Cell Polarity Signaling: Methods and Protocols 1071620347, 9781071620342

Table of contents :
Preface
Contents
Contributors
Chapter 1: Use of Fluorescence Recovery After Photobleaching (FRAP) to Measure In Vivo Dynamics of Cell Junction-Associated Po...
1 Introduction
1.1 The Use of Fluorescent Proteins in FRAP
1.2 FRAP on In Vivo Tissue Samples
1.3 FRAP on Planar Polarity Proteins
2 Materials
2.1 Materials for Mounting Live Pupae for Imaging of Pupal Wings
2.2 Microscope Equipment
2.3 Software for Analysis of FRAP Raw Data
3 Methods
3.1 Collecting and Staging Drosophila melanogaster White Prepupae
3.2 Exposing the Drosophila Pupal Wing for Imaging
3.3 Mounting the Drosophila Pupae on an Imaging Dish
3.3.1 Mounting Method 1
3.3.2 Mounting Method 2
3.4 Live-Imaging Settings
3.5 Optimizing the FRAP Imaging Method
3.5.1 Choosing ROI Size, Shape, and Number
3.5.2 Selecting Control Unbleached Regions and Background Regions
3.5.3 Choosing the Number of Pre-bleach Images to Record
3.5.4 Specifying the Scanning Conditions Required for the Initial Bleach
3.5.5 Selecting the Number of Replicate Samples
3.5.6 Summary of FRAP Settings Used for Junctionally Associated Planar Polarity Proteins in the Drosophila Pupal Wing
3.6 Performing the FRAP Experiment
3.7 FRAP Processing Method
3.7.1 FRAP Data Collecting
3.7.2 FRAP Data Normalization
3.7.3 Graph Plotting to Check the Results
3.7.4 Curve Fitting and Extraction of Summary Data
3.7.5 Comparing Curves Between Data Sets
3.7.6 Calculating Mobile and Immobile Amounts
3.7.7 (Optional) Correcting Mean Intensity Data if the Samples Have a Variable Distance from the Coverslip
3.8 Publishing FRAP Data
4 Notes
References
Chapter 2: FRET Imaging of Rho GTPase Activity with Red Fluorescent Protein-Based FRET Pairs
1 Introduction
2 Materials
2.1 Plasmid
2.2 Reagents for Ser-FRET Imaging
2.3 Reagents for FLIM-FRET Imaging
2.4 Fluorescence Microscope
3 Methods
3.1 Ser-FRET Imaging
3.1.1 Hippocampal Neuron Preparation
3.1.2 Image Acquisition
3.1.3 Image Analysis
3.2 FLIM-FRET Imaging
3.2.1 Hippocampal Slice Preparation
3.2.2 Image Acquisition
3.2.3 Image Analysis
4 Notes
References
Chapter 3: Live-Cell Total Internal Reflection Fluorescence (TIRF) Microscopy to Investigate Protein Internalization Dynamics
1 Introduction
2 Materials
3 Methods
3.1 Laser Alignment and Focusing of Through-the-Objective TIRF
3.2 Preparing the Microscope
3.3 Imaging
3.4 Data Analysis
4 Notes
References
Chapter 4: Single-Cell Single-Molecule Pull-Down (sc-SiMPull) for Detection of Protein Complexes from Embryonic Lysates
1 Introduction
2 Materials
2.1 Microfluidic Device Master Molds
2.2 Microfluidic Devices
2.3 PEG Functionalization of Devices
2.4 C. elegans Strains and Fluorescent Labeling
2.5 Microscope
2.6 Analysis Software
3 Methods
3.1 Mold Fabrication for Microfluidic Devices
3.2 SiMPull Device Fabrication
3.2.1 Making PDMS Devices
3.2.2 Cleaning Coverslips with UV Ozone Cleaner
3.2.3 Cleaning Coverslips with Piranha Solution
3.2.4 Assembling PDMS Devices
3.3 PEGylation
3.4 Antibody Functionalization
3.5 Halo Dye Labeling
3.6 Embryo Dissection
3.7 Sealing and Lysis
3.7.1 Tape Sealing Method
3.7.2 Valap Sealing Method
3.7.3 Lysis
3.7.4 Lysis Before Sealing (for Colocalization Controls/Highly Abundant Proteins)
3.8 Data Acquisition
3.9 Automated Data Analysis
3.9.1 Analysis Software Installation
3.9.2 Automated Data Processing
3.10 Post Processing and Data Visualization
3.10.1 Sort Images and Discard Artifacts: colocalization_inspector_GUI
3.10.2 Evaluate Experiment Quality and Lysis: ``spotcount_1dplotter´´
3.10.3 Determine Labeling Efficiency and Protein Colocalization
3.10.4 Stoichiometry and Photobleaching Step Counting: See Note 10
4 Notes
References
Chapter 5: Biochemical Assays to Detect Activation of Small GTPases Rho, Rac, and Cdc42 during Morphogenesis
1 Introduction
2 Materials
2.1 Bacteria Cell Culture
2.2 Mammalian Cell Culture
2.3 Embryos
2.4 GST-PRD and GST-PBD Binding Assay Buffers
2.5 Western Blotting
3 Methods
3.1 Rho and Rac/Cdc42 Assays
3.1.1 Preparation of Recombinant GST-RBD Protein
3.1.2 Preparation of Recombinant GST-PBD Fusion Protein
3.1.3 Extraction of GST-RBD
3.1.4 Extraction of GST-PBD
3.1.5 Sample Preparations for Rho Pull-Down Assays Using Mammalian Cells
3.1.6 Sample Preparations for Rac/Cdc42 Assay Using Mammalian Cells
3.1.7 Sample Preparations for Rho/Rac/Cdc42 Assay Using Xenopus Embryos and Explants
3.1.8 GST-RBD and GST-PBD Binding Assay
3.2 Western Blot Analysis
4 Notes
References
Chapter 6: Analysis of Planar Cell Polarity Complexes by Proximity Biotinylation in Xenopus Embryos
1 Introduction
2 Materials
2.1 Xenopus Culture and Manipulation
2.2 Proximity Biotinylation and Protein Detection
3 Methods
3.1 Xenopus Embryo Culture and Microinjections
3.2 Biotinylation
3.3 Immunoprecipitation
3.4 Immunoblotting
3.5 Enhanced Chemiluminescence
4 Notes
References
Chapter 7: Regulation of Cell Polarity by Posttranslational Protein Palmitoylation
1 Introduction
2 Materials
2.1 Cell Culture Medium and Transfection Reagent
2.2 Metabolic Labeling and Click Reaction
2.3 Streptavidin Bead Pull-Down
2.4 Streptavidin Blot and Western Blot
2.5 Antibody
2.6 Constructs
2.7 3D Culture and Immunofluorescence
3 Methods
3.1 Chemical Proteomic Profiling of Palmitoylated Proteins
3.1.1 Metabolic Labeling
3.1.2 Protein Extraction
3.1.3 Click Reaction
3.1.4 Streptavidin Bead Pull-Down
3.1.5 Mass Spectrometry Analysis
3.2 Validation of SCRIB Palmitoylation
3.2.1 Transfection
3.2.2 Metabolic Labeling
3.2.3 Protein Extraction
3.2.4 Click Reaction
3.2.5 Streptavidin Blot
3.2.6 Western Blot
3.3 Identification of SCRIB Palmitoylation Sites
3.4 Exploration of the Mechanisms Regulating SCRIB Palmitoylation
3.4.1 ZDHHC-PAT Screen
3.4.2 ZDHHC-PAT and SCRIB Interactions by Co-IP Assay
3.4.3 ZDHHC7 Inactivation Analysis
3.4.4 ZDHHC7 Knockdown and Rescue Assay
3.5 Evaluation of SCRIB Palmitoylation in Regulating Cell Polarity
3.5.1 Culture MCF-10A Cells Under 3D Culture Conditions
3.5.2 Immunofluorescence Analysis of Apical-Basal Polarity
4 Notes
References
Chapter 8: Enrichment and Detection of Wnt Proteins from Cell Culture Media
1 Introduction
2 Materials
2.1 Preparation of Sample
2.2 Wnt Precipitation Using BS
2.3 Immunoblotting to Detect BS-Precipitated Wnt Proteins
3 Methods
3.1 Preparation of Sample (See Note 5)
3.2 Wnt Precipitation Using BS
3.3 Immunoblotting to Detect BS-Precipitated Wnt Proteins
4 Notes
References
Chapter 9: Using Live Imaging to Examine Early Cardiac Development in Zebrafish
1 Introduction
2 Materials
2.1 Mounting
2.2 Imaging
3 Methods
3.1 General Guidelines
Box 1 General Guidelines and Tips
3.1.1 Tailoring
3.1.2 Minimize Obstacles
3.2 Cardiomyocyte Specific Protocol
3.2.1 Preparation
3.2.2 Mounting
3.2.3 Imaging
3.2.4 Preprocessing of Images
3.2.5 Cell Tracking Analysis
4 Notes
References
Chapter 10: Imaging Planar Cell Polarity Proteins in Xenopus Neuroectoderm
1 Introduction
2 Materials
2.1 Xenopus Embryo Culture and Manipulation
2.2 PCP Protein Imaging
2.2.1 Plasmids and Antibodies
Plasmids
2.2.2 Buffers and Chemicals
2.2.3 Equipment
2.2.4 Embryo Handling Tools
3 Methods
3.1 Xenopus Embryo Culture, Microinjections, and Phenotypic Analysis
3.2 Whole Mount Immunostaining for Vangl2
3.3 Detection of Polarized Ectopic GFP-Pk3 and HA-Vangl2 Complexes in the Neural Plate (Fixed Samples)
3.3.1 Direct Fluorescence
3.3.2 Double Immunostaining
3.4 Live Imaging of Embryos Expressing Fluorescent PCP Proteins
3.5 Quantitative Analysis of PCP Based on Live or Fixed Tissue Images
4 Notes
References
Chapter 11: In Ovo Gain- and Loss-of-Function Approaches to Study Gut Morphogenesis
1 Introduction
1.1 Dorsal Mesentery as a Model to Study Gut Morphogenesis
1.2 In Ovo Electroporation of Plasmid DNA
1.3 In Ovo Electroporation of Morpholinos
1.4 In Ovo Pharmacological Targeting of Cellular Processes Through Surgical Insertion of Chemical-Soaked Resin Beads
2 Materials
2.1 In Ovo Electroporation of Plasmid DNA
2.1.1 Reagents
2.1.2 Tools
2.2 In Ovo Electroporation of Morpholinos
2.2.1 Reagents
2.2.2 Tools
2.3 In Ovo Pharmacological Targeting of Cellular Processes Through Surgical Insertion of Chemical-Soaked Resin Beads
2.3.1 Reagents
2.3.2 Tools
3 Methods
3.1 In Ovo Electroporation of Plasmid DNA
3.1.1 Egg Setup and Incubation
3.1.2 Windowing Eggs
3.1.3 DNA Electroporation
3.2 In Ovo Electroporation of Morpholinos
3.2.1 Nature of Oligos to Be Ordered
3.2.2 Making Stock Solution of Morpholino and Storage
3.2.3 Electroporating Morpholinos
3.3 In Ovo Pharmacological Targeting of Cellular Processes Through Surgical Insertion of Chemical-Soaked Resin Beads
4 Conclusion
5 Notes
References
Chapter 12: Avian Embryos as a Model to Study Vascular Development
1 Introduction
1.1 Dorsal Mesentery as a Model to Study Vascular Development
1.2 Live Imaging Using Transgenic Quails
1.3 Quail-Chick Transplantation
2 Materials
2.1 Live Imaging Using Transgenic Quails
2.1.1 Reagents
2.1.2 Tools
2.2 Quail-Chick Transplantation
2.2.1 Reagents
2.2.2 Tools
3 Methods
3.1 Live Imaging Using Transgenic Quails
3.1.1 Preparation of Embryonic Quail Slices
3.1.2 Embedding Tissue Slices
3.2 Quail-Chick Transplantation
3.2.1 Preparation of Neutral Red/Agarose to Mark the Site of Engraftment
3.2.2 Preparing the Donor (Quail) Graft Tissue
3.2.3 Preparing the Host (Chicken) Embryos to Receive the Grafts
3.2.4 Transplanting
4 Conclusion
5 Notes
References
Chapter 13: BAC Recombineering and Transgenesis to Study Cell Polarity and Polarized Tissue Morphogenesis in Mice
1 Introduction
2 Materials
2.1 Media and Antibiotics for BAC Culture
2.2 Solutions, Materials, and Equipment for BAC DNA Mini Prep
2.3 Restriction Digest and Pulse Field Gel Electrophoresis (PFGE)
2.4 Electroporation of BAC DNA into Recombineering Competent Strains
2.5 BAC Recombineering with Linear Template
2.6 Purification of BAC DNA for Pronuclear Injection
2.7 Use BAC Transgenic Mice to Study Polarized Tissue Morphogenesis
3 Methods
3.1 Identify and Acquire the Desired BAC Clones
3.2 DNA Purification from BAC Clones
3.3 Restriction Digest and Pulse Field Gene Electrophoresis (PFGE) to Characterize the Purified BAC DNA
3.4 Electroporate BAC DNA into Recombineering Competent Bacterial Strains
3.5 BAC Recombineering with Linear Template
3.6 Purification of BAC DNA for Pronuclear Injection
3.7 Use BAC Transgenic Mice to Study Polarized Tissue Morphogenesis
3.7.1 Specimen Preparation
3.7.2 Whole Mount Phalloidin Staining and RIMS Clearing
3.7.3 Confocal Imaging and 3D Reconstruction
4 Notes
References
Chapter 14: Two-Photon Cell and Tissue Level Laser Ablation Methods to Study Morphogenetic Biomechanics
1 Introduction
2 Materials
2.1 Dissection Materials
2.2 Confocal Microscope Specifications
3 Methods
3.1 Before Starting the Experiment
3.2 Embryo Collection
3.3 Embryo Positioning
3.3.1 Embryo Positioning for Cell Border Ablations
3.3.2 Embryo Positioning for Tissue-Level Ablations
3.4 Imaging and Ablation Settings
3.4.1 Imaging Settings for Cell Border Ablations
3.4.2 Imaging Settings for Tissue-Level Ablations
3.5 Image Analysis
3.5.1 Image Analysis of Cell Border Ablations
3.5.2 Image Analysis of Tissue-Level Ablations
4 Notes
References
Chapter 15: Protocols for Investigating the Epithelial Properties of Cardiac Progenitor Cells in the Mouse Embryo
1 Introduction
2 Materials
2.1 Immunofluorescence on Paraffin Sections and Whole Mount Preparations of the Dorsal Pericardial Wall Epithelium
2.1.1 Whole-Embryo Paraffin Embedding
2.1.2 Paraffin-Embedded Tissue Sectioning
2.1.3 Immunofluorescence of Paraffin-Embedded Sections
2.1.4 Immunofluorescence and Image Analysis of Whole Mount DPW Epithelium
2.2 Ex Vivo Mouse Embryo Culture and Dorsal Pericardial Wall Electroporation
2.2.1 Mouse Embryo Culture
2.2.2 Mouse Embryo Electroporation
2.3 Culture and Time-Lapse Imaging of Thick Transverse Embryo Sections
2.4 Mechanical Stress Assessment Using an Epithelial Wounding Assay
3 Methods
3.1 Immunofluorescence on Paraffin Sections and Whole Mount Preparations of the Dorsal Pericardial Wall Epithelium
3.1.1 Whole-Embryo Paraffin Embedding
3.1.2 Paraffin-Embedded Tissue Sectioning
3.1.3 Immunofluorescence of Paraffin-Embedded Sections
3.1.4 Immunofluorescence and Image Analysis of Whole Mount DPW Epithelium
3.2 Ex Vivo Mouse Embryo Culture and Dorsal Pericardial Wall Electroporation
3.2.1 Mouse Embryo Collection and Ex Vivo Culture
3.2.2 Dorsal Pericardial Wall Electroporation
3.3 Culture and Time-Lapse Imaging of Thick Transverse Embryo Sections
3.4 Mechanical Stress Assessment Using an Epithelial Wounding Assay
4 Notes
References
Chapter 16: Methods to Investigate Cell Polarity of Inner Ear
1 Introduction
2 Materials
2.1 Tools
2.2 Equipment
2.3 Reagents
2.3.1 For Dissection and Fixation
2.3.2 For Paint-Filling
2.3.3 For Frozen Section
2.3.4 For Scanning Electron Microscopy (SEM)
2.3.5 For Organ Culture
2.3.6 For EdU Detection
2.3.7 For Staining
3 Methods
3.1 Dissection of Inner Ear
3.1.1 Cochlea Dissection
3.1.2 Vestibule Dissection
3.2 Paint-Filling for Gross Development of Cochlea
3.2.1 Preparation for Paint-Filling
3.2.2 Paint-Filling
3.3 Frozen Section
3.3.1 Inner Ear Sample Preparation
3.3.2 Frozen Tissue Sectioning
3.4 SEM
3.4.1 Preparation for SEM
3.4.2 SEM
3.4.3 SEM Examination and Analysis
3.5 Whole Embryo or Organ Culture
3.5.1 Preparing for Sterile Devices
3.5.2 Preparing Complex Medium of Cell Culture and Coatings
3.6 EdU Injection in Pregnant Females for Visualization of Proliferating Cells
3.6.1 Preparation of EdU Solution
3.6.2 Injecting EdU to Pregnant Mice
3.6.3 EdU Staining
3.6.4 DNA Staining
3.7 Whole Mount or Frozen Sections Antibody Staining
3.7.1 Sample Preparation
3.7.2 Immunostaining
3.7.3 Antibody Selection
3.7.4 Mounting and Confocal Image
3.7.5 Fluorescence Observation and Image Acquisition
3.8 Image Processing and Quantification
3.8.1 Superimpose Image (Merge)
3.8.2 Image Stack Z Projection
3.8.3 Quantification of Cochlea
4 Notes
References
Chapter 17: Characterization of Axon Guidance Phenotypes in Wnt/PCP Mutant Mice
1 Introduction
1.1 Axon Guidance Phenotypes
2 Materials
2.1 Genetic Mouse Models
2.2 Immunofluorescence
2.3 In Situ Hybridization
2.4 DiI Tracing
2.5 Ex Utero Electroporation
2.6 Tamoxifen-Induced Sparse Labeling of Commissural Axons
3 Methods
3.1 Obtain or Create a Mouse Model of Your Gene of Interest
3.2 Immunofluorescence
3.3 In Situ Hybridization
3.4 DiI Tracing of Open-Book Explants
3.5 Ex Utero Electroporation
3.6 Tamoxifen-Induced Sparse Labeling of Commissural Axons
4 Notes
References
Chapter 18: In Vitro Explant Assays and Cultures to Study PCP Signaling in Axon Guidance
1 Introduction
2 Materials
2.1 Open-Book Cultures
2.1.1 Dissection, Culture, and Staining
2.1.2 Pharmacological Treatment
2.2 Pre- or Postcrossing Explant Cultures
2.2.1 Explant Dissection and Culture
2.2.2 Cell-Line Aggregates
2.3 Dissociated Neuronal Culture
2.3.1 Culture Preparation
2.3.2 Dunn Chamber Assay
2.3.3 Filopodia Tips and Vesicle Trafficking
2.3.4 Growth Cone-Growth Cone Interaction
3 Methods
3.1 Open-Book Culture
3.1.1 Dissection
3.1.2 Rat Tail Collagen Preparation
3.1.3 Culture
3.1.4 Fixation and Staining
3.1.5 Pharmacologic Treatment
3.1.6 Quantification of the Phenotype
3.2 Explant Culture
3.2.1 Explant Dissection
3.2.2 Culture Setting
3.2.3 Fixation and Staining
3.2.4 Cell-Line Aggregates
3.3 Dissociated Neuronal Culture
3.3.1 Dissociation and Culture
3.3.2 Dunn Chamber Assay
3.3.3 Filopodia Tips and Vesicle Trafficking
3.3.4 Growth Cone-Growth Cone Interaction
4 Notes
References
Chapter 19: Biochemical and Cellular Assays to Study Mechanisms of PCP Signaling in Axon Guidance
1 Introduction
2 Materials
2.1 Coimmunoprecipitation
2.2 Signaling Assays for PCP Pathway
2.3 Transcellular Interaction Assay
2.4 AP Binding Assay
3 Methods
3.1 Coimmunoprecipitation
3.2 Signaling Assays for PCP Pathway
3.3 Transcellular Interaction Assay
3.4 AP Binding Assay
4 Notes
References
Chapter 20: Dissection, Fixation, and Immunostaining of the Drosophila Midgut
1 Introduction
2 Materials
2.1 Generation of Mosaic Clones in Adult Drosophila Midgut (see Note 1)
2.1.1 Fly Stocks
2.1.2 Fly Food in Vials
2.1.3 Incubators
2.2 Fly Gut Dissection
2.3 Fixation
2.3.1 Heat Fixation Equipment (Fig. 2)
2.3.2 Buffers
2.4 Immunostaining
2.4.1 Buffers and Antibodies
2.4.2 Specimen Mounting
2.5 Imaging
3 Methods
3.1 Generation of Mutant Clones in the Adult Drosophila Midgut
3.1.1 Larval Heat Shock (LHS)
3.1.2 Adult Heat Shock (AHS)
3.2 Adult Fly Midgut Dissection
3.2.1 Preparation for Sample Dissection and Fixation
3.2.2 Sample Dissection
3.3 Heat Fixation
3.4 Immunostaining
3.5 Imaging
4 Notes
References
Chapter 21: In Vivo Analysis of Pathways Regulating Epithelial Polarity and Secretion Using Drosophila Salivary Glands
1 Introduction
2 Materials
2.1 Fly Stocks
2.2 Larvae Collection and SG Dissection
2.3 Fixation and Immunostaining of SGs
2.4 Food Intake Assay
3 Methods
3.1 Fly Crossings, Egg Collection, and Larval Staging
3.2 Preparation of Larvae for Dissections (See Note 6)
3.3 Dissection of SGs for Confocal Live Imaging
3.4 Confocal Live Imaging of Larval SG
3.5 Dissection and Fixation of Larval SGs for Immunostaining
3.6 Image Quantifications
3.7 Generating Movies from Time-Lapse Recordings
3.8 Food Intake Assay
4 Notes
References
Chapter 22: Imaging Epidermal Cell Rearrangement in the C. elegans Embryo
1 Introduction
2 Materials
2.1 Mounting Embryos for Imaging of Cell Rearrangement
2.1.1 Reagents
2.1.2 Equipment
2.2 4D Nomarski Imaging of Morphogenesis
2.3 Fluorescence Imaging of Morphogenesis
2.3.1 Phalloidin Staining for Analyzing Morphogenesis
2.4 Inducible Expression of Rho Family GTPase Constructs Using the NMD System
2.5 Laser Killing of Blastomeres
3 Methods
3.1 Mounting Embryos for Imaging of Cell Rearrangement
3.1.1 Agar Mounts for Imaging Morphogenesis; Modified from
3.1.2 Other Mounting Methods
3.2 4D Nomarski Imaging of Morphogenesis
3.2.1 Imaging Setup
3.2.2 Setting Up a 4D (or 5D) Acquisition Sequence
3.2.3 Viewing 4D/5D Datasets
3.2.4 Introducing Pharmacological Agents During 4D Acquisition
3.2.5 Creating Colorized Overlays on 4D Nomarski Movies
3.3 Fluorescence Imaging of Morphogenesis
3.3.1 General Considerations for 4D Fluorescence Imaging of Cell Rearrangement
3.3.2 Probes for Visualizing Morphogenesis
3.3.3 Quantifying Protrusive Behavior During Dorsal Intercalation
3.4 Inducible Expression of Rho Family GTPase Constructs Using the NMD System
3.5 Laser Killing of Blastomeres in Caenorhabditis elegans
3.6 Summary
4 Notes
References
Chapter 23: Methods for the Study of Apical Constriction During Ascidian Gastrulation
1 Introduction
1.1 Contributions of the Study of Fixed Embryos
1.2 Contribution of the Study of Live Embryos
1.3 Imaging Ascidian Embryos
2 Materials
2.1 Embryo Preparation
2.1.1 General Reagents
2.1.2 Ciona Work
2.1.3 Phallusia Work
2.2 Introducing Pharmacological Agents During 4D Acquisition
2.2.1 Phospho-Myosin Immunostaining
2.2.2 Phalloidin Actin Labeling
2.3 Analysis of Apical Constriction in Live Embryos Using Light Sheet Microscopy
3 Methods
3.1 Embryo Preparation
3.1.1 Gamete Collection
3.1.2 Egg Dechorionation
For Ciona
For Phallusia
3.1.3 Egg Microinjection
Preparation and Storage of Microinjection Needles
Preparing for Microinjection Under a Stereoscope
Preparation of a Microinjection Chamber to Use with an Inverted Microscope
Preparing for Microinjection Under an Inverted Microscope
Microinjection Under a Stereoscope or Inverted Microscope
3.1.4 Fertilization of Ascidian Eggs
Phallusia
Ciona
3.1.5 Embryo Staging
3.2 Analysis of Apical Constriction in Fixed Embryos
3.2.1 Phospho-Myosin Immunostaining
3.2.2 Preparation of Poly-l-Lysine-Coated Coverslips
3.2.3 Fluorescent Phalloidin Staining of Actin Cytoskeleton and Sample Mounting
3.3 Analysis of Apical Constriction in Live Embryos Using Light Sheet Microscopy
3.3.1 Fluorescent Reporters of Cell Membranes and Actomyosin Network and Optical Perturbation Tools
3.3.2 Preparation of the Sample Holder for Multiangle SPIM Imaging
3.3.3 Mounting and Imaging of Live Samples in a Multiangle SPIM
3.4 Pharmacological and Optogenetic Tools to Alter Cytoskeleton Activity
3.5 Morphological Measurements
3.5.1 Measuring Shape in Fixed Samples Imaged by Confocal Microscopy
3.5.2 Measuring Shape in Live Samples Imaged with Multiangle Light Sheet Microscopy
3.6 Myosin Activity Measurements
4 Notes
References
Chapter 24: Assays for Apical Constriction Using the Xenopus Model
1 Introduction
2 Materials
2.1 Obtaining and Culturing Xenopus Embryos
2.2 Microinjection of Early Xenopus Embryos
2.3 Staining of Actin and Activated Myosin II
2.4 Live Imaging of Actomyosin Dynamics During Gastrulation and Neurulation
3 Methods
3.1 Analyze Ectopic Induction of Apical Cell Constriction in the Animal Region (See Note 12)
3.1.1 Assessing Apical Constriction by Cell Morphology
3.1.2 Quantification of Apical Cell Surface Reduction
3.1.3 Visualization of Apical Cell Surface by Fluorescent Imaging
3.2 Inspect Actomyosin Cytoskeleton in Cells Undergoing Apical Constriction
3.2.1 Examine F-Actin Distribution in Fixed Samples
3.2.2 Analyze the Patterns of Activated Myosin in Fixed Samples
3.2.3 Explore Actomyosin Dynamics in Live Samples
3.3 Investigate Gene Function in Apical Constriction of Bottle Cells During Xenopus Gastrulation
3.4 Assess Apical Constriction During Neural Tube Closure
3.4.1 Preparation of Embryos for Confocal Imaging of NTC
4 Notes
References
Chapter 25: The Use of Three-Dimensional Cell Culture to Study Apicobasal Polarization and Lumen Formation
1 Introduction
2 Materials
2.1 MDCK Cell Culture
2.2 Generation of Stable Cell Lines
2.3 Immunofluorescence (IF) Method 1
2.4 IF Method 2
2.5 Imaging and Analysis
3 Methods
3.1 Generation of Stable Cell Lines
3.1.1 Lentiviral Production
3.1.2 Infection of MDCK Cells with Lentivirus
3.2 3D Overlay Culture of MDCK Cells
3.3 Immunofluorescence (IF) Method 1
3.4 Immunofluorescence (IF) Method 2 (Use for PIP Antibodies)
4 Notes
References
Chapter 26: Studying Cell Polarity Dynamics During Cancer Initiation Using Inducible 3D Organotypic Cultures
1 Introduction
2 Materials
2.1 Cell Line
2.2 Primary Cells
2.3 Lentivirus Production
2.4 Materials for Growing Cells in 3D Cultures
2.5 Immunostaining
3 Methods
3.1 Caco2 Cell Culture
3.1.1 Maintenance of Cultures
3.1.2 Infect Cells with Lentivirus
3.1.3 Growing 3D Cysts from Caco2 Cells (See Notes 4 and 5)
3.2 Primary Mouse Mammary Epithelial Cells
3.2.1 Isolation of Primary Mouse Mammary Epithelial Cells (See Note 6)
3.2.2 Infect Primary Cells with Lentivirus
3.2.3 Growing 3D Cysts from Primary Epithelial Cells (See Notes 4 and 5)
3.3 Induction of Gene Expression: Optional (See Note 7)
3.3.1 Caco2 Cysts
3.3.2 Primary Murine Organotypic Cultures
3.4 Immunostaining (See Notes 8-10)
3.4.1 Caco2 Cell Line
3.4.2 Mouse Mammary Organoids
3.5 Anticipated Results
4 Notes
References
Chapter 27: Under-Agarose Chemotaxis and Migration Assays for Dictyostelium
1 Introduction
2 Materials
2.1 For Dictyostelium Cell Growth
2.2 For Preparing Agarose Plates
2.3 Chemoattractants
2.4 Preparation of cAMP Competent Cells
2.5 Imaging
2.6 Image Analysis
3 Methods
3.1 Growth of Dictyostelium Cells for Chemotaxis Assays
3.2 Folate Under-Agarose Chemotaxis Assay
3.2.1 Folate Under-Agarose Chemotaxis: Preparation of Agarose Plates
3.2.2 Folate Under-Agarose Chemotaxis Assay: One Well Assay
3.2.3 Folate Under-Agarose Chemotaxis Assay: Two Well Assay
3.3 cAMP Under-Agarose Chemotaxis Assay
3.3.1 cAMP Under-Agarose Chemotaxis Assay: Preparation of cAMP Responsive Cells
3.3.2 cAMP Under-Agarose Chemotaxis Assay: Preparing Agarose
3.4 Imaging
3.5 Quantitative Image Analysis
3.5.1 Analysis and Measurement of Cell Speed, Directionality, and Chemotactic Index
3.5.2 Image Analysis: Cell Morphology
3.5.3 Image Analysis: Pseudopod Dynamics
4 Notes
References
Chapter 28: Mapping Asymmetry in Collective Cell Migration: Lessons from Border Cells in Drosophila Oogenesis
1 Introduction
2 Materials
2.1 Live Cell Imaging
2.1.1 Reagents and Dissection Tools
2.1.2 Equipment for Live Imaging
2.2 Tissue Immunohistochemistry
2.2.1 Reagents and Dissection Tool
2.3 Microscopy and Image Analysis
3 Methods
3.1 Time-Lapse Imaging and Analysis of Actin-Based Protrusions in Migrating Border Cell Clusters
3.2 Quantitative Analysis of Border Cell Clusters Captured by Live Cell Time-Lapse Imaging
3.2.1 Speed Calculation
3.2.2 Length of Protrusion
3.2.3 Direction of Protrusion
3.2.4 Number of Protrusions
3.2.5 Stability of Protrusion
3.3 Immunohistochemistry of Polarity Markers and Analysis of Distribution of Apical Basal Proteins
3.3.1 Immunostaining
3.3.2 Distribution of Apical Basal Polarity Proteins
4 Notes
References
Chapter 29: A Toolbox to Study Tissue Mechanics In Vivo and Ex Vivo
1 Introduction
2 Materials
2.1 Microinjection
2.1.1 Embryo Medium
2.1.2 Plasmids and mRNA
2.1.3 Microinjection Equipment
2.1.4 Immunofluorescence
2.2 AFM Measurements
2.3 Polyacrylamide Hydrogels
2.4 Neural Crest Dissection Tools and Protocol
2.5 Image and Statistical Analysis
3 Methods
3.1 Mesoderm Targeted Injections and Validation
3.1.1 Preparation for Microinjection
3.1.2 Setting Up the Microinjector
3.1.3 Microinjection
3.1.4 Confirmation of Injection Accuracy by Immunofluorescent Staining of Fibronectin
3.2 Analyze the Impact of PCP Inhibition on Mesoderm Cell Density In Vivo
3.3 Measuring the Impact of PCP Inhibition on Mesoderm Stiffening by In Vivo Atomic Force Microscopy (iAFM)
3.3.1 Initializing the Nanosurf Flex-ANA System (See Note 14)
3.3.2 Calibrating the Cantilever in the Nanosurf Flex-ANA System
3.3.3 Set Up the Parameters and Perform a Test Measurement
3.3.4 Determining Mesodermal Apparent Elasticity of Control and Dsh-DEP+ Embryos
3.3.5 In Vivo Atomic Force Microscopy (iAFM) Data Analysis and Presentation
3.4 Ex Vivo Analyses of NC Polarity Using Hydrogels
3.4.1 Glass Coating
3.4.2 Polyacrylamide Hydrogels Preparation
3.4.3 Gel Activation
3.4.4 Ex Vivo Analyses of Neural Crest Polarity
4 Notes
References
Chapter 30: Quantitative Analysis of Directional Neural Crest Cell Migration
1 Introduction
2 Materials
2.1 Embryo Injection
2.1.1 Embryo Medium
2.1.2 Microinjection
2.2 Explant Dissection and Culture
2.2.1 Dissection Tools
2.2.2 Matrix Preparation
2.2.3 Explant Culture
2.3 Imaging and Analysis
3 Methods
3.1 Microinjection to Label Neural Crest Cells
3.2 Prepare FN-Coated Dish
3.3 Neural Crest Explant Dissection
3.4 Neural Crest Explant Culture
3.5 Imaging and Analyzing Neural Crest Cell Migration
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2438

Chenbei Chang Jianbo Wang Editors

Cell Polarity Signaling Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-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 Polarity Signaling Methods and Protocols

Edited by

Chenbei Chang and Jianbo Wang Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA

Editors Chenbei Chang Department of Cell, Developmental and Integrative Biology University of Alabama at Birmingham Birmingham, AL, USA

Jianbo Wang Department of Cell, Developmental and Integrative Biology University of Alabama at Birmingham Birmingham, AL, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-0716-2034-2 ISBN 978-1-0716-2035-9 (eBook) https://doi.org/10.1007/978-1-0716-2035-9 © Springer Science+Business Media, LLC, part of Springer Nature 2022 The chapter 29 is 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 chapter. This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Cellular polarity refers to asymmetry within a cell and can be manifested in several forms. Apicobasal (AB) polarity in the epithelium segregates cell surface from internal cell environment, whereas planar cell polarity (PCP) coordinates asymmetric molecular organization within the plane of tissues. Both polarity cues can also be transferred into front-to-rear polarity in directional migrating cells. Proper establishment and maintenance of cell polarity is essential for the formation and morphogenesis of tissues and organs during embryonic development and is also required for integrity, homeostasis, and function of adult organs. Hence, in-depth knowledge of cell polarity will not only yield profound insight into basic biology of cell asymmetry in all aspects, including molecular, cellular, and tissue-level regulation and function, but also help provide understanding about disease processes that result from impaired cell polarity, such as congenital birth defects and cancer. To facilitate the research effort in cell polarity investigation, we have assembled a collection of a wide range of protocols that cover the commonly used methods to interrogate and manipulate cell polarity both in cultured cell systems and in animal models in this special topic issue of Methods in Molecular Biology. All the chapters in this volume are contributed by leading researchers studying different aspects of biology related to cell polarity. The first eight chapters cover advanced imaging and biochemical methods that are employed to investigate dynamics, conformation, interaction, and modification of proteins involved in cell polarity. The following eleven chapters focus on PCP signaling in morphogenesis in diverse developmental contexts, such as that of heart, vasculature, neural tube, gut, inner ear, and axon guidance. The next seven chapters describe experimental techniques used in the investigation of biological regulation of properties linked to AB cell polarity in development and diseases. The final four chapters are centered around the issues of directional cell migration and biomechanics involved in the process. The collection includes the usage of a variety of model systems, such as Dictyostelium, Drosophila, C. elegans, Ascidian, zebrafish, Xenopus, chick, and mouse, in addition to 3-dimensional (3D) cultured cell systems. An array of techniques are covered, including genetics, imaging, biochemical, and biomechanical. Diverse biological processes are explored using these techniques, such as organ morphogenesis, apical constriction, directional cell migration, and tumorigenesis. We hope that these methods will enable additional researchers to delve into the stimulating field of cell polarity and contribute to our understanding of how coordinated control of protein stability, trafficking, membrane retention, post-translational modification, and dynamic organization leads to active regulation of cell polarity. This book is made possible only because all the contributors have devoted their precious time and effort to provide a concise and detailed account of their experimental procedures with helpful notes to overcome technical hurdles. We are grateful to them for their excellent contributions and their enthusiasm to share their technical expertise and insight with both seasoned and novice researchers in the field. Birmingham, AL, USA

Chenbei Chang Jianbo Wang

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

1 Use of Fluorescence Recovery After Photobleaching (FRAP) to Measure In Vivo Dynamics of Cell Junction–Associated Polarity Proteins . . . . . . . . . . . . . . Samantha J. Warrington, Helen Strutt, and David Strutt 2 FRET Imaging of Rho GTPase Activity with Red Fluorescent Protein-Based FRET Pairs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bryce T. Bajar, Xinmeng Guan, Amy Lam, Michael Z. Lin, Ryohei Yasuda, Tal Laviv, and Jun Chu 3 Live-Cell Total Internal Reflection Fluorescence (TIRF) Microscopy to Investigate Protein Internalization Dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tejeshwar C. Rao, Tomasz J. Nawara, and Alexa L. Mattheyses 4 Single-Cell Single-Molecule Pull-Down (sc-SiMPull) for Detection of Protein Complexes from Embryonic Lysates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Naomi Stolpner and Daniel J. Dickinson 5 Biochemical Assays to Detect Activation of Small GTPases Rho, Rac, and Cdc42 during Morphogenesis . . . . . . . . . . . . . . . . . . . . . . Mark L. Berns and Raymond Habas 6 Analysis of Planar Cell Polarity Complexes by Proximity Biotinylation in Xenopus Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ilya Chuykin and Sergei Y. Sokol 7 Regulation of Cell Polarity by Posttranslational Protein Palmitoylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Baoen Chen, Carla Guarino, Abdelhalim Azzi, Hannah Erb, and Xu Wu 8 Enrichment and Detection of Wnt Proteins from Cell Culture Media. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pooja R. Sonavane and Karl Willert 9 Using Live Imaging to Examine Early Cardiac Development in Zebrafish. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tess McCann, Rabina Shrestha, Alexis Graham, and Joshua Bloomekatz 10 Imaging Planar Cell Polarity Proteins in Xenopus Neuroectoderm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Olga Ossipova, Pamela Mancini, and Sergei Y. Sokol 11 In Ovo Gain- and Loss-of-Function Approaches to Study Gut Morphogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bhargav D. Sanketi and Natasza A. Kurpios 12 Avian Embryos as a Model to Study Vascular Development . . . . . . . . . . . . . . . . . . Bhargav D. Sanketi and Natasza A. Kurpios

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BAC Recombineering and Transgenesis to Study Cell Polarity and Polarized Tissue Morphogenesis in Mice. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Deli Yu and Jianbo Wang Two-Photon Cell and Tissue Level Laser Ablation Methods to Study Morphogenetic Biomechanics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abigail R. Marshall, Eirini Maniou, Dale Moulding, Nicholas D. E. Greene, Andrew J. Copp, and Gabriel L. Galea Protocols for Investigating the Epithelial Properties of Cardiac Progenitor Cells in the Mouse Embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Claudio Cortes, Christopher De Bono, Charlotte Thellier, Alexandre Francou, and Robert G. Kelly Methods to Investigate Cell Polarity of Inner Ear . . . . . . . . . . . . . . . . . . . . . . . . . . . Jihan Lyu, Xiaoqing Qian, Binjun Chen, and Dongdong Ren Characterization of Axon Guidance Phenotypes in Wnt/PCP Mutant Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kathryn M. Miller, Pau Marfull-Oromı´, and Yimin Zou In Vitro Explant Assays and Cultures to Study PCP Signaling in Axon Guidance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pau Marfull-Oromı´, Kathryn M. Miller, and Yimin Zou Biochemical and Cellular Assays to Study Mechanisms of PCP Signaling in Axon Guidance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pau Marfull-Oromı´, Kathryn M. Miller, and Yimin Zou Dissection, Fixation, and Immunostaining of the Drosophila Midgut. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jia Chen and Daniel St Johnston In Vivo Analysis of Pathways Regulating Epithelial Polarity and Secretion Using Drosophila Salivary Glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . Johanna Lattner, Marko Brankatschk, and David Flores-Benitez Imaging Epidermal Cell Rearrangement in the C. elegans Embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jeff Hardin, Joel Serre, Ryan King, Elise Walck-Shannon, and David Reiner Methods for the Study of Apical Constriction During Ascidian Gastrulation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ulla-Maj Fiu´za and Patrick Lemaire Assays for Apical Constriction Using the Xenopus Model . . . . . . . . . . . . . . . . . . . . Austin T. Baldwin, Ivan K. Popov, John B. Wallingford, and Chenbei Chang The Use of Three-Dimensional Cell Culture to Study Apicobasal Polarization and Lumen Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ´ lvaro Roma´n-Ferna´ndez, Emma Sandilands, and David M. Bryant A Studying Cell Polarity Dynamics During Cancer Initiation Using Inducible 3D Organotypic Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rachel Catterall, Reem Kurdieh, and Luke McCaffrey

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Under-Agarose Chemotaxis and Migration Assays for Dictyostelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shashi Prakash Singh and Robert H. Insall Mapping Asymmetry in Collective Cell Migration: Lessons from Border Cells in Drosophila Oogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martina Felix and Mohit Prasad A Toolbox to Study Tissue Mechanics In Vivo and Ex Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sofia Moreira, Jaime A. Espina, Joana E. Saraiva, and Elias H. Barriga Quantitative Analysis of Directional Neural Crest Cell Migration . . . . . . . . . . . . . Shuyi Nie

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

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Contributors ABDELHALIM AZZI • Cutaneous Biology Research Center, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA BRYCE T. BAJAR • Department of Biological Chemistry, Medical Scientist Training Program, David Geffen School of Medicine, University of California, Los Angeles, Los Angeles, CA, USA AUSTIN T. BALDWIN • Department of Molecular Biosciences, University of Texas at Austin, Austin, TX, USA ELIAS H. BARRIGA • Mechanisms of Morphogenesis Lab, Gulbenkian Institute of Science (IGC), Oeiras, Portugal MARK L. BERNS • Department of Biology, College of Science and Technology, Temple University, Philadelphia, PA, USA JOSHUA BLOOMEKATZ • Department of Biology, University of Mississippi, University, MS, USA MARKO BRANKATSCHK • The Biotechnological Center of the TU Dresden (BIOTEC), Dresden, Germany DAVID M. BRYANT • Institute of Cancer Sciences, University of Glasgow, Glasgow, UK; The CRUK Beatson Institute, Glasgow, UK RACHEL CATTERALL • Rosalind and Morris Goodman Cancer Institute, McGill University, Montreal, QC, Canada; Division of Experimental Medicine, Department of Medicine, McGill University, Montreal, QC, Canada CHENBEI CHANG • Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA BAOEN CHEN • Cutaneous Biology Research Center, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA BINJUN CHEN • ENT Institute and Department of Otorhinolaryngology, Eye & ENT Hospital, Fudan University, Shanghai, China; NHC Key Laboratory of Hearing Medicine (Fudan University), Shanghai, China JIA CHEN • The Gurdon Institute and the Department of Genetics, University of Cambridge, Cambridge, UK JUN CHU • Guangdong Provincial Key Laboratory of Biomedical Optical Imaging Technology & Center for Biomedical Optics and Molecular Imaging, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China ILYA CHUYKIN • Department of Cell, Developmental and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA ANDREW J. COPP • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK CLAUDIO CORTES • Aix-Marseille Universite´, CNRS UMR 7288, IBDM, Marseille, France CHRISTOPHER DE BONO • Aix-Marseille Universite´, CNRS UMR 7288, IBDM, Marseille, France; Department of Genetics, Albert Einstein College of Medicine, Bronx, NY, USA DANIEL J. DICKINSON • Department of Molecular Biosciences, University of Texas at Austin, Austin, TX, USA HANNAH ERB • Cutaneous Biology Research Center, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA

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JAIME A. ESPINA • Mechanisms of Morphogenesis Lab, Gulbenkian Institute of Science (IGC), Oeiras, Portugal MARTINA FELIX • Department of Biological Sciences, Indian Institute of Science Education and Research—Kolkata, Nadia, West Bengal, India ULLA-MAJ FIU´ZA • CRBM, University of Montpellier, CNRS, Montpellier, France DAVID FLORES-BENITEZ • The Biotechnological Center of the TU Dresden (BIOTEC), Dresden, Germany ALEXANDRE FRANCOU • Aix-Marseille Universite´, CNRS UMR 7288, IBDM, Marseille, France; Memorial Sloan Kettering Cancer Center, SKI, Developmental Biology Department, New York, NY, USA GABRIEL L. GALEA • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK; Comparative Bioveterinary Sciences, Royal Veterinary College, London, UK; Birth Defects Research Centre, UCL GOS ICH, London, UK ALEXIS GRAHAM • Department of Biology, University of Mississippi, University, MS, USA NICHOLAS D. E. GREENE • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK XINMENG GUAN • Guangdong Provincial Key Laboratory of Biomedical Optical Imaging Technology & Center for Biomedical Optics and Molecular Imaging, Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China CARLA GUARINO • Cutaneous Biology Research Center, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA RAYMOND HABAS • Department of Biology, College of Science and Technology, Temple University, Philadelphia, PA, USA JEFF HARDIN • Department of Integrative Biology, University of Wisconsin-Madison, Madison, WI, USA ROBERT H. INSALL • CRUK Beatson Institute, Glasgow, UK; Institute of Cancer Sciences, University of Glasgow, Glasgow, UK DANIEL ST JOHNSTON • The Gurdon Institute and the Department of Genetics, University of Cambridge, Cambridge, UK ROBERT G. KELLY • Aix-Marseille Universite´, CNRS UMR 7288, IBDM, Marseille, France RYAN KING • Department of Biology, St. Norbert College, De Pere, WI, USA REEM KURDIEH • Rosalind and Morris Goodman Cancer Institute, McGill University, Montreal, QC, Canada; Division of Experimental Medicine, Department of Medicine, McGill University, Montreal, QC, Canada NATASZA A. KURPIOS • Department of Molecular Medicine, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA AMY LAM • Departments of Neurobiology and Bioengineering, Stanford University, Stanford, CA, USA JOHANNA LATTNER • Max-Planck Institute of Molecular Cell Biology and Genetics (MPICBG), Dresden, Germany TAL LAVIV • Neuronal Signal Transduction Group, Max Planck Florida Institute for Neuroscience, Jupiter, FL, USA; Department of Physiology and Pharmacology, Sackler School of Medicine and Sagol School of Neuroscience, Tel Aviv University, Tel Aviv, Israel PATRICK LEMAIRE • CRBM, University of Montpellier, CNRS, Montpellier, France MICHAEL Z. LIN • Departments of Neurobiology and Bioengineering, Stanford University, Stanford, CA, USA

