Methods in Cell Biology VOLUME 74 Development of Sea Urchins, Ascidians, and Other Invertebrate Deuterostomes: Experimental Approaches
Series Editors Leslie Wilson Department of Biological Sciences University of California Santa Barbara, California
Paul Matsudaira Whitehead Institute for Biomedical Research Department of Biology Massachusetts Institute of Technology Cambridge, Massachusetts
Methods in Cell Biology Prepared under the Auspices of the American Society for Cell Biology
VOLUME 74 Development of Sea Urchins, Ascidians, and Other Invertebrate Deuterostomes: Experimental Approaches Edited by
Charles A. Ettensohn Department of Biological Sciences Carnegie Mellon University Pittsburgh, Pennsylvania
Gregory A. Wray Department of Biology Duke University Durham, North Carolina
Gary M. Wessel Department of Molecular and Cell Biology and Biochemistry Brown University Providence, Rhode Island
Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK
This book is printed on acid-free paper. Copyright ß 2004, Elsevier Inc. All Rights Reserved.
No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2004 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0091-679X/2004 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail:
[email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Customer Support’’ and then ‘‘Obtaining Permissions.’’ For all information on all Academic Press publications visit our Web site at www.academicpress.com ISBN: 0-12-480278-8 Case ISBN: 0-12-480279-6 Paperback PRINTED IN THE UNITED STATES OF AMERICA 04 05 06 07 08 9 8 7 6 5 4 3 2 1
CONTENTS
Contributors
xvii
Preface
xxiii
Dedication
xxv
1. The Invertebrate Deuterostomes: An Introduction to Their Phylogeny, Reproduction, Development, and Genomics Charles A. Ettensohn, Gary M. Wessel, and Gregory A. Wray I. II. III. IV. V.
PART I
Introduction Phylogeny Reproduction Development Genomics References
1 2 5 8 10 11
Procurement, Maintenance & Culture of Oocytes, Embryos, Larvae and Adults
2. Care and Maintenance of Adult Echinoderms S. Anne Bo¨ttger, Charles W. Walker, and Tatsuya Unuma I. II. III. IV. V. VI.
Overview Introduction Adult Echinoderm Models: Their Reproductive Cycles and Gametogenesis Obtaining Adult Echinoderms Maintenance of Adult Echinoderms in Land-Based Systems Care and Handling of Adult Echinoderms References
18 18 18 24 26 28 33
v
vi
Contents
3. Echinoderm Eggs and Embryos: Procurement and Culture Kathy R. Foltz, Nikki L. Adams, and Linda L. Runft I. II. III. IV. V. VI. VII.
Introduction Method for Sea Urchins (Class Echinoidea) Method for Sea Stars (Class Asteroidea) Method for Brittle Stars (Class Ophiuroidea, Order Ophiurida) Method for Sea Cucumbers (Class Holothuroidea) Method for Interspecific Crosses Summary References
40 41 53 60 64 68 70 71
4. Culture of Echinoderm Larvae through Metamorphosis Gregory A. Wray, Chisato Kitazawa, and Benjamin Miner I. II. III. IV. V.
Introduction Materials for Culturing Algae and Larvae Establishing Algal Cultures Culturing Echinoderm Larvae Metamorphosis and Beyond References
75 76 78 81 84 85
5. Obtaining and Handling Echinoderm Oocytes Gary M. Wessel, Ekaterina Voronina, and Jacqueline M. Brooks I. II. III. IV. V. VI. VII.
Introduction Experimental Preparation Methods of Oocyte Collection Sea Urchin Oocytes Cultured In Vitro Labeling of Oocyte Components Introduction of Experimental Substances into the Oocytes Concluding Remarks/Outlook References
88 88 92 98 101 110 113 114
6. Procurement and Culture of Ascidian Embryos Billie J. Swalla I. II. III. IV. V.
Overview Ascidian Development and Metamorphosis Larval Tissue Specification Experimental Techniques Protocols References
116 116 126 129 132 139
vii
Contents
7. Culture of Adult Ascidians and Ascidian Genetics Carolyn Hendrickson, Lionel Christiaen, Karine Deschet, Di Jiang, Jean-Ste´phane Joly, Laurent Legendre, Yuki Nakatani, Jason Tresser, and William C. Smith I. II. III. IV. V.
Overview Introduction Culturing Ascidians Induced Developmental Mutants and Natural Variants/Mutants Linkage Analysis and Mapping Genes in Ascidians References
143 144 145 153 160 169
8. Hemichordate Embryos: Procurement, Culture, and Basic Methods Christopher J. Lowe, Kuni Tagawa, Tom Humphreys, Marc Kirschner, and John Gerhart I. II. III. IV. V. VI. VII.
Introduction Procurement, Spawning, and Culture of S. kowalevskii Procurement, Spawning, and Culture of Ptychodera flava Removal of Vitelline Envelope in S. kowalevskii Whole-Mount In Situ Hybridization Preparation of Blocking Reagent for P. flava Materials and Reagents References
172 173 179 182 183 190 191 192
9. Cephalochordate (Amphioxus) Embryos: Procurement, Culture, and Basic Methods Linda Z. Holland and Ju-Ka Yu I. II. III. IV. V. VI. VII.
Overview Introduction Obtaining Gametes of Branchiostoma floridae Raising Embryos Manipulating Embryos Amphioxus Resources Available Concluding Remarks References
195 196 200 202 206 212 213 213
viii
Contents
PART II Embryological Approaches 10. Quantitative Microinjection of Oocytes, Eggs, and Embryos Laurinda A. Jaffe and Mark Terasaki I. Introduction II. Methods III. Equipment and Supplies (Prices are as of December, 2002) References
219 220 230 242
11. Blastomere Isolation and Transplantation Hyla Sweet, Shonan Amemiya, Andrew Ransick, Takuya Minokawa, David R. McClay, Athula Wikramanayake, Ritsu Kuraishi, Masato Kiyomoto, Hiroki Nishida, and Jonathan Henry I. II. III. IV. V. VI. VII. VIII.
Introduction Preparation of Mouth Pipettes and Needles Removal of Fertilization Envelope of Sea Urchin Eggs Isolation and Recombination of Blastomeres in Sea Urchin Embryos Blastomere Isolation and Transplantation in Starfish Embryos Blastomere Isolation and Transplantation in Amphioxus Blastomere Isolation and Transplantation in Ascidians Lineage Tracing, Blastomere Isolation, and Transplantation in Hemichordates References
244 245 248 249 263 265 266 267 270
12. Isolation and Culture of Micromeres and Primary Mesenchyme Cells Fred H. Wilt and Stephen C. Benson I. Introduction II. Isolation and Culture of Micromeres III. Isolation and Culture of Primary Mesenchyme Cells References
273 275 282 284
13. Rapid Microinjection of Fertilized Eggs Melani S. Cheers and Charles A. Ettensohn I. Introduction II. Equipment III. Experimental Protocols References
287 289 292 310
ix
Contents
14. Methods for Embryo Dissociation and Analysis of Cell Adhesion David R. McClay I. II. III. IV.
Introduction Gaining Access to the Egg and Embryo Cell Adhesion Assays: Nonquantitative and Semiquantitative Quantitative Centrifugation Assay References
311 312 317 320 328
PART III Cell Biological Approaches 15. Analysis of Sea Urchin Embryo Gene Expression by Immunocytochemistry Judith M. Venuti, Carmen Pepicelli, and Vera Lynn Flowers I. II. III. IV. V.
Introduction Immunocytochemistry on Whole Embryos Immunostaining Small Numbers of Embryos Co-localization Troubleshooting References
334 335 351 359 364 366
16. Light Microscopy of Echinoderm Embryos Laila Strickland, George von Dassow, Jan Ellenberg, Victoria Foe, Peter Lenart, and David Burgess I. II. III. IV.
