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Modeling human gene variants that affect WNT signalling in the chicken embryo Gignac, Sarah J. 2019

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    Modeling human gene variants that affect WNT signalling in the  chicken embryo  by  Sarah J. Gignac  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSPHY  in  The Faculty of Graduate and Postdoctoral Studies  (Cell and Developmental Biology)   THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2019  © Sarah J. Gignac, 2019    ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled:  Modeling human gene variants affecting non-canonical WNT signaling in the chicken embryo  submitted by Sarah J. Gignac  in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Cell and Developmental Biology  Examining Committee: Joy Richman, Oral Health Sciences Supervisor  Michael Underhill, Cellular & Physiological Sciences Supervisory Committee Member  Timothy O’Connor, Cellular & Physiological Sciences Supervisory Committee Member Anna Lehman, Medical Genetics University Examiner Cheryl Gregory-Evans, Ophthalmology and Visual Sciences University Examiner   Sigmar Stricker, Biochemistry and Genetics Additional Examiner  Additional Supervisory Committee Members: Janel Kopp, Cellular & Physiological Sciences Supervisory Committee Member       iii Abstract  The study of rare genetic diseases provides valuable insights into human gene function. The chicken embryo was used as a model to investigate the role of WNT signaling in skeletogenesis and to elucidate the functional consequences of mutations in dominant Robinow Syndrome (RS). RS mutations affect non-canonical WNT signaling that controls a variety of developmental events to regulate convergent extension, cell polarity, and cytoskeletal rearrangement. RS is characterized by short stature, mesomelic limb shortening, hypertelorism, and mandibular hypoplasia. Mutations in dominant RS occur in several components of the non-canonical WNT signaling pathway, and this study is focusing on two mutations in WNT5A ligand (WNT family member) and three mutations in Dishevelled1 (DVL1), a protein that relays WNT signals intracellularly. We delivered the human genes to the chicken embryo using replication competent retroviruses (RCAS) and analyzed morphologic, cellular, and molecular effects in the forelimbs and mandible. Misexpression of mutants in dominant RS led to a shortening of the forelimb and mandible and caused polarity disruptions in the chondrocytes that were not seen in the GFP virus controls. The variants were unable to activate canonical WNT signaling and over-activated non-canonical WNT signaling, demonstrating the importance of non-canonical WNT signaling in skeletogenesis. Dominant RS mutations have dominant neomorphic effects on chondrogenesis that interfere with the function of the wild-type protein. This work establishes that the dominant effect of the mutations leads to elevated non-canonical WNT signaling and randomizes the distribution of planar cell polarity molecules of which produces shortened skeletal elements.      iv Lay Summary  Disorders affecting growth of the skeleton often result from a genetic or medical condition with >450 known disorders. Skeletal disorders often have differences in the size/shape of affected bones and are classified based on which part of the skeleton is involved. The discovery of genes that affect skeletal development is progressing, though there is still a need to understand how these genes function. We are studying a rare syndrome called Robinow Syndrome (RS) and patients have reduced outgrowth of limbs and facial abnormalities. Mutated genes that cause RS affect a cascade of chemical reactions that instruct a cell how to behave. These mutant genes are related to the ‘Wingless’ gene and they regulate cell proliferation, migration and shape. We put the human mutated genes of RS into the chicken embryo and found that limb and lower jaw outgrowth were disrupted along with disorganized cartilage cells, creating misshapened and shortened bones.        v Preface All of the work presented hereafter was conducted in the Life Sciences Institute at the University of British Columbia, Point Grey campus.   I was the lead investigator for the limb project located in Chapter 3 where I was responsible for all major areas of concept formation, data collection and analysis, as well as the majority of manuscript composition. The limb project was initially started by Sara Hosseini-Farahabadi with work conducted during her PhD thesis (ubc_2014_spring_hosseinifarahabadi_sara). Sara HF performed preliminary experiments for skeletal analysis. Nathan Schuck also worked alongside Sara HF during the initial stages of the limb project and aided with some of the histological work. Katherine Fu was involved in cloning constructs for plasmids and viruses as well as helped with histology. The western blot data in the thesis is my own data, but another student (Takashi Akazawa) increased the blot sample size for densitometry analysis. Joy Richman was the supervisory author on this project and was involved throughout the project in concept formation and manuscript edits. The WNT5A limb project in chapter 3 has been accepted for publication [Gignac, S. J., Hosseini-Farahabadi, S., Akazawa, T., Schuck, N.J., Fu, K., & Richman, J. M. (2019). Robinow syndrome skeletal phenotypes caused by the WNT5AC83S variant are due to dominant interference with chondrogenesis. Human Molecular Genetics, ddz071.] and includes work conducted on the limb with wtWNT5A and the C83S variant. I am sole first author on this manuscript.   A section of Chapter 3 has been published [Hosseini-Farahabadi, S., Gignac, S. J., Danescu, A., Fu, K., & Richman, J. M. (2017). Abnormal WNT5A Signaling Causes Mandibular Hypoplasia in Robinow Syndrome. Journal of Dental Research, 96(11), 1265–1272.] and includes the work conducted solely on the mandible. I was first co-author responsible for increasing sample size of some of the experiments as well as performing cell proliferation, cell polarity and cell shape experiments which included analysis of data that I conducted, as well as manuscript composition. Sara Hosseini-Farahabadi was involved in the early stages of this project during her PhD thesis (ubc_2014_spring_hosseinifarahabadi_sara) and contributed to performing the initial experiments in the chicken embryo which also included cell migration and luciferase assays as well as manuscript edits. Adrian Danescu also contributed to the project with morphometric analysis, as    vi well as work from Katherine Fu who performed all the viral cloning and qPCR as well as assisted with immunofluorescence experiments to drive the project forward. Both AD and KF also contributed to manuscript edits. Joy Richman was the supervisory author on this project and was involved throughout the project in concept formation and manuscript composition.   I was the lead investigator for the project located in Chapter 4 where I was responsible for all major areas of concept formation, data collection and analysis, as well as the majority of manuscript composition. I will be sole first author on the DVL1 manuscript. Katherine Fu was involved in the early stages of concept formation and performed all cloning of constructs for plasmids and viruses as well as helped with initial histology. Joy Richman was the supervisory author on this project and was involved throughout the project in concept formation and manuscript edits.         vii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary ............................................................................................................................... iv Preface ............................................................................................................................................ v Table of Contents ........................................................................................................................ vii List of Tables ................................................................................................................................. x List of Figures ............................................................................................................................... xi List of Abbreviations ................................................................................................................. xiii Acknowledgements ................................................................................................................... xvii 1. Introduction .................................................................................................... 1 1.1 Overview........................................................................................................................ 1 1.2 Limb development in the chicken embryo ................................................................. 2 1.2.1 Limb bud outgrowth and patterning in the limb ............................................................. 2 1.2.2 Molecular regulation of the embryonic limb .................................................................. 3 1.2.3 Chondrogenesis and skeletogenesis in the limb ............................................................. 5 1.2.4 Facial development in the chicken embryo and molecular regulation of the mandible . 6 1.2.5 Chondrogenesis and skeletogenesis in the face .............................................................. 7 1.3 Genetic Skeletal Dysplasias ......................................................................................... 8 1.3.1 Robinow Syndrome ........................................................................................................ 9 1.4 Many branches of WNT signaling ............................................................................ 10 1.4.1 WNT ligands ................................................................................................................. 10 1.4.2 WNT synthesis and secretion ....................................................................................... 11 1.4.3 WNT receptors .............................................................................................................. 12 1.4.4 WNT signaling intermediates (Dishevelled) ................................................................ 14 1.4.5 WNT signaling .............................................................................................................. 15 1.4.5.1 Canonical WNT signaling................................................................................................. 16 1.4.5.2 Non-canonical WNT signaling ......................................................................................... 17 1.4.6 Endpoints for WNT signaling ....................................................................................... 20 1.5 Animal Models to study skeletal dysplasias ............................................................. 22 1.6 Local transgenesis in chicken embryos .................................................................... 24 1.7 Rationale...................................................................................................................... 26 1.8 Approach ..................................................................................................................... 27 1.9 Hypotheses .................................................................................................................. 27 1.10 Novelty ......................................................................................................................... 28 1.11 Aims and Objectives ................................................................................................... 28 2. Materials and Methods ................................................................................29 2.1 Chicken embryos ........................................................................................................ 29 2.2 Cloning of WNT5A virus and plasmid constructs ................................................... 29 2.3 Cloning of DVL1 virus and plasmid constructs ....................................................... 30 2.4 Retrovirus construction of WNT5A and DVL1 variants ......................................... 30 2.5 Retrovirus injection into the limb and face.............................................................. 31 2.6 Whole-mount skeletal staining .................................................................................. 32    viii 2.7 Histology ...................................................................................................................... 33 2.8 Immunofluorescence .................................................................................................. 33 2.9 Quantification of chondrocyte polarity and shape .................................................. 38 2.10 BrdU, TUNEL staining and analysis ........................................................................ 38 2.11 Quantification of forelimb cell density and morphometrics .................................. 39 2.12 Contrast enhanced micro CT of limb buds and volumetric analysis .................... 39 2.13 Cell culture and Luciferase reporter assay .............................................................. 40 2.14 Immunocytochemistry with transfected WNT5A or DVL1 variants ..................... 42 2.15 Western blot ................................................................................................................ 43 2.16 Statistical analysis....................................................................................................... 44 3. Investigations of WNT5A variants that cause Robinow Syndrome ........45 3.1 Introduction ................................................................................................................ 45 3.2 Results.......................................................................................................................... 47 3.2.1 Mutant WNT5AC83S variant shortens the long bone and delay ossification .................. 47 3.2.2 Mutant WNT5A shortens and widens dimensions of the developing long bone ......... 49 3.2.3 WNT5AC83S virus showed increased cell density in the diaphysis of the ulna .............. 54 3.2.4 Phenotypic changes are first visible 4 days post-infection with RCAS viruses ........... 56 3.2.5 Volume of cartilage is not changed by the RCAS::WNT5AC83S ................................... 57 3.2.6 wtWNT5A and WNT5AC83S show increased cell proliferation in the cartilage ............. 58 3.2.7 WNT5AC83S disrupts chondrocyte polarity and cell elongation in forelimbs ................ 61 3.2.8 WNT5A mutations cause shortening of the mandible ................................................... 64 3.2.9 WNT5A mutations do not affect cell proliferation in the mandible or in Meckel’s cartilage ......................................................................................................................... 66 3.2.10 WNT5A mutations randomize chondrocyte polarity in Meckel’s cartilage .................. 68 3.2.11 WNT5A mutations disrupt chondrocyte cell shape in Meckel’s cartilage .................... 69 3.2.12 WNT5AC83S is secreted much less efficiently than wtWNT5A .................................... 70 3.2.13 Transient transfections of WNT5A plasmids show similar transfection rates ............... 72 3.2.14 WNT5A variants super activate JNK-PCP signaling ..................................................... 73 3.2.15 Mutant WNT5A is just as effective at antagonizing canonical WNT signaling as wtWNT5A .................................................................................................................... 76 3.2.16 WNT5A variants are unable to antagonize WNT3A stimulated signaling unless Ror2 is provided ........................................................................................................................ 77 3.2.17 wtWNT5A and mutant WNT5A do not activate calcium WNT signaling .................... 78 3.3 Discussion .................................................................................................................... 80 3.3.1 Defects during chondrogenesis combine to give shorter limbs in RS .......................... 80 3.3.2 Mutant and Wild-type forms of WNT5A reduce outgrowth of Meckel’s cartilage ..... 82 3.3.3 Comparisons between limb and mandible phenotypes ................................................. 83 3.3.4 Genotype-phenotype correlations of WNT5A variants in the chicken model .............. 83 3.3.5 The C83S mutation decreases protein levels intra and extracellularly ......................... 84 3.3.6 Antagonism of canonical WNT signaling may not play a role in the dominant RS phenotype ...................................................................................................................... 86 3.3.7 Dominant RS WNT5A variants are the result of a neomorphic function ..................... 87    ix 4. DVL1 variants disrupt chondrogenesis and overactivate PCP signaling ....................................................................................................90 4.1 Introduction ................................................................................................................ 90 4.2 Results.......................................................................................................................... 93 4.2.1 DVL1 variants affect limb development ....................................................................... 93 4.2.2 DVL1 variants create a pinching phenotype of the perichondrium and delay chondrocyte hypertrophy .............................................................................................. 97 4.2.3 DVL1 variants do not affect cell proliferation ............................................................ 102 4.2.4 DVL1 variant 1529ΔG randomizes cell polarity ......................................................... 104 4.2.5 Transient transfections of DVL1 plasmids show functional variants ......................... 109 4.2.6 DVL11529ΔG overactivates JNK-PCP activity .............................................................. 110 4.2.7 DVL11529ΔG does not activate WNT calcium signaling ............................................... 111 4.2.8 DVL11529ΔG does not activate endogenous canonical WNT signaling ........................ 112 4.3 Discussion .................................................................................................................. 115 4.3.1 Defects during chondrogenesis combine to give limb phenotype in RS .................... 115 4.3.2 Genotype-phenotype correlations in the chicken model for DVL1 variants .............. 117 4.3.3 Planar cell polarity signaling is disrupted in the chicken model for RS ..................... 117 4.3.4 Biochemical defects caused by DVL1 variants are a complex mixture of gain-of-function in the JNK-PCP pathway concomitant with a loss-of-function in the canonical WNT signaling pathway and no change in the Calcium signaling pathway .............. 118 5. General Discussion .................................................................................121 5.1 Strengths and weaknesses of the chicken model system as compared to the mouse .................................................................................................................................... 121 5.2 Comparison of the effects of the WNT5A and DVL1 mutations and how these inform us about the pathogenesis of RS ................................................................. 122 5.3 Differences between WNT5A and DVL1 mutations suggest other mediators may be involved................................................................................................................. 123 5.4 Debunking the myth that Dominant Robinow Syndrome, is caused by hypomorphic effects of the WNT5A gene mutations ............................................ 126 5.5 Overall significance .................................................................................................. 127 5.6 Future directions ...................................................................................................... 127 5.7 Concluding remarks ................................................................................................. 131 References .................................................................................................................................. 132       x List of Tables Table 1.1. WNT ligands, receptors and components of WNT signaling. ..................................... 15 Table 2.1. Primer sequences for WNT5A variants. ...................................................................... 29 Table 2.2. Primer sequences for DVL1 variants. .......................................................................... 30 Table 2.3. Embryo stage and fixative post-injection. ................................................................... 32 Table 2.4. Antibodies and immunofluorescence treatments for forelimb WNT5A assays. ......... 35 Table 2.5. Antibodies and immunofluorescence treatments for mandible WNT5A assays. ........ 36 Table 2.6. Antibodies and immunofluorescence treatments for forelimb DVL1 assays. ............. 37 Table 2.7. Plasmids and stimulates used for luciferase assays on HEK293 cells. ........................ 42 Table 3.1. Qualitative analysis of WNT5A forelimb phenotype at stage HH38. ......................... 49 Table 3.2. Qualitative analysis of mandibular phenotype. ............................................................ 66 Table 3.3. Summary of the dominant effects of WNT5A mutations on biological function. ...... 88 Table 4.1. Qualitative analysis of DVL1 forelimb phenotype at stage HH38. ............................. 96 Table 4.2. Qualitative analysis of DVL1 forelimb phenotype at stage HH34. ........................... 100 Table 4.3. Qualitative analysis of DVL1 forelimb phenotype at stage HH29. ........................... 102 Table 4.4. Summary of the dominant effects of DVL1 mutations on biological function. ........ 120       xi List of Figures Figure 1.1. Canonical WNT signaling pathways. ......................................................................... 17 Figure 1.2. Non-canonical WNT signaling pathways. .................................................................. 20 Figure 3.1. Skeletal phenotypes obtained from misexpression of WNT5A retroviruses in the forelimb. ..................................................................................................................... 48 Figure 3.2. Mutant WNT5A shortens and widens dimensions of the developing long bone. ....... 52 Figure 3.3. WNT5A viruses widen the ulna in chicken embryos. ................................................. 54 Figure 3.4. Variant WNT5A viruses showed increased cell density in both the diaphysis and epiphysis of the ulna. ................................................................................................. 56 Figure 3.5. Phenotypic changes are first visible 4-5 days post infection with WNT5AC83S. ......... 56 Figure 3.6. Embryo forelimb skeletal elements show similar volume four days post-injection. . 57 Figure 3.7. wtWNT5A or WNT5AC83S injected forelimbs do not affect cell proliferation at stage HH29 in the developing ulna. .................................................................................... 59 Figure 3.8. wtWNT5A or WNT5AC83S injected forelimbs have increased cell proliferation in the cartilage at stage HH30. ............................................................................................. 60 Figure 3.9. wtWNT5A or WNT5AC83S injected forelimbs do not affect apoptosis in the cartilage.61 Figure 3.10. WNT5AC83S virus randomizes chondrocyte polarity and causes rounder chondrocyte shape in the forelimb. ................................................................................................. 63 Figure 3.11. WNT5AC83S virus randomizes chondrocyte PCP molecule, Prickle in the forelimb. 64 Figure 3.12. Skeletal phenotypes obtained from misexpression of WNT5A retroviruses in the mandible. .................................................................................................................... 65 Figure 3.13. WNT5A variants do not affect cell proliferation in the mandible. ........................... 68 Figure 3.14. WNT5A mutant viruses alter chondrocyte polarity in the mandible. ........................ 69 Figure 3.15. WNT5A mutant viruses alter chondrocyte shape in Meckel’s cartilage. .................. 70 Figure 3.16. WNT5AC83S shows less protein in the cell lysate than wtWNT5A. ......................... 71 Figure 3.17. WNT5AC83S is secreted much less efficiently than wtWNT5A. .............................. 72 Figure 3.18. WNT5A labeling of transfected cells in HEK293 cells. .......................................... 73 Figure 3.19. WNT5A variants activate ATF2 luciferase reporter. ................................................. 75 Figure 3.20. WNT5A variants do not activate STF luciferase reporter. ........................................ 76 Figure 3.21. WNT5AC83S partially mediates inhibitory activity in canonical WNT signaling. ..... 78 Figure 3.22. WNT5A constructs do not activate calcium WNT signaling. ................................... 79 Figure 4.1. Skeletal phenotypes obtained from misexpression of DVL1 retroviruses in the forelimb. ..................................................................................................................... 95    xii Figure 4.2. Mutant DVL1 delays ossification and creates a pinching phenotype in the perichondrium. ........................................................................................................... 98 Figure 4.3. Near-adjacent sections of forelimbs injected with DVL1 viruses. ............................. 99 Figure 4.4. Mutant DVL1 injected forelimbs are unable to form intact cartilage elements. ....... 101 Figure 4.5. Wild-type DVL1 and DVL1 variants injected forelimbs do not affect cell proliferation at stage HH29 in the developing ulna. ..................................................................... 104 Figure 4.6. DVL11529ΔG virus randomizes chondrocyte polarity in the forelimb. ....................... 106 Figure 4.7. DVL1 variants randomize chondrocyte PCP molecule, Prickle in the forelimb. ..... 108 Figure 4.8. DVL1 labeling of transfected cells in HEK293 cells. .............................................. 109 Figure 4.9. DVL11529ΔG overactivates non-canonical PCP signaling. ......................................... 110 Figure 4.10. DVL11529ΔG partially activates calcium WNT signaling. ........................................ 112 Figure 4.11. DVL11529ΔG does not activate STF luciferase reporter. ........................................... 