Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocol [2219, 2 ed.] 9781071609736, 9781071609743

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Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocol [2219, 2 ed.]
 9781071609736, 9781071609743

Table of contents :
Preface
Contents
Contributors
Chapter 1: Laboratory Culture and Mutagenesis of Amphioxus (Branchiostoma floridae)
1 Introduction
2 Materials
2.1 Materials for Aquaculture of Amphioxus
2.2 General Materials for Transgenesis
2.2.1 Materials for TALEN Construct Assembly
2.2.2 Materials for TALEN mRNA Synthesis and Egg Injection
2.2.3 Materials for Tol2 Mutagenesis
3 Methods
3.1 Method for Amphioxus Aquaculture (Figs. 2, 3, and 4)
3.1.1 Obtaining Adult Amphioxus and Establishing a Culture
3.1.2 Salinity
3.1.3 Parasites and Diseases
3.1.4 Laboratory Breeding Culture (see Note 1)
3.1.5 Feeding Adults and Algal Culture (Fig. 2)
3.1.6 Breeding (Fig. 3)
3.1.7 Raising Larvae of B. floridae (Fig. 4)
3.2 Generation of Amphioxus Mutants with TALEN Method (Figs. 5, 6, and 7)
3.2.1 TALEN Target Site Design
3.2.2 TALEN Construct Assembly (Figs. 5 and 6)
3.2.3 Before Golden Gate Assembly
3.2.4 Golden Gate Assembly: Day 1
3.2.5 Golden Gate Assembly: Day 2 (Fig. 6)
3.2.6 Golden Gate Assembly: Day 3
3.2.7 Golden Gate Assembly: Day 4
3.2.8 Golden Gate Assembly: Day 5
3.2.9 TALEN mRNA Synthesis, Microinjection and Analysis of Somatic Mutations (Figs. 6 and 7)
3.2.10 Detection of Mutations in Founder and F1 Amphioxus (Fig. 7)
3.2.11 Obtaining and Analyzing Homozygous Mutant Embryos (Fig. 7)
3.3 Generation of Amphioxus Transgenic Line with Tol2 Transposon System
3.3.1 Constructs
3.3.2 Preparation of the Injection Reagent and Embryonic Injection
3.3.3 Detection of Germline Transmission in F0 Founders
3.3.4 Identification of F1 Transgenic Amphioxus
4 Notes
References
Chapter 2: Handling and Manipulation of Gametes and Embryos of the Annelidan Worm Pseudopotamilla occelata
1 Introduction
2 Materials
2.1 Culture of Adults and Collection of Gametes
2.2 Induction of Oocyte Maturation and Fertilization
2.3 Microinjection
2.4 Ca2+ Imaging at Fertilization
3 Methods
3.1 Culture of Adults and Collection of Gametes
3.2 Induction of Oocyte Maturation and Fertilization
3.3 Microinjection
3.4 Ca2+ Measurements at Fertilization
4 Notes
References
Chapter 3: The Starfish Asterina pectinifera: Collection and Maintenance of Adults and Rearing and Metamorphosis of Larvae
1 Introduction
2 Materials
2.1 Collection and Selection of Starfish
2.2 Laboratory Aquaria
2.3 Seawater for Aquaria
2.4 Oocyte Maturation, Insemination, and Embryonic Development
2.5 Collection of Normal Embryos
2.6 Diatom Culture
2.7 Induction of Metamorphosis
3 Methods
3.1 Collection of Adult Starfish
3.2 Getting Collection Permits from Local Agencies
3.3 SCUBA Diving for Starfish
3.4 Selection of Starfish
3.5 Packing and Transport
3.6 Maintenance of Adult Starfish in Laboratory Aquaria
3.7 Managing Laboratory Populations of Starfish
3.8 Quality Monitoring of Aquarium Seawater
3.9 Seawater Temperature
3.10 Feeding Adults
3.11 Preparation of ASW for Embryonic and Larval Cultures
3.12 Oocyte Maturation, Fertilization, and Culture of Embryos
3.13 Collection of Normally Developing Embryos
3.14 Culture of Diatoms (Chaetoceros Sp.) as Food for Larvae
3.15 Rearing Larvae
3.16 Induction of Metamorphosis by Pebbles Associated with Adults
4 Notes
References
Chapter 4: Experimental Tools to Study Regeneration in the Sea Anemone Nematostella vectensis
1 Introduction
2 Materials
2.1 Animal Care
2.2 Induction of Regeneration
2.3 Fixation, Permeabilization, Coating, and Washing Buffer for Immunostaining and EdU/EU Staining
2.4 Membrane and Nuclei Staining
2.5 EdU (5-Ethynyl-2′-Deoxyuridine) and EU (5-Ethynyl-Uridine) Staining
2.6 TUNEL Staining
2.7 Immunostaining
3 Methods
3.1 Animal Care
3.2 Induction of Regeneration
3.3 In Vivo Analysis of Wound Healing and Pharynx Reformation
3.4 Tissue Morphology, Cellular Proliferation, Neotranscription, and Apoptosis Staining
3.4.1 Tissue Morphology
3.4.2 Cellular Proliferation (EdU) and Neotranscription (EU)
3.4.3 Apoptosis
3.4.4 Immunostaining
4 Notes
References
Chapter 5: Staining and Tracking Methods for Studying Sponge Cell Dynamics
1 Introduction
2 Materials
2.1 Biological Material
2.1.1 Amphimedon queenslandica Biological Samples
2.1.2 Lycopodina hypogea Biological Samples
2.1.3 Oscarella lobularis Biological Samples
2.2 Antibody-Based Staining on Whole Mount Biological Samples (See Note 2)
2.2.1 A. queenslandica
2.2.2 L. hypogea
2.2.3 O. lobularis
2.3 EDU Incorporation Assays
2.4 Live Cell Staining and Imaging Assays in O. lobularis and A. queenslandica
3 Methods
3.1 Obtaining Amphimedon queenslandica Juveniles
3.2 Stalling and Nutrition of Lycopodina hypogea Adults
3.3 Obtaining Oscarella lobularis Buds
3.4 Detection of Phosphohistone 3 (PH3) by Immunofluorescence (See Note 6)
3.4.1 A. queenslandica
3.4.2 L. Hypogea
3.4.3 O. lobularis
3.5 EdU Incorporation Assays
3.5.1 A. queenslandica
3.5.2 L. hypogea
3.5.3 O. lobularis
3.6 TUNEL Assays (See Note 7)
3.6.1 O. lobularis and L. hypogea
3.7 MAPLC3 Staining in L. hypogea (See Note 8)
3.8 Staining Choanocytes with the Lipidic Marker CM-DiI Dye
3.8.1 A. queenslandica
3.8.2 O. lobularis
3.9 Staining O. lobularis Choanocytes with Fluorescent Microspheres (See Note 9)
3.10 Staining O. lobularis Choanocytes with Fluorescent Lectins (See Note 9)
3.11 Live Imaging in White Light
3.11.1 A. queenslandica
4 Notes
References
Chapter 6: Microscopy Studies of Placozoans
1 Introduction
2 Materials
2.1 Culturing and Maintenance
2.2 High Pressure Freezing for Thin Section EM
2.3 Freeze Substitution for Thin Section EM
2.4 Chemical Fixation for SEM
2.5 Freezing, Freeze Substitution, and Immunolabeling for Light Microscopy
3 Methods
3.1 Collection and Maintenance
3.2 Algal Cultures
3.3 Trichoplax Cultures
3.4 High Pressure Freezing for Thin Section EM
3.5 Freeze Substitution for Thin Section EM
3.6 Freezing, Freeze-Substitution and Immunolabeling for Light Microscopy
3.7 Fixation for SEM
3.8 Freeze Fracturing Tissues for SEM
3.9 Critical Point Drying and SEM
4 Notes
References
Chapter 7: Identification of SH2 Domain-Mediated Protein Interactions that Operate at Fertilization in the Sea Star Patiria mi...
1 Introduction
2 Materials
2.1 Starfish Egg and Sperm Preparation
2.2 Mutagenesis
2.3 GST Fusion Protein Expression
2.4 Affinity Interactions, Gel Electrophoresis, and Western Blotting
3 Methods
3.1 Obtaining Oocytes
3.2 Obtaining Sperm
3.3 Oocyte Maturation and Fertilization
3.4 Mutagenesis of GST-Wild-Type SH2 Domain Fusion Protein Constructs
3.5 Fusion Protein Expression and Purification
3.6 Affinity Interactions
3.7 SDS-PAGE and Silver Staining
3.8 Western Blotting
4 Notes
References
Chapter 8: Marine Nemertean Worms for Immunoblotting Studies of Oocyte Aging
1 Introduction
2 Materials
2.1 General Solutions
2.2 SDS-Polyacrylamide Gel Components
2.3 Transfer Components
2.4 Supplies to Take to the Darkroom When Developing Film
3 Methods
3.1 Maintaining Adult Specimens
3.2 Obtaining Gametes and Fertilized Embryos
3.3 General Cell Biological Applications
3.4 Incubating Oocytes During Aging Experiments
3.5 Sample Processing for Immunoblotting Analyses
3.6 Gels
3.7 Electrophoresis
3.8 Transfer
3.9 Antibody Probing
3.10 Film Developing
3.11 Reprobing Stripped Membranes
3.12 Quantification of Blot Intensities
4 Notes
References
Chapter 9: Recovery of Sea Star Egg Cell Surface Proteins Released at Fertilization
1 Introduction
2 Materials
2.1 Obtaining and Culturing Oocytes/Testes
2.2 Biotinylation
2.3 Streptavidin-Biotin Affinity Interaction
2.4 SDS-PAGE Gel and Western Blotting
2.5 Miscellaneous
3 Methods
3.1 Oocyte/Sperm Removal and Preparation
3.2 Biotinylation of Eggs
3.3 Fertilization
3.4 Affinity Interaction Using Streptavidin Mag Sepharose Beads (See Note 11)
3.5 SDS-PAGE Gel and Western Blotting
4 Notes
References
Chapter 10: Quantifying Cell Proliferation During Regeneration of Aquatic Worms
1 Introduction
2 Materials
2.1 Buffers and Relaxation Solutions
2.2 Immunohistochemistry and Nuclear Counterstains
2.3 Thymidine Analogues
2.4 Peroxidase Development
2.5 Click Chemistry
3 Methods
3.1 Worm Relaxation and Fixation
3.2 PH3/PCNA Detection
3.3 BrdU Proliferation Assay
3.4 EdU Proliferation Assay
4 Notes
References
Chapter 11: In Situ Hybridization Techniques in the Homoscleromorph Sponge Oscarella lobularis
1 Introduction
2 Materials
2.1 Biological Material
2.2 Probe Synthesis
2.3 Sample Fixation
2.4 Paraffin Fixation and Cuts
2.5 In Situ Hybridization
2.6 Embedding and Sectioning Tissues
3 Methods (see Notes 2 and 3)
3.1 Probe Synthesis
3.2 Fixation (See Note 4)
3.2.1 To Fix Adults
3.2.2 To Fix Buds and Larvae
3.3 Paraffin Embedding and Sectioning (for SISH) (See Note 5)
3.4 In Situ Hybridization: Section ISH (SISH)
3.5 In Situ Hybridization: Whole mount ISH (WISH) on buds and larvae (See Note 8)
3.6 In Situ Hybridization: Whole Mount ISH on Adult Tissue (See Note 8)
3.7 In Situ Hybridization: Whole Mount ISH Using InsituPro VSi (INTAVIS)
3.8 Embedding and Sectioning for Light Microscopy after WMISH (See Note 9)
3.8.1 Epoxy Embedding
3.8.2 L.R. White
4 Notes
References
Chapter 12: Methodology for Whole Mount and Fluorescent RNA In Situ Hybridization in Echinoderms: Single, Double, and Beyond
1 Introduction
2 Materials
3 Methods
3.1 Fixation Protocol
3.1.1 Harvest of Embryos and Larvae
3.1.2 Fixation
3.2 Riboprobe Synthesis
3.3 Sea Urchin WMISH Protocols
3.4 Sea Urchin FISH Protocol
3.5 Second Probe Detection for Double-FISH
3.6 P. miniata WMISH and FISH Protocols
3.7 Positive and Negative Controls
3.8 Combination of Immunolabeling and In Situ Hybridization
3.9 Conclusions
4 Notes
References
Chapter 13: Gene Editing in the Ascidian Phallusia mammillata and Tail Nerve Cord Formation
1 Introduction
2 Materials
2.1 Microinjection of mRNA or CRISPR-Cas9 Protein Mixtures
2.2 CRISPR-Cas9 Protein or mRNA Microinjection
3 Methods
3.1 Microinjection
3.2 Preparing and Injecting CRISPR-Cas9
3.3 Imaging the Nerve Cord
3.4 Analyzing the Nerve Cord Imaging Data Using Freeware: Fiji, Ilastik, and ICY
4 Notes
References
Chapter 14: Transcriptomic Analysis in the Sea Anemone Nematostella vectensis
1 Introduction
2 Materials
2.1 RNA Isolation
2.2 Quantification of Transcriptomic Data
3 Methods
3.1 RNA Isolation and Quality Control
3.2 Quantification of Transcriptomic Data
4 Notes
References
Chapter 15: RNA Interference on Regenerating Holothurian Gut Tissues
1 Introduction
2 Materials
2.1 Tissue Selection
2.2 Gut Rudiments Culture
2.3 Gut Rudiment Electroporation
2.4 Determining Electroporation Efficiency Using Histological Approaches
2.5 DsiRNAs
2.6 Determination of RNA Abundance Using qPCR
3 Methods
3.1 Sea Cucumber Collection and Evisceration
3.2 Sea Cucumber Disinfection and Explant Preparation
3.3 Optimization of the Electroporation Protocol
3.4 Histological Studies for Electroporation Efficiency
3.5 siRNA Design
3.6 Electroporation of Gut Rudiments with DsiRNAs
3.7 RNA Extraction
3.8 cDNA Synthesis
3.9 qPCR to Determine the Efficiency of the Interference
4 Notes
References
Chapter 16: ATAC-Seq for Assaying Chromatin Accessibility Protocol Using Echinoderm Embryos
1 Introduction
2 Materials
2.1 Animal Origin
2.2 Equipment
2.2.1 In Vitro Fertilization and Embryo Collection
2.2.2 ATAC-Seq Library Preparation
2.3 Reagents
2.4 Downloading Genome and Annotation
3 Methods
3.1 In Vitro Fertilization
3.2 Collection of Embryos
3.3 Cell Lysis
3.4 Tagmentation
3.5 PCR Amplification
3.6 Concentration Check and Electrophoresis Gel
3.7 Data Analysis
3.8 Anticipated Results
4 Notes
References
Chapter 17: Usage of the Sea Urchin Hemicentrotus pulcherrimus Database, HpBase
1 Introduction
2 Materials
3 Methods
3.1 Gene Search Page
3.2 Homology Search Page
3.3 Genome Viewer Page (Genome Explorer)
3.3.1 How to Display a Specific Genome Position
3.3.2 How to Search for a Gene
3.3.3 How to Retrieve a Sequence
3.3.4 How to Change Color and Thickness of Boxplot
3.3.5 How to Obtain the Image of Genome Map
3.4 Data Download Page
3.5 Protocol Page
3.5.1 Protocol for Immunohistochemistry
3.5.2 Protocol for In Situ Hybridization
3.5.3 Reagents for Microinjection
3.5.4 Protocol for qPCR
3.5.5 Protocol for Luciferase Assay (TOPFlash Assay)
4 Notes
References
Chapter 18: Functional Studies of Trichoplax adhaerens Voltage-Gated Calcium Channel Activity
1 Introduction
2 Materials
2.1 Extraction of RNA from Trichoplax adhaerens
2.2 Cloning of the Trichoplax T-Type Voltage-Gated Calcium Channel (TCaV3) cDNA into the pIRES2-EGFP Vector
2.3 Transfection of TCaV3-pIRES-EGFP Vectors into HEK-293T Cells
2.4 Whole-Cell Patch Clamp Recording of TCaV3-pIRES2-EGFP Transfected HEK-293T Cells
3 Methods
3.1 Extraction of RNA from Trichoplax adhaerens
3.2 Cloning of the Trichoplax T-Type Voltage-Gated Calcium Channel (TCaV3) cDNA into the pIRES2-EGFP Vector
3.3 Transfection of the pTCaV3-IRES2-EGFP Vector into HEK-293 T Cells
3.4 Whole-Cell Patch Voltage Clamp Recording of pTCaV3-IRES2-EGFP Transfected HEK-293T Cells
4 Notes
References
Chapter 19: A Bioinformatics Tutorial for Comparative Development Genomics in Diverse Meiofauna
1 Introduction
2 Materials
2.1 Computers
2.2 Required Software
2.3 Database
3 Methods
3.1 Genome Assembly and Assessment Tutorial Overview
3.2 Examining the Raw Data
3.3 Adapter and Quality Trimming
3.4 Genome Assembly
3.5 Genome Assessment
3.5.1 Part 2: Comparative Genomics
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2219

David J. Carroll Stephen A. Stricker Editors

Developmental Biology of the Sea Urchin and Other Marine Invertebrates Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Developmental Biology of the Sea Urchin and Other Marine Invertebrates Methods and Protocols Second Edition

Edited by

David J. Carroll Department of Biochemistry and Molecular Genetics, Midwestern University, Glendale, AZ, USA

Stephen A. Stricker Department of Biology, University of New Mexico, Albuquerque, NM, USA

Editors David J. Carroll Department of Biochemistry and Molecular Genetics Midwestern University Glendale, AZ, USA

Stephen A. Stricker Department of Biology University of New Mexico Albuquerque, NM, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0973-6 ISBN 978-1-0716-0974-3 (eBook) https://doi.org/10.1007/978-1-0716-0974-3 © Springer Science+Business Media, LLC, part of Springer Nature 2014, 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Marine invertebrates have contributed greatly to our understanding of many biological processes—from ecological and environmental issues to cell, developmental, and molecular biological problems. There are many reasons for this, including the fact that many of these organisms are easily accessible and have long stimulated the curiosity of humankind. You can probably remember back to a time in your childhood when you were fascinated with some interesting creature—a sea urchin at the beach or an earthworm in your backyard, for example. Even Charles Darwin was captivated by invertebrates, publishing a monograph on barnacles [1] before pondering the immense diversity of marine organisms he found during his voyage on the HMS Beagle, no doubt contributing to his ideas on natural selection [2]. In some cases, the relative simplicity of marine invertebrates has allowed breakthroughs that would be more difficult to obtain in research involving complicated higher vertebrates. The discovery of cyclin is a prime example of this. Rosenthal, Hunt, and Ruderman noted the change in synthesis of proteins in the surf clam after fertilization, precisely because these eggs are easily accessible, can be fertilized in vitro, and divide rapidly [3]. The sea urchin provided an even better model for studying these early cleavage stages and led to the naming of these proteins as the cyclins [4]. Also contributing to the usefulness of marine invertebrates is the fact that a wide variety of natural products are produced by marine invertebrates and have been exploited for applications related to human health, including compounds with anticancer, antimicrobial, and other therapeutic properties [5]. In this volume, a variety of marine invertebrate model systems are described along with novel experimental protocols for taking advantage of their unique properties. The techniques range from culturing the organisms to modifying their DNA. We have much to learn about the natural world, and these organisms will play a large role in helping us do so. In Chapter 1 on the “Laboratory Culture and Mutagenesis of Amphioxus (Branchiostoma floridae),” Holland and Li describe techniques for continuous breeding of the cephalochordate Branchiostoma floridae in the laboratory and methods for mutagenesis using TALEN and Tol2. Amphioxus occupies a special place in the hierarchy of organisms, being a sister group to vertebrates and tunicates. Studies on these organisms could provide novel insights into how vertebrates evolved. In Chapter 2 on the “Handling and Manipulation of Gametes and Embryos of the Annelidan Worm Pseudopotamilla occelata,” Deguchi and Nakano provide detailed methods for utilizing Pseudopotamilla occelata as a model system for oocyte maturation, signaling at fertilization and early development. This polychaete worm is widely distributed in the northern Pacific coast making it likely to be universally useful. They have developed methods for the care and experimental manipulation of gametes and embryos, including methods for microinjection and calcium imaging. Continuing with the theme of providing methods for scientists to adopt these useful animals as novel model systems is Chapter 3 on “The Starfish Asterina pectinifera: Collection and Maintenance of Adults and Rearing and Metamorphosis of Larvae” by Murabe et al. This chapter takes us from collecting these starfish during their breeding period, to maintaining gravid adults, to inducing larvae to undergo metamorphosis in the lab. Not only will these methods allow for analysis of development all the way through the starfish life

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cycle, it could also open the possibility of using this animal for studies of genetics, which has been largely missing from the Phylum Echinodermata. In Chapter 4, “Experimental Tools to Study Regeneration in the Sea Anemone Nematostella vectensis” by Amiel and Ro¨ttinger addresses the phenomenon of animal regeneration using the sea anemone Nematostella vectensis as a whole-body regeneration model. Amazingly, regeneration in this sea anemone occurs within 7 days! The chapter describes methods of inducing regeneration in this organism, along with the accompanying methods of fixation and staining to allow detailed morphological, cellular, and molecular studies of the process. This chapter should be enjoyed along with Chapter 14, which details methods for transcriptome analysis using the same organism. In Chapter 5, “Staining and Tracking Methods for Studying Sponge Cell Dynamics,” Borchiellini et al. present protocols for staining cells of three phylogenetically different sponge species to detect dynamic states of cell death, cell proliferation, and cell migration. They apply these methods to study different cell behaviors and fates in these sponges during development. Since sponges occupy such a strategic phylogenetic position in the tree of life, this comparative work can provide fundamental knowledge about cell behavior and physiology during these dynamic processes. Chapter 6, “Microscopy Studies of Placozoans” by Smith et al., describes advanced techniques for light and electron microscopic studies of Trichoplax adhaerens and is a good companion for Chapter 18 on genomics in Placozoans. This organism is a fascinating animal with very simple cellular organization and morphology. These microscopy techniques have increased our knowledge regarding the diversity and function of different cell types in placozoans. Understanding this phylogenetic group promises to provide insight into the evolution, development, differentiation, and physiology of cells and tissues. In Chapter 7, “Identification of SH2 Domain-Mediated Protein Interactions That Operate at Fertilization in the Sea Star Patiria miniata,” Bates et al. present methods for the isolation of tyrosine phosphorylated proteins that interact with a specific signaling molecule in the sea star egg during fertilization. The signaling mechanisms controlling internal calcium release at fertilization in animals are still largely unknown. This chapter begins with methods for preparing the protein samples from the eggs and describes a method for producing fusion proteins for affinity interactions with these egg lysates. Methods for mutagenesis of the fusion protein and resolution of the bound proteins by gel electrophoresis and western blotting are also described. Chapter 8, “Marine Nemertean Worms for Immunoblotting Studies of Oocyte Aging” by Stricker, outlines detailed methods for immunoblotting analysis of oocyte protein samples using phospho-specific antibodies. The nemertean worm is an established model system for understanding oocyte maturation and fertilization. In this chapter, the use of this organism as a model system for understanding the roles played by protein kinases as oocytes age and lose their ability to undergo normal fertilization is described. Methods for obtaining adult worms and for handling their gametes in experiments assessing oocyte aging are also included. In Chapter 9, “Recovery of Sea Star Egg Cell Surface Proteins Released at Fertilization” by Wiseman et al., we return to the sea star Patiria miniata with a method for visualizing egg cell surface proteins that are released during fertilization. There is a lot of interest in understanding the proteins on the surface of gametes and how they may function during fertilization. However, only a few have been described in any model system. This method describes the collection and analysis of proteins (or peptides) that are either naturally released by the egg or cleaved by sperm proteases. These released proteins can be

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concentrated by an affinity interaction and subsequently analyzed by western blotting and mass spectrometry to determine the identity of egg membrane proteins. Chapter 10, “Quantifying Cell Proliferation During Regeneration of Aquatic Worms” ¨ zpolat, presents methods for identifying and quantifying cell proliferation by Zattara and O by several methods, including very clever “click” chemistry. These methods are used to address the role of cell proliferation in animal regeneration, particularly for visualization in the whole worm. While they have developed these methods for application in whole mounts of small marine and freshwater worms, they could also be applied to larger samples. Chapter 11, “In situ Hybridization Techniques in the Homoscleromorph Sponge Oscarella lobularis” by Fierro-Constain et al., addresses methods for understanding gene expression over space and time at different stages of development in the sponge Oscarella lobularis. The methods are applied to both whole mount individuals and paraffin sections. The Porifera are widely considered as the sister group to all other metazoans. Therefore, studies on this phylum are useful for understanding the early evolution of important morphological or organismal features. In Chapter 12, “Methodology for Whole Mount and Fluorescent RNA In Situ Hybridization in Echinoderms: Single, Double, and Beyond,” Perillo et al. detail methods for localization of RNA by in situ hybridization in echinoderms. Unique to this chapter is the use of co-labeling techniques to identify multiple RNAs in one experiment. The authors also include an extensive “troubleshooting” section that should prove invaluable to investigators adopting these techniques into their labs. In Chapter 13, “Gene Editing in the Ascidian Phallusia mammillata and Tail Nerve Cord Formation” by McDougall et al., the powerful CRISPR-Cas9 system is applied to the ascidian Phallusia mammillata for the study of nerve cord development. The CRISPR-Cas9 system is employed to knock-out specific genes of interest and is applied along with imaging techniques for visualizing the nerve cord. Ascidians are a valuable model system for combining cell and molecular biology methods for understanding cell specification and morphogenesis during development because of their relative simplicity and rapid development. Chapter 14, “Transcriptomic Analysis in the Sea Anemone Nematostella vectensis” by Warner and Ro¨ttinger, is a companion chapter for Chapter 4. In this chapter, recent procedures for generating high-quality RNA for next generation sequencing are presented along with methods for quantification of this data. The sea anemone Nematostella vectensis is being developed as a model system for studying embryonic development and tissue regeneration at the multiple experimental levels. This organism shares similarities with mammalian genomes and has long been useful for evolutionary developmental studies. In Chapter 15, “RNA Interference on Regenerating Holothurian Gut Tissues,” AliceaDelgado et al. describe a useful transfection protocol for the introduction of Dicer-substrate interference RNAs into tissue explants for the study of gut regeneration. This adds a muchneeded protocol to the toolkit of echinoderm biologists and expands the use of this technology past embryos and into adult animals. Chapter 16, “ATAC-Seq for Assaying Chromatin Accessibility Protocol Using Echinoderm Embryos” by Magri et al., describes a novel method to determine chromatin accessibility called Assay for Transposase-Accessible Chromatin with high-throughput sequencing (ATAC-seq). This allows for identification of putative cis-regulatory elements for understanding the regulation of gene expression. The method has been specifically adapted for use in echinoderms and will be extremely useful for understanding the regulation of genes that have complicated temporal and morphological expression patterns during development.

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In Chapter 17, “Usage of the Sea Urchin Hemicentrotus pulcherrimus Database, HpBase” by Kinjo et al., an overview of the very useful HpBase (http://cell-innovation. nig.ac.jp/Hpul/) is provided along with a description of some of the useful analysis tools included. HpBase provides both genome and transcriptome resources for the sea urchin Hemicentrotus pulcherrimus. In addition to these resources, HpBase also presents protocols for biological experiments that have been assembled using H. pulcherrimus. This resource should be valuable for investigators in a wide range of fields. Chapter 18, “Functional Genomic Studies of Trichoplax adhaerens Neuronal Activity” by Gauberg et al., details methods for studying cellular and molecular activities in the placozoan Trichoplax adhaerens. Trichoplax has been generating a lot of interest recently because of its position as a basal member of the animal kingdom and its array of interesting behavioral responses to environmental stimuli. Great progress has been made on the development of cellular and molecular methods for studying this behavior. In Chapter 19, “A Bioinformatics Tutorial for Comparative Development Genomics in Diverse Meiofauna,” Sevigny et al. provide a detailed bioinformatics tutorial, with a specific focus on genome assembly, genome comparison, and the characterization of Hox clusters in meiofaunal species. These methods will be broadly applicable to many different miniaturized benthic invertebrates which are not yet well represented in the typical databases, but which will be very useful for addressing unique biological and evolutionary questions. Glendale, AZ Albuquerque, NM

David J. Carroll Stephen A. Stricker

References 1. Darwin C (1852–1854) A monograph on the sub-class Cirripedia, with figures of all the species, vol 1, 2. The Ray Society, London 2. Darwin C (1859) On the origin of species by means of natural selection, or preservation of favoured races in the struggle for life. John Murray, London 3. Rosenthal ET, Hunt T, Ruderman JV (1980) Selective translation of mRNA controls the pattern of protein synthesis during early development of the surf clam, Spisula solidissima. Cell 20:487–494 4. Evans T, Rosenthal ET, Youngblom J, Distel D, Hunt T (1983) Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33:389–396. 5. Blunt JW, Copp BR, Keyzers RA, Munro MHG, Prinsep MR (2012) Marine natural products. Nat Prod Rep 29:144–222

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

1 Laboratory Culture and Mutagenesis of Amphioxus (Branchiostoma floridae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Linda Z. Holland and Guang Li 2 Handling and Manipulation of Gametes and Embryos of the Annelidan Worm Pseudopotamilla occelata . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ryusaku Deguchi and Takeshi Nakano 3 The Starfish Asterina pectinifera: Collection and Maintenance of Adults and Rearing and Metamorphosis of Larvae . . . . . . . . . . . . . . . . . . . . . . . . Naoyuki Murabe, Ei-ichi Okumura, Kazuyoshi Chiba Enako Hosoda, Susumu Ikegami, and Takeo Kishimoto 4 Experimental Tools to Study Regeneration in the Sea Anemone Nematostella vectensis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aldine R. Amiel and Eric Ro¨ttinger 5 Staining and Tracking Methods for Studying Sponge Cell Dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carole Borchiellini, Sandie M. Degnan, Emilie Le Goff Caroline Rocher, Ame´lie Vernale, Stephen Baghdiguian Nina Se´journe´, Florent Marschal, Andre´ Le Bivic, Nelly Godefroy Bernard M. Degnan, and Emmanuelle Renard 6 Microscopy Studies of Placozoans. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carolyn L. Smith, Tatiana D. Mayorova, Christine A. Winters Thomas S. Reese, Sally P. Leys, and Andreas Heyland 7 Identification of SH2 Domain-Mediated Protein Interactions that Operate at Fertilization in the Sea Star Patiria miniata . . . . . . . . . . . . . . . . . Lauren Bates, Emily Wiseman, Jamie Kitson, and David J. Carroll 8 Marine Nemertean Worms for Immunoblotting Studies of Oocyte Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stephen A. Stricker 9 Recovery of Sea Star Egg Cell Surface Proteins Released at Fertilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emily Wiseman, Lauren Bates, and David J. Carroll 10 Quantifying Cell Proliferation During Regeneration of Aquatic Worms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ zpolat Eduardo E. Zattara and B. Duygu O

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In Situ Hybridization Techniques in the Homoscleromorph Sponge Oscarella lobularis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Laura Fierro-Constaı´n, Caroline Rocher, Florent Marschal Quentin Schenkelaars, Nina Se´journe´, Carole Borchiellini, and Emmanuelle Renard Methodology for Whole Mount and Fluorescent RNA In Situ Hybridization in Echinoderms: Single, Double, and Beyond . . . . . . . . . . . . . . . . . Margherita Perillo, Periklis Paganos, Maxwell Spurrell Maria I. Arnone, and Gary M. Wessel Gene Editing in the Ascidian Phallusia mammillata and Tail Nerve Cord Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alex McDougall, Celine Hebras, Isa Gomes, and Remi Dumollard Transcriptomic Analysis in the Sea Anemone Nematostella vectensis . . . . . . . . . . . Jacob F. Warner and Eric Ro¨ttinger RNA Interference on Regenerating Holothurian Gut Tissues . . . . . . . . . . . . . . . . Miosotis Alicea-Delgado, Samir A. Bello-Melo, and Jose´ E. Garcı´a-Arrara´s ATAC-Seq for Assaying Chromatin Accessibility Protocol Using Echinoderm Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marta S. Magri, Danila Voronov, Jovana Ranđelovic´ Claudia Cuomo, Jose Luis Gomez-Skarmeta, and Maria I. Arnone Usage of the Sea Urchin Hemicentrotus pulcherrimus Database, HpBase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sonoko Kinjo, Masato Kiyomoto, Takashi Yamamoto Kazuho Ikeo, and Shunsuke Yaguchi Functional Studies of Trichoplax adhaerens Voltage-Gated Calcium Channel Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julia Gauberg, Adriano Senatore, and Andreas Heyland A Bioinformatics Tutorial for Comparative Development Genomics in Diverse Meiofauna . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Joseph L. Sevigny, Jon L. Norenburg, and Francesca Leasi

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

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Contributors MIOSOTIS ALICEA-DELGADO • University of Puerto Rico, San Juan, Puerto Rico ALDINE R. AMIEL • Universite´ Coˆte d’Azur, CNRS, INSERM, Institute for Research on Cancer and Aging, Nice (IRCAN), Nice, France MARIA I. ARNONE • Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Naples, Italy STEPHEN BAGHDIGUIAN • ISEM, University of Montpellier, CNRS, EPHE, IRD, Montpellier, France LAUREN BATES • Department of Biomedical and Chemical Engineering and Sciences, Florida Institute of Technology, Melbourne, FL, USA SAMIR A. BELLO-MELO • University of Puerto Rico, San Juan, Puerto Rico CAROLE BORCHIELLINI • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France DAVID J. CARROLL • Department of Biochemistry and Molecular Genetics, Midwestern University, Glendale, AZ, USA KAZUYOSHI CHIBA • Department of Biological Sciences, Ochanomizu University, Bunkyo-ku, Tokyo, Japan CLAUDIA CUOMO • Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Naples, Italy BERNARD M. DEGNAN • Centre for Marine Science, School of Biological Sciences, The University of Queensland, Brisbane, QLD, Australia SANDIE M. DEGNAN • Centre for Marine Science, School of Biological Sciences, The University of Queensland, Brisbane, QLD, Australia RYUSAKU DEGUCHI • Miyagi University of Education, Sendai, Miyagi, Japan REMI DUMOLLARD • Laboratoire de Biologie du De´veloppement de Villefranche-sur-mer (LBDV), Sorbonne Universite´/CNRS, Villefranche-sur-Mer, France LAURA FIERRO-CONSTAI´N • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France JOSE´ E. GARCI´A-ARRARA´S • University of Puerto Rico, San Juan, Puerto Rico JULIA GAUBERG • Cell and Systems Biology, University of Toronto Mississauga, Mississauga, ON, Canada NELLY GODEFROY • ISEM, University of Montpellier, CNRS, EPHE, IRD, Montpellier, France ISA GOMES • Laboratoire de Biologie du De´veloppement de Villefranche-sur-mer (LBDV), Sorbonne Universite´/CNRS, Villefranche-sur-Mer, France JOSE LUIS GO´MEZ-SKARMETA • Centro Andaluz de Biologı´a del Desarrollo, CSIC/ Universidad Pablo de Olavide, Sevilla, Spain CELINE HEBRAS • Laboratoire de Biologie du De´veloppement de Villefranche-sur-mer (LBDV), Sorbonne Universite´/CNRS, Villefranche-sur-Mer, France ANDREAS HEYLAND • Department of Integrative Biology, College of Biological Science, University of Guelph, Guelph, ON, Canada LINDA Z. HOLLAND • Marine Biology Research Division, Scripps Institution of Oceanography, University of California San Diego, La Jolla, CA, USA

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ENAKO HOSODA • Department of Biological Sciences, Ochanomizu University, Bunkyo-ku, Tokyo, Japan SUSUMU IKEGAMI • Program of Food and AgriLife Science, Graduate School of Integrated Sciences for Life, Hiroshima University, Higashi-Hiroshima City, Hiroshima, Japan KAZUHO IKEO • Center for Information Biology, National Institute of Genetics, Shizuoka, Japan SONOKO KINJO • Department of Genomics and Evolutionary Biology, National Institute of Genetics, Shizuoka, Japan TAKEO KISHIMOTO • Science & Education Center, Ochanomizu University, Bunkyo-ku, Tokyo, Japan JAMIE KITSON • Department of Biomedical and Chemical Engineering and Sciences, Florida Institute of Technology, Melbourne, FL, USA MASATO KIYOMOTO • Marine and Coastal Research Center, Ochanomizu University, Chiba, Japan FRANCESCA LEASI • Department of Biology, Geology and Environmental Science, University of Tennessee at Chattanooga, Chattanooga, TN, USA ANDRE´ LE BIVIC • Aix Marseille University, CNRS, UMR 7288, IBDM, Marseille, France EMILIE LE GOFF • ISEM, University of Montpellier, CNRS, EPHE, IRD, Montpellier, France SALLY P. LEYS • Department of Biological Sciences, University of Alberta, Edmonton, AB, Canada GUANG LI • State Key Laboratory of Cellular Stress Biology, School of Life Sciences, Xiamen University, Xiamen, Fujian, China MARTA S. MAGRI • Centro Andaluz de Biologı´a del Desarrollo, CSIC/Universidad Pablo de Olavide, Sevilla, Spain FLORENT MARSCHAL • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France TATIANA D. MAYOROVA • National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA ALEX MCDOUGALL • Laboratoire de Biologie du De´veloppement de Villefranche-sur-mer (LBDV), Sorbonne Universite´/CNRS, Villefranche-sur-Mer, France NAOYUKI MURABE • Department of Physiology, Teikyo University School of Medicine, Itabashiku, Tokyo, Japan TAKESHI NAKANO • Sendai Daisan High School, Sendai, Miyagi, Japan JON L. NORENBURG • Department of Invertebrate Zoology, Smithsonian National Museum of Natural History, Washington, DC, USA EI-ICHI OKUMURA • School of Life Science and Technology, Tokyo Institute of Technology, Midori-ku, Yokohama, Japan ¨ ZPOLAT • Marine Biological Laboratory, Woods Hole, MA, USA B. DUYGU O PERIKLIS PAGANOS • Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Naples, Italy MARGHERITA PERILLO • Department of Molecular and Cellular Biology and Biochemistry, Brown University, Providence, RI, USA JOVANA RANĐELOVIC´ • Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Naples, Italy THOMAS S. REESE • National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA

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EMMANUELLE RENARD • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France; Aix Marseille University, CNRS, UMR 7288, IBDM, Marseille, France CAROLINE ROCHER • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France ERIC RO¨TTINGER • Universite´ Coˆte d’Azur, CNRS, INSERM, Institute for Research on Cancer and Aging, Nice (IRCAN), Nice, France QUENTIN SCHENKELAARS • Department of Genetics and Evolution, Faculty of Sciences, Institute of Genetics and Genomics in Geneva (IGe3), University of Geneva, Geneva, Switzerland; Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France NINA SE´JOURNE´ • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France ADRIANO SENATORE • Cell and Systems Biology, University of Toronto Mississauga, Mississauga, ON, Canada JOSEPH L. SEVIGNY • Hubbard Center for Genome Studies, Department of Molecular, Cellular, and Biomedical Sciences, University of New Hampshire, Durham, NH, USA CAROLYN L. SMITH • National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA MAXWELL SPURRELL • Department of Molecular and Cellular Biology and Biochemistry, Brown University, Providence, RI, USA STEPHEN A. STRICKER • Department of Biology, University of New Mexico, Albuquerque, NM, USA AME´LIE VERNALE • Aix Marseille Univ, Avignon Universite´, CNRS, IRD, IMBE, Marseille, France; Aix Marseille University, CNRS, UMR 7288, IBDM, Marseille, France DANILA VORONOV • Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Naples, Italy JACOB F. WARNER • Universite´ Coˆte d’Azur, CNRS, INSERM, Institute for Research on Cancer and Aging, Nice (IRCAN), Nice, France; Department of Biology and Marine Biology, University of North Carolina at Wilmington, Wilmington, NC, USA GARY M. WESSEL • Department of Molecular and Cellular Biology and Biochemistry, Brown University, Providence, RI, USA CHRISTINE A. WINTERS • National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD, USA EMILY WISEMAN • Department of Biomedical and Chemical Engineering and Sciences, Florida Institute of Technology, Melbourne, FL, USA SHUNSUKE YAGUCHI • Shimoda Marine Research Center, University of Tsukuba, Shizuoka, Japan TAKASHI YAMAMOTO • Department of Mathematical and Life Sciences, Graduate School of Science, Higashi-Hiroshima, Japan EDUARDO E. ZATTARA • Instituto de Investigaciones en Biodiversidad y Medio Ambiente (INIBIOMA), CONICET-Universidad Nacional del Comahue, Bariloche, Rio Negro, Argentina

Chapter 1 Laboratory Culture and Mutagenesis of Amphioxus (Branchiostoma floridae) Linda Z. Holland and Guang Li Abstract Cephalochordates (amphioxus) are invertebrate chordates closely related to vertebrates. As they are evolving very slowly, they are proving to be very appropriate for developmental genetics studies aimed at understanding how vertebrates evolved from their invertebrate ancestors. To date, techniques for gene knockdown and overexpression have been developed, but methods for continuous breeding cultures and generating germline mutants have been developed only recently. Here we describe methods for continuous laboratory breeding cultures of the cephalochordate Branchiostoma floridae and the TALEN and Tol2 methods for mutagenesis. Included are strategies for analyzing the mutants and raising successive generations to obtain homozygotes. These methods should be applicable to any warm water species of cephalochordates with a relatively short generation time of 3–4 months and a life span of 3 years or more. Key words Cephalochordate, Amphioxus, Branchiostoma, Aquaculture, Germ-line mutagenesis, Tol2, TALEN, Transgenics

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Introduction There are three genera of cephalochordates (amphioxus): Branchiostoma with 33 named species, Asymmetron with seven named species and Epigonichthys with two named species. The last two genera likely have additional cryptic species. Once thought to be the nearest invertebrate relative of vertebrates, molecular phylogenetic studies with nuclear genes have shown they are the sister group of vertebrates plus the fast-evolving tunicates [1]. Because of its resemblance to vertebrates, amphioxus species have long been the focus of embryological and evolutionary studies designed to give insights into how vertebrates evolved from their invertebrate ancestors. All three genera are exclusively marine, generally inhabiting near-shore waters. In nature, adults live burrowed in sand with just their extreme anterior ends exposed to the seawater, only emerging after sunset when they spawn. For many

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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years, research on developing embryos of Branchiostoma was hampered by inability to control spawning in the laboratory (Fig. 1). Female gametes are stored in the gonad as primary oocytes; on spawning days, which occur every ~10–14 days in summer, meiotic divisions commence in the early afternoon. Meiosis arrests at second metaphase and spawning occurs about 30 min after sundown. On the same day, the sperm acquire the ability to become motile on dilution into seawater. After fertilization, the zygote completes the meiotic divisions. Semicontrolled laboratory spawning of Branchiostoma floridae during the breeding season was developed in 1988 [2]. It was found that spawning could be delayed if animals that would normally spawn that day were collected in the afternoon and kept in the light; spawning could then be induced by a mild electric shock or by putting the animals in the dark. This allowed the development of techniques for manipulating embryos such as microinjection of antisense morpholino oligonucleotides for gene knockdowns, DNA constructs for analyzing gene regulatory sequences and mRNAs for gene overexpression [3–5]. The B. floridae genome is sequenced and assembled [6, 7], and a cDNA library of 210,000 clones has been end-sequenced (ESTs) [8]; individual clones are available from L. Z. H. In the laboratory, the life span of B. floridae is at least 3 years. Until recently, however, one essential technique for understanding how the genotype makes a phenotype—the generation of germline mutants—was missing. The first roadblock to obtaining germline mutants was that while several species of Branchiostoma (B. belcheri, B. japonicum, B. lanceolatum, B. floridae) as well as Asymmetron lucayanum (see Note 1) had been bred in the laboratory [9–12], most of the Branchiostoma species studied are cool-water species with a larval life of 3 months or more and relatively low survival to metamorphosis; A. lucayanum has not been raised to metamorphosis. B. floridae, a warm-water species, proved easy to raise through metamorphosis in the laboratory with nearly 100% survival (Figs. 1 and 4). B. floridae embryos and larvae develop normally from 21  C to 30  C. This is a major advantage as microinjection and manipulations of eggs and embryos can all be done at room temperature. Moreover, at the higher temperature, the life cycle of B. floridae from egg to ripe adult is only 3–4 months. A given individual will spawn repeatedly throughout its lifetime of at least 3 years. This means that the multiple generations necessary to obtain homozygous mutants require only 6–8 months for two generations and 9–12 months for three generations. This time frame is similar to that of the zebrafish. Because of these advantages, L. Holland established continuous breeding cultures of B. floridae in in the laboratory, and G. Li has adopted this species in developing techniques for transgenesis. Methods for generating germline transgenics include Tol2 transposons, TALEN and CRISPR/Cas9 [13]. Tol2 cannot be

Laboratory Culture and Mutagenesis of Amphioxus

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Fig. 1 Ripe female and ripe male specimens of Branchiostoma floridae. Anterior to the right. As the gonads are transparent, their color is that of the gametes. Each animal has 26 gonads per side. In the wild, most females have yellow oocytes, although a few have rather white ones. In the laboratory, the oocytes are relatively white. The difference in color between gonads from animals in the field versus the laboratory is presumably due to differences in the food. The males all have white to slightly grey gonads. Maximum length of adults is about 5 cm

directed to mutate a specific gene but has been used to insert exogenous DNA fragments at 10–30% ratios in amphioxus [14, 15]. To date, however, only TALEN has been adapted for genome editing in amphioxus [16]. Transcription activator-like (TAL) effectors were initially isolated from the bacterium Xanthomonas, a plant pathogen. The principle is that two peptides are constructed. Each consists of a TALE backbone including a central region with 12 to 31 nearly identical amino acid repeats, each composed of 34 amino acids differing from one another by only two amino acids at the 12th and 13th positions. These repeatvariable diresidues (RVD) determine the single nucleotides recognized by the TAL effector. Thus, NI binds to adenine, NH or NN to guanine, HD to cytosine, and NG to thymine. Two of these TALE proteins are engineered to opposite strands of DNA. At the C-terminal, each peptide has a spacer and the FokI restriction enzyme. Since FokI works as a dimer (it cleaves between any two bases to create double-stranded breaks), two arms containing assembled modules and a FokI domain are required to introduce double strand breakage at the target locus (Fig. 5). The only constraint is that the 50 bases to which the TALENs bind must be T. In most genomes, target sites for TALENs occur about every 35 bp. The repeat modules can be assembled in a specific order upstream of the FokI nuclease in a preconstructed vector, and the assembled modules can then bring a FokI nuclease to the target site. The TALEN method has been shown to be highly efficient in introducing indel mutations through nonhomologous end joining (NHEJ) pathway at the target loci in amphioxus. However, in amphioxus, it is currently almost impossible to insert exogenous

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DNA fragments at the target site with the TALEN method ( Import > Image Sequence” command. Select the necessary files and set up the file type in the next “Sequence Options” window (see Note 13). This creates a multipage image stack (here, each image file is called a “slice”), which can be saved as a single file through the “File > Save” or “File > Save As” command. It can then be opened with the “File > Open” command. Even after the construction of a stack file, unnecessary slices can be removed with the “Image > Stacks > Delete Slice” or “Image > Stacks >Tools > Slice Remover/ Slice Keeper” command. For trimming images, select the necessary region of the slices, which is termed a region of interest (ROI), with a rectangular area selection tool and trim the other region using the “Image > Scale” command. In this “Scale” window, the resolution of images can also be changed by entering appropriate values into the “X Scale” and “Y Scale” cells. 5. Measure the mean background fluorescence intensity by creating a ROI outside the target egg and executing the “Analyze > Measure” command (see Note 14). Next, subtract the background value from all slices with the “Process > Math > Subtract” command (see Note 15). To create an averaged baseline image, execute the “Image > Stacks > Z Project” command. In this window, enter the numbers of “Start Slice” and “Stop Slice” to specify the images immediately before a Ca2 + increase at fertilization with the selection of “Average Intensity.” To make a stack of “normalized” images, divide all slices by the averaged baseline image through the window opened by the “Process > Image Calculator” command and input data as follows: Image 1: a stack of images after background subtraction; Operation: “Divide”; and Image 2: an averaged baseline image, selecting “Create new window” and “32-bit (float) result” checkboxes. Change the color tone of the newly created normalized images through the “Image > Lookup Tables” command to select an appropriate LUT file, which can be edited and saved in a window following the “Image > Color > Edit LUT” command (see Note 16). Further, set the minimum and maximum values with the “Image > Adjust > Brightness/Contrast” command by pressing the “Set” button to enter the minimum and maximum displayed values. Remove

Handling and Manipulation of Gametes and Embryos of the Annelidan Worm. . .

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noise from the images if desired. We routinely perform noise removal using the “Process > Smooth” and/or “Process > Noise > Despeckle” command. 6. If desired, add timestamp information (e.g., the starting time of Ca2+ increase ¼ zero) to the slices with the “Image > Stacks > Label” command by entering the appropriate values in the next “Label Stacks” window (see Note 17). A scale bar can also be added using the “Analyze > Tools > Scale Bar” command after setting the scale (e.g., pixels/μm) in the window following the “Analyze > Set Scale” command. Then, create a montage of normalized images with the “Image > Stacks > Make Montage” command (Fig. 11A) (see Note 18). Display a LUT bar with the “Analyze > Tools > Calibration Bar” command. 7. Temporal changes in fluorescence intensity (F/F0) can be quantified by the “Image > Stacks > Measure Stack” command after setting a ROI inside the target egg. Numerical intensity values (mean gray values) appearing in the “Results” window can be copied to a spreadsheet. To measure fluorescence intensities in more than one region of the same image (Fig. 11B), open the ROI Manager using the “Analyze > Tools > ROI Manager” command. In this window, multiple ROIs can be recorded in the list by successively pushing the “Add” button or the “T” key on the keyboard. The regions and order of ROIs can be confirmed by selecting “Show All” and “Labels” checkboxes. Then, calculate the fluorescence intensities in the ROIs using the “More >> Multi Measure” command by selecting “Measure all slices” and “One row per slice” checkboxes (see Note 19). Similarly, use the numerical values in the “Results” window. 8. The stack file can also be saved as an AVI video file using the “File > Save As > AVI” command while entering the frame rate. Before saving as a video file, we sometimes draw arrows indicating noteworthy structures such as the cytoplasmic protrusion at fertilization. The shape and color of arrows can be edited using the “Edit > Options > Arrow Tool” and “Edit > Options > Colors” commands, respectively. To embed arrows into the slices, use the “Edit > Draw” command.