Introduction (D. Burgess and L. Strickland) Formaldehyde Fixation of Cleavage Stage Sea Urchin Embryos Staining and Imaging Fixed Embryos Simultaneous Fixation and Visualization of the Actin and Microtubule Cytoskeletons (G. von Dassow and V. Foe) V. Observation of Live Embryos (D. Burgess and L. Strickland) VI. 4-D Imaging of Fluorescent Markers in Live Starfish Oocytes ( J. Ellenberg and P. Lenart) References
372 372 382 385 393 399 407
17. TEM and SEM Methods Bruce J. Crawford and Robert D. Burke I. II. III. IV.
Introduction Basic Fixation and Preparation Embedding Media Preservation of Extracellular Matrix
412 412 421 424
x
Contents
V. VI. VII. VIII. IX. X.
Fixation for SEM Quick Freezing and Freeze-Substitution Fixation for Immunohistochemistry and Immunocytochemistry Immunogold Methods Ultrastructural Immunoperoxidase Preparation of Colloidal Gold Reagents (Horisberger, 1981; Slot and Geuze, 1985) as modified by Campbell (1990) and Reimer (1994) References
426 428 434 434 436
437 440
18. Calcium Imaging Michael Whitaker I. II. III. IV. V. VI. VII. VIII.
Introduction Calcium Signals in Embryos Getting Calcium Sensors into Embryos Measuring Emitted Light Fluorescence Lifetime Imaging Calibration Manipulating Intracellular Calcium Conclusions and Perspectives References
443 444 450 454 459 459 460 462 463
19. Labeling of Cell Membranes and Compartments for Live Cell Fluorescence Microscopy Mark Terasaki and Laurinda A. Jaffe I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII.
Introduction Labeling of Extracellular Space/Endocytosis/Exocytosis Labeling of Plasma Membrane/Exocytosis/Endosomes Labeling of the Cytosol Yolk Platelets/Reserve Granules Mitochondria Other Organelles Endoplasmic Reticulum Golgi Apparatus Nucleus Microscopy Considerations Future Directions References
469 471 472 475 476 478 478 479 483 485 486 487 487
xi
Contents
20. Isolation of Organelles and Components from Sea Urchin Eggs and Embryos Gary M. Wessel and Victor D. Vacquier I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII. XIII. XIV. XV. XVI.
Overview Egg Jelly Molecules Affecting Sperm Isolation of the Vitelline Layer from Sea Urchin Eggs Isolation of the Cell Surface Complex and the Plasma Membrane–Vitelline Layer (PMVL) Complex from Sea Urchin Eggs The Cytolytic Isolation of the Egg Cortex Isolation of Cortical Granules Isolation of Yolk Platelets Isolation of Mitochondria from Eggs and Embryos Isolation of Plasma Membranes and Lipid Rafts from Eggs and Zygotes Isolation of Microsomes Containing the Endoplasmic Reticulum Nuclear Isolation Procedures Removal and Isolation of the Fertilization Envelope Isolation of Cilia from Embryos Isolation of Extracellular Matrices from Embryos and Larvae Isolation of Sea Urchin Larval Skeletons Resources for the Isolation of Additional Organelles References
492 492 496 498 500 501 503 504 505 507 507 508 511 512 517 518 519
21. Sea Urchin Spermatozoa Victor D. Vacquier and Noritaka Hirohashi I. II. III. IV. V. VI. VII. VIII. IX. X. XI.
Introduction Obtaining Sperm and Removing Coelomocytes Isolating Sperm Heads, Flagella, and Chromatin The Isolated Sperm Flagellum as a Sealed Compartment Extracting Intact Sperm or Isolated Flagella and Heads with Detergents Wheat Germ Agglutinin (WGA) Affinity Chromatography of Sperm Membrane Proteins Isolating Sperm Plasma Membranes Isolation of Acrosome Reaction Vesicle Membranes (ARV) Isolating Lipid Rafts from Sperm Scoring the Acrosome Reaction by Phase Contrast Microscopy Sea Urchin Sperm Bindin References
524 528 529 530 531 532 532 535 536 537 539 540
22. Measuring Ion Fluxes in Sperm Alberto Darszon, Christopher D. Wood, Carmen Beltra´n, Daniel Sa´nchez, Esmeralda Rodrı´guez, Julia Gorelik, Yuri E. Korchev, and Takuya Nishigaki I. Overview II. Introduction
546 546
xii
Contents
III. Sperm Physiology is Deeply Influenced by Ion Channels and Transporters IV. Strategies to Study Sperm Ion Channels and Transporters References
547 549 573
PART IV Molecular Biological Approaches 23. Isolating DNA, RNA, Polysomes, and Protein Bruce P. Brandhorst I. II. III. IV. V. VI.
Overview Introduction Purification of Total RNA Purification of Cytoplasmic and Polysomal RNA Purification of Genomic DNA Preparation of Protein Samples for One- and Two-Dimensional Gel Electrophoresis References
580 580 580 586 589 592 598
24. Detection of mRNA by In Situ Hybridization and RT-PCR Andrew Ransick I. Introduction II. In Situ Hybridization III. Quantitative PCR References
601 602 613 618
25. Using Reporter Genes to Study cis-Regulatory Elements Maria I. Arnone, Ivan J. Dmochowski, and Christian Gache I. II. III. IV. V. VI. VII.
Overview Introduction Using Chloramphenicol Acetyltransferase (CAT) Reporter Gene Using Green Fluorescent Protein (GFP) Reporter Gene Quantitative Imaging of GFP in Living Embryos Using Luciferase (luc) Reporter Gene Using lacZ Reporter Gene Summary, Prospects, Concluding Remarks References
622 622 627 632 633 640 643 648 649
xiii
Contents
26. Identification of Sequence-Specific DNA Binding Proteins James A. Coffman and Chiou-Hwa Yuh Overview I. Introduction II. Preparation of Nuclear Extracts III. Identification of Transcription Factor Target Sites by Footprint and Electrophoretic Mobility Shift Analysis (EMSA) IV. Affinity Purification of Sequence-Specific DNA Binding Proteins V. Identification of Affinity-Purified DNA-Binding Proteins in SDS Gels VI. Preparation of Affinity-Purified Transcription Factors for Protein Sequencing VII. Prospects References
654 654 655 658 662 666 671 673 674
27. Expression of Exogenous mRNAs to Study Gene Function in the Sea Urchin Embryo Thierry Lepage and Christian Gache Overview I. Introduction II. Methods for Expressing Synthetic mRNA in the Sea Urchin Embryo III. Summary, Prospects, Concluding Remarks References
677 678 688 695 696
28. Disruption of Gene Function Using Antisense Morpholinos Lynne M. Angerer and Robert C. Angerer I. II. III. IV.
Introduction Morpholino-Mediated Loss-of-Function: Advantages and Limitations Morpholino Methods Concluding Remarks References
699 700 704 709 710
29. Generation and Use of Transgenic Ascidian Embryos Robert W. Zeller I. II. III. IV. V. VI.
Overview Ascidian Embryos as Model Chordate Embryos Transgenic Ascidian Embryos Fundamentals of Electroporation Transgene Construction Detailed Protocols for Electroporation of Ascidian Zygotes List of Required Chemicals and Supplies References
713 714 716 719 720 721 725 726
xiv
Contents
PART V Genomics 30. Genomic Resources for the Study of Sea Urchin Development R. Andrew Cameron, Jonathan P. Rast, and C. Titus Brown I. II. III. IV.
Overview Introduction Sea Urchin Arrayed Library Resources Computational Tools Sea Urchin Genome Project Web Site References
734 734 736 746 750 755
31. Genomic Resources for Ascidians: Sequence/Expression Databases and Genome Projects Nori Satoh I. II. III. IV.
Overview Introduction The cDNA Projects: cDNA Sequence and Expression Databases The Genome Project Conclusion References
759 760 762 769 771 772
32. Gene Regulatory Network Analysis in Sea Urchin Embryos Paola Oliveri and Eric H. Davidson I. II. III. IV.
Introduction Perturbation Analysis Assembling and Testing the GRN Logic Map Summary References
775 777 787 792 793
PART VI Echinoderm Eggs and Embryos in the Teaching Lab 33. Sea Urchin Gametes in the Teaching Laboratory: Good Experiments and Good Experiences David Epel, Victor D. Vacquier, Margaret Peeler, Pam Miller, and Chris Patton Overview I. Introduction II. Obtaining Adult Urchins and Gametes for the Classroom
798 798 799
xv
Contents
III. IV. V. VI.
Some Guidelines for Using Sea Urchin Gametes in the Classroom Basic Introductory Labs Inquiry-Based Experiments Using the Basic Fertilization Protocol Inquiry-Based Experiments in Later Development: Experiments on Differential Gene Expression VII. Experiments Using Morphology as an Endpoint VIII. Epilogue References
Appendix
801 802 810 819 821 822 822
825
Index
841
Volumes in Series
877
This Page Intentionally Left Blank
CONTRIBUTORS
Numbers in parentheses indicate the pages on which authors’ contributions begin.