114 Figure 5.1. Summary of the dominant effects of WNT5A and DVL1 mutations on skeletal morphogenesis. ........................................................................................................ 126         xiii List of Abbreviations AER   Apical Ectodermal Ridge ACAN   Aggrecan AlkPO4  Alkaline Phosphatase ANOVA  Analysis of variance AP    Anterior-posterior AP1   Activator protein 1 APC    Adenomatous polyposis coli ASLV    Avian sarcoma-leukosis virus ATF    Activating transcription factor  BAT-GAL  β-catenin/TCF/LEF reporter transgenic mice BMP   Bone morphogenetic protein BrdU   Bromodeoxyuridine BSA   Bovine serum albumin Ca2+    Calcium CAMKII   Calcium/calmodulin-dependent protein kinase II caNFAT  Constitutively active NFAT cDNA   Complementary DNA CK1    Casein kinase 1α COL    Collagen type CRD    Cysteine-rich binding domain CREB    cAMP response element-binding protein CRISPR/Cas  Clustered regularly interspaced short palindromic repeats/CRISPR-associated CTTNB1  β-catenin DAAM1   Dishevelled associated activator of morphogenesis 1 DAPI   4′,6-diamidino-2-phenylindole DAX    DIX Axin domain DEP    DVL, EGL-10, Pleckstrin DIX    DVL/Axin Dsh   Dishevelled DV    Dorsal-ventral DVL   Dishevelled ECM    Extracellular matrix EDTA   Ethylenediaminetetraacetic acid EN1   Engrailed-1 ENV    Envelope ER    Endoplasmic reticulum FBS   Fetal bovine serum FGF    Fibroblast Growth Factors FGFR   FGF receptor FL   Forelimb FZD   Frizzled GAG    Group-associated antigens GAPDH  Glyceraldehyde 3-phosphate dehydrogenase  GFP   Green fluorescent protein    xiv GLI3   Glioma-associated oncogene family zinc finger 3 GM130  Golgi bodies GPC4   Glypican-4 GPR124   G protein coupled receptor 124 GREM1  Gremlin GS   Goat serum GSK3β   Glycogen synthase kinase 3β GTPase  Guanosine triphosphatase HAND2  Heart and neural crest derivatives expressed 1 HCl   Hydrochloric acid Hes1   Hairy and enhancer of split 1 HH    Hamburger Hamilton HOX    Homeobox HSPG    Heparan sulphate proteoglycans IHH    Indian hedgehog Int1    Integration  iPSC   Induced pluripotent stem cell IRE1   Inositol requiring 1 JNK    Jun N-terminal kinase KCNJ2  Potassium voltage-gated channel subfamily J member 2 kDa    Kilodalton  KOH   Potassium hydroxide kVp   Peak kilovoltage LEF    Lymphoid enhancer factor LiCl   Lithium chloride LMNA  Laminin A/C LMX1B  LIM Homeobox Transcription Factor 1 Beta LRP    Low-density lipoprotein receptor-related protein LSD   Least significant difference LTR    Long terminal repeat microCT  Micron-scale computed tomography MMTV   Mouse mammary tumour virus MSX2   MSH homeo box homolog 2 MUSK   Muscle skeletal receptor Tyr kinase n   Sample size N-CAM  Neural cell adhesion molecule NFAT    Nuclear factor associated with T cells NFκB   Nuclear factor kappa-light-chain-enhancer of activated B cells NKD   Naked NRH1   Neurotrophin receptor homolog 1 NXN   Nucleoredoxin OMIM   Online Mendelian Inheritance in Man OSX    Osterix p   Probability (p value) PANX3  Pannexin 3 PBS   Phosphate buffered saline    xv PCP    Planar cell polarity PD    Proximal-distal PDZ    Postsynaptic density 95, Discs Large, Zonula occludens-1 PERK   PKR-like ER kinase PFA   Paraformaldehyde PGC    Primordial germ cells PITX2   Pituitary homeobox 2 PKC    Protein kinase C PLC   Phospholipase C POL    Polymerase PORCN   Porcupine PTA   Phosphotungstic acid PTCH1  Patched homolog 1 PTH   Parathyroid hormone PTHLH  Parathyroid hormone related-like protein PTHRP  Parathyroid hormone related protein PTK7    Protein Tyr kinase PZ    Progress zone qPCR   Quantitative polymerase chain reaction qRT-PCR  Reverse transcriptase PCR Rac   Ras-Related C3 Botulinum Toxin Substrate 1 RCAN   Replication-Competent, ASLV LTR, No splice acceptor RCAS    Replication-Competent ASLV LTR with a Splice acceptor RCASBP   RCAS Bryan Polymerase RCASBPY   RCASBP Gateway RECK   Reversion inducing cysteine rich protein with kazal motifs  Rho   Rhodopsin RIA   Replication incompetent avian retrovirus RIPA   Radioimmunoprecipitation assay ROCK   Rho-associated kinase ROR    Receptor Tyr kinase-like orphan receptor RS    Robinow Syndrome  RSV    Rous sarcoma virus RTK    Receptor tyrosine kinases RUNX   Runt related transcription factor RYK    Receptor Tyr kinase Satb2   Special AT-rich sequence-binding protein 2 SD   Standard deviation SDS   Sodium dodecyl sulfate sFRP   Secreted Fzd related protein SHH    Sonic hedgehog Siah2   Seven in absentia homolog, E3 ubiquitin protein ligase 2 SOST   Sclerostin SOX   Sex determining region Y-box STF    SuperTopFlash SWIM   Secreted Wg-interacting molecule    xvi TBX   T-box transcription factor  TCF    T-cell factor TCOF1  Treacher Collins Syndrome Protein 1 TOP-GAL  T cell factor (TCF)βcatenin, X-gal TUNEL  Terminal deoxynucleotidyl transferase dUTP nick end labeling TWIST1  Twist Basic Helix-Loop-Helix Transcription Factor 1 Tyr   Tyrosine UPR   Unfolded protein response Vangl    Van Gogh-like WLS   Wntless WIF1   WNT inhibitory factor 1 WNT    Wingless wt   Wild-type ZMPSTE24  Zinc Metallopeptidase STE24 ZPA    Zone of Polarizing Activity      xvii Acknowledgements Special mention goes to my supervisor, Dr. Joy Richman. My PhD has been an amazing experience and I thank Joy wholeheartedly, not only for her tremendous academic support, but also for giving me so many wonderful opportunities. I thank her for providing the freedom to pursue my own interests and make my own mistakes, for encouragement and support, and most importantly, fostering a love for science. Thank you also to my supervisory committee members Drs. Michael Underhill, Tim O’Connor and Janel Kopp for your valuable time and insight, and also for the hard questions which incented me to widen my research from various perspectives.  I am extremely grateful for the financial support from the UBC Four Year Fellowship that I received during my time here, as well as the various awards received and opportunities to present my research within the Cell and Developmental Biology program.  I would also like to thank each and every one of the Richman Lab members (including the fantastic work study students!) for their stimulated discussions and encouragement. I met some truly amazing people in the Richman lab, particularly Katherine Fu for her support, advice, encouragement and willingness to listen to any problems I encountered. I always looked forward to her company in the lab. I would also like to thank Adrian Danescu for the help and suggestions throughout my time in the Richman lab. I could not have done this degree without the both of you! Thank you to all my fellow grad students for providing scientific insight, who were of great support in deliberating over our problems and findings, as well as providing happy distraction to rest my mind outside of my research. I am so lucky to have found such incredible friends here. A special gratitude to my partner, Robbie, thank you for listening to all my scientific experiments and continuing to love and support me.  Lastly, I would like to express my deepest gratitude to my family and friends. My parents are the most positive, encouraging, inspirational and supportive people in my life. Their generosity and support is truly amazing and I would have not been able to do this without their help.   Thank you for all your encouragement!      1  1. Introduction 1.1 Overview Rare medical diseases are difficult for patients, families, caretakers, and for researchers studying them. Rare diseases affect approximately 6-8% of the population (Foley, 2015; Hieter and Boycott, 2014; Taruscio et al., 2014; Taruscio et al., 2003; Wangler et al., 2017) with the majority of diseases being genetic that affect the patient’s entire life (Plaiasu et al., 2010). The diseases are characterized by a diverse range of signs and symptoms that can vary in severity from patient to patient suffering from the same disease (Groft and Posada de la Paz, 2017). The field of rare diseases lacks scientific and medical knowledge, thus leaving many patients undiagnosed. Science can help provide some answers about understanding the genesis or mechanisms causing the disease. A single gene causing a rare disease indicates that the genetic variant is sufficient to produce physiological disruption during development. We can observe differences between unaffected genes (wild-type) and those affected (mutant) to hypothesize how the gene and its protein product affect the biochemical pathway(s). This can enhance our knowledge of the normal and disease states of the biochemistry and the pathology.  Several rare diseases affect development of the skeleton. When endochondral or intramembranous bone formation is disrupted, dysplasias can affect the bones in the arms, legs or face (Krakow and Rimoin, 2010). Cartilage is an important structural component that provides the foundation for developing long bones, vertebrae, and bones of the posterior cranial base (Amini et al., 2012; Parada and Chai, 2015; Young et al., 2006). Abnormal development of the cartilage can result in numerous birth defects including achondroplasias, dwarfism, and craniofacial abnormalities (Kornak and Mundlos, 2003; Krakow and Rimoin, 2010; Ornitz and Legeai-Mallet, 2017). Growth factor signaling is disrupted in most of the skeletal dysplasias and several different signaling pathways are involved such as Fibroblast Growth Factors (FGF), Wingless-related (WNT), Parathyroid hormone related peptide (PTHRP) and hedgehog signaling to name a few.  The studies presented in this thesis are aimed at identifying the molecular mechanisms that regulate cartilage formation in autosomal dominant Robinow Syndrome (RS). RS is a rare skeletal dysplasia disorder (1:500,000 live births) and patients present an array of phenotypes affecting the face and limbs (Mazzeu et al., 2007). RS affects non-canonical WNT (β-catenin independent)    2 signaling and a better understanding of this less-studied WNT signaling pathway is imperative as it is relevant to a large number of developmental mechanisms including planar cell polarity and convergent extension.  1.2 Limb development in the chicken embryo The vertebrate limb is an excellent model for studying the fundamental aspects of embryonic development. Limb buds appear as small bulges in the flank of the embryo and consist of homogenous undifferentiated lateral plate mesodermal cells covered by an ectoderm (Davey et al., 2018b; Zuniga, 2015). Development of the limb requires crosstalk of signaling pathways that involve the interaction between different molecules to transfer positional information along the proximo-distal, anterior-posterior and dorso-ventral axes. Additionally, limbs contain similar genes involved in other developmental contexts such as craniofacial development; therefore, the information gained from studying the limb can be applied to other tissues/structures during development (Anderson and Stern, 2016; Lopez-Rios, 2016; Melrose et al., 2016; Ornitz and Marie, 2015; Suzuki and Morishita, 2017; Tao et al., 2017; Tickle, 2015; Tickle and Towers, 2017; Verheyden and Sun, 2017; Zuniga, 2015). Seeing that limbs are not necessary for embryonic survival; the limbs can be experimentally and molecularly manipulated to study cellular and molecular pattern formation. 1.2.1 Limb bud outgrowth and patterning in the limb Limb outgrowth is initiated from the lateral plate mesoderm and epithelial-mesenchymal interactions drive the limb bud outward from the flank of the embryo (Gros and Tabin, 2014; Tanaka, 2013; Verheyden and Sun, 2017). Limb bud outgrowth is mediated by specific signaling centers such as the progress zone (discussed later) that is located in mesenchymal cells that lie just under the apical ectodermal ridge (AER) (Verheyden and Sun, 2017; Wolpert, 2002). The AER is initiated from a signaling center along the distal edge of the limb that provides patterning instructions to the underlying mesenchyme (Tickle, 2015). The AER primarily mediates proximal-distal (PD) outgrowth (shoulder to finger tips) of the limb via FGF signaling (Cohn et al., 1995; Kawakami et al., 2001; Lewandoski et al., 2000). Another signaling center is the zone of polarizing activity (ZPA), which secretes Sonic Hedgehog (SHH) and is located at the posterior margin of the limb. The ZPA determines the anterior-posterior (AP) axis (thumb to little finger) and maintains the AER (Lopez-Rios, 2016; Riddle et al., 1993; Tickle and Towers, 2017). The third    3 axis in the limb is the dorsal-ventral (DV) axis (back of hand to palm) and is controlled by the ectoderm, with the AER developing at the boundary separating the dorsal and ventral ectoderms (Altabef et al., 1997; Tickle, 2015).  The limb is organized into three regions along the PD axis: stylopod, zeugopod, and autopod (Zuniga, 2015). The stylopod comprises the humerus/femur, the zeugopod includes the radius-ulna/fibula-tibia and the autopod involves the most distal part of the limb with the carpal-metacarpal/tarsal-metatarsal and digits (Abzhanov et al., 2007). There are several models of vertebrate limb patterning and most involve outgrowth of the limb in the PD axis. In the PD axis, cells acquire positional values, which they interpret to form appropriate structures (differentiation) (Towers and Tickle, 2009a). Two main scenarios for positional information in the limb bud have been explained by the classic French flag model (Wolpert, 1969). The first scenario is known as the progress zone (PZ) model, which uses timing to determine cell fate, and is located in mesenchymal cells that lie just under the AER (Niswander et al., 1993; Wolpert, 2002). Cells that leave the PZ differentiate in a PD order, and those that leave early on in development form more proximal structures (e.g., stylopod); whereas cells that leave the PZ later in development form more distal structures (e.g., autopod) (Towers and Tickle, 2009b; Wolpert, 1969). The second scenario is known as the early specification model, which proposes that the PD pattern is established very early in development, via morphogen gradient (Dudley et al., 2002). It is now thought that these two models work together and control early limb development (e.g., stylopod and zeugopod) (Tabin and Wolpert, 2007); whereas another model, known as the two-signal model, proposes that two opposing signals (retinoic acid from the paraxial mesoderm and FGF from the AER) pattern the skeletal elements along the PD axis (Benazet and Zeller, 2009). It is tricky to integrate the models that pattern the limb and it is suggested that there is also an intrinsic timer in accompaniment with the other models, which specifies the distal limb structures (Saiz-Lopez et al., 2015; Zuniga, 2015).  1.2.2 Molecular regulation of the embryonic limb  In the chicken embryo, limbs initiate during organogenesis at Hamburger-Hamilton stage (HH) 12 (45-49 hours) (Kawakami et al., 2001), but do not bud out until around stage HH16-17 (51-64 hours) (Hamburger and Hamilton, 1951; Wyngaarden et al., 2010). The limb buds form in discrete positions along the flank of the embryo determined by the level of HOX (Homeobox) gene expression (Burke et al., 1995; Cohn et al., 1997). Limb identity is specified within areas of the    4 flank of the embryo either as fore- or hindlimb fields via the induction of specific transcription factors (TBX4, TBX5, and PITX2) (Tickle, 2015). Specific instances of crosstalk between WNTs and FGFs control limb bud initiation and AER induction. At stage HH12-14, WNT2B and WNT8C are expressed in the lateral plate mesoderm near the fore- and hindlimb regions, respectively, and activate FGF10. Mesenchymal progenitors of the nascent limb bud arise at stage HH15 by epithelial-to-mesenchymal transition that is regulated by TBX5 and FGF10 (Gros and Tabin, 2014). The limb bud begins to form at stage HH16 with regulation of WNT3A to induce FGF8 signaling in the overlying limb ectoderm. A regulatory loop between FGF10-WNT3A-FGF8 is now formed (Kawakami et al., 2001). The PD axis is maintained by SHH which is induced by FGF8 expression in the AER. This is maintained by the SHH/AER-FGF module (Sheeba et al., 2016).  The AP axis is formed by mutual antagonism between posteriorly positioned HAND2 and anteriorly located GLI3. This antagonistic interaction posteriorly restricts SHH in the limb mesenchyme, which defines the ZPA signaling centre (Riddle et al., 1993; Zuniga, 2015). At around stage HH17, SHH forms a morphogen gradient involved in AP patterning of the limb to control proliferative expansion of mesenchymal progenitors (Tickle, 2015; Tickle and Towers, 2017; Towers et al., 2008). Increased SHH signaling inhibits GLI3 repressor (GLI3R), creating an accumulation of GLI3 activator (GLI3A) (Tickle and Towers, 2017; Zuniga, 2015). The anterior limb bud contains increased levels of GLI3R and low levels of GLI3A (Zeller et al., 2009). These opposing signals help establish the AP axis of the limb (Sheeba et al., 2016).  Establishment of the DV axis occurs through epithelial-mesenchymal interactions at around stage HH17, which are directed by signals covering the ectoderm of both dorsal and ventral sides of the limb. WNT7A is expressed in the dorsal ectoderm which induces LMX1B in the underlying dorsal mesoderm (Altabef and Tickle, 2002; Tickle, 2015). BMP signaling in the ventral ectoderm induces EN1 expression that also helps establish the AER (Ahn et al., 2001). The AER creates a boundary between the dorsal and ventral ectoderm. Termination of limb bud outgrowth requires the SHH/GREM1/FGF feedback loop. SHH upregulates GREM1 which relays the signal to FGFs in the AER. AER-FGF transmits the signal back to SHH in the ZPA, establishing an epithelial-mesenchymal feedback loop. BMP4, located in the mesenchyme, also upregulates GREM1 in a negative feedback loop. The higher levels of GREM protein block BMP signaling and thus maintain FGF in the AER. As the limb grows the    5 GREM domain is separated more and more from the SHH domain. SHH is required to maintain GREM expression. When SHH signals are decreased, GREM downregulates and this allows BMP4 to increase in activity. BMP4 then inhibits FGF4 and FGF8 in the epithelium, leading to flattening of the AER and a cessation of outgrowth of the undifferentiated mesenchyme) (Zuniga, 2015).  1.2.3 Chondrogenesis and skeletogenesis in the limb  Endochondral ossification occurs in the appendicular skeleton where a cartilage template is replaced by bone. Initiation of pre-cartilaginous condensations results from aggregation of mesenchymal progenitor cells from the lateral plate mesoderm, and the failure of cells to disperse, probably by increased adhesiveness (Hall and Miyake, 1995, 2000). N-CAM, tenascin and fibronectin mediate calcium dependent and independent cell-cell adhesion during condensation formation (Chimal-Monroy and Diaz de Leon, 1999; Tavella et al., 1994). This cellular compaction is an early morphogenetic process that organizes chondrocytes into clusters to form the cartilage template (Barna and Niswander, 2007). The first evidence of histological changes in mesenchyme condensations in the chicken embryo occur at stage HH24 (4 days) (Gould et al., 1972). Once the cartilage template has formed, the mesenchyme differentiates into immature chondrocytes where transcription factors modulate cell proliferation (Long and Ornitz, 2013). SOX9 is a transcription factor expressed in cartilage progenitor cells and precedes expression of extracellular matrix (ECM) proteins such as collagen type II (COL2A1) and aggrecan (ACAN) (Akiyama et al., 2005; Kozhemyakina et al., 2015). The perichondrium is also forming, which is a fibroblastic layer of flattened cells that surrounds the cartilage. Markers of the perichondrium include FZD1, CTNNB1, BMP4, LEF1, PTCH2, PTH/PTHrP-receptor, WNT5A (Hartmann and Tabin, 2000), and RUNX2 (Ducy et al., 1997; Hinoi et al., 2006). Afterward, chondrocytes mature into zones of prehypertrophic and hypertrophic chondrocytes with different profiles. The prehypertrophic chondrocytes proliferate and express parathyroid hormone like hormone (PTHLH or PTHRP) and IHH (Hartmann and Tabin, 2000; Vortkamp et al., 1996). Cells located within the centre cartilaginous core undergo maturation and increase in volume and produce hypertrophic chondrocytes that express COL10A1 (Hartmann and Tabin, 2000) and MMP13 (Bond et al., 2016; Cooper et al., 2013; Hartmann and Tabin, 2000). Hypertrophic chondrocytes begin secreting ECM components at the center of the cartilage (Behonick and Werb, 2003). As COL10A1 expression increases, SOX5, SOX6, SOX9, COLII and ACAN begin to decrease and form late hypertrophic chondrocytes (Kozhemyakina et al., 2015). Recent data has shown that hypertrophic chondrocytes    6 can become osteoblasts in a cartilage-to-bone transition, thereby transdifferentiating directly into osteoblasts (Park et al., 2015; Tsang et al., 2015; Yang et al., 2014; Zhou et al., 2014). This will form the primary ossification center and will gradually replace the remaining cartilage within the diaphysis of the long bone and enable formation of a bone collar (Mackie et al., 2008). The production of new chondrocytes is next restricted to the ends of the long bones at the epiphysis in the growth plate creating a secondary ossification centre. The growth plate comprises of three zones: (1) resting, round chondrocytes that give rise to (2) flattened proliferating chondrocytes that stack and arrange in columns, (3) which will differentiate into hypertrophic chondrocytes (Kozhemyakina et al., 2015). Resting chondrocytes divide at arbitrary planes, whereas more mature chondrocytes, such as the flattened proliferative chondrocytes involve oriented cell division to generate daughter cells that are displaced laterally and pivot around each other to form a single column (Li and Dudley, 2009; Li et al., 2017). As the center cartilage region in the diaphysis begins to hypertrophy the remaining chondrocytes in the epiphysis will continue to proliferate, allowing further growth of the long bone. As new bone material is added the internal region of the bone gets hollowed out via resorption of osteoclasts from the peripheral blood, forming the bone marrow cavity (Clarke, 2008). It takes 7 days after a wing bud appears for the complete cartilage skeleton to be laid down and about 14 days for the diaphysis to ossify (Hamburger and Hamilton, 1951). Full ossification of the chicken skeleton occurs post-hatching. 1.2.4 Facial development in the chicken embryo and molecular regulation of the mandible Development of the skull is a complex and precisely timed morphogenetic event. The skeletal tissues derive from paraxial mesoderm and neural crest derived mesenchyme (Chai et al., 2000; Gross and Hanken, 2008; Jiang et al., 2002; Le Douarin, 2012; McBratney-Owen et al., 2008; Yoshida et al., 2008). Initially, in the chicken embryo facial development begins with the migration of cranial neural crest cells, which are a unique group of cells derived from the developing neural folds that pinch off and transform into mesenchymal cells (Dupin and Le Douarin, 2014; Le Douarin et al., 2004). Cranial neural crest cells give rise to a diverse cell lineage such as melanocytes, smooth muscle, some neurons, glia, and craniofacial cartilage and bone (Dupin and Le Douarin, 2014). Cranial neural crest migration occurs between stages HH7-10 in the chicken embryo (Kontges and Lumsden, 1996). Initially, neural crest cells migrate from the fore- mid- and    7 anterior part of the hindbrain into the pharyngeal arches (Kontges and Lumsden, 1996). The pharyngeal arch is patterned by HOX genes located along the AP axis of the embryo, with the more anterior regions being HOX-negative (Creuzet et al., 2005). HOX-negative neural crest cells are mainly patterned by the distal-less genes (Beverdam et al., 2002; Depew et al., 2002; Depew et al., 2005).  Once neural crest cells have entered the presumptive face, the facial prominences begin to form. The prominences are swellings of neural crest-derived mesenchyme that flank the primitive mouth or stomodeum. The upper jaw is formed by the medial nasal, lateral nasal and maxillary prominences whereas the lower jaw is formed by the mandibular prominences (Jiang et al., 2006). In the chicken embryo the frontonasal mass is equivalent to the medial nasal prominences (Abramyan et al., 2015). Work in the mouse embryo has shown that the mandibular prominence is patterned along the proximo-distal and oral-aboral axes by the expression of several transcription factors and signaling molecules. These include tightly controlled regulation of BMP and FGF expression (Chai and Maxson, 2006), and Satb2 which is important for PD growth (Fish, 2016; Fish et al., 2011). Unlike the limb, many craniofacial defects can be traced back to neural crest cell deficiencies. For example, failure to migrate or proliferate in sufficient numbers can lead to first arch abnormalities such as Treacher-Collins or Goldenhar syndrome (Dixon et al., 2007; Manocha et al., 2018; Passos-Bueno et al., 2009; Sakai et al., 2016). 1.2.5 Chondrogenesis and skeletogenesis in the face The chondrocranium is part of the cranial base and is a cartilaginous structure that grows to envelope the growing brain. In primitive vertebrate animals where only a cartilaginous skeleton exists, there is a cranial base, nasal, orbital and otic capsules as well as an upper (palatoquadrate) and lower jaw cartilage (Meckel’s cartilage). All vertebrates begin development of the skull with the formation of the chondrocranium. The cartilaginous skull is the first skeletal tissue to develop from cranial neural crest cells and cervical somites (Couly et al., 1993; Sperber and Sperber, 2018).  The craniofacial complex is comprised of the dermatocranium and viscerocranium which are formed by intramembranous ossification (Sperber and Sperber, 2018). Specifically, the calvaria and facial bones are formed by intramembranous ossification where osteoblasts arise directly from the mesenchymal condensations (Ornitz and Marie, 2015). This process is first initiated by RUNX2 followed by its downstream target Osterix (OSX) (Nakashima et al., 2002; Takarada et al., 2016). Transcriptional regulators (MSX2 and TWIST1) maintain both proliferation and repress    8 commitment towards an osteogenic fate by repressing the RUNX2 gene (Komori, 2006). In the mandible, Meckel’s cartilage is surrounded by ossification centres for the ramus and body of the mandible. Most of Meckel’s cartilage is resorbed but, in the midline, the fate of chondrocytes is to become osteoblasts in mammals (Ishizeki et al., 1999). In birds, Meckel’s cartilage persists throughout life (Havens et al., 2008).  1.3 Genetic Skeletal Dysplasias  Skeletal dysplasias can be congenital where no specific genetic cause is identified or may be heritable where discrete mutations within one gene or other DNA alterations have been diagnosed. There are over 450 recognized genetic conditions that affect primarily bone and cartilage (Krakow and Rimoin, 2010; Offiah, 2015; Warman et al., 2011). The skeletal dysplasias commonly affect morphology of multiple bones, leading to abnormal shape and size of the appendicular, axial and craniofacial skeleton (Manocha et al., 2018; Tao et al., 2017). These forms of skeletal dysplasias are separate from hormonal disorders such as lack of growth hormone (Divall, 2016; Divall and Radovick, 2013).  