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Copyright and use of this thesis This thesis must be used in accordance with the provisions of the Copyright Act 1968. of material protected by copyright may be an infringement of copyright and copyright owners may be entitled to take legal action against persons who infringe their copyright. Section 51 (2) of the Copyright Act permits an authorized officer of a university library or archives to provide a copy (by communication or otherwise) of an unpublished thesis kept in the library or archives, to a person who satisfies the authorized officer that he or she requires the reproduction for the purposes of research or study. The Copyright Act grants the creator of a work a number of moral rights, specifically the right of attribution, the right against false attribution and the right of integrity. You may infringe the author’s moral rights if you: - fail to acknowledge the author of this thesis if you quote sections from the work - attribute this thesis to another author - subject this thesis to derogatory treatment which may prejudice the author’s reputation For further information contact the University’s Director of Copyright Services sydney.edu.au/copyright The Morphological and Genetic Characterisation of Pelvic Finless Zebrafish

Emily Kate Don

The Laboratory of Development and Disease Models

A thesis submitted in fulfilment of the requirements for the

degree Doctor of Philosophy (Medicine)

in the

Discipline of and Histology, Bosch Institute

The University of Sydney

2013

i Declaration

I hereby certify that this thesis does not contain, without appropriate acknowledgement, any material previously submitted for a degree or diploma in any university, except where due reference is made. I also certify that this thesis does not contain, to the best of my knowledge, any material previously published or written by another person, except where due reference is made.

Emily Kate Don

Dr Nicholas Cole

ii Acknowledgments

I would like to say thank you to all the people from the Anderson Stuart and zebrafish communities who have assisted me during my time as a PhD student.

Firstly, I would like to deeply thank my supervisor Dr. Nicholas Cole who has lived and breathed this project alongside me, through all its ups and downs. In particular, thank you for allowing me this opportunity and the invaluable training you have provided. Thank you for your ongoing enthusiasm, input and support (both professionally and personally). It has been an interesting ride from setting up a new laboratory all the way through to changing institutions. I wish you the best of luck in your new endeavour over at ASAM.

I would also like to thank the Australian government for awarding me an Australian Postgraduate Award to allow me to undertake my doctoral research.

A big thank you needs to go to my mapping team: Dr. Tanya de Jong-Curtain and Associate Professor Joan Heath. Thank you for taking me into your lab and teaching positional to me. Thank you for your ongoing assistance and input.

To Dr. Louise Cole, Dr. Donna Lai, Dr. Dane King, Dr. Errin Johnson and Dr. Ian Kaplin, thank you for your technical knowledge that made this project possible.

To the lab, and in particular, Claire and Andy, I’m not sure that thank you is a big enough word. You have shared this project with me, significantly contributed, been there for my little victories, listened to my complaints and most of all, put up with me on a daily basis!

My time in the Anderson Stuart building would not have been the same without the social club. Thank you to all who have made my Friday evenings over the past few just that little bit more entertaining. Keep up the tradition, we all know that the best collaborations are made over a nice cold beer!

I also owe a huge thank you to my . Mum and Dad, thank you for instilling in me your love of education and knowledge. Thank you for teaching me, by example, that you can have anything you want in , if you are prepared to work hard enough for it. Thank you for the proofreading of this thesis. If that doesn’t show unconditional love, I don’t know what does. To Claire and Hannah, thank you for your support during this project and always inspiring me to finish and get a real job 

My biggest and most heartfelt thanks must go to Kjetil. This would not have been possible without you, and your love and support. You have always been there to celebrate

iii the highs with me and provide a hug that fixed everything during the lows. I’m sorry you now know so much more about pelvic fins than you ever wanted to know. I am so excited for our wedding in February and cannot wait to begin of the rest of our together. I love you.

iv Table of contents

Declaration...... ii

Acknowledgments ...... iii

Table of contents ...... v

Abstract ...... ix Research publications and awards ...... xi List of abbreviations ...... xiii Nomenclature ...... xv List of Tables and Figures ...... xvi List of Tables ...... xix

1. Introduction ...... 1 1.1. Introduction to development ...... 1 1.1.1. Introduction to zebrafish as developmental models ...... 2 1.1.2. Advantages of utilising zebrafish as developmental models ...... 3 1.1.3. Zebrafish fins as a model for development ...... 6

2. Literature Review ...... 8 2.1. Updates to literature review...... 29 2.1.1. Examination of Euphanerops ...... 29 2.1.2. Publication of the African genome...... 29

3. Materials and methods ...... 30 3.1. Production and manipulation of zebrafish ...... 30 3.2. Molecular methods ...... 34 3.3. Staining ...... 42 3.4. Microscopy ...... 46 3.5. Statistical methods ...... 46 3.6. Sequence analysis and alignments ...... 47

4. Morphological characterisation of pelvic finless zebrafish ...... 48 4.1. Introduction ...... 48 4.2. Results ...... 49 4.2.1. Embryonic morphology of pelvic finless zebrafish ...... 49 4.2.2. Larval morphology of pelvic finless zebrafish ...... 52 4.2.3. Adult morphology of pelvic finless zebrafish ...... 54 4.2.4. Pelvic finless zebrafish display abnormal development ...... 57 4.2.5. Pelvic fins of pelvic finless zebrafish lack an apical ectodermal ridge ...... 60 4.2.6. Incomplete penetrance of the pelvic finless phenotype ...... 64

v 4.3. Discussion ...... 66

5. Analysis of pelvic fin gene expression ...... 69 5.1. Introduction ...... 69 5.2. Results ...... 71 5.2.1. Sense probes...... 71 5.2.2. pitx1 expression ...... 71 5.2.3. tbx4 expression ...... 73 5.2.4. tbx5 expression ...... 75 5.2.5. fgf10a expression ...... 77 5.2.6. sp8 expression ...... 79 5.2.7. fgf8 expression ...... 81 5.2.8. shh expression ...... 83 5.3. Discussion ...... 85 5.3.1. Overview of results ...... 85 5.3.1.1. Gene expression in wild-type zebrafish ...... 85 5.3.1.2. Gene expression in pelvic finless zebrafish ...... 86 5.3.2. Do pelvic finless zebrafish establish an apical ectodermal thickening? ...... 86 5.3.3. Consideration of genetic pathways that lead to apical ectodermal thickening formation ...... 87 5.3.3.1. Specification/initiation of pelvic fins ...... 87 5.3.3.2. Formation of the apical ectodermal thickening in pelvic fins ...... 89 5.3.3.3. Genetic pathways that lead to the onset of fgf10 expression...... 90 5.3.4. Limitations of the gene expression analysis ...... 90 5.3.5. Linking phenotype to genotype ...... 91

6. Positional cloning of pelvic finless zebrafish ...... 93 6.1. Introduction ...... 93 6.2. General overview of positional cloning ...... 94 6.2.1. Meiosis, genetic recombination and linkage ...... 94 6.2.2. Positional cloning resources ...... 96 6.2.2.1. Polymorphic zebrafish strains ...... 96 6.2.2.2. Positional markers ...... 97 6.2.2.3. Genomic Resources ...... 99 6.2.3. Positional cloning workflow ...... 99 6.2.3.1. Creating heterogeneous zebrafish...... 99 6.2.3.2. Low resolution mapping ...... 100 6.2.3.3. Intermediate resolution mapping ...... 100 6.3. Chapter specific methods ...... 102 6.3.1. Generation of mapping pedigrees ...... 102 6.4. Results ...... 103 6.4.1. Low resolution mapping ...... 103 6.4.1.1. Genome scan ...... 103 6.4.1.2. Individual PCR analysis ...... 106 6.4.2. Intermediate resolution mapping ...... 108 6.5. Discussion ...... 120 6.5.1. Overview of results ...... 120

vi 6.5.2. Identifying pelvic finless genomic lesion ...... 120

7. Targeted sequencing of the pelvic finless locus ...... 121 7.1. Introduction ...... 121 7.2. Introduction to genetic mapping by next generation sequencing ...... 122 7.2.1. Region-specific targeted enrichment next generation sequencing ...... 124 7.3. Chapter specific methods ...... 127 7.3.1. Extraction of genomic DNA ...... 127 7.3.2. Targeted enrichment ...... 127 7.3.3. Next generation sequencing ...... 128 7.3.4. Bioinformatics workflow...... 128 7.3.4.1. Detection of large insertions or deletions ...... 128 7.3.4.2. Detection of single nucleotide polymorphisms by BGI ...... 129 7.3.4.3. Detection of small insertions and deletions by BGI ...... 129 7.3.4.4. Analysis of SNP and Indel data ...... 129 7.3.5. Tbx4 transcript and analysis ...... 130 7.3.5.1. Splice site enhancer analysis ...... 130 7.3.5.2. Profile comparison ...... 130 7.3.5.3. Stability comparison ...... 131 7.4. Results ...... 132 7.4.1. Identifying candidate polymorphisms ...... 132 7.4.1.1. SNPs and Indels...... 132 7.4.1.2. Large deletions or insertions ...... 132 7.4.2. Sequencing candidate genomic lesions ...... 133 7.4.2.1. Nonsense SNPs ...... 133 7.4.2.2. Frameshift Indels ...... 135 7.4.2.3. Missense SNPs ...... 137 7.4.3. Investigation of tbx4 transcript ...... 139 7.4.3.1. Sequence ...... 139 7.4.3.2. Splicing ...... 141 7.4.4. Investigations of the Tbx4 protein ...... 143 7.4.4.1. Amino acid sequence ...... 143 7.4.4.2. Bioinformatic protein analysis...... 147 7.5. Discussion ...... 149 7.5.1. Overview of results ...... 149 7.5.2. Limitations of region-specific targeted next generation sequencing ...... 150 7.5.2.1. Exome sequencing by next generation sequencing ...... 151 7.5.2.2. Whole genome sequencing by next generation sequencing ...... 151 7.5.3. tbx4 as a candidate gene to underlie the pelvic finless phenotype ...... 153 7.5.3.1. Analysis Tbx4 protein and mRNA transcript ...... 154 7.5.3.2. Do the identified amino acid substitutions affect Tbx4 function? ...... 155

8. Ongoing Strategies Towards Linking Genotype to Phenotype...... 158 8.1. Intro ...... 158 8.2. Chapter specific methods ...... 159 8.2.1. DNA injections ...... 159 8.2.2. The Pel2.5 kb-GFP construct ...... 159

vii 8.2.2.1. Description of the Pel2.5 kb-GFP construct ...... 159 8.2.2.2. Analysis of the Pel2.5kb GFP construct ...... 160 8.2.3. Tbx4 rescue constructs ...... 160 8.2.3.1. Generation of Tbx4 rescue constructs...... 160 8.2.3.2. Analysis of rescue experiments ...... 163 8.3. Results ...... 164 8.3.1. Pel2.5kb enhancer drives GFP expression in zebrafish pelvic fins ...... 164 8.3.2. Transgenic rescue experiments ...... 168 8.4. Discussion ...... 172 8.4.1. Overview of results ...... 172 8.4.2. Limitations of rescue experiments ...... 172

9. Final discussion and concluding remarks ...... 175 9.1. Overview of results ...... 175 9.2. Tbx4 ...... 175 9.2.1. Is Tbx4 critical for pelvic fin development? ...... 177 9.2.2. Missense mutations in tbx4 ...... 177 9.2.3. A hypomorphic allele or compensation? ...... 178 9.2.4. Could mutations in tbx4 cause the loss of the apical ectodermal thickening in pelvic finless zebrafish? 179 9.2.5. Implications of a viable mutant Tbx4 model ...... 184 9.3. Conclusion and future directions ...... 185

References ...... 187 Appendices ...... 215 Appendix A ...... 215 Appendix B ...... 224 Appendix C ...... 232 Appendix D ...... 234 Appendix E ...... 239 Appendix F ...... 243 Appendix G ...... 244 Appendix H ...... 247

viii Abstract

Historically, studies of limb development have relied heavily on mouse and chick as models to understand the genetic mechanism of limb induction, identity and outgrowth. Recently, the paired pectoral fins of zebrafish have emerged as an excellent model for dissecting the genetic mechanisms of initiation and early outgrowth. The vertebrate’s hindlimb is evolutionarily derived from pelvic fins, and therefore pelvic fin of zebrafish should provide an excellent model to understand early hindlimb development. Despite this, there have only been a handful of studies that have included pelvic fins and the genetic basis of pelvic fin development is virtually undefined. The goal of this research is to make a novel contribution to the molecular understanding of pelvic fin development in the hope that the zebrafish pelvic fin can complement other traditional models of limb development.

The pelvic finless zebrafish strain represents a naturally occurring phenotype where the adult zebrafish lack pelvic fins. Pelvic finless zebrafish develop in a similar fashion to wild- type siblings and display normal fin structure in the pectoral, dorsal, ventral and caudal fins. The differences between pelvic finless zebrafish and wild-type siblings become apparent during the early stages of pelvic fin development, around 21 days post fertilisation (dpf). Mesenchymal bulges in the prospective pelvic fin regions are observed in pelvic finless zebrafish, however these bulges lack an overlying ectodermal structure. In contrast to wild- type zebrafish, these bulges do not develop an apical ectodermal thickening or an apical fold and cease to continue development. Examination of gene expression in the pelvic regions of pelvic finless zebrafish revealed early expression of the key hindlimb development genes, pitx1 and tbx4, but the expression of the key limb development genes fgf10, fgf8 and shh was never observed.

The positional cloning and targeted next generation sequencing of the genomic lesion responsible for the pelvic finless phenotype is at an advanced stage. A genome scan positioned the pelvic finless locus on chromosome 15 and the closest flanking SSLP markers were subsequently identified. The pelvic finless locus was unambiguously located between SSLP markers z13310/z3760 and z11320 at 54.1 cM and 63.7 cM on chromosome 15. This region was subsequently sequenced by targeted next generation sequencing to reveal any polymorphisms between pelvic finless zebrafish and wild-type siblings which could underlie the pelvic finless phenotype. Bioinformatic analysis and subsequent Sanger sequencing of

ix possible candidates identified three candidate missense mutations in the pelvic finless sequence of tbx4. Ongoing transgenic rescue experiments, DNA binding assays and knockdown experiments are currently being pursued to confirm that these candidate mutations truly underlie the pelvic finless phenotype. On the basis of preliminary results, a mutation in tbx4 appears to be the most promising contender to underlie the pelvic finless phenotype. This novel finding in the field of pelvic fin development is hoped to contribute to the understanding of the molecular mechanisms of hindlimb and pelvic fin development and loss.

x Research publications and awards

Peer reviewed journal articles

Don EK, Currie P, Cole NJ (2012). “The evolutionary history of the development of the pelvic fin/hindlimb.” Journal of Anatomy 222(1): 114-133.

Cole NJ, Hall TE, Don EK, Berger S, Boisvert CA, Neyt C, Ericsson R, Joss J, Gurevich D.B, Currie PD (2011) Development and of the Muscles of the Pelvic Fin. PLoS Biol 9(10).

Don EK, Hall TE, Currie P, Cole NJ (2011). “Morphology of Pelvic Fin Loss in a Zebrafish (Danio rerio).” Journal of Morphology. 272(5): 583-589

Veilleux HD, van Herwerden L, Cole NJ, Don EK, De Santis C, Dixson DL, Wenger AS, Munday PL (2013). “Otx2 expression and implications for olfactory imprinting in the anemonefish, Amphiprion percula.” Open. In press.

Oral presentations

2013 and New Zealand Zebrafish Meeting, Queenstown, New Zealand. Don EK, deJong-Curtain T, Hall T, Winnick C, Heath J, Currie PD, Cole NJ. Pelvic fin loss in a novel zebrafish model.

2012 Bosch Young Investigators Symposium, Sydney Australia. Don EK, deJong-Curtain T, Hall T, Heath J, Currie PD, Cole NJ. Pelvic fin loss in a novel zebrafish model.

2012 Australia and New Zealand Zebrafish Meeting, Victoria, Australia. Don EK, deJong- Curtain T, Hall T, Heath J, Currie PD, Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

2011 Bosch Young Investigators Seminar Series, Sydney, Australia. Don EK, deJong- Curtain T, Hall T, Heath J, Currie PD, Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

2011 Australia and New Zealand Zebrafish Meeting, Queensland, Australia. Don EK, Hall T, Currie PD, Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

2010 Australia and New Zealand Zebrafish Meeting, Sydney, Australia. Don EK and Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

xi Poster presentations

2011 Bosch Young Investigators Symposium, Sydney, Australia. Don EK, deJong-Curtain T, Hall T, Heath J, Currie PD, Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

2011 Bosch Institute Annual Scientific Meeting, Sydney, Australia. Don EK, deJong- Curtain T, Hall T, Heath J, Currie PD, Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

2010 Bosch Young Investigators Symposium, Sydney Australia, Don EK and Cole NJ. The morphological and genetic basis of pelvic fin development and loss in zebrafish.

Awards

2013 Genetics Otago Travel Scholarship, for travel to the 14th annual Australian and New Zealand conference

2012 Bio-Rad Prize for Best Oral Presentation, Bosch Young Investigators Symposium, Sydney, Australia

2012 Bosch Advanced Microscopy Facility, Micrograph of the , 8th place

2012 Postgraduate Research Student Support, Bosch Institute, University of Sydney

2011 Bosch ASM Best Poster Prize, Bosch Institute Annual Scientific Meeting, University of Sydney

2011 Prize for Outstanding Oral Presentation, Australia and New Zealand Zebrafish Meeting, Brisbane, Australia

2011 Faculty of Medicine Travel Fellowship for a collaborative project at the Ludwig Institute for Cancer Research

2011 NWG Macintosh Memorial Fund Department of Anatomy and Histology, University of Sydney

2011 Postgraduate Research Student Support Bosch Institute, University of Sydney

2010 Lastek Prize for Outstanding Poster Presentation, Bosch Young Investigators Symposium, Sydney, Australia

2010 Postgraduate Research Student Support Bosch Institute, University of Sydney

2010 Australian Postgraduate Award (APA) Australian Federal Government, competitive PhD Scholarship

xii List of abbreviations

AER apical ectodermal ridge aldh1a2 aldehyde dehydrogenase 1 family, member A2 BGI Beiging Genomics Institute bp base pairs cDNA complementary DNA cM centimorgans cRNA complementary RNA DIG digoxigenin dpf days post fertilisation E3 medium EL embryonic length eng1a engrailed 1a eng2 engrailed 2 eng3 engrailed 3 ENU N-ethyl-N-nitrosourea ESE exonic splicing enhancer EST expressed sequence tag st F1 1 generation nd F2 2 generation FGF fibroblast growth factor fgf10 fibroblast growth factor 10 fgf24 fibroblast growth factor 24 fgf8a fibroblast growth factor 8a fgfr2 fibroblast growth factor receptor 2 Gb gigabase pairs gDNA genomic DNA GFP green fluorescent protein H & E Haematoxylin and eosin hnRNP1 heterogeneous nuclear ribonucleoprotein 1 HOXB9 HOMEOBOX B9 hoxc10a homeobox c10a HOXC9 HOMEOBOX C9 hoxd9a homeobox d9a hpf hours post fertilisation Indel Small insertion or deletion kb kilobase pairs LB Luria-Bertani broth lef1 lymphoid enhancer-binding factor 1 macf1 microtubule-actin crosslinking factor 1 Mb megabase pairs morpholinos antisense morpholino oligonucleotides mpf months post fertilisation mRNA messenger

xiii NEB New England Biolabs npat nuclear protein, ataxia-telangiectasia locus pax2.1 paired box 2.1 PCR polymerase chain reaction pitx1 paired-like homeodomain transcription factor 1 pitx2 paired-like homeodomain transcription factor 2 prdm1 PR domain containing 1 ptbp1a polypyrimidine tract binding protein 1 rap1gap2 rap1 GTPase activating protein rnh1 ribonuclease/angiogenin inhibitor 1 sall1a sal-like 1a sall3 sal-like 3 sall4 sal-like 4 scarf1 scavenger receptor F, member 1 shh sonic hedgehog SNP single nucleotide polymorphism sp8 specificity protein 8 spg20b spastic paraplegia 20 SSLP Simple sequence length polymorphism SVM support vector machine TALENs transcription activator-like effector nucleases Tbx2 T-box transcription factor 2 Tbx3 T-box transcription factor 3 tbx4 t-box transcription factor 4 tbx5 t-box transcription factor 5 tcf7 transcription factor 7 TL total length tRNA total RNA Tü Tübingen wnt3a wingless-type MMTV integration site family, member 3a wnt7a wingless-type MMTV integration site family, member 7a wnt8c wingless-type MMTV integration site family, member 8c wpf weeks post fertilisation

xiv Nomenclature

In this study, the nomenclature for zebrafish genes and follows the Zebrafish Nomenclature Guidelines (http://zfin.org/zf_info/nomen.html). Gene names are lower case and in italics. Protein names are lower case with the first letter capitalised and in normal case. The nomenclature for human and mouse genes and proteins is based on the Trends in Genetics Genetic Nomenclature Guide (March, 1995). When referring to human and chicken genes and proteins, all letters are capitalised. Gene names are italicised and protein names are in normal case. When referring to mouse genes and proteins, the first letter is capitalised. Gene names are italicised and protein names are in normal case. For all other organisms, the zebrafish nomenclature is adopted.

Examples:

Gene and protein symbol conventions for “sonic hedgehog” Gene Protein Human SHH SHH Mouse Shh SHH Chicken SHH SHH Zebrafish shh Shh

xv List of figures

Figure 1.1. The increasing number of publications utilising zebrafish for developmental studies ...... 3

Figure 1.2. The fins of zebrafish ...... 7

Figure 4.1. Wild-type vs. pelvic finless total length ...... 50

Figure 4.2. Images of wild-type and pelvic finless embryos ...... 51

Figure 4.3. Images of wild-type and pelvic finless larvae ...... 53

Figure 4.4. Images of all fins in pelvic finless zebrafish and wild-type siblings ...... 55

Figure 4.5. Wild-type vs. pelvic finless fin ray number ...... 56

Figure 4.6. Development of the pelvic fins in wild-type zebrafish ...... 58

Figure 4.7. Comparison of pelvic fin development in pelvic finless zebrafish and wild-type siblings ...... 59

Figure 4.8. Scanning electron microscopy images of developing pelvic fins in wild-type and pelvic finless zebrafish ...... 61

Figure 4.9. Sections of wild-type and pelvic finless pelvic regions ...... 63

Figure 4.10. Incomplete penetrance of the pelvic finless phenotype ...... 65

Figure 5.1. The hierarchy and interactions of genes examined by in situ hybridisation...... 70

Figure 5.2. pitx1 expression ...... 72

Figure 5.3. tbx4 expression ...... 74

Figure 5.4. tbx5 expression ...... 76

Figure 5.5. fgf10a expression ...... 78

Figure 5.6. sp8 expression ...... 80

Figure 5.7. fgf8a expression ...... 82

Figure 5.8. shh expression ...... 84

Figure 6.1. Diagram of recombination during meiosis ...... 95

Figure 6.2. Modes of SSLP inheritance ...... 98

xvi Figure 6.3. Diagram and time line of mapping pedigrees ...... 102

Figure 6.4. Example of polymorphic markers ...... 104

Figure 6.5. Genome scan results for chromosome 15 ...... 105

Figure 6.6. Individual PCR for chromosome 15 SSLP marker z21982...... 107

Figure 6.7. Additional chromosome 15 markers ...... 109

Figure 6.8. The recombination gradient ...... 111

Figure 6.9. Individual PCR for chromosome 15 SSLP marker z11320...... 113

Figure 6.10. Individual PCR for chromosome 15 SSLP marker z13310...... 115

Figure 6.11. Individual PCR for chromosome 15 SSLP marker z3760...... 117

Figure 6.12. Genetic interval containing the pelvic finless locus ...... 119

Figure 7.1. The deceasing cost of sequencing ...... 123

Figure 7.2. Nonsense mutations are in rap1gap2 and scarf1 in zebrafish ...... 134

Figure 7.3. Frameshift Indels identified in npat and rnh1 are not present in zebrafish ...... 136

Figure 7.4. Candidate mutations in tbx4 are present in pelvic finless zebrafish ...... 138

Figure 7.5. tbx4 cDNA sequence ...... 140

Figure 7.6. tbx4 transcript length ...... 142

Figure 7.7. Tbx4 amino acid substitutions ...... 144

Figure 7.8. Tbx4 protein sequence analysis ...... 146

Figure 7.9. Change in total free energy graphs for the predicted A79V and G80A mutations...... 148

Figure 8.1. The Pel2.5kb-GFP construct...... 159

Figure 8.2. The pDestTol2pA2-Pel2.5kb-Gata2-Tbx4-pA rescue construct...... 162

Figure 8.3. Wild-type Pel2.5kb-GFP embryos ...... 165

Figure 8.4. The developing pelvic fins wild-type Pel2.5kb GFP zebrafish ...... 166

Figure 8.5. The developing pelvic regions of wild-type and pelvic finless Pel2.5kb-GFP zebrafish ...... 167

xvii Figure 8.6. Pelvic fin rescue at low doses ...... 169

Figure 8.7. High dose mosaic transgenic rescue results ...... 171

Figure 9.1. A diagram of the pathways leading to the onset of fgf10 expression ...... 183

xviii List of Tables

Table 7.1. Nonsense SNPs ...... 133

Table 7.2. Frameshift Indels ...... 135

Table C.1. Table of length measurements of pelvic finless zebrafish and wild-type siblings...... 232

Table C.2. Fin ray number ...... 233

Table D.1. Recombinants at chromosome 15 markers ...... 234

Table F.1. A table of the results from the low dose mosaic transgenic rescue experiments . 243

Table F.2. Tables of the results from the high dose mosaic transgenic rescue experiments . 243

xix 1. Introduction The specific aims of this body of work are to morphologically and genetically characterise pelvic fin development in zebrafish (Danio rerio) and to identify the genomic lesion underlying pelvic fin loss in a naturally occurring zebrafish strain which lacks pelvic fins, known from herein as “pelvic finless zebrafish”. This project was conceived in the spirit of the original theme of research in our laboratory, which aimed to utilise the advantages of the zebrafish to study limb and muscle development in to better understand developmental processes and mechanisms in all . This project was conducted on the premise that it would set the foundation for establishing the zebrafish pelvic fin as a developmental model and build upon this foundation by discovering a novel developmental gene or pathway through the genetic characterisation of a pelvic finless zebrafish.

1.1. Introduction to development Development is the orderly combination of processes that progressively add complexity to the relative simplicity of a fertilised to result in the creation of a complex organism. The complexity of the organism is achieved by the combination of many processes that increase cell number and type and control the number, function and position of organs and structures that determine the overall body plan and life of the organism. It is through the precise timing and coordination of these processes, at a molecular and cellular level, that the structures that comprise the new organism are built. By gaining a better understanding into the mechanisms that occur at the molecular and cellular level during development, a greater knowledge of essential developmental processes can be achieved.

In vivo and in vitro developmental models can be utilised in order to investigate the mechanisms that occur at the molecular and cellular level during development. To better understand the potential of research conducted in developmental models, it is important to first understand the basics of development. The development of many structures and organs begins by the induction of a primordium (a precursor structure) at a specific time and location in the embryo by a combination of pre-existing cues. The primordium is comprised of a group of embryonic cells that have originated from the endoderm, mesoderm or , or from a combination of these embryonic layers. The development of a structure or organ is generally driven by cell-to-cell communication between the cells in the mesenchymal and epidermal layers of the primordium, however there are exceptions to this. These cell-to-cell molecular interactions and cellular events are performed by an evolutionarily conserved ‘genetic toolkit’ which operates in the development of a wide array of organs and structures.

1 In fact, organs and structures as diverse as teeth, kidneys, limbs, wings, fins, hair follicles and all utilise similar genetic pathways and mechanisms during development. It is only the timing of the pathways and downstream genes that make the final organs and structures so different, in both appearance and function. This wonderful evolutionary conservation allows for knowledge gained from one specific developmental model to be applied to the understanding of many different developmental processes.

1.1.1. Introduction to zebrafish as developmental models Over the past few decades, zebrafish have moved into place amongst the other classical models of development, such as mouse and chick (Grunwald and Eisen, 2002; Lele and Krone, 1996; Veldman and Lin, 2008). Zebrafish have been established as a mainstream developmental model because they represent a quickly developing vertebrate which is easily amenable to invertebrate-like genetic approaches (such as forward genetic screens) and therefore present many advantages over other classic models of development.

Zebrafish are a vertebrate which belong to the class of ray-finned fish (). The of zebrafish develop externally allowing easy observation and manipulation of early developmental processes. The zebrafish embryo also develops rapidly; a beating is present in the first 24 hours post fertilisation (hpf) and all major organ systems are developed by 5 days post fertilisation (dpf) (Kimmel et al., 1995). As the zebrafish is a vertebrate, most of these organ systems are similar to those found in humans and other , with some exceptions (such as the uterus). In addition, zebrafish are optically transparent for the first six weeks of development which allows for the easy observation of organ systems and later developmental processes. In addition, once sexual maturity is reached at 3-4 months post fertilisation (mpf), female zebrafish are capable of producing a large number of offspring which allows for the production of several hundred samples each week.

Although zebrafish have been utilised for developmental studies for over eighty years, their popularity has dramatically increased over the past decade (Figure 1.1). Zebrafish have been utilised for embryonic and developmental studies as early as the 1930s and are now used in ‘forward’ and ‘reverse’ genetic studies of developmental biology (Laale, 1977; Veldman and Lin, 2008). Since the 1980s zebrafish have been used to perform large scale ‘forward’ genetic screens in order to identify novel regulators of developmental processes of interest as techniques such as cloning, mutagenesis, transgenesis and mapping approaches became available in zebrafish (Chakrabarti et al., 1983; Grunwald and Streisinger, 1992;

2 Solnica-Krezel et al., 1994; Streisinger et al., 1981; Streisinger et al., 1986; Stuart et al., 1988; Walker and Streisinger, 1983). During the 1990s, the zebrafish began to truly emerge as a powerful and mainstream developmental model due to two large scale mutagenesis screens which resulted in thousands of zebrafish mutants with specific developmental abnormalities (Driever et al., 1996; Haffter et al., 1996). In the twenty-first century, zebrafish are an established mainstream developmental model with a fully sequenced and annotated genome and provide an excellent platform from which a multitude of new developmental discoveries can be made.

Figure 1.1. The increasing number of publications utilising zebrafish for developmental studies A line graph demonstrating the number of publications utilising zebrafish for developmental studies based on a PubMed search using the key terms ‘zebrafish’ and ‘development’. The number of publications utilising zebrafish for development studies has steadily increased since the late 1990’s.

1.1.2. Advantages of utilising zebrafish as developmental models Although not as closely related to humans as the mouse and chick, the zebrafish offers numerous advantages as a developmental model. As a teleost, zebrafish have undergone several rounds of genome duplication, the latest of which occurred at the base of the teleost radiation, before the divergence of the zebrafish, pufferfish and medaka lineages, around 300 million years ago (Amores et al., 1998; Aparicio, 2000; Hedges and Kumar, 2002; Meyer and Mélaga-Trillo, 1999; Meyer and Schartl, 1999; Naruse et al., 2000; Santini and Tyler, 1999;

3 Taylor et al., 2001). After this latest round of genome duplication, it is believed that chromosome re-arrangements resulted in the inactivation of some duplicated genes and the subfunction partitioning of paralogous genes. Therefore, there are sometimes two or more copies of essentially the same gene in the zebrafish genome with each performing a part of the role of the original gene. This trait can be utilised to examine the role of complex genes during the development in zebrafish by avoiding the early lethality that some genetic manipulations cause in mice and chicks due to compound phenotypes (Postlethwait, 2006). For example, the brom bones mutant zebrafish has a nonsense mutation in the gene heterogeneous nuclear ribonucleoprotein (hnRNP1), also known as polypyrimidine tract binding protein 1a (ptbp1a) and displays an intestinal phenotype (Yang et al., 2009). However in other systems, this hnRNP1 is critical for the formation of embryonic polarity which is crucial for development and any mutations in the gene are lethal at an early stage (Lewis et al., 2008). In zebrafish, this gene has undergone duplication and subfunctionalisation, and therefore the brom bones mutant zebrafish is viable. This is because ptbp1b controls the embryonic function of this pair of paralogous genes and allows for successful (Mei et al., 2009; Yang et al., 2009). This allowed for the novel discovery of the role of ptbp1a in Notch signalling during the development of the zebrafish intestine (Yang et al., 2009).

Another advantage of using zebrafish as developmental models is the availability of so many specific developmental mutant phenotypes, produced from forward genetic screens. These allow for an unbiased approach to identifying the function of a gene based on a phenotype of interest. Two large scale (and many small scale) mutagenesis screens have been performed in zebrafish (Driever et al., 1996; Haffter et al., 1996). At the time of writing, these screens have produced thousands of mutant zebrafish with developmental defects (Bradford et al., 2011). Although many of the genes responsible are yet to be identified, several important discoveries have come from these mutants. For example, studies of the zebrafish t-box transcription factor 5 (tbx5) mutant, known as heartstrings (hstm21), confirmed an early and necessary role for tbx5 in pectoral fin development (Garrity et al., 2002), which was later confirmed in mice and chicks (Agarwal et al., 2003; Rallis et al., 2003; Takeuchi et al., 2003). In addition to identifying genes that are essential for development, these numerous zebrafish mutants can be used to address the genetic hierarchy of development processes. For example, zebrafish fibroblast growth factor 24 (fgf24), fibroblast growth factor10 ( fgf10), tbx5, aldehyde dehydrogenase 1 family, member A2

4 (aldh1a2) mutants were all used to place the newly discovered gene PR domain containing 1 (prdm1) just upstream of fgf10 signalling in the pectoral fin development genetic cascade (Mercader et al., 2006; Wilm and Solnica-Krezel, 2005). These are just two examples of developmental discoveries that were made utilising the advantages of the zebrafish model, however, with the large number of developmental mutants available, there is great potential for many more discoveries.

Zebrafish can also be used for ‘reverse’ genetic approaches and many of these exciting new technologies are now emerging in the mainstream. For almost twenty-five years, transgenic zebrafish have been utilised to examine gene function (Stuart et al., 1988). Investigating gene function in vivo and in real-time in zebrafish during development has lead to many exciting new discoveries which would have not been feasible in other model organisms. For example a study utilising zebrafish paired box 2.1 (pax2.1) promoter transgenic lines revealed a novel positive transcriptional feedback loop in - hindbrain boundary development involving pax2.1, engrailed 2 (eng2) and engrailed 3 (eng3) which allows for the continuation of pax2.1 expression to become independent of the patterning machinery of the gastrula embryo (Picker et al., 2002).

In the past, one drawback of using zebrafish as a developmental model was the lack of gene deletion and modification technologies in zebrafish; however, this is now changing. Until recently, antisense morpholino oligonucleotides (morpholinos; Genetools) have been used to transiently knockdown gene expression in zebrafish (Nasevicius and Ekker, 2000). Although the application of morpholinos in zebrafish is easier than in other organisms, as morpholinos can be injected by hand under a dissecting microscope, they are only functional for a short amount of time and unlike a true mouse knockout, morpholinos only offer a transient knockdown that lasts approximately around 3-5 dpf (Nasevicius and Ekker, 2000). However, morpholinos have been used widely and successfully in studies of structures which develop within this timeframe. For example, morpholinos have been utilised in studies of the development of the pectoral fins, as the pectoral fins of zebrafish develop within the first 48 hours after fertilisation (Ahn et al., 2002; Garrity et al., 2002; Grandel and Schulte-Merker, 1998; Harvey and Logan, 2006). Morpholinos have been used not only to phenocopy zebrafish pectoral fin mutants and thus confirm the responsible gene (ex. tbx5/heartstrings mutant; Ahn et al., 2002; Garrity et al., 2002), but also to determine the location and function of genes in the fin development cascade (ex. sal-like 4 (sall4); Harvey and Logan, 2006).

5 The drawbacks of morpholinos occur when examining later or long-term developmental processes. However, new breakthrough technologies are offering a solution to this problem. Transcription activator-like effector nucleases (TALENs) offer the ability to perform targeted gene knockout and modification in zebrafish. This new technology works by targeting a gene of interest and creating a double standard break. When this break is repaired, a mutation can be induced which can create a null version of the gene of interest (Sander et al., 2011). In addition, it is now possible to fill the excision with a new sequence of interest, in order to create specific mutations in the gene of interest (Zu et al., 2013). TALENs are well adapted to use in the zebrafish as they can be simply injected by hand into the one-cell stage embryo. The embryos are then raised to adulthood and screened to identify founder fish in which targeted gene modification is present in the germ-line cells. Once founder fish have been identified, the fish can be crossed to create a stable knockout line (if the knockout is non- lethal) or kept as heterozygotes and incrossed to produce homozygous knockout offspring. With these new technologies, what was once a drawback for zebrafish developmental models has been overcome and in fact, is now another advantage of the zebrafish developmental model. 1.1.3. Zebrafish fins as a model for development Many developmental studies have focused on the developing limbs of tetrapods because the limbs of vertebrates offer a good model in which to investigate and understand the genetic processes that govern organ development. This is because the shared ‘genetic toolkit’ is utilised during limb development and therefore any knowledge gained from these models can be applied to other developmental systems. In addition to the conservation of developmental pathways, limbs develop externally to the embryo and offer relative ease of access and are not essential for survival. The developing fins of zebrafish offer all of these same advantages in combination with the numerous advantages of the zebrafish model system, and therefore present a fantastic model in which to investigate developmental mechanisms at biological, disease and evolutionary levels. This is possible due to the evolutionary conservation between limbs and fins. The and hindlimbs of tetrapods are evolutionarily derived from the paired fins of ancestral fish (Carroll, 1988; Coates, 1994) and the shared origins of limbs and the fins of modern fish can be observed in the similar gene and protein expression patterns present during limb and fin development (Don et al., 2012; Mercader, 2007). As zebrafish share high genomic and molecular similarities with other vertebrates, important developmental discoveries made in zebrafish can be applied to all vertebrates. 6 Adult zebrafish have two sets of paired fins and several other singular fins (Figure 1.2). The paired fins of zebrafish are the pectoral and the pelvic fins which are derived from the lateral plate mesoderm (Grandel and Schulte-Merker, 1998). The pectoral fins of zebrafish are homologous to the forelimb (arms) of tetrapods and the pelvic fins of zebrafish are homologous to the hindlimb (legs) of tetrapods. The other fins are the , the anal fin and the caudal fin which are derived from the somatic mesoderm and partially from the neural crest (Grandel and Schulte-Merker, 1998).

Figure 1.2. The fins of zebrafish A lateral view of the fins of a wild-type zebrafish. Zebrafish have two pairs of paired fins (the pectoral and pelvic fins) and several singular fins (the dorsal, ventral and caudal fins). c: caudal or tail fin; d: dorsal fin; pec: pectoral fin; pel: pelvic fin; v: ventral or anal fin.

Despite the evolutionary conservation of gene and protein expression during development between pelvic fins of zebrafish and the hindlimbs of tetrapods (for a full description of the evolution and development of pelvic fins please see Chapter 2), the potential of pelvic fins of zebrafish as a developmental model has been relatively ignored. We aim to change this by describing pelvic fin development in zebrafish and characterising a pelvic fin mutant, the pelvic finless zebrafish. It is hoped that this will shed light on the previously uncharacterised mechanisms of pelvic fin development and therefore set the foundation for further developmental studies in zebrafish pelvic fins.

7 2. Literature Review

The evolutionary history of the development of the pelvic fin/hindlimb

Emily K. Don, Peter J. Currie, Nicholas J. Cole

Journal of Anatomy (2012)

This paper was written and published during the course of this degree. This paper provides a review of the literature surrounding the evolution and development of the pelvic fins and hindlimbs. The author contributions were as follows;

Emily Don provided intellectual input, researched and wrote the manuscript.

Nicholas Cole contributed to the original idea, offered intellectual input and assisted with the production of the manuscript.

Peter Currie contributed to the work in the form of intellectual discussions, guidance and proof reading of the manuscript.

Signatures from co-authors:

Nicholas Cole Peter Currie

8 Journal of Anatomy

J. Anat. (2012) doi: 10.1111/j.1469-7580.2012.01557.x

REVIEW ARTICLE The evolutionary history of the development of the pelvic fin/hindlimb Emily K. Don,1 Peter D. Currie2 and Nicholas J. Cole1

1Department of Anatomy & Histology, School of Medical Sciences and Bosch Institute, University of Sydney, Sydney, NSW, Australia 2Australian Regenerative Medicine Institute, Monash University, Clayton, Vic., Australia

Summary

The arms and legs of man are evolutionarily derived from the paired fins of primitive jawed fish. Few evolutionary changes have attracted as much attention as the origin of tetrapod limbs from the paired fins of ancestral fish. The hindlimbs of tetrapods are derived from the pelvic fins of ancestral fish. These evolutionary origins can be seen in the examination of shared gene and protein expression patterns during the development of pelvic fins and tetrapod hindlimbs. The pelvic fins of fish express key limb positioning, limb bud induction and limb outgrowth genes in a similar manner to that seen in hindlimb development of higher vertebrates. We are now at a point where many of the key players in the development of pelvic fins and vertebrate hindlimbs have been identified and we can now readily examine and compare mechanisms between species. This is yielding fascinating insights into how the developmental programme has altered during evolution and how that relates to anatomical change. The role of pelvic fins has also drastically changed over evolutionary history, from playing a minor role during swimming to developing into robust weight-bearing limbs. In addition, the pelvic fins/hindlimbs have been lost repeatedly in diverse species over evolutionary time. Here we review the evolution of pelvic fins and hindlimbs within the context of the changes in anatomical structure and the molecular mechanisms involved. Key words: development; evolution; hindlimb; pelvic fin.

Introduction to pelvic fins and hindlimbs vic fins and hindlimbs and we are now able readily to com- pare between species. We will be presenting evidence for The transition of vertebrates from to land was one of the evolution of pelvic fins and hindlimbs organised by inte- the greatest steps in evolutionary history. This required the gration of anatomy and molecular mechanisms. development of paired pelvic fins and their muscles to eventually form weight-bearing hindlimbs (Goodrich, Pelvic fin and hindlimb morphology 1930). A number of forms have shed light on the evo- lution of pelvic fins and hindlimbs within ancestral fish and During the evolutionary history of vertebrates, paired pelvic early tetrapods (Andrews & Westoll, 1970; Coates, 1996; appendages have undergone many changes since their first Jarvik, 1996; Clement et al. 2004; Ahlberg et al. 2005, 2008; appearance. In fish, pelvic fins vary greatly in morphology, Boisvert, 2005; Callier et al. 2009; Niedzwiedzki et al. 2010; function and position, whereas in tetrapods they have Zhu et al. 2012). Studies of these have shown that evolved into robust weight-bearing hindlimbs necessary for the transition from paired pelvic fins to tetrapod hindlimbs . is characterised by a gradual progression from posterior Although the morphology of the fish fin differs greatly and ventrally placed slender pelvic fins articulated to a pel- from that of the tetrapod limb, a clear connection in the vic girdle to a dorsally located robust and hindlimb. evolution of limbs from fins can be seen in the anatomy of Recent genetic approaches haveledustoapointwherethe the paired fins of chondrichthyans, actinopterygians, sar- key developmental mechanisms have been identified in pel- copterygians and the limbs of tetrapods (Janvier, 1996). The pelvic fin of chondrichthyans (sharks and rays) is composed

Correspondence of a propterygium, a mesopterygium and a metapterygium. Nicholas J. Cole, Department of Anatomy & Histology, School of Whereas actinopterygians (e.g. paddlefish) have Medical Sciences and Bosch Institute, University of Sydney, New maintained all three elements of the pelvic fin, (e.g. South Wales 2006, Australia. E: [email protected] zebrafish) have maintained the propterygium and mesop- Accepted for publication 27 July 2012 terygium, and have lost the metapterygium (Coates, 1994;

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 9 2 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

Coates & Cohn, 1998) and sarcopterygians (lobe-finned fish) present but can be found on the dorsal side of the girdle have maintained the metapterygium. The tetrapod limb is (Winterbottom, 1974; Stiassny & Moore, 1992) (Fig. 2). The thought to originate from the metapterygium of sarcop- positioning of the adult pelvic fin has shifted during teleost terygians (reviewed in Wagner & Chiu, 2001) (Fig. 1). fish evolution, ranging from an abdominal position in the The pelvic fins of chondrichthyans and basal ray-finned ventral body wall near the cloacae to an anterior position fish are usually located at an approximately mid-body at either a thoracic or a jugular level in more derived teleost position which is posterior to the centre of mass. The pelvic groups, although there are many exceptions to this condi- girdle of sharks is embedded in the body wall and is articu- tion (Greenwood et al. 1966; Rosen, 1982; Nelson, 1994). lated to a number of elongated cartilaginous elements, Amongst teleosts, there are many exceptional pelvic fin which support the pelvic fin rays (Liem & Summers, 1999). structures such as dorsally placed pelvic fins (Bathophilus, The distinctive pelvic fins of teleosts consist of three main Fink, 1985), unpaired structures (such as the fused pelvic skeletal elements; the basipterygium (pelvic girdle), a spine of triggerfish and the dewlap of filefishes; Matsuura, reduced number of radials (fin base) and the lepidotrichs 1979; Tyler, 1980) and even the complete loss of pelvic fins (slender bony fin rays) (Stiassny & Moore, 1992; Cubbage & (some sticklebacks and some zebrafish, Cole et al. 2003; Mabee, 1996; Coates & Cohn, 1998). The pelvic fins are sup- Shapiro et al. 2004; Chan et al. 2010; Don et al. 2011). It is ported by a cartilaginous endoskeleton which ossifies dur- possible that as the pelvic fins are not absolutely necessary ing development. The endoskeleton consists of a pelvic for survival, this diverse range of morphologies is possible. girdle which articulates with the endoskeleton of the pelvic The pelvic fins of lobe-finned fish are the evolutionary fin, the pattern of which differs greatly between species forerunners of the hindlimbs of tetrapods. The pelvic fins of (Goodrich, 1930; Zangerl, 1981; Shubin, 1995). The pelvic lobe-finned fish are derived from the metapterygium of fins of fish consist of a proximal muscular component and a chondrichthyans and basal actinopterygian fish (Mabee, distal non-muscularised dermal fin fold. The fin folds are 2000; Raff, 2007). The pelvic girdle of lobe-finned fish artic- supported by long fin rays which are ossified (Grandel & ulates with the ; at its distal tip, the femur articulates Schulte-Merker, 1998). Teleosts usually have six pelvic fin with the tibia and fibula, which may be fused in basal sar- muscles, three pairs lying on each side of the pelvis (Winter- copterygians (Vorobyeva & Hinchliffe, 1995). The mesopodi- bottom, 1974). The arrector ventralis pelvicus, the abductor um of lobe-finned fish is composed of an incomplete and superficialis pelvicus and the abductor profundus pelvicus variable arrangement of bones, which makes up the ankle are found on the ventral side of the pelvic girdle. The arrec- in tetrapods (Wagner & Chiu, 2001). The more distal acrop- tor dorsalis pelvicus, adductor superficialis pelvicus and the odium is absent in the pectoral and pelvic fins of living and adductor profundus pelvicus are located on the medial side extinct sarcopterygian fish (Johanson et al. 2007). The mus- of the pelvic girdle. Infracarinalis anterior is connected to cles of the pelvic fin of an extant sarcopterygian fish, the the basipterygium and the cleithrum, while the infracarinal- Australian lungfish (Neoceratodus forsteri) have been is medius is connected to the basipterygium and the first described previously (Young et al. 1989 and Boisvert et al. anal-fin pterygiophores. The extensor proprius is not always 2009). The adductor muscles of the Australian lungfish

A

B Fig. 1 Schematic representation the appendicular endochondral skeleton of a basal fish and a tetrapod. (A) A Polyodon (paddlefish) pectoral fin (adapted from Grande & Bemis 1991). (B) A human arm. Each appendage is orientated with anterior (preaxial) upward and distal to the right. Figures are not to scale. Reproduced with permission from Wagner & Larsson (2007) and Grande & Bemis (1991).