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Notes 1. Place the air stone above or beside the colony because worms directly exposed to air bubbles can be injured. 2. Wear a pair of close-fitting disposable gloves during this procedure to prevent fingers from being stung by the polyps of P. flavicirrata, which specifically live on the tip of the P. occelata tube.

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Fig. 11 Two-step Ca2+ increase in a fertilized egg of P. occelata. (A) Fluorescence images every 2 s are normalized by dividing them by an averaged baseline image just before the initiation of Ca2+ increase (time zero), and expressed as pseudocolor images. A cytoplasmic protrusion, which is manifested as a white region (arrowhead), is formed by the first-step localized Ca2+ increase and terminated by the second-step global Ca2 + increase. Scale bar: 100 μm. (B) Changes in relative fluorescence intensities (F/F0) measured in the three ROIs indicated

3. In the case of an upright microscope, observation at high magnification needs a long-focus objective lens to obtain sufficient working distance. In either inverted or upright microscope, adjust the angle formed by the microinjection needle and the microscope stage to ~10 . This enables a nearly “twodimensional” movement of the needle tip, which is desired for the microinjection with the chamber described in Fig. 9. 4. Select the thickness of the double-sided tape between the bottom and middle coverslips in accordance with the diameter and characteristics of the target oocyte or egg. We usually use double-sided tape of 70-μm thickness for immature oocytes of P. occelata, which have an irregular shape and become spherical in shape (~150 μm in diameter) after oocyte maturation, because successful microinjection cannot be accomplished without relatively strong pressure on them. The same spacer of double-sided tape can also be used for microinjection into mature eggs of P. occelata. By contrast, oocytes or eggs of the hydrozoan jellyfish Cytaeis uchidae (~100 μm in diameter) [13, 14] should never be compressed because of their fragile nature; double-sided tape of a 100-μm thickness is suitable. For oocytes or eggs of most other marine invertebrates, using a thickness slightly thinner (10–20 μm) than the oocyte or egg diameter may yield the desirable result.

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5. The use of low Ca2+ seawater facilitates microinjection into oocytes or eggs of P. occelata. However, the medium can lower the repair ability of oocytes or eggs in some other species. In such cases, microinjection should be performed in normal seawater. 6. Vigorously eject the oocytes from the mouth-operated pipette toward the opening of the space between bottom and middle coverslips. Otherwise, push the oocytes into the space using a fine metal needle. 7. Confirm the proper flow rate of the injection solution. This is easy when fluorescing substances (such as Ca2+ indicators) are injected under a fluorescence microscope. In this case, be careful not to permit long-term irradiation of excitation light. The flow of the injection solution can also be checked by brightfield observation when the tip of the glass needle is located at a forcibly ruptured oocyte, where particles of the cytoplasm are moved by the flow of the injection solution. 8. Tapping is a prerequisite for microinjection into oocytes or eggs of P. occelata and the hydrozoan jellyfish Cladonema pacificum [13, 15]. By contrast, the action is unnecessary for, and even injurious to, oocytes or eggs of some other animals, such as bivalves and limpets [16]. 9. Regulate the injection amount according to the size of the white spot. For calibration, we measured fluorescence intensities of oocytes injected with fluorescent dyes (such as FITC dextran) of known concentration and compared them with those of diluted dyes inserted into the space of the measurement chamber. In our experiments, the estimated injection amount was 4–8% of the oocyte volume. 10. Although intracellular Ca2+ changes can also be measured after loading of acetoxymethyl (AM) ester forms of Ca2+ dyes, this method fails to detect a rapid shift of fluorescence intensity in oocytes or eggs of some marine invertebrates. Recently developed blue light–exciting Ca2+ indicators such as Fluo-8, Cal-520, and Calbryte 520 show a greater increase in fluorescence intensity upon Ca2+ binding, with a higher signal-tonoise ratio. Among them, dextran conjugates are manufactured only in Cal-520, which may be most suitable for Ca2+ measurements in marine invertebrates, owing to the reduced compartmentalization and lower rate of dye leakage. 11. Prevent the fluorescence intensities from reaching saturation levels after fertilization. Saturated signals will underestimate the rate of increase. Once the gain and offset levels are determined, do not change the settings unless necessary. 12. Transmit light through a red band-pass filter (e.g., 600–650 nm), simultaneously using a beam splitter that can

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divide the light into two directions (an eyepiece and a camera) and a green band-pass filter (e.g., 520–560 nm) placed immediately in front of the camera head, facilitates bright-field observation through the eyepiece during the fluorescence recording. 13. With the “File > Import” command, various file types, including video file formats such as AVI, can be opened as a stack of images. 14. A mean value in the ROI is displayed in the “Results” window only when the “Mean gray value” checkbox is selected in the window opened by the “Analyze > Set Measurements” command. 15. The background subtraction is an essential step to examine the amount of increase in fluorescence intensity from the baseline level. When the background intensity is not uniform because of nonuniform illumination, obtain background images from a region under the same microscope other than from the eggs and subtract the averaged background image from all slices through the window opened by the “Process > Image Calculator” command. Input the data as follows: Image 1: a stack of original images; Operation: “Subtract”; and Image 2: a background image, selecting “Create new window” and “32-bit (float) result” checkboxes. Uneven background intensities can also be subtracted using the “Process > Subtract Background” command. If there is a significant autofluorescence in a target egg, the fluorescence intensity measured from a noninjected egg should also be subtracted. 16. A modified “royal” LUT bar is applied to the pseudocolor images in Fig. 11A. The edited LUT bar (file name: xxx.lut) can be applied to any images when the LUT file is opened using the “File > Open” command. 17. When the “Use overlay” checkbox is selected in the “Label Stacks” window, the timestamp can be edited using the submenu following the “Image > Overlay” command (e.g., “Hide Overlay,” “Show Overlay,” and “Remove Overlay”). However, the timestamp will be lost when a montage is constructed. To create a stack in which the timestamp is embedded, use the “Image > Overlay > Flatten” command. Alternatively, create the embedded timestamp directly using the “Image > Stacks > Series Labeler” command. 18. The montage of Fig. 11A is constructed from the fluorescence images captured every 1 s. The input data in the “Make Montage” window are as follows: columns: 7, rows: 7, first slice: 59 (an image 2 s before the beginning of Ca2+ increase), and increment: 2. A scale bar is added to the first slice.

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19. Note that information about the locations of ROIs will be lost when the image is copied and pasted into a new image file. To create the image where ROIs are embedded, push the “Flatten” button in the ROI Manager window and select “No” in the next window that asks if all images are processed. Alternatively, take a screenshot through other applications such as Preview (Macintosh).

Acknowledgments This work was supported by Japan Society for the Promotion of Science (JSPS) KAKENHI under grant numbers 26440177 and 17K07482 for R.D. References 1. Weigert A, Helm C, Meyer M et al (2014) Illuminating the base of the annelid tree using transcriptomics. Mol Biol Evol 31 (6):1391–1401. https://doi.org/10.1093/ molbev/msu080 2. Struck TH, Golombek A, Weigert A et al (2015) The evolution of annelids reveals two adaptive routes to the interstitial realm. Curr Biol 25(15):1993–1999. https://doi.org/10. 1016/j.cub.2015.06.007 3. Nakano T, Kyozuka K, Deguchi R (2008) Novel two-step Ca2+ increase and its mechanisms and functions at fertilization in oocytes of the annelidan worm Pseudopotamilla occelata. Dev Growth Differ 50(5):365–379. https:// doi.org/10.1111/j.1440-169X.2008. 01022.x 4. Nakano T, Deguchi R, Kyozuka K (2014) Intracellular calcium signaling in the fertilized eggs of Annelida. Biochem Biophys Res Commun 450(3):1188–1194. https://doi.org/10. 1016/j.bbrc.2014.06.056 5. Nakano T, Kyozuka K (2015) Soluble sperm extract specifically recapitulates the initial phase of the Ca2+ response in the fertilized oocyte of P. occelata following a G-protein/PLCβ signaling pathway. Zygote 23(6):821–835. https:// doi.org/10.1017/S0967199414000501 6. Ishii T, Otake T, Okoshi K et al (1994) Intracellular localization of vanadium in the fan worm Pseudopotamilla occelata. Mar Biol 121 (1):143–151. https://doi.org/10.1007/ BF00349483 7. Yamaguchi N, Yoshinaga M, Kamino K, Ueki T (2016) Vanadium-binding ability of nucleoside diphosphate kinase from the vanadium-rich fan worm, Pseudopotamilla occelata. Zool Sci 33

(3):266–271. https://doi.org/10.2108/ zs150188 8. Hirai E, Kakinuma Y (1973) Differentiation and symbiosis in two hydrozoans. Publ Seto Mar Biol Lab 20:257–273 9. Deguchi R, Takeda N, Stricker SA (2011) Comparative biology of cAMP-induced germinal vesicle breakdown in marine invertebrate oocytes. Mol Reprod Dev 78 (10–11):708–725. https://doi.org/10.1002/ mrd.21346 10. Momose T, Concordet JP (2016) Diving into marine genomics with CRISPR/Cas9 systems. Mar Genomics 30:55–65. https://doi.org/10. 1016/j.margen.2016.10.003 11. Takahashi A, Camacho P, Lechleiter JD et al (1999) Measurement of intracellular calcium. Physiol Rev 79(4):1089–1125. https://doi. org/10.1152/physrev.1999.79.4.1089 12. Bootman MD, Rietdorf K, Collins T et al (2013) Ca2+-sensitive fluorescent dyes and intracellular Ca2+ imaging. Cold Spring Harb Protoc 2013(2):83–99. https://doi.org/10. 1101/pdb.top066050 13. Deguchi R, Kondoh E, Itoh J (2005) Spatiotemporal characteristics and mechanisms of intracellular Ca2+ increases at fertilization in eggs of jellyfish (Phylum Cnidaria, Class Hydrozoa). Dev Biol 279(2):291–307. https://doi.org/10.1016/j.ydbio.2004.11. 036 14. Arakawa M, Takeda N, Tachibana K et al (2014) Polyspermy block in jellyfish eggs: collaborative controls by Ca2+ and MAPK. Dev Biol 392(1):80–92. https://doi.org/10. 1016/j.ydbio.2014.04.020

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15. Kondoh E, Tachibana K, Deguchi R (2006) Intracellular Ca2+ increase induces postfertilization events via MAP kinase dephosphorylation in eggs of the hydrozoan jellyfish Cladonema pacificum. Dev Biol 293 (1):228–241. https://doi.org/10.1016/j. ydbio.2006.02.005

16. Deguchi R (2007) Fertilization causes a single Ca2+ increase that fully depends on Ca2+ influx in oocytes of limpets (Phylum Mollusca, Class Gastropoda). Dev Biol 304(2):652–663. https://doi.org/10.1016/j.ydbio.2007.01. 017

Chapter 3 The Starfish Asterina pectinifera: Collection and Maintenance of Adults and Rearing and Metamorphosis of Larvae Naoyuki Murabe, Ei-ichi Okumura, Kazuyoshi Chiba, Enako Hosoda, Susumu Ikegami, and Takeo Kishimoto Abstract Here we describe methods for (a) collecting starfish during their breeding period; (b) maintaining adults with fully grown gonads in laboratory aquaria; (c) rearing fertilized eggs to brachiolaria larvae, and (d) inducing larvae to metamorphose into juveniles under laboratory conditions. Such protocols should facilitate various analyses of starfish development throughout the entire life cycle of these model organisms. Key words Starfish, Asterina pectinifera, Breeding Period, Laboratory Aquarium, Fertilization, Embryonic and Larval Development, Metamorphosis, Oocytes, Eggs, Brachiolaria Larvae

1

Introduction The year 2019 marks the 50th anniversary of the identification of 1-methyladenine (1-MeAde) in starfish by Haruo Kanatani (1930–1984) and colleagues as the first bona fide maturationinducing hormone in metazoan oocytes [1, 2]. A decade prior to the discovery of 1-MeAde, Chaet and McConnaughy (1959) found that the starfish radial nerve contains a gonadotropin-like substance that was later named gonad-stimulating substance (GSS) [3]. Seven years after 1-MeAde was characterized, MPF (maturationpromoting factor) was demonstrated in starfish oocytes [4, 5]. Retracing these steps, GSS opened the window to studies of reproductive endocrinology in invertebrates, whereas 1-MeAde enabled in vitro maturation of isolated immature oocytes, and MPF connected starfish oocytes to universal M-phase control in eukaryotic cells [2, 5]. Historically, analyses of starfish have made enormous contributions to classical embryology and more modern aspects of

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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developmental biology. In addition, recent progress made in genome editing techniques should allow for molecular developmental analyses to be conducted throughout the life cycle of starfish. In support of such efforts, Takehiko Itoh of Tokyo Institute of Technology [6] and our group have nearly finished the starfish genome project and are now preparing to publish well-assembled haplotype genome sequence for Asterina pectinifera (N50: 2.1 Mbp) along with transcriptome data. Thus, to complement ontogenetic investigations employing genomic methods, it would be helpful to review how the starfish life cycle can be maintained in the laboratory, particularly emphasizing the induction of larval metamorphosis under laboratory conditions. Here we focus on Asterina pectinifera (renamed Patiria pectinifera in the 2007 NCBI Taxonomy Browser) and, as an introduction for those planning to begin analyses of starfish describe first how to obtain adult starfish in the coastal sea and maintain them in laboratory aquaria. Although some methods may be specific for researchers in Japanese institutions, the general methodology described here can be also easily adapted to other locales throughout the world. Thereafter, methods are introduced for induction of oocyte maturation, fertilization, embryonic development from fertilized eggs to blastula, gastrula, bipinnaria, and brachiolaria larvae, and finally induction of metamorphosis to obtain juvenile starfish.

2

Materials

2.1 Collection and Selection of Starfish

1. An international marine signal A flag, approx. L60  W90 (cm). 2. Two diving signboards, approx. L30  W120 (cm), with ropes. 3. Three red flags, approx. L30  W40 (cm), with wooden handles. 4. A wooden hammer and a hand loudspeaker. 5. Six catch bags. 6. Two ropes with hooks. 7. Ten bread trays, approx. L40  W60  H8 (cm). 8. Forceps and scissors. 9. Thin gloves. 10. Ten net cage hangers with ropes. 11. Marker pens and a packing tape roll. 12. Papers such as newspapers. 13. A bucket with a rope. 14. Plastic bags. 15. Several cold packs and a cooler box. 16. A rental minivan.

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1. Each unit of the aquarium comprises (a) three holding tanks (each ~100 L of aerated seawater) to keep starfish, (b) a filtration tank (~200 L) containing filtration stones, (c) a temperature controller, (d) a pump to circulate seawater, (e) an aeration pump, and (f) a heat exchanger (separated outside of the room) to regulate temperature of the circulating seawater (Fig. 1a–d) (see Note 1). 2. Temperature-controlled seawater is delivered to each starfish tank along with air. Thereafter, seawater overflowing from the starfish tanks enters the filtration tank (Fig. 1a). Such circulation along with aeration via air stones supply sufficient O2 while also stirring the seawater in each starfish tank. 3. Each tank is made of transparent acrylic plates, while the frame (blue in Fig. 1b, c) is made of FRP (fiber-reinforced plastics). 4. Coral sands are recommended as filtration stones. Alternatively, sand gravel is used as filtration sands. “Gravel” of two types of

Fig. 1 Laboratory aquarium for starfish. (a) One unit of aquarium. Major components are three starfish tanks (each ~100 L with aeration), a filtration tank (~200 L), a pump to circulate seawater, and a heat exchanger outside of the room (see right side in d). Each starfish tank size is W43 cm, D50 cm, and H50 cm; and the filtration tank size is W90 cm, D50 cm, and H50 cm. (b) Actual view of one unit of aquarium. Regulator panel (pale cream color) for seawater temperature is seen in the middle right. (c) Whole view of the aquarium room. Several units are equipped. (d) Seawater tank is built outside of the aquarium room. To supply natural seawater, pipeline is connected from the seawater tank (4 t) to each unit of aquarium inside of the room

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particle size (3.5–7 mm and 2–3.5 mm) are available from “NIHON GENRYO” (http://www.genryo.co.jp/sandgravel.html). Each type is kept in a mesh bag of ~5 cm height, and is piled up as two layers (see Note 2). 5. Preferably, more than one unit of aquarium should be equipped. Each unit should be independent, that is, seawater circulates only within each unit, and its temperature is controlled separately (Fig. 1b, c) (see Note 3). 6. When you use natural seawater, a seawater tank (approx. 4 tons) is built outside of the aquarium room (Fig. 1d). Each unit of aquarium can be supplied with seawater from this seawater tank. 2.3 Seawater for Aquaria

1. Natural seawater can be purchased from Tokai Kisen Co. Ltd. (https://www.tokaikisen.co.jp/), which ships to the Izu Islands. When the ship comes back to Tokyo, its tank is filled for the balance with natural seawater obtained near the coast of Hachijojima Island. Usually, 2–4 tons of natural seawater is purchased per transaction and kept in a seawater tank (Fig. 1d). 2. Alternatively, artificial seawater for aquarium can be purchased. For example, “REI-SEA marine II 500L” is available from “REI-SEA IWAKI” (https://rei-sea.iwakipumps.jp/ products/filter/marine/), and “SEALIFE 250” from “MARINE-TECH” (http://www.marine-tech.co.jp/sealife/) (see Note 4).

2.4 Oocyte Maturation, Insemination, and Embryonic Development

[Prepare all solutions using ultrafiltered water purified by Milli-Q HR-7000 High- Throughput Central Water (Merck KGaA, Darmstadt, Germany) and analytical grade reagents.] 1. Commercial ASW: Marine Art SF-1 (Tomita Pharmaceutical, Tokushima, Japan) (see Note 5). 2. 90-mm dishes. 3. Iridectomy scissors. 4. Tweezers and 35-mm dishes. 5. Graduated centrifuge tubes (15 mL and 50 mL). 6. Pipettes (10 mL). 7. Pasteur pipettes and pipette bulbs. 8. 1-Methyladenine (1-MeAde; Acros Organics, Geel, Belgium) dissolved in ASW. 9. Inverted microscope objective.

and

stereomicroscope

10. 1-L beakers. 11. Stirring motors (60 rpm) (see Note 6).

with

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12. Hand centrifuge. 13. Roller 6 Digital Shaker model 4011 000, IKA CO., Wilmington, North Carolina, U.S.A. 2.5 Collection of Normal Embryos

1. 200 mm dishes. 2. Pipettes (10 mL). 3. Nylon mesh (100 μm mesh size).

2.6

Diatom Culture

1. Diatoms Chaetoceros gracilis and C. calcitrans, food for starfish larvae, were purchased from Bivalve Culture Institute, Sasebo, Nagasaki, Japan, and MarineTec Co., Yokohama, Japan, respectively. C. gracilis and C. calcitrans strains can be obtained from The University of Texas at Austin (www.utex.org), and Bigelow Laboratory for Ocean Sciences (ncma.bigelow.org), East Boothbay, Maine, USA, respectively. 2. Chaetoceros medium stock solution A in 100 mL ultrafiltered water: 10 g NaNO3, 1.0 mg sodium β-glycerophosphate, 0.9 mg EDTA-Fe, 1.3 g tris (hydroxymethyl) aminomethaneHCl (pH 7.0), 20 mg vitamin B12, and 6 g Clewat 32 (Nagase Chemitex Co., Osaka, Japan), a mixture of EDTA chelates of Co, B, Mn, Fe, Cu, Zn and Mo. 3. Chaetoceros medium stock solution B in 100 mL ultrafiltered water: 0.31 g Na2SiO3  9H2O. 4. Chaetoceros medium stock solution C in 100 mL ultrafiltered water: 7 g NaNO3, 0.45 g KH2PO4, 0.38 g EDTA-Fe, 0.5 g Clewat 32, 3 g Na2SiO3  9H2O, 100 μg biotin, 10 mg thiamin, and 1.2 mg vitamin B12. The solution is kept in the dark to prevent degradation of thiamin and vitamin B12. 5. Silicosen vent stoppers, Shin-Etsu Polymer Co. Ltd., Tokyo, Japan. 6. Silicone stoppers. 7. Air pump, air stones, airline tubing, and connectors to bubble air through diatom culture. 8. Desktop waving shaker model Wave-PR, Taitec CO., Koshigaya, Saitama, Japan. 9. Thoma-Zeiss hemocytometer. 10. Conical flasks (0.5 L, 0.3 L, 0.1 L). 11. 50 mL centrifuge tubes. 12. 0.5 L glass Roux culture bottles with offset neck. 13. Milex filter unit (0.22 μm), Millipore/Merck KGaA, Darmstadt, Germany.

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2.7 Induction of Metamorphosis

1. 24-well plates. 2. Pebbles obtained from an aquarium in which adult starfish are maintained. 3. Disposable hypodermic needles (27 gauge).

3

Methods

3.1 Collection of Adult Starfish

3.2 Getting Collection Permits from Local Agencies

Shallow-water starfish typically have a single breeding period each year. In Japan, Asterina pectinifera breeds in early May in Tokyo Bay, early June in Ise Bay (Mie Prefecture), early July in Choushi (Chiba Prefecture), and early September in Mutsu Bay (Aomori Prefecture). If you plan to collect starfish in new area, you should first establish when peak breeding occurs, which can be estimated based on examination of relative gonad sizes (see Subheading 3.4). Although Japan is surrounded by the sea, it can be difficult to obtain sufficient numbers of starfish during the breeding period from marine biological stations. As far as we know, there are no commercial dealers for starfish in Japan. Thus, along with small numbers of starfish that we personally collect, we, sometimes in collaboration with several laboratories, hire professional divers to collect several hundreds or thousands of starfish (see following sections). Usually, we collect starfish with fully grown gonads just before spawning, that is, at the beginning of the breeding period. Immediately after the collection, starfish are sorted into female, male, or “empty” (poor gonad) at the seashore. Female and male starfish are brought back to laboratory aquaria, whereas empty starfish are returned to the sea for preservation. 1. Because whole coast around the Japanese Archipelago is officially controlled, more than a month prior to the collection date, contact the corresponding local office of the Japan Fisheries Cooperative to obtain a collecting permit and to have other concerned parties notified of the planned collection (see Note 7). 2. Contact the corresponding local office of the Navigation Safety Division, Maritime Traffic Department of Japan Coast Guard and send an application for starfish collection (see Note 8). For this application, it is also necessary to notify the local fire station and yacht harbor office regarding the collection. 3. Contact the Fisheries Adjustment Commissions of the local prefectural government office and get permission for the collection of starfish (see Note 9). 4. Contact the Port Authority of the local prefectural government office and get permission to use a place of shoreline for starfish collection.

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1. Arrange with a local diving company for two divers and with a local office of the Japan Fisheries Cooperative for a boat and a pilot more than a month prior to the collection date. 2. On the collection day, lift the international marine signal flag A and put diving signboards on the both sides of the boat. 3. Appoint three lifeguards who board the boat with SCUBA (self-contained underwater breathing apparatus) divers and watch divers from the boat during diving. 4. Move the boat to the chosen dive locale. 5. Give a catch bag to each diver, and verify their diving safety through watching bubbles on the sea surface. If other boats approach the dive boat, lifeguards must wave red flag and warn them of the danger with a hand loudspeaker. Divers should also be alerted by knocking the boat edge with a wooden hammer (see Note 10). 6. Receive from the diver a catch bag filled with starfish by lifting its attached rope with a hook. 7. Transfer starfish from the catch bag into a net cage hanger, and hang it with ropes at the boat edge. 8. Come back to the pier when SCUBA tanks are empty, and, if needed, collect more starfish after the tanks are replaced with charged ones and the divers have taken their required break.

3.4 Selection of Starfish

1. Label three bread trays respectively as “female” and “male” with a marker and a tape. 2. After the initial dive, put collected starfish into unlabeled trays. 3. Pick up each starfish with a gloved hand and make a small hole on the ventral side by a forceps or scissors. 4. Push the dorsal side softly with fingers and watch inside of the hole carefully. 5. When an orange colored ovary is observed, put the starfish into “female” tray. When a white colored testis is observed, put the starfish into “male” tray. If nothing or poor gonad is observed, put the starfish outside of trays as an “empty” starfish. 6. Count the number of starfish of “female”, “male” and “empty”. 7. Put the sorted starfish separately into net cage hangers with ropes labeled “female,” “male,” or “empty.” 8. Repeat 1–7 for subsequent dives. 9. After finishing the starfish sorting, return “empty” starfish to sea for conservation.

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3.5 Packing and Transport

1. Share the collected starfish between laboratories depending on their required number of starfish. 2. Put newspapers into bread trays filled with seawater which is drawn by a bucket with a rope. 3. Wrap the starfish in wet newspaper. Usually, put 5–10 starfish per a page. 4. Put each pack of the wrapped starfish into a plastic bag labeled with “female” or “male”. 5. Put the plastic bags containing the wrapped starfish into a cooler box with several cold packs (see Note 11). 6. Transport the cooler box to the aquarium in each lab by car or train.

3.6 Maintenance of Adult Starfish in Laboratory Aquaria

Female and male starfish collected from the sea are kept in laboratory aquaria with circulating cold seawater. A major purpose is to maintain fully grown gonads (just before spawning) for experimental use (but not to breed starfish) as long as possible. Typically, fully grown gonads appropriate for experiments are available for a couple of months and sometimes half a year or more. Adult starfish can usually survive for several months without feeding even in a simple aquarium with an aeration system, such as used for keeping small pet fishes at home. A key to prolonging survival under such conditions is keeping starfish at relatively low population densities and at 10–11  C. To optimize the bacterial flora needed in aquaria, small amounts of stone or sand from the shallow sea or the sea shore should be added to the aquarium setup (see Note 2).

3.7 Managing Laboratory Populations of Starfish

1. Each 100-L starfish tank can theoretically hold up to ~30 Asterina pectinifera, and hence, keeping 99%. The vessel was a 0.5-L glass Roux bottle in which the opening was closed with an air-tight silicon

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stopper. The diatom cells were collected by centrifugation and the cell lysate was subjected to SDS-polyacrylamide gel electrophoresis. The major ribulose bisphosphate carboxylase large chain band was cut out and subjected to in-gel digestion with trypsin. Tryptic peptides were fractionated and analyzed by nanoflow LC-MS/MS, and database searches were performed with Mascot. It was found that 50–85% of 36 carbon atoms of an oligopeptide containing 7 amino acid residues was 13C (unpublished results). These data show that highly efficient in vivo stable isotopic labeling of diatom cells can be achieved in a relatively simple way. Combined with a spectrometric technique, labeled diatom cells will be useful for quantitative analysis of feed intake, digestion, absorption and utilization of nutrients, especially for individual proteins produced during embryonic and larval growth. 18. The bipinnaria larva develops to the brachiolaria gradually. The brachiolaria is characterized by the presence of the brachiolar arms in the anterior, which are used for exploring the surface of benthic environment (Fig. 3b, c). The larva develops the brachiolar arms in the anterior preoral lobe and the adult rudiment in the posterior concomitantly. Although the transition from bipinnaria to brachiolaria is gradual, once the brachiolar arm is visible under a stereomicroscope, the larva is competent to metamorphose. Brachiolaria larvae keep growing further in their body mass if they do not encounter environmental cue (s) for metamorphosis [7]. 19. When changing ASW, wipe all inside surfaces of the beaker with Kimwipes moistened with alcohol in order to prevent the growth of bacteria and the formation of primary microbial “biofilm,” which could be cues for metamorphosis. 20. Competent brachiolaria larvae initiate metamorphosis in response to 3 h incubation with environmental cues from pebbles taken from the adult habitat. After exploration of pebble surfaces by means of brachiolar arms, the larva irreversibly attaches to the surface through the adhesive disk located in the base of the brachiolar arms. The anterior body of the larva is absorbed inward, while the adult rudiment in the posterior transforms into the body of the juvenile. Since the juvenile podium forms only after metamorphosis in A. pectinifera, this organ is a reliable marker for metamorphosis. Juvenile podia can be found around 24 h after triggering metamorphosis, although it is hard to distinguish them from degenerating larval structures. Practically, it is easy to identify juvenile podia 48 h after incubation, because the absorption of the larval body is almost completed [8]. Yamakawa et al. [9] showed that exogenous application of 1 μM all-trans retinoic acid to 8- to 12-day-old brachiolaria induced metamorphosis

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commencement, suggesting that retinoic acid signaling functions as an endogenous mediator of environmental cues received by adhesive papillae on the brachiolar arm upon attachment to a substrate [8]. 21. Juvenile A. pectinifera starts to use tube feet to move around 7 days after metamorphosis. The authors have not reared juveniles further than this. Yamaguchi [10] reported the growth to sexual maturity of laboratory-reared crown-of-thorns starfish, Acanthaster planci, which is phylogenetically closely related to A. pectinifera [11]. Although adults feed on corals, early juveniles are herbivorous, feeding on coralline algae. A. planci juveniles complete transformation from algae-feeding to coral-eating within 5 months after metamorphosing. Sexually mature adults were obtained 17–20 months after metamorphosis, that is, 18–21 months after fertilization.

Acknowledgments N.M. and S.I. gratefully acknowledge Professor Hiroyuki Kaneko, Department of Biology, Keio University, in whose laboratory most of larval rearing and metamorphosis research was conducted. This work was supported by grants-in-aid to S.I. from JSPS [grant numbers 15 K14806, 19580129], to T.K. from JSPS [grant numbers 25291043, 16H04783] and the Takeda Science Foundation. The authors declare no competing financial interests. Note added in proof Interested readers may contact Ei-ichi Okumura at for starfish collection, Kazuyoshi Chiba at for starfish maintenance in aquarium, Susumu Ikegami at for starfish larvae and metamorphosis; and Takeo Kishimoto at for the rest. References 1. Kanatani H, Shirai H, Nakanishi K, Kurokawa T (1969) Isolation and identification of meiosis-inducing substance in starfish, Asterias amurensis. Nature 221:273–274 2. Kishimoto T (2018) MPF-based meiotic cell cycle control: half a century of lessons from starfish oocytes. Proc Jpn Acad Ser B 94:180–203 3. Chaet AB, McConnaughy RA (1959) Physiologic activity of nerve extracts. Biol Bull 117:407–408 4. Kishimoto T, Kanatani H (1976) Cytoplasmic factor responsible for germinal vesicle

breakdown and meiotic maturation in starfish oocyte. Nature 260:321–322 5. Kishimoto T (2015) Entry into mitosis: a solution to the decades-long enigma of MPF. Chromosoma 124:417–428 6. Kajitani R, Yoshimura D, Okuno M et al (2019) Platanus-allee is a de novo haplotype assembler enabling a comprehensive access to divergent heterozygous regions. Nat Commun 10:1702 7. Murabe N et al (2010) Competent larva in the starfish, Asterina pectinifera shifts from larval growth to juvenile formation by using environmental cue(s). In: Harris LG, Boetger SA,

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Walker CW, Lesser MP (eds) Echinoderms: Durham - Proceedings of the 12th International Echinoderm Conference, Durham, pp 463-466 8. Murabe N et al (2007) Adhesive papillae on the brachiolar arms of brachiolaria larvae in two starfishes, Asterina pectinifera and Asterias amurensis, are sensors for metamorphic inducing factor(s). Develop Growth Differ 49:647–656

9. Yamakawa S et al (2018) The role of retinoic acid signaling in starfish metamorphosis. EvoDevo 19:1–10 10. Yamaguchi M (1974) Growth of juvenile Acanthaster planci (L.) in the laboratory. Pacific Sci 28:123–138 11. Matsubara M et al (2004) Close relationship between Asterina and Solasteridae (Asteroidea) supported by both nuclear and mitochondrial gene molecular phylogenies. Zool Sci 21:785–793

Chapter 4 Experimental Tools to Study Regeneration in the Sea Anemone Nematostella vectensis Aldine R. Amiel and Eric Ro¨ttinger Abstract Animal regeneration is a biological process leading to the reformation of injured or lost tissues/body parts. One of the most fascinating regenerative phenomena is the so-called whole-body regeneration, leading to the reformation of fully functional organisms within days after bisection. The sea anemone Nematostella vectensis is currently emerging as novel whole-body regeneration model. Here we describe the methods of inducing the regenerative process in this cnidarian as well as the fixation and staining protocols for morphological, molecular, and cellular analysis. Key words Whole body regeneration, Bisection, Fixation, Permeabilization, Immunohistochemistry, Staining

1

Introduction Animal regeneration has intrigued scientists and humankind for centuries. This biological process leads to the reformation of injured or lost tissues/body and occurs at different levels from the cell, tissue, organ, structure to even the entire organism [1]. The phenomenon is called whole body regeneration, and Hydra as well as planarians are the historical models in this line of research. Since a couple of decades, the field of regeneration biology has seen the emergence of a variety of new research models, one of which is the cnidarian sea anemone Nematostella vectensis [2–5]. As cnidarians are the sister group to all bilaterian animals [6, 7], this marine invertebrate holds an interesting phylogenetic position and provides new insights into the evolution of this intriguing developmental trajectory. After bisection, regeneration in adult and juvenile Nematostella occurs within 7 days. Developing cellular and molecular tools in order to study the dynamics of the tissues as well as cellular response/behaviour during regeneration, is crucial for our understanding of this intriguing phenomenon

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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[5]. Using four-tentacle juveniles that are more amenable for staining and imaging approaches we have adapted/optimized protocols to highlight cellular and molecular mechanisms involved in the regeneration process in Nematostella that are based on (a) small molecules, such as DAPI (40 ,6-diamidine-20 -phenylindole dihydrochloride) or HOECHST (Bisbenzimide) to label nuclei, EdU (5-ethynyl-20 -deoxyuridine) to label cell proliferation/S-phase, EU (5-ethynyl-uridine) to label neosynthesized RNA (i.e., hypertranscription) or phalloidin to label cell membranes (actin filaments) as well as (b) antibody staining. Combined with molecular information (e.g., transcriptomic time courses [8], see Chapter 14) these approaches are useful tools to determine the phenotypes of perturbation experiments (i.e., pharmacological or gene-specific knockdowns/knockouts) [5] and predict gene network interactions.

2

Materials Prepare and store all reagents at room temperature (unless indicated otherwise). Follow all national waste disposal regulations when discarding waste materials.

2.1

Animal Care

1. Nematostella vectensis (Nv) polyps. 2. One-third strength artificial sea water (1/3 ASW), pH 8.2, density 10.10, KH 7, prepared from artificial sea water (ASW pH 8.2, density 10.30, KH 7) diluted to 1:2 with reverse osmosis (RO) water KH7 (4.5 g sodium bicarbonate in 20 L RO water). 3. Stackable glass bowls for approx. 200 ml 1/3 ASW (e.g., Carolina, 250 ml, #741004). 4. Incubator to maintain animals at 22  C (e.g., POL-EKOAPARATURA, ST-700). 5. Artemia salina—nauplius stages. 6. Scalpels.

2.2 Induction of Regeneration

1. Relaxing solution—magnesium chloride 7.14% in 1/3 Microfiltered ASW—0.2 μm, or 1/3 MFASW): 7.14 g of MgCl2 in 100 ml of 1/3 ASW. Store at room temperature (RT). 2. Petri dishes for approximately 40 ml of 1/3 ASW (e.g., Greiner, #633185). 3. Plastic pipette, 2 ml (e.g., Biosigma, #390509). 4. Scalpel handle (Swann-Morton n 7) with blade (SwannMorton n 15).

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5. P200 pipette (e.g., Gilson P200) and 200 μl tips. 6. Microfiltered 1/3 artificial sea water (1/3 MFASW). 2.3 Fixation, Permeabilization, Coating, and Washing Buffer for Immunostaining and EdU/EU Staining

1. Fixative IS: 4% paraformaldehyde (from 32% PFA without methanol stock solution, e.g., EMS, #15714, see Note 1) in 1/3 MFSW. Can be kept at 4  C for 1 week. 2. Ice. 3. Phosphate buffered saline 10 (PBS 10): for 1 l solution add 2.23 g NaH2PO4 (anhydrous 18.6 mM); 11.94 g Na2HPO4 (81.1 mM); 102.2 g NaCl; adjust pH to 7.4. Autoclave. 4. Tween 20 20% stock solution (autoclaved MilliQ water). 5. Tween 20 0.1% in PBS 1 (0.1% PBTw): For 50 ml add 1 ml PBS 10; 0.1250 ml Tween 20 20%; 48.75 ml MilliQ water. 6. Triton X-100 0.2% and 0.5% in PBS 1 (0.2% and 0.5% PBT, respectively). 7. Methanol (MeOH).

2.4 Membrane and Nuclei Staining

All products are stored at 20  C. 1. Alexa Fluor® 488 Phalloidin (Thermo Fisher Scientific, #A12379, see Note 1). 2. DAPI (Thermo Fisher Scientific, #D3571, see Note 1) 10.9 mM stock solution or Hoechst (Thermo Fisher Scientific, #H1399, see Note 1) 10 mg/ml stock solution: use at 1/5000 from stock solution in PBS1.

2.5 EdU (5-Ethynyl20 -Deoxyuridine) and EU (5-Ethynyl-Uridine) Staining

Protocol adapted from the Click-iT™ kits, Thermo Fisher Scientific—EdU # C10337, EU # C10329 1. EdU solution (see Note 1): dilute EdU powder in 2 ml of deionized water for 10 mM stock solution. Aliquot and store at 20  C. 2. EU solution (see Note 1): dilute EU powder in 373 μl of deionized water for 100 mM stock solution. Aliquot and store at 20  C. 3. Deionized water. 4. 10 Click it reaction buffer (Click-iT™ kits, stored at 4  C) 5. CuSO4 100 mM stock solution (Click-iT™ kits, stored at 4  C). 6. Alexa Fluor Azide 488 (Click-iT™ kits, stored at 20  C). 7. Reaction Buffer Additive (Click-iT™ kits, stored at 20  C).

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TUNEL Staining

Protocol adapted #11772457001)

from

the

TUNEL

AP

kit

(Roche,

1. Proteinase K 0.01 mg/ml in PBS1. 2. TUNEL Enzyme solution (Kit Roche, stored at 20  C). 3. TUNEL label (Kit Roche, stored at 20  C). 4. DNase at 0.1 U/μl for positive control. 5. Fixative IS: 4% Paraformaldehyde (from 32% PFA without Methanol stock solution, e.g., EMS, #15714, see Note 1) in 1/3 MFSW. Can be kept at 4  C for 1 week. 6. Secondary fixative: 4% Paraformaldehyde (from 32% PFA without Methanol stock solution, e.g., EMS, #15714, see Note 1) in PBS 1. Can be kept at 4  C for 1 week. 2.7

Immunostaining

1. Normal Goat Serum (NGS, Sigma #G9023) inactivated: Heat NGS stock solution at 56  C for 30 min; while still warm, filter the inactivated NGS through 0.22 μm using a filter and syringe; aliquot in sterile tubes. Store at 20  C. 2. Dimethylsulfoxide (DMSO, Sigma #D4540) 100% stock solution. 3. Blocking buffer PA for primary antibody: 10% NGS in 0.1% PBT + 1% DMSO. Prepare fresh. 4. Blocking buffer SA for secondary antibody: 10% NGS in 0.1% PBT. Prepare fresh. 5. Primary antibody: rabbit anti-phospho-histone H3 antibody (e.g., Abcam, #14955). Store at 20  C. 6. Secondary antibody: Goat anti-rabbit or donkey anti-rabbit Alexa 488 antibody. Store at 20  C. 7. 80% glycerol in 1 PBS final concentration.

3 3.1

Methods Animal Care

Nematostella culture, care and the induction of spawning is done according to previous publications [9–11]. Adult animals (Fig. 1) are kept at 17  C under dark conditions in glass bowls (~40 animal/ bowl/200 ml) and fed daily with freshly hatched Artemia nauplii. Change the water and clean glass bowls every 2 weeks. Induce spawning through a temperature (increase the temperature to 22  C) and light stimulus by placing the animals on a light table for 12 h. After spawning and fertilization let the embryos develop, metamorphose and raise Nematostella polyps in 1/3 ASW at 22  C until they reach the desired size for regeneration experiments (e.g., 4 tentacle juveniles, ~6 weeks post fertilization with a smashed Artemia feeding regime, Fig. 1 [5]).

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Fig. 1 General anatomy of adult and juvenile Nematostella vectensis. Photographs illustrating the adult (left) and juvenile (right) morphology of Nematostella; (a) Polyps are oriented toward the oral region to the top and aboral region to the bottom. Adult is on the left, juvenile is on the right. Red dotted lines indicate the amputation site. Ten, tentacles; pha, pharynx; m, mesenteries; asterisk indicates the location of the mouth (Figure and legend reproduced from ref. 5, under the terms and conditions of the Creative Commons by Attribution (CC-BY) license) 3.2 Induction of Regeneration

1. Coat all plastic equipment you will use (petri dishes, tips, pipettes. . .) with 0.1% PBTw to avoid that the animals stick to the plastic and carry out the experiment at room temperature. 2. Add the Nematostella polyps (~100 for the 4 tentacles juveniles and ~ 30 for the sub-adults or sexually mature adult polyps) into a petri dish in 40 ml of 1/3 ASW using a plastic pipette. 3. Place the petri dish on a light table to allow the Nematostella polyps to relax for 10–15 min and add 2 ml of relaxing solution (7.14% MgCl2) to the petri dish. Allow the Nematostella polyps to relax 10–15 more minutes.

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4. Place the petri dish with the relaxed Nematostella under a binocular macroscope. 5. Use a scalpel to cut the polyp below the pharynx, perpendicularly to the oral–aboral axis of the body (Fig. 1). This microdissection results in two Nematostella parts, the isolated oral region and the main body part. 6. Replace and rinse the MgCl2 of the used petri dish three times with 1/3 ASW. 7. Place the isolated Nematostella parts of interest in a newly coated petri dish in 40 ml 1/3 ASW and let the animals regenerate at 22C until the desired step [4, 5]. 3.3 In Vivo Analysis of Wound Healing and Pharynx Reformation

1. To study wound healing in vivo, use the compression assay developed in [5], (Fig. 2). This assay assesses the state of the opening at the amputation site following bisection on the fourtentacle juvenile polyps. It uses nematosomes (free circulating aggregates of cnidocytes) as a marker to follow the fluid dynamics present in the gastric cavity of Nematostella. (a) Use tweezers (e.g., Outils Rubis SA, #3C GRIP, Stabio, Switzerland) to compress laterally, along the oral–aboral axis, the relaxed and freshly bisected or regenerating juveniles.

Fig. 2 Wound healing/compression assay. (A) Diagram of the compression assay during regeneration. The purple dots represent the nematosomes. The red dotted line represents the amputation site. The forceps are laterally compressing the regenerating polyp body. (B) Time series of the compression assay in an opened (Ba–Bc) or a wound-closed (Bd–Bf) polyp. The dotted double arrow in (Ba) indicates the axial orientation of the animals shown in (Ba–Bf). O oral, AbO aboral (Figure and legend reproduced from ref. 5, under the terms and conditions of the Creative Commons by Attribution (CC-BY) license)

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Fig. 3 (A) Image of 488+ detection in the pharynx. Fed (a, a’), starved (b–c’), regenerating 72 hpa (d–e’) Nematostella polyp juveniles. DIC optic images (a–d). Epifluorescent images (a’–d’). The red dotted line labels the amputation site under the pharynx (c–d’). The green line in (c, c’) shows the 488+ fluorescence localized in the basal part of the pharynx. The area delimitated with the dotted line in d and d’ is the region where the 488 + reemerged in the polyp at 72 hpa. (e, e’) Confocal images at 72 hpa labeled for DNA (nuclei in cyan) on the 488-negative (e) and 488-positive (e’) polyp juvenile. The area delimitated with the white dotted line in (e’) (488+ regenerating polyps) shows the pharyngeal lip/pharynx in formation that is absent from the 488-negative regenerating polyps in which only the contact between the two mesenteries is visible. pha pharynx, m mesentery. Scale bar in (Aa’) is 20 μm and applies to (Aa-Ae, Ab’-Ae’) (Figure and legend reproduced from ref. 5, under the terms and conditions of the Creative Commons by Attribution (CC-BY) license)

(b) With an open wound, the nematosomes will be expelled at the amputation site. With a closed wound the nematosomes will either remain in the gastric cavity or leak out of the body cavity through the aboral pore. 2. To study pharynx formation, use the pharynx reformation assay develops in [5] (Fig. 3) that takes advantage of naturally occurring autofluorescence in the pharynx. (a) Mount living freshly bisected or regenerating fourtentacle juveniles under a slide and coverslip. Use clay at the four corners of the coverslip to avoid smashing the fragile tissues of the bisected juvenile. (b) Before any observation orient the juvenile polyp with the two mesenteries laterally by using a binocular and moving/pressuring the border of the coverslips with forceps. (c) Place the slide under a fluorescent microscope and observe the emergence of the endogenous 488 nm autofluorescence in the body cavity near the amputation site. The emergence of the 488 nm autofluorescence is correlated with the emergence of pharyngeal tissues [5].

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3.4 Tissue Morphology, Cellular Proliferation, Neotranscription, and Apoptosis Staining 3.4.1 Tissue Morphology

1. Place the petri dish on an illuminated surface (e.g., light table) to allow the Nematostella polyps to relax for 10–15 min and add 2 ml of relaxing solution (7.14% MgCl2) to the petri dish. Allow the Nematostella polyps to relax 10–15 more minutes. 2. When relaxed, collect the regenerating polyps and add them into a coated 1.5 or 2 ml Eppendorf tube. Remove as much liquid as possible before fixation. 3. Fix relaxed polyps (controls and regenerates) by adding the fixative IS in the Eppendorf tube. Fix animals for 1 h at room temp (RT) or overnight (ON) at 4  C. Then, wash the fixed polyps 3 times using 0.2% PBT (see Note 2). 4. Dilute Alexa Fluor® 488 Phalloidin stock solution to 1/200 and DAPI or Hoechst stock solution to 1/5000 in 0.1% PBTw. 5. In the Eppendorf tube, containing the bisected juvenile, remove as much liquid as possible and add the solution containing Alexa Fluor® 488 Phalloidin 1/200 and DAPI or Hoechst 1/5000. Let incubate ON at 4  C. 6. The next morning, rinse by performing 3 washes with 0.1% PBTw. 7. Remove as much liquid as possible and add 80% glycerol– PBS1. Let settle at least half a day, as the glycerol will partly clear the polyp tissue. 8. Mount the fixed and stained polyps between slide and coverslips using clay “feet” at its four corners. Press carefully to slightly compress the polyp and visualize properly the tissue detail under the microscope. 9. Use 488 nm and 405 nm filter under a fluorescent microscope to observe Phalloidin 488 and DAPI (or Hoechst), respectively.

3.4.2 Cellular Proliferation (EdU) and Neotranscription (EU)

1. Prepare 300 μM EdU (to address cell proliferation) or 1 mM EU (to address neotranscription) working solution in 1/3 ASW (see Note 3). 2. Collect the polyps (controls and regenerates) and place them into a 1.5 or 2 ml Eppendorf tube for the four-tentacle juveniles or in a 24-well plate for sub-adult or sexually mature adult polyps. 3. Remove as much liquid as possible and add the EdU or EU solution to the polyps. Let settle and incubate in the dark (e.g., wrapped foil around tube or plate) on a rocking table between 30 and 60 min. 4. Place the Eppendorf tube or the 24-well plate containing the polyps on an illuminated surface (e.g., light table) to allow the animals to relax for 10–15 min. Add a few drops of relaxing

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solution (7.14% MgCl2) to the tube or petri dish. Allow the polyps to relax for 10–15 more minutes. 5. When relaxed, remove as much liquid as possible before fixation. 6. Fix relaxed polyps by adding the fixative IS. Fix the animals for 1 h at room temp (RT) or over-night (ON) at 4  C (if working with the 24-well plate, do not forget to place the plate under a fume hood during the fixation). Wash the fixed polyps three times using 0.2% PBT (see Note 2). 7. Permeabilized the animal tissue by removing as much liquid as possible and add 0.5% PBT to the fixed polyps for 20–30 min on a rocking table. Nb. For sub-adult or sexually mature adult tissues, before developing the staining (see next steps), cut the fixed polyp in multiple identifiable parts to improve the efficiency/accessibility of the staining (see Note 5). In the dark, incubate the polyps for 30 min in the following Click-iT EdU/EU cocktail: For 100 μl use 75.8 μl deionized water; 10 μl 10 Click-iT Reaction buffer, 4 μl CuSO4 (100 mM stock solution); 0.25 μl Alexa Fluor Azide 488; 10 μl Reaction buffer additive. 8. Wash 3 times in PBS1. 9. At this step, the fixed and labeled polyps can also be stained with DAPI or Hoechst using the protocol described in Subheading 3.4.1 (see Note 6). 3.4.3 Apoptosis

1. Collect the polyps (controls and regenerates) and add them into a 1.5 or 2 ml Eppendorf tube for the four-tentacle juveniles or in a 24-well plate for the sub-adult or sexually mature adult polyps (see Note 4). 2. Place the Eppendorf tube or the 24-well plate containing the bisected polyp on an illuminated surface (e.g., light table) to allow the Nematostella polyps to relax for 10–15 min. Add a few drops of relaxing solution (7.14% MgCl2) to the tube or petri dish. Allow the polyps to relax for 10–15 more minutes. 3. When relaxed, remove as much liquid as possible before fixation. 4. Fix relaxed polyps by adding the fixative IS. Fix relaxed polyps by adding the fixative IS solution. Fix the animals for 1 h at room temp (RT) or over-night (ON) at 4  C (if working with the 24-well plate, do not forget to place the plate under a fume hood during the fixation). Wash the fixed polyps 3 times using PBT 0.2% (see Note 2).