Diagnosis is becoming easier especially since the cost of sequencing exomes has decreased in recent years. If the gene variant occurs in a non-coding region, then it takes more effort to find the genetic change that causes the disease. Once a patient with a phenotype is identified, sequencing of multiple family members is necessary to confirm the cause (Offiah, 2015).  Types of genetic dwarfism that directly affect the skeleton include hypochondroplasia which affects bone growth (mutations in FGFR3) (Foldynova-Trantirkova et al., 2012), diastrophic dysplasia caused by mutations in the SLC26A2 gene and leads to short-limbed dwarfism plus deformities of the feet and thumb (Haila et al., 2001), spondyloepiphyseal dysplasias characterized by a shortened trunk with mutations in COL2A1 (Lee et al., 1989) and achondroplasia, which is the most common form of dwarfism (Horton et al., 2007). Achondroplasia is similar to hypochondroplasia, but features are more severe. Achondroplasia results in short arms/legs often featuring midfacial hypoplasia and is caused by mutations in FGFR3 (Horton et al., 2007; Rousseau et al., 1994; Shiang et al., 1994). Thus one of the predominant signaling pathways involved in skeletal dysplasias is the Fibroblast Growth Factor pathway. Depending on the location of bone shortening in the limbs (e.g., stylopod, zeugopod etc.) dwarfism can be classified as:    9 rhizomelia (shoulder/hip), mesomelia (forearms/lower legs), acromelia (hands/feet), or micromelia (abnormally small and short limbs) (Panda et al., 2014).  Many forms of skeletal dysplasia also affect the face. The mandible can produce a variety of developmental malformations such as micrognathia or undersized jaw (Paladini, 2010). Micrognathia occurs in a variety of syndromes such as: Pierre Robin sequence, generally causing mandibular hypoplasia that might be a result of mutations in SOX9 and KCNJ2 genes (Gangopadhyay et al., 2012), Goldenhar syndrome, which usually affects one side of the face with an unknown etiopathogenesis (Bogusiak et al., 2017), mandibuloacral dysplasia caused by mutations in LMNA or ZMPSTE24 (Cenni et al., 2018), or Treacher-Collins that affects many bones and tissues of the face with mutations most commonly in TCOF1 (Trainor et al., 2009). In this thesis I am studying a skeletal disorder with limb and craniofacial defects called Robinow Syndrome (RS), with both mesoderm and neural crest-derived skeleton affected. However, unlike some skeletal dysplasias, RS affects the Wingless-related or WNT signaling pathway.  1.3.1 Robinow Syndrome RS is a rare skeletal dysplasia disorder (1:500,000 live births) and patients present an array of phenotypes affecting the face and limbs (Mazzeu et al., 2007). The syndrome was originally described in 1969 in a family that had short stature (Robinow et al., 1969). There are three forms of inheritance of RS, autosomal recessive, autosomal dominant and X-linked. All forms of RS are characterized by skeletal dysplasias affecting primarily the face and limbs (Afzal et al., 2000; Mazzeu et al., 2007; Person et al., 2010; Roifman et al., 2015; White et al., 2015; White et al., 2018; White et al., 2016b). Patients typically present with short stature and mesomelic limb shortening as well as hypertelorism, wide nasal bridge and midface hypoplasia (Person et al., 2010; White et al., 2018). All cases of RS are caused by mutations in a specific branch of the WNT signaling pathway, non-canonical WNT signaling that occurs independently of β-catenin. I used RS as an entry point for my research on the role of non-canonical WNT signaling in skeletogenesis.  The recessive forms of RS are either caused by loss-of-function mutations in the ROR2 receptor (OMIM#26310) (Afzal et al., 2000; Aglan et al., 2015; Tamhankar et al., 2014; van Bokhoven et al., 2000), or NXN (Nucleoredoxin), a negative regulator of WNT signaling (White et al., 2018). The dominant forms of RS (RS), are caused by missense or non-frameshift mutations in the WNT5A ligand (OMIM#180700) (Person et al., 2010; Roifman et al., 1993), frameshift mutations in the Dishevelled mediator proteins, DVL1 or DVL3 (DVL1, OMIM#616331; DVL3,    10 OMIM#616894) (Bunn et al., 2015; White et al., 2015; White et al., 2018; White et al., 2016b), missense and truncating variants in the FZD2 receptor, missense mutations in RAC3, and hemizygous missense mutations in glypican-4 (GPC4) (weak evidence for X-linked RS) (White et al., 2018). Dominant RS can be inherited or may occur due to de novo mutations (White et al., 2018).  1.4 Many branches of WNT signaling Wingless (WNT) signaling is a conserved pathway in metazoan animals comprising a multifaceted signal transduction pathway that is essential for development (Holstein, 2012; Wiese et al., 2018). The history of Wnt signaling can be traced back approximately 40 years ago from studies in both mice and fruit flies (Nusse and Varmus, 1982; Nusslein-Volhard and Wieschaus, 1980; Sharma, 1973; Sharma and Chopra, 1976). The genesis of the field of Wnt signaling in vertebrates began with the discovery of the first mammalian Wnt gene in 1982 by studies involving both mouse cancer models along with oncogenic retroviruses. In 1982, Nusse and Varmus conducted proviral tagging to discover proto-oncogenes in mice via retroviruses. They located a putative proto-oncogene that was transcriptionally activated as a result of the MMTV-induced breast cancer (mouse mammary tumour virus) proviral DNA insertion in tumours. The located gene was named int1 to signify the first common integration site (Nusse and Varmus, 1982). Previously, during the 1970s, genetic screens in Drosophila melanogaster presented segment polarity genes required for patterning during embryogenesis. One of the genes was termed Wingless, with mutant flies resulting in the loss of wings (Sharma and Chopra, 1976). In the late 1980s int1 and Wingless genes were cloned, and it was found that both genes had restriction maps matching the same regions, indicating that the int1 homologue in Drosophila was Wingless (Baker, 1987; Rijsewijk et al., 1987). This break-through provided opportunities to uncover mechanisms of int1 function and generated an initial outline of canonical int1/Wingless signaling with combined data from mouse, Drosophila, and Xenopus (Orsulic and Peifer, 1996). To settle any confusion with nomenclature, genes belonging to int1/Wingless family were renamed to ‘Wnt1’, denoting the Wingless-related integration site (Nusse et al., 1991).  1.4.1 WNT ligands WNTs are secreted glycoproteins involved in short-range cell-cell communication important for patterning, morphogenesis, proliferation and cell survival. WNT proteins are approximately 40    11 kDa in size and contain 22-24 conserved cysteine residues that are involved in forming disulfide bridges to maintain a globular secondary structure (Janda et al., 2012). There are 19 WNTs (Table 1.1) and each of these genes appear to have a specific role in development or regulation on the control of gene expression and cell fate changes including: cell migration, cell polarity, and organogenesis (Yang and Mlodzik, 2015). Roles for WNTs have been defined in many organs and at several stages of development. For example WNTs are involved in early body axis formation (Hikasa and Sokol, 2013; Yamaguchi et al., 1999), somite patterning (Geetha-Loganathan et al., 2008), cardiac development (Brade et al., 2006), craniofacial development (Brugmann et al., 2007; Geetha-Loganathan et al., 2014; Hosseini-Farahabadi et al., 2017), limb bud initiation (Yang, 2003), and DV limb patterning (Altabef et al., 1997). At later stages of development WNT signaling in the limb appear to be critical for outgrowth, skeletogenesis, and muscle differentiation (Lerner and Ohlsson, 2015).  Moreover, of the 19 WNT ligands, at least ten are expressed at various stages throughout limb development in the chicken embryo (WNT2B, WNT3A, WNT4, WNT5A, WNT5B, WNT6, WNT7A, WNT9A [formerly WNT14], WNT10A, and WNT11) (Baranski et al., 2000; Geetha-Loganathan et al., 2005; Hartmann and Tabin, 2000; Kawakami et al., 2001; Kawakami et al., 1999; Narita et al., 2005; Summerhurst et al., 2008; Witte et al., 2009). There are also several WNTs expressed in the face including WNT2B, WNT5A, WNT5B, WNT11 and WNT16 (Geetha-Loganathan et al., 2009). 1.4.2 WNT synthesis and secretion WNT ligands require a set of post-translational modifications for secretion and receptor binding (Clevers et al., 2014; Langton et al., 2016; Willert and Nusse, 2012). WNT proteins are hydrophobic, generally found associated with the cell membrane or ECM, and they are lipid modified by the attachment of a palmitoleic acid to a conserved cysteine residue (Langton et al., 2016; Willert et al., 2003). Lipid modification is essential for WNT function to enable binding to FZD receptors (Cong et al., 2004; Kurayoshi et al., 2007; Willert et al., 2003). Prior to secretion, WNT proteins associate with several molecules that accompany WNT before release to the extracellular space. WNTs first enter the endoplasmic reticulum (ER) and receive a lipid modification (palmitoylation chain) from transmembrane protein, Porcupine (PORCN) (Kadowaki et al., 1996). Additionally, WNTs are glycosylated; however, the number of glycosylation sites varies depending on the WNT ligand (Smolich et al., 1993). It has been shown that glycosylation is required for WNT secretion, as unglycosylated WNT5A in culture is not detected in the ECM    12 or conditioned media (Kurayoshi et al., 2007). After binding to PORCN, WNTs leave the ER to enter the Golgi and bind to transmembrane protein Wntless (WLS) (Bartscherer et al., 2006; Langton et al., 2016). WLS will accompany WNT to the cell membrane for secretion. Once WNTs are secreted from the cell, WLS is recycled by endosomes and a retromer complex back to Golgi to start the secretion process all over again (Langton et al., 2016; Yang et al., 2008).  After secretion to the extracellular space, WNTs encounter several protein-binding partners to protect the fatty acid chain from the aqueous environment. The lipid modification renders the ligand hydrophobic; thus, WNT is generally chaperoned by heparan sulphate proteoglycans (HSPG) (Fuerer et al., 2010). HSPG can regulate WNT signaling distribution of the ligand throughout the tissue. WNTs show restricted diffusion along the surface of receiving cells, where morphogens are transferred from one HSPG to the next, moving from high concentration to low concentration. HSPGs can enable a signaling platform for morphogen gradient formation as HSPGs are able to bind to several cell surface co-receptors (Yan and Lin, 2009). Another WNT-binding protein includes the secreted wingless-interacting molecule (SWIM) that facilitates WNT distribution to promote long range signaling (Mulligan et al., 2012). In addition to WNT binding partners, WNTs can be transported extracellularly via lipid containing membranous transporters (Greco et al., 2001) or exosomes, which are derived from the budding of multivesicular bodies to secrete and carry WNT proteins (Gross et al., 2012). Adding to the complex interactions with the various WNT transporters, WNTs can encounter several antagonistic binding partners that hinder their activity (Table 1.1). 1.4.3 WNT receptors There are an array of receptors that Wnt ligands can bind that comprise up to 15 different receptors and co-receptors (Table 1.1; (Niehrs, 2012; Stricker et al., 2017). This includes the transmembrane protein Frizzled (Fzd), which was the first WNT receptor identified. Fzd was discovered in 1944 in a mutant fly that contained disorganized bristles and ommatidia (Lindsley et al., 1967). Fzd was later identified in Drosophila as a planar polarity gene to orient a field of cells in a tissue (Bhanot et al., 1996; Gubb and Garcia-Bellido, 1982). It was then found that Wingless could bind to Drosophila Fzd2, thereby increasing β-catenin levels (Bhanot et al., 1996). FZD is a non-traditional G-protein coupled receptor that is comprised of a conserved extracellular N-terminus cysteine-rich binding domain (CRD) and a seven-pass transmembrane segment (Janda et al., 2012; Nichols et al., 2013). The intracellular C-terminus contains a PDZ (Postsynaptic    13 density 95, Discs Large, Zonula occludens-1)-binding domain (Schulte and Bryja, 2007), and a highly conserved KTxxxW motif, which is crucial for recruitment of DVL (Wu et al., 2008). There are 10 FZD family members (from FZD1 through 10), and genetic redundancy has been seen within subfamilies of the receptors (Wang et al., 2016).  There are several other receptors and co-receptors involved in Wnt signaling in addition to FZD. In canonical WNT signaling, FZD receptor works together with co-receptors low-density lipoprotein receptor-related protein (LRP5/6), forming a dimeric structure (MacDonald and He, 2012; Pinson et al., 2000; Tamai et al., 2000). LRP cannot work alone with WNT; thus, upon WNT ligand signaling, a ternary complex is formed between FZD-LRP-WNT (Bourhis et al., 2010).  Other WNT receptors include the receptor tyrosine kinases (RTK), such as receptor Tyr kinase-like orphan receptors (ROR1 and ROR2), receptor Tyr kinase (RYK), protein Tyr kinase 7 (PTK7) and muscle skeletal receptor Tyr kinase (MUSK) (Stricker et al., 2017). ROR1/ROR2 were originally considered “orphan” receptors because their ligands were unknown, but we have learned that they contain WNT extracellular binding domains (Mikels and Nusse, 2006; Oishi et al., 2003). ROR1/ROR2 RTKs are type 1, single-pass transmembrane receptors that contain an extracellular CRD, presenting close homology with FZD CRDs (Masiakowski and Carroll, 1992). Multiple WNT ligands are able to bind to ROR1/ROR2 receptors (Brinkmann et al., 2016; Green et al., 2007; Hikasa et al., 2002; Liu et al., 2008a; Oishi et al., 2003; Paganoni et al., 2010; Winkel et al., 2008), and it is well known that WNT5A acts as the primary ligand for the ROR2 receptor (Grumolato et al., 2010; Mikels et al., 2009). WNT5A can cause the dimerization of ROR2 and FZD receptors, or induce homodimerization of ROR2, independent of FZD (Feike et al., 2010; Liu et al., 2007; Liu et al., 2008b). The function of ROR1 in WNT signaling has not been investigated as thoroughly; although it has been shown that WNT5A can also bind to ROR1 (Fukuda et al., 2008). RYK is a type 1, single pass transmembrane protein (Halford and Stacker, 2001) that likely acts as a co-receptor, as it has been shown to interact with FZD8 (Wong et al., 2003). Other RTKs that bind WNTs include MUSK and PTK7, which are single-pass transmembrane proteins. MUSK acts as a WNT receptor in muscle, initiating the formation of neuromuscular junctions, and PTK7 binds FZD in the presence of WNT and is involved in PCP signaling (Niehrs, 2012). Two receptors reported are involved in establishing the blood brain barrier. These receptors comprise the G-protein coupled receptor, GPR124 and the glycosylphosphatidylinositol-anchored glycoprotein, RECK (Eubelen et al., 2018). The receptors form a complex (GPR124-RECK) and    14 are specific to bind only WNT7 ligand. It was found that WNT7 has a linker domain from binding to the CRD on FZD and the binding site on RECK, with the simultaneous intracellular binding of DVL targeting both FZD and GPR124 (Eubelen et al., 2018). Other receptors associated with WNT signaling are listed in Table 1.1. Deciphering which WNT will bind to each FZD remains enigmatic. On top of that, WNT receptors and co-receptors continue to be discovered. WNT ligands have the ability to bind to multiple FZDs/co-receptors; and conversely, an FZD receptor/co-receptor has the ability to bind multiple WNTs, adding to the complexity of this signaling pathway (Niehrs, 2012).  1.4.4 WNT signaling intermediates (Dishevelled) Once signaling has been initiated by WNT ligand binding to a receptor complex, the key downstream mediator, Dishevelled (Dvl or Dsh) gets recruited to the FZD receptor. DVL is an intracellular adaptor/scaffold protein that was first identified based on randomized hair on the body and wings in Drosophila (dsh) (Fahmy and Fahmy, 1959). DVL is located intracellularly, associated with actin fibers, and/or forming protein assemblies such as puncta (often seen when there are high levels of DVL) (Axelrod, 2001; Schwarz-Romond et al., 2005). Three DVL homologs have been identified, DVL1, DVL2 and DVL3 in humans and mice (Klingensmith et al., 1996; Lijam and Sussman, 1995; Sussman et al., 1994; Yang et al., 1996) and Xenopus (Gray et al., 2009), two in C. elegans, two in chickens (DVL1, DVL3) (Gray et al., 2009), one paralog in flies (dsh) (Fahmy and Fahmy, 1959), and >four in zebrafish (Mlodzik, 2016).  Once DVL proteins are activated and recruited to FZD, DVL can signal to either the canonical or non-canonical WNT pathways. This “switch-like” mechanism acts through distinct regions of the DVL protein domains (Rothbacher et al., 2000; Wallingford and Habas, 2005). DVL protein contains three conserved domains: an amino terminal DIX (DVL/Axin) domain, a central PDZ domain, and a carboxyl-terminal DEP (DVL, EGL-10, Pleckstrin) domain (Wallingford and Habas, 2005; Wynshaw-Boris, 2012). DVL also contains a cluster of positively charged (basic) residues located between the DIX and PDZ domains, and a proline-rich region located between the PDZ and DEP domains (Penton et al., 2002). The three domains in DVL provide docking sites for many interacting proteins (Wang and Malbon, 2012). The PDZ and DIX domains are important for canonical WNT signaling (Clevers and Nusse, 2012), and for the most part, the DEP domain is utilized to activate the non-canonical WNT pathway (Wallingford and Habas, 2005). In canonical signaling, DVL forms a complex with Axin via its own DIX domain (Clevers and Nusse,    15 2012). DVL and Axin each contain a DIX domain, (named DAX in Axin to distinguish it from DVL DIX), and DIX and DAX domains form a heteropolymer complex when WNT ligand is present (Gammons et al., 2016). This complex causes Axin to become inactivated, thus increasing canonical WNT signaling (discussed in more detail in section 1.4.5.1). Moreover, DVL also promotes signalosome formation via DEP dimerization, which is important for WNT signal transduction to the nucleus in canonical signaling (Gammons et al., 2016). A conformational switch of the DEP domain from a monomer to a swapped dimer prompts DIX-dependent polymerization and signaling to β-catenin. First, monomeric binding of DEP to FZD is followed by domain swapping of DEP. This facilitates polymerization of the DIX domains and stimulates relocation of DVL into clathrin-coated pits (Gammons et al., 2016).   Table 1.1. WNT ligands, receptors and components of WNT signaling. Components and Function Family members WNT ligands WNT1 (Wingless), WNT2, WNT2B, WNT3, WNT3A, WNT4, WNT5A, WNT5B, WNT6, WNT7A, WNT7B, WNT8A, WNT8B, WNT9A, WNT9B, WNT10A, WNT10B, WNT11, WNT16 Alternative ligands R-spondin, Norrin Agonists/Antagonists sFRP, WIF1, DKK, SOST Factors required for secretion PORCN, WI FZD receptors FZD1, FZD2, FZD3, FZD4, FZD5, FZD6, FZD7, FZD8, FZD9, FZD10 LRP receptors LRP5, LRP6 RTKs receptors ROR1/ROR2, RYK, PTK7, MUSK Alternative receptors GPR124, RECK, NRH1, Syndecan, Glypican Signaling intermediates DVL1, DVL2, DVL3 β-catenin destruction complex Axin, APC, GSK3β, CK1  Cellular trafficking and distribution HSPG, SWIM Transcription factors TCF1, TCF3, TCF4, LEF1, ATF2, AP1, NFAT Intracellular modulators Groucho, Siah2, NXN  1.4.5 WNT signaling After WNTs are secreted they bind to the CRD of the FZD receptor. FZD is the principle receptor WNT ligands signal through, and FZD is involved in both canonical and non-canonical WNT signaling pathways. WNT-FZD interaction has been revealed by the co-crystal structure of Xenopus Wnt8 (XWnt8) complex with mouse Fzd8 (Janda et al., 2012). XWnt8 forms a “hand-like” structure, with a ‘palm’ that extends a ‘thumb’—aka the fatty acid palmitoylation site—and    16 an ‘index finger’ to grasp Fzd8 on both sides of its CRD. The ‘thumb’ comprises of a palmitoleic acid that fits into the hydrophobic groove of the CRD on Fzd8 (Janda et al., 2012). After WNT ligand binds to the receptor DVL gets recruited to the cytoplasmic region of FZD. DVL is the first intracellular protein that is a critical component of canonical and non-canonical WNT signaling pathways. WNT ligands trigger two categories of intracellular signaling, the β-catenin-dependent pathway (canonical) or the β-catenin-independent pathway (non-canonical) (Niehrs, 2012). 1.4.5.1 Canonical WNT signaling Canonical signaling which is obligated to use β-catenin as a signal transducer, was the first WNT pathway to be characterized. The canonical WNT pathway plays roles in determining cell fate during gastrulation (Harland and Gerhart, 1997), regulating posterior patterning (Yamaguchi, 2001), controlling the formation of several organ systems (Clevers, 2006; Logan and Nusse, 2004; Yamaguchi, 2001), neural patterning (Kawano and Kypta, 2003), and regulating cell proliferation (Teo and Kahn, 2010). In the absence of a WNT ligand, cytoplasmic β-catenin is bound to the destruction complex: Axin, glycogen synthase kinase 3β (GSK3β), and adenomatous polyposis coli (APC). β-catenin gets phosphorylated by GSK3β and casein kinase 1α (Ck1α) then becomes ubiquitylated and destroyed via proteasomal degradation (Figure 1.1 A) (Clevers and Nusse, 2012; Wiese et al., 2018). Therefore, in the absence of nuclear β-catenin, the nuclear repressor (Groucho) is bound to T-cell factor/lymphoid enhancer factor (TCF/LEF), preventing transcription (Figure 1.1 A) (Daniels and Weis, 2005).  When a WNT ligand is present, the ligand will bind to the CRD of FZD and co-receptor, LRP5/6 (Figure 1.1 B). WNT binding to FZD will sequester components of the β-catenin destruction complex. The cytoplasmic tail of LRP5/6 becomes phosphorylated, causing Axin (along with GSK3β) to be relocated from the β-catenin destruction complex, to the intracellular region of LRP5/6. This results in DVL protein stimulation and translocation to the cytoplasmic tail of the Frizzled receptor. Therefore, the number of ‘β-catenin destruction complexes’ in the cytoplasm decrease leaving greater amounts of β-catenin in the cytoplasm (Figure 1.1 B). Stabilized β-catenin can now enter the nucleus and act as a ‘transcriptional activator’ to displace Groucho and form complexes with TCF/LEF transcription factors (Daniels and Weis, 2005; Grumolato et al., 2010; Komiya and Habas, 2008; Niehrs, 2012). These steps comprise the immediate response of WNT-canonical signaling. The delayed response involves sequestering β-catenin in multivesicular    17 bodies to protect β-catenin from phosphorylation and proteasomal degradation (not shown) (Taelman et al., 2010).    Figure 1.1. Canonical WNT signaling pathways. (A) Canonical WNT signaling in the absence of WNT ligand.  No ligand is present, the receptors are not dimerized and dishevelled is not recruited to the FZD receptor.  The β-catenin destruction complex (AXIN, APC, and GSK3β) is now activated.  β-catenin gets phosphorylated and cannot enter the nucleus.  Groucho is a transcription repressor and blocks transcription of target genes. (B) Canonical WNT signaling pathway in the presence of WNT ligand.  WNT binds to the CRD of the FZD receptor and co-receptor LRP5/6.  DVL relocates to FZD and AXIN moves to the cytoplasmic tail of co-receptor LRP.  β-catenin accumulates in the cytoplasm and translocates to the nucleus,  thereby replacing Groucho and activating the target genes.  1.4.5.2 Non-canonical WNT signaling The non-canonical WNT pathway is also referred to as the β-catenin-independent pathway and uses other downstream signaling to activate intracellular kinases. This pathway is further split into two branches, the planar cell polarity (PCP) pathway and the less studied WNT-calcium pathway.    18 The PCP pathway refers to the alignment of cell polarization across a tissue plane. It first emerged from studies in Drosophila whereby mutations in Wnt components such as Fzd or Dvl disoriented epithelial structures including bristles or cuticle hairs (Fahmy and Fahmy, 1959; Lindsley et al., 1967; Seifert and Mlodzik, 2007). It was later found that PCP is also involved in orientation of structures such as stereocilia in the ear (Ezan and Montcouquiol, 2013; Wang et al., 2006), hair follicle organization (Guo et al., 2004), migratory cells in gastrulation (Gubb and Garcia-Bellido, 1982; Zallen and Wieschaus, 2004), and organization of chondrocytes (Gao et al., 2011). The endpoint of this pathway leads to the rearrangement of actin organization. PCP provides positional information to orient cells in a directed fashion. Like the canonical pathway, PCP signaling involves WNT ligand typically binding to an FZD receptor along with a co-receptor. Although, in Drosophila it is unclear if a Wnt molecule regulates the PCP pathway, as this has only been found in vertebrate models (Maung and Jenny, 2011; Strutt and Strutt, 2005; Tree et al., 2002).  Upon ligand binding to the receptors, DVL is recruited to the intracellular part of FZD for both PCP and Calcium signaling (Figure 1.2). NXN is a negative regulator of PCP signaling by blocking ubiquitination and degradation of DVL (Funato et al., 2008). GPC4 is a heparin sulphate proteoglycan and is a positive regulator of PCP signaling via promoting the accumulation of DVL at the plasma membrane (Ohkawara et al., 2003).  In the PCP pathway, the PDZ domain of DVL can activate the small GTPases Rho and Rac (i.e., RAC3) (Wallingford and Habas, 2005) (Figure 1.2 A). Rho activation creates a DVL-DAAM1 (Dishevelled associated activator of morphogenesis 1) complex, which causes activation of Rho-associated kinase (ROCK) (Marlow et al., 2002). Activation of ROCK can modify the actin cytoskeleton. The other component of PCP pathway includes the DEP domain of DVL, which can activate Rac GTPases that will stimulate the Jun N-terminal kinase (JNK) pathway (Habas et al., 2003). JNK will then translocate to the nucleus and activate transcription factors such as activating transcription factor 2 (ATF2) (Weston and Davis, 2007). There are several key components in the PCP pathway that play a role in convergent extension and mediating polarity that include the participation of membrane proteins such as Celsr, Van Gogh-like proteins (Vangl), as well as intracellular proteins including Prickle and Diversin (Yang and Mlodzik, 2015). To initiate cell polarity, Prickle and Vangl will form a complex to antagonize FZD-DVL complex on the opposite side of the cell. The two antagonizing complexes will stabilize each other and establish planar polarization (Yang and Mlodzik, 2015).     