Medaka biology, management, and experimental protocols [Second edition] 9781119575344, 1119575346, 9781119575306, 9781119575290

Table of contents :
Content: List of Contributors xvPreface xxi1 Medaka Management 11.1 Introduction 11.2 Medaka Management for Scientific Research 11.2.1 Outline of medaka life?cycle in the wild 21.2.2 Preparation of normal rearing conditions of medaka in the laboratory and procedures for breeding 21.2.2.1 Breeding system set?up 21.2.2.2 Obtaining medaka 31.2.2.3 Collecting eggs in a laboratory setting 31.2.2.4 Daily care and maintenance of eggs 41.2.2.5 Rearing medaka from the larval stage to adulthood 41.2.2.6 Anesthesia and euthanasia 41.3 Standardized Culture and Growth Curve 71.3.1 Characteristics and selection of strains 71.3.2 Management of medaka eggs and fish 81.3.2.1 Mating 81.3.2.2 Management of embryos 81.3.2.3 Management of embryos before hatching 131.3.2.4 Rearing from the larval stage to adulthood (to induce earlier maturation) 141.3.3 Maintenance of breeding tanks during breeding 231.3.3.1 Judgment of water quality 231.3.3.2 Maintenance of breeding water 241.3.4 Anesthesia 251.3.4.1 Behavior under each anesthesia stage 261.3.4.2 Difference in sensitivity to anesthesia among strains 261.3.4.3 Growth stage specificity in sensitivity to MS?222 271.3.4.4 Eugenol is recommended as an anesthetic reagent 281.3.4.5 Euthanasia 281.3.4.6 Important reminders for euthanasia 292 Medaka and Oryzias Species as Model Organisms and the Current Status of Medaka Biological Resources 312.1 Introduction 312.2 Common and Unique Futures of Medaka and Related Species as Model Organisms 312.3 Phylogenetic Relationships of Medaka and Related Species 352.3.1 The javanicus species group 352.3.2 The latipes species group 402.3.3 The celebensis species group 422.4 BAC Resources of Species Related to Medaka 432.5 National Bio?Resource Project Medaka (NBRP Medaka) 432.5.1 Support for visiting researchers 453 Looking at Adult Medaka 493.1 General Morphology 493.1.1 Secondary sexual characters 493.1.1.1 Dorsal fin 493.1.1.2 Anal fin 493.1.1.3 Papillar processes 503.1.1.4 Urogenital papillae 503.1.2 Body color 513.1.2.1 Pigment cells (chromatophores) 513.1.2.2 Structures of the chromatophores 513.1.2.3 Chromatophores in medaka 513.1.2.4 Chromatophore distribution in medaka 553.1.2.5 See?through medaka 563.2 Anatomy and Histology 563.2.1 Observations of internal organs 563.2.1.1 Observations of internal organs in the live see?through medaka 563.2.1.2 Dissection of adult medaka 583.2.2 Horizontal and sagittal sections of juvenile medaka 583.2.3 Nervous system 583.2.3.1 Adult central nervous system 583.2.3.2 Adult peripheral nervous system 673.2.4 Endocrine system 743.2.4.1 Hypothalamo?pituitary system 763.2.4.2 Pineal organ (epiphysis) 783.2.4.3 Thyroid gland 793.2.4.4 Heart 813.2.4.5 Interrenal gland and chromaffin cells 813.2.4.6 Gonads 813.2.4.7 Endocrine pancreas (islets of Langerhans) 813.2.4.8 Gastrointestinal tract 813.2.4.9 Ultimobranchial gland 823.2.4.10 Corpuscle of Stannius 823.2.4.11 Urophysis 833.2.4.12 Thymus 833.2.5 Gonads 833.2.5.1 Ovary 833.2.5.2 Testis 853.2.6 Kidney 853.2.6.1 Pronephros 863.2.6.2 Mesonephros 863.2.6.3 Histology of the kidney 86Column 3.1 How to make sections of a mature ovary for histological analysis 884 Looking at Medaka Embryos 974.1 Development of Various Tissues and Organs 974.1.1 Developmental stages 974.1.1.1 Stage 0: unfertilized egg (Figure 4-1) 974.1.1.2 Stage 1: activated egg stage (3 minutes) (Figure 4-1) 994.1.1.3 Stage 2: blastodisc stage (Figure 4-1) 994.1.1.4 Stage 3: two?cell stage (1 hour 5 minutes) (Figure 4-1) 994.1.1.5 Stage 4: four?cell stage (1 hour 45 minutes) (Figure 4-1) 1004.1.1.6 Stage 5: eight?cell stage (2 hours 20 minutes) (Figure 4-1) 1004.1.1.7 Stage 6: 16?cell stage (2 hours 55 minutes) (Figure 4-2) 1004.1.1.8 Stage 7: 32?cell stage (3 hours 30 minutes) (Figure 4-2) 1004.1.1.9 Stage 8: early morula stage (4 hours 5 minutes) (Figure 4-2) 1004.1.1.10 Stage 9: late morula stage (5 hours 15 minutes) (Figure 4-2) 1004.1.1.11 Stage 10: early blastula stage (6 hours 30 minutes) (Figure 4-2) 1004.1.1.12 Stage 11: late blastula stage (8 hours 15 minutes) (Figure 4-2) 1024.1.1.13 Stage 12: pre?early gastrula stage (10 hours 20 minutes) (Figure 4-3) 1024.1.1.14 Stage 13: early gastrula stage (13 hours) (Figure 4-3) 1024.1.1.15 Stage 14: pre?mid?gastrula stage (15 hours) (Figure 4-3) 1024.1.1.16 Stage 15: mid?gastrula stage (17 hours 30 minutes) (Figure 4-3) 1024.1.1.17 Stage 16: late gastrula stage (21 hours) (Figure 4-3) 1024.1.1.18 Stage 17: early neurula stage (1 day 1 hour) (Figure 4-3) 1034.1.1.19 Stage 18: late neurula stage (1 day 2 hours) (Figure 4-4) 1044.1.1.20 Stage 19: two?somite stage (1 day 3 hours 30 minutes) (Figure 4-4) 1044.1.1.21 Stage 20: four?somite stage (1 day 7 hours 30 minutes) (Figure 4-4) 1044.1.1.22 Stage 21: six?somite stage (1 day 10 hours) (Figure 4-4) 1044.1.1.23 Stage 22: nine?somite stage (1 day 14 hours) (Figure 4-4) 1044.1.1.24 Stage 23: 12?somite stage (1 day 17 hours) (Figure 4-4) 1044.1.1.25 Stage 24: 16?somite stage (1 day 20 hours) (Figure 4-5) 1064.1.1.26 Stage 25: 18-19?somite stage (2 days 2 hours) (Figure 4-5) 1074.1.1.27 Stage 26: 22?somite stage (2 days 6 hours) (Figure 4-5) 1074.1.1.28 Stage 27: 24?somite stage (2 days 10 hours) (Figure 4-5) 1074.1.1.29 Stage 28: 30?somite stage (2 days 16 hours) (Figure 4-5) 1074.1.1.30 Stage 29: 34?somite stage (3 days 2 hours) (Figure 4-5) 1084.1.1.31 Stage 30: 35?somite stage (3 days 10 hours) (Figure 4-6) 1084.1.1.32 Stage 31: gill blood vessel formation stage (3 days 23 hours) (Figure 4-6) 1084.1.1.33 Stage 32: somite completion stage (4 days 5 hours) (Figure 4-6) 1084.1.1.34 Stage 33: stage at which notochord vacuolization is completed (4 days 10 hours) (Figure 4-6) 1084.1.1.35 Stage 34: pectoral fin blood circulation stage (5 days 1 hour) (Figure 4-6) 1104.1.1.36 Stage 35: stage at which visceral blood vessels form (5 days 12 hours) (Figure 4-6) 1104.1.1.37 Stage 36: heart development stage (6 days) (Figure 4-7) 1104.1.1.38 Stage 37: pericardial cavity formation stage (7 days) (Figure 4-7) 1104.1.1.39 Stage 38: spleen development stage (8 days) (Figure 4-7) 1104.1.1.40 Stage 39: hatching stage (9 days) (Figure 4-7) 1104.1.1.41 Stage 40: first larval stage (Figure 4-8) 1124.1.1.42 Stage 41: second larval stage (Figure 4-8) 1134.1.1.43 Stage 42: third larval stage (Figure 4-8) 1134.1.1.44 Stage 43: first juvenile stage (Figure 4-8) 1134.1.1.45 Stage 44: second juvenile stage (Figure 4-8) 1134.1.1.46 Stage 45 (Figure 4-8) 1134.1.2 Brain 1134.1.2.1 Gastrula step (stages 13-17) 1144.1.2.2 Neurula step (stages 17-18) 1144.1.2.3 Neural rod step (stages 19-22) 1164.1.2.4 Neural tube step (stages 23-27) 1164.1.2.5 Late embryonic brain step (stages 28-34) 1174.1.2.6 Larval brain step (stages 35-42) 1194.1.3 Hatching gland 1194.1.3.1 Origin of fish hatching gland cells 1194.1.3.2 Secretion of hatching enzymes from hatching gland cells 1224.1.4 Eye development 1244.1.4.1 Specification of the anterior neural plate 1244.1.4.2 Eye field determination and establishment of retinal identity 1254.1.4.3 Splitting of the retinal anlage into two retinal primordia 1254.1.4.4 Morphogenesis I: evagination of the optic vesicle 1254.1.4.5 Morphogenesis II: formation of the optic cup 1274.1.4.6 Retinal differentiation I: central retina 1274.1.4.7 Retinal differentiation II: Ciliary Marginal Zone 1274.1.4.8 Retinotectal projection 1284.1.5 Branchial arch and jaws 1284.1.5.1 Skeletal development 1284.1.5.2 Muscle development 1304.1.6 Vasculature 1314.1.6.1 Vascular anatomy of the developing medaka 1314.1.6.2 Origin of the medaka endothelial lineage 1424.1.6.3 Abbreviations 1424.1.6.4 Acknowledgment 1434.1.7 Blood cells (hematopoiesis) 1434.1.7.1 Overview 1434.1.7.2 Observation of embryonic and adult blood cells 1444.1.8 Heart 1464.1.8.1 Overview 1464.1.8.2 Heart architecture 1464.1.8.3 Heart morphogenesis 1474.1.8.4 Observation of the developing heart 1564.1.9 Kidney 1594.1.9.1 Overview 1594.1.9.2 Nephrogenesis 1594.1.9.3 Pronephros 1604.1.9.4 Mesonephros 1604.1.10 Thymus 1604.1.10.1 Overview 1604.1.10.2 Early development of the thymus 1604.1.10.3 Cortex and medulla 1614.1.10.4 Involution of the thymus 1624.1.11 Gut and liver 1624.1.12 Bones 1644.1.12.1 Vertebral column 164Column 4.1 Key words in bone formation 1724.1.13 Fins 1734.1.13.1 Overview 1734.1.13.2 Fin anatomy 1734.1.13.3 Embryonic fin development (from fertilization to stage 39 [hatching stage]) 1744.1.13.4 Fin development after hatching (after stage 39) 1754.1.13.5 Gene expression during fin development 1754.1.14 Gonads 1764.1.14.1 Overview 1764.1.14.2 PGC specification 1774.1.14.3 Formation of gonadal primordium (Figure 4-60b) 1774.1.14.4 Sexual dimorphism in germ cell proliferation (Figure 4-61) 1794.1.14.5 Posthatching period in XX gonads 1804.1.14.6 Posthatching period in XY gonads 1804.2 Medaka EGG Envelope and Hatching Enzyme 1814.2.1 Overview 1814.2.2 Preparation of a hatching enzyme solution from hatching liquid 1824.2.2.1 Procedure 1824.2.3 Simple method for preparing hatching enzyme solution 1834.2.3.1 Procedure 1834.2.4 Solubilization of the egg envelope using hatching enzyme 183Column 4.2 Easy method for preparation of a small amount of hatching enzyme solution (see website for figure) 1844.3 Observation of Embryos (Embedding Embryos) 1854.3.1 Anesthesia of embryos using MS?222 1854.3.1.1 Equipment and reagents 1854.3.2 Observation of embryos (mounting) 1854.3.2.1 Living embryos 1854.3.2.2 Processed embryos 1884.4 Whole?Mount In Situ Hybridization (see section 4.1.8.4 for a similar protocol) 1894.4.1 Fixation and storage 1894.4.1.1 Procedure 1 1894.4.2 Rehydration, proteinase K treatment, and postfixation at RT 1904.4.2.1 Procedure 2 1904.4.3 Hybridization and washing 1904.4.3.1 Procedure 3 1904.4.4 Immunoreaction and washing antibodies 1914.4.4.1 Procedure 4 1914.4.5 Staining 1914.4.5.1 Procedure 5 1914.5 Embedding in a Plastic Resin (Technovit 7100) 1924.5.1 Equipment and reagents 1924.5.2 Agarose mounting (Figure 4-68) 1924.5.2.1 Procedure 1 1924.5.3 Dehydration and infiltration (Figure 4-68) 1924.5.3.1 Procedure 2 1924.5.4 Polymerization (Figure 4-68) 1934.5.4.1 Procedure 3 193Column 4.3 Pigment Cells (Figure 4-69) 194Column 4.4 Kupffer's Vesicle 1955 Reproductive Behavior of Wild Japanese Medaka 2055.1 Wild Japanese Medaka 2055.2 Reproductive Behavior of Wild Medaka 2065.2.1 Aggressive behavior 2075.2.2 Spawning behavior 2075.2.3 Egg deposition behavior 2105.2.4 Egg discarding behavior 2105.2.5 School and aggregation 2115.3 Conclusion 2116 Cryopreservation and Transplantation of Medaka Germ Cells 2156.1 Introduction 2156.2 Cryopreservation of Medaka Testes 2156.2.1 Solutions 2166.2.2 Materials 2176.2.3 Procedures 2176.3 Transplantation of Thawed Testicular Cells into Recipient Larvae 2186.3.1 Solutions 2186.3.2 Materials 2196.3.3 Procedures 219Column 6.1 Production of triploid medaka 2227 Genome Editing 2257.1 Introduction 2257.2 Outline of Targeted Genome Editing Using Nucleases 2257.3 Preparation of CRISPR/Cas9 Genome Editing Tools 2267.3.1 Materials 2277.3.2 Production of custom?designed sgRNA 2287.3.2.1 Preparation of the bsai?digested sgRNA backbone 2287.3.2.2 Design and production of customized sgRNA 2287.3.3 Production of capped RNA encoding a Cas9 nuclease 2327.4 Preparation of Custom?Designed TALENs 2347.4.1 Materials 2347.4.2 Preparation of the TALEN assembly system 2367.4.2.1 Preparation of TAL modules (HD1?6, NG1?6, NI1?6, and NN1?6) 2367.4.2.2 Preparation of array backbone plasmids (pFUS vectors) 2367.4.2.3 Preparation of last repeat modules (LR?HD, NG, NI, and NN) 2377.4.2.4 Preparation of TALEN backbone plasmids (pCS2TAL3DD and RR vectors) 2387.4.3 Design and construction of custom?designed TALENs 2397.4.3.1 Design of TALEN using TALE?NT 2397.4.3.2 First assembly: construction of 6?modules array vectors 2407.4.3.3 Second assembly: construction of TALEN expression vectors 2427.5 Heteroduplex Mobility Assay - A Simple Method to Detect Targeted Genome Modification 2447.5.1 Materials 2467.5.2 Procedure 2467.5.2.1 Identification of the wild type, heterozygotes, and homozygotes 2467.5.2.2 Evaluation of the efficiency of targeted genome modifications 2467.6 How to Establish Gene Knock?out Strains 2477.6.1 Design and synthesis of genome?editing tools 2477.6.2 Evaluation of genome?editing activity with fertilized medaka eggs 2477.6.3 Microinjection of the selected genome?editing tool(s) 2487.6.4 Selection of founder fish by genotyping F1 embryos 2497.6.5 Selection of F1 fish carrying the same mutation and the establishment of mutant strain 2497.6.6 Selection of homozygous mutant fish in the F2 family 2517.7 How to Establish Gene Knock?in Strains 2527.7.1 Design and synthesis of CRISPR/Cas9 components 2537.7.2 Evaluation of genome?editing activity with fertilized medaka eggs 2537.7.3 Construction of donor plasmid with homology arms (Ca. 0.5 kbp) and bait sequences 2537.7.4 Microinjection for establishing knock?in strains 2547.7.5 Selecting G0 founders harboring the insert gene in the genomic target site 254Column 7.1 Utilization of crRNA, tracrRNA, Cas9 Protein 2557.A Simple Genomic DNA Preparation by an Alkaline Lysis Method 2567.A.1 Materials 2567.A.2 Procedure 2568 Photo?Inducible Gene Expression in Medaka 2618.1 Outline of IR?LEGO 2618.2 Practical Strategies of IR?LEGO in Medaka Study 2628.2.1 Selection of heat shock promoters and application studies 2628.3 Laser Irradiation Conditions and Sample Preparation 2658.4 Caution in Maintaining Strains 2678.5 Other Uses of IR?LEGO 2678.6 Summary and Future Prospects 2689 Screening and Testing Methods of Endocrine?Disrupting Chemicals Using Medaka 2719.1 Applied Toxicity Tests for Endocrine Disruptors 2719.2 Detection of Androgenic and Antiandrogenic Chemicals Using Medaka 2759.2.1 The formation of papillary processes on anal fin rays as an indicative phenotype for exposure of androgenic and/or antiandrogenic chemicals 2759.2.2 Candidate biomarkers for assessing the action of androgenic and antiandrogenic chemicals 2769.2.3 Visualization of androgenic and antiandrogenic activity as green fluorescence with spiggin?GFP medaka 27610 Application of the Seawater Medaka Oryzias melastigma (McClelland) for Marine Ecotoxicology 28110.1 Background and Development of Oryzias melastigma for Marine Ecotoxicology 28110.2 Marine Medaka Developmental Staging 28310.3 Standard Breeding and Rearing Conditions 28410.3.1 Seawater 28510.3.2 Temperature 28510.3.3 Photoperiod 28510.3.4 Feeding 28610.3.5 Embryo collection and rearing 28610.3.6 Hatching and larvae collection 28710.3.7 Larvae rearing 28810.3.8 Larvae feeding 28810.4 Raising Marine Medaka for Experimental Use 28910.4.1 Experiments using adult fish 28910.4.2 Experiments using larvae 28910.5 Troubleshooting 28910.5.1 Mass mortality 28910.5.2 Low egg production 28910.5.3 Extensive algal growth 29010.6 How to Obtain Marine Medaka O. melastigma 29010.7 Experimental Protocols Using Marine Medaka 29010.8 Immunotoxicity Assessment: Bacteria Challenge Assays 29010.8.1 SOP for adult bacterial challenge assay 29110.8.2 SOP for larval bacterial challenge assay 29210.8.3 Age selection for larval bacterial challenge 29310.9 Fish Dissection and the Whole Adult Histoarray 29410.9.1 SOP for fish dissection 29510.9.2 SOP for adult medaka histoarray 29510.10 Embryo Chip 29710.10.1 SOP for embryo and larvae histoarray 29710.A Materials for SOP for Adult Medaka Histoarray (see section 10.9.2) 29911 Telomerase and Telomere Biology in Medaka 30311.1 Introduction 30311.2 SOP for Quantification of Telomerase Activity Using the Real?Time Quantitative Telomeric Repeat Amplification Protocol (RTQ?TRAP) 30811.2.1 Procedures for sample extraction 30811.2.2 Procedures for determination of protein concentration 30811.2.3 Procedures for RTQ?TRAP linearity test 30811.2.4 Calculation of telomerase activity 30911.3 SOP for Quantification of Telomere Length Using Southern Blotting Analysis 30911.3.1 Procedures for genomic DNA extraction and digestion with restriction enzymes 30911.3.2 Procedures for probe preparation 31111.3.3 Procedures for electrophoresis and southern blotting 31111.3.4 Procedures for hybridization and detection 31211.3.5 Procedures for computerized telomere analysis 31211.4 SOP for Quantification of Telomere Length Using Fluorescence In Situ Hybridization 31311.4.1 Procedures for fluorescence in situ hybridization 31311.4.2 Procedures for confocal microscopy detection 31311.4.3 Procedures for ImageJ analysis 31412 Assessments of Medaka Skeletal Toxicity 31712.1 Introduction 31712.2 Methods 31812.2.1 Embryonic exposures: dioxin 31912.2.2 Embryonic exposure: dithiocarbamates 31912.2.3 Whole?mount alcian blue staining of hatchlings/larvaea 32012.2.4 Whole?mount Alizarin red S staining of hatchlings/larvaec 32012.2.5 In vivo Alizarin complexone fluorescent staining for mineralized bone matrix 32112.2.6 In vivo calcein fluorescent staining for mineralized bone matrix 32112.2.7 Confocal imaging of embryo/hatchling medaka 32112.2.8 Morphological assessments 32312.3 Results and Discussion 32412.3.1 Dithiocarbamates 32412.3.2 Dioxin 325Appendix A Solutions 329Attributions 331Index 335

Citation preview

Medaka

­Medaka Biology, Management, and Experimental Protocols Volume 2 Edited by

Chief Editors Kenji Murata University of California, Davis, CA, USA

Masato Kinoshita Kyoto University, Sakyo-ku, Kyoto, Japan

Kiyoshi Naruse National Institute for Basic Biology, Okazaki, Aichi, Japan

Editors Minoru Tanaka Nagoya University, Nagoya, Aichi, Japan

Yasuhiro Kamei National Institute for Basic Biology, Okazaki, Aichi, Japan

This edition first published 2020 © 2020 John Wiley & Sons Ltd All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/ go/permissions. The right of Kenji Murata, Masato Kinoshita, Yasuhiro Kamei, Minoru Tanaka and Kiyoshi Naruse to be identified as the authors of the editorial material in this work has been asserted in accordance with law. Registered Offices John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Editorial Office The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print-on-demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Cataloging-in-Publication Data Names: Murata, Kenji, 1961- editor. | Kinoshita, Masato, editor. Title: Medaka : biology, management, and experimental protocols / edited by Chief editors, Kenji Murata, University of California Davis, CA, USA, Masato Kinoshita, Kyoto University, Sakyo-ku, Kyoto, Japan ; editors, Yasuhiro Kamei, National Institute for Basic Biology, Okazaki, Aichi, Japan, Minoru Tanaka, Nagoya Uiversity, Nagoya, Aichi, Japan, Kiyoshi Naruse, National Institute for Basic Biology, Okazaki, Aichi, Japan. Description: Second edition. | Hoboken, NJ : Wiley-Blackwell, [2020] | Includes bibliographical references and index. | Identifiers: LCCN 2019011589 (print) | LCCN 2019011759 (ebook) | ISBN 9781119575344 (Adobe PDF) | ISBN 9781119575306 (ePub) | ISBN 9781119575290 (hardcover) Subjects: LCSH: Oryzias latipes. Classification: LCC QL638.O78 (ebook) | LCC QL638.O78 M43 2020 (print) | DDC 597.53–dc23 LC record available at https://lccn.loc.gov/2019011589 Cover Design: Wiley Cover Images: courtesy of Kiyoshi Naruse. Medaka with rainbow color image courtesy of Lazaro Centanin and Jochen Wittbrodt Set in 10/12pt Times LT Std by SPi Global, Chennai, India 10 9 8 7 6 5 4 3 2 1

Contents List of Contributors xv Preface xxi 1  Medaka Management 1.1 ­Introduction 1.2 ­Medaka Management for Scientific Research 1.2.1  Outline of medaka life‐cycle in the wild 1.2.2  Preparation of normal rearing conditions of medaka in  the laboratory and procedures for breeding 1.2.2.1  Breeding system set‐up 1.2.2.2  Obtaining medaka 1.2.2.3  Collecting eggs in a laboratory setting 1.2.2.4  Daily care and maintenance of eggs 1.2.2.5  Rearing medaka from the larval stage to adulthood 1.2.2.6  Anesthesia and euthanasia 1.3 ­Standardized Culture and Growth Curve 1.3.1  Characteristics and selection of strains 1.3.2  Management of medaka eggs and fish 1.3.2.1 Mating 1.3.2.2  Management of embryos 1.3.2.3  Management of embryos before hatching 1.3.2.4  Rearing from the larval stage to adulthood (to induce earlier maturation) 1.3.3  Maintenance of breeding tanks during breeding 1.3.3.1  Judgment of water quality 1.3.3.2  Maintenance of breeding water 1.3.4 Anesthesia 1.3.4.1  Behavior under each anesthesia stage 1.3.4.2  Difference in sensitivity to anesthesia among strains 1.3.4.3  Growth stage specificity in sensitivity to MS‐222 1.3.4.4  Eugenol is recommended as an anesthetic reagent 1.3.4.5 Euthanasia 1.3.4.6  Important reminders for euthanasia

1 1 1 2 2 2 3 3 4 4 4 7 7 8 8 8 13 14 23 23 24 25 26 26 27 28 28 29

2  Medaka and Oryzias Species as Model Organisms and the Current Status of Medaka Biological Resources 31 2.1 ­Introduction 31 2.2 ­Common and Unique Futures of Medaka and Related Species as Model Organisms 31 2.3 ­Phylogenetic Relationships of Medaka and Related Species 35 2.3.1 The javanicus species group 35 2.3.2 The latipes species group 40 2.3.3 The celebensis species group 42 v

vi

Contents 2.4 ­BAC Resources of Species Related to Medaka 2.5 ­National Bio‐Resource Project Medaka (NBRP Medaka) 2.5.1  Support for visiting researchers

3  Looking at Adult Medaka 3.1 ­General Morphology 3.1.1  Secondary sexual characters 3.1.1.1  Dorsal fin 3.1.1.2  Anal fin 3.1.1.3  Papillar processes 3.1.1.4  Urogenital papillae 3.1.2  Body color 3.1.2.1  Pigment cells (chromatophores) 3.1.2.2  Structures of the chromatophores 3.1.2.3  Chromatophores in medaka 3.1.2.4  Chromatophore distribution in medaka 3.1.2.5  See‐through medaka 3.2 ­Anatomy and Histology 3.2.1  Observations of internal organs 3.2.1.1  Observations of internal organs in the live see‐through medaka 3.2.1.2  Dissection of adult medaka 3.2.2  Horizontal and sagittal sections of juvenile medaka 3.2.3  Nervous system 3.2.3.1  Adult central nervous system 3.2.3.2  Adult peripheral nervous system 3.2.4  Endocrine system 3.2.4.1  Hypothalamo‐pituitary system 3.2.4.2  Pineal organ (epiphysis) 3.2.4.3  Thyroid gland 3.2.4.4 Heart 3.2.4.5  Interrenal gland and chromaffin cells 3.2.4.6 Gonads 3.2.4.7  Endocrine pancreas (islets of Langerhans) 3.2.4.8  Gastrointestinal tract 3.2.4.9  Ultimobranchial gland 3.2.4.10  Corpuscle of Stannius 3.2.4.11 Urophysis 3.2.4.12 Thymus 3.2.5 Gonads 3.2.5.1 Ovary 3.2.5.2 Testis 3.2.6 Kidney 3.2.6.1 Pronephros 3.2.6.2 Mesonephros 3.2.6.3  Histology of the kidney Column 3.1  How to make sections of a mature ovary for ­histological analysis

43 43 45 49 49 49 49 49 50 50 51 51 51 51 55 56 56 56 56 58 58 58 58 67 74 76 78 79 81 81 81 81 81 82 82 83 83 83 83 85 85 86 86 86 88

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4  Looking at Medaka Embryos 97 4.1 ­Development of Various Tissues and Organs 97 4.1.1  Developmental stages 97 4.1.1.1   Stage 0: unfertilized egg (Figure 4-1) 97 4.1.1.2   Stage 1: activated egg stage (3 minutes) (Figure 4-1) 99 4.1.1.3   Stage 2: blastodisc stage (Figure 4-1) 99 4.1.1.4   Stage 3: two‐cell stage (1 hour 5 minutes) (Figure 4-1) 99 4.1.1.5   Stage 4: four‐cell stage (1 hour 45 minutes) (Figure 4-1) 100 4.1.1.6   Stage 5: eight‐cell stage (2 hours 20 minutes) (Figure 4-1) 100 4.1.1.7   Stage 6: 16‐cell stage (2 hours 55 minutes) (Figure 4-2) 100 4.1.1.8   Stage 7: 32‐cell stage (3 hours 30 minutes) (Figure 4-2) 100 4.1.1.9   Stage 8: early morula stage (4 hours 5 minutes) (Figure 4-2) 100 4.1.1.10  Stage 9: late morula stage (5 hours 15 minutes) (Figure 4-2) 100 4.1.1.11  Stage 10: early blastula stage (6 hours 30 minutes) (Figure 4-2) 100 4.1.1.12  Stage 11: late blastula stage (8 hours 15 minutes) (Figure 4-2) 102 4.1.1.13  Stage 12: pre‐early gastrula stage (10 hours 20 minutes) (Figure 4-3) 102 4.1.1.14  Stage 13: early gastrula stage (13 hours) (Figure 4-3) 102 4.1.1.15  Stage 14: pre‐mid‐gastrula stage (15 hours) (Figure 4-3) 102 4.1.1.16  Stage 15: mid‐gastrula stage (17 hours 30 minutes) (Figure 4-3) 102 4.1.1.17  Stage 16: late gastrula stage (21 hours) (Figure 4-3) 102 4.1.1.18  Stage 17: early neurula stage (1 day 1 hour) (Figure 4-3) 103 4.1.1.19  Stage 18: late neurula stage (1 day 2 hours)  (Figure 4-4) 104 4.1.1.20  Stage 19: two‐somite stage (1 day 3 hours 30 minutes) (Figure 4-4) 104 4.1.1.21  Stage 20: four‐somite stage (1 day 7 hours 30 minutes) (Figure 4-4) 104 4.1.1.22  Stage 21: six‐somite stage (1 day 10 hours) (Figure 4-4) 104 4.1.1.23  Stage 22: nine‐somite stage (1 day 14 hours) (Figure 4-4) 104 4.1.1.24  Stage 23: 12‐somite stage (1 day 17 hours) (Figure 4-4) 104 4.1.1.25  Stage 24: 16‐somite stage (1 day 20 hours) (Figure 4-5) 106 4.1.1.26  Stage 25: 18–19‐somite stage (2 days 2 hours) (Figure 4-5) 107 4.1.1.27  Stage 26: 22‐somite stage (2 days 6 hours) (Figure 4-5) 107 4.1.1.28  Stage 27: 24‐somite stage (2 days 10 hours) (Figure 4-5) 107 4.1.1.29  Stage 28: 30‐somite stage (2 days 16 hours) (Figure 4-5) 107 4.1.1.30  Stage 29: 34‐somite stage (3 days 2 hours) (Figure 4-5) 108 4.1.1.31  Stage 30: 35‐somite stage (3 days 10 hours)  (Figure 4-6) 108 4.1.1.32  Stage 31: gill blood vessel formation stage (3 days 23 hours) (Figure 4-6) 108

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Contents 4.1.1.33  Stage 32: somite completion stage (4 days 5 hours) (Figure 4-6) 108 4.1.1.34  Stage 33: stage at which notochord vacuolization is completed (4 days 10 hours) (Figure 4-6) 108 4.1.1.35  Stage 34: pectoral fin blood circulation stage (5 days 1 hour) (Figure 4-6) 110 4.1.1.36  Stage 35: stage at which visceral blood vessels form (5 days 12 hours) (Figure 4-6) 110 4.1.1.37  Stage 36: heart development stage (6 days) (Figure 4-7) 110 4.1.1.38  Stage 37: pericardial cavity formation stage (7 days) (Figure 4-7) 110 4.1.1.39  Stage 38: spleen development stage (8 days) (Figure 4-7) 110 4.1.1.40  Stage 39: hatching stage (9 days) (Figure 4-7) 110 4.1.1.41  Stage 40: first larval stage (Figure 4-8) 112 4.1.1.42  Stage 41: second larval stage (Figure 4-8) 113 4.1.1.43  Stage 42: third larval stage (Figure 4-8) 113 4.1.1.44  Stage 43: first juvenile stage (Figure 4-8) 113 4.1.1.45  Stage 44: second juvenile stage (Figure 4-8) 113 4.1.1.46  Stage 45 (Figure 4-8) 113 4.1.2 Brain 113 4.1.2.1  Gastrula step (stages 13–17) 114 4.1.2.2  Neurula step (stages 17–18) 114 4.1.2.3  Neural rod step (stages 19–22) 116 4.1.2.4  Neural tube step (stages 23–27) 116 4.1.2.5  Late embryonic brain step (stages 28–34) 117 4.1.2.6  Larval brain step (stages 35–42) 119 4.1.3  Hatching gland 119 4.1.3.1  Origin of fish hatching gland cells 119 4.1.3.2  Secretion of hatching enzymes from hatching gland cells 122 4.1.4  Eye development 124 4.1.4.1  Specification of the anterior neural plate 124 4.1.4.2  Eye field determination and establishment of retinal identity 125 4.1.4.3  Splitting of the retinal anlage into two retinal primordia 125 4.1.4.4  Morphogenesis I: evagination of the optic vesicle 125 4.1.4.5  Morphogenesis II: formation of the optic cup 127 4.1.4.6  Retinal differentiation I: central retina 127 4.1.4.7  Retinal differentiation II: Ciliary Marginal Zone 127 4.1.4.8  Retinotectal projection 128 4.1.5  Branchial arch and jaws 128 4.1.5.1  Skeletal development 128 4.1.5.2  Muscle development 130 4.1.6 Vasculature 131 4.1.6.1  Vascular anatomy of the developing medaka 131 4.1.6.2  Origin of the medaka endothelial lineage 142 4.1.6.3 Abbreviations 142 4.1.6.4 Acknowledgment 143

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4.1.7  Blood cells (hematopoiesis) 143 4.1.7.1 Overview 143 4.1.7.2  Observation of embryonic and adult blood cells 144 4.1.8 Heart 146 4.1.8.1 Overview 146 4.1.8.2  Heart architecture 146 4.1.8.3  Heart morphogenesis 147 4.1.8.4  Observation of the developing heart 156 4.1.9 Kidney 159 4.1.9.1 Overview 159 4.1.9.2 Nephrogenesis 159 4.1.9.3 Pronephros 160 4.1.9.4 Mesonephros 160 4.1.10 Thymus 160 4.1.10.1 Overview 160 4.1.10.2  Early development of the thymus 160 4.1.10.3  Cortex and medulla 161 4.1.10.4  Involution of the thymus 162 4.1.11  Gut and liver 162 4.1.12 Bones 164 4.1.12.1  Vertebral column 164 Column 4.1  Key words in bone formation 172 4.1.13 Fins 173 4.1.13.1 Overview 173 4.1.13.2  Fin anatomy 173 4.1.13.3  Embryonic fin development (from fertilization to stage 39 [hatching stage]) 174 4.1.13.4  Fin development after hatching (after stage 39) 175 4.1.13.5  Gene expression during fin development 175 4.1.14 Gonads 176 4.1.14.1 Overview 176 4.1.14.2  PGC specification 177 4.1.14.3  Formation of gonadal primordium (Figure 4-60b) 177 4.1.14.4  Sexual dimorphism in germ cell proliferation (Figure 4-61) 179 4.1.14.5  Posthatching period in XX gonads 180 4.1.14.6  Posthatching period in XY gonads 180 4.2 ­Medaka EGG Envelope and Hatching Enzyme 181 4.2.1 Overview 181 4.2.2  Preparation of a hatching enzyme solution from hatching liquid 182 4.2.2.1 Procedure 182 4.2.3  Simple method for preparing hatching enzyme solution 183 4.2.3.1 Procedure 183 4.2.4  Solubilization of the egg envelope using hatching enzyme 183 Column 4.2  Easy method for preparation of a small amount of hatching enzyme solution (see website for figure) 184

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Contents 4.3 ­Observation of Embryos (Embedding Embryos) 4.3.1  Anesthesia of embryos using MS‐222 4.3.1.1  Equipment and reagents 4.3.2  Observation of embryos (mounting) 4.3.2.1  Living embryos 4.3.2.2  Processed embryos 4.4 ­Whole‐Mount In Situ Hybridization (see section 4.1.8.4 for a similar protocol) 4.4.1  Fixation and storage 4.4.1.1  Procedure 1 4.4.2  Rehydration, proteinase K treatment, and postfixation at RT 4.4.2.1  Procedure 2 4.4.3  Hybridization and washing 4.4.3.1  Procedure 3 4.4.4  Immunoreaction and washing antibodies 4.4.4.1  Procedure 4 4.4.5 Staining 4.4.5.1  Procedure 5 4.5 ­Embedding in a Plastic Resin (Technovit 7100) 4.5.1  Equipment and reagents 4.5.2  Agarose mounting (Figure 4-68) 4.5.2.1  Procedure 1 4.5.3  Dehydration and infiltration (Figure 4-68) 4.5.3.1  Procedure 2 4.5.4  Polymerization (Figure 4-68) 4.5.4.1  Procedure 3 Column 4.3  Pigment Cells (Figure 4-69) Column 4.4  Kupffer’s Vesicle

185 185 185 185 185 188 189 189 189 190 190 190 190 191 191 191 191 192 192 192 192 192 192 193 193 194 195

5  Reproductive Behavior of Wild Japanese Medaka 5.1 ­Wild Japanese Medaka 5.2 ­Reproductive Behavior of Wild Medaka 5.2.1  Aggressive behavior 5.2.2  Spawning behavior 5.2.3  Egg deposition behavior 5.2.4  Egg discarding behavior 5.2.5  School and aggregation 5.3 ­Conclusion

205 205 206 207 207 210 210 211 211

6  Cryopreservation and Transplantation of Medaka Germ Cells 6.1 ­Introduction 6.2 ­Cryopreservation of Medaka Testes 6.2.1 Solutions 6.2.2 Materials 6.2.3 Procedures 6.3 ­Transplantation of Thawed Testicular Cells into Recipient Larvae

215 215 215 216 217 217 218

Contents 6.3.1 Solutions 6.3.2 Materials 6.3.3 Procedures Column 6.1  Production of triploid medaka

xi 218 219 219 222

7  Genome Editing 225 7.1 ­Introduction 225 7.2 ­Outline of Targeted Genome Editing Using Nucleases 225 7.3 ­Preparation of CRISPR/Cas9 Genome Editing Tools 226 7.3.1 Materials 227 7.3.2  Production of custom‐designed sgRNA 228 7.3.2.1  Preparation of the bsai‐digested sgRNA backbone 228 7.3.2.2  Design and production of customized sgRNA 228 7.3.3  Production of capped RNA encoding a Cas9 nuclease 232 7.4 ­Preparation of Custom‐Designed TALENs 234 7.4.1 Materials 234 7.4.2  Preparation of the TALEN assembly system 236 7.4.2.1  Preparation of TAL modules (HD1‐6, NG1‐6, NI1‐6, and NN1‐6) 236 7.4.2.2  Preparation of array backbone plasmids (pFUS vectors) 236 7.4.2.3  Preparation of last repeat modules (LR‐HD, NG, NI, and NN) 237 7.4.2.4  Preparation of TALEN backbone plasmids (pCS2TAL3DD and RR vectors) 238 7.4.3  Design and construction of custom‐designed TALENs 239 7.4.3.1  Design of TALEN using TALE‐NT 239 7.4.3.2  First assembly: construction of 6‐modules array vectors 240 7.4.3.3  Second assembly: construction of TALEN expression vectors 242 7.5 ­Heteroduplex Mobility Assay – A Simple Method to Detect Targeted Genome Modification 244 7.5.1 Materials 246 7.5.2 Procedure 246 7.5.2.1  Identification of the wild type, heterozygotes, and homozygotes 246 7.5.2.2  Evaluation of the efficiency of targeted genome modifications 246 7.6 ­How to Establish Gene Knock‐out Strains 247 7.6.1  Design and synthesis of genome‐editing tools 247 7.6.2  Evaluation of genome‐editing activity with fertilized medaka eggs 247 7.6.3  Microinjection of the selected genome‐editing tool(s) 248 7.6.4  Selection of founder fish by genotyping F1 embryos 249 7.6.5  Selection of F1 fish carrying the same mutation and the establishment of mutant strain 249 7.6.6  Selection of homozygous mutant fish in the F2 family 251

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Contents 7.7 ­How to Establish Gene Knock‐in Strains 7.7.1  Design and synthesis of CRISPR/Cas9 components 7.7.2  Evaluation of genome‐editing activity with fertilized medaka eggs 7.7.3  Construction of donor plasmid with homology arms (Ca. 0.5 kbp) and bait sequences 7.7.4  Microinjection for establishing knock‐in strains 7.7.5  Selecting G0 founders harboring the insert gene in the genomic target site Column 7.1  Utilization of crRNA, tracrRNA, Cas9 Protein 7.A  Simple Genomic DNA Preparation by an Alkaline Lysis Method 7.A.1 Materials 7.A.2 Procedure

8  Photo‐Inducible Gene Expression in Medaka 8.1 ­Outline of IR‐LEGO 8.2 ­Practical Strategies of IR‐LEGO in Medaka Study 8.2.1  Selection of heat shock promoters and application studies 8.3 ­Laser Irradiation Conditions and Sample Preparation 8.4 ­Caution in Maintaining Strains 8.5 ­Other Uses of IR‐LEGO 8.6 ­Summary and Future Prospects 9  Screening and Testing Methods of Endocrine‐Disrupting Chemicals Using Medaka 9.1 ­Applied Toxicity Tests for Endocrine Disruptors 9.2 ­Detection of Androgenic and Antiandrogenic Chemicals Using Medaka 9.2.1  The formation of papillary processes on anal fin rays as an ­indicative phenotype for exposure of androgenic and/or ­antiandrogenic chemicals 9.2.2  Candidate biomarkers for assessing the action of androgenic and ­antiandrogenic chemicals 9.2.3  Visualization of androgenic and antiandrogenic activity as green fluorescence with spiggin‐GFP medaka

252 253 253 253 254 254 255 256 256 256 261 261 262 262 265 267 267 268 271 271 275 275 276 276

10 Application of the Seawater Medaka Oryzias melastigma (McClelland) for Marine Ecotoxicology 281 10.1 ­Background and Development of Oryzias melastigma for Marine Ecotoxicology 281 10.2 ­Marine Medaka Developmental Staging 283 10.3 ­Standard Breeding and Rearing Conditions 284 10.3.1 Seawater 285 10.3.2 Temperature 285 10.3.3 Photoperiod 285 10.3.4 Feeding 286 10.3.5  Embryo collection and rearing 286 10.3.6  Hatching and larvae collection 287

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10.3.7  Larvae rearing 288 10.3.8  Larvae feeding 288 10.4 ­Raising Marine Medaka for Experimental Use 289 10.4.1  Experiments using adult fish 289 10.4.2  Experiments using larvae 289 10.5 ­Troubleshooting 289 10.5.1  Mass mortality 289 10.5.2  Low egg production 289 10.5.3  Extensive algal growth 290 10.6 ­How to Obtain Marine Medaka O. melastigma 290 10.7 ­Experimental Protocols Using Marine Medaka 290 10.8 ­Immunotoxicity Assessment: Bacteria Challenge Assays 290 10.8.1  SOP for adult bacterial challenge assay 291 10.8.2  SOP for larval bacterial challenge assay 292 10.8.3  Age selection for larval bacterial challenge 293 10.9 ­Fish Dissection and the Whole Adult Histoarray 294 10.9.1  SOP for fish dissection 295 10.9.2  SOP for adult medaka histoarray 295 10.10  Embryo Chip 297 10.10.1  SOP for embryo and larvae histoarray 297 10.A  Materials for SOP for Adult Medaka Histoarray (see section 10.9.2) 299 11  Telomerase and Telomere Biology in Medaka 303 11.1 ­Introduction 303 11.2 ­SOP for Quantification of Telomerase Activity Using the Real‐Time Quantitative Telomeric Repeat Amplification  Protocol (RTQ‐TRAP) 308 11.2.1  Procedures for sample extraction 308 11.2.2 Procedures for determination of protein concentration 308 11.2.3  Procedures for RTQ‐TRAP linearity test 308 11.2.4  Calculation of telomerase activity 309 11.3 ­SOP for Quantification of Telomere Length Using Southern Blotting Analysis 309 11.3.1 Procedures for genomic DNA extraction and digestion with restriction enzymes 309 11.3.2  Procedures for probe preparation 311 11.3.3  Procedures for electrophoresis and southern blotting 311 11.3.4  Procedures for hybridization and detection 312 11.3.5  Procedures for computerized telomere analysis 312 11.4 ­SOP for Quantification of Telomere Length Using Fluorescence In Situ Hybridization 313 11.4.1  Procedures for fluorescence in situ hybridization 313 11.4.2  Procedures for confocal microscopy detection 313 11.4.3  Procedures for ImageJ analysis 314