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5. Permeabilize the animal tissue by removing as much liquid as possible and add Proteinase K 0.101 mg/ml in PBS1 for 20 min at RT (see Note 5). 6. Wash 3 times in PBS1. 7. To maintain tissue integrity, fix with secondary fixative for 1 h at RT. 8. Wash 5 times in PBS1. 9. Prepare 50 μl of TUNEL reaction mixture using 5 μl of TUNEL-enzyme solution and 45 μl of TUNEL-Label solution contained in the kit. 10. Remove as much liquid as possible and add 50 μl of TUNEL reaction mixture to the polyps and let incubate for 60 min at RT in the dark. 11. Wash 3 times in PBS1. 12. Observe Cell death using a 488 nm filter on a fluorescent microscope. The fixed and TUNEL stained polyps can also be costained at this step with Alexa Fluor® 594 Phalloidin and DAPI or Hoechst using the protocol described in Subheading 3.4.1. 3.4.4 Immunostaining

1. Perform Subheading 3.4.1, steps 1–3. 2. Permeabilize the animal tissue by removing as much liquid as possible and add 0.5% PBT to the fixed polyps for 20–30 min on a rocking table. 3. For sub-adult or sexually mature adult tissues, before developing the staining (next steps), cut the fixed polyp in multiple identifiable parts to improve the efficiency/accessibility of the staining (see Note 5). 4. Prepare blocking buffer PA for the primary antibody, remove as much liquid as possible and incubate the fixed polyps with the blocking buffer PA for 2–3 h on a rocking table at RT. 5. Prepare the primary Ab at the desired concentration diluted in the blocking buffer + DMSO 1%. Remove as much liquid as possible from the tube with the polyps and replace with the primary antibody solution. Incubate ON at 4  C on a rocking table. If a first purified primary Ab is tested, use it at 1/100. 6. Wash in 0.1% PBTw quickly two times, then, three times for 20–30 min each at RT. 7. Prepare the secondary Ab at the desired concentration diluted in the blocking buffer SA WITHOUT DMSO 1%. Remove as much liquid as possible and add the secondary Ab solution to the polyps. Incubate ON at 4  C on a rocking table. Usually the secondary antibody is used at 1/250–1/500.

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8. Wash in 0.1% PBTw quickly two times, then, three times for 20–30 min each at RT. 9. Mount the immunostained polyps in glycerol 80%–PBS1 under a slide and coverslip and observe the staining using the corresponding filters with a fluorescent microscope. The fixed and immunostained polyps can also be costained at this step with Alexa Fluor® 594 Phalloidin and DAPI or Hoechst using the protocol described in Subheading 3.4.1.

4

Notes 1. EdU, EU, DAPI, HOECHST, Phalloidin and Paraformaldehyde are CMR (Carcinogenic, Mutagenic and Reprotoxic) agents. Wear gloves when handling and discard properly according to health and safety rules. 2. When fixed, Nematostella polyp can be store for several weeks at 4  C in PBS1. 3. For negative EdU controls, at the Subheading 3.4.2, step 8 use the developing reaction WITHOUT adding the Click-iT Reaction buffer and/or Reaction buffer additive OR use 3–4 weeks starved Nematostella polyps, as their proliferative activity is strongly reduced in all tissues [3]. In latter condition, use the Reaction buffer and Reaction buffer additive as mentioned in Subheading 3.4.2, step 8. For positive EdU controls, use freshly fed but 2-days starved polyps, as feeding is a strong inducer of cell proliferation [3]. 4. For TUNEL negative control, at the Subheading 3.4.3, step 9 use only the 45 μl of TUNEL-Label solution contained in the kit WITHOUT adding the Enzyme-TUNEL solution. For TUNEL positive control, add an additional step between Subheading 3.4.3, step 8 and 9 by incubating the polyps with 0.1 U/μl TURBO DNase (Applied Biosystems/Ambion, Austin, TX) in 1 TURBO DNase buffer for 10 min prior to the TUNEL staining (see Subheading 3.4.3, step 8). 5. Optional—For adult tissue (thick tissues), to optimized the permeabilization and any labeling, cut the polyp in several pieces keeping a morphological characteristic in each of these pieces to orient your analysis after labeling. 6. According to the manufacturer, Phalloidin staining is not compatible with the Click-iT® detection reaction.

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References 1. Bely AE, Nyberg KG (2010 Mar) Evolution of animal regeneration: re-emergence of a field. Trends Ecol Evol (Amst) 25(3):161–170 2. Reitzel A, Burton P, Krone C, Finnerty J (2007) Comparison of developmental trajectories in the starlet sea anemone Nematostella vectensis: embryogenesis, regeneration, and two forms of asexual fission. Invertebr Biol 126(2):99–112 3. Passamaneck YJ, Martindale MQ (2012) Cell proliferation is necessary for the regeneration of oral structures in the anthozoan cnidarian Nematostella vectensis. BMC Dev Biol 12 (1):1–1 4. Bossert PE, Dunn MP, Thomsen GH (2013) A staging system for the regeneration of a polyp from the aboral physa of the anthozoan cnidarian Nematostella vectensis. Dev Dyn 242:1320–1331 5. Amiel AR, Johnston HT, Nedoncelle K, Warner JF, Ferreira S, Ro¨ttinger E (2015) Characterization of morphological and cellular events underlying oral regeneration in the sea anemone, Nematostella vectensis. Int J Mol Sci 16(12):28449–28471 6. Zapata F, Goetz FE, Smith SA, Howison M, Siebert S, Church SH et al (2015)

Phylogenomic analyses support traditional relationships within Cnidaria. PLoS One 10 (10):e0139068 7. Chang ES, Neuhof M, Rubinstein ND, Diamant A, Philippe H, Huchon D et al (2015) Genomic insights into the evolutionary origin of Myxozoa within Cnidaria. Proc Natl Acad Sci U S A 112(48):14912–14917 8. Warner JF, Guerlais V, Amiel AR, Johnston H, Nedoncelle K, Ro¨ttinger E (2018) NvERTx: a gene expression database to compare embryogenesis and regeneration in the sea anemone Nematostella vectensis. Development 145(10): dev162867 9. Hand C, Uhlinger KR (1992) The culture, sexual and asexual reproduction, and growth of the sea anemone Nematostella vectensis. Biol Bull 182(2):169–176 10. Fritzenwanker JH, Technau U (2002) Induction of gametogenesis in the basal cnidarian Nematostella vectensis (Anthozoa). Dev Genes Evol 212(2):99–103 11. Stefanik DJ, Friedman LE, Finnerty JR (2013) Collecting, rearing, spawning and inducing regeneration of the starlet sea anemone, Nematostella vectensis. Nat Protoc 8(5):916–923

Chapter 5 Staining and Tracking Methods for Studying Sponge Cell Dynamics Carole Borchiellini, Sandie M. Degnan, Emilie Le Goff, Caroline Rocher, Ame´lie Vernale, Stephen Baghdiguian, Nina Se´journe´, Florent Marschal, Andre´ Le Bivic, Nelly Godefroy, Bernard M. Degnan, and Emmanuelle Renard Abstract To better understand the origin of animal cell types, body plans, and other morphological features, further biological knowledge and understanding are needed from non-bilaterian phyla, namely, Placozoa, Ctenophora, and Porifera. This chapter describes recent cell staining approaches that have been developed in three phylogenetically distinct sponge species—the homoscleromorph Oscarella lobularis, and the demosponges Amphimedon queenslandica and Lycopodina hypogea—to enable analyses of cell death, proliferation, and migration. These methods allow for a more detailed understanding of cellular behaviors and fates, and morphogenetic processes in poriferans, building on current knowledge of sponge cell biology that relies chiefly on classical (static) histological observations. Key words Differentiation, Proliferation, Programmed cell death, Migration, Morphogenesis, Nutrition, Transdifferentiation, Juvenile, Porifera, Evo-devo

1

Introduction Most animals are classified as bilaterians, and possess a head, gut, central nervous system, and bilateral symmetry [1]. An understanding of the origin and evolution of these morphological features, and of the full girth of animal diversity, requires comparative evo-devo analyses on earlier-branching, non-bilaterian metazoan phyla, namely, Cnidaria, Placozoa, Ctenophora, and Porifera. Although the Cnidaria—sister group of Bilateria—has been studied in detail [1, 2], the other three phyla remain less explored. For example,

Carole Borchiellini, Sandie M. Degnan, Emilie Le Goff and Caroline Rocher equally contributed to this work. Nelly Godefroy, Bernard M. Degnan and Emmanuelle Renard equally supervised this work. David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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modern experimental tools such as transgenesis and CRISPR-Cas9 have yet to be developed in sponges, placozoans or ctenophores. Sponges (Porifera) are morphologically simple and were traditionally relegated to a separate subkingdom from the rest of the animal kingdom, the Parazoa (literally “beside animals”). The monophyletic Porifera is widely regarded as the likely sister group to all other metazoans [3–6], which means that comparative studies between sponges and other metazoans can provide valuable insights into early animal evolution. Currently, however, insights are constrained by the fact that many aspects of sponge biology are not well understood at physiological, cellular, or even morphological levels. The phylum consists of four classes (Hexactinellida, Demospongiae, Calcarea, and Homoscleromorpha) characterized by contrasting features at developmental, tissue, organizational and ecological levels. Even within the phylum, comparative studies among these four classes are required to determine which features are ancestral [7]. The gene content of sponges has been characterized by transcriptomic or genomic approaches. These gene inventories and analyses have yielded two surprising results: (1) despite their simplicity, sponges contain most of the developmental gene families present in bilaterians; and (2) inconsistencies between gene content and morphological features make it difficult to transpose gene functions described in bilaterians directly to sponges. Therefore, one of the next challenges for biologists is to understand how similar genomic content, structure and organization yield such markedly different body plan organization, dynamics, and life histories. Given their relatively simple body organization, sponges provide unique opportunities for understanding how genomes instruct the formation of different body plans. To address this challenge, four main axes of research are currently ongoing: (a) the sequencing, assembly, annotation and analysis of more poriferan genomes [7]; (b) the analysis of gene expression patterns and profiles [8, 9] (Fierro-Constaı´n et al., in this volume); (c) the development of reproducible knock-down and knock-in transfection protocols [10–12]; and (d) the reinterpretation of sponge cell structures, behaviors, and mechanisms with advanced cell analysis tools. This chapter focuses on the fourth aspect. Present knowledge of sponge cell biology primarily relies on classical (static) observations, which fail to reveal dynamic cellular interactions, movements, and decisions that are essential for deciphering morphogenetic processes. Here we present methods for evaluating primordial cellular processes in three sponge species with contrasting body plans and life histories: Amphimedon queenslandica (class Demospongiae: order Haplosclerida: family Niphatidae) [13, 14], Lycopodina hypogea (class Demospongiae: order Poecilosclerida: family Cladorhizidae) [15] and Oscarella lobularis (class Homoscleromorpha, family Oscarellidae) [16–19]. Protocols for three cellular processes are described: (a) cell death, using TUNEL staining to detect apoptosis and MAPLC3 staining to

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detect autophagy; (b) cell proliferation, using phosphohistone H3 antibody labeling to detect mitotically active cells in fixed samples, and 5-ethynyl-20 -deoxyuridine (EdU) stain to achieve the same in vivo; and (c) cell fate and migration, using cell dyes such as DiI, Lectin kits, FluoSpheres, and live imaging. These protocols provide a means to understand some of the cellular mechanisms underlying sponge morphogenesis and the so-called lability of sponge cells [20].

2

Materials

2.1 Biological Material 2.1.1 Amphimedon queenslandica Biological Samples

1. Amphimedon queenslandica inhabits shallow Indo-Pacific coral reef habitats often attached to decaying coral rubble. This species can be collected by breaking dead coral rubble surrounding the sponge; a hammer and chisel are handy. The sponge is best transported and maintained in aquaria while still attached to the rubble. 2. A. queenslandica embryos, larvae and juveniles can be produced in the laboratory year-round. An individual A. queenslandica will produce larvae for about 4 months when maintained in a closed aquarium system at 25  C and fed daily with a mixture of dead phytoplankton. In our system, we feed suspension-feeding marine invertebrates an equal proportion of the following marine microalgal concentrates from Reed Mariculture: Nanno3600 (CCMP525/Nannochloropsis sp.); Isochrysis1800 (CCMP1324/T-Isochrysis sp.); Pavlova1800 (CCMP359/Pavlova sp.); and TW1200 (CCMP1051/Thalassiosira weissflogii). 3. The coralline alga Amphiroa fragilissima induces high rates of A. queenslandica larval settlement and metamorphosis. We collect A. fragilissima at the same time as A. queenslandica. It can also be maintained in a recirculating closed aquarium system for more than 4 months.

2.1.2 Lycopodina hypogea Biological Samples

1. Lycopodina hypogea was first observed in a shallow marine cave in La Ciotat (France) in 1992 at around 20 m deep. L. hypogea biotope is equivalent to seabed depths of about 1000 m: constant temperature of 13–14  C, no light, no movement. L. hypogea was the first carnivorous sponge described in 1995 [21]. 2. L. hypogea possesses a basal peduncle of 1 or 2 cm attached to rocky substrate and an apical ovoid head with long thin filaments covered with spicules that are used to capture prey. 3. L. hypogea does not possess any choanocytes or an aquiferous system.

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4. L. hypogea is maintained in the laboratory and fed Artemia larvae; most studies on this sponge have focused on feeding mechanisms and nutrition [21–24]. Other aspects of its biology (e.g., development and reproduction) are less well explored. 2.1.3 Oscarella lobularis Biological Samples

1. Oscarella lobularis adults are commonly found on the rocky and coralligenous substrates of the north-western Mediterranean Sea. O. lobularis is the type species of the family Oscarellidae (see Note 1). Its phylogenetic position [16–19], histology, embryology, and ecology [25, 26] are well known. 2. This species is capable of both sexual and asexual reproduction. Asexual reproduction by budding can be triggered in the laboratory. The resultant asexual buds can be maintained in Petri dishes at 17  C for several months [27]. The following protocols were developed on buds because of their abundance and transparency.

2.2 Antibody-Based Staining on Whole Mount Biological Samples (See Note 2) 2.2.1 A. queenslandica

1. 0.2 μm filtered sea water (FSW) 2. Saline buffer stock solutions: (A) MOPS Buffer: 100 mM MOPS, 500 mM NaCl, 2 mM MgSO4; (B) Phosphate buffer saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7. 3. Fixative stock solutions: (A) 25% glutaraldehyde; (B) 16% paraformaldehyde (PFA). 4. 70% ethanol. 5. Detergent: Tween 20. 6. 10% bovine serum albumin (BSA). 7. 2 mg/mL fluorescent dye 40 ,6-diamidino-2-phenylindole (DAPI) for counterstaining (Thermo Fisher Scientific). 8. ProLong™ Scientific).

Gold

Antifade

Mountant

(Thermo

Fisher

9. Glass slides or chamber glass slide (Ibidi) and coverslips. 10. For Immunofluorescent staining of phosphohistone H3 (PH3): (a) Primary antibodies against phospho-histone H3 [pSer10] (Abcam ab5176 - rabbit, 1:500). (b) Secondary antibodies: Alexa Fluor 488, 568, or 647 goat anti-rabbit IgG (Thermo Fisher Scientific). 2.2.2 L. hypogea

1. Natural sea water (NSW). 2. Saline buffer stock solution: Tris-Buffered Saline (TBS): 150 mM NaCl, 25 mM Tris, pH 7.5. 3. Fixative stock solution: 32% PFA.

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4. Detergent: 10% Triton-X-100. 5. 10% BSA in TBS. 6. 1 mg/mL DAPI for counterstaining (Sigma-Aldrich). 7. Glass slides or chamber glass slide (Ibidi) and coverslips. 8. Milli-Q H2O. 9. Fluorescence mounting medium (Dako). 10. For Immunofluorescent staining of phosphohistone H3 (PH3): (a) 10% BSA. (b) Mouse monoclonal anti-PH3 antibody (MA5-15220 from Invitrogen). (c) Donkey TRITC secondary antibody against mouse (Jackson ImmunoResearch). 11. For TUNEL assays: (a) In situ Cell Death Detection Kit, Fluorescein or TMR red (Roche). (b) 1 mg/mL DNase I recombinant (Sigma Aldrich). 12. For immunofluorescent staining of MAPLC3 assays: (a) Anti-MAPLC3 polyclonal antibody (sc-28266 Santa Cruz biotechnology). (b) Mouse secondary antibody against rabbit–FITC (Jackson ImmunoResearch). 2.2.3 O. lobularis

1. Saline buffer stock solutions: for TUNEL only. 2. Fixative: 16% PFA. 3. Detergents: 10% Saponin; 10% Triton X-100 for TUNEL only. 4. 10% Blocking reagent (BR) (Roche) in maleic acid buffer (MAB): 100 mM maleic acid, 150 mM NaCl, pH 7.5; or 10% BSA for TUNEL only. 5. Nucleic acid staining: 2 mg/mL DAPI for counterstaining (Thermo Fisher Scientific), or 10 mg/mL Hoechst 33342 (for EdU only). 6. Mountant: ProLong™ Diamond Antifade Mountant or ProLong™ Gold Antifade Mountant (Thermo Fisher Scientific), or Dako fluorescence mounting medium (Dako) (for TUNEL only). 7. Glass slides or chamber glass slide (Ibidi) and coverslips. 8. For Immunofluorescent staining of phosphohistone H3 (PH3).

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(a) Antibodies: Anti-PH3 primary antibody Abcam ab5176 (rabbit); Alexa Fluor 488 goat anti-rabbit IgG secondary antibody (Thermo Fisher Scientific). (b) Actin staining: Phalloidin-Alexa 647 (Santa Cruz Biotechnology or Thermo Fisher Scientific). 9. In addition, for TUNEL assays. (a) FSW to dilute PFA stock. (b) In situ Cell Death Detection Kit, Fluorescein or TMR red (Roche). (c) 1 mg/mL recombinant DNase I (Sigma-Aldrich). 2.3 EDU Incorporation Assays

2.4 Live Cell Staining and Imaging Assays in O. lobularis and A. queenslandica

Click-iT™ or Click-iT™ Plus EdU Alexa Fluor™ 488 Imaging Kit (Thermo Fisher Scientific) in addition to reagents and materials listed above in Subheading 2.2 (see Note 3). 1. CellTracker™ CM-DiI Dye (Molecular Probes) 2. Lectin kits (I, II and III)—fluorescein or rhodamine labeled (Vector laboratories). 3. FluoSpheres™ Carboxylate-Modified Microspheres, 0.2 μm (Thermo Fisher Scientific). 4. DMSO. 5. MOPS Buffer: 100 mM MOPS, 500 mM NaCl, 2 mM MgSO4. 6. Tween 20. 7. DAPI (2 mg/mL) for counterstaining (Thermo Fisher Scientific).

3

Methods

3.1 Obtaining Amphimedon queenslandica Juveniles

1. Place adults in a dark-colored aquarium or tub in ambient sea water [21–27  C]. 2. Induce the emergence of A. queenslandica larvae from adults by increasing the ambient sea water temperature by 1–2  C over 1 h using either solar energy or electric aquarium heating bar [27]. 3. Collect swimming larvae with a hand or mouth pipette; larvae are white in color, >500 μm, and can be seen by eye. 4. Transfer swimming larvae into a 2 L beaker of NSW. Allow larvae to develop for at least 6 h under ambient temperature and light conditions until a majority are competent to settle and metamorphose [28].

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5. Place 8–10 competent larvae in a well of a sterile 6-well plastic plates with 10 mL of FSW and pieces of live coralline algae Amphiroa fragilissima covering 30–50% of the bottom of the well; A. fragilissima strongly induces A. queenslandica settlement and metamorphosis. 6. Leave the 6-well plates in the dark for about 4 h. 7. Remove postlarvae that have settled onto the A. fragilissima using fine forceps (e.g., Dumont #5); transfer immediately onto round coverslips placed in the bottom of a sterile 24-well plastic plate with 2 mL of FSW; place three postlarvae on each coverslip. The translocated postlarvae tend to initially ball up and look similar to a newly settled larva. 8. Incubate the plates at 25  C in the dark until metamorphosis is complete and filter feeding juveniles with a pronounced osculum are observed; time to metamorphosis is variable but typically takes 3–4 days. About 50% of the post larvae transferred onto coverslips will develop into filter feeding juveniles, compared to 80–90% of postlarvae reared continuously on A. fragilissima [29]. 3.2 Stalling and Nutrition of Lycopodina hypogea Adults 3.3 Obtaining Oscarella lobularis Buds [27]

1. Place individuals in aquarium of NSW at 13  C (see Note 4). 2. Renew NSW once a month. 3. Feed them every week or less (they can stay unfed for several months) with Artemia franciscana nauplii. 1. Cut sampled adults into fragments (about 1 cm3). 2. Place each fragment in a well of a 6-well culture plates in 8 mL of NSW at 17  C. 3. Renew NSW each day until the formation of free buds (between 10 and 15 days depending the sample). 4. Transfer free buds in Petri dishes (14 cm diameter) filled with NSW and maintain at 17  C (see Note 5). 5. Renew NSW once a week.

3.4 Detection of Phosphohistone 3 (PH3) by Immunofluorescence (See Note 6) 3.4.1 A. queenslandica

All steps are performed on juveniles attached to the coverslips they settled upon (see above). Juveniles are fixed according to Larroux et al. [30]. Immunofluorescence is undertaken following the protocol described in Nakanishi et al. [29]. 1. Remove FSW from wells containing juveniles and add 0.5 mL of fixative (4% PFA and 0.05% glutaraldehyde in MOPS buffer). Gently swirl the plate for 5 s and immediately replace with 1 mL of fresh fixative. Incubate on a slow rocker at room temperature for 60 min.

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2. Exchange half of the fixative with 70% ethanol. Wash for 5 min. Repeat three times with the last wash being for 10 min. At this point, fixed juveniles can be stored at 20  C. 3. Rehydrate fixed juveniles by washing twice in 1 mL of PBST (PBS + 0.1% Tween 20), each time for 5 min. Block with 1 mL 1% BSA in PBST for 1 h. Replace the 1% BSA in PBST with 0.4 mL 1% BSA in PBST that includes 2 μL anti-PH3 [pSer10] as the primary antibody; incubate juveniles overnight at 4  C. 4. Next day, repeat washes in PBST and blocking in 1% BSA in PBST as described above. Replace the 1% BSA in PBST with 0.4 mL 1% BSA in PBST that includes 2 μL Alexa Fluor 488, 568, or 647 goat anti-rabbit IgG as the secondary antibody; incubate juveniles overnight at 4  C and wash in PBST the next day as described above. 5. Label juvenile sponge nuclei with DAPI (1:1.000) for 30 min. Wash in PBST for 5 min prior to mounting on a slide using Prolong Gold antifade reagent. 6. Observe and record data using confocal microscopy. 7. Analyze images using ImageJ. 3.4.2 L. Hypogea

1. Prepare 3.7% PFA in NSW. 2. Transfer individuals from aquarium to chamber glass slide. 3. Discard NSW. 4. Fix samples in 500 μL of 3.7% PFA. 5. Incubate for 30 min at room temperature (RT). 6. Wash 3 times with 1 mL TBS with 1% BSA. 7. Incubate samples in 0.2% Triton in TBS with 1% BSA (20 min; RT). 8. Wash 3 times with 1 mL TBS with 1% BSA. 9. Prepare a solution containing anti-PH3 mouse antibody (1:200) diluted in TBS with 1% BSA. 10. Incubate for 2 h at 37  C in humid chamber. 11. Wash 3 times with 1 mL TBS with 1% BSA. 12. Dilute donkey secondary antibody against mouse TRITC (2:100) in a solution containing DAPI (1:1000) in TBS with 1% BSA. 13. Incubate for 1 h at 37  C in humid chamber. 14. Wash 2 times with 1 mL TBS with 1% BSA. 15. Wash 2 times with 1 mL TBS. 16. Wash quickly 1 time in Milli-Q H2O. 17. Discard Milli-Q H2O.

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18. Add 15 μL of mounting medium. 19. Discard silicon chamber structure. 20. Cover with a 22  22 coverslip. 21. Store at 4  C without light. 22. Observe with confocal microscope. 3.4.3 O. lobularis

1. Prepare 3% PFA in PBS. 2. Transfer buds from Petri dishes where they are cultured to 2 mL microtube with a glass Pasteur pipette. 3. Discard NSW. 4. Put 1 mL of 3% PFA on the buds. 5. Incubate at 4  C overnight. 6. Rinse 1 time with 2 mL PBS at RT. 7. Wash 3 times for 5 min in 2 mL PBS at RT. 8. Prepare blocking solution: 1 mL composed of 2% Blocking reagent (BR) and 0.1% saponin in PBS. 9. Incubate buds in the blocking solution (30 min to 1 h) at RT. 10. Incubate buds in anti-PH3 antibody diluted in blocking solution (1:500) overnight at 4  C. 11. Rinse with PBS at RT. 12. Wash 3 times for 5 min in 2 mL PBS at RT. 13. Dilute alexa-labeled goat secondary antibody against mouse IgG (1:500) in blocking solution. 14. Incubate buds in this solution for 1–2 h at RT, in obscurity. 15. Add DAPI (1:500) and phalloidin (1:1000) and incubate at RT for 15 min in the dark. 16. Rinse with PBS in the dark at RT. 17. Wash 2 times for 5 min in 2 mL PBS at RT. 18. Place buds on a slide with a glass Pasteur pipette and discard exceeding PBS (by aspiration and absorbing with a tiny piece of absorbent paper if necessary). 19. Add 15 μL of mounting medium. 20. Cover with a 22  22 coverslip. 21. Allow to polymerize for 40 min at 37  C in the dark or at room temperature overnight. 22. Observe with confocal microscopy (Leica LSM 510 or Leica 780 or Leica 880). Image analysis is performed using ImageJ.

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3.5 EdU Incorporation Assays 3.5.1 A. queenslandica

1. Culture postlarvae and juveniles in 2 mL FSW in 24-well plates as described in Subheading 3.1. 2. Add EdU to the 2 mL of FSW to a final concentration of 200 μM and incubate postlarvae and juveniles for a period of time, typically 6 h, to label S-phase nuclei. 3. Wash labeled specimens in FSW and immediately add 0.5 mL of fixative (4% PFA and 0.05% glutaraldehyde in MOPS buffer). Gently swirl the plate for 5 s and immediately replace with 1 mL of fresh fixative. Incubate on a slow rocker at room temperature for 60 min. 4. Conduct fluorescent labeling of incorporated EdU according to the manufacturer’s recommendations. 5. DAPI labeling, mounting, and microscopy are as described in Subheading 3.4.

3.5.2 L. hypogea

1. Prepare stock solutions according to the manufacturer’s instructions. 2. Transfer individuals from aquarium to chamber glass slide. 3. Incubate for 24 h at 13  C with EdU 50 μM in 500 μL of NSW. 4. Wash 3 times with 1 mL NSW. 5. Samples are fixed in 500 μL of 3.7% PFA in NSW. 6. Incubate for 30 min at RT. 7. Wash 3 times with 1 mL TBS with 3% BSA. 8. Incubate samples for 20 min at RT in the Triton 0.2% in TBS with 3% BSA. 9. Wash 3 times with 1 mL TBS with 3% BSA. 10. Incubate in 500 μL of revelation mix as manufacturer’s instructions added with DAPI (1:1000) during 30 min at RT without light. 11. Wash 2 times with 1 mL TBS with 3% BSA. 12. Wash 2 times with 1 mL TBS. 13. Wash quickly 1 time in Milli-Q H2O. 14. Discard Milli-Q H2O. 15. Add 15 μL of mounting medium. 16. Discard silicon chamber structure. 17. Cover with a 22  22 coverslip. 18. Store at 4  C without light. 19. Observe with confocal microscope.

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1. Prepare a 50 μM EdU solution in FSW in 15 mL Falcon Vials and vortex the solution. 2. Use a glass Pasteur pipette to transfer buds in a 24 well-culture plate and carefully remove water overflow. 3. Dispense 1 mL of 50 μM EdU solution on each well (negative control: a well filled with 1 mL FSW). 4. Incubate for 6 or 24 h at 17  C. 5. Prepare 3% PFA in PBS. 6. Fix treated buds in 3% PFA over night at 4  C. 7. Carefully remove the PFA from the vial with a Pasteur pipette and add 1 mL of PBS. Incubate for 15 min at RT and repeat operation twice again. 8. Prepare blocking solution composed of 0.1% Blocking reagent (BR) and 0.1% saponin in PBS: after removal of PBS from each vial, dispense 1 mL of this solution per vial. 9. Incubate for 1 h at RT. 10. Prepare the Alexa Fluor azide solution according to manufacturer’s instruction, maximum 15 min before use. 11. Remove blocking solution and add 250 μL of Alexa Fluor azide solution. 12. Protect vial from the light and incubate for 30 min at RT. 13. Prepare 1/2000e Hoechst solution in PBS. 14. Wash Alexa Fluor azide solution with 1 mL PBS. 15. Remove PBS and add 1 mL Hoechst solution. 16. Protect vial from the light and incubate for 30 min at RT. 17. Remove Hoechst solution and finally add 1 mL PBS. 18. Buds are immediately mounted with ProLong® Antifade Mountant.

3.6 TUNEL Assays (See Note 7) 3.6.1 O. lobularis and L. hypogea

1. Prepare PFA 3.7% in NSW. 2. Use a glass Pasteur pipette to transfer buds in a 24 well-culture plate and carefully remove water overflow (for O. lobularis) or transfer individuals from aquarium to chamber glass slide Ibidi (for L. hypogea). 3. Discard NSW. 4. Fix samples in 500 μL of 3.7% PFA. 5. Incubate for 30 min at RT. 6. Wash 3 times with 1 mL TBS with 1% BSA. 7. Incubate samples in the Triton 0.2% in TBS with 1% BSA (20 min at RT). 8. Wash 3 times with 1 mL TBS with 1% BSA.

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9. For positive control: put one sample with DNase treatment 1 mg/mL in Milli-Q H2O for 30 min to 1 h at RT. 10. Prepare a labeling solution according to the manufacturer’s instructions added with DAPI (1:1000). 11. Incubate for 1 h at 37  C in humid chamber. 12. Wash 2 times with 1 mL TBS with 1% BSA. 13. Wash 2 times with 1 mL TBS. 14. Wash quickly 1 time in Milli-Q H2O. 15. Discard Milli-Q H2O. 16. Add 15 μL of mounting medium. 17. Discard silicon chamber structure (for L. hypogea). 18. Cover with a 22  22 coverslip. 19. Store at 4  C without light. 20. Observe with confocal microscope. 3.7 MAPLC3 Staining in L. hypogea (See Note 8)

1. Prepare PFA 3.7% in NSW. 2. Transfer individuals from aquarium to chamber glass slide Ibidi. 3. Discard NSW. 4. Fix samples in 500 μL of 3.7% PFA. 5. Incubate for 30 min at RT. 6. Wash 3 times with 1 mL TBS with 1% BSA. 7. Incubate samples 20 min at RT in TBS with 0.2% Triton and 1% BSA. 8. Wash 3 times with 1 mL TBS with 1% BSA. 9. Prepare a solution containing (2:100) MAPLC3 rabbit antibody diluted in TBS with 1% BSA. 10. Incubate for 2 h at 37  C in humid chamber. 11. Wash 3 times with 1 mL TBS with 1% BSA. 12. Dilute mouse secondary antibody against rabbit FITC (2:100) in a solution containing DAPI (1:1000) in TBS with 1% BSA. 13. Incubate for 1 h at 37  C in humid chamber. 14. Wash 2 times with 1 mL TBS with 1% BSA. 15. Wash 2 times with 1 mL TBS. 16. Wash quickly once in Milli-Q H2O. 17. Discard Milli-Q H2O. 18. Add 15 μL of mounting medium. 19. Discard silicon chamber structure. 20. Cover with a 22  22 coverslip.

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21. Store at 4  C without light. 22. Observe with confocal microscope. 3.8 Staining Choanocytes with the Lipidic Marker CMDiI Dye 3.8.1 A. queenslandica

1. Incubate juveniles in 1 μM CM-DiI in FSW for 30 min to 1 h, and then wash for 5 min in FSW. These incubation conditions should minimize the labeling of cells other than choanocytes. 2. Inspect CM-DiI labeled juveniles by epifluorescence microscopy immediately after washing out the CM-DiI to determine if nonchoanocyte cells are labeled; discard any juveniles that show any CM-DiI labeled cells outside of choanocyte chambers. 3. Develop remaining juveniles in FSW for a period of time (e.g., 2, 6, or 24 h postincubation (hpi) in CM-DiI), then wash in FSW three times for 5 min. 4. Remove FSW from wells containing juveniles and add 0.5 mL of fixative (4% PFA and 0.05% glutaraldehyde in MOPS buffer). Gently swirl the plate for 5 s and immediately replace with 1 mL of fresh fixative. Incubate on a slow rocker at room temperature for 60 min. 5. Wash fixed juveniles three times in MOPST (MOPS buffer +0.1% Tween). 6. DAPI labeling, mounting, and microscopy are as described in Subheading 3.4.

3.8.2 O. lobularis

1. Prepare the stock solution of 1 mM CM-Dil in DMSO. 2. Place Oscarella buds into 1 mL of FSW (either in microtubes or in 24 well culture plates). 3. Add 2 μL CM-DiI (1:500 dilution). 4. Incubate at 17  C overnight. 5. Rinse 3 times with FSW (thoroughly mix with a glass Pasteur pipette). 6. Wash once for 5 min with FSW and put the buds in fresh FSW before following experiments or fixation.

3.9 Staining O. lobularis Choanocytes with Fluorescent Microspheres (See Note 9)

1. Place O. lobularis buds into 5 mL of NSW (in a 6 well-culture plate). 2. Add 0.5 μL of FluoSpheres (1:10000 dilution). 3. Incubate at 17  C at least 1 h in the dark. 4. Rinse 3 times with NSW and thoroughly mix with a glass Pasteur pipette to remove the spheres glued to the surface. 5. Put the buds in fresh NSW before following experiments or fixation.

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3.10 Staining O. lobularis Choanocytes with Fluorescent Lectins (See Note 9)

1. Place O. lobularis buds into 1 mL of NSW (either in microtubes or in 24 well culture plates). 2. Add 2 μL of fluorescein labeled- Phaseolus vulgaris erythroagglutinin (1: 500 dilution). 3. Incubate at 17  C overnight in the dark. 4. Rinse 2 times with fresh NSW before following experiments or fixation.

3.11 Live Imaging in White Light 3.11.1

A. queenslandica

Live imaging was developed by creating a stable environment that allowed for visualization of cells in transparent A. queenslandica juveniles for multiple hours to days. Cells and structures are best visualized on the edge of the juvenile by light microscopy using high numerical aperture (NA) objectives with Nomarski techniques (differential interference contrast, DIC). Video-enhanced DIC is capable of resolving intracellular structures as small as 20 nm by means of digitally enhancing image contrast and detecting object edge movements [31, 32]. The resolution potential was increased further by positioning an optical short-pass filter (450 nm) in the epi-illumination system before the condenser. 1. Transfer juveniles cultured on coverslips into a perfusion chamber (Warner Instruments, Model RC-30WA), which is mounted on a Nikon TiE inverted microscope comprised of a set of Plan Achromat objectives (10–60), Plan Apochromat TIRF 1.49 NA 100 objective, 1.5 intermediate magnification adjustment, two condensers (0.52 and 0.85 NA), and a full set of Nomarski prisms, including the Nikon high resolution version for the 100 objective. 2. Acquire images using a Nikon DS-Qi1 16-bit monochrome camera, complemented by a Canon EOS 5d Mark iii for larger field-of-view images where appropriate. The Nikon DS-Qi1 camera allows for low-light visualization. Control both the Nikon camera and multiple elements of the microscope using NIS-Elements AR software (Nikon) on an associated desktop computer. 3. Equip microscope with a precision XY motorized stage and motorized computer-controlled Z adjustment. Z-series are analyzed by Zerene Stacker software (Zerene Systems) implementing the PMax algorithm for image analysis, and on occasion for generating 3d-imitating images and stereo pairs (Longson et al. 2010). 4. Process raw images first in NIS-Elements AR for levels adjustment and file-format conversion, with final adjustments made in ImageJ and Adobe Photoshop CC. Use batch function in Photoshop CC to correct alignment errors introduced by Zerene Stacker for the long sequences needed to generate video

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files. Generate videos from processed image sequences using Sony Vegas 12 software with uncompressed avi output and compress with online stream-ready algorithms (H.264) of Handbrake software. All videos have a fps value of 23.976.

4

Notes 1. Oscarella lobularis can be morphologically confused with other Oscarella species, particularly its sister species O. tuberculata (with different cellular features); taxonomic expertise is needed. 2. Direct light should be avoided or minimized for all fluorescent labeled molecules; they have to be stored in the dark. 3. Phalloidin staining is not compatible with click-it EDU kit staining. 4. L. hypogea are maintained in a controlled chamber to maintain a good temperature and hygrometry. 5. O. lobularis buds are easier to aspirate and manipulate with glass Pasteur pipette because they tend to stick in plastic pipettes or tips. 6. For L. hypogea and O. lobularis, EdU is more efficient than PH3 staining. 7. For TUNEL positive control, DNase is very volatile. Be careful to not incubate in the same incubator used for TUNEL tests. 8. For L. hypogea, Z-VAD-fmk (V116 Sigma Aldrich) treatment might be used at 10 μM to inhibit caspases during apoptosis. 9. To make sure that all choanocyte chambers of O. lobularis buds are stained, check staining with fluorescent microscopy and do not reduce incubation time under 1 h for microspheres and under 10 h for PhaE lectin to avoid incomplete staining of choanocyte chambers. In contrast, prolonging incubation with CMDil can lead to extraspecific staining. Staining choanocytes with PhaE is cleaner and more specific than with CMDiI.

Acknowledgments All the authors acknowledge the Centre National de la recherche Scientifique (CNRS) for funding the project for international scientific cooperation (PICS) STraS, supporting their collaboration since January 2018. The IMBE and IBDM researchers thank the Amidex foundation for funding the Spongex project (n ANR-11IDEX-0001-02) and the imaging facilities of the France

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Bioimaging infrastructure and in particular Cedric Matthews for his help and advice as to developing imaging in O. lobularis buds. We also acknowledge the diving facilities of the Institute OSU Pytheas (Laurent Vanbostal) and divers from the IMBE lab (Christian Marschal, Thierry Perez, Virgile Calvert, and Sandrine Chenesseau) for collecting O. lobularis and L. hypogea. The ISEM researchers thank Jean Vacelet for his help and advices for handling and stalling L. hypogea, Camille Martinand-Mari for having initiated cell staining in L. hypogea and the Montpellier RIO imaging platform. SMD and BMD are supported by grants from the Australian Research Council, and acknowledge the assistance and efforts of Nagayasu Nakanishi, Shunsuke Sogabe, and Daniel Stoupin in developing the techniques for A. queenslandica. References 1. Gilbert SF (2010) Developmental biology, 9th edn. Sinauer Associates, Inc., Sunderland, MA 2. Genikhovich G, Technau U (2017) On the evolution of bilaterality. Development 144:3392–3404 3. Dunn CW (2017) Ctenophore trees. Nat Ecol Evol 1:1600 4. Pett W, Adamski M, Adamska M, Francis WR, Eitel M, Pisani D, Wo¨rheide G (2019) The role of homology and orthology in the phylogenomic analysis of metazoan gene content. Mol Biol Evol 36:643–649 5. Simion P, Philippe H, Baurain D, Jager M, Richter DJ, Di Franco A, Roure B, Satoh N, Que´innec E´, Ereskovsky A et al (2017) A large and consistent phylogenomic dataset supports sponges as the sister group to all other animals. Curr Biol 27:958–967 6. Whelan NV, Kocot KM, Moroz TP, Mukherjee K, Williams P, Paulay G, Moroz LL, Halanych KM (2017) Ctenophore relationships and their placement as the sister group to all other animals. Nat Ecol Evol 1:1737–1746 7. Renard E, Leys SP, Wo¨rheide G, Borchiellini C (2018) Understanding animal evolution: the added value of sponge transcriptomics and genomics. BioEssays 40:e1700237 8. Sebe´-Pedro´s A, Chomsky E, Pang K, LaraAstiaso D, Gaiti F, Mukamel Z, Amit I, Hejnol A, Degnan BM, Tanay A (2018) Early metazoan cell type diversity and the evolution of multicellular gene regulation. Nat Ecol Evol 2:1176–1188 9. Sogabe S, Hatleberg WL, Kocot KM, Say T, Stoupin D, Roper KE, Fernandez-Valverde SL, Degnan SM, Degnan BM (2019) Pluripotency

and the origin of animal multicellularity. Nature 570:519–522 10. Hall C, Rodriguez M, Garcia J, Posfai D, DuMez R, Wictor E, Quintero OA, Hill MS, Rivera AS, Hill AL (2019) Secreted frizzled related protein is a target of PaxB and plays a role in aquiferous system development in the freshwater sponge, Ephydatia muelleri. PLoS One 14:e0212005 11. Rivera A, Winters I, Rued A, Ding S, Posfai D, Cieniewicz B, Cameron K, Gentile L, Hill A (2013) The evolution and function of the Pax/six regulatory network in sponges. Evol Dev 15:186–196 12. Rivera AS, Hammel JU, Haen KM, Danka ES, Cieniewicz B, Winters IP, Posfai D, Wo¨rheide G, Lavrov DV, Knight SW et al (2011) RNA interference in marine and freshwater sponges: actin knockdown in Tethya wilhelma and Ephydatia muelleri by ingested dsRNA expressing bacteria. BMC Biotechnol 11:67 13. Hooper JNA, van Soest RWM (2006) A new species of Amphimedon (Porifera, Demospongiae, Haplosclerida, Niphatidae) from the Capricorn-Bunker Group of Islands, Great Barrier Reef, Australia: target species for the “sponge genome project.” Zootaxa 1314 14. Leys SP, Larroux C, Gauthier M, Adamska M, Fahey B, Richards GS, Degnan SM, Degnan BM (2008) Isolation of Amphimedon developmental material. CSH Protoc 2008:pdb. prot5095 15. Hestetun JT, Vacelet J, Boury-Esnault N, Borchiellini C, Kelly M, Rı´os P, Cristobo J, Rapp HT (2016) The systematics of carnivorous sponges. Mol Phylogenet Evol 94:327–345

Tracking Sponge Cell Dynamics 16. Gazave E, Lape´bie P, Renard E, Vacelet J, Rocher C, Ereskovsky AV, Lavrov DV, Borchiellini C (2010) Molecular phylogeny restores the supra-generic subdivision of Homoscleromorph sponges (Porifera, Homoscleromorpha). PLoS One 5:e14290 17. Gazave E, Lape´bie P, Ereskovsky AV, Vacelet J, Renard E, Ca´rdenas P, Borchiellini C (2012) No longer Demospongiae: Homoscleromorpha formal nomination as a fourth class of Porifera. Hydrobiologia 687:3–10 18. Gazave E, Lavrov DV, Cabrol J, Renard E, Rocher C, Vacelet J, Adamska M, Borchiellini C, Ereskovsky AV (2013) Systematics and molecular phylogeny of the family Oscarellidae (Homoscleromorpha) with description of two new Oscarella species. PLoS One 8:e63976 19. Boury-Esnault N, Sole-Cava AM, Thorpe JP (1992) Genetic and cytological divergence between colour morphs of the Mediterranean sponge Oscarella lobularis Schmidt (Porifera, Demospongiae, Oscarellidae). J Nat Hist 26:271–284 20. Simpson TL (1984) The cell biology of sponges. Springer-Verlag, New York 21. Vacelet J, Boury-Esnault N (1995) Carnivorous sponges. Nature 373:333 22. Godefroy N, Le Goff E, Martinand-Mari C, Belkhir K, Vacelet J, Baghdiguian S (2019) Sponge digestive system diversity and evolution: filter feeding to carnivory. Cell Tissue Res 377(3):341–351. https://doi.org/10. 1007/s00441-019-03032-8 23. Martinand-Mari C, Vacelet J, Nickel M, Wo¨rheide G, Mangeat P, Baghdiguian S (2012) Cell death and renewal during prey capture and digestion in the carnivorous sponge Asbestopluma hypogea (Porifera: Poecilosclerida). J Exp Biol 215:3937–3943 24. Vacelet J, Duport E (2004) Prey capture and digestion in the carnivorous sponge Asbestopluma hypogea (Porifera: Demospongiae). Zoomorphology 123:179–190

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25. Ereskovsky AV (2010) The comparative embryology of sponges. Springer Netherlands, Dordrecht 26. Ereskovsky AV, Borchiellini C, Gazave E, Ivanisevic J, Lape´bie P, Perez T, Renard E, Vacelet J (2009) The Homoscleromorph sponge Oscarella lobularis, a promising sponge model in evolutionary and developmental biology. BioEssays 31:89–97 27. Caroline R, Ame´lie V, Laura F-C, Nina S, Sandrine C, Christian M, Emilie LG, Morgan D, Ce´dric M, Florent M, Nicolas B, Dominique M-H, Alexander E, Andre´ LB, Carole B, Emmanuelle R (2020) The buds of Oscarella lobularis (Porifera): a new convenient model for sponge cell and developmental biology. bioRxiv. https://doi.org/10.1101/2020.06. 23.167296 28. Leys SP, Larroux C, Gauthier M, Adamska M, Fahey B, Richards GS, Degnan SM, Degnan BM (2008) Isolation of Amphimedon developmental material. CSH Protoc 2008:pdb. prot5095 29. Degnan SM, Degnan BM (2010) The initiation of metamorphosis as an ancient polyphenic trait and its role in metazoan life-cycle evolution. Philos Trans R Soc Lond Ser B Biol Sci 365:641–651 30. Nakanishi N, Sogabe S, Degnan BM (2014) Evolutionary origin of gastrulation: insights from sponge development. BMC Biol 12:26 31. Larroux C, Fahey B, Adamska M, Richards GS, Gauthier M, Green K, Lovas E, Degnan BM (2008) Whole-mount in situ hybridization in Amphimedon. CSH Protoc 2008:pdb. prot5096 32. Stephens DJ, Allan VJ (2003) Light microscopy techniques for live cell imaging. Science 300:82–86 33. Weiss DG (1992) Video-microscopic analysis of the motility of sub-resolution sized cell constituents. Fresen J Anal Chem 343:39–40

Chapter 6 Microscopy Studies of Placozoans Carolyn L. Smith, Tatiana D. Mayorova, Christine A. Winters, Thomas S. Reese, Sally P. Leys, and Andreas Heyland Abstract Trichoplax adhaerens is an enigmatic animal with an extraordinarily simple morphology and a cellular organization, which are the focus of current research. Protocols outlined here provide detailed descriptions of advanced techniques for light and electron microscopic studies of Trichoplax. Studies using these techniques have enhanced our understanding of cell type diversity and function in placozoans and have provided insight into the evolution, development, and physiology of this little understood group. Key words Placozoa, Electron microscopy, Immunohistochemistry, Trichoplax, Cryopreservation, Ultrastructure

1

Introduction Trichoplax adhaerens was described over 100 years ago by Schulze [1] and assigned by Grell [2] to the phylum Placozoa in which it and two other recently named species, Hoilungia hongkongensis and Polyplacotoma mediterranea [3, 4], are the only members to date. Renewed interest in these enigmatic organisms is due to their basal phylogenetic position within the animal kingdom as well as their uniquely simple morphological organization (for recent reviews see 5, 6). Trichoplax appears as a flattened disk a few millimeters in diameter (Fig. 1a). Its lower and upper surfaces are ciliated with monociliated cells (Figs. 1b–j and 2a, b) and are coated with a layer of mucus (Fig. 1b, e). The edge typically appears upturned in preserved specimens (Fig. 1c, d), as does the edge of living animals as they glide across a surface. Though no more than a small disc, Trichoplax is nevertheless polarized with a thick ventral layer of ciliated and secretory cells, a thin middle layer, and an equally thin upper ciliated layer (Figs. 1f–h and 2a). The upper surface cells are held tightly together by intercellular junctions, and cilia

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 (a) View of the upper surface of a whole Trichoplax adhaerens. (b) Magnification of a portion of the upper surface of the specimen in (a) showing bacteria (arrow) caught in mucus. (c) A view of the lower surface of a fractured specimen. The animal is mounted upper surface down. The portion in the rectangle is shown in (d). (d) A fracture of the upturned edge of an animal showing the thin layer of cells on the upper surface and thicker columnar layer of cells on the lower surface. (e, f) Magnified views of the upper surface of the specimen in (d) showing the single layer of ciliated cells that are tightly juxtaposed and the apparent absence of any extracellular matrix below that layer. Fiber cells lie just above the columnar cells that form the lower surface of the animal. (g) The upper surface of Trichoplax showing tightly adhered cells with membrane protrusions from the ciliated cells. (h) A fracture of an animal showing the contrast between ciliated cells of the upper and lower surface. A thin layer of mucus (arrowhead) lies below the cilia on the lower surface. (i, j) The cells of the lower surface appear to have large numbers of microvilli-like extensions seen in cross section (i, arrow) and in surface view (j, arrow). Abbreviations: us upper surface, ls lower surface, mu mucus, fc fiber cell

arise from slightly raised ridges on the surface of the cells (Fig. 1g). The cells are contractile and behave as a dynamic elastic sheet [7]. The ciliated ventral epithelial cells (VECs) on the lower surface propel the gliding movements of the animal. VECs also have

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Fig. 2 Electron microscopy of Trichoplax prepared by high pressure freezing and freeze substitution with hafnium staining. (a) The edge of the animal showing ventral (V) and dorsal (D) epithelial cells, gland cells (G), fiber cells (F) and lipophil (L) cells. Ventral epithelial cells bear a cilium (arrow) and microvilli (arrowhead) and contain an electron dense inclusion in cytoplasm (asterisks). TEM of an ultrathin section counterstained with lead and uranyl acetate. (The image modified after [15]). (b) Ventral epithelial cells (V), gland cells (G) and a fiber cell (F) viewed by scanning electron microscopy; increased concentration of HfCl during freeze substitution, but no section counterstaining. (c) Adherens junction between ventral epithelial cells. Arrow indicates actin cytoskeleton of the microvillus. TEM of an ultrathin section counterstained with phosphotungstic acid. Scale bars: A, B—2 μm; C—200 nm

numerous microvilli (Figs. 1h–j and 2). Interspersed among the VECs are lipophil cells (Figs. 2a and 3d), which are packed with large lipophilic secretory granules that contain digestive enzymes that the animal uses for extracellular digestion of microorganisms [8]. The ventral epithelium also includes several different types of peptidergic gland cells (that are thought to have regulatory functions 9–12). All epithelial cells are joined by adherens junctions (Fig. 2c) that fix the positions of cells relative to one another but provide only a leaky barrier to diffusion of molecules into the intercellular clefts [13]. Sandwiched between the dorsal and ventral epithelium is a layer of fiber cells that have long branching processes that contact the other cell types (Fig. 3a, c). Fiber cells likely serve as a scaffold to hold the dorsal and ventral epithelia together [10] and are thought to be contractile [14]. Crystal cells, so called because they contain a birefringent crystal [10], are arrayed in a row around the edge of the animal and are reported to be functional statocysts [15]. Recent studies have introduced improved microscopic techniques for investigating placozoans [7, 10–12, 16]. Scanning electron microscopy (SEM) of Trichoplax wholemounts is useful for revealing structures on the outer surfaces of the epithelial cells or, following removal of the dorsal epithelium or fracturing the animal, the organization of cells in the interior [10, 17]. Early studies of Trichoplax by transmission electron microscopy (TEM) showed that cellular fine structure was not well preserved by chemical fixation, as membranes were broken and some cytoplasmic

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Fig. 3 Immunolabeling in frozen, freeze-substituted Trichoplax. (a, b) Oblique cross section from an animal stained with anti-alpha tubulin (red), anti-TaCDH (cadherin, green) and Hoechst stain (blue). Abundant tubulincontaining cilia protrude from the thick ventral epithelium (VE) while the dorsal epithelium (DE) is thin and has sparse cilia. Green TaCDH staining outlines fiber cells (FC), which have large, weakly red inclusions in their interiors. TaCDH also stains the apical surfaces of epithelial cells. The red and blue channel image (b) shows the basal apparatus (BA) at the bases of the ventral cilia and microtubules in cell interiors. (c, d) En face views of an animal stained with anti-pan MAGUK (red), anti-TaCDH (green) and Hoechst stain (blue). Projection of three, 0.5 μm optical sections in the fiber cell layer (c) shows fiber cells outlined by TaCDH staining and, around the edge, epithelial cells outlined by MAGUK staining. Note the absence of MAGUK staining at the

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structures were damaged [18–20]. Application of freezing and freeze substitution yielded significant improvement in tissue preservation [21, 22] and helped resolve structures such as cytoskeletal elements and intercellular contacts. Recent TEM studies of Trichoplax tissue showed that optimal results were achieved by combining freeze substitution with high pressure freezing of living animals [8, 10, 13, 16]. High pressure freezing followed by a finely honed freeze substitution allows investigators to minimize the artifacts seen with room temperature fixation and the sample processing that follows. These techniques were also adapted to prepare animals for analysis by serial sectioning and by SEM [15] and are suitable for sophisticated electron microscopy (EM) techniques such as tomography [23]. Poor preservation of Trichoplax cells following chemical fixation also hampered early investigations with light microscopy and fluorescent labels [9, 21]. In this chapter, we provide a simple protocol for freezing and freeze substituting Trichoplax prior to chemical fixation that provides superior preservation of cell structure and more intense fluorescent labeling [10].