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 10 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 3

ABC

Fig. 2 Schematic diagram and bone and cartilage staining of the musculature of the pelvic girdle of teleost. Ventral (A, C) and dorsal (B) views of the pelvic fin muscles (anterior to the top). ABSP, abductor superficialis pelvicus (red); ABPP, abductor dorsalis pelvicus (orange); ADPP, adductor profundus pelvicus (light blue); ADSP, adductor superficialis pelvicus (blue); ARRDP, arrector dorsalis pelvicus (green); ARRVP, arrector ventralis pel- vicus (yellow); EXTP, extensor proprius (purple); ICARA, infracarinalis anterior (light grey); ICARM, infracarinalis medius (dark grey). A tendonis shown in black. Reproduced with permission from Winterbottom (1974); and Yamanoue et al. (2010); and modified with permissions from Don et al. (2011). pelvic fin are the superficial ventromesial adductor, the The muscles of the ( maculosus) hind- superficial ventrolateral adductor, the deep ventral adduc- limb have been previously described and compared with tor depressor, the dorsomesial adductor levator and the the pelvic fin muscles of the Australian lungfish (Young mesial adductor (Young et al. 1989; Boisvert et al. 2009). et al. 1989; Walker & Homberger, 1992; Boisvert et al. The pelvic fin abductor muscles of the Australian lungfish 2009). Most of the muscles of the salamander hindlimb and are the superficial ventromesial abductor, the superficial pelvic girdle can be matched in both insertion points and ventrolateral abductor, the deep ventral abductor depressor function to their equivalents in the Australia lungfish. The and the dorsolateral abductor levator (Young et al. 1989; developmental mechanisms of how the pelvic fin muscles Boisvert et al. 2009). The radial flexors of the lungfish pelvic of ancient fish evolved to become more robust musculature fin functions as both an adductor and abductor (Young of the tetrapod hindlimb will be discussed below. et al. 1989; Boisvert et al. 2009). The supinator and prona- tor pelvic fin muscles of the Australian lungfish are possibly Pelvic fin and hindlimb function the precursors to tetrapod muscles as the radial–axial muscles of the Australian lungfish attach to the distal radi- In addition to the varied morphology of pelvic fins and tet- als and it is thought that the distal radials of ancient fish rapod hindlimbs, the paired pelvic appendages have a vari- were the precursors of digits (Young et al. 1989; Johanson ety of functions. The slender pelvic fins of fish mostly play et al. 2007; Boisvert et al. 2008, 2009). The hypaxial muscles minor roles during swimming and manoeuvring, and the of the Australian lungfish do not attach to the pelvis hindlimbs of tetrapods are usually employed as the major (Young et al. 1989; Boisvert et al. 2009). form of locomotion. Fossil evidence suggests that tetrapod hindlimb origi- Dominate propulsion by the body and caudal fin during nated in the 370 million years ago and that it swimming is a common feature throughout the evolution has its origins in the metapterygium of sarcopterygians of fish; however, the pelvic fins do play a role during steady (Vorobyeva, 1991; Ahlberg & Milner, 1994; Wagner & Chiu, swimming and manoeuvres, especially in teleosts (Gosline, 2001). Most tetrapod hindlimbs follow a general plan of a 1980; Webb, 1982; Standen, 2008). Early work on the func- proximal stylopodium, a zeugopodium and a distal autopo- tion of pelvic fins in dogfish concluded that the pelvic fins dium. The stylopodium of modern tetrapods consists of a had a very limited and mostly passive, stabilising function single elongated skeletal element called the femur. The during locomotion and were mainly concerned with the modern tetrapod zeugopodium consists of two parallel production of vertical forces (Harris, 1937, 1938). It was skeletal elements, the fibula and the tibia. The autopodium shown that, in sharks, the pelvic fins increase the static sta- of modern tetrapods consists of the more proximal mesopo- bility for pitching movements, but only to a small extent dium and the distal acropodium. A full complement of tar- (Harris, 1938). sal bones makes up the mesopodium or the ankle, whereas In more derived ray-fin fishes, the pelvic fins have moved the more distal acropodium usually consists of five radiating to beneath the centre of mass and have a greater degree of digits (Fig. 1B). The pelvis of modern tetrapods is made up mobility when compared with the pelvic fins of sharks of three elements, the ilium, ischium and pubis, and is fused (Harris, 1938; Gosline, 1980; Rosen, 1982; Schrank et al. to the though a sacral (Clack, 2000). 1999). Early studies suggested that this adaptation leads to

© 2012 The Authors © Journal of Anatomy 2012 Anatomical Society 11 4 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

the pelvic fins of bony fish having little effect on body yaw functions reflect the evolutionary changes that have when used simultaneously but being capable of inducing occurred in pelvic fins and hindlimbs since their first appear- rolling movements (Harris, 1938; Gosline, 1980; Rosen, 1982; ance. Schrank et al. 1999). More recent studies have suggested that the pelvic fins of teleost play an even greater role dur- Evolution of pelvic fins and hindlimbs ing swimming (Drucker & Lauder, 2003; Lauder & Drucker, 2004; Standen, 2008). It has been shown that teleosts Origin of paired limbs actively use their pelvic fins as control surfaces during turn- ing manoeuvres and in combination with other fins (anal For over a century the origin of vertebrate paired fins and and dorsal) compensate for pitching movements during limbs has been fiercely debated. One of the first theories breaking (Drucker & Lauder, 2003; Lauder & Drucker, 2004). put forward proposed that fins evolved from the arches A recent three-dimensional kinematics study of the rainbow of the early limbless vertebrates (Gegenbaur, 1878). Recent trout (Oncorhynchus mykiss) has shown that during steady theory proposes that the fins of vertebrates evolved from swimming, pelvic fins oscillate in regular contralateral cycles continuous stripes of competency for appendage formation and during manoeuvres act as trimming foils. The cyclic located ventrally and laterally along the embryonic flank oscillation, involving active and passive components, may (Yonei-Tamura et al. 2008) (Fig. 3A). A continuation of this function to dampen body oscillation and stabilise body theory proposed that the paired appendages of jawed ver- position (Standen, 2008). When acting as trimming foils tebrates evolved with a shift in the zone of competency to during manoeuvres, the pelvic fins move variably, which the lateral plate mesoderm (LPM) in conjunction with the helps to stabilise and return the body to a steady swimming establishment of the lateral somitic frontier, which allowed posture (Standen, 2008). Despite the new-found roles for for the formation of limb/fin buds with internal supporting the pelvic fins of teleosts during swimming and manoeu- skeletons (Freitas et al. 2006; Durland et al. 2008; reviewed vring, the pelvic fins are still considered to be the least Johanson, 2010) (Fig. 3A). The conservation of genetic important fin for swimming because they have been lost mechanisms (Hox and Tbx expression patterns) between frequently during evolution and their amputation does not median fins and paired fins of shark and embryos greatly change body motion during swimming (Harris, supports this theory (Freitas et al. 2006). 1938; Gosline, 1980; Standen, 2008). To explain the emergence of ‘two sets’ of paired fins, sev- There are also cases where the pelvic fins of teleosts are eral theories have been put forward. The Thacher–Mivart– used for different kinds of locomotion. Some fish groups Balfour fin fold theory of the origin of the paired fins sug- use their pelvic fins to move, as if over aquatic and gests that tetrapod forelimbs and hindlimbs evolved by the terrestrial substrata (Peters, 1985; Webb, 1996), whereas splitting of a single lateral fin (Thacher, 1877; Mivart, 1879; others use hypertrophied pelvic fins, in combination with Balfour, 1881) (Fig. 3B). This theory has been contested due pectoral fins to fly (Davenport, 1992, 1994, 2003). to the inconsistency with the fossil record and a lack of Although tetrapods display a large diversity of function embryonic evidence. In contrast, Tabin & Laufer (1993) sug- in the hindlimb, most often, the robust weight-bearing gested that pelvic fins were acquired before pectoral fins in hindlimbs of tetrapods are used for hindlimb-propelled the ‘pelvic before pectoral’ fin model due to the collinear locomotion. Most modern tetrapods predominantly employ expression pattern of the Hox genes along the embryonic hindlimb-powered symmetrical gaits such as lateral flank and in developing limb buds (Fig. 3C). It was thought sequence walking where the hindlimb footfall is followed that the pattern of Hox gene expression along the flank of by the ipsilateral forelimb (Rewcastle, 1981). The hindlimb- the embryo was co-opted into the pelvic fins and then propelled walk of most modern tetrapods is facilitated by passed to the pectoral fins, which is why only the posterior several features unique to tetrapods, such as a weight- Hox genes are expressed during limb/fin development bearing pelvis that is attached to the vertebral column, a (Tabin & Laufer, 1993). However, to date, no fossils have laterally orientated leg, an elongated femur for longer been described which possess only pelvic fins (Coates, 1993; stride length and a flexible ankle. It is important to note Thorogood & Ferretti, 1993) and these authors suggested that not all tetrapods use the hindlimb only for locomotion, that based on fossil and developmental evidence that pec- examples of other functions of hindlimbs in tetrapods are toral fins were acquired before pelvic fins (Fig. 3D). Ruvin- the prehensile foot of monkeys and apes, and the loss of sky & Gibson-Brown (2000) proposed that an ancestral hindlimbs in cetaceans (whales and dolphins) for stream- Tbx4/5 cluster was initially co-expressed in the first pair of lined swimming (Fleagle, 1999; Bejder & Hall, 2002). It is fins to evolve. In modern jawed vertebrates, Tbx4 is possible that the variety of functions and morphologies of expressed in the pelvic appendages and Tbx5 is expressed in the tetrapod hindlimb is more constricted than in the pelvic the pectoral appendages (Gibson-Brown et al. 1996; fins of fish, as the robust weight-bearing hindlimb of most Tamura et al. 1999; Ruvinsky et al. 2000). To explain this tetrapod provides locomotion which is necessary for sur- expression pattern, it was suggested that the ancestral vival. However, the vast array of various morphologies and Tbx4/5 cluster underwent a duplication event, either before

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 12 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 5

A Parayunnanolepis xitunensis, from the early Devonian (approximately 430 Mya), which provided evidence for the presence of pelvic girdles in these phylogenetically basal placoderms (Zhu et al. 2012). Antiarch placoderms (extinct armoured jawed fishes) first appeared in the and as antiarchs are placed at the base of the gnathostome radia- tion, the far-reaching implications of this study are that all jawed vertebrates (including antiarch placoderms) primi- tively possess both pectoral and pelvic fins and that the pel- vic fins did not arise within gnathostomes at a point subsequent to the origin of (Coates, 1994; Janvier, 1996; Goujet, 2001; Zhu et al. 2012). In contrast, these results imply that paired pelvic appendages (pelvic fins/ hindlimbs) appeared within gnathostomes before the development of moveable jaws (Zhu et al. 2012).

B C D Pelvic fins and the tetrapod transition

The fish-to-tetrapod transition involved a gradual shift towards more coastal and terrestrial environments (Clack, 2000) and with it came a change in pelvic fin function. This transition involved the shift from paired pectoral and pelvic fins to the development of weight-bearing fore and hind- limbs for locomotion on land. It is thought that the fin to limb transition first began in the pectoral fins and that the evolution of pelvic fins into hindlimbs occurred in a rela- Fig. 3 Diagram of the evolution of paired fins. (A) The evolution of tively brief period of time between and paired fins from the ‘zone of competence’ (Freitas et al. 2006; Yonei- Acanthostega (Coates, 1996, 2002; Boisvert, 2005). Unfortu- Tamura et al. 2008; Johanson, 2010). A dorsal zone of competence for unpaired fins is the first to arise in jawless vertebrates (Johanson, nately, there is a real paucity of fossils in this interval with 2010). The dorsal zone of competence was then duplicated and intact pelvic fins. However, the insights gained from recent co-opted to a ventrolateral position along the flank (Freitas et al. fossil finds, re-examination of older fossils and evidence 2006). The ventrolateral zone of competence was then shifted to the obtained from developmental biology challenge the old LPM, which coincided with the evolution of the abaxial region and the ideas and suggest that the pelvic fin to hindlimb transition lateral somitic frontier (Johanson, 2010). (B) The ‘lateral fin fold’ theory was evolving even before early tetrapods moved out of the suggests that two paired fins evolved from a single continuous lateral water and colonised land. Two key developmental break- fin (Thacher, 1877; Mivart, 1879; Balfour, 1881). (C) Based on the collinear expression pattern of Hox gene expression, the ‘pelvic before throughs during this time were the elaboration of the distal pectoral’ theory suggests that the pelvic fins evolved before the pec- skeleton and the development of a robust weight-bearing toral fins (Tabin & Laufer, 1993). (D) Based on fossil evidence and the pelvis (Boisvert, 2005; Johanson et al. 2007). anterior–posterior pattern of development, the ‘pectoral before pelvic’ Evidence from the fossil record and developmental stud- theory suggests that pectoral fins evolved first and were then dupli- ies of living sarcopterygians suggest that during the evolu- cated to form pelvic fins (Coates, 1993; Thorogood & Ferretti, 1993). tion of the distal pelvic fin skeleton the digits appeared before the full complement of ankle elements (Wagner & or Tbx4 became localised at the pelvic level or after the clus- Chiu, 2001; Coates, 2003; Clack & Ahlberg, 2004; Johanson ter became localised at the pelvic level (Ruvinsky & Gibson- et al. 2007). It is currently thought that digits were not an Brown, 2000). In this model, Tbx4 acted in conjunction with evolutionary novelty of tetrapods, as previously believed, Pitx1 to modify the morphology of the developing limb to but evolved from the pre-existing distal radials of sarcop- a pelvic fin/hindlimb identity (Ruvinsky & Gibson-Brown, terygians (Johanson et al. 2007; Boisvert et al. 2008). During 2000). this stage of evolution, was plesiomorphic amongst Tetrapodomorpha. Fossil evidence from early Devonian tetrapods indicates that had Origin of pelvic fins seven , Acanthostega had eight toes, and Tulerpeton Although the of the origin of paired fins remains had at least six toes (Coates & Clack, 1990; Lebedev & controversial, it is now clear from the fossil record that pel- Coates, 1995; Coates, 1996). It is thought that pentadactyly vic fins arose within jawed fish (Fig. 4). Zhu et al. (2012) of later tetrapods did not evolve until the recently re-examined a fossil of a primitive antiarch, period (Coates, 1994, 1996).

© 2012 The Authors © Journal of Anatomy 2012 Anatomical Society 13 6 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

Fig. 4 Simplified phylogeny of the living relatives of tetrapods and the evolution of paired appendages based on the results of Inoue et al. (2003).

The evolution of the full complement of the central Most Carboniferous tetrapods have three to four central bones of the ankle (the mesopodium) came after the evolu- elements in the mesopodium, which allows for the ankle tion of digits (Wagner & Chiu, 2001; Coates, 2003; Clack & flexibility necessary for walking on land (Coates, 1996). Ahlberg, 2004; Johanson et al. 2007). The pelvic fins of The development of a robust weight-bearing pelvis was a ancestral sarcopterygians possessed the long bones equiva- key step in the evolution of the hindlimb during the tetra- lent to a femur, tibia and fibula, and distal radials from pod transition onto land. To walk on land, the relatively which digits would evolve, but did not possess the full com- gracile unattached pelvic girdle of fish gradually trans- plement of bones of the mesopodium (Andrews & Westoll, formed into a large tripartite weight-bearing structure con- 1970; Wagner & Chiu, 2001; Coates et al. 2002; Johanson nected to the vertebral column (Fig. 5). The pelvic girdle of et al. 2007; Boisvert et al. 2008). Recent re-examination of lobe-finned fish is composed of a crescentric pubis often Panderichthys has revealed that the pelvic fin of this tetra- connected through cartilage at the midline, but lacks an podomorph fish has a proximal mesopodium element, the ilium and is not connected to the vertebral column (Fig. 5A) fibulare, but lacks the central bones of the mesopodium (Ahlberg, 1989). In contrast, the pelvis of tetrapods has an (Boisvert, 2005). Two of the earliest tetrapods with well pre- ilium that is fused to the vertebral column and an ischium served hindlimbs, Ichthyostega and Acanthostega,had that is posterior to the pubis. In addition, the ischium and hindlimbs that had more derived characteristics, but still the pubis from both halves of the pelvis are fused along had very few central bones of the mesopodium (Jarvik, their midline, which creates a weight-bearing pelvis 1980, 1996; Coates, 1996; Johanson et al. 2007). The full (Fig. 5B) (Clack, 2000). There is much evidence from both complement of the central bones of the ankle seems to sides of this transition, but little information about how this appear in Tulerpeton, which has 12 preserved tarsal bones, evolution occurred due to the paucity of fossils from this including three central elements (Lebedev & Coates, 1995). period with intact pelvic girdles. On one side of the

AB

Fig. 5 (A) In sarcopterygian fish the pelvic girdle is supported by the hypaxial musculature and consists of a pubis (pb) with a caudally oriented acetabulum (ac) (articulation to the fin) (redrawn from Andrews & Westoll, 1970). (B) In early tetrapods the pelvic girdle consists of a pubis, an ischium (ish), and an ilium (il), which connects to the vertebral column through the sacral rib (sr). The acetabulum is placed laterally (redrawn from Coates, 1996). Figure modified from Cole et al. (2011).

© 2012 The Authors 14 Journal of Anatomy © 2012 Anatomical Society Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 7 transition, the pelvic girdle of the tetrapodomorph Pande- understanding of the evolution of fin/limb motor circuitry richthys is small, flat, club-shaped and distinctly fish-like necessary for hindlimb dominated locomotion in verte- (Boisvert, 2005). Unfortunately, the pelvic girdle and fin of brates (Murakami & Tanaka, 2011). the more crownward tetrapodomorph has not In addition to insights gained from recent fossil finds and been preserved, but the early tetrapods, Ichthyostega and the re-examination of older fossils, discoveries of preserved Acanthostega, had already evolved a distinctively tetrapod- pelvic fins and girdles of more crownward transitional like pelvis with an ilium and ischium (Jarvik, 1980, 1996; Devonian tetrapodomorph fish are eagerly awaited to shed Coates, 1996). light on the evolution of the vertebrate hindlimb from the With the evolution of the distal pelvic appendage skele- pelvic fins of ancestral fish. ton and the pelvis, came a shift in locomotory from ‘front wheel drive’ to ‘rear wheel drive’ during the tet- Development rapod transition (Boisvert, 2005). Non-sarcopterygian fish predominately use body muscle undulations and pectoral Recent examination of the mechanisms involved in the initi- fins for locomotion, whereas tetrapods use their hindlimbs ation, outgrowth and patterning of hindlimbs among dif- for this function (Coates et al. 2002). Recent evidence from ferent classes of extant vertebrates have given insights into African lungfish (Protopterus annectens)hasshownthat the evolution of vertebrate hindlimbs. Approaches using this sarcopterygian fish can use a range of pelvic fin-driven model organisms, such as mouse, chick and teleosts, in addi- gaits such as walking and bounding and use their pelvic fins tion to approaches using extant non-model organisms posi- to lift their body clear of the substrate in an aquatic envi- tioned at strategically important points in the vertebrate ronment (King et al. 2011). Descriptions of the paired pec- phylogeny have shed light upon the important players in toral and pelvic fins of fossils such as Panderichthys and hindlimb development. These studies have shown the high Ichthyostega also offer insights into the evolution of tetra- degree of conservation in the genetic mechanisms of fin pod locomotion. Panderichthys probably employed an and limb formation between fish and tetrapods but some intermediate ‘front-wheel drive’ mode of locomotion, using species-specific differences are present. its pelvic fins as minor anchors while body-flexion propul- Hindlimb development has been mainly investigated in sion pushed the fish forward (Boisvert, 2005). A recent study chick and mouse, whereas there have been relatively few of limb joint mobility of Ichthyostega has shown that this studies of pelvic fin development in teleosts and cartilagi- early tetrapod had terrestrially ineffectual hindlimbs, as it nous fish. The zebrafish (Danio rerio), a powerful, geneti- lacked the necessary rotary motions in its hindlimbs to lift cally tractable , has recently been utilised to its body off the ground and therefore could not employ lat- study pectoral fin/forelimb developmental mechanisms; eral sequence walking (Pierce et al. 2012). This new study however, only a few studies have focused on the pelvic fins. indicates that early tetrapods went through a stage of hip- Zebrafish pelvic fin buds develop around 3 weeks post- joint restriction before they evolved the locomotory behav- fertilisation as two small mesenchymal bulges that emerge iours of modern tetrapods (Pierce et al. 2012). Recently, from the lateral plate mesoderm (LPM) of the abdominal Swartz (2012) described a well preserved fossil specimen of flank of the fish (Grandel & Schulte-Merker, 1998). The mes- the extinct of sarcopterygian fish from the Middle enchymal bulges in the pelvic fin region proliferate to cre- Devonian, Tinirau clackae. Tinirau shares many advanced ate two pelvic fins bud covered at their distal edge by an features with later tetrapodomorphs in the pelvic elements. apical ectodermal thickening (Grandel & Schulte-Merker, Tinirau is the earliest known stem tetrapod to have a signif- 1998). This structure, which is thought to be analogous to icantly reduced postaxial process, and a fibula more like the tetrapod apical ectodermal ridge (AER; Grandel & those of later tetrapods. Caudally, the pelvis articulates with Schulte-Merker, 1998), is also seen during trout pelvic fin a femur that is preserved in association with the acetabu- development, where it was termed the pseudoapical ecto- lum. The postaxial fibular process is highly reduced and dis- dermal ridge (Geraudie, 1978). The apical ectodermal thick- plays a similar ‘lip’ overhanging the postaxial edge of the ening of the pelvic fins is only transient and becomes the fibulare. The lack of a prominent postaxial process in the apical fold which is formed by a layer of dorsal and ventral fibula of Tinirau is more similar to the condition observed cylindrical cells (Grandel & Schulte-Merker, 1998). As the in crownward taxa. This pattern underscores previous phy- apical fold of the pelvic fin is infiltrated by the migrating logenetic reconstructions of the appendicular skeleton in mesenchyme it morphs into a fin fold, which gives rise to which conventional limb characteristics first the adult fin (Grandel & Schulte-Merker, 1998). originate in the pelvic fins. The early stages of hindlimb development in fish fins are Historically, the evolution of the neural control in the pel- similar to those of tetrapod (mouse and chick) hindlimbs. In vic fins and hindlimbs associated with this transition has not the chick embryo, the LPM in the prospective hindlimb received much attention. However, a recent review has region thickens and an AER forms in the overlying compared the organization of the motor neurons in the ectoderm around 3 days post-fertilisation (reviewed in spinal cord of various vertebrates which aids in the Saunders, 1977). The hindlimb then continues to grow and

© 2012 The Authors © Journal of Anatomy 2012 Anatomical Society 15 8 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

elongate. In the mouse embryo, the hindlimb buds first known that Hox genes have a unique ability to establish appear around 10 days post-fertilisation, but the AER morphologies along the anterior–posterior axis of an appears later than in the chick (Martin, 1990). Like the embryo by establishing a pre-pattern of the embryonic axis chick, the mouse limb continues to grow and elongate; (Lewis, 1978; reviewed Garcia-Fernandez, 2004;). It is also however, there is a difference in shape between developing known that positional differences along the body axis, such chick and mouse hindlimbs. as the positions of the limbs, are specified by the staggered boundaries of Hox gene expression (Cohn et al. 1997; Akam, 1998; Marshall et al. 1996; Krumlauf, 1994). Noro Positioning et al. (2011) showed that the regionalisation of the preso- Development of paired appendages at appropriate levels mitic mesoderm along the embryonic axis by the Hox code along the primary body axis is a hallmark of the body plan is responsible for the specification of the limb and flank of jawed vertebrates. In all jawed vertebrates, the paired fields in the chick embryo (Noro et al. 2011). Through trans- appendages arise from a region known as the lateral com- plant and ablation experiments it was shown that the petent stripe (Yonei-Tamura et al. 2008). The lateral compe- presomitic mesoderm adjacent to the prospective limb field tent stripe is defined as a region along the flank that is is permissive to the development of a limb field and affects competent for paired appendage development if the cor- the size of the limb field while the paraxial mesoderm adja- rect signals are received (Yonei-Tamura et al. 2008). The cent to the flank suppresses limb bud development (Noro entire length of the lateral competent stripe of all jawed et al. 2011). In addition, it has been suggested that Hox vertebrates studied has the competency to form paired transcription factors may be responsible for the restricted appendages (Yonei-Tamura et al. 2008). The utilisation of patterns of Tbx4 (Minguillon et al. 2005). The position of the lateral stripe differs greatly, with each species having a the pelvic fins/hindlimbs in jawed vertebrates is thought to different ratio for the forelimb/pectoral fin, interlimb and be determined by Hoxb9, Hoxc9, Hoxd9 and Hoxc10 activity hindlimb/pelvic fin regions. Differences in regulation of the in the developing embryo (Cohn et al. 1997; Lance-Jones Hox genes in the paraxial mesoderm are thought responsi- et al. 2001; Tanaka et al. 2005; Choe et al. 2006; Murata ble for the variations in the positioning of paired append- et al. 2010). ages (Burke et al. 1995; Sordino et al. 1995; Burke & There is a phylogenetic correlation between Hoxd9 Nowicki, 2003; Noro et al. 2011). expression and hindlimb/pelvic fin positioning. During very Hox genes are a family of transcriptional regulators that early chick development and teleost development, the pri- are involved in axial patterning of many structures in verte- mary pattern of Hoxb9, Hoxc9 and Hox9d expression is stag- brates (Gruss & Kessel, 1991; Krumlauf, 1994; Burke et al. gered in the prospective forelimb/pectoral fin region, and 1995; Deschamps et al. 1999), including fins and limbs strong expression is also observed in the interlimb/fin and (Yokouchi et al. 1991; Sordino et al. 1995; Nelson et al. prospective hindlimb/pelvic fin region (Cohn et al. 1997; 1996). In jawed vertebrates, the Hox genes are classified Tanaka et al. 2005) (Fig. 6). At later stages of development, into 13 paralogous groups and are tightly clustered at four during tetrapod limb bud formation, Hoxd9 expression dis- loci: HoxA to HoxD. A clear correspondence between partic- appears from the interlimb region and is restricted to ular Hox groups and defined morphological boundaries domains adjacent to the prospective limb buds (Cohn et al. along the antero-posterior axis of jawed vertebrates has 1997) (Fig. 6A). During pelvic fin budding in the threespine been documented (Gaunt, 1994; Burke et al. 1995; Cohn & stickleback fish (Gasterosteus aculeatus), the expression Tickle, 1999; Kmita & Duboule, 2003). It has long been domain of Hoxd9 extends from the prospective pelvic

AB

Fig. 6 In situ hybridisation of Hoxd9 during pelvic fin/hindlimb positioning in chick (A) and threespine stickleback (B). (A) In limb budding stages in chick Hoxd9 expression disappears from the interlimb region and is restricted to domains adjacent to the prospective limb buds (image reproduced with permission from Cohn et al. 1997). (B) During pelvic fin development stages in threespine stickleback, Hoxd9 expression extends from the prospective pelvic region and is maintained in the interlimb region (image reproduced with permissions from Tanaka et al. 2005).

© 2012 The Authors 16 Journal of Anatomy © 2012 Anatomical Society Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 9

region and is maintained in the interlimb region ABChick Zebrafish (Tanaka et al. 2005) (Fig. 6B). It is thought that Hoxd9 activ- Tbx 5 ity is particularly important for the development of pelvic fins/hindlimbs as its absence is thought to be the cause of the loss of pelvic fins in fugu (Takifugu rubripes), in a mech- anism analogous to the loss of pectoral axial patterning cues in python snakes (Pythonidae) (Cohn & Tickle, 1999; Tanaka et al. 2005). Recent work has also showed a conserved role for Hoxc10 in the positioning of hindlimbs/pelvic fins. In tetrapods, the hindlimb buds arise in the body wall at the level of Hox10 Tbx4 expression in the spinal cord (Lance-Jones et al. 2001; Choe et al. 2006). However, it was observed that in teleosts the level of Hoxc10a expression did not correlate to the position of pelvic fins in the adult fish (Murata et al. 2010). How- ever, Murata et al. (2010) have shown that pelvic fin precur- sor cells do, in fact, lie next to the posterior expression boundary of Hoxc10a and that these cells migrate to the Fig. 7 In situ hybridisation of Tbx4 and Tbx5 in chick (A) and zebra- pelvic fin field due to trunk-tail protrusion of the growing fish (B). (A) Tbx5 is expressed in the developing wing buds of chick embryo. In addition, it has been shown in knockout mice embryos; Tbx4 is expressed in the developing leg buds (figure repro- duced with permission from Logan, 2003). (B) In zebrafish, Tbx5 is and zebrafish morphants that removal of the TGF beta fam- expressed in the developing pectoral fins and Tbx4 is expressed in the ily member, Gdf11, changes the expression domain of developing pelvic fins (figure reproduced with permission from Ruvin- Hoxc10 and subsequently changes the axial position of the sky et al. 2000). hindlimb/pelvic fin (McPherron et al. 1999; Murata et al. 2010). In mice, gene deletion-gene replacement studies using knockout and transgenic mice have shown that in the Pelvic fin/hindlimb-type specification absence Tbx5 in the forelimb, Tbx4 does not confer hind- An important step during the development of pelvic fins/ limb-like morphology to the limb (Minguillon et al. 2005, hindlimbs is the establishment of a fin/limb-type specifica- 2009). If Tbx5 is ablated in the forelimb of mice and tion. In the past, two genes from the T-box transcription replaced with ectopic Tbx4, the forelimbs have a normal factor family, Tbx4 and Tbx5, were thought to determine forelimb-type pattern of gene expression and a forelimb- limb identity, due to their expression patterns (Gibson- type skeletal morphology (Minguillon et al. 2005). These Brown et al. 1996; Tamura et al. 1999; Ruvinsky et al. 2000). results suggested that other factors must determine limb- In , teleosts and anurans Tbx4 is expressed in type morphology. In addition, it has been shown that there developing hindlimbs and Tbx5 is expressed in developing is a low level of expression of Tbx4 in the developing fore- forelimbs (Gibson-Brown et al. 1996; Logan et al. 1998; limbs of mouse (Gibson-Brown et al. 1996; Naiche et al. Tamura et al. 1999; Ruvinsky et al. 2000; Takabatake et al. 2011). The forelimb expression domain of Tbx4 has been 2000) (Fig. 7). Historically, this expression pattern was shown to be unnecessary for forelimb morphology in mice thought to be conserved across vertebrates with paired and the enhancer that drives this expression is not evolu- limbs. tionarily conserved outside of the mammalian class (Menke However, there is conflicting evidence about the ability et al. 2008; Naiche et al. 2011). However, this is similar to of Tbx4 and Tbx5 to confer limb-type identity. Ectopic gene the situation in urodeles, where both Tbx4 and Tbx5 are misexpression studies in chick embryos have shown that expressed in the developing fore and hindlimbs of N. viri- Tbx4 and Tbx5 play a role in determining limb-type mor- descens during limb development (Khan et al. 2002). These phology. Ectopic expression of Tbx5 in the developing chick results shed doubt on the specific role of Tbx4 in determin- hindlimb bud can confer forelimb-type morphology to the ing hindlimb-like identity. In contrast, studies by Ouimette developing limb and induce ectopic expression of forelimb et al. (2008) showed that while Tbx4 does not confer hind- markers (Rodriguez-Esteban et al. 1999; Takeuchi et al. limb-like morphology when ectopically expressed in the 2003). Equally, it has been shown that misexpression of forelimb, it is sufficient to rescue hindlimb-like morphology Tbx4 in developing chick forelimb buds gives the develop- to Pitx1 null hindlimbs, which lack hindlimb-like morphol- ing limb hindlimb-type morphology and induces ectopic ogy. Ouimette et al. (2008) also demonstrated that Tbx4 expression of hindlimb markers (Takeuchi et al. 2003). How- has a unique transcriptional repressor site which is possibly ever, this is in direct contrast to some results obtained from responsible for hindlimb-like morphology, and that this other species as detailed below. transcriptional repressor site is absent in Tbx5 and the

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 17 10 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

ancestral Tbx4/5 cluster. It is important to note that due to hindlimb bud development in chick and mouse. More the coupling of limb initiation and limb-type morphology recently, a few of these initiation factors have been investi- the rescued hindlimb-like morphologies may be due to the gated in teleosts, showing that many of the same molecules activity of Tbx4 on hindlimb-like morphology or the return seem to be employed (Gibson-Brown et al. 1996; Tamura of the normal hindlimb development initiation cascade. et al. 1999; Ruvinsky et al. 2000; Cole et al. 2003; Shapiro Although the studies on Tbx4 and Tbx5 appear to be et al. 2004). The origin of the limb/fin bud induction signal inconclusive concerning their role in limb-type specification, remains controversial and there is evidence to show that it it is clear that there are other factors involved in this pro- begins in either the presomitic/paraxial mesoderm or the cess. The paired-like homeodomain transcription factor, intermediate mesoderm. Historic studies on chick forelimb Pitx1, is expressed in the region forming the pelvic append- bud induction have shown that a foil barrier placed age and in the pelvic appendage buds, but not in the fore- between the somites and the LPM blocks forelimb bud limb regions (Lamonerie et al. 1996; Cole et al. 2003). Pitx1 induction (Murillo-Ferrol, 1965; Sweeney & Watterson, is thought to be involved in both hindlimb identity specifi- 1969; Stephens & McNulty, 1981). However, these results cation and hindlimb development (Lamonerie et al. 1996; have been reinterpreted by Crossley et al. (1996) as blocking Lanctot et al. 1999). In tetrapods, expression of Pitx1 is an Fgf8 signal from the intermediate mesoderm, which sub- observed before the onset of Tbx4 expression (Lamonerie sequently caused a failure of chick limb bud induction. et al. 1996). In both mouse and chick, misexpression of Pitx1 More recent work in zebrafish fins has shown that the fin in the forelimb imparts hindlimb-like characteristics to the bud induction signal begins in the paraxial mesoderm and muscles, skeleton and (Logan & Tabin, 1999; Takeuchi is transferred by secondary signals to the LPM and the over- et al. 1999; DeLaurier et al. 2006). In addition, Pitx1 null lying ectoderm (Begemann et al. 2001; Grandel et al. 2002; mice develop hindlimbs in which the skeleton has lost some Gibert et al. 2006; Mercader et al. 2006; Grandel & Brand, of its hindlimb-like characteristics (Lanctot et al. 1999; Szeto 2011). As the paraxial mesoderm and the LPM are separated et al. 1999; Marcil et al. 2003). However, as Pitx1 null mice by the intermediate mesoderm, it seems likely that mole- do develop hindlimbs and not all hindlimb-like features are cules from the paraxial mesoderm could signal to the LPM affected, it is possible that other factors are involved in the through the intermediate mesoderm, where both the par- specification of hindlimb-type morphologies in tetrapods axial mesoderm and the intermediate mesoderm would (Fig. 8). The role of Pitx1 in specifying pelvic fin identity still have a role in limb/fin bud initiation. needs to be determined in fish to confirm whether its hind- limb identity role is conserved amongst vertebrates. The role of Fgf signalling and the intermediate mesoderm in pelvic fin/hindlimb bud induction Previous work in tetrapods has suggested that Fgf activity Initiation in the intermediate mesoderm might trigger the induction Several extracellular signalling molecules and transcription of limb budding by transferring a signal to the limb fields factorshavebeenreportedtobeinvolvedininitiating of the LPM (Crossley et al. 1996; Vogel et al. 1996). The Fgf8 gene product was thought to play this role, as it is expressed in the intermediate mesoderm adjacent to the AB limb-forming regions before and during limb bud induction and can maintain cells in a proliferative state (Crossley et al. 1996; Vogel et al. 1996). Furthermore, it has been shown that the application of an Fgf8-soaked bead to the inter- limb region of tetrapods can induce an ectopic limb to form (Cohn et al. 1995; Mahmood et al. 1995; Ohuchi et al. 1995; CD Crossley et al. 1996; Vogel et al. 1996; Yonei-Tamura et al. 1999). However, recent studies have demonstrated that condi- tional removal of certain Fgf activity from the mesoderm has little effect on limb bud induction in mice (Boulet et al. 2004; Perantoni et al. 2005). When Fgf8 activity is transgeni- cally removed from the mesoderm of mice, the mutant’s Fig. 8 Pelvic appendage skeletal preparations from wild-type (A) and limbs form at the normal position and time (Perantoni et al. Pitx1 null (B) mice and wild-type (C) and Pitx1 null threespine stickle- 2005). Likewise, transgenic mice lacking both Fgf4 and Fgf8 back (D). (A, B) Mice lacking Pitx1 develop small hindlimbs that have lost some hindlimb characteristics (figure reproduced with permission expression in the mesoderm also develop limbs which form from Duboc & Logan, 2011). (C, D) Threespine stickleback which lack at the normal position and time (Boulet et al. 2004). Addi- Pitx1 in the pelvic regions do not develop pelvic spines (figure tionally, in zebrafish, it has been shown that Fgf signalling is reproduced with permission from Cole et al. 2003). not required for pectoral fin bud initiation (Mercader et al.

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 18 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 11

2006). Zebrafish embryos treated with the Fgf receptor Downstream pelvic fin/hindlimb bud induction cascade antagonist, SU5402, display normal pectoral fin bud induc- Although the specific mechanisms of the pelvic fin/hindlimb tion in the face of a lack of Fgf signalling (Mercader et al. bud induction cascade remain unknown, secondary signals 2006). Despite controversy over a role of Fgf activity during are thought to trigger the expression of Tbx4 in pelvic fins/ limb/fin bud induction, Fgf activity is certainly necessary dur- hindlimbs. Recent studies in mice have suggested that Tbx4 ing later stages of limb/fin budding and development. and Tbx5 play a shared role in limb initiation (Minguillon et al. 2005, 2009). Tbx4 is one of the first genes known to The role of retinoic acid and the somites in pelvic fin/ be expressed in the prospective pelvic fin/hindlimb region hindlimb bud induction during pelvic fin/hindlimb bud induction (Gibson-Brown Recently, it has been proposed that a retinoic acid- et al. 1996; Tamura et al. 1999; Ruvinsky et al. 2000). In controlled signal is responsible the onset of Tbx5 expression jawed vertebrates, Tbx4 isexpressedinthedeveloping pel- in pectoral fins and the induction of pectoral fin buds vic fin/hindlimb just before and during the pelvic fin/hind- (Gibert et al. 2006; Mercader et al. 2006; Grandel & Brand, limb bud induction stages (Gibson-Brown et al. 1996; 2011). Evidence from these studies suggests that retinoic Tamura et al. 1999; Ruvinsky et al. 2000). Tbx4 and its fore- acid and Tbx5 signalling is upstream of Fgf function which limb counterpart Tbx5 have been shown to share a tran- is recently thought to play only a local role in the limb bud scriptional activator site which has been suggested to be during induction, at least in zebrafish (Gibert et al. 2006; responsible for the shared limb initiation activity of these Mercader et al. 2006; Grandel & Brand, 2011). genes (Ouimette et al. 2008). Under normal conditions, In zebrafish, studies of retinoic acid signalling have shown Tbx4 activity leads to the initiation of Fgf10 signalling in that its effect on pectoral fin bud induction is probably the bud mesoderm and the subsequent establishment of indirect and likely to be mediated by secondary signals the FGF positive signalling feedback loop between the bud (Mercader et al. 2006). It has been shown that wnt2b relays mesenchyme and the overlying ectoderm (see later) (Fig. 9). the retinoic acid fin bud induction signal to the pectoral fin Mouse and chick studies have found that Tbx4 is necessary field and induces Tbx5 expression (Mercader et al. 2006). In for proper hindlimb bud induction and subsequent hind- chick embryos, wnt2b in forelimbs and wnt8c (previously limb outgrowth (Naiche & Papaioannou, 2003, 2007; Takeu- known as wnt8a) in hindlimbs are thought be involved in chi et al. 2003). Conditional mice knockout models have limb bud induction (Kawakami et al. 2001). In the chick shown that a loss of Tbx4 activity results in a drastically embryo, wnt8c is expressed in a domain that suggests it reduced hindlimb bud which fails to complete limb bud may be involved in hindlimb bud induction (Kawakami induction, while overexpression of a dominant negative et al. 2001). It addition, ectopic expression of wnt8c has form of Tbx4 in the prospective hindlimb fields of chick been shown to induce hindlimb buds along the flank of embryos results in a complete legless phenotype (Naiche & chick embryos (Kawakami et al. 2001). Papaioannou, 2003, 2007; Takeuchi et al. 2003). However, neither wnt2b nor wnt8c have been detected in the limb buds of early mice embryos (Agarwal et al. 2003). This may be due to the fact that Tbx5 is not necessary fortheestablishmentofthelimbfieldinthisspecies,asitis in zebrafish, or that the limb bud induction signal may come from the intermediate mesoderm and not the somitic mesoderm in mice (Crossley et al. 1996; Agarwal et al. 2003; Ahn et al. 2002; Gibert et al. 2006; Mercader et al. 2006; Grandel & Brand, 2011). Recent studies from mice have con- firmed that although retinoic acid promotes mouse fore- limb bud induction, it is not necessary for hindlimb bud induction, as retinoic acid-deficient mice develop normal hindlimb buds (Zhao et al. 2009). These authors concluded that the role of retinoic acid in mouse limb development was to suppress Fgf8 activity along the flank, allowing for a Fig. 9 A simplified version of some of the key genes involved in pelvic permissive domain for forelimb bud induction to occur; fin/hindlimb development. It should be noted that some of these however, this mechanism is not necessary for Fgf8 regula- genes have not yet been investigated in pelvic fins. Both Hoxd9a and tion at the time the hindlimbs develop (Zhao et al. 2009). Hoxc10a have been shown to be involved in the positioning of the pelvic appendage in amniotes and teleosts. Wnt8c has the ability to The difference between permissive signals for fore and induce ectopic hindlimbs in chick. The hindlimb identity genes Tbx4 hindlimb induction may account for the ability of fish to and Pitx1 have both been shown to play roles in hindlimb specification develop pelvic fins at such a relatively late stage compared and initiation. A positive feedback loop between Fgf10 in the meso- to pectoral fins. To date, it remains uncertain which mecha- derm, Fgf8 in the apical ectoderm and Shh in the ZPA controls the nisms lead to the induction of hindlimb/pelvic fin buds. outgrowth and patterning of the pelvic appendage.

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 19 12 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

In addition to Tbx4 signalling, it is possible that the hind- signalling centres; the AER, the zone of polarising activity limb-related gene Pitx1 may play an important role in hind- (ZPA) and the non-ridge ectoderm. These signalling centres limb/pelvic fin bud induction. In tetrapods, Pitx1 is neither are conserved in jawed vertebrates, are interdependent and necessary nor sufficient for hindlimb bud induction, as Pitx1 are established early through communication between the null mice possess small hindlimbs and Pitx1 misexpression limb/fin bud mesoderm and the overlying ectoderm. cannot induce ectopic hindlimbs (Lanctot et al. 1999; Szeto et al. 1999; Minguillon et al. 2005)(Fig. 8A and B). However, The establishment of the apical ectodermal ridge Pitx1 is known to partially regulate Tbx4 in developing The establishment of an AER is essential to the develop- hindlimb buds, as Pitx1 knockout mice display reduced Tbx4 ment of hindlimbs and teleost pelvic fins. The AER expression (Lamonerie et al. 1996; Lanctot et al. 1999). Pitx1 is a thickened layer of ectodermal cells at the distal tip of a is thought to regulate Tbx4 via a Tbx4 upstream enhancer developing limb/fin bud (Fig. 10). In the forelimb/pectoral region in tetrapods (Menke et al. 2008). Tetrapods have fin, specific gene expression in the dorsal and ventral ecto- two Tbx4 enhancer regions, 5′ HLEA and 3′ HLEB, and only derm is thought to induce the distal limb ectoderm to the 3′ HELB is conserved in fish (Menke et al. 2008). Only become ridge-like (see Fernandez-Teran & Ros, 2008 for a the 5′ HLEA has Pitx1 putative binding sites and it is detailed review; Hatta et al. 1991; Norton et al. 2005). It thought that in tetrapods, Pitx1 regulates Tbx4 expression remains unknown whether a similar process occurs in hind- through this enhancer region (Menke et al. 2008). It limbs and pelvic fins and additional experiments are remains to be determined whether Pitx1 regulates Tbx4 needed. However, in hindlimbs it is known that as the expression in fish, as it is unknown if the 3′ HLEB enhancer developing hindlimb bud reaches a sufficient size, Tbx4 sig- region interacts with Pitx1. nalling triggers Fgf10 expression in the mesenchyme of the In contrast to mouse hindlimbs, pelvic fin bud induction bud, which in turn is necessary to activate Fgf8 expression in teleosts seems to be dependent on Pitx1 activity. It has in the overlying ectoderm (see Fernandez-Teran & Ros, 2008 been shown that the absence of Pitx1 activity in the pro- for a detailed review). In developing amniote hindlimb spective pelvic regions of certain populations of threespine buds, mesenchymal Fgf10 signals to Fgf8 in the overlying sticklebacks is linked to the failure of pelvic spine formation ectoderm via wnt3a (chick) or wnt3 (mouse) (Barrow et al. at an early stage (Cole et al. 2003; Shapiro et al. 2004) 2003; Narita et al. 2005, 2007) (Fig. 9). In mouse, the ongo- (Fig. 8C and D). The absence of Pitx1 activity and subse- ing activity of Fgf8 in the AER is necessary for proper out- quent failure of pelvic spine development has been attrib- growth of the hindlimb (Lewandoski et al. 2000; Moon & uted to regulatory mutations deleting a tissue-specific Capecchi, 2000) (Fig. 11A). enhancer of Pitx1, which causes specific pelvic region loss of Based on evidence from zebrafish pectoral fin develop- Pitx1 expression in certain populations of threespine stickle- ment, it is thought that similar Fgf signalling establishes the backs (Chan et al. 2010). Due to the differences in morphol- pseudoapical ectodermal ridge in teleost pelvic fins. How- ogy that loss of Pitx1 in tetrapods and teleost causes, it ever, in contrast to mouse and chick, in the developing seems possible that Pitx1 plays a slightly different role in tel- zebrafish pectoral fin buds the expression of Fgf8 is not eost pelvic fin development than it does in amniote hind- observed until after the establishment of the AER (Fischer limb development. It would be of great interest to examine et al. 2003). In fact, Fgf8 is not absolutely necessary for pec- the role of Pitx1 in the induction of hindlimbs in other ver- toral fin development, as the Fgf8 mutant zebrafish acere- tebrates situated phylogenetically between teleosts and bellar develops normal pectoral fins (Reifers et al. 1998) amniotes. (Fig. 11B). Instead, in developing zebrafish pectoral fin buds, expression of Fgf16 and Fgf24 is seen before the expression of Fgf8 (Fischer et al. 2003; Nomura et al. 2006). Outgrowth In addition, Fgf8 expression is dependent on Fgf16 activity The growth from a bulge in the body wall to a bud that (Fischer et al. 2003; Nomura et al. 2006). The zebrafish grows out independently is a critical step in limb develop- mutant, ikarus, which has a point mutation in the coding ment and involves interaction between the three main region of Fgf24, fails to develop pectoral fins which

AB

Fig. 10 Scanning electron microscopy of the AER of a developing mouse hindlimb bud (A) and a developing zebrafish pelvic fin (B) (figure reproduced with permission from Martin, 1990 and Don et al. 2011).

© 2012 The Authors 20 Journal of Anatomy © 2012 Anatomical Society Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 13

A domain of Shh is conserved between the developing hind- limb/pelvic fin buds of many jawed vertebrates, including mouse, chick and zebrafish (Echelard et al. 1993; Krauss et al. 1993; Roelink et al. 1994; Don EK & Cole NJ, unpublished). However, the evolutionary origin of the ZPA remains unclear, as studies by Tanaka et al. (2002), Dahn B et al. (2007) and Yonei-Tamura et al. (2008) have shown key spatial and temporal differences in Shh expression in cartilaginous fish from those found in tetrapods and teleo- sts. In cartilaginous fish (Chondrichthyes), Shh expression is only seen during later stages of post-budding pelvic fin development and the expression domain does not map to Fig. 11 Skeletal preparations of hindlimbs of wild-type and Fgf8 null the ZPA of the developing pelvic fins (Tanaka et al. 2002; mice (A) and wild-type and Fgf8 mutant zebrafish pectoral fins (B). Dahn et al. 2007; Yonei-Tamura et al. 2008). While the hindlimbs of Fgf8 null mice have skeletal outgrowth defects, the pectoral fins of Fgf8 mutant zebrafish develop normally (figure reproduced with permission from Lewandoski et al. 2000 and Reifers Development of the muscles of the pelvic fins/ et al. 1998). hindlimbs Recently, the development and evolution of the muscula- demonstrate that Fgf24 is necessary for proper pectoral fin ture of the pelvic fin/hindlimb has been examined in several outgrowth. However, Fgf24 is not necessary for pelvic fin species occupying key phylogenetic positions (Cole et al. development, as ikarus mutants develop normal pelvic fins 2011). In amniotes, and the pectoral fins of (Fischer et al. 2003). bony fish, the limb and pectoral fin muscles are generated by limb myoblasts which are derived from the migration of AER maintenance mesenchymal precursor cells (Nicolas et al. 1998; Neyt et al. Despite the differences in morphology, studies of teleosts, 2000; Satoh et al. 2005; Vasyutina & Birchmeier, 2006; Sabo amphibians and amniotes have shown that the formation et al. 2009). These cells, under the direction of Lbx and maintenance of an AER/pseudoapical ectodermal ridge/ de-laminate from the hypaxial region of the hindlimb-level apical thickening is necessary for proper hindlimb out- somites and migrate to the developing hindlimb mesen- growth and patterning along the proximal distal axis. As chyme (Nicolas et al. 1998; Satoh et al. 2005; Vasyutina & the limb bud grows outward, the ridge maintains a zone of Birchmeier, 2006; Sabo et al. 2009) (Fig. 12A). undifferentiated mesenchymal cells at the distal tip of the In contrast, within chondrichthyan species, the pectoral bud, whereas the more proximal cells differentiate. The role and pelvic fin muscles are formed by a migrating epithelial of the ridge in promoting limb bud outgrowth was first bud tipping a direct myotome extension (Neyt et al. 2000 demonstrated in chick embryos. It was shown that when and references within). The migrating epithelial buds the ridge is cut away, limb bud outgrowth ceases and a extend ventrally from the myotome and enter the develop- truncated limb develops (Saunders, 1948). Unlike teleosts ing fin mesenchyme to generate the paired fin muscles. This and amniotes, direct developing , such as the treefrog mechanism is not under the control of Lbx as no expression ( coqui), lack a morphological AER, but do of Lbx is seen during this process (Neyt et al. 2000; Cole have an apical ectodermal thickening that is necessary from et al. 2011) (Fig. 12C). proper limb patterning, as its removal results in anterior A distinct process which incorporates both primitive and defects in skeletal patterning (Richardson et al. 1998). How- derived characteristics of vertebrate appendicular muscle ever, the defects seen are not as severe as those seen in formation has been described in the pelvic fins of bony fish amniotes, which may be due to the ability of the apical (Cole et al. 2011). A recent study by Cole et al. (2011) thickening of amphibians to regenerate (Tschumi, 1957; showed that in zebrafish, paddlefish and lungfish pelvic Richardson et al. 1998; Stopper & Wagner, 2005). fins, an epithelial myotomal extension tipped by an epithe- lial bud, extends towards the pelvic fin bud but fails to enter The AER and the ZPA the developing mesenchyme of the pelvic fin bud. Instead, While AER maintenance requires continuous positive Wnt the pelvic fin muscle precursors are carried ventrally by the and Fgf signalling between the mesoderm and the overly- myotome extension and, once in position, undergo an epi- ing ectoderm, another feedback loop involving the ZPA is thelial–mesenchymal transition. These cells are then induced also necessary for proper AER maintenance and outgrowth. to express Lbx and migrate into the fin mesenchyme to form The ZPA is a group of cells located in the posterior mesen- the individual pelvic fin muscles (Cole et al. 2011) (Fig. 12B). chyme of the developing hindlimb bud that express Shh This mode of muscle formation represents an important (Saunders & Gasseling, 1968; Riddle et al. 1993). This step in the evolution of the tetrapod hindlimb muscle

© 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 21 14 Evolution and development of the pelvic fin/hindlimb, E. K. Don et al.