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Contents

12  Assessments of Medaka Skeletal Toxicity 12.1 ­Introduction 12.2 ­Methods 12.2.1  Embryonic exposures: dioxin 12.2.2  Embryonic exposure: dithiocarbamates 12.2.3  Whole‐mount alcian blue staining of hatchlings/larvae a 12.2.4  Whole‐mount Alizarin red S staining of hatchlings/larvaec 12.2.5  In vivo Alizarin complexone fluorescent staining for ­mineralized bone matrix 12.2.6  In vivo calcein fluorescent staining for mineralized bone matrix 12.2.7  Confocal imaging of embryo/hatchling medaka 12.2.8  Morphological assessments 12.3 ­Results and Discussion 12.3.1 Dithiocarbamates 12.3.2 Dioxin

317 317 318 319 319 320 320 321 321 321 323 324 324 325

Appendix A  Solutions 329 Attributions 331 Index 335

List of Contributors Satoshi Ansai Laboratory of Bioresources, National Institute for Basic Biology, Japan

Misato Fujita Department of Biological Sciences, Faculty of Science, Kanagawa University, Japan

Doris W.T. Au State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong

Shoji Fukamachi Laboratory of Evolutionary Genetics, Department of Chemical and Biological Sciences, Japan Women’s University, Japan

Napo K.M. Cheung State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong

Hisashi Hashimoto Bioscience and Biotechnology Center, Nagoya University, Japan

Michael W.L. Chiang State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong Shin-ich Chisada Department of Preventive Medicine and Public Health, Kyorin University, School of Medicine, Japan Tomonori Deguchi Biomedical Research Institute, National Institute of Advanced Industrial Science and Technology, Japan

Narisato Hirai National Research Institute of Fisheries and Environment of Inland Sea, Japan Fisheries Research and Education Agency, Japan Taisen Iguchi Nanobioscience, Yokohama City University, Japan Masayuki Iigo Department of Applied Biological Chemistry, Faculty of Agriculture, Utsunomiya University, Japan Keiji Inohaya School of Life Science and Technology, Tokyo Institute of Technology, Japan

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List of Contributors

Yuji Ishikawa Neurobiology Lab, Japan Yasuko Isoe Graduate School of Natural Science and Technology, Okayama University, Japan Ichiro Iuchi Department of Materials and Life Sciences, Faculty of Science and Technology, Sophia University, Japan

Fumi Kezuka Department of Aquatic Biosciences, Tokyo University of Marine Science and Technology, Japan Masato Kinoshita Department of Applied Biosciences, Graduate School of Agriculture, Kyoto University, Japan Shin-ichi Kitamura Centre for Marine Environmental Studies, Ehime University, Japan

Norimasa Iwanami Department of Regenerative Medicine, National Center for Geriatrics and Gerontology, Japan

Daisuke Kobayashi Department of Anatomy and Developmental Biology, Kyoto Prefecture University of Medicine, Japan

Eri Iwata Laboratory of Veterinary Ethology, Faculty of Veterinary Medicine, Okayama University of Science, Japan

Makito Kobayashi Laboratory of Aquatic Biology, Department of Natural Sciences, International Christian University, Japan

Yasuhiro Kamei Spectrography and Bioimaging Facility, NIBB Core Facilities, National Institute for Basic Biology, Japan

Akira Kudo International Frontier, Tokyo Institute of Technology, Japan

Sakurako Kamide Laboratory of Aquatic Biology, Department of Natural Sciences, International Christian University, Japan Takashi Kawasaki Medical and Biological Engineering Research Group, Biomedical Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), Japan

Seth W. Kullman Department of Biological Sciences, North Carolina State University, USA Rie Kusakabe Laboratory for Evolutionary Morphology, RIKEN Center for Biosystems Dynamics Research, Japan

List of Contributors Sungki Lee Biological and Genetic Resources Assessment Division, National Institute of Biological Resources, South Korea Kouichi Maruyama National Institute of Radiological Sciences, National Institutes for Quantum and Radiological Science and Technology, Japan Shinichi Miyagawa Department of Biological Science and Technology, Faculty of Industrial Science and Technology, Tokyo University of Science, Japan Helen O.L. Mok State Key Laboratory of Marine Pollution, Department of Biomedical Science, City University of Hong Kong, Hong Kong Yu Murakami Department of Applied Biosciences, Graduate School of Agriculture, Kyoto University, Japan Kenji Murata Center for Health and the Environment, University of California Davis, USA Yuki Nakatani Department of Biological Sciences, Tokyo Institute of Technology, Japan Kiyoshi Naruse Laboratory of Bioresources, National Institute for Basic Biology, Japan

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Yukiko Ogino Centre for Promotion of International Education and Research, Faculty of Agriculture, Kyushu University, Japan Kataaki Okubo Department of Aquatic Bioscience, Graduate School of Agricultural and Life Sciences, University of Tokyo, Japan Drew R. Reterson State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong Daisuke Saito Research Center for Systems Immunology, Kyushu University, Japan Anthony Sèbillot Département Adaptations du vivant, "Évolution des régulations endocriniennes", Muséum National d’Histoire Naturelle, France Shinsuke Seki Bioscience Education and Research Support Center, Akita University, Japan Eriko Shimada Laboratory of Cell Biology, Cellevolt, Japan Ai Shinomiya Division of Seasonal Biology, National Institute for Basic Biology, Japan

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List of Contributors

Ian T. Stancil Department of Biological Sciences, North Carolina State University, USA Yoshiro Takano Department of Cell Biology and Neuroscience, Juntendo University School of Medicine, Japan Yusuke Takehana Department of Animal Bioscience, Nagahama Institute of Bio‐Science and Technology, Japan Minoru Tanaka Division of Biological Science, Nagoya University, Japan Yoshihito Taniguchi Department of Preventive Medicine and Public Health, School of Medicine, Kyorin University, Japan Norihisa Tatarazako Graduate School of Agriculture, Department of Science and Technology for Biological Resources and Environment, Ehime University, Japan Yuko Wakamatsu Physiological Chemistry I, Biocenter, University of Würzburg, Germany AtLee T.D. Watson Department of Biological Sciences, North Carolina State University, USA

Joachim Wittbrodt COS Heidelberg, Heidelberg University, Germany Rudolf S.S. Wu Department of Science and Environmental Studies, Education University of Hong Kong, Hong Kong Kazunori Yamahira Tropical Biosphere Research Center, University of the Ryukyus, Japan Shigeki Yasumasu Department of Materials and Life Sciences, Faculity of Science and Technology, Sophia University, Japan Ryohei Yatsu Department of Integrative Biology, University of Texas, USA Roy R. Ye State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong Bill W.P. Yip State Key Laboratory of Marine Pollution, Department of Chemistry, City University of Hong Kong, Hong Kong Hiroki Yoda Developmental Biology Unit, European Molecular Biology Laboratory, Germany

List of Contributors Hirofumi Yokota Department of Biosphere Sciences, School of Human Sciences, Kobe College, Japan

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Goro Yoshizaki Department of Aquatic Biosciences, Tokyo University of Marine Science and Technology, Japan

Preface Ten years after publishing our first book (Volume 1) in 2009, this second book (Volume 2) of Medaka: Biology, Management, and Experimental Protocols presents significant progress in technological innovation and development in the fields of biological and medical science. The purpose of Volume 2 is to familiarize scientists worldwide with the advantages of using medaka in experimental designs, to facilitate research using medaka, and address the value to science of adopting medaka as a model animal. In Volume 2, the authors provide additional information and current protocols that have been recently developed, or modified, to successfully raise medaka under stable laboratory culture conditions and how to use medaka as a model animal. This Volume 2 describes new technologies developed after 2009, using the fish as a molecular tool in the fields of life science, evolution, ecology, and toxicology. It provides an informational bridge that spans the varied research disciplines and abilities ranging from undergraduate education through senior researchers. Contributing authors were chosen because of their expertise and demonstrated ability to conduct experiments involving medaka, and because they are recognized pioneers in the use of medaka as a model animal in their scientific fields. The authors were also asked to describe their experimental protocols in detail, and explain their rationale for the chosen protocols in achieving their conceptual goals. The editors also recommend that users read the procedures described in the first edition that describe the maintenance of medaka, and use that information to create or modify the current fish maintenance systems. Chapter 1: Dr. Chisada and colleagues describe contemporary procedures used to maintain medaka in culture facilities. Chapter  2: This chapter covers the current phylogenetic relationship of medaka and other Oryzias species, and the geographical distributions of each species. In this chapter, Dr. Naruse and colleagues also describe and update the present status of medaka resources available through the National Bio‐Resource Project Medaka (NBRP Medaka) that has been supported by the Japanese government since 2002. Chapters 3 and 4 introduce the reader to the medaka fish: Chapter  3 covers general information about adult medaka, including secondary sexual characters, body color, and internal organs. Section  3.2, “Anatomy and Histology,” provides details of the nervous system, endocrine system, kidneys, and gonads. Chapter 4 covers the characteristics of the developing embryos. It includes brief outlines of the development of the fish’s organs and tissues, with an emphasis on histology rather than developmental mechanisms. Chapter 4 also discusses the basic procedures for preparing and mounting embryos, and performing in situ hybridization. These similar chapters appear as Chapter 5 and 6 in Volume1; however, the authors retained the material in Chapter 3 and 4 of volume2, because of the importance of the information. To address medaka reproduction, two groups of researchers were chosen to describe procedures and applications for the preservation of genetic resources of the fish, and for the fish’s reproductive ecology.

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Chapter 5: Dr. Kobayashi and colleagues, experts on wild medaka fish, address sexual behavior and reproduction of wild populations of medaka. The chapter’s focus is on wild populations as opposed to laboratory‐maintained fish. Chapter 6: Dr. Kezuka and colleagues introduce methods of cryopreservation of whole fish testes, the preparation of testicular cell suspension, and testicular cell transplantation into recipient fish. This method will potentially contribute not only to saving and recovering populations of endangered species, but also to increasing the numbers of commercially important, high‐quality food fishes. Since publication of the first Volume, new technologies using medaka have been developed and these advances contribute to the identification and function of genes and their products in the body. Two useful technologies are addressed as follows: Chapter 7: In this chapter, Dr. Ansai and colleagues describe recent developments and progress in the amazing application of molecular biology used in gene editing using medaka. They address the basic theory and applications used to create and apply the establishment of transgenic, gene knock‐out medaka, and conditional knock‐out medaka. Chapter 8: In this chapter, Drs. Isoe and Kamei describe newly developed technology using medaka and the infrared laser‐evoked gene operator (IR‐LEGO) system. They explain how to use the system not only for research using medaka, but also with cultured cells and plants. The authors also address how to analyze the function of genes in vivo, in developmental biology and related fields, and the development of a new microscopic technology used to control the expression of genes using the combination of the heat shock promoter system and an infrared laser, by focusing on the targeted cells. Chapters 9–12 introduce the use of medaka in the fields of toxicology and medicine; and address recently developed methodologies and new technologies. Chapter  9: Dr. Iguchi and colleagues describe several test guidelines used by the Organization of Economic Cooperation and Development (OECD) employing medaka in standardized testing methods to screen and/or access potential endocrine‐disrupting chemicals, and how the fish is used to identify adverse effects of toxic chemicals. They also describe how toxicologists are using medaka to assess hazardous chemicals in both drinking water and sewage effluent, and to identify endocrine disruptors in natural waters. The authors address endocrine disruptors, not only restricted to chemicals with estrogenic activities but also those with androgenic activities. Dr. Ogino and colleagues also describe the use of medaka to detect androgenic and antiandrogenic chemicals. Chapter 10: This chapter introduces the basic biology of the marine medaka, Oryzias melastigma (McClelland), and how its use has been applied to research in marine ecotoxicology. Dr. Peterson and colleagues describe the fish’s biological characterization, how to maintain the species in the laboratory, and how to use it in research on marine ecotoxicology. Chapter 11: Dr. Au and colleagues, experts in telomere biology using medaka, introduce their SOPs and procedures, which have great potential for research in the fields of toxicology, senescence, and aging. They address medaka as a unique, alternative vertebrate model for studying telomere and telomerase function in a cross‐disciplinary range from environmental toxicology to biomedical research on aging, as well as for cancer and tissue regeneration research. Chapter 12: Dr. Watson and colleagues describe the application of medaka in research on human skeletal biology and toxicity and as a human disease model.

Preface

xxiii

The format of Volume 2 is designed to capture the thoughts and methods of researchers that use medaka as a model animal and to make this expertise accessible to students, beginning researchers, and senior researchers who might become intrigued with using medaka as the model animal in their own work. To accomplish this, and following a reading of Volume 1, the reader is given step‐by step specifics for each protocol that allows application of the fish in their own work. The information includes specific information to facilitate ease of adoption, including details such as reagents used, instrumentation, and other essential requirements. It is anticipated that this highly practical format will encourage the reader to use medaka as a model animal and permit the reader to bring new concepts into personal practice in a more efficient manner. The use of medaka fish as a model animal requires experimental insight and practical troubleshooting of experimental designs. Of equal importance is an overall appreciation of both the power and limits of using medaka as a model animal. To assist in visualizing and understanding medaka and the research protocols used, the editors strongly suggest that readers refer to Volume 1 of Medaka: Biology, Management, and Experimental Protocols (2009) (ebook: http://onlinelibrary.wiley.com/book/10.1002/9780813818849) as a reference. All figures and videos in both Volumes are shown at https://medaka-book.org/. As a final note, the preparation of this book would not have been possible without the dedication of the excellent array of contributing authors. We also thank the staff of Wiley‐ Blackwell Publishers, specifically Justin Jeffryes, Shelby Hayes, Rebecca Ralf, Antony Sami, Holly Regan-Jones and Vivek Jagadeesan as they have demonstrated great patience with our efforts and provided excellent guidance and assistance. Finally, we also express our thanks to Ms. Robin Lee Kingsley (Deceased) and Dr. Fred S. Conte (University of California Davis) for their assistance in editing each chapter. Kenji Murata, Masato Kinoshita (Chief Editors) Yasuhiro Kamei, Minoru Tanaka, Kiyoshi Naruse (Editors)

Chapter 1

Medaka Management

1.1 ­Introduction This chapter provides supplemental information about medaka breeding described in Chapter 2 Medaka Management of the first edition of Medaka: Biology, Management, and Experimental Protocols (2009). In this decade, the biological usefulness and convenience of medaka as a model animal have been recognized and the number of research fields and researchers using medaka has been expanding. In these situations, strict and sophisticated rearing methods are required to evaluate the effects of the gene of interest, chemicals, ­environmental conditions, and so on. Well‐designed and regulated feeding is especially critical to evaluate growth and appetite. Additionally, ethical issues such as animal welfare have been emphasized including the “3Rs” concept: Replacement, Reduction, and Refinement. Therefore, it has become important to understand anesthesia and euthanasia methods. Understanding the life‐cycle of medaka in the wild is important to construct a research plan for successful breeding in the laboratory. First, in this chapter, the life‐cycle of medaka in the wild is described. Then, the outline of the breeding process is mentioned before the detailed breeding procedure is explained. Finally, anesthesia methods are discussed.

1.2 ­Medaka Management for Scientific Research In breeding model animal strains, the most critical point is to maintain the phenotypical characteristics of the various strains in order to obtain experimental reproducibility. Although the same model animal strain is subjected to a certain experiment, the results may differ due to different breeding and maintenance conditions. Therefore, it is very important to describe the proper breeding and maintenance conditions as well as the significant results in the research paper so that other researchers can reproduce these procedures as accurately as possible. In this section, the standard breeding procedures based on the life‐ cycle of medaka in the wild are described; they are helpful in setting up a medaka breeding system as a first step to begin a research project.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

1

2

Medaka

1.2.1  Outline of medaka life‐cycle in the wild Medaka (Japanese medaka, Oryzias latipes) has a widespread distribution in Japan, primarily in small ponds and rice paddies (See 1.2 Phylogeny in Chapter 1 in the first edition of Medaka: Biology, Management, and Experimental Protocols (2009) and Chapter 2 in this book). Medaka can live throughout the year in Japan (Shima and Mitani 2004), but has a limited spawning period in the wild. More than 12 hours of daylight and water temperature higher than 13 °C are required for oogenesis and spermatogenesis to take place. Then, the fish start to mate and the females spawn eggs. These climate conditions match those during spring and summer in Japan. Actually, the combination of a 14 hour light/10 hour dark (14L–10D) cycle and temperatures of 25–28 °C (which are consistent with those of early summer in Japan) and sufficient food provides the best conditions for spawning eggs in the wild. In spring, in the wild, when the water temperature is lower than 25 °C, larvae hatch in 10 or more days. The larvae reach maturity in the early summer. They mate and spawn eggs until the hours of daylight become shorter than the hours of darkness (late summer to autumn). A small number of the next generation survives the winter and a smaller number may survive two winters. The average lifespan in the wild is considered to be around one year because predation and/or seasonal environment change are the leading causes of death of medaka in the wild. Egami reported that the average lifespan is less than three years and the maximum lifespan is approximately five years under experimental conditions (Egami 1971).

1.2.2  Preparation of normal rearing conditions of medaka in the laboratory and procedures for breeding The outline of the rearing schedule is shown in Figure 1-1. To obtain eggs, creating the conditions that are required for successful mating and spawning is important. Details of these conditions are described in 3.5 Necessary Conditions for Spawning in Chapter 3 in the first version of this book. Briefly, these conditions consist of feeding three times a day, a 14L–10D light cycle at 25–28 °C, and avoiding keeping the fish at a high density.

1.2.2.1  Breeding system set‐up Select the water system (flowing, recycled, or static water) according to the laboratory situation or the requirements of the experiment, and prepare the water to fill the fish tanks. Soft water (this means the water contains a concentration of calcium and magnesium ions lower than 120 mg/L) without chlorine is recommended. The light/ dark cycle (14L–10D) and the room temperature (25–28 °C) of the breeding room should be controlled respectively using a timer and an air conditioner, before bringing medaka into the room. In case it is difficult to control the lighting conditions of the room where they have been set for the experiments and the regulation of the water temperatures in each breeding tank in the room as a single unit, the light/dark cycle and the temperatures of the water in the tanks can be independently regulated at each breeding tank or aquarium system. (See Chapter 2 Medaka Management in the first version of this book.)

Medaka Management

Tools Fish Tanks and Dishes Water Foods Light and Timer

3

Daily care of medaka culture Embryo culture (9-day incubation) Eggs

Larva culture (A month nursery)

Next generation

hatching

Adult culture (a pair) Continue daily care

3 pairs of adult

Stock center (e.g. NBRP Medaka)

12

New strain

Embryos

Embryos are cultured in a small dish in 0.1 x sea water under 27°C incubator. Embryos will hatch at day 9.

hatching

Larvae are cultured in a small tank in 0.1 x sea water under 25–27°C. Frequent feeding and water change. Nursery will be last for a month.

A month nursery

Water change etc.

Eggs

9-day incubation

Adults

9

Young fish

18

Daily cycle for medaka culture

Larvae Feeding:

24 Egg collection:

Figure 1-1.  Outline of the rearing schedule.

1.2.2.2  Obtaining medaka It is strongly recommended that medaka be obtained from researchers who are culturing medaka or from the National Bio‐Resource Project (NBRP) Medaka (https://shigen.nig. ac.jp/medaka) to prevent the contamination and/or introduction of pathogens or parasites into the aquarium(s). The NBRP Medaka at the National Institute for Basic Biology in Japan distributes medaka strains, including wild, inbred, transgenic, and mutant strains, domestically and internationally. In the wild, medaka habitats are located in areas that encompass tropical, temperate, and cold climate zones within various types of aquatic habitats, with water conditions ranging from still, freshwater lakes, flowing freshwater streams, and rice paddies to brackish and sea water; medaka have adapted to all these conditions. However, to avoid problems such as the contamination of pathogens and/or parasites from the environment as described above, the medaka used for research should be kept away from all wild aquatic animals, and water taken directly from the natural environment should never be used, since untreated water from natural habitats may harbor pathogens and/or parasites. If it should be necessary to use medaka obtained from the wild or pet shops, these fish should be reared in an independent tank to collect more than 20 eggs from them. After pasteurizing (see section 1.3.2.2) these eggs, the larvae that hatch from these eggs can be introduced into the aquarium. Also see Chapter 2 Medaka Management in the first version of this book.

1.2.2.3  Collecting eggs in a laboratory setting In order to obtain eggs from medaka to use for your experiment, introduce a pair of adult medaka into a half‐full breeding tank. The volume of the tank should be at least 1 L

4

Medaka

(10 × 10 × 10 cm). The fish will spawn eggs (10–30 eggs a day per a female) within two weeks with enough food and proper water temperature (25–28 °C).

1.2.2.4  Daily care and maintenance of eggs The collected eggs are cultured with embryo culture medium* (ECM) or 0.3% (W/V) artificial sea water in a plastic dish (a, b, and c in Table 1-1) at 25–28 °C. Since a lack of oxygen causes delay of ontogeny and/or abnormal development of the embryos, an excessive number of embryos in a dish should be avoided. The proper number of embryos in a dish is shown in rows a, b, and c in Table 1-1. The dish should be examined every day for abnormal and/or dead embryos which should be promptly removed; otherwise fungi and bacteria will multiply sufficiently to kill the embryos. The details of sorting and cleaning methods of embryos are described in sections 1.3.2.2 and 1.3.2.3. * 0.1% (W/V) NaCl, 0.003% (W/V) KCl, 0.004% (W/V) CaCl2‐2H2O, 0.016%(W/V) MgSO4‐7H2O.

1.2.2.5  Rearing medaka from the larval stage to adulthood After bringing the newly acquired larvae to the breeding facility, transfer them to a culture tank (see d or e in Table 1-1) as soon as possible. Larvae can start eating in one or two days after hatching, therefore it is necessary to start feeding them just after the transfer of hatched larvae into the tanks. The amount of food and frequency of feeding depend on the type of food. The feeding methods using paramecia and commercial powdered food are described in section 1.3.2.4, and Table 1-2. Leftover food on the surface and bottom causes deterioration of water quality. In breeding with static water, leftover food and debris should be removed every day and half of the breeding water should be changed every day. When breeding fish in a tank that has a water recycling system with filtration, it is recommended that leftover food and debris are removed every day. One method for removing them from the bottom of the tank is described in section 1.3.3.2. The amount of food should be increased as the larvae grow. The size of the food particles is another important matter and should be changed as the fish grow. Information about the particle size and the amount of commercial powdered food needed is described in Tables 1-2 and 1-4. Additionally, since the density of larvae in the tank/ aquarium affects the growth rate, the size of the tank and the number of fish should be adjusted to the optimal conditions as shown in Table 1-3. The fish grow to sexual maturity within three months in proper breeding conditions  and they can continue spawning for at least for three months. Rearingmultigeneration will allow to obtain eggs throughout the year (Figure 1-2). It is helpful to keep in mind that a lower density of fish in a breeding tank results in a shorter time between generations.

1.2.2.6  Anesthesia and euthanasia The ethics of animal welfare have become an important issue in daily life and even affect conditions in scientific fields. Therefore, the fish used in experiments should be treated properly after the experiments have finished. Principally, these fish should be treated according to the guidelines of experimental animals established by each institute or

Medaka Management Table 1-1.  Tools for medaka embryonic/larval culture and treatments. Tools

Size, company, and remarks

a

Polystyrene dish (without coating)

b

Polystyrene dish (without coating)

c

Polystyrene dish (without coating)

d

Round plastic cup (deep type)

e

Round plastic cup (deep type)

f

Komagome pipette (glass pipette)

f’

Silicone nipple (for 10 mL)

g

Transfer pipette (polypropylene)

h i

Transfer pipette (polypropylene) Fine forceps

j

Steel needle with handle

k

Medaka spoon

l

Nonslip forceps (serrated, straight, fine tip)

m

Small scissors

n

Micro scissors (spring type)

35–40 × 10–13 [diameter × height mm] (similar products) To culture 10.0%, fiber 1.50% Particle size: 0.36–0.65 mm For 13 wk stage to adulthood Nutrition: protein >50.0%, fat >10.0%, fiber 1.50%

Table 1-3.  Optimal tank size and density (numbers of fish). Stagea

Round container

Circulation system rack

(body length)

d

e

2–3 L tank

5 L tank

9 L tank

First to second larval (25 mm)

1~15

10~60 ~20

~50

~100

~10

~20

~30

~5

~15

~20

~5

~10

~15

 The photos of round containers d and e are in Figure 1-3.  The definition of stages is described in Chapter 6 of the 1st edition. b  The fish display signs of secondary sexual characteristics but haven’t produced eggs yet. 2 a

government. Although some governments have guidelines for welfare for mammals, birds, and reptiles, they ­generally lack guidelines for fish and amphibians. Nevertheless, the scientific community expects that experimental model fish are controlled by appropriate guidelines and that each research project using fish should be approved by local or regional ethics committees or related organizations involved in handling animal welfare before each project is started. Therefore, considering this situation, knowledge of the effects of anesthesia on fish is required not only in regard to the welfare of the animals, but also in order to humanely

Medaka Management

Tools Fish tanks Water Foods and so on

7

Rearing schedule for a year-round egg collection

Eggs for experiments

Adult culture (starting pair: G0)

Eggs for experiments

Eggs for the next generation

Next generation (F1)

Embryos

Larvae Young fish Eggs for the next generation (F2)

Figure 1-2.  Outline of generating medaka eggs year round.

euthanize the fish at the end of experiments. The details of anesthesia methods are described in section 1.3.4.

1.3 ­Standardized Culture and Growth Curve 1.3.1  Characteristics and selection of strains Medaka, as a model animal, has been used for more than 100 years. To date, many strains (such as the wild population in various regions in Japan and the Far East, color mutants, artificially induced mutants, and transgenic fishes) have been collected. Most of these strains have been kept in small‐scale breeding facilities, for example keeping 10 females and 10 males of each generation of each strain. As a result of small‐scale and long‐term breeding, the genetic background has become unified in each strain. These strains are called “closed colonies” and each fish in a colony has homogeneous morphological characters, behaviors, and personalities. Moreover, the internal crossings of brother–sister (sib) mating for more than 20 generations has resulted in the establishment of “inbred strains.” At present, about 10 inbred strains are kept in the NBRP Medaka stock center. These fish have no differences in their genetic background. Among experimental model animals, except for the mouse and rat, only medaka has multiple inbred strains. On the other hand, it is difficult to breed inbred strains. For example, the most famous inbred strain, Hd‐rR, which was used for the Medaka Genome Project (https://shigen.nig.ac.jp/medaka/genome/ top.jsp), has some disadvantages for experiments. The obvious disadvantages are the smaller numbers of spawned eggs and shorter spawning period compared with Cab which has been kept with mass mating and possesses polymorphism in its genome. This defect is often seen in many inbred strains, so at the beginning of research, it is important to select medaka strains that are suitable for the aim of the research. Although inbred strains retain some defects such as the smaller numbers of spawned eggs, their homogeneous genetic background is advantageous and attractive for specific studies, such as the assembly of genome sequence and fine quantitative trait locus (QTL) analysis. Refer to the first edition of this book and the NBRP Medaka HP (https://shigen.nig.ac.jp/medaka) for more information about inbred strains and other medaka lines, such as transgenic lines, natural populations, and mutants.

8

Medaka

1.3.2  Management of medaka eggs and fish The standard conditions of light and temperature are the 14L–10D cycle and 25–28 °C for medaka embryos, larvae, and adults.

1.3.2.1 Mating A mature female prefers a familiar male to an unfamiliar male for mating and spawning (Okuyama et al. 2014). Therefore, it is better to breed a female and a male in a tank at least one day before the scheduled egg collection day. In the laboratory, under regulated lighting conditions, medaka spawn within one hour after the light is turned on. Occasionally, a female will not accept the male in the same tank. Therefore, if the pair does not spawn for a few days, replace the male fish with another one. A good pair spawns every day. Usually, females spawn 10–20 eggs a day, but this number is reduced in females of inbred strains (the number differs slightly according to the type of inbred strain). When many eggs are required, increase the number of breeding tanks where a male and a female are breeding or add one more female to the breeding tank to make a male to female ratio of 1:2. It is not recommended that more than one male fish is bred in a single tank for the following reasons: male fish fight each other for females, making certain males vulnerable; and the spawning performance of a male and a female is interrupted by another male. These behaviors reduce the number of eggs spawned. It is also important to provide enough food to the breeding pairs to obtain many eggs. Details for feeding are given below.

1.3.2.2  Management of embryos Keeping the ECM clean is one of the most critical matters in order to culture and maintain healthy embryos; otherwise bacteria and/or fungi will grow and cause harmful effects on embryonic development. Another critical matter is the number of embryos in a culture dish. Too many embryos in one dish disrupt normal embryonic development. The recommended number of embryos per culture dish is shown in Table 1-1. Under the appropriate culture conditions, the date of hatching can be controlled. Tools The tools used in embryo/larva culture are listed in Table 1-1 and Figure 1-3. Alternatives are listed below. Reagents and Solutions 1. Sea water (SW): Dilute 3 g of artificial sea salt powder with 100 mL of reverse osmotic water (RO water) or distilled water (DW). 2. Embryonic water (EW): Dilute SW by 10% with RO water or DW (The final concentration of sea salt is 0.3%). This medium can be replaced by a balanced salt solution (BSS), Yamamoto’s Ringer solution, or ECM (BSS: 0.65% NaCl, 0.04% KCl, 0.02% MgSO4‐7H2O, 0.02% CaCl2‐2H2O, 0.001% phenol red [sterilize and adjust to pH 8.3 with 5% NaHCO3]. Yamamoto’s Ringer solution: 0.75% NaCl, 0.02% KCl, 0.02% CaCl2, 0.002% NaHCO3 [adjust to pH 7.3 with 5% NaHCO3]. ECM: 0.1% NaCl, 0.003% KCl, 0.016% MgSO4‐7H2O, 0.004% CaCl2‐2H2O).

Medaka Management

9

Figure 1-3.  Tools for medaka embryo/larva culture and treatments. Standard use of each tool is explained in Table 1-1.

3. Methylene blue (MB): CAS number 61‐73‐4. 4. MB stock: 0.1% methylene blue stock solution in RO water or DW. 5. MB‐EW*: Dilute MB stock with 1000 times volume of EW. 6. Sodium hypochlorite (SH) stock solution: Some chemical companies provide hypochlorite solution, for example Wako 194‐02216. Be sure not to use certain hypochlorite products (for example, breach reagent for clothes) that contain NaOH and detergents. 7. SH‐EW: Sodium hypochlorite working solution for the pasteurization of eggs. Dilute 150 μL SH stock with 1.5 L EW. * For more information on methylene blue, refer to the last part of section 1.3.2.3. Collecting and separating eggs A female holds eggs on the outside of her cloaca region in her abdomen after spawning. To collect eggs, first scoop the female with a small net and then, using a pipette, suck the eggs off from the body (Refer to movies M3‐2a and M3‐2c in the first version of this book). The eggs form a cluster with their attaching filaments (see Figure 1-5). In order to more easily manipulate and/or move the eggs during experiments, the attaching filament on the egg envelope (chorion) should be removed and the clustered eggs should be separated individually. It is also important to remove unfertilized and abnormal eggs which may become feeding grounds for bacteria and/or fungus growth. Since the egg envelope is very soft just after spawning, it is better to collect the eggs after the envelope has hardened (wait for 1 hour) unless a very early stage of fertilized eggs is required; for example, one‐cell stage eggs are required for microinjection. These eggs are firm enough to handle with fingers. In this section, two methods for removing attaching filaments and separating the eggs from the cluster are described. One is rubbing the egg cluster with a finger onto a piece of paper or into a culture dish (Figure 1-4). The other is twisting each egg with forceps (Figure 1-5).

10

Medaka

Figure 1-4.  Egg separation with a finger (method 1).

Separation method 1 (rubbing the eggs with a finger) (Figure 1-4)  This method is simple but be sure to use it only after the egg envelope has become hard enough (approximately one hour after fertilization). This method is demonstrated in Movie M3‐3a in the first version of this book. Materials Plastic dishes (tool c in Table 1-1) Two transfer pipettes (tool g or h in Table 1-1) Fine forceps (tool i in Table 1-1) EW (appropriate volume) Disposable rubber gloves (if necessary) Procedure 1. Transfer egg clusters into a dish filled with EW. 2. Remove debris using a pipette. 3. Gather the clusters in the center, and then push and rub eggs with a finger (with or without a rubber glove). 4. Keep rubbing gently by rotating the finger over the top of the egg cluster, just like drawing a small circle; the eggs will detach from the clutch one by one. 5. Stop rotating when each cluster contains a few eggs and then, clip a bundle body (a small clot of attaching filaments) directly with forceps and pull apart the remaining bundles to remove them from the eggs using the other forceps. 6. Clean the eggs by the method described in next page. Modified Method It is also possible to rub the eggs onto the surface of a newspaper or paper towel in order to separate the eggs, but some eggs may fly off the paper. Separation method 2 (twisting the eggs with forceps) (Figure 1-5)  This method requires two fine forceps and is suitable for eggs with soft egg envelopes (within 30 minutes after

Medaka Management

11

Figure 1-5.  Egg separation with two forceps (method 2).

fertilization). Therefore, this method is useful to prepare fertilized eggs in the early stage required for microinjection. Refer to movies M3‐3b to 3d and M7‐1c in the first edition. Materials Fine forceps (× 2) (tool i in Table 1-1) Plastic cup (tool d or e in Table 1-1) Plastic dish (tool a, b, or c in Table 1-1) EW (appropriate volume) Method 1. Transfer the egg clusters onto a plastic dish or a plastic cup filled with EW. Use a stereo microscope to carry out the following procedures. 2. Gather the clusters into one big cluster as if tangling the attaching filaments of each cluster. 3. Pinch thick bundles of attaching filaments using one pair of forceps, and pinch other ­bundles with another pair of forceps. 4. Tangle or wind the attaching filaments around the bundles using the forceps. 5. The eggs will drop down onto the bottom of the dish or cup one by one (keep ­tangling the filaments). 6. Stop tangling the filaments when the cluster is reduced to a few eggs. Remove the remaining bundles from eggs with forceps. 7. Clean eggs using the method described in next section. Egg cleaning It is necessary to clean the eggs soon after their separation from the bundle in order to avoid the growth of microorganisms and fungi. Rinsing the eggs three to five times with EW is sufficient to clean the eggs. When the eggs are collected from the bottom of the tank (eggs that have already detached from the female’s abdomen), it is likely that they will be contaminated with microorganisms and fungi, which will adversely affect incubation. In that situation, stricter cleaning methods using bleach (the pasteurization method) are recommended and will reliably maintain higher viability of the embryos.

12

Medaka

This procedure can also be used to pasteurize eggs derived from the wild or another laboratory. Simple rinsing method Materials Plastic dish, (tool a, b or c in Table 1-1) Transfer pipette (tool g or h in Table 1-1) EW (appropriate volume) Method 1. Discard as much EW as possible from the dish containing eggs by decantation or use a pipette. 2. Fill the dish with new EW appropriately. 3. Repeat steps 1 and 2 for two to five times. Pasteurization method (see movie M4‐1 in the first version of this book) (Figure 1-6) Materials 500 mL scale beaker (× 5) Tea strainer (with smaller mesh than the eggs and that fits with the 500 mL beaker) (a)

(b)

(c)

(d)

(e)

Figure 1-6.  Egg cleaning (pasteurization method). (a) Set‐up for pasteurization, (b) tea strainer, (c) eggs on the tea strainer, (d) eggs in the SH‐EW, and (e) collection as the last step.

Medaka Management

13

2.5 L EW 1.5 L Sodium hypochlorite solution (SH‐EW) (1/10 000 diluted stock solution [8.5–13.5% SH] with EW) Plastic dishes Method 1. Put a series of solutions in five beakers as follows: #1 SH‐EW, #2 EW, #3 SH‐EW, #4 EW, #5 EW. 2. Transfer the eggs onto a strainer using a pipette. (minimize the volume of carry‐in culture medium). 3. Move the strainer containing eggs into #1 and keep the eggs in the solution for five minutes. 4. Move the strainer from #1 to #2 beaker and keep the eggs in the solution for five minutes. 5. Move the strainer from #2 to #3 beaker and keep the eggs in the solution for five minutes. 6. Move the strainer from #3 to #4 beaker and keep the eggs in the solution for five minutes. 7. Move the strainer from #4 to #5 beaker and keep the eggs in the solution for five minutes. 8. Move the eggs from #5 beaker to a new dish with a small amount of solution. 9. Pour new EW into the dish and culture the eggs. For more information, see Chapter 3.8 Embryo Collection in the first version of this book.

1.3.2.3  Management of embryos before hatching Separated and cleaned eggs are kept in a plastic dish at 28 °C in an incubator. The embryos should be checked every day; abnormal embryos must be removed. If there are moldy eggs in the dish, remove them and replace the contaminated dish and all the EW with a new dish and fresh EW. Although methylene blue is effective in inhibiting growth of microbes, the addition of methylene blue into the culture medium is not necessary to culture embryos (refer to the following additional information). The rate of embryo development depends on the temperature. At temperatures ranging from 25 to 30 °C, embryos progress normally throughout embryogenesis and hatch in about 10 days after fertilization. At temperatures higher than 32 °C, the risk of abnormal development increases and the expression of sex‐ related genes is also affected, resulting in production of reverse‐sex individuals (Sato et al. 2005). At temperatures lower than around 10 °C, the developmental process will not be completed and the embryos will die. In order to ensure healthy development of the embryos, it is recommended that they are cultured at a constant temperature of 28 °C. Under this condition, the embryos will hatch within nine days after fertilization. Moderate increases or decreases in temperature can lead to faster or slower development, respectively. Just before hatching, the embryo secretes a hatching enzyme from a gland located in its mouth cavity. This enzyme decomposes the egg envelope (refer to 6.1.3. Hatching Gland and 6.2. Medaka EGG Envelope and Hatching Enzyme in the first version of this book). It is better to incubate the embryos, as well as to rear fish, under a 14L–10D lighting cycle because the light stimulation triggers the secretion of the hatching enzyme. After hatching,

14

Medaka

the hatching enzyme diffuses into the medium and acts on other embryos which are not ready to hatch because of their earlier developmental stage. Therefore, mixing embryos of totally different developmental stages should be avoided, otherwise the less mature embryos will come out of the egg envelope. Within a day after hatching, the larva should be transferred into a larval culture tank (refer to section 1.3.2.4). Additional Information: Use of Methylene Blue in the Embryo Culture Methylene blue is advantageous in the embryo culture for two reasons. 1. The reduction of microorganisms (protoctista) and fungi. The chemical structure of MB creates a clear blue color when dissolved in water and absorbs visible light to generate peroxides (reactive oxygen species). MB penetrates microbes through plasma membranes and works as a sterilization agent generating peroxides. For medaka embryo culture, 0.0001% MB solution diluted with EW (MB‐EW) is often used instead of EW. 2. Labeling of dead embryos. Living embryos can actively excrete MB that flows into the  chorions from the culture medium. On the other hand, in dead embryos MB is ­accumulated and the dead embryos look blue (arrows in Figure 1-7). This phenomenon makes it easy to distinguish dead embryos from living ones and is helpful when removing dead embryos from culture dishes. However, since MB generates reactive oxygen species, its use must be avoided in some experiments, such as those using fluorescent observations and pharmacological evaluation of reagents in embryos.

1.3.2.4  Rearing from the larval stage to adulthood (to induce earlier maturation) Scientific studies need to be reproduced even in aspects such as the growth curve, which is the record of body weight and body length congruent with the age of the animal. Controlling the breeding conditions (for example, frequency of feeding during a 24‐hour period, the kind of food, the size of the tank, and so on) can achieve reproducibility with few errors, resulting in the creation of the required number of fish of the same age that are almost the same size.

Figure 1-7.  An easy method to identify dead eggs using MB.

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15

Furthermore, rapid growth and maturation of medaka are required in studies dealing with multigenerations, for example, evaluating the effects of chemicals in subsequent generations and establishing homozygous transgenic strains which is usually completed in the F2 generation. In our experience, the history of nutrition (history of growth rate) in the larval stage and the history of breeding density strongly affect the growth rate in later stages and the rate of sexual maturation. Here we provide a description of the breeding method that will achieve rapid growth as the result of an ideal feeding program. The key to this method is to create an environment where the fish can eat constantly. For this purpose, there are two important points: one is feeding in a proper way and the other is maintaining good water quality. In this subsection, an ideal feeding program (how to feed at each growth stage of medaka) is described. The way to maintain water quality is described in section 1.3.3. There is a good correlation between an ideal feeding program and growth in the length and weight of the fish. The feeding program is also important to achieve the maximum survival ratio and early maturation at two month post hatch. In this subsection, feeding is separated into five stages, depending on the amount of growth. 1. Primary larval stage (25 mm). Choice of food for larvae and adult medaka It is important to obtain an appropriate diet that contains the nutrients necessary for optimal growth and also fits into the different sizes of the gape of the growing fish. Paramecia and brine shrimp provide lower nutrition than commercial powdered food (Goolish et al. 1999; Moren et al. 2006; Carvalho et al. 2006), but still lead to normal growth of medaka. The size of paramecia is smaller than the size of the gape at each stage. It is better to change the diet from paramecia to brine shrimp as soon as possible in order for the fish to grow faster because brine shrimp provide higher nutrition than paramecia (Goolish et  al. 1999). Powdered food can be used instead of live food (for example, paramecia and brine shrimp) through all stages but leftover food should be removed using a pipette or the flow of the water recycling system. Furthermore, the different sizes of powdered food should be used according to each growth stage of the fish (Table 1-2). In the primary larval stage, it is better to mash the smallest commercial foods using a mortar and muddler and then feed the fish with the much smaller powders (2 hours. 5. Dilute an appropriate amount (depending on probe) of DIG‐ (or fluorescein‐) labeled probe with 200 μL of HB. 6. Heat the diluted probe at 80 °C for 10 minutes and quickly chill on ice. 7. Remove the HB from the tubes containing the embryos and add the denatured probe to the tubes. 8. Hybridize at 65 °C for >15 hours. 9. Preheat the following washing reagents (see below) to 65 °C. 10. Wash the embryo in the tube with the following solutions (2 mL each) at 65 °C. 11. Wash twice with 50% formamide/2× SSC* (300 mM NaCl, 30 mM Na‐citrate) containing 0.1% Tween‐20 for 30 minutes each time. 12. Wash twice with 2× SSC containing 0.1% Tween‐20 for 15 minutes each time. 13. Wash twice with 0.2× SSC containing 0.1% Tween‐20 for 30 minutes each time.

4.4.4  Immunoreaction and washing antibodies 4.4.4.1  Procedure 4 1. Quickly rinse embryo with 1× PBST. 2. Quickly rinse with 300 μL of 0.2% blocking reagent (Roche)/1× PBST. 3. Incubate in 300 μL of 0.2% blocking reagent (Roche)/1× PBST for >2 hours. Quickly rinse with 500 μL of 1/7000 anti‐DIG alkaline phosphatase (AP) (Roche)/0.2% blocking reagent/1× PBST. 4. Incubate in 500 μL 1/7000 of anti‐DIG AP/0.2% blocking reagent/1× PBST at 4 °C overnight. 5. Transfer embryos to a new six‐well plate. 6. Quickly rinse with 1× PBST. 7. Wash six times with 4 mL of 1× PBST for 10 minutes each time. 8. Wash twice with 4 mL of AP buffer for five minutes each time. Stock 1 M Tris–HCl, pH 9.5 5 M NaCl 1 M MgCl2 H2O Total

Final concentration (mM) 100 100  50

Volume (mL) 5.0 1.0 2.5 41.5 50

4.4.5 Staining 4.4.5.1  Procedure 5 1. Add 3.5 μl NBT (100 mg/mL; Roche) and 4.5 μL BCIP (50 mg/mL, Roche) per 1 mL of AP buffer.

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2. Add 3 mL of this NBT/BCIP solution to the embryo. 3. Keep in the dark as much as possible and allow the color reaction to proceed until the signal is strongest without producing background staining. 4. Stop the staining by washing five times with PBST (four quick washes, then one five‐ minute wash). The following procedures are necessary to remove the background signal. Note, however, that longer treatment may remove a weak positive signal. 5. Add 1× PBS containing 1% Tween‐20 and keep overnight at RT (do not agitate). 6. Rinse with 1× PBST.

