3.5 Establishment of Xenografted Tumors
3.6 Histological Analysis of Tumors Derived from Cells Expressing Fucci
3.7 (Optional) Optical Imaging of Fluorescence Kinetics After Irradiation (See Note 9)
4 Notes
References
Chapter 17: Dynamic Behavior of Inactive X Chromosome Territory During the Cell Cycle as Revealed by H3K27me3-Specific Intrace...
1 Introduction
2 Materials
2.1 Plasmids
2.2 Cell Line and Medium
2.3 Transfection Reagents and Inhibitor for Selection
2.4 Microscopy and Image Analysis
3 Methods
3.1 Establishing Cell Lines Stably Expressing H3K27me3-NLS-Mintbody and Halo-PCNA
3.2 Image Acquisition
3.3 Data Analysis
4 Notes
References
Chapter 18: Analyzing Centrioles and Cilia by Expansion Microscopy
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Expansion Reagents, Buffers, and Polymerization Mixture
2.3 Expansion Tools and Equipment
2.4 Immunofluorescent Labeling
3 Methods
3.1 Sample Preparation and Fixation
3.2 Incubation with Acrylamide and Formaldehyde
3.3 Polymerization
3.4 Cutting (Punching) the Sample into Multiple Smaller Samples
3.5 Denaturation by Boiling in SDS Buffer
3.6 SDS Washout
3.7 Immunolabeling
3.8 Expansion of Immunolabeled Samples in dH2O
3.9 Sample Mounting and Imaging
4 Notes
References
Chapter 19: Analysis of Cell Cycle Progression in the Budding Yeast S. cerevisiae
1 Introduction
2 Materials
2.1 Media and Chemicals
2.2 Equipment
2.3 Consumables
3 Methods
3.1 Cell Synchronization Methods in Budding Yeast
3.1.1 Pheromone-Induced Arrest of MATa Cells
3.1.2 Pheromone-Induced Arrest of MATα Cells
3.1.3 Mitotic Arrest by Nocodazole Treatment
3.2 Analysis of Cell Cycle Progression in Time Course Experiments
3.2.1 Sample Collection
3.2.2 Flow Cytometry: Analysis of DNA Content
3.2.3 Budding Index as a Marker of Cell Cycle Progression
3.2.4 Sample Preparation for Western Blotting and Detection of the Cell Cycle Marker Proteins
3.2.5 Fluorescence Microscopy to Determine Cell Cycle Stage by Spindle Staining
4 Notes
References
Chapter 20: Application of PALM Superresolution Microscopy to the Analysis of Microtubule-Organizing Centers (MTOCs) in Asperg...
1 Introduction
2 Materials
2.1 Preparing Hyphae for Live Cell Imaging
2.2 General Microscopy Equipment for Live Cell Imaging
2.3 PALM Set-Up
3 Methods
3.1 Live Cell Imaging of MTOC Components in A. nidulans
3.2 PALM Experiments
4 Notes
References
Chapter 21: Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci Too...
1 Introduction
2 Materials
2.1 Cell Culture
2.2 Confocal Microscopy
2.3 Image Analysis
3 Methods
3.1 Generation of Stable Arl13bCerulean-Fucci2a Expressing Cell Lines
3.2 Live Cell Imaging of Arl13bCerulean-Fucci2a Expressing Cells
3.3 Experimental Setup
3.4 Imaging Setup
3.5 Image Analysis with Fiji and Fucci Tools
3.6 Installation of Fucci_Tools.ijm and Initial Definition of an Experimental Profile
3.7 Performing an Analysis with Fucci Tools
3.8 Plotting of the Results Table Data
3.9 Concluding Remarks
4 Notes
References
Chapter 22: Calorimetric Heat Dissipation Measurements of Developing Zebrafish Embryos
1 Introduction
2 Materials
2.1 Fish
2.2 Solutions
2.3 Calorimeter and Accessories
2.4 Software
3 Methods
3.1 Set-Up and Preparation
3.2 Workflow on a Day-to-Day Basis
3.3 Egg Collection
3.4 Calorimeter Cleaning
3.5 Set Up of the Reference and Experimental Cell
3.6 Software Settings
3.7 Calorimeter Equilibration, Initial Baseline Recording, Embryo Injection and Heat Dissipation Measurements
3.8 Raw Data Retrieval and Initial Data Curation
3.9 Data Analysis
3.10 Troubleshooting
4 Notes
References
Chapter 23: The Conditional Knockout Analogous System: CRISPR-Mediated Knockout Together with Inducible Degron and Transcripti...
1 Introduction
2 Materials
2.1 Stock Solutions and Reagents
2.2 Cell Culture
2.3 Equipment
3 Methods
3.1 CRISPR and the Conditional-Off Expression Construct
3.2 Generation of the Conditional Inactivation Cell Line
3.3 Assessment of Degron-Degradation Kinetics
3.4 Genomic DNA Sequencing of CRISPR Targeting Region
4 Notes
References
Correction to: Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis
Correction to: Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci
Index

Citation preview

Methods in Molecular Biology 2329

Amanda S. Coutts Louise Weston Editors

Cell Cycle Oscillators Methods and Protocols Second Edition

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-bystep 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.

Cell Cycle Oscillators Methods and Protocols Second Edition

Edited by

Amanda S. Coutts School of Science and Technology, Nottingham Trent University, Nottingham, UK

Louise Weston Scientific Writer, Newcastle upon Tyne, UK

Editors Amanda S. Coutts School of Science and Technology Nottingham Trent University Nottingham, UK

Louise Weston Scientific Writer Newcastle upon Tyne, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-0716-1537-9 ISBN 978-1-0716-1538-6 (eBook) https://doi.org/10.1007/978-1-0716-1538-6 © Springer Science+Business Media, LLC, part of Springer Nature 2021, Corrected Publication 2021 Chapters 19 and 21 are licensed under the terms of the Creative Commons Attribution 4.0 International License (http:// creativecommons.org/licenses/by/4.0/). For further details see license information in the chapters. Methods in Molecular Biology 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. Cover Illustration Caption: U2OS cell in the process of division, in telophase. The cell was expanded ~4 fold, labeled, and imaged using a conventional wide field microscope. Two groups of chromosomes are labeled with DAPI (magenta). Immunolabeling for acetylated tubulin (green) reveals acetylated microtubules and two pairs of centrioles. Image credit: Dong Kong and Jadranka Loncarek, NIH/NCI/CCR. 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 series of highly ordered and regulated events that take place in a cell leading it to divide into two daughter cells are collectively known as the cell cycle, a process that in addition to driving both reproduction and the development of living systems also facilitates proliferative diseases and cancer. Oscillatory networks, which are inextricably linked, underlie each cycle of cell division. Understanding the dynamic interaction of small molecules, genes, and proteins that facilitate such a sophisticated biological process remains a challenging scientific problem. In this new edition of Cell Cycle Oscillators: Methods and Protocols, we bring together a range of expert researchers who discuss recent progress in the field from both holistic and reductionist perspectives. The edition provides a space for researchers to highlight and explore the latest developments in molecular biology and biochemical techniques for studying oscillatory networks and to share these across the research community to facilitate further progress. Written in the highly successful Methods in Molecular Biology series format, chapters include introductions to their respective topics, lists of the necessary materials and reagents, step-by-step, readily reproducible laboratory protocols, and key tips on troubleshooting and avoiding known pitfalls. This book aims to bring together a unique collection of protocols that cover novel and specialized techniques as well as updated and improved adaptations of more standard procedures. Because of this range, the protocols will be useful for those new to the field as well as the more experienced scientist. Importantly, we hope these techniques will be used to gain further insight into the complex and incompletely understood processes that are involved in the cell cycle and its regulation by oscillatory networks. Lastly, we would also like to thank all the authors for their excellent contributions, John Walker for his expert advice and assistance, and Springer Nature for all their efforts. Nottingham, UK Newcastle upon Tyne, UK

Amanda S. Coutts Louise Weston

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

v ix

1 Cell Cycle Control: A System of Interlinking Oscillators . . . . . . . . . . . . . . . . . . . . . 1 Randy Y. C. Poon 2 Substrate Phosphorylation Rates as an In Vivo Measurement of Kinase Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 Matthew P. Swaffer 3 Real-Time Monitoring of APC/C-Mediated Substrate Degradation Using Xenopus laevis Egg Extracts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 Julia Kamenz, Renping Qiao, Qiong Yang, and James E. Ferrell Jr 4 Fluorescent Peptide Biosensors for Probing CDK Kinase Activity in Cell Extracts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 Morgan Pellerano and May C. Morris 5 Phosphatase and Kinase Substrate Specificity Profiling with Pooled Synthetic Peptides and Mass Spectrometry. . . . . . . . . . . . . . . . . . . . . . 51 Andrew G. DeMarco, Pete E. Pascuzzi, W. Andy Tao, and Mark C. Hall 6 Whole-Mount Immunostaining for the Identification of Histone Modifications in the S-Phase Nuclei of Arabidopsis Roots . . . . . . . . . . . . . . . . . . . . 71 Hirotomo Takatsuka and Masaaki Umeda 7 Construction of a Full-Length 30 UTR Reporter System for Identification of Cell-Cycle Regulating MicroRNAs. . . . . . . . . . . . . . . . . . . . . . 81 Dominika Kaz´mierczak and Per Hydbring 8 Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Mardo Ko˜ivom€ a gi 9 Optimizing Cell Synchronization Using Nocodazole or Double Thymidine Block. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Arif A. Surani, Sergio L. Colombo, George Barlow, Gemma A. Foulds, and Cristina Montiel-Duarte 10 Highly Synchronous Mitotic Progression in Schizosaccharomyces pombe Upon Relief of Transient Cdc2-asM17 Inhibition. . . . . . . . . . . . . . . . . . . . . 123 Pawan Singh, Lenka Halova, and Iain Michael Hagan 11 Elucidating Human Mitosis Using an Anaphase-Like Cell-Free System. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 Danit Wasserman, Sapir Nachum, Meirav Noach-Hirsh, Naomi Auerbach, Evelin Sheinberger-Chorni, Taylor P. Enrico, Roxane Lahmi, Michael J. Emanuele, and Amit Tzur

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Contents

EDU (5-Ethynyl-20 -Deoxyuridine)-Coupled Fluorescence-Intensity Analysis: Determining Absolute Parameters of the Cell Cycle . . . . . . . . . . . . . . . . ˜ o A. Ferreira, Marco Neves, Miguel Alpalha ˜ o, Pedro Pereira, Joa Daniela Cunha, Fernando Ferreira, Rene´ Santus, Ana E. Sousa, and Paulo L. Filipe 13 High-Resolution Analysis of Centrosome Behavior During Mitosis . . . . . . . . . . . Vanessa Nunes, Margarida Dantas, Joana T. Lima, and Jorge G. Ferreira 14 Assaying Cell Cycle Progression via Flow Cytometry in CRISPR/Cas9-Treated Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan M. Geisinger and Tim Stearns 15 Use of Mitotic Protein Kinase Inhibitors and Phospho-Specific Antibodies to Monitor Protein Phosphorylation During the Cell Cycle. . . . . . . . Isha Nasa, Greg B. Moorhead, and Arminja N. Kettenbach 16 Visualization of Radiation-Induced Cell Cycle Kinetics with a Fluorescent Ubiquitination-Based Cell Cycle Indicator (Fucci) . . . . . . . . . Atsushi Kaida and Masahiko Miura 17 Dynamic Behavior of Inactive X Chromosome Territory During the Cell Cycle as Revealed by H3K27me3-Specific Intracellular Antibody . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuko Sato and Hiroshi Kimura 18 Analyzing Centrioles and Cilia by Expansion Microscopy . . . . . . . . . . . . . . . . . . . . Dong Kong and Jadranka Loncarek 19 Analysis of Cell Cycle Progression in the Budding Yeast S. cerevisiae . . . . . . . . . . Deniz Pirincci Ercan and Frank Uhlmann 20 Application of PALM Superresolution Microscopy to the Analysis of Microtubule-Organizing Centers (MTOCs) in Aspergillus nidulans . . . . . . . . Xiaolei Gao, Reinhard Fischer, and Norio Takeshita 21 Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci Tools. . . . . . . . . . . . . . . . . Melinda Van Kerckvoorde, Matthew J. Ford, Patricia L. Yeyati, Pleasantine Mill, and Richard L. Mort 22 Calorimetric Heat Dissipation Measurements of Developing Zebrafish Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan Rodenfels and Karla M. Neugebauer 23 The Conditional Knockout Analogous System: CRISPR-Mediated Knockout Together with Inducible Degron and Transcription-Controlled Expression. . . . . . . . . . . . . . . . . . . . . . . . . . . Hoi Tang Ma Correction to: Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Correction to: Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci Tools . . . . . . . . . . . . . . . . . . . . . 12

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

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C1 C3 337

Contributors MIGUEL ALPALHA˜O • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal NAOMI AUERBACH • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel GEORGE BARLOW • The John van Geest Cancer Research Centre, School of Science and Technology, Nottingham Trent University, Nottingham, UK SERGIO L. COLOMBO • Centre for Diabetes, Chronic Diseases and Ageing, School of Science and Technology, Nottingham Trent University, Nottingham, UK DANIELA CUNHA • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal MARGARIDA DANTAS • Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de (i3S), Porto, Portugal; BiotechHealth PhD Program, Instituto de Cieˆncias Biome´dicas Abel Salazar (ICBAS), Porto, Portugal ANDREW G. DEMARCO • Department of Biochemistry, Purdue University, West Lafayette, IN, USA MICHAEL J. EMANUELE • Lineberger Comprehensive Cancer Center, Department of Pharmacology, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA TAYLOR P. ENRICO • Lineberger Comprehensive Cancer Center, Department of Pharmacology, The University of North Carolina at Chapel Hill, Chapel Hill, NC, USA FERNANDO FERREIRA • CIISA—Centro de Investigac¸a˜o Interdisciplinar em Sanidade Animal, Faculdade de Medicina Veterina´ria, Universidade de Lisboa, Lisbon, Portugal ˜ JOAO A. FERREIRA • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal JORGE G. FERREIRA • Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de (i3S), Porto, Portugal; Departamento de Biomedicina, Faculdade de Medicina do Porto, Porto, Portugal JAMES E. FERRELL JR • Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, CA, USA; Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, USA PAULO L. FILIPE • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal REINHARD FISCHER • Department of Microbiology, Institute for Applied Biosciences, Karlsruhe Institute of Technology (KIT), Karlsruhe, Germany MATTHEW J. FORD • Goodman Cancer Research Centre, Department of Human Genetics, McGill University, Montreal, QC, Canada GEMMA A. FOULDS • The John van Geest Cancer Research Centre, School of Science and Technology, Nottingham Trent University, Nottingham, UK XIAOLEI GAO • Department of Microbiology, Institute for Applied Biosciences, Karlsruhe Institute of Technology (KIT), Karlsruhe, Germany JONATHAN M. GEISINGER • Department of Biology, Stanford University, Stanford, CA, USA; Department of Genetics, Stanford University, Stanford, CA, USA IAIN MICHAEL HAGAN • Cell Division Group, CRUK Manchester Institute, The University of Manchester, Alderley Park, UK

ix

x

Contributors

MARK C. HALL • Department of Biochemistry, Purdue University, West Lafayette, IN, USA; Center for Cancer Research, Purdue University, West Lafayette, IN, USA LENKA HALOVA • Cell Division Group, CRUK Manchester Institute, The University of Manchester, Alderley Park, UK PER HYDBRING • Department of Oncology and Pathology, Karolinska Institutet, Stockholm, Sweden ATSUSHI KAIDA • Division of Oral Health Sciences, Department of Oral Radiation Oncology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan JULIA KAMENZ • Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, CA, USA; Molecular Systems Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands DOMINIKA KAZ´MIERCZAK • Department of Oncology and Pathology, Karolinska Institutet, Stockholm, Sweden ARMINJA N. KETTENBACH • Department of Biochemistry and Cell Biology, Geisel School of Medicine at Dartmouth, Hanover, NH, USA; Norris Cotton Cancer Center, Geisel School of Medicine at Dartmouth, Lebanon, NH, USA HIROSHI KIMURA • Cell Biology Center, Institute of Innovative Research, Tokyo Institute of Technology, Yokohama, Japan MARDO KO˜IVOMA€ GI • Department of Biology, Stanford University, Stanford, CA, USA DONG KONG • Laboratory of Protein Dynamics and Signaling, NIH/NCI/CCR, Frederick, MD, USA ROXANE LAHMI • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel JOANA T. LIMA • Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de (i3S), Porto, Portugal JADRANKA LONCAREK • Laboratory of Protein Dynamics and Signaling, NIH/NCI/CCR, Frederick, MD, USA HOI TANG MA • Division of Life Science, The Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong PLEASANTINE MILL • MRC Human Genetics Unit, MRC Institute of Genetics & Molecular Medicine, Western General Hospital, University of Edinburgh, Edinburgh, UK MASAHIKO MIURA • Division of Oral Health Sciences, Department of Oral Radiation Oncology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Tokyo, Japan CRISTINA MONTIEL-DUARTE • The John van Geest Cancer Research Centre, School of Science and Technology, Nottingham Trent University, Nottingham, UK GREG B. MOORHEAD • Department of Biological Sciences, University of Calgary, Calgary, AB, Canada MAY C. MORRIS • Institut des Biomole´cules Max Mousseron, CNRS, UMR 5247, Faculte´ de Pharmacie, Universite´ de Montpellier, Montpellier, France RICHARD L. MORT • Division of Biomedical and Life Sciences, Faculty of Health and Medicine, Lancaster University, Lancaster, UK SAPIR NACHUM • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel ISHA NASA • Department of Biochemistry and Cell Biology, Geisel School of Medicine at Dartmouth, Hanover, NH, USA; Norris Cotton Cancer Center, Geisel School of Medicine at Dartmouth, Lebanon, NH, USA

Contributors

xi

KARLA M. NEUGEBAUER • Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA MARCO NEVES • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal MEIRAV NOACH-HIRSH • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel VANESSA NUNES • Instituto de Investigac¸a˜o e Inovac¸a˜o em Sau´de (i3S), Porto, Portugal; BiotechHealth PhD Program, Instituto de Cieˆncias Biome´dicas Abel Salazar (ICBAS), Porto, Portugal PETE E. PASCUZZI • Department of Biochemistry, Purdue University, West Lafayette, IN, USA; School of Information Studies, Purdue University, West Lafayette, IN, USA; Center for Cancer Research, Purdue University, West Lafayette, IN, USA MORGAN PELLERANO • Institut des Biomole´cules Max Mousseron, CNRS, UMR 5247, Faculte´ de Pharmacie, Universite´ de Montpellier, Montpellier, France PEDRO PEREIRA • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal DENIZ PIRINCCI ERCAN • Chromosome Segregation Laboratory, The Francis Crick Institute, London, UK RANDY Y. C. POON • Division of Life Science, The Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong; State Key Laboratory of Molecular Neuroscience and Center for Cancer Research, Hong Kong University of Science and Technology, Clear Water Bay, Hong Kong RENPING QIAO • Research Institute of Molecular Pathology (IMP), Vienna Biocenter (VBC), Vienna, Austria JONATHAN RODENFELS • Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA; Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany RENE´ SANTUS • De´partement RDDM, Muse´um National d’Histoire Naturelle, Paris, France YUKO SATO • Cell Biology Center, Institute of Innovative Research, Tokyo Institute of Technology, Yokohama, Japan EVELIN SHEINBERGER-CHORNI • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel PAWAN SINGH • Cell Division Group, CRUK Manchester Institute, The University of Manchester, Alderley Park, UK ANA E. SOUSA • Instituto de Medicina Molecular-Joa˜o Lobo Antunes, Faculdade Medicina da Universidade de Lisboa, Lisbon, Portugal TIM STEARNS • Department of Biology, Stanford University, Stanford, CA, USA; Department of Genetics, Stanford University, Stanford, CA, USA ARIF A. SURANI • The John van Geest Cancer Research Centre, School of Science and Technology, Nottingham Trent University, Nottingham, UK MATTHEW P. SWAFFER • Department of Biology, Stanford University, Stanford, CA, USA HIROTOMO TAKATSUKA • Graduate School of Science and Technology, Nara Institute of Science and Technology, Ikoma, Nara, Japan; School of Biological Science and Technology, School of Science and Technology, Kanazawa University, Kanazawa, Ishikawa, Japan NORIO TAKESHITA • Microbiology Research Center for Sustainability (MiCS), Faculty of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Japan

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Contributors

W. ANDY TAO • Department of Biochemistry, Purdue University, West Lafayette, IN, USA; Center for Cancer Research, Purdue University, West Lafayette, IN, USA; Department of Chemistry, Purdue University, West Lafayette, IN, USA AMIT TZUR • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel FRANK UHLMANN • Chromosome Segregation Laboratory, The Francis Crick Institute, London, UK MASAAKI UMEDA • Graduate School of Science and Technology, Nara Institute of Science and Technology, Ikoma, Nara, Japan MELINDA VAN KERCKVOORDE • Division of Biomedical and Life Sciences, Faculty of Health and Medicine, Lancaster University, Lancaster, UK DANIT WASSERMAN • Faculty of Life Sciences and Institute of Nanotechnology and Advanced Materials, Bar-llan University, Ramat-Gan, Israel QIONG YANG • Department of Biophysics, University of Michigan, Ann Arbor, MI, USA PATRICIA L. YEYATI • MRC Human Genetics Unit, MRC Institute of Genetics & Molecular Medicine, Western General Hospital, University of Edinburgh, Edinburgh, UK

Chapter 1 Cell Cycle Control: A System of Interlinking Oscillators Randy Y. C. Poon Abstract The cell cycle is the sequence of events through which a cell duplicates its genome, grows, and divides. Key cell cycle transitions are driven by oscillators comprising of protein kinases and their regulators. Different cell cycle oscillators are inextricably linked to ensure orderly activation of oscillators. A recurring theme in their regulation is the abundance of autoamplifying loops that ensure switch-like and unidirectional cell cycle transitions. The periodicity of many cell cycle oscillators is choreographed by inherent mechanisms that promote automatic inactivation, often involving dephosphorylation and ubiquitin-mediated protein degradation. These inhibitory signals are subsequently suppressed to enable the next cell cycle to occur. Although the activation and inactivation of cell cycle oscillators are in essence autonomous during the unperturbed cell cycle, a number of checkpoint mechanisms are able to halt the cell cycle until preconditions or defects are addressed. Together, these mechanisms orchestrate orderly progression of the cell cycle to produce more cells and to safeguard genome stability. Key words Anaphase-promoting complex, Cell cycle, Cell division, Cell growth, Checkpoints, Cyclin-dependent kinases, Cyclin, DNA replication, Mitosis, Phosphorylation, pRb, Proteolysis, Ubiquitin-mediated degradation

1

Introduction The cell cycle is the sequence of events through which a cell duplicates its genome, grows, and divides into two daughter cells. The cell cycle is divided into four phases (Fig. 1). After cell division, daughter cells undergo a period of growth (G1) when cellular macromolecules including proteins, RNA, and membranes are synthesized. G1 is followed by a period of DNA synthesis (S). After another period of growth (G2), cells undergo mitosis (M), during which the DNA is divided equally into two daughter cells. Most nondividing cells exit the cell cycle at G1 into quiescence (G0). Progress in the past several decades has revealed that the eukaryotic cell cycle is driven by an evolutionarily conserved engine composed of a family of protein kinases called cyclin-dependent kinases (CDKs). Although the orderly progression of the cell cycle

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Randy Y. C. Poon CDK1 Cyclin B CDK1

M

Cyclin A

G0

G2

CDK4/6 Cyclin D

G1 R

S

CDK2 Cyclin E

CDK2 Cyclin A

Fig. 1 The cell cycle and cyclin–CDK complexes. The cell cycle is divided into four phases: Gap 1 (G1), DNA synthesis (S), Gap 2 (G2), and mitosis (M). Most nondividing mammalian cells exit the cell cycle at G1 into a quiescent state (G0). After passing the restriction point (R), a cell is committed to another round of cell cycle and becomes independent of proliferation stimulants. The cyclin–CDK complexes involved in different periods of the cell cycle are shown (isoforms of the different cyclins are not shown)

depends on a number of factors, a good approximation is that different stages of the cell cycle are promoted by the sequential switching on and off of different CDKs (Fig. 1). Accordingly, the activities of CDKs are stringently regulated by mechanisms including protein–protein interaction, phosphorylation, and ubiquitinmediated proteolysis. This introduction summarizes the fundamental concepts of cell cycle oscillators. Although the basic mechanisms of cell cycle control are conserved in all eukaryotic cells, details such as the complexity of protein families involved do vary between organisms and between embryonic and somatic cells. Here the emphasis is placed on the somatic cell cycle of mammalian cells. 1.1 Anatomy of a Cell Cycle Oscillator

The cell cycle is steered by successive waves of cell cycle oscillators. Myriad mechanisms have developed to ensure that cell cycle regulators are turned on and off at the correct time and in proper order. These oscillators are characterized by several features, including (a) an activating mechanism; (b) an autoamplifying loop to ensure switch-like cell cycle transitions; an additional kickstarting mechanism may also be involved; (c) an autoinactivating mechanism that automatically turns off the oscillator; (d) a mechanism to prevent the reactivation of the oscillator during the same cell cycle, and a way to remove this inhibitory signal during the next cell cycle;

Cell Cycle Oscillators Auto-amplifying loop

Activator

Next cell cycle Inhibitor

Activity

Kick-starter

3

Time

Fig. 2 Cell cycle oscillators. Due to the periodic nature of the cell cycle, an activating mechanism of a cell cycle oscillator is followed by an inactivating one. The latter then has to be suppressed to enable a subsequent cell cycle. Several features are frequently found in cell cycle oscillators, including (a) an activating mechanism; (b) an autoamplifying machinery and a kickstarting mechanism; (c) an autoinactivating mechanism; (d) means to prevent the reactivation of the oscillator during the same cell cycle, and ways to remove these inhibitory signals during the next cell cycle; as well as (e) a stimulator of the next oscillator in the cell cycle (blue-grey)

and (e) a stimulator of the next oscillator in the cell cycle (Fig. 2). Not all of these features are present in every cell cycle oscillator. Emphasis is placed in the subsequent sections to identify these components in each cell cycle oscillator. 1.2 Checkpoints: Putting a Break on Cell Cycle Oscillators

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Once passed the restriction point, the cell cycle can be viewed as a succession of autonomous oscillators (see Subheading 2). However, the free running of the cell cycle engine is restrained by surveillance mechanisms termed checkpoints. By temporarily halting the cell cycle, checkpoints ensure that each stage of the cell cycle is completed before the next stage is initiated. In general, checkpoints include a sensor that monitors cell cycle defects, a transducer that transmits and amplifies the signal, and an effector that stops the cell cycle. Several major checkpoints, including those that monitor proper spindle assembly, completion of DNA replication, and DNA damage will be discussed here.

Entering the Cell Cycle and G1–S Whether a cell stays in the cell cycle depends on the integration of extracellular signals from cell surface receptors responding to mitogenic growth factors and growth inhibitory factors. This decision is made at a transition in G1 called the restriction point (R). Cells exit the cell cycle into G0 if insufficient mitogenic signals are present to overcome the restriction point. After passing the restriction point, a

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Randy Y. C. Poon Mitogenic cues

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pRb E2F

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CDK4/6 Cyclin D

G1

CDK2

p27

Cyclin E

S

CDK2

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Fig. 3 Regulation of G1–S. Both the transcription and stability of cyclin D increase when quiescent cells are stimulated to enter the cell cycle by extracellular growth signals. Hyperphosphorylation of pRb by cyclin D–CDK4/6 and other cyclin–CDK pairs releases pRb from E2F, allowing E2F to activate transcription. Increased E2F-dependent transcription enables cells to pass through the restriction point (R). E2F increases the expression of cyclin E and cyclin A (among other genes), which activate CDK2 and further boost the phosphorylation of pRb in a positive feedback loop. Sequestering of p21CIP1/ WAF1 and p27KIP1 by cyclin D–CDK4/6 further increases the activity of cyclin E–CDK2. Cyclin E–CDK2 also reduces p27KIP1 by targeting it to SCFSKP2dependent degradation. Resetting pRb to a hypophosphorylated state is carried out by phosphatases at the end of mitosis

cell is committed to another round of the cell cycle and becomes independent of external stimuli. Mechanistically, the restriction point involves phosphorylation of the retinoblastoma gene product pRb by G1 cyclin–CDK complexes (Fig. 3). After DNA damage, the G1–S cell cycle engine is suppressed by the G1 DNA damage checkpoint. 2.1 Cyclin D as a Mitogenic Sensor for the Cell Cycle

Transcription of D-type cyclins (D1, D2, and D3) increases when quiescent cells are stimulated with growth factors. The strong dependence of cyclin D expression on extracellular mitogenic cues, coupling to the relative short half-life of the protein (~30 min), enables cyclin D to act as an effective mitogenic sensor that conveys extracellular signals to the cell cycle [1]. The promoters of D-type cyclins are under the control of multiple cell surface receptors and signaling pathways [2]. For example, activation of the RAS—RAF—MEK—ERK signaling cascade, either in response to soluble growth factors binding to cell surface tyrosine kinase receptors or extracellular matrix (ECM) binding to integrins, activates the transcription of cyclin D1. This is mediated by the downstream transcription factors AP-1

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(including members of the FOS, JUN, and ATF families) of the RAS signaling pathway. In addition, RAS also activates AKT/PKB through phosphoinositide 3-kinase (PI3K). AKT/PKB then phosphorylates and inactivates GSK-3β, thereby preventing β-catenin from degradation; the accumulated β-catenin then recruits the TCF/LEF family of transcription factors to activate cyclin D1 transcription. In this connection, activation of β-catenin by the canonical WNT signaling pathway also increases the transcription of cyclin D1. As degradation of cyclin D1 involves phosphorylation by GSK-3β (at residue threonine (Thr) 286, generating a phosphodegron that is recognized by the ubiquitin ligase SCFFBX4), inhibition of GSK-3β by AKT/PKB also has an additional effect of stabilizing cyclin D1 protein [3]. 2.2 Phosphorylation of pRb Underlies the Restriction Point

Once cyclin D is synthesized, it binds and activates two cyclindependent kinases, CDK4 and CDK6. The cyclin D–CDK4/6 complexes then phosphorylate pRb (and the related p107 and p130) [4]. One of the key functions of pRb (and related proteins) before it is phosphorylated by cyclin D–CDK4/6 (hypophosphorylated form) is sequestering of E2F. Several members of the E2F family (E2F1–3) bind DP proteins (DP1 or DP2), forming transcription factors critical for transcribing genes important for entry into S phase [5]. Hypophosphorylated pRb inhibits E2F by both blocking the transactivating domain as well as recruiting other proteins to repress E2F-mediated transcription. One mechanism involves the association of pRb with chromatin remodeling enzymes including histone deacetylase (HDAC), thereby indirectly targeting HDAC to the promoters bound by E2F–DP [4]. This represses the transactivation of the promoter through chromatin remodeling. Phosphorylation of pRb by cyclin D–CDK4/6 releases pRb from E2F (removing HDAC at the same time), liberating E2F–DP complexes to activate transcription. The classic model is that hyperphosphorylation of pRb is initiated by cyclin D–CDK4/6, but is then maintained by cyclin E–CDK2 and cyclin A–CDK2. Unlike that of cyclin D, the expression of cyclin E and cyclin A is independent of extracellular signals. A large number of genes, many of which are needed for S phase, are transcriptionally activated by E2F–DP complexes. Among these are cyclin E and cyclin A, which activate CDK2 and further increase the phosphorylation of pRb. The pRb– E2F pathway therefore functions as a switch to convert graded growth factor stimulations into an all-or-none E2F response. Several members of the E2F family including E2F4 and E2F5 are transcriptional repressors. During G0, E2F4 and E2F5 repress E2F-responsive genes that promote entry into G1. Following mitogenic stimulation, phosphorylation of pRb by cyclin–CDK complexes results in the release of E2F repressors and the accumulation of newly synthesized E2F activators (E2F1–3) [5].

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Negative regulators of the G1 cyclin–CDK complexes including CDK inhibitors can modulate the threshold of the restriction point. Binding of cell surface receptors by TGF-β stimulates a signaling pathway involving SMAD proteins, eventually leading to the synthesis of the CDK inhibitor p15INK4B [6]. Members of the INK4 family of CDK inhibitors (p16INK4A, p15INK4B, p18INK4C, and p19INK4D) are specific for CDK4 and CDK6. Unlike the p21CIP1/ WAF1 family of CDK inhibitors (p21CIP1/WAF1, p27KIP1, and p57KIP2), the INK4 proteins act by binding to monomeric CDK4/6 and blocking the formation of complexes with cyclin D. They also have an additional effect of displacing the p21CIP1/ WAF1 /p27KIP1 that normally associates with cyclin D-CDK4/6 (see below) to redistribute to other cyclin-CDK complexes. The protein of p27KIP1 is further stabilized by TGF-β signaling through destruction of SKP2 [7]. The levels of some of the CDK inhibitors are modulated during the cell cycle. For example, p27KIP1 is degraded by the ubiquitin ligase SCFSKP2 complex. SKP2 itself is destroyed by APC/CCDH1 during G1 [8]. The accumulation of p27KIP1 conferred by APC/ CCDH1 therefore contributes to the inhibition of CDK2 activity during G1. When cyclin D accumulates during G1, it drags p21CIP1/ WAF1 and p27KIP1 away from cyclin E–CDK2 complexes, thereby liberating cyclin E–CDK2 from the CDK inhibitors. Intriguingly, the kinase activity of cyclin D–CDK4/6 is actually unaffected by p21CIP1/WAF1 and p27KIP1. In fact, p21CIP1/WAF1 and p27KIP1 play a role in stimulating the formation of cyclin D–CDK4/6 complexes. After cyclin E–CDK2 complexes are turned on, they phosphorylate p27KIP1 and stimulate SCFSKP2-dependent degradation of p27KIP1. This in turn allows more cyclin E–CDK2 to be activated to promote G1–S. As described above, signaling by RAS activates AKT/PKB. AKT/PKB also phosphorylates p21CIP1/WAF1 and p27KIP1 and blocks their nuclear accumulation, thereby preventing these CDK inhibitors from acting on cyclin E–CDK2 complexes [9]. More recent evidence increases the complexity of the classic model of the control of restriction point by pRb described above [10]. Cyclin D–CDK4/6 appears to be able to only monophosphorylate pRb, on one of the 14 CDK sites, which still allows pRb to inactivate E2F. Additional phosphorylation events by cyclin E– CDK2 are required to hyperphosphorylate pRb and release E2F. Phosphorylation of pRb is reset to the hypophosphorylated state by the phosphatase PP1 at the end of mitosis, at a time when the levels of cyclin D, cyclin E, and cyclin A are at their lowest during the cell cycle [11]. Consequently, the overcoming of the restriction point becomes once again dependent on extracellular cues and the accumulation of cyclin D. Another interesting recent discovery is that for cycling cells, molecular events during the

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previous cell cycle (such as the RAS activity) may influence the passage through the restriction point [10]. This suggests that the pRb pathway may not be completely reset at the beginning of each new cell cycle as once thought. 2.3 The G1 DNA Damage Checkpoint

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DNA damage occurring during G1 phase activates a checkpoint that pauses the cell cycle to allow time for DNA repair. The molecular mechanism underlying this checkpoint comprises of a p53-dependent mechanism that feeds into the pRb pathway [12]. In the absence of DNA damage, p53 is suppressed by one of its own transcriptional targets called MDM2 in a negative feedback loop. MDM2 binds to the N-terminal transactivation domain of p53 and inhibits p53-mediated transcription, shuttles p53 out of the nucleus, and promotes ubiquitin-dependent degradation of p53. The last effect is due to the fact that MDM2 is itself a ubiquitin ligase. DNA damage activates sensors that facilitate the activation of the PI3K-related protein kinases ATM and ATR. They in turn activate the checkpoint kinases CHK1 and CHK2. ATM/ATR, CHK1/CHK2, and other DNA damage-activated protein kinases phosphorylate the N-terminal region of p53. Phosphorylation of these sites abolishes the MDM2–p53 interaction, leading to a rise in p53 level and transcriptional activity. One of the transcriptional targets of p53 is the CDK inhibitor p21CIP1/WAF1. The accumulated p21CIP1/WAF1 then binds and inhibits cyclin A/E–CDK2. This diminishes the phosphorylation of pRb, thereby stopping the cell cycle in G1 phase (see Subheading 2.2). Another important control of the p53 pathway comes from the CDKN2A gene, which encodes both the CDK inhibitor p16INK4A and a protein called p14ARF [13]. The inhibition of p53 by MDM2 is interrupted by p14ARF because it sequesters MDM2 to the nucleolus. The CDKN2A gene is generally activated in response to oncogenic stresses. The ensuing increase in p16INK4A and p14ARF reduces cyclin D–CDK4/6 activity and elevates p53 expression, respectively. Both of these events eventually suppress pRb phosphorylation and arrest the cell cycle in G1.

Control of S Phase The key issues concerning the regulation of S phase are (1) how DNA replication occurs only in S phase, and (2) how replication is initiated once and once only per cell cycle (Fig. 4). Centrosome duplication also occurs during S phase and is in part coupled with the mechanisms that govern DNA replication. In addition, the replication checkpoint is responsible for delaying S phase progression and preventing mitosis in the presence of stalled replication forks.

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P

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MCM2-7

GINS

(Geminin)

CDC45

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CDT1 Geminin

ORC

(CDC6)

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Prereplicative complex MCM2-7

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CDT1 CDC6 ORC

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CDC7

Cyclin A/E

DBF4 P

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GINS MCM2-7 CDC45

MCM2-7

CDT1 CDC6

CDT1 CDC6

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ORC

Fig. 4 Regulation of S phase. Initiation of DNA replication occurs at origins of replication. During G1 phase, the origins are licensed by binding to the prereplication complex. During G1–S transition, cyclin A, cyclin E, and DBF4 are transcriptionally activated by E2F. Prereplication complex components including MCM2–7 are phosphorylated by cyclin A/E–CDK2 and DBF4–CDC7, thereby stimulating the recruitment of CDC45 and GINS. This activates the MCM2–7 helicase to unwind the origin. Finally, the unwound DNA allows for the recruitment of DNA polymerases and other components of the DNA synthesis machinery to initiate DNA synthesis. After DNA replication, several mechanisms including degradation of CDC6 and binding of CDT1 to newly synthesized geminin prevent rereplication. These mechanisms are reset later in G1 (including the removal of geminin by APC/C shown in the Figure) 3.1 Initiation of S Phase

Initiation of DNA replication occurs at chromosomal locations known as origins of replication. The budding yeast S. cerevisiae is the only known eukaryote with a defined initiation sequence. Several proteins, including origin recognition complex (ORC, which is composed of ORC1–6), CDC6, and CDT1 are assembled at the origins of replication during G1. This facilitates the loading of double hexamers of the MCM2–7 core helicase, forming the so-called prereplication complex (pre-RC). The formation of pre-RC on origins is called origin licensing [14]. During G1–S transition, the origins are activated by CDK2 and another kinase called CDC7. CDK2 is activated by cyclin A and cyclin E and CDC7 is activated by a protein called DBF4 (ASK in humans) [15]. Higher eukaryotes contain a second DBF4-like protein called DRF1 or ASKL1. Similar to cyclin–CDK pairs, while the level of CDC7 remains relatively constant during the

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cell cycle, the level of DBF4/ASK oscillates during the cell cycle, being absent during G1 and accumulating during S and G2. Similar to cyclin A and cyclin E, transcription of DBF4/ASK1 is activated during late G1 and S phase by E2F. CDK2 and CDC7 phosphorylate components of the pre-RC including the MCM2–7 complex, triggering the recruitment of two helicase coactivators, CDC45 and GINS. The MCM2–7 helicase is then activated and unwinds the origin. The single-stranded DNA is subsequently stabilized by binding to replication protein A (RPA). Finally, the unwound DNA facilitates the recruitment of DNA polymerases and other components of the DNA synthesis machinery to initiate DNA synthesis. Cyclin A/E–CDK2 also coordinates the initiation of DNA replication with the centrosome cycle [16]. The centrosome is located near the nucleus and contains the microtubule organizing center, playing important roles in the establishment of the interphase cytoplasmic microtubule network and bipolar mitotic spindles. Since each daughter cell receives just one centrosome after cell division, the centrosome has to duplicate once before the next mitosis. Centrosome duplication occurs during S phase and is coupled to the cell cycle, at least in part, by the activity of cyclin A/E–CDK2. 3.2 Prevention of Rereplication

Once the genome has been replicated, formation of the pre-RC is inhibited by multiple mechanisms until the next cell cycle [17]. Cyclin E is degraded by the ubiquitin ligase SCFFBW7 after S phase, thereby turning off the cyclin E-CDK2 kinase activity. The phosphodegron recognized by SCFFBW7 is created by CDK2dependent autophosphorylation as well as by GSK-3β. On the other hand, DBF4/ASK1 is degraded by APC/C only after mitosis. The accumulation of CDK activity during late G1, S, and G2 prevents the reassembly of the pre-RC through several mechanisms. Although cyclin E is degraded during S phase, the expression of cyclin A persists till mitosis. CDK-dependent phosphorylation excludes MCM2–7 from the nucleus, targets CDT1 and CDC6 for degradation, and dissociates ORC from chromatin. Furthermore, accumulation of geminin during S and G2 results in the formation of a tight geminin–CDT1 complex, thereby preventing CDT1 from loading onto the pre-RC. Cyclin A–CDK2 also phosphorylates E2F1 and E2F3, decreasing their DNA binding capability and terminating the transcription of genes involve in S phase control [18]. SCFSKP2-dependent degradation of E2F1 during S and G2 further limits the activity of E2F after S phase [19]. Several members of the E2F family including E2F7 and E2F8 are transcriptional repressors. After G1–S, these transcriptional repressor E2Fs attenuate the transcription of genes activated earlier by E2F1–3. They also directly repress the

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expression of transcriptional activator E2Fs such as E2F1 [5]. Together, these mechanisms prevent the expression of E2F-activating genes after G1–S. Assembly of the pre-RC can occur again in early G1 because destruction of cyclin A and cyclin B during mitosis provides an environment of low CDK activity. Proteolysis of geminin by APC/C during mitosis also unleashes CDT1 to form the pre-RC. Hence APC/C resets the mechanisms that safeguard rereplication during the previous cell cycle. 3.3 Replication Checkpoint

4

Stalled replication forks activate a checkpoint involving ATR [20]. Replication fork progression is disrupted by either an insufficient supply of nucleotides or lesions and obstacles on the DNA. Several proteins including ATRIP, Claspin, and TopBP1 are involved in recruiting ATR to single-stranded DNA present at the stalled replication forks. Specifically, ATR is activated by binding to the single-strand binding protein RPA-coated single-stranded DNA. The activated ATR then phosphorylates and activates CHK1. CHK1 subsequently activates WEE1 and inactivates CDC25 (see Subheading 4.2). Consequently, this alters the inhibitory phosphorylation status of CDKs, tipping the balance toward inhibition of the cyclin–CDK complexes involved in both replication (see Subheading 3.1) and mitotic entry (see Subheading 4). Hence the replication checkpoint regulates origin firing, replication forks progression, as well as preventing untimely mitosis. These mechanisms provide the cell with time to restart or repair the stalled replication forks. Significantly, the checkpoint is essential during unperturbed S phase even in the absence of exogenous stresses.

G2–Mitosis Arrays of dramatic events occur during mitosis, including chromosome condensation, nuclear envelope breakdown, formation of mitotic spindles, attachment of chromosomes to the mitotic spindles, and separation of sister chromatids. In essence, mitosis is driven by the activation of CDK1 as well as inactivation of phosphatases that dephosphorylate CDK1 substrates (Fig. 5). Working in concert with a number of kinases and phosphatases, the activation of CDK1 is characterized by feedback mechanisms that ensure CDK1 is activated rapidly.

4.1 Cyclin B–CDK1 as the Engine of Mitosis

The key event for mitotic entry is the activation of CDK1. Although CDK1 is present at a constant level throughout the cell cycle, it is active only during mitosis due to regulation by several mechanisms including binding to cyclins and phosphorylation. The mitotic cyclins (cyclin A and cyclin B) are synthesized and destroyed around the time of mitosis (cyclin A also functions in S

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Separase MAD2

Aurora A BORA

(Securin)

PLK1

APC/C

CDC25

CDK1

WEE1

Cyclin B

CDC20

Separase

CDK1

M

Securin

(Cyclin B)

G0

G2

G1 S

R

Fig. 5 Regulation of mitotic entry and exit. After cyclin B is synthesized and associates with CDK1, the complex is kept in an inactive state by WEE1/MYT1dependent phosphorylation of CDK1 (Thr14 and Tyr15). Dephosphorylation of CDK1 by members of the CDC25 phosphatase family during G2–M transition activates cyclin B–CDK1. Cyclin B–CDK1 activation is autocatalytic because CDK1 activates CDC25 and inactivates WEE1/MYT1. Initial activation of CDC25 is believed to be carried out by PLK1, which in turn is activated by Aurora A and BORA. During early mitosis, APC/CCDC20 is activated by cyclin B–CDK1 and other mitotic kinases. However, its activity is suppressed by the SAC through the binding of MAD2 to APC/CCDC20. Once all kinetochores are properly attached, the SAC is silenced to allow APC/CCDC20 activation, which then targets several proteins including cyclin B, PLK1, Aurora A, and securin to ubiquitin-mediated degradation. Proteolysis of securin releases separase, which in turn cleaves cohesin to allow sister chromatid separation. Reactivation of CDK1 during G1 is safeguarded by activation of another APC/C complex involving CDH1 (not shown)

phase) [21]. Mammalian cells contain two A-type cyclins (A1 and A2) and three B-type cyclins (B1, B2, and B3). While cyclin A2 is present in all proliferating somatic cells, cyclin A1 is critical only during spermatogenesis. Cyclin B1 is the major mitotic cyclin partner of CDK1. Cyclin B2 is coexpressed with cyclin B1 in the majority of dividing cells but is less abundant. The expression of

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cyclin B3 is restricted to developing germ cells and in the adult testis. A salient characteristic of the mitotic cyclins is their periodicity. Cyclin A starts to accumulate during late G1, continues through S phase and G2, before being rapidly destroyed during mitosis. Cyclin B is synthesized and destroyed slightly later than cyclin A. The cell cycle expression of cyclin A and cyclin B is regulated at the levels of transcription and proteolysis [21]. For example, several transcription factors, including B-MYB, FOXM1, MuvB complexes, E2F, and NF-Y, regulate the mRNAs of the cyclin B1 so that they accumulate during G2 and diminish after mitosis [22]. The sharp decrease of the mitotic cyclins at the end of mitosis, however, is caused mainly by proteolysis involving APC/C-dependent mechanisms (see Subheading 5). 4.2 Feedback Control Ensures Biphasic Activation of CDK1

The defining characteristic of CDK1 activation is a system of feedback loops that converts the slow accumulation of cyclin B into an abrupt activation of CDK1. Monomeric CDK1 is inactive and unphosphorylated. On binding to cyclin B, the kinase activity of CDK1 is initially suppressed by inhibitory phosphorylation of CDK1Thr14/Tyr15 by MYT1 and WEE1 [23]. WEE1 is a dualspecificity kinase that phosphorylates tyrosine (Tyr) 15 (but not Thr14). MYT1, a kinase that is normally bound to the endoplasmic reticulum and Golgi complex, can phosphorylate both the Thr14 and Tyr15, but has a stronger preference for Thr14. At the end of G2, the stockpile of inactive cyclin B–CDK1 complexes is rapidly activated by members of the CDC25 phosphatase family (A, B and C) [24]. CDC25B is believed to be the initial activator of cyclin B–CDK1 at the centrosomes. This is followed by the complete activation of cyclin B–CDK1 by CDC25A and CDC25C in the nucleus. Significantly, active CDK1 activates more CDC25 and inactivates WEE1 by directly phosphorylating these proteins. Hence, a small amount of active cyclin B–CDK1 can lead to a rapid and complete activation of all the complexes by this autocatalytic loop. Phosphorylation of WEE1 by CDK1 (as well as by PLK1) also creates a phosphodegron for SCFβTrCP-dependent degradation. Thus, cyclin B–CDK1 is essentially a biphasic switch system that becomes autocatalytic once a critical portion is activated.

4.3 Kickstarting the CDK1 Activating Loops

Given that the activation of cyclin B–CDK1 is autocatalytic, how the initial batch of cyclin B–CDK1 is activated becomes a salient issue. The available data indicate that the multifunctional protein kinase PLK1 may initiate the system by activating CDC25 and inactivating WEE1/MYT1 [25]. PLK1 also promotes the translocation of cyclin B into the nucleus during prophase [26]. During G2, binding of the export mediator CRM1 to the nuclear exporting sequence (NES) of cyclin

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B1 promotes cytoplasmic localization of cyclin B1. Phosphorylation of residues in the NES by kinases including CDK1 and PLK1 is important for the nuclear translocation of cyclin B1, presumably by disrupting the CRM1–cyclin B1 interaction. This mechanism enables the localization of cyclin B1–CDK1 to the nucleus (involving binding of cyclin B1 to importin-β) when the complexes are active. The activation of PLK1 involves phosphorylation by Aurora A, an event that is assisted by a protein called BORA [27]. Cyclin B– CDK1 then phosphorylates BORA to promote PLK1 activation in another feedback loop. PLK1 also phosphorylates BORA, generating a phosphodegron motif that is recognized by the ubiquitin ligase SCFβTrCP to trigger BORA destruction [28]. Aurora A is activated by different mechanisms at centrosomes and spindle microtubules, involving distinct cofactors, localization, and phosphorylation [29]. The spindle microtubule-associated pool of Aurora A is activated allosterically by binding to TPX2 to promote microtubule nucleation. By contrast, the centrosomal pool of Aurora A is activated by binding to CEP192. Interestingly, while activation of Aurora A by CEP192 involves stimulation of autophosphorylation of Aurora A at the activation loop (Thr288), activation of Aurora A by TPX2 is independent on Thr288 phosphorylation. Both Aurora A and PLK1 are also important for various centrosome functions, including centrosome separation, maturation, and mitotic spindle formation. Separation of duplicated and matured centrosomes in late G2 is crucial for the formation of bipolar mitotic spindles. At the end of mitosis, both Aurora A and PLK1 are degraded by APC/CCDH1-mediated ubiquitination. 4.4 Greatwall (MASTL) Helps Maintain the Mitotic State

Phosphorylation events during mitosis are reversible. To ensure that mitotic cells do not reverse to G2, it is important both to maintain the activities of the mitotic kinases as well as to suppress the phosphatases that counteract the kinases’ actions. Greatwall (MASTL in humans) is a kinase that phosphorylates ARPP19 and α-endosulfine (ENSA), promoting their inhibition of the phosphatase PP2A–B55 [30]. As PP2A–B55 is a major phosphatase that dephosphorylates cyclin B–CDK1 substrates, Greatwall activity is important for maintaining the phosphorylation of CDK1 substrates during mitosis [31]. Greatwall also regulates the activation of CDK1 by maintaining the phosphorylation of CDC25, thereby keeping CDK1 in a Thr14/Tyr15-dephosphorylated state [30]. Greatwall itself appears to be activated during mitosis by CDK1 in a feedback loop. At the end of mitosis, Greatwall is reset to an inactive state by PP2A–B55-dependent dephosphorylation of an essential CDK phosphorylation site (Thr194).

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4.5 The G2 DNA Damage Checkpoint

5

The G2 DNA damage checkpoint involves the activation of the protein kinases ATM/ATR followed by CHK1/CHK2 similarly to the G1 DNA damage checkpoint [32]. CHK1/CHK2 then activates WEE1 and inactivates all three isoforms of the CDC25 family (CDC25A, CDC25B, and CDC25C) [33]. CDC25C is inactivated by CHK1/CHK2-dependent phosphorylation both directly and indirectly through the creation of a 14-3-3 binding site. Binding of 14-3-3 to CDC25C masks a proximal nuclear localization sequence, thereby anchoring CDC25C in the cytoplasm and preventing efficient access to cyclin B–CDK1. The centrosomal CDC25B is also phosphorylated by CHK1, creating a docking site for 14-3-3 that disrupts access to CDK1. In contrast to other CDC25 isoforms, CDC25A is targeted for rapid degradation by CHK1/CHK2. CDC25A stability is usually controlled by APC/CCDH1 complexes during early G1 and by SCFβTrCP complexes during interphase. Importantly, the SCFβTrCP-mediated degradation of CDC25A is enhanced after DNA damage through phosphorylation by CHK1. In addition to acting on CDC25, CHK1/CHK2 also appears to phosphorylate and activate WEE1 by promoting 14–3-3 binding. Together, these mechanisms promote CDK1Thr14/Tyr15 phosphorylation, leading to the inactivation of CDK1 and cell cycle arrest in G2.

Mitosis–G1 The key event in mitotic exit is the onset of anaphase, which is driven by APC/C-dependent ubiquitination. Degradation of APC/C substrates including cyclin B and securin promotes several events during mitotic exit, including sister chromatid separation, spindle disassembly, chromosome decondensation, cytokinesis, and reformation of the nuclear envelope. How to keep APC/C inactivate before all the chromosomes are attached to the spindle correctly is the major feedback that orchestrates mitotic exit (Fig. 5).

5.1 APC/C Drives Mitotic Exit

Bipolar spindle formation and proper attachment of chromosomes are highly regulated to ensure that chromosomes are segregated equally to the daughter cells. Several kinases that are targeted to unattached kinetochores, including CDK1, PLK1, and NEK2, phosphorylate key kinetochore proteins such as HEC1 and contribute to the stabilization of microtubule–kinetochore interactions [23]. The chromosomal passenger complex (CPC, composed of aurora B, borealin, INCENP, and survivin) plays a major role in spindle assembly and cytokinesis [34]. CPC localizes to the kinetochores and chromosomes during early mitosis and functions in microtubule–kinetochore interactions, sister chromatid cohesion, and the spindle-assembly checkpoint (SAC). CPC corrects mis-attachments of chromosomes until they are bioriented and

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under tension. CPC is relocated to the central spindle at anaphase and subsequently to the midbody to promote cytokinesis. Once all kinetochores are properly attached, the ubiquitin ligase APC/C is activated to degrade the mitotic cyclins and other proteins [35]. Both A- and B-type cyclins contain a short sequence at the N-terminal region known as the destruction box (D-box). The D-box targets the mitotic cyclins to the multisubunit ubiquitin ligase APC/C. Two targeting subunits called CDC20 and CDH1 are involved in facilitating the ubiquitination of cyclins by APC/C. While CDC20 is present only during mitosis, CDH1 remains constant during the cell cycle, but only associates with APC/C during G1 [36]. The ubiquitinated cyclins are then rapidly degraded by a constitutively active proteasome complex. Importantly, activated cyclin B–CDK1 stimulates the activity of APC/CCDC20 through phosphorylation of several subunits of APC/C and CDC20. APC/C is also phosphorylated and activated by PLK1. Hence, cyclin B primes its own destruction and ensures that APC/CCDC20 is activated only after mitotic entry. However, the activity of APC/CCDC20 is suppressed by the SAC until all the all kinetochores are properly attached (see Subheading 5.2). In addition to cyclin B, APC/CCDC20 also degrades several substrates including securin and geminin [37]. Degradation of securin is important for sister chromatid separation during anaphase. After DNA is replicated, sister chromatids are tethered together by cohesin, a ring-shaped complex consisting of four SMC subunits. This involves a cohesin-interacting protein called sororin, which protects the removal of cohesin by PDS5 and WAPL. Cohesin is removed from chromosomes in a two-step manner during mitosis: while cohesin at the chromosome arms is removed during early mitosis, the centromeric cohesin is protected until anaphase. During prophase, CDK1, PLK1, and Aurora B collaborate to phosphorylate cohesin and sororin, inducing WAPL-dependent removal of cohesin from chromosome arms. However, a pool of cohesin at centromeres is protected by Shugoshin (SGO1). The primary signal for localizing SGO1 to centromeres is BUB1 (a component of the SAC)-dependent phosphorylation of histone H2ASer121. SGO1 interacts with the phosphatase PP2A, thereby keeping cohesin and sororin in a hypophosphorylated state and maintaining centromeric cohesion. During metaphase-anaphase transition, proper kinetochore–microtubule attachment creates tension across sister kinetochores and triggers the removal of SGO1. Kinetochore tension also silences the SAC. This allows the APC/CCDC20 to degrade securin, leading to activation of the protease separase. Separase then cleaves centromeric cohesin to facilitate sister-chromatid separation. Another substrate of APC/CCDC20 is geminin. Degradation of geminin by APC/CCDC20 releases CDT1, a subunit required for the initiation of DNA replication (see Subheading 3). Hence by

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simultaneously destroying the mitotic cyclins, securin, and geminin, APC/CCDC20 coordinates several important processes during the mitosis–G1 transition and prepares the cell for the next S phase. In addition to CDC20, APC/C can also associate with a targeting subunit called CDH1 [37]. In contrast to APC/CCDC20, APC/CCDH1 is turned off during mitosis because phosphorylation by cyclin B–CDK1 alters the conformation of CDH1 and prevents its interaction with APC/C. Destruction of cyclin B at anaphase therefore relieves the inhibition of APC/CCDH1, enabling it to degrade CDC20 and take over the task of degrading any remaining or newly synthesized cyclin B during G1. Finally, APC/CCDH1 is also responsible for destroying several important mitotic regulators including PLK1, CDC25A, Aurora A, and SGO1. During late G1, E2F is released from pRb and activates the transcription of cyclin A (see Subheading 2). Cyclin A–CDK complexes then phosphorylates CDH1 and inhibits its association with APC/C. APC/CCDH1 and APC/CCDC20 are also turned off by binding to EMI1, which begins to accumulate at late G1 (also transcriptionally activated by E2F). EMI1 has to be removed subsequently to allow APC/C to function in mitotic exit. This is achieved by PLK1-dependent phosphorylation, targeting EMI to ubiquitin-mediated degradation by SCFβTrCP. 5.2 The Spindle-Assembly Checkpoint

The spindle-assembly checkpoint (SAC) is activated by either the presence of unattached kinetochores or the absence of tension between paired kinetochores [38]. Consequently, the SAC ensures that chromosomes have achieved correct bipolar attachment to the mitotic spindles before cyclin B and other proteins are degraded by APC/C (Fig. 5). Unattached kinetochores attract several components of the checkpoint sensors (including BUB1, BUBR1, BUB3, CENP-E, MAD1, MAD2, and MPS1), catalyzing the formation of diffusible complexes called mitotic checkpoint complexes (MCC, components include MAD2, BUBR1, and BUB3). These checkpoint components act as signal transducers, resulting in the inhibition APC/CCDC20 through the sequestration of CDC20 by MAD2. Binding to CDC20 requires a conformational change of MAD2 from an open conformation (known as O-MAD2) to the more stable close conformation (C-MAD2). Although the mechanism remains incompletely understood, several lines of evidence suggest that C-MAD2 can convert more C-MAD2 from O-MAD2 in an autocatalytic manner [39]. Once all kinetochores are properly attached, the SAC is silenced to allow APC/CCDC20 to drive anaphase. Several mechanisms have been implicated in switching off the SAC, including those that involve PP1 [40] and a MAD2-binding protein called p31comet, a protein with a similar structure as C-MAD2 [38]. In one mechanism, p31comet assists in converting C-MAD2 back to O-MAD2 by a p31comet-interacting AAA+-ATPase called TRIP13 [41].

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Acknowledgments Related works in my laboratory are supported in part by grants from the Innovation and Technology Commission (ITCPD/17-9) and the Research Grants Council (16100417, 16102919, 16103020).

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Chapter 2 Substrate Phosphorylation Rates as an In Vivo Measurement of Kinase Activity Matthew P. Swaffer Abstract Measuring kinase activity in different in vivo contexts is crucial for understanding the mechanism and functions of protein kinases, such as the cyclin-dependent kinases (Cdks) and other cell cycle kinases. Here, I present the rationale and the experimental framework for an alternative approach to measure kinase activity that is based on estimating substrate phosphorylation rates in vivo. The approach presented was first developed for experiments performed to measure Cdk1 activity at different stages of the fission yeast S. pombe’s cell cycle [Swaffer et al., Cell 167:1750–1761, 2016]. However, it also affords a more generalizable framework that can be adaptable to other systems and kinases, as long as specific, rapid, and reversible kinase inhibition is possible. Briefly this involves transient and reversible kinase inhibition to dephosphorylate kinase substrates in vivo, followed by quantitative measurements of phosphorylation after inhibition is removed. Key words Phosphorylation, Kinases, Kinase activity, Cdk1

1

Introduction

1.1 Kinases and Their Activity

Changes in kinase activity underpin a wide range of cellular decision-making processes. For instance, a major fraction of the phosphoproteome is dynamically regulated during the cell cycle [1–4]. A naı¨ve view of such processes might consider the total cellular activity of any given kinases as either on or off, but in reality, the activities of many key kinases are dynamically modulated across a range of values. The fission yeast cell cycle control is the perfect illustration of this. Cdk1 is the only cell cycle Cdk in the fission yeast S. pombe and only the one mitotic cyclin (Cdc13) is required to bind and activate Cdk1 for cell division [5–7]. As cells progress through the cell cycle, Cdk1 activity continuously increases, and in doing so, sequentially passes through a series of substrate-specific activity thresholds. At each threshold different subsets of substrate become phosphorylated, resulting in the initiation of the downstream events that their phosphorylation regulates.

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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In this way the different events of the cell cycle can be ordered in time simply by tuning the correct Cdk activity level, because the substrates responsible for later events are phosphorylated at higher activity thresholds than those that function early in the cell cycle [8]. It is simple to imagine how other kinases could function in a similar manner to modulate different biological outputs in response to changing inputs as required. As such, to properly understand the functions and mechanism of kinases such as Cdk1 it is critical to be able to directly measure and compare their protein kinase activity levels in different biological situations. 1.2 Caveats to Current Approaches to Measuring Kinase Activity

Although simply stated, measuring the activity of a kinase is easier said than done. Various estimates of how Cdk activity changes during the cell cycle have been made across different model organisms. Historically, these have been focused on measuring the rate of phosphorylation of a model substrate in cellular extracts or after purifying Cdk1 [9–11]. More recently a number of groups have tried to address this issue more directly by the developing synthetic fluorescent reporters expressed in vivo. The design of these reporters is based on a fragment of a model substrate, the phosphorylation of which is then coupled to some detectable property of the fluorescent reporter, such as changes in its localization or FRET signal [12–14]. While highly informative, these sensors are limited because they are measuring the net phosphorylation of a single “model” substrate, not the substrate’s phosphorylation rate. Consider again the different activity thresholds in the fission yeast cell cycle. As Cdk1 activity passes through different thresholds the net phosphorylation of any given substrate will not necessarily be an accurate reflection of the upstream kinase activity. This is because the phosphorylation state is not a linear function of activity—any given substrate can remain net phosphorylated or net unphosphorylated over a wide range of kinase activity levels above or below their in vivo threshold. For instance, at a low level of Cdk1 activity, the net phosphorylation of early substrate phosphorylation will change in response to changes in kinase activity in the cell. However, despite the fact that Cdk1 activity is increasing, late substrates do not become net phosphorylated. Conversely, when Cdk1 activity is increasing from G2 into mitosis, late substrates’ phosphorylation will change significantly while early substrates are already close to being fully phosphorylated so their phosphorylation will not increase in a manner proportional to the changes in activity [1]. Taking a substrate whose phosphorylation changes over a wider dynamic range would be best but even then, the changes in net phosphorylation are unlikely to be linearly proportional to the changes in kinase activity. Ultimately, such activity sensors cannot always be regarded as quantitative readouts of all changes in kinase activity because the

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read out reflects the net phosphorylation state of a single given substrate. More closely related to kinase activity is the phosphorylation rate of a substrate—more specifically the initial rate of phosphorylation for a previously unphosphorylated substrate. To date such measurements have been restricted to in vitro experimental approaches. 1.3 Direct Measurement of Initial Substrate Phosphorylation Rates In Vivo Can Be Used to Estimate Kinase Activity

To determine how Cdk1 activity changes during the fission yeast cell cycle we developed an approach to measure the phosphorylation rates of substrates in vivo. Phosphorylation rates are a more accurate reflection of kinase activity in the regime in which an unphosphorylated substrate is initially phosphorylated. To get in vivo estimates of phosphorylation rates we developed the following strategy: transiently inhibit kinase activity until all substrates are dephosphorylated and then rapidly remove the inhibition. Substrate phosphorylation can then be measured over a time course after reactivation (Fig. 1). This allows the measurement of the initial increase in phosphorylation by a kinase in vivo on a previously unphosphorylated substrate. Importantly, it is the initial phosphorylation rates that should be used to most closely approximate the kinase’s activity (i.e., the period in which phosphorylation increases linearly) (Fig. 1). While we have typically used mass-spectrometry based phosphoproteomics to quantify substrate phosphorylation during such experiments [1], more conventional approaches using phospho-specific antibodies can also be used. Alternatively, this approach could readily be used in conjunction with the fluorescence-based phosphorylation sensors discussed above. Phosphoproteomics, however, is an excellent approach as it allows for the simultaneous measurement of a large number of substrates at single phosphosite resolution.

Fig. 1 Schematic illustrating the experimental approach for measuring substrate phosphorylation rates as an in vivo estimate of kinase activity. (a) In any given condition the kinase of interest is inactivated for a short period of time by addition of a small molecule inhibitor to dephosphorylate its substrate(s) (grey dots). Then after substrates are dephosphorylated the inhibitor is washed out and removed allowing substrates to be rephosphorylated. Initially the increase in phosphorylation will be linear. In this linear regime the increase in phosphorylation can be used to estimate a phosphorylation rate by calculating the slope of the linear fit to phosphorylation values as a function of time. (b) Changes in the slope reflect changes in kinase activity in vivo: lower slopes represent lower kinase activities

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Materials

2.1 Strains and Media

Yeast strains or cell lines are as is appropriate for the specific nature of the experiment and depend on which kinase and which biological conditions are under investigation. To analyze Cdk1 substrates in S. pombe, a strain harboring an analog sensitive (as) Cdk1 allele was used. The Cdk1-as has a mutation in the active site (F84G) that results in the kinase being sensitive to bulky ATP-analogs [6, 15, 16]. This means that treatment with such bulky ATP-analogs will selectively and reversibly inhibit Cdk1 activity (see Note 1). The same chemical genetic approach has been reported to work with numerous other kinases [17, 18]. Cell media is as appropriate for the specific nature of the experiment and depend on which organisms and which biological conditions are under investigation. For S. pombe experiments Edinburgh Minimal Medium (EMM) should be used [19]. For SILAC based mass-spectrometry labeling a SILAC-compatible strain and modified EMM media is required [20–22].

2.2

Kinase Inhibitor

For Cdk1-as inhibition the bulky ATP-analog 1-NMPP1 is used at 10 μM. For other kinases, if no analog sensitive allele is available, any other characterized small molecule ATP-competitive inhibitor of the respective kinases should work as long as inhibition is specific and reversible upon withdrawal of the drug.

2.3

Filtration Units

For yeast cultures it is essential to remove the inhibitor on a filter membrane as it provides the most rapid method for washing cells. This can be attached to a vacuum pump and is ideally ~90 mm in diameter and has sufficiently large filter pore sizes for rapid filtration (0.22–0.45 μm is optimal for working with S. pombe).

2.4 Yeast Protein Extraction

1. 100% (w/v) Trichloroacetic acid (TCA). 2. 100% Acetone (cold). 3. Lysis buffer: 8 M urea, 50 mM ammonium bicarbonate + cOmplete Mini EDTA-free protease inhibitor cocktail (Roche, Basal, Switzerland) + PhosSTOP phosphatase inhibitor cocktail (Roche). 4. Acid washed glass beads (0.4 mm, Sigma-Aldrich, St. Louis, MO, USA). 5. FastPrep24 or equivalent bead-breaking device. 6. Bradford reagent.

Quantifying Kinase Activity In Vivo

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Method Below is a more detailed explanation of the experimental setup and a point-by-point protocol for the actual inhibition, release and time course experiment as well as subsequent protein extraction.

3.1

Growing Cells

3.2 Inhibition, Release, and Time Course

Grow cells (in the case of S. pombe cells should be grown until mid-exponential phase in EMM media at 25
C in a shaking water bath (see Note 2)) and synchronize them in the relevant stage of the cell cycle or whichever other biological conditions are under examination (see Note 3). 1. Throughout the process of inhibition, release, and time course, exact timings are critical to allow for comparisons between different conditions. Once started, the process requires nearcontinuous work by the experimenter so it is necessary to have everything prepared and close to the culture before starting the experiment, including the following. • 100 ml culture per condition • Receiver culture flask with 75 ml fresh media in water bath. • Filter unit attached to a strong vacuum pump. • 3 aliquot of 100 ml media for washes • 5 ml aliquot of 100% TCA in 50 ml tube on ice (one per timepoint being collected).

3.2.1 Inhibition

1. Once cells are at the appropriate condition, add the inhibitor to 100 ml culture to fully inhibit kinase activity. 10 μM 1-NMPP1 is sufficient to result in the dephosphorylation of nearly all Cdk1 substrates in S. pombe after 15 min at 25
C [1], but for other kinases the exact concentration and time frame required to result in substrate dephosphorylation may have to be experimentally optimized using the conditions for Cdk1 as a sensible starting point.

3.2.2 Release

After 15 min inhibition cells are then washed three times on a filter to remove 1-NMPP1 and then reinoculated into fresh media. For the experiment to work this entire process has to be done extremely fast, that is, ~30 s per wash, which necessitates filtration and resuspension to be completed very quickly (see Note 4). 1. Apply the initial 100 ml culture to the filter membrane attached to the pump. 2. Once the membrane is almost dry (< ~0.5 ml left on the membrane) detach the vacuum, pour 100 ml fresh media onto the membrane and start the clock to time each wash.

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3. Quickly resuspend cells from the filter surface by pipetting 1–2 times with a 25 ml serological pipette. 4. Reattach the vacuum until the membrane is once again almost dry (at this point the clock should read 20–30 s). 5. Repeat steps 2–4 until the cells have been washed in total three times (clock should read ~90 s). 6. Once the third wash is complete resuspend cells in 25 ml media and transferred to a receiver culture flask holding 75 ml fresh media (100 ml final). 3.2.3 Time Course

Once the washes are complete samples are collected at timepoints following the release. 1. For yeast cultures pour 45 ml culture into a 50 ml tube containing 5 ml ice cold 100% TCA at each of the relevant timepoints (final concentration ¼ 10% TCA). 2. Invert the tube and leave on ice for >20 min (see Notes 5 and 6).

3.3 Protein Extraction

Once cells are fixed in 10% TCA, protein can be extracted. The fixation and extraction protocol here are appropriate for either budding or fission yeast studies. If alternative methods are used it is important to ensure they preserve the phosphorylation state of most phosphoproteins. 1. Once all samples have been collected in 10% TCA and incubated for >20 min, pellet the cells (3000  g, 5 min, 4
C) and wash once in cold acetone (3000  g, 5 min, 4
C). 2. Discard off the acetone and retain the pellet (this can now be stored at 80
C until lysis). 3. Resuspend the pellet in 1 ml lysis buffer, transfer to a screw cap 1.5 ml tube and pellet (13,000  g, 2 min, 4
C). 4. Discard the supernatant, retaining the pellet. 5. Resuspend the pellet in 200 μl lysis buffer and add ~1 ml of acid washed glass beads. 6. Lyse cells 3 on a fastprep24 (or equivalent bead-beating machine) for 30 s, setting 6. Incubate on ice for 5 min between beating cycles (check cell breakage is >99% under a microscope after final lysis cycle). 7. Recover lysate from beads by piercing the bottom of the screw cap tube with a red-hot needle and then place in a 1.5 ml tube. Spin (2000  g, 2 min, 4
C) to transfer the lysate into the 1.5 ml tube, thereby separating the lysate from the beads. Discard the beads and keep the lysate.

Quantifying Kinase Activity In Vivo

25

8. Pellet cell debris (13,000  g, 5 min, 4
C) and retain supernatant as protein extract. 9. Determine protein concentration by Bradford assay. 10. Quantify substrate phosphorylation either by Western blotting or mass spectrometry–based phosphoproteomics. Detailed protocols for phosphoproteomic analysis of yeast samples include the SILAC based method [1, 5] and the TMT-labelling based method [23]. 3.4

Data Analysis

1. Once phosphorylation values are quantified, a simple linear fit can be used to model phosphorylation as a function of time from inhibition removal (Fig. 1a). 2. The differences between the slopes of this linear fit in different conditions then gives the estimate of the kinase activity in different conditions (Fig. 1b).

4

Notes 1. Recently an improved Cdk1 analog sensitive allele has been reported that compensates for the structural aberrations caused by the original analog sensitive mutation [16]. Where such rescue mutations are available for different kinases they are preferable. 2. Culturing cells in a water-bath is preferable as it allows easy access to the culture without it being constantly removed from and replaced in an incubator. The experiment and prior culture growth can be performed at 25
C to reduce the reaction rates inside the cell, compared to typical culture conditions at 30–32
C. This should result in the dephosphorylation and rephosphorylation happening slower and therefore afford more accuracy in relative timings and better resolution over the timeframe of rephosphorylation. Growing cells at 25
C also has the added benefit that cells will experience less changes in temperature as they are removed from the water bath to be washed and then again as they are replaced in the water bath. 3. For cell cycle synchronization in S. pombe G2 block and release is well suited as it is compatible with large enough cultures for downstream phosphoproteomics analysis. 4. Washing the culture very quickly requires a strong vacuum pump, a large filter membrane (typically 90 mm in diameter) and sufficiently large filter pore sizes; 0.45 μm is optimal for working with S. pombe. It also requires the volume being filtered to be small enough—we found 100 ml culture to have the best cost-benefit in terms of allowing fast filtration and also sufficient protein yield at the end of the experiment.

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5. Given the close spacing between early timepoints it is critical to collect samples in a manner than instantaneously stops any further phosphorylation or dephosphorylation of proteins in the samples. Fixation in a final concentration of 10% TCA, as above, is excellent for this. 6. Choosing the timepoints for protein sample collection is critical to collecting informative data. Ultimately this has to be empirically optimized. Taking samples just before the washes (i.e., 0 s) and 30, 60, 120, and 600 s after the washes gives a good range for Cdk1 substrates in fission yeast cells grown at 25
C. In this range most substrates analyzed could be fitted to a linear function for at least some of the timepoints. References 1. Swaffer MP, Jones AW, Flynn HR, Snijders AP, Nurse P (2018) Quantitative phosphoproteomics reveals the signaling dynamics of cell-cycle kinases in the fission yeast Schizosaccharomyces pombe. Cell Rep 24(2):503–514. https://doi.org/10.1016/j.celrep.2018.06. 036 2. Olsen JV, Vermeulen M, Santamaria A, Kumar C, Miller ML, Jensen LJ, Gnad F, Cox J, Jensen TS, Nigg EA, Brunak S, Mann M (2010) Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis. Sci Signal 3(104): ra3. https://doi.org/10.1126/scisignal. 2000475 3. Sharma K, D’Souza RC, Tyanova S, Schaab C, Wisniewski JR, Cox J, Mann M (2014) Ultradeep human phosphoproteome reveals a distinct regulatory nature of Tyr and Ser/Thrbased signaling. Cell Rep 8(5):1583–1594. https://doi.org/10.1016/j.celrep.2014.07. 036 4. Carpy A, Krug K, Graf S, Koch A, Popic S, Hauf S, Macek B (2014) Absolute proteome and phosphoproteome dynamics during the cell cycle of Schizosaccharomyces pombe (Fission Yeast). Mol Cell Proteomics 13 (8):1925–1936. https://doi.org/10.1074/ mcp.M113.035824 5. Coudreuse D, Nurse P (2010) Driving the cell cycle with a minimal CDK control network. Nature 468(7327):1074–1079. https://doi. org/10.1038/nature09543 6. Gutierrez-Escribano P, Nurse P (2015) A single cyclin-CDK complex is sufficient for both mitotic and meiotic progression in fission yeast. Nat Commun 6:6871. https://doi.org/10. 1038/ncomms7871

7. Fisher DL, Nurse P (1996) A single fission yeast mitotic cyclin B p34cdc2 kinase promotes both S-phase and mitosis in the absence of G1 cyclins. EMBO J 15(4):850–860 8. Swaffer MP, Jones AW, Flynn HR, Snijders AP, Nurse P (2016) CDK substrate phosphorylation and ordering the cell cycle. Cell 167 (7):1750–1761. e1716. https://doi.org/10. 1016/j.cell.2016.11.034 9. Draetta G, Beach D (1988) Activation of cdc2 protein kinase during mitosis in human cells: cell cycle-dependent phosphorylation and subunit rearrangement. Cell 54(1):17–26. https://doi.org/10.1016/0092-8674(88) 90175-4 10. Labbe JC, Lee MG, Nurse P, Picard A, Doree M (1988) Activation at M-phase of a protein kinase encoded by a starfish homologue of the cell cycle control gene cdc2+. Nature 335 (6187):251–254. https://doi.org/10.1038/ 335251a0 11. Moreno S, Hayles J, Nurse P (1989) Regulation of p34cdc2 protein kinase during mitosis. Cell 58(2):361–372. https://doi.org/10. 1016/0092-8674(89)90850-7 12. Gavet O, Pines J (2010) Progressive activation of CyclinB1-Cdk1 coordinates entry to mitosis. Dev Cell 18(4):533–543. https://doi.org/ 10.1016/j.devcel.2010.02.013 13. Gavet O, Pines J (2010) Activation of cyclin B1-Cdk1 synchronizes events in the nucleus and the cytoplasm at mitosis. J Cell Biol 189 (2):247–259. https://doi.org/10.1083/jcb. 200909144 14. Spencer SL, Cappell SD, Tsai FC, Overton KW, Wang CL, Meyer T (2013) The proliferationquiescence decision is controlled by a bifurcation in CDK2 activity at mitotic exit. Cell 155

Quantifying Kinase Activity In Vivo (2):369–383. https://doi.org/10.1016/j.cell. 2013.08.062 15. Bishop AC, Ubersax JA, Petsch DT, Matheos DP, Gray NS, Blethrow J, Shimizu E, Tsien JZ, Schultz PG, Rose MD, Wood JL, Morgan DO, Shokat KM (2000) A chemical switch for inhibitor-sensitive alleles of any protein kinase. Nature 407(6802):395–401. https://doi.org/ 10.1038/35030148 16. Aoi Y, Kawashima SA, Simanis V, Yamamoto M, Sato M (2014) Optimization of the analogue-sensitive Cdc2/Cdk1 mutant by in vivo selection eliminates physiological limitations to its use in cell cycle analysis. Open Biol 4(7). https://doi.org/10.1098/ rsob.140063 17. Lopez MS, Kliegman JI, Shokat KM (2014) The logic and design of analog-sensitive kinases and their small molecule inhibitors. Methods Enzymol 548:189–213. https://doi.org/10. 1016/B978-0-12-397918-6.00008-2 18. Gregan J, Zhang C, Rumpf C, Cipak L, Li Z, Uluocak P, Nasmyth K, Shokat KM (2007) Construction of conditional analog-sensitive kinase alleles in the fission yeast Schizosaccharomyces pombe. Nat Protoc 2(11):2996–3000. https://doi.org/10.1038/nprot.2007.447 19. Moreno S, Klar A, Nurse P (1991) Molecular genetic analysis of fission yeast

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Schizosaccharomyces pombe. Methods Enzymol 194:795–823. https://doi.org/10.1016/ 0076-6879(91)94059-l 20. Macek B, Carpy A, Koch A, Bicho CC, Borek WE, Hauf S, Sawin KE (2017) Stable isotope labeling by amino acids in cell culture (SILAC) technology in fission yeast. Cold Spring Harb Protoc 6:pdb top079814. https://doi.org/10. 1101/pdb.top079814 21. Carpy A, Koch A, Bicho CC, Borek WE, Hauf S, Sawin KE, Macek B (2017) Stable isotope labeling by amino acids in cell culture (SILAC)-based quantitative proteomics and phosphoproteomics in fission yeast. Cold Spring Harb Protoc 6:pdb prot091686. https://doi.org/10.1101/pdb.prot091686 22. Koch A, Bicho CC, Borek WE, Carpy A, Macek B, Hauf S, Sawin KE (2017) Construction, growth, and harvesting of fission yeast stable isotope labeling by amino acids in cell culture (SILAC) strains. Cold Spring Harb Protoc 6:pdb prot091678. https://doi.org/ 10.1101/pdb.prot091678 23. Touati SA, Hofbauer L, Jones AW, Snijders AP, Kelly G, Uhlmann F (2019) Cdc14 and PP2A phosphatases cooperate to shape phosphoproteome dynamics during mitotic exit. Cell Rep 29(7):2105–2119. e2104. https://doi.org/ 10.1016/j.celrep.2019.10.041

Chapter 3 Real-Time Monitoring of APC/C-Mediated Substrate Degradation Using Xenopus laevis Egg Extracts Julia Kamenz, Renping Qiao, Qiong Yang, and James E. Ferrell Jr Abstract The anaphase promoting complex/cyclosome (APC/C), a large E3 ubiquitin ligase, is a key regulator of mitotic progression. Upon activation in mitosis, the APC/C targets its two essential substrates, securin and cyclin B, for proteasomal destruction. Cyclin B is the activator of cyclin-dependent kinase 1 (Cdk1), the major mitotic kinase, and both cyclin B and securin are safeguards of sister chromatid cohesion. Conversely, the degradation of securin and cyclin B promotes sister chromatid separation and mitotic exit. The negative feedback loop between Cdk1 and APC/C—Cdk1 activating the APC/C and the APC/C inactivating Cdk1—constitutes the core of the biochemical cell cycle oscillator. Since its discovery three decades ago, the mechanisms of APC/C regulation have been intensively studied, and several in vitro assays exist to measure the activity of the APC/C in different activation states. However, most of these assays require the purification of numerous recombinant enzymes involved in the ubiquitylation process (e.g., ubiquitin, the E1 and E2 ubiquitin ligases, and the APC/C) and/or the use of radioactive isotopes. In this chapter, we describe an easy-to-implement method to continuously measure APC/C activity in Xenopus laevis egg extracts using APC/C substrates fused to fluorescent proteins and a fluorescence plate reader. Because the egg extract provides all important enzymes and proteins for the reaction, this method can be used largely without the need for recombinant protein purification. It can also easily be adapted to test the activity of APC/C mutants or investigate other mechanisms of APC/C regulation. Key words Cell cycle, Anaphase promoting complex/cyclosome (APC/C), Xenopus laevis, Frog egg extracts, Plate reader assay, Enzymatic activity

1

Introduction The anaphase promoting complex/cyclosome (APC/C) is a 1.2 MDa E3 ubiquitin ligase. In vertebrates the complex comprises 14 subunits, some in multiple copies (reviewed in [1]). APC/Cmediated ubiquitylation and subsequent proteasomal degradation of distinct substrates at different times during the cell cycle is instrumental in coordinating cell cycle events (reviewed in [2, 3]) and also plays a role in nonproliferative cells [4, 5]. The activity of the APC/C is tightly regulated by multiple mechanisms, including association

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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with coactivators, posttranslational modifications, and inhibitory interactions. In mitosis, the APC/C in association with its coactivator Cdc20 promotes sister chromatid separation and the initiation of mitotic exit [6–9]. At the end of mitosis, APC/C in association with a second coactivator, Cdh1, facilitates cytokinesis [10]. In G1 phase, APC/CCdh1 activity opposes cell cycle progression by mediating the degradation of S-phase and mitotic cyclins [11, 12]. A third coactivator, Ama1, has a specific role in meiosis [13, 14]. APC/C activity is further regulated by phosphorylation. During mitosis, APC/C phosphorylation of an unstructured loop in the APC1 subunit promotes the recruitment of Cdc20 [15–17], whereas phosphorylation of Cdh1 prevents its association with the APC/C [18–20]. Cdh1 dephosphorylation at the end of mitosis promotes the switch from the APC/CCdc20 to the APC/CCdh1 with Cdc20 itself becoming a target of the APC/CCdh1. In somatic cells, APC/CCdc20 activity is additionally controlled by the mitotic checkpoint, a signaling network that delays APC/C activation until all chromosomes have become attached to the mitotic spindle (reviewed in [21]). However, in many organisms, during early embryogenesis this control seems to be absent or unreliable at best [22–24]. Another inhibitor, the early mitotic inhibitor 1 (Emi1), prevents APC/CCdh1 activity during S and G2 phase to allow for DNA replication and cell cycle progression [25, 26]. A close relative, Emi2, has been found to play a role in controlling the APC/CCdc20 in Xenopus laevis eggs and early embryos [26, 27]. Substrate recognition is achieved through the interaction of the coactivators and the APC/C with short linear motifs (SLIMs) within the substrate (reviewed in [28]). The most common motifs are the destruction box (D box, consensus RxxLx[D/E][Ø]xN [N/S]) and the KEN box. Both securin and cyclin B possess a D box in their unstructured N-terminus; securin furthermore has a KEN box. Abolishing these motifs or truncating the N-terminus of cyclin B and securin prevent their APC/C-mediated degradation (the so-called nondegradable versions). Other SLIMs provide additional specificity but are found less frequently; for example, Cyclin A and Nek2A have additional motifs, the ABBA ([ILVF]x[ILMVP] [FHY]x[DE]) motif and the MR-tail, respectively, that facilitate their APC/C-dependent degradation even in the presence of an active mitotic checkpoint. The combinatorial, multivalent interactions of the different motifs as well as the adherence or divergence from the core consensus motif are likely to define substrate specificity and temporal order of substrate degradation. Two kinds of assays are commonly used to measure APC/C activity. The first one directly measures the ubiquitylation of a fluorescence-labeled or radiolabeled model substrate in vitro (e.g., [29–31]). This assay constitutes the most direct measurement of the enzymatic activity of the APC/C, but requires the purification of the APC/C, either after recombinant expression or by

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immunoprecipitation, as well as all other regulators and enzymes involved in the reaction (e.g., E1 and E2 ligase, ubiquitin and the model substrate). On the other hand, APC/C activity has also been measured by proxy, following the APC/C-mediated proteasomal degradation of an APC/C substrate in a cytoplasmic extract that provides all the necessary components [32, 33]. These extracts are often derived from frog eggs. For this, a purified recombinant labeled substrate is added to the extract, samples taken every few minutes, resolved using SDS polyacrylamide gel electrophoresis and detected by immunoblotting or autoradiography. However, this assay has been difficult to scale without losing accuracy due to the necessity to analyze numerous time points for each condition as well as the somewhat limited time resolution. Here we present an alternative way of measuring APC/C activity, which is based on measuring the degradation of an APC/C substrate fused to a fluorescent protein (substrate-FP) in Xenopus laevis egg extracts using a fluorescence plate reader. This method was presented in Yang and Ferrell [34]. By using the frog egg extract, the method preserves the advantage of not having to purify every single component of the ubiquitin ligation reaction, but streamlines the detection to enable a higher throughput of measurements. Although we have mainly used the method to gain insight into the relationship between APC/C activity and its activating kinase Cdk1 [34], we envision that the method can be used in a variety of contexts: (1) studying substrate recognition by using different substrates or different substrate mutants, (2) studying the properties of APC/C with its different coactivators, and (3) studying APC/C regulation directly by using recombinant mutant APC/C. The protocol can be divided into three main steps: (1) Xenopus laevis egg extract preparation and manipulation, (2) substrate preparation by in vitro transcription and translation, and (3) the actual measurement and analysis of APC/C activity. Frog egg extracts can be manipulated in multiple ways in order to accommodate the different research interests. As a starting material we use cytostatic factor (CSF)-arrested or low-speed interphase supernatant (LSS) extracts. Excellent protocols on how to prepare either of these extracts are available [35, 36]. These extracts can then be prepared to resemble interphase or mitosis. Although frog egg extracts only contain the coactivator Cdc20, APC/C activity in the context of other coactivators can be studied by adding the purified recombinant coactivator of interest. Cdc20, the APC/C or other regulators can be immunodepleted and substituted by recombinant variants (for a protocol on how to purify the APC/C and its coactivators see [37]). Different substrate-FPs or mutant variants thereof can easily be cloned into the expression vector and tested using this method.

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Materials Prepare all solutions using ultrapure water and store all reagents at room temperature (unless indicated otherwise).

2.1 Xenopus laevis Egg Extract Preparation and Manipulation

If starting from CSF-arrested extracts: 1. 40 mM CaCl2. 2. 10 mg/ml cycloheximide (stored at
20  C). If mitotic extracts are desired (e.g., if APC/CCdc20 activity should be measured): 3. 10 mg/ml cycloheximide (stored at
20  C). 4. Nondegradable cyclin B (see Note 1, stored at
80  C). 5. Optional: Wee1 inhibitor (e.g., PD0166285, dissolved in DMSO, stored at
20  C).

2.2 Immunodepletion

1. Antibody raised against the protein of interest (for APC/C immunodepletion antibodies against the APC3/Cdc27 subunit are commonly used). An irrelevant antibody (e.g., purified IgG) as a control. 2. Protein A or Protein G magnetic beads (depending on the antibody). 3. HEPES buffered saline (HBS): 50 mM HEPES, 1.5 mM Na2HPO4, 140 mM NaCl. 4. The identical extract buffer used to crush the Xenopus laevis eggs (e.g., CSF-XB or ELB).

2.3 Preparation of Fluorescently Labeled Substrate by In Vitro Translation

1. Plasmid encoding the APC/C substrate of interest fused to a fluorescent protein (substrate-FP) usually under the control of a T7 or SP6 promoter (see Note 2, store at
20  C). 2. Coupled in vitro transcription/translation system or cell-free protein expression system compatible with the plasmid promoter (see Notes 3 and 4, usually stored at
80  C). 3. Temperature block (with mixing function) depending on the chosen system for protein expression.

2.4 Measuring APC/C Activity Using a Fluorescence Plate Reader

1. Fluorescence plate reader with fluorescence excitation and emission detection capabilities in the range of the selected fluorescent protein and compatible with 384-well plates. Temperature control around 20–25  C is an advantage. 2. 384-Well microwell plate, flat bottom, suitable for fluorescence measurements (see Note 5).

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Methods Carry out all procedures at room temperature, unless otherwise specified.

3.1 Egg Extract Preparation

3.1.1 Preparing Interphase-Arrested Extract from CSF Extract

Depending on the experiment, APC/C activity can be measured in interphase (e.g., APC/CCdh1) or in mitosis (e.g., APC/CCdc20). CSF-arrested or interphase-arrested frog egg extracts can be used as the starting material. These extracts can be stored at
80  C for at least one month. Many protocols add an energy mix, though we have performed experiments successfully without it. 1. Thaw extract(s) for 5 min at 20  C and if necessary pool the aliquots. 2. Add cycloheximide to a final concentration of 100 μg/ml. 3. Add CaCl2 to a final concentration of 0.8 mM. 4. Mix thoroughly by carefully pipetting up and down (at least ten times). 5. Incubate for 50 min at 20  C. Stir the extract with a pipet tip every 10 min to keep the extract well mixed.

3.1.2 Preparing Mitotic Extract from Interphase Extract

1. If you have not added it already, add cycloheximide to 100 μg/ ml. 2. Add nondegradable cyclin B to about 150 nM (see Note 6). 3. Mix thoroughly by carefully pipetting up and down (at least ten times). 4. Incubate for 1 h at 20  C. Stir the extract with a pipet tip every 10 min to keep the extract well mixed. Optional: Take samples from the CSF, interphase, and mitotic extract to analyse the phosphorylation status of the APC/C and other proteins later by immunoblotting.