A (claspers) of sharks. In addition, secondary loss of hind- limbs/pelvic fins has occurred repeatedly throughout the evolution of many different lineages, including , and fish. In some species, hindlimbs are completely B absent (Fugu), whereas in others, remnants remain (pythons, whales) (Cohn & Tickle, 1999; Bejder & Hall, 2002; Santini & Tyler, 2003; Tanaka et al. 2005). Further to this, an asymmetry of pelvic fin loss is observed in other species, such as stickleback and manatees (Bell & Orti, 1994; Cole et al. 2003; Shapiro et al. 2004; Shapiro et al. 2006). Pelvic fin loss is documented in one or more species of C 92 teleostean families; the pelvic fin has been lost about 50 independent times, excluding multiple losses within fami- lies (Nelson, 1993). Many freshwater populations of three- spine sticklebacks have undergone pelvic spine (a modified pelvic fin) loss, ranging from complete loss to asymmetric loss, from an anadromous ancestor (Bell & Orti, 1994; Cole Fig. 12 Diagram of the different modes of fin/limb muscle formation. et al. 2003; Shapiro et al. 2004). The genetic basis of pelvic (A) Amniotes and anurans use long-range migration of individual mes- spine loss has been examined in freshwater Scottish and enchymal migratory myoblasts in both the fore- and hindlimbs for freshwater Canadian populations of threespine sticklebacks limb muscle formation. Bony fish deploy this mode of fin muscle for- mation in the pectoral fins. (B) Zebrafish utilise the long-range migra- and has been determined to be caused by an upstream tion of individual mesenchymal migratory myoblasts from an epithelial pelvic region regulator of Pitx1 (Chan et al. 2010). Pelvic myotomal extension to make the muscle of the pelvic fin. (C) Chondri- fin loss in threespine sticklebacks was also shown to be chthyans utilise the primitive mechanism of direct epithelial extension asymmetric, with greater reduction of the right than the to generate the muscle of pectoral and pelvic fins (figure reproduced left (Cole et al. 2003; Shapiro et al. 2004). This right–left from Cole et al. 2011). asymmetry has been attributed to the partial functional compensation of a closely related gene Pitx2, which is pref- erentially expressed on the left side during development developmental mechanisms which evolved during the oste- (Shapiro et al. 2004). However, no Pitx2 transcripts were ichthyan radiation. By adopting a more derived mode of detected in the pelvic regions of pelvic spine-deficient pelvic fin muscle formation, the pelvic fin could be located threespine sticklebacks during pelvic spine development anywhere in the dorsoventral body axis, as well as allowing stages (Cole et al. 2003). The same right–left asymmetry is for earlier deployment of muscle precursors to the pelvic fin also seen in the hindlimbs of Pitx1 knockout mice and it environment, both of which may have facilitated the devel- has been suggested that the right–left asymmetry seen in opment of more robust, weight-bearing hindlimbs. the hindlimb remnants of manatees may also be due to The later stages of development of muscles of the hind- the same Pitx1/Pitx2 mechanism (Lanctot et al. 1999; Marcil limbs of tetrapods are thought to be under the influence of et al. 2003; Minguillon et al. 2005; Szeto et al. 1999; Shap- Pitx1 and Tbx4.IfTbx4 activity is absent during later stages iro et al. 2006). of hindlimb development, there is no effect on the out- In teleosts, different mechanisms account for pelvic fin growth of the limb skeleton but muscle patterning is loss in different species. Adult pufferfish completely lack a affected (Hasson et al. 2010). Although, an absence of Tbx4 bony pelvic apparatus and display no evidence of pelvic fin expression affects hindlimb muscle patterning, misexpres- bud development (Santini & Tyler, 2003; Tanaka et al. sion of Tbx4 in the developing forelimb does not confer 2005). The cause of pelvic fin loss in this species is thought hindlimb-like morphologies to the muscles of the forelimb to be caused by an absence of late onset Hox gene expres- (DeLaurier et al. 2006). It seems that Pitx1 mayplayagreater sion, specifically Hoxd9a (Tanaka et al. 2005). Some popula- role in determining hindlimb-like muscle morphology, as tions of zebrafish have also undergone secondary hindlimb misexpression of Pitx1 in the forelimb of mice gives the mus- loss. Pelvic finless zebrafish initiate pelvic fin development cles more hindlimb-like morphologies (DeLaurier et al. and form mesenchymal bulges in the pelvic region but fail 2006).Themechanismsofthelaterstagesofdevelopmentof to form a morphological pseudoapical ectodermal ridge the muscles of the pelvic fins are yet to be investigated fully. (Don et al. 2011). The pelvic fin buds then regress and no pelvic fins or pelvic skeletal apparatus is present in the adult fish (Don et al. 2011). Pelvic fin/hindlimb loss Secondary loss of hindlimbs has also occurred in many tet- Hindlimbs and pelvic fins show a wide range of morpho- rapod lineages, such as snakes, and cetaceans. All logies from the legs of man to the modified pelvic fins limbless tetrapods are descended from limbed ancestors.

© 2012 The Authors 22 Journal of Anatomy © 2012 Anatomical Society Evolution and development of the pelvic fin/hindlimb, E. K. Don et al. 15

Extant snakes have evolved from limbed ancestors, as Conclusions demonstrated by extinct snakes with hindlimbs, such as Pachyrhachis problematicus, Haasiophis terrasanctus and Fascinating new insights into the field of pelvic fin/hindlimb Podophis descouensi (Haas, 1980; Rieppel, 1988; Lee & Cald- developmental evolution have shed light on both develop- well, 1998; Coates & Ruta, 2000; Greene & Cundall, 2000; mental and evolutionary mechanisms. Studies of the pelvic Rage & Escuillie´, 2000; Tchernov et al. 2000). In more primi- fins/hindlimbs of model and non-model organisms in higher tive snakes, such as pythons, a rudiment of a hindlimb bud and lower vertebrates have shown that hindlimb develop- forms in the embryo (Cohn & Tickle, 1999). The early hind- mental mechanisms are conserved throughout evolution limb bud is initiated and begins to develop, but fails to con- with species specific differences accounting for the varied tinue outgrowth and a developed AER and ZPA are never morphologies and functions observed. We now know more formed (Cohn & Tickle, 1999). After a series of experiments than ever before about the evolution of hindlimb develop- demonstrating that the mesoderm of developing python mental mechanisms and are at a point that we can readily hindlimbs is capable of inducing both an AER and a ZPA in compare organisms. chick limb buds, it was suggested that the cause of hindlimb loss in pythons is due to changes in mesodermal Hox gene Acknowledgements expression (Cohn & Tickle, 1999). Whereas primitive snakes initiate hindlimb development, more advanced snakes com- We would like to acknowledge Dr Zerina Johanson, Dr Greg Edge- pletely lack hindlimb buds (Cohn & Tickle, 1999). Similar to combe and Claire Winnick for revision of the manuscript and also hindlimb loss in primitive snakes, many limbless lizards also two anonymous reviews whose comments significantly improved initiate hindlimb development (Camp, 1923; Essex, 1927; this manuscript. Gans, 1975; Presch, 1975). However, unlike primitive snakes, the hindlimb buds of slow and green lizards do References develop an AER, which then regresses (Raynaud et al. 1974; Raynaud, 1990). Agarwal P, Wylie JN, Galceran J, et al. (2003) Tbx5 is essential for forelimb bud initiation following patterning of the limb Although all modern cetaceans lack external hindlimbs, field in the mouse embryo. Development 130, 623–633. vestigial elements of the hindlimb skeleton can be found in Ahlberg PE (1989) Paired fin skeletons and relationships of the many adult cetaceans (Andrews, 1921; Howell, 1970). This is fossil group Porolepidormes (: ). 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28 © 2012 The Authors Journal of Anatomy © 2012 Anatomical Society 2.1. Updates to literature review Since the publication of this literature review, there have been several newly published works on the subject of the evolutionary history of the development of the pelvic fin/hindlimb.

2.1.1. Examination of Euphanerops The recent examination of the fossil of Euphanerops, a jawless fish from the Devonian has shown that this fish possessed paired anal (ventral) fin radials, but no pectoral or pelvic fins (Sansom et al., 2013). This unique anatomy is thought to have occurred at an early stage on the stem-gnathostome lineage (Sansom et al., 2013; Sansom et al., 2010; Janvier and Arsenault, 2007). This discovery has suggested to the authors that there was a large amount of morphological plasticity during this episode and that at this evolutionary time point a number of different were experimenting with a range of different body plans before settling on the two ‘arms’ and two ‘legs’ body plan of modern vertebrates (Sansom et al., 2013).

2.1.2. Publication of the African coelacanth genome Recently the genome of the African coelacanth ( chalumnae) has been published and analysed (Amemiya et al., 2013). The African coelacanth is the only living member of an ancient group of lobe-fined fishes (Smith, 1939) and is of interest to evolutionary biologists due to its controversial relationship to our ancient fish ancestors (Zimmer, 1999). This relationship is controversial as it has long been debated whether the (Neoceratodus forsteri) or the coelacanth is our closest living fish relative (Schultze and Trueb, 1991). Phylogenomic comparison of the genomes of the African coelacanth and the lungfish has demonstrated that the lungfish is in fact the closest living fish to the tetrapod ancestor (Amemiya et al., 2013). However, as the lungfish genome is so large (estimated at 50-100 Gb; Gregory, 2011) that it is difficult to work with, the publication of the African coelacanth genome remains critical to our understanding of the transition of vertebrates from water to land.

29 3. Materials and methods The recipes for buffers and solutions and a list of suppliers mentioned in this thesis can be found in Appendix A.

3.1. Production and manipulation of zebrafish The use and treatment of in the project were in accordance with the National Health and Medical Research Council guidelines. All protocols were approved by the Ethics Review Committee, University of Sydney (N.S.W., Australia) and the Animal Ethics Committee, Macquarie University (N.S.W., Australia).

Methods described herein were the standard operating procedures of the Cole Lab Facility, which were based on those in the Zebrafish Book (Westerfield, 2000), or were taken directly from the Zebrafish Book.

3.1.1. Generation and collection of embryos Zebrafish used for the experiments presented in this study were obtained from the natural spawning of several different breeding pairs. The strains used in this study were the pelvic finless zebrafish on a golden background (Don et al., 2011; Streisinger et al., 1986), wild- type zebrafish of either a WIK or Tübingen (Tü) background (Westerfield, 2000). In addition the pelvic finless/WIK heterozygous strain was utilised for positional cloning.

For all breeding pairs and colonies, the controlled environment in the aquarium was the same. The day/night cycle was controlled with an automatic timer (11 hr light/13 hr dark). The breeding fish were fed a varied diet including pellet, live artemia and frozen sterile bloodworms. The temperature (28°C), pH (neutral) and conductivity (800 µS/m) were checked daily. Embryos were collected by placing one male and one female zebrafish on each side of a false bottom pair tank the night before. With the onset of light in the morning the divider was removed, the male and female zebrafish were allowed to come into contact and . After spawning, the eggs were collected by straining the tank water containing the embryos through a plastic tea strainer. The eggs were then rinsed with system water and transferred into E3 embryo medium with 0.5% methylene blue (Sigma-Aldrich®) in a Petri dish (embryo medium). To help prevent mould, the faeces and other residue were removed by plastic disposable pipette. The eggs were them examined under a microscope to determine if they were fertilised. Fertilised eggs were left in the Petri dish and placed in an incubator at 28°C.

30 3.1.2. Microinjection Microinjection into zebrafish one-cell stage embryos is an established technique and was utilised in this project for both mRNA and DNA injections.

Zebrafish mating pairs were set up in a pair-breeding tank at night with a plastic divider separating the animals. At the addition of light in the morning, the plastic divider was removed and the pair was allowed to come into contact. Pairs were left for an average of 20 minutes before the eggs were collected. Eggs were screened for fertilisation and fertilised eggs were immediately used for microinjection. Eggs were lined up against a glass slide in a Petri dish and excess water was removed.

Microinjection was performed by hand under a steromicroscope (Leica) with a loaded glass needle and a Picospritzer™ II (General Valve™ corporation). Glass needles [1.0 mm outer diameter, 0.78 mm inner diameter, 150 mm length (Harvard Apparatus, GC100TF- 15SDR)] were prepared using a vertical pipette puller (Model 700D, David Kopf instruments) and the tips broken by contact with the glass slide.

3.1.3. Raising embryos to adulthood Embryos were maintained in embryo medium in a Petri dish in an incubator at 28 °C with regular cleaning for 3 days. From 3 dpf, hatched larvae were transferred to plastic baby tanks containing 2 cm of embryo medium and fed live paramecium cultures daily. From 10 dpf onwards, fry were fed live artemia once daily. At approximately 21 dpf, fry were placed on the main system and fed a diet including small pellet and live artemia.

3.1.4. Chorion removal Where required, small numbers of embryos were dechorionated manually using Dumont #5 watchmaker forceps (Simga-Aldrich®). Larger numbers of embryos were dechorionated enzymatically using pronase digestion. Pronase (2 mg/ml; Simga-Aldrich®) was added to embryo medium containing embryos and incubated for 10 minutes at 28ºC. To ease dechorionation, embryos were periodically passed through a plastic pasture pipette. Digestion was terminated when the majority of embryos had emerged from their chorions by several washes with embryo medium.

3.1.5. Fixation of zebrafish All zebrafish were anesthetised using tricaine (4 g/L, pH 7; Simga-Aldrich®), and were then washed twice quickly in phosphate buffered saline (PBS; Amresco). Samples were fixed and stored according to the intended purpose.

31 3.1.5.1. For in situ hybridisation Samples were fixed overnight in 4% paraformaldehyde (PFA, Amresco) in PBS at room temperature. After fixation, embryos were washed in PBS and dehydrated by a methanol (Amresco) series and stored at 4°C until needed.

3.1.5.2. For histology Samples were fixed overnight in 4% PFA in PBS at room temperature. After fixation, the heads and tails were removed from larger fish to allow for easier penetration of solutions. Fish were dehydrated via an ethanol series to 70% ethanol (Amresco) in preparation for immediate paraffin embedding. Paraffin embedding was processed overnight in a Tissue-Tek automatic tissue processing machine (Sakura). Samples were washed twice in 70% ethanol, and then further dehydrated through an ethanol series into 100% ethanol. Samples were then washed three times in xylene (Amresco), before being transferred through a 60 ºC xylene- paraffin series into 100% paraffin (Simga-Aldrich®). Samples were left at 60ºC in 100% paraffin until morning. In the morning, samples were processed on a Tissue-Tek embedding machine (Sakura). Samples in hot paraffin were placed into moulds and orientated. The blocks were allowed to cool and stored at room temperature until needed.

3.1.5.3. For skeletal staining Samples were fixed overnight in 4% PFA in PBS at room temperature. Samples were then washed with PBS to remove PFA residues and transferred to 50% ethanol. Samples were stored in 50% ethanol at 4°C until needed.

3.1.5.4. For electron microscopy Samples were fixed in 1% gluteraldehyde (Simga-Aldrich®) and 1% PFA in 0.1 M PBS (pH 7.2) for 2 hours at room temperature and postfixed in 1% OsO4 in 0.1 M PBS for 1 hour. Samples were then dehydrated through an ethanol series (2 x 50%; 70%; 2 x 90%; 95%; 3 x 100%) for 10 minutes for each step. Samples were then critical point dried using liquid to remove ethanol. Samples were attached to a stage using adhesive pads and then sputter coated with 15 nm of at 25 mA for 2 minutes. Samples were stored at room temperature until needed for imaging.

3.1.5.5. For extraction of genomic DNA Zebrafish intended for gDNA extraction were processed two different ways. Samples that were needed for morphological characterisation and rescue experiments were anesthetised after imaging and tail clips were taken. Tail clips were stored in 100% methanol at -20ºC in preparation for gDNA extraction.

32 Samples for mapping and positional cloning were sorted for the pelvic finless phenotype around 6 weeks post fertilisation (wpf) using the steromicroscope (Leica, M165FC stereo dissection microscope). Samples were either snap frozen in liquid nitrogen and stored in individual sterile labelled microfuge tubes at -80°C in preparation for extraction of genomic DNA or processed immediately.

3.1.5.6. For extraction of total RNA

After being euthanised, samples were washed twice with dH2O and stored in RNAlater (Qiagen) at 4°C overnight and then transferred to -20°C for long-term storage. Embryos were process whole, whereas the pelvic sections of older zebrafish were dissected prior to processing.

33 3.2. Molecular methods 3.2.1. Agarose gel electrophoresis Electrophoresis of DNA was performed as described (Sambrook and Russell, 2001). DNA samples were separated on 0.8-3% (weight/volume) DNA grade agarose (Sigma- Aldrich®) in 1x TBE or TAE buffer with the addition of 0.05 µg/ml of ethidium bromide. The gel loading buffer with dye was added in appropriate volume to the DNA samples and electrophoresis was carried out at 25-100 V. DNA bands were visualised via UV transillumination (λ: 302 nm) and photographed using the UVP White/UV transilluminator with camera (UVP). Ethidium bromide waste was disposed of by the University's hazardous waste system. 3.2.2. Extraction of genomic DNA from adult zebrafish Extraction of gDNA from individual stored fish for mapping and positional cloning was performed using the standard Invitrogen Purelink™ Genomic DNA Kit (Cat no. K1820- 01, Life Technologies™) according to the manufacturer’s instructions. Each fish was removed from storage at -80° and kept on ice. A 25 mg section of tail was cut from each fish using a sterile scalpel blade and processed according to the manufacturer’s protocol for Mammalian Tissue and Mouse/Rat Tail Lysate. The extracted gDNA was eluted twice in 25 µl sterile water for a total volume of 50 µl. Extraction of gDNA from individual fresh adult samples was performed using a quick genotyping kit (Bioland). Fin clips from the caudal fin were obtained and placed in 100 µl of lysis buffer. Samples were incubated at 95°C for 20 minutes. Samples were placed on ice and 10 µl of DNA stabilising solution was added. Extracting of gDNA from individual embryos and juveniles from the morphological analysis or after transgenic rescue was performed by using a sodium hydroxide (NaOH) protocol. Methanol was removed from frozen samples and evaporated on ice. Fresh samples were directly processed. 100 µl of 50 mM NaOH (Simga-Aldrich®) was added to the sample and then incubated at 95°C for 20 minutes. Samples were then immediately transferred to ice and 10 µl of 1 M Tris (pH 8; Simga-Aldrich®) was added to stabilise the sample. All gDNA samples were analysed prior to use to obtain concentration and purity data. Samples were analysed on a NanoDrop ND-1000 Spectrophotometer (Thermo Scientific) and only samples obtaining a concentration of > 50 ng/µl and a 260/280 nm purity ration of 1.7-1.9 were utilised during this project. Samples were stored at -20°C until needed and if required, diluted to a concentration of approximately 50ng/µl for downstream use. 34 3.2.3. Extraction of total RNA Total RNA (tRNA) was extracted from pooled embryos or the pooled developing pelvic regions of wild-type and pelvic finless zebrafish. Samples were removed from RNAlater (Qiagen) and tRNA was extracted with an RNeasy mini kit (Qiagen). Samples were processed according to the manufacturer’s instructions for tissue samples and disrupted with a mechanical tissue homogeniser. Concentration and quality measurements were taken on a NanoDrop ND-1000 Spectrophotometer (Thermo Scientific). Samples were then immediately processed to make cDNA.

3.2.4. Generation of cDNA Samples of extracted tRNA were processed immediately to generate cDNA with ThermoScript™ RT-PCR System for First-Strand cDNA Synthesis (Cat no. 11146-024, Invitrogen™, Life Technologies™). Samples were processed using oligo-dt primers (supplied) and 1 µg tRNA template according to the manufacturer’s protocol. For cDNA synthesis, samples were incubated at 50ºC for 50 minutes. Samples were then either stored at -20ºC or immediately processed for PCR

3.2.5. Oligodeoxyribonucleotide primers and primer design Oligonucleotide primers for PCR and DNA Sanger sequencing were synthesised by Invitrogen (Life Technologies™ or Sigma-Aldrich®). Simple sequence length polymorphism (SSLP) primers as suggested by (http://zebrafish.mgh.harvard.edu) for the genome scan were shared by our collaborators at the Laboratory for Colorectal Cancer Research at the Walter and Eliza Hall Institute of medical research, Melbourne.

Primers for the genes and sequences of interest were designed using Primer3 (Rozen & Skaletsky, 2000). Primers were selected to be of 21-28 base pairs (bp) of length, and highly selective for the target as determined by NCBI primer blast tool for the zebrafish genome (Ye et al., 2012). In addition, primers were chosen that were compatible for the generation of PCR products with minimal energy (less than -3.6kc/m) for self-dimers and pair dimers and no hairpins as indicated by Sigma-Genosys DNA calculator (http://www.sigma- genosys.com/calc/DNACalc.asp). The sequences for selected SSLP primers and genes of interest are listed in Appendix A. Genotyping reactions were performed using the z11320 SSLP primer.

35 3.2.6. Polymerase Chain Reaction Polymerase Chain Reaction (PCR) was carried out to amplify DNA according to published methods (Mullis et al., 1992). The PCR conditions utilised varied depending on the DNA template (gDNA, complementary DNA (cDNA) or plasmid DNA) and whether the reaction was for a SSLP or a gene/sequence of interest (please see Appendix A for specific PCR conditions).

Reaction mixtures were assembled with ~50 ng gDNA/cDNA template or < 1 ng plasmid DNA template. The final concentration of other reagents depended on the type of DNA polymerase utilised. For standard reactions, the final concentrations were; 250 µM of each dNTP (pre-mixed, New England Biolabs [NEB]), 2 µM for each oligonucleotide primer and 1 U of Thermopol Taq, (NEB) in 1x PCR buffer. For high-fidelity reactions, the final concentrations were, 250 µM of each dNTP (pre-mixed, NEB), 400 µM of dimethyl sulfoxide (DMSO), 0.75 µM for each oligonucleotide primer and 1 U of Phusion high-fidelity DNA polymerase (NEB) in 1x PCR buffer. Thermal cycling was performed using a variety of different conditions (please see Appendix A for specific cycling conditions).

Standard PCR products were visualised by 1-1.5% agarose/TAE buffer gel electrophoresis at 100 V for an average of 25 minutes. SSLP PCR products were analysed on 3% low electroendosmosis agarose (Sigma-Aldrich®) /TBE buffer gels after electrophoresis at 100 V for 1-3 hours.

3.2.7. Purification of DNA fragments Amplified PCR products were purified by column with a QIAquick Gel Extraction Kit (Qiagen) following manufacturer’s instructions with slight modifications. Five times the reaction volume of buffer QC was added to the PCR reaction tube and then the protocol was

strictly followed. PCR products were eluted twice into 30 µl H2O.

Gel electrophoresis was also utilised to purify DNA fragments. After separation by gel electrophoresis, products were quickly cut from the agarose on a UV illuminator under λ: 365 nm UV transillumination. Products were purified using a QIAquick Gel Extraction Kit

(Qiagen) according to manufacturers’ instructions and eluted twice into 40 µl of H20 prior to immediate downstream processing.

3.2.8. Restriction enzyme digests All digests were performed using restriction enzymes from NEB utilising the suggested buffers and reactions. For vector digestion, 5 µg of plasmid DNA was digested for 3 hours

36 with 1 Unit (U) of the required enzyme. Purified DNA fragments were digested with 1 U of the required enzymes for 1 hour. If possible, heat inactivation of the restriction enzyme was performed at the required temperature for 15 minutes.

3.2.9. Klenow blunting Klenow blunting of sticky-ended DNA fragments was performed with 1 µl of DNA polymerase I, large (Klenow) fragment (NEB). Approximately 5 µg of ligated vector was blunted in a 50 µl reaction suggested by the manufacturer. Reactions were incubated at 25ºC for 15 minutes for blunting and then subjected to 75ºC for 20 minutes for heat inactivation.

3.2.10. Phosphorylation of PCR fragments Phosphorylation of PCR fragments (150 ng) was performed utilising 0.5 µl of polynucleotide kinase (NEB) in a 9 µl reaction in ligation buffer (NEB). The reaction was incubated for 37ºC for 20 minutes for phosphorylation of insert and then incubated at 65ºC for 10 minutes for heat inactivation.

3.2.11. Vector dephosphorylation Dephosphorylation of the linearised vectors was performed to avoid self-ligation during subsequent ligation reactions. 2 µl of Antarctic phosphatase (NEB) was utilised to dephosphorylate approximate 5 µg of vector template in Antarctic phosphatase buffer (NEB). Reactions were incubated at 37ºC for 15 minutes.

3.2.12. Ligations Ligations of DNA fragments and vectors were performed according to standard protocols. The amount of insert needed was determined according the following formula, with an insert vector molar ration of 3:1 ng of vector × kb of insert ng of insert = × insert/vector molar ratio kb of vector However, typically 150 ng of insert was used in a 20 µl reaction with 50-75 ng of vector. If multiple inserts required ligation, 100 ng of each insert was utilised. The ligation was performed with 1µl T4 DNA ligase (NEB) in ligation buffer (NEB) in a 20 µl reaction according to the manufacturer’s instructions. Reactions were incubated overnight at 16ºC.

37 3.2.13. TOPO® cloning TOPO® cloning (Invitrogen™) uses TA cloning® combined with the enzymatic activity of Topoisomerase I to efficiently clone PCR products. Standard techniques for TOPO® cloning can be found in the manual for TOPO® TA Cloning® Kit for Sequencing. Listed below is the typical protocol used during this project for TOPO® cloning. For this project, DNA fragments were cloned into pCR®4-TOPO® vector (Invitrogen™, Life Technologies™).

DNA fragments were generated by amplification with gene specific primers by PCR with standard Taq polymerase in a 50 µl reaction in order to obtain DNA fragments with Adenine overhangs (see 3.2.6). Reactions were optimised to eliminate non-specific products and the need for gel purification. PCR products were purified utilising the on-column method described in 3.2.7.

Purified PCR products were immediately used for TOPO® cloning as Adenine overhangs are very fragile and can be removed by freeze/thaw cycles. 2 µl of purified PCR product was used in the standard TOPO® cloning reaction (according to the manufacturer’s instructions) and allowed to incubate for a minimum of 5 minutes before proceeding to transformation (see 3.2.15).

3.2.14. MultiSite Gateway® cloning MultiSite Gateway® three-fragment vector construction cloning (Invitrogen™) utilises uses site-specific recombinational cloning. This allows simultaneous cloning of multiple DNA fragments in a defined order and orientation based on MultiSite Gateway® technology (Hartley et al., 2000).

Throughout this project, expression clones were created using the MultiSite Gateway® LR recombination reaction according to the manufacturer’s instructions from version D (2007) of the MultiSite Gateway® Three-Fragment Vector Construction Kit manual (Invitrogen™, Life Technologies™). Various entry clones (an attL4 and attR1-containing entry clone [p5’E], an attL1 and attL2-containing entry clone [pME] and an attR2 and attL3- containing entry clone [p3’E]) were obtained and combined into a pDestTol2 destination vector (a derivative of the pDEST™R4-R3 vector (Invitrogen™, Life Technologies™) (Kwan et al., 2007). The MultiSite Gateway® LR recombination reaction was performed according to the manufacturer’s instructions using 60 ng of each vector. Reactions were allowed to incubate

38 for 20 hours at 25ºC. Reactions were stopped with 10 minutes Protinase K (Invitrogen™, Life Technologies™) digestion at 37ºC before proceeding to transformation (see 3.2.15).

3.2.15. Transformations Plasmids were transformed into competent bacteria by the heat-shock method as follows; OneShot® Top10 chemically competent E.coli (Invitrogen™, Life Technologies™) were thawed on ice and incubated on ice for a minimum of 5 minutes with approximately 2 µl of plasmid DNA. The cells were then heat-shocked at 42ºC for exactly 30 seconds and immediately returned to ice. 250 µl of room temperature S.O.C. medium was added to the cells which were then allowed to recover for 1 hour at 37ºC at 200 rpm. After recovery, 30- 250 µl of cells were spread over the surface of an LB agar plate containing the appropriate antibiotic. Plates were inverted and incubated at 37ºC overnight.

3.2.16. PCR screening of colonies Colonies of interest were screening by PCR amplification to determine the presence and direction of the inserts. Colonies were picked from the growth plate by a barrier tip which was then used to inoculate a PCR microtube and a 5 ml aliquot of LB. Standard PCR amplification was performed with a gene specific primer and either a forwards or reverse vector specific primer. PCR products were visualised by gel electrophoresis. Colonies which harboured the DNA fragment of interest in the correct orientation were purified (see 3.2.17) and stored at -20 ºC until needed.

3.2.17. Purification of plasmid DNA from E. coli bacteria Preparation of plasmid DNA from E. coli bacteria was performed using the standard Jetquick plasmid purification spin kit protocol according to the manufacturer’s instructions (Genomed). A few modifications were made as follows: the starter culture was inoculated into 5 ml of Luria-Bertani broth (LB) containing a selective antibiotic and incubated for an average of 20 hours at 37°C at 200 rpm. After incubation, 0.5 ml of culture was used for a glycerol stock, while the remaining culture was centrifuged at 2000 rpm, at room temperature for 10 minutes. The pellet was then processed according to the manufacturer’s instructions. The final plasmid DNA was eluted twice into 30 µl sterile water. The concentration was determined on the NanoDrop ND-1000 Spectrophotometer and 5µl of the preparation was assessed by 1.5% agarose:TAE buffer gel electrophoresis. The samples were stored at -20ºC until needed.

39 3.2.18. Generation of DIG labelled cRNA probes 3.2.18.1. Generation of templates Digoxigenin (DIG) labelled complementary RNA (cRNA) antisense probes for the genes paired-like homeodomain transcription factor 1 (pitx1), t-box transcription factor 4 (tbx4), tbx5, gf10a, specificity protein 8 (sp8), fibroblast growth factor 8a (fgf8a) and sonic hedgehog (shh) were synthesised for in situ hybridisation analysis from plasmids containing fragments of the gene of interest.

The plasmids containing fragments of fgf10a and sp8 (Nagayoshi et al., 2008), fgf8a (Komisarczuk et al., 2009), tbx4 (Tamura et al., 1999) tbx5 (Garrity et al., 2002) and shh (Currie and Ingham, 1996) were kindly donated for use in this project. The plasmid containing a fragment of pitx1 was generated in the lab with gene specific primers designed to the published cDNA sequence. The amplified pitx1 PCR product was directly TOPO™ cloned into the pCR®4-TOPO® vector (see section 3.2.13).

Once the plasmid containing the fragment gene of interest was obtained, the plasmids were linearised by restriction digestion and transcribed in-vitro to create sense and antisense cRNA probes.

3.2.18.2. DIG labelled cRNA probe synthesis As each plasmid was different, please see Appendix A for a list of restriction enzymes and RNA polymerase used.

To generate DIG labelled cRNA sense and antisense probes, two restriction digest reactions were set up in parallel to linearise the plasmid. Approximately 5 µg of plasmid was linearised by digestion for 1 hour with 1 U of the necessary enzyme. 5 µl of the reaction was checked for completion of digestions by 1% agarose: TAE gel electrophoresis. Reactions were purified via the on-column method described in (3.2.7). Linearised plasmids were eluted twice into 30 µl milliQ water.

DIG labelled cRNA sense and antisense probes were created by transcribing the fragment of interest from 1 µg linearised plasmid with the appropriate RNA polymerase (T3, T7 or SP6). Reactions were assembled according to the manufacturer’s instructions (Roche).

Reactions were incubated at 37ºC for T3 and T7 RNA polymerase or 40 ºC for SP6 RNA polymerase for 2 hours or overnight. A 2 µl aliquot was subjected to 1% agarose:TAE gel electrophoresis to ensure the presence of a cRNA transcript. 80 µl of hybridisation buffer

40 was added to the remaining reaction and the cRNA probe was stored at -20ºC until needed for in-situ hybridisation.

3.2.19. Generation of Transposase mRNA Transposase messenger RNA (mRNA) was generated from the PCS2FA-transposase plasmid (Kwan et al., 2007). Approximately 5 µg of the plasmid was linearised by digestion with NotI restriction enzyme. The linearised plasmid template was purified via the on- column method described in 3.2.7. The linearised plasmid template was eluted twice into 30

µl H2O.

Transposase mRNA was generated using a mMessage machine SP6 kit (Ambion®, Life Technologies™). Approximately 1 µg of linear plasmid template was utilised in the reaction which was assembled according to the manufacturer’s instructions. The reaction was incubated for two hours. The mRNA was then purified using an RNeasy mini kit (Qiagen) following the manufacturer’s instructions for RNA purification. Aliquots of Transposase mRNA were stored at -80ºC until needed.

3.2.20. Sanger sequencing Both PCR products and plasmids were sequenced during this project. In general, the sense primer used in the original reaction was used for the direct sequencing of PCR products. The primers used to sequence DNA fragments contained in plasmids varied and depended on the plasmid. The primers used to sequence DNA fragments in plasmids were the standard M13 forward, M13 reverse or SP6 transcription start site primers.

Samples were sequenced at the Sydney Node of the Australian Genome Research Facility (AGRF). Samples were sequenced by the Sanger method using Applied Biosystems 3730 and 3730xl sequencers. These were automated platforms which utilised Big Dye Terminator (BDT) chemistry version 3.1 (Applied Biosystems) under standardised cycling PCR conditions. The raw chromatogram trace file (designated at *.ab1 files) were analysed using Geneious® 6.0.3 software (Biomatters).

41 3.3. Staining A variety of staining techniques were used during this project. Many of the standard methods were adapted for use in juvenile and adult zebrafish.

3.3.1. Bone and cartilage staining The protocol used for double bone and cartilage staining was adapted from Walker and Kimmel, 2007 with small changes. All steps were performed on a gently rocking platform. Samples were retrieved from storage in 50% ethanol and stained in 0.04% Alcian Blue

(Sigma-Aldrich®)/ 0.01% Alizarin Red (Sigma-Aldrich®)/10 mM MgCl2 (Sigma-Aldrich®)/ 80% ethanol for 5 days. Samples were then rinsed with 80% ethanol/10 mM MgCl2/water overnight, with 50% ethanol/water for 5 minutes and in 25% ethanol/water for 5 minutes. For digestion the samples were washed in 0.5 M sodium borate (Sigma-Aldrich®) for 5 minutes and digested in 0.5 M sodium borate with 0.25% Trypsin (Invitrogen™, Life Technologies™) for 6 hours. To bleach samples, the samples were wash in 2% potassium hydroxide (Sigma-Aldrich®) for 10 minutes and bleached in 3% hydrogen peroxide (Sigma- Aldrich®) / 0.5% potassium hydroxide for 10 minutes with the lids open. To prepare for imaging samples were rinsed in 25% glycerol/ 0.1% potassium hydroxide for 1 hour, in 50% glycerol/ 0.1% potassium hydroxide overnight. Finally, the solution was replaced with new 50% glycerol/ 0.1% potassium hydroxide and samples were stored 4°C until needed.

3.3.2. Acridine orange staining for confocal microscopy Samples for acridine orange staining were fixed and stored according to protocols described in 3.1.5.1. Samples were rehydrated from methanol into PBS. Samples were then stained in a solution of 5 µg/ml acridine orange (Sigma-Aldrich®) in PBS for 15 minutes at 37°C. Samples were then washed twice in PBS prior to confocal microscopy.

3.3.3. Haematoxylin and eosin (H & E) staining Paraffin blocks containing samples were cut into 10-15 µM sections using a microtome. Sections were dried overnight at room temperature before further processing. Staining was employed according to standard protocols described in Kiernan, 1999. After deparaffinisation, slides were immersed in haematoxylin (Sigma-Aldrich®) for 1 minute. In addition, slides were dipped twice in PBS to ‘blue’ and stained in Eosin (Sigma-Aldrich®) for 30 seconds. Samples were mounted and coverslipped with DePex mounting medium (Sigma-Aldrich®), dried overnight and stord at room temperature until needed.

42 3.3.4. In situ hybridisation The following procedure required subjecting the samples to many changes of solutions. In order to facilitate the process, samples were processed in Costar Netwells (Crown Scientific) and transferred between solutions. Unless otherwise stated, the procedures listed were carried out in 3 ml of solution.

As in situ hybridisation in juvenile and adult fish can be variable, all cRNA probes were first tested on embryonic stages to determine the validity of the probe by the observation of the expected gene expression pattern.

Day 1:

REHYDRATION

Samples that had been fixed and stored for in-situ hybridisation were rehydrated into PBS through a methanol series in reverse.

1. 15 min in 80% methanol: 20% PBS. 2. 15 min in 60% methanol: 40% PBS. 3. 15 min in 40% methanol: 60% PBS. 4. 15 min in 20% methanol: 80% PBS. 5. 2 x 5 min in PBST.

At this stage, the pelvic regions were dissected posterior to the pectoral fins and anterior to the cloacae and the intestines were removed to prevent trapping of the probe in larger samples. In addition, as the fin tissue was so fragile, typical proteinase K digestion was omitted from the protocol.

HYBRIDISATION

For the next steps, samples were processed in probe specific batches in 1.5 ml microfuge tubes. Care was taken so as to not allow the temperature of the samples and solutions to drop after the addition of the probes.

1. Replace PBST with 1 ml pre-warmed hybridisation buffer. 2. Incubate for 1 hour at 65ºC. 3. Replace hybridisation buffer with fresh pre-warmed hybridisation buffer with probe at 1:200 (probe solution to hybridisation buffer). 4. Incubate at 65ºC for 24-72 hours in rocking hybridisation oven.

43 Day 2

PROBE REMOVAL

After hybridisation, the probe solutions were removed at stored for re-use at -20ºC. The post- hybridisation washes were performed with pre-warmed solutions in a hybridisation oven on a rocking platform unless otherwise stated. Special care was taken so that the temperature of the samples and solutions did not drop during solution changes.

1. Wash in 2 x SSC at 65°C for 15min. 2. 2x (wash in 2 x SSC + 0.1% CHAPS (Sigma-Aldrich®) at 65°C for 15min). 3. 2x (wash in 0.2 x SSC + 0.1% CHAPS at 65°C for 15min).

ANTIBODY

All steps were performed in Netwell racks on a gently rocking platform at room temperature.

1. Wash in Tris buffer (pH 7.5) for 30 minutes at room temperature on rocking platform. 2. Block in blocking solution for 1 hour at 4ºC. 3. Replace with fresh blocking solution. 4. Add DIG antibody (Roche) at 1:5000 (DIG antibody to blocking solution). 5. Incubate overnight at 4ºC.

Day 3

COLOUR DEVELOPMENT

All steps were performed in Netwell racks on a gently rocking platform at room temperature in the dark unless otherwise stated.

1. Wash in Tris buffer (pH 7.5) 4 x 30 minutes. 2. Wash in Tris buffer (pH 9) for 30 minutes. 3. Carefully transfer samples from Netwell sieves into wells containing 3 ml fresh Tris buffer (pH 9.5). 4. Add 30 µl NBT/BCIP (Roche) stock solution into each well containing 3 ml fresh Tris buffer (pH 9). 5. Develop in the dark at 4ºC whilst periodically monitoring colour development.

44 Colour development was monitored under the steromicroscope. Development time was variable depending on the probe used. Some probes required only 2 hours at 4ºC for development while other required 3-4 days at room temperature. Samples were post- fixed in 4% PFA in PBS when desired level of staining was achieved.

45 3.4. Microscopy 3.4.1. Light microscopy Bright field illumination microscopy was performed on a Leica M165FC stereo dissection microscope (Leica) using a ProRes CJ cool camera (Jenoptic). Any samples imaged before the development of pelvic fins was observed were genotyped after imaging. The majority of samples were imaged in a 3% solution of methyl cellulose (Sigma-Aldrich®) in PBS. Skeletal staining samples were imaged in an 80% glycerol (Sigma-Aldrich®) solution.

Bright field illumination microscopy of histological slides was performed on a Leica Manual Inverted Microscope (DMI 3000 B) using a ProRes CJ cool camera (Jenoptic).

3.4.2. Florescence microscopy Florescence microscopy was performed on a Leica M165FC stereo dissection microscope (Leica) using a ProRes CJ cool camera (Jenoptic). UV illumination was provided by an external light source for excitation (EL6000, Leica) using a GFP filter cube (Leica).

3.4.3. Confocal microscopy Images were taken on a Zeiss LSM 510 Meta confocal microscope (Carl Zeiss) using the 488 nm argon laser and a 505 nm long pass emission filter. Samples were embedded in a solution of 1.5% agarose in water on a circular glass bottom culture dish.

3.4.4. Scanning electron microscopy Samples were imaged in high vacuum mode on a XL30 CP scanning electron microscope (Philips) at 5 to 15 kV in the Australian Centre for Microscopy and Microanalysis of Sydney University under the expert guidance of Dr Errin Johnson and Dr Ian Kaplin.

3.5. Statistical methods Quantitative data presented in this body of work derives from at least three independent experiments or individuals. Descriptive statistics are described as mean ± standard error of difference (SE of difference) of data for ‘n’ individuals or ‘n’ independent experiments. Statistical analysis, including unpaired two-tailed students t-tests, two way ANOVA and Pearson’s Chi-squared tests were conducted utilising GraphPad Prism 6.0 (GraphPad Software). A value of p < 0.05 was used to determine statistically significant difference. Throughout this body of work, the asterisks refer to the following significant vaules: p < 0.05 (*), p < 0.01 (**) and p < 0.001 (***).

46 3.6. Sequence analysis and alignments Geneious® 6.0.3 software (Biomatters) was utilised to perform sequence analysis and alignments. Nucleotide sequence alignments were performed in the software either using the ‘Map to reference’ or the ‘Pairwise/Multiple Geneious Alignment’ functions with the manufacturer’s set conditions. Protein sequence alignments were performed utilising the ‘Pairwise/Multiple Geneious Alignment’ function with the manufacturer’s set conditions.

47 4. Morphological characterisation of pelvic finless zebrafish 4.1. Introduction The absence of pelvic fins in the pelvic finless strain of zebrafish (Don et al., 2011, Appendix B) is particularly attractive due to its potential to shed light on the previously undescribed genetic mechanisms of pelvic fin development. The aim of this chapter is to extend the morphological characterisation of pelvic finless zebrafish through a comparison to wild-type siblings utilising a combination of microscopy and histology. The investigation of pelvic fin development in pelvic finless zebrafish was performed in conjunction with gene expression analysis, genetic mapping and deep sequencing in order to identify the genetic basis underlying the loss of pelvic fins. This approach was adopted with the aim of gaining a more complete understanding of the developmental mechanism behind pelvic fin development and loss.

In this chapter, pelvic finless zebrafish were compared in terms of length, overall morphology and pelvic fin development to their wild-type siblings. Embryonic and larval length and morphology was compared in 140 individuals. As the pelvic finless phenotype does not become apparent until around 21 dpf, 20 individuals were sampled and genotyped (see 3.2.2 and 3.2.5) at each time point, which on average, gave 4 pelvic finless individuals for each time point. The measurements of length were then subjected to two-way ANOVA testing to determine significance (see 3.5). Pelvic fin development was examined in adult zebrafish by light microscopy (see 3.4.1), bone and cartilage staining (see 3.1.5.3 and 3.3.1), histology (see 3.1.5.2 and 3.3.3) and scanning electron microscopy (see 3.1.5.4 and 3.4.4).

48 4.2. Results 4.2.1. Embryonic morphology of pelvic finless zebrafish Bright field illumination microscopy of embryos demonstrated that both pelvic finless and wild-type siblings developed normally and at the same rate. A number of measurements of embryonic length (EL; longest linear direction) identified no significant difference between pelvic finless embryos and wild-type siblings (Figure 4.1, two-way ANOVA, p < 0.05). At 24 hpf, pelvic finless embryos displayed an average EL of 1.80 mm (n=4) whereas wild-type siblings had achieved an EL of 1.87 mm (n=4). Both pelvic finless zebrafish and wild-type siblings had reached the prim-5 stage of the pharyngula period (Kimmel et al., 1995), and accordingly displayed early pigmentation in retina and skin, possessed the beginnings of the median fin fold and had a heartbeat (Figure 4.2). Pelvic finless and wild-type sibling embryos continued normal development and by 48 hpf had straightened along the body axis, possessed pigmented and had developed the early pectoral fins of the long-pec hatching stage (Kimmel et al., 1995) (Figure 4.2). At this stage, pelvic finless embryos had reached an EL of 2.3 mm (n=5) and wild-type siblings had an EL of 2.3 mm (n=5). At 72 hpf, the majority of pelvic finless zebrafish and wild-type siblings had hatched. Both pelvic finless zebrafish (n=3) and wild-type siblings (n=3) had reached an EL of 2.9 mm and had reached the normal developmental milestones of blade shaped pectoral fins and a fully developed median fin fold of the protruding mouth stage (Kimmel et al., 1995) (Figure 4.2).

49 Figure 4.1. Wild-type vs. pelvic finless total length A graph demonstrating the increase in size of pelvic finless zebrafish and wild-type siblings from data presented in the Appendix C. Overall, pelvic finless zebrafish and wild-type siblings show no significant difference in embryonic or total length during development. Error bars represent the standard deviation from the mean.

50 Figure 4.2. Images of wild-type and pelvic finless embryos Bright field illumination microscopy images of lateral views of wild-type siblings and pelvic finless embryos at 24hpf, 48hpf and 72hpf. The images demonstrate that both pelvic finless zebrafish and wild-type siblings develop normally and at the same rate. Scale bar: 1mm. Anterior to left in all panels.

51 4.2.2. Larval morphology of pelvic finless zebrafish At 7 dpf, both pelvic finless larvae and wild-type siblings had fully inflated swim bladders and had reabsorbed the majority of the yolk sack (Figure 4.3). Pelvic finless zebrafish had reached a total length (TL; tip of the snout to the tip of the caudal fin) of 3.0 mm (n=4), whereas wild-type siblings had a TL of 3.3 mm (n=4) (Figure 4.3). By this stage both larvae and wild-type siblings demonstrated the active swimming and the prey seeking behaviour of the early larval period. By 14 dpf, both the pelvic finless larval and wild-type siblings possessed flattened caudal fins with developing lepidotrichia. Pelvic finless zebrafish had reached a TL of 4.8 mm (n=4) whereas wild-type siblings had a TL of 4.9 mm (n=4) (Figure 4.3).

The first signs of pelvic fins were observed around 21 dpf, although the exact timing was highly variable, ranging from 21 dpf to 28 dpf. However, there was no significant difference in the TL at which pelvic finless larvae and wild-type siblings began to develop pelvic fins, (pelvic finless TL=5.3 mm, n=4, wild-type siblings TL=5.1 mm, n=4, p=0.59). Additionally, at 21 dpf, the dorsal and ventral fins had begun to protrude from the median fin fold and had developed lepidotrichia, however, the median fin fold still remained in both genotypes (Figure 4.3). By 28 dpf, in both pelvic finless larvae and wild-type siblings the majority of the median fin fold had regressed and the pectoral, dorsal, ventral and caudal fins of these fish were fully developed (Figure 4.3). Although displaying no significant difference in TL (pelvic finless TL=7.9 mm, n=4, wild-type siblings TL=7.7 mm, n=4, p=0.51), the pelvic fins of the wild-type siblings could be observed whereas pelvic fins were notably absent in the pelvic finless larvae (Figure 4.3).

52 Figure 4.3. Images of wild-type and pelvic finless larvae Bright field illumination microscopy images of lateral views of wild-type siblings and pelvic finless larvae at 7dpf, 14dpf, 21dpf and 28dpf. The images demonstrate that both pelvic finless zebrafish and wild-type siblings develop normally and at the same rate apart from the absence of pelvic fins in pelvic finless zebrafish. Scale bar: 1mm. Anterior to left in all panels.

53 4.2.3. Adult morphology of pelvic finless zebrafish The skeletal morphology of the paired and unpaired fins of adult pelvic finless zebrafish was compared to wild-type siblings. Bone and cartilage stained samples were observed under bright field illumination microscopy to record the number of fin rays of the pelvic, pectoral, dorsal, ventral and caudal fins of pelvic finless zebrafish (n=4) and wild-type siblings (n=4) (Figure 4.4 and Figure 4.5). Both pelvic finless zebrafish and wild-type siblings demonstrated the presence of the paired pectoral fins, and unpaired dorsal, ventral and caudal fins. There was no significant difference in the number of fins rays between pelvic finless zebrafish and wild-type siblings in pectoral (wt=10.25; pFL=10.25), dorsal (wt=9.25; pFL=9.5), ventral (wt=15.75; pFL=14.75) and caudal (wt=28.75; pFL=29.5) fins (unpaired two-tailed Student’s t-test, p < 0.05). However, all pelvic finless zebrafish examined lacked all skeletal pelvic fin elements (wt=8; pFL=0).

54 Figure 4.4. Images of all fins in pelvic finless zebrafish and wild-type siblings

Bright field illumination microscopy images of bone and cartilage stained fins in wild-type siblings and pelvic finless zebrafish. The pectoral, dorsal, ventral and caudal fins of pelvic finless zebrafish and wild-type siblings are similar in terms of fin ray number. Examination of the pelvic fins of pelvic finless zebrafish and wild-type siblings demonstrated that the pelvic finless zebrafish did not possess pelvic fins or the associated fin rays. Anterior to left in all panels. Scale bar 1 mm.

55 Figure 4.5. Wild-type vs. pelvic finless fin ray number

A graph demonstrating the number of fin rays present in all fins between pelvic finless zebrafish and wild-type siblings from data presented in Appendix C. There was no significant difference between the number of fin rays between pelvic finless zebrafish and wild-type siblings for the pectoral, dorsal, ventral and caudal fins..

56 4.2.4. Pelvic finless zebrafish display abnormal pelvic fin development The pelvic fins of wild-type s iblings first became visible at a posterior position along the flank, anterior to the , on either side of the median fin fold and could be seen as a distinct paddle-like shape protruding from the flank (Figure 4.6, 21dpf). Conversely, at the same stages, pelvic finless zebrafish had merely developed two bulge-like structures in the pelvic fin regions (Figure 4.7, 21dpf). This bulge-like structure of the pelvic finless zebrafish was notably smaller than the distinct paddle-like shaped pelvic fin bud which had developed in the wild-type siblings (Figure 4.7, 21dpf). By 32dpf, the pelvic fins of wild- type zebrafish displayed adult morphology with lepidotrichia (fin rays) (Figure 4.6, 32dpf), whereas the bulge-like structures in the pelvic regions of pelvic finless zebrafish failed to increase in size or structure, did not develop lepidotrichia (Figure 4.7, 28dpf).

57 Figure 4.6. Development of the pelvic fins in wild-type zebrafish Lateral views of a wild-type sibling developing pelvic fins. The pelvic fin buds of wild-type zebrafish are first visible around 21 dpf on either side of the flank, anterior to the cloaca. Between 21 dpf and 28 dpf, the pelvic fins of wild-type fish continue to develop into a distinct paddle like shape and elongate. At 32 dpf, the pelvic fins of wild-type fish take on the adult pelvic fin shape with visible fin rays. The asterisks marks the position of the cloaca and the arrowheads mark the position of the fin rays. Scale bar: 0.1mm . Anterior to left in all panels.

58 Figure 4.7. Comparison of pelvic fin development in pelvic finless zebrafish and wild-type siblings Lateral views comparing the pelvic fin developing in pelvic finless zebrafish and wild-type siblings. At 21 dpf, a small bulge can be observed in the pelvic fin regions of both pelvic finless zebrafish and wild-type siblings. At 28 dpf, the pelvic fins of the wild-type siblings have grown and elongated into a paddle-like shape, whereas the pelvic fins of pelvic finless zebrafish remain as a small bulge. Scale bar: 0.1mm. Anterior to left in all panels.

59 4.2.5. Pelvic fins of pelvic finless zebrafish lack an apical ectodermal ridge Around 21 dpf, the pelvic fin buds of wild-type fish were observed as a distinct narrow band of cells, distally capped by an apical ectodermal thickening, protruding from the flank of the fish (Figure 4.8). In contrast, at the same developmental stage, the pelvic fin buds of pelvic finless fish were observed as mesenchymal bulges with no evidence of any distal ectodermal structure (Figure 4.8). Observations at 28 dpf revealed that pelvic fins of wild- type zebrafish (n=3) continued normal development, whereas the pelvic fin buds of pelvic finless zebrafish (n=3) never formed an apical fold or its derivative, the fin fold, and did not increase in size (Figure 4.8). By 5 wpf, the pelvic fins of wild-type zebrafish were fully formed, whereas the pelvic fin buds of pelvic finless did not develop further (data not shown).