4.5 ­Embedding in a Plastic Resin (Technovit 7100) This procedure can be applied to WISH samples that have a chromogenic reaction.

4.5.1  Equipment and reagents TC65 tungsten carbide disposable blades and a microtome (Leica, Germany) Technovit 7100 (Heraeus Kulzer, Germany) Bacto agar (BD, NJ, USA) 0.2 ml polymerase chain reaction (PCR) tube Plastic Petri dish Glass vial (5 mL) 70%, 90%, 95% ethanol Entelan New (mounting media, Merck) 1× PBST Solution I: Technovit 7100 100 mL plus Hardener I (1 g) Solution II (should be freshly prepared): add Solution I/Hardener II in a 15:1 ratio. Mix well.

4.5.2  Agarose mounting (Figure 4-68) 4.5.2.1  Procedure 1 1. Drop 2% Bacto agar solution (dissolved in water) onto a plastic Petri dish. 2. Transfer the embryo to the agar solution in the smallest possible amount of 1× PBST. 3. Orient the embryo as desired. 4. After the agar hardens, cut the agar block containing the embryo to fit into a PCR tube. 5. Transfer the embryo in agarose to a glass vial.

4.5.3  Dehydration and infiltration (Figure 4-68) 4.5.3.1  Procedure 2 1. Add 70% ethanol to the glass vial containing the embryo and incubate at RT with gentle shaking for two hours.

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Agar block containing embryo

Embryo

Dehydration & infiltration in a glass vial.

cut 2% bacto agar

Glass vial. Cut agarose to fit PCR tube Plastic Petri dish 0.2 ml PCR tube

Cut shoulder Overlay mineral oil Solution II

Section

Take out

Embryo

Fit to the tip of a PCR tube Pap pen

Pap pen

Degassed water

Slide glass

Figure 4-68.  Method for preparing sections in plastic resin (Technovit 7100).

2. Incubate in 95% ethanol at RT with gentle shaking for two hours. 3. Incubate in 100% ethanol at RT with gentle shaking for one hour. 4. Incubate in EtOH/Solution I (1/1) at RT with gentle shaking for two hours. 5. Incubate in Solution I overnight.

4.5.4  Polymerization (Figure 4-68) 4.5.4.1  Procedure 3 1. Transfer the embryo into a PCR tube. 2. Add freshly prepared Solution II. 3. Overlay with mineral oil. 4. Incubate at RT for one hour. 5. After the polymer has hardened, hold the plastic‐embedded block to a microtome. 6. Cut the corners of the block to indicate their orientation (see below). 7. Slice the embryo blocks at the desired thickness (usually 4–10 μm; 4 μm is the easiest thickness to cut) using Leica TC65 tungsten carbide disposable blades.

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Neural crest Neural tube Dermamyotome Sclerotome Notochord

Aorta

Figure 4-69.  Migration pathways for neural crest cells during embryogenesis. Black, dorsolateral pathway; gray, ventromedial pathway.

8. Line the slide (see Figure 4-68). 9. Float the sections in degassed water on the slide. 10. Orient the sections as shown in Figure 4-68. 11. Dry the sections on a warm plate at 40 °C overnight. 12. Mount with Entelan New.

Column 4.3  Pigment Cells (Figure 4-69) The body surfaces of animals are often pigmented with species‐specific colors and patterns. The cells responsible for the body coloration in vertebrates are called chromatophores, which are distributed in the skin and contain visible pigments or light‐reflecting substances (see section 3.1.2 for more information). The chromatophores develop from the neural crest, the fourth germ layer of vertebrates (Ota et al. 2007). The neural crest cells arise along the dorsal neural tube and differentiate into various cell types, including chromatophores, retinal pigment epithelial cells, neurons, glia connective tissue, and sympathoadrenal cells (Knecht and Bronner‐Fraser 2002). The precursors of chromatophores are called chromatoblasts, which migrate extensively along the dorsolateral pathway (between the ectoderm and somites in the trunk; see Figure 4-69) before colonizing at the final destination in the skin. If this migration does not occur correctly, white/colorless patches of skin or hypopigmentation of the entire surface of the body will occur (e.g., piebaldism Waardenburg syndromes). Because some chromatophore genes also regulate other cells derived from the neural crest, individuals with these mutations often have other defects (e.g., mental retardation, deafness, and colonic aganglionosis) (Bennett and Lamoreux 2003).

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Whereas mammals have only one type of chromatophore (melanocytes), up to five types are known in lower vertebrates: melanophores, leukophores, erythroxanthophores, cyanophores, and andiridophores. In fish, the homologs of some mammalian melanocyte genes that control the development of melanophores also affect other types of chromatophores. For example, the zebrafish colorless mutant (Sox10−) loses all chromatophores and the nacre (Mitf−) mutant has no melanophores, a decreased number of xanthophores, and an increased number of iridophores (Lister et al. 1999; Dutton et al. 2001). A few genes (e.g., Csf1r, somatolactin) that are not involved in pigmentation in mammals have been found to regulate chromatophore development in fish (Parichy et al. 2000; Fukamachi et al. 2004). These examples indicate that orthologous genes do not necessarily have identical functions, which is not surprising considering the highly divergent color patterns on the bodies of vertebrates. A simple model has been proposed in order to explain the development and evolution of these various color patterns: the reaction–diffusion system (Kondo 2002). Genes that might be involved in this mechanism are being identified via studies of zebrafish mutants (Watanabe et al. 2006). Zebrafish provide a good opportunity to study stripe formation (Parichy 2006). Because medaka do not have stripes, it would be very interesting to assess the functions of the zebrafish stripe formation genes in medaka. The results of such a comparative study would provide important clues to understanding how genes have evolved to achieve organismal diversity.

Column 4.4  Kupffer’s Vesicle Kupffer’s vesicle is a fish‐specific organ that is transiently attached to the tailbud and appears as a clear spherical bubble (Figure 4-70a,b, arrowhead). Research on zebrafish and medaka has revealed that Kupffer’s vesicle is functionally equivalent to the node in mice, in that they both establish the L–R axis (Essner et al. 2005; Kramer‐Zucker et al. 2005; Okada et al. 2005). Motile monocilia extend from cells lining Kupffer’s vesicle (Figure 4-70e) and create directional fluid flow. Disruption of cilia movement perturbs L–R development, suggesting that the Kupffer’s vesicle cilia play a similar role in establishing the L–R axis as do node monocilia in mice. In the medaka embryo, Kupffer’s vesicle first appears at stage 16/17.Thevesicle appears on the ventral side, near the posterior end of the embryo, and results from the consolidation of a number of small vesicles (Figure 4-70c,d, arrowhead) (S. Takashima, unpublished observation). The vesicle increases in size up to stage 22 (Figure 4-70a,b, arrowhead). When fully grown, the vesicle is almost spherical in shape and has a diameter of about 100–150 μm. The roof of Kupffer’s vesicle consists of epithelial columnar cells, each of which possesses a long single cilium (monocilium) (Figure  4-70e,f) (Hojo et  al. 2007). From stage 22 onward, the vesicle decreases in size and can no longer be seen at stage 25.

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(a)

(d)

(b)

(e)

(f)

(c)

NT

A KVE

KV

PSM

P

Figure 4-70.  Kupffer’s vesicle. (a) Lateral view at stage 22. (b) Dorsal view at stage 22. (c) Dorsal view at stage 16/17. (d) Posterior view at stage 16/17. (e) Monocilia in Kupffer’s vesicle, stained with antiacetylated‐α‐tubulin antibody. (f) Cross‐section of Kupffer’s vesicle. NT, neural tube; NC, notochord; KV, Kupffer’s vesicle; KVE, KV epithelium; PSM, presomitic mesoderm. KV is indicated by an arrowhead in (a–d). Source: Reproduced from Hojo et al. (2007) with permission of John Wiley & Sons, Inc., and S. Takashima, unpublished data.

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Chapter 5

Reproductive Behavior of Wild Japanese Medaka

5.1 ­Wild Japanese Medaka The wild Japanese medaka is a freshwater fish inhabiting the marshes, ponds, and ­irrigation canals of rice fields. Genetically, two distinct populations of medaka have been recognized in Japan: the northern population, Oryzias sakaizumii, and the southern population, Oryzias latipes (Asai et  al. 2011). Many artificial varieties have been ­cultured in Japan; one of the artificial varieties, an orange‐red variety that is a color mutant of O. latipes, is widely used for research and education, and as aquarium pets. The population of wild Japanese medaka has decreased because of changes in their aquatic environment caused by human activity, such as the modification of river and pond conformations, water pollution, and the introduction of invasive species. Also, less attention has been paid to wildlife by Japanese biologists and ordinary people. These two species of medaka have been listed as “Vulnerable” in the Red List of Threatened Animals of Japan (Ministry of the Environment 2019). Therefore, the conservation of wild Japanese medaka and their habitat is one of the most urgent conservation concerns in Japan. Although there are a large number of studies using the orange‐red type of medaka as a model animal, biological studies on wild Japanese medaka are quite limited. For the efficient conservation of Japan’s native medaka species, information on the ecology of wild medaka, especially the reproductive ecology, is essential. However, until recently there have been no reports on how wild medaka spawn in their natural environment. Wild Japanese medaka spawn seasonally from spring to autumn under natural conditions (Awaji and Hanyu 1987). Gonadal maturation and the subsequent spawning of medaka are initiated in spring by the rise of water temperature (over 12 °C) regardless of the length of photoperiod (Awaji and Hanyu 1988, 1989a). Based on studies with the orange‐red type of medaka, it is considered that female medaka ovulate and spawn almost every day during the spawning period. Interestingly, the shorter photoperiod in autumn suppresses the gonadal activity and terminates the spawning season even if the water temperature is warm enough for spawning (Awaji and Hanyu 1989b). The neural mechanism instigating the medaka’s response to this seasonal reduction of the p­ hotoperiod is unknown.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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5.2 ­Reproductive Behavior of Wild Medaka In order to conserve wild medaka, it is necessary to know how they reproduce and what types of environment should be protected for their reproductive activity. We have been studying the reproductive behavior of wild Japanese medaka, O. latipes, in outdoor ponds and rivers where they have been reproducing naturally for generations, and we were able to successfully observe their reproductive behavior in outdoor ponds for the first time (Kobayashi et al. 2012). We also compared the reproductive behavior of wild medaka with that of orange‐red medaka to confirm whether the behaviors of the orange‐red medaka in the laboratory are biologically normal or artificially induced. Our observations of the reproductive behavior of wild medaka have been mainly conducted in two artificial outdoor ponds where wild medaka are released and have reproduced for several generations without any human care (Kobayashi et al. 2012; Iwata et al. 2015). Based on an ethogram of the orange‐red medaka reported by Ono and Uematsu (1957, 1968), we made an ethogram of wild Japanese medaka (Figure 5-1). There are many ethological studies of the reproductive behavior of the orange‐red medaka, and each researcher uses different words for the repertoire of its behavior (Ono and Uematsu 1957, 1968; Hamilton et al. 1969; Walter and Hamilton 1970; Uematsu 1990; Grant and Green 1994; Grant et al. 1995; Howard et al. 1998; Koya and Onchi 2002; Kitamura and Kobayashi 2. Following

1. Approaching

4. Quick circle 6. Contact & Wrapping (Head up I)

(Head up II)

5. Floating 7. Quivering Egg release Sperm release

3. Positioning 8. Leaving

Figure 5-1.  Sequence of spawning behavior of wild medaka modified from Kamide et al. (2016). The dark color of the ventral fin is a secondary sexual characteristic of wild males. See text and Table  5-1 for details. Source: Kamide et  al. (2016). Reproduced with permission of the Hiraoka Environmental Science Laboratory.

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2003; Oshima et al. 2003; Kinoshita et al. 2009; Hayakawa et al. 2012; Nakao and Kitagawa 2015). In this chapter, we propose establishing a database of common terminology for the repertoire of reproductive behaviors exhibited by medaka among biologists. The reproductive behavior of wild medaka consists of three parts: (i) aggressive behavior by a male toward other males to increase chances to mate with females, (ii) spawning behavior for fertilization by male and female, (iii) egg deposition behavior by spawned female to deposit fertilized eggs on the substrate. As a daily cycle, the reproductive behavior of wild medaka starts at dawn. Males perform spawning behavior with females or aggressive behavior toward other males. Spawning by males and females occurs in the morning and after spawning, eggs adhere to the female’s vent by filaments for a few hours. Females deposit fertilized eggs on the substrate such as aquatic vegetation from morning to early afternoon. In the late afternoon until dusk, medaka seldom show reproductive behavior and aggressive behavior and often display schooling and/or aggregation. No reproductive behavior or aggressive behavior was observed during hours of darkness.

5.2.1  Aggressive behavior Male medaka show aggressive behavior in the morning during the spawning period (Kobayashi et al. 2012). Since this aggressive behavior seems not to be related to social dominance or defending the territory, we think it is a part of reproductive behavior for males to increase their chances to spawn with females (Iwamatsu 2006). We observed three types of aggressive behaviors: chase, attack, and disruption. Males search for females in order to spawn with them, but when a male encounters other males, he chases and repels them for a distance of 20–30 cm (chase). Sometimes males attack by making contact by butting or hitting other males with their tails (attack). Nonpaired males often disrupt paired males and females for spawning by accessing the pair (disruption).

5.2.2  Spawning behavior Repertoire of spawning behavior of wild medaka is shown in Figure  5-1 and listed in Table 5-1 (Kobayashi et al. 2012). The sequence of spawning behavior consists of the following activities. A male medaka approaches a female (approaching) and swims behind her (following). Then the male keeps his distance slightly below the female (positioning). If the female rejects the male (head‐up I), the female escapes from the male by swimming away. When the female accepts the male, the male performs a typical courtship behavior called the “quick circle” in which he swims from a lateral position in an arc in front of the female (Grant and Green 1994) (Figure 5-2a). After the quick circle by the male, when the female rejects the male (head‐up II), the male tends to repeat the “quick circle” several times, and if the female accepts the male, the sequence of the behaviors extends to further steps. The male floats up from beneath the female (floating). Then, the male and the female contact their urogenital pores (contact), and the male wraps the female with his dorsal fin and anal fin (wrapping). The male and the female release their gametes (sperm release and egg release) by quivering their body (quivering). The male adopts an S‐shaped posture during the early phase of quivering (Figure  5-2b) and then, the male and the

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Table 5-1.  Reproductive behavior of wild Japanese medaka, Oryzias latipes. Aggressive behavior Chasing Attack Disrupting

Actions of male to male Male repels other males Butting or hitting with tail Disruption of spawning by nonpaired male

Spawning behavior Approaching (1)a Following (2) Positioning (3) Head‐up I Quick circle (4) Head‐up II Floating (5) Contact (6) Wrapping (6) Quivering (7) Egg release Sperm release Leaving (8) Sneaking

Actions by male and female for fertilization Access of male to female Male swims behind female Male keeps a certain distance not too far but not too close to the female Rejection of male by female Male swims in a circle in front of female Rejection of male by female Male floats up from beneath female Contact of urogenital pores of male and female Male wraps female with dorsal and anal fins Male and female quiver their bodies to release gametes

Egg deposition behavior Picking Attaching

Actions by female to attach fertilized eggs on substrate Female picks at substrate with snout Female brushes eggs off onto substrate

Egg discarding behavior Rubbing on Head‐up III Shedding

Actions by female to detach eggs Female detaches eggs by rubbing her body on the bottom of the vessel Detachment of eggs with quick movement Female shakes her body to detach eggs

Others School Aggregation

Gathering with some interaction among individuals Gathering without interaction among individuals

Male leaves female Sperm release by nonpaired male

a The numbers correspond to those in Figure 5-1.

female place some distance between their heads during the late phase of quivering, making a V‐shaped ­position (Figure  5-2c). After the gametes have been released, the male leaves the female (leaving). After spawning, fertilized eggs adhere to the female’s vent by means of filaments for a few hours until she deposits the eggs onto the substrate (Figure 5-2d). No special type of substrate is necessary to accommodate this spawning behavior. There seems to be no preference for a specific water depth for the spawning behavior, but there is a tendency for a spawning pair to sink down slowly to the deeper water column during quivering. In orange‐red medaka, it is reported that there are actions of rejection by females called head‐up I and head‐up II (Ono and Uematsu 1957; Walter and Hamilton 1970). These are elevations of the rostral part of the body. The action after male positioning is called head‐up I and that after quick circle is called head‐up II. However, these behaviors seem to be very subtle and we could not confirm these behaviors in wild medaka.

Reproductive Behavior of Wild Japanese Medaka

(a)

(b)

(c)

(d)

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Figure 5-2.  Spawning behavior of wild medaka in outdoor ponds. (a) Quick circle (male courtship). Male swims from lateral position in an arc in front of a female. Yellow arc indicates the trace of male’s movement. (b) Early phase of quivering. Male shows S‐shaped posture. (c) Late phase of quivering. Fish make a V‐shaped position. (d) A spawned female with fertilized eggs adhering to her vent. Red arrow, female; blue arrow, male; white arrowhead, fertilized eggs. See text and Table 5-1 for details. Source: (a,d) Iwata et al. (2015). Reproduced with permission of the Hiraoka Environmental Science Laboratory. (b,c) Kobayashi et  al. (2012). Reproduced with permission of the Japanese Society of Fisheries Science.

Sneaking (parasitic male mating behaviors) (Koya et al. 2013) is known as an alternative reproductive tactic males use to increase their reproductive success (Grant et al. 1995). Koya et  al. (2013) report two types of sneaking in orange‐red medaka: (i) an unpaired male (sneaker) rushes toward a pair of quivering male and female, and emits sperm directed toward the paired female on the opposite side from the paired male at the same time as the paired male is emitting his sperm (simultaneous emission); (ii) just after spawning by a paired male and female, an unpaired male (sneaker) rushes toward the spawned female and emits sperm (postspawn emission). We have observed the second type of sneaking by wild medaka in a pond, indicating that sneaking is a naturally occurring reproductive behavior of wild medaka and not an abnormal behavior by the orange‐red medaka under stressed conditions in the laboratory. The body size of the sneakers was not smaller than that of paired males (Kobayashi, unpublished observation). Koya et al. (2013) report that the body sizes of

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sneakers of the orange‐red medaka do not differ from those of paired males and that one individual male was observed to play the roles of a paired male and the two types of sneaker in a laboratory experiment. Although we have not observed the simultaneous emission type of sneaking, it may be possible that the two types of sneaking are performed by wild medaka.

5.2.3  Egg deposition behavior After spawning, females with fertilized eggs deposit the eggs onto various types of substrate, such as aquatic plants or aquatic moss (Figure 5-3) (Kobayashi et al. 2012). The female picks at the substrate with its snout a few times to determine whether the substrate is appropriate for the deposition of eggs (picking). Some females skip the picking at egg deposition. Wild medaka seem to prefer stable substrates with soft surfaces. If the substrate moves due to a female’s picking, she does not try to attach the eggs onto that substrate. After picking, the female brushes the eggs off with a quick turn near the substrate (attaching). A mass of eggs is assembled and adheres to the female’s vent with filaments and the filaments are severed by this action. The female repeats the picking and attaching many times so that the eggs are attached onto the substrate little by little. Egg deposition behavior continues until all the eggs are attached to the substrate. The egg deposition behavior of females is often disrupted by males as males perform courtship behaviors directed to females with eggs or it may be possible that the male is trying to eat the eggs. The sites of egg deposition are mostly located on the underwater banks of ponds or rivers with slow or no water flow (0–3.0 cm/s) and near the surface of the water (0–15 cm depth) where appropriate aquatic vegetation grows for egg deposition (Kobayashi et  al. 2012; Iwata et al. 2015). Female medaka do not deposit eggs in deep areas even if aquatic plants appropriate for egg deposition grow there.

5.2.4  Egg discarding behavior Ono and Uematsu (1957) and Kinoshita et al. (2009) describe that spawned female orange‐red medaka with fertilized eggs detach the eggs in three ways: rubbing on, head‐up III, and ­shedding. These actions are often observed in the orange‐red medaka in a laboratory vessel when there is no appropriate substrate for egg deposition (Kamide et  al. 2016) (Figure  5-4, Table 5-1). 2. Attaching 1. Picking

Figure 5-3.  Egg deposition behavior. Picking and attaching. See text and Table  5-1 for details. Source: Kamide et al. (2016). Reproduced with permission of the Hiraoka Environmental Science Laboratory.

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Shedding Rubbing-on

Figure 5-4.  Egg discarding behavior in an aquarium in the absence of substrate for egg deposition. Rubbing on and shedding. See text and Table 5-1 for details. Source: Kamide et al. (2016). Reproduced with permission of the Hiraoka Environmental Science Laboratory.

A female medaka with eggs picks the bottom or the wall of the vessel with her snout and rubs the ventral side of her body against the vessel to detach the eggs by cutting the filaments (rubbing on). In the absence of substrate for egg deposition, sometimes the female keeps the eggs attached to her body for a longer time. In that situation, the filaments elongate and some of the eggs dangle from the fish. The female in this condition tries to cut the filament to detach the eggs using quick body shaking (head‐up III and shedding). Higher frequency of body movement is called shaking (Ono and Uematsu 1957). Since these three types of egg detaching actions have not been observed in wild medaka, they are considered to be induced under artificial conditions. Females with elongated filaments have not been observed in wild medaka (Kobayashi et al. 2012). Interestingly, when wild female medaka with eggs were placed in an experimental vessel without substrate for egg deposition, they performed rubbing on and shedding (Kamide et al. 2016). Since eggs that have been detached with these actions do not attach to any part of the vessel but stay on or sink to the bottom, it is considered that these three actions are a kind of egg discarding behavior in preparation for spawning of the following morning.

5.2.5  School and aggregation Medaka form schools and aggregations. A school is a gathering of fish with some interaction among individuals. An aggregation is a gathering of fish but there is no special interaction among individuals (Kobayashi et al. 2012). In medaka, a school is mostly observed during the nonspawning seasons and during the afternoons in spawning season when spawning and aggressive behaviors are not performed. Aggregation was sometimes observed during the day in spawning season. In observations of wild medaka, when an observer comes close to the water, the medaka often escape in the same direction as a group. This behavior is considered to be an aggregation of escape rather than schooling behavior.

5.3 ­Conclusion It is important to know the reproductive behavior of wild medaka in field conditions since it is natural behavior acquired during evolution. When the orange‐red medaka and other artificial varieties are used for laboratory experiments, it is essential to understand whether

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the observed behaviors are the natural behavior of medaka or nonnatural behavior induced under artificial laboratory conditions. Also, in order to conserve wild medaka, more information about their reproductive ecology is indispensable. In addition to a decrease in population of wild Japanese medaka, a recent problem in their conservation is a genetic disturbance caused by the introduction of the orange‐red medaka to the natural environment of Japan (Nakai et al. 2011; Yokota et al. 2014; Nakao and Kitagawa 2015; Nakao et al. 2017). Genetic introgressions from the orange‐red medaka into the wild medaka populations were confirmed in wild medaka caught in the natural environment. For the conservation of the natural genetic diversity of wild Japanese medaka, field study is an urgent concern.

­References Asai, T., Senou, H., and Hosoya, K. (2011). Oryzias sakaizumii, a new ricefish from northern Japan (teleostei: Adrianichthydae). Ichthyol. Explor. Freshw. 22: 289–299. Awaji, M. and Hanyu, I. (1987). Annual reproductive cycle of the wild type medaka. Nippon Suisan Gakkaishi 53: 959–965. Awaji, M. and Hanyu, I. (1988). Effects of water temperature and photoperiod on the beginning of spawning season in the orange‐red type medaka. Zool. Sci. 5: 1059–1064. Awaji, M. and Hanyu, I. (1989a). Seasonal changes in ovarian response to photoperiods in orange‐red type medaka. Zool. Sci. 6: 945–950. Awaji, M. and Hanyu, I. (1989b). Temperature‐photoperiod conditions necessary to begin the spawning season in wild type medaka. Nippon Suisan Gakkaishi 55: 747. Grant, J.W.A. and Green, L.D. (1994). Mate copying versus preference for actively courting males by female Japanese medaka (Oryzias latipes). Behav. Ecol. 7: 165–167. Grant, J.W.A., Bryant, M., and Soos, C. (1995). Operational sex ratio, mediated by synchrony of female arrival, alters the variance of male mating success in Japanese medaka. Anim. Behav. 49: 367–375. Hamilton, J.B., Walter, R., Daniel, R. et al. (1969). Competition for mating between ordinary and supermale Japanese medaka fish. Anim. Behav. 17: 168–176. Hayakawa, Y., Takita, S., Kikuchi, K. et al. (2012). Involvement of olfaction in spawning success of medaka Oryzias latipes. Jpn. J. Ichthyol. 59: 111–124. (in Japanese). Howard, R.D., Martens, R., Innis, S. et al. (1998). Mate choice and mate competition influence male body size in Japanese medaka. Anim. Behav. 55: 1151–1163. Iwamatsu, T. (2006). The Integrated Book for the Biology of the Medaka. Tokyo: Daigaku Kyoiku Shuppan (in Japanese). Iwata, E., Sakamoto, K., Ohkouchi, T. et al. (2015). Egg deposition sites of the wild minami‐medaka Oryzias latipes in an aquatic area of a biotope. Nat. Environ. Sci. Res. 28: 11–21. (in Japanese). Kamide, S., Shimizu, A., Koido, M. et al. (2016). Environmental conditions suitable for egg deposition of female medaka, Oryzias latipes. Nat. Environ. Sci. Res. 29: 31–39. (in Japanese). Kinoshita, M., Murata, K., Naruse, K. et al. (2009). Medaka: Biology, Management, and Experimental Protocols. Chichester: Wiley‐Blackwell. Kitamura, W. and Kobayashi, M. (2003). The effect of water flow on spawning in the medaka, Oryzias latipes. Fish Physiol. Biochem. 28: 429–430. Kobayashi, M., Yoritsune, T., Suzuki, S. et  al. (2012). Reproductive behavior of wild Japanese medaka in an outdoor pond. Nippon Suisan Gakkaishi 78: 922–933. (in Japanese). Koya, Y. and Onchi, R. (2002). Observation method for mating and spawning behavior in medaka fish. Sci. Rep. Fac. Educ. Gifu Univ. (Nat. Sci.) 26: 19–22. (in Japanese). Koya, Y., Koike, Y., Onchi, R. et al. (2013). Two patterns of parasitic male mating behaviors and their reproductive success in Japanese medaka, Oryzias latipes. Zool. Sci. 30: 76–82.

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Ministry of the Environment (2019). Revision of the Red List of Freshwater and Brackish Water Fish. Tokyo: Ministry of the Environment, Government of Japan (in Japanese). Nakai, K., Nakao, R., Fukamachi, S. et  al. (2011). Genetic analysis of Medaka (Oryzias latipes) populations in the Yamato River, Nara Prefecture, Japan: detection of the b allele responsible for the himedaka phenotype. Jpn. J. Ichthyol. 58: 189–193. (in Japanese). Nakao, R. and Kitagawa, T. (2015). Differences in the behavior and ecology of wild type medaka (Oryzias latipes complex) and an orange commercial variety (himedaka). J. Exp. Zool. 323A: 349–358. Nakao, R., Iguchi, Y., Koyama, N. et al. (2017). Current status of genetic disturbances in wild medaka populations (Oryzias latipes species complex) in Japan. Ichthyol. Res. 64: 116–119. Ono, Y. and Uematsu, T. (1957). Mating ethogram in Oryzias latipes. J. Fac. Sci. Hokkaido Univ. Ser. VI Zool. 13: 197–202. Ono, Y. and Uematsu, T. (1968). Sequence of the mating activities in Oryzias latipes. Jpn. J. Ecol. 18: 1–10. Oshima, Y., Kang, I., Kobayashi, M. et al. (2003). Suppression of sexual behavior in male Japanese medaka (Oryzias latipes) exposed to 17β‐estradiol. Chemosphere 50: 429–436. Uematsu, K. (1990). An analysis of sufficient stimuli for the oviposition in the medaka Oryzias latipes. J. Fac. Appl. Biol. Sci. Hiroshima Univ. 29: 109–116. Walter, R. and Hamilton, J.B. (1970). Head‐up movements as an indicator of sexual unreceptivity in female medaka, Oryzias latipes. Anim. Behav. 18: 125–127. Yokota, H., Kuwahara, N., Nakano, E. et al. (2014). Genotype distribution in wild populations of medaka inhabiting the Muko River system and the actual state of genetic introgression from the domesticated orange‐red fish (hi‐medaka). Bull. Kansai Org. Nat. Conserv. 36: 53–58. (in Japanese).

Chapter 6

Cryopreservation and Transplantation of Medaka Germ Cells

6.1 ­Introduction Cryobanking of gametes could be an effective measure for the preservation of various genetic resources in medaka. However, past attempts at cryopreservation of fish embryos and mature oocytes have been unsuccessful as a result of their large size, high yolk content, and low membrane permeability. Although sperm cryopreservation is a powerful tool for long‐term preservation of transgenic or mutant strains of medaka, it cannot be used to preserve maternally inherited cytoplasmic compartments, such as mitochondria. More importantly, in order to produce live offspring, eggs are always indispensable. Since, by transplanting germ cells retrieved from frozen whole testes into newly hatched larvae, the recipient larvae, depending on their sex, produce either functional sperm or eggs, cryopreservation of testes can be an effective alternative for cryopreservation of medaka eggs. In this protocol, the methods of cryopreservation of whole testes, the preparation of testicular cell suspension, and their transplantation into recipients are introduced (Figure 6-1). Although the protocols introduced here still need some fine‐tuning, their use in salmonid species is well established (Lee et al. 2013) and has already been applied to several practical conservation projects.

6.2 ­Cryopreservation of Medaka Testes Since we previously found that spermatogonial stem cells possess quite a high level of sexual plasticity in several gonochoristic teleosts (Okutsu et  al. 2006; Morita et  al. 2012) and the number of transplantable stem cells is much larger than that naturally occurring in the ovaries, in this protocol the cryopreservation of whole testes is introduced.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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Figure 6-1.  Overall experimental scheme to obtain functional sperm and eggs derived from cryopreserved testes via germ cell transplantation.

6.2.1 Solutions Eagle’s medium Eagle’s minimum essential medium (Nissui) HEPES (Sigma, H4034) 1.95% l‐glutamine (Sigma, G8540) in distilled water (DW) NaHCO3 Fetal bovine serum (FBS) (Gibco) Total (pH 7.8)

4.7 g 2.38 g 7.5 mL 2.5 g 25 mL 500 mL

HEPES, N‐2‐hydroxyethylenepiperazine‐N’‐2‐ethanesulfonic acid.

10 × Extender stock HEPES (Sigma, H4034) NaCl KCl KH2PO4 Na2HPO4 Sodium pyruvate (Sigma, P8574) Total (pH 7.8)

4.61 g 7.68 g 0.19 g 1.1 g 0.19 g 0.14 g 70 mL

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2 × Extender working solution 10 × extender 1 M CaCl2·2H2O 1 M MgCl2·6H2O DW Total

294 μL 3.9 μL 2.1 μL 1.2 mL 1.5 mL

Cryomedium 2 × Extender 15% bovine serum albumin (BSA) (Sigma) 1 M trehalose Dimethylsulfoxide (DMSO) DW Total

176 μL 50 μL 50 μL 46.2 μL 177.8 μL 500 μL

6.2.2 Materials Two sharp forceps (Dumont No. 5) Dissecting scissors Twenty‐four well plates 1.2 mL cryotube (TPP Cryo Tube 12) Slow‐freezing vessel (Bicell, Nihon Freezer Co.) Deep freezer (−80 °C) Liquid nitrogen Water bath

6.2.3 Procedures Freezing of Isolated Whole Testes 1. Isolate testes from adult males by using sharp forceps. 2. Transfer the testes isolated from each medaka into 500 μLof ice‐cold Eagle’s medium filled in 24‐well plates kept on crashed ice. 3. Transfer the testis into the cryotube containing 500 μLof prechilled cryomedium. 4. Equilibrate the testicular tissue with the cryomedium by keeping it on ice for 60 minutes. 5. Transfer the cryotubes into the slow‐freezing vessel and store them at −80 °C for exactly 90 minutes. The slow‐freezing vessel should be prechilled on ice for at least 30 minutes before use. The maximum number of tubes that can fit into each slow‐freezing vessel is eight. 6. Plunge the cryotubes into liquid nitrogen. This process should be done quickly in order to avoid the increase of temperature of the samples once the temperature reaches −80 °C. 7. Store the cryotubes in a liquid nitrogen tank until needed.

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Thawing of Whole Testes 1. Set the water bath to 10 °C. 2. Put 1 mL of Eagle’s medium into 24‐well plates and chill by keeping them on crashed ice. 3. Thaw testicular tissue in the cryotubes by shaking them in the 10 °C water bath. This takes approximately 60–70 seconds. 4. Take the testicular tissue out from the cryotube and transfer to Eagle’s medium in the 24‐well plates (see step 2). 5. Rehydrate the testicular tissue in Eagle’s medium for 15 minutes. The medium should be changed every five minutes (i.e. a total of three times). 6. The testes are then ready for dissociation.

6.3 ­Transplantation of Thawed Testicular Cells into Recipient Larvae The immature germ cells (most likely spermatogonial stem cells) that are transplanted into the peritoneal cavities of larvae around the time of hatching can migrate to the recipient gonads by following the migration pathways of the endogenous primordial germ cells (Okutsu et al. 2006) and begin either spermatogenesis or oogenesis, depending on the sex of the recipient larva. Since this method is just transplanting dissociated testicular cells into peritoneal cavity of the recipient larvae, it is quite simple and not time‐consuming. In this section, the protocol to prepare the donor cell suspension using frozen whole testes and their transplantation method will be introduced.

6.3.1 Solutions Collagenase solution: Collagenase H (Roche, 11 074 059 001), 40 mg/mL in phosphate buffered saline (PBS) Dispase solution: Dispase II (Wako Chemicals, GD81070), 33.3 mg/mL in PBS DNase solution: deoxyribonuclease I from bovine pancreas (Sigma‐Aldrich, D5025), 15 units/μL in PBS Penicillin solution: benzylpenicillin potassium (Wako, 023‐07731), 50 units/μL in PBS Streptomycin solution: streptomycin sulfate (Wako, 196‐08511), 50 mg/mL in PBS Ampicillin solution: ampicillin sodium (Wako, 018‐10372), 50 mg/mL in PBS. Dissociation medium

PBS(+) FBS Collagenase solution Dispase solution DNase solution

370 μL 50 μL 25 μL 25 μL 30 μL

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L‐15/10% FBS Leibovitz L‐15 medium (Gibco, 41300–070) HEPES (Sigma, H4034) DW Total (pH 7.8) FBS (Gibco, 26140–079) Penicillin solution Streptomycin solution Ampicillin solution

1.374 g 0.6 g 90 mL 10 mL 100 μL 100 μL 100 μL

Medaka balanced salt solution (BSS) 0.65% 0.04% 0.02% 0.02% 0.001%

NaCl KCl MgS04 7H2O CaCl2 2H2O Phenol red

Sterilize and adjust to pH 8.3 with 5% NaHCO3.

3% agarose: suspend 3 g of agarose (Agar powder, Wako, 010–15815) in 100 mL of DW and microwave to dissolve completely.

6.3.2 Materials Sharp forceps (Dumont No. 5) Dissecting scissors Forty‐eight well plates Microtube (1.5 mL) Polymerase chain reaction (PCR) tube (0.2 mL) 42 μm pore size nylon screen (NL‐screen, 130‐035/330TW, NBC Industries) Mineral oil (Sigma, mineral oil light, 330779‐1L) Micropipette puller (PC‐10, Narishige) Micropipette grinder (EG‐400, Narishige) Micromanipulator (right hand, MP‐2R, Narishige) Microinjector (IM‐9A, Narishige) Micro capillary (GD‐100, Narishige) Plastic Petri dish (3 and 9 cm) Triploid medaka larvae (see Column 6.1 for a method to make triploids)

6.3.3 Procedures Dissociation of Whole Testes 1. Transfer the thawed testes into a well filled with 500 μL of the dissociation medium. 2. Mince the thawed testes into small pieces using a pair of scissors until each piece becomes less than approximately 1 mm3 (Figure 6-2a).

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(a)

(b)

42 μm mesh

0.2 mL PCR tube

(c)

1.5 mL microtube

(d)

Figure 6-2.  Preparation of testicular cell suspension. Testes from two adult medaka were minced in a well using a pair of scissors (a) and dissociated using collagenase and dispase. The remaining cell clumps were removed by filtration (d). The filter is made by fixing the 42 μm mesh between the PCR tube and its tube cap (c) followed by cutting the center of the tube cap and the bottom of the PCR tube.

3. Pipette 50 times very gently using 1 mL autopipette with blue tips and incubate for 30 minutes at 26 °C. 4. Pipette 50 times again and incubate for another 30 minutes at 26 °C. 5. Pipette 50 times again (Figure 6-2b) and transfer the resulting cell suspension into a microtube. 6. Add 500 μL of ice‐cold L‐15/10% FBS to each well, rinse the well carefully in order to recover all dissociated cells, and transfer to a microtube. 7. Centrifuge at 1000 rpm for 10 minutes at 4 °C. 8. Discard the supernatant slowly using a 1 mL autopipette. 9. Add 1000 μL of ice‐cold L‐15/10% FBS and resuspend the cell pellet by gently flicking the bottom of the microtube with the fingers. 10. Remove the remaining cell clumps by filtering the cell suspension with 42 μm pore size nylon screen (Figure 6-2c,d) and transfer the filtered cell suspension to a new microtube. Then centrifuge at 1000 rpm for 10 minutes at 4 °C. 11. Discard the supernatant and add 20 μL of ice‐cold L‐15/10% FBS. 12. Flick the bottom of the microtube gently in order to resuspend the testicular cells. 13. Transfer the whole cell suspension to a small Petri dish (3 cm). To prevent evaporation, cover the drop of the resulting cell suspension with mineral oil. Transplantation of Dissociated Testicular Cells into Peritoneal Cavity of Larvae 1. Transplantation micropipettes are prepared by pulling glass microcapillaries (GD‐100, Narishige Co., Japan) using an electric puller (PC‐10, Narishige).

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(a) Fishing line

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(b)

Scotch tape

3 cm Petri dish

Groove to support larvae 3% Agar

3% Agar

(d) (c)

Figure 6-3.  Preparation of a tray for testicular cell transplantation into larvae. First, a mold is made using a 3 cm Petri dish, monofilament fishing line (diameter approximately 0.5 mm), and Scotch tape (a,b). The fishing line is fixed on the outside of the Petri dish bottom using Scotch tape (b,c top). The mold is used (b) to make a groove on the surface of the agar plate (c bottom, d).

2. The tips of the micropipettes are sharpened with a grinder (EG‐400, Narishige) until their openings reach approximately 120 μm. 3. Set up a microinjector by inserting the micropipette into an injection holder and fill the micropipette with mineral oil. 4. Pour the microwaved 3% agarose into a Petri dish (9 cm) until it covers the entire ­bottom of the dish. Before solidifying the agarose, use the mold (Figure  6-3a–c) to make small grooves to hold the recipient larvae (Figure 6-3d). 5. Dilute 12 μL of 2‐phenoxyethanol with 30 mL of medaka BSS in a plastic Petri dish (9 cm). 6. Anesthetize recipient triploid medaka larvae (newly hatched) in the 2‐phenoxyethanol solution. 7. When the medaka larvae are fully anesthetized, transfer a larva to the Petri dish (9 cm) with the bottom covered with 3% agarose using a disposable pipette with a wide opening. 8. Suck up approximately 10 000 dissociated cells into the micropipettes prepared in step 1–3 (Figure 6-4a,b). 9. Inject them into the peritoneal cavity of the medaka larva through the dorsal muscle (Figure 6-4c) using a micromanipulator under a dissection microscope. 10. Transfer the recipient madaka larva into the medaka BSS to help it recover from the anesthesia.

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Micropipette Mineral oil Micropipette

Air bladder

Cell suspension Cell suspension (a)

(b)

(c)

Figure 6-4.  Intraperitoneal transplantation of testicular cells into recipient larvae. Testicular cell suspension is sucked up using a glass micropipette (a,b) and transplanted into the body cavity of newly hatched medaka larvae. The micropipette is penetrated just posterior to the air bladder (c). Approximately 10 000 testicular cells are delivered into one recipient.

Column 6.1  Production of triploid medaka Male and female medaka were separated before crossing at day 1. Males and females were crossed on day 2 and after spawning, the fertilized eggs were immediately isolated. Within 2–3 minutes post fertilization, eggs were immersed in 42 °C water for two minutes to prevent second polar body extrusion (Naruse et al. 1985). The rate of triploid formation ranged from 20% to 100%. To increase the success rate of triploid medaka production, it will be necessary to optimize the timing of the application of heat shock to the fertilized eggs. Ploidy analysis will also be essential to determine the ratio of triploidization. Interspecific crosses between 2n male Oryzias luzonensis and 2n female O. latipes are mostly lethal. F1 hybrids (2n) develop abnormally and their hatching ratio ranges between 0.0% and 2.3% (Formacion & Uwa 1985). Hybrid lethality has been observed in crossings between O. latipes females and O. luzonensis males. Other combinations among species closely related to O. latipes are generally viable. So, we believe that the use of O. luzonensis males is essential for interspecific triploid formation without ploidy analysis. Sato et al. (2001) used O. luzonensis males crossed with female O. latipes in their production of triploid hybrids. When female O. latipes (F1 hybrid between Hd‐rR and HNI) were crossed with male O. luzonensis, and then heat shock was applied to fertilized eggs using the regimen described above, 59% of the embryos hatched and all the fry were triploid (Sato et al. 2001). The surviving triploid hybrids developed into adults (Sato, personal communication). Although the hatching ratio of heat‐treated embryos may have varied, most of the hatched fry can be distinguished as triploid without performing ploidy analysis. 11. Raise the recipients as usual until they reach maturity and use them for mating. The triploid females and males usually produce some aneuploid gametes. However, the offspring produced using triploid recipient‐derived gametes cannot survive after the

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hatching stage and only the donor‐derived diploid larvae can grow up normally. It should be easy to identify the donor‐derived offspring if body color mutants or GFP (green florescence protein) transgenics are used for either donors or recipients.

­References Formacion, M.J. and Uwa, H. (1985). Cytogenetic studies on the origin and species differentiation of the Philippine medaka, Oryzias luzonensis. J. Fish Biol. 27: 285–291. Lee, S., Iwasaki, Y., Shikina, S. et al. (2013). Generation of functional eggs and sperm from cryopreserved whole testes. Proc. Natl. Acad. Sci. U.S.A. 110: 1640–1645. Morita, T., Kumakura, N., Morishima, K. et  al. (2012). Production of donor‐derived offspring by allogeneic transplantation of spermatogonia in the yellowtail (Seriola quinqueradiata). Biol. Reprod. 86: 1–11. Naruse, K., Ijiri, K., Shima, A., and Egami, N. (1985). The production of cloned fish in the medaka (Oryzias latipes). J. Exp. Zol. 236: 335–341. Okutsu, T., Suzuki, K., Takeuchi, Y. et al. (2006). Testicular germ cells can colonize sexually undifferentiated embryonic gonad and produce functional eggs in fish. Proc. Natl. Acad. Sci. U.S.A. 103: 2725–2729. Sato, T., Yokomizo, S., Matsuda, M. et  al. (2001). Gene centromere mapping of medaka sex chromosomes using triploid hybrids between Oryzias latipes and O. luzonensis. Genetica 111: 71–75.

Chapter 7

Genome Editing

7.1 ­Introduction With the advent of the omics era, reverse genetics approaches that are designed to manipulate the genetic sequence and observe the impact of genetic modification on the organism have been of increasing importance in demonstrating gene functions and the understanding of complex biological processes. Genome editing has become a versatile and powerful tool for genetic manipulation. It induces site-specific DNA double-strand breaks (DSBs) using engineered nucleases, resulting in genome modifications, such as targeted mutagenesis with a small insertion and/or deletion or targeted gene integration during the subsequent repair process. In this chapter, we describe a method for targeted genome editing in medaka using two families of nucleases, clustered, regularly interspaced, short palindromic repeats (CRISPR)/CRISPR-associated (Cas)-based RNA-guided nucleases and transcription activator-like effector nucleases (TALENs), whose target sequence can be flexibly and easily customized for any desired endogenous loci in medaka. First, we describe how to prepare the targetable endonucleaes for the endogenous genomic loci in medaka. Then, a simple method, called heteroduplex mobility assay (HMA), used to detect the mutation induced by the targetable endonucleases in treated embryos and in later generations, is described. Finally, the methods to produce knock-out (KO) and knock-in (KI) medaka strains are introduced.