3.2 Immunodepletion

The exact amount of antibody used for the immunodepletion needs to be established for each protein–antibody pair. Antibodies can either be covalently coupled to the beads, which allows for the possibility of eluting the protein and reusing the beads, or simply bound to the beads overnight as we describe here. In addition to the antibody coupled beads, prepare control IgG coupled beads in the same way. 1. Use about 10 μl magnetic beads per round of depletion for each 30 μl of extract. As two rounds of depletion are performed, prepare a total of 20 μl of magnetic beads per 30 μl of extract.

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2. Remove supernatant and wash the magnetic beads twice with two bead volumes of HBS. 3. Resuspend beads in two bead volumes of HBS and add the empirically determined amount of antibody. 4. Rotate overnight at 4  C. 5. Collect the beads using a magnet and wash five times with two bead volumes of extract buffer (e.g., CSF-XB and ELB). Remove all buffer. 6. Add the egg extract to one half of the antibody beads. Rotate for 40 min at 4  C. Collect the beads using a magnet. 7. Transfer the extract to a fresh tube with the other half of the antibody beads. Rotate for 40 min at 4  C. Collect the beads using a magnet. 8. Transfer immunodepleted extract to a fresh tube. Store on ice until further use (typically no longer than 1 h). Optional: Take extract samples for immunoblotting to confirm efficient protein depletion. 3.3 Preparation of Fluorescently Labeled Substrate by In Vitro Transcription/ Translation

The fluorescently labelled substrate can be prepared in parallel to the (immunodepleted) extract or ahead of time. The substrate-FB is best used fresh, but can be stored 2–3 days at 4  C protected from light if necessary. 1. For best results ensure an RNase-free environment (e.g., wear gloves, clean pipettes with RNase AWAY, and use RNase-free consumables). 2. Add the plasmid to the coupled transcription/translation system following manufacturer’s instructions. In addition, prepare a blank control for measuring the baseline fluorescence of the extract later. 3. During the incubation, protect your fluorescent substrate from light by covering the temperature block or wrapping aluminium foil around the tube. 4. Confirm the successful transcription and translation of the substrate-FP by measuring the fluorescence relative to the blank control using a plate reader (see Subheading 3.4). The fluorescence should be at least 50-fold above the blank control for good results in the assay. 5. No further purification of the protein is necessary.

3.4 Measuring APC/C Activity Using a Fluorescence Plate Reader

A typical measurement is performed at least as a duplicate using about 20 μl of extract, manipulated beforehand in the desired way, per measurement. Degradation might start right after mixing the substrate-FP and the extracts; therefore, all steps are performed on ice and in proximity to the plate reader. The plate reader should be

Monitoring APC/C-Mediated Substrate Degradation

Interphase M phase Background

225 200 175 150 125 100 75 50 0

20

40

60

80 100 120

Time (min)

B

Interphase M phase

1.0 Normalized securin-CFP

Securin-CFP fluorescence (a.u.)

A

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0.8 0.6 0.4 0.2 0

0

20

40

60

80 100 120

Time (min)

Fig. 1 (a) Measured fluorescence of an interphase or M phase (mitotic) extract supplemented with securin-CFP as a function of time. An interphase extract supplemented with the blank control in vitro transcription/ translation reaction was used as a background control. Shown are data from two technical replicates (full or empty circles respectively). (b) Data shown in (a) after background subtraction and normalization. The dashed or solid lines, respectively, show the exponential fit of the data

equilibrated to the desired temperature and all settings adjusted before the mixing step. 1. Add the substrate-FP to the extract in the ratio 1:16–1:20 (see Note 7). Additionally, set up at least one sample with the blank control to get a good background reading. 2. Mix vigorously by pipetting up and down (at least ten times). Do not vortex! Try to avoid air bubbles as much as possible. 3. Carefully pipette 20 μl of extract into each well of the microplate avoiding air bubbles as much as possible. 4. Load plate into the plate reader and start measurement (top read). We commonly measure for 120 min, with 1 min intervals and mixing before every measurement. Excitation and emission wavelengths have to be determined for the specific fluorescent protein; for CFP we use 435 nm/475 nm (see Fig. 1a for an example measurement). 3.5

Data Analysis

We are using the apparent first order rate constant (k) as a measure for APC/C activity. To estimate the first order rate constant, the following steps are undertaken: 1. Subtract the background measurement from all measurements. 2. Normalize the measurements to the maximum (see Note 8). 3. Fit the data to fluorescence (t) ¼ A0  e
k  t + C (see Note 9, see Fig. 1b for an example of the normalized and fitted data).

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Notes 1. We commonly use recombinant nondegradable cyclin B from sea urchin purified from bacteria (for a protocol see [38]) or nondegradable cyclin B1 from Xenopus laevis purified from baculovirus-infected insect cells (for a protocol see [39]). 2. We commonly use SP6-securin-CFP but have also successfully used cyclin B-CFP or cyclin A-CFP as well as other fluorescent proteins (e.g., mCherry). 3. Do not use transcription/translation systems based on rabbit reticulocytes as their pink color interferes with the fluorescence measurements. We commonly use transcription/translation systems based on wheat germ. Depending on the system and promoter used, the plasmid might have to be linearized and purified before used for transcription/translation (though this was not applicable for our system–promoter combination). 4. So far, we have not been able to perform consistent measurements using purified recombinant substrate-FPs and therefore advise using the in vitro transcribed and translated protein. 5. Several plate suppliers exist. We use black 384-well fluotrac 200 plates from Greiner. 6. The exact concentration of nondegradable cyclin B necessary for promoting the mitotic state varies from batch to batch and also depends on the exact construct used (e.g., we have observed that higher concentrations of bacterially expressed sea urchin nondegradable cyclin B are necessary relative to insect cell expressed Xenopus laevis nondegradable cyclin B). 7. Extracts should be diluted as little as possible and more than 10% dilution should be avoided. Depending on the yield of the in vitro translation/transcription reaction 1:16–1:20 works well for us to avoid extract dilution while still having a good signal-to-noise ratio for the measurement. 8. To account for some variability in the starting concentration of the substrate-FP, the data is normalized. 9. In theory, due to the normalization and the background subtraction, A0 should be close to 1 and C should be close to 0 in the fit and we normally constraint these parameters for the fit (A0 > 0.8 and C < 0.2).

Acknowledgments We thank Jan-Michael Peters and members of the Peters lab from the Research Institute of Molecular Pathology (IMP, Vienna, Austria) for their support and helpful discussions. The work was assisted by grants from the National Institutes of Health (R35

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GM131792 to J.E.F., and R35 GM119688 to Q.Y.), the National Science Foundation (Early CAREER Grant #1553031 and MCB #1817909 to Q.Y.) and a postdoctoral fellowship from the German Research Foundation (KA 4476/1-1, J.K.). References 1. Alfieri C, Zhang S, Barford D (2017) Visualizing the complex functions and mechanisms of the anaphase promoting complex/cyclosome (APC/C). Open Biol 7(11). https://doi.org/ 10.1098/rsob.170204 2. Peters JM (2006) The anaphase promoting complex/cyclosome: a machine designed to destroy. Nat Rev Mol Cell Biol 7(9):644–656. https://doi.org/10.1038/nrm1988 3. Pines J (2011) Cubism and the cell cycle: the many faces of the APC/C. Nat Rev Mol Cell Biol 12(7):427–438. https://doi.org/10. 1038/nrm3132 4. Delgado-Esteban M, Garcia-Higuera I, Maestre C, Moreno S, Almeida A (2013) APC/C-Cdh1 coordinates neurogenesis and cortical size during development. Nat Commun 4:2879. https://doi.org/10.1038/ ncomms3879 5. Eguren M, Porlan E, Manchado E, GarciaHiguera I, Canamero M, Farinas I, Malumbres M (2013) The APC/C cofactor Cdh1 prevents replicative stress and p53-dependent cell death in neural progenitors. Nat Commun 4:2880. https://doi.org/10.1038/ncomms3880 6. Irniger S, Piatti S, Michaelis C, Nasmyth K (1995) Genes involved in sister chromatid separation are needed for B-type cyclin proteolysis in budding yeast. Cell 81(2):269–278. https://doi.org/10.1016/0092-8674(95) 90337-2 7. King RW, Peters JM, Tugendreich S, Rolfe M, Hieter P, Kirschner MW (1995) A 20S complex containing CDC27 and CDC16 catalyzes the mitosis-specific conjugation of ubiquitin to cyclin B. Cell 81(2):279–288. https://doi. org/10.1016/0092-8674(95)90338-0 8. Sudakin V, Ganoth D, Dahan A, Heller H, Hershko J, Luca FC, Ruderman JV, Hershko A (1995) The cyclosome, a large complex containing cyclin-selective ubiquitin ligase activity, targets cyclins for destruction at the end of mitosis. Mol Biol Cell 6(2):185–197. https:// doi.org/10.1091/mbc.6.2.185 9. Tugendreich S, Tomkiel J, Earnshaw W, Hieter P (1995) CDC27Hs colocalizes with CDC16Hs to the centrosome and mitotic spindle and is essential for the metaphase to anaphase transition. Cell 81(2):261–268. https:// doi.org/10.1016/0092-8674(95)90336-4

10. Floyd S, Pines J, Lindon C (2008) APC/C Cdh1 targets aurora kinase to control reorganization of the mitotic spindle at anaphase. Curr Biol 18(21):1649–1658. https://doi. org/10.1016/j.cub.2008.09.058 11. Li M, Shin YH, Hou L, Huang X, Wei Z, Klann E, Zhang P (2008) The adaptor protein of the anaphase promoting complex Cdh1 is essential in maintaining replicative lifespan and in learning and memory. Nat Cell Biol 10 (9):1083–1089. https://doi.org/10.1038/ ncb1768 12. Sigrist SJ, Lehner CF (1997) Drosophila fizzyrelated down-regulates mitotic cyclins and is required for cell proliferation arrest and entry into endocycles. Cell 90(4):671–681. https:// doi.org/10.1016/s0092-8674(00)80528-0 13. Cooper KF, Mallory MJ, Egeland DB, Jarnik M, Strich R (2000) Ama1p is a meiosisspecific regulator of the anaphase promoting complex/cyclosome in yeast. Proc Natl Acad Sci U S A 97(26):14548–14553. https://doi. org/10.1073/pnas.250351297 14. Okaz E, Arguello-Miranda O, Bogdanova A, Vinod PK, Lipp JJ, Markova Z, Zagoriy I, Novak B, Zachariae W (2012) Meiotic prophase requires proteolysis of M phase regulators mediated by the meiosis-specific APC/CAma1. Cell 151(3):603–618. https:// doi.org/10.1016/j.cell.2012.08.044 15. Fujimitsu K, Grimaldi M, Yamano H (2016) Cyclin-dependent kinase 1-dependent activation of APC/C ubiquitin ligase. Science 352 (6289):1121–1124. https://doi.org/10. 1126/science.aad3925 16. Qiao R, Weissmann F, Yamaguchi M, Brown NG, VanderLinden R, Imre R, Jarvis MA, Brunner MR, Davidson IF, Litos G, Haselbach D, Mechtler K, Stark H, Schulman BA, Peters JM (2016) Mechanism of APC/CCDC20 activation by mitotic phosphorylation. Proc Natl Acad Sci U S A 113 (19):E2570–E2578. https://doi.org/10. 1073/pnas.1604929113 17. Zhang S, Chang L, Alfieri C, Zhang Z, Yang J, Maslen S, Skehel M, Barford D (2016) Molecular mechanism of APC/C activation by mitotic phosphorylation. Nature 533 (7602):260–264. https://doi.org/10.1038/ nature17973

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18. Blanco MA, Sanchez-Diaz A, de Prada JM, Moreno S (2000) APC(ste9/srw1) promotes degradation of mitotic cyclins in G(1) and is inhibited by cdc2 phosphorylation. EMBO J 19(15):3945–3955. https://doi.org/10. 1093/emboj/19.15.3945 19. Kramer ER, Scheuringer N, Podtelejnikov AV, Mann M, Peters JM (2000) Mitotic regulation of the APC activator proteins CDC20 and CDH1. Mol Biol Cell 11(5):1555–1569. https://doi.org/10.1091/mbc.11.5.1555 20. Zachariae W, Schwab M, Nasmyth K, Seufert W (1998) Control of cyclin ubiquitination by CDK-regulated binding of Hct1 to the anaphase promoting complex. Science 282 (5394):1721–1724. https://doi.org/10. 1126/science.282.5394.1721 21. Lara-Gonzalez P, Westhorpe FG, Taylor SS (2012) The spindle assembly checkpoint. Curr Biol 22(22):R966–R980. https://doi. org/10.1016/j.cub.2012.10.006 22. Chenevert J, Roca M, Besnardeau L, Ruggiero A, Nabi D, McDougall A, Copley RR, Christians E, Castagnetti S (2020) The spindle assembly checkpoint functions during early development in non-chordate embryos. Cell 9(5). https://doi.org/10.3390/ cells9051087 23. Galli M, Morgan DO (2016) Cell size determines the strength of the spindle assembly checkpoint during embryonic development. Dev Cell 36(3):344–352. https://doi.org/10. 1016/j.devcel.2016.01.003 24. Minshull J, Sun H, Tonks NK, Murray AW (1994) A MAP kinase-dependent spindle assembly checkpoint in Xenopus egg extracts. Cell 79(3):475–486. https://doi.org/10. 1016/0092-8674(94)90256-9 25. Dong X, Zavitz KH, Thomas BJ, Lin M, Campbell S, Zipursky SL (1997) Control of G1 in the developing Drosophila eye: rca1 regulates cyclin A. Genes Dev 11(1):94–105. https://doi.org/10.1101/gad.11.1.94 26. Reimann JD, Freed E, Hsu JY, Kramer ER, Peters JM, Jackson PK (2001) Emi1 is a mitotic regulator that interacts with Cdc20 and inhibits the anaphase promoting complex. Cell 105(5):645–655. https://doi.org/10. 1016/s0092-8674(01)00361-0 27. Tischer T, Hormanseder E, Mayer TU (2012) The APC/C inhibitor XErp1/Emi2 is essential for Xenopus early embryonic divisions. Science 338(6106):520–524. https://doi.org/10. 1126/science.1228394 28. Davey NE, Morgan DO (2016) Building a regulatory network with short linear sequence motifs: lessons from the degrons of the

anaphase-promoting complex. Mol Cell 64 (1):12–23. https://doi.org/10.1016/j. molcel.2016.09.006 29. Carroll CW, Morgan DO (2005) Enzymology of the anaphase-promoting complex. Methods Enzymol 398:219–230. https://doi.org/10. 1016/S0076-6879(05)98018-X 30. Izawa D, Pines J (2015) The mitotic checkpoint complex binds a second CDC20 to inhibit active APC/C. Nature 517(7536):631–634. https://doi.org/10.1038/nature13911 31. Kraft C, Gmachl M, Peters JM (2006) Methods to measure ubiquitin-dependent proteolysis mediated by the anaphase-promoting complex. Methods 38(1):39–51. https://doi. org/10.1016/j.ymeth.2005.07.005 32. Yamaguchi M, Yu S, Qiao R, Weissmann F, Miller DJ, VanderLinden R, Brown NG, Frye JJ, Peters JM, Schulman BA (2015) Structure of an APC3-APC16 complex: insights into assembly of the anaphase-promoting complex/cyclosome. J Mol Biol 427(8):1748–1764. https:// doi.org/10.1016/j.jmb.2014.11.020 33. Yamano H, Trickey M, Grimaldi M, Kimata Y (2009) In vitro assays for the anaphasepromoting complex/cyclosome (APC/C) in Xenopus egg extracts. Methods Mol Biol 545:287–300. https://doi.org/10.1007/ 978-1-60327-993-2_18 34. Yang Q, Ferrell JE Jr (2013) The Cdk1-APC/C cell cycle oscillator circuit functions as a timedelayed, ultrasensitive switch. Nat Cell Biol 15 (5):519–525. https://doi.org/10.1038/ ncb2737 35. Banaszynski LA, Allis CD, Shechter D (2010) Analysis of histones and chromatin in Xenopus laevis egg and oocyte extracts. Methods 51 (1):3–10. https://doi.org/10.1016/j.ymeth. 2009.12.014 36. Murray AW (1991) Cell cycle extracts. Methods Cell Biol 36:581–605 37. Jarvis MA, Brown NG, Watson ER, VanderLinden R, Schulman BA, Peters JM (2016) Measuring APC/C-dependent ubiquitylation in vitro. Methods Mol Biol 1342:287–303. https://doi.org/10.1007/9781-4939-2957-3_18 38. Glotzer M, Murray AW, Kirschner MW (1991) Cyclin is degraded by the ubiquitin pathway. Nature 349(6305):132–138. https://doi.org/ 10.1038/349132a0 39. Ha SH, Kim SY, Ferrell JE Jr (2016) The prozone effect accounts for the paradoxical function of the Cdk-binding protein Suc1/Cks. Cell Rep 14(6):1408–1421. https://doi.org/ 10.1016/j.celrep.2016.01.033

Chapter 4 Fluorescent Peptide Biosensors for Probing CDK Kinase Activity in Cell Extracts Morgan Pellerano and May C. Morris Abstract Fluorescent biosensors can report on the relative abundance, activity, or conformation of biomolecules and analytes through changes in fluorescence emission. A wide variety of genetically-encoded and synthetic biosensors have been developed to monitor protein kinase activity. We have focused on the design, engineering and characterization of fluorescent peptide biosensors of cyclin-dependent kinases (CDKs) that constitute attractive cancer biomarkers and pharmacological targets. In this chapter, we describe the CDKACT fluorescent peptide biosensor technology and its application to assess the relative kinase activity of CDKs in vitro, either using recombinant proteins or cell extracts as a more complex source of kinase. This technology offers a straightforward means of comparing CDK activity in different cell lines and evaluating the specific impact of treatments intended to target kinase activity in a physiologically relevant environment. Key words Fluorescent biosensor, Cyclin-dependent kinase, Kinase activity, Peptide, Cancer biomarker, Cell extracts

1

Introduction Studies of protein kinase signaling pathways have traditionally relied on the use of antigenic approaches to detect protein kinase expression levels and their phosphorylated substrates, through Western blotting, immunohistochemistry, and immunofluorescence. However, while antibodies allow to detect the presence and quantify the relative protein levels of a kinase of interest, they do not convey any information regarding its functional status, which is dependent on a network of activating and inhibitory posttranslational signals. The development of genetically-encoded and synthetic fluorescent biosensors that report on kinase activities has addressed this challenge providing a functional readout and offering opportunities to develop diagnostic strategies [1–4]. Genetically-encoded kinase activity reporters (KARs) are most often expressed in living cells for imaging purposes, while peptide-based biosensors tend to be more useful reporters

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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in vitro, to probe kinase activities in cell extracts for instance, although they may also be delivered into living cells for imaging purposes [3, 5–7]. Fluorescent peptide biosensors (FPBs) can be easily synthesized by chemical synthesis and can be readily incubated with recombinant proteins or cell lysates to evaluate kinase activity in vitro. Over the past decade a wide variety of FPBs that report on protein kinase activity have been developed for Abl, Akt1, MK2, Pim2, PKA, PKC, Src, and CDKs [2–4, 8–10]. The design of a performant biosensor requires several criteria be met, most importantly specificity/selectivity and sensitivity [11]. It is therefore essential to select and validate a specific substrate sequence for the kinase of interest for each activity biosensor, either through rational design of a sequence derived from a specific substrate described in the literature or through screening of a library of peptide sequences [2]. The choice of the fluorescent dye to be incorporated in the peptide scaffold will determine the sensitivity and the dynamic range of biosensor response to kinase activity. Biosensor response should be dose-dependent and robust, with a sufficiently high signal-to-noise ratio. When developing a new biosensor, it is advisable to compare several dyes in an in-depth study to identify which one is the most appropriate. In this respect solvatochromic or environmentally-sensitive fluorophores such as Dansyl, FITC, TAMRA, cyanine and merocyanine dyes constitute the most suitable and commercially available dyes [12, 13]. Over the past 5 years, we have focused on the design, engineering and validation of fluorescent peptide biosensors that report on CDK activities [14–18], key enzymes that play a central role in coordinating cell growth and division [19, 20]. Since many CDKs are hyperactivated in human cancers and contribute to sustain oncogenic hyperproliferation they are considered relevant biomarkers and established pharmacological targets for cancer therapeutics [21–25]. CDKs are poorly abundant in mammalian cells (subnanomolar concentrations reported in the literature [26]) and their relative abundance does not reflect their activity status. CDK activation relies on their interaction with a regulatory cyclin or cyclinlike partner, and further regulation by a network of structural inhibitors and posttranslational modifications (phosphorylation/ dephosphorylation of the CDK) [19]. CDKACT fluorescent peptide biosensor technology was initially developed as a proof of concept for overall detection of CDK activities and further derivatives were then engineered specifically for CDK4, CDK5, CDK6 [14–18]. In this chapter we describe how the CDKACT fluorescent peptide biosensor technology can be used in vitro using recombinant proteins or cell extracts, so as to assess relative kinase activity of CDKs in a complex and physiologically relevant environment and compare functional differences in different cell lines or following treatment with drugs.

Fluorescent Peptide Biosensors for Probing CDK Kinase Activity in Cell Extracts

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Materials Prepare all solutions using ultrapure water (unless otherwise stated) and analytical grade reagents.

2.1 Peptide Biosensor Preparation and Labeling

CDKACT peptide biosensors combine a sequence derived from the WWdomain of Pin1 which serves as a phospho–amino acid binding domain (PAABD) with a peptide sequence derived from a protein substrate which is specifically recognized and phosphorylated by the kinase of interest. A unique cysteine is included in the substrate sequence at position
2 relative to the serine or threonine of the S/ T-P phosphorylation site (Fig. 1). Peptide biosensors described in this chapter are CDKACT4, CDKACT5, and CDKACT6 [15, 17, 18]. 1. Lyophilized CDKACT peptide. 2. Phosphate buffer saline (PBS). 3. Dimethyl sulfoxide (DMSO). 4. Precision balance. 5. Ultrasound bath. 6. Environmentally-sensitive dyes such as TAMRA, fluorescein, dansyl, cyanine dyes etc. 7. NAP-5 column or NAP-10 column. 8. Spectrofluorimeter.

Fig. 1 Schematic representation of CDKACT peptide biosensors

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2.2 Source of Kinases: Recombinant Protein or Cell Extracts

1. Recombinant proteins: either commercially purchased or expressed in E. coli and purified by fast protein liquid chromatography (FPLC). 2. Cell extracts prepared from cultured mammalian cell lines. 3. PBS lysis Buffer: 2 mM phenylmethylsulfonyl fluoride (PMSF), 0.2% Nonidet-P40, 1 mM ethylenediaminetetraacetic acid (EDTA) diluted with PBS. 4. Complete™, Mini, EDTA-free Protease Inhibitor Cocktail from Roche. 5. Refrigerated centrifuge. 6. Ultrasound bath. 7. Spectrophotometerr.

2.3 Activity Assay in Black 96-Well Plates

1. PBS. 2. PBS lysis buffer. 3. A 10  ATP/MgCl2 stock solution: 50 mM MgCl2 and 5 mM ATP diluted with PBS. 4. Black 96-well Polystyrene microplate with flat bottom. 5. Fluorescently labeled CDKACT peptide at 10
4 M. 6. Recombinant protein at 10
6 M or cell extract. 7. Protein kinase inhibitor at 10
2 M: Roscovitine or Abemaciclib (LY2835219). 8. 96-well plate reader fluorimeter (see Note 1).

3

Methods

3.1 Fluorescent Biosensor Preparation

1. CDKACT peptides are synthesized and purified to 95% and lyophilized as described in [15, 17, 18]. 2. Lyophilized CDKACT peptides are stored at
20  C (see Note 2).

3.1.1 Prelabeled CDKACT

1. Take the lyophilized TAMRA-labeled CDKACT peptide out of the freezer and let it reach room temperature (15 min minimum). 2. Weigh 1 mg using a precision balance. 3. Dissolve the powder in PBS in the appropriate volume to achieve 10
4 M final concentration. 4. Place the sample in an ultrasound bath for 10 min. 5. Check the optical density (O.D) with an UV spectrophotometer at the wavelength corresponding to TAMRA: 550 nm.

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6. Calculate the concentration with the Beer-Lambert law: O.D ¼ ε  l  c (ε: molar extinction coefficient, l: optical path length, and c: concentration of the sample; see Note 3). 7. Keep the solution at room temperature for immediate use or store at
20  C (see Note 4). 3.1.2 Unlabeled CDKACT

1. Take the lyophilized CDKACT peptide out of the freezer and let it reach room temperature (15 min minimum). 2. Weigh 1 mg using a precision balance. 3. Dissolve the powder in PBS in the appropriate volume to achieve 10
3 M final concentration. 4. Place the sample in an ultrasound bath for 10 min. 5. Centrifuge at 16200  g for 10 min in a microcentrifuge to check if the peptide is correctly dissolved. 6. In a 1.5 ml microtube mix by vortexing: 400 μl PBS, 50 μl peptide at 10
3 M, 50 μl of an environmentally-sensitive dye (TAMRA, FITC, Cy3, Cy5, etc.) at 2.10
3 M. 7. Incubate overnight at 4  C with constant agitation and protection from light with aluminum paper (see Note 5). 8. Separate excess probe from labeled peptide on a NAP-5 column and collect five tubes with 250 μl (see Note 6). 9. Check the O.D of each sample with an UV spectrometer at the wavelength corresponding to the absorbance of the dye used to label the peptide. 10. Calculate the concentration with the Beer–Lambert law: O. D ¼ ε  l  c (ε: molar extinction coefficient, l: optical path length and c: concentration of the sample; see Note 7). 11. Store the solution at room temperature for quick use or store at
20  C (see Note 8).

3.2 Cell Extract Sample Preparation

1. Resuspend the cell pellets in 150 μl PBS lysis buffer including Protease inhibitors (3 ul Complete™, Mini, EDTA-free Protease Inhibitor Cocktail from Roche). 2. Centrifuge in a microcentrifuge for 16200  g at 4  C for 10 min. 3. Recover the supernatant. 4. Determine the total protein concentration at 280 nm and adjust it to 1 μg/μl (see Note 9). 5. Store the sample on ice (see Note 10).

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Fig. 2 Schematic representation of a typical plate layout for kinase activity assay 3.3 Performing Kinase Assays with CDKACT Biosensors

1. Dilute the labeled CDKACT solution from 10
4 to 10
6 M in PBS (see Note 11). 2. In the 96-well plate, prepare the following plate plan (Fig. 2): – One well containing 140 μl PBS, 20 μl 10  ATP/MgCl2, 40 μl labeled CDKACT at 10
6 M (i.e., 200 nM final concentration) (see Note 12). – One well containing 100 μl PBS, 20 μl 10 ATP/MgCl2, 40 μl labeled CDKACT at 10
6 M, 40 μl PBS lysis buffer (see Note 12). – One well which should be replicated at least three times, each containing 100 μl PBS, 20 μl 10  ATP/MgCl2, 40 μl labeled CDKACT at 10
6 M, 40 μl cell extract at 1 μg/μl (i.e., 40 μg final or a more diluted concentration if desired) (see Notes 12 and 13). – It is strongly advised to prepare another well to be replicated at least three times containing 80 μl PBS, 20 μl 10  ATP/MgCl2, 40 μl labeled CDKACT at 10
6 M, 40 μl cell extract at 1 μg/μl (i.e., 40 μg final or a more diluted concentration if desired) (see Notes 12 and 13) and 20 μl 20  inhibitor concentration from 10 mM stocks (generally between 5 and 20 μM CDK inhibitors are used to ensure complete CDK inhibition). – Prepare this series of wells in triplicate (Fig. 2). – The outer wells of the plate should not be used to avoid side effects and either left empty or filled with PBS or ultrapure H2O. 3. Place the plate in the spectrofluorimeter thermostated at 30  C. 4. Measure the fluorescence emission following excitation of the corresponding dye every 30 s for 30–45 min (see Note 14–16).

Fluorescent Peptide Biosensors for Probing CDK Kinase Activity in Cell Extracts

3.4 Determination of Kinase Curves

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1. Recover the raw data acquired by the plate-reader in an excel sheet and plot fluorescence emission values against time on a graph (see Note 17) (Fig. 3a). 2. Calculate relative fluorescence values for the biosensor alone, for biosensor incubated with lysis buffer and biosensor incubated with cell extracts prepared in lysis buffer by normalizing fluorescence emission values (i.e., dividing values at each timepoint by the initial fluorescence emission value) (Fig. 3b). 3. In order to obtain a typical kinase activity curve, subtract normalized fluorescence emission values corresponding to CDKACT biosensor with lysis buffer from fluorescence values corresponding to CDKACT biosensor incubated with cell extracts (Fig. 3c). Kinase activity is represented as the increase in the percentage of relative fluorescence emission. 4. When characterizing a peptide biosensor for the first time, it is wise to compare several fluorophores and evaluate the sensitivity of response so as to select the most appropriate dye to achieve the greatest dynamic range and the most robust response. For instance, CDKACT4 biosensor responds to CDK4 kinase activity in cell extracts with much greater sensitivity when it is conjugated to TAMRA compared to Cy3 (Fig. 4).

Fig. 3 Fluorescence emission of CDKACT4-TAMRA to A375 cell extracts. (a) Raw data; (b) Relative data; (c) Final activity curve

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Fig. 4 Response of CDKACT4-TAMRA and CDKACT4-Cy3 to CDK4 activities in A375 cell extracts 3.5 Histogram Representation of Relative Maximal Activity

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1. Histograms can be generated to represent relative fluorescence intensity values at a given time point, generally corresponding to maximal fluorescence response of the biosensor. 2. Histogram representations generally compare the relative maximal activity of different samples at 1800 s (30 min) or 2400 s (40 min). For example maximal biosensor response to a kinase of interest in cell extracts from different cell lines (Fig. 5a) or maximal biosensor response of kinase samples incubated with different concentrations of kinase inhibitors (Fig. 5b) (see Note 18).

Notes 1. The Clariostar™ spectrofluorimeter plate-reader (BMG) was used for all experiments described in this chapter. 2. Lyophilized peptides can be stored at
20  C in the freezer for a couple of months. It is not wise to store them longer than 6 months. 3. If the concentration is greater than 10
4 M, adjust it with PBS. If it is lower, use the measured concentration value, rather than the calculated one. 4. The peptide diluted to 10
4 M can be stored for 2 weeks at
20  C. 5. The best incubation condition is constant rotation. 6. For a 1 ml solution, use NAP-10 column.

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Fig. 5 Histogram representation of relative maximal biosensor response. (a) Response of CDKACT6 to CDK6 kinase activity in different cell lines (activity curves on the left panel, histogram representing relative fluorescence emission at 2400 s on the right panel); (b) Maximal CDKACT5 response to CDK5 activity in U87 cell extracts treated with 20 μM CDK inhibitor Roscovitine (activity curves on the left panel, histogram representing relative fluorescence emission at 2400 s on the right panel)

7. Generally, the yield of labeled peptide is between 20 and 50% of the initial unlabeled peptide concentration. If the ratio of labeled/unlabeled peptide yield is too low, excess unlabeled peptide may perturb the reaction. 8. Labeled peptide in solution can be stored for 2 weeks at
20  C. 9. The total protein concentration is determined relative to a standard BSA concentration curve from 0.1 to 10 μg/μl. 10. The sample should be used within the same hour to avoid protein degradation and ensure kinase activity is preserved. 11. When preparing a large number of samples it is advisable to calculate the volume to be prepared in advance. 12. It is preferable to add the solutions in this order so as to ensure all reagents are present prior to addition of CDKACT peptide biosensor and finally the protein or cell extracts. To save time, a master mix can be prepared and divided between wells. 13. When using the biosensor for the first time, it is advisable to establish the best working conditions by performing a couple

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of dose-dependent experiments with different concentrations of kinase or cell lysates expressing the kinase of interest (10, 20, 30, 40, 50, 100, 150, 200 μg) to determine both the linear range of response and the concentration that yields the greatest biosensor response. Conversely for a fixed concentration of kinase (or cell lysate expressing the kinase of interest), it is advisable to perform an experiment with varying concentrations of biosensor to determine the optimal concentration for the greatest fluorescent response. 14. The excitation and fluorescence emission wavelengths may vary slightly depending on the spectrofluorimeter models and filters. 15. Evaluate fluorescence emission from the well containing the biosensor alone, and adjust acquisition parameters so as to avoid signal saturation by anticipating an estimated 25–50% increase in fluorescence. 16. It is recommended to shake the plate for 10 s at 300 rpm before the first measurement. 17. Generally the lysis buffer has a quenching effect on CDKACT biosensor fluorescence emission over time. 18. It is important to use kinase inhibitors to ensure the biosensor responds and reports appropriately to kinase activity. Prior to the experiment it is advisable to verify whether the inhibitor affects biosensor fluorescence itself, in which case cells should be treated with inhibitor prior to preparation of cell extracts, instead of mixing the inhibitor with untreated cell extracts and biosensor in the well. Alternatively, control experiments can be performed with samples which have been treated with siRNA to silence the kinase of interest or samples in which the kinase is absent or has been knocked out.

Acknowledgments This work was supported by the CNRS (Centre National de la Recherche Scientifique) and a grant from the Region Occitanie and European Regional Development Fund (READYNOV/ FEDER n 2018-003539-01) to M.C. Morris. We acknowledge GL Biochem (Shanghai, China) Ltd. and GENEPEP (St-Jean de Ve´das, France) for synthesis of peptide biosensors. We thank all former members of the “Kinase Biosensors & Inhibitors” group for their contributions to development of fluorescent peptide biosensors: T.N.N. Van, C. Pre´vel, M. Peyressatre, J.A. Gonzalez Vera, C. Tilmaciu, Sebastien Diot, Jessica Soamalala and Arthur Laure.

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References 1. Morris MC (2014) Spotlight on fluorescent biosensors—tools for diagnostics and drug discovery. ACS Med Chem Lett 5:99–101. https://doi.org/10.1021/ml400472e 2. Nhu Ngoc Van T, Morris MC (2013) Fluorescent sensors of protein kinases: from basics to biomedical applications. In: Morris MC (ed) Progress in molecular biology and translational science. Academic, Cambridge, MA, pp 217–274 3. Gonza´lez-Vera JA, Morris MC (2015) Fluorescent reporters and biosensors for probing the dynamic behavior of protein kinases. Proteomes 3:369–410. https://doi.org/10. 3390/proteomes3040369 ˜ as JL, Va´zquez 4. Pazos E, Va´zquez O, Mascaren ME (2009) Peptide-based fluorescent biosensors. Chem Soc Rev 38:3348–3359. https:// doi.org/10.1039/B908546G 5. Zhang J, Campbell RE, Ting AY, Tsien RY (2002) Creating new fluorescent probes for cell biology. Nat Rev Mol Cell Biol 3:906–918. https://doi.org/10.1038/ nrm976 6. Ibraheem A, Campbell RE (2010) Designs and applications of fluorescent protein-based biosensors. Curr Opin Chem Biol 14:30–36. https://doi.org/10.1016/j.cbpa.2009.09.033 7. Palmer AE, Qin Y, Park JG, McCombs JE (2011) Design and application of genetically encoded biosensors. Trends Biotechnol 29:144–152. https://doi.org/10.1016/j. tibtech.2010.12.004 8. Wang Q, Zimmerman EI, Toutchkine A et al (2010) Multicolor monitoring of dysregulated protein kinases in chronic myelogenous leukemia. ACS Chem Biol 5:887–895. https://doi. org/10.1021/cb100099h 9. Lukovic´ E, Gonza´lez-Vera JA, Imperiali B (2008) Recognition-domain focused chemosensors: versatile and efficient reporters of protein kinase activity. J Am Chem Soc 130:12821–12827. https://doi.org/10. 1021/ja8046188 10. Yeh R-H, Yan X, Cammer M et al (2002) Real time visualization of protein kinase activity in living cells. J Biol Chem 277:11527–11532. https://doi.org/10.1074/jbc.M111300200 11. Turner APF (2013) Biosensors: sense and sensibility. Chem Soc Rev 42:3184–3196. https://doi.org/10.1039/C3CS35528D 12. Lavis LD, Raines RT (2008) Bright ideas for chemical biology. ACS Chem Biol 3:142–155. https://doi.org/10.1021/cb700248m

13. Loving GS, Sainlos M, Imperiali B (2010) Monitoring protein interactions and dynamics with solvatochromic fluorophores. Trends Biotechnol 28:73–83. https://doi.org/10.1016/ j.tibtech.2009.11.002 14. Van TNN, Pellerano M, Lykaso S, Morris MC (2014) Fluorescent protein biosensor for probing CDK/cyclin activity in vitro and in living cells. Chembiochem 15:2298–2305. https:// doi.org/10.1002/cbic.201402318 15. Pre´vel C, Pellerano M, Gonza´lez-Vera JA et al (2016) Fluorescent peptide biosensor for monitoring CDK4/cyclin D kinase activity in melanoma cell extracts, mouse xenografts and skin biopsies. Biosens Bioelectron 85:371–380. https://doi.org/10.1016/j.bios.2016.04.050 16. Gonza´lez-Vera JA, Bouzada D, Bouclier C et al (2017) Lanthanide-based peptide biosensor to monitor CDK4/cyclin D kinase activity. Chem Commun 53:6109–6112. https://doi.org/ 10.1039/C6CC09948C 17. Peyressatre M, Laure A, Pellerano M et al Fluorescent biosensor of CDK5 kinase activity in glioblastoma cell extracts and living cells. Biotechnol J 15:e1900474. https://doi.org/10. 1002/biot.201900474 18. Soamalala J, Diot S, Pellerano M et al Fluorescent peptide biosensor for probing CDK6 kinase activity in lung cancer cell extracts. ChemBioChem 22(6):1065-1071.https:// doi.org/10.1002/cbic.202000677 19. Morgan DO (1997) Cyclin-dependent kinases: engines, clocks, and microprocessors. Annu Rev Cell Dev Biol 13:261–291. https://doi. org/10.1146/annurev.cellbio.13.1.261 20. Malumbres M, Barbacid M (2005) Mammalian cyclin-dependent kinases. Trends Biochem Sci 30:630–641. https://doi.org/10.1016/j.tibs. 2005.09.005 21. Malumbres M, Barbacid M (2007) Cell cycle kinases in cancer. Curr Opin Genet Dev 17:60–65. https://doi.org/10.1016/j.gde. 2006.12.008 22. Asghar U, Witkiewicz AK, Turner NC, Knudsen ES (2015) The history and future of targeting cyclin-dependent kinases in cancer therapy. Nat Rev Drug Discov 14:130–146. https:// doi.org/10.1038/nrd4504 23. Peyressatre M, Pre´vel C, Pellerano M, Morris MC (2015) Targeting cyclin-dependent kinases in human cancers: from small molecules to peptide inhibitors. Cancers (Basel) 7:179–237. https://doi.org/10.3390/cancers7010179

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24. Lapenna S, Giordano A (2009) Cell cycle kinases as therapeutic targets for cancer. Nat Rev Drug Discov 8:547–566. https://doi. org/10.1038/nrd2907 25. Malumbres M, Barbacid M (2001) To cycle or not to cycle: a critical decision in cancer. Nat

Rev Cancer 1:222–231. https://doi.org/10. 1038/35106065 26. Arooz T, Yam CH, Siu WY et al (2000) On the concentrations of cyclins and cyclin-dependent kinases in extracts of cultured human cells. Biochemistry 39:9494–9501. https://doi. org/10.1021/bi0009643

Chapter 5 Phosphatase and Kinase Substrate Specificity Profiling with Pooled Synthetic Peptides and Mass Spectrometry Andrew G. DeMarco, Pete E. Pascuzzi, W. Andy Tao, and Mark C. Hall Abstract Reversible phosphorylation is a pervasive regulatory event in cellular physiology controlled by reciprocal actions of protein kinases and phosphatases. Determining the inherent substrate specificity of kinases and phosphatases is essential for understanding their cellular roles. Synthetic peptides have long served as substrate proxies for defining intrinsic kinase and phosphatase specificities. Here, we describe a high throughput protocol to simultaneously measure specificity constants (kcat/KM) of many synthetic peptide substrates in a single pool using label-free quantitative mass spectrometry. The generation of specificity constants from a single pooled reaction provides a rigorous and rapid comparison of substrate variants to help define an enzyme’s specificity. Equally applicable to kinases and phosphatases, as well as other enzyme classes, the protocol consists of three general steps: (1) reaction of enzyme with pooled peptide substrates, each ideally with a unique mass and at concentrations well below KM, (2) analysis of reaction products using liquid chromatography-coupled mass spectrometry (LC-MS), and (3) automated extraction and integration of elution peaks for each substrate/product pair. We incorporate an ionization correction strategy allowing direct calculation of reaction progress, and subsequently kcat/KM, from substrate and product peak areas in a single sample, obviating the need for stable isotope labeling. Peptide consumption is minimal, and high peptide purity and accurate concentrations are not required. Access to a high-resolution LC-MS system is the only nonstandard equipment need. We present an analysis pipeline consisting entirely of established open-source software tools, and demonstrate proof of principle with the highly selective cell cycle phosphatase Cdc14 from Saccharomyces cerevisiae. Key words Synthetic peptides, Phosphatases, Kinases, Quantitative mass spectrometry, Substrate specificity, Label-free mass spectrometry, High throughput enzyme assay, Multiplex substrate reaction, LC-MS assay

1

Introduction Reversible protein phosphorylation regulates virtually all physiological processes, including the cell division cycle, and is controlled by opposing kinase and phosphatase activities. Protein kinases transfer the γ phosphate from ATP to specific serine, threonine, and/or tyrosine side chains within a target protein, while protein phosphatases catalyze the reverse hydrolysis reaction to restore the

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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original amino acid and release inorganic phosphate. The addition or removal of phosphate can have diverse effects on protein structure and function that are key to dynamic control of biological processes. A persistent question is how kinases and phosphatases select specific substrate proteins and amino acid positions to act on. The existence of hundreds of distinct kinase and phosphatase activities in eukaryotic species suggests that these enzymes must act in a highly selective manner by recognizing and binding specific substrate structures. Indeed, the structural and mechanistic basis for substrate selectivity has been defined in detail for some kinases and phosphatases [1–5]. Defining specificity and identifying the direct substrates of kinases and phosphatases continue to be important for understanding their cellular functions. Using full-length protein substrates to characterize kinase and phosphatase specificity is challenging. However, short synthetic peptides are easier to make and assay, particularly for phosphatases where phosphoamino acids can be stoichiometrically incorporated at any desired position. Synthetic peptides and phosphopeptides have been critical reagents in the systematic characterization of kinase and phosphatase specificity over the past several decades [6–9]. Since kinase specificity is largely defined by interactions with substrate features adjacent to the phosphorylation site [1, 10], short synthetic peptides are useful substrate proxies. This is also true for some phosphatases, including the protein tyrosine phosphatase family [11, 12], although the major Ser/Thr phosphatase families often rely heavily on substrate interactions involving regulatory subunits distal from the catalytic site [4, 13]. Nonetheless, past use of synthetic phosphopeptides has still been useful in revealing specificity information for these enzymes [6] and recent studies, for example of the PP2A-B55 complex [14], suggest that they also can recognize substrate features relatively close to the phosphorylation site that could be accommodated within synthetic peptides. Synthetic peptide libraries are now routinely obtained commercially, with lengths up to ~25 amino acids. A standard format is the positional scanning library, in which individual positions of a starting sequence, often from a known substrate, are systematically changed to different amino acids. Other formats are possible as well, including randomization of select positions or sequence collections derived from naturally existing proteins. All of these formats can help identify preferred substrate recognition features and consensus motifs. Synthetic peptides have been used to profile kinase and phosphatase specificities in both low and high throughput assay formats. Low-throughput assays include spectroscopic monitoring of phosphate release by phosphatases using malachite green dye or γ[32P] incorporation by kinases [7, 15, 16]. High throughput methods fall

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into two general categories. In one, immobilized peptide arrays are assayed using imaging systems to detect radioactivity, fluorescence, or chemiluminescence as indicators of enzyme activity on each substrate in the array [9]. In the other, LC-MS is used to detect products after reaction of pooled substrate peptides in solution based on the resulting mass changes [17–20]. Peptide arrays require that substrates be anchored to a surface, potentially constraining enzyme binding, and rarely provide rigorous rate measurements for substrate comparisons. The specificity constant, kcat/ KM, is an effective and widely used measure of substrate preferences for an enzyme [21]. LC-MS methods can provide specificity constants for quantitative substrate comparisons under appropriate conditions but are rarely used to do so. The method we describe here is based on prior LC-MS assay development by several labs and allows simultaneous measurement of kcat/KM for many peptide substrates in a single pooled reaction using label-free quantitation and freely available software tools. Craik and colleagues initially made specificity constant measurements from label-free LC-MS extracted ion chromatograms (XICs) of pooled synthetic peptide substrate libraries reacted with peptidases [22]. This required generating reaction progress curves for each substrate, and the method was subsequently applied to kinases by the same group [23]. The group of Minliang Ye extended this work by formally establishing pooled substrate reaction conditions that render competition negligible and allow calculation of accurate kcat/KM values from the measurement of the fractions of substrates consumed at a given time point [24]. In this study, dimethyl stable isotope labeling was employed to calculate substrate consumption by comparing substrate signals between time points. We wanted to avoid stable isotope labeling to make our method simple to implement. The Kirschner lab described a method for calculating, and correcting for, ionization differences between phosphorylated and unphosphorylated peptide pairs that allows direct calculation of the phosphorylated fraction from the corresponding XICs [25]. Combining these approaches, it is possible to calculate specificity constants for each peptide substrate in complex pools from single time points without the need for isotope labeling. Our protocol consists of the following steps. First, the mixed substrate pool is reacted with enzyme. Then, reaction products are desalted and subjected to LC-MS/MS analysis with datadependent acquisition and database searching to create a spectral library. Finally, LC-MS peaks are extracted, integrated and matched to the spectral library sequences. Output substrate and product signals are corrected for ionization differences and used to calculate kcat/KM values, which can be directly compared to reveal information on intrinsic specificity.

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Materials We recommend HPLC-grade water and solvents for all steps, and low retention microcentrifuge tubes and pipet tips to minimize sample adsorption. 1. Synthetic peptides or phosphopeptides (see Note 1). 2. Purified phosphatase or kinase of interest and an appropriate reaction buffer (see Note 2). 3. Centrifugal evaporator or lyophilizer. 4. Broad specificity phosphatase or kinase preparation (see Note 3). 5. 10% formic acid. 6. HPLC solvent A: 3% acetonitrile 0.3% formic acid. 7. HPLC solvent B: 80% acetonitrile, 0.3% formic acid. 8. Methanol. 9. Desalting wash solvent: 0.1% trifluoroacetic acid. 10. Desalting elution solvent: 0.1% trifluoroacetic acid in 60% acetonitrile. 11. C18 desalting tips (10 μg binding capacity) with microcentrifuge adaptors (e.g., Thermo Fisher Scientific # PI84850). 12. NanoHPLC-coupled mass spectrometer with high-resolution mass analyzer (see Note 4). 13. MaxQuant software (https://www.maxquant.org/). 14. Skyline software (https://skyline.ms/project/home/soft ware/Skyline/begin.view). 15. R (https://cloud.r-project.org/) and R studio (https:// rstudio.com/products/rstudio/download/) or comparable spreadsheet data analysis software.

3

Methods Subheadings 3.1 and 3.2 only need to be performed once, prior to first assay implementation, but require some of the full assay instructions from Subheadings 3.3–3.5 below.

3.1 Defining Instrument Linear Response Range

It is useful to initially define the linear dynamic range for substrate and product detection in the mass spectrometer. The upper limit establishes the maximum amount of peptide for injection. The lower limit reflects the minimal amount of product formation required for accurate measurement. 1. Create a twofold dilution series of the peptide substrate library in HPLC solvent A or ultrapure water. The appropriate range

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1x1010

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Fig. 1 Dynamic range determination. Mean integrated MS1 peak areas for 13 phosphopeptides used as a substrate set for Cdc14 were plotted as a function of the injected amount of each peptide. Linear regression was performed on the data points from 2.5 to 1000 fmol. The horizontal line essentially represents the instrument noise level and helps approximate the minimal product amount and signal required for detection. Note that individual peptide intensities may vary substantially from the mean values

may vary with the instrument. We suggest 1–5000 fmol of total peptide as a starting point. 2. Analyze each dilution using the LC-MS/MS method and data analysis procedures described below in Subheadings 3.4 and 3.5. 3. Plot the mean integrated peptide area as a function of injection amount. To thoroughly define the dynamic range, the plot should begin to plateau at the top and bottom (Fig. 1). If not, adjust the dilution series until the full linear range is visible. 3.2 Calculate Ionization Correction Factors for Substrate and Product Peptide Pairs

Potential ionization differences between phosphorylated and unphosphorylated forms of a peptide prevent reliable quantitative comparison of their intensities. These ionization differences can be measured empirically, and correction factors derived as described previously [25], allowing direct quantitative comparison and calculation of the fraction of substrate consumed in a reaction. The ionization correction factor (ICF) is calculated just once for each phosphopeptide/peptide pair provided the LC-MS conditions remain constant. ICF calculation requires measurement of the

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ICF =

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Intensity

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Fig. 2 Ionization correction. Representative XICs for a substrate peptide (black) and its product (red) at two different intensity ratios, A and B, generated from partial enzymatic conversion. S and P are the integrated peak areas for substrate and product species, respectively. The equation for using S and P from conditions A and B to calculate ICF, which is the ratio of the change in the linked S and P signals between the two states, is adapted from Steen et al. 2005 [25]

relative signals of phosphorylated and unphosphorylated forms of substrate/product pairs at two different abundance ratios differing by at least 10% (Fig. 2) [25]. We recommend using ratios between 5:95 and 95:5. Since the starting peptide libraries exclusively contain substrates, the best way to achieve the required ratios is by partial enzymatic conversion to products, for example, with a broad specificity enzyme activity. Alternatively, the target enzyme itself might be suitable with an appropriate time course and/or enzyme titration to achieve the required product conversion for all substrates. Perform in triplicate to assess variation. 1. Treat the peptide library with enzyme and stop aliquots of the reaction mixture at a series of time points, as described in Subheading 3.3. This may require optimization (enzyme concentration and/or time) to achieve two different substrate– product values in the 5:95 to 95:5 range. 2. Desalt reaction products as described in Subheading 3.4. 3. Collect LC-MS/MS data using the same method that will be used in Subheading 3.4. 4. Identify peptides as described in Subheading 3.5.1 and export integrated peak areas for each using Skyline as described in Subheading 3.5.2. 5. Calculate ICF for each peptide pair using the equation in Fig. 2 [25] that relates the change in the phosphorylated peptide signal to the change in the unphosphorylated peptide signal. ICF values are then used in the final calculations as directed in Subheading 3.5.3.

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Enzyme Assay

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Choose substrate concentrations far below the lowest KM, if known. When the KM values are not known, try starting with 100 nM each peptide, well below the KM values for good peptide substrates of many kinases and phosphatases. If desired, test if substrate concentrations are appropriate by measuring specificity constants at several different substrate concentrations (e.g., 50, 100, 200 nM). If substrate concentrations are appropriate, the specificity constant values will not change [24]. Three independent trials are recommended. 1. Create a mixture of all peptide substrates at 10
their final concentration in reaction buffer. 2. Make a 10
enzyme solution in reaction buffer. Keep on ice. 3. For each reaction being performed, label a clean microcentrifuge tube. Include a “no enzyme” control reaction. Decide how many time points to collect (see Note 5). Label a microcentrifuge tube for each time point and add 20 μl of 10% formic acid as a quenching reagent. Calculate a total reaction volume as 60 μl per time point plus 20 μl extra (e.g., if three time points are to be collected then total reaction volume ¼ 200 μl). 4. For the “no enzyme” control, mix 72 μl reaction buffer and 8 μl 10
peptide mix. 5. For each reaction, mix reaction buffer, 10
peptide substrate mixture, and 10
enzyme solution at an 8:1:1 ratio to give the total reaction volume. Combine the buffer and 10
peptide mixture first, then initiate reactions by adding the 10
enzyme solution. Place reactions in an incubator for temperature control. 6. Stop each reaction at the selected time points by transferring 60 μl aliquots into the labeled microcentrifuge tubes containing formic acid and mixing well (see Note 6).

3.4

LC-MS Analysis

Samples must be desalted before LC-MS analysis to ensure good LC performance and electrospray ionization. We recommend C18 resin packed into pipet tips, available from many vendors, for simplicity and speed. They are used as mini spin columns. All centrifugation steps are performed at 1000
g in a standard microcentrifuge. 1. Place a C18 desalting spin tip for each sample in a 2 ml microcentrifuge collection tube using plastic tube adaptors and wet resin by adding 20 μl of 100% methanol and spinning for 1 min (see Note 7). 2. Equilibrate the C18 tip with 20 μl of wash solvent and spin for 1 min. 3. Load sample peptides from quenched reactions, 40 μl at a time, onto the C18 tip and spin each time until the entire sample has

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passed through. If desired, repeat this loading step a second time. Discard the flow-through. 4. Wash the C18 tip twice with 20 μl of desalting wash solvent, spinning for 1 min each time. Transfer the tip and adaptor to a clean collection tube. 5. Elute peptides by adding 20 μl of desalting elution solvent to the C18 tip and spinning for 1 min. Repeat a second time with an additional 20 μl of elution solvent. 6. Dry the eluted peptides in a centrifugal evaporator and store at 80  C. 7. Immediately prior to LC-MS/MS, reconstitute dried peptides in 10 μl HPLC solvent A. 8. Set up a data-dependent LC-MS/MS acquisition method. Methods will vary depending on peptide library complexity and sequence content. A generally useful method is ten MS/MS product ion scans per MS survey scan with a 30–60 min LC gradient of 3–40% acetonitrile using HPLC solvents A and B (see Note 8). 9. Run the method, injecting enough sample to generate signals near the upper end of the dynamic range. 3.5

Data Analysis

3.5.1 Substrate and Product Peak Identification

Data analysis consists of three steps: (1) substrate and product peak identification, (2) integration of MS peaks identified in step 1, and (3) kcat/KM calculation for each substrate. Various programs can be used to accomplish these steps. We describe a protocol that uses MaxQuant (MQ) for step 1, Skyline for step 2 (see Note 9), and R studio for step 3. While it is formally possible to set up simultaneous analyses of multiple experiments (e.g., time points or enzyme concentrations), each with replicate trials, for simplicity our instructions are written just for the simultaneous analysis of all replicates of individual reactions. 1. Unless indicated below, use default settings for MQ database search. 2. Create a custom protein sequence database in Fasta format. Begin with a Saccaromyces cereivisae .fasta proteome file (see Note 10). Open in a text editor like Notepad, and add all peptide sequences as individual protein entries, giving each a unique name or number as the entry header (Fig. 3). The S. cerevisiae sequences act as a background proteome to facilitate meaningful search statistics. 3. Copy raw MS files and the custom sequence database file to a new folder for the MQ output. 4. Open MQ and load raw MS files for each replicate reaction by clicking tab ! < Load>, and then navigating to the raw files and selecting them.

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>peptide_1 HTSPIKSIG >peptide_2 HTTPIKSIG >peptide_3 HTYPIKSIG >peptide_4 HTSAIKSIG >peptide_5 HTSGIKSIG >peptide_6 HTSGIKRIG >peptide_7 HTSPIASIG >peptide_8 HTSPIRSIG >peptide_9 HTSPKASIG >peptide_10 HTSPRKSIG >peptide_11 HTSPIAKIG >peptide_12 HTSPIKKIG >peptide_13 HTSPPKSIG >sp|P36166|PXL1_YEAST Paxillin-like protein 1 OS=Saccharomyces cerevisiae (strain ATCC 204508 / S288c) OX=559292 GN=PXL1 PE=1 SV=3 MYNSIYGSPFPKINPKVRYKTALERAGFDTKPRNPFSSQRNASTGSLQASVKSPPITRQR NVSAAPSVPVTMKSAYTASSKSAYSSVKGESDIYPPPVLENSERRSVTPPKNSNFTSSRP SDISRSISRPSERASQEDPFRFERDLDRQAEQYAASRHTCKSPANKEFQAADNFPFNFEQ EDAGNTEREQDLSPIERSFMMLTQNDTASVVNSMNQTDNRGVLDQKLGKEQQKEESSIEY ESEGQQEDENDIESLNFEPDPKLQMNLENEPLQDDFPEAKQEEKNTEPKIPEINVTRESN TPSLTMNALDSKIYPDDNFSGLESSKEQKSPGVSSSSTKVEDLSLDGLNEKRLSITSSEN VETPYTATNLQVEQLIAQLDDVSLSRNAKLDMNGNCLNAVDRKASRFKKSSAYLSGYPSM DIPVTQQTSIVQNSNTNLSRQTILVDKGDVDEDAPSESTTNGGTPIFYKFKQSNVEYSNN EGMGSQETFRTKLPTIEALQLQHKRNITDLREEIDNSKSNDSHVLPNGGTTRYSSDADYK ETEPIEFKYPPGEGPCRACGLEVTGKRMFSKKENELSGQWHRECFKCIECGIKFNKHVPC YILGDEPYCQKHYHEENHSICKVCSNFIEGECLENDKVERFHVDCLNCFLCKTAITNDYY IFNGEIPLCGNHDMEALLKEGIDNATSSNDKNNTLSKRRTRLINFN >sp|P03962|PYRF_YEAST Orotidine 5'-phosphate decarboxylase OS=Saccharomyces cerevisiae (strain ATCC 204508 / S288c) OX=559292 GN=URA3 PE=1 SV=2 MSKATYKERAATHPSPVAAKLFNIMHEKQTNLCASLDVRTTKELLELVEALGPKICLLKT HVDILTDFSMEGTVKPLKALSAKYNFLLFEDRKFADIGNTVKLQYSAGVYRIAEWADITN AHGVVGPGIVSGLKQAAEEVTKEPRGLLMLAELSCKGSLATGEYTKGTVDIAKSDKDFVI GFIAQRDMGGRDEGYDWLIMTPGVGLDDKGDALGQQYRTVDDVVSTGSDIIIVGRGLFAK GRDAKVEGERYRKAGWEAYLRRCGQQN

Fig. 3 Custom Fasta sequence database file. Example Fasta file format, shown in Notepad, for adding substrate peptide sequences as protein entries to facilitate their identification in the MQ LC-MS/MS database search and subsequent assembly into a Skyline spectral library. The peptides listed at top were used with budding yeast Cdc14 phosphatase to generate the data shown in Fig. 5. The proteins at bottom are the beginning of the S. cerevisiae proteome Fasta database file, obtained from www.uniprot.org

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5. Load the custom Fasta file by selecting tab ! ! < Add>, navigating to the file, and clicking (see Note 11). 6. In most cases, a new protease cleavage specificity must be added to the MQ Proteases table to ensure the peptide sequence entries in the Fasta file are not cleaved in the database search. This would prevent their identification. Because Cys is rarely used in synthetic peptides, adding a hypothetical protease that cleaves only after Cys is often a suitable solution. In MQ, select tab ! ! to open a specificity table and initiate a new entry. Provide a name and description and, importantly, click the C+ button in the far-left column of the table to set Cys.Xxx as the cleavage specificity. Click ; a new row should appear in the protease table. Finally, click to permanently save the entry. If synthetic peptides do contain Cys, then a different protease specificity must be designed that is not represented in the peptide library. 7. Set the following parameters in the MQ < Group-specific parameters> tab: (a) —remove any preselected proteases and select the custom protease from step 6 above. (b) —remove any preexisting Variable modification selections from the right window and add Phospho (ST) or Phospho (STY) as a Variable modification. For Fixed modifications, select any modifications to standard peptide termini, for example, Amidated (C-term) if an amide group was included at peptide C-termini during synthesis. This is essential for MQ to compute correct masses of the database peptide sequences (see Note 12). (c) —choose the closest match to the instrument used for the assay. Check the First search peptide tolerance and make sure it is appropriate based on the instrument resolution and accuracy. If there is an exact instrument type match and the instrument is properly calibrated this setting should not need editing. 8. Set the following parameters in the MQ tab: (a) —make sure “Min. peptide length” is lower than the shortest library peptide. Deselect the box for “Include contaminants.” 9. Click at bottom left to initiate the MQ search. When the search is done, find and open the peptides.txt file in the “combined” folder ! “txt” subfolder using spreadsheet software to evaluate the results. Sort the entries by the PEP column, which provides a p-value-like probability score for the match significance. Most of the entries should be the synthetic

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peptides and have PEP scores well below 0.01 if LC-MSMS data quality is high. Confirm that all expected library peptides are represented in the search results. Note that any peptides not identified in the MQ search will not by quantified by Skyline. 10. Find and copy the msms.txt file in the “combined” folder ! “txt” subfolder and mqpar.xml file from the parent folder. Paste these into a new folder along with the raw MS files for Skyline analysis. 3.5.2 MS1 Peak Integration

1. Open Skyline, select to create a new file, and save it in the same new folder containing the two MQ output files and MS raw data files. 2. Click ! ! , make sure the “Build” radio button is selected, click and select the msms.txt file. Make sure the “iRT standard peptides” pull-down menu is set to “None” and the “DDA with MS1 filtering” radio button is selected. Click . 3. In the resulting Extract Chromatograms window, the raw MS files should be listed. Confirm that “Many” is selected in the drop-down menu at botom and click . 4. If the “Add Modifications” window appears, check each modification used in the MQ search and click . 5. In the “Configure Full-Scan Settings” window set “Precursor charges:” to “2,3” at a minimum (4 can be included if warranted, depending on the size and sequence content of peptides). “Count” should be selected in the “Isotope peaks included:” drop-down menu. In the “Precursor mass analyzer:” drop-down menu select the mass analyzer used. The “Resolving power:” window should automatically populate with a suitable resolution. This can be adjusted if necessary. Set “Peaks:” to 3. Leave the “Use high-selectivity extraction” box unchecked. “Retention time filtering” should have the top radio button selected with 5 in the input window. Click . 6. In the Import FASTA window use to select a Fasta file with only the peptide substrate sequences as entries (create from the custom Fasta MQ database file by removing the background proteome). Select the same enzyme specificity used in the MQ search. If a custom cleavage specificity was used, select in the “Enzyme” drop-down menu and enter the required information to define the custom specificity matching the MQ search and click (see Note 13). 7. In the next window set “Min peptides per protein” to 1. Click to import data from the raw MS files. 8. XICs are generated from the raw MS file(s) for each spectral library entry (generated from the MQ msms.txt file) that matches an entry from the provided Fasta file (Fig. 4) and can be reviewed by cycling through the identified targets list in the

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Time point X

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0 18.1

18.3

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18.7

18.9

Retention Time (min)

17.7

17.9

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18.3

Retention Time (min)

Fig. 4 Calculation of substrate consumption and kcat/KM. Representative Skyline screenshots showing the conversion of starting substrate at time 0 (top left) into product at arbitrary time X (top right). The MS1 XICs for substrate and product species are overlaid in the top panels. In the bottom panels the substrate and product XICs from time X are shown separately (+2 charge state only) with the first three isotope peaks (colored lines) overlaid, and MS/MS ID markers linking the peak and its retention time to a spectral library match. Black arrowheads (with LC retention time and precursor mass error in ppm labeled above) indicate that Skyline has

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left-hand panel (each independent sample will have a tab in the main XIC window). For this method, precursor to product ion transitions are ignored; only the precursor XICs are needed. In principle, each integrated XIC peak should be manually validated for correct isotopic distribution ( menu ! Peak Areas), a unique spectral library match ( menu ! Library Match ! click on ID markers in XIC window), and appropriate integration boundaries (vertical dotted lines flanking peaks in XIC window). However, this may be impractical for large libraries. XIC peak integration boundaries can be adjusted by moving the dotted lines (see Note 14). 9. Export data in comma-separated values (.csv) format as follows. Initially, use ! ! ! ! to create a custom report format called “Precursor Results.” It should include the following data columns: Protein Name, Peptide Modified Sequence, Replicate, Precursor Mz, Total Area MS1, and Total Background MS1. Check the boxes for Protein Name and Replicates when the Edit Report window first opens. Find the Peptide Modified Sequence checkbox by clicking “+” to open the Peptides category. The Precursor Mz checkbox is found under Precursors. Further opening Precursor Results will reveal checkboxes for Total Area MS1 and Total Background MS1. Click OK to save the new Report format then select it and click and choose a file name and destination. Once the Report format is created it can be directly selected in future experiments without editing. 3.5.3 kcat/KM Calculation

The general steps needed are described first and then specific instructions for using our method in R follow. 1. Subtract background MS1 peak area signal (time 0) from MS1 area of each product peptide. 2. Sum the MS1 peak areas for each peptide charge state. 3. Correct for ionization difference between substrate and product species by multiplying product MS1 peak area by the appropriate ICF value calculated in Subheading 3.2. 4. Calculate the fraction of substrate remaining (Sr) from the substrate and product MS1 areas using the first equation in Fig. 4.

ä Fig. 4 (continued) chosen and integrated the peak. Dotted vertical lines indicate integration boundaries. The equations in the red box are used to calculate the fraction of substrate remaining (Sr) from the integrated, ionization-corrected areas of substrate (S) and product (P) species at time X (top equation) and subsequently kcat/KM from Sr, total molar enzyme concentration ([E]tot), and reaction time (t) in seconds (bottom equation)

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5. Calculate the kcat/KM for each substrate using the second equation in Fig. 4, Sr from the previous step, the total molar enzyme concentration, and reaction time in seconds. 6. Determine mean values and variation (e.g., standard deviation) and display results as desired. An example dataset using a small phosphopeptide collection with the budding yeast Cdc14 phosphatase is presented in Fig. 5. Cdc14 exhibits strong selectivity for certain substrate sequence features [26, 27]. Consequently, kcat/ KM varies by >4 orders of magnitude for substrates in this collection, despite the similar overall sequences. This example highlights the broad range of activities accessible with this method and its power in comparing many substrates simultaneously. 7. Specific instructions for using R for kcat/KM calculation (see Note 15): (a) Download and install R and R studio. Refer to the materials section for links to download these programs. (b) In R Studio, install the tidy verse package by typing the following into the console: install.packages(“tidyverse”). This is the only package needed to run the calculations script. (c) Download a script for performing the kcat/KM calculations in R studio here: https://github.com/ thehalllabpurduebchm?tab¼repositories. There is one for use with phosphatases and one for kinases that differ only by the definition of substrate as the unphosphorylated or phosphorylated species. (d) Create a new directory folder for each enzyme concentration and reaction time data set. Save a copy of the R script in each. For ICF calculations, continue to step e. If ICF calculations exist from previous experiments, skip to step f. (e) Open the R script in R Studio. Create an ICF template from your skyline export file by clicking the green run arrow at right at line 12 (“Run Current Chunk”). When prompted, load the Skyline export file. A file titled “ICF_template.csv” in folder “ICF_Template” will be created. Open the file and enter the calculated ICF value from Subheading 3.2 for each product and 1 for its cognate substrate in the “ICF” column. Save the file. (f) In the script, enter reaction time (min) at line 66 and enzyme concentration (nM) at line 68. (g) Create a new output file.csv name at line 76 for the calculation results. (h) Click the green run arrow at right on line 46. When prompted, select the Skyline export file and then the ICF_template.csv file.

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A

B

Substrate

kcat/KM (M-1 s-1)

1 2 3 4 5 6 7 8 9 10 11 12 13

25000 ± 4300 4.4 ± 0.7 155 ± 12 35 ± 4 16 ± 3 31 ± 1 173 ± 26 3400 ± 800 90 ± 7 384 ± 20 20300 ± 6000 56000 ± 5000 170 ± 26

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1000

100

10

1

1

2

3

4

5

6

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8

9 10 11 12 13

Substrate Fig. 5 Application of method to study substrate preference of Cdc14 phosphatase. The method was applied in full to the Cdc14 phosphatase from S. cerevisiae using the peptide set shown in Fig. 3 to illustrate typical final results as numerical values (a) and graphical comparison (b). This peptide collection begins with a sequence from a known Cdc14 substrate, the Yen1 endonuclease [27] and varies individual amino acids, including the phosphoamino acid itself, at positions known to be important for recognition by Cdc14. Data are means with standard deviations from three independent reactions. Note that kcat/KM values cover a 4+ log range (14,000-fold difference between best and worst substrates), demonstrating the high selectivity of Cdc14 and the broad

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(i) The R script will export a file with calculations to the folder named “output,” containing individual Sr and kcat/KM values and the mean, standard deviation, and coefficient of variation for Sr and kcat/KM for each substrate.

Notes 1. Desalted synthetic peptide libraries can be obtained commercially from many vendors. Crude, desalted material is generally acceptable. Store high concentration peptide stocks in ultrapure water or buffered solutions at 80  C (e.g., 10 mM in 10 mM HEPES pH 7.5). Organic solvents like DMSO or acetonitrile may be needed to aid the solubilization of some peptides. Follow vendor recommendations on solubilization conditions. Peptide concentrations can be estimated based on the mass yield. The specificity constant calculations do not require accurate substrate concentrations. Pooled peptide mixes for reactions can be stored in small aliquots at 80  C or dried for long-term storage. Peptide length, number, and sequence variations are up to the investigator. We offer the following guidelines. For kinases, peptides should contain a single possible phosphorylation site (Ser, Thr, or Tyr) to avoid problems identifying which position is phosphorylated. We generally synthesize peptides with an amide group at the C-terminus. Design peptides with unique masses whenever possible. Identical or near-identical masses are not required, as long as LC retention times are unique. Retention times will be determined in the LC-MS/MS analysis. In cases where isobaric peptides cannot be distinguished chromatographically, the library can be split into multiple pools for parallel reactions. ä

4

Fig. 5 (continued) measurement range of the method. Reactions were performed in 25 mM HEPES pH 7.5, 2 mM tris(2-carboxyethyl)phosphine, 1 mM EDTA, and 150 mM NaCl. [E]Total varied between 1 nM and 1 μM. Each substrate was 375 nM (Lowest KM value ¼ ~50 μM). LC-MS settings: LTQ Orbitrap Velos Pro coupled to EASY-nano LC 1000 (Thermo Fisher Scientific). 750 fmol each peptide were injected on a homemade Prontopearl C18 (2.2 μm, 100 Å pores; Bischoff Chromatography) column (45 cm
360 μm OD
75 μm ID) in HPLC solvent A and eluted at 250 nL/min with a multistep acetonitrile gradient (0–35% solvent B over 10 min, 35–50% over 5 min, 50–95% over 5 min). MS1 mass range ¼ 200–1100 m/z; 60,000 Hz resolution at 400 m/z; ten MS/MS scans per cycle using normalized collision energy of 35% and 12 s dynamic exclusion. Note also that in our experience low throughput phosphatase assays using malachite green-based phosphate detection are not sensitive enough to directly measure the second order rate constant kcat/KM at substrate concentrations well below KM.

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2. Optimal buffer conditions will be unique for each enzyme and should be determined by the investigator. Avoid nonionic detergents that are difficult to remove and interfere with LC-MS analysis. 3. Broad specificity phosphatase or kinase preparations can be obtained commercially. Lambda protein phosphatase or alkaline phosphatase work well. We used lambda phosphatase (New England Biolabs) in 50 mM HEPES pH 7.5, 100 mM NaCl, 2 mM DTT, 1 mM MnCl2. For kinases, custom blends of different recombinant kinases can be created. The key is that significant conversion of each peptide substrate to product is achieved. Data collected with the enzyme being studied can also be used if sufficient product can be generated for each substrate. 4. In the absence of LC-MS experience mass spectrometry core facilities, available at many institutions, are a good option for assistance with method development and sample analysis. No specific instrument type is required; however, TOF and orbitrap analyzers are preferable because of their high resolution. 5. Appropriate enzyme concentrations and times must be empirically determined. Multiple reactions with different enzyme concentrations may be needed to ensure sufficient product formation for all substrates (e.g., see Fig. 5). Collect a wide range of time points. While a single time point is sufficient to calculate kcat/KM, a range of timepoints will help ensure appropriate reaction progress is obtained for all substrate peptides and will allow for validation of kcat/KM measurements. A time 0 sample (i.e., no enzyme control) is important to establish baseline product signals in the substrate mixture that must be subtracted from reaction time points. 6. Reaction and aliquot volumes can be varied to ensure that sufficient peptide is available after desalting for triplicate LC-MS injections with peptide signal near the top of the dynamic range defined in Subheading 3.1. The 60 μl aliquots used here were appropriate for our reactions with 375 nM each substrate peptide and our LC-MS system. 7. Extra spins or extended spin time may be required to completely move the sample through the desalting tips. Ensure the liquid is completely cleared from the tip before moving to the next step. 8. The MS/MS method must generate product ion spectra for all substrate and product peptides so that they are included in the Skyline spectral library and can be matched to XICs. Depending on library complexity, the LC gradient can be shortened or lengthened and other method parameters changed, including number of product ion scans per cycle, acquisition times for product ion scans, and dynamic exclusion window.

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9. A detailed protocol guide for MQ use has been published [28]. Refer to the Skyline tutorial “MS1 scan filtering” for a useful tutorial that explains the settings and parameters listed here: https://skyline.ms/wiki/home/software/Skyline/page. view?name¼tutorial_ms1_filtering. These instructions are valid as of Skyline version 4.2.0 and MQ version 1.6.10. Software formats change over time and these instructions may require modification for newer versions. 10. Fasta proteome sequence files can be obtained from www. uniprot.org. Any species will work as the background proteome source. 11. To ensure correct formatting of the custom Fasta file, test MQ’s ability to read it first by selecting the option in the tab where the Fasta library file is loaded. 12. If a fixed modification is not present in the MQ list, it can be added in the tab ! . Note that if a new modification is added for MQ analysis, it will be saved in a modifications.xml file (MaxQuant folder ! bin ! conf) and must be provided to Skyline along with the other MQ output files. 13. The original custom Fasta file can be used, but any hits from the background proteome will also be included in the Skyline target list and will need to be manually deleted. In addition, the target list may include modified forms of peptides that do not exist in the peptide library (e.g., different phosphorylation sites on phospopeptides). These can be manually selected in the target window and deleted if desired. 14. Identity of each peptide can be confirmed based on (a) parent mass, (b) sequence-specific product ions from MSMS spectra, (c) LC retention time, and (d) correct isotopic distribution for the parent. Manual confirmation is recommended initially for new assays but is not required with repeated use of libraries provided LC-MS/MS conditions do not change. 15. Our R scripts are one option for performing calculations that help automate some of the steps. The scripts are currently built to analyze three replicate trials of a single experiment. The number of replicates can be edited, if desired, in lines 133 and 152. For example, add a “T4” after “T3” if a fourth replicate exists. Be sure to maintain the same format and syntax. References 1. Songyang Z, Lu KP, Kwon YT, Tsai LH, Filhol O, Cochet C, Brickey DA, Soderling TR, Bartleson C, Graves DJ, DeMaggio AJ, Hoekstra MF, Blenis J, Hunter T, Cantley LC (1996) A structural basis for substrate

specificities of protein Ser/Thr kinases: primary sequence preference of casein kinases I and II, NIMA, phosphorylase kinase, calmodulin-dependent kinase II, CDK5, and Erk1. Mol Cell Biol 16(11):6486–6493

Phosphatase & Kinase Specificity Profiling by LC-MS 2. Brown NR, Noble MEM, Endicott JA, Johnson LN (1999) The structural basis for specificity of substrate and recruitment peptides for cyclin-dependent kinases. Nat Cell Biol 1 (7):438–443 3. Jia Z, Barford D, Flint A, Tonks N (1995) Structural basis for phosphotyrosine peptide recognition by protein tyrosine phosphatase 1B. Science 268(5218):1754–1758 4. Peti W, Nairn AC, Page R (2013) Structural basis for protein phosphatase 1 regulation and specificity. FEBS J 280(2):596–611 5. Gray CH, Good VM, Tonks NK, Barford D (2003) The structure of the cell cycle protein Cdc14 reveals a proline-directed protein phosphatase. EMBO J 22(14):3524–3535 6. Pinna LA, Donella-Deana A (1994) Phosphorylated synthetic peptides as tools for studying protein phosphatases. Biochim Biophys Acta 1222(3):415–431 7. Kemp BE, Pearson RB (1991) Design and use of peptide substrates for protein kinases. Methods Enzymol 200:121–134 8. Songyang Z, Blechner S, Hoagland N, Hoekstra MF, Piwnica-Worms H, Cantley LC (1994) Use of an oriented peptide library to determine the optimal substrates of protein kinases. Curr Biol 4(11):973–982 9. Thiele A, Stangl GI, Schutkowski M (2011) Deciphering enzyme function using peptide arrays. Mol Biotechnol 49(3):283 10. Ubersax JA, Ferrell JE Jr (2007) Mechanisms of specificity in protein phosphorylation. Nat Rev Mol Cell Biol 8(7):530–541 11. Cho H, Krishnaraj R, Itoh M, Kitas E, Bannwarth W, Saito H, Walsh CT (1993) Substrate specificities of catalytic fragments of protein tyrosine phosphatases (HPTP beta, LAR, and CD45) toward phosphotyrosylpeptide substrates and thiophosphotyrosylated peptides as inhibitors. Protein Sci 2(6):977–984 12. Zhang Z-Y (2002) Protein tyrosine phosphatases: structure and function, substrate specificity, and inhibitor development. Annu Rev Pharmacol Toxicol 42(1):209–234 13. Slupe AM, Merrill RA, Strack S (2011) Determinants for substrate specificity of protein phosphatase 2a. Enzyme Res 2011:398751 14. Cundell MJ, Hutter LH, Nunes Bastos R, Poser E, Holder J, Mohammed S, Novak B, Barr FA (2016) A PP2A-B55 recognition signal controls substrate dephosphorylation kinetics during mitotic exit. J Cell Biol 214(5):539–554 15. McAvoy T, Nairn AC (2010) Serine/threonine protein phosphatase assays. Curr Prot Mol Biol 92(1):18.18.1–18.18.11

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16. Powers BL, Melesse M, Eissler CL, Charbonneau H, Hall MC (2016) Measuring activity and specificity of protein phosphatases. In: Coutts AS, Weston L (eds) Cell cycle oscillators: methods and protocols. Springer, New York, NY, pp 221–235 17. Kubota K, Anjum R, Yu Y, Kunz RC, Andersen JN, Kraus M, Keilhack H, Nagashima K, Krauss S, Paweletz C, Hendrickson RC, Feldman AS, Wu C-L, Rush J, Ville´n J, Gygi SP (2009) Sensitive multiplexed analysis of kinase activities and activity-based kinase identification. Nat Biotechnol 27(10):933–940 18. Barber KW, Miller CJ, Jun JW, Lou HJ, Turk BE, Rinehart J (2018) Kinase substrate profiling using a proteome-wide serine-oriented human peptide library. Biochemistry 57 (31):4717–4725 19. Kettenbach AN, Wang T, Faherty BK, Madden DR, Knapp S, Bailey-Kellogg C, Gerber SA (2012) Rapid determination of multiple linear kinase substrate motifs by mass spectrometry. Chem Biol 19(5):608–618 20. Huang Y, Thelen JJ (2012) KiC assay: a quantitative mass spectrometry-based approach. In: Marcus K (ed) Quantitative methods in proteomics. Humana Press, Totowa, NJ 21. Fersht A (1985) Enzyme structure and mechanism, 2nd edn. W.H. Freeman, New York, NY 22. O’Donoghue AJ, Eroy-Reveles AA, Knudsen GM, Ingram J, Zhou M, Statnekov JB, Greninger AL, Hostetter DR, Qu G, Maltby DA, Anderson MO, Derisi JL, McKerrow JH, Burlingame AL, Craik CS (2012) Global identification of peptidase specificity by multiplex substrate profiling. Nat Methods 9 (11):1095–1100 23. Meyer NO, O’Donoghue AJ, SchulzeGahmen U, Ravalin M, Moss SM, Winter MB, Knudsen GM, Craik CS (2017) Multiplex substrate profiling by mass spectrometry for kinases as a method for revealing quantitative substrate motifs. Anal Chem 89 (8):4550–4558 24. Deng Z, Mao J, Wang Y, Zou H, Ye M (2017) Enzyme kinetics for complex system enables accurate determination of specificity constants of numerous substrates in a mixture by proteomics platform. Mol Cell Proteomics 16 (1):135–145 25. Steen H, Jebanathirajah JA, Springer M, Kirschner MW (2005) Stable isotope-free relative and absolute quantitation of protein phosphorylation stoichiometry by MS. Proc Natl Acad Sci U S A 102(11):3948–3953 26. Bremmer SC, Hall H, Martinez JS, Eissler CL, Hinrichsen TH, Rossie S, Parker LL, Hall MC,

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Charbonneau H (2012) Cdc14 phosphatases preferentially dephosphorylate a subset of cyclin-dependent kinase (Cdk) sites containing phosphoserine. J Biol Chem 287 (3):1662–1669 27. Eissler CL, Mazon G, Powers BL, Savinov SN, Symington LS, Hall MC (2014) The

Cdk/Cdc14 module controls activation of the Yen1 holliday junction resolvase to promote genome stability. Mol Cell 54(1):80–93 28. Tyanova S, Temu T, Cox J (2016) The MaxQuant computational platform for mass spectrometry-based shotgun proteomics. Nat Protoc 11(12):2301–2319

Chapter 6 Whole-Mount Immunostaining for the Identification of Histone Modifications in the S-Phase Nuclei of Arabidopsis Roots Hirotomo Takatsuka and Masaaki Umeda Abstract This chapter describes a method used to analyze the behavior of histone modifications in S phase in Arabidopsis using a whole-mount immunostaining technique. Previous studies have demonstrated that dramatic changes in local chromatin structure are required for the initiation and progression of DNA replication, and that histone modifications play an essential role in the determination of chromatin structure in S phase. Since euchromatic and heterochromatic regions are replicated in distinct S-phase stages, it is important to identify histone modifications at each stage. Here, we introduce a protocol for whole-mount immunostaining combined with 5-ethynyl-20 -deoxyuridine (EdU) staining, which enables the visualization of spatial patterns in histone modifications in the early and late S-phase nuclei of Arabidopsis roots. Key words Cell cycle, S phase, Histone modification, Euchromatin, Heterochromatin, Immunostaining, EdU staining, Root, Arabidopsis

1

Introduction Proper progression of DNA replication followed by mitosis is essential for the maintenance of genome integrity in somatic cells [1, 2]. During the S phase of the cell cycle, each genome compartment is replicated in distinct stages; specifically, euchromatic and heterochromatic regions are replicated in the early and late S phases, respectively [3]. Loading of the prereplication complex (pre-RC) and fork progression require changes in local chromatin structure, which are precisely controlled in each genome compartment, enabling faithful and efficient DNA replication with proper timing [4]. Histone modifications, such as methylation and acetylation, determine chromatin structure, and thus are known to be involved in the control of DNA replication [5, 6]. Indeed, several reports have demonstrated that, in Arabidopsis cultured cells, specific histone modifications are highly associated with S-phase progression [7, 8]. However, due to technical difficulties, the

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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involvement of histone modifications in DNA replication has not been studied in living plant tissues. Here, we introduce a method of immunostaining for histone modifications in S-phase nuclei using Arabidopsis roots. This method will facilitate understanding of the mechanisms underlying the epigenetic regulation of DNA replication in plant cells.

2

Materials

2.1

Plant Material

1. Arabidopsis thaliana ecotype Col-0.

2.2

Seed Sterilization

1. 70% ethanol. 2. Sterilization solution: 4% sodium hypochlorite and 0.05% Triton X-100. 3. 0.1% agar solution. 4. Sterilized distilled water. 5. Microtube rotator.

2.3

Media

1. MS media: Dissolve 4.6 g Murashige and Skoog (MS) Plant Salt Mixture in 900 mL distilled water. Add 0.5 g 2-(N-morpholino)ethanesulfonic acid (MES) and completely dissolve the powder. Adjust pH to 6.3 with 1 M KOH. Adjust the volume to 1 L with distilled water. Autoclave at 121  C for 15 min. On a clean bench, pour 40 mL autoclaved medium into a 100 mL flask and put a silicon cap on the flask. 2. For solid MS media, add 4 g Phytagel™ (Sigma Aldrich, St. Louis, MO, USA) to MS media before autoclaving. Pour 50 mL autoclaved medium into a square petri dish and solidify for 30 min.

2.4

Seed Sowing

1. Aluminum foil. 2. Surgical tape.

2.5

Reagents

1. Microtubule-stabilizing buffer (MTSB): Dissolve 1.5 g piperazine-1,4-bis(2-ethanesulfonic acid) (PIPES), 0.19 g ethylenediaminetetraacetic acid (EDTA), 0.13 g MgSO4·7H2O, and 0.5 g KOH in 800 mL distilled water. Adjust the pH to 7.0 with KOH and make up to 1 L with distilled water. 2. Phosphate-buffered saline (PBS): Dissolve 7.6 g NaCl, 0.724 g Na2HPO4, and 0.218 g KH2PO4 in 900 mL distilled water. Adjust the pH to 7.4 with NaOH and make up to 1 L with distilled water. 3. Fixation solution: Add 0.3 g paraformaldehyde (PFA) to 9.5 mL heated MTSB. Slowly raise the pH to 7–8 by adding 1 M NaOH dropwise using a pipette. Adjust the volume to 10 mL by adding MTSB.

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4. Click-iT™ Plus EdU Cell Proliferation Kit for Imaging, Alexa Fluor™ 488 dye (Thermo Fisher Scientific, Waltham, MA, USA) (see Note 1). Just before use, prepare 300 μL EdU detection cocktail per sample according to the manual. 5. Pectolyase solution: Add 5 mg pectolyase and 15 μL Triton X-100 to 9 mL distilled water and dissolve the powder completely. Make up to 10 mL with distilled water. 6. 4 Tris–NaCl–Blocking reagent buffer (TNB): Mix 4 mL distilled water, 4 mL 1 M Tris–HCl (pH 7.5), 0.35 g NaCl, and 0.2 g blocking reagent (Roche, Basel, Switzerland) and completely dissolve at 60  C. Make up to 10 mL with distilled water. 7. Antibody solution: Mix 75 μL 4 TNB and 225 μL distilled water. Add 3 μL primary or secondary antibody and mix well. Prepare 300 μL antibody solution per sample. Primary antibodies are anti-histone H3 antibody (Abcam, Cambridge, UK), anti-H3K9me2 antibody (Abcam), and anti-H4K12ac antibody (Abcam). Secondary antibodies are Alexa 647 Donkey Anti-Rabbit IgG (H+L) (Jackson ImmunoResearch, West Grove, PA, USA) and Alexa 647 Donkey Anti-Mouse IgG (H +L) (Jackson ImmunoResearch). 8. DAPI stock solution: Dissolve 1 mg 40 ,6-diamidino-2-phenylindole (DAPI) in 1 mL distilled water. 2.6

Immunostaining

1. Water-repellent glass slide (Matsunami Glass, Osaka, Japan). 2. Coverslip (Matsunami Glass).

2.7 Moisture Chamber

1. Lay a paper towel at the bottom of a square petri dish. 2. Wet the paper towel with distilled water. 3. Place four or five plastic bars on the wet paper towel in parallel (see Note 2). 4. Close the petri dish with lid.

2.8

3 3.1

Observation

1. Imaging is conducted with an Olympus FV1000 confocal microscope. Images are acquired and analyzed using FV10-ASW software with a Kalman filter (Olympus, Tokyo, Japan).

Methods Seed Sterilization

1. Mix seeds with 1 mL of the sterilization solution in a 1.5-mL microtube. 2. Rotate with a microtube rotator at room temperature for 10 min and remove the sterilization solution with a pipette. 3. Add 1 mL sterilized water.

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4. Gently shake the tube a few times and remove sterilized water with a pipette. 5. Repeat steps 3 and 4 three times. 6. Remove sterilized water with a pipette and add 1 mL 0.1% agar solution. 3.2

Seed Sowing

1. Sow sterilized seeds on the surface of solidified MS media. 2. Seal the dish with surgical tape and cover with aluminum foil to provide shading. 3. Incubate the petri dish at 4  C for 2 days.

3.3

Cultivation

1. Place the petri dish at a 90 angle in a plant growth chamber and incubate at 22  C for 5 days under continuous light conditions.

3.4

EdU Staining

1. Add 80 μL 10 mM EdU to 40 mL liquid MS media in a 100 mL flask to obtain a final concentration of 20 μM. 2. Soak 4-day-old seedlings in the MS media containing EdU (Fig. 1). 3. Put a silicon cap on the flask and wrap the top part with aluminum foil. 4. Rotate the flask at 60 rpm for 30 min under continuous light conditions.

Fig. 1 EdU treatment of Arabidopsis seedlings. Four-day-old seedlings are immersed in a liquid medium containing EdU and incubated for 30 min with shaking

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5. Remove the media and replace with 40 mL MS media without EdU. 6. Shake the flask gently. 7. Repeat steps 5 and 6 three times. 8. Collect root tips of approximately 1 cm in a 1.5 mL tube containing 1 mL fixation solution (see Note 3). 9. Fix the root tips in a vacuum for 2 min. Repeat three times. 10. Incubate the sample for 14 min at room temperature with gentle shaking. 11. Remove the fixation solution. 12. Wash the root tips with 1 mL PBS for 10 min. Repeat three times. 13. Soak the root tips in 300 μL EdU detection cocktail and incubate for 30 min in the dark. 14. Remove the detection cocktail. 15. Wash the root tips with 1 mL PBS for 10 min. Repeat three times. 3.5

Immunostaining

1. Put 300 μL PBS into the well of a water-repellent glass slide. 2. Carefully transfer EdU-stained roots to the well of the waterrepellent glass slide (Fig. 2; see Note 4). 3. Remove PBS and add 300 μL pectolyase solution. 4. Incubate at 30  C for 2 h and remove the pectolyase solution. 5. Wash with 300 μL PBS for 10 min. Repeat three times. 6. Remove PBS and add 300 μL primary antibody solution. 7. Incubate in the moisture chamber at 4  C for at least 8 h (Fig. 3; see Note 5), then remove the primary antibody solution (see Note 6). 8. Wash with 300 μL PBS for 10 min. Repeat three times. 9. Remove PBS and add 300 μL secondary antibody solution.

Fig. 2 Transfer of EdU-stained roots to a water-repellent glass slide

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Fig. 3 Reaction with antibodies in a moisture chamber. Water-repellent glass slides are put on plastic bars in a petri dish dampened with wet paper towels. Plastic bars are made from 1 mL plastic pipettes

10. Incubate in the moisture chamber at 37  C for 1 h (Fig. 3), then remove the secondary antibody solution. 11. Wash with 300 μL PBS for 10 min. Repeat three times. 12. Remove PBS and stain nuclei with PBS containing DAPI at the final concentration of 1 μg/mL for 15 min. 13. Remove DAPI-containing PBS and wash with PBS for 10 min. 14. Mount roots onto a glass slide with a drop of ProLong® Diamond (Thermo Fisher Scientific) and gently put on a coverslip. 3.6 Microscopic Observation

1. Epidermal cells in the root tip are observed under a confocal microscope (see Note 7). The excitation wavelengths for Alexa 488, Alexa 647, and DAPI are 488, 647, and 405 nm, respectively. Due to variation in the fluorescence intensity of immunostained nuclei, the laser strength needs to be optimized for each sample. 2. To view a single confocal section of the whole root tip, a 20 objective lens is used. Representative images are shown in Fig. 4. To obtain high-resolution images, a 100 objective lens is used. Z-stack images are taken around the nucleus of interest at 0.2 μm intervals. The maximum projection of the Z-stack view is generated with ImageJ software.