Nikki L. Adams (39), Department of Biological Sciences, California Polytechnic State University, San Luis Obispo, California 93407 Shonan Amemiya (243), Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa, Chiba 277-8562, Japan Lynne M. Angerer (699), Department of Biology, University of Rochester, Rochester, New York 14627 Robert C. Angerer (699), Department of Biology, University of Rochester, Rochester, New York 14627 Maria I. Arnone (621), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121 Napoli, Italy Carmen Beltra´n (545), Department of Developmental Genetics and Molecular Physiology, Institute of Biotechnology, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Me´xico Stephen C. Benson (273), Department of Biological Sciences, California State University, Hayward, Hayward, California 94542 S. Anne Bo¨ttger (17), Department of Zoology and Marine Biology, Biomedical Research Group, Durham, New Hampshire 03824 Bruce P. Brandhorst (579), Department of Molecular Biology and Biochemistry, Simon Fraser University, Burnaby, British Columbia, V5A 1S6, Canada C. Titus Brown (733), Division of Biology and the Center for Computational Regulatory Genomics, Beckman Institute, California Institute of Technology, Pasadena, California 91125 Jacqueline M. Brooks (87), Department of Molecular and Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912 David Burgess (371), Department of Biology, Boston College, Chestnut Hill, Massachusetts 02167 Robert D. Burke (411), Departments of Biology and Biochemistry/Microbiology, University of Victoria, Victoria, British Columbia V8W 3P6, Canada R. Andrew Cameron (733), Division of Biology and the Center for Computational Regulatory Genomics, Beckman Institute, California Institute of Technology, Pasadena, California 91125 Melani S. Cheers (287), Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213 Lionel Christiaen (143), INRA Junior Group, UPR 2197 DEPSN, CNRS, Institut de Neurobiologie A. Fessard, Gif-sur-Yvette, France
xvii
xviii
Contributors
James A. Coffman (653), Stowers Institute for Medical Research, Kansas City, Missouri 64110 Bruce J. Crawford (411), Department of Anatomy and Cell Biology, University of British Columbia, Vancouver, British Columbia V6T 1Z4, Canada Alberto Darszon (545), Department of Developmental Genetics and Molecular Physiology, Institute of Biotechnology, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Me´xico Eric H. Davidson (775), Division of Biology, California Institute of Technology, Pasadena, California 91125 Karine Deschet (143), Neuroscience Research Institute and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 Ivan J. Dmochowski (621), Department of Chemistry, University of Pennsylvania, Philadelphia, Pennsylvania 19104 Jan Ellenberg (371), European Molecular Biological Lab, Heidelberg, Germany David Epel (797), Hopkins Marine Station of Stanford University, Pacific Grove, California 93950 Charles A. Ettensohn (1, 287), Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213 Vera Lynn Flowers (333), Department of Cell Biology and Anatomy, Louisiana State University Health Sciences Center, New Orleans, Louisiana 70112 Victoria Foe (371), Friday Harbor Laboratories, University of Washington, Seattle, Washington 98195 Kathy R. Foltz (39), Department of Molecular, Cellular, and Developmental Biology, and the Marine Science Institute, University of California, Santa Barbara, California 93106 Christian Gache (621, 677), Laboratory of Developmental Biology, CNRS-Universite´ Pierre et Marie Curie (Paris VI), Observatoire Oce´anologique, 06230 Villefranchesur-Mer, France John Gerhart (171), Department of Molecular and Cellular Biology, University of California Berkeley, Berkeley, California 94720 Julia Gorelik (545), Division of Medicine, Imperial College London, MRC Clinical Sciences Centre, London W12 0NN, United Kingdom Carolyn Hendrickson (143), INRA Junior Group, UPR 2197 DEPSN, CNRS Institut de Neurobiologie A. Fessard, Gif-sur-Yvette, France Jonathan Henry (243), Department of Cell and Structural Biology, University of Illinois, Urbana, Illinois 61801 Noritaka Hirohashi (523), Department of Biology, Ochanomizu University, Tokyo, 112-8610 Japan Linda Z. Holland (195), Marine Biology Research Division, Scripps Institution of Oceanography, University of California San Diego, La Jolla, California 92093 Tom Humphreys (171), Kewalo Marine Laboratory, Honolulu, Hawaii 96813 Laurinda A. Jaffe (219, 469), Department of Cell Biology, University of Connecticut Health Center, Farmington, Connecticut 06032
Contributors
xix Di Jiang (143), Neuroscience Research Institute and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 Jean-Ste´phane Joly (143), INRA Junior Group, UPR 2197 DEPSN, CNRS, Institut de Neurobiologie A. Fessard, Gif-sur-Yvette, France Marc Kirschner (171), Systems Biology, Harvard Medical School, Boston, Massachusetts 02115 Chisato Kitazawa, (75), Department of Biology, Duke University, Durham, North Carolina 27708 Masato Kiyomoto (243), Tateyama Marine Laboratory, Ochanomizu University, Tateyama, Chiba 294-0301, Japan Yuri E. Korchev (545), Division of Medicine, Imperial College London, MRC Clinical Sciences Centre, London W12 0NN, United Kingdom Ritsu Kuraishi (243), Marine Biological Station, Graduate School of Science, Tohoku University, Aomori 039-3501, Japan Laurent Legendre (143), INRA Junior Group, UPR 2197 DEPSN, CNRS, Institut de Neurobiologie A. Fessard, Gif-sur-Yvette, France Peter Lenart (371), European Molecular Biological Lab, Heidelberg, Germany Thierry Lepage (677), Laboratory of Developmental Biology, CNRS-Universite´ Pierre et Marie Curie (Paris VI), Observatoire Oce´anologique, 06230 Villefranchesur-Mer, France Christopher J. Lowe (171), Department of Organismal Biology and Anatomy, University of Chicago, Chicago, Illinois 60612 and Department of Molecular and Cellular Biology, University of California Berkeley, Berkeley, California 94720 David R. McClay (243, 311), Department of Biology, Duke University, Durham, North Carolina 27708 Pam Miller (797), Seaside High School, Seaside, California 93955 Benjamin Miner (75), Department of Zoology, University of Florida, Gainesville, Florida 32611 Takuya Minokawa (243), Division of Biology, California Institute of Technology, Pasadena, California 91125 Yuki Nakatani (143), Neuroscience Research Institute and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 Hiroki Nishida (243), Department of Biological Sciences, Tokyo Institute of Technology, Nagatsuta, Midori-ku, Yokohama 226-8501, Japan Takuya Nishigaki (545), Department of Developmental Genetics and Molecular Physiology, Institute of Biotechnology, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Me´xico Paola Oliveri (775), Division of Biology, California Institute of Technology, Pasadena, California 91125 Chris Patton (797), Hopkins Marine Station of Stanford University, Pacific Grove, California 93950
xx
Contributors
Margaret Peeler (797), Department of Biology, Susquehanna University, Selinsgrove, Pennsylvania 17870 Carmen Pepicelli (333), Curis, Inc., Cambridge, Massachusetts 02138 Andrew Ransick (243, 601), Division of Biology, California Institute of Technology, Pasadena, California 91125 Jonathan P. Rast (731), Division of Biology and the Center for Computational Regulatory Genomics, Beckman Institute, California Institute of Technology, Pasadena, California 91125 Esmeralda Rodrı´guez (545), Department of Developmental Genetics and Molecular Physiology, Institute of Biotechnology, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Me´xico Linda L. Runft (39), Department of Molecular, Cellular, and Developmental Biology, and the Marine Science Institute, University of California, Santa Barbara, California 93106 Daniel Sa´nchez (545), Division of Medicine, Imperial College London, MRC Clinical Sciences Centre, London W12 0NN, United Kingdom Nori Satoh (759), Department of Zoology, Graduate School of Science, Kyoto University, Kyoto 606-8502, Japan William C. Smith (143), Neuroscience Research Institute and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 Laila Strickland (371), Department of Biology, Boston College, Chestnut Hill, Massachusetts 02167 Billie J. Swalla (115), Biology Department and Friday Harbor Laboratories, University of Washington, Seattle, Washington 98195 Hyla Sweet (243), Department of Biological Sciences, College of Science, Rochester Institute of Technology, Rochester, New York 14623 Kuni Tagawa (171), Kewalo Marine Laboratory, Honolulu, Hawaii 96813 Mark Terasaki (219, 469), Department of Cell Biology, University of Connecticut Health Center, Farmington, Connecticut 06032 Jason Tresser (143), Neuroscience Research Institute and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 Tatsuya Unuma (17), National Research Institute of Aquaculture, Nansei, Mie 516-0193, Japan Victor D. Vacquier (491, 523, 797), Center for Marine Biotechnology and Biomedicine, Scripps Institution of Oceanography, University of California, San Diego, La Jolla, California 92093 Judith M. Venuti (333), Department of Cell Biology and Anatomy, Louisiana State University Health Sciences Center, New Orleans, Louisiana 70112 George von Dassow (371), Friday Harbor Laboratories, University of Washington, Seattle, Washington 98195 Ekaterina Voronina (87), Department of Molecular and Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912