Contributors

xiii

JIHAN LYU • ENT institute and Department of Otorhinolaryngology, Eye & ENT Hospital, Fudan University, Shanghai, China; NHC Key Laboratory of Hearing Medicine (Fudan University), Shanghai, China PAMELA MANCINI • Department of Cell, Developmental and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA EIRINI MANIOU • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK PAU MARFULL-OROMI´ • Neurobiology Section, Biological Sciences Division, University of California San Diego, La Jolla, CA, USA ABIGAIL R. MARSHALL • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK ALEXA L. MATTHEYSES • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA LUKE MCCAFFREY • Rosalind and Morris Goodman Cancer Institute, McGill University, Montreal, QC, Canada; Gerald Bronfman Department of Oncology, McGill University, Montreal, QC, Canada; Division of Experimental Medicine, Department of Medicine, McGill University, Montreal, QC, Canada; Department of Biochemistry, McGill University, Montreal, QC, Canada TESS MCCANN • Department of Biology, University of Mississippi, University, MS, USA KATHRYN M. MILLER • Neurobiology Section, Biological Sciences Division, University of California San Diego, La Jolla, CA, USA SOFIA MOREIRA • Mechanisms of Morphogenesis Lab, Gulbenkian Institute of Science (IGC), Oeiras, Portugal DALE MOULDING • Developmental Biology and Cancer, UCL GOS Institute of Child Health, London, UK TOMASZ J. NAWARA • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA SHUYI NIE • School of Biological Sciences, Georgia Institute of Technology, Atlanta, GA, USA; Petit Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, GA, USA OLGA OSSIPOVA • Department of Cell, Developmental and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA IVAN K. POPOV • Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA MOHIT PRASAD • Department of Biological Sciences, Indian Institute of Science Education and Research—Kolkata, Nadia, West Bengal, India XIAOQING QIAN • ENT institute and Department of Otorhinolaryngology, Eye & ENT Hospital, Fudan University, Shanghai, China; NHC Key Laboratory of Hearing Medicine (Fudan University), Shanghai, China TEJESHWAR C. RAO • Department of Cell, Developmental, and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA DAVID REINER • Texas A&M Health Science Center, Houston, TX, USA DONGDONG REN • ENT institute and Department of Otorhinolaryngology, Eye & ENT Hospital, Fudan University, Shanghai, China; NHC Key Laboratory of Hearing Medicine (Fudan University), Shanghai, China ´ LVARO ROMA´N-FERNA´NDEZ • Institute of Cancer Sciences, University of Glasgow, Glasgow, A UK; The CRUK Beatson Institute, Glasgow, UK

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Contributors

EMMA SANDILANDS • Institute of Cancer Sciences, University of Glasgow, Glasgow, UK; The CRUK Beatson Institute, Glasgow, UK BHARGAV D. SANKETI • Department of Molecular Medicine, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA JOANA E. SARAIVA • Mechanisms of Morphogenesis Lab, Gulbenkian Institute of Science (IGC), Oeiras, Portugal JOEL SERRE • Program in Genetics, University of Wisconsin-Madison, Madison, WI, USA RABINA SHRESTHA • Department of Biology, University of Mississippi, University, MS, USA SHASHI PRAKASH SINGH • CRUK Beatson Institute, Glasgow, UK SERGEI Y. SOKOL • Department of Cell, Developmental and Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA POOJA R. SONAVANE • Department of Cellular & Molecular Medicine, University of California San Diego, La Jolla, CA, USA NAOMI STOLPNER • Department of Molecular Biosciences, University of Texas at Austin, Austin, TX, USA DAVID STRUTT • School of Biosciences, University of Sheffield, Sheffield, UK HELEN STRUTT • School of Biosciences, University of Sheffield, Sheffield, UK CHARLOTTE THELLIER • Aix-Marseille Universite´, CNRS UMR 7288, IBDM, Marseille, France ELISE WALCK-SHANNON • Department of Biology, Washington University in St. Louis, St. Louis, MO, USA JOHN B. WALLINGFORD • Department of Molecular Biosciences, University of Texas at Austin, Austin, TX, USA JIANBO WANG • Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA SAMANTHA J. WARRINGTON • School of Biosciences, University of Sheffield, Sheffield, UK KARL WILLERT • Department of Cellular & Molecular Medicine, University of California San Diego, La Jolla, CA, USA XU WU • Cutaneous Biology Research Center, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA RYOHEI YASUDA • Neuronal Signal Transduction Group, Max Planck Florida Institute for Neuroscience, Jupiter, FL, USA DELI YU • Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL, USA YIMIN ZOU • Neurobiology Section, Biological Sciences Division, University of California San Diego, La Jolla, CA, USA

Chapter 1 Use of Fluorescence Recovery After Photobleaching (FRAP) to Measure In Vivo Dynamics of Cell Junction–Associated Polarity Proteins Samantha J. Warrington, Helen Strutt, and David Strutt Abstract Here, we present a detailed protocol for fluorescence recovery after photobleaching (FRAP) to measure the dynamics of junctional populations of proteins in living tissue. Specifically, we describe how to perform FRAP in Drosophila pupal wings on fluorescently tagged core planar polarity proteins, which exhibit relatively slow junctional turnover. We provide a step-by-step practical guide to performing FRAP, and list a series of controls and optimizations to do before conducting a FRAP experiment. Finally, we describe how to present the FRAP data for publication. Key words FRAP, In vivo imaging, Live-imaging, Fluorescence, Photobleaching, Drosophila, Pupal wing, Planar polarity, Planar cell polarity, PCP

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Introduction Fluorescence recovery after photobleaching (FRAP) is a commonly used live-imaging technique for measuring protein dynamics and turnover within living cells and tissues. To permit visualization of protein turnover, a fluorescent tag is attached to the protein of interest, this most commonly being achieved via a genetically encoded fusion to a fluorescent protein such as EGFP. A proportion of the fluorescent protein present within a region of interest (ROI) is irreversibly bleached using high-intensity illumination, and the ROI is imaged over time to reveal the recovery of the fluorescence intensity within the region compared to the same region before bleaching (Fig. 1a, b). A crucial conceptual point to note is that the bleaching of the fluorophore does not result in the tagged protein being removed from the bleached region. Instead the protein is still present and subjected to its normal rates of diffusion, protein

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_1, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Principles of FRAP in cells and tissues. (a) Diagram of a simple FRAP experiment, showing a region within a cell that is selected for bleaching (blue), a control region (orange) for correcting for acquisition bleaching during imaging and a background region outside of the cell (black). (b) Diagram of a FRAP recovery curve showing the initial bleach time (blue arrow) and the mobile and immobile protein fractions. (c) An example of in vivo FRAP on the junction between cells in an epithelium (blue). A similar junction can be used as the control region (orange) for measuring acquisition bleaching. The background value can be difficult to obtain, if the tissue fills the whole field of view, so “laser off” background is measured. The two cells affected by the bleach ROI are shown in green. (d-d0 ) Representation of which part of the fluorescent sample is bleached in the XY plane (d) and in the XZ plane (d0 ). The location of the protein within the cell membrane should be considered before designing the experiment; (e) shows a situation where the tagged protein is spread along the cell membrane in the Z plane and some of it is not included within the bleach region. Consequently, the unbleached population could contribute to a faster rate of recovery than anticipated, compared to a tagged junctional protein that was completely bleached

exchange and degradation. It is this dynamic exchange of bleached for unbleached protein which is measured in FRAP experiments (for general reviews see, e.g., [1–7]). Protein exchange within a cell can occur via several different processes including rapid ones such as local diffusion and slower ones such as endocytosis and motor-mediated transport. Many of these exchange methods occur simultaneously, collectively accounting for the recovery of the signal in the bleached region, with the contributions of the different processes varying depending on the properties of the protein [5, 8, 9].

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Analysis of FRAP data is conducted by plotting the normalized ROI fluorescence intensity over time (Fig. 1b). The data can be explored further to identify the different components that contribute to protein exchange, by fitting a curve using an appropriate mathematical model. The most common curves fitted are a one-phase or two-phase exponential recovery [10, 11]. However, note there is no consensus as to which curve is best for a given situation, so ideally several should be tried and the results compared [3, 8–10, 12]. Curve fitting allows the calculation of the rate of recovery and the curve plateau. From the latter it is possible to determine the size of the mobile fraction (the fraction of fluorescence intensity that has recovered, indicating protein turnover) and the immobile fraction (the fraction that has not recovered). FRAP measurements to study protein diffusion were initially performed in the 1970s to look at protein diffusion (e.g., [1, 11]). Since then the method has been refined and adapted as technology has advanced, enabling more information to be extracted and more weakly expressed proteins to be studied. FRAP has become a standard method to analyze protein dynamics within cells and tissues, but drawing accurate conclusions from the data requires knowledge of appropriate controls and data analysis that cannot always be acquired from published experimental work. In this chapter we present an in-depth method for performing FRAP and interpreting the resulting data for junctionally localized proteins in the Drosophila pupal wing, using the core planar polarity proteins as an example. We present the method we have used to study the dynamics of clusters of protein complexes using “puncta” FRAP [13], “junctional” FRAP to look at dynamics on a junction between a pair of cells [14] and “hub and spoke” FRAP for studying whole-cell junctional turnover [15]. However, the method is applicable for use with other proteins that are localized to, or very near to, the cell junctions in Drosophila and other in vivo tissues. Special consideration is given to setting up the imaging conditions and the controls required to ensure the reliability of the data collected. 1.1 The Use of Fluorescent Proteins in FRAP

The easiest way to attach a fluorophore to a protein of interest is via fusion to a genetically encoded fluorescent protein. To avoid artefacts in measuring fluorescence recovery the ideal fluorescent protein should be (1) bright enough to achieve a good signal-to-noise ratio, (2) photostable and not prone to photo-reversible bleaching [16], and (3) monomeric to avoid dimerization of the fluorophore itself adversely affecting protein exchange [17]. The fluorescent protein EGFP is most commonly used [1], although there are published examples using RFP derived fluorescent proteins such as mCherry [18–21], RFP [22], and other GFP derivatives such as Venus [23, 24]. However, some fluorescent proteins used for liveimaging are often unsuitable for FRAP as they photoconvert or reversibly bleach [25, 26].

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The fluorescent tag can be attached to any part of the protein as long as it does not affect its function. Attaching the tag to either the N- or C-terminus of the protein is the simplest strategy, although tagging at either location can affect the protein’s properties. If possible, the protein should be independently tagged at both ends and assays performed to check that the function and cellular localization is like that of the endogenous protein. Note, however, that attaching a 27 kDa fluorescent tag to a protein of interest will inevitably lead to changes, however subtle, in protein dynamics. Ideally the fluorescent tag should be introduced into the endogenous gene locus, to allow the protein product to be expressed as at close to normal levels as possible, and checks should be performed to verify this. The intensity of fluorescent proteins can vary depending on the pH of the environment. This is important as the pH of intercellular vesicles such as lysosomes, endosomes, and Golgi compartments can be as low as 4.6 [27]. GFP fluorescence is reduced to 50% at less than pH 5.5 [16, 28], so GFP variants may not be a suitable marker for proteins that localize to regions of low pH. As different fluorescent proteins exhibit different cellular behaviors, only FRAP data obtained using the same fluorescent tag should be compared. 1.2 FRAP on In Vivo Tissue Samples

FRAP has been used extensively on in vitro tissue culture and ex vivo samples. However, studies are increasingly performing FRAP on in vivo samples, for example to examine the role of various pathways in setting up cell polarity in Drosophila [13, 29–31]. As with all in vivo imaging approaches, FRAP on live samples is more challenging than the equivalent experiment on cultured cells. However, the benefits of doing FRAP in vivo is that the protein is in its natural context and is subjected to all the normal influences it would receive during the timecourse of the experiment. Key considerations when undertaking an in vivo FRAP experiment include the following. 1. Being able to image the sample for a sufficiently long period under normal physiological conditions, while keeping sample movement to a minimum (for instance by being suitably adhered to the mounting dish). Ideally the recovery of the fluorescence should be observed until it reaches a plateau with no more recovery observed, otherwise the rate of recovery and size of the mobile and immobile fractions cannot be accurately calculated. 2. In vivo samples often have low fluorescence intensity and poor signal-to-noise ratio. This can cause imaging problems and is often due to low endogenous protein expression or imaging at an increased distance from the coverslip. Both of these can lead to an increase in acquisition bleaching during imaging as the laser power is increased to increase the detection of the fluorescence signal.

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3. The three-dimensional nature of tissues means that fluorescence bleaching with a standard laser occurs in the Z plane as well as the XY plane (Fig. 1d, e). This can mean for example that the degree of bleaching of a population of a fluorescent protein localized to the apical junctions will be different compared to that of a lateral membrane localized protein that is present in deeper Z planes (Fig. 1d, e), which in turn can lead to differences in the rate and amount of recovery observed. However, the spreading of the bleaching within the Z plane can be minimized if a 2-photon laser is used [12, 32, 33]. 4. Bleaching of a large proportion of the cellular population of a fluorescent protein will result in changes in the resulting perceived protein dynamics. The analysis of FRAP data assumes that there is an infinitely large pool of unbleached protein for use in fluorescence recovery. Therefore, avoid bleaching all of the fluorescent protein within a cell, and only compare results from experiments where similar degrees of bleaching are used. 1.3 FRAP on Planar Polarity Proteins

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Planar polarity is essential to coordinate the formation of an axis of polarity perpendicular to the apical–basal axis of epithelial cells. The proteins required to establish and maintain planar polarity accumulate into membrane domains at the apical cell junctions, forming asymmetric complexes spanning the cell junctions between neighboring cells. There are two main pathways involved in setting up planar polarity in animal tissues (reviewed in [34–36]). In Drosophila, the “Fat-Dachsous pathway” consists primarily of the atypical cadherins Fat and Dachsous, while the “core pathway” is composed of the transmembrane proteins Frizzled, Strabismus, and the atypical cadherin Flamingo, as well as the cytoplasmic proteins Prickle, Dishevelled, and Diego. FRAP has been used extensively to understand core pathway protein dynamics in Drosophila [13–15, 37–42], in mice [43, 44] and also in Xenopus [45–48]. Fat and Dachsous dynamics have also been studied in flies [49] and in mammalian tissue culture [50].

Materials

2.1 Materials for Mounting Live Pupae for Imaging of Pupal Wings

1. Dissecting microscope with light. 2. Glass bottom imaging dishes (35 mm). 3. 10 cm Petri dishes. 4. 2 pairs of dissection forceps (size 4 or 5 is preferred) one pair sharper than the other. 5. Double sided tape cut into 4–5 cm strips. 6. Plastic Pasteur pipettes. 7. 200 μl pipettor and tips.

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8. Razor blades. 9. double distilled (dd) H2O. 10. Halocarbon 700 oil (mounting method 1 only). 11. Heptane glue (mounting method 2 only). To make, place sticky tape (Sellotape) in a glass jar with 10 ml of heptane. Shake and leave to settle for a few hours. The heptane should be yellow in colour and slightly thickened but still liquid. It can be kept for a few months in the sealed jar and topped up with more heptane and tape if required. 2.2 Microscope Equipment

1. A fluorescence, laser scanning confocal (e.g., Nikon A1 with gallium arsenide phosphide (GaAsP) detectors). 2. A high power objective lens (e.g., 60 plan Apochromat, NA 1.4). 3. Lasers and filters for imaging your fluorophore of interest, for example, for imaging EGFP we use a 488 nm diode laser and a 525–550 nm Band Pass filter or a GFP long pass filter. 4. Immersion oil.

2.3 Software for Analysis of FRAP Raw Data

To analyze the raw data various software packages can be used. We list below the software we currently use but any software that can perform the tasks could be used instead and the process could be simplified by coding it on a platform such as R or MATLAB to reduce the time spent on analysis. We recommend using the following. 1. ImageJ for collecting the raw mean intensity values from the images. 2. Microsoft Excel (or other spreadsheet software) to collect together the raw data and process it. 3. Prism (version 8, http://www.graphpad.com) to plot the FRAP graphs and fit exponential equations 4. G*Power to calculate the Power of the experiment and the sample size required.

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Methods This FRAP method is optimized for Drosophila pupal wings.

3.1 Collecting and Staging Drosophila melanogaster White Prepupae

1. For staging, collect Drosophila white prepupae with a paintbrush or forceps from the wall of the fly bottle and transfer to the wall of a fresh vial containing fly food so they stay slightly damp (Fig. 2a).

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Fig. 2 Drosophila pupal wing dissection and mounting methods (a) Collect the pupae and place in a moist environment. A husbandry vial filled with fly food is easiest. (b) Move the pupae using forceps onto double sided tape placed inside the lid of a petri dish. Line them up anterior away and ventral up. (c) Cut the pupa along the midline. Start posterior to the notum and stop just anterior to the end of the wing. Peel back the cuticle using forceps. (d) An example of an exposed region of pupa (outlined by a dotted line) and the wing outlined in blue, anterior left and posterior right. The anterior spiracles and larval mouth parts are also

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2. Note down the time and genotype of the flies, and age the prepupae until the desired age. Live-imaging and FRAP can be easily performed on pupal wings aged for 20 h at 25  C after puparium formation (APF) onward (see Note 1). 3. Pupae can be imaged for extended times over many hours and should not dry out or die (see Note 2, Fig. 2h). 3.2 Exposing the Drosophila Pupal Wing for Imaging

1. Cut a 5 cm long piece of double-sided tape, stick it to the inside of a lid of a 10 cm Petri dish and remove the double-sided tape backing. 2. Pick up pupae by the anterior spiracles, using the blunter pair of forceps and being careful not to induce damage. Place the pupae in the Petri dish lid. Then position the pupae on the tape in a row with their anterior ends pointing away from you and the ventral sides facing up (see Note 3, Fig. 2b). 3. Using the corner of a razor blade make a cut in the puparium cuticle along the midline approximately 1/3 of the way along the pupa, starting where the notum ends and the wing hinge starts. Continue the cut along the midline between the legs and stop just before the wing ends (see Note 4, Fig. 2c). 4. To remove a square of cuticle from over the wing, slide the sharper pair of forceps in between the puparium cuticle case and the pupa. Then take hold of the cuticle case and pull toward the tape, uncovering the wing, and then pull the cuticle posteriorly to remove the square of pupal casing (Fig. 2d). 5. Dissect 2–5 pupae in succession before releasing them from the tape. 6. To remove the pupae from the tape, use a Pasteur pipette to place a bead of ddH2O next to the pupae on the opposite side to the hole in the puparium case. Do not pull the pupae off the tape, but wait until the pupae are released from the tape by the action of the water and then remove by taking hold of an anterior spiracle and transfer to the imaging dish (Fig. 2e).

ä Fig. 2 (continued) indicated. (e) Transfer the pupae to the edge of the imaging dish. (f) For mounting using mounting method 1: place the pupae on the dish, wing up, and place a small blob of Halocarbon oil on the surface of each wing. Rotate the pupae until the oil and wing are facing the dish. For mounting method 2: make sure the dish is coated with heptane glue, pick up the pupa with the wing facing down and place directly onto the imaging dish, wing facing down. Align pupae in a vertical row to make imaging easier. (g) Turn over the dish to observe the pupae from underneath. Wings should be stuck to the imaging dish and should be flat. (h) After imaging, age the pupae for a few days to ensure they develop normally. (i, j) Examples of pupae mounted with mounting method 1, anterior to the right. Note the oil meniscus around the pupa. (k, l) Examples of pupae mounted with mounting method 2, anterior to the right. Note there is no oil around the pupa, instead gently press on the pupa to get it to stick to the heptane glue

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7. At this point the pupae are wet and need to be dried slightly before mounting (see Note 5), by placing all the dissected pupae around the edge of the imaging dish for a few minutes (Fig. 2e). 3.3 Mounting the Drosophila Pupae on an Imaging Dish

3.3.1 Mounting Method 1

The mounting methods here could be applied to other fly tissues, particularly if using mounting method 2. Here we describe two methods that we use routinely to mount Drosophila pupae on a glass bottom dish to be imaged on an inverted microscope (see Note 6). 1. Move the dissected pupae to the center of the imaging dish and align in a vertical row with the hole in the cuticle facing up. 2. Cover each exposed pupal wing with a small amount of halocarbon oil. To do this, dip a 200 μl pipettor tip into a 1.5 ml microfuge tube containing the halocarbon oil. Only dip in the very end of the tip and avoid picking up a large amount of oil. 3. Carefully touch the wing with the oily tip (see Notes 7 and 8). Then, use two pairs of forceps to roll the pupa until the exposed wing is face down (Fig. 2f). Press the pupa down firmly. Do this to all of the pupae you have transferred into the imaging dish. Check the orientation of the wings/pupae by turning over the dish and observing down the dissecting microscope (the wings remain stuck to the dish when turned over, Fig. 2g). The wings should be flat against the dish surrounded by oil (see Notes 8 and 9, Fig. 2i–j).

3.3.2 Mounting Method 2

1. Prior to dissecting the pupae coat the imaging dish in a fine coating of heptane glue. Use a Pasteur pipette to transfer 500 μl of very runny heptane glue into the imaging dish, spread it around to cover the glass coverslip then remove as much as possible with a 200 μl pipettor or disposable Pasteur (see Note 10). 2. Allow it to dry for 10 min or until tacky. This can be tested by touching a pipette tip to the surface and on removing listening for a sticky noise. 3. Once coated, the dishes can be stored for several weeks, and after imaging can be cleaned with 70% ethanol, recoated and reused. 4. To mount the dissected pupae, use a pair of forceps to pick up a pupa by its spiracles, with the open window of cuticle facing down. Place the pupa wing side down onto the heptane glue coated dish and press (although not too firmly) to adhere the pupa to the glue (Fig. 2f, Note 11). Check the orientation of the wings/pupae by turning over the dish and observing down the dissecting microscope (the wings remain stuck to the dish when turned over, Fig. 2g, k, l).

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3.4 Live-Imaging Settings

The aim of fluorescence imaging over time is to capture the most representative image possible and is a trade-off between achieving maximum resolution while minimizing acquisition bleaching and/or photodamage. To do a successful FRAP experiment, live-imaging conditions should be defined that minimize bleaching during imaging (to, e.g., less than 20%) while providing sufficient imaging resolution, over a long enough time frame to obtain a useable fluorescence recovery curve that either recovers to 100% of initial fluorescence or reaches a plateau. Therefore, there will be a degree of parameter testing for live-imaging and FRAP before a proper experiment can be performed. Initial starting points for settings to live-image junctionally associated polarity proteins can be found in the literature, for example the live-imaging and FRAP settings for core planar polarity proteins [13, 37], and for the Fat and Dachsous cadherins [49]. There are several parameters that can be varied: 1. The image capture settings. These should be set to be able to see signal above the noise with minimal bleaching at a sufficient resolution. These include the laser power and detector gain, and also the pixel size, pixel dwell time and scan area. 2. The interval between images. This can only be determined once the rate of recovery during FRAP has been observed, to avoid missing an initial fast phase of recovery. Hence, initial time intervals should be chosen to minimize acquisition bleaching over the timecourse of the experiment, and altered if necessary once the recovery has been observed. For proteins with a fast turnover, fluorescence will recover quickly so must be sampled with short time intervals. Failing to capture the initial fast recovery phase will result in inaccurate curve fitting and calculation of the rate of recovery (see Note 12). Time intervals of post-bleach images can be kept constant or can be varied to allow the capture of the initial faster phase and then slower intervals as a recovery plateau is approached (see Note 13). For the core planar polarity proteins, we use variable intervals: 5 images  5 s, 10 images  10 s, 10 images  15 s and 8 images  30 s, although others use uniform 5 s or 10 s intervals [13, 15, 40]. 3. The time of the experiment. Imaging should be continued for 2–3 times longer than the time taken to achieve the plateau for the control condition. This should ensure that mutant or experimental conditions can be imaged using the same settings, even if they take longer for recovery to reach a plateau. To check that the settings selected limit any acquisition bleaching, take a time series of images with a particular time interval and image capture settings. Measure the mean intensity of the whole

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image for the start and end time points and calculate the percentage change in intensity (see Subheading 3.7.1, steps 1–5 for instructions for measuring intensity with ImageJ). Below we give detailed parameter settings for live-imaging and FRAP of core planar polarity proteins in the Drosophila pupal wing using mounting method 1 and imaged on a Nikon A1 microscope: 1. Objective: 60 1.4NA (see Note 14). 2. Pinhole setting: Use the manufacturer’s recommended pinhole size for optimal spatial resolution. For a Nikon A1 microscope, this is 1.2 AU (35.8 μm) (see Note 15). 3. Pixel size: 100 nm (see Note 16). 4. Scan region: 256  256 pixels (see Note 17). 5. Zoom factor: this is interrelated with the pixel size and scan region. On our system a 100 nm pixel size and scan region of 256  256 pixels using the 60 objective is equivalent to a zoom factor of 8 (see Note 18). 6. Frames per second: 2 frames per second (see Note 19). 7. Z sections: A single Z section is captured, to minimize acquisition bleaching. 8. Averaging of images: No averaging (see Note 20). 9. Laser power: For EGFP imaging a 488 nm laser is used. We use a Nikon A1 with a GaAsP detector, a 525/550 filter cube dichroic and 525–550 nm bandpass filter. Our 488 nm laser at 0.8% laser power gives a power of ~12 μW at the lens. Using these settings, we can image EGFP-tagged core proteins in live pupal wings with negligible acquisition bleaching for 10 min. 10. Gain: A gain of 80–100 Hv is sufficient in our hands to image planar polarity proteins fused to EGFP (see Notes 21–23). 11. Offset: This is used to specify the minimum cutoff point for the fluorescence intensity detected. If the offset is increased, then more noise and signal is removed. We keep the offset at 0, to capture as much signal as possible (see Note 23). 12. Time of experiment: We image Frizzled, Strabismus and Flamingo transmembrane proteins tagged with EGFP for 8 min. This is ~2–3 times longer than the time taken to reach the plateau in control conditions. 3.5 Optimizing the FRAP Imaging Method

There are three steps to a FRAP experiment: 1. Selecting the bleach ROIs and collecting pre-bleach images by imaging the sample with low illumination (control ROIs are selected postprocessing). 2. Bleaching the illumination.

bleach

ROIs

with

high-intensity

laser

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3. Monitoring the ROIs over time using the low illumination settings described in Subheading 3.4. 3.5.1 Choosing ROI Size, Shape, and Number

The essential point to note when choosing a bleach ROI is to not bleach all of the fluorescent protein within a cell or even within an entire cell membrane. Otherwise the pool of unbleached fluorescent protein available to recover from will be limited, and this will affect the FRAP results. In addition, when using FRAP on membrane proteins the bleach ROI should not cover a large proportion of the cytoplasm as this will bleach a large part of the recovery pool of protein and inhibit fluorescence recovery. The ROI should consist of at least tens of pixels, to allow it to be tracked over time, and allow a mean intensity value to be measured that is not influenced by excessive noise [2]. The core planar polarity and Fat and Dachsous proteins are not uniformly localized to the cell junctions and instead are asymmetrically localized into junctional puncta of varying sizes, the largest being approximately 200 nm in diameter (Fig. 3a and [13–15, 49]). Therefore, we have used different shaped bleach regions to sample the dynamics of different proteins populations within the cell junctions. For the core proteins we used small oval regions (2 μm2) to look at protein dynamics within a punctum [13], larger membrane regions (3–5 μm2 oval regions) to study the dynamics within a particular cell junction [14, 40] and “hub and spoke” regions to try to sample the entire junctional population of a cell while not bleaching large amounts of the cytoplasm (Fig. 3b and [15]). Others have also used FRAP on small membrane regions to determine local protein stability [30, 51–53]. For proteins that are spread throughout the membrane rather than in discrete domains, the proportional contributions of protein dynamics (i.e., local diffusion and intracellular transport) can be further elucidated by bleaching the fluorescence of different sized regions [54]. Common confocals enable the user to select multiple ROI regions. For FRAP on pupal wings we select 4 bleach regions in a 256  256 pixel image (see Note 24).

3.5.2 Selecting Control Unbleached Regions and Background Regions

Even if the imaging conditions have been carefully optimized, acquisition bleaching can still occur and must be corrected. Hence, non-bleached regions are selected within the same image and the mean intensity is used to normalize the mean intensity of the bleach regions over time. In a pupal wing we select a minimum of 4 non-bleached regions per image, or the same number as for the bleach ROIs. The background intensity should also be subtracted. Unlike FRAP in cultured cells, in vivo samples often fill the field of view so this measurement cannot easily be directly collected (Fig. 1c). Instead, we take a “laser off” image using the same settings, to allow us to subtract background noise created by the detector.

Fig. 3 Core proteins localized to cell junctions within Drosophila pupal wing cells. (a) Core proteins localize into discrete membrane subdomains described as puncta (white arrowheads). Junctions not containing obvious puncta are indicated by yellow arrowheads. (a0 ) Frizzled-EGFP and (a00 ) Strabismus-EGFP, proximal is to the left, dorsal is up. (b) Examples of different ROI shapes used to study at core polarity proteins dynamics (left to right: puncta, junctional and “hub and spoke”). Smaller ROI regions (~2 μm2) are good for targeting small punctate structures. Larger ROIs are more suited to looking at protein stability on a single junction. The “hub and spoke” region is used to look at general junctional stability in a whole cell, and spoke regions avoid bleaching too much of the surrounding cytoplasm and inhibiting fluorescent protein recovery (note that these irregular shapes cannot be performed on all confocals). (c) A double bleach experiment to ensure that the mobile fraction recovery is not inhibited due to excessive initial bleaching causing crosslinking of the protein. The first bleach FRAP experiment is performed and the data is normalized between 1 (pre-bleach intensity) and 0 (intensity after the initial bleach) and the fluorescence (mobile fraction) is allowed to recover (blue line). Then the FRAP experiment is repeated with the same settings (second bleach) and the mobile fraction is again allowed to recover. As most of the immobile fraction was already bleached in the first bleaching step, the majority of the fluorescence bleached and recovering in the second experiment is only the mobile fraction. Therefore, the recovery should approach 100% of the mobile fraction measured in the first experiment (dashed blue lines between the graphs). As with a normal FRAP experiment it is not essential or desirable for all the fluorescence within a region to be bleached, instead we aim for 50–75% bleaching in each cycle and extrapolate to calculate the effects of 100% bleaching in postprocessing

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3.5.3 Choosing the Number of Pre-bleach Images to Record

A number of pre-bleach images should be taken (e.g., 3) to record a fluorescence intensity baseline (these images can be taken with no time interval) (see Note 25).

3.5.4 Specifying the Scanning Conditions Required for the Initial Bleach

Most confocal microscopes have the ability to separately specify laser settings for bleaching (e.g., laser power, scan rate, and how many times the laser will scan the ROIs) independently of the acquisition settings. The initial bleach parameters chosen should result in a fast bleach time (e.g., 1 s), achieve 50–75% loss of fluorescence and not inhibit fluorescence recovery (see Notes 26 and 27). The initial bleach conditions will need to be defined by trial and error. Try to keep the bleach time length to 1 s by adjusting the scan speed and the number of scans for the initial bleach settings. Often repeated quick scans will bleach faster than one slow scan over the ROI. Then systematically vary the laser power (see Note 28), keeping the scan rate and the number of scans the same, and check the fluorescence that remains after bleaching and the shape of the FRAP recovery curve. Once a set of parameters has been chosen, a double bleach experiment is a good control to check if the initial bleaching negatively affects the recovery due to the bleached protein being unable to leave the ROI due to protein crosslinking (Fig. 3c). Carry out a FRAP experiment (see Subheading 3.6) and allow the fluorescence (the mobile fraction) to recover after the initial bleach. The fraction that does not recover is the immobile fraction. Then bleach the same regions again with the same parameters and allow the fluorescence to recover again (Fig. 3c). As most of the immobile fraction has already been bleached in the first FRAP procedure, the majority of the bleaching in the second experiment will be of the mobile fraction [4]. Consequently, the recovery after the second bleach will be close to 100%. If not, this suggests that photodamage has occurred due to excessive bleaching.

3.5.5 Selecting the Number of Replicate Samples

A question that should be asked at the beginning of a series of FRAP experiments is: what is the minimum sample size required to obtain meaningful statistical data? Sample size (“n”) is deemed to be the number of independent biological replicates, for example, number of animals or tissue samples. This value should be irrespective of the number of technical replicates that are measured per sample. We consider different ROIs in the same time series of the same wing to be technical replicates, and the number of different wings to be the true “n” number (i.e., the biological replicates). A small pilot experiment (e.g., on a scale of 8–10 samples) should be conducted to see how many replicates are required. Establish the Ymax values and then use a Power calculation to establish if the sample size is large enough to determine if the statistical conclusion is acceptable (see Note 29).

Using FRAP to Measure Polarity Protein Dynamics 3.5.6 Summary of FRAP Settings Used for Junctionally Associated Planar Polarity Proteins in the Drosophila Pupal Wing

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The FRAP settings outlined here are suitable for core planar polarity junctional proteins that are tagged with EGFP in their endogenous locus, in Drosophila wing discs and pupal wings. These settings should be used as a guide for imaging on a Nikon A1 with GaAsP detectors and should be further optimized after initial pilot experiments. Only then embark on gathering a large dataset. 1. Bleaching laser power: 80% of the available laser power (see Notes 28 and 30). 2. Laser scan time for bleaching: 8 passes per sec for 1 s to obtain between 50% and 75% bleaching. 3. Time intervals: 3 pre-bleach images, and post-bleach images captured at variable intervals: 5  5 s, 10  10 s, 10  15 s, and 8  30 s. 4. ROI size: A 2 μm2 oval sized region for FRAP of junctional protein clusters in pupal wing cell junctions. 5. ROI number per image: 2–4 bleach regions per image. Once the imaging and FRAP settings are optimized the same settings should be used for consistency, where applicable and possible [54].

3.6 Performing the FRAP Experiment

Most confocals have settings built in to perform FRAP and are similar in setup—see relevant manufacturer’s manuals. All require the ROIs to be drawn on an initial image and specified as “bleach” or “stimulation” regions. We select only the bleach ROIs before imaging, and the non-bleach control ROIs are selected postimaging. In addition, as our samples are prone to movement, we manually track the ROIs over time using ImageJ (see Subheading 3.7.1) and so do not use the microscope software to record the ROI intensities. To begin the FRAP experiment: 1. Place the mounted sample on the microscope and ensure the sample is in focus. The pre-bleach imaging setting will need to be defined, inputting the number of images and the time interval between them (Subheading 3.5.3). Then the bleach settings specifying the number of laser passes, the scan speed and the laser power for the bleach ROIs will have to be selected (Subheading 3.5.4). Finally input the post-bleach image settings, specifying the number of images and the time interval between them (Subheading 3.4). 2. Choose the ROIs (see Subheading 3.5.1). Check that there is enough tissue to select the non-bleach ROIs during postprocessing, and try to choose the same number of non-bleach ROIs as bleach ROIs. Record an image of where the ROIs are located relative to the cells for use in the reselecting of the ROIs during postprocessing; for example, use print screen and save the resulting picture (see Note 31).

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3. Start the FRAP experiment. Check that a loss of fluorescence has occurred post-bleach. If not check the settings and ensure the ROIs are set to bleach/stimulate. 4. At this point it is useful to watch the images over time to ensure the sample does not go out of focus (see Note 32) and adjust the focus if necessary. 5. At the end of image capture make sure the images have been saved. 6. Take a “laser off” background image or other background ROI and save it. 3.7 FRAP Processing Method 3.7.1 FRAP Data Collecting

In FRAP of cultured cells the sample usually does not move, and therefore the data from each ROI can be automatically collected on the confocal during acquisition. However, in tissue FRAP the cells are attached to each other and often shrink and extend their junctions. This makes automated intensity data collection for junctional proteins more challenging as there is no location in the image that is stationary to use as a reference point. Therefore, it is nearly always necessary to manually reselect the ROIs. The reselection of the ROIs is unavoidably inexact, but this is acceptable because the bleached spot achieved is never exactly the same size as the ROI selected due to light diffraction as the region is bleached (see Note 33) and due to protein diffusion during the bleach [55]. Intensity measurements of the bleached and control non-bleached regions can be measured using open source software for example Fiji (Fiji Is Just ImageJ, https://fiji.sc). To process the FRAP data: 1. Open the time series in Fiji (using the Bio-Formats plugin). 2. Reselect one of the bleach ROIs by hand using the freehand selection or oval tools found on the main ImageJ window. 3. Measure the properties of the selected bleach ROI by pressing the “M” shortcut button on the keyboard, measurements will appear in a new results window that should display: Area of the ROI, mean intensity, min and max intensity (these measurements can be selected using Analyze>Set measurements). 4. Then use the “> “button on the keyboard to move to the next image in the time series. 5. If the tissue in the image has moved, reposition the ROI before measuring the intensity. Discard the time series if any images move out of focus or if regions undergo substantial junctional rearrangements (see Note 34). 6. Once all mean intensities for an ROI have been collected, copy and paste the mean intensities into a spreadsheet. 7. Repeat steps 2–6 for all the bleach ROIs.

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8. Select a set of non-bleach ROIs one at a time and repeat the process above to measure their intensities across the whole time series. These intensity measurements can then be pasted into a different region of the spreadsheet. 9. Open the “laser off” background image in ImageJ, select the whole image (Ctrl A on a PC, Command A on a Mac) and measure its mean intensity (by pressing the “M” shortcut). Record this background value in your spreadsheet. 10. Calculate the amount of acquisition bleaching that has occurred by comparing the intensities of the non-bleach ROIs at the start and end of the experiment (see Note 35). This should be no more than 20%. 11. Further calculate the amount of experimental bleaching that has occurred by comparing the intensities of the bleach ROIs between the last pre-bleach image and the first post-bleach image (see Note 36). This should fall between 50% and 75%. 3.7.2 FRAP Data Normalization

FRAP data should be normalized to correct for any acquisition bleaching that occurs during imaging. In addition, the data is rescaled, so the first post-bleach intensity is set to 0 and the pre-bleach intensity at 1. Perform the normalization either in the existing spreadsheet or in a new sheet in the same workbook, see Fig. 4 for an example layout. Fill in the raw data section (left panel) with the intensities that have been collected for each of the bleach ROIs and the average intensity of the non-bleach ROIs for each imaging timepoint. Average the intensities of the three pre-bleach images for each ROI (blue row in left panel). Also add the “laser off” background measurement and the recorded time values. The normalization is conducted in two steps: 1. The first step is to adjust the bleach ROIs for any acquisition bleaching that has occurred (Fig. 4, middle panel), which can be done with the equation:
I ½non  bleachROI Prebleach  I ½background 
I ½non  bleach ROI n  I ½background 
 I ½bleach ROI n  I ½background  where I[bleach ROI]n is the intensity of the bleached ROI at timepoint n, I[non-bleach ROI]n is the intensity of the non-bleach ROI at timepoint n and I[non-bleach ROI]Pre-bleach is the intensity of a non-bleach ROI averaged for the three pre-bleach timepoints. I[background] is the laser off background intensity.

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Fig. 4 An example of a sheet for normalizing the FRAP raw data values. The sheet is divided into three sections. The left section is the raw data imported from other sheets in the workbook and includes the absolute time in seconds, the background intensity value, the bleach ROIs raw data (orange columns) and an average of the non-bleach control ROIs (cyan column). The middle panel normalizes the data for any loss of fluorescence in each ROI due to acquisition bleaching. The right panel rescales the data to set the first postbleach intensity value at 0 (yellow highlighted cells) and the initial pre-bleach values at 1. The average of these normalized regions (cyan column) and the average standard deviations of the regions (blue column) are also shown. The blue-colored cells at the top of each column are the average intensity of the three pre-bleach values for each ROI

2. Secondly the data is rescaled to take into account 50–25% of the initial fluorescence still being present (i.e., the first postbleach value is not 0). The remaining fluorescence intensity in the first post-bleach image of each bleach ROI is subtracted from all the timepoints for that bleach ROI such that the first post-bleach ROI intensity becomes 0. The data are then normalized so that the average of the pre-bleach values is set at 1 (Fig. 4 right panel).
I ½bleach ROI n  I ½bleach ROI 0
I ½bleach ROI Prebleach  I ½bleach ROI 0 where I[bleach ROI]n is the intensity of the bleached ROI at timepoint n, I[bleach ROI]0 is the intensity of the first timepoint after the bleach, and I[bleach ROI]Pre-bleach is the intensity of the bleach ROI averaged for the three pre-bleach timepoints. 3. Calculate the average of the bleach ROIs for each time point (Fig. 4 right panel cyan column).

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Before curve fitting, the data can be plotted to check for any anomalies. Use scatter plots to plot: 1. The raw data for each bleach ROI and non-bleach ROI on a graph as a function of time, to check the intensity changes as expected (Fig. 5a). 2. The normalized data for each bleach ROI on a graph as a function of time (Fig. 5b). The values on the y axis should range from 0 (intensity of the time point after the bleach) to 1 (normalized to the pre-bleach intensity).

3.7.4 Curve Fitting and Extraction of Summary Data

Once the bleach ROI intensities have been normalized for each image, then the averaged data per time series is transferred from the spreadsheet into GraphPad Prism. We normally fit a curve for each experimental sample (i.e., one time series with multiple bleach ROIs) and then a final curve which averages the data from multiple time series (where n is the number of image series from different wings), together with 95% confidence intervals (Fig. 5c). There are many different exponential curves that can be fitted to the data, and we recommend testing a few of the ones built into GraphPad Prism (see Note 37). These include one-phase and two-phase exponential curves. The simplest model is best, so fit a one-phase exponential curve first and check the R square value for goodness of fit, using GraphPad Prism’s analysis tools. However, with care a two-phase model can be used which may help to reveal rates of two processes. In theory a more than two-phase model could be fitted, but in practice this is difficult to achieve due to the difficulty in obtaining sufficiently good signal-to-noise ratio in liveimaging and with sample movement [10]. To empirically measure which model is the best fit nonlinear regression analysis can be used [56]. For the core polarity proteins, we fit a one-phase recovery curve to our data and check it by calculating the R-squared value.

3.7.5 Comparing Curves Between Data Sets

Two or more curves can also be compared within GraphPad Prism by using the Extra sum-of-squares F test, which determines if there is a significant difference between the curve plateaux (YMax) and the rates (t1/2) of recovery. The half-time or rate of recovery (t1/2) is described as the time it takes for half of the recovery to occur. For this measurement to be accurate the recovery of the fluorescence needs to approach a plateau. Note that calculated half-times using an exponential function depend on the size and shape of the ROI and the amount of photobleaching. Therefore, only experiments performed using the same settings and the same ROI size and shape can be compared.

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Fig. 5 Workflow of data analysis and graphing. (a) Graph of raw data intensities over time, showing simulated results for the bleach ROIs and the average of the control non-bleach ROIs. (b) After background correction the data is normalized and rescaled, and each bleach ROI for a time series is plotted on a single graph. (c) The data are then averaged between all the ROIs within a time series and then across time series. A curve is fitted to the data, for example a one-phase exponential curve (black line), and 95% confidence intervals are added (dashed lines). (d) The mobile and immobile fractions are then converted into mobile and immobile amounts by multiplying the Ymax by the pre-bleach intensity. Standard error of the means are indicated 3.7.6 Calculating Mobile and Immobile Amounts

If comparing two experimental conditions, it may be worthwhile calculating the mobile and immobile amounts of protein for each condition (Fig. 5d). This will then compare not only the fraction of protein that is mobile and immobile but also take into account the overall amount of the protein present, which may vary between mutant background or conditions. The mobile fraction is the same value as the Ymax, and the immobile fraction is the mobile fraction subtracted from 1. Calculate these per time series, then multiply these mobile and immobile

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fractions separately by the average pre-bleach intensity of the bleach ROIs, to get the mobile and immobile amounts in arbitrary units of fluorescence intensity. These values can be averaged across multiple time series of the same genotype/experimental condition and plotted as a stacked bar charts in GraphPad Prism (Fig. 5d). 3.7.7 (Optional) Correcting Mean Intensity Data if the Samples Have a Variable Distance from the Coverslip

For some in vivo tissues the position of the sample relative to the coverslip can vary between samples. For example, when imaging Drosophila pupal wings, they are at variable distances from the coverslip depending on how they are mounted. The fluorescent intensity of the sample was shown to decrease exponentially with distance from the coverslip [39]. To correct for this, the intensity is normalized so that it resembles levels of intensity that would be seen if the wing was touching the coverslip (see Note 38). If this step is required, then do this step before calculating the mobile and immobile amounts.

3.8 Publishing FRAP Data

Published FRAP experiments do not always describe the basic conditions and settings used, making it difficult to replicate the findings. Others have reported on the lack of clarity and have sought ways of establishing uniformity in the method [54]. We suggest providing a list of protocol and imaging settings that should be stated in Subheading 3 to ensure others can reliably reproduce the FRAP results. We suggest including (as a minimum): 1. For the image settings: microscope, objective (e.g., 60 1.4 NA Plan-Apochromat), laser type and power for imaging and bleaching, any filters in the light path, gain and offset settings, pixel size, pixel dwell time, time intervals and bleaching time, size of the ROIs, and what control regions and background corrections were used. 2. For the image processing: What amount of bleaching was achieved, the amount of acquisition bleaching observed, what controls were used, what normalization was performed, which curves were tried and fitted, the experimental replicate number (n) and what software was used to analyze the data.

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Notes 1. Prior to 20 h APF the two surfaces of the wing are not apposed, and imaging is more challenging. For analysis of asymmetric core planar polarity proteins we collect white prepupae and age for 28–31 h APF (e.g., for imaging 28 h APF wings collect at 10 am and dissect at 1:30 pm the next day, to start imaging at 2 pm).