2

Materials

2.1 Culturing and Maintenance

1. Cryptomonas sp. (from UTEX at the University of Texas at Austin, stock number LB 2423 - http://web.biosci.utexas. edu/utex/). 2. f/2 growth medium (Sigma-Aldrich G0154). 3. Pyrex® 90 mm  50 mm evaporating dishes (part # 3140). 4. Disposable borosilicate glass Pasteur pipettes.

2.2 High Pressure Freezing for Thin Section EM

1. High pressure freezing machine Baltec 010 (Techno Trade). 2. Liquid Nitrogen (RLN160DG, Roberts Oxygen Co, Inc. Gaithersburg MD). 3. Pure nitrogen ultra-pure carrier gas (R104A3, Roberts Oxygen Co, Inc. Gaithersburg MD. 4. Isopropanol (9084-1, JT Baker, Avantor Performance Materials, Center Valley, PA). 5. Hexadecene (H2131 Sigma-Aldrich, St. Louis, MO).

 Fig. 3 (continued) apical ends of the epithelial cells. Optical section near the lower surface of the ventral epithelium (d, red and blue channel) shows the uniform layer of MAGUK surrounding epithelial cells. The large profiles (L) that have no nucleus inside likely represent lipophil cells, which are larger than the more numerous ciliated ventral epithelial cells and have a nucleus located deeper in the interior

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6. Gold specimen chambers 3.0  0.8, indentation 1.4 mm dia., 0.3 mm deep (16LZ02125VN, Leica microsystems, Buffalo Grove, IL). 7. 1 ml Cryotubes, (36656 Nunc, Daigger, Vernon Hills, IL) 8. A pointed tool, for example, an awl. 9. Aluminum cans. 10. Liquid nitrogen resistance twine. 11. Ethanol. 12. Low Melting Point Agarose, Life Technologies, Gaithersburg, MD dissolved in artificial seawater preheated to 30  C to keep it liquid. 13. 1,2-Dipalmitoyl-sn-Glycero-3-Phosphocholine Avanti Polar Lipids, Alabaster, AL).

(850355P

14. Chloroform. 15. Appropriate forceps for transfer and handling under liquid nitrogen (16701955 Leica Microsystems, Buffalo Grove, IL) and Dumont Tweezers (No 5223, Ted Pella, Redding, CA). 16. Wrench or other device to allow one to remove chambers from specimen holder and foam container. 2.3 Freeze Substitution for Thin Section EM

1. AFS machine with temperature tubes, (AFS, Leica Buffalo Grove, IL). 2. 100 glass vials 20 ml size with Urea/Polyseal Cone cap (EF28410K 74515-20 20, Daigger Vernon Hills, IL 60061). 3. HPLC grade acetone (270725-100ML, Sigma-Aldrich, St. Louis MO). 4. Uranyl Acetate- (21447, Polysciences, Warrington, PA). 5. Acrolein (110221-25, Sigma-Aldrich, St. Louis MO. 6. Osmium tetroxide (19130 Electron Microscopy Sciences, Hatfield, PA). 7. Hafnium (IV) St. Louis MO).

Chloride

(590592-5G,

Sigma-Aldrich,

8. Micro Spoon Heyman Type A Heyman type micro-spatula made from nickel stainless steel with a glazed finish. The wire diameter is 0.103 in. (2.62 mm). Overall length: 6¼ in. (159 mm). Flat end: 1¼ in. (31.35 mm) long. Spoon end: ½ in. (12.5 mm) long  1/8 in. (3 mm) wide. 9. HPLC-grade Methanol (34860-100ML-R, Sigma-Aldrich, St. Louis MO). 10. SPI-Pon 812 Embedding Kit (SPI supplies, West Chester, PA). 11. Disc Block Embedding mold, blue (10530, Ted Pella, Redding, CA).

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12. Sonicator (2510, Branson, Danbury, CT). 13. Leica plastic capsules—small acetone resistant basket that allows easy sample transfer (16702734, Leica Buffalo Grove, IL). 14. Appropriate forceps for transfer and handling under liquid nitrogen (16701955 Leica Microsystems, Buffalo Grove, IL and Dumont Tweezers prod No 5223, Ted Pella, Redding, CA). 15. Jeweler’s saw and saw blades 79.40.008 Size 8/0 Herkules “White Label” Saw Blades (Starr Gems Inc. Silver Supplies). 2.4 Chemical Fixation for SEM

1. Osmium tetroxide (19130 Electron Microscopy Sciences, Hatfield, PA). 2. Glutaraldehyde (Electron Microscopy Sciences, Hatfield, PA). 3. Sodium acetate (Sigma-Aldrich). 4. “Glass Shell Vials”—Fisher Scientific (03-339-30). 5. Appropriate forceps for transfer and handling under liquid nitrogen (16701955 Leica Microsystems, Buffalo Grove, IL and Dumont Tweezers prod No 5223, Ted Pella, Redding, CA). 6. Aluminum stubs for mounting specimens; model and size depends on the scanning electron microscope used. Stub holders. 7. Nail polish. 8. Eye-brow tool, made by attaching an eye-brow hair to the end of a small wooden rod.

2.5 Freezing, Freeze Substitution, and Immunolabeling for Light Microscopy

1. Cover glasses. 22 mm square #1.5 German glass cover glasses. Warner Instruments #64-0721. Carl Zeiss Microscopy LLC #474030-9020-000. 2. Cover glass staining rack. Electron Microscopy Sciences (EMS)m# 72238. 3. Glass beakers. 4. Glass staining jars (2 oz.) with lids. Fisher Scientific #13-756722. 5. Nitric acid. 6. 3-Aminopropyltriethoxysilane #A3648-100 ml.

(APES).

Sigma-Aldrich

7. Tetrahydrafuran. Sigma-Aldrich # A3648-100 ml. Alternatives: acetone or methanol. 8. Ethanol, methanol, acetone. 9. D-mannitol 0.97 M aqueous solution.

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10. Paraformaldehyde 16% aqueous solution. EMS #30525-89-4. 11. Phosphate buffered saline (PBS). 12. Artificial seawater (ASW). 13. Dry ice. 14. Styrofoam box. 15. Drying oven. 16. Humid staining chamber. 17. Hoechst 3342 20 mM solution. ThermoFisher #62249. 18. Antibodies: mouse anti-pan MAGUK clone K28/86, Millipore Sigma #MABN72; mouse anti-α tubulin, Sigma-Aldrich #T9026; rabbit anti-TaCHD custom affinity-purified antibody from New England Peptide. 19. Secondary antibodies: Alexa 555 goat anti-mouse IgG, Thermo Fisher #21422; Atto 488 goat anti-rabbit IgG, Sigma-Aldrich #18772. 20. Blocking buffer: PBS with 3% normal goat serum, 2% horse serum, 1% BSA. 21. Mounting medium with anti-fade reagent, such as Prolong Gold. ThermoFisher # P36930. 22. Glass slides. 23. Thin (35 ppt) better than lower salinities (12 h. 2. Rinse in distilled water 5. 3. Bake in drying oven 200  C degrees for 2–3 h. Store in a dustfree container. 4. Load cover glasses into racks and submerge in a beaker containing APES. Soak for 2 min. 5. Pour APES back into a stock bottle and purge with nitrogen gas. 6. Rinse cover glasses in acetone 2 for 2 min. 7. Rinse in distilled water 5. 8. Dry at 40  C overnight. 9. Pour tetrahydrofuran (THF) into glass staining jars. This procedure should be done in a fume hood because THF is toxic and highly flammable. Loosely replace the lids on the jars. Acetone or MeOH can be used instead; however, the latter produces more autofluorescence, and neither solvent preserves cell structure as well as THF does. 10. Place the jars in a Styrofoam box containing ~10 cm of dry ice. Allow the jars to chill for at least 2 h. 11. Mix equal volumes of 0.97 M mannitol and ASW and pipet 200 μl of the solution to the center of each cover glass. 12. Transfer ~6 animals into each drop of mannitol/ASW. Allow the animals to flatten and adhere to the substrate (~2 h). 13. Pick up cover glass with forceps, tip vertically and blot edge and the back of the coverslip with a tissue to remove excess ASW.

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14. Quickly plunge each cover glass into a jar containing THF. Keep your face well away from jar because the THF sometimes splatters when the cover glass is submersed. Store overnight, making sure that there is sufficient dry ice to last. 15. Add to a separate set of staining jars 9 ml MeOH and 1 ml aqueous paraformaldehyde solution (16%). Store in a 20  C freezer overnight. These jars can be stored in the freezer and reused in subsequent experiments. 16. The next day, transfer the coverslips from the jars containing THF to jars containing MeOH/paraformaldehyde. Store at 20  C for 2 h. 17. Remove the jars from the freezer and keep at room temperature for ~1 h. 18. Transfer the coverslips to a rack in a beaker containing 100% EtOH. Then rehydrate the coverslips in a graded series of EtOH/PBS mixtures beginning with 90% EtOH, followed by 70% and 50% each for 5–10 min. Rinse in PBS. The EtOH/PBS solutions can be reused in subsequent experiments. 19. Place the cover glasses in a humid staining chamber and add to each 200 μl PBS. Incubate 5 min and then remove PBS. 20. Add 200 μl blocking buffer. Incubate 15 min and then remove buffer. 21. Prepare primary antibody solution in blocking buffer, using 200 ml for each cover glass. Dilute mouse anti-pan MAGUK or anti-tubulin 1:500. Dilute rabbit anti-TaCDH 1:200. Briefly spin the solution in a mini-centrifuge to sediment particulate material and then transfer it to the coverslips. 22. Tightly close the humid chamber and store at 4 overnight. 23. Rinse cover glasses 3 in PBS each for 5 min (see Note 21). 24. Prepare secondary antibodies and Hoechst stain in blocking buffer. Dilute Atto 488 anti-rabbit and Alexa 555 anti-mouse 1:500. Dilute Hoechst 1:1000. Sediment by spinning in a mini-centrifuge. 25. Transfer secondary antibody solution to the cover glasses and incubate in a dark place for 2–4 h at room temperature. 26. Rinse 3 with PBS each for 5 min. 27. Mount the wafers (EM grids) onto the glass slides using silicon grease as the adhesive and placing three wafers in a triangle to support each cover glass. 28. Put one drop of mounting medium in the center of each triangle of wafers.

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29. Remove excess PBS from the coverslip, wipe the back of it with a tissue and then mount on top of the wafers. If the mounting medium is a type that solidifies, then allow it to harden before viewing. Or, if you want to look at the samples immediately, put a small drop of fingernail polish at each corner of the cover glass to secure it to the slide. If the mounting medium is the type that remains liquid, then completely seal the edges of the cover glass with fingernail polish. 30. To observe the results of the antibody labeling, view the samples with a confocal microscope (Fig. 3; see Notes 22–24). 3.7

Fixation for SEM

Fracturing specimens for scanning electron microscopy provides a three-dimensional view of cells and tissues. This method is adapted for chemical fixation that optimizes structural preservation. The fixative is a cocktail in which osmium and glutaraldehyde are mixed with a buffer immediately prior to preserving the tissues. 1. Submerge sterile coverslips (ethanol cleaned and flamed) into seawater in a Petri dish and transfer several Trichoplax individuals onto the coverslips. Allow the animals to adhere and crawl over several hours. 2. The final concentrations of fixative will be 2% glutaraldehyde, 1% osmium tetroxide in 0.45 M sodium acetate buffer pH 6.4, with 10% sucrose in the final solution. The method involves mixing two solutions, a glutaraldehyde buffer solution and osmium tetroxide in a ratio of 3:1 immediately prior to immersing tissues. 3. Glutaraldehyde solution: In a fume hood, pour 20 ml of 0.45 M sodium acetate into a 50 ml tube. Add 10 ml of 8% electron microscopy grade glutaraldehyde. Add 4 g sucrose, cap tightly and mix by inversion. Allow 2 h for the sucrose to dissolve and for the solution to be entirely mixed. 4. For two coverslips with Trichoplax attached, in a fume hood, put 3 ml of the glutaraldehyde solution into a tube, add 1 ml of 4% osmium tetroxide solution, and mix by pipetting. Add 2 ml of this to each 35 mm Petri dish. 5. Using sterile forceps remove the coverslips with Trichoplax from their culture dish and immerse them directly into fixative. Place the Petri dishes into a container with sealing lid (e.g., Ziploc® box) and keep cool for 6 h. Cooler temperatures slow the fixation process. 6. Rinse fixative from tissues by moving the coverslips to a Petri dish with distilled water for 1 min. Change the water twice for 1 min each. 7. Dehydrate by replacing half the volume with 70% ethanol; leave 5 min. Replace half the volume another three times, each time

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leaving tissues in the new solution for 5 min. Coverslips can be kept in 70% ethanol overnight. 3.8 Freeze Fracturing Tissues for SEM

1. To lift Trichoplax from the coverslips without damaging the tissue it is necessary to dissolve away the silica of the coverslip. Make a solution of 4% hydrofluoric acid in 70% ethanol. 2. Add 2 ml of the solution to a new 35 mm Petri dish. Immerse the coverslip with Trichoplax directly into the hydrofluoric acid–ethanol solution for 1–5 min. The animals appear as dark patches. 3. After 5 min using a transfer pipette, gently squirt the solution over the placozoan to dislodge it from the coverslip and pipet it into a new dish with 70% ethanol. 4. Transfer the animals to a glass vial (e.g., “Glass Shell Vial”) and dehydrate to 95% and then 100% ethanol. 5. To fracture the tissues, fill a styrofoam cooler 2 cm depth with liquid nitrogen. Remove ethanol from the glass vial with animals in the bottom, until only 0.5 cm height covers the Trichoplax. Best results are obtained if the animals are in the middle of the bottom of the vial. 6. Using protective glasses and plastic forceps carefully transfer the vial with ethanol and animals into the liquid nitrogen, ensuring it stays vertical. With time the ethanol will freeze solid and fracture. Approaching the vial with metal such as a scalpel with metal blade can speed up fracture. 7. After fracture, remove the glass vial to a Petri dish to thaw. Once it has thawed, check the extent of fracture by looking at the pieces under a stereo microscope. If insufficiently fractured, repeat step 6. If sufficiently fractured continue to the next steps immediately or store in 70% ethanol for subsequent processing.

3.9 Critical Point Drying and SEM

1. Use a Bal-Tec 030 critical point dryer or equivalent to dry the samples. Mesh size of the sample holder should be 30–50 μm and can be adjusted by using Nitex® mesh screwed into the cover of the holder or by making small mesh pockets in which to hold the tissue fragments. 2. Pipet the pieces of Trichoplax into the holder while in 100% ethanol. Transfer to the Critical Point Dryer and begin to exchange the ethanol for CO2. Ensure exchange of CO2 fully happens (often 1 h is required for full exchange). 3. After reaching critical point, reduce pressure slowly, often over 1 h, and after reaching atmospheric pressure remove specimens to a clean Petri dish lined with filter paper. 4. Using a stereomicroscope, mount fractured specimens on aluminum SEM stubs using nail polish and an “eye-brow” brush

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made from a single eye-brow hair attached to the end of a wooden rod. 5. Allow the nail polish to dry overnight. Sputter coat with a Nanotek SEMprep2 sputter coater or equivalent, and keep in a desiccator until viewed by SEM (for example of such a specimen see Fig. 1).

4

Notes 1. The amount of time needed to maintain Trichoplax largely depends on whether filtered seawater is readily available and whether the cultures are fed with live algae or rice (see Collection and Maintenance). 2. The salinity can be measured with a handheld salinity meter or refractometer. 3. Trichoplax cultures can be easily maintained for several months using uncooked rice. Add 2–3 rice grains to the culture at each water change and remove the old ones. However, if cultures are maintained for longer periods, the use of live algae is recommended. 4. All glassware that is used for Trichoplax or algae cultures needs to be embryologically clean. This means that it must be autoclaved and not exposed to any soap, detergent, or bleach at any time. It is generally recommended that a separate area with dedicated tools is set aside in the lab for cleaning. 5. Trichoplax can be shipped in 50 ml Falcon tubes. Rinse the tube with deionized water then fill it with FSW. Place the desired number of Trichoplax inside the tube and wait 10 min for the Trichoplax to attach themselves. Gently add more water until the tube starts to overflow, at this point stretch Parafilm™ over the top of the tube and tighten the lid on the tube. 6. We tested different algae species, and the best results were achieved with Cryptomonas sp. (stock number LB 2423 from UTEX): The Culture Collection of Algae; The University of Texas at Austin; 1 University Station A6700; Austin, TX 78712-0183; USA; (ph) 512-471-4109; (fax) 512-471-0354. 7. If cultures are supplemented with CO2 it is recommended to keep it at a concentration of approximately 1% which can be achieved by gently bubbling CO2 into the algae cultures. This cannot be done with the axenic stocks. 8. Recommended protocol for maintaining axenic algae cultures: First, open the sterile algae culture immediately after arrival and transfer 1 ml of culture into two autoclaved tall disposable borosilicate glass test tubes with either a breathable plastic lid

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or a sterile cotton plug. To the same tubes add 4 ml of sterile FSW containing 0.05% f/2 component A and B (i.e., 2 μl component A and 2 μl component B; note that this stock can also be prepared in larger quantities and kept at 4  C). Care should be taken not to cross-contaminate component A and B (use different pipette tips). This and all subsequent transfers should be done under a laminar flow hood using sterile pipette tips and glassware. The axenic culture needs to be changed once every 2 weeks by transferring 1 ml of the original culture into a new tube with 4 ml f/2. One such culture (5 ml total volume) can also be used to setup a new working stock culture (see below) after at least 1 week of growth. 9. To estimate algal density take a small sample from these cultures and count the number of cells every 2 days using a hemocytometer. Do not use the culture to feed Trichoplax once the growth curve reaches a plateau (the maximum cell density ranges from 1000 cells/μl under low light conditions to 3000 cell/μl under high light conditions). Once the working stock culture has reached this point, transfer 100 ml into 500 ml growth medium (250 μl of f/2 component A and 250 μl component B in 500 ml of FSW) to setup a new culture. Note that this transfer should be done maximally four times before a new axenic culture sample is used to inoculate the working stock culture. 10. While cultures can be maintained in plastic dishes, we have better success with glass dishes. 11. Alternatively, cultures can be inoculated with algae up to a week before adding Trichoplax so that a biofilm can grow. 12. Gold specimen chambers 3.0  0.8 mm, indentation 1.4 mm dia., 0.3 mm deep (16LZ02125VN, Leica Microsystems, Buffalo Grove, IL) are no longer sold by Leica. 13. One can also put an animal into the chamber directly from the pipette tip, but the water flow may remove animals put into the chamber previously. Up to five animals one by one could be transferred into one chamber. 14. AFS machine has been replaced by the AFS2 from Leica. 15. All solutions and rinses should be dry; do not expose solutions to air for a long time since this will cause water absorption; work quickly when preparing solutions or transferring samples. For the same reason, use freshly open acetone bottles for staining solutions. 16. Acrolein is no longer sold by Sigma. 17. Avoid touching the specimen gold chamber with forceps or otherwise during freeze substitution; if upside down, shake the basket carrier to flip it over; do this so that the sample is submerged into solution.

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18. The 40  C and 90  C tubes are devices which go together with a FS machine. They are connected to the FS chamber with samples. An appropriate temperature tube should be used according to the programmed temperature range. 19. Hafnium powder absorbs water from room air in seconds and becomes useless, as seen by failure to dissolve in acetone. 20. Do not use plastic Pasteur pipettes since this also produces bubbles. 21. In the following steps, be gentle while removing and replacing solutions to avoid detaching the animals from the cover glass. 22. The overall shapes of the animals should remain largely unchanged after processing, although occasionally animals detach from the substrate or suffer varying amounts of damage. Examples of animals immunolabeled with anti-TaCDH and anti-tubulin or anti-pan MAGUKs are shown in Fig. 3. Tubulin labeling (Fig. 3a, b) reveals microtubule-containing cilia protruding from the surface of the epithelium and bundles of microtubules in cell interiors. Anti-TaCDH stains the external surfaces of epithelial cells (Fig. 3a) and brightly outlines fiber cells located in zone between the dorsal and ventral epithelia (Fig. 3a, c). Large inclusions inside the fiber cells autofluoresce weakly in the red channel. Staining for pan MAGUKs lines epithelial cell junctions (Fig. 3c, d). Nuclei stained with Hoechst (Fig. 3a–d) have smooth outlines. The empty pockets near the edge of the animal are invaginations that may have formed during freezing. 23. In our experience, most intracellular organelles, including mitochondria, lysosomes, endoplasmic reticulum, Golgi complex, are well preserved by freezing and freeze substitution. However, lipophil cells in the ventral epithelium lose their large lipophilic granule during processing. Also, the secretory granules in gland cells tend to aggregate at the apical ends of the cells, whereas these granules appear more dispersed when examined by electron microscopy after having been prepared by freezing at high pressure. Conventional fixation with 4% paraformaldehyde provides better preservation of the secretory granules of gland cells, but not lipophil cells. 24. Interpreting the results of immunolabeling experiments requires appropriate control experiments to establish the specificity of the primary and secondary antibodies. A recommended procedure for experiments employing a polyclonal antibody is to process three samples in parallel, staining one with the primary antibody, the second with the same concentration of primary antibody preabsorbed with the antigen used for immunization, and the third with no primary antibody.

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References 1. Schulze FE (1883) Trichoplax adhaerens. Zool Anz 6:92–97 2. Grell KG (1971) Trichoplax adhaerens, F.E. Schulze und die entstehung der metazoen. Nat wiss Rundsch 24:160–161 3. Eitel M, Osigus HJ, Desalle R et al (2013) Global diversity of the Placozoa. PLoS One 8 (4):e57131. https://doi.org/10.1371/jour nal.pone.0057131 4. Osigus H-J, Rolfes S, Herzog R et al (2019) Polyplacotoma mediterranea is a new ramified placozoan species. Curr Biol 29:R148–R149 5. Syed T, Schierwater B (2002) Trichoplax adhaerens: discovered as a missing link, forgotten as a hydrozoan, re-discovered as a key to metazoan evolution. Vie Milieu 52:177–187 6. Schierwater B (2005) My favorite animal, Trichoplax adhaerens. BioEssays 27:1294–1302 7. Armon S, Storm Bull M, Aranda-Diaz A et al (2018) Ultra-fast cellular contractions in the epithelium of T. adherens and the “active cohesion” hypothesis. bioRxiv 258103 8. Smith CL, Pivovarova N, Reese TS (2015) Coordinated feeding behavior in Trichoplax, an animal without synapses. PLoS One 10(9): e0136098. https://doi.org/10.1371/journal. pone.0136098 9. Schuchert P (1993) Trichoplax adherens (phylum Placozoa) has cells that react with antibodies against the neuropeptide Rfamide. Acta Zool 74:115–117 10. Smith CL, Varoqueaux F, Kittelmann M et al (2014) Novel cell types, neurosecretory cells, and body plan of the early-diverging metazoan Trichoplax adhaerens. Curr Biol 24:1565–1572 11. Senatore A, Reese TS, Smith CL (2017) Neuropeptidergic integration of behavior in Trichoplax adhaerens, an animal without synapses. J Exp Biol 220:3381–3390 12. Varoqueaux F, Williams EA, Grandemange S et al (2018) High cell diversity and complex peptidergic signaling underlie placozoan behavior. Curr Biol 28:3495–3501.e2 13. Smith CL, Reese TS (2016) Adherens junctions modulate diffusion between epithelial cells in Trichoplax adhaerens. Biol Bull 231:216–224

14. Grell KG, Ruthmann A (1991) Placozoa. In: Harrison FW, Westfall JA (eds) Microscopic anatomy of invertebrates. Wiley-Liss, New York, pp 13–28 15. Mayorova TD, Smith CL, Hammar K et al (2018) Cells containing aragonite crystals mediate responses to gravity in Trichoplax adhaerens (Placozoa), an animal lacking neurons and synapses. PLoS One 13(1): e0190905. https://doi.org/10.1371/journal. pone.0190905 16. Eitel M, Francis WR, Varoqueaux F et al (2018) Comparative genomics and the nature of placozoan species. PLoS Biol 16(7): e2005359. https://doi.org/10.1371/journal. pbio.2005359 17. Rassat J, Ruthmann A (1979) Trichoplax adhaerens Schulze, F.E. (Placozoa) in the scanning electron microscope. Zoomorphologie 93:59–72 18. Guidi L, Eitel M, Cesarini E et al (2011) Ultrastructural analyses support different morphological lineages in the phylum Placozoa Grell, 1971. J Morphol 272:371–378 19. Ruthmann A, Terwelp U (1979) Disaggregation and reaggregation of cells of the primitive metazoan Trichoplax adhaerens. Differentiation 13:185–198 20. Grell KG, Benwitz G (1974) Elektronenmikroskopische beobachtungen uber das wachstum der eizelle und die bildung der “befruchtungsmembran” von Trichoplax adhaerens F. E. Schulze (Placozoa). Zeitschrift fur Morphol der Tiere 79:295–310 21. Behrendt G, Ruthmann A (1986) The cytoskeleton of the fiber cells of Trichoplax adhaerens (Placozoa). Zoomorphology 106:123–130 22. Ruthmann A, Behrendt G, Wahl R (1986) The ventral epithelium of Trichoplax adhaerens (Placozoa) - cytoskeletal structures, cell contacts and endocytosis. Zoomorphology 106:115–122 23. Gruber-Vodicka HR, Leisch N, Kleiner M et al (2019) Two intracellular and cell type-specific bacterial symbionts in the placozoan Trichoplax H2. Nat Microbiol 4(9):1465–1474

Chapter 7 Identification of SH2 Domain-Mediated Protein Interactions that Operate at Fertilization in the Sea Star Patiria miniata Lauren Bates, Emily Wiseman, Jamie Kitson, and David J. Carroll Abstract The signaling mechanisms controlling internal calcium release at fertilization in animals are still largely unknown. Echinoderms, such as the sea star Patiria miniata, produce abundant and easily accessible sperm and eggs. In addition, eggs are naturally synchronized at the same cell cycle stage, collectively making these animals an attractive model to study the signaling proteins controlling fertilization. However, the lack of antibodies to identify proteins in this model system has slowed progress in identifying key signaling molecules. With the advances in mass spectrometry, we present a method for identifying tyrosine phosphorylated proteins binding to GST-tagged SH2 domains in sea star cell lysates for downstream mass spectrometry analysis. Key words Fertilization, Egg activation, Src Family Kinase, Ca release, Phospholipase C gamma, Affinity interaction, GST fusion protein

1

Introduction Many somatic cell signaling pathways have been well characterized; however, the identification of signaling molecules controlling fertilization has proven to be more elusive. Sea stars are an excellent model system for studying cell signaling at fertilization due to several features. For example, millions of sperm and eggs are easily obtained from each adult. Moreover, their large and clear eggs are synchronized at the same stage of the cell cycle, collectively making these cells amenable to a variety of biochemical and live cell assays [1, 2]. One of the major challenges in studying the eggs and sperm of sea stars and other model organisms used in developmental biology is the lack of available antibodies and genomic resources to manipulate and identify proteins. However, recent advancements in RNA sequencing and mass spectrometry should greatly facilitate such research by expanding protein identification beyond simple antibody-based methods of detection.

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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A universal mechanism critical to egg activation in all species is a rise in intracellular calcium [3–6]. Based on previous work, phospholipase C (PLC) enzymes are important regulators of calcium release at fertilization in several species [7–9]. Specifically in the sea star Patiria miniata, PLCγ is the key PLC subtype needed for calcium release, given that GST-tagged tandem SH2 domains of PLCγ inhibit calcium release at fertilization [10, 11]. Additionally, a sea star Src Family Kinase binds to these GST-PLCγSH2 proteins and is required for calcium release upstream of PLCγ [12–14]. This chapter describes proper GST-tagged control proteins for use in affinity interactions with sea star cell lysates using mass spectrometry to characterize interacting proteins, thereby providing an alternative to antibodies for protein identifications.

2

Materials All solutions are prepared in MilliQ water (18 MΩ) at room temperature and are listed with the final concentrations of the reagents, unless otherwise indicated.

2.1 Starfish Egg and Sperm Preparation

1. The Pacific bat star Patiria miniata (taxonomic ID 46514 in the National Center for Biotechnology Information database) can obtained from Marinus Scientific, LLC in Long Beach, CA. 2. Artificial sea water (ASW) [1]: 28.32 g/l NaCl, 0.77 g/l KCl, 1.18 g/l CaCl2, 5.41 g/l MgCL2-6H2O, 7.13 g/l MgSO47H2O, 0.2 g/l NaHCO3, pH 8.0. 3. Nitex Mesh of different opening sizes (Fisher Scientific). 4. 3 mm sample corer (Fine Science Tools) 5. Microsurgical scissors (Fine Science Tools). 6. 1-methyladenine (ACROS Organics) 7. 30 ml clean glass beakers 8. Small capacity incubator to keep oocytes at 16–18  C.

2.2

Mutagenesis

1. Q5®Site-Directed Mutagenesis Kit from New England Biolabs®, Inc. 2. High Speed Plasmid Mini Kit from IBI Scientific. 3. pGEX-6p vectors from GE Healthcare Life Sciences, 4. BL21(DE3)pLysS Corporation.

competent

cells

from

Promega

5. Luria Broth (10 g/l NaCl, 10 g/l peptone, 5 g/l yeast extract, pH 7.5–8.0). 6. Ampicillin from ThermoFisher Scientific.

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1. Isopropyl β-D-1-thiogalactopyranoside (IPTG) from USBiological Life Sciences. 2. Bacterial lysis buffer: 0.2% Triton X-100 in phosphate buffered saline, pH 7.3–7.5. 3. Luria Broth (LB): 10 g/l NaCl, 10 g/l peptone, 5 g/l yeast extract, pH 7.5–8.0. 4. Ampicillin from ThermoFisher Scientific. 5. Glutathione-Sepharose 4B® beads from GE Healthcare. 6. 2 sample buffer: 125 mM tris, pH 6.8, 4% SDS, 20% glycerol, 30 min. No soap should ever be used to clean these tools. 2. The oocytes are minced until the water appears orange and cloudy and the ovary is in tiny pieces. The scissors will not harm the oocytes. 3. An estimated volume of ASW is added in all of the steps. For example, if the ovary is originally placed in a 20 ml beaker, ~10–15 ml of ASW is added originally and for each washing. 4. Small (6–18 bottle) wine cooler refrigerators are excellent table top 16  C incubators. These can be found at many department stores and are more reasonably priced than a chiller incubator. 5. The easiest way to make a 10% solution of eggs is to pour a well-mixed beaker of eggs into a conical tube, allow the eggs to settle by gravity, then read the volume level on the tube where the eggs have settled to determine the volume of eggs. Next, remove or add ASW to create a 10% solution (e.g., if 1 ml of eggs, pour ASW up to the 10 ml mark). 6. The oocytes or eggs will settle by gravity to the bottom of the beaker. They can be resuspended by gentle swirling.

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Fig. 2 Tyrosine phosphorylated proteins detected in F5 samples at different stages of the affinity interaction procedure. Affinity interactions were performed using GST-PLCγSH2SH2 or GST in F5 eggs (eggs fertilized for 5 min) following the procedure indicated in 1.2. Samples were prepared in 2 sample buffer and loaded onto a 10% SDS-PAGE gel. The proteins were transferred to nitrocellulose and probed for tyrosine phosphorylation using Cell Signaling Technology® Phospho-Tyrosine Mouse mAb (P-Tyr-100) (1:1000). The blot was developed using Clarity™ Western Enhanced Chemiluminescence (Bio-Rad Laboratories, Inc.) and imaged using a ChemiDoc™ MP Imaging System under Chemi-Hi Sensitivity and exposed for 6 min. Lanes 1 and 6 contains the Bio-Rad Precision Plus Protein™ All Blue Prestained Protein Standard (3 μl). Lanes 2 and 7 contain 10 μl of the cell pellet left after cell lysis. Lanes 3 and 8 contain unbound proteins (20 μg protein per lane) after affinity interactions with GST-PLCγSH2SH2 or GST, respectively. Lanes 4 and 9 contain proteins bound to GST-PLCγSH2SH2 or GST, respectively. Lanes 5 and 10 contain 5 μg of GST-PLCγSH2SH2 or GST, respectively

7. Protein expression from pGEX6p is controlled by the IPTG inducible tac promoter. Additionally, pGEX6p vectors contain a lacIq gene that encodes a repressor protein that prevents protein expression until IPTG is added by binding to the operator region of the tac promoter (GE Healthcare Life Sciences). 8. For the initial mutagenesis PCR, the recommended annealing temperatures from NEBaseChanger® v1.2.9 are used; however, sometimes no product is generated, and they need to be lowered. 9. BL21(DE3)pLysS cells are used because they are deficient in proteases Ion and OmpT, increasing the stability of expressed proteins. 10. Aliquots of bacteria are taken before induction and at each hour after induction to monitor protein expression. These

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Fig. 3 Tyrosine phosphorylated proteins bound to GST-PLCγSH2SH2 at different time points before and after fertilization. Unfertilized eggs (2), and eggs fertilized for 1 min (3), 2.5 min (4), 5 min (5), 7.5 min (6), and 10 min (7) were lysed in TX100 lysis buffer. The samples were equilibrated to 1 mg/ml and 5 μg of wildtype GST-PLCγSH2SH2 were added to each. The samples incubated with the fusion protein for 4 h at 4  C with constant rocking (end-over-end). Samples were washed three times in TX100 lysis buffer, an equal volume of 2 sample buffer was added to each, and samples were heated at 95  C for 5 min. Samples were separated on a 10% SDS-PAGE gel, then transferred to nitrocellulose. The blot was probed for tyrosine phosphorylation using Cell Signaling Technology® Phospho-Tyrosine Mouse mAb (P-Tyr-100) #9411 (1:1000) and Cell Signaling Technology® Anti-mouse IgG, HRP-linked Antibody #7076S (1:10000)

samples are important when a new protein is being expressed to troubleshoot when there is no expression; however, once an expression method has been established, it is not necessary to collect such samples. 11. When sonicating, ensure the probe is ~1 cm below the surface of the lysate. Additionally, the number of sonication intervals will need to be empirically determined to get the optimal amount of cell lysis without degrading the fusion protein. The times in this protocol work for us and are a good starting point for establishing a protocol. 12. When using the Bio-Rad Protein Assay, we always run three replicates of standards and three replicates of samples. The 0.2% Triton X-100 in the bacterial lysis buffer has not interfered with this assay.

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13. The TX100 lysis buffer should be added to each sample immediately without starting the lysis by needle aspiration. This is so that as the samples thaw prior to needle aspiration, they are met with the protease inhibitors and phosphatase inhibitors in the lysis buffer. Once the buffer has been added to every sample, you can lyse by needle aspiration one at a time. Alternatively, one sample at a time can be removed from the freezer and lysed. 14. Pull up the sample to the ~0.8 ml mark on the syringe, then eject down to the ~0.2 ml mark on the syringe and repeat until homogenous. Never fully pull up the entire sample or eject the entire sample until finished lysing. The more bubbles that are created, the more likely it is that you are degrading proteins. 15. The samples need to be diluted 1:10 to reduce the concentration of Triton X 100 so it does not interfere with the assay. Add 10 μl of sample to 90 μl of water and pipet to mix. This 100 μl of diluted sample is plenty for three replicates in the protein assay. Be sure to multiply all the concentrations by 10 to get the real concentration. 16. The fusion proteins and egg lysates we have used have all resulted in values within the 0.5–0.05 mg/ml concentration range, thus we follow the microtiter plate procedure for this concentration range in the Bio-Rad Protein Assay protocol. 17. The incubation time for the affinity interaction with a particular wild-type construct should be empirically determined. We perform the affinity interactions for several different time periods using the same time point egg sample and decide on the shortest interaction time that detects interacting proteins. 18. The ratio of fusion protein to cell lysate will need to be empirically determined. We use the least amount of fusion protein that results in detectable interacting proteins. 19. Typically, the egg cell lysates contain ~3 mg/ml of protein. Therefore, we make two to three 1 mg/ml samples and perform the affinity interactions in each. We save one for mass spectrometry and use the remaining two for analysis by electrophoresis—one gel is subjected to Silver stain or Coomassie stain and the other is used for western blotting.

Acknowledgments We are very grateful for funding provided by the Eunice Kennedy Shriver National Institute of Child Health and Human Development of the National Institutes of Health under award number R15HD094324.

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References 1. Wessel GM, Reich AM, Klatsky PC (2010) Use of sea stars to study basic reproductive processes. Syst Biol Reprod Med 56:236–245 2. Wiseman E, Bates L, Dube´ A et al (2018) Starfish as a model system for analyzing signal transduction during fertilization. In: Kloc M, Kubiak J (eds) Marine organisms as model systems in biology and medicine, vol 65. Springer, Cham, pp 49–67 3. Stricker SA (1999) Comparative biology of calcium signaling during fertilization and egg activation in animals. Dev Biol 211:157–176 4. Runft LL, Jaffe LA, Mehlmann LM (2002) Egg activation at fertilization: where it all begins. Dev Biol 245:237–254 5. Whitaker M (2006) Calcium at fertilization and in early development. Physiol Rev 86:25–88 6. Townley IK, Roux MM, Foltz KR (2006) Signal transduction at fertilization: the Ca2+ release pathway in echinoderms and other invertebrate deuterostomes. Semin Cell Dev Biol 17:293–302 7. Wessel GM, Wong JL (2009) Cell surface changes in the egg at fertilization. Mol Reprod Dev 76:942–953 8. Parrington J, Davis LC, Galione A et al (2007) Flipping the switch: how a sperm activates the egg at fertilization. Dev Dyn 236:2027–2038 9. Swann K, Lai FA (2015) Egg activation at fertilization by a soluble sperm protein. Physiol Rev 96:127–149 10. Carroll DJ, Ramarao CS, Mehlmann LM et al (1997) Calcium release at fertilization in starfish eggs is mediated by phospholipase Cγ. J Cell Biol 138:1303–1311 11. Runft LL, Carroll DJ, Gillett J et al (2004) Identification of a starfish egg PLC-γ that

regulates Ca2+ release at fertilization. Dev Biol 269:220–236 12. Giusti AF, Carroll DJ, Abassi YA et al (1999) Requirement of a Src family kinase for initiating calcium release at fertilization in starfish eggs. J Biol Chem 274:29318–29322 13. Giusti AF, Carroll DJ, Abassi YA et al (1999) Evidence that a starfish egg Src family tyrosine kinase associates with PLC-1 SH2 domains at fertilization. Dev Biol 208:189–199 14. Giusti AF, Xu W, Hinkle B et al (2000) Evidence that fertilization activates starfish eggs by sequential activation of a Src-like kinase and phospholipase Cγ. J Biol Chem 275:16788–16794 15. Kanatani H, Shirai H, Nakanishi K et al (1969) Isolation and identification of meiosis inducing substance in starfish Asterias amurensis. Nature 221:273–274 16. Meijer L, Guerrier P (1984) Maturation and fertilization in starfish oocytes. Int Rev Cytol 86:129–196 17. Stoica B, DeBell KE, Graham L et al (1998) The amino-terminal Src homology 2 domain of phospholipase C gamma 1 is essential for TCR-induced tyrosine phosphorylation of phospholipase C gamma 1. J Immunol 160:1059–1066 18. Chattopadhyay A, Vecchi M, Ji QS et al (1999) The role of individual SH2 domains in mediating association of phospholipase C-γ1 with the activated EGF receptor. J Biol Chem 274:26091–26097 19. Merril C, Goldman D, Sedman S et al (1981) Ultrasensitive stain for proteins in polyacrylamide gels shows regional variation in cerebrospinal fluid proteins. Science 211:1437–1438

Chapter 8 Marine Nemertean Worms for Immunoblotting Studies of Oocyte Aging Stephen A. Stricker Abstract Immunoblotting analyses employing phospho-specific antibodies can help elucidate potential roles played by protein kinases as oocytes age and lose their ability to undergo normal fertilization. This chapter updates a previously published protocol for conducting immunoblotting analyses of oocyte maturation in marine nemertean worms by adding general methods for obtaining adult worms and for handling their gametes in experiments assessing oocyte aging. Key words Nemertea, Oocyte Aging, MAPK, MPF

1

Introduction Most oocytes must first initiate a process of maturation before they can be fertilized. Such maturation begins with the oocyte’s prophase-I–arrested nucleus (¼ the “germinal vesicle,” or GV), undergoing rapid disassembly during a phase referred to as germinal vesicle breakdown (GVBD). If left uninseminated, post-GVBD mature oocytes have a finite period of viability before their capacity to complete normal fertilization is lost and death is triggered. At the ~18  C, aged mature oocytes of starfish echinoderms lose fertilizability and end up disintegrating via an apoptotic form of programmed cell death beginning about a day after being removed from the ovary [1]. Similarly, at the higher culture temperatures used for mammals, aging-induced deterioration over a period of ~6–12 h can prevent mature mammalian oocytes from being properly fertilized, and in the absence of fertilization, aged oocytes eventually undergo apoptosis [2, 3]. To help prolong oocyte viability and optimize assisted reproductive technologies (ARTs), several factors potentially mediating oocyte aging and death have been analyzed [4]. For example, cellcycle-related kinases, such as the Cdc2 kinase of maturation-

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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promoting factor (MPF) and several kinds of mitogen-activated protein kinases (MAPKs) have been shown to affect the rates of oocyte aging. In most cases, prolonged MPF activation in mature oocytes retards oocyte aging, whereas extracellular signal regulated kinases (ERKs), c-Jun-N-terminal kinases (JNKs), and p38 types of MAPKs can either slow down or accelerate apoptotic death in aged oocytes [2, 5–11]. In most cases, the activities of these kinases in aging oocytes have been analyzed by means of immunoblotting protocols utilizing phospho-specific antibodies, and such studies have typically focused on species of deuterostome animals (e.g., echinoderms and vertebrates). However, unlike echinoderms and vertebrates, the vast majority of animals actually belong to protostome lineages (e.g., arthropods and worms), and to date, relatively few oocytes from protostomes have been analyzed for their patterns and mechanisms of aging. One group of protostomes that provide a suitable supply of oocytes for studies of oocyte aging are the predominantly marine worms belonging to the phylum Nemertea [12]. Approximately 1500 species of nemerteans, or “ribbon worms,” have been described, and most of these live near seashores where they prey on other invertebrates by means of an eversible proboscis [13]. Sexes are typically separate in nemerteans, and each ripe adult usually forms multiple gonads that discharge their gametes freely into surrounding seawater during the breeding season [14]. Previously, the roles of Cdc2 kinase and several kinds of mitogen-activated protein kinases have been analyzed in aging nemertean oocytes [15, 16]. To promote the use of nemerteans in further embryological analyses, this contribution supplements a previous protocol that outlines immunoblotting analyses of maturing nemertean oocytes [17] by adding information regarding the collection or purchase of North American species and practical tips for handling oocytes during aging experiments.