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Fig. 4 Arabidopsis root tips after immunostaining (Alexa 647), EdU staining (Alexa 488), and DAPI staining. Immunostaining was conducted using antibodies against histone H3, H3K9me2, and H4K12ac. Bar ¼ 100 μm

3. S-phase nuclei can be identified as those stained with EdU. Nuclei in the early and late S phases can be distinguished based on EdU staining patterns; those with an EdU signal over the whole nucleus are in the early S phase, during which euchromatin replication occurs, while those showing speckled

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Fig. 5 Magnified images of nuclei after immunostaining (Alexa 647), EdU staining (Alexa 488), and DAPI staining. Nuclei in the early and late S phases are distinguished according to EdU staining patterns. Immunostaining was conducted using antibodies against histone H3, H3K9me2, and H4K12ac. Bar ¼ 5 μm

EdU signaling are in the late S phase, which is characterized by the exclusive replication of centromeric heterochromatin [9]. Representative images of early and late S-phase nuclei are shown in Fig. 5. Note that heterochromatic H3K9me2 marks completely overlap with EdU signals in the late S phase.

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Notes 1. An alternative kit, Click-iT™ EdU Cell Proliferation Kit for Imaging, Alexa Fluor™ 488 dye, interferes with fluorescent signals of immunostaining, and thus is not appropriate for this experiment. 2. Bars are needed to support the glass slide on the wet paper towel to prevent it from becoming wet. We use 1 mL plastic pipettes cut into small pieces. 3. The addition of Triton® X-100 to the fixation solution may facilitate penetration of antibodies into cells. Test 0.1-0.5% Triton® X-100 in case that immunostaining does not work well. 4. We usually put about 20 seedlings in one well of a waterrepellent glass slide. 5. Incubation with primary antibodies can be extended up to 48 h. 6. The success of immunostaining depends heavily on the quality of the primary antibodies. If immunostaining does not work well, we recommend testing other antibodies purchased from different companies. 7. This protocol is optimized to observe nuclei in the root epidermis, into which antibodies can easily penetrate. To observe other cell types or tissues, it will be necessary to establish optimal immunostaining conditions accordingly.

Acknowledgments This work was supported by MEXT KAKENHI Grant Numbers 17H06470 and 17H06477 to M.U., and JSPS KAKENHI Grant Number 17K15415 to H.T. References 1. Schuermann D, Fritsch O, Lucht JM, Hohn B (2009) Replication stress leads to genome instabilities in Arabidopsis DNA polymerase δ mutants. Plant Cell 21:2700–2714 2. Domenichini S, Benhamed M, de Jaeger G et al (2012) Evidence for a role of Arabidopsis CDT1 proteins in gametophyte development and maintenance of genome integrity. Plant Cell 24:2779–2791 3. Concia L, Brooks AM, Wheeler E et al (2018) Genome-wide analysis of the Arabidopsis replication timing program. Plant Physiol 176:2166–2185

4. Belsky JA, Macalpine HK, Lubelsky Y et al (2015) Genome-wide chromatin footprinting reveals changes in replication origin architecture induced by pre-RC assembly. Genes Dev 29:212–224 5. Rivera C, Gurard-Levin ZA, Almouzni G, Loyola A (2014) Histone lysine methylation and chromatin replication. Biochim Biophys Acta Gene Regul Mech 1839:1433–1439 6. Sugimoto N, Fujita M (2017) Molecular mechanism for chromatin regulation during MCM loading in mammalian cells. In: Advances in experimental medicine and biology. Springer, Heidelberg, pp 61–78

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7. Lee TJ, Pascuzzi PE, Settlage SB et al (2010) Arabidopsis thaliana chromosome 4 replicates in two phases that correlate with chromatin state. PLoS Genet 6:1–18 8. Jiang D, Berger F (2017) DNA replication–coupled histone modification maintains

Polycomb gene silencing in plants. Science 357:1146–1149 9. Hayashi K, Hasegawa J, Matsunaga S (2013) The boundary of the meristematic and elongation zones in roots: endoreduplication precedes rapid cell expansion. Sci Rep 3:1–8

Chapter 7 Construction of a Full-Length 30 UTR Reporter System for Identification of Cell-Cycle Regulating MicroRNAs Dominika Kaz´mierczak and Per Hydbring Abstract Three prime untranslated region (30 UTR) reporter constructs are widely used by the scientific community to functionally link microRNAs (miRNAs) to suppression of mRNA expression. However, full-length 30 UTR vectors are rarely employed due to labor-intensive cloning work. Instead, 30 UTR fragments containing putative miRNA binding sites are commonly utilized to mechanistically validate miRNAs. Assaying truncated 30 UTRs may falsely validate miRNAs due to altered positioning of binding sites in respect to 30 UTR length and RNA secondary structure. Here we present a detailed protocol for the construction of full-length 30 UTR luciferase reporter constructs that was used to unveil miRNAs regulating multiple cell-cycle factors. Key words 30 UTR, Dual-luciferase vector, Nested PCR, Genomic DNA, Restriction enzyme digestion, G418 selection, U2OS cells

1

Introduction MiRNAs are small noncoding RNAs regulating the expression of target genes through binding to complementary sequences in mRNA transcripts. A single miRNA may regulate hundreds of mRNA transcripts [1]. MiRNAs have been shown to bind to multiple regions of mRNA transcripts, including 50 UTRs, open reading frames, and 30 UTRs. Binding of miRNAs to 30 UTRs is generally accepted as the main interaction resulting in mRNA degradation and/or protein translational inhibition [1, 2]. There are numerous molecular techniques to monitor suppression of selected mRNA transcripts by specific miRNAs [3–7]. However, due to various challenges, most techniques are not suitable for large-scale screening. Since miRNAs are vastly pleiotropic in their targeting pattern, the screening technology requires a measuring output that is not affected by indirect biological events. 30 UTR reporter assays have

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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been around since the dawn of miRNA biology [8]. The major advantage with 30 UTR reporter assays is the ability to assess a direct effect of a particular miRNA on a chosen 30 UTR. The drawback is the requirement of using transfection methods, with the consequence of nonphysiological miRNA expression. While the usage of miRNA-inducible cell lines can circumvent this problem to some extent, such approaches are unfeasible in screening studies. Due to the significant size of many 30 UTRs, making cloning work cumbersome, scientists often opt for 30 UTR fragments limited to the target sites of specific miRNAs. This does not only create a completely artificial system, but is also incompatible with systematic screening methodologies. In order to uncover miRNAs regulating cell-cycle factors on a systematic basis, we developed a full-length 30 UTR reporter system to specifically identify miRNAs regulating cell cycle cyclins and cyclin-dependent kinases. Uncovered cell cycletargeting miRNAs displayed a substantial enrichment in targeting multiple cell cycle factors [9, 10]. Although we so far only used this system to uncover miRNAs targeting cyclins and cyclin-dependent kinases, the system can be adapted to any 30 UTR of interest, and hence any cell cycle factor of interest [11]. Since our 30 UTR screening system relies on full-length 30 UTRs, thereby preserving the secondary structure of the 30 UTR as well as miRNA target site positioning in relation to the stop codon [12–18], it is less prone to generate false positive or false negative miRNA candidates. Moreover, in addition to the luciferase reporter gene upstream of the 30 UTR, constructed 30 UTR reporter vectors contain a reference luciferase reporter gene for normalization of results. Following completion of cloning, constructed 30 UTR constructs are inserted stably into the genome of U2OS cells, through G418 selection, providing a physiological environment for expression of the 30 UTR. Clones emerging from each 30 UTR construct transfection are monitored for detectable reporter gene expression, and validated for miRNA functionality. Selected clones are screened for miRNA functional interaction by subjecting the 30 UTR reporter cell lines to miRNA mimic libraries through transient transfection. Luciferase reporter gene expression is monitored 28 h post miRNA-library transfection (Fig. 1). Importantly, we have validated the suitability of the host cell line, U2OS, by profiling it for endogenous expression of cyclins and cyclin-dependent kinases, as well as endogenous expression of approximately 1000 human miRNAs [9, 10]. The outcome of performing full-length 30 UTR reporter assays in U2OS cells is not affected by endogenous expression of the target gene, or by endogenous expression of the investigated miRNA [9, 10].

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Fig. 1 Schematic overview of the construction of a full-length 30 UTR reporter system for microRNA library screening. Cell-cycle factors indicated in the figure refers to 30 UTRs previously cloned and systematically assayed for functional targeting by microRNA [9, 10]. MCS multiple cloning site, SV40 Simian virus 40 promoter, Ren-Luc-NeoR Renilla luciferase–neomycin resistance cassette fusion gene, AmpR ampicillin resistance cassette, PGK phosphoglycerate kinase promoter, FF-Luc firefly luciferase gene, FF/Ren-Luc firefly–Renilla luciferase expression ratio

2 2.1

Materials Cloning

1. 100 bp and 1 kb DNA ladder. 2. pmirGLO dual luciferase vector (Promega, #E1330). 3. Cloning primers: Make a stock solution (100 μM) of each primer by adding the appropriate volume of nuclease-free water depending on amounts of mole synthesized. For example, if 100 nmol of primer, add 1 mL of nuclease-free water to obtain a stock solution of 100 μM. Dilute stock solution 1:10 to obtain working solution for PCR reaction. 4. Restriction enzymes: Preferably XhoI (20 U/μL) and XbaI (20 U/μL). If site/s already present in genomic 30 UTR sequence, use Pme I (20 U/μL), EcoICRI (20 U/μL), NheI (20 U/μL), SalI (20 U/μL), or SbfI (20 U/μL), depending on cloning strategy. 5. Human genomic DNA: 100 ng/μL in 10 mM Tris, 0.1 mM EDTA. 6. PCR purification kit. 7. Tris–acetate–EDTA buffer: For a 50 stock solution, mix 242 g tris base in double distilled water with 57.1 mL glacial

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acetic acid and 100 mL of 0.5 M EDTA solution (pH 8.0). Adjust volume to 1 L with ultrapure water. To make 1 working solution, dilute stock solution 1:50 using ultrapure water. 8. SYBR Safe DNA gel stain. 9. 1% agarose Tris–acetate–EDTA gel: 1 g of molecular biology grade agarose in 50 mL of 1 Tris–acetate–EDTA buffer. Heat solution in microwave until agarose is completely dissolved. Add 50 mL of 1 Tris–acetate–EDTA buffer to bring total volume to 100 mL. Add 20 μL of SYBR safe DNA gel stain and swirl bottle gently to mix. 10. CutSmart buffer (New England Biolabs, #B7204S). 11. 0.4 U/μL T4 ligase. 12. 10 T4 ligase buffer. 13. Competent cells: Subcloning efficiency DH5 alpha. 14. LB medium: Mix 10 g of tryptone, 10 g NaCl, and 5 g yeast extract. Add 900 mL of double distilled water and mix until content is completely dissolved. Adjust the pH to 7.0 by slowly adding, drop-wise, 1 M of NaOH solution. Make up to 1 L with double distilled water. Autoclave the solution using liquid cycle setting for 20 min. 15. LB-Amp medium: Add ampicillin to cooled sterile LB medium to 100 ug/mL final. 16. LB-Amp-plates: Add 3 g of bacteriological agar to 200 mL of LB media. Autoclave the solution using liquid cycle setting for 20 min. Let solution cool down to approximately 50  C before adding 200 μL of ampicillin (100 mg/mL). 17. Column miniprep kit. 18. PfuUltra II Hotstart PCR 2 master mix (Agilent, #600850). 19. 100 mg/mL Ampicillin: Prepare in sterile, nuclease-free water; store at 20  C. 2.2 DNA Transfection and Selection of Stable Cell Lines

Human U2OS cells are grown at 37  C, 5% CO2. 1. U2OS cells. 2. Lipofectamine 2000 transfection reagent. 3. Opti-MEM I. 4. McCoy’s 5A complete medium: For a 500 mL bottle, add 50 mL Fetal Bovine Serum and 5 mL of 100 PenicillinStreptomycin solution. 5. McCoy’s 5A complete medium G418: 1 g of G418 in 10 mL of nuclease-free water to make a 100 mg/mL stock solution. Dilute to 800 μg/mL working solution in McCoy’s 5A complete medium. Filter medium using a 0.2 micron filter before use. Working solution can be stored at +4  C.

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6. Cloning discs (Sigma-Aldrich, #Z374431-100EA). 7. Trypsin–EDTA (0.05%), phenol red. 8. Fetal bovine serum. 9. Cryopreserving media: 90% FBS, 10% DMSO (cell culture, hybridoma grade). 10. 1 PBS. 11. 70% EtOH. 2.3 miRNA Transfection and Assaying of Luciferase Expression

1. Lipofectamine RNAiMAX (see Note 1). 2. Opti-MEM I. 3. McCoy’s 5A complete medium. 4. McCoy’s 5A #10358633).

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5. Dual-Glo Luciferase Assay System (Promega, #E2920). 6. Assay plate: 96-well, white, flat bottom. 7. Costar sterile disposable reagent reservoirs (Corning, #4870 (10320551)). 8. mirVana miRNA mimic customized library (Thermo Fisher Scientific, #4464066). Resuspend customized miRNA mimic library in nuclease-free water to a stock solution of 25 μM for long-term storage in 96-well plates in 80  C. 9. Nuclease-free water (not DEPC-treated). 10. Pre-miR miRNA precursor molecule negative control #2 (Thermo Fisher Scientific, #AM17111). 11. Generic luminescence plate reader for 96-well plates.

3

Methods Carry out all procedures at room temperature unless otherwise specified. Human U2OS cells are grown at 37  C, 5% CO2.

3.1 Cloning of 30 UTR from Human Genomic DNA into pmirGLO Vector

1. Select the most prevalent 30 UTR of your gene of interest from TargetScanHuman database. For example, for CCND1, select identifier ENST00000227507.2 (see Note 2). 2. Go to the Ensembl database to find out the Ref-seq number that corresponds to your Transcript ID identifier number. 3. Go to the NCBI website, select “Nucleotide” in the dropdown menu and paste your identified Ref-seq number. 4. Download the 30 UTR sequence and manually inspect it for presence of XhoI (CTCGAG) and XbaI (TCTAGA) sites. If not present, these restriction enzyme sites can be used for cloning.

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Fig. 2 Amplification of genomic 30 UTR regions through nested PCR. Forward and reverse nest out primers are designed to bind upstream of the stop codon and downstream of the 30 UTR, respectively. Following a first round of PCR, forward and reverse nest in primers are designed to locate exactly at the start and end of the 30 UTR, respectively. A second round of PCR is performed on the product from the first PCR. F forward primer, R reverse primer

5. Select “graphics” to visualize your transcript in the genome view. Identify the direction of the transcript and “zoom to sequence” at the start of the 30 UTR. Manually assess the nucleotide sequence around the start and end of the 30 UTR. 6. Design your PCR nest out and nest in primers. Nest out primers should be approximately 20 nucleotides long with a GC content of 40–60%. Forward nest out primer should be located before the stop codon of your gene of interest while the reverse nest out primer is located downstream of the end of the 30 UTR. Optimally, nest in primers should perfectly overlap with the start and end of the 30 UTR (Fig. 2). For nest in primers, use XhoI as flanking site for your forward primer, and XbaI as flanking site for your reverse primer. In order for the restriction enzyme to cut the PCR product, add “CCG” in front of the XhoI (CTCGAG) site and “CCA” in front of the XbaI (TCTAGA) site. Design your primers using the oligo ordering service through Integrated DNA Technologies, or similar online software. 7. Mix 100 μL of genomic DNA, 1 μL forward primer (10 μM concentration), 1 μL reverse primer (10 μM concentration), 25 μL PfuUltra II 2 master mix and nuclease-free water to a total volume of 50 μL. Run the following PCR program:

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95  C, 2 min (Hot start), 95  C, 20 s (denaturing), Primer pair average melt point, 5  C, 20 s (annealing), 72  C, 15 s/kb (extension). Repeat the denaturing, annealing and extension for 32 cycles. End the PCR program with an additional 3 min at 72  C followed by hold at 4  C (see Note 3). 8. Clean up the PCR product using the Qiagen PCR purification kit, or similar, according to the manufacturer’s instructions. 9. Run 5 μl of the PCR product on a 1% agarose Tris-Acetate EDTA gel stained with SYBR safe DNA intercalator. Depending on the expected size of the PCR product, visualize the product with a 100 bp, or 1 kb DNA ladder. 10. If PCR run proves successful, set up an XhoI/XbaI restriction digestion of the PCR product as well as the parental vector pmirGLO. Use 20 U of each restriction enzyme for each digestion. Use 1 μg of pmirGLO for the vector digest. Use a total volume of 10 μL in each restriction enzyme digestion. Digest for 3 h at 37  C. Following completion of digestion, clean up the reaction using the Qiagen PCR purification kit, or similar, according to the manufacturer’s instructions (see Note 4). 11. Set up ligation of the PCR fragment into the parental vector using 3, 5, and 8 excess in molarity of fragment in relation to vector. Each ligation reaction should contain 0.4 U of T4 DNA ligase, 1 ligase buffer, and 20 ng of vector. To calculate amount of PCR fragment, based on excess in molarity, use the formula, mass ¼ molarity weight  molarity. The molecular weight of a base pair is 650 g/mol. The size of the parental vector is 7350 bp. Set the ligation reactions at 16  C for a minimum of 4 h. 12. Take competent cells from 80  C and thaw on ice for 30 min. Set a water-bath to 42  C. Mix 2 μL of ligation reaction with 50 μL of competent cells in an Eppendorf tube. Gently mix by flicking the tube and incubate on ice for 30 min. Heat-shock the mixture at 42  C for exactly 45 s. Put the tubes back on ice for 2 min. Add 500 μL of LB medium to the mixture and incubate at 37  C, shaking, for 45 min. 13. Plate all of the transformation mixture reactions to LB-Amp-plates. Incubate plates at 37  C for 16 h. 14. Pick a minimum of eight bacterial colonies for inoculation into 2.5 mL LB-Amp medium. Do not close lids tightly. Incubate tubes with inoculated bacteria for 16 h at 37  C, shaking at 250 rpm. 15. Perform a DNA plasmid mini preparation using the Qiagen miniprep kit, or similar, according to the manufacturer’s instructions.

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16. Perform a test restriction digestion by mixing 1 μL of plasmid DNA, 10 U of XhoI, 10 U of XbaI, and 1 μL of 10X restriction enzyme buffer in a total volume of 10 μL. Incubate for at least 1 h at 37  C. 17. Run the entire restriction enzyme reaction on a 1% agarose Tris-Acetate EDTA gel stained with SYBR safe DNA intercalator. Depending on the expected size of the PCR product, visualize the product with a 100 bp, or 1 kb DNA ladder. 18. If two bands appear at the expected size (vector and cloned 30 UTR), send plasmid for DNA sequencing using a forward sequencing primer located approximately 100 bp upstream of the start of the 30 UTR (see Note 5). 3.2 Establishment of 30 UTR Dual Luciferase Reporter Cell Lines

1. Seed approximately one million U2OS cells on a 100 mm diameter cell culture plate in 8 mL McCOY’s 5A complete medium and return to incubator for 24 h before performing transfection. 2. Mix 5 μL Lipofectamine 2000 with 250 μL Opti-MEM I in an Eppendorf tube. Mix up to 1000 ng of 30 UTR vector (in maximum 5 μL) with 250 μL Opti-MEM I in a separate Eppendorf tube. Incubate both tubes for 5–10 min at room temperature. 3. Mix the contents of both tubes together and incubate for another 15–20 min. 4. Remove all the McCOY’s 5A complete medium from the cell culture plate and replace with 3.5 mL of Opti-MEM I. Add the transfection mixture from step 3 to the cell culture plate (total volume 4 mL). Swirl the plate gently to make sure the entire plate is covered with medium. Incubate for 4–6 h. 5. Aspirate transfection medium from the cell culture plate and replace with McCOY’s 5A complete medium and incubate for a further 48 h. 6. Add McCOY’s 5A complete medium G418 (see Note 6). 7. Replace G418-containing medium every 48–72 h until the emergence of G418-resistant colonies (see Note 7). 8. Mark G418-resistant colonies with a marker pen under an inverted microscope with 4 magnification. Mark at least twenty colonies for each 30 UTR-vector cell line. 9. Aspirate all medium from the cell culture plate. Wash cells once by gently adding 3 ml of 1 PBS. Aspirate PBS from the cell culture plate. Using a tweezer, dip a cloning disc in TrypsinEDTA and gently place it on top of a marked up G418-resistant cell colony. Wash the tweezer in 70% EtOH, to avoid cell contamination, and repeat the procedure for all marked up colonies on the plate (see Note 8).

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10. Incubate the plate for approximately 5 min. 11. Transfer each cloning disc from the 100 mm cell culture plate to a 24-well cell culture plate with 500 μL G418-containing medium per well. Wash the tweezer in 70% EtOH, to avoid cell contamination, between transfer of each cloning disc. 12. Amplify all G418-resistant U2OS cell clones to approximately five million cells per clone by transferring cells, through multiple trypsin–EDTA mediated passages, from 24-well cell culture plates to 6-well cell culture plates to 100 mm cell culture plates. 13. Cryopreserve all G418-resistant U2OS cell clones by pelleting trypsinized cells at 200–300  g, for 5 min. Wash cell pellets by adding 3 mL of PBS followed by another spin at 200–300  g for 5 min. Resuspend each cell pellet in 1 mL of cryopreserving media followed by transfer to 80  C. 3.3 miRNA Library Screening of 30 UTR Reporter Targeting

1. Prepare sufficient amount of U2OS 30 UTR-reporter cells in a subconfluent condition. Each library plate requires 900,000 cells per replicate covering a maximum of 60 miRNA mimics. Three replicates of a single library plate require 2,700,000 cells (see Note 9). 2. Dilute library, in nuclease-free water, to 1 μM for transfection. Manually add scrambled negative miRNA mimic controls, as well as positive controls, to selected wells (see Note 10). 3. Mix Opti-MEM I and Lipofectamine RNAiMAX in a ratio of 90:1 in an Eppendorf tube. Calculate the total amount needed depending on size of the miRNA library. 13.5 μL of OptiMEM–RNAiMAX mixture will be used for each well in a 96-well plate. Include at least 1 mL of excess Opti-MEM– RNAiMAX mixture. 4. Vortex the Opti-MEM–RNAiMAX solution for 10 s followed by a 10-min incubation at room temperature. 5. Pipet the miRNA mimic library, prediluted to 1 μM, into three separate tissue culture treated 96-well plates with flat bottom, that is, the content of each library plate is transferred into three separate tissue culture treated 96-well plates with flat bottom. 1.5 μL of each miRNA is transferred into each well of a 96-well plate. 6. Pour the Opti-MEM–RNAiMAX mixture into a conical sterile disposable reagent reservoir. Using a multichannel pipette aid, pipet 13.5 μL of the mixture into each well of the three 96-well plates already containing the miRNA mimics, except for wells in the outer columns and rows. 7. Incubate for 20 min at room temperature.

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8. While incubating the miRNA-mimic–Opti-MEM–RNAiMAX mixture, trypsinize the U2OS 30 UTR reporter cells for 5 min at 37  C, 5% CO2. Once detached, resuspend the cells with 5 the volume of McCoy’s 5A complete medium. 9. Count cells using a Burker chamber or automatic cell counter. Adjust cell concentration to 333,333 cells/mL by diluting cells further in McCoy’s 5A complete medium in a conical sterile disposable reagent reservoir. 10. Pipet 45 μL of cells, corresponding to 15,000 cells, into each well of the three 96-well plates, already containing 15 μL miRNA-mimic–Opti-MEM–RNAiMAX mixture, except for wells in the outer columns and rows. 11. Pipet 60 μL of McCoy’s 5A cell culture medium, not containing any cells, into the wells of the outer columns and rows (see Note 11). 12. Carefully look through each plate in an inverted microscope, 10 magnification, to make sure cells are evenly dispersed. If not, tap the plates gently and check again under microscope. 13. Incubate for 28 h before assaying luciferase activity. 14. Aspirate miRNA-containing cell culture medium from the 96-well plates using a multichannel pipette. 15. Add 45 μL of McCOY’s 5A, without phenol red and without FBS, to each well (see Note 12). 16. Lyse cells by adding 45 μL of Firefly luciferase substrate solution (see the manufacturer’s instructions, Dual-Glo Luciferase Assay) to each well, cover the plate with aluminum foil, and incubate for 10 min at room temperature. 17. Transfer 85 μL of lysed cell solution to the white 96-well assay plates with flat bottoms. Carefully pipet the solution up and down a few times without introducing any air bubbles in the solution (see Note 13). 18. Read Firefly luminescence on a 96-well luminescence reader. Set the reading time of each well to 1 s. 19. Add 45 μL of freshly made Renilla luciferase solution (see the manufacturer’s instructions, Dual-Glo Luciferase Assay) to each well of the white 96-well assay plates, cover the plates with aluminum foil, and incubate for 10 min. 20. Read Renilla luminescence on a 96-well luminescence reader. Set the reading time of each well to 1 s. 21. Calculate Firefly–Renilla ratios. Compare luciferase ratios from wells transfected with scrambled negative miRNA mimic control with experimental miRNA mimic wells and wells transfected with positive miRNA mimic controls.

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Notes 1. When comparing a few different lipid-based transfection reagents for 30 UTR reporter knockdown efficiency in U2OS cells, expressing a full-length 30 UTR of cyclin D1, Lipofectamine RNAiMAX resulted in the most pronounced 30 UTR knockdown efficiency. However, all tested reagents led to some knockdown of reporter construct. If using a different transfection reagent than Lipofectamine RNAiMAX, it is advisable to compare the knockdown efficiency in the stable 30 UTR reporter cell line before initiating a full library screen. 2. Many genes have several 30 UTRs listed in this database. The different 30 UTRs will be listed according to prevalence as well as 30 UTR length. In case a gene has several listed 30 UTRs, always select the most prevalent 30 UTR. In the vast majority of cases, the most prevalent 30 UTR is also the longest 30 UTR. 3. Using the indicated PCR program and polymerase, maximum amplification is around 5500 bp. If your 30 UTR of interest is longer than 5500 bp you need to subclone it in multiple steps. For example, the 30 UTR of CDK6 (10,219 bp), should be subcloned in two steps. When subcloning 30 UTRs, you would need a total of three restriction enzyme sites. Make sure you have three restriction enzyme sites in the multiple cloning site (MCS) of pmirGLO that are unique for the MCS and not present in the 30 UTR sequence. 4. A 3-h digestion should be efficient enough to completely cut the parental vector. In case you expect restriction enzymes to not be fully operational, scale up the restriction enzyme reactions to a total of 50 μL, and run the digested vector and fragment on a preparative 1% agarose Tris-Acetate EDTA gel, with two separate wells for the vector and two separate wells for the fragment. Cut the gel using a scalpel and only expose one well each of the vector and fragment. Carefully cut out the region with digested vector and fragment, reassemble the gel, and cut the corresponding regions for the other wells. Extract the DNA using Qiagen Gel Extraction kit, or similar, according to the manufacturer’s instructions. 5. Sanger sequencing normally gives about 700–1000 nucleotides of accurate reading. Depending on the size of your 30 UTR, you may need more than one sequencing primer. Some 30 UTRs may even require ten or more sequencing primers to optimally document the full sequence of the cloned 30 UTR. Place each primer approximately 100 nucleotides upstream of the region of interest. Keep GC content of primer to 40–60%.

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6. 800 μg/mL G418 will give a good killing effect on U2OS cells not expressing the 30 UTR vector. However, if G418 solution is freshly made, 400 μg/mL should also work. 7. During G418 selection it is essential the cells never become confluent. If confluent, or nearly confluent, the killing effect of G418 will decrease dramatically with the result of no isolated resistant clones. If cells become too dense at the beginning of the selection process, simply passage cells into multiple plates, before continuing G418 selection. 8. It is imperative to work fast but accurate during this process. After aspiration of PBS, you only have a few minutes before cells will become completely devoid of liquid. The tip of the tweezer can be quickly dipped in 70% EtOH as the residual EtOH will evaporate very fast. Do not exceed 20 colonies per plate during this procedure. 9. Before starting the full library screen, G418-resistant clones should have been evaluated for Firefly and Renilla expression as well as 30 UTR functionality. Clone validation is conducted by using the identical protocol as the screening protocol. Instead of filling each well with a distinct miRNA, half of the 96-well plate is filled with a scrambled miRNA mimic control, for example, pre-miR miRNA precursor molecule negative control #2, Thermo Fisher Scientific, #AM17111. The other half of the plate is filled with a positive control. For example, miR-193a-3p can be used if screening for miRNA regulating CCND1. If no miRNAs are reported in the literature, use TargetScanHuman as a prediction data source to select appropriate miRNAs. A positive control should reduce normalized luciferase activity by at least 50%. Moreover, it is crucial to monitor the expression levels of Firefly and Renilla luciferase. Optimally, raw expression should be within twofold differential expression between Firefly and Renilla luciferase with luciferase raw counts exceeding 50,000. If raw expression diverts substantially between Firefly and Renilla luciferase, it may be an indication of truncation of the gene during the insertion process into the U2OS genome. 10. Depending on the ambition of the project, a miRNA-library may range from 40 to >2000 human miRNA mimics. Multiple vendors, including Thermo Fisher Scientific, sell miRNA libraries. If you customize your library, it is easier to include positive and negative controls in specific wells of the library plate. If using a premade library, positive controls are usually included while negative controls need to be discussed upfront. 11. There should never be any cells in the outer columns and rows. If placing cells in the outer columns and rows, edge effect, due to increased evaporation, will result in altered results compared

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to cells placed in row B-G, column 2–11. Always fill outer columns and rows with cell culture medium to avoid edge effects on subsequent columns and rows. 12. Lysing cells in medium without phenol red and without FBS, preequilibrated to room temperature, will increase luciferase raw expression and reduce variability between library plate readings. To make sure medium is completely equilibrated to room temperature, put medium bottle at room temperature at least 1 h before starting the assay. 13. Dual-Glo solution is highly susceptible to air bubbles. Excess amounts of air bubbles will ruin the reproducibility of your luciferase readings. After adding the Firefly luciferase solution to the 96-well tissue culture plate the total volume is 90 μL. By transferring 85 μL to the assay plates, you minimize the risk of introducing air into the pipette tip, and thereby into the assay plate. References 1. Bartel DP (2009) MicroRNAs: target recognition and regulatory functions. Cell 136 (2):215–233. https://doi.org/10.1016/j.cell. 2009.01.002 2. Guo H, Ingolia NT, Weissman JS, Bartel DP (2010) Mammalian microRNAs predominantly act to decrease target mRNA levels. Nature 466(7308):835–840. https://doi. org/10.1038/nature09267 3. Karginov FV, Conaco C, Xuan Z, Schmidt BH, Parker JS, Mandel G, Hannon GJ (2007) A biochemical approach to identifying microRNA targets. Proc Natl Acad Sci U S A 104 (49):19291–19296. https://doi.org/10. 1073/pnas.0709971104 4. Lal A, Thomas MP, Altschuler G, Navarro F, O’Day E, Li XL, Concepcion C, Han YC, Thiery J, Rajani DK, Deutsch A, Hofmann O, Ventura A, Hide W, Lieberman J (2011) Capture of microRNA-bound mRNAs identifies the tumor suppressor miR-34a as a regulator of growth factor signaling. PLoS Genet 7(11): e1002363. https://doi.org/10.1371/journal. pgen.1002363 5. Chi SW, Zang JB, Mele A, Darnell RB (2009) Argonaute HITS-CLIP decodes microRNAmRNA interaction maps. Nature 460 (7254):479–486. https://doi.org/10.1038/ nature08170 6. Selbach M, Schwanhausser B, Thierfelder N, Fang Z, Khanin R, Rajewsky N (2008) Widespread changes in protein synthesis induced by microRNAs. Nature 455(7209):58–63. https://doi.org/10.1038/nature07228

7. Baek D, Villen J, Shin C, Camargo FD, Gygi SP, Bartel DP (2008) The impact of microRNAs on protein output. Nature 455 (7209):64–71. https://doi.org/10.1038/ nature07242 8. Doench JG, Sharp PA (2004) Specificity of microRNA target selection in translational repression. Genes Dev 18(5):504–511. https://doi.org/10.1101/gad.1184404 9. Hydbring P, Wang Y, Bogorad RL, Yin H, Anderson DG, Li C, Sicinski P (2017) Identification of cell cycle-targeting microRNAs through genome-wide screens. Cell Cycle 16 (23):2241–2248. https://doi.org/10.1080/ 15384101.2017.1380132 10. Hydbring P, Wang Y, Fassl A, Li X, Matia V, Otto T, Choi YJ, Sweeney KE, Suski JM, Yin H, Bogorad RL, Goel S, Yuzugullu H, Kauffman KJ, Yang J, Jin C, Li Y, Floris D, Swanson R, Ng K, Sicinska E, Anders L, Zhao JJ, Polyak K, Anderson DG, Li C, Sicinski P (2017) Cell-cycle-targeting microRNAs as therapeutic tools against refractory cancers. Cancer Cell 31(4):576–590. e578. https:// doi.org/10.1016/j.ccell.2017.03.004 11. Hydbring P, Malumbres M, Sicinski P (2016) Non-canonical functions of cell cycle cyclins and cyclin-dependent kinases. Nat Rev Mol Cell Biol 17(5):280–292. https://doi.org/10. 1038/nrm.2016.27 12. Robins H, Li Y, Padgett RW (2005) Incorporating structure to predict microRNA targets. Proc Natl Acad Sci U S A 102

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(11):4006–4009. https://doi.org/10.1073/ pnas.0500775102 13. Kertesz M, Iovino N, Unnerstall U, Gaul U, Segal E (2007) The role of site accessibility in microRNA target recognition. Nat Genet 39 (10):1278–1284. https://doi.org/10.1038/ ng2135 14. Long D, Lee R, Williams P, Chan CY, Ambros V, Ding Y (2007) Potent effect of target structure on microRNA function. Nat Struct Mol Biol 14(4):287–294. https://doi. org/10.1038/nsmb1226 15. Grimson A, Farh KK, Johnston WK, GarrettEngele P, Lim LP, Bartel DP (2007) MicroRNA targeting specificity in mammals: determinants beyond seed pairing. Mol Cell 27 (1):91–105. https://doi.org/10.1016/j. molcel.2007.06.017

16. Zhao Y, Samal E, Srivastava D (2005) Serum response factor regulates a muscle-specific microRNA that targets Hand2 during cardiogenesis. Nature 436(7048):214–220. https:// doi.org/10.1038/nature03817 17. Zhao Y, Ransom JF, Li A, Vedantham V, von Drehle M, Muth AN, Tsuchihashi T, McManus MT, Schwartz RJ, Srivastava D (2007) Dysregulation of cardiogenesis, cardiac conduction, and cell cycle in mice lacking miRNA-1-2. Cell 129(2):303–317. https://doi.org/10.1016/j. cell.2007.03.030 18. Lewis BP, Burge CB, Bartel DP (2005) Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell 120 (1):15–20. https://doi.org/10.1016/j.cell. 2004.12.035

Chapter 8 Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis Mardo Ko˜ivom€agi Abstract Protein kinases are common elements in multiple signaling networks, influencing numerous downstream processes by directly phosphorylating specific target proteins. During the cell cycle, multiple complexes, each comprising one cyclin and one cyclin-dependent kinase (Cdk), function to regulate the orderly progression of cell cycle events. The mechanisms of cyclin–Cdk mediated control have, in part, been established through biochemical experiments involving the purification of cyclin and Cdk proteins to evaluate the activity of a given complex toward its target substrate proteins. Here I present a detailed procedure to simplify the preparation of cyclin–Cdk complexes by purifying them as a single fusion molecule with a 1:1 molar ratio and a detailed protocol for performing reconstituted kinases assays with the purified complexes. This methodology has allowed us to measure the activity and specificity of all budding yeast cyclin–Cdk1 complexes toward the model substrate histone H1. In addition, it has allowed us to perform kinase assays with a panel of purified human cyclin–Cdk complexes to analyze their specificity toward the retinoblastoma protein (Rb) and map the substrate cyclin-Cdk kinase docking interactions between Rb and human G1–Cdk complex. This chapter is focused on purification of cell cycle cyclin–Cdk complexes, but also affords a generalizable framework that can be adapted to other cyclin-dependent kinases like transcriptional cyclin–Cdks or any other multisubunit enzyme complexes. Taken together, the described workflow is a powerful and flexible biochemical platform for solving long-standing biological questions and has potential value in synthetic biology and in therapeutic discovery. Key words Kinase purification, Kinase fusion, Kinase assay, Cyclin, Cdk, Cell cycle

1

Introduction

1.1 Cyclin–Cdk Complexes in Cell Cycle Control

The cyclin-dependent kinases are a family of proline-directed serine (S) or threonine (T) protein kinases distinguished mainly by their association with cyclins [1]. Cyclin binding causes conformational changes in Cdk that confer kinase activity to the cyclin–Cdk

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-1538-6_24 Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021, Corrected Publication 2021

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complex [2]. Active kinase complexes are able to phosphorylate S or T residues in optimal S/T-P-x-K/R (where x is any amino acid) or suboptimal S/T-P consensus motifs [3, 4]. Unlike in higher organisms, in budding yeast a single Cdk (Cdk1) regulates all phases of the cell division cycle. Cdk1 is activated by different cyclins at different cell cycle phases [5]. In G1, Cln3, 2 and 1 bind to Cdk1. During the S and G2 phase Cdk1 is activated by Clb5/6 or Clb3/4, respectively. In M phase Cdk1 is activated by Clb2/1. In higher eukaryotes, at least six Cdks have been shown to be involved directly in cell cycle control [6, 7]. Contrary to budding yeast, each Cdk interacts with a specific subset of cyclins. For example, Cdk1 and Cdk2 both show wider preference in their choice of cyclin partners, binding with cyclins A, B and E, whereas Cdk4 and Cdk6 are activated by D-type cyclins [8]. 1.2 Purification of Cyclin–Cdk Complexes for In Vitro Analysis

Protein kinase activity studies require sensitive assays. Many protein kinases are multisubunit complexes (like cyclin–Cdk) and a substrate can be phosphorylated by more than one kinase, which makes it difficult to assay for the activity of a specific protein kinase. Therefore, the ability to purify and assay the activity of kinase complexes in the test tube has been useful in identifying new substrates and comparing the specificity between different complexes. So far cyclin– Cdk complexes have been purified using different expression systems and purification methods like affinity chromatography with yeast p13suc1 protein [9], by expressing cyclins in bacteria and the Cdk subunit in yeast [10] or in insect cells followed by one-step affinity purification using affinity tagged Cdk subunit [11, 12]. Tagging the Cdk subunit for affinity purification can result in complexes with different cyclins. Attaching an affinity tag to the cyclin subunit can overcome this limitation and yield more homogeneous preparations [4, 13–15]. Purification procedure described in this chapter allows production of cyclin–Cdk complexes with one molecule of cyclin and one molecule of Cdk from budding yeast, using affinity tag attached to the cyclin subunit and a linker region between the cyclin and Cdk subunit. This makes the protocol suitable for the purification of any specific cyclin–Cdk complex of interest.

1.3 Use of Suitable Linker Between Cyclin and Cdk

Linkers are short peptide sequences between protein domains. In an artificial setting it is possible to engineer linkers to fuse two or more proteins. Linkers are often composed of flexible residues like glycine and serine allowing the protein domains to move relative to one another. Linker length can determine if two proteins sterically interfere with each other. Flexible glycine and serine rich linkers have been used to create fusion proteins between cyclin D1 and Cdk4 [16, 17], cdc13 and cdc2 [18, 19] and Cdc28 and Cln3 [20]. In theory it might be possible to fuse budding yeast G1 cyclins Cln1–3 to Cdk1 without any linker region as they have a long intrinsically disordered C-terminal tail. I have tested different linker versions of Cln3-L-Cdk1 (where L is a 0
, 1
, 3
and 6
glycine-

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Fig. 1 The experimental approach for measuring substrate phosphorylation by purified cyclin- L–Cdk1 complexes. (a) A schematic illustrating the design principles of cyclin–Cdk complex fused via linker region, generating cyclin- L–Cdk. (b) A schematic of one step affinity chromatography procedure resulting in near homogeneous purification of cyclin–Cdk complex. (c) A schematic of in vitro kinase assay with purified cyclinL–Cdk complex and model substrate histone H1. (d) The autoradiograph of in vitro phosphorylation of a Cdk model substrate histone H1 by the indicated budding yeast cyclin- L–Cdk1 complexes

serine linker (GGGGS) between Cln3 and Cdk1) and have not noticed any differences compared to wild-type Cln3-Cdk1 function. For S/G2/M cyclin- L–Cdk1 complexes I have used a GGGGS linker with three repeats. 1.4 Advantages of the Cyclin–Cdk1 Fusion Strategy in Purification

Purification of cyclin–Cdk complexes has been described previously by many groups. Cyclin–Cdk complexes have been purified from bacteria, yeast, insect or mammalian cells using different experimental methodologies as described above. The method described in this chapter employs the use of a flexible linker region between cyclin and Cdk to achieve a ratio of 1:1 between cyclin and Cdk subunit (Fig. 1a). Using the cyclin–Cdk1 fusion purification strategy we have been able to purify all budding yeast cyclin–Cdk1 complexes and measure their activity towards histone H1, a model substrate for measuring kinase activity. Moreover, we have been able to purify human cyclin–Cdk complexes from yeast cells to analyze their specificity toward Rb and model substrate histone H1 [21].

1.5 Cyclin–Cdk Fusion Complex Purification and Activity Measurement

The basic workflow for cyclin–Cdk fusion complex purification and activity measurement is as follows. l

Construct a DNA sequence encoding the fusion complex of interest.

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Integrate the constructed DNA into an expression vector in a manner suitable for the expression of cyclin–Cdk complex.

l

Transform the constructed expression vector into an appropriate yeast strain.

l

Culture the yeast strain in a manner to express the fusion complex of interest.

l

Purify the recombinantly produced fusion complex of interest.

l

Determine the activity of the purified fusion complex in a kinase assay.

Materials Buffers

Prepare all solutions using ultrapure water and analytical grade reagents. Prepare and store all reagents at room temperature (unless indicated otherwise).

2.1.1 Lysis Buffer for Controlling Cyclin- L– Cdk1 Expression

1. Urea lysis buffer: 20 mM TRIS–HCl, pH 7.5, 7 M urea, 2 M thiourea, 65 mM CHAPS, 70 mM DTT, 50 mM sodium fluoride, 110 mM β-glycerophosphate, 1 mM sodium orthovanadate, pH 10, 1 mM phenylmethylsulfonyl fluoride.

2.1.2 Buffers for the Purification of Cyclin- L–Cdk1 Complex (See Note 1)

1. Cell lysis buffer (CLB): 50 mM Hepes–KOH, pH 7.6, 1 M KCl, 1 mM MgCl2, 1 mM EGTA, 5% (w/v) glycerol, 0.25% Tween 20, 1 TIU (trypsin inhibitor unit) aprotinin, 1 mM phenylmethylsulfonyl fluoride, 1 μg/mL pepstatin A, 1 μg/ mL mM bestatin, 1 μg/mL leupeptin, 1 μM benzamidine, 1 μM sodium orthovanadate, pH 10, 5 mM sodium fluoride, 80 mM β-glycerophosphate. 2. Wash buffer 1 (WB1): 50 mM Hepes–KOH, pH 7.6, 1 M KCl, 1 mM MgCl2, 1 mM EGTA, 5% (w/v) glycerol, 0.25% Tween 20. 3. Wash buffer 2 (WB2): 50 mM Hepes–KOH, pH 7.6, 1 M KCl, 1 mM MgCl2, 1 mM EGTA, 5% (w/v) glycerol. 4. Elution buffer (EB): 50 mM Hepes–KOH, pH 7.6, 0.25 M KCl, 1 mM MgCl2, 1 mM EGTA, 5% (w/v) glycerol, 0.2 mg/ mL competitor peptide.

2.1.3 Kinase Assay Buffers

1. 5
Kinase buffer + ATP: 425 mM HEPES, pH 7.4, 750 mM NaCl, 25 mM MgCl2, 2.5 mM ATP. 2. Kinase assay composition: 50 mM HEPES, pH 7.4, 150 mM NaCl, 5 mM MgCl2, 20 mM imidazole. 0.01 mg/mL FLAG peptide, 2% glycerol, 0.05 mM EGTA, 0.2 mg/mL BSA, 500 μM ATP (with 2 μCi of [γ-32P] ATP added per reaction (PerkinElmer), 1–10 nM recombinant cyclin- L–Cdk1 purified

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from yeast, 1–5 μM model substrate H1 from bovine or recombinant substrate purified from bacteria, 500 nM recombinant Cks1 purified from bacteria. 3. Coomassie Staining Solution: 10% acetic acid, 40% methanol, 1 g/L Coomassie R250 powder (0.1%). 4. Coomassie Destaining Solution: 10% acetic acid, 40% methanol. 2.2

Reagents

1. Anti-FLAG M2 Affinity Gel. 2. 3FLAG competitor peptide for cyclin- L–Cdk complex elution with the sequence of MDYKDHDGDYKDHDIDYKDDDDK (see Note 2). 3. Monoclonal anti-Flag M2 mouse antibody. 4. Anti-phospho-cdc2 (Tyr15) rabbit antibody. 5. Anti-phospho-cdc2 (Thr161) rabbit antibody. 6. ATP, [γ-32P]-adenosine triphosphate, labeled on the gamma phosphate group with 32P (PerkinElmer). This is a radioactive product for kinase assay. 7. Bovine histone H1. 8. Bovine serum albumin (BSA). 9. Silver Stain Plus Kit.

2.3

Media

1. YPD media: peptone 20 g/L, yeast extract 10 g/L, dextrose 2% w/v, (Agar 2% w/v for plates). 2. SC-Leucine media: yeast nitrogen base without amino acids 1.7 g/L, ammonium sulfate 5 g/L, amino acid mix—leucine 1.3 g/L, dextrose 2% w/v, or raffinose 2% w/v, and galactose 2% w/v (agar 2% w/v for plates).

2.4 Expression Vectors

1. pMK9 pGAL1-pRS425. 2. pMK178 pGAL1–3Flag-CLN3-L-CDK1-pRS425. 3. pMK184 pGAL1–3Flag-CLN2-L-CDK1-pRS425. 4. pMK186 pGAL1–3Flag-CLN1-L-CDK1-pRS425. 5. pMK361 pGAL1–3Flag-CLB5-L-CDK1-pRS425. 6. pMK362 pGAL1–3Flag-CLB6-L-CDK1-pRS425. 7. pMK363 pGAL1–3Flag-CLB3-L-CDK1-pRS425. 8. pMK364 pGAL1–3Flag-CLB4-L-CDK1-pRS425. 9. pMK366 pGAL1–3Flag-CLB2-L-CDK1-pRS425.

2.5

Yeast Strains

1. MKy9 (W303) MATa ksp1::TRP1. 2. MKy12 MKy9 [pGAL1–3Flag-CLN3-L-CDK1-pRS425]. 3. MKy38 MKy9 [pGAL1–3Flag-CLN2-L-CDK1-pRS425]. 4. MKy226 MKy9 [pGAL1–3Flag-CLN1-L-CDK1-pRS425].

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5. MKy770 MKy9 sic1::URA3. 6. MKy759 MKy770 [pGAL1–3Flag-CLB5-L-CDK1-pRS425]. 7. MKy760 MKy770 [pGAL1–3Flag-CLB6-L-CDK1-pRS425]. 8. MKy761 MKy770 [pGAL1–3Flag-CLB3-L-CDK1-pRS425]. 9. MKy762 MKy770 [pGAL1–3Flag-CLB4-L-CDK1-pRS425]. 10. MKy763 MKy770 [pGAL1–3Flag-CLB2-L-CDK1-pRS425]. 2.6

Equipment

1. Poly-Prep Gravity Flow Chromatography Columns. 2. Amersham Typhoon system. 3. Amersham Typhoon storage phosphor screen cassette.

2.7

Software

1. ImageQuant TL software. 2. Prism software.

3

Methods The following workflow describes the purification of budding yeast Clb5-L-Cdk1 complex. The same workflow is easily adaptable to any cyclin-dependent kinase complex of interest.

3.1 Construction of Yeast Expression Plasmid

For expressing Clb5-L-Cdk1 complex in budding yeast, plasmid pMK361 was constructed using the pRS42
series backbone [22]. Using the pMK9 based construct the GAL1 promoter is used to express the cyclin- L–cdk1 complex in yeast under galactose inducible control. The coding regions of the Saccharomyces cerevisiae Clb5 and Cdk1 were amplified from genomic DNA. N-terminal 3 FLAG affinity tag and linker region between the coding regions of cyclin and Cdk1 were introduced by overlap extension polymerase chain reaction (OE-PCR). Example oligonucleotides for generating 3FLAG-Clb5-L-Cdk1 sequence to insert it into pGAL1-pRS425 vector cut with BamHI and XhoI are as follows (the underlined sequence is the gene specific part of the oligonucleotide): 3FLAG-Clb5_forward 50 ACAGAAGGATCCATGGATTATAAAGATCATGATGGCGAT TATAAAGATCATGATATTGATTATAAAGATGATGATGA TAAGATGGGAGAGAACCACGACCATGAGCAGAG 30 Clb5-L_reverse 50 GCATAGCAACTTTCAAAATCTATTTAATCTTAAG GGTGG CGGTGGCAGCGGCGGTGGCGG TAGCGGTGGCGGTGGCAGC 30 L-Cdk1_forward 50 GGTGGCGGTGGCAGCGGCGGTGGCGGTAGCGGTGGC GGTGGCAGCATGAGCGGTGAATTAGCAAATTAC 30

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Cdk1_reverse 50 TTCTGTGAGCTCTTATGATTCTTGGAAGTAGGGGTGG 30 This step leads to the generation of pMK361 (pGAL1–3FlagClb5-L-Cdk1-pRS425) vector to express the Clb5-L-Cdk1 complex (see Note 3). 3.2 Yeast Transformation and Characterization of Expression

pMK361 plasmid was introduced into budding yeast MKy770 strain by lithium acetate transformation to generate MKy759 (see Notes 4 and 5). For that, cells were plated on a selective SCD-Leucine plate and incubated at 30  C for 48 h. Single clones were streaked out to a new selective plate and incubated at 30  C overnight. To test the expression of the cyclin L–Cdk1 complex, cells were inoculated into 5 mL of SC without leucine media with 2% raffinose. Clb5-L-Cdk1 expression was induced by adding 2% galactose to the media after culture reached to OD600 0.6. Cells were harvested by centrifugation (4100
g at 4  C for 5 min), lysed using urea lysis buffer, and analyzed with Western blot using anti-Flag and anti-Cdk1 antibodies. Positive clones, which showed Clb5-L-Cdk1 expression when compared to the empty vector (pGAL-pRS425) control was picked for large-scale induction (see Note 6).

3.3 One Potential Strategy for Inducing Large-Scale Cultures

1. A single colony of positive transformant from MKy759 transformation is inoculated into 50–100 mL of SC without leucine media with 2% raffinose and grown at 30  C for about 24 h before use. 2. 4 L of SC without leucine with 2% raffinose is inoculated with cultures from step 1 and the cells are grown at 30  C until OD600 is 0.6–0.8. 3. To start the induction, weigh and add an appropriate amount of galactose (20 g/L) powder to the cultures. 4. Grow the induced culture at 30  C for 2–4 h, depending on the protein stability. For example, for yeast complexes, the optimal time for G1 cyclin- L–Cdk1 complexes is 2 h, while the optimal time for S/G2/M-L-Cdk1 complexes is 4 h. 5. The cells are harvested by centrifugation (4100
g at 4  C for 10 min) using large centrifuge bottles in a Thermo Scientific Sorvall centrifuge or similar. 6. The harvested cells are washed once with ice-cold 1
PBS and transferred into 50 mL conical tubes. The cells are pelleted by centrifugation (4100
g at 4  C for 10 min), supernatant removed and pellets snap-frozen in liquid nitrogen. 7. The pellet is weighed before storing at 80  C. Alternatively, in the case of smaller culture volumes, a filter membrane attached to a vacuum pump may be used to harvest cells.

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3.4 Purification Procedure of Cyclin-Dependent Kinase Complex from Budding Yeast

The one-step FLAG-tag affinity chromatography purification protocol presented in this chapter is visualized in Fig. 1b and is based on the previously published protocol, used for HA-tagged Cln2Cdk1 purification [14]. 1. Prepare the cell lysis buffer by adding the phosphatase and protease inhibitors immediately before use. It is recommended to use ~20–25 mL of CLB per 15 g pellet (approximately 3–5 volumes of buffer to yeast powder). 2. Frozen MKy759 pellets from step 7, Subheading 3.3 are ground into a powder using a mortar and pestle, that were prechilled overnight at 80  C. Liquid nitrogen is added between successive rounds of grinding to ensure cells stay frozen until a light-colored powder is obtained. This can be stored indefinitely at 80  C. Alternative mechanical disruption methods can be used, but it is important to maintain temperature control and minimize bubble formation in the lysate to avoid enzyme inactivation. 3. Scoop powder from the mortar into a cooled glass beaker (stored overnight at 20  C) and thaw on bench until the light-colored yeast powder begins to darken and melt at the edges. Add CLB and stir vigorously on a magnetic stirrer at 4  C for 10 min or until all powder is dissolved. 4. Collect crude lysate sample for Western blot analysis (see Note 7). 5. To separate different biological materials, the homogenate is centrifuged twice. First 10 min at 17,000
g at 4  C using a Thermo Scientific Sorvall centrifuge with SS-34 fixed angle rotor or similar. For the second spin pour the supernatant into the ultracentrifuge tubes and centrifuge 60 min at 180,000
g at 4  C using Beckman Ultracentrifuge with Ti-70 fixed angle rotor. In case ultracentrifugation is not available, spinning 60 min at 38,000
g at 4 using Thermo Scientific Sorvall centrifuge with SS-34 fixed angle rotor or similar can be performed. 6. Before the end of the second spin prepare the beads. To equilibrate beads, wash the bead slurry twice, resuspending in CLB and collected at 500
g, room temperature for 2 min. In the case of 3FLAG affinity tag, Anti-FLAG M2 Affinity Gel from Sigma-Aldrich can be used. The volume of affinity gel to use in any particular purification procedure depends on the expression level of the complex, the collected pellet size, and the binding capacity specified by the manufacturer. 7. Collect cleared lysate sample for Western blot analysis. 8. After the second spin, remove supernatant to a new tube using a Pasteur glass pipette to avoid mixing the lipid layer on top and

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residual cell debris at the bottom and add equilibrated affinity beads to the cleared lysate. Rotate the tube for 2–4 h end over end at 4  C. 9. Wash the beads in the tube two times with 15 mL of WB1, by centrifuging at 500
g 4  C for 2 min. After the second spin, remove WB1 conservatively, leaving about 5 mL liquid on the beads. 10. Load the column with a 5 mL mixture of WB1 and beads from the previous step and let drain by gravity flow. Make sure that all the affinity gel is transferred from the tube and in the column. 11. Collect flow through sample for Western blot analysis. 12. Wash the beads in the column with WB1, using 10 gel volumes of wash buffer 1 at 4  C. 13. Collect column wash 1 sample for Western blot analysis. 14. Repeat procedure step 12. 15. Wash the beads in the column with WB2, using 5 gel volumes of wash buffer 2 at 4  C. 16. Collect column wash 2 sample for Western blot analysis. 17. Add 1 affinity bead volume of EB to equilibrate beads with an elution buffer. Bring the column to room temperature and wait 30 min. 18. Collect elution 0 sample for Western blot analysis. 19. Move the column back to 4  C. Add 1 affinity bead volume of EB to elute the enzyme complexes and collect the flow through into a prechilled low protein binding tube. 20. Collect elution 1 sample for Western blot analysis. 21. Aliquot cyclin–Cdk1 complexes into prechilled 0.5 mL low protein binding tubes and immediately snap-freeze using liquid nitrogen. Aliquot volume depends on the purpose. For in vitro kinase assays aliquots can be between 12–25 μL. It is not recommended to aliquot enzymes in less than 10 μL because stability and activity is reduced when these smaller volumes are stored at 80  C, which will result in tube-totube variability (see Note 8). 22. Close the column and add 1 volume of EB to elute the enzyme complexes a second time. Incubate the column with EB on top of the beads for at least 1 h or overnight. To collect the second elution, open the column and collect the flow through into a low protein binding tube. 23. Collect elution 2 samples for Western blot analysis. 24. Repeat procedure step 21. 25. Collect some of the affinity gel from the column, add an equal amount of sample buffer and boil.

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26. Run a gel with all the collected samples (see Note 9) and analyze by Western blot using antibodies against Flag affinity tag, cyclin, or Cdk subunit. This will help to estimate the total amount of purified cyclin- L–Cdk1 complex. In addition, it is possible to evaluate the amount of active/inactive enzyme in the preparation using antibodies against the two known regulatory phosphorylation sites of Cdk1 in budding yeast: the inhibitory site at position Tyr19 and activating site at position Thr169. 3.5 In Vitro Kinase Assay to Measure the Activity of Purified Cyclin- L–Cdk1 Complexes

The activity of purified enzyme complexes can be tested by performing an in vitro kinase assay (Fig. 1c, d). Before starting to analyze purified cyclin L–Cdk1 complex activity toward a substrate of interest, it is recommended to test its activity toward a model substrate (see Note 10). The kinase assay protocol presented in this chapter is based on the previously published protocol [4, 23]. Step-by-step protocol. This step-by-step protocol is for an in vitro kinase assay performed in final volume of 20 μL at room temperature. 1. Prepare tubes for the enzyme mix, ATP mix, 5
kinase buffer and the reaction tubes for substrates of interest. When performing kinase assays with multiple time points, prepare sample collection tubes for each time point with 8 μL of 2
SDS-PAGE loading buffer (see Note 11). 2. Mix the enzyme mixture, which consists of 5
kinase buffer + ATP, BSA, recombinant Cks1, recombinant cyclinL–Cdk1 enzyme and H2O (see Table 1). Do not add the enzyme at this point (see Note 12). 3. Mix the ATP mix, which consists of 5
kinase buffer, ATP [γ-P32] and H2O (see Table 1). Do not add the ATP [γ-P32] at this point. 4. Add substrate protein to the reaction tubes, for example Cdk1 model substrate histone H1 from bovine (see Table 1). 5. Take all the tubes to the radioactive work area behind a protective screen. 6. Retrieve enzyme aliquots from 80  C and ATP [γ-P32] from 4  C. 7. Add an appropriate amount of enzyme (determined by Coomassie G 250 staining and Western blot with antibodies against the inhibitory site at position Tyr19 and the activating site at position Thr169) to the enzyme mix and start a timer for 3 min (this time can vary depending on the type of enzyme). This step is required if the enzyme of interest autophosphorylates itself and the size of enzyme overlaps with the size of substrate protein (see Note 13).

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Table 1 The example composition of an in vitro kinase assay using the purified cyclin- L–Cdk1 complexes Kinase assay reaction Master mix Enzyme mix (8 μL)

ATP mix (4 μL)

Substrate (8 μL)

Enzyme/elution buffer

5
Kinase buffer +ATP

Substrate protein/elution buffer

BSA

ATP [γ-32]

H 2O

5
Kinase buffer + ATP

H2O

Cks1 H2O

8. During the 3 min incubation add an appropriate amount of radioactive ATP [γ-P32] to the ATP mix (see Note 14). 9. After a 3 min incubation, add an appropriate amount of the ATP mix to the enzyme mix. This generates the master mix (see Table 1). 10. Add an appropriate amount of the master mix to the reaction tube with substrate to start the in vitro kinase assay (see Table 1). 11. Collect timepoints of interest by taking 8 μL out of the reaction tube and mixing it into sample tubes with 2
-SDS-PAGE loading buffer. 12. Run the samples in an SDS-PAGE gel. 13. Stain the gel(s) with Coomassie R250 staining solution, then destain the gel(s) using Coomassie destaining solution. 14. Dry gel(s) on top of Whatman filter paper. 15. Expose the gel(s) by placing it inside the Typhoon Storage Phosphorimager cassette with Phosphor screen (both from GE Healthcare). Exposure time will vary from hours to days depending on the enzyme of interest and the amount and freshness of ATP [γ-P32]. 16. Scan the Phosphor (GE Healthcare).

screen

using

Typhoon

machine

17. Quantify the results by using ImageQuant TL (GE Healthcare) software or similar. See Fig. 2a and Table 2 for an example quantification of a gel from an in vitro kinase assay with different budding yeast cyclin- L–Cdk1 complexes. Quantified results can be visualized using Prism software (GraphPad) or similar (Fig. 2b).

Fig. 2 The example quantification of an autoradiograph from the in vitro kinase assay with purified cyclinL–Cdk1 complexes. (a) The autoradiograph of in vitro phosphorylation of a model substrate histone H1 by the indicated budding yeast cyclin L–Cdk1 complexes. ImageQuant (GE Healthcare) software, included with Typhoon (GE Healthcare) scanner, was used to quantify the phosphorylated histone H1 signal intensity. Boxes 1–8 drawn around the phosphorylated histone H1 signal allow the band quantitation. For background subtraction the box 9 average intensity was used. For data analysis see Table 2. (b) Quantified data blotted from Table 2. Equal amounts of cyclin L–Cdk1 complexes were used in the kinase assay with model substrate histone H1. The amounts of enzyme used were calculated from total Western blot signal against Flag-tag affinity tag Table 2 The example data analysis of an in vitro kinase assay using the purified cyclin L–Cdk1 complexes

Box Name 1

Cln3-LCdk1

2

Cln2-LCdk1

3

Volume 370,497.37

b

Area

Background subtractiona

12,556

1,368,693.8

95.37

14,352

Cln1-LCdk1

1,136,835.49

78.95

14,400

860,355.49

0.28

4

Clb5-LCdk1

4,450,086.96 269.31

16,524

4,132,826.16

1.32

5

Clb6-LCdk1

2,459,680.53 158.16

15,552

2,161,082.13

0.69

6

Clb3-LCdk1

7,318,537.15 382.13

19,152

6,950,818.75

2.23

7

Clb4-LCdk1

5,895,612.36 309.64

19,040

5,530,044.36

1.77

8

Clb2-LCdk1

15,797,486.16 591.75

9

Background

1,075,236.74

19.2

129,422.17

Normalized valuesb

29.51

Volume-Area
Average Intensity Normalized to Clb2-Cdk1 values [4]

a

Average Intensity

1,093,135.4

26,696 15,284,922.96 56,000

0.04 0.35

4.9

Cyclin-Dependent Kinase Fusion Purification

4

107

Notes 1. CLB without protease and phosphatase inhibitors, WB1 and WB2 can be prepared in advance and stored at 4  C. It is recommended to prepare the EB before use, but it can be prepared in advance and stored at 20  C. 2. In case of HA-tag affinity purification the following competitor peptide can be used: 2HA peptide with the sequence of CYPYDVPDYAGY PYDVPDYAG. 3. The LexA promoter system derived from Ottoz et al., 2014 (pLexA-pRS425) can be used to express cyclin- L–Cdk1 complexes under estradiol inducible control [24]. 4. All Saccharomyces cerevisiae strains used in this protocol were derived from the W303 background. The specific nature of the enzyme complex of interest determines the purification method and specific strain to be used. For example, if the activity of the kinase complex of interest is weak, then a strain harboring an analog sensitive (as) Cdk1 allele (Cdk1as) at the endogenous locus can be used to test in a subsequent in vitro kinase assay if all the activity of the purified complex originates from the affinity purified enzyme. The Cdk1as has a mutation in the active site of the kinase (F84G) that results in the kinase being sensitive to bulky ATP-analogs [25]. This means that incubation with such bulky ATP-analogs in an in vitro kinase assay will selectively and irreversibly inhibit any potential contaminating Cdk1 activity. 5. The purification of B-type (Clb1–6) cyclin–Cdk1 complexes requires the deletion of SIC1 gene. Sic1 is a stoichiometric cyclin-dependent kinase inhibitor that regulates the G1/S transition in budding yeast, by binding to and inhibiting Clb-Cdk1 complexes. 6. Specific amounts needed for purification vary depending on the cyclin–Cdk1 complex and the amount of enzyme needed. For example, Cln2-L-Cdk1 purification requires pellets obtained from 10 L of induced culture, whereas Clb5-L-Cdk1 can be purified from pellets obtained from 4 L of induced culture. 7. When combined, KCl and SDS sample buffer can cause precipitation, which makes it difficult to load the gel. I have found it helpful to keep samples on a 100  C heat block before loading the gel. 8. The purified kinase complexes can be stored without detectable loss of activity for several months at 80  C. Although I have not thoroughly examined the long-term stability, in general I have found the enzyme to be stable provided repeated freezing

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and thawing is avoided. To keep the subsequent in vitro kinase assays constant, I have avoided freeze/thaw cycles and use each aliquot only once. 9. Samples can be saved from the following steps: crude lysate, cleared lysate, flow through, column wash 1, column wash 2, elution 0, elution 1, elution 2, and beads after elution. The enzyme purity can be analyzed by SDS-PAGE (8% or 10% gels). Gels can be stained with colloidal Coomassie Brilliant Blue G-250 or complexes can be detected by Silver Stain Plus Kit (Bio-Rad). The extent of purity can be analyzed by quantification of signal after scanning the stained gel with a Typhoon scanner or similar machine. 10. Histone H1 or H1 peptide with the sequence (PKTPKKAKKL) derived from the H1 protein is the best in vitro model substrate at present for measuring the biochemical activity of different purified cyclin- L–Cdk1 complexes. 11. If possible, reaction aliquots should be taken at two time points, for example at 8 and 16 min. This makes it easier to determine if the reaction progression is linear. 12. Budding yeast recombinant Cks1 protein can be purified following the procedure described previously [26]. 13. In the case of measuring phosphorylation of the substrate protein of interest, concentrations should be kept in the range of 0.5–3 μM (in the linear [S] versus v0 range, severalfold below the estimated KM value). Usually it is enough for the signal strength if the low nanomolar range (0.5–2 nM) concentration of the cyclin- L–Cdk1 complex is used in the assay. 14. ATP [γ-P32] has a half-life of 14.3 days. If ATP is fresh then 0.2 μL per 20 μL reaction is enough for signal detection. With older ATP, more can be added to the reaction or the exposure time can be raised.

Acknowledgments I am grateful for Benjamin Topacio, who helped to optimize several steps in the workflow described here and for giving feedback. I am thankful for Sirle Saul and Matthew Swaffer for reading the manuscript. References 1. Morgan DO (2007) The cell cycle principles of control. New Science Ltd, London 2. De Bondt HL, Rosenblatt J et al (1993) Crystal structure of cyclin-dependent kinase 2. Nature 363(6430):595–602

3. Songyang Z, Blechner S et al (1994) Use of an oriented peptide library to determine the optimal substrates of protein kinases. Curr Biol 4 (11):973–982

Cyclin-Dependent Kinase Fusion Purification ˜ ivom€agi M, Valk E, Venta R et al (2011) 4. Ko Dynamics of Cdk1 substrate specificity during the cell cycle. Mol Cell 42(5):610–623. https://doi.org/10.1016/j.molcel.2011.05. 016 5. Hartwell LH, Mortimer RK et al (1973) Genetic control of the cell division cycle in yeast: V. Genetic analysis of cdc mutants. Genetics 74(2):267–286 6. Malumbres M, Harlow E et al (2009) Cyclindependent kinases: a family portrait. Nat Cell Biol 11(11):1275–1276 7. Satyanarayana A, Kaldis P (2009) Mammalian cell-cycle regulation: several Cdks, numerous cyclins and diverse compensatory mechanisms. Oncogene 28(33):2925–2939 8. Hochegger H, Takeda S et al (2008) Cyclindependent kinases and cell-cycle transitions: does one fit all? Nat Rev Mol Cell Biol 9 (11):910–916 9. Brizuela L, Draetta G, Beach D (1987) p13suc1 acts in the fission yeast cell division cycle as a component of the p34cdc2 protein kinase. EMBO J 6(11):3507–3514 10. Jackson LP, Reed SI, Haase SB (2006) Distinct mechanisms control the stability of the related S-phase cyclins Clb5 and Clb6. Mol Cell Biol 26(6):2456–2466. https://doi.org/10.1128/ MCB.26.6.2456-2466.2006 11. Costanzo M, Nishikawa JL, Tang X, Millman JS, Schub O, Breitkreuz K, Dewar D, Rupes I, Andrews B, Tyers M (2004) CDK activity antagonizes Whi5, an inhibitor of G1/S transcription in yeast. Cell 117(7):899–913. https://doi.org/10.1016/j.cell.2004.05.024 12. Nash P, Tang X, Orlicky S et al (2001) Multisite phosphorylation of a CDK inhibitor sets a threshold for the onset of DNA replication. Nature 414:514–521 13. Honey S, Schneider BL, Schieltz DM, Yates JR, Futcher B (2001) A novel multiple affinity purification tag and its use in identification of proteins associated with a cyclin-CDK complex. Nucleic Acids Res 29(4):E24. https:// doi.org/10.1093/nar/29.4.e24 14. McCusker D, Denison C, Anderson S et al (2007) Cdk1 coordinates cell-surface growth with the cell cycle. Nat Cell Biol 9(5):506–515. https://doi.org/10.1038/ncb1568 ˜ ivom€agi M, Ord M, Iofik A, Valk E, Venta R, 15. Ko Faustova I, Kivi R, Balog ER, Rubin SM, Loog M (2013) Multisite phosphorylation networks as signal processors for Cdk1. Nat Struct Mol Biol 20(12):1415–1424 16. Konstantinidis AK, Radhakrishnan R, Gu F, Rao RN, Yeh WK (1998) Purification,

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characterization, and kinetic mechanism of cyclin D1. CDK4, a major target for cell cycle regulation. J Biol Chem 273 (41):26506–26515. https://doi.org/10. 1074/jbc.273.41.26506 17. Rao RN, Stamm NB, Otto K et al (1999) Conditional transformation of rat embryo fibroblast cells by a cyclin D1-cdk4 fusion gene. Oncogene 18(46):6343–6356. https:// doi.org/10.1038/sj.onc.1203009 18. Coudreuse D, Nurse P (2010) Driving the cell cycle with a minimal CDK control network. Nature 468(7327):1074–1079. https://doi. org/10.1038/nature09543 19. Swaffer MP, Jones AW, Flynn HR, Snijders AP, Nurse P (2016) CDK substrate phosphorylation and ordering the cell cycle. Cell 167 (7):1750–1761.e16. https://doi.org/10. 1016/j.cell.2016.11.034 20. Yahya G, Parisi E, Flores A, Gallego C, Aldea MA (2014) Whi7-anchored loop controls the G1 Cdk-cyclin complex at start. Mol Cell 53 (1):115–126. https://doi.org/10.1016/j. molcel.2013.11.015 21. Topacio BR, Zatulovskiy E, Cristea S et al (2019) Cyclin D-Cdk4,6 drives cell-cycle progression via the retinoblastoma protein’s C-terminal helix. Mol Cell 74(4):758–770.e4. https://doi.org/10.1016/j.molcel.2019.03. 020 22. Christianson TW, Sikorski RS, Dante M, Shero JH, Hieter P (1992) Multifunctional yeast high-copy-number shuttle vectors. Gene 110 (1):119–122. https://doi.org/10.1016/ 0378-1119(92)90454-w ˜ ivom€agi M, Valk E, Venta R et al (2011) 23. Ko Cascades of multisite phosphorylation control Sic1 destruction at the onset of S phase. Nature 480(7375):128–131. https://doi.org/10. 1038/nature10560 24. Ottoz DS, Rudolf F, Stelling J (2014) Inducible, tightly regulated and growth conditionindependent transcription factor in Saccharomyces cerevisiae. Nucleic Acids Res 42(17): e130 25. Bishop AC, Ubersax JA, Petsch DT et al (2000) A chemical switch for inhibitorsensitive alleles of any protein kinase. Nature 407(6802):395–401. https://doi.org/10. 1038/35030148 26. Reynard GJ, Reynolds W, Verma R, Deshaies RJ (2000) Cks1 is required for G(1) cyclincyclin-dependent kinase activity in budding yeast. Mol Cell Biol 20(16):5858–5864. https://doi.org/10.1128/mcb.20.16.58585864.2000

Chapter 9 Optimizing Cell Synchronization Using Nocodazole or Double Thymidine Block Arif A. Surani, Sergio L. Colombo, George Barlow, Gemma A. Foulds, and Cristina Montiel-Duarte Abstract Cell synchronization is crucial when studying events that take place at specific points of the cell cycle. Several chemical agents can be used to achieve the cell culture synchronization but not all type of cells respond equally to a given concentration of these drugs. Here we describe a simple optimization method to select concentrations and timings for nocodazole or thymidine treatments using fluorescence staining. In addition, we provide detailed protocols to arrest an asynchronous culture of either suspension or adherent cells in G1/S or in G2/M. Key words Cell cycle, Synchronization, Nocodazole, Thymidine, Suspension cells, G1/S, G2/M

1

Introduction Cell proliferation is a highly regulated process, key for the growth and development of any organism. Cell proliferation involves the division of one cell into two identical daughter cells, in a process termed “mitosis” and the stages a cell undergoes for this to happen are called “phases of the cell cycle.” During the cell cycle, the phases involving duplication of the genetic material (S phase) and segregation into two daughter cells (M phase) are intermitted by three gap phases (G0, G1, and G2). The sequential transition through these five phases ([G0] ! G1 ! S ! G2 ! M ! [G0]) defines the eukaryotic cell cycle [1], and a complex transcriptional and posttranslational regulatory system coordinates critical molecular and biochemical events for the progression through cell division. There are many reasons to study the cell cycle and having an interest in gene and protein expression is one of them: a periodic gene expression pattern has been confirmed in different eukaryotic cells including yeast [2], primary human fibroblast [3], and immortal human HeLa cells [4]. Furthermore, chemotherapy has been

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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shown to modulate the global gene expression profile and cell cycle kinetics [5]. But in order to detect and study these temporal changes in the transcriptomic and proteomic cells’ profile, it is imperative to use a synchronous population of cells. Cell synchronization is a method that brings cultured cells at different phases of the cell cycle to the same stage. The synchronized cells progress through the phases of the cell cycle as a relatively uniform cohort, aiding in the understanding of the changes occurring during a particular phase. Different approaches have been used for batch synchronization of a cell culture and could be broadly classified into physical and chemical methods. Physical methods separate the cells in a particular phase of the cell cycle based on cell size, density, fluorescent labeling, light scattering, and/or attachment to the growth matrix (e.g., flask). Commonly used physical methods for cell synchronization include centrifugal elutriation, fluorescent-activated cell sorting, mitotic detachment, and contact inhibition. These physical methods, unlike chemical methods, have minimal effects on cell metabolism and allow cells to progress through different cell cycle phases without any perturbations. However, these methods are not universally applicable to different cell lines and synchronising the cells based on size (centrifugal elutriation) is limited by variability in cell synchrony, expensive instrumentation and technical complexities [6, 7]. Chemical methods involve the use of pharmacological agents which block the progression of the cells at a specific phase. Common drugs that arrest the cells in S phase include excess thymidine, aphidicolin, and methotrexate. These drugs affect the synthesis of DNA and inhibit DNA replication [8]. Excess thymidine has been shown to allosterically inhibit ribonucleotide reductase, altering the deoxyribonucleotide pool and halting DNA replication [9]. It arrests the cells at the G1–S boundary and synchronizes the cell cycle at early S phase following release [10]. Consecutive exposure to thymidine increases the population of synchronous cells [8]. The initial exposure to excess thymidine for 24 h halts the cells at the S phase of cell cycle. Following the release, the cells arrested in G1/S and early S phase would progress through G2/M phase and cells blocked in late S phase would reenter G1 phase. The repeated exposure to thymidine would collect most of the cell population at G1–S interface and the release would result in cells progressing through the cell cycle synchronously. Another category of chemical blockade is mediated by drugs that disrupt polymerization of microtubules that form the mitotic spindle, and is comprised of nocodazole, colcemid, and colchicine. These agents prevent cytokinesis, arresting the cells in G2/M phase [11, 12]. Cells could be also arrested at G0/G1 phase by serum starvation, depletion of isoleucine in the medium or using lovastatin (HMG-CoA reductase inhibitor) [13].

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It has been argued that the use of chemical agents for synchronization is associated with undesirable side effects such as cytotoxicity, growth imbalance, disruption of metabolic processes and deregulated expression of cyclin proteins [14–16]. It is therefore of upmost importance, that cell survival is asserted when choosing any of these chemical agents, assessing their concentration and time of exposure. Here, we describe the steps necessary to optimize cell cycle synchronization using nocodazole or double thymidine block in a concise manner so the readers can adapt these to their particular cells of interest. We use the chronic myeloid leukemia cell line TCC-S that grows in suspension, and later provide modifications to the protocol necessary when using adherent cells, using the prostate cancer cell line DU145 and the epithelial HeLa S3 cells as examples.

2 2.1

Materials Cell Cycle Drugs

1. 100 mM Thymidine: Dissolve 242.2 mg of thymidine in 100 mL of autoclaved double-distilled water, place in a water bath at 37  C to ensure complete dissolution of thymidine, sterilize the solution by passing through a 0.2 μm filter and store in aliquots at 20  C. 2. 1 mg/mL Nocodazole: Dissolve 10 mg of nocodazole in 10 mL cell-culture grade DMSO. Aliquot and freeze (20  C) for up to 1 year.

2.2 DNA Staining for Viability or FACS Analysis

1. ReadyProbes® Cell Viability Imaging Kit, NucBlue/NucGreen (ThermoFisher Cat. No. R37609). 2. 70% (v/v) ethanol: Take 70 mL of absolute ethanol (see Note 1) and add distilled water to bring the volume to 100 mL. Store at 20  C. 3. 1 mg/mL RNase A solution: Dissolve 10 mg of RNase A (DNase-free RNase A) in 10 mL of nuclease-free water. Aliquot in 1.5 mL microcentrifuge tubes and store at 20  C. 4. 1 mg/mL Propidium Iodide (PI, see Note 2): Prepared in double-distilled water, store in the dark at 4  C. 5. PI staining solution: 250 μL of staining solution is required per sample. It is made by mixing 17.5 μL of PI solution, 35 μL of RNase A solution and 197.5 μL of PBS. The solution should be prepared fresh and used immediately. The final concentration of PI and RNase A used per sample are 50 and 100 μg/mL, respectively. 6. FACS tubes (Sarstedt, catalog number: 55.1578). 7. Beckman Coulter™ ISOTON™ II Diluent (Fischer Scientific, product code: 12754878).

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Cell Culture

Cells are routinely grown in a humidified atmosphere at 37  C with 5% CO2. 1. Cell lines: TCC-S, suspension cell line grown in complete RPMI medium; DU145 and Hela S3, adherent cell lines grown in complete DMEM medium. 2. Complete RPMI medium: RPMI 1640 with 25 mM HEPES and L-glutamine (e.g., Lonza Cat. No. LZBE12-115F) supplemented with 10% FBS. 3. Complete DMEM medium: Dulbecco’s Modified Eagle’s Medium (DMEM) with high glucose and GlutaMAX™ Supplement (Thermo Fischer Scientific Cat. No. 10566016), supplemented with 10% FBS. 4. Sterile 1 Phosphate-Buffered Saline (PBS). 5. Trypsin-EDTA (0.25%).

3

Methods Whilst the thymidine concentration used for cell cycle arrest is quite standard (2 mM) for a variety of cells, the range of concentrations for nocodazole is wider (40–200 ng/mL) as nocodazole exposure can affect cell viability. Therefore, it is essential that suitable concentrations are determined on each cell line of interest prior to attempting cell synchronization.

3.1 Preliminary Assessment of Nocodazole Concentrations

1. Seed TCC-S cells in a 24-well plate at a concentration of 0.5  106 cells/mL, using 500 μL per well. 2. Leave cells incubating overnight. 3. Treat three wells with nocodazole to a final concentration of 100 ng/mL and another three wells to a final concentration of 200 ng/mL (see Note 3). Add also the equivalent volume of DMSO to three wells (controls). This will be the 24 h treatment time. 4. Four hours later, treat another six wells in the same manner. This will be the 20 h point. 5. Four hours later, treat another six wells in the same manner. This will be the 16 h point. 6. The next day (24 h after the first treatments started) add 1 drop of each NucBlue® Live and NucGreen® Dead dyes to individual wells. 7. Incubate for 30 min at room temperature, covering the plate with foil to protect from light. 8. Assess cell staining under a fluorescence microscope using standard DAPI (live cells) and GFP (dead cells) filters (Fig. 1). Choose the concentration with the least effects on viability (lower proportion of green signal).

Optimising Cell Synchronisation Nocodazole: 100 ng/mL

16h

20h

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200 ng/mL

24h

24h

Fig. 1 Nocodazole treatments. TCC-S were treated with nocodazole at 100 or 200 ng/mL for 16–24 h and the cell viability was assessed with the ReadyProbes™ Cell Viability Imaging Kit. Green cells represent cells with a compromised cell membrane (dead) whilst live cells are stained blue. Cell viability clearly decreases with prolonged times of exposure and with the higher nocodazole concentration

A

B G2/M G1

G1 G2/M

Nocodazole treatment

Thymidine treatment

Fig. 2 Optimizing (a) nocodazole and (b) thymidine treatment time. In TCC-S cells, treating the cells with nocodazole (100 ng/mL) for 18 h was the option that produced a lower G1 peak and arrested the highest number of cells in the G2/M phase. In the case of thymidine, the treatment time of 24 h was the most successful at arresting the cells at the G1–S boundary 3.2 Selection of Nocodazole and Thymidine Timings

All centrifugations are performed at room temperature unless otherwise stated. The steps described correspond to one set of experiments (see Fig. 2)—be sure to perform in duplicate or triplicate, as needed. 1. Seed TCC-S cells at a concentration of 1  106 cells/mL in four T25 flasks with 8 mL of complete RPMI medium and incubate overnight (see Note 4). 2. In two of the flasks, add either 8 μL of DMSO (control, asynchronous culture) or 8 μL of 1 mg/mL nocodazole to achieve a final concentration of 100 ng/mL in the flask, and return to incubator. Remove 2 mL of the cell suspension from each flask 14, 16 and 18 h after treatment, transferring cells to individual 15 mL tubes and continuing the process at step 9.

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3. To the remaining two flasks, add either 160 μL of autoclaved water (control, asynchronous cells) or 160 μL of the thymidine stock (100 mM) to achieve a 2 mM thymidine concentration in the culture (first block), and incubate for 24 h. 4. After 24 h, collect the cell suspension from the flask and centrifuge at 200  g for 5 min to pellet cells. Discard supernatant. 5. Wash the cells with sterile PBS and centrifuge at 200  g for 5 min to pellet cells. Discard supernatant. 6. Reseed the entire cell pellet in the same volume of fresh media (8 mL) without thymidine (block release) and incubate for 12 h. 7. Add 160 μL of the thymidine stock (100 mM) to the synchronized flask (second block) and 160 μL autoclaved water to the control flask and incubate for 24 h. 8. Remove 2 mL of the cell suspension from control and treated cell culture flasks and transfer to individual 15 mL tubes. 9. Centrifuge at 200  g for 5 min to pellet the cells then decant supernatant and flick to resuspend cells. 10. Add 2 mL of PBS, mix cells, centrifuge at 200  g for 5 min, decant supernatant and flick to resuspend cells. 11. Add 500 μL of ice-cold 70% ethanol to all tubes to fix the cells and store at 20  C for a minimum of 30 min (see Notes 5 and 6). 12. Centrifuge to pellet the cells at 300  g for 5 min. Then carefully remove the ethanol supernatant with a pipette. 13. Wash with 2 mL PBS then centrifuge at 300  g for 5 min, decant the supernatant and flick to resuspend cells. 14. Repeat wash with 2 mL PBS, then centrifuge (300  g, 5 min) decant supernatant and flick to resuspend cells. 15. Add 100 μL PBS, resuspend the cells and transfer 100 μL to a labeled FACS tube. 16. Add 250 μL of PI staining solution to the cell suspension in the FACS tube and incubate for 30 min at room temperature in the dark (see Notes 7 and 8). 17. Transfer the cells to FACS tubes. 18. Add 100 μL ISOTON™ diluent and analyze on a flow cytometer (in our case, Beckman Coulter Gallios). 19. Optimize flow cytometer acquisition settings using unstained cells and asynchronous PI stained cells. The flow rate should be set to low and the acquisition rate should not exceed 500 cells/s.

Optimising Cell Synchronisation

Seed 3 x 105 in 3 mL required cell culture medium and incubate overnight at 37°C and 5% CO2

Add 2 mM thymidine and incubate the cells for 24 hours

Release the cells from thymidine arrest and incubate with complete medium for 12 hours

Repeat 2 mM thymidine treatment and incubate again for 24 hours

Release the cells from thymidine arrest and collect cells at 0, 3, 6, 12 and 24 hours

0 hr

Add 100 µL ISOTON™ diluent and analyse on FACS instrument

Incubate for 30 min at room temperature in dark (covered with foil) and transfer the cells to FACS tubes

Centrifuge the cells to remove ethanol and wash twice with 2 mL PBS. Resuspend in 1 mL PBS and add 250 µL of PI staining solution

Centrifuge the cells to remove media, gently disperse the cell pellet and cells with 1 mL of ice-cold 70% ethanol

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3 hr

6 hr

12 hr

24 hr

Trypsinise to deatch cells and collect in 15 mL falcon tube

Fig. 3 Graphical representation of the double thymidine block cell synchronization process for adherent cell line DU145 3.3 Collection of Cells After Release

Once the cell culture is synchronized and the chemical removed (release step), cells can be collected every 2–3 h, or as often as desired, and analyzed following steps 8–19 in the protocol above to assess the progression of the cells through the cell cycle or to collect RNA or protein samples. Here, we provide an adapted protocol for adherent cells synchronization (see Fig. 3) and we assess the outcome of the cell cycle progression in Fig. 4 (after double thymidine block) and in Fig. 5 (after nocodazole treatment). 1. Prepare the required cell culture medium and warm it up to 37  C before use. 2. For DU145, plate the cells at 3  105 cells per well in 3 mL of complete prewarmed medium. This should result in approximately 30–40% confluency the following day. The seeding density needs to be optimized for the cell line under investigation. 3. Leave the cells in the incubator overnight. 4. Add 2 mM thymidine (6 μL from 100 mM stock) in each sample well. Add 6 μL of autoclaved double distilled water in the control well as vehicle control. Incubate the cells for 24 h. 5. After incubation, release the cells from thymidine by washing twice with 2 mL of prewarmed PBS. Incubate with complete medium for 12 h.

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n sy

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Fig. 4 Synchronization of DU145 cells after double thymidine block. DU145 were arrested at the G1–S border after a double thymidine block and their progression through the different cycle phases after the release was determined using the fluorescence intensity and can be observed in (a). The percentage of cells in each phase of the cell cycle at different time points is presented in (b). Sample data was acquired on a Beckman Coulter Gallios and subsequent analysis was performed using Kaluza analysis software. Thy thymidine

Optimising Cell Synchronisation

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Release: 0 h

Release: 2 h

Release: 4 h

Release: 6 h

Release: 8 h

Release: 10 h

Release: 12 h

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Cell cycle distribution (%)

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100 75 50 25 0 0

2

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8 10 12 24

Hours after Noc release Fig. 5 HeLa S3 cell cycle arrest with nocodazole. HeLa S3 cells were treated following the protocol described in this chapter, with the modification of maintaining the 100 ng/mL nocodazole treatment for 14 h only [17]. After release, it can be observed in (a) how these cells transitioned through the cell cycle, with the percentages of cells in each phase of the cycle represented in (b). Noc nocodazole

6. Following incubation, repeat second thymidine treatment by adding 2 mM thymidine to each well and incubate again for 24 h. 7. After incubation, release the cells by washing twice with 2 mL of prewarmed PBS and incubate in prewarmed fresh complete medium.

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8. Collect the cells at selected times, for example at 0, 3, 6, 12 and 24 h after release: remove the complete medium and rinse the well with 2 mL of prewarmed PBS. Add prewarmed TrypsinEDTA (0.2 mL/well) and incubate at 37  C for 5 min to detach the cells. Neutralize Trypsin-EDTA by adding 1 mL of complete medium and collect the detached cells in a 15 mL tube. 9. Centrifuge the cells at 300  g for 5 min at room temperature, discard the supernatant and fix the cells by resuspending the pellet in 1 mL of ice-cold 70% ethanol. 10. Follow from step 12 in Subheading 3.2.

4

Notes 1. There is no need to use molecular biology grade ethanol for fixation. 2. As PI causes irritation, a good option is to buy an already-made PI solution (for example, Sigma-Aldrich, catalog number: P4864-10ML). 3. Depending on the information available in the literature, this range of concentrations can be varied or expanded. 4. This initial overnight incubation in complete media after splitting the cells has the purpose to get the culture in the exponential phase, with cells dividing. The length of this incubation can be anything between 16 and 24 h and it is worth considering the timing of the treatments given afterward. For example, if an 18 h nocodazole treatment is going to be given, cells could be split in the late afternoon (not the morning) and the treatment given the day after at 3 pm, to be collected at 9 am the following day. 5. The incubation with ethanol can be prolonged overnight or over the weekend. It is a good stop in the protocol to collect all different tubes in the experiment together and then continue with the protocol. 6. To avoid precipitation and clumping, it is important to add ethanol gradually (dropwise) whilst vortexing the cells. 7. Cover the tubes with foil. Propidium iodide is a red-fluorescent counterstain that is photosensitive. 8. The PI staining solution contains the RNase. Reduced times of incubation with RNase will make your peaks less differentiated when plotting fluorescence intensity against cell count. In our hands, 30 min is the optimal time for clearly defined peaks.