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2. Keep pupae after imaging to see if they continue to develop and eclose (Fig. 2h). 3. You should be able to see through the cuticle if using a bright light on a dissecting microscope, and observe the head, wings and legs (Fig. 2d). 4. It is important to only uncover the wing part of the pupa. Do not cut further than the posterior end of the wing as the wing will not lie flat. You can remove more of the cuticle anterior to the wing but if using mounting method 1 you will need more halocarbon oil to mount the pupae. Also avoid cutting into the pupa itself, although a nick in the legs is usually acceptable. This dissection technique will need some practice to perfect. 5. Do not dry the pupae completely as they are more difficult to mount; you just want to make sure there is not a large meniscus of water remaining around the pupa. 6. Other mounting methods exist. One such method requires the use of Scotch tape to hold the pupae in position. This works well but also inhibits the repositioning of the pupa after mounting [57]. 7. If the pupa is very dry at this point then the oil coated tip will stick to the pupa. If this occurs then use another pair of forceps to place the pupa back on the dish (this can be fiddly). To stop this from happening, do not dry out the pupa for too long in Subheading 3.2, step 7. Alternatively, some people prefer to use a needle to add the oil. 8. The amount of oil is critical, as too much causes the pupa to roll around in it, but there needs to be sufficient oil for imaging. To check if you have used enough oil, once the pupa has been turned and mounted, turn the dish over and observe the pupa from underneath. The oil spread should cover the base of the pupa but no further. 9. Mounting the wings like this is tricky and takes practice to perfect but the benefit is that the pupa can be repositioned once mounted to allow imaging of another part of the wing. 10. Use only a very fine coating of the heptane glue, as a thicker coating or drops of glue stop the pupae from lying flat. The heptane glue made using Sellotape is not autofluorescent; we have not tried other brands of sticky tape. 11. The heptane glue mounting method is easier to use. However, the pupae cannot be repositioned after mounting. Therefore, experience in how to position the pupae is useful. Practice mounting pupae at different angles to find the best way of positioning the pupae for imaging.

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12. Initial tests should be performed with different time intervals to check if the time interval selected is sufficient to capture the fast rate of recovery. 13. Not all confocal software can perform variable time intervals. 14. The 60 objective is suitable for imaging the small cells of pupal wings, but other tissues may require a lower magnification. 15. Most other microscopes have a recommended pinhole of 1 AU due to the shape of the pinhole aperture. If the fluorescent sample being imaged is particularly weak then the pinhole can be opened to let more light through, although this will lead to a decrease in the resolution achieved. 16. The pixel size is affected by the objective chosen, the scan area and the number of pixels within that area. The chosen pixel size should reflect the size of the smallest objects to be observed. If for example the object you wish to observe is 200 nm in diameter then you will want to use a 100 nm pixel size to sample that diameter twice, to achieve Nyquist sampling. Standard confocal microscopes can typically resolve objects about 200 nm apart. 17. Keep the size of the scan region to a minimum. A larger scan region takes longer to image so can be a problem if require short time intervals, and for weakly fluorescent samples a large scan region can increase the amount of acquisition bleaching occurring. 18. The zoom factor is dependent on the chosen pixel size and scan region. 19. Once the frame size is set then the scan speed determines the pixel dwell time. Reducing the speed at which the sample is scanned allows more photons to be detected. Faster scanning leads to less light being absorbed per fluorophore and a weaker signal but reduces acquisition bleaching. 20. Frame or pixel averaging is not recommended for FRAP as timepoints should not be merged. In addition, averaging would also increase any acquisition bleaching that is occurring. 21. Mounting method 2 results in the tissue to be imaged being nearer the coverslip compared to mounting method 1 and produces brighter intensity images, so imaging settings should be optimized for each method and the mounting methods should not be changed within an experiment. 22. The gain is the voltage passed across the PMT and regulates its sensitivity. Therefore, a high gain increases the sensitivity and detects more photons of light but it will also amplify any noise in the system. A low gain will result in a higher signal-to-noise

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ratio. The Nikon A1 software specifies the gain as Hv and not as the Voltage. The gain we suggest is about 40% of the maximum gain. 23. Gain and offset must be optimized to capture the full range of intensity levels produced by the image. To check your data falls within the intensity limits of 0 to 4095 for a 12 bit raw image, use the maximum and minimum look up table found in the software of most microscopes. Apply it to the image to see if any pixels are over or under-saturated then adjust the gain and offset to keep the intensities slightly more than 0 and less than 4095. However, the maximum intensity can be much lower than 4095 if needed to minimize acquisition bleaching. 24. The scan region size is chosen to limit acquisition bleaching, so the number of bleach ROIs that can be selected within a single image may be limited. The number of regions selected per image will depend on (1) how many points of interest there are, but also (2) how many of those potential bleach ROIs will have to be selected as control non-bleach regions. ROIs should not overlap or be located within the same cell. If the regions were in the same cell then multiple regions would be recovering from the same pool of fluorescent protein, which would reduce the amount of recovery achieved in all of the regions. 25. The number of pre-bleach values should be more than one to provide an average baseline of intensity. 26. The amount of bleaching of the fluorescent signal achieved should lie between 50% and 75% of the initial fluorescence. Less than 50% bleach of the initial fluorescence will result in low signal-to-noise ratio and recovery may be difficult to measure. However, achieving a bleach of more than 80% can lead to crosslinking of proteins and reduced protein turnover [58]. Try varying the bleach amount within the 50–75% range and check that differences in the initial bleach do not change the results. 27. The time taken to achieve a 50–75% bleaching of the initial fluorescence should be as short as possible, as the curve fitting models assume that no movement of molecules is occurring during the bleach phase. Continual bleaching for many seconds means that any fluorescence recovery that is occurring during the bleaching phase is also being bleached, depleting the pool of fluorescent protein available. We aim for a bleaching time that is less than 5% of the time taken for recovery to approach a plateau. Consider shortening the frame rate during the initial bleach to allow more passes of the laser to occur. Observing the recovery curves obtained with different bleach times can help with deciding if the initial bleach is too long and inhibiting protein recovery.

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28. We use less than 100% of the available laser power, typically set to 80%. This allows us to increase the power and maintain the same level of bleaching if the laser power reduces as the laser gets older. If 80–100% of the available laser power still does not bleach to 50% of the initial fluorescence, then the scan rate and/or the number of scans should be altered until the desired loss of fluorescence is achieved. 29. A power calculation should be performed on a small trial dataset prior to the main experiment to calculate how big an n number is required (the sample size). To be confident in your statistical analysis you must be sure that you analysed enough samples to say that the statistically significant result is not due to chance, and you have not made a type I error (which says the mean values are different when they are the same) or a type II error (which says that the mean values are the same when they are different). If the sample size was small then the two samples may look like they are from the same population even if they are not. To check for type I and type II errors a power calculation is performed to calculate the achieved power of the dataset, which should be above 0.80, therefore 80% probability of not making an error. G*Power or a similar open source program can be used for this purpose. The means and the standard deviation of each sample group are required to calculate the effect size and the α value. The effect size is defined as the number of standard deviations between the null mean and the experimental mean, the α value is the probability cut off value of making a type I error. The α value is normally 0.05 (which is the same as for the cut off point for a t-test). There are different types of Power analysis which allows the calculation of: (a) The achieved Power of the experiment given the effect size, the α value and the sample size. This should be done on small sample of trial data before the main experiment is imaged. (b) The sample size required given the effect size, the α value and the power (usually putting the value at 0.80, so that 80% of the time the result will be correct). As well as the power calculation informing the appropriate sample size to be used, it can also test if two samples “not significantly” different from each other have been sampled enough to say they are likely to be from the same population. 30. Ideally the laser power should be checked at the beginning of each imaging session (or at least once a week) to check if the power output has changed. This can be done using a power meter measuring the laser power at the lens using the favored scan settings. If an Argon laser is used it should be warmed up

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for 30 min before use as it can fluctuate for a period of time after it is turned on. Argon lasers deteriorate over a period of months/years and this may not be immediately noticeable. A diode 488 nm laser performs better over time. If FRAP experiments are to be carried out over an extended period of time then a control FRAP experiment should be conducted using identical settings and sample material every 6–12 months to check that results are not altering due to changes in the microscope or laser. 31. Most common microscope software allows you to reopen the images later to see the ROI locations, although they often require the proprietary software and this cannot be done in ImageJ. 32. In vivo samples are prone to moving out of focus either due to sample drift, temperature changes, or developmental movements. If the movement is not too great, XYZ positions can be manually adjusted during image acquisition. Discard FRAP images that lose fluorescence and/or move out of focus. If a bleached protein is in low abundance in the cell, it may be useful to use another reference marker emitting fluorescence of a longer wavelength to track the ROIs and movement of the sample. 33. When light passes through a small circular aperture it forms an Airy disc pattern. This pattern consists of a central Gaussian distributed spot surrounded by less intense concentric rings. These diffraction rings also participate in the photobleaching process and this causes a broadening of the bleached area. 34. Cellular rearrangements and cell division in tissues may result in the intracellular redistribution of junctional proteins. Therefore, the rates and amounts of recovery measured will not be representative of steady state protein exchange. This may result in the apparent recovery of the fluorescence increasing past 100% of the initial fluorescence. 35. To calculate the amount of acquisition bleaching, determine the average intensity of the non-bleach ROIs for the three pre-bleach images. Subtract from this the background value as obtained from the “laser off” image. Do the same for the non-bleach ROIs in the last image of the experiment. Check that the background subtracted intensity at the end of the experiment is at least 80% of the value at the start. We find that any more than 20% acquisition bleaching leads to reduced fluorescent recovery. Acquisition bleaching should have been optimized earlier in Subheading 3.4. However, checking this parameter now can additionally indicate if the sample is suffering from Z drift and is moving out of focus.

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36. To calculate the amount of experimental bleaching, determine the average intensity of the bleach ROIs in the last pre-bleach image and subtract from this the background value. Do the same for the bleach ROIs in the first post-bleach image. Check that the background subtracted intensity after the bleaching step is at least 50–75% lower than the pre-bleach value. 37. To fit an exponential curve in GraphPad prism, input the mean intensities of the bleach ROIs over time in an XY data table. Fit the curve to the XY table data by selecting Analyze, selecting XY analysis and clicking on nonlinear-regression (curve fit). Select the Exponential section and choose the appropriate curve to fit. Select the Diagnostics tab at the top of the current window and choose the R-square to quantify the goodness of fit and the D’Agostino–Pearson omnibus normality test to check the data fits a Gaussian (normal) curve. Also select the Runs test to see if the curve follows the data points. If fitted correctly, the curve should ideally have one data point above the curve and the next data point below the curve. If data points cluster either above or below the curve then that suggests the curve does not fit correctly. The resulting table will indicate how well the curves were fitted. The R value should be about 0.7 but this can vary between experiments. The data should pass the normality test and the Runs test should return a not-significant result. If the data do not pass the tests then other curves should be tried, and the tests repeated. 38. To correct for the variable distance between the coverslip and the Drosophila pupal wing membrane, we extrapolate the fluorescent intensity to what would have been observed if the sample was touching the coverslip (I0) using the light attenuation equation: I ¼ I 0  exp ð  X Þ where I is the intensity at X distance from the coverslip, I0 is intensity at the coverslip, and α is the attenuation coefficient [39]. This equation specifies that light intensity decreases exponentially with distance from the coverslip. To calculate the attenuation coefficient an initial dataset of approximately 10–20 images should be collected. The sample-coverslip distance should be plotted on a graph as a function of mean image intensity. Then a best fit curve should be fitted through the data and a best fit value extracted. In the case of Drosophila pupal wings the best fit value was α ¼ 0.01864 [39], but it might vary depending on the mounting method and sample. To correct for the change in intensity you need to know X (distance from the coverslip). Record the Z position of your sample (either written down from the microscope while imaging or collected from the metadata later), and also the

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Z position of the coverslip. Beware: the coverslip Z position value can ONLY be collected when imaging, if you fail to measure it then, you cannot obtain it later. Calculate the wing membrane distance then correct the intensity and use this to calculate the mobile and immobile amounts. References 1. Lippincott-Schwartz J, Snapp EL, Phair RD (2018) The development and enhancement of FRAP as a key tool for investigating protein dynamics. Biophys J 115(7):1146–1155. https://doi.org/10.1016/j.bpj.2018.08.007 2. Day CA, Kraft LJ, Kang M, Kenworthy AK (2012) Analysis of protein and lipid dynamics using confocal fluorescence recovery after photobleaching (FRAP). Curr Protoc Cytom. Chapter 2:Unit2 19. https://doi.org/10. 1002/0471142956.cy0219s62 3. Kang M, Day CA, DiBenedetto E, Kenworthy AK (2010) A quantitative approach to analyze binding diffusion kinetics by confocal FRAP. Biophys J 99(9):2737–2747. https://doi. org/10.1016/j.bpj.2010.09.013 4. Lippincott-Schwartz J, Altan-Bonnet N, Patterson GH (2003) Photobleaching and photoactivation: following protein dynamics in living cells. Nat Cell Biol:S7–S14. https://doi.org/ 10.1038/ncb1032 5. Phair RD, Misteli T (2001) Kinetic modelling approaches to in vivo imaging. Nat Rev Mol Cell Biol 2(12):898–907. https://doi.org/10. 1038/35103000 6. Lippincott-Schwartz J, Snapp E, Kenworthy A (2001) Studying protein dynamics in living cells. Nat Rev Mol Cell Biol 2(6):444–456. https://doi.org/10.1038/35073068 7. Reits EA, Neefjes JJ (2001) From fixed to FRAP: measuring protein mobility and activity in living cells. Nat Cell Biol 3(6): E145–E147. https://doi.org/10.1038/ 35078615 8. Mueller F, Mazza D, Stasevich TJ, McNally JG (2010) FRAP and kinetic modeling in the analysis of nuclear protein dynamics: what do we really know? Curr Opin Cell Biol 22(3):403–411. https://doi.org/10.1016/j. ceb.2010.03.002 9. Sprague BL, Pego RL, Stavreva DA, McNally JG (2004) Analysis of binding reactions by fluorescence recovery after photobleaching. Biophys J 86(6):3473–3495. https://doi. org/10.1529/biophysj.103.026765 10. Sprague BL, McNally JG (2005) FRAP analysis of binding: proper and fitting. Trends Cell Biol

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Chapter 2 FRET Imaging of Rho GTPase Activity with Red Fluorescent Protein-Based FRET Pairs Bryce T. Bajar, Xinmeng Guan, Amy Lam, Michael Z. Lin, Ryohei Yasuda, Tal Laviv, and Jun Chu Abstract With the development of fluorescent proteins (FPs) and advanced optical microscopy techniques, Fo¨rster or fluorescence resonance energy transfer (FRET) has become a powerful tool for real-time noninvasive visualization of a variety of biological processes, including kinase activities, with high spatiotemporal resolution in living cells and organisms. FRET can be detected in appropriately configured microscopes as changes in fluorescence intensity, lifetime, and anisotropy. Here, we describe the preparation of samples expressing FP-based FRET sensors for RhoA kinase, intensity- and lifetime-based FRET imaging, and postimaging data analysis. Key words Fluorescent protein, FRET, Sensitized emission FRET, Fluorescence lifetime, FLIMFRET, Rho GTPase, RhoA

1

Introduction In Fo¨rster or fluorescence resonance energy transfer (FRET), a donor fluorophore in an excited state nonradiatively transfers its excitation state energy to a nearby acceptor fluorophore in a ground state via dipole-dipole coupling, leading to fluorescence emission from the acceptor. The efficiency of FRET (EFRET), defined as the percent of energy transfer from the donor to acceptor fluorophore, is dependent on the inverse sixth power of the distance between donor and acceptor and occurs only over a distance shorter than 10 nm, making it a very sensitive tool for reporting biochemical activities that produce changes in molecular proximity [1]. Fluorescent protein (FP)-based FRET biosensors, mainly composed of a donor and red-shifted acceptor FPs and a sensing module, have been developed to monitor a variety of biological processes such as

Bryce T. Bajar, Xinmeng Guan and Amy Lam contributed equally to this work. Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_2, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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protein–protein interactions, conformational changes of proteins, ion concentrations, and enzyme activities with a high spatiotemporal resolution in intact living cells and organisms [2]. Unlike dye- or nanomaterial-based FRET reporters, FP-based sensors are genetically encoded and can be easily integrated into various cell types and subcellular structures, allowing for long-term tracking of molecular activities in specific cellular populations within an animal or in specific subcellular compartments [3]. A key component of designing and optimizing FRET-based biosensors is the selection of donor and acceptor fluorophores. Generally, donors and acceptors should be chosen to maximize the spectral overlap between donor emission and acceptor excitation, minimize the spectral overlap between the respective emission and excitation of the donor and acceptor, and maximize the quantum yield of the donor and absorption coefficient of the acceptor [3]. Historically, cyan fluorescent protein (CFP) donors and yellow fluorescent protein (YFP) acceptors have been commonly used due to the availability of high quantum yield fluorescent proteins with emission spectra in the 450–550 nm range, derived primarily from Aequorea victoria green fluorescent protein (GFP). However, CFP-YFP FRET pairs exhibit certain disadvantages compared to red-shifted pairs, including fast photobleaching of YFPs, photoconversion of YFPs, phototoxicity from near-ultraviolet excitation, severe spectral cross talk, and relatively small dynamic range [4]. By contrast, green fluorescent protein (GFP) donors and red fluorescent protein (RFP) acceptors offer less phototoxicity and greater spectral emission separation. Moreover, GFPs are more photostable than CFPs under both one-photon and two-photon excitation [5]. Additionally, when imaging in cells, the red-shifted excitation and emission in green–red pairs offer higher signal-tonoise due to reduced autofluorescence from flavoproteins [6]. In recent years, development of improved GFPs (mClover3 [7] and mNeonGreen [8]) and RFPs (mRuby3 [7], mRuby4 [9], mScarletI [10]) have made GFP-RFP FRET increasingly attractive as FRET pairs. Indeed, recently engineered GFP-RFP FRET pairs outperform bright CFP-YFP FRET pairs with respect to dynamic range, the range of EFRET in which a given FRET reporter operates [4]. In addition, cyan-excitable RFP donors and far-red fluorescent protein acceptors exhibit minimal spectral cross talk and less phototoxicity. More importantly, they are spectrally compatible with GFP-based sensors, allowing for simultaneous imaging of two biological events under single excitation in living cells and organisms. FRET induces changes in the fluorescence intensity and polarization of donor and acceptor and the fluorescence lifetime of donor, which can be detected in various ways [3, 11]. Of these, two methods are most commonly used: sensitized emission ratiometric FRET (Ser-FRET) and fluorescence lifetime FRET (FLIMFRET). In Ser-FRET, the EFRET is calculated as the ratio of the

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uncorrected FRET signal from the acceptor (increase) to the signal from directly excited donor (decrease), thus boosting small changes in FRET. In FLIM-FRET, the EFRET can be achieved by comparing fluorescence lifetimes of the donor in the presence and absence of the acceptor. Compared to Ser-FRET, FLIM-FRET has several advantages and disadvantages. First, FLIM-FRET is more suited for in vivo imaging with intermolecular sensors because the lifetime is independent of donor fluorescence (significantly attenuated in vivo) and the acceptor-to-donor ratio (unknown under cotransfection) [12, 13]. Second, FLIM-FRET enables true FRET efficiency measurements, which allows for the identification of fractions of molecules involved in FRET [13]. Third, FLIMFRET can use a less photostable FP as donor and a dark FP (high extinction coefficient and negligible quantum yield) as acceptor, thus potentially increasing multiplexing of FRET pairs [14]. Fourth, FLIM-FRET has lower temporal resolution (sec to min) because enough photons need to be collected to achieve lifetime decay kinetics. Fifth, FLIM requires expensive and complicated instrumentation, preventing its wide use in most laboratories. However, any lab with a standard epifluorescence microscope equipped with appropriate filters for a particular FRET pair can perform live-cell Ser-FRET imaging. In summary, Ser-FRET is optimal for intramolecular FRET sensors in living cells while FLIM-FRET is superior for intermolecular FRET sensors in vivo. Here we outline the use of GFP-RFP FRET to measure RhoA activity in cultured dissociated neurons with Ser-FRET and in organotypic hippocampal slices with FLIM-FRET techniques. For Ser-FRET, we use the intramolecular Raichu-RhoA-CR reporter, in which a GFP-RFP FRET pair, Clover-mRuby2, replaces the original CFP-YFP FRET pair in Raichu-RhoA FRET reporter, which has been used to analyze RhoA activation in dorsal root ganglion neurons [15], hippocampal neurons [4, 16], in addition to imaging applications for the cell cycle [17]. Raichu-RhoA-CR has been used to study RhoA-dependent axonal growth cone dynamics in hippocampal neurons [4, 18]. For FLIM-FRET, we use mCyRFP1-RhoA and mMaroon1-Rhotekin-mMaroon1, an intermolecular RhoA FRET sensor that uses cyan-excitable red (mCyRFP1) and far-red (mMaroon1) fluorescent proteins as donor and acceptor fluorophores [14]. Both Ser-FRET and FLIM-FRET approaches outlined in this protocol enable live imaging of RhoA activity with high spatiotemporal precision.

2 2.1

Materials Plasmid

1. pCAGGS-Raichu-RhoA-CRs (Addgene #40258). 2. mCyRFP1-RhoA and mMaroon1-Rhotekin-mMaroon1 (Addgene #84358 and 84359).

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2.2 Reagents for Ser-FRET Imaging

1. Papain (Life Technologies). 2. DNase I (Life Technologies). 3. Rat neuron nucleofector kit (Lonza). 4. Eight-well chamber slides (Nunc). 5. Poly-D-lysine (Sigma) in borate buffer. 6. Laminin (BD Biosciences). 7. Plating medium: Neurobasal medium (Life Technologies) supplemented with GlutaMAX (Thermo Fisher), penicillin–streptomycin, and 5% fetal bovine serum (Life Technologies). 8. Phenol red-free Neurobasal medium supplemented with GlutaMAX (Thermo Fisher) and B27. 9. Preclustered Ephrin-A solution: 10 μg mL
1 of ephrin-A4-Fc and ephrin-A5-Fc (R&D Systems) are separately preclustered with 50 μg mL
1 of goat anti-human IgG (H + L) (Jackson ImmunoResearch) in Hank’s Balanced Salt Solution (Thermo Fisher) for 1 h and then mixed. 10. Imaging solution: Hank’s Balanced Salt Solution (Thermo Fisher).

2.3 Reagents for FLIM-FRET Imaging

1. Millicell membranes (Millipore). 2. Plate medium: minimal essential medium (Thermo Fisher) supplemented with 20% horse serum (Sigma), 1 mM L-glutamine (Sigma), 1 mM CaCl2, 2 mM MgSO4, 12.9 mM Dglucose, 5.2 mM NaHCO3, 30 mM HEPES, 0.075% ascorbic acid and 1 μg/mL insulin (Sigma). 3. Imaging solution: Mg2+-free artificial cerebral spinal fluid (ACSF; 127 mM NaCl, 2.5 mM KCl, 4 mM CaCl2, 25 mM NaHCO3, 1.25 mM NaH2PO4 and 25 mM glucose) containing 1 μM tetrodotoxin (TTX, Sigma) and 4 mM 4-Methoxy-7nitroindolinyl-caged-L-glutamate (MNI-caged L-glutamate, Tocris) aerated with 95% O2 and 5% CO2. 4. Biolistic gene transfer system (Bio-Rad) with gold beads.

2.4 Fluorescence Microscope

1. Inverted epifluorescence microscope (Zeiss Axiovert 200 M). (a) 40  water-immersion objective (Zeiss 40  1.2-numerical aperture (NA) C-Apochromat lens) (b) Optical filters (ex, excitation, em, emission, dm, dichroic mirror): ex HQ470/30 nm (Chroma) dm 565 nm, and em 505AELP nm (Omega) for GFP, and ex HQ470/ 30 nm (Chroma) dm 565 nm, and em BA575IF nm (Olympus) for FRET. (c) Cooled charge-coupled device camera (Hamamatsu OrcaER).

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(d) Environmental chamber with temperature and CO2 control (Live Cell Instrument, Korea). (e) Microscopy software (Micromanager 1.4.18 together with ImageJ version 1.49 or later). 2. Custom-built two-photon FLIM setup. (a) 60  water-immersion objective (Olympus 1.0 NA Apochromat objective). (b) Optical filters (dm, em): dm 565 nm (Chroma) and em 520/60 nm (Chroma) for GFP, and dm 565 nm (Chroma) and em 620/20 nm (Chroma) for CyRFP. (c) Two photomultiplier tubes (PMTs) with low transfer time spread (H7422-40p, Hamamatsu). (d) Chameleon Ti:sapphire laser (Coherent) tuned at 720 or 920 nm. (e) Temperature controller (TC-324C, Warner Instrument).

3

Methods

3.1 Ser-FRET Imaging 3.1.1 Hippocampal Neuron Preparation

1. Hippocampal neurons are dissected from embryonic day 18 (E18) rats (Charles River Labs), dissociated with papain and DNase I. 2. Transfect neurons by electroporation with a rat neuron Nucleofector kit and plated at a density of 30,000 per cm2 in 8-well chamber slides. Prior to plating, coat slides by incubating with 0.25 mg mL
1 poly-D-lysine in borate buffer for 12–24 h, washing three times with water for 15 min each, incubating with 18 μg mL
1 laminin in Neurobasal medium for 12–16 h, and washing three times with water for 15 min each. 3. Neurons are plated in Neurobasal medium with GlutaMAX, penicillin and streptomycin, and 5% FBS. 4. Plating medium is replaced 12 h later with phenol red–free Neurobasal medium with GlutaMAX and B27. 5. Neurons are imaged 1–2 days after transfection in imaging solution (see Subheading 2.2) on an Zeiss Axiovert 200M microscope (see Subheading 2.4) at 37  C.

3.1.2 Image Acquisition

1. Cells with appropriate expression level are chosen for FRET imaging (see Note 1). 2. Baseline measurements are taken sequentially in FRET donor and acceptor channels every 3 min for 21 min (see Note 2). 3. Add preclustered ephrin-A and IgG dropwise to chambers at a final concentration of 5 μg mL
1 of ephrin-A and 25 μg/mL of IgG (see Note 3). Continue FRET imaging every 3 min for another 21 min (see Note 4).

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To quantitate the FRET response from the reporter, we measure the ratios (R) of background-subtracted intracellular green (Ig
Igb) and red fluorescence (Ir
Irb) under cyan light excitation in cells of interest and monitor its change over time. Analysis of the FRET response is performed on Fiji/ImageJ, with additional analysis performed on the data processing and visualization software of choice. The FRET response can be represented as follows. 



 ð1Þ ðRt
R0 Þ=R0 ¼ 1
ðI tr
I trb Þ= I tg
I tgb = ðI 0r
I 0rb Þ= I 0g
I 0gb

3.1.3 Image Analysis

where the subscript characters 0 and t represent time 0 and time t after addition of ephrin-A, respectively. 1. Open FRET and GFP image stacks separately on ImageJ. 2. Background subtract both stacks, through either of the methods indicated below. (a) Built-in rolling ball background subtraction (Process>Background Subtraction. . .>Rolling Ball Radius ¼ 50.0 pixels. (b) Manual background subtraction. l

Draw a region of interest (ROI) in the background area of the field of view, at least 15  15 pixels in size.

l

Measure the average (Analyze>Measure).

l

Subtract that value from (Process>Math>Subtract).

value

of

the

the

entire

ROI image

3. To measure the same regions in each cell, we want to use the same ROIs on cells between both stacks. However, the GFP and FRET stacks may not be perfectly aligned depending on the optical alignment between channels in the imaging system. We will first need to align the stacks using one of the methods indicated below (see Note 5): (a) Manual registration l

Find a reference point in the GFP stack.

l

Note the x–y coordinates of that reference point.

l

l

l

In the FRET stack, find the same reference point and note the x–y coordinates. Subtract those coordinates to find the x–y translation required to align each stack. Align images by translating the FRET stack by the identified x–y distance to match the GFP stack using Image>Transform>Translate. . .

4. Regions of interest are manually segmented. Using the ROI Manager (Analyze>Tools>ROI Manager. . .) and the “Polygon selections” option, create ROIs around the desired

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features to analyze (e.g., neuronal growth cones) on each of the cells in the field of view. Click “Add” on the ROI Manager to save the selected ROI (shortcut: t). This enables retrieval of the ROI in either channel. 5. For each ROI, measure the intensities in each cell across all slices in the stack, for both FRET and GFP stacks. Two simple methods in ImageJ include: (Image>Stacks>Measure Stack. . .) or (Image>Stacks>Plot Z-axis Profile. . .) (see Note 6). 6. In your desired analysis software (e.g., Microsoft Excel, MATLAB, R), open the measured values in each channel. 7. Normalize measurements to baseline values in each stack. 8. Divide normalized GFP values by normalized FRET values at each timepoint. This gives the ratiometric FRET response at each timepoint (see Note 7). Imaging results are presented in Fig. 1. 3.2 FLIM-FRET Imaging 3.2.1 Hippocampal Slice Preparation

1. Hippocampal slices are prepared from postnatal 4- to 6-day-old C57BL/6 mice. 350 μm-thick hippocampal slices were dissected using a tissue chopper and then plated on Millicell membranes in minimal essential medium supplemented with 20% horse serum, 1 mM L-glutamine, 1 mM CaCl2, 2 mM MgSO4, 12.9 mM D-glucose, 5.2 mM NaHCO3, 30 mM HEPES, 0.075% ascorbic acid and 1 μg/mL insulin, with fresh medium replaced every other day. 2. After 7–10 days in culture, neurons are transfected by ballistic gene gun (Bio-Rad) using gold beads (8–12 mg). Bullets are coated with plasmids containing mCyRFP1–RhoA and mMaroon1–Rhotekin–mMaroon1 (20 μg each). 3. Slices are imaged 2–5 days after transfection in imaging solution (see Subheading 2.3) on the two-photon FLIM imaging system (see Subheading 2.4) at 24–26  C.

3.2.2 Image Acquisition

1. Hippocampal slices are chosen based on presence of CA1 cells positive for RhoA-sensor expression at least 2 days after transfection. Sensor expression is typically sparse and averages at around 1 cell per slice. 2. Baseline imaging is performed for a single secondary or tertiary dendritic branch of CA1 pyramidal cells every 1 min for 10 min (see Note 8). 3. To activate a single dendritic spine in the dendritic branch, MNI-caged glutamate (Tocris) is uncaged near (~1 μm) the spine of interest by two-photon 720 nm light with a train of 4–6 ms, 2.5–3 mW laser pulses (30 times at 0.5 Hz) (see Note 9).

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Fig. 1 Reporting of fast local RhoA activation in neurons with Raichu-RhoA-CR. (a) Design of the Raichu-RhoACR reporter, based on Raichu-RhoA. PKN, protein kinase N. (b) Ephrin-A stimulation locally activated RhoA in a hippocampal growth cone (asterisks) from the first time point after stimulation. Scale bar, 10 μm. (c) RaichuRhoA-CR acceptor–donor emission ratio change (ΔR/RAD) graphed as mean  S.E.M. Peak ratio change (asterisk) was significantly different from baseline by two-tailed t-test (P ¼ 0.019, n ¼ 5 cells). (c) With Raichu-RhoA, peak ratio changes were not statistically different from baseline at any time (P > 0.1, n ¼ 8 cells). (d) With Raichu-RhoA, peak ratio change was not significantly different from baseline (P ¼ 0.468, n ¼ 8 cells). (Adapted from Ref. 4)

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4. Imaging during uncaging is performed with 2  2 binning at each time-point every 8 s, and continued every minute following uncaging for a period of 25–30 min (see Note 10). 5. To test the specificity of the sensor response to RhoA activity, we recommend performing control experiments by repeating steps 1–3 with the following changes: (a) control imaging solution using 4 mM MgCl2 instead of CaCl2, and (b) using a control sensor that does not target RhoA (e.g., Cdc42 sensor, as per Ref. 14) (see Note 11). 3.2.3 Image Analysis

For measurement of fluorescence lifetime, we use a custom script written in MATLAB or C++. The following explanation is intended for code generation and calculation of fluorescence lifetime. Briefly, the fluorescence lifetime decay curve F(t) is fit with a double exponential function. This is used to measure FRET efficiency as it is directly proportional to the lifetime of the donor fluorescent protein, as described in: FRET¼ 1
τDA/τD, where τDA is the fluorescence lifetime of the donor in the presence of acceptor, and τD is the fluorescent lifetime of the donor alone. τD is obtained by fitting the fluorescent lifetime of mCyRFP1-RhoA alone with a mono-exponential convolved with the Gaussian instrument response function: 

H ðt, t 0 , τD , τG Þ ¼ § exp τG 2 = 2τD


2

A ðt Þ ¼ A0 H ðt, t 0 , τD , τG Þ ð2Þ   2 
ðt
t 0 Þ=τD erfc τG
τD ðt
t 0 Þ=ð2 τG τD Þ ð3Þ

in which τG is the width of the Gaussian pulse response function, t0 is the time offset, and erfc is the complementary error function. A0 is the initial fluorescence before convolution. The instrument response function represents the characteristic measurement feature for each microscope system. Fitting is performed by calculating weighted residuals, E(t) ¼ (F(t)
A(t))2/F(t), by minimizing the error summed over time δ2 ¼ Σt E(t) for fitting parameters t0, τD, and τG. To analyze RhoA activity using FLIM, we fix the value of τD to the fluorescence lifetime obtained from RhoA-mCyRFP1 (3.55 ns) alone. Then we obtain τDA by fitting fluorescence lifetime with: A ðt Þ ¼ A 0 ½P D H ðt, t 0 , τD , τG Þ þ P DA H ðt, t 0 , τDA , τG Þ

ð4Þ

where PD and PDA is the fractional population with the decay time constant of τD and τDA. It should be noted that reliable fitting requires high binding fraction and number of photons. When this condition is not available, τDA ¼ ½ τD provides a good approximation. For experiments with small number of photons, we will fix τDA and τDA, and obtain binding fraction (PDA). To generate the fluorescence lifetime images, we calculate the weighted sum of fluorescence decay in each pixel.

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τm ¼ Σt tF ðt Þ=Σt F ðt Þ
t 0

ð5Þ

in which t0 is obtained by a curve fitting to the fluorescence lifetime decay averaged over the whole image. 1. We mark an ROI over the stimulated spine, as well as an ROI over a nearby unstimulated spine in the same image, which can account for nonspecific changes in lifetime due to repeated imaging. A background ROI is marked for background subtraction. 2. We calculate and average baseline lifetime and binding fraction for the stimulated spine. 3. We calculate absolute changes in binding fraction during glutamate uncaging, and following stimulation. 4. Using the fluorescent intensity changes, we can compare changes in spine volume indicating structural changes induced by glutamate uncaging. Imaging results are presented in Fig. 2.

4

Notes 1. A high expression level may lead to low sensitivity because a fraction of sensors are unresponsive. However, a low expression level can lead to a low signal-to-noise ratio if low excitation power is used, or fast photobleaching if high excitation power is used. 2. Switching filters or filter cubes inevitably introduces a delay between acquisition of donor and FRET signals, which limits its use in monitoring rapid FRET changes. To overcome this limitation, we recommend using image splitters such as DualView (Photometrics) or Optosplit (Cairn Research) or ORCAD2 CCD camera (Hamamatsu) devices that allow simultaneous recording of fluorescence in two channels. 3. Preclustered ephrin-A should be added one-to-one to achieve final concentration. For example, if baseline imaging is performed in 200 μL HBSS, then 200 μL preclustered ephrin-A can be added after the final baseline timepoint. Slow, dropwise addition of the stimulation solution is necessary to minimize agitation of the imaged cells. 4. The interval and total imaging times are dependent on the kinetics and dynamics of a given biological process. 5. In our experience, manual registration gave reliable results and was quick to implement, since we find small and repeatable x–y translations between channels. As a result, we do not commonly use automated registration methods to align individual channels.

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Fig. 2 Measurements of RhoA activity using 2pFLIM. (a) Representative fluorescent lifetime decay curves of the red-shifted RhoA sensor (RhoA CyRM) expressed in HEK293 cells. Counts of number of photons along with the decay curve double exponential fit allow to quantitatively discriminate activity levels of dominant negative (DN, blue) and constitutive active (CA, red) RhoA mutants. (b) Representative pseudocolored lifetime images of HEK293 cells showing different basal activity levels of the RhoA sensor with WT, DN and CA versions. Scale bar: 20 μm, (c) Quantification of binding fraction differences of RhoA CyRM sensor showing quantification of differential basal activity of with WT (wild type), DN (dominant negative) and CA (constitutively active) versions. (d) Representative fluorescence lifetime images of RhoA sensor activity in a dendritic branch of CA1 pyramidal neuron in an organotypic hippocampal slice. Images were acquired with 2pFLIM and show the spread of RhoA activity before (
16 s) and after (+16 s, +2 min +20 min) glutamate uncaging at the dendritic spine indicated with arrowhead. Scale bar: 2 μm. (Adapted from Ref. 14)

6. While either approach allows values to be exported to a spreadsheet, we most commonly use the first approach (Image>Stacks>Measure Stack. . .) and plot the resultant values on Microsoft Excel. However, if the region of interest is dynamic and needs to be adjusted between timepoints, we typically draw manual ROIs for each time point and measure them individually across both channels. This manual approach is possible because relatively few time points are captured. 7. We find it useful to visualize the signal in individual channels in addition to the ratiometric signal in order to identify the amount of baseline noise in the sample, and determine whether FRET signals have the expected proportional decrease in donor signal.

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8. To improve signal-to-noise ratio, each image is averaged from 24 frames. PMT gain is tuned to 0.82 V. To avoid photobleaching and reduction in fluorescence lifetime, average imaging power used is set at 1.5–2.0 mW, as measured under the objective. 9. We use an uncaging protocol that consists of 30 pulses of 4–6 ms, at 2.5–3 mW 720 nm laser power. The pulse dwell time can be altered depending on Z position of uncaging in the slices. Successful uncaging should result in spine volume increases that last for more than 30 min. 10. Photobleaching/photoconversion: The advantage of using a red-shifted FLIM RhoA sensor is that it allows simultaneous imaging in a green emission channel. However, RFPs are inherently less photostable then GFPs and can undergo photoconversion, especially under prolonged illumination by widefield fluorescent lamps. Therefore, care must be given to reduce un-necessary examination under fluorescence lamp to minimum. 11. Control measurements: to validate the specificity and interpretation of changes in FRET as activation of RhoA, several control experiments can be performed. For FLIM, a nonspecific acceptor can be used to account for nonspecific donor only changes in lifetime due to repeated imaging and photobleaching. In addition, uncaging experiments should include mock uncaging with extracellular solution containing no Ca2+ for examining nonspecific effects of 720 nm laser pulses during uncaging.

Acknowledgments This work was supported by National Key Research and Development Program of China (2017YFA0700403), National Natural Science Foundation(NSFC) of China (Grant 81927803, 31670872, 21874145), and NSFC of Guangdong Province Shenzhen Science and Technology Innovation Committee (Grant KQJSCX20170331161420421, JCYJ20170818163925063, JCYJ20170818164040422). We would like to acknowledge support from Human Frontiers Science Program (HFSP) for a longterm postdoctoral fellowship (TL), support from the Max Planck Florida Institute for Neuroscience (TL and RY), support from National Institutes of Health (NIH) grants (R01MH080047, DP1DP1NS096787, and R35NS116804 for RY, and F30EY029952 and T32GM008042 for BTB).

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References 1. Piston DW, Kremers GJ (2007) Fluorescent protein FRET: the good, the bad and the ugly. Trends Biochem Sci 32(9):407–414. https://doi.org/10.1016/j.tibs.2007.08.003 2. Sample V, Mehta S, Zhang J (2014) Genetically encoded molecular probes to visualize and perturb signaling dynamics in living biological systems. J Cell Sci 127(Pt 6):1151–1160. https://doi.org/10.1242/jcs.099994 3. Bajar BT, Wang ES, Zhang S, Lin MZ, Chu J (2016) A guide to fluorescent protein FRET pairs. Sensors (Basel) 16(9). https://doi.org/ 10.3390/s16091488 4. Lam AJ, St-Pierre F, Gong Y, Marshall JD, Cranfill PJ, Baird MA, McKeown MR, Wiedenmann J, Davidson MW, Schnitzer MJ, Tsien RY, Lin MZ (2012) Improving FRET dynamic range with bright green and red fluorescent proteins. Nat Methods 9(10):1005–1012. https://doi.org/10.1038/ nmeth.2171 5. Stoltzfus CR, Barnett LM, Drobizhev M, Wicks G, Mikhaylov A, Hughes TE, Rebane A (2015) Two-photon directed evolution of green fluorescent proteins. Sci Rep 5:11968. https://doi.org/10.1038/srep11968 6. Reinert KC, Gao W, Chen G, Ebner TJ (2007) Flavoprotein autofluorescence imaging in the cerebellar cortex in vivo. J Neurosci Res 85(15):3221–3232. https://doi.org/10. 1002/jnr.21348 7. Bajar BT, Wang ES, Lam AJ, Kim BB, Jacobs CL, Howe ES, Davidson MW, Lin MZ, Chu J (2016) Improving brightness and photostability of green and red fluorescent proteins for live cell imaging and FRET reporting. Sci Rep 6: 20889. https://doi.org/10.1038/srep20889 8. Shaner NC, Lambert GG, Chammas A, Ni Y, Cranfill PJ, Baird MA, Sell BR, Allen JR, Day RN, Israelsson M, Davidson MW, Wang J (2013) A bright monomeric green fluorescent protein derived from Branchiostoma lanceolatum. Nat Methods 10(5):407–409. https:// doi.org/10.1038/nmeth.2413 9. Xu Y, Deng M, Zhang S, Yang J, Peng L, Chu J, Zou P (2019) Imaging neuronal activity with fast and sensitive red-shifted electrochromic FRET indicators. ACS Chem Neurosci 10(12):4768–4775. https://doi.org/10. 1021/acschemneuro.9b00501 10. Bindels DS, Haarbosch L, van Weeren L, Postma M, Wiese KE, Mastop M,

Aumonier S, Gotthard G, Royant A, Hink MA, Gadella TW Jr (2017) mScarlet: a bright monomeric red fluorescent protein for cellular imaging. Nat Methods 14(1):53–56. https:// doi.org/10.1038/nmeth.4074 11. Skruzny M, Pohl E, Abella M (2019) FRET microscopy in yeast. Biosensors (Basel) 9(4):122. https://doi.org/10.3390/ bios9040122 12. Laviv T, Scholl B, Parra-Bueno P, Foote B, Zhang C, Yan L, Hayano Y, Chu J, Yasuda R (2020) In vivo imaging of the coupling between neuronal and CREB activity in the mouse brain. Neuron 105(5):799–812. e795. https://doi.org/10.1016/j.neuron.2019. 11.028 13. Yasuda R (2012) Imaging intracellular signaling using two-photon fluorescent lifetime imaging microscopy. Cold Spring Harb Protoc 2012(11):1121–1128. https://doi.org/10. 1101/pdb.top072090 14. Laviv T, Kim BB, Chu J, Lam AJ, Lin MZ, Yasuda R (2016) Simultaneous dual-color fluorescence lifetime imaging with novel red-shifted fluorescent proteins. Nat Methods 13(12):989–992. https://doi.org/10.1038/ nmeth.4046 15. Nakamura T, Kurokawa K, Kiyokawa E, Matsuda M (2006) Analysis of the spatiotemporal activation of rho GTPases using Raichu probes. Methods Enzymol 406:315–332. https://doi. org/10.1016/S0076-6879(06)06023-X 16. Duman JG, Mulherkar S, Tu YK, Erikson KC, Tzeng CP, Mavratsas VC, Ho TS, Tolias KF (2019) The adhesion-GPCR BAI1 shapes dendritic arbors via Bcr-mediated RhoA activation causing late growth arrest. Elife 8:e47566. https://doi.org/10.7554/eLife.47566 17. Yoshizaki H, Ohba Y, Kurokawa K, Itoh RE, Nakamura T, Mochizuki N, Nagashima K, Matsuda M (2003) Activity of rho-family GTPases during cell division as visualized with FRET-based probes. J Cell Biol 162(2):223–232. https://doi.org/10.1083/ jcb.200212049 18. Takano T, Wu M, Nakamuta S, Naoki H, Ishizawa N, Namba T, Watanabe T, Xu C, Hamaguchi T, Yura Y, Amano M, Hahn KM, Kaibuchi K (2017) Discovery of long-range inhibitory signaling to ensure single axon formation. Nat Commun 8(1):33. https://doi. org/10.1038/s41467-017-00044-2

Chapter 3 Live-Cell Total Internal Reflection Fluorescence (TIRF) Microscopy to Investigate Protein Internalization Dynamics Tejeshwar C. Rao, Tomasz J. Nawara, and Alexa L. Mattheyses Abstract The establishment of apicobasal or planar cell polarity involves many events that occur at or near the plasma membrane including focal adhesion dynamics, endocytosis, exocytosis, and cytoskeletal reorganization. It is desirable to visualize these events without interference from other regions deeper within the cell. Total internal reflection fluorescence (TIRF) microscopy utilizes an elegant optical sectioning approach to visualize fluorophores near the sample–coverslip interface. TIRF provides high-contrast fluorescence images with limited background and virtually no out-of-focus light, ideal for visualizing and tracking dynamics near the plasma membrane. In this chapter, we present a general experimental and analysis TIRF pipeline for studying cell surface receptor endocytosis. The approach presented can be easily applied to study other dynamic biological processes at or near the plasma membrane using TIRF microscopy. Key words Total internal reflection, Fluorescence, Evanescent wave, Dynamics, Clathrin-mediated endocytosis

1

Introduction Fluorescence microscopy is an essential tool in diverse research fields including biology, biomedical engineering, chemistry, physics, and materials sciences [1–3]. The advent of modern cutting-edge light microscopy techniques in combination with the development of the plethora of fluorescent labeling technologies have made studying dynamic processes in living cells routine [4–7]. One challenge in fluorescence microscopy is the relatively small signal compared to background [8, 9]. For example, processes involving vesicular transport such as endocytosis and exocytosis have been investigated using epifluorescence microscopy. In epifluorescence the excitation light propagates through the sample, resulting in excitation of fluorophores both in and out of the focal plane (Fig. 1, left) [10]. While this approach can allow visualization of vesicle recycling, accurate quantitative assessments are

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_3, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Comparison of Epifluorescence (EPI) and Total Internal Reflection Fluorescence (TIRF) microscopy. Cos-7 cells were transfected with clathrin light chain tagged with GFP (CLC-GFP) and imaged with EPI and TIRF. In EPI the excitation light passes though the sample, leading to the excitation of all the fluorescently tagged proteins, regardless of where they are in the cell (left panel). It is difficult to distinguish clathrin at the plasma membrane from clathrin deeper in the cell at the trans-Golgi network. In TIRF the excitation light is incident on the coverslip–sample interface at an angle θ greater than the critical angle. This generates an exponentially decaying evanescent field which selectively excites fluorophores at or near the plasma membrane (right panel). TIRF provides excellent optical sectioning allowing selective imaging of clathrin at or near the plasma membrane, while eliminating background fluorescence from clathrin deeper in the cell. Green proteins ¼ CLC-GFP following excitation, orange proteins ¼ CLC-GFP without excitation, black arrows ¼ direction of the 488 nm laser, θ ¼ incidence angle, scale bars ¼ 10 μm

challenged by the small vesicles and high fluorescence background. An additional hurdle is that resolution along the optical axis, or z axis, is limited by diffraction to approximately 500 nm. Given the variability in vesicle size (50–300 nm), some of which are below the diffraction limit of light microscopy, the uncertainty in their localization and distribution densities in the plasma membrane make dynamic measurements even harder [11]. Epifluorescence microscopy muddles details at the basal surface of the cell with background fluorescence from the entire depth of the cell. Consequently, it restricts quantitative reporting of protein localization and turnover at the cell surface, as these measurements are obscured by a large depth of focus and fluorescence from proteins distributed within the cell volume.