19 The WNT/calcium (Ca2+) pathway is another branch of non-canonical WNT signaling (Figure 1.2 B). This signaling pathway became evident when researchers found that certain WNTs injected into zebrafish embryos could double the levels of Ca2+ (Slusarski et al., 1997; Westfall et al., 2003). Additionally, Ca2+ waves have also been shown in both zebrafish and Xenopus embryos undergoing gastrulation (Gilland et al., 1999; Wallingford et al., 2001). This pathway is involved in regulation of dorsal axis formation and convergent extension movements during gastrulation. Both WNT5A (Slusarski et al., 1997) and WNT11 (Westfall et al., 2003) ligands are capable of releasing intracellular Ca2+ leading to the activation of protein kinase C (PKC) and calcium/calmodulin-dependent protein kinase II (CAMKII) (Kohn and Moon, 2005; Slusarski and Pelegri, 2007). PKC and CAMKII can prompt NFκB, and CREB to translocate to the nucleus and activate nuclear factor associated with T cells (NFAT) (De, 2011).       20   Figure 1.2. Non-canonical WNT signaling pathways. (A)  Planar cell polarity is activated upon interaction of non-canonical WNT with FZD receptors and co-receptor (ROR, RYK or PTK7).  Signal is transmitted intracellularly through DVL to small GTPases, Rho and Rac.  Small GTPases will phosphorylate JNK, then translocate to the nucleus and  activate transcription factors. This eventually leads to cytoskeletal rearrangement. (B)  Activated calcium signaling via ligand-receptor binding increases  the intracellular Ca2+ mediated by phospholipase C (PLC).  Increased Ca2+ leads to activation of CAMKII, calcineurin, and PKC, which then activates transcription factor, NFAT . Note: WNT (WNT5A) signaling can also activate other receptors without FZD, such as the homodimerization of ROR2 that antagonizes canonical signaling or activate non-canonical signaling.  1.4.6 Endpoints for WNT signaling The general readout for WNT signaling is activation of specific transcription factors, dependent on the particular WNT pathway. In canonical signaling, WNT/β-catenin activity is generally mediated by TCF/LEF family of transcription factors (Molenaar et al., 1996). Several groups have designed transgenic reporters containing multiple TCF binding sites, which entails a luciferase-   21 expressing transfection construct that responds to WNT/β-catenin, named TopFlash. The TopFlash reporter has three TCF binding sites upstream of a c-fos promoter driving luciferase expression (Korinek et al., 1997); whereas, an advanced reporter construct named SuperTopFlash includes eight copies of the TCF/LEF binding sites (Veeman et al., 2003). This assay is simple, robust, quantitative, and scalable allowing for rapid identification in components of the WNT/β-catenin pathway. Luciferase reporters to monitor PCP activity have been reported to measure activity of the AP-1 (activator protein-1) response elements (Le Floch et al., 2005); however, the reporters do not appear to be well characterized so they are not used as frequently. Other assays have been described in the literature to monitor JNK-phosphorylation (Yamanaka et al., 2002) or observe for cell polarity (Antic et al., 2010), but polarity defects may not provide information about whether there is too much or too little activation of the pathway. The JNK signal transduction pathway is involved in activating transcription factors jun and ATF2 (Weston and Davis, 2007). A group designed a readout for ATF2 luciferase to monitor JNK-PCP signaling and they found that high doses of WNT5A or WNT11 activated ATF2 luciferase reporter (Ohkawara and Niehrs, 2011). Calcium WNT signaling is activated by nuclear factor of activated T-cells (NFAT) luciferase activity that is regulated by calcineurin (Rao et al., 1997).  Additionally, reporter transgene mice for canonical WNT signaling have been generated and are based on containing multiple TCF binding sites that either drive expression of LacZ or GFP—instead of driving luciferase (TOP-GAL; BAT-GAL) (Barolo, 2006; Barolo and Posakony, 2002; Brugmann et al., 2007; DasGupta and Fuchs, 1999; Maretto et al., 2003; Topol et al., 2003). A few other reporter lines have been established in zebrafish or Drosophila involving canonical WNT signaling (Chang et al., 2008; DasGupta and Fuchs, 1999; Dorsky et al., 2002), but are not as reliable as the mouse reporter lines. However, these in vivo reporters have revealed discrepancies in certain contexts in expression of tissue type, with weak or restricted patterns (Barolo, 2006; Barolo and Posakony, 2002). Also, it has been reported that some transgenic TCF sites do not recognize regions of known WNT/β-catenin signaling in the animal (Barolo, 2006). Thus, results obtained from these reporter animals should not be considered conclusive and further experiments should be conducted, or the limitations should be acknowledged. Additionally, TCF reporters can also respond to false negative responses such as crosstalk from another signaling pathway that might increase β-catenin levels independent of WNT signaling (Barolo, 2006; Brantjes et al., 2001; Cavallo et al., 1998). As mentioned above, transgenic constructs used for in vitro work including    22 the TopFlash reporter contains multiple TCF/LEF binding sites, whereby cultured cell lines may be subjected to increased levels of WNT signaling, then in the whole animal (Barolo and Posakony, 2002). The availability of quantitative reporters as reliable endpoints for signaling have aided in the study of WNT signaling. 1.5 Animal Models to study skeletal dysplasias Animal experiments are essential to understand the fundamental mechanisms governing disease and to aid with possible treatment. Bone disorders have a great impact on the general population and a better understanding of the disorder by use of animal models could mimic the pathologic condition and help test hypotheses and disease states. Studies in mice have contributed to understanding the mechanism and genetic basis of skeletal dysplasias. An in vivo system is necessary to study the 3D morphogenesis of the bones, as full ossification will not occur in culture systems. The best studied syndromes with skeletal phenotypes are those affecting FGF signaling. Mutations in FGF receptors cause Achondroplasia (Fgfr3; (Wilkie, 2005). Several of the human mutations have been knocked into the mouse genome and recapitulate the craniosynostosis phenotypes  Even though the mouse is a powerful genetic model, some human diseases cannot be fully modeled in the mouse. Some mutations have variable penetrance, mutations that cause haploinsufficiency in humans often give no phenotypes in mouse heterozygotes, the strain background also impacts the expression of the phenotype. It is well accepted that variability in expressivity of a phenotype is due to the interaction of the gene of interest with the rest of the genome (modifier genes) and environmental factors (Fisch, 2017; Orgogozo et al., 2015; Symonds and Zuberi, 2018). Consequently, there is a need to test the variants of a particular syndrome in relevant tissue. The chicken provides a unique opportunity to isolate the dominant effects of mutation, with the ability to study several variants of a particular syndrome while minimizing the contributions of genetic background. The chicken embryo offers a multitude of advantages for studying the function of genes during development that have helped reveal fundamental mechanisms of human diseases (Mok et al., 2015). Many genes have crucial roles in development thus, basic research on chickens has implications for understanding human health and disease. In the early stages of chicken development, chicken embryo morphology is similar to human; both are amniotes and their    23 development is similar (Streit et al., 2013). Experiments using the chicken embryo are highly reproducible and regulated, and we can easily access and observe chicken embryos in vivo at various stages of development. We can also observe complex processes of development such as cell migration and cell movement using time-lapse video microscopy (Chuai et al., 2009; Dean and Palmer, 2014; Li et al., 2017). Now that the chicken genome has been sequenced (International Chicken Genome Sequencing, 2004) forward and reverse genetics can be applied to phenotypes in chicken development and mutant chickens.  The chicken embryo is an ideal model to study as they are relatively easy to manipulate by the use of a window made through the shell of the egg. Furthermore, chicken embryonic development is fast and highly reproducible. The eggs can be incubated to any stage of interest and the embryos are large enough to allow for easy manipulation (Abramyan and Richman, 2018; Darnell and Schoenwolf, 2000; Davey et al., 2018b; Tickle, 2000). Chicken embryos have demonstrated a myriad of cell-cell interactions between neighboring tissue including epithelial-mesenchymal interactions, which are involved in DV outgrowth of the limb (Gros and Tabin, 2014), or mesenchymal-mesenchymal interactions involved in AP patterning of the limb (Riddle et al., 1993). The patterning mechanisms in limb development are conserved between vertebrates (Capdevila and Izpisua Belmonte, 2001). Research on the limb system is well established in the chicken limb bud including the identification of limb morphogens (Baranski et al., 2000; Gao et al., 2011; Pitts et al., 1986; Riddle et al., 1993), digit patterning (MacCabe et al., 1974; Summerbell et al., 1973; Wolpert, 1969), and fate mapping (Vargesson et al., 1997). Additionally, the chicken provides an excellent model for studying craniofacial development. The avian face provides highly conserved organization and growth of facial prominences (Abramyan et al., 2015; Schock et al., 2016). Much of what we know about facial patterning and growth was conducted in the chicken embryo including cranial neural crest cell migration (Couly et al., 1996; Couly et al., 1998; Couly et al., 1993; Le Douarin et al., 2004; Minoux and Rijli, 2010), fate mapping (Couly et al., 1993; Kontges and Lumsden, 1996; Noden, 1978, 1983), and facial patterning (Noden, 1983; Trainor et al., 2002; Tucker and Lumsden, 2004). The chick embryo has been beneficial for classical approaches such as the ‘cut and paste’ experiments including: grafting tissue to different regions of the embryo (Richman and Tickle, 1992; Van Alten and Fennell, 1959), creating chick:quail chimeras via grafting (Eames and Schneider, 2008; Fish et al., 2014; Le Douarin et al., 1996; Schneider and Helms, 2003), or bead implants (Ashique et al., 2002; Cela et al., 2016; Higashihori    24 et al., 2010; Lee et al., 2001; Mohammed and Sweetman, 2016; Richman and Delgado, 1995; Song et al., 2004; Szabo-Rogers et al., 2008). These experiments elucidated signaling interactions and pathways in vivo to directly test the effects of particular growth factors or inhibitors. The weakness of the chicken embryo is that it is not a genetic model. Producing transgenic chickens is a complex and laborious process. There are however, methods to introduce foreign DNA into a chicken embryo in a targeted manner (Gordon et al., 2009; Logan and Tabin, 1998; Mozdziak and Petitte, 2004). 1.6 Local transgenesis in chicken embryos The use of transgenic chickens has lagged behind that of the mouse model. However, recent advances in science have shown the ability to edit the chicken genome using several different techniques. Genome editing using the CRISPR/Cas methodologies can be used to modify endogenous genes, generally conducted with in vivo electroporation in somatic cells of the developing chicken embryo (Morin et al., 2017). However, the use of electroporation to deliver genes into the developing chicken limb could cause truncation (Suzuki and Ogura, 2008), making this study difficult as we are observing truncated/shortened limbs. CRISPR chickens can also be produced by injecting genome-edited primordial germ cells (PGCs) into a developing embryo until hatching (Han and Lee, 2017), but they run the risk of lower transfection efficiency than other methods such as retroviral vectors (Davey et al., 2018a).  Due to some setbacks with genome editing in the chicken embryo, we turn our studies to implementing retroviral vectors to generate transgenic chickens. Retroviral vectors are a vehicle used to introduce foreign genes into the host DNA (Nishijima and Iijima, 2013). Recently, researchers have been able to develop transgenic chickens via injection of retroviral vectors into chicken embryos at the blastoderm stage in PGCs (Nishijima and Iijima, 2013). However, these approaches remain challenging, as it is difficult to successfully target PGCs in the chicken embryo (Nakamura et al., 2013; Nishijima and Iijima, 2013). The first genetically modified chicken was established using foreign retroviral DNA into the germline via avian leukosis viruses (Salter et al., 1987). Since then, several retroviral vectors have been routinely used to develop transgenic chickens. Lentivirus are a class of retroviruses that are able to introduce transgenes into host DNA even in non-dividing cells, which can lead to reduced efficiency (Hughes, 2004; Naldini et al.,    25 1996). Most other retroviruses require the breakdown of the nucleus in order to integrate foreign DNA into the host, thereby involving cell division (Nishijima and Iijima, 2013).  The avian-specific RCAS retroviruses stands for: Replication-Competent Avian sarcoma-leukosis virus (ASLV) long terminal repeat (LTR) with a Splice acceptor. RCAS vectors are derived from a Rous sarcoma virus (RSV) from the ASLV family (Hughes, 2004). RSV acquired a cellular oncogene, src, thus rendering it replication competent. The src site was replaced with a unique restriction site (ClaI) to insert foreign genes. Replication competent retroviruses contain an intact viral genome and packaging genes to permit continuous production of infectious particles to infect other cells (Hughes, 2004). The viral envelope protein binds to specific receptors on the target cell creating a fusion of the viral membrane and membrane of the target cell. This fusion introduces the virion core into the cytoplasm of the target cell (Mozdziak and Petitte, 2004). Viral RNA is converted into cDNA by reverse transcriptase, which gets incorporated into the host genome. Expression of the inserted gene is driven by the viral LTR. The integration of viral DNA into the host genome occurs at many places, and rarely does inserted viral DNA affect a critical role in regulating cell growth; most oncogene-containing retroviruses are replication-defective (Hughes, 2004). Proviral DNA can get transcribed into viral RNA for synthesis of viral proteins including: polymerase (POL), group-associated antigens (GAG), and envelope (ENV). The POL region affects replication and expression in avian cells and substituting the POL region from the Bryan high-titer strain generated a virus that replicated one log better than RCAS, named RCASBP for RCAS Bryan Polymerase (Petropoulos and Hughes, 1991). RCASBP derivatives are also available that allow insertion of genes using the Gateway system named, RCASBPY (Loftus et al., 2001).  Use of cross-species genes offers the ability to follow the transgene in the chicken embryo. Point mutations (missense or nonsense) can be introduced into the human gene prior to cloning into the retrovirus, and the use of microinjecting the virus into the embryo allows us to target the gene of interest temporally and spatially (Gordon et al., 2009; Logan and Tabin, 1998).       26 1.7 Rationale Understanding particular gene functions and genotype-phenotype relationships involved in dominant RS require an appropriate animal model. There are no appropriate animal models for dominant RS and there is no informative data about the cell behaviour impacted by these mutations. Currently, the only animal models that represent RS is overexpression of mutant ROR2 in chicken (Stricker et al., 2006), and the Ror2-/- and Wnt5a-/- mice (Oishi et al., 2003). With the ROR2 chicken model, Stricker et al. (2006) used the RCAS expressing system with a human mutation in ROR2. Overexpression of the truncated ROR2 gene in the chicken limb led to disruption of the growth plate with reduced skeletal elements (Stricker et al., 2006). However, the full-length wild-type ROR2 could not be used in their system due to size restraints of the RCAS vector. It is unclear whether the effects of the gene truncation are due to increased levels of ROR2 or due to the mutation itself. The Wnt5a-/- and Ror2-/- mice both consist of similar phenotypes consisting of PD limb defects with truncated stylopod and zeugopod, as well as outgrowth defects in the face (DeChiara et al., 2000; Oishi et al., 2003; Schwabe et al., 2004; Yamaguchi et al., 1999; Yang et al., 2003). However, phenotypes are not the same as in RS. The missense mutation in Ror2 (W745X) knocked into mouse more successfully recapitulated the recessive RS phenotypes (Raz et al., 2008). There are no appropriate animal models that represent the DVL1 RS phenotypes as the conditional knockouts of Dvl1 mice show redundancy (Wynshaw-Boris, 2012). The proposed experiments define an appropriate animal model to study dominant RS. In this study we will use retroviruses to deliver the genes to the chicken embryo. Other methods for gene delivery into the chicken embryo have been established (Sato et al., 2007; Suzuki and Ogura, 2008; Ueda et al., 2017). It is common to use electroporation where plasmids are driven into cells in a localized region (usually next to a body cavity to contain the DNA). The most successful use of electroporation has been for the neural crest cells (Krull, 2004; Sauka-Spengler and Barembaum, 2008), or the somites (Scaal et al., 2004). However, electroporation of genes into the limb field gives patchy expression and may result in truncated or shorter limb buds (Suzuki and Ogura, 2008). This method is not suitable for my study as I am observing a type of dwarfism. Moreover, a major disadvantage of electroporation is that most vectors only cause transient transfection. Even with constructs that can be integrated stably, it is hard to get enough cells to express the transgene to see a phenotype. RCAS retroviruses will be used to introduce misexpression of both wild-type and variant genes into the developing chicken embryo, which has    27 worked proficiently for our lab (Bond et al., 2016; Geetha-Loganathan et al., 2014; Hosseini-Farahabadi et al., 2013; Hosseini-Farahabadi et al., 2017) and many others in the past (Abzhanov et al., 2007; Hartmann and Tabin, 2000; Stricker et al., 2006). The results will show how the mutations in WNT5A or DVL1 induce a phenotype and will help discover the underlying mechanisms involved in dominant RS.  1.8 Approach This project was initiated by the identification of two cases of a novel disorder, dominant RS from the original study with missense mutations in WNT5A (Person et al., 2010). First, I will be studying the effects of two mutations in WNT5A (C83S and C182R) in both the limb and the face. This will help elucidate the functions of the mutations during limb and mandibular morphogenesis by using retroviral misexpression to locally modify expression of mutant WNT5A in vivo. Reproducibility of the phenotypes will allow for analysis of cellular defects at earlier stages of chondrogenesis. I will compare the results obtained in the limb to results in the face. I will also conduct in vitro analysis using cell lines by using genetic reporters to study gene expression of the mutations in order to identify specific WNT signaling pathways that the mutations are interfering with.  My second approach is to study three genetic mutations involved in dominant RS that are the result of mutations in the adaptor protein of the WNT pathway, DVL1. I will study the effects of three DVL1 mutations (1519∆T, 1529∆G, and 1615∆A) in limb development, similar to experiments conducted with the WNT5A mutations. I will look at several stages of embryonic development in attempt to pinpoint when the mutations initially affect chondrogenesis. I will also conduct luciferase assays to understand the mechanism behind the mutations as DVL1 is involved in all WNT signaling pathways.  1.9 Hypotheses 1) Dominant RS mutations affect elongation of cartilage by directly affecting the arrangement of chondrocytes. 2) RS mutations increase the activity of the JNK-PCP pathway. 3) RS mutations indirectly affect canonical signaling since WNT5A is a well-known antagonist of the canonical pathway.    28 4) DVL1 and WNT5A mutations have similar biochemical effects since people with mutations in these two genes have a similar phenotype.  1.10 Novelty  This study will clarify the skeletogenic mechanisms of mutations involved in dominant RS through molecular and phenotypical studies using the chicken model system. These mutations affect the less studied, non-canonical WNT signaling pathway. In a broader sense, this project can aid in understanding limb deficiencies as well as those affecting other developmental systems such as the face. The methods developed here (whole animal, tissue and biochemical) pave the way for functional testing of human gene variants involved in a variety of diseases. 1.11 Aims and Objectives Aim 1. Analysis of skeletal morphogenesis in avian limbs and mandible using retroviruses that contain dominant RS mutations in WNT5A. This involves morphometric characterization of the phenotypes by looking at shape changes of the developing limb and mandible. I will also observe how the mutations influence cell polarity in the developing chondrocytes by immunostaining with cell polarity markers. Aim 2. Biochemical characterization of dominant RS mutations in WNT5A. I will look at how the RS mutations in WNT5A affect its secretion. I will characterize the activity of dominant RS mutations using luciferase assay with a canonical and non-canonical WNT reporter plasmid. Aim 3. Analysis of skeletal morphogenesis in avian limbs using retroviruses that contain dominant RS mutations in DVL1. This will be done by observing chondrogenesis at various stages of development and looking at cell polarity changes in the limb.  Aim 4. Biochemical characterization of dominant RS mutations in DVL1. I will characterize the activity of dominant RS mutations using luciferase assays with canonical and non-canonical WNT reporter plasmids.      29 2. Materials and Methods 2.1 Chicken embryos White leghorn eggs received from the University of Alberta were incubated to the appropriate embryonic stages, according to the Hamburger and Hamilton (HH) staging guide (Hamburger and Hamilton, 1951). Neutral Red stain (0.33%, Fisher #N129) was used to enhance visualization of the embryos while staging. This study on prehatching embryos was considered exempt by the University of British Columbia Animal Care Committee and the Canadian Council on Animal Care.  2.2 Cloning of WNT5A virus and plasmid constructs The open reading frame containing human WNT5A (Genbank Ref seq: NM_003392.4) was purchased from Invitrogen (#IOH39817). We used site direct mutagenesis with restriction-free cloning (Bond and Naus, 2012) to knock-in two mutations that cause autosomal dominant RS2 (OMIM: 180700): (1) the 248G-C mutation in exon 3 of the WNT5A gene, resulting in a cysteine 83 to serine conversion (C83S) and (2) the 544/545CT-TC inversion in exon 4 resulting in cysteine 182 to arginine (C182R) (Table 2.1). A C-terminal Flag-tag was cloned after the coding sequence and a stop codon was added to the 3’ end of the tag. Gateway cloning (Invitrogen) was used to move the mutant or wild-type sequences from pENTRY into pcDNA3.2 for plasmid transfection and RCASBPY for retroviral expression (referred to as RCAS for simplicity) as described (Hosseini-Farahabadi et al., 2017). The human FZD2 coding sequence in a shuttle vector was purchased from GeneCopoeia (S01933) and recombined into pcDNA3.2 using Gateway recombinase (Invitrogen). Other viruses containing GFP (green fluorescent protein) (A. Gaunt) or Alkaline Phosphatase (AlkPO4) (L. Niswander) were generously provided by other investigators.   Table 2.1. Primer sequences for WNT5A variants.  WNT5A Mutation Primer Sequence C83S (284GC) Fw: GACAGAAGAAACTGTCCCACTTGTATC  Rev: GTGGTGAACGCCATGAGC  C182R (544-545CTTC)  Fw: GGCGGTCGCGGCGACAAC  Rev: GGCCTGCAAGTGCCATGGG      30 2.3 Cloning of DVL1 virus and plasmid constructs The open reading frame containing human DVL1 (Origene #RC217691) came in vector pCMV6 with full coding sequence and two tags, myc and analog. We moved DVL1 to pDONR221 with Gateway cloning, where all mutations were performed in and then got recombined into destination vector with LRclonase2 (Life Technologies). We created three mutations that cause autosomal dominant RS 2 (OMIM: 616331) using site direct mutagenesis with restriction-free cloning: (1) the 1519∆T mutation of the DVL1 gene, resulting in a nonsense-frameshift mutation, (2) the 1529∆G mutation of the DVL1 gene, resulting in a nonsense-frameshift mutation, and (3) the 1615∆A mutation of the DVL1 gene, resulting in a nonsense-frameshift mutation (Table 2.2). Each mutation was created using site directed mutagenesis with STOP added to the C-terminus. A Kozak sequence followed by an N-terminal Flag-tag were cloned upstream of the coding sequence. A STOP codon was added to the 3’ end of the coding sequences for the wtDVL1. The mutant constructs resulted in a natural stop codon due to the frameshift.  Gateway cloning (Invitrogen) was used to move the mutant or wild-type sequences from pENTRY into pcDNA3.2 for plasmid transfection and RCASBPY for retroviral expression (RCAS) as described. GFP virus (A. Gaunt) was generously provided by another investigator (S. Loftus).   