60 Figure 4.8. Scanning electron microscopy images of developing pelvic fins in wild-type and pelvic finless zebrafish Scanning electron microscopy images of the developing pelvic regions of wild-type and pelvic finless zebrafish. At 21 dpf, the pelvic fins of wild-type zebrafish have developed an apical ectodermal thickening, whereas the pelvic bulges of pelvic finless zebrafish lack overlying ectodermal structure. By 28 dpf, the pelvic fins of wild- type zebrafish have elongated and developed a paddle-like shape whereas the structure in the pelvic regions of pelvic finless zebrafish remains as a mesenchymal bulge. Black arrowheads indicate the apical ectodermal thickening, whereas white arrowheads demonstrate the absence of this structure. Scale bar: 100µm. Anterior to left.

61 The presence of an apical ectodermal ridge was also investigated by histological examination of pelvic finless zebrafish and wild-type siblings. This analysis confirmed that the pelvic fin buds of wild-type fish progressed to form an apical fold, with two opposing layers of ectodermal cells of the basal stratum at the distal tip of the fin. Subsequently, in the pelvic fins of wild-type zebrafish, a fin fold consisting of the separating layers of the basal stratum infiltrated by migrating mesenchyme emerges at the distal tip of the fin. In contrast, the pelvic fin buds of pelvic finless fish remained as mesenchymal bulges, 3-4 layers thick protruding from the ventrolateral body wall in the prospective pelvic fin region. The bulges retained a small population of mesenchymal cells which possessed little distal ectodermal structure (Figure 4.9).

62 Figure 4.9. Sections of wild-type and pelvic finless pelvic regions Histological sections of the developing pelvic fin regions of wild-type and pelvic finless zebrafish. At 25 dpf, the apical fold (circled) of the developing pelvic fin is present in wild-type individuals and absent in pelvic finless individuals (indicated by arrow). At 28 dpf, the fin fold (circled) of the developing pelvic fin is present in wild-type individuals and absent in pelvic finless individuals (indicated by arrow). In addition, the pelvic fins of wild-type individuals posses proliferating mesenchymal cells, whereas the pelvic bulges of pelvic finless zebrafish maintain a small population of mesenchymal cells. White asterisks mark the presence of the mesenchymal cells. Scale bar 50 µm.

63 4.2.6. Incomplete penetrance of the pelvic finless phenotype During examinations of pelvic finless zebrafish it was noted that the pelvic finless phenotype exhibits incomplete penetrance. A partial phenotype was observed in one in every one hundred pelvic finless fish around 32 dfp (n=8⁄800). These fish displayed a range of partial phenotypes from one, either abnormal or complete pelvic fin on either the right or left side, or two abnormal pelvic fins (Figure 4.10). Although the majority of adult pelvic finless zebrafish demonstrated a total absence of pelvic fins (n=396/400, Figure 4.10 B'), a partial phenotype was occasionally observed in adult pelvic finless zebrafish on the golden background (n=4⁄400) (Figure 4.10). Observations of pelvic finless zebrafish on the WIK background revealed a higher level of incomplete penetrance with 3 out of 41 fish displaying partial pelvic fin development. Of these fish, one individual displayed one normal pelvic fin with 8 lepidotrichia on the left pelvic fin (Figure 4.10 C'), while another individual developed one abnormal pelvic fin on the right hand side consisting of 2 lepidotrichia. The third individual displayed two abnormal pelvic fins consisting of 1 lepidotrichia on the left pelvic fin and 2 lepidotrichia on the right pelvic fin (Figure 4.10 D').

64 Figure 4.10. Incomplete penetrance of the pelvic finless phenotype Confocal microscopy and bright field illumination microscopy images of wild-type and pelvic finless zebrafish during pelvic fin development and adult fish stages. A, A') Wild-type zebrafish develop two paired pelvic fins with 7-9 lepidotrichia. B, B') The majority of pelvic finless zebrafish do not develop pelvic fins (n=1,188/1200). C, C') An individual developed a complete pelvic fin on the left side with 8 lepidotrichia. D, D') An individual developed two abnormal pelvic fins consisting of 1 lepidotrichia on the left pelvic fin and 2 lepidotrichia on the right pelvic fin. White arrows indicate the position of the developing pelvic fins. Anterior to left in all panels. Scale bar 1 mm. 65 4.3. Discussion During this project, the morphology of pelvic fin development in zebrafish was examined at various developmental stages and compared to wild-type siblings in order to characterise the pelvic finless phenotype. The developmental morphology of pelvic finless zebrafish and wild-type siblings was compared from embryonic stages through to adulthood by a combination of microscopy, histology and bone and cartilage staining, with a particular emphasis on pelvic fin development.

The main feature associated with the pelvic finless phenotype was the loss of pelvic fins. Bright field illumination microscopy revealed that pelvic finless zebrafish displayed normal and similar embryonic and larval development and morphology to wild-type siblings. The morphological analysis of adult zebrafish by bone and cartilage staining revealed that no other fins were affected in terms of size, position or number of fin rays and that pelvic finless zebrafish possessed a specific pelvic fin phenotype.

Further investigations revealed that early pelvic fin development is initiated in pelvic finless zebrafish. This was demonstrated by the presence of mesenchymal bulges which formed on either side of the median fin fold in the prospective pelvic fin regions of pelvic finless zebrafish. However, the differences between wild-type pelvic fin development and that of pelvic finless zebrafish became evident during later stages of pelvic fin development. In contrast to wild-type zebrafish, pelvic bulges of pelvic finless zebrafish never developed a morphologically distinguishable apical ectodermal thickening. This narrow band of cells, that is present in wild-type zebrafish and absent in pelvic finless zebrafish, has been previously observed in both trout (Oncorhynchus) and killifish (Cyprinodon) pelvic fins where it was termed the apical ectodermal pseudo-ridge and it has been presumed to play the same role as the apical ectodermal ridge (AER) does in tetrapods (Geraudie, 1978; Wood, 1982; Grandel and Schulte-Merker, 1998). It is feasible that the lack of a ridge-like structure and subsequent apical fold may be the cause of the failure of pelvic fin outgrowth, as the apical ectodermal ridge is essential in pectoral fin and tetrapod limb outgrowth (as will be discussed in chapters 5 and 9).

In addition, some pelvic finless individuals did develop one or two, normal or abnormal pelvic fins. This raises the possibility that the genomic lesion underlying the pelvic finless phenotype is not fully penetrant. This could be due to an hypomorphic allele or that another gene may be partially compensating during the development of the pelvic fins in these individuals (as has been suggested for paired-like homeodomain transcription factor 2 (pitx2) 66 in threespine stickleback fish (Gasterosteus aculeatus) which lack pitx1 activity (Cole et al., 2003; Shapiro et al., 2004)).

The arrested development of the pelvic fins of pelvic finless zebrafish at the mesenchymal bulge stage and the absence of a morphologically distinct apical ectodermal ridge suggests that two distinct possibilities that could underlie the loss of pelvic fins in these fish. One possibility is that pelvic fin development is arrested at a stage prior to the induction of the apical ectodermal thickening and thus pelvic fins do not develop beyond mesenchymal bulges. The other possibility is that the process which results in the establishment and maintenance of the apical ectodermal thickening is disrupted and therefore the pelvic fins cannot continue to develop.

However, the pectoral, dorsal, ventral and caudal fins of pelvic finless zebrafish all develop normally which suggests that the genomic lesion underlying the pelvic finless phenotype lies in gene or regulatory element that is specific to early pelvic fin development. An example of this is the genomic lesion in the pitx1 pelvic region regulatory element, Pel, which underlies the loss of pelvic fins in some stickleback fish populations (Cole et al., 2003; Shapiro et al., 2004; Chan et al., 2010). It is also possible that the genomic lesion underlying the pelvic finless phenotype lies in a pelvic fin specific pathway. As the pelvic fins of zebrafish develop approximately three weeks after the development of the pectoral fins, it is possible that different pathways are needed for the late development of pelvic fins. Studies of retinoic acid deficient mice have shown that retinoic acid is not necessary for hindlimb development (Duester, 2008; Zhao et al., 2009), but retinoic acid signalling has been shown to be necessary for limb and fin bud induction in the forelimbs of mice and the pectoral fins of zebrafish (Begemann et al., 2001; Gibert et al., 2006; Grandel and Brand, 2011; Grandel et al., 2002; Mercader et al., 2006; Sandell et al., 2007; Zhao et al., 2009; Cunningham et al., 2011; Cunningham et al., 2013). Therefore this implies that different mechanisms may be at work during the development of forelimbs and hindlimb. For example, the genes Pitx1 and Tbx4 are expressed during the early stages of hindlimb and pelvic fin development, but are not expressed during the early stages of forelimb development, whereas Tbx5 is expressed during forelimb (and not hindlimb) development. It is possible that different signalling pathways are necessary for the induction of Pitx1 and Tbx4 in the hindlimbs of vertebrates (Agarwal et al., 2003; Ahn et al., 2002; Cole et al., 2003; Garrity et al., 2002, Logan and Tabin, 1999; Lanctôt et al., 1999; Naiche and Papaioannou, 2003; Marcil et al., 2003; Rallis

67 et al., 2003; Ruvinsky et al., 2000; Shapiro et al., 2004; Szeto et al., 1999; Takeuchi et al., 2003; Tamura et al., 1999).

In order to explore why pelvic finless zebrafish do not establish a morphological distinct apical ectodermal thickening and where in the genetic pathway of pelvic fin development the pelvic finless zebrafish is defective, the expression of a multitude of key hindlimb development genes was examined by in situ hybridisation and is presented in Chapter 5.

68 5. Analysis of pelvic fin gene expression 5.1. Introduction Although adult pelvic finless zebrafish do not have pelvic fins, they do develop the beginnings of a pelvic fin bud which appears as a mesenchymal bulge (Chapter 4). These pelvic fin buds do not develop beyond mesenchymal bulges and appear to lack a morphologically distinguishable apical ectodermal thickening. However, the exact point in the genetic cascade that leads to pelvic fin development at which the pelvic fin buds arrest development remains unknown. Determining the genetic point at which fin development is disrupted could yield insights into the genetic cause underlying the pelvic finless phenotype. The aim of this chapter is to examine the expression of genes with well established roles in limb development in order to better characterise pelvic fin development in both wild-type and pelvic finless zebrafish. Therefore the expression of the genes pitx1, tbx4, tbx5, fgf10a, sp8, fgf8a and shh were each examined by in situ hybridisation (Figure 5.1). The expression patterns of these genes were examined by in situ hybridisation (see 3.3.4) at the first visible signs of pelvic fin development at 21 dpf and during later pelvic fin development at 28 dpf in a minimum of 20 pelvic finless or wild-type zebrafish at each time point. The pecotral and pelvic fins were dissected prior to imaging (see 3.4.1).

69 Figure 5.1. The hierarchy and interactions of genes examined by in situ hybridisation. A diagram depicting a simplified version the hierarchy and interactions of some of the key genes known to be involved in limb development which were examined during pelvic fin development. The expression of pitx1 and tbx4 were examined due to their known necessary roles in hindlimb and pelvic fin development, while the expression of tbx5 was examined due to its known role in forelimb and pectoral fin development (Agarwal et al., 2003; Ahn et al., 2002; Cole et al., 2003; Garrity et al., 2002, Logan and Tabin, 1999; Lanctôt et al., 1999; Naiche and Papaioannou, 2003; Marcil et al., 2003; Rallis et al., 2003; Ruvinsky et al., 2000; Shapiro et al., 2004; Szeto et al., 1999; Takeuchi et al., 2003; Tamura et al., 1999). The expression pattern of fgf10a was examined due to its known role downstream of tbx4 during hindlimb development and its necessary role in the establishment and maintenance of the AER (Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006; Min et al., 1998; Naiche and Papaioannou, 2003; Norton et al., 2005; Ohuchi et al., 1997; Sekine et al., 1999; Te Welscher et al., 2002; Yonei-Tamura et al., 1999). The expression pattern of the genes sp8 and fgf8a were examined due to their necessary roles in the function of the AER and limb outgrowth (Kawakami et al., 2004; Ohuchi et al., 1997). Finally, the expression pattern of shh was examined due to its role in the outgrowth and patterning of the limb (Bell et al., 1998). *The tbx4 homologue, tbx5, replaces tbx4 in the forelimb/pectoral fin.

70 5.2. Results 5.2.1. Sense probes In situ hybridisation against sense probes for pitx1, tbx4, tbx5, fgf10a, sp8, fgf8a and shh was performed as a control for wild-type and pelvic finless samples and all revealed no background staining (data not shown).

5.2.2. pitx1 expression In situ hybridisation performed against pitx1 mRNA in wild-type and pelvic finless zebrafish detected pitx1 mRNA transcripts in both pelvic finless and wild-type zebrafish (Figure 5.2). No pitx1 mRNA transcripts were detected in the developing pectoral fins of wild-type or pelvic finless zebrafish embryos (Figure 5.2 A and B). At 21 dpf, pitx1 mRNA transcripts were observed in the early pelvic fin buds of wild-type zebrafish and in the pelvic bulges of pelvic finless zebrafish (Figure 5.2 C and D). By 28 dpf, pitx1 mRNA transcripts were restricted to the proximal portion of wild-type pelvic fins and pitx1 mRNA transcripts were present in the pelvic bulges of pelvic finless zebrafish (Figure 5.2 E and F).

71 Figure 5.2. pitx1 expression

In situ hybridisation against pitx1 mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A,B) and developing pelvic fin stages (C-F). A-B) No pitx1 mRNA transcripts were detected in the developing pectoral fins of wild-type or pelvic finless zebrafish. C-F) pitx1 mRNA transcripts were detected in the proximal portion of the pelvic fins of wild-type and pelvic finless zebrafish.

72 5.2.3. tbx4 expression tbx4 mRNA transcripts were detected in the developing pelvic region of both pelvic finless and wild-type zebrafish (Figure 5.3). tbx4 mRNA transcriptions were not detected in the pectoral fins of both pelvic finless and wild-type zebrafish embryos (Figure 5.3 A and B). At 21 dpf, tbx4 mRNA expression was observed in the developing pelvic fin regions of both pelvic finless and wild-type zebrafish (Figure 5.3 C and D). By 28 dpf tbx4 expression was confined to the proximal mesenchyme of the pelvic fins of wild-type zebrafish, whereas tbx4 mRNA continued to be expressed in the pelvic bulges of pelvic finless zebrafish (Figure 5.3 E and F).

73 Figure 5.3. tbx4 expression In situ hybridisation against tbx4 mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A, B) and developing pelvic fin stages (C-F). A-B) No tbx4 mRNA transcripts were detected in the developing pectoral fins of wild-type or pelvic finless zebrafish. C-F) tbx4 mRNA transcripts were detected in the proximal portion of the pelvic fins of wild-type and pelvic finless zebrafish

74 5.2.4. tbx5 expression The expression pattern of tbx5 mRNA was examined in both pelvic finless and wild-type zebrafish. tbx5 mRNA transcripts were detected in the mesenchyme of the developing pectoral fins of both pelvic finless and wild-type sibling embryos at 36 hpf (Figure 5.4 A-D). At 21 dpf, no tbx5 mRNA transcripts were detected in the developing pelvic fin regions of pelvic finless or the pelvic fins wild-type zebrafish (Figure 5.4 E and F). Additionally, no tbx5 mRNA transcripts were detected in the pelvic regions of pelvic finless or the pelvic fins wild-type zebrafish at 28 dpf (Figure 5.4 G and H).

75 Figure 5.4. tbx5 expression

In situ hybridisation against tbx5 mRNA in wild-type (A, C, E, G) and pelvic finless (B, D, F, H) zebrafish in the pectoral fins (A-D) and developing pelvic fin stages (E-H). A-D) tbx5 mRNA transcripts were detected in the developing pectoral fins of wild-type or pelvic finless zebrafish at 36 hpf. E-H) No tbx5 mRNA transcripts were detected in the pelvic fins of wild-type and pelvic finless zebrafish. Black arrowheads mark the position of the pectoral fins at 36 hpf.

76 5.2.5. fgf10a expression fgf10a mRNA transcripts were detected in the mesenchyme of the developing pectoral fins of both pelvic finless and wild-type embryos (Figure 5.5 A and B). At 21 dpf, fgf10a mRNA transcripts were detected in the developing pelvic fins of the wild-type zebrafish but were not detected in the pelvic bulges of pelvic finless zebrafish (Figure 5.5 C and D). fgf10a mRNA transcripts were detected in the pelvic fins of wild-type zebrafish at 28 dpf, but were not detected in the pelvic bulges of pelvic finless zebrafish at this stage (Figure 5.5 E and F).

77 Figure 5.5. fgf10a expression In situ hybridisation against fgf10a mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A,B) and developing pelvic fin stages (C-F). A-B) fgf10a mRNA transcripts were detected in the proximal portions of the pectoral fin buds of both wild-type and pelvic finless zebrafish at 36 hpf. C-D) fgf10a mRNA transcripts were detected in the proximal portions of the pelvic fins of wild-type, but not pelvic finless zebrafish at 21 dpf and 28 dpf.

78 5.2.6. sp8 expression During pectoral fin development, sp8 mRNA transcripts were detected in the apical ectodermal thickening of the pectoral fin of both pelvic finless and wild-type sibling embryos at 36 hpf (Figure 5.6 A and B). At 21 dpf, sp8 mRNA transcripts were observed in the distal edge of the developing pelvic fins of wild-type zebrafish, whereas transcripts were observed in the proximal-to-middle portion of the pelvic bulge of pelvic finless zebrafish (Figure 5.6 C and D). At 28 dpf in wild-type zebrafish, sp8 mRNA transcripts were detected near the distal edge of the developing pelvic fin (Figure 5.6 E). In contrast, in pelvic finless zebrafish, expression of sp8 mRNA transcripts was observed throughout the pelvic bulge (Figure 5.6 F).

79 Figure 5.6. sp8 expression In situ hybridisation against sp8 mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A, B) and developing pelvic fin stages (C-F). A-B) sp8 mRNA transcripts were detected in the distal edge of the pectoral fin buds of both wild-type and pelvic finless zebrafish at 36 hpf. C-F) sp8 mRNA transcripts were detected in the distal edge of the pelvic fins of wild-type zebrafish at 21 dpf and 28 dpf, but were detected throughout the pelvic bulges of pelvic finless zebrafish at the same stages.

80 5.2.7. fgf8 expression At embryonic stages, fgf8a mRNA expression could be detected in the distal edge of the developing pectoral fins of both pelvic finless and wild-type embryos (Figure 5.7 A and B). However, at 21 dpf, fgf8a mRNA transcripts could only be detected in the distal tip of the pelvic fin buds of the wild-type zebrafish, whereas fgf8a mRNA transcripts were not detected in the pelvic bulges of pelvic finless zebrafish (Figure 5.7 C and D). This pattern was still evident at 28 dpf, with no evidence of fgf8a mRNA transcripts in the pelvic bulges of pelvic finless zebrafish, despite the continued presence of fgf8a mRNA expression in the distal edge of the pelvic fins of wild-type zebrafish (Figure 5.7 E and F). It should be noted that the fgf8a expression in the wild-type fish is weak, however even weak signal was never detected in pelvic finless individuals.

81 Figure 5.7. fgf8a expression In situ hybridisation against fgf8a mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A, B) and developing pelvic fin stages (C-F). A-B) fgf8a mRNA transcripts were detected in the distal edge of the pectoral fin buds of both wild-type and pelvic finless zebrafish at 36 hpf. C-F) fgf8a mRNA transcripts were detected in the distal edge of the pelvic fins of wild-type zebrafish at 21 dpf and 28 dpf, but were not detected in the pelvic bulges of pelvic finless zebrafish at the same stages. Black arrowheads mark the location of the fgf8a mRNA transcripts.

82 5.2.8. shh expression In the developing pectoral fins of both pelvic finless and wild-type embryos, shh mRNA transcripts were detected in the posterior portion of the pectoral fin bud at 36 hpf (Figure 5.8 A and B). At 21 dpf, shh mRNA transcripts could not be detected in the pelvic fin regions of either wild-type or pelvic finless zebrafish (Figure 5.8 C and D). However, at 28 dpf, shh mRNA transcripts were detected in the posterior region of the pelvic fins of wild-type zebrafish (Figure 5.8 E). In contrast, at the same stage, no shh mRNA transcripts were ever detected in the pelvic fin regions of pelvic finless zebrafish (Figure 5.8 F).

83 Figure 5.8. shh expression

In situ hybridisation against shh mRNA in wild-type (A, C, E) and pelvic finless (B, D, F) zebrafish in the pectoral fins (A, B) and developing pelvic fin stages (C-F). A-B) shh mRNA transcripts were detected in the posterior portion of the pectoral fin buds of wild-type and pelvic finless zebrafish at 36 hpf. C-D) No shh mRNA transcripts were detected in the developing pelvic fins of wild-type or pelvic finless zebrafish at 21dpf. E-F) shh mRNA transcripts were detected in the developing pelvic fins of wild-type individuals at 28 dpf, but not in the pelvic bulges of pelvic finless zebrafish at the same stage. Black arrowheads mark the location of shh mRNA expression.

84 5.3. Discussion 5.3.1. Overview of results In order to better characterise the pelvic finless phenotype, the expression patterns of key genes known to be involved in limb development were examined in pelvic finless zebrafish. There is little published data on gene expression in the developing pelvic fins of teleosts, therefore this analysis was conducted in combination with the investigation of normal gene expression patterns in the pelvic fins of wild-type zebrafish. A particular emphasis was placed on examining the several genes known to be involved in early limb and AER development as the pelvic fins of pelvic finless zebrafish were shown to arrest development at an early stage and morphologically lack an apical ectodermal thickening.

The mRNA expression patterns of pitx1, tbx4, tbx5, fgf10a, sp8, fgf8a and shh were examined by in situ hybridisation in pelvic finless and wild-type zebrafish with a particular emphasis on the pelvic regions and the pelvic fins. This analysis revealed that each of these genes, with the exception of tbx5, is expressed in the developing pelvic fins of wild-type zebrafish. In addition, all of these genes were expressed in similar patterns to those observed in the hindlimbs of tetrapods and in the pectoral fins of zebrafish as discussed in 5.3.1.1.

5.3.1.1. Gene expression in wild-type zebrafish The expression of pitx1 in the mesenchyme of the developing pelvic fins of wild-type fish is similar to published expression patterns in the hindlimbs of mice and the pelvic fins of stickleback fish (Szeto et al., 1999; Cole et al., 2003; Shapiro et al., 2004). The expression of tbx4 has previously been reported in the pelvic fins of zebrafish in a similar domain as observed in this study (Tamura et al., 1999; Ruvinsky et al., 2000). In addition, this matches the Tbx4 expression domain described in the hindlimbs of mouse and chick (Chapman et al., 1996; Gibson-Brown et al., 1996; Takeuchi et al., 1999). Similar to the forelimb specific expression of Tbx5 in tetrapods, tbx5 expression was observed in the developing pectoral fins of zebrafish and not the developing pelvic fins (Chapman et al., 1996; Gibson-Brown et al., 1996; Takeuchi et al., 1999). Expression of fgf10a was also observed in the mesenchyme of the developing pelvic fins of zebrafish which is similar to reported expression patterns of Fgf10 in tetrapods (Ohuchi et al., 1997). The published expression pattern of Sp8 is the pre- AER and AER cells of the developing limb buds of tetrapods and a similar expression pattern for sp8 was observed in the pelvic fins of wild-type zebrafish (Casanova et al., 2011; Kawakami et al., 2004). Although difficult to detect, fgf8a transcripts were detected in the distal ridge of the developing pelvic fins of zebrafish matching the reported expression of fgf8

85 in tetrapods (Crossley and Martin, 1995). In wild-type zebrafish, shh expression was observed in the posterior domain of the developing pelvic fins which is similar to the published expression domain of Shh in tetrapods (Riddle et al., 1993).

5.3.1.2. Gene expression in pelvic finless zebrafish In addition to the conserved gene expression patterns between the hindlimbs of tetrapods and the pelvic fins of zebrafish, the gene expression analysis revealed some similarities and some differences between gene expression in the developing pelvic fins of pelvic finless and wild-type zebrafish. The results demonstrated that both wild-type and pelvic finless zebrafish expressed both the hindlimb/pelvic fin associated initiation genes, pitx1 and tbx4, during the early stages of pelvic fin development. Further analysis revealed that fgf10a, a possible downstream target of tbx4 (Agarwal et al., 2003; Ng et al., 2002), was never observed in the pelvic bulges of pelvic finless zebrafish, despite the expression of fgf10 transcripts in the staged-matched pelvic fin buds of wild-type zebrafish. Although evidence of sp8 expression was observed in the pelvic bulges of pelvic finless zebrafish, no expression of fgf8a was ever observed in the overlying ectoderm of pelvic finless zebrafish. In addition, expression of the gene shh was also shown to be absent in the pelvic bulges of pelvic finless zebrafish.

5.3.2. Do pelvic finless zebrafish establish an apical ectodermal thickening? Evidence from the morphological analysis of pelvic finless zebrafish demonstrates that these fish do not establish an apical ectodermal thickening, but the genetic evidence is less clear cut. This is because the results from gene expression analysis show that only one of the two apical ectodermal ridge markers examined is expressed in the distal ectoderm of the pelvic bulges of pelvic finless zebrafish. The results from in situ hybridisation revealed that sp8 and not fgf8a is expressed in the pelvic bulges of pelvic finless zebrafish.

Traditionally, either sp8 or fgf8a have been used to mark the presence of an apical ectodermal thickening in the paired fins of zebrafish (Kawakami et al., 2004; Murata et al., 2010). However, using the presence of sp8 expression alone in the distal ectoderm of a developing fin as evidence of an apical ectodermal thickening can be misleading. This is because although sp8 does mark the cells of the apical ectodermal thickening in the pectoral fins of zebrafish (Kawakami et al., 2004), it is also known to mark pre-apical ectodermal ridge cells in chicken limbs (Casanova et al., 2011). However, in zebrafish pectoral fins, fgf8a expression is observed at a relatively late stage in the apical ectodermal thickening cells and is not expressed in the pre-apical ectodermal thickening cells (Reifers et al., 1998). It is therefore important to gain evidence of both sp8 and fgf8a expression before the presence of

86 an actively signalling apical ectodermal thickening can be determined. As only the expression of sp8, and not fgf8a, is observed in the distal ectoderm of the pelvic bulges of pelvic finless zebrafish, it is most likely that this expression pattern is marking the pre-apical ectodermal thickening cells and not the cells of a true apical ectodermal thickening.

This suggests that the pelvic bulges of pelvic finless zebrafish develop pre-apical ectodermal thickening cells, but that these cells do not progress to form an apical ectodermal thickening. In addition, the combination of genetic and morphological analyses suggests that pelvic finless zebrafish do not establish an apical ectodermal ridge. This evidence provides insights into the stage at which the pelvic fin buds arrest development, therefore the pathways which lead to the formation of an apical ectodermal thickening should be considered in order to determine the genetic cause of the pelvic finless phenotype.

5.3.3. Consideration of genetic pathways that lead to apical ectodermal thickening formation The morphological and genetic analyses of pelvic fin development in pelvic finless zebrafish have demonstrated that pelvic finless zebrafish lack an apical ectodermal thickening. Previous studies have shown that the establishment and maintenance of the apical ectodermal thickening (in fish) or the AER (in mice and chicks) is crucial for continued limb and fin development (Boulet et al., 2004; Crossley et al., 1996; Lewandoski et al., 2000; Moon and Capecchi, 2000; Barrow et al., 2003; Narita et al., 2005, Narita et al., 2007; Naiche and Papaioannou, 2003; Min et al., 1998; Sekine et al., 1999; Fischer et al., 2003; Norton et al., 2005). These studies have shown that if the apical ectodermal thickening or AER is not established, properly maintained or removed, the fin or limb will cease to develop further. This indicates that the absence of an apical ectodermal thickening may be responsible for pelvic fin loss in pelvic finless zebrafish by causing the cessation of further pelvic fin development. Therefore, it is important to consider the genetic pathways involved in the establishment and maintenance of an apical ectodermal thickening in zebrafish pelvic fins as possible causes for the loss of pelvic fins in pelvic finless zebrafish.

5.3.3.1. Specification/initiation of pelvic fins The first step in the development of hindlimbs and pelvic fins is the specification and initiation of a hindlimb/pelvic fin field from which the limbs and fins develop. Two genes, long known to be involved in the specification and initiation of pelvic fin development are pitx1 and tbx4 (Ruvinsky et al., 2000; Cole et al., 2003; Shapiro et al., 2004).

87 Pitx1 is one of the first genes known to be expressed during pelvic fin formation (Cole et al., 2003; Shapiro et al., 2004) and is thought to partially control the expression of Tbx4 (Logan and Tabin, 1999; Lanctôt et al., 1999; Szeto et al., 1999; Marcil et al., 2003). It has been established that pitx1 activity in the developing pelvic fins is essential to pelvic fin development (Cole et al., 2003; Shapiro et al., 2004). In some stickleback fish populations, the absence of pitx1 activity in the pelvic regions has been shown to result in the loss of pelvic fins in these fish (Cole et al., 2003; Shapiro et al., 2004; Chan et al., 2010).

Tbx4 has been shown to be essential during hindlimb development in mice, and has also been shown to be expressed in zebrafish pelvic fins (Naiche and Papaioannou, 2003; Ruvinsky et al., 2000, Tamura et al., 1999). Unfortunately, up to this point there have been no reported examples of loss of tbx4 in pelvic fins. However in Tbx4 knockout mice, the absence of Tbx4 activity has been shown to cause a failure of hindlimb development and the subsequent absence of hindlimbs in these animals (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). The results of these studies suggest that both pitx1 and tbx4 must be considered when investigating the pelvic finless phenotype.

The results demonstrated that both pitx1 and tbx4 are expressed in a similar fashion to wild-type zebrafish during early pelvic fin development in pelvic finless zebrafish. From this expression data, it is possible to rule out a genomic sequence variant associated with pitx1 as the cause of the pelvic finless phenotype. This is due to the fact that pitx1 is expressed normally during the early stages of pelvic fin development in pelvic finless fish and more importantly, that tbx4 is expressed normally at these stages. When excluding pitx1 as a candidate, the normal expression pattern of tbx4 is important because studies in mice and stickleback fish have shown that if Pitx1/pitx1 activity is removed the expression pattern of Tbx4/tbx4 is altered (Cole et al., 2003; Logan and Tabin, 1999; Lanctôt et al., 1999; Szeto et al., 1999; Marcil et al., 2003). When Pitx1 activity is removed in knockout mice, the expression of Tbx4 is reduced (Lanctôt et al., 1999; Szeto et al., 1999). Similarly in stickleback fish lacking pitx1 expression, the expression of tbx4 is not observed (Cole et al., 2003). These results were thought to be observed because Tbx4 expression is, at least, partially controlled by Pitx1 activity (Logan and Tabin, 1999; Lanctôt et al., 1999; Szeto et al., 1999; Marcil et al., 2003; DeLaurier et al., 2006; Duboc and Logan, 2011). Ectopic misexpression of Pitx1 is sufficient to induce Tbx4 expression in the developing forelimbs of transgenic mice (Logan and Tabin, 1999; DeLaurier et al., 2006). Additionally, in a series of gene replacement experiments in Pitx1 -/- mice, it has been shown that positive transcriptional input of Pitx1 88 ensures the appropriate levels of Tbx4 necessary for limb development are reached (Duboc and Logan, 2011). As tbx4 expression has been shown to be present in pelvic finless zebrafish, the results presented in this chapter provide strong evidence that a genomic sequence variation associated with pitx1 is not responsible for the failure of pelvic finless zebrafish to establish an apical ectodermal thickening and the subsequent loss of pelvic fins. However, a mutation associated with tbx4 and/or any downstream genetic pathways cannot be ruled out as will be demonstrated in chapters 7 and 8.

5.3.3.2. Formation of the apical ectodermal thickening in pelvic fins Although the pathways that lead to the establishment and outgrowth of the apical ectodermal thickening have not been described in the pelvic fins of zebrafish, there are some basic mechanisms which can be taken from the limbs of tetrapods and pectoral fins of zebrafish. In forelimbs and pectoral fins, specific gene expression in the dorsal and ventral ectoderm is thought to induce the distal limb/fin ectoderm to become ridge-like (see Fernandez-Teran and Ros, 2008 for a detailed review; Hatta et al., 1991; Norton et al., 2005). In zebrafish pectoral fins, once the distal ectoderm has become ridge-like, it has been demonstrated that tbx5 signalling indirectly induces fgf10a expression in the fin bud mesenchyme which, in turn, is thought to eventually trigger fgf8a activity in the overlying apical ectodermal thickening (Harvey and Logan, 2006; Mercader et al., 2006; Norton et al., 2005). It is then thought that interaction between fgf10 and fgf8 signalling drives the maintenance of the apical ectodermal thickening and the outgrowth of the pectoral fin (Kawakami et al., 2004). Although functional evidence is yet to be provided, it is possible that this mechanism is also at work in zebrafish pelvic fins due to the similarities between the known gene expression patterns in both zebrafish pectoral fins and tetrapod hindlimbs and the pelvic fin expression patterns presented in this chapter and in the literature.

As all of the abovementioned genes can be used as markers to examine the progress of fin development, the expression pattern of these genes was analysed in pelvic finless zebrafish to determine at which point of the genetic cascade that apical ectodermal thickening formation fails in the pelvic fins of pelvic finless zebrafish. The results demonstrated that although tbx4 is expressed in the pelvic bulges of pelvic finless fish, expression of fgf10a and fgf8a is never observed. This leads to the possible conclusion that the signalling pathways that lead from the onset of tbx4 expression to the regulation and expression fgf10a may be affected by the genomic sequence variant that is responsible for the pelvic finless phenotype. Therefore, the

89 genes involved in these signalling pathways must be considered as possible candidates responsible for the failure of apical ectodermal ridge formation in pelvic finless zebrafish.

5.3.3.3. Genetic pathways that lead to the onset of fgf10 expression As pelvic finless zebrafish demonstrate the early presence of tbx4 expression and not fgf10a expression, it is important to consider the genetic pathways that lead to the regulation and expression of fgf10a. This is important because, it has been previously shown that the absence of Fgf10 activity causes the cessation of limb formation in mice and pectoral fin formation in zebrafish due to a failure to form a functional AER/apical ectodermal thickening (Min et al., 1998; Sekine et al., 1999; Norton et al., 2005). It is therefore possible that the loss of fgf10a expression in the pelvic fin buds of pelvic finless zebrafish may account for the arrested development of the pelvic fins via the failure of apical ectodermal thickening formation.

While in tetrapod forelimbs, it has been shown that the onset of Fgf10 expression is directly regulated by Tbx5 activity, both mouse hindlimb and zebrafish pectoral fin studies have shown that there may be a complex regulatory mechanism downstream of Tbx4/tbx5 signalling that leads to the onset of Fgf10/fgf10a expression in the developing limb/fin (Naiche and Papaioannou, 2003; Te Welscher et al., 2002; Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006). In addition, as there is a Tbox binding element in the fgf10a promoter, it is possible that Tbx4 may directly regulate fgf10a expression (Agarwal et al., 2002; Duboc and Logan, 2011). Either way, the onset and maintenance of Fgf10 expression that is necessary for proper limb/fin development seems to be dependent on Tbx4. If tbx4 does regulate the expression of fgf10a in the zebrafish pelvic fin, then the dysregulation of tbx4 activity must be considered as a candidate to underlie the pelvic finless phenotype and will be investigated in chapters 7, 8 and 9.

5.3.4. Limitations of the gene expression analysis Although the expression patterns of several key limb/fin development genes were examined, the genetic characterisation of pelvic fin development was limited. In order to better characterise pelvic fin development and pinpoint the stage at which development is arrested in pelvic finless zebrafish it would have been beneficial to examine the expression patterns of additional genes known to be involved in limb and fin development. Attempts were made to examine the expression patterns of the early limb field markers (homeobox d9a (hoxd9a), homeobox c10a (hoxc10a) and wingless-type MMTV integration site family, member 8c (wnt8c)) and apical ectodermal ridge establishment and maintenance factors

90 (distal-less homeobox 2a (dlx2a), engrailed 1a (eng1a), fgf24, fibroblast growth factor receptor 2 (fgfr2), lymphoid enhancer-binding factor 1 (lef1), pdrm1, sal-like 3 (sall3), transcription factor 7 (tcf7), wingless-type MMTV integration site family, member 3a (wnt3a), and wingless-type MMTV integration site family, member 7a (wnt7a)), however reliable expression patterns were not achieved for these genes (Ahn et al., 2002; Cohn et al., 1997; Cygan et al., 1997; Davis et al., 1991; Fischer et al., 2003; Gardner and Barald, 1992; Harvey and Logan, 2006; Kawakami et al., 2001; Logan et al., 1997; Loomis et al., 1998; Mercader et al., 2006; Nagayoshi et al., 2008; Pizette and Niswander, 1999; Tanaka et al., 2005) . In addition, it would have also been beneficial to examine protein expression patterns in pelvic finless zebrafish to determine if the proteins of interest were expressed in the expected domains and shown no difference in expression level when compared to wild- type pelvic fins. However, this was beyond the scope of this project due to the unavailability of antibodies in zebrafish for many of the proteins of interest.

5.3.5. Linking phenotype to genotype Previous studies from tetrapod hindlimbs and zebrafish pectoral fins alongside the genetic and morphological evidence presented in this thesis have yielded insights into the genetic pathways that may be disrupted in pelvic finless zebrafish. The data presented here indicates that the developmental pathways downstream of tbx4 are affected in pelvic finless zebrafish as no expression of fgf10a, fgf8a or shh is observed in the pelvic fins of these fish. However, the basis for the lack of fgf10a, fgf8a or shh remains to be determined. It is possible that expression of these genes is never observed in the pelvic fin regions of pelvic finless zebrafish due to the early stage at which the pelvic fin buds arrest development. Another possibility is that the expression of these genes is absent in the pelvic fin regions of pelvic finless zebrafish due to a specific disruption in the genetic cascades which lead to the onset of expression for these genes.

Studies from both mice and zebrafish reveal that the loss of fgf10a expression could be directly responsible for the loss of the apical ectodermal thickening and the subsequent loss of pelvic fins (Min et al., 1998; Norton et al., 2005; Sekine et al., 1999). Some studies have indicated that Tbx4 may directly regulate Fgf10 expression in the developing hindlimb bud (Duboc and Logan, 2011; Ouimette et al., 2008), which suggests the hypothesis that a disruption in tbx4 DNA binding to the fgf10 promoter may be the cause of the loss of fgf10a expression in pelvic finless zebrafish. In contrast to this, recent studies in mice argue that Tbx4 does not solely regulate the onset of Fgf10 expression and that Fgf10 regulatory

91 pathways must be considered when investigating the absence of fgf10a expression in pelvic finless zebrafish. When determining the cause of pelvic fin loss in zebrafish, it will be important to consider all the genetic pathways which lead to the induction of fgf10 expression and the formation of an apical ectodermal thickening in zebrafish pelvic fins. However, as no other fins are affected in pelvic finless zebrafish, the evidence suggests that a genomic lesion in a pelvic fin specific element such as tbx4, a pelvic fin region regulatory element or a previously uncharacterised gene which affects the expression of fgf10a could underlie the pelvic finless phenotype.

Insights gathered from the morphological and genetic characterisation of pelvic finless zebrafish are crucial when considering candidate genes and genomic sequence variants that may be responsible for the pelvic finless phenotype. In particular, the pathways which lead to the onset and regulation of fgf10a should be considered when examining candidates identified during genetic mapping and deep sequencing of pelvic finless zebrafish. As pelvic finless zebrafish only manifest defects in pelvic fin formation, the identification of the genomic sequence variant responsible for the phenotype could yield insights into the developmental mechanisms of limb development. Genetic mapping and deep sequencing of the pelvic finless locus is described and discussed in Chapters 6, 7 and 8.

92 6. Positional cloning of pelvic finless zebrafish 6.1. Introduction As detailed in Chapters Four and Five, pelvic finless zebrafish were deemed to be of great interest due to their potential to contribute to the understanding of pelvic fin development and loss. Of particular interest were the pelvic fin specific loss of the apical ectodermal thickening and the loss of fgf10a expression in the developing pelvic fins and the subsequent arrested development of the pelvic fins. Pelvic finless zebrafish likely harboured defects which specifically affected the formation or maintenance of the apical ectodermal thickening of the pelvic fins and consequently caused the loss of further pelvic fin development. However, the pathways that lead to apical ectodermal thickening formation and maintenance in the fore or hindlimbs are not fully understood in any model organism and are yet to be explored in pelvic fin development. It was hoped that by utilising the tools available in zebrafish research to determine the cause of the loss of the apical ectodermal thickening in pelvic finless zebrafish, an essential component of the pelvic fin development pathways could be identified. Therefore, in order to gain a greater knowledge of the mechanisms underlying pelvic fin development and loss, positional cloning of pelvic finless zebrafish was conducted to identify the genomic lesion responsible for the pelvic finless phenotype.

93 6.2. General overview of positional cloning The goal of positional cloning is to identify a link between an abnormal phenotype and the genomic lesion which underlies the phenotype. To achieve this, positional cloning identifies a genomic locus responsible for an abnormal phenotype by locating its position on a chromosome. This is achieved by detecting strain specific polymorphisms and rare recombination events that occur during meiosis. However, before discussing positional cloning in detail, it is first necessary to introduce the concepts of meiosis, genetic recombination and linkage.

6.2.1. Meiosis, genetic recombination and linkage Meiosis, genetic recombination and linkage are all concepts that are commonly used when presenting positional cloning. Meiosis is the type of cell that occurs in the ovaries and testes to produce haploid gametes ( and eggs). In these cells, prior to the first meiotic division, homologous maternally and paternally derived chromosomes become aligned and joined in pairs. This creates an opportunity for the arms of the four homologous chromatids to come in contact with one another, in structures called chiasmata. During meiosis, in the chiasmata, breakages and chromosomal rearrangements can occur on and between the chromosomes (Figure 6.1). These breakages and chromosomal rearrangements result in the variation of alleles via the swap of genetic information between the homologous maternally and paternally derived chromosomes. This process of the swapping of alleles is called genetic recombination. As a result of genetic recombination, each individual chromosome will contain a unique combination of genetically recombined material from the maternally and paternally derived chromosomes and this will be propagated to the offspring. Therefore, each resulting zygote will possess maternal and paternal chromosomes which have undergone unique genetic recombination events and display a unique set of alleles.

94 Figure 6.1. Diagram of recombination during meiosis Meiosis is the type of cell division that occurs in the ovaries and testis to produce haploid gametes. Prior to meiosis I, the chromosomes are replicated which produces two exact copies of each maternal and paternal chromosome. During meiosis I, the maternal and paternal homologous chromosomes align and pair to each other. When the maternal and paternal homologs align, chiasmata are formed where the chromatids join. At the chiasmata, it is possible for genetic recombination to occur which results in the crossover of alleles between maternal and paternal homologous chromosomes. At the end of meiosis I, the homologous pairs divide and the diploid cell divides into two haploid daughter cells. Each daughter cell posses a haploid number of chromosomes composed of two chromatids. During meiosis II, a second round of cell division occurs. The two sister chromatids which make up each homologous pair are separated and passed onto one of the four resulting gamete cells. At the end of meiosis, each gamete cell posses a haploid number of chromosomes which have each undergone unique recombination events.

95 The concept of linkage is also commonly used during positional cloning to explain the relative chromosomal coordinates of two separate loci, which can be markers, genes or polymorphisms. Genetic linkage is the tendency of two loci that are located proximal to each other on a chromosome to be inherited together during meiosis. Loci that are located proximal to each other on a chromosome tend to be inherited together because statistically there is less chance of chiasmata formation (and subsequent genetic recombination) occurring between two proximally positioned loci than two distally positioned loci. Loci that are linked and inherited together during meiosis are said to co-segregate. In contrast, loci that are not linked are not necessarily inherited together due to random segregation.

The concepts of linkage and genetic recombination are utilised during positional cloning to position the mutant locus on a chromosome. These concepts are utilised by performing positional cloning in heterogeneous mutant offspring which are derived from two different inbred strains of zebrafish. This process creates heterozygous mutant individuals in which the chromosome harbouring the mutant polymorphism contains polymorphisms that differ from the chromosome harbouring the wild-type polymorphism. In these individuals, the number of recombination events decreases closer to the mutant locus. This occurs because of progressively decreasing levels of genetic recombination that occurs between closely linked loci. Therefore it is possible to locate the mutant locus by first excluding genomic regions which display non-linkage to the mutant phenotype and then identifying chromosomal regions of decreasing levels of genetic recombination.

6.2.2. Positional cloning resources 6.2.2.1. Polymorphic zebrafish strains In order to obtain heterogeneous offspring, two different inbred strains of zebrafish must be utilised. Using such strains generates genetic polymorphisms between the chromosomes harbouring the mutation and the chromosome harbouring the wild-type polymorphism. Although, no zebrafish strains are truly isogenic, there are several inbred zebrafish lines which show a relatively high degree of genomic sequence similarity on both homologous chromosomes, which can be used in positional cloning mapping pedigrees.

The Tü and WIK strains of zebrafish are the mostly commonly used backgrounds for positional cloning because historically, most zebrafish mutants have been induced on the Tü background due to the two large scale N-ethyl-N-nitrosourea (ENU) forward genetic screens performed in Tübingen and Boston (Driever et al., 1996; Haffter et al., 1996). Therefore, Tü is the most widely used background for mutants during positional cloning. The WIK strain of

96 zebrafish is derived from the wild Indian zebrafish and has a relatively high degree of polymorphism compared to the Tü line (Rauch et al., 1997). Approximately, 68% of scorable microsatellite markers used in a genome scan (see 6.2.2.2) will show polymorphisms between the WIK and Tü lines (Geisler, 2002). Furthermore, the WIK strain is more robust, requiring less intensive fish husbandry practises when conducting positional cloning projects than some other inbred zebrafish lines (such as SJD).

6.2.2.2. Positional markers In order to identify the mutant locus, polymorphisms which consistently co-segregate with the mutant phenotype need to be identified and assigned a chromosomal location in a process known as a genome scan. To achieve this, positional markers are used to amplify and score polymorphisms in heterogeneous mutant individuals and siblings. The most commonly used positional markers in zebrafish are SSLPs (Litt and Luty, 1989; Tautz, 1989). The most commonly used SSLPs during positional cloning are genomic sequences containing tandem CA repeats of varying length which can be of differing lengths between different inbred strains of zebrafish (Litt and Luty, 1989; Tautz, 1989). During positional cloning, these differences between different SSLPs are examined by PCR amplification of the repeats and analysis by gel electrophoresis. The SSLPs which are polymorphic between the two original zebrafish strains are utilised for further analysis. There are multiple modes of SSLP inheritance and these are explained in Figure 6.2.

97 Figure 6.2. Modes of SSLP inheritance This diagram depicts the possible polymorphic systems that were used in this project. The size differences

shown are represented by the position of the bands. The F0 generation represents the homozygous wild-type individual of a WIK background crossed to a homozygous mutant on the Tü background. The F1 generation are

the heterozygous individuals in which genetic recombination occurs. The F2 generation consists of mutants and their phenotypic wild-type siblings. The WIK polymorphims of theoretical SSLP PCR products are represented by red lines, whereas blue lines represent those from the Tü background. Purple lines indicate the presence of both a WIK and Tü polymorphism. Black arrows indicate recombinant mutant individuals. A) In this ideal polymorphic system, recombinant individuals are easy to identify in the F2 mutant population as they display two distinct bands, representing the WIK and Tü polymorphisms. B) This is a non-ideal polymorphic system as the longer polymorphism is shared between the WIK and Tü genetic backgrounds. Recombinant individuals in the F2 generation can be missed as there is no way to identify if a mutant which displays only the upper band is harbouring both Tü polymorphisms or one Tü and the WIK polymorphism. C) This is also a non-ideal polymorphic system as the shorter polymorphism is shared by both the WIK and Tü genetic backgrounds. In this system, it is possible to miss recombinant individuals as there is no way to identify if a mutant individual is harbouring both Tü polymorphisms or one Tü and one WIK polymorphism at this locus. D) Multiple polymorphic systems can also occur, however it is possible to identify recombinant mutant individuals in this system.

98 6.2.2.3. Genomic Resources The zebrafish genome sequencing project conducted by the Sanger centre allows identification of genomic polymorphisms and their position on the individual chromosomes. In addition, genetic maps such as the ZMAP meiotic panel (Day et al., 2009) which combines data from two large radiation hybrid mapping panels (Chevrette M, 2000; Geisler, 1999; Hukriede, 1999; Kwok et al., 1998) and the double haploid heat shock meiotic mapping panel (Kelly, 2000; Postlethwait, 2000; Woods, 2000) also provide genetic markers with which to positionally clone mutations in zebrafish. These invaluable resources make positional cloning in zebrafish a viable method for identifying mutations generated by forward genetic screens, or those that arise naturally, such as the finless zebrafish described in this thesis.

6.2.3. Positional cloning workflow 6.2.3.1. Creating heterogeneous zebrafish The first step of the positional cloning process is to obtain heterogeneous mutant and wild- type offspring from which genomic DNA can be extracted and analysed. This is achieved by creating mapping pedigrees from fish of originally distinct genetic backgrounds. For the purposes on this discussion, it should be assumed that the mutagenesis was performed on the Tü strain and the WIK strain was the polymorphic reference strain used for mapping. In addition, the reader should assume that the mutation of interest is homozygous viable and the line can be maintained as homozygotes.

Definitions

Wild-type: WIK/WIK Mutant: Tü (mutant)/ Tü (mutant) Heterozygote: Tü (mutant)/ WIK or WIK/ Tü (mutant) Sibling: Tü (mutant)/ WIK or WIK/ Tü (mutant) or WIK/WIK

To create a mapping pedigree, two rounds of crosses must be performed over several generations of zebrafish. In the first round of crosses, individual wild-type fish are pair

mated to individual mutant zebrafish. These fish are the F0 generation. The resulting offspring from these crosses (the F1 generation) are raised and brother-sister mated. In turn,

the offspring from these crosses are the F2 generation which are the heterogeneous mutant

and sibling individuals which are used during positional cloning. The F2 generation is raised and sorted on the basis of phenotype. They are sorted into mutants (which display the abnormal phenotype) and siblings (which demonstrate a wild-type phenotype, but will consist

99 of homozygous wild-type and heterozygous fish). It is from this F2 generation that genomic DNA is extracted and used during positional cloning.