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Total internal reflection fluorescence (TIRF) microscopy allows selective excitation of fluorophores in the cellular environment closest to the coverslip interface without exciting molecules deeper in the cell. The intensity of the TIRF excitation field, also called the evanescent wave decays exponentially creating a very thin excitation field extending only 100 nm into the sample [12, 13]. TIRF offers a major practical advantage over other fluorescence imaging modalities for processes that happen at the cell surface, such as endocytosis and exocytosis (Fig. 1, right) [14, 15]. TIRF provides an increased contrast, limited out-of-focus fluorescence, and reduced photo damage making it ideal for imaging dynamic processes at or near the plasma membrane. The depth of the evanescent field provides improved axial resolution for single vesicles approaching, docking, and fusing with the plasma membrane. Fluorescence is detected with a camera allowing for fast acquisition compared to conventional laser scanning confocal microscopy. TIRF microscopy allows selective visualization of cell–substrate contact regions and quantification of the position, amount, composition, and dynamics of labeled protein or lipid molecules in this contact zone. This is ideal for spatially distinguishing intracellular processes from those occurring at or near the plasma membrane such as in cell adhesion, endocytosis, exocytosis, and cytoskeletal dynamics, all of which are crucial for proper establishment of cell polarity. TIRF microscopy can be implemented in several ways including using a prism, lightguide, or through-the-objective [16–20]. There are multiple excellent papers and reviews highlighting the strengths and weaknesses of each design. Here, we will focus on through-theobjective TIRF which is the most commonly employed geometry in cell biology studies. There are many commercially available through-the-objective TIRF systems (including those from Agilent Technologies, Nikon, Leica, Carl Zeiss, Olympus, and others). If you do not have access to a commercial system, TIRF can be homebuilt with a laser excitation source, an inverted microscope, and a high numerical aperture (NA) objective. Mattheyses et al. have published an excellent primer detailing the background, methodology, and pitfalls of TIRF [21]. In through-the-objective TIRF the evanescent field is created by the process of total internal reflection at the coverslip–sample interface. The excitation light is focused on the back focal plane of the objective so it emerges parallel. It is then incident on coverslip– sample interface at an angle θ. If θ is less than the critical angle as defined by Snell’s law, the light is refracted as it enters the sample. If θ is larger than the critical angle, the light is totally internally reflected at the sample interface and an exponentially decaying evanescent field is generated in the sample (Fig. 2). The depth of the evanescent field depends on the refractive indices of the sample and the coverslip, the angle of incidence (θ), and the wavelength of the excitation light.

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Fig. 2 Microscope setup for TIRF imaging. Through-the-objective TIRF is centered around an inverted microscope with a high NA objective lens. Laser excitation is achieved through a laser launch, illustrated here as a 488 nm laser reflected from a mirror into an optical fiber. The optical fiber brings the laser to a commercial TIRF arm. On the TIRF arm there are manipulators to control the angle (x and y) and focus of the laser. The laser reflects off the dichroic and passes though the objective. The focus manipulator allows you to focus the laser beam on the objective back focal plane. The x and y manipulators control how the laser enters the objective, and ultimately the incidence angle. When the incidence angle is above the critical angle, it is totally internally reflected at the coverslip–sample interface and an exponentially decaying field (the evanescent wave or evanescent field) is generated in the sample

In TIRF, fluorescence is excited with an exponentially decaying evanescent field penetrating only 100 nm into the sample. This facilitates imaging of membrane receptor recycling during exocytosis and endocytosis, lipid-raft dynamics, single molecule detection, real-time kinetics of biomolecular interactions, nanoengineering, and superresolution microscopy among others [22–26]. The decay of the evanescent field intensity functions as an excellent “optical sectioning platform” ideal for imaging fluorescence of cellular features at the cell–substrate interface, such as focal adhesions,

TIRF Microscopy

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secretory vesicles, and endocytic pits [27, 28]. Additionally, the combination of high signal-to-noise (SNR) and high signal-tobackground (SBR) ratios has led to a variety of methods that depend on single-molecule detection on the basolateral surface of cells [29]. These include single-molecule tracking, which can highlight the patterns of protein movement in the membrane and their kinetic rates of association and dissociation; fluorescence resonance energy transfer (FRET) to characterize short range interactions or conformational changes, and localization-based super-resolution techniques to obtain high-resolution maps of protein localizations [30–32]. Additionally, each of the three spatial axes within the evanescent field can be individually polarized. This in combination with the use of orthogonally oriented membrane dyes further provides unique opportunities to determine local cell membrane orientation [33, 34]. In this chapter we focus on how to align your TIRF microscope, collect image, and analyze data with a focus on vesicle internalization in clathrin-mediated endocytosis.

2

Materials 1. 25 mm #1.5 coverslips (Example: CG15XH, Thorlabs, NJ, USA). 2. Attofluor chambers (Thermo Fisher Scientific, MA, USA). 3. Fluorescein. 4. DiI (optional). 5. Tetraspek microspheres, 0.1 um (optional). 6. Inverted fluorescence microscope equipped with a camera (CCD, sCMOS, or EMCCD), epi illumination source, and appropriate filter cubes (see Note 1). 7. High numerical aperture (NA) oil immersion objective lens (NA > 1.4) (Example: Nikon APO TIRF 60 1.49 MRD01691, Tokyo, Japan). 8. TIRF illuminator, lasers, and appropriate dichroics and emission filters. Commercial TIRF systems have accompanying laser launches coupled to an optical fiber and introduced directly to the TIRF illuminator. The specifications for dichroics and emission filters are more stringent than for epifluorescence lamp illumination to preserve the reflected wave front and block the excitation. (e.g., for 488 nm excitation, TRF49904; Chroma). 9. Laser safety goggles—certified for the laser source. 10. Stage top environmental control to maintain temperature and CO2 for live-cell imaging.

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11. Image analysis software: Fiji (ImageJ prepackaged with plugins) (free download: fiji.sc/Fiji) or equivalent software. The experimental sample is adherent cells expressing a fluorescently tagged protein of interest. Before transfection with a fluorescently tagged protein construct, cells should be seeded onto an appropriate substrate for live-cell imaging such as 25 mm #1.5 coverslips placed into individual wells of a 6-well plate (see Note 2 for additional sample details). Mount the coverslip in an Attofluor chamber immediately before imaging. There are two control samples. The required control sample for flat-field correction is a soluble dye matched to the excitation/ emission spectra of the fluorophore in the sample (e.g., GFP and fluorescein). The optional control sample allows you to validate the quality of the evanescent field. This can be either DiI on a coverslip with fluorescein in solution or tetraspeck microspheres.

3

Methods

3.1 Laser Alignment and Focusing of Through-the-Objective TIRF

1. Carefully clean the TIRF objective using lens paper. Place a drop of immersion oil on your high NA TIRF objective. Mount a bare coverslip into an Attofluor chamber, clean the bottom with ethanol, and place it on the microscope stage (see Notes 3 and 4). For TIRF it is always critical to clean both the objective and the bottom of the sample coverslip. Focus on the upper surface of the coverslip using brightfield illumination (see Note 5). 2. Remove all obstructions between the coverslip and ceiling. Turn the laser on and set it to the lowest possible power. The laser illumination will be seen on the ceiling, roughly straight up. To make sure the laser beam is collimated when exiting the objective, adjust the focus to minimize the size of the spot on the ceiling (in our set-up using the manual manipulator, see Fig. 2). 3. Use the x and y angle manipulators to center the laser beam so that it hits the ceiling directly above the objective (Fig. 2). This ensures the beam is aligned straight along the optical axis. Go back to step 2 to refine the focus if needed. 4. The quality of the TIRF field should be confirmed before imaging the experimental sample. You want to check for elimination of propagating excitation light and uniformity of the evanescent field. This is most easily confirmed using a control samples such as a combination of DiI on a coverslip with fluorescein in solution above or tetraspeck microspheres (see Note 6).

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5. Next, using laser excitation view the sample fluorescence through the camera. The illuminated region should be circular and centered in the field of view (see Note 7). Use the x angle manipulator to change the angle of incidence. This will move the laser beam to one side across the ceiling and then down the wall. 6. Continue to increase the incidence angle until you pass the critical angle and are imaging in TIRF. If you are using 488 nm excitation, in TIRF mode (above the critical angel), DiI (red) fluorescence will primarily be visible. If critical angle is not yet reached (incident angle below critical angle), fluorescein (green) will primarily be visible. The DiI fluorescence can then be used to evaluate the uniformity and purity of the evanescent excitation. For microspheres there will be a clear shift to seeing primarily immobile microspheres interacting with the coverslip, and not much background fluorescence from the microspheres in solution (see Note 8). 3.2 Preparing the Microscope

1. Preheat and humidify the stage chamber and switch on the lasers to allow them to stabilize for at least 30 min before imaging (see Note 9). 2. Place the coverslip of transfected cells in an Attofluor Chamber and add 1 ml of live-cell imaging media such as FluoroBrite DMEM media supplemented with 10% FBS. Clean the bottom of the coverslip with ethanol to remove any fingerprints, oil, media residue, or other debris. 3. Clean the objective lens, apply a small drop of immersion oil, and position the sample chamber on the stage over the center of the objective (see Notes 3 and 4). 4. Allow the sample to acclimatize and stabilize in the humidity chamber for 15 min prior to imaging.

3.3

Imaging

1. Using the epifluorescence lamp focus on the sample and find a cell to image. Find a cell expressing the fluorescent construct with a nice contact region. 2. Switch to laser excitation and TIRF imaging. You may need to fine tune the incidence angle using the x manipulator (see Note 10). You should be able to clearly see the transition from Epifluorescence to TIRF while you are adjusting the incidence angle. Choose an incidence angle that eliminates background fluorescence and highlights the features critical for your experiment (Fig. 1). 3. For imaging endocytosis, set the exposure time for approximately 30–200 ms and the interval between frames for 0–300 ms. Acquire image stacks with a duration of anywhere between approximately 1–5 min. All times are estimates and

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should be adjusted based on your biological system, intensity of the illumination, sensitivity of the camera, fluorescence brightness, and photobleaching. 4. After imaging the experimental sample, collect the flat field normalization data. Do not change the TIRF alignment or incidence angle. 5. Prepare the control sample to collect flat field data for 488 nm excitation by dissolving 1 mg/ml of fluorescein in 1 M NaOH. Then dilute 5 μl to 1 ml of fresh NaOH and mix. Place a 25 mm #1.5 coverslip into an Attofluor Chamber and add the diluted fluorescein. 6. Place the control sample onto the microscope and focus on this coverslip. Move to a field of view with no scratches or dirt and acquire an image. Acquire 10 images this way moving the sample between images being careful to maintain focus and avoid contaminants in the field of view. 3.4

Data Analysis

1. Open the acquired data file in Fiji. Each image analysis step below has a description followed by the specific steps in Fiji. Before data analysis, three major corrections must be performed (Fig. 3): (a) Background subtraction. (b) Flat field correction. (c) Bleach correction (only for live cell imaging). 2. For background subtraction pick an area outside the cell and draw a region of interest (ROI) at least 200 by 200 pixels (see Note 11). Measure the mean fluorescence value within the ROI (BGmean) and subtract this value form the raw data (RD) single image or every frame of the image stack: BS ¼ RD  BGmean where BS is the background subtracted image, RD is the raw data, and BGmean is the mean value of the background ROI. In Fiji: Draw a rectangle outside the cell area (background ROI) ! Press M to measure ! note the mean value. Then go to the menu Process ! Math ! Subtract. Enter the mean value of background ROI and click OK. If correcting a stack make sure “Apply to all pictures in the stack” is checked. 3. The experimental data was collected with an inhomogeneous excitation field (some areas had brighter excitation than others). This can be corrected with the flat field correction (FFC). The flat field data is a stack of 10 images of FITC (or a soluble fluorescent dye for the appropriate wavelength). This image stack is averaged to a single image (flat field, FF).

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Fig. 3 Pipeline for data processing. (a) Flowchart representing data processing steps prior to data analysis. The white arrow indicates the orientation of the line scans displayed in (b). The white box shows the off-cell area used to measure the mean background intensity. RD Raw Data, BS Background subtracted data, FF Flat field, corrected data. White circles highlight three clathrin accumulation events at the plasma membrane. Scale bar ¼ 5 μm. (b) Intensity line scans for each of the images displayed above

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In Fiji: Open the flat field correction data. Then go to Image ! Stack ! Z projection. Enter number of frames to be averaged (e.g., 1–10). Select “based on average intensity” and click OK ! Press CTRL+A on the averaged picture ! Press M ! Note the max value from the averaged flat field image (FFmax). 4. Apply the flat field correction. To do so, change your background subtracted data (BS) to 32-bit (BS32-bit), divide by FF and multiply by the max intensity (FFmax): FFC ¼

BS32bit  FFmax FF

In Fiji: Click on the BS image. Go to Image ! Type ! 32-bit. Then go to Process ! Calculator plus ! Set parameters as follow: i1 ¼ BS32-bit, i2 ¼ FF, Operation ¼ divide, k1 ¼ FFmax, k2 ¼ 0. Click OK to compute the flat field corrected image (FFC). 5. Optional: Bleach correction (for live-cell time lapse imaging data set with significant photobleaching). There are different ways to correct your data for photobleaching. Here we use the simple ratio method which corrects for bleaching by normalizing the images in a stack to the same mean intensity resulting in the corrected image Norm. FFC

Bleach correction

!

Norm

In Fiji: Image ! Adjust ! Bleach correction ! Simple ratio ! OK ! Background value ¼ 0.0 ! OK. 6. The data, Norm, is ready for analysis. 7. Identify puncta and generate traces of intensity as a function of time (Fig. 4). A circular ROI is used to measure the intensity. The ROI should encompass the entire puncta, but not much background. The size of the ROI should be constant for all analyses (i.e., diameter ¼ 7 pixels). In Fiji: Draw a circular ROI around a fluorescent puncta and measure the intensity as a function of time ! Image ! Stack ! Plot Z profile. Due to the heterogeneity of endocytic events, one needs to be careful when identifying puncta. One possibility to eliminate bias is to select all puncta visible within a certain frame of the image data. Use ROI manager to keep track of all the puncta and “multi-measure” to measure all intensities at the same time. Quantitative criteria can be set and used to distinguish endocytic events based on colocalization of multiple proteins and/or the dynamics of the disappearance as is appropriate for the question at hand. For example, in Fig. 4, puncta 2 shows clathrin accumulation followed by rapid disappearance, which

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Fig. 4 Quantification of individual endocytotic events using live-cell TIRF. (a) A Cos-7 cell expressing clathrin light chain tagged with GFP (CLC-GFP) was imaged with TIRF for 5 min with a 100 ms exposure time at 300 ms intervals. Scale bar ¼ 5 μm. The white box indicates the inset, scale bar ¼ 1 μm. The empty carrot indicates the location in this frame of (Puncta 1) for which the endocytosis event has completed. The filled carrot indicates an active clathrin event (Puncta 2). (b) Normalized intensity traces for the two representative events (time scale is relative to the endocytosis event), red dashed boxes highlight the entire duration of the event. (c) Two representations of the individual puncta dynamics. Montage constructed using images taken every 6 s. Kymographs represent frame-by-frame clathrin accumulation during the endocytosis event

suggests vesicle scission and internalization while clathrin disappearance in puncta 1 is much slower and can suggest progressive clathrin dispersion, not vesicle scission.

4

Notes 1. This protocol describes through-the-objective TIRF imaging on a commercial TIRF microscope. It can be easily adapted for a home-built microscope [13, 18, 21, 35]. see Fig. 2 for key components. 2. We used Cos-7 cells transfected with clathrin light chain tagged with GFP in the experiments shown here. Cos-7 cells transfect well and have a flat and spread out morphology making them ideal for TIRF imaging of endocytosis. The cell type and fluorescent construct can be selected to suit any experimental need, with the requirement that the cells are adherent on the coverslip as only the cell–substrate interface will be imaged. 3. When lasers, which are coherent light sources, are used for illumination imperfections such as dust or debris in the excitation light path can lead to interference fringes. Careful cleaning of the objective and coverslip can eliminate some but usually not all of the interference fringes in the excitation field. 4. It is critical that there are no bubbles in the immersion oil. Bubbles can lead to scatter of the incident laser beam and a

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disrupted evanescent field. If the image appears obscured or it is difficult to focus: remove the sample, clean the objective and the coverslip, reapply oil, and try again. 5. Finding the focus can be aided by using a small piece of Scotch magic tape (preferably white) affixed on the coverslip or a line drawn with a sharpie to more easily find the surface. If these methods are used, move the tape or sharpie line out of the field of view before you continue with the alignment. 6. One control sample to help evaluate if you are in TIRF is a combination of two different fluorescent dyes excited by the same wavelength. One dye is preferentially stuck to the coverslip surface while the other is suspended free in solution. If the sample is imaged in TIRF, only the dye on the coverslip will primarily be excited. For example, with a 488 nm excitation, fluorescein (green) in solution can be used in combination with DiI (red) on the surface. To prepare the sample, first incubate DiI (0.5 mg/ml in ethanol) on the coverslip for 10 min. Then wash with water and add fluorescein (0.1 mg/ml in 1 M NaOH). When imaging with 488 nm laser, fluorescein will primarily be visible when the incident angle of the excitation light is below the critical angle. Above the critical angle, DiI will primarily be visible [35]. 7. If the illuminate region is not centered in the field of view or if it moves dramatically as the incidence angle is changed, this indicates the alignment of the laser is incorrect. This can be addressed though a more complex alignment protocol not detailed here or a service visit. 8. If you do not see a clear boundary between Epi and TIRF, it may indicate the laser is not focused properly on the back focal plane. Bring the laser beam to the “straight up” position and repeat the focusing step, trying to minimize the size of the laser spot on the ceiling. 9. Instead of using a humidifying chamber to prevent media evaporation a layer of mineral oil can be placed on top of the imaging buffer or the sample chamber can be sealed. 10. Changing the incidence angle should not result in major intensity fluctuations until the critical angle is reached. In cells expressing clathrin light chain GFP, a rapid disappearance of clathrin localized at the trans-Golgi network will be evident as you pass the critical angle into TIRF. The incidence angle may need to be different than when you imaged the control samples. This is because of the different refractive index of the experimental cell sample and the control sample. 11. When selecting the background ROI be sure to exclude any other cells or fluorescent debris present in the image.

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Acknowledgments This work was supported by funding to A.L.M. from the National Institutes of Health/National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIH/NIAMS) (R01AR072697) and the National Science Foundation (NSF) CAREER (1832100). References 1. Combs CA (2010) Fluorescence microscopy: a concise guide to current imaging methods. Curr Protoc Neurosci. Chapter 2:Unit2.1Unit2.1. https://doi.org/10.1002/ 0471142301.ns0201s50 2. Shashkova S, Leake MC (2017) Singlemolecule fluorescence microscopy review: shedding new light on old problems. Biosci Rep 37(4):BSR20170031. https://doi.org/ 10.1042/BSR20170031 3. Renz M (2013) Fluorescence microscopy—a historical and technical perspective. Cytometry A 83(9):767–779. https://doi.org/10.1002/ cyto.a.22295 4. Liu Z, Lavis Luke D, Betzig E (2015) Imaging live-cell dynamics and structure at the singlemolecule level. Mol Cell 58(4):644–659. https://doi.org/10.1016/j.molcel.2015.02. 033 5. Frigault MM, Lacoste J, Swift JL, Brown CM (2009) Live-cell microscopy – tips and tools. J Cell Sci 122(6):753–767. https://doi.org/10. 1242/jcs.033837 6. Jensen EC (2013) Overview of live-cell imaging: requirements and methods used. Anat Rec 296(1):1–8. https://doi.org/10.1002/ar. 22554 7. Halpern AR, Howard MD, Vaughan JC (2015) Point by point: an introductory guide to sample preparation for single-molecule, super-resolution fluorescence microscopy. Curr Protoc Chem Biol 7(2):103–120. https://doi.org/10.1002/9780470559277. ch140241 8. Waters JC (2009) Accuracy and precision in quantitative fluorescence microscopy. J Cell Biol 185(7):1135–1148. https://doi.org/10. 1083/jcb.200903097 9. Lee J-Y, Kitaoka M (2018) A beginner’s guide to rigor and reproducibility in fluorescence imaging experiments. Mol Biol Cell 29 (13):1519–1525. https://doi.org/10.1091/ mbc.E17-05-0276 10. Gaidarov I, Santini F, Warren RA, Keen JH (1999) Spatial control of coated-pit dynamics

in living cells. Nat Cell Biol 1(1):1–7. https:// doi.org/10.1038/8971 11. Mettlen M, Stoeber M, Loerke D, Antonescu CN, Danuser G, Schmid SL (2009) Endocytic accessory proteins are functionally distinguished by their differential effects on the maturation of Clathrin-coated pits. Mol Biol Cell 20(14):3251–3260. https://doi.org/10. 1091/mbc.e09-03-0256 12. Axelrod D (1981) Cell-substrate contacts illuminated by total internal reflection fluorescence. J Cell Biol 89(1):141–145. https:// doi.org/10.1083/jcb.89.1.141 13. Axelrod D, Thompson NL, Burghardt TP (1983) Total internal reflection fluorescent microscopy. J Microsc 129(1):19–28. https:// doi.org/10.1111/j.1365-2818.1983. tb04158.x 14. Paige MF, Bjerneld EJ, Moerner WE (2001) A comparison of through-the-objective Total internal reflection microscopy and epifluorescence microscopy for single-molecule fluorescence imaging. Single Mol 2(3):191–201. https://doi.org/10.1002/1438-5171( 200110)2:33.0.co;2-k 15. Oheim M, Loerke D, Stu¨hmer W, Chow RH (1998) The last few milliseconds in the life of a secretory granule. Eur Biophys J 27(2):83–98. https://doi.org/10.1007/s002490050114 16. Martin-Fernandez ML, Tynan CJ, Webb SED (2013) A “pocket guide” to total internal reflection fluorescence. J Microsc 252 (1):16–22. https://doi.org/10.1111/jmi. 12070 17. Fish KN (2009) Total internal reflection fluorescence (TIRF) microscopy. Curr Protoc Cytom. Chapter 12:Unit12.18-Unit12.18. https://doi.org/10.1002/0471142956. cy1218s50 18. Axelrod D (1989) Chapter 9 Total internal reflection fluorescence microscopy. In: Taylor DL, Wang Y-L (eds) Methods in cell biology, vol 30. Academic Press, Cambridge, Massachusetts, pp 245–270. https://doi.org/ 10.1016/S0091-679X(08)60982-6

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19. Ramachandran S, Cohen DA, Quist AP, Lal R (2013) High performance, LED powered, waveguide based total internal reflection microscopy. Sci Rep 3(1):2133. https://doi. org/10.1038/srep02133 20. Stout AL, Axelrod D (1989) Evanescent field excitation of fluorescence by epi-illumination microscopy. Appl Opt 28(24):5237–5242. https://doi.org/10.1364/AO.28.005237 21. Mattheyses AL, Simon SM, Rappoport JZ (2010) Imaging with total internal reflection fluorescence microscopy for the cell biologist. J Cell Sci 123(21):3621–3628. https://doi. org/10.1242/jcs.056218 22. Rao TC, Passmore DR, Peleman AR, Das M, Chapman ER, Anantharam A (2014) Distinct fusion properties of synaptotagmin-1 and synaptotagmin-7 bearing dense core granules. Mol Biol Cell 25(16):2416–2427. https://doi. org/10.1091/mbc.E14-02-0702 23. Mashanov GI, Tacon D, Knight AE, Peckham M, Molloy JE (2003) Visualizing single molecules inside living cells using total internal reflection fluorescence microscopy. Methods 29(2):142–152. https://doi.org/ 10.1016/S1046-2023(02)00305-5 24. Cai D, Verhey KJ, Meyho¨fer E (2007) Tracking single kinesin molecules in the cytoplasm of mammalian cells. Biophys J 92 (12):4137–4144. https://doi.org/10.1529/ biophysj.106.100206 25. Mattheyses AL, Atkinson CE, Simon SM (2011) Imaging single endocytic events reveals diversity in clathrin, dynamin and vesicle dynamics. Traffic 12(10):1394–1406. https:// doi.org/10.1111/j.1600-0854.2011. 01235.x 26. Asanov A, Zepeda A, Vaca L (2012) A platform for combined DNA and protein microarrays based on total internal reflection fluorescence. Sensors (Basel) 12(2):1800–1815. https:// doi.org/10.3390/s120201800 27. Steyer JA, Almers W (2001) A real-time view of life within 100 nm of the plasma membrane.

Nat Rev Mol Cell Biol 2(4):268–275. https:// doi.org/10.1038/35067069 28. Krylyshkina O, Anderson KI, Kaverina I, Upmann I, Manstein DJ, Small JV, Toomre DK (2003) Nanometer targeting of microtubules to focal adhesions. J Cell Biol 161 (5):853–859. https://doi.org/10.1083/jcb. 200301102 29. Kudalkar EM, Davis TN, Asbury CL (2016) Single-molecule total internal reflection fluorescence microscopy. Cold Spring Harb Protoc 2016(5):pdb.top077800-pdb.top077800. https://doi.org/10.1101/pdb.top077800 30. Rust MJ, Bates M, Zhuang X (2006) Subdiffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat Methods 3(10):793–795. https://doi.org/ 10.1038/nmeth929 31. Bates M, Jones SA, Zhuang X (2013) Stochastic optical reconstruction microscopy (STORM): a method for superresolution fluorescence imaging. Cold Spring Harb Protoc 2013(6):pdb.top075143. https://doi.org/10. 1101/pdb.top075143 32. Weiss S (1999) Fluorescence spectroscopy of single biomolecules. Science 283 (5408):1676–1683. https://doi.org/10. 1126/science.283.5408.1676 33. Anantharam A, Onoa B, Edwards RH, Holz RW, Axelrod D (2010) Localized topological changes of the plasma membrane upon exocytosis visualized by polarized TIRFM. J Cell Biol 188(3):415–428. https://doi.org/10.1083/ jcb.200908010 34. Passmore DR, Rao TC, Peleman AR, Anantharam A (2014) Imaging plasma membrane deformations with pTIRFM. J Vis Exp 86: 51334. https://doi.org/10.3791/51334 35. Johnson DS, Jaiswal JK, Simon S (2012) Total internal reflection fluorescence (TIRF) microscopy illuminator for improved imaging of cell surface events. Curr Protoc Cytom 61 (1):12.29.11–12.29.19. https://doi.org/10. 1002/0471142956.cy1229s61

Chapter 4 Single-Cell Single-Molecule Pull-Down (sc-SiMPull) for Detection of Protein Complexes from Embryonic Lysates Naomi Stolpner and Daniel J. Dickinson Abstract Mapping how proteins form complexes and change binding partners is central to understanding cell signaling. Bulk biochemistry can provide a summary of what complexes are present in a cell, but information about the diversity of individual protein complexes is lost. Here, we describe single-cell, singlemolecule pull-down (sc-SiMPull), a TIRF microscopy-based coimmunoprecipitation method, to visualize thousands of individual proteins, their binding partners, and protein complex stoichiometry directly from single-cell lysate. By iterating sc-SiMPull over time, temporal dynamics of protein complexes in response to signaling can be constructed. Key words Single molecule, Single cell, Cell polarity, TIRF, Microfluidics, Development, Biochemistry

1

Introduction Protein complexes and their dynamics play a fundamental role in cell signaling. Proteins oligomerize, change binding partners, and assemble diverse complexes in response to signals that regulate cell behavior and organismal development. Bulk biochemistry assays provide a useful overview of protein complexes, but these methods have limited time resolution and are difficult to apply in vivo when large amounts of starting material may be unavailable. Here we describe single-cell single molecule pull-down (sc-SiMPull), an imaging-based method to detect and analyze protein complexes from single-cell lysates. We developed sc-SiMPull for C. elegans embryos in order to measure developmentally regulated changes in protein complex composition [1]. In sc-SiMPull, an embryo is trapped and lysed in a microfluidic chip functionalized with antibodies against a protein of interest (Fig. 1). This immobilizes individual protein complexes in a near-native state, and these complexes are immediately detected by multicolor TIRF microscopy. The images are analyzed

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_4, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Illustration of sc-SiMPull microfluidic device and pull down. (a) sc-SiMPull device containing twelve microfluidic channels. (b) Detailed view of a single microfluidic channel. A staged embryo is placed in the inlet well, flows to the constriction point and is trapped. (c) Cross sectional view of a trapped embryo (gray oval) in microfluidic channel. Coverslip is functionalized with antibodies against fluorescent tags. Fluorescent tags (magenta and green circles) are fused to proteins of interest (“protein X; “protein Y”). (d) After the trapped embryo is lysed, antibodies bind to fluorescent tag on the bait protein (“protein X”) and pull down complexes containing the bait protein, including oligomers of “protein X” and heterodimers of “protein X” and “protein Y,” which is labeled with a different fluorophore. Complexes are imaged by TIRF microscopy

with a MATLAB package that quantitatively examines thousands of spots per experiment in an automated and unbiased way to determine protein complex composition by colocalization and protein complex stoichiometry by counting photobleaching steps. By comparing SiMPull data from precisely staged cells, this approach quantitatively shows how complexes change over time [1]. This method has been used to study anterior-posterior polarity in C. elegans, where we showed that PAR-3 oligomerizes and binds to other anterior proteins dynamically during zygote polarization. This approach can be applied to investigate how other protein complexes change during development, or how protein complexes behave when polarity or other developmental signaling is perturbed.

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Materials

2.1 Microfluidic Device Master Molds

1. SU-82015 Photoresist (Kayaku Advanced Materials, Westborough, MA). 2. SU-8 Developer (Kayaku Advanced Materials, Westborough, MA). 3. 300 (76.2 mm) test-grade silicon wafers (University Wafer, South Boston, MA). 4. Photomask for soft lithography. Custom masks are available from many sources; we use CAD/Art Services (Bandon, OR; https://www.outputcity.com/). 5. Spin coater. 6. Two level hot plates with digital temperature setting/display. 7. Mask aligner/UV exposure system with long-pass filter (per wavelength recommendations in the SU-8 documentation provided by Kayaku). 8. Trichlorooctyl silane.

2.2 Microfluidic Devices

1. Polydimethylsiloxane (PDMS) (Sylgard 184 silicone elastomer kit, Dow Corning, Midland, MI). 2. Lint free wipes (Technicloth TX604, Texwipe, Kernersville, North Carolina). 3. Vacuum chamber/desiccator and vacuum. 4. Spin coater. 5. N2 tank with regulator and nozzle. 6. Laminar flow hood (optional but recommended). 7. Convection Oven set at 85  C. 8. Self-healing cutting mat. 9. Razor blades/scalpel. 10. Forceps. 11. 2 mm Biopsy punches. 12. Plasma cleaner. 13. 24  60 mm coverslips. 14. UV/Ozone cleaner machine (optional but strongly recommended; see Subheading 3.2.3 for alternative method). 15. Tupperware. 16. Drierite desiccant. 17. Plastic petri dish. 18. Dissecting stereomicroscope. 19. Lab tape.

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2.3 PEG Functionalization of Devices

1. HPLC water. 2. PEG solutions and storage: (a) 600 Da mPEG-Silane (Gelest cat. SIM6492.72) is kept in a desiccator at room temperature. Backfill the bottle with Nitrogen after each use. (b) 3400 Da Biotin-PEG-Silane (Laysan Bio) is stored as a 1% solution in EtOH at 20  C. The PEG precipitates during storage but should dissolve readily upon warming to room temp. The powder is kept in a desiccator at 20  C. Allow it to warm to room temperature before opening, and backfill with nitrogen after use. 3. SiMPull Buffer: 10 mM Tris pH 8.0, 50 mM NaCl, 0.1% TX-100, 0.1 mg/mL BSA. Prepare stock solutions with HPLC water. Filter-sterilize and store at 4  C for up to 1 month. 4. Clear, nonfluorescent tape: such as 0.7500 Crystal Clear Mini tape (Hampton Research, Aliso Viejo, CA). 5. Electronic repeating recommended).

pipettor

(optional

but

highly

6. Vacuum aspirator attached to 1/800 OD Tygon tubing. 7. Antibodies: We have successfully used the following. 8. Anti-mNeonGreen nanobody: mNeonGreen VHH, recombinant binding protein, code nt-250 (ChromoTek). 9. Anti-HaloTag polyclonal antibody, cat. no. G9281 (Promega). 10. Biotin-anti-Flag monoclonal antibody, cat. no. F9291 (Millipore-Sigma). 11. For antibody biotinylation: (Thermo-Fisher). 2.4 C. elegans Strains and Fluorescent Labeling

EZlink

NHS-PEG4-Biotin

1. Control worm strain(s): Control strains: A SiMPull control strain contains a single transgenic fusion construct of the bait and prey fluorophores. For example, mNeonGreen::HaloTag (strain LP539, available from the Caenorhabditis Genetics Center (CGC)) is a fusion of the mNeonGreen fluorescent protein and HaloTag. This acts as a constitutively colocalized control in SiMPull colocalization experiments, and controls for Halo dye labeling efficiency and fluorophore maturation efficiency. Choose fluorophores with rapid maturation, reliable photobleaching with minimal blinking, and minimal aggregation propensity. See Note 1 for recommended best-performing fluorescent tags. 2. Experimental C. elegans strains: Endogenous tagging of bait and prey proteins using CRISPR/Cas9-mediated homologous repair is necessary to eliminate overexpression artifacts and to

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gather accurate stoichiometry data. Highly efficient and streamlined protocols have been developed for endogenous tagging in C. elegans [2–4]. 3. HaloTag ligand dyes: JF646 is commercially available as a HaloTag ligand (Promega). To prepare HaloTag dyes for storage, dissolve at 1 mM in acetonitrile, make 2 μL aliquots in PCR tubes and dry in speedvac. Store desiccated at 20  C. 4. Cell culture-grade DMSO in ampules (e.g., Millipore-Sigma cat. no. D2650-5X5ML) to dissolve HaloTag ligand dyes prior to feeding to worms. After opening an ampule, make 200 μL aliquots and store at 20  C. Keep one aliquot at room temperature as a working stock (do not freeze–thaw). Discard the working stock and thaw a fresh aliquot every 2 weeks. 5. HB101 (preferred) or OP50 E. coli feeding strain. 6. S medium for liquid culture. Prepare by mixing the following solutions. (a) 10 mL S Basal solution (5.85 g/L NaCl, 1 g/L K2HPO4, 6 g/L KH2PO4, 5 μg/L cholesterol). (b) 100 μL 1 M potassium citrate pH 6.0. (c) 100 μL trace metals solution (1.86 g/L EDTA, 0.69 g/L FeSO4·7H2O, 0.2 g/L MnCl2·4H2O, 0.29 g/L ZnSO4·7H2O, 0.025 g/L CuSO4·5H2O). (d) 30 μL each of 1 M CaCl2 and MgCl2. 7. 96 well plate with round-bottom wells. 8. 27–22 gauge metal needles. 9. Watch glass slides for worm dissection. 10. Sutter P-97 micropipette puller, glass capillaries (World Precision Instruments cat. no. 1B100F-4) and aspirator tubes (Millipore-Sigma cat. no. A5177) for cell/embryo manipulation. 11. Valap: Mix equal amounts of petroleum jelly (Vaseline), lanolin and paraffin wax. Heat gently to melt and stir to combine. Store at room temperature. Melt Valap in a tube in a heat block before use. 2.5

Microscope

1. TIRF microscope with appropriate excitation lasers, objectives, filters, and camera. A detailed description of microscope optics is beyond the scope of this chapter, but most commercial TIRF systems are readily capable of single-molecule detection. 2. Micro-manager microscope control software (available free at https://micro-manager.org/). 3. Micro-manager plugin for SiMPull acquisition (available free at https://github.com/dickinson-lab/SiMPull-Acquisition).

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2.6 Analysis Software

3

1. MATLAB. 2. Analysis software for SiMPull data (available free at https:// github.com/dickinson-lab/SiMPull-Analysis-Software).

Methods

3.1 Mold Fabrication for Microfluidic Devices

Microfluidic devices are fabricated using a standard soft lithography procedure. 1. Design a photomask corresponding to the desired channel shape and order from a mask supplier (see above). 2. Deposit a 25–30 μm-thick layer of SU-82015 photoresist on a plasma-treated silicon wafer by spin coating for 10 s at 500 rpm followed by 30 s at 875 rpm. Soft-bake on hot plates for 3 min at 65  C followed by 10 min at 95  C. 3. Expose to 150 mJ UV light through the photomask. 4. Postexposure bake on hot plates for 3 min at 65  C followed by 10 min at 95  C. Develop molds in SU-8 developer and rinse with isopropanol. 5. Hard bake at 95  C for 30 min and then at 120  C for 1–2 h. Finally, treat with vapor-phase trichlorooctyl silane overnight to reduce stickiness.

3.2 SiMPull Device Fabrication 3.2.1 Making PDMS Devices

1. Mix 10:1 ratio by weight of PDMS elastomer base and curing agent in a weigh boat. Mix thoroughly by stirring with a spatula. 2.5 g of this mixture will cover a mold that is 300 in diameter. 2. Pour about a 100 -diameter drop of PDMS onto the center of each mold. Molds can be placed on lint-free fabric wipes for easier handling. Place molds in a vacuum desiccator. Degas under vacuum until there are no more visible air bubbles (about 1–5 min, depending on the strength of your vacuum source). Turn off vacuum and release air slowly, covering the valve with a gloved hand to prevent dust from entering the desiccator. 3. Spin coat molds at 300 rpm for 30 s. 4. Cure in an 85  C oven for at least 20 min (can leave up to overnight).

3.2.2 Cleaning Coverslips with UV Ozone Cleaner

1. Inspect 24  60 mm coverslips for damage and blow off any dust with a stream of compressed nitrogen. Place coverslips on UV ozone cleaner tray. Clean for 20 min. Always use freshly cleaned coverslips; UV ozone cleaning activates the glass surface to allow PDMS adherence.

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This is an alternative method to Subheading 3.2.2. We strongly recommend purchasing a UV/Ozone cleaner, as these machines are relatively inexpensive but save enormous amounts of time and chemical waste while yielding more reproducible results. However, if a UV/Ozone cleaner is unavailable, or for initial experiments, glass slides can be cleaned using Piranha solution as follows: ***Caution: Piranha solution is highly corrosive and can explode if it comes into contact with plastic, metal, or organic solvents. Handle with extreme care.*** 1. Take a bottle of 30% H2O2 out of the fridge and let it warm to room temperature. Do not use cold H2O2—this can lead to coverslip cracking. 2. Blow coverslips with N2 to remove dust. Inspect and discard any that are cracked or visibly dirty. DO NOT clean with organic solvents. 3. Put coverslips in Teflon or glass containers and place in a glass secondary container. A wide dish such as a crystallizing dish (used in organic chemistry labs) is best for this purpose so that coverslips can be laid out in a single layer, not touching each other. 4. Transfer coverslip container to a clean, uncluttered fume hood where they can sit undisturbed for the remainder of the procedure. 5. To prepare piranha solution, pour 50 mL of 30% H2O2 followed by 100 mL of concentrated H2SO4 into the container holding the coverslips (adjust volume as needed depending on your container so that coverslips are fully submerged. Maintain a 2:1 H2SO4:H2O2 ratio). Pour in the acid carefully, but somewhat quickly to encourage mixing. You should see lots of small bubbles, like an electrophoresis chamber. 6. Close the fume hood and leave for at least 4 h or up to overnight. 7. Using Teflon forceps, transfer coverslips one at a time to a container full of ultrapure water. Sonicate for 5 min in a bath sonicator. DO NOT USE METAL FORCEPS FOR THIS STEP, they will dissolve in the piranha solution and can cause an explosion. 8. Transfer coverslips to a new container of water and repeat sonication. Repeat a total of four times to remove all of the acid. Then, sonicate three times in ethanol (to remove the water) and three times in chloroform (to remove the ethanol). Each time the solution is changed, ensure that the coverslips are separate and not sticking to each other. 9. Remove coverslips from chloroform one at a time and dry with N2. If a coverslip is clean, the chloroform should sheet off

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cleanly and should dry almost instantly. If it is difficult to dry or if you see spots of water or oily residue remaining after drying, the coverslip is not clean; throw it out and don’t use it. 10. After drying, put coverslips in a clean glass container and fabricate devices within 24 h. Plasma treat the coverslip along with the PDMS in Subheading 3.2, step 11. 11. Disposal of piranha solution: (a) Pour the used solution into a 4 L beaker containing 2 L of cold water. Label and let stand in a fume hood for at least 24 h (the longer the better) to allow the H2O2 to dissipate naturally. Do not proceed immediately to the next step. (b) In a fume hood, very slowly and carefully add 150 g of solid NaOH per 100 mL of H2SO4 used above, to neutralize the acid. Add the NaOH a little at a time, and wait for the bubbling and fizzing to stop each time before adding more. The neutralization reaction is very energetic; if the NaOH is added too quickly, the solution will boil over. Stirring is not necessary at this step; if you do stir, do so very slowly, gently and carefully. (c) Once all of the NaOH is dissolved, transfer to a stir plate and add a healthy shake of Tris base. Once the Tris is dissolved, check the pH using pH paper. Adjust to neutral using additional NaOH or acid as needed. Universal indicator solution can be added to make this process easier. (d) Once the solution has a pH between 5 and 8, set it aside and let it cool to room temperature. Then check the pH again and adjust as needed. Once the solution is cool and has a neutral pH, it can be safely poured down the sink along with plenty of water. 3.2.4 Assembling PDMS Devices

Recommended: Proceed with the following steps in laminar flow hood to reduce dust. 1. While glass is being cleaned, cut out PDMS devices and punch out wells. Use a razor blade or scalpel to cut through the PDMS on the mold in a rectangle around the entire row of 12 channels—if using the provided mold template, there are two devices per mold. Be sure to allow enough space around the circular wells to punch holes later. Be careful not to nick any of the features on the mold while cutting as they can come off if scraped. 2. Starting at one corner, use forceps to gently peel the PDMS device off of the mold. It can be helpful to peel diagonally from one corner of the device to avoid ripping channels. 3. Place PDMS device on the self-healing mat with the channel side—the side that was in contact with the mold—facing

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up. Avoid touching the channel side of the PDMS device— handle with forceps or gloved hands on edges/corners. 4. Use a 2 mm biopsy punch to punch out the inlet and outlet well for each channel. Align the biopsy punch directly on top of the circle imprinted from the mold, then firmly press down all the way through the PDMS. 5. Once all PDMS devices are cut out and the wells are punched out, place each device channel side up on plasma cleaner tray. Make sure PDMS devices are not overlapping or touching. 6. Place tray into plasma cleaner and plasma clean at approximately 150 W power and 200–250 mTorr vacuum pressure for 30 s. As different plasma cleaners vary in their power levels, you may need to adjust settings to achieve good PDMS-glass bonding. 7. Immediately adhere PDMS devices to the clean side of the glass coverslip with the PDMS channel side down, touching the glass (i.e., the side of the PDMS that was up in the plasma cleaner should be touching the side of the coverslip that was facing up in the UV/Oz machine). Using gloved hands, pick up the PDMS from both ends and hold it so that it forms a U-shape. Then, slowly lower the PDMS onto the clean glass (Fig. 1a). Touch the center of the PDMS (the bottom of the “U”) to the glass first, then roll slowly out toward the ends to avoid introducing air bubbles. The PDMS should cling readily to the glass since both surfaces are activated. Do not lift and realign PDMS on glass. Use tip of forceps to gently flatten any air bubbles. 8. Place assembled devices into a sealed plastic Tupperware container with Drierite desiccant for 10 min. This absorbs water molecules that are generated by the chemical reaction between the glass and PDMS, facilitating a permanent bond. Transfer devices to petri dishes for storage and tape down for easier handling. Proceed immediately to next step. 3.3

PEGylation

PEGylation with PEG-Biotin prepares the microfluidic device for adding antibodies and blocks nonspecific binding. PEGylated devices can be stored in the dark at room temperature for 1–2 months without loss of passivation [5]. Use HPLC-grade water for wash steps. 1. Immediately before use, transfer 250 μL of mPEG-Silane to a tube. Add 2.5 μL of Biotin-PEG-Silane solution (in EtOHbring to room temperature and vortex before using to ensure no precipitate remains) and 2.5 μL of HPLC water. Mix thoroughly by vortexing and pipetting up and down.

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2. Apply 2 μL of the PEG mixture to one side of each microfluidic channel. The PEG should slowly flow into the channel; check under a dissecting scope. Once the channel is filled, add 0.5 μL to the outlet well to allow flow while ensuring that both wells get coated. Incubate at room temp for 30–60 min. An electronic repeating pipettor is useful for these steps. 3. Aspirate the PEG using a vacuum with 1/800 OD size tubing attachment. Wash 2 with HPLC water by applying 1.5 μL water to both wells, then vacuuming the water through from one side. This ensures that the wells, which constitute the majority of the surface area, as well as the channel are washed. After washing, use vacuum to completely dry the lanes—check that no liquid remains under dissecting scope. 4. Place the PEGylated devices in a Tupperware container with desiccant to cure the PEG layer. Cure at least overnight, and ideally 24 h, in the dark at room temperature. The curing step is essential for maturation of the PEG layer and for maximal resistance to nonspecific binding. Cured devices may be stored desiccated for at least 1–2 months without loss of passivation [5]. 3.4 Antibody Functionalization

Devices functionalized with antibody can be stored at 4  C for up to a week. 1. When ready to use a device, wash once with SiMPull buffer to re-hydrate the channels. Aspirate most of the liquid from the wells but leave buffer inside channels. 2. Prepare a solution of 0.2 mg/mL Neutravidin in SiMPull buffer and apply 1.5 μL to each outlet well. Incubate 10 min., then wash 4 with 1.5 μL SiMPull buffer (see Note 2). 3. Prepare a solution of 100 nM of the appropriate biotinylated antibody in SiMPull buffer. Add 1.5 μL to each outlet well. Incubate 10 min, then wash 4 with 1.5 μL SiMPull buffer. 4. If any lanes will not be used immediately, put 0.5 μL of SiMPull buffer in each well and seal with clear tape to prevent evaporation. Label petri dish containing the devices and use immediately, or store at 4  C for up to 1 week, depending on the antibody used.