2

Materials

2.1 General Solutions

2.2 SDS– Polyacrylamide Gel Components

Solutions should be mixed with high-quality (e.g., MilliQ) water, and unless stated otherwise, prepared fresh from newly dissolved reagents or from stocks maintained at either room temperature or 4  C (see specific comments accompanying the various recipes). 1. Lysis buffer: 100 μl 0.5 M HEPES, pH 7, 750 μl 0.5 M NaCl, 1250 μl 2% NP-40, 150 μl 1 M β-glycerophosphate, 150 μl 0.25 M EGTA, pH 8, and 100 μl 1 M NaF. Just before vortexing and subsequent use on samples, add to the above the following: 4.5 mg sodium orthovanadate, 25 μl 100 mM PMSF (from aliquots dissolved in DMSO and stored at 20  C), and ¼ tablet of Complete Protease Inhibitor Mini

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Tablet (Roche Diagnostics, Mannheim, Germany). Combine all nine components listed above and vortex for 2 min; let sit on ice for 10 min and transfer fluid component to microcentrifuge tube to avoid froth on top. Formulation is based on a recipe from reference 13, using stocks stored at 4  C, and makes 2.5 ml (i.e., enough for at least 50 protein-assay samples). 2. 10% resolving gel: 16.4 ml MilliQ water, 13.2 ml premixed acrylamide solution (37:5:1, Bio-Rad Corp., Hercules, CA, #161–0158), 10 ml 1.5 M Tris–HCl resolving buffer, pH 8.8, 400 μl 10% SDS (w/v), 200 μl 10% ammonium persulfate (freshly made), and 20 μl TEMED. In a side-arm Erlenmeyer flask, add solutions in the order listed and gently swirl ~10 times in each direction to mix without generating bubbles. 3. 4% stacking gel: 12.2 ml MilliQ water, 2.6 ml premixed acrylamide solution (37:5:1, Bio-Rad Corp., Hercules, CA, #161–0158), 5 ml 0.5 M Tris–HCl stacking buffer, pH 6.8, 200 μl 10% SDS (w/v), 100 μl 10% ammonium persulfate (freshly made), and 20 μl TEMED. Mix as described above for the resolving gel. The above “quadruple-batch” gel recipes make enough monomer for four 4%/10% gels that can be used to separate various proteins in the 20–250 kD molecular weight (MW) range when the four gels are run simultaneously as two pairs connected to two electrophoresis power packs. For running just two gels, halve the amounts, and when analyzing relatively high- or low-MW proteins that are not suitably separated with such a 4/10 formulation, gel concentrations can be altered to optimize protein separations (see Bio-Rad Instruction Manual for Mini Protean Tetra Cell, for further details). Note: acrylamide is toxic and should be handled with care. Allow leftover monomer mixes to polymerize overnight so that the less toxic solid form of the gel can be disposed more safely. 4. Sample buffer: 3.55 ml MilliQ water, 1.25 ml 0.5 M Tris–HCl, pH 6.8, 2.5 ml glycerol, 2 ml 10% SDS (w/v), and 0.5% 0.2 ml Bromophenol Blue (w/v). This formulation is based on a recipe in ref. 14, using stock solutions stored at room temperature. Just before use on samples, add appropriate amount of DTT (see Subheading 3.5). 5. 10 Running buffer: 30.3 g Tris, 144 g glycine, and 10 g SDS. Bring all three to 1 l in MilliQ water. Stock solution is stored at room temperature; 1 working solutions can also be used after several months of storage at 4  C. 2.3 Transfer Components

1. Transfer buffer: 3.03 g Tris, 14.4 g glycine, 800 ml MilliQ water, and 200 ml methanol (spectroscopy grade or higher quality). After mixing the above on a stirrer, add 0.5 g SDS

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and stir for at least an additional 15 min. For each run, make 1 l for 2 transfers or 2 l for 4 transfers. 2. 10X TBS (Tris-buffered saline) washing solution: 80 g NaCl, 24.2 g Tris, and ~800 ml MilliQ water. Dissolve and pH to 7.6 with concentrated HCl before bringing up total to 1 l. Stock solution is stored at room temperature. Make at least 250 ml of 1 working solution for each run. 3. TTBS (Tween–Tris-buffered saline): to each liter of 1 TBS solution, add 500 μl of Tween 20 and stir for at least 15 min. Make 2 l of TTBS for each run. Stored at room temperature, this TTBS working solution is used for all washes of membranes except for the last (plain TBS) washes before conducting the ECL protocol. 4. Blocking solution: 5% nonfat dry milk freshly dissolved in TTBS. 5. Cold stripping buffer (made fresh just before use): 3 g glycine, 0.2 g SDS, and 80 ml MilliQ water. Put on stirrer and pH to 2.2 with 6 N HCl (takes ~6 ml). Bring up to 98 ml in a graduated cylinder and add 2 ml Tween 20. Mix up and down numerous times to dissolve the Tween. 2.4 Supplies to Take to the Darkroom When Developing Film

1. Clear plastic wrap (e.g., Saran wrap). 2. White reflective screen (Fold an 8.5  11 in. white paper in half, and have the 5.5  8.5 in. doubled sheet laminated at a print shop. Make several screens for multiple ECL incubations). 3. Forceps. 4. Laboratory tape. 5. Plastic transfer pipettes. 6. A book with a black nonreflective cover. 7. Timer. 8. Sharpie marker. 9. X-ray film, 5  7 in for mini-blots (Sterling X-ray film from Life Science Products, Frederick, CO has better sensitivity and costs less than comparable Kodak film). 10. Enhanced chemiluminescence solution (keep in dark and combine the two components of the ECL in the darkroom immediately before use on the blots).

3

Methods

3.1 Maintaining Adult Specimens

For many nemertean species, intact adults or large body pieces can survive at least several weeks in the laboratory if maintained in continuously flowing natural seawater (NSW) or if supplied with

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at least daily exchanges of NSW or artificial seawater (ASW). Worms should be kept at temperatures typically encountered in natural habitats either by use of cooled circulating SW or in regularly refreshed containers of SW maintained in incubators or cooling baths. Storage containers for adult worms should be of embryological grade (“E ware”) glass or plastic that had not held fixatives or harsh solvents, and if purchased new, the containers should ideally be preconditioned for several days in NSW to generate a beneficial microbial coating. Depending on the particular size of the worm, adults kept in storage containers should be maintained in uncrowded conditions (e.g., ~200–1000 ml SW/ adult), and every few days, the container should be rinsed in fresh water to avoid contamination from waste products and the fragments of the worm that can accelerate further degradation. 3.2 Obtaining Gametes and Fertilized Embryos

With small adults (e.g., Maculaura alaskensis), gametes can be obtained by puncturing ripe specimens with jeweler’s forceps to release sperm or oocytes from the many dozens of serially repeated gonads occurring along the length of the body, whereas similar methods can be used on ~0.5–1-cm-long pieces cut from larger specimens (e.g., Cerebratulus), which in turn allows a single worm to be used multiple times for experiments. Ranging in size from ~70–200 μm in most nemertean species, fully grown oocytes stripped from ovaries typically lack surrounding follicle cells and are initially arrested at the prophase-I stage of meiosis, as indicated by their prominent GV. Following incubation in either NSW or calcium-containing ASW, prophase-I specimens usually begin to mature by initiating GVBD within ~30–60 min at 12–15  C. Subsequently, post-GVBD oocytes usually arrest at metaphase I of meiosis within 1–2 h of GVBD onset and can be reliably fertilized for up to several hours after maturing to metaphase I. For in vivo imaging of membrane-impermeable probes, such as dextran-conjugated calcium ion indicators, oocytes are best microinjected as prophase-arrested specimens or soon after GVBD has begun [18]. Alternatively, maturation can be reversibly prevented if oocytes stripped from ovaries are rapidly transferred to ice-cold calcium-free seawater (CaFSW) or NSW solutions containing various pharmacological agents, including inhibitors of MPF activation and some blockers of MAPKs [19–24]. Depending upon the particular maturation inhibitor that was used, immature oocytes can be kept for up to 1–2 days before being rinsed in several changes of NSW and thereby triggered to mature. Regardless of whether GVBD is initiated directly after removal from the ovary or following a prolonged prophase arrest, the jelly coats surrounding oocytes can be mechanically removed to optimize packing during subsequent steps in processing [18]. Moreover, all mature oocytes should be gently washed in three changes in SW prior to fertilization.

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Sperm used in inseminations should be quickly collected by pipetting from the surface of punctured males and can be kept undiluted for up to a day at 4  C. Just prior to fertilization, sperm needs to be diluted in SW to obtain concentrations that yield ~5–50 sperm around each inseminated oocyte. Zygotes are routinely kept in loosely arranged monolayers to avoid anoxia, and once cleavages begin, the overlying seawater should be changed on a daily basis. Depending on species and incubation temperature, embryos tend to remain stationary and easily observed for about a day postfertilization before ciliated stages begin to swim toward the top of the culture dish. 3.3 General Cell Biological Applications

Once fully grown oocytes are obtained from ripe females, such material can be subjected to various cell biological protocols according to standard methods that are described in detail elsewhere. For example, microinjection of nemertean oocytes with calcium-sensitive fluorescent dyes and the subsequent monitoring of dye-loaded specimens by confocal imaging [25].

3.4 Incubating Oocytes During Aging Experiments

In order to minimize aging-independent variability, control oocytes vs. drug-treated specimens need to be incubated over the duration of each experiment using similar temperature and illumination levels. Aging oocytes should also be spread throughout the container surface in nonpatchy monolayers to help avoid subpopulations that are subjected to substantially different cellular densities. In addition, it may be possible to extend the viability of oocytes considerably by adding antibiotics to the surrounding SW, according to methods successfully used on sea urchin eggs [26, 27]. Once the appropriate culturing conditions have been established for aging experiments, kinase activities can be monitored via the following immunoblotting protocol that does not require specialized equipment other than a darkroom and an image-processing system for conducting densitometric quantification of band intensities.

3.5 Sample Processing for Immunoblotting Analyses

1. Make lysis buffer as directed in Subheading 2. Add 36 μl to each sample and vortex for 25 s so that froth is visible in each tube. Centrifuge samples at 4  C and 14,000 rpm for 12 min in a microcentrifuge. Maintain samples throughout the procedure on ice and after protein assays have been completed, store at 20  C (see Note 1). 2. After centrifugation, transfer 2 μl of the supernatant from each sample into a cuvette containing 798 μl of MilliQ water for the protein assay. Set aside the cuvettes and continue processing the samples as described below. 3. After removing the 2-μl aliquot for a protein assay, add 26 μl of freshly made sample buffer (850 μl of stock solution from Subheading 2 plus 150 μl of 1 M DTT added just before use)

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to the remaining 34 μl of lysed sample. Mix by briefly vortexing and snap spin the tubes to bring down the sample. Boil the tubes for 4 min and subsequently store them at 20  C (see Note 2). 4. After boiling the samples in sample buffer, add 200 μl of Bradford reagent (Bio-Rad, Hercules, CA) to each cuvette. Mix gently to avoid excessive bubbles by pipetting up and down with a plastic transfer pipette until the solution is a uniform color (see Note 3). 5. Read the absorption of each cuvette at 595 nm, using a cuvette containing 798 μl of MilliQ water, 2 μl of lysis buffer, and 200 μl of Bradford reagent as the blank. 6. To convert absorbance into total protein per sample, construct a standard curve based on serial dilutions of bovine serum albumin. 3.6

Gels

1. Use either precast gels (e.g., Bio-Rad Mini-PROTEAN TGX) or pour gels as follows: first, prepare resolving gel solution according instructions in Subheading 2 (see Note 4). 2. Using a pipette, add the gel fluid slowly and steadily to the side of the glass plates, to avoid introducing bubbles. In order to leave room for the stacking gel, the resolving gel should extend to 1 cm below the level of where the comb of the wells will protrude once inserted into the overlying stacking gel (see Note 5). 3. After the gel has been poured, gently add a MilliQ water cap until the water reaches the top of the short plate. Let the gel polymerize for at least 45 min at room temperature (see Note 6). 4. Following gel polymerization, pour out the water cap by tilting the glass plates, and use a paper towel to remove the remaining water without touching the gel. 5. Mix the stacking gel according to the recipe in Subheading 2 and pipette it on top of the resolving gel, filling to the top of the short glass plate. To avoid squirting out liquid acrylamide, tilt the comb and slowly insert from one side rather than rapidly putting it in straight down on the stacking gel. 6. Let stacking gel polymerize for 45 min, then cover with plastic wrap and keep in the refrigerator for use within the next few days. 7. Mix up 1 running buffer and store in refrigerator overnight so it will be cold when used.

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Fig. 1 Proper assembly of the gel apparatus showing downward pressure while clamping with thumbs to minimize chances of leakage 3.7

Electrophoresis

1. Thaw frozen samples that had been boiled in sample buffer and vortex them briefly to mix. 2. Place the gel into its holder ensuring there is a good seal (Fig. 1). Once assembled, put the holder with its two gels in a gel rig and add cold running buffer (see Subheading 2) into the central area between the two sets of gels. Pour remaining running buffer outside the gels to fill the container (see Note 7). 3. Based on the protein assays, load the equivalent of 25 μg of total protein per well using a gel-loading pipette with its tip placed against the edge of the short glass to ensure that the fluid flows into the well. It is important not to overload the wells. To determine well capacities in gels of varying thicknesses, consult the Bio-Rad Instruction Manual for Mini Protean Tetra Cell. Fill all wells; if one is to be left blank, add sample buffer. Also, add molecular weight markers to one or two of the lanes (see Note 8). 4. Plug the gel apparatus into an appropriate power supply that is set to a constant voltage of 100 V and run the gel until the dye front is ~0.5 cm from the bottom (about 2–2.5 h) (see Note 9).

3.8

Transfer

1. After electrophoresis has been completed, turn off the power supply and carefully separate the gel plates. Using a clean razor blade, remove the stacking portion of the gel and rinse the remaining resolving gel in MilliQ water for a few seconds.

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2. After rinsing, equilibrate the gel in transfer buffer (see Subheading 2) for 15 min on an orbital shaker. 3. While the gel is equilibrating, manually agitate for 90 s in methanol PVDF membranes (Bio-Rad, Hercules, CA), which had been precut to ~5.5  8.5 cm rectangles for use in a miniblot apparatus. Discard the methanol and equilibrate the membranes in transfer buffer for 15 min on an orbital shaker. 4. Load the gels into the transfer setup in the correct order. For example, with Bio-Rad mini-blot transfer setups, start with the black side of holder on bottom, followed by sponge, filter paper, gel, membrane, filter paper, sponge, and clear side of holder. It is important to assemble this “sandwich” submerged in transfer buffer in order to minimize trapped bubbles. To maintain the same order of the gel lanes on the blot, flip the gel horizontally so that the last lane of the gel is switched to the left side of the overlying membrane and thus will be on the right once the transfer is complete and the membrane is flipped over. 5. Place the loaded sandwich into the transfer apparatus with the black side of the sandwich oriented toward the back of the chamber. Add a magnetic stir bar and place the transfer unit on a stirrer. Put in an ice pack and fill the unit with transfer buffer until the sandwich is completely covered. 6. Turn on the stirrer, snap on the lid, and run the transfer at 100 constant volts for 75 min. For two blots, the current should start around 230–270 mA and will gradually rise to 300–380 mA by the end of the run (see Note 10). 3.9

Antibody Probing

1. When the transfer is complete, peel away the membrane (you should see the molecular weight markers on the membrane). If probing with multiple antibodies for proteins of divergent MWs, it may be possible to cut the blot into horizontal strips that can be incubated individually in smaller containers (Fig. 2). In any case, with full or cut blots, put the membrane directly in blocking solution (see Subheading 2) with the side that had been next to the gel face-up. Shake at room temperature on an orbital shaker for 60 min. Do not block for shorter or longer periods as this might result in increased background or reduced signal (see Note 11). 2. Briefly wash the blocked membrane with TTBS (see Subheading 2) and add the primary antibody solution. Cover and agitate overnight at 4  C (see Note 12). 3. Decant the primary antibody solution and retain for future use after freezing at 20  C. Rinse the probed membranes with TTBS for 30 s; then, wash 3 times with minimal agitation for 5 min each in TTBS at room temperature (see Note 13).

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Fig. 2 A topside and oblique view of an incubation chamber with a glued-down glass slide divider allowing for two blot strips to be probed simultaneously

4. d) After washing, incubate with secondary antibody dissolved in blocking solution for 90 min at room temperature (see Note 14). 5. Remove the secondary antibody, rinse for 30 s in TTBS and wash twice for 10 min each in TTBS. After washing in TTBS, perform 2 15-min washes in TBS (see Note 15). 3.10

Film Developing

1. In addition to the probed membranes incubating in TBS, take to the darkroom items listed in Subheading 2, and set up two pieces (~6  8 in.) of clear plastic wrap. Place the washed blots face-up on one sheet of wrap and quickly incubate them for 90 s in the ECL solution. It is important to make sure that there are no dry spots on the blots. 2. Remove the ECL solution and place the membranes face-down on the other sheet of wrap. Put a white reflective screen on top of the membranes and neatly fold the wrap around the back of the screen before taping down the wrap. Make sure no bubbles or leaks are present. 3. Turn off the room lights and turn on red safety lights. Take out unexposed film, keeping it a safe distance from the safety light, and place it on the ECL-loaded blots. Using a Sharpie, quickly make several small marks that extend across the edge of the film to the reflective sheet. This will allow subsequent orientation of the developed film relative to the molecular weight markers. 4. Cover with a book and expose. Exposure times vary depending on the intensity of the signal but are typically in the range of 15 s to 5 min (see Note 16). 5. Place the film in an automated film processor or develop the film manually using Kodak D19 developer and fixer according to the manufacturer’s instructions.

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1. Keep the membranes in the refrigerator still wrapped in plastic on their white reflective screens in order to avoid drying of the blots. For reprobing, remove the blots and quickly place them in stripping buffer (see Subheading 2) with agitation for 30 min. Replace with fresh stripping buffer and agitate for another 30 min. 2. Give the membranes three washes with TTBS for 15 min each. 3. Incubate the membranes in freshly made blocking solution for 1 h, transfer to primary antibody solution, and repeat the rest of the protocol as described above to get a second set of signals from the blot (see Note 17).

3.12 Quantification of Blot Intensities

1. Create a digital image of the blot signal that was recorded on film. 2. Invert the contrast so that background is dark, and the blot bands are light (this will allow higher-density bands to be quantified as higher numbers, as compared to lower-density bands) (Fig. 3). 3. Place a region of interest around each band to be quantified and obtain an integrated intensity for that band using morphometric software (e.g., MetaMorph, Molecular Devices, Sunnyvale, CA). 4. After obtaining band densities, shift each region of interest to a background region of the film and obtain a background reading that is to be subtracted from the band intensity

Fig. 3 Example of an original version of a blot (top image) and its inverted-contrast view (lower image) that, in turn, allows for high-density bands to be recorded as high values during densitometric analyses

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(thereby accounting for opacity in the film itself). Such quantification assumes that the blot signals are recorded in the linear range of the film and are not either saturated or below a threshold detection value for the film.

4

Notes 1. To adapt this protocol for nonoocyte cells or tissues, it may be possible to modify the lysis buffer and/or protein extraction steps. For example, longer incubations in lysis buffer or more stringent extractions employing sonication might be tried. 2. When boiling samples, set the Bunsen burner/hot plate to the minimum required to obtain weak boiling, thereby reducing the chance of getting water into the sample should a tube pop open. 3. The same pipette can be used for more than one sample, as long as very little, if any, fluid is transferred from one cuvette to the other. Also, the Bradford reagent loses signal over time, so the faster the readings are completed, the better. 4. To mix the gel solution, gently swirl the container; if bubbles continue to appear in the blots, try degassing gel solutions for 10 min in a vacuum prior to addition of the polymerizing agents (ammonium persulfate and TEMED). 5. To determine the appropriate height of the resolving gel prior to gel pouring, place a comb within the gel plate setup and mark 1 cm below the bottom of the comb. 6. Pour the water cap slowly to avoid mixing with the gel solution, which in turn can result in a curved gel front. 7. Smaller quantities of running buffer can be used (e.g., as little as 450 ml), but ~850 ml per gel setup ensures that any leakage from the central chamber will not lead to a significant current decrease and an ensuing stoppage of the gel run. 8. To reduce the amount of antibody solution needed, it is possible to cut the blot into strips that contain the proteins of the desired molecular weight for each antibody. These strips can fit in containers that require as little as 8 ml to cover the blots entirely (Fig. 2). In order to help cutting horizontal strips for multiple antibody incubations, load two lanes of MW markers with one near either end of the gel. 9. The current at the beginning of the run should be around 45–55 mA and by the end of the run, about half of starting value. Because of this decrease, the E9 safety feature of certain Bio-Rad power supplies should be disabled prior to the start of the run to avoid having the power turn off as the current drops.

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10. It is possible to regulate the current by modulating the speed of the magnetic stirrer. Faster stirring will result in lower currents; adjust the speed until you reach the desired values. 11. Thoroughly mix the milk into TTBS at least 30–60 min prior to blocking in order to avoid uneven blocking. 12. Depending upon the strength of the primary antibody, try initial dilutions of 1:2500 to 1:500 in TTBS containing 1–5% bovine serum albumin as a blocking agent. For full blots, ~18 ml of solution are required to cover the entire blot (see Note 8). 13. It is possible to save some primary antibody solutions (e.g., phospho-ERK 1/2, phospho-Cdc2 Y15, phospho-AMPK T172) and freeze them at 20  C for subsequent use on another blot. If the signal begins to fade, boost the solution by adding a few more microliters of concentrated antibody. 14. Following incubation in primary antibody, an appropriate secondary antibody conjugated to horseradish peroxidase (e.g., from Santa Cruz Biotechnology, Santa Cruz, CA). Initially try a 1:5000 dilution in blocking solution and adjust accordingly. 15. TTBS should be avoided in the final washes in order not to degrade the ECL signal. 16. Keep the book still over the blot and film in order to avoid blurring the signal. Use a book with a black nonreflective cover. The ECL solution fades over time, with optimal results being achieved within 25 min of mixing. Because of this, later exposures take longer to get equivalent results; so, whenever possible, do longer exposures first. 17. Strong bands usually do not get fully stripped. Thus, it is better to probe for a weak band first, and then probe the stronger band after stripping. References 1. Yuce O, Sadler KC (2001) Postmeiotic unfertilized starfish eggs die by apoptosis. Dev Biol 237:29–44 2. Fissore RA, Kurokawa M, Knott J, Zhang M, Smyth J (2002) Mechanisms underlying oocyte activation and postovulatory ageing. Reproduction 124:745–754 3. Miao Y-L, Kikuchi K, Sun Q-Y, Schatten H (2009) Oocyte aging: cellular and molecular changes, developmental potential and reversal possibility. Hum Reprod Update 15:573–585 4. Tiwari M, Prasad S, Tripathi A, Pandey AN, Ali I, Singh AK, Shrivastav TG, Chaube SK (2015) Apoptosis in mammalian oocytes: a review. Apoptosis 20:1019–1025

5. Kikuchi K, Izaike Y, Noguchi J, Furukawa T, Daen FP, Naito K, Toyoda Y (1995) Decrease of histone H1 kinase activity in relation to parthenogenetic activation of pig follicular oocytes matured and aged in vitro. J Reprod Fertil 105:325–330 6. Kikuchi K, Naito K, Noguchi J, Shimada A, Kaneko H, Yamashita M, Aoki F, Tojo H, Toyoda Y (2000) Maturation/M-phase promoting factor: a regulator of aging in porcine oocytes. Biol Reprod 63:715–722 7. Wu B, Ignotz G, Currie BW, Yang X (1997) Dynamics of maturation-promoting factor and its constituent proteins during in vitro

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maturation of bovine oocytes. Biol Reprod 56:253–259 8. Ono T, Mizutani E, Li C, Yamagata K, Wakayama T (2011) Offspring from intracytoplasmic sperm injection of aged mouse oocytes treated with caffeine or MG132. Genesis 49:460–471 9. Sasaki K, Chiba K (2004) Induction of apoptosis in starfish eggs requires spontaneous inactivation of MAPK (extracellular signal-regulated kinase) followed by activation of p38 MAPK. Mol Biol Cell 15:1387–1396 10. Sadler KC, Yuce O, Hamaratog˘lu F, Verge´ V, Peaucellier G, Picard A (2004) MAP kinases regulate unfertilized egg apoptosis and fertilization suppresses death via Ca2+ signaling. Mol Reprod Dev 67:366–383 11. Ebeling S, Labudda A, Meinecke B (2010) In vitro ageing of porcine oocytes: changes in phosphorylation of the mitogen-activated protein kinase (MAPK) and parthenogenetic activability. Reprod Dom Anim 45:e398–e404 12. Kvist S, Laumer CE, Junoy J, Giribet G (2014) New insights into the phylogeny, systematics, and DNA barcoding of Nemertea. Invertebr Syst 28:287–308 13. Stricker SA, Cloney RA (1983) The ultrastructure of venom-producing cells in the nemertean Paranemertes peregrina. J Morphol 177:89–107 14. Stricker SA, Smythe TL, Miller LA, Norenburg JL (2001) Comparative biology of oogenesis in nemertean worms. Acta Zool 82:213–230 15. Stricker SA, Beckstrom B, Mendoza C, Stanislawski E, Wodajo T (2016) Oocyte aging in a marine protostome worm: the roles of maturation promoting factor and extracellular signal regulated kinase form of mitogenactivated protein kinase. Develop Growth Differ 58:250–259 16. Stricker SA, Ravichandran N (2017) The potential roles of c-Jun N-terminal kinase (JNK) during the maturation and aging of oocytes produced by a marine protostome worm. Zygote 25:686–696 17. Escalona JR, Stricker SA (2014) Immunoblotting analyses of changes in protein phosphorylations during oocyte maturation in marine nemertean worms. In: Carroll DJ, Stricker SA (eds) Methods in molecular biology:

developmental biology of the sea urchin and other marine invertebrates. Humana Press, New York, pp 237–247 18. Stricker SA, Silva R, Smythe T (1998) Calcium and endoplasmic reticulum dynamics during oocyte maturation and fertilization in the marine worm Cerebratulus lacteus. Dev Biol 203:305–322 19. Smythe TL, Stricker SA (2005) Germinal vesicle breakdown is not fully dependent on MAPK activation in maturing oocytes of marine nemertean worms. Mol Reprod Dev 70:91–102 20. Stricker SA, Smythe TL (2006) Differing mechanisms of cAMP- versus seawater-induced oocyte maturation in marine nemertean worms II. The roles of tyrosine kinases and phosphatases. Mol Reprod Dev 73:1564–1577 21. Stricker SA (2009) Roles of protein kinase C isotypes during seawater- versus cAMPinduced oocyte maturation in a marine worm. Mol Reprod Dev 76:693–707 22. Stricker SA (2011) Potential upstream regulators and downstream targets of AMP-activated kinase signaling during oocyte maturation in a marine worm. Reproduction 142:1–12 23. Stricker SA (2012) Inhibition of germinal vesicle breakdown by antioxidants and the roles of signaling pathways related to nitic oxide and cGMP during meiotic resumption in oocytes of a marine worm. Reproduction 143:261–270 24. Stricker SA, Escalona JR, Abernathy S et al (2010) Pharmacological analyses of protein kinases regulating egg maturation in marine nemertean worms: a review and comparison with mammalian eggs. Mar Drugs 8:2417–2434 25. Stricker SA (2000) Confocal microscopy of intracellular calcium dynamics during fertilization. BioTechniques 29:492–498 26. Epel D, Vacquier VD, Peeler M, Miller P, Patton C (2004) Sea urchin gametes in the teaching laboratory: good experiments and good experiences. Methods Cell Biol 74:797e823 27. Kiyomoto M, Hamanaka G, Hirose M, Yamaguchi M (2013) Preserved echinoderm gametes as a useful and ready-to-use bioassay material. Mar Environ Res 93:102–105. https://doi.org/10.1016/j.marenvres.2013. 08.014

Chapter 9 Recovery of Sea Star Egg Cell Surface Proteins Released at Fertilization Emily Wiseman, Lauren Bates, and David J. Carroll Abstract To provide a better understanding of the composition of the egg cell membrane, we describe a method in which proteins and peptides that are either naturally released by the egg or cleaved by sperm proteases can be collected, analyzed, and identified. Such molecules are captured and isolated from the surrounding seawater via biotinylation, before being concentrated by an affinity interaction and subsequently analyzed by western blotting and mass spectrometry. Key word Signal transduction, Fertilization, Egg activation, Cell surface, Biotinylation, Blotting

1

Introduction Fertilization has long been a challenging field to study due, in part, to the evolutionary divergence of many of the proteins needed for gamete recognition, binding, fusion, and activation [1]. Such a high evolutionary rate of change is a contributor to species variation, which is necessary for diversity; however, this also makes it difficult to find the elusive membrane orthologs that are critical for fertilization [2]. In fact, given the long-term interest in fertilization mechanisms, particularly within echinoderms (e.g., sea urchins and starfish), our understanding of the molecular signaling cascade at the moment when the sperm and egg meet remains surprisingly meager. One of the more important discoveries about the molecular events of echinoderm fertilization was the finding that the egg generates a calcium wave that is critical for restarting the cell cycle and initiating the slow block to polyspermy [3–7]. How this calcium wave is triggered has only been partially elucidated. In essence, calcium is released from internal stores in the endoplasmic reticulum (ER) via the hydrolysis of phosphatidylinositolbisphosphate (PIP2) by phospholipase C (PLC). Such hydrolysis yields diacylglycerol (DAG), which remains in the membrane, and

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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inositol triphosphate (IP3), which binds to the IP3 receptor on the ER to release the Ca2+ stores. Up to this point, the pathway seems to be well conserved in most animals, including mammals and echinoderms; however, the isoform of PLC that hydrolyzes PIP2 and how PLC is activated varies depending on the species. In mammals, phospholipase C zeta (PLC ζ) is supplied from the sperm cytosol and produces IP3 following sperm–egg fusion [6, 8, 9]. In ascidians and echinoderms, the PLC-gamma (PLCγ) isoform in the egg is activated after sperm and egg interaction by Src-family kinases (SFKs) via phosphorylation [10]. How the SFKs are activated is less well understood. One theory is that surface receptors may send a signal to activate the pathway, but little is known about the cell surface of gametes and what might send the activating signal. So far, only a handful of gamete surface proteins have been identified. These include the mammalian sperm protein Izumo1 and egg protein Juno [11], whose interactions are considered to be critical for successful fertilization [12, 13]. Once the egg is fertilized, Juno is released from the surface in vesicles rather than endocytosed. It is hypnotized that these Juno enriched vesicles are shed in an effort to generate “phony eggs” that bind free acrosome reacted sperm, thereby decreasing the number of sperm that bind the actual egg which may serve as another potential mechanism garnishing a more robust block to polyspermy [14]. In Xenopus, a protein called Uroplakin III is a Src substrate localized to lipid rafts within egg cell surface. Uroplakin III may be necessary for fertilization [15], since following disruption of lipid rafts, the efficiency of fertilization is decreased [16, 17]. In C. elegans, two proteins, EGG1 and EGG2, possess LDL extracellular receptor domains, a transmembrane domain, and a small cytoplasmic tail, all of which may play a role in signaling [18]. In echinoderms, two proteins have been discovered—the aptly named sperm protein, Bindin, and egg protein, Egg Bindin Receptor 1 (EBR1) [19, 20]. All of these above-mentioned molecules, however, seem to be necessary only for gamete binding or recognition rather than for signal cascades involved in calcium release. In echinoderms, eggs can be activated by proteases without sperm being present [21], and as a way of identifying potential surface proteins critical for fertilization, various exogenous proteases have been used to mimic the effects of proteases naturally produced by sperm. For example, eggs that are treated with a protease, called lysyl endoproteinase C (LysC), which cleaves cell surface proteins, release a 70 kDa fragment from a larger 350 kDa protein [22]. Although found to not be required for fertilization, pretreating the eggs with LysC prior the addition of sperm, lowered the fertilization efficiency [19, 22]. A nonchemical approach to visualize what is released at fertilization can be obtained through the use of biotin labeling.

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Fig. 1 (a) Confocal image of a live biotinylated egg confirms that the biotin does not cross the egg cell membrane and only labels the surface. Sample was biotinylated then incubated with Streptavidin conjugated with DyLight 488 and viewed with a confocal microscope. (b) As a control, live unbiotinylated oocyte was incubated with Streptavidin conjugated with DyLight 488 and viewed by confocal microscopy to identify natural fluorescence at this wavelength

Biotinylation labels free amines on the surface of cells, and following such labeling, does not cross the cell membrane [23] (Fig. 1). The biotinylation method is one of the more useful approaches to separate membrane proteins from cytosolic proteins, which tend to be significantly overrepresented compared to membrane proteins [24, 25]. Biotinylation of the egg cell membrane has been used before as well [26]; however, the vast number of proteins that can be pulled down can be challenging to wade through to find any that may be vital for fertilization. This can be addressed, in part, by looking at what is released into the seawater. After fertilization, anything that is biotinylated and released into the seawater can be pulled down using an avidin affinity interaction and either visualized using western blotting (Fig. 2) or identified via tandem mass spectrometry. Using the Pacific bat sea star Patiria miniata as a model organism, this chapter outlines a way to help identify essential membrane proteins involved in fertilization by analyzing biotinylated peptides that are released into seawater postfertilization

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Fig. 2 To see if there are any the peptides that are released naturally or cleaved via sperm proteases at fertilization, eggs were matured, biotinylated, fertilized, then spun down to transfer the seawater into a new tube. The seawater samples were then frozen in liquid nitrogen at their appropriate time point to terminate all proteolytic function. Once thawed and spun down to remove any egg particulate, the biotinylated proteins were isolated from the seawater using a Streptavidin Mag Sepharose affinity interaction then analyzed via western blotting using streptavidin-HRP. “UF” indicates unfertilized samples, the numbers indicate fertilized samples and how many minutes after the introduction of sperm that the sample was frozen in liquid nitrogen. Molecular weight marker is in kDa

naturally either by the egg or by cleavage mediated by a sperm protease. This animal is an ideal model organism for this kind of discovery-based science as they have a vast number of large nearly transparent oocytes and eggs, which are all meiotically synchronized and are naturally fertilized in vitro [27].

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Materials

2.1 Obtaining and Culturing Oocytes/ Testes

1. P. miniata can be shipped overnight from Marinus Scientific LLC (Long Beach, CA) and in a seawater tank with circulation and filtration at 17  C. 2. 1-Methyladenine from Fisher Scientific (item number: AC201310250). Prepare a 5 mM stock in diH2O (see Note 1) and aliquot into 20 μl divisions and store at 80  C for long term storage or at 20  C for 6 months. 3. A 3 mm sample corer with plunger (item 18035-02), 10 cm serrated curved Graege forceps (item number 11051–10), and Noyes spring scissors (item number 15012–12) can be

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purchased from Fine Science Tools, Inc. (https://www. finescience.com/en-US/). 4. Nylon mesh filter sheet, 210 μm pore size can be purchased from BioQuip Products, Inc. (https://www.bioquip.com/). 5. Miscellaneous tools, such as 50 ml glass beakers, Pasteur pipets, Eppendorf tubes, conical tubes, etc., can be purchased from your supplier (see Note 2). 6. To keep samples at appropriate temperature, we use a small tabletop wine cooler without the rack set at 16  C. 2.2

Biotinylation

1. EZ-Link™ Sulfo-NHS-LC-Biotin (ThermoFisher A39257). Remove from freezer and allow it to reach room temperature before opening (see Note 3). 2. Glycine at a 0.5 M concentration in filtered natural seawater (FNSW) solution. This should be kept at 16  C and will be usable for a month.

2.3 Streptavidin– Biotin Affinity Interaction

1. Streptavidin Mag Sepharose from Sigma-Aldrich (product number: 28–9857-38) stored at 4  C. 2. Binding buffer known as Tris-buffered saline (TBS), 50 mM Tris–HCl, 150 mM NaCl, pH 7.5. TBS can be kept at room temperature; however, the life span can be prolonged if kept at 4  C. 3. Washing buffer which is standard TBS with 2 M urea added as well. 4. Elution buffer is 2% SDS in nuclease-free water. This solution should be stored at room temperature as the SDS falls out of solution at 4  C.

2.4 SDS-PAGE Gel and Western Blotting

1. Pierce™ High Sensitivity Streptavidin-HRP (ThermoFisher 21,130) kept at 4  C. The probe is sensitive enough to be used at 1:10,000 dilution in blotto. 2. Tris-based Saline Solution with Tween 20 (TBST). Final concentrations: Tris-HCl 20 mM, pH 7.6, 150 mM NaCl, and 0.1% Tween 20 (see Note 4). TBST is used for washes on the western blots as well as for making blotto. It is recommended to make a liter at a time. Make sure to pH the solution prior to adding the Tween 20. 3. Nonfat dry milk, biotin-free (LabScientific M-0841) (see Note 5). Blotto is TSBT with 5% of the dried milk mixed well. 4. Clarity™ Western ECL substrate (Bio-Rad 1705060S). 5. Nitrocellulose 0.45 μm (Bio-Rad 1620115). The roll can be cut into 8 cm  6.5 cm rectangles to fit the SDS-PAGE gel.

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Miscellaneous

1. Liquid nitrogen is needed to flash freeze samples at specific times to stop cellular processes and proteolytic activity. 2. Standard light microscope to check the quality of eggs or oocytes, and to check the success of fertilization.

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Methods

3.1 Oocyte/Sperm Removal and Preparation

1. Remove gonads through a 3 mm hole made by a punch biopsy tool in the aboral side of the sea star. Testes, which are white, should be transferred to a clean, dry 1.7 ml tube and kept on ice. The orange or yellow ovary should be transferred to a 50 ml beaker containing FNSW. 2. Mince the ovaries using spring microdissection scissors to release the oocytes into the seawater. 3. Swirl the oocytes and ovary pieces to resuspend; then pour through a 210 μm nylon mesh filter into a new 50 ml beaker to separate the ovary pieces from the oocytes. The oocytes will pass through the filter into the new beaker. 4. Allow the oocytes to settle by gravity, replace as much seawater as possible with fresh FNSW. Repeat with rinses three times. Oocytes are maintained at 16  C. 5. Check a small aliquot of oocytes for quality and uniformity by light microscope. Healthy oocytes should be approximately 180–190 μm in diameter. The germinal vesicle should be large and be visible as well as the nucleolus. 6. Create a 10% (v/v) slurry of oocytes and place in a clean 50 ml beaker. Add 1-MA to a final concentration of 3–5 μM. Swirl to resuspend the oocytes. Every 5 min take a small aliquot to view under the microscope. An oocyte is considered mature when germinal vesical has broken down entirely and can no longer be seen. This process typically takes anywhere from 45 min to 1 h at 16  C. 7. Rinse the mature oocytes (eggs) three times with FNSW.

3.2 Biotinylation of Eggs

1. Prepare the biotin solution immediately before use. Mix with 100 μl of FNSW to a final concentration of 10 mg/ml. Vortex tube to ensure that the biotin is completely dissolved. 2. Add 10 ml of 10% slurry of eggs to a 50 ml beaker (see Note 6). Remove 100 μl of seawater from the beaker. For a final concentration of 0.1 mg/ml, add all 100 μl of the 10 mg/ml of the biotin solution to the beaker—swirl eggs to resuspend and evenly distribute the biotin. 3. Incubate at room temperature for 30 min while resuspending the eggs by swirling every 5 min.

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4. After 30 min, add 5 ml of 0.5 M glycine in FNSW to the beaker and swirl to resuspend the eggs. This will quench the remaining free biotin. 5. Allow eggs to settle by gravity, replace FNSW, and resuspend eggs. Repeat washes four times to ensure removal of all biotin and glycine. Incubate biotinylated eggs at 16  C until needed. 6. Check eggs using light microscopy to ensure they are still healthy. 3.3

Fertilization

1. Ensure your beaker contains an ~10% (v/v) slurry of eggs, tubes are prelabeled for both the eggs and seawater samples, liquid nitrogen is ready and available, and a timer is ready for use. 2. Resuspend and distribute eggs evenly by swirling. Remove a 1 ml aliquot into a 1.7 ml tube labeled “unfertilized” (see Note 7). Briefly spin down at 2000  g using a personal tabletop microcentrifuge to concentrate eggs to the bottom of the tube. Using a clean pipette tip, carefully remove as much seawater as possible, without disrupting the pellet of eggs, and place supernatant into a new tube. In order to avoid contaminating the seawater with eggs, it is acceptable to leave some of the seawater with the egg sample, or remove and discard the remainder if time allows. Spin down the seawater supernatant sample with a microcentrifuge for a few seconds to concentrate any possible egg contamination. Ensure tops are securely closed and freeze by careful immersion into liquid nitrogen (see Note 8). 3. To prepare sperm and facilitate pipetting, use the fine scissors to mince the testis in the Eppendorf (you will not see anything being released). Cut the end of a p200 pipette tip to widen the opening. Pipet 10 μl of sperm and add to an Eppendorf tube with 1 ml with FNSW. Vortex the tube briefly to create a homogenous 1:100 dilution of sperm. 4. Add the 1 ml of sperm dilution to the 9 ml of eggs. Start the timer as soon as the sperm is added. Resuspend eggs by swirling to ensure equal distribution of eggs. Sperm are now at a 1:1000 dilution. 5. Aliquot 1 ml of resuspended eggs into a new tube for a fertilization sample at 1-min. Approximately 15 s before the 1-min mark, briefly spin down to concentrate eggs (and sperm) to the bottom of the tube. Using a clean pipette tip, carefully pipette off as much of the seawater as possible into a new tube (see Note 9). Ensure tops are securely closed and carefully place into the liquid nitrogen when the timer reaches 1-min (see Note 10). 6. Prior to the next time point, take a small aliquot and place it on a slide to visualize on a compound microscope. The sperm and

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fertilized eggs should be visible at 10 and 25. Around 2 min, the fertilization envelope should begin to be visible. Document the number of successfully fertilized eggs vs. unfertilized eggs. 7. Repeat step 5 for remaining time increments. It is recommended to sample at least every 2 min. 8. Once complete, store all samples at further analysis. 3.4 Affinity Interaction Using Streptavidin Mag Sepharose Beads (See Note 11)

80  C until ready for

1. Defrost the seawater samples on ice. Once thawed, spin the samples down for 10 min at 4  C at 12,128  g. Remove the supernatant to a new tube. This initial spin is to concentrate any eggs or egg particulate that may have been left in the seawater. 2. Resuspend the Mag Sepharose beads and aliquot 100 μl into a new tube and magnetize the beads. Remove the storage solution. 3. Add 500 μl of binding buffer and resuspend the beads. Immobilize the beads using the magnet and remove the liquid. 4. Add the 600 μl of the seawater samples to the beads and resuspend. Rotate tubes over end on a bench top rotator for 1 h at room temperature (see Note 12). 5. Magnetize the beads and remove the unbound liquid (see Note 13). Add 500 μl of washing buffer and resuspend the beads. Magnetize the beads and remove and discard the liquid. Repeat three times. 6. Add 100 μl of elution buffer and resuspend the beads. Incubate at 95–100  C for 5 min. 7. Magnetize the beads and transfer the eluate to a new tube. The eluate can be analyzed by western blotting for biochemical analysis, or by tandem mass spectrometry for identification; the latter protocol is not described here.

3.5 SDS-PAGE Gel and Western Blotting

1. Make a 7% polyacrylamide:bis-acrylamide (29:1 ratio) resolving gel with a 5% stacking gel (see Note 14). 2. Vortex thawed samples briefly to ensure homogenous distribution. Aliquot 10–15 μl into a new tube and add 2 sample buffer to a final concentration of 1. Heat sample on a heating block to 95  C for 5 min. Load entire sample on the gel and run at 110v for approximately 1.5 h, or until bromophenol blue in sample buffer reaches the bottom. 3. Transfer proteins from the gel to nitrocellulose overnight for a total of 800–1000 mA-hours (see Note 15). 4. Remove blot from transfer buffer and block in 5% blotto for 1 h at room temperature on a rotary tabletop shaker at low speed. Prepare the streptavidin-HRP probe by diluting it in the blotto

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at a 1:10,000 dilution. Pour off the blocking blotto and replace it with the blotto with the probe present. Incubate at room temperature for 1 h with gentle shaking to keep the blotto moving across the membrane. 5. Remove the blotto and wash the blot three times in TBST for approximately 10 min each time. 6. Prepare the ECL in a 1:1 ratio. While holding the blot at its corner by forceps, lay the blot face-up across plastic wrap after touching the corner to a paper towel to remove any excess TBST. Coat the blot in the ECL and leave standing without any shaking for 5 min. 7. Pick up the blot using the forceps and remove the ECL. Touch the edge to a paper towel to remove any extra. Place on clean plastic wrap and capture the signal using a BioRad ChemiDoc Imaging System (Fig. 2).

4

Notes 1. Given that 1-MA has to be dissolved in diH2O rather than FNSW, it may be prudent to make relatively high stock concentration of 1-MA (5 mM), in order to introduce as little diH2O as possible when adding the stock to FNSW. Aliquots of stock 1-MA can be stored at 20 for 6 months and at 80 for more extended periods. 2. Any glassware contacting oocytes or eggs should either be new or not washed with soap. Gametes are highly sensitive to soap, including soap residue, and such soap-based toxicity will affect the experimental outcome. 3. Sulfo-NHS-Biotin is stored desiccated because the NHS-ester reagents hydrolyze rapidly after introduction of water (including natural humidity in a laboratory), thereby rendering them unusable; if a 100 mg vial of biotin is purchased, ensure it is wrapped tightly with parafilm after each use and stored with a desiccant. Before using, remove from freezer and allow it to reach room temperature prior to use. To avoid having to do this and to circumvent unnecessary exposure to moisture in the air, purchase of individually packaged 1 mg samples is recommended (A39257, ThermoFisher Scientific, Waltham, MA). This chapter is based on use of the individual 1 mg packages. 4. Tween 20 is exceptionally viscous. To obtain correct amounts, either cut the tip on a p1000 pipette to widen the opening or mix a 10% solution of Tween 20 in diH2O. Tween 20 also tends to degrade after about 2 years, which can contribute to high backgrounds on the western blots.

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5. Ensure that the milk for the blotto is biotin-free (M-0841, LabScientific, Highlands, NJ); otherwise, the milk may bind the streptavidin probe. 6. A 1 ml aliquot of a 10% slurry of eggs typically gives about 100 μl of concentrated eggs which is approximately 1 mg of protein. 7. To avoid damaging eggs, cut approximately 3 mm off of the tip of a p1000 pipette tip to widen the opening. 8. In this protocol, seawater subjected to analyses, but egg pellets can be frozen as well for use in other applications. 9. It may be difficult to remove all of the seawater while trying to keep to the time frame. Remove as much as possible from the eggs because the seawater will be spun down to remove egg particulate prior to streptavidin affinity interaction. 10. The tubes will “sizzle” once they are placed into the liquid nitrogen. Once the “sizzling” stops, they are completely frozen and can be removed using tongs or a slotted spoon and stored in an ultrafreezer. 11. There are several methods such as spin dialysis tubes that can be used to remove biotinylated peptides; however, using the Streptavidin Mag Sepharose from Sigma-Aldrich (GE289857-38) provides the cleanest preparation of the biotinylated peptides. 12. If incubated at a lower temperature, such as 4  C, the samples should be incubated overnight. 13. For the first few times running this protocol, it may be worthwhile to keep the unbound fraction and run a western blot. This will allow for adjustment of the sample volume added to the beads to ensure there is enough sample or the beads are not being overloaded. 14. A 7% SDS-PAGE gel was used in Fig. 2; however, the percentage of the gel can be lowered to 5% for the resolving layer to spread the proteins out farther, thereby making it easier to see any changes. 15. You can write on the nitrocellulose using a pencil before its immersion in the transfer buffer as a means to ensure you have the correct orientation, or if you have more than one blot.

Acknowledgments We are very grateful for funding provided by the Eunice Kennedy Shriver National Institute of Child Health and Human Development of the National Institutes of Health under award number R15HD094324.