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References 1. Lodish H, Berk A, Zipursky SL, Matsudaira P, Baltimore D, Darnell J (2000) Molecular cell biology, 4th edn. WH Freeman, New York, NY 2. Spellman PT, Sherlock G, Zhang MQ, Iyer VR, Anders K, Eisen MB, Brown PO, Botstein D, Futcher B (1998) Comprehensive identification of cell cycle-regulated genes of the yeast Saccharomyces cerevisiae by microarray hybridization. Mol Biol Cell 9:3273–3297 3. Cho RJ, Huang M, Campbell MJ, Dong H, Steinmetz L, Sapinoso L, Hampton G, Elledge SJ, Davis RW, Lockhart DJ (2001) Transcriptional regulation and function during the human cell cycle. Nat Genet 27:48–54 4. Whitfield ML, Sherlock G, Saldanha AJ, Murray JI, Ball CA, Alexander KE, Matese JC, Perou CM, Hurt MM, Brown PO et al (2002) Identification of genes periodically expressed in the human cell cycle and their expression in tumors. Mol Biol Cell 13:1977–2000 5. Kokkinakis DM, Liu X, Neuner RD (2005) Modulation of cell cycle and gene expression in pancreatic tumor cell lines by methionine deprivation (methionine stress): implications to the therapy of pancreatic adenocarcinoma. Mol Cancer Ther 4:1338–1348 6. Banfalvi G (2008) Cell cycle synchronization of animal cells and nuclei by centrifugal elutriation. Nat Protoc 3:663–673 7. Cooper S (2003) Rethinking synchronization of mammalian cells for cell cycle analysis. Cell Mol Life Sci 60:1099–1106 8. Banfalvi G (2011) Overview of cell synchronization. Methods Mol Biol (Clifton, NJ) 761:1–23

9. Bjursell G, Reichard P (1973) Effects of thymidine on deoxyribonucleoside triphosphate pools and deoxyribonucleic acid synthesis in Chinese hamster ovary cells. J Biol Chem 248:3904–3909 10. Chen G, Deng X (2018) Cell synchronization by double thymidine block. Bio Protoc 8: e2994 11. Sonoda E (2006) Synchronization of cells. Subcell Biochem 40:415–418 12. Samake´ S, Smith LC (1996) Synchronization of cell division in eight-cell bovine embryos produced in vitro: effects of nocodazole. Mol Reprod Dev 44:486–492 13. Jackman J, O’Connor PM (1998) Methods for synchronizing cells at specific stages of the cell cycle. Curr Protoc Cell Biol 8:3.1–3.20 14. Urbani L, Sherwood SW, Schimke RT (1995) Dissociation of nuclear and cytoplasmic cell cycle progression by drugs employed in cell synchronization. Exp Cell Res 219:159–168 15. Kung AL, Zetterberg A, Sherwood SW, Schimke RT (1990) Cytotoxic effects of cell cycle phase specific agents: result of cell cycle perturbation. Cancer Res 50:7307–7317 16. Kung AL, Sherwood SW, Schimke RT (1993) Differences in the regulation of protein synthesis, cyclin B accumulation, and cellular growth in response to the inhibition of DNA synthesis in Chinese hamster ovary and HeLa S3 cells. J Biol Chem 268:23072–23080 17. Tudzarova S, Colombo SL, Stoeber K, Carcamo S, Williams GH, Moncada S (2011) Two ubiquitin ligases, APC/C-Cdh1 and SKP1-CUL1-F (SCF)-β-TrCP, regulate glycolysis during the cell cycle. Proc Natl Acad Sci U S A 108(5278):5283

Chapter 10 Highly Synchronous Mitotic Progression in Schizosaccharomyces pombe Upon Relief of Transient Cdc2-asM17 Inhibition Pawan Singh, Lenka Halova, and Iain Michael Hagan Abstract Synchronized progression of a cell population through the cell division cycle supports the biochemical and functional dissection of cell cycle controls and execution. The concerted behaviour of the population reflects the attributes of each cell within that population. The reversible imposition of a block to cell cycle progression at the G2–M boundary through transient inactivation of the Cdk1-Cyclin B activating phosphatase, Cdc25, with the temperature sensitive cdc25-22 mutant, has been widely used to study fission yeast mitosis and DNA replication. However, the biology of the compromised Cdc25-22 phosphatase generates significant division abnormalities upon release from mitotic arrest. We show how reversible inhibition of Cdc2-asM17, with the ATP analog 3-BrB-PP1, generates higher levels of synchrony with timing and morphology much more reminiscent of a normal division. We also describe a version of the H1 kinase assay of Cdk1-Cyclin B activity that is widely used to monitor mitotic progression which does not require radiolabeled ATP. Key words Fission yeast, Cell cycle, Mitosis, Synchronization, G2–M, Cdc2, Cdc25, P-Thr-Pro-101, Histone H1 kinase assay, S. pombe

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Introduction A unicellular S. pombe cell divides by medial fission to generate two equally sized daughter cells that grow by tip extension until they reach the same size as their parent, whereupon these daughters, themselves, divide [1]. This mode of growth makes fission yeast an excellent model system for cell cycle studies [2] because a cell’s length reflects its position in the cell cycle. Consequently, the isolation of small, recently divided cells from asynchronous cultures by density gradient, or elutriation centrifugation, generates secondary cultures that progress synchronously through the cell division cycle [3–5]. Biochemical and functional changes within these synchronized cultures reflect the changes accompanying the division of a single cell within that population.

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Size selection has been complemented by induction synchronization procedures, in which transient exposure to a cell cycle arrest accumulates the entire population at a defined point in the cell cycle, before relief of the block releases the population to progress synchronously through the next phases of the cycle. Successful approaches for fission yeast include transient mating pheromone arrest at the start of the cycle, early S phase arrest through nucleotide depletion, and reversible inactivation of cell cycle regulators [6–8]. In this chapter, we focus upon transient arrest at the G2–M transition as a means of synchronizing cell cycle progression. Passage through the fission yeast G2–M transition is determined by the activity of Cdc2-Cyclin B kinase. Phosphorylation of the Cdc2 catalytic subunit by Wee1 kinase restrains Cdc2-Cyclin B activity until the inhibitory phosphate is removed by Cdc25 phosphatase [9]. The newly activated Cdc2-Cyclin B triggers a feedback loop that engages polo kinase to repress Wee1 and enhance Cdc25 activities to ensure that the commitment to mitosis is a rapid and irreversible, bistable, switch [10, 11]. Temperature sensitive mutants, in which a shift from 25 to 36
C inactivates products of cell division cycle (cdc) genes, underpinned the definition of this switch [12]. The transient arrest of cdc2-33 to synchronize mitotic progression was an early exploitation of the reversibility of these mutants [13, 14]. However, this innovation in mitotic synchronization coincided with the realization that, alongside its pivotal role in G2–M control, Cdc2 acted in G1 to control passage through the first rate limiting step in cell cycle progression: START [15]. Thus, release from cdc2-33 arrest simultaneously promotes progression through synchronized S (25% of the population) and M (75% of the population) phases, to superimpose S phase data upon the mitotic dataset [15]. We therefore turned to the Cdc2-Cyclin B phosphatase Cdc25 because it exclusively regulates Cdc2-Cyclin B control of the G2–M boundary. Transient inhibition of Cdc25 function resulted in highly synchronous progression through mitosis [16, 17] . We demonstrate the utility of this approach here with a nonradioactive adaptation of the longstanding histone H1 kinase assay of Cdk1-Cyclin B activity [18] (in other organisms, Cdc2 is referred to as Cdk1) (Fig. 1) and by staging mitotic progression with spindle morphology (Fig. 2a, b). The temperature shifts in cdc25-22 induction synchronization promote heat/cold stress responses during the initial transfer from 25 to 36
C and the return to 25
C that triggers mitotic commitment [19]. These temperature stresses induce transcription programs that enable the cell to accommodate the insult. This specific temperature response is superimposed upon a generic, core environmental stress response (CESR), that confers cross-stress protection against alternative insults [20]. Thus, cdc25-22 released M and S phases do not entirely reflect an unperturbed context, because

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Fig. 1 cdc25-22 induction synchronized mitosis. cdc25-22 temperature induction synchronization (Subheading 3.1) of a 500 mL culture grown overnight to 5  105 cells/mL in EMM2 with sampling for the p13suc1 bead precipitate histone H1 kinase assay (Subheading 3.4) and immunofluorescence microscopy (Fig. 2b). (a) The septation index (Subheading 3.3) and quantification of the Histone H1 kinase assay. (b) The three replicates used to calculate the activity plot in a: for each set the phospho-TP blot is upper and ponceau S stain to reveal the quantity of histone H1 is lower. These assays were quantified using the mask in Image J 1.53a, whereby a large rectangle covering the band and a significant area of background membrane above and below the band are captured. Standard processing removes the background noise from the signal. This value was normalized to the Histone H1 level in the ponceau S image. Arbitrary values of 1 were assigned to each time 0 dataset and the units represent fold change over this value of 1. The plot shows the average of three technical replicates  SD error bars. “+Ros 20” lane indicates a duplicate of the 20 min sample that includes 2 mM roscovitine (Cdc2 inhibitor) to demonstrate the specificity of the assay for Cdc2 activity. Basal activity at the 0 time point (compare t ¼ 0 with t ¼ 70–100) reflects the requirement for a modest level of Cdc2 activity in G2 to trigger new end take off (NETO, the activation of the growth of the cell tip that was generated by the preceding cytokinesis) and the activation of homologous recombination repair in G2 by low levels of Cdc2 after cells have exited S phase [1, 39, 40]

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Fig. 2 A comparison of division kinetics of cdc25-22 and cdc2-asM17 induction synchronization. cdc25-22 temperature and cdc2-asM17 induction synchronization in EMM2 with sampling for both calcofluor staining to monitor septation and immunofluorescence staining with the monoclonal antibody TAT-1 to stain the α-tubulin in the microtubules and DAPI to stain chromatin [41, 42]. The frequency of each mitotic stage was individually scored in counts of 200 cells according to the following criteria: metaphase defines all cells from spindle formation to the metaphase spindle that spans the nucleus, anaphase, a cell in which the spindle is longer than the interphase nucleus and yet there are no cytoplasmic microtubules and the post anaphase array (PAA) in which microtubules are nucleated from the equatorial microtubule organizing center, at the cytokinetic ring, as the spindle disassembles [41, 43, 44]. (a, b) Samples from the same 500 mL culture used for the histone H1 activity assays in Fig. 1. Panel (a) shows the septation profile, while (b) shows the frequency of different mitotic stages through the time course. Images from this dataset are shown in Figs. 3 and 5. (c) cdc2-asM17 septation profile in the EMM2 culture (squares) from which immunofluorescence is used to monitor mitotic progression in (d), alongside a septation profile of the same strain cultured for cdc2-asM17 induction synchronization in rich YES (circles) medium for comparison. For the EMM2 sample a 120 mL culture was grown and processed as outlined in the text whereas a 100 mL YES culture was exposed to 2M 3-BrB-PP1 for 3 h, rather than the 4 h 15 min experienced by the EMM2 culture. Septa were scored by calcofluor staining (Subheading 3.3). (d) The frequency of different mitotic stages through the cdc2-asM17 EMM2 synchronization shown in (c). A representative field from the 20 min time point is shown in Fig. 5d-f

changes associated with the stress responses are superimposed upon those arising from mitotic and S phase transit. Furthermore, the identical consensus SP/TP phosphorylation motif shared by MAP kinases and CDK-Cyclins [21], means that temperature induced bursts of stress-MAP kinase SP/TP phosphorylation will be

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superimposed upon the Cdc2-Cyclin B driven modifications at other SP/TP sites, making it hard to assign responsibility for specific phosphorylation events. After heat stress, the mitotic kinase polo is engaged in a recovery specific pathway to reverse the stress imposed arrest of mitotic commitment and so enable cells to return to mitosis [22]. Thus, the clarity of a dataset that is generated by the high levels of synchronous mitotic progression in the cdc25-22 induction synchrony approach will be compromised by the superimposition of the stress and recovery responses. We therefore exploited the transformative technology developed by Kevan Shokat, in which the catalytic pocket of a protein kinase of choice is enlarged by mutation/s to accommodate bulky ATP analogs. As the modified kinase becomes the only kinase in the cell with a catalytic pocket large enough to accept the bulky, nonhydrolyzable ATP analogs, these analogs can then be used to specifically inhibit the targeted kinase, while having minimal impact upon the rest of the proteome [23]. Sensitization of Wee1 kinase to analog inhibition in a cdc25-22 background generated a strain in which the temperature induced arrest of cell cycle progression at the G2–M boundary was rapidly reversed at the restrictive temperature by analog addition [24]. This approach avoids the cold shock associated with the return to permissive temperature in the traditional release regimen. Here we show how the mitotic fidelity in the approach to synchronizing mitotic progression by transient inhibition of Cdc2-asM17 introduced by Swaffer et al, for phosphoproteomic analysis of Cdc2 impact on the proteome [25], is a superior approach to cdc25-22 based mitotic synchronization (Fig. 2c, d). The cdc2-asM17 (K79E F84G) allele was isolated to overcome the temperature sensitivity and compromised sexual differentiation that accompanied analog sensitization for the canonical F84G cdc2-as1 allele [26, 27]. cdc2-asM17 grows normally in the absence of analog. Because the level of Cdc2 activity that is required to drive early cell cycle events is lower than that required to drive later events [25, 28, 29], cells can complete S at levels of ATP analog inhibition of Cdc2 that block mitotic commitment [29]. Thus, unlike cdc2-33 inactivation, that compromises commitment to both S and M [15], modest cdc2-asM17 analog inhibition can specifically block commitment to M [25]. Subsequent release from this arrest drives highly synchronous progression of the culture through mitosis [25], without either temperature stress associated with cdc25-22 induction synchronization (Fig. 2c, d). The ostensibly normal growth of cdc2-asM17, in the absence of analog, contrasts with the increased cell size at division of cdc25-22 at 25
C. Consequently, cdc2-asM17 cells do not reach the excessive lengths attained by cdc25-22 cells upon imposition of the G2 arrest before release [17]. This may account for the more efficient synchronization (peak spindles of 89% as opposed to 62% for cdc25-22

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Fig. 3 Double spindles in cdc25-22 released mitosis. Widefield epifluorescence microscopy images from the cdc25-22 samples processed 37 min after release for immunofluorescence microscopy in Fig. 2b showing the abnormal double spindles that are seen in around 30% of released divisions. In the absence of threedimensional reconstruction by high resolution microscopy, this figure remains a crude estimate of the frequency at which double spindles form as the range of angles of observation within which both spindles are clearly noted is limited. (a–c) metaphase, (d–f) anaphase. The large boxes in a and d show enlargements of the area highlighted by the smaller square. Scale bar 10m

(Fig. 2)) and the closer resemblance of a cdc2-asM17 released mitosis to a wild type division. Specifically, “double” spindles (Fig. 3) that are present in at least 30% of cdc25-22 released divisions are not seen in cdc2-asM17 mitoses and the frequency of double/multiple actin rings/septa (Fig. 4) declines from 32% to 8% in cdc2-asM17. Furthermore, extended metaphase spindles that straddle a clearly defined metaphase plate (Fig. 5a–c) are unique to cdc25-22 release, because cdc2-asM17 release mitoses (Fig. 5d–f) emulate wild-type divisions (Fig. 5g–l) in being well into anaphase B when cdc25-22 cells are yet to initiate it (compare the lengths of the spindles of the metaphase cdc25-22 release cells labeled “m” in Fig. 5a–c, with the length of the metaphase and early anaphase spindles labeled “m” and “a”, respectively, in both the cdc2asM17 synchronized release cells in Fig. 5d–f and wild type asynchronous cells in Fig. 5g–l). The striking normality of a cdc2-asM17 release mitosis begs the question as to why cdc25-22 release divisions are so deviant? Can we entirely attribute the abnormalities to the temperature stresses? Perhaps the extended duration of G2 in cycling cdc25-22 cells at 25
C combines with the G2–M arrest at 36
C to elevate levels of cyclin B, securin, and other anaphase promoting complex/cyclosome (APC/C) targets, and so thereby reduce the efficacy of the ensuing division. The need to degrade excessive levels of Cyclin B could delay the metaphase–anaphase transition to support

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Fig. 4 Double septa in cdc25-22 released divisions. (a) Calcofluor stained (Subheading 3.3) cdc25-22 cells 85 min after release from the G2–M block in the culture shown in Fig. 1. (b) Cells from the 60 min time point of the cdc2-asM17 EMM2 induction synchronized culture in Fig. 2c. Bright division septa bisect the cells at their equator [1]. The abnormal morphology of the double structures in the cdc25-22 cells in (a), contrasts with the normal, single, linear, septa bisecting the cdc2-asM17 cells in (b). Scale bar 10m

overextension of both the metaphase and anaphase B spindles [17]. Elevated Cyclin B levels could also trigger excessive Cdc2Cyclin B activity that could overwhelm the balance with the counteracting PP2A, PP1 and Cdc14 phosphatases [30]. However, the origins of the double spindles may lie elsewhere, because key elements of spindle pole body (SPB) duplication occur during the latter stages of anaphase [31]. As anaphase occurs after the release from the cdc25-22 block, it is hard to see how a G2–M arrest would support repeated rounds of SPB duplication. Perhaps the SPB duplication cycle mirrors the behavior of respiration. Meticulous studies from Novak and Mitchison showed how both carbon dioxide production and oxygen consumption continue to cycle in a cdc2-33 cell cycle arrest, even though the DNA duplication and segregation cycle has been arrested. Thus, respiration is normally entrained with cell cycle progression, however when the functions that we consider to be the heartbeat of the cell cycle are arrested by Cdc2 inactivation, respiration continues to cycle with its normal periodicity [32, 33]. Perhaps the SPB duplication cycle follows an independent cycle, or one that requires very low levels of Cdc2 activity that persist in a cdc25-22 arrest? Whatever the reason, the increase in SPB number is likely to impact upon the timing and execution of cytokinesis as the septum initiation network regulates the timing of cytokinesis from one of the two anaphase B SPBs [34]. Perhaps altered SIN signaling from the SPB is the cause of the cytokinetic abnormalities (Fig. 4)? cdc2-asM17 and cdc25-22 induction synchronization approaches each have specific attributes and issues. The need to avoid stress on the filter pad limits the scale of cdc2-asM17 synchronization cultures, whereas cdc25-22 temperature and cdc25-22 wee1-as8 inhibition synchronizations can be done on any scale. cdc25-22 temperature dependent synchronization avoids the

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Fig. 5 Over-extended metaphases spindles with a clear metaphase plate in a cdc25-22 released mitoses. Widefield epifluorescence microscopy images from the synchronized cdc25-22 (a–c), cdc2-asM17 (d–f) 27 and 20 min samples from Fig. 2b, d, respectively and wild type cells from a mid-log phase asynchronous culture (g–l). The yellow letter “m” identifies cells at metaphase, while “a” indicates early anaphase B and “a*” indicates late anaphase B. Three images of the same field are shown with the indicated fluorescence to identify the microtubules or chromatin. For wild type, two separate fields are shown (g–i and j–l). Note the highly condensed chromosomes at the metaphase plates of the cdc25-22 cells in (a) that are of equivalent length to the cdc2-asM17 early anaphase spindles in (d) and wild type (g, j). In fission yeast, the centromeres have already moved to the SPBs when the spindle elongates beyond the span of an interphase nucleus [45]. Scale bar 10m

expense of an ATP analog and its simplicity will maintain it as the first choice when superimposition of stress responses is not a concern. However, the size of cdc25-22 cells does mean that aspects of physiology that will change in response to this increase in size, will be better studied in a cdc2-asM17 release. Thus, cytoskeletal dynamics, or nuclear structure, which scales with cell size in fission

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yeast [35], would be better studied in the shorter, more normal cdc2-asM17 release division. Although cdc25-22 wee1-as8 synchronization needs the initial temperature shift to 36
C, it does avoid the second stress of the cold shock, which may account for the superior DNA replication profiles following wee1-as8 analog relief of the cdc25-22 arrest [24]. A major appeal of cdc25-22 wee1-as8 is the ability to combine one of the many temperature sensitive (ts) mutations with this background in order to perturb function in a highly synchronized S or M phase [24, 36]. However, cdc2-asM17 arrest may provide this as an additional option too, all be it with a superimposed heat shock. It should be possible to arrest cells at the G2–M boundary with cdc2-asM17 inhibition and shift the culture to 36
C for an hour before removal of the inhibitor at 36
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2

Materials Prepare all solutions using ultrapure water (prepared by purifying deionized water, to attain a sensitivity of 18 MΩ-cm at 25
C) and analytical grade reagents. Prepare all reagents at room temperature (unless indicated otherwise). Diligently follow all waste disposal regulations when disposing waste materials.

2.1 Cell Culture and Filtration

1. wild type (972 h), cdc25-22 h, cdc2-asM17 h (Laboratory stock numbers IH5975, IH13659, IH13757, respectively). 2. Edinburgh Minimal Medium 2 (EMM2) [37]. 3. Supplemented yeast extract (YES) [37]. 4. Bacterial Luria–Bertani (LB) broth. 5. LB broth +100 μg/mL ampicillin. 6. 3-BrB-PP1 (3-[(3-bromophenyl)methyl]-1-(1,1-dimethylethyl)-1H-pyrazolo[3,4-d]pyrimidin-4-amine) Toronto Research Chemicals: 50 mM stock in methanol; store at 80
C. 7. Two gyrating water baths: one at 36
C; the other at 25
C. 8. A 25
C incubator large enough to store filtration equipment and replacement media and flask. 9. Glass microfiber filters, binder free, GF/C Diameter: 47 mm (Whatman: WHA1822047). 10. Millipore filtration system: Top container (XX1014704), glass frit membrane support (XX1014702), aluminum spring clamp (XX1004703). 11. Heavy walled Buchner flask (1 L). 12. Vacuum source to filter at 120 mL/min. 13. Digital timer.

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2.2 Calcofluor Staining to Score the Septation Index

1. Phosphate Buffered Saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4.2H2O, 2 mM KH2PO4, pH 7.4. 2. Formalin. 3. 5 mg/mL Calcofluor (saturated solution) (see Note 1). 4. Coverslips (18  18 mm thickness 1.5) and microscope slides. 5. Epifluorescence microscope to visualize Calcofluor (excitation 300–412 nm with a peak at 347 nm and emission at 475 nm). 6. Forceps: Style 5 Dumont tweezers.

2.3 Generation of p13Suc1 Beads

1. Bacterial Lysis Buffer A (BLBA): 50 mM Tris-HCl (from a 1 M pH 8.0 stock solution), 100 mM NaCl, 15 mM MgSO4, 0.1 mM DTT, 0.1 mM PMSF. 2. 1 M Isopropyl β-D-1-thiogalactopyranoside (IPTG). 3. Lysozyme: 10 mg/mL, store at 4
C. 4. DNase1: 1 mg/mL prepared fresh on ice. 5. 100 mM PMSF: Prepare in methanol and store at 20
C. 6. 1 M DTT: store at 20
C. 7. Glutathione Sepharose 4B (GE Healthcare: 17-0756-01). 8. 20 mL bed volume chromatography column (Bio-Rad: 7321010) and stand to secure the column. 9. GST Sepharose elution buffer: 50 mM Tris–HCl (from a 1 M pH 8.0 stock solution), 10 mM reduced glutathione pH 8.0. 10. GST fusion protein storage buffer: 50 mM Tris-HCl (from a 1 M pH 7.4 stock solution), 25% glycerol. 11. Magnetic rack for 15 mL tubes (Thermo Fisher Scientific: DynaMag-15). 12. Rotating wheel. 13. Magnetic stirrer. 14. Dewar flask for liquid nitrogen. 15. Zeba spin desalting columns 7k MWCO 5 mL (Thermo Scientific: 89891). 16. Dialysis tubing (Bio Design dialysis tubing; Fisher Scientific: BID-010-020H) and clamps. 17. Pierce NHS-Activated magnetic beads (5 mL; Thermo Scientific: 88827). 18. Magnetic bead coupling buffer: 50 mM borate, pH 8.5. 19. Magnetic bead wash buffer A: ice-cold 1 mM HCl. 20. Magnetic bead wash buffer B: 0.1 M glycine, pH 2.0. 21. Magnetic bead quenching buffer: 3 M ethanolamine, pH 9.0. 22. Magnetic bead storage buffer: 50 mM borate buffer, pH 8.5, 0.05% sodium azide.

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1. Stop buffer: 10 mM EDTA, 150 mM NaCl, 50 mM NaF, 1 mM NaN3. 2. Cell lysis buffer (CLB): 25 mM Tris–HCl (from a 1 M pH 7.5 stock), 15 mM EGTA (from a 0.5 M pH 8.0 stock), 0.1 mM activated Na3VO4, 0.1 mM NaF, 15 mM MgCl2, 0.1% NP40. Just before use the following are added: 1 mM PMSF, 1 mM DTT, 60 mM Na-β-glycerol phosphate, protease inhibitor cocktail (complete mini EDTA free tablet, Roche: 11836170001) and 15 mM PNP (4-Nitrophenyl phosphate disodium salt hexahydrate, Sigma: P4744). 3. Okadaic acid: 250μM in DMSO. Light sensitive. 4. Recombinant Human Histone H1 (Sigma: H1917) 1 mg/mL. 5. 0.3 mM ATP. 6. Kinase assay buffer (KAB): 50 mM Tris-HCl (from a 1 M pH 7.5 stock), 10 mM MgCl2, 20 mM EGTA (from a 0.5 M pH 8.0 stock) and 5 mM KCl. Just before use add Na–βglycerol phosphate to 1 mM (from a freshly prepared 100 mM stock solution in water) and DTT to 1 mM. 7. Acid washed glass beads (Sigma: G8772). 8. Microfuge tubes: 1.7 mL prelubricated (Costar: 3207), screw cap 1.5 mL tubes (Star lab: E1415-2230), standard 1.5 mL microfuge tubes. 9. Yasui Kikai Multi-beads shocker. 10. Magnetic rack for 1.5 ml microfuge tubes: DynaMag™-2 Magnet (Thermo Fisher Scientific: 12321D). 11. 21 G needles and heat source. 12. Roscovitine: 50 mM stock in DMSO. 13. 4 LDS sample buffer with 10% β-mercaptoethanol. 14. 12% NuPAGE™ Bis-tris gel. 15. NuPAGE™ 20 MES buffer. 16. NuPAGE™ antioxidant. 17. 0.45μm nitrocellulose membrane. 18. Transfer buffer: 25 mM Tris, 192 mM glycine, 20% methanol. 19. Antibodies: Phospho-Threonine-Proline Mouse mAb (P-Thr-Pro-101) (Cell Signaling Technology: 9391S), antimouse IgG–alkaline phosphatase antibody (Sigma: A3688). 20. Nitro blue tetrazolium (NBT): prepare 50 mg/mL in 70% dimethylformamide (DMF). 21. 5-Bromo-4-chloro-3-indolyl 50 mg/mL in 30% DMF.

phosphate

(BCIP):

prepare

22. Alkaline phosphatase buffer: 0.1 M Tris–Cl pH 9.5, 0.1 M NaCl, 5 mM MgCl2.

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23. TBS-T: Tris-buffered saline (25 mM Tris, 2.7 mM KCl, 137 mM NaCl, pH 7.4) with 0.05% Tween 20. 24. Blocking buffer: 5% BSA, 100 mM lysine hydrochloride (Sigma: L5626) in TBS-T. 25. Ponceau S: 0.1%(w/v) in 5% acetic acid. 26. Rocker (for westerns): Stuart see-saw rocker SSL4.

3

Methods

3.1 cdc25-22 Induction Synchronization

1. Inoculate cdc25-22 in EMM2 from a fresh exponentially growing starter culture to generate a culture at 5  105 cells/mL after overnight growth in the 25
C water bath shaker at 250 rpm (see Notes 2 and 3). 2. Transfer culture to the 36
C water bath shaker at 250 rpm for 4 h 15 min. 3. Aliquot 100 μL formalin into labeled 1.5 mL microfuge tubes for sampling every 10 min from 0 to 120 min. 4. Mix a 50:50 mixture of ice and water in a container to accommodate the culture flask as you swirl its contents. 5. At 4 h 15 min transfer 900 μL of culture into tube 0 and mix with formalin by tube inversion. 6. Take the culture from the 36
C water bath shaker, insert thermometer into the culture and swirl the flask in the ice bath until temperature drops to 27
C, then return to the 25
C water bath shaker and rotate at 250 rpm (the culture will be 25
C within 1–2 min). 7. Start sampling regime, including removal of 900 μL for calcofluor staining (see Subheading 3.3) every 10 min.

3.2 cdc2-asM17 Induction Synchronization (See Note 4)

1. Place filtration apparatus and EMM2 (for both washing (50 mL) and growth after release from arrest (this volume is the same volume as the original culture volume)) in a 25
C incubator overnight (see Note 5). 2. Inoculate cdc2-asM17 from a fresh exponentially growing starter culture in EMM2 to generate a culture at 1  106 cells/mL after overnight growth in the 25
C water bath shaker at 250 rpm. 3. Add 3-BrB-PP1 to the culture to a final concentration of 2 μM and continue to rotate at 250 rpm for 4 h 15 min in 25
C. 4. To 1.5 mL microfuge tubes, labeled from 0 to 100 at intervals of 10, add 100 μL formalin, close and leave at room temperature. 5. Just before filtration, decant 40 mL from the flask of prewarmed release EMM2 medium (that was put in the incubator overnight in step 1) into a screw cap centrifuge tube and leave

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this tube alongside the rest of the release medium in the flask in a 25
C incubator. The medium in this tube will be used to resuspend cells from the filter in step 7. 6. At 4 h 15 min remove filtration assembly parts from 25
C incubator and assemble filtration unit on a polystyrene sheet (see Note 6). Connect the vacuum tubing/line to the Buchner flask and pour the culture into the top of the funnel at a rate of 120 mL/min. Do not allow the filter to dry out completely (see Notes 7 and 8), before immediately passing 50 mL of prewarmed EMM2 medium through the cells on the filter, disconnecting the vacuum and starting the timer. 7. Unclamp the top funnel from the glass frit membrane support and remove the glass fiber filter with forceps to drop it in a 40 mL aliquot from the prewarmed EMM2 medium (step 5), screw the lid on the tube and invert the tube 4–5 times to resuspend cells (see Note 9). 8. Pour the cell suspension (leaving the filter behind) from the tube into the remaining EMM2 medium in the conical flask that had been kept at 25
C overnight (steps 1 and 5). 9. Rotate the flask in the 25
C water bath shaker at 250 rpm (see Note 5) and commence sampling regime, including removal of 900 μL for calcofluor staining (see Subheading 3.3) every 10 min. 3.3 Calcofluor Staining

1. Add 900 μL of culture to 100 μL of formalin in a 1.5 mL microfuge tube and mix by inversion and place samples on ice until all samples have been collected at the end of the experiment. 2. Centrifuge at 16,000  g for 1 min, resuspend in PBS and repeat this wash twice more before resuspending pellet in 20 μL PBS at room temperature. 3. Centrifuge saturated calcofluor stock solution at 16,000  g at room temperature for 5 min. 4. Place 1–2 μL of cell suspension on the microscope slide next to 1–2 μL of 5 mg/mL calcofluor solution, mix with the pipette tip and then overlay with a coverslip. 5. Score the frequency of septa in 200 cells with an epifluorescence microscope.

3.4 Histone H1 Kinase Assay 3.4.1 Preparation of Magnetic p13Suc1 Beads

1. Transform E. coli Rosetta host cells with pGEX-4T-3-Spsuc1+ (encoding N-terminally GST-tagged S. pombe p13Suc1 (see Notes 10 and 11) and, the next morning, inoculate one colony into 1 L of LB broth + 100 μg/mL ampicillin and grow the 1 L culture in a 5 L flask with vigorous shaking at 37
C. When OD595 reaches 0.4 (usually around 5–6 h) add 1 mM IPTG at

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37
C. Two hours later harvest by centrifugation (3500  g, 20 min, 4
C), add 20 mL BLBA to the bacterial pellet and freeze in 20
C (see Note 12). 2. Next day, de-frost sample at 37
C for 30 min, add 4 mg lysozyme (see Note 13) and incubate at 37
C for 30 min, or until the solution becomes viscous (indicative of cell lysis). 3. Add 0.2 mg DNase I. Leave at room temperature for 30 min before centrifugation at 12,000  g for 15 min at 4
C (see Note 14). 4. Transfer the supernatant into a screw cap 50 mL centrifuge tube, dilute 50:50 with PBS and incubate for 45 min at room temperature on a rotating wheel with 2 mL of Glutathione Sepharose 4B beads (prewashed in PBS). 5. Pour the contents of the screw cap 50 mL centrifuge tube into an empty 20 mL polypropylene chromatography column (secured in a stand), allow the unbound fraction to flowthrough by gravity flow and wash the GST Sepharose beads three times with 5 mL ice-cold PBS by gravity flow. 6. Cap the column at the bottom, add 1 mL GST Sepharose elution buffer, cap the top of the column and incubate on a rotating wheel for 10 min at room temperature. Collect eluate by gravity flow and repeat elution four times more. 7. Pool the elution fractions together and dialyze against 1.5 L GST fusion protein storage buffer (in three successive rounds of 500 mL with stirring). 8. Dilute the purified p13Suc1-GST fusion protein in GST fusion protein storage buffer to a concentration of 1.8 mg/mL, snapfreeze in liquid nitrogen in 100 μL aliquots before storing at 80
C (see Note 15). 9. Desalt the purified GST-p13Suc1 in Zeba spin desalting columns to exchange the storage buffer for the coupling buffer to couple 5 mL of the purified p13Suc1-GST fusion protein (1.8 mg/mL) to 5 mL Pierce NHS-activated magnetic beads according to the manufacturer’s protocol. 3.4.2 Histone H1 Kinase Assay: Sample Collection (See Note 16)

1. Collect 10 mL of culture by centrifugation at 5500  g for 1 min at room temperature. 2. Resuspend the cell pellet in 1 mL stop buffer and pipet into a 1.5 mL screw cap microfuge tube. 3. Centrifuge the cell resuspension at 16,000  g for 30 s at room temperature and remove supernatant. 4. Snap-freeze cell pellet in liquid nitrogen and store at 80
C.

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1. Place cell pellets on ice and immediately add 500 μL of cold cell lysis buffer (CLB: see Note 17), 1 mL of cold glass beads and lyse cells with precooled Yasui Kikai Multi-bead shocker: 2500 rpm for 30 s (see Notes 18 and 19). 2. Pierce the bottom of the screw cap microfuge tubes with a hot 21 G needle (see Note 20). Place the perforated screw cap tubes into the tops of standard 1.5 mL microfuge tubes and place the “piggy backed” tube pair in a 50 mL screw cap centrifuge tube and spin at 400  g for 2 min at 4
C. The cell extract in the upper screw cap tubes will be forced into the standard 1.5 mL microfuge tubes. Discard the upper screw cap tubes. 3. Equilibrate the magnetic p13Suc1 beads with cell lysis buffer to give enough beads for 5 μL per time point. 4. Centrifuge the lower, retained, microfuge tubes at 16,000  g for 5 min at 4
C in order to clear the extract. Transfer the supernatant into prelubricated tubes containing equilibrated 5 μL chilled magnetic p13Suc1 beads. 5. Incubate for 1 h at 4
C in a rotating wheel at around 10 rpm to capture the Cdc2-Cyclin complexes. 6. Place tubes in magnetic rack on ice (see Note 21). Allow the solution to clear, before aspirating the liquid from the beads and removing tubes from the magnetic rack to resuspend the beads in 0.5 mL cell lysis buffer. Repeat this wash step with 0.5 mL cell lysis buffer once more. 7. Wash magnetic beads twice with 0.5 mL KAB and finally resuspend in 10 μL of kinase reaction (KAB + 1 μM Okadaic acid just before use), 2 μg Histone H1, and incubate at 30
C for 5 min (see Note 22). 8. Start the kinase reaction by adding 5 μL 0.3 mM ATP (Final concn 0.1 mM) and incubate at 30
C for 5 min. 9. Stop the reaction by adding 5 μL 4 LDS sample buffer with 10% β-mercaptoethanol and heating the samples at 70
C for 10 min.

3.4.4 Histone H1 Kinase Assay: Detection

1. Immediately after completion of the kinase assay, run the samples on 12% NuPAGE Bis-tris gel (running conditions: 1 MES buffer (with antioxidant) at 170 V for 35 min) (see Note 23). 2. Blot onto a 0.45 μm nitrocellulose membrane using a wet transfer apparatus in transfer buffer at 100 V for 1 h at 4
C. 3. Rinse the membrane with water, place in Ponceau S stain for 30 s, before washing the stained membrane with water once more. Capture an image of the membrane and then re-immerse in blocking buffer at room temperature.

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4. One hour later, replace blocking buffer with fresh blocking buffer containing the P-Thr-Pro-101 antibody (1:1000 in blocking buffer) at 4
C overnight, with rocking. 5. Wash blots with TBS-T three times (10 min each) before incubation with the secondary alkaline phosphatase conjugated anti-mouse antibody (1:1000 in blocking buffer) for 1 h at room temperature. 6. After three more 10 min washes in TBS-T, develop by rinsing and immersion in alkaline phosphatase buffer containing 0.33 mg/mL NBT and 0.165 mg/mL BCIP (see Note 24).

4

Notes 1. Some suppliers call Calcofluor, Calcofluor white, Blankofluor, or Fluorescent Brightner 28. 2. Temperature is more accurately controlled and maintained in a water bath shaker, rather than shaking in an air incubator, however if a shaking water bath is not available, an air shaker will suffice. 3. If setting up cdc25-22 cultures by using cell number rather than OD595 to gauge density, it is important to accommodate the increased cell size at 25
C, otherwise cell cycle and metabolic controls will differ from the mid-log phase cultures that are studied as a standard growth state in the field [37]. 4. The timings and cell densities described here are for growth at 25
C in EMM2, however, as shown in Fig. 2c cdc2-asM17 induction synchronization is equally effective in other media. Note that the arrest time must be changed to accommodate alterations in cell cycle timings in different growth contexts [37]. For example, to accommodate the faster growth in YES in the experiment for Fig. 2c the arrest was limited to 3 h instead of the 4 h 15 min required in EMM2. We routinely use 25
C for cdc2-asM17 induction synchronization for two reasons. Firstly, it minimizes temperature stress during filtration by operating close to room temperature. Secondly, transit of a synchronized mitosis is slower at 25
C than at 36
C [24] to provide greater resolution of individual steps. 5. Ensuring that the release medium volume matches the original culture volume and the same size flask is rotated at the same speed as the original culture is vital. If the volume or rate of aeration differs after the shift, there will be metabolic changes and consequential adjustments in growth rate, to induce changes that will be coincident with the changes that arise as a consequence of the mitotic synchronization [37].

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6. The polystyrene sheet (e.g., the lid of a “goods on dry ice” delivery box) insulates the culture from a cold shock stress from the cool bench surface. 7. Stress responses will be increased by drying on the filter. Disconnection of the vacuum line, rather than turning off the vacuum source is the most rapid and efficient way to ensure that the residual vacuum does not dry the filter. 8. At culture volumes greater than 500 mL, excessive accumulation of cells on the filter can slow filtration rates to impose stress, so it is important to frequently resuspend the cells during filtration to avoid stressing the first cells trapped by the filter. 9. Avoid vigorous agitation as this may release glass microfibers into the culture medium. 10. The E. coli BL21 host strain can be used as an alternative to Rosetta. 11. The pGEX-4T-3-Spsuc1+ plasmid has been deposited to Addgene (ID 162558). 12. BLBA may be prepared ahead and stored at room temperature, but the DTT and PMSF must be added fresh, immediately prior to use. 13. The lysozyme stock can be stored in 4
C for up to 1 month. 14. If a centrifuge capable of 12,000  g is not available a standard benchtop centrifuge can be used instead; the centrifugation period should then be extended to 30 min for speeds between 3200 and 3500  g. 15. Protein concentration can be measured by a variety of ways, including Nanodrop. 16. To gauge the background signal of a zero activity control for each experiment, we routinely take a duplicate sample from the at which we anticipate high levels of activity (e.g., a cdc25-22 20 min release point) and run the assay with and without the Cdk1-Cyclin B inhibitor Roscovitine (see Fig. 1b). 17. Add PMSF, DTT, protease inhibitor tablet, PNP, and Na-β-glycerol phosphate to the cell lysis buffer just before use. 18. After adding cell lysis buffer and glass beads to the cell pellet, proceed immediately and do not defrost the cell pellets. 19. Other forms of glass bead-based lysis such as manual agitation with a bench top vortex machine or the fast prep machine are also effective. However, in our hands, the Yasui Kikai Multibeads shocker gives the best lysis, while preserving the integrity of complexes in immunoprecipitation reactions and enzymatic activities, of any system we have tried. If using alternative systems, care should be taken to ensure that kinase activity is

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maintained upon lysis. For example, when we were developing Polo kinase activity assays we found that the activity seen when lysis was 70% efficient was lost upon 100% lysis [38]. 20. Heat 21 G needle in a flame until red hot to ease the piercing of the tube. 21. It is vital to use a strong magnetic rack to ensure rapid and efficient washing and isolation of the target p13Suc1-Cdc2Cyclin B complex. 22. Key steps in the kinase assay are the duration of the reaction, the number of yeast cells sampled and the quantity of the p13Suc1 beads. The conditions described here give a reproducible assessment of Cdc2 Histone H1 activity during cdc25-22 synchronous release. 23. Using precast, rather than self-made, gels makes it much easier to optimize phosphorylation specific antibodies. Because the protein resolution is better, the band for an individual protein is much thinner, such that the antigen is more concentrated. This tighter focus of the band enhances the differential between the signal to noise ratio of the specific signal from this band and background, nonspecific, signals. 24. We favor the “old fashioned” NBT BCIP colorimetric assay for developing blots, even though other forms of detection are effective. Extensive experience with many phosphorylation specific antibodies has taught us that the lower sensitivity of this system makes it easier to find a broad “sweet zone” in which the detection is phosphorylation specific. When using more sensitive detection systems, this window of utility can be so narrow that the reproducibility of the phosphorylation dependency can be challenged.

Acknowledgments Daniel Mulvihill (University of Kent, UK) and Janni Petersen (Flinders University, Australia) for critical reading of the manuscript and CRUK (A27336 and A24458) and Wellcome (200847/Z/16/Z) for funding. References 1. Mitchison JM, Nurse P (1985) Growth in cell length in the fission yeast Schizosaccharomyces pombe. J Cell Sci 75:357–376 2. Hayles J, Nurse P (2018) Introduction to fission yeast as a model system. Cold Spring Harb Protoc. https://doi.org/10.1101/pdb. top079749

3. Mitchison JM (1970) Physiological and cytological methods for Schizosaccharomyces pombe. Meth Cell Physiol 4:131–165 4. Hagan IM, Grallert A, Simanis V (2016) Cell cycle synchronization of Schizosaccharomyces pombe by lactose gradient centrifugation to

Synchronization and Assessment of S. pombe Mitotic Progression isolate small cells. Cold Spring Harb Protoc. https://doi.org/10.1101/pdb.prot091249 5. Hagan IM, Grallert A, Simanis V (2016) Cell cycle synchronization of Schizosaccharomyces pombe by centrifugal elutriation of small cells. doi: https://doi.org/10.1101/pdb. prot091231 6. Hagan IM, Grallert A, Simanis V (2016) Analysis of the Schizosaccharomyces pombe cell cycle. doi: https://doi.org/10.1101/pdb. top082800 7. Petersen J, Hagan IM (2003) S. pombe aurora kinase/survivin is required for chromosome condensation and the spindle checkpoint attachment response. Curr Biol 13:590–597 8. Nielsen O (2016) Synchronization of S phase in Schizosaccharomyces pombe cells by transient exposure to M-factor pheromone. Cold Spring Harb Protoc. https://doi.org/10.1101/pdb. prot091272 9. Nurse P (1990) Universal control mechanism regulating onset of M-phase. Nature 344:503–508 10. Ferrell JE (2008) Feedback regulation of opposing enzymes generates robust, all-ornone bistable responses. Curr Biol 18:R244– R245 11. Hagan IM, Grallert A (2013) Spatial control of mitotic commitment in fission yeast. Biochem Soc Trans 41:1766–1771 12. Nurse P, Thuriaux P, Nasmyth K (1976) Genetic control of the cell division cycle in the fission yeast Schizosaccharomyces pombe. Mol Gen Genet 146:167–178 13. Fantes PA (1977) Control of cell size and cycle time in Schizosaccharomyces pombe. J Cell Sci 24:51–67 14. King SM, Hyams JS (1982) Synchronisation of mitosis in a cell division cycle mutant of Schizosaccharomyces pombe released from temperature arrest. Can J Microbiol 28:261–264 15. Nurse P, Bissett Y (1981) Gene required in G1 for commitment to cell cycle and in G2 for control of mitosis in fission yeast. Nature 292:558–560 16. Hagan IM (1988) A study of the behaviour of microtubules and the mitotic spindle in the fission yeast Schizosaccharomyces pombe. PhD thesis, University of London, London 17. Hagan IM, Riddle PN, Hyams JS (1990) Intramitotic controls in the fission yeast Schizosaccharomyces pombe: the effect of cell size on spindle length and the timing of mitotic events. J Cell Biol 110:1617–1621 18. Booher RN, Alfa CE, Hyams JS et al (1989) The fission yeast cdc2/cdc13/suc1 protein kinase: regulation of catalytic activity and nuclear localization. Cell 58:485–497

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19. Hagan IM, Grallert A, Simanis V (2016) Synchronizing progression of Schizosaccharomyces pombe cells from G2 through repeated rounds of mitosis and S phase with cdc25–22 arrest release. doi: https://doi.org/10.1101/pdb. prot091264-6 20. Chen D, Toone WM, Mata J et al (2003) Global transcriptional responses of fission yeast to environmental stress. Mol Biol Cell 14:214–229 21. Nigg EA (1993) Cellular substrates of p34cdc2 and its companion cyclin-dependent kinases. Trends Cell Biol 3:296–301 22. Petersen J, Hagan IM (2005) Polo kinase links the stress pathway to cell cycle control and tip growth in fission yeast. Nature 435:507–512 23. Bishop AC, Ubersax JA, Petsch DT et al (2000) A chemical switch for inhibitorsensitive alleles of any protein kinase. Nature 407:395–401 24. Tay YD, Patel A, Kaemena DF et al (2013) Mutation of a conserved residue enhances the sensitivity of analogue-sensitised kinases to generate a novel approach to the study of mitosis in fission yeast. J Cell Sci 126:5052–5061 25. Swaffer MP, Jones AW, Flynn HR et al (2016) CDK substrate phosphorylation and ordering the cell cycle. Cell 167:1750–1761 26. Aoi Y, Kawashima SA, Simanis V et al (2014) Optimization of the analogue-sensitive Cdc2/ Cdk1 mutant by in vivo selection eliminates physiological limitations to its use in cell cycle analysis. Open Biol 4:140063–140063 27. Dischinger S, Krapp A, Xie L et al (2008) Chemical genetic analysis of the regulatory role of Cdc2p in the S. pombe septation initiation network. J Cell Sci 121:843–853 28. Stern B, Nurse P (1996) A quantitative model for the cdc2 control of S phase and mitosis in fission yeast. Trends Genet 12:345–350 29. Coudreuse D, Nurse P (2010) Driving the cell cycle with a minimal CDK control network. Nature 468:1074–1079 30. Nilsson J (2018) Protein phosphatases in the regulation of mitosis. J Cell Biol 218:395–409 31. Bouhlel IB, Ohta M, Mayeux A et al (2015) Cell cycle control of spindle pole body duplication and splitting by Sfi1 and Cdc31 in fission yeast. J Cell Sci 128:1481–1493 32. Novak B, Mitchison JM (1986) Change in the rate of CO2 production in synchronous cultures of the fission yeast Schizosaccharomyces pombe: a periodic cell cycle event that persists after the DNA-division cycle has been blocked. J Cell Sci 86:191–206 33. Novak B, Mitchison JM (1990) Changes in the rate of oxygen consumption in synchronous

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cultures of the fission yeast Schizosaccharomyces pombe. J Cell Sci 96:429–433 34. Simanis V (2015) Pombe’s thirteen – control of fission yeast cell division by the septation initiation network. J Cell Sci 128:1465–1474 35. Neumann FR, Nurse P (2007) Nuclear size control in fission yeast. J Cell Biol 179:593–600 36. Birot A, Eguienta K, Vazquez S et al (2017) A second Wpl1 anti-cohesion pathway requires dephosphorylation of fission yeast kleisin Rad21 by PP4. EMBO J 36:1364–1378 37. Petersen J, Russell P (2016) Growth and the environment of Schizosaccharomyces pombe. Cold Spring Harb Protoc. https://doi.org/ 10.1101/pdb.top079764 38. Tanaka K, Petersen J, MacIver F et al (2001) The role of Plo1 kinase in mitotic commitment and septation in Schizosaccharomyces pombe. EMBO J 20:1259–1270 39. Caspari T, Murray JM, Carr AM (2002) Cdc2cyclin B kinase activity links Crb2 and Rqh1topoisomerase III. Genes Dev 16:1195–1208 40. Grallert A, Patel A, Tallada VA et al (2013) Centrosomal MPF triggers the mitotic and

morphogenetic switches of fission yeast. Nat Cell Biol 15:88–95 41. Hagan IM (2016) Immunofluorescence microscopy of Schizosaccharomyces pombe using chemical fixation. https://doi.org/10. 1101/pdb.prot091017 42. Woods A, Sherwin T, Sasse R et al (1989) Definition of individual components within the cytoskeleton of Trypanosoma brucei by a library of monoclonal antibodies. J Cell Sci 93:491–500 43. Hagan IM, Hyams (1988) The use of cell division cycle mutants to investigate the control of microtubule distribution in the fission yeast Schizosaccharomyces pombe. J Cell Sci 89:343–357 44. Heitz MJ, Petersen J, Valovin S et al (1989) MTOC formation during mitotic exit in fission yeast. J Cell Sci 114:4521–4532 45. Funabiki H, Hagan IM, Uzawa S et al (1993) Cell cycle-dependent specific positioning and clustering of centromeres and telomeres in fission yeast. J Cell Biol 121:961–976

Chapter 11 Elucidating Human Mitosis Using an Anaphase-Like Cell-Free System Danit Wasserman, Sapir Nachum, Meirav Noach-Hirsh, Naomi Auerbach, Evelin Sheinberger-Chorni, Taylor P. Enrico, Roxane Lahmi, Michael J. Emanuele, and Amit Tzur Abstract A balanced progression through mitosis and cell division is largely dependent on orderly phosphorylation and ubiquitin-mediated proteolysis of regulatory and structural proteins. These series of events ultimately secure genome stability and time-invariant cellular properties during cell proliferation. Two of the core enzymes regulating mitotic milestones in all eukaryotes are cyclin dependent kinase 1 (CDK1) with its coactivator cyclin B, and the E3 ubiquitin ligase anaphase promoting complex/cyclosome (APC/C). Discovering mechanisms and substrates for these enzymes is vital to understanding how cells move through mitosis and segregate chromosomes with high fidelity. However, the study of these enzymes has significant challenges. Purely in vitro studies discount the contributions of yet to be described regulators and misses the physiological context of cellular environment. In vivo studies are complicated by the fact that each of these enzymes, as well as many of their regulators and downstream targets, are essential. Moreover, longterm in vivo manipulations can result in cascading, indirect effects that can distort data analysis and interpretation. Many of these challenges can be circumvented using cell-free systems, which have historically played a critical role in identifying these enzymes and their contributions under quasicellular environments. Here, we describe the preparation of a newly developed human cell-free system that recapitulates an anaphase-like state of human cells. This new toolkit complements traditional cell-free systems from human cells and frog eggs and can be easily implemented in cell biology labs for direct and quantitative studies of mitotic signaling regulated by phosphorylation, APC/C-mediated proteolysis, and beyond. Key words Cell-free system, Cell extracts, Ubiquitin-mediated degradation, Nondegradable cyclin B, Mitosis, Anaphase, APC/C, Cdc20, Cdh1, Cdk1

1

Introduction The unidirectional nature of the cell cycle is first and foremost achieved by a highly regulated series of protein modifications— most notably phosphorylation, and ubiquitination-mediated proteolysis [1]. Orderly (de)phosphorylation and proteolysis is perhaps best demonstrated during mitosis and cell division. Much of the

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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protein degradation controlling mitotic progression and exit is regulated by an E3 ubiquitin ligase, the anaphase-promoting complex/cyclosome (APC/C), and its two substrate adaptors—Cdc20 and Cdh1. Cdc20-bound APC/C (APC/CCdc20) is active from mitotic metaphase until late anaphase, mediating the proteolysis of mitotic cyclins, Securin, and other key cell cycle proteins [2– 5]. Cdh1-bound APC/C (APC/CCdh1) is active from telophase through G1-phase of the cell cycle. The switch from APC/CCdc20 to APC/CCdh1 is regulated by temporal (de)phosphorylation and proteolysis, and is a central event in the overall mechanism underlying orderly cell division in all eukaryotes [6–8]. Cell-free systems are known for their capacity to recapitulate complex cellular processes in vitro while maintaining a physiologically relevant context. These systems are optimal for direct and quantitative assays, circumventing caveats associated with time-sensitive assays and long-term in vivo manipulations. Cellfree systems can either capture a certain physiological state (e.g., interphase) or reproduce biochemical and structural dynamics characterizing transitions between cell cycle phases (e.g., phosphorylation wave, chromatin condensation, and spindle formation during mitotic entry). Much of the core machinery of the vertebrate cell cycle was originally resolved in cell-free systems derived from frog eggs and early embryos. The unmatched simplicity of obtaining near physiological protein concentration in vitro and the synchronous two-phase cell cycles during early embryogenesis provide a homogenous biochemical ‘soup’ for elucidating the cell cycle machinery [9–12]. Mitotic entry and exit, metaphase-to-anaphase transition and cytokinesis were all demonstrated in frog egg extracts [13– 15]. Moreover, the APC/C and some of its key targets were both discovered in egg extracts, including Cyclin B [3], Securin [4] and Geminin [16]. With respect to Cdc20 vs. Cdh1 specificity of APC/ C targets, egg extracts are optimal. Vertebrate Cdh1 is undetectable pre–mid-blastula transition, and thus, APC/C activity in mitotic egg extracts is mediated solely by Cdc20, whereas interphase egg extracts supplemented with recombinant Cdh1 recapitulate APC/ CCdh1-specific activity. Systematic screens for APC/C targets in both mitotic and interphase egg extracts have shed enormous light on vertebrate cell cycle [4, 16–19]. Currently, frogs are no longer considered a popular animal model and thus, frog colonies with sufficient size are scarce. Moreover, the high cost of frog facilities is a limiting factor for many labs. In the last 20 years, human cell-free systems have been gradually integrated into cell cycle research. Extracts from synchronous cell populations provide quasi-cellular environments for analyzing orderly protein degradation, phosphorylation and other signaling events in a somatic 4-stage cell cycle context, which is lacking in egg extracts [20–23]. These environments can be manipulated either

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genetically in the source cells prior to extract preparation, or biochemically before, during or after extract preparation. Extracts from human cells arrested at the G1 phase exhibit APC/CCdh1mediated proteolysis, much like interphase egg extracts. Human mitotic cell extracts are typically obtained from nocodazole arrested cells. These extracts are not equivalent to mitotic egg extracts because the activated mitotic checkpoint complex halts APC/ CCdc20 activity. Removal of this checkpoint apparatus can be induced thermodynamically and/or biochemically, but then, mitotic exit is triggered and both APC/CCdc20 and Cdk1 are gradually inactivated [24]. We recently developed a human cell-free system which recapitulates an early anaphase-like state where both Cdk1 and APC/ CCdc20 remain stably active [24]. The system is named ‘NDB’ and is based on HEK293 cells inducibly expressing a nondegradable mutant variant of Cyclin B1 (NDB). In this method chapter we describe the preparation of NDB extracts and source cells. The potential use of the system in studying mitotic phosphorylation and degradation and the Cdc20 vs. Cdh1 specificity of APC/C substrates is demonstrated.

2

Materials

2.1 Buffers and Reagents

1. Unless otherwise is noted, all solutions made with double deionized water (DDW); 18 MΩ.

2.1.1 Cell Culture Maintenance

2. Culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS), 2 mM L-glutamine, and 1% of X100 penicillin–streptomycin solution (10,000 U/mL penicillin, 10,000 μg/mL streptomycin). 3. Phosphate buffered saline (PBS), pH 7.4. 4. Trypsin–EDTA Solution: 0.25% Trypsin, 0.05% EDTA. 5. Antibiotics for cell selection: zeocin™ stock solution (100 mg/ mL); blasticidin stock solution (10 mg/mL).

2.1.2 Generation of NDB Cell System

NDB cell system is based on human 293T-REx™ expressing a nondegradable mutant variant of human Cyclin B1 under Tetracycline-regulated CMV promoter. 1. Mammalian cell line: 293T-REx™ (Thermo Fisher Scientific; #R71007). 2. Mammalian expression vector pcDNA™4/TO (Thermo Fisher Scientific, #V102020). 3. A primer set for cloning human Cyclin B1 from a source vector or genomic cDNA into pcDNA™4/TO vector:

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Forward— GACTGGATCCATGGCGCTCCGAGTCAC CAG. Reverse complement—GACTCTCGAGTTACAC CTTT GCCACAGCCTT. 4. Restriction enzymes BamHI and XhoI. 5. High-fidelity DNA polymerase (e.g., Agilent’s Herculase II; #600675). 6. T4 DNA Ligase (New England Biolabs; #M0202S). 7. QuikChange Lightning™ site directed mutagenesis kit (Agilent, #210519). 8. A primer set for generating nondegradable mutant Cyclin B1 using pcDNA4/TO-Cyclin B1 as a template (see Note 1). Forward— CCAATGTCCCCAA CAGCTGTTCCTGGCCTCAGTCCG. Reverse complement— CGGACTGAGGCCAGGAA CAGCTGTTGGGGACATTGG. 9. A plasmid DNA miniprep kit (e.g., Qiagen; #27104). 10. Competent bacteria DH5α™ (Thermo Fisher Scientific; #18263012). 11. Agarose–TAE powder blends (1%) for Gel electrophoresis (Sigma-Aldrich; #A6236-100G). Follow the manufacturer’s protocol for gel preparation. 12. LB Agar carbenicillin plates: 100 μg/mL carbenicillin. 13. 2
HeBS solution: 50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 1.5 mM Na2HPO4, 280 mM NaCl. Adjust pH to 7.05 with NaOH. 14. 2.5 M CaCl2 stock solution. Dissolve 11 g of CaCl2∙6H2O in a final volume of 20 mL of DDW. Sterilize the solution by passing it through a 0.22-μm filter. Store at 4  C. 2.1.3 Validating Mitotic Arrest in NDB Cells Expressing Nondegradable Cyclin B1

1. Tetracycline (Tet) stock solution: 2.5 mg/mL Tet in 95% ethanol absolute and 5% DDW. 2. PBS. 3. Ethanol absolute. Store at 20  C. 4. Propidium iodide (PI) staining solution: 0.02 mg/mL PI, 0.05 mg/mL RNAse A in PBS. 5. Anti-human Cyclin B1 antibody (Cell Signaling Technology; #4138). 6. Standard reagents for SDS-PAGE and Western blotting (see Subheading 2.1.8, items 6–11). 7. Nocodazole stock solution: 1 mM in DMSO. Store at 20  C. 8. Prewarmed hypotonic solution (37  C): 5.6 g KCl in 1000 mL DDW (0.075 M). 9. Fresh fixative solution: 3:1 (v/v) methanol–glacial acetic acid.

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10. Mounting solution: Mounting medium (Richard-Allan Scientific, #4112APG) supplemented with 5 μg/mL 40 ,6-diamidino-2-phenylindole (DAPI) stain. 11. Clear nail polish. 2.1.4 Preparation of NDB Mitotic Protein Lysate

1. Tet stock solution. 2. PBS. 3. 0.5 M Ethylenediaminetetraacetic acid (EDTA) stock solution: Dissolve 93.05 g EDTA disodium in 300 mL DDW. Adjust pH to 8.0 with NaOH. Adjust volume to 500 mL with DDW. Store at room temperature (RT). 4. Cell lysis solution: 50 mM Tris–HCl (pH 7.6), 150 mM NaCl, 5 mM EDTA (pH 8.0 with NaOH), 0.5% NP-40, supplemented with a protease inhibitor cocktail (Roche; #4693159001), phosphatase inhibitor cocktails (Sigma-Aldrich; #P5726 and #P0044), 1 mM phenylmethylsulfonylfluoride (PMSF), 10 mM NaF, 20 mM β-glycerophosphate, 1 mM Na3VO4, 20 mM P-nitrophenylphosphate. 5. Bradford reagent (Bio-Rad #500-0006). 6. Bovine serum albumin (BSA).

2.1.5 Preparation of NDB Mitotic Extracts

1. Tet stock solution. 2. PBS. 3. Swelling Buffer: 20 mM HEPES (pH 7.5), 2 mM MgCl2, 5 mM KCl, 1 mM dithiothreitol [DTT], and protease inhibitor cocktail (Roche; #4693159001). 4. E-mix (Energy regeneration mixture): 20 mM ATP, 2 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N0 ,N0 -tetraacetic acid (EGTA), 20 mM MgCl2, 150 mM creatine phosphate, 1 mg/ mL creatine phosphokinase. Store 50 μL aliquots in 80  C. 5. Liquid N2.

2.1.6 Degradation and Mobility-Shift Assays in NDB Mitotic Extracts

1. NDB mitotic extract. 2. SP6 expression vector carrying an ORF of interest. 3. TNT® SP6 coupled reticulocyte lysate system (Promega; #L2080). 4.

35

S-methionine/35S-L-cysteine #NEG772002MC).

mix

(PerkinElmer;

5. Recombinant Ubiquitin (Boston Biochem; #U-100H). 6. E. coli-derived recombinant UbcH10 and dominant negative UbcH10 (UbcH10DN). Commercial version of these reagents are available (Boston Biochem; #E2-650, #E2-654, #E2-652). 7. MG-132 proteasome inhibitor stock solution: 5 mg/mL in DMSO.

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8. 2
Laemmli sample buffer (Bio-Rad Laboratories; #161-0737). 9. Dithiothreitol (DTT). 1 M stock solution in DDW. Store at -20  C. 10. Polyacrylamide gels (e.g., Mini-Protean TGX precast gels [Bio-Rad Laboratories]). Recommended polyacrylamide concentrations: 7.5%, 10%, and 4–15% (#456-1-23, #456-1033, #456-1083, respectively). 11. Destain solution: 10% methanol, 7.5% acetic acid, in DDW. 2.1.7 G1-Like NDB Extracts

1. NDB mitotic extracts. 2. Cdk1 inhibitor stock solution: 10 mM RO-3306 (Cdk1 inhibitor) in DMSO. Store 20  C. 3. DMSO (dimethyl sulfoxide).

2.1.8 Immunoprecipitation of APC/C from NDB Extracts and Immunoblot of Cdc20/ Cdh1

1. Wash buffer I: 150 mM NaCl, 20 mM Tris–HCI (pH 7.5), 10% glycerol, 0.1% Triton, 1 mM EDTA. 2. Wash buffer II: 75 mM NaCl, 20 mM Tris-HCI (pH 7.5), 10% glycerol, 0.1% Triton, 1 mM EDTA. 3. Agarose-conjugated monoclonal anti-Cdc27 antibody: clone AF3.1 (Santa Cruz Biotechnology; SC-9972AC). 4. Anti-Cdc20 antibody (Santa Cruz Biotechnology; #SC-8358). 5. Anti-Cdh1 antibody [Clone DH01]).

(MilliporeSigma™;

#CC43100UG

6. Secondary antibodies conjugated to horseradish peroxidase (HRP) from Jackson ImmunoResearch: goat anti-rabbit IgG (#111-035-144), goat anti-Mouse IgG (#115-035-003), mouse anti-rabbit IgG, light chain specific (#211-032-171), goat anti-mouse IgG light chain specific (#115-035-174). 7. TBST buffer: 150 mM NaCl, 50 mM Tris–HCl, 0.1% Tween 20, in DDW. pH adjusted to 7.6 with HCl. 8. Transfer buffer: 25 mM Tris, 192 mM glycine, 20% methanol. 9. Running buffer: 25 mM Tris, 192 mM glycine, 0.1% sodium dodecyl sulfate (SDS). 10. Antibody solution: 5% BSA, 0.05% sodium azide in TBST (see item 7 above). 11. Reagents for Enhanced Chemiluminescence (ECL) Western detection (e.g., Thermo Fisher Scientific; #34095). 2.2

Equipment

2.2.1 General Use

1. Pipettes for 2, 10, 200, and 1000 μL, and matching pipette tips. 2. Pipette-aid and serological pipettes: 5, 10, and 25 mL. 3. Tubes: 0.2, 1.5, 2, 15, and 50 mL.

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4. Refrigerated centrifuges and adaptors for 1.5 and 2 mL tubes, and 15, 50, and 500 mL conical tubes. 5. PCR machine. 6. Temperature controlled water bath. 7. Thermomixer or dry block incubator for 1.5 mL tubes. 8. Spectrophotometer for DNA quantification and Bradford protein assay. 9. 4  C environment. 10. Freezers: 20 and 80  C. 11. Apparatuses and power supplies for preparing and running agarose and acrylamide gels. 12. Gel loading tips, 1–200 μL (e.g., Fisherbrand™ H01096RSFIS). 13. Autoclave. 2.2.2 Cell Culture Maintenance

1. 37  C, 5% CO2 controlled incubator. 2. Laminar flow hood. 3. Tissue culture plates: 60, 100, and 150 mm/diameter. 4. Multiwell tissue culture dishes: 24- and 96-well dishes.

2.2.3 Microscopy

1. Inverted phase contrast microscope equipped with 20
and 40
objectives. 2. Epifluorescence microscope equipped with a light source and a filter set for imaging DAPI-stained chromosomes, and
40-
100 oil objectives. 3. Glass slides (1 mm thick, 25 mm
75 mm). 4. Coverslips (No. 1 thickness, 22 mm
22 mm).

2.2.4 Flow Cytometry

1. Optical flow cytometer with a 488 nm laser and filter/detector for Phycoerythrin (PE). 2. Falcon® round-bottom polypropylene 5 mL tubes with cell strainer cap.

2.2.5 Extract Preparation

1. Liquid N2 dewar. 2. 250 or 500 mL conical tubes. 3. 10 mL beaker. 4. 21 G needle. 5. 3 or 5 mL syringe. 6. Cooled centrifuge for 15–500 mL conical tubes, and a swing bucket rotor. 7. Cooled centrifuge for 1.5 to 2 mL tubes.

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2.2.6 Degradation/ Phosphorylation Assays

1. Whatman™ paper (46 cm
57 cm; Whatman grade: 3MM). 2. Gel drying machine (e.g., Bio-Rad, model #583). 3. Phosphoimager. 4. Phosphor screen and cassette. 5. A phosphorscreen eraser. 6. Image analysis software for signal quantification (e.g., ImageJ).

2.2.7 Immunoblot and Immunoprecipitation

1. Rocking shaker. 2. Tube rotator. 3. Western blot imaging system.

3

Methods

3.1 Generation of NDB Cell Line

1. Culture 293T-REx™ cells in a medium containing 5 μg/mL Blasticidin (see Notes 2–4). 2. Amplify human Cyclin B1 ORF using (a) a source plasmid or genomic cDNA as a template; (b) a primer set (see Subheading 2.1.2); (c) restriction enzymes BamHI and XhoI; and (d) a PCR reaction with a high-fidelity DNA polymerase (see Subheading 2.1.2, items 4 and 5). 3. Clone human Cyclin B1 ORF into pcDNA™4/TO expression vector using a standard DNA ligation protocol. 4. Generate a nondegradable mutant variant of Cyclin B1 by introducing two point mutations for substituting Arg 42 and Leu 45 at the destruction box of Cyclin B1 with Gly and Val, respectively [5, 25]. QuikChange Lightning™ site-directed mutagenesis kit is optimal for this application. Use the pcDNA™4/TO-Cyclin B1 plasmid as a template. A primer set for mutagenesis is indicated in Subheading 2.1.2. 5. Digest the PCR product with DpnI restriction enzyme (DpnI is supplied with the site-directed mutagenesis kit) and transform into DH5α E. coli. Select transformants bacteria on agar plates containing 100 μg/mL Carbenicillin. 6. Select a transformed colony for plasmid preparation using a standard miniprep protocol. 7. Validate mutagenesis by sequencing the miniprep product. 8. Transfect 293T-REx™ cells with pcDNA™4/TO-Cyclin B1-DM plasmid using a standard transfection method (see Note 5). 9. Forty-eight hours post transfection split and reculture cells at 25% confluency. Start cell selection by adding 200 μg/mL Zeocin™ in addition to 5 μg/mL blasticidin.

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10. Change selection medium every 3–4 days until single colonies are formed. 11. Pick at least 24 colonies and expand them separately (see Note 6). 12. Culture colonies in 24-well dishes. 13. Once cells reach 50% confluency, treat cells with 1 μg/mL Tet for 18–24 h (see Note 7). 14. Assess synchronization quality by phase contrast microscopy. Cells in which the ORF encoding nondegradable cyclin B1 was integrated successfully into the genome and expression is properly induced arrest in mitosis, become round, form aggregates and partly detach from the surface (Fig. 1). From this point onward, maintain about five positive colonies for further validation. In addition, cryofreeze two vials for each colony. 15. In addition, assess synchronization quality by DNA quantification before and after 22 h treatment with 1 μg/mL Tet (see Subheading 3.2). Maintain colonies in which G2/M index following mitotic arrest reaches ~95% (Fig. 1). 16. Validate expression of nondegradable Cyclin B1 by Western blotting with anti-Cyclin B1 before and 22 h after induction by Tet (see Note 8). 17. Validate sister chromatid separation in mitotic-arrested NDB cells. Tet-induced NDB cells arrest in mitosis post metaphaseto-anaphase transition. At this anaphase-like stage, full or partial sister chromatid separation has already taken place [24]. A standard protocol for chromosome spreads can be used to validate this feature of the arrested cells (see Subheading 3.3 and refs. 24, 26, 27). As a control, nocodazole-arrested NDB cells should be analyzed in parallel for visualizing unseparated sister chromatids for comparison. To this end, treat NDB cells with 100 nM nocodazole for ~16 h to enrich for prometaphase cells. Chromosomes can be visualized by DAPI staining on glass slides using an epifluorescence microscope equipped with a 405 nm light source and 40–100
oil-immersion objectives. NDB cell colonies exhibiting a high fraction of Tet-induced cells with separated sister chromatids should be prioritized for further experiments. 18. For all further experiments, select a cell colony whose proliferating time, DNA distribution in steady state culture conditions, overall shape and size range are most similar to the source cell line (293T-REx™) [24]. 3.2 PI Staining Protocol for Quantifying DNA Content by Flow Cytometry

1. Harvest about one million cells and transfer to a 10 mL conical tube. 2. Centrifuge cells for 5 min (220
g, 4  C), discard media. 3. Resuspend cells with 10 mL cold PBS. Centrifuge cells for 5 min (220
g, 4  C), discard supernatant.

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Fig. 1 NDB cell- and cell-free systems for elucidating human mitosis. (a) NDB cell system is based on human 293-T-REx™ cells stably expressing nondegradable (ND) mutant of Cyclin B1 under a tetracycline (Tet)/ doxycycline (Dox)-regulated CMV promoter. Expression of ND-Cyclin B1 induces mitotic arrest in an anaphase-like state. The constant activity of the Cdk1–Cyclin B1 complex prevents mitotic exit and maintains a peak level of APC/CCdc20 activity for hours. NDB extracts are typically prepared from 20 plates of 150 mm/ diameter. Recommended cell confluency for Tet-induced expression is 75–80%. Following 20–22 h treatment with Tet, cells acquire a spherical shape. This archetypal morphology indicates mitotic arrest. Cell synchronization should be further validated by standard DNA quantification. The arrested cells will be loosely attached to the surface, if at all. Thus, cells can be collected directly by gentle pipetting (without trypsinization) into a single 500 mL conical tube. Once cells are pelleted and washed, they are ready for extract preparation. In vitro translated (IVT) protein products are generated in reticulocyte lysate supplemented with radiolabeled methionine (35S-Met). pCS2-based expression vectors and reticulocyte lysates supporting transcription from SP6 promoter are recommended. A typical reaction mix contain 10–20 μL NDB mitotic extracts and 0.5–1.5 μL IVT

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4. Resuspend cells with 250 μL cold PBS. 5. Fix cells by adding 750 μL absolute Ethanol (20  C), dropby-drop, while agitating tube with your finger. Store in 20  C for 2 h or longer. 6. For PI staining, centrifuge fixed cells for 5 min (220
g, 4  C), discard media. 7. Resuspend cells with 1 mL PI staining solution, mix and incubate for 30 min at RT in the dark. 8. Transfer cells through a cell strainer cap into a round-bottom polypropylene 5 mL tubes. 9. Analyze cells using a flow cytometer equipped with a 488 nm laser. 3.3 Chromosome Spreads

As a control, nocodazole-arrested NDB cells should be analyzed in parallel for visualizing unseparated sister chromatids for comparison. To this end, treat NDB cells with 100 nM nocodazole for ~16 h to enrich for prometaphase cells. NDB cell colonies exhibiting a high fraction of Tet-induced cells with separated sister chromatids should be prioritized for further experiments. 1. Harvest Tet-induced and nocodazole-arrested NDB cells from a 10 cm/diameter dish by gentle pipetting. 2. Transfer cell to a 15 mL conical tube. Centrifuge cells for 5 min (220
g, RT). Discard media. 3. Resuspend cells with 10 mL PBS, centrifuge cells for 5 min (220
g, RT). 4. Aspirate PBS, leaving 0.5 mL of buffer in the tube. Gently resuspend the pellet by flicking the tube with your fingers. 5. Add 2 mL of prewarmed (37  C) hypotonic solution (0.56% KCl), drop-by-drop, while agitating tube.

ä Fig. 1 (continued) product. Extracts already contain energy-regeneration mix (E-mix), as well as endogenous Ubiquitin (Ub) and the E2 enzyme UbcH10. As such, extracts are highly active. Yet adding 0.5–1 μL of
20 E. mix, recombinant Ub, and UncH10 into the reaction mix can facilitate proteolysis mediated by APC/CCdc20. Conversely, proteasome inhibitors (e.g., MG132), dominant negative UbcH10 (UbcH10DN), or APC/C specific inhibitors (e.g., Emi1 or TAME) can be added for validating the potency and specificity of the assay. Assays are typically performed in a temperature range of 23–30  C. During degradation/mobility-shift assays, 3–5 μL of the reaction mix is sampled each time point, mixed with Laemmli buffer, boiled and frozen at 80  C. Protein sampled are resolved by SDS-PAGE. Gels are then soaked in a distain solution, heat/vacuum dried, and exposed to a Phosphorimager screen for 1 h to 1 day. Longer exposure might be helpful in case of weak signals. The potency and specificity of NDB mitotic extracts is demonstrated; Geminin, an APC/CCdc20 substrate, but not Tome-1, an APC/CCdh1 substrate, is degraded. This degradation is blocked by Emi1. Orderly phosphorylation of Tome-1 is evident in minute-scale resolution by a gradual mobility shift. The high mobility shift of Tome-1 observed after 140 min indicates for the stability and potency of this cell-free system

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6. Add additional 10 mL hypotonic solution and incubate for 10 min in 37  C. Centrifuge for 5 min (220
g, RT) and aspirate supernatant, leaving 0.5 mL solution. Gently resuspend pellet. 7. Prepare fresh fixative solution (3:1 v/v methanol–glacial acetic acid). 8. Add 5 mL fixative solution (3:1 v/v methanol–glacial acetic acid) from the side of the tube. Mix by gently inverting the tube. Centrifuge cells for 5 min (220
g, RT) and discard supernatant. Repeat step 8. 9. Add 500 μL fixative solution, mix well by flicking tube. Let large clumps settle. 10. Immerse precleaned slides in fixative solution for 10 min. Air-dry the slide. 11. Drop cell droplets on tilted glass slides from 1-m height. 12. Air-dry slides, mount with a mounting/DAPI solution, place a coverslip, and seal with clear nail polish. 13. Image chromosomes using an epifluorescence microscope equipped with a 405 nm light source, a filter set for imaging DAPI, and
40–
100 oil objectives. 3.4 Harvesting Mitotic NDB Cells for Whole-Cell Protein Lysate Preparation and Immunoblot Assays

1. Split NDB cells into a desired number of plates. One 10 cm plate is sufficient for ~1 mg total protein. 2. The following day, when cells reach ~75% confluency, add 1 μg/μL Tet for 22 h. 3. Tet-induced NDB cells will be fully or partially detached from the surface. Collect medium with a 10 mL serological pipette. Pipet up and down gently across the plate in a serpentine manner to facilitate cell detachment. 4. Transfer cells to a 15 mL conical tube. Rewash the plate with 5 mL fresh media and collect remaining cells. 5. Centrifuge cells for 5 min (220
g, 4 supernatant.



C). Remove

6. Resuspend cell pellet gently with 10 mL cold PBS and centrifuge again. Remove supernatant. 7. Resuspend cell pellet gently with 1 mL cold PBS, transfer cells to a 1.5 mL tube and centrifuge for 5 min (220
g, 4  C). Remove supernatant. 8. Resuspend pellet with 150–200 μL cell lysis solution and incubate 30 min on ice. 9. Centrifuge for 40 min (20,000
g, 4  C) to remove cell debris. Collect protein lysate (supernatant). 10. Transfer protein lysate into a new precooled tube.

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11. Determine protein concentration by a standard Bradford assay. 12. Store protein lysate in 80  C. For immunoblot applications, storing protein lysate in aliquots of 10–20 μL is recommended. 3.5 Preparation of NDB Mitotic Extracts

1. Split NDB cells into 21 plates of 150 mm/diameter (by trypsinization). We recommend preparing extracts from 20 plates. Expected extract yield is 2–3 mL. Keep at least one extra dish for backup or maintenance. We typically split cells from seven plates of ~90% cell confluency in a 1:3 ratio 24 h before adding Tet (see Note 9). The following day, at ~75% cell confluency, add freshly prepared 1 μg/mL Tet for 22 h. 2. Assess synchronization quality by phase microscopy. Nearly all cells should be rounded, in clusters, and either fully or partially detached from the surface (Fig. 1). 3. Collect cells from each plate by pipetting the medium up-and down 2–3 times across the plate in a serpentine manner using 25–50 mL serological pipettes. Transfer all media into a single 500 mL conical tube (see Note 10). 4. Rewash all plates with 25 mL medium by transferring medium from plate to plate. Collect remaining cells. 5. Place the 500 mL tube in a swing bucket rotor with a matching adaptor, balance weight, and centrifuge 5 min (220
g, 4  C). 6. Discard supernatant by pouring the medium gently into a chilled beaker in case the cell pellet is inadvertently detached. 7. Resuspend cell pellet with 40 mL ice-cold PBS and transfer into 50 mL conical tube. Centrifuge for 5 min (220
g, 4  C). Discard supernatant. Repeat step 7. 8. Resuspend cell pellet with 2–3 mL ice-cold PBS and transfer into precooled 2 mL tubes. Tube number should be minimized. Centrifuge for 5 min (220
g, 4  C). Discard supernatant. Pellet volume should be approximately 1 mL per tube. 9. For each 1 mL cell pellet, resuspend with 750 μL swelling buffer and 50 μL E-mix. Incubate 30 min on ice. Every 5 min mix by inverting the tube three times. 10. To break cells, flash-freeze the tubes in liquid N2 and thaw quickly in 30  C water bath. Minimize thawing time. Repeat step 10. 11. Combine the contents of all tubes into a 10 mL precooled beaker. To facilitate lysis, pass liquid ten times through a 21 G needle using a precooled 3–5 mL syringe. 12. Divide liquid into precooled 2 mL tubes and centrifuge for 10 min at 20,000
g (4  C).

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13. Collect supernatant from all tubes with a 200 μL pipette and transfer to new precooled 2 mL tubes. Fill tubes all the way up. Centrifuge for 40 min at 20,000
g (4  C). Expect to see (a) cell pellet at the bottom of the tube; (b) a clear phase of cell extracts; and (c) a thin layer of floating lipids. 14. Collect supernatant gently with a 200 μL pipette. Place the tip well above the pellet phase. Aspirate protein extracts slowly and with minimal mixing. Avoid taking up pellet or lipids floating at the top. Typically, 1–1.2 mL extracts can be safely collected per full 2 mL tube. 15. Combine cell extracts in new precooled tubes. Set aside a few μL of the extracts to measure protein concentration by a Bradford assay. Extract concentration typically ranges from 18 to 25 mg/mL (see Note 11). 16. Aliquot extracts in 0.2 mL tubes. Strips of thin-wall PRC tubes are recommended. Aliquots of 45–65 μL are convenient. 17. Snap-freeze tubes in liquid N2, and store at 80  C (see Notes 12 and 13). 3.6 Degradation and Mobility Shift Assays in NDB Extracts

1. Express protein of interest in reticulocyte lysate following manufacturer’s protocol. We recommend having the desired ORF in a pCS2-based vector and using an SP6 premixed version of Promega’s TNT-coupled transcription–translation reaction kit (see Note 14). Add 35S-Met to reaction mix to generate a radiolabeled in vitro translated (IVT) protein product (Fig. 1; see Note 15). Degradation and mobility shift assays are typically performed with 0.5–1.5 μL IVT product. Based on our experience, IVT products can be refrozen five times without noticeable damage. 2. Prepare reaction mix on ice (Fig. 1). A default reaction mix for degradation assay contains (a) 20 μL NBD extract; (b) 1 μL radiolabeled IVT product; (c) 10 μg ubiquitin; and (d) 1 μL E-mix (see Note 16). 3. Reaction mix can contain additional reagents that either facilitate or block APC/CCdc20-mediated proteolysis (Fig. 1). These supplements are critical controls for testing the potency and specificity of the assay. For example, APC/CCdc20-mediated proteolysis is facilitated by recombinant UbcH10 but blocked by UbcH10DN, Emi1 (APC/C inhibitor), Securin (competitive inhibitor), or TAME (small molecule drug) [28]. Control experiments with proteasome inhibitors (e.g., 20 μM MG-132) are recommended. 4. For phosphorylation-mediated mobility shift assays, ubiquitin is not required. Freshly added E-mix is also dispensable in assays shorter than 1 h. Instead, control assays can contain the Cdk1 inhibitor RO-3306 (15 μM) to couple mobility

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shift of a tested protein to Cdk1-Cyclin B1-dependent phosphorylation [22–24]. Small molecule inhibitors of other mitotic kinases (e.g., Plk1) can also be used. 5. Mix all reagents in a 0.2 mL PCR tube on ice or in a precooled metal rack on ice. Add IVT product last. 6. Homogenize reaction mix by pipetting up and down 4–5 times with a 10 μL pipette. Avoid foaming. 7. Prepare 1.5 mL tubes with 10 μL 2
Laemmli sample buffer to stop the reaction; one tube per time point. The volume allocated for each time point may vary according to the desired number of samples and the overall volume of the reaction mix (see Note 17). 8. While reaction mix is on ice, take the first sample and mix with 2
Laemmli buffer. Snap-freeze in liquid N2. This sample represents time point 0. 9. Place the reaction mix tube in a prewarmed PCR machine. Recommended temperature range for the assay is 23–30  C (see Fig. 2 and Notes 18 and 19). 10. For each time point, transfer 3–5 μL reaction mix into a tube containing 10 μL 2
Laemmli buffer and snap-freeze in liquid N2 (see Note 17). 11. Store all samples in 80  C.

Fig. 2 Characterization of APC/CCdc20-mediated degradation in NDB mitotic extracts. (a) Time-dependent degradation of Securin (35S-labeled, IVT product) in mitotic NDB extracts was assayed in three different temperatures. Extracts were supplemented with E-mix and Ub (0.5 mg/mL). The impact of adding recombinant UbcH10 or UbcH10DN (0.5 mg/mL) on the degradation of Securin also tested at 28  C. Degradation was assayed by SDS–PAGE and autoradiography. Quantification of the depicted raw data (left) are plotted on the right. Data are normalized to max signal at t ¼ 0. (b) Time-dependent degradation of Securin in diluted NDB mitotic extracts are shown (28  C). See (a) for details. NDB extracts were diluted two- or fourfold in swelling buffer

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12. Denature proteins in 95  C for 10 min just before gel loading. Quick-spin samples and load the entire sample on a gel. 13. Resolve sample by SDS-PAGE. We typically use fresh gels of 1 mm thick and 8% or 10% acrylamide and a Tris-Glycine running buffer. 14. Soak gel in destain solution for 20 min. 15. Lay gels on a wet Whatman paper. Add a piece of dry Whatman paper underneath and a piece of saran wrap on top (the gel should remain unwrapped). Dry the gel using a vacuum/heat gel drying system (90 min, 80  C). 16. Expose the dry gel to a phosphor screen for 1 day and scan by phosphorimager (see Note 20). 17. Quantify 35S-signal by ImageJ software. 3.7 Recapitulating a G1-Like State in NDB Extracts

3.7.1 Mitotic Exit in Tet-Induced NDB Cells

Both NDB cells and extracts override mitotic arrest in the presence of small molecule inhibitors of Cdk1 (e.g., RO-3306) (Fig. 3). Inactivation of Cdk1 effectively generates a synchronous population of NDB cells in a quasi G1 state. In vitro, this treatment signals mitotic exit into a G1-like state (see Notes 21 and 22). The end result is a complementary extract system in which mitotic vs. G1 signaling of target proteins can be tested, including the Cdc20 vs. Cdh1 specificity of APC/C targets [24]. 1. Culture NDB cells in plates until they reach 75–80% confluency. 2. Treat cells with 1 μg/mL Tet for 22 h. 3. Collect and transfer mitotic cells into a 50 mL tube. Centrifuge for 5 min (room temperature [RT], 220
g) and discard supernatant. 4. Resuspend with fresh, Tet-free, prewarmed media and reculture cells on plates. 5. Treat cells with 15 μM RO-3306 FOR 2–2.5 h. 6. Validate mitotic exit by monitoring cell morphology (phase contrast microscopy) and DNA quantification (see Subheading 3.2). 7. Harvest cells for immunoblot or other assays.

3.7.2 Mitotic Exit in NDB Extracts

1. Prepare fresh mitotic NDB extracts or thaw premade extracts on ice (see Subheading 3.5 for details). 2. Incubate NDB mitotic extracts with 15–30 μM RO-3306 for 15–30 min at 28  C. Place tube on ice. 3. For Western blotting or protein immunoprecipitation assays one can either use fresh RO-3306-activated NDB extracts or snap-freeze extracts and conduct the assay later.

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Fig. 3 Mitotic NDB cells and extracts can exit mitosis by Cdk1 inhibition. NDB cells induced by Tet are static in an anaphase-like state. These cells can, however, progress into G1 following treatment with the Cdk1 inhibitor RO-3306. Phase images and DNA distributions are shown before and after 2.5 h treatment with RO-3306. Similarly, RO-3306 induces mitotic exit in NDB extracts. The resulting G1-like extracts support the degradation of Tome-1 (APC/CCdh1 substrate). Moreover, the mitotic—Cdk1-dependent—mobility shift of this protein (top right panel) is no longer apparent (bottom right panels). Tome-1 degradation is G1-like NDB extracts is facilitated by UbcH10 and blocked by MG132. These results demonstrate the shift from APC/CCdc20- to APC/ CCdh1-specific activity in mitotic vs. G1-like NDB extracts. Degradation of Tome-1 (35S-labeled, IVT product) was assayed in 28  C, and resolved by SDS-PAGE and autoradiography

4. For Western blotting, we typically mix 50–80 μg of the extracts with 2
Laemmli buffer, denature samples for 10 min in 95  C, and resolve by SDS-PAGE. 5. For degradation or mobility-shift assays, assemble a reaction mix on ice and conduct the assay (see Subheading 3.6). 3.7.3 Immunoprecipitation of APC/C in Mitotic vs. G1-Like NDB Extracts

1. Wash 15 μL agarose-conjugated anti-Cdc27 monoclonal antibodies (AF3.1 clone) with 500 μL PBS. 2. Centrifuge beads for 1 min (250
g, 4  C). Aspirate supernatant using a fine protein-loading tip while tilting the tube sideways to minimize contact with the beads. 3. Wash agarose-conjugated antibodies with 500 μL wash buffer I (150 mM NaCl). Centrifuge beads for 1 min (250
g, 4  C). Aspirate supernatant. 4. Prepare or thaw 200–250 μL of mitotic and/or G1-like NDB extracts (see Subheading 3.5).

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5. Put aside 10 μL of the extracts for input sample. Mix this input sample with 2
Laemmli buffer, denature 10 min at 95  C and store at 80  C. 6. Mix cell extracts with (a) 100 μL Wash buffer I; (b) protease and phosphatase inhibitor cocktails; and (c) the prewashed agarose-conjugated antibodies. Rotate tube for 4 h at 4  C. 7. Spin-down agarose-conjugated antibodies (1 min, 220
g, 4  C), aspirate supernatant, resuspend with 500 μL wash buffer I, rotate tube 5 min at 4  C, and centrifuge for 1 min (250
g, 4  C). Repeat step 7. 8. Resuspend agarose-conjugated antibodies with 500 μL wash buffer II (75 mM NaCl), Rotate tube 5 min at 4  C, and centrifuge for 1 min (250
g, 4  C). 9. Resuspend agarose-conjugated antibodies with 15 μL 2
Laemmli buffer, denature 10 min at 95  C and resolve by SDS-PAGE. 10. Process the gel for Western blotting (see Note 23).

4

Notes 1. Primers for generating nondegradable mutant of human Cyclin B1 were designed using the Agilent QuikChange Primer Design tool. Two missense mutations are introduced for substituting Arg 42 to Gly and Leu 44 to Val. 2. NDB cell system were generated based on 293T-REx™. Flp-In™ T-Rex™-293 cell system can be used instead (Thermo Fisher Scientific; #R78007). 3. Repetitive attempts to generate NDB cells based on T-REx™-HeLa cell line failed. 4. Either Tet or doxycycline can be used for activating expression of nondegradable Cyclin B1. 5. Derivates of HEK293 cells are easy to transfect. We used standard HeBSx2 CaCl2 transfection protocol for generating NDB cells. However, we expect any other transfection reagent to be successful. 6. Colony picking can be performed manually using cloning cylinder (e.g., PYREX® 6
8 mm Cloning Cylinders; # 3166-6). To this end, aspirate media, wash the plate with PBS, and dip the end of the cloning cylinder into sterile silicone grease before pressing to the bottom of a culture dish to create an isolated well around the colony. Trypsinize the cell colony with 2–3 μL Trypsin solution. After 1–2 min, collect cells and transfer into a 24-well-dish filled with 0.5 mL media and selection antibiotics. Alternatively, trypsinize cell colonies and utilize single cell sorting pipeline for sorting individual cells into a

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96 well-dish filled with conditioned media (generated from 293T-REx™ cell culture) and selection antibiotics. 7. Tet solution is ethanol based. To minimize ethanol volume in the media, prepare a stock solution of 2.5–5 mg/mL. Higher Tet concentration might be more challenging to dissolve. After mixing Tet with 95% absolute ethanol, pipette vigorously or vortex. Centrifuge tube for 30 s (>250
g, RT) to assure there is no pellet of undissolved material. If so, incubate in 37  C water bath for 5 min before repeating the process until no pellet is detected. Note, Tet is considered a light-sensitive material. For preparation of NDB mitotic extract, we recommend using a freshly made Tet solution. As a default, we add 10 μL of 2.5 mg/mL Tet solution into a 150 mm/diameter plate containing 25 mL medium. 8. Tet-induced NDB cells arrest with high APC/CCdc20 activity. Consequently, the level of endogenous Cyclin B1 is low and the overall signal of Cyclin B1 at this stage results almost exclusively from the exogenous variant. Cell colonies exhibiting highest levels of nondegradable Cyclin B1 following mitotic arrest should be prioritized for further characterization. 9. NDB cells are always maintained in the presence of blasticidin and Zeocin™. However, to avoid unnecessary additional costs, there is no need to add these antibiotics during culture expansion for extract preparation. 10. Harvest mitotic arrested NDB cells by gentle pipetting using 25 or 50 mL pipette. Pipettes with smaller orifices may damage the cells. After centrifugation, resuspend pellet by pipetting slowly 2–3 times using 25 or 50 mL pipette. 11. During extract preparation, cells are shredded in a 10 mL beaker on ice. To minimize foaming: (a) tilt the beaker while shredding the cells; (b) pull syringe plunger back gently; (c) always leave a quarter of the liquid in the beaker; (d) maintain needle tip in liquid during the entire process; and (e) use blunt tip needles. 12. NDB extract cannot be reused once defrosted. We recommend storing extracts in aliquots of 45–65 μL in 0.2 mL PCR tubes. 13. NDB extracts remain active up to 6 months in 80  C. 14. For expressing protein substrates in vitro we recommend using pCS2 or pCS-based vectors and the TNT® SP6 coupled reticulocyte lysate system. Protein translation was satisfactory in both premixed and non-premixed lysate systems (Promega; #L4600). Expression by T3 or T7 promoters is possible by matching TNT® systems from Promega. 15. Degradation and mobility shift assays can be performed with tagged IVT products (e.g., Myc or FLAG). Then, detection is based on immunoblot and primary antibodies coupled to a

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fluorophore, rather than autoradiography. A designated imaging system is required for signal quantification (e.g., Odyssaey®; LI-COR). 16. Mitotic NDB extracts are highly active. We often find recombinant ubiquitin and freshly added E-mix superfluous. Moreover, degradation assays in 10–15 μL extracts (instead of 20 μL) are almost equally informative (data not shown). In fact, extracts diluted two and even fourfold in swelling buffer can still support targeted proteolysis, though with slower kinetics (Fig. 2). These qualities save cell extracts. 17. For degradation and mobility shift assays we recommend sampling 3–5 μL for each time point. Lower amount saves reagents but with the cost of a fainter signal. On the flip side, higher sample volume may damage protein separation by SDS-PAGE due to overloading (5 μL reaction mix is about 100 μg total protein). 18. Enzymatic assays in cell free systems are highly sensitive to temperature shifts (Fig. 2). Higher temperature will shorten the assay but may lower the resolution of protein dynamics (Fig. 2). This can be critical in phosphorylation-mediated mobility shift assays (Fig. 1). On the flip side, increased temperature may compensate for lower activity associated with imperfect batch preparation, diluted extracts, or the lack of facilitating reagents (e.g., ubiquitin or E-mix). 19. We find mitotic NDB extracts to be more active than equivalent mitotic extracts from HeLa S3. In optimal assay conditions (see above), there is no real reason to extend degradation assays beyond 1 h. Within this time frame, time-intervals of 15 min are recommended (Figs. 1 and 2). Phosphorylation-mediated mobility shifts are considerably faster than proteolysis; a maximal mobility shift can be obtained within 15–30 min, if not faster. Oversampling during this time period is recommended for monitoring orderly phosphorylation (Fig. 1). 20. The default exposure time for autoradiography is one day. In case of a faint signal, increasing exposure time to 3 days is helpful. For the most part, short exposure for 1–2 h is informative though provide a less esthetic image. 21. Mitotic exit in mitotic arrested NDB cells and extracts is induced by RO-3306, that is, the most selective Cdk inhibitor to date. However, we expect roscovitine and flavopiridol to be equally effective for this purpose [5, 29]. 22. Incubation time and RO-3306 concentration must be calibrated per extract batch by testing (a) the level of endogenous Cdc20 (an APC/CCdh1 target) in extracts before and after Cdk1 inactivation by RO-3306 using immunoblot. Cdc20 levels should drop during mitotic exit as APC/CCdh1 becomes active;

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(b) the potency of the extracts to support APC/CCdh1-mediated degradation of Tome-1 IVT product following mitotic exit (see Subheading 3.6); and (c) the switch between Cdc20 bound APC/C to Cdh1 bound APC/C by immunoprecipitation of Cdc27 (a core APC/C subunit) followed by immunoblot with anti-Cdc20/Cdh1 antibodies (see Subheading 3.7.3). 23. We recommend over-night incubation (4  C) with anti-Cdc20 and anti-Cdh1 primary antibodies, and the use of IgG light chain-specific HRP-coupled secondary antibodies.