3.5 Halo Dye Labeling

1. To prepare for use, dissolve an aliquot of dye in 2 μL of DMSO to reconstitute a 1 mM stock. Replace DMSO stock every 2 weeks (see Note 3). 2. Spin down 0.5 mL HB101 or OP50 bacteria (grown overnight and stored at 4  C) and resuspend pellet in 100 μL S media.

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3. Add 1–1.5 μL dye to resuspended bacteria for 10–15 μM final concentration. 4. In each well of a 96-well plate add 30 μL OP50–dye mixture and 20–30 L4 worms. 5. Add water to adjacent wells to prevent evaporation. 6. Shake at 20  C overnight. 3.6 Embryo Dissection

1. Pull glass needles for embryo manipulation see Note 4. These will be used with a mouth pipette to move individual embryos into the SiMPull device. 2. Prepare two depression slides—one with 50 μL egg buffer and the other with 50 μL SiMPull buffer. 3. Prepare the SiMPull channel. If the channel was previously sealed with clear tape, remove the tape from the channel that will be used, leaving the remaining wells taped. Wash the channel once with SiMPull buffer. Add 1.5 μL SiMPull buffer to the inlet well. 4. Use a worm pick to transfer 2–3 gravid adult worms into the egg buffer drop in the first watch glass. Use two metal needles in a scissor motion to bisect the worms. Continue dissecting each half of the worm until all embryos are dissected. Younger embryos will be near the head and tail ends of the worm. 5. Use a mouth pipette with a glass needle to transfer your embryo of choice to the SiMPull buffer in the second watch glass. Wash the embryo a few times in the SiMPull buffer by moving it to different spots in the buffer drop by mouth pipetting. 6. Use the mouth pipette to transfer the embryo to the drop of buffer in the inlet well of the prepared SiMPull channel. Using the mouth pipette or a clean metal needle, gently nudge the embryo toward the opening of the channel. The embryo should flow into the channel toward the constriction point. If not, gently touch the vacuum tubing to the outlet well to encourage flow. The orientation of the embryo in the channel is not important because upon lysis, mixing and diffusion of the cellular contents will occur.

3.7

Sealing and Lysis

Embryo lysis can be altered to control whether there is flow in the channel during lysis. Sealing the channel before lysis eliminates flow and concentrates the lysate into a smaller volume. Alternatively, lysing the embryo before sealing allows flow and dilutes the lysate. Sealing before lysis is our typical strategy because it captures the majority of the protein of interest, which is important when using endogenously tagged proteins that are expressed at normal levels. In some cases, such as in mNG::Halo controls, where the protein is

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expressed under a strong promoter, the bait protein is too abundant and lysis without flow results in images with spot density that is too high for reliable colocalization measurements. If analysis shows that protein is too dense, the cell can be lysed before sealing instead (Subheading 3.7.4), in order to dilute the lysate and reduce the spot density. Channels can be sealed using two methods. We find that different members of our lab have different preferences, so we recommend that new users try both to decide which they prefer. 3.7.1 Tape Sealing Method

1. Aspirate extra buffer out of inlet and outlet wells; add just under 0.5 μL buffer to inlet and then outlet well so there is a concave film of buffer in each well. 2. Pull out ~100 of clear crystallography tape from the end of the tape dispenser so that the tape hangs freely without touching anything. Using forceps and a pair of small scissors, cut out a small square of tape that is just large enough to cover one well of the device. 3. While watching under the dissecting scope, use forceps to gently place the tape square over the outlet of the device (the opposite side of the constriction from where the embryo is. Use forceps to gently press around the well to form a seal. Placing the tape creates a small amount of suction force, which tends to pull the embryo toward the well that was just taped. For this reason, it is critical to tape the outlet well first; this way, the constriction stops the embryo flowing out of the channel. 4. Cut a second small piece of tape (as in step 2) and place it over the inlet well, pressing gently around the edges to form a seal. 5. Untape the device from the petri dish. Use a razor or scalpel to cut any clear tape that is adhering the SiMPull device to the petri dish.

3.7.2 Valap Sealing Method

1. Aspirate extra buffer out of inlet and outlet wells; add just under 0.5 μL buffer to inlet and then outlet well so there is a concave film of buffer in each well. 2. Untape the device from the petri dish. Use a razor or scalpel to cut any clear tape that is adhering the SiMPull device to the petri dish. 3. Add a drop of melted valap to cover the outlet well, then a drop of melted valap onto the inlet well to cover. The embryo will continue to develop in the sealed chamber since the PDMS is oxygen permeable. Channel should be free of air bubbles for proper TIRF.

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3.7.3 Lysis

1. While watching under the dissecting scope, use the tip of a pencil or a previously melted end of a glass Pasteur pipette to firmly press on the PDMS above the embryo to crush the eggshell. Successful lysis should be clearly visible, as the embryo will disappear and be replaced by a small “cloud” of debris. Immediately proceed to acquiring imaging data.

3.7.4 Lysis Before Sealing (for Colocalization Controls/Highly Abundant Proteins)

1. For lower flow: Aspirate extra buffer out of inlet and outlet wells; add just under 0.5 μL buffer to inlet and then outlet well so there is a concave film of buffer in each well. Lyse as in Subheading 3.7.3. Seal with tape or valap as in Subheadings 3.7.1 and 3.7.2. 2. For higher flow: Aspirate extra buffer out of inlet and outlet wells; add 1uL buffer to inlet well then 0.5 μL buffer to outlet well—the volume difference will encourage flow. Lyse as in Subheading 3.7.3. Seal with tape or valap as in Subheadings 3.7.1 and 3.7.2.

3.8

Data Acquisition

For each microfluidic channel containing a single embryo lysate, a series of TIRF images are acquired at several positions along the length of the microfluidic channel. Each image field will contain an array of diffraction-limited spots which are the pulled-down bait protein and any associated prey proteins. The acquisition program will acquire consecutive frames at each position along the channel. The laser power and exposure can be adjusted to photobleach fluorescent proteins to gain stoichiometric information about the pulled-down protein complexes. Since each microfluidic chip has 12 separate channels, one can easily collect several biological replicates in a single experimental session. A typical experiment will include one positive control sample (ex. Halo::mNG), which will be used to determine or verify the labeling efficiency of dyes, and replicates of experimental samples. 1. Open Micro-manager and launch the SiMPull Acquisition plugin. 2. Place SiMPull device on the TIRF objective so that the microfluidic channel is in view. Use minimal oil, since excess oil can cause oscillations in Z that appear as signal fluctuation. Make sure the coverslip side, not the PDMS side, of the device is in contact with the objective. 3. Use brightfield to find and focus on the edge of the channel near the inlet. With the channel containing lysate in focus and in the center of the field of view, select a region of interest (ROI) inside the channel, not including the edges. PDMS is autofluorescent, so try to avoid including channel edges in the ROI (but accidental inclusion of edges can be corrected later in data processing).

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Fig. 2 Determining X-offset and configuring SiMPull data acquisition. (a) Brightfield image of SiMPull channel (edges of channel are thick black lines on sides) with yellow region of interest (ROI) centered in channel. Overlaid blue bars highlight that ROI is centered in the channel in X. Inset shows stage position coordinates with X position circled in blue. (b) After moving the stage down 150 μM in Y, the ROI is not centered in the X dimension, indicated by overlaid red bars on either side of the ROI. Inset of stage position X coordinate did not change because stage was only moved in the Y direction. (c) To correct for the misalignment, the stage was manually adjusted in the Y direction until the ROI appears centered, as indicated by overlaid blue bars.

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4. Set laser powers for relevant wavelengths in Micro-manager (see Note 5). 5. Switch to the fluorescent channel corresponding to the bait protein and focus. A field of spots should be visible; if not, move along the microfluidic channel toward the place where the cell was lysed until fluorescent molecules are seen. Focus the microscope. If spots are bleaching too quickly and not allowing time to adjust focus, temporarily decrease laser power. Enable Perfect Focus (for Nikon microscopes) or other focus maintenance system. 6. Due to the manual nature of PDMS device fabrication, it is common for the long axis of the microfluidic channel not to be perfectly aligned with the Y axis of the motorized stage. Therefore, determine the X-offset, which is the distance the stage needs to move in X in order to stay in the center of the channel (Fig. 2). 7. Align the ROI in the center of the channel and note the X position (Fig. 2a). 8. Move down the channel (in the Y direction) 150 μm (Fig. 2b) and realign the ROI in the center of the channel (Fig. 2c) so channel edges are not in view. 9. Note the new X position value (Fig. 2c). The X-offset is the difference between these two X position values. Enter the Xoffset in the corresponding field in the SiMPull Acquisition dialog box (Fig. 2d). 10. Set the number of stage positions to take along the channel. Acquiring twenty images is usually sufficient but can be adjusted as desired. 11. Select the appropriate imaging channels and enter the appropriate frame number and exposure time for each channel in the top half of the SiMPull acquisition GUI/dialog box (Fig. 2d). We typically acquire 500 frames per field of view, at 50 ms exposure. Acquire channels in reverse wavelength order (starting with the reddest wavelengths and moving to the bluest ones) to avoid crosstalk between channels. 12. Select saving directory and prefix for files. To save files with the organization that the analysis software expects, make a top-level folder for each experiment containing subfolders of images corresponding to each lane of the microfluidic device ä Fig. 2 (continued) Inset of stage position coordinates shows new X coordinate of the stage. (d) The X offset is the difference between the starting X coordinate (a inset; b inset) and final corrected X coordinate (c inset). This value is entered into the SiMPull dialog box for the “Distance to move in X (corrects for nonvertical channel) value. Select channels and enter number of frames and exposure at the top half of the dialog box

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(Example: Create a folder “Exp001” with subfolders: “Exp001_control; Exp001_embryo1; Exp001_embryo2, etc.”) 3.9 Automated Data Analysis

The automated analysis software [1] will find spots in each image, determine spot colocalization, and count photobleaching steps (see Note 6). To accomplish this, the software generates a spot detection image for each position along the channel (Fig. 3a). The spot detection image is an average of first several frames which increases the signal to noise ratio for improved spot detection by the probabilistic segmentation algorithm (see Note 7 [6]. After spot detection, colocalization is counted by looking for detected spots at the same location in each fluorescence channel. Stoichiometry is determined by using a Bayesian algorithm to count photobleaching steps [7]. Then, a summary matrix is generated that tabulates spots counted, colocalization percentages and photobleaching step counts for the entire experiment. Analysis generates three file types in the experiment directory location: a spot detection image for each position and color, a MATLAB data file for each embryo channel, and an overall summary file for all channels in the experiment.

3.9.1 Analysis Software Installation

1. To simplify subsequent steps, start by installing the Git version control on your system. For an introduction to using Git, see [8]. (a) Follow the instructions at https://git-scm.com/book/ en/v2/Getting-Started-Installing-Git/ to install Git. (b) Download the MATLAB git wrapper from https://www. mathworks.com/matlabcentral/fileexchange/29154-athin-matlab-wrapper-for-the-git-source-control-system which will allow you to run Git commands from within MATLAB. Place the downloaded file ‘git.m’ into your main MATLAB folder. 2. Launch MATLAB and import SiMPull Analysis software package using the command ‘git clone https://github.com/ dickinson-lab/SiMPull-Analysis-Software’. 3. Right-click on the newly created ‘SiMPull-Analysis-Software’ folder and select ‘Add to Path > Selected Folders and Subfolders.’ This allows MATLAB to utilize the functions you just installed. Then, type ‘savepath’ at the command prompt to save these settings for future sessions of MATLAB. Our lab periodically makes updates and improvements to the software (see Note 8).

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Fig. 3 sc-SiMPull data analysis workflow. MATLAB functions for each step are in the blue boxes. (a) Automated Analysis. Raw images (outlined in green and red) are analyzed in MATLAB to produce averaged spot detection images used for colocalization detection and photobleaching step counting. (b) Post Processing. Images are inspected and imaging artifacts, such as out of focus images and images containing the edges of PDMS channels are discarded. Shown is a screenshot from the colocalization viewing tool within the analysis software. mNeonGreen spots are circled in green, HaloTag JF646 (far red) spots are circled in red, and colocalized spots are circled in yellow. Note that in the image presented here, colocalized spots are

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3.9.2 Automated Data Processing

1. Run the “analyze_batch” function. 2. Select the experiment folder containing subfolders to analyze (remember, your entire experiment should be in one folder, with each subfolder containing images from one sample). 3. Select image file type. 4. Check boxes for wavelength(s) to be detected. 5. Define frames to be averaged for spot detection for each wavelength see Note 7. 6. Click ‘Continue’ to run the analysis. Depending on the speed of your computer and the size of the dataset, processing may take a few hours, so we typically set it up at the end of the day and let it run overnight.

3.10 Post Processing and Data Visualization

After analysis, images need to be manually sorted to eliminate artifacts such as dirt, edges of the PDMS channel, and out-offocus images. After sorting, built-in data visualization tools can be used to plot and analyze data quality and interpret results. This section of the protocol outlines a typical data processing workflow. It is not an exhaustive list of the tools in the SiMPull software package; see software documentation for descriptions of the full list of functions available.

3.10.1 Sort Images and Discard Artifacts: colocalization_inspector_GUI

Run “colocalization_inspector_GUI.m” Select the .mat data file corresponding to the experiment. The averaged images, which were generated by analyze_batch during data processing, will open in the GUI window. Detected spots will be outlined in color coordinated circles, and colocalized spots will be outlined in a merged color (Fig. 3b). Toggle these spot outlines on/off in the “Show” box in the upper left. Use the keyboard shortcuts to navigate through the images (N ¼ next, P ¼ previous, R ¼ random). Images containing artifacts, such as channel edges, dirt or abnormal spots, and/or out of focus spots can either be completely deleted, or portions of the image can be selected to be kept or removed (Fig. 3b). 1. To remove entire images containing imaging artifacts, navigate to the image to be deleted and click “Toss Image” in the “Artifact Removal” menu in the upper right.

ä Fig. 3 (continued) circled on all three images (not just the merged image); the user can change this behavior by toggling which spots are circled. (c) Data Visualization. (i) Scatterplot of spots per image along the channel. (ii) Colocalization bubble plots. Each circle represents data from one embryo, and the size of the circle represents how many molecules were detected in the dataset. (ask dan what the error bars are). (iii) Individual photobleaching traces (red trace) and tabulated photobleaching step counts (inset bar graph) are produced to determine protein complex stoichiometry

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2. To select portions of images containing imaging artifacts, click “Draw Region” in the “Artifact Removal” menu in the upper right. Using the options in the “Apply Region” box, choose if this ROI manipulation should be applied only to the open image, or if it should apply to all images in the experiment. Either draw an ROI around the area to keep and click “Define ROI” or draw an ROI around the area to discard and click “Exclude ROI.” 3.10.2 Evaluate Experiment Quality and Lysis: “spotcount_1dplotter”

To generate a scatterplot of spots per image (Fig. 3c-i), use spotcount_1dplotter. This is useful for determining if the pulldown was successful, as well as the amount of flow after lysis. A curve with a peak in the center corresponds to lysis with minimal flow, whereas a flatter curve indicates more evenly dispersed lysate (see Note 9).

3.10.3 Determine Labeling Efficiency and Protein Colocalization

1. In the summary table, refer to columns 3 and 4 for “% colocalization with (color)” to determine colocalization. The colocalization percentages for the control labeling strain (fusion protein of bait and prey fluorophores ex. mNG::Halo) represent fluorophore maturation and dye labeling efficiency, and will vary depending on which fluorescent proteins and HaloTag ligand dyes are used. In our hands, the mNG::Halo control strain should give 60–70% of far red spots colocalizing with green, and 70–80% of green spots colocalizing with far red. 2. Next, experimental data can be analyzed. Colocalization for experimental channels will vary widely depending on the bait and prey proteins, along with any other perturbations performed in the experiment. Generally, colocalization below 5–10% may represent background and should be interpreted with caution. A negative control experiment using only prey protein can help determine background binding levels due to the tags alone. 3. Sometimes, one wishes to consider only certain images when calculating colocalization, since including images that contain only background signal will compromise the accuracy of the measurement. To calculate colocalization for only a specific region, first use spotcount_1dplotter.m to identify the portion of each dataset that contains signal above background. Then, use subregion_stats.m to calculate colocalization for only the desired images. 4. Use plotspreadbubble.m to generate a “bubble plot” (Fig. 3cii). In these plots, each data point represents a single replicate experiment (i.e., a single lysed cell or embryo), while the size of the circle represents the number of molecules that were counted in that experiment. These plots are useful for displaying colocalization in different conditions or different timepoints/stages.

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3.10.4 Stoichiometry and Photobleaching Step Counting: See Note 10

1. The summary table displays the fraction of spots photobleaching with 1, 2, 3. . ., 10 steps in separate columns. A final column lists “rejected” spots, which are those that either (1) do not show any photobleaching steps, or (2) exhibit an increase in intensity during the experiment, which is not readily explained by a typical photobleaching process. 2. To visually inspect photobleaching traces for individual spots, run trace_inspector_GUI.m (Fig. 3c-iii). 3. subregion_stats.m returns photobleaching step counts in addition to colocalization, allowing the user to tabulate photobleaching data from only the portion of the dataset that contains signal above background.

4

Notes 1. In our hands, the best-performing fluorescent tags for SiMPull are: (a) Green/Yellow channel: mNeonGreen [9] (first choice) or mEYFP (second choice) fluorescent proteins. They provide good maturation, are readily detectable at singlemolecule levels, and have reliable bleaching. They are suitable for stoichiometry determination [1]. (b) Red channel: mScarlet-I [10] fluorescent protein. This is the best available red-emitting FP, but performs less well than mNG (see above) or HaloTag (see below). We therefore recommend using mScarlet-I only in 3-color experiments, when the green and far-red channels are occupied by mNG and HaloTag. mScarlet-I has reasonable maturation, although not as good as mNG or EYFP. It has some propensity to aggregate but can be used with caution for stoichiometry determination [1]. (c) Far red channel: HaloTag labeled with JaneliaFluor dyes [11, 12]. In our hands, JF646 dye performs best for in vivo labeling and exhibits excellent labeling efficiency [1]. However, we find that ~25% of molecules exhibit two-state photobleaching, making this dye unsuitable for stoichiometry determination. In summary, our recommendations are to use mNeonGreen plus HaloTag-JF646 for routine colocalization experiments, and mNeonGreen for stoichiometry experiments. 2. Thorough washing is essential for reproducible results. If residual unbound neutravidin or antibody is left in the channels at the end of the functionalization, it will compete with the desired binding to the surface in later steps.

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3. DMSO that is old, or lower-grade, contains impurities that are toxic to C. elegans. In our hands this is the most common cause of failed HaloTag labeling in liquid culture. 4. Needle puller program settings for embryo manipulation. (a) Heat ¼ Ramp Value (from needle puller ramp test). (b) Pull ¼ 155. (c) Velocity ¼ 100. (d) Time ¼ 150. 5. Optimizing laser power and exposure time is necessary for accurate spot detection and photobleaching step counting. Prepare a control sample from the mNG::Halo strain (LP539). Select a laser power and exposure time that allow single molecules to be visible without bleaching immediately. We typically use 20–50 ms exposures and 70% power when operating a 100 mW laser diode. Acquire 500 frames of data at a single stage position using the SiMPull acquisition plugin (set number of positions to (1). Then, scroll through the stack of 500 images and evaluate how quickly the spots are photobleaching by eye. The spots should photobleach steadily throughout the 500 frames. If a significant number of spots remain at the end, photobleaching may be incomplete. Increase the power and/or exposure to get accurate photobleaching step counts. Conversely, if the spots bleach too quickly (i.e., bleaching is complete far before the end of the acquisition) decrease laser power/exposure. Repeat for several iterations for each fluorophore until the timing looks to be in the correct range. Then, to evaluate photobleach time quantitatively, collect a complete dataset (several stage positions) with the settings you choose, and analyze the data by running analyze_batch.m. Then run bleachhalftime.m and select the . mat file for the relevant test dataset. This will plot a curve of spots in each frame over time. The bleach t1/2 should be near 100 frames (20 frames) for complete bleaching in 500 frames. Make further adjustments to laser power and exposure time accordingly. 6. When performing TIRF, the excitation light does not have uniform intensity across the field of view due to diffraction and the Gaussian profile of the laser beam. Additionally, TIRF excitation intensity declines exponentially with distance from the coverslip surface, and thus molecules that are not exactly equidistant from the glass will experience different illumination intensities. For these reasons, spot brightness/intensity is not a reliable measure of the number of molecules in a spot and is not used to report spot stoichiometry in our analysis.

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7. Creating an averaged image for spot counting amplifies signal to noise and eliminates transient spots for accurate spot detection. The range of frames to be averaged is specified in the “analyze_batch” dialogue window under “Frames to average for spot detection.” We typically exclude the first frame to eliminate artifacts caused by, for example, incomplete synchronization of laser light emission with camera exposure. We have empirically found that averaging frames 2–50 gives good spot detection results when the laser power and exposure time have been calibrated as described in Note 5. 8. To check if your copy of the SiMPull Analysis software is up to date, run the command ‘git status -uno’. If new updates are available, you’ll see a message like “Your branch is behind ‘origin/master’ by ## commits.” If you want to update to the latest version, run the command ‘git pull origin master.’ 9. Lysis is inherently variable, so expect some differences in the peak profile for each channel in an experiment. When lysing without flow, the best way to keep flow to a minimum is to ensure that the channel is well sealed with tape or valap. 10. Although stoichiometry determination via photobleaching is reproducible and biologically informative, it carries some caveats that a user should be aware of; in particular, due to incomplete fluorophore maturation and simultaneous bleaching of multiple molecules, photobleaching systematically underreports the actual stoichiometry of molecular complexes. For more detailed information about this effect, see [1, 13].

Acknowledgments This work was supported by NIH K99/R00 GM115964 and R01 GM138443 (DJD) and by grants from the Helen Hay Whitney Foundation and Mallinckrodt Foundation (DJD). DJD is a CPRIT Scholar supported by Cancer Prevention and Research Institute of Texas grant RR170054. References 1. Dickinson DJ, Schwager F, Pintard L, Gotta M, Goldstein B (2017) A single-cell biochemistry approach reveals PAR complex dynamics during cell polarization. Developmental Cell 42:416–434.e11. https://doi. org/10.1016/j.devcel.2017.07.024

2. Dickinson DJ, Pani AM, Heppert JK, Higgins CD, Goldstein B (2015) Streamlined genome engineering with a self-excising drug selection cassette. Genetics 200:1035–1049. https:// doi.org/10.1534/genetics.115.178335

Single-Cell Single Molecule Pull-Down (sc-SiMPull) 3. Dickinson DJ, Goldstein B (2016) CRISPRbased methods for Caenorhabditis elegans genome engineering. Genetics 202:885–901. https://doi.org/10.1534/genetics.115. 182162 4. Ghanta KS, Mello CC (2020) Melting dsDNA donor molecules greatly improves precision genome editing in Caenorhabditis elegans. Genetics 216:643–650. https://doi.org/10. 1534/genetics.120.303564 5. Sui G, Wang J, Lee C-C, Lu W, Lee SP, Leyton JV, Wu AM, Tseng H-R (2006) Solution-phase surface modification in intact poly(dimethylsiloxane) microfluidic channels. Anal Chem 78: 5543–5551. https://doi.org/10.1021/ ac060605z 6. Padeganeh A, Ryan J, Boisvert J, Ladouceur A-M, Dorn JF, Maddox PS (2013) Octameric CENP-A nucleosomes are present at human centromeres throughout the cell cycle. Curr Biol 23:764–769. https://doi.org/10.1016/ j.cub.2013.03.037 7. Ensign DL, Pande VS (2010) Bayesian detection of intensity changes in single molecule and molecular dynamics trajectories. J Phys Chem B 114:280–292. https://doi.org/10.1021/ jp906786b 8. Blischak JD, Davenport ER, Wilson G (2016) A quick introduction to version control with Git and GitHub. PLoS Comput Biol 12: e1004668. https://doi.org/10.1371/journal. pcbi.1004668

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9. Shaner NC, Lambert GG, Chammas A, Ni Y, Cranfill PJ, Baird MA, Sell BR, Allen JR, Day RN, Israelsson M, Davidson MW, Wang J (2013) A bright monomeric green fluorescent protein derived from Branchiostoma lanceolatum. Nat Methods 10:407–409. https://doi. org/10.1038/nmeth.2413 10. Bindels DS, Haarbosch L, van Weeren L, Postma M, Wiese KE, Mastop M, Aumonier S, Gotthard G, Royant A, Hink MA, Gadella TWJ (2017) mScarlet: a bright monomeric red fluorescent protein for cellular imaging. Nat Methods 14:53–56. https://doi. org/10.1038/nmeth.4074 11. Grimm JB, English BP, Chen J, Slaughter JP, Zhang Z, Revyakin A, Patel R, Macklin JJ, Normanno D, Singer RH, Lionnet T, Lavis LD (2015) A general method to improve fluorophores for live-cell and single-molecule microscopy. Nat Methods 12:244–250. https://doi.org/10.1038/nmeth.3256 12. Grimm JB, Muthusamy AK, Liang Y, Brown TA, Lemon WC, Patel R, Lu R, Macklin JJ, Keller PJ, Ji N, Lavis LD (2017) A general method to fine-tune fluorophores for live-cell and in vivo imaging. Nat Methods 14: 987–994. https://doi.org/10.1038/nmeth. 4403 13. Hines KE (2013) Inferring subunit stoichiometry from single molecule photobleaching. J Gen Physiol 141:737–746. https://doi.org/ 10.1085/jgp.201310988

Chapter 5 Biochemical Assays to Detect Activation of Small GTPases Rho, Rac, and Cdc42 during Morphogenesis Mark L. Berns and Raymond Habas Abstract Wnt/Frizzled (Fz) signaling controls developmental, physiological, and pathological processes through several distinct pathways. Wnt/Fz activation of the small GTPases Rho, Rac, and Cdc42, is one key mechanism that regulates cell polarity and migration during vertebrate gastrulation. In this chapter, we describe biochemical assays for detection of Wnt/Fz-mediated activation of Rho, Rac and Cdc42 in both mammalian cells and Xenopus embryo explants. Key words Noncanonical Wnt signaling, Rac, Rho, Cdc42, Gastrulation, Xenopus

1

Introduction The Wnt family of secreted proteoglycans play central roles during embryonic development and is deregulated in a number of human diseases [1, 2]. Although Wnt signaling via the canonical β-catenin pathway has been most intensively studied for cell fate determination, cell proliferation, and breast and colorectal cancers [3, 4]. Wnt activation of the Rho family of GTPases has received increased attention. This so-called noncanonical Wnt signaling pathway or β-catenin independent pathway is essential for cell polarity and movements during vertebrate gastrulation and shares common components similar to those involved in the establishment of planar cell polarity (PCP) in Drosophila, thus it is also often referred to as the Wnt/PCP pathway (Fig. 1) [5]. In addition, Wnt proteins can also activate other noncanonical signaling pathways, including calcium calmodulin–dependent kinase 2 (CaMK2), protein kinase C (PKC) [6], and protein kinase A (PKA) [7]. It is now accepted that the Fz family of seven-pass transmembrane receptors are cell surface receptors for Wnt proteins [8]. There are 19 Wnt genes and 10 Fz genes in both the mouse and human genomes. These sheer numbers and general lack of

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_5, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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Fig. 1 Schematic of several known noncanonical Wnt signaling cascades. In noncanonical or planar cell polarity (PCP), Wnt signaling is transduced through Fz independently of LRP 5/6. This pathway through Dishevelled mediates cytoskeletal changes through the small GTPases Rho and Rac, which in turn activate Rho kinase (ROCK) and Jun N-terminal K (JNK), respectively. For the Wnt/Ca2+ pathway, Wnt signaling by way of Fz mediates the activation of G-proteins to activate PLC and PKC. There is disagreement about the involvement of Dishevelled in this pathway

purified soluble Wnt proteins (except for Wnt-3a and Wnt-5a) have hindered the study of Wnt-Fz interaction and specificity, which is poorly understood in vertebrates [9, 10]. In addition to Fz, several other families of cell surface receptors, including low-density lipoprotein (LDL) receptor-related proteins, 5 and 6 (LRP5/6), Ryk (an atypical receptor tyrosine kinase) and Strabismus are also important for Wnt signal transduction, highlighting the complexity and versatility of the Wnt signaling system [11–13]. Although LRP5/6 are established as coreceptors for the Fz proteins and required specifically for the Wnt/β-catenin pathway (Fig. 2) [11], proteoglycans seem to be involved in extracellular Wnt ligand transport and distribution, thus they potentially affect many Wnt pathways [14]. Wnt-11/Fz-7-mediated endocytosis requires Ryk to regulate convergent extension movements in Xenopus [15]. Attempts have been made to classify Wnt molecules into one of two groups: (1) the canonical subfamily that includes Wnt-1, Wnt-3a, and Wnt-8 “only” activates the β-catenin pathway or (2) the noncanonical subfamily that activates β-catenin–independent pathways, such as Wnt-4, Wnt-5a, and Wnt-11. However, accumulating evidence suggests that such a classification may be oversimplified, and many Wnt proteins, including Wnt-1, Wnt-3a,

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Fig. 2 Schematic of Canonical Wnt signaling. Signaling through the Fz receptor, LRP 5/6 coreceptor complex, and Disheveled induce stabilization of β catenin, which then translocates into the nucleus via its interaction with Custos where it complex with LEF/TCF family members to mediate transcriptional induction of target genes

Wnt-5a, and Wnt-11, can activate multiple pathways in different experimental contexts, likely depending on receptor complements (including Fz and other receptors) and other cofactors with which these Wnt proteins may interact [16–21]. Thus, with perhaps a few exceptions, there are no clear rules yet that faithfully predict Wnt or Fz specificity for the activation of a particular pathway in vertebrates. Despite these caveats, it is well established that activation of the Rho family of GTPases by Wnt/Fz signaling is critical for vertebrate development and is involved in tumor development and progression [22, 23]. The Rho family of GTPases plays important roles in regulating cytoskeletal architectures associated with cell polarity and motility. The spatiotemporal activation of Rho GTPases is coordinated by intricate epigenetic networks and protein-protein interactions. Phosphorylation of Rho GTPases signals various regulatory mechanisms, such as sequestration, subcellular localization, and protein degradation [22]. The prototypes of this family, Rho, Rac, and Cdc42, act as bimolecular switches cycling between the active guanosine triphosphate (GTP)-bound form and the inactive guanosine diphosphate (GDP)-bound form, with the GTP-bound form interacting with downstream effectors

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[24, 25]. Rho, Rac, and Cdc42 are responsible for the regulation, assembly, and organization of the actin cytoskeleton in eukaryotic cells. Rho controls the assembly of actin/myosin filaments to generate contractile forces, whereas Rac and Cdc42 promote actin polymerization at the cell periphery to generate protrusive forces, in the form of lamellipodia and filopodia, respectively [26]. ROCK1, a downstream effector of GTP-bound Rho, phosphorylates the small actin-binding protein, profilin 1 (PFN1) at Ser-137, maintaining it in its active state [27]. The roles of PFN1 and profilin 2 (PFN2) in tumorigenesis are nonredundant and the basis of continuing study as small-molecule drug targets. Knockdown experiments of PFN1 and PFN2 in breast cancer cells produced contrary effects on metastasis and invasion; suppression of PFN1 yielded decrease metastasis and knockdown of PFN2 increased metastasis [28]. Overexpression of PFN1 resulted in increased invasion and protrusive activity, whereas overexpression of PFN2 diminished protrusive activity and suppressed migration and invasion into the matrix [28]. PFN1 and PFN2 have vital nonredundant roles during gastrulation; PFN1 regulates neural tube closure whereas PFN2 controls convergent extension [25, 29]. Rearrangement of the actin cytoskeleton via Wnt/Fz activation of the Rho family of GTPases is vital not only in its role in tumorigenesis but also in mediation of cell movement during early development. During Xenopus embryogenesis, Wnt-11 and its corresponding receptor, Fz7, activate both RhoA and Rac to regulate convergent extension (CE) movements that are a major driving force of gastrulation. During CE movements, cells in the dorsal mesoderm polarize, elongate, and align along the mediolateral axis and intercalate among one another, resulting in the mediolateral narrowing (convergence) and anteroposterior lengthening (extension) of the axis of the embryo [30, 31]. Wnt-11 activation of RhoA and Rac can be demonstrated in dorsal embryo explants, in which interfering with Wnt-11 or Fz function prevents RhoA and Rac activation [17, 32]. Conversely, overexpression of Wnt-11 or Xenopus Fz7 (Xfz7) in the embryo ventral region, which neither exhibits CE movements nor expresses Wnt-11 and Xfz7, is sufficient to activate RhoA and Rac [17, 32]. Wnt/Fz activation of Rho and Rac also has been observed in several commonly used mammalian cell lines [16, 20, 32]. In these experiments, transfection of Wnt-1 or Wnt-3a cDNA, of certain (but not all) Fz cDNA, or treatment with Wnt-1- or Wnt-3a–conditioned medium results in RhoA and Rac activation. These observations in vertebrate embryos/mammalian cells followed and paralleled earlier genetic studies in Drosophila that showed that Fz function in PCP relies on RhoA and Rac gene function [33]; thus, Wnt/Fz signaling to RhoA and Rac is conserved, although it remains unknown whether a Wnt ligand is required for PCP in flies. Wnt/Fz activation of RhoA and Rac

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requires the cytoplasmic phosphoprotein Dishevelled (Dsh), which often acts downstream of Fz [34]. In this chapter, we describe biochemical assays used to investigate the activation of Rho, Rac, and Cdc421 GTPases in mammalian culture cells and in the Xenopus embryo. These assays utilize a glutathione S-transferase (GST) pull-down strategy using fusion proteins that specifically bind to the activated/GTP-bound forms of Rho, Rac, and Cdc42. For Rho assays, a Rho-binding fragment of the Rho-effector Rhotekin is fused to GST and termed GSTRBD. For the Rac and Cdc42 assays, the Rac and Cdc42 binding fragment of p21 (PAK) is fused to GST and termed GST-PBD (Fig. 3). The GST-RBD and GST-PBD fusion proteins are produced in bacterial cells, purified and incubated with cell lysates derived from either mammalian cells or Xenopus embryo explants (Fig. 4). GST-RBD and GST-PBD bind specifically to the GTP-bound forms of Rho, Rac, or Cdc42, respectively, which are precipitated using glutathione-agarose beads and detected by conventional immunoblotting. The following protocols provide an efficient method to study the activation of the small GTPases Rho, Rac, and Cdc42 both in vitro and in vivo.

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Materials

2.1 Bacteria Cell Culture

1. BL21 bacterial cells. 2. 1 M IPTG. 3. LB-ampicillin plates: 1% Bacto tryptone, 0.5% yeast extract, 1% NaCl, 100 mg/mL ampicillin, 1.5% Bacto agar. 4. LB-ampicillin growth media: 1% Bacto tryptone, 0.5% yeast extract, 1% NaCl, 100 μg/mL ampicillin. 5. 1 PBS: 1.54 mM KH2PO4, 155.17 mM NaCl, 2.71 mM Na2HPO4·7H2O (pH 7.2). 6. Glutathione Sepharose beads. 7. 1 PBS/10 mM DTT/1% Triton X-100. 8. Protease inhibitor cocktail. 9. 50 mg/mL lysozyme; 10% Triton X-100. 10. 1 M MgCl2. 11. 10 mg/mL DNase1.

2.2 Mammalian Cell Culture

1. HEK293T cells. 2. 10% FBS in DMEM supplemented with 1% penicillin and streptomycin 3. Polyfect Transfection Reagent (Qiagen).

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Fig. 3 Schematic representation of the Rho, Rac, and Cdc42 assays. For in vitro experiments, cells were transfected and cultured for 12–24 h and were subsequently lysed for binding studies. For in vivo experiments, Xenopus embryos are injected with RNA at stage 3 and explants are removed at stage 10.5. They are lysed and incubated with GST proteins. Samples are then incubated with either GST-RBD or PBD and a GST pulldown assay is performed. Samples are then resolved on an SDS-PAGE gel and subject to Western blotting

4. 1 Trypsin 5. 1 PBS. 2.3

Embryos

1. Xenopus laevis embryos. 2. 10 MMR: 1 M NaCl, 20 mM KCl, 20 mM CaCl2, 10 mM MgCl2, 50 mM HEPES, pH to 7.6.

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Fig. 4 Rho, Rac, and Cdc42 assays in Xenopus. (a) Embryos are injected at the four-cell stage into the two dorsal or ventral cells and allowed to develop until stage 10.5. The DMZ and VMZ are then explanted and subjected to Rho, Rac, or Cdc42 pull-down assays. (b) Examples of Rho, Rac, and Cdc42 assays performed from DMZs and VMZs. Rho, Rac, and Cdc42 activation detected from GST-RBD or GST-PBD pull-down assays show Rho and Rac but not Cdc42 being preferentially in the DMZ. Endogenous Rho, Rac, and Cdc42 levels are shown in the lysate samples for comparison

3. 3% Ficoll–0.5 MMR. 4. 1% BSA–0.5 MMR. 2.4 GST-PRD and GST-PBD Binding Assay Buffers

1. Rho Lysis Buffer: 50 mM Tris–HCl pH 7.2, 500 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholic acid, 0.1% SDS, 10 mM MgCl2, and 1  protease inhibitors (added fresh each time). 2. Rho Wash Buffer: 50 mM Tris–HCl pH 7.2, 1% Triton X-100, 150 mM NaCl, 10 mM MgCl2, and 1  protease inhibitors. 3. Rac and Cdc42 Lysis Buffer: 50 mM Tris, pH 7.5, 200 mM NaCl, 2% NP40, 10% glycerol, 10 mM MgCl2, and 1  protease inhibitors. 4. Rac and Cdc42 Wash Buffer: 25 mM Tris, pH 7.5, 40 mM NaCl, 1% NP40, 30 mM MgCl2, and 1  protease inhibitors.

2.5

Western Blotting

1. 12% SDS-PAGE gel. 2. Running Buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS. 3. Transfer Buffer: 25 mM Tris, 192 mM glycine, 20% methanol. 4. 1 PBST: 1  PBS, 0.5% Tween 20.

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5. 2 sample buffer; 125 mM Tris–HCl (pH 6.8), 10% 2-mercaptoethanol, 4% SDS, 20% glycerol. 6. 5% Nonfat dry milk. 7. Rho monoclonal and polyclonal antibodies (Santa Cruz). 8. Rac/Cdc42 monoclonal antibodies (Transduction Labs). 9. SuperSignal PicoWest ECL (Pierce).

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Methods

3.1 Rho and Rac/ Cdc42 Assays 3.1.1 Preparation of Recombinant GST-RBD Protein

1. Grow an overnight culture of a single colony of BL21 bacterial cell containing the GST-RBD plasmid in 20 mL LB-amp (100 mg/L) at 30  C. 2. Dilute the culture into 1 L of LB-amp (100 mg/mL) and grow at 30  C until the optical density at 600 nm is 1.0. This takes 5–7 h, depending on starting optical density. 3. Induce the bacterial culture with 1 mL of 1 M IPTG and incubate for 3–4 h at 30  C. 4. Aliquot the bacteria into 50-mL Falcon tubes and spin at 4000 rpm for 10 min to pellet the bacteria. Discard supernatant and flash freeze the pellets in liquid nitrogen. 5. Store the pellets at 1 year.

3.1.2 Preparation of Recombinant GST-PBD Fusion Protein

80  C. The pellets are stable for up to

1. Grow an overnight culture of a single colony of BL21 bacterial cell containing the GST-PBD plasmid in 20 mL LB-amp (100 mg/mL) at 30  C. 2. Dilute the culture into 1 L of LB-amp (100 mg/mL) and grow at 30  C until the optical density at 600 nm is 1.0. This takes 5–7 h, depending on starting optical density. 3. Induce the bacterial culture with 1 mL of 1 M IPTG and incubate for 3–4 h at 30  C. 4. Lyse the cells in 1 PBS and protease inhibitors using either sonication or a French press. 5. Pellet the lysate by spinning at 15,000 rpm for 15 min and isolate the supernatant. 6. Aliquot the supernatant into 1.5-mL Eppendorf tubes and flash freeze in liquid nitrogen. 7. Store at

3.1.3 Extraction of GSTRBD

80  C.

1. Prepare the glutathione Sepharose beads by swelling approximately 100 μL with 1 PBS–10 mM DTT–1% Triton X-100 for at least 30 min on ice, then wash three times with 500 μL of 1 PBS–10 mM DTT–1% Triton-X 100. After the final wash,

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the beads can be stored on ice as a 1 slurry. Do not spin beads higher than 3000 rpm during the pelleting and washing stages because this will damage the beads. Also, prepare two tubes of beads, because you will have 2 mL of lysate. 2. Thaw one aliquot of frozen GST-RBD pellet on ice and resuspend in 2 mL 1 PBS. 3. Add 20 μL 1 M DTT, 20 μL protease inhibitor cocktail (Boehringer-Mannheim), and 40 μL of 50 mg/mL lysozyme. 4. Vortex briefly to mix well and incubate on ice for 30 min. 5. Add 225 μL, 10% Triton X-100, 22.5 μL, 1 M MgCl2, and 22.5 μL of 10 mg/mL DNase1. 6. Vortex briefly to mix well and incubate on ice for 30 min. 7. Spin at 14,000 rpm at 4  C for 2 min, and add 1 mL supernatant to each of the two tubes of the preswollen beads. 8. Incubate on a Nutator at 4  C for 45 min (do not exceed 1 h). 9. Spin and wash beads three times with 500 μL of 1 PBS–10 mM DTT–1% Triton X-100. After the final wash, store on ice in 1 slurry with the final volume approximately 500 μL (see Notes 1–3). 3.1.4 Extraction of GSTPBD

1. Prepare the glutathione Sepharose beads by swelling approximately 100 μL with 1 PBS–10 mM DTT–1% Triton X-100 for at least 30 min on ice, then wash three times with 500 μL of 1 PBS–10 mM DTT–1% Triton X-100, and after final wash, store on ice as a 1 slurry. Do not spin beads higher than 3000 rpm during the pelleting and washing stages, because this will damage the beads. Also prepare two tubes of beads, because you will have 1 mL of bacterial lysate. 2. Thaw one aliquot of frozen bacterial supernatant on ice. 3. Add 500 μL supernatant to each of the two tubes of the preswollen beads. 4. Incubate on a Nutator at 4  C for 45 min and do not exceed 1 h. 5. Spin and wash beads three times with 500 μL of 1 PBS– 10 mM DTT–1% Triton X-100. After the final wash, store on ice in 1 slurry with the final volume approximately 500 μL (see Notes 1–3).

3.1.5 Sample Preparations for Rho PullDown Assays Using Mammalian Cells

1. Mammalian HEK293T cells are cultured in 6-well plates (30 mm) in 10% fetal bovine serum and DMEM media supplemented penicillin–streptomycin until 60% confluency. 2. One microgram of cDNA is transfected into the cells using standard calcium phosphate method. 3. Media is changed 12 h after transfection, and cells are incubated for 12–24 h.

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3.1.6 Sample Preparations for Rac/Cdc42 Assay Using Mammalian Cells

1. Mammalian HEK293T cells are cultured in 6-well plates (30 mm) in 10% fetal bovine serum and DMEM media supplemented penicillin–streptomycin until 60% confluency. At this point the cells are changed into 0.5% fetal bovine serum and DMEM media supplemented penicillin–streptomycin. This step helps to reduce the basal level of activated Rac/ Cdc42. 2. Six hours after media is changed to 0.5% sera, 1 μg of cDNA is transfected into the cells using standard calcium phosphate method. 3. Media is changed 12 h after transfection, and cells are incubated for an additional 12–24 h in 0.5% fetal bovine serum and DMEM media supplemented with penicillin–streptomycin.

3.1.7 Sample Preparations for Rho/Rac/ Cdc42 Assay Using Xenopus Embryos and Explants

1. Xenopus embryos are injected at the 4-cell stage into the two dorsal cells (for dorsal marginal zone [DMZ] explants) or into the two ventral cells (for ventral marginal zone [VMZ] explants) in 3% Ficoll/0.5  MMR. 2. Two hours after injections, embryos are changed into 0.1  MMR and cultured to stage 10.5. 3. The vitelline membrane is removed from the embryo, and DMZ or VMZ is explanted using forceps. Explants are pooled and stored on ice until they are lysed. All embryos are dissected on agarose-coated culture dishes in a solution of 0.5  MMR/ 1% BSA. Xenopus embryos and explants are handled as described elsewhere [35].

3.1.8 GST-RBD and GSTPBD Binding Assay

1. Lyse the cells in 500 μL of Rho or Rac lysis buffer and to each sample add 10 μL of 10 mg/mL DNase1 solution. Incubate on ice for 10 min and then spin samples at 14,000 rpm at 4  C. For Xenopus explants, we typically use 50 explants (DMZ or VMZ) for each sample. 2. Remove 25 μL of supernatant and add 25 μL of 2 sample buffer, heat at 90  C for 5 min and store. This is your whole cell lysate for control immunoblotting. 3. Remove the remaining 475 μL of supernatant and add to tubes containing approximately 50 μL of GST-beads coupled to the RBD or to PBD (see Note 3). 4. Incubate on a nutator at 4  C for 1 h, and wash three times with Rho or Rac wash buffer. After final wash, resuspend in 50 μL of 2 sample buffer and heat at 90  C for 5 min. 5. Perform Western blotting.

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3.2 Western Blot Analysis

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1. Resolve samples on a 12% SDS-PAGE gel and run until the bromophenol dye is approximately 1 in. from the bottom of the gel. 2. Transfer to nitrocellulose membrane, which is incubated with 10% nonfat dry milk in 1 PBST for 30 min at room temperature. 3. Wash thrice for 5 min with 1 PBST. 4. Incubate with the primary antibody (Rho monoclonal, Santa Cruz, for mammalian cell extracts or Rho polyclonal, Santa Cruz, for Xenopus explant extracts; Rac or Cdc42 monoclonal, Transduction Labs, for both mammalian cell and Xenopus explant extracts) at a 1:500 dilution for 1 h at room temperature or overnight at 4  C. 5. Wash once for 15 min and four times for 5 min with 1 PBST. 6. Incubate with secondary antibody at 1:5000 solution for 1 h at room temperature. 7. Wash once for 15 min and four times for 5 min with 1 PBST. 8. Perform ECL reaction. 9. The endogenous Rho/Rac/Cdc42 are detectable within 1 min in total lysates or 5 min in the GST-RBD/PBD pull-down samples for mammalian cell extracts. For Xenopus explant extracts, the endogenous Rho/Rac/Cdc42 are detectable within 3–5 min in lysates or 10 min in the GST-RBD/PBD pull-down samples.