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References 1. Clark NL, Gasper J, Sekino M et al (2009) Coevolution of interacting fertilization proteins. PLoS Genet 5:el000570 2. Wilburn DB, Swanson WJ (2016) From molecules to mating: rapid evolution and biochemical studies of reproductive proteins. J Proteome 135:12–25 3. Sardet C, Dumollard R, McDougall A (2006) Signals and calcium waves at fertilization. Semin Cell Dev Biol 17:223–225 4. Kumano M, Carroll DJ, Denu JM et al (2001) Calcium-mediated inactivation of the MAP kinase pathway in sea urchin eggs at fertilization. Dev Biol 236:244–257 5. Rongish BJ, Wu W, Kinsey WH (1999) Fertilization-induced activation of phospholipase C in the sea urchin egg. Dev Biol 215:147–154 6. Stricker SA (1999) Comparative biology of calcium signaling during fertilization and egg activation in animals. Dev Biol 211:157–176 7. Steinhardt RA, Epel D (1974) Activation of sea-urchin eggs by a calcium ionophore. Proc Natl Acad Sci U S A 71:1915–1919 8. Sanders JR, Swann K (2016) Molecular triggers of egg activation at fertilization in mammals. Reproduction 152:41–50 9. Swann K, Lai FA (2016) The sperm phospholipase C- and Ca2+ signalling at fertilization in mammals. Biochem Soc Trans 44:267–272 10. Abassi YA, Carroll DJ, Giusti AF et al (2000) Evidence that Src-type tyrosine kinase activity is necessary for initiation of calcium release at fertilization in sea urchin eggs. Dev Biol 218:206–219 11. Inoue N, Ikawa M, Isotani A et al (2005) The immunoglobulin superfamily protein Izumo is required for sperm to fuse with eggs. Nature 434:234–238 12. Ohto U, Ishida H, Krayukhina E et al (2016) Structure of IZUMO1–JUNO reveals sperm–oocyte recognition during mammalian fertilization. Nature 534:566–569 13. Kato K et al (2016) Structural and functional insights into IZUMO1 recognition by JUNO in mammalian fertilization. Nat Commun 7:12198. https://doi.org/10.1038/ ncomms12198 14. Bianchi E, Doe B, Goulding D et al (2014) Juno is the egg Izumo receptor and is essential for mammalian fertilization. Nature 508:483–487

15. Hasan AKMM et al (2005) Uroplakin III, a novel Src substrate in Xenopus egg rafts, is a target for sperm protease essential for fertilization. Dev Biol 286:483–492 16. Simons K, Toomre D (2000) Lipid rafts and signal transduction. Nat Rev Mol Cell Biol 1:31–39 17. Belton RJ, Adams NL, Foltz KR (2001) Isolation and characterization of sea urchin egg lipid rafts and their possible function during fertilization. Mol Reprod Dev 59:294–305 18. Kadandale P et al (2005) The EGG surface LDL receptor repeat-containing proteins EGG-1 and EGG-2 are required for fertilization in Caenorhabditis elegans. Curr Biol 15:2222–2229 19. Giusti AF, Hoang KM, Foltz KR (1997) Surface localization of the sea urchin egg receptor for sperm. Dev Biol 184:10–24 20. Glabe CG, Vacquier VD (1978) Egg surface glycoprotein receptor for sea urchin sperm bindin. Cell Biol 75:881–885 21. Carroll DJ, Jaffe LA (1995) Proteases stimulate fertilization-like responses in starfish eggs. Dev Biol 170:690–700 22. Foltz KR, Lennarz WJ (1990) Purification and characterization of an extracellular fragment of the sea urchin egg receptor for sperm. J Cell Biol 111:2951–2959 23. Avidin-Biotin Technical Handbook (2009) Thermo Scientific. http://assets.thermofisher. com/TFS-Assets/LSG/brochures/1601675Avidin-Biotin-Handbook.pdf 24. Ho¨rmann K et al (2016) A surface biotinylation strategy for reproducible plasma membrane protein purification and tracking of genetic and drug-induced alterations. J Proteome Res 15:647–658 25. Smolders K, Lombaert N, Valkenborg D et al (2015) An effective plasma membrane proteomics approach for small tissue samples. Sci Rep 5:10917 26. Cihal S, Carroll DJ (2014) Biotinylation of oocyte cell surface proteins of the starfish Patiria miniata. In: Carroll DJ, Stricker S (eds) Developmental biology of the sea urchin and other marine invertebrates, Methods in molecular biology (Methods and protocols), vol 1128. Humana Press, New York, NJ 27. Wiseman E, Bates L, Dube´ A et al (2018) Starfish as a model system for analyzing signal transduction during fertilization. Results Probl Cell Differ 65:49–67

Chapter 10 Quantifying Cell Proliferation During Regeneration of Aquatic Worms Eduardo E. Zattara and B. Duygu O¨zpolat Abstract Many species of aquatic worms, including members of the phyla Nemertea, Annelida, Platyhelminthes, and Xenacoelomorpha, can regenerate large parts of their body after amputation. In most species, cell proliferation plays key roles in the reconstruction of lost tissues. For example, in annelids and flatworms, inhibition of cell proliferation by irradiation or chemicals prevents regeneration. Cell proliferation also plays crucial roles in growth, body patterning (e.g., segmentation) and asexual reproduction in many groups of aquatic worms. Cell proliferation dynamics in these organisms can be studied using immunohistochemical detection of proteins expressed during proliferation-associated processes or by incorporation and labeling of thymidine analogues during DNA replication. In this chapter, we present protocols for labeling and quantifying cell proliferation by (a) antibody-based detection of either phosphorylated histone H3 during mitosis or proliferating cell nuclear antigen (PCNA) during S-phase, and (b) incorporation of two thymidine analogues, 50 -bromo-20 -deoxyuridine (BrdU) and 50 -ethynyl-20 -deoxyuridine (EdU), detected by immunohistochemistry or inorganic “click” chemistry, respectively. Although these protocols have been developed for whole mounts of small (2 cm), it is easier to remove as much of the current solution as before adding the next solution to the container. The choice of container is also guided by specimen size: 1.5 mL microcentrifuge tubes can be used for smaller worms, allowing incubation in smaller volumes of each solution (which can result in considerable savings in reagent use), while small petri dishes (35 mm diameter and 15 mm high) can be used for larger worms. 6. In contrast to organisms with internal or external hard skeletons, many marine worms depend on soft tissues—particularly muscles and connective tissue—to keep their shape. As a result, muscle contraction and tissue dehydration during fixation might cause both morphological distortion and breakage, obscuring anatomical details and potentially generating fixation artifacts. Thus, adequate relaxation of specimens before fixation is fundamental to attain high quality material for morphological studies. Relaxation of specimens can be achieved through sedation or inhibition of muscle contraction: usually, specimens are submerged for a period in relaxing solutions isotonic to their regular medium (from sea to fresh water) that contain sedative or anesthetic reagents like ethanol, potassium chloride, chlorobutanol, magnesium chloride, nicotine etc., and allowed to fully relax before adding fixative agents. Different species and life stages might react differently to each relaxation technique; thus researchers may need to experiment with reagent combinations that works best with their study system. The amount of time needed for proper relaxation is species- and size-specific. Smaller worms usually relax more quickly than larger ones, which might take several minutes to become immobilized. Lower temperatures usually help in lowering the worm’s activity; in such cases, using cold (4  C) solutions and keeping the petri dish or other containers on ice might help the relaxation process. However, in some species excessive cold causes severe contraction, fragmentation and premature death. Continual or sporadic gentle agitation of the container might enhance specimen relaxation. It is highly recommended for researchers to test and optimize conditions to find an ideal relaxation state before fixation. 7. Although fixative is usually 4% paraformaldehyde, other reagents might be used for specialized ends—like

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glutaraldehyde for specimens that will be examined using electron microscopy. This worm relaxation and fixation protocol below can be used for either small (1 cm) specimens. The remaining protocols have been optimized for use with small worms and 1.5 mL centrifuge tubes, as this size range is also adequate for marine embryos, larvae and adults of other infaunal and meiofaunal species, and allows for considerable savings in the costliest reagents. These protocols can be adapted to larger specimens by increasing the working volume of incubation solutions and increasing incubation times. The optimal fixation time will depend of the size and consistency of each specimen, and downstream application. If incubation in fixative solution is too short, the specimen will suffer degradation over time that will be more pronounced in deeper tissues. If incubation in fixative is too long, excessive fixation might result in decreased antigen availability due to extensive crosslinking, tissue hardening and breakage, and DNA degradation. 8. This protocol assumes specimens will be kept in the same container throughout the process and change of solutions is done by removing most of the current solution from the container with a pipette and then adding the new solution. We find that using hand-held regular glass pipettes manually operated with a rubber bulb gives the best compromise between convenience, precision and operator fatigue. However, pipetting can also be done using automated pipettes, plastic Pasteur pipettes, mouth pipettes or vacuum-operated pipettes. Pipettes with glass, transparent ends are especially recommended when the specimens are small enough to fit through the tip opening. 9. Constant or frequent, mild agitation of the tubes is useful to homogenize the washing and incubation solutions, promote tissue penetration and prevent multiple specimens stained in the same tube from sticking together or to the walls of the tube. Energetic agitation by vortexing, shaking, or inverting the tube is to be avoided, as this might cause specimen breakage or sticking of the specimens to the walls of the tube on areas not covered by liquid. Gentle agitation is best achieved by gently tapping the bottom of the tube with a fingertip while keeping the tube vertical from its upper rim, by making the tube spin around its vertical axis while sitting on a tube rack, or by placing it on an orbital shaker set at low speed. 10. The purpose of this Blocking step is to incubate the sample with a high concentration of a protein mix that lack antigens that could cause off-target antibody binding. Serum proteins in the Blocking solution will bind to any “sticky” molecules present in the sample and block them from binding nonspecifically to the primary or secondary antibodies. Ideally, serum from the host species that generated the secondary antibodies are used

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(e.g., with secondary antibodies produced in goats, normal goat serum is used). Do not use serum from the same host species used to produce any of the primary antibodies, as it will compete for sites with the secondary antibody and introduce non-specific signal. Other Blocking reagents are available. The correct Blocking reagent can greatly reduce noisy staining due to nonspecific binding of primary or secondary antibodies; thus, tweaking the blocking step is crucial for optimizing the protocol when analyzing different species. 11. Both the working antibody concentration and incubation times are quite flexible. As with most other parameters, antibody concentration can be tuned to the specific properties of each species. Lower concentrations will reduce costs and might decrease background or nonspecific staining. Higher concentrations might result in stronger signal and shorter incubation times but might cause more noise and require additional washes. Shorter incubation times can speed up the protocol, but might result in lower signal, especially deeper within the tissues. In general, longer incubation times have no negative effects besides the additional time spent. Antibody incubations are ideal breakpoints of this protocol, as they can be done overnight or even over a few days if kept at 4  C in a fridge. 12. PBTx washes are meant to remove unbound antibodies. They can also help remove or decrease antibodies bound with low affinity to off-target molecules, thereby reducing noise. Larger/thicker specimens should be washed longer, as large molecules take longer to diffuse out of tissues. Washes are also a potential breakpoint for this protocol, although it is more useful to break at the next step. 13. There is a wide variety of conjugated secondary antibodies to choose from. Besides being attached to numerous fluorescent molecules, antibodies can be conjugated to enzymes, like alkaline phosphatase or horseradish peroxidase, which enable highly sensitive chromogenic reactions. Fluorescent labels are ideal for use with epifluorescence and confocal laser microscopes, but can sometimes result in noisy images, especially if the primary antibody cross-reacts with off-target molecules. Chromogenic enzyme labels, on the other hand, are highly sensitive since signal builds up over time, but the chemical cocktail used to reveal their presence can also react with endogenous enzymes; thus, appropriate negative controls (without primary antibodies and without secondary antibodies) are recommended when using such labels. Try minimizing spectral emission overlap when choosing fluorescent labels for multichannel imaging.

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14. Storing the stained specimens in glycerol helps to clear tissues and improves imaging when mounting with glass slides and coverslips, as the refractive index of glycerol is closer to glass than that of water. Stained worms with no hard parts and other delicate species might shrivel or deform if moved directly from the PBS aqueous medium to a pure glycerol solution but passing them through a graded concentration series will minimize such problems. At concentrations >90%, glycerol becomes too viscous for handling specimens with pipettes; samples can be mounted in glycerol/n-propyl gallate mounting medium (9 mL glycerol, 1 mL 1 M Tris–HCl pH 8.0, and 0.05 g n-propyl gallate). 15. The optimal concentration of BrdU or EdU for each species and purpose should be determined experimentally. Aquatic worms might incorporate the thymidine analogues directly through the skin or by ingestion of the incubation medium followed by gut absorption and tissue distribution. For larger species in which epidermal and endodermal absorption is insufficient for adequate marker incorporation, direct injection of the thymidine analogues into the body cavities is often a viable alternative. 16. Do not use automatic pipettors to handle concentrated HCl as its vapors can corrode their internal metallic components. 17. This DNA denaturation step is critical to expose the incorporated BrdU as an epitope for the anti-BrdU antibody. Unfortunately, this step also denatures many proteins in the sample, thus interfering with other immunolabeling aims. Thus, if labeling other proteins is desired, then it is advisable to test if milder denaturation conditions (i.e., a lower HCl concentration) still yield reliable signal while not abolishing other epitopes. For some species, HCl treatment may result in nonspecific background signal, sometimes to the point of completely obscuring to BrdU signal. An alternative approach is to replace acid treatment with a DNase enzyme incubation [26, 27]. 18. After the last of these washes, the pH should approximate the 7.4 value of PBS. Use pH paper to determine how many washes are needed to reach this neutral point. 19. Some species and tissues might have endogenous peroxidases that react with the chromogenic solution to generate high levels of nonspecific background staining. Presence of endogenous peroxidases can be assessed by developing a negative control (samples of the same species not incubated with the secondary antibody) in the chromogenic solution. If present, endogenous peroxidases can be eliminated by saturating concentrations of hydrogen peroxide (H2O2) through an optional

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pretreatment step. Usually incubating between 10 to 30 min in 0.3% H2O2 is enough, but the optimal time and concentration should be optimized experimentally. 20. Addition of NiCl is optional: it increases sensitivity and results in darker coloring. However, NiCl might cause the sample to overdevelop and yield more background in the presence of endogenous peroxidases. 21. Development time should be determined experimentally, preferably while looking at the samples with known cell proliferation patterns using a magnifying glass or a dissecting microscope. Ideally, developed samples should show clearly localized nuclear signal. Underdeveloped samples have faint or no signal, whereas overdeveloped samples tend to have diffuse background staining not localized to nuclei. 22. The EdU labeling and detection system was initially released commercially as the Click-iT® reagent kit (ThermoFisher). This kit is still the most widely used option to perform this assay. More recently, Andrew Brumm and Joachim Goedhart shared an alternative protocol that used specific reagents, but otherwise employed materials similar to the commercial version [28]; the protocol presented here is based on their protocol. 23. The EdU 10 mM stock solution can be prepared as described in Subheading 2.3, item 3, or by adding 2 mL of dH2O to the vial of Component A of the Click-iT® commercial kit. 24. Although most reagents in this protocol have low molecular weights, permeabilization of tissues with a detergent might improve penetration and thereby shorten protocol times. Whether permeabilization is necessary should be determined for each species of interest. 25. In contrast to the BrdU development assay with its harsh denaturation step, the EdU development reaction is usually compatible with immunohistochemical protocols (for example, the one described in Subheading 2.3). When combining this EdU protocol with immunohistochemistry procedures, the recommended sequence of steps is to perform EdU incubation, relaxation and fixation (see Subheading 3.4, steps 1–4), perform the immunohistochemistry steps (see Subheading 3.2, steps 2–8) and then perform EdU development (see Subheading 3.4, steps 5–9) [4]. For some species, the permeabilization step (see Subheading 3.4, step 5) might require an additional treatment with a proteinase. 26. If using the Click-iT® commercial kit, add instead the following to prepare the EdU Reaction Mix: 86 μL of 1 EdU reaction buffer (prepare stock by adding 0.1 mL of Component D to 0.9 mL of dH2O) + 4 μL of CuSO4

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(component E) + 8 μL of dH2O + 1 μL of the dye azide (prepare 0.24 stock by adding 290 μL of Component C to the vial of Component B, aliquot and store at  20  C) + 1 μL of 10 EdU buffer additive (prepare stock by adding 2 mL of dH2O to the vial of Component F, aliquot and store at 4  C; if the solution develops a brown color, it has degraded and should be discarded) + 1 μL of 1 mg/mL DAPI. References 1. Gline SE, Kuo D-H, Stolfi A, Weisblat DA (2009) High resolution cell lineage tracing reveals developmental variability in leech. Dev Dyn 238:3139–3151 2. Zantke J, Bannister S, Rajan VBV, Raible F, Tessmar-Raible K (2014) Genetic and genomic tools for the marine annelid Platynereis dumerilii. Genetics 197:19–31 3. Zielke N, Edgar BA (2015) FUCCI sensors: powerful new tools for analysis of cell proliferation. Wiley Interdiscip Rev Dev Biol 4:469–487 ¨ zpolat BD, Handberg-Thorsager M, 4. O Vervoort M, Balavoine G (2017) Cell lineage and cell cycling analyses of the 4d micromere using live imaging in the marine annelid Platynereis dumerilii. elife 6:e30463 5. Tadokoro R, Sugio M, Kutsuna J, Tochinai S, Takahashi Y (2006) Early segregation of germ and somatic lineages during gonadal regeneration in the annelid Enchytraeus japonensis. Curr Biol 16:1012–1017 6. Tweeten KA, Anderson A (2008) Analysis of cell proliferation and migration during regeneration in Lumbriculus variegatus (Clitellata: Lumbriculidae). Bios 79:183–190 7. Bely AE, Sikes JM (2010) Latent regeneration abilities persist following recent evolutionary loss in asexual annelids. Proc Natl Acad Sci 107:1464–1469 8. Bely AE, Sikes JM (2010) Acoel and platyhelminth models for stem-cell research. J Biol 9:14 9. Demilly A, Steinmetz P, Gazave E, Marchand L, Vervoort M (2013) Involvement of the Wnt/β-catenin pathway in neurectoderm architecture in Platynereis dumerilii. Nat Commun 4:1915 10. Zattara EE, Bely AE (2013) Investment choices in post-embryonic development: quantifying interactions among growth, regeneration, and asexual reproduction in the annelid Pristina leidyi. J Exp Zoolog B Mol Dev Evol 320:471–488

11. Srivastava M, Mazza-Curll KL, van Wolfswinkel JC, Reddien PW (2014) Whole-body acoel regeneration is controlled by Wnt and Bmp-Admp signaling. Curr Biol 24:1107–1113 12. Bird AM, von Dassow G, Maslakova SA (2014) How the pilidium larva grows. EvoDevo 5:13 13. Szabo´ R, Ferrier DEK (2014) Cell proliferation dynamics in regeneration of the operculum head appendage in the annelid Pomatoceros lamarckii. J Exp Zoolog B Mol Dev Evol 322:257–268 14. de Jong DM, Seaver EC (2017) Investigation into the cellular origins of posterior regeneration in the annelid Capitella teleta. Regeneration 5:61–77 15. Buchwalow IB, Bo¨cker W (2010) Immunohistochemistry: basics and methods. SpringerVerlag, Berlin, Heidelberg 16. Nowak SJ, Corces VG (2004) Phosphorylation of histone H3: a balancing act between chromosome condensation and transcriptional activation. Trends Genet 20:214–220 17. Gurley LR, D’anna JA, Barham SS, Deaven LL, Tobey RA (1978) Histone phosphorylation and chromatin structure during mitosis in Chinese hamster cells. Eur J Biochem 84:1–15 18. Hendzel MJ, Wei Y, Mancini MA et al (1997) Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106:348–360 19. Miyachi K, Fritzler MJ, Tan EM (1978) Autoantibody to a nuclear antigen in proliferating cells. J Immunol 121:2228–2234 20. Boehm EM, Gildenberg MS, Washington MT (2016) The many roles of PCNA in eukaryotic DNA replication. The Enzymes 39:231–254 21. Niwa N, Akimoto-Kato A, Sakuma M, Kuraku S, Hayashi S (2013) Homeogenetic inductive mechanism of segmentation in polychaete tail regeneration. Dev Biol 381:460–470

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22. Taylor JH, Woods PS, Hughes WL (1957) The organization and duplication of chromosomes as revealed by autoradiographic studies using tritium-labeled thymidine. Proc Natl Acad Sci U S A 43:122–128 23. Gratzner HG (1982) Monoclonal antibody to 5-bromo- and 5-iododeoxyuridine: a new reagent for detection of DNA replication. Science 218:474–475 24. Dean PN, Dolbeare F, Gratzner H, Rice GC, Gray JW (1984) Cell-cycle analysis using a monoclonal antibody to Brdurd. Cell Prolif 17:427–436 25. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci 105:2415–2420

26. Gonchoroff NJ, Katzmann JA, Currie RM et al (1986) S-phase detection with an antibody to bromodeoxyuridine: role of DNase pretreatment. J Immunol Methods 93:97–101 27. Ye W, Mairet-Coello G, DiCicco-Bloom E (2007) DNAse I pre-treatment markedly enhances detection of nuclear cyclindependent kinase inhibitor p57Kip2 and BrdU double immunostaining in embryonic rat brain. Histochem Cell Biol 127:195–203 28. Can anyone recommend an alternative to Click-it EdU labeling kit from Invitrogen? https://www.researchgate.net/post/Can_any one_recommend_an_alternative_to_Click-it_ EdU_labeling_kit_from_Invitrogen

Chapter 11 In Situ Hybridization Techniques in the Homoscleromorph Sponge Oscarella lobularis Laura Fierro-Constaı´n, Caroline Rocher, Florent Marschal, Quentin Schenkelaars, Nina Se´journe´, Carole Borchiellini, and Emmanuelle Renard Abstract The Porifera are one of the best candidates as the sister group to all other metazoans. Studies on this phylum are therefore expected to shed light on the origin and early evolution of key animal features. Transcriptomic or genomic data acquired during the last 10 years have highlighted the conservation of most of the main genes and pathways involved in the development of the other metazoans. The next step is to determine how similar genetic tool boxes can result in widely dissimilar body plan organization, dynamics, and life histories. To answer these questions, three main axes of research are necessary: (1) conducting more gene expression studies; (2) developing knockdown protocols; and (3) reinterpreting sponge cell biology using modern tools. In this chapter we focus on the in situ hybridization (ISH) technique, needed to establish the spatiotemporal expression of genes, both on whole mount individuals and paraffin sections, and at different stages of development (adults, embryos, larvae, buds) of the homoscleromorph sponge Oscarella lobularis. Key words Porifera, Evo-devo, RNA probe, Gene expression

1

Introduction Sponges (Porifera) represent one of the earliest animal lineages. As for the three other non-bilaterian lineages (Ctenophora, Placozoa, and Cnidaria), a more in-depth study of their biology in a comparative evo-devo approach is needed to understand the origin and evolution of animal body plan features. Most of the studies performed so far on this phylum are aimed at characterizing the gene content of their genomes. These useful transcriptomic and genomic data have challenged previous views on these animals but we still need to determine the functions of these genes in sponges (for

Laura Fierro-Constaı´n and Caroline Rocher contributed equally to this work. Emmanuelle Renard and Carole Borchiellini equally supervised this work. David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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review see [1]. To study gene function, we need to determine their expression pattern (in time and space), the location of the protein produced, and explore the effects of knockdown or overexpression of the targeted genes. These types of data still remain scarce in sponges, probably because of various technical difficulties: cell transcriptomics were performed in Ephydatia fluviatilis and Amphimedon queenslandica [2–4], immunolocalization of proteins in Ephydatia muelleri and Amphimedon queenslandica [5–8], Oscarella pearsei [9], and Oscarella lobularis (Caroline et al. [10], Borchiellini et al., in this volume), gene knockdown by pharmacological or RNAi approaches in Ephydatia muelleri [11–14] and Oscarella lobularis [15]. In this chapter, we focus on in situ hybridization (ISH), a useful tool to explore gene expression at cell, tissue, or organism levels. Although this technique is routinely used in model organisms since the 1960s [16], it is quite sensitive; therefore, it is sometimes delicate to perform in emergent model organisms and even more so in marine animals [17, 18]. The first ISH realized on sponges only dates back to 2000s on two demosponges Aplysina cavernicola and Suberites domuncula [19, 20]. Since then, ISH studies has been developed and used in only five species: the demosponges Amphimedon queenslandica [21–25] and much more recently in Ephydatia muelleri [5, 11] and Halisarca dujardinii [26], in the Calcarea Sycon ciliatum [27– 32] and in the homoscleromorph Oscarella lobularis [15, 33, 34]. In order to make this technique routinely reliable and reproducible, each spongiologist team had to face different problems, probably linked to the different histological characteristics harbored by each class [35]; in the marine sponge Oscarella lobularis (Porifera—Homoscleromorpha) we faced problems of probe penetration, tissue integrity, and background staining. This chapter details the protocol of the in situ hybridization ISH technique from digoxygenin-labeled RNA probes synthesis, tissues fixation, hybridization, and postfixation to perform both whole mount in situ hybridization WMISH (either manually or using In situPro VSi (Intavis AG)) and in situ hybridization on serial paraffin sections SISH (manually). This technique is essential to study gene expression at every stages of development (adult, embryos, larvae, and buds (Fig. 1). Our protocols were validated on more than 20 genes [33].

2

Materials

2.1 Biological Material

Oscarella lobularis is a marine sponge species that has been already well described, (for more information see [36]). The initial protocol for in situ hybridization in O. lobularis was published in Gazave

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Fig. 1 Life cycle of O. lobularis showing different developmental stages

et al. [34], and secondarily improved to obtain fully reproducible results with numerous probes at more numerous developmental stages [33]. 2.2

Probe Synthesis

1. DIG-RNA Labeling Kit T7/Sp6 (Roche®). 2. Wizard® SV Gel and PCR clean-up system (Promega).

2.3

Sample Fixation

1. Diethylpyrocarbonate (DEPC in H2O: 1 mL for 1 L H2O autoclaved 2 times successive) (Sigma-Aldrich). 2. Methanol (100%). 3. Natural sea water, 0.2 μm filtered (FNSW). 4. Paraformaldehyde (PFA, 16% solution, pH 7.5) (see Note 1). 5. Phosphate-buffered saline (PBS, 10: 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4 in DEPC-treated H2O, pH 7.4, autoclaved). 6. Phosphate-buffered saline with 0.01% Tween 20® (PBT). 7. Tween 20® (20%, Sigma-Aldrich).

2.4 Paraffin Fixation and Cuts

1. Ethanol (100%). 2. Glass vials 10 mL (e.g., Wheaton® Snap-Cap® Specimen Vials). 3. NeoClear® (Millipore) or Tissue-Clear Xylene Substitute (Sakura). 4. Paraffin.

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5. Paraffin embedding molds. 6. Superfrost plus® slides (Thermo scientific). 2.5 In Situ Hybridization

1. Antidigoxigenin (anti-DIG alkaline AP-conjugated, Fab fragments Roche®).

phosphatase

2. Blocking reagent (BR, 10% in MAB, autoclaved, Roche®). Prepare aliquots and store at 20  C. 3. Blocking solution (BS, 1% Blocking reagent in MABT). 4. Deionized formamide 100%. 5. Diethyl pyrocarbonate (DEPC-H2O). Prepare DEPC (1:1000) in H2O and autoclave 2 times (Sigma-Aldrich). 6. Ethanol, Absolute. 7. Glycine (40 mg/mL). Only for whole mount ISH. 8. Heparin sodium salt (50 mg/mL in DEPC-treated H2O, Sigma-Aldrich). Prepare aliquots and store at 20  C. 9. Hybridization Solution (HS): 50% deionized formamide, 5 SSC, 50 μg/mL tRNA, 100 μg/mL heparin, 0.1%Tween 20 in DEPC-treated H2O. 10. Levamisole (50 mg/mL) (Sigma-Aldrich). 11. Maleic acid buffer (MAB, 0.1 M maleic acid, 0.15 M NaCl in DEPC-treated H2O, pH 7.5, autoclaved). 12. Maleic acid buffer with Tween (MABT: 0.01% Tween 20 in MAB). Store at 4  C NaCl (5 M pH 7.6, autoclaved). 13. NBT/BCIP Ready to Use Tablets (Roche®). 14. NeoClear® (Millipore) or Tissue-Clear Xylene Substitute (Sakura). 15. Neo-mount® (Merck Millipore). 16. NTMT 0.1 M NaCl, 0.1 M Tris–HCl, 50 mM MgCl2, 0.5 mg/mL Levamisole, 0.1% Tween 20 in DEPC-treated H2O. 17. NTT 0.1 M NaCl, 0.1 M Tris–HCl, 0.1% Tween 20 in DEPCtreated H2O. 18. MgCl2 (1 M, autoclaved). 19. Proteinase K (1 mg/mL in PBT) (Euromedex). Only for whole mount ISH. 20. Phosphate-buffered saline with Tween (PBT: 0.01% Tween 20 in PBS). Store at 4  C. 21. Saline–sodium citrate buffer (SSC, 20: 0.3 M sodium citrate, 3 M NaCl in DEPC-treated H2O, pH 7, autoclaved). 22. SSC 2 Solution 0.01% Tween 20. Only for whole mount ISH using INTAVIS robot.

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23. Slide jar, slide staining tray, slide incubation chamber. 24. Tris–HCl (1 M, pH 7.6, autoclaved). 25. tRNA (10 mg/mL in DEPC-treated H2O, Roche®). Prepare aliquots and store at 4  C). 26. Tween 20 (20%, Bio-Rad). 27. WASH 1 Solution: 50% deionized formamide, 4 SSC, 0.1% Tween 20 in DEPC-treated H2O. Only for whole mount ISH. 0.01% Tween 20 for whole mount ISH using INTAVIS robot. 28. WASH 2 Solution: 50% deionized formamide, 2 SSC, 0.1% Tween 20 in DEPC-treated H2O. Only for whole mount ISH. 0.01% Tween 20 for whole mount ISH using INTAVIS robot. 29. WASH 3 Solution: 50% deionized formamide, 1 SSC, 0.01% Tween 20 in DEPC-treated H2O. Only for whole mount ISH using INTAVIS robot. 2.6 Embedding and Sectioning Tissues

1. Acetone. Only for Epoxy embedding. 2. Embedding gelatin (or polyethylene) capsules. 3. Epoxy Embedding Kit (Sigma) or LR White Resin (EMS). 4. Ethanol, Absolute.

3 3.1

Methods (see Notes 2 and 3) Probe Synthesis

1. To facilitate the linearization of the vector when the gene of interest has been cloned, perform a PCR with T7-SP6 primers and purify the PCR reaction or the excised DNA agarose bands following the electrophoresis (e.g., Wizard® SV Gel and PCR clean-up system, Promega). If necessary, pool several bands/ PCR reactions from a same gene to obtain a high DNA concentration. This allows to obtain a T7-Sp6 flanking DNA template with a known orientation. 2. Use 1000 ng of template and other reactive to synthetize the RNA probe (e.g., DIG-RNA Labeling Kit T7/Sp6 (Roche®) according to the manufacturer’s recommendations. 3. When the probe synthesis is achieved, just after the DNase treatment, store at 80  C 1 μL of the mix in a PCR tube (to dose the RNA, see step 5). 4. Once the RNA pellet is diluted, keep 2.5 μL of the RNA probe in a new PCR tube. Store, meanwhile, RNA probe at 80  C. 5. Denature 3 min at 95  C PCR tubes in a thermocycler, chill 3 min on ice, add the loading dye and rapidly transfer them in a 1.5% agarose electrophoresis gel (30–40 min at 110 V). Use a DNA/RNA ladder designed for nucleic acid quantification (e.g., Smart ladder) to evaluate the RNA concentration.

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6. Aliquot in PCR tubes the volume needed to a RNA probe concentration of 50 ng in each aliquot. Store at 80  C (for up to one year), avoid thawing and refreezing. 3.2 Fixation (See Note 4) 3.2.1 To Fix Adults

1. Wash several times and carefully clean adult specimens with forceps under a magnifier in FNSW (avoiding thermal shocks), to remove potential epi and endobionts and cut in small pieces. 2. Transfer the pieces in a clean Falcon tube containing 4% PFA-FNSW and fix during 3 h at room temperature or overnight at 4  C on a nutator (slowest speed to avoid damaging the tissues). 3. Wash in 50% and 75% PBT-FNSW for 15 min each, in a nutator. 4. Wash twice in 100% PBT for 15 min each, on a nutator. 5. Wash in 25%, 50%, and 75% MeOH-PBT for 15 min each, in a nutator. 6. Wash twice in 100% MeOH for 15 min each, on a nutator. 7. Store with fresh MeOH 100% at

3.2.2 To Fix Buds and Larvae

20  C for several years.

1. Transfer buds or larvae in a six-well plate (Image Sequence and double click on the first image of the entire dataset saved at step 11 to open the stack (see Note 2). 12. Fiji: The Segmentation data once opened in Fiji will be displayed as a digital array of 1 s and 2 s. Typically, we subtract 1, then multiply the result by 255 to create a saturated image. 13. Fiji: With the 8-bit image, choose the color. 14. Fiji: Then transform to RGB color images, and merge using the color command (merged with the “Filtered Input” image stack for example). (Save and open the “Filtered Input” following the same procedure. However, you will not need to subtract and multiply since this image will appear with pixel values that are appropriate.) 15. ICY (http://icy.bioimageanalysis.org/): Save the merged RGB color image as a Tiff and open using the freeware ICY program for 3D rendering. Use the VTK icon to render. (Be sure to reassign the voxel parameters from the original confocal dataset before rendering otherwise the rendered image may appear flat.) 16. ICY: Adjust intensity values of each channel so that both are visible in the rendered 3D view and save using the snap-shot icon (see resultant image, Fig. 1) (see Note 3).

4

Notes 1. Ilastik is good for simple and fast 3D segmentation using the Carving module. The segmented object can also be saved using the “Save current object” command (this creates an “X.obj” file), that can be opened in the freeware animation program Blender (https://www.blender.org/) using the “Import” command for further processing if desired. Alternatively, the “segmented” image and “filtered input” image can be opened in Imaris (which is not freeware) for further analysis.

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2. Fiji is used here for converting the binary Ilastik segmented image (composed of values 1 and 2 only) into an image composed of 0 and 255, and for selecting the image colors for merging. 3. ICY is useful for creating a 3D image from a stack of images. It is also capable of making a 3D plus time rendering of 3D timelapse data. 4. CRISPOR protocol and sequence of oligonucleotide generated for Phallusia Brachyury and Tyrosinase: Step 1. Using CRISPOR [22] copy and paste gene exon sequence using the following instructions (use 12,000 rcf (see Note 4). 4. Add 500 μl of Tri-reagent and mix by pipetting up and down. The sample should be thawed and no longer translucent. 5. Add 200 μl of chloroform to the sample. If using Phase Lock tubes (optional), transfer the sample to a prespun Phase Lock tube. Shake the sample for 15–20 s. Do not vortex, especially if using Phase Lock tubes! 6. Incubate the samples for 3 min on ice. 7. Spin the samples at >12,000 rcf for 15 min at 4  C. 8. Carefully pipet the aqueous (upper) phase into a new 1.7 ml tube or prespun Phase Lock tube. 9. Perform a second extraction adding 500 μl of phenol–chloroform (pH 6.0) and repeating steps 6–8 (see Notes 5 and 6). 10. Precipitate the RNA by adding 2 μl of glycogen (20 mg/ml), followed by 500 μl of 100% isopropanol. Shake tube for 10 s and incubate at room temperature for 2 min. 11. Spin samples at >12,000 rcf for 15 min at 4  C. 12. Remove the liquid from the tube by carefully pipetting. Avoid disturbing the translucent whitish-tan pellet (see Notes 7 and 8). Once the liquid is removed, spin briefly at >12,000 rcf for 10 s and remove remaining liquid with a p10 pipette. 13. Wash pellet by adding 1 ml of 70% ethanol. Mix by inverting the tube 3 times. 14. Spin samples at >12,000 rcf for 15 min at 4  C. 15. Repeat steps 12–14. 16. Remove the remaining liquid as described in step 12. Invert tube on clean surface or delicate task wipe and allow to dry for approximately 5 min (see Note 9). 17. Resuspend in 15–30 μl of nuclease free water (see Note 10). 18. Optional: treat RNA with DNase (see Notes 11 and 12). 19. Assess RNA integrity by running 1 μl on an RNA gel or using a microfluidic electrophoresis machine (e.g., Agilent Bioanalyzer, Fig. 1) and quantify using a spectrophotometer (e.g., NanoDrop) (see Note 13). 20. Proceed with Next Generation Sequencing library preparation protocol that is appropriate for the downstream application (see Note 14). 3.2 Quantification of Transcriptomic Data

The following describes a standard workflow for quantifying Next Generation Sequencing (NGS) reads from a transcriptomic analysis. As programs, tools, and file-formats are ever changing we include here a general workflow describing tools that are current

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Fig. 1 Bioanalyzer results from total RNA extraction of Nematostella vectensis. The capillary electrophoresis gel is pictured on the left with the fluorescence intensity quantification on the right. The 5S, 18S, and 28S ribosomal subunits show up as clear, well separated bands indicating a high-quality extraction

at the time of writing. The programs below are intended to be run on a Linux server or Linux/Unix based PC. 1. Obtain reference for read alignment: Download a Nematostella vectensis genome or transcriptome assembly from one of the sources in Table 1. 2. Trim reads (optional): While read trimming is not required, it can increase the read mapping rate of lesser quality sequence data. Both Trimmomatic and Cutadapt work very well for trimming NGS sequence reads. Ensure that the appropriate sequencing adapters are used to trim (see Note 15). 3. Align reads to the reference: Read alignment can be performed with any number of popular programs including Bowtie2 for end-to-end alignment (e.g., for alignment to a transcriptome) or HISAT2 for splice aware alignments (e.g., for alignment to a genome). If the alignment rate is less than 50%, assess read quality with fastQC and consider trimming low quality reads. 4. Count successfully mapped reads: Read quantification can be performed with a variety of tools including RSEM [27], or Featurecounts [26]. The former works well with alignments to complex transcriptomes while the later works very well with genomic alignments. 5. Gene level quantification: When aligning to a transcriptome in which multiple transcripts per gene are present, gene level quantification can be estimated by summing the mapped reads of all transcripts per gene. The R functions aggregate (), and summarize() are very useful for this. NvERTx, includes Nemve1 annotations that can be used for gene level quantification (see Note 16). De novo transcriptome assemblies can be annotated using a draft genome or closely related species to identify putative genes.

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6. Remove lowly expressed genomic features (optional but recommended): Before normalizing, transcripts or genes with very low counts should be removed as these features, depending on the number of replicates, will often have very high variance and affect dispersion estimates. A detection limit can be calculated to determine the cutoff threshold. As a rule of thumb, when analyzing a result done in triplicate, genomic features with 5 counts or fewer in more than 25% of the samples will be too low to accurately quantify. 7. Normalization and differential testing: The Bioconductor R packages both DEseq2 and edgeR perform very well with transcriptomic data and can accommodate a variety of experimental designs. For both of these, there are extensive tutorials, documentation, and knowledgebase (see Note 17). We refer the reader to these resources to develop a workflow appropriate to their experimental design.

Tagwise Common Trend

PC2

0.6

1 2 3 0

Condition 1 2 3

0.2

0.4

Replicate

40

-40 0.0

Biological coefficient of variation

0.8

8. Quality assurance: While the DEseq2 and edgeR workflows produce differential expression analyses with corrected pvalues, it is still recommended to ensure the global structure of your results are reasonable. Two methods for doing this are to use principal component analysis (PCA) or hierarchical clustering to visualize sample clustering. The R functions prcomp () and hclust() work well for these types of analyses (see Note 18). Ideally the results would show tight clustering of the replicates, with separation of samples by other biological conditions as is reasonable, for example treatment and control samples grouping together (see Fig. 2).

-5

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5 Average log CPM

10

15

-80

0

-40

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PC1

Fig. 2 (a) Mean–variance plot of reanalyzed RNAseq data from Warner et al. 2018. Y-axis is the Biological coefficient of variation (BCV), x-axis shows the average log counts per million of each feature CPM across all samples. The common BCV is ~0.21. Note the higher variance of lowly expressed features (average log CPM < 0). (b) A principal component analysis (PCA) of the same samples with PC1 shown on the x-axis and PC2 on the Y-axis. Note the grouping of the three replicates and clear separation of the samples by condition

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9. Batch correction (optional): If the samples being analyzed were sequenced on different platforms, or if they were produced from different research groups, the results can be strongly skewed by batch effects. These effects will be evident in the quality assurance step above. If this is the case, batch effect correction can be employed. The R package sva [30] includes tools for batch correction including the function ComBat() (see Notes 19 and 20). 10. GO term enrichment: Differential testing will yield a list of genes enriched in a sample of interest (e.g., timepoint or condition). Browsing this list can help develop hypotheses but often these lists are large and difficult to interpret. These gene lists can be further mined by testing for significant enrichment of gene ontology (GO) terms. To do this, the gene IDs or transcript IDs must be mapped to GO terms. Nemve1 includes associated GO terms on the download tab of the web browser (https://mycocosm.jgi.doe.gov/Nemve1/ Nemve1.home.html). An updated GO terms list for Nemve1 gene IDs, based on best blast hit to the Uniprot databases can be found at (http://ircan.unice.fr/ER/ER_plotter/about). These GO terms can be used as a background set for enrichment testing where the differentially expressed gene list is used as a test set. The R package topGO can be used to calculate GO term enrichment (see Note 21).

4

Notes 1. Tri reagent, phenol, and chloroform should be used under an appropriate fume extraction hood. 2. If harvesting RNA from multiple samples, work each sample one at a time, not in parallel. This helps minimize the time between Tri-reagent addition, lysis, and freezing helping to prevent RNA degradation. 3. If harvesting RNA from adult tissues, use a larger amount of Tri-reagent. The sample should not exceed 10% of the Tri-reagent volume. 4. Phase Lock tubes are optional but can increase yield and reduce contamination. 5. This second extraction increases RNA purity. 6. For certain commercial phenol–chloroform preparations, not adding the supplied buffer results in the required pH of 6.0. 7. An RNA pellet will appear as a whitish-tan opaque speck on the bottom of the tube. Holding the tube up to a window while removing the liquid can help avoid aspirating the pellet.

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8. Excessive white precipitate can indicate contamination by salts or other debris during the extraction. These samples may be rescued by resuspending in nuclease free water and performing an additional phenol–chloroform extraction followed by precipitation. 9. When dry, the pellet will appear glass-like and lose its color. Hold tube up to a window or light to help visualize the pellet. 10. Pellets can occasionally be difficult to resuspend if dried for longer than necessary. Be patient and pipet until small particles are no longer observed when pipetting up and down. 11. For DNase treatment we strongly recommend using a DNase kit with bead-based inactivation as heat inactivation can result in RNA degradation. 12. If using a DNAse treatment set aside 1 μl of untreated RNA for quality assurance. This hold back sample can be analyzed during trouble shooting in the event the final RNA is degraded and rule out the DNAse treatment as a source of the degradation. 13. Typically formaldehyde-based gels are used to assess RNA quality. An appropriate substitute, however, is to use a “Bleach gel” as described in [31]. 14. There are many Next Generation Sequencing library preparation kits commercially available. The major considerations in selecting a kit include whether or not the kit produces “stranded” libraries, whether or not the kit selects for mRNA or other types of RNA, the size of the fragments produced, and the compatibility with sequencing technologies. 15. If the transcriptomic data is to be used for a de novo transcriptome assembly, read trimming is essential. 16. Transcript clustering algorithms that aim to reduce redundancy such as Cd-hit can be used to approximate “gene level” quantification [32]. It should be noted, however, that these programs rarely reduce transcriptome redundancy to the number of “true genes,” often resulting in several contigs per gene. This can be problematic for gene level quantification as it inflates gene copy number and will skew downstream results. 17. As of writing, there are two excellent tutorials for DEseq2 and edgeR which include detailed explanations and code examples: (http://bioconductor.org/packages/release/bioc/vignettes/ DESeq2/inst/doc/DESeq2.html; https://www.bio conductor.org/packages/release/bioc/vignettes/edgeR/ inst/doc/edgeRUsersGuide.pdf). 18. Log2 normalization of the data can help reduce the distances/ heights in PCA and hierarchical clustering analyses.

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19. Any batch correction should be performed as conservatively as possible to avoid introducing inappropriate bias to the data. For example, batches should be assigned as categorical covariates and other phenotypic information including timepoint, treatment or condition should not be included. 20. The ComBat() function does not perform well with excessive missing or 0 values. These should be removed prior to batch correction. 21. topGO includes related GO terms for testing. As such, terms may be identified as significant that are not included in the GO mappings list. The “depth” of these related terms can be adjusted in the analysis. For more information please see: http://geneontology.org/docs/ontology-relations/. References 1. Hand C, Uhlinger KR (1992) The culture, sexual and asexual reproduction, and growth of the sea anemone Nematostella vectensis. Biol Bull 182(2):169–176 2. Reitzel A, Burton P, Krone C, Finnerty J (2007) Comparison of developmental trajectories in the starlet sea anemone Nematostella vectensis: embryogenesis, regeneration, and two forms of asexual fission. Invertebr Biol 126(2):99–112 3. Passamaneck YJ, Martindale MQ (2012) Cell proliferation is necessary for the regeneration of oral structures in the anthozoan cnidarian Nematostella vectensis. BMC Dev Biol 12 (1):1–1 4. Bossert PE, Dunn MP, Thomsen GH (2013) A staging system for the regeneration of a polyp from the aboral physa of the anthozoan cnidarian Nematostella vectensis. Dev Dyn 242:1320–1331 5. Amiel AR, Johnston HT, Nedoncelle K, Warner JF, Ferreira S, Ro¨ttinger E (2015) Characterization of morphological and cellular events underlying oral regeneration in the sea anemone, Nematostella vectensis. Int J Mol Sci 16(12):28449–28471 6. Putnam NH, Srivastava M, Hellsten U, Dirks B, Chapman J, Salamov A et al (2007) Sea anemone genome reveals ancestral eumetazoan gene repertoire and genomic organization. Science 317(5834):86–94 7. Schwaiger M, Schonauer A, Rendeiro AF, Pribitzer C, Schauer A, Gilles AF et al (2014) Evolutionary conservation of the eumetazoan gene regulatory landscape. Genome Res 24 (4):639–650

8. Rentzsch F, Fritzenwanker JH, Scholz CB, Technau U (2008) FGF signalling controls formation of the apical sensory organ in the cnidarian Nematostella vectensis. Development 135(10):1761–1769 9. Ro¨ttinger E, Dahlin P, Martindale MQ (2012) A framework for the establishment of a Cnidarian Gene Regulatory Network for “Endomesoderm” specification: the inputs of ß-catenin/ TCF signaling. PLoS Genet 8(12):e1003164 10. Layden MJ, Ro¨ttinger E, Wolenski FS, Gilmore TD, Martindale MQ (2013) Microinjection of mRNA or morpholinos for reverse genetic analysis in the starlet sea anemone, Nematostella vectensis. Nat Protoc 8(5):924–934 11. Servetnick MD, Steinworth B, Babonis LS, Simmons D, Salinas-Saavedra M, Martindale MQ (2017) Cas9-mediated excision of Nematostella brachyury disrupts endoderm development, pharynx formation and oral-aboral patterning. Development 144(16):2951–2960 12. Kraus Y, Aman A, Technau U, Genikhovich G (2016) Pre-bilaterian origin of the blastoporal axial organizer. Nat Commun 7:11694 13. Ikmi A, McKinney SA, Delventhal KM, Gibson MC (2014) TALEN and CRISPR/Cas9mediated genome editing in the earlybranching metazoan Nematostella vectensis. Nat Commun 5:5486 14. Fischer A, Smith J. Nematostella high-density RNAseq time-course. 2013 15. Tulin S, Aguiar D, Istrail S, Smith J (2013) A quantitative reference transcriptome for Nematostella vectensis early embryonic development: a pipeline for de novo assembly in emerging model systems. EvoDevo 4(1):16

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16. Helm RR, Siebert S, Tulin S, Smith J, Dunn CW (2013) Characterization of differential transcript abundance through time during Nematostella vectensis development. BMC Genomics 14(1):266 17. Warner JF, Guerlais V, Amiel AR, Johnston H, Nedoncelle K, Ro¨ttinger E (2018) NvERTx: a gene expression database to compare embryogenesis and regeneration in the sea anemone Nematostella vectensis. Development 145(10): dev162867 18. Fredman D, Schwaiger M, Rentzsch F, Rentzsch F, Technau U (2013) Nematostella vectensis transcriptome and gene models v2.0. 1–1 19. Wingett SW, Andrews S (2018) FastQ Screen: A tool for multi-genome mapping and quality control. F1000Res 7:1338 20. Bolger AM, Lohse M, Usadel B (2014) Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30 (15):2114–2120 21. Martin M (2011) Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBNetjournal 17:10–12 22. Kim D, Langmead B, Salzberg SL (2015) HISAT: a fast spliced aligner with low memory requirements. Nat Methods 12(4):357–360 23. Dobin A, Davis CA, Schlesinger F, Drenkow J, Zaleski C, Jha S et al (2013) STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29 (1):15–21 24. Langmead B, Salzberg SL (2012) Fast gappedread alignment with Bowtie 2. Nat Methods 9 (4):357–359

25. Li H, Durbin R (2009) Fast and accurate short read alignment with burrows-wheeler transform. Bioinformatics 25(14):1754–1760 26. Liao Y, Smyth GK, Shi W (2014) featureCounts: an efficient general purpose program for assigning sequence reads to genomic features. Bioinformatics 30(7):923–930 27. Li B, Dewey CN (2011) RSEM: accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC Bioinformatics 12(1):323 28. Love MI, Huber W, Anders S (2014) Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol BioMed Central 15(12):550 29. Robinson MD, McCarthy DJ, Smyth GK (2010) edgeR: a bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26 (1):139–140 30. Leek JT, Johnson WE, Parker HS, Jaffe AE, Storey JD (2012) The sva package for removing batch effects and other unwanted variation in high-throughput experiments. Bioinformatics 28(6):882–883 31. Aranda PS, LaJoie DM, Jorcyk CL (2012) Bleach gel: a simple agarose gel for analyzing RNA quality. Electrophoresis 33(2):366–369 32. Li W, Godzik A (2006) Cd-hit: a fast program for clustering and comparing large sets of protein or nucleotide sequences. Bioinformatics 22(13):1658–1659

Chapter 15 RNA Interference on Regenerating Holothurian Gut Tissues Miosotis Alicea-Delgado, Samir A. Bello-Melo, and Jose´ E. Garcı´a-Arrara´s Abstract Functional studies on echinoderms have been reduced to the use of pharmacological treatments. The ability to modulate the genetic expression of regenerating tissues can elucidate potential effectors during this process. Here we describe an effective transfection protocol that allows the introduction of Dicer-substrate interference RNAs (DsiRNAs) for the modulation of gene expression and its characterization during regeneration. Key words Echinoderm, Sea Cucumber, DsiRNA, RNAi, Regeneration, Electroporation, Gut rudiment

1

Introduction RNA interference (RNAi) is a well-conserved silencing mechanism found in most eukaryotes [1]. Originally, this mechanism was recognized as an endogenous antiviral protector and gene regulator, but now it has been harnessed as an experimental tool for gene knockdowns [2]. The formal effectors of this silencing mechanism are the double-stranded RNAs (dsRNAs), which are processed into short-interfering RNAs (siRNAs), ~21–22 nucleotides in length, by the protein Dicer. Then, these siRNAs are incorporated into the RNA-induced silencing complex (RISC), which recognizes, cleaves, and/or represses complementary RNA [3, 4]. Previously, many researchers synthesize 21-mer siRNAs, which mimic the natural siRNAs obtained from Dicer processing. IDT researchers have advanced this technology by designing a more effective RNAi technology called Dicer-substrate interference RNA (DsiRNA). DsiRNAs are 27-mers RNA duplexes processed by Dicer into 21-mer siRNAs [5]. With this technology scientists provide Dicer with a substrate instead of a product, thus increasing RNAi potency up to 100-fold when compared with usual 21-mers [5].

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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There are few techniques available to knockdown specific gene expression in members of the phylum Echinodermata, thus limiting the type of studies that can be performed on these animals. Echinoderms, with their key phylogenetic position and their impressive regeneration potential have much to offer to studies of evolution and developmental interest. At present, most studies can only be performed in embryos but not in adult animals [6]. Gene knockdown techniques have been performed recently in certain tissues/ organs of adult echinoderms but still remain highly limited [7]. We have now implemented DsiRNA technology in tissue explants of the sea cucumber Holothuria glaberrima. For this, gut rudiments were electroporated with tetramethylrhodamineconjugated anionic dextran 3000 MW lysine fixable using different parameters to elucidate the efficiency of the transfection method. After selecting the optimal parameter, DsiRNA-targeting β-catenin was introduced into the intestinal tissue explants to reduce its abundance and characterize its role on different cellular events during intestinal regeneration.

2

Materials Prepare all solutions using ultrapure water and analytical-grade reagents. Diligently follow waste disposal regulations when disposing of waste materials. All cell culture procedures should be carried out in a laminar flow hood.