6.2.3.2. Low resolution mapping The second phase in genetic mapping is to use low resolution mapping to identify the chromosome which harbours the mutant locus. Low resolution mapping utilises genome scanning to identify the chromosome which is linked to the mutant locus by identifying SSLP polymorphisms which co-segregate with the mutant phenotype. As SSLPs occur relatively frequently and evenly throughout the genome, they provide a good system of reference points and are therefore useful tools for assigning the mutation to a chromosome by genome scanning.

Genome scanning involves the analysis of two hundred and forty different SSLPs spread throughout the zebrafish genome. The zebrafish genome consists of approximately 1.7 x 109 base pairs of DNA (Hinegardner, 1972), which corresponds to a genomic length of 2500cM in the female zebrafish and 950cM in the male zebrafish due to the increased frequency of meiotic cross over events in female zebrafish (Postlethwait, 1994; Singer, 2002). Therefore, using two hundred and forty SSLPs gives approximately ten SSLPs per chromosome, or approximately one SSLP per 10 cM (in the female). This gives enough coverage to assign a genomic mutation to a chromosome after genome scanning analysis. The SSLPs used during this project were those suggested by http://zebrafish.mgh.harvard.edu.

Genome scanning analysis, also known as bulk segregant analysis, is performed by pooling gDNA from forty mutant and forty wild-type sibling individuals from a single mapping pedigree (Michelmore, 1991). Each SSLP is PCR amplified from mutant and wild- type pools of gDNA and the amplified bands are compared by agarose gel electrophoresis. Each SSLP is analysed to determine if it is polymorphic between the background strains, which can be visualised by the presence of different sized bands or by bands of different intensities on the gel. If a SSLP is polymorphic, it is then analysed to determine if it is linked to the mutant phenotype. A SSLP is said to be linked to the mutant phenotype when one of the polymorphisms (represented by a band of different size or intensity) consistently co- segregates with the mutant pool. Linkage of the mutation to a chromosome is confirmed when several SSLPs from the same chromosome co-segregate with the mutant phenotype.

6.2.3.3. Intermediate resolution mapping Once low resolution mapping has identified a chromosome that harbours the mutant locus, intermediate resolution mapping is used to narrow down the approximate chromosomal

100 coordinates of the mutant locus. To do this, intermediate resolution mapping identifies mutant individuals in which rare recombination events have occurred.

Mutant individuals in which these rare recombination events have occurred are known as recombinants. Recombinant individuals are observed as harbouring wild-type polymorphisms which are linked to the mutant locus. Recombinant individuals are identified by amplifying SSLPs from the gDNA of mutant individuals and analysing for the presence of wild-type polymorphisms. The majority of mutant individuals will harbour the mutant polymorphism, whereas recombinant individuals will display the wild-type polymorphism at the locus for which they are recombinant. If a recombinant individual is a single recombinant, it will only display recombination on one side of the mutant locus. However, it is possible to identify double recombinants which display recombination on both sides of the mutant locus. As markers are identified progressively closer to the mutant locus the number of recombination events diminish. The genetic interval containing the mutant locus is defined by flanking markers at which at least one recombination event has been identified. It is in this way that the mutant locus is approximately positioned on a chromosome and progressively narrowed down until either a list of candidate genes can be identified for sequencing analysis or the information is used as a springboard for additional mapping strategies.

101 6.3. Chapter specific methods 6.3.1. Generation of mapping pedigrees Two mapping pedigrees were created for this positional cloning project. Female pelvic finless zebrafish were crossed to male WIK zebrafish (Figure 6.3). The offspring from these crosses were raised to sexual maturity and brother-sister mated (Figure 6.3). The resulting heterogeneous offspring from these crosses were raised until the presence or absence of pelvic fins was clearly visible, around 6 wpf, and then sorted into phenotypic mutants (lacking pelvic fins, n=122) and wild-type siblings (with pelvic fins, n=40) (Figure 6.3). It was from these heterogeneous fish that genomic DNA was extracted for positional cloning.

Genomic DNA was extracted from individuals as described in section 3.2.2. Genome scanning was performed by PCR (see 3.2.6) on gDNA pooled from 40 pelvic finless fish and compared to gDNA pooled from 40 wild-type siblings utilising 240 SSLP markers (see 3.2.5) spaced approximately every 10 cM on all chromosomes. Intermediate resolution mapping was performed by PCR (see 3.2.6) on 122 pelvic finless individuals (unless otherwise stated) and 8 wild-type siblings at each SSLP marker (see 3.2.5) described in the text.)

Figure 6.3. Diagram and time line of mapping pedigrees This diagram depicts the mapping pedigrees that were generated and used in this project. Pelvic finless individuals were crossed to wild-type zebrafish maintained on the WIK background. The heterozygous individuals were brother-sister mated to result in heterogeneous wild-type (n=40) and pelvic finless offspring (n=122) to be utilised during positional cloning. Two separate mapping pedigrees were created and used during this project.

102 6.4. Results 6.4.1. Low resolution mapping 6.4.1.1. Genome scan Genome scanning by bulk segregant analysis revealed that polymorphisms were present between the WIK wild-type reference strain and the pelvic finless strain. The PCR product(s) generated from the two hundred and forty SSLP prr paime irs demonstrated that many of the SSLPs were polymorphic in mapping pedigree 1 (Figure 6.4). The polymorphisms were indicated by multiple PCR products generated for an individual SSLP which were identified by either a difference in size or intensity of the band on the agarose gel .

Analysis of the genome scan results revealed that the pelvic finless mutation was possibly linked to one of two chromosomes. Analysis of the agarose gels revealed that several amplified SSLPs demonstrated consistent differences between the mutant and wild-type pools and that these SSLPs reside on chromosome 15 (Figure 6.5) or chromosome 21 (data not shown). Chromosome 15 SSLP markers z10289, z6312, z6712, z21982, z4396, z11320, z13230, z7381, z6024 and GOF15 and chromosome 21 SSLP markers z4492, z4425 and z4074 all displayed polymorphisms and certain polymorphisms appeared to co-segregate with the pelvic finless phenotype (Chromosome 15: Figure 6.5; Chromosome 21, data not shown).

103 Figure 6.4. Example of polymorphic markers An image taken after gel electrophoresis of PCR products from PCR amplification of pooled genomic DNA at various genomic markers. Each group of four represents genomic DNA from two mutant pools (white) and two sibling pools (yellow) after PCR amplification with a particular SSLP primer pair. SSLPs z4053 and z6661 display band of different sizes and are therefore polymorphic.

104 Figure 6.5. Genome scan results for chromosome 15 An image taken after gel electrophoresis of PCR products from PCR amplification of pooled genomic DNA at chromosome 15 SSLPs loci. Each group of four represents genomic DNA from two mutant pools (white) and two sibling pools (yellow) after PCR amplification with a particular SSLP primer pair. Cyan asterisks (*) mark SSLPs which demonstrate a difference in band size or intensity between the mutant and wild-type pools. Chromosome 15 markers z10289, z21982, z4396, z11320, z13230, z7381 and z6024 show a difference between the wild-type and mutant pools which indicates possible linkage of the pelvic finless phenotype to chromosome 15.

105 6.4.1.2. Individual PCR analysis Amplification of chromosome 15 marker z21982 was performed on individual pelvic finless zebrafish and confirmed that chromosome 15 was truly linked to the mutant phenotype. Amplification of z21982 revealed alleles which consistently co-segregated with the pelvic finless phenotype indicating that the pelvic finless locus is linked to chromosome 15 (Figure 6.6). The results of individual PCR for chromosome 15 marker z21982 revealed that this marker was an ideal polymorphic system in mapping pedigree number 1 (Figure 6.6). The results revealed that this marker had two alleles. The lower band on the gel represented the pelvic finless allele and the allele for the wild-type background was represented by the upper band visible on the gel. The majority (n=32/33) of mutant individuals harboured the lower band, indicating that this allele consistently co-segregated with the pelvic finless phenotype. Amplification of chromosome 21 markers z4492, z4425 and z4074 on individual pelvic finless zebrafish revealed that none of these markers had alleles which cosistantly co- segregated with the pelvic finless phenotype (data not shown). Therefore, the pelvic finless locus was unambiguously linked to chromosome 15.

106 Figure 6.6. Individual PCR for chromosome 15 SSLP marker z21982 An image taken after gel electrophoresis of PCR products from PCR amplification of genomic DNA from individual siblings (yellow) and pelvic finless zebrafish (white) at the z21982 locus on chromosome 15. At this locus, the wild-type allele is represented by the upper band (yellow asterisk) whereas the pelvic finless allele is represented by the lower band (white asterisk). The majority (n=32/33) of mutant individuals harboured the lower band, indicating that this allele consistently co-segregated with the pelvic finless phenotype and that the pelvic finless phenotype is linked to chromosome 15.

107 6.4.2. Intermediate resolution mapping Low resolution mapping of pelvic finless zebrafish revealed that the pelvic finless locus was unambiguously linked to chromosome 15. To identify and narrow the pelvic finless locus, intermediate resolution mapping was performed by testing additional chromosome 15 SSLPs and identifying the number of recombinant individuals at each marker.

Additional chromosome 15 SSLP markers were tested by individual PCR on pelvic finless zebrafish. The markers z6312, z6712, z4396, z11320, z13310, z7381, z13230, z21982 and z10289 were evaluated for segregation by individual PCR to determine if they were polymorphic markers that could be used to identify recombinant individuals. Representative gels for these markers are displayed in Figure 6.7 and demonstrate that additional SSLP markers z4396, z11320, z13310, z7381, z13230, z21982 and z10289 all displayed alleles specific to the pelvic finless background and could therefore be used to identify recombinant individuals (Figure 6.7).

108 Figure 6.7. Additional chromosome 15 markers An image taken after gel electrophoresis of PCR products from PCR amplification of genomic DNA from individual siblings (yellow) and pelvic finless (white) zebrafish at additional chromosome 15 loci. z6312 is non-polymorphic in this system. z6712 is non-polymorphic in this system. z4396 is non-ideal polymorphic in this system. z11320 is a non-ideal polymorphic system. z13310 is non-ideal polymorphic in this system. z7381 is a polymorphic ideal system. z13230 is a non-ideal polymorphic system. Z21982 is an ideal polymorphic system. z10289 is a non-ideal polymorphic system. The polymorphic systems represent informative markers that can be utilised to identify recombinant individuals. The wild-type alleles are represented by the yellow asterisks. The pelvic finless alleles are represented by the white asterisks. White arrows indicate recombinant individuals.

109 Individual PCR on 122 pelvic finless zebrafish at informative chromosome 15 markers revealed that a decreasing recombination gradient centred on the middle of chromosome 15. The markers at the telomeric edges of chromosome 15, such as z10289 at 7.3 cM and z7381at 79.6 cM, displayed the greatest number of recombinant individuals, with at least 23 recombinants out of 60 individuals at the z10289 locus and 50 recombinants out of 122 individuals at the z7381 locus. Additional markers were tested and showed a decreasing number of recombinant individuals when approaching 50-70 cM on chromosome 15: z21982 at 34.4 cM (n=19/59), z4396 at 49.3 cM (n=9/59) and z13230 at 67.4 cM (n=9+/122) (Figure 6.8).

110 Figure 6.8. The recombination gradient A diagram demonstrating the number of recombinant individuals at each informative marker. The number of recombinants at non-ideal polymorphic markers are represented with plus signs (+) as it is possible that a number of recombinant individuals were missed at these markers. A decreasing recombination gradient was observed when approaching 50-70 cM on chromosome 15. This indicates that the pelvic finless locus is located in this region.

111

T a p A T Analysis of recombinant individuals and the decreasing recombination gradient revealed that z11320 at 54 cM and z13310/z3760 at 64 cM were the closest flanking markers to the pelvic finless locus in which recombinants could be identified. Individual PCR at z11320 revealed that this marker was an ideal polymorphic system which displayed two alleles (Figure 6.9). The bright upper band on the gel represented the pelvic finless background allele and the allele for the wild-type background was represented by the bright lower band visible on the gel (Figure 6.9). The results of individual PCR revealed that the majority of mutant individuals harboured the lower band at this marker and that 8 out of 122 mutants were recombinant at this locus (Appendix D).

112 Figure 6.9. Individual PCR for chromosome 15 SSLP marker z11320

An image taken after gel electrophoresis of PCR products from PCR amplification of genomic DNA from individual siblings (yellow) and pelvic finless (white) zebrafish at the z11320 locus on chromosome 15. At this locus, the wild-type allele is represented by the lower band and marked with a yellow asterisk, whereas the pelvic finless allele is represent by the upper band and marked with the white asterisk. The white arrow indicates a rare recombinant pelvic finless individual. Only 8 out of 122 individuals were recombinant at this locus indicating the proximity to the pelvic finless locus.

113 Individual PCR at z13310 revealed that this marker was a complex polymorphic system in mapping pedigree one, but that it could be used to identify recombinant individuals in pedigree one (Figure 6.10). It was not informative in mapping pedigree two. In mapping pedigree one, the pelvic finless background possessed two different alleles at the z13310 locus, whereas the wild-type background possessed two alleles at this locus. Although this system was a complex polymorphic system, it was possible to discern that the pelvic finless zebrafish possessed both the upper and lower bands on the gel, which represented the pelvic finless alleles, whereas the wild-type alleles were represented by the upper band and a middle band (Figure 6.10). Analysis of the PCR products from z13310 revealed that both pelvic finless alleles consistently co-segregated with the pelvic finless phenotype in the majority of mutant individuals (Figure 6.10). Recombinants were identified by the presence of the middle band and 2 recombinants were identified out of 59 mutants (Appendix D).

114 Figure 6.10. Individual PCR for chromosome 15 SSLP marker z13310 An image taken after gel electrophoresis of PCR products from PCR amplification of genomic DNA from individual siblings (yellow) and pelvic finless (white) zebrafish at the z13310 locus on chromosome 15. At this locus, the wild-type alleles (yellow asterisks) are represented by the upper and middle bands whereas the pelvic finless alleles (white asterisks) are represented by the upper and lower bands. The white arrow indicates an identifiable rare recombinant pelvic finless individual. Only 2 out of 59 individuals were recombinant at this locus indicating the proximity to the pelvic finless locus.

115 SSLP marker z3760 was positioned at the same locus as z13310 and individual PCR at z3760 revealed that this marker was an ideal polymorphic system in mapping pedigree two (Figure 6.9). The upper band on the gel represented the pelvic finless background allele and the allele for the wild-type background was represented by the lower band visible on the gel (Figure 6.9). The results of individual PCR revealed that the majority of mutant individuals harboured the lower band at this marker, and that 12 out of 63 mutants were recombinant at this locus (Appendix D). Therefore, a total of 14 out 122 pelvic finless zebrafish were recombinants at the z11310/z3760 locus (Appendix D).

116 Figure 6.11. Individual PCR for chromosome 15 SSLP marker z3760 An image taken after gel electrophoresis of PCR products from PCR amplification of genomic DNA from individual siblings (yellow) and pelvic finless (white) zebrafish at the z3760 locus on chromosome 15. At this locus, the wild-type allele is represented by the lower band and marked with a yellow asterisk, whereas the pelvic finless allele is represent by the upper band and marked with the white asterisk. The white arrows indicate rare recombinant pelvic finless individuals. Only 12 out of 63 individuals were recombinant at this locus indicating the proximity to the pelvic finless locus.

117 As none of the individuals that were recombinant at z11320 were recombinant at 13310/z3760, these genetic markers flank the mutant/finless locus (Appendix D). Therefore, the pelvic finless locus was unambiguously located between z13310/z3760 and z11320 at 54.1 cM and 63.7 cM on chromosome 15 (Figure 6.12). Additional markers, z22430 at 55.4 cM and z21165 at 57.9 cM were tested to further narrow down the interval. z21165 revealed no recombinate individuals in both mapping families (n= 0/122) whereas z22430 was only polymorphic in mapping family 1 and revealed no recombinate individuals in this family (n=0/59). Therefore, both markers revealed no recombinant individuals and could not be utilised to further narrow down the pelvic finless locus.

118 Figure 6.12. Genetic interval containing the pelvic finless locus A diagram of the genetic interval containing the pelvic finless locus The closest markers at which recombinant individuals could be identified were z11320 and z13310. This identifies the genomic interval containing the pelvic finless locus as between z11320 (54.1 cM) and z13310 (63.7 cM) on chromosome 15.

119 6.5. Discussion 6.5.1. Overview of results Low and intermediate resolution mapping of pelvic finless zebrafish determined the approximate genomic location of the pelvic finless mutant locus. Low resolution genomic mapping was performed by genomic scanning and unambiguously linked the pelvic finless locus to chromosome 15 while intermediate resolution genomic mapping generated a recombination gradient which positioned the pelvic finless locus between z11320 at 54.1 cM and z13310/z3760 at 63.7 cM. This region contains 99 genes (see appendix H).

6.5.2. Identifying pelvic finless genomic lesion Positional cloning of pelvic finless zebrafish identified a 12 Mb genomic interval which contains the genomic lesion responsible for the pelvic finless phenotype. In the past, traditional mapping would be continued to further narrow down the interval containing the pelvic finless locus. It would have been normal to test the closest flanking markers and additional markers on up to 2000 mutant individuals to further narrow the region of interest. Once the interval had been sufficiently narrowed down, fine mapping using SNP analysis, chromosomal walks and gene of interest sequencing would have been initiated (Zhou and Zon, 2011). These processes are labour and resource intensive and time consuming. For example, the pelvic finless phenotype is not identifiable until around 3-4 wpf, therefore to obtain 2000 pelvic finless zebrafish would have involved generating, rearing and sorting

8000 F2 individuals at 4 wpf. This represented a practical impossibility with resources available in the timeframe of this project.

As this project was conducted on the cusp of the emergence of new sequencing technologies, the traditional mapping was halted and an approach utilising a next generation sequencing method was adopted. The application of next generation sequencing allowed for a non-traditional approach to the genetic mapping of pelvic finless zebrafish, which negate some of the obstacles encountered during traditional fine genetic mapping (Zhou and Zon, 2011). As the genomic interval containing the pelvic finless locus had been identified, deep sequencing technologies could now be employed to obtain the entire genomic sequence of the identified region and thus reveal the genomic lesion underlying the pelvic finless phenotype. For these reasons, the genetic mapping of pelvic finless zebrafish continued via region-specific targeted next generation sequencing. The progress of this strategy will be presented in Chapters 7 and 8.

120 7. Targeted sequencing of the pelvic finless locus 7.1. Introduction As detailed in Chapter 6, positional cloning had positioned the genomic region containing the pelvic finless locus on chromosome 15 in the region flanked by z11320 at 54 cM and z13310 at 64 cM. However, due to the inherent difficulties of traditionally mapping the pelvic finless phenotype, as outlined in chapter 6, positional cloning was halted at this point and a non-traditional approach utilising emerging deep sequencing technology was adopted. As the interval containing the pelvic finless locus had already been identified, a region- specific sequencing strategy was employed to determine the underlying genetic lesion responsible for the pelvic finless phenotype. The region-specific targeted enrichment next generation sequencing and bioinformatic analysis of the genomic region containing the pelvic finless locus will be presented in this chapter.

121 7.2. Introduction to genetic mapping by next generation sequencing Over the last decade, the methods for mapping zebrafish mutations have not drastically changed (Zhou and Zon, 2011). This has meant that until recently, mapping zebrafish mutants required large amounts of resources and labour hours. This has left a sizeable portion (62%) of potential development and disease models molecularly uncharacterised (Bradford et al., 2011). As a result, this has limited the effectiveness of previous forward genetic screens and has discouraged further screens.

Recent advances in next generation sequencing technologies and the high quality genome sequence assembly available for zebrafish, present novel opportunities for more efficient strategies for the genetic mapping of mutant zebrafish. These new strategies also have the advantage that they can be employed after intermediate resolution mapping, or initiated from the start of a mapping project. Although an emerging technology, at the time of writing, these strategies have allowed for the identification of the genomic lesion underlying five different zebrafish mutants (Bowen et al., 2012; Gupta et al., 2010; Leshchiner et al., 2012; Obholzer et al., 2012; Voz et al., 2012).

Utilising new sequencing technologies to map zebrafish mutants is possible due to the ongoing decrease in the cost of sequencing per megabase and per genome (Figure 7.1). In September 2001, the cost per raw megabase of DNA sequence was around $5,292 while the cost per genome was $95,263,072 utilising Sanger-based sequencing methods (Wetterstrand, 2013). Improvements in computer technologies following Moore’s law meant that by October 2007, the cost per raw megabase of DNA sequence was around $397.09 while the cost per genome was $7,147,571 utilising Sanger-based sequencing methods (Wetterstrand, 2013). However, in 2008 next generation sequencing technologies came online and the decrease in the cost of sequencing began to outpace the decrease in cost anticipated due to improvements in computer technologies (Wetterstrand, 2013). In January 2008, the average cost per raw megabase of DNA sequence was around $102.13 while the cost per genome was $3,063,820 (Wetterstrand, 2013). In stark contrast, in January 2013, the cost per raw megabase of DNA sequence had dropped to $0.06 while the cost per genome had dropped to $5,671. Due to these decreasing costs, it is now possible to map a zebrafish mutant in just two weeks at the cost of USD $2000 (Obholzer et al., 2012).

122 Figure 7.1. The deceasing cost of sequencing Graphs depicting the decreasing cost of sequencing per raw megabase of DNA sequence (A) and per genome (B). With the introduction of next generation sequencing technologies, the cost of sequencing has dramatically decreased. Figure created by author from data in Wetterstrand, 2013.

123 To date, there are three main strategies for high-throughput sequencing and genetic mapping in zebrafish which are; region-specific targeted enrichment coupled with next generation sequencing (7.2.1), exome capture coupled with next generation sequencing (7.5.2.1) and whole genome sequencing by next generation sequencing (7.5.2.2). While exome capture and whole genome sequencing can be employed from the outset of a zebrafish mapping project, region-specific targeted enrichment coupled with next generation sequencing is useful once a genomic interval has been defined by positional cloning (Bowen et al., 2012; Gupta et al., 2010; Leshchiner et al., 2012; Obholzer et al., 2012; Voz et al., 2012; Zhou and Zon, 2011). As positional cloning had previously identified the genomic interval which contained the pelvic finless locus, region-specific targeted enrichment coupled with next generation sequencing was employed for this project

7.2.1. Region-specific targeted enrichment next generation sequencing Region-specific targeted enrichment coupled with next generation sequencing reveals the entire genomic sequence of a previously identified genetic interval. This strategy can be employed after traditional low and intermediate resolution mapping has identified the genomic interval containing the mutant locus and the region has been significantly narrowed. It has been previously published that this strategy can work for the region of interest of around 300 kb (Gupta et al., 2010). The region of interest is then sequenced in both homozygous wild-type siblings and non-recombinant mutant zebrafish and the sequences compared for genetic lesions.

As this strategy uses region-specific next generation sequencing, the region of interest must be enriched compared to the rest of the genome and then sequenced. To do this, targeted enrichment is utilised where the region of interest is selectively captured from a DNA sample and amplified before sequencing. There are several methods currently employed to enrich the region of interest before sequencing which are PCR, molecular inversion probes and hybrid capture. PCR has been the most widely used pre-sequencing sample preparation technique for the past 20 years (Saiki et al., 1988). This is because PCR sample preparation is well suited to Sanger sequencing, where the upper limit of sequencing is around the same size as amplicons easily generated by PCR (around 200-700 bp). However, PCR sample preparation is not well suited to high-throughput sequencing technologies because high-throughput sequencing is best used for simultaneous sequencing of many products, or large regions of the genome. To combat this problem, it is possible to

124 multiplex PCR sample preparation (using many different primer pairs designed to different amplicons) or perform long range PCR (of up to 30 kb), but these solutions involve their own drawbacks. Many primer pairs are needed for multiplex PCR and this can lead to a high level of background amplification and even a failure to amplify the sequence of interest (Cho et al., 1999; Wang et al., 1998). It is also possible to perform long range PCR to cover large regions of the genome. However for reliable amplicons for use in next generation sequencing, overlapping products of no more than 10 kb are recommended (Mamanova et al., 2010). Therefore, to sequence large amounts of the genome, many reactions are needed and this can be time consuming and expensive.

Despite some draw backs to targeted enrichment by PCR, it is recommended for regions up to several hundred kilobases long, when the sequencing run is performed in conjunction with many other bar-coded and pooled samples. This is because it is feasible and efficient in terms of cost, workload and amount of DNA involved (Mamanova et al., 2010). Molecular inversion probes offer another method of targeted enrichment of a region of interest for next generation sequencing. These probes are based on an enzymatic method for target region amplification based on target circularisation, which works well with multiplexing (Faruqi et al., 2001; Antson et al., 2000; Hardenbol et al., 2003; Hardenbol et al., 2005; Lizardi et al., 1998). In this method, single stranded oligonucleotides which consisted of target-specific sequences flanked by a common linker are designed to target the region of interest (Nilsson et al., 1994; Landegren et al., 2004). These oligonucleotides bind to their specific target sequence and are circularised by a ligase. The DNA species which have not been circularised are then digested to reduce background, while the circularised species are amplified by PCR with primers directed at the common linker.

This method of target capture has both advantages and disadvantages for use with next generation sequencing. One of the advantages of targeted enrichment by molecular inversion probes is that if this method is used for a small number of targets over many samples, it is very cost effective. In addition, if the oligonucleotides are designed well, the products can be directly sequenced, eliminating the need for shotgun library construction. On the other side, although there have been recent improvements in capture uniformity (Turner et al., 2009), the use of molecular inversion probes for targeted enrichment still compares poorly to recent capture results from capture by hybridisation (Mamanova et al., 2010). This method of targeted enrichment for next generation sequencing is recommended for projects involving a small number of targets, but a large number of samples. This is because only a small amount

125 of sample DNA is required, it is relatively easy to multiplex these reactions, and the samples can easily be pooled for sequencing runs (Mamanova et al., 2010).

The third method for targeted enrichment of a region of interest for next generation sequencing is capture by hybridisation. This method can be employed either on-array or in solution. On-array capture was the method employed to enrich the genomic interval containing the pelvic finless locus. The direct selection method for on-array hybridisation capture is well established (Lovett et al., 1991; Parimoo et al., 1991). During this method a shotgun fragment library is hybridised on a microarray slide to immobilise probes which have been designed to target the region of interest. The non-specific hybrids are then washed off and the targeted DNA is eluted. Using this method, a target region of up to 34 Mb can be captured for next generation sequencing (Mamanova et al., 2010). This method offers several advantages over PCR based methods. On-array hybridisation capture is far quicker and less laborious than PCR based methods and offers the ability to capture huge regions of genomic DNA. The drawback of on-array hybridisation capture is that it uses large amounts of DNA with a small number of probes and requires specialised equipment.

In-solution hybridisation capture overcomes many of these disadvantages by favouring large amounts of probes and a small amount of DNA, and being in solution, can be used in a 96-well plate and therefore does not require specialised equipment. In addition, for relatively small target regions (around 3.5 Mb range), in-solution hybridisation capture yields slightly superior sequence coverage, however, this difference is not observed for whole exome capture (Mamanova et al., 2010).

126 7.3. Chapter specific methods 7.3.1. Extraction of genomic DNA

Individual F2 mutants and wild-type siblings from the positional cloning mapping pedigrees (see 6.3.1) were utilised for the experiments described in this chapter. Before extracting gDNA for use in region-specific targeted enrichment next generation sequencing, the fish were first geneotyped as described in 3.2.5 to identify homozygous non-recombinant mutant individuals (n=4) and homozygous wild-type individuals (n=4).

High quality, high molecular weight gDNA was extracted from identified individuals using a Blood and Cell Culture DNA midi kit (Qiagen). Whole fish were homogenised directly after euthanasia using a mechanical tissue homogeniser and incubated with protease (Qiagen). High molecular weight gDNA was extracted following the manufacturer’s instructions for tissue and diluted into 300 µl MilliQ H2O.

To obtain concentration and purity data, samples were analysed on a NanoDrop ND-1000 Spectrophotometer (Thermo Scientific) and only samples obtaining a concentration of > 50 ng/µl and a 260/280 nm purity ration of 1.8 were utilised during this project. To check the quality, 200 ng of gDNA from each sample was run on a 0.6% agarose:TAE gel for 18 hours, at 25V at 4°C to identify the samples with an average length of greater than 50 kb. The high molecular weight gDNA samples (wt n=1; pFL n=1) were sent directly for enrichment and sequencing. 7.3.2. Targeted enrichment Region-specific targeted enrichment was performed using a hybrid array capture system by the Beijing Genomics Institute (BGI) (Hong Kong). Low and intermediate resolution mapping had previously positioned the genomic region containing the pelvic finless locus between 54 and 64 cM on chromosome 15 which corresponded to Ch15:24168360 - 36060966 (Ensembl Zv9). Working with BGI, probes were designed to the region of interest for theoretical 90% coverage of the region. This was the maximum coverage possible while remaining specific to sequences from chromosome 15. Hybrid array capture was performed with a Roche NimbleGen HD2 11.8 Mb sequence capture array (Roche NimbleGen). Qualified gDNA samples were randomly fragmented by the Covaris AFA process and the size of the library fragments was mainly distributed between 250 bp and 300 bp. Adapters were then ligated to both ends of the resulting fragments. Extracted DNA was then amplified by ligation-mediated PCR (LM-PCR), purified, and hybridized to the NimbleGen sequence capture array for enrichment. Non-

127 hybridized DNA fragments were then washed out. Both non-captured and captured LM-PCR products were subjected to quantitative PCR to estimate the magnitude of enrichment.

7.3.3. Next generation sequencing Next generation sequencing of the enriched pelvic finless locus was performed by the BGI (Hong Kong). Each captured library was loaded onto a Hiseq2000 platform (Illumina), and high-throughput sequencing for each captured library was performed to ensure that each sample met the desired average sequencing depth. Raw image files were processed by Illumina basecalling Software 1.7 for base-calling with default parameters and the sequences of each individual were generated as 90 bp pair-end reads.

7.3.4. Bioinformatics workflow Two main approaches were employed to analyse the data generated from the high- throughput sequencing and identify polymorphisms unique to the pelvic finless genotype. Polymorphism were detected by the BGI and subsequently analysed in the laboratory while consensus scanning was employed under the guidance of Dr Dane King at the Bosch Institute research computing and engineering facility. For each analysis, the reference genome was provided by Ensembl (Zv9) ftp://ftp.ensembl.org/pub/release-68/fasta/danio_rerio/dna/ the RefSeq genome was extracted from the refGene.txt file from USCS http://hgdownload.cse.ucsc.edu/goldenPath/danRer7/database/refGene.txt.gz and the Ensembl database was also utilised ftp://ftp.ensembl.org/pub/release- 68/gtf/danio_rerio/Danio_rerio.Zv9.68.gtf.gz.

SOAPaligner/SOAP2 software was used to map reads onto the reference genome. SOAPaligner (soap2.21) software was used to align the clean reads to the zebrafish reference genome with maximum 3 mismatches, the parameters were set as -a -b -D -o -u -p -2 -m -x -s 40 -l 35 -v 3 . The consensus genotypes for the wild-type and pelvic finless individuals was then made available by the BGI in data files.

7.3.4.1. Detection of large insertions or deletions In order to detect any large insertions or deletions, the assembled genomes were scanned for regions of a lack of consensus. The pelvic finless and wild-type sequences generated by the BGI were utilised. Two columns of data (the reference genome base and the sequenced base) were extracted from the consensus genotype files, base by base, filling gaps with (-). This data was extracted as .txt files for the wild-type and pelvic finless sequences. These files were reformatted as .fasta files for subsequent analysis.

128 The reference genome for the wild-type and the pelvic finless files were compared and shown to be the same. The three files (reference, wild-type and pelvic finless) were then mashed together (from column format) with a paste command and compared. Reference to wild-type, reference to mutant and wild-type to mutant sequences were compared. An asterisk (*) was used to show consensus between all three sequences and a space ( ) was used to show a lack of consensus. This information was then analysed in Geneious® 6.0.3 software (Biomatters).

7.3.4.2. Detection of single nucleotide polymorphisms by BGI Based on results from SOAPaligner, SOAPsnp software was used to assemble the consensus sequence and call genotypes in the target regions. The following parameters were set: -i -d -o -r 0.0005 -e 0.001 -u -L 150 -T -s -2. Candidate SNPs were filtered with the following criterion: SNP quality was equal or larger than 20, sequencing depth was between 4 and 200, estimated copy number was no more than 2 and the distance between two SNPs was larger than 5. The consensus genotypes were called by SOAPsnp and they were defined as the diploid genotype for the genomic positions that were subjected to SNP calling.

Furthermore, all SNPs were annotated by an automated pipeline. The fraction of target positions where high-confidence consensus genotype were called was comparable to the target regions covered in the on-array capture. The genotypes that were different to the reference genome were extracted as candidate SNPs, then an unreliable proportion of SNPs were filtered out which finally resulted in high-confidence SNPs for the samples. These SNPs were then further annotated and categorised.

7.3.4.3. Detection of small insertions and deletions by BGI Small insertion or deletion (Indel) analysis was performed using BWA to map reads onto the reference genome. Indels in the targeted regions were identified through the sequencing reads. The sequencing reads were aligned to the reference genome by BWA, and then the alignment result was passed to GATK to identify the breakpoints. Finally, the genotypes of insertions and deletions were annotated.

7.3.4.4. Analysis of SNP and Indel data The SNP and Indel analysis resulted in data that required further analysis. The provided files were opened in and analysed using Microsoft Office Excel 2007 for Windows (Microsoft). Heterozygous polymorphisms were filtered out using an Excel sort function. The polymorphism data for each annotated category (nonsense single nucleotide polymorphisms (SNPs), missense SNPs, intronic SNPs, splice SNPs, coding region Indels,

129 splice Indels, intergenic SNPs or intergenic Indels) was extracted with an Excel sorting function and both the pelvic finless and wild-type data for each polymorphism category was complied into one file. The polymorphism data was then filtered with an Excel matching command to remove any polymorphisms which matched in both genotypes. .

Identified candidate genetic lesions were investigated in additional pelvic finless zebrafish and wild-type populations. Sanger sequencing (as described in 3.2.20) was utilised to investigate the identified candidate genetic lesions in additional pelvic finless zebrafish (n=3). In addition, expressed sequence tags (ESTs) from wild-type zebrafish populations were examined for the presence of the identified candidate genetic lesions.

7.3.5. Tbx4 transcript and protein analysis 7.3.5.1. Splice site enhancer analysis To identify possible exonic splicing enhancer (ESE) sites within tbx4, the RESCUE-ESE web server was utilised (Yeo et al., 2004). This server allows a sequence to be checked for the presence of one of 447 hexamers (6 nucleotide sequences) that have been shown to be candidate ESEs in zebrafish. The wild-type tbx4 sequence was subjected to analysis alongside the pelvic finless tbx4 sequence to compare the effect of sequence variation on ESE content.

7.3.5.2. Profile comparison A protein profile comparison was performed in order to investigate the effect of the predicted amino acid substitutions on the Tbx4 protein. However, no programs were available to perform this task on zebrafish protein sequences, so therefore the equivalent mutations were tested on human protein sequences. This was possible due to the high degree of sequence homology between human and zebrafish Tbx4 proteins (Figure 7.8). Human TBX4 A92V was utilised in the place of zebrafish Tbx4 A78V while human TBX4 G93A was utilised in the place of zebrafish Tbx4 G79A.

The SNPs3D web server was utilised to generate a support vector machine (SVM) sequence profile model score (SVM profile score) based on the analysis of homology sequence families related to human proteins (Yue and Moult, 2006). A positive SVM profile score indicates a variant classified as non-deleterious whereas a negative score indicates a deleterious case. The larger the score, the more confident the classification and the accuracy is significantly higher for scores greater 0.5 or less than -0.5 (Yue and Moult, 2006).

130 7.3.5.3. Stability comparison A stability comparison was performed to identify any differences in protein stability and folding caused by the predicted amino acid substitutions in Tbx4. No crystalline x-ray structure was available for Tbx4, so therefore the equivalent amino acid substitutions were investigated in the Tbx4 homolog, Tbx5. In addition, it should be noted that this analysis was performed on human Tbx5 from the PDB file (SwissProt accession number Q99593) described in Stirnimann et al., 2010. Human Tbx5 A79V was used in place of zebrafish Tbx4 A78V while human Tbx5 G80A was used in place of zebrafish Tbx4 G79A (Figure 7.8). The total energy of the wild-type Tbx5 protein was first computed using FoldX 3.0 software using a repair PDB function (Guerois et al., 2002; Schymkowitz et al., 2005). Protein mutagenesis and subsequent stability analysis was also performed in FoldX 3.0 software using a position scan command.

131 7.4. Results 7.4.1. Identifying candidate polymorphisms Region-specific targeted enrichment coupled with next generation sequencing successfully sequenced the region of interest to a sufficient depth and coverage to allow for the identification of polymorphisms between the pelvic finless and wild-type samples. In the target region, 11,075,935 bp were sequenced to an average depth of 67.15x (pelvic finless) and 67.56x (wild-type) with a coverage of the target region of 95.04% in pelvic finless sample and 93.83% in the wild-type sample. The fraction of unique mapped bases on, or near target was 88.70% for the pelvic finless sample and 89.63% for the wild-type sample. The captured region followed a Poisson distribution which revealed that the captured region was evenly sampled. Only mapped reads were used for subsequent analysis.

Bioinformatic analysis of the region of interest revealed that polymorphisms were present in the region of interest. Compared to the reference genome, there were 114,588 polymorphisms present in the pelvic finless individual and 123,028 polymorphisms present in the wild-type individual. Further analysis, revealed that 22,919 of the polymorphisms identified in the pelvic finless individual were homozygous and not present in the wild-type sibling and therefore possible candidate genomic lesions.

7.4.1.1. SNPs and Indels Out of the 22, 919 polymorphisms identified as unique to the pelvic finless individual, 22,708 polymorphisms were identified outside the coding regions whereas 212 polymorphisms were identified in coding regions. These were further categorised into 3 SNPs that resulted in premature stop codons, 2 Indels which led to frameshift mutations, 115 SNPs which resulted in missense mutations and 92 polymorphisms in intronic regions.

7.4.1.2. Large deletions or insertions Scanning for a lack of consensus between the pelvic finless and wild-type sequence determined that the largest insertion or deletion was a deletion that occurred at Ch:15 32,183,352 and was only 7 bp long. Therefore, no large insertions, deletions or microdeletions were identified in the region of interest.

132 7.4.2. Sequencing candidate genomic lesions 7.4.2.1. Nonsense SNPs Bioinformatic analysis identified 3 nonsense SNPs in the sequenced pelvic finless individual. The identified SNPs were in the genes rap1 GTPase activating protein (rap1gap2), spastic paraplegia 20 (spg20b) and scavenger receptor class F, member 1 (scarf1) (Table 7.1).

Gene Mutation Location Change Mutation in Mutation Type pelvic finless in wild- population type Exon mRNA Codon population rap1gap2 SNP – 14 1441 481 CAA Yes Yes premature =>TAA; STOP codon Q=>STOP Scarf1 SNP – 10 2356 786 AGA=>TGA; Yes Yes premature R=>STOP STOP codon Spg20b SNP – 2 808 270 CAA Heterozygous No premature =>TAA; STOP codon Q=>STOP

Table 7.1. Nonsense SNPs A table summarising the nonsense SNPs identified in coding regions during bioinformatic analysis.

Bioinformatic analysis revealed that in exon 14, at mRNA position 1441 of rap1gap2 a thymine replaced a cytosine, which was predicted to result in an in-frame premature stop codon (Figure 7.2A). Sanger sequencing confirmed this SNP was present in additional pelvic finless zebrafish (Figure 7.2 A). Additional analysis of EST EE213219.1 demonstrated that this was a background SNP present in wild-type zebrafish.

Analysis of scarf1 identified a SNP present at mRNA position 2356 in exon 10 which replaced an adenine with a thymine to code for an in-frame stop codon (Figure 7.2 B). Sanger sequencing revealed that this SNP was also present in additional pelvic finless zebrafish (Figure 7.2 B). Additional analysis of EST CT726283.2 revealed that this was a background SNP present in wild-type zebrafish (Figure 7.2 B).

Additionally, bioinformatic analysis identified a SNP in exon 2 of spg20b at mRNA position 808 which replaced a cytosine with a thymine (Figure 7.2 C). Sanger sequencing of this gene revealed that additional pelvic finless zebrafish harboured both polymorphisms at this locus (Figure 7.2 C).

133 Figure 7.2. Nonsense mutations are present in rap1gap2 and scarf1 in zebrafish The results obtained from next generation sequencing and subsequent Sanger-based sequencing for A) rap1gap2 B) scarf1 and C) spg20b. A) In rap1gap2 a thymine replaces a cytosine at position 1441, coding for a premature stop codon in the sequencing results, however this polymorphism is also present in the sequence of EST clone of rap1gap1 (EE213219.1). B) In scarf1 a thymine replaces a adenine at position 2356, coding for a premature stop codon in the sequencing results, however this polymorphism is also present in the sequence of EST clone of scarf1 (CT726283.2). C) In spg20b a thymine replaces a cytosine in some, but not all of the sequencing results. The reference genome was taken from Ensembl Zv9. WT NGS: sequence obtained from next generation sequencing of the homozygous wild-type individual. pFL NGS: sequence obtained from next generation sequencing of the non-recombinant homozygous pelvic finless individual. pFL Sanger: consensus sequence obtained from Sanger-based sequencing of additional pelvic finless zebrafish.

134 7.4.2.2. Frameshift Indels Bioinformatic analysis additionally identified two candidate Indels which were predicted to cause frameshift mutations in the genes ribonuclease/angiogenin inhibitor 1 (rnh1) and nuclear protein, ataxia-telangiectasia locus (npat) (Table 7.2).

Gene Mutation Type Location Change Mutation in pelvic Exon mRNA Codon finless population rnh1 Indel 6 780 260 + 2 bp No frameshift npat Indel 14 1227 409 -2bp No frameshift

Table 7.2. Frameshift Indels A table summarising Indels identified in coding regions during bioinformatic analysis.

In the sequenced pelvic finless individual, the addition of two adenines was observed at mRNA position 1227, in exon 14 of NPAT (Figure 7.3 A). Sanger sequencing demonstrated that this polymorphism is in fact not present in additional pelvic finless zebrafish (Figure 7.3 A)

Analysis of the gene RNH1 revealed the deletion of a cytosine and a thymine starting at mRNA position 780 in exon 6 (Figure 7.3 B). Sanger sequencing demonstrated that this polymorphism was also not present in additional pelvic finless zebrafish (Figure 7.3 B).

135 Figure 7.3. Frameshift Indels identified in npat and rnh1 are not present in zebrafish The results obtained from next generation sequencing and subsequent Sanger-based sequencing for A) npat and B) rnh1. A) An insertion of two adenine residues at position 1227 of npat was observed in the next generation sequencing results for a pelvic finless individual but was not confirmed by Sanger sequencing. B) A deletion of a cytosine and a thymine at position 780 of rnh1 was observed in the next generation sequencing results for a pelvic finless individual but was not confirmed by Sanger sequencing. The reference genome was taken from Ensembl Zv9. WT NGS: sequence obtained from next generation sequencing of the homozygous wild-type individual. pFL NGS: sequence obtained from next generation sequencing of the non-recombinant homozygous pelvic finless individual. pFL Sanger: consensus sequence obtained from Sanger-based sequencing of additional pelvic finless zebrafish.

136 7.4.2.3. Missense SNPs Region-specific targeted next generation sequencing also identified 115 missense polymorphisms in the genomic interval containing the pelvic finless locus. Analysis of the 115 missense polymorphisms revealed several SNPs in a striking candidate gene. Three candidate missense mutations were located in the coding region of tbx4.

Further analysis revealed that the three SNPs were all located in exon 3 of tbx4 (Figure 7.4). The first candidate mutation was located at mRNA position 233 and resulted in a thymine in place of a cytosine. The other two candidate mutations were located at mRNA positions 236 and 237. At mRNA 236 a cytosine took the place of a guanine and at mRNA position 237 an adenine replaced a cytosine. Sanger sequencing of exon 3 of tbx4 in four additional pelvic finless zebrafish demonstrated that all three candidate mutations were present in the gDNA of additional pelvic finless zebrafish (Figure 7.4). In addition, examination of tbx4 ESTs revealed that these candidate mutations are not background polymorphisms present in wild-type zebrafish (Figure 7.4).

137 Figure 7.4. Candidate mutations in tbx4 are present in pelvic finless zebrafish The results obtained from next generation sequencing and subsequent Sanger-based sequencing of gDNA for tbx4. The reference genome was taken from Ensembl Zv9. Three missense SNPs of a thymine replacing a cytosine at position 233 of tbx4, a cytosine replacing a guanine at position 237 of tbx4 and an adenine replacing a cytosine at position 238 of tbx4 were observed by next generation sequencing of a pelvic finless individual and confirmed by Sanger sequencing of additional pelvic finless zebrafish. WT NGS: sequence obtained from next generation sequencing of the homozygous wild-type individual. ESTs: sequences obtained from tbx4 EST clones BC162554.1 and AF179406.1. pFL NGS: sequence obtained from next generation sequencing of the non-recombinant homozygous pelvic finless individual. pFL Sanger: consensus sequence obtained from Sanger-based sequencing of additional pelvic finless zebrafish.

138 7.4.3. Investigation of tbx4 transcript 7.4.3.1. Sequence Sanger-based sequencing of pooled cDNA extracted from wild-type (n=30) and pelvic finless (n=30) embryos demonstrated that the three candidate mutations were all present in cDNA obtained from pelvic finless embryos and absent from cDNA obtained from wild-type embryos (Figure 7.5). Sanger-based sequencing of pooled cDNA extracted from the developing pelvic regions of wild-type (n=30) and pelvic finless (n=30) zebrafish demonstrated that tbx4 transcripts were present in the developing regions of both wild-type and pelvic finless zebrafish. This analysis also revealed that the three candidate mutations were present in pelvic finless zebrafish but absent in the wild-type zebrafish (Figure 7.5).

139 Figure 7.5. tbx4 cDNA sequence Sequence of tbx4 exon 3 cDNA sequence. Notice the candidate mutations at positions 233, 236 and 237 (highlighted in green, blue and red) in the pelvic finless cDNA sequence obtained and amplified from embryos and the pelvic regions of pelvic finless zebrafish.

140 7.4.3.2. Splicing Additional sequence analysis revealed that the three candidate mutations were located adjacent to the exon 3 3’splice site and within a predicted ESE sequence (Figure 7.6 A). The candidate mutations were not predicted to affect the splice site, but could possibly disrupt the function of the predicted ESE (see 7.3.5.1).

It was predicted that if the identified candidate mutations disrupted the function of the predicted ESE, exclusion of exon 3 of the pelvic finless tbx4 transcript would be observed and the amplified pelvic finless tbx4 transcript would be 95 bp shorter than the wild-type tbx4 transcript (pelvic finless tbx4: 1,643bp; wild-type tbx4: 1,738 bp) (Figure 7.6 B). Additional analysis of the pelvic finless tbx4 transcript revealed no difference in length when compared to the wild-type tbx4 transcript. Full length cDNA amplified from wild-type and pelvic finless embryos revealed that both transcripts were around 1,738 bp in length in embryonic zebrafish (Figure 7.6 C). In addition, full length cDNA amplified from the developing pelvic regions of wild-type and pelvic finless zebrafish revealed no difference in length between wild-type and pelvic finless tbx4 transcripts in the developing pelvic regions. The transcripts were also shown to be around 1,738 bp in length (Figure 7.6 C). Full length cDNA amplified and sequenced from the developing pelvic regions of wild-type and pelvic finless zebrafish also showed no consistent difference in sequence apart from the identified candidate mutations (Appendix E).

141 Figure 7.6. tbx4 transcript length A) Diagram of the Tbx4 exon 3 intron/exon boundary and splice site details. The splice site is shaded in red and the predicted ESE is shaded in blue. The identified candidate mutations lie in the ESE but not the splice site itself. B) A diagram of the intron/exon structure of tbx4 pre-mRNA and the predicted outcome of a loss of function of the predicted ESE on the pelvic finless mRNA. If the predicted ESE was affect, the pelvic fins mRNA transcript should be 95 bp shorter due to the exclusion of exon 3 C) Gel image of Tbx4 cDNA amplified from wild-type and pelvic finless embryos and developing pelvic regions. All transcripts are around 1,738 bp and no difference in transcript length was observed between samples obtained from wild-type and pelvic finless zebrafish.

142 7.4.4. Investigations of the Tbx4 protein 7.4.4.1. Amino acid sequence Bioinformatic analysis of the pelvic finless Tbx4 protein sequence revealed that the three identified candidate mutations were predicted to cause two amino acid substitutions in the Tbx4 peptide sequence. Bioinformatic analysis revealed that the candidate mutation located at mRNA position 233 corresponded to amino acid 78 and was predicted to substitute the amino acid Alanine to a Valine at this position. Additionally, the candidate mutations located at mRNA positions 236 and 237 were together predicted to substitute a Glycine to an Alanine at amino acid 79. However, the candidate mutation located at mRNA position 237 was predicted to be silent if it occurred individually, while individually the candidate mutation located at mRNA position 236 was predicted to substitute a Glycine to an Alanine at amino acid 79 (Figure 7.7).

143 Figure 7.7. Tbx4 amino acid substitutions Chromatograms of the DNA sequence and the predicted amino acid substitutions in exon 3 of tbx4. The three identified candidate mutations are predicted to result in two amino acid substitutions (A78V and G79A).

144 Additional bioinformatic analysis revealed that these amino acid substitutions occurred in important residues. Residues 78 and 79 of the Tbx4 protein are located in a highly conserved domain known as the T-box, or the DNA binding domain (Figure 7.8 A). Additional analysis revealed that Tbx4 residues 78 and 79 are perfectly conserved amongst all vertebrates with hindlimbs with known Tbx4 protein sequences (Figure 7.8 B). The only vertebrate species which showed variation at the same residues was the bottlenose dolphin (Tursiops truncatus), which also lacks hindlimbs (Figure 7.8 B). Alignment of the Tbx4 protein sequence to its homolog, Tbx5, revealed that residues 78 and 79 are also located in a predicted nuclear localisation sequence and that residue 79 is likely directly involved in DNA binding (Figure 7.8 C).

145 Figure 7.8. Tbx4 protein sequence analysis A) Diagram of the Tbx4 protein with the A78V and G79V amino acid substitutions. Note that the amino acid substitutions occur in the T-box, or the DNA binding domain. B) Multiple species alignment of the protein sequence adjacent to the A78V and G79V amino acid substitutions. These residues are highly conserved amongst vertebrates with hindlimbs/pelvic fins. C) Tbx4 and Tbx5 protein alignment of the residues adjacent to the A78V and G79V amino acid substitutions. The residues affected by the identified candidate SNPs are underlined. The nuclear localisation sequence is shaded in blue. Residue 80 of Tbx5 is circled as it is known to be directly involved in DNA binding.