Table 2.2. Primer sequences for DVL1 variants. DVL1 Mutation Primer Sequence 1519 (∆T) Fw: GCTGCCCCCGGCCTCTGGGTCAG Rev: CTGACCCAGAGGCCGGGGGCAGC 1529 (∆G)   Fw: CTGGCCTCTGGTCAGGGCTACCCCTAC Rev: GTAGGGGTAGCCCTGACCAGAGGCCAG 1615 (∆A)   Fw: CTATGGCAGCGGCGCACCGGGAG Rev: CTCCCGGTGCGCCGCTGCCATAG  2.4 Retrovirus construction of WNT5A and DVL1 variants Retroviruses were propagated by using DF1 chicken cell line (from ATCC #CRL-12203). DF1 cells were maintained at 37°C in DMEM (Life Technologies #1967497) medium supplemented with 10% fetal bovine serum (FBS), (Sigma #F1051), and 1% penicillin/streptomycin (Life Technologies #15070-063). Cells were kept in 100 mm culture dishes, and media changed every    31 other day, with cultures passaged 1:2 two to three times/week using trypsin-EDTA (0.25%, Life Technologies 25200-072). Human WNT5A, two human WNT5A variants (C83S and C182R), human DVL1 and three human DVL1 variants (1519∆T, 1529∆G, 1615∆A), as well as GFP control RCAS vector constructs were transfected into DF1 chicken fibroblast cell lines using Lipofectamine3000 (Invitrogen #L3000-008), according to the manufacturers protocol with 2.5 µg DNA. After one month of culture, conditioned media were collected from confluent cultures x2 and transferred to a 1x3½ inch Polyallomer centrifuge tube (Beckman Coulter #326823) and spun at 25 000 rpm at 4°C using swing bucket rotor SW28 (Beckman #97U 9661) in a Beckman centrifuge for 2.5h (no break) to pellet the viral particles. Supernatant were removed and 50-100 µl Optimem (Life Technologies #319850962) were added to the pellet and incubated overnight at 4°C. Concentrated viral particles were mixed, aliquoted, flash frozen in methanol + dry ice, then stored at -80°C.  2.5 Retrovirus injection into the limb and face Eggs were incubated in a humified incubator at 38°C until stage HH15 and concentrated RCAS retrovirus viral particles (~5-10 µl) combined with Fast Green FCF stain (0.42%, Sigma #F7252) (1-2 µl) were injected into the forelimb region (between somites 15 and 16); or injected into the first pharyngeal arch with glass filament needles (thin-wall borosilicate capillary glass with microfilament, A-M systems #615000) using a Picospritzer microinjector (General Valve Corp.). We chose this stage in order to give the virus sufficient time to replicate prior to key events in limb and mandibular patterning and overt differentiation. Table 2.3 outlines the embryos studied for forelimb and mandible analyses in both chapters 3 and 4 with the fixative described.        32 Table 2.3. Embryo stage and fixative post-injection.  Thesis Chapter Forelimb/ Mandible Days post-injection Stage euthanized (HH) Fixative 3 Forelimb 4 29 4% PFA1   4.5 30 4% PFA   10 36 4% PFA   12 38 100% ethanol 3 Mandible 3 28 4% PFA   4 29 4% PFA   12 38 100% ethanol 4 Forelimb 4 29 4% PFA   8 34 4% PFA   12 38 100% ethanol 1PFA=Paraformaldehyde  2.6 Whole-mount skeletal staining  Injected embryos were grown to appropriate stages. Skulls were dissected from the embryo and forelimbs dissected from the flank of the embryo and processed to 100% ethanol for 4 days for bone and cartilage staining. Limbs were then transferred to acetone for 4 days, then stained for 10 days at room temperature on a shaker; stain: 1 volume 0.3% Alcian blue 8GX (Sigma #A5268) in 70% ethanol, 1 volume, 0.1% Alizarin Red S (Sigma #A5533) in 95% ethanol, 1 volume Acetic acid, 17 volume 70% ethanol. Following staining forelimbs and skulls are cleared for 24h in 2% potassium hydroxide (KOH), followed by several days in 2% KOH, 20% glycerol, then transferred to 50% glycerol and stored in 100% glycerol at room temperature. Specimen were photographed in 50% glycerol using a glass petri dish with agarose, images were taken with a Leica M125 stereomicroscope. Measurements of limbs were made using ImageJ on the photographs. The length of the radius and ulna (humerus also included in the DVL1 project) and the width (AP diameter through the diaphysis) was measured in the WNT5A injected forelimbs. Mandible length was measured in wholemount skulls.     33 2.7 Histology  Embryos fixed in 4% PFA were used for histological analysis and were in fixative for 2-3 days. Limbs at stage HH34 or 36 were decalcified in 12% EDTA (Fisher #S312-212) at 4°C on a shaker for 4-5 days prior to processing into wax; changing solution every day. Specimen were processed to 1X Phosphate Buffered Saline (PBS; 137 mM NaCl, 8.1 mM Na2HPO4, 2.7 mM KCl, 1.5 mM KH2PO4; pH 7.3) for 30 min 2x, 50% Ethanol 2x, then 70% Ethanol 2x. Specimen were then transferred to histological cassettes and were either sent to the Biomedical Research Centre for tissue processing or placed in a wax tissue processor (Leica #ASP300S), from 70% ethanol to 70% isopropanol to absolute isopropanol, with several changes of xylene and followed by paraffin wax. Wax-embedded tissue were positioned in paraffin wax and sectioned using a microtome, sectioned at 7 µm. Selected sections (sagittal or transverse for forelimb; frontal for mandible) were stained to see the differentiated cartilage and bone. Sections were first dewaxed in xylene and rehydrated from 100% ethanol to water, then stained with 1% Alcian blue 8GX (in 1% acetic acid) for 30 minutes. After staining sections were rinsed in 1% acetic acid, then rinsed in water. Following that, sections were stained in Picrosirius Red (0.1% Sirius Red F3B in saturated picric acid) for 1h, then rinsed in 1% acetic acid and dehydrated through ethanol, back to xylene, followed by Shandon Consul-mount (Thermo Scientific #9990441). 2.8 Immunofluorescence Immunofluorescence analysis was carried out on forelimbs and mandibles at various stages of development. Table 2.4, Table 2.5 and Table 2.6 outline specific antibodies and treatments performed for each assay, listing specific antibody markers regarding: viral expression (anti-GAG), hypertrophic cartilage (COL10A), cell polarity (Golgi, Prickle), cell shape (non-muscle myosin II, phalloidin), and chondrocytes (SOX9). Primary antibodies were incubated overnight at 4°C and secondary antibodies were incubated at room temperature for 1h unless otherwise stated. Phalloidin staining was carried out in wholemount 200 μm thick slices of limbs since this reagent is incompatible with ethanol. Limb slices were fixed for 48h at 4°C in 4% paraformaldehyde. Sections were either counterstained with Hoescht (10 μg/ml #33568, Sigma) and incubated for 30 min at room temperature, then mounted with Prolong Gold antifade (Life Technologies #P36930); or counterstained with DAPI mount (Molecular Probes #P36935). Fluorescence images were    34 collected with a Leica SP5 confocal microscope or with a 20X objective on a slide scanner (3DHISTECH Ltd., Budapest, Hungary).    35 Table 2.4. Antibodies and immunofluorescence treatments for forelimb WNT5A assays.   Stage (HH) Antigen Retrieval (steam 15min 95°C) Permeabilization/ Pre-treatment Block Primary antibody Secondary Antibody Counterstain Forelimb (7 µm sections) 29, 30, 36 10 mM sodium citrate   10% GS1, 0.5% tween-20/PBS 1h 3C2/GAG Developmental Studies Hybridoma bank (DSHB), 1:4 #AMV-3C2 Invitrogen, anti-mouse A11029 1:200 Hoechst  36 Diva Decloaker Biocare #DV2004MX 1:10 0.5% hyaluronidase in Hank’s Balanced Salt Solution, 30min (prior to antigen retrieval) 10% GS, 0.5% tween-20/PBS 1h Collagen X (COL10A1) DSHB, 1:250 #X-AC9-c Invitrogen, anti-mouse A11029 1:200 Hoechst /  TO-PRO-3 iodide (Life technologies, #T3605) 29 Diva Decloaker 1:10 0.5% tween-20, 10 min 10% GS, 0.5% tween-20/PBS 1h Pan-Prickle  Abcam, 1:50,  #15577 Invitrogen, anti-rabbit A11034, 1:200 Hoechst Forelimb (200 µm whole-mount sections) 29  0.5% triton-X,  30 min 10% fetal bovine serum, 0.5% triton-X/ PBS 30min 2x Golgi antibody (GM130)  BD labs, 1:100, #610822 4°C, 96h, rocker Invitrogen, anti-mouse,  A-21236, 1:200 In 0.5% triton-X/PBS, 4°C, 96h, rocker Phalloidin-tagged-568 (Molecular Probes, 1:50, #A12380) 4°C, 48h, rocker;  Hoechst (2h) SOX9  Sigma-Aldrich, 1:200, #HPA001758 4°C, 48h, rocker Invitrogen, anti-rabbit A11034,1:200 In 0.5% triton-X/PBS, 4°C, 48h, rocker 1GS = goat serum (Sigma #G9023)    36 Table 2.5. Antibodies and immunofluorescence treatments for mandible WNT5A assays.   Stage (HH) Antigen Retrieval (steam 15min 95°C) Permeabilization Block Primary antibody Secondary Antibody Counterstain Mandible (7 µm sections) 29 10 mM sodium citrate  10% GS,  0.5% tween-20/PBS, 1h 3C2/GAG  DSHB, 1:4 #AMV-3C2 Invitrogen, anti-mouse A11029 1:200 DAPI 29 Diva Decloaker Biocare #DV2004MX 1:10 0.5% tween-20, 10 min 10% GS, 0.05% tween-20/PBS, 1h Non-muscle myosin II, 5 μg/ml  DSHB #CMII 23 Invitrogen, anti-mouse A11029 1:200 DAPI 29 Diva Decloaker 1:10 0.5% tween-20, 10 min 10% GS, 0.05% tween-20/PBS, 1h SOX9  Sigma-Aldrich, 1:200, #HPA001758 Invitrogen, anti-rabbit A11034, 1:200 DAPI  29 Diva Decloaker 1:10 0.5% tween-20, 10 min 10% GS, 0.05% tween-20/PBS, 1h Golgi antibody (GM130), 1:100, BD labs, #610822 Invitrogen, anti-mouse,  A-21236, 1:200 DAPI       37 Table 2.6. Antibodies and immunofluorescence treatments for forelimb DVL1 assays.  Stage (HH) Antigen Retrieval (steam 15min 95°C) Permeabilization/ Pre-treatment Block Primary antibody Secondary Antibody Counterstain Forelimb (7 µm sections) 29, 34 10 mM sodium citrate  10% GS,  0.5% tween-20/PBS, 1h 3C2/GAG  DSHB, 1:4 #AMV-3C2 Invitrogen, anti-mouse A11029 1:200 Hoechst 34 Diva Decloaker  1:10 0.5% hyaluronidase in Hank’s Balanced Salt Solution, 30min (prior to antigen retrieval) 10% GS, 0.5% tween-20/PBS 1h Collagen X (COL10A1) DSHB, 1:250  #X-AC9-c,  Invitrogen, anti-mouse A11029 1:200 Hoechst 29 Diva Decloaker 1:10 0.5% tween-20, 10 min 10% GS, 0.05% tween-20/PBS, 1h Golgi antibody (GM130), 1:100, BD labs, #610822 Invitrogen, anti-mouse A10524 1:200 Hoechst 29 Diva Decloaker 1:10 0.5% tween-20, 10 min 10% GS, 0.5% tween-20/PBS 1h Pan-Prickle  Abcam, 1:50,  #15577 Invitrogen, anti-rabbit A11034, 1:200 Hoechst 29 Diva Decloaker 1:10 0.5% triton-X,  10 min 10% GS, 0.2% triton-X/PBS 1h SOX9  Sigma-Aldrich, 1:200, #HPA001758 Invitrogen, anti-rabbit A10523, 1:200 Hoechst    38 2.9 Quantification of chondrocyte polarity and shape Embryos were fixed at stage HH29 and immunofluorescence analysis for forelimb and mandibles were performed for Golgi and phalloidin or non-muscle myosin described in section 2.8. ImageJ software was used to measure cell polarity and shape. Golgi angles between 0° and 90° (Golgi-nucleus angle relative to the long axis of the cartilage in the diaphysis region of the forelimb zeugopod, or the long axis of Meckel’s cartilage in the mandible) were recorded for 70 chondrocytes in forelimb with WNT5A injected viruses (section 3.2.7), 100 chondrocytes in mandible with WNT5A injected viruses (section 3.2.10), or 75 cm2 analyzed in forelimb with DVL1 injected viruses (section 4.2.4). Angle orientation was plotted with Rose.net software. To quantify cell shape, ImageJ software was used to measure feret diameters (the longest and shortest diameters of an ellipse on perpendicular axes) for 50 chondrocytes in the forelimb and 75 chondrocytes in the mandible to assess cell roundness. Statistical analysis was conducted with analysis of variance and Tukey’s post hoc testing (Statistica 6.0), with an n=3 specimen per virus/analysis.  2.10 BrdU, TUNEL staining and analysis Apoptosis was looked at using TUNEL (Terminal deoxynucleotidyl transferase dUTP nick end labeling) assay on chick forelimbs stage HH29 and HH30 for Chapter 3. TUNEL was carried out using the ApopTag Apoptosis Kit (Chemicon #S7101) and was detected using anti-digoxigenin tagged with fluorescein. For cell proliferation studies, chicken embryos 48, 72 or 96h post-virus injection (stage HH28, 29 or 30) were labeled with 10 mM BrdU (bromodeoxyuridine) (Sigma #B5002), by injecting into the heart 2h prior to euthanasia. Forelimbs were studied at stages 29 and 30, antigen retrieval was performed with 2N HCl for 30 min at room temperature and treated with 0.1% trypsin/PBS for 10 min at 37°C. Tissue sections were blocked with 10% goat serum (Sigma #G9023) and 0.1% tween-20 for 1h then incubated overnight at 4°C in blocking solution with anti-BrdU (Developmental Studies Hybridoma bank, 1:20, #G3G4). Sections were then probed for anti-SOX9 (Sigma #HPA001758) (1:100) in block overnight at 4°C. The proportion of BrdU-labeled cells in the SOX9-positive region was counted using ImageJ. Sections were counterstained with Hoescht (10 μg/ml, Sigma #33568) and incubated for 30 min at room temperature, then mounted with Prolong Gold antifade (Life Technologies #P36930); HH29    39 forelimb, n=4 GFP, n=5 wtWNT5A and WNT5AC83S; DVL1 wild-type and variants n=3; at HH30 forelimb, n=4 GFP, n=5 wtWNT5A and WNT5AC83S. Mandibles were studied at stages HH28 and 29, and DNA was digested with Sau3AI (Roche, Mannheim, Germany) in Buffer A (Roche) at 37°C for 30 min. Tissue sections were blocked with 10% goat serum and 0.1% tween-20 for 1 hour then incubated overnight at 4°C in blocking solution with anti-BrdU (GE Healthcare #RPN202). Sections were counterstained with DAPI (Prolong Gold Antifade mountant with DAPI, Molecular Probes #P36935). Fluorescence images were collected with a Leica SP5 confocal microscope or with a 20X objective on a slide scanner (3DHISTECH Ltd., Budapest, Hungary). The proportion of BrdU positive cells to total cells was determined in half of the infected mandible (right side), or the entire ulna of the developing forelimb (n=4 virus/stage). 2.11 Quantification of forelimb cell density and morphometrics  To measure cell density in the forelimb at stage HH36 for embryos injected with WNT5A viruses regional cell density in the ulna was quantified in sagittal sections with nuclear stain. Nuclei located in a 200 μm2 region in the epiphysis and diaphysis of the ulna, as well as cell density in half of the developing ulna (closest to the digits) were manually quantified using ImageJ software (n=4 GFP, wtWNT5A, WNT5AC83S and WNT5AC182R). Sagittal measurements of the ulna were conducted at stage HH29, 30 and 36, which included the distance from the epiphysis near the digits to epiphysis near the humerus, the width in the centre of the diaphysis and area (around entire ulna) (for stages 29 and 30, n=4 GFP; n=5 wtWNT5A and WNT5AC83S; for stage HH36 n=4 for all conditions). Transverse sections performed on stage HH36 limbs perpendicular to the long axis of the cartilage, through the ulna spanning the epiphysis (closest to the digits) and the diaphysis (n=4). 2.12 Contrast enhanced micro CT of limb buds and volumetric analysis  To measure skeletal condensations in 3D, embryos injected with WNT5A viruses (Chapter 3), fixed at HH29 in 10% formaldehyde and stained with phosphotungstic acid (PTA). Specimens were washed several times in water and dehydrated from 30, 50 and 70% ethanol (each concentration per day). Then transferred to ethanol-methanol-water mixture (4:4:3) for 1h, 80% methanol for 1h, 90% methanol for 1h, then to 0.7% PTA-methanol solution (0.7% PTA dissolved in 90% methanol). Specimen are kept in this solution for 6 days, with solution changed every day. Specimens then get rehydrated through a methanol series (90%, 80%, 70%, 50% and 30%)    40 finishing in sterile distilled water. Limbs are supported in 1% agarose in a 2 ml Eppendorf. Specimens in agarose are next dehydrated back into 70% ethanol. Limbs were scanned with a Scanco100 microCT scanner at a resolution of 10 µm (55 kVp, 60 µA exposure, 0.5-mm Al filter, calibrated with 1,200 mg of hydroxyapatite/cm3). Dicom files were imported into Amira v. 6.3 (Thermofisher/ FEI) and cartilage condensations were segmented. Volumes were exported from Amira (n=3). 2.13 Cell culture and Luciferase reporter assay Micromass cultures were established to initially test transfection of WNT5A variants. Stage 24 forelimb buds were used for micromass cultures as described (Underhill et al., 2014). Limbs are dissected in Hank’s balanced saline solution (without calcium and magnesium), with 10% FBS. Limbs were cut into smaller pieces (~100 µm2), and then put on a bacterial shaker at 70 rpm, 37°C for 1h in 10% Dispase II (Sigma #D4693). Media was added to inhibit the Dispase and centrifuged at 1000 g, 4°C for 5 minutes. Supernatant was removed and add fresh micromass media were added: 10% FBS (Sigma #F1051), 52% F12 (Life Technologies #11765-054), 34% DMEM (Life Technologies #1967497), 1%, Antibiotic-antimycotic (Life Technologies #15240-062), 1% L-Glutamine (Life Technologies #25030), 0.5% Ascorbic acid (Life Technologies #850-3080IM)). Cell solution were then passed through a cell strainer (Falcon #2340) to remove epithelial tissue. Micromass cultures were seeded in 10 µl droplets/well in a 24 well TC plate (NUNC #142475) that contain 2x105 cells/droplet. Plates were incubated for 30 minutes at 37°C to allow cultures to attach, and transfected with plasmids using Lipofectamine3000 (Invitrogen #L3000-008), then incubated overnight at 37°C. The following plasmids were used: Empty (pcDNA3.2/V5-DEST), WNT5A, WNT5AC83S, WNT5AC182R at 0.3 μg. Additionally, Renilla luciferase reporter plasmid was transfected for normalization (0.05 μg), along with Firefly reporter plasmid: Activating Transcription Factor 2 (ATF2, (Ohkawara and Niehrs, 2011)) at 0.2 μg. After 24h, media was changed replaced and 100 ng/ml WNT5A protein were added. Luciferase assays were performed after 48h of culture using Dual-luciferase reporter assay system (Promega #E1910).  Luciferase assay were also performed on cell lines using HEK293 cells. HEK293 were maintained at 37°C in DMEM (Life Technologies #1967497) medium supplemented with 10% FBS, (Sigma #F1051), and 1% penicillin/streptomycin (Life Technologies #15070-063). Cells were kept in 100 mm culture dishes, and media changed every other day, with cultures passaged    41 1:4 twice/week using trypsin-EDTA (0.25%, Life Technologies 25200-072). Transient transfections for luciferase assays were performed on 30-40% confluent HEK293 cells (0.17-0.18x106 cells/ml). Cells were transfected with Lipofectamine3000 (Invitrogen #L3000-008) 24h after plating in 24-well plates (Invitrogen #L3000-008; Nunc #142475). Table 2.7 outlines the plasmids (pcDNA3.2/V5-DEST) used for transfection along with Firefly reporter plasmids. Luciferase assays were performed after 48h of culture using Dual-luciferase reporter assay system (Promega #E1910). A Tecan luminometer (Spark® multimode Tecan plate reader) was used to read luminescence activity at one second reading with OD1 filter. At least 3 technical and 3 biological replicates were carried out for each transfection mixture and the experiment was repeated on 2 different days.       42 Table 2.7. Plasmids and stimulates used for luciferase assays on HEK293 cells.   Chapter Firefly assay (0.2 µg) Plasmid (Gene of interest)* Gene of interest concentration (µg) Renilla StimulantΨ 3 ATF2  STF Empty WNT5A 0.3 0.05 STF assay: LiCl (6 mM) or WNT3Aω (100ng/ml)  NFAT¥ WNT5AC83S      WNT5AC182R  mRor2†   NFAT assay: caNFAT plasmid 3 ATF2 Empty 0.6 0.01    WNT5A      WNT5AC83S      WNT5AC182R FZD2†    4 ATF2 Empty DVL1 0.6 0.01 ATF2 assay: WNT5A§ (100 ng/ml)   DVL11529∆G    4 NFAT  STF Empty DVL1 DVL11529∆G 0.3 0.05 STF assay: LiCl (24 mM) or WNT3A (200 ng/ml)        NFAT assay: caNFAT plasmid *Not all plasmids are used for each firefly assay, e.g., Ror2 is only used in ATF2 reporter assay. ΨStimulates for STF assays are added 24h before lysing—except for NFAT reporter assay, caNFAT plasmid is added at time of transfection.  †Assays with receptors FZD2 (Genecopoeia, clone GC-S0193-B) or mROR2 (Addgene #22613) are in combination. Half the amount of each plasmid is used; e.g., Chapter 3 with ATF2 assay using FZD2 receptor: 0.3 µg of each plasmid is used (WNT5A 0.3 µg + FZD2 0.3 µg), totalling 0.6 µg. ωR&D, #5036-WN-010  §R&D, #645-WN  ¥Addgene #10959  2.14 Immunocytochemistry with transfected WNT5A or DVL1 variants  HEK293 cells were cultured as described in section 2.13. Coverslips were first washed in 70% ethanol 2x, PBS 2x, then coated with Poly-L-Lysine for 15 min. Cells were seeded onto coated coverslips (18 mm2, Corning; Poly-L-Lysine, Sigma #RNBC8085). Cultures were grown to 40% confluency and transfected using Lipofectamine3000 (Invitrogen #L3000-008) following manufacturer’s instructions. Transfected plasmids include: WNT5A, WNT5AC83S, WNT5AC182R or DVL1, DVL11519∆T, DVL11529∆G, DVL11615∆A (pcDNA3.2), at 2.5 µg. Cells were fixed in 4% PFA    43 48h-post transfection for 20 min and then stored in PBS + 0.01% sodium azide at 4°C. Cultures were blocked in 10% normal horse serum (Sigma #H0146) with 0.2% triton-X for 1h then incubated overnight at 4°C with primary antibody (anti-Wnt5a R&D, 2 μg/ml, #AF645; anti-Flag Sigma 1:1000 #F7425). Secondary antibody was applied for 1h at room temperature (anti-goat Sigma #11C1215) and counterstained with 10 µg/ml Hoechst. Cells were imaged using a Leica SP5 confocal microscope. Three random fields of view per treatment were analyzed; assay performed 3x.  2.15 Western blot To generate cells that stably express WNT5A and WNT5AC83S proteins, chicken DF1 cells were transfected with RCAS constructs as in the initial stages of preparing virus for embryo injections, as described in section 2.4. After one month of passaging, viruses have been spread to all cells in the culture and peak expression has been reached. At 24h before collection, wtWNT5A- and WNT5AC83S infected cells along with GFP-infected control DF1 cells were treated with 1% FBS and 100 µg/ml heparin. Conditioned media was collected and centrifuged 1000 g at 4°C for 10 minutes. The supernatant was transferred to a centrifugal filter (Amicon Ultra Cell 10K #UFC901024) and concentrated at 5000 g at 4°C in a fixed rotor for 95 minutes. To prepare cell lysates, the same cultures from which conditioned media was collected were washed 2x in cold PBS then cell lysate buffer was added (RIPA buffer with SDS containing mini protease inhibitor cocktail, Roche #04693124001) and phosphatase inhibitor cocktail (phosSTOP, Roche #04906845001). Lysed cells were removed with a cell scraper (Corning, 3010) then transferred to a 1.5 ml Eppendorf tube and held on ice for 15 minutes. Cell lysates were spun at 14000 g at 4°C for 15 minutes and the supernatants were collected and stored at -20°C. For conditioned media, n=3 blots. Cell lysate samples were mixed with sample buffer at 1.25 mg/ml with 13.33% β-Mercaptoethanol (BDH #UN2966). A total of 40 μg protein was added per lane for the lysate and 90 μg for conditioned media. Samples were resolved on SDS-10% acrylamide gels and wet-transferred (50V for 90 min) to 0.45-micron nitrocellulose membrane (ThermoFisher #88018). Membranes were incubated for 1 h in blocking solution (according to R&D protocol) and primary antibodies incubated overnight at 4°C in 0.5% bovine serum albumin (BSA) in blotting buffer (R&D blotting buffer group 8; anti-Wnt5a R&D, 2 μg/ml, #AF645; anti-Flag Sigma 1:1000    44 #F7425; anti-GAG DSHB 1:50 #AMV-3C2; anti-GAPDH Thermo Fisher 1:1000 #AM4300). Licor secondary antibodies were incubated at 1:10,000 for 1h at room temperature. Membranes were scanned using near-infrared Licor scanner (Odyssey). For cell lysate, n=5 blots. 2.16 Statistical analysis All statistical analyses for cell density, BrdU, Golgi orientation, cell shape, limb morphometrics, were done using one-way analysis of variance (ANOVA), followed by Tukey’s or Fisher’s least significant difference (LSD) post-hoc test for multiple comparisons. Statistica software version 6.0 was used. For luciferase assays, data were normalized to control Empty pcDNA3.2 plasmid values and were analyzed by one-way ANOVA, followed by Tukey’s post hoc test using Statistica. For densitometry, software from the Licor scanner was used and values were normalized to GAPDH for the cell lysate. The media values were normalized to 2 non-specific bands that were present in all samples (n=3 blots).      45 3. Investigations of WNT5A variants that cause Robinow Syndrome 3.1 Introduction In this study we explore the connection between WNT signaling and a human disease, dominant Robinow syndrome (RS). There are eight genes that are linked to RS (WNT5A, ROR2, FZD2, DVL1, DVL3, NXN, RAC3, GPC4) but we focused on WNT5A since it triggers the WNT pathway and there have been detailed studies on the expression of WNT5A as well the function of the wild-type gene in several animal models. In the developing limb bud, prior to skeletogenesis, Wnt5a RNA is expressed in a gradient with greater expression at the distal-most end of the limb bud decreasing proximally (seen in both mouse and chick embryos) (Baranski et al., 2000; Gros et al., 2010). As well, WNT5A protein is also in a distal to proximal gradient as shown in western blots from different regions of the mouse limb (Gao et al., 2018). At later stages WNT5A expression is in the perichondrium, surrounding the diaphysis of long bones (Hartmann and Tabin, 2000). In the chicken mandible, WNT5A expression is highest in Meckel’s cartilage at stage HH29 and decreases by stage HH35 (Hosseini-Farahabadi et al., 2013).  Germline knockouts of Wnt5a have major shortening of the body axis, appendicular skeleton and jaws (Ho et al., 2012; Wang et al., 2011; Yamaguchi et al., 1999; Yang et al., 2003). Conditional overexpression of Wnt5a also causes moderate shortening of the long bones, absence of distal digits (Hartmann and Tabin, 2000; Yamaguchi et al., 1999; Yang et al., 2003) and a delay in ossification in both intramembranous and endochondral bones (van Amerongen et al., 2012). The limb defects in Wnt5a-/- mouse were due to decreased/delayed expression of genes that are involved in chondrocyte differentiation and ossification (Gao et al., 2018; Gao et al., 2011; Yamaguchi et al., 1999; Yang, 2003), decreased cell proliferation (Kuss et al., 2014; Yamaguchi et al., 1999; Yang, 2003), and altered cell orientation (Gao et al., 2018; Gao et al., 2011; Gros et al., 2010; Kuss et al., 2014; Qian et al., 2007). Furthermore, Wnt5a-/- mice have truncating defects that disrupt all outgrowing structures including the face, ears, limbs, genitals, and rostro-caudal axis of the body (Yamaguchi et al., 1999). These phenotypes are not consistent with dominant RS as RS patients do not have a shortened body axis and the severity of the limb phenotypes is more extreme in the mouse than that of RS (Mazzeu et al., 2007). For instance, patients with dominant RS have limb mesomelia; whereas Wnt5a-/- mice have shortening in all skeletal elements of the limbs in addition to lacking digits. As well, knockout mice have other facial phenotypes including    46 truncated tongue and reduced outgrowth of the external ears (Yamaguchi et al., 1999). Thus, Wnt5a knockout mice are not beneficial to understand the functional impact of WNT5A missense mutations in dominant RS. Using retroviral misexpression of Gallus WNT5A in chicken limbs showed a general shortening of posterior skeletal elements in the autopod and zeugopod (Hartmann and Tabin, 2000; Kawakami et al., 1999). Thus, gain or loss of function of WNT5A has profound effects on skeletal development, some of which resemble those caused by dominant RS mutations. WNT5A is thought to signal primarily via the JNK-PCP pathway (Kikuchi et al., 2011, 2012), which mediates tissue polarity (Ho et al., 2012). The JNK-PCP pathway in vertebrates involves the participation of membrane proteins such as VANGL, as well as intracellular proteins including PRICKLE and DVL (Yang and Mlodzik, 2015). The loss of PCP genes including Vangl2 or Prickle causes chondrocytes to lose their orientation and causes similar skeletal phenotypes to the knockout of Wnt5a (Gao et al., 2018; Gao et al., 2011; Liu et al., 2014; Wang et al., 2011; Yang et al., 2013). Little is known about the domain structure of WNT5A, so it is not clear what effect the missense mutations will have on protein function. The defects caused by missense mutations in human WNT5A have only been studied twice. The C83S and C182R variants were studied in Xenopus and zebrafish embryos by others, but only in very restricted studies targeted at gastrulation stages (Person et al., 2010). We expanded the present study to explore the effects on limb and mandible development in the chicken embryo. Limb mesenchyme is derived from lateral plate mesoderm as opposed to the facial mesenchyme which is largely neural crest cell derived (Creuzet et al., 2005). Furthermore, in the limb, bones form by endochondral ossification whereas in the mandible cartilages persist to adulthood (Eames and Helms, 2004). Here we use the chicken model to misexpress/overexpress wild-type or variant human WNT5A in the limb bud over top of the Gallus expression. Using this strategy, we can detect dominant effects of the mutation. In addition, comparison to the wtWNT5A gene will allow us to distinguish general effects of raising the level of WNT5A from those of the mutation. The sequence homology between human and chicken WNT5A is 91% identical at the protein level. Importantly, all 24 cysteine residues are conserved between species. Therefore, the human protein in wild-type and mutated forms is expected to be functionally equivalent to the chicken orthologues.     