2.1

Tissue Selection

2.2 Gut Rudiments Culture

In our case, we selected the gut rudiments to perform our experiments and followed the procedure of the sea cucumber evisceration, disinfection and gut dissection found in Bello et al. [8], but this protocol can be applied to any tissue of interest. 1. L-15 Medium (Leibovitz, Sigma): conditioned for marine species by adding salts to the original composition [9]. Weigh 6.9 g L-15 powder, 6.25 g NaCl, 3.12 g glucose, 1.58 g magnesium sulfate (MgSO4), 172 mg KCl, 96 mg sodium bicarbonate (NaHCO3), 1.33 g magnesium chloride (MgCl2), 150 mg L-glutamine, 745 mg calcium chloride dihydrate (CaCl22H2O) and transfer to a 500 mL beaker. Complete to 500 mL with ultrapure water. Mix with a magnetic stirrer and sterilize by filtration (0.22 μm filters). Store in a bottle wrapped in aluminum foil at 4  C. 2. Gut rudiment medium: L-15 medium conditioned for marine species supplemented with antibiotics (100 U/mL Penicillin, 100 μg/mL Streptomycin, 50 μg/mL Gentamicin), antifungal (2.5 μg/mL amphotericin B), 1  MEM nonessential amino acids, 1 mM sodium pyruvate, 1.75 mg/mL α-tocopherol

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acetate. Add 500 μL of Penicillin/Streptomycin stock solution (10,000 U Penicillin and 10 mg/mL Streptomycin, respectively), 250 μL of Gentamicin stock solution (10 mg/mL), 50 μL of amphotericin B stock solution (2.5 mg/mL), 500 μL of MEM nonessential amino acid stock solution (100), 500 μL of sodium pyruvate stock solution (100 mM), and 50 μL of α-tocopherol acetate stock solution (17.5 mg/mL) in a 50 mL tube. Complete 45 mL of the 50 mL with L-15 medium conditioned for marine species. Mix by inversion of the tube and adjust to pH 7.7–7.8. Complete to 50 mL adding L-15 medium conditioned for marine species and sterilize by filtration (0.22 μm filters). Store in a bottle wrapped in aluminum foil at 4  C (see Note 1). 3. 24-well plate: tissue culture treated polystyrene, flat bottom with lid. 2.3 Gut Rudiment Electroporation

1. Phosphate Buffered Saline (PBS) 0.1 M: Weigh 8 g NaCl and 13.4 g phosphate dibasic heptahydrate (NaH2PO47H2O) and transfer to a 1 L beaker. Add 800 mL of ultrapure water while stirring. Adjust to pH 7.4 and complete to 1 L with ultrapure water. 2. Tetramethylrhodamine-conjugated anionic dextran (TMRAD) 3000 MW, lysine fixable: TMRAD 50 mg/mL stock solution. Dilute to 5 μg/μL TMRAD in 0.01 M PBS for every sample (see Note 2). 3. BTX Harvard Apparatus ECM 830 Square Wave Electroporation System and electroporation cuvettes: 4 mm electroporation cuvettes. 4. Safety Stands for BTX Cuvettes & Chambers.

2.4 Determining Electroporation Efficiency Using Histological Approaches

1. 4% paraformaldehyde (PFA, Sigma; see Note 3): Weigh 4 g of PFA and dissolve it on 50 mL of ddH2O. Heat the solution up to 60  C in a chemical hood. Add sodium hydroxide (NaOH), pellet by pellet, until the solution becomes transparent. Add 50 mL of 0.2 M PB. Put the solution on ice and adjust to pH 7.4. 2. Sucrose: 40% sucrose in 0.1 M PBS. 3. Mounting media: buffered glycerol solution containing 2 μg/μ L 40 ,6-diamidino-2-phenylindole (DAPI). To prepare 5 mg/ mL DAPI stock solution (14.3 mM), dissolve one vial (10 mg) in 2 mL of ddH2O. Then prepare a 1 mg/mL DAPI working solution adding 200 μL of the stock solution to 800 μL ddH2O. To prepare the mounting media add 50 μL of the working solution to 24.95 mL of 0.1 M PBS in a 50 mL. Then, add 50 mL of glycerol to the 50 mL tube and mix. Wrap the tube with aluminum paper and store at 4  C.

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DsiRNAs

1. Nuclease-Free Duplex Buffer from Integrated DNA Technologies (IDT). 2. DsiRNAs 2 and/or 10 nmol (see Note 4): For 2 nmol and 10 nmol of DsiRNAs, add 20 μL and 100 μL of Nuclease-Free Duplex Buffer, respectively to obtain a stock solution of 100 μM. Heat at 94  C for 2 min. Dilute the oligonucleotides to 10 μM using Nuclease-Free Duplex Buffer. Store at 20  C.

2.6 Determination of RNA Abundance Using qPCR

1. iQ SYBR Green Supermix (Bio Rad): Add 12.5 μL of iQ SYBR Green Supermix. 2. Nuclease-free water. 3. qPCR primers 10 μM: For 100 μM stock solution, add Nuclease-Free water to the lyophilized primers (see Note 5). Heat the oligonucleotide at 55  C for 5 min, then vortex thoroughly. Prepare a 10 μM working solution and store at 20  C.

3

Methods

3.1 Sea Cucumber Collection and Evisceration

1. Collect the sea cucumbers from the species Holothuria glaberrima in the Northeast coast of Puerto Rico and keep in indoor saltwater aquaria at 22–24  C. 2. Eviscerate the sea cucumbers by injecting 0.35 M KCl (3–5 mL) into the coelomic cavity. During the process of evisceration, H. glaberrima expels out most of its internal organs, including its intestine, through the cloaca. After evisceration, sea cucumbers are placed in an artificial aquarium for 4 days.

3.2 Sea Cucumber Disinfection and Explant Preparation

1. Immerse the sea cucumbers (see Note 6) that have regenerated their intestinal tract for 4 days in a dish preparation containing 250 mL of anesthetic solution (RT) for 35 min. 2. Decontaminate the exterior of the sea cucumbers by immersing the animals one at a time in a 10% sodium hypochlorite solution for 1 min, 95% ethanol for 1 min, and a quick rinse in sterile ultrapure water. 3. Spray a vinyl pad in a dissecting pad with 70% ethanol and place the sea cucumbers ventral side down. The ventral side of the sea cucumbers can be identified by the presence of the ambulacral tubes. Cut the anterior region (head) near the calcareous ring and then cut with scissors along the longitudinal line that separates the dorsal and ventral axis. Once the body wall is opened, hold down the sea cucumber using pushpins. Dissect the gut rudiments using fine tweezers and microdissection scissors. On day 4 of regeneration, the gut rudiment appears as a pink thickening at the free end of the mesentery.

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4. Collect the gut rudiment (see Note 7) and place each, individually, in 1 mL of the antibiotic/antifungal solution in a 2 mL tube. 5. Wash the gut rudiments with fresh antibiotic/antifungal solution by placing the tubes in a shaker (moving slowly) for 1 h twice. 3.3 Optimization of the Electroporation Protocol

1. Prepare the 24-well plate with 1 mL of the L-15 supplemented with antibiotics and antifungal. 2. Add 25 μL of 5 μg/μL TMRAD on each electroporation cuvette. 3. Remove the gut rudiment from the antibiotic/antifungal solution with a tweezer or a pipette tip (see Note 8) and place it inside the electroporation cuvette. 4. Place the electroporation cuvette in the Safety Stands for Cuvettes and apply the square electric pulses ten times using the following parameters (see Note 9): 15 V, time pulse: 45 ms, interval pulse: 955 ms; 20 V, time pulse: 40 ms, interval pulse: 960 ms; 27 V, time pulse: 30 ms, interval pulse: 970 ms; 35 V, time pulse: 25 ms, 975 ms. 5. Remove the gut rudiment from the electroporation cuvette and place it in a 24-well plate prepared on step 1. 6. Incubate the gut rudiment for 2 days in a modular incubator chamber at RT. Check your culture daily to evaluate its morphology and possible contamination.

3.4 Histological Studies for Electroporation Efficiency

1. After culturing the gut rudiments for 2 days, use a tweezer or a pipette tip to remove the gut rudiment from the 24-well plate. Place the gut rudiments into the 2 mL tubes with 1 mL of 4% PFA for 24 h and store at 4  C. 2. After fixation, remove the 4% PFA from the 2 mL tubes and wash the gut rudiments with 0.1 M PBS three times for 10 min/each. 3. Add 1 mL of 30–40% sucrose to each 2 mL tube containing the gut rudiments for cryoprotection and store at 4  C for 24 h (see Note 10). 4. Add Tissue-Tek O. C. T in a metal base mold for the gut rudiment embedding (see Note 11). 5. Use a tweezer to remove the gut rudiments from the 30–40% sucrose and place it in a petri dish to cut ~2.5 cm of the gut rudiment. Place the remaining of the gut rudiments back to the 30–40% sucrose and store at 4  C as backup. 6. Use a tweezer to put the gut rudiment that has been cut in the metal base mold with the Tissue-Tek O. C. T. (see Note 12).

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Fig. 1 Electroporation of tetramethylrhodamine-conjugated anionic dextran in explants of the sea cucumber H. glaberrima. Nuclei are stained with DAPI (green) and tetramethylrhodamine-conjugated anionic dextran stained the cytosol (red). (a) Electroporation of the explant with 0.1 M PBS. (b) Tetramethylrhodamineconjugated anionic dextran alone (without electroporation). (c) Electroporation of the explant with tetramethylrhodamine-conjugated anionic dextran. (d) Higher magnification of (c)

7. Place the metal base mold with the gut rudiments into the cryostat (see Note 13) until the Tissue-Tek O. C. T. solidifies. 8. Place the block of Tissue-Tek O. C. T. in the microtome and perform 20 μm cryosections. Place the cryosections on poly-Llysine–treated slides; 5 or 6 sections per slide. 9. After cryosectioning, dry the slides for 1 h before doing any histological study. 10. Mount each slide in buffered glycerol containing DAPI and cover with a 24  55 mm coverslip. Seal the slides with nail polish and dry under a blower. DAPI stains the cell nuclei. 11. Evaluate the slides using a fluorescent microscope or a microscope equipped with the appropriate filters (see Note 14) (Fig. 1) [10]. 3.5

siRNA Design

1. Access siDirect version 2.0 (e.g., at http:sidirect2.rnai.jp) and paste the nucleotide sequence of interest to choose the target sequence and design the siRNA. This program will retrieve different effective siRNA candidates and a graphical view of these candidates [10]. 2. Select the candidates with a seed-duplex stability Tm < 21.5  C. These are off-target reduced siRNAs, which are siRNAs that interfere specifically the sequence of interest and have a less probability to interfere other sequences. 3. After selecting the siRNA duplex, use the guidelines of Integrated DNA Technologies (IDT), Inc. to design the DsiRNA-targeting β-catenin: (a) Target sequence and siRNA duplexes:

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50 – GGGTGTAACAACTTGACAATTAC–30 . 5’–GUGUAACAACUUGACAAUUAC–3’ (sense) strand.

Passenger

3’–CCCACAUUGUUGAACUGUUAA–50 Guide (antisense) strand. (b) To create a Dicer substrate RNAi, add four bases to the 30 -end of the sense strand and six bases to the 50 -end of the antisense strand to create an asymmetric blunt ended and 30 overhang molecule. 5’–GGGTGTAACAACTTGACAATTAC–30 5’–GUGUAACAACUUGACAAUUACAGGA–30 . 3’–CCCACAUUGUUGAACUGUUAAUGUCCU–5’ (c) Replace two bases of the 30 -end of the sense strands with DNA bases when necessary. 50 – GGGTGTAACAACTTGACAATTAC– 30 . 5’–GUGUAACAACUUGACAAUUACAGGA– 30 . 3’–CCCACAUUGUUGAACUGUUAAUGUCCU– 50 . 3.6 Electroporation of Gut Rudiments with DsiRNAs

1. Follow Subheading 3.2. 2. Prepare the 24-well plate with 1 mL of the L-15 supplemented with antibiotics and antifungal. 3. Add 5 μL of three different concentrations of DsiRNAtargeting β-catenin on each electroporation cuvette: 1 μM, 10 μM, and 100 μM. Use biological replicates. 4. Remove the gut rudiment from the antibiotic/antifungal solution with a tweezer or a pipette tip (see Note 8) and place it inside the electroporation cuvette. 5. Place the electroporation cuvette in the Safety Stands for Cuvettes and apply the square electric pulses ten times using the following parameters: 35 V, time pulse: 25 ms, 975 ms. 6. Remove the gut rudiment from the electroporation cuvette and place it in a 24-well plate prepared in step 2. 7. Incubate the gut rudiment for 2 days in a modular incubator chamber at RT. Check your culture daily to evaluate its morphology and possible contamination. 8. After 2 days in culture, remove the gut rudiments from the 24-well plate and store at 20  C in 1 mL of RNAlater™ in a 2 mL tube.

3.7

RNA Extraction

1. Place the 2 mL tubes with the gut rudiments on ice. 2. Prepare five 15 mL tubes: three 15 mL tubes with 10 mL ddH2O, one with 10 mL 70% ethanol and one with 10 mL

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ZAP. Place the tubes in the following order: 2 ddH2O, 70% ethanol, ZAP, and ddH2O. 3. Take out the RNA later from the 2 mL tubes of step 8 from Subheading 3.6 and add 1 mL of Trizol (maintain the tubes on ice). 4. Homogenize the gut rudiments at low/medium level using a homogenizer until the tissue is completely homogenized. 5. After homogenizing the gut rudiments, place the tubes on ice for 30 min. 6. After 30 min on ice, add 200 μL of chloroform and mix vigorously. 7. Incubate the tubes at RT for 10 min. 8. Centrifuge the tubes for 15 min at 12,000 rpm (14167  g) at 4  C (see Note 15). 9. Remove the aqueous phase (RNA is here!—see Note 16) and place it in a 1.5 mL RNase-free tube. 10. Follow the Qiagen RNeasy Protocol for RNA purification, including the DNase treatment (Cat. Num. 74104). 11. Place the 1.5 mL tubes with the purified RNA on ice and determine the RNA concentration using the NanoDrop 1000 spectrophotometer. Write down the concentration, the 260/280, and the 260/230 ratios. 3.8

cDNA Synthesis

1. Calculate the volume needed for 1 μg of RNA extracted from the gut rudiments starting from the RNA concentration obtained in Methods 3.7. 2. In a 0.2 mL tube add 2 μL of Oligo dT, the volume calculated for 1 μg of RNA and complete to 10 μL with nuclease-free water. 3. Mix, spin briefly and heat for 5 min at 70  C. 4. Add the remaining components to the 0.2 mL tube (see Note 17): 4 μL Improm-2 5 Reaction Buffer, 3 μL MgCl2, 1 μL dNTPs, 1 μL RNasin Ribonuclease inhibitor, and 1 μL Improm-2 Reverse transcriptase to obtain a final volume of 20 μL. 5. Mix by pipetting up and down, incubate at 25  C (RT) for 5 min, 42  C for 1 h and 70  C for 15 min (see Note 18).

3.9 qPCR to Determine the Efficiency of the Interference

1. Place the cDNA samples, the primers (see Note 19), the iQ SYBR Green Supermix on ice. 2. Add 12.5 μL iQ SYBR Green Supermix, 0.4 μL primer forward, 0.4 μL primer reverse, 5.7 μL nuclease-free water, and 1 μL of the cDNA sample on each well on a 96-well plate (see

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Dsi -catenin Log2 fold change

4

1 µM 10 µM 100 µM

2 0 -2 -4

**

Fig. 2 RNA interference promotes β-catenin downregulation. To evaluate the efficiency of the interference of β-catenin transcript, qRT-PCR of intestinal explants was performed after 2 days of DsiRNA electroporation. Expression values were plotted as fold change relative to DsiRNA-targeting GFP (negative control) treatment in log2 scale. N ¼ 4

Note 20). Perform technical replicates of each biological replicates. Use primers for β-catenin and NADH (reference gene). 3. Put the 96-well plate in a Mastercycler Ep realplex and use the following parameters: (a) Step 1: 95  C for 3:00 min. (b) Step 2: 95  C for 15 s. (c) Step 3: 57  C for 30 s. (d) Step 4: 60  C for 40 s. (e) Melting curve. 4. Analyze the data using the Livak Method or the analysis of relative gene expression. Perform a one-way ANOVA for statistical analyses (see Note 21) (Fig. 2) [11].

4

Notes 1. Antibiotics and antifungals lose their activity in ~3–5 days in culture medium. Also, other supplements lose their activity in a few days in culture. Thus, it is recommended to supplement the media one day before the experiment and change the media every 3–5 days. The L-15 supplemented medium can be stored at 4  C up to 2 weeks. 2. This dilution corresponds to one gut rudiment. For more samples, take into account the total volume needed on each electroporation cuvette for each gut rudiment and prepare a master mix.

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3. For 4% PFA preparation wear gloves, laboratory glasses and a face mask. Prepare the solution inside a chemical hood. For 0.2 M Phosphate Buffer (PB), prepare Solution A dissolving 13.8 g of NaH2PO4H2O in 500 mL ddH2O and for Solution B dissolve 14.2 g of Na2HPO4 -anhydrous in 500 mL ddH2O. Mix 19 mL of Solution A and 81 mL of Solution B to obtain 0.2 M PB. 4. Purchase the amount of DsiRNAs according to the number of samples and the concentration needed. Centrifuge the tube with DsiRNA previous to its resuspension to avoid any loss of material. 5. The amount of Nuclease-free water depends on the nmol of the primers. The formula used is: nmol  10, according to IDT. 6. The number of sea cucumbers is not established in this part of the protocol because it depends on the requirements of the experiments. 7. In this particular experiment, we used the gut rudiment, but this protocol can be applied to the tissue of interest. 8. Gut rudiments are delicate during this regenerative stage. Thus, use a pipette tip as a hook to transport the gut rudiments from one place to another. If using a tweezer, try not to squeeze the gut rudiment. 9. In this part of the experiment, we used 18 sea cucumbers, three sea cucumbers per parameter. We were trying to elucidate the optimal parameter to accomplish the transfection of the TMRAD and, eventually the transfection of the DsiRNAs. This part of the protocol is very important for elucidating the optimal parameter of a particular tissue. It is very important to mention that every tissue has a different response to the process of electroporation. So, these parameters can be used as a reference for the electroporation of other tissues. 10. Gut rudiments are placed in 30–40% sucrose for cryoprotection. 24 h is enough for the sucrose to be absorbed by the gut rudiments, but another signal is when the tissues sink to the bottom of the 2 mL tube containing the 30–40% sucrose. 11. Gut rudiments embedding are prepared at RT because TissueTek O. C. T. freezes at lower temperatures. 12. As gut rudiments are long, but thin, several replicates can be embedded in a metal base mold. 13. Set the cryostat temperature on 35  C. Perform the cryosections from 27 to 35  C to obtain better results. Set the temperature 1 h before starting to cut. 14. In our laboratory, histological studies are examined and photographed using a Nikon Eclipse Ni fluorescent microscope

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equipped with DAPI, FITC and R/D2 filters and a DS-Qi2 camera. Images were obtained using the software NIS-Elements. 15. Set the temperature of the centrifuge to 4  C before starting the RNA extraction protocol. After this centrifugation, set the temperature to 22–24  C and press the “fast temp” button; RNA purification is performed at room temperature. 16. The RNA extraction using TRIzol/chloroform produce three phases after the centrifugation. Proteins are extracted to the organic phase (bottom of the tube), DNA resolves at the interface and RNA remains in the aqueous phase (top of the tube). 17. In our laboratory, we prepare master mixes for the remaining reagents of the cDNA synthesis. Thus, we multiply the volume of each reagent by the total number of samples including an extra sample. 18. We set these parameters in a thermocycler. We add our first 10 μL of the protocol (Oligo dT, nuclease-free water, and cDNA), put the tubes in the thermocycler to start the first part of the protocol. Then, we add the remaining components to each tube and put the tubes in the thermocycler to finish the protocol. 19. The primers were designed to flank the target sequence of the RNAi to assure that the knockdown was efficiently done. 20. To prepare the 96-well plate, we prepare master mixes for each pair of primers. In this case, we prepare master mixes for β-catenin and NADH. The master mixes are prepared taking into account the total number of samples and technical replicates of the biological replicates. We prepare 3 technical replicates for each sample. Thus, if we are going to calculate the total volume for the forward and/or reverse primers for 12 samples, we multiply 0.4 μL  3.5 (technical replicates)  13 (extra sample). This applies also for the iQ SYBR Green Supermix and the nuclease-free water. After that, we prepare twelve 0.2 mL tubes (one per sample) and add 66.5 μL of the master mix for β-catenin and/or NADH, and 3.5 μL of the cDNA samples. We have now 12 mini master mixes with 70 μL corresponding to the technical replicates of each biological sample. At the end, each well will have 20 μL of three technical replicates for each biological sample. 21. Results present three different concentration treatments, 1, 10, and 100 μM. By doing this gradient, we wanted to elucidate the optimal DsiRNA concentration for β-catenin knockdown.

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Acknowledgments This project was supported by NIH (Grants R15GM124595 and R21AG057974), and the University of Puerto Rico. References 1. Hannon GJ (2002) RNA interference. Nature 418:244–251 2. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC (1998) Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391:806–811 3. Elbashir SM, Lendeckel W, Tuschl T (2001) RNA interference is mediated by 21- and 22nucleotide RNAs. Genes Dev 15:188–200 4. Meister G, Tuschi T (2004) Mechanisms of gene silencing by double-stranded RNA. Nature 431:343–349 5. Kim DH, Behlke MA, Rose SD, Chang MS, Choi S, Rossi JJ (2005) Synthetic dsRNA dicer substrates enhance RNAi potency and efficacy. Nat Biotechnol 23:222–226 6. Bogarad L, Arnone MI, Chang C, Davidson EH (1998) Interference with gene regulation in living sea urchin embryos: transcription factor knock out (TKO), a genetically controlled vector for blockade of specific transcription factor. PNAS 25:14827–14832 7. Mashanov VS, Zueva OR, Garcı´a-Arrara´s JE (2015) Myc regulated programmed-cell death

and radial glia dedifferentiation after neural injury in an echinoderm. BMC Dev Biol 15:1–9 8. Bello SA, Abreu-Irizarry RJ, Garcı´a-Arrara´s JE (2015) Primary cell cultures of regenerating holothurian tissues. Methods Mol Biol 1189:283–297 9. Schacher S, Proshansky E (1983) Neurite regeneration by Aplysia neurons in dissociated cell culture: modulation by Aplysia hemolymph and the presence of the initial axonal segment. J Neurosci 3:2403–2413 10. Naito Y, Yoshimura J, Morishita S, Ui-Tei K (2009) siDirect 2.0: updated software for designing functional siRNA with reduced seed-dependent off-target effect. BMC Bioinformatics 10:392 11. Alicea-Delgado M (2019) Functional role of β-catenin and Myc as active players in the canonical Wnt signaling pathway during intestinal regeneration of the sea cucumber Holothuria glaberrima. Thesis, University of Puerto Rico – Rı´o Piedras Campus

Chapter 16 ATAC-Seq for Assaying Chromatin Accessibility Protocol Using Echinoderm Embryos Marta S. Magri, Danila Voronov, Jovana Ranđelovic´, Claudia Cuomo, Jose Luis Go´mez-Skarmeta, and Maria I. Arnone Abstract Cis-regulatory elements (CREs) and transcription factors (TFs) associated with them determine temporal and spatial domains of gene expression. Therefore, identification of these CREs and TFs is crucial to elucidating transcriptional programs across taxa. With chromatin accessibility facilitating transcription factor access to DNA, the identification of regions of open chromatin sheds light both on the function of the regulatory elements and their evolution, thus allowing the recognition of potential CREs. Buenrostro and colleagues have developed a novel method for exploring chromatin accessibility: assay for transposaseaccessible chromatin with high-throughput sequencing (ATAC-seq), which can be used for the purpose of identifying putative CREs. This method was shown to have considerable advantages when compared to traditional methods such as sequence conservation analyses or functional assays. Here we present the adaptation of the ATAC-seq method to echinoderm species and discuss how it can be used for CRE discovery. Key words Cis-regulatory element, ATAC-seq, Echinoderm, Development, Evolution

1

Introduction During embryonic development, cell proliferation and morphogenetic movements shaping the embryo are orchestrated by cascades of cell-signaling pathways resolved in the expression of developmental genes well conserved in the animal kingdom. A regulatory code of interactions between proteins that act as transcription factors (TFs) and cis-regulatory elements (CREs) placed in the non-coding DNA, such as enhancers and promoters, determines whether target genes are transcribed in a strict spatial and temporal domain. When TFs are codified in the nucleus and bind specific binding sites in accessible CREs, coordination of the expression of genes occurs in a highly regulated manner in order to avoid non-specific activation of other genes and, thus, allowing the correct development of an

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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organism [1, 2]. Moreover, genes with complex expression patterns have been shown to have several CREs that enhance specific expression in a particular territory or time during development [3–7]. Common morphological traits are related to a common ancestral gene tool kit and involve comparable transcriptional programs, where specific CREs have binding sites for homologous TFs [7– 12]. The sequence of CREs are characterized to be more exposed and therefore to be subjected to high rates of sequence turnover, compared to coding DNA, but less, compared to non-regulatory sequences in non-coding DNA. The generation of new TF binding sites due to random mutations in the DNA are likely to be established only in chromatin regions that are accessible to the corresponding TF proteins, suggesting that these regions constitute particularly favorable environments for studying regulatory innovations. Changes in the chromatin accessibility may modify the transcription status during development with repercussions on the body plan and might generate novel morphological traits. From an evolutionary point of view, to find ancestral CREs and study their functions would allow us to define gene regulatory networks shared across phyla and to unveil the basic developmental processes common to all metazoans. Identifying CREs is extremely challenging, and till recently, it was only possible through indirect and limited methods such as synteny and sequence conservation analysis or through laborintensive functional assays by cloning multiple regions to an expression vector and injecting into embryos [10, 13]. This has rapidly changed thanks to the advent of new techniques based on highthroughput sequencing. Among these techniques, the Assay for Transposase-Accessible Chromatin with next-generation sequencing (ATAC-seq) from Buenrostro et al., Nat. Methods 2013 [14] is a remarkably simple, flexible and powerful technique to profile chromatin regions genome-wide. It can be used in cell culture, dissected tissues and even in whole embryos. As the name suggests, this technique is based on the use of the hyperactive Tn5 transposase, which cuts exposed chromatin DNA into fragments (Fig. 1a). Because Tn5 is highly active, only a small number of cell nuclei are needed for the assay, as little as less than 105 cells (see Note 1). Another benefit of using the Tn5 transposase is that it simultaneously attaches Illumina sequencing adapters to the ends of the produced DNA fragments (Fig. 1b), thereby facilitating sequencing of libraries. Unlike similar methods, which can take up to 3 days to complete, ATAC-seq library preparation can be completed in 1 day. Sequencing reads can then be used to infer regions of increased accessibility, which can then be used in a range of downstream analyses [7, 12, 15, 16], such as to map regions of transcription factor binding sites with nucleosome position resolution (Fig. 1c). Active regulatory sites are, indeed, located in open chromatin

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Fig. 1 ATAC-seq reaction cartoon. (a) The Tn5 transposase (dark pink) inserts two sequencing adapters (red and blue) preferentially in accessible regions such as regions between nucleosomes (yellow), promoters (P) and enhancers bound by transcription factors (TF). (b) Tn5 generates sequencing fragments that can be amplified by PCR. (c) Mapped sequenced reads (black) identify open chromatin regions and nucleosome positions

regions, whereas the inactive ones are tightly wrapped around histones and are, therefore are protected from the transposase. Thanks to ATAC-seq data we can obtain a more precise and realistic picture of how regulatory information is organized in the genome. Comparing ATAC-seq data generated from pools of embryos at specific developmental stages we are able to study the dynamics of regulatory activity across the different developmental stages. Likewise, by applying this method to isolated tissues we are able to obtain specific regulatory information of discrete populations of cells. We present an ATAC-seq protocol performed in sea urchin embryos of Strongylocentrotus purpuratus, Paracentrotus lividus and in sea star Patiria miniata embryos, at different early developmental stages (Fig. 2). The handling of animals, gamete collection, in vitro fertilization and embryo culturing are described for each organism. Otherwise, the main steps of the protocol are common for all the organisms used and exceptions will be indicated. By employing this technique in echinoderm embryos, we propose this protocol as a tool to understand how changes in CREs have impacted the evolution of gene expression and body plans of this phylum.

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Embryo culture

Embryo collection

Cold fASW washes

Embryo Lysis

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3 times

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Gel Electrophoresis BioAnalyzer

Fig. 2 ATAC-seq protocol outline. Steps are numbered to emphasize the sequence of steps performed. The colours represent blocks of actions performed to obtain main intermediate results

2 2.1

Materials Animal Origin

1. Strongylocentrotus purpuratus and Patiria miniata animals are provided by Patrick Leahy (Kerchoff Marine Laboratory, California Institute of Technology, Pasadena, CA, USA) and kept in a closed tank system with circulating diluted Mediterranean Sea water at 36.02 ppt and 14  C at Stazione Zoologica Anton Dohrn until use. Pacific Ocean water at 35.5 ppt can also be used to keep Pacific animals and embryo cultures. 2. Paracentrotus lividus animals are obtained from the Bay of Naples and kept in open tank systems with circulating Mediterranean Sea water at 37.8 ppt salinity and 16  C at Stazione Zoologica Anton Dohrn.

Equipment

1. Glassware: 50 mL and 5 L glass beakers for in vitro fertilization.

2.2.1 In Vitro Fertilization and Embryo Collection

2. Petri dishes for P. miniata oocyte maturation and embryo collection.

2.2

3. Nitex meshes of 40 and 200 μm pore size. 4. 0.22 μm filters. 5. Pasteur pipettes. 6. Sea water at appropriate concentrations: 37.8 ppt for P. lividus and 34.02 ppt for S. purpuratus and P. miniata.

2.2.2 ATAC-Seq Library Preparation

1. Gloves and filter tips for micropipettes are to be used throughout the protocol in order to avoid possible contamination with exogenous DNA.

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2. All centrifugations for sample washings and for the precipitation of the nuclei are carried out at 4  C; therefore, a refrigerated centrifuge is needed. 3. ATAC-seq sample are generated from amplify DNA fragments PCR thermocycler and PCR tubes are needed. 4. Rocker table. 5. Heat block. 6. Neubauer or Malassez chambers are useful to count lysed nuclei in a fluorescent microscope. 7. Clear thin-walled 500 μL tubes for Qubit. 2.3

Reagents

Prepare all solutions using autoclaved milliQ water. 1. Filtered artificial sea water (fASW): 28.3 g NaCl, 0.77 g KCl, 5.41 g MgCl2·6H2O, 3.42 g MgSO4 or 7.13 g MgSO4·7H2O, 0.2 g NaHCO3, and 1.56 g CaCl2·2H2O per 1 L of autoclaved milliQ water. Adjust pH to 8.2. Filter through a 0.22 μm filter. The salinity of the resulting fASW is around 34–36 ppt which corresponds to Pacific Ocean water and is used for all echinoderm species described in this protocol for washing. 2. Prepare fresh lysis buffer by mixing 10 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, and 0.2% IGEPAL CA-630 (see Note 2) and keep it on ice. It is possible to use the same lysis buffer for consecutive ATAC-seq reactions on the same day. 3. Use DAPI 1000 diluted 1: 100 to count the nuclei by using a fluorescent microscope. 4. Tagmented DNA (TD) buffer and enzyme are provided by the Nextera Illumina kit (Illumina Cat #FC-121-1030). Alternatively, a 2 TD buffer can be prepared as follows: 20 mM Tris– HCl, 10 mM MgCl2, and 20% (vol/vol) dimethylformamide. This solution can be stored at 20  C for at least 6 months. 5. Use Qiagen MinElute kit (Qiagen, 28004) following the instructions for all DNA purifications (see Note 3). 6. Use NEB Next High-Fidelity 2 PCR Master Mix (New England Labs Cat #M0541) in order to amplify the libraries. Complete list of primers is available in Supplementary Information Table 1 Buenrostro et al., Nat. Methods 2013. 7. Final concentrations of purified samples are measured with Qubit dsDNA BR Assay Kit (Molecular Probes #Q32850). 8. Prepare 2% agarose gel in order to check sample quality.

2.4 Downloading Genome and Annotation

Genome and the annotation files for mapping of the ATAC-seq data can be downloaded at Echinobase (echinobase.org) (Kudtarkar and Cameron, 2017). S. purpuratus genome and gff3 annotation files can be found at http://www.echinobase.org/

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Echinobase/SpDownloads. P. miniata corresponding files can be found at http://www.echinobase.org/Echinobase/PmDownload. P. lividus genome data is not currently publicly available. Note that the genome version and annotation (gff3 or gtf) version must be the same.

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Methods

3.1 In Vitro Fertilization

Echinoderms do not display sexual dimorphism; thus, their sex can only be determined by examination of their gametes. For the species in question, eggs are of yellow/orange color, while sperm is white and much more viscous than the egg suspension. 1. Gametes for both sea urchin species (S. purpuratus and P. lividus) are obtained by vigorously shaking the animals until they spawn. Eggs are collected by placing the spawning female over a beaker with filtered sea water of appropriate salinity placed on ice, so that the animal aboral side would be partially submerged in the filtered sea water to allow gamete shedding into it and depositing at the bottom of the beaker. The salinity suitable for P. lividus embryo cultures is 37.8 ppt corresponding to Mediterranean Sea water salinity. The salinity for S. purpuratus and P. miniata embryo culture is 34.02 ppt obtained by diluting Mediterranean Sea water 9:1 with distilled water. 2. Sperm is collected from the aboral surface of male sea urchins using a P200 micropipette and pipette tips into a 1.5 mL Eppendorf tube placed on ice (dry sperm). 3. Gametes for the sea star P. miniata are obtained surgically by making a V-shaped incision next to gonads on the aboral side of the animal, prying open the incision carefully with forceps and using another forceps to take part of the gonad. The female gonads are placed into petri dishes with filtered sea water of appropriate salinity. Male gonads are collected into 1.5 mL Eppendorf tube placed on ice. Female gonads are then torn apart using a pair of forceps under a dissecting microscope to release oocytes into the sea water. The oocytes are then treated with 10 μM 1-methyl-adenine to mature until the germinal vesicle disappears. 4. Eggs of all species are then passed through a 200 μm Nitex mesh to remove broken spines, tube feet, pieces of algae and gonad debris (in case of P. miniata). 5. For in vitro fertilization, sperm is diluted by mixing 5 μL of dry sperm with 13 mL of filtered sea water of appropriate salinity for the species (see above) in a 15 mL Falcon tube; excess water is removed from the beaker containing eggs leaving a small

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amount of filtered sea water covering the eggs. Using a Pasteur pipette 10 to 20 drops of diluted sperm is added to the eggs. Volume of diluted sperm required to fertilize the eggs is dependent on the number of the eggs and the volume of water they are in. Fertilization can be confirmed by the elevation of the vitelline membrane. 6. After fertilization, the embryos are cultured in 3 L of filtered sea water of appropriate salinity in glass 5 L beakers at 15  C for the Pacific species: S. purpuratus and P. miniata, or at 18  C for the Mediterranean species: P. lividus, until they reach the required developmental stage. 3.2 Collection of Embryos

1. When the embryos reach required developmental stage they are filtered using a 40 μm Nitex mesh placed in a 50 mL glass beaker. 2. The embryos are larger than 40 μm so they will not pass through the filter and can be washed with 1 L of 4  C fASW on the filter and then collected into a Petri dish using a Pasteur pipette. Washing with fASW is important to remove potential contaminants and unwanted ions found in sea water which cannot be removed by filtering. 3. Required number of embryos are then collected from the Petri dishes into 1.5 mL Eppendorf tubes placed on ice under a dissecting microscope using either a P200 micropipette or a mouth pipette.

3.3

Cell Lysis

1. Immediately place the tube in a microcentrifuge at 4  C and spin at 500  g for 5 min to pellet the embryos. Carefully remove the supernatant with a P200 micropipette. 2. Wash twice with 200 μL of ice cold fASW spinning down the cells at 500  g for 5 min at 4  C. 3. Resuspend the pellet in 50 μL of cold lysis buffer and pipette up and down with P200; this may take 3–5 min of pipetting. The lysate should be clear. 4. Transfer 50 μL, 20 μL, and 10 μL of the lysate into new tubes and centrifuge at 500  g 10 min at 4  C. Meanwhile, use the remaining 20 μL of lysate to count the nuclei using 2.5 μL of DAPI 1000 diluted 1: 100 with the help of a cell counting chamber under a fluorescent microscope (see Note 1). 5. Discard the supernatant, and immediately continue with Subheading 3.4.

3.4

Tagmentation

1. Make sure the cell pellet is set on ice.

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2. Prepare 50 μL of tagmentation reaction mix, combining 25 μL 2 TD buffer, 2.5 μL TD enzyme, and 22.5 μL nuclease-free H2O. 3. Use the appropriate combination of the different aliquots (50 μL, 20 μL, 10 μL), according to nuclei count, to get 50,000–90,000 nuclei. 4. Gently resuspend the pelleted nuclei in 50 μL of tagmentation mix and if more than one aliquot has to be used, use the same prepared 50 μL of mix. 5. Incubate nuclei in the tagmentation reaction for 30 min at 37  C. 6. Immediately after transposition, purify using the Qiagen MinElute Kit following the instructions (see Note 4). Elute transposed DNA in 10 μL Elution Buffer (see Note 5). Be sure to dry the column before adding the Elution Buffer. 7. Purified DNA can be stored at Subheading 3.5. 3.5 PCR Amplification

20  C or proceed with the

1. To generate the ATAC library, combine the following 50 μL mix in a PCR tube: 10 μL Tagmented DNA, 10 μL nucleasefree H2O, 2.5 μL Nextera PCR 25 μM Primer 1, 2.5 μL Nextera PCR 25 μM Primer 2, 25 μL NEBNext High-Fidelity 2 PCR Master Mix. 2. Use PCR cycle as follows: (1) 72  C, 5 min (2) 98  C, 30 s (3) 98  C, 10 s (4) 63  C, 30 s (5) 72  C, 1 min (6) Repeat steps 3–5, 15 (7) Hold at 4  C (see Note 6). 3. Purify amplified DNA using Qiagen MinElute Kit (see Note 4). Elute the purified library in 20 μL Elution Buffer (see Note 5). Be sure to dry the column before adding Elution Buffer. 4. Purified DNA can be stored at

3.6 Concentration Check and Electrophoresis Gel

20  C.

1. Use 1 μL of the amplified DNA library to determine the concentration using the Qubit dsDNA BR Assay Kit. Follow instructions provided by the kit. A minimum concentration of 20 ng/mL is preferred for NGS. 2. For quality control analysis of the samples, run 2–5 μL (depending on sample concentration) of the amplified library on a 2% agarose gel. A smear should be observed and depending on the quality of the gel, mono- (~200 bp) and di-nucleosome (~400 bp) fragments might be observed.

3.7

Data Analysis

1. ATAC-seq libraries from S. purpuratus wild type whole embryo samples were sequenced to produce an average of 57 million reads.

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2. Adapter sequences and bad quality bases can be trimmed from the resulting reads using Trimmomatic [17]. 3. The resulting pairs of fastq files can be aligned on genome with bowtie2 [18]. 4. In order to represent the center of the transposon event that correspond to accessible chromatin, the read start sites need to be adjusted aligned reads on the + strand have to be offset by +4 bp while reads on the - strand by 5 bp. 5. Reads corresponding to nucleosome-free pairs (insert”. Search for multiple sequences is not supported. 2. Select a BLAST program from the pull-down menu. Available programs are blastn, blastp, blastx, tblastn, and tblastx.

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3. Select a Database from the pull-down menu. Available databases are listed in Table 1. 4. Specify additional optional parameters of the BLAST program if needed: expect value (e-value), score, number of alignments, word size, number of descriptions, and query filtering option (default is OFF). 5. Click the search button below the search box. 6. BLAST result appears on the same page. Clicking the “DATA download” button placed in the upper part of the page allows users to download the BLAST result with text format. 3.3 Genome Viewer Page (Genome Explorer)

Genome viewer page (Fig. 1d) shows the visualized genome assembly (HpulGenome_v1), gene structures, and transcriptome positions with a genome viewing program, Genome Explorer (GE) [3]. GE has a number of functions, such as displaying the position of a gene, searching a gene with gene ID, gene name, synonym, and other keywords, and retrieving the sequence of a gene. The followings are the explanation how to use those tools. GE is composed of four components; (a) Genome box, (b) Chromosome/Scaffold/Contig bar, (c) Control menu bar, and (d) Mapping view (Fig. 2).

3.3.1 How to Display a Specific Genome Position

1. Select a scaffold from a list of scaffolds in the Genome box (Fig. 2, i). Alternatively, user can narrow down the list by entering the scaffold number in the “Filter” box (Fig. 2, Arrow). 2. Enter a numeric value in the “Position” box (Fig. 2, Arrowhead). 3. Clicking “Move” button displays the specified genomic position in the mapping view.

3.3.2 How to Search for a Gene

1. Click the search tab in the Genome box (Fig. 2, i’). 2. Enter a keyword in the “Keyword” box. Available keywords are HPU gene ID (e.g., HPU_19385) and gene name (e.g., Hp-Bmp11). 3. Clicking “Search” button returns a list of genes containing the keyword in the “Result” box. 4. Clicking one of the listed genes displays the specified gene position in the mapping view.

3.3.3 How to Retrieve a Sequence

Using “Copy to sequence” button 1. Display a gene on the mapping view with reference to 2.3.1 or 2.3.2.

2. Clicking anywhere in the gene region indicated by boxplot displays “Copy to sequence” button.

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Fig. 1 Screenshots of HpBase (http://cell-innovation.nig.ac.jp/Hpul/). HpBase consists of six pages; Home (a), Gene Search (b), Homology Search with BLAST (c), Genome Viewer with Genome Explorer (d), Data Download (e), and Protocols (f)

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Fig. 2 Graphical view of H. pulcherrimus genome assembly and gene positions displayed by genome viewing tool, Genome Explorer (GE). GE is composed of four components; (i) Genome box, (ii) Chromosome/Scaffold/ Contig bar, (iii) Control menu bar, and (iv) Mapping view. A specific gene position can be displayed by entering scaffold number in the “Filter” box (Arrow) and a numeric value in the “Position” box (Arrowhead)

3. Click “Copy to sequence” button to copy whole sequence of a gene including exons and introns or transcriptome to the clipboard. Using “From” and “To” boxes 1. Enter numeric values in the “From” and “To” boxes (Fig. 2, iii). The value in the “To” box should be larger than the one in the “From” box.

2. Click “get reference sequence” button to copy the genomic sequence of the specified region to the clipboard. 3.3.4 How to Change Color and Thickness of Boxplot

1. Click the “Option” icon on the right side of control menu bar (Fig. 2, iii) to open the option setting page. 2. Select “Plot style” tab in the option setting page. 3. Click target track(s) (Gene or Transcript). Multiple tracks are selectable. 4. Choose plot color from the color chart to change the color of boxplot. 5. Specify the plot thickness using horizontal slider. 6. Click “OK” button to reflect the changes.

3.3.5 How to Obtain the Image of Genome Map

1. Click the “Capture” icon on the right side of control menu bar (Fig. 2, iii), the save screen will appear.

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2. Save the image of the genomic region currently displayed in the mapping view in PNG format. 3.4 Data Download Page

1. This page (Fig. 1e) allows users to download the sequence data and annotation files listed in Table 1 by clicking the “download” button to the right of each dataset. 2. The md5, which is the file that tracks all changes to the data, for each dataset are also downloadable and can be used to verify the data integrity.

3.5

Protocol Page

1. This page (Fig. 1f) contains PDFs showing a brief protocol for the experiment in each laboratory. 2. The contents of this page will help researchers using H. pulcherrimus by providing experimental protocols and additional information based on the experiences of each scientist and/or each laboratory (see Note 3).

3.5.1 Protocol for Immunohistochemistry

1. This protocol shows how to stain sea urchin embryos/larvae using specific antibodies. 2. The conditions for fixation, blocking, and buffers are posted. 3. With reference to the condition shown here, readers can modify their own protocols.

3.5.2 Protocol for In Situ Hybridization

1. This protocol shows how to detect gene expression in whole mount condition using specific RNA probes. 2. The conditions and buffers refer to previous reports [4–7]. 3. With reference to the condition shown here, readers can modify their own protocols.

3.5.3 Reagents for Microinjection

1. This protocol shows how to prepare the reagents for microinjection into the eggs of H. pulcherrimus. 2. Based on the published data for H. pulcherrimus, the effective concentrations of morpholino oligonucleotides (MO) to knockdown genes are between 100 μmol/L (e.g., NodalMO1) [8] and 3.8 mmol/L (e.g., Wnt7-MO1) [9] so far. These experimental concentrations shown in the protocol produced healthy morphants. 3. To show the specificity of morpholino, using the second nonoverlapped morpholino should be the first choice. 4. The second choice is using different species of sea urchins to confirm the same phenotype. 5. The third choice is using control morpholinos, but universal control-MO, GFP-MO, and randomized-MO obtained from Gene Tools are not available at high concentration for H. pulcherrimus because 3.8 mmol/L aliquots of these

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morpholinos killed the embryos for unknown reasons while some morpholinos like Wnt-7 mentioned above produced healthy embryos with the same concentration (Fig. 1f). 3.5.4 Protocol for qPCR

1. This protocol shows how to detect the quantity of expressed genes using quantitative PCR (qPCR). SYBR green method is shown. 2. The protocol of cDNA synthesis from injected embryos and designing primers are included. 3. The detailed conditions for PCR depend on the target. 4. With reference to the condition shown here, readers can modify their own protocols.

3.5.5 Protocol for Luciferase Assay (TOPFlash Assay)

1. This protocol shows how to detect the activity of Wnt/TCF quantitatively in sea urchin embryos/larvae. 2. The protocol includes the amplification of required fragment from plasmids obtained from Addgene, reagents for microinjection, and detecting procedures. Gene expression in whole mount condition using specific RNA probes. 3. The conditions and buffers refer the previous reports [9, 10]. 4. With reference to the condition shown here, readers can modify their own protocols.

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Notes 1. Whole genome sequencing has become more common even in nonmodel organism in recent years. On the other hand, bioinformatics skills required to access individual genes from enormous data sets are not yet common. In order to allow such researchers to utilize the genomic data, it is necessary to provide tools to easily access individual genes. For this purpose, we have created the website HpBase. 2. In addition to providing download links to bulk data, HpBase equips multiple tools to browse, search, and retrieve individual gene data. 3. Additionally, experimental protocols that have been accumulated in the field of developmental and cell biology, so that many researchers can utilize genomic data for their subsequent experiments. Therefore, we expect that HpBase will assist efficient use of genome resources for researchers and promote researches in the various fields such as evolutionary, developmental, and cell biology.

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Acknowledgments This work was supported by a project of the Joint Usage/Educational Center (to MK) and Grant-in-Aid for Scientific Research (B) (26290070) (to TY and SY), (C) (25440101) (to SY), and Grant-in-Aid for Young Scientists (B) (23770241) (to SY) by the Ministry of Education, Culture, Sports, Science and Technology in Japan. This work also partially supported by the Ministry of Education, Culture, Sports, Science and Technology, Japan (MEXT) grant [code 09002013] and by Agency for Medical Research and Development (AMED) grant [JP16am0101058j0003] and [JP17am0101102]. We thank Masafumi Muraoka, Kazutoshi Yoshitake, Sadahiko Misu, and Norikazu Monma for their help in the analyses and HpBase construction. References 1. Kinjo S, Kiyomoto M, Yamamoto T et al (2018a) HpBase: a genome database of a sea urchin, Hemicentrotus pulcherrimus. Develop Growth Differ 60:174–182 2. Sodergren E, Weinstock GM, Davidson EH et al (2006) The genome of the sea urchin Strongylocentrotus purpuratus. Science 314:941–952 3. Kinjo S, Monma N, Misu S et al (2018b) Maser: one-stop platform for NGS big data from analysis to visualization. Database 2018: bay027 4. Arenas-mena C, Cameron AR, Davidson EH (2000) Spatial expression of Hox cluster genes in the ontogeny of a sea urchin. Development 127:4631–4643 5. Yaguchi S, Katow H (2003) Expression of tryptophan 5-hydroxylase gene during sea urchin neurogenesis and role of serotonergic nervous system in larval behavior. J Comp Neurol 229:219–229

6. Minokawa T, Rast JP, Arenas-mena C et al (2004) Expression patterns of four different regulatory genes that function during sea urchin development. Gene Expr Patterns 4:449–456 7. Yaguchi S, Yaguchi J, Burke RD (2006) Specification of ectoderm restricts the size of the animal plate and patterns neurogenesis in sea urchin embryos. Development 133:2337–2346 8. Yaguchi S, Yaguchi J, Angerer RC et al (2010) TGFβ signaling positions the ciliary band and patterns neurons in the sea urchin embryo. Dev Biol 347:71–81 9. Yaguchi J, Takeda N, Inaba K, Yaguchi S (2016) Cooperative Wnt-nodal signals regulate the patterning of anterior neuroectoderm. PLoS Genet 12(4):e1006001 10. Veeman MT, Slusarski DC, Kaykas A et al (2003) Zebrafish prickle, a modulator of noncanonical Wnt/Fz signaling, regulates gastrulation movements. Curr Biol 13:680–685

Chapter 18 Functional Studies of Trichoplax adhaerens Voltage-Gated Calcium Channel Activity Julia Gauberg, Adriano Senatore, and Andreas Heyland Abstract Trichoplax adhaerens is a member of the phylum Placozoa, an enigmatic group of benthic animals with remarkably simple morphology. While initial work on these organisms has primarily focused on their morphology and the development of genomic resources, Trichoplax has received increased attention as a model for studying the evolution of nervous and sensory systems. This work is motivated by the fact that Trichoplax features distinct behaviours and responses to environmental stimuli. Therefore, much progress has been made in recent years on the molecular, cellular, and behavioral understanding of this organism. Methods outlined here provide hands-on approaches to cutting edge molecular and cellular techniques to record cellular activities in Trichoplax. Key words Placozoa, Patch clamp, Calcium channels, Cloning

1

Introduction Trichoplax adhaerens is a species in the phylum Placozoa that has been the subject of considerable investigation in recent years. The newly emerged interest in this enigmatic creature arose from its basal position within the animal kingdom as well as its extraordinarily simple morphological organization (for reviews see 1, 2). Trichoplax shows a remarkable array of behavioral responses to its environment [3–7], reproduces both sexually [8–10] and asexually [8] and defends itself chemically against predators [11]. It also relies on a broad array of developmental and physiological signaling pathways for essential cellular and molecular functions [4, 12]. The cellular organization of Trichoplax remains an area of active investigation and some approaches that can facilitate the morphological and immunohistochemical studies of these cells are further outlined in Chapter 6 by Smith et al. in this same volume. However, from a physiological perspective it is critical to gain insight into responses to inter- and intracellular signaling systems. While the

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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cellular organization of Trichoplax is simple, it provides all essential functions for a free-living lifestyle, thus almost certainly excluding the possibility that Trichoplax is a parasitic form. Individuals are characterized by a dorsal and ventral ciliated epithelium. The cell arrangement between the two ciliated epithelia is relatively loose. Trichoplax does not have a coelom, and the only specialized cell type in this region is the fiber cell which may be used for a diversity of functions, including locomotion and cell communication. Studies suggest that fiber cells can produce cytoplasmic extensions in vitro [13] and may contact other cells in vivo [14]. Trichoplax also has regular cell–cell junctions between epithelial cells. However, the basal lamina appears to be missing and a true extracellular matrix (ECM) has not been identified to date, although the genome contains genes that encode ECM-interacting proteins [12, 15]. Trichoplax incorporates food through specialized digestive cells in the ventral epithelium which secrete digestive enzymes and can phagocytose digested food particles [5]. The dorsal epithelium has been shown to exhibit ultrafast contractions in response to calcium [16]. Gland cells located around the periphery of the animal contain neuropeptides that can alter the behavior of the animal [4]. Trichoplax expresses genes necessary for regulated exocytosis, as well as many voltage-gated calcium channels [17, 18]. In fact, Trichoplax is the most early-diverging animal to express all three types of voltage-gated calcium channels found in animals with true nervous systems [17]. These observations suggest that despite its cellular simplicity, Trichoplax expresses proteins for rapid communication and coordinated movement. Finally, shiny spheres are additional components of Trichoplax that have recently been shown to induce paralysis in would-be predators [11]. The methods outlined in this chapter provide a basis for molecular and functional characterization of Trichoplax voltage-gated calcium channels. Linking these aspects will contribute to our understanding of their physiology and evolution.