4

Notes 1. For each preparation of GST-RBD or GST-PBD bacteria, use one aliquot and follow the protocol to extract the fusion protein and resolve a fraction of the sample on a 12% SDS-PAGE gel and Coomassie blue stain. GST-RBD runs at approximately 35 kDa and GST-PBD approximately 37 kDa. If there is extensive degradation, redo the protein preparation. 2. Each 50-mL bacterial aliquot of GST-RBD yields approximately 200–300 μg of fusion protein, and each is enough for 10 samples for the GST-RBD assay. Each 1 mL aliquot of GST-PBD yields approximately 150–250 μg of protein, and each is enough for 10 samples for the GST-PBD assay. 3. Always keep samples on ice whenever possible and do not exceed wash times.

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15. Kim GH, Her JH, Han JK (2008) Ryk cooperates with frizzled 7 to promote Wnt11mediated endocytosis and is essential for Xenopus laevis convergent extension movements. J Cell Biol 182:1073–1082 16. Endo Y, Wolf V, Muraiso K, Kamijo K, Soon L, Uren A, Barshishat-Kupper M, Rubin JS (2005) Wnt-3a-dependent cell motility involves RhoA activation and is specifically regulated by dishevelled-2. J Biol Chem 280: 777–786 17. Habas R, Dawid IB, He X (2003) Coactivation of Rac and rho by Wnt/frizzled signaling is required for vertebrate gastrulation. Genes Dev 17:295–309 18. He X, Saint-Jeannet JP, Wang Y, Nathans J, Dawid I, Varmus H (1997) A member of the frizzled protein family mediating axis induction by Wnt-5A. Science 275:1652–1654 19. Kishida S, Yamamoto H, Kikuchi A (2004) Wnt-3a and Dvl induce neurite retraction by activating Rho-associated kinase. Mol Cell Biol 24:4487–4501 20. Qiang YW, Endo Y, Rubin JS, Rudikoff S (2003) Wnt signaling in B-cell neoplasia. Oncogene 22:1536–1545 21. Tao Q, Yokota C, Puck H, Kofron M, Birsoy B, Yan D, Asashima M, Wylie CC, Lin X, Heasman J (2005) Maternal Wnt11 activates the canonical Wnt signaling pathway required for axis formation in Xenopus embryos. Cell 120: 857–871 22. Hodge RG, Ridley AJ (2016) Regulating rho GTPases and their regulators. Nat Rev Mol Cell Biol 17:496–410 23. Mardilovich K, Olson MF, Baugh M (2012) Targeting Rho GTPase signaling for cancer therapy. Future Oncol 8:165–177. https:// doi.org/10.2217/fon.11.143 24. Raftopoulou M, Hall A (2004) Cell migration: Rho GTPases lead the way. Dev Biol 265: 23–32 25. Sato A, Khadka DK, Liu W, Bharti R, Runnels LW, Dawid IB, Habas R (2006) Profilin is an effector for Daam1 in non-canonical Wnt signaling and is required for vertebrate gastrulation. Development 133:4219–4231 26. Jaffe AB, Hall A (2005) Rho GTPases: biochemistry and biology. Annu Rev Cell Dev Biol 21:247–269 27. Shao J, Welch WJ, Diprospero NA, Diamond MI (2008) Phosphorylation of profilin by ROCK1 regulates polyglutamine aggregation. Mol Cell Biol 17:5196–5208

Biochemical Assay for Rho Family Small GTPases 28. Mouneimne G, Hansen SD, Selfors LM, Petrak L, Hickey MH, Gallegos LL, Simpson KJ, Lim J, Gertler FB, Hartwig JH, Mullins RD, Brugge JS (2012) Differential remodeling of actin cytoskeleton architecture by profilin isoforms leads to distinct effects on cell migration and invasion. Cancer Cell 5:615–630 29. Khadka DK, Liu W, Habas R (2009) Non-redundant roles for profilin 2 and profilin 1 during vertebrate gastrulation. Dev Biol 332: 396–406 30. Keller R (2002) Shaping the vertebrate body plan by polarized embryonic cell movements. Science 298:1950–1954 31. Wallingford JB, Fraser SE, Harland RM (2002) Convergent extension: the molecular control of polarized cell movement during embryonic development. Dev Cell 2:695–706

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32. Habas R, Kato Y, He X (2001) Wnt/frizzled activation of rho regulates vertebrate gastrulation and requires a novel Formin homology protein Daam1. Cell 107:843–854 33. Mlodzik M (2002) Planar cell polarization: do the same mechanisms regulate drosophila tissue polarity and vertebrate gastrulation? Trends Genet 18:564–571 34. Wharton KA Jr (2003) Runnin’ with the Dvl: proteins that associate with Dsh/Dvl and their significance to Wnt signal transduction. Dev Biol 253:1–17 35. Sive H, Grainger RM, Harland RM (2000) Dejellying embryos. Early development of Xenopus laevis: a laboratory manual, 1st edn. Cold Spring Harbor laboratory press, Cold Spring Harbor, Chapter 6

Chapter 6 Analysis of Planar Cell Polarity Complexes by Proximity Biotinylation in Xenopus Embryos Ilya Chuykin and Sergei Y. Sokol Abstract Understanding signaling processes operating in cells during development and disease requires extensive knowledge of protein interactions. Proximity-dependent biotinylation mediated by a promiscuous bacterial biotin ligase is a sensitive approach for evaluating protein interactions under physiological conditions. This technique allows for assessing protein association when conventional pull-down assays are not applicable due to high background or transient nature of the interaction. In contrast to many studies of proximity biotinylation in cultured cells, this protocol has been adapted to detect protein interactions in Xenopus embryos. Here, we apply this technique to evaluate planar cell polarity (PCP) complexes formed by Prickle3 and Vangl2, and show that Prickle3 fused to the N-terminal fragment of the biotin ligase from Aquifex aeolicus efficiently biotinylates Vangl2 in vivo. We present our step-by-step proximity biotinylation protocol that provides a reliable semiquantitative assay for protein interactions and highlights the use of Xenopus embryos as a model for biochemical studies. Key words Xenopus, BioID, Modified biotin ligase, Biotin, PCP, Vangl2, Prickle3

1

Introduction An important goal of modern experimental biology is to define physical protein interactions and reveal their roles at the cellular and organismal levels. To face this challenge, many protein–protein association assays have been developed that are based on direct physical binding. These approaches, however, are often restricted to binary interactions or require the preservation of the protein complex during in vitro affinity capture. To overcome these limitations, proximity-based approaches that explore the protein interactome directly in live cells have been developed. The BioID (biotin identification) method utilizes the ability of the mutated bacterial biotin ligase BirA* (BirA R118G) to promiscuously biotinylate proteins in its immediate vicinity [1–5]. First, it identifies proteins located within nanometer distances from known protein complexes [6]. Second, it is not restricted to binary interactions and can be

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_6, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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used to assess the composition of complexes containing multiple components potentially at the proteome-wide scale. Third, it detects weak and transient interactions in live cells and tissues. This technology has a potential to define interactomes for the spatially localized signaling complexes [7]. Genetic approaches and directed evolution of new biotinylation enzymes have extended this approach from in vitro experiments using cultured cells to the organismal level [8, 9]. However, it has remained unclear whether this technique can be efficiently applied to cold-blooded vertebrates, such as Danio rerio or Xenopus laevis. One example of an intricate protein interaction network are protein complexes that regulate planar cell polarity (PCP), a common property of developing epithelia. Initially discovered in Drosophila, PCP proteins have been also implicated in early morphogenetic processes in vertebrate embryos and causally connected to human syndromes and congenital defects [10– 13]. Being essential for tissue polarity, core PCP proteins segregate to different sides of each epithelial cell. Detailed mechanistic understanding of this process is lacking, awaiting the characterization of the composition and the dynamics of the PCP interactome. Previous work revealed striking colocalization of the core PCP proteins Vangl2 and Prickle3 at anterior cell boundaries in the Xenopus neural plate [14]. Although these core PCP proteins have been shown to associate in vitro [15, 16], our attempts to detect their binding in vivo by immunoprecipitation have not been successful. To overcome this problem, we have adapted proximity biotinylation to Xenopus embryos and used it to study spatially localized PCP signaling complexes in the neural plate [17] (Fig. 1). However, overexpressed BirA* constructs revealed considerable toxicity in Xenopus embryos (data not shown, and [18]). We then fused the core PCP protein Prickle3 to the N-terminal fragment of Aquifex aeolicus biotin ligase (BL), which is more efficient than E. coli BirA* in proximity biotinylation and lacks the N-terminal DNA-binding domain (Fig. 2a) [19]. The fused fragment (amino acids 3–185) also lacks the conserved C-terminal domain of unknown function (amino acids 186–233) (Fig. 2a, b). When the C-terminal domain of BL has been replaced by Prickle3, the fusion protein (referred to as BLN-Pk3) remained enzymatically active when assessed by self-biotinylation in Xenopus embryos from late blastula stages in the presence of biotin [17] (Fig. 3). Moreover, BLN-Pk3 biotinylated its known binding partner Vangl2 and Par3/Pard3, a newly identified Prickle3-interacting protein [17]. Despite concerns related to potentially low activity of bacterial biotinylation enzymes at low temperatures and the long time required for labeling with biotin in vitro, Vangl2 was efficiently biotinylated by BLN-Pk3 at temperatures ranging from 13  C to 24  C (Fig. 3). Supporting specificity, Vangl2 lacking the Prickle3binding domain has remained unmodified (data not shown). These

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Fig. 1 Proximity biotinylation in Xenopus embryos. RNAs encoding BLN-Pk3 and HA-Vangl2 are coinjected with 0.2–0.4 mM biotin into 2–4 animal blastomeres at the 4–8-cell stage. Alternatively, 20 nl of 0.4–0.8 mM biotin can be injected into the blastocoel of embryos at the midblastula stage. BLN-Pk3 biotinylates the associated proteins including Vangl2. Embryos can be cultured until neurula stages at various temperatures 13–25  C and protein biotinylation is analyzed in the lysates after the pull-down and immunoblotting

observations demonstrate that proximity biotinylation can be applied for the analysis of the composition and dynamics of PCP protein complexes in vivo. This chapter describes our step-by-step biotinylation protocol that assesses the interaction between BLN-Prickle3 and Vangl2 in Xenopus embryos.

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Materials

2.1 Xenopus Culture and Manipulation

1. In vitro fertilized Xenopus embryos at various developmental stages. 2. Marc’s Modified Ringer’s solution (MMR): 100 mM NaCl, 2 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 5 mM HEPES, pH 7.5 [20]. Prepare 10 MMR stock and adjust pH with sodium hydroxide to 7.5, sterilize by autoclaving. 3. Injection solution: 3% Ficoll 400 (GE Healthcare) in 0.6 MMR. 4. Gentamicin sulfate (Sigma, stock 50 mg/ml); the working concentration is 50 μg/ml in 0.1 MMR. 5. Dissecting stereomicroscopes (Leica, Nikon, Zeiss). 6. Nitrogen gas–driven microinjector IM-300 (Narishige).

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Fig. 2 The enzyme fragment used for the biotinylation assay in Xenopus embryos. (a) Domain structures of biotin ligases (BL) from A. aeolicus and E. coli (also known as BirA*). A. aeolicus BL lacks the region corresponding to the N-terminal 82 amino acid DNA binding domain of BirA*. (b) The pairwise sequence alignment (EMBL-EBI) of the BL from A. aeolicus and E. coli shows identical amino acids (|) or those with low (.) or high similarity (:); spaces indicate gaps or mismatches. The conserved R mutated to G is indicated by red asterisk. The N-terminal fragment (BLN), highlighted in red, does not contain amino acids 185–232 corresponding to a conserved domain of unknown function

7. Micromanipulator (Narishige M152 or Singer instruments Mk1). 8. Low-temperature embryo incubator (VWR, Model 2005). 9. Glass capillaries, thin wall, 1 mm in diameter (World Precision Instruments, TW100F-4) for injection needle preparation. 10. Pipet puller (David Kopf Instruments, Model 730). 11. Micropipette grinder (Narishige). 12. Petri plates and tissue culture trays (6-well and 12-well) for embryo culture. 2.2 Proximity Biotinylation and Protein Detection

1. FLAG-BLN-Pk3 RNA synthesized in vitro from pCS2-FLAGBLN-Pk3. This plasmid encodes a fragment (N3-L185) of promiscuous BL from A. aeolicus [19] fused in-frame to the N-terminus of Xenopus laevis Prickle3.

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Fig. 3 Biotinylation of Vangl2 by BLN-Pk3 in Xenopus embryos. (a) Vangl2 is biotinylated by BLN-Pk3 in stage 13 embryos. (b) Vangl2 RNA was coinjected with different amounts of BLN-Pk3 RNA as indicated. Biotindependent self-biotinylation of BLN-Pk3 is visible. Vangl2 biotinylation signal is higher at stage 13 as compared to stage 9. Embryos were injected with 500 pg or 50 pg of HA-Vangl2 mRNA to achieve equal HA-Vangl2 protein levels at stages 9 and 13, respectively. Total levels of BLN-Pk3 protein in (b) were assessed with anti-Pk3 antibody. Protein levels are shown by immunoblotting with anti-biotin, anti-HA, and anti-Pk3 antibodies. Endogenous protein bands recognized in lysates by the anti-biotin (a) and anti-HA antibody (b) indicate equal loading (asterisks)

2. HA-Vangl2 RNA synthesized in vitro from pCS2-HA-Vangl2 encoding Xenopus laevis HA-tagged Vangl2 [21]. 3. Biotin (Sigma), stock in DMSO 80 mM, store at

20  C.

4. Protein A Sepharose (GE Healthcare). 5. Goat anti-biotin antibody, conjugated to horseradish peroxidase (HRP) (Cell Signaling, 7075, 1:1000–1:3000). 6. Anti-FLAG mouse monoclonal antibody (Sigma, M2, 1: 1000). 7. Anti-HA mouse monoclonal antibody (12CA5, 1:1000). 8. Anti-HA rabbit polyclonal antibody (Bethyl Labs, A190-108A, 1:5000). 9. HRP-conjugated secondary antibodies; goat anti-mouse IgG (Jackson ImmunoResearch, 709-036-1, 1:5000), goat antirabbit IgG (Jackson ImmunoResearch, 115-035-003, 1: 5000), mouse anti-goat IgG (Santa Cruz, sc2354, 1:5000). 10. Embryo lysis buffer: 50 mM Tris–HCl at pH 7.5, 50 mM NaCl, 1 mM EDTA, 1% Triton X-100, 1 mM Na3VO4, 10 mM NaF, containing cOmplete Mini EDTA-free protease inhibitor cocktail (Roche).

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11. 2 Sample buffer: 0.125 M Tris–HCl at pH 6.8, 10% β-mercaptoethanol, 20% glycerol, 4% SDS, 0.004% bromophenol blue. 12. Transfer buffer (TB): 25 mM Tris; 190 mM Glycine; 20% methanol. 13. Tris-buffered saline (TBS): 50 mM Tris–HCl pH 7.5 and 150 mM NaCl. 14. TBST: TBS with 0.05% Tween 20 (Santa Cruz). 15. Immobilon P membrane (PVDF, Millipore). 16. Luminol: 250 mM stock in DMSO (store at

20  C).

17. Paracoumaric acid: 90 mM stock in DMSO (store at

20  C).

18. Hydrogen peroxide (30%). 19. 0.1 M Tris–HCl, pH 8.8. 20. Standard polyacrylamide gels and the running buffer as described [22]. 21. Rotating shaker. 22. Power supply. 23. Bio-Rad minigel apparatus for protein separation by electrophoresis. 24. Bio-Rad mini-TransBlot cell for protein transfer. 25. ChemiDoc MP imager (Bio-Rad) for chemiluminescence.

3

Methods

3.1 Xenopus Embryo Culture and Microinjections

3.2

Biotinylation

Detailed protocols for egg in vitro fertilization and injections of Xenopus embryos have been described previously [23]. In vitro– fertilized eggs were cultured in 0.1 MMR [20] as described [24]. Staging is according to [25]. RNAs for microinjections are synthesized in vitro using mMessage mMachine kit (ThermoFisher). 1. Prepare injection needles from glass capillaries, 1 mm in diameter (World Precision Instruments, TW100F-4) using a pipet puller (David Kopf Instruments, Model 730). Each capillary makes two injection needles. 2. For microinjections, transfer four-cell embryos into 1 ml drops of the injection solution (3% Ficoll, 0.6x MMR) on a flipped 10 cm petri dish. 3. Inject each animal blastomere of 4–8-cell embryos using Narishige IM300 microinjector with 10 nl of the solution containing 0.1–0.5 ng of FLAG-BLN-Pk3 and HA-Vangl2 RNAs (see Note 1) and 0.2–0.4 mM biotin (see Notes 2 and 3) (Fig. 1) [18].

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4. Keep the embryos in the injection solution for 2 h at 13  C or 22  C until the injection wounds heal, then transfer them into 0.1% MMR until the desired stage (stages 10–16). No differences have been observed in biotinylation efficiency for embryos cultured at 13  C versus 22  C. 3.3 Immunoprecipitation

1. Lyse the injected embryos in the lysis buffer (50 mM Tris–HCl at pH 7.5, 50 mM NaCl, 1 mM EDTA, 1% Triton X-100, 1 mM Na3VO4, 10 mM NaF), containing cOmplete Mini EDTA-free protease inhibitor cocktail (Roche). To prepare lysates, use 10–20 μl of the lysis buffer per embryo. Typically, there are 20–40 injected embryos per experimental group. 2. Clear the lysates by centrifugation at 13, 500  g for 5 min in an Eppendorf centrifuge at room temperature (RT). Save 20–40 μl of lysates for immunoblotting before immunoprecipitation (see Note 4). 3. Add 10–20 μl of anti-HA antibodies (12CA5 hybridoma supernatant) to the lysates. Incubate for 2 h at RT or overnight at 4  C and then with 4 μl of Protein A Sepharose (GE Healthcare) for another 2 h at 4  C on a rotating shaker. 4. Wash the beads in the lysis buffer two times, 5 min each, and pellet by centrifugation in an Eppendorf centrifuge at low speed (900  g ) for 2 min. 5. Add an equal volume of 2 gel loading buffer, heat samples for 5 min at 95  C and separate proteins on a polyacrylamide gel using standard protocols [22]. Avoid boiling the samples in the sample buffer to detect transmembrane proteins.

3.4

Immunoblotting

1. Cut out a piece of 9  7 cm2 of the Immobilon P membrane (PVDF, Millipore) and prewet it by immersing into methanol for a few seconds. Place it into the transfer buffer (TB). 2. Remove the gel from the chamber and rinse it in TB for 1–5 min. 3. Assemble the transfer “sandwich”; place the gel on the top of the Whatman 3MM blotting paper prewetted in TB on a fiber pad. Place wet PVDV membrane on the top and remove any bubbles between the membrane and the gel. Put a second sheet of the 3MM paper on the top of the “transfer sandwich.” 4. Insert the “sandwich” into the TransBlot cell in the correct orientation (with the membrane proximal to the anode), fill it up with chilled TB (see Note 5). 5. Transfer proteins from the gel to the membrane at 40 V overnight or 100–110 V for 1–2 h.

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6. Disassemble the “sandwich” and block the PVDV membrane with 3% BSA (A709, Sigma) (see Note 6) in TBS for 1 h. Briefly wash with TBS. 7. Incubate with primary anti-tag antibody or HRP-conjugated goat anti-biotin antibody (1:1000–1:3000) in TBST containing 0.1% BSA for 2 h at RT or overnight at 4  C. Anti-HA and anti-FLAG antibody solutions do not include BSA, milk or other blocking reagents (see Note 7). 8. Wash with TBST 3–5 times (10 min each). 9. Incubate with the appropriate secondary antibody conjugated with HRP: goat anti-mouse IgG, goat anti-rabbit IgG, or mouse anti-goat IgG (to enhance the signal produced by HRP-conjugated goat anti-biotin antibody). Each antibody is added at 1:5000 dilution in 1% skim milk in TBST for 1 h at RT. Wash n TBST three times (8–10 min each) to prepare for enhanced chemiluminescence (ECL). 3.5 Enhanced Chemiluminescence

1. Add 44 μl of 90 mM stock of paracoumaric acid and 100 μl of 250 mM stock of luminol to a tube with 10 ml of 0.1 M Tris– HCl, pH 8.8. 2. Add 6 μl of 30% H2O2 to another tube with 10 ml of 0.1 M Tris–HCl, pH 8.8. 3. Quickly mix the contents of the two tubes in a container, place the membrane into the container, and agitate for 1 min. 4. Acquire chemiluminescence signal in the next 2–15 min in the ChemiDoc MP imager (Bio-Rad). Quantify band intensities by the accompanying software (Bio-Rad).

4

Notes 1. Injecting RNAs into all four blastomeres reduces background and increases sensitivity of immunoblotting. 2. Occasionally needles get clogged by precipitated biotin. To avoid this problem, do not inject biotin in concentrations higher than 0.2–0.4 mM. Use sharply beveled needles with large openings and keep spare calibrated needles for replacement. 3. If more convenient, 20 nl of 0.4 mM biotin solution in water can be directly injected into the blastocoel, independently of mRNA injections. 4. When comparing the degree of biotinylation, ensure that the BLN-fusion protein is expressed equally in different experimental groups.

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5. Adding 0.05–0.1% SDS to the transfer buffer is recommended for the transfer of large proteins. 6. For biotin detection, use BSA as a blocking reagent, instead of skim milk, which may contain biotin and would interfere with the detection. 7. Vangl2 biotinylation is poorly detected in embryo lysates, but it is easy to visualize in pull-downs (Fig. 3).

Acknowledgments We are grateful to Bo Xiang and Chih-Wen Chu, who contributed to the development of this assay in our group. The work in the Sokol laboratory has been supported by the NIH grants GM122492, NS100759, and HD092990. References 1. Cronan JE (2005) Targeted and proximitydependent promiscuous protein biotinylation by a mutant Escherichia coli biotin protein ligase. J Nutr Biochem 16(7):416–418. https://doi.org/10.1016/j.jnutbio.2005. 03.017 2. Roux KJ, Kim DI, Raida M, Burke B (2012) A promiscuous biotin ligase fusion protein identifies proximal and interacting proteins in mammalian cells. J Cell Biol 196(6):801–810. https://doi.org/10.1083/jcb.201112098 3. Choi-Rhee E, Schulman H, Cronan JE (2004) Promiscuous protein biotinylation by Escherichia coli biotin protein ligase. Protein Sci 13(11):3043–3050. https://doi.org/10. 1110/ps.04911804 4. Gupta GD, Coyaud E´, Gonc¸alves J, Mojarad BA, Liu Y, Wu Q, Gheiratmand L, Comartin D, Tkach JM, Cheung SWT, Bashkurov M, Hasegan M, Knight JD, Lin Z-Y, Schueler M, Hildebrandt F, Moffat J, Gingras A-C, Raught B, Pelletier L (2015) A dynamic protein interaction landscape of the human centrosome-cilium Interface. Cell 163(6):1484–1499. https://doi.org/10. 1016/j.cell.2015.10.065 5. Gingras AC, Abe KT, Raught B (2019) Getting to know the neighborhood: using proximity-dependent biotinylation to characterize protein complexes and map organelles. Curr Opin Chem Biol 48:44–54. https://doi. org/10.1016/j.cbpa.2018.10.017 6. Kim DI, Birendra KC, Zhu W, Motamedchaboki K, Doye V, Roux KJ (2014) Probing nuclear pore complex architecture

with proximity-dependent biotinylation. Proc Natl Acad Sci U S A 111(24):E2453–E2461. https://doi.org/10.1073/pnas.1406459111 7. Zhang Y, Song G, Lal NK, Nagalakshmi U, Li Y, Zheng W, P-j H, Branon TC, Ting AY, Walley JW, Dinesh-Kumar SP (2019) TurboID-based proximity labeling reveals that UBR7 is a regulator of N NLR immune receptor-mediated immunity. Nat Commun 10(1):3252. https://doi.org/10.1038/ s41467-019-11202-z 8. Branon TC, Bosch JA, Sanchez AD, Udeshi ND, Svinkina T, Carr SA, Feldman JL, Perrimon N, Ting AY (2018) Efficient proximity labeling in living cells and organisms with TurboID. Nat Biotechnol 36(9):880–887. https://doi.org/10.1038/nbt.4201 9. Rudolph F, Fink C, Hu¨ttemeister J, Kirchner M, Radke MH, Lopez Carballo J, Wagner E, Kohl T, Lehnart SE, Mertins P, Gotthardt M (2020) Deconstructing sarcomeric structure-function relations in titinBioID knock-in mice. Nat Commun 11(1): 3133. https://doi.org/10.1038/s41467020-16929-8 10. Gray RS, Roszko I, Solnica-Krezel L (2011) Planar cell polarity: coordinating morphogenetic cell behaviors with embryonic polarity. Dev Cell 21(1):120–133. https://doi.org/10. 1016/j.devcel.2011.06.011 11. Goodrich LV, Strutt D (2011) Principles of planar polarity in animal development. Development 138(10):1877–1892. https://doi. org/10.1242/dev.054080

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12. Butler MT, Wallingford JB (2017) Planar cell polarity in development and disease. Nat Rev Mol Cell Biol 18(6):375–388. https://doi. org/10.1038/nrm.2017.11 13. Vladar EK, Konigshoff M (2020) Noncanonical Wnt planar cell polarity signaling in lung development and disease. Biochem Soc Trans 48(1):231–243. https://doi.org/10.1042/ BST20190597 14. Ossipova O, Kim K, Sokol SY (2015) Planar polarization of Vangl2 in the vertebrate neural plate is controlled by Wnt and myosin II signaling. Biol Open 4(6):722–730. https://doi. org/10.1242/bio.201511676 15. Bastock R, Strutt H, Strutt D (2003) Strabismus is asymmetrically localised and binds to prickle and Dishevelled during drosophila planar polarity patterning. Development 130(13): 3007–3014. https://doi.org/10.1242/dev. 00526 16. Jenny A, Darken RS, Wilson PA, Mlodzik M (2003) Prickle and strabismus form a functional complex to generate a correct axis during planar cell polarity signaling. EMBO J 22(17): 4409–4420. https://doi.org/10.1093/ emboj/cdg424 17. Chuykin I, Ossipova O, Sokol SY (2018) Par3 interacts with Prickle3 to generate apical PCP complexes in the vertebrate neural plate. Elife 7:e37881. https://doi.org/10.7554/eLife. 37881 18. Reis AH, Xiang B, Itoh K, Sokol SY (2021) Identification of the centrosome maturation factor SSX2IP as Wtip-binding partner by targeted proximity biotinylation. PLoS ONE 16

(10):e0259068. https://doi.org/10.1371/ journal.pone.0259068 19. Kim DI, Jensen SC, Noble KA, Kc B, Roux KH, Motamedchaboki K, Roux KJ (2016) An improved smaller biotin ligase for BioID proximity labeling. Mol Biol Cell 27(8): 1188–1196. https://doi.org/10.1091/mbc. E15-12-0844 20. 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(3): 675–686 21. Chu C-W, Ossipova O, Ioannou A, Sokol SY (2016) Prickle3 synergizes with Wtip to regulate basal body organization and cilia growth. Sci Rep 6:24104. https://doi.org/10.1038/ srep24104 22. Sambrook J, Fritsch E, Maniatis T (1989) In: molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, p 1546 23. Sive H, Grainger R, Harland R (2000) Early development of Xenopus laevis: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York 24. Itoh K, Brott BK, Bae G-U, Ratcliffe MJ, Sokol SY (2005) Nuclear localization is required for Dishevelled function in Wnt/beta-catenin signaling. J Biol 4(1):3. https://doi.org/10. 1186/jbiol20 25. Nieuwkoop PD, Faber J (1967) Normal table of Xenopus laevis. North-Holland Publishing Company, Amsterdam

Chapter 7 Regulation of Cell Polarity by Posttranslational Protein Palmitoylation Baoen Chen, Carla Guarino, Abdelhalim Azzi, Hannah Erb, and Xu Wu Abstract Cell polarity is a common feature of many living cells, especially epithelial cells, and plays important roles in development, tissue homeostasis, and diseases. Therefore, the signaling pathways involved in establishing and maintaining cell polarity are tightly controlled. Protein S-palmitoylation has been recently recognized as an important posttranslational modification involved in cell polarity, via dynamic covalent attachment of fatty acyl groups to the cysteine residues of cell polarity proteins. Here, we describe the methods to study the function and regulation of S-palmitoylation of cell polarity proteins. Key words Cell polarity, Protein palmitoylation, Chemical probe, Metabolic labeling, Click chemistry

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Introduction Cell polarity is the asymmetric distribution of the cellular components along a defined axis, which allows cells to organize specific biological functions in a spatially controlled manner [1, 2]. Polarized epithelial cells have an apical membrane directly exposed to the environment, a lateral membrane in constant contact and interaction with the neighboring cells, and a basal membrane anchored to the extracellular matrices. Such organization of cell structures is termed apical–basolateral polarity [3, 4]. In addition, the orientation of cells in a given tissue in a plane vertical to the apical-basal axis is called planar cell polarity (PCP) [5, 6]. Cell polarity is highly dynamic, and tightly regulated by specific signaling pathways sensing external and internal cues. Establishment and maintenance of cell polarity requires several molecular complexes, which are specifically localized to be functional in polarity. Three polarity complexes have been reported. Partitioning defective protein (PAR) complexes are widely distributed and involved in asymmetric cell division during embryogenesis and maintenance of cell polarity. Crumbs (CRB) complexes mainly localize at the apical membranes. Scribble (SCRIB) complexes, including Scrib, lethal giant larvae

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_7, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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(Lgl), and Discs large (Dlg), are highly conserved across species from Drosophila to mammals. SCRIB complexes are predominantly localized to the lateral membranes and regulate apicobasal polarity as well as asymmetric cell division during embryogenesis [7–9]. Mutations of SCRIB in Drosophila alter morphogenesis and lead to embryonic lethality. Mislocalization of SCRIB in mammalian cells has been linked to tumor growth, metastasis and neuronal transmission [10–13]. Protein lipidation is an important co- or posttranslational modification in which lipid moieties covalently attach to proteins. Lipidation plays a crucial role in diverse cell signaling, including cell polarity, by dynamically regulating protein localization and functions. S-Palmitoylation (or S-fatty acylation) refers to the covalent attachment of palmitic acid or other long-chain fatty acid to a cysteine residue, which can be dynamically regulated by ZDHHC (zinc finger DHHC-domain) family of palmitoyl acyltransferases (PAT), and depalmitoylating enzymes, including acyl-protein thioesterases (APTs) and ABHD (alpha/beta-hydrolase domain) family of serine hydrolases. In addition, several proteins, including Wnt ligands, are known to be fatty acylated on Ser residues (O-fatty acylation) through ester linkage. Recent studies using chemoproteomics and biochemical methods have revealed that many cell polarity regulators are S-palmitoylated (Table 1). As many of the cell polarity proteins need to be precisely localized to membrane junctions, and palmitoylation is known to be a key regulator of protein trafficking and localization, deregulation of palmitoylation might disrupt proper localization of polarity signaling proteins and lead to various human diseases, including cancers. Due to hydrophobicity of fatty acyl groups and the lack of a specific antibody, it has been challenging to detect fatty acylated proteins directly and robustly [24]. The first detection method for protein palmitoylation is based on the metabolic incorporation of [3H], [14C] or [125I] radiolabeled fatty acyl derivatives followed by fluorography [25], which requires radiation protection and long exposure time due to low sensitivity. Over the past years, many new methods have been developed, including Acyl-Biotin or PEG exchange (ABE or APE) and acyl-Resin-Assisted Capture (acylRAC) assay [26–29] to study S-fatty acylation. Bioorthogonal chemical reporters can be used to analyze various types of lipid modifications, including palmitoylation, myristoylation and prenylation [30, 31]. Based on our study and those from other labs [14, 15], here we describe the methods of using bioorthogonal chemical reporters to study S-palmitoylation of cell polarity protein SCRIB (Fig. 1). The methods described herein can also be applied to the study of other palmitoylated polarity proteins.

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Table 1 Palmitoylation of cell polarity regulators Protein

Modification sites

Enzymes

Function

hSCRIB

Cys4/10 [14]

ZDHHC7 [14]; APT2 [15]

Apicobasal cell polarity

rDLG4/PSD95

Cys3/5 [16]

ZDHHC2 [17]; ABHD17 [18]

Neuronal cell polarity

dFat

Cys4938/4987 [19]

App [19]

Planar cell polarity

dWnt1/Wingless

Ser239 [20]

Porcupine [20]; Notum [21]

Planar cell polarity

mWnt3a

Ser209 [22]

PORCN [22]; Notum [21]

Planar cell polarity

mWnt11

Ser215 [23]

PORCN

Planar cell polarity

h Homo sapiens, r rat, d Drosophila melanogaster, m mouse

Fig. 1 Scheme of bioorthogonal methods for detection of protein palmitoylation

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Materials

2.1 Cell Culture Medium and Transfection Reagent

1. HEK293A medium: Dulbecco’s Eagle’s Minimal Essential Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 50 μg/ml penicillin–streptomycin. 2. MCF-10A complete growth medium: DMEM/F12 (Gibco, Cat. No. 11320033) supplemented with 5% horse serum, 100 ng/ml cholera toxin (Sigma-Aldrich, Cat. No. C8052), 500 ng/ml hydrocortisone (Sigma-Aldrich, Cat. No. H0888), 20 ng/ml EGF (Gibco, Cat. No. PHG0311), 10 μg/ml insulin (Sigma-Aldrich, Cat. No. I1882), and 50 μg/ml penicillin– streptomycin. 3. MCF-10A low serum medium: DMEM/F12 supplemented with 2% horse serum, 100 ng/ml cholera toxin, 500 ng/ml hydrocortisone, 10 μg/ml insulin, and 50 μg/ml penicillin– streptomycin. 4. MCF-10A neutralizing medium: DMEM/F12 supplemented with 20% horse serum and 50 μg/ml penicillin–streptomycin.

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5. MCF-10A three-dimensional (3D) growth medium: DMEM/ F12 supplemented with 2% Matrigel, 2% horse serum, 100 ng/ ml cholera toxin, 500 ng/ml hydrocortisone, 5 ng/ml EGF, 10 μg/ml insulin, and 50 μg/ml penicillin–streptomycin. 6. Dulbecco’s phosphate buffered saline No. 14190250, Life Technologies).

(DPBS)

(Cat.

7. jetPRIME transfection reagent (Polyplus transfection SA, Cat. No. 114-07). 8. INTERFERin siRNA transfection transfection SA, Cat. No. 40910).

reagent

(Polyplus-

9. siRNA control (GE Dharmacon). 10. siRNA targeting ZDHHC7 No. M-021091-00-0005). 2.2 Metabolic Labeling and Click Reaction

(GE

Dharmacon,

Cat.

1. Dialyzed FBS (Bio-Techne, Cat. No. S11150). 2. Alk-16C (15-hexadecynoic acid) (Cayman Chemical, Cat. No. 13266), 50 mM in DMSO. 3. Lysis buffer: 50 mM TEA-HCl (pH 7.4), 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 1 mM PMSF, 1 cOmplete, EDTAfree protease inhibitor cocktail (Roche, Cat. No. 11873580001), 1 PhosSTOP, phosphatase inhibitor cocktail (Roche, Cat. No. 04906845001). 4. Biotin azide (Click Chemistry Tools LLC, Cat. No. 1167), 10 mM in DMSO. 5. TCEP (Tris(2-carboxyethyl)phosphine hydrochloride) (Sigma-Aldrich, Cat. No. C4706) 50 mM in pure water. 6. TBTA (Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine) (Sigma-Aldrich, Cat. No. 678937), 2 mM in DMSO/t-butanol (1/4). 7. CuSO4·5H2O (Sigma-Aldrich, Cat. No. 469130), 50 mM in pure water.

2.3 Streptavidin Bead Pull-Down

1. Streptavidin agarose beads (EMD No. 69203-3).

MILLIPORE , Cat.

2. Methanol. 3. Dissolve buffer: 50 mM Tris–HCl pH 7.4, 150 mM NaCl, 10 mM EDTA, 2% SDS. 4. Dilution buffer: 50 mM Tris–HCl pH 7.4, 150 mM NaCl, 10 mM EDTA, 1% NP-40. 5. Wash buffer-1: 50 mM Tris–HCl pH 7.4, 150 mM NaCl, 10 mM EDTA. 6. Wash buffer-2: 50 mM Tris–HCl, pH 7.4, 150 mM NaCl.

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2.4 Streptavidin Blot and Western Blot

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1. 6 SDS-Sample Buffer. 2. 4–12% precast polyacrylamide gel. 3. PVDF membrane. 4. Nonfat dry milk. 5. BSA. 6. Hydroxylamine (NH2OH) (Sigma-Aldrich, Cat. No. 467804). 7. HRP-conjugated No. N100). 8. Enhanced substrate.

2.5

Antibody

streptavidin

chemiluminescent

(Thermo (ECL)

Scientific, western

Cat.

blotting

1. Anti-FLAG (Sigma-Aldrich, Cat. No. F1804). 2. Anti-FLAG Magnetic No. M8823).

Beads

(Sigma-Aldrich,

Cat.

3. Anti-HA (Cell Signaling, Cat. No. 3724). 4. Anti-SCRIB (Santa Cruz, Cat. No. sc-374139). 5. Anti-β-Actin (Abcam, Cat. No. ab6276). 6. Anti-GAPDH (Santa Cruz, Cat. No. SC-47724). 7. Anti-GM130 (BD Biosciences, Cat. No. 610822). 8. Alexa Fluor Plus 488 secondary antibody (Invitrogen, Cat. No. A32766). 2.6

Constructs

1. SCRIB cDNA (Homo sapiens) was requested from Dana-Farber/Harvard Cancer Center DNA Resource Core at Harvard Medical School and cloned into pCMV-3Tag-6 (N-terminal FLAG). 2. SCRIB mutants (C4S, C10S, C22S, C4/10S, C4/22S, C10/22S, C4/10/22S, P305L) are generated by site-directed mutagenesis kit (Agilent, Cat. No. 200523). 3. HA-tagged ZDHHC3, 4 and 7 are gifts from M. Fukata at National Institute for Physiological Sciences, Japan.

2.7 3D Culture and Immunofluorescence

1. Matrigel (Biosciences, Cat. No. 356234). 2. Falcon chambered culture slides (Thermo Scientific, Cat. No. 08-774-26). 3. 16% paraformaldehyde (PFA). 4. Triton X-100. 5. 3D wash buffer: DPBS buffer containing 0.1% BSA, 0.2% Triton-X 100, 0.04% Tween 20, NaN3 (0.05%), pH 7.4. 6. Blocking buffer (3D wash buffer + 10% goat serum). 7. Hoechst 33342 (Thermo Scientific, Cat. No. H3570).

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8. ProLong Antifade No. P10144).

Mountant (Molecular

Probes, Cat.

9. Glycine buffer: 100 mM glycine in DPBS, pH 7.4.

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Methods All procedures are performed at room temperature, unless otherwise specified.

3.1 Chemical Proteomic Profiling of Palmitoylated Proteins 3.1.1 Metabolic Labeling

1. Seed HEK293A cells (~3  106) in 10-cm dish (see Note 1). 2. After around 24 h incubation, wash cells once with DPBS and gently aspirate off. 3. Add 5 ml of prewarmed DMEM supplemented with 10% dialyzed FBS (see Note 2). 4. Two hours later, cells are treated with 100 μM of Alk-16C probe or the equivalent volume of DMSO overnight.

3.1.2 Protein Extraction

1. Put the cell culture dish directly on ice after taking them out from incubator (see Note 3). 2. Add 1 ml of lysis buffer to each dish after gentle wash three times with prechilled DPBS and aspiration off the residues (see Note 4). 3. Harvest cells with a cell lifter and transfer cell lysates into 15 ml tube. 4. After 30 min incubation on ice, cell lysates are centrifuged at >15,000  g  20 min, 4  C. 5. Transfer the supernatant into a new tube without disturbing the pellet. 6. Measure protein concentration with bicinchoninic acid (BCA) method. 7. Adjust protein concentrations to ~1 mg/ml using lysis buffer.

3.1.3 Click Reaction

1. Transfer 4.5 ml of supernatant into 15 ml tube. 2. Add 500 μl of click reagents (50 μl biotin azide, 100 μl TCEP, 250 μl TBTA and 100 μl CuSO4) to each tube (see Note 5). Vortex briefly to mix. 3. Incubate for 1 h on rotator (see Note 6).

3.1.4 Streptavidin Bead Pull-Down

1. Transfer click reaction product into 50 ml tube. 2. Add 45 ml methanol to each tube, yielding 90% methanol final concentration. 3. Incubate for >2 h at

80  C.

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4. Centrifuge at >15,000  g 10 min, 4  C. 5. Gently remove supernatant without disturbing pellets. 6. Air-dry for 15 min (see Note 7). 7. Add 2.5 ml dissolve buffer to pellets. 8. Vortex or rotate until fully dissolved. 9. Transfer dissolved pellets into 15 ml tube. 10. Dilute with 2.5 ml dilution buffer. 11. During dissolving pellets, transfer 500 μl streptavidin agarose beads slurry into 2 ml tube. 12. Add 1.5 ml wash buffer-1 to beads. 13. Centrifuge at 1500  g 4 min. 14. Gently remove wash buffer-1 without disturbing beads. 15. Repeat steps 10–12 three times. 16. Resuspend beads with wash buffer with a final volume of 550 μl. 17. Add 250 μl beads to each sample and incubate overnight on rotator. 18. Wash beads 6–10 times with wash buffer-2 (5 ml/time) to completely remove EDTA and SDS (see Note 8). 3.1.5 Mass Spectrometry Analysis

3.2 Validation of SCRIB Palmitoylation 3.2.1 Transfection

3.2.2 Metabolic Labeling

The pulldown samples are submitted to the Taplin Biological Mass Spectrometry facility at Harvard Medical School to identify potential palmitoylated proteins. 1. HEK293A cells are seeded at a density of ~2  105 cells/well in 6-well plate (see Note 1). 2. About 24 h later, cells are transfected with 2 μg of FLAGSCRIB construct and empty vector control using jetPRIME transfection reagent, respectively. 1. At 36 h posttransfection, wash cells once with DPBS and gently aspirate off. 2. Add 1 ml of prewarmed DMEM supplemented with 10% dialyzed FBS (see Note 2). 3. Two hours later, treat cells with 50 μM of Alk-16C probe or equivalent volume of DMSO for 2–4 h (see Note 9).

3.2.3 Protein Extraction

1. Put 6-well plate directly on ice after taking them out from incubator (see Note 3). 2. Wash cells three times with prechilled DPBS and gently aspirate off. 3. Add 120 μl of lysis buffer to each well (see Note 4).

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4. Harvest cells with cell lifter. 5. Transfer cell lysates into1.5 ml tube. 6. Incubate for 30 min on ice. 7. Centrifuge at >15,000  g 20 min, 4  C. 8. Transfer supernatant into new tube without disturbing pellets. 9. Measure protein concentration with BCA method. 10. Adjust protein concentrations to ~1 mg/ml using lysis buffer. 3.2.4 Click Reaction

1. Transfer 90 μl supernatant to a new tube. 2. Add 10 μl of click reaction reagents (1 μl biotin azide, 2 μl TCEP, 5 μl TBTA, and 2 μl CuSO4) to each tube (see Note 5). Vortex briefly to mix. 3. Incubate for 1 h on rotator (see Note 6). 4. Stop click reaction by adding 20 μl of 6 SDS-Sample Buffer and boiling at 95  C for 5 min (see Note 10).

3.2.5 Streptavidin Blot

1. Load 10 μl of each sample into 15-well 4–12% precast polyacrylamide gel. 2. Run the gel according to manufacture instructions. 3. Transfer protein to PVDF membrane. 4. Block membrane with 5% BSA in TBST overnight at 4  C (see Note 11). 5. Wash membrane once with TBST for 10 min. 6. Incubate membrane with streptavidin-HRP (1:20,000) for 1 h. 7. Wash membrane three times, 10 min each time in TBST. 8. Detect membrane with ECL (Fig. 2).

3.2.6 Western Blot

3.3 Identification of SCRIB Palmitoylation Sites

After streptavidin blot, sequentially perform western blot using Anti-FLAG (1:5000) and Anti-GAPDH (1:1000) antibody on the same membrane (Fig. 2). 1. Conservation analysis of cysteine residues. Multiple sequence alignment (ClustalW) identifies three highly conserved cysteine residues of SCRIB: Cys4, Cys10, and Cys22. 2. Single, double, and triple Cys-to-Ser mutants are generated by site-directed mutagenesis kit. 3. HEK293A cells are transfected with 1 μg each of wild-type (WT) and mutant SCRIB constructs using jetPRIME transfection reagent. 4. The detection methods for SCRIB palmitoylation are the same as previously described.

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Fig. 2 Validation of SCRIB palmitoylation using chemical reporters and streptavidin blot. FLAG-tagged SCRIB was transfected into HEK293A cells and labeled with probe Alk-16C for different time points (0, 0.5, 1, 2, and 4 h). Click reaction followed by streptavidin blot showing palmitoylation of SCRIB at each time point 3.4 Exploration of the Mechanisms Regulating SCRIB Palmitoylation 3.4.1 ZDHHC-PAT Screen

3.4.2 ZDHHC-PAT and SCRIB Interactions by Co-IP Assay

1. 1 μg of FLAG-SCRIB and 1 μg of HA-tagged ZDHHCs or HA-tagged GST as control are cotransfected into HEK293A cells using jetPRIME transfection reagent in 6-well plate. 2. The detection methods for SCRIB palmitoylation are the same as previously described. ZHDDC7 substantially promotes SCRIB palmitoylation (Fig. 3). 1. Cotransfect 1 μg of FLAG-SCRIB and 1 μg of HA-tagged ZDHHC7 into HEK293A cells using jetPRIME transfection reagent in 6-well plates. 2. Detect ZDHHC7 level after co-IP with 30 μl of Anti-FLAG Magnetic Beads and immunoblotting using Anti-HA antibody (1:1000). 3. Detect SCRIB level after co-IP with 30 μl of Anti-HA Magnetic Beads and immunoblotting using Anti-FLAG antibody (1:5000) (see Note 12).