Acknowledgments This study was by the Israel Cancer Research Fund (ICRF), Grant no. RCDA00102, and the Israel Science Foundation (ISF) Grant no. 659/16 and 2038/19. The Emanuele lab is supported by funds from the UNC University Cancer Research Fund, National Institutes of Health (R01GM120309, R01GM134231), American Cancer Society (RSG-18-220-01-TBG) and donations from the Brookside Foundation. References 1. Morgan DO (2007) The Cell Cycle, Principles of Control. New Science Press, 2007. 297 pp. ISBN: 978-0-9539181-2-6 2. Sudakin V, Ganoth D, Dahan A et al (1995) The cyclosome, a large complex containing cyclin-selective ubiquitin ligase activity, targets cyclins for destruction at the end of mitosis. Mol Biol Cell 6:185–197 3. King RW, Peters JM, Tugendreich S et al (1995) A 20s complex containing CDC27 and CDC16 catalyzes the mitosis-specific conjugation of ubiquitin to cyclin B. Cell 81:279–288 4. Zou H, McGarry TJ, Bernal T et al (1999) Identification of a vertebrate sister-chromatid separation inhibitor involved in transformation and tumorigenesis. Science 285:418–422 5. Zur A (2001) Securin degradation is mediated by fzy and fzr, and is required for complete chromatid separation but not for cytokinesis. EMBO J 20:792–801 6. Kernan J, Bonacci T, Emanuele MJ (2018) Who guards the guardian? Mechanisms that restrain APC/C during the cell cycle. Biochim Biophys Acta Mol Cell Res 1865(12):1924–1933 7. Barford D (2020) Structural interconversions of the anaphase-promoting complex/

cyclosome (APC/C) regulate cell cycle transitions. Curr Opin Struct Biol 61:86–97 8. Yamano H (2019) APC/C: current understanding and future perspectives. F1000Res 8: F1000 Faculty Rev-725 9. Vorlaufer E, Peters JM (1998) Regulation of the cyclin B degradation system by an inhibitor of mitotic proteolysis. Mol Biol Cell 9:1817–1831 10. Buendia B, Draetta G, Karsenti E (1992) Regulation of the microtubule nucleating activity of centrosomes in Xenopus egg extracts: Role of cyclin A-associated protein kinase. J Cell Biol 116:1431–1442 11. Shamu CE, Murray AW (1992) Sister chromatid separation in frog egg extracts requires DNA topoisomerase II activity during anaphase. J Cell Biol 117:921–934 12. Masui Y (1992) Towards understanding the control of the division cycle in animal cells. Biochem Cell Biol 70(10–11):920–945 13. Funabiki H, Murray AW (2000) The Xenopus chromokinesin Xkid is essential for metaphase chromosome alignment and must be degraded to allow anaphase chromosome movement. Cell 102:411–424

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14. Murray AW, Desai AB, Salmon ED (1996) Real time observation of anaphase in vitro. Proc Natl Acad Sci U S A 93:12327–12332 15. Nguyen PA, Groen AC, Loose M et al (2014) Spatial organization of cytokinesis signaling reconstituted in a cell-free system. Science (80) 346:244–247 16. McGarry TJ, Kirschner MW (1998) Geminin, an inhibitor of DNA replication, is degraded during mitosis. Cell 93:1043–1053 17. Ayad NG, Rankin S, Ooi D, et al (2005) Identification of ubiquitin ligase substrates by in vitro expression cloning. Methods Enzymol. 399:404–414 18. Ayad NG, Rankin S, Murakami M et al (2003) Tome-1, a trigger of mitotic entry, is degraded during G1 via the APC. Cell 113:101–113 19. Rankin S, Ayad NG, Kirschner MW (2005) Sororin, a substrate of the anaphase-promoting complex, is required for sister chromatid cohesion in vertebrates. Mol Cell 18:185–200 20. Nguyen H, Gitig DM, Koff A (1999) Cell-free degradation of p27 kip1 , a G 1 cyclindependent kinase inhibitor, is dependent on CDK2 activity and the proteasome. Mol Cell Biol 19:1190–1201 21. Rape M, Kirschner MW (2004) Autonomous regulation of the anaphase-promoting complex couples mitosis to S-phase entry. Nature 432:588–595

22. Pe’er T, Lahmi R, Sharaby Y et al (2013) Gas2l3, a novel constriction site-associated protein whose regulation is mediated by the APC/CCdh1 complex. PLoS One 8(2): e57532 23. Cohen M, Vecsler M, Liberzon A et al (2013) Unbiased transcriptome signature of in vivo cell proliferation reveals pro- and antiproliferative gene networks. Cell Cycle 12:2992–3000 24. Wasserman D, Nachum S, Cohen M et al (2020) Cell cycle oscillators underlying orderly proteolysis of E2F8. Mol Biol Cell 2020: mbcE19120725. 25. Glotzer M, Murray AW, Kirschner MW (1991) Cyclin is degraded by the ubiquitin pathway. Nature 349:132–138 26. Zur A, Brandeis M (2002) Timing of APC/C substrate degradation is determined by fzy/fzr specificity of destruction boxes. EMBO J 21:4500–4510 27. Panet E, Ozer E, Mashriki T et al (2015) Purifying cytokinetic cells from an asynchronous population. Sci Rep 5:13230 28. Zeng X, King RW (2012) An APC/C inhibitor stabilizes cyclin B1 by prematurely terminating ubiquitination. Nat Chem Biol 8:383–392 29. Potapova TA, Daum JR, Pittman BD et al (2006) The reversibility of mitotic exit in vertebrate cells. Nature 440:954–958

Chapter 12 EDU (5-Ethynyl-20 -Deoxyuridine)-Coupled Fluorescence-Intensity Analysis: Determining Absolute Parameters of the Cell Cycle Joa˜o A. Ferreira, Marco Neves, Miguel Alpalha˜o, Pedro Pereira, Daniela Cunha, Fernando Ferreira, Rene´ Santus, Ana E. Sousa, and Paulo L. Filipe Abstract The principles and practice of a methodology of cell cycle analysis that allows the estimation of the absolute length (in units of time) of all cell cycle stages (G1, S, and G2) are detailed herein. This methodology utilizes flow cytometry to take full advantage of the excellent stoichiometric properties of click chemistry. This allows detection, via azide-fluorochrome coupling, of the modified deoxynucleoside 5-ethynyl-20 -deoxyuridine (EDU) incorporated into replicated DNA through incremental pulsing times. This methodology, which we designated as EdU-Coupled Fluorescence Intensity (E-CFI) analysis, can be applied to cell types with very distinct cell cycle features, and has shown excellent agreement with established techniques of cell cycle analysis. Useful modifications to the original protocol (Pereira et al., Oncotarget, 8:40514–40,532, 2017) have been introduced to increase flexibility in data collection and facilitate data analysis. Key words E-CFI, Flow cytometry, Cell cycle phases, Absolute length (units of time)

1

Introduction The E-CFI (EDU-coupled fluorescence intensity) analysis method allows for the accurate estimation of the absolute duration (in units of time) of S phase, as well as the absolute durations of G1 and G2 phases of the cell cycle in asynchronously growing cells. This contrasts with most techniques described thus far for cell cycle analyses using flow cytometry. Indeed, most of the described approaches provide data on relative distributions of cells along the different cell cycle phases [1, 2]. These frequencies of distribution only allow conclusions restricted to the relative duration of each phase. Absolute estimates (in time units) for the duration of each cell cycle stage

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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are achieved indirectly by combining data on doubling times of cell populations, which reflect the full duration of the cell cycle; doubling times are typically assessed during the phase of exponential growth. This poses limitations to experiments that are conducted at other growth phases. Also, these approaches do not allow the extraction of accurate data on absolute durations of the distinct cell cycle phases in experiments in which cell cycle progression is perverted; for instance, by interference with progression through one or more of its phases. For example, a higher proportion of cells in G1 phase does not necessarily mean a delayed progression through G1 phase; it could, alternatively, mean a shortening of S and/or G2 phases. E-CFI, by yielding absolute estimates of the different cell cycle phases, may thus provide a useful alternative for refined analysis in studies involving the cell cycle. E-CFI utilizes the excellent stoichiometric properties of click chemistry [3] for detecting, via an azide-fluorochrome complex, the modified deoxynucleoside 5-ethynyl-20 -deoxyuridine (EDU) incorporated into replicated DNA [4, 5]. When using E-CFI to estimate the absolute duration of S phase (in units of time) asynchronously growing cells need to be pulsed with EDU (5-ethynyl-20 -deoxyuridine) for incremental periods of time (typically 1-h increments). Pulses that are shorter than the duration of S phase have no chance for full labelling of replicating genomes, and therefore no chance of yielding EDU-coupled maximal fluorescence intensities. As pulsing times equal (or come close to) the duration of S phase, most cells in asynchronous populations are expectedly not exposed to EDU for the full length of their S phases. However, for an elite cohort of cells, this may not be the case. In these cells the beginning of the pulse may coincide with the initiation of their S phase, and thus the end of the pulse shall coincide with their ending of S. These cells shall show maximal EDU-coupled fluorescence. Predictably, increasing pulsing times shall not increase maximal fluorescent signals but, instead, the number of cells featuring maximal fluorescence intensities. Therefore, the minimal pulsing time to achieve maximal EDU-coupled fluorescence intensity corresponds to the mean duration of S phase for the peculiar cell type under study ([3]; see also Fig. 1). Importantly, E-CFI also allows determination of absolute durations (in units of time) of G1 and G2 phases. Since all cell cycle stages are estimated simultaneously, the duration of an E-CFI experiment is typically much shorter than that of a full cell cycle. As previously mentioned, quantitation of the absolute duration of S phase relies on analyses of fluorescent signals originating from EDU-positive cells. By contrast, to estimate the absolute durations of G1 and G2 phases E-CFI resorts on analysis of data obtained exclusively from EDU-negative cell populations.

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Fig. 1 Determination of absolute length of S phase in U2OS (osteosarcoma) cells using E-CFI. Cells were exposed for incremental pulsing periods to EDU (10 μM; 1 h increments) before collection. (a) Control cells not exposed to EDU and incubated with EDU detection mix. This allows identification of the background staining introduced by the EDU detection system. This is relevant to identify the EDU-negative zone in histograms of EDU-pulsed cells. (b, d) Cells pulsed with EDU (10 μM) for 1 h yielding peaks of background and EDU-coupled fluorescence that are clearly identified in the histogram (b); in (d) a dual parameter plot (DNA/PI vs EDU-Alexa 488) of the same sample is shown. Note the discrimination between cells in G1 and G2 stages, and the values of their relative proportions; the intensity maxima of EDU-coupled fluorescence (green double arrow) for this specific experiment have not yet been reached. (c, e) Cells pulsed with EDU (10 μM) for 7 h. In the histogram (c) note that the maximum of EDU-coupled fluorescence has been reached by EDU-positive cells (green double arrow); the same can be seen in the corresponding dual parameter plot (e). Pulsing times greater than 7 h did not yield higher intensity maxima of EDU-coupled fluorescence, establishing S phase length as 7 h. Also, in the plot image (e) note the abnormally high DNA content—denoted by the red double arrow to the right—featured by cells with the highest EDU-coupled fluorescence. This is a recurrent artifact that, importantly, is absent from EDU-negative cells in G1 (2n) and G2 (4n) phases (d, e). Therefore, accurate estimates of their relative proportions within whole cell populations remain possible throughout the increasing pulsing periods with EDU required for E-CFI experiments

As EDU pulsing times increase, the EDU-negative cells initially in G2 phase progressively move through G2 heading a trail of EDU-positive G2 cells; this trail of EDU-positive G2 cells stems from cells that incorporated EDU during the preceding S phase. Importantly, the time that the target EDU-negative G2 population takes to decline to reach nearly null levels, via progressive

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Fig. 2 Determination of the absolute lengths of G1 and G2 phases in U2OS (osteosarcoma) cells. (a) In a typical E-CFI experiment, the proportions (relative to the whole cell population) of EDU-negative cells in G2 and G1 phases were estimated after staining for EDU plus DNA (DAPI). The time of decline of the EDU-negative G2 population until near-null levels are reached denotes the absolute duration of G2 phase (~7–8 h). After this time point is reached (red arrow), the decline in the percentage of EDU-negative G1 cells becomes obvious; the time these cells take to reach near-null levels equals the absolute length of G1 phase. However, in a typical E-CFI experiment (10–12 h) near-null levels may not be reached in cells with a long G1 phase. (b) Shown are regressions of the linear decrease of the proportion of cells in G1 and G2 in each time period. Note that linear regression analysis for judging G1 length initiates only after G2 cells reach near-null levels (red arrow). In this specific case the estimated length of G1 phase was ~16 h, a period of time that is longer than that of a typical E-CFI experiment (10–12 h)

incorporation into the subsequent G1 compartment, shall be monitored. Indeed, this time corresponds to the absolute duration of G2 phase. Of note, the decline of this target EDU-negative G2 population follows a kinetics consistent with linear regression (Fig. 2). A similar reasoning, that is, appreciation of the progressive decline of the EDU-negative population, also applies for assessing the absolute duration of G1 phase. In this case, however, the progressive loss of G1 cells through their incorporation into the subsequent S phase compartment is initially compensated by a concurrent incoming of EDU-negative cells originating from the preceding G2 phase. This incoming period equals the duration of G2. This explains why, during the initial EDU pulsing times, the EDU-negative G1 population changes little for a period of time that is similar to the duration of the preceding G2 stage (Fig. 2a). Therefore, for the decline in the EDU-negative G1 population to reflect the duration of G1 stage it should be judged from time points that stand after the length of G2 stage (Fig. 2). Again, the decline of this target EDU-negative G1 population follows a kinetics that is amenable to analysis by a linear regression procedure (Fig. 2). To maintain EDU pulsing periods within limits of the practical, that is, 10–12 h of maximum pulsing times, for cell types with unusually long G1 phases (e.g., prequiescent), we devised an additional experimental group. This group, which we termed “guide,”

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is simple and fast to process. The guide experiment provides more accurate estimates of the relative proportions (percentages) of cells at distinct cell cycle phases than canonical cell cycle histograms. In the latter histograms poorly defined borders between G1 and S phases, and between S and G2 phases, may pose problems in determining the percentage of cells at each cell cycle phase; these problems may persist despite the use of dedicated algorithms incorporated into FlowJo platforms (e.g., Watson Pragmatic, Dean Jet Fox) [3, 6, 7]. As detailed in Methods, the guide group is useful for cells with long G1 phases, by allowing calculation of the absolute duration of G1 (in units of time) from its relative duration, that is, percentage in the whole cell population. This avoids extending EDU-pulsing periods. In summary, using incremental periods of exposure to EDU the E-CFI methodology allows determination of the absolute durations (in units of time) of (1) S phase, given by the minimal pulsing times to achieve maximal EDU-coupled fluorescence intensities, and of (2) G1 and G2 phases, assessed by the time of disappearance of, respectively, G1 and G2 cells that are EDU-negative. Analyses of G1 and G2 phases is further aided by simple linear regression procedures. For cells with unusually long G1 phases, the use of a brief pulsing with EDU (“guide” sample) may help in the accurate estimation of G1 phase length while keeping pulsing times within limits of feasibility.

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Materials Prepare all solutions using deionised water and analytical grade reagents, unless otherwise indicated.

2.1 Cell Culture and EDU Supplementation

1. Complete culture medium: cell culture medium (e.g., DMEM) plus supplements (fetal calf serum, antibiotics, nonessential amino acids, glutamine, etc.) prepared according to specific cell line requirements. 2. EDU stock solution: Dissolve EDU to 10 mM in DMSO and keep at 20  C until use. 3. Culture medium supplemented with EDU: Dissolve EDU stock solution to a final concentration of 10 μM in complete culture medium just before use and maintain at 37  C. Plan total volume of EDU-supplemented medium such that ~3 mL medium is added per sample (typically, per 60 mm diameter petri dish). Also, prepare complete culture medium containing 20–30 μM EDU if a guide sample is planned. 4. TrypLE Express: This provides milder trypsinization than more classical trypsin solutions.

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2.2 Cell Collection and Flow Cytometry

1. Phosphate-buffered saline (PBS), pH 7.4. 2. Triton X-100 stock solution: Prepare a 10% (vol/vol) of Triton X-100 in water and keep at 4  C for a maximum of 4 weeks. 3. PBS-Triton X-100: PBS, 0.05%. Triton X-100. 4. Sodium azide stock solution: Dissolve sodium azide to a final concentration of 20% (w/vol) in water; store at 4  C in the dark. This solution is stable for at least 1 year. 5. PBS-FCS: PBS, 5% fetal calf serum, 0.05% Triton X-100, 0.1% sodium azide. 6. BSA (bovine serum albumin) stock solution: Dissolve BSA to a final concentration of 20% (w/vol) in water, by gentle stirring at 4  C until fully dissolved. Aliquot and store at 20  C. 7. Flow cytometry buffer: PBS, 0.5% BSA, 0.1% sodium azide. According to users experience other buffer formulations may be tested. 8. Formaldehyde fixative: 2% formaldehyde in PBS (pH 7.4) prepared from a commercially available solution at 37–40%. 9. Formaldehyde-Triton X-100 fixative: 2% formaldehyde in PBS (pH 7.4), 0.2% Triton X-100. 10. DAPI (4,6-diamidino-2-phenylindole): 300 ng/mL, 0.05% Triton X-100, in PBS (pH 7.4). DAPI is diluted from a stock solution at 1 mg/mL in water, kept at 20  C. 11. PI (propidium iodide): 10 μg/mL, RNase A (190 μg/mL), 0.1% Triton X-100, in PBS (pH 7.4). PI is diluted from a stock solution at 1 mg/mL in water, kept at 20  C. 12. EDU Cell Proliferation Kit: kits containing azide coupled to Alexa 488 or 6-FAM (6-carboxyfluorescein) have been extensively used by us; other azide-coupled fluorochromes may, however, be tested. 13. Hemocytometer or automated cell counter. 14. Flow cytometer equipped with appropriate laser lines.

3

Methods

3.1 Cell Culture and Labelling with EDU

1. Plan the following experimental groups: (a) cells to be exposed to EDU (10 μM) for increasing periods of time; hourly increments are generally utilized; maximal incorporation times should be within the range of 10–12 h for most cell types, thus yielding 10–12 samples (i.e., 10–12 petri dishes or tissue culture flasks); (b) one sample (“guide sample”) of cells to be exposed to a higher concentration of EDU (20–30 μM) for a brief period of time (10–15 min); this shall provide useful

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information on the distribution (percentage) of cells per cell cycle stage, particularly important for estimates on the duration of G1 and G2 phases; (c) two samples of cells not exposed to EDU (no-EDU); one of the samples is to be incubated with EDU detection mix in parallel with the EDU-exposed samples, to determine the background (nonspecific) fluorescence, that is, fluorescent signals not stemming from EDU-DNA; this sample shall be further stained for DNA (e.g., DAPI staining) to provide a typical cell cycle histogram. The second sample of cells not exposed to EDU shall remain unstained, to be later used for calibration purposes in preparation for imaging by flow cytometry. 2. Split cells at identical numbers per culture dish. This facilitates subsequent adjustments between cell numbers and volumes of EDU detection mix per sample (see Note 1). 3. When cells reach exponential growth (see Note 1), at 1-h intervals replace the regular culture medium by prewarmed EDU-supplemented medium (3 mL/culture dish) (see Note 2). Add EDU-supplemented medium (20–30 μM; 10–15 min) to guide sample just before the planned first (or second) moment of collection (see Note 2). Remember to keep at least two additional culture dishes of cells EDU-free. 4. For cell collection wash cells twice in sterile PBS, pH 7.4. Add 1 mL TrypLE Express per dish and incubate at 37  C until cells detach. Add 9 mL of complete culture medium per dish. Resuspend cells by vigorous pipetting to disrupt cell aggregates. 5. Transfer cells to 14 mL Falcon tubes. Centrifuge in a benchtop centrifuge at 300  g, 7 min at room temperature. Discard all supernatant and gently scrape the tubes over a rough surface to disaggregate cell pellets. Place cells on ice. 6. Add 200 μL of ice-cold formaldehyde fixative to each cell pellet. Pipette vigorously (20–30 times) with a micropipette (yellow tips preferred) to obtain single cell suspensions. Fix pellets for 3–5 min on ice. 7. Add 1 mL of formaldehyde–Triton X-100 fixative and homogenize by gentle pipetting (harsh pipetting at this stage can result in disruption of cell membranes). Fix-permeabilize samples for an additional 10 min on ice. 8. Add ~12 mL of PBS–Triton X-100 to each sample. Centrifuge at 680  g, 9 min (see Note 3). Discard supernatant. 9. Resuspend cell pellets in 200 μL of PBS–Triton X-100 by vigorous pipetting (20–30 times), and then add an additional 1 mL of PBS–Triton X-100.

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10. Take a 30 μL aliquot of cell suspension per sample for cell counting purposes (hemocytometer; automated cell counter). Adjust cell suspension volumes to obtain identical cell concentrations per group (see Note 4). 11. Add ~12 mL of PBS–Triton X-100 to each volume-adjusted sample. Centrifuge at 680  g, 9 min at room temperature. Discard supernatant. Do another short spin at 680  g and remove any remaining buffer with a micropipette. Gently scrape the tubes over a rough surface to loosen the cell pellets. Samples are now ready for detection of EDU-DNA. 3.2 EDU-DNA Detection

1. For detection of EDU-DNA prepare the appropriate volume of EDU-click reaction mix according to manufacturer’s instructions immediately before usage (see Note 5). Place reaction mix on ice in the dark. Incubations with reaction mix, and subsequent washing and incubation steps are performed in the dark unless stated otherwise. 2. Add to each sample 150–200 μL of reaction mix/2  106 cells (see Note 4). Resuspend cells in reaction mix by vigorous pipetting (20–30 times; yellow micropipette tips). Incubate for 35–40 min at 37  C in the dark (see Note 6). Resuspend cell pellets by gentle vortexing (20 Hertz, 10 s), at least once, during the incubation period. 3. Add ~12 mL PBS-Triton X-100 per sample. Mix by inversion and vortex briefly (20 Hz, 20 s). Centrifuge at 680  g, 9 min at room temperature. Discard supernatant. 4. Resuspend cell pellets in 200 μL of PBS-FCS by vigorous pipetting (20–30 times) (see Note 7.) Add 1 mL of PBS-FCS and keep at 4  C, for a minimum of 30 min. 5. Add ~12 mL of PBS-Triton X-100 and vortex gently (20 Hz, 10 s). Centrifuge at 680  g, 9 min at room temperature. Discard supernatant. 6. Proceed to a prolonged storage procedure (see Note 8) when samples cannot be processed within 2–3 days of staining. Otherwise, proceed to next step. 7. Add to each sample 400 μL of DAPI. Resuspend samples by vigorous pipetting (20–30 times). Incubate at 37  C for 30–45 min. Alternatively, 400 μL of a PI solution can be used, but if using PI, incubate samples with PI for 30 min on ice prior to the 37  C incubation. Resuspend cell pellets by gentle vortexing (20 Hertz, 10 s), at least once, during incubation at 37  C (see Note 9). 8. Add ~12 mL of PBS–Triton X-100 per sample at end of the incubation period. Vortex gently (20 Hz,10 s). Centrifuge at 680  g, 9 min at room temperature. Discard supernatant. 9. Resuspend cell pellets in 250–400 μL of flow cytometry buffer. Note that other currently used formulations may be used.

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Samples stained for EDU (coupled to Alexa 488 or 6-FAM/6carboxyfluorescein), and DAPI (or PI) are analyzed using a three laser (blue—488 nm; red—640 nm; violet—605 nm) flow cytometer (see Note 10). 1. Measure EdU–Alexa 488 and PI signals via excitation by the blue laser using 530/30 and 695/40 bandpass filters, respectively. Measure DAPI signals through excitation by the violet laser (525/50 bandpass filter) (see Note 10). Acquire a minimum of 30,000 events per sample, in slow rate mode to avoid doublets. 2. Perform sample measurements using a dedicated software; for example, FACSDiva Software; BD CellQuest, or ModFit LT Software provide appropriate alternatives. 3. Perform data analysis, such as mean fluorescence intensity (MFI) measurements, with FlowJo Software (or equivalent alternative). Exclude cell debris and aggregates from the analysis, namely by using pulse processing FSC-A vs FSC-H, FSC-H vs FSC-W, SSC-H vs SSC-W, and FL2-A vs Fl2-W when appropriate. 4. Using FACSDiva Software (or equivalent) build histograms of EDU-coupled fluorescence and dual parameter plots (DNA versus EDU-coupled fluorescence) (Fig. 1). In EDU-less controls identify regions of nonspecific background staining introduced by the EDU detection system. Using this information identify similar regions in EDU-pulsed cells. Identify in EDU-pulsed cells (fluorescence histograms, dual parameter plots) the shorter time point at which the maximum EDU-coupled fluorescence is reached. This time point/pulsing period corresponds to the absolute length of S phase (Fig. 1). 5. Export to Excel files data on DNA content (2n, 4n) of EDU-negative (non-S phase) cells present in background peaks of EDU-pulsed samples. Use Excel analytical tools to build graphs featuring, across time points, relative proportions of EDU-negative G1 (2n) and G2 (4n) cells in the whole cell population (Fig. 2). The time these G1 and G2 cells take to decline to near-null levels corresponds, respectively, to the absolute lengths of G1 and G2 phases. Note, however, that estimation of G1 phase initiates only after completion of the G2 decline (Fig. 2). Use regression analysis when EDU-pulsing periods are not sufficient to judge G1 absolute length in full; typically, this analysis requires a minimum of four time points (Fig. 2). When four successive time points of decline of EDU-negative cells in G1 phase cannot be achieved (very long G1 phase) use data obtained from guide samples, as detailed next.

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Fig. 3 The guide experiment—a short EDU pulse to facilitate distinction between phases of the cell cycle. After a short pulse with EDU (20–23 μM; 10–15 min), S phase cells (EDU-positive) can be clearly distinguished from the EDU-negative G1 and G2 cells in fluorescence intensity histograms. Concurrent DNA staining (DAPI, propidium) allows the proportion of G1 (2n DNA content) and G2 (4n) to be accurately estimated at steady state, that is, at any time point during an E-CFI experiment

6. In cells with long G1 phases, typically nontransformed cell lines, we advise to use a guide sample to estimate the absolute length of G1 phase (Subheading 3.1, step 1). The guide sample provides a modified histogram that shows clearer separation of cells at different stages of the cell cycle when compared to classical histograms (Fig. 3). Specifically, G1 and G2 cells are separated from S phase cells by dispatching the EDU-positive S phase cells to regions of higher EDU-coupled fluorescence intensities (Fig. 3). After calculating the percentage of G1 cells (2n DNA content; DAPI or PI staining) within the whole cell population (FACSDiva, or similar software), and by combining this value with the previously obtained absolute durations of G2 and S phases (Subheadings 3.3, steps 4 and 5) estimates of the absolute duration of G1 (in units of time) can then be assessed. To this end, the following equation can be used: tG1 ¼ % cells in G1/% cells in G2  tG2, where tG1 is the duration of G1 in time units, and tG2 the duration of G2 in time units. Also, for increased robustness of data, the absolute length of S phase can also be also included in the estimates of the absolute length of G1, according to the following equation: tG1 ¼ % cells in G1/(% cells in G2 + % cells in S)  (tG2 + tS), where tG2 and tS correspond, respectively, to the absolute durations of G2 and S phases in time units. Note that all percentages are referred to the whole cell population.

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Notes 1. To obtain similar numbers of cells per petri dish adherent cells shall be split at the most appropriate dilution for the specific cell line such that the E-CFI experiment is initiated during exponential growth. Typically, exponential growth is achieved at sub-confluency (~60–70% confluency) and is highlighted by an increased fraction of mitotic cells. Prior knowledge of doubling times for the particular cell line under scrutiny is desirable but not essential. For a typical E-CFI experiment 12–14 petri dishes (60 mm diameter) shall be seeded by dispensing equal volumes of a well homogenized cell suspension per petri dish; this will ensure similar numbers of cells per dish. Six-well plates provide an appropriate alternative to petri dishes. 2. An alternative way to supplement cell cultures with EDU is by adding medium with a higher concentration of EDU to reach a final concentration of 10 μM. This is the preferred method for suspension cultures. Also, optimization of times of EDU addition to cell cultures and of cell collection is highly desirable to simplify the whole procedure. Thus, in the exemplary case of a maximum incorporation time of 10 h, EDU can be added at 1-h intervals, in parallel, to samples to be exposed for 10 and 5 h, and subsequently to those to be exposed for 9- and 4-h, then to the 8- and 3-h samples, and so forth, in decreasing order of exposure times. This reduces the moments of EDU supplementation to cultures to 5 and moments of sample collection to two. Cells exposed to EDU for 1–5 h are collected first, and cells exposed for 6–10 h later. EDU-negative controls and guide sample can be collected at any of these moments. Compared to sequential single supplementations with EDU at hourly intervals, the scheme of parallel supplementations proposed above also reduces the number of samples to be handled per collection session. We finally note that shorter increments in EDU pulsing periods, of for example 30 min, do not significantly improve temporal resolution in most cell types, with the possible exception of cells with very short cell cycles [3]. 3. Note that centrifugation conditions for living cells (300  g, 7 min) and fixed-permeabilized cells (680  g, 9 min) are different. 4. For the sake of data robustness in E-CFI experiments it is mandatory to keep constant, for each sample, the proportion of total cells versus volumes of EDU detection mix. E-CFI is a quantitative method using click chemistry which shows excellent stoichiometry [3]. This is only achieved, however, when the balance between cell number and detection mix volume is

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strictly maintained. We recommend using 150–200 μL of reaction mix/2  106 cells. Counting the number of cells per sample is thus a key issue. We note that little variation in total cell numbers are expected among groups since the whole experiment typically lasts only for 10–12 h, and cells are split at identical densities. 5. We have thus far tested EDU detection kits from two different suppliers with similar results. 6. With respect to detection of EDU-DNA, proceeding with the reaction to saturation such that most (if not all) ethynyl groups of EDU-DNA become coupled to fluorochrome- azide is relevant for quantitative purposes. We have systematically used incubation times of 35–40 min, at 37  C, independently of the supplier of the EDU-Click kit. 7. Regarding the use of azide in solutions, the click reaction involved in EDU detection requires the covalent coupling of an azide-fluorochrome complex to the alkyne group of EDU. Therefore, the presence of azide in solutions that are utilized before the detection reaction is to be avoided. Azide containing solutions can, however, be used afterward. 8. For prolonged storage of EDU stained samples, after detection of EDU-DNA and washing in PBS-Triton (step 6, in Subheading 3.2) cell pellets can be resuspended by pipetting in 200 μL of distilled water, 0.05% TX-100 (precipitation of salts in presence of ethanol may occur if PBS is used). Add 1 mL of 70% ethanol per sample; gently, resuspend the pellet. Samples can then be kept in the dark at 4  C for up to 3 months without loss of fluorescence. When required, add 12 mL of H2O, Tx-100 0.05% to each sample, vortex (2k Hz; 20 s) and centrifuge (680  g, 9 min). After discarding the supernatant, resuspend the pellets again in 200 μL of water, Tx-100 0.05%, then add 12 mL of water, Tx-100 0.05% to each sample, and centrifuge again (680  g, 9 min). These washing steps are intended at eliminating any trace of ethanol before proceeding for DNA staining with DAPI or PI (Subheading 3.2, step 6). 9. Prolonged incubations with EDU (10 μM; above 4–5 h) systematically introduce artifacts in DNA staining with either DAPI or PI. This is obvious in cells whose DNA is more heavily substituted with EDU and thus feature higher intensities of EDU-coupled fluorescence (Fig. 1c, e). Typically, DAPI tends to lead to underestimation of DNA amounts, with the appearance of sub-G1 regions in some histograms, whereas PI tends to generate overestimates (Fig. 1). The reason for these distortions remains to be elucidated. Importantly, these distortions are restricted to EDU-positive cells and do not interfere with estimates of DNA content in EDU-negative cells (Fig. 1, dual

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parameter plots). Therefore, accurate assessment of the lengths of G1 and G2 phases, which is made using exclusively EDU-negative cells, remains unperturbed by this phenomenon. Also, this phenomenon does not affect EDU-coupled fluorescence intensities, and thus estimates of S phase length. 10. For cell cycle studies using E-CFI only basic flow cytometry settings and data analysis tools are required. We have used systematically EDU coupled to either Alexa-488 or 6-FAM (6-carboxyfluorescein), both of which excite at 488 nm. When using other fluorochromes some of the parameters provided in Subheading 3 (step 1) may require adjustment. References 1. Rabinovitch P (1994) Introduction to cell cycle analysis. . Basics of DNA cell cycle analysis. Phoenix Flow Systems, Inc, San Diego, CA 2. Pozarowski P, Darzynkiewicz Z (2004) Analysis of cell cycle by flow cytometry. Methods Mol Biol 281:301–311 3. Pereira PD, Serra-Caetano A, Cabrita M, Bekman E, Barga J, Rino J, Santus R, Filipe PL, Sousa AE, Ferreira JA (2017) Oncotarget 8:40514–40532 4. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci U S A 105:2415–2420

5. Cavanagh BL, Walker T, Norazit A, Meedeniya AC (2011) Thymidine analogues for tracking DNA synthesis. Molecules 16:7980–7993 6. Dressler LG (1993) DNA flow cytometry measurements as surrogate endpoints in chemoprevention trials : clinical, biological, and quality control considerations. J Cell Biochem 17G:212–218 7. Baldetorp B, Bendahl PO, Ferno M, Alanen K, Delle U, Falkmer U, Hansson-Aggesjo B, Hockenstrom T, Lindgren A, Mossberg L, Nordling S, Sigurdsson H, Sta˚l O, Visakorpi T (1995) Reproducibility in DNA flow cytometric analysis of breast cancer: comparison of 12 laboratories’ results for 67 sample homogenates. Cytometry 22:115–127

Chapter 13 High-Resolution Analysis of Centrosome Behavior During Mitosis Vanessa Nunes, Margarida Dantas, Joana T. Lima, and Jorge G. Ferreira Abstract Cell division requires a dynamic reorganization of cytoskeletal and nuclear components. One essential step is the separation of centrosomes, which allows the assembly of a microtubule-based mitotic spindle. This has to be spatially and temporally coordinated with other events such as adhesion complex disengagement, assembly of an actin-rich cell cortex and nuclear envelope breakdown (NEB), to ensure chromosome segregation fidelity. Previous methodologies often focused on a single event and failed to provide an integrated view of the process. In this chapter, we describe a method to study mitosis with high resolution, by analyzing the dynamic interplay between centrosomes, nucleus, and cell membrane, using a combination of live-cell imaging and micromanipulation with custom-designed computational tools. Key words Centrosome, Mitosis, Automated tracking, Nucleus, Cell membrane

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Introduction Mitosis is the process by which cells segregate duplicated chromosomes between two daughter cells. Chromosome segregation is accomplished through a chain of highly regulated events, which include extensive reorganization of the cytoskeleton and cell nucleus [1, 2]. Many of these changes take place during the first stage of mitosis, termed “prophase.” During this stage, cells lose adhesion [3–5], and their volume changes [6–8] to allow the assembly of a microtubule-based mitotic spindle [9–12]. At the same time, within the nucleus chromosomes condense [13, 14] and the nuclear lamina disassembles [15, 16]. How these events are coordinated to ensure efficient chromosome segregation remains unclear. In animal cells, the duplicated centrosomes migrate in opposite directions along the nuclear envelope (NE), to originate the two spindle poles [17]. The molecular players involved in centrosome separation and positioning have been previously identified and include microtubules and microtubule-associated motors, such as

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Eg5 and dynein [12, 18–20], but also myosin II [21], actin [22] and Arp2/3 [12]. However, a comprehensive understanding of how these molecular factors cross talk in space and time to regulate centrosome behavior during prophase is absent. The study of highly dynamic events such as cell division has greatly benefited from live-cell imaging techniques, because they allow us to probe the spatiotemporal characteristics of the process [23]. This has been facilitated by the development of new microscopy techniques [24] and improved fluorescent probes [25], which are less phototoxic to cells [26, 27]. Importantly, live-cell imaging can be used in combination with microfabrication techniques that aim to recapitulate or mimic the complexity of the cellular microenvironment [28–30], or with cell confinement systems, that allow us to physically constrain the height and the stiffness applied onto cells [31, 32]. Used separately or in combination, these methods have provided a more realistic picture of the process of mitosis [12, 29, 33]. The ability to extract quantitative data from live-cell imaging datasets using image analysis software, has allowed the detailed characterization of key events that take place during mitosis [12, 34, 35]. In particular, analysis of centrosome motion has been extensively studied using both commercial and custom-designed tools [36–38]. However, current approaches fail to analyze the changes that occur at the cell membrane, nucleus and cytoskeleton in an integrated manner. In this chapter, we describe a methodology established in our laboratory to study centrosome, nucleus and cell membrane behavior during mitosis in human cells, using state-of-the-art, live-cell microscopy in combination with cell micropatterning and/or cell confinement techniques and posterior bio-image reconstruction and analysis. Using a custom-designed computation tool [39], we are able to precisely measure and correlate the dynamic changes that occur during mitosis.

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Materials Prepare, use, store, and dispose all the reagents according to the Material Safety Data Sheet (MSDS) guidelines. All the mediums and buffers used in the cell culture procedures should be prewarmed, unless otherwise stated.

2.1 Cell Lines and Culture Conditions

Unless otherwise noted, all human cell lines are cultured in complete medium and grown in a 37
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1. Human cell lines of interest. 2. Disposable 25 and 75 cm2 tissue culture flasks and 6-well plates. 3. Complete medium: Dulbecco’s Modified Eagle Medium supplemented with 10% heat-inactivated fetal bovine serum (FBS). 4. Trypsin: Trypsin-EDTA solution 0.25%. Can be prepared by adding 2.5 g porcine trypsin and 0.2 g EDTA.4Na to 1 L of Hanks’ Balanced Salt Solution. 5. Sterile phosphate-buffered solution (PBS), pH 7.4. 6. Disposable sterile pipettes (1, 5, and 10 mL). 7. Disposable conical tubes (15 and 50 mL). 2.2 Transient Plasmid DNA and Small Interfering RNAs (siRNAs) Transfections

1. Complete medium. 2. Reduced serum medium: DMEM supplemented with 5% FBS. 3. Opti-MEM. 4. Lipofectamine 2000. 5. Lipofectamine RNAiMAX. 6. Plasmid DNA of interest or siRNA oligos. 7. Solution A: (a) For DNA: Dilute 5 μL of Lipofectamine 2000 in 250 μL of Opti-MEM. (b) For siRNAs: Dilute 5 μL of Lipofectamine RNAiMAX in 250 μL of Opti-MEM. 8. Solution B: (a) For DNA: Dilute 1 ng/mL of the plasmid DNA of interest in 250 μL of Opti-MEM. (b) For siRNAs: Dilute 2 μL of the siRNA of interest (Stock Concentration ¼ 20 mM; Final Concentration ¼ 20 nM) in 250 μL of Opti-MEM. 9. Selection antibiotics, depending on the plasmid to be used (e.g., for HeLa cells, G418 at 200–400 μg/mL, puromycin at 1–2 μg/mL).

2.3 Micropatterning on Glass Coverslip with Deep UV Light

1. Deep UV lamp (185 nm wavelength; Novascan). 2. Synthetic quartz mask (Delta Mask B.V.). 3. Plasma cleaner (Zepto Plasma System, Diener Electronic). 4. Glass coverslips: 22  22 mm No. 1.5. 5. Milli-Q water. 6. 10 mM Hepes, pH 7.4: Dilute 238.30 g of Hepes (MW ¼ 238.30) in 1 L of ddH2O to make a 1 M stock. To adjust the pH, use HCl 37% or 0.1 M NaOH inside a flow hood. Filter-sterilize or autoclave the buffer. Dilute 5 mL of stock solution in 500 mL of dH2O for 10 mM solution.

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7. PLL-g-PEG: 0.1 mg/mL PLL(20)-g[3.5]-PEG(2) (SuSoS) in 10 mM Hepes pH 7.4. Dissolve PLL-g-PEG powder in 10 mM Hepes pH 7.4 to obtain a stock solution of 1 mg/mL. Before usage, dilute the stock solution in 10 mM Hepes pH 7.4 to obtain the working concentration of 0.1 mg/mL. 8. 100 mM NaHCO3, pH 8.6: Dilute 84 mg of NaHCO3 in 10 mL ddH2O. To adjust the pH, use HCl 37% or 0.1 M NaOH, inside a flow hood. 9. Fibronectin (FBN) solution: 25 μg/mL, diluted in 100 mM NaHCO3 (pH 8.6). 10. Alexa546 or 647-conjugated fibrinogen: 5 μg/mL, diluted in 100 mM NaHCO3 (pH 8.6). 11. Air gun. 12. Isopropanol. 13. Stainless steel forceps, 0.4 mm with angled tip. 14. PTFE forceps, 145 mm with pointed ends. 2.4

Cell Confinement

1. Microfluidic pressure pump and generator (AF1 Dual, ElveFlow). 2. PDMS cell confiner [custom-designed as described in [32] or obtained from 4Dcell]. The size of the PDMS confiner device will depend on the size of the imaging chamber. 3. Cell confinement slides (custom-designed or obtained from 4Dcell). To prepare custom-designed confinement slides, use round cover glass, 10 mm, 0.17 mm thickness, coated with PDMS micropillars according to [32]. 4. FluoroDishes 35 mm well, glass-bottom 0.17 mm thickness (World Precision Instruments). 5. Plasma cleaner (Zepto, Diener Electronics). 6. FBN solution. 7. Stainless steel forceps, 0.4 mm with angled tip.

2.5 Live-Cell Imaging

1. Spinning disk microscope. 2. Magnetic imaging chamber (Live Cell Instrument). 3. 6-well plates. 4. FluoroDishes 35 mm well, glass-bottom 0.17 mm thickness (World Precision Instruments). 5. Glass coverslips: 22  22 mm No. 1.5. 6. Complete medium. 7. Imaging medium: Leibovitz’s L-15 medium supplemented with 10% FBS. 8. Antibiotics (e.g., penicillin–streptomycin solution, antibiotic– antimycotic solution).

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1. Live-Cell Imaging datasets. 2. Bio-image analysis software. (a) ImageJ/Fiji. (b) MATLAB 2018 or newer version. (c) Tracking and reconstruction tools (e.g., Trackosome: available at https://github.com/Trackosome).

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3.1 Transient Plasmid DNA and Small Interfering RNAs (siRNAs) Transfections

Day 0 1. Seed the human cell line of interest in a 6-well plate well with 2 mL of complete medium (see Note 4). 2. Grow cells in a 37
C humidified incubator with 5% CO2, until they are 50–75% confluent. Day 1 3. Rinse the cells with sterile phosphate-buffered solution (PBS) and replace the cell culture medium with 1.5 mL of reduced serum medium. 4. Incubate cells in a 37
C humidified incubator with 5% CO2. 5. For cell transfection, prepare solutions A and B (see Notes 5 and 6). 6. Incubate solutions A and B separately for 5 min at room temperature (RT). 7. Mix solutions A and B and incubate for 30 min at RT. 8. Add the oligo-lipid complexes to the cells, dropwise. 9. Incubate the cells for 5–6 h in a 37
C humidified incubator with 5% CO2. 10. Replace the culture medium with 2 mL of complete medium. 11. Grow cells in a 37
C humidified incubator with 5% CO2 for 24, 48, or 72 h (see Note 7). 12. Analyse cells (see Note 8). 1. Activate glass coverslips with air plasma for 2 min (Fig. 1a1).

3.2 Micropatterning on Glass Coverslip with Deep UV Light

2. Incubate the activated side of the coverslip in 50–200 μL of PLL-g-PEG for 30 min-1 h at RT (see Note 9) (Fig. 1a2).

3.2.1 Coating of Glass Coverslips with PLL-g-PEG

3. Rinse the coverslips with dH2O, dry and store them at RT (see Note 10).

3.2.2 UV Illumination of PLL-g-PEG-coated Glass Coverslips Using a Quartz Photomask

1. Wash the quartz mask with isopropanol and air-dry it if necessary (see Notes 11 and 12). 2. Illuminate the brown side of the quartz mask with the UV lamp for 5 min (see Note 13) (Fig. 1a3).

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3. Seal the pegylated side of the coverslip onto the brown side of the quartz mask using 2–3 μL of Milli-Q water (see Notes 9 and 14) (Fig. 1a4). 4. Illuminate the silver side of the quartz mask with the UV lamp for 5 min (see Notes 13 and 15) (Fig. 1a5). 5. To detach the coverslips from the quartz mask, pour 2 to 5 mL of dH2O onto the coverslips and incubate for 2–3 min. 6. Collect the coverslips and stored them at RT after drying (see Note 16). 3.2.3 Seeding Cells on FBN-Patterned Glass Coverslips

1. Incubate the patterned face of the coverslip with 50–200 μL of FBN solution and Alexa-conjugated fibrinogen for 1 h, at RT (see Notes 9, 17, and 18) (Fig. 1a6). 2. Rinse the coverslips with PBS (see Notes 19 and 20). 3. Harvest human cell line of interest by washing with PBS and incubating with the Trypsin solution for 2 min. Trypsin volume

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will vary according to the surface area (e.g., for a 25mm2 flask, use 2 mL of Trypsin and for a 6-well plate well, use 1 mL of Trypsin). 4. Seed 50,000–200,000 cells in the FBN-patterned coverslips with 2 mL of complete medium supplemented with antibiotics (see Note 21 and 22) (Fig. 1a7). 5. Incubate the cells in a 37
C humidified incubator with 5% CO2 for 1–2 h (see Note 22). 6. Gently wash the unattached cells with complete medium. 7. Grow cells in a 37
C humidified incubator with 5% CO2 for 12–24 h. 8. Image cells on the microscope (see Subheading 3.4). 3.3

Cell Confinement

3.3.1 Cell Seeding for Confinement

1. Activate the FluoroDishes with air plasma for 2 min. 2. Coat the activated FluoroDishes with FBN solution for 30 min to 1 h, at RT. 3. Rinse FluoroDishes three times with dH2O. 4. Seed cells in the coated FluoroDishes with 2 mL complete medium supplemented with antibiotics. Number of cells to be seeded will depend on the cell type used (see Note 21). 5. Incubate the cells overnight at 37
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3.3.2 Using the Dynamic Cell Confiner

1. Replace the cell culture medium with 500 μL of imaging medium supplemented with antibiotics (see Note 23). 2. Add the confinement slide to the PDMS piston in the confiner cell (Fig. 2a). Assemble the dynamic confiner setup by connecting the PDMS pillar to the pressure/vacuum generator (see Note 24). 3. Gently place the PDMS pillar on top of the FluoroDishes with the cells. Gently press it down to ensure that the PDMS confiner is attached to the bottom and turn on the vacuum generator. 4. Lower the confinement slide onto the cells by generating a vacuum (Fig. 2b). 5. Proceed with live-cell image acquisition. Confinement can be modulated by increasing or decreasing the pressure on the vacuum line.

3.4 Live-Cell Imaging

Day 0 1. 12–24 h prior to each experiment, seed 50,000–200,000 cells (depending on cell type; see Note 21) in FBN-patterned coverslips with 2 mL of complete medium supplemented with antibiotics (see Note 1).

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Day 1 2. Approximately 30 min to 2 h prior to each experiment, assemble the magnetic imaging chamber and replace the cell culture medium with 1–2 mL of imaging medium supplemented with antibiotics. Add SiR-dyes or drugs of interest (acute pharmacological inhibition) to the cell culture medium at this time point (see Notes 1 and 25). 3. Turn ON all the imaging equipment and set the appropriate temperature and CO2 levels (37
C and 5% CO2) for the microscope chamber (see Note 26). 4. Set up your experimental conditions, namely the appropriate illumination wavelengths, laser power intensities, exposure times, and time-lapse conditions (see Notes 27 and 28). 5. If using a confinement setup, additional adjustments to your imaging setup are required at this stage. Refer to step 2. 6. Start imaging. Representative frames from a movie of a human U2-OS cell expressing H2B-GFP/tubulin-RFP, seeded on a line micropattern can be seen in Fig. 3a, b. 3.5 Analysis and Data Extraction from Live-Cell Imaging Datasets

Bio-image analysis and data extraction can be performed with our custom-designed tool [39]. For the purpose of this chapter, we will focus on the use of the Centrosome Dynamics and Compile Data modules.

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3.5.1 Single-Cell Analysis

1. Open 4D dataset files (e.g., *.nd2, *.tiff) using ImageJ/Fiji software. 2. Split channels (Image!Color!Split Channels) (see Note 29). 3. Adjust image brightness and contrast ge!Adjust!Brightness/Contrast!Reset).

(Ima-

4. Convert image to 8-bit (Image - > Type - > 8-bit). 5. Save each channel as a *.tiff file (e.g., Centrosomes.tiff, Nucleus.tiff, and Membrane.tiff). 6. Install the analysis tool in the MATLAB working directory. 7. Open MATLAB and run the command “GUI_Main_Menu”. 8. Select “Load Data.” 9. Load “Centrosomes Projection” (e.g., Centrosomes.tiff). 10. Set parameters (e.g., define the pixel and z-step sizes) (see Note 30). 11. Repeat procedure to load additional files containing the nucleus labeling and cell membrane labeling using “Nuclear Membrane Projection” and “Cell Membrane Projection,” respectively. 12. Select “Crop,” adjust window size on the “Centrosome Projection” image and click “Save.” 13. Select “Next.” 14. Select the frames of interest and click “Ok.” 15. Manually locate both centrosomes in the projection and click “Confirm.” 16. Select Filter and then “Next.” 17. Select “Run” (see Note 31). 18. Click “Results” (Fig. 3c–j). 19. Save results and figures (see Note 32). 20. The same procedure can be applied to cells seeded on regular dishes (i.e., without any surface modification) or subjected to micromanipulation or confinement (Fig. 4). 3.5.2 Compile Results

1. Open MATLAB “GUI_Compile_Data”.

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Fig. 4 (a) Time-lapse images of an RPE-1 cell expressing H2B-GFP/tubulin-RFP, dividing without confinement. (b) 3D reconstruction of the RPE-1 cell in (a), showing cell membrane (green), nucleus (yellow), and the two centrosomes (blue and red dots). The vector connecting both centrosomes is shown as a blue line. (c) Timelapse images of an RPE-1 cell expressing H2B-GFP/tubulin-RFP, dividing under confinement. (d) 3D reconstruction of the RPE-1 cell in (c). Time zero corresponds to NEB (nuclear envelope breakdown). Time is in seconds and time-lapse is 20 s. Scale bar corresponds to 10 m. (e) Reconstruction of centrosomes trajectories in 4D. Each point corresponds to a time-point extracted from a live-cell imaging movie. (f) Reconstruction and correlation between centrosomes vector (blue arrows), long cellular axis (large green arrows), and long nuclear axis (large yellow arrows). Each set of arrows corresponds to a time-point extracted from a live-cell imaging movie. Right panel shows a temporal projection of left panel. (g) Correlation of the angles between nucleus and cell membrane (red), centrosomes and cell membrane (green) and centrosomes and nucleus (blue) over time for the cell shown in (c). (h) Cell membrane (blue) and nucleus (black) eccentricity over time for the cell shown in (c). (i) Cell (blue) and nuclear (black) irregularity over time for cell shown in (c). Dashed lines correspond the NEB

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Notes 1. The continuous use of antibiotics in the cell culture medium should be avoided as it may select resistant subclones. When necessary (e.g., imaging the cells), the use of antibiotic–antimycotic solutions (e.g., penicillin–streptomycin) is recommended. 2. A sterile laminar flow hood should be used to perform all cell manipulation procedures. 3. Cells should be used at a low-passage number. 4. It is advised to grow cells for experimental and control conditions in parallel. 5. Prepare solutions A and B for the control condition in parallel. For controls, add 2 μL of Scrambled RNAi or perform a mock transfection. 6. Adjust the concentration according to the oligo and the type of cells used. 7. When the transiently transfected plasmid contains a selection marker, positive cells can be selected by adding the appropriate selection drug to the cell culture medium for 1–2 weeks. 8. When siRNAs are transiently transfected, monitor protein depletion efficiency by collecting cell extracts for Western blot analysis and by fixing cells for phenotypic analysis. 9. For 22  22 mm2 coverslips, use 100 μL of PLL-g-PEG or FBN. For other coverslip sizes, adjust the volume accordingly. 10. PLL-g-PEG-coated glass coverslips can be store dried, at RT, for 3–5 weeks. 11. The micropattern designs on the quartz mask should be customized according to requirements (e.g., size and shape of the micropatterns) (Fig. 1b–b0 ). 12. Use soft tissues (e.g., Kimwipes) to avoid scratching the mask during the cleaning process. Repeat washing steps if necessary. Occasionally, wash the mask with soap and tap water to remove greasy debris. 13. Use plastic or glass spacers to avoid contact of the mask with the bottom of the lamp box. 14. Try to remove all the air bubbles using a plastic tip to press the coverslip against the mask. The presence of air bubbles will enlarge and distort the underlying pattern. 15. Illumination time might need to be adjusted for different cell types. 16. Patterned glass coverslips can be store dried, at RT, for 2–3 weeks.

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17. Never allow the coverslips to dry after FBN incubation. 18. The use of fibrinogen is not mandatory but advised to evaluate micropatterning efficiency and quality. 19. To store the coverslips after FBN incubation (no longer then 1–2 days), incubate the coverslips with PBS at 4
C. The best results are obtained when the FBN incubation step is performed immediately before seeding the cells. 20. It is possible to UV-sterilize the coverslips in the Laminar Flow Hood, at this point. Keep the coverslips in PBS during UV exposure. 21. Optimal seeding density will vary depending on cell size. Typical number of cells per 22  22 mm2 coverslip or 6-well plate well are: U2-OS, approximately 50,000 cells; HeLa and RPE-1, approximately 75,000 cells; HCT116, approximately 150,000 cells; MDA-MB-468, approximately 200,000 cells. 22. Optimal seeding time should be predetermined for each cell line, taking into account its properties. For more adherent cells such as RPE-1 or U2-OS, a 15–20 min incubation is sufficient, whereas HCT116 and MDA-MB-468 require 20–30 min and HeLa cells typically require 30–40 min. 23. Using a small volume of cell culture medium avoids suction of the culture medium into the vacuum line attached to the pressure/vacuum generator. 24. The cell confiner and the cell confinement slides can be bought or microfabricated in the lab, as previously described [31, 32]. There are different devices (e.g., static vs dynamic cell confiner) and types of confinement coverslips (e.g., slides with PDMS pillars or with hydrogels with tunable stiffness) commercially available (e.g., 4DCell, France). Adjust the equipment according to the requirements of your experiment. 25. Leibovitz’s L-15 is a phenol red-free and a CO2-independent medium. The phenol red dye usually used as a pH indicator in the cell culture medium should be avoided. CO2-independent medium should be used when pH cannot be controlled by the standard NaHCO2/CO2-buffering system. 26. Mitosis is tremendously sensitive to temperature; minor temperature changes might induce mitotic delay [40]. It is crucial that the appropriate temperature is set prior to mounting the imaging chamber on the microscope stage. The temperature should be kept stable during the acquisition time. 27. To precisely track centrosome, nucleus, and cell membrane behavior during mitosis, cells should be expressing suitable fluorescent tags (e.g., tubulin, actin, and H2B). Acquisition time will vary according to the temporal resolution required. For high spatiotemporal resolution such as centrosome

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tracking during mitotic entry, cells can be filmed every 20 s, capturing 17–21 z-stacks with a 0.5–0.7 μm step. For lower temporal resolution (e.g., follow a complete mitosis), cells can be filmed every 2–5 min, capturing 10–15 z-stacks with a 0.7–1 μm step. 28. To keep the cells healthy during the acquisition process, cells must be protected from light-induced damage. Excessive illumination often leads to cell cycle arrest at G2, or delayed G2– M transition. Reduce light exposure time and laser intensity to a minimum, just enough to guarantee the appropriate temporal and spatial resolutions required for the study. 29. The Centrosome Dynamics module of Trackosome was designed to track centrosomes in 4D and to correlate their trajectories with cell and nuclear membrane metrics. Therefore, live-cell imaging datasets where the three cellular components of interest are independently labeled are required. 30. When metadata information is available on the 4D dataset, the image parameters (e.g., pixel size, time step, and z step) are automatically loaded. 31. Using the option “Correct Centrosomes Coordinates” in Trackosome, allows the manual editing of the centrosomes coordinates in specific time frames. 32. It is recommended to save both the excel and MATLAB files. MATLAB files can be used to edit or reanalyze the data at later time points. 33. Results include key metrics for centrosome, nucleus, and cell membrane such as intercentrosome distances; angles between long nuclear axis/long cell axis, angles between centrosomes/ long cell axis and centrosomes/long nuclear axis; and nucleus and cell membrane eccentricity and irregularity. References 1. Champion L, Linder MI, Kutay U (2017) Cellular reorganization during mitotic entry. Trends Cell Biol 27:26–41. https://doi.org/ 10.1016/j.tcb.2016.07.004 2. McIntosh JR (2016) Mitosis. Cold Spring Harb Perspect Biol 8. https://doi.org/10. 1101/cshperspect.a023218 3. Cramer LP, Mitchison TJ (1997) Investigation of the mechanism of retraction of the cell margin and rearward flow of nodules during mitotic cell rounding. Mol Biol Cell 8:109– 119. https://doi.org/10.1091/mbc.8.1.109 4. Dao VT, Dupuy AG, Gavet O, Caron E, de Gunzburg J (2009) Dynamic changes in Rap1 activity are required for cell retraction and

spreading during mitosis. J Cell Sci 122:2996–3004. https://doi.org/10.1242/ jcs.041301 5. Marchesi S, Montani F, Deflorian G, D’Antuono R, Cuomo A, Bologna S, Mazzoccoli C, Bonaldi T, Di Fiore PP, Nicassio F (2014) DEPDC1B coordinates de-adhesion events and cell-cycle progression at mitosis. Dev Cell 31:420–433. https://doi.org/10. 1016/j.devcel.2014.09.009 6. Matthews HK, Delabre U, Rohn JL, Guck J, Kunda P, Baum B (2012) Changes in Ect2 localization couple actomyosin-dependent cell shape changes to mitotic progression. Dev Cell 23:371–383. https://doi.org/10.1016/j. devcel.2012.06.003

Centrosome Analysis During Mitosis 7. Stewart MP, Helenius J, Toyoda Y, Ramanathan SP, Muller DJ, Hyman AA (2011) Hydrostatic pressure and the actomyosin cortex drive mitotic cell rounding. Nature 469:226–230. https://doi.org/10.1038/ nature09642 8. Zlotek-Zlotkiewicz E, Monnier S, Cappello G, Le Berre M, Piel M (2015) Optical volume and mass measurements show that mammalian cells swell during mitosis. J Cell Biol 211:765–774. https://doi.org/10.1083/jcb.201505056 9. Kunda P, Pelling AE, Liu T, Baum B (2008) Moesin controls cortical rigidity, cell rounding, and spindle morphogenesis during mitosis. Curr Biol 18:91–101. https://doi.org/10. 1016/j.cub.2007.12.051 10. Lancaster OM, Le Berre M, Dimitracopoulos A, Bonazzi D, ZlotekZlotkiewicz E, Picone R, Duke T, Piel M, Baum B (2013) Mitotic rounding alters cell geometry to ensure efficient bipolar spindle formation. Dev Cell 25:270–283. https://doi. org/10.1016/j.devcel.2013.03.014 11. Mchedlishvili N, Matthews HK, Corrigan A, Baum B (2018) Two-step interphase microtubule disassembly aids spindle morphogenesis. BMC Biol 16(14). https://doi.org/10.1186/ s12915-017-0478-z 12. Nunes V, Dantas M, Castro D, Vitiello E, Wang I, Carpi N, Balland M, Piel M, Aguiar P, Maiato H, Ferreira JG (2020) Centrosome-nuclear axis repositioning drives the assembly of a bipolar spindle scaffold to ensure mitotic fidelity. Mol Biol Cell 2020:mbc.E2001-0047. https://doi.org/10.1091/mbc. E20-01-0047 13. Antonin W, Neumann H (2016) Chromosome condensation and decondensation during mitosis. Curr Opin Cell Biol 40:15–22. https://doi.org/10.1016/j.ceb.2016.01.013 14. Belmont AS (2006) Mitotic chromosome structure and condensation. Curr Opin Cell Biol 18:632–638. https://doi.org/10.1016/ j.ceb.2006.09.007 15. Gu¨ttinger S, Laurell E, Kutay U (2009) Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat Rev Mol Cell Biol 10:178–191. https://doi.org/10. 1038/nrm2641 16. Kutay U, Hetzer MW (2008) Reorganization of the nuclear envelope during open mitosis. Curr Opin Cell Biol 20:669–677. https://doi. org/10.1016/j.ceb.2008.09.010 17. Khodjakov A, Rieder CL (2001) Centrosomes enhance the fidelity of cytokinesis in vertebrates and are required for cell cycle

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progression. J Cell Biol 153:237–242. https://doi.org/10.1083/jcb.153.1.237 18. Whitehead CM, Winkfein RJ, Rattner JB (1996) The relationship of HsEg5 and the actin cytoskeleton to centrosome separation. Cell Motil Cytoskeleton 35:298–308. https://doi.org/10.1002/(SICI)1097-0169( 1996)35:43.0.CO;2-3 19. Bolhy S, Bouhlel I, Dultz E, Nayak T, Zuccolo M, Gatti X, Vallee R, Ellenberg J, Doye V (2011) A Nup133-dependent NPC-anchored network tethers centrosomes to the nuclear envelope in prophase. J Cell Biol 192:855–871. https://doi.org/10. 1083/jcb.201007118 20. Kotak S, Busso C, Go¨nczy P (2013) NuMA phosphorylation by CDK1 couples mitotic progression with cortical dynein function. EMBO J 32:2517–2529. https://doi.org/10. 1038/emboj.2013.172 21. Rosenblatt J, Cramer LP, Baum B, McGee KM (2004) Myosin II-dependent cortical movement is required for centrosome separation and positioning during mitotic spindle assembly. Cell 117:361–372. https://doi.org/10. 1016/s0092-8674(04)00341-1 22. Cao J, Crest J, Fasulo B, Sullivan W (2010) Cortical actin dynamics facilitate early-stage centrosome separation. Curr Biol 20:770– 776. https://doi.org/10.1016/j.cub.2010. 02.060 23. Khodjakov A, Rieder CL (2006) Imaging the division process in living tissue culture cells. Methods San Diego Calif 38:2–16. https:// doi.org/10.1016/j.ymeth.2005.07.007 24. Walther N, Ellenberg J (2018) Quantitative live and super-resolution microscopy of mitotic chromosomes. Methods Cell Biol 145:65–90. https://doi.org/10.1016/bs.mcb.2018.03. 014 25. Shcherbo D, Murphy CS, Ermakova GV, Solovieva EA, Chepurnykh TV, Shcheglov AS, Verkhusha VV, Pletnev VZ, Hazelwood KL, Roche PM, Lukyanov S, Zaraisky AG, Davidson MW, Chudakov DM (2009) Far-red fluorescent tags for protein imaging in living tissues. Biochem J 418:567–574. https://doi. org/10.1042/BJ20081949 26. Douthwright S, Sluder G (2017) Live cell imaging: assessing the phototoxicity of 488 and 546 nm light and methods to alleviate it. J Cell Physiol 232:2461–2468. https://doi. org/10.1002/jcp.25588 27. Magidson V, Khodjakov A (2013) Circumventing photodamage in live-cell microscopy. Methods Cell Biol 114:545–560. https://doi.org/10. 1016/B978-0-12-407761-4.00023-3

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28. Azioune A, Storch M, Bornens M, The´ry M, Piel M (2009) Simple and rapid process for single cell micro-patterning. Lab Chip 9:1640–1642. https://doi.org/10.1039/ b821581m 29. The´ry M, Jime´nez-Dalmaroni A, Racine V, Bornens M, Ju¨licher F (2007) Experimental and theoretical study of mitotic spindle orientation. Nature 447:493–496. https://doi.org/ 10.1038/nature05786 30. Velve-Casquillas G, Le Berre M, Piel M, Tran PT (2010) Microfluidic tools for cell biological research. Nano Today 5:28–47. https://doi. org/10.1016/j.nantod.2009.12.001 31. Le Berre M, Zlotek-Zlotkiewicz E, Bonazzi D, Lautenschlaeger F, Piel M (2014) Methods for two-dimensional cell confinement. Methods Cell Biol 121:213–229. https://doi.org/10. 1016/B978-0-12-800281-0.00014-2 32. Le Berre M, Aubertin J, Piel M (2012) Fine control of nuclear confinement identifies a threshold deformation leading to lamina rupture and induction of specific genes. Integr Biol Quant Biosci Nano Macro 4:1406–1414. https://doi.org/10.1039/c2ib20056b 33. Fink J, Carpi N, Betz T, Be´tard A, Chebah M, Azioune A, Bornens M, Sykes C, Fetler L, Cuvelier D, Piel M (2011) External forces control mitotic spindle positioning. Nat Cell Biol 13:771–778. https://doi.org/10.1038/ ncb2269 34. Harder N, Mora-Bermu´dez F, Godinez WJ, Wu¨nsche A, Eils R, Ellenberg J, Rohr K (2009) Automatic analysis of dividing cells in live cell movies to detect mitotic delays and correlate phenotypes in time. Genome Res 19:2113–2124. https://doi.org/10.1101/gr. 092494.109

35. Neumann B, Walter T, He´riche´ J-K, Bulkescher J, Erfle H, Conrad C, Rogers P, Poser I, Held M, Liebel U, Cetin C, Sieckmann F, Pau G, Kabbe R, Wu¨nsche A, Satagopam V, Schmitz MHA, Chapuis C, Gerlich DW, Schneider R, Eils R, Huber W, Peters J-M, Hyman AA, Durbin R, Pepperkok R, Ellenberg J (2010) Phenotypic profiling of the human genome by time-lapse microscopy reveals cell division genes. Nature 464:721– 727. https://doi.org/10.1038/nature08869 36. Collins E, Mann BJ, Wadsworth P (2014) Eg5 restricts anaphase B spindle elongation in mammalian cells. Cytoskeleton (Hoboken NJ) 71:136–144. https://doi.org/10.1002/cm. 21158 37. De Simone A, Ne´de´lec F, Go¨nczy P (2016) Dynein transmits polarized actomyosin cortical flows to promote centrosome separation. Cell Rep 14:2250–2262. https://doi.org/10. 1016/j.celrep.2016.01.077 38. Boudreau V, Chen R, Edwards A, Sulaimain M, Maddox PS (2019) PP2A-B55/SUR-6 collaborates with the nuclear lamina for centrosome separation during mitotic entry. Mol Biol Cell 30:876–886. https://doi.org/10.1091/mbc. E18-10-0631 39. Castro D, Nunes V, Lima JT, Ferreira JG, Aguiar P (2020) Trackosome: a computational toolbox to study the spatiotemporal dynamics of centrosomes, nuclear envelope and cellular membrane. J Cell Sci 133: jcs252254. https:// doi.org/10.1242/jcs.252254 40. Rieder CL, Maiato H (2004) Stuck in division or passing through: what happens when cells cannot satisfy the spindle assembly checkpoint. Dev Cell 7:637–651. https://doi.org/10. 1016/j.devcel.2004.09.002

Chapter 14 Assaying Cell Cycle Progression via Flow Cytometry in CRISPR/Cas9-Treated Cells Jonathan M. Geisinger and Tim Stearns Abstract CRISPR/Cas9 system is a powerful technique for genome editing and engineering but obtaining a sizeable population of edited cells can be challenging for some cell types. CRISPR/Cas9-induced cell cycle arrest is a possible cause of this barrier to efficient editing; thus, it is desirable to know the cell cycle progression profile of any given cell line or type of interest resulting from CRISPR/Cas9 treatment. Here we describe a flow cytometry-based assay that enables the determination of cell cycle progression in the presence of CRISPR/ Cas9 treatment, in addition to the transfection and expression efficiencies of Cas9 vectors. This assay can also easily determine the effect of various interventions on obtaining a larger pool of Cas9-treated cells. Key words Genome engineering, CRISPR/Cas9, Flow cytometry, Click chemistry, Immunofluorescence, Gene editing, Cell cycle

1

Introduction The ability to edit and engineer the genome of a cell in a defined manner is an immensely powerful tool, but still depends on the capability to isolate the engineered cell of interest. This capability is particularly relevant in the current age of designer nucleases in which zinc-finger nucleases [1], TAL effector nucleases [2], and the CRISPR/Cas9 system [3–5] are all commonly used to generate edited cells in a largely selection-free manner. Thus, the ease of isolating an edited cell derived using these tools is strongly reliant on the proportion of edited cells in the population relative to unedited cells at the time of isolation. Such an issue would not usually be a concern, but several studies have found that the CRISPR/Cas9 system can induce a TP53-dependent cell cycle arrest or delay (or even cell death) in some biologically relevant cell types [6–9]. Another study has reached the conclusion that such an effect may also be dependent on the copy number of the target locus in question [10]. This effect can be quite detrimental to obtaining edited cells without selectable markers, so knowing

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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the degree of arrest for any given cell type or line would be very useful. In the case of generating specific edits or “knockouts,” such a cell cycle arrest can greatly increase the difficulty of obtaining such cells. This effect would be particularly apparent in the absence of selection, resulting in the need, in some cell lines, for a large number of clones to be analyzed to obtain the desired edit. Using some form of selection, either antibiotic selection [11] or a highthroughput, direct identification method, such as fluorescenceactivated cell sorting based on modified surface markers [12], may sufficiently mitigate the effects of CRISPR/Cas9-associated cell cycle arrest for some cell lines. However, knowing the degree of cell cycle arrest is still valuable as some cell lines may prove recalcitrant even with the aid of selection and may require the use of inducible Cas9 vectors, as one study has demonstrated [8]. In considering the generation and isolation of knockout lines in the presence of CRISPR/Cas9-associated cell cycle arrest, one should bear in mind that, in the absence of specific selection for the desired allele, it may be very difficult to obtain knockouts. This difficulty may be due to the arrest selecting against Cas9-treated cells, in addition to the known effects of some Cas9-generated mutations resulting in still-functioning proteins [13, 14]. Depending on the severity of the arrest, one may also unintentionally select for clones that bear additional mutations that suppress the arrest phenotype. Such additional mutations may not be desirable due to the potential confounding of phenotypic analysis. Knowledge that cell cycle arrest is a potential problem is also of particular importance for the proper design and interpretation of genome-wide CRISPR-Cas9 screens. Depending on the chosen cell line, the recovered candidates may be biased toward having mutations in TP53 and other cell cycle control genes, as observed in one relatively recent screen [6]. Performing such a screen using a wild-type Cas9 as opposed to a CRISPR-inactivation or activation screen using a dead Cas9 may lead to confounding of the results by selecting for such mutations. Furthermore, the existence of such an arrest calls for a more judicious interpretation of identifying “essential” genes from such screens (discussed in [15, 16]). This situation is similar to how such mutations can arise and overtake induced pluripotent stem cell cultures over time [17]. To address these issues, we present here a simple, flow cytometric-based assay utilizing the thymidine analog EdU, which is visualized via click chemistry [9, 18] to quantify S-phase progression of CRISPR/Cas9-treated cells. This assay is easily applied to a wide-range of cell types and can be quickly used to determine the effect of various interventions to reduce CRISPR/Cas9-associated cell cycle arrest.

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Materials Prepare all solutions using deionized water and analytical grade reagents, unless otherwise indicated.

2.1 Cell Culture and Transfection

1. Growth media: Dulbecco’s modified Eagle’s medium/Ham’s F-12 50/50 mix (DMEM/F-12) with L-glutamine supplemented with 10% Cosmic Calf Serum (HyClone Laboratories, Logan, UT). 2. Phosphate-buffered saline (PBS), pH 7.4. 3. 1 mg/mL polyethylenimine (PEI), pH 7.0 (see Note 1). 4. HyQ DMEM-RS or Opti-MEM I reduced serum medium. 5. 0.05% trypsin–EDTA. 6. An SpCas9 expression plasmid, preferably also encoding an sgRNA of interest and a separate expression cassette encoding GFP, Clover, mClover3, or any other GFP-derived fluorescent protein (see Note 2). 7. A GFP-derived fluorescent protein expression plasmid. 8. 10 cm tissue culture dishes. 9. 24-Well tissue culture plates. 10. 5-Ethynyl-20 -deoxyuridine (EdU) at 10 mM in DMSO. 11. RPE-1 hTERT cells (see Note 3). 12. CO2 incubator set to 37  C. 13. Hemocytometer. 14. Tissue culture laminar flow hood. 15. 15 mL conical tubes.

2.2 Flow Cytometry Assay

1. Primary antibodies: goat IgG anti-GFP; (Rockland, Immunochemicals, Limerick, PA), mouse IgG1 anti-Cas9 (BioLegend, San Diego, CA) (see Note 4). 2. Secondary antibodies: donkey anti-goat IgG Alexa Fluor 488, donkey anti-mouse IgG Alexa Fluor 680. 3. 5 mg/mL 40 ,6-diamidino-2-phenylindole (DAPI). 4. 4% formaldehyde in PBS. 5. PBS-BT: PBS, pH 7.4, 5% bovine serum albumin, 0.2% Tween 20. 6. PBS-T: PBS, pH 7.4, 0.2% Tween 20. 7. 10 mM Alexa Fluor 594 azide in DMSO. 8. Tris-buffered saline (TBS), pH 7.4. 9. 100 mM Cu(II)SO4. 10. 100 mM ascorbic acid.

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11. 1.75 mL microcentrifuge tubes. 12. 5 mL round-bottom polystyrene tubes. 13. Microcentrifuge. 14. Flow cytometer equipped with (a) 405 nm violet, 488 nm blue, 532 nm green, and 640 nm red lasers, (b) 505, 600, and 685 nm LP splitters, and (c) 450 50 nm, 525 50 nm, 610 20 nm, and 710 50 nm BP filters.

3

Methods All procedures should be carried out at room temperature, unless otherwise specified.

3.1 Transfection and EdU Treatment of Cells

1. Grow RPE-1 hTERT cells in growth media in 10 cm tissue culture plates in a CO2 incubator set to 37  C until they are approximately 80–90% confluent. 2. Aspirate medium and wash with PBS once before adding 2 mL of 0.05% trypsin–EDTA per plate. 3. Incubate the cells in a CO2 incubator set to 37  C for 10 min. 4. Add 8 mL of growth medium to each plate and wash the plate with the cell suspension twice to remove cells from the plate. Transfer suspension to a 15 mL conical tube. 5. Calculate the concentration of cells using 10 μL of the suspension and a hemocytometer, or alternative cell counting method. 6. Plate 50,000 cells per well in a 24-well plate in 0.5 mL of growth medium. For each intended transfection, as well as for the negative control, plate three wells. Also, plate an additional three wells each for negative control, fluorescent protein control, and Cas9 control to serve as staining controls for flow cytometry. 7. Incubate the plated cells in a CO2 incubator set to 37  C for 24 h. 8. After 24 h, prepare the transfection reactions in the flow hood in 1.5 mL microcentrifuge tubes. Each reaction volume is 50 μL consisting of 1.5 μL 1 mg/mL PEI solution and 500 ng of either the fluorescent protein plasmid or the SpCas9 plasmid with the remaining volume consisting of DMEM-RS. Prepare a total number of reaction volumes equal to the number of wells (18 total volumes across three conditions in this case). 9. Briefly vortex each reaction mixture tube. 10. Incubate the transfection reactions for 30 min.

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11. Aspirate medium from the cells in the 24-well plate and replace with fresh medium. Remove 50 μL of medium from each well. 12. Add 50 μL of each appropriate reaction mixture dropwise to each appropriate well and briefly shake the plate in a crosspattern to mix. 13. Incubate plate in a CO2 incubator set to 37  C for 24 h. 14. After 24 h, aspirate medium and replace with fresh growth medium supplemented with EdU to a final concentration of 10 μM for all wells except for two negative control wells. Replace in the medium in one well with growth medium and replace the medium in the other with growth medium supplemented with a volume of DMSO equivalent to one well’s volume of EdU. 15. After a further 24 h, repeat step 14. 3.2 Click Chemistry Labeling Flow Cytometry-Based Cell Cycle Progression Assay

1. 24 h after the previous step, aspirate medium and wash cells with PBS before adding 200 μL of 0.05% trypsin–EDTA to each well. 2. Place the plate in a CO2 incubator set to 37  C for 10 min. 3. In a tissue culture flow hood, add 800 μL of growth medium to each well. 4. Using a P1000 micropipette, transfer each well’s cell suspension to a separate 1.75 mL microcentrifuge tube. 5. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g. 6. Carefully aspirate supernatants and resuspend each cell pellet in 1 mL of cold PBS. Repeat step 5. 7. Aspirate supernatants and resuspend each cell pellet in 200 μL of 4% formaldehyde. Incubate cells at room temperature for 10 min. 8. Add 800 μL of cold PBS to each tube and centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g. 9. Aspirate supernatant and resuspend each cell pellet in 200 μL of cold PBS. Incubate cells at room temperature for 10 min. 10. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g. 11. Aspirate supernatant and resuspend each cell pellet in 50 μL of PBS-BT. Incubate cells at room temperature for 30 min. 12. Prepare the azide labeling solution in a 1.75 mL microcentrifuge tube using the following components (volume per 50 μL reaction): 43 μL TBS, 2 μL 100 mM Cu(II)SO4, 5 μL 100 mM ascorbic acid, 0.025 μL 10 mM Alexa Fluor 594 azide. Vortex

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Table 1 Suggested control tubes for accurate analysis Replicate #1

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WT-Cas9 and fluorescent protein

– No click chemistry – Primary antibodies – Alexa Fluor 680

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to mix and briefly centrifuge. Protect solution from light until ready to use. 13. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 14. Resuspend one sample from each of the additional negative, Clover-transfected, and Cas9-transfected wells from step 6 of Subheading 3.1 in 50 μL PBS-BT. 15. Resuspend all other cell pellets in 50 μL of azide labeling solution. 16. Incubate the tubes for 30 min at room temperature protected from light (see Note 5). 17. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 18. Resuspend cell pellet in 300 μL PBS-T, and then centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 19. Repeat step 18. 20. Prepare the primary antibody staining cocktail in a 1.75 mL microcentrifuge tube using 50 μL PBS-BT per reaction volume and 1:500 dilutions of both goat IgG anti-GFP and mouse IgG1 anti-Cas9. See Table 1 for details on the controls. Vortex cocktails and centrifuge briefly. 21. Resuspend cell pellets in 50 μL of the appropriate antibody cocktail. Incubate for 30 min at room temperature protected from light.

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22. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 23. Resuspend cell pellet in 300 μL PBS-T, and then centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 24. Repeat step 23. 25. Prepare the secondary antibody staining cocktail in a 1.75 mL microcentrifuge tube using 50 μL PBS-BT per reaction volume and 1:1000 dilutions of both donkey anti-goat IgG Alexa Fluor 488 and donkey anti-mouse IgG Alexa Fluor 680 and DAPI at a final concentration of 3 μg/mL. See Table 1 for setting up the controls. Vortex cocktails and centrifuge briefly. 26. Resuspend cell pellets in 50 μL of the appropriate antibody cocktail. Incubate for 30 min at room temperature protected from light. 27. Centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 28. Resuspend cell pellet in 300 μL PBS-T, and then centrifuge tubes in a benchtop microcentrifuge for 5 min at 300  g and then aspirate supernatant. 29. Repeat step 28. 30. Resuspend each cell pellet in 200 μL PBS and transfer each suspension to a 5 mL round-bottom polystyrene tube. Protect tubes from light (see Note 6). 3.3

Flow Cytometry

Collect 10,000 total events for each sample and control (see Note 7). All analysis is assumed to be done afterward in flow cytometry analysis software of choice. 1. Carry out any required start-up and calibration operations relevant to your flow cytometer (see Note 8). 2. Collect data from the completely unstained control first, using it to both bring the cells on scale for forward and side scatter, as well as to set the initial gates for transfected, EdU, and Cas9 staining. The order of gating is Scatter!Singlets!Transfected +![Cas9 EdU], [Cas9 EdU+], [Cas9+ EdU], and [Cas9+ EdU+]. 3. Collect data from the remaining controls, proceeding in order from negative to fluorescent-protein-alone to Cas9-and-fluorescent-protein. 4. After collecting data from each of the controls, collect data from each from each of the samples. 5. Export the data in accordance with local procedure and carry out any required maintenance and shutdown procedures.

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Fig. 1 An example of the gating strategy on fully stained cells. Cells depicted are RPE-1 cells. Arrows indicate which population is further analyzed

6. Analyze the data using flow cytometry analysis of your choice (see Note 9). The initial gates should be set in the following order using the completely unstained control sample: Scatter!Singlets!Transfected+![Cas9 EdU], [Cas9 EdU +], [Cas9+ EdU], and [Cas9+ EdU+] (see Note 10 and Fig. 1 for an example of gating). 7. Adjust the gates as necessary using the other control samples to adjust for residual Clover fluorescence and nonspecific binding of the secondary antibodies. 8. Apply the corrected gates to all actual samples and extract the cell numbers and percentages of parental population statistics for the Transfection+ and all four [Cas9 EdU] subpopulation gates for each sample. 9. In statistical analysis software, normalize the [Cas9 EdU] subpopulations’ percentages to the sum of the four subpopulations (see Note 11).

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Notes 1. We used PEI transfection here, but any standard transfection protocol known to work for the cells in question would be acceptable. 2. The separate fluorescent protein expression cassette aids in efficient identification of transfected cells. However, an IRES or 2A element connecting the Cas9 and fluorescent protein reading frames may work as a substitute. 3. Any cell line is suitable for this assay. If using a cell line other than RPE-1, use appropriate medium and growth conditions. 4. Although we have had success with these particular antibodies, it is likely that other similar antibodies would work equally well. 5. While the incubated cells for both click chemistry and immunostaining could be placed on a nutating mixer, we have found

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that nutation leads to additional cell loss. Thus, we recommend benchtop incubation. 6. As long as the tubes are protected from light and stored at 4  C, flow cytometry can be carried out up to 3 days later. However, we have observed some swelling of the cells, which only affects the initial scatter gate dimensions. 7. We have compared data between collecting on the initial scatter gate versus collecting on the transfected/fluorescent-protein+ gate and have observed no difference. Collecting on the transfected+ gate will obtain more events, but correctly using the controls to draw the gates, especially the [Cas9 EdU] gates has a greater impact on data analysis. 8. We have performed this assay using a BD Biosciences LSR II flow cytometer, but any make and model of flow cytometer is compatible with this assay as long as it has the detectors, filters, and lasers outlined in Subheading 2. 9. Our software of choice for analysis is FlowJo (Becton, Dickinson and Company, Ashland, OR), but any flow cytometry analysis software will be sufficient. 10. The use of the control samples detailed in Table 1 are instrumental in setting accurate gates, especially for the [Cas9 EdU] subpopulations. Of particular importance is the untreated replicate #2 control (Table 1), as this control defines the nonspecific fluorescence of the secondary antibodies. 11. We recommend this normalization step because, in our experience FlowJo does not permit enough flexibility with its Quadrants and Curly Quadrants tool to ensure accurate gating. Because of this, we have chosen to err on the side of caution with our gates, resulting in a small number of cells not being counted. To us, this small loss of cells between gates is more acceptable than double-counting cells. If your software of choice enables more flexibility in setting quadrants via a tool, quadrants can be used in place of drawing freehand gates. References 1. Kim YG, Cha J, Chandrasegaran S (1996) Hybrid restriction enzymes: zinc finger fusions to Fok I cleavage domain. Proc Natl Acad Sci 93:1156–1160 2. Christian M, Cermak T, Doyle EL, Schmidt C, Zhang F, Hummel A, Bogdanove AJ, Voytas DF (2010) Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186:757–761 3. Cong L, Ran FA, Cox D, Lin S, Barretto R, Habib N, Hsu PD, Wu X, Jiang W, Marraffini LA et al (2013) Multiplex genome engineering

using CRISPR/Cas systems. Science 339:819–823 4. Mali P, Yang L, Esvelt KM, Aach J, Guell M, DiCarlo JE, Norville JE, Church GM (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826 5. Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337:816–821

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6. Haapaniemi E, Botla S, Persson J, Schmierer B, Taipale J (2018) CRISPR-Cas9 genome editing induces a p53-mediated DNA damage response. Nat Med 24:927–930 7. Ihry RJ, Worringer KA, Salick MR, Frias E, Ho D, Theriault K, Kommineni S, Chen J, Sondey M, Ye C et al (2018) p53 inhibits CRISPR–Cas9 engineering in human pluripotent stem cells. Nat Med 24:939–946 8. van den Berg J, Manjo´n GA, Kielbassa K, Feringa FM, Freire R, Medema RH (2018) A limited number of double-strand DNA breaks is sufficient to delay cell cycle progression. Nucleic Acids Res. https://doi.org/10.1093/ nar/gky786 9. Geisinger JM, Stearns T (2020) CRISPR/ Cas9 treatment causes extended TP-53 dependent cell cycle arrest in human cells. Nucleic Acids Res 48(16):9067–9081 10. Aguirre AJ, Meyers RM, Weir BA, Vazquez F, Zhang C-Z, Ben-David U, Cook A, Ha G, Harrington WF, Doshi MB et al (2016) Genomic copy number dictates a gene-independent cell response to CRISPR/Cas9 targeting. Cancer Discov 6:914–929 11. Kaulich M, Lee YJ, Lo¨nn P, Springer AD, Meade BR, Dowdy SF (2015) Efficient CRISPR-rAAV engineering of endogenous genes to study protein function by allelespecific RNAi. Nucleic Acids Res 43:e45–e45 12. Byrne SM, Ortiz L, Mali P, Aach J, Church GM (2015) Multi-kilobase homozygous targeted

gene replacement in human induced pluripotent stem cells. Nucleic Acids Res 43:e21–e21 13. Makino S, Fukumura R, Gondo Y (2016) Illegitimate translation causes unexpected gene expression from on-target out-of-frame alleles created by CRISPR-Cas9. Sci Rep 6:39608 14. Rodriguez-Rodriguez J-A, Lewis C, McKinley KL, Sikirzhytski V, Corona J, Maciejowski J, Khodjakov A, Cheeseman IM, Jallepalli PV (2018) Distinct roles of RZZ and Bub1KNL1 in mitotic checkpoint signaling and kinetochore expansion. Curr Biol 28:3422–3429.e5 15. Brown KR, Mair B, Soste M, Moffat J (2019) CRISPR screens are feasible in TP53 wild-type cells. Mol Syst Biol 15(8):e8679 16. Haapaniemi E, Botla S, Persson J, Schmierer B, Taipale J (2019) Reply to “CRISPR screens are feasible in TP53 wild-type cells”. Mol Syst Biol 15(8):e9059 17. Merkle FT, Ghosh S, Kamitaki N, Mitchell J, Avior Y, Mello C, Kashin S, Mekhoubad S, Ilic D, Charlton M et al (2017) Human pluripotent stem cells recurrently acquire and expand dominant negative P53 mutations. Nature 545:229–233 18. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci 105:2415–2420

Chapter 15 Use of Mitotic Protein Kinase Inhibitors and Phospho-Specific Antibodies to Monitor Protein Phosphorylation During the Cell Cycle Isha Nasa, Greg B. Moorhead, and Arminja N. Kettenbach Abstract Reversible protein phosphorylation regulates the transitions between different phases of the cell cycle ensuring proper segregation of the duplicated genome into two daughter cells. Protein kinases and protein phosphatases establish the appropriate phosphorylation stoichiometries in diverse substrates maintaining genomic stability as a cell undergoes this complex process. Along with regulating common substrates, these opposing enzymes regulate one another by fine-tuning each other’s activity both spatially and temporally throughout mitosis. Protein phosphatase catalytic subunits work together with regulatory proteins, which control their localization, activity, and specificity. Protein phosphatase 1 (PP1) recognizes its regulatory proteins via a short linear interaction motif (SLIM) called the “RVxF” motif. A subset of proteins with these “RVxF” motifs contain a phosphorylatable amino acid (S/T) at the ‘x’ position. Here, we describe methods to generate, affinity purify and utilize phospho-specific antibodies to monitor phosphorylation sites during the cell cycle and the appropriate use of mitotic kinase inhibitors. More specifically, we employ phospho-specific antibodies, which recognize phosphorylated RVp[S/T]F motifcontaining proteins, to monitor the phosphorylation status of these motifs throughout the cell cycle. Furthermore, we use mitotic kinase inhibitors to examine the effect of kinase inhibition on the phosphorylation status of multiple RV[S/T]F motifs using these phospho-specific antibodies. Key words Kinase inhibitors, Cell cycle, Phospho-specific antibodies, Aurora A, Aurora B, CDK1, Polo-like kinase, Protein phosphatase one

1

Introduction The cell division cycle is a highly regulated and complex process that results in the generation of two genetically identical daughter cells from one precursor cell. The transitions into and out of mitosis are controlled by posttranslational modifications of proteins, including reversible protein phosphorylation, which ensures proper spatial and temporal order of mitotic events to maintain high fidelity throughout the process.