146 7.4.4.2. Bioinformatic protein analysis Two separate bioinformatic approaches were utilised to analyse affects of the zebrafish A78V and G79A amino acid substitutions. Analysis of homology sequence families revealed that the human A92V amino acid substitution (corresponding to zebrafish A78V) generated a SVM profile (see 7.3.5.2) score of -1.71 while the human G93A amino acid substitution (corresponding to zebrafish G79V) generated a SVM profile score of -2.92. Both amino acid substitutions were determined to be deleterious.

The stability of the human Tbx5 protein was examined by comparing the change in free folding energy between the wild-type and mutant proteins (see 7.3.5.3). It was first determined that the folded human wild-type Tbx5 protein had a total energy of -1.64 kcal/mol. The folded human Tbx5 protein with the A79V mutation resulted in a change in total energy of 0.14 kcal/mol (Figure 7.9 A) whereas the folded human Tbx5 protein with the G80A mutation resulted in a change in total energy of 5.15 kcal/mol (Figure 7.9 B). Respectively, these represented some of the smallest changes in free energy caused by amino acid substitutions at these positions. Additionally, the folded human Tbx5 protein with both of the predicted amino acid substitutions resulted in a change in total energy of 5.42 kcal/mol.

147 Figure 7.9. Change in total free energy graphs for the predicted A79V and G80A mutations. Histograms of the change in total free energy as a result of each amino acid substitution at residues 79 (A) and 80 (B). The A79V and G80A represent some of the smallest predicted changes in total free energy.

148 7.5. Discussion 7.5.1. Overview of results Region-specific targeted enrichment coupled with next generation sequencing was employed to obtain the entire sequence of the interval containing the pelvic finless locus and subsequently identify candidate genetic lesions in the pelvic finless sequence. Using hybrid array capture, a 12 Mb region containing the pelvic finless locus was captured and enriched. The region was then sequenced to a depth and coverage that allowed for detailed bioinformatic analysis. A total of 114,588 polymorphisms were identified in the sequenced pelvic finless individual and therefore, further bioinformatic analysis was necessary before candidate genetic lesions responsible for the pelvic finless phenotype could be identified.

Additional bioinformatic analysis identified 22,919 candidate polymorphisms that were homozygous and specific to the pelvic finless individual. It was only necessary to investigate these candidate polymorphisms as the pelvic finless phenotype was a homozygous recessive mutation which followed Mendelian inheritance (Don et al., 2011; Chapter 6). Therefore, the candidate polymorphism would have to be homozygous in the pelvic finless individual and absent from the homozygous wild-type sibling. Of the identified candidate polymorphisms, 22,708 were located outside of coding regions. As these did not account for a large insertion, deletion or microdeletion, and could not easily be investigated further, they were set aside and further analysis focused on candidate polymorphisms identified in coding regions.

Polymorphisms that resulted in premature stop codons and frameshifts were first investigated. Three nonsense SNPs and two Indels were identified in the coding region of the sequenced pelvic finless individual and were sequenced in additional pelvic finless zebrafish to determine if they could be responsible for the pelvic finless phenotype. The two Indels were found to be absent in additional pelvic finless zebrafish. In addition, the SNP identified in spg20b was found to be absent in certain pelvic finless individuals. As any candidate genetic lesions would have to be present in all pelvic finless zebrafish, these polymorphisms could not be responsible for the pelvic finless phenotype.

The remaining two nonsense SNPs in rap1gap2 and scarf1 were present in additional pelvic finless zebrafish, however, the same SNPs were identified in published EST sequences. This indicated that these were background SNPs present in the wild-type zebrafish from which the ESTs were obtained. As the pelvic finless zebrafish has not been reported to spontaneously occur in other laboratories, it is unlikely that background

149 polymorphisms present in wild-type zebrafish could be the cause of the pelvic finless phenotype.

Analysis then shifted to the 115 candidate missense mutations that were identified in the sequenced pelvic finless zebrafish. Of these, one striking candidate was identified. Bioinformatic analysis revealed that there were three missense SNPs in exon 3 of tbx4. Additional analysis demonstrated that these SNPs were present in additional pelvic finless zebrafish and were not present in any sequences obtained from wild-type zebrafish. It is therefore possible that one of these candidate mutations may be responsible for the pelvic finless phenotype. The implications of these three candidate mutations and the role of tbx4 in pelvic fin and hindlimb development are discussed in more detail in section 7.5.3.

7.5.2. Limitations of region-specific targeted next generation sequencing Region-specific targeted next generation sequencing has been shown previously to be a viable option for the genetic mapping of zebrafish (Gupta et al., 2010). This strategy has previously been successfully employed to identify a genomic lesion responsible for the magellan mutant zebrafish (Gupta et al., 2010). However, in this study positional cloning had identified a 300 kb genomic interval and region-specific targeted enrichment coupled with next generation sequencing revealed only 4 candidate polymorphisms in this region (Gupta et al., 2010). After further investigation, the genomic lesion underlying the magellan mutant phenotype was identified as a 31 base pair deletion in the microtubule-actin crosslinking factor 1 (macf1) gene, demonstrating that region-specific targeted enrichment next generation sequencing is a viable strategy for genetic mapping in zebrafish (Gupta et al., 2010).

During the course of this project, it was found that while utilising region-specific targeted enrichment next generation sequencing for genetic mapping is a viable option, it does have some drawbacks and limitations. The main drawback of using region-specific targeted enrichment next generation sequencing to identify the genetic lesion responsible for pelvic finless phenotype stemmed from sequencing such a large region. The region-specific targeted enrichment next generation sequencing for pelvic finless zebrafish covered a 12 Mb region and returned a monumentally large amount of polymorphisms in the region. The large return of data necessitated additional bioinformatic analysis, literature review and potentially represented additional years of further experimental analysis. Despite this, the sequencing and bioinformatic analysis of pelvic finless zebrafish did identify candidate genetic lesions in a striking candidate gene. Therefore, region-specific targeted enrichment coupled with next

150 generation sequencing can be employed as a useful genetic mapping strategy for projects which cannot progress beyond intermediate resolution mapping using traditional methods, however, for future mapping projects in may be beneficial to employ a next generation sequencing strategy from the outset of the project (see 7.5.2.1 and 7.5.2.2).

7.5.2.1. Exome sequencing by next generation sequencing New technologies are allowing next generation sequencing mapping strategies to be employed from the start of a mapping project, thus eliminating the need for traditional mapping strategies. For example, exome capture coupled with next generation sequencing allows for the whole exome (all coding regions) of a mutant zebrafish to be sequenced. Using this strategy, the exome of a mutant zebrafish and its homozygous wild-type sibling are sequenced and compared. Several companies offer readymade exome capture arrays (SureSelect from Agilent and Sequence Capture Arrays or SeqCap libraries from NimbleGen, Roche) that can be used to capture the exome of the zebrafish of interest. All known exons are then sequenced in a single run using a next generation sequencing platform. Although this strategy eliminates the need for traditional mapping, it does present some disadvantages. The obvious disadvantage is that this method will not identify any genomic sequence variants outside of the exome and therefore genetic lesions in promoters, enhancers or introns will be missed. In addition, the amount of sequence information recovered will necessitate heavy bioinformatic analysis and many false positive candidates could be identified due to naturally occurring polymorphisms.

7.5.2.2. Whole genome sequencing by next generation sequencing Perhaps the most promising new mapping strategy is whole genome sequencing followed by bioinformatic analysis. Whole genome sequencing coupled with bioinformatic analysis is revolutionising the world of positional cloning in zebrafish. This approach eliminates the need for traditional mapping by essentially performing polymorphism analysis in-silico, which results in a cheaper and quicker option (Bowen et al., 2012; Leshchiner et al., 2012; Obholzer et al., 2012; Voz et al., 2012).

One way to use whole genome sequencing is to use variations of bulk-segregant linkage analysis to genetically map mutant zebrafish (Bowen et al., 2012; Leshchiner et al., 2012; Voz et al., 2012). This approach involves generating zebrafish mapping pedigrees, similar to traditional mapping, and collecting gDNA samples from the F2 mutant individuals. The gDNA is then pooled and subjected to whole genome sequencing followed by bioinformatic filtering for background strain specific polymorphisms. The bioinformatic analysis for the

151 approach involves calculating the mutant and reference strain polymorphism frequency and determining the mutant peak, which is reported to result in a genomic interval that contains the mutant locus (Obholzer et al., 2012). This method essentially performs the polymorphism analysis of traditional mapping without the need for thousands of PCR reactions. However, this method can still take up to six months of time and labour due to the generation time of zebrafish.

Another approach that uses whole genome sequencing to map and identify zebrafish mutations is to use homozygosity mapping. In this time saving approach, multiple heterozygous map crosses carrying the mutation of interest are performed and gDNA is

extracted from the F1 mutant individuals. The gDNA is subjected to whole genome sequencing, and the mutant locus is determined by identifying the chromosomal region with the largest region of homogeneity (Obholzer et al., 2012). This method works based on the fact that the region closest to the mutant locus will undergo the least amount of genetic recombination and therefore display a large degree of homogeneity.

Using either method, once the genomic region containing the mutant locus has been identified, the region is subjected to further bioinformatic analysis. The region is filtered for known polymorphisms and then bioinformatically characterised into homozygous polymorphisms such as nonsense, splicing, missense and frameshift mutations which are unique to the mutant population (Obholzer et al., 2012). In the past, these approaches and the bioinformatic analysis involved have been project specific and are not yet widely adopted. To combat this, a freely accessible software pipeline has now been created to allow other labs to adopt these novel mapping approaches (MegaMapper based on the Galaxy toolkit; Obholzer et al., 2012). Genetic mapping by whole genome sequencing represents a great reduction in time, cost and labour when compared to traditional positional cloning and it appears that this strategy could become the standard strategy employed to map mutant zebrafish.

It is important to note that when utilising any next generation sequencing technology, care must be taken as the technology has in-built limitations due to the short sequencing read lengths produced. By utilising short sequencing read lengths, mutations arising from inversions, repeat sequences and duplications can be missed by next generation sequencing and subsequent bioinformatic analysis (Alkan et al., 2010; Hert et al., 2008; Shendure and Ji, 2008). To combat this, high-quality sequencing and assembly approaches must be adopted to avoid generating incomplete sequence data. It is hoped that in the future, there will be

152 standardised guidelines for the generation and publication of next generation sequencing data that will provide a template for next generation sequencing mapping strategies for zebrafish mutants.

7.5.3. tbx4 as a candidate gene to underlie the pelvic finless phenotype Despite some drawbacks to genetic mapping by region-specific targeted enrichment next generation sequencing, candidate mutations in tbx4 were identified as possible contenders to underlie the pelvic finless phenotype. A mutant version of tbx4 is a striking candidate for the loss of pelvic fins in zebrafish for several reasons (a full review of the literature surrounding tbx4 can be found in chapters 2 and 9). In short Tbx4 is known to play a vital role in limb development and its absence is known to cause the loss of hindlimbs in mice (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). In zebrafish, tbx4 has also been shown to be expressed in the developing pelvic fins, suggesting that it plays a similar role in the pelvic fins of zebrafish as it does in the hindlimbs of other tetrapods (Ruvinsky et al., 2000; Tamura et al., 1999; Figure 5.3). In addition, it should be noted that tbx4 is pelvic fin specific and is not expressed during the development of pectoral fins, nor does its absence affect the development of pectoral fins or tetrapod forelimbs (Ahn et al., 2002; Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007; Ruvinsky et al., 2000; Tamura et al., 1999; Figure 5.3). These examples from the literature provide the first suggestions that candidate mutations in tbx4 may underlie the loss of pelvic fins in pelvic finless zebrafish.

The specificity of tbx4 expression in the pelvic fins and the specificity of the pelvic finless phenotype provide a compelling reason for further investigation of tbx4 as a candidate gene to underlie the pelvic finless phenotype. Observations of pelvic finless zebrafish morphology provide additional indications that candidate mutations in tbx4 could be responsible for the pelvic finless phenotype. Morphological observations revealed that only the pelvic fins, and no other fins, are affected in pelvic finless zebrafish (Chapter 4). This indicates that a factor specific to hindlimb/pelvic fin development is responsible for the loss of pelvic fins in these fish and tbx4 is pelvic fin specific (Ruvinsky et al., 2000; Tamura et al., 1999). In addition, pelvic finless zebrafish were shown to display defects in establishing or maintaining an apical ectodermal thickening (Chapter 4). Although there is no direct data, studies of Tbx4 in tetrapod hindlimbs and studies of its homolog, tbx5, in zebrafish pectoral fins suggest that tbx4 could have a role in the establishment and maintenance of the apical ectodermal thickening in zebrafish pelvic fins (Agarwal et al., 2003; Duboc and Logan, 2011; Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007; Ruvinsky et al., 2000; Tamura et al.,

153 1999). This would subsequently imply that any genetic lesion affecting the function of tbx4 could result in defects in the establishment and maintenance of the apical ectodermal ridge in the pelvic fins of pelvic finless zebrafish.

The mRNA expression patterns observed in the pelvic regions of pelvic finless zebrafish also suggest that mutations in tbx4 could be responsible for the pelvic finless phenotype (Chapter 5). In the pelvic regions of pelvic finless zebrafish, while the expression of tbx4 is observed in the early pelvic fin bulges, expression of fgf10a, fgf8a and shh is never observed. These results are consistent with missense candidate mutations in tbx4 underlying the pelvic finless phenotype. If missense candidate mutations were responsible for the pelvic finless phenotype, it is possible that tbx4 mRNA transcripts would still be observed as missense mutations can affect the amino acid sequence of the translated protein, and not necessarily the mRNA transcript (Belgrader and Maquat, 1994; Byrne et al., 2009). Therefore, the candidate missense mutations would not be predicted to affect expression of tbx4 mRNA, unlike some nonsense mutations which target mRNA for nonsense-mediated decay (Wittkopp et al., 2009), unless Tbx4 function was required for its own expression. As such, the detection of tbx4 mRNA expression by in situ hybridisation in pelvic finless zebrafish shown in this thesis (Figure 5.3) does not rule out tbx4 as a candidate gene to underlie the pelvic finless phenotype. However, missense mutations in tbx4 could affect the function of the Tbx4 protein (Needham et al., 2006; Reva et al., 2011; Zhang et al., 2012) and therefore, the expression genes downstream of tbx4, such as fgf10a, fgf8a and shh, would be expected to be altered. Therefore, the absence of fgf10a, fgf8a and shh expression in the pelvic regions of pelvic finless zebrafish is also consistent with a loss-of-function of Tbx4 activity, strengthening the argument that candidate missense mutations in tbx4 underlie the pelvic finless phenotype.

7.5.3.1. Analysis Tbx4 protein and mRNA transcript Several techniques were employed to determine if the candidate mutations identified in tbx4 could be responsible for the loss of pelvic fins in pelvic finless zebrafish. Examination of the pelvic finless tbx4 transcript revealed that all three candidate mutations were present in the mRNA extracted from the embryos and developing pelvic regions of pelvic finless zebrafish. Additional analysis revealed that although these candidate mutations were in a predicted exonic splice enhancer sequence, they did not affect the splicing or transcript length or other sequence of the pelvic finless tbx4 transcript.

154 Bioinformatic analysis of the predicted wild-type and pelvic finless Tbx4 protein sequence was then conducted. Analysis of the identified candidate SNPs revealed that they were predicted to cause amino acid substitutions in highly conserved amino acid residues. The candidate mutation at mRNA position 233 was predicted to cause an A78V mutation in the Tbx4 amino acid sequence whereas the candidate mutations at mRNA positions 236 and 237 were predicted to cause a G79A mutation in the Tbx4 amino acid sequence. These amino acid substitutions were shown to have only a small affect on the total free folding energy of the protein, however, additional analysis revealed that these amino acid substitutions occurred in a predicted nuclear localisation sequence as well as in residues predicted to be directly involved in DNA binding.

7.5.3.2. Do the identified amino acid substitutions affect Tbx4 function? The three candidate mutations identified in the pelvic finless sequence of tbx4 were shown to have no effect on the length, splicing or other sequence of the tbx4 transcript in pelvic finless zebrafish. Therefore the candidate mutations must alter the function of the Tbx4 protein if these candidate mutations truly underlie the pelvic finless phenotype. Evidence that these candidate mutations may result in at least a partial loss of function of the Tbx4 protein has been gained from protein sequence analysis, and published literature surrounding human TBX5 mutations (Collavoli et al., 2003; Kispert et al., 1995; Kispert and Herrmann, 1993; Papaioannou and Silver, 1998; Papaioannou, 2001; Smith, 1999; Stirnimann et al., 2010; Wilson and Conlon, 2002).

The identified candidate mutations result in two amino acid substitutions that occur in highly conserved residues. These residues are perfectly conserved in all vertebrate species with known Tbx4 protein sequences, with the exception of dolphins (which lack developed hindlimbs). In addition, these residues are well conserved amongst the T-box family of proteins, with amino acid residue 79 remaining perfectly conserved across the family (Ruvinsky et al., 2000). This degree of conservation at the protein level suggests that most amino acid substitutions at these residues have been selected against (Kimura, 1968; Jukes and King, 1971; Zuckerkandl and Pauling, 1962). This implies that most missense mutations at these residues give rise to a functionally impaired Tbx4 protein. Therefore, the high degree of conservation of these amino acids residues supplies evidence to suggest that the identified candidate mutations could affect the function of the Tbx4 protein and subsequently underlie the pelvic finless phenotype.

155 The question remains of how these identified candidate mutations could cause impaired function of the pelvic finless Tbx4 protein. Bioinformatic folding analysis demonstrated that the A78V and G79A amino acid substitutions had the some of the smallest effects of any amino acid substitutions at these residues. These amino acid substitutions resulted in a small change of total free energy and were therefore not likely to substantially affect the folding and stability of the Tbx4 protein. Additional bioinformatic analysis demonstrated that the A78V and G79A amino acid substitutions are located in a predicted nuclear localisation sequence (77KAGRRMFPSYKVK89) for Tbx4 (Collavoli et al., 2003). However, it has been previously shown in the Tbx4 homolog, Tbx5, that it is only the basic residues at Tbx4 positions K77, R80, R81 and K89 that are essential components of the nuclear localisation signal (Collavoli et al., 2003; Hodel et al., 2001). This line of evidence suggests that the identified candidate mutations and the resulting amino acid substitutions would not alter the function of the pelvic finless Tbx4 protein through a decrease in protein stability, but the subcellular localisation of pelvic finless Tbx4 cannot yet be excluded without further testing. Although it is beyond the scope of this project, it would be interesting to empirically test this hypothesis by GFP tagging the pelvic finless Tbx4 protein, or designing an antibody to the zebrafish Tbx4 protein, and examining the subcelluar localisation of the mutant protein. If these mutations in the predicted nuclear localisation signal of Tbx4 resulted in the mislocalisation of Tbx4 in the presumptive pelvic fin cells in pelvic finless zebrafish, this would then be of upmost interest in order to determine the mechanism behind the role of mutant Tbx4 in the loss of pelvic fins in pelvic finless zebrafish.

The remaining evidence suggests that if the identified candidate mutations in tbx4 were to underlie the pelvic finless phenotype, it would be through interfering or impeding Tbx4 DNA binding. The amino acid substitutions caused by the identified candidate mutations occur in the DNA binding domain of Tbx4 and, in particular, the G79A substitution occurs in a residue known to be directly involved in Tbx5 DNA binding (Kispert et al., 1995; Kispert and Herrmann, 1993; Papaioannou and Silver, 1998; Papaioannou, 2001; Smith, 1999; Stirnimann et al., 2010; Wilson and Conlon, 2002). In addition, it has been previously shown that mutations in the equivalent residue of Tbx5 decreases the affinity of the mutant protein to bind specific target DNA sequences and increases the likelihood of unspecific DNA binding (Stirnimann et al., 2010). Therefore, this line of evidence suggests that the identified candidate mutations in pelvic finless zebrafish could cause amino acid substitutions in a crucial DNA binding domain which would subsequently affect Tbx4 DNA binding. As it is

156 thought that Tbx4 protein DNA binding of downstream targets is essential for pelvic fin development, this line of evidence suggests that the identified candidate mutations could underlie the loss of pelvic fins by impeding Tbx4 from specifically binding target DNA sequences and thus disrupting the pelvic fin development genetic cascade.

Although beyond the scope of this project, conclusive evidence that amino acid substitutions in Tbx4 at residues 78 and 79 alter Tbx4 DNA binding could be obtained via a luciferase DNA binding affinity assay. A similar assay has been performed to test the difference in DNA binding affinity between Tbx4 and Tbx5 (Duboc and Logan, 2011). This was investigated by determining the affinity of Tbx4 and Tbx5 to bind the promoter sequence of Fgf10, a suspected downstream target of these proteins (Duboc and Logan, 2011). In order to provide mechanistic evidence that the candidate mutations identified in the pelvic finless tbx4 sequence affect the DNA binding function of the Tbx4 protein, a luciferase assay should be performed to compare the DNA binding affinity of wild-type and pelvic finless Tbx4 protein. If the identified candidate mutations were shown to decrease the DNA binding affinity of Tbx4 to its downstream targets, then this would give great insight into the mechanism underlying pelvic fin loss in pelvic finless zebrafish.

Positional cloning, bioinformatic analysis, morphology and gene expression analysis all suggest that the candidate missense mutations in tbx4 could be responsible for the pelvic finless phenotype. Therefore the three candidate missense mutations identified in the pelvic finless tbx4 sequence warrant further investigation to determine if one, or the combination, of these candidate mutations is the cause of pelvic fin loss in pelvic finless zebrafish. The continued investigation of these candidate missense mutations will be presented in Chapter 8. However, it remains possible that additional experiments may provide evidence that the identified candidate mutations did not truly underlie the pelvic finless phenotype. If this becomes the case, the investigation must turn back to the additional bioinformatic data and continued analysis of the additional SNPs and Indels must be undertaken.

157 8. Ongoing Strategies Towards Linking Genotype to Phenotype 8.1. Intro The combination of morphological and genetic analysis of pelvic fins coupled with positional cloning and region-specific targeted enrichment next generation sequencing of pelvic finless zebrafish revealed three candidate genetic lesions in tbx4 which could underlie the pelvic finless phenotype. As pelvic finless zebrafish are of interest due to their potential to contribute to the field of limb development, these candidate genetic lesions deserve further investigation. As the pelvic finless phenotype only becomes apparent around 21 dpf, the traditional approaches such as morpholino targeted knockdown (to replicate the phenotype) and mRNA injection (to rescue the phenotype) which only persist for the first few days after procedure are not applicable to pelvic finless zebrafish (Nasevicius and Ekker, 2000; Nusslein-Volhard and Dahm, 2002). Therefore, the identified candidate genomic lesions are currently being examined by a combination of transgenic rescue experiments, DNA binding assays and protein localisation experiments. The specific aims of this chapter are:

-To investigate whether a pelvic fin specific driver element can be utilised to drive protein expression in the developing pelvic regions of pelvic finless zebrafish.

-To engineer a construct to drive the expression of wild-type tbx4 in the developing pelvic regions of pelvic finless zebrafish.

-To determine if the restoration of wild-type Tbx4 can rescue pelvic fin development in pelvic finless zebrafish.

158 8.2. Chapter specific methods 8.2.1. DNA injections For DNA microinjections, 160 ng of plasmid was combined with 160 ng of transposase mRNA and 1 µl of phenol red in 3 µl of sterile water for an approximate final concentration of plasmid of 40 ng/µl. This solution was injected neat or diluted into the animal pole of 1- cell stage embryos according to the protocol outlined in 3.1.2. Injections were always performed in conjunction with several different plasmids to ensure that the injections and transposase were performing as desired.

8.2.2. The Pel2.5 kb-GFP construct 8.2.2.1. Description of the Pel2.5 kb-GFP construct A construct containing the Pel2.5kb pelvic fin enhancer element described in Chan et al., 2010 was obtained from the Kingsley Lab (Stanford, California, USA). This construct contained the Pel2.5kb element and green fluorescent protein (GFP) in a pBH-mcs-YFP vector backbone (Figure 8.1). The plasmid was propagated according to protocols outlined in 3.2.15 and 3.2.16.

Figure 8.1. The Pel2.5kb-GFP construct. A diagram of the Pel2.5kb-GFP construct as described in Chan et al., 2010.

159 8.2.2.2. Analysis of the Pel2.5kb GFP construct The Pel2.5kb pelvic fin specific enhancer element (Chan et al., 2010) was tested in wild- type and pelvic finless zebrafish to determine if it could drive pelvic fin specific expression of GFP in zebrafish.

The Pel2.5kb-GFP construct was injected and integrated into the zebrafish genome by microinjection with transposase mRNA. The injected embryos were screened for the presence of a cardiac GFP reporter and raised to adulthood. To create wild-type zebrafish Pel2.5kb-GFP lines, transgenic founders were identified and crossed to wild-type zebrafish. The resulting offspring were then examined for the expression of GFP (n=100). To create pelvic finless Pel2.5kb-GFP lines, wild-type background transgenic founders were crossed to pelvic finless zebrafish, to create heterozygous transgenic offspring carrying the pelvic finless genomic sequence variant. These offspring were screened for the presence of the cardiac GFP reporter and were crossed again to pelvic finless zebrafish. These fish were screened for the absence of pelvic fins and those identified were then incrossed again. The resulting pelvic finless Pel2.5kb-GFP zebrafish were then examined for the expression of GFP in the pelvic regions (n=10).

8.2.3. Tbx4 rescue constructs 8.2.3.1. Generation of Tbx4 rescue constructs. In order to generate the wild-type tbx4 rescue construct (pDestTol2pA2- Pel2.5kb-Gata2- Tbx4WT-pA) and the pelvic finless tbx4 construct (pDestTol2pA2- Pel2.5kb-Gata2- Tbx4WT-pA) several constructs needed to be first be generated and combined using MultiSite Gateway® technology (Invitrogen™).

Firstly, three entry clones (the 5’ entry, the middle entry and the 3’ entry) needed to be obtained. For this experiment, the 5’ entry clone needed to contain the Pel2.5b enhancer element and a minimal promoter. As the construct described in Figure 8.1 was difficult to work with, it was decided to re-engineer this construct. A DNA fragment containing the Pel2.5kb enhancer element (Chan et al., 2010) and a Gata2 minimal promoter (Yang et al., 2007) was ordered from GeneArt® (Invitrogen™, Life Technologies™) alongside sub- cloning of this DNA fragment into the SalI and BxtXI sites of the p5’E-MCS entry vector (Kwan et al., 2007). This became known as p5’E-MCS-Pel2.5kb-Gata2.

A middle entry clone containing either the wild-type or pelvic finless coding region of tbx4 cDNA was created. The wild-type and pelvic finless tbx4 cDNAs were created by amplification of pelvic region derived cDNA using gene specific primers to the tbx4 coding 160 region. The forward primer was designed to an area just outside of the tbx4 coding region 5’- GCGTTCTGGAATGATATGGAGG-3’. The reverse primer was designed to an area just outside of the tbx4 coding region 5’-CAGTTGTTGCTCTGAGAGAG-3’. The reaction was amplified according to the high-fidelity DNA polymerase protocol outlined in 3.2.6 under standard conditions in order to create blunt ended PCR products. The PCR products were then phosphorylated according to the protocol outlined in 3.2.10.

The tbx4 cDNAs were then blunt-end cloned into the EcoRI cloning sites of a pME-MCS vector (Kwan et al., 2007). The pME vector was digested with EcoRI, blunted and dephosphorylated according to protocols outlined in 3.2.8, 3.2.9 and 3.2.11. The tbx4 cDNAs were ligated into the pME vector according to 3.2.12. The plasmids containing the correct inserts were then propagated according to protocols outlined in 3.2.15 and 3.2.16. These became known as pME-Tbx4WT or pME-Tbx4pFL.

The p5’E-MCS-Pel2.5kb-Gata2 construct was then combined with either the pME- Tbx4WT or pME-Tbx4pFL constructs and a 3’ entry clone with a poly-A signal described by Kwan et al., 2007 (p3’E-pA) into a pDestTol2pA2 destination vector with an α-crystallin GFP reporter (Kwan et al., 2007) using MultiSite Gateway® technology according the protocol described in 3.2.14 and the resulting plasmids containing the correct inserts were propagated according to protocols outlined in 3.2.15 and 3.2.16. The plasmids were then checked by digestion with BamHI according to 3.2.8 and PCR amplification of tbx4 according to 3.2.6 for the presence of the correct inserts. The final constructs are referred to as either the wild-type tbx4 construct (pDestTol2pA2- Pel2.5kb-Gata2-Tbx4WT-pA) or the pelvic finless tbx4 construct (pDestTol2pA2- Pel2.5kb-Gata2-Tbx4WT-pA) (Figure 8.2).

161 Figure 8.2. The pDestTol2pA2-Pel2.5kb-Gata2-Tbx4-pA rescue construct. Diagram of the pDestTol2pA2-Pel2.5kb-Gata2-Tbx4-pA rescue construct.

162 8.2.3.2. Analysis of rescue experiments Pelvic finless individuals were injected with either wild-type or pelvic finless tbx4 rescue constructs to determine if the expression of wild-type tbx4, and not pelvic finless tbx4, could restore pelvic fin development. The rescue constructs were injected into pelvic finless

embryos which originated from in-crosses of the F2 pelvic finless zebrafish utilised during positional cloning (see 6.3.1). 1-cell embryos from the same clutch were injected at either a low dose (20 pg) or a high dose (40 pg) of either the wild-type of pelvic finless tbx4 rescue constructs or left as uninjected controls. The low dose injections were screened at 3 wpf. The high dose experiments were performed on four separate clutches. The embryos were then raised until 8 wpf and screened for the presence or absence of pelvic fins under low power bright field illumination microscopy (n=150) and compared to uninjected individuals (n=61). Individuals were subsequently genotyped as described in 3.2.5 to ensure that they were of the pelvic finless background.

163 8.3. Results Several attempts to rescue the pelvic finless phenotype with wild-type, and not pelvic finless, tbx4 were undertaken during this project. It was determined that the best way to rescue the phenotype would be transgenically, with either a pelvic fin specific promoter or enhancer driving either wild-type of pelvic finless tbx4 in the developing pelvic regions of pelvic finless zebrafish. The progress made towards transgenically rescuing the pelvic finless phenotype is described in this section.

8.3.1. Pel2.5kb enhancer drives GFP expression in zebrafish pelvic fins In order to identify a pelvic fin specific driver element that could be utilised in this project, the stickleback pelvic fin specific enhancer element, Pel2.5kb (Chan et al., 2010) was tested to examine its ability to drive GFP expression in the pelvic fins of zebrafish. Therefore, wild- type Pel2.5kb-GFP and pelvic finless Pel2.5kb-GFP transgenic lines were created and examined.

In wild-type Pel2.5kb-GFP transgenic lines, GFP expression was observed in embryos and in the developing pelvic fins (Figure 8.3, Figure 8.4). At 24 hpf, no GFP expression was observed in the Pel2.5kb-GFP transgenic embryos (Figure 8.3). However, at 48 hpf, GFP expression from the cardiac GFP reporter was observed and faint GFP expression was observed in the developing pectoral fin buds of wild-type Pel2.5kb-GFP embryos (Figure 8.3). At 5 dpf, observations revealed that GFP expression from the cardiac GFP reporter continued alongside GFP expression in the (Figure 8.3).

Later examination of the wild-type Pel2.5kb-GFP transgenic lines revealed that the Pel2.5kb enhancer element could drive GFP expression in the developing pelvic fins of these zebrafish. In the wild-type Pel2.5kb-GFP zebrafish, the earliest expression of GFP in the pelvic regions was observed in the pelvic fin region of the flank of the fish just prior to the onset of pelvic fin development at 20 dpf (Figure 8.4). At 25 dpf, GFP expression was observed in the developing pelvic fins of wild-type Pel2.5kb-GFP zebrafish (Figure 8.4). Observation at 32 dpf revealed that GFP continued to be expressed in the pelvic fins during the later pelvic fin development (Figure 8.4). Additionally, the pelvic finless Pel2.5kb-GFP lines were also examined at pelvic fin development stages for the presence of GFP. Examination of the pelvic finless Pel2.5kb-GFP transgenic lines revealed that GFP was also expressed in the pelvic fin regions of pelvic finless zebrafish in the correct location and as the pelvic fins were formed (Figure 8.5). Therefore, the stickleback pelvic fin specific enhancer element, Pel2.5kb, could be utilised as a driver element during transgenic rescue experiments. 164 Figure 8.3. Wild-type Pel2.5kb-GFP embryos Bright-field and GFP fluorescence microscopy images of the wild-type Pel2.5kb-GFP at embryonic stages. At 24 hpf, there is no observable expression of GFP. At 48 hpf, GFP expression is present in the heart and the pectoral fin buds. At 5 dpf, GFP expression is present in the heart and the eye. Sites of GFP expression are marked by the white arrowheads. Scale bar: 0.1 mm

165 Figure 8.4. The developing pelvic fins wild-type Pel2.5kb GFP zebrafish GFP fluorescence microscopy images of the developing pelvic fins of wild-type Pel2.5kb-GFP zebrafish at 20 dpf, 25 dpf and 32 dpf. The Pel2.5kb element drives GFP expression in the developing pelvic fins of wild-type zebrafish. Sites of GFP expression are marked by the white arrowheads and the outline of the developing pelvic fin is marked by the dotted line. Scale bar: 0.1 mm

166 Figure 8.5. The developing pelvic regions of wild-type and pelvic finless Pel2.5kb-GFP zebrafish GFP fluorescence microscopy images of the developing pelvic regions of wild-type and pelvic finless Pel2.5kb- GFP zebrafish at 32 dpf. The Pel2.5kb element drives GFP expression in the developing pelvic fins of both wild-type and pelvic finless zebrafish Sites of GFP expression are marked by the white arrowheads. Scale bar: 0.5 mm

167 8.3.2. Transgenic rescue experiments Early observations of individuals injected with either the wild-type or pelvic finless tbx4 rescue constructs revealed that the α-crystallin GFP reporter was not functioning in any injected fish (n=150). This made subsequent analysis difficult as it was impossible to identify fish which had been successfully injected with the rescue constructs. However, in some cases, individuals injected with the wild-type tbx4 rescue construct showed enhanced, but variable pelvic fin development.

Observations of individuals injected with a low dose (20 ng) of the wild-type tbx4 rescue construct revealed the presence of a large pelvic fin bud in these fish (n=10/20) (Appendix F). The pelvic fin bud was observed on either the right flank (n=4), left flank (n=5) or both flanks (n=1) of the fish, in the pelvic fin regions (Figure 8.6). In contrast, uninjected individuals and individuals injected with a low dose of the pelvic finless tbx4 rescue construct never developed a large pelvic fin bud and maintained the pelvic finless phenotype (n=0/20 and n=0/20 respectively) (Figure 8.6).

168 Figure 8.6. Pelvic fin rescue at low doses Representative bright-field microscopy images of pelvic fin bud rescue after injection with a low dose (20 pg) of rescue construct. Uninjected and individuals injected with the pelvic finless tbx4 rescue construct display only small pelvic bulges. Individuals injected with the wild-type tbx4 rescue construct developed pelvic fin buds. Black arrowhead indicates the presence of a pelvic fin bud in a wild-type tbx4 injected individual. Scale bar: 0.1 mm

169 At higher doses (40 ng), mosaic transgenic individuals injected with the wild-type tbx4 rescue construct showed enhanced, but variable pelvic fin development when compared to siblings injected with pelvic finless tbx4 rescue constructs or uninjected siblings (Figure 8.7 A). A total of 14 out of 61 pelvic finless individuals displayed signs of pelvic fin development when injected with the wild-type tbx4 construct (Figure 8.7 B). These individuals developed pelvic fins on either side of the flank with 8 individuals developing a fin on the left flank, 4 individuals developing a fin on the right flank and 2 individuals developing fins on both sides of the flank. A total of 7 out of 69 pelvic finless individuals displayed signs of pelvic fin development when injected with the pelvic finless tbx4 rescue construct. However, analysis of uninjected pelvic finless individuals revealed a background level of pelvic fin development as 3 out of 41 uninjected pelvic finless individuals also displayed signs of pelvic fin development.

Despite the background level of pelvic fin development, statistical analysis (Pearson’s Chi-squared test, p <0.05) determined that there was a significant difference in the number of individuals possessing pelvic fins after injection with the wild-type tbx4 rescue construct when compared to either the individuals injected with the pelvic finless tbx4 rescue construct (p= 0.048) or the uninjected siblings (p= 0.038). In addition, the individuals injected with the pelvic finless tbx4 rescue construct showed no significant difference in the number of pelvic fins when compared to uninjected individuals (p= 0.618).

170 Figure 8.7. High dose mosaic transgenic rescue results A) A graph generated from data provided in Appendix F of the percentage of individuals with pelvic fin development after injection with wild-type or pelvic finless tbx4 rescue construct. There was a statistically significant difference in the percentage of individuals with pelvic fin development after injection with wild-type tbx4 rescue construct when compared to siblings injected with pelvic finless tbx4 rescue construct or uninjected siblings. Un: Uninjected siblings; WT/Tbx4 WT: siblings injected with wild-type tbx4 rescue construct; pFL/Tbx4 pFL: siblings injected with pelvic finless tbx4 rescue construct. B) A representative image of rescued pelvic fin development in a mosaic transgenic wild-type tbx4 rescued individual. Scale bar: 1 mm

171 8.4. Discussion 8.4.1. Overview of results Ongoing strategies to rescue the pelvic finless phenotype with wild-type tbx4 driven from a transgenic element have so far shown variable results. Mosaic transgenic rescue attempts revealed a statistically significant increase in the presence of pelvic fins in individuals injected with the wild-type tbx4 rescue construct when compared to control injected and uninjected siblings. However, these results must be assessed with caution as the α-crystallin GFP reporter was not observed to function in any injected individuals and some background level of pelvic fin development was observed in uninjected individuals (discussed in more detail in section 8.4.2).

8.4.2. Limitations of rescue experiments To determine if the identified candidate mutations truly underlie the pelvic finless phenotype, it is necessary to either replicate or rescue the loss of pelvic fins in pelvic finless zebrafish. As the pelvic finless phenotype does not become apparent until around 3 wpf, traditional phenocopy and rescue experiments utilising morpholinos and mRNA injections were not possible for this project. Therefore, a non-traditional strategy of transgenic rescue was adopted (Takeuchi et al., 2010). The aim was to utilise a pelvic fin specific promoter/ enhancer element to drive expression of tbx4 in the pelvic regions to investigate if pelvic fin development could be restored. However, if the actual underlying genetic lesion was upstream of tbx4, the restoration of Tbx4 activity could inadvertently rescue the phenotype. Therefore, the experiment was controlled by examining whether the restoration of wild-type and not pelvic finless Tbx4 could rescue the pelvic finless phenotype.

A pelvic fin specific enhancer element was utilised to transgenically drive the expression of tbx4 in the pelvic regions. This experimental design is the first limitation of the rescue experiments presented in this chapter. Although the Pel2.5kb pelvic fin enhancer element can drive GFP expression in a similar expression pattern as is seen for tbx4, is not the endogenous enhancer or promoter of tbx4. In addition, in first generation transgenic individuals, there will only be mosaic expression of the injected construct as it will not integrate into every cell (Stuart et al., 1990). Therefore, in the described experiments the expression of tbx4 is driven in a slightly different expression pattern than endogenous tbx4 and in only a small subset of cells. Although the development of pelvic fins has been reported to be rescued by transgenic mosaic expression (Chan et al., 2010), it is possible that

172 these limitations could account for the relatively low level of rescued pelvic fin development observed in these experiments.

Another limitation of the rescue experiments presented in the chapter was the absence of the α-crystallin GFP reporter in any of the injected individuals. It is possible that either silencing of this reporter occurred or a small mutation in the GFP DNA sequence had accumulated over time (Goll et al., 2009; Akitake et al., 2011). It was not possible to test for silencing of the reporter before the injection of the construct into the fish, but had the constructs been engineered with a GFP tag on the Tbx4 protein, successfully injected individuals could have been identified. However, as the experiment was conducted, the absence of the α-crystallin GFP reporter resulted in the inability to exclude unsuccessfully injected individuals. Therefore, these individuals were included in the treatment groups thought to harbour the rescue constructs. The presence of unsuccessfully injected individuals in the rescue groups was another limitation which could account for the relatively low level of rescued pelvic fin development observed in these experiments.

A further limitation was the incomplete penetrance of the pelvic finless phenotype. At this stage it is uncertain if the pelvic finless individuals which developed pelvic fins due the introduction of wild-type tbx4 or if these individuals naturally developed pelvic fins. In addition, as these rescue experiments were conducted on WIK background pelvic finless zebrafish and these fish were shown to exhibit a higher degree of incomplete penetrance of the phenotype than pelvic finless zebrafish on the golden background (4.2.6), it is possible that all ‘rescued’ individuals may have naturally developed pelvic fins. Although beyond the scope of this project, to successfully prove that the restoration of wild-type, and not pelvic finless, Tbx4 activity can rescue pelvic fin development in pelvic finless zebrafish, additional rescue experiments are necessary. To better conduct these experiments, it would be prudent to tag the Tbx4 proteins with a florescent protein so that the ectopic expression of Tbx4 could be observed and conduct the experiments on pelvic finless zebrafish of the golden background. Although tagging with a florescent protein may interfere with protein function, Takeuchi et al., 1999, demonstrated that limb development can be induced in chicks utilising a C-terminal tagged TBX4 protein. However, it would be prudent to experiment with N versus C-terminal tagging or incorporating a site between TBX4 and the florescent protein. It could also prove beneficial to drive tbx4 expression from its endogenous promoter or enhancer elements, however at the time of writing, these remain unknown.

173 Using these strategies, mosaic transgenic rescue may restore pelvic fin development, but stable transgenic lines should also be created to increase the chances of pelvic fin rescue. In addition, to determine which of the identified candidate mutations truly underlies the pelvic finless phenotype, it would be necessary to test each individual candidate mutation to determine which amino acid residue is essential for the restoration of normal Tbx4 protein function. Traditionally, rescue experiments of mutant zebrafish are conducted in combination with phenotype replication experiments in order to gain proof that a candidate genetic lesion truly underlies the mutant phenotype (Nasevicius and Ekker, 2000). As traditional morpholinos cannot be used to target a phenotype which occurs at such a late stage of development (Nasevicius and Ekker, 2000), traditional phenotype replication experiments are not possible for pelvic finless zebrafish. However TALENs, which are an emerging technology, could be utilised to determine if genetic lesions in tbx4 can replicate the pelvic finless phenotype. A TALEN can modify a genomic sequence of interest and thus cause selected genomic lesions in a gene of interest, resulting in an impaired protein of interest (Sander et al., 2011). Such a TALEN could be designed to the tbx4 sequence to generate a Tbx4 null zebrafish in order to determine if the absence of Tbx4 activity produces a similar phenotype to pelvic finless zebrafish. In addition, in order to obtain genetic proof that the identified candidate mutations in tbx4 underlie the pelvic finless phenotype, a complementation cross could be performed between the pelvic finless zebrafish and the TALEN injected individuals. This would involve crossing homozygous individuals harbouring the TALEN induced genetic lesion to homozygous pelvic finless zebrafish. If the resulting offspring lacked pelvic fins, and the other additional experiments also pointed towards the identified candidate mutations in tbx4, this would provide unambiguous proof that the identified candidate mutations truly underlie the pelvic finless phenotype.

174 9. Final discussion and concluding remarks 9.1. Overview of results Characterisation of pelvic fin development in both wild-type and pelvic finless zebrafish has been achieved, while significant progress has been made towards identifying the genomic lesion responsible for the pelvic finless phenotype. Positional cloning initially identified a 12Mb region on chromosome 15 that harbours the pelvic finless locus. In this region, bioinformatic analysis after deep sequencing has identified three specific candidate missense mutations that occur in tbx4 to be the likely candidates for the cause of pelvic fin loss in pelvic finless zebrafish. Further analysis of mosaic transgenic zebrafish that express zebrafish tbx4 under the control of the stickleback Pel enhancer provided additional evidence that these missense candidate mutations are a possible cause of pelvic fin loss as replacement of wild-type, and not mutated tbx4, was shown to induce enhanced but variable pelvic fin development in pelvic finless zebrafish.

9.2. Tbx4 Tbx4 belongs to the T-box family of transcription factors that contain a conserved DNA- binding motif known as the T-box domain (Papaioannou, 2001). Throughout the development of many organisms, members of the T-box family are expressed dynamically and are vital for many developmental processes. Mutations in the T-box genes cause developmental defects in a range of organisms ranging from C. elegans to humans (Papaioannou, 2001). Of particular interest for this project is the T-box gene Tbx4 which is part of the T-box transcription factor 2 (Tbx2) subfamily consisting of Tbx2, T-box transcription factor 3 (Tbx3), Tbx4 and Tbx5 (Ruvinsky et al., 2000). In vertebrates, these genes are highly conserved and are arranged as two pairs of linked genes (Tbx3/Tbx5 and Tbx2/Tbx4) which are thought to have arisen from an ancestral tandem duplication followed by duplication and dispersion of the two gene clusters (Agulnik et al., 1996). Mutations in the first cluster of genes, Tbx3/Tbx5 have been shown to cause human development disorders. Ulnar-mammory syndrome is caused by mutations in TBX3 (Bamshad et al., 1997) and Holt- Oram syndrome is caused by mutations in TBX5 (Basson et al., 1999; Li et al., 1997). Mutations in the second cluster of genes Tbx2/Tbx4 have also been shown to cause developmental disorders as it has been shown that mutations in TBX4 cause lower limb deformities in humans (Alvarado et al., 2010; Ballif et al., 2010; Bongers et al., 2001; Bongers et al., 2002; Bongers et al., 2004; Lu et al., 2012; Mangino et al., 1999; Wang et al.,

175 2010) and it is the lower limb phenotype arising from TBX4 mutations that make this gene particularly interesting when examining pelvic finless zebrafish.

Numerous studies have shown that tbx4 is involved in hindlimb/pelvic fin development. Tbx4 was first described in limb development as being specifically expressed in hindlimbs and not forelimbs (Chapman et al., 1996; Gibson-Brown et al., 1996). This expression pattern is thought to be evolutionarily conserved in vertebrates with such diverse species as mouse, chick, opossum, zebrafish and shark showing similar Tbx4/tbx4 expression patterns (Gibson-Brown et al., 1996; Gibson-Brown et al., 1998; Tamura et al., 1999; Ruvinsky et al., 2000; Tanaka et al., 2002; Horton et al., 2008; Keyte and Smith, 2010). While it is known that Tbx4/tbx4 is expressed during hindlimb/pelvic fin development, direct evidence for the essential role of Tbx4 in hindlimb development has been demonstrated by Tbx4 knockout mice and TBX4 misexpression studies in chicks.

Direct evidence for the essential role of TBX4 in hindlimb development has been provided in chicks as TBX4 has been shown to be sufficient for hindlimb bud induction in this species (Takeuchi et al., 2003). Misexpression of TBX4 in the flank of chicks at E10 induces an additional limb (Takeuchi et al., 2003). The additional limbs where shown to express the key hindlimb development markers WNT8C, HOMEOBOX B9 (HOXB9), HOMEOBOX C9 (HOXC9), FGF10 AND FGF8 and posses partial hindlimb-like identity (Takeuchi et al., 2003). Although the role of TBX4 in determining hindlimb identity suggested in this study remains controversial (Duboc and Logan, 2011; Ouimette et al., 2008; Takeuchi et al., 2003), this study did demonstrate that during the early stages of hindlimb bud induction, TBX4 is sufficient to induce hindlimb bud induction in chicks (Takeuchi et al., 2003).

In addition, genetic manipulation experiments conducted in mice have shown that Tbx4 is necessary for proper hindlimb development. Mice in which Tbx4 has been deleted do not develop proper hindlimbs. It has been demonstrated that Tbx4 null mice do initiate hindlimb development, as a tiny hindlimb bud is observed, however it is drastically reduced in size and although hindlimb buds of these mice display early faint Fgf10 expression, it is maintained at later stages (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). It has been proposed that the hindlimbs of these mice fail to grow due to improper limb initiation and the failure to maintain Fgf10 expression (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). This study provided direct evidence that Tbx4 is necessary for hindlimb development in mice.

176 9.2.1. Is Tbx4 critical for pelvic fin development? The role of Tbx4 in hindlimb development is well established in tetrapods and all studies conducted to date agree that Tbx4 is necessary for hindlimb development and that this function is likely to be conserved amongst gnathostomes (Chapman et al., 1996; Duboc and Logan, 2011; Gibson-Brown et al., 1996, Gibson-Brown et al., 1998; Horton et al., 2008; Keyte and Smith, 2010; Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007; Ruvinsky et al., 2000; Tamura et al., 1999; Tanaka et al., 2002). However, in teleosts, the role of tbx4 in pelvic fin development is less well characterised, but there is a large body of literature that suggests that tbx4 could be vital for pelvic fin development.

As previously mentioned, the sequence and expression pattern of Tbx4 is highly conserved amongst vertebrates (Chapman et al., 1996; Gibson-Brown et al., 1996, Gibson-Brown et al., 1998; Ruvinsky et al., 2000; Tamura et al., 1999). In particular, the amino acid sequence of T-box DNA binding domain of Tbx4 is very similar between zebrafish and tetrapods (Agulnik et al., 1996; Chapman et al., 1996; Isaac et al., 1998; Gibson-Brown et al., 1996, Gibson-Brown et al., 1998; Ruvinsky et al., 2000; Simon et al., 1997; Tamura et al., 1999). In addition, the spatiotemporal expression pattern of tbx4 in the developing pelvic fins is very similar to that observed for Tbx4 in the developing hindlimbs of tetrapods (Gibson-Brown et al., 1996, Gibson-Brown et al., 1998; Horton et al., 2008; Keyte and Smith, 2010; Ruvinsky et al., 2000; Tamura et al., 1999, Figure 5.3). Due to this high degree of conservation of the DNA binding domain and the shared expression patterns, it is likely that the function of tbx4 is also conserved between tetrapods and zebrafish. Therefore, it is likely that tbx4 plays an important role during pelvic fin development in zebrafish.

9.2.2. Missense mutations in tbx4 During this project, three candidate missense mutations were identified in the tbx4 sequence of pelvic finless zebrafish. These three candidate missense mutations were shown to occur in the T-box DNA binding domain of tbx4 and cause two amino acid substitutions, A78V and G79A. It has been previously shown that missense mutations in the T-box DNA binding domain of human TBX4 cause the lower limb deformities of small patella syndrome (Bongers et al., 2004). The p.Q62X, p.G248V and p.Q531R amino acid substitutions have all been shown to be linked to small patella syndrome (Bongers et al., 2004). These amino acid substitutions are thought to interfere with the DNA binding capacity of the human TBX4 protein and thus cause the lower limb deformities observed in small patella syndrome (Bongers et al., 2004).