Contributors
xxi Charles W. Walker (17), Department of Zoology and Marine Biology, Biomedical Research Group, Durham, New Hampshire 03824 Gary M. Wessel (1, 87, 491), Department of Molecular and Cell Biology and Biochemistry, Brown University, Providence, Rhode Island 02912 Michael Whitaker (443), School of Cell & Molecular Biosciences, Faculty of Medical Sciences, University of Newcastle upon Tyne, Framlington Place, NE2 4HH, United Kingdom Athula Wikramanayake (243), Department of Zoology, University of Hawaii at Manoa, Honolulu, Hawaii 96822 Fred H. Wilt (273), Department of Molecular Cell Biology, University of California, Berkeley, California 94720 Christopher Wood (545), Department of Developmental Genetics and Molecular Physiology, Institute of Biotechnology, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Me´xico Gregory A. Wray (1, 75), Department of Biology, Duke University, Durham, North Carolina 27708 Ju-Ka Yu (195), Marine Biology Research Division, Scripps Institution of Oceanography University of California, San Diego, La Jolla, California 92093 Chiou-Hwa Yuh (653), Division of Biology 156-29, California Institute of Technology, Pasadena, California 91125 Robert W. Zeller (713), Department of Biology, San Diego State University, San Diego, California 92182
This Page Intentionally Left Blank
PREFACE
The past decade has been one of the most exciting periods in the history of developmental biology. Numerous genomes have been sequenced, powerful new experimental approaches are rapidly being developed, and we are now able to visualize processes in living embryos in ways that that were inconceivable a short while ago. From these powerful new experimental tools has emerged a much richer view of developmental processes across scales of biological organization, as well as a growing appreciation for the shared genetic and molecular underpinnings of development in diverse species. Within this broad research effort, the invertebrate deuterostomes are playing an important role. As the closest living relatives of vertebrates, the invertebrate deuterostomes occupy a uniquely informative phylogenetic position, providing a crucial perspective on the organization and evolution of vertebrate genomes and clues concerning origin of the vertebrate body plan. As experimental subjects, the embryos of these animals provide a singular and powerful combination of advantages: they are simple, optically transparent, develop externally, are robust to physical manipulation, and can be raised in large quantities. Invertebrate deuterostomes have genomes that are similar to those of vertebrates in many respects, but contain fewer gene duplications. With the completion of an ascidian genome project in 2003 and the imminent completion of a sea urchin genome project in 2004, we are entering the ‘‘postgenome’’ era for the invertebrate deuterostomes. As the pace of research accelerates, there is a clear need for accessible protocols to guide experimental studies with the embryos of these animals. The most recent compendium of experimental methods was published 18 years ago. ‘‘Echinoderm Gametes and Embryos,’’ edited by Thomas Schroeder, has been an invaluable resource at the bench, but many essential new methods have been developed since that book was published and an updated collection of methods is clearly needed. Moreover, while the focus of the earlier volume was on echinoderms, a growing number of researchers are working with other invertebrate deuterostomes: urochordates, hemichordates, and cephalochordates. Our aim with the present volume is to bring together a set of protocols for rearing and working with these embryos. This book would not have been possible without the collective efforts of the research community to which we belong: the scientists who study the development of invertebrate deuterostomes. We would like to extend our thanks and appreciation to the many people who developed and tested the methods described in the following chapters, and who have inspired us by applying these methods to a wide range of exciting problems in developmental biology. We also thank our xxiii
xxiv
Preface
many colleagues who wrote, reviewed, and otherwise contributed to the chapters in this book. To the extent that this book proves useful, it will be thanks to their generous efforts to distill years of experience into brief, practical guidelines. Finally, we extend our special thanks to Mica Haley and Kristi Savino at Elsevier, who patiently guided us through the process of assembling this book. Charles A. Ettensohn Gregory A. Wray Gary M. Wessel
The editors dedicate this volume to their scientific mentor, Dr. David R. McClay, the Arthur S. Pearse Professor of Biology at Duke University.
xxv
CHAPTER 1
The Invertebrate Deuterostomes: An Introduction to Their Phylogeny, Reproduction, Development, and Genomics Charles A. Ettensohn,* Gary M. Wessel,{ and Gregory A. Wray{ *Department of Biological Sciences Carnegie Mellon University Pittsburgh, Pennsylvania 15213 {
Department of Biology Duke University Durham, North Carolina 27708
{
Department of Molecular Biology, Cell Biology, and Biochemistry Brown University Providence, Rhode Island 02912
I. II. III. IV.
Introduction Phylogeny Reproduction Development A. Historical Contributions B. Experimental Characteristics C. Shared Features of Development V. Genomics References
I. Introduction The invertebrate deuterostomes comprise approximately 10,000 species of marine animals distributed throughout the oceans of the world. Early studies on the eggs and embryos of these animals played a critical historical role in embryology, METHODS IN CELL BIOLOGY, VOL. 74 Copyright 2004, Elsevier Inc. All rights reserved. 0091-679X/04 $35.00
1
2
Charles A. Ettensohn et al.
cell biology, and genetics. They have continued to be widely used as model organisms for developmental and cell biological studies for more than a century. In recent years, the traditional strengths of these systems for developmental studies have been supplemented by a variety of powerful methodologies for analyzing and perturbing gene expression and function. Moreover, with the advent of modern genomics and the re-emergence of evolutionary–developmental biology, this group of organisms promises to yield critical new insights in many areas. Analysis of early embryonic patterning and gene regulatory networks is advancing rapidly. The invertebrate deuterostomes are also proving to be a rich resource for analysis of the evolution of developmental programs. There are two major reasons for this: (1) embryos of a diverse collection of species can be obtained and studied with relative ease, and (2) there are examples of remarkable variations in life cycles, developmental patterns, and adult body plans within the invertebrate deuterostomes, notwithstanding the many shared features of their development. Finally, the invertebrate deuterostomes are the closest living relatives of the vertebrates, and analysis of their development promises to shed light on the origins of the vertebrates, the chordates, and the entire deuterostome lineage.