2

Materials

2.1 Extraction of RNA from Trichoplax adhaerens

1. TRI Reagent® (Millipore Sigma). 2. Chloroform (ACS Grade, Caledon Laboratories Ltd). 3. Ethanol (Caledon Laboratories Ltd). 4. DEPC-treated water (BioShop® Canada, Inc.). Prepare a 0.1% DEPC solution in double-distilled water. Autoclave before use. 5. Deoxyribonuclease (DNase) I, Amplification Grade—1 U/mL and 10 DNase I Reaction buffer (Invitrogen Canada Inc). 6. RiboLock RNase inhibitor (Thermo Fisher Scientific). 7. Glycogen, molecular biology grade, RNase free (Thermo Fisher Scientific).

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Table 1 Primer sequences used for cloning TCaV3 cDNA Primer

Sequence (50 to 30 )

NT cDNA

CTTTAGGTAGTATGAGCAAGGAATG

CT cDNA

TTTTTTTTTTTTTTTTTTVN

NT 50 1 (NT F1)

TGA TGT TTT ATT CAA GTC ATGC

0

CAT CAG GCG TCTTAACTTTGC

0

NT 5 2 (NT F2)

TACTTAGCTAGCCGCGGGAGCCACCATGGATATTTCGTTCGCTATCAT

NT 30 2 (NT R2)

TGCTCTGTGGTGGATCTCGAG

CT 50 1 (CT F1)

AAATCTGGTAGATCCTAATGAAGTC

NT 3 1 (NT R1)

0

CTTGATATTGATTAAATTTATCCAGATG

0

CT 5 2 (CT F2)

TCG CGT TCC AAT TGG TCCTCGAG

CT 30 2 (CT R2)

ACTCATCCCGGGTTATACTAATGTTTGAATCAT

CT 3 1 (CT R1)

NT N-terminal, CT C-terminal

2.2 Cloning of the Trichoplax T-Type Voltage-Gated Calcium Channel (TCaV3) cDNA into the pIRES2-EGFP Vector

1. Anchored Oligo (dT)18 primer (Table 1). 2. Superscript III Reverse Transcriptase—200 U/mL (Invitrogen Canada, Inc.). 3. DTT—0.1 M and 5 First-Strand Buffer (Invitrogen Canada Inc.). 4. dNTP—10 mM (Invitrogen Canada, Inc.). 5. Phusion® High-Fidelity DNA Polymerase (New England Biolabs). 6. pIRES2-IR-enhanced green fluorescent protein (EGFP) vector (Clontech laboratories), 7. T4 DNA ligase 5 U/μL and 10 T4 DNA ligase buffer (New England Biolabs). 8. Shrimp Alkaline Phosphatase (rSAP) 1 U/μL (New England Biolabs). 9. Kanamycin solution: 35 mg/mL Kanamycin sulfate (BioShop) in ddH2O. Filter sterilize twice and freeze at 20  C in 200 μL aliquots. 10. Super broth: 32 g Tryptone, Bacteriological (BioShop), 20 g Bacto Yeast Extract (BD), 5 g NaCl (BioShop) in 1 L of double-distilled water. Autoclave and store at 4  C. 11. Kanamycin-agar plates: Add 5 g of Agar, Bacteriological Grade (BioShop) to 333 mL super broth. Autoclave, cool to 55  C and add 300 μL of kanamycin solution. Pour 20 mL of kanamycin-agar solution into individual petri dishes and let

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them dry, covered, overnight. Kanamycin-agar plates can be stored at 4  C for up to 2 weeks. 12. NEB Stable Competent E. coli (New England Biolabs). 2.3 Transfection of TCaV3-pIRES-EGFP Vectors into HEK-293T Cells

1. Eppendorf Cell Culture Flasks T-25 (Eppendorf Canada Ltd). 2. Dulbecco’s Modified Eagle Media with 4500 mg/L glucose, sodium bicarbonate, L-glutamine, without sodium pyruvate (D-MEM; Millipore Sigma). Store at 4  C until use. 3. Penicillin-Streptomycin (Millipore Sigma). 10,000 units/mL penicillin and 10 mg/mL streptomycin. Divide into 1 mL aliquots and store at 20  C until use. 4. Fetal bovine serum (FBS; Millipore Sigma). Divide into 50 mL aliquots and store at 20  C until use. 5. Trypsin-EDTA solution (Millipore Sigma). 0.05% porcine trypsin and 0.53 mM EDTAl4Na in Hanks’ Balanced Salt Solution with phenol red. Divide into 10 mL aliquots and store at 20  C until use. Single aliquots can be stored at 4  C for several days and are temperature equilibrated to 37  C prior to use. 6. Supplemented D-MEM: 500 mL sterile D-MEM supplemented with 20 IU/mL penicillin, 20 μg/mL streptomycin, and 50 mL FBS. Complete media is kept at 4  C and equilibrated to 37  C prior to use. 7. Dulbecco’s phosphate buffered saline (PBS; Millipore Sigma). This solution is kept at 4  C and equilibrated to 37  C prior to use. 8. 2 M CaCl2 solution. Filter-sterilize and store at 4  C. 9. 2x HEPES buffered saline (HBS): 274 mM NaCl, 55 mM HEPES, 1.5 mM Na2HPO4. Filter-sterilize and store at 4  C until use. 10. Rat α2δ1 pMT2 vector (Addgene #58726). 11. Rat β1b pMT2 vector (Addgene #107423). 12. 13 mm plastic, tissue culture treated coverslips (Sarstedt Inc). 13. 60 mm cell culture dishes (Eppendorf Canada Ltd).

2.4 Whole-Cell Patch Clamp Recording of TCaV3-pIRES2-EGFP Transfected HEK-293T Cells

1. External recording solution:140 mM tetraethylammonium (TEA)-Cl2, 2 mM MgCl2, 3 mM CaCl2, 10 mM glucose, 10 mM HEPES, pH 7.4 with TEA-OH, adjust to 320 mOsm with glucose. Store at 4  C and warm up to room temperature before performing electrophysiological recodings. Internal recording solution:120 mM CsCl, 1 mM MgCl2, 10 mM HEPES, 10 mM EGTA, 4 mM ATP-Mg, 0.3 mM GTP-Li, pH 7.2 with CsOH, adjust to 300 mOsm with glucose. Store in 1 mL aliquots at –20  C and defrost on ice before use.

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2. 60 mm cell culture dishes (Eppendorf Canada Ltd) 3. Borosilicate glass capillaries (1.5 mm O.D., 0.86 mm I.D., 10 cm length, with filament; Sutter instruments). 4. P-1000 micropipette puller (Sutter instruments). A program is optimized to generate pipettes with resistances of 2–5 MΩ between the patch pipette electrode and the reference electrode (after fire polishing). 5. Fire polisher (Micro Forge MF-830; Narishige, Japan). Fire polishing smooths the pipette tip surfaces and allows for better seals between the patch pipette and cell membranes. 6. Patch pipette holder (1-HL-U; Molecular Devices, Sunnyvale, California). 7. Digidata 1440A analog-to-digital converter. 8. Axopatch 200B (Axon Instruments, Union City, CA). 9. pClamp10.1 California),

software

(Molecular

Devices,

Sunnyvale,

10. Dual pipette manipulator system (MPC-385-2, Sutter Instrument Company). 11. Epifluorescence microscope (Axiovert 40 CFL, Zeiss Canada). 12. Vibraplane air table (Kinetic systems, Boston, MA). 13. Microelectrode Holder (Holder 90 F Pellet 1.5 mm; #MEH3RF15; World Precision Instruments Inc.) filled with 3 M CsCl solution. 14. Performance Series Vertical Power Bar, Surge Protector (NOMA). 15. Valvelink 8.2 8 channel perfusion system (Automate Scientific). 16. Perfusion pencil 8 channel manifold + removable tip (Automate Scientific).

3

Methods

3.1 Extraction of RNA from Trichoplax adhaerens

1. 30 Trichoplax are transferred into a new glass dish filled with artificial seawater 2 h before extracting RNA. For extraction, the animals are transferred to a 1.5 mL tube and the seawater is aspirated away. 500 μL of TRI Reagent® is added to the tube and the cells are dissociated by pipetting up and down. Total RNA extraction is performed according to the manufacturer’s instructions (see Note 1). RNA concentration is assessed by spectrophotometry and integrity by the A260/A280 ratio. 2. An aliquot of total RNA (1 μg) is treated with DNase I and RiboLock for 15 min at room temperature followed by a 5 min

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incubation at 65  C. RNA is precipitated by ethanol precipitation with glycogen according to manufacturer’s instructions (see Note 2). Ethanol is aspirated and RNA is resuspended in DEPC-treated water. RNA can be flash-frozen and stored at -80  C until use. 3.2 Cloning of the Trichoplax T-Type Voltage-Gated Calcium Channel (TCaV3) cDNA into the pIRES2-EGFP Vector

1. Large channels (~6000 bp) require independent cloning of two ~3000 bp fragments (N-terminal and C-terminal) around a restriction site. We have found that it is easier to amplify DNA under 2.5 kbp and to clone smaller fragments. 2. First-strand cDNA is synthesized by using 1 μg of DNase I-treated total RNA with the reverse transcriptase. Two cDNA libraries are made: the C-terminal half of TCaV3 is cloned with an anchored oligo-(dT)18 primer, and the N-terminus half is cloned with a primer targeting a central region of the TCaV3 coding sequence (Table 1). 3. The TCaV3 N- and C-terminal coding sequences are amplified three times from the cDNA, via nested PCR using Phusion® DNA polymerase. This allows for amplicons to be compared in order to differentiate between naturally occurring polymorphisms and PCR errors. Amplification of the N- and C-terminal fragments is performed with two sets of nested primers (Table 1, Fig. 1). The first set of nested primers (NT/CT F1 and R1) is complementary to the TCaV3 N- and C-terminal untranslated regions and sequences around the internal XhoI site. In the second set of nested primers, two primers (NT R2 and CT F2) are complementary to the existing XhoI site present in the middle of the TCaV3 cDNA (see Note 3). The other two primers are complementary to the start codon (NT F2) and stop codon (CT R2) of TCaV3 and have NheI and XmaI restriction enzyme sites incorporated into them, respectively. These restriction enzymes were chosen because they are present in the multiple cloning site of the pIRES2-EGFP vector and absent from the TCaV3 sequence. The second nested N-terminal primer (NT F2) also has a mammalian Kozak translation initiation site (i.e., 50 -GCC ACC-30 ) sequence incorporated immediately upstream of the start codon to facilitate expression of the TCaV3 channel in mammalian cells. 4. TCaV3 N- and C-terminal DNA fragments are cloned into the pIRES2-EGFP plasmid, which allows for the identification of positively transfected cells via EGFP fluorescence. The EGFP is produced in transfected cells from the same transcript as the cloned insert cDNA, due to an internal ribosome entry site just downstream of the insert cDNA and upstream of the EGFP coding sequence. The PCR amplified DNA fragments and pIRES2-EGFP vector are incubated with NheI-XhoI or XhoIXmaI restriction enzymes, and the 50 ends of vector DNA are

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Fig. 1 Cloning the TCaV3 channel cDNA and ligation into the pIRES2-EGFP vector. (a) Schematic of cDNA synthesis and nested PCR primers for cloning of TCaV3 cDNA. The coding sequence is split into N-terminal (NT) and C-terminal (CT) fragments that are amplified individually in triplicate. (b) The two fragments are ligated into the pIRES2-EGFP vector, where they can be amplified individually and sequenced. Multiple cloning site: MSC, Cytomegalovirus: CMV; Internal ribosome entry site: IRES, Enhanced green fluorescence protein: EGFP, Kanamycin resistance gene: KanR. (c) The CT fragment is ligated into the plasmid containing the NT fragment to produce the full-length coding sequence (CDS)

dephosphorylated by adding rSAP according to manufacturer instructions. Digested TCaV3 and pIRES-EGFP products are resolved on a 1% agarose gel (100 V for 1.5 h) and ethidium bromide is used for visualization. TCaV3 and pIRES2-EGFP DNA are extracted and TCaV3 DNA is ligated into the pIRES2-EGFP vector using T4 ligase in a 3:1 ratio according

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to manufacturer instructions (see Note 4). N- and C-terminal DNA fragments are sequenced and compared with each other and the Trichoplax genome (JGI Genome Portal, Grell-BS1999 v1.0, scaffold _2: 6781672–6793175) to generate a consensus coding sequence. 5. The full-length TCav3 coding sequence is then prepared by ligating the XhoI-XmaI C-terminal subclone into the pIRES2EGFP vector containing the N-terminal TCav3 fragment using T4 ligase, producing pTCav3-IRES2-EGFP. 5 μL of ligated product is added to 50 μL of NEB stable competent E. coli, which are plated onto kanamycin-agar plates. Large-scale plasmid prep is then performed to isolate the pTCaV3-IRES2EGFP vector (see Notes 5 and 6). 3.3 Transfection of the pTCaV3-IRES2-EGFP Vector into HEK-293 T Cells

1. Human embryonic kidney 293 T cells (HEK-293T) are cultured in vented 25 cm2 flasks in supplemented media at 37  C with 5% CO2. Cells are split biweekly from a ~90% confluency in a 1:8 dilution (2.6  104 cells/cm2) on Mondays and a 1:12 dilution (1.71  104 cells/cm2) on Thursdays. All work with cells is done in a laminar flow hood. 2. 90% confluent cells are havested before transfection by trypsinization and replated at a 1:4 dilution (5.2  104 cells/cm2) in a 25 cm2 flask with 6 mL of supplemented D-MEM. The culture is incubated for 4–8 h at 37  C with 5% CO2 before transfection. 3. 36.6 μL of 2 M CaCl2 is combined with 4 μg of pTCaV3IRES2-EGFP plasmid DNA and 2 μg of the rat α2δ and β plasmid DNA (see Note 7) in a sterile 1 mL tube and sterile water is used to bring the volume to 300 μL. Add one volume of this solution, drop by drop, to an equal volume of 2 HBS. Mix thoroughly and push bubbles through the mixture 6 times using a micropipette. 4. The calcium phosphate–DNA suspension is added to the media above the cultured cells (see Note 8). The flask is rocked back and forth to mix the media. 5. The cells are incubated in a 37  C incubator with 5% CO2 overnight then the media and DNA are removed by aspiration. The cells are washed gently three times with 4 mL of PBS (see Note 9). The wash solution is aspirated away and 6 mL of fresh supplemented media is added to the flask. The flask is returned to the incubator for 1–3 days to allow for expression of ectopic cDNA (see Note 10). Successful transfection can be confirmed using an epifluorescence microscope. Cells that fluoresce green when exposed to UV light are positively transfected and should express the heterologous ion channel and EGFP proteins.

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6. To transfer cells onto coverslips for whole-cell patch clamp recording, the media is aspiated away and 500 μL of trypsin (see Note 11) (warmed to 37  C) is added. The flask is rocked back and forth and the trypsin and any cell debris are aspirated away. Another 200 μL of trypsin is added and incubated for 2–4 min at 37  C. The cells are then resuspended in 4 mL of supplemented media. 7. Three cover slips are added into two sterile 60 mm cell culture dishes and 4.5 mL of supplemented media is transferred into each dish. 500 μL of the cell suspension is added to the cell culture dishes (see Note 12). Dishes are placed into the 37  C incubator for 4 h and then transferred to 28  C for 1–3 days prior to electrophysiological recordings (see Note 13). 3.4 Whole-Cell Patch Voltage Clamp Recording of pTCaV3-IRES2-EGFP Transfected HEK-293T Cells

1. One coverslip containing transfected cells is transferred to a 35 mm dish filled with 2 mL of external recording solution (see Note 14). Whole-cell patch voltage clamp recordings are done at room temperature. Whole-cell patch clamp of TCaV3-transfected HEK-293T cells is conducted as described in [19]. Figure 2 illustrates calcium currents recorded from an HEK-293T cell expressing the TCaV3 channel. The cell was held at a voltage of 110 mV, and incremental depolarisations between 90 mV and +40 mV were taken to elicit inward calcium currents. Different voltage step protocols can be performed to understand the biophysical properties of TCaV3 channels. 2. In addition to varying the voltage step protocols, blockers and various external solutions can be perfused over the surface of the cells. The perfusion pencil to cell distance must be consistent between patches, and cells must be equilibrated to the perfusion solution before recordings are performed (see Note 15). To prevent overflow of solution into the dish, a 60 mL syringe with a blunt needle bent to 90 can be used to manually aspirate excess solution (see Note 16). After each perfusion, fresh external solution should be pipetted into the dish, so that other cells can be recorded (see Note 17). Figure 3 shows calcium and barium currents recorded from one HEK-293T cell.

4

Notes 1. Trichoplax RNA is unstable and degrades quickly, so always keep extracted RNA on ice and use RNase inhibitors when removing DNA with DNase I. 2. We noticed that precipitating RNA with glycogen and ethanol results in better 260/230 ratios. An RNA cleanup kit can be used as an alternative to this step.

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Fig. 2 Representative trace of calcium currents through TCaV3 channels ectopically expressed in HEK-293T cells. Inset shows voltage protocol used to elicit calcium into the cell. Currents were obtained by incremental depolarization from a holding potential of 110 mV to between 90 and + 40 mV. Scale bar indicates time in milliseconds (ms) and current in picoamperes (pA)

Fig. 3 Representative traces of calcium and barium currents through TCaV3 channels ectopically expressed in HEK-293T cells. External solutions containing 2 mM calcium or barium were perfused over the cell. Currents were obtained by incremental depolarization from a holding potential of 110 mV to between 90 and +40 mV. Calcium currents are indicated in black and barium currents are indicated in red. Scale bar indicates time in milliseconds (ms) and current in picoamperes (pA)

3. The restriction site must also be present in the multiple cloning site of the pIRES2-EGFP vector. If no restriction enzyme site can be found, a silent mutation can be introduced in the CT F2 and NT R2 primers to create a restriction enzyme site.

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4. Run a vector-only negative control without the insert to see if the vector is able to ligate on itself. You should get little to no bacterial colonies on negative control plates. 5. When quantifying pTCaV3-IRES2-EGFP concentration, add sterile water to dilute DNA concentration to 1 μg/μL. This is the most convenient concentration for transfections. 6. Trichoplax DNA can be codon-optimized for better expression in mammalian cells. This requires synthesis of the cDNA sequence by a company such as GenScript. 7. Typically, 4 μg of pTCaV3-IRES2-EGFP DNA and 2 μg of each subunit produce enough positively transfected cells for 2–3 days of recording. However, more or less DNA can be added to optimize transfection efficiency. 8. Add transfection solution slowly and evenly into media (drop by drop over the surface), trying to cover as much of the plate as possible. 9. PBS should be pipetted onto the back wall of the flask, not directly onto the cells. 10. Usually 1 day at 37  C is enough to see green fluorescence, however, keeping the cells at 37  C for an additional day or two can increase the expression of TCaV3 DNA. 11. The first trypsin “wash” should not be pipetted directly onto the cells because they will detach and be removed during aspiration. 12. If the coverslips float when media is added, push them down with the pipette tip before expelling the cell suspension. 13. Dishes with transfected cells can typically be kept in the 28  C incubator for up to 3 days for whole-cell patch clamp recording. If kept in the 28  C incubator for longer periods of time, the health of the cells declines and obtaining good patches becomes more difficult. 14. Touch one edge of the coverslip to a Kimwipe to remove excess media before transferring to the dish with external recording solution. 15. For perfusion of various blocker concentrations, we have found that 4–5 sweeps (voltage steps) should be performed at one blocker concentration before moving on to a higher blocker concentration. This allows for the cell to be completely “bathed” in the blocker solution and results in the most reliable current recordings. 16. When aspirating, be careful not to aspirate too quickly or forcefully because that may alter the calcium currents or disrupt the patch on the cell.

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17. A new coverslip with transfected HEK cells may need to be used for each recording, especially when perfusing pharmacological agents that irreversibly bind to CaV channels. References 1. Syed T, Schierwater B (2002) Trichoplax adhaerens: discovered as a missing link, forgotten as a hydrozoan, re-discovered as a key to metazoan evolution. Vie Milieu 52:177–187 2. Schierwater B (2005) My favorite animal, Trichoplax adhaerens. BioEssays 27:1294–1302 3. Fortunato A, Aktipis A (2019) Social feeding behavior of Trichoplax adhaerens. Front Ecol Evol 7:19 4. Senatore A, Reese TS, Smith CL (2017) Neuropeptidergic integration of behavior in Trichoplax adhaerens, an animal without synapses. J Exp Biol 220:3381–3390 5. Smith CL, Pivovarova N, Reese TS (2015) Coordinated feeding behavior in Trichoplax, an animal without synapses. PLoS One 10(9): e0136098 6. Smith CL, Reese TS, Govezensky T et al (2019) Coherent directed movement toward food modeled in Trichoplax, a ciliated animal lacking a nervous system. Proc Natl Acad Sci U S A 116:8901–8908 7. Varoqueaux F, Williams EA, Grandemange S et al (2018) High cell diversity and complex peptidergic signaling underlie placozoan behavior. Curr Biol 28(21):3495–3501 8. Grell KG (1984) Reproduction of placozoa. In: Engels W (ed) Advances in invertebrate reproduction. Elsevier, Amsterdam, pp 541–546 9. Signorovitch AY, Dellaporta SL, Buss LW (2005) Molecular signatures for sex in the Placozoa. Proc Natl Acad Sci U S A 102:15518–15522 10. Eitel M, Guidi L, Hadrys H et al (2011) New insights into placozoan sexual reproduction and development. PLoS One 6(5):e19639

11. Jackson AM, Buss LW (2009) Shiny spheres of placozoans (Trichoplax) function in antipredator defense. Invertebr Biol 128:205–212 12. Srivastava M, Begovic E, Chapman J et al (2008) The Trichoplax genome and the nature of placozoans. Nature 454:955–959 13. Ruthmann A, Terwelp U (1979) Disaggregation and reaggregation of cells of the primitive metazoan Trichoplax adhaerens. Differentiation 13:185–198 14. Smith CL, Varoqueaux F, Kittelmann M et al (2014) Novel cell types, neurosecretory cells, and body plan of the early-diverging metazoan Trichoplax adhaerens. Curr Biol 24:1565–1572 15. Grell KG, Ruthmann A (1991) Placozoa. In: Harrison FW, Westfall JA (eds) Microscopic anatomy of invertebrates. Wiley-Liss, New York, pp 13–28 16. Armon S, Storm Bull M, Aranda-Diaz A et al (2018) Ultrafast epithelial contractions provide insights into contraction speed limits and tissue integrity. PNAS 115(44):E10333–E10341 17. Senatore A, Raiss H, Le P (2016) Physiology and evolution of voltage gated calcium channels in early diverging animal phyla: Cnidaria, Placozoa, Porifera and Ctenophora. Front Phys 7:481 18. Wong YY, Le P, Elkhatib W et al (2019) Transcriptome profiling of Trichoplax adhaerens highlights its digestive epithelium and a rich set of genes for fast electrogenic and slow neuromodulatory cellular signaling. Preprint at https://doi.org/10.21203/rs.2.14504/v1 19. Senatore A, Boone A, Lam S et al (2011) Mapping of dihydropyridine binding residues in a less sensitive invertebrate L-type calcium channel (LCav1). Channels 5:173–187

Chapter 19 A Bioinformatics Tutorial for Comparative Development Genomics in Diverse Meiofauna Joseph L. Sevigny, Jon L. Norenburg, and Francesca Leasi Abstract Miniaturization, which is a common feature in animals, is particularly manifest in meiofauna—animals sharing peculiar phenotypic features that evolved as adaptations to the highly specialized aquatic interstitial habitat. While revealing much about the extreme phyletic diversity of meiofauna, the genome structure of meiofaunal species could also characterize the phenotype of ancestral states as well as explain the origin and evolution of miniaturization. Here, we present a practical bioinformatics tutorial for genome assembly, genome comparison, and characterization of Hox clusters in meiofaunal species. Key words Bioinformatics, Convergent evolution, Gene architecture, Gene function, Genomics, Hox genes, Interstitial habitat, Meiofauna, Miniaturization

1

Introduction Marine interstitial meiofauna are small (35 to phred scores ~25). This is common for Illumina reads in general and is especially true for the reverse reads. You may also notice that the fastqc report “failed” for adapter content. 5. These output files are cleaned reads, which have removed the adapters from the sequences and should retain the sequences with high quality. Some sequences will be lost entirely, some will lose a few bases off the ends, and some will not be trimmed at all. When a reverse read is lost but a forward read is maintained, the forward read will be written to the unpaired_forward.fastq.gz file (and vice versa). You can run FASTQC again with your new trimmed reads, comparing the original html and the new one, and you should note the differences. You can also count the number of reads for each of your files like you did for the raw reads. How does this compare to the original count? What percentage of your reads did you lose? How many reads are unpaired? 6. There are many programs that are used for genome assembly and different assemblers work well with certain genomes (how large the genome is, how complex it is, whether or not it is a eukaryote, etc.) (alternative tools: [ABySS](http://www.bcgsc. ca/platform/bioinfo/software/abyss), [MaSuRCA](http:// masurca.blogspot.com/). We use SPAdes (http://cab.spbu. ru/software/spades/), this program works well for most genomes and is straightforward to use. No matter what assembler you decide to use, the programs are usually run with the same sort of syntax. We encourage to try out different assemblers and compare the results.

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7. In the case of SPAdes, the FASTA headers are named in a common format. Something like “NODE_1_length_263127_cov_73.826513”. The first field is a unique name for the contig (just a numerical value), the next field is the length of the sequence, and the last field is the kmer coverage of that contig. Furthermore, the contigs are organized by length where the longest contigs are first. 8. The FASTA format is similar to the FASTQ format except it does not include quality information. Each sequence is also delimited by a “>” symbol instead of a “@”. In addition, all the sequences will be much larger (since they were assembled). Instead of all the sequencing being 250 bp they could be the size of an entire chromosome. Each of the sequence entries in the FASTA file are typically referred to as a “contig,” which means contiguous sequence. Ideally, the assembler would work perfectly, and we would have one contig per chromosome in the genome. However, this is very rarely the case (unless we add some sort of long-read technology like PacBio or Nanopore sequencing). How fragmented your reconstructed genome is, that usually depends on how many reads you put into your assembler, how large the genome is, and what is the architecture and complexity of the genome like. We typically see a genome split into hundreds to thousands of contigs for a typical run. 9. This type of information is something you would typically provide in a publication or to assess different assemblers/ options you may use. 10. There are different datasets for various taxonomic groups (Eukaryotes, Metazoa, Bacteria, Gammaproteobacteria, etc.). The idea is that a newly sequenced genome should contain most of these highly conserved genes. If your target genome does not contain a large portion of these single-copy orthologs it may indicate that the target genome is not complete. 11. We are hopeful that the majority of our genes will be found as “complete single-copy.” Duplicated genes could indicate that the particular gene underwent a gene duplication event or that we have had a misassembly, and essentially have two copies of a region of our target genome. Fragmented genes are an artifact of the fact that our genome did not assemble perfectly. Some of our genome is fragmented into multiple contigs, and with that some of our genes are going to be fragmented as well. Therefore, it is important to inspect the N50 of the genome with QUAST. We want the majority of our contigs to be at least as big as a gene, otherwise, we will have many fragmented genes as a result.

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12. Keep in mind that BLAST has options that span books (literally). There are different “flavors” of BLAST. For example, “blastn” is used for searching a nucleotide query against a nucleotide reference, “blastp” is used to searching a protein query against a protein reference, and so on. These different types of BLASTs have different sorts of advantages. For example, at the protein level sequences tend to be more conserved, this means we can likely identify more distantly related sequences compared to a standard blastn. The BLAST flavor you choose is largely dependent on what sort of data you have. Using the command line BLAST we can specify options like output formats as well as use a local database. Moreover, it is very useful for pipelines and workflows since it can be automated, as such, you do not need to open a webpage and fill out any forms. You may download sequences of genes related to an experiment and check if they correspond to your genome assembly. 13. Many programs perform read mapping. The recommended program depends on what you are trying to do. We recommend “BWA mem” (http://bio-bwa.sourceforge.net/bwa. shtml) which balances performance and accuracy well. The input to the program is a reference assembly and reads to map (forward and reverse). The output is a SAM file. By default, BWA writes the SAM file to standard output. There are lots of options, please see the manual to understand what is being using in this tutorial. 14. There are hundreds of programs that use SAM files as a primary input. A BAM file is the binary version of a SAM and can be converted very easily using SAMtools. 15. To get any sort of meaningful phylogenetic tree we need to be sure to include at least four different genome datasets. The number and taxonomic breadth of genomes you compare will be based on the experimental question. For example, if you want to investigate the presence and absence of developmental genes across the phylum Annelida you will want to download the protein FAA files for all Annelida species. We will use a reduced set of genomes for this tutorial. Next, we will determine what genomes we want to download and go over the best ways to retrieve them from NCBI. 16. Examine the output files. We will review some of them. See the manual for extensive details. (a) Orthogroups.csv—Each orthogroup is a raw, and each column is a different sample. The table provides all of the data for orthogroups that are in at least two different samples. If a sample has more than one protein for that

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particular orthogroup than it will have a comma separated list for the entry. (b) Orthogroups_UnassignedGenes.csv—The same style table. Instead this one contains Orthogroups that are not belonging to an orthogroup, they are unique to a single sample. As you scroll down you should notice the proteins belong to different samples. (c) Orthogroups.GeneCount.csv—Orthogroups are the rows, columns are gene counts per species. This can be easily parsed to see what orthogroups are specific to what species. It provides total gene counts for each sample. (d) Statistics_Overall.csv—A file containing the overall statistics for the analysis (total number of genes in the dataset etc.). (e) Statistics_PerSpecies.csv—Probably the most important statistics output file. It provides details for each sample such as how many genes were specific to a sample. (f) SpeciesTree_rooted.txt—A rooted-species tree. Orthofinder computes a root for the tree automatically. This file is in Newick format and you can view this in any tree viewing program like FigTree or TreeView (macs). 17. In addition to this orthofinder analyses, I also recommend utilizing blastp to further examine potential Hox genes within your species. To do this you will utilize NCBI to search for protein sequences and BLAST them similar to how we obtained taxonomic classifications. This will allow you to identify genes not present in the annotation files. References 1. Artois T, Fontaneto D, Hummon W et al (2011) Ubiquity of microscopic animals? Evidence from the morphological approach in species identification. In: Fontaneto D (ed) Biogeography of microscopic organisms, is everything small everywhere? Systematics Association & Cambridge University Press, UK, Cambridge 2. Curini-Galletti M, Artois T, Delogu V et al (2012) Patterns of diversity in soft-bodied meiofauna: dispersal ability and body size matter. PLoS One 7:e33801 3. Dunn CW, Hejnol A, Matus DQ et al (2008) Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452:745–749 4. Fontaneto D, Flot J-F, Tang CQ (2015) Guidelines for DNA taxonomy, with a focus on the meiofauna. Mar Biodivers 45:433–451

5. Ga˛siorowski L, Hejnol A (2019) Hox gene expression in postmetamorphic juveniles of the brachiopod Terebratalia transversa. EvoDevo 10:1 6. Giere O (2009) The microscopic motile Fauna of aquatic sediments. Springer, Berlin 7. Gross V, Treffkorna S, Reichelt J et al (2019) Miniaturization of tardigrades (water bears): morphological and genomic perspectives. Arthropod Struct Dev 48:12–19 8. Hanken J, Wake DB (1993) Miniaturization of body size: organismal consequences and evolutionary significance. Annu Rev Ecol Syst 24:501–519 9. Higgins RP, Thiel H (1988) Introduction to the study of Meiofauna. Smithsonian Institution Press, Washington D.C. 10. Jo¨rger KM, Norenburg JL, Wilson NG et al (2012) Barcoding against a paradox?

Hox Clusters Characterization in Meiofauna Combined molecular species delineations reveal multiple cryptic lineages in elusive meiofaunal sea slugs. BMC Evol Biol 12:245 11. Koonin EV (2009) Evolution of genome architecture. Int J Biochem Cell Biol 41:298–306 12. Laumer CE, Bekkouche N, Kerbl A et al (2015) Spiralian phylogeny informs the evolution of microscopic lineages. Curr Biol 25:2000–2006 13. Leasi F, Andrade SCDS, Norenburg JL (2016) At least some meiofaunal species are not everywhere. Indication of geographic, ecological and geological barriers affecting the dispersion of species of Ototyphlonemertes (Nemertea, Hoplonemertea). Mol Ecol 25:1381–1397 14. Leasi F, Norenburg JL (2014) The necessity of DNA taxonomy to reveal cryptic diversity and spatial distribution of meiofauna, with a focus on Nemertea. PLoS One 9:e104385 15. Leasi F, Tang CQ, De Smet WH et al (2013) Cryptic diversity with wide salinity tolerance in the putative euryhaline Testudinella clypeata (Rotifera, Monogononta). Zool J Linnean Soc 168:17–28 16. Leasi F, Sevigny JL, Laflamme EM (2018) Biodiversity estimates and ecological interpretations of meiofaunal communities are biased by the taxonomic approach. Comm Biol 1:112 17. Mills SJ, Alca´ntara-Rodrı´guez A, Ciros-Pe´rez J et al (2016) Fifteen species in one: deciphering the Brachionus plicatilis species complex (Rotifera, Monogononta) through DNA taxonomy. Hydrobiologia 796:39–58 18. Neves RC, Bailly X, Leasi F et al (2013) A complete three-dimensional reconstruction of the myoanatomy of Loricifera: comparative

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morphology of an adult and a Higgins larva stage. Front Zool 10:19 19. Niven JE, Farris SM (2012) Miniaturization of nervous systems and neurons. Curr Biol 22: R323–R329 20. Pfenninger M, Nowak C, Kley C et al (2007) Utility of DNA taxonomy and barcoding for the inference of larval community structure in morphologically cryptic chironomus (Diptera) species. Mol Ecol 16:1957–1968 21. Rundell RJ, Leander BS (2010) Masters of miniaturization: convergent evolution among interstitial eukaryotes. BioEssays 32:430–437 22. Smith FW, Boothby TC, Giovannini I et al (2016) The compact body plan of tardigrades evolved by the loss of a large body region. Curr Biol 26:224–229 23. Tang CQ, Leasi F, Urbertegger U et al (2012) The widely used small subunit 18S rDNA molecule greatly underestimates true diversity in biodiversity surveys of the meiofauna. PNAS 109:16208–16212 24. Schuelke T, Pereira TJ, Hardy SM et al (2018) Nematode-associated microbial taxa do not correlate with host phylogeny, geographic region or feeding morphology in marine sediment habitats. Mol Ecol 27:1930–1951 25. Treffkorn S, Kahnke L, Hering L et al (2018) Expression of NK cluster genes in the onychophoran Euperipatoides rowelli: implications for the evolution of NK family genes in nephrozoans. EvoDevo 9:17 26. Yoshida Y, Koutsovoulos G, Laetsch DR et al (2017) Comparative genomics of the tardigrades Hypsibius dujardini and Ramazzottius varieornatus. PLoS Biol 15:1–40

INDEX A Affinity interactions................... 120–122, 124, 126–128, 130–132, 134, 153–155, 158, 160 Aging .................................................................... 138, 142 Amphimedon queenslandica, see Sponge Amphioxus................................................................... 1–27 Amphiroa fragilissima, see Coralline alga Annelida..........................................................31, 290, 303 Antibodies ........................ 22, 72, 78, 83–86, 88, 89, 92, 106, 112, 113, 117, 119, 120, 122, 128, 130, 133, 138, 145–149, 163–165, 168–171, 175–177, 187, 189, 191, 197, 198, 201–203, 205, 206, 208–210, 212, 215, 273 Antisense...................................2, 25, 200, 201, 209, 247 Apoptosis ...................................... 13, 75–79, 82, 95, 137 Aquaculture ....................................................................... 4 Artificial seawater (ASW) ................................7, 8, 33, 52, 58–64, 66, 70, 72–75, 104, 106, 109, 120, 122, 123, 131, 141, 281 Ascidians ............................................................... 217–228 Assay for transposase-accessible chromatin with high-throughput sequencing (ATAC-seq)............................................... 253–263 Asterina pectinifera, see Starfish

B Bioconductor................................................................. 236 Bioinformatics ............................ 201, 232, 274, 289–304 Biotinylation ........................................152, 153, 155–157 Blotting...........................vi, vii, 121, 126, 128, 129, 134, 153–155, 158 Brachiolaria larvae ....................................... 50, 61, 63, 66 Branchiostoma floridae, see Amphioxus Breeding period............................................................... 54 5’-Bromo-2’-deoxyuridine (BrdU).................... 165, 168, 170–172, 177, 178

C Ca2+ indicators ............................................ 32, 40, 41, 45 Calcium.........................33, 58, 120, 141, 151, 152, 242, 278, 283, 285–287 Calcium channels ........................................ 278, 282, 284 Ca release ................................................ vi, 120, 151, 152

Cell cycle...........................119, 151, 163, 164, 166, 217, 218, 221, 222, 226 Cell surface ...................................................151–160, 218 Cephalochordates.............................................................. 1 Cerebratulus, see Nemertea Cis-regulatory elements (CREs) .................253–255, 262 Click chemistry..................................................... 168, 173 Cloning .........................16, 22, 124, 201, 254, 279, 280, 282–284, 286 Cnidarian ................................................................ 69, 231 Confocal microscopy .............................. 88, 89, 153, 197 Coralline algae .................................................... 67, 83, 87 CRISPR-Cas9........................................ 82, 221, 223–225

D Databases ............................ 66, 120, 224, 237, 267–274, 290, 292, 295, 296, 303 De-novo assembly ......................................................... 296 Developments..................... 2, 11, 12, 22, 27, 36, 62, 65, 82, 84, 134, 160, 165, 168, 178, 182, 188–191, 196, 197, 210, 253, 254, 262, 275, 289–304 40 ,6-Diamidino-2-phenylindole (DAPI) .........70, 71, 75, 77–79, 84–86, 88–90, 92, 93, 168, 170, 171, 173, 179, 205, 208, 212, 243, 246, 251, 257, 259 Diatoms .......................................... 53, 58, 60–62, 65, 66 Dicer-substrate interference RNAs (DsiRNAs) ...............................241, 244, 247, 250 Differentiation............................................................... 261 DNA ...................2–6, 14–17, 19–25, 75, 125, 163–166, 175, 177, 185, 196, 201–203, 210, 224, 225, 244, 246, 247, 251, 253, 254, 256, 257, 260, 263, 279, 282–285, 287, 290, 292, 296

E Echinoderms ............................. 137, 138, 151, 195–215, 242, 253–263 Egg activation................................................................ 120 Eggs .............. 2, 4, 8, 11, 12, 15, 20, 22, 24, 32, 35–37, 41–46, 50, 58, 119, 120, 122–124, 127, 130–134, 142, 151–160, 196, 200, 219, 220, 222–225, 258, 259, 273 Electron microscopy (EM) ........................ 101, 103–106, 108–112, 117, 175, 199 Electroporation .......................... 243, 245–247, 249, 250

David J. Carroll and Stephen A. Stricker (eds.), Developmental Biology of the Sea Urchin and Other Marine Invertebrates: Methods and Protocols, Methods in Molecular Biology, vol. 2219, https://doi.org/10.1007/978-1-0716-0974-3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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AND

Embryonic development ........................... 22, 50, 52, 53, 209, 224, 226, 231, 253 Embryos ...................2, 8, 12, 20–25, 31–47, 53, 58–60, 64, 65, 83, 141, 142, 166, 175, 196–200, 205–210, 212, 214, 215, 218, 220–226, 233, 242, 253–263, 269, 273, 274 Enhanced chemiluminescence (ECL)................ 122, 130, 132, 140, 146, 149, 155, 159 5’-Ethynyl-2’-deoxyuridine (EdU) ..........................86, 95 Evo-devo.........................................................81, 181, 262 Evolution .........................69, 81, 82, 181, 242, 255, 278 Experimental controls................72, 76, 77, 79, 203, 209

F Fertilization in amphiox ............................................................. 1–27 in polychaetes .........................................................v, 31 in starfish.............................................................49–67, 137, 151 Fixation ..............71, 75, 77, 93, 94, 101, 103, 112, 117, 169, 174, 175, 178, 182, 186, 198–200, 208, 210, 214, 215, 245, 273 Fluorescence ..................... 24, 25, 32, 34, 41–46, 75, 85, 153, 197, 209, 235, 282, 283, 287 Fluorescence microscopy ......................24, 25, 34, 41, 45 Freeze substitution.................... 101, 103–105, 108–111, 116, 117 Fusion proteins.......................... 121, 124–128, 130, 133, 134, 164

G Gametes ........................2, 3, 15, 21, 22, 31–47, 58, 130, 138, 141, 142, 152, 159, 255, 258 Gel electrophoresis....................................... 66, 121, 122, 127, 128 Gene-editing...................................................32, 217–228 Gene expression ................. 82, 182, 195–197, 209, 231, 242, 249, 255, 273 Gene function...................................................22, 82, 182 Gene ontology............................................................... 237 Genomes.................. 2, 3, 12, 16, 22, 24, 25, 27, 50, 82, 181, 224, 228, 231–233, 235, 255, 257, 258, 261, 267–274, 278, 284, 290–303 Genomics .............................. 50, 82, 124, 125, 181, 201, 231, 233, 235, 236, 261, 267–270, 272–274, 277–304 Genotyping................................................................15, 22 Germinal vesicle breakdown (GVBD) ................... 59, 65, 123, 137, 141 Germline ......................................................................2, 24 Glutathione S-transferase (GST)............... 121, 124–127, 130–132 Gut rudiments ...................................................... 242–250

OTHER MARINE INVERTEBRATES: METHODS

AND

PROTOCOLS

H Hemicentrotus pulcherrimus, see Sea urchin Hox genes...................................290, 291, 299, 300, 304 HpBase ................................................................. 267–275

I Immunohistochemistry ......................168, 178, 210, 273 Immunolabeling........................102, 105, 106, 109, 112, 117, 177, 209–212 In situ hybridization (ISH) ................... 22, 25, 181–193, 196, 197, 201, 209–212, 214 Interstitial habitat................................................. 290, 291

J Juveniles............................13, 22, 25, 32, 37, 38, 50, 63, 66, 67, 69, 70, 72–75, 77, 83, 86–88, 90, 93, 94, 167, 232, 233

L Laboratory aquaria................................ 50–52, 54, 56, 64 Laboratory culture ...................................................... 1–27 Larvae ............................... 2, 7, 9, 12, 13, 25, 32, 36, 37, 49–67, 83, 84, 86, 87, 167, 175, 182, 186, 189, 190, 192, 196, 198–200, 204, 205, 208, 210, 273, 274, 290 Larval development.............................................. 289, 290 Live imaging ..............................................................83, 94 Lysates .........................66, 120, 125–127, 133, 134, 259

M Maculaura alaskensis, see Nemertea Maturation promoting factor (MPF)............49, 138, 141 Meiofauna............................................................. 289–304 Metamorphosis in amphioxus ............................................2, 12, 13, 22 in starfish.......................................................... v, 49–67 Metazoans.................................................. 49, 81, 82, 254 1-methyladenine (1-MeAde)..................... 49, 52, 59, 62, 65, 120, 123, 154 Microinjection in amphioxus ..............................................2, 8, 11, 20 in ascidians...................................................... 223, 224 in eggs....................2, 8, 11, 20, 32, 44, 45, 224, 273 in polychaetes ...............................................................v in sea urchin ............................................................ 273 Miniaturization..................................................... 290, 291 Mitogen-activated protein kinase (MAPKs)....... 138, 218 Morphogenesis ......................................83, 218, 221, 263 Morpholino oligo-nucleotides (MO) .......................... 273 mRNA synthesis .............................................................. 14 Multiplexing .................................................................. 210 Mutagenesis..............................1–27, 120, 124, 125, 132

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AND

OTHER MARINE INVERTEBRATES: METHODS

AND

PROTOCOLS Index 309

N

R

Nematostella vectensis, see Cnidarian Nemertea .............................................................. 138, 290 Nerve cord......................................................12, 217–228 Neurulae .......................................................................... 12 Nitrocellulose ............................122, 126, 128, 132, 133, 155, 158, 160 Nutrition....................................................................65, 84

Regeneration ........ 69–80, 163–179, 231, 232, 242, 244 RNA .......... 70, 119, 182, 185, 195–215, 232–235, 238, 241–252, 278, 281, 282, 285 RNA interference (RNAi) .........182, 241, 247, 249, 251 RNA isolation................................................................ 232 RNA probes................................185, 186, 203, 273, 274

O Oocyte aging ........................................................ 137–149 Oocyte maturation ........................ 33, 36, 37, 41, 44, 50, 58, 59, 123, 124, 256 Oocytes .....................2, 3, 32, 34–37, 39–41, 44, 45, 49, 52, 53, 57, 59, 65, 120, 122, 123, 131, 137, 138, 141, 142, 153, 154, 156, 159, 196, 258 Oscarella lobularis, see Sponge

P Paracentrotus lividus, see Sea urchin Paraformaldehyde (PFA) ..................................71, 72, 79, 84–93, 106, 112, 117, 167, 174, 183, 198, 199, 243, 245, 250 Patch clamp .......................................................... 285, 287 Patiria miniata, see Sea star Phallusia mammillata, see Ascidian Phosphate-buffered saline (PBS) .....................23, 71, 72, 84, 85, 88, 89, 91, 106, 112, 113, 167, 169–173, 177, 183, 184, 192, 199, 210–212, 243, 245, 246, 280, 283, 287 Phospholipase C gamma (PLC)................. 120, 130, 151 Placozoa................................................... 81, 99, 181, 277 Plasmids ............... 4–6, 14, 16, 17, 19, 20, 24, 120, 124, 125, 201, 209, 274, 282–284 Polychaetes ...................................................................... 31 Polymerase chain reaction (PCR) ................5, 14, 16–21, 23–26, 124, 132, 183, 185, 186, 196, 201, 225, 255, 257, 260, 263, 274, 282, 283 Porifera, see Sponge Programmed cell death................................................. 137 Proliferation....................... 70, 75, 79, 83, 163–179, 253 Pseudopotamilla occelata, see Polychaete

S Scanning electron microscopy (SEM) ............... 101, 103, 105, 111, 112, 114, 115 Sea cucumbers .....................................242, 244, 246, 250 Sea stars.....................119–134, 151–160, 197–199, 204, 205, 208, 211, 255, 258 Sea urchins................142, 197–201, 204–208, 210, 211, 255, 258, 261, 262, 267–274 Signal transduction .............................................. 119–134 Silver stain.....................................................129–131, 134 Spawnings ........................ 2, 7, 9–12, 26, 54, 56, 72, 258 Sponges......................34, 58, 81–95, 128, 145, 181–193 Src Family Kinase .......................................................... 120 Starfish, see Sea star Strongylocentrotus purpuratus, see Sea urchin

T TALENs............................................................ 2–6, 12–23 Tol2................................................................... 2, 6, 22–26 Transcription factors (TFs).................................. 253, 255 Transcriptomes ............................50, 232, 233, 235, 238, 268–270, 272, 296 Transfections .............. 82, 242, 250, 280, 283, 285, 287 Transgenics ..................................... 2, 22, 24, 25, 27, 262 Transmission electron microscopy (TEM) .................101, 103, 111 Transposons........................................................ 2, 22, 261 Trichoplax, see Placozoa Troubleshooting ..........................................196–198, 209 Tyrosine phosphorylation..........126, 128, 130, 132, 133

W Western blotting........................121, 122, 126, 128, 130, 134, 153–155, 158, 159