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Mitosis involves remarkable cellular and structural reorganization, including nuclear envelope breakdown, chromosome condensation, chromosome segregation, and division of two sister chromatids into two new daughter cells. The initiation of mitosis is accompanied by the activation of several mitotic protein kinases including the mitotic master kinase Cdk1 in complex with cyclin B1, Aurora kinases (AURKA and AURKB), and Polo-like kinase 1 (PLK1), which phosphorylate thousands of proteins at serine and threonine residues [1–4]. Mitotic exit requires the resetting of the phosphorylation level to interphase levels and is mediated by protein phosphatases, most of which belong to the phosphoprotein phosphatase (PPP) family of serine/threonine phosphatases [5–7]. Serine/threonine protein phosphatases from the PPP family work either as heterodimers consisting of a catalytic subunit and a regulatory subunit (e.g., Protein Phosphatase 1- PP1), or as heterotrimers with a catalytic, a regulatory and a scaffolding subunit (e.g., Protein Phosphatase 2A-PP2A). The interactions between the phosphatase catalytic subunits and regulatory proteins are often mediated by Short Linear Interaction Motifs (SLIMs) [7– 10]. In the case of PP1, the catalytic subunit has a binding pocket that recognizes and binds to a degenerate “RVxF” motif ([K/R] [K/R][V/I]{FIMYDP}[F/W], braced residues are excluded), which is present in about 90% of all validated PP1-interacting proteins [11]. Thus, the “RVxF” SLIM acts as a PP1-recruiting motif that promotes its holoenzyme assembly, substrate specificity, and proper cellular localization. Protein kinases and protein phosphatases work antagonistically to achieve the proper phosphorylation stoichiometry on a cellular level [12]. However, this antagonism does not act as a binary switch. Protein phosphorylation and protein dephosphorylation are coordinated in time and space to achieve a defined response for each protein that fine-tunes its phosphorylation status throughout the cell cycle (Fig. 1). One approach to study the antagonistic relationship between protein kinases and protein phosphatases is to monitor the phosphorylation stoichiometry of common substrates using phospho-specific antibodies. In addition, protein kinases and phosphatases can not only regulate common substrates but can also regulate each other’s activity through the phosphorylation/dephosphorylation of SLIM motifs. A subset of PP1-interacting proteins contain a phosphorylatable amino acid (serine/threonine) at the “x” position within the “RVxF” motif, that is, a RV[S/T]F motif. Phosphorylation at the serine/threonine residue within the RV[S/T]F motif disrupts the binding of the PP1 catalytic subunit to regulatory proteins, thereby reducing PP1 activity [13–16]. The phosphorylation of these RV[S/T]F motifs can be monitored using

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Nucleus/cytoplasm/spindle CDK1-cyclin B Centrosome Aurora A/PLK1 PP4/PP1-CEP192 PP1/NEK2

Centromeres Aurora B/Haspin PP2A-B56

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Fig. 1 Regulation of the antagonistic activities of protein kinases and protein phosphatases during cell cycle. The active pool of protein kinases (indicated in pink) localized to the same cellular structures as the active protein phosphatases (indicated at blue) dynamically regulate each other in a spatially controlled manner. As a cell undergoes mitosis, there is a spike in phosphorylation stoichiometry on most of the mitotic proteins due to the activation of multiple mitotic kinases including CDK1-cyclin B, Aurora kinases A and B, PLK1, MPS1, Greatwall, and Haspin. Specific holoenzyme complexes of phosphatase catalytic proteins with their regulatory proteins (including PP1, PP2A-B56, and PP2A-B55) as indicated, target the dephosphorylation of mitotic substrates thereby fine-tuning the regulation of reversible phosphorylation during cell cycle

phospho-specific antibodies that recognize these RV[S/T]F motifs in the human proteome [13]. Furthermore, when used in combination with specific kinase inhibitors, phospho-specific antibodies can act as valuable tools to decipher the effect of protein kinases on phosphorylation stoichiometry, particularly in the context of mitosis. In this protocol, we describe the use of p-RV[S/T]F phospho-specific antibodies to monitor the phosphorylation of diverse RV[S/T]F motifs during the cell cycle and upon inhibition of multiple mitotic protein kinases. The combined use of phospho-specific antibodies and mitotic kinase inhibitors enables monitoring of specific phosphorylation sites while simultaneously providing information on whether certain protein kinases control these events.

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Materials

2.1 Generation and Purification of Phospho-Specific Antibodies

1. Unless stated otherwise, Milli-Q grade water is used throughout. 2. Phosphorylated and nonphosphorylated version of peptides (at >98% purity) against which antibody is to be raised (25 mg). 3. Keyhole limpet hemocyanin (KLH) (Imject Maleimideactivated KLH kit, Pierce). Follow kit instructions. 4. Glutaraldehyde Solution: 25% in water (see Note 1). 5. 0.5 M sodium phosphate buffer pH 7.5. 6. CNBr-activated Sepharose. 7. Coupling buffer: 0.1 M NaHCO3 with 0.5 M NaCl, pH 8.0. 8. Column wash buffers: 10 mM Tris–HCl, pH 7.5 with 500 mM NaCl (high salt) and 10 mM Tris–HCl, pH 7.5 (no salt). 9. Elution buffer: 100 mM glycine, pH 2. 10. 12–14 kDa cutoff dialysis membrane 11. 10 kDa cutoff Amicon Centrifugal Filter Units (EMD Millipore). 12. Nitrocellulose membrane. 13. TBST: 150 mM NaCl, 20 mM Tris–HCl pH 7.5, and 0.2% Tween 20. 14. Blocking buffer: 5% milk (w/v) in 1
TBST. 15. 1 M Tris–HCl, pH 7.5. 16. 1 mM HCl. 17. 0.1 M Tris–HCl, pH 8.0. 18. 0.2 M Tris–HCl, pH 8.0 19. Protein quantitation reagent: Bradford reagent (or other, e.g., BCA).

2.2 Cell Culture Reagents and Drugs

1. HeLa cells (American Type Culture Collection, Manassas, VA, USA). 2. Cell culture media: Dulbecco’s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FBS) and penicillin–streptomycin (100 U/mL and 100 μg/mL, respectively). 3. Phosphate buffered saline (PBS), sterile tissue-culture grade; store at RT. 4. Dimethyl sulfoxide (DMSO); store at RT (see Note 2). 5. Trypsin–EDTA solution: 0.25% trypsin containing 1 mM EDTA; store at 20  C (see Note 3).

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6. Thymidine stock solution: 100 mM in water; sterile filter and store in aliquots at 20  C (see Note 4). 7. 5 mg/mL nocodazole solution in DMSO. Store at 20  C (see Note 5). 8. 20 mM MG132 stock solution in DMSO. Store at 20  C (see Note 6). 9. 5 mM MLN8054 solution in DMSO. Store at 20  C. 10. 5 mM ZM447439 solution in DMSO. Store at 20  C. 11. 0.1 mM hesperadin solution in DMSO. Store at 20  C. 12. 0.1 mM BI2536 solution in DMSO. Store at 20  C. 13. 2 mM flavopiridol solution in DMSO. Store at 20  C. 14. 100 mM Thymidine solution in water. Store at 20  C. 2.3 Cell Lysis, SDS-PAGE and Immunoblotting

1. Lysis buffer: 150 mM NaCl, 50 mM Tris–HCl pH 7.5 and 0.5% Triton X-100 (see Note 7). 2. Protease inhibitors: Complete EDTA-free protease inhibitor tablets (Roche). 3. Phosphatase inhibitor stock solutions: 0.5 M β-glycerophosphate pH 7.0; 0.5 M NaF; 1 mM microcystinLR (MC-LR) dissolved in 95% EtOH. Unless otherwise stated, use at the following concentrations: 5 mM β-glycerophosphate; 50 mM NaF; and 0.5 μM microcystin-LR (MC-LR). 4. SDS-PAGE solutions: 30% acrylamide–bis-acrylamide solution 37.5:1; 1.5 M Tris–HCl pH 8.8; 10% SDS; 0.5 M Tris–HCl pH 6.8; 10% ammonium persulfate (APS); N-,N-,N0 -,N0 -tetramethylethylenediamine (TEMED). 5. 10
SDS-PAGE running buffer stock: 250 mM Tris, 1.92 M glycine, and 10% SDS (w/v); dilute to 1
before use with distilled water. 6. 2
Laemmli buffer: 40 mL SDS (10%), 20 mL glycerol, 12 mL Tris–HCl pH 6.8, 28 mL water, and 2 mL bromophenol blue. 7. Transfer set-up: Nitrocellulose membrane; Whatman 3MM chromatography paper. 8. 10
Transfer buffer stock: 250 mM Tris, 1.92 M glycine; dilute to 1
including 200 mL methanol per 1 L before use. 9. TBS: 150 mM NaCl, 20 mM Tris–HCl pH 7.5. 10. TBST: 150 mM NaCl, 20 mM Tris–HCl pH 7.5, and 0.2% Tween 20. 11. Blocking buffer: 5% milk (w/v) in 1
TBST. 12. Primary antibodies: In-house generated phospho-specific p-RV[S/T]F antibody, commercially available mitotic markers, e.g., phospho-Histone 3 Ser-10 and phospho-PP1 Thr-320 and

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loading control antibodies, e.g., α-tubulin can be used. All antibodies are diluted in blocking buffer with the addition of phosphatase-inhibitors (5 mM β-glycerophosphate; 50 mM NaF; and 0.5 μM microcystin-LR (MC-LR) for phospho-specific antibodies. All antibodies, sources, and dilutions are listed in Table 2. 13. Secondary antibodies: Commercially available HRP-conjugated goat anti-rabbit and goat anti-mouse diluted in blocking buffer. 14. 95% Ethanol.

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3.1 Generation, Purification, and Testing of Phospho-Specific Antibodies 3.1.1 Generation of Phospho-Specific Antibodies

A schematic workflow of this section is shown in Fig. 2.

1. Synthetic phospho-peptides (>98% purity) are used for the generation of phospho-specific antibodies (see Note 8). In this protocol, we use the phosphorylated version of the RV [S/T]F containing peptide ‘KRRVpSFADK’ to raise a phospho-specific antibody against this antigen. 2. Couple the phospho-peptide to carrier protein KLH (see Note 9). For coupling, aim at 4 mg peptide per injection. To get a good antibody titer, we use one primary injection with 3 booster injections (total injections—four), and therefore, we couple a total of 16 mg peptide. For coupling reaction, dissolve the phospho-peptide in 1 mL PBS and add 370 μL 0.2 M sodium phosphate buffer pH 7.5 and 1 mL KLH (10 mg/mL) solution. This is followed by very slow addition of 100 μL of 2.5% (1:10 (v/v) dilution of stock in water) glutaraldehyde in a fume hood. 3. Let the reaction take place for several hours at room temperature in the fume hood with constant slow stirring. The reaction mixture turns yellow. 4. After the reaction is complete, add 150 μL of 1 M Tris–HCl pH 7.5 to block the excess glutaraldehyde. 5. Dialyze the conjugated peptide against PBS overnight at 4  C. 6. Aliquot the dialyzed peptide into four tubes for four injections and freeze at 80  C until ready to be injected in a rabbit. 7. Follow the injection schedule as depicted in Fig. 2. 8. The antibody titer reaches a maximum around 12–14 days after each booster injection. Obtain crude serum 12–14 days after each

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Fig. 2 Approach used for generation and purification of phospho-specific antibodies. Peptides containing the desired phosphorylated residue are synthesized and coupled to carrier protein KLH to be injected into a rabbit in the Stage 1 of this process. For the successful generation of anti-peptide antibodies, immunization schedule as depicted in Stage 2 is followed to illicit the desired immune response. Serum from the immunized rabbit is collected at Day 96 after the primary injection. Stage 3 consists of affinity purification and validation of phospho-specific antibody. CNBr-activated Sepharose beads coupled to the phospho-peptide are used to specifically enrich for the desired phospho-specific antibodies from the antibody mix in rabbit sera. This is followed by elution of the phospho-specific antibodies from the matrix using low pH (pH 2.0) glycine. The purified phospho-specific antibody is then tested and validated using dot blots with phosphorylated and nonphosphorylated versions of the peptide used as the antigen

injection and test for antibody titer. To test for antibody titer, perform dot blots using phosphorylated and non-phosphorylated peptides as described in Subheading 3.1.4. Wait additional 14 days after serum collection for the next booster (maximum of 28 days wait between booster injections). 3.1.2 Preparing Phosphopeptide Affinity Column

1. Swell 0.5 g CNBr-activated Sepharose beads in 200 mL of 1 mM ice-cold HCl for 15 min. 2. Wash the resin with coupling buffer (0.1 M NaHCO3 with 0.5 M NaCl, pH 8.0). Spin down the beads in a centrifuge at

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2000
g for 1 min. Let the beads settle and remove the wash solution. Repeat the wash step twice. 3. To prepare 1 mL packed affinity column resin, resuspend 2 mg phospho-peptide in 5 mL coupling buffer with phosphatase inhibitor (50 mM NaF) and add 1 mL washed activated CNBrSepharose beads from step 2. Couple end over end for 4 h at room temperature or overnight at 4  C. 4. Block with 0.1 M Tris–HCl, pH 8.0 for 1 h end over end at room temperature. 5. Stop the reaction overnight by incubating beads in 0.2 M Tris– HCl, pH 8.0 at 4  C. 3.1.3 Purification of Phospho-Specific Antibodies

1. Dilute the serum obtained from rabbit 1:10 (v/v) with 10 mM Tris–HCl, pH 7.5, and load onto the phosphopeptide coupled Sepharose beads from Subheading 3.1.2. Let them incubate end over end for 4 h at 4  C. 2. Wash the column with at least 100 column volumes (CVs) of high salt wash buffer (10 mM Tris–HCl, pH 7.5 and 500 mM NaCl) with phosphatase inhibitor (50 mM NaF), followed by 100 CVs of no salt wash buffer (10 mM Tris–HCl, pH 7.5) with phosphatase inhibitor (50 mM NaF). 3. Elute the antibody from the column using 5 CVs of elution buffer (100 mM glycine, pH 2) in a tube containing 1 CV of 1 M Tris–HCl, pH 8, to immediately neutralize the pH of the eluate. Check the pH of the eluate and use additional Tris– HCl, pH 8 if needed, to bring the pH of the eluate close to 7.5. 4. Dialyze the antibody against PBS overnight and concentrate the antibody next day using 10 kDa cut-off Amicon Centrifugal Filter Units. The total protein (antibody concentration) can be estimated using any protein quantification assay like Bradford or BCA with BSA as standard. 5. Test the purified phospho-specific antibodies for purity and specificity using dot blots (see Subheading 3.1.4) and Western blots (see Subheadings 3.2.6 and 3.2.7).

3.1.4 Validation of Purified Phospho-Specific Antibodies Using Dot Blots

1. Draw a grid using a pencil on a nitrocellulose membrane, indicating the region to be blotted. 2. Spot on the membrane different amounts of phosphorylated and nonphosphorylated versions of the peptide used to raise the phospho-specific antibody ranging from 1000 to 1 ng. If available, a random, but unrelated, phosphorylated and nonphosphorylated peptide can be used as a negative control. 3. Let the membrane dry before blocking the membrane with blocking buffer with phosphatase inhibitor (50 mM NaF) for 1 h rocking at room temperature.

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4. Incubate the membrane with purified phospho-specific antibody (1–2 μg/mL) diluted in blocking buffer with phosphatase inhibitor (50 mM NaF) for 1 h at room temperature. Phospho-specific antibody preincubated with nonphosphorylated peptide (5 μg/mL) to quench epitopes against nonphosphorylated peptide and phospho-specific antibody preincubated with phosphorylated peptide (5 μg/mL) to quench epitopes against phosphorylated peptide can be used as controls to determine the phospho-specificity of the antibody as depicted in Fig. 2. 5. Wash the membrane three times with TBST for 5 min each, rocking at room temperature. 6. Incubate the membrane with goat anti-rabbit secondary antibody conjugated with HRP for 30 min at room temperature. 7. Wash the membrane three times with TBST for 5 min each, rocking at room temperature. 8. Rinse the membrane with TBS and develop using ECL on a ChemiDoc or using an X-ray film. An example dot-blot to validate p-RV[S/T]F antibody is shown in Fig. 2. 3.2 Cell-Cycle Synchronization, Kinase Inhibition, and Immunoblotting

A schematic workflow of this section is shown in Fig. 3.

3.2.1 Maintaining Cell Lines

Grow adherent HeLa cells at 37  C in a humidified incubator with 5% CO2 in cell culture media.

3.2.2 Mitotic Cell Synchronization

1. Freshly seed HeLa cells in 10 cm diameter tissue culture dishes to achieve a confluency of 50–60% the next day (around 5
106 cells per dish). Also seed cells for the asynchronous control sample. 2. Synchronize cells in G1/S phase of cell cycle by the addition of 2 mM thymidine for 16–24 h on day 1. 3. On day 2, remove thymidine containing media after 16–24 h of treatment, wash cells twice with 10 mL PBS and release cells into fresh media for 3 h. 4. 3 h after release, add 40 ng/mL of nocodazole for collecting cells throughout cell cycle and 100 ng/mL of nocodazole for kinase inhibition experiment for 16 h (see Note 10). Proceed with Subheading 3.2.3 for cell cycle experiment and Subheading 3.2.4 for kinase inhibitor experiment.

3.2.3 Sample Collection for the Duration of Cell Cycle

1. On day 3, collect the cells treated with 40 ng/mL nocodazole by mitotic shake-off (see Note 11). Spin down the supernatant media to collect mitotic cells and wash the cells twice with PBS.

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Asynchronous HeLa cells Thymidine

20 h

G1/S arrest Release

3h

Nocodazole 40 ng/mL

Nocodazole 100 ng/mL

Prometaphase arrest

Prometaphase arrest Release Monitor cell-cycle phosphorylation Harvest cells after 30 mins at indicated time points for 24 h

MG132

30 mins

MLN8054 ZM447439 Hesperadin BI2536 Flavopiridol

30 mins

Kinase inhibition

Collect cells

kDa 290 240 170 116

p-RV[S/T]F

0.5 1 2 4 10 12 13 15 17 19 22

Async p-RV[S/T]F

Time after release from T/N block (hrs)

Tubulin

35 66

BI2536

ZM447439

Hesperadin

MLN8054 (5 µM)

Flavopiridol 250 150 100 75 50 37

76 PP1 p-Thr 320

Mitosis

Nitrocellulose membrane Async

SDS-PAGE gel

MLN8054 (1 µM)

Lyse in 2X SDS-PAGE Laemelli buffer and boil

H3 p-Ser 10

15

AurA p-Ser 288 AurB p-Ser 232 PP1 p-Thr 320

50

TCTP p-Ser 46

25

Actin

37

37

Fig. 3 Workflow for monitoring cell-cycle phosphorylation and mitotic kinase dependent phosphorylation using phospho-specific antibodies. HeLa cells were synchronized in prometaphase using a thymidine/nocodazole

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2. Release the cells into fresh culture media. Plate the cells in a 6-well plate (11 wells for 11 time points over 24 h). 3. Collect the cells at indicated time points (Fig. 3) starting at 30 min after release from nocodazole by centrifugation for 2 min at 300
g (see Note 12). 4. Wash the cells once with PBS and lyse as in Subheading 3.2.5. 5. Repeat steps 3 and 4 for each time point over the next 22 h. 3.2.4 Kinase Inhibitor Treatment in Mitotic Cells and Sample Preparation

1. Pretreat the mitotic cells with 20 μM proteasome inhibitor, MG132, for 30 min. 2. After preincubation with MG132, add the respective kinase inhibitors at indicated concentrations (Table 1) for 30 min. 3. Collect cells by mitotic shake-off. Gently knock the plates on the bench to detach mitotic cells. Wash cells once with PBS after collection.

3.2.5 Cell Lysis and Sample Preparation

1. Lyse the cell pellets in lysis buffer by sonication. For this, add 200 μL of lysis buffer with protease inhibitors to each cell pellet. Sonicate the lysate three times with pulses of 30 s each at mid-range intensity setting. 2. After sonication, clear the lysates in a tabletop centrifuge for 15 min at 14,000
g. Add an equal volume of 2
Laemmli buffer to the supernatant in a fresh microfuge tube and boil the sample at 95  C for 5 min.

3.2.6 SDS-PAGE and Western Blotting

1. For SDS-PAGE analysis, we use the Bio-Rad Mini-PROTEAN electrophoresis set up. Wash the glass plates and clean them with 70% ethanol and dry before use. Place the clean glass plates in the gasket and fit into the gel casting stand system. Mark the glass plate to indicate the height of resolving gel (usually 7.5 mL). 2. For a 10% resolving gel, combine 3.3 mL 30% acrylamide, 2.5 mL 1.5 M Tris–HCl pH 8.8, 0.1 mL 10% SDS, and 4 mL water.

ä Fig. 3 (continued) block and either released into fresh culture media to monitor cell cycle phosphorylation of mitotic proteins or treated with indicated kinase inhibitors to determine the role of mitotic kinases in phosphorylation of specific proteins. The cells were collected and lysed at indicated time points followed by SDS-PAGE and Western blot analysis. In this method, we use a p-RV[S/T]F antibody to monitor the phosphorylation of RV[S/T]F motif containing proteins throughout the cell cycle as indicated in the immunoblot. Furthermore, we also use the p-RV[S/T]F antibody to determine the effect of mitotic kinase inhibition on this phosphorylation event in HeLa cells. p-H3 Ser 10, p-AurA Thr 288, p-AurB T232, p-PP1 Thr 320, p-TCTP Ser 46, and actin were used as controls to illustrate the efficiency of kinase inhibitors (Western blots from Isha Nasa, Scott F. Rusin, Arminja N. Kettenbach, & Greg B. Moorhead (2018) Aurora B opposes PP1 function in mitosis by phosphorylating the conserved PP1-binding RVxF motif in PP1 regulatory proteins. Science Signaling 11, eaai8669. Reprinted with permission from AAAS)

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Table 1 List of mitotic protein kinase inhibitors and their concentrations used in this method

Stock concentration

Working concentration

Volume per 10 mL media

Kinase inhibitor

Target kinase (IC50)

MLN8054

Aurora A (IC50 4 nM) [17] Aurora B (IC50 172 nM) [17]

5 mM

1–5 μM

2–10 μL

ZM447439

Aurora A (IC50 110 nM) [18] Aurora B (IC50 130 nM) [18]

5 mM

5 μM

10 μL

Hesperadin

Aurora B (IC50 250 nM) [19]

0.1 mM

100 nM

10 μL

BI2536

PLK1 (IC50 0.83 nM) [20]

0.1 mM

100 nM

10 μL

Flavopiridol

CDK1 (IC50 30 nM) [21] CDK2 (IC50 40 nM) [21] CDK4 (IC50 20–40 nM) [22] CDK6 (IC50 60 nM) [22] CDK7 (IC50 875 nM) [21] CDK9 (IC50 20 nM) [21]

2 mM

2 μM

10 μL

3. When ready to pour, add 100 μL APS and 10 μL TEMED. Overlay the solution with 1 mL of 100% ethanol. Let the gel polymerize for at least 15 min. 4. Once the resolving gel is polymerized, drain the ethanol out and wash with water. Prepare the stacking gel solution by adding 3.1 mL water, 1.3 mL 0.5 M Tris–HCl pH 6.8, 0.6 mL 30% acrylamide, 0.05 mL 10% SDS. When ready to pour add 50 μL 10% APS and 5 μL TEMED. Pour the stacking over the polymerized resolving gel and insert the comb at an angle to avoid getting bubbles in the wells. Let the stacking gel polymerize for at least 15 min. 5. Place the gel in the buffer tank and setup for loading. Add 1
SDS-PAGE running buffer to the tank, slowly remove the comb, and carefully wash the wells with running buffer using a syringe. 6. Load the boiled samples on the gel along with appropriate protein markers. Run the gel at 150 V for about 1 h to resolve the proteins on the gel. 7. After running the gel, carefully remove the gel from the plates. Remove and discard the stacking gel and soak the gel in cold transfer buffer until ready to use. 8. Set up the transfer cassette by laying a wetted sponge followed by Whatman filter paper and roll out the paper to remove bubbles. Place the presoaked gel on the paper followed by the nitrocellulose membrane. Place another Whatman filter paper,

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roll out any bubbles and place a wet sponge on top. Carefully close and lock the cassette. 9. Add a frozen ice-block and a stir bar to the transfer container. Fill the transfer container with transfer buffer and carefully place the transfer cassette in the tank avoiding any spill. 10. Connect the anode to anode and cathode to cathode (red to red side and black to black side) and run the transfer at 100 V for 1 h with stirring. 11. After the transfer is complete, carefully take out the nitrocellulose membrane and add blocking buffer to block the membrane for 1 h at room temperature with rocking. Add fresh phosphatase inhibitor (50 mM NaF) to blocking buffer for all phospho-specific antibodies to abolish the activity of trace phosphatases in the blocking solution. 3.2.7 Immunoblotting with Phospho-Specific Antibodies

1. Discard the blocking buffer and add the primary phosphospecific antibody (dilution dependent on the respective antibody) diluted in blocking buffer (plus phosphatase inhibitors) to the membrane overnight at 4  C. For the p-RV[S/T]F antibody, any epitopes not specific for the phospho-peptide are quenched by preincubating the antibody with 5 μg/mL nonphosphorylated peptide “KRRVSFADK” for 30 min before adding it to the membrane. Antibodies and dilutions used in this protocol are listed in Table 2. 2. After overnight incubation, wash the membrane three times with TBST for 5 min each with gentle rocking.

Table 2 List of primary antibodies used in this method Primary antibody

Source

Dilution used

p-RV[S/T]F

In-house generated

5 μg/mL antibody with 5 μg/mL dephosphorylated peptide

PP1 p-Thr 320

Cell Signaling Technology (Catalog #2581S)

1:1000

α-Tubulin

Sigma-Aldrich (Catalog #T9026)

1:2000

H3 p-Ser 10

Cell Signaling Technology (Catalog #9701)

1:1000

Aur A (p-Thr 288)/Aur B (p-Thr 232)/ Aur C(p-Thr 198)

Cell Signaling Technology (Catalog #2914S)

1:500

TCTP p-Ser 46

Cell Signaling Technology (Catalog #5251)

1:1000

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3. Discard the wash solution and add the respective secondary antibody in fresh blocking buffer plus phosphatase inhibitor (50 mM NaF) incubating the membrane for 1 h at room temperature with rocking. 4. Wash the membrane again after the secondary antibody incubation three times with TBST for 5 min each with rocking. 5. Place the membrane in TBS and prepare the enhanced chemiluminescence (ECL) developing solution. 6. Incubate the membrane with enough volume of ECL solution to cover the membrane (usually 1–2 mL) for 1 min. Put the membrane in a plastic sheet protector and develop either using a ChemiDoc or a X-ray film. Examples of gels probed with phospho-RV[S/T]F antibody and developed using ChemiDoc is shown in Fig. 3.

4

Notes 1. Glutaraldehyde is acutely toxic, harmful if swallowed or inhaled, can cause severe skin burns or eye damage. Keep containers tightly sealed and upright. Always use a fresh vial of glutaraldehyde for peptide coupling. 2. DMSO is a highly polar aprotic organic reagent and can be used as a solvent for both organic and inorganic chemicals. It can also act as a cryoprotectant. Use cell culture grade DMSO as a solvent for most kinase inhibitors used in this method. 3. Trypsin-EDTA is used for dislodging adherent cells from the surface. This EDTA-containing solution can be irritating to the eyes, respiratory system and skin. Store frozen at 20  C and thaw before use. Avoid repeated freeze–thaw cycles. 4. Thymidine is a pyrimidine nucleoside, high concentrations of which can block DNA replication by interfering in the deoxynucleotide-metabolism pathway, thereby arresting cells in G1/S phase of cell cycle. Thymidine can be dissolved for a 100 mM stock by incubation at 37  C followed by filter sterilization. 5. Antimitotic agent, nocodazole, is a member of class of benzimidazoles and is an inhibitor of microtubule polymerization. The structure, chemical safety and toxicity of this compound can be found at https://pubchem.ncbi.nlm.nih.gov/com pound/4122. 6. MG132 is a cell permeable proteasome inhibitor which should be stored in tightly sealed container in a cool area. Avoid contact with eyes and skin and avoid inhalation.

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7. Prepare the lysis buffer fresh each time. Add fresh protease (see Subheading 2.3.2) and phosphatase inhibitors (see Subheading 2.3.3) to a small aliquot based on number of cells to be lysed. 8. Coupling of peptides utilizes primary amines for immobilization. Use peptide sequences containing lysine residues or modify the N or C termini of the peptide sequences to include additional lysine residues to help in the coupling process. 9. KLH can serve as a carrier in peptide-protein conjugates and generate a strong immune response ensuring the successful generation of anti-peptide antibodies. 10. Lower concentrations of nocodazole (up to 40 ng/mL) do not affect the release of cells from prometaphase block. When cells are released from this block, they can form the spindle microtubules and progress through mitosis. At higher concentrations of nocodazole (up to 100 ng/mL), the efficiency of prometaphase arrest is better. However, at high concentrations of nocodazole, the cells inherit many microtubule anomalies and cannot form the spindle microtubules and thus cannot progress through mitosis even after release into fresh media. 11. To perform mitotic shake-off, gently tap the tissue culture plate on a hard surface and collect the supernatant media. Additional PBS can be added to the plate to repeat the process and get most of the rounded mitotic cells off the plate. 12. On average HeLa cells take 20–22 h to complete a cell cycle, in which they spend more than 8–9 h in G1, 8–9 h in S, 2–3 h in G2, and 1 h in mitosis. The time-points chosen for cell-cycle analysis are solely based on the cell type and the duration of cell cycle in the respective cell type. If using other cells than HeLa, adjust the time-points for the experiment accordingly.

Acknowledgments GBM is funded by the Natural Sciences and Engineering Research Council of Canada. A.N.K. is funded by NIH/NIGMS R35GM119455 and NIH/NCI R33CA225458. References 1. Bayliss R, Fry A, Haq T, Yeoh S (2012) On the molecular mechanisms of mitotic kinase activation. Open Biol 2(11):120136. https://doi. org/10.1098/rsob.120136 2. Dephoure N, Zhou C, Villen J, Beausoleil SA, Bakalarski CE, Elledge SJ, Gygi SP (2008) A quantitative atlas of mitotic phosphorylation.

Proc Natl Acad Sci U S A 105 (31):10762–10767. https://doi.org/10.1073/ pnas.0805139105 3. Olsen JV, Vermeulen M, Santamaria A, Kumar C, Miller ML, Jensen LJ, Gnad F, Cox J, Jensen TS, Nigg EA, Brunak S, Mann M (2010) Quantitative phosphoproteomics

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reveals widespread full phosphorylation site occupancy during mitosis. Sci Signal 3(104): ra3. https://doi.org/10.1126/scisignal. 2000475 4. Sharma K, D’Souza RCJ, Tyanova S, Schaab C, Wisniewski JR, Cox J, Mann M (2014) Ultradeep human phosphoproteome reveals a distinct regulatory nature of Tyr and Ser/Thrbased signaling. Cell Rep 8(5):1583–1594. https://doi.org/10.1016/j.celrep.2014.07. 036 5. Barr FA, Elliott PR, Gruneberg U (2011) Protein phosphatases and the regulation of mitosis. J Cell Sci 124(Pt 14):2323–2334. https://doi. org/10.1242/jcs.087106 6. Cundell MJ, Hutter LH, Nunes Bastos R, Poser E, Holder J, Mohammed S, Novak B, Barr FA (2016) A PP2A-B55 recognition signal controls substrate dephosphorylation kinetics during mitotic exit. J Cell Biol 214 (5):539–554. https://doi.org/10.1083/jcb. 201606033 7. Wu JQ, Guo JY, Tang W, Yang C-S, Freel CD, Chen C, Nairn AC, Kornbluth S (2009) PP1-mediated dephosphorylation of phosphoproteins at mitotic exit is controlled by inhibitor-1 and PP1 phosphorylation. Nat Cell Biol 11(5):644–U451. https://doi.org/ 10.1038/ncb1871 8. Bollen M, Peti W, Ragusa MJ, Beullens M (2010) The extended PP1 toolkit: designed to create specificity. Trends Biochem Sci 35 (8):450–458. https://doi.org/10.1016/j. tibs.2010.03.002 9. Hertz EPT, Kruse T, Davey NE, LopezMendez B, Sigurethsson JO, Montoya G, Olsen JV, Nilsson J (2016) A conserved motif provides binding specificity to the PP2A-B56 phosphatase. Mol Cell 63(4):686–695. https://doi.org/10.1016/j.molcel.2016.06. 024 10. Wang X, Bajaj R, Bollen M, Peti W, Page R (2016) Expanding the PP2A interactome by defining a B56-specific SLiM. Structure (London, England: 1993) 24 (12):2174–2181. https://doi.org/10.1016/j. str.2016.09.010 11. Heroes E, Lesage B, Gornemann J, Beullens M, Van Meervelt L, Bollen M (2013) The PP1 binding code: a molecularlego strategy that governs specificity. FEBS J 280(2):584–595. https://doi.org/10.1111/j. 1742-4658.2012.08547.x 12. Nasa I, Kettenbach AN (2018) Coordination of protein kinase and phosphoprotein phosphatase activities in mitosis. Front Cell Dev

Biol 6:30. https://doi.org/10.3389/fcell. 2018.00030 13. Nasa I, Rusin SF, Kettenbach AN, Moorhead GB (2018) Aurora B opposes PP1 function in mitosis by phosphorylating the conserved PP1-binding RVxF motif in PP1 regulatory proteins. Sci Signal 11(530). https://doi.org/ 10.1126/scisignal.aai8669 14. Dent P, Campbell DG, Caudwell FB, Cohen P (1990) Identification of three in vivo phosphorylation sites on the glycogen-binding subunit of protein phosphatase 1 from rabbit skeletal muscle, and their response to adrenaline. FEBS Lett 259(2):281–285 15. Liu D, Vleugel M, Backer CB, Hori T, Fukagawa T, Cheeseman IM, Lampson MA (2010) Regulated targeting of protein phosphatase 1 to the outer kinetochore by KNL1 opposes Aurora B kinase. J Cell Biol 188 (6):809–820. https://doi.org/10.1083/jcb. 201001006 16. Kim Y, Holland AJ, Lan W, Cleveland DW (2010) Aurora kinases and protein phosphatase 1 mediate chromosome congression through regulation of CENP-E. Cell 142(3):444–455. https://doi.org/10.1016/j.cell.2010.06.039 17. Manfredi MG, Ecsedy JA, Meetze KA, Balani SK, Burenkova O, Chen W, Galvin KM, Hoar KM, Huck JJ, LeRoy PJ, Ray ET, Sells TB, Stringer B, Stroud SG, Vos TJ, Weatherhead GS, Wysong DR, Zhang M, Bolen JB, Claiborne CF (2007) Antitumor activity of MLN8054, an orally active small-molecule inhibitor of Aurora A kinase. Proc Natl Acad Sci U S A 104(10):4106–4111. https://doi. org/10.1073/pnas.0608798104 18. Ditchfield C, Johnson VL, Tighe A, Ellston R, Haworth C, Johnson T, Mortlock A, Keen N, Taylor SS (2003) Aurora B couples chromosome alignment with anaphase by targeting BubR1, Mad2, and Cenp-E to kinetochores. J Cell Biol 161(2):267–280. https://doi.org/ 10.1083/jcb.200208091 19. Hauf S, Cole RW, LaTerra S, Zimmer C, Schnapp G, Walter R, Heckel A, van Meel J, Rieder CL, Peters JM (2003) The small molecule Hesperadin reveals a role for Aurora B in correcting kinetochore-microtubule attachment and in maintaining the spindle assembly checkpoint. J Cell Biol 161(2):281–294. https://doi.org/10.1083/jcb.200208092 20. Steegmaier M, Hoffmann M, Baum A, Le´na´rt P, Petronczki M, Krssa´k M, Gu¨rtler U, Garin-Chesa P, Lieb S, Quant J, Grauert M, Adolf GR, Kraut N, Peters JM, Rettig WJ (2007) BI 2536, a potent and selective

Mitotic Protein Kinase Inhibitors and Phospho-Specific Antibodies inhibitor of polo-like kinase 1, inhibits tumor growth in vivo. Curr Biol 17(4):316–322. https://doi.org/10.1016/j.cub.2006.12.037 21. Montagnoli A, Valsasina B, Croci V, Menichincheri M, Rainoldi S, Marchesi V, Tibolla M, Tenca P, Brotherton D, Albanese C, Patton V, Alzani R, Ciavolella A, Sola F, Molinari A, Volpi D, Avanzi N, Fiorentini F, Cattoni M, Healy S, Ballinari D, Pesenti E,

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Chapter 16 Visualization of Radiation-Induced Cell Cycle Kinetics with a Fluorescent Ubiquitination-Based Cell Cycle Indicator (Fucci) Atsushi Kaida and Masahiko Miura Abstract Among the methods for detecting cell cycle kinetics in tumor cells, fluorescent ubiquitination-based cell cycle indicator (Fucci) is innovative because it allows observation in live cells without losing spatiotemporal information. We succeeded in using the Fucci system to visualize radiation-induced G2 arrest in tumor cells with deficient p53 function. Here we describe protocols for establishing Fucci-expressing cell lines and analyzing radiation-induced G2 arrest kinetics in three different models: monolayer cell cultures, spheroids, and xenografted solid tumors in mice. Key words Cell cycle, Radiation, Fluorescent ubiquitination-based cell cycle indicator (Fucci), Timelapse imaging

1

Introduction Ionizing radiation causes DNA damage in cells, leading to induction of the DNA damage response, such as DNA repair, activation of the cell cycle checkpoint, and apoptosis [1, 2]. The importance of G2 arrest, especially in tumor cells with deficient p53 function, has been recognized as “redistribution” during fractionated radiotherapy, as G2/M boundary is a radiosensitive phase; consequently, accumulation at this phase results in radiosensitization [3]. Therefore, understanding the detailed kinetics of radiation-induced G2 arrest is also crucial in the clinic. The first approach to understanding this issue was autoradiography using 3H-thymidine [4], but this was replaced by flow cytometry based on DNA content analysis [5]. However, this method loses spatiotemporal information due to the requirement for preparing a single-cell suspension that can pass through the flow cell. Given that features of the tumor microenvironment including hypoxia, low pH, and anchorage-independent

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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a

mKO2-hCdt1(30/120)

mAG-hGeminin(1/110) 1

mAG b

110

30

hGem

mKO2

120

hCdt1

c

M G1

G2

S early S Fig. 1 Principle of Fucci. (a) The components of Fucci, mAG-hGeminin (1/110) and mKO2-hCdt1 (30/120). (b) Fluorescence color according to cell cycle phase in cells expressing Fucci. (c) Merged image of HeLa-Fucci cells

three-dimensional growth [6, 7] can greatly influence G2 arrest, next-generation methods are required to visualize cell cycle kinetics without losing spatiotemporal information under live conditions. Since the green fluorescent protein was first discovered in the 1960s [8], fluorescent proteins have become powerful tools for visualizing various cellular phenomena and have been used in many studies [9]. The fluorescent ubiquitination-based cell cycle indicator (Fucci) was developed by taking advantage of two types of fluorescent proteins and the cell cycle-dependent ubiquitination of Cdt1 and Geminin (Fig. 1; ref. 10). This system consists of two components: mKO2, a red fluorescent protein, fused with human Cdt1 (a.a. 30–120) expressed in G1 phase; and mAG, a green fluorescent protein, fused with human Geminin (a.a. 1–110) expressed in S/G2/M phases (Fig. 1a). As a result, cells expressing Fucci emit red and green fluorescence in the G1 and S/G2/M phases, respectively, allowing us to visualize cell cycle phase in individual live cells (Fig. 1b, c; ref. 10). We speculated that when combined with time-lapse imaging, this method would enable visualization of radiation-induced G2 arrest kinetics in p53-deficient tumor cells. Indeed, we succeeded for the first time in such visualization using monolayer cultures of cancer cell lines carrying mutant p53 or in which p53 function was disrupted by human papillomavirus infection. The kinetics of accumulation of cells emitting green fluorescence, and release accompanied by the appearance of red fluorescence, coincided almost perfectly with the timing of DNA content alterations observed by flow cytometry [11–14]. Next, we applied the approach to spheroids, which mimic many aspects of the in vivo tumor microenvironments. Confocal laser scanning made it possible to observe inside

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the spheroid; due to the optical limitations, however, depth was restricted to 60–100 μm below the surface under live conditions [13, 15]. The method was also applied to xenografted tumors in mice. In this case, histology sections had to be prepared in order to see inside of the tumor [16]. Nevertheless, we could also quantify fluorescence intensity from tumor cells in surface areas of solid tumors, and it was possible to obtain information about radiation-induced G2 arrest kinetics in real time [16]. Comparison of results from each model revealed that radiation-induced G2 arrest is significantly extended as growth conditions approach those in vivo. In addition to radiation, we found that Fucci is also useful for visualizing cell cycle kinetics after treatment with various kinds of drugs and under specific conditions [11, 12, 17, 18]. Hence, the availability of Fucci is of immense value to cancer biology. In this chapter, we introduce protocols for establishing Fucciexpressing cell lines and analyzing radiation-induced cell cycle kinetics using tumor cells expressing Fucci in three different models: monolayer cell cultures, spheroids, and xenografted solid tumors in mice.

2 2.1

Materials Plasmids

1. Lentivirus vectors CFII-EF-mKO2-hCdt1(30/120) and CFII-EF-mAG-hGeminin(1/110) [available from RIKEN (URL: https://cfm.brc.riken.jp/lentiviral-vectors/plasmidlist/)]. 2. Packaging plasmid psPAX2 (Addgene). 3. Envelope plasmid pMD2.G (Addgene).

2.2

Cell Lines

1. HEK293T cells. 2. Cells expressing Fucci: HeLa cells expressing Fucci (HeLaFucci) and SAS cells (a human tongue squamous cell carcinoma cell line) were used for the experiments described in this chapter. SAS cells expressing Fucci were established by our group as described below. HeLa-Fucci cells and SAS cells are available from RIKEN (URL: https://cell.brc.riken.jp/en/) with permission from Dr. Atsushi Miyawaki and the Health Science Research Resource Bank (Sendai, Japan), respectively.

2.3

Reagents

1. Complete medium: Dulbecco’s Modified Eagle Medium (DMEM) with 100 units/mL penicillin and 100 μg/mL streptomycin, supplemented with 10% fetal bovine serum (FBS). This complete medium was used in all procedures shown in this chapter. 2. jetPRIME (Polyplus Transfection).

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3. Hexadimethrine bromide (Polybrene): To prepare the stock solution, 10 mg of polybrene should be dissolved in 1 mL of sterilized water, and the stock should be stored at 20  C. 4. Phosphate buffer saline (PBS) containing 1% FBS. 5. 2 DMEM: 26.8 g DMEM powder (D5648: Sigma) and 7.4 g sodium carbonate are dissolved in 1 L distilled water. Filtersterilize. 6. 1% agar solution: 100 mg agar (molecular biology grade) in 10 mL sterile water. Boil to dissolve. The 1% agar solution can be stored at 4  C and reused after boiling. 7. 0.5% agar DMEM: Mix 1 mL of 2-DMEM (warmed at 42  C) to 1 mL of 1% agar solution (before gelation) and apply to coat the dish or plate. 8. 0.25% trypsin in PBS. 9. 4% paraformaldehyde in PBS. 10. 20% sucrose in PBS. 11. Optimal cutting temperature (OCT) compound. 12. Hoechst33342: To prepare the stock solution, 10 mg of Hoechst33342 powder should be dissolved in 1 mL of sterilized water, and the stock should be stored at 20  C. To stain cellular DNA, the stock solution is diluted to 10 μg/mL with DMEM. The stock solution is diluted with 0.9% saline for intravenous injection. 13. Pimonidazole HCl (Hypoxyprobe-1; Hypoxyprobe): Prepare stock of 100 mg/mL in 0.9% saline. Store at 4  C. To inject intraperitoneally, dilute the stock solution to 60 mg/mL with 0.9% saline. 14. Target Retrieval Solution (Dako). 15. Protein Block, Serum Free (Dako). 16. PBS containing 0.1% Triton-X100. 17. Hypoxyprobe ™ PAb2627 Antibody (Hypoxyprobe). 18. Alexa Fluor 647 goat anti-rabbit IgG (H + L). 19. ProLong Gold Antifade Reagent. 20. Isoflurane: When mice are anesthetized, isoflurane must be administered with a vaporizer and the gas concentration should be adjusted between 1% and 4%. 2.4

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1. 5-mL round-bottom tube with cell strainer cap. 2. 10-mL syringe and 0.45- μm polyether sulfone (PES) filter. 3. 35-mm glass-based dish or plastic dish. 4. HydroCell 24- or 96-well plate (CellSeed). 5. 1.0-mL syringe with 26 G needle.

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6. BIOREVO BZ-9000 fluorescence microscope (Keyence). 7. FV10i-LIV confocal laser scanning microscope (Olympus). 8. X-irradiator: This equipment was used to yield DNA double strand breaks that subsequently lead to G2 arrest. 9. Photon imager (BIOSPACE LAB).

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1. Seed HEK293T cells on a 60 mm dish and start transfection once confluence reaches 80–90%. 2. Use jetPRIME transfection reagents for DNA complex formation. Add the appropriate volumes of packaging plasmid and envelope plasmid with each lentivirus vector in 300 μL of jetPRIME buffer in a 1.5 mL tube. Mix gently and incubate for 10 min at room temperature (see Note 1). 3. Change the medium on the HEK293T cells during the 10 min incubation described in the previous step. 4. Add the complex to cells in fresh media and mix gently by shaking the dish. 5. Incubate for 24 h at 37  C. The dish should be shaken gently every 30 min for the first 2 h to increase transfection efficiency. 6. Remove the medium containing DNA complexes and add 8 mL of fresh medium 24 h after transfection. 7. Incubate for more 24 h at 37  C. 8. Collect conditioned medium containing viruses and filter it through a 0.45-μm PES filter to get rid of any cells.

3.2 Establishment of Cells Expressing Fucci (Fig. 2a)

1. For Fucci transduction, seed 1–3  104 cells in a 12-well plate a day before transduction. 2. Add media containing two kinds of virus, mKO2-hCdt1 (30/120) and mAG-hGem(1/110), simultaneously to target cells. Adding 8 μg/mL of Polybrene may increase the efficiency of transduction, but it may also kill some cells. Incubate for 24 h at 37  C (see Note 2). 3. After 24 h replace medium with fresh complete medium. 4. Check expression of the Fucci constructs with a fluorescence microscope 48–72 h after the infection. If the transduction efficiency is too low, more infections must be performed (see Note 3). 5. Once the number of infected cells becomes sufficient to perform cell sorting (at least 80–90% confluence on a 60-mm dish), harvest cells and make a single-cell suspension in PBS

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containing 1% FBS. Before sorting, the cell suspension should be filtered through a 35 μm nylon mesh. 6. Sort green-positive cells into a collection tube (Fig. 2c). mAG-hGem(1/110) should be in sorted cells. 7. Seed the sorted cells on an appropriately sized dish or plate, depending on the number of cells obtained in the previous step. Incubate at 37  C. 8. Check for green and red fluorescence by microscopy. If cells expressing red fluorescence are found, it means both vectors, mAG-hGem(1/110) and mKO2-hCdt1(30/120), should be integrated in some cells because only green-positive cells were sorted in step 6. If red cells are not observed, another infection with mKO2-hCdt1(30/120) virus must be performed. 9. Once the number of cells becomes sufficient to perform further cell sorting for red fluorescence, harvest the cells in PBS containing 1% FBS and filter to make a single-cell suspension through a 35 μm nylon mesh. 10. Sort red-positive cells into a collection tube or a single cell into each well of a 96-well plate. If cells are collected in a tube, plate single cells onto a 96-well plate by the limiting dilution method. Incubate at 37  C for 7–10 days until colonies form (Fig. 2d). 11. After colony formation, check fluorescence expression. Pick colonies in which two different populations, green and red

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cells, are observed and transfer to a larger-scale plate to expand the cells. 12. To confirm that Fucci fluorescence is correlated with cell-cycle phase, perform flow cytometry or time-lapse imaging. In flow cytometry, expression of each fluorescence signal should be correlated with DNA content, as determined by Hoechst staining, but the whole population should be distributed in an inverse U shape (Fig. 2e). To prepare samples for flow cytometry, cells are trypsinized, and cell suspension in PBS is filtrated through the mesh after Hoechst staining is performed by incubating living cells with 10 μg/mL of Hoechst33342 at 37  C for 30 min. In time-lapse imaging, confirm the order of fluorescence, as well as the doubling time (see Note 4). 3.3 Time-Lapse Imaging in Monolayer Culture After Irradiation

1. Before starting the time-lapse imaging, turn on and prepare the imaging equipment. The incubator system on the microscope needs to be prewarmed and equilibrated with 5% CO2 and 95% air. The appropriate place in the chamber should be filled with distilled water to maintain humidity in the chamber. 2. Seed cells expressing Fucci at the appropriate density on either 35-mm glass-bottom or plastic dishes, according to the purpose (see Note 5). 3. At least 1 day after seeding, irradiate cells with an X-irradiator to cause DNA double strand breaks. Any drugs or other types of radiation such as γ-rays depending on the purpose can be applied instead of X-rays. 4. Place the dish with irradiated cells on the warmed stage in the chamber in the fluorescence microscope. 5. Place the lid on the chamber. 6. Decide the magnification and exposure time for phase-contrast and fluorescence images. If you want to observe cell cycle kinetics in each cell, a 20 lens is recommended. The phasecontrast objective lenses are better for distinguishing the characteristic round shape of mitotic cells from interphase cells. 7. Register as many fields as you want to observe (see Note 6). 8. Register a Z-stack. 9. Set the interval and total time for this experiment. 10. Start the time-lapse imaging. 11. After time-lapse imaging, merge the phase-contrast and fluorescence images, and make a series of images following irradiation. Adjust brightness and contrast as necessary (Fig. 3a). 12. Record the change in fluorescence color as the durations of red and green phases in each cell in any software (e.g., Excel). Based on the result, measure the durations of red and green

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phases in each cell, and determine whether mitosis occurs normally after irradiation (Fig. 3b, c). 3.4 Time-Lapse Imaging in Spheroids After Irradiation

1. Seed cells expressing Fucci on HydroCell 24- or 96-well plates. The number of cells seeded depends on cell lines and purpose. 2. Incubate at 37  C for 7–10 days and replace media every 3 days. Incubation period is decided according to cell lines and experimental purpose. 3. Irradiate spheroids with an X-irradiator. 4. For time-lapse imaging after irradiation, use a 35-mm glassbased dish coated with DMEM containing 0.5% agar to prevent the cells from attaching to the dish (see Note 7). 5. Transfer only spheroids for observation on the coated dish. Use a glass Pasteur pipette or a pipette tip and take a spheroid carefully as you aspirate the media slowly by the tip.

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6. Place the dish containing the irradiated spheroids on a warmed stage. 7. Place the lid on the chamber. 8. Decide on the magnification and exposure time for mAG and mKO2. 9. Register as many fields as you want to observe. 10. Register a range of depth. Due to differences in transparency, the range you can visualize depends on the cell line. When light cannot be delivered at a given depth and the signal is not detected, a center of the spheroid becomes dark; this will help decide the range. 11. Set an interval and total time for this experiment (see Note 6). 12. Start time-lapse imaging. 13. After time-lapse imaging, adjust the brightness and contrast and arrange sequential images (Fig. 4a). 14. To perform a line profile for each image, use the analysis software for the FV10i-LIV. Choose a sectional image in which you want to quantify each fluorescence intensity and draw a line. Export the quantification result as an Excel file, and calculate relative intensity by adjusting the maximum fluorescence intensity to 1.0 (Fig. 4b). 15. To quantify the ratio of green to red, measure the integrated area for green and red using the ImageJ software (available from http://rsbweb.nih.gov/ij/). Calculate the ratio by dividing the integrated green area by the integrated red area in each section and plot the ratio as a function of the depth (Fig. 4c). 3.5 Establishment of Xenografted Tumors

1. Trypsinize exponentially growing cells and harvest into a 15 mL conical tube. 2. Count the cells. 3. Wash cells with 1 PBS. 4. Resuspend cells in 1 PBS at an appropriate concentration (e.g., 1  106 cells per 50 μL of 1 PBS). 5. Inject cells subcutaneously into the hind leg of a nude mouse using a 1.0 mL syringe with a 26 G needle. Once the xenografted tumor grows to the appropriate size, prepare to irradiate the tumors. 6. Before irradiation, shield the whole body of the mouse, other than the tumor, using an appropriate thickness of lead. 7. Irradiate the tumors using an X-irradiator. 1. Excise tumors at appropriate times after irradiation. As a control, also prepare tumors without irradiation. For the detection of blood perfusion, inject 16 mg/kg Hoechst 33342

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3.6 Histological Analysis of Tumors Derived from Cells Expressing Fucci

intravenously 1 min before sacrifice. As a hypoxia marker, inject 60 mg/mL pimonidazole HCl intraperitoneally 30 min before sacrifice. 2. Wash with 1 PBS and remove unnecessary tissue from the tumors. 3. Fix tumors in 4% paraformaldehyde in PBS overnight. 4. Immerse in 20% sucrose in PBS until the tumors sink to the bottom of the tube. 5. Embed tumors in OCT compound and store at

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6. Cut tumors into 8- to 10- μm sections using a cryostat. 7. Wash sections with 1 PBS. For observation of mAG, mKO2, and blood perfusion, go to step 16.

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8. For detection of pimonidazole with mAG and mKO2, boil sections in Target Retrieval Solution for 20 min at 70  C (see Note 8). 9. Wash sections with 1 PBS. 10. Block nonspecific antigens in Protein Block, Serum Free for 10 min. 11. Incubate sections with anti-Hypoxyprobe ™ PAb2627 Antibody (dilution 1:50 in 1 PBS containing 0.1% Triton X-100) for 60 min at room temperature. 12. Wash three times with 1 PBS. 13. Incubate sections with Alexa Fluor 647 Goat Anti–rabbit IgG (H + L) (dilution 1:1000 in 1 PBS containing 0.1% Triton X-100) for 30 min at room temperature. 14. Wash three times with 1 PBS. 15. For counterstaining, incubate sections with 1 μg/mL Hoechst 33342 diluted in 1 PBS for 10 min at room temperature. 16. Mount sections with ProLong Gold Antifade Reagent. 17. Observe sections using a fluorescence microscope (Fig. 5a, b). 3.7 (Optional) Optical Imaging of Fluorescence Kinetics After Irradiation (See Note 9)

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Notes 1. The best ratio of plasmids and incubation time should be found by performing several trials. 2. Each plasmid can be added separately but not simultaneously. If Polybrene induces too much cell death, decrease the concentration or change the media at an early time (6–8 h) after infection. 3. Perform spinoculation to more efficiently infect target cells with virus. 4. Determine whether the cellular characteristics (doubling time, fraction in each cell cycle phase, behavior after treatment, etc.) are similar to those in parental cells, and choose appropriate clones for your study.

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Fig. 5 Fluorescence kinetics after irradiation in xenografted tumors derived from HeLa-Fucci cells in living mice. (a) Fluorescence images of tumor sections at the indicated times after irradiation. Bar: 500 μm. (b) Fluorescence images of sections from nonirradiated tumors. Upper image shows localization of hypoxia (yellow) with mAG (green), mKO2 (red), and nucleus (blue). The lower image shows blood perfusion (blue). Bar: 200 μm. (c) Optical images of mAG and mKO2 in xenografted tumors at the indicated time after irradiation (IR). NT indicates fluorescence kinetics in nonirradiated tumors. (d) Ratio of green fluorescence intensity to red fluorescence intensity (mAG/mKO2) in live nude mice bearing tumor xenografts after no treatment (NT) or irradiation (IR). mAG/mKO2 ratio was calculated by dividing photon counts of green fluorescence by those of red fluorescence and is plotted as a function of time after irradiation [16]

5. A glass-bottom dish provides better fluorescence images than a plastic one; however, the material to which cells attach may affect cell cycle kinetics. You should confirm which material is more suitable for observing cell cycle kinetics under your conditions. Coating solutions like poly-L-lysine may be also useful for promoting the attachment of cells to the glass-bottom dish. 6. Light exposure can damage cells, which may cause unexpected cell cycle kinetics because the damage activates the cell cycle checkpoint. Optimize parameters such as the exposure time, number of fields, distance between fields, and interval time to observe real cell cycle kinetics without any artifacts.

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7. A spheroid is easy to move on the dish and can leave the field if the dish is shaken even a little bit. Therefore, a smaller space such as a well or the center space on a multiwell glass-based dish is better for time-lapse imaging. 8. Fluorescence is sensitive to temperature. Heating at too high of a temperature (90  C) causes quenching of mAG and mKO2. To avoid this, a lower temperature (70  C) than usual should be applied in order to simultaneously observe mAG and mKO2 along with any antigens. 9. In this assay, only fluorescence signals from the surface of the tumor are detectable because optical limitations make it difficult to detect them inside the tumor. Therefore, the result of this experiment will not necessarily be consistent with the results of histological analysis.

Acknowledgments We thank Dr. A. Miyawaki and Dr. A. Sakaue-Sawano for providing the Fucci plasmids and the HeLa cells expressing Fucci. References 1. O’Connor MJ (2015) Targeting the DNA damage response in cancer. Mol Cell 60:547–560 2. Belka C, Jendrossek V, Pruschy M, Vink S, Verheij M, Budach W (2004) Apoptosismodulating agents in combination with radiotherapy-current status and outlook. Int J Radiat Oncol Biol Phys 58:542–554 3. Hall EJ, Giaccia AJ (eds) (2019) Radiobiology for the radiologist, 8th edn. Wolters Kluwer, Philadelphia, PA 4. Howard A, Pelc SR (1953) Synthesis of deoxyribonucleic acid in normal and irradiated cells and its relation to chromosomal breakage. Heredity 6(Suppl):261–273 5. Pozarowski P, Darzynkiewicz Z (2004) Analysis of cell cycle by flow cytometry. Methods Mol Biol 281:301–311 6. Al Tameemi W, Dale TP, Al-Jumaily RMK, Forsyth NR (2019) Hypoxia-modified cancer cell metabolism. Front Cell Dev Biol 7:4 7. Huber V, Camisaschi C, Berzi A, Ferro S, Lugini L, Triulzi T, Tuccitto A, Tagliabue E, Castelli C, Rivoltini L (2017) Cancer acidity: an ultimate frontier of tumor immune escape and a novel target of immunomodulation. Semin Cancer Biol 43:74–89 8. Shimomura O (2005) The discovery of aequorin and green fluorescent protein. J Microsc 217:1–15

9. Specht EA, Braselmann E, Palmer AE (2017) A critical and comparative review of fluorescent tools for live-cell imaging. Annu Rev Physiol 79:93–117 10. Sakaue-Sawano A, Kurokawa H, Morimura T, Hanyu A, Hama H, Osawa H, Kashiwagi S, Fukami K, Miyata T, Miyoshi H, Imamura T, Ogawa M, Masai H, Miyawaki A (2008) Visualizing spatiotemporal dynamics of multicellular cell-cycle progression. Cell 132:487–498 11. Kaida A, Sawai N, Sakaguchi K, Miura M (2011) Fluorescence kinetics in HeLa cells after treatment with cell cycle arrest inducers visualized with Fucci (fluorescent ubiquitination-based cell cycle indicator). Cell Biol Int 35:359–363 12. Kaida A, Miura M (2012) Visualizing the effect of hypoxia on fluorescence kinetics in living HeLa cells using the fluorescent ubiquitination-based cell cycle indicator (Fucci). Exp Cell Res 318:288–297 13. Onozato Y, Kaida A, Harada H, Miura M (2017) Radiosensitivity of quiescent and proliferating cells grown as multicellular tumor spheroids. Cancer Sci 108:704–712 14. Jiaranuchart S, Kaida A, Onozato Y, Harada H, Miura M (2018) DNA damage response following X-irradiation in oral cancer cell lines HSC3 and HSC4. Arch Oral Biol 90:1–8

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15. Kaida A, Miura M (2013) Visualizing the effect of tumor microenvironments on radiationinduced cell kinetics in multicellular spheroids consisting of HeLa cells. Biochem Biophys Res Commun 439:453–458 16. Kaida A, Miura M (2015) Unusual prolongation of radiation-induced G2 arrest in tumor xenografts derived from HeLa cells. Cancer Sci 106:1370–1376

17. Homma H, Nojima H, Kaida A, Miura M (2019) Induction of endomitosis-like event in HeLa cells following CHK1 inhibitor treatment. Biochem Biophys Res Commun 520:492–497 18. Nojima H, Homma H, Onozato Y, Kaida A, Harada H, Miura M (2020) Differential properties of mitosis-associated events following CHK1 and WEE1 inhibitor treatments in human tongue carcinoma cells. Exp Cell Res 386:111720

Chapter 17 Dynamic Behavior of Inactive X Chromosome Territory During the Cell Cycle as Revealed by H3K27me3-Specific Intracellular Antibody Yuko Sato and Hiroshi Kimura Abstract Posttranslational histone modifications are critical for the regulation of genome function. The levels of histone modifications oscillate during the cell cycle. Most modifications are diluted after DNA replication and then their levels are restored during the rest of the cell cycle with different kinetics depending on the modification. Some modifications, like histone H4 Lys20 monomethylation (H4K20me1), exhibit cell cycle-dependent dynamic changes. To track histone modifications in living cells, we have developed genetically encoded probes termed modification specific intracellular antibodies, or “mintbodies.” As mintbodies shuttle between the cytoplasm and nucleus by diffusion, their nuclear concentration depends on the target modification level. By measuring the nuclear to cytoplasmic intensity ratio of H4K20me1specific mintbody, we have monitored the increase of H4K20me1 in the G2 phase. Here we describe how the mintbody-based methods can be applied to track a specific chromosome, such as the inactive X chromosome (Xi), on which genes are repressed through histone H3 Lys27 trimethylation (H3K27me3). When H3K27me3-specific mintbodies are expressed in cells that harbor Xi, the mintbodies are concentrated on Xi and the dynamic behavior of Xi can be tracked using a confocal microscope. After acquiring 3D time-lapse images, an image analysis allows measuring the volume, shape and H3K27me3 level of Xi during the cell cycle. Key words X chromosome inactivation, Chromosome territory, Histone modification, Intracellular antibody, Live-cell imaging

1

Introduction Posttranslational histone modifications play a fundamental role in gene regulation and chromosome integrity [1–3]. Some modifications can persist over cell divisions being epigenetic marks to maintain the states of gene activation and inactivation. In cycling cells, histone modifications are diluted during DNA replication and their levels are restored before the next round of cell cycle. During DNA replication in the S phase, parental histone H3 and H4 are

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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transferred to either daughter DNA, thereby their modifications can be maintained in roughly half of the nucleosomes on daughter chromatin, while another half of the nucleosomes are assembled with newly synthesized H3 and H4. In general, active marks are soon restored after DNA replication with little contribution of parental modification states, while the restoration of inactive marks occurs slowly and often requires the G1 phase in the next cell cycle [4–8]. For example, mass spectrometry [5] and immunofluorescence [8] have revealed that levels of histone H3 Lys27 trimethylation (H3K27me3), which is associated with gene silencing, decrease during the S phase and increase during the G2 phase through the next G1 phase. Apart from marks associated with gene regulation, some modifications exhibit cell cycle-specific oscillations. The global levels of histone H4 Lys20 monomethylation (H4K20me1) and H3 Ser10 phosphorylation (H3S10ph) drastically increase during G2 and M phases, respectively [9, 10], suggesting possible involvement in chromosome condensation and segregation. These dynamic changes are mediated through the balance between modifying and de-modifying enzymes. Various approaches have been used to detect the global changes in histone modifications during the cell cycle, including mass spectrometry using synchronized cells [4, 9] and quantitative multicolor immunofluorescence using asynchronous cells [8]. In addition, live-cell imaging methods have recently been applied to monitor the modification dynamics in single cells. In order to detect changes in the level of histone modifications in living cells, we have developed genetically encoded modificationspecific intracellular antibodies, termed “mintbodies.” Mintbodies consist of a single-chain variable fragment (scFv) tagged with a fluorescent protein [11, 12] (Fig. 1a). Since the molecular size is small enough to freely pass through the nuclear pores, mintbodies synthesized in the cytoplasm can diffuse into the nucleus and bind to the target modification of endogenous histones. When the target modification level is high, mintbodies accumulate more in the nucleus since the free mintbodies can shuttle between the nucleus and cytoplasm. Based on the localization dynamics, relative changes in the modification level can be measured as the nucleus-to-cytoplasm intensity ratio. As a proof of concept, we have monitored dynamic changes of H4K20me1 during the cell cycle using the H4K20me1-specific mintbody [12]. H4K20me1 is an essential modification in mammals, involved in chromosome condensation, segregation, replication, and repair, as well as gene regulation [13, 14]. The level of H4K20me1 increases during the G2 to M phases mediated through a methyltransferase, PR-Set7/Set8, and the depletion of this enzyme causes mitotic defects [9, 15]. To monitor the cell cycle stage during interphase, proliferating cell nuclear antigen (PCNA) tagged with a fluorescent protein has been extremely useful because

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it is concentrated in DNA replication foci formed during the S phase [16]. Using cells expressing both the H4K20me1-mintbody and mCherry-PCNA, changes of H4K20me1 level were monitored by measuring the nucleus to cytoplasmic intensity ratios during the whole cell cycle (Fig. 1b) [12]. The level of H4K20me1 rapidly increased during the G2 and gradually decreased during the G1

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(Fig. 1b). The method to monitor global changes based on the nuclear to cytoplasmic intensity ratio of mintbody has been described in detail [17]. We here describe the usage of mintbodies to detect dynamic behavior of a specific structure, that is, the inactive X chromosome (Xi), during the cell cycle. Xi is a facultative heterochromatin known to be enriched in histone H3 Lys27 trimethylation (H3K27me3) and H4K20me1 [18–20]. While Xi can be visualized using the H4K20me1-mintbody, another mintbody specific for H3K27me3 is more enriched in Xi with high signal-to-background ratio. Using the H3K27me3-specific mintbody, we tracked the behavior of Xi territory during the cell cycle by 3D time-lapse imaging, combining with mCherry-PCNA as a cell cycle marker (Fig. 2). Image analysis then revealed the changes in the volume, shape, and H3K27me3 enrichment level of Xi during the cell cycle (Fig. 3). The H3K27me3 levels appeared to decrease during Xi replication and then increased late in the S phase though the G2 phase. The volume of Xi appeared to increase late in the S phase after Xi replication, while the changes in its shape did not appear to depend on the specific cell cycle. Thus, mintbodies can be useful for tracking specific chromosome structures in addition to the global changes of modifications during the cell cycle.

2 2.1

Materials Plasmids

1. PB533-2E12LI-NLS-sfGFP: a PB533-based plasmid expression vector that harbors a gene encoding the super-folder GFP (sfGFP) version of H3K27me3-mintbody with a nuclear localization signal [21]. PB533 is a PiggyBac Transposon system vector (System Biosciences). 2. PB533-Halo-PCNA: a PB533-based plasmid expression vector that harbors a gene encoding the proliferating cell nuclear antigen (PCNA) tagged with HaloTag. 3. PB210PA-1: a super PiggyBac Transposase expression vector (System Biosciences).

2.2 Cell Line and Medium

1. Mouse embryonic carcinoma MC12 cells [22]: cells harboring one (diploid) or two (tetraploid) inactive X chromosomes. A part of chromosome 2 is translocated near the telomere region of the inactive X chromosome. Cells are maintained at 37  C with 5% CO2 in a humidified incubator. 2. Culture medium: Dulbecco’s modified Eagle’s medium, highglucose (DMEM, high-glucose), supplemented with 10% fetal bovine serum, 100 U/mL streptomycin, and 100 μg/mL penicillin.

Xi Territory Tracked by H3K27me3-Mintbody

241

(A) Nuc Xi H3K27me3

(B)

Elapsed time (hh:mm:ss) Binary objects H3K27me3

0:31:38

1:51:43

7:41:34

10:01:41

13:21:40

14:11:50

Nuc Xi Early S

Mid S

Late S

G2

Prophase

PCNA

G1

(C) Xi (H3K27me3 foci) 30

3.5

Volume (µm³)

600

Elongation

20

2.5

500

10

1.5

400

G₁

0 0

G₁

0.5

2

4

6

8

10

12

14

Intensity (a.u.)

G₁

300

0

2

4

Time (hour)

6

8

10

12

14

0

2

4

Time (hour)

6

8

10

12

14

Time (hour)

Nucleus 1500

3

Volume (µm³)

1000

2

500

1

205

Elongation

Intensity (a.u.)

195 185

G₁

0 0

175

G₁

0 2

4

6

8

10

Time (hour)

12

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G₁

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4

6

8

10

Time (hour)

12

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Time (hour)

Fig. 2 Three-dimensional time-lapse imaging and analysis. (a) Representative volume view of H3K27me3NLS-mintbody signal in an MC12 cell and binary objects. Twenty-six z-planes with 0.5 m intervals were acquired. Three-dimensional binary objects for the inactive X chromosome (Xi, pink) and the nucleus (Nuc, Blue) were semi-automatically created. (b) Time-lapse images of H3K27me3-NLS-mintbody and JF646labeled Halo-PCNA. Twenty-six z-stack images were acquired every 10 minutes. Binary objects (3D) for the Xi

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3. Trypsin: 2.5 mg/mL trypsin dissolved in phosphate-buffered saline (PBS) containing 1 mM EDTA and sterilized by filtration. 2.3 Transfection Reagents and Inhibitor for Selection

1. FuGENE HD.

2.4 Microscopy and Image Analysis

1. 24-well glass bottom plate.

2. Opti-MEM I. 3. G418: 50 mg/mL G418 dissolved in water and sterilized by filtration.

2. 35-mm Glass Base Dish (Glass 12φ, No. 1-S). 3. Low-fluorescent medium: FluoroBrite™ DMEM (ThermoFisher Scientific), supplemented with 10% fetal bovine serum, 100 U/mL streptomycin, and 100 μg/mL penicillin. 4. Dimethylsulfoxide (DMSO; Molecular Biology Grade). 5. 200 μM Janelia Fluor 646 (JF646) HaloTag Ligand (Promega): Dissolve in DMSO.

Volume (a.u.)

Mean Intensity (a.u.)

Elongation (a.u.)

2

2

1.4

1.5

1.5

1.2

1

1

1

0.5

0.5

0.8

0

0.6

0

S phase

S phase

S phase

Fig. 3 Xi dynamics during the S phase. The volume, elongation, and mean intensity were measured as in Fig. 2. The relative values to the average and relative time points during the S phase were plotted (ten cells). The values were normalized using the average during the S phase. The relative time points during the S phase were obtained according to the method described in Subheading 3.3, step 11. Polynomial trend lines are shown as red dashed lines. The volume of Xi appears to increase late in the S phase after Xi replication, while the shape shows little fluctuation. The intensity appears to decrease during the replication of Xi ä Fig. 2 (continued) and Nuc were defined using H3K27me3-NLS-mintbody intensity. The cell cycle stages were judged by the Halo-PCNA localization pattern in the nucleus. Scale bar ¼ 10 m. (c) Quantified data. The volume, elongation (the ratio of the long and short axes of oval), and mean intensity of Xi and Nuc were measured and plotted. Moving averages of 15 time points are shown as red dashed lines

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6. A spinning disk confocal system (Yokogawa Electric; CSU-W1) attached to an inverted fluorescence microscope (Nikon Ti-E) equipped with a Plan Apo VC 100 Oil (NA 1.4) lens, an electron multiplying charge-coupled device (EM-CCD), a laser unit (488- and 640-nm) and a heat-stage chamber (Tokai Hit). 7. Operating and analysis software: NIS-Element ver. 4.30 for image acquisition and 5.21 for analysis (Nikon). 8. Excel (Microsoft).

3

Methods

3.1 Establishing Cell Lines Stably Expressing H3K27me3-NLSMintbody and Halo-PCNA

1. Plate MC12 cells in a well of a 6-well plate at ~20% confluency in 2 mL culture medium on the day before transfection. 2. Aliquot 100 μL of Opti-MEM I into an Eppendorf tube. 3. Add each 0.8 μg of PB533-2E12LI-NLS-sfGFP and PB533Halo-PCNA plasmids and 0.4 μg of PB210PA-1 to Opti-MEM I in a tube, mix well using a vortex mixer, and briefly spin down. 4. Add 6 μL FuGENE HD to Opti-MEM I–DNA mixture, mix well using a vortex mixer, and briefly spin down to collect the mixture at the bottom of the tube. 5. Leave the mixture for 5 min at room temperature and drop the mixture on to a well where cells were plated. 6. Incubate cells for a day. 7. Trypsinize cells and replate them in a 100 mm dish in cell culture medium containing 1 mg/mL G418. 8. After 1–2 weeks, pick up single colonies using a micropipette tip and transfer to wells containing 0.5 mL cell culture medium in a 24-well glass bottom plate. 9. Grow cells in the 24-well plate for 3–4 days. 10. Replace the medium with 0.5 mL Low-fluorescent medium containing 100 nM JF646 HaloTag Ligand and incubate for 1 h. 11. Wash the medium three times with 0.5 mL Low-fluorescent medium and add 0.5 mL Low-fluorescent medium after the final washing step. 12. Set the plate onto a heated stage on an inverted microscope with a confocal system (see Note 1). 13. Select the clones that show both green and far-red fluorescence under the microscope (see Note 2). 14. Expand cells and make frozen stock vials.

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3.2 Image Acquisition

1. Plate MC12 cells that express H3K27me3-NLS-mintbody and Halo-PCNA in 2 mL culture medium on a 35 mm glass base dish at 10–20% confluency on the day before observation. 2. Replace the medium with 1 mL Low-fluorescent medium containing 100 nM JF646 HaloTag Ligand and incubate for 1 h. 3. Wash the medium three times with 2 mL Low-fluorescent medium and add 2 mL Low-fluorescent medium. 4. Set the dish onto a heated stage on an inverted microscope with a confocal system. 5. Acquire 3D time-lapse images for sfGFP and JF646 labeled HaloTag using 488- and 640-nm laser lines combined with a dichroic mirror DM405/488/561/640 and emission filters 520/30 and 685/40, respectively. 6. To cover the whole nucleus in z-dimension, acquire 26 z-stack images with 0.5 μm intervals every 10 min for 24 h (see Note 3).

3.3

Data Analysis

1. Open 3D time-lapse image in NIS-elements AR. 2. Select the background ROI by choosing an area without cells. 3. Obtain the net intensity images by background subtraction. 4. Segment the nuclear region by thresholding and binarization using nuclear signals of H3K27me3-NLS-mintbody (see Note 4). 5. Segment the region of inactive X chromosome (Xi) by thresholding and binarization using signals of H3K27me3-NLSmintbody enriched on Xi (see Note 5). 6. Make 3D binary objects for nuclei and Xi foci using “Connect 3D object” tool in “3D object measurement” module (Fig. 2a). 7. Measure the Volume, Elongation (the ratio of long and short axes of oval), and Mean intensity using “3D object measurement”. 8. Export the data to Excel. 9. Open the exported data in Excel and create scatterplot graph. 10. Determine the cell cycle stage assisted by the localization pattern of Halo-PCNA signal in the nucleus. Halo-PCNA becomes accumulated in numerous foci in the Early S phase. As the cell progresses to the Mid S phase, the foci become larger and localize preferentially around the nuclear periphery. During this stage Halo-PCNA is also concentrated on Xi. In the Late S phase, the Halo-PCNA foci become much larger and separate from Xi foci. The Halo-PCNA foci disappear in G2 phase (Fig. 2b, c).

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11. To analyze the behavior of Xi during S phase in multiple cells with different S phase lengths, normalize the S phase period using the S-phase beginning time (at which Halo-PNCA foci first appeared) and the S-phase ending time (at which HaloPCNA foci disappeared) using the following formula. T0 ¼ (T  Tini)/(Tend  Tini), where T0, normalized time; T, actual time; Tini, a time point when the S phase initiated; and Tend, a time point when the S phase ended. 12. Plot all relative values to see the changes in individual cells and trends (Fig. 3).

4

Notes 1. As a confocal microscope is eventually used for time-lapse imaging, we screen cells that exhibit fluorescence using the same microscope. However, for the initial screening to quickly check if cells exhibit fluorescence signals, a convenient widefield microscope can be used. 2. Select cells that exhibit bright fluorescence and look healthy. Although mintbody expression is usually not toxic to cells, very bright cells may show slow growth. Such slowly growing cells should not be chosen. Three to four out of 24 clones are double-positive on average. 3. The time duration and intervals depend on the purpose. Since doubling time of MC12 cells is less than 20 h, time-lapse imaging for ~24 h is enough to obtain the whole cell cycle. To analyze the dynamics during the interphase, imaging every 10 min is usually sufficient. To follow chromosome movements during mitosis in detail, rapid imaging with short intervals are needed. 4. If the shape of the binary object is not accurate, modify it using drawing tools on “Binary Toolbar”. “segment.ai” of NIS.ai module can be helpful. 5. The intensity of H3K27me3 enriched on Xi is ~2 times higher than that of the nucleus.

Acknowledgments We thank Akito Ohi at Kimura lab (Tokyo Tech) for constructing H3K27me3-specific mintbody, M.C. Cardoso (Technische Universita¨t Darmstadt) for providing the PCNA-GFP construct, N. Takagi (Hokkaido University) for supplying MC12 cells, and the Open Facility Center, Tokyo Institute of Technology for DNA sequencing analysis. The authors work was supported by Japan

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Society for the Promotion of Science KAKENHI grants JP17KK0143 and JP20K06484 (to Y.S.) and JP17H01417 and JP18H05527 (to H.K.), and Japan Science and Technology Agency (JST) CREST JPMJCR16G1 (to H.K.) and JPMJCR20S6 (to Y.S.). References 1. Kimura H (2013) Histone modifications for human epigenome analysis. J Hum Genet 58 (7):439–445. https://doi.org/10.1038/jhg. 2013.66 2. Zhao Y, Garcia BA (2015) Comprehensive catalog of currently documented histone modifications. Cold Spring Harb Perspect Biol 7 (9):1–21. https://doi.org/10.1101/ cshperspect.a025064 3. Gorkin DU, Barozzi I, Zhao Y et al (2020) An atlas of dynamic chromatin landscapes in mouse fetal development. Nature 583 (7818):744–751. https://doi.org/10.1038/ s41586-020-2093-3 4. Zee BM, Levin RS, Xu B, Leroy G, Wingreen NS, Garcia BA (2010) In vivo residue-specific histone methylation dynamics. J Biol Chem 285(5):3341–3350. https://doi.org/10. 1074/jbc.M109.063784 5. Xu M, Wang W, Chen S, Zhu B (2012) A model for mitotic inheritance of histone lysine methylation. EMBO Rep 13(1):60–67. https://doi.org/10.1038/embor.2011.206 6. Revero´n-Go´mez N, González-Aguilera C, Stewart-Morgan KR, Petryk N, Flury V, Graziano S, Johansen JV, Jakobsen JS, Alabert C, Groth A (2018) Accurate recycling of parental histones reproduces the histone modification landscape during DNA replication. Mol Cell 72(2):239–249.e5. https:// doi.org/10.1016/j.molcel.2018.08.010 ˜ a-Meyer R, 7. Escobar TM, Oksuz O, Saldan Descostes N, Bonasio R, Reinberg D (2019) Active and repressed chromatin domains exhibit distinct nucleosome segregation during dna replication. Cell 179(4):953–963.e11. https://doi.org/10.1016/j.cell.2019.10.009 8. Hayashi-Takanaka Y, Kina Y, Nakamura F, Becking LE, Nakao Y, Nagase T, Nozaki N, Kimura H (2020) Histone modification dynamics as revealed by a multicolor immunofluorescence-based single-cell analysis. J Cell Sci 133:jcs243444. https://doi.org/10. 1242/jcs.243444 9. Pesavento JJ, Yang H, Kelleher NL, Mizzen CA (2008) Certain and progressive methylation of histone H4 at lysine 20 during the cell

cycle. Mol Cell Biol 28(1):468–486. https:// doi.org/10.1128/MCB.01517-07 10. Hendzel MJ, Wei Y, Mancini MA, Van Hooser A, Ranalli T, Brinkley BR, BazettJones DP, Allis CD (1997) Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106(6):348–360. https://doi.org/10.1007/s004120050256 11. Sato Y, Mukai M, Ueda J, Muraki M, Stasevich TJ, Horikoshi N, Kujirai T, Kita H, Kimura T, Hira S, Okada Y, Hayashi-Takanaka Y, Obuse C, Kurumizaka H, Kawahara A, Yamagata K, Nozaki N, Kimura H (2013) Genetically encoded system to track histone modification in vivo. Sci Rep 3:2436. https:// doi.org/10.1038/srep02436 12. Sato Y, Kujirai T, Arai R, Asakawa H, Ohtsuki C, Horikoshi N, Yamagata K, Ueda J, Nagase T, Haraguchi T, Hiraoka Y, Kimura A, Kurumizaka H, Kimura H (2016) A genetically encoded probe for live-cell imaging of H4K20 monomethylation. J Mol Biol 428(20):3885–3902. https://doi.org/10. 1016/j.jmb.2016.08.010 13. Beck DB, Oda H, Shen SS, Reinberg D (2012) PR-Set7 and H4K20me1: at the crossroads of genome integrity, cell cycle, chromosome condensation, and transcription. Genes Dev 26 (4):325–337. https://doi.org/10.1101/gad. 177444.111 14. Jørgensen S, Schotta G, Sorensen CS (2013) Histone H4 lysine 20 methylation: key player in epigenetic regulation of genomic integrity. Nucleic Acids Res 41(5):2797–2806. https:// doi.org/10.1093/nar/gkt012 15. Houston SI, McManus KJ, Adams MM, Sims JK, Carpenter PB, Hendzel MJ, Rice JC (2008) Catalytic function of the PR-Set7 histone H4 lysine 20 monomethyltransferase is essential for mitotic entry and genomic stability. J Biol Chem 283(28):19478–19488. https://doi.org/10.1074/jbc.M710579200 16. Leonhardt H, Rahn HP, Weinzierl P, Sporbert A, Cremer T, Zink D, Cardoso MC (2000) Dynamics of DNA replication factories

Xi Territory Tracked by H3K27me3-Mintbody in living cells. J Cell Biol 149(2):271–280. https://doi.org/10.1083/jcb.149.2.271. 17. Sato Y, Kimura H (2017) Semi-quantitative analysis of H4K20me1 levels in living cells using mintbody. Bio-Protocol 7(10):e2276. https://doi.org/10.21769/BioProtoc.2276 18. Silva J, Mak W, Zvetkova I, Appanah R, Nesterova TB, Webster Z, Peters AHFM, Jenuwein T, Otte AP, Brockdorff N (2003) Establishment of histone H3 methylation on the inactive X chromosome requires transient recruitment of Eed-Enx1 polycomb group complexes. Dev Cell 4(4):481–495. https:// doi.org/10.1016/s1534-5807(03)00068-6. 19. Plath K, Fang J, Mlynarczyk-Evans SK, Cao R, Worringer KA, Wang H, de la Cruz CC, Otte AP, Panning B, Zhang Y (2003) Role of histone H3 lysine 27 methylation in X inactivation. Science 300(5616):131–135. https:// doi.org/10.1126/science.1084274 20. Kohlmaier A, Savarese F, Lachner M, Martens J, Jenuwein T, Wutz A (2004) A

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chromosomal memory triggered by xist regulates histone methylation in X. PLoS Biol 2(7): E171. https://doi.org/10.1371/journal. pbio.0020171 21. Tjalsma SJD, Hori M, Sato Y, Bousard A, Ohi A, Raposo AC, Roensch J, Saux AL, Nogami J, Maehara K, Kujirai T, Handa T, Bage´s-Arnal S, Ohkawa Y, Kurumizaka H, da Rocha ST, Z˙ylicz JJ, Kimura H, Heard E (2021) H4K20me1 and H3K27me3 are concurrently loaded onto the inactive X chromosome but dispensable for inducing gene silencing. EMBO Rep 22:e51989 https://doi. org/10.15252/embr.202051989 22. Abe K, Takagi N, Sasaki M (1988) Nonhistone nuclear proteins specific to certain mouse embryonal carcinoma clones having an inactive X chromosome. Exp Cell Res 179 (2):590–594. https://doi.org/10.1016/ 0014-4827(88)90297-2

Chapter 18 Analyzing Centrioles and Cilia by Expansion Microscopy Dong Kong and Jadranka Loncarek Abstract Expansion microscopy is an imaging method based on isotropic physical expansion of biological samples, which improves optical resolution and allows imaging of subresolutional cellular components by conventional microscopes. Centrioles are small microtubule-based cylindrical structures that build centrosomes and cilia, two organelles essential for vertebrates. Due to a centriole’s small size, electron microscopy has traditionally been used to study centriole length and ultrastructural features. Recently, expansion microscopy has been successfully used as an affordable and accessible alternative to electron microscopy in the analysis of centriole and cilia length and structural features. Here, we describe an expansion microscopy approach for the analysis of centrioles and cilia in large populations of mammalian adherent and nonadherent cells and multiciliated cultures. Key words Expansion microscopy, Centriole, Centrosome, Centriole length, Cilia

1

Introduction The size of most cellular organelles is beyond the attainable spatial resolution of light microscopy, which is ~220 nm in lateral and ~550 nm in axial dimensions. This obstacle prevents studying the morphology of most organelles and the spatial and dynamic properties of their components. To decipher the ultrastructural features of cellular organelles, electron microscopy has been widely used in biological sciences since the 1950s. However, electron microscopy is a tedious and specialized technique that does not provide information about protein localization. Thus, a class of superresolution imaging techniques has been invented to overcome the resolution barrier of conventional light microscopy and to enable imaging of fluorescently labeled cellular components in nanoscale resolution [1–3]. Recently expansion microscopy has been developed to circumvent the problem of limited optical resolution, by physically increasing the size of the specimen [4–8]. Expansion microscopy is

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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based on embedding a specimen in a swellable polymer and crosslinking the specimen components or fluorescent moieties to it. The polymer is then physically expanded in water, resulting in isotropic expansion of the components of the specimen linked to the polymer. Increased physical distance between components of the expanded specimen allows their imaging by conventional microscopes with increased resolution. Expansion protocols follow one of the two different original approaches developed by the Boyden [5] and Chung [7] laboratories. In the first approach, the sample is first immunolabeled, immunofluorescent molecules are chemically crosslinked to the polymer, and proteins are enzymatically digested before expansion. In the second approach, specimen components are cross-linked to the polymer, proteins are denatured by high temperature and sodium dodecyl sulfate (SDS), immunolabeled, and the polymer linked to immunolabeled proteins is expanded. Centrioles are cellular structures that build centrosomes and cilia. They are organized as ninefold symmetrical cylinders comprising nine sets of microtubule singlets, doublets or triplets (Fig. 1) [9–11]. They can range from 200 nm to several microns long and are ~230 nm wide. In addition to microtubules, centrioles are comprised of dozens of proteins and dozens of associated pericentriolar components, which extend further from the centriole (Fig. 1) [12–15]. Structural, numerical, and biochemical aberrations of centrioles are associated with pathologies such as cancer and a group of hereditary diseases known as ciliopathies [16–19]. Centriole and centrosome size are close to or below the diffraction limit of classical light microscopy, and they are challenging objects for classical optical microscopy, which cannot be used to precisely determine their length or the intricate patterns of their components. Here we describe an expansion protocol (based on the original protocol from Chung laboratory), adapted for the analysis of centrioles and cilia in large populations of cultured cells using conventional microscopy [20]. This approach can be affordably employed in any typical cell biology laboratory and can be used as an alternative to electron and superresolution microscopy for studying centriole length, number, duplication, structural features, ciliation, and localization of various centrosomal components [20, 21].

2 2.1

Materials Cell Culture

1. Mammalian cells grown in suspension or adherent to a coverslip or petri dish or Transwell-Clear permeable filter supports. 2. 25 mm round glass coverslips, thickness # 1.5.

Centriole Analysis by Expansion Microscopy

Mother centriole

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Centriole nine triplet microtubules; cross section, view from the distal end.

Distal appendage Sub distal appendage

~500 nm

Centriole lumen; distal end Centriole triplet microtubule; longitudinal section Procentriole formed adjacent to the mother centriole Cartwheel Pericentriolar material Centriole lumen; proximal end

An interphase cell with two duplicated centrioles positioned near the nucleus (blue) and associated with microtubules (black lines).

Fig. 1 Organization of a human centriole/centrosome. Scheme shows organization of a typical duplicated mother centriole associated with a procentriole. Human centrioles are built of nine microtubule triplets organized in a perfect rotational symmetry. Mother centrioles are, on their proximal end, associated with a proteinaceous complex called pericentriolar material, which is the site of many centrosomal functions. A centriole with associated pericentriolar material is called a centrosome. On their distal end, fully assembled mother centrioles harbor distal appendages and subdistal appendages, which mediate cilia assembly and microtubule anchoring, respectively. Centrioles propagate by duplication, during which a new procentriole forms in association with the proximal end of the mother centriole. Proximal ends of procentriole contains a cartwheel, which promotes ninefold organization of procentrioles. A typical somatic cell has only two mother centrioles, which can be unduplicated (in G1), or duplicated (in S, G2, and mitosis)

3. Poly-L-lysine: Reconstitute poly-L-lysine in dH2O to 1 mg/ mL. To coat coverslips, add poly-L-lysine solution to the coverslips, incubate for 5 min at room temperature (RT), remove poly-L-lysine, wash coverslips in dH2O, and allow them to dry at RT. Solution can be reused 3–4 times. For long-term storage aliquot and store at  20  C. 4. Cell culture incubator with 5% CO2. 2.2 Expansion Reagents, Buffers, and Polymerization Mixture

1. 10 PBS, pH 7.4. 2. 37% Formaldehyde (Electron Microscopy Sciences; 15686). 3. Fixing solution: 4% formaldehyde in 1 PBS. Prepare fresh. 4. 40% acrylamide solution (Sigma; A4058). 5. Formaldehyde–acrylamide solution: 6 mL 40% acrylamide solution, 0.86 mL 37% formaldehyde, 0.8 mL 10 PBS, and 0.34 mL dH2O. Prepare fresh.