II. Phylogeny Recent molecular phylogenetic studies suggest that the deuterostomes include only the echinoderms, hemichordates, and chordates (Adoutte et al., 2000; Aguinaldo et al., 1997) (Fig. 1). Certain groups that were previously considered to be deuterostomes, notably, the chaetognaths and lophophorates, are instead likely to be protostomes. The invertebrate deuterostomes consist of all deuterostomes outside Subphylum Vertebrata; i.e., the echinoderms (Phylum Echinodermata), hemichordates (Phylum Hemichordata), and the members of two invertebrate subphyla within the chordates, Urochordata and Cephalochordata (Fig. 1). Among the invertebrate deuterostomes, morphological and molecular data indicate that echinoderms and hemichordates are closely related to one another and represent a distinct clade, that the urochordates are monophyletic and could be considered a separate phylum, and that the cephalochordates are most closely related to vertebrates (Bromham and Degnan, 1999; Cameron et al., 2000; Swalla et al., 2000; Fig. 1). A close relationship between hemichordates and echinoderms is also supported by recent studies showing similarities in patterns of gene expression during early embryogenesis (Gross and McClay, 2001; Harada et al., 2002; Nakajima et al., 2004; Shoguchi et al., 1999; Tagawa et al., 1998). Most present-day species of invertebrate deuterostomes are members of Phylum Echinodermata (7000 species). Adult echinoderms are bottom-dwelling, radially symmetrical animals, but their larvae are free-swimming and bilaterally symmetrical (Fig. 2A, B, Fig. 3I). Modern echinoderms are grouped into five classes: Echinodea (sea urchins and sand dollars), Holothuroidea (sea cucumbers), Asteroidea (starfish), Ophiuroidea (brittle stars), and Crinoidea (feathers stars and
1. Invertebrate Deuterostomes
3
Fig. 1 A phylogenetic tree of the bilaterally symmetrical animals (Bilateria). These are usually grouped into three multiphyletic clades: Ecdysozoa, Lophotrochozoa, and Deuterostomia. Taxa that comprise the invertebrate deuterostomes (the organisms considered in this book) are underlined. They consist of Phylum Echinodermata and Phylum Hemichordata, and two chordate subphyla (Subphylum Urochordata and Subphylum Cephalochordata).
sea lilies). Among the echinoderms, sea urchins have been most widely used for developmental studies. Indeed, the developmental biology of sea urchins is better understood than that of any other invertebrate deuterostome. Starfish have also been widely used for studies of gametogenesis and embryonic development, but far less attention has been paid to members of the remaining three classes of echinoderms. The urochordates, or tunicates (Subphylum Urochordata), are represented by some 3000 extant species. They are usually grouped into three classes: Ascidiacea (ascidians), Thaliacea, and Larvacea. The ascidians, the most numerous group (2300 species) have been the most thoroughly studied from a developmental perspective. They are benthic, sessile, filter feeders that develop via a bilaterally symmetrical, nonfeeding tadpole larva (Figs. 2C, Fig. 3K). Some live as individuals (solitary, or simple, ascidians) while others form colonies (colonial, or compound, ascidians). The solitary ascidians have been the most widely used for developmental studies (Satoh, 1994). Unlike ascidians, thalaceans and larvaceans remain free-swimming throughout their life cycle (Bone, 1998). The hemichordates (Phylum Hemichordata) are represented by relatively few (85) extant species and are grouped into the classes Enteropneusta, Pterobranchia, and Planctosphaeroidea. The enteropneusts (acorn worms) are likely to be basal within the group (Cameron et al., 2000) and are solitary, sedentary, burrowing worms (Fig. 2D). The pterobranchs are colonial, sessile organisms that live in secreted tubes. The monotypic Planctosphaeroidea form spherical, transparent larvae, the adult forms of which have not yet been described (see Cameron et al., 2000).
4
Charles A. Ettensohn et al.
Fig. 2 Representative adult invertebrate deuterostomes. (A) Sea urchin (Lytechinus variegatus, Phylum Echinodermata). (B) Starfish (Linkia lavigata, Phylum Echinodermata). (C) Ascidians (Boltenia villosa and Styela gibbsii, Subphylum Urochordata). Photograph courtesy of Billie Swalla. (D) Hemichordate (Saccoglossus kowalevskii, Phylum Hemichordata). Photograph courtesy of Chris Lowe. (E) Amphioxus (Branchiostoma floridae, Subphylum Cephalochordata). Photograph courtesy of Linda Holland. (See Color Insert.)
The cephalochordates, commonly known as amphioxus (or the lancelet), include 28 diVerent species, grouped into two genera. They are the closest living relatives of the vertebrates and share a dorsal hollow nerve cord, notochord, and phayngeal
1. Invertebrate Deuterostomes
5
gill slits, but lack paired limbs, paired ears, and image-forming eyes (Fig. 2E). They are relatively sedentary, sand-dwelling filter feeders. Most developmental studies have been carried out on three species of the genus Branchiostoma (B. lanceolatum, B. floridae, and B. belcheri) (Holland and Yu, Chapter 9).
III. Reproduction Most species of invertebrate deuterostomes have separate sexes. The major exceptions are the ascidians, which are hermaphrodites. Sexual reproduction is by far the most common mode within the group. Most species have a distinct reproductive season during which sperm and eggs are released under appropriate environmental conditions. Invertebrate deuterostomes are typically broadcast spawners, and fertilization and development occur externally. Colonial ascidians and some echinoderms, however, brood their eggs within body cavities of the adult. Although sexual reproduction is predominant, reproduction by asexual budding occurs in a number of groups that have colonial forms, including pterobranch hemichordates and the colonial ascidians (Cameron et al., 2000; Satoh, 1994). In addition, among the echinoderms, some solitary forms are capable of asexual reproduction through the splitting of adult animals (McGovern, 2003) or larvae (Bosch et al., 1989; Eaves and Palmer, 2003). Many invertebrate deuterostomes develop through a free-swimming, feeding larval stage that bears little anatomical resemblance to the adult form (maximal indirect development) (Fig. 3). There is a complex nomenclature associated with these larval forms, particularly those of hemichordates and echinoderms, which reflects variations in certain aspects of their morphology, such as the distribution of cilia and the organization of calcified skeletal elements. For example, the larva of enteropneust hemichordates is known as a tornaria, the sea urchin larva is called a pluteus, and larvae of asteroids, holothuroids, ophiuroids, and crinoids are known as bipinneria, auricularia, ophiopluteus, and doliolaria larvae, respectively. These various ciliated, feeding larvae often undergo dramatic morphological changes at metamorphosis to give rise to the adult. For example, the radially symmetrical, adult sea urchin is formed from a rudiment (the echinus rudiment) derived from the left coelomic pouch (hydrocoel) and the overlying ectoderm (vestibule) of the bilaterally symmetrical larva, while most of the other larval tissues degenerate (Burke, 1989). Metamorphosis in indirect developing enteropneust hemichordates results in less pronounced anatomical changes, and the body plan of the tornaria is more directly inherited by the adult (Lowe et al., Chapter 8). Indirect development was clearly the ancestral mode of development within the echinoderms (Nakano et al., 2003; Peterson et al., 1999; RaV, 1987), and morphological similarities between the larvae of echinoderms and hemichordates suggest that this was the ancestral mode within the echinoderm þ hemichordate clade. Direct development (i.e., the loss of a feeding larval stage) has arisen independently many times, however, within the group (RaV, 1987; Strathmann, 1978). In its
1. Invertebrate Deuterostomes
7
extreme form, direct development is associated with the formation of a late-stage embryo that has few morphological features of the feeding larva and develops rapidly and directly into a juvenile adult. As others have noted, however, many species of echinoderms show intermediate modes of development that challenge a simple ‘‘indirect’’ vs ‘‘direct’’ classification scheme (Chia et al., 1993; McEdward and Janies, 1997). These include larvae that are morphologically very similar to feeding larvae but do not feed (or feed facultatively) as well as various late embryonic forms with morphologies intermediate between those of extreme indirect and direct developing forms (Chia et al., 1993; RaV, 1987). Direct development has also arisen within the enteropneust hemichordates. Again, however, there are examples of intermediate forms, including species with swimming, tornaria-like larvae that do not feed (Barrington, 1965). A simple categorization of urochordates and cephalochordates as either direct or indirect developers is also somewhat problematic, given the specific characteristics of their larval forms. In cephalochordates, embryogenesis gives rise to a ciliated, pelagic larva that can swim and feed for many weeks prior to metamorphosis. In these organisms, the morphological changes at metamorphosis are considerably less pronounced than in echinoderms, however. They include the generation of a second row of gill slits, movement of gills slits and the mouth, and changes in the digestive tract (Holland and Yu, Chapter 9). Most ascidian species develop via a swimming tadpole larva and are considered to undergo indirect development. Ascidian tadpole larvae diVer from the larvae of other invertebrate deuterostomes, however, in that they are uniformly nonfeeding and swim by muscular contraction rather than ciliary beating. Ascidian larvae usually swim for only minutes or a few hours before settling and undergoing metamorphosis. Extensive anatomical changes take place at that time, including resorption of the tail, loss of the outer layer of tunic and some organs, and the repositioning of visceral organs (Satoh, 1994). This indirect mode of development is ancestral within the ascidians, but in so-called ‘‘tailless’’ or ‘‘aneural’’ species, it has given way to direct development, in which various features of the tadpole, such as diVerentiated tail muscle cells, notochord, and the neural sensory organ, have been lost (JeVery and Swalla, 1992; JeVery et al., 1999).