7.2 ­Outline of Targeted Genome Editing Using Nucleases Genome editing is a technology used to modify genomic sequences at the targeted locus by utilizing the DSB which is induced by targetable nucleases. This technology was first proposed using zinc finger nucleases (ZFNs), which consist of an engineered zinc finger DNA-binding domain and the FokI DNA-cleavage domain (Kim et  al. 1996). Despite the widespread applicability of this technology, ZFNs have not become common for the following reasons: it is not easy to construct ZFNs that can be targeted toward the gene of interest, and the generated ZFNs do not always induce a DNA DSB on the target site.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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Transcription activator-like effector nucleases are alternative artificial nucleases that were developed by replacing the zinc finger domain of ZFNs into a DNA recognition domain of a transcription activator-like (TAL) effector found in the plant pathogenic bacteria of the genus Xanthomonas (Christian et al. 2010). The TAL effector domain contains repetitive modules of 33–35 amino acids. Two polymorphic residues at position 12 and 13 in the module, known as repeat variable do-residues (RVDs), determine the base specificity to interact (Boch et al. 2009; Moscou and Bogdanove 2009). This modularity of base recognition enables us to engineer the TAL effector domain for the sequence of interest by using a modular assembly strategy (Cermak et al. 2011), which promotes widespread usage of TALENs for targeted genome editing. The CRISPR/Cas system is a recently developed nuclease system for genome editing, which in its natural state functions as an adaptive immune system in bacteria and archaea to detect and silence foreign genetic elements such as viruses and plasmids (Wiedenheft et al. 2012). In type II CRISPR systems, a Cas9 endonuclease forms a complex with each CRISPR RNAs (crRNAs) hybridized with a trans-activating crRNA (tracrRNA). The Cas9 endonuclease then cleaves to the target DNA sequences that are complementary to the crRNAs and adjacent to short sequences known as protospacer adjacent motifs (PAMs) (Gasiunas et al. 2012; Jinek et al. 2012). A synthetic, single guide RNA (sgRNA) consisting of a fusion of crRNA and tracrRNA can also program the recognition specificity of the Cas9 nuclease from Streptococcus pyogenes. Cas9 nuclease acts as an RNA-guided endonuclease (RGEN) that has a cleavage site that can be easily altered by only the engineered 5′ sequence of the sgRNA. These targetable nucleases introduce site-specific double-strand DNA breaks that are repaired by either of two major pathways: nonhomologous end joining (NHEJ) and homology-directed repair (HDR) (Urnov et al. 2010). NHEJ joins the end of the broken DNA strands without any templates, frequently inducing targeted gene disruption with a small insertion and/or deletion (indel) around the cleaving site or a site-specific addition of tags such as GFP (green florescence protein). HDR in the presence of DNA templates with homologous sequences can repair the DSBs through the replacement of flanking homology arms, resulting in the precise integrations of a DNA fragment or a targeted gene correction. To date, these nucleases have been effective tools for targeted genomic engineering in a wide range of organisms (Peng et al. 2014). We have established an efficient method for the generation of a knock-out medaka mutant strain with NHEJ-induced small indels that have been successfully demonstrated using ZFNs (Ansai et  al. 2012), TALENs (Ansai et  al. 2013), and the CRISPR/Cas system (Ansai and Kinoshita 2014). Recently, we also established an effective knock-in method via HDR (Murakami et al. 2017). In contrast to zebrafish, in medaka HDR is preferable in the knock-in process rather than microhomology-mediated end joining (MMEJ) (Kawahara et al. 2016).

7.3 ­Preparation of CRISPR/Cas9 Genome Editing Tools This section describes how a Cas9 nuclease and engineered sgRNAs are prepared for microinjection into fertilized medaka eggs. Capped RNA encoding for a Cas9 nuclease is transcribed from an expression vector constructed for fish. The sgRNAs are engineered by cloning a pair of custom-ordered oligonucleotides and therefore are transcribed using an RNA polymerase.

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7.3.1 Materials Plasmids • pCS2 + hSpCas9 (Addgene Plasmid #51815): a Cas9 expression plasmid is used to create a capped RNA by in vitro transcription using a SP6 RNA polymerase for microinjection into fish embryos (Ansai and Kinoshita 2014). This vector contains a human codon-optimized Cas9 gene derived from Streptococcus pyogenes (Cong et al. 2013) and an ampicillin-resistant gene. • pDR274 (Addgene Plasmid #42250): an empty gRNA plasmid is used to create a targetspecific gRNA by in vitro transcription using a T7 RNA polymerase (Hwang et al. 2013). This vector contains a kanamycin-resistant gene. • Polymerase chain reaction (PCR) primers for colony PCR checking. • M13 forward primer: 5′-GTAAAACGACGGCCAGT-3′. Molecular Biology Reagents • Restriction enzymes  –  BsaI, DraI, and NotI: restriction enzymes with lower star activity are preferable. For example, BsaI-HF (high-fidelity version of BsaI, NEB, Cat. No. R3535) is more suitable for overnight digestion than BsaI without “HF” (NEB, Cat. No. R0535). • DNA ligase: ligation reagents with high efficiency (e.g., Ligation high Ver. 2, Toyobo, Cat. No. LGK-201) are recommended. • PCR enzyme: high-fidelity polymerase is not necessarily required (e.g., HybriPol DNA Polymerase, Bioline, Cat. No. BIO-21080; TaKaRa Ex Taq, Takara Bio, Cat. No. RR001A are available). • T7 RNA polymerase: a high-yield in vitro transcription kit with T7 RNA polymerase (e.g., AmpliScribe T7-Flash Transcription Kit, Lucigen, Cat. No. ASF3507) is recommended for the stable transcription of sgRNAs. However, the standard T7 RNA polymerase (e.g., Promega, Cat. No. P2075) is also available. • Capped RNA transcription kit with SP6 RNA polymerase: mMessage mMachine SP6 Transcription Kit (Life Technologies, Cat. No. AM1340) is recommended for highyield production of Cas9 RNA. • Proteinase K: prepare a 20 mg/mL of stock solution. • Plasmid purification kit: a plasmid purification kit (spin column or higher grade) should be used (e.g., NucleoSpin Plasmid QuickPure, MACHEREY-NAGEL, Cat. No. 740615). • Gel extraction kit: a gel extraction kit (spin column type) should be used (e.g., NucleoSpin Gel and PCR Clean-up, MACHEREY-NAGEL, Cat. No. 740609). • RNA extraction kit: a RNA extraction kit with spin columns (e.g., RNeasy Mini Kit, Qiagen, Cat. No. 74104) should be used for purification of transcribed RNAs. • 10× annealing buffer: 400 mM Tris-HCl (pH 8.0) containing 200 mM MgCl2 and 500 mM NaCl. • 10% sodium dodecyl sulfate (SDS). • Phenol-chloroform mixture: mixture of equal volumes of phenol saturated by TE buffer and chloroform. • Chloroform. • 3 M sodium acetate (CH3COONa) (pH 5.2). • 0.5 M EDTA (pH 8.0).

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• Co-precipitated reagent: glycogen (e.g., Roche, Cat. No. 10901393001) or other coprecipitated reagent is used for high-yield recovery of DNA in ethanol precipitation. • Ethanol (100% and 70%). • Competent E. coli: DH5α and any other competent cells can be used. • LB broth: both the LB plates and the liquid medium containing appropriate antibiotics at the concentrations indicated below should be prepared. For a liquid culture, higher nutrient medium such as 2X YT and Plusgrow II (Nacalai Tesque, Cat. No. 08202-75) is more suitable for plasmid DNA recovery with a high yield. • Ampicillin (amp): a stock solution is 50 mg/mL ampicillin sodium salt in H2O or 70% ethanol. The final concentration is 50 μg/mL in the medium. • Kanamycin (kan): a stock solution (20 mg/mL kanamycin sulfate in H2O) should be used at a final concentration of 20 μg/mL in the medium according to standard protocols.

7.3.2  Production of custom-designed sgRNA 7.3.2.1  Preparation of the bsai-digested sgRNA backbone 1. E. coli containing the pDR274 vector is spread on the LB plate containing 20 μg/mL kanamycin and is incubated at 37 °C overnight. 2. Pick an isolated colony from the plate, inoculate it into the LB or any liquid medium with 20 μg/mL kanamycin, and grow the cultures overnight at 37 °C. Prepare the amount of medium that is enough to recover more than 5 μg of plasmid DNA. 3. Purify the plasmid from the cultures using the plasmid purification kit with spin columns and then quantify the concentration of each elution with the spectrophotometer or any other method for DNA quantification. 4. Purified plasmid is digested with the restriction enzyme BsaI. Assemble the reaction as described below and then incubate at 37 °C overnight. pDR274 (5 μg) CutSmart Buffer (10×) BsaI-HF (NEB) H2O Total

X μL 5 μL 2 μL Up to 50 μL 50 μL

5. All the 50 μL of reaction is individually loaded into 1% agarose gels and separated by electrophoresis at 135 V for 25 minutes or the same separation condition. 6. Excise each DNA fragment (ca. 2.1 kb) with minimized UV exposure and then retrieve the DNA using the spin column gel extraction kit. Elute the DNA with 30 μL of the elution buffer from each column whose residual ethanol is removed by using a longer duration of centrifuge and incubating the columns for several minutes at 70 °C prior to elution. 7. Eluted DNAs are stored at −20 °C.

7.3.2.2  Design and production of customized sgRNA Day 1: Design sgRNA for medaka endogenous loci and order oligonucleotides. 1. Find the “5′-N21GG-3′” or “5′-CCN21-3′” sequence in the target locus because the sgRNA can recognize the sequence followed by a PAM (5′-NGG-3′) in both strands.

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2. Choose two or more target sites in a target locus with reference to (a) potential off-target sequences and/or (b) microhomology sequences. (a) Off-target alterations are sometimes identified in the genomic sequences harboring up to several base pairs of mismatches adjacent to a NGG PAM (Ansai and Kinoshita 2014). Target sequences with fewer off-target candidates in the medaka genome can be predicted using the CRISPR/Cas9 target online predictor (CCTop; http://crispr. cos.uni-heidelberg.de) (Stemmer et al. 2015) or CRISPRscan (www.crisprscan.org) (Moreno-Mateos et al. 2015). Potential off-target candidates of each target can also be found using the pattern match tool for CRISPR/Cas (http://viewer.shigen.info/ medakavw/crisprtool). (b) We previously found that microhomologous sequences striding over the DSB point frequently induce specific patterns of deletion between the homologous sequences (Ansai et al. 2014). Target candidates with microhomologies whose deletion patterns are frequently observed in RNA-injected fish could be predicted and are identified using the program to search for CRISPR target sites with microhomology sequences (http://viewer.shigen.info/cgi-bin/crispr/crispr.cgi). 3. Enter the selected target sequences as “5′-N21GG-3′” into the file “sgRNA_design.xls” for Microsoft Excel or “sgRNA_design.ods” for Apache OpenOffice Calc, which produces each pair of 22-mer oligonucleotide sequences for subcloning into the pDR274 vector. These files can be downloaded at http://satoshi-ansai.github.io/en/misc.html. 4. Order the designed oligonucleotides from a supplier. Purification using the PCR primer grade is sufficient for the experiments described in Day 2 and Day 3. Day 2: Clone the annealed oligonucleotides into the sgRNA expression vector. 1. Anneal each pair of the ordered oligonucleotides. Prepare the reaction solution as described below. The mixture is heated to 95 °C for two minutes and then cooled slowly to 25 °C in one hour using a thermal cycler. Oligonucleotide Sense (100 μM) Oligonucleotide Anti-Sense (100 μM) 10× Annealing buffer H2O Total

1 μL 1 μL 1 μL 7 μL 10 μL

2. Assemble the ligation mixture using the annealed oligonucleotides as below. pDR274 (BsaI digested) Annealed oligonucleotides Ligation High Ver. 2 (Toyobo) Total

1 μL 1 μL 2 μL 4 μL

3. Incubate at 16 °C for 30 minutes. 4. Transform 2 μL of the ligation solution into competent E. coli cells according to standard protocols (e.g., on ice for five minutes, 42 °C for 45 seconds, and on ice for five minutes). 5. Add 100 μL of the SOC medium into the transformed E. coli solution and then incubate at 37 °C for 30–40 minutes, continuously shaking vigorously. 6. Spread E. coli cells on the LB plate containing 20 μg/mL of kanamycin. 7. Incubate at 37 °C overnight.

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Day 3: Colony PCR and miniculture of the sgRNA vector. 1. Make a large master mix by mixing together the following reagents and aliquot 10 μL in each PCR tube. Prepare 4–8 reactions for each sgRNA vector. The sense strand oligonucleotide used for construction of each sgRNA vector is added as a primer. Distilled water (DW) 10× Reaction buffer 100 mM dNTP mix 50 mM MgCl2 Oligonucleotide Sense (2 μM) M13 forward primer (2 μM) HybriPol DNA Polymerase Total

5.85 μL 1 μL 0.8 μL 0.3 μL 1 μL 1 μL 0.05 μL 10 μL

2. Pick up a small amount of white colony, poke the colony into a replica LB plate containing ampicillin, and then add to each PCR reaction. Do not take such a large amount of the colony that it could inhibit the PCR reaction. The replica plates are incubated at 37 °C until it is possible to get a clear view of the colonies in order to pick them up in a later step (step 6). 3. Run PCR with the following thermal cycle conditions: 95 °C for two minutes; 30 cycles of 95 °C for 20 seconds, 55 °C for 30 seconds, and 72 °C for 30 seconds. 4. 5 μL of each PCR reaction with an appropriate volume of loading dye is subjected to electrophoresis (135 V, 25 minute) on a 2% agarose gel. 5. Identify the 269 bp of the PCR product which should be detected if the cloning process has been done successfully in the colony. 6. Take the identified colony out of the replica plate, inoculate it into 2–5 mL of the LB or any liquid medium with 20 μg/mL kanamycin, and then grow the cultures overnight at 37 °C. Day 4: Minipreparation of the sgRNA vector and linearization for RNA synthesis. 1. Purify the plasmid from each culture grown overnight using a plasmid purification kit with spin columns. 2. Prepare the reaction solution for digestion with DraI as below. sgRNA plasmid (5 μg) 10X M buffer (Takara Bio) DraI (Takara Bio) H2O Total

X μL 10 μL 1 μL Up to 100 μL 100 μL

3. Incubate at 37 °C overnight. Day 5: Purify the linearized plasmid DNA and in vitro transcription of sgRNA. Purification of the linearized plasmid to use as a template of RNA transcription. 1. 5 μL of each digested aliquot is subjected to electrophoresis with a 2% agarose gel. Complete digestion with DraI yields two fragments 1862 and 282 bp. 2. The remaining digested solution is subjected to proteinase K treatment to eliminate the remaining RNase activity. Add 5 μL of 10% SDS and 1 μL of proteinase K (20 mg/mL) into 95 μL of the DraI-digested solution, and then incubate at 55 °C for 30 minutes.

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3. After this step, all reagents should be handled with RNase-free grade to avoid degradation of transcribed RNA. 4. Add 100 μL of the phenol-chloroform mixture. Mix the organic and aqueous phase by vortexing and then centrifuge the solution at 12 000 rpm or more for five minutes. Transfer the aqueous upper layer to a new tube. 5. Add 100 μL of chloroform. Mix the organic and aqueous phase by vortexing and then centrifuge the solution at 12 000 rpm or more for five minutes. Transfer the aqueous upper layer to a new tube. 6. Add 10 μL of 3 M sodium acetate, 5 μL of 0.5 M EDTA, and 1 μL of glycogen (20 mg/mL). Mix the solution by vortexing. 7. Add 250 μL of 100% ethanol and then mix the solution vigorously by vortexing. Collect the precipitate DNA by centrifugation at 12 000 rpm or more for 10 minutes at room temperature. 8. Remove the supernatant by pipette. Add 300 μL of 70% ethanol to the pellet and mix by vortexing. Retrive the DNA by centrifugation at 12 000 rpm or more for five minutes at room temperature. 9. Remove all of the supernatant by pipette. Store the open tube on a heat block at 70 °C until the remaining ethanol has evaporated (3–5 minutes). 10. Dissolve the pellet in 5 μL of RNase-free water by vortexing and spinning down, and then use it as a template DNA for the following RNA transcription. This solution can be stored at −20 °C. sgRNA synthesis by in vitro transcription using T7 RNA polymerase. 11. Prepare the reaction solution for in vitro transcription using the AmpliScribe T7-Flash Transcription Kit (Epicenter). Make a large master mix by mixing together the following reagents and aliquot 9 μL in each 0.2 mL PCR tube. DO NOT assemble the solution on ice. RNase-free water 10× Reaction buffer 100 mM ATP 100 mM CTP 100 mM GTP 100 mM UTP 100 mM DTT RiboGuard RNase Inhibitor AmpliScribe T7-Flash Enzyme Solution Total

2.15 μL 1 μL 0.9 μL 0.9 μL 0.9 μL 0.9 μL 1 μL 0.25 μL 1 μL 9 μL

12. Add 1 μL of template DNA in each tube and then mix the solution by gently pipetting the solution up and down. Incubate at 37 °C for 3–4 hours using a thermal cycler. 13. Add 1 μL of RNase-free DNase I (or “TURBO DNase” included in the mMessage mMachine Kit) and mix the solution gently, pipetting it up and down. Incubate at 37 °C for 15 minutes using the thermal cycler. Purification of transcribed sgRNA using the spin column. 14. For purification using the RNeasy Mini Kit, according to the RNA Clean-up protocol, adjust to a volume of 100 μL by adding 89 μL of RNase-free water.

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15. Add 350 μL of Buffer RLT and mix by vortexing. 16. Add 250 μL of 100% ethanol and then mix well, gently pipetting the solution up and down. Immediately transfer the sample (700 μL) to an RNeasy Mini spin column. 17. Centrifuge at 8000 × g for 15 seconds. Discard the flow-through. 18. Add 500 μL of Buffer RPE to the spin column, and centrifuge at 8000 × g for 15 seconds. Discard the flow-through. 19. Add 500 μL of Buffer RPE to the spin column again. Centrifuge at 8000 × g for two minutes, and discard the flow-through. 20. Place the RNeasy spin column in a new 2 mL collection tube, and centrifuge at 10 000 × g or more for five minutes. 21. Remove the remaining Buffer RPE on the upper edge of the column membrane with a pipette. Place the spin column in a new 1.5 mL tube and store the open column on a heat block at 70 °C for two minutes to eliminate any possible carryover of Buffer RPE. 22. Place the spin column in a new 1.5 mL tube. Add 30 μL of RNase-free water directly to the center of the spin column membrane. Centrifuge at 8000 × for two minutes to elute the RNA. 23. The eluted solution is immediately stored at −80 °C. Confirmation of synthesized RNA by electrophoresis and quantification. 24. A 1 μL aliquot of the eluted RNA is subjected to electrophoresis using a 2% agarose gel (nondenaturing gel is available for simple identification of successful sgRNA synthesis). Successfully synthesized sgRNA shows a dense and broad band around a 100 bp band of the ladder marker DNA. 25. Quantify the eluted RNA solution with a spectrophotometer or other methods. Adjust to an appropriate concentration using the RNase-free water and store at −80 °C until microinjection.

7.3.3  Production of capped RNA encoding a Cas9 nuclease 1. E. coli harboring the pCS2 + hSpCas9 vector is cultured in a liquid medium containing 50 μg/mL of ampicillin. Purify the plasmid using a plasmid purification kit. If the CRISPR/Cas experiment is performed constantly, purification using a midiprep kit (e.g., NucleoBond Xtra Midi, MACHEREY-NAGEL, Cat. No. 740410.50) is recommended for high-yield recovery of the plasmid. 2. Prepare the reaction solution for digestion with NotI as described below. pCS2 + hSpCas9 (10 μg) 10× M buffer (Takara Bio) Triton X-100 (0.1%) Bovine serum albumin (BSA) (0.1%) NotI (Takara Bio) H2O Total

X μL 10 μL 10 μL 10 μL 1 μL Up to 100 μL 100 μL

3. Incubate at 37 °C, overnight. 4. 5 μL of each digested aliquot is subjected to electrophoresis using a 1% agarose gel. Complete digestion by NotI shows a single band of 8343 bp.

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5. The remaining digested solution is subjected to proteinase K treatment to eliminate any remaining RNase activity. Add 5 μL of 10% SDS and 1 μL of proteinase K (20 mg/mL) into 95 μL of the NotI-digested solution, and then incubate at 55 °C for 30 minutes. 6. After this step, all reagents should be handled with RNase-free grade to avoid degradation of transcribed RNA. 7. Add 100 μL of the phenol-chloroform mixture. Mix the organic and aqueous phase by vortexing and then centrifuge the solution at 12 000 rpm or more for five minutes. Transfer the aqueous upper layer to a new tube. 8. Add 100 μL of chloroform. Mix the organic and aqueous phase by vortexing and then centrifuge the solution at 12 000 rpm or more for five minutes. Transfer the aqueous upper layer to a new tube. 9. Add 10 μL of 3 M sodium acetate, 5 μL of 0.5 M EDTA, and 1 μL of glycogen (20 mg/ mL). Mix the solution by vortexing. 10. Add 250 μL of 100% ethanol and then mix the solution vigorously by vortexing. Collect the precipitate DNA by centrifugation at 12 000 rpm or more for 10 minutes at room temperature. 11. Remove the supernatant with a pipette. Add 300 μL of 70% ethanol to the pellet and mix by vortexing. Retrive the DNA by centrifugation at 12 000 rpm or more for five minutes at room temperature. 12. Remove all of the supernatant with a pipette. Store the open tube on a heat block at 70 °C until the remaining ethanol has evaporated (3–5 minutes). 13. Dissolve the pellet in 10 μL of RNase-free water by vortexing and spining down, and use it as a template DNA for the following RNA transcription. This solution can be stored at −20 °C. 14. Prepare the reaction solution for in vitro transcription using the mMessage mMachine SP6 Transcription Kit (Life Technologies). Make a mixture in a 0.2 mL PCR tube as described below. DO NOT assemble the solution on ice. RNase-free water 2X NTP/CAP 10× Reaction buffer Template DNA Enzyme mix Total

2 μL 5 μL 1 μL 1 μL 1 μL 10 μL

15. Incubate at 37 °C for 3–4 hours using a thermal cycler. 16. Add 1 μL of TURBO DNase and mix the solution by pipetting. Incubate at 37 °C for 15 minutes using the thermal cycler. 17. For purification using the RNeasy Mini Kit according to the RNA Clean-up protocol, adjust to a volume of 100 μL by adding 89 μL of RNase-free water. 18. Add 350 μL of Buffer RLT and mix by vortexing. 19. Add 250 μL of 100% ethanol and then mix well with a pipette. Immediately transfer the sample (700 μL) to an RNeasy Mini spin column. 20. Centrifuge at 8000 × g for 15 seconds. Discard the flow-through. 21. Add 500 μL of Buffer RPE to the spin column, and centrifuge at 8000 × g for 15 seconds. Discard the flow-through. 22. Add 500 μL of Buffer RPE to the spin column again. Centrifuge at 8000 × g for two minutes, and discard the flow-through.

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23. Place the RNeasy spin column in a new 2 mL collection tube, and centrifuge at 10 000 × g or more for five minutes. 24. Remove the remaining Buffer RPE on the upper edge of the column membrane with a pipette. Place the spin column in a new 1.5 mL tube, and store the open column on a heat block at 70 °C for two minutes to eliminate any possible carryover of Buffer RPE. 25. Place the spin column in a new 1.5 mL tube. Add 30 μL of RNase-free water directly to the center of the spin column membrane. Centrifuge at 8000 × g for two minutes to elute the RNA. The eluted solution is immediately stored at −80 °C. 26. A 1 μL aliquot of the eluted RNA is subjected to electrophoresis using a 1% agarose gel (it is not necessary to use a denaturing agarose gel). 27. Quantify the eluted RNA solution with a spectrophotometer or other methods. Adjust to an appropriate concentration using the RNase-free water, and store at −80 °C until microinjection.

7.4 ­Preparation of Custom-Designed TALENs This protocol describes a method to construct the expression vectors and capped RNA of custom-designed TALENs for microinjection in medaka embryos. The assembly method in this protocol basically follows the guidelines according to the Golden Gate TALEN and TAL Effector Kit (Cermak et al. 2011) but there are some added modifications, such as ligation using the digested and purified module fragments.

7.4.1 Materials Plasmids • All plasmids used in this protocol are available via Addgene (www.addgene.org). • Golden Gate TALEN and TAL Effector Kit 2.0 (Addgene Kit #1000000024): a TALEN construction kit contains 86 plasmids for the Golden Gate assembly of TALENs (Cermak et  al. 2011). In this protocol, we only use 32 plasmids (pHD1–6, pNG1–6, pNI1–6, pNN1–6, pLR-HD, pLR-NG, pLR-NI, pLR-NN, and pFUS_B2–5). Contents of this kit are described in detail at the Addgene website (www.addgene.org/TALeffector/ goldengateV2). • Yamamoto Lab TALEN Accessory Pack (Addgene Kit #1000000030): this is a supplemental package containing modified pFUS array vectors for 6-modules assembly. These vectors are designed for use with the Golden Gate TALEN Kit as replacements for the pFUS_A vectors (pFUS_A, pFUS_A30A, pFUS_A30B) (Sakuma et al. 2013). Only the pFUS_A2A and pFUS_A2B are used in this protocol. • pCS2TAL3DD and pCS2TAL3RR (Addgene Plasmid #37275 and #37276): these vectors are TALEN backbone vectors designed for use with the Golden Gate TALEN kit as replacements of pTAL1, 2, 3, or 4 (Dahlem et al. 2012). The partial DNA fragment of pTAL3 resulting in the truncated TAL protein (136 and 63 a.a. on the N- and C-terminal, respectively) and an obligate heterodimerized FokI domain (DD or RR) were cloned into a pCS2 expression vector for capped RNA transcription with a SP6 RNA polymerase.

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PCR Primers for Colony PCR Checking • For the 1st assembly: pCR8_F1 (5′-TTGATGCCTGGCAGTTCCCT-3′) and pCR8_R1 (5′-CGAACCGAACAGGCTTATGT-3′). • For the 2nd assembly: TAL_F1 (5′- TTGGCGTCGGCAAACAGTGG-3′) and TAL_ R2 (5′- GGCGACGAGGTGGTCGTTGG-3′). Molecular Biology Reagents • BsaI-HF (NEB, Cat. No. R3535): the HF version is more suitable for overnight digestion than the BsaI without “HF” (NEB, Cat. No. R0535). • FastDigest Esp3I (Thermo Fisher Scientific, Cat. No. FD0454): the reaction of this enzyme requires a 20 mM DTT solution that is not supplied with this enzyme. • Restriction enzymes (XbaI, XhoI, and HindIII): these enzymes are used for confirmation of the assembled vectors. Other enzymes capable of digesting the assembled vectors are also available. • DNA ligase: high-efficiency ligation reagents (e.g., Ligation High Ver. 2, Toyobo, Cat. No. LGK-201) are recommended. • PCR enzyme: generally all DNA polymerases used for PCR are available. • Bacterial alkaline phosphatase (BAP): a commonly used enzyme (e.g., alkaline phosphatase [E. coli C75], Takara Bio, Cat. No. 2120A) is available. • Plasmid purification kit: a plasmid purification kit (spin column type) should be used (e.g., NucleoSpin Plasmid QuickPure, MACHEREY-NAGEL, Cat. No. 740615). Plasmids purified by phenol/chloroform extraction or polyethylene glycol (PEG) precipitation are not suitable for complete digestion using either BsaI or Esp3I. • Gel extraction kit: a gel extraction kit (spin column type) should be used (e.g., NucleoSpin Gel and PCR Clean-up, MACHEREY-NAGEL, Cat. No. 740609). • Competent E. coli: DH5α and other preferred competent cells without the Tetr allele can be used. XL1-Blue and other strains harboring the Tetr allele are not suitable to amplify the module plasmids (pHD1-6, pNG1-6, pNI1-6, and pNN1-6) with tetracycline selection. • LB broth: both the LB plates and the liquid medium containing appropriate antibiotics at concentrations shown below should be prepared. For the liquid culture, higher nutrient mediums such as 2XYT and Plusgrow II (Nacalai Tesque, Cat. No. 08202-75) are more suitable for plasmid DNA recovery with high yields. • Ampicillin (amp): a stock solution (50 mg/ml ampicillin sodium salt in H2O or 70% ethanol) should be used at a final concentration of 50 μg/mL of the medium according to standard protocols. • Tetracycline (tet): a stock solution (5 mg/ml tetracycline hydrochloride in 100% ethanol) should be used at a final concentration of 10 μg/mL of the medium according to standard protocols. • Spectinomycin (spec): 10  mg/ml of spectinomycin dihydrochloride pentahydrate (Sigma-Aldrich, cat. No. S4014; Wako, cat. No. 191-11533) in H2O is prepared as a stock solution and stored at −20 °C. This solution should be at a final concentration of 100 μg/mL of the medium. • X-Gal (5-bromo-4-chloro-3-indolyl-β-D-galactoside): prepare a 20 mg/mL solution of X-Gal in N,N-dimethylformamide (DMF) according to standard protocols. • IPTG (isopropylthio-β-D-galactoside): prepare a 20% IPTG in H2O solution according to standard protocols.

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7.4.2  Preparation of the TALEN assembly system 7.4.2.1  Preparation of TAL modules (HD1-6, NG1-6, NI1-6, and NN1-6) The E. coli cells that have been transformed by one of the 24 module plasmids (pHD1-6, pNG1-6, pNI1-6, and pNN1-6) are spread on the LB plate containing 10 μg/mL tetracycline, and then the plate is incubated at 37 °C overnight. 1. Pick an isolated colony from the plate, inoculate the colony into the LB or another liquid medium with 10 μg/mL tetracycline, and grow the cultures overnight at 37 °C. Prepare an amount of medium that is sufficient to recover more than 5 μg of plasmid DNA. 2. Purify plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentration of each elution with the spectrophotometer or other methods for DNA quantification. 3. Purified module plasmids are individually digested with BsaI. Assemble the reaction as described below and then incubate at 37 °C overnight (14–18 hours). Plasmid (5 μg) CutSmart Buffer (10×) BsaI-HF (NEB) H2O Total

X μL 3 μL 1 μL Up to 30 μL 30 μL

4. All the 30 μL of reactions are individually loaded into 2% agarose gels and separated by electrophoresis at 100 V for 20 minutes. 5. Excise each DNA fragment (ca. 100 bp) with minimized UV exposure and then retrieve the DNA using the spin column gel extraction kit. Elute the DNA with 30 μL of the elution buffer from each of the columns. Before the elution step, be sure to remove as much of the residual ethanol as possible in the column increasing the duration of the centrifuge time and then by incubating the columns for several minutes at 70 °C prior to elution. 6. Eluted DNAs are stored at −20 °C and organized with colored labels corresponding to each module.

7.4.2.2  Preparation of array backbone plasmids (pFUS vectors) 1. E. coli cells containing one of six array backbone plasmids which consist of single pFUS_A2A, A2B, B2, B3, B4, or B5 are spread onto the LB plate containing 100 μg/mL spectinomycin and the plate is incubated at 37 °C overnight 2. Pick an isolated colony from the plate, inoculate it into the LB or another liquid medium with 100 μg/mL spectinomycin, and grow the cultures overnight at 37 °C. Prepare the amount of medium that is enough to obtain more than 5 μg of plasmid DNA. 3. Purify the plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentrations of each elution with the spectrophotometer or any other methods for DNA quantification. 4. Purified module plasmids are individually digested with BsaI. Assemble the reaction as described below and then incubate at 37 °C overnight.

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Plasmid (5 μg) CutSmart Buffer (10×) BsaI-HF (NEB) H2O Total

237

X μL 5 μL 1 μL Up to 50 μL 50 μL

5. Exchange the buffer with the ethanol precipitation and then each precipitation is dissolved in 44.5 μL of H2O. Assemble the BAP reaction as described below and incubate at 37 °C for 30 minutes. This treatment enhances the success rate of the modules assembly by inhibiting self-ligation of the backbone plasmids. DNA fragment digested by BsaI 10× Alkaline phosphatase buffer Alkaline phosphatase (E. coli C75) (Takara Bio) Total

44.5 μL 5 μL 0.5 μL 50 μL

6. All the 50 μL of reactions are individually loaded into 1% agarose gels and separated by electrophoresis at 100 V for 40 minutes. 7. Excise each DNA fragment (ca. 2.5 kb) with minimized UV exposure and then retrieve the DNA using the spin column gel extraction kit. Elute the DNA with 30 μL of the elution buffer as described in section 7.4.2.1. 8. Eluted DNAs are stored at −20 °C and arranged with colored labels that are different for each of the module plasmids.

7.4.2.3  Preparation of last repeat modules (LR-HD, NG, NI, and NN) 1. E. coli cells that have been transformed by one of the four last repeat plasmids (pLRHD, NG, NI, and NN) are spread on the LB plate containing 100 μg/mL spectinomycin and incubated at 37 °C overnight. 2. Pick an isolated colony from the plate, inoculate it into the LB or liquid medium with 100 μg/mL spectinomycin, and grow the cultures overnight at 37 °C. Prepare the amount of medium that is enough to obtain more than 3 μg of plasmid DNA. 3. Purify plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentration of each elution with the spectrophotometer or any other methods for DNA quantification. 4. Purified module plasmids are individually digested with Esp3I. Assemble the reaction as described below and then incubate at 37 °C for one hour. Plasmid (3 μg) 10× Tango buffer 20 mM DTT Esp3I (Thermo Scientific) H2O Total

X μL 2 μL 1 μL A1 μL Up to 20 μL 20 μL

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5. The entire volume (20 μL) of each reaction is loaded into each well of 2% agarose gels and the electrophoresis is run at 100 V for 20 minutes. 6. Excise each DNA fragment (ca. 100 bp) with minimized UV exposure and then obtain the DNA using the spin column gel extraction kit. Elute the DNA with 15 μL of the elution buffer as described in section 7.4.2.1. 7. Eluted DNAs are stored at −20 °C and arranged with a colored labels that are different for each of the different module plasmids.

7.4.2.4  Preparation of TALEN backbone plasmids (pCS2TAL3DD and RR vectors) 1. E. coli cells containing one of the TALEN expression plasmids (pCS2TAL3DD and pCS2TAL3RR) are spread on the LB plate containing 50 μg/mL ampicillin and the plate is incubated at 37 °C overnight. 2. Pick up E. coli cells from an isolated colony from the plate with the yellow tip of a micropipette or some kind of thin rod, inoculate the cells into the LB or other liquid medium containing 50 μg/mL ampicillin, and culture the cells at 37 °C overnight. Prepare an amount of medium that is enough to obtain more than 3 μg of plasmid DNA. 3. Purify plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentration of each elution with the spectrophotometer or other methods for DNA quantification. 4. Purified module plasmids are individually digested with Esp3I. Assemble the reaction as described below and then incubate at 37 °C for one hour. Plasmid (3 μg) 10× Tango buffer 20 mM DTT Esp3I (Thermo Scientific) H2O Total

X μL 2 μL 1 μL 1 μL Up to 20 μL 20 μL

5. Exchange the buffer with the ethanol precipitation and then each precipitate is dissolved in 44.5 μL of H2O. Assemble the BAP reaction as described below and incubate at 37 °C for 30 minutes. This treatment enhances the success rate of the modules assembly by inhibiting self-ligation of the backbone plasmids. DNA fragment digested by BsaI 10× Alkaline phosphatase buffer Alkaline phosphatase (E. coli C75) (Takara Bio) Total

44.5 μL 5 μL 0.5 μL 50 μL

6. All the 50 μL of reactions are individually loaded into 1% agarose gels and separated by electrophoresis at 100 V for 40 minutes. 7. Excise each DNA fragment (ca. 5 kb) with minimized UV exposure and then retrieve the DNA using the spin column gel extraction kit. Elute the DNA with 15 μL of the elution buffer as described in section 7.4.2.1.

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8. Eluted DNAs are stored at −20 °C and arranged with colored labels that are different for each of the module plasmids.

7.4.3  Design and construction of custom-designed TALENs 7.4.3.1  Design of TALEN using TALE-NT 1. Obtain the genomic sequence including a target gene and select a target exon considering the desired features of the targeted gene, such as the position of the start codon and the domain structure. If needed, perform sequence analysis of the target genomic locus to identify polymorphisms. This analysis will become an important step, especially when nonstandard strains, such as strains derived from wild populations and northern Japanese populations, are used. 2. Open a web browser, access the TALE-NT website (https://tale-nt.cac.cornell.edu), and then click on the tab labeled “TALEN Targeter.” 3. Enter the target genomic sequence formatted with FASTA into the “sequence” box, set all parameters as below, and then click on “submit.”

Parameter

Input value

(Parameters appear after clicking “Provide Custom Spacer/RVD Length”) Minimum spacer length: Maximum spacer length: Minimum repeat array length: Maximum repeat array length:

14 17 16 18

G Substitute

NN

There is no need to change other parameters. (The default setting is useful.) 4. After being redirected to the appropriate page, the candidate target sequences and the corresponding TAL amino acid sequences are shown as a list. This list can be exported as a tab-delimited TXT file and opened by Microsoft Excel or any other spreadsheet software. When the list of candidates contains too many or too few entries, change the entry length of the target genomic sequence and then resubmit. 5. Select which TALENs to construct among the candidates. We usually select the target based on three criteria as described below. • Avoid using repeated modules containing the same RVD, especially when the “NN” module that is regarded as having relatively lower affinity for the target DNA is repeated more than three times. • Spread the “HD” modules over the entire modules because of their relatively higher affinity to the target DNA. • Examine for the presence or absence of two stretches of several nucleotides of homologous sequences (microhomology) across the cleavage site of the TALENs.

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6. Determine the modules and array vectors required for the following assembly steps using an accompanying Microsoft Excel file “TALEN_assembly_sheet.xlsx.” This file generates a list of modules and array vectors corresponding to the designed TALENs by entering both left and right recognition sequences into the respective entry boxes.

7.4.3.2  First assembly: construction of 6-modules array vectors Day 1: Assembly of modules and transformation into E. coli cells. 1. Dissolve the stock of modules and array backbones on ice. 2. Assemble the ligation solutions as described below. It is recommended that researchers arrange each module according to the assembly sheet in order to assemble the ligation solutions accurately and rapidly. For pFUS_A2A and A2B (6 modules assembly) pFUS_A2A or A2B vector (BsaI digested) Each module (BsaI digested) Ligation High Ver. 2 (Toyobo) Total

0.5 μL 0.5 μL × 6 3.5 μL 7 μL

For pFUS_B2–5 (2–5 modules assembly) DW pFUS_B2-B5 vector (BsaI digested) Each module (BsaI digested) Ligation High Ver. 2 (Toyobo) Total

0.5 μL × (6 − n) 0.5 μL 0.5 μL × n 3.5 μL 7 μL

3. Incubate at 16 °C for 30 minutes. 4. During incubation, prepare the IPTG/X-gal solution as described below and then spread 50 μL of the solution over each of the LB plates containing 100 μg/mL spectinomycin. These plates are dried naturally to remove water drops on each plate. 20% IPTG 25 μL 20 mg/mL X-gal 25 μL Total 50 μL

5. Transform the E. coli cells with the ligation solution according to standard protocols. 6. Spread the transformed E. coli cells on the LB + spec + IPTG/X-gal plates and incubate the plates at 37 °C overnight. Day 2: Colony PCR and miniculture for array vectors. 1. Take the plates out of the incubator. A very small number of white colonies often indicates a failure of the assembly, in which case this assembly should be performed again. 2. Make a large master mix by mixing together the following reagents and aliquot 10 μL in each PCR tube. Prepare four reactions for each array vector.

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DW 10× Reaction buffer 100 mM dNTP mix 50 mM MgCl2 Primer (pCR8_F1; 2 μM) Primer (pCR8_R1; 2 μM) HybriPol DNA Polymerase Total

241

5.85 μL 1 μL 0.8 μL 0.3 μL 1 μL 1 μL 0.05 μL 10 μL

3. Pick up a small amount of white colony, poke it onto a replica LB plate with spectinomycin, and then add it to each PCR reaction. Do not pick a large amount of colony that could inhibit the PCR reaction. Replica plates are incubated at 37 °C until it is possible to get a clear view of the colonies. 4. Run PCR with the following thermal cycle conditions: 95 °C for two minutes; 30 cycles of 95 °C for 20 seconds, 55 °C for 30 seconds, and 72 °C for 30 seconds. 5. Subject 5 μL of each PCR reaction with an appropriate volume of loading dye to electrophoresis (135 V, 25 minutes) on a 2% agarose gel. 6. Identify the clones harboring the successfully assembled array vector. The expected length of each PCR product is as follows: 845 bp for pFUS_A2A, 833 bp for pFUS_ A2B, 455 bp for pFUS_B2, 557 bp for pFUS_B3, 659 bp for pFUS_B4, 761 bp for pFUS_B5. A representative result is shown in Figure 7-1. 7. Pick up the identified colony from the replica, inoculate it into 2–5 mL of the LB liquid medium with 100 μg/mL spectinomycin, and then grow the cultures overnight at 37 °C. Day 3: Mini preparation and confirmation of the assembled array vectors. 1. Purify plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentrations of each elution. 2. The retained plasmid DNA is digested with the restriction enzymes XbaI and XhoI and then subjected to electrophoresis. The expected lengths of the digested fragments are as follows: 1938 and 1153 bp for pFUS_A2A, 1926 and 1153 bp for pFUS_A2B, 1548 and

pFUS_A2A

pFUS_A2B

pFUS_B4

–1000 bp –500 bp

6 modules

4 modules

Figure 7-1.  Representative electrophoretic profile of colony PCR for the first assembly of TAL modules. This example shows an assembly of 6 (pFUS_A2A and pFUS_A2B) or 4 modules (pFUS_ B4). A green check above each lane indicates a correctly assembled repeat.

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1153 bp for pFUS_B2, 1650 and 1153 bp for pFUS_B3, 1752 and 1153 bp for pFUS_ B4, 1854 and 1153 bp for pFUS_B5. 3. Sequence the array vectors using the primer pCR8_F1.

7.4.3.3  Second assembly: construction of TALEN expression vectors Day 4: Assembly of module arrays and transformation of E. coli cells. 1. Each array vector retrieved on day 3 is digested with the restriction enzyme Esp3I at 37 °C for 60 minutes as below. Plasmid (1 μg) 10× Tango buffer 20 mM DTT Esp3I (Thermo Scientific) H2O Total

X μL 2 μL 1 μL 0.5 μL Up to 20 μL 20 μL

2. Subject the digested DNA to electrophoresis (135 V, 20 minute) in a 1% agarose gel. 3. Excise each digested fragment (ca. 200–600 bp) from agarose gel under minimized UV exposure. A representative result of electrophoresis is shown in Figure 7-2. Put three array fragments consisting of a TAL effector domain together into a tube and then purify using the spin column gel extraction kit. Elute the DNA with 15 μL of the elution buffer. Prepare the ligation mixture as described below. The backbone vector pCS2TAL3DD and RR are used for the left and right TALENs, respectively. pCS2TAL3DD or pCS2TAL3RR (Esp3I digested) Last repeat (Esp3I digested) Module array mix (Esp3I digested) Ligation High ver. 2 Total

0.5 μL 0.5 μL 1 μL 2.5 μL 5 μL

4. Incubate at 16 °C for 30 minutes.

–3000 bp –1000 bp 6 modules– 4 modules–

Figure 7-2.  Representative electrophoretic profile of assembled pFUS vectors that were digested by Esp3I for the second assembly of TAL modules. Digested fragments (approximately 700 or 500 bp for 6 or 4 modules, respectively) are purified from the agarose gel.

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5. During the incubation, prepare the IPTG/X-gal solution as described below and then spread 50 μL of the solution over each of the LB plates containing 50 μg/mL ampicillin These plates are dried naturally to remove water drops on each plate. 25 μL 25 μL 50 μL

20% IPTG1 20 mg/mL X-gal Total

  IPTG is required for E. coli strains with the lac lq allele such as XL1-Blue, but not for other strains without lac lq allele such as DH5α.