47 3.2 Results 3.2.1 Mutant WNT5AC83S variant shortens the long bone and delay ossification  To investigate the effects of WNT5A mutations on limb development, we compared misexpression of wtWNT5A to the effects of two variants of WNT5A that cause dominant RS (C83S and C182R). Embryos were injected at stage 15 in order to capture the lineage that will give rise to chondrocytes (Pearse et al., 2007). Later injections would have resulted in limited infection of sub compartments of the limb with lower levels of infection in chondrocyte progenitor cells.  Results of the experimental viruses were compared to the GFP control virus which does not affect development. Animals were grown until osteogenesis had taken place (stage HH38) and processed for whole-mount bone and cartilage staining using Alizarian red and Alcian blue, respectively. This staining procedure is not compatible with visualizing live GFP expression, however at the time of fixation all GFP infected limbs had fluorescence (data not shown). Variant C83S shows a shorter forelimb compared to wtWNT5A or WNT5AC182R (Figure 3.1, Table 3.1). The majority of the specimens injected with the C83S virus qualitatively affected all skeletal elements including delayed or absent ossification in some bones (Figure 3.1 C).        48  Figure 3.1. Skeletal phenotypes obtained from misexpression of WNT5A retroviruses in the forelimb. Embryos injected with RCAS::GFP, RCAS::wtWNT5A, RCAS::WNT5AC83S, or RCAS::WNT5AC182R into the region of the developing forelimb at stage HH15 (see inset in A), and fixed 12 days later at stage HH38 showing lateral view. (A) Control GFP forelimb. (B) Embryo injected with RCAS::wtWNT5A. Radius is slightly affected in length compared to GFP control (arrow). (C) Forelimb injected with RCAS::WNT5AC83S shows a shortened forelimb, much thicker diameter cartilage in the humerus, radius and ulna (arrows). There is delayed ossification as shown by the lack of Alizarin red stain. (D) RCAS::WNT5AC182R injected forelimb show only slight decrease in the overall forelimb. (E,F) Quantification of bone length and AP diameter from photographs. Wild-type and C83S bones are shorter and have increased AP diameter compared to GFP control limbs. Only C182R variant was shorter in the radius length. Scale bar=5 mm. Key: h – humerus, r – radius, u – ulna. Dark blue/purple = Alizarin red (stains for bone), light blue = Alician blue (stains cartilage).         49 Table 3.1. Qualitative analysis of WNT5A forelimb phenotype at stage HH38. Biological replicates and detailed phenotype analysis of embryos injected with viruses into prospective limb region. Limbs can be both short/wide and can have a phenotype affected the stylopod, zeugopod, or autopod.Genotype Normal Stylopod and/or zeugopod abnormalities1 Delayed or absent ossification in any bone RCAS::GFP (n=22) 22 100% 0 0 RCAS::wtWNT5A (n=12) 2 17% 10 83% 0 RCAS::WNT5AC83S (n=17) 0 17 100% 16 94% RCAS::WNT5AC182R (n=7) 4 57% 3 42% 0 1abnormalities refer to thicker or shorter skeletal element  3.2.2 Mutant WNT5A shortens and widens dimensions of the developing long bone  For the remainder of our analysis, we turned to histological sections rather than wholemount stained embryos. The main reasons are that we wanted to know exactly which cells had been infected by the virus so that we could correlate molecular changes with viral expression. We fixed embryos at several stages from 4 to 10 days post-injection, by the time when endochondral ossification had begun, and the full pattern of the limb was present (stage HH36). We used an antibody to the viral coat, GAG, and found that the viruses had spread throughout the stylopod and zeugopod. However, at earlier stages, the autopod was not always infected so we focused our analysis on the zeugopod (Figure 3.2 A’-D’; Figure 3.3 A-H insets). The morphology of the wtWNT5A limbs was indistinguishable from GFP controls in terms of length and AP width at stage HH36 (Figure 3.2 A,B,E,F, n=4 for each virus). Also, in adjacent sections stained for the chondrocyte hypertrophy marker, COL10A1, there was similar staining in controls and limbs treated with wtWNT5A virus (Figure 3.2 A”. B”). Contrastingly, there was an absence of staining for COL10A1 in WNT5AC83S infected limbs (Figure 3.2 C’’). The WNT5AC182R virus infected limbs had very light staining for COL10A1 (Figure 3.2 D’’) suggesting a lesser effect on hypertrophy. Also, viruses expressing mutant human WNT5AC83S or WNT5AC182R resulted in shorter limb elements particularly affecting the stylopod (humerus) and zeugopod (radius and ulna) (Figure 3.2 C,D-F). The degree of PD shortening was far more severe in the C83S variant (Figure 3.2 E).  We wanted to test whether width was increased in a DV axis, so we examined injected forelimbs    50 at stage 36 in the perpendicular axis (cut transversely). In transverse sections, the diameter of the epiphysis and diaphysis (measured from outermost edge of the bone collar if present) was significantly increased by both C83S and C182R variants (Figure 3.3 C,D,G,H,I). We also measured the cartilage and bone thickness separately. The DV diameter of the cartilage was increased for all WNT5A viruses while the bone collar was decreased only for wtWNT5A and the C83S variant (Figure 3.3 J). However, unlike wtWNT5A virus, the WNT5AC83S virus reduction in bone collar thickness was correlated with a lack of hypertrophy (Figure 3.2 C”). The skeletal muscles were present in their normal positions but were proportionately shorter and displaced, especially in the C83S infected limbs (Figure 3.2 C,D; Figure 3.3 G,H). Since the changes in cartilage morphology caused by mutant viruses, particularly the C83S variant, are different than wtWNT5A, it appears that the variants caused dominant, neomorphic functional changes that are not simply explained by generally higher levels of WNT5A gene expression.     51     52 Figure 3.2. Mutant WNT5A shortens and widens dimensions of the developing long bone. Embryos were injected into the presumptive limb field at stage 15 (inset in A) and fixed at stage 36. Sagittal sections of injected forelimbs were stained with Alcian blue and Picrosirius red (A-D) and adjacent sections were stained with anti-GAG to locate the virus in the tissues (A’-D’), or with anti-COL10A1 to mark hypertrophic chondrocytes (A”-D”). (A) GFP virus has no effect on development. The onset of endochondral bone formation in normal limbs is seen first in the diaphysis with larger chondrocytes (a and A” white arrow). Muscles are differentiated and attach to perichondrium. (B) wtWNT5A virus has minimal effects on limb development with normal muscle development and show larger chondrocytes in (b), and normal COL10A1 staining (B”, white arrow). (C) The C83S variant virus causes significantly shorter and wider cartilages to develop with smaller chondrocytes (b) and lacked COL10A1 staining (C”). The limb musculature is present but displaced and shorter than normal. (D) The C182R virus has caused shortening of the bones and thickening in diameter. The muscles seem thinner than normal, displaced by the larger cartilage elements. Larger chondrocytes are present (d) and COL10A1 is weakly expressed (D”, white arrows). The dimensions of the ulna were measured: (E) length and (F) diameter. Mutant WNT5A variants showed significantly shorter length (E) and wider diameter (F) of the developing ulna as compared to the wtWNT5A virus. Diagram indicates measurements along the hatched lines in the ulna. One-way ANOVA, Tukey’s post hoc test, *p<0.05, ***p<0.001, n=4. Scale bar=1 mm. Lower case letters represent inset of diaphysis. Key: anc – anconeus, edc – extensor digitorum communis, eil – extensor indicis longus, emr – extensor metacarpi radialis, emu – extensor metacarpi ulnaris, h – humerus, r – radius, u – ulna.     53       54 Figure 3.3. WNT5A viruses widen the ulna in chicken embryos. (A-H) Serial transverse sections of the injected forelimbs fixed 10 days post-injection (stage 36). Insets show viral (anti-GAG) expression. The width of the ulna bone was measured in the epiphysis and diaphysis (see double-headed arrows in A and E). (A-D) Transverse sections of epiphysis of the forelimb. WNT5AC83S and WNT5AC182R injected forelimbs were significantly wider in their epiphyses compared to both GFP control and wtWNT5A viruses. (E-H) Transverse sections of the diaphysis of the forelimb. wtWNT5A and mutant WNT5A variants were significantly wider in the diaphysis of the ulna compared to GFP control virus. (I) Ulna width measuring the epiphysis and diaphysis, diagram represents the transverse section through the forelimb. (J) Width of cartilage and bone collar located in the diaphysis, diagrams represent region measured. Wild-type and mutant WNT5A injected forelimbs showed significantly more cartilage in the diaphysis, and GFP controls along with WNT5AC182R had a thicker bone collar. One-way ANOVA followed by Tukey post hoc test. *p<0.05, **p<0.01, ***p<0.001; n=4 for all viruses. Scale bar=500 µm and applies to all brightfield images. Key: An – Anterior, D – Dorsal, dia – diaphysis, e – epiphysis, edc – extensor digitorum communis, eil – extensor indicis longus, emr – extensor metacarpi, fdp – flexor digitorum profundus, fcu – flexor carpi ulnaris, h – humerus, Po – Posterior, r – radius, u – ulna, V – ventral.  3.2.3 WNT5AC83S virus showed increased cell density in the diaphysis of the ulna The chondrocytes appeared to be smaller and more numerous in the WNT5AC83S infected cartilage (as shown in Figure 3.2 C). DAPI stained nuclei were quantified in the epiphysis and diaphysis of stage HH36 forelimbs. There was no change in cell density in the epiphysis of the developing long bones between controls and treated. However, there was increased cell density in the diaphysis of WNTAC83S (Figure 3.4 C’, E) as compared to the control GFP (Figure 3.4 A’,E), wtWNT5A (Figure 3.4 B’,E) or the C182R variant (Figure 3.4 D’, E). We also quantified cell density in half of the ulna (diaphysis to epiphysis closest to the digits). Interestingly, the total number of chondrocytes over a defined area was not different between mutant (C83S or C182R) expressing limbs compared to wtWNT5A or GFP controls (Figure 3.4 F). Due to the fact that the C83S variant gave rise to a more penetrant phenotype compared to GFP control and wtWNT5A viruses we pursued this mutation in more detail in subsequent analyses.    55       56 Figure 3.4. Variant WNT5A viruses showed increased cell density in both the diaphysis and epiphysis of the ulna. (A-D) Sagittal sections of injected forelimbs fixed 10 days post-injection (stage 36) and stained with DAPI for nuclei. Cell density was quantified in the same area size in the diaphysis (A’-D’) and epiphysis (A”-D”) of the ulna. (D,E) WNT5AC182R showed increased cell density in the epiphysis, which was significantly different than GFP and wtWNT5A. (C,E) There was a significant increase in cell density in diaphysis of WNT5AC83S injected limbs compared to all other treatments. One-way ANOVA post hoc Tukey, *p<0.05 compared to GFP control, n=4. (F) Cell count was quantified in half of the ulna bone closest to the digits. WNT5AC83S
 and WNT5AC182R injected limbs showed no significant difference in cell count compared to GFP control or wtWNT5A injected limbs (n=4). Diagram represents ulna and the area region indicates the region quantified. Scale bar=500 µm. Key: h – humerus, r – radius, u – ulna.   3.2.4 Phenotypic changes are first visible 4 days post-infection with RCAS viruses  To determine when the phenotypes first appeared, we looked at earlier stages of development, just after chondrogenesis has taken place. We measured the ulna at 4 and 5-days post-injection, (stage HH29, 30) (Figure 3.5). Initially, there is no difference in length of the cartilage condensations at stage 29. However, the AP diameter of the cartilage was significantly wider for the wtWNT5A compared to the other conditions (Figure 3.5 B, p<0.05). By stage 30, the wtWNT5A and C83S variant caused a significant shortening of the cartilage compared to GFP controls (Figure 3.5 A). The AP diameter was significantly increased by the C83S variant (Figure 3.5 B) similar to stage 36 embryos (Figure 3.2 F).     Figure 3.5. Phenotypic changes are first visible 4-5 days post infection with WNT5AC83S. The ulna was measured on sections used in Figure 3.7 (stage HH29) and Figure 3.8 (stage HH30) (For both stages n=4 GFP; n=5 wtWNT5A and WNT5AC83S). (A) The length of the ulna was significantly shorter in the treated limbs at stage HH30. There was no difference in the length between wtWNT5A and WNT5AC83S. (B) The AP diameter of the ulna was significantly wider at stage HH29 in wtWNT5A injected limbs. However, at stage HH30 there was already a large increase in diameter produced by the WNT5AC83S virus. *p<0.05, ***p<0.001 compared to GFP control.    57  3.2.5 Volume of cartilage is not changed by the RCAS::WNT5AC83S  We looked at the initial stages of chondrogenesis to investigate the earliest changes in the condensations. The shape changes first become apparent at stage 29 which is 96h post-infection. Using microCT scanning with contrast enhancement, segmented skeletal elements displayed similar decreases in length and increases in width to that seen in histology (Figure 3.6 A-C). We found that there was no difference in total volume of the different skeletal elements, humerus, ulna, and radius (Figure 3.6 D), which correlates with the overall cell number not being different at later stages.    Figure 3.6. Embryo forelimb skeletal elements show similar volume four days post-injection. Embryos were injected with viruses into the right developing limb bud at stage 15 and fixed four days post-injection (stage HH29). (A-C) Segmented volumes of the humerus (blue), ulna (red), and radius (yellow) were created in Amira (FEI) for each specimen (n=3); entire forelimb is green. (D) No changed in volume were detected with each skeletal element.    58 3.2.6 wtWNT5A and WNT5AC83S show increased cell proliferation in the cartilage At stage HH29, the normal pattern of proliferation consists of slightly higher proliferation in the future epiphyses compared to the diaphysis (Figure 3.7 B). There was no significant difference in the proliferation index at stage HH29 between the different viral treatments (Figure 3.7 B,D,F,G). However, at 5 days post-injection (stage HH30), proliferation was 5 times higher in the diaphysis region in the wtWNT5A injected embryos and 2.5 times higher in the WNT5AC83S virus infected limbs compared to the GFP control (Figure 3.8 B,D,F,G). Wild-type WNT5A had significantly higher proliferation in the epiphysis compared to WNT5AC83S (Figure 3.8 G). Only GFP virus showed increased proliferation in the epiphysis compared to its diaphysis, whereas the wild-type and variant WNT5A showed an evenness of cell proliferation throughout the cartilage (Figure 3.8 B,D,F,G). The increased proliferation in the wtWNT5A was correlated with a relatively higher cell density in these cartilage elements at stage HH30. Thus, the extra cells were packed more tightly in a shorter cartilage rod (data not shown). This increased density caused by wtWNT5A was transient; however, since by stage HH36 we had shown the diaphysis and epiphysis had similar densities to GFP controls. The viruses did not affect the level of apoptosis (TUNEL at HH29 or 30; Figure 3.9). We can therefore conclude that higher levels of WNT5A do not induce stress in the cells that would lead to cell death. Thus, continuing proliferation in the diaphysis partially accounts for the increased diameter of the diaphysis. The mutation C83S may be hypomorphic relative to wtWNT5A since proliferation was stimulated to a lesser degree (Figure 3.8 G). Nevertheless, the unique combination of delayed hypertrophy plus increased proliferation caused by WNT5AC83S virus could account for the increase in diameter of the diaphysis.      59  Figure 3.7. wtWNT5A or WNT5AC83S injected forelimbs do not affect cell proliferation at stage HH29 in the developing ulna. Embryos were injected with viruses into the right limb bud at stage HH15 and fixed four days post-injection. (A,C,E) Sagittal sections of the limbs show viral (anti-GAG) expression (green), and (B,D,F) neighboring sections probed for anti-BrdU labeling (green) and anti-SOX9 expression (red). There were very few BrdU positive cells in the centre of the ulna or the future diaphysis. (G) BrdU positive cells in the entire ulna were quantified and normalized to control GFP virus. No change in cell proliferation was detected. One-way ANOVA, Fisher’s LSD post hoc test, n=4 GFP; n=5 wtWNT5A and WNT5AC83S. Scale bar=500 µm, Key: r – radius, u – ulna.    60  Figure 3.8. wtWNT5A or WNT5AC83S injected forelimbs have increased cell proliferation in the cartilage at stage HH30. (A,C,E) Sagittal sections of injected forelimbs fixed 5 days post-injection show viral (anti-GAG) expression (green), and (B,D,F) neighboring sections probed for anti-BrdU labeling. Note that the normal limbs diaphysis is devoid of proliferating cells (B). In contrast there are labeled cells in the wtWNT5A or WNT5AC83S injected ulna diaphysis (D,F). (G) Percentage of BrdU positive cells were similar in the epiphysis regardless of treatment. In the diaphysis, wtWNT5A and WNT5AC83S showed significantly increased cell proliferation compared to GFP. One-way ANOVA and Fisher’s LSD post hoc test, **p<0.01, ***p<0.001; n=4 GFP; n=5 wtWNT5A and WNT5AC83S. Scale bar=500 µm. Key: dia – diaphysis, epi – epiphysis, r – radius, u – ulna.    61   Figure 3.9. wtWNT5A or WNT5AC83S injected forelimbs do not affect apoptosis in the cartilage. Apoptotic cell death by TUNEL assay was performed at 4- and 5- days post injection (stage HH29 (A-C’), stage HH30 (D-F’)). Apoptotic cells are marked as light green dots, and no difference was detected between the different treatment groups. n=4, scale bars=100 µm Key: r – radius, u – ulna.  3.2.7 WNT5AC83S disrupts chondrocyte polarity and cell elongation in forelimbs  The polarity of chondrocytes is intimately connected to the growth and elongation of cartilage rods. We used Golgi as a marker for cell polarity as Golgi complexes in chondrocytes are found oriented perpendicular to the long axis of the developing cartilage (Hosseini-Farahabadi et al., 2017; Kuss et al., 2014). The angle between the Golgi-nucleus axis and the long axis of the bones in the zeugopod cartilage was measured at stage HH29 when the phenotypes were first apparent. Here, we found that in Alkaline phosphatase (AlkPO4) controls Golgi are oriented perpendicular to the long axis of the developing cartilage (65.28° ± 3.2°) (Figure 3.10 A,D). Overexpression of wtWNT5A was not significantly different from our control forelimbs (Figure 3.10 B,E); however, WNT5AC83S mutation was significantly more randomized compared to AlkPO4 controls (47.7° ± 0.7°, p<0.005) (Figure 3.10 C,F). Using phalloidin staining to mark the chondrocyte cell membranes, we found that AlkPO4 virus infection did not change the typical flattened and elongated shape of the cells (Figure 3.10 G). In contrast, chondrocytes were rounder in both    62 RCAS::wtWNT5A and RCAS::WNT5AC83S infected limbs (p=0.01 for both viruses) (Figure 3.10 G).  Since we observed several signs that polarity was disrupted, we next wanted to see if one of the core PCP molecules was similarly redistributed. Prickle1 is concentrated on opposite ends of the flattened chondrocyte cells (Kuss et al., 2014). We found that in the developing ulna at stage HH29, AlkPO4 control and wtWNT5A limbs showed pan-Prickle molecules concentrated on opposite ends of the developing chondrocytes (Figure 3.11 A-A”), whereas in WNT5AC83S injected limbs the pan-Prickle expression showed diffuse distribution (Figure 3.11 B-B”,C-C”). Taken together, the cell morphology and polarity analyses provide additional evidence for the C83S variant causing a gain of behaviour, the same as wtWNT5A.       63  Figure 3.10. WNT5AC83S virus randomizes chondrocyte polarity and causes rounder chondrocyte shape in the forelimb. (A-C) Sagittal slices (200 µm) of injected forelimbs fixed 4 days post-injection (stage HH29) and stained in wholemount with anti-Golgi (GM130, pink) and actin filaments with Phalloidin-568. Expression in chondrocytes (SOX9-positive) was identified in adjacent sections (not shown). The angle between the Golgi-nucleus axis and the long axis of the bones in the ulna was measured (inset in A). (D-F) Graphical representation of angular data for each virus type (70 cells measured/specimen). Blue line is the average of each specimen, red line is the overall mean for the group and green lines represent the standard deviation. Colored blocks (1, 2 and 3) represent the specimens. Numbers 10, 20 and 30 indicate the number of cells at that particular angle. The cells were more randomly oriented with WNT5AC83S compared to AlkPO4 control virus (One-way ANOVA and Tukey’s post hoc test, p<0.005; n=3). (G) Chondrocyte shape was analyzed using Phalloidin stained cell membranes (from A-C). Feret diameter was measured in perpendicular axes in 50 cells/specimen (length/width). Significantly rounder cells were observed in both wtWNT5A and WNT5AC83S viruses (closer to 1) compared to AlkPO4. One-way ANOVA post hoc Tukey **p<0.01, n=4. Scale bar=50 µm. Key: L – length, W – width.     64  Figure 3.11. WNT5AC83S virus randomizes chondrocyte PCP molecule, Prickle in the forelimb. (A-C) Sagittal sections of injected forelimbs fixed 4 days post-injection (stage HH29) show staining for pan-Prickle and counterstained with Hoechst for nuclei; viral expression (GAG) shown in the inset. (A’-C”) Increased magnification of box region in the zeugopod cartilage of the forelimb showing combined signal for nuclei and prickle as well as the prickle stain alone. (A’,A”) Double arrows indicate Prickle distribution at the ends of the elongated chondrocytes in the control forelimbs. (B’,B”) The wtWNT5A has not affected Prickle distribution; however (C’,C”) WNT5AC83S caused a general reduction in staining and a more diffuse pattern. n=4. Scale bar=50 µm. Key: Ant – anterior, Di – distal, Pos – posterior, Pr – proximal.  3.2.8 WNT5A mutations cause shortening of the mandible  We also studied both WNT5A mutations (C83S and C182R) in the face, targeting the mandible (lower jaw). The overexpression of wtWNT5A caused mandibular hypoplasia on the injected side (6 of 8 embryos) as reported (Table 3.2) (Hosseini-Farahabadi et al., 2013). The mutant WNT5A viruses also caused shortening of the lower jaw; the C83S mutation has a more penetrant phenotype than the C182R mutation (C83S, 11 of 13; C182R, 5 of 12; Table 3.2). The wtWNT5A- and WNT5AC83S-infected mandibles were 20% shorter than GFP controls (Figure 3.12 E). Although    65 the wild-type and mutant forms of WNT5A gave similar phenotypes (Figure 3.12 B-D), we predicted that there might be differences at the molecular/cellular level.    Figure 3.12. Skeletal phenotypes obtained from misexpression of WNT5A retroviruses in the mandible. Embryos injected with RCAS::GFP, RCAS::wtWNT5A, RCAS::WNT5AC182R, or RCAS::WNT5AC83S into the mandibular arch at stage 15 (inset, A with GFP viral expression in the mandible) and fixed 12 days post-injection at stage 38. (A–D) Dorsal views of lower beaks stained with alizarin red and alcian blue reveal that many skeletal elements were reduced in size in the treated embryos (B–D). (E) Mandibular length measured on the injected side at stage 38 with the Alcian blue stain, staining Meckel’s cartilage. Arrowheads point to shortening of injected side of mandible (B,C,D). Significant shortening was caused by the 3 WNT5A viruses (**p<0.01). Tukey’s post hoc test showed that the C83S mutation caused significant shortening of the right mandible compared to the C182R mutation. Scale bar=5 mm, key: a – angular bone (orange), d – dentary bone (yellow), mc – Meckel’s cartilage (blue), s – splenial bone (green), sa – surangular bone (red).       66 Table 3.2. Qualitative analysis of mandibular phenotype. Virus type Severity1 Bones affected Normal Mild Severe Angular Surangular Dentary Splenial Meckel’s Cartilage GFP (n = 11) 100% (11) 0% 0% 0% 0% 0% 0% 0% wtWNT5A (n = 8) 12.5% (1) 12.5% (1) 75% (6) 75% (6) 87.5% (7) 75% (6) 62.5% (5) 87.5% (7) WNT5AC182R (n = 12) 41.6% (5) 16.6% (2) 41.6% (5) 41.6% (5) 41.6% (5) 41.6% (5) 41.6% (5) 50% (6) WNT5AC83S (n = 13) 0% 15.4% (2) 85% (11) 85% (11) 100% (13) 79% (10) 85% (11) 100% (13) 1Mandibular phenotype severity of specimens grouped into three categories: normal, mild, or severe. The degree of severity is determined by shortness or loss of mandibular bones: angular, surangular, dentary, splenial, Meckel’s cartilage (see Figure 3.12 for bone and cartilage location in the mandible). If ≥ 2 bones are short/missing, then the specimen is considered severe.   3.2.9 WNT5A mutations do not affect cell proliferation in the mandible or in Meckel’s cartilage We measured cell proliferation in the mandible at two stages, when Meckel’s cartilage is condensing (stage 28) and when it is actively elongating (stage 29) (Figure 3.13 A-J). At stage 28, proliferating cells were quantified in half the developing mandible (where viral spread was located). At stage 29, proliferating cells were quantified in Meckel’s cartilage (see hatched lines in Figure 3.13 F,G,H,I). We found no detectable difference in cellular dynamics at stage HH28 and 29 despite high levels of expression of the virus. In wholemount preparations, we did not see a wider diameter of the cartilage rod in the mandible like we did in the limb. Therefore, it is likely that even at later stages, we would not have detected any proliferation effects from the WNT5A viruses.    