3.4.3 ZDHHC7 Inactivation Analysis

1. Generation of catalytically dead mutant C160S of ZDHHC7 by using site-directed mutagenesis kit. 2. Cotransfect 1 μg of FLAG-SCRIB and 1 μg of HA-ZDHHC7 WT, C160S mutant or empty vector control into HEK293A cells using jetPRIME transfection reagent in 6-well plate. 3. Detect SCRIB palmitoylation levels as previously described.

3.4.4 ZDHHC7 Knockdown and Rescue Assay

1. Transfect 30 nM of siRNA targeting ZDHHC7 or mock siRNA into HEK293A cells using INTERFERin siRNA transfection reagent in 6-well plate. 2. After about 24 h, cotransfect 1 μg of FLAG-SCRIB and 1 μg of HA-ZDHHC7 WT or C160S mutant construct into cells.

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Fig. 3 ZDHHC7 is the major acyltransferase regulating SCRIB palmitoylation. FLAG-SCRIB and HA-ZDHHCs (4, 3, and 7) or empty vector control were cotransfected into HEK293A cells. About 36 h later, cells were labeled with probe Alk-16C for 4 h. Click reaction followed by streptavidin blot showing overexpression of ZDHHC7 or ZDHHC3 substantially increases SCRIB palmitoylation

3. Around 48 h later, analyze SCRIB palmitoylation level as previously described. 3.5 Evaluation of SCRIB Palmitoylation in Regulating Cell Polarity 3.5.1 Culture MCF-10A Cells Under 3D Culture Conditions

1. Generation of stable MCF-10A cells expressing FLAG-SCRIB WT (pBABE-puro vector) and palmitoylation-deficient mutant C4/10S by using 1 ml of virus with a titer of ~5  105 CFU/ ml in 6-well plate. 2. At 48 h posttransduction, cells are selected with 1 μg/ml puromycin for 2 weeks. 3. Split cells and add fresh puromycin to medium every 3–4 days. 4. Before 3D culture, remove puromycin and passage one time (see Note 13). 5. Thaw Matrigel on ice and add 45 μl of Matrigel to each well in a prechilled 8-well chamber slide (see Note 14). 6. Put chamber slide in CO2 incubator at 37  C for ~45 min to solidify Matrigel. 7. Harvest cells by trypsinization. 8. Add 5 ml neutralizing medium. 9. Centrifuge at 500  g, 5 min. 10. Remove neutralizing medium and resuspend cells in low serum medium.

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11. Pipette up and down to achieve single cell suspension before counting them. 12. During trypsinization, prepare 3D growth medium. 13. Dilute cells to 12,500 cells/ml in 3D growth medium and mix thoroughly. 14. Once Matrigel solidified in chamber slide, gently transfer 400 μl of mixed cells into each well. 15. Put chamber slide in CO2 incubator at 37  C. 16. Change fresh growth medium every 4 days (see Note 15). 3.5.2 Immunofluorescence Analysis of Apical-Basal Polarity

1. At day 14, gently remove 3D growth medium without disturbing Matrigel. 2. Fix cells with 400 μl of 2% PFA in DPBS for 20 min. 3. Permeabilize cells with 0.5% Triton X-100 in DPBS for 10 min at 4  C. 4. Wash three times with Glycine buffer, 10 min each time. 5. Incubate with 200 μl Blocking buffer for 1.5 h. 6. Gently remove Blocking buffer. 7. Incubate with primary antibody (Anti-GM130) in Blocking buffer (1:100, 120 μl/well) at 4  C overnight (see Note 16). 8. Leave slide at room temperature for 1 h to heat up. 9. Wash three times with Glycine buffer, 20 min each time. 10. Incubate with secondary antibody (Alexa Fluor Plus 488) in Blocking buffer (1:2000, 120 μl/well) for 1 h (see Note 17). 11. Wash once with 3D wash buffer for 20 min. 12. Wash twice with DPBS, 10 min each time. 13. Incubate with Hoechst 33342 in DPBS (1:2000) for 20 min. 14. Wash one time with DPBS. 15. Gently pull up the chambers with lifter. 16. Mount slide with ProLong Antifade Mountant. 17. Air-dry overnight (see Note 18). 18. Collect images using Leica TCS-NT 4D confocal microscope (Fig. 4).

4

Notes 1. Cell confluency should be around 75% before probe labeling. If confluency is too low or too high, the labeling efficiency may decrease.

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Fig. 4 Representative confocal images of 3D immunofluorescence showing the alterations of acini morphology and cis-Golgi marker GM130 in wild-type (WT) and palmitoylation deficient mutant of SCRIB (C4/10S). Scale bar, 20 μm

2. To minimize the effect of fatty acid from normal FBS, dialyzed FBS should be used for probe labeling. 3. Use cells immediately or quick-freeze them in liquid nitrogen and store at 80  C. Cells should be placed on ice as much as possible during cell lysis. 4. Metal chelating reagents, such as EDTA, must be excluded from lysis buffer. EDTA can substantially affect coppercatalyzed azide–alkyne cycloaddition reaction. Copper-free Click reaction can be used as a substitute if metal chelating reagents are necessary. 5. Click reagent solutions must be aliquoted and stored at 20  C. Freeze-thaw cycles may substantially reduce the efficiency of the reagents. The final concentration of reagents: 100 μM biotin azide, 1 mM TCEP, 100 μM TBTA, and 1 mM CuSO4. 6. During click reaction, mix the sample thoroughly by shaking or rotating the tube. 7. Do not overdry the pellets. Overdried pellets are hard to dissolve. 8. For mass spectrometry analysis, the streptavidin beads must be washed thoroughly with pure water or 10 mM Tris–HCl pH 7.4. EDTA, Triton X-100, and NP-40 may substantially disrupt mass spectrometry analysis. 9. Metabolic labeling time varies from protein to protein. For SCRIB, around 4-h labeling is enough. 10. Hydroxylamine treatment assay can be performed to test the formation of thioester bond between Alk-16C probe and cysteine residues. Add 2.5% (v/v) hydroxylamine (NH2OH) to

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the sample and boil at 95  C for 5 min. The intensity of palmSCIRB band should substantially decrease on the following streptavidin blot. 11. 5% w/v BSA in TBST is the ideal blocking buffer. Milk must be excluded. After blocking, wash membrane once with DPBS or TBST for 10 min. Incubate with streptavidin-HRP diluted in TBST at 1:10,000–1:20,000. Low concentration of streptavidin-HRP might reduce background during detection. 12. To analyze endogenous SCRIB and ZDHHC7 interactions, transfect 1.5 μg of HA-tagged ZDHHC7 into HEK293A cells. Detect endogenous SCRIB level after co-IP with 30 μl of AntiHA Magnetic Beads and immunoblotting using Anti-SCRIB antibody (1:1000). 13. After 2 weeks puromycin selection, MCF-10A stable cells should be passaged one time to allow the cells recovery from puromycin and used for 3D culture as soon as possible. Continuous passage may result in SCRIB level decrease or loss. 14. During 3D culture preparation, always keep Matrigel on ice. Once thawed, Matrigel should be gently mixed and buried in ice. Gently transfer Matrigel into prechilled chamber using prechilled tips. Prevent bubbles as much as possible. 15. Use tips instead of vacuum to remove top medium without disrupting cells on Matrigel. 16. Antibody specificity is critical for 3D immunofluorescent staining. 17. From this point on, keep slide in darkness to prevent fluorochrome bleaching. 18. Once the mountant is completely dried, the slide can be stored at 4  C for 2 weeks or at 80  C for several months.

Acknowledgments The authors thank the Taplin Mass Spec Core Facility of Harvard Medical School, and the confocal core at CBRC, Massachusetts General Hospital. Wu lab is supported by Melanoma Research Alliance, NIH R01 (R01CA181537 and R01DK107651-01). References 1. Bryant DM, Mostov KE (2008) From cells to organs: building polarized tissue. Nat Rev Mol Cell Biol 9(11):887–901. https://doi.org/10. 1038/nrm2523 2. St Johnston D, Ahringer J (2010) Cell polarity in eggs and epithelia: parallels and diversity.

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cell polarity. Nat Chem Biol 12(9):686–693. https://doi.org/10.1038/nchembio.2119 15. Hernandez JL, Davda D, Cheung See Kit M, Majmudar JD, Won SJ, Gang M, Pasupuleti SC, Choi AI, Bartkowiak CM, Martin BR (2017) APT2 inhibition restores scribble localization and S-Palmitoylation in snailtransformed cells. Cell Chem Biol 24(1): 87–97. https://doi.org/10.1016/j.chembiol. 2016.12.007 16. Topinka JR, Bredt DS (1998) N-terminal palmitoylation of PSD-95 regulates association with cell membranes and interaction with K+ channel Kv1.4. Neuron 20(1):125–134 17. Fukata M, Fukata Y, Adesnik H, Nicoll RA, Bredt DS (2004) Identification of PSD-95 palmitoylating enzymes. Neuron 44(6):987–996. https://doi.org/10.1016/j.neuron.2004. 12.005 18. Yokoi N, Fukata Y, Sekiya A, Murakami T, Kobayashi K, Fukata M (2016) Identification of PSD-95 Depalmitoylating enzymes. J Neurosci 36(24):6431–6444. https://doi.org/10. 1523/JNEUROSCI.0419-16.2016 19. Matakatsu H, Blair SS, Fehon RG (2017) The palmitoyltransferase approximated promotes growth via the hippo pathway by palmitoylation of fat. J Cell Biol 216(1):265–277. https://doi.org/10.1083/jcb.201609094 20. Herr P, Basler K (2012) Porcupine-mediated lipidation is required for Wnt recognition by Wls. Dev Biol 361(2):392–402. https://doi. org/10.1016/j.ydbio.2011.11.003 21. Kakugawa S, Langton PF, Zebisch M, Howell S, Chang TH, Liu Y, Feizi T, Bineva G, O’Reilly N, Snijders AP, Jones EY, Vincent JP (2015) Notum deacylates Wnt proteins to suppress signalling activity. Nature 519(7542):187–192. https://doi.org/10. 1038/nature14259 22. Takada R, Satomi Y, Kurata T, Ueno N, Norioka S, Kondoh H, Takao T, Takada S (2006) Monounsaturated fatty acid modification of Wnt protein: its role in Wnt secretion. Dev Cell 11(6):791–801. https://doi.org/10. 1016/j.devcel.2006.10.003 23. Yamamoto H, Awada C, Hanaki H, Sakane H, Tsujimoto I, Takahashi Y, Takao T, Kikuchi A (2013) The apical and basolateral secretion of Wnt11 and Wnt3a in polarized epithelial cells is regulated by different mechanisms. J Cell Sci 126(Pt 13):2931–2943. https://doi.org/10. 1242/jcs.126052 24. Chen B, Sun Y, Niu J, Jarugumilli GK, Wu X (2018) Protein Lipidation in cell signaling and diseases: function, regulation, and therapeutic opportunities. Cell Chem Biol 25(7):817–831.

Polarity Regulation by Protein Palmitoylation https://doi.org/10.1016/j.chembiol.2018. 05.003 25. Jackson CS, Magee AI (2001) Metabolic labeling with fatty acids. Curr Protoc Cell Biol. Chapter 7:Unit 7 4. https://doi.org/10. 1002/0471143030.cb0704s05 26. Wan J, Roth AF, Bailey AO, Davis NG (2007) Palmitoylated proteins: purification and identification. Nat Protoc 2(7):1573–1584. https:// doi.org/10.1038/nprot.2007.225 27. Percher A, Ramakrishnan S, Thinon E, Yuan X, Yount JS, Hang HC (2016) Mass-tag labeling reveals site-specific and endogenous levels of protein S-fatty acylation. Proc Natl Acad Sci U S A 113(16):4302–4307. https://doi.org/ 10.1073/pnas.1602244113 28. Drisdel RC, Green WN (2004) Labeling and quantifying sites of protein palmitoylation. BioTechniques 36(2):276–285

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Chapter 8 Enrichment and Detection of Wnt Proteins from Cell Culture Media Pooja R. Sonavane and Karl Willert Abstract Wnt proteins are secreted, lipid-modified growth factors with a wide range of activities across all metazoan species. Their production, secretion, and signaling range are under tight cellular control such that detection of Wnt proteins in biological samples is often extremely difficult. In this chapter, we describe a protocol to detect secreted Wnt proteins in the culture medium of cell lines that ectopically or endogenously express Wnt genes. This protocol uses an affinity resin, called Blue Sepharose, that binds and thereby enriches Wnt proteins, followed by immunoblotting for the Wnt protein of interest. This method for detecting Wnt proteins will aid in the isolation of biologically active Wnt proteins, provide an assay to study the molecular basis of Wnt secretion, and potentially offer a means to detect trace amounts of Wnt proteins associated with pathological states. Key words Wnt signaling, Lipid modification, Hydrophobic protein, Wnt secretion, Blue Sepharose, Wnt purification

1

Introduction Wnt glycoproteins are a group of secreted ligands, with the mammalian genome encoding 19 Wnt genes. These proteins undergo a series of modifications in the endoplasmic reticulum (ER), including glycosylation and lipidation, before release into the extracellular space. The covalently attached lipid renders Wnt proteins hydrophobic [1, 2], presenting a solubility problem that has captivated the attention of many researchers: how does this highly hydrophobic Wnt protein traffic in an extracellular space that is largely aqueous? Part of the answer to this conundrum is that Wnts act locally (reviewed in [3]). In fact, signaling by Wnts is more akin to Notch than to Insulin signaling, often requiring direct cell-cell contact. Nonetheless, Wnt proteins have been detected long distances from their site of secretion and in conditioned media (CM) of cell lines, indicating that some Wnts can detach from cells. Several

Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_8, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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mechanisms have been proposed and investigated to explain how these hydrophobic proteins may transit through a largely aqueous environment from their site of secretion to their target cells. For example, actin-based membrane protrusions, referred to as filipodia or cytonemes, can facilitate the spread of Wnts several cell diameters from their site of synthesis [4]. Wnts may also associate with chaperones as they transit through the secretory pathway and exit the cell. Such chaperone activity has been described for Swim [5] and Afamin [6], which bind Wnts and shield their hydrophobic moiety. Additionally, several studies have detected Wnts on lipid vesicles, such as exosomes [7, 8], and lipid particles, such as lipoprotein complexes [9, 10]. Collectively, such mechanisms disseminate Wnt proteins in the extracellular environment in vivo and in vitro. When purifying Wnt proteins, it is done in the presence of detergents to disrupt protein complexes, leaving Wnt as a monomer in solution. However, the amount of cell-free Wnt protein is often quite low and difficult to detect using antibodies. Even the most robust overexpression systems yield abysmal amounts of protein, with the concentrations of Wnt3a or Wnt5a estimated at 0.1 mg per liter of CM (in contrast, antibody expression systems routinely yield thousandfold more recombinant proteins). The dearth of Wnt protein released from cells may be due to bottlenecks in the secretory pathway where essential Wnt processing and trafficking enzymes, such as PORCN and WLS, are limiting or saturated, with a vast excess of misfolded protein remaining in the ER. The established protocol to purify biologically active Wnt3a from CM [2, 11] can be adapted to other Wnts, likely requiring some modifications in salt and detergent concentrations. The key step in this purification involves the use of Blue Sepharose (BS), a resin to which the aromatic anionic ligand, Cibacron Blue F3G-A, has been coupled. BS, which traditionally is used as an ion exchanger to remove albumin from samples, exhibits an extraordinary capacity to bind Wnts. In our application, described here and illustrated in Fig. 1, BS binds Wnt proteins under physiological salt concentrations and high detergent concentrations, both conditions that do not favor BS-albumin binding. Because Wnt CM should generally be collected from cells cultured in high serum (e.g., 10% fetal bovine serum), which contains high concentrations of lipid to facilitate Wnt solubility, this single fractionation step provides a remarkable enrichment of the Wnt protein. The BS precipitated Wnt protein can then be analyzed by immunoblotting with a suitable Wnt-specific antibody. Here we provide several examples in which we applied this BS-precipitation protocol to detect various recombinant Wnt proteins, including tagged versions (Fig. 2). We also provide an example to detect endogenously expressed Wnt proteins, in this case, secreted WNT3/WNT3A and WNT5A in the CM of

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Fig. 1 Schematic representation of Wnt precipitation using Blue Sepharose. Briefly, the sample, often conditioned medium (CM), is collected from Wnt expressing cells and filtered to remove cell debris. After addition of detergent (Triton X-100) and buffer (Tris–Cl pH 7.5) and filtering to remove insoluble material, prewashed Blue Sepharose (BS) is added and incubated on a rotator for 15–60 min. Proteins bound to BS are then analyzed by immunoblotting

differentiating human embryonic stem (hES) cells (Fig. 3). With the availability of Wnt-specific antibodies, this protocol can be readily adapted to detect the presence of Wnt proteins in any biological sample.

2

Materials

2.1 Preparation of Sample

1. Wnt-containing sample (see Note 1), 1–100 mL (see Note 2). 2. Syringe (1, 5, 10, 30, a or 50 mL). 3. Syringe filter with 0.22-μm pore size. 4. 1.5 mL, 15 mL, and 50 mL tubes. 5. 20% (v/v) Triton X-100. 6. 1 M Tris–HCl, pH 7.5. 7. 10% (w/v) NaN3.

2.2 Wnt Precipitation Using BS

1. Blue Sepharose (BS) 1:1 slurry (see Note 3). 2. Wash Buffer: 150 mM KCl, 50 mM Tris–HCl, pH 7.5, 1% (w/v) CHAPS.

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Fig. 2 Detection of recombinant Wnt proteins. (a) Four-day conditioned media from CHO cells engineered to overexpress Wnt3a, Flag-WNT3A, Flag-WNT5A and WNT9A were collected and processed as described in Subheading 3. BS pellets were washed with Wash Buffer containing different salt concentrations (50, 150 and 500 mM KCl) to demonstrate that the precipitated Wnt proteins are eluted at higher ionic strength. The Ponceau S stained samples demonstrate that significantly less total protein is bound to BS at 150 mM KCl, whereas the majority of each Wnt protein is still bound, highlighting the selectivity of BS for Wnt over other proteins. (b) Four-day conditioned media from CHO cells engineered to overexpress WNT3 and EGFP-tagged WNT3 (N-terminally tagged: EGFP-WNT3; C-terminally tagged: WNT3-EGFP) were collected and processed as described in Subheading 3. The tagged WNT3 proteins exhibit the same binding to BS as untagged WNT3

3. Vortex mixer. 4. 1.5 mL, 15 mL tubes. 5. Bench-top centrifuge capable of spinning 1.5 mL or 15 mL tubes. 6. Tube Rotator. 7. Protein Sample Loading Dye (4): 250 mM Tris–HCl, pH 6.8, 8% (w/v) sodium dodecyl sulfate (SDS), 40% (v/v) glycerol, 20% (v/v) 2-mercaptoethanol, 10 mg bromo phenol blue. 8. Heating block.

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Fig. 3 Detection of endogenously expressed Wnt proteins. Human embryonic stem cells (H1, WA01) were cultured overnight in Essential 8 (E8) medium [16] on Matrigel® after which the media were replaced with fresh E8 (Untreated), E8 containing 10% (v/v) fetal bovine serum (FBS), or E8 containing 10% FBS and 500 nM CHIR98014 (FBS + CHIR), and cells were incubated for 7 days without media changes. Addition of FBS promotes nondirected differentiation of hES cells, whereas addition of CHIR, a GSK3 inhibitor and potent Wnt agonist, promotes mesendodermal-directed differentiation of hES cells. CM were collected after 7 days of culturing and processed and analyzed as described in Subheading 3. Ponceau S staining of BS samples after transfer to a nitrocellulose membrane demonstrates that a large number of proteins bind to BS. Immunoblotting with Wnt3a- and WNT5A-specific antibodies reveals the presence of these proteins in the CM of hES cells. Note that WNT3 and WNT3A are highly similar and the antibody used here reacts with both proteins such that we cannot distinguish whether WNT3 or WNT3A is detected here

2.3 Immunoblotting to Detect BS-Precipitated Wnt Proteins

1. Suitable SDS-polyacrylamide gel electrophoresis (PAGE) setup. 2. Nitrocellulose or polyvinyl difluoride (PVDF) membrane. 3. Suitable electroblotting apparatus. 4. Ponceau S solution. 5. TBST: 50 mM Tris–HCl pH 7.5, 150 mM NaCl, and 0.2% (v/v) Tween. 6. Blocking Buffer: 3% (w/v) nonfat milk powder (NFMP) and 1% (w/v) bovine serum albumin (BSA) in TBST. 7. Wnt antibody (see Note 4). 8. Appropriate secondary antibody conjugated for detection. 9. Suitable instrumentation for detection of immunoblot signal.

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Methods

3.1 Preparation of Sample (See Note 5)

1. Collect conditioned media from cells suspected to express and secrete a Wnt protein of interest (see Note 6). 2. Filter the sample using the appropriate size syringe fitted with a 0.22-μm syringe filter (see Note 7). 3. To the filtrate, add Triton X-100 to 1% (v/v), Tris–HCl, pH 7.5 to 20 mM, and NaN3 to 0.01% (w/v) (e.g., to 9.3 mL sample, add 0.5 mL 20% (v/v) Triton X-100, 0.2 mL 1 M Tris–HCl, pH 7.5, and 10 μL 10% (w/v) NaN3). 4. Filter through a 0.22-μm syringe filter as in Subheading 3.1, step 1.

3.2 Wnt Precipitation Using BS

1. Prepare BS by washing an appropriate amount of beads three times with Wash Buffer (e.g., centrifuge 1 mL BS at 2000  g for 1 min to pellet beads, aspirate supernatant, add 1 mL Wash Buffer, vortex, and repeat twice more). 2. To sample from Subheading 3.1, step 3 add 20 μL washed BS. 3. Rotate sample/BS mixture on tube rotator for 15–60 min. 4. Centrifuge sample/BS mixture to pellet beads, 2000  g for 2 min. 5. Aspirate supernatant leaving BS pellet undisturbed. 6. Add 1 mL Wash Buffer to BS pellet and vortex briefly. 7. Centrifuge at 2000  g for 2 min. 8. Repeat steps 5–7 twice more. 9. To the washed and aspirated (nearly dry) BS add 10 μL Protein Loading Dye and 30 μL H2O and vortex. 10. Incubate in heating block at 95–100  C for 5 min.

3.3 Immunoblotting to Detect BS-Precipitated Wnt Proteins

1. Electrophorese 20 μL sample from step 10 in Subheading 3.2 by SDS-PAGE (see Note 8). 2. Transfer separated proteins from the polyacrylamide gel to nitrocellulose or PVDF membrane using an electroblotter (see Note 9). 3. Rinse membrane carrying immobilized proteins in water. 4. Submerge membrane in Ponceau S solution for 30 s. 5. Remove Ponceau S solution and wash the membrane in water to visualize proteins. If needed, photograph the Ponceau S– stained membrane. 6. Wash membrane in TBST for 5 min. 7. Pour off TBST, add 20–50 mL Blocking Buffer and incubate for 1 h.

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8. Incubate the membrane in Blocking Buffer containing primary antibody, diluted to the appropriate concentration, 1 h at room temperature or overnight at 4  C on a shaker. 9. Wash the membrane three times with TBST, 5 min per wash. 10. Incubate the membrane with appropriate secondary antibody diluted in blocking solution at room temperature for 1 h. 11. Wash the membrane three times with TBST, 5 min per wash. 12. Use the appropriate instrumentation for detection of the secondary antibody.

4

Notes 1. There are many potential sources of samples that could be examined for the presence of a Wnt protein. Conditioned media (CM) from cells overexpressing a Wnt transgene (either by transient transfection or stable transfection) serve as a relatively abundant source of Wnt protein. Several cell lines stably overexpressing Wnt proteins are available from ATCC, including Wnt3a (ATCC# CRL-2647 and CRL-3276) and Wnt5a (ATCC# CRL-2814). Endogenously expressed Wnt proteins can also be detected in CM of a variety of cell lines, as we previously showed for WNT5A in patient-derived fibroblasts [12]. Here we provide an example of detecting endogenously expressed WNT3A and WNT5A in the CM of differentiating human embryonic stem cells (Fig. 3). 2. The volume of sample is dictated by its availability. Generally speaking, 1–10 mL of CM from cells overexpressing Wnt is sufficient to detect the Wnt protein of interest. However, volumes of certain samples, such as patient derived samples, may be limiting. The goal of this protocol is to precipitate all Wnt protein in the sample on BS such that the Wnt protein of interest can be analyzed and detected by SDS-PAGE followed by immunoblotting. 3. Several types of Cibacron Blue F3G-A conjugated resins are commercially available. We have confirmed that Blue Sepharose® (6 Fast Flow and HP) and Capto™ Blue are effective for this protocol. 4. Many Wnt-specific antibodies are commercially available and described in the scientific literature. Here, we use antibodies specific to Wnt3a (Figs. 2a and 3) [2], WNT5A (Figs. 2a and 3) [13], WNT9A (Fig. 2a) [14], and Flag (mouse monoclonal anti-Flag® M2 antibody, Sigma-Aldrich; Fig. 2a). Attention should be given to the choice of Wnt-specific antibody to be used. For example, we previously showed using RNA-seq that differentiating human embryonic stem (hES) cells express

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WNT3, WNT3A, and WNT5A [15], and in Fig. 3 we confirm the presence of WNT3/WNT3A and WNT5A in the CM of hES cells. 5. In the specific cases shown here (Figs. 2 and 3) all procedures were performed at room temperature. However, in cases where little is known about the stability of a Wnt protein it would be advisable to perform these steps on ice. Note that the WNT9A immunoblot shows a prominent lower band (Fig. 2a), which may be the result of degradation. Such degradation may be minimized by performing all manipulations on ice. 6. In the case of overexpressed Wnt proteins, we generally condition the media for 3–4 days once cells have reached confluency. The time frame for conditioning without media changes may need to be adjusted for different cell types depending on their growth rates, metabolic activity, and overall health at high cell densities. 7. Depending on the viscosity of the CM, the syringe filter may clog. If this occurs, use multiple filters or use a filter with a 0.45 μm pore size. 8. Most Wnt proteins migrate at an apparent molecular weight of 40–50 kD on SDS-PAGE. Therefore, a polyacrylamide concentration of 10–12% is suitable for these gels. However, tagged Wnt proteins are accordingly larger as demonstrated in Fig. 2b for EGFP-tagged WNT3. 9. Transfer of Wnt3a and Wnt5a to nitrocellulose using electroblotting is complete after 1 h at 70 V. This parameter may need to be optimized for different Wnt proteins. References 1. Takada R, Satomi Y, Kurata T, Ueno N, Norioka S, Kondoh H, Takao T, Takada S (2006) Monounsaturated fatty acid modification of Wnt protein: its role in Wnt secretion. Dev Cell 11(6):791–801. https://doi.org/10. 1016/j.devcel.2006.10.003 2. Willert K, Brown JD, Danenberg E, Duncan AW, Weissman IL, Reya T, Yates JR 3rd, Nusse R (2003) Wnt proteins are lipid-modified and can act as stem cell growth factors. Nature 423(6938):448–452. https://doi.org/10. 1038/nature01611 3. Loh KM, van Amerongen R, Nusse R (2016) Generating cellular diversity and spatial form: Wnt signaling and the evolution of multicellular animals. Dev Cell 38(6):643–655. https:// doi.org/10.1016/j.devcel.2016.08.011 4. Hsiung F, Ramirez-Weber FA, Iwaki DD, Kornberg TB (2005) Dependence of

drosophila wing imaginal disc cytonemes on decapentaplegic. Nature 437(7058):560–563. https://doi.org/10.1038/nature03951 5. Mulligan KA, Fuerer C, Ching W, Fish M, Willert K, Nusse R (2012) Secreted winglessinteracting molecule (swim) promotes longrange signaling by maintaining wingless solubility. Proc Natl Acad Sci U S A 109(2):370–377. https://doi.org/10.1073/ pnas.1119197109 6. Mihara E, Hirai H, Yamamoto H, TamuraKawakami K, Matano M, Kikuchi A, Sato T, Takagi J (2016) Active and water-soluble form of lipidated Wnt protein is maintained by a serum glycoprotein afamin/alpha-albumin. eLife 5:e11621. https://doi.org/10.7554/ eLife.11621 7. Gross JC, Chaudhary V, Bartscherer K, Boutros M (2012) Active Wnt proteins are secreted

Detection of Wnt Proteins on exosomes. Nat Cell Biol 14(10):1036–1045. https://doi.org/10. 1038/ncb2574 8. Korkut C, Ataman B, Ramachandran P, Ashley J, Barria R, Gherbesi N, Budnik V (2009) Trans-synaptic transmission of vesicular Wnt signals through Evi/Wntless. Cell 139(2):393–404. https://doi.org/10.1016/j. cell.2009.07.051 9. Kaiser K, Gyllborg D, Prochazka J, Salasova A, Kompanikova P, Molina FL, Laguna-Goya R, Radaszkiewicz T, Harnos J, Prochazkova M, Potesil D, Barker RA, Casado AG, Zdrahal Z, Sedlacek R, Arenas E, Villaescusa JC, Bryja V (2019) WNT5A is transported via lipoprotein particles in the cerebrospinal fluid to regulate hindbrain morphogenesis. Nat Commun 10(1):1498. https://doi.org/10.1038/ s41467-019-09298-4 10. Panakova D, Sprong H, Marois E, Thiele C, Eaton S (2005) Lipoprotein particles are required for hedgehog and wingless signalling. Nature 435(7038):58–65. https://doi.org/ 10.1038/nature03504 11. Willert KH (2008) Isolation and application of bioactive Wnt proteins. Methods Mol Biol 468:17–29. https://doi.org/10.1007/978-159745-249-6_2 12. Ross J, Busch J, Mintz E, Ng D, Stanley A, Brafman D, Sutton VR, Van den Veyver I, Willert K (2014) A rare human syndrome provides genetic evidence that WNT signaling is

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required for reprogramming of fibroblasts to induced pluripotent stem cells. Cell Rep 9(5):1770–1780. https://doi.org/10.1016/j. celrep.2014.10.049 13. Bauer M, Benard J, Gaasterland T, Willert K, Cappellen D (2013) WNT5A encodes two isoforms with distinct functions in cancers. PLoS One 8(11):e80526. https://doi.org/10. 1371/journal.pone.0080526 14. Richter J, Stanley EG, Ng ES, Elefanty AG, Traver D, Willert K (2018) WNT9A is a conserved regulator of hematopoietic stem and progenitor cell development. Genes (Basel) 9(2):66. https://doi.org/10.3390/ genes9020066 15. Huggins IJ, Bos T, Gaylord O, Jessen C, Lonquich B, Puranen A, Richter J, Rossdam C, Brafman D, Gaasterland T, Willert K (2017) The WNT target SP5 negatively regulates WNT transcriptional programs in human pluripotent stem cells. Nat Commun 8(1):1034. https://doi.org/10.1038/ s41467-017-01203-1 16. Chen G, Gulbranson DR, Hou Z, Bolin JM, Ruotti V, Probasco MD, Smuga-Otto K, Howden SE, Diol NR, Propson NE, Wagner R, Lee GO, Antosiewicz-Bourget J, Teng JM, Thomson JA (2011) Chemically defined conditions for human iPSC derivation and culture. Nat Methods 8(5):424–429. https://doi.org/10. 1038/nmeth.1593

Chapter 9 Using Live Imaging to Examine Early Cardiac Development in Zebrafish Tess McCann, Rabina Shrestha, Alexis Graham, and Joshua Bloomekatz Abstract Visualizing dynamic cellular behaviors using live imaging is critical to the study of cell movement and to the study of cellular and embryonic polarity. Similarly, live imaging can be vital to elucidating the pathology of genetic disorders and diseases. Model systems such as zebrafish, whose in vivo development is accessible to both the microscope and genetic manipulation, are particularly well-suited to the use of live imaging. Here we describe an overall approach to conducting live-imaging experiments with a specific emphasis on investigating cell movements during the early stages of heart development in zebrafish. Key words Zebrafish, Cardiac development, Collective movement, Cell migration, Time-lapse, Live imaging

1

Introduction Dynamic processes such as polarized protrusions and cellular movements are critical to embryonic development, physiological homeostasis and the progression of disease [1]. However, techniques which rely on fixation are of limited use in studying these dynamic behaviors since they provide only a snap shot of cellular behaviors and fragile transient structures can be lost during fixation and processing [2, 3]. Live-imaging approaches which visualize events as they occur in real-time are therefore essential for studying dynamic cellular processes. Zebrafish is a vertebrate model system uniquely suited to live imaging due to their external fertilization, rapid development, and the wide range of available genetic and transgenic tools [4, 5]. Indeed, several protocols have been developed to use zebrafish to visualize in vivo cell movements during gastrulation [6], posterior body elongation [7], somitogenesis [8], and cranial

Supplementary Information: The online version of this chapter (https://doi.org/10.1007/978-1-0716-20359_9) contains supplementary material, which is available to authorized users. Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_9, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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neural crest cell migration [9], among others. Differences between these protocols reveal that live-imaging experiments often must be tailored to the developmental stage and tissue being analyzed. During zebrafish heart tube formation, cardiomyocytes which originate in bilateral domains within the anterior lateral plate mesoderm move to the midline to form a ring of cells, in a process known as cardiac fusion [10]. This process of medial movement occurs in all vertebrates [11], is essential for heart development and provides a model for understanding collective cellular movement and tissue– tissue communication [12–16]. We have used live imaging to identify mechanisms underlying the movement of both cardiomyocytes and endocardial cells during cardiac fusion [13, 17]. In this chapter, we identify a few broad guidelines for conducting live-imaging experiments that can be applied to the study and visualization of a variety of dynamic processes. And we present a specific protocol for the live imaging of cardiomyocyte movement during the early stages of heart tube formation.

2 2.1

Materials Mounting

1. Coverslip bottom D35-20-1.5N).

dish

(CellVis,

D60-30-1.5N

or

2. 50
E3 medium: 14.61 g NaCl, 0.63 g KCl,1.83 g CaCl2·H2O, 1.99 g MgSO4·7H2O in 1 L dH2O. Dilute to 1
with dH2O. 3. 2% (g/ml) agarose (GoldBio, A-201-500) in 1
E3 medium. 4. 0.8% low melting temperature agarose (GoldBio, A-204-25) in 1
E3 medium (see Note 1). 5. Heat block with multiple holding spots (VWR, 10153-348). 6. Small-bore Pasteur pipette (Fisherbrand, 13-678-6A). 7. Wide-bore Pasteur pipette (Fisherbrand, 13-678-30). 8. Pipette pump (SP Scienceware, 37898-0000). 9. 4% Tricaine: 2 g Tricaine-S (Western Chemical, A02F01G), 10.6 ml 1 M Tris pH 9 in 500 ml of dH2O. Adjust pH to 7.4. 10. Tricaine/E3 solution: 1.2 ml 4% Tricaine-S, 30 ml 1
E3 medium. 11. Forceps for dechorionation and embryo orientation (Fisherbrand 12-000-122 or similar). 2.2

Imaging

1. Stereo dissecting microscope scope for mounting embryos in the mold (e.g., Zeiss Stemi508). 2. Inverted confocal microscope with a motorized stage and applicable software. This protocol was developed using a Leica TCS SP8 X confocal microscope, although it is also

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suitable for use with other inverted microscopes. Upright microscopes, however, require a different mounting technique [18]. The system must be configured with lasers capable of exciting the fluorophores in the samples. 3. Objectives such as HC PL APO CS2 20
/0.72 or HC PL APO 40
/1.10 with long working distances, 0.68 mm and 0.65 mm, respectively, are encouraged because cardiac cells during the early stages of heart development are located deep within the embryo between the yolk and neural tube. 4. Temperature control devices fitted to the microscope for maintaining 28.5  C, the optimal temperature for zebrafish growth [19]. For example, the Okolab temperature controller H201T-UNIT-BL with H201-Enclosure.

3

Methods

3.1 General Guidelines

In a live-imaging experiment there are at least four main phases to consider: experimental design and preparation, mounting of the sample, imaging, and analysis (see Fig. 1 for workflow). Throughout these phases, following a few guidelines has helped us conduct successful experiments (summarized in Box 1).

Box 1 General Guidelines and Tips 1. Design and tailor the experiment including the imaging choices to the requirements of your specific developmental or cellular process. 2. Break down a complicated developmental process into small parts to image first and then build incrementally toward trying to capture the entire process. 3. Make the experiment easy by minimizing obstacles. 4. Ensure images from healthy embryos are analyzed by culturing after imaging. 5. Use semiautomated segmentation software.

3.1.1 Tailoring

Many decisions occur while conducting a live-imaging experiment and tailoring these choices toward a specific question or process rather than trying to visualize everything can help to ensure success. For example, individual light microscopes from compound microscopes to confocal microscopes (spinning disk, point-scanning, or light sheet) to multiphoton microscopes are each designed for a specific imaging challenge related to requirements of resolution, tissue depth, and duration or speed of imaging. Often fulfilling one

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Fig. 1 Experimental workflow for a live-imaging experiment focused on cardiac fusion in zebrafish. (a) Experimental design—Embryos are collected from mating two homozygous Tg(myl7:eGFP) fish, which express eGFP in myocardial cells. (b) Mounting—Embryos at the 12 somite stage are mounted head down in low melting temperature agarose in an agarose mold in which holes have been punched out. A mark on the dish (shown in red) allows one to identify embryos

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imaging requirement comes at the expense of another. Spinning disk microscopes, for example, are designed for capturing very quick events with minimal light damage but can lack sufficient axial resolution for processes that occur in the Z-plane [20]. Similarly, within a microscope there are many parameters that can be adjusted. For example, on point-scanning confocal microscopes, one can adjust parameters such as laser power, image size, scan speed, pinhole size, and line/frame average and accumulation, among others. Changing these parameters often involves a tradeoff between image quality, speed and embryo or cellular health. Thus, tailoring these decisions to a specific developmental or cellular process can help to ensure success. Furthermore, if the developmental process is particularly complicated it may be easier to conduct a live-imaging experiment in parts and build incrementally toward a comprehensive visualization. 3.1.2 Minimize Obstacles

Taking steps to minimize the obstacles of a live-imaging experiment, even if they are small, can also improve the probability of success. For example, during the experimental design phase one can maximize the imaging capability of the microscope and the model system by considering how the different properties of the fluorophores including relative brightness and maturation speed (see Note 2) [21] complement the microscope and model system. Using animals that are homozygous for a transgene can also minimize imaging obstacles since one does not have to prescreen the embryos and all embryos have the same brightness. Similarly, using a light-box to time shift a developmental process away from an inconvenient time period can make it easier to repeat the liveimaging experiment multiple times.

3.2 Cardiomyocyte Specific Protocol

Below we describe our protocol for the live imaging of cardiomyocyte movement during cardiac fusion using a Leica SP8 confocal microscope (see Fig. 1 for workflow).

ä Fig. 1 (continued) via their position in the mold. (c) Imaging—Embryos in the best orientation are imaged with an inverted confocal microscope containing an incubation chamber. (d, d0 ) Preprocessing—Stacks of optical sections from both the brightfield (d) and eGFP (d0 ) channels are collected approximately every 3 min over a 4 h time-period. The tip of the notochord (indicated by the arrow) as visualized in the brightfield image (d) is used as a reference point to correct for embryo drift that may occur during imaging. (e, e0 ) Segmentation and Tracking—Using images of eGFP, cells are identified in the first timepoint (e) and then their positions are tracked (e0 ) in subsequent timepoints. (f, f0 ) Analysis—Graphs depict cell movement properties including average displacement (f) and overall directionality (f0 ) of individual cardiomyocytes

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3.2.1 Preparation

1. Set up adult zebrafish (see Note 3) containing transgenes that fluorescently label cardiomyocytes (e.g., Tg(myl7:eGFP)) [22] or endocardial cells (e.g., Tg(flk1:eGFP)) [23]. Use dividers to separate males and females in order to control the timing of breeding. 2. The next morning lift the dividers and collect embryos every 30 min in separate dishes to ensure all embryos in a dish are of similar age. 3. To visualize cell membranes or other intracellular structures one can inject DNA or mRNA into embryos at the one-cell stage. For example, in Fig. 2 we have injected plasmid DNA containing nkx2.5:lck-mScarlet-i into Tg(myl7:eGFP) embryos in order to visualize the cell membrane of cardiomyocytes. See [24] for microinjection procedures. 4. Place embryos into an incubator set to 28.5  C and leave them to develop until they reach the age of interest. 5. Dissolve 0.8% low melting temperature agarose in 5 ml of 1
E3 medium in a 15 ml conical tube by heating to 70  C in a water bath and occasionally inverting. Then aliquot the 0.8% low melting temperature agarose/E3 solution into 1.5 ml microfuge tubes and place on a heat block set at 42  C (see Fig. 3). Do this at least an hour before mounting to allow the solution to equilibrate at 42  C.

Fig. 2 Mosaic labeling via plasmid DNA injection. (a, a0 ) 3D projection (a) and single optical section (a0 ) of cardiomyocytes from a 20 somite stage Tg(myl7: eGFP) (green) embryo which was injected (Inj) with a plasmid containing nkx2.5: lck-mScarlet-i (magenta) when it was at the 1-cell stage. (b, b0 ) Magnification showing only nkx2.5:lck-mScarlet-i expression (black) from the area in the white boxes in a, a0 respectively. Scale bar: 20 μm. See also Electronic Supplementary Video 1

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Fig. 3 Mounting technique for visualizing cardiac fusion in zebrafish embryos. (a) Wells in a 2% agarose/E3 layer on a coverslip bottom dish are made using a small-bore Pasteur pipette. One side of the coverslip is marked (indicated by the red dot) to identify the position of the mounted embryos. (b) Microfuge tubes with 0.8% low melting temperature agarose/E3 are maintained at 42  C on a heat block. (c) A single dechorionated embryo is transferred to and then reacquired from the low melting temperature agarose/E3 solution in a widebore Pasteur pipette. The embryo and the low-melting temperature agarose/E3 is discharged from the pipet into a well on the coverslip bottom dish. (d, e) The embryo is positioned in such a way that the cardiomyocytes (green) are perpendicular to the coverslip. (f) The low-melting temperature agarose is left to solidify and then Tricaine/E3 solution is added to the dish

6. Prepare 5 ml of a 2% regular agarose/E3 solution and use this to coat a coverslip bottom dish (see Note 4). Do this 1 h before mounting to allow it to solidify completely. 7. Turn on the temperature controller for the imaging chamber and leave it to equilibrate at 28.5  C (see Note 5). 3.2.2 Mounting

1. Use a small-bore Pasteur pipette to create a well in the 2% agarose/E3 layer. Remove any residual agarose with forceps. Ensure the well reaches all the way down to the coverslip bottom (see Fig. 3). 2. Once embryos have reached the 12 somite stage start mounting (see Note 6). 3. Transfer embryos to a petri dish with Tricaine/E3 solution (see Note 7). 4. Use forceps to manually dechorionate embryos. Do this carefully to avoid damaging them. Discard embryos in which the yolk has been pierced (see Note 8). 5. Use a wide-bore Pasteur pipette to transfer a single embryo into a tube containing low melting temperature agarose. Try to transfer as little liquid as possible. Once the embryo is in the

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low melting temperature agarose, discharge the rest of the liquid from the pipette. Quickly, reacquire the embryo from the low melting temperature agarose with the same pipette (see Fig. 3) and place it in one of the wells in the agarose mold. Place the tube with low melting temperature agarose back in the heat block (see Note 9). 6. Position the embryo using forceps as close to the coverslip as possible. For imaging cardiac fusion, the embryo should be perpendicular to the coverslip with the head down, at a slight angle to align the ALPM to the coverslip (see Fig. 3 and Notes 10–12). 7. Let the low melting temperature agarose/E3 solution solidify. Repeat steps 5 and 6 to mount other embryos. Be careful not to disrupt the solidifying wells. During the process of mounting, if wells that have already been mounted look like they are drying out, add a small amount of E3 over these wells. 8. Once all embryos have been mounted and the agarose solidified, cover all embryos and the dish with the Tricaine/E3 solution. 3.2.3 Imaging

1. At the confocal, examine the mounted embryos to identify the embryo(s) in the best orientation. The goal is to find ones that are not tilted to one side, that are close to the coverslip and have the brightest GFP intensity (see Note 13). 2. Use the Mark-and-Find function on the confocal to select these embryos. Using a mark on the imaging dish, keep track of the identity of the embryo(s) for later verification and genotyping. 3. Use as little laser power as possible for a good image (see Note 14). 4. When adjusting the parameters of the microscope, use the Quick LUT mode to visualize pixel intensities. Avoid both too low and too high pixel intensities. 5. Create a Z-stack that covers the depth of the cardiomyocytes in all embryos. This is typically 30 Z-planes of 1 μm thickness. A wide Z-stack accommodates slight drift of the embryo in the axial direction without having to readjust the settings during imaging. 6. For assessing overall cell movements a Z-stack from an individual embryo is recorded approximately every 3 min. However, visualization of highly dynamic processes such as protrusions can require shorter time-intervals and high scan speeds. 7. For cardiac fusion the total running time is usually 3–4 h (see Notes 15 and 16). 8. For cell tracking and image analysis ensure the files are saved in a format which conserves all the metadata.

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9. After imaging, gently remove the embryo(s) from the mold and incubate in 1
E3 for 24 h to confirm imaging did not damage the embryos. Embryos are incubated individually in 6 well plates. After incubation embryos can be genotyped. Individual embryos are matched to their 4D imaging stack via their unique location in the imaging dish. 3.2.4 Preprocessing of Images

A preprocessing step is often necessary before segmentation and image analysis. This preprocessing step improves the image quality and corrects for artifacts introduced during imaging, such as embryo drift. 1. To correct for embryo drift the ‘Correct 3D drift’ plugin [25] available through FIJI [26] can be used. For cardiomyocyte imaging, the tip of the notochord identified in brightfield images (see Fig. 1) is used as the reference point. 2. Depending on the processing power of the computer, this program can take several minutes to perform the correction. Typically, a 3.5 h video of 45 timepoints with 25 slices each takes about 30 min to correct.