1

6. Transform the competent E. coli cells with the ligation solution according to standard protocols. 7. Spread transformed E. coli cells onto the LB + amp + IPTG/X-gal plate and incubate at 37 °C overnight. Day 5: Colony PCR and miniculture for TALEN expression vectors. 1. Take out the plates from the incubator. A very small number of white colonies often indicates a failure of the assembly, in which case this assembly should be performed again. Pick up a small amount of the white colony with the tip of the micropipette, poke it onto a replica LB plate containing ampicillin, and then put the tip into a 1.5 mL microcentrifuge tube containing 50 μL of DW to suspend the E.coli cells. The aliquot of the suspension is used as a PCR template in the next step. (We recommend using the aliquot as a PCR template instead of the E. coli cells because using the E. coli cells sometimes causes failure of the PCR amplification of template DNA containing long and highly repeated sequences such as TALEN expression vector.) The replica plates are incubated at 37 °C. 2. Make a large master mix by mixing together the following reagents and aliquot 9 μL in each PCR tube. Prepare four reactions for each array vector. DW 10× Reaction buffer 100 mM dNTP mix 50 mM MgCl2 Primer (TAL_F1; 2 μM) Primer (TAL_R2; 2 μM) HybriPol DNA Polymerase Total

4.85 μL 1 μL 0.8 μL 0.3 μL 1 μL 1 μL 0.05 μL 9 μL

3. Add 1 μL of the E. coli suspension into each PCR tube. 4. Run PCR with the following thermal cycle conditions: 95 °C for two minutes, 30 cycles of 95 °C for 20 seconds, 63 °C for 30 seconds, and 72 °C for 90 seconds. 5. 5 μL of each PCR reaction with an appropriate volume of loading dye is subjected to electrophoresis (135 V, for 25 minutes) on a 1% agarose gel. 6. Identify the clones harboring the successfully assembled array vector. The expected length of each PCR product varies depending on the number of repeats as follows: 1745 bp for 15 repeats, 1847 bp for 16 repeats, 1949 bp for 17 repeats, and 2051 bp for 18 repeats. A representative result of electrophoresis is shown in Figure 7-3. 7. Select and remove the identified colony from the replica, inoculate it into 2–5 mL of the LB or any liquid medium with 50 μg/mL ampicillin, and then grow the cultures overnight at 37 °C.

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Medaka pCS2TAL3DD

pCS2TAL3RR

17 modules

16 modules

3000 bp – 1000 bp –

Figure 7-3.  Representative electrophoretic profile of colony PCR for the second assembly of TAL modules. This example shows an assembly of 17 (pCS2TAL3DD) or 16 modules (pCS2TAL3RR). A green check above each lane indicates a correctly assembled repeat.

Day 6: Mini preparation and confirmation of the assembled expression vectors. 1. Purify plasmids from the cultures using the plasmid purification kit with spin columns and then quantify the concentration of each elution. 2. Retrieved plasmid DNA is digested with the restriction enzyme HindIII and then subjected to electrophoresis. The expected lengths of the digested fragments vary depending on the numbers of repeats as follows: 4347 and 2509/2539 (DD/RR) bp for 15 repeats, 4347 and 2611/2641 (DD/RR) bp for 16 repeats, 4347 and 2713/2743 (DD/RR) bp for 17 repeats, and 4347 and 2815/2845 (DD/RR) bp for 18 repeats. 3. After confirming with HindIII digestion, the plasmids are subjected to RNA synthesis. 4. The expressed plasmids are linearized by NotI digestion, and produce capped RNA according to the protocol for Cas9 RNA production described in section 7.3.3.

7.5 ­Heteroduplex Mobility Assay – A Simple Method to Detect Targeted Genome Modification Simple and rapid detection of insertions and deletions (indels) is important for efficient targeted gene disruption using targetable nucleases. Thus far, a number of methods, such as restriction length fragment polymorphism (RFLP) analysis, DNA-cleaving assay with mismatch sensitive nucleases, high-resolution melting analysis (HRMA), and LacZ disruption/ recovery assay, have been used to detect indels induced by targetable nucleases; however, each method has advantages and disadvantages. In this section, we describe the estimation of the efficiency of targeted genome modifications and the identification of mutant individuals using HMA, one of the simplest ways to detect nuclease-induced indels with gel electrophoresis after PCR amplification. The principle of HMA is described in Figure 7-4. In gel electrophoresis, the mobility of completely complementary double-stranded DNA (homoduplex) depends on its length but the motility of heteroduplex DNA containing some mismatched nucleotides is much less than that of homoduplex DNA because of its open single-strand structure. Therefore, PCR products amplified with a template containing both the wild-type and mutated sequences show

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Homozygous mutant

PCR

PCR

Homoduplex (wild type)

Homoduplex (mutant)

245

Heterozygous mutant

PCR Homoduplex (wild & mutant)

Heteroduplex

Electrophoresis

WT (b)

Wild type

Homo

Hetero

Nuclease-injected fish

PCR

PCR

Homoduplex (wild type)

Many types of homo- and heteroduplexes

Electrophoresis

WT

Nuclease-injected fish

Figure 7-4.  Schematic illustration of the principle of heteroduplex mobility assay (HMA). (a) Completely complementary double-stranded DNA (homoduplex), which is amplified from the wild-type or homozygous mutants, is segregated on the basis of their molecular weight and as a result, the amplicons are detected as a single band in electrophoresis. On the other hand, heteroduplex DNA containing some mismatched nucleotides usually moves more slowly than homoduplex DNA, resulting in the patterns appearing as multiple bands of PCR products amplified from heterozygous mutants. (b) Fish injected with the targetable nucleases usually have various types of insertions and/ or deletions, which exhibit a number of bands with different mobilities from the wild-type product derived from many types of homo- and heteroduplexes.

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a multiple banding pattern, while PCR products with either the wild-type or the modified sequence show a single banding pattern. This method is useful for all mutation detection steps in the process of establishing gene knock-out strains as described in the next section.

7.5.1 Materials Equipment • Polyacrylamide gel electrophoresis (PAGE) apparatus: prepare mini gel PAGE apparatus, power supply, and 15% polyacrylamide gels (e.g., SuperSep DNA, 15%, 17 wells, Wako, Cat. No. 190-15481). Electrophoresis is performed according to standard DNA-PAGE protocols. • Automated electrophoresis system: a microchip electrophoresis system MCE-202 with DNA-500 reagent kit (Shimadzu) or other automated electrophoresis systems suitable for fractionation of small DNA fragments (~250 bp) can be used as a substitute for PAGE apparatus. Solutions • PCR enzymes: generally all DNA polymerases used for PCR are available. • PCR primers: design and order pairs of oligonucleotides to amplify 80–250 bp of the genomic region containing each target site of the custom-designed nucleases.

7.5.2 Procedure 1. Amplify the genomic region containing the target sequence with PCR according to the standard method. 2. Apply the PCR amplicons to the 15% PAGE or automatic electrophoresis system. 3. Analyze the HMA profiles by checking the electrophoretic results.

7.5.2.1  Identification of the wild type, heterozygotes, and homozygotes The wild type, heterozygotes, and homozygotes can be identified according to the banding pattern resulting from electrophoresis. While the wild type and the homozygous mutants show a single band with small differences in their mobility, the heterozygous mutants exhibit four bands, including two bands of homoduplexes consisting of either the wild type or the homozygous products, and two bands of heteroduplexes consisting of both the wild type and mutated sequences (Figure  7-4a). Generally, heterozygotes show four banding patterns. However, two or three banding patterns are sometimes observed. Such variation of banding patterns depends on the modified pattern of the target DNA sequence.

7.5.2.2  Evaluation of the efficiency of targeted genome modifications Fish injected with active targetable nucleases show a number of bands with different levels of mobilities, which are derived from many types of homo- and heteroduplexes containing

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the wild type and/or various types of indel sequences induced by the nucleases (Figure 7-4b). The degree of mutation efficiency can be evaluated according to the number of the bands derived from the wild-type sequence. Higher modification efficiency leads to a fainter intensity of the wild-type band.

7.6 ­How to Establish Gene Knock-out Strains In this section, we describe how to establish gene knock-out (KO) medaka strains using targetable nucleases. Figure 7-5 shows a graphical abstract of a workflow.

7.6.1  Design and synthesis of genome-editing tools For details, see sections 7.3 and 7.4.

7.6.2  Evaluation of genome-editing activity with fertilized medaka eggs One of the keys to establishing genome-edited strains is the selection of efficient genome-editing tools (TALENs, sgRNA, or crRNA). Actually, researchers should evaluate several genome-editing tools targeting the gene of interest and select the more efficient one(s).

1. Design and synthesis of genome-editing tools

Microinjection

Cas9 RNA/sgRNA TALEN RNA

RNA-injected fish (1 dpf ~ hatching)

2. Evaluation of genome-editing activity with fertilized medaka eggs 3. Microinjection of the selected genome-editing tool(s)

G0 founder

Wild-type

4. Selection of founder fish by genotyping F1 embroyos

F1 fish

(1 dpf ~ hatching)

F1 heterozygote (+/m)

F1 heterozygote (+/m)

5. Selection of F1 fish carrying the same mutations and mating each other 6. Selection of homozygous mutant fish in F2 family

F2 homozygote (m/m)

:

F2 heterozygote (+/m)

:

F2 wild-type (+/+)

Figure 7-5.  Schematic illustration of a workflow to establish gene knock-out strains using targetable nucleases.

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Procedure 1. Introduce a genome-editing tool into the cytoplasm of 10 or more fertilized medaka eggs at the one-cell stage using the microinjection method. (For details see Chapter 7 Transgenesis in the first edition of Medaka: Biology, Management, and Experimental Protocols (2009).) 2. Incubate these eggs at 26–28 °C for 3–5 days. 3. Prepare the crude genome DNA from each embryo with the method described above in Appendix 7.A at the end of this chapter and in Figure 7-A.2. 4. Perform PCR to amplify the target sequence. The size of the amplicon should be designed to be less than 250 bp because the resolution of HMA becomes lower with a longer amplicon. 5. Apply the amplicons onto 15% polyacrylamide gel or into an automatic electrophoresis system. Figure 7-6 shows HMA estimation of the genome-modifying activity of TALENs and a sgRNA targeted at a gene using the automatic electrophoresis system. With TALEN-2, PCR products of the wild-type genome (red arrowhead) are more diminished than those of TALEN-1, indicating that TALEN-2 has more activity than TALEN-1, so it is better to use TALEN-2.

7.6.3  Microinjection of the selected genome-editing tool(s) Microinject approximately 2–4 nL of the selected genome-editing tool(s) mixture into the cytoplasm of a one-cell stage embryo at the concentrations indicated below. Usually ~30 adult fish are enough to establish several KO strains. Therefore, when standard strains are used (Cab and d-rR) as background, a sufficient number of injected egg is 30–50. TALEN 1

TALEN 2

CRISPR/Cas

gDNA extraction from each injected embryo and PCR amplification Evaluation of genome editing efficiency by HMA WT

WT

WT

Figure 7-6.  Evaluation of genome-editing efficiency of targetable nucleases in the injected embryos. Genomic DNA is extracted from each injected embryo, and the target genomic region is analyzed by PCR amplification followed by heteroduplex mobility assay (HMA). If the injected nuclease has genome-editing activity, each injected embryo shows multiple bands in HMA whereas wild-type embryos show a single band (red arrowhead).

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Genome-editing system

Genome-editing tool

Final injected concentration

TALEN CRISPR/Cas9

TALEN RNA sgRNA Cas9 RNA crRNA tracrRNA Cas9 RNA

50–300 ng/μL for each TALEN 25–50 ng/μL 100–200 ng/μL 10–25 ng/μL 40 ng/μL 100–200 ng/μL

CRISPR/Cas9

249

If unwanted, off-target activity is observed, it is recommended to decrease the concentration of each genome-editing tool to diminish this activity. If biallelic disruption of the target gene has lethal effects on the injected fish, it is also recommended to decrease the concentration of each genome-editing tool to obtain single allele mutated fish.

7.6.4  Selection of founder fish by genotyping F1 embryos Even if genome-editing tools are introduced into one-cell stage eggs, not all cells in the adults are mutated. Therefore, we have to screen individuals that potentially harbor mutations in their germ cells. Procedure 1. Culture the eggs injected with the genome-editing tool. After these eggs hatch and the fish reach maturity, mate the adult fish with their wild-type counterparts. 2. Extract gDNA from 8–16 of the resultant offspring (F1 embryo at 3–5 dpf) as described in Appendix 7.A and in Figure 7-A.2. 3. Perform HMA as described in section 7.5. 4. Identify the mutant DNA sequences by direct sequencing of the PCR product that shows heteroduplex in HMA. Because the PCR product is a mixture of wild and mutant DNA sequences, the raw data contain two peaks in each position, one from the wildtype and the other from the mutant type (Figure 7-7). By subtracting wild-type data from these mixed data, the mutant sequences can be identified. A freeware tool (for example, “POLY PEAK PARSER”; http://yosttools.genetics.utah.edu/PolyPeakParser) is also available to identify the mutant sequences. 5. Select the founder fish that produce the mutant offspring desired (Figure 7-8). When loss-of-function mutants are required, select founder fish that produce offspring carrying frameshift mutations that generate nonsense amino acid sequences and/or stop codons to generate shorter proteins that have no function.

7.6.5  Selection of F1 fish carrying the same mutation and the establishment of mutant strain 1. Rear F1 fish and let them grow until they have reached a body length longer than 1 cm. 2. Cut off the tail fins from each F1 fish and extract genomic DNA as described in Appendix 7.A and Figure 7-A.1. Rear each fish in a small cup (Figure 7-A.1j).

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Figure 7-7.  Direct sequencing in a heterozygous mutant that carries a wild-type allele and an 8 bp deletion allele. In case of Sanger sequencing, the heterozygous mutant shows double waves from the deletion point in a raw data file. The deletion sequence can be identified by subtracting the wild-type sequence from the double waves.

TALEN -or RGEN-injected

#1

#2

Wild-type

#3

#4

#5

HMA profiles

Figure 7-8.  Screening of founder fish by genotyping their F1 embryos. Each fish injected with targetable nucleases mates with wild-type fish, and then their F1 embryos are individually subjected to heteroduplex mobility assay (HMA) followed by direct sequencing. This step is important to identify preferable founders, which produce F1 progeny harboring desirable types of mutations with high efficiencies.

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3. Perform PCR and subsequent HMA using the extracted DNA from tail fin and classify F1 individuals according to the HMA profile (banding pattern). Principally, individuals that share the same HMA profile harbor the same mutation sequences. Therefore, the simple way to identify F1 fish with the desired mutation sequence is as follows: compare the HMA profile of F1 fish and those of F1 embryos in section 7.5 (and Figure 7-8) and choose F1 fish which show the same HMA profile as the F1 embryo harboring the desired mutation sequence. 4. Select both male and female fish harboring the desired mutation and then mate them with each other to obtain the F2 generation (Figure 7-9).

7.6.6  Selection of homozygous mutant fish in the F2 family The selected F1 generation consists of heterozygous mutants. Therefore, the F2 family obtained from the F1 crossing consists of wild-type, heterozygous, and homozygous mutants at a ratio of 1:2:1. To analyze the KO individuals, we should identify the homozygous mutant fish in the F2 fish family. Procedure 1. Breed the F2 fish when they have reached a body length of more than 1 cm. 2. Cut off the tail fins from each F2 fish and extract genomic DNA as described in Appendix 7.A and Figure 7-A.1. Rear each fish in a small cup (Figure 7-A.1j). 3. Identify the homozygous mutants using HMA. Heterozygous mutants show m ­ ultiple bands and should be selected out (Figure 7-10: 1, 4, and 5). In contrast, ­homozygous

Figure 7-9.  Genotyping in F1 families to identify pairs of F1 fish carrying the same sequence of mutations. F1 fish are obtained by mating between the selected founder and wild-type fish. Each F1 fish is raised to adulthood and is genotyped with the type of fin clip according to heteroduplex mobility assay (HMA) followed by direct sequencing.

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Figure 7-10.  Genotyping in a F2 family to identify homozygous mutant fish. The F2 fish family obtained by crossing among F1 fish harboring the same pattern of mutation usually consists of wild-type, heterozygous, and homozygous mutants in a ratio of 1:2:1. Heterozygous mutants (Lane 1, 4, and 5) can be easily distinguished by multiple band patterns in heteroduplex mobility assay (HMA), while both wild-type fish (Lane 2, 7, and 8) and homozygous mutants (Lane 3 and 6) show a single band, and therefore they are sometimes difficult to distinguish (“1st HMA” in the panel). To clearly distinguish between wild-type fish and homozygous mutants, the aliquot of the PCR product subjected to the HMA is mixed with PCR product amplified from the wild-type genome, and subsequently, the mixture is heat-denatured and annealed using a thermal cycler and then subjected to HMA again (“2nd HMA” in the panel). Homozygous mutants (Lane 3′ and 6′) show multiple bands similar to heterozygous mutants in the 1st HMA, whereas wild-type fish (Lane 2′, 7′, and 8′) show a single band.

mutants (Figure 7-10: 3 and 6) show a single band with shifted mobility from that of the wild type (Figure 7-10: 2, 3, and 8). Then, the homozygous mutants should be selected. As an option, to clarify the difference between the wild-type and homozygous mutants, subsequent second HMA analysis is simple. Separately ­prepared amplicons from the wild-type individuals are added to the amplicons that showed single banding pattern in the first HMA. After reannealing, the solution is subjected to HMA again. In the second HMA, the homozygous mutants show multiple bands (Figure 7-10: 3′ and 6′), whereas the wild types show a single band (Figure 7-10: 2′, 7′, and 6′).

7.7 ­How to Establish Gene Knock-in Strains In this section, we describe how to establish gene knock-in (KI) medaka strains using the CRISPR/Cas9 system. This protocol helps to perform advanced genome editing such as reporter gene tagging, site-specific introduction of point mutations in genes of interest, and

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BaitD

Left HA

23 bp

500 bp

XhoI

Linker

BamHI

EGFP

PolyA

Right HA

BaitD

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Figure 7-11.  The mapping of the donor plasmid with homology arms and BaitD sequences. Left and right homology arms (HA) are homologous sequences to the 5′ and 3′ genomic target regions. Lightning marks show the cleavage sites of Cas9, and the simultaneous cleavage of genomic target site and BaitD induces highly efficient gene knock-in events in medaka. When the inserted GFP gene generates fusion protein with the targeted gene product, a short linker sequence is required to meet the codon frame.

spatiotemporal regulation of gene expression, and so on. For gene KI, the donor plasmid (Figure 7-11) and sgRNA for BaitD are required in addition to Cas9 RNA (or protein) and sgRNA for the target site.

7.7.1  Design and synthesis of CRISPR/Cas9 components In addition to the sgRNA for the target site in the medaka genome, the sgRNA for BaitD is required. The sequence of BaitD is 5′-GATCTTCGGCCTAGACTGCGAGG-3′. For details on how to prepare sgRNA and Cas9 RNA, see section 7.3.

7.7.2  Evaluation of genome-editing activity with fertilized medaka eggs To achieve high KI efficiency, the DSB-inducing activity of sgRNA for the target site is critical. For details on how to select efficient sgRNAs, see section 7.3.

7.7.3  Construction of donor plasmid with homology arms (Ca. 0.5 kbp) and bait sequences To achieve precise and high efficient gene KI in medaka, homology arms (ca. 0.5 kbp) and bait sequences are needed to prepare the donor plasmid. The homology arms are upstream and downstream regions at the genomic target site, and play a role in inducing gene KI events mediated by homologous recombination. The latter are the target sequences of Cas9 for linearization of donor plasmid, and enable an increase in gene integration efficiency. BaitD sequence, one of the bait sequences, is reported to be effectively cleaved by the CRISPR/Cas9 system in mammalian cultured cells (Xu et al. 2015) and does not have any off-target sequences in the medaka genome. In addition, our study demonstrated

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that BaitD could induce the highest efficiency in gene KI events in medaka (Murakami et  al. 2017). The donor plasmid can be customized using pBaitD-gap43-linker-EGFP, which contains BaitD sequence and is obtained from RIKEN DNA BANK (http://dna.brc. riken.jp/en) (order #RDB15409). For details of plasmid mapping, see Figure 7-11. Procedure In this case, the knockiin of GFP gene into the target site is described. 1. Extract genomic DNA from embryo or tail fins by the method described in Appendix 7.A. 2. Amplify left and right homology arms (ca. 0.5 kbp) by PCR using the genomic DNA as templates with primer pairs containing the restriction enzyme site of XhoI/BamHI or EcoRI/SpeI. 3. Digest the PCR amplicons with XhoI/BamHI or EcoRI/SpeI. 4. Clone the PCR amplicons and the BamHI/EcoRI-digested GFP gene into the XhoI/ SpeI-digested backbone plasmid containing BaitD sequences (Figure 7-11). 5. Transform E. coli with the donor plasmid, and incubate the E.coli in 2 mL of the liquid medium (for example, LB medium) with 50 μg/mL ampicillin at 37 °C overnight. 6. Purify the donor plasmid from the liquid medium using a plasmid miniprep kit. 7. Adjust the volume of the plasmid solution to 100 μL using sterile water. 8. Add 5 μL of 10% SDS solution and 2 μL of 20 mg/mL proteinase K to the plasmid solution. 9. Incubate the mix solution at 55 °C for 30 minutes. This procedure helps to remove RNase in the plasmid solution and prevents disruption of CRISPR/Cas9 components (Cas9 RNA and gRNA). 10. Purify the SDS/proteinase K-treated plasmid using NucleoSpin Gel and PCR Clean-up Kit (Macherey-Nagel) with Buffer NTB according to the manufacturer’s instructions.

7.7.4  Microinjection for establishing knock-in strains 1. Prepare the injection solution containing the following components: 100 ng/μL of Cas9 RNA, 50 ng/μL of two gRNAs (one is for the genomic target site and the other for BaitD), and 2.5 ng/μL of the donor plasmid. To avoid random integration into the genome and decrease the toxicity of DNA, injection of a high concentration of donor plasmid is not recommended. 2. Inject the solution into eggs at the one-cell stage (see Chapter 7 Transgenesis in the first edition of this book). For establishing a gene KI line, it is necessary to inject more than 100 eggs. 3. Incubate the injected eggs at around 28 °C, and collect embryos expressing GFP.

7.7.5  Selecting G0 founders harboring the insert gene in the genomic target site 1. Rear the collected embryos until they reach a body length longer than 1 cm, and extract genomic DNA from their tail fins (see Appendix 7.A). 2. PCR-amplify the genomic regions of the 5′ and 3′ junctions between the genomic target site and GFP gene. 3. Perform sequence analysis of the amplicons, to identify G0 founders harboring the precise integrated insert gene at the genomic target site.

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4. Rear the identified G0 fish until sexual maturation, and mate them with wild-type fish to collect more than 20 F1 eggs. 5. Extract genomic DNA from the F1 eggs and identify germline transmitted G0 fish by PCR genotyping. 6. Mate the identified G0 fish with wild-type fish and collect F1 embryos with GFP expression. 7. Rear the collected F1 embryos until they reach a body length longer than 1 cm, and confirm the precise integration by PCR genotyping with genome DNA extracted from their tail fins as described in Appendix 7.A. 8. Select the confirmed F1 KI fish with the insert gene at the genomic target site to establish KI strains.

Column 7.1  Utilization of crRNA, tracrRNA, Cas9 Protein The popular CRISPR/Cas9 tool consists of two components. One is a single “guide” RNA (sgRNA or gRNA), which has a complement nucleotide sequence with a target and functions as an interface with Cas9 protein. The other component is Cas9 RNA which encodes nuclease (Hwang et al. 2013). The original CRISPR/Cas9 bacterial system uses two short RNAs, CRISPR-RNA (crRNA) and trans-activating crRNA (tracrRNA), instead of sgRNA (Figure 7-12). Recently, the efficacy of crRNA and tracrRNA in genome engineering has been demonstrated in zebrafish (Kotani et al. 2015) and in medaka (unpublished data). The advantages of using crRNA and traceRNA are as follows: because of the shorter size of both crRNA (42 nt) and tracrRNA (69 nt), compared to that of sgRNA (102 nt), they are easy to synthesis. There is no need to synthesize tracrRNA for each target sequence because it is common in every target. Additionally, it has been discovered that the induction of the Cas9 protein instead of the Cas9 RNA is more effective in genome editing (Kotani et al. 2015). crRNA, tracrRNA, and Cas9 protein are customizable and commercially available, indicating that these tools open the door of genome engineering to researchers who are not familiar with molecular biology. Cas9 nuclease

Cas9 nuclease

PAM 5ʹ 3ʹ

PAM

Target DNA 5ʹ

crRNA

3ʹ

5ʹ

5ʹ

3ʹ

Target DNA

3ʹ 5ʹ

5ʹ

3ʹ 5ʹ

3ʹ sgRNA

tracrRNA

(a)

(b) 3ʹ

Figure 7-12.  Structure of crRNA, tracrRNA, and sgRNA. (a) In type II CRISPR/Cas9 acquired immune system in bacteria, CRISPR RNA (crRNA) recognizes its target sequence and trans-activating crRNA (tracrRNA) functions as a interface between crRNA and Cas9 nuclease protein. (b) As a genome editing tool, crRNA and tracrRNA were fused to form a single RNA (sgRNA or gRNA).

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7.A  Simple Genomic DNA Preparation by an Alkaline Lysis Method This section provides a simple genomic DNA extraction method from the tail fins, hatched larvae, and embryos, which are convenient for genotyping in screening of mutants.

7.A.1 Materials • Alkaline lysis buffer: 25 mM NaOH and 0.2 mM EDTA. Prepare by diluting 40× NaOH stock solution (1 M NaOH) and 50× EDTA stock solution (10 mM EDTA, pH 8.0) with sterile water. • Neutralization buffer: 40 mM Tris-HCl, pH 8.0. • 0.2 mL PCR tube or 1.5 mL microcentrifuge tube: as for 0.2 mL PCR tube, both single and multitube strips are available. For finclip of adult fish • 200 mL cup: both plastic and glass cups are available. • Forceps. • Scissors. • Paper towel. • Tricaine (MS-222): for details see Chapter 1, section 1.3.4. For embryo • Forceps. • Scissors. • 200 μL or 1000 μL micropipette tips. • Stereomicroscope.

7.A.2 Procedure 1. Put 25 μL of alkaline lysis buffer into 0.2 mL PCR tubes or 1.5 mL microcentrifuge tubes. 2. Put one sample into each tube. • Adult fish individual. – Preparation of breeding cup: assign a number for each cup and pour 100 mL of breeding water into each cup. – Anesthetize fish (see Chapter 1, section 1.3.4). – Put an anesthetized fish on a paper towel (Figure 7-A.1a). – Cut off less than half of each tail fin (Figure 7-A.1b,c). Be sure not to cut muscle. If muscle is cut, an infectious disease may occur. – Pick up a piece of the detached fin using forceps and put it into a tube containing alkaline lysis buffer (Figure 7-A.1d–f). – Put each fish into a breeding cup. The fish can be kept alive until genotyping has been completed. (Fish can be kept alive more than two weeks with feeding.) • Hatched larva. The whole body of an anesthetized larva is put into the tube.

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(b)

(c)

(d)

(e)

(h)

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(g)

(i)

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Figure 7-A.1.  Tail fin clipping for adult fish genotyping. (a–f) Procedure to cut off the tail fin. (a) Anesthetized fish is put on a paper towel. (b,c) Less than half of the tail fin is removed with scissors. (d–f) The clipped fin is transferred into the tube containing the alkaline lysis buffer with forceps. (g) Both standard scissors (bottom) and ophthalmic scissors (top) can be used to cut the tail fin. (h,i) Even if the tail fin is cut off (i), the treated fish can swim the same as before (h). (j,k) Fish with their tail fins removed are individually kept in numbered plastic cups.

• Embryo with egg envelope. – Preparation of micropipette tip. Cut the end of a 200 μL micropipette tip to give it a 1 mm outside diameter (Figure 7-A.2a,b). Use a 1000 μL micropipette tip as it is. – Put each egg into a separate tube. (using 0.2 mL PCR tubes makes it easier to break the egg envelope than using a 1.5 mL tube).

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(b)

(a)

ca. 1 mm

(e)

(c)

(d)

(f)

(g)

(h)

(i)

Figure 7-A.2.  Breaking the egg envelope for genotyping. (a,b) Either a 1000 μL micropipette tip (blue) or a 200 μL tip (yellow) is used to break the egg envelope. If the 200 μL tip is used, its end should be expanded to approximately 1 mm in outside diameter by cutting the head of tip (b). (c,d) After transfering each embryo into a 0.2 mL PCR tube, break the egg envelope by pushing with the tip of the micropipette. (e) Under a stereomicroscope, it is easy to confirm the process of breaking the egg envelope. (f–i) Stereomicroscopic images of breaking the egg envelope. (f) Touch the egg with the micropipette tip. (g) Press the egg firmly. (h) Break the egg envelope. (i) The embryonic body emerges from the egg envelope.

– Press the end of the micropipette tip firmly against the embryo. – Break the egg envelope to bring the embryonic body outside the egg envelope (Figure 7-A.2c–i).

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3. After spinning down the lysis buffer and sample, incubate at 95 °C for 10 minutes with several episodes of mixing. 4. Neutralize by adding 25 μL of neutralization buffer. 5. Mix with a vortex and then spin down. This aliquot can be used as a template for PCR.

­References Ansai, S. and Kinoshita, M. (2014). Targeted mutagenesis using CRISPR/Cas system in medaka. Biol. Open 3: 362–371. Ansai, S., Ochiai, H., Kanie, Y. et al. (2012). Targeted disruption of exogenous EGFP gene in medaka using zinc-finger nucleases. Dev. Growth Differ. 54: 546–556. Ansai, S., Sakuma, T., Yamamoto, T. et al. (2013). Efficient targeted mutagenesis in medaka using custom-designed transcription activator-like effector nucleases. Genetics 193: 739–749. Ansai, S., Inohaya, K., Yoshiura, Y. et al. (2014). Design, evaluation, and screening methods for efficient targeted mutagenesis with transcription activator-like effector nucleases in medaka. Dev. Growth Differ. 56: 98–107. Boch, J., Scholze, H., Schornack, S. et al. (2009). Breaking the code of DNA binding specificity of TAL-type III effectors. Science 326: 1509–1512. Cermak, T., Doyle, E.L., Christian, M. et al. (2011). Efficient design and assembly of custom TALEN and other TAL effector-based constructs for DNA targeting. Nucleic Acids Res. 39: e82. Christian, M., Cermak, T., Doyle, E.L. et al. (2010). Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186: 757–761. Cong, L., Ran, F.A., Cox, D. et al. (2013). Multiplex genome engineering using CRISPR/Cas systems. Science 339: 819–823. Dahlem, T.J., Hoshijima, K., Jurynec, M.J. et al. (2012). Simple methods for generating and detecting locus-specific mutations induced with TALENs in the Zebrafish genome. PLos Genet. 8: e1002861. Gasiunas, G., Barrangou, R., Horvath, P., and Siksnys, V. (2012). Cas9-crRNA ribonucleoprotein complex mediates specific DNA cleavage for adaptive immunity in bacteria. Proc. Natl. Acad. Sci. U.S.A. 109: E2579–E2586. Hwang, W.Y., Fu, Y., Reyon, D. et al. (2013). Efficient genome editing in zebrafish using a CRISPRCas system. Nat. Biotechnol. 31: 227–229. Jinek, M., Chylinski, K., Fonfara, I. et al. (2012). A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337: 816–821. Kawahara, A., Hisano, Y., Ota, S., and Taimatsu, K. (2016). Site-specific integration of exogenous genes using genome editing technologies in zebrafish. Int. J. Mol. Sci. 17: 727. Kim, Y.G., Cha, J., and Chandrasegaran, S. (1996). Hybrid restriction enzymes: zinc finger fusions to Fok I cleavage domain. Proc. Natl. Acad. Sci. U.S.A. 93: 1156–1160. Moreno-Mateos, M., Vejnar, C., Beaudoin, J. et al. (2015). CRISPRscan: designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nat. Methods 12: 982–988. Moscou, M.J. and Bogdanove, A.J. (2009). A simple cipher governs DNA recognition by TAL effectors. Science 326: 1501. Murakami, Y., Ansai, S., Yonemura, A., and Kinoshita, M. (2017). An efficient system for homologydependent targeted gene integration in medaka (Oryzias latipes). Zoo. Lett. 3: 10. Peng, Y., Clark, K.J., Campbell, J.M. et al. (2014). Making designer mutants in model organisms. Development 141: 4042–4054. Sakuma, T., Hosoi, S., Woltjen, K. et  al. (2013). Efficient TALEN construction and evaluation methods for human cell and animal applications. Genes Cells 18: 315–326.

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Stemmer, M., Thumberger, T., del Sol Keyer, M. et al. (2015). CCTop: an intuitive, flexible and reliable CRISPR/Cas9 target prediction tool. PLoS One 10: e0124633. Urnov, F.D., Rebar, E.J., Holmes, M.C. et al. (2010). Genome editing with engineered zinc finger nucleases. Nat. Rev. Genet. 11: 636–646. Wiedenheft, B., Sternberg, S.H., and Doudna, J.A. (2012). RNA-guided genetic silencing systems in bacteria and archaea. Nature 482: 331–338. Xu, H., Xiao, T., Chen, CH. et al. 2015. Sequence determinants of improved CRISPR sgRNA design. Genome Res., 25, 1147–1157.

Chapter 8

Photo‐Inducible Gene Expression in Medaka

This chapter is an introduction to a new microscopic technology, the infrared laser‐evoked gene operator (IR‐LEGO) method, which enables single‐cell/local gene induction in medaka embryos or larvae.

8.1 ­Outline of IR‐LEGO The single‐cell gene induction method satisfies various research requirements, such as analysis of cell fate, cell–cell interaction, and gene function, especially in developmental biology. Traditionally, the heat shock promoter system, which is one of the endogenous stress response systems in almost all organisms, has been used to achieve temporal gene induction. In the case of medaka, the heat shock protein 70.1 (hsp70.1) promoter region is often used to induce the expression of the targeted genes by using the bath to heat up the temperature of the whole body (in this chapter, we call this procedure heat shock) (Oda et al. 2010). In medaka, many organs responded to this heat shock and were stimulated to overexpress the ectopic genes but this system was not able to control the expression in a specific manner. Therefore, we utilized a microscope technology that can realize spatially controlled heating using an infrared laser by focusing on the targeted cells (Figure 8-1). An infrared light (wavelength 1480 nm) matches the absorption wavelength of the O‐H vibrational mode of water molecules, and cells contain significant amount of water. The irradiation of a cell by infrared light results in a temperature rise of the target cell. Thus, appropriate irradiation results in the induction of heat shock of the target cell. This system is called the infrared laser‐evoked gene operator system (IR-LEGO) (Kamei et al. 2009). To date, using this system, the induction of genes in a single cell or local area has been applied to many model species such as nematode (Caenorhabditis elegans) (Kamei et al. 2009; Suzuki et al. 2013), fruit fly (Drosophila melanogaster) (Miao et al. 2015), zebrafish (Danio rerio) (Deguchi et  al. 2009; Kimura et  al. 2013), medaka (Oryzias latipes) (Deguchi et al. 2009; Kobayashi et al. 2013; Okuyama et al. 2013; Shimada et al. 2013), frog (Xenopus laevis) (Hayashi et  al. 2014; Kawasumi‐Kita et  al. 2015), and newt (Pleurodeles waltl) (Kawasumi‐Kita et  al. 2015), and model plants such as thale cress (Arabidopsis thaliana) (Deguchi et  al. 2009; Kurihara et  al. 2013) and liverwort (Marchantia polymorpha) (Nishihama et al. 2016). Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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IR-LEGO Box (Relay lenses, Shutter)

Objective

Optical Fiber

Dichroic Mirror (IR/VIS)

Microscope system

Observation

IR (1480 nm)

Figure 8-1.  Optical diagram of IR‐LEGO microscope system. An IR laser beam (1480 nm) is introduced into an inverted microscope through the IR‐LEGO box which enables the introduction of parallel beams using relay lenses. The IR beam is reflected by a dichroic mirror which reflects IR and transmits visible light, and is focused onto the target cell in a living embryo with a custom‐made objective. The embryo is held with viscous gel (2–3% nitrocellulose solution or 0.5–1% low melting agarose) to maintain a constant depth in the target cell. Because this wavelength is effectively absorbed by water, the IR beam intensity is gradually decreased as the depth increases. The heated spot is restricted near the focal point and then a single cell (or a small number of cells) expresses the target gene due to the heat shock response.

The limitation for the application of IR‐LEGO is that for each species, there needs to be a certain method to establish transgenic lines in which expression of the targeted genes is controlled using the promoter region of hsp70.1 specifically in the targeted organs. But there are heat shock promotor induction systems for most model organisms that can be directly applied to the IR‐LEGO system. Recent studies using the IR‐LEGO system tested its application, such as to confirm that fluorescent proteins were induced in a target cell or a target region in the organs. These reports show that the IR‐LEGO system is one of the most promising methods to analyze the function of genes in vivo in developmental biology and related fields.

8.2 ­Practical Strategies of IR‐LEGO in Medaka Study 8.2.1  Selection of heat shock promoters and application studies The heat shock response is characterized by the transient induction of a set of proteins upon temperature upshift and occurs in all organisms. A downstream gene of heat shock promoter may be induced in a transient manner by a single heat shock treatment. Thus, the gene product is expressed transiently in cells. The duration of the gene product depends on

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its stability in each tissue and cell. In the case of enhanced green fluorescent protein (EGFP), fluorescence produced by heat induction appears 8–12 hours after induction and lasts only for a few days in medaka embryos and larvae. The delay in the initial appearance of fluorescence is caused by the maturation of the EGFP chromophore formation. So, in order to induce a gene expression that has a longer duration, for example to use for long‐ term cell labeling for cell fate analysis, it is necessary to choose another strategy. To solve this problem, Cre recombinase and its recognition sequence, loxP, are highly effective. Cre recombinase catalyzes site‐specific recombination between the loxP sequences (Nagy 2000). By utilizing this system, the heat shock gene induction system can realize long‐term gene expression by recombination of the loxP site in a transgenic construct. The “Cre driver line” consists of the heat shock promoter, the Cre (hsp::Cre), and the “effector line” that has a constitutive promoter (or tissue specific promoter) and a loxP cassette (loxP [gene A] loxP and gene B) (Figure 8-2). To confirm the induction of long‐term gene expression, we established a double transgenic line by crossing the Cre driver line and effector line. The promoter of hsp70.1 was used to drive the expression of Cre, and a β‐actin promoter was used to induce fluorescent proteins (DsRed and EGFP) to check the recombination of the genome. Before heat shock induction is performed, only DsRed should be expressed in the double transgenic line. We expected that

A: Transient expression hsp prom.

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ON

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After HS Figure 8-2.  Heat shock promoter constructions. The heat shock applied to a direct drive transgenic line results in a transient expression of a target gene X (a). The Cre‐mediated heat shock expression system is effective for long‐term gene expression (b). The system is a combination of Cre driver and effector constructs. Gene A is expressed using the constitutive promoter, while gene B is inhibited by the poly A (pA) sequence located upstream of gene B. Once the heat shock induces the expression of Cre recombinase, the IR‐LEGO microscope system enables induction of the expression of the genes in the single cell because this recombination reaction unit exists in the cell. After heat shock treatment, the cell starts expressing gene B, and this expression will permanently carry into its daughter cells. This means that we can track the fate of the cell by a fluorescence derived from the product of gene B. Usually, we use the 8xHSEs as the DNA construct of Cre driver to prevent leaky expression of Cre.

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only DeRed would be expressed in the embryos of the double transgenic line without heat shock. However, we found that embryos of the double transgenic line slightly expressed EGFP in a mosaic manner. This result showed that some cells expressed Cre without heat shock induction (A. Shimizu and Y. Kamei, unpublished data ). We believed this was because hsp70.1 might perform a house‐keeping function to maintain protein quality in cells. In the case of direct EGFP induction using the hsp70.1 promoter, a trace of EGFP may be expressed without heat shock, but we were not able to detect EGFP with a fluorescent microscope due to microscope sensitivity (Oda et al. 2010). On the other hand, a Cre‐mediated system can enhance the EGFP signals because EGFP was produced by a strong promoter (β‐actin), immediately after Cre excised the loxP sites. To reduce this leaky expression, we changed the Cre driver promoter to an artificially synthetic heat shock promoter that has eight tandem heat shock elements (HSE) (Bajoghli et  al. 2004). We established the 8xHSE::Cre transgenic line and then cross‐bred them with some loxP effector lines to determine the existence or otherwise of the leaky EGFP expression (or to check the condition of the leaky EGFP expression). We found no evidence of leaky EGFP expression in both the neurospecific HuC promoter (HuC::loxP[DsRed]‐GFP) line and the glyceraldehyde‐3‐phoshpate dehydrogenase (gapdh) promoter (gapdh::loxP[DsRed]‐GFP) line unless they were accompanied by heat shock induction (Kobayashi et  al. 2013; Okuyama et  al. 2013). Of course, these lines can also be used for IR‐LEGO induction. The Cre‐mediated long‐term gene expression in medaka using the IR‐LEGO system has already been used for clarifying two biological questions. In the developmental biology field, it was believed that exoskeletal tissues originated from neural crest cells. Indeed, transplantation of labeled neural crest cells exhibited invasive behavior into fins in zebrafish and swordtail fish (Smith et al. 1994; Hirata et al. 1994). However, no direct evidence of the origin of exoskeletal tissues in the trunk region has been reported and the origins of scales and fin rays have been unclear. Based on the data obtained using an excellent cell/ tissue transplantation technique (see section 10.5 in Chapter 10 of the first edition), it was hypothesized that exoskeletal tissues, such as scales and fin rays in the trunk region, do not originate from neural crest cells but are derived from mesodermal cells due to preliminary transplantation experiments. So the IR‐LEGO system was used to test this hypothesis by long‐term cell labeling using 8xHSE::Cre and β-act::loxP[None]‐EGFP lines (“None” means a nonsense protein without fluorescence). We irradiated IR to somite cells or lateral plate mesodermal cells in 3 dpf embryos to label these cells by green fluorescent protein (GFP). Then the GFP expression was observed from 4 dpf (one day after irradiation) to maturity (Figure  8-3), and finally, we obtained evidence that the trunk exoskeleton was mesodermal in origin (Shimada et al. 2013). Furthermore, this long‐term gene expression system in medaka is also advantageous in performing cell lineage analyses. The IR‐LEGO system was used to visualize the structure of a clonal unit of young neurons, which is composed of cells derived from the same neural stem cells in early developmental stages. It was found, for the first time, that the structure of clonal units of the neural network in the teleost’s telencephalon is compartmentalized (Okuyama et al. 2013) (Figure 8-4). Also, the Gaudi system, which is the brainbow system in medaka, can be utilized for clonal analysis (Centanin et al. 2014). Additionally, the site‐ specific recombination systems, such as VCre/VloxP and SCre/SloxP, are available in medaka (Kishimoto et al. 2016). Combinations of these new methods and the IR‐LEGO system will enable more precise investigation of cell lineage analyses.

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Cre driver line (male) 4 dpf

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Figure 8-3.  The combination of IR‐LEGO and Cre‐mediated cell labeling system enables long‐ term fate analysis. Embryos were obtained by crossing the driver and effector lines. To prevent effect of maternal Cre mRNA, we usually used a male of the Cre driver line. At three days post fertilization (3 dpf), the IR irradiation operation was done. In order to check the labeled cell, we observed the EGFP expression using a fluorescent microscope at 4 dpf (one day after irradiation). Some cells at the target region (in this case, the target was somite) expressed EGFP (a). Those embryos were cultivated until they became juveniles (three weeks or more) to analyze the fates of the target cells (b). Source: Photographs are modified from the cover art: Saida, M. and Kamei, Y. Introduction of Infrared Laser Evoked Gene Operator (IR‐LEGO) Technique. Kagaku to Seibutu, 53, 580–585. 2015 (in Japanese).