177 In addition it has been shown that amino acid substitutions at the equivalent residues of human TBX5 underlie familiar cases of Holt-Oram syndrome which presents with upper limb deformities (Basson et al., 1999; Fan et al., 2003; Stirnimann et al., 2010). A missense mutation in human TBX5 which results in a Glycine to Arginine amino acid substitution at residue 80 of human TBX5 (the equivalent of residue 79 in zebrafish Tbx4) has been shown to be linked to patients with Holt-Oram syndrome (Basson et al., 1999; Fan et al., 2003; Stirnimann et al., 2010). The G80R amino acid substitution has been shown to affect the DNA binding affinity of human TBX5 to its target DNA sequence and it has been suggested that this is the cause of the upper limb deformities observed in Holt-Oram syndrome (Basson et al., 1999; Fan et al., 2003; Stirnimann et al., 2010).

It is therefore possible that missense mutations in pelvic finless tbx4 which are predicted to result in an amino acid substitution in the T-box DNA binding domain of pelvic finless Tbx4 protein could also affect DNA binding affinity of the Tbx4 protein and subsequently disrupt pelvic fin development in pelvic finless zebrafish. The likely role of tbx4 in pelvic fin development (see 9.2.1) in combination with the morphological and genetic characterisation of pelvic finless zebrafish presented in this thesis support the hypothesis that the candidate missense mutations identified in pelvic finless tbx4 underlie the loss of pelvic fins in pelvic finless zebrafish.

9.2.3. A hypomorphic allele or compensation? In the light of all the evidence pointing towards the loss of function of Tbx4 in pelvic finless zebrafish, it does seem possible that pelvic finless zebrafish either harbour a hypomorphic tbx4 allele or undergo compensation for the loss of Tbx4, to account for the incomplete penetrance of the phenotype. Either hypothesis is plausible as pelvic finless zebrafish are viable and some pelvic finless individuals do undergo pelvic fin development.

A minority of pelvic finless zebrafish were shown to undergo partial pelvic fin development with one or two, normal or abnormal pelvic fins observed in some individuals. It is therefore possible that the loss of Tbx4 activity could be compensated by the activity another member of the T-box family which is expressed during pelvic fin development. This is similar to a situation suggested in another model of pelvic fin loss in some populations of threespine stickleback fish (Chan et al., 2010; Cole et al., 2003; Shapiro et al., 2004). It has been shown that pelvic fin loss in certain populations of threespine is caused by a lack of Pitx1 activity during pelvic fin development (Chan et al., 2010; Cole et al., 2003; Shapiro et al., 2004). Some of these individuals do display partial or complete pelvic fin development

178 and it has been suggested that compensatory activity by Pitx2 may be responsible for the restoration of pelvic fin development in these individuals (Cole et al., 2003; Shapiro et al., 2004). However, the only additional T-box genes examined in the context of zebrafish pelvic fin development, tbx2 and tbx5, have been shown to be absent during pelvic fin development (Ruvinsky et al., 2000, Figure 5.4). It is possible that another member of the T-box family could compensate for the loss of tbx4 activity in the pelvic regions of pelvic finless zebrafish, but at the time of writing there is no published evidence supporting this hypothesis.

It seems more likely that the Tbx4 A78V G79A candidate mutation in pelvic finless zebrafish could result in a hypomorphic Tbx4 protein with a partial loss of function. This conclusion is based on evidence gained during this study, and the literature surrounding Tbx4 null mice and human cases of TBX4 mutations. Tbx4 null mice are embryonic lethal, due to the necessary function of Tbx4 in other developmental processes, such as and allantois development and vasculogenesis (Naiche and Papaioannou, 2003). In contrast, pelvic finless zebrafish and humans carrying TBX4 missense mutations develop into adults and display a specific phenotype of lower limb or pelvic fin defects (Alvarado et al., 2010; Ballif et al., 2010; Bongers et al., 2001; Bongers et al., 2002; Bongers et al., 2004; Lu et al., 2012; Mangino et al., 1999; Wang et al., 2010). This suggests that if the candidate missense mutations identified in tbx4 of pelvic finless zebrafish truly underlie the pelvic finless phenotype, they could result in a hypomorphic protein which retains partial function. This line of evidence suggests that the retained partial function of the Tbx4 protein would be sufficient for survival and incomplete penetrance of the phenotype, but would normally result in the arrested development of the pelvic fins of zebrafish or the lower limb deformities observed in human patients.

9.2.4. Could mutations in tbx4 cause the loss of the apical ectodermal thickening in pelvic finless zebrafish? It has been shown that pelvic finless zebrafish lack a morphologically and genetically distinct apical ectodermal thickening (Chapters 4 and 5) and it has been previously shown that the disruption of the apical ectodermal thickening (in fish) or the AER (in tetrapods) causes a failure of limb/fin development (Boulet et al., 2004; Crossley et al., 1996; Lewandoski et al., 2000; Moon and Capecchi, 2000; Barrow et al., 2003; Narita et al., 2005, Narita et al., 2007; Naiche and Papaioannou, 2003; Min et al., 1998; Sekine et al., 1999; Fischer et al., 2003; Norton et al., 2005). In additional, it has been shown that although pelvic finless zebrafish display early tbx4 expression in the developing pelvic regions, there is

179 no evidence of fgf10a expression. This is important as the absence of Fgf10 has been shown to result in the failure of the apical ectodermal thickening/AER (Min et al., 1998; Sekine et al., 1999; Norton et al., 2005) and therefore the absence of fgf10a in the pelvic regions of pelvic finless zebrafish could cause the failure of the pelvic fin apical ectodermal ridge and subsequently cause the arrested development of the pelvic fins in pelvic finless zebrafish. However, the question remains whether and how the candidate missense mutations identified in tbx4 could cause the lack of fgf10a expression in pelvic finless zebrafish.

A multitude of evidence indicates Fgf10 expression is dependent on Tbx4. It has been previously demonstrated that Tbx4 directly influences Fgf10 expression as Fgf10 is a likely direct target of the Tbx4 gene product (Duboc and Logan, 2011; Ouimetter et al., 2008). It has been shown that there are predicted T-box binding sites located in the Fgf10 promoter region and Tbx4 is capable of activating reporter gene expression due to binding to these sites in vitro (Duboc and Logan, 2011; Ouimette et al., 2008). In addition, tbx4 has been shown to activate reporter gene expression downstream of the mouse Fgf10 promoter region (Duboc and Logan, 2011). Other studies argue that, in mice, the onset of Fgf10 expression may be Tbx4 independent, while the maintenance of Fgf10 expression is Tbx4 dependent. Mouse Tbx4 knockout studies have shown that in the absence of Tbx4 activity, the beginnings of hindlimb buds are able to form, but these show abnormal Fgf10 expression and do not continue development in vivo or in vitro (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). The observation of any Fgf10 expression in the absence of Tbx4 activity suggested to the authors that Tbx4 activity may not be the sole regulator of Fgf10 expression in the mouse hindlimb bud (Naiche and Papaioannou, 2003; Naiche and Papaioannou, 2007). The authors hypothesised that other regulators of the onset of Fgf10 expression are responsible for the presence of Fgf10 expression in the mutant hindlimbs, but that Tbx4 activity is required to achieve the threshold levels of Fgf10 expression that are required for subsequent limb development. Recent studies in zebrafish pectoral fins provide evidence of other regulators of the onset of fgf10 expression (Figure 9.1). In zebrafish pectoral fins, there is no evidence that tbx5 binds to the fgf10 promoter and directly influences fgf10 expression (Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006). In addition, there appears to be an additional level of signalling required to regulate the genetic pathway leading from tbx5 to fgf10a in zebrafish pectoral fins. These signalling pathways have been shown to involve at least four other genes aside from tbx5; fgf24, sall4/sall1a, and prdm1 and interestingly, all of these genes have been shown to play a role in pectoral fin bud compaction 180 during development and are necessary for the regulation of fgf10 expression in zebrafish pectoral fin buds (Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006). Limb/fin bud compaction occurs due to cell proliferation and possibly cell migration from the lateral plate mesoderm to the limb/fin producing region, however this process is more pronounced in zebrafish than in tetrapods (Fischer et al., 2003; Mercader, 2007). The paired fins of teleosts may have needed to evolve a separate level of genetic signalling necessary for fin bud compaction and fgf10 expression as the lateral plate mesoderm is much thinner in zebrafish than in tetrapods (Mercader, 2007). It remains unknown if this additional level of signalling is necessary for pelvic fin development in zebrafish, however, these pathways must be considered when investigating if the candidate missense mutations identified in pelvic finless tbx4 could result in the loss of fgf10a expression.

In zebrafish pectoral fins, it is thought that tbx5 induces the expression of fgf10 indirectly through the regulation of fgf24 activity via sall4/sall1a and prdm1 (Figure 9.1) (Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006). Fgf24 is the first fibroblast growth factor (FGF) expressed in the mesenchyme of developing pectoral fins and its activation is necessary for the induction of fgf10 expression (Fischer et al., 2003; Harvey and Logan, 2006). This is known from studies of mutant fgf24 zebrafish, where fgf24 activity is ablated, due to a truncation mutation, and there is no evidence of fgf10 expression (Fischer et al., 2003; Harvey and Logan, 2006). In zebrafish pectoral fins, the induction of fgf10 by fgf24 is thought to be under the control of Tbx5 activity via, at least, two separate pathways, the sall4/sal-like 1a (sall1a) pathway and the prmd1 pathway (Harvey and Logan, 2006; Mercader et al., 2006).

Tbx5 and sall4/sall1a ultimately regulate the induction of fgf10 expression via both the regulation of fgf24 and its receptor fgfr2 (Figure 9.1) (Harvey and Logan, 2006). In the developing zebrafish pectoral fin, fgf24 signals to fgf10 through the fgfr2 receptor (Harvey and Logan, 2006). It is through this pathway that tbx5 is able to regulate the expression to fgf10 in two ways. Firstly, tbx5 is able to directly induce the expression of fgf24 and thus control the levels of fgf24 available in the developing pectoral fin bud (Fischer et al., 2003; Harvey and Logan, 2006). A second level of regulation is present through the tbx5 controlled induction of sal4 and sal1a, which are thought to be two transcriptional activators which are able to directly regulate the expression of fgfr2 (Fischer et al., 2003; Harvey and Logan, 2006). In

181 this way, tbx5 is able indirectly regulate the induction of fgf10 by directly regulating the levels of fgf24 and the levels the receptor through which fgf24 signals to fgf10.

In addition to tbx5 regulation of the induction of fgf10 via fgf24, a feedback loop involving prdm1 adds another level of regulation to the onset of fgf10 expression in zebrafish pectoral fins (Figure 9.1). It has previously been shown that prdm1 is necessary for pectoral fin development in zebrafish (Lee and Roy, 2006; Mercader et al., 2006). Studies of zebrafish pectoral fins have shown that the onset of prdm1 expression is regulated by fgf24 activity in the fin bud mesenchyme, but that subsequently, fgf24 expression quickly becomes prdm1 dependent (Mercader et al., 2006). It has additionally been shown that the induction of fgf10 expression is dependent on the fgf24/prdm1 positive feedback loop in the developing pectoral fin bud (Lee and Roy, 2006; Mercader et al., 2006). These results suggest that fgf24, sall4/sal1la and prdm1 all work together to regulate the induction of fgf10 by tbx5. Therefore, tbx5 induced fgf10 expression in the developing zebrafish pectoral fin bud mesenchyme is regulated by at least three additional pathways and that these pathways should be considered when investigating the link between candidate missense mutations in tbx4 and the loss of fgf10a expression.

Although an additional level of mesenchymal FGF signalling has not been described in tetrapod limb development, studies from zebrafish indicate that these pathways are necessary for the induction of fgf10 in the developing pectoral fins and that they may also be necessary for the development of zebrafish pelvic fins. When considering an additional level of FGF signalling in zebrafish pelvic fins, it is important to note that homozygous fgf24 mutant zebrafish are viable and develop normal pelvic fins (Fischer et al., 2003). As zebrafish fgf24 mutants result from a mutation in the coding region of fgf24, and not a region specific regulatory enhancer, this indicates that either the additional level of FGF signalling involving fgf24 is not necessary in pelvic fins or that a different FGF family member may play the role of fgf24 in pelvic fins (Fischer et al., 2003).

It is therefore possible that the candidate mutations in tbx4 of pelvic finless zebrafish may directly cause the loss of fgf10 expression by the disruption of direct binding to the fgf10 promoter sequence or through a previously undescribed regulatory network that operates during pelvic fin development and is necessary for apical ectodermal ridge formation and the subsequent development of pelvic fins.

182 Figure 9.1. A diagram of the pathways leading to the onset of fgf10 expression The pathway leading to the onset of fgf10 expression in the developing zebrafish pectoral fin bud is complex and involves multiple levels of regulation. It has been suggested that tbx5 regulates the induction of fgf10 expression via two different pathways by regulating the expression of fgf10 itself and the levels of its receptor fgfr2 through interactions with sall4/1a, fgf24 and prdm1 (Fischer et al., 2003; Harvey and Logan, 2006; Lee and Roy, 2006; Mercader et al., 2006). These pathways remain undescribed in zebrafish pelvic fins, but it is possible that similar pathways exist. In zebrafish pelvic fins fgf24 may be replaced by an unknown FGF family member. However, in zebrafish pelvic fins it is also possible that Tbx4 directly regulates Fgf10 due to its ability to bind the Fgf10 promoter sequence (Duboc and Logan, 2011).

183 9.2.5. Implications of a viable mutant Tbx4 model If the mutations in tbx4 underlie the pelvic finless phenotype, then this could provide a foundation for future studies of pelvic fins and hindlimbs in the fields of limb development and evolutionary biology. Pelvic finless zebrafish could provide a base for future pelvic fin and hindlimb development studies as these fish could represent a loss of Tbx4 activity specifically during limb/fin development in a viable model organism.

In other model organisms, the complete absence of Tbx4 activity is lethal at early embryonic stages. The first mouse Tbx4 knockout models were embryonic lethal at E10.5, which prevented later in vivo investigation of the hindlimb phenotype (Naiche and Papaioannou, 2003). To overcome this problem, a complicated conditional deletion strategy was employed which effectively knocked-out Tbx4 from E9.5 (Naiche and Papaioannou, 2007). This strategy allowed later in vivo investigation of the hindlimb phenotype, however, this complicated strategy does not lend itself to high throughput analysis for future experiments. Additional strategies which involved the removal of the Tbx4 hindlimb enhancer element (HLEA) did not result in a failure of hindlimb development, due to the residual Tbx4 expression in the developing hindlimb (Menke et al., 2008). Therefore, if the pelvic finless phenotype is due to mutations in tbx4 which specifically target the hindlimb/pelvic fin function of Tbx4, then pelvic finless zebrafish represent a novel developmental model in which to examine the hindlimb/pelvic fin development cascade. For example, a chromatin precipitation next generation sequencing experiment could be performed on the pelvic regions of wild-type and pelvic finless zebrafish to determine the downstream binding targets of Tbx4. The different binding targets between wild-type and mutant Tbx4 would reveal the promoters and genes necessary for pelvic fin development. Pelvic finless zebrafish could also be utilised as a springboard from which to examine the evolutionary processes of hindlimb/pelvic fin loss. Human mutation studies have identified recurrent microdeletions at the TBX4 locus which demonstrates that this locus is an area of segmental duplication (Ballif et al., 2010). Segmental duplications are known to mediate many kinds of chromosomal mutations, such as Indels, polymorphisms, duplications and inversions and are therefore ‘hot spots’ for chromosome evolution (Koolen et al., 2006; Sharp et al., 2006; Shaw-Smith et al., 2006). As tbx4 is located in a ‘hot spot’ for chromosome evolution, chromosomal mutations at this locus could be responsible for the secondary loss of hindlimbs or pelvic fins in other species and therefore pelvic finless zebrafish could represent a platform from which to investigate the loss of hindlimbs or pelvic fins in other species.

184 The secondary loss of hindlimbs/pelvic fins has occurred repeatedly throughout the evolution of many different lineages, including mammals, reptiles and fish. In some species, hindlimbs are completely absent (Fugu), whereas in others, remnants remain (pythons, whales) (Cohn and Tickle, 1999; Bejder and Hall, 2002; Santini and Tyler, 2003; Tanaka et al., 2005). Further to this, an asymmetry of pelvic fin loss is observed in other species, such as stickleback and manatees (Bell and Orti, 1994; Cole et al., 2003; Shapiro et al., 2004; Shapiro et al., 2006). Pelvic fin loss is documented in one or more species of 92 teleostean families; the pelvic fin has been lost about 50 independent times, excluding multiple losses within families (Nelson, 1993). However, all of these cases of hindlimb/ pelvic fin loss have occurred in non-model organisms. As the loss of pelvic fins in pelvic finless zebrafish has occurred in an established model organism which boasts a range of advantages for the study of limb/fin development, pelvic finless zebrafish represent a real and novel opportunity to examine the role of the tbx4 locus as the cause underlying the evolution of hindlimb/pelvic fin loss in other diverse species. For example, dolphins harbour a similar mutation in TBX4 as pelvic finless zebrafish, a deletion of A92 (which corresponds to A87 in zebrafish). The pelvic finless zebrafish and transgenic pelvic fin rescue experiments could be utilised to determine if the dolphin mutation also results in a less functional TBX4 protein and therefore a possible cause of hindlimb loss in this species. It will be very exciting to determine if the identified candidate mutations in tbx4 underlie the pelvic finless phenotype as this fish has so much potential to reveal insights into the biology and evolution of limb and fin development. 9.3. Conclusion and future directions This project represents an extension to the scientific knowledge of pelvic fin development in zebrafish. Prior to this research, there had never been a genetic analysis of pelvic fin development in zebrafish nor had a pelvic fin specific mutant, such as pelvic finless zebrafish, been previously described and genetically mapped. Although much work remains to be done, this project has identified three candidate missense mutations as the possible cause of pelvic fin loss in pelvic finless zebrafish. The confirmation of one, or all of these candidate mutations in tbx4 as the cause of pelvic fin loss is likely to provide key insights into the function of tbx4 in hindlimb/pelvic fin development. As mutations in tbx4 are such a strong candidate to underlie the pelvic finless phenotype, several key experiments should be performed to determine the role of the identified candidate mutations in pelvic fin loss.

185 These experiments include addtional rescue, phenocopy, protein localisation and luciferase assay experiments. Additional rescue experiments should be performed with a fluorescently tagged Tbx4 protein so that the ecoptic expression of Tbx4 can be observed in real time in the pelvic fins and the generation of these constructs is already underway. In addition, these constructs could be utilised to examine the subcellular localisation of the Tbx4 protein. Experiments to investigate a tbx4 knockdown zebrafish are also already underway in collaboration with our colleague Dr Daniel Hesselson at the Garvan Institute (Sydney). A TALEN targeting tbx4 will create a tbx4 null zebrafish, which may produce a similar phenotype to pelvic finless zebrafish. In addition, this fish will be crossed to pelvic finless zebrafish in a complementation study to unambiguously prove that the identified candidate mutations in tbx4 underlie the pelvic finless phenotype. Finally, to provide mechanistic evidence whether the missense polymorphisms identified in the pelvic finless tbx4 sequence effect the DNA binding function of the Tbx4 protein, a luciferase assay should be performed to determine if mutant Tbx4 is capable of binding its reported downstream target. The constructs necessary for these experiments have been generated and will be sent to our collaborators, under the supervision of Associate Professor Joan Heath at the Walter and Eliza Hall Institute of Medical Research (Melbourne), to conduct the luciferase assay. If these experiments all continued to point towards Tbx4, would then be possible to conclude that the candidate mutations identified in the pelvic finless tbx4 sequence are the cause of pelvic fin loss in zebrafish.

The foundation for utilising the pelvic fins of zebrafish as a novel developmental model has now been established. It is hoped that this study will provide the groundwork for future pelvic fin investigations and for using the zebrafish pelvic fin as a developmental model. It is an exciting time in the field of pelvic fin development and we hope that this research will enable many wne di scoveries tha t will significantly enhance the scientific knowledge in the wide and important field of development.

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214 Appendices

Appendix A

List of suppliers

Ambion® (Mulgrave, Victoria, Australia)

Amresco (Solon, Ohio, USA)

Applied Biosystems (Mulgrave, Victoria, Australia)

Bioland (Paramount, California, USA)

Biomatters (Auckland, New Zealand)

Carl Zeiss (North Ryde, New South Wales, Australia)

Crown Scientific Pty Ltd. (Moorebank, New South Wales, Australia)

David Kopf instruments (Tujunga, California, USA)

General Valve™ corporation (Fairfield, New Jersey, USA)

Genomed (Lӧnhe, Germany)

GraphPad Software (San Diego, California, USA)

Harvard Apparatus (Holliston, Massachusetts, USA)

Illumina (San Diego, California, USA)

Invitrogen™ (Mulgrave, Victoria, Australia)

Jenoptic (Jena, Thuringia, Germany)

Leica (North Ryde, New South Wales, Australia)

Life Technologies™ (Mulgrave, Victoria, Australia)

Microsoft (North Ryde, New South Wales, Australia)

New England Biolabs (NEB) (Ipswich, Massachusetts, USA)

Philips (Hillsboro, Oregon, USA)

Qiagen (Chadstone, Victoria, Australia)

Roche (Castle Hill, New South Wales, Australia)

RocheNimbleGen (Madison, Wisconsin, USA)

215 Sakura (The Netherlands)

Sigma-Aldrich® (Castle Hill, New South Wales, Australia)

Thermo Fisher Scientific (Scoresby, Victoria, Australia)

UVP (Upland, California, USA)

216 List of recipes for buffers and solutions

Solutions and buffers Components

Antibody blocking solution 0.1% DMSO 0.1% FBS in PBS E3 Embryo medium 60 mg/L Instant 0.5% Methylene blue In-situ hybridisation blocking solution 2% FBS 2 mg/ml BSA 1% DMSO in Tris buffer (pH 7.5) Hybridisation buffer 50% Deionised formamide 5x SSC 0.1% Tween 20 1mM EDTA 0.1% CHAPS 20g/L Blocking agent (Roche) 50ug/L Yeast tRNA 100ng/L Heparin Luria Bertani broth 1.0% Tryptone 0.5% Yeast Extract 1.0% Sodium Chloride (NaCl) PBS 0.8% NaCl 0.02% KCl 0.02 M PO4 pH 7.3 PBST 0.1% Tween 20 in PBS Proteinase K solution 2 μg/μl in: 10 mM Tris-HCl (pH 7.5) 20 mM CaCl2 50% glycerol S.O.C 2% tryptone 0.5% yeast extract 10 mM NaCl 2.5 mM KCl 10 mM MgCl2 10 mM MgSO4 20 mM glucose Sodium citrate buffer 10mM Sodium Citrate 0.05% Tween 20 pH 6.0 SSC 150 mM NaCl 15 mM Na3-citrate pH to 7.0

217 TAE buffer 40 mM Tris-base 20 mM Acetic acid 1 mM EDTA TBE buffer 90 mM Tris-base 90 mM Boric acid 2 mM EDTA Tris buffer (pH 7.5) 0.1 M Tris (pH 7.5) 0.1 M NaCl 1% Tween20 Tris buffer (pH 9) 0.1 M Tris (pH 9) 0.1 M NaCl 1% Tween20

218 PCR primers and protocols

Primer name Primer Sequence (5’-3’) PCR conditions z1408 F: AATTCTGAAATTAAGATGTTGACCG SSLP R: GAAAACGCACATCACGTGAC 60ºC anneal z3558 F: GTTATTGGCACCAGCCAGGC SSLP R: TGATAGTCCTGTCGCCGCAA 60ºC anneal z11403 F: TTTGTTTTAAGCATTGCCGA SSLP R: GCATTTTGACTCCACTGGGT 60ºC anneal z4053 F: TGGCGACATCTGCGATGTTT SSLP R: GGAAAGCGTCTGCCTCACGT 60ºC anneal z9154 F: AACCAGGTTTTGGTGTCTGC SSLP R: CGTGACGAGGAGACTGTTCA 60ºC anneal z6661 F: CCTCCAGGATCCTGCAGAAA SSLP R: TTCTCTGACTGCCCTCGG 60ºC anneal z10289 F: CACCATATATGTGTTGTTCTGCA SSLP R: CGACACATCACGCCATTTAC 60ºC anneal z6312 F: TCCTCCTGTGTGTGAACATCA SSLP R: CAGCCATCAGAGAACATCGA 60ºC anneal z6712 F: GACAAATTGCACCCTCACCT SSLP R: GCTCGCAGATGTATTGAAAGG 60ºC anneal z21982 F: TGGAAAAACATGAATGCTGC SSLP R: AACACCTGCTGGGTAAGTGC 60ºC anneal z4396 F: GGGATTGTGGTTCTCCACGC SSLP R: AGGCAGCCCTTTCCTAAAGGC 60ºC anneal z11320 F: ACAGGGTAAGCAGGGAAGCT SSLP R: GCAGAGTCCTTCTGTTTTCTCA 60ºC anneal z13230 F: TATGAAAAGGCACTTGATGTGG SSLP R: TCTGCGCAGTAATAGCAAACA 60ºC anneal z7381 F: TCCAGTCCACAAAAATGCAA SSLP R: TGACCACAGCAACTCCATTC 60ºC anneal z6024 F: CTGCAGGCTTAATGCATTCA SSLP R: GGCTCAGCAGTTACATGCAA 60ºC anneal GOF 15 F: ACATGTTCAGAGACGCCAGA SSLP R: ATTCTGAAACCTGAGGCCCT 60ºC anneal z13310 F: TGGTGATCGTAAATAAAAGAGCTG SSLP R: CACCAGATCCAACCCGTATC 60ºC anneal z3760 F: ACATGGGTGCTCAATCCGCT SSLP R: TCCTTCCCGAACCATCCAGA 60ºC anneal z21165 F: TGTCAGTTCTCTCAGTGGCG SSLP R: AATGGCGCATATAGAAACGC 60ºC anneal z22430 F: AGTAGAGCCAAGCGCTTCTG SSLP R: GGGTTATGTGGACTCGCAGT 60ºC anneal M13 F: GTAAAACGACGGCCAGT Standard R: AACAGCTATGACCATG 60ºC anneal

219 T7 TAATACGACTCACTATAGGG Standard 60ºC anneal T3 ATTAACCCTCACTAAAGGGA Standard 60ºC anneal SP6 ATTTAGGTGACACTATAG Standard 60ºC anneal Pitx1-ISH F: GGACTCACTTCACNAGCCAGCAG Standard R: TAGGCTGGAGTTGCAVGTGTCCCGGTA 60ºC anneal Tbx4mRNASTOP F: CGGGATCCATGCTTCAGGAGAAG High-fidelity R: ATATCTCGAGACTGTCAGTCCAGTGTG 60ºC anneal Tbx4minigene F: GCGTTCTGGAATGATATGGAGG High-fidelity R: CAGTTGTTGCTCTGAGAGAG 60ºC anneal Tbx4 cDNA F: GCGTTCTGGAATGATATGGAGG High-fidelity R: CAGTTGTTGCTCTGAGAGAG 60ºC anneal Tbx4 exon1 seq F: GCTCGTTTGGAGTCGTGTGAGT Standard R: TCTAATCGCATTACCGCCCTCC 60ºC anneal Tbx4 exon2 seq F: TGCACCAAAATACATTGCCTA Standard R: TGAGTCATTGTCGGTGGAATC 60ºC anneal Tbx4 exon3(2) F: GGCTCTTGCCATGTAGGGTA Standard seq R: GAATGGGTGGATGAAAATGG 60ºC anneal Tbx4 exon4(2) F: TGTTTTGACTTTATTTACGCCATT Standard seq R: AAATTTTCCAAGGGCTTTCGA 60ºC anneal Tbx4 exon5 seq F: CGGGATGATGTCAGAGCTG Standard R: 60ºC anneal GAAATGGACACCACTAACAGAGCAATTACC Tbx4 exon6/7 seq F: TCGAAGGCCGATTACAGTTT Standard R: CTAGGAAAGGGGGCACTTGATGAGTA 60ºC anneal Tbx4 exon7/8 seq F: TAAAGCCTCCCCAAAACACA Standard R: TTTGAAGTGTTCGGCAAAAA 60ºC anneal Tbx4 exon9.1 seq F: GCTGACAGTCTGCAGTCATTTT Standard R: TGTTTGGTACTGATAGGCTGGTT 60ºC anneal Tbx4 exon9.2 seq F: AAAGCTGGGTCTCCATTACGTTCA Standard R: TTCCAAACTGCTGTGCCATAC 60ºC anneal RAP1GAP2 F: TCCTTACAGTGTGTGTTTGTTTGCTCT High-fidelity R: CCTGGGAATCGGGAACCCACCT 60ºC anneal Spg20b F: TCCCAACAGACCACCTGCCT High-fidelity R: ATTCATCTCACAACTAGGCCCAACG 60ºC anneal Scarf1 F: CGGCGGACAGGACGCAGATT High-fidelity R: CACAGTGTCAGGCAGAGCCACA 60ºC anneal RNH1 F: GCTCTGAGATCAAACCCTTCACACC High-fidelity R: TGGCTGAGGACAGCACCATCA 60ºC anneal NPAT F: GGCACAGACGGAGTCAGACCCT High-fidelity R: GGCCAGCCTCGTCACTCTCT 60ºC anneal

220 SSLP PCR reagents and cycling conditions

1 µl 10x PCR buffer

1 µl 2.5mM (each) dNTPS (NEB)

4 µl primers (5 µM each)

4 µl gDNA template (~50 ng)

0.1 µl Thermopol Taq (NEB)

10 µl

95ºC for 2 min

(95 ºC for 30 sec; 60 ºC for 30 sec; 72 ºC for 1 min) x 40

72 ºC for 10 min

Standard PCR reagents and cycling conditions

2.5 µl 10x PCR buffer

1 µl 10 mM (each) dNTPS (NEB)

5 µl primers (10 µM each) x µl template (~50 ng gDNA/cDNA template or < 1 ng plasmid DNA)

0.1 µl Thermopol Taq (NEB) x µl milliQ H2O

25 µl

95 ºC for 2 min

(95 ºC for 30 sec; 60 ºC for 30 sec; 72 ºC for 1 min per kb) x 35

72 ºC for 2 min

221 High-fidelity PCR reagents and cycling conditions

5 µl 5x PCR buffer 1 µl 10 mM (each) dNTPS (NEB) 0.6 µl DMSO 4 µl primers (10 µM each) x µl template (~50 ng gDNA/cDNA template or < 1 ng plasmid DNA) 0.25 µl Phusion DNA polymerase (NEB) x µl milliQ H2O 25 µl

98 ºC for 3 min

(98 ºC for 10 sec; 60 ºC for 30 sec; 72 ºC for 30 sec per kb) x 35

72 ºC for 10 min

222 In situ hybridisation probes

Probe Restriction enzyme RNA polymerase Pitx1 Sense: NotI Sense: T3 Antisense: SpeI Antisense: T7 Tbx4 Sense: SalI Sense: SP6 Antisense: NotI Antisense: T7 Tbx5 Sense: SalI Sense: SP6 Antisense: NotI Antisense: T7 Fgf10a Sense: BamHI Sense: T7 Antisense: NotI Antisense: SP6 Sp8 Sense: NotI Sense: SP6 Antisense: KpnI Antisense: T7 Fgf8a Sense: SalI Sense: SP6 Antisense: NotI Antisense: T7 Shh Sense: SspI Sense: T3 Antisense: HindIII Antisense: T7

223 Appendix B

Morphology of Pelvic Fin Loss in a Zebrafish Strain (Danio rerio)

Emily K. Don, Thomas E. Hall, Peter J. Currie, Nicholas J. Cole

Journal of Morphology (2011)

This paper was written and published during the course of this degree. It contains work researched during my honours degree (approximately 60%) and during this degree (approximately 40%). This paper provides a preliminary morphological description of pelvic finless zebrafish. The author contributions were as follows.

Emily Don performed all experiments described, wrote the manuscript and offered intellectual input into the experimental design.

Nicholas Cole developed the original concept, offered intellectual input and guidance into the experimental design, and assisted the production of the manuscript.

Thomas Hall and Peter Currie contributed to the work in the form of development of the original concept, intellectual discussions, guidance and proof reading of the manuscript.

Signatures from co-authors:

Nicholas Cole Thomas Hall Peter Currie

224 JOURNAL OF MORPHOLOGY 272:583–589 (2011)

Morphology of Pelvic Fin Loss in a Zebrafish Strain (Danio rerio)

Emily K. Don,1 Thomas E. Hall,2 Peter D. Currie,2 and Nicholas J. Cole1*

1Department of Anatomy & Histology, School of Medical Sciences and Bosch Institute, University of Sydney, New South Wales 2006, Australia 2Australian Regenerative Medicine Institute, Monash University, Victoria 3800, Australia

ABSTRACT We describe the morphology of a zebrafish been only a few studies on the pelvic fins of tele- strain which lacks pelvic fins but no other abnormalities. osts (Geraudie, 1978; Nelson, 1993; Gibson-Brown This description is the first step of analyzing hindlimb et al. 1996; Tamura et al., 1999; Cole et al., 2003; loss in an established model organism. By combining Shapiro et al., 2004; Tanaka et al., 2005; Chan light microscopy, bone and cartilage staining, scanning et al., 2010; Murata et al., 2010) and the genetic electron microscopy and histological sections we were able to comprehensively describe the morphology of the basis of pelvic fin development remains relatively developing pelvic fins of a pelvic finless zebrafish in con- unexplored. This zebrafish will have a significant trast with the developing pelvic fins of wild-type zebra- impact upon our knowledge of both pelvic fin/hind- fish. We have shown that although adult pelvic finless limb developmental mechanisms and the evolu- zebrafish completely lack pelvic fins, they do develop tionary mechanisms of hindlimb loss. It is there- mesenchymal bulges in the pelvic regions at the pelvic fore of some importance to characterize pelvic fin fin development stage. Understanding the morphology loss in this genetically tractable model species. and the subsequent genetic analysis of this fish will lead to important insights into both pelvic fin/hindlimb devel- opmental mechanisms and the evolution of hindlimb MATERIALS AND METHODS loss. It is for this reason that we present a morphologi- Fish Stocks cal analysis of pelvic fin development and loss in this The use and treatment of animals in the project were in genetically tractable model species. J. Morphol. 272:583– accordance with the National Health and Medical Research 589, 2011. Ó 2011 Wiley-Liss, Inc. Council guidelines. All protocols were approved by the Animal Ethics Review Committee, University of Sydney (NSW, KEY WORDS: zebrafish; limb; fin; development; Australia). This research was conducted at the University of evolution Sydney between March 2009 and September 2010. Fish maintenance and crossing was performed as described in The Zebrafish Book (Westerfield, 2000). WIK and Tubingen were used as wild-type strains. A small number of pelvic finless zebrafish were found in a school of zebrafish obtained from a INTRODUCTION local store in Sydney, Australia. These fish were incrossed and the offspring were used for this study. We describe the morphology of a zebrafish strain which lacks pelvic fins but no other abnormalities. Describing the morphology of this pelvic finless Histology zebrafish strain is a first step in analyzing hind- Wild-type (n 5 10) and pelvic finless zebrafish larvae (n 5 limb loss in an established model organism. 10) were fixed in 4% paraformaldehyde in phosphate buffered Normally, adult zebrafish have two sets of paired saline overnight and dehydrated via an ethanol series. The l fins, the pectoral and the pelvic fins (Grandel and larvae were then embedded in paraffin and 12 m sections were cut on a microtome. Haematoxylin and eosin staining was per- Schulte-Merker, 1998). The pectoral fins of zebra- formed on the sections which were then mounted and cover fish are homologous to the forelimbs of tetrapods slipped. Sections were then imaged on a Leica DM1 300 D and the pelvic fins of zebrafish are homologous to inverted microscope. the hindlimbs (review: Carroll, 1988; Coates, 1994). This evolutionary origin is indicated by the *Correspondence to: Nicholas J. Cole, Department of Anatomy pattern of gene and protein expression during the and Histology, Anderson Stuart Building (F13), Eastern Ave, Uni- development of paired fins (review: Mercader, versity of Sydney, NSW 2006, Australia. 2007). This makes the paired fins of fish a directly E-mail: [email protected] relevant model to investigate vertebrate limb for- mation. In the particular zebrafish strain Received 23 September 2010; Revised 11 November 2010; Accepted 13 November 2010 described here, there are no pelvic fins, which is equivalent to an animal with no hindlimbs. Published online 22 February 2011 in Despite the important conservation between Wiley Online Library (wileyonlinelibrary.com) pelvic fin and hindlimb development, there have DOI: 10.1002/jmor.10938

Ó 2011 WILEY-LISS, INC. 225 584 E.K. DON ET AL.

Fig. 1. Pelvic finless zebrafish (A–C) and wild-type zebrafish (D–F). a, anal fin; c, caudal fin; d, dorsal fin; pec, pectoral fin; pel, pelvic fin; scale bars 5 1 mm. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

100%) for 10 minutes each step. Larvae were critical point dried Skeletal Staining to remove ethanol. The larvae were sputter coated and imaged A two-color acid free cartilage and bone stain for adult zebra- in high vacuum mode on a XL30 scanning electron microscope at 5–15 kV. fish was adapted from Walker and Kimmel (2007), but CaCl2 was used in place of MgCl2. Whole mount skeletal staining (Wild-type adults n 5 10 and pelvic finless adults n 5 10) were imaged in 3% methyl cellulose using a Leica M165FC stereo RESULTS dissection microscope. Pelvic finless zebrafish lack any gross external Scanning Electron Microscopy pelvic fin structures (Fig. 1A–C). In contrast, the paired pelvic fins of wild-type adult zebrafish are Larvae (Wild-type n 5 10 and pelvic finless n 5 10) were positioned on the ventrolateral body wall just ante- fixed in 1% gluteraldehyde and 1% paraformaldehyde in 0.1 M PBS (pH 7.2) for 2 hours at room temperature and postfixed in rior to the cloacae at the posterior end of the 1% OsO4 in 0.1 M PBS for 1 hour. Larvae were dehydrated abdominal cavity (Fig. 1D–F). Bone and cartilage through an ethanol series (2 3 50%; 70%; 2 3 90%; 95%; 3 3 staining was performed on pelvic finless zebrafish

Journal of Morphology 226 MORPHOLOGY OF PELVIC FIN LOSS 585

Fig. 2. Bone and Cartilage staining showing a comparison of the skeletal elements of pelvic fins in pelvic finless (A–C) and wild-type zebrafish (D–F). lep, lepidotrichs; r, radials; b, basip- terygium; scale bars 5 1 mm. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.] to determine if the strain developed any pelvic these developmental stages pelvic finless zebrafish skeletal structures. This staining revealed that the did in fact have tiny bulges in the presumptive adults of the pelvic finless strain of zebrafish lack pelvic fin regions of equal size on both sides in ev- all pelvic skeletal elements (Fig. 2A–C). Usually, ery case examined (Fig. 3A,B). In contrast, wild- wild-type adult zebrafish pelvic fins consist of type zebrafish at the same total length had fully three main skeletal elements; the basipterygium developed adult pelvic fins (Fig. 3C,D). (pelvic girdle), the three radials (fin base) and the In pelvic finless zebrafish, each mesenchymal lepidotrichs (fin rays; Fig. 2D–F; Cubbage and bulge develops ventral to the ninth and tenth myo- Mabee, 1996). tome; however the ectodermal cells of the bulges To see the fine detail of the presumptive pelvic do not seem to be any different from the surround- fin regions, we performed scanning electron ing ectoderm. In contrast, a narrow, clearly organ- microscopy of 8–14 mm larvae to determine if ized band of cells which are morphologically differ- there were any early morphological signs of pelvic ent from their surroundings is seen protruding fins which are subsequently absent in the adult. from the flank in this position in wild-type zebra- Scanning electron microscopy revealed that at fish (Fig. 4A,D).

Journal of Morphology 227 586 E.K. DON ET AL.

Fig. 3. Scanning electron micrographs showing the position and morphology of developing pelvic fins in both pelvic finless (A, B) and wild-type zebrafish (C, D). pec, pectoral fin; pel, pelvic fin.

In wild-type zebrafish, the narrow band of cells pelvic bulges of pelvic finless zebrafish revealed continues to develop as a pelvic fin bud. The pelvic that the bulges in the pelvic region of pelvic finless fin bud of wild-type fish forms an apical ectoder- zebrafish do not have the same developmental mal thickening (Fig. 4E). However, in pelvic finless morphology as wild-type pelvic fin buds. There is zebrafish, the mesenchymal bulge grows but never no morphological evidence of an apical ectodermal forms a narrow band of cells and does not form an thickening or apical fold in the pelvic bulges of apical ectodermal thickening (Fig. 4B). As develop- pelvic finless zebrafish. ment continues the wild-type pelvic fins take on a Small mesenchymal bulges form in the pelvic paddle like shape and as the wild-type zebrafish region of pelvic finless zebrafish but these bulges reach 14 mm standard length the pelvic fins are lack the features of wild-type pelvic fin buds. The fully formed (Fig. 4F). When the pelvic finless pelvic bulges of the pelvic finless zebrafish do not zebrafish develop further, the bulges in the pelvic form into a narrow band of cells that protrude region have arrested development and true pelvic from the flank of the larvae, and seem to lack any fins never form. morphologically distinct apical ectodermal thicken- Histological sections stained with haematoxylin ing. In fact, the cells of the overlying ectoderm of and eosin also reveal the pelvic bulges of pelvic the bulge show no morphological difference in finless zebrafish in comparison to wild-type zebra- appearance compared to the surrounding nonbulge fish (Fig. 5A,C). The overlying ectodermal cells of ectoderm. Grandel and Schulte-Merker (1998) the pelvic bulges show no morphological difference described the formation of this narrow band of compared to the surrounding ectoderm (Fig. 5B) cells protruding from a mesenchymal bulge on the while the ectodermal cells in the wild-type pelvic flank of the zebrafish and its subsequent de- fins have formed an apical fold (Fig. 5D). The pel- velopment into the apical fold as one of the key vic bulges of the pelvic finless zebrafish do not steps in zebrafish pelvic fin development. This nar- form the apical ectodermal thickening or the apical row band of cells has been previously observed in fold seen in wild-type pelvic fins (Fig. 5B,D). both trout and killifish pelvic fins when it was termed the apical ectodermal pseudo-ridge and it has been presumed to play the same role as DISCUSSION the apical ectodermal ridge does in tetrapods We have discovered a novel pelvic finless zebra- (Geraudie, 1978; Wood, 1982; Grandel and fish strain, a well established, genetically tractable Schulte-Merker, 1998). It is feasible that the lack model vertebrate, and performed a detailed mor- of a ridge-like structure and subsequent apical fold phological examination of the pelvic region of this may be the cause of the failure of pelvic fin out- zebrafish in comparison to wild-type zebrafish with growth, as the apical ectodermal ridge is essential normal pelvic fin formation. Comparison between in pectoral fin and tetrapod limb outgrowth the pelvic fin buds of wild-type zebrafish and the (review: Robert and Lallemand, 2006). We are

Journal of Morphology 228 MORPHOLOGY OF PELVIC FIN LOSS 587

Fig. 4. Scanning electron micrographs showing pelvic fin development in pelvic finless (A–C) and wild-type zebrafish (D–F). (A, D) white arrows indicate first visible evidence of devel- oping pelvic fin buds. (B, E) Black arrows indicate the presence (E) and absence (B) of an apical ectodermal thickening.

currently performing a comprehensive analysis of This secondary loss of pelvic structures has the signaling mechanisms at these developmental occurred independently in several lineages of fish, stages in both wild-type and pelvic finless zebra- including pufferfish, cowfish and some three-spine fish in an effort to determine what is responsible stickleback fish populations (Bell et al., 1993; for this loss of pelvic fins. This knowledge may Santini and Tyler, 2003; Tanaka et al., 2005). In lead to insights into genes and regulators responsi- fact, pelvic fin loss is documented in one or more ble for vertebrate hindlimb development and their species of 92 teleostean families; the pelvic fin has secondary loss. been lost about 50 independent times, excluding

Journal of Morphology 229 588 E.K. DON ET AL.

Fig. 5. Histological sections showing the developing pelvic fins of both pelvic finless (A, B) and wild-type zebrafish (C, D). (B, D) Note that the pelvic finless bulges do not form an apical fold that is seen in the wild-type. pb, pelvic bulge; af, apical fold; scale bars 5 500 lm (A, C); 50 lm (B, D). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

multiple losses within families (Nelson, 1993). Many species that have undergone secondary However, these examples of hindlimb loss have hindlimb loss show different developmental mor- occurred in nonmodel organism species and this phologies of the hindlimb/pelvic fin buds. Hindlimb has lead to difficulties in establishing the exact loss has also occurred repeatedly over evolutionary mechanisms responsible for the loss of hindlimbs. time in diverse lineages such as mammals (e.g., In stickleback absence of Pitx1 activity has been whales and dolphins), snakes and teleost fish. Pel- reported to be responsible for pelvic spine loss in vic spine deficient stickleback fish and pufferfish some three spine stickleback populations (Cole (which lack pelvic fins) show no morphological de- et al., 2003; Shapiro et al., 2004; Chan et al., velopment of pelvic fins at any stage (Cole et al., 2010). However, the pelvic finless zebrafish is dif- 2003; Shaprio et al., 2004; Tanaka et al., 2005). ferent from the spine deficient sticklebacks as The dolphin (Stenella attenuate) develops hindlimb scanning electron microscopy performed in Cole buds that do form an apical ectodermal ridge et al., 2003 clearly showed that there is no bulge which regresses and pythons do not develop an ap- present in the pelvic regions of spine deficient ical ectodermal ridge but do develop the begin- sticklebacks. nings of hindlimb buds (Cohn and Tickle, 1999;

Journal of Morphology 230 MORPHOLOGY OF PELVIC FIN LOSS 589 Thewissen et al., 2006). The pelvic finless zebrafish cation of threespine stickleback. Current Biology 13:R951– strain described here appears to have hindlimb R952. Cubbage CC, Mabee PM. 1996. Development of the cranium loss that is more similar to dolphins and snakes, and paired fins in the zebrafish Danio rerio (Ostariophysi, where it is the outgrowth phase of hindlimb devel- Cyprinidae). J Morphol 229:121–160. opment that is affected. It will be interesting to DeLaurier A, Schweitzer R, Logan M. 2006. Pitx1 determines discover the different mechanisms responsible for the morphology of muscle, tendon, and bones of the hindlimb. secondary loss of hindlimbs and these mechanisms Dev Biol 299:22–34. Geraudie J. 1978. Scanning electron microscope study of the may shed light on the differences between fore and developing trout pelvic fin bud. Anat Rec 191:391–396. hindlimb development in vertebrate species. Gibson-Brown J, Agulnuk S, Chapman D, Alexiou M, Garvey As this pelvic loss has occurred in a model orga- N, Silver L, Papaioannou V. 1996. Evidence of a role for T-box nism there are many options available for further genes in the evolution of limb morphogenesis and the specifi- cation of forelimb/hindlimb identity. Mech Dev 56:93–101. investigation. It is hoped that the genetic mapping Grandel H, Schulte-Merker S. 1998. The development of the of this pelvic finless strain of zebrafish will shortly paired fins in the Zebrafish (Danio rerio). Mech Dev 79:99– reveal potential candidates responsible for pelvic 120. fin loss and may identify a key component in the Mercader N. 2007. Early steps of paired fin development in specification and/or development of vertebrate hin- zebrafish compared with tetrapod limb development. Dev Growth Differ 49:421–437. dlimbs. It is exciting to think that through this Murata Y, Tamura M, Aita Y, Fujimura K, Murakami Y, Okabe previously undescribed, model zebrafish, we may M, Okada N, Tanaka M. 2010. Allometric growth of the trunk be able to obtain novel insights into the mecha- leads to the rostral shift of the pelvic fin in teleost fishes. Dev nisms and signaling centers involved in vertebrate Biol 347:236–245. Nelson JS. 1993. Analysis of the multiple occurrence of pelvic limb development. fin absence in extant fishes. Matsya 1989/90; 15/16:21–38. Robert B, Lallemand Y. 2006. Anteroposterior patterning in the limb and digit specification: Contribution of mouse genetics. ACKNOWLEDGMENTS Dev Dyn 235:2337–2352. The authors thank the contributions of the Santini F, Tyler JC. 2003. A phylogeny of the families of fossil and extant tetraodontiform fishes (Acanthomorpha, Tetrao- Australian Centre for Microscopy and Micro- dontiformes), upper cretaceous to recent. Zoo J Linn Soc analysis and in particular the expertise of Dr. 139:565–617. Errin Johnson and Dr. Ian Kaplin for assistance Shapiro MD, Marks ME, Peichel CL, Blackman BK, Nereng with the scanning electron microscopy. KS, Jonsson B, Schluter D, Kingsley DM. 2004. Genetic and developmental basis of evolutionary pelvic reduction in threespine sticklebacks. Nature 428:717–723. LITERATURE CITED Tamura K, Yonei-Tamura S, Belmonte JCI. 1999. Differential expression of Tbx4 and Tbx5 in Zebrafish Fin buds. Mech Bell MA, Orti G, Walker JA, Koenings JP. 1993. Evolution of Dev 87:181–184. pelvic reduction in threespine stickleback fish: A test of com- Tanaka M, Hale LA, Amores A, Yan Y-L, Cresko WA, Suzuki T, peting hypotheses. Evolution 47:906–914. Postlethwait JH. 2005. Developmental genetic basis for the Carroll R, editor. 1988. Vertebrate Paleontology. San Francisco: evolution of pelvic fin loss in the pufferfish Takifugu rubripes. Freeman. Dev Biol 281:227–239. Chan YF, Marks ME, Jones FC, Villarreal G Jr, Shapiro MD, Thewissen JGM, Cohn MJ, Stevens LS, Bajpai S, Heyning J, Brady SD, Southwick AM, Absher DM, Grimwood J, Schmutz Horton WE. 2006. Developmental basis for hind-limb loss in J, Myers RM, Petrov D, Jonsson B, Schluter D, Bell MA, dolphins and origin of the cetacean bodyplan. Proc Natl Acad Kingsley DM. 2010. Adaptive evolution of pelvic reduction in Sci USA 103:8414–8418. sticklebacks by recurrent deletion of a pitx1 enhancer. Sci- Walker M, Kimmel C. 2007. A two-color acid-free cartilage and ence 327:302–305. bone stain for zebrafish larvae. Biotech Histochem 82:23–28. Coates MI. 1994. The origin of vertebrate limbs. Dev Westerfield M,editor. 2000. The Zebrafish Book. A Guide for the Suppl:169–180. Laboratory Use of Zebrafish (Danio rerio), 4 ed. Eugene: Uni- Cohn MJ, Tickle C. 1999. Developmental basis of limblessness versity of Oregon Press. and axial patterning in snakes. Nature 399:474–479. Wood A. 1982. Early pectoral fin development and morphogene- Cole NJ, Tanaka M, Prescott A, Tickle C. 2003. Expression of sis of the apical ectodermal ridge in the Killifish. Anat Rec limb initiation genes and clues to the morphological diversifi- 204:349–356.