Fig. 3 Representative embryonic and larval stages of selected invertebrate deuterostomes. (A, E, I) Sea urchin (Lytechinus variegatus). (B, F, J) Enteropneust hemichordate (B, Saccoglossus kowalevskii; F, J, Ptychodera flava). (C, G, K) Ascidian (C, G, Ciona intestinalis; K, Boltenia villosa). (D, H, L) Amphioxus (Branchiostoma floridae). The upper panels (A–D) show early cleavage stage embryos (4–8 cell stage) and illustrate the holoblastic pattern of cleavage common to the group. The middle panels (E–H) show gastrula stage embryos. Endoderm and mesoderm are internalized through a roughly circular blastopore (arrowhead) positioned at the vegetal pole of the spherical gastrula. The lower panels (I–L) show larval stages (1–3 d larvae except for J, which is much older). There is considerable diversity in larval morphology within the group. Scale bars ¼ 50 m except for J (300 m). Photographs courtesy of Chris Lowe (B, J), Nori Satoh (C, G), Linda Holland (D, H, L), Jonathan Henry (F), Rachel Fink (I), and Billie Swalla (K).
8
Charles A. Ettensohn et al.
These variations in developmental modes and larval and adult morphologies among the extant, invertebrate deuterostomes raise several questions: Did the ancestral chordate and the ancestral deuterostome develop in an ‘‘indirect’’ fashion, via a free-swimming, feeding larva? If so, what did the ancestral larva look like? How diVerent was its anatomy from that of the adult? These questions are obviously diYcult to answer in light of the many independent evolutionary changes that have taken place in the various lineages since they first diverged. One hypothesis is that the ancestral deuterostome developed via a ciliated, pelagic, feeding larva that may have given rise to a sessile, wormlike adult. Garstang (1928) suggested that vertebrates arose through neoteny, the sexual maturation of a swimming larva that may have resembled the ascidian tadpole larva (see Barrington, 1965; Gee, 1996).
IV. Development A. Historical Contributions Experimental work with invertebrate deuterostomes has played an important role in developmental biology, cell biology, and genetics. Descriptive and experimental studies on the embryonic development of these organisms date back to the mid-nineteenth century. The earliest papers describing in vitro fertilization and embryonic development in sea urchins were published by Derbes and von Baer in 1847 (see Ho¨rstadius, 1973). The blastomere separation experiments of Chabry (1887), who worked with the ascidian Ciona intestinalis, are often considered the first work in the field of experimental embryology (cited in Fischer, 1992). These were soon followed by the pioneering studies of Driesch on sea urchin embryos, which led to the discovery of regulative development (Driesch, 1892). Other early work with sea urchins pointed to the central role of the nucleus in development and heredity (Hertwig, 1875) and revealed the nonequivalence of chromosomes (Boveri, 1907). Several decades later, the famous cell isolation and transplantation studies of Sven Ho¨rstadius provided early evidence of inductive interactions among blastomeres and helped define the notion of developmental gradients (reviewed by Ho¨rstadius, 1939). Early experimental studies on ascidian embryos were central in revealing cytoplasmic regionalization of the egg, the role of ooplasmic constituents in regulating cell fates, and the relationship between the egg and larval body axes (Castle, 1896; Conklin, 1905; van Beneden and Julin, 1884). Observations on hemichordate and cephalochordate embryos also date back to the mid-nineteenth century (Bateson, 1884; Kowalevsky, 1866; Mu¨ller, 1841). B. Experimental Characteristics The characteristics of invertebrate deuterostomes that attracted many early experimental embryologists remain extremely useful today. The ease with which gametes can be collected and the external fertilization and development of the
1. Invertebrate Deuterostomes
9
embryos originally made it possible to study their early development. Today, the ability to produce vast numbers of synchronously developing embryos (using methods that are best developed with sea urchins) facilitates a wide variety of biochemical and molecular biological approaches, from protein purification to cDNA library construction. The unsurpassed transparency of many invertebrate deuterostome eggs and embryos originally made it possible to directly observe developmental processes using only a light microscope. The clarity of these embryos now facilitates the application of a wide variety of modern light optical technologies, including fluorescence-based methods for monitoring gene expression, protein localization, protein–protein interactions, and biochemical activity. The embryos of many invertebrate deuterostomes are highly amenable to physical manipulations, including embryo dissociation, cell isolation, and cell transplantation. These approaches continue to be invaluable in analyzing cellular interactions that pattern the embryo. Molecular biological approaches including expression of exogenous mRNAs, injection of reporter DNA constructs for cisregulatory analysis, and injection of morpholino antisense oligonucleotides, make it possible to dissect gene regulation and function. The development of genomicsbased resources, including the assembly of draft genomic sequences of several organisms, is spurring work of all kinds (see Section V). C. Shared Features of Development Several features of early development are shared among the invertebrate deuterostomes, as might be expected from their close phylogenetic relationships. The eggs of invertebrate deuterostomes are isolecithal (i.e., they have sparse, evenly distributed yolk). They are usually 100 to 200 m in diameter, but can range up to 4 mm, with larger, more yolky eggs typical of direct-developing species (Chia et al., 1993; RaV, 1987). Primary oocytes typically undergo germinal vesicle breakdown shortly before spawning. Depending upon the organism, at the time of fertilization the oocyte may be in metaphase of the first meiotic division (starfish and ascidians), metaphase of the second meiotic division (amphioxus), or may have completed meiosis (sea urchins). The unfertilized egg is polarized along a single maternal axis; the animal–vegetal axis. In ascidians, other axes are established immediately after fertilization, as reflected by movements of the cytoplasm and sperm aster (Satoh, 1994). In sea urchins, a secondary axis (the oral–aboral axis) is probably entrained soon after fertilization, although the initial cues have not been identified (Duboc et al., 2004; Ettensohn and Sweet, 2000). Cleavage is almost always holoblastic (i.e., the cleavage furrows pass completely through the egg) and generally radial (Fig. 3A–D). Urochordates, however, exhibit bilateral cleavage, which produces an early embryo with mirror-image right and left halves. Early cleavages are also typically equal (or nearly equal), with the bestknown exception being the unequal divisions of the vegetal blastomeres of the 8-cell stage sea urchin embryo, which generate the micromeres. Gastrulation begins 5 to 10 hours after fertilization in warm water species, when the embryo consists of
10
Charles A. Ettensohn et al.
between 100 (ascidians) and 800 cells (amphioxus and sea urchins). As deuterostomes, all these organisms gastrulate through a vegetally positioned blastopore, which subsequently becomes the anus (Fig. 3E–H). The mouth opening forms secondarily, and body cavities form from mesodermal pouches that balloon outward from the primitive gut (enterocoely). Gastrulation involves a process of invagination, by which mesoderm and endoderm move inward through a circular blastopore. Typically, there is some involution of cells near the margins of the blastopore and limited epibolic spreading of animal cells. The animal region of the embryo (opposite the blastopore) invariably gives rise to the ectoderm and the mesoderm and endoderm are derived from the vegetal half of the egg. In amphioxus and ascidians, the cells at the vegetal pole give rise to endoderm, and the mesoderm arises from more equatorial cells. In echinoderms, however, the orientation of these two prospective territories is reversed. Embryonic development is often extremely rapid, particularly in warm water species, and the swimming, feeding larva may form in less than a day in some species. If the ascidian tadpole and sea urchin pluteus are representative of the group, the swimming larva contains 2000 to 3000 cells and some 10 to 15 basic cell types, including pigment cells, muscle cells, neurons, other mesenchyme cells, and gut cells.