8.3 ­Laser Irradiation Conditions and Sample Preparation For the IR‐LEGO system, we used a continuous wave (CW) 1460 nm wavelength laser diode (LD). The laser can heat cells effectively because this wavelength matches the O‐H vibrational mode of water. The temperature of IR‐irradiated cells increases rapidly, quickly reaches a plateau, and then is kept at a constant level during irradiation; however, the plateau temperature depended on the laser power (Kamei et al. 2009). We have experimentally determined that the relation between temperature and duration for heat shocking is in reverse proportion, that is, high temperature requires short duration. A heat shock experiment using a water bath requires 15 minutes or more to induce the gene expression to achieve stable reproducibility, since the volume of sample (whole body and medium) requires a long duration to reach the target temperature by heat conduction from the preheated water bath. On the other hand, the IR‐LEGO system can directly heat a quite small

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Neural stem cells Neural progenitor cells Neurons

Figure 8-4.  Cell lineage analysis of the neural network in the medaka telencephalon using the IR‐LEGO system. (a) Schematic drawing of Cre‐loxP recombination used in this experiment. (b) Cell lineage analysis of young neurons in the adult stage derived from the neural stem cells in the embryo (stage 24). (c) Heat shock is performed on the neural stem cells at the red points in the embryo at stage 24. (d) Fluorescent proteins were detected in the adult telencephalon using cross‐sectional analysis (right photos). The figure on the left is a schematic drawing of the lateral view of an adult brain. Scale bars 100 μm. Source: Modified from Okuyama et al. 2013. https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0066597. Licensed under CC BY 4.

amount of water (only in the target cell), so it can induce heat shock response with the shortest duration. Indeed, a one‐second irradiation can induce heat shock gene expression (Kamei et al. 2009). On the other hand, of course, excess heating (excess duration or intensity of irradiation) results in cell death or damage. Therefore, it is necessary to find the optimal duration and intensity of IR irradiation levels. In order to determine the optimal conditions for gene expression without cell damage, it is important to fix the position of the sample, especially in regard to the depth of the target cell from the cover glass. Depth is the most critical parameter in IR‐LEGO experiments because IR laser energy is absorbed by water before reaching the target cell, so target cells located in deep tissue require more laser power. Through experiments for various target cells and various species, we found that laser power of 10–30 mW on the side of the objective facing the sample seemed to be the best condition for one‐second laser heat shocking. And we also learned that it is important to find a stable sample‐holding technique to maintain the depth in the targeted cell.

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8.4 ­Caution in Maintaining Strains The heat shock response can be caused not only by temperature increase but also by other stresses, such as high‐density cultivation and hypoxia. Hence, embryos, larvae, and adult fish should be kept in the best rearing conditions, that is, 20 embryos or fewer in a P10 dish, 20 larvae or fewer in a 1 L tank, and five adult fish or less in a 1 L tank. In any case, it is important to check for unexpected expressions of target genes before using the line for further experiments. Moreover, in the case of long‐term expression system, two transgenic lines (Cre driver line and effector line) should be handled and maintained separately. If a loxP cassette in the double transgenic line is accidentally excised from the genome of germline cells, the offspring inherits the excised genome, causing it to become unusable because the line has already lost the potential for recombination by Cre. Also, we recommend not crossing a female fish of the Cre driver line with a male fish of the effector line, because the maternal mRNA of Cre can be retained when females are subjected to strong stress (Figure 8-3).

8.5 ­Other Uses of IR‐LEGO As mentioned above, excess heating results in cell death so the system can also be used for single‐cell ablation. Of course, other laser techniques can also be used for cell ablation, such as the micropoint microscope system that uses a dye laser (440 nm) to focus on target cells. In the case of the IR‐LEGO microscope system, high power (i.e., 50 mW or more) and short irradiation (i.e., 1/60–1/125 seconds) are effective for single‐cell ablation (Kimura et al. 2013; Okuyama et al. 2014; Zeng et al. 2016). This technique has been applied to analyze the function of neuronal networks. The high‐ power IR laser beam was used to irradiate gonadotropin‐releasing hormone 3 (GnRH3) neurons labeled by EGFP at the early stage of the medaka embryo. The embryos were cultivated to adulthood (or maturity) and their sexual behaviors observed. Then, the neuronal networks related to GnRH3 neurons of the ablated fish were disturbed and the mate choice behaviors of this experimental fish were seen to be abnormal compared with normal control fish (Okuyama et al. 2014). This cell ablation technique was also applied to research in zebrafish on endothelial cells and neurons (Kimura et al. 2013; Zeng et al. 2016). On the other hand, high‐power and long‐duration irradiation resulted not only in cell death, but also heat induction around the target cells (Kawasumi‐Kita et al. 2015). The IR‐LEGO system can also be used for heating a single cell (the aim is not gene expression) in individuals. In general, heat stimulus is applied to specimens using a heated needle, so it was difficult to separate the effects of heat and contact stimuli. With the IR‐LEGO system, the contact stimuli were distinguished from heat stimuli; there is a specific sensory neuron which receives blue light, heat and contact stimuli, and transmits these signals to the nerve system in a different manner depending upon the type of stimuli (Terada et  al. 2016). There are still many unanswered questions about temperature and heat effects at the cellular level (Nakano et al. 2017) and the IR‐LEGO microscope may be a useful tool to help provide some clarity on these issues.

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8.6 ­Summary and Future Prospects An infrared laser with a microscope is a multimodal technique for biological studies. As an infrared laser can heat cells rapidly, the system enables cell ablation, cell heating to create a temperature gradient and single‐cell heat shock gene expression by employing the heat shock promoter system. The temporal gene induction using heat shock can be applied to many species because most species exhibit the heat shock response. Furthermore, the IR‐ LEGO system, the single‐cell gene induction system, can combine with other molecular biological techniques, such as the Cre‐loxP system to achieve long‐term gene expression. In medaka, the Cre‐loxP system works well and we have already established a Cre driver medaka line for distribution to medaka researchers. The IR‐LEGO system will be a useful tool to improve not only developmental studies but also cellular physiological studies regarding heat and temperature. The microscope add‐on system is available from Sigma‐Koki Co. Ltd. (Saitama, Japan). Also there are facilities in research organizations where the system has been set up for collaboration. The National Institute for Basic Biology welcomes collaboration using the IR‐ LEGO system. Furthermore, some transgenic lines of Cre driver and effector of fluorescent proteins are deposited in the National Bio‐Resource Project Medaka in Japan, and are available for use by researchers.

­References Bajoghli, B., Aghaallaei, N., Heimbucher, T. et al. (2004). An artificial promoter construct for heat‐ inducible misexpression during fish embryogenesis. Dev. Biol. 271: 416–430. Centanin, L., Ander, J., Hoeckendorff, B. et  al. (2014). Exclusive multipotency and preferential asymmetric divisions in post‐embryonic neural stem cells of the fish retina. Development 141: 3472–3482. Deguchi, T., Itoh, M., Urawa, H. et al. (2009). Infrared laser‐mediated local gene induction in medaka, zebrafish and Arabidopsis thaliana. Dev. Growth Differ. 51: 769–775. Hayashi, S., Ochi, H., Ogino, H. et  al. (2014). Transcriptional regulators in the Hippo signaling pathway control organ growth in Xenopus tadpole tail regeneration. Dev. Biol. 396: 31–41. Hirata, M., Ito, K., and Tsuneki, K. (1994). Migration and colonization patterns of HNK‐1‐immunoreactive neural crest cells in lamprey and swordtail embryos. Zool. Sci. 14: 305–312. Kamei, Y., Suzuki, M., Watanabe, K. et al. (2009). Infrared laser‐mediated gene induction in targeted single cells in vivo. Nat. Methods 6: 79–81. Kawasumi‐Kita, A., Hayashi, T., Kobayashi, T. et al. (2015). Application of local gene induction by infrared laser‐mediated microscope and temperature stimulator to amphibian regeneration study. Dev. Growth Differ. 57: 601–613. Kimura, E., Deguchi, T., Kamei, Y. et al. (2013). Application of infrared laser to the zebrafish vascular system. Arterioscler. Thromb. Vasc. Biol. 33: 1264–1270. Kishimoto, K., Nakayama, M., and Kinoshita, M. (2016). In vivo recombination efficiency of two site‐specific recombination systems, VCre/VloxP and SCre/SloxP, in medaka (Oryzias latipes). Dev. Growth Differ. 58 (6): 516–521. Kobayashi, K., Kamei, Y., Kinoshita, M. et al. (2013). A heat‐inducible CRE/LOXP gene induction system in medaka. Genesis 51: 59–67. Kurihara, D., Hamamura, Y., and Higashiyama, T. (2013). Live‐cell analysis of plant reproduction: live‐cell imaging, optical manipulation, and advanced microscopy technologies. Dev. Growth Differ. 55: 462–473.

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Miao, G. and Hayashi, S. (2015). Manipulation of gene expression by infrared laser heat shock and its application to the study of tracheal development in Drosophila. Dev. Dyn. 244: 479–487. Nagy, A. (2000). Cre recombinase: the universal reagent for genome tailoring. Genesis 26: 99–109. Nakano, M., Arai, Y., Kotera, I. et al. (2017). Genetically encoded ratiometric fluorescent thermometer with wide range and rapid response. PLoS One 12 (2): e0172344. Nishihama, R., Ishida, S., Urawa, H. et al. (2016). Conditional gene expression/deletion systems for Marchantia polymorpha using its own heat‐shock promoter and Cre/loxP‐mediated site‐specific recombination. Plant Cell Physiol. 27: 271–280. Oda, S., Mikami, S., Urushihara, Y. et al. (2010). Identification of a functional medaka heat shock promoter and characterization of its ability to induce exogenous gene expression in medaka in vitro and in vivo. Zool. Sci. 27: 410–415. Okuyama, T., Isoe, Y., Hoki, M. et al. (2013). Controlled Cre/loxP site‐specific recombination in the developing brain in medaka fish, Oryzias latipes. PLoS One 8: e66597. Okuyama, T., Yokoi, S., Abe, H. et al. (2014). A neural mechanism underlying mating preferences for familiar individuals in medaka fish. Science 343 (6166): 91–94. Shimada, A., Kawanishi, T., Kaneko, T. et al. (2013). Trunk exoskeleton in teleosts is mesodermal in origin. Nat. Commun. 4: 1639. Smith, M., Hickman, A., Amanze, D. et al. (1994). Trunk neural crest origin of caudal fin mesenchyme in the Zebrafish Brachydanio rerio. Proc. R. Soc. London, Ser. B 256: 137–145. Suzuki, M., Toyoda, N., Shimojou, M. et al. (2013). Infrared laser‐induced gene expression in targeted single cells of Caenorhabditis elegans. Dev. Growth Differ. 55: 454–461. Terada, S., Matsubara, D., Onodera, K. et al. (2016). Neuronal processing of noxious thermal stimuli mediated by dendritic Ca2+ influx in Drosophila somatosensory neurons. eLife 5: e12959. Zeng, C.W., Kamei, Y., Wang, C. et al. (2016). Subtypes of hypoxia‐responsive cells differentiate into neurons in spinal cord of zebrafish embryos after hypoxic stress. Biol. Cell 108: 357–377.

Chapter 9

Screening and Testing Methods of Endocrine-Disrupting Chemicals Using Medaka

9.1 ­Applied Toxicity Tests for Endocrine Disruptors The Validation and Management Group for Ecotoxicity Testing (VMG-eco) in the Organization of Economic Cooperation and Development (OECD) has been developing various standardized testing methods to screen and/or access potential endocrine-­disrupting chemicals (EDCs), and identify adverse effects of toxic chemicals using invertebrates, freshwater fish, frogs, birds, and mammals. In addition to in vivo testing, we have established reporter gene assay systems using estrogen receptors (ERs) from various fish species, including Japanese medaka (Oryzias latipes), and found that the responses to endogenous estrogen, 17β-estradiol (E2), are very similar among fish species; however, responses to estrogenic chemicals such as bisphenol A, nonylphenol, and o,p′-DDT show that species differ in sensitivity (Miyagawa et al. 2014). The in vivo response of vitellogenin (vtg) mRNA expression in the liver to estrogenic chemicals also showed sensitivity differences among different species (Lange et al. 2012). The OECD has developed several test methods in wildlife to detect the endocrine-disrupting activities of chemicals. Test Guideline No. 230 (TG230): 21-Day Fish Assay (OECD 2009a) uses freshwater fish such as Japanese medaka, fathead minnow (Pimephales promelas), and zebrafish (Danio rerio) to screen for chemicals with estrogenic, androgenic, and aromatase inhibitory activity. TG229: Fish Short-Term Reproduction Assay (OECD 2012a) uses medaka and fathead minnow for assessing the adverse effects of chemicals on the gonadal axis, including aromatase inhibition. TG231: Amphibian Metamorphosis Assay (OECD 2009b) uses the African clawed frog (Xenopus laevis) to screen for chemicals which cause thyroid hormone disruption and adverse effects on the thyroid axes. And TG211: The Daphnia magna Reproduction Test (OECD 2012b) uses the water flea to identify chemicals having adverse effects on reproduction. The United States Environmental Protection Agency (US EPA) and the Ministry of the Environment, Japan (MOE), have cooperatively developed comprehensive tests: the Larval Amphibian Growth and Development Assay (LAGDA; TG241, OECD 2015a) and the Medaka Extended One Generation Reproduction Test (MEOGRT; TG240, OECD 2015b). The LAGDA describes the protocol for a toxicity test using X. laevis that evaluates growth Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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and development from fertilization through the early juvenile period. It is an assay (­typically 16 weeks) that assesses early development, metamorphosis, survival, growth, and partial reproductive maturation. It also enables the measurement of a suite of other endpoints that allows for the diagnostic evaluation of suspected EDCs or other types of developmental and reproductive toxicants. The MEOGRT describes the protocol for a toxicity test using Japanese medaka that are exposed over multiple generations to provide data relevant to ecological hazards and risk assessment of any chemical, including suspected EDCs. In the second generation (F2), under the MEOGRT protocol, exposure continues until hatching (two weeks post fertilization – wpf). Additional investigations would be needed to justify the utility of extending the F2 generation beyond hatching since there is not sufficient information to provide relevant conditions or criteria to warrant extension of the F2 generation at this time. However, this Test Guideline may be updated as new information and data are considered. For example, guidance on extending the F2 generation through reproduction may be potentially useful under certain circumstances (e.g., chemicals with high bioconcentration potential or indications of transgenerational effects in other taxa). This test can be used to evaluate the potential chronic effects of chemicals on fish, including potential EDCs. This method gives primary emphasis to potential population-relevant effects: namely, adverse impacts on survival, development, growth, and reproduction. Several TGs for potential EDCs in the OECD using Japanese medaka will be explained in this section. TG230 (21-Day Fish Assay) is an in vivo screening assay for certain endocrine active substances where sexually mature male and spawning female fish are kept together and exposed to a chemical for 21 days. This assay covers the screening of estrogens, antiestrogens, androgens, antiandrogens, aromatizable androgen, and aromatase inhibitors. However, the statistical ability to identify antiandrogenic activity is low. This assay was validated for medaka, fathead minnow, and zebrafish but it did not provide for the detection of androgenic activity of chemicals in zebrafish. In medaka, on termination of the 21-day exposure period, one or two biomarker endpoint(s) such as vitellogenin (VTG) and papillary processes in the anal fin and secondary sexual characteristics (SSC) are measured in males and females. In fathead minnow, tubercles on the head in both sexes are used as SSC. SSC is a male characteristic induced by androgens in females and VTG is induced by estrogens in males. Regarding culture conditions, a flow-through system is recommended. The water temperature (25 ± 2 °C) should not vary by more than ±1.5 °C throughout the test. Three differential concentrations of the test substances, one control (water) and, if needed, one solvent control, are used. Brine shrimp nauplii, commercially available food or a combination of these will be fed ad libitum two or three times daily. For medaka, on day 21 of the experiment, males and females from each treatment level (five males and five females in each of the two replicates) and from the control(s) are sampled to measure VTG and SSC. For the VTG measurement, species-specific enzyme-linked immunosorbent assay (ELISA) methods using blood and/or liver can be used. The mortality in the water (or solvent) controls should not exceed 10% by the end of the exposure period. The data need to be analyzed to determine statistically significant differences between the treatment and control responses. The highest concentration should be set at the maximum tolerated concentration determined from a range finder or from other toxicity data, 10 mg/L, or maximum solubility in water, whichever is lowest. A range of spacing factors between 3.2 and 10 is recommended. The actual chemical concentrations of the test need to be measured in all vessels at the start

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of the test and at weekly intervals thereafter. Detailed explanations can be found in the OECD Guideline for the Testing of Chemicals (TG230, OECD 2009a). TG229 (Fish Short-Term Reproduction Assay) is an in vivo screening assay for fish reproduction where sexually mature male and spawning female fish are kept together and exposed to a chemical for 21 days. The short-term reproduction assay was validated in the fathead minnow and medaka. This assay covers the screening of chemicals with estrogenic, antiestrogenic, androgenic, and antiandrogenic activity, aromatizable androgens, aromatase inhibitors, chemicals with nonspecific effects on the hypothalamo-pituitary-gonadal axis, and other reproductive toxicants. However, its statistical ability to identify antiandrogenic activity is low. The assay is run using three test chemical concentrations and the necessary controls, including a carrier control if necessary. Four replicate test vessels are used for each treatment level and the control(s). Daily, quantitative measurements of spawning are taken in each test vessel. At the termination of the 21-day exposure period, two biomarker endpoints, VTG and SSC, are measured in males and females separately, as indicators of the endocrine activity of the test chemical. Gonads of both sexes are also preserved and histopathology may be evaluated to assess the reproductive fitness of the test animals and to add to the weight of evidence of other endpoints. A flow-through system is recommended. Other methods are the same as TG230. For screening the antiandrogenic activity of chemicals, the androgenized female stickleback screen (Jolly et al. 2009; Katsiadaki et al. 2006; OECD 2010, 2011a) has been developed as a specialized method using the three-spined stickleback (Gasterosteus aculeatus) (Guidance Document No. 140: Androgenized Female Stickleback Screen, OECD 2010, 2011a). However, this method includes difficulties in formulating uniform test conditions because of the use of testosterone and detecting the androgen antagonizing effect of chemicals; therefore, Nakamura et al. (2014) developed a method to screen antiandrogenic activity produced by chemicals using juvenile medaka. Significant decreases in male papillary processes were observed in the juvenile medaka treated with the highest concentration of antiandrogens, vinclozolin and flutamide. This study indicates that the papillary processes can be used as an endpoint for screening the antiandrogenic activity of chemicals using juvenile medaka for a specific period based on the existing short-term reproduction assay. Ogino et al. (2014) demonstrated expression of bone morphogenetic protein 7 (Bmp7) and lymphoid enhancer-binding factor-1 (Lef1) in female anal fin by androgen exposure. These genes are essential for bone nodule outgrowth, leading to the formation of SSC (papillary processes) if female medaka are exposed to androgen and androgenic chemicals. TG234 (Fish Sexual Development Test) is an in vivo assay that assesses the early lifestage effects and potential adverse consequences of putative EDCs (estrogens, androgens, steroidogenesis inhibitors) on fish sexual development (OECD 2011b). This method has been validated using medaka, zebrafish, fathead minnow, and stickleback. In principle, it is an enhancement of TG210 (Fish, Early-Life Stage Toxicity Test) (OECD 1992). Newly fertilized eggs are exposed to at least three concentrations of the test substance dissolved in water until the completion of sexual differentiation at about 60 days post hatching. A minimum of four replicates, including control(s), is recommended. On termination of the test, the VTG, sex ratio, intersex (testis/ova) and undifferentiated fish should be analyzed through gonadal histology. In medaka, the genetic sex is identified to determine sex reversal in individual fish. The combination of VTG, phenotypic and genotypic (Matsuda et  al. 2002) sex ratio enables the test to indicate the mode of action of the test chemical. Microarray analysis of estrogen-exposed testes revealed upregulation of genes related to the zona pellucida (ZP) and the oocyte marker gene 42Sp50 in male medaka. Using quantitative

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reverse transcriptase polymerase chain reaction (RT-PCR), the Zpc5 gene was confirmed to be useful as a marker for the detection of testis/ova in male medaka (Hirakawa et al. 2012). The MEOGRT (OECD 2015b) requires an exposure period for the F2 until hatching (2 wpf). For chemicals not requiring assessment over “multigenerations” or chemicals that are not potential endocrine disruptors, other tests may be more appropriate (OECD 2012a). The medaka is an appropriate species for use in this test guideline, given its short life-cycle and the ability to determine its genetic sex (Matsuda et al. 2002). Other small laboratory fish species might also be suitable for use in “multigeneration” tests where they have a well-established historical database for endpoints measured in this test. The MEOGRT measures several biological endpoints (Table 9-1). Primary emphasis is given to potential adverse effects on population-relevant parameters including survival, gross development, growth, and reproduction. Secondarily, in order to provide mechanistic information and to provide linkage between results from other kinds of field and laboratory studies, where there is a priori evidence for a chemical having potential endocrine-disrupting activity (e.g., androgenic or estrogenic activity in other tests and assays), then other useful information is obtained by measuring vtg mRNA (or VTG), phenotypic SSC, genetic sex, and evaluating histopathology (Table 9-1). The test is started by exposing sexually mature males and females in breeding pairs of the parental generation (F0) to the target chemical for three weeks. On the first day of the fourth week, the eggs are collected to start the F1 generation. During rearing of the F1 generation (a total of 14 wpf), hatchability and survival are assessed. In addition, the fish are sampled at 12 wpf for developmental endpoints, and spawning is assessed for three weeks from 12 through 14 wpf. An F2 generation is started after the third week of the reproduction assessment and reared until the completion of hatching (Table 9-1). Optionally, the F2 generation may be reared to evaluate the transgenerational effects on survival, growth, development, reproduction, and other relevant endpoints if the substance has an accumulative property. The adverse effects of some chemicals was demonstrated using former versions (Nakamura et al. 2015; Flynn et al. 2017) and the final version of MEOGRT (Watanabe et al. 2017). Table 9-1.  Endpoint overview of the MEOGRT.a Life-stage

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Hatch (% and time to hatch) Survival Survival Growth (length and weight) Vitellogenin (mRNA or protein) Secondary sex characteristics (anal fin papillae) External sex ratio Time to 1st spawn Reproduction (fecundity and fertility) Survival Growth (length and weight) Secondary sex characteristics (anal fin papillae) Histopathology (gonad, liver, kidney)

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a These endpoints are to be statistically analyzed.

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These assays can be used in the context of the OECD Conceptual Framework for the Testing and Assessment of Endocrine-Disrupting Chemicals. Medaka researchers studying chemical effects on development, sexual differentiation, and reproduction are advised to follow the protocols of TGs in the OECD (OECD 2018).

9.2 ­Detection of Androgenic and Antiandrogenic Chemicals Using Medaka 9.2.1  The formation of papillary processes on anal fin rays as an indicative phenotype for exposure of androgenic and/or antiandrogenic chemicals The modification of the anal fin followed by papillary processes, which is developed in mature males but absent in females (Oka 1931; Uwa 1971), is a prominent masculine sex characteristic in the appendage development of Japanese medaka (O. latipes) (Figure 9-1a–d). The mating male embraces the posterior part of the female’s body with the anal fin for efficient external fertilization (Yamamoto and Egami 1974). The development of this characteristic is highly sensitive to androgen because the transplantation of a testis into the body of an adult female or the administration of androgens to

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Figure 9-1.  Induction of the formation of papillary processes in the female by androgen treatment. (a,b) Tetracycline/ calcein staining of anal fins of control female (a) and 32 nM 17α-methyltestosterone (17 MT)-treated female at 10 days (b). (c) Higher magnification of anal fin. (d) Masson/ trichrome staining of longitudinal section of developing papillary processes. Papillary processes are formed as a branching bone nodule (bn) from lepidotrichia (lep, stained with blue) in the posterior anal fin rays. A thickened mesenchyme is observed adjacent to the bone nodule (m, stained with red). Scale bars in (a,b) and (c,d) represent 0.5 mm and 0.1 mm, respectively. Source: Adapted from Ogino et al. (2014).

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females induces papillary processes formation on their anal fin rays (Okada and Yamashita 1944; Hishida and Kawamoto 1970). Therefore, the number and size of the papillary processes in females have been used as an important detector of exogenous androgenic compounds (OECD 2004), although other sexual differences such as teeth, bones, and  body shape were also reported (Egami 1959; Egami and Ishii 1956). Exposure to  both  natural (11-ketotestosterone: 11KT) and synthetic androgens (trenbolone, 17α-methyltestosterone: 17MT) causes females to develop papillary processes in their anal fins (Hishida and Kawamoto 1970; OECD 2004; Seki et al. 2004; Ogino et al. 2014). For the screening of chemicals with antiandrogenic activity, a short-term reproduction assay using juvenile medaka has been developed (Nakamura et al. 2014). Juvenile fishes at 40 days post fertilization, before the initiation of papillary process formation, were exposed to the antiandrogens vinclozolin and flutamide for 28 days. The validity of this short-term assay method was confirmed using fenitrothion (Horie et al. 2017). The papillary processes development was inhibited by these antiandrogenic chemicals in males. Papillary process development starts when total body length reaches 2.1 cm (Ogino et al. 2014). The growth rate varies depending upon the environmental conditions, such as the quality and quantity of food, temperature, and population density. Before starting the assay, it is necessary to establish and then maintain consistent growth conditions for each breeding colony.

9.2.2  Candidate biomarkers for assessing the action of androgenic and antiandrogenic chemicals Androgen-dependent augmentation of Bmp7 and Lef1 was detected in the posterior part of the anal fin in female medaka within two days of 17MT treatment (Ogino et al. 2014). Both Bmp7 and Lef1 were also highly expressed in the posterior part of the anal fin of papillary processdeveloping males. These gene expressions were inhibited by an antiandrogen, flutamide, indicating the androgen dependency of these genes during papillary processes development (Ogino et al. 2014). In other teleosts, several genes were identified as the putative effector genes that can  potentially interact with androgen signaling. The expression of Sonic hedgehog (Shh) induced by androgen was required for formation of the gonopodium that develops as a copulatory organ through elongation of the anterior region of the anal fin in mosquitofish (Ogino et  al. 2004; Brockmeier et  al. 2013). Androgen also induced the expression of muscle segment homeobox C (MsxC) and fibroblast growth factor receptor 1 (Fgfr1) in anal fin rays during the anal fin to gonopodium transition in mosquitofish (Brockmeier et al. 2013) and swordtail fish (Zauner et al. 2003; Offen et al. 2008). MsxC expression is associated with growth of the sword and gonopodium in swordtail fish (Zauner et al. 2003).

9.2.3  Visualization of androgenic and antiandrogenic activity as green fluorescence with spiggin-GFP medaka Antiandrogens induce feminizing effects in teleosts (Jobling et al. 2009). Robust screening models have been needed to identify androgenic and antiandrogenic activities among the thousands of currently untested chemicals in the environment. Two distinct paralogs of the androgen receptor (AR), ARα and ARβ, have evolved in the teleost lineage (Ogino et al. 2016).

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Four distinct steroid receptors for androgen, glucocorticoid, progesterone, and mineralocorticoid recognize identical DNA response elements (ARE/GRE) (Horie-Inoue et  al. 2006), which has hampered the development of a specific tool for AR modulation. The Spiggin production in three-spined stickleback (G. aculeatus) has been used as a biomarker for androgen axis disruption (Jolly et al. 2009). In the breeding season, the male stickleback produces the Spiggin glue protein in the kidney for the building of nests. 11KT is effective in inducing Spiggin production (Jakobsson et al. 1999). In 2011, the Guidance Document on the Androgenised Female Stickleback Screen (GD 148) was established by the OECD. The fishes are simultaneously treated with an androgen (5α-dihydrotestosterone: DHT, at 5 μg/L or 17MT 0.5 μg/L) and putative antiandrogens at a range of concentrations. The antiandrogenic activity produced by chemicals is detected by the degree of the reduction/ inhibition of androgen-induced Spiggin expression. Spiggin is measured by an ELISA (OECD 2009b). Sébillot et al. (2014) developed an alternative in vivo test by generating Spiggin-GFP transgenic medaka that contained the green fluorescence protein (GFP) gene driven by the Spiggin promoter (Figure 9-2). GFP induction is exclusive to the kidney. Significant GFP expression is induced in the kidney of just-hatched fry after three or four days of androgen treatment, with the highest sensitivity to 17MT (1.5 μg/L). Importantly, such GFP expression is dose-dependently induced by androgen. No significant GFP expression is detected by estrogens, glucocorticoids, progesterones, and mineralocorticoids, indicating a specific response to androgens. By exposing this transgenic fish that expresses the 17MT-inducing GFP to flutamide and other biocides, the antiandrogenic activities of these substances can be detected by the resulting suppression of GFP expression.

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Figure 9-2.  GFP expression pattern of Spiggin-GFP transgenic medaka. This transgenic strain contains 4.2 kbp upstream region of three-spined stickleback Spiggin 1 gene. (a,b) The expression of GFP is restricted in the kidney of an adult male (white arrow). (b) Dissected untreated adult male. (c) Spiggin-GFP fry treated for four days with vehicle (control) or 17MT (302 μg/L). The white arrow points to the 17MT-induced GFP expression in the kidney. Source: Adapted with permission from Sébillot et al. (2014). Copyright (2014) American Chemical Society.

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­References Brockmeier, E.K., Ogino, Y., Iguchi, T. et  al. (2013). Effects of 17β-trenbolone on Eastern and Western mosquitofish (Gambusia holbrooki and G. affinis) anal fin growth and gene expression patterns. Aquat. Toxicol. 128–129: 163–170. Egami, N. (1959). Note on sexual difference in the shape of the body in the fish, Oryzias latipes. Annot. Zool. Japon. 32: 59–64. Egami, N. and Ishii, S. (1956). Sexual differences in the shape of some bones in the fish, Oryzias latipes. J. Fac. Sci., Tokyo Univ. IV 7: 563–571. Flynn, K., Lothenbach, D., Whiteman, F. et  al (2017). Summary of the development the US Environment Protection Agency’s Medaka Extended One Generation Reproduction Test (MEOGRT) using data from nine multigenerational medaka tests. Environ. Toxicol. Chem., 36: 3387-3403. Hirakawa, I., Miyagawa, S., Katsu, Y. et al. (2012). Gene expression profiles in the testis associated with testis-ova in adult Japanese medaka (Oryzias latipes) exposed to 17α-ethinylestradiol. Chemosphere 87: 668–674. Hishida, T.O. and Kawamoto, N. (1970). Androgenic and male inducing effects of 11 keto testosterone on a teleost the medaka Oryzias latipes. J. Exp. Zool. 173: 279–284. Horie, Y., Watanabe, H., Takanobu, H. et  al. (2017). Development of an in vivo anti-androgenic activity detection assay using fenitrothion in Japanese medaka (Oryzias latipes). J. Appl. Toxicol. 37: 339–346. Horie-Inoue, K., Takayama, K., Bono, H. et al. (2006). Identification of novel steroid target genes through the combination of bioinformatics and functional analysis of hormone response elements. Biochem. Biophys. Res. Commun. 339: 99–106. Jakobsson, S., Borg, B., Haux, C. et al. (1999). An 11-ketotestosterone induced kidney-secreted protein: the nest building glue from male three-spined stickleback, Gasterosteus aculeatus. Fish Physiol. Biochem. 20: 79–85. Jobling, S., Burn, R., Thorpe, K. et al. (2009). Statistical modeling suggests that antiandrogens in effluents from wastewater treatment works contribute to widespread sexual disruption in fish living in English rivers. Environ. Health Perspect. 117: 797–802. Jolly, C., Katsiadaki, I., Morris, S. et al. (2009). Detection of the anti-androgenic effect of endocrine disrupting environmental contaminants using in vivo and in vitro assays in the three-spined stickleback. Aquat. Toxicol. 92: 228–239. Katsiadaki, I., Morris, S., Squires, C. et al. (2006). Use of the three-spined stickleback (Gasterosteus aculeatus) as a sensitive in vivo test for detection of environmental antiandrogens. Environ. Health Perspect. 114: 115–121. Lange, A., Katsu, Y., Miyagawa, S. et al. (2012). Comparative responsiveness to natural and synthetic estrogens of fish species commonly used in the laboratory and field monitoring. Aquat. Toxicol. 109: 250–258. Matsuda, M., Nagahama, Y., Shinomiya, A. et  al. (2002). DMY is a Y-specific DM-domain gene required for male development in the medaka fish. Nature 417: 559–563. Miyagawa, S., Lange, A., Hirakawa, I. et al. (2014). Differing species responsiveness of estrogenic contaminants in fish is conferred by the ligand binding domain of the estrogen receptor. Environ. Sci. Technol. 48: 5254–5236. Nakamura, A., Takanobu, H., Tamura, I. et al. (2014). Verification of responses of Japanese medaka (Oryzias latipes) to antiandrogens, vinclozolin and flutamide, in short-term assays. J. Appl. Toxicol. 34: 545–553. Nakamura, A., Tamura, I., Takanobu, H. et al. (2015). Fish multi-generation test with preliminary short-term reproduction assay for estrone using Japanese medaka (Oryzias latipes). J. Appl. Toxicol. 35: 11–23.

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OECD (1992). OECD Guidelines for the Testing of Chemicals, Section 2, Test No. 210: Fish, EArlyLife Stage Toxicity Test. Paris: OECD. OECD (2004). OECD Draft Report of the Initial Work Towards the Validation of the Fish Screening Assay for the Detection of Endocrine Active Substances: Phase 1A. Paris: OECD. OECD (2009a). Guidelines for the Testing of Chemicals, Test No. 230: 21-Day Fish Assay: A ShortTerm Screening for Oestrogenic and Androgenic Activity, and Aromatase Inhibition. Paris: OECD. OECD (2009b). Guideline for the Testing Chemicals, Test No. 231: Amphibian Metamorphosis Assay. Paris: OECD. OECD (2010). Draft Guidance Document No. 140: Androgenized Female Stickleback Screen. Paris: OECD. OECD (2011a). Guidance Document on the Androgenised Female Stickleback Screen (AFSS) (GD148). Paris: OECD. OECD (2011b). Test No. 234: Fish Sexual Development Test. Paris: OECD. OECD (2012a). Guidelines for the Testing of Chemicals, Test No. 229: Fish Short Term Reproduction Assay. Paris: OECD. OECD (2012b). Guidelines for the Testing of Chemicals, Test No. 211: Daphnia magna Reproduction Test. Paris: OECD. OECD (2015a). Guidelines for the Testing of Chemicals, Test No. 241: Larval Amphibian Growth and Development Assay. Paris: OECD. OECD (2015b). Guidelines for the Testing of Chemicals, Test No. 240: Medaka Extended One Generation Assay. Paris: OECD. OECD (2018). Revised Guidance Document 150 on Standardized Test Guidelines for Evaluating Chemicals for Endocrine Disruption. Paris: OECD. Offen, N., Blum, N., Mayer, A. et  al. (2008). Fgfr1 signalling in the development of a sexually selected trait in vertebrates, the sword of swordtail fish. BMC Dev. Biol. 8: 98. Ogino, Y., Katoh, H., and Yamada, G. (2004). Androgen dependent development of a modified anal fin, gonopodium, as a model to understand the mechanism of secondary sexual character expression in vertebrates. FEBS Lett. 575: 119–126. Ogino, Y., Hirakawa, I., Inohaya, K. et al. (2014). Bmp7 and Lef1 are the downstream effectors of androgen signaling in androgen-induced sex characteristics development in medaka. Endocrinology 155: 449–462. Ogino, Y., Kuraku, S., Ishibashi, H. et al. (2016). Neofunctionalization of androgen receptor by gainof-function mutations in teleost fish lineage. Mol. Biol. Evol. 33: 228–244. Oka, T.B. (1931). On the processes on the fin-rays of the male of Oryzias latipes and other sex characters of this fish. J. Fac. Sci. Imp. Univ. Tokyo. IV2: 209–218. Okada, Y.K. and Yamashita, H. (1944). Experimental investigation of the manifestation of secondary sexual characters in fish, using the medaka, (Oryzias latipes) (Temminck & Schlegel) as material. J. Fac. Sci. Imp. Univ. Tokyo. IV6: 383–437. Sébillot, A., Damdimopoulou, P., Ogino, Y. et al. (2014). Rapid fluorescent detection of (anti)androgens with spiggin-gfp medaka. Environ. Sci. Technol. 48: 10919–10928. Seki, M., Yokota, H., Matsubara, H. et al. (2004). Fish full life-cycle testing for androgen methyltestosterone on medaka (Oryzias latipes). Environ. Toxicol. Chem. 23: 774–781. Uwa, H. (1971). The synthesis of collagen during the development of anal-fin processes in ethisterone-treated females of Oryzias latipes. Dev. Growth Differ. 13: 119–124. Watanabe, H., Horie, Y., Takanobu, H. et al. (2017). Medaka Extended One-Generation Reproduction Test (MEOGRT) evaluating 4-nonylphenol. Environ. Toxicol. Chem. 36: 3254–3266. Yamamoto, M. and Egami, N. (1974). Fine structure of the surface of the anal fin and the processes on its fin rays of male Oryzias latipes. Copeia 262–265. Zauner, H., Begemann, G., Mari-Beffa, M. et al. (2003). Differential regulation of msx genes in the development of the gonopodium, an intromittent organ, and of the “sword,” a sexually selected trait of swordtail fishes (Xiphophorus). Evol. Dev. 5: 466–477.

Chapter 10

Application of the Seawater Medaka Oryzias melastigma (McClelland) for Marine Ecotoxicology 10.1 ­Background and Development of Oryzias melastigma for Marine Ecotoxicology In past decades, small‐sized fish, such as the Japanese medaka (Oryzias latipes), zebrafish (Danio rerio), fathead minnow (Pimephales promelas), mosquitofish (Gambusia affinis), and guppy (Poecilia reticulata), have been widely used as sentinel vertebrate models for ecotoxicology. These fish models offer many advantages for ecotoxicological studies due to their small size, generally easy maintenance, and ability to breed under laboratory conditions. However, these are all freshwater species, which are not desirable for evaluating the toxic effects of pollutants in marine ecosystems. The freshwater/estuarine species sheepshead minnow (Cyprinodon variegatus) and mummichog (Fundulus heteroclitus) have been commonly adapted to seawater for toxicological studies due to their hardiness in captivity. Yet both have varying rates of growth, which compromises equilibrating toxic responses. Furthermore, the mummichog takes two years to reach sexual maturity, rendering it not time‐effective to carry out whole life‐cycle toxicity studies. Then, in the mid‐2000s, the Centre for Coastal Pollution and Conservation (CCPC), City University of Hong Kong (now the State Key Laboratory of Marine Pollution, SKLMP; www6.cityu.edu.hk/sklmp/sklmp_en/index.asp), took the initiative to establish the Indian medaka Oryzias melastigma (McClelland) as a seawater fish model for marine ecotoxicology. In 2009, the Environmental Protection Department (EPD) of the Hong Kong Special Administrative Region (HKSAR) Government employed the CCPC as consultant to develop standard tests for assessing the effect of chronic toxicity of sewage effluents on marine biota in coastal waters (CCPC 2009a). The CCPC developed the “Standard Operating Procedure (SOP): 14-Days Survival and Growth Test using O. melastigma” (CCPC 2009a) to assess the effects of chronic exposure to chemicals on the survival and growth of juvenile O. melastigma. The findings highly recommended the application of the O. melastigma toxicity test for effluent characterization and impact assessment for marine waters (CCPC 2009b). Oryzias melastigma has been well described in the literature (Yamamoto 1975; Iwamatsu. 1985, Iwamatsu et  al. 1993). Phylogenetically, O. melastigma (synonyms: O. dancena, O. melastigmus, O. melanostigma) is closely related to its freshwater counterpart O. latipes, of which the entire genome has been analyzed (http://dolphin.nig.ac.jp/medaka/

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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Table 10‐1.  A summary of  general features and  basic biology of  the marine medaka Oryzias melastigma. Body length (adult) Time to sexual maturation Secondary sex characteristics (phenotypic sex) Lifespan Spawning behavior Daily egg production Egg diameter Fertilization rate Hatching rate Density tolerance Seawater maintenance Behavior

3.5–4 cm Around three months Males with larger and longer caudal and anal fins, which are distinguishable as early as one month of age Median lifespan 18 months; live up to three years Daily 5–15 eggs per female ~1000 μm >90% >90% 0.3 L of seawater per adult fish Low; change seawater 1–3 times per week Calm

index.php). Details of the ecology and taxonomical information of O. melastigma can be found the Fishbase website (www.fishbase.org/summary/Oryzias‐melastigma.html). The general features and basic biology of O. melastigma are summarized in Table 10-1. This small fish, often referred to as the “marine medaka,” completes its whole life‐cycle in seawater. It is easy to culture and breed in captivity, and stock is available on demand all year round. Importantly, it exhibits fast and uniform growth, the generation time is relatively short (2–3 months) and females can produce eggs daily, hence providing a variety of developmental and reproductive endpoints for whole life‐cycle and multigeneration assessments. The male O. melastigma is characterized by having enlarged anal and dorsal fins and less prominent urogenital papilla, whereas the female has smaller anal and dorsal fins but well‐developed urogenital papilla (Iwamatsu. 1985) (Figure 10-1). The anal fin of

1000 μm

Figure 10-1.  Adult marine medaka Oryzias melastigma. Male (bottom) and female (top) display distinct secondary sex characteristics, which can be visually distinguished as early as one month of age. The male has longer, more flowing dorsal and anal fins.

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males often develops very distinct white tips and continues to elongate as the fish ages. Such a distinct phenotypic sex dimorphism of marine medaka confers an additional advantage for studying gender‐dependent responses in toxicity assessments. The first paper proposing O. melastigma as a marine ecotoxicological model was published in 2008 (Kong et al. 2008). It described the establishment of a high‐throughput, cost‐effective whole adult medaka histoarray for quantitative in situ hybridization (ISH) and immunohistochemistry (IHC) of mRNA and protein expressions simultaneously in multiple organs of a single adult medaka, using chronic hypoxia as an environmental stressor (Kong et al. 2008). Subsequently, over 91 papers were published in 10 years (Keyword: Oryzias melastigma, PubMed, 2008–2017), using this marine fish model for ecotoxicological studies on ubiquitous marine pollutants, including brominated flame retardants (Ye et al. 2011; Deane et al. 2014; Fong et al. 2014), water‐repellent perfluorooctanesulfonic acids, polycyclic aromatic hydrocarbons from oil (Kim et al. 2014; Mu et al. 2012, 2014), endocrine‐disrupting compounds (Lee et al. 2014; Ye et al. 2017), heavy metals (Wang et al. 2015), and antifouling compounds (Chen et al. 2014a, 2016a,b, 2017a) as well as aquatic hypoxia (Wang et al. 2016). O. melastigma was also used for environmental biomonitoring to assess the water quality of Cape d’Aguilar Marine Reserve (CAMR) in Hong Kong (Xu et al. 2015). The recent advent and popularity of next generation sequencing (NGS) technology and proteomic tools greatly facilitate the use of O. melastigma as an in vivo model for molecular toxicology (Bo et al. 2012; Hwang et al. 2012; Chen et al. 2014b, 2015, 2017b; Lau et al. 2014; Lai et al. 2015; Kim et al. 2015, 2016a,b; Wang et al, 2016).

10.2 ­Marine Medaka Developmental Staging The O. melastigma embryos share highly similar developmental stages as those described for O. latipes by Iwamatsu (2004). The overall developmental rate is relatively slower for O. melastigma compared to O. latipes. However, at the optimal temperature of 26 °C, embryos of both species develop precisely at the same rate, starting from the early Stage 1 to the midgastrula Stage 15. At Stage 16 (with the first visible formation of the embryonic body), O. melastigma starts to develop more slowly. The delay in the development of O. melastigma becomes more obvious with time: when O. melastigma is still at Stage 22, O. latipes is at Stage 24. Subsequently, when O. melastigma is at Stage 31, O. latipes is already at Stage 35. The difference between the two developmental rates is in agreement with a longer period from fertilization until hatching for O. melastigma: 11 ± 2 days compared to that of O. latipes at 8 ± 2 days (at 26 °C). There are a few distinct phenotypic differences in embryonic development between the two species. Compared to O. latipes, O. melastigma embryos at Stage 1 (Figure 10-2a) are smaller and more translucent, and the divided cells at Stages 2b to 6 (Figure 10-2b–d) protrude more prominently from the yolk sphere. At Stage 15 (Figure 10-2e), the yolk sphere of O. melastigma is more completely enveloped by the dividing cells. At Stages 16–21 (Figure 10-2f,g), during early embryonic body development, the indentation of the body in the yolk sphere is very pronounced. At Stage 26 (Figure 10-2h), the melanophores on the dorsal viscera are dark and numerous in O. melastigma. Although the size of O. melastigma embryos is smaller compared to those of the O. latipes, the newly hatched larvae of both species are of similar size, approximately 4.5 mm in body length (Figure 10-2i).