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6. 40% acrylamide–bisacrylamide A9926).

solution

(19:1)

(Sigma;

7. Sodium acrylate (Sigma; 408220): Once opened, keep in desiccator. 8. Ammonium persulfate. 9. TEMED. 10. Polymerization mixture (20% acrylamide, 0.04% bisacrylamide, 7% sodium acrylate, 0.5% TEMED, 0.5% ammonium persulfate, in 1 PBS). Prepare fresh. This solution is prepared in four stages. Stage 1. Weigh 280 mg of sodium acrylate in a 2 mL microcentrifuge tube 1, add 1.440 mL of 2 PBS, vortex, and keep on ice (the final volume is ~1.580 mL). Stage 2. Prepare a fresh stock of 5% ammonium persulfate: weigh 50 mg of ammonium persulfate into a 2 mL microcentrifuge tube, add 1 mL of 2 PBS. Vortex and keep on ice. Stage 3. In a precooled 15 mL tube, combine the entire content of tube 1 (~1.580 mL of sodium acrylate solution) with 1.920 mL of 40% acrylamide solution, 0.08 mL of 40% acrylamide–bisacrylamide solution, and 0.02 mL of TEMED. Mix and keep on ice. Stage 4. Finalize the polymerization mixture by adding 0.4 mL of a cold 5% ammonium persulfate stock. Vortex, and keep on ice to slow polymerization. At this point the polymerization mixture is complete and needs to be added to the precooled coverslip with cells as soon as possible. 11. SDS buffer (for boiling): 129.3 mL dH2O, 20 mL 4 N NaCl, 20 mL 1 M Tris–HCl (pH 9.0), and 230.7 mL 10% SDS (346.8 mM). Keep at RT. 2.3 Expansion Tools and Equipment

1. 4 mm biopsy puncher (Integra; 33-34-P/25). 2. Vacuum pick-up tool (Ted Pella; 528-2). 3. Reverse tweezers. 4. Flat-tip tweezers. 5. Cell strainer that fits a 50 mL conical tube, no specific pore size. 6. Incubator at 40  C. 7. Plasticware: 35, 60, and 100 mm petri dishes, 2 and 5 mL microcentrifuge tubes, 15 and 50 mL conical tubes. 8. Parafilm. 9. 2 and 0.5 L glass beaker. 10. Thermometer.

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11. Timer. 12. Ice bucket with a lid. 2.4 Immunofluorescent Labeling

1. Immunofluorescence (IF) buffer: 1% BSA, 0.05% Tween-20 in 1 PBS. Filter using 0.22 μm filter, and store at 4  C. 2. DAPI (ThermoFisher Scientific; D1306): Dissolve 10 mg in 2 mL of dH2O. Vortex well. Aliquot and store for long-term storage at  20  C. Take one aliquot and further dilute tenfold to prepare 1 stock solution (0.5 mg/mL), which can be kept at 4  C for several months. 3. Primary and secondary antibodies: listed in Tables 1 and 2.

3

Methods In this chapter, we describe a procedure for expansion and microscopy analysis of centrioles and centrosomes in cultured cells attached to coverslips (due to their self-adhering properties, or after their interaction with a poly-L-lysine coating). However, the same procedure can be used for cells grown in Transwells or any petri dish (see Note 1). Expansion of samples includes nine major steps which are delineated in Fig. 2. Each step of the protocol illustrated in Fig. 2 is described in detail in one of the nine sections of this chapter.

3.1 Sample Preparation and Fixation

1. Plate self-adherent cells on 25 mm round glass coverslips and prepare them according to experimental needs. Or collect nonadherent or mitotic cells and adhere them to the round 25 mm coverslip precoated with poly-L-lysine. To do this, centrifuge cells 3 min at 160  g, resuspend them in a small volume of growth medium or 1 PBS, load them to poly-L-lysine coated coverslips, and fix ~1 min later. 2. Fix cells in freshly prepared fixing solution for 1 h at RT. After fixation, samples can be stored for several hours in 4% formaldehyde before proceeding to Subheading 3.2. Note that previously immunolabeled and imaged samples can also be expanded using this method (see Note 1). 3. For expansion of multiciliated cells grown on Transwell-Clear permeable filter supports, remove the filter from the well using a scalpel and place it on the surface of the coverslip with the cells facing up. Add fixing solution directly to the cells. Alternatively, cells on the filters can be fixed by immersing the filter in the fixing solution in a tube (remain mindful which side contains cells).

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Table 1 A list of commercially available primary antibodies with tested compatibility with this protocol Protein

Dilution

Vendor (host species, catalog number)

Acetylated tubulin

1:4000

Sigma (mouse, T7451)

Polyglutamylation

1:800

Adipogen (rabbit, AG-25B-0030-C050)

γ-Tubulin

1:500

Sigma (mouse, T5326)

α-Tubulin

1:500

Sigma (mouse, T6074)

β-Tubulin

1:250

Abcam (rabbit, ab15568)

SAS-6

1:200

Santa Cruz (mouse, sc-81431)

CP110

1:1000

Proteintech (rabbit, 12780-1-AP)

Cep290

1:600

Abcam (rabbit, ab84870)

Cep152

1:1000

Bethyl (rabbit, A302-479A)

Cep63

1:200

Millipore (rabbit; 06-1292)

Pericentrin

1:400

Abcam (rabbit, ab4448)

Centrobin

1:200

Abcam (mouse, ab70448)

CCDC41

1:100

Sigma (rabbit, HPA038161)

SCLT1

1:100

Sigma (rabbit, HPA036561)

FBF1

1:100

Sigma (rabbit, HPA023677)

Cep164

1:500

Proteintech (rabbit, 22227-1-AP)

ANKRD26

1:200

GeneTex (rabbit, GTX128255)

ODF2

1:150

Proteintech (rabbit, 12058-1-AP)

ARL13B

1:150

Abcam (rabbit, ab83879)

RPGRIP1L

1:150

Proteintech (rabbit, 55160-1-AP)

IFT88

1:150

Proteintech (rabbit, 13967-1-AP)

Rootletin

1:50

Santa Cruz (mouse, sc-374056)

3.2 Incubation with Acrylamide and Formaldehyde

1. After fixation, transfer each coverslip to a 35 mm petri dish and add 2 mL of freshly made formaldehyde–acrylamide solution (see Note 2). Scale up the amount of formaldehyde–acrylamide solution depending on the number of coverslips that need to be covered, 8 mL suffices for four coverslips. 2. Swirl the petri dishes with coverslips several times. Formaldehyde–acrylamide solution should evenly cover the coverslips. 3. Seal petri dishes with Parafilm and incubate the samples in a humid incubator at 40  C for 16 h.

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Table 2 A list of goat secondary antibodies with tested compatibility with this protocol Protein

Dilution

Vendor; catalog number

Anti-mouse Alexa Fluor 488

1:800

Thermo Fisher Scientific; A11029

Anti-rabbit Alexa Fluor 488

1:800

Thermo Fisher Scientific; A11034

Anti-mouse Alexa Fluor 555

1:800

Thermo Fisher Scientific; A28180

Anti-rabbit Alexa Fluor 555

1:800

Thermo Fisher Scientific; A21429

CF647 anti-mouse antibody

1:800

Biotium; 20042

CF647 anti-rabbit antibody

1:800

Biotium; 20045

Step 1. Fixation

Step 2. Incubation in acrylamide and formaldehyde

Step 3. Polymerization

Step 4. Cutting into smaller pieces

Step 5. Denaturation in hot SDS solution

Step 9. Sample mounting and Imaging

Step 8. Expansion in dH2O

Step 7. Immunolabeling

Step 6. SDS washout

Fig. 2 Key steps in sample preparation for expansion microscopy. A scheme illustrates major steps of the protocol, as described in the subheadings of Subheading 3. Cells growing on the coverslip are fixed and then incubated in a mixture of acrylamide and formaldehyde solution, which establishes chemical groups required for crosslinking of the sample components to the polymer during subsequent polymerization step. After polymerization, the sample is cut to several smaller samples. Individual samples (or ‘punches’ if a biopsy puncher is used to excise smaller pieces of the gel) are boiled in the presence of high SDS concentration, to denature proteins and to allow expansion of various cellular structures. During this time, the gel, and the biological specimen within, expands ~2-fold. In the subsequent step, SDS is thoroughly washed out of the gel, which is required for efficient immunolabeling. Proteins of interest are then immunolabeled with primary and secondary antibodies, and DNA is labeled using DAPI. After immunolabeling, samples are gradually expanded by incubation in dH2O 3.3

Polymerization

1. After incubation at 40  C, transfer each coverslip(s) containing cells to a clean 60-mm petri dish. 2. Wash the coverslips three times with 1 PBS, 10 min each time at RT.

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Fig. 3 Polymerization and punching. (a) A 100-mm petri dish with its bottom layered by Parafilm on the top of an ice–water bath. Two coverslips with cells facing up are covered with polymerization mixture. Samples are kept on ice during the first 20 min of polymerization, followed by incubation at RT for another 1–2 h (b). (c) A coverslip containing cells and polymerized gel before (left) and after (right) samples have been excised using a 4-mm biopsy puncher (green, below). Multiple individual samples can be excised from one gel. (d) Punching and transferring of a punch to a 50 mL conical tube

3. While washing the cells, layer the bottom of a 100-mm petri dish with Parafilm (Fig. 3a). 4. Transfer the coverslip(s) with cells to Parafilm-coated petri dish with cells facing up. 5. Add ~0.5 mL of 1 PBS to each coverslip to prevent drying of the sample. 6. Prepare ice–water bath by filling an ice bucket with ice and water (Fig. 3a). 7. Place the Parafilm-coated petri dish into the ice–water bath to cool down coverslips with cells for ~20 min (Fig. 3a). The ice– water bath ensures good contact with the bottom of the dishes and thus efficient cooling. 8. Add complete polymerization mixture (see Note 3) to the precooled coverslip with cells following steps 9–12. Try to complete steps 9–12 as quickly as possible. Samples need to remain cold. 9. Remove 1 PBS from the sample by tilting the Parafilm-coated dish with samples. 10. Aspirate residual 1 PBS from the edge of the coverslip. Try to remove as much 1 PBS as possible.

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11. Add 1 mL of complete polymerization solution on the top of each coverslip, and quickly pipette up and down at three different spots along the coverslip edge, to mix the polymerization solution with residual 1 PBS. 12. Remove ~0.5 mL of polymerization mixture, leaving behind ~0.5 mL on each coverslip (Fig. 3a). 13. Keep the samples with the polymerization mixture on ice for 20 min with the lid on the petri dish (polymerization should start in about 5 min). Put the lid on the ice bucket to maintain cold environment. 14. Transfer the petri dish containing samples to a flat surface and additionally incubate for ~1–2 h at RT (Fig. 3b). If polymerization was successful, coverslips will be coated with a layer of a flexible gel that does not break and is easy to handle (see Note 4). 3.4 Cutting (Punching) the Sample into Multiple Smaller Samples

1. With a 4-mm biopsy puncher, carefully cut out several gel punches from each coverslip (Fig. 3c, d). Cells will be detached from the coverslip and remain in the gel. 2. Place punches to the bottom of an empty 50 mL conical tube (Fig. 3d) (see Notes 5 and 6). Gel punches should be sturdy enough for gentle handling with a flat-tip tweezer or by a vacuum pick-up tool (our recommended way of handling punches). 3. Gently push punches to the bottom of the 50 mL tube (Fig. 4a). 4. Multiple punches can be cut out from one 25-mm round coverslip (Fig. 3c), if the cells were evenly plated across the coverslip. After polymerization, cells can be easily observed within the gel using a cell-culture microscope.

3.5 Denaturation by Boiling in SDS Buffer

1. Assemble the boiling equipment (Fig. 4b) as shown in Fig. 4c: place a smaller 500 mL beaker inside a larger, 2 L, beaker with a plastic tip-rack underneath the small beaker. 2. Fill both beakers with water to ~50% of their volumes. 3. Cover the large beaker (for instance with a polystyrene tube rack) and bring the temperature of the water bath >90  C. 4. To monitor the real-time temperature inside, put a thermometer through the polystyrene tube rack into the water of the small beaker (Fig. 4c). 5. Preheat ~25 mL SDS buffer in a 50 mL conical tube in the water bath. 6. When the temperature in the water bath reaches >90  C, place the 50 mL conical tube with gel punches in the water bath and incubate for 10 min (dry heat the punches).

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Fig. 4 Denaturation by boiling in SDS buffer. (a) One 50 mL conical tube with multiple gel punches before boiling. (b) Equipment used for boiling of the samples. (c) Assembled boiling equipment. Large and small beaker are filled with water. A polystyrene tube-rack and weight serve to prevent evaporation and splashes during boiling. A thermometer is used to monitor the real-time temperature of the water bath. Samples are heated in 50 mL conical tubes

7. Add preheated 25 mL SDS buffer to the 50 mL conical tube containing punches and swirl. 8. Place the 50 mL tube containing punches and SDS buffer into the water bath. Boil the punches in SDS buffer for 1 h, with gentle swirling every 10 min. 9. After boiling, remove 50 mL tubes from water bath and allow the SDS buffer with punches to cool down to RT. 3.6

SDS Washout

1. Remove SDS buffer by pipetting or using a cell strainer. 2. Add ~40 mL 1 PBS to 50 mL tube. 3. Place the tube on a shaking platform and gently shake for 20 min at RT. Repeat washing nine more times at RT. After SDS washout, the size of punches will be ~2-fold larger than before boiling (Fig. 5a). 4. Leave the punches in 1 PBS overnight on a shaking platform at 4  C and follow by several more washes the next day at RT (see Note 7).

3.7

Immunolabeling

1. Transfer washed punches to 60-mm petri dish containing 1.5 mL of 1 PBS (for instance, collect them using a cell strainer and flip the strainer over the petri dish). 2. Using a vacuum pick-up tool or a flat-tip tweezer, carefully transfer one punch into a 2 mL round bottom microcentrifuge tube (or if more punches are to be immunolabeled with the same antibodies, transfer 2–3 punches to a 5 mL microcentrifuge tube).

Centriole Analysis by Expansion Microscopy

a

b

After gelation and punching

After SDS washout in 1xPBS

After overnight expansion in dH2O

3.9 mm (1x)

8.6 mm (~2.2x)

16.2 mm (~4.2x)

d

c Attofluor cell chamber with expanded gel

Glass bottom dish with expanded gel

e

Upper coverslip Upper metal plate

259

Screw Expanded gel

Expanded gel on lower coverslip

Rubber gasket Lower metal plate

Top view

Lower coverslip

Objective Side view

Top view

Fig. 5 Expansion of immunolabelled samples and mounting of the samples for imaging. (a) Examples of gel punches at three different stages of the protocol. Left: After punching. Middle: after SDS wash-out. Right: after expansion in dH2O. (b) Attofluor chamber with an expanded punch. (c) A glass bottom dish with an expanded punch. (d) Left: A Rose chamber with an expanded punch and filled with dH2O assembled for imaging. Right: A scheme illustrates an assembled Rose chamber as viewed from the side (for more details see Subheading 3.9). Rose chambers are ideal for imaging of expanded gels because both surfaces of the gel can be accessible for imaging by flipping the chamber. This is particularly useful if there is ambiguity as to which surface contains a layer of cells. In addition, the gel is secured between two coverslips, which prevents moving of the gel during imaging. (e) Example of an expanded punch placed in the Rose chamber before addition of dH2O and closing of the chamber with the upper coverslip and the upper metal plate. Please note that the volume of polymerization solution (Subheading 3.3) needs to be adjusted to yield the gel thickness optimal for immunostaining and imaging. Alternatively, excess gel can be cut off using a scalpel

3. Add ~0.5 mL of IF buffer per punch to the microcentrifuge tube and block for 1–2 h at RT with gentle shaking. 4. Dilute primary antibody (or multiple antibodies if costaining) in IF buffer. Then add 0.3–0.5 mL of the antibody solution per punch and incubate at RT for ~2 h, followed by incubation at 4  C for 24–48 h, with gentle shaking (see Note 8). 5. Wash primary antibody in 1 PBS for 1–2 h with gentle shaking, exchanging 1 PBS every 10 min. 6. Dilute secondary antibody(ies) at 1:800 in IF buffer. Use ~0.5 mL of diluted antibody per punch. 7. Add DAPI from 1 stock solution to final dilution of 1:1000.

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8. Incubate punches with secondary antibodies/DAPI solution for 1–2 h at RT, followed by incubation at 4  C for 24 h, with gentle shaking. 3.8 Expansion of Immunolabeled Samples in dH2O

1. Start expanding immunolabeled gel punches by transferring punches to a petri dish or 50 mL tube. 2. Add dH2O to wash and expand punches for the first 2 h at RT, exchanging dH2O every 10 min. 3. Continue expanding overnight at 4  C with gentle agitation (see Note 9). Monitor expansion by measuring the size of the punches (Fig. 5a).

3.9 Sample Mounting and Imaging

4

To prevent gels from shrinking, imaging needs to be performed in dH2O and all imaging dishes need to be thoroughly cleaned and rinsed with dH2O. For imaging, punches can be mounted in Attofluor cell chambers or in any glass-bottom petri dishes (Fig. 5b, c). However, we prefer mounting expanded samples in a homemade variation of a Rose Chamber (Fig. 5d) [22]. The chamber consists of two metal hollow plates, and one silicone rubber gasket. In the assembled chamber, above and below the rubber gasket is a coverslip (Fig. 5d, e). The space between two coverslips creates an imaging chamber which, in this case, will hold an expanded gel and dH2O. To prevent sliding of the expanded punches during imaging, coverslips or petri dishes can be coated with poly-L-lysine. Imaging of expanded samples can be performed on any conventional microscope [20, 21]. Oil objectives of high magnifications (60 and 100) can also be used (Fig. 6), especially if the structure of interest is localized close to the coverslip. Imaging of centrioles and cilia localized deeper in cells and mitotic cells is also possible using oil immersion, but it is facilitated using water immersion objectives. To achieve increased resolution, other imaging modalities such as structured illumination microscopy can be used [20].

Notes 1. The protocol described here can also be used for expansion of previously immunolabeled cells that were previously fixed in cold methanol for 5 min or fixed in 1.5% formaldehyde for 4 min followed by cold methanol for 5 min [20]. Previously immunolabeled cells should not be embedded in hardening mounting medium prior to expansion. The best expansion results will be obtained if immunolabeled samples are kept in 1 PBS during imaging and are expanded as soon as possible. Before expansion, immunolabeled samples need to be postfixed with 4% formaldehyde in 1 PBS for 1 h at RT (that is, starting

Centriole Analysis by Expansion Microscopy

a

b

c

Prophase, mIMCD3 S phase, mIMCD3 DNA Acetylated tubulin

261

d

Prometaphase, mIMCD3 Telophase, U2OS

Fig. 6 Examples of expanded centrioles and cilia using herein described method. Cells were grown on coverslips, expanded ~4.2, and immunolabeled using an antibody recognizing acetylated tubulin. DNA is labeled with DAPI. (a) An interphase cell with two mother centrioles (pink arrow) associated with procentrioles (green arrow). One mother centriole is ciliated (blue arrow is pointing to the ciliary axoneme). (b–d) Examples of cells in various stages of mitosis. Each mother centriole (pink arrows) is associated with one procentriole (green arrows). Centrioles are in various orientations. Scale bars: 20 and 2 μm for the inserts

from Subheading 3.1). As an alternative to round 25 mm coverslips, cells can be grown on the coverslips of any size and in various types of petri dishes. However, the volume of polymerizing mixture (Subheading 2.2) will need to be adjusted, to achieve an optimal gel thickness of ~1–2 mm (before expansion). 2. After fixation in 4% formaldehyde, do not wash the samples. Remove the fixative and directly add freshly made formaldehyde–acrylamide solution. 3. Please note that the recommended polymerization mixture (20% acrylamide, 0.04% bisacrylamide, 7% sodium acrylate, 0.5% TEMED, 0.5% ammonium persulfate, in 1 PBS) will result in ~4.2-fold expansion of the gel/sample. The expansion factor can be changed by varying the concentration of bisacrylamide while, keeping the concentrations of other components constant. In our hands, the polymerization mixtures with the final percentages of bisacrylamide of 0.02%, 0.04%, 0.1%, and 0.2%, result in the ~5-, 4.2-, 3.6-, and 3.2-fold expansion, respectively [20]. Scale up the amount of polymerization mixture depending on the number of coverslips (4 mL is enough for four coverslips). However, try not to handle more than four coverslips at a time during the polymerization step of the protocol (Subheading 3.3). 4. All the reagents (2 PBS, 40% acrylamide solution, 40% acrylamide–bisacrylamide, sodium acrylamide, ammonium persulfate, and TEMED) and tubes must be precooled before finalizing the polymerization mixture, otherwise the gel may polymerize too fast and unevenly, causing uneven expansion of

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the samples later on. After polymerization, a gel may have ruffled edges, and sometimes may detach from the coverslip and curl. This is not unusual, and it will not affect expansion. Flat-tip tweezers and a biopsy puncher can be used to gently unravel curled gels before punching or cutting. 5. If cells were evenly plated across the coverslip, multiple punches/pieces can be punched/cut out of one 25-mm round coverslip (Fig. 3d). Since the distance between cells also increases during expansion, it is important to start the experiment with optimal cell confluency. Low confluency will require immunolabeling of multiple punches and require more material (such as antibodies) and labor. On the other hand, a cell confluency that is too high may result in rounding of the cells and in centrioles being positioned further from the coverslip, which is not optimal for subsequent imaging. Thus, cell density needs to be carefully optimized. 6. Transwell-Clear permeable filter can be punched/cut together with gel, since the filter usually remains attached to the gel after polymerization. Also, do not peel off the filter immediately after punching, as it might damage the gel. After boiling in SDS buffer (Subheading 3.5), the filter can easily be peeled off. 7. It is critical to remove all SDS before immunostaining. Poor washing will result in poor immunolabeling. After SDS washout, extra punches can be stored in 1 PBS at 4  C and can be immunolabeled even a week later. However, prolonged storage will diminish the quality of immunostaining. 8. Table 1 provides primary antibodies compatible with our protocol. Please note that for each cell type or a specific batch of antibody, additional troubleshooting may be needed for optimal immunolabeling results. Duration of immunolabeling can be shortened, although it may result in a weaker signal. 9. There is no a strict protocol as to how to expand gel punches. Duration of expansion has to be determined empirically and will depend on the size of the punches, and on the volume and the frequency of exchanged dH2O.

Acknowledgments We thank members of LPDS and Dr. Valentin Magidson for critical reading of the manuscript, and Dr. Catherine Sullenberger for acquiring images used in Fig. 3d. This work was supported by the Intramural Research Program of the National Institutes of Health (NIH), National Cancer Institute to J.L.

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References 1. Schermelleh L et al (2019) Super-resolution microscopy demystified. Nat Cell Biol 21 (1):72–84 2. Bates M, Huang B, Zhuang X (2008) Superresolution microscopy by nanoscale localization of photo-switchable fluorescent probes. Curr Opin Chem Biol 12(5):505–514 3. Schermelleh L, Heintzmann R, Leonhardt H (2010) A guide to super-resolution fluorescence microscopy. J Cell Biol 190(2):165–175 4. Asano SM et al (2018) Expansion microscopy: protocols for imaging proteins and RNA in cells and tissues. Curr Protoc Cell Biol 80(1): e56 5. Chen F, Tillberg PW, Boyden ES (2015) Optical imaging. Expansion microscopy. Science 347(6221):543–548 6. Chozinski TJ et al (2016) Expansion microscopy with conventional antibodies and fluorescent proteins. Nat Methods 13(6):485–488 7. Ku T et al (2016) Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues. Nat Biotechnol 34(9):973–981 8. Tillberg PW, Chen F (2019) Expansion microscopy: scalable and convenient superresolution microscopy. Annu Rev Cell Dev Biol 35:683–701 9. Gupta A, Kitagawa D (2018) Ultrastructural diversity between centrioles of eukaryotes. J Biochem 164(1):1–8 10. Vorobjev IA, Chentsov Yu S (1982) Centrioles in the cell cycle. I Epithelial cells. J Cell Biol 93 (3):938–949 11. Anderson RGW (1972) The three-dimensional structure of the basal body from the Rhesus monkey oviduct. J Cell Biol 54(2):246–265 12. Mennella V et al (2012) Subdiffractionresolution fluorescence microscopy reveals a domain of the centrosome critical for

pericentriolar material organization. Nat Cell Biol 14(11):1159–1168 13. Mennella V et al (2014) Amorphous no more: subdiffraction view of the pericentriolar material architecture. Trends Cell Biol 24 (3):188–197 14. Lawo S et al (2012) Subdiffraction imaging of centrosomes reveals higher-order organizational features of pericentriolar material. Nat Cell Biol 14(11):1148–1158 15. Sonnen KF et al (2012) 3D-structured illumination microscopy provides novel insight into architecture of human centrosomes. Biol Open 1(10):965–976 16. Nigg EA, Raff JW (2009) Centrioles, centrosomes, and cilia in health and disease. Cell 139 (4):663–678 17. Nigg EA, Schnerch D, Ganier O (2017) Impact of centrosome aberrations on chromosome segregation and tissue architecture in cancer. Cold Spring Harb Symp Quant Biol 82:137–144 18. Bettencourt-Dias M et al (2011) Centrosomes and cilia in human disease. Trends Genet 27 (8):307–315 19. Wang L, Dynlacht BD (2018) The regulation of cilium assembly and disassembly in development and disease. Development 145(18): dev151407 20. Sahabandu N et al (2019) Expansion microscopy for the analysis of centrioles and cilia. J Microsc 276(3):145–159 21. Kong D et al (2020) Prolonged mitosis results in structurally aberrant and over-elongated centrioles. J Cell Biol 219(6):e201910019 22. Rose GG et al (1958) A cellophane-strip technique for culturing tissue in multipurpose culture chambers. J Biophys Biochem Cytol 4 (6):761–764

Chapter 19 Analysis of Cell Cycle Progression in the Budding Yeast S. cerevisiae Deniz Pirincci Ercan and Frank Uhlmann Abstract The cell cycle is an ordered series of events by which cells grow and divide to give rise to two daughter cells. In eukaryotes, cyclin–cyclin-dependent kinase (cyclin–Cdk) complexes act as master regulators of the cell division cycle by phosphorylating numerous substrates. Their activity and expression profiles are regulated in time. The budding yeast S. cerevisiae was one of the pioneering model organisms to study the cell cycle. Its genetic amenability continues to make it a favorite model to decipher the principles of how changes in cyclin-Cdk activity translate into the intricate sequence of substrate phosphorylation events that govern the cell cycle. In this chapter, we introduce robust and straightforward methods to analyze cell cycle progression in S. cerevisiae. These techniques can be utilized to describe cell cycle events and to address the effects of perturbations on accurate and timely cell cycle progression. Key words Cell cycle, Cell synchronization, Time course experiment, Flow cytometry, Budding index, Western blotting, Immunofluorescence microscopy, S. cerevisiae

1

Introduction The eukaryotic cell cycle is a highly regulated process that coordinates the doubling of all cellular content, especially genome duplication during DNA replication, with chromosome segregation and cell division. The timing of events and their unidirectional progression are controlled by the master cell cycle regulator, the cyclin– Cdk complexes. Cdk is a serine/threonine protein kinase that is activated by forming a complex with non-enzymatic cell cycle regulatory subunits, called cyclins. The cyclical expression of cell cycle stage-specific cyclins results in oscillations of cyclin-Cdk activity. This allows the step by step phosphorylation of many cyclinCdk substrates throughout the cell cycle [1]. The overall activity of cyclin–Cdk complexes increases from the time when cells are born in the G1 phase of the cell cycle until

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_19, © The Author(s) 2021

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metaphase, when cyclin-Cdk activity reaches its peak. Then, a ubiquitin ligase complex, the APC/C (anaphase promoting complex/ cyclosome) is activated that ubiquitinates cyclins and thereby targets them for degradation. This causes cyclin-Cdk activity to fall, facilitating the dephosphorylation of cyclin-Cdk substrates by counteracting phosphatases and leading to completion of cell division and return of the cell cycle to G1. APC/C activation also results in degradation of securin, which liberates the protease separase to cleave the cohesive links between sister chromatids to trigger chromosome segregation. The abovementioned events sketch out an overview of cell cycle control, however many crucial questions remain open. How are the phosphorylation and dephosphorylation timings of each cyclinCdk substrate controlled to coordinate the multitude of molecular events that are required for successful cell growth and division, and for maintaining the integrity of the genome? Here we report a series of techniques which allow the analysis of cell cycle progression in the unicellular model eukaryote S. cerevisiae. Human cells contain a similar set of G1, S phase, and mitotic cyclins to those found in this yeast. Studying the role of genes and pathways that are part of the cell cycle control network is of great importance to understand how cell cycle regulation goes wrong in human disease, notably in cancer. In this chapter, we describe detailed protocols for cell synchronization and time course experiments for the analysis of the budding yeast cell cycle. We begin by describing cell cycle synchronization in a G1-like state using mating pheromone, from which cells can be released to synchronously progress through S phase and mitosis. We describe the use of flow cytometry to analyze DNA content. Together with the budding index, this is a useful marker to follow DNA replication and cell division. This is followed by a protocol for reliable protein extraction for Western blotting, used to visualize both protein abundance changes and electrophoretic mobility shifts that often go hand in hand with cyclin-Cdk phosphorylation. We then turn to the cytological observation of nuclei and the microtubule cytoskeleton, which yields additional information on the progression through mitosis.

2

Materials

2.1 Media and Chemicals

1. Yeast peptone (YP) media: weigh 20 g peptone, 10 g yeast extract and 0.05 g Adenine-HCl and make up to 1 L with water (water is Milli-Q ultrapure water unless otherwise stated). Autoclave before use.

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2. Yeast peptone dextrose (YPD) media: YP is prepared as described above, dextrose is added at a final concentration 2% (w/v) from 20% (w/v) dextrose stock dissolved in water and sterilized by filtration. 3. a-factor, stock: 0.2 mg/ml in methanol, stored at
20  C. 4. α-factor, stock: 5 mg/ml in methanol, stored at
20  C. 5. Nocodazole (Sigma-Aldrich), stock: 2 mg/ml in DMSO, stored at
20  C. 6. Flow cytometry buffer: 200 mM Tris-HCl pH 7.5, 210 mM NaCl, 78 mM MgCl2. 7. Propidium iodide (Sigma-Aldrich), stock: 1 mg/ml in water, stored at 4  C. 8. 50 mM Tris-HCl pH 7.5. 9. RNase A (Sigma-Aldrich), stock: 10 mg/ml in 50% glycerol, 10 mM Tris-HCl pH 8.0, stored at
20  C. 10. Fixation buffer: 100 mM KH2PO4 pH 6.4, 0.5 mM MgCl2, 3.7% formaldehyde (w/v, added freshly). 11. Wash buffer: 100 mM KH2PO4 pH 7.4, 0.5 mM MgCl2, 1.2 M Sorbitol. 12. 1 M Tris base, dissolved in water without pH adjustment. 13. Trichloroacetic acid (TCA), stock: 20%. 14. 70% Ethanol. 15. 37% Formaldehyde solution. 16. Methanol. 17. Acetone. 18. Poly-L-lysine (Sigma-Aldrich), stock: 0.1% (w/v) in water. 19. Lyticase (Sigma-Aldrich), stock: 20000 units/ml in 20% glycerol, 50 mM Tris-HCl pH 8.0, stored at
20  C. 20. ß-mercaptoethanol. 21. PBS: 2.7 mM KCl, 137 mM NaCl, 10 mM Na2HPO4, 1.8 mM KH2PO4. 22. Blocking buffer: 1% Bovine serum albumin (BSA) in PBS. 23. VECTASHIELD HardSet Antifade Mounting Medium containing 40 ,6-diamidino-2-phenylindole (DAPI). 24. Dithiothreitol (DTT), stock: 1 M in water stored at
20  C. 25. SDS-PAGE loading buffer: 100 mM Tris-HCl pH 6.8, 4% SDS (w/v), 0.2% bromophenol blue (w/v), 20% glycerol (v/v), 200 mM DTT (added freshly). 26. Protein assay dye reagent concentrate (Bradford solution).

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Equipment

1. Spectrophotometer. 2. Light microscope with 40 phase contrast objective and 10 eyepieces. 3. Flow cytometer. 4. Sonicator, MSE Soniprep 150. 5. Vortex. 6. Centrifuge for 50 ml centrifuge tubes. 7. Tabletop centrifuge for 1.5 ml microcentrifuge tubes and 2 ml screw-cap tubes. 8. 200 μl PCR tube. 9. Shaking water bath. 10. Vacuum filtration system. 11. FastPrep-24 cell breaker, MP Biomedicals. 12. Fluorescence microscope.

2.3

Consumables

1. 1.5 ml microcentrifuge tubes. 2. 2 ml screw-cap tubes. 3. 15 ml and 50 ml centrifuge tubes. 4. Mixed cellulose esters membrane filter, 1.2 μm pore size. 5. Flow cytometer tubes. 6. Acid-washed glass beads. 7. 23G needles. 8. Slides and coverslips.

3

Methods

3.1 Cell Synchronization Methods in Budding Yeast 3.1.1 PheromoneInduced Arrest of MATa Cells

1. Culture cells overnight in YPD at 25  C. 2. The following morning measure OD600 (see Note 1). 3. When OD600 reaches 0.2 (approximately 4  105 cells/ml) add α-factor to a final concentration 5 μg/ml and grow cells at 25  C for 1 h (see Note 2). 4. Continue adding the same amount of α-factor to cultures every hour until 2.5 h after the initial α-factor addition. 5. Retrieve a small volume of the culture, briefly sonicate (see Note 3) and check the cells under a phase contrast microscope. More than 90% of cells are expected to have accumulated as unbudded cells with a mating projection tip called shmoo [2].

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3.1.2 PheromoneInduced Arrest of MATα Cells

1. Follow steps 1–5 in Subheading 3.1.1. but use 0.02 μg/ml afactor instead of 5 μg/ml α-factor (see Note 4).

3.1.3 Mitotic Arrest by Nocodazole Treatment

1. Arrest cells in G1 by following the steps in Subheading 3.1.1 or 3.1.2. 2. Filter the culture and wash cells with 5 volumes of YP. 3. Resuspend the cells in a new flask containing the required amount of prewarmed YPD with nocodazole at a final concentration of 8 μg/ml. 4. Incubate at 25  C for 90 min to 2 h. 5. Observe the cells under microscope after incubation. More than 90% cells are expected to have accumulated as large budded cells and arrested in mitosis.

3.2 Analysis of Cell Cycle Progression in Time Course Experiments 3.2.1 Sample Collection

1. Grow and arrest cells using one of the methods described in Subheading 3.1. Meanwhile, prepare the following: – Two 1.5 ml microcentrifuge tubes, for flow cytometry and fluorescence microscopy samples, one 15 ml centrifuge tube and one 2 ml screw-cap tube, for protein extraction, per time point (see Note 5). – 70% ethanol and 20% TCA stocks, kept on ice. – A new flask containing an equal culture volume of YPD placed in a shaking water bath that is set at 25  C. 2. Collect 17 ml culture from the arrested culture using a 25 ml pipette and partition 1 ml each into the two microcentrifuge tubes and the remainder into a 15 ml centrifuge tube. 3. Spin down cells in microcentrifuge tubes at 6000 rpm (3430  g), 1 min and the 15 ml centrifuge tube at 3500 rpm (14,130  g) for 3 min at 4  C. 4. Remove supernatant from all tubes by aspiration. 5. Resuspend the two small cell aliquots, to be analyzed by flow cytometry and fluorescence microscopy, in 1 ml 70% ethanol and 1 ml fixation buffer containing 3.7% formaldehyde, respectively. 6. Resuspend the large aliquot, to be used for protein extraction, in 1 ml 20% TCA and transfer to the 2 ml screw-cap tube. 7. Filter the culture and wash cells with 5 volumes of YP (see Notes 6 and 7). 8. Transfer the filter with the cells into the flask with fresh medium (containing nocodazole in case of a mitotic arrest), prepared prior to release (see step 1), and start timer.

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9. Collect and process samples at regular intervals (e.g., every 15 min) following steps 2–6 until the end of experiment. 10. Upon bud formation (approximately 60 min following the release) α- or a-factor should be readded to the culture so that cells arrest again following complete passage through one cell cycle (this step is not required in mitotic arrest experiments using nocodazole). 11. End the experiment when all cells have returned to G1, or once all cells have reached mitosis in case of an experiment using nocodazole. 12. Sample processing at the end of the time course experiment: – Store flow cytometry samples in 70% ethanol for at least 2 h at room temperature or overnight at 4  C. – Incubate fluorescence microscopy samples in fixation buffer for 2 h at 30  C or overnight at 4  C for fixation. – Leave Western blotting samples on ice in 20% TCA for at least 30 min (see Note 8). 3.2.2 Flow Cytometry: Analysis of DNA Content

Continue with the samples stored in 70% ethanol. 1. Pellet cells at 6000 rpm (3430  g), 2 min and remove supernatant. 2. Resuspend samples in 1 ml 50 mM Tris–HCl pH 7.5, containing 0.1 mg/ml RNase A. 3. Incubate samples at 37  C for at least 2 h or overnight. 4. Spin down again at 6000 rpm (3430  g), 2 min and aspirate supernatant (see Note 9). 5. Resuspend pellets in 400 μl flow cytometry buffer containing 50 μg/ml propidium iodide. 6. Sonicate samples for 5 s to disrupt cell aggregates. 7. Transfer 50 μl from cell suspension to flow cytometry tubes containing 400 μl 50 mM Tris–HCl pH 7.5 and vortex briefly. 8. Use blue (488 nm) or yellow-green (561 nm) flow cytometer lasers to immediately read 10,000 events per sample prepared in step 7. A sample analysis that shows transition from 1c to 2c DNA content during DNA replication and regeneration of 1c daughter cells during cell division is shown in Fig. 1a. 9. Sonicated samples from step 6 can be stored at 4  C for further analysis, for example budding index counts.

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Fig. 1 Cell cycle analysis in budding yeast. (a) MATa cells were arrested by the addition of α-factor with 1c DNA content and then released to progress through a synchronous cell cycle. α-factor was re-added at 60 min (upon visible bud formation) to rearrest cells after completing one cell division cycle. Cell cycle progression was monitored by flow cytometry analysis of DNA content. (b) The fraction of visibly budded cells was counted in aliquots of the flow cytometry samples over time. One hundred cells were counted at each time point. The means of three independent experiments and the standard error are shown. (c) Protein extracts were prepared at the indicated times and analyzed by SDS-PAGE and Western blotting. Clb2, Clb5, and Orc6 were detected using specific antibodies (see Note 13). Tubulin served as a loading control and was detected using monoclonal antibody clone Tat1 3.2.3 Budding Index as a Marker of Cell Cycle Progression

1. Place 10 μl of sonicated cell suspension (from step 6 in Subheading 3.2.2) on a slide. 2. Cover with a coverslip and count the fraction of cells that display a visible bud, small or large, in a population of at least one hundred cells under the phase contrast microscope. 3. Plot the percentage of budded cells over time to obtain the budding index profile (Fig. 1b). Visible buds emerge at the time DNA replication.

3.2.4 Sample Preparation for Western Blotting and Detection of the Cell Cycle Marker Proteins

Continue with the Western blotting samples stored in 20% TCA. 1. Spin down samples at 13,000 rpm (16,060  g), 1 min at 4  C. 2. Wash cells with 1 ml 1 M Tris base (without pH adjustment). 3. Repeat step 1 and resuspend the cells completely in 100 μl SDS-PAGE loading buffer using a vortex mixer (see Note 10). 4. Measure 200 μl of glass beads using a 200 μl PCR tube and add to the screw-cap tubes. 5. Break cells in the Fast Prep 24 according to the manufacturer’s instructions (see Note 11). 6. Boil samples at 95  C for 5 min.

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Meanwhile, prepare and label new microcentrifuge tubes for all the samples. 7. Release pressure in the screw-cap tubes and tighten the lids back. 8. Puncture the bottom of the screw-cap tubes twice with a 23 G needle and place them tightly onto new microcentrifuge tubes, prepared at step 6. 9. Loosen the screw caps slightly once again and place both the screw caps and the microcentrifuge tubes into a 50 ml centrifuge tube. 10. Spin at 1100 rpm (1400  g) for 2 min (see Note 12). 11. Discard the screw-cap tubes, which should contain only glass beads, boil the microcentrifuge tubes containing the samples at 95  C for 5 min. 12. Spin down samples at 13,000 rpm (16,060  g), 3 min to sediment debris, then determine protein concentration using a Bradford protein assay. 13. Load 10 μg protein per sample for analysis by typical SDSPAGE methodology [3]. The remaining samples can be stored at
20  C. 14. Following protein transfer to a nitrocellulose membrane, probe against cell cycle regulated proteins, for example the S phase cyclin Clb5 and the mitotic cyclin Clb2, or proteins that show a cell cycle regulated mobility shift [4] (e.g., Orc6 [5]) (see Note 13). An example analysis is shown in Fig. 1c. 3.2.5 Fluorescence Microscopy to Determine Cell Cycle Stage by Spindle Staining

Continue with the samples stored in fixation buffer containing 3.7% formaldehyde. 1. Pellet cells at 6000 rpm (3430  g), 2 min and remove supernatant. 2. Resuspend cells in 1 ml fixation buffer without formaldehyde. 3. Repeat step 1 and resuspend pellets in 1 ml wash buffer. 4. Repeat step 1 once again and resuspend cells in 200 μl wash buffer containing 2 μl/ml ß-mercaptoethanol and 2 μl/ml lyticase solution to digest the cell wall (spheroplasting). 5. Incubate for 40 min at 37  C. Meanwhile, coat the wells of a 15-well slide with 10 μl poly-L-lysine solution and leave it in a moist chamber for 30 min at room temperature. Next, wash the slide under flowing water briefly and dry by aspirating the remaining water drops. The slide should always be kept in a moist chamber from now on.

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6. Check spheroplasts under the phase contrast light microscope. Spheroplasts appear dark, while intact cells remain bright (see Notes 14 and 15). 7. Centrifuge cells at 3500 rpm (1160  g), 2 min and aspirate supernatant carefully. 8. Resuspend cells in 0.5 ml wash buffer and repeat step 7. 9. Add 200 μl wash buffer to the pellets (see Note 16). 10. Carefully pipette up and down five times to mix and transfer 5 – 10 μl cell suspension to a well on the 15-well slide. The remaining samples can be stored at
20  C. 11. Let cells adhere to the slide for 30 min in a moist chamber at room temperature. 12. Aspirate the excess of cell suspension using a fine tip (see Note 17) and plunge the slide into
20  C cold methanol for 3 min. 13. Transfer slide to
20  C cold acetone for 10 s. 14. Let the acetone evaporate at room temperature quickly, then place the slide back into the moist chamber. 15. Add 10 μl blocking buffer to the wells for at least 20 min. 16. Aspirate blocking buffer and add 8 μl of primary antibody diluted in blocking buffer onto the wells (see Note 18). 17. Incubate for 1 h at room temperature. 18. Wash cells three times with 10 μl blocking buffer. 19. Add 8 μl of secondary antibody diluted in blocking buffer into the wells (see Note 18). 20. Repeat step 17 in the dark, as well as step 18. 21. Aspirate as much as possible of the blocking buffer after the final wash and mount slides using 0.5 μl mounting media containing DAPI. 22. Cover the slide with a coverslip and seal the edges with a nail polish. 23. Slides can be imaged immediately, or can be stored in the dark at
20  C and imaged later. 24. Count the appearance of short (2 μm, metaphase) spindles, as well as long (>2 μm, anaphase) spindles in a population of at least a hundred cells under a fluorescence microscope. 25. Plot the fraction of cells with metaphase and anaphase spindles over time to mark the onset of mitosis when short spindles form, anaphase when spindles elongate and mitotic exit when spindles disassemble (see Fig. 2 for examples of cells with metaphase and anaphase spindles).

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Fig. 2 Immunofluorescence analysis of the microtubule cytoskeleton to identify mitotic stages. Asynchronously proliferating cells were fixed and stained using an anti-α-tubulin antibody. DNA was counterstained with DAPI. The cell outlines were marked with a dashed line. Examples of a metaphase cell with a short spindle (arrowhead 1) and an anaphase cell with an elongating spindle and separating nuclei (arrowhead 2) are highlighted (Adapted with permission from Borges et al. (2010) Mol. Cell 39:677–688 [8])

4

Notes 1. In the morning, if the culture is overgrown but cells are still budding, back dilute to OD600 ¼ 0.1 and allow one generation doubling to reach OD600 ¼ 0.2 before starting the experiment. 2. A lower concentration of α-factor might be sufficient to achieve cell cycle arrest, depending on its purity. On the other hand, if most cells are still budded after 2.5 h in medium containing α-factor, the concentration may need to be increased. 3. All the sonication steps in this chapter are performed using an exponential microprobe and the amplitude meter is set at 6 microns. 4. MATα cells treated with a-factor show less prominent shmoo formation compared to α-factor treated MATa cells [2]. 5. Keep microcentrifuge tubes and screw-cap tubes on ice prior to release and during the experiment. 6. If the experiment requires filtration of more than one culture, membranes should be folded and kept on a sterile foil until filtration is completed for all cultures. Cells can then be simultaneously resuspended in new flasks. 7. If more than two cultures are analyzed, the use of more than one filtration system is recommended. 8. Western blotting samples can be stored as pellets at
80  C prior to protein extraction by simply spinning down the samples at 13,000 rpm (16,060  g), 1 min and removing 20% TCA.

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9. After RNase treatment, pellets become slightly translucent and adhere to the microcentrifuge wall less tightly. The supernatant should be carefully aspirated in order to prevent loss of the pellets. 10. If SDS-PAGE loading buffer turns yellow after mixing, this indicates an acidic pH due to the presence of residual TCA. In this case, samples can be neutralized by adding 1–2 drops of 1 M Tris base. 11. Running the manufacturer’s S. cerevisiae program (6 m/s for 40 s) once is enough to break the cells. 12. It is possible to fit two screw-cap microcentrifuge tube combinations into one 50 ml centrifuge tube [6]. These 50 ml centrifuge tubes can be stored and reused. 13. Clb5 and Clb2 can be detected using specific primary antibodies (anti-Clb5, Santa Cruz sc20170, anti-Clb2 Santa Cruz sc9071, Fig. 1c) or by fusing small epitope tags (e.g., HA or Pk) to their C terminus [7]. Orc6 can be detected with monoclonal antibody clone SB49. 14. If a majority of the cells appear dark, continue with the next step. Otherwise, incubate cells longer at 37  C. 15. Spheroplasts are fragile; therefore, subsequent centrifuge steps will be performed at a lower speed. 16. At this step, the samples can be stored longer term at
20  C. 17. Always aspirate from the same side of the wells to reduce the risk of scratch marks on the slide. 18. For spindle staining, we recommend using an anti-α-tubulin monoclonal primary antibody clone YOL1/34, Abcam ab6161, diluted 1:200 in blocking buffer. Similarly, a secondary antibody conjugated to a fluorescent dye should be diluted in blocking buffer according to manufacturer’s instructions before use.

Acknowledgments We would like to thank Florine Chre´tien and Hon Wing Liu for their help. This work was supported by The Francis Crick Institute, which receives its core funding from Cancer Research UK (FC001198), the UK Medical Research Council (FC001198), and the Wellcome Trust (FC001198). D.P.E. was supported by a Boehringer Ingelheim Fonds Ph.D. Fellowship.

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References 1. Morgan D (2007) The cell cycle: principles of control. New Science Press, London 2. O’Reilly N, Charbin A, Lopez-Serra L, Uhlmann F (2012) Facile synthesis of budding yeast a-factor and its use to synchronize cells of a mating type. Yeast 29:233–240 3. Sambrook J, Fritsch EF, Maniatis T (1989) Section 18.47-18.54. In: molecular cloning: a laboratory manual, 2nd edn. Cold Spring Harbor Laboratory Press, New York, NY, p 1659 4. Sambrook J, Fritsch EF, Maniatis T (1989) Section 18.64-18.66. In: molecular cloning: a laboratory manual, 2nd edn. Cold Spring Harbor Laboratory Press, New York, NY, p 1659 5. Nguyen VQ, Co C, Li JJ (2001) Cyclindependent kinases prevent DNA re-replication

through multiple mechanisms. Nature 411:1068–1073 6. Kakui Y, Uhlmann F (2019) Efficient depletion of fission yeast condensin by combined transcriptional repression and auxin-induced degradation. Methods Mol Biol 2004:25–33 7. Knop M, Siegers K, Pereira G, Zachariae W, Winsor B, Nasmyth K, Schiebel E (1999) Epitope tagging of yeast genes using a PCR-based strategy: more tags and improved practical routines. Yeast 15(10B):963–972 8. Borges V, Lehane C, Lopez-Serra L, Flynn H, Skehel M, Rolef Ben-Shahar T, Uhlmann F (2010) Hos1 deacetylates Smc3 to close the cohesin acetylation cycle. Mol Cell 39 (5):677–688. https://doi.org/10.1016/j. molcel.2010.08.009

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 20 Application of PALM Superresolution Microscopy to the Analysis of Microtubule-Organizing Centers (MTOCs) in Aspergillus nidulans Xiaolei Gao, Reinhard Fischer, and Norio Takeshita Abstract Photoactivated localization microscopy (PALM), one of the super resolution microscopy methods improving the resolution limit to 20 nm, allows the detection of single molecules in complex protein structures in living cells. Microtubule-organizing centres (MTOCs) are large, multisubunit protein complexes, required for microtubule polymerization. The prominent MTOC in higher eukaryotes is the centrosome, and its functional ortholog in fungi is the spindle-pole body (SPB). There is ample evidence that besides centrosomes other MTOCs are important in eukaryotic cells. The filamentous ascomycetous fungus Aspergillus nidulans is a model organism, with hyphae consisting of multinucleate compartments separated by septa. In A. nidulans, besides the SPBs, a second type of MTOCs was discovered at septa (called septal MTOCs, sMTOC). All the MTOC components appear as big dots at SPBs and sMTOCs when tagged with a fluorescent protein and observed with conventional fluorescence microscopy due to the diffraction barrier. In this chapter, we describe the application of PALM in quantifying the numbers of individual proteins at both MTOC sites in A. nidulans and provide evidence that the composition of MTOCs is highly dynamic and dramatically changes during the cell cycle. Key words PALM, Aspergillus nidulans, Ascomycete, Spindle pole body, Septal MTOC

1

Introduction The filamentous fungus Aspergillus nidulans is an ideal model system to study the cell biology of lower eukaryotes due to the well-established toolboxes in genetics, biochemistry, cell biology and molecular biology. Many studies in A. nidulans focused on the analysis of polarity establishment, polar growth, or the function of the cytoskeleton [1–4]. Highlights were the discovery of γ-tubulin or the molecular analysis of cell cycle mutants [5–8]. The arrangement and interplay of the microtubule (MT) and the actin cytoskeletons is crucial for hyphal growth and the maintenance of polarity [9–11]. In A. nidulans, the MT cytoskeleton delivers the so-called cell-end marker proteins to the hyphal tips, which in turn polarize

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_20, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Scheme of a hyphal compartment and the arrangement of MTOCs and MTs in A. nidulans. Hyphal compartments are separated by a septum and contain several nuclei. SPBs are the main MTOCs located at nuclei and septal MTOCs polymerize MTs bidirectionally from septa into the cytoplasm

the actin cytoskeleton [11, 12]. MTs are nucleated from large protein complexes, microtubule-organizing centers (MTOCs) in vivo. In A. nidulans, MTs are nucleated from at least two sites, the spindle-pole bodies (SPBs) which are embedded in the nuclear envelope and septal MTOCs (Fig. 1). Whereas the plus ends of MTs are highly dynamic, the minus ends are anchored at the MTOCs. The arrangement of MTOCs hence leads to a mixture of MT orientations in hyphal compartments, although in the tip region MTs are mainly oriented with their plus ends toward the apex (Fig. 1). The simplest, but best characterized MTOC is the SPB of the budding yeast Saccharomyces cerevisiae which is a multilayered structure embedded into the nuclear envelope [13, 14]. The SPB is composed of at least 18 different proteins with a minimal complex for MT polymerization. This minimal complex is called γ-tubulin small complex (γ-TuSC) and consists of γ-tubulin plus two other γ-tubulin complex proteins (named GCP in humans). The protein names differ between organisms, because they were often discovered in mutagenesis approaches. In S. cerevisiae γ-tubulin is Tub2, GCP2 is Spc97 and GCP3 is Spc98 [15]. The fission yeast Schizosaccharomyces pombe, fungi like A. nidulans, and higher eukaryotes, contain a complex with more subunits GCP4, GCP5, and GCP6, which is called γ-tubulin ring complex (γ-TuRC). The understanding of γ-TuRC assembly into a spiral geometric structure for MTs nucleation is highly improved with cryo-electron microscopy reconstructions in a recent study of Xenopus laevis [16]. It highlights the significance of GCP6-specific insertion domain and actin as important structures of γ-TuRC. In A. nidulans MipA (γ-tubulin), GcpB and GcpC constitute the core γ-TuSC, and three additional GCPs GcpD, GcpE and GcpF are called γ-TuRC specific components or noncore components [17]. Although the SPB in S. cerevisiae and in S. pombe are very well studied, it was discovered that in S. pombe noncentrosomal MTOCs exist. Three different MTOCs were described, the spindle pole body, temporal MTOCs in the division plane (eMTOCs) and nuclear-envelope associated MTOCs in interphase cells (iMTOCs)

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Fig. 2 Proposed model for septal MTOCs of A. nidulans. In the center of a mature septum Spa10 concentrates into a central disk structure and acts as an anchor for sMTOC components. ApsB interacts with Spa18 and both attach to the periphery of Spa10. From there they recruit γ-TuRCs and together polymerize microtubules (left, Adapted from [23]). Notably, γ-TuRC at septa contains GcpD, GcpE, GcpF, and MztA but with two outer plaque receptors ApsB and Spa18, defining a different structure from SPBs (right, Adapted from [22])

[18, 19]. We discovered that A. nidulans has two types of MTOCs, SPBs and septal MTOCs (sMTOCs). Although all γ-TuRC components were found at SPBs and at sMTOCs, other proteins were specific for the SPB or sMTOCs [20–23]. γ-TuRC is regulated by various factors that recruit it to different cellular sites, spatially controlling microtubule nucleation activities. In fission yeast, the outer plaque receptors of the γ-TuRC Mto1 and Mto2 play essential roles in nucleating cytoplasmic MTs, and the inner plaque receptor Pcp1 is responsible for spindle MTs nucleation [24–26]. Similarly, the outer plaque receptors of A. nidulans, ApsBMto1 and Spa18Mto2, were found to be at SPBs and sMTOCs and play essential roles for septal MTs nucleation [23]. At septa, an intrinsically disordered protein Spa10 was found to recruit ApsB and Spa18 (Fig. 2). At forming septa, Spa10 colocalized with the tropomyosin ring TpmA and at mature septa, where the actin ring disappeared, Spa10 concentrated into a stable central disk, while ApsB and Spa18 were only targeted to mature septa through attachment to the periphery of the Spa10 disk [23]. Recently, a new component of γ-TuRC was discovered in humans and named mitotic-spindle organizing protein associated with a ring of γ-tubulin (MOZART1) [27]. MOZART1 is conserved in most eukaryotes and its ortholog in A. nidulans is MztA. MztA interacted and colocalized with PcpAPcp1, the inner plaque receptor of the γ-TuRC, at SPBs. On the other hand, neither interaction nor colocalization was detected between MztA and ApsB [22]. MztA was also required to recruit GcpD and was located only at inner plaques of SPBs (Fig. 3). Taken together, the results suggested an asymmetry of the composition of the

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Fig. 3 Proposed model for the spindle pole body (SPB) of A. nidulans. The spindle pole body is composed of three plaques and a half bridge. The outer plaque harbours the small γ-TuSCs and the two receptor proteins ApsB and Spa18. The inner plaque harbours the big γ-TuRCs with MztA, and its receptor PcpA spans the central and inner plaque. SepK is one of the ApsB recruiters at the outer plaque. The only known component of the central plaque is CaM which binds to PcpA. Ndc1 is supposed to be a SPB component but its exact position still needs to be determined. Notably, it is not an anchor for SPBs. Cdc31 and Sfi1 are two orthologs of proteins resident at the half bridge and required for SPB duplication (Adapted from [22])

SPB in A. nidulans. Whereas the outer plaque of the SPBs is composed of γ-TuSCs, the inner plaque is composed of γ-TuRCs [22] (Fig. 3). Similarly, heterogeneity of γ-TuRCs was found in sperm cells of Drosophila melanogaster [28]. MztA was also found at sMTOCs in A. nidulans (Fig. 2,4). Hence, in A. nidulans three different MTOCs are established. The outer plaque with γ-TuSCs lacking MztA and the inner plaque with γ-TuRC containing MztA

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Fig. 4 Localization of MztA at SPBs and at sMTOCs as observed with epifluorescence microscopy in A. nidulans. MztA was fused to GFP and alpha tubulin (TubA) to mCherry. (Left) A single mitotic spindle with the two SPBs at their ends. (Right) Interphase MTs emanate from sMTOCs at septa. Asterisks indicate the septum position. Scale bar, 2 μm (Adapted from [22])

(Fig. 3). sMTOCs appear to be hybrids of the two SPB MTOCs with γ-TuRCs including MztA (like the inner plaque) but with the anchor proteins ApsB and Spa18 (Fig. 2), which are typical components of the SPB outer plaques. Using epifluorescence microscopy, all proteins of MTOCs in A. nidulans appeared as dots at SPBs and sMTOCs making it hard to resolve any further details of the molecular structures (Fig. 4). Light microscopy is restricted to an optical resolution of roughly 250 nm due to the diffraction limit [29]. Superresolution microscopy has been invented to bypass the physical barrier and enabled visualization of previously invisible molecular details in many biological systems [30]. Based on different principles of overcoming the diffraction limit, two categories of super-resolution approaches have been developed, (1) structured illumination microscopy (SIM) and stimulated emission depletion microscopy (STED) using patterned illumination to differently modulate the fluorescence emission, and (2) photoactivated localization microscopy (PALM) or stochastic optical reconstruction microscopy (STORM) by stochastically turning on individual molecular at different time points. These methods have been applied to answer questions involving the organization, interaction, stoichiometry, and dynamics of cellular components. However, there are only a few applications of superresolution microscopy in filamentous fungi. One application of SIM in filamentous fungi is the localization of a STRIPAK subunit PRO45 at the nuclear envelope, endoplasmic reticulum, and mitochondria in Sordaria macrospora [31]. STED has revealed the size of lipid rafts to around 70 nm by visualizing flotillin FloA in A. nidulans [32, 33]. Expansion microscopy (ExM) is a sample preparation tool for biological samples that enables high-resolution imaging by expanding them using a polymer system on a conventional fluorescence microscopy [34]. This method was very recently applied to ascomycetes and basidiomycetes [35].

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Here, we describe the application of photoactivated localization microscopy (PALM). It dramatically improves the spatial resolution to 10–20 nm [36, 37]. The basic principle behind PALM is the use of photoactivatable or photoconvertible fluorescent proteins to selectively switch on thousands of sparse subsets of molecules in a sequential manner. Most of the molecules are in an inactive state before a small fraction (less than 1%) is photoactivated or photoconverted using a brief pulse of ultraviolet or violet light to render that subset fluorescent. The activated molecules are then imaged and precisely localized followed by photobleaching to remove the unactivated molecules. The process is repeated many thousand times until all of the labeled molecules are obtained which allow the construction of super-resolution images [36]. One example of photoconvertible fluorescent protein is mEosFP which can be photoconverted from green to red fluorescence by 400 nm light [38]. First applications of PALM in filamentous fungi revealed a dynamic picture of the membrane-associated polarity marker TeaR in A. nidulans [39]. The PALM analyses clearly showed TeaR clusters near the apex of the hyphae, along the plasma membrane, with average sizes of approximately 120 nm. It was estimated that about 20 TeaR proteins composed each of these clusters. Another PALM example is chitin synthase ChsB in A. nidulans [40]. ChsB is mainly located at the Spitzenko¨rper near the hyphal tip and produces chitin, a key component of the cell wall. The quantitative and time-lapse PALM visualized the pulsatory dynamics of the Spitzenko¨rper, reflecting vesicle accumulation before exocytosis and their subsequent fusion with the apical plasma membrane [41, 42]. Similar PALM analyses contribute to reveal a MT guidance mechanism depending on actin cables by MigA in A. nidulans, an ortholog of the karyogamy protein Kar9 from Saccharomyces cerevisiae [43], and dynamics of actin cables in A. nidulans [44]. In this chapter, we describe mEosFP based PALM application to the large protein complex, the microtubule organizing center (MTOCs), in A. nidulans. Through quantifying the numbers of MztA, GcpC, GcpD and ApsB, we revealed that their molecular numbers at SPBs and sMTOCs are very dynamic (Fig. 5a, b). GcpC is the essential component of γ-TuRCs and each γ-TuRC contains 13–14 γ-tubulin and 6–7 GcpC molecules. We observed different numbers of GcpC molecules at different MTOCs. Interestingly, the numbers differed by 6–7 or multiples of that. Taking into account that each γ-TuRC contains 6–7 GcpC molecules, these results suggested different numbers of entire γ-TuRCs or entire γ-TuSCs at different MTOCs during interphase (Fig. 5c). During metaphase, GcpC numbers (around 60) at SPBs appeared to be constant, suggesting inner plaque nucleation centres are constant to ensure proper assembly of spindles (Fig. 5c).

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Fig. 5 PALM microscopy images of MztA-mEos and GcpC-mEos at two MTOC sites. (a, b) Images of MztAmEos (a) and GcpC-mEos (b) at two distinct SPBs and two sMTOCs. Scale bar, 5 μm. (c) GcpC numbers are dynamic in interphase and stable in metaphase. A total of 40 SPBs in interphase and ten SPBs during mitosis were included in the distribution analysis. The numbers of GcpC molecules per SPB were sorted ascendingly. The red line highlights discrete steps of GcpC numbers (Adapted from [22])

This is the first direct evidence for changing numbers of γ-TuRCs/γ-TuSCs in filamentous fungi during interphase of the cell cycle and shows the power of PALM imaging. PALM imaging allows determination of the number of single molecule and is a

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powerful tool to answer the questions of dynamics of individual molecular building blocks as well as elucidating stoichiometry of functional protein complexes in live cells.

2

Materials

2.1 Preparing Hyphae for Live Cell Imaging

1. Minimal medium: 5% (v/v) salt solution, 0.1% (v/v) trace elements, 2% (w/v) glucose or 2% (v/v) glycerol (see Note 1), add H2Obid., adjust pH with 10 M NaOH to 6.5. 2. Salt solution: 12% NaNO3, 1% KCl, 1% MgSO4∙7H2O, 3% KH2PO4, add H2Obid. 3. Trace elements: 2.2% ZnSO4 x 7H2O, 1.1% H3BO3, 0.5% MnCl2 x 4H2O, 0.5% FeSO4∙7H2O, 0.16% CoCl2∙5H2O, 0.16% CuSO4∙5H2O, 0.11% (NH4)6Mo7O24∙4H2O, 5% Na4EDTA, adjust pH with 10 M KOH to 6.5. 4. U-slide 8-well glass-bottom dishes for time-lapse images. 5. Strains: SXL36 (mztA::GFP; alcA(p)::mCherry::tubA); SXL85 (mztA::mEoSFP); SXL98 (gcpC::mEoSFP).

2.2 General Microscopy Equipment for Live Cell Imaging

1. Widefield-fluorescence microscope, AxioImager Z1 (Zeiss, Jena, Germany). 2. TL Halogen lamp HXP 120 for excitation of eGFP (488 nm/ 50 or 100 mW) or mCherry (561 nm/50 or 150 mW). 3. Filter sets: eGFP BP 470/40, FT 495, BP 525/50 and mCherry BP 545/25, FT 570, FT 570, BP 605/70 (Zeiss, Jena, Germany). 4. Zeiss objectives: Plan-Apochromat (63, NA 1.4). 5. Zeiss AxioCam MR camera. 6. Zeiss software package, Axio Vision v4.8.1, ZEN 2012 Blue Edition v1.20. 7. Microscope slides 76  26 mm, 1 mm thickness. 8. Coverslips 18  18 mm # 1.

2.3

PALM Set-Up

1. Super-resolution PALM/STROM inverted (ELYRA 3D-PALM, Zeiss, Jena, Germany).

microscope

2. High numerical aperture oil immersion objective (α-planApochromat, 100, numerical aperture, 1.46). 3. Multiple excitation laser lines (405, 473, and 561 nm). 4. An electron-multiplying charge-coupled device (EMCCD) camera (iXon Ultra 897; Andor Technology, Ltd., Belfast, Northern Ireland). 5. Zeiss software package.

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Methods To enable live cell imaging in a fungal model system, construction of transgenic strains expressing the proteins of interest fused N- or C-terminally to fluorescent proteins (e.g., the green and red fluorescent protein; eGFP or mCherry) is the first step. In this chapter, MTOC components were all tagged with GFP or mEosFP C-terminally via integration at the endogenous locus (knock-in). Therefore, the fusion proteins are expressed from their natural promoter to avoid overexpression artefacts. Construction details are described here [22]. We also verified that the fusion proteins are biologically functional and do not cause any defective phenotype. For nonessential genes such as mztA, which causes a mutant phenotype, the test can be easily done by complementation of the deletion mutant with the construct of interest. Full complementation of the mutant phenotype is desired.

3.1 Live Cell Imaging of MTOC Components in A. nidulans

1. Inoculate spores from strain of MztA-GFP; mCherry-tubA into 0.5 ml minimal medium (about 103 spores/ml) with 2% glycerol mounted on a sterile coverslip. Incubate the cells for 16–20 h at 28  C for germination and hyphal growth (see Note 2). 2. Mount the coverslips upside down on a microscope slide and avoid air bubbles inside (see Note 3). 3. For observation use the 63x objective and an appropriate camera to visualize the whole hyphae. 4. Choose appropriate illumination for excitation of fluorophores, filter sets (excitation/emission/dichroic beam splitter), and exposure time (see Note 4). 5. Set up a Z-stack series to get a full set of images according to the thickness of the hyphae (see Note 5). 6. Use camera in stream mode to obtain fastest acquisition. 7. Convert resulting images into maximum projection. This results in a 2D projection of 3D data (Fig. 4). 8. Alternatively, quantification of signal intensities between wild type and mutant can be performed (see Note 6).

3.2 PALM Experiments

1. Inoculate spores of A. nidulans strains expressing GcpC-mEos or MztA-mEos under natural promoter in a glass-bottom chamber with minimal media. Incubate at 25  C for 12 h. 2. With low 473 nm illumination, identify a cell that expresses a high level of fluorophores. 3. Adjust laser intensity to image mEos. Initially, illuminate the sample with the 561 nm laser to bleach the small population of mEos that is already converted, for example, emits red fluorescence (see Note 7).

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4. Fluorescent proteins are converted one by one from their green to their red emitting forms using low intensity 405 nm light and excited at 561 nm simultaneous illumination. 5. Use a 607/50 band-pass filter after passing through the excitation dichroic (z 405/473/561/635) to detect the red fluorescence. 6. Detect and record the signals from individual red fluorescent mEosFP molecules with a back illuminated EMCCD camera typically at 50 ms time resolution (see Note 8). 7. Analyse the PALM data with Zen software with PALM method (see Note 9). Use the molecule identification and quantification IMARIS 9 64 9.3 software (BITPLANE) alternatively.

4

Notes 1. For choosing the appropriate carbon source in microscopy medium, one has to consider that glucose represses the expression of alcA promoter and glycerol de-represses the expression. In another point, glucose helps strains grow faster than glycerol but increases the darkness of the medium. In this chapter, strains are controlled with native promoters; thus, glucose and glycerol are both applied. Considering better visibility, glycerol has priority. 2. For time-lapse images, use glass bottom dishes and hyphae will stay healthy for an extended period of time. 3. After overnight incubation, samples are taken out and put at room temperature for half an hour to adapt to room temperature. Fluorescent protein can be visualized better at room temperature than at 28  C. 4. We recommend exposure times no longer than 100 ms for eGFP and 200 ms for mCherry. Check the frame rate limit of the camera and choose an appropriate region of interest. 5. Before performing Z-stack images, the center of the hyphae should be focused through the DIC channel to assure that acquisition started in the correct focal plane. 6. Identify a background region of a certain size that can be used in all experiments to correct for background artifacts. Number of Z-stack series should be maintained for all the running and not exceed 20. Shuttle level, laser intensities, and exposure time should be set up identically. One should try to reduce the fluorescence photobleaching as much as possible. Replicates of the experiments need to be done to create data set. 7. A small population of mEosFP emits red fluorescence already before irradiation with 405 nm light. That is typical for mEos.

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8. As the pool of nonconverted mEosFP is depleted during data acquisition, it is advisable to increase the power of the activation laser to keep the number of activated fluorophores per frame constant. 9. The PALM micrograph in Fig. 5a is constructed by 1000 frames with a time resolution of 50 ms. ZEN 2.1, use PALM method, settings; discard overlapping molecules. Peak finder; peak mask size, 9 pixel, peak intensive to noise, 6. Localizer; x,y 2D Gauss Fit. Pixel resolution XY; 83 nm/pixel.

Acknowledgments Our research was in part financed by grants from the German Science Foundation (DFG Fi 459/20-1) and Japan Science and Technology Agency (JST, ERATO JPMJER1502). References 1. Takeshita N, Manck R, Gru¨n N, de Vega SH, Fischer R (2014) Interdependence of the actin and the microtubule cytoskeleton during fungal growth. Curr Opin Microbiol 20:34–41 2. Takeshita N (2016) Coordinated process of polarized growth in filamentous fungi. Biosci Biotechnol Biochem 80(9):1693–1699 3. Riquelme M, Aguirre J, Bartnicki-Garcı´a S, Braus GH, Feldbru¨gge M, Fleig U, Hansberg W, Herrera-Estrella A, K€amper J, Ku¨ck U (2018) Fungal morphogenesis, from the polarized growth of hyphae to complex reproduction and infection structures. Microbiol Mol Biol Rev 82(2):e00068–e00017 4. Fischer R, Zekert N, Takeshita N (2008) Polarized growth in fungi–interplay between the cytoskeleton, positional markers and membrane domains. Mol Microbiol 68(4):813–826 5. Oakley CE, Oakley BR (1989) Identification of γ-tubulin, a new member of the tubulin superfamily encoded by mipA gene of Aspergillus nidulans. Nature 338(6217):662 6. Morris NR, Enos AP (1992) Mitotic gold in a mold: Aspergillus genetics and the biology of mitosis. Trends Genet 8(1):32–33 7. Osmani SA, Engle DB, Doonan JH, Morris NR (1988) Spindle formation and chromatin condensation in cells blocked at interphase by mutation of a negative cell cycle control gene. Cell 52(2):241–251 8. Osmani AH, McGuire SL, Osmani SA (1991) Parallel activation of the NIMA and p34cdc2 cell cycle-regulated protein kinases is required

to initiate mitosis in A. nidulans. Cell 67 (2):283–291 9. Horio T, Oakley BR (2005) The role of microtubules in rapid hyphal tip growth of Aspergillus nidulans. Mol Biol Cell 16(2):918–926 10. Taheri-Talesh N, Horio T, Araujo-Baza´n L, ˜ alva MA, Osmani SA, Dou X, Espeso EA, Pen Oakley BR (2008) The tip growth apparatus of Aspergillus nidulans. Mol Biol Cell 19 (4):1439–1449 11. Takeshita N, Higashitsuji Y, Konzack S, Fischer R (2008) Apical sterol-rich membranes are essential for localizing cell end markers that determine growth directionality in the filamentous fungus Aspergillus nidulans. Mol Biol Cell 19(1):339–351 12. Higashitsuji Y, Herrero S, Takeshita N, Fischer R (2009) The cell end marker protein TeaC is involved in growth directionality and septation in Aspergillus nidulans. Eukaryot Cell 8 (7):957–967 13. Jaspersen SL, Winey M (2004) The budding yeast spindle pole body: structure, duplication, and function. Annu Rev Cell Dev Biol 20:1–28 14. Cavanaugh AM, Jaspersen SL (2017) Big lessons from little yeast: budding and fission yeast centrosome structure, duplication, and function. Annu Rev Genet 51:361–383 15. Knop M, Schiebel E (1997) Spc98p and Spc97p of the yeast γ-tubulin complex mediate binding to the spindle pole body via their interaction with Spc110p. EMBO J 16 (23):6985–6995

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16. Liu P, Zupa E, Neuner A, Bo¨hler A, Loerke J, Flemming D, Ruppert T, Rudack T, Peter C, Spahn C, Gruss OJ, Pfeffer S, Schiebel E (2020) Insights into the assembly and activation of the microtubule nucleator γ-TuRC. Nature 578(7795):467–471 17. Xiong Y, Oakley BR (2009) In vivo analysis of the functions of γ-tubulin-complex proteins. J Cell Sci 122(22):4218–4227 18. Heitz MJ, Petersen J, Valovin S, Hagan IM (2001) MTOC formation during mitotic exit in fission yeast. J Cell Sci 114(24):4521–4532 19. Ho¨o¨g JL, Schwartz C, Noon AT, O’Toole ET, Mastronarde DN, McIntosh JR, Antony C (2007) Organization of interphase microtubules in fission yeast analyzed by electron tomography. Dev Cell 12(3):349–361 20. Konzack S, Rischitor PE, Enke C, Fischer R (2005) The role of the kinesin motor KipA in microtubule organization and polarized growth of Aspergillus nidulans. Mol Biol Cell 16(2):497–506 21. Zekert N, Veith D, Fischer R (2010) Interaction of the Aspergillus nidulans microtubuleorganizing center (MTOC) component ApsB with gamma-tubulin and evidence for a role of a subclass of peroxisomes in the formation of septal MTOCs. Eukaryot Cell 9(5):795–805 22. Gao X, Schmid M, Zhang Y, Fukuda S, Takeshita N, Fischer R (2019) The spindle pole body of Aspergillus nidulans is asymmetrical and contains changing numbers of γ-tubulin complexes. J Cell Sci 132(24): jcs234799 23. Zhang Y, Gao X, Manck R, Schmid M, Osmani AH, Osmani SA, Takeshita N, Fischer R (2017) Microtubule-organizing centers of Aspergillus nidulans are anchored at septa by a disordered protein. Mol Microbiol 106 (2):285–303 24. Samejima I, Lourenc¸o PC, Snaith HA, Sawin KE (2005) Fission yeast mto2p regulates microtubule nucleation by the centrosominrelated protein mto1p. Mol Biol Cell 16 (6):3040–3051 25. Samejima I, Miller VJ, Rincon SA, Sawin KE (2010) Fission yeast Mto1 regulates diversity of cytoplasmic microtubule organizing centers. Curr Biol 20(21):1959–1965 26. Flory MR, Morphew M, Joseph JD, Means AR, Davis TN (2002) Pcp1p, an Spc110prelated calmodulin target at the centrosome of the fission yeast Schizosaccharomyces pombe. Cell Growth Differ 13(2):47–58 27. Hutchins JR, Toyoda Y, Hegemann B, Poser I, He´riche´ J-K, Sykora MM, Augsburg M, Hudecz O, Buschhorn BA, Bulkescher J

(2010) Systematic analysis of human protein complexes identifies chromosome segregation proteins. Science 328(5978):593–599 28. Tovey CA, Tubman CE, Hamrud E, Zhu Z, Dyas AE, Butterfield AN, Fyfe A, Johnson E, Conduit PT (2018) γ-TuRC heterogeneity revealed by analysis of Mozart1. Curr Biol 28 (14):2314–2323.e2316 29. Schermelleh L, Ferrand A, Huser T, Eggeling C, Sauer M, Biehlmaier O, Drummen GP (2019) Super-resolution microscopy demystified. Nat Cell Biol 21(1):72–84 30. Sigal YM, Zhou R, Zhuang X (2018) Visualizing and discovering cellular structures with super-resolution microscopy. Science 361 (6405):880–887 31. Nordzieke S, Zobel T, Fr€anzel B, Wolters DA, Ku¨ck U, Teichert I (2015) A fungal sarcolemmal membrane-associated protein (SLMAP) homolog plays a fundamental role in development and localizes to the nuclear envelope, endoplasmic reticulum, and mitochondria. Eukaryot Cell 14(4):345–358 32. Dempwolff F, Schmidt FK, Herva´s AB, Stroh A, Ro¨sch TC, Riese CN, Dersch S, Heimerl T, Lucena D, Hu¨lsbusch N (2016) Super resolution fluorescence microscopy and tracking of bacterial flotillin (Reggie) paralogs provide evidence for defined-sized protein microdomains within the bacterial membrane but absence of clusters containing detergentresistant proteins. PLoS Genet 12(6): e1006116 33. Takeshita N, Diallinas G, Fischer R (2012) The role of flotillin FloA and stomatin StoA in the maintenance of apical sterol-rich membrane domains and polarity in the filamentous fungus Aspergillus nidulans. Mol Microbiol 83 (6):1136–1152 34. Chen F, Tillberg PW, Boyden ES (2012) Expansion microscopy. J Phys Oceanogr 42:1445–1460 35. Go¨tz R, Panzer S, Trinks N, Eilts J, Wagener J, Turra` D, Di Pietro A, Sauer M, Terpitz U (2020) Expansion microscopy for cell biology analysis in fungi. Front Microbiol 11:574 36. Betzig E, Patterson GH, Sougrat R, Lindwasser OW, Olenych S, Bonifacino JS, Davidson MW, Lippincott-Schwartz J, Hess HF (2006) Imaging intracellular fluorescent proteins at nanometer resolution. Science 313 (5793):1642–1645 37. Sahl SJ, Moerner W (2013) Super-resolution fluorescence imaging with single molecules. Curr Opin Struct Biol 23(5):778–787 38. Wiedenmann J, Gayda S, Adam V, Oswald F, Nienhaus K, Bourgeois D, Nienhaus GU

PALM Microscopy of MTOCs (2011) From EosFP to mIrisFP: structurebased development of advanced photoactivatable marker proteins of the GFP-family. J Biophotonics 4(6):377–390 39. Ishitsuka Y, Savage N, Li Y, Bergs A, Gru¨n N, Kohler D, Donnelly R, Nienhaus GU, Fischer R, Takeshita N (2015) Superresolution microscopy reveals a dynamic picture of cell polarity maintenance during directional growth. Sci Adv 1(10):e1500947 40. Zhou L, Evangelinos M, Wernet V, Eckert AF, Ishitsuka Y, Fischer R, Nienhaus GU, Takeshita N (2018) Superresolution and pulse-chase imaging reveal the role of vesicle transport in polar growth of fungal cells. Sci Adv 4(1): e1701798 41. Takeshita N, Evangelinos M, Zhou L, Serizawa T, Somera-Fajardo RA, Lu L,

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Chapter 21 Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci Tools Melinda Van Kerckvoorde, Matthew J. Ford, Patricia L. Yeyati, Pleasantine Mill, and Richard L. Mort Abstract The cell and cilia cycles are inextricably linked through the dual functions of the centrioles at both the basal body of cilia and at mitotic centrosomes. How cilia assembly and disassembly, either through slow resorption or rapid deciliation, are coordinated with cell cycle progression remains unclear in many cell types and developmental paradigms. Moreover, little is known about how additional cilia parameters including changes in ciliary length or frequency of distal tip shedding change with cell cycle stage. In order to explore these questions, we have developed the Arl13bCerulean-Fucci2a tricistronic cilia and cell cycle biosensor (Ford et al., Dev Cell 47:509–523.e7, 2018). This reporter allowed us to document the heterogeneity in ciliary behaviors during the cell cycle at a population level. Without the need for external stimuli, it revealed that in several cell types and in the developing embryo cilia persist beyond the G1/S checkpoint. Here, we describe the generation of stable cell lines expressing Arl13bCerulean-Fucci2a and open-source software to aid morphometric profiling of the primary cilium with cell cycle phases, including changes in cilium length. This resource will allow the investigation of multiple morphometric questions relating to cilia and cell cycle biology. Key words Cell cycle, Cilia, ImageJ, Image analysis, Live cell imaging, Fucci2A, Biosensor, Morphometrics, Cell division, Ciliogenesis

1

Introduction Cilia are small microtubule-based protrusions projecting from the surface of most mammalian cell types [1]. Akin to cellular “antennae,” primary cilia play critical roles in sensing and eliciting cellular responses to environmental and developmental signals, from embryogenesis through to adult tissue homeostasis. These specialized signaling “organelles” are compartmentalized with a distinct protein and lipid composition maintained by dedicated

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-1538-6_25 Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_21, © The Author(s) 2021, Corrected Publication 2021

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selective trafficking modules and structural elements [2]. Defects in cilia structure and/or function result in a group of genetic disorders termed the ciliopathies, where the alterations in gross cilia morphology observed in patient cells have been proposed to underlie the clinical phenotypes [3]. However, it is also possible that biochemical changes in the cilia compartment may affect cilia function, at a transient stage in cell cycle progression or without altering length or stability at all. Therefore, better tools are required to resolve cilia dynamics in order to understand gene function. Moreover, cilia are not static structures. The assembly, elongation and resorption of the ciliary axoneme is regulated by cytoskeletal dynamics that define centrosome positioning across the cell cycle [4]. Ciliogenesis is a highly regulated process that is led by the mother centriole, destined to become the basal body of the nascent cilium. The basal body acts as a cilia microtubule organizing center (MTOC) nucleating the positive end of the microtubules and orientating them out of the cell into the soon-to-be ciliary tip. Subsequent elongation of cilia requires anterograde and retrograde intraflagellar transport (IFT) along the axoneme and is critical for its function [5]. In order to divide, cells must dismantle their cilia to free up the centrioles and form the mitotic spindle. How this process occurs in asynchronous populations like the rapidly growing embryo, versus synchronized cells in culture which reenter the cell cycle following serum addition, remains unclear [6]. In contrast to synchronized culture experiments, where there are two distinct phases of cilia disassembly [7, 8], the precise timing of cilia assembly and disassembly in relation to the cell cycle in actively cycling cells is still ill-defined. We and others have previously shown that whilst the initial cilia assembly time is more variable in terms of kinetics, cilia disassembly during interphase occurs in a tight window ahead of mitosis across the asynchronous population [1, 9]. In an actindependent process they term “decapitation,” Phua et al. demonstrated that the release of vesicles from the distal ciliary tip in response to serum was required for ciliary disassembly and cell cycle reentry, including from quiescence [10]. Recent work by Mirvis et al. [11] showed that in IMCD3 cells a rapid deciliation which involved shedding of the ciliary axoneme and membrane was the predominant mode of deciliation, one dependent on katanin activity, in response to serum. There is much to be learned about the heterogeneity in cell behaviors exhibited during cilia disassembly in asynchronous populations. For example, how does cilia length change during cell cycle progression? Does decapitation occur at specific times during the cell cycle? And do cilia disassemble in a cell-type and context-dependent manner? To better understand these processes, we used Fucci2a (Fluorescent Ubiquitination-based Cell Cycle Indicator 2a), a multicistronic construct that encodes two cell cycle biosensors fused using the Thosea asigna virus 2A (T2A) self-cleaving peptide sequence [12]. hCdt1-mCherry incorporates a truncated form of human

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CDT1 (amino acids 30–120) fused to the mCherry fluorescent protein, whereas hGem-mVenus includes a truncated form of human Geminin (amino acids 1–110) fused to mVenus. Fucci2a allows discrimination between cell cycle phases in live cells using imaging of mCherry and mVenus abundance [12]. CDT1 abundance peaks during the G1 cell cycle phase in order to recruit the replicative DNA helicase, whereas Geminin is stable from S until M phase as an inhibitor of CDT1 ensuring that DNA replication happens only once per cell cycle [13]. Hence, the advantage of coexpressing these fluorescent markers is the ability to image their reciprocal abundance in proliferating cells without artificially perturbing cell cycle progression. To this construct we added a marker to image the primary cilium across the different cell cycle oscillations, by fusing the ciliary GTPase ARL13B to the fluorescent protein mCerulean [1]. This fusion protein was then fused with Fucci2a using the porcine teschovirus-1 P2A self-cleaving peptide sequence (Fig. 1a). We demonstrated in asynchronously cycling NIH-3 T3 cells that cilia assemble across a wide window of cell cycle stages in interphase, and that these cilia persist until just prior to mitosis [1]. Further comparisons tracking the fate of daughter cells revealed that daughter cells from ciliated mother cells assemble their primary cilia faster than from nonciliated mothers and that one daughter cell tends to initiate ciliogenesis earlier than the other, confirming previous studies [1, 14]. We also showed with the development of a Cre-inducible R26Arl13b-Fucci2aR reporter mouse that we could generate high-resolution images enabling tracking of the cell cycle stage and ciliation state of individual cells in culture and in all tissues examined, both embryonic and adult [1]. Here, we present detailed methods and novel analytical tools to streamline monitoring cilia behaviors across cell cycle transitions. We describe the generation of stable cell lines expressing Arl13bCerulean-Fucci2a, their live imaging by confocal microscopy and the simultaneous analysis of cell cycle and cilia dynamics using Fucci Tools software to generate high quality tracking data and multifactorial plots of cell cycle and cilia cycle kinetics.

2

Materials Prepare all solutions using ultrapure water and store at room temperature (unless indicated otherwise).

2.1

Cell Culture

1. Mouse Flp-In™-3T3 cells (ThermoFisher, cat. no. R76107) or Arl13bCerulean-Fucci2a 3T3 cells ([1] - Riken BRC accession #RCB5029) (see Note 1). 2. Culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM), 10% Fetal bovine serum (FBS), 1% Penicillin/Streptomycin, 25 mM D-glucose, 4 mM L-glutamine, 1 mM sodium pyruvate. Store 4
C.

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Fig. 1 Arl13bCerulean-Fucci2a—a tricistronic cell and cilia cycle biosensor. (A) Schematic of the Arl13bCerulean-Fucci2a biosensor. A full-length mouse Arl13b cDNA was fused to the Cerulean fluorescent protein and concatenated to Fucci2a cell cycle probes hCdt1-mCherry and mVenus-hGem, with viral 2a peptide sequences in a multicistronic construct. (B) Cell cycle phase specificity of the probes. mCherry abundance peaks in G1 and is lost at the G1–S transition, whereas mVenus is present through S/G2/M phases and is lost at mitosis. Primary cilia localization of ARL13B-Cerulean can be observed from G1 though S/G2 until M phase (see Note 13). (C) Asynchronous, Flp-In™ 3T3 cells stably expressing Arl13bCerulean-Fucci2a. (D) The Flp-In™ system is used to make a single integration of the pcDNA5-CAG-Arl13bCerulean-Fucci2a plasmid into the genomic Flp-In™ locus of a compatible cell line. (E) Integration of the plasmid switches the selection from zeocin to hygromycin. Scale bar in C ¼ 100 μm

3. Imaging medium: Phenol red-free Dulbecco’s Modified Eagle’s Medium (DMEM), 10% fetal bovine serum (FBS), 1% Penicillin/Streptomycin, 25 mM D-glucose, 4 mM L-glutamine, 1 mM sodium pyruvate. Store 4
C. 4. Freezing medium: 45 mL culture medium, 5 mL dimethyl sulfoxide (DMSO). Store 4
C. 5. Opti-MEM (ThermoFisher, cat. no. 31985062). Store 4
C.

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6. Phosphate buffered saline (1 PBS). 7. 0.25% Trypsin-EDTA (1). Store 4
C. 8. Zeocin: 100 mg/mL, store at -20
C. 9. Hygromycin B: 50 mg/mL, store at 4
C. 10. 1 TE buffer: 10 mM Tris–HCl pH 7.4, 1 mM EDTA. 11. 15 mL sterile, conical tubes. 12. 1.5 mL sterile, tubes. 13. 2 mL cryovials. 14. Cell culture flasks 75 cm2. 15. Parafilm. 16. 24-well glass bottom plates such as ibidi μ-Plates (Ibidi, cat no. 82406) or Greiner Sensoplates (Greiner, cat. no. 662892). 17. Tissue culture incubator (37
C, 5% CO2). 18. Benchtop centrifuge. 19. Hemocytometer. 20. Tissue culture microscope with 10 objective to count cells. 21. Neon Transfection no. MPK5000).

System

(ThermoFisher,

cat.

22. Neon 100-μL electroporation tips (ThermoFisher, cat. no. MPK10096). 23. Neon electroporation tube (ThermoFisher, cat. no. MPT100). 24. Neon E2 electrolytic no. MPK10096).

buffer

(ThermoFisher,

cat.

25. Neon Buffer R (ThermoFisher, cat. no. MPK10096). 26. Arl13bCerulean-Fucci2a expressing Flp-In™ compatible pcDNA™5-CAG-Arl13bFucci2a plasmid: 1 μg/μL in TE (Riken BRC accession #RDB16057 (see [1]). 27. pOG44 Flp recombinase expressing plasmid: 1 μg/μL in TE (ThermoFisher, cat. no. V600520). 28. pcDNA™5/FRT/CAT positive control plasmid: 1 μg/μL in TE (ThermoFisher, cat. no. V601020). 2.2 Confocal Microscopy

1. Microscope: Laser scanning confocal platform (e.g., Nikon A1R) or spinning disc confocal platform (e.g., Andor Dragonfly) (see Note 2). 2. Hardware autofocus (e.g., Nikon Perfect Focus System (PFS)) (see Note 3). 3. Environmental chamber to maintain a constant temperature of 37
C and deliver humidified 5% CO2 to the sample (see Note 4). 4. Stage insert for microscope that accommodates multiwell plates.

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5. High numerical aperture dry 20 or 40 objective for multiwell imaging (see Note 5). 6. Acquisition software (e.g., Nikon NIS Elements or Andor Fusion). 2.3

Image Analysis

1. Fiji distribution of ImageJ. 2. Fucci_Tools.ijm macro toolset and fucci_tools_profile.txt (https://github.com/richiemort79/fucci_tools).

3

Methods

3.1 Generation of Stable Arl13bCeruleanFucci2a Expressing Cell Lines

Given the toxicity of the Fucci2a probes when they are overexpressed and the variability in expression levels generated between cells, transient transfection techniques are not recommended when imaging Arl13bCerulean-Fucci2a. Therefore, we describe the generation of a stable cell line using the Flp-In™ system (ThermoFisher). The Flp-In™ system allows for the integration of a single copy of a Flp-In™ compatible plasmid (harboring a single FRT site) into the FRT site of a Flp-In™ locus in a compatible cell line (Fig. 1D–E). Integration of the plasmid results in a switch of selection cassette allowing for selection of successful integrants. 1. Untargeted Flp-In™ NIH 3 T3 cells (or other Flp-In™ compatible cells, see Note 1) are maintained in culture media supplemented with 100 ng/mL Zeocin in 75 cm2 flasks (see Note 6). Approximately 1  107 cells are required for each transfection. 2. Prewarm 15 mL of Opti-MEM in 3  75 cm2 tissue culture flasks. 3. Add 3 mL of electrolytic buffer E2 to the electroporation tube and insert into the Neon pipette station. 4. Label three 1.5 mL tubes: negative control, positive control, and Arl13bCerulean-Fucci2a. Into each tube, prealiquot in 27 μL of pOG44 plasmid. In the negative control tube, add 3 μL TE. In the positive control tube, add 3 μL pcDNA 5/FRT/CAT plasmid. In the experimental tube, add 3 μL of pcDNA 5-CAG-Arl13bFucci2a (see Note 7). 5. Wash the Flp-In™ cells with PBS, trypsinize, pellet in a 15 mL tube, wash with PBS and count with a hemocytometer. Resuspend pellet in a volume of Neon Buffer R required for the electroporation (i.e., aliquots of 5  107 cells each in 90 μL Neon Buffer R, at a final concentration of ~5  107 cells/mL). For example, this workflow requires nine aliquots total for the three conditions each requiring two experimental and one backup (see Note 8).