V. Genomics Because of the utility of invertebrate deuterostomes for developmental analysis and their close phylogenetic relationship to vertebrates, genomic resources are being developed from a variety of these organisms. The draft genomic sequences of two invertebrate deuterostomes, the related ascidians C. intestinalis and C. savignyi, have been released (see Cameron et al., Chapter 30; Satoh, Chapter 31. The assembly of the C. intestinalis genome is considerably more complete, although in neither species is it yet possible to assign scaVolds of assembled genomic sequence to specific chromosomes. The draft genomic sequence of a sea urchin, Strongylocentrotus purpuratus, is nearing completion, with a release expected in late 2004. The genome of S. purpuratus is large (800 megabases, or about 25% the size of the human genome), while the genomes of C. intestinalis and C. savignyi are considerably smaller (150–180 megabases). Gene density in the ascidians is therefore likely to be higher. It seems almost assured that as these three initial genome sequencing projects are completed, eVorts will be made to sequence the complete genomes of other invertebrate deuterostomes. In addition to the whole genome sequencing projects, many other genomic resources are being developed. Vast EST databases have been generated for C. intestinalis, including EST collections from many developmental stages and several adult tissues (Satoh, Chapter 31). Considerable progress has also been made in this organism in determining full-length cDNA sequences, developing DNA microarrays, and in carrying out large-scale, whole mount in situ hybridization analyses to determine the spatial patterns of expressions of genes. In the sea
1. Invertebrate Deuterostomes
11
urchin, S. purpuratus, extensive EST databases have been developed for whole embryos of various developmental stages and for specific embryonic cell types (Cameron et al., Chapter 30; Poustka et al., 2003; Zhu et al., 2001). Robotically arrayed cDNA and genomic libraries have also been developed for a number of other sea urchin species and these are proving to be extremely useful for comparative studies. Computational tools have been developed that facilitate the use of these genomic resources; for example, the FamilyRelations program facilitates comparisons of genomic sequences for the purpose of identifying conserved regions that may function as cis-regulatory elements (Cameron et al., Chapter 30). A large EST database is currently being developed for the direct developing, enteropneust hemichordate, Saccoglossus kowalevskii (Lowe et al., 2003). A useful feature of invertebrate deuterostome genomes is that many genes appear to be present in single copies that are found in multiple copies in vertebrate genomes. A frequently cited example is the Hox gene cluster, which is found in four copies in mammals but in only one copy in cephalochordates and sea urchins (Arenas-Mena et al., 2000; Bailey et al., 1997). Models of genome evolution within the vertebrate lineage posit sequence duplications on scales ranging from individual genes to multiple duplications of entire genomes (Durand, 2003; Holland, 1999, 2003; Panopoulou et al., 2003; Wagner, 2001). The ‘‘simplified’’ nature of invertebrate deuterostome genomes may prove very useful in dissecting developmental pathways, gene networks, and the functions of conserved gene products that are more challenging to study in vertebrates because of the proliferation of the genes involved. Undoubtedly, analysis of the genomes of invertebrate deuterostomes will also reveal genes and pathways that are specific to particular lineages of deuterostomes. This may provide insight into the genetic changes that led to the diversification of the deuterostomes, including the innovations that gave rise to the vertebrates. The prospects are bright, indeed, that this group of organisms, which played such a central role in the emergence of developmental biology, will yield important and exciting new insights in the future.
References Adoutte, A., Balavoine, G., Lartillot, N., Lespinet, O., Prud’homme, B., and de Rosa, R. (2000). The new animal phylogeny: Reliability and implications. Proc. Natl. Acad. Sci. USA 97, 4453–4456. Aguinaldo, A. M., Turbeville, J. M., Linford, L. S., Rivera, M. C., Garey, J. R., RaV, R. A., and Lake, J. A. (1997). Evidence for a clade of nematodes, arthropods, and other moulting animals. Nature 387, 489–493. Arenas-Mena, C., Cameron, A. R., and Davidson, E. H. (2000). Spatial expression of Hox cluster genes in the ontogeny of a sea urchin. Development 127, 4631–4643. Bailey, W. J., Kim, J., Wagner, G. P., and Ruddle, F. H. (1997). Phylogenetic reconstruction of vertebrate Hox cluster duplications. Mol. Biol. Evol. 14, 843–853. Barrington, E. J. (1965). ‘‘The Biology of Hemichordata and Protochordata.’’ W. H. Freeman and Co., San Francisco. Bateson, W. (1884). The early stages in the development of Balanoglossus (sp. uncert.). Quart. J. Micros. Sci. 24, 209–235.
12
Charles A. Ettensohn et al. Bone, Q. (1998). ‘‘The Biology of Pelagic Tunicates.’’ Oxford University Press, Oxford, UK. Bosch, I., Rivkin, R. B., and Alexander, S. P. (1989). A sexual reproduction by oceanic planktotrophic echinoderm larvae. Nature 337, 169–170. Boveri, T. (1907). Zellenstudien VI: Die Entwicklung dispermer Seeigelier. Ein Beitrag zur Befruchtungslehre und zur Theorie des Kernes. Jena Zeit. Naturw. 43, 1–292. Bromham, L. D., and Degnan, B. M. (1999). Hemichordates and deuterostome evolution: Robust molecular phylogenetic support for a hemichordate þ echinoderm clade. Evol. Dev. 1, 166–171. Burke, R. D. (1989). Echinoderm metamorphosis: Comparative aspects of the change in form. In ‘‘Echinoderm Studies’’ (M. Jangoux and J. Lawrence, eds.), pp. 81–108. A. A. Balkema Pub., Rotterdam, Netherlands. Cameron, C. B., Garey, J. R., and Swalla, B. J. (2000). Evolution of the chordate body plan: New insights from phylogenetic analyses of deuterostome phyla. Proc. Natl. Acad. Sci. USA 97, 4469–4474. Castle, W. E. (1896). The early embryology of Ciona intestinalis Flemming (L.). Bull. Mus. Comp. Zool. 27, 203–280. Chia, F. S., Oguro, C., and Komatsu, M. (1993). Sea-star (asteroid) development. Oceanogr. Mar. Biol. Annu. Rev. 31, 223–257. Conklin, E. G. (1905). The organization and cell lineage of the ascidian egg. J. Acad. Nat. Sci. Phila. 13, 1–119. Driesch, H. (1892). The potency of the first two cleavage cells in echinoderm development. Experimental production of partial and double formations. In ‘‘Foundations of Experimental Embryology’’ (B. H. Willier and J. M. Oppenheimer, eds.). Hafner, New York, 1974. Duboc, V., Rottinger, E., Besnardeau, L., and Lepage, T. (2004). Nodal and BMP2/4 signaling organizes the oral–aboral axis of the sea urchin embryo. Dev. Cell. 6, 397–410. Durand, D. (2003). Vertebrate evolution: Doubling and shuZing with a full deck. Trends Genet. 19, 2–5. Eaves, A. A., and Palmer, A. R. (2003). Reproduction: Widespread cloning in echinoderm larvae. Nature 425, 146. Ettensohn, C. A., and Sweet, H. (2000). Patterning the early sea urchin embryo. Curr. Top. Dev. Biol. 50, 1–44. Fischer, J. L. (1992). The embryological oeuvre of Laurent Chabry. Roux’s Arch. Dev. Biol. 201, 125–127. Garstang, W. J. (1928). The morphology of the Tunicata, and its bearing on the phylogeny of the Chordata. Quart. J. Micros. Sci. 72, 51–187. Gee, H. (1996). ‘‘Before the Backbone: Views on the Origins of the Vertebrates.’’ Chapman and Hall, London, UK. Gross, J. M., and McClay, D. R. (2001). The role of Brachyury (T) during gastrulation movements in the sea urchin Lytechinus variegatus. Dev. Biol. 239, 132–147. Harada, Y., Shoguchi, E., Taguchi, S., Okai, N., Humphreys, T., Tagawa, K., and Satoh, N. (2002). Conserved expression pattern of BMP-2/4 in hemichordate acorn worm and echinoderm sea cucumber embryos. Zoolog. Sci. 19, 1113–1121. Hertwig, O. (1875). Beitra¨ge zur Kenntnis der Bildung, Befruchtung, und Teilung des tierischen Eies, I. Morphologische Jahrbuch 1. Holland, P. W. (1999). Gene duplication: Past, present, and future. Semin. Cell Dev. Biol. 10, 541–547. Holland, P. W. (2003). More genes in vertebrates? J. Struct. Funct. Genomics 3, 75–84. Ho¨rstadius, S. (1939). The mechanics of sea urchin development, studied by operative methods. Biol. Rev. 14, 132–179. Ho¨rstadius, S. (1973). ‘‘Experimental Embryology of Echinoderms.’’ Clarendon Press, Oxford, UK. JeVery, W. R., and Swalla, B. J. (1992). Evolution of alternate modes of development in ascidians. Bioessays 14, 219–226. JeVery, W. R., Swalla, B. J., Ewing, N., and Kusakabe, T. (1999). Evolution of the ascidian anural larva: Evidence from embryos and molecules. Mol. Biol. Evol. 16, 646–654.