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Figure 10-2.  Stages of normal development for marine medaka O. melastigma. Staging is based on that identified by Iwamatsu (2004) for O. latipes. Embryos were reared at 26 °C, and the approximate time post fertilization is given for each stage. (a) Stage 1: activated egg stage (1–3 minutes). (b) Stage 2b: blastodisc stage (30 minutes). (c) Stage 3: two‐cell stage (1 hour). (d) Stage 5: eight‐cell stage (2 hours 20 minutes). (e) Stage 15: mid‐gastrula stage (17 hours 30 minutes). Until this stage, the O. melastigma developmental timing is consistent with O. latipes reared under the same conditions. (f) Stage 17: early neurula stage (1 day 3 hours). (g) Stage 24: 16‐somite stage (2 days 5 hours). (h) Stage 26: 22‐somite stage (3 days). (i) Stage 40: first larval stage (11 days). Source: Iwamatsu (2004). Reproduced with permission of Elsevier.

10.3 ­Standard Breeding and Rearing Conditions The main differences between O. melastigma and O. latipes regarding rearing conditions are the levels of water salinity and the need for air stones (or a glass Pasteur pipette attached

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to an air pump) to increase the movement of seawater and improve oxygen exchange, especially for large tanks. For stock culture, 30–100 L tanks can maintain fish at a density of ca. 3 adults per liter (density: 0.3 l water per fish; sex ratio ca. 1:1). For adult breeding, more space (0.5 L water per fish) and a sex ratio of 1 male:1.5 female are recommended. Avoid overcrowding which may stress the fish and facilitate disease transmission. It is very important to maintain optimal conditions for mass culture of marine medaka in order to ensure a high hatching rate, reduce larval mortality during development, and allow rapid and uniform growth into adulthood.

10.3.1  Seawater Oryzias melastigma can withstand up to 60‰ salinity, but the optimal level of salinity is 30‰ under laboratory conditions. Either artificial seawater or natural seawater may be used for stock keeping and/or toxicity tests. Artificial seawater is preferable to natural seawater, which may contain unknown substances and/or contaminants. Artificial seawater can be prepared by adding sea salt (e.g., Seatreasure, DeepOcean) to UV sterilized, dechlorinated tap water to a salinity level of 30‰ (~7 kg sea salt per 300 L water). The salt should be completely dissolved and the seawater aged in aerated holding tanks overnight prior to use. The salinity levels should be checked using a calibrated refractometer and adjusted to 30‰ with artificial sea salt or dechlorinated tap water if necessary. To maintain clean tanks, stock adult culture tanks should have ca. 75% of the seawater changed every other day to remove excess food and feces. For juvenile holding tanks, change 75% of the seawater at least once a week. The bottom and sides of the tanks should be periodically scrubbed clean with a hard brush (2–4 times per month) to prevent excess growth of fungi and algae. Air tubes and stones should be cleaned monthly using ethanol or bleach and soaked overnight in dechlorinated tap water. If a dead fish is found, remove it immediately and change the water.

10.3.2  Temperature The optimal temperature for maintaining egg production is 25 °C. However, the marine medaka continue to produce eggs, in fewer numbers, at temperatures as low as 21 °C. Water heaters should be used if the water temperature is below 21 °C or too low for the desired egg production. An ambient room temperature of 25–30 °C is required to keep the water temperature within the egg production conditions. The specific water and rearing conditions for maintaining marine medaka are summarized in Table 10-2.

10.3.3  Photoperiod Oryzias melastigma live in tropical or subtropical surface waters, so it prefers plenty of sunlight. However, continuous lighting would upset the rhythm of reproduction and the circadian rhythm of the fish. A photoperiod of 12 hours light/12 hours dark is the most desirable to maintain daily egg production. Full‐spectrum overhead lighting (e.g., white‐ light fluorescent tubes, 100–500 lux) should be used for rearing and breeding.

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Table 10‐2.  Seawater conditions used for rearing marine medaka. Parameter

Range

Optimal

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Diurnal cycle Density Temperature Salinity Dissolved oxygen Nitrate Nitrite Ammonia pH KH GH

12:12 2–4 24 ± 3 25–60 6 ± 1  T) AAAGTTTTCA mutation, which corresponds to K195X at the amino acid level. The K195X mutation eliminates the telomerase active site. The optimized RTQ-TRAP assay (see SOP I) has confirmed the lack of TA in the TERT-KO. The TERT-KO line is available from the NBRP (National Bio-Resource Project) at the NIBB (National Institute for Basic Biology, Japan) (www.shigen.nig. ac.jp/medaka).

Telomerase and Telomere Biology in Medaka

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Figure 11-3.  Southern blotting measurement of telomere length in the liver of adult Japanese medaka O. latipes (OL), adult marine medaka O. melastigma (OM), and fry using the telomere restriction fragment (TRF) length, Southern blot assay. Source: Au et al. (2009). Reproduced with permission of Elsevier.

An age conversion model has been established to enable age translation between medaka (in months) and human (in years) (Gopalakrishnan et al. 2013). Moreover, a hypothetical in vivo “critical” telomere length of approximately 4 kb was deduced in the medaka liver as a model for the prediction of organismal mortality, which is highly comparable with that of human cells (see reference in Gopalakrishnan et al. 2013). In summary, the medaka is a unique, alternative vertebrate model for studying telomeres and telomerase function in a cross-disciplinary range from environmental toxicology to biomedical research on aging, as well as for cancer and tissue regeneration research. To facilitate research in these disciplines, three standard operating procedures (SOPs) are provided here to detail quantification of TA and TL optimized for medaka. I.  SOP for quantification of telomerase activity using the RTQ-TRAP. II.  SOP for quantification of telomere length using Southern blotting analysis. III.  SOP for quantification of telomere length using fluorescence in situ hybridization (q-FISH).

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11.2 ­SOP for Quantification of Telomerase Activity Using the Real-Time Quantitative Telomeric Repeat Amplification Protocol (RTQ-TRAP) 11.2.1  Procedures for sample extraction 1. Extract medaka tissue in 250 μL CHAPS lysis buffer with the aid of a polypropylene pellet pestle (Sigma Aldrich, Z359947) or a cordless motor for pellet pestles (Sigma Aldrich, Z359971). Note: • CHAPS (3-[(3-Cholamidopropyl)-dimethyl-ammonio]-1-propanesulfonate)-containing lysis buffer (10 mM Tris-HCl, pH 7.5, 1 mM MgCl2, 1 mM EGTA, pH 8.0, 0.5% (v/v) CHAPS, 10% glycerol, 0.1 mM PMSF, 5 mM β-mercaptoethanol). • Sample pooled from five individuals is required for tiny organs, e.g., kidney, testis. For muscle tissue, isolate approximately 30–50 mg from a single fish. 2. Incubate the lysed extract on ice for 30 minutes before centrifuging it at 14 000 rpm for 30 minutes at 4 °C. 3. Collect the supernatant, snap-frozen in liquid nitrogen (inside a Dewar flask) and store at −80 °C until use.

11.2.2  Procedures for determination of protein concentration 1. Use bovine serum albumin (BSA) (Sigma Aldrich P-5619) as the protein standard and measure the protein concentration with a protein assay (e.g., Bio-Rad) according to the manufacturer’s instructions. 2. Dilute the tissue lysate with CHAPS lysis buffer so that the absorbance falls within the BSA standard curve. 3. Aliquot 10 μL of diluted protein standard or tissue lysate into the wells of a clear 96-well plate and add 200 μL of the diluted Bio-Rad protein assay buffer into each well. All samples were analyzed in triplicates. 4. Incubate the reaction mixture at room temperature for five minutes and scan the 96-well plate at a wavelength of 595 nm using a spectrophotometer (Spectra MAX340, Molecular Device). 5. Determine the protein concentration in the tissue lysate from the BSA standard curve.

11.2.3  Procedures for RTQ-TRAP linearity test 1. For each tissue type, set up a series of 25 μL RTQ-TRAP reactions containing 1× homemade SYBR Green Buffer supplemented with 0.1 μg TS primer, 0.08 μg ACX primer and serially diluted protein lysate. Dilution factor is tissue specific so a linearity test has to be carried out for each tissue type. Note: • Home-made SYBR Green buffer (20 mM Tris-HCl, pH 8.3, 63 mM KCl, 1 mM EGTA at pH 8.0, 0.1 mg/mL BSA, 0.005% Tween-20, 10 nM fluorescein, SYBR

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Green I (1:250) and 1.25 U Qiagen HotStar Taq Polymerase) supplemented with 3.5 mM MgCl2 and 100 μM dNTPs. • TS primer (HPLC grade) (5′ to 3′ ): AATCCGTCGAGCAGAGTT. • ACX primer (HPLC grade) (5′ to 3′): GCGCGGCTTACCCTTACCCTTACCCTAACC. 2. Run reactions of each dilution in triplicate and include a lysate-free control (containing lysis buffer alone) as a null template control (NTC) on every run. 3. Perform RTQ-TRAP assay in iCycler or ABI 7500 Fast Real-Time PCR System. PCR cycling profile: 25 °C for 30 minutes (for primer extension by telomerase); 95 °C for 15 minutes (to inactive telomerase and activate the HotStar Taq Polymerase); 40 cycles of 95 °C for 30 seconds, 60 °C for 30 seconds and 72 °C for 1 minute 30 seconds; melt curve analysis is automatically carried out after the cycling. Note: If the reactions are carried out using the ABI 7500 Fast Real-Time PCR System, replace the reference dye fluorescein in the reaction mix with reporter dye ROX. Amplification plot, linearity test, and melt curve analysis using muscle protein as template are shown in Figure 11-4a–c, respectively. 4. Determine the Ct values from the semi-log amplification plots (log increase in fluorescence signal versus cycle number) for each sample. 5. Determine the PCR efficiency and correlation coefficient between Ct values and the protein input from the standard curve generated for each sample type. 6. For routine RTQ-TRAP using the same tissue type, follow the steps in the linearity test using the same amount of protein input for every sample so that comparison of quantitative change can be made.

11.2.4  Calculation of telomerase activity 1. The relative telomerase activity is calculated by the 2−ΔCt method (Livak and Schmittgen 2001) using the equation:

2

Ct of fish 1 Ct of fish 2



2. Arbitrarily set the telomerase activity of the first sample (the control) as 1 and the rest of samples are calculated with reference to this sample.

11.3 ­SOP for Quantification of Telomere Length Using Southern Blotting Analysis 11.3.1  Procedures for genomic DNA extraction and digestion with restriction enzymes 1. Extract the genomic DNA from isolated medaka tissue using the DNeasy® Blood and Tissue Kit (Qiagen) or other available DNA extraction kits according to the manufacturer’s instructions.

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(c)

Figure 11-4.  Optimized RTQ-TRAP assay for fish muscle. (a) Amplification plot and (b) standard curve showing the negative correlation between Ct value and protein loading in the assay and the linear range of muscle protein input between 8–1000 ng. Amount of muscle extract: (1) 1000 ng; (2) 200 ng; (3) 40 ng; (4) 8 ng and null-template control (NTC). (c) The broad base of the melting curve suggests a mixture of PCR amplicons.

2. If the genomic DNA concentration is below 75 ng/μL, it can be concentrated by DNA precipitation. • Mix 400 μL of extracted genomic DNA with 40 μL of 3 M sodium acetate (pH 5.2) and 1 mL of ice-cold absolute ethanol. • Incubate the mixture at −20 °C for two hours, and then centrifuge at 14 000 rpm for 15 minutes at 4 °C.

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• Discard the supernatant and add 1 mL of 70% ethanol to resuspend the DNA. • Centrifuge the mixture again at 14 000 rpm for 15 minutes at 4 °C. • Discard the supernatant and air-dry the DNA pellet for five minutes before dissolving in 40 μL of nuclease-free water at 65 °C (heat block) for 10 minutes (the DNA extract). • Wash the precipitated genomic DNA with 1 mL 70% ethanol and resuspend the DNA pellet in 40 μL of nuclease-free water at 65 °C for 10 minutes. 3. Digest 3 μg genomic DNA with HinfI and RsaI at 37 °C (water bath) overnight until complete digestion occurs. 4. 2 μL of digested DNA is used for electrophoresis in 1% agarose gel to ascertain complete digestion. 5. The table below shows the restriction sites of HinfI and RsaI, respectively (source: New England BioLabs; www.neb.com). Restriction enzyme

Restriction site

HinfI

5′…G A N T C…3′ 3′…C T N A G…5′

RsaI

5′…G T A C…3′ 3′…G A T C…5′

11.3.2  Procedures for probe preparation 1. DIG-labeled TTAGGG5 oligonucleotide probes are used to detect the DNA fragments with telomere sequence (TTAGGG repeats). 2. Label the telomere TTAGGG5 oligonucleotide probes (Invitrogen) using the DIG Oligonucleotide 3′-End Labeling Kit 2nd Generation (Roche Applied Science) according to the manufacturer’s instructions.

11.3.3  Procedures for electrophoresis and southern blotting 1. Resolve 3 μg of completely digested genomic DNA in 1% agarose gel electrophoresis in 1× Tris-acetate-EDTA (TAE) buffer (40 mM Tris-acetate, 1 mM EDTA, pH 8.0) with 10% of 0.5 μg/mL ethidium bromide at 70–80 V for 6–8 hours. 2. Run in parallel with 10 μL of 0.1 μg/μL of GeneRuler™ DNA Ladder Mix and 2 μL of 0.5 μg/μL of λDNA/EcoRI + HindIII (Fermentas) as the DNA molecular weight markers. 3. Depurinate the DNA by soaking the gel in 250 mM HCl solution for five minutes, denaturation solution (0.5 M NaOH and 1.5 M NaCl) for 30 minutes, and neutralization solution (0.5 M Tris-HCl, pH 7.2, 1.5 M NaCl, 100 mM EDTA) for 30 minutes.

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4. Blot the DNA on the gel onto a positively charged nylon membrane (Hybond XL, Amersham) by capillary transfer using 10× saline-sodium citrate (SSC) buffer (1.5 M NaCl, 150 mM sodium citrate, pH 7.0) overnight. 5. UV cross-link the nylon membrane at 150 mJ UV-C for 150 seconds in a GS Gene Linker UV Chamber (Bio-Rad).

11.3.4  Procedures for hybridization and detection 1. Rinse the DNA blotted membrane with 10× SSC. 2. Prehybridize the membrane with ExpressHyb solution at 42 °C for 30 minutes in a hybridization incubator (combi-H12, FINEPCR). 3. Denature 10 μL DIG-labeled TTAGGG5 oligonucleotide probes at 95 °C for five minutes in a thermal cycler, quench on ice for one minute, and then add to 10–15 mL ExpressHyb solution (in which the DNA blotted nylon membranes have been incubated overnight at 42 °C). 4. Carry out the hybridization at 42 °C overnight. 5. After hybridization, wash the membrane twice for five minutes in 2× SSC/0.1% sodium dodecyl sulfate (SDS) at 25 °C and twice for 15 minutes in 0.1× SSC/0.1% SDS at 42 °C. 6. Wash the membrane further in washing buffer (0.1 M NaCl, 0.15 M maleic acid, pH 7.5 and 0.3% Tween-20) at room temperature for five minutes before blocking. 7. Block the membrane for 30 minutes with blocking solution (0.1 M NaCl, 0.15 M maleic acid, pH 7.5 and 1% w/v blocking reagent from Roche Applied Science) and incubate in anti-DIG-AP conjugate (Roche, 1:10 000 in blocking solution) for 2–4 hours. 8. Incubate the membrane in detection buffer (0.1 M Tris-HCl, pH 9.5 and 0.1 M NaCl) for five minutes, and add 10 mL CDP-star (Roche, 1:200 in the detection buffer) to the membrane according to the manufacturer’s instructions. 9. Expose the membrane to Hyperfilm™ ECL (GE Healthcare) in the dark for five minutes.

11.3.5  Procedures for computerized telomere analysis 1. Scan the exposed films with a scanner (e.g., Duoscan HiD scanner, AGFA, with software Agfa FotoLook 3.60.00) and save as TIFF files. 2. Analyze the files with ImageJ (NIH, freely available at https://imagej.nih.gov/ij/) to determine the distribution of the lengths of the TRFs in each sample. 3. TeloRun is used to quantify TRFs. TeloRun is freely available at Telorun:https://www. utsouthwestern.edu/labs/shay-wright/methods and/or www4.utsouthwestern.edu/ cellbio/shay-wright/research/1UTSWTelorunforweb.xls 4. Quantify the proportion of TRFs greater than 5 kb, 5–4 kb, 4–3 kb, 3–2 kb and less than 2 kb with TeloRun using the following equation:

Proportion of TRFs

OD i /

OD total

where ODi refers to the optical density above background within interval i ( 2 kb) and ODtotal refers to the total optical density above background.

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11.4 ­SOP for Quantification of Telomere Length Using Fluorescence In Situ Hybridization 11.4.1  Procedures for fluorescence in situ hybridization 1. Prepare medaka tissue sections using the SOP for whole adult medaka histoarray (Kong et al. 2008) as described in section 10.9 in Chapter 10 (Figure 11-5). 2. Deparaffinization: melt the paraffin on the medaka tissue section slide at 55 °C. Then, perform tissue rehydration and antigen retrieval. 3. Permeabilization: digest the section with 1 mg/mL proteinase K at room temperature for 10–20 minutes. 4. Incubate the section in 95% ethanol for five minutes, and allow it to air-dry. 5. Denature the tissue sections on a slide in 70% formamide, 10 mM Tris, pH 7.5 and 0.5% blocking reagent at 84 °C for four minutes in the dark. 6. Hybridize denatured sections with Cy3-labeled telomere-specific PNA probes (0.3 ng/μL) at room temperature for 2–4 hours in the dark. 7. Wash the hybridized slides twice in PNA wash solution (70% formamide, 10 mM Tris, pH 7.5 and 0.1% blocking reagent). 8. Wash three more times with Tris-buffered saline (TBS), five minutes each wash. 9. Counterstain the slides with DAPI and examine the sections under confocal laser scanning microscope.

11.4.2  Procedures for confocal microscopy detection 1. Capture the images of the slide with confocal laser scanning microscope (e.g., Leica TCS SPE confocal laser scanning microscope) at 63× magnification.

Fixation Embedding Sectioning Fluorescence in situ hybridization (FISH) Confocal microscopy detection ImageJ analysis Statistical analysis

Figure 11-5.  Procedure of telomere length measurement by FISH.

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DAPI

Cy3- probed telomeres

Figure 11-6.  Representative image of testis of adult medaka. Cy3-probed telomeres are red and DAPI-stained chromosomes are blue.

2. Take the images with bright field, Cy3 signal (excitation at 556 nm and emission at 574 nm) and DAPI signal (excitation at 372 nm and emission at 456 nm) and save the images in TIFF format (1024 × 1024 pixels) (Figure 11-6). 3. The number of Cy3 pixels represents the amount of telomere repeats and the number of DAPI pixels represents the amount of DNA.

11.4.3  Procedures for ImageJ analysis 1. Determine the number of pixels of Cy3 and DAPI using ImageJ with the plugin Telometer (freely available at http://demarzolab.pathology.jhmi.edu/telometer) or with a macro script written by Napo Cheung, City University of Hong Kong. The instructions and macro script can be obtained for free by email ([email protected]) to Doris W.T. Au. 2. Calculate the relative amount of telomere repeats according to the equation:

Relative amount of telomere repeats

no. of pixels of Cy3 no. of pixels of DAPI

­References Au, D.W.T., Mok, H., Elmore, L. et al. (2009). Japanese medaka: a new vertebrate model for studying telomere and telomerase biology. Comp. Biochem. Physiol. C: Toxicol. Pharmacol. 149: 161–167. Blackburn, E.H. (1991). Structure and function of telomere. Nature 350: 569–573. Blasco, M.A., Lee, H., Hande, M. et al. (1997). Telomere shortening and tumor formation by mouse cells lacking telomerase RNA. Cell 91: 25–34. Bodnar, A.G., Ouellette, M., Frolkis, M. et  al. (1998). Extension of life-span by introduction of telomerase into normal human cells. Science 279: 349–352.

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Cherif, H., Tarry, J., Ozanne, S. et al. (2003). Ageing and telomeres: a study into organ- and genderspecific telomere shortening. Nucleic Acids Res. 31: 1576–1583. Gopalakrishnan, S., Cheung, N., Yip, B. et al. (2013). Medaka fish exhibits longevity gender gap, a natural drop in estrogen and telomere shortening during aging: a unique model for studying sex dependent longevity. Front. Zool. 10: 78. Harley, C.B., Futcher, A., and Greider, C. (1990). Telomeres shorten during ageing of human fibroblasts. Nature 345: 458–460. Hatakeyama, H., Nakamura, K., Izumiyama-Shimomura, N. et al. (2008). The teleost Oryzias latipes shows telomere shortening with age despite considerable telomerase activity throughout life. Mech. Ageing Dev. 129: 550–557. Hayflick, L. (1965). The limited in vitro lifetime of human diploid cell strains. Exp. Cell. Res. 37: 614–636. Hou, M., Xu, D., Björkholm, M. et al. (2001). Real-time quantitative telomeric repeat amplification protocol assay for the detection of telomerase activity. Clin. Chem. 47: 519–524. Kim, N.W., Piatyszek, M., Prowse, K. et al. (1994). Specific association of human telomerase activity with immortal cells and cancer. Science 266: 2011–2015. Kirkwood, T.B. (2008). A systematic look at an old problem. Nature 451: 644–647. Kong, R.Y.C., Giesy, J., Wu, R. et al. (2008). Development of a marine fish model for studying in vivo molecular responses in ecotoxicology. Aquat. Toxicol. 86: 131–141. Livak, K.J. and Schmittgen, T.D. (2001). Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta Ct) method. Methods 25: 402–408. Meyne, J., Ratliff, R., and Moyzis, R. (1989). Conservation of the human telomere sequence (TTAGGG)n among vertebrates. Proc. Natl. Acad. Sci. U.S.A. 86: 7049–7053. Peterson, D.R., Mok, H., and Au, D. (2015). Modulation of telomerase activity in fish muscle by biological and environmental factors. Comp. Biochem. Physiol. C: Toxicol. Pharmacol. 178: 51–59. Pfennig, F., Kind, B., Zieschang, F. et al. (2008). Tert expression and telomerase activity in gonads and somatic cells of the Japanese medaka (Oryzias latipes). Dev. Growth Differ. 50: 131–141. Southern, E. (2006). Southern blotting. Nat. Protoc. 1: 518–525. Tan, W.H., Witten, P., Winkler, C. et al. (2017). Telomerase expression in the medaka (Oryzias melastigma) pharyngeal teeth. J. Dent. Res. 96: 678–684. Taniguchi, Y., Takeda, S., Furutani-Seiki, M. et  al. (2006). Generation of medaka gene knockout models by target-selected mutagenesis. Genome Biol. 7: R116. Wege, H., Chui, M., Le, H. et al. (2003). SYBR Green real-time telomeric repeat amplification protocol for the rapid quantification of telomerase activity. Nucleic Acids Res. 31: e3. Wienholds, E., Schulte-Merker, S., Walderich, B. et  al. (2002). Target-selected inactivation of the zebrafish rag1 gene. Science 29: 99–102. Yip, B.W.P., Mok, H., Peterson, D. et  al. (2017). Sex-dependent telomere shortening, telomerase activity and oxidative damage in marine medaka Oryzias melastigma during aging. Mar. Pollut. Bull. 124 (2): 701–709. Yu, R.M.K., Chen, E., Kong, R. et  al. (2006). Hypoxia induces telomerase reverse transcriptase (TERT) gene expression in non-tumor fish tissues in vivo: the marine medaka (Oryzias melastigma) model. BMC Mol. Biol. 7: 27.

Chapter 12

Assessments of Medaka Skeletal Toxicity

12.1 ­Introduction Skeletal development is a highly conserved process across vertebrate phyla. Tetrapods (mammals) and teleosts share many aspects of bone and cartilage development, as their earliest shared vertebrate ancestor (Osteoicthyes, i.e., bony fish) already possessed the most basic elements of the endoskeleton – cartilage and bone. Thus, mammals and teleosts share homology of many of the signaling pathways that govern cell migration, proliferation, and/or differentiation during skeletal development. In recent decades, Japanese medaka and other teleost fish models have experienced increasing use as models to understand mechanisms of human skeletal development due to the conservation of common skeletal networks (Apschner et al. 2011; Hall 2005; Witten et al. 2017). Proper skeletal development is critical to the survival of the individual as it serves as a scaffold for further growth and development. Given the strict coordination of molecular and cellular events required for normal skeletal development, it is no surprise that skeletal defects represent one of the most common toxicological endpoints observed in mammals and teleosts (Tyl et al. 2007). Genetic and chemical perturbations in key skeletal developmental pathways have the potential to cause morphological and functional defects during early growth (Gavaia et al. 2014). In this chapter, we demonstrate how embryonic exposure of medaka embryos to prototypic environmental contaminants results in skeletal maladies associated with alterations in osteochondral development. Using wild‐type (southern orange‐red) medaka and specific medaka transgenic reporter models, we illustrate several methods of enhanced in vivo imaging to assess spatiotemporal changes of skeletal dysmorphogenesis during medaka embryonic development. Dithiocarbamates (DTCs) are an important class of compounds with many applications in industry, medicine, and agriculture (US Environmental Protection Agency 2004a). Two widely used DTC fungicidesare tetramethylthiuram disulfide (thiram) and zinc dimethyldithiocarbamate (ziram). Thiram is applied to prevent crop damage in fields and to protect harvested crops from deterioration during storage/transport (US Environmental Protection

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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Agency 2004b). Similarly, ziram is an agricultural fungicide used to control fungal diseases on a wide range of crops. While beneficial in many respects, DTCs may pose a potential risk to aquatic organisms, specifically fish and amphibians, due to the transport and fate of DTCs within aquatic ecosystems following fungicidal and industrial applications. 2,3,7,8‐Tetrachlorodibenzo‐p‐dioxin (TCDD or dioxin) is a widely studied polychlorinated aromatic compound known to impact reproductive, developmental, and cardiovascular systems in mammalian and model vertebrate species (Carney et al. 2006; Matsumura 2009). TCDD mediates numerous biological effects via its interaction with the aryl hydrocarbon receptor (AhR). AhR is a basic‐helix‐loop‐helix Per‐ARNT‐Sim (bHLH‐PAS) transcription factor and functions as a master regulator of drug metabolism and cell regulatory signaling pathways including cell proliferation, differentiation, and apoptosis (Abel and Haarmann‐Stemmann 2010; Chopra and Schrenk 2011). Fish embryos are highly sensitive to TCDD at low parts per trillion (ppt) to parts per billion (ppb) concentrations and exhibit significant developmental abnormalities including pericardial edema, reduced peripheral blood flow, craniofacial malformations, growth retardation, and premature death (Carney et al. 2006; Teraoka et al. 2003). Skeletal dysmorphogenesis, in particular disruption of craniofacial and axial skeletal development, is among the earliest and most sensitive endpoints of TCDD toxicity in fish models (Henry et al. 1997; Hill et al. 2004; Hornung et al. 1999; Dong et al. 2012; Watson et al. 2017).

12.2 ­Methods For many of the same reasons mentioned in previous chapters, medaka offer numerous advantages over traditional rodent models of skeletal development. Daily spawning (20–30 embryos per female), and oviparous and optically transparent development are important considerations that elevate the use of medaka as models of skeletal development. Taken together, these advantages enable detection of delays and malformations at various stages of skeletal development. Skeletal assessment can be conducted at the organismal and structural level through whole‐mount staining for bone and cartilage. Alizarin red S is one of the most commonly used stains to label the presence of ossified bone as it selectively stains hydroxyapatite mineral that comprises bone. Similarly, Alcian blue is widely used to stain the glycosaminoglycans characteristic of cartilaginous structures. For decades, Alizarin red and Alcian blue staining have been the gold standards used to assess skeletal development in rodent toxicity testing. The smaller size of Japanese medaka and other teleosts permits shorter staining and washing steps relative to traditionally used rodent models. The advent of transgenic medaka strains has also enabled a more detailed assessment of cells involved in bone and cartilage development (Hammond and Moro 2012). In these strains, the expression of fluorescent protein is driven by a promoter for a cell‐specific gene of interest. For example, in the transgenic model tg(col10a1:nlGFP) used in the experiments below, expression of a green fluorescent protein containing a nuclear localization signal (nlGFP) is present in cells containing an active promoter for the collagen 10 alpha 1 gene. These cells are involved in the segmental mineralization of the notochordal sheath, an event that precedes perichordal ossification of the vertebral bodies (Renn et al. 2013). An additional transgenic model tg(osx:mCherry) shown below uses mCherry (a red fluorescent protein) to label differentiating osteoblasts expressing osterix (osx) along the

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perichordal centrum which contribute to perichordal ossification (Renn and Winkler 2009). In addition, the use of the fluorescent stains calcein (green) or Alizarin complexone (red) permits a detailed assessment of transgene‐labeled cell populations within the context of mineralized bone. In the following section, we describe how to conduct a developmental exposure of medaka embryos to TCDD and ziram. It is important to note that these protocols can be adapted or modified for use with other chemicals suspected of causing delays or defects in skeletal development. With each experiment, individuals should be assessed daily for signs of overt toxicity (mortality) and to identify the onset of specific skeletal deficits.

12.2.1  Embryonic exposures: dioxin 1. Collect masses of fertilized medaka embryos (20–30/mass) from females and separate (disruption of attachment filaments joining individual eggs) by gentle rolling on a moistened surface. (Note: TCDD is highly lipophilic and will pass through the chorion and bioaccumulate within the embryo mass and oil droplet; there is no need to remove the chorion.) 2. At 4–5 hours post fertilization (hpf) (stages 8–9), transfer embryos to six‐well flat‐ bottomed tissue culture plates containing 3 mL of 1× medaka embryo rearing medium (ERM; 17.1 mM NaCl, 272 μM CaCl2 • 2H2O, 402 μM KCl, 661 μM MgSO4 • 7H2O in 18 MΩ‐cm water, pH 7.4). For each treatment, use up to 15 embryos per well with three or more replicate experimental wells. Each experiment is replicated three or more times. 3. Embryos are treated with 0.01–1 ppb TCDD by adding 3 μL of 1000× TCDD stock concentrations in DMSO, for a duration of one hour. Concurrent DMSO vehicle controls (0.1% v/v) are run with each exposure. 4. Rinse treated embryos four times with 1× ERM. 5. Maintain embryos in 1× ERM at 26 ± 1 °C under 14 hours:10 hours light:dark conditions with renewal of ERM every other day for the duration of the experiment. 6. Process embryos at the desired stage for morphological analysis by whole‐mount Alizarin red S and/or Alcian blue staining for light microscopy, or in vivo Alizarin complexone staining for confocal imaging.

12.2.2  Embryonic exposure: dithiocarbamates 1. Collect masses of medaka embryos (20–30/mass) from females and separate (disruption of attachment filaments joining individual eggs) by gentle rolling on a moistened surface. 2. At 4–5 hpf (stages 8–10), transfer embryos to six‐well flat‐bottomed tissue culture plates containing 3 mL of 1× ERM. For each treatment, use up to 15 embryos per well with three or more replicate experimental wells. Each experiment is replicated three or more times. 3. Expose embryos to thiram or ziram by adding 3 μL of 1000× ziram or thiram working stocks (dissolved in DMSO) to 3 mL of 1× ERM. Concurrent DMSO vehicle controls (0.1% v/v) are run with each exposure.

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4. Maintain embryos at 26 ± 1 °C under 14 hours: 10 hours light:dark conditions for the duration of the experiment (10 days, or until hatching), with static renewal of exposure media every other day. 5. Process embryos for morphological analysis by whole‐mount Alizarin red S and Alcian blue staining for light microscopy, or in vivo Alizarin complexone staining for confocal imaging.

12.2.3  Whole‐mount Alcian blue staining of hatchlings/larvaea 1. Euthanize medaka in 0.125% tricaine methanesulfonate (MS‐222), and fix in 4% paraformaldehyde (PFA) in phosphate‐buffered saline/Tween (PBST) for four hours or overnight. 2. Rinse specimens two to three times with PBST, and wash sequentially in 25% and 50% ethanol/PBST solution for 15 minutes each (R).bb 3. Stain specimens in 0.4% Alcian blue solution overnight (R) (0.4% Alcian blue dissolved in 50% ethanol/3% glacial acetic acid). 4. Rinse two to three times with 50% ethanol, and rehydrate sequentially in 25% ethanol/ PBST and PBST for 15 minutes each step (R). 5. Bleach embryos/larvae in 2% H2O2/0.5% KOH for 30 minutes–2 hours. 6. Wash twice in 0.25% KOH for five minutes (R). 7. Wash with PBST briefly. 8. Digest in 0.05% trypsin in 30% saturated sodium tetraborate solution for 10 or 20 minutes for hatchings and larvae, respectively (R). 9. Wash with 0.25% KOH. 10. Infiltrate and clear samples in a graded glycerol series of 25%, 50%, 75% glycerol in 0.1% KOH (25% glycerol/0.1% KOH. The first step of 25% glycerol in 0.1% KOH is for two hours to overnight (R); the remaining steps in the series shall require overnight infiltration (R). 11. Store at 4 °C in 75% glycerol for long‐term storage.

12.2.4  Whole‐mount Alizarin red S staining of hatchlings/larvaec This protocol describes staining for mineralized bone in 10–20 dpf medaka, and follows the same steps as whole‐mount Alcian blue staining with the exception of Step 3. Here, stain medaka with a 0.05% Alizarin red S solution in 70% ethanol. Dilute from a 0.5% Alizarin red stock (dissolved in water) and adjust final volumes of absolute ethanol and water to reach 70% ethanol. Following staining, wash and rehydrate the samples, and proceed with the remainder of the protocol. Take extra care not to overbleach or overdigest samples. Trypsin concentrations and incubation times may need to be adjusted.

  Times may vary with size of the fish. Those described here pertain to medaka at 10–20 dpf.   (R) = rotating/orbital shaker. c   Times may vary with size of the fish. Those described here pertain to medaka at 10–20 dpf. a

b

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12.2.5  In vivo Alizarin complexone fluorescent staining for mineralized bone matrix 1. Prepare 0.1% Alizarin complexone solution.d Dissolve 50 mg Alizarin‐3‐methylimino‐ diacetic acid (Sigma, USA) in 50 mL of 1× ERM. Adjust pH to 4.5, filter through a 0.22 μM membrane, and store at room temperature. 2. Transfer hatchlings (or newly hatched fry) to a new dish or well. 3. Remove 1× ERM and replace with 0.1% Alizarin complexone solution. 4. Incubate at 26 °C for two hours or overnight. Concentration and incubation time can be adjusted. 5. Wash three times with 1× ERM, five minutes. 6. Proceed with confocal/epifluorescence imaging. Excitation/emission spectrum is 453 nm/560 nm at pH 4.5.

12.2.6  In vivo calcein fluorescent staining for mineralized bone matrix 1. Prepare 0.02–0.05% calcein solution.e For a 0.02% solution, dissolve 10 mg calcein (Sigma, USA) in 50 mL of 1× ERM. Adjust pH to 8.5, filter through 0.22 μM ­membrane, and store at room temperature until use. 2. Transfer hatched embryos to a new dish or well. 3. Remove 1× ERM and replace with 0.02% calcein solution. 4. Incubate at 26 °C for two hours. 5. Wash three times with 1× ERM for five minutes. 6. Proceed with confocal/epifluorescence imaging steps. Excitation/emission spectrum is 495 nm/515 nm at pH 8.5.e

12.2.7  Confocal imaging of embryo/hatchling medaka 1. Dechorionate embryos. a. Score the chorion surface by rolling embryos on p2000 fine grit sandpaper. Take extra care not to rupture the later stage embryos; if embryos have reached 8–10 dpf, it may not necessary to score the chorion. b. Gently wash three times in 1× ERM. c. Transfer embryos to new petri dishes or wells. Wash once more with 1× ERM. d. Perform pronase digestion. Tilt dish or plate at a 45° angle and remove residual ERM. Add 20 mg/mL pronase solution to completely immerse the embryos (100– 150 μL/10 embryos). Incubate at 26 °C for 45 minutes. Carefully remove pronase for reuse, and wash embryos five times in 1× ERM (some individuals may hatch out during pronase digestion). Incubate embryos in hatching enzyme. As before, tilt plate at a 45° angle and remove all residual ERM. Immerse embryos in 1:1 hatching

  Light sensitive, keep covered when not in use.   Light sensitive, keep covered when not in use.

d e

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enzyme:1× ERM solution. Incubate at 26 °C for up to 90 minutes, checking periodically (every 15 minutes) to monitor hatching. Carefully remove the hatchingenzyme for reuse and gently wash embryos three times in 1x ERM. 2. Stain embryos with 0.1% Alizarin complexone or 0.02% calcein as previously described. 3. Anesthetize individuals in 0.02% MS‐222 in 1× ERM. 4. Orient and embed individuals in 0.3–1.0% low‐melt agarose in 0.02% MS‐222. Transfer anesthetized individuals to a glass‐bottomed Petri dish (MatTek Corporation, Ashland, USA). Carefully remove any residual MS‐222 from the dish, and replace with 0.5– 1.0 mL of low‐melt agarose. Using trimmed gel‐loading pipette tips, quickly position individuals into the sagittal or other desired orientation. Allow 2–3 minutes for the gel to harden. 5. Once the gel has hardened, add approximately 1 mL of 0.02% MS‐222 to keep individuals anesthetized. 6. Proceed with confocal imaging (the steps below are general guidelines for imaging on a Zeiss LSM 710 confocal microscope). a. Place the mounted specimen on a stage and identify the region of interest using wide‐field epifluorescence. b. Proceed to set up tracks, channels, and light paths. i.  Select laser line for excitation. ii.  Select dichroic beam splitter to match laser lines. iii.  Select emission range. iv.  If there is emission overlap (bleed‐through), i.e., cross‐excitation between the chosen fluorophores, then use sequential scanning with multiple tracks. Set settings for each track individually. c. Image acquisition settings. i.  Set the number of pixels in the X and Y dimensions. Most imaging software will have an “Optimal” setting that will calculate pixel size based on the diffraction limited resolution of your optics to optimize the spatial resolution, i.e., Nyquist sampling. ii.  Select scan mode. “Frame” averaging is better for fluorophores that are sensitive to photobleaching. iii.  Select scan speed (a.k.a. pixel dwell time). Lower speeds will enhance the signal:noise ratio. iv.  Select gray level resolution (8‐bit, 12‐bit, or 16‐bit). v.  Select the number of scans that are averaged to generate the image. vi.  Set pinhole to 1 Airy Unit to offer the best compromise between signal strength and section thickness. d. Individual image settings. i.  Use range indicator to adjust laser power, photomultiplier (PMT) gain, and offset for maximum dynamic range. ii.  Begin with low laser power and increase PMT gain (up to 70% of maximum voltage). iii.  Adjust the laser power if the signal is still too low; note that high laser power may cause rapid photobleaching or cell death. iv.  If doing sequential scanning, repeat for each track.

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v.  If taking serial sections in the z‐dimension, ensure that each signal is within the dynamic range throughout the sectioned area. e. Capture the image and save as an original .lsm file. f. Process images and save as a new file.

12.2.8  Morphological assessments 1. Open maximum intensity projections .lsm files in ImageJ software (Figure 12-1). 2. (Optional) Split image into individual channels. 3. Adjust the brightness, contrast, and sharpness to desired levels for Alizarin complexone (ALC) (or Calcein) channel 4. Using a combination of drawing/tracing tools: a. Trace the border of the mineralized centra to measure the two‐dimensional area via the “Polygon Section” tool. b. Repeat the same process with the intervertebral ligament area in between the mineralized centra. c. Trace the neural and hemal arches using the “Segmented Line” tool. d. Measure the lateral width and dorsoventral height of each centrum of interest with the “Straight Line” tool. e. For each structure, take the mean of three measurements. 5. Normalize measurements from control fish to 1.0 and report corresponding measurements in treated animals as a proportional increase or decrease relative to the controls.

NA17

IVL 17

C17

HA17

NA18

IVL 18

C18

HA18

NA19

IVL 19

C19

HA19

Figure 12-1.  Morphometric assessment of vertebrae structures in ImageJ software. Centra (C) and intervertebral ligament (IVL) area, length of neural arches (NA) and hemal arches (HA) are measured for vertebrae 17–19. The image shown is a confocal maximum intensity z‐stack projection of a 20‐ dpf tg(col10a1:nlGFP) medaka stained with Alizarin complexone (20× magnification, scale bar 50 μm). Source: Adapted from Watson et al. (2017).

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12.3 ­Results and Discussion 12.3.1 Dithiocarbamates Though widely used, only a handful of studies have investigated the toxicological effects of DTCs on fish embryonic development. Here, we demonstrate that exposure to low μM concentrations of thiram results in severe skeletal deficits. The most dramatic effects are observed in the developing notochord 10 days post initiation of exposures, and they manifest as lordosis, kyphosis, and/or scoliosis of the spine. Additionally, severe undulation or kinking of the notochord is observed with embryonic DTC exposures. Phenotypes are not mutually exclusive as medaka display multiple skeletal abnormalities at these concentrations. Alcian blue staining of whole‐mount medaka reveals impaired deposition of axial and hypural cartilage/proteoglycans at 10 days post exposure (see protocol above) (Figure 12-2). Assessment of notochord mineralization through staining with Alizarin red (see protocol above) demonstrates a significant loss of calcified mineral matrix in DTC‐ treated animals compared with age‐matched controls (data not presented). In vivo confocal imaging of the medaka tg(osx:mCherry) transgenic reporter line illustrates severe undulation of the notochord with a diminished osx + expression and calcein staining (Figure 12-3). These results are consistent with previous reports demonstrating the sensitivity of the

(a)

+DMSO (b)

+DMSO (c)

(d)

+ 5 μM Ziram

+ 5 μM Ziram

Figure 12-2.  Images from DMSO‐ or 5 μM ziram‐treated medaka at 10 dpf. (a, c) Bright‐field images. (b, d) Representative images of Alcian blue‐stained medaka. Images are 2× magnification captured using Nikon SMZ1500 dissecting microscope; scale bars 500 μm.

Assessments of Medaka Skeletal Toxicity

+ DMSO

(a)

+ 5 μM Ziram

325

(b)

osx:mCherry/calcein

+ DMSO

(c)

+ 5 μM Ziram

(d)

Figure 12-3.  Confocal images of tg(osx:mCherry) medaka stained with Calcein at 10 dpf. (a, b) Whole‐mount medaka treated with DMSO and 5 μM ziram, respectively; 10× magnification; scale bars 200 μm. (c, d) 20× Magnification of regions from (a) and (b), respectively; scale bars 50 μm. Images are representative serial z‐stack projections of 6.74 μm, EC‐Plan Neofluar 10/0.3 M27 objective at 20× magnification (10× objective, 2× zoom). Scale bars 50 μm.

developing skeletal system to exposure of these pesticides, including formation of craniofacial abnormalities, wavy distortions of the notochord, and disorganized somites (Leonardus van Boxte et al. 2010). While the exact mechanism of these developmental skeletal maladies is not known, it appears that dithiocarbamates maintain the ability to disrupt critical skeletal regulatory networks associated with notochord development, organization, and maturation. The ability to directly observe individual embryos and select skeletal phenotypes with the combined illumination of specific skeletal cell types in medaka illustrates the utility of this comparative model for assessment and mechanistic determination of chemically induced skeletal dysmorphogenesis during development.

12.3.2 Dioxin Using 2,3,4,7‐tetrachlorodibenzo‐p‐dioxin as a prototypic osteochondral disruptor, we demonstrate modulation of axial osteochondral development through attenuation of mesenchymal cell recruitment, cell migration, and mineralization (Dong et al. 2012; Watson et al. 2017). The observed effects on mineralization are consistent with modifications in cell number and cell localization of transgene‐labeled osteoblast and osteoblast progenitor cells. For example, transgenic tg(col10a1:nlGFP) medaka counterstained for mineralized bone matrix (Aliziran complexone) and imaged in vivo using confocal microscopy demonstrates skeletal alterations within medial and rostral vertebrae in relation to qualitative

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Medaka

s­ patiotemporal analysis of osteoblast and osteoblast precursor cell populations. Detailed examination of morphological structures is additionally possible using a defined morphological matrix (see section 12.2; Figure 12-1) and can be quantified through simple measurements of skeletal structures. Here, we demonstrate that TCDD impacts axial bone development through an overall attenuation of skeletal mineralization, resulting in truncated centra, increased intervertebral ligament area, and decreased hemal and neural arch structures at 10 dpe (Figure 12-4). Overall, we demonstrate that sublethal TCDD exposure impacts osteoblast proliferation and/or differentiation during critical periods of bone growth and mineralization in medaka.

col10a1:n1GFP/ALC

+ DMSO

(a)

+ DMSO

(b) + 0.3 nM TCDD

(c)

+ 0.3 nM TCDD

(d)

Figure 12-4.  Confocal images from tg(col10a1:nlGFP) medaka stained with Alizarin complexone (ALC) for mineralized bone matrix at 10 dpf. Shown are vertebrae 17–19 (a, c) and caudal fin structures (b, d). Compared to DMSO‐treated animals (a, b), medaka exposed to 0.3 nM TCDD (c, d) demonstrate overall fewer col10a1:GFP+ osteoblasts localized along the centra and caudal fin. As a result, we observe overall deficits in mineralization of these structures. Images are representative serial z‐stack projections of 6.74 μm, EC‐Plan Neofluar 10×/0.3 M27 objective at 20× magnification (10× objective, 2× zoom). Scale bars 50 μm.