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6. For each electroporation combine 10 μL of the DNA from step four with one 90 μL aliquot of cells from step 5. Use the Neon electroporation system (2 pulses: 1350 V, 20 ms for Flp-In™ 3 T3) to electroporate the Flp-In™ cells. For each condition use 2  100 μL electroporations (use 1 100 μL Neon tip twice per tube, change between conditions) and dispense both into the same 75 cm2 flask containing prewarmed Opti-MEM. 7. Place flasks in the incubator for 24 h. 8. After 24 h, remove the Opti-MEM and replace with 15 mL of culture media supplemented with 50 ng/mL Hygromycin B. 9. Grow cells under Hygromycin B selection changing the media every 3–4 days for 14 days and regularly inspect for clones. 10. Following 14 days of Hygromycin B selection or when well defined clonal colonies can be observed, trypsinize the cells and passage to generate a polyclonal Arl13bCerulean-Fucci2a expressing cell line. Using the Flp-In™ system, generated cells are isogenic (i.e., integration occurs into the same genomic locus in every clone, such that all clones should be identical). This can be confirmed by determining loss of Zeocinresistance, loss of lacZ expression as well as expression of Arl13bCerulean-Fucci2a biosensor cassette. 11. Cryopreserve early passage cells in 1 mL aliquots of ~106 cells/ mL in freezing medium using 2 mL cryovials. Freeze cells following standard procedures and store long-term in liquid nitrogen. 3.2 Live Cell Imaging of Arl13bCeruleanFucci2a Expressing Cells

3.3 Experimental Setup

Successful imaging of Arl13bCerulean-Fucci2a expressing cells requires striking a balance between image acquisition speeds, laser exposure and the number of parallel experiments one wishes to conduct. Consider the time it will take to image each position on the plate when deciding how many conditions to include in an experiment. When analyzing cilia and cell cycle dynamics in parallel, we find it best to image each position every 10–20 min for 48–72 h in order to capture multiple mitosis-to-mitosis transitions. However, catching rarer or transient events, such as decapitation, may require more frequent intervals, consequently decreasing the number of parallel experiments or positions which can be imaged for each time point. We find that confocal systems that incorporate an environmental enclosure that encompasses the entire stage and that employ hardware autofocus are the most reliable for long term time-lapse because they are best at minimizing temperature variations as well as air currents or vibrations and therefore sample drift across long imaging sessions. 1. Wash the Flp-In™ Arl13bCerulean-Fucci2a cells with PBS, trypsinize, pellet in a 15 mL tube, wash with PBS and count with a hemocytometer.

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2. Seed the cells into a 24-well glass bottomed plate containing imaging media in densities ranging from 1  105 to 5  105 cells per well (see Note 9). Use at least 1000 μL media per well to account for evaporation across the imaging session. Incubate the plate for at least 4 h prior to imaging to give the cells time to attach. 3. Fill the unused surrounding wells with 1000 μL of sterile PBS to further minimize evaporation of the medium in adjacent wells. 4. If gassing with a needle, make a hole in the side of the 24-well plate with a hot hypodermic needle. 5. If not using a stage-top enclosure, seal the outside of the plate with a strip of Parafilm (see Note 10). 6. Prewarm the environmental chamber and microscope stage for at least 1 h prior to imaging to prevent stage drift. 7. Insert the glass bottomed 24-well plate into the microscope stage plate insert and if gassing with a needle, insert the needle into the hole in the plate, inside the chamber at 37
C. 3.4

Imaging Setup

1. Choose a lens—if acquiring data at multiple positions, dry lenses allow for ease of movement between positions over long time periods (see Note 5). 2. In order to visualize Cerulean, Venus, and Cherry, set the pinhole size according to the longest wavelength channel (Cherry) being recorded. This ensures that all channels have the same optical section thickness and that no channel has a pinhole size less than 1 AU. 3. For each channel, check pixel saturation and adjust to minimize over-saturation in order to maintain the full dynamic range. 4. To image multiple positions and/or wells, define the XY coordinates for each independent position using your imaging platform’s acquisition software. 5. We recommend including a small z-stack at each acquisition point to account for stage drift. This does not need to be the optimal “step size” calculated by the image acquisition software, larger step sizes will increase the rate of acquisition and decrease the amount of light exposure to the sample. Capture a time series overnight at ~20-min intervals for upward of 48 h in order to ensure that many complete mitoses are captured.

3.5 Image Analysis with Fiji and Fucci Tools

ImageJ is a freeware image analysis platform based on NIH Image [15]. Fiji is a distribution of ImageJ that includes many useful plugins and imaging libraries preinstalled (https://fiji.sc/) [16]. In order to investigate the dynamics of the cilia and cell cycles, the fluorescence intensity of the two Fucci2a probes must be

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monitored and morphological measurements of the cilia must be made in parallel. We present “Fucci Tools” a macro toolset written for Fiji that allows users to manually track cell nuclei and cilia in order to capture this information. Fucci Tools allows for the easy tracking of the daughters of a cell division, summarizes the tracking data, fluorescence intensities and morphometric data in a comprehensive results table and allows plotting of the data in several useful formats. Substacks cropped to the nucleus of interest are generated dynamically during tracking. The tracking data can be parsed to the .mdf2 format allowing compatibility with MTrackJ, an ImageJ plugin to visualize and measure cell tracking statistics. Users can define an experimental profile that captures the key parameters from their image acquisition to streamline their workflow. 3.6 Installation of Fucci_Tools.ijm and Initial Definition of an Experimental Profile

Fucci Tools is free software provided under a Massachusetts Institute of Technology (MIT) license. Please read the license file included in the Fucci Tools GitHub repository. In order to help users set up and troubleshoot Fucci Tools, we have included the example dataset used in this chapter [1], along with an example results table of tracking data that can be imported to test the plotting functions. 1. Comprehensive instructions on the installation of Fiji can be found on the Fiji website (https://fiji.sc/). 2. Once Fiji has been successfully installed, download fucci_toolsmaster.zip from GitHub (https://github.com/richiemort79/ fucci_tools) and extract the archive. 3. Place Fucci_Tools.ijm in the Fiji/macros/toolsets folder of your Fiji installation. 4. Place fucci_tools_profile.txt in the Fiji/macros folder of your Fiji installation. Restart Fiji. 5. Next define the image acquisition parameters that were used to capture the dataset. Open fucci_tools_profile.txt in a Text editor and edit the listed parameters (for an explanation see Table 1).

3.7 Performing an Analysis with Fucci Tools

1. Launch Fiji and open the dataset generated by your image acquisition software above. Fiji supports a wide number of proprietary image formats (see Note 11). 2. If a z-stack was captured during image acquisition, first flatten the data by either selecting an individual z-plane for analysis or by using a maximum intensity projection. 3. On the main ImageJ toolbar click the “More Tools” menu button “>>” and select “Fucci Tools” from the dropdown menu.

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Table 1 Summary of the parameters defined by fucci_tools_profile.txt to establish an experimental profile Parameter

Description

Example

pro_time_step

Time interval between frames in minutes

20

pro_scale

Image resolution in microns/pixel

0.42

pro_number_channels Number of channels acquired in the dataset

4

pro_channels

The five available LUTS for visualization are: “Cyan, Green, Red, Magenta, Grays”

static

pro_channel_order

Map the above LUTS to the channels acquired in the dataset (1 ¼ Cyan, 2 ¼ Green, 3 ¼ Red, 4 ¼ Magenta, 5 ¼ Grays)

12304

pro_view

Define which channels are visible during the tracking analysis (1 ¼ visible, 0 ¼ hidden)

11100

pro_norm

Write 1 for channels for which the intensity values must be normalized for plotting, 0 if not

11111

pro_crop

Defines the dimensions of the required substack window in pixels

50

pro_track

Write “true” if you would like to track and measure a cilium “false” if true not

pro_track_step

Define how often you would like to make measurements of the cilum 1 (1 ¼ measure every frame)

pro_view2

Define which channels are visible during the cilia analysis (1 ¼ visible, 10000 0 ¼ hidden)

Use this file to define the key parameters used during image acquisition. These parameters will then be loaded automatically when Fucci Tools is initialized.

4. Click the “Initialize” Tool (Fig. 2A-1). The required parameters are loaded automatically from the fucci_tools_profile. txt file but can also be manually refined at this point. If you wish to track a cilium, remember to tick the “Track Cilia” checkbox in the initialize dialog. 5. Scan through the time-lapse data to identify a cell that divides twice during the time lapse (see Note 12). Fucci Tools allows one to track the mother cell into the first division and then track both daughters. When a mother cell of interest has been identified click the “Interactive Measure Tool” (Fig. 2A-2). The tool button becomes greyed out when it is active. 6. With “Interactive Measure Tool” (Fig. 2A-2) selected click on the cell nucleus. A new track (Track 1) is added to the results table. The intensities of the individual channels will be recorded in the results table and a cropped version of the dataset will be recorded in the Substack window (Fig. 3C). 7. If “Track Cilia” was selected in the initialize dialog, Fucci Tools will prompt the user to make this measurement. Simply click

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Fig. 2 Fucci Tools toolset and initialize dialog. (A) The Fucci Tools appear in the main Fiji window as a toolset consisting of seven buttons. The function of which is as follows: (1) “Initialize Tool” to initialize the analysis and load the key parameters; (2) “Interactive Track Tool” for manually tracking cells; (3) “Add Track Tool ” to add a new track to the analysis; (4) “Add Mitosis Tool” to record mitotic events; (5) “Switch Daughter Tool” to record the end of a complete mitosis and return to its origin; (6) “Fucci Plot Tool” for plotting the acquired data; (7) “Parse to mdf2 Tool” to export the tracking data to MTrackJ for further analysis. (B) The initialize dialog captures the key parameters for the analysis. Parameters are explained fully in Table 1. In order to track a cilium as well as analyzing the Fucci probe intensities, one must check “Track Cilia,” enter how often a cilium measurement is required to be made and which channels to view while measuring

“OK” in the dialog if no cilia are present at this time point otherwise use the “Segmented Line” tool to define the cilia. Double click to close the line and then press “OK” in the dialog window. 8. The time lapse will then advance to the next frame so that the user can click and make measurements again. 9. At the point that the mother cell undergoes cytokinesis click the “Add Mitosis Tool” (Fig. 2A-4) and then continue to track one of the daughters of the division. A new track (Track 1a) is added to the results table and a new Substack is opened. 10. When the first daughter cell divides click the “Switch Daughter” tool (Fig. 2A-5). This will return the time lapse to the timepoint at which the mother underwent mitosis so that second daughter cell can be tracked. The mitosis point will be circled. A new track (Track 1b) is added to the results table and a new Substack is opened. 11. At any time, a user can click the “Add Track Tool” (Fig. 2A-3) to start tracking a new cell. 3.8 Plotting of the Results Table Data

The manner in which one might plot the data generated by Fucci Tools will depend on the experimental question. However, we have included a number of plotting methods that will help in the initial

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Fig. 3 Fucci Tools toolbar and tracking features. (A) ImageJ main window and Fucci Tools toolset. (B) A mitosis is tracked by following the mother followed by the two daughters the positions of the nuclei in each frame are recorded and shown as an overlay (daughter a ¼ yellow, daughter b ¼ red). (C) Substacks of the nuclei are recorded for each track to make figure preparation and data validation easier. (D) A comprehensive tracking table is generated that records the nuclei positions and fluorescence intensity in each channel as well as morphological features of cilia. Scale bar In B ¼ 100 μm, scale bar in C ¼ 10 μm

exploration of datasets generated (Figs. 4 and 5, see Note 13). As well as the plotting functions described below, montages (Fig. 5A) can be easily generated from the substacks collected during tracking (Fig. 3C) using standard ImageJ commands. Four types of plots are available in Fucci Tools: (1) “Single” plots of each complete mitosis (Fig. 5B) are generated either as a stack of images or as a montage (Fig. 4B). These plots included the normalized intensity of each channel chosen in the plot dialog (Fig. 4A) and an overlay that shows the normalized length of the cilia (if present) with time. Use these plots to examine how cilia assembly varies between individual cell cycles. (2) “Interpolated Mean” plots (Fig. 5C) attempt to provide a method to examine the mean behavior of each probe across the length of a cell cycle. They are generated by interpolating the probe intensity and cilia length data using linear resampling so that a complete mitosis is represented by 100 arbitrary time-points. A mean and 95% confidence interval is then generated for each

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Fig. 4 The Fucci Tools plotting dialog and montage view. (A) The plotting dialog allows you to check and amend acquisition parameters as well as choose the plot types you wish to visualize. (B) Selecting “Single” and “Montage” displays a montage of all the complete mitosis tracks in the results table. Use the dialog to define the channels and features that you wish to plot

fluorescent channel and for cilia length at each time point. (3) “Interpolated Fucci with Cilia Overlay” plots allow plotting of cilia assembly and disassembly times over mean fluorescence intensity plots (Fig. 5D). (4) In order to examine the heterogeneity in assembly and disassembly times, “Interpolated Cilia Length” plot trims the cilia length data to remove time points before assembly and after disassembly and then interpolates cilia length using linear resampling so that time is represented as 100 arbitrary points. A mean and 95% confidence interval is then generated for cilia length at each timepoint. This allows examination of the mean behavior during cilia assembly maintenance and disassembly across all the tracks in the results table. 3.9 Concluding Remarks

Careful observation at cellular and subcellular resolution with live imaging allows elucidation of the underlying intercellular heterogeneity of biological processes. Isogenic cell lines (or mouse tissues) expressing our Arl13bCerulean-Fucci2a biosensor allow users to capture a variety of cilia parameters in parallel to precise cell cycle stages. Here, we describe Fucci Tools, an analysis package, designed to enable users to easily elucidate multiple cilia and cell cycle parameters from imaging files and explore the data in a variety of informative formats. Fucci Tools allows examination of cilia assembly/disassembly and length mapped to a standardized cell cycle across many individual cells (Fig. 5B–E). We observe three phases in cilia dynamics during cell cycle progression: (1) fast assembly, with a heterogenous start time, up to ~50% of final length; (2) growth (+50% length) and maintenance until just before M phase; and (3) rapid disassembly at M-phase. We examined cell

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Fig. 5 Advanced plotting of cilia and cell cycle dynamics. (A) Substacks of a single Arl13bCerulean-Fucci2a nucleus progressing through the cell and cilia cycles. (B) Individual plot showing normalized fluorescence intensity of the probes. The reciprocal mCherry-hCdt1 (magenta) and mVenus-hGem (yellow) Fucci2a peaks indicate the G1 and S/G2/M phases of the cell cycle respectively. Normalized cilia length is indicated in cyan. (C) Aggregated, interpolated plot of n ¼ 16 cell cycles. (D) Aggregated Fucci2a intensities with cilia assembly and disassembly times overlaid showing the diversity in assembly time with respect to cell cycle phase. (E) Aggregated plot of the normalized cilia plot across one cilia cycle. Error bars in B–D ¼ 95% Confidence Intervals. Note: adjacent nuclei in A were masked for presentation purposes. A ¼ ARL13B, C ¼ hCdt1, G ¼ hGem. Scale bar in A ¼ 10 μm

cycle progression times for adjacent cells in asynchronous cultures from two experiments to ask whether cell cycle times differ in ciliated cells (Fig. 6). While mean cell cycles varied between experiments (experiment A: ciliated ¼ 37.7  8.7 h vs. unciliated ¼ 30.9  8.1 h; experiment B: ciliated ¼ 44.1  5.0 h vs. unciliated ¼ 38.1  9.0 h, nonsignificant differences), it was intriguing that ciliated cells consistently took around 6 h longer to divide (Fig. 6A). When we asked what stage of the cycle they spent more time in, there was no clear trend (Fig. 6B). In our analysis (Fig. 7), we struggled to find many examples (n ¼ 1/25 ciliated cells) of decapitation prior to cilia

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Fig. 6 Cell cycle kinetics in adjacent ciliated versus unciliated cells. (A) Total cell cycle times (mitosis-tomitosis) in hours (h) from ciliated (cyan) versus unciliated (white) cells from two independent experiments. Mean and standard deviation (error bars) displayed. No significant difference between conditions (Student ttest, P > 0.05). Differences such as cell density between experiments may explain differences in cell cycle times. (B) Individual proportions of cell cycle phases (G1: magenta vs. S/G2/M: yellow) are represented grouped as ciliated versus unciliated cells. Mean proportion in G1 per cohort plotted as dotted black line. No significant difference between conditions (Student t-test P > 0.05). Expt experiment

disassembly [10], but this could reflect differences operating in asynchronous cycling populations versus those stimulated by serum. It may also be possible that our time acquisition intervals are not frequent enough to capture these transient events. Adjusting image acquisition parameters to meet the requirements of your biological question is key to a successful experiment. These tools will be important for comparing effects on cell cycle and cilia dynamics between genotypes, but also more fundamental questions. For example, how does a transient organelle which is required for mitogenic signals, like Hedgehog signaling, respond if it is only present for a portion of the cell cycle. Intriguing work from Ho, Tsai and Stearns using a medulloblastoma cell line, as well as cerebellar granular precursors cells from which they are derived [17], revealed that cilia also persisted into S phase where they mediate robust Hh signaling. If and how cilia content may change with the cell cycle is another open question- with evidence suggesting this may be the case for phosphoinositide content [10] and signaling competence in subsequent interphases [17]. Indeed, a recent study from the Mick lab, revealed specific and rapid remodeling of the cilia proteome in the timescale of 10s of minutes in response to Hh ligand [18]- although how this occurred in the background of the cell cycle remains unclear. These observations raise questions as to the physiological consequence of having a cilium at different stages of the cell cycle. Can they function as

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Fig. 7 Ectocytosis/decapitation is a rare event in exponentially expanding cells. Out of the 25 ciliated full mitosis-to-mitosis events analyzed in these two data sets, only one decapitation event was observed. (A) Montage at 60 min timepoints reveals a cilium present from early in G1 through to just prior to nuclear envelope breakdown. (B) At 20 min timepoints the ectocytosis event can be observed, the tip vesicle persisted for 2.2 h. (C) Individual plot showing normalized fluorescence intensity of the probes. Normalized cilia length is indicated in cyan. Decapitation occurred in G2, but cilia persisted for another 6.6 h. Scale bar in A, B ¼ 10 μm

effective ‘signaling organelles’ if present regardless of cell cycle stage or are their contents and signaling competence regulated in a cell cycle specific manner? Our tools which allow simultaneous analysis of cell cycle stage, cilia kinetics and morphology provide a very powerful means to investigate these open questions.

4

Notes 1. It is also possible to use other Flp-In™ compatible cell lines, which for cilia research include RPE-1 Flp-In™ [19] or IMCD3 Flp-In™ [20], as long as they include an FRT site integrated at a single genomic locus. Flp-In™ cell lines can also be generated de novo by integration of the pFRT/lacZeo construct (ThermoFisher).

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2. These require excitation and emission filters to allow imaging of Cerulean (433 nm/475 nm), Venus (515 nm/528 nm) and mCherry (587 nm/610 nm) fluorescent proteins. 3. Hardware autofocus (i.e., Nikon’s PFS) is highly advantageous for eliminating stage drift especially in Z greatly reducing the number of failed multiday imaging experiments. 4. We have successfully used both microscope stage mounted systems that heat and provide CO2 (e.g., Okolabs stage-top incubator) and full environmental chambers that enclose the stage and objective lenses (e.g., Solent Scientific enclosures) in combination with either CO2 delivery by hypodermic needle to a Parafilm sealed plate or using a passive Okolabs stage-top chamber for CO2 delivery. 5. On most microscopes, multiposition acquisitions over long periods necessitate use of dry objectives as spreading of immersion media (oil or water) across the surface of a multiwell plate over time will interrupt focus of a high magnification lens. Some manufacturers like Leica offer a Water Immersion Micro Dispenser which overcomes this potential problem during long-term live-cell imaging experiments like these. If using a dry 40, check the correction collar is set for imaging through a standard #1.5 cover glass found on the bottom of Ibidi plates. 6. It is imperative that Flp-In™ compatible cell lines are maintained during passaging according to the specific guidelines for retaining the FRT site integrated at a single genomic locus. In the case of untargeted Flp-In™ NIH 3T3 cells, this requires culturing in culture media supplemented with 100 ng/mL Zeocin in order to retain ability to target efficiently. 7. To avoid arcing, the total volume of DNA must not exceed 10% of the volume of cells. For transfection grade plasmid DNA, we recommend using commercial midi or maxi DNA prep kits which allow elution in minimal volumes of endotoxin-free buffer. 8. Prealiquoted plasmid reagents will help minimize the time cells spend in Neon Buffer R. A quick workflow to get cells back into medium is important for a successful transfection with optimal viability. When using the Neon transfection system, we recommend preparing transfections mixes (n + 1), as cells can be quite viscous in Neon Buffer R and there is a need to avoid bubbles taken up in the tip to avoid arcing (i.e., a spark). Discard cells if arcing occurs. 9. Experiment with seeding density in order to get a density that allows easy tracking of cells. Especially for NIH 3T3 cells, we recommend plating at low density to allow imaging of exponentially growing cultures for tracking rates of ciliation in

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relation to cell cycle. Moreover, some cell types including Flp-In™ 3T3 tend to migrate over one another and can be hard to track. We find that mixing them 1:3–1:2 with unlabeled parental cells can help with this. 10. Take care to avoid pulling the Parafilm too tight as it may then fail overnight. Ensure no Parafilm underlaps the plate as this will prevent it from sitting flat on the microscope stage. 11. Fiji comes with the Open Microscopy Environment (OME) Bio-formats plugins (https://www.openmicroscopy.org) preinstalled. Bio-formats supports the opening of many proprietary file types (e.g., Nikon .nd2, Zeiss .czi, Olympus .vsi) as multidimensional “Hyperstacks” in Fiji/ImageJ. If your Imaging format is not supported export the data from your acquisition software as a .tif stack. 12. It is often easiest to scan backward from the end of the timelapse to identify cells to track. 13. Whilst data from the three channels in the biosensors are associated with real spectra and these are used for Fucci Tools, for accessible representation purposes we have altered the use of “false colour” in the three colour confocal images so that they are not red, green, and cyan but rather magenta, yellow, and cyan. Similarly, we use this palette to represent dynamics in our line art graphs too. These color combinations that are more favourably viewed by about 10% of the population who are red/green colour blind (https://imagej.nih.gov/ ij/docs/guide/146-9.html and https://www.ascb.org/sci ence-news/how-to-make-scientific-figures-accessible-toreaders-with-color-blindness/).

Acknowledgments The authors are grateful to the IGMM Advanced Imaging Resource for imaging support and the BSCB for summer studentship support for MVK. PLY and PM were supported by core funding from the UKRI Medical Research Council (MC_UU_00007/14). RLM was supported by funding from North West Cancer Research (CR1132) and the NC3Rs (#NC/ T002328/1). References 1. Ford MJ, Yeyati PL, Mali GR et al (2018) A cell/cilia cycle biosensor for single-cell kinetics reveals persistence of cilia after G1/S transition is a general property in cells and mice. Dev Cell 47:509–523.e7

2. Garcia-Gonzalo FR, Reiter JF (2017) Open sesame: how transition fibers and the transition zone control ciliary composition. Cold Spring Harbor Perspect Biol 9(2):a028134

Analysis of Cilia and Cell Cycle Dynamics 3. Reiter JF, Leroux MR (2017) Genes and molecular pathways underpinning ciliopathies. Nat Rev Mol Cell Biol 18:533–547 4. Bernabe´-Rubio M, Alonso MA (2017) Routes and machinery of primary cilium biogenesis. Cell Mol Life Sci 74:4077–4095 5. Ishikawa H, Marshall WF (2017) Intraflagellar transport and ciliary dynamics. Cold Spring Harbor Perspect Biol 9(3):a021998 6. Wang L, Dynlacht BD (2018) The regulation of cilium assembly and disassembly in development and disease. Development 145(18): dev151407 7. Tucker RW, Scher CD, Stiles CD (1979) Centriole deciliation associated with the early response of 3T3 cells to growth factors but not to SV40. Cell 18:1065–1072 8. Pugacheva EN, Jablonski SA, Hartman TR et al (2007) HEF1-dependent Aurora A activation induces disassembly of the primary cilium. Cell 129:1351–1363 9. Anderson CT, Stearns T (2009) Centriole age underlies asynchronous primary cilium growth in mammalian cells. Curr Biol 19:1498–1502 10. Phua SC, Chiba S, Suzuki M et al (2017) Dynamic remodeling of membrane composition drives cell cycle through primary cilia excision. Cell 168:264–279.e15 11. Mirvis M, Siemers KA, Nelson WJ et al (2019) Primary cilium loss in mammalian cells occurs predominantly by whole-cilium shedding. PLoS Biol 17:e3000381 12. Mort RL, Ford MJ, Sakaue-Sawano A et al (2014) Fucci2a: a bicistronic cell cycle reporter that allows Cre mediated tissue specific expression in mice. Cell Cycle 13:2681–2696

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13. Sakaue-Sawano A, Kurokawa H, Morimura T et al (2008) Visualizing spatiotemporal dynamics of multicellular cell-cycle progression. Cell 132:487–498 14. Paridaen JTML, Wilsch-Br€auninger M, Huttner WB (2013) Asymmetric inheritance of centrosome-associated primary cilium membrane directs ciliogenesis after cell division. Cell 155:333–344 15. Schneider CA, Rasband WS, Eliceiri KW (2012) NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9:671–675 16. Schindelin J, Arganda-Carreras I, Frise E et al (2012) Fiji: an open-source platform for biological-image analysis. Nat Methods 9:676–682 17. Ho EK, Tsai AE, Stearns T (2020) Transient primary cilia mediate robust hedgehog pathway-dependent cell cycle control. Curr Biol 30:2829–2835.e5 18. May EA, Kalocsay M, Galtier D’Auriac I et al (2020) Time-resolved proteomic profiling of the ciliary Hedgehog response reveals that GPR161 and PKA undergo regulated co-exit from cilia. BioRxiv. https://doi.org/10.1101/ 2020.07.29.225797 19. Klebig C, Korinth D, Meraldi P (2009) Bub1 regulates chromosome segregation in a kinetochore-independent manner. J Cell Biol 185:841–858 20. Mukhopadhyay S, Wen X, Chih B et al (2010) TULP3 bridges the IFT-A complex and membrane phosphoinositides to promote trafficking of G protein-coupled receptors into primary cilia. Genes Dev 24:2180–2193

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 22 Calorimetric Heat Dissipation Measurements of Developing Zebrafish Embryos Jonathan Rodenfels and Karla M. Neugebauer Abstract Living cells, tissues and organisms are open, metabolically active systems that constantly exchange matter and energy with their environment in the form of heat. The heat exchanged is equal to the net enthalpy of all chemical reactions taking place within the system. Thus, heat dissipation can inform on the energetic costs of the constellation of cellular processes that contribute to physiology and address unanswered questions about development, responses to the environment, signaling and metabolic pathways, and the roles of morphological substructures. Here, we describe the methods we established to measure the heat dissipated by early zebrafish embryos undergoing synchronous cell cycles of cleavage stage embryogenesis, using isothermal calorimetry. The non-invasive nature of calorimetry and the versatility of these methods enables the investigation of the energetic costs of embryonic development and of the cellular processes associated with the early embryonic cell cycles. Key words Calorimetry, Heat dissipation, Energetics, Embryogenesis, Development, Cell cycle, Zebrafish, Oscillations, Metabolism

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Introduction After fertilization, most species undergo cleavage stage development, a cellular program to exponentially expand the number of cells. These embryonic cleavage cycles differ from the canonical four-phased cell cycle in somatic cells [1, 2]. They are rapid, autonomous, and lack cell cycle checkpoints and gap (G1 and G2) phases [1–6]. The large zygote divides without significant cytoplasmic growth and, thus, is progressively cleaved into smaller cells giving rise to the blastula, a cluster of cells of the same overall volume as the original zygote. During these reductive cleavage divisions, cells progress through a coordinated and tightly regulated sequence of processes—DNA replication during S-phase and mitosis and cytokinesis during M-phase—that define the embryonic cell cycle. Cleavage stage and the exponential increase in embryonic cell number requires matter and energy to fuel cell cycle progression.

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Precursors such as nucleotides, fatty acids and amino acids are needed for DNA replication, the increase in plasma membranes, and protein synthesis. Each cell of the embryo must expend energy to assemble these precursors into macromolecules and to build cellular machinery, such as chromatin and mitotic spindles. Furthermore, energy is required to generate forces needed to segregate the chromosomes, divide the cell, and change the activity of signaling pathways that enforce coordinated cell cycle progression [7– 11]. These requirements are satisfied by cellular metabolism, and work of the last decades shows that proliferating cells can use a variety of metabolic strategies to fulfill these demands. For example, the shift from oxidative metabolism to aerobic glycolysis (Warburg metabolism) promotes growth in many cancers and has also been implicated in normal tissue growth [12–14]. Interestingly, the metabolic requirements of the embryonic cleavage stage of embryogenesis (e.g., preimplantation mammalian development) are met by a different metabolic strategy, namely oxidative phosphorylation [15–17]. These studies have highlighted the principle(s) of how metabolism allows cells to proliferate at the level of biochemical networks within cells. However, it remains unclear how metabolic energy is partitioned among cellular processes driving the cell cycle. The embryo can be considered as an open system exchanging energy and matter with its environment. Thus, its metabolic and energetic profile can be understood from first principles, such as mass and energy conservation, stoichiometric constraints on production, homeostasis and morphometric parameters [18]. To investigate overall energetics, we and others have measured the energy exchanged between embryos undergoing development and their environment in the form of heat [19–22]. This heat dissipation is equal to the net enthalpy change associated with all biochemical reactions taking place in the embryo. The measurements show that embryogenesis is exothermic, meaning that the medium surrounding the embryos heats up. Recent work showed that the heat dissipation oscillates in early zebrafish and frog embryos undergoing early cleavage divisions, with an amplitude of ~2% of the total heat dissipation [19, 22]. We have shown that the oscillations have a period equal to the cell cycle time and arise from heat dissipated by the biochemical reactions associated with the cell cycle oscillator, which coordinates events that take place during the cell division cycles [19]. This analysis, based on non-invasive isothermal calorimetry measurements, shows the value of an approach that associates cellular events during development with energetics. In this chapter, we describe how to measure heat dissipation by early zebrafish embryos undergoing synchronous cell cycles using isothermal calorimetry (see Fig. 1 for workflow). The zebrafish system offers several experimental advantages. As noted earlier, the embryo volume remains approximately constant during

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Fig. 1 Workflow. The calorimeter is cleaned and the experimental and reference cells are prepared. The experiment is started, the system equilibrates and the initial baseline is recorded and inspected for drift, noise and spikes. Individual pairs of fish are bred and laid embryos are collected and staged at the 2-cell stage. Thirty 2-cell stage embryos are injected into the experimental cell of the calorimeter which results in an endothermic injection peak after which the heat dissipation by developing embryos is measured. Finally, the raw data is saved, curated and analyzed

cleavage stage, allowing one to study proliferative, developmental and metabolic processes independent of the overall growth of the embryo. Second, pharmacological perturbations can be made by adding agents to the embryo media. Third, the large size of the eggs, which undergo ten reductive cleavage divisions, allow for sufficient material to measure comparatively small amounts of heat flow. And finally, the high temporal precision of the cell divisions allows for the synchronization of many cells so that heat flows can be analyzed with respect to the phases of the cell cycle. The protocol steps downstream of fish breeding include calorimeter set up, embryo staging, embryo calorimeter injection, heat dissipation measurements, troubleshooting, and raw data retrieval and curation. This protocol uses a Malvern VP-ITC calorimeter but can be adapted for heat dissipation measurements using other calorimetry platforms. Moreover, the non-invasive nature of calorimetry and the versatility of these methods enables the investigation of heat dissipation by many other aquatic model systems used to study cell cycle associated processes.

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Materials

2.1

Fish

2.2

Solutions

Danio rerio (D. rerio): Wild-type AB (Zebrafish International Resource Center (ZIRC), Cat#:ZL1). 1. 60 E3 Medium: To prepare a 60 stock, dissolve 24.8 g NaCl, 1.6 g KCl, 5.8 g CaCl2 ∙2H2O and 9.78 g MgCl2∙6H2O in H2O, to a final volume of 2 L. Adjust the pH to 7.2 with NaOH. Autoclave. Store at 4  C. 2. 1 E3 Medium: 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2 0.33 mM MgSO4, 10 5% methylene blue. To prepare 1 medium: dilute 16.5 mL of the 60 E3 medium to 1 L. Add 100 μL of 1% methylene blue. 3. Calorimeter cleaning solution: 5% (v/v) Contrad (Decon-90) in pure H2O. To prepare 1 L, dissolve 50 mL of Contrad in 950 mL of H2O.

2.3 Calorimeter and Accessories

1. Malvern VP-ITC. 2. Hamilton syringe: 3 mL, for calorimeter cell rising and filling. 3. ThermoVac: facilitates degassing samples and cleaning of the cells. 4. ThermoVac cleaning accessories for vacuum-assisted cleaning.

2.4

Software

1. Generic text editor. 2. R (R Development Team, 2008). 3. R-Studio.

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Methods

3.1 Set-Up and Preparation

In this protocol, heat dissipation measurements are carried out using a Malvern MicroCal VP isothermal titration (ITC) calorimeter. Maintenance of the system should be performed routinely according to the manufacturer’s instructions. This includes cleaning of the calorimeter cells, blank baseline recordings, and blank injections (see Note 1). For embryonic heat dissipation measurements, the calorimeters titration syringe and stirrer are omitted and the opening to the reference and the experimental cell is covered by a plastic lid. This protocol can be set up on Day 1 for experiments on Day 2, extending through multiple days thereafter. In other words, you can set up for a whole week of experiments (see below).

3.2 Workflow on a Day-to-Day Basis

Day 1: 1. Rinse and prepare a fresh reference cell. 2. Set up individual pairs of fish for breeding.

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Day 2: 1. Clean the calorimeter experimental cell. 2. Prepare the desired experimental cell. 3. Start calorimeter equilibration followed by a minimum of 30–45 min initial baseline recording. 4. Collect eggs and stage early embryos. 5. Injection of embryos and heat dissipation measurement. 3.3

Egg Collection

1. The day before the experiment, set up individual pairs of fish in breeding tanks and keep them separated overnight (see step 3, Fig. 1). 2. The morning of the experiment. Breed fish and note the time when the fish start laying eggs. 3. Collect eggs from a 10 min time window of egg-laying in fresh 1 E3 medium. 4. Allow the fertilized eggs to develop for ~30 min at 28.5  C. 5. Collect 30 eggs with an apparent first cleavage furrow in fresh 1 E3 medium. 6. Proceed with heat dissipation measurements outlined in Subheading 3.7.

3.4 Calorimeter Cleaning

For an overview of the workflow see Fig. 1. 1. Rinse the reference cell 5–10 with de-ionized H2O using a long Hamilton syringe (see Note 2). 2. Rinse experimental cells with 100–200 mL calorimeter cleaning solution using a ThermoVac and the provided cleaning device. 3. Rinse the experimental cell with 400–500 mL of H2O. 4. Draw up and discard residual H2O in the experimental cell using a long Hamilton syringe.

3.5 Set Up of the Reference and Experimental Cell

1. Degas 3 mL of de-ionized H2O for 5 min. 2. Draw 2 mL of degassed H2O into a long Hamilton syringe. Take care that no air bubbles are introduced while drawing. Snap or flick the syringe with your fingernails to float potential air bubble to the top. 3. Insert the syringe into the reference cell and fill the reference cell until it slightly overflows in 3–4 dispersion steps while rotating the syringe to avoid the formation of air bubbles. 4. Draw up excess water from the reference cell. 5. Repeat steps 2–4 for the experimental cell using 1 E3 medium (see Note 3).

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3.6 Software Settings

In this protocol, heat dissipation measurements are carried out using a Malvern MicroCal VP isothermal titration (ITC) calorimeter. The systems software is intended to control classical isothermal titration experiments and was not designed for longer heat dissipation measurements. The following software settings have been empirically determined to facilitate embryonic measurements by performing short virtual injections (i.e., nothing is really being injected) with long measurement time windows in between injections; this is what is happening in step 4 below. The total number of “injections” determines the amount of time you are measuring heat dissipation. 1. Launch the VP-Viewer software and the Main Window of the VP Viewer is displayed (see manufacturer’s instructions for a detailed description of each parameter). 2. Use the following settings in VP-Viewer software’s tabs (see manufacturer’s manual for details): 3. Thermostat/Calib. tab: Thermostat Controls: Set point ( C)—set to desired experimental temperature minus 2 C. 4. ITC control tab: Experimental parameters: Total # of injections (virtual, see above): 3—for cleavage stage development and initial baseline recordings—change for longer/shorter heat dissipation measurements. See Injection parameters (below) for details. Temperature: desired temperature of your experiment. Set to 28.5 C for zebrafish development. The reference power (μcal/s): 11.5. Initial delay (sec): 240. Syringe concentration (mM): 0. Cell concentration (mM): 0. Stirring speed: 0. File name: yourexperiment.itc. 5. Feedback Mode/Gain: High. 6. ITC equilibration option: Fast equilibration & auto. 7. Injection parameters: The ITC experiments are performed without the injection syringe and stirring and the calorimeter cells and injection ports were covered with a plastic lid. Due to operational software requirements, the injection syringe parameters are set as follows (see Note 4): Injection volume (μL); 2 μL. Duration (s): 2 s.

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Spacing (s): 14,400 s (240 min): The total measurement time is determined by the number of injections (see Subheading 3.6, step 4) X Spacing (14,400 s). Filter period (s): 2 s. 8. Edit mode: All the same. This is a repeated loop as defined by the injection settings (above step 7). 3.7 Calorimeter Equilibration, Initial Baseline Recording, Embryo Injection and Heat Dissipation Measurements

For each newly prepared reference cell, perform blank baseline recording for at least 1 h followed by a blank media injection (see step 2 in Figs. 1 and 2b). A newly prepared reference is usually good for about a week (see Notes 1 and 2). 1. Start the experiment, equilibrate.

the

system

will

automatically

2. After equilibration, the initial baseline around the set reference power (see Subheading 3.6, step 4) will be recorded (Fig. 2). Record baseline for ~45–60 min (Fig. 1, step 2). This time can be used to start breeding and collect staged 2-stage embryos (Fig. 1, step 3). 3. Assess the system’s stability by inspecting the initially recorded baseline for drift, noise, and spikes. If the baseline is stable proceed with embryo or blank media injection (Fig. 1, step 4). Should baseline issues arise see the troubleshooting section for further instructions (Subheading 3.10). 4. Cut a 200 μL pipette tip to increase the diameter of the tip to fit individual embryos. 5. Draw up 10–15 individual embryos in 150–200 μL using the cut pipet tip and 200 μL pipet. 6. Pipet the embryos on top of the small opening of the experimental cell. Make sure to insert the tip opening to the neck of the opening. The embryos naturally sink down the neck of the experimental cell. 7. Repeat steps 5–7 until the desired number of embryos are injected. 8. Use the long Hamilton syringe to push embryos stuck in the neck down in the experimental cell by inserting the syringe into the neck and carefully push down about 1/3 of the way to the bottom of the cell. 9. Draw up access media using the Hamilton syringe. 10. The injection will result in an endothermic injection peak (Fig. 2b, c) lasting about 2–3 min. For blank media injections and metabolically inactive material such as dead embryos, the baseline should return to the initial readings (Fig. 2b). For metabolically active embryos, the baseline will stabilize around a new more negative reading (Fig. 2c). This indicates an

Fig. 2 Schematic of an isothermal calorimetry experiment. (a) The calorimeter has two cells, one of which contains water and acts as a reference cell, the other contains the sample and sample buffer. The calorimeter keeps these two cells at exactly the same temperature. When a reaction occurs within the experimental cell, the temperature difference between the cells is sensed. The calorimeter gives feedback to the heaters in the form of differential power, which is needed to compensate for the temperature difference and return the cells to baseline. Calorimeter equilibration of H2O in the reference cell against 1 E3 medium alone in the experimental cell. Thirty zebrafish embryos are staged at the 2-cell stage and injected into the experimental cell which causes a reproducible endothermic injection peak prior to the heat dissipation measurement by developing embryos. (b) Raw heat dissipation data trace of pre-injection baseline recording, 1 E3 media injection and measurement. (c) Raw heat dissipation data trace of pre-injection baseline recording, 30  2cell stage embryo injection and measurement

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exothermic reaction is taking place within the calorimeter chamber and the embryos dissipate heat. 11. Cover the cell openings with a lid. 12. Record heat dissipation for the desired time. For cleavage stage development, record heat dissipation measurements for 3–4 h (Fig. 1, step 5). 3.8 Raw Data Retrieval and Initial Data Curation

1. The raw data files are saved as *.itc by the VP Viewer software. 2. Use any text editor to open *itc file. 3. Remove the metadata from the *.itc file, retaining the columns, time, differential power (DP, in μcal/s) and temperature in the dataset. 4. Search for the “@” sign containing rows. “@” indicates the time of a virtual injection. Delete each “@” containing row. 5. Save the file as *.csv.

3.9

Data Analysis

3.10 Troubleshooting

Custom R scripts for data analysis of heat dissipation oscillations concurrent with the cleavage stage cell cycle as well as example files can be found at https://github.com/rodenfels/zfish-calorimetry. For an overview of the data processing steps (Fig. 1, step 6) see Fig. 3. Possible issues (not a comprehensive list): 1. Baseline/measurement contains spikes/abrupt changes in heat dissipation: These spikes can be caused by the burst of air bubbles, which were likely introduced during reference/experimental cell preparation. Discard run, clean, prepare new reference/experimental cell and restart. 2. Baseline noise is larger than usual. Discard run, clean calorimeter, prepare new reference/experimental cell and restart.

download *itc file

convert to *.csv structure data as time, differential power (µcal s-1)

substract baseline convert differential power (µcal s-1) to nJ s-1 per embryo

discard injection data & normalize time

Fig. 3 Heat dissipation data analysis workflow. *.itc file is downloaded, then a generic text editor is used to remove the metadata and virtual injection time points (marked by @) from VP-ITC *.itc file and to converted it into *.csv leaving the raw data structured as time, differential power (μcal/s). Use your data analysis software of choice to subtract the initial baseline and convert the differential power to nJ/s per embryo. Delete data associated with the embryo injection and time-normalize heat dissipation data to post-injection measurements

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3. Strong baseline drift: Undesired chemical reactions may occur within the calorimeter. Discard run, clean experimental/reference cell and restart. 4. Reproducible baseline patterns such as oscillations occur. These can be caused by the room’s air condition/temperature control. Move the calorimeter to a different location. Cover the calorimeter, especially the lid with a styrofoam box. Do not cover the calorimeter exhaust fan. Adjust room temperature to 5 C of the desired experimental temperature. Turn off room temperature controls.

4

Notes 1. A blank injection of media and a blank baseline recording should be performed for each newly prepared reference cell. This will document baseline stability and noise, which can be influenced by environmental conditions. See troubleshooting Subheading 3.10 for details. 2. A newly prepared reference is usually good for about a week. Under normal circumstances, the reference cell does not require intensive cleaning. Should baseline stability or noise issues arise after troubleshooting steps for the experimental cell have been taken, intensive cleaning of the reference cell should be performed according to manufacturer’s instructions. 3. The experimental cell buffer (usually 1 E3 medium) can be modified to contain desired concentrations of drugs for chemical perturbation experiments. Control heat dissipation measurements of embryos developing under mock-treated conditions (e.g., 1 E3 medium containing solvents such as DMSO or Ethanol) need to be performed. 4. Make sure the injection syringe is drawn up and has a “virtual” volume in order for the software to execute “virtual” injections. This is achieved by using the Pipet control settings in the ITC control tab.

Acknowledgements We thank Dr. Jonathon Howard and the participants of the 2015 Physiology course at the Marine Biology Laboratory in Woods Hole, MA for helpful discussion during the inception of this project. This work was supported by funding from an EMBO Longterm Fellowship ALTF 754-2015 (to J.R.) and the NIH R21 HD094013 (to K.M.N.). Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the NIH.

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References 1. Kane DA, Kimmel CB (1993) The zebrafish midblastula transition. Development 119:447–456 2. Newport J, Kirschner M (1982) A major developmental transition in early Xenopus embryos: I. Characterization and timing of cellular changes at the midblastula stage. Cell 30:675–686 3. Ikegami R, Rivera-Bennetts AK, Brooker DL et al (1997) Effect of inhibitors of DNA replication on early zebrafish embryos: evidence for coordinate activation of multiple intrinsic cellcycle checkpoints at the mid-blastula transition. Zygote 5:153–175 4. Kimelman D, Kirschner M, Scherson T (1987) The events of the midblastula transition in Xenopus are regulated by changes in the cell cycle. Cell 48:399–407 5. Newport JW, Kirschner MW (1984) Regulation of the cell cycle during early Xenopus development. Cell 37:731–742 6. Paranjpe SS, Veenstra GJC (2015) Establishing pluripotency in early development. Biochim Biophys Acta 1849:626–636 7. Loog M, Morgan DO (2005) Cyclin specificity in the phosphorylation of cyclin-dependent kinase substrates. Nature 434:104–108 8. Novak B, Tyson JJ (1993) Numerical analysis of a comprehensive model of M-phase control in Xenopus oocyte extracts and intact embryos. J Cell Sci 106(Pt 4):1153–1168 9. Murray AW, Hunt T (1993) The cell cycle: an introduction. Oxford University Press, Oxford 10. Tsai TY-C, Theriot JA, Ferrell JE Jr (2014) Changes in oscillatory dynamics in the cell cycle of early Xenopus laevis embryos. PLoS Biol 12:e1001788 11. Kamenz J, Ferrell JE Jr (2017) The temporal ordering of cell-cycle phosphorylation. Mol Cell 65:371–373

12. Miyazawa H, Aulehla A (2018) Revisiting the role of metabolism during development. Development 145:dev131110–dev131111 13. Ward PS, Thompson CB (2012) Metabolic reprogramming: a cancer hallmark even warburg did not anticipate. Cancer Cell 21:297–308 14. Agathocleous M, Harris WA (2013) Metabolism in physiological cell proliferation and differentiation. Trends Cell Biol 23:484–492 15. Gardner DK, Leese HJ (1990) Concentrations of nutrients in mouse oviduct fluid and their effects on embryo development and metabolism in vitro. J Reprod Fertil 88:361–368 16. Gardner DK (1998) Changes in requirements and utilization of nutrients during mammalian preimplantation embryo development and their significance in embryo culture. Theriogenology 49:83–102 17. Houghton FD, Thompson JG, Kennedy CJ et al (1996) Oxygen consumption and energy metabolism of the early mouse embryo. Mol Reprod Dev 44:476–485 18. Jusup M, Sousa T, Domingos T et al (2017) Physics of metabolic organization. Phys Life Rev 20:1–39 19. Rodenfels J, Neugebauer KM, Howard J (2019) Heat oscillations driven by the embryonic cell cycle reveal the energetic costs of signaling. Dev Cell 48:646–658.e6 20. Rodenfels J, Sartori P, Golfier S et al (2020) Contribution of increasing plasma membrane to the energetic cost of early zebrafish embryogenesis. Mol Biol Cell 31:520–526 21. Song Y, Park JO, Tanner L et al (2019) Energy budget of Drosophila embryogenesis. Curr Biol 29:R566–R567 22. Nagano Y, Ode KL (2014) Temperatureindependent energy expenditure in early development of the African clawed frog Xenopus laevis. Phys Biol 11:046008

Chapter 23 The Conditional Knockout Analogous System: CRISPR-Mediated Knockout Together with Inducible Degron and Transcription-Controlled Expression Hoi Tang Ma Abstract The revolutionary CRISPR technology opens a new era of cell biology in mammalian cells. The InDel mutation is induced by CRISPR and results in the frameshift mutation of the gene. Owing to the nature of CRISPR induced knockout, the conditional knockout using CRISPR technology is not common. With the recent development of the small molecule-inducible degron system, an analogous system to the classical genetic conditional knockout has become feasible. By integrating CRISPR-knockout, the tetracyclinecontrolled transcriptional and auxin-induced degradation post-translational control of protein expression, a method imitating the conditional knockout is developed. We herein describe the detailed protocol for the generation of a conditional protein inactivation in human cancer cells. The system is especially useful to study essential gene function in aneuploidy cancer cells where gain in copy number is common. Key words Auxin induced degron, Conditional knockout, CRISPR

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Introduction Direct gene disruption represents the most effective approach to study gene function by investigating the loss-of-function phenotype. With the revolutionary CRISPR technology, it can be achieved significantly easier than using classical genetic methods. As many of the cell cycle oscillator genes are essential, the lethality phenotype abrogates the generation of cells carrying the mutated gene. The inducible degron system allows the rapid proteolysis of proteins of interest, and is analogous to the conditional knockout genetic system, when the endogenous protein of interest is targeted. Inducible degron systems are composed of at least three components: inducer, degron and degradation system. The inducer is usually a small cell permeable ligand molecule (such as the plant hormone auxin, and thalidomide-like molecules), which facilitates the destruction of the degron via the degradation machinery.

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Notably, the inducer is not necessarily a chemical; a change in temperature, such as heat stock, was one of the earliest inducers for controlling the conditional allele to be degraded [1]. Other forms of physical energy such as light [2], can be used as an inducer in controlling protein degradation. The degron refers to the transferable protein motif (such as Auxin Induced Degron, AID [3]), crafted from the native substrate of the degradation system. It is physically linked to the protein of interest via an N-terminal or C-terminal fusion. To avoid unnecessary interference with the protein of interest’s function, the size of the degron should be minimized. The smallest degron reported is 25 amino acids, derived from IKZF3 in the IMiD-induced degradation system [4]. In general, the degradation system is made out of a ubiquitin ligase and proteasome. The ubiquitin ligase can be endogenously expressed (such as CRBN/CRL4 in the IMiD-sysem) or chimeric exogenous-endogenously expressed (such as the exogenously expressed TIR1 with endogenously expressed SKP1-CUL1). The ubiquitinated protein of interest is then targeted to degradation by the endogenous proteasome. One exception of the system has been reported by Matouschek [5]; they utilize a rapalog as the inducer, to trigger the dimerization of FKBP (degron) and chimeric FRB-proteasome adaptor Rad23B, which directly targets to proteasome-mediated degradation without ubiquitination [5]. The features of different inducible degradation systems are summarized in Table 1 and the details of the AID system will be discussed. The AID system was first described as a conditional protein depletion system in S. pombe [3]. The system has been adopted in mammalian cells to provide temporal and quantitative control of protein expression [6]. In plants, the phytohormone auxin (indole-3-acetic acid; IAA) induces degradation of the AID domain containing transcriptional repressors by targeting to the substrate recognition domain of an F-box protein called TIR1 [7]. TIR1 forms an SCF-type ubiquitin ligase (E3) with cullin, RBX1, and SKP1, and promotes the ubiquitination and subsequent proteasomal degradation of the AID-containing protein. In mammalian cells, ectopically expressed TIR1 can form a functional E3 ligase with the remaining SCF components from endogenous proteins and degrade AID-tagged proteins [3]. In order to deplete the protein of interest in mammalian cells, the TIR1 need to be ectopically expressed while both endogenous loci need to be targettagged with the AID-domain. Endogenous AID-tagging is facilitated by CRISPR mediated recombination and has been successfully implemented to the nearly diploid cell lines such as HCT116 [8]. However, the AID-tagging of all endogenous loci in aneuploidy cells, which are quite commonly used in cell cycle studies, can be challenging. In fact, the most widely used cell line, HeLa, contains hypertriploid genome with 3n + set of chromosomes and

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Table 1 Summary of different inducible degradation systems

Inducer

Feature

Original organism References

Degron

Adaptor

AID, mAID

Plant based degron Transport inhibitor Auxin response 1 (TIR1) (indole-3acetic acid)

O. sativa

[3]

sJAZ

Chimeric receptor JA-Ile Plant based degron (TIR1 (jasmonateF-box-COI1BLRR) isoleucine)

O. sativa

[17]

A. sativa

[2]

cODCAsLOV2)

Blue light

Ubiqutination independent

minimal Cereblon IMiDresponsive IKZF3 degron

IMiD (pomali Degron only work with domide) C-terminal tag, no adaptor expression is required

FKBP Proteasome adaptor (T2098L) (Rad23Bb UbL-mCherryFRB-MBP)

Rapalog

H. sapiens [4]

Direct proteasome H. sapiens [5] mediated degradation

many loci are with three or more copies [9], indicating that multiple rounds of AID-tagging with different antibiotic selection markers may be required. Moreover, another limitation of the AID system is that rapid degradation of AID-fusion proteins requires relatively high levels of TIR1, which may not be attainable in many mammalian cell lines. In fact, residue levels of AID-fusion proteins could often be detected even after prolonged incubation with IAA. Strategies involving different AID and F-box adaptors from different plant species as well as employing different auxin derivatives as inducers do improve the residual level issue [10, 11], however, we believe that suppressing AID-expression at the transcriptional level together with inducible degron control can be a simpler solution [12]. A widely used system for downregulating gene expression in mammalian cells involves placing the gene of interest under tetracycline-controlled promoters (Tet-Off), which are based on the Tet repressor protein (TetR) and tet operator (tetO) DNA elements that control the Tn10-encoded tetracycline resistance operon in Escherichia coli [13]. TetR is fused to the transcription activation domain of VP16 from herpes simplex viruses, resulting in a tetracycline-controlled transcriptional activator (tTA). Tetracycline response element (TRE) is constructed by fusing seven tetO sequences to a minimal TATA-box containing a eukaryotic promoter derived from the CMV immediate early gene. In the absence

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of tetracycline, tTA dimers bind TRE and activate the expression of the downstream transgene. Binding of tetracycline or derivatives such as doxycycline (Dox) induces a conformational change in the TetR of tTA, inducing the dissociation from tetO, thus turning off gene expression. By putting the Tet-Off, AID system and CRISPR mediated knockout together, we are able to generate conditional gene inactivation cell lines, which are analogous to the conditional knockout. Conceptually, the AID-tagged protein of interest under the control of a Tet-Off promoter and the TIR1 protein under the constitutively active promoter are stably expressed in cells; the loss of function mutation, usually frameshift, at the endogenous locus is introduced by CRISPR. In the normal culture condition, the AID-tagged protein of interest is stable and complements the mutated endogenous gene. With the addition of Dox and IAA, the AID-tagged protein will rapidly be degraded via TIR1dependent ubiquitination and subsequently degraded by the proteasome (see Fig. 1). In this protocol, we will outline the generation of the “turn-off” cell line step by step using cyclin A2 as an example. Notably, antibodies capable of recognizing endogenous protein expression and CRISPR capable of effectively targeting the endogenous protein expression are the most critical factors. To facilitate the integration of exogenous genetic material in to the cell line genome, we utilized the recombinant retrovirus system, however, alternative methods such as the Flp-In™ system can be modified accordingly to fit the same purpose.

2

Materials

2.1 Stock Solutions and Reagents

1. Cell lysis buffer: 50 mM Tris–HCl, pH 7.5, 250 mM NaCl, 5 mM EDTA, and 50 mM NaF, and 0.2% NP40. Add fresh: 1 mM PMSF, 1 μg/ml leupeptin, 2 μg/ml aprotinin, 10 μg/ml soybean trypsin inhibitor, 15 μg/ml benzamidine, 10 μg/ml chymostatin, and 10 μg/ml pepstatin. 2. PBS (phosphate-buffered saline): 137 mM NaCl, 2.7 mM KCl, 10 mM sodium phosphate dibasic, and 2 mM potassium phosphate monobasic, pH 7.4. 3. SDS sample buffer: 10% w/v SDS, 1 M Tris–HCl, pH 6.8, 50% v/v glycerol, and bromophenol blue (as required). Add 50 μl/ ml 2-mercaptoethanol before use. 4. 5 μg/ml Blasticidin. 5. 2 μg/ml Doxycycline hydrochloride (Dox). 6. 200 μg/ml Hygromycin B. 7. 50 μg/ml Indole-3-acetic acid (IAA). 8. 0.6 μg/ml Puromycin.

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Fig. 1 Combining the tetracycline-controlled transcriptional system, the auxin-inducible degron (AID) system, and CRISPR-mediated knockout. The transcription of AID-tagged genes (X) is placed under the control of a Tet-Off promoter. In Tet-Off cell lines, the tetracycline-controlled transcriptional activator (tTA) binds to the tetracycline response element in the promoter and activates the transcription of AID-tagged gene in the absence of doxycycline (Dox); the endogenous loci of X are targeted by CRISPR to introduce the loss of function frameshift mutation. Addition of Dox turns off the transcription of the promoter. In response to IAA, AID-fusion protein is rapidly targeted for degradation in cells expressing the ubiquitin ligase SCFTIR1

9. 10 mg/ml Polybrene. 10. Plasmid DNA: pREVTRE-AID (Addgene #121067), pUMVC (Addgene # 8449), pCMV-VSV-G (Addgene # 8454), TIR1-myc retrovirus constructs [14], Histone H2B-GFP/Bsd construct was a gift from Tim Hunt (Cancer Research UK, UK). 11. Annealing buffer (10
): 100 mM, Tris pH 7.5 to 8, 500 mM sodium chloride, and 50 mM EDTA. 2.2

Cell Culture

All solutions and equipment that will contact with cells must be sterile and proper sterile technique should be used accordingly: 1. A Tet-off clone, of HeLa cells (American Type Culture Collection, Manassas, VA, USA). Cells are grown in a humidified incubator at 37  C in 5% CO2 (see Note 1). 2. Growth medium: Dulbecco’s Modified Eagle Medium (DMEM) containing 10% heat-inactivated calf serum and 30 U/ml penicillin–streptomycin. 3. Trypsin: 0.25% with EDTA. 4. Tissue culture plates and standard tissue culture consumables.

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Equipment

1. Standard tissue culture facility. 2. Centrifuge that can accommodate 15 and 50 ml centrifuge tubes. 3. Microcentrifuge that can reach 16,000
g at 4  C. 4. Sterile disposable syringe with 0.45 PVDF membrane filter unit. 5. Hemocytometer.

3

Methods

3.1 CRISPR and the Conditional-Off Expression Construct

The CRISPR target can be designed with various online resources (see Note 2). In general, two types of CRISPR target can be used to introduce the frameshift mutation to disrupt the gene function: one targeting the ORF region of cDNA and another one targeting the exon–UTR/intron region of cDNA (see Note 3). In our showcase example, we have designed two CRISPR sequences targeting the UTR-exon and the exon–intron region of cyclin A2 gene (Fig. 2a). One advantage of this kind of CRISPR is that they will not target the exogenous cDNA expression DNA, which will be critical in complementation of the essential function when the endogenous gene is disrupted. However, this type of CRISPR is relatively limited as it is restricted by the sequence of the limited number of UTR–exon and exon–intron regions. Alternatively, a silence mutation in the exogenous cDNA expression DNA will be introduced if the CRISPR targeting the ORF region of cDNA is used (see Note 4). 1. Synthesize the following oligonucleotides through a commercial service (see Note 5): CCNA2_CRISPR1_sense:

50 -caccgCCCTTTACCCGTCTCGTCTT30

CCNA2_CRISPR1_antisense: 50 -aaacAAGACGAGACGGGTAAAGGGC -30 CCNA2_CRISPR2_sense:

50 -caccgCTCCCGGGAGCAGTGATGTT30

CCNA2_CRISPR2_antisense: 50 -aaacAACATCACTGCTCCCGGGAGC -30

2. Set up the annealing reaction in a microfuge tube as follows: 10 μl sense oligo (100 μM). 10 μl antisense oligo (100 μM). 5 μl of 10
annealing buffer. 25 μl of MQH2O.

The Conditional Knockout Analogous System: CRISPR-Mediated Knockout. . . ATG

A

STOP

cyclin A2

CRISPR1

cy cli n

AI D

- 1 2

1+ 2

cy cli n

He La

C

AI D

He La

A2 -T m ir1 yc

B

CRISPR2

A2 -T ir1

ccatttcaatagtcgcgggatacttgaactgcaagaacagccgccgctccggcgggctgctcgctgcatctctgggcgtcttt ggctcgccacgctgggcagtgcctgcctgcgcctttcgcaacctcctcggccctgcgtggtctcgagctgggtgagcgagcgg gcgggctggtaggctggcctgggctgcgaccggcggctacgactattctttggccgggtcggtgcgagtggtcggctgggcag agtgcacgctgcttggcgccgcaggctgatcccgccgtccactcccgggagcagtgATGTTGGGCAACTCTGCGCCGGGGCCT GCGACCCGCGAGGCGGGCTCGGCGCTGCTAGCATTGCAGCAGACGGCGCTCCAAGAGGACCAGGAGAATATCAACCCGGAAAA GGCAGCGCCCGTCCAACAACCGCGGACCCGGGCCGCGCTGGCGGTACTGAAGTCCGGGAACCCGCGGGGTCTAGCGCAGCAGC AGAGGCCGAAGACGAGACGGgtaaagggatgcgggatatctgcaggagggtgggtcgagcagggtttggcattggcttagcag gcagagaagagtggcgagaaggcatgtcctgggcttttggaagggggccagggtctggagtgcagttggtgctggtgtgtggt ttgccacagtaggagttctcccatattagcatcaggaccccgccaacactgtgtgtatgctgccggtctgctctctatagggg gcatctgcctgtacataatggggtcaaaccaagctctaaaacgtgtagtcttcgctcagctccgcgcctttctctccactttt aaaccccaggcactgctgtaggactctgacccctatcctcctcacgcttaagagatgacctctactttagaaaagcgtgtaca aaatactttgcttttggcaaattccgccattttagccggatttgctctgttgctccaccctcgggcgacagtggtgaaccaac aatttttttgctgtctttcctaaacttgtcacgtattggcctgtcacccgacccttcttgtggccctgataagttttgcataa ttccacctagtgttatctattgatagcctttgtgggaatgcctgtgacaaatgggaacatcccttctcttttgaatactgaaa ctcttctttgtcccagaaagtttaattcctgatagagtatttgggagaaaaagcaaaggccaacacccataagagaaagaatg caagactagtaggctcaaagccagttattaatttttttttagGTTGCACCCCTTAAGGATCTTCCTGTAAATGATGAGCATGT CACCGTTCCTCCTTGGAAAGCAAACAGTAAACAGCCTGCGTTCACCATTCATGTGGATGAAGCAGAAAAAGAAGCTCAGAAGA AGCCAGCTGAATCTCAAAAAATAGAGCGTGAAGATGCCCTGGCTTTTAATTCAGCCATTAGTTTACCTGGACCCAGAAAACCA TTGGTCCCTCTTGATTATCCAATGGATGGTAGTTTTG

DOX IAA:

329

− −

+

cyclin A2 actin

myc actin Fig. 2 Inducible degron, transcription-controlled expression of AID-cyclin A2. (a) A schematic diagram of the CCNA2 gene. The start codon “ATG” and stop codon are indicated. The two CRISPR targets in the UTR-ATG boundary and the exon–intron boundary are highlighted by the dotted underline and the Cas9 cut sites are indicated by the black triangle (~). (b) The HeLa Tet-Off cells infected with AID-cyclin A2 and TIR1-myc expression retrovirus were generated as described in Subheading 3.2, step 7. The cells were treated with IAA and Dox for 16 h. The expression of AID-cyclin A2, endogenous cyclin A2, TIR1-myc, and actin were detected with immunoblotting. (c) The mixed-population culture of the AID-cyclin A2 and TIR1-myc expressing HeLa Tet-Off cells transfected with CRISPR targeted CCNA2 were generated as described in Subheading 3.2, step 8. The expression of AID-cyclin A2, endogenous cyclin A2 and actin were detected. The endogenous- but not the AID- cyclin A2 expression were suppressed by both CRISPRs

3. Put the annealing reaction in hot boiling water (at least 300 ml) within a floating rack. Let the boiling water cool down gradually to room temperature. 4. Set up the ligation of the 1:1000 diluted annealedoligonucleotides to the BbsI digested gel-extracted

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pX330 to generate CCNA2_CRISPR2.

the

CCNA2_CRISPR1

and

5. The ORF of the gene of interest is subcloned downstream to the AID sequence to generate the AID-ORF fusion expression construct. In our showcase example, the cyclin A2 ORF is subcloned into the pREVTRE-AID (see Subheading 2.1). The pREVTRE-AID is the retrovirus vector carrying the Hygromycin-B-phosphotransferase for hygromycin selection upon infection. The AID-fusion expression is driven by the TRE promoter, which can be suppressed by Dox treatment. 6. Amplify the cDNA of cyclin A2 with the following oligonucleotides using cyclin A2 cDNA as the template: cyclin A2_NcoI_FOR:

30 -GCAGCCATGGTGGGCAACTCT-30

cyclin A2_EcoRI_REV:

30 -TGAATTCTTACAGATTTAGTGTCTCTGG-30

7. Gel purify the PCR product and digest it with restriction enzymes (NcoI and EcoRI). 8. Set up the ligation of the gel-purified digested PCR product to the NcoI-EcoRI digested pREVTRE-AID to generate the pREVTRE-AID (cyclin A2). 3.2 Generation of the Conditional Inactivation Cell Line

The retroviruses carrying the constitutive expression of TIR1 and the Dox-controlled expression of AID-fused gene of interest are prepared as follows (see Note 6): 1. Transfect the HEK293 with the retrovirus vectors, pREVTREAID (cyclin A2) or pBabe-TIR1 together with the retrovirus packaging constructs (see Subheading 2.1) using the calcium phosphate transfection procedure [15]. 2. Replace the transfection medium with fresh medium after 12–16 h incubation. 3. Collect the virus-containing medium at 36–48 h posttransfection and filter through a 0.45 μm PDVF filter unit. Store the filtered virus-containing medium at 4  C (stable for 1–2 weeks). The AID-fused gene of interest and TIR expressing cells are generated as follows: 4. Transduce the Tet-OFF cell line (HeLa Tet-Off) by addition of the filtered viruses with polybrene at low seeding density (about 5% confluency). 5. Coinfect both viruses, leave for 6–12 h and then reinfect (repeat 3–4 times; see Note 7).

The Conditional Knockout Analogous System: CRISPR-Mediated Knockout. . .

331

6. Add the antibiotics to select the cells infected by both type of virus (the Hygromycin for the pREVTRE-AID and puromycin for the pBabe-TIR1, see Note 8). 7. Upon selection, the expression of the AID-fusion protein and TIR1 can be verified by Western blotting (see Note 9). In our showcase example, HeLa Tet-OFF transduced with the AID-cyclin A2 and TIR1-myc retroviruses were generated as described in Subheading 3.2. The cells were treated with Dox and IAA for 16 h to evaluate the suppression of the AID-cyclin A2 expression and the constitutive expression of TIR1 (Fig. 2b). (a) Prepare the cells as described in Subheading 3.2. (b) Treat the cells with Dox and IAA for 16 h and treat the control cells with DMSO. (c) Harvest the cells by trypsin treatment and centrifugation. (d) Store the cell pellets at 80  C. (e) Add2 pellet volumes of cell lysis buffer into the microfuge tube. Vortex to mix. (f) Incubate on ice for 30 min. (g) Centrifuge at 16,000
g at 4  C for 30 min. (h) Transfer the supernatant to a new tube. (i) Measure the protein concentration of the lysates. Dilute to 1 mg/ml with SDS sample buffer (see Note 10). (j) Run the samples on SDS-PAGE and analyze by immunoblotting with specific antibodies against cyclin A2, myc, and actin (see Note 11). 8. The mixed population cells expressing the AID-fusion protein of interest and the TIR1 are then transfected with the CRISPR targeting the endogenous loci to introduce the loss of function frameshift mutation. In our showcase example, the abovedescribed cells were transfected with two CRISPRs followed by transient antibiotic selection. In our example, both CRISPRs were able to reduce the expression of endogenous cyclin A2 but not the virus transduced AID-cyclin A2 (Fig. 2c). (a) Transfect the cells prepared as in Subheading 3.2 with the cyclin A2 CRISPR1 and/or CRISPR2 together with Histone H2B-GFP/Bsd construct (see Subheading 2.1) using the calcium phosphate transfection procedure [15]. (b) Replace the transfection medium with fresh medium after 12–16 h incubation. (c) Add blasticidin at 8 h after replacement of the transfection medium. (d) Replace the selection medium by fresh medium after 36 h and further recover the cells in the fresh medium for 36 h.

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(e) Analyze the protein expression as described as Subheading 3.2, step 7. 9. The single cell derived cell clones are then isolated from the mixed population cell lines generated in Subheading 3.2, step 8 by limited dilution in 96 well plates (see below). The expression of the AID-fused protein of interest, TIR1 and the knockout of endogenous expression are estimated by Western blotting. Cell clones expressing the AID-fused protein of interest, TIR1 and KO of endogenous expression will be selected. In our showcase example, we tested 11 clones and various combinations of endogenous cyclin A2, AID-cyclin A2 and TIR1-myc expression were found (Fig. 3a). (a) Count the cells density with hemocytometer. (b) Dilute the cells to 75 cells/15 ml. Add 150 μl of diluted cells to each of the wells in 96-well tissue culture plates and allow them to grow for 12–14 days (see Note 12). (c) Identify the wells containing the single cell derived cell clone by light microscopy (see Note 13). (d) Expand the single cell derived culture step-wise from 96 well plate, to 24 well plate and then 60 mm plate. (e) Analyze the proteins expression of the clones as in Subheading 3.2, step 7. 3.3 Assessment of Degron-Degradation Kinetics

The single cell derived cell clones are treated with Dox and IAA for various time points to evaluate the degradation kinetics of the AID-fusion protein of interest (see Note 14). In our showcase example, we tested two clones by treating with Dox and IAA for 0, 4, 8, 12 and 16 h. The expression of AID-cyclin A2 were greatly reduced after 4 h of DOX and IAA treatment, and become undetectable after 8 h of treatment (Fig. 3b).

3.4 Genomic DNA Sequencing of CRISPR Targeting Region

Although the Western blotting method described in Subheading 3.2, step 7 provides the best estimation of the knockout of endogenous protein expression, this can be further verified by the InDel analysis of the CRISPR targeting genomic region by DNA sequencing. 1. Harvest the cells as described in Subheading 3.2, step 7. The parent cell line is needed to serve as the control for the CRISPR targeted cell line. 2. Extract the genomic DNA by overnight proteinase K treatment followed with phenol-chloroform extraction. 3. Further purify the genomic DNA by ethanol precipitation. 4. Amplify the CRISPR targeting genomic region with the primers flanked about 300–500 bp around the CRISPR targeting

The Conditional Knockout Analogous System: CRISPR-Mediated Knockout. . .

333

A H

1

2

3

4

5

6

7

8

9 10 11

cyclin A2

9E10 actin

B cyclin A2 CCNA2KO #9 0 4 8 12 16

AID

HeLa

DOX IAA:

HeLa

AID

cyclin A2 CCNA2KO #11 0 4 8 12 16 (h)

cyclin A2

cyclin A2

actin

Fig. 3 Efficient depletion of AID-cyclin A2 using IAA and Dox. (a) The single cell derived cell clone 2 were generated as described in Subheading 3.2, step 9. The expression of AID-cyclin A2, endogenous cyclin A2, TIR1-myc, and actin were detected with immunoblotting. Equal loading of lysates was confirmed by immunoblotting for actin. (b) The clones #9 and #11 were grown in the presence of IAA and Dox, and harvested at the indicated time. The expression of cyclin A was detected using immunoblotting. Control HeLa cells were included in the analysis to indicate the position and abundance of endogenous cyclin A. The extended exposure of cyclin A Western blotting indicated the complete elimination of both endogenous- and AID-cyclin A2 after 8 h of IAA and Dox treatment

genomic region and 1–10 ng genomic DNA as template (see Note 15). 5. Gel purify the PCR and sequence the gel purified product with one of the flanking primers by Sanger sequencing methods. 6. Analyze the Sanger sequencing result using the Tracking of Indels by Decomposition (TIDE) method [16].

334

4

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Notes 1. Various Tet-off cell lines are also commercially available (Takara Bio USA, Inc). 2. https://portals.broadinstitute.org/gpp/public/analysistools/sgrna-design. 3. To introduce the frameshift mutation, the Cas9 cutting site should be located in the exon region. The CRISPR targeting the UTR–ORF and exon–intron boundary, which cut at the ORF–exon region, should be selected. 4. The silence mutation is introduced by replacing the original codon with the degenerate codons. We usually introduce at least three mismatches in the cDNA to prevent targeting by the CRISPR. 5. The standard desalting oligonucleotide is sufficient for CRISPR cloning. 6. Retroviruses are classified as a Biosafety Level 2 (BSL-2) organisms. Retroviruses require BSL-2 practices and procedures for all work with the virus. 7. The retrovirus infection efficiency can be improved by multiple rounds of virus addition. We usually add the virus for four times at 6–12 h interval. 8. The antibiotics sensitivity of the cells should be tested empirically. The general rule is that the lowest concentration of antibiotic which kills all the cells in 5 days should be used. 9. An antibody, which can detect the protein of interest at the endogenous level, is needed. Notably, the N-terminal fusion of AID may block the epitope recognized by a monoclonal antibody. This can be solved by C-terminal tagging the or by using another monoclonal antibody. 10. Many reagents are available for measuring the concentration of the lysates. We use BCA protein assay reagent from Pierce (Rockford, IL, USA), using BSA as standards. 11. Cyclin A2, TIR1-myc, and actin are readily detectable using commercially available monoclonal antibodies: cyclinA2 (AT10), myc (9E10), and actin (AC-74). 12. The 96-well plates should be kept undisturbed and in a wellhumidified incubator to prevent the evaporation of the medium. 13. The single cell derived cell clone should be grown as one colony in the well, and a microscope equipped with low power objective Len (4
) should be used. 14. The cell clone expresses the AID-fusion protein close to the endogenous expression level, with complete elimination of endogenous protein and can be rapidly degraded should be chosen.

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15. The primers for genomic DNA PCR should have a high melting temperature (~62–70  C). The primer pair should be verified by a tool such as UCSC In-Silico PCR (https://genome. ucsc.edu/cgi-bin/hgPcr). References 1. Verghese J, Abrams J, Wang Y, Morano KA (2012) Biology of the heat shock response and protein chaperones: budding yeast (Saccharomyces cerevisiae) as a model system. Microbiol Mol Biol Rev 76:115–158 2. Sun W, Zhang W, Zhang C, Mao M, Zhao Y, Chen X et al (2017) Light-induced protein degradation in human-derived cells. Biochem Biophys Res Commun 487:241–246 3. 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 6:917–922 4. Koduri V, McBrayer SK, Liberzon E, Wang AC, Briggs KJ, Cho H et al (2019) Peptidic degron for IMiD-induced degradation of heterologous proteins. Proc Natl Acad Sci U S A 116:2539–2544 5. Wilmington SR, Matouschek A (2016) An inducible system for rapid degradation of specific cellular proteins using proteasome adaptors. PLoS One 11(4):e0152679 6. 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:E3350–E3357 7. Hayashi KI, Neve J, Hirose M, Kuboki A, Shimada Y, Kepinski S et al (2012) Rational design of an auxin antagonist of the SCF TIR1 auxin receptor complex. ACS Chem Biol 7:590–598 8. 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 15:210–218

9. Adey A, Burton JN, Kitzman JO, Hiatt JB, Lewis AP, Martin BK et al (2013) The haplotype-resolved genome and epigenome of the aneuploid HeLa cancer cell line. Nature 500:207–211 10. Li S, Prasanna X, Salo VT, Vattulainen I, Ikonen E (2019) An efficient auxin-inducible degron system with low basal degradation in human cells. Nat Methods 16:866–869 11. Uchida N, Takahashi K, Iwasaki R, Yamada R, Yoshimura M, Endo TA et al (2018) Chemical hijacking of auxin signaling with an engineered auxin-TIR1 pair. Nat Chem Biol 14:299–305 12. Ng LY, Ma HT, Liu JCY, Huang X, Lee N, Poon RYC (2019) Conditional gene inactivation by combining tetracycline-mediated transcriptional repression and auxin-inducible degron-mediated degradation. Cell Cycle 18:238–248 13. Gossen M, Bujard H (1992) Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc Natl Acad Sci U S A 89:5547–5551 14. Ma HT, Poon RYC (2018) TRIP13 functions in the establishment of the spindle assembly checkpoint by replenishing O-MAD2. Cell Rep 22:1439–1450 15. Kingston RE, Chen CA, Okayama H (1999) Calcium phosphate transfection. Curr Protoc Immunol 31:10.13.1–10.13.9 16. Brinkman EK, Chen T, Amendola M, Van Steensel B (2014) Easy quantitative assessment of genome editing by sequence trace decomposition. Nucleic Acids Res 42(22):e168 17. Brosh R, Hrynyk I, Shen J, Waghray A, Zheng N, Lemischka IR (2016) A dual molecular analogue tuner for dissecting protein function in mammalian cells. Nat Commun 7:11742

Correction to: Purification of Cyclin-Dependent Kinase Fusion Complexes for In Vitro Analysis Mardo Ko˜ivom€agi

Correction to: Chapter 8 in: Amanda S. Coutts (ed), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_8 In the original version of this book, chapter 8 was published with a typographical error in chapter title. This has been updated in the revised version of this book.

The updated online version of these chapters can be found at https://doi.org/10.1007/978-1-0716-1538-6_8 Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_24, © Springer Science+Business Media, LLC, part of Springer Nature 2021

C1

Correction to: Live Imaging and Analysis of Cilia and Cell Cycle Dynamics with the Arl13bCerulean-Fucci2a Biosensor and Fucci Tools Melinda Van Kerckvoorde, Matthew J. Ford, Patricia L. Yeyati, Pleasantine Mill, and Richard L. Mort

Correction to: Chapter 21 in: Amanda S. Coutts and Louise Weston (ed), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_21 In the original version of this book, chapter 21 was published non-open access. It has now been changed to open access under a CC BY 4.0 license and the copyright holder has been updated to “The Author(s).” This book has also been updated with these changes.

The updated online version of this chapter can be found at https://doi.org/10.1007/978-1-0716-1538-6_21 Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6_25, © Springer Science+Business Media, LLC, part of Springer Nature 2021

C3

INDEX A Absolute length (units of time)..........165–169, 173, 174 Anaphase......................... 14–16, 29, 126, 128–130, 144, 266, 273, 274 Anaphase-promoting complex (APC) ................... 6, 8, 9, 11–16, 29–36, 128, 144, 145, 148, 152, 153, 156, 157, 159–163, 266 Arabidopsis.................................................................71–79 Ascomycete .................................................................... 281 Aspergillus nidulans ............................................. 277–287 Aurora A .................................................... 11, 13, 16, 216 Aurora B .................................................. 14, 15, 215, 216 Auxin induced degron (AID).............324–327, 330, 334 Azide labeling solution ........................................ 199, 200

B Biomarker ........................................................................ 40 Biopsy puncher....................................252, 255, 256, 262 Biosensor ................................................. 39–48, 291–308 Budding index ............................................. 266, 270, 271 Budding yeast mitotic arrest ........................................................... 269 pheromone-induced arrest ................... 124, 268, 269

C Calorimetry ................................................. 312, 313, 318 Cdc2..................................... 96, 124, 125, 127, 129, 140 Cdc20 ............................... 15, 16, 30, 31, 144, 145, 148, 157, 162, 163 Cdc25 ..................................................10, 11, 13, 14, 124 Cdk1 ........................... 10–16, 19–23, 25, 26, 31, 96–98, 100, 101, 103–108, 124, 145, 148, 152, 156, 157, 159, 162, 206, 216 CDKACT......................................................40–45, 47, 48 Cell confinement .................................180, 182, 185, 191 Cell culture conditional knockout .............................................. 196 EDU supplementation............................................ 169 fixation ..................................................................... 269 spheroids.................................................................. 225 stable cell line generation .............................. 293, 296 transfection .............................................................. 197 Cell cycle anaphase................................................................... 129

arrest .........................6, 14, 112, 114, 119, 124, 127, 128, 138, 192, 195, 196, 223, 224, 270, 274 budding yeast ........................................... 96, 265–275 checkpoint ...........................3, 4, 6, 14, 30, 112, 129, 145, 166, 174, 213, 223, 224, 234, 266, 271, 293, 294, 304, 305, 311 G1 ......................... 1–12, 14–16, 111, 112, 118, 119, 144, 165–169, 171, 173, 174, 177, 213, 219, 224, 238, 265, 266, 269, 270, 293, 294, 304–306, 311 G2 ................................. 1, 2, 4, 9–14, 20, 25, 30, 96, 111, 124, 165–169, 171, 173, 174, 177, 192, 219, 223–225, 227, 230, 238–240, 244, 294, 304, 305, 311 interphase............................... 14, 144, 145, 228, 238, 239, 283, 292, 293, 305 kinetics .................112, 223–235, 292, 293, 305, 306 metaphase .............................................. 128, 144, 265 mitosis.......................................................1, 2, 6, 9, 12 mitotic shake-off ................................... 213, 215, 219 protein phosphorylation ......................................... 206 radiation-induced ........................................... 223–235 restriction point................................................... 2–4, 6 S ............................1–6, 9, 12, 30, 96, 111, 112, 124, 127, 165, 169, 174, 207, 215, 219, 224, 237, 239, 240, 244, 266, 272, 293, 294, 304, 305 synchronization ........................................25, 266, 313 Cell division ............................ 1, 9, 19, 51, 96, 111, 123, 124, 143, 144, 180, 205, 237, 265, 266, 270, 271, 299, 312, 313 Cell extract............ 39–48, 137, 145, 156, 160, 162, 190 Cell-free system .................................................... 143–163 Cell growth............................................................. 40, 266 Cell lines 293 T-REx™ ........................ 145, 150, 151, 160, 161 U2OS............................................. 83–85, 88–92, 168 Cell lysis .............................. 98, 102, 133, 136, 137, 139, 147, 154, 209, 210, 215, 326, 331 Cell membrane .......................... 115, 171, 180, 187–189, 191, 192 Cell micro-patterning ................................................... 180 Cell synchronization mitotic.................................................... 112, 152, 213 mitotic shake-off ............................................ 214, 215 nocodazole .................................... 111–120, 213–215 thymidine....................................... 111–120, 213, 214

Amanda S. Coutts and Louise Weston (eds.), Cell Cycle Oscillators: Methods and Protocols, Methods in Molecular Biology, vol. 2329, https://doi.org/10.1007/978-1-0716-1538-6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

337

CELL CYCLE OSCILLATORS: METHODS

338 Index

AND

PROTOCOLS

Centriole length ....................................................................... 250 Centrosome ...........................6, 9, 11, 13, 179–192, 250, 251, 253, 292 Checkpoints DNA ...........112, 166, 174, 223, 224, 266, 271, 293 G1/S ............................................................... 112, 213 G2/M........................... 112, 223, 224, 294, 304, 305 Chromosome territory......................................... 237–245 Cilia ARL3B ..................................................................... 293 Click chemistry........................... 166, 175, 196, 199–202 CRISPR/Cas9 conditional knockout ..................................... 323–335 Cyclin A2..................................................... 11, 326, 328–333 Clb2 .................................................................. 96, 272 Clb5 .................................................................. 96, 272 Cyclin dependent kinase (CDK) CDK1 .................................................................. 10–16

D Daughter cell ...................... 1, 9, 14, 111, 123, 179, 205, 206, 270, 293, 301 Dephosphorylation ................ 11, 13, 23, 25, 26, 30, 40, 206, 207, 266 Development .................... 39, 48, 53, 67, 111, 180, 293, 311, 312, 314, 316, 319 DNA replication ............... 3, 6, 8, 9, 15, 30, 71, 72, 112, 131, 218, 237–239, 265, 266, 270, 271, 293, 311, 312 Dual-luciferase vector ..................................................... 83 Dynamic cell confiner .......................................... 185, 191

E EDU-Coupled Fluorescence Intensity (E-CFI) ........................... 165–169, 174, 175, 177 Embryogenesis ......................................30, 144, 291, 312 Energetics ...................................................................... 312 Enzymatic activity .................................................. 30, 139 5-Ethynyl-20 -deoxyuridine (EDU) ..................... 166–177 Euchromatin.................................................................... 77

F Fibrinogen ...........................................182, 184, 187, 191 Fibronectin .................................................. 182, 184, 187 Fission yeast ...................... 19–21, 24, 26, 123, 124, 130, 278, 279 Flow cytometry .........................149, 151, 153, 165, 170, 171, 173, 174, 177, 195–203, 223, 224, 228, 229, 266, 267, 269–271 Fluorescent biosensor ........................................ 39, 42, 43 Fluorescent label ........................................................... 253

Fluorescent ubiquitination-based cell cycle indicator (Fucci) Fucci2A........................................................... 291–308 FluoroDishes ........................................................ 182, 185 Frog egg extracts..............................................31, 33, 144

G Geminin .........................8, 9, 15, 16, 144, 153, 224, 293 Gene editing .................................................................. 195 Genome engineering .................................................... 195 Genomic DNA ................ 83, 85–88, 100, 331, 333, 335 Glass coverslips FBN-patterned ............................................... 184–185 PLL-g-PEG-coated ............................... 183, 184, 190 G418 selection ..........................................................82, 92

H Heat dissipation.................................................... 311–320 Heterochromatin ................................................... 78, 240 High throughput enzyme assay ..................................... 52 Histone H1 ....................................97, 99, 104, 106, 124–126, 133–134, 136–138, 140 H2B ...................................... 186, 187, 189, 191, 327 H3K27me3 .................................................... 238, 240 Histone H1 kinase ............ 124, 125, 133, 134, 136–138 Histone modification .............................. 71–79, 237, 238

I Image analysis software Fucci tools ...................................................... 291–308 ImageJ/Fiji............................ 76, 150, 158, 183, 188, 231, 296, 298, 299, 301, 302, 308 MATLAB ............................................... 183, 188, 192 Trackosome .................................................... 183, 192 Immunoblotting ............................. 31, 33, 34, 209, 210, 213, 215–218, 329, 331, 333 Immunodepletion .....................................................32–34 Immunofluorescence labelling .......................................................... 184, 253 Immunostaining............................. 71–79, 202, 259, 262

K Kinase activity ..........6, 9, 11, 19–26, 39–48, 95–97, 140 Kinase assay ............................. 44, 97–99, 103, 105–108, 124, 125, 133, 134, 136–138 Kinase fusion ...........................................................95–108 Kinase inhibitor ................. 22, 42, 46, 48, 107, 205–219 Kinase purification...................................................95–108

L Label-free mass spectrometry ......................................... 53 Linker................................................................96, 97, 100

CELL CYCLE OSCILLATORS: METHODS Liquid chromatography-mass spectrometry (LC-MS) ..............................53, 55–59, 64, 66–68 Live-cell imaging spinning-disk ........................................................... 182

M Meiosis ............................................................................. 30 Metabolism........................................................... 112, 312 Micromanipulation ....................................................... 188 Micro-patterning........................................................... 191 MicroRNAs (miRNAs) .............................................81–93 Microscopy confocal.................................................................... 293 expansion ................................................249–262, 281 fluorescence ........ 130, 269, 270, 272, 273, 281, 282 immunofluorescence ...................................... 125, 128 live cell ............................................................ 180, 284 PALM superresolution................................... 277–287 spinning-disk ........................................................... 182 3D time-lapse ................................................. 240, 244 time-lapse................................................................. 245 Microtubule-organizing centers (MTOCs)........ 277–287 Mintbody ......................................................238–240, 245 Mitosis .............................. 9–16, 20, 30, 31, 33, 71, 111, 124, 125, 127, 128, 131, 138, 143–163, 179–192, 205–207, 215, 219, 230, 245, 251, 261, 266, 269, 270, 273, 283, 292–294, 301–303, 311

N Nested PCR..................................................................... 86 Nocodazole ............. 111–119, 145, 146, 151, 153, 209, 213–215, 218, 219, 267, 269, 270 Non-degradable cyclin B ................................... 32, 33, 36 Nucleus ........... 6, 9, 11, 13, 76, 77, 126, 130, 179, 180, 187–189, 191, 192, 234, 238, 239, 241, 242, 244, 245, 299, 300, 304

O Oscillations ............... 238, 239, 265, 293, 312, 319, 320

P Peptide ........................ 39–48, 53–61, 63–68, 96, 98, 99, 107, 108, 208, 210–213, 216–219, 292–294 Pheromone-induced Arrest ................................. 268, 269 Phosphatase inhibitors .........................22, 107, 147, 160, 209, 210, 212, 213, 216–219 Phospho-peptide affinity column ........................ 211, 212 Phosphorylation .................. 2, 4–6, 9–11, 13–16, 19–26, 30, 33, 40, 41, 51, 52, 64, 68, 97, 104, 106, 108, 124, 126, 127, 140, 143–145, 150, 153, 157, 162, 205–219, 238, 265, 266, 312 Phospho-specific antibodies generation....................................................... 210–213

AND

PROTOCOLS Index 339

purification ..................................................... 210–213 testing ............................................................. 210–213 Phospho-Threonine-Proline Mouse mAb (P-Thr-Pro-101) ...................................... 133, 138 Plate reader ........................ 31, 32, 34, 35, 42, 45, 46, 85 Polo-like kinase ............................................................. 206 Posterior bio-image reconstruction and analysis......... 180 pRb ..........................................................................4–7, 16 Proliferating cell nuclear antigen (PCNA) ......... 238, 240 Proteasome inhibitor .................147, 153, 156, 215, 218 Protein kinases...................... 1, 6, 11, 14, 20, 39, 40, 42, 51, 95, 96, 127, 205–219, 265 Protein phosphatase one (PP1)............... 6, 16, 129, 206, 207, 209, 215, 218 Proteolysis..............................2, 9, 11, 12, 143–145, 153, 156, 162, 323

Q Quantitative mass spectrometry ..................................... 53

R R..................................................54, 63, 64, 66, 314, 319 Radiation ..................................................... 223, 225, 228 Root ...........................................................................71–79 R-studio ..................................................... 54, 58, 64, 314

S Saccharomyces cerevisiae........................8, 58, 59, 65, 100, 107, 265–275, 278, 282 Schizosaccharomyces pombe.......................... 19, 22, 23, 25, 123–140, 278, 324 SDS-PAGE103, 105, 108, 146, 153, 158–160, 162, 209, 210, 215–217, 267, 271, 272, 331 Single-cell analysis ......................................................... 188 SLIM motifs .................................................................. 206 Specificity constant............................................. 53, 57, 64 Spindle-pole body (SPB) ....................129, 278–281, 283 Spindle staining ........................................... 272, 273, 275 Substrate specificity ................................... 30, 51–68, 206 Suspension cells ............................................................. 114 Synchronization .......................... 25, 111–120, 125–127, 129, 134–135, 138, 139, 151, 152, 155, 213–218, 266, 268–269, 313 Synthetic peptides .....................................................51–68

T Thymidine ................ 111–121, 196, 209, 213, 214, 218 Transfections ...................... 82, 84–86, 89, 91, 150, 160, 181, 183, 190, 198, 199, 202, 225, 227, 242, 243, 295, 296, 307, 330, 331

U Ubiquitin-mediated degradation .............................11, 16 3’ UTR ............................................................................ 82

CELL CYCLE OSCILLATORS: METHODS

340 Index

AND

PROTOCOLS

W

Z

Western blotting...........25, 39, 146, 151, 157, 159, 160, 215–217, 266, 270–272, 274, 331–333

Zebrafish breeding................................................................... 313 embryo calorimeter injection ................................. 313 embryos .......................................................... 311–320 embryo staging........................................................ 313

X X chromosome inactivation................................. 237–245 Xenografts...................................................................... 234 Xenopus laevis ................................................... 29–36, 278