1. Invertebrate Deuterostomes
13
Kowalevsky, A. (1866). Entwicklungsgeschichte des Amphioxus lanceolatus. Me´moires de l’Acade´mie des Sciences de St. Pe´tersbourg 7(11), No. 4, 1–17. Lowe, C. J., Wu, M., Salic, A., Evans, L., Lander, E., Stange-Thomann, N., Gruber, C. E., Gerhart, J., and Kirschner, M. (2003). Anteroposterior patterning in hemichordates and the origins of the chordate nervous system. Cell 113, 853–865. McEdward, L. R., and Janies, D. A. (1997). Relationships among development, ecology, and morphology in the evolution of echinoderm larvae and life cycles. Biol. J. Linnean Soc. 60, 381–400. McGovern, T. M. (2003). Plastic reproductive strategies in a clonal marine invertebrate. Proc. R. Soc. Lond. B Biol. Sci. 270, 2517–2522. Mu¨ller, J. (1841). Mikroskopische Untersuchungen u¨ber den Bau und die Lebenserscheinungen des Branchiostoma lubricum Costa, Amphioxus lanceolatus Yarrell, pp. 396–411. Ber. Preuss. Akad. Wissensch. Berlin, Germany. Nakajima, Y., Humphreys, T., Kaneko, H., and Tagawa, K. (2004). Development and neural organization of the tornaria larva of the Hawaiian hemichordate, Ptychodera flava. Zoolog. Sci. 21, 69–78. Nakano, H., Hibino, T., Oji, T., Hara, Y., and Amemiya, S. (2003). Larval stages of a living sea lily (stalked crinoid echinoderm). Nature 421, 158–160. Panopoulou, G., Hennig, S., Groth, D., Krause, A., Poustka, A. J., Herwig, R., Vingron, M., and Lehrach, H. (2003). New evidence for genome-wide duplications at the origin of vertebrates using an amphioxus gene set and completed animal genomes. Genome Res. 13, 1056–1066. Peterson, K. J., Harada, Y., Cameron, R. A., and Davidson, E. H. (1999). Expression pattern of Brachyury and Not in the sea urchin: Comparative implications for the origins of mesoderm in the basal deuterostomes. Dev. Biol. 207, 419–431. Poustka, A. J., Groth, D., Hennig, S., Thamm, S., Cameron, A., Beck, A., Reinhardt, R., Herwig, R., Panopoulou, G., and Lehrach, H. (2003). Generation, annotation, evolutionary analysis, and database integration of 20,000 unique sea urchin EST clusters. Genome Res. 13, 2736–2746. RaV, R. A. (1987). Constraint, flexibility, and phylogenetic history in the evolution of direct development in sea urchins. Dev. Biol. 119, 6–19. Satoh, N. (1994). ‘‘Developmental Biology of Ascidians.’’ Cambridge University Press. Shoguchi, E., Satoh, N., and Maruyama, Y. K. (1999). Pattern of Brachyury gene expression in starfish embryos resembles that of hemichordate embryos but not of sea urchin embryos. Mech. Dev. 82, 185–189. Strathmann, R. R. (1978). The evolution and loss of feeding larval stages of marine invertebrates. Evolution 32, 894–906. Swalla, B. J., Cameron, C. B., Corley, L. S., and Garey, J. R. (2000). Urochordates are monophyletic within the deuterostomes. Syst. Biol. 49, 52–64. Tagawa, K., Humphreys, T., and Satoh, N. (1998). Novel pattern of Brachyury gene expression in hemichordate embryos. Mech. Dev. 75, 139–143. van Beneden, E., and Julin, C. H. (1884). La segmentation chez les ascidens dans ses rapportes avec l’organization de la larve. Arch. Biol. 5, 111–126. Wagner, A. (2001). Birth and death of duplicated genes in completely sequenced eukaryotes. Trends Genet. 17, 237–239. Zhu, X., Mahairas, G., Illies, M., Cameron, R. A., Davidson, E. H., and Ettensohn, C. A. (2001). A large-scale analysis of mRNAs expressed by primary mesenchyme cells of the sea urchin embryo. Development 128, 2615–2627.
This Page Intentionally Left Blank
PART I
Procurement, Maintenance & Culture of Oocytes, Embryos, Larvae, and Adults
This Page Intentionally Left Blank
CHAPTER 2
Care and Maintenance of Adult Echinoderms S. Anne Bo¨ttger,* Charles W. Walker,* and Tatsuya Unuma{ *Department of Zoology and Marine Biology Biomedical Research Group Durham, New Hampshire 03824 {
National Research Institute of Aquaculture Nansei, Mie 516-0193
I. Overview II. Introduction III. Adult Echinoderm Models: Their Reproductive Cycles and Gametogenesis A. Sea Urchin Models from North and South America, Europe, and Japan B. Sea Star Models from North America, Europe, and Japan C. Echinoderm Gametogenesis in Plastic Sections for Light and Electron Microscopy IV. Obtaining Adult Echinoderms A. Collection and Transport of Adult Echinoderms B. Commercial Suppliers V. Maintenance of Adult Echinoderms in Land-Based Systems A. Culture in Small Aquaria B. Large-Scale, Land-Based Facilities VI. Care and Handling of Adult Echinoderms A. Physical and Chemical Factors B. Biological Factors References
METHODS IN CELL BIOLOGY, VOL. 74 Copyright 2004, Elsevier Inc. All rights reserved. 0091-679X/04 $35.00
17
18
S. Anne Bo¨ttger et al.
I. Overview This chapter addresses the care and maintenance of adult echinoderms that produce gametes and embryos commonly used for molecular, cellular, and developmental biology (see Chapter 1). Emphasis is placed on sea urchin and sea star species from North and South America, Europe, and Japan. Sea cucumbers, brittle stars, and crinoids are not discussed because they are less amenable to fertilization in the laboratory and are thus less widely used for experimental manipulation of their eggs and embryos. Specific topics addressed in this chapter are: (a) adult echinoderm models: their reproductive cycles and gametogenesis; (b) methods for obtaining adult echinoderms; (c) maintenance of adult echinoderms in land-based systems; and (d) methods for care and handling of adult echinoderms.
II. Introduction Sea urchins and sea stars have been employed by generations of molecular, cellular, and developmental biologists as models in studies of gametogenesis, egg and sperm interaction and activation, fertilization, and early development (see Chapter 1). Echinoderms that produce large numbers of gametes do so using annual cycles of reproduction in which the germinal epithelium may contain amitotic germ-line stem cells (oogonia or spermatogonia) for one to several months. This period is followed by a gametogenic phase lasting several months during which germ-line stem cells initiate mitosis, increase in numbers in both sexes to form resulting gonocytes (primary and secondary oocytes or spermatocytes) that complete gametogenesis, and produce gametes (fully mature ova in sea urchins, primary oocytes in sea stars, and spermatozoa in both). Depending upon the stage of gametogenesis and the degree of spawning, gonads in these echinoderms may contain few or copious numbers of gametes. An understanding of the cell biology of gametogenesis in sea urchins and sea stars is vital in determining when one can expect to obtain ‘‘ripe’’ eggs (fully mature ova in female sea urchins and primary oocytes in female sea stars) that can be successfully fertilized to yield embryos for experimentation. An understanding of gametogenesis in these two echinoderm groups can be matched with available data on the reproductive cycles for particular species in diVerent parts of the world to determine when each will be most useful in experimental applications (Tables I and II).
III. Adult Echinoderm Models: Their Reproductive Cycles and Gametogenesis Annually, within the gonads of both sexes of most echinoderms, millions of gonial cells (oogonia and spermatogonia) originate by mitosis, produce gonocytes (primary and secondary) that undergo gametogenesis and result in gametes that
19
2. Care of Echinoderms
Table I Summary of Species Information Relating to Collection of Echinoids (Sea Urchins) and Their Breeding Seasona Species
Size
Depth
Substrate
Anthocidaris crassispina Arbacia punctulata