Assessments of Medaka Skeletal Toxicity

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­References Abel, J. and Haarmann‐Stemmann, T. (2010). An introduction to the molecular basics of aryl hydrocarbon receptor biology. Biol. Chem. 391: 1235–1248. Apschner, A., Schulte‐Merker, S., and Witten, P.E. (2011). Not all bones are created equal – using zebrafish and other teleost species in osteogenesis research. Methods Cell Biol. 105: 239–255. Carney, S.A., Prasch, A.L., Heideman, W., and Peterson, R.E. (2006). Understanding dioxin developmental toxicity using the zebrafish model. Birth Defects Res. A Clin. Mol. Teratol. 76: 7–18. Chopra, M. and Schrenk, D. (2011). Dioxin toxicity, aryl hydrocarbon receptor signaling, and apoptosis‐persistent pollutants affect programmed cell death. Crit. Rev. Toxicol. 41: 292–320. Dong, W., Hinton, D.E., and Kullman, S.W. (2012). TCDD disrupts hypural skeletogenesis during medaka embryonic development. Toxicol. Sci. 125 (1): 91–104. Gavaia, P.J., Cancela, M.L., and Laize, V. (2014). Fish: a suitable system to model human bone disorders and discover drugs with osteogenic or osteotoxic activities. Drug Discovery Today: Dis. Models 13: 29–37. Hall, B.K. (2005). Bones and Cartilage: Developmental and Evolutionary Skeletal Biology. New York: Academic Press. Hammond, C.L. and Moro, E. (2012). Using transgenic reporters to visualize bone and cartilage signaling during development in vivo. Front. Endocrinol. (Lausanne) 3: 1–8. Henry, T.R., Spitsbergen, J.M., Hornung, M.W. et al. (1997). Early life stage toxicity of 2,3,7,8‐tetrachlorodibenzo‐p‐dioxin in zebrafish (Danio rerio). Toxicol. Appl. Pharmacol. 142: 56–68. Hill, A.J., Bello, S.M., Prasch, A.L. et al. (2004). Water permeability and TCDD‐induced edema in zebrafish early‐life stages. Toxicol. Sci. 78: 78–87. Hornung, M.W., Spitsbergen, J.M., and Peterson, R.E. (1999). 2,3,7,8‐Tetrachlorodibenzo‐p‐dioxin alters cardiovascular and craniofacial development and function in sac fry of rainbow trout (Oncorhynchus mykiss). Toxicol. Sci. 47 (1): 40–51. Leonardus van Boxte, A., Pieterse, B., Cenijn, P., and Kamstra, J.H. (2010). Dithiocarbamates induce craniofacial abnormalities and downregulate sox9a during zebrafish development. Toxicol. Sci. 117 (1): 209–217. Matsumura, F. (2009). The significance of the nongenomic pathway in mediating inflammatory signaling of the dioxin‐activated Ah receptor to cause toxic effects. Biochem. Pharmacol. 77: 608–626. Renn, J. and Winkler, C. (2009). Osterix‐mCherry transgenic medaka for in vivo imaging of bone formation. Dev. Dyn. 238: 241–248. Renn, J., Büttner, A., To, T.T. et al. (2013). A col10a1: NlGFP transgenic line displays putative osteoblast precursors at the medaka notochordal sheath prior to mineralization. Dev. Biol. 381: 134–143. Teraoka, H., Dong, W., Tsujimoto, Y. et al. (2003). Induction of cytochrome P450 1A is required for circulation failure and edema by 2,3,7,8‐tetrachlorodibenzo‐p‐dioxin in zebrafish. Biochem. Biophys. Res. Commun. 304: 223–228. Tyl, R., Chernoff, N., and Rogers, J. (2007). Altered axial skeletal development. Birth Defects Res. B Dev. Reprod. Toxicol. 80: 451–472. US Environmental Protection Agency (2004a). EPA R.E.D. FACTS: Thiram. www3.epa.gov/ pesticides/chem_search/reg_actions/reregistration/fs_PC‐079801_1‐Sep‐04.pdf US Environmental Protection Agency (2004b). EPA R.E.D. FACTS: Ziram. www3.epa.gov/ pesticides/chem_search/reg_actions/reregistration/fs_PC‐034805_1‐Jul‐04.pdf Watson, A., Planchart, A., Mattingly, C. et al. (2017). Embryonic exposure to TCDD impacts osteogenesis of the axial skeleton in Japanese medaka, Oryzias latipes. Toxicol. Sci. 155 (2): 485–496. Witten, P.E., Harris, M.P., Huysseune, A., and Winkler, C. (2017). Small teleost fish provide new insights into human skeletal disease. Methods Cell Biol. 138: 321–346.

Appendix A

Solutions

Balanced salt solution (BSS) 0.65% NaCl 0.04% KCl 0.02% MgSO4•7H2O 0.02% CaCl2•2H2O 0.001% Phenol red Sterilize and adjust to pH 8.3 with 5% NaHCO3.

Yamamoto’s Ringer’s solution 0.75% NaCl 0.02% KCl 0.02% CaCl2 0.002% NaHCO3 Adjust to pH 7.3 with NaHCO3 (Yamamoto 1939)

Iwamatsu’s balanced salt solution (Iwamatsu’s BSS) 20× Solution A: 130 g NaCl 8 g KCl 4 g CaCl2•2H20 4 g MgSO4•7H20 Adjust to 1 L with Milli Q water and autoclave. Solution B: 5% NaHCO3 in Milli Q water Sterilize by filtration. Dilute 20× Solution A 20‐fold with Milli Q water, autoclave, and adjust pH to 7.4 by addition of Solution B (Iwamatsu 1983). NOTE: There is not much difference between Yamamoto’s Ringer’s solution and Iwamatsu’s BSS. Therefore, in many cases, these solutions are interchangeable.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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Embryo culture medium 1 g NaCl 0.03 g KCl 0.04 g CaCl2•2H2O 0.16 g MgSO4•7H2O Adjust to 1 L with distilled water. 100× stock embryo culture solution prepared as follows is convenient for daily use. 100 g NaCl 3 g KCl 4 g CaCl2•2H2O 16 g MgSO4•7H2O Adjust to 1 L with distilled water. NOTE: Conveniently, embryo culture solution can be replaced by 0.03% (W/V) artificial sea water. Artificial sea salt is commercially available (for example, Red Sea, www.redseafish. com).

Phosphate-buffered saline (PBS) 137 mM NaCl 2.68 mM KCl 8.10 mM Na2HPO4 1.47 mM KH2PO4

PBST PBS containing 0.1% Tween‐20.

­References Iwamatsu, T. (1983). A new technique for dechorination and observations on the development of the naked egg in Oryzias latipes. J. Exp. Zool. 228: 83–89. Yamamoto, T. (1939). Changes of the cortical layer of the egg of Oryzias Iatipes at the time of fertilization. Proc. Imp. Acad. (Tokyo) 15: 269–271.

Attributions

Chapter

Name

Contributed

Chapter 1

Yasuhiro KAMEI Masato KINOSHITA Shin‐ich CHISADA

Edit 1, 1.0., 1.1., 1.2. Edit 1. 1.0., 1.1., 1.2.

Chapter 2

Kiyoshi NARUSE Kazunori YAMAHIRA Yusuke TAKEHANA

Edit 2, 2.0., 2.1., 2.2., 2.3., 2.4., 2.5. 2.1., 2.2. 2.1., 2.2., 2.3., 2.4.

Chapter 3

Ai SHINOMIYA Minoru TANAKA Daisuke KOBAYASHI Shoji FUKAMACHI Yuko WAKAMATSU Tomonori DEGUCHI Hisashi HASHIMOTO Masayuki IIGO Narisato HIRAI

Edit 3, 3.2.1., 3.2.2., 3.2.5. Edit 3, 3.2.5. 3.1.1. 3.1.2. 3.2.1. 3.2.1., 3.2.3. 3.2.1., 3.2.6. 3.2.4. 3.2.5., Column 3.

Chapter 4

Minoru TANAKA Kataaki OKUBO Yuji ISHIKAWA Shigeki YASUMASU Keiji INOHAYA Hiroki YODA Joachim WITTBRODT Rie KUSAKABE Misato FUJITA Akira KUDO Kouichi MARUYAMA Ichiro IUCHI Kenji MURATA Eriko SHIMADA Hisashi HASHIMOTO Norimasa IWANAMI Daisuke KOBAYASHI Yoshiro TAKANO Yuki NAKATANI Daisuke SAITO

Edit 4 4.1.1., 4.1.2. 4.1.2. 4.1.3., 4.2. 4.1.3., 4.1.12., Column 4.1. 4.1.4. 4.1.4. 4.1.5. 4.1.6. 4.1.6., 4.1.12., 4.1.13. 4.1.7. 4.1.7. 4.1.8. 4.1.8. 4.1.9. 4.1.10. 4.1.11., 4.3., 4.4., 4.5., Column 4.4. 4.1.12. 4.1.13. 4.1.14.

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

331

332

Attributions

Chapter

Name

Contributed

Takashi KAWASAKI Shoji FUKAMACHI

Column 4.2. Column 4.3.

Chapter 5

Kenji MURATA Makito KOBAYASHI Sakurako KAMIDE Hirofumi YOKOTA Eri IWATA

Edit 5 5.1., 5.2., 5.3. 5.1., 5.2., 5.3. 5.1., 5.2., 5.3. 5.1., 5.2., 5.3.

Chapter 6

Kiyoshi NARUSE Fumi KEZUKA Shinsuke SEKI Sungki LEE Goro YOSHIZAKI

Edit 6, Column 6. 6.1., 6.1., 6.3. 6.1., 6.1., 6.3. 6.1., 6.1., 6.3. 6.1., 6.1., 6.3.

Chapter 7

Masato KINOSHITA Satoshi ANSAI Yu MURAKAMI

Edit 7, 7.0., 7.1., 7.2., 7.3., 7.4., 7.5., 7.6., Column 7., Appendix 7. 7.0., 7.1., 7.2., 7.3., 7.4., 7.5., Appendix 7. 7.6.

Chapter 8

Yasuhiro KAMEI Yasuko ISOE

Edit 8., 8.0., 8.1., 8.2., 8.3., 8.4., 8.5. 8.1., 8.2., 8.3., 8.4., 8.5.

Chapter 9

Kenji MURATA Taisen IGUCHI Yukiko OGINO Shinichi MIYAGAWA Ryohei YATSU Norihisa TATARAZAKO Anthony SEBILLOT

Edit 9. 9.1., 9.2. 9.1., 9.2. 9.1., 9.2. 9.1. 9.1., 9.2. 9.2.

Chapter 10

Kenji MURATA Drew PETERSON

Edit 10. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10. 10.8. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10. 10.1., 10.2., 10.3., 10.4., 10.5., 10.6., 10.7., 10.8., 10.9., 10.10.

Roy YE Napo KM CHEUNG Michael WL CHIANG Shin‐ichi KITAMURA Rudolf SS WU Doris WT AU

Attributions

Chapter

Name

Contributed

Chapter 11

Kiyoshi NARUSE Doris WT AU Bill WP YIP Helen OL MOK Yoshihito TANIGUCHI Kenji MURATA Seth William KULLMAN AtLee Taylor Darr WATSON Ian Thomas STANCIL Masato KINOSHITA

Edit 11. 11.1., 11.2., 11.3., 11.4. 11.2., 11.3., 11.4. 11.2., 11.3., 11.4. 11.2., 11.3., 11.4. Edit 12. 12.1., 12.2., 12.3.

Chapter 12

Appendix: Solutions

12.2., 12.3. 12.2., 12.3.

333

Index abdominal cavity, 56–58, 74, 83, 99, 104, 117, 146, 148, 149, 295 ablation, 267, 268 acquired immune system, 160, 255 actinotrichia, 174 ACV see anterior cardinal vein adaptation, 31, 33, 41–43 adaptive immune organ, 293 Adrianichthys spp., 34, 40–42 A. oophorus, 36, 42 A. poptae, 36, 42 A. roseni, 36, 42 agarose, 185–187, 192, 219, 221, 228, 230, 232, 234, 236–238, 241–243, 262, 297–298, 311, 322 agglutinin, 162 aggregation, 51, 78, 207, 211 aggressive behavior, 207, 211 aging, 304, 306, 307 aging process, 303 alcian blue, 174–176, 318–320 alcian blue staining, 318–320, 324 algal growth, 290 Alizarin complexone, 319–323, 326 alizarin red staining, 320 alkaline phosphatase, 170, 172, 191, 235, 237, 238, 304 4‐allyl‐2‐methoxyphenol, 26 5alpha‐dihydrotestosterone, 277 17alpha‐methyltestosterone (17MT), 276, 277 AMA see anterior mesenteric artery amplicon, 245, 246, 248, 252, 254, 310 anal fin, 24, 49–50, 99, 113, 173–175, 207, 272, 273, 275–276, 282, 295 androgen, 50, 272, 273, 275–277 androgen receptor (AR), 276 anesthesia, 1, 4–7, 25–29, 185, 186, 221 anterior cardinal vein (ACV), 134, 140, 142, 143, 151 anterior mesenteric artery (AMA), 136, 137, 143 anterior musculature, 108

anti‐androgen, 272, 273, 276, 277 anti‐DIG, 157, 191, 304, 312 anti‐estrogen, 272 aorta, 58, 86, 99, 107, 108, 133, 143, 146, 151 aortic arch, 133, 140, 142 approaching, 179, 207 AR see androgen receptor ARE/GRE, 277 aromatase, 180, 271–273 aromatizable androgen, 272, 273 artificial insemination, 44 artificial sea water, 4, 285–288, 290, 292, 330 aryl hydrocarbon receptor (AhR), 318 asteriscus, 113 Atonal‐homolog 5 (Ath5), 127 atrium, 58, 99, 108, 110, 147, 151–156 attaching, 131, 210 filament, 9–11, 97, 99, 181, 182 attack, 207 Axin 1, 125

BAC, 31, 43, 47 bacteria challenge assay, 290–294 BaitD, 253, 254 balanced salt solution (BSS), 8, 219, 221, 329 basilar artery (BA), 139, 143 beak‐like cell mass, 99, 104, 116 beta ig‐h3, 176 17beta‐methytestosterone, bf1, 116 bf2, 116 biglycan, 176 biological resources, 31–45 biomarker, 272, 273, 276, 277 bisphenol A, 271 blastoderm, 102–104, 114, 115 blastodisc, 99, 284 blastomere, 99, 100 blastopore, 99, 104 blastula, 100, 102, 177, 182

Medaka: Biology, Management, and Experimental Protocols, Volume 2, First Edition. Edited by Kenji Murata, Masato Kinoshita, Kiyoshi Naruse, Minoru Tanaka, and Yasuhiro Kamei. © 2020 John Wiley & Sons Ltd. Published 2020 by John Wiley & Sons Ltd.

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336

Index

blood cell, 107, 133, 135, 137, 138, 140–146, 151, 160 blood circulation, 110, 141, 142, 144, 151 blood island, 99, 104, 107, 133, 151 blood vessel, 55, 57, 80, 82–85, 99, 104, 108, 110, 172, 173, 175 body color, 31, 42, 49, 51–56, 77, 78, 194, 223 bone and cartilage development, 317, 318 matrix, 165–170, 172, 321, 325, 326 bone morphogenetic protein (BMP), 124 bone morphogenic protein 2 (bmp2), 176 bone morphogenic protein 7 (bmp7), 273, 276 Bony fish, 317 Bowman’s capsule, 158, 160 brain heart infusion culture medium, 291 branchial arch, 58, 68, 74, 76, 121, 128–131, 140, 142 breeding, 1–8, 14, 15, 17, 19–21, 23–25, 28, 32, 35, 45, 53, 162, 256, 274, 276, 277, 284–288 breeding system, 1, 2 brine shrimp, 15–21, 23, 272, 286, 288 BSS see balanced salt solution

calcein, 319, 321–325 cardiac sac, 110, 155 cardiomyocyte, 146, 148, 149 cardiovascular system, 146, 148, 318 cartilage, 128–130, 165, 174, 317, 318, 324 Cas, 225, 226, 229, 232 Cas9 protein, 225 caudal artery (CA), 135, 143 caudal division (CaDI), 133, 143 caudal fin, 99, 110, 173–175, 295, 326 caudal vein (CV), 99, 107, 135, 143, 155 cave fish, 32 CCTop, 44, 229 cDNA, 31, 44, 74, 77, 120, 157, 181 Cdt1, 128 cell death, 266, 267, 303, 322 cell‐fate, 261, 263 cell labeling, 125, 263–265 cell migration, 317, 325 cell senescence, 303 Centra, 165, 168, 170, 323, 326 central nerve system, 29, 58–67, 102 centrum, 165–170, 172, 173, 319, 323 cerebelli, 67, 117

chamber, 147, 148, 151, 154, 155, 160, 161, 188, 189, 312 CHAPS lysis buffer, 308 chase, 18, 207 chokh, 125 Chordin, 124 choriogenin H, 181 choriogenin Hm, 181 choriogenin L, 181 choriolytic enzyme, 119–122, 182 chorion, 9, 44, 84, 97, 99, 157, 190, 287, 288, 297, 319, 321 choroidea, 107, 108, 110 chromosome number, 35 chronic effect, 272 circadian rhythm, 31, 32, 66, 78, 80, 285 circle of Willis, 139 circumstance, 23, 272 closed colony, 7 cloudy water, 23, 25 clustered, regularly interspaced, short palindromic repeat (CRISPR), 31, 32, 44, 225–234, 249, 252–255 c‐myb, 143 collagen 10 alpha 1 gene, 318 collagenase, 218, 220 commissural fiber, 117 common cardinal vein (CCV), 133, 143 confocal microscopy, 125, 313–314, 325 conservation, 205, 212, 215, 317 contact, 69, 104, 148, 166, 207, 267 conus arteriosus, 147 cornea, 99, 108, 125 cortical alveoli, 97, 99 cranial division, 133, 143 cranial nerve, 58, 67, 68, 70, 116 craniofacial malformations, 318 Cre‐driver medaka line, 268 Cre‐loxP system, 268 Cre recombinase, 263 CRISPR see clustered, regularly interspaced, short palindromic repeat CRISPR/Cas9, 31, 32, 44, 226–234, 252–255 CRISPR‐RNA (crRNA), 226, 247, 255 CRISPRscan, 229 crisprtool, 229 cryobanking, 215 cryopreservation, 44, 215–223 Cuvierian duct, 99, 104, 106–108, 110, 143, 174 CXCR4, 177 cyst, 85, 179, 180

Index DAV see dorsal anastomotic vein deletion, 225, 226, 229, 244, 245, 250 density of larvae, 4 developmental staging, 283–284 dHAND, 176 DHT, 277 diagnostic evaluation, 272 dimorphism, 42, 43, 179–180, 283 diode laser, 265 dioxin, 318, 319, 325–326 disorganized somites, 325 dispase, 218, 220 disruption, 195, 207, 226, 244, 249, 254, 271, 277, 290, 318, 319 dissection, 26, 58, 113, 221, 290, 292, 294–297 dithiocarbamates (DTCs), 317–320, 324–325 DLAV see dorsal longitudinal anastomotic vessel dmrt1, 180 dmrt1bY, 177, 179, 180 DMY, 177, 179, 180 DNA preparation, 256–259 DNA replication, 303 donor, 218, 223, 253–254 dorsal anastomotic vein (DAV), 137, 138, 141, 143 dorsal aorta (DA), 86, 99, 108, 133, 143, 151 dorsal ciliary vein (DCV), 136, 137, 143 dorsal lip (DL), 99, 102, 114, 115 dorsal longitudinal anastomotic vessel (DLAV), 137, 140, 143 double‐strand break (DSBs), 225, 226, 229 driver line, 263, 265, 267 d‐rR, 248 DTCs see dithiocarbamates ducts of Cuvier (DC), 133

Eagle’s minimum essential medium, 216 ECM see embryo culture medium ecosystem, 281, 318 ecotoxicology, 33, 281–300 EDS Edwardsiella tarda, 291–293 effector line, 263–265, 267 efferent duct (ED), 85, 180 egg deposition, 207, 210, 211 egg discarding behavior, 208, 210–211 egg envelope, 9, 10, 13, 14, 44, 84, 97, 99, 111, 117, 119, 145, 157, 181–185, 187, 189, 257–259 egg release, 207

337

embedding, 88, 185–189, 192–196, 296–299 embryo and larvae histoarray, 297–298 embryo chip, 290, 297–298 embryo culture medium (ECM), 4, 8, 185, 187, 330 embryonic body, 99, 102–104, 106–108, 114, 115, 117, 119, 120, 133, 150, 151, 177, 258, 283 embryonic exposure, 317, 319–320 embryonic shield, 99, 102, 114, 115, 121, 122 emx1, 116 en2, 116 endocrine disrupting chemical, 271–277 endocrine disruptor, 271–275 endoderm, 102, 121, 161–164 endoskeleton, 317 endpoint, 272–274, 282, 290 ENTPD5, 176 Environmental Protection Department (EPD), 281 epiblast, 102 epiboly, 102, 104, 114 epidermis, 51, 55, 102, 128, 173 erythrocyte, 51, 142 esophagus, 162, 163 EST, 44 estrogen, 50, 84, 181, 271, 306 estrogenic chemical, 271 estrogen receptor E2, 271 estuarine species, 281 ethical issue, 1 ethogram, 206 eugenol, 26–29 euthanasia, 1, 4–7, 28–29 evagination, 78, 116, 124–127 eversion, 117 excess feeding program, 21, 22 expression profile, 44 Exp variable do‐residue (VD), 44 eyeless, 125

FA‐100, 26 fathead minnow, 271–273, 281 feeding, 1, 2, 4, 8, 9, 14–24, 32, 78, 256, 286, 288, 289, 293 fgf8, 116 fibroblast growth factor receptor 1 (fgfr1), 276 fin fold, 99, 108, 173 form, 24, 107, 176 ray, 49, 50, 99, 110, 112, 113, 173–175, 264, 275–276

338

Index

fish Early‐Life Stage Toxicity Test, 273, 297 Sexual Development Test, 273 Short‐Term Reproduction Assay, 271, 273, 276 fixation, 88, 147, 156–157, 189–190, 297 floating, 207 fluorescent in situ hybridization (FISH), 304, 313–314 flutamide, 273, 276, 277 FokI, 225, 234 follicle, 79, 80, 83, 84, 88, 180 folliculogenesis, 180 following, 8, 11, 13, 20, 23–25, 35, 58, 67, 113, 148, 150, 157, 170, 187, 191, 192, 207, 211, 218, 225, 230, 231, 233, 240, 241, 243, 254, 286, 287, 290, 312, 318–320 food, 2, 4, 6, 8, 14–19, 21–24, 58, 272, 276, 285, 286, 288 food particles, 4, 288 fosmid clone, 44 foxA2, 114, 118, 162, 163 foxl2, 180 frame‐shift, 249 freshwater snail, 24 frontal bone, 110 ftz‐f1, 178 fugu, 32 fungi, 4, 8, 11, 14, 24, 285, 288 fungicides, 317, 318

gallbladder, 58, 99, 108, 110 ganglion, 64, 69, 117, 118, 127, 128 gastrula, 102, 114, 115, 120, 122, 143, 177 gastrulation, 102, 114, 121, 124, 125, 146, 177 gata‐1, 143, 144 geminin, 148 gender, 283, 291 generation cycle, 32 genetic and chemical perturbations, 317 genetic introgression, 212 genetic manipulation, 225 genome editing, 31, 32, 44, 225–259 germ cell, 84, 85, 176, 179–180, 215–223, 249 transplantation, 215–223 germ ring, 102 gill arch, 108 glomerulus, 86, 87, 158, 160 glyceraldehyde‐3‐phosphate dehydrogenase (gapdh) promoter, 264 glycosaminoglycans, 318

gonadal maturation, 205 gonadal somatic precursor, 177 gonopodium, 276 goosecoid, 114, 115 GPHS fixative, 299 granulosa cell, 84, 180 growth curve, 7–29 retardation, 318 guanophore, 54, 99, 107, 108, 110 guideline, 4, 6, 88, 234, 271, 272, 274, 292, 322 gum acacia, 299 guppy, 281 gut, 57, 58, 74, 83, 99, 107, 110, 113, 135, 137, 144, 162–164

hatching enzyme, 13, 14, 31, 44, 108, 119, 120, 122, 123, 181–184, 321, 322 hatching gland, 13, 14, 31, 44, 108, 119, 120, 122–123, 181–184, 321, 322 hazard, 272 Hd‐rR, 7, 17, 19, 21, 28, 54, 222 head‐up, 207, 208, 210, 211 heartbeat, 106, 133, 142, 145, 148, 149, 151, 152, 293 heart tube, 146–151, 153 heart valve, 151, 156 heat shock, 222, 261–268 promoter, 261–265, 268 heat shock element (HSE), 264 heat shock protein 70.1 (hsp70.1), 261–264 hemal arch, 165–167, 173, 323 hemangioblast, 133, 142 hematopoiesis, 143–146 hematopoietic cell, 142, 143 hepatic portal vein, 138 hepatic vein (HeV), 108, 137, 138, 140, 141, 143 heteroduplex mobility assay (HMA), 225, 244–252 HNI‐II, 17, 19, 21 HO5, 17, 19, 21, 28 holocrine, 122 homoduplex, 244–246 homology‐directed repair (HDR), 226 hoxAa, 176 hoxAb, 176 Hoxb8a, 176 hoxDa, 176 HuC promoter, 264

Index husbandry, 26, 44 hyaloid artery (HyA), 137, 139, 143 hyaloid vein (HyV), 137, 139, 143 hydroxyapatite mineral, 318 hypoblast, 102, 114, 115, 120–122 hypothalamo‐pituitary‐gonadal axis, 273 hypoxia, 267, 283

ICM see intermediate cell mass ImageJ, 304, 312, 314, 323 immune system, 291 immunohistochemistry (IHC), 188, 283, 294, 304 immunoreaction, 191–192 immunotoxicity, 290–291 inbred strain, 7, 8 infrared laser, 261, 268 infrared laser‐evoked gene operator (IR‐LEGO), 261, 265 infrared light, 261 insertion, 225, 226, 244, 245 in situ hybridization (ISH), 80, 97, 120, 122, 157, 161, 162, 283, 294, 304 intermediate cell mass (ICM), 133, 142–144 internal ear, 72, 73, 108 interneuron, 116, 117 intersegmental vein, 140 intersex, 49, 273 intervertebral ligament area, 323, 326 in the wild, 1–3, 53 in vivo confocal imaging, 324 IR‐LEGO system see infrared laser‐evoked gene operator ISH see in situ hybridization

jaw, 99, 113, 119–121, 123, 128–132 juvenile, 15, 23, 58, 60, 61, 86, 88, 113, 170, 265, 272, 273, 276, 281, 285, 288, 294

Kaga, 17, 19, 21, 28, 29, 265 11 ketotestosterone (11KT), 276 kinking of the notochord, 324 knock‐in (KI), 225, 226, 252–255 knock‐out (KO), 44, 225, 226, 246–252, 306 krox20, 116 Kupffer’s vesicle, 99, 104, 107, 195–196 Kyoto‐Cab, 17, 19, 21, 26–29 kyphosis, 324

339

labyrinth, 72, 73, 99, 108 LADGA lagena, 73, 74, 113 Lakritz, 127 larva, 8, 9, 14, 18, 20, 21, 121, 167, 168, 170, 174, 175, 218, 221, 256, 292, 293 Larval Amphibian Growth and Development Test (LAGDA), 271 latipes species, 36–38, 40, 42 leaving, 19, 139, 182, 208 Leibovitz L‐15 medium, 219 lens, 57, 99, 104, 108, 119, 121, 124, 125, 127, 152, 153, 155, 185, 197 life‐cycle, 1, 2, 33, 274, 281, 282 lifespan, 2, 303 light/dark cycle (14L–10D), 2, 8, 13, 78, 80, 287, 290 lighting conditions, 2, 8, 290 lighting cycle, 13 lineage analysis, 122, 266 linear eukaryotic chromosomes, 303 lipophilic, 319 lobule, 82, 180 local gene induction, 261 long‐term cell labeling, 263, 264 lordosis, 324 loxP, 263, 264, 266, 267 L–R axis, 195 lymphocyte, 83, 160–162, 294 lymphoid enhancer‐binding factor‐1 (lef1), 273, 275, 276 lymphoid organ, 160

marginal blastomere, 100 marine biota, 281 marine ecotoxicology, 33, 281–300 marine medaka, 282–294, 307 masculine sex characteristic, 275 maternal mRNA, 267 mating, 2, 7, 8, 78, 209, 222, 251, 275 maturity, 2, 4, 24, 222, 249, 264, 267, 281 Mauthner cell, 67, 117 MB‐EW, 9, 14 MB stock, 9 medaka cardiac myosin light chain 2 (mcmlc2), 146, 148–151, 153–157 Medaka Extended One Generation Reproduction Test (MEOGRT), 271, 272, 274 medaka hemopexin‐like protein (mWap65‐1), 176 medaka‐related species, 31–44

340

Index

medial longitudinal fasciculus, 67, 116 meiosis, 84, 179, 180 membranous fin, 99, 108, 173–176 menopause, 306 MEOGRT see Medaka Extended One Generation Reproduction Test mesenchymal cell, 128, 158, 173, 176, 325 mesoderm, 102, 131, 142–144, 146–148, 160, 176–178, 196, 264 metamorphosis, 51, 272 metencephalic artery (MtA), 136, 137, 142, 143 methylcellulose, 185–188 methylene blue (MB), 9, 13, 14, 25 microangiography, 133, 135, 142 microhomology mediated endo joining (MMEJ), 226 microinjection, 9, 11, 45, 133, 226, 227, 232, 234, 248–249, 254, 292 micromanipulator, 219, 221 microorganisms, 11, 14 micropipette, 145, 219–222, 238, 243, 256–258, 292, 293 micropyle, 97, 99 middle cerebral vein (MCeVs), 136, 137, 140, 143 mineralization, 165–168, 170, 172, 318, 324–326 mineralocorticoids, 81, 277 The Ministry of the Environment, Japan, 205, 271 Ministry of the Environment, Japan (MOE), 271 Mitf, 195 mitochondria, 33, 35, 215 mitotic division, 179 mMYH, 131 modular assembly, 226 molten agarose, 298 morphology, 42, 49–56, 82, 86, 119, 126 morula, 100 mosquitofish, 276, 281 mounting, 97, 185–189, 192, 296–297 MS‐222, 27–28, 185, 186, 256, 292, 320, 322 Müller glial cell, 127 multiple generation, 272 multipotent, 127 mummichog, 281 muscle, 56, 58, 67, 68, 70, 71, 80, 105, 130–132, 153, 169, 174, 221, 256, 276, 304, 310 muscle segment homeobox C (MsxC), 276 myoD, 176 myosin light polypeptide 2 (mylz2), 131 myotome, 105, 168, 170

nanos, 177, 178 nasal ciliary artery (NCA), 135, 137, 143 National Bio‐Resource Project (NBRP) Medaka, 3, 31, 43–45, 268, 306 nephrogenesis, 158–160 neural arch(es), 165–167, 173, 323, 326 neural crest, 51, 102, 128, 130, 194, 264 neural groove, 114 neural keel, 114, 116, 126 neural spine, 113 neurocoel, 104, 116, 117 neuroectoderm, 125 neurula, 103–104, 114–115, 128, 177, 284 neurulation, 113, 116, 125 nodal, 125 Noggin, 124 nonhomologous end joining (NHEJ), 226 nonylphenol, 271 notochord, 99, 107–110, 129, 133, 137, 142, 165–170, 172, 324, 325

off‐target, 229, 249, 253 oil droplet, 97, 99, 100, 105, 151–153, 155, 319 ojo plano (opo), 127 olfactory primordium, 118 oligonucleotide, 125, 226, 228–230, 246, 303, 311, 312 OlMA, 131 omnivorous, 286 oogenesis, 2, 83, 218 o,p' ,‐DDT, 271 optic artery (OA), 135, 137, 143 optic bud, 103–104, 114, 126 optic tectum, 58, 59, 66, 113, 115–117, 119, 124, 128, 129 optic vein (OV), 137, 143 optic vesicle, 104, 114, 124–127 orange‐red variety, 205 Organization of Economic Cooperation and Development (OECD), 271–277 orifice, 99, 122 Oryzias spp. O. asinua, 36, 42 O. celebensis, 42, 43 O. curvinotus, 33, 36, 40, 41 O. dancena, 33, 35, 36, 43, 281 O. eversi, 36, 42 O. hadyatyae, 42 O. haugiangensis, 35, 40 O. hubbsi, 35, 40

Index O. javanicus, 33–35, 40, 42, 43 O. latipes, 2, 31, 33, 40, 41, 43, 205, 206, 208, 222, 261, 271, 275, 281, 283, 284, 303, 306 O. marmoratus, 34, 42 O. matanensis, 34, 42 O. mekongnensis, 40, 41 O. melastigma, 33, 35, 281–300, 303, 304, 306, 307 O. minutillus, 35, 41 O. nebulosus, 42 O. nigrimas, 42 O. orthognathus, 42 O. pectoralis, 40–42 O. profundicola, 42 O. sakaizumii, 40, 205 O. sarasinorum, 36, 42 O. setnai, 35 O. sinensis, 40, 41 O. soerotoi, 42 O. songkhramensis, 40–42 O. wolasi, 42 O. woworae, 42 oscillation, 152 ossification, 129, 130, 165, 318, 319 osteoblast, 165, 169, 170, 172, 173, 318–319, 325, 326 osteocalcin, 172, 173 osteochondral development, 317, 325 osteochondral disruptor, 325 osteocyte, 165, 172 os‐terix, 176 osx, 318–319 otic placode, 99, 104 Otohime, 6, 288 otolith, 73, 74, 99, 107, 113, 294, 295 otx1, 116 outdoor pond, 206, 209 ovarian cavity, 83, 84, 180 oviduct, 83, 180

PAGE see polyacrylamide gel electrophoresis pancreas, 81, 82, 107, 218 papillary process, 50, 113, 272, 273, 275–276 paraformaldehyde (PFA), 157, 295, 297, 299, 320 paramecium, 16–18 particle size, 4, 6, 15, 23 pasteurization, 9, 11, 12 pathogenic bacterial infection, 290 pax, 54, 114, 116, 118, 125, 127, 172 P450c17, 180

341

PCNA marker, 303, 304 pCS2+hSpCas9, 227, 232 PCV see posterior cardinal vein pDR274, 227–229 pectral fin, 38 pellet pestles, 308 pelvic fin, 36, 42, 173–175, 295 peptide nucleic acid (PNA), 304, 313 periblast, 99, 100, 102 pericardial edema, 318 perichordal centrum, 165, 167–170, 172 peritoneal cavity, 110, 218, 220–223, 292 peritoneum, 55, 56, 58, 110 perivitelline space, 97, 99 peroxide, 14 pharyngeal teeth, 110 pharynx, 119, 120, 162 PHBC see primordial hindbrain channel photoperiod, 31, 205, 285–286 phylogenetic relationship, 31, 35–44 physical map, 44 physiology, 26, 42, 55 picking, 208, 210 pillow, 99, 104, 115, 121 pineal gland, 99, 108–109, 115, 118 pit organ, 110 plastic resin, 192–196 ploidy, 222 PMBC see primordial midbrain channel PNA see peptide nucleic acid pollutant, 281, 283 polster, 99, 104, 115, 116, 121, 122 polyacrylamide gel electrophoresis (PAGE), 246, 303 polychlorinated aromatic compound, 318 positioning, 155, 207, 208 posterior cardinal vein (PCV), 133–137, 140, 143 postspawn emission, 209 powdered food, 4, 6, 15–17, 19–23 prechordal hypoblast, 114, 115 premature death, 318 primary head sinus (PHS), 140 primary intersegmental vessel, 136 primary mechanosensory neuron, 116 primordial germ cells (PGCs), 176–179, 218 primordial hindbrain channel (PHBC), 134, 137, 140, 141 primordial midbrain channel (PMBC), 134–136, 139, 141 progesterone, 277 prohibitin, 176

342

Index

pronase, 321 protospacer adjacent motif (PAMs), 226

quick circle, 207–209 Quintet, 19, 21, 28 quivering, 207–209

rag1, 161, 162 ray node, 113 real‐time quantitative telomeric repeat amplification protocol (RTQ‐TRAP), 303, 308–309 re‐annealing, 252 rearing, 1–4, 14–16, 18, 25, 32, 267, 274, 284–290, 292, 319 rearing schedule, 2–7 recipient, 215, 218–223 the Red List, 205 reduced peripheral blood flow, 318 rejection, 160, 208 repeat variable diresidues (RVDs), 226, 239 reproductive behavior, 205–212 reproductive maturation, 272 reticular formation, 116, 117 reverse‐sex, 13 rhombencephalic isthmic construction, 116, 117 rhombomere, 115, 117 rib, 173 ribonucleoprotein enzyme, 303 risk, 13, 272, 293, 304, 318 RNA‐guide endonuclease (RGEN), 226 Robo, 128 Rohon‐Beard cell, 116 roof plate, 114, 116, 117 rug1, 176 rug2, 176 rx, 125, 126

salinity, 31–34, 41, 284–286 scale, 54, 55, 57, 113, 264 school, 208, 211 scleroblast, 174 sclerotomal cell, 165, 170, 172 scoliosis, 324 Scre/SloxP, 264 sdf‐1a, 177, 178, 180 seawater medaka, 281–300 secondary intersegmental vessels, 137, 143 secondary sexual characteristics, 6, 206, 272

second polar body, 99, 222 semicircular canal, 73, 108 sensory placode, 102 sewage effluent, 281 sex characteristics, 275 sex differentiation, 177, 180 sex hormones and telomere length, 306 sex ratio, 273, 274, 285 sexual dimorphism, 42, 43, 179–180 sexually maturity, 4, 272, 273, 281 shedding, 208, 210, 211 sheepshead minnow, 281 SH‐EW, 9, 12, 13 signal peptide peptidase‐like‐2, 176 simultaneous emission, 209, 210 single‐cell gene induction, 261, 268 single guide RNA (sgRNA), 45, 226, 228–232, 247, 253, 255 sinus venosus (SV), 108, 134, 143, 146, 147, 153, 155 Six, 125, 128 skeletal development, 128–130, 317–319 maladies, 317, 325 skull, 130, 173, 294, 295 Slit, 128 sneaker, 209, 210 sneaking, 208–210 social dominance, 207 sodium hypochlorite (SH) stock, 9 soft water, 2 Sonic hedgehog (shh), 114, 125, 127, 276 Southern blotting, 304, 307, 309–312 sox9, 176 sox10, 195 sox9b, 178, 179 spawning, 2, 4, 7–9, 31, 32, 97, 181, 205, 207–211, 222, 272–274, 286, 290 spawning behavior, 206–210, 282 Spemann’s organizer, 102, 114 spermatogenesis, 2, 180, 218 spermatogonial stem cell, 215, 218 sperm cryopreservation, 32, 44–45, 215 sperm release, 207, 208 spiggin, 277 spleen, 57, 58, 99, 110, 144 sppl2, 176 staining, 74, 84, 191–192, 320, 324 Standard Operating Procedure (SOP), 281, 290–293, 295–297, 299–300, 307–314 steroidogenic cell, 85, 180

Index stickleback, 32, 273, 277 STII, 19, 21, 26–29 subintestinal vein (SIV), 133, 136–138, 141, 143 supraintestinal artery (SIV), 133, 136, 137, 141, 143 surface ectoderm, 125, 127 survival, 15, 19, 28, 29, 272, 274, 281, 317 swim bladder, 73, 82, 108, 110, 137, 295 swordtail fish, 264, 276 SYBR Green, 304, 308, 309

tank size, 6 Targeting Induced Local Lesion IN Genome (TILLING) technology, 306 tbx, 161 tbx5a, 175 Technovit 7100, 192–196 teeth, 110, 113, 130, 276 telomerase activity (TA), 303, 306–309 TElomerase Reverse Transcriptase (TERT), 303 telomere(s) lengths, 303, 306, 307, 309–313 maintenance, 303, 306 probe, 304 telomeric DNA, 303, 304 temperate zone, 32 terminology, 207 territory, 207 Test Guideline, 272, 274 testis‐ova, 273, 274 testosterone, 273 2,3,7,8‐tetrachlorodibenzo‐p‐dioxin (TCDD), 318, 319, 326 tetramethylthiuram disulfide, 317 tg(col10a1:nlGFP), 318, 323, 325, 326 tg(osx:mCherry), 318, 324, 325 theca, 84, 180 thiram, 317, 319, 324 thrombospondin‐2, 176 thymus, 74, 83, 160–162, 293, 294 tissue homeostasis, 303 T lymphocyte, 83, 160, 294 tolerance, 31, 33, 34 toxicity, 254, 272, 283, 285, 290 toxicity test, 271–275, 281, 285, 290, 297 trans‐activating crRNA (tracrRNA), 226, 255 transcription activator‐like effector nucleases (TALENs), 45, 225, 226, 234–244, 247, 248 Targeter, 239

343

transgenerational effect, 272, 274 transgenic strain, 15, 172, 173, 277 transglutaminase, 181 transplantation, 122, 125, 215–223, 264, 275 transverse vessel, 133, 143 trenbolone, 276 tricaine methanesulfonate, 26, 320 triploid, 219, 221, 222 tropical fish, 31, 33 TTAGGG, 304, 311 tubule, 85, 86, 88, 158, 160 the 2‐ΔCt method, 309

ubiquitous expressions, 303 The United States Environmental Protection Agency (US EPA), 271 upper jaw, 99, 113 urogenital pore, 207, 208 urogenital protuberance, 113

Validation and Management Group for Ecotoxicity Testing (VMG‐eco), 271 vascular endothelial cell, 133, 142 vasculature, 131–143 vasculogenesis, 133 Vcre/VloxP, 264 vegetal pole, 97, 99, 103, 104, 114, 181 ventral aorta (VA), 133, 143, 146 ventricle, 58, 59, 108, 110, 116, 118, 147, 151–156 vertebral column, 58, 113, 164–173 vertebrate, 33, 51, 58, 76, 77, 81, 83, 113, 116, 117, 127, 130, 131, 133, 134, 136, 137, 139, 140, 142, 143, 146, 160, 164, 165, 173, 176, 181, 194, 195, 303, 306, 318 vinclozolin, 273, 276 vitello‐caudal vein, 104, 106, 107, 134, 151

water quality, 4, 15, 16, 19, 23–25, 283 wavy distortions of the notochord, 325 whole adult histoarray, 294–297 whole adult medaka histoarray, 283, 294, 313 whole‐mount immunohistochemistry, 153, 154, 157–158, 175 whole‐mount in situ hybridization, 120, 148, 149, 154, 156, 157, 163, 189–192 whole‐mount staining, 318 wild Japanese medaka, 205–212

344

Index

wild medaka, 44, 205–212 Wnt, 124, 125 wnt5a, 176 wrapping, 207, 208

Xanthomonas, 226

Yamamoto’s Ringer solution, 8, 184–186, 329 yolk plug, 103, 104 yolk syncytial layer (YSL), 143, 144, 163

zebrafish, 26, 31, 32, 54, 69, 72, 122, 124, 127, 128, 133, 134, 140, 143, 148, 151, 156, 161, 172, 177, 195, 261, 264, 271–273, 292 ZI‐1, 181 ZI‐2, 181 ZI‐3, 181 zinc dimethyldithiocarbamate, 317 zinc‐finger nuclease (ZFNs), 225, 226 ziram, 317–319 zona pellucida (ZP), 273

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