231 Journal of Morphology Appendix C

Tables from Morphology Chapter

Table C.1. Table of length measurements of pelvic finless zebrafish and wild-type siblings. Figure 1.3. The data from length measures of pelvic finless zebrafish and wild-type siblings. A two-way ANOVA with the null hypothesis of µ1≠µ2 was performed on length measurements to determine if a difference in length was present between pelvic finless embryos and wild-type siblings. Overall, there was no significant difference in length (p=0.8).

Stage Genotype Number Average length (mm) SE of difference

24hpf WT 4 1.87 ±0.1

pFL 4 1.80

48hpf WT 5 2.3 ±0.2

pFL 5 2.3

72hpf WT 3 2.9 ±0.0

pFL 3 2.9

7dpf WT 4 3.3 ±0.1

pFL 4 3.0

14dpf WT 4 4.8 ±0.3

pFL 4 4.9

21dpf WT 4 5.1 ±0.3

pFL 4 5.3

28dpf WT 4 7.7 ±0.7

pFL 4 7.9

232 Table C.2. Fin ray number A table describing the number of fin rays present in all fins between pelvic finless zebrafish and wild-type siblings. No difference were significant, except of the pelvic fins.

Pelvic Pectoral Ventral Dorsal Caudal

WT (n=4) 8 11 15 8 28

8 11 15 10 29

9 10 17 9 30

7 9 16 10 28

pFL (n=4) 0 10 14 9 30

0 11 15 10 30

0 10 15 9 28

0 10 15 10 30

Mean WT 8 10.25 15.75 9.25 28.75

Mean pFL 0 10.25 14.75 9.5 29.5

SE of difference ±0.41 ±0.54 ±0.54 ±0.56 ±0.69

p-value <0.0001*** >0.9999 0.11 0.67 0.32

233 Appendix D

Table D.1. Recombinants at chromosome 15 markers A table of the number of recombinant individuals at tested chromosome 15 markers is presented on the following pages.

234 z10289 (7.3cM)PNI z21982 (34.4cM)PI z4396 (49cM)PNI z11320 (54cM)PI mutation z13310 (63.7cM) PNI z13230 (67.4cM)PNI z7381 (79cM) PI M2 R M3 R M4 R R R R missort? M5 [R] R M6 R R R R missort? M7 M8 R M9 R R R R M10 M11 M12 M13 R M14 R R R R M15 [R] R R M16 [R] R R M17 R M18 M19 R M20 R R R [R] R missort? M21 [R] R M22 M23 [R] R R M24 R M25 R R M26 [R] R M27 R M28 M29 M30 [R] R M31 [R] R R M32 R R [R] R R [R] R missort? M33 M34 R M35 M36 M37 M38 R M39 R z10289 (7.3cM)PNI z21982 (34.4cM)PI z4396 (49cM)PNI z11320 (54cM)PI mutation z13310 (63.7cM) PNI z13230 (67.4cM)PNI z7381 (79cM) PI M40 [R] R M41 [R] R M42 R M43 M44 R M45 RR R M46 RR M47 M48 M49 R M50 [R] R missort? M51 R R [R] R R M52 R R R M53 M54 M55 M56 [R] R M57 RR R M58 M59 R M60 M61 [R] R M62 [R] R [R] R Total MF1 13+ [13] (PNI) 22 (PI) 12 + [1](PNI) 3 (PI) 0 4+ (PNI) 4+ [2] (PNI) 21 (PI) minus missorts 10 + [13] (PNI) 19 (PI) 9 (PNI) 2 (PI) 0 2 (PNI 4+ (PNI) 18 (PI) z10289 (7.3cM)PNI z21982 (34.4cM)NP z4396 (49cM)NP z11320 (54cM)PNI mutation z3760 (63.7cM) PI z13230 (67.4cM) PNI z7381 (79cM) PNI M63 R R missort? M64 1 fin M65 [R] M66 M67 M68 R M69 M70 M71 R [R] [R] 1 fin M72 M73 M74 M75 R M76 R R missort? M77 [R] M78 M79 R [R] [R] M80 R [R] M81 M82 R [R] [R] M83 M84 R [R] [R] 1 fin M85 R M86 M87 M88 R M89 [R] M90 R [R] 1 fin M91 M92 R 1 fin M93 R [R] [R] M94 R [R] [R] M95 M96 M97 M98 M99 [R] M100 [R] z10289 (7.3cM)PNI z21982 (34.4cM)NP z4396 (49cM)NP z11320 (54cM)PNI mutation z3760 (63.7cM) PI z13230 (67.4cM) PNI z7381 (79cM) PNI M101 [R] M102 M103 M104 M105 R [R] [R] 1 fin M106 M107 M108 R R [R] M109 M110 R [R] R M111 R M112 RR M113 M114 R [R] [R] M115 R R [R] M116 M117 R [R] [R] M118 M119 R [R] [R] M120 M121 M122 M123 R [R] M124 R R [R] M125 Total MF1 26 (PNI) 22 (PI) 12 (PNI) 2 (PI) 2+ (PNI) 4+ (PNI) 18 (PI) Total MF2 6+ (PNI) 13 (PI) 5+ (PNI) 6 + [16] minus missorts 23 19 9 8 12 (PI) Total 23/59 19/59 9/59 8/122 12/122 9+/122 Total (PI) 19/59 8/122 12/63 40/122 Appendix E tbx4 cDNA sequence

Complete alignment of Tbx4 cDNA transcript from mRNA extracted from Wild-type embryos (We), Wild-type developing pelvic fins (Wp), pelvic finless embryos (Fe) and pelvic finless developing pelvic regions (Fp). The coding region of Tbx4 is represented in uppercase. Discrepancies in the sequence are highlighted.

1 10 20 30 40 50 60 | | | | | | |

WeF_D12.ab1 ------ccgagctacaatatttatgagaATGCTTCAGGAG WpF_E04.ab1 ------gagggccactcactgtatgagaATGCTTCAGGAG FeF_E02.ab1 ------cgcatcgcatattttatgagaATGCTTCAGGAG FpF_E06.ab1 ------cggcccgactatattttatgagaa-TGCTTCAGGAG FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 AAGGCCTCAGTTGTGGCTGATGAAGGAATGACCGTTGCCCAGTCGGGTGGGCGTCCTGAG WpF_E04.ab1 AAGGCCTCAGTTGTGGCTGATGAAGGAATGACCGTTGCCCAGTCGGGTGGGCGTCCTGAG FeF_E02.ab1 AAGGCCTCAGTTGTGGCTGATGAAGGAATGACCGTTGCCCAGTCGGGTGGGCGTCCTGAG FpF_E06.ab1 AAGGCCTCAGTTGTGGCTGATGAAGGAATGACCGTTGCCCAGTCGGGTGGGCGTCCTGAG FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 CTGGCCAGCGATTCCTCACATCTGGGCCTGCCTACAACTCCCAGCAATCCCCAGAACAAT WpF_E04.ab1 CTGGCCAGCGATTCCTCACATCTGGGCCTGCCTACAACTCCCAGCAATCCCCAGAACAAT FeF_E02.ab1 CTGGCCAGCGATTCCTCACATCTGGGCCTGCCTACAACTCCCAGCAATCCCCAGAACAAT FpF_E06.ab1 CTGGCCAGCGATTCCTCACATCTGGGCCTGCCTACAACTCCCAGCAATCCCCAGAACAAT FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 GAACCTGACCAGAGCATTGAAAACATCAAAGTGGTTCTTCATGACAGGGAATTATGGAAG WpF_E04.ab1 GAACCTGACCAGAGCATTGAAAACATCAAAGTGGTTCTTCATGACAGGGAATTATGGAAG FeF_E02.ab1 GAACCTGACCAGAGCATTGAAAACATCAAAGTGGTTCTTCATGACAGGGAATTATGGAAG FpF_E06.ab1 GAACCTGACCAGAGCATTGAAAACATCAAAGTGGTTCTTCATGACAGGGAATTATGGAAG FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 AAGTTTCACGAGGCTGGAACCGAGATGATCATCACCAAAGCAGGCAGGCGGATGTTTCCT WpF_E04.ab1 AAGTTTCACGAGGCTGGAACCGAGATGATCATCACCAAAGCAGGCAGGCGGATGTTTCCT FeF_E02.ab1 AAGTTTCACGAGGCTGGAACCGAGATGATCATCACCAAAGTAGCAAGGCGGATGTTTCCT FpF_E06.ab1 AAGTTTCACGAGGCTGGAACCGAGATGATCATCACCAAAGTAGCAAGGCGGATGTTTCCT FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 AGCTACAAAGTTAAAGTCACAGGAATGAACCCCAAGACCAAATACATTCTATTAACTGAC WpF_E04.ab1 AGCTACAAAGTTAAAGTCACAGGAATGAACCCCAAGACCAAATACATTCTATTAACTGAC FeF_E02.ab1 AGCTACAAAGTTAAAGTCACAGGAATGAACCCCAAGACCAAATACATTCTATTAACTGAC FpF_E06.ab1 AGCTACAAAGTTAAAGTCACAGGAATGAACCCCAAGACCAAATACATTCTATTAACTGAC FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

239 WeF_D12.ab1 ATTGTTCCAGCTGATGATCATCGGTACAAGTTCTGTGACAACAAGTGGATGGTGGCCGGA WpF_E04.ab1 ATTGTTCCAGCTGATGATCATCGGTACAAGTTCTGTGACAACAAGTGGATGGTGGCCGGA FeF_E02.ab1 ATTGTTCCAGCTGATGATCATCGGTACAAGTTCTGTGACAACAAGTGGATGGTGGCCGGA FpF_E06.ab1 ATTGTTCCAGCTGATGATCATCGGTACAAGTTCTGTGACAACAAGTGGATGGTGGCCGGA FpR_E07.ab1 ------FeR_E03.ab1 ------WpR_E05.ab1 ------WeR_E01.ab1 ------

WeF_D12.ab1 AAAGCAGAACCTGCCATGCCTGGAAGACTCTATGTCCACCCAGACTCTCCTGCAACAGGC WpF_E04.ab1 AAAGCAGAACCTGCCATGCCTGGAAGACTCTATGTCCACCCAGACTCTCCTGCAACAGGC FeF_E02.ab1 AAAGCAGAACCTGCCATGCCTGGAAGACTCTATGTCCACCCAGACTCTCCTGCAACAGGC FpF_E06.ab1 AAAGCAGAACCTGCCATGCCTGGAAGACTCTATGTCCACCCAGACTCTCCTGCAACAGGC FpR_E07.ab1 ------TTTAAATCTGCCGGC FeR_E03.ab1 ------GACACCCCCTGCCTGC WpR_E05.ab1 ------GGGACATTCTGCCTCGC WeR_E01.ab1 ------CAAGATCTGCATG

WeF_D12.ab1 GCACATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT WpF_E04.ab1 GCGCATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT FeF_E02.ab1 GCGCATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT FpF_E06.ab1 GCGCATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT FpR_E07.ab1 GCGCATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT FeR_E03.ab1 GCGCATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT WpR_E05.ab1 GCGCATTGGATGAGACA-TTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT WeR_E01.ab1 CGCAATTGGATGAGACAATTGGTTTCTTTTCAGAAGCTCAAGCTCACAAACAACCACCTT

WeF_D12.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT WpF_E04.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT FeF_E02.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT FpF_E06.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT FpR_E07.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT FeR_E03.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT WpR_E05.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT WeR_E01.ab1 GATCCTTTTGGACATATTATCTTGAACTCGATGCACAAATATCAGCCCAGGTTACACATT

WeF_D12.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT WpF_E04.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT FeF_E02.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT FpF_E06.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT FpR_E07.ab1 GTGAAAGCTGATGATAACAATGCCTTTGGTTCCAAGAACACAGCCTACAGCACTCATGTT FeR_E03.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT WpR_E05.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT WeR_E01.ab1 GTGAAAGCTGATGAGAACAATGCCTTTGGTTCCAAGAACACGGCCTACTGCACTCATGTT

WeF_D12.ab1 TTCCACGAGACGGCTTTCATCTCTGTTACATCTTATCAGAACCATAAGATCACGCAATTA WpF_E04.ab1 TTCCACGAGACGGCTTTCATCTCCGTTACATCTTATCAGAACCATAAGATCACGCAATTA FeF_E02.ab1 TTCCACGAGACGGCTTTCATCTCCGTTACATCTTATCAGAACCATAAGATCACGCAATTA FpF_E06.ab1 TTCCACGAGACGGCTTTCATCTCCGTTACATCTTATCAGAACCATAAGATCACGCAATTA FpR_E07.ab1 TTCCACGAGACGGCTCTATCACCCTTGTGATTGTTAAAAACCTTAGAAAGAACCCAATAG FeR_E03.ab1 TTCCACGAGACGGCTTTCATCTCCGTTACATCTTATCAGAACCATAAGATCACGCAATTA WpR_E05.ab1 TTCCACGAGACGGCTTTCATCTCCGTTACATCTTATCAGAACCATAAGATCACGCAATTA WeR_E01.ab1 TTCCACGAGACGGCTTTCATCTCTGTTACATCTTATCAGAACCATAAGATCACGCAATTA

WeF_D12.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT WpF_E04.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT FeF_E02.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT FpF_E06.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT FpR_E07.ab1 ACATTACCAGCGCGCGTTTGGGCACATCATCTGTAGGCAGTTTTACTTTCCCCCGACGAC FeR_E03.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT WpR_E05.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT WeR_E01.ab1 AAAATTGAGAACAATCCTTTTGCCAAAGGTTTTCGCGGCAGTGATGAGGGTGATTTGCGT

WeF_D12.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG WpF_E04.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG FeF_E02.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG FpF_E06.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG FpR_E07.ab1 AACCAACACCTCCTTTATGTAAGCCCTTCGGGGGGCTGCAACTTAGATGGTGAATGAAAG FeR_E03.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG WpR_E05.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG WeR_E01.ab1 GTGTCTAGACTTCAAGGGAAAGATTACCCAGTGATTTCTAAGAACATGGTCCGGCAGCGG

240 WeF_D12.ab1 CTTATCTCCTCACATGGCCATCTGTCAAGAAAACTGATGCAGGTTGTGCTGAGTACCACC WpF_E04.ab1 CTTATCTCCTCACATGGCCATCTGTCAAGAAAACTGAGTGC-GGTGTGCTGAGTAGCCAC FeF_E02.ab1 CTTATCTCCTCACATGGCCATCTGTCAG-AAAACTGAGTGCAGGTGTGCTGAGTAGCCAC FpF_E06.ab1 CTTATCTCCTCACATGGCCATCTGTCAG-AAAACTGAGTGCAGGTGTGCTGAGTAGCCAC FpR_E07.ab1 ATCATAACCGAAATCCGCCCCTAGATTTTGTACCAAATAGATTGAGCGAACACGATAATG FeR_E03.ab1 CTTATCTCCTCACATGGCCATCTGTCAGGAAAACTGAGTGCAGGTGTGCTGAGTAGCCAC WpR_E05.ab1 CTTATCTCCTCACATGGCCATCTGTCAGGAAAACTGAGTGCAGGTGTGCTGAGTAGCCAC WeR_E01.ab1 CTTATCTCCTCACATGGCCATCTGTCAGGAAAACTGAGTGCAGGTGTGCTGAGTAGCCAC

WeF_D12.ab1 CCGCAGTGCTTTCTCATTATCATATGACGTGGTGTCCCCTTGCCAAACTCAGACTCCAGG WpF_E04.ab1 CCGCAG-TGCTTTCTCATTATCAATATGACAGTG-TGTCCCCTTGCCAAACTCAGACTCC FeF_E02.ab1 CCGCAG-TGCTTTCTCATTATCAATATGACAGTGGTGTCCCCTTGCCAAACTCAGACTCC FpF_E06.ab1 CCGCAGGTGCTTTCTCATTATCAATATGACAGTGGTGTCCCCTTGCCAAACTCAGACTCC FpR_E07.ab1 GGCTTTTGTTGGGAACACCAGAAAATGCAAATATGAAAGAAATTCTTCAAATCGACTCCT FeR_E03.ab1 CCGCAGGTGCTTTCTCATTATCAATATGACAGTGGTGTCCCCTTGCCAAACTCAGACTCC WpR_E05.ab1 CCGCAGGTGCTTTCTCATTATCAATATGACAGTGGTGTCCCCTTGCCAAACTCAGACTCC WeR_E01.ab1 CCGCAGGTGCTTTCTCATTATCAATATGACAGTGGTGTCCCCTTGCCAAACTCAGACTCC

WeF_D12.ab1 AAGCCCTTTCCACTCCTTACATCTAGCAGGAACCAGCTTCTTTACCACTGCTCAGCAAAG WpF_E04.ab1 CAGGAAGCCCTTTCCACTCCTTTACATCTAGCCAGGCAACCCAGCTTCTTTACCACTGCT FeF_E02.ab1 CAGGAAGCCCTTTCCA-CTCCTTTACATCTAGCCGGGAACCCAGCCTTCTTTACCACTGC FpF_E06.ab1 CAGGAAGCCCTTTCCAACTCCTTTACATCTAGCCGGGA-CCCAGCCTTCTTTACCACTGC FpR_E07.ab1 AATGCCTTTTTCCGAAGCTAACACAACAGCTATACAACAAAATCA------FeR_E03.ab1 CAGGAAGCCCTTTCCAACTCCTTTACATCTAGCCGGGAACCCAGCCTTCTTTACCACTGC WpR_E05.ab1 CAGGAAGCCCTTTCCAACTCCTTTACATCTAGCCGGGAACCCAGCCTTCTTTACCACTGC WeR_E01.ab1 CAGGAAGCCCTTTCCAACTCCTTTACATCTAGCCGGGAACCCAGCCTTCTTTACCACTGC

WeF_D12.ab1 AGATACCCAAGTAGTGGAGCTGGATGCAACGACTTATCTTGACCGACTCATCTGCAGTTC WpF_E04.ab1 TCAAGCACAGAAGATACCCAAGTACAGTTGTAGCTGGATGCAACGATCCCTATCTGACAC FeF_E02.ab1 TTCAGCACAGAGATACCCAGACATTTGAGCTGGGATGCAACGACCTATTCTGACACGACA FpF_E06.ab1 TTCAAGCACAGAGATA-CCCAAGACATTTGGAGCTGGGATGCAA-CGACCCTATTCTGAC FpR_E07.ab1 ------FeR_E03.ab1 TTCAAGCACAGAGATAACCCAAGACATTTGGAGCTGGGATGCAAACGACCCTATCTTGAC WpR_E05.ab1 TTCAAGCACAGAGATAACCCAAGACATTTGGAGCTGGGATGCAAACGACCCTATCTTGAC WeR_E01.ab1 TTCAAGCACAGAGATAACCCAAGACATTTGGAGCTGGGATGCAAACGACCCTATCTTGAC

WeF_D12.ab1 CGAAGAAATACTCGCTCCCTCTTCATGATCCCCTGCTGTCCACCAACTGATGAGCTGAGC WpF_E04.ab1 GACATCATCTGCAGTTCGAGAACCCTACTTCGCTTCCTCTCATGATCCGCTGCGTCACTA FeF_E02.ab1 TCATCTGCAGTTCGGAGGAGCATACTTTCGCCTCTCCCTCTCTATGATCCCCCTGCTGTC FpF_E06.ab1 ACGACATCATCTGCAGTTTCGGAGGAGCACTACTTTCGCTCTCCTCCTCTATGATCCCCC FpR_E07.ab1 ------FeR_E03.ab1 ACGACATCATCTGCAGTTTCGGAGGAGCACTACTTTCGCTCTCCTCCTTCCTATGATTCC WpR_E05.ab1 ACGACATCATCTGCAGTTTCGGAGGAGCACTACTTTCGCTCTCCTCCTTCCTATGATTCC WeR_E01.ab1 ACGACATCATCTGCAGTTTCGGAGGAGCACTACTTTCGCTCTCCTCCTTCCTATGATTCC

WeF_D12.ab1 TAAAAGCATGCAGTACCAGCTGGAAGAAAAGAGGGTGCATGTACAGAATAACTCCGGAT- WpF_E04.ab1 CTGCATGAGCTTAGCCTAGAGGCATGCATGTACGAGCTGGAAGGAAAGAGAGCGTGCATT FeF_E02.ab1 CACCCATACTGCAATGAGCTTAGCCTCTAAGAGCCATGCATGTACGAAGCTGAGAAGAGG FpF_E06.ab1 TGCTGTCCCACCATACTGCATGAGCTTTAGCTCTAGAAGCATGCATGTACGAGCTGAGCA FpR_E07.ab1 ------FeR_E03.ab1 CCCCTGCTGTCCCACCCATACTGCAATGAGGCTTTAGGCTCTAGAGAGGCATGCATGTAC WpR_E05.ab1 CCCCTGCTGTCCCACCCATACTGCAATGAGGCTTTAGGCTCTAGAGAGGCATGCATGTAC WeR_E01.ab1 CCCCTGCTGTCACACCCATACTGCAATGAGGCTTTAGGCTCTAGAGAGGCATGCATGTAC

WeF_D12.ab1 ------WpF_E04.ab1 GGTTACCGGAGTACTCGTCATCCTGAACTGCAATGTGCACTATGCAACCGTACCTCGTAG FeF_E02.ab1 GGCATGTACGATGACTCGTCATCTGACTGCAATTGGCATAGTACGTACCCTCGGCATGCA FpF_E06.ab1 AAGAGGGTTCAATGGTACGAGACTCGATCATCTGACTGCAATGTGGCATCAGGCCACCGG FpR_E07.ab1 ------FeR_E03.ab1 GGAGGCCTGGAGGGAGAAGGAGGGGGTGCAGTGGGTACGGATGACCTCCCGGCTCCATCC WpR_E05.ab1 GGAGGCCTGGAGGGAGAAGGAGGGGGTGCAGTGGGTACGGATGACCTCCCGGCTCCATCC WeR_E01.ab1 GGAGGCCTGGAGGGAGAAGGAGGGGGTGCAGTGGGTACGGATGACCTCCCGGCTCCATCC

WeF_D12.ab1 ------WpF_E04.ab1 TCA------FeF_E02.ab1 TGCCACAGT------FpF_E06.ab1 TACCTCGCTATGG------FpR_E07.ab1 ------FeR_E03.ab1 CTGAACTGCAATATGTGGGCATCAGTGCAGCCGTACCCTCGCTATGGCATGCAAACAGTA WpR_E05.ab1 CTGAACTGCAATATGTGGGCATCAGTGCAGCCGTACCCTCGCTATGGCATGCAAACAGTA WeR_E01.ab1 CTGAACTGCAATATGTGGGCATCAGTTCAGCCGTACCCTCGCTATGGCATGCAAACAGTA

241 WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 GAGGCTATGCAATACCAGCCTTTCACAGCCCACTTCAACAGCACAGCCTCTGCAGCATCG WpR_E05.ab1 GAGGCTATGCAATACCAGCCTTTCACAGCCCACTTCAACAGCACAGCCTCTGCAGCATCG WeR_E01.ab1 GAGGCTATGCAATACCAGCCTTTCACAGCCCACTTCAACAGCACAGCCTCTGCAGCATCG

WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 ATGGTTTCCCATCACTCTC-A-TCCATGCAGCGTCCACATCCAACACCACCCGACTGTCA WpR_E05.ab1 ATGGTTTCCCATCACTCTCCA-TCCATGCAGCGTCCACATCCAACACCACCCGACCTTGT WeR_E01.ab1 ATGGTTTCCCATCACTCTC-A-TCCATGCAGCGTCCACATCCACACCACCCGACTTGTCA

WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 CATTTACCACACAGCGGGTACTGCCACCCACGCATCTCAGCTTCCACCTCCCCGTCTGGT WpR_E05.ab1 CACATTTACCACACAGCGGGTACTGCCACCCACGCCATCTTCAGCTTCCACCTCCCCGTC WeR_E01.ab1 CATTTACACACAGCGGGTACTGCACCCACGCATCTCAGCTCCACCTCCCCGTCTGGTCTG

WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 TCTGTCACCATGACAGAGCACATCTCTCTGTTTCATAGAAAGCTGGGTCTCAATACGTCA WpR_E05.ab1 TGTTCTGTCACATGACAGAGCACATTCCTCTCTGTTTCATAAAAAAGCTGGGTCTCAATA WeR_E01.ab1 TCACAATGAACGAGCACATCCTCTGGTTCATAGAAAGCTGATCTCCATACGTTCACAGAT

WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 CAAGGGATTTCACTGCTATCACCCAAGTCCTACCAAACGGAAACAGCTATCAGTACAACC WpR_E05.ab1 CGTTCACGAGGATTTCACTGCTATTCAACCCACAGTCCTACCCACCGGAACAGCTATCCG WeR_E01.ab1 TGCATGCCTAGTCAACCTCAGTCTACTCAACGGAACAGCTTCGTACACGGTTGGACAGTT

WeF_D12.ab1 ------WpF_E04.ab1 ------FeF_E02.ab1 ------FpF_E06.ab1 ------FpR_E07.ab1 ------FeR_E03.ab1 GTCTGACAGTTGCACTGACTGACGTTAACGTATCCACATAAAatcctcacatatctatc- WpR_E05.ab1 ATACAACAGTCTTGACCATATGCCCACTGACTTGACAGTTAAacgtatcgactataactt WeR_E01.ab1 GCATGACTGACGATAAACGTCTCCGACCATAATAAGTCCGTGacaaactctgtg------

242 Appendix F

Results from mosaic transgenic rescues.

Table F.1. A table of the results from the low dose mosaic transgenic rescue experiments

Clutch 1 Treatment n fin buds Left fin Right fin 2 fins no fins rescues 9.1.12 Uninj 20 0 0 0 0 20 0/20 Tbx4 FL 20 0 0 0 0 20 0/20 Tbx4 WT 20 10 5 4 1 10 10/20

Table F.2. Tables of the results from the high dose mosaic transgenic rescue experiments

# # fin Left # fin Right fin 2 rays No Treatment Individuals fin rays fin rays fins (l, r) fins Rescues Clutch 1 Uninj 13 0 0 0 13 0/13 16.1.13 Tbx4 FL 13 0 0 1 12 1/13 Tbx4 WT 13 3 1 0 9 4/13 Clutch 2 Uninj 18 1 8 1 2 1 1,2 15 3/18 31.1.13 Tbx4 FL 27 1 8 1 3 1 4,4 24 3/27 Tbx4 WT 24 1 3 2 2;0 2 8,8; 8,1 19 5/24 Clutch 3 Uninj 10 0 0 0 10 0/10 31.1.13 Tbx4 FL 11 0 1 8 1 8,3 9 1/11 Tbx4 WT 10 0 0 0 10 0/10 Clutch 4 Tbx4 FL 18 2 3;4 0 0 16 2/18 9.1.12 Tbx4 WT 14 4 2;3;8;8 1 8 0 9 5/14

Uninjected Wild-type tbx4 Pelvic finless tbx4 Total

Fins 3 14 7 24

No fins 38 47 62 147

Total 41 61 69 171

Comparison P-value

Uninjected vs. wild-type tbx4 0.038*

Pelvic finless tbx4 vs. wild-type tbx4 0.048*

Uninjected vs. pelvic finless tbx4 0.618

243 Appendix G

A list of all genes in the 12 Mb region identified by positional cloning.

244 Genes in Region

Gene Expression Limb/Fin Role in Limb Reference gosr1 ubiquitous Y N DEM1 unknown N mettl 16 unknown N pafah1b1a ubiquitous Y N zgc: 152873 ubiquitous Y (muscle) N (U1C alternative splicing) hif1al ubiquitous Y N tsr1 unknown ? hic1 ubiquitous Y Y (loss leads to stunted HL) Carter (2000) npat ubiquitous Y ? dph1 unknown Y (polydactyly in HL) (Webb 2008) rpa1 ubiquitous Y N sepinf2 unknown ppm1d ubiquitous Y N bcas3 unknown N tbx2b various Y Y tbx4 eye, pelvic fin Y Y (loss leads to loss of hindlimbs (Naiche and Papaioannou, 2007) brip1 (fancaj) various Y (AER) Y (loss leads to limb malformations), fanca g and i in AER not j (Titus 2009) lhx1a various N N dhrs13 ubiquitous Y N foxn1 thymus N N cryba1a NN crybb1l3 lens NN sarm1 immune N N zgc: 101731 brain N vtna unknown N dharma various N N scarf1 endothelial Y N naif1 unknown N prpf8 ubiquitous Y N slc43a2a unknown N pitpna various Y N slc6a4a brain N N blmh ubiquitous Y N si:ch211‐137a8.2 unknown xaf1 unknown N hsp47 ubiquitous Y Y (severly twisted) (Masago 2012) tsku ubiquitous Y Y (autopod) (Uejima 2010) hspa13 unknown N nrip1a ubiquitous Y N gucy2f retina N N samsn1a various N N nlk2 unknown N nos2b various Y N mrps23 ubiquitous Y N msi2b unknown N akaplb various Y (muscle) N lgals9l1 blood cells N N zgc: 92326 unknown zgc: 171951 unknown wsb1 ubiquitous Y Y (regulated by Shh) (Vasiliauskas 1999) arfip2a various N N zgc: 136924 unknown vezf1 unknown N N or116‐1 unknown N brca2 ubiquitous N Y (limb malformations) (Howlett 2002) arl4aa blood/brain N N vwde unknown N tmem106ba ubiquitous N N mag brain N N lsr various Y N zgc: 73340 ubiquitous Y mab21l1 brain Y N

245 Gene Expression Limb/Fin Role in Limb Reference postn muscle N Y (regeneration of HL muscle) (Goetsch 2003) mrpl44 unknown N agmo brain N N mff ubiquitous Y N sostdc1a unknown Y (polydactyly, KO interferes with Fgf and Shh) (Yee, 2011) ankmy2a unknown N tnfb ubiquitous Y N dhx16 ubiquitous Y N zgc: 66024 ubiquitous Y zgc: 114034 ubiquitous N agfg1a various N N sst1.1 various N N fam101b unknown Y (controled by sonic in limb artery (Bangs 2010) vps53 ubiquitous Y N tlcd2 unknown N rtn4rl1 brain, spine Y N myo1c unknown N nova2 unknown N ssh2a unknown N eml2 various N N capns1b various Y N hmgb1b various Y Y (in cartilage ‐> bone) (Taniguchi 2007) pds5b unknown Y (short limbs) (Zhang 2007) mmp13b unknown N frya unknown N etv1 brain N N trim3a unknown N fremb median fin Y Y (fin blistering), frem2a in pectoral fin AER, not frem2b (Carney 2010) tspan13a unknown N sh3bp5la ubiquitous Y N bag6 ubiquitous Y N Ita various N TNF gene mecom various Y N sesn3 YSL NN ispd head N N gabbr1a brain N N col4a4 skin NN

246 Appendix H

A list of all missense SNPs identified at the pelvic finless locus. Please note that these SNPs have not been verified in the pelvic finless population and represent missense SNPs identified in one pelvic finless individual.

247 name=AIPL1 24227138 ref=A alleles=C/C exonNum=6 mRNA_pos=1328 codonNum=443 codonChange='ATG=>AGG' residueChange='M=>R' name=AIPL1 24227148 ref=G alleles=C/C exonNum=6 mRNA_pos=1318 codonNum=440 codonChange='CCA=>GCA' residueChange='P=>A' name=AIPL1 24227240 ref=T alleles=A/A exonNum=6 mRNA_pos=1226 codonNum=409 codonChange='GAG=>GTG' residueChange='E=>V' name=protein_coding 24313445 ref=G alleles=T/T exonNum=1 mRNA_pos=157 codonNum=53 codonChange='CAA=>AAA' residueChange='Q=>K' name=protein_coding 24313564 ref=A alleles=G/G exonNum=1 mRNA_pos=38 codonNum=13 codonChange='TTA=>TCA' residueChange='L=>S' name=BX908796.1 24435626 ref=C alleles=T/T exonNum=3 mRNA_pos=103 codonNum=35 codonChange='GAA=>AAA' residueChange='E=>K' name=DEM1 24445159 ref=C alleles=T/T exonNum=3 mRNA_pos=230 codonNum=77 codonChange='CGG=>CAG' residueChange='R=>Q' name=hif1al 24836097 ref=G alleles=C/C exonNum=15 mRNA_pos=1706 codonNum=569 codonChange='TCC=>TGC' residueChange='S=>C' name=hif1al 24840754 ref=G alleles=C/C exonNum=9 mRNA_pos=1028 codonNum=343 codonChange='GCC=>GGC' residueChange='A=>G' name=hif1al 24840935 ref=C alleles=G/G exonNum=9 mRNA_pos=847 codonNum=283 codonChange='GAC=>CAC' residueChange='D=>H' name=NPAT 24864573 ref=C alleles=G/G exonNum=13 mRNA_pos=1136 codonNum=379 codonChange='GCC=>GGC' residueChange='A=>G' name=NPAT 24864582 ref=A alleles=T/T exonNum=13 mRNA_pos=1145 codonNum=382 codonChange='CAG=>CTG' residueChange='Q=>L' name=NPAT 24865101 ref=C alleles=A/A exonNum=14 mRNA_pos=1787 codonNum=596 codonChange='ACA=>AAA' residueChange='T=>K' name=NPAT 24865376 ref=A alleles=G/G exonNum=14 mRNA_pos=2062 codonNum=688 codonChange='AGT=>GGT' residueChange='S=>G' name=NPAT 24869594 ref=A alleles=G/G exonNum=20 mRNA_pos=3334 codonNum=1112 codonChange='ACA=>GCA' residueChange='T=>A' name=MMP20 24887070 ref=T alleles=G/G exonNum=1 mRNA_pos=92 codonNum=31 codonChange='CAA=>CCA' residueChange='Q=>P' name=TP53I13 24918667 ref=A alleles=G/G exonNum=3 mRNA_pos=188 codonNum=63 codonChange='ATT=>ACT' residueChange='I=>T' name=tsr1 24975790 ref=C alleles=T/T exonNum=5 mRNA_pos=797 codonNum=266 codonChange='GCG=>GTG' residueChange='A=>V' name=tsr1 24976010 ref=G alleles=T/T exonNum=5 mRNA_pos=1017 codonNum=339 codonChange='GAG=>GAT' residueChange='E=>D' name=rtn4rl1b 25736883 ref=G alleles=A/A exonNum=3 mRNA_pos=1193 codonNum=398 codonChange='GGA=>GAA' residueChange='G=>E' name=rpa1 25841836 ref=C alleles=A/A exonNum=16 mRNA_pos=1621 codonNum=541 codonChange='GCT=>TCT' residueChange='A=>S' name=rpa1 25870544 ref=G alleles=A/A exonNum=7 mRNA_pos=472 codonNum=158 codonChange='CCC=>TCC' residueChange='P=>S' name=WDR81 25903157 ref=C alleles=T/T exonNum=2 mRNA_pos=1015 codonNum=339 codonChange='GAC=>AAC' residueChange='D=>N' name=WDR81 25903246 ref=A alleles=G/G exonNum=2 mRNA_pos=926 codonNum=309 codonChange='ATG=>ACG' residueChange='M=>T' name=WDR81 25904367 ref=G alleles=A/A exonNum=1 mRNA_pos=253 codonNum=85 codonChange='CGT=>TGT' residueChange='R=>C' name=CR853282.3 25946538 ref=C alleles=A/A exonNum=15 mRNA_pos=1318 codonNum=440 codonChange='CTT=>ATT' residueChange='L=>I' name=CR853282.4 25952926 ref=T alleles=C/C exonNum=17 mRNA_pos=1714 codonNum=572 codonChange='ATC=>GTC' residueChange='I=>V' name=CR853282.2 25993681 ref=T alleles=A/A exonNum=1 mRNA_pos=22 codonNum=8 codonChange='AGG=>TGG' residueChange='R=>W' name=protein_coding 26209719 ref=G alleles=A/A exonNum=19 mRNA_pos=2288 codonNum=763 codonChange='CCA=>CTA' residueChange='P=>L' name=protein_coding 26211496 ref=T alleles=A/A exonNum=17 mRNA_pos=2027 codonNum=676 codonChange='CAA=>CTA' residueChange='Q=>L' name=protein_coding 26211515 ref=T alleles=C/C exonNum=17 mRNA_pos=2008 codonNum=670 codonChange='ACT=>GCT' residueChange='T=>A' name=protein_coding 26212866 ref=G alleles=T/T exonNum=16 mRNA_pos=1950 codonNum=650 codonChange='CAC=>CAA' residueChange='H=>Q' name=protein_coding 26215201 ref=T alleles=G/G exonNum=14 mRNA_pos=1812 codonNum=604 codonChange='AAA=>AAC' residueChange='K=>N' name=protein_coding 26215256 ref=G alleles=A/A exonNum=14 mRNA_pos=1757 codonNum=586 codonChange='GCA=>GTA' residueChange='A=>V' name=protein_coding 26215825 ref=T alleles=C/C exonNum=12 mRNA_pos=1671 codonNum=557 codonChange='ATA=>ATG' residueChange='I=>M' name=protein_coding 26219767 ref=G alleles=C/C exonNum=5 mRNA_pos=1061 codonNum=354 codonChange='CCA=>CGA' residueChange='P=>R' name=protein_coding 26219902 ref=A alleles=G/G exonNum=5 mRNA_pos=926 codonNum=309 codonChange='ATG=>ACG' residueChange='M=>T' name=protein_coding 26221011 ref=T alleles=C/C exonNum=2 mRNA_pos=559 codonNum=187 codonChange='AAT=>GAT' residueChange='N=>D' name=protein_coding 26221011 ref=T alleles=C/C exonNum=2 mRNA_pos=559 codonNum=187 codonChange='AAT=>GAT' residueChange='N=>D' name=CCDC9 26257896 ref=A alleles=G/G exonNum=9 mRNA_pos=1058 codonNum=353 codonChange='ATG=>ACG' residueChange='M=>T' name=CCDC9 26257907 ref=A alleles=G/G exonNum=2 mRNA_pos=76 codonNum=26 codonChange='TGG=>CGG' residueChange='W=>R' name=CCDC9 26262136 ref=C alleles=A/A exonNum=5 mRNA_pos=542 codonNum=181 codonChange='GGT=>GTT' residueChange='G=>V' name=CCDC9 26262410 ref=A alleles=C/C exonNum=5 mRNA_pos=268 codonNum=90 codonChange='TTT=>GTT' residueChange='F=>V' name=bcas3 26293092 ref=C alleles=A/A exonNum=6 mRNA_pos=385 codonNum=129 codonChange='CAT=>AAT' residueChange='H=>N' name=tbx4 26740752 ref=C alleles=T/T exonNum=3 mRNA_pos=233 codonNum=78 codonChange='GCA=>GTA' residueChange='A=>V' name=tbx4 26740755 ref=G alleles=C/C exonNum=3 mRNA_pos=236 codonNum=79 codonChange='GGC=>GCA' residueChange='G=>A' name=tbx4 26740756 ref=C alleles=A/A exonNum=3 mRNA_pos=237 codonNum=79 codonChange='GGC=>GCA' residueChange='G=>A' name=cryba1a 27441491 ref=C alleles=A/A exonNum=3 mRNA_pos=102 codonNum=34 codonChange='CAG=>CAT' residueChange='Q=>H' name=crybb1l3 27449575 ref=G alleles=A/A exonNum=4 mRNA_pos=290 codonNum=97 codonChange='CGC=>CAC' residueChange='R=>H' name=foxn1 27483701 ref=G alleles=T/T exonNum=7 mRNA_pos=1240 codonNum=414 codonChange='CCC=>ACC' residueChange='P=>T' name=foxn1 27495555 ref=T alleles=C/C exonNum=2 mRNA_pos=181 codonNum=61 codonChange='AAA=>GAA' residueChange='K=>E' name=slc46a1 27524723 ref=A alleles=T/T exonNum=1 mRNA_pos=85 codonNum=29 codonChange='ACA=>TCA' residueChange='T=>S' name=slc46a1 27527140 ref=A alleles=G/G exonNum=2 mRNA_pos=578 codonNum=193 codonChange='GAA=>GGA' residueChange='E=>G' name=sarm1 27544496 ref=C alleles=G/G exonNum=3 mRNA_pos=1188 codonNum=396 codonChange='GAG=>GAC' residueChange='E=>D' name=scarf1 27559937 ref=A alleles=G/G exonNum=10 mRNA_pos=2413 codonNum=805 codonChange='TGT=>CGT' residueChange='C=>R' name=scarf1 27559982 ref=T alleles=C/C exonNum=10 mRNA_pos=2368 codonNum=790 codonChange='AGA=>GGA' residueChange='R=>G' name=scarf1 27560439 ref=G alleles=A/A exonNum=3 mRNA_pos=578 codonNum=193 codonChange='TCC=>TTC' residueChange='S=>F' name=scarf1 27560781 ref=G alleles=A/A exonNum=3 mRNA_pos=236 codonNum=79 codonChange='GCC=>GTC' residueChange='A=>V' name=scarf1 27566044 ref=A alleles=C/C exonNum=7 mRNA_pos=1256 codonNum=419 codonChange='GTT=>GGT' residueChange='V=>G' name=scarf1 27566407 ref=A alleles=T/T exonNum=5 mRNA_pos=1059 codonNum=353 codonChange='CAT=>CAA' residueChange='H=>Q' name=scarf1 27566491 ref=G alleles=T/T exonNum=5 mRNA_pos=975 codonNum=325 codonChange='CAC=>CAA' residueChange='H=>Q' name=myo1c 27692574 ref=C alleles=T/T exonNum=26 mRNA_pos=2783 codonNum=928 codonChange='TCA=>TTA' residueChange='S=>L' name=CR478288.1 27705016 ref=G alleles=A/A exonNum=1 mRNA_pos=257 codonNum=86 codonChange='GGG=>GAG' residueChange='G=>E' name=CR478288.1 27705284 ref=T alleles=A/A exonNum=1 mRNA_pos=525 codonNum=175 codonChange='AGT=>AGA' residueChange='S=>R' name=ssh2a 27898756 ref=A alleles=C/C exonNum=14 mRNA_pos=2709 codonNum=903 codonChange='TTT=>TTG' residueChange='F=>L' name=ssh2a 27899015 ref=T alleles=C/C exonNum=14 mRNA_pos=2450 codonNum=817 codonChange='CAT=>CGT' residueChange='H=>R' name=ssh2a 27900217 ref=G alleles=C/C exonNum=13 mRNA_pos=1664 codonNum=555 codonChange='TCT=>TGT' residueChange='S=>C' name=eml2 28243314 ref=C alleles=A/A exonNum=6 mRNA_pos=522 codonNum=174 codonChange='GAG=>GAT' residueChange='E=>D' name=eml2 28256801 ref=T alleles=C/C exonNum=1 mRNA_pos=8 codonNum=3 codonChange='GAA=>GGA' residueChange='E=>G' name=TMEM87A 28449717 ref=G alleles=A/A exonNum=6 mRNA_pos=494 codonNum=165 codonChange='CCA=>CTA' residueChange='P=>L' name=TMEM87A 28461057 ref=C alleles=T/T exonNum=4 mRNA_pos=335 codonNum=112 codonChange='AGC=>AAC' residueChange='S=>N' name=TMEM87A 28461063 ref=C alleles=T/T exonNum=4 mRNA_pos=329 codonNum=110 codonChange='AGC=>AAC' residueChange='S=>N' name=NAIF1 28497911 ref=A alleles=T/T exonNum=3 mRNA_pos=473 codonNum=158 codonChange='CAA=>CTA' residueChange='Q=>L' name=NAIF1 28498046 ref=C alleles=T/T exonNum=3 mRNA_pos=608 codonNum=203 codonChange='TCT=>TTT' residueChange='S=>F' name=XAF1 28505519 ref=G alleles=T/T exonNum=5 mRNA_pos=311 codonNum=104 codonChange='GCC=>GAC' residueChange='A=>D' name=gucy2f 28715041 ref=A alleles=T/T exonNum=2 mRNA_pos=973 codonNum=325 codonChange='ATC=>TTC' residueChange='I=>F' name=GDPD5 28800698 ref=A alleles=T/T exonNum=3 mRNA_pos=313 codonNum=105 codonChange='ATG=>TTG' residueChange='M=>L' name=samsn1a 28916408 ref=T alleles=A/A exonNum=15 mRNA_pos=1617 codonNum=539 codonChange='GAA=>GAT' residueChange='E=>D' name=samsn1a 28916835 ref=C alleles=G/G exonNum=14 mRNA_pos=1524 codonNum=508 codonChange='GAG=>GAC' residueChange='E=>D' name=nrip1a 29060453 ref=T alleles=A/A exonNum=1 mRNA_pos=1978 codonNum=660 codonChange='TTA=>ATA' residueChange='L=>I' name=nrip1a 29060463 ref=A alleles=G/G exonNum=1 mRNA_pos=1988 codonNum=663 codonChange='AAA=>AGA' residueChange='K=>R' name=nrip1a 29061291 ref=G alleles=A/A exonNum=1 mRNA_pos=2816 codonNum=939 codonChange='AGT=>AAT' residueChange='S=>N' name=nrip1a 29061654 ref=T alleles=C/C exonNum=2 mRNA_pos=3179 codonNum=1060 codonChange='TTG=>TCG' residueChange='L=>S' 248 name=NUFIP2 29512547 ref=C alleles=G/G exonNum=2 mRNA_pos=1531 codonNum=511 codonChange='CCA=>GCA' residueChange='P=>A' name=nos2b 29716647 ref=C alleles=T/T exonNum=13 mRNA_pos=1421 codonNum=474 codonChange='GCA=>GTA' residueChange='A=>V' name=mrps23 29731619 ref=G alleles=C/C exonNum=2 mRNA_pos=84 codonNum=28 codonChange='GAG=>GAC' residueChange='E=>D' name=or102-1 30760507 ref=C alleles=T/T exonNum=1 mRNA_pos=104 codonNum=35 codonChange='ACG=>ATG' residueChange='T=>M' name=or102-1 30760598 ref=C alleles=G/G exonNum=1 mRNA_pos=195 codonNum=65 codonChange='TTC=>TTG' residueChange='F=>L' name=or102-1 30760641 ref=G alleles=T/T exonNum=1 mRNA_pos=238 codonNum=80 codonChange='GCA=>TCA' residueChange='A=>S' name=or102-3 30770293 ref=T alleles=G/G exonNum=1 mRNA_pos=470 codonNum=157 codonChange='GTG=>GGG' residueChange='V=>G' name=or102-4 30775312 ref=G alleles=A/A exonNum=1 mRNA_pos=921 codonNum=307 codonChange='ATG=>ATA' residueChange='M=>I' name=AL929092.4 30800582 ref=T alleles=A/A exonNum=4 mRNA_pos=885 codonNum=295 codonChange='CAA=>CAT' residueChange='Q=>H' name=frya 31078674 ref=C alleles=T/T exonNum=8 mRNA_pos=734 codonNum=245 codonChange='ACG=>ATG' residueChange='T=>M' name=brca2 31163139 ref=G alleles=C/C exonNum=10 mRNA_pos=994 codonNum=332 codonChange='GAA=>CAA' residueChange='E=>Q' name=brca2 31166562 ref=T alleles=C/C exonNum=11 mRNA_pos=4334 codonNum=1445 codonChange='TTA=>TCA' residueChange='L=>S' name=brca2 31170482 ref=G alleles=A/A exonNum=21 mRNA_pos=7298 codonNum=2433 codonChange='AGG=>AAG' residueChange='R=>K' name=brca2 31174436 ref=T alleles=G/G exonNum=27 mRNA_pos=8514 codonNum=2838 codonChange='AAT=>AAG' residueChange='N=>K' name=arfip2a 31512258 ref=T alleles=C/C exonNum=5 mRNA_pos=638 codonNum=213 codonChange='ATG=>ACG' residueChange='M=>T' name=TEKT1 31999229 ref=C alleles=T/T exonNum=7 mRNA_pos=887 codonNum=296 codonChange='CGA=>CAA' residueChange='R=>Q' name=INPP5K 32010491 ref=C alleles=T/T exonNum=13 mRNA_pos=1357 codonNum=453 codonChange='GCT=>ACT' residueChange='A=>T' name=STARD13 32403584 ref=A alleles=T/T exonNum=15 mRNA_pos=4118 codonNum=1373 codonChange='TAT=>TTT' residueChange='Y=>F' name=postnb 33140184 ref=T alleles=C/C exonNum=17 mRNA_pos=2098 codonNum=700 codonChange='AAG=>GAG' residueChange='K=>E' name=postnb 33140205 ref=T alleles=C/C exonNum=17 mRNA_pos=2077 codonNum=693 codonChange='ACG=>GCG' residueChange='T=>A' name=postnb 33161298 ref=G alleles=T/T exonNum=2 mRNA_pos=215 codonNum=72 codonChange='ACA=>AAA' residueChange='T=>K' name=frem2b 33402331 ref=A alleles=G/G exonNum=7 mRNA_pos=5889 codonNum=1963 codonChange='ATA=>ATG' residueChange='I=>M' name=frem2b 33405596 ref=G alleles=A/A exonNum=9 mRNA_pos=6274 codonNum=2092 codonChange='GGA=>AGA' residueChange='G=>R' name=frem2b 33428755 ref=C alleles=G/G exonNum=23 mRNA_pos=8846 codonNum=2949 codonChange='ACA=>AGA' residueChange='T=>R' name=agmo 33631498 ref=C alleles=G/G exonNum=13 mRNA_pos=1301 codonNum=434 codonChange='TGT=>TCT' residueChange='C=>S' name=agmo 33631532 ref=T alleles=C/C exonNum=13 mRNA_pos=1267 codonNum=423 codonChange='ATC=>GTC' residueChange='I=>V' name=CT573389.1 34230880 ref=A alleles=T/T exonNum=1 mRNA_pos=58 codonNum=20 codonChange='ACA=>TCA' residueChange='T=>S' name=col4a4 35263287 ref=C alleles=T/T exonNum=19 mRNA_pos=1226 codonNum=409 codonChange='AGC=>AAC' residueChange='S=>N' name=col4a4 35263330 ref=T alleles=A/A exonNum=19 mRNA_pos=1183 codonNum=395 codonChange='ATG=>TTG' residueChange='M=>L' name=RNH1 35633561 ref=G alleles=A/A exonNum=5 mRNA_pos=644 codonNum=215 codonChange='ACA=>ATA' residueChange='T=>I' name=RNH1 35636054 ref=C alleles=T/T exonNum=3 mRNA_pos=334 codonNum=112 codonChange='GAT=>AAT' residueChange='D=>N' name=RNH1 35636088 ref=C alleles=T/T exonNum=3 mRNA_pos=300 codonNum=100 codonChange='ATG=>ATA' residueChange='M=>I'

249