67     68 Figure 3.13. WNT5A variants do not affect cell proliferation in the mandible. Frontal sections of mandibles injected with RCAS::GFP, RCAS::wtWNT5A, RCAS::WNT5AC182R, and RCAS::WNT5AC83S viruses at stage HH15. The first column are sections from stage 28 embryos (3 days post-injection) stained for proliferation (BrdU, A-D). There was no significant difference in number or distribution of the percentage of proliferating cells (E). The right column contains stage 29 embryos (4 days post-injection F-I). Dashed white line indicates Meckel’s cartilage. There was no change in proliferation within Meckel’s cartilage (J). Scale bar=250 μm for all sections.  3.2.10  WNT5A mutations randomize chondrocyte polarity in Meckel’s cartilage  We tested whether the shortened mandible (lower jaw) caused by the WNT5A viruses was due to changes in the polarity and shape of Meckel’s chondrocytes. To measure polarity, we used the position of the Golgi apparatus relative to the nucleus and long axis of the cartilage rod. In control embryos, the Golgi apparatus was located at 60° to the long axis of Meckel’s cartilage similar to the limb. In the wtWNT5A- injected embryos, Golgi appeared to be a bit more random compared to GFP control (Figure 3.14 E,F,I,J); similarly, the virus with the C182R mutation was not significantly different from controls (Figure 3.14 G,K). In contrast, the WNT5AC83S mutation caused significantly greater randomization when compared with the control embryos or wtWNT5A (46° ± 1.0°, p=0.025) (Figure 3.14 H,L), again similar to the limb.        69  Figure 3.14. WNT5A mutant viruses alter chondrocyte polarity in the mandible. Embryos were injected with viruses into the right mandibular arch at stage 15 and fixed 4 days post-injection at stage HH29. (A–D) Serial frontal sections of mandible with anti-GAG. Dashed lines outline Meckel’s cartilage on injected side. (E–H) Localized staining for Golgi bodies (anti-GM130) was observed in each chondrocyte. (I–L) The angle between the Golgi-nucleus axis and the long axis of Meckel’s cartilage was measured (see inset in E). Graphical representation of angular data for the 4 biological replicates for each virus type. Red line is the overall mean for the group; individual means for each specimen are shown in blue lines. The curved orange lines represent ±1 SD for each biological replicate. (E, I) In GFP controls, the Golgi-nucleus axis is oriented 60° ± 2.82° to the cartilage rod. (F, J) In wild-type WNT5A, the average angle is not significantly different from GFP controls; however, there is a high degree of variability. (G, K) In WNT5AC182R-injected embryos, the Golgi apparatus tended to be more randomized, but this was not significant when compared with GFP controls. (H, L) In WNT5AC83S, the cells were significantly more randomly oriented (46° ± 1.0; P = 0.025). Scale bars=500 µm for A–D; 25 µm for E–H.  3.2.11 WNT5A mutations disrupt chondrocyte cell shape in Meckel’s cartilage To measure cell roundness, we stained the chondrocytes with non-muscle myosin II antibody. In wtWNT5A, cells were elongated, similar to GFP controls (p=0.122 (Figure 3.15 A,B,E)). Mutated versions of WNT5A caused chondrocytes to be rounder than GFP controls (C182R, p=0.006; C83S, p=0.0009 (Figure 3.15 C-E)). In addition, WNT5AC83S produced significantly rounder chondrocytes as compared with wtWNT5A (p=0.03). Therefore, both WNT5A mutations are interfering with proper stacking and flattening of chondrocytes within Meckel’s cartilage, which potentially causes shortening of mandible.      70  Figure 3.15. WNT5A mutant viruses alter chondrocyte shape in Meckel’s cartilage. Embryos were injected with viruses into the right mandibular arch at stage 15 and fixed 4 days post-injection at stage 29. Serial frontal sections were used for anti-non-muscle myosin II and SOX9. (A-D) Chondrocyte shape was analyzed by staining the overall cytoskeleton with non-muscle myosin II and SOX9 to identify chondrocytes. Feret diameter was measured in 75 cells (length/width, inset A; n=3 GFP, wtWNT5A, WNT5AC182R; n=4 WNT5AC83S). (E) Significantly rounder cells were observed in both C182R and C83S viruses as compared with GFP or wtWNT5A (**p<0.01, ***p<0.001). GFP control cells were clearly oval in shape when compared with the mutant WNT5A viruses (C,D). Scale bar=25 µm.  3.2.12 WNT5AC83S is secreted much less efficiently than wtWNT5A  We wanted to address whether the mutation in WNT5A altered synthesis or secretion of the protein. Since the phenotypes from the C83S were more consistent and more severe than for the C182R variant, we focused our protein expression work on the C83S variant. We first examined cell lysate from chicken fibroblasts infected with C-terminal, Flag-tagged WNT5A and WNT5AC83S RCAS retroviruses. The virus results in a long-term stable incorporation of the gene without the need for antibiotic selection. We detected a 40% reduction in the levels of WNT5AC83S compared to wtWNT5A using WNT5A antibody (Figure 3.16 A,B). We compared viral levels across the cultures by probing with the anti-GAG antibody normalized to GAPDH (Figure 3.16 C). Levels of viral expression were equal in all cultures, therefore the lower level of WNT5A protein in the C83S cell lysate represented a true difference. We wanted to make sure that the missense mutation in our WNT5A variant had not prevented the WNT5A antibody from recognizing the protein on the blot, so we also used Flag antibody. The same results of lower levels in the lysate were detected with the Flag antibody. Also, the band migrated at the same size as on blots probed with the    71 WNT5A antibody (Figure 3.17 A). Next, we collected conditioned media from mutant and wtWNT5A expressing cultures. To ensure we could detect even small amounts of protein, the conditioned media was concentrated 600-fold prior to loading on the gel. Significantly reduced levels of mutant WNT5A were also observed in conditioned media (20% of wtWNT5A; Figure 3.17 B,C). We were surprised to see less protein in both the conditioned media and in the lysate. If secretion was less efficient, we expected to see increased levels of protein retained in the secretory pathway and these should show up as higher levels in the lysate. The lower levels of WNT5A in the lysate and media could mean that in vivo infection led to slightly lower levels of WNT5A in the mutant infected limbs. Viral sequences were confirmed for all constructs using internal and external primers with the sequencing primer in the RCAS 5’ insert.    Figure 3.16. WNT5AC83S shows less protein in the cell lysate than wtWNT5A. Western blot analysis of WNT5A protein. DF1 (chicken fibroblast) cells were transfected with RCAS retroviruses (GFP, WNT5A or WNT5AC83S) for 1 month and cell lysate collected. Blots were probed with anti-human WNT5A polyclonal antibody, anti-GAPDH and anti-GAG. (A) The lysate from wtWNT5A-infected cells shows a strong band at 45 kDa, the predicted size of WNT5A protein (40 µg protein loaded per lane). The lane with WNT5AC83S variant protein shows considerably lighter staining. (B) Densitometry readings were made for 3 blots and normalized to GAPDH. Significantly lower values for WNT5AC83S suggests that protein may be synthesized at lower efficiency. (C) Viral (GAG) levels normalized by GAPDH. Students T-test, *p < 0.05, ***p<0.001.    72  Figure 3.17. WNT5AC83S is secreted much less efficiently than wtWNT5A. Western blot analysis of FLAG protein. DF1 (chicken fibroblast) cells were transfected with RCAS retroviruses (FLAG tagged-GFP, -WNT5A or -WNT5AC83S) or no virus, for 1 month. Heparin (100μg/ml) was added 24h prior to collecting conditioned media. After media was collected cells were lysed and protein extracted. Blots were probed with anti-Flag polyclonal antibody, then stripped and reprobed with antibody to GAPDH. (A) The lysate from wtWNT5A infected cells show a strong band at 45 kDa and WNT5AC83S lane shows a slightly lighter staining. Secretion of wtWNT5A into media was detected but levels of WNT5AC83S were considerably lower. (B) Conditioned media probed for anti-WNT5A shows similar bands for wild-type and mutant protein as with anti-FLAG. (C) Densitometry readings of conditioned media were made for 3 blots and normalized to non-specific bands. Students T-test, *p<0.05, ***p<0.001. CM – conditioned, concentrated media, DF1 – DF1 fibroblasts, parent cell line, GFP – cells infected with GFP virus, wt - wtWNT5A.   3.2.13 Transient transfections of WNT5A plasmids show similar transfection rates  Examination of transfected HEK293 cells with wtWNT5A and WNT5A variants were visualized 48h post-transfection after immunocytochemical labeling (Figure 3.18 A-C). Transfected cells    73 encompassed the entire cytoplasm. Wildtype WNT5A and both WNT5A variants showed similar amount of transfected cells, around 10%. Thus, we could compare readouts in other assays of WNT signaling between cells transfected with different plasmids.    Figure 3.18. WNT5A labeling of transfected cells in HEK293 cells. Cells were transfected using pCDNA3.2 plasmids and fixed 48h post-transfection. (A-C) Percent of transfected cells (green) per field of view were quantified in 5 fields, repeated 3 times. (D) No difference in transfected cells were detected. One-way ANOVA, Tukey’s post hoc.  3.2.14 WNT5A variants super activate JNK-PCP signaling  We tested whether missense variants were able to activate the JNK-PCP pathway using the ATF2 construct (Ohkawara and Niehrs, 2011). We initially performed the assay on HH24 chick forelimb micromass cultures that were transfected with the plasmids. We saw an increase in ATF2 reporter activation with wtWNT5A and the WNT5A variants (Figure 3.19 A,C), but only the C182R variant was able to significantly activate the luciferase reporter; however, there is a trend of increased activation with C83S variant (Figure 3.19 A). Next, we tested the variants using HEK293 cells and saw similar results to those obtained with micromass cultures. We found that    74 WNT5AC83S and WNT5AC182R mutant proteins derived from plasmids significantly activated ATF2 reporter activity (Figure 3.19 B). In addition, we found that the C182R mutation showed significantly more activation compared to wtWNT5A (Figure 3.19 B). These ATF2 results are compelling evidence for a molecular gain-of-function caused by the missense variants. In addition, for the second time we were able to discern differences between the C83S and C182R variants (first occurrence was in the skeletal phenotypes).  Next, we looked at whether the activation of the ATF2 reporter was facilitated by either the FZD2 or ROR2 receptors, both of which are mutated in dominant RS or recessive RS respectively (Afzal et al., 2000; White et al., 2018). As expected, providing FZD2 plasmid significantly activated the reporter due to the presence of endogenous WNT ligands (Figure 3.19 D). When we combined FZD2 with WNT5A plasmids, the results were similar to the experiments where no FZD2 was provided (compare Figure 3.19 B,D). There was no significant gain in activity by the addition of wtWNT5A or WNT5AC83S suggesting that the endogenous WNTs were the main source of ligand for FZD2. However, when WNT5AC182R plasmid was combined with FZD2, there was significant increased activity compared to FZD2 alone or in combination with wtWNT5A and the C83S variant plasmids (Figure 3.19 D). In this case, the C182R variant protein is likely to be binding to the exogenous FZD2 receptor and dimerizing with endogenous ROR2 (Figure 3.19 D’). ROR2 is thought to be one of the main receptors for WNT5A (Gao et al., 2011; Ho et al., 2012; Mikels et al., 2009; Nishita et al., 2010), so we examined both plasmids concomitantly with ATF2 reporter. Interestingly mouse Ror2 plasmid did not significantly increase activation of the ATF2 reporter (Figure 3.19 E). Instead, the presence of wtWNT5A or WNT5AC83S DNA was required (Figure 3.19 E). There was no significant difference between wild-type or mutant DNA suggesting that C83S variant does not alter binding to ROR2 receptor or recruitment of endogenous FZD co-receptor (Figure 3.19 E’).     75  Figure 3.19. WNT5A variants activate ATF2 luciferase reporter. Luciferase assays conducted on primary limb mesenchyme (A), or HEK293 cells (B,D,E). Plasmids were transiently transfected 48h prior to lysing the cells. (A,C) Micromass cultures of forelimb with wild-type and mutant WNT5A plasmids activated non-canonical JNK-PCP signaling, the most potent of the plasmids was the C182R variant. (B) HEK293 cells also responded to transfection of wtWNT5A or the two variant forms and show activation of non-canonical JNK-PCP signaling. Mutant WNT5A plasmids showed increased ATF2 activity compared to wtWNT5A. (D,D’) FZD2 receptor enhanced activity of the ATF2 reporter compared to empty plasmid. Thus, endogenous WNTs present in the HEK293 cells were able to utilize the transfected receptor. The addition of plasmids expressing WNTs further increased activity of ATF2. The C182R variant is significantly more active than the C83S or wtWNT5A. (E) No change in activity occurred with Ror2 on its own suggesting there are insufficient endogenous ligands to bind to this receptor. (E,E’) Adding WNT5A or C83S variant combined with Ror2 significantly increased activity when compared to Ror2 alone. (F) Schematic diagrams represent predicted signaling pathway reflecting the luciferase results. Unknown endogenous receptors in (C) indicated by question mark. One-way ANOVA, Tukey’s post hoc *p<0.05, **p<0.01, ***p<0.001. Key: mutWNT5A – mutant WNT5A, wtWNT5A – wild-type WNT5A, enWNT – endogenous WNT.     76 3.2.15 Mutant WNT5A is just as effective at antagonizing canonical WNT signaling as wtWNT5A WNT5A is involved in both the canonical and non-canonical WNT signaling pathways (Grumolato et al., 2010; Mikels and Nusse, 2006). In luciferase assays, WNT5A is a strong inhibitor of canonical WNT signaling stimulated by WNT3A protein or a GSK3β antagonist, LiCl (Klein and Melton, 1996; Thorne et al., 2010). Here, HEK293 cells were transfected with STF luciferase reporter which measures canonical WNT activity. The endogenous canonical WNT signaling was repressed by wtWNT5A and both variants (Fig. 3.20A). The wtWNT5A or the C83S variant DNA were equally able to antagonize the actions of LiCl (Figure 3.20 C). LiCl acts downstream of the receptors and inhibits GSK3β (Freland and Beaulieu, 2012) so it is likely WNT5A acts at this level.   Figure 3.20. WNT5A variants do not activate STF luciferase reporter. Luciferase assays conducted on HEK293 cells (A,C). Plasmids were transiently transfected 48h prior to lysing the cells. LiCl were added 24h prior to lysing. (A,B) Wild-type WNT5A plasmid significantly inhibits endogenous canonical WNT signaling. The two mutant forms of WNT5A also inhibit canonical signaling. (C,D) LiCl activated the STF reporter and both wtWNT5A and C83S variant were similarly able to antagonize signaling. Schematic diagrams represent predicted signaling pathway reflecting the luciferase results. Unknown endogenous receptors in (B) indicated by question mark. One-way ANOVA, Tukey’s post hoc *p<0.05, **p<0.01, ***p<0.001. Key: mutWNT5A – mutant WNT5A, wtWNT5A – wild-type WNT5A.     77 3.2.16 WNT5A variants are unable to antagonize WNT3A stimulated signaling unless Ror2 is provided  Wild-type WNT5A inhibition of canonical signaling is mediated by the ROR2 receptor (Mikels et al., 2009), so we wanted to see whether binding to ROR2 is impaired by the mutations in WNT5A. We first looked at the effects of adding a stimulus to increase canonical signaling using WNT3A ligand. Wild-type WNT5A plasmids significantly decreased STF in the presence of WNT3A protein. Interestingly, the C83S variant was unable to antagonize STF activity (Figure 3.21 A). When wtWNT5A plasmid was transfected along with mRor2 plasmid in the presence of WNT3A protein, the levels of STF activity were significantly decreased (Figure 3.21 A). The trend of C83S variant protein being less able to antagonize canonical signaling appeared to be rescued by the addition of mRor2 plasmid. Thus, WNT5AC83S appears to be able to bind ROR2 when provided in excess.       78  Figure 3.21. WNT5AC83S partially mediates inhibitory activity in canonical WNT signaling. Luciferase assays conducted on HEK293 cells. Plasmids were transiently transfected 48h prior to lysing the cells. WNT3A was added 24h prior to lysing. (A) WNT5A constructs in combination with Ror2 did not activate STF. When combined with WNT3A, STF activity was enhanced significantly compared to Ror2 alone. WNT5AC83S was less effective at antagonizing STF activity compared to wtWNT5A. (B) Schematic diagram representing predicted signaling pathway reflecting the luciferase results for wtWNT5A + WNT3A. (C) Schematic diagram representing predicted signaling pathway reflecting the luciferase results for WNT5AC83S + WNT3A. Unknown endogenous receptors in (B,C) indicated by question mark. (D) Schematic diagram representing predicted signaling pathway reflecting the luciferase results for WNT5AC83S or wtWNT5A + WNT3A + Ror2. One-way ANOVA, Tukey’s post hoc *p < 0.05, **p<0.01, ***p<0.001.  3.2.17 wtWNT5A and mutant WNT5A do not activate calcium WNT signaling  We measured the ability of WNT5A variants to activate the calcium Wnt signaling pathway using the NFAT luciferase reporter (Bradley and Drissi, 2010; Yang et al., 2003). HEK293 cells transiently transfected with the positive control plasmid expressing constitutively active NFAT    79 (caNFAT) strongly activated the reporter (Figure 3.22). In contrast, neither wild-type nor mutant WNT5A plasmids activated the calcium reporter. Our results are in line with previous studies that found Wnt5a only weakly activates the NFAT reporter (Topol et al., 2003). Therefore, there is no evidence for an involvement of the Ca2+ pathway in the dominant RS phenotypes; however, we cannot rule out that Ca2+ is involved in vivo.    Figure 3.22. WNT5A constructs do not activate calcium WNT signaling. NFAT luciferase reporter was used to assess non-canonical WNT Ca2+ signaling activity. HEK293 cells were transiently transfected (48h) with WNT5A plasmids (wild-type, C83S, and C182R). WNT5A plasmids failed to activate NFAT reporter (see left side of graph). The positive control, caNFAT, activated NFAT reporter showing that the necessary signal transduction molecules are present in HEK293 cells. No further increase or decrease was seen with the addition of WNT5A constructs. One-way ANOVA, Tukey’s post hoc ***p<0.001 compared to Empty plasmid alone.      80 3.3 Discussion Chondrocyte organization in the developing limb and face is a key process during embryonic development (Gao et al., 2018; Gao et al., 2011; Kuss et al., 2014; Le Pabic et al., 2014). By delivering the mutations in dominant RS to the chicken embryo we can learn how these mutations affect cell orientation and shape, and how these molecular factors can translate into phenotypic features. The retroviruses delivered to the limb or mandible are ubiquitously expressed; however, the main phenotypes are in the cartilage. This misexpression model has been very useful for understanding the cellular responses where we can measure differences. WNT5A is expressed in the early limb bud and developing face (Baranski et al., 2000; Yamaguchi et al., 1999) and is critical to establish chondrogenic elongation, as Wnt5a-/- mice show truncation defects in all outgrowing tissues (Yamaguchi et al., 1999). Herein, we have identified diverse roles for WNT5A mutations in dominant RS, whereby a single point mutation impacts chondrogenesis in the limb and mandible by disrupting outgrowth and chondrocyte polarity. Studies over the last few years have shown that PCP is required for organized orientation of chondrocytes (Gao et al., 2018; Gao et al., 2011; Kuss et al., 2014; Le Pabic et al., 2014). These cell polarity activities are linked to the non-canonical WNT signaling pathway where WNT5A is a main ligand that activates this pathway (Grumolato et al., 2010). Cell polarity in cartilage involves chondrocyte stacking to create columns that drives the elongation and shaping of the skeletal primordia. Loss of proper intercalation, flattening and stacking of chondrocytes within the cartilage could disrupt outgrowth and lead to dwarfism in the limb or mandibular hypoplasia. In addition to cell polarity in the limbs, long bone elongation is also influenced by the rate of ECM deposition (Li et al., 2015). Our experiments are similar to the human situation where normal protein is present and yet a major phenotype is produced by a single base-pair change in the other allele. This study shows a shortening in both the forelimb and mandible as well as disrupted cell polarity in the limb skeletal elements and in Meckel’s cartilage of the mandible. This data reflects the increased activation found with the non-canonical JNK-PCP signaling in our luciferase results, where others have also found that enhanced JNK-PCP signaling disrupts convergent extension (Lee et al., 2015; Qi et al., 2017). 3.3.1 Defects during chondrogenesis combine to give shorter limbs in RS  The morphology of cartilage rods in the C83S mutant viral infected limbs was greatly changed compared to the wtWNT5A and controls. These differences in direction of growth could be due to    81 changes in cell proliferation, supported by an even distribution of BrdU in the cartilage in the limb compared to GFP limbs, which had regional differences of proliferation (epiphysis vs diaphysis). The epiphysis region of the long bone regulates elongation with a large component including enhanced proliferation (van der Eerden et al., 2003). Wild-type and mutant WNT5A limbs showed an even expression of proliferation in the epiphysis + diaphysis which could contribute to a shortening of the skeletal element. Other studies in chickens at slightly later stages, have shown that once chondrocytes proliferate, the daughter cells slip underneath each other to form columns aligned with the long axis of the bone (Li and Dudley, 2009; Li et al., 2017). At later stages we found that wtWNT5A injected limbs recovered and are able to elongate and undergo hypertrophy. Furthermore, we noted that as early as stage HH29, the C83S variant caused morphology changes in the chondrocytes that are likely incompatible with elongation of the skeleton. Chondrocytes exposed to the mutant WNT5A were rounder and randomly oriented. Therefore, at the cellular level there were 3 additional changes only induced by the C83S variant that could explain the distorted morphogenesis: 1) randomized orientation of the chondrocytes, 2) round rather than ellipsoidal cell shape and 3) diffuse Prickle expression. All the aforementioned effects dominate over those of the endogenous Gallus WNT5A gene and support the idea of interference by the mutant version of WNT5A with normal functions of the protein. It is these disruptions in cell polarity including the failure of chondrocytes to flatten and stack in the proximodistal axis that likely contribute to the shortening of the limbs seen in dominant RS.  Once the chondrocytes in a long bone align into columns, the diaphyseal cells are the first to hypertrophy and it is this increase in cell size that is timed with the most rapid period of growth (Breur et al., 1991). In normal GFP or wtWNT5A infected limbs, the progression to hypertrophy occurs in the diaphysis on schedule which correlates with normal length bones. It is interesting that although there is significantly higher proliferation caused by the wtWNT5A virus, the ultimate length of the bone is no different than controls. We have seen that the C83S mutation prevents both the first phase of chondrocyte alignment and the second phase of hypertrophy while maintaining high a rate of proliferation. Also, the C182R variant reached about 75% of control length, which is correlated with a small proportion of the chondrocytes still being able to enter the hypertrophic phase. The progress to hypertrophy is confirmed by the weak COL10A1 expression and low cell density in the diaphysis of C182R infected limbs, similar to GFP or wtWNT5A viruses. In mutant infected limbs, there are also increases in the size of the epiphyses (morphology,    82 proliferation). Therefore, there could be phenotypes involving chondrocyte alignment within the growth plate. Translating to a human limb phenotype, it is likely that a delay in progressing to hypertrophy has long lasting effects on bone length that may be exacerbated by a poorly functioning growth plate. Furthermore, we did not notice any disruptions to the muscle fibers in the injected limbs with either wild-type or variant WNT5A. These results were also repeated by another group in the chicken forelimb using RCAS misexpression of WNT5A which did not affect slow or fast twitch muscle fibers in the autopod or zeugopod (Anakwe et al., 2003).  Another possible mechanism underlying the increased diameter of limb cartilages is an abnormality of the perichondrium. It has been stated that the perichondrium in the limb acts as a constraining sheath such as a ‘corset’ to restrict radial expansion and favour longitudinal growth (Rooney and Archer, 1992). The perichondrium appears in the avian ulna at stage HH28-30, when chondrocyte hypertrophy is initiated (Rooney and Archer, 1992; von der Mark et al., 1976). Regions of the limb that lack the perichondrium (such as the epiphysis) show proportionally greater radial expansion compared to the diaphysis; thus, it is envisaged that the perichondrium in the limb restricts radial growth. It is possible, based on the histological phenotypes of the WNT5AC83S variant, that the perichondrium is disrupted as compared to the wtWNT5A virus. Further characterization of the perichondrial gene expression is needed to confirm this hypothesis.     3.3.2 Mutant and Wild-type forms of WNT5A reduce outgrowth of Meckel’s cartilage While growth of the cartilage is likely inhibited due to abnormal chondrocyte stacking in the presence of mutant WNT5A, how does the mandible become shorter when wtWNT5A is introduced? We have determined that there is no difference in the cell density within a defined region of Meckel’s cartilage, nor are there changes in the diameter of Meckel’s cartilage suggesting that the condensations are misshapen (data not shown, n=6 or 7 per virus type). The explanation that we favor is that the phenotypes are due to abnormally high levels of JNK-PCP signaling, which also disrupt PCP mechanisms. This is consistent with the fact that two of the four wtWNT5A-infected embryos had disrupted cell polarity. In addition, there was a trend for cells to be rounder in wtWNT5A-infected Meckel’s cartilage. We conclude that increased JNK-PCP signaling from either the wtWNT5A or variant forms of WNT5A protein can cause disruption of tissue morphogenesis.     