3.2.5 Cell Tracking Analysis

1. To track cells along the XY axis a maximum intensity projection of the eGFP channel is created and this is used for segmentation and tracking analysis. 2. The mTrackJ plugin available through FIJI is particularly useful for tracking cardiomyocytes (see Fig. 4) [27]. A complete user manual for the Plugin can be found online [28] (see Note 17). 3. Properties of cell tracks such as displacement, duration, velocity, directionality, coherence and neighbor exchange can be analyzed via MATLAB (MathWorks) or other software programs (see Fig. 1). 4. These properties can be assessed and compared to cell tracks from embryos treated with pharmacological agents or from mutant embryos. Pilot studies are recommended for determining the sample sizes required to achieve sufficient statistical power. If embryos generated from heterozygous parents are genotyped after imaging and analysis, the researcher is naturally blinded from bias.

4

Notes 1. Several different compounds can be used for mounting including, low-melting temperature agarose, methylcellulose, fluorinated ethylene propylene [29] or a combination thereof [12]. 2. Due to the quick development of zebrafish embryos, fluorophore maturation speed is an important property to consider.

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Fig. 4 Live-imaging of cardiac fusion in a wild-type Tg(myl7:eGFP) embryo. (a–a000 ) 3D reconstructions of confocal slices at select timepoints during the live-imaging of cardiac fusion in a wild-type Tg(myl7:eGFP) embryo. (b–b000 ) Tracks of cardiomyocyte movement created using the mTrackJ FIJI plugin show the early movement of cardiomyocytes toward the embryonic midline. (a, b) Initial position of visible cardiomyocytes (arrows in a). Images show the phases of cardiomyocyte movement including the early medial movement of cardiomyocytes (a0 , b0 ), merging of posterior cardiomyocytes (a00 , b00 ), and finally the merging of anterior cardiomyocytes (a000 , b000 ) to form a ring. Tracks displayed show the paths travelled by a subset of the cardiomyocytes during a–a000 . Scale bar: 60 μm. See also Electronic Supplementary Video 2

3. To increase the brightness of the signal, use animals that are homozygous for the fluorophores. This maximizes what can be observed at the confocal. 4. When making the 2% agarose/E3 layer, it is helpful to tilt the dish back and forth, because it is quite viscous. The taller the wells the more difficult it is to orientate the embryos correctly. The layer should be 1.2 mm thick, use 500 μl for a 35 mm dish or 1 ml for a 55 mm dish. 5. Equilibrating the imaging chamber at the appropriate temperature prevents focal drift and ensures embryonic development continues as expected. Check that it is working correctly before starting the recording. 6. As the mounting process will take time, start before the time frame you wish to image. It can take approximately 1 h to mount 20–30 embryos. 7. Avoid media containing methylene blue or phenol red due to their autofluorescent properties. 8. Use glass bottom or agarose coated dishes to hold dechorionated embryos, if embryos have not finished gastrulation ( 30 exonuclease that can degrade double strand (ds) DNA template to expose ssDNA ends for recombination. Working together, they can promote highly efficient homologous recombination to induce any desired genetic changes in the BAC using either ssDNA or dsDNA as templates [34, 35]. Finally, a temperature sensitive λ repressor (allele CI857) was incorporated to control Gam, Beta, and Exo transcription to avoid their toxic effect. The repressor is active at low temperature (~32  C), but can be rapidly inactivate with a 15 min incubation at 42  C to provide a pulse of Gam, Beta, and Exo expression. We will provide a step-by-step protocol on how to (1) identify the appropriate BAC clone, (2) introduce the BAC into the recombination competent strains, (3) design the strategy to create the desired changes, (4) carry out the BAC recombineering, and (5) purify the engineered BAC for pronuclear injection to acquire transgenes (Fig. 1).

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Table 1 Summary of three commonly used recombineering bacterial strains List of recombineering competent strains Name

Genotype

Note

DY380 DH10B [λcl857 (cro-bioA < > tet]

DH10B cell with integrated λ prophage that expresses Gam, Beta, and Exo under the control of temperature sensitive cl857 repressor. Tetracycline (12.5 μg/mL) resistant

EL250 DH10B [λcl857 (cro-bioA < > araC-PBADflpe]

Same as DY380 except also containing an arabinose-inducible flippase that can excise sequence flanked by FRT sites, such as FRT-neo-FRT introduced through construct PL451 (see Note 8). Not tetracycline resistant

EL350 DH10B [λcl857 (cro-bioA < Same as DY380 except also containing an arabinose-inducible > araC-PBADcre] Cre recombinase that can excise sequence flanked by LoxP sites, such as LoxP-neo-LoxP introduced through construct PL452 (see Note 8). Can also be used to test any BACs that are designed to function as conditional transgenes with floxed sequence. Not tetracycline resistant These strains can be requested from NCI through the following link: https://frederick.cancer.gov/Science/ BrbRepository/#/productDataSheets/Bacteria PL451 and 452 plasmids can be requested from NCI through the following link: https://frederick.cancer.gov/Science/BrbRepository/#/productDataSheets/Plasmid

Fig. 1 Workflow of BAC recombineering and transgenesis

2

Materials

2.1 Media and Antibiotics for BAC Culture

1. Chloramphenicol stock: make 25 mg/mL stock in ethanol; aliquot and store at 20  C. 2. LB (Luria-Bertani) media and agar. 3. LB/chloramphenicol plate: LB plate with 12.5 μg/mL chloramphenicol. 4. LB/chloramphenicol media: LB media with 12.5 μg/mL chloramphenicol.

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1. P1 solution: 50 mM Tris–HCl pH 8.0; 10 mM EDTA; 100 μg/mL RNaseA. 2. P2 solution: 200 mM NaOH; 1% SDS. 3. P3 solution: 3.0 M potassium acetate, pH 5.5. 4. Ethanol. 5. TE: 10 mM Tris, 1 mM EDTA, pH 8.0. 6. 15 and 50 mL conical tubes. 7. Eppendorf tubes. 8. Centrifuge.

2.3 Restriction Digest and Pulse Field Gel Electrophoresis (PFGE)

1. Restriction enzyme and buffer. 2. PFGE system: for instance, the CHEF-DR II system from Bio-Rad. 3. PFGE marker (such as MidRange PFG Marker from NEB cat.# N0342S). 4. Agarose. 5. 0.5 TBE (Tris/Boric Acid/EDTA) running buffer: 45 mM Tris; 45 mM boric acid; 1 mM EDTA, pH 8.3. 6. Nucleic acid dye such as ethidium bromide for DNA staining.

2.4 Electroporation of BAC DNA into Recombineering Competent Strains

1. Autoclaved water, chilled to 4  C. 2. 1 mm electroporation cuvettes. 3. An electroporator such as the BIO-RAD Gene Pulser Xcell system. 4. Recombineering competent strains (Table 1). The strains can be requested from NCI through the following link: https:// frederick.cancer.gov/Science/BrbRepository/#/ productDataSheets/Bacteria

2.5 BAC Recombineering with Linear Template

1. Linear recombineering template (see Subheading 3.5 and Note 9).

2.6 Purification of BAC DNA for Pronuclear Injection

1. Qiagen Large Construct Kit; cat.# 12,462.

2. Autoclaved water, electroporation cuvettes and electroporator as in Subheading 2.4.

2. Ethanol. 3. Phenol–chloroform–isoamyl alcohol (25:24:1). 4. Chloroform. 5. Isoamyl alcohol. 6. TE: 10 mM Tris, 1 mM EDTA, pH 8.0. 7. Sodium acetate, pH 5.2.

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8. Low TE (10 mM Tris, 0.1 mM EDTA, pH 7.5). 9. Injection buffer: 10 mM Tris–HCl, pH 7.5, 0.5 mM EDTA, 30 nM spermine, 70 nM spermidine, 100 mM NaCl. 10. Fluorometer and/or spectrometer. 11. Centrifuges. 2.7 Use BAC Transgenic Mice to Study Polarized Tissue Morphogenesis

1. 4% PFA: 4% paraformaldehyde in PBS, pH 7.5. 2. PBS with 15% sucrose. 3. PBS with 15% sucrose. 4. OCT: Optimal cutting temperature compound. 5. Ethanol or butanol and dry ice for snap-freezing the specimen. 6. 1 PBS, 0.5% Tween 20. 7. FITC-conjugated phalloidin. 8. PBST: 1 PBS, 0.1% Tween 20. 9. RIMS (Refractive index matching solution): 88% (weight by volume) Histodenz (Sigma); 0.02 M phosphate buffer; 0.01% sodium azide; 0.1% Tween 20 (Sigma); 2.5% 1,4-Diazabicyclo [2.2.2]octane (Sigma). 10. Superglue for making imaging chamber. 11. Cryostat and blades for trimming the embryo. 12. Petri dish (with cover slip bottom for inverted microscope; or regular to make imaging chamber for upright microscope). 13. Confocal microscope.

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Methods

3.1 Identify and Acquire the Desired BAC Clones

1. Go to the Ensembl genome browser Archive 88 (http:// mar2017.archive.ensembl.org/index.html) (see Note 1). 2. Under “Search,” select the desired species and enter the gene name (for instance, select “mouse” and enter “Rosa26”), then click “Go”. 3. From the search results displayed, click the link of the desired gene name (for instance for Rosa26, click the link “Gt(ROSA) 26Sor (Mouse Gene, Strain: reference (CL57BL6))“). 4. On the gene summary page, scroll down and click “Region in Detail”; then click the “Configure this page” pane on the left hand side. 5. An “Active tracks” window will appear. Click the “Clones & misc. regions” pane on the left hand side. 6. Check the square boxes for the clones to be displayed. For mouse BAC clones, click on “RP23” and “RP24 mouse

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clones”; then click the check mark on the upper right hand corner to confirm the selection and close the window (see Note 2). 7. Scroll down to “Contigs” to view the BAC clones. To better view the gene in the context of the BAC clone, it might be necessary to zoom out by clicking the arrows in the “Location” pane, or by dragging the zoom bar to the right (toward “-”). 8. Choose the clone(s) in which the gene of interest is located in the middle. Click on the clone to display the information on its genome location and size. For instance, when clicking on the BAC clone RP24-85 L15 that contain the Rosa26 gene, it will display its genome coordinates as “Range: 112952746–113,158,583” and size as “Length: 205838 bps”. 9. Copy the clone name. Go to the website “bacpacresources. org” (a distributor of BAC clones). Paste or enter the clone name under the “Clone Information Search” pane, click “Search”. 10. Click “LINK” under “ORDERING & PRICING” and follow the instruction displayed to order the BAC clone(S). 3.2 DNA Purification from BAC Clones

BAC clones are shipped as bacterial stab cultures (usually DH10B cells) instead of DNA. This is because due to its large size, purified BAC DNA cannot be stored as frozen stock but only temporarily at 4  C. For long-term storage, bacteria stock of BAC clones are kept at 80  C. Therefore, upon receiving the BAC clone culture from BACPAC, it is necessary to amply the bacteria for restriction digest to confirm that they contain the correct BAC DNA. The bacterial culture containing the confirmed BAC DNA should then be frozen down and store at 80  C. The purified DNA will also need to be electroporated into the recombinant competent strains (Table 1) for BAC recombineering. 1. Using an inoculation loop, steak the received BAC culture on an LB/chloramphenicol plate (Subheading 2.1, step 2), incubate at 37  C overnight. 2. Pick 3 different colonies, and inoculate them each into 11 mL LB/chloramphenicol media (Subheading 2.1, step 3) in a 50 mL conical tube, shake at 250 rpm at 37  C overnight. 3. Save 1 mL of the bacterial culture in an autoclaved Eppendorf tube to make frozen cell stock (see Subheading 3.3, step 5). Spin down the rest of the bacterial culture. Resuspend the pellet in 250 μL P1 solution and transfer to an 1.7 mL Eppendorf tube. 4. Add 250 μL P2 solution to each Eppendorf tube. Invert the tubes several times to mix P2 with the resuspended bacterial culture. Incubate 5 min at room temperature.

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5. Add 350 μL P3 to each tube. Invert to mix. Spin 3 min at 14,000  g or above (~17,900  g or above). 6. Transfer supernatant to a new Eppendorf tube, spin again for 3 min to further clear the extract. 7. Transfer supernatant to a new Eppendorf tube (~800 μL), add 750 μL isopropanol. Invert the tubes to mix. Allow DNA to precipitate for 10 min at room temperature. 8. Centrifuge the tubes for 5 min to collect DNA pellets. Discard the solution and wash the DNA pellet with 1 mL 70% ethanol. 9. Spin again and discard ethanol. Do a quick spin and carefully suck up any remaining ethanol solution with a pipet. Let the pellets dry for 3 min at room temperature with the lid open (see Note 3). 10. Add 30 μL TE buffer to each tube. Let the solution soak the pellet for 10 min at room temperature to resuspend DNA. Gently tap the tube to help dissolving the DNA, or allow the DNA continuing to dissolve at 4  C for a few hours or overnight. 11. Spin down the DNA solution at 14,000  g or above for 5 min to clear the solution. Transfer the cleared supernatant to a new tube (see Note 4). 12. Store the DNA at 4  C, or continue with Subheading 3.3 for restriction digest. 3.3 Restriction Digest and Pulse Field Gene Electrophoresis (PFGE) to Characterize the Purified BAC DNA

The standard method to determine the integrity of the BAC clone is to digest the purified DNA with rare-cutter enzymes (such as NotI). The resulting DNA fragments from the restriction digest are usually between ~20 and 100 kb and will require running PFGE for 20–24 h to resolve them. 1. Use 10–15 μL of the purified DNA to set up overnight restriction digest with a rare-cutter such as NotI (see Note 5). 2. Set up PFGE with a system such as CHEF-DR II (Bio-Rad). Load the restriction digests and PFGE marker into each well of a 1% agarose gel made with 0.5 TBE, and run the gel in chilled 0.5 TBE with the following parameter: 5–25 s switch; 6 volts/cm; 22 h. Use a chiller to keep the buffer cool, or run the PFGE in a cold room. 3. After PFGE, stain the gel in 0.5 μg/mL EtBr solution for 10 min and visualize under UV. Destain the gel in water if necessary. 4. Compare the observed restriction pattern to the predicated pattern (see Note 6), and calculate the size of the BAC clone based on the restriction fragments observed (see Note 7).

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5. Make a frozen cell stock by adding 150 μL of glycerol to the 1 mL bacterial culture saved from step 3 of Subheading 3.2. Vortex to mix and store at 80  C. 3.4 Electroporate BAC DNA into Recombineering Competent Bacterial Strains

Upon confirming the BAC DNA from Subheading 3.3, it will need to be electroporated into one of the recombineering competent strains (Table 1). We will use EL350 as an example in the subsequent procedures, but the protocol will be identical for the other two strains. 1. Inoculate 1–2 μL of frozen EL350 cell stock in 1 mL LB media in a 15 mL conical tube. Shake at 250 rpm at 32  C overnight (see Note 8). 2. The next morning, add 200 μL of the overnight culture in 10 mL LB media in a 50 mL conical tube, shake at 250 rpm at 32  C until OD600 reaches 0.5–0.7 (~4 h). 3. Chill the culture on ice for 20 min. 4. Spin down the cells at 5500  g for 5 min at 4  C. Remove the culture media and resuspend the cells in 1.5 mL of autoclaved water chilled to 4  C. Transfer the cells to an autoclaved Eppendorf tube. 5. Spin down the cells as in step 4, and repeat the wash with ice cold water for two more times. 6. After the last wash, resuspend the cells in 50 μL ice cold water. Add 1–3 μL of miniprepped BAC DNA from Subheading 3.2. Swirl the solution and tap the tube to help mixing BAC DNA with the cells. 7. Transfer the DNA-cell mixture to a 1 mm electroporation cuvette. Make sure the solution is dispensed into the slot between the two aluminum electrode plates. Incubate the cuvette on ice for 5–10 min. 8. Electroporate the cuvette with the following setting: 1.75 kV, 25 μF, 200 ohms. 9. Add 1 mL LB to the cuvette, and transfer the electroporated cells to a 15 mL conical tube, shake at 250 rmp at 32  C for 1.5 h. 10. Spin down the cells at 5500  g for 5 min. Resuspend the cells in 100 μL LB and plate them on an LB/chloramphenicol plate. Incubate at 32  C overnight. 11. Pick three colonies and culture each of them in 11 mL LB/chloramphenicol media. Save 1 mL of the culture for frozen cell stock, and use the remaining 10 mL to purify BAC DNA as in Subheading 3.2. Characterize the DNA as in Subheading 3.3 to confirm that the BAC DNA in EL350 cells is the same as the original clone.

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12. Make a frozen cell stock by adding 150 μL of glycerol to the 1 mL bacterial culture saved from step 11 above. Vortex to mix and store at 80  C. 3.5 BAC Recombineering with Linear Template

The design of the linear template for recombineering will depend on what type of modification one wants to engineer, and how precise the modification needs to be. See Note 9 for a general discussion of two main strategies that are commonly used to engineer different modifications ranging from insertions, deletions and point mutations. Once a strategy is chosen, either PCR or cloning can be used to add homology arms to flank the modification cassette (for instance, LoxP sites, Cre or fluorescent protein sequence that are to be inserted). If using the PCR method, primers can be designed to directly add homology arms up to 80 bp on each end to generate the template. Alternatively, for increased recombineering efficiency, longer homology arms (200–500 bp) can be added to each end of the modification cassette through PCR mediated cloning [38], and the template is then excised out of the cloning vector through restriction digest. We have found that 200 bp homology arms are sufficient for inserting a 9 kb cassette into the Rosa26 locus in a BAC through recombineering [31]. Finally, if the modification to be made is a point mutation or deletion, the template can also be a single strand oligo with up to 50 base homology arms flanking the point mutation or deletion breakpoints. If the linear recombineering template is made via PCR or restriction digest of a construct after cloning, it should be gel purified with Qiagen Gel Extraction Kit (or something similar) and eluted in sterile water before proceeding to the following step. It is important that the template is free of contamination from primers, nucleotides and other DNA, and ideally with a concentration above 50 ng/μL. Oligos can be directly diluted in water to ~100 ng/μL. 1. Inoculate 1–2 μL of frozen cell stock from step 12 of Subheading 3.4 in 1 mL LB/chloramphenicol media in a 15 mL conical tube. Shake at 250 rpm at 32  C overnight. 2. The next morning, add 200 μL of the overnight culture in 10 mL LB/chloramphenicol media in a 50 mL conical tube, shake at 250 rpm at 32  C for ~4 h until OD600 reaches 0.5–0.7. 3. Place the culture tube in a 42  C shaking water bath for 15 min to induce the expression of Gam, Beta, and Exo. 4. Chill the culture on ice for 20 min. 5. Spin down the cells at 5500  g for 5 min at 4  C. Remove the culture media and resuspend the cells in 1.5 mL of autoclaved water chilled to 4  C. Transfer the cells to an autoclaved Eppendorf tube.

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6. Spin down the cells as in step 5, and repeat the wash with ice cold water for two more times. 7. After the last wash, resuspend the cells in 50 μL ice cold water. Add 100–300 ng linear DNA template. Mix the template DNA with the cells. 8. Transfer the template DNA-cell mixture to a 1 mm electroporation cuvette. Incubate the cuvette on ice for 5–10 min. 9. Electroporate the cuvette with the following setting: 1.75 kV, 25 μF, 200 ohms. 10. Add 1 mL LB/chloramphenicol to the cuvette, and transfer the electroporated cells to a 15 mL conical tube, shake at 250 rmp at 32  C for 1.5 h. 11. Spin down the cells at 5500  g for 5 min. Resuspend the cells in 100 μL LB and plate them on an LB/chloramphenicol plate. Incubate at 32  C overnight. 12. Pick colonies and culture them in 100 μL LB/chloramphenicol in a 96 well plate. Shake at 125 rpm at 32  C overnight. Use 1–2 μL of the bacterial culture as PCR template to screen for recombined clones. 13. Once candidate clones are identified, streak the culture on LB/chloramphenicol plates and incubate at 32  C overnight. Pick single colonies and grow in 11 mL LB/chloramphenicol media. Save ~1 mL to make frozen cell stock, and to further confirm that they contain the correct modification(s) by PCR and/or sequencing of the PCR reaction. Use the remaining 10 mL to purify BAC DNA as in Subheading 3.2. Characterize the DNA as in Subheading 3.3 to confirm that the modified BAC has the correct size and restriction digest pattern in comparison to the original clone. 3.6 Purification of BAC DNA for Pronuclear Injection

The easiest method to acquire high quality BAC DNA suitable for pronuclear injection in mice is to first perform purification with Qiagen Large Construct Kit (cat.# 12,462), and then extract DNA with phenol–chloroform [39]. It requires no special equipment such as ultracentrifuge, and can be carried out within 3 days in any labs with basic molecular biology set up. 1. Streak frozen cell stock from step 12 of Subheading 3.5 onto an LB/chloramphenicol plate. Incubate at 32  C overnight. 2. Inoculate a single colony in 2 mL LB/chloramphenicol media. Shake at 250 rpm at 32  C overnight. 3. Dilute the overnight culture 1:1000 into 500 mL LB/chloramphenicol media. Shake at 250 rpm at 32  C for 20–24 h (see Note 10). 4. Proceed with BAC DNA purification with Qiagen Large Construct Kit. After ethanol precipitation of the BAC DNA eluted

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from column, skip the 70% ethanol wash and instead dissolve the BAC DNA pellet in 500 μL TE. There is no need to dry the pellet before dissolving in TE either. 5. Determine the DNA concentration with a spectrometer or fluorometer. Typical yield of BAC DNA from the large construct kit is 10–50 μg/500 mL culture, so the expected concentration usually ranges from 20 to 100 ng/μL. 6. Transfer the DNA solution to a clean Eppendorf tube and mix DNA solution with equal volume (500 μL) of phenol–chloroform–isoamyl alcohol (25:24:1). Do not vortex but invert tube multiple time to gently mix DNA with phenol–chloroform. Centrifuge the tube at 8600  g for 3 min. Transfer the upper DNA solution to a clean autoclaved Eppendorf tube. 7. Repeat step 3 once with phenol–chloroform–isoamyl alcohol (25:24:1), and once with chloroform–isoamyl alcohol (24:1). 8. Add 1/10 volume (~50 μL) of 3 M sodium acetate and two volume of ethanol (~1 mL). Invert the tube to mix. Centrifuge at 13,000  g for 15 min at room temperature to precipitate DNA. 9. Wash the pellet with 70% ethanol. Centrifuge at 13,000  g for 3 min and remove the liquid. Centrifuge again briefly and use a pipette and pipette tip to suck out any residual liquid. 10. Air dry for 3–5 min. Add 100 μL low TE to the tube. Gently tap the tube and let the pellet dissolve overnight at 4  C. 11. Determine DNA concentration using a spectrometer or fluorometer. The concentration is usually between 50 and 300 ng/μL. Store the DNA at 4  C. 12. Characterize the DNA as in Subheading 3.3 to confirm that the purified BAC has the correct size and restriction digest pattern. 13. The day before the scheduled injection, spin the DNA solution at 13,000  g for 15 min to precipitate any particles that may interfere with pronuclear injection. Carefully transfer the upper half of the DNA solution into a clean autoclaved Eppendorf tube. Measure the DNA concentration in the upper fraction. 14. Depending on the preference of the transgenic facility, dilute the DNA in either injection buffer or low TE. 200–300 μL of the diluted DNA solution is usually enough. Invert the tube to mix. 15. Spin at 13,000  g for 15 min at room temperature to precipitate any residual particles/aggregates. Carefully transfer the upper half of the DNA solution to a clean autoclaved Eppendorf tube. 16. Store the DNA at 4  C before providing to the transgenic facility for pronuclear injection.

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3.7 Use BAC Transgenic Mice to Study Polarized Tissue Morphogenesis

Core PCP genes and presumptive PCP ligands such as Wnt5a/11 regulates diverse morphogenetic events in the mouse. To elucidate their function at the tissue level, we created BAC transgenes in which the tamoxifen-inducible CreERT2 is inserted into BAC clones containing Wnt5a and Wnt11. We used these BAC transgenes in conjunction with the Cre reporter Rosa26tdTomato for genetic labeling to (1) map the expression of Wnt5a and Wnt11 with high spatial and temporal resolution, (2) lineage trace the contribution of Wnt5a- and Wnt11-expressing cells in different organs and tissues, and (3) study the origin and impact of morphogenetic defects in PCP mutants [31–33]. We present below a method to visualize these genetically labeled embryos using wholemount imaging with confocal microscopy and three dimensional reconstruction.

3.7.1 Specimen Preparation

For most conventional confocal microscopes, although objectives can have working distance up to 2 mm and fluorescent signal can be observed as deep as 1 mm into the specimen after clearing, the image resolution often drops significantly after 400–500 μm. Therefore, it is often necessary to first trim the specimens close to the plane where the imaging will start. 1. After tamoxifen induction, collect embryos at the desired times points and fix in 4% PFA at 4  C for 30 min to overnight. 2. Place the fixed embryo in a 10 cm petri dish on its lateral side, and slowly pour melted 2% agarose to fill the entire dish and fully overed the embryo (Fig. 2a) (see Note 11). 3. After the agarose is completely solidified, use a razor or microtome blade to roughly trim the embryo close to the scanning plane. For the remainder of the protocol, we will use imaging the aortic arch as an example so the cuts are around the lower jaw and behind the forelimb (Fig. 2a). 4. Take the specimen out of the agarose and remove the forelimb. Wash it with PBS for 5 min. 5. Transfer the specimen into PBS with 15% sucrose and shake at room temperature until it sinks to the bottom. 6. Transfer the specimen into PBS with 30% sucrose and shake at room temperature until it sinks to the bottom (see Note 12). 7. Transfer the specimen into a mold and cover it completely with OCT (see Note 13). Leave the specimen submerged in OCT at room temperature for 15–30 min, then snap-freeze the specimen by immersing the mold into dry ice chilled ethanol or butanol. 8. Mount the specimen into a cryostat to continue trimming until the surface is close to the desired scanning plane. Check the sections under the microscope frequently during sectioning.

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Fig. 2 Whole-mount imaging of genetically labeled mouse embryo and 3-D reconstruction. (a) 4% PFA fixed embryo is embedded in 2% agarose in a 10 cm petri dish. The dashed line indicates the cutting position to image Wnt5a-CreER labeled cells in the aortic arch region. (b) Schematic diagram of mounting the specimen in the RIMS in the imaging chamber for confocal microscopy. (c) A set of aortic arch images taken by z-stack scanning. (d) 3-D reconstructed aortic arch (grey) from an E14.5 Wnt5a-CreER; Rosa26td-Tomato embryo. Wnt5a-CreER labeled cells are marked in light pink

9. Melt the OCT embedded specimen in PBS at room temperature then wash with PBS five times. 3.7.2 Whole Mount Phalloidin Staining and RIMS Clearing

1. Permeabilize the specimen in 1 PBS, 0.5% Tween 20 at 4  C overnight. 2. Dilute FITC-conjugated phalloidin 1:2000 in PBST. Stain the specimen with diluted phalloidin at 4  C for 24 h. 3. Wash the specimen with PBST at room temperature four times, 1–2 h each time with sharking. Then transfer the specimen into fresh PBST and wash on a shaker at 4  C overnight. 4. To clear the specimen, transfer it into a 2 mL Eppendorf tube with RIMS. Wrap the tube with foil and rotate it at room temperature for 24 h or until the specimen becomes transparent.

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1. To image with an upright microscope, it will be necessary to make an imaging chamber to mount the cleared specimen with the scanning surface facing up (Fig. 2b). To make the chamber, cut a 5 mm height ring from a 15 mL tube and glue it to a petri dish. Check that the seal is tight and there is no leakage. 2. Gently transfer the specimen to the imaging chamber filled with RIMS, with the cryostat trimmed surface facing up. Carefully place a coverslip on the top (Fig. 2b) without introducing any bubbles. 3. Place the mounted specimen on the stage of a confocal microscope. Determine the start and end position for the z-stack, set the step size to 2–4 μm (for 5–10 objectives). Adjust the laser power and PMT settings (see Note 14). 4. After scanning, the z-stack images (Fig. 2c) can be import into Amira or other imaging software for 3D reconstruction. 5. If using Amira, split the channels if the sample is imaged for two or more fluorophores. 6. Perform “z-drop correction” to normalize the fluorescent intensity on all the images within the z-stack. 7. Run automatic alignment mode.

alignment

with

the

least-squares

8. Use the “segmentation” tool to label and define the area of interest. For instance, in our images in the green channel (FITC-phalloidin), the blood vessel lumen can be labeled and defined as the aorta/aortic arch and pulmonary trunk. 9. Create another material in the segmentation tool. Use the “grow” function to mark the vessel wall of the labelled aorta and pulmonary trunk. 10. Go to the red channel and use the “threshold tool” to identify Wnt5a-Cre lineage cells (labeled by td-Tomato) on the blood vessel wall. 11. Generate the surface from segmented blood vessel in the green channel and Cre lineage in the red channel to create the tetra grid and 3D view of the z-stack (Fig. 2d).

4

Notes 1. We have noticed that the more current versions of Ensembl browser display few choices of BAC clones, whereas the archives from 2017 have more BAC clone information. 2. RP23 (RPCI-23) BAC library is constructed from genomic DNA of female C57BL/6 J mice using pBACe3.6 as cloning vector (https://bacpacresources.org/library.php?id¼11),

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whereas RP24 (RPCI-24) BAC library is constructed from genomic DNA of male C57BL/6 J mice using pTARBAC1 as cloning vector (https://bacpacresources.org/library.php? id¼12). 3. Do not overdry the pellet, otherwise the DNA may be difficult to dissolve. 4. Because of their large size, BAC DNA will be more easy to break during pipetting. Therefore cautions should be taken throughout the protocol to avoid repeated pipetting of BAC DNA solution. 5. To be able to visualize all the bands, usually ~0.5–1 μg BAC DNA and 1 μL of enzyme will be needed for the restriction reaction. 6. The predicted restriction fragments can often be determined by analyzing the sequence of the BAC clones if their genome coordinates are given in Ensembl during the initial search in Subheading 3.1. For instance, BAC clone RP24-85 L15 that contain the Rosa26 gene, has genome coordinates of 112,952,746–113,158,583. To download its sequence, go to http://useast.ensembl.org/index.html, click on “Mouse,” enter “6: 112952746–113,158,583” in the “Search all categories,” then click “Go” (6 stands for moue chromosome 6 on which Rosa26 is located). On the next page, click on “Export data” on the left hand to download the 205,838 bp sequence. If NotI is used for restriction digest, note that there is usually 1–2 NotI sites on the cloning vector besides the genomic sequence in the BAC DNA. 7. To further confirm that the BAC clone contains the gene(s) of interest, PCR can also be performed. The PCR reaction covering the region to be modified should also be sequenced and compare to the genome sequence in the database. This is critical for designing the primers that will serve as the homology arms for recombineering, as even minor difference between the designed homology arm and the actual sequence within the BAC will reduce the recombineering efficiency. 8. In the recombineering competent strains, transcription of Gam, Beta, and Exo is controlled by a temperature sensitive λ repressor in order to minimize their toxic effect. The repressor is active at low temperature (~32  C), but can be rapidly inactivate with a 15 min incubation at 42  C to provide a pulse of Gam, Beta, and Exo expression. Therefore, when growing these cells either in media or on plates, the temperature should always be at or lower than 32  C. 9. There are two general strategies to make the desired modifications. The first is a two-step strategy using a dual-selectable marker. The marker is first selected for integration into the

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location to be modified because it provides resistance to an antibiotic, and is subsequently replaced (selected against) with the desired modification. This strategy is more difficult and time-consuming, but can create precise, seamless modifications such as point mutations or in-frame insertions/deletions with minimal perturbation to the flanking genomic sequence. The most commonly used dual selectable markers include SacBNeo [40, 41] and rpsL-Neo [42, 43], which confer resistance to kanamycin when inserted into the BAC, and sensitivity to sucrose (~7%) and streptomycin (60 μg/mL), respectively, in the subsequent step when selecting for their replacement by the desired modification. The second strategy is to directly link the desired modification, such as Cre or EGFP, with a selection marker, such as Neo. PCR or cloning is then used to place homology arms around them. This strategy is considerably easier, but often leaves behind exogenous sequence around the desired modification. PL451 and PL452 [38] are constructs that contain Neo flanked by FRT and LoxP sites, respectively. They can be used to link a desired modification, such as Cre or EGFP, with Neo. Once the Cre-Frt-Neo-Frt or Cre-LoxP-Neo-LoxP cassette has been integrated into a desired location in the BAC through recombineering, Cre or Flippase expression can be induced by arabinose (0.1% for 1 h) in EL350 or EL250 cells (Table 1) to remove Neo [33, 38]. This strategy can also be used to easily insert LoxP sites in introns to engineer conditional BAC transgenes [16, 38]. Similar to the recombineering cells lines, PL451 and 452 constructs can be requested from NCI through the following link: https://frederick.cancer.gov/Sci ence/BrbRepository/#/productDataSheets/Plasmid. 10. The handbook of Qiagen Large Construct Kit recommends culturing the bacterial for ~16 h. The modified BAC in the recombineering strains, however, is culture at 32  C instead of standard 37  C, and therefore a longer culturing period is needed to achieve the same level of cell growth. 11. Regular agarose can be used but make sure the temperature is lower than 30  C before pouring. Shaking the agarose during cooling to keep it from solidifying. 12. Can also leave the specimen in 30% sucrose at 4  C overnight. 13. It is important to remove any residual PBS from the surface of the specimen with Kimwipes. Otherwise, the PBS on the surface will form a sheet of ice during the snap-freezing processes, causing the specimen to detach from OCT during sectioning. 14. If the specimen to be scanned is too thick or specimen clearing is not optimal, it may be difficult to image the entire sample with one laser power setting. If this is the case, the imaging can

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be divided into several z-stacks with increasing laser power settings for deeper tissues. Images from different z-stacks can be stitched together during 3D reconstruction with various software.

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Chapter 14 Two-Photon Cell and Tissue Level Laser Ablation Methods to Study Morphogenetic Biomechanics Abigail R. Marshall, Eirini Maniou, Dale Moulding, Nicholas D. E. Greene, Andrew J. Copp, and Gabriel L. Galea Abstract Laser ablation is routinely performed to infer mechanical tension in cells and tissues. Here we describe our method of two-photon laser ablation at the cellular and tissue level in mouse embryos. The primary outcome of these experiments is initial retraction following ablation, which correlates with, and so can be taken as a measure of, the tensile stress that structure was under before ablation. Several experimental variables can affect interpretation of ablation tests. Pre-test factors include differences in physical properties such as viscoelasticity between experimental conditions. Factors relevant during the test include viability of the cells at the point of ablation, image acquisition rate and the potential for overzealous ablations to cause air bubbles through heat dissipation. Post-test factors include intensity-biased image registration that can artificially produce apparent directionality. Applied to the closing portion of the mouse spinal neural tube, these methods have demonstrated long-range biomechanical coupling of the embryonic structure and have identified highly contractile cell populations involved in its closure process. Key words Laser ablation, Two photon, Biomechanics, Mouse, Neural tube

1

Introduction Primary neurulation is a fundamentally biomechanical event during which the relatively flat neural plate folds to form a closed neural tube [1, 2]. The mechanical forces required for this change in shape are generated by embryonic cells, primarily the neuroepithelial cells themselves [3, 4] but also with biomechanical influences from adjacent tissues. Many cellular processes can generate morphogenetic mechanical forces, but the most extensively studied are those mechanisms which require cells to change their own shape through actomyosin contractility. The forces they generate are withstood as

Abigail R. Marshall and Eirini Maniou contributed equally. Chenbei Chang and Jianbo Wang (eds.), Cell Polarity Signaling: Methods and Protocols, Methods in Molecular Biology, vol. 2438, https://doi.org/10.1007/978-1-0716-2035-9_14, © Springer Science+Business Media, LLC, part of Springer Nature 2022

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mechanical stress (force/area) and deform tissues, causing mechanical strain (change in dimension, for example, the percentage increase in length) [2]. Numerous techniques have been developed to quantify forces and material properties of cells and tissues [2, 5]. The mechanobiological parameter which is most commonly measured experimentally is mechanical tension. At equilibrium, cells withstanding tension are resisting being stretched; that is, they are constricting against the source of tension. This is equivalent to a rubber band stretched between two fingers (see ref. [2] for a more detailed physical model). If, hypothetically, you cut off one of the fingers stretching the rubber band, the band will constrict to a shorter length. In contrast, if you cut the band itself, the two ends will recoil away from each other. The initial velocity at which the cut ends recoil away from each other is proportional to the tension they withstood. The same principles can be applied to embryonic tissues [6]. If a cell constricts, it will stretch the cells it is attached to. In mammalian epithelia, these stretching forces are transmitted by cell-cell junctions including adherens and tight junctions [7]. If the constricted cell is cut, the stretched cell will retract to a shorter length, and if a stretched cell is bisected its two ends will recoil away from each other. Many insightful studies have been reported in which cuts were made in embryonic tissues using fine needles [8–10]. Over the past decade, high powered laser pulses have been increasingly used to achieve very precise cuts with subcellular accuracy. Laser ablation is a destructive assay which can only meaningfully be applied to healthy, living cells or tissues as fixation dissipates tissue tension [4]. The physics of plasma-mediated laser ablation of biological tissues are explained elsewhere [11, 12]. Many methods have been described to perform highly accurate ablations using pulsed UV lasers in biological contexts [13–15]. Two-photon lasers can also be used to perform laser ablations [3, 16], providing the dual benefit of being combined with confocal or two-photon imaging of complex 3D structures. Some two-photon laser ablation protocols are limited by the slower imaging speed of laser-scanning confocal microscopy compared with wide-field imaging, and heat dissipation away from the ablation site as evidenced by formation of air bubbles from boiling [12]. These air bubbles can be inconsequential or indeed desirable, for example if attempting to thermally destroy a cell population [17]. However, they confound experiments in which the primary endpoint is inference of tension by measuring initial retraction velocity. Factors which influence retraction velocity include cell–cell and cell–substrate adhesiveness, viscoelasticity, and rheological properties [5, 18, 19]. Nonetheless, when comparing equivalent cell types, ablation remains the most common method to infer tension in biological systems. We routinely employ two-photon laser

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ablation to identify tissue load-bearing structures and compare cell border mechanical tensions in mouse embryos. Applying these techniques to study closure of the mammalian neural tube, we have demonstrated that abnormal tension at the neural tube fusion points precedes failure of closure in many [3, 4, 20–22], but not all [22, 23] models of spina bifida. Here, we provide detailed descriptions of the methods involved in generating these data. We focus on methods relevant to the study of mouse primary neurulation, specifically closure of the open neural tube in the future spinal region, called the posterior neuropore (PNP).

2

Materials

2.1 Dissection Materials

1. Microsurgical needles, purchased as swaged needles on 11–0 Mersilene (TG140-6; Ethicon) and 10–0 Prolene (BV75-3; Ethicon). Cut away the suture material. 2. Agarose: regular melting point, molecular biology grade. 3. Stainless steel watchmaker forceps (#5). 4. 5 mL sterile bijou containers with screw caps. 5. 50 mL sterile falcon tubes. 6. DMEM buffered with HEPES. 7. Fetal bovine serum (FBS), heat-inactivated for 30 min at 56
C. 8. Dissection medium made up of 10% FBS v/v in DMEM. 9. CellMask Deep Red plasma membrane (C10046 Invitrogen). 10. 60 mm cell culture dishes. 11. 1.5 mL sterile Eppendorf tubes. 12. 3 mL plastic Pasteur pipettes. Cut the end to avoid embryo damage and use for transferring the embryos. 13. Dissecting stereoscope. 14. 10% CO2–20% O2 in nitrogen compressed gas mixture and regulator to allow sample gassing. 15. Water bath or hot block preset to 37
C.

2.2 Confocal Microscope Specifications

Upright confocal multiphoton microscope (in our case Zeiss LSM 880) equipped with: 1. SpectraPhysics Mai Tai eHP DeepSee multiphoton laser 690–1040 nm. 2. Laser lines for excitation including 633 nm. 3. 10/NA0.5 W-Plan Apochromat Water dipping objective. WD 3.7 mm.

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4. 20/NA1.0 Plan Apochromat Water dipping objective with correction collar. WD 2.4 mm. 5. Temperature controlled chamber at 37
C.

3

Methods

3.1 Before Starting the Experiment

1. The day before, prepare agarose plates on which to position the embryos. Microwave 4% w/v agarose powder in PBS, swirling intermittently to prevent excessive bubbling (see Note 1). Continue until the agarose is fully dissolved (approximately 5 min including swirling to dissolve a 50 mL solution in an 800 W microwave). Pour ~4 mL solution into a 60 mm dish and allow to cool. Using the blunt end of a p20 pipette tip, cut a cylindrical hole in the middle of the dish. Store the dish in PBS to avoid desiccation. 2. Prior to collecting the embryos, switch on the confocal microscope and the MaiTai multiphoton laser. Set the temperature of the microscope chamber to 37
C. Allow at least an hour for the chamber temperature to equilibrate before use. Set the water bath/hot block to 37
C. This needs to be close to the confocal microscope and is needed to keep the embryos alive before ablations. 3. For cell border ablations, dilute CellMask 1:500 in DMEM without FBS and transfer the solution to the water bath (see Note 2). 500 μL is enough for ~10 embryos. 4. Prepare all materials for dissection prior to embryo collection and place them next to the stereomicroscope. Prepare 50 mL of dissection medium and prewarm the medium in a 37
C water bath. Transfer 2 mL of this dissection medium to a 5 mL bijou container.

3.2 Embryo Collection

1. After sacrificing the pregnant female (see Notes 3, 4), collect the uterus in the 5 mL bijou tube containing prewarmed dissection medium. 2. Transfer the uterus and medium to a 60 mm dish and under a stereomicroscope, remove the muscular uterine lining exposing the decidua around each implantation. Then, carefully separate the implantations while keeping them intact. Detailed dissection procedures have been described previously [24]. 3. Place each implantation into an individual 1.5 mL Eppendorf tube containing fresh dissection medium. 4. Equilibrate each tube with 5% CO2, 20% O2, 75% N2 by flowing the gas mixture over the top of the medium for ~30 s. The gas should not be bubbled through the medium. 5. Place the Eppendorf tubes in the 37
C water bath/hot block next to the confocal microscope.

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1. Immediately before ablation, fill a new 60 mm dish (without agarose) with dissection medium. This will be used to dissect embryos out of their decidua. 2. Using a disposable Pasteur pipette with the end cut off to provide a suitable size aperture, retrieve and implantation from its 1.5 mL tube and transfer it into the dissection dish. 3. Under a stereomicroscope, dissect away the decidua, followed by the extraembryonic membranes: the mural trophoblast and attached Reichert’s membrane are removed, to produce embryos that are enclosed within an intact yolk sac, with underlying amnion, and with the ectoplacental trophoblasts attached (see Note 5). 4. Continue with the steps below, which are modified for each type of ablation. The key difference between them is that, for tissue level ablations, the entire embryo is kept intact and alive, whereas for the cell-level ablations the rostral half of the embryo including the beating heart is dissected off immediately before positioning for ablation.

3.3.1 Embryo Positioning for Cell Border Ablations

1. After removing the extraembryonic membranes, use a Pasteur pipette to transfer the embryo to the CellMask solution and stain for 5 min at 37
C. Aim to minimize the volume of dissection medium transferred with the embryo. 2. Fill an agarose plate with prewarmed dissection medium and transfer the stained embryo into it. 3. Separate the caudal from the rostral half of the embryo (Fig. 1a) and discard the latter or use it for genotyping. This is to remove the effect of the beating heart, which causes too much movement to accurately ablate single cell borders. 4. Position the caudal region by piercing a curved suture needle through the most ventral part of the tissue (see Note 6), ensuring the PNP is pointing upwards. As a guide, aim to insert the needle below the level of the somites (Fig. 1a, see Note 7), ensuring the neuroepithelium and dorsal surface ectoderm are not touched. 5. Fix both ends of the needle in the agarose (Fig. 1a). This is to ensure there are no sharp metal tips pointing towards the objective.

3.3.2 Embryo Positioning for Tissue-Level Ablations

1. Use an agarose plate with a central hole and fill it with prewarmed dissection medium. 2. After removing the extraembryonic membranes, transfer the embryo to the agarose plate and guide it into the hole (see Note 8).

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Fig. 1 Schematic to demonstrate positioning of embryos for single cell and tissue laser ablation. (a) For single cell ablations, the caudal region of the embryo is cut using forceps and positioned by piercing the ventral half of the tissue using a curved suture needle. The tissue is then held in place with both ends of the suture needle fixed in the agarose. (b) For tissue level ablation, whole embryos are positioned by creating a hole in the agarose and piercing the embryo through the body to the walls of the hole using a suture needle

3. Pin a suture needle dorsal to the heart and into the side of the hole. The embryo should be oriented so that the PNP is visible above the surface of the agarose (Fig. 1b). The heart beat should continue steadily throughout the experiment. 4. If the heart beat interferes with imaging, additional needles can be added to support and stabilize the caudal part of the embryo (see Note 9).

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3.4 Imaging and Ablation Settings

The imaging and ablation settings need to be optimized for the equipment used. Those described here achieve the desired objectives in our hands. Carefully transfer the agarose dish with the embryo pinned on it into the microscope incubation chamber and place it into an appropriate stage inset, ensuring it does not wobble. Find the part of the embryo to be ablated and continue with the steps below, which are modified for each type of ablation.

3.4.1 Imaging Settings for Cell Border Ablations

1. Image CellMask stained live embryo cell borders with a 20 dipping objective using the 633 nm wavelength, at maximum scan speed without averaging. Identify a cell border to ablate and rotate the sample so that border is along the Y axis, such that the laser travels along the X axis (faster). 2. Zoom in 4 (0.1  0.1 μm x/y pixel size, see Note 10) and adjust the Z-plane so the border signal is clear in the plane of view. Be careful when focusing to avoid contact between the objective and the embryo or needle. 3. Using the microscope (ZEN) software in continuous imaging mode, draw a Region Of Interest (ROI) as a 0.1 μm wide line, which does not overlay other cell borders. 4. To perform an ablation, adjust the MaiTai multiphoton laser to 710 nm, 80% laser power, 0.34 μs pixel dwell time for 20 iterations. In our hands, these settings reliably ablate superficial cell borders without producing an air bubble. 5. Image the field of view before and immediately after ablation. You can reimage repeatedly after ablation to generate kymographs of border movement over time. 6. Only one ablation is analyzed per embryo to avoid potential confounding effects of local reactions on nearby cells, although multiple ablations could potentially be performed in different regions.

3.4.2 Imaging Settings for Tissue-Level Ablations

1. For tissue ablations, PNPs are imaged using a 10 dipping objective and reflection mode imaging at 633 nm with a MBS T80/R20 beam splitter [4]. 2. Acquire a pre-ablation Z-stack of the PNP. For embryonic day 9.5 embryos, 150–200 slices of 1.4 μm Z-step are required to go through the whole tissue. With these settings, total acquisition time for each stack is