83 3.3.3 Comparisons between limb and mandible phenotypes During development cartilage in long bones and in Meckel’s cartilage are aligned into columns of discoid cells that arrange into stacks or rows parallel to the long axis. Cell-cell intercalations in the cartilage are regulated by the PCP pathway which is seen in chondrogenesis (Kuss et al., 2014; Li and Dudley, 2009; Sisson et al., 2015; Topczewski et al., 2011). Many of the core PCP molecules are expressed in cranial neural crest cells (Bekman and Henrique, 2002) and in the limb (Gao and Yang, 2013), which are required for proper development (Hartmann, 2007). Our data shows obvious shortening in both the limb and mandible with WNT5A variants, as well as moderate phenotypic changes with wtWNT5A. We found that the C83S variant in the limbs showed a more severe phenotype than C182R variant. Whereas in the mandible the both C83S and C182R appeared to disrupt development; although, cell polarity was only significantly disrupted with the C83S variant. These results are interesting, as comparisons made to the human clinical features show that both variants affected limb development in a similar manner, whereas C83S does not have as strong of a phenotype in the face compared to C182R (Roifman et al., 2015).  Phenotypes that were similar between the limb and face included disrupted cell polarity and cell shape. Conversely, a major difference detected between the limb and mandible in the mutant injected embryos is this overall shape change of the skeletal elements seen in the limb. The developing appendicular skeleton developed a wider diameter cartilage that is not characteristic of the mandible phenotype. One reason is that the cartilage in the avian mandible is persistent and never undergoes endochondral ossification. There are differences in the transcription factors expressed in the cartilage condensations of the limb versus Meckel’s cartilage in the mandible (Eames and Helms, 2004). Meckel’s cartilage does not express RUNX2 unlike similarly staged limb cartilage condensations (Eames and Helms, 2004). Therefore, the context in which the WNT5A viruses are operating is different in the face and limb.   3.3.4 Genotype-phenotype correlations of WNT5A variants in the chicken model The data suggests that skeletal mesomelia in patients with the C83S variant would be more severe than other variants such as C182R. The chicken provides a unique opportunity to isolate the dominant effects of the mutation while minimizing the contributions of genetic background. It is well accepted that variability in expressivity of a phenotype is due to the interaction of the gene of interest with the rest of the genome (modifier genes) and environmental factors (Fisch, 2017;    84 Orgogozo et al., 2015; Symonds and Zuberi, 2018). Our results suggest that if all genetic and epigenetic factors are held equal, that C83S is a more disruptive sequence change than C182R. As previously mentioned, the C182R variant showed greater activation of JNK-PCP compared to C83S variant in the ATF2 luciferase readout. The increased JNK-PCP activity caused by C182R does not seem to be affecting cell differentiation as judged by normal hypertrophy and bone elongation. Thus, it is not possible to predict the cellular response based solely on biochemical data. Taken together, when making conclusions about pathogenicity, multiple experimental approaches are needed and use of prediction algorithms such as those in Polyphen or Sift is inadequate (Richards et al., 2015). Unfortunately, due to the rarity of the syndrome and lack of detailed phenotypic data from radiographs it is not possible to determine whether subjects with different variants of WNT5A have greater or lesser severity in the skeletal phenotypes. Still, our results pave the way for genotype-phenotype studies carried out in the chicken that are more costly and time-consuming in the mouse model.  3.3.5 The C83S mutation decreases protein levels intra and extracellularly The direct effect of the point mutation on protein structure is unknown, however we have several pieces of evidence that can b used to suggest a mechanism. First, we can rule out an effect of the mutation on antibody binding. In ICC we see that the WNT5A polyclonal antibody recognizes similar numbers of cells expressing the mutant and wild-type proteins. The intensity of staining of the native protein is similar. Typically, antibodies have more difficulty recognizing fully folded proteins as compared to those in which only primary structure remains (reducing and denaturing conditions). Thus, for subsequent western blot analysis, any differences in levels of expression were not due to effects on antibody binding.    In the western blots, we have shown that WNT5AC83S variant protein, is less efficiently secreted than wtWNT5A. We expected to see mutant WNT5A trapped in the WNT secretion pathway (Langton et al., 2016). However, there was a decrease rather than an increase in protein in cell lysates as shown on western blots. There are three possibilities, 1) that the protein is degraded via the proteosomal pathway, 2) that the unfolded protein feedback loop has resulted in lower levels of translation of the mutant protein or 3) that RNA transcription levels are reduced. The unfolded protein response (UPR) keeps the balance between amount of protein translation for a given message and the amount of ER being made (Korennykh and Walter, 2012; Walter and Ron, 2011). Unfolded peptides feedback to sensors on the ER that in turn activate a complex response, resulting    85 in a restoration of homeostasis. Since we did not see any signs of increased apoptosis in vivo and cell passaging was carried out at a similar frequency in all DF1 cells, it appears that cell stress was not induced by the mutated protein; however, we did not look into ER stress markers including ATF6, inositol requiring 1 (IRE1) or PKR-like ER kinase (PERK) (Oslowski and Urano, 2011). If the UPR has been activated and leads to lower protein translation (Korennykh and Walter, 2012; Walter and Ron, 2011) we should detect changes in the aforementioned markers plus a decrease in the amount of ER. These hypotheses will be tested in future studies.  WNT5A has an important palmitoylation site (C104) required for FZD binding to CRD (Kurayoshi et al., 2007). The palmitoylation site is required for receptor internalization which is part of triggering downstream signaling events including antagonism of canonical signaling (Kurayoshi et al., 2007). Since mutant WNT5A ligands do not affect the palmitoylation site, it is likely the variant proteins are able to bind to cognate receptors. WNT5AC83S likely binds to the CRD on both Ror2 and FZD2 receptors in our canonical and non-canonical luciferase assays where the receptors were supplied exogenously. It is possible that the mutations introduced novel affinity for a specific receptor type or a gain in affinity for unknown receptors (Nusse and Clevers, 2017). Either of these effects would explain a dominant effect of WNT5A on development in RS. The glycosylation of WNT5A occurs on asparagines 114, 120, 311 and 325 (Kurayoshi et al., 2007). These glycans control protein folding, oligomerization and they are required for secretion (Kurayoshi et al., 2007). We demonstrated a major deficit in secretion in the C83S variant form of the protein. This evidence could not have been obtained without having a strong expression system such as the RCAS virus. Thus, we predict that the conversion of cysteine 83 to serine may indirectly affect glycosylation although the mechanism cannot be inferred from available data.  In our previous study we demonstrated that WNT5AC83S failed to activate non-canonical WNT signaling via ATF2 luciferase reporter (Hosseini-Farahabadi et al., 2017). We had performed luciferase assays using conditioned media collected from RCAS-infected cells. Conversely, in our current study, we now find an increase in ATF2 non-canonical WNT signaling (C83S or C182R) using expression plasmids. The new discovery in the present study is that the mutant protein is secreted at very low levels likely explains the previous observation (Hosseini-Farahabadi et al., 2017). In our previous study we also tested the effect of the C83S protein in an in vitro cell migration scratch test. We observed a delay in the closure of the scratch with C83S conditioned media compared to wtWNT5A or GFP. It is unlikely that the low amount of C83S protein available    86 in the conditioned media is responsible for the slower rate of cell migration. Instead it is likely that the WNT5AC83S conditioned media exerts novel inhibitory effects mediated by expression of an unidentified protein.  Overall, the lower levels of protein in a patient with WNT5A missense mutations may be lower from the mutant allele. However, there is still a normal copy of WNT5A present, so the levels of protein may not be low enough to cause haploinsufficiency. We showed that even if protein levels of WNT5AC83S are predicted to be lower in the viral infected limbs there is still a very obvious skeletal phenotype compared to wtWNT5A. This leads to the conclusion that it is not protein levels that are important for the phenotype. Instead it is disruption of chondrocyte polarity and stacking that is the main reason for shorter and wider bones. These dominant PCP effects preside over the wild-type Gallus gene that is also present in all of the limb cartilage.  3.3.6 Antagonism of canonical WNT signaling may not play a role in the dominant RS phenotype One of the main findings of our study is that even though the main signaling pathway of WNT5A is considered to be the JNK-PCP pathway, there are inhibitory effects on the canonical signaling pathway. The C83S variant is significantly impaired when it comes to antagonizing the canonical stimulus, Wnt3a. We do not feel that the increase in canonical activity affected chondrogenesis in our study. Previous work by our group and others has amply demonstrated that over-activation of canonical WNT signaling blocks the initiation of chondrogenesis including in the chicken limb using retroviruses with activated β-catenin (Hosseini-Farahabadi et al., 2013; Li and Dudley, 2009; Topol et al., 2003). Cartilages still differentiated in all the virus treatments used here regardless of whether canonical signaling might have been inhibited by wtWNT5A or slightly increased by C83S and C182R variants. Thus, the level of change in canonical signaling in vivo is likely too low to be of functional importance or may be compensated by other local factors. We have learned more about the pathogenesis of dominant RS and the complex nature of WNT signaling through our work in the chicken embryo. The avian embryo is a low cost, relatively high throughput system in which we can investigate autosomal dominant human genetic disorders. One of the mysteries that remain to be solved is why the missense mutations in WNT5A give rise to an autosomal dominant disease. If we imagine that protein levels are lower, then it would make sense    87 that the mechanism is haploinsufficiency. Dominant RS mutations in WNT5A may to a derepression of targets that are normally degraded by the wild-type WNT5A.  3.3.7 Dominant RS WNT5A variants are the result of a neomorphic function  Wild-type and variant WNT5A are able to activate JNK-PCP signaling as well as inhibit canonical WNT signaling. However, individuals with dominant RS are heterozygous and have only a single copy of the mutant allele so levels of expression will not exceed those of the wtWNT5A. We showed that the expression levels of C83S variant protein is less in the mutant allele and that the mutant protein is less efficiently secreted. We can exclude a gain-of-function as the reason for craniofacial and limb phenotypes produced in chicken embryos as well as seen in patients with dominant RS (Table 3.3).  Part of the data described with the WNT5A variants are similar to the control results, or quantitatively less than that observed with wtWNT5A (Table 3.3). The C83S variant showed similar even proliferation results in the diaphysis compared to wtWNT5A and C83S was unable to antagonize canonical WNT signaling induced by WNT3A; unless in the presence of ROR2. In dominant RS it is possible that there are slightly lower levels of WNT5A protein in the extracellular domain than normal. Furthermore, the local, short-range disruption of cell polarity and hypertrophy are more important for the manifestation of the syndrome. Other non-canonical WNTs that bind to similar receptors (WNT9A, (Weissenbock et al., 2019)) do not compensate for the presence of mutated WNT5A.       88 Table 3.3. Summary of the dominant effects of WNT5A mutations on biological function. Biological function Experimental Readout Loss of function mutWNT5A similar to control or lower levels than wtWNT5A Gain of function mutWNT5A similar to wtWNT5A Neomorphic function  mutWNT5A different than wtWNT5A Cell cycle Proliferation  1. Increase in diaphyseal proliferation at stage 30 compared to GFP (forelimb (FL))  Signaling Activation of the JNK pathway in ATF2 luciferase assays  2. Increase in ATF2 in micromass and HEK cells 3. Increase in ATF2 in HEK cells when Ror2 was added 1.  Greater increase in ATF2 when FZD2 was added   Inhibition of the canonical WNT pathway 1.  Complete inability to antagonize WNT3A, just like empty plasmid 2.  Less effective at antagonizing endogenous canonical signaling compared to wtWNT5A 4. Able to inhibit canonical signaling induced by LiCl  5. Adding Ror2 helps to improve antagonism of WNT3A  Translation Protein synthesis and secretion 3.   Less secretion in media than wtWNT5A 4.   Less synthesis in lysate than wtWNT5A 6. Similar expression pattern in ICC  Morphogenesis Skeletal morphology  7. Increase in diameter of diaphysis in DV plane just like wtWNT5A (FL) 8. wtWNT5A shortens bones by 20%, C83S causes a 50-60% reduction (FL) 2.   2X diameter of diaphysis in AP plane in the FL 3.   significantly shorter length than wtWNT5A or GFP (FL) 4.   Thinner bone collar (FL) 5.   Increase in cell density in diaphysis (FL)    89 Biological function Experimental Readout Loss of function mutWNT5A similar to control or lower levels than wtWNT5A Gain of function mutWNT5A similar to wtWNT5A Neomorphic function mutWNT5A different than wtWNT5A Morphogenesis    6.   Delay in COL10A1 expression (FL) Planar Cell Polarity PCP phenotypes   8. Rounder cell morphology (FL and mandible) 7.   Randomized orientation of chondrocytes (FL and mandible)  8.   Prickle expression diffuse (FL)        90 4. DVL1 variants disrupt chondrogenesis and overactivate PCP signaling  4.1 Introduction Dishevelled (DVL) proteins are highly conserved and are involved in both canonical and non-canonical WNT signaling pathways. There are three DVL genes (DVL1-3) in humans that may have stemmed from two rounds of genome duplication (Dillman et al., 2013; Kasahara, 2007). Mutations in DVL result in dominant Robinow Syndrome (RS) with thirteen mutations located in DVL1 and six mutations in DVL3 (Bunn et al., 2015; Murali et al., 2018; White et al., 2015; White et al., 2018; White et al., 2016b). Mutations associated with DVL1 or 3 in dominant RS are clustered in the penultimate or final exons resulting in a frameshift variant (White et al., 2018). Our lab chose to study dominant RS in DVL1 variants in addition to mutations in WNT5A (Chapter 3), as hypomorphic missense alleles in WNT5A have only been reported in a small number of families (<10, with only six variants) (Person et al., 2010; Roifman et al., 2015; White et al., 2018). Moreover, DVL1 is more compelling as it is a key player in all WNT signaling pathways. We are focusing on three of the thirteen described DVL1 variants (1519∆T, 1529∆G, 1615∆A) that cause dominant RS. These variants had the highest severity of clinical features affected in the initial report of DVL1 mutations (White et al., 2015). Clinical features of individuals with DVL1 mutations in dominant RS typically present with facial phenotypes including midface hypoplasia and hypertelorism, as well as limb phenotypes in the autopod and zeugopod, with mesomelia seen in 100% of the affected individuals (White et al., 2015).  All thirteen DVL1 variant alleles result in a C-terminally truncated protein with a premature STOP codon in the final exon (exon 15) (White et al., 2015). DVL1 variants have an identical premature termination codon and escape nonsense-mediated decay suggesting the protein is expressed albeit with an abnormal peptide of that retains 109 amino acids shared by all subjects substituting for the normal C-terminal (Mansour et al., 2018; White et al., 2015). DVL C-terminus is highly conserved from invertebrates to vertebrates (Wallingford and Habas, 2005).  Recent structural biology data show that Dvl C-terminus is important for WNT signaling. The extreme end of the C-terminus (last 40 amino acids) contains a PDZ-binding motif whereby the DVL C-terminus can bind intrinsically onto its own PDZ domain (Lee et al., 2015). This PDZ-binding motif is present in all isoforms of DVL and is also evolutionarily conserved from invertebrates to vertebrates (Qi et al., 2017; Wang and Malbon, 2012).    91  The activated state of DVL is determined by its conformation; without a WNT ligand DVL acquires a closed confirmation, whereby WNT signaling opens the conformation of DVL into its ‘active’ state. The closed conformation is known as DVL autoinhibition and causes reduced activity in the WNT pathway (Qi et al., 2017). However, the closed conformation is still able to activate canonical activity compared to an empty plasmid control. The looping of the C terminal only modulates the level of canonical signaling but does not eliminate it. The non-canonical WNT pathway was also moderately reduced (as shown by xenopus embryo phenotypes) when the DVL1 protein formed a closed conformation (Qi et al., 2017). This autoinhibition might be mediated by the C-terminus since in experiments by Witte et al. (CITATION) the C terminus could antagonize canonical signaling stimulated by CTNNB1. The C terminus alone is unable to activate canonical signaling, thus it is only the PDZ and DIX domains that are needed. The RS mutations in DVL1 and 3 replace almost the entire C terminus with a novel peptideso it is likely that signal transmission is disrupted. Intracellularly, DVL is thought to direct signaling to either the canonical or non-canonical pathways; however, the exact mechanism of the switching of signals is not understood. DVL likely activates a particular WNT pathway depending on the interacting partners, such as the available receptors at present. DVL1 associates with the cytoplasmic tails of FZD and LRP co-receptors and in this situation canonical signaling is promoted. The binding of DVL via its PDZ domain to FZD prompts signalosome formation (Bilic et al., 2007; Wong et al., 2003). The DIX and DAX domains on DVL and Axin, respectively, will interact and activate canonical WNT signaling and stabilize the signalosome (Bienz, 2014). In the presence of FZD-ROR2 dimers, it is thought DVL activates JNK-PCP signaling via its DEP domain (Nishita et al., 2010). An open conformation of DVL allows the DEP domain to be more accessible for DVL interacting partners, such as DAAM1 (Habas et al., 2001; Qi et al., 2017). The function of Dvl genes has been examined in compound mouse knockouts. There is redundancy among the Dvl genes since more novel/severe phenotypes are seen in double mutant Dvl mice (Gao and Chen, 2010; Gentzel and Schambony, 2017; Hamblet et al., 2002; Lijam et al., 1997). Skeletal defects have been observed for Dvl2−/−, Dvl1−/−;Dvl2−/−, and Dvl2+/−;Dvl3−/− mice indicating that Dvl genes are involved in development of the axial skeleton. The mouse models are not consistent with the phenotypes seen in dominant RS. The differences are likely due to the fact that human mutations in DVL1 or 3 are probably not causing a haploinsufficiency or loss-of-    92 function. Instead of attempting to knockdown DVL1, we wanted to study the exact same mutations as seen in the human genetic disease. Here, we tested the effects of several DVL1 variants on limb development in the chicken embryo. We used the RCAS retroviral system to deliver genes to the limb bud in a spatial and temporally restricted manner.  We hypothesized that cell polarity will be randomized, as our group has seen disruption in chondrocyte polarity in the mandible or limb with WNT5A mutations involved in dominant RS (section 3.2.10 and section 3.2.7) (Hosseini-Farahabadi et al., 2017). We are testing the effect of the mutations on WNT signaling using luciferase reporter assays for the various WNT pathways. We hypothesize that non-canonical WNT signaling will increase with mutant DVL1 because the open conformation should super activate this pathway.      93 4.2  Results 4.2.1 DVL1 variants affect limb development We compared the effects of expressing wtDVL1 to three variants of DVL1 that cause dominant RS (variants: 1519ΔT, 1529ΔG and 1615ΔA) (White et al., 2015). Results were compared to the GFP control virus. We first observed whole-mount cleared skeletal stained specimen at stage HH38 (12 days post-injection) and noticed a shortening in all DVL1 injected forelimbs, with a clear shortening of the stylopod (humerus), and generally the ulna as well (Figure 4.1, Table 4.1). All skeletal elements were present. The virus appeared to mostly affect the development of the stylopod and zeugopod; however, some of the skeletal elements had not started ossification yet, mostly in the autopod. In the zeugopod, it appears that the virus is mostly affecting the ulna, with the developing radius creating a bent phenotype to fit the shortened limb (Figure 4.1 F,H,J). Even though there appeared to be shortening of the limb skeletal elements, there were only statistical differences with 1615ΔA and GFP in the humerus length (Figure 4.1 K).    94     95 Figure 4.1. Skeletal phenotypes obtained from misexpression of DVL1 retroviruses in the forelimb. Embryos injected with RCAS::GFP, RCAS::wtDVL1, RCAS::DVL11519ΔT, or RCAS:: DVL11529ΔG and RCAS:: DVL11615ΔA into the region of the developing forelimb at stage HH15 (see inset in B), and fixed 12 days later at stage HH38 showing lateral view of injected (left column) and contralateral (uninjected; right column) forelimbs. The most severely affected specimens are shown in the right column. (B) Control GFP forelimbs. (D) Embryo injected with RCAS::wtDVL1. Humerus is shorter and bent (arrow), with slightly short ulna. (F) Forelimb injected with RCAS::DVL11519ΔT shows a shortened forelimb affecting the stylopod and zeugopod (arrows). Digit II and IV have delayed ossification (arrowheads). (H) RCAS:: DVL11529ΔG shows a shortened forelimb and affecting the stylopod and zeugopod (arrows). Digit IV has delayed ossification (arrowhead). (J) RCAS::DVL11615ΔA shows a shortened forelimb and affecting the stylopod and zeugopod (arrows). (K) Quantification of bone length from photographs. The only bone that had significant shortening was the humerus in the 1615 variant. Scale bar=5 mm. Key: h – humerus, r – radius, u – ulna. Digits are labeled from AP axis as II, III and IV. Dark blue/purple = Alizarin red (stains for bone), light blue = Alician blue (stains cartilage).    96 Table 4.1. Qualitative analysis of DVL1 forelimb phenotype at stage HH38. Biological replicates and detailed phenotype analysis of embryos injected with viruses into prospective limb region. Limbs can be both short/wide and can have a phenotype affecting the stylopod, zeugopod, or autopod. Abnormalities were either a limb shortening and/or pinching phenotype of the perichondrium. Virus Normal Only stylopod abnormalities Stylopod plus zeugopod abnormalities Stylopod, zeugopod and autopod abnormalities Specimens with a phenotype Delayed or absent ossification in any bone Short/ wide RCAS::GFP (n=24) 24  100% 0  0  0  0  0  0  RCAS::wtDVL1 (n=23) 7  30.43% 9  39.13% 2  8.70% 5  21.74% 16  69.57% 4  17.39% 14 60.87% RCAS::DVL1 1519∆T (n=11) 5 45.45% 0  1 9.09% 5 45.45% 6 54.55% 3 27.27% 6 54.55% RCAS::DVL1 1529∆G (n=30) 11 36.67% 9 30.00% 6 20.00% 4 13.33% 19 63.33% 5 16.66% 19 63.33% RCAS::DVL1 1615∆A (n=17) 7 41.18% 8 47.06% 2 11.76% 0  10 58.82% 2 11.76% 10 58.82%    97  4.2.2 DVL1 variants create a pinching phenotype of the perichondrium and delay chondrocyte hypertrophy We decided to look at earlier stages before osteogenesis had taken place and looked at histological sections of the limbs in order to observe the viral spread. The viruses spread throughout the limb including the cartilage by stage HH34 (8 days post injection) (Figure 4.2, inset in A-I). Wild-type DVL1 limbs were wider, slightly shorter, had compact chondrocytes as well as reduced hypertrophy in the cartilage, compared to GFP control limbs (Figure 4.2 A-D’). The DVL1 variants had a different phenotype. The cartilage had regions with mature-looking, hypertrophic chondrocytes; however, these cells also had reduced COL10A1, with pockets of COL10A1 expression (Table 4.2, Figure 4.2 E-J’). The variants also appeared to create a pinching of the perichondrium (Figure 4.2 E’,G’,I’). We looked at serial sections and we noticed that indeed, this phenotype appears to be a pinching of the cartilage rod (Figure 4.3). Instead of being a smooth cylinder, the cartilaginous elements became highly irregular. Sectioning through the DV axis of the limb reveals that the areas of cartilage are all connected to each other, rather than being separate nodules (Figure 4.3). To understand what had happened to create such irregular cartilages we examined earlier stages just after chondrogenesis had taken place, at stage HH29 to see when the phenotype first arises (Figure 4.4). We noticed that at stage HH29, 4 days post-injection, the cartilage elements are prevented from creating smooth cartilage condensations and instead the cartilage forms clusters of cells, which lack expression of SOX9 (Figure 4.4; Table 4.3). SOX9 negative regions within the cartilage contained viral GAG expression that appeared to be pockets/islands of undifferentiated mesenchyme.      98  Figure 4.2. Mutant DVL1 delays ossification and creates a pinching phenotype in the perichondrium. Embryos were injected into the presumptive limb field at stage 15 and fixed at stage 34. Sagittal sections of injected forelimbs were stained with Alcian blue and Picrosirius red (A-I) and adjacent sections were stained with anti-GAG to locate the virus in the tissues (insets), or with anti-COL10A1 to mark hypertrophic chondrocytes (B,D,F,H,J). (A) GFP virus has no effect on development. The onset of endochondral bone formation in normal limbs is seen first in the diaphysis (B,B’ arrow). (C) wtDVL1 virus shortens and widens the limb, with small chondrocytes (C’). GAG is weakly expressed in the radius, with complete expression in the ulna (inset in C). COL10A1 was not expressed in the ulna, only in the radius (D,D’, arrow). (E) 1519ΔT variant virus causes shorter and wider cartilages to develop, with a pinching phenotype in the perichondrium (E’) and showed reduced COL10A1 staining (F,F’ arrow). (G) 1529ΔG causes shorter and wider cartilages to develop, with a pinching phenotype in the perichondrium (G’) and showed delay in COL10A1 staining with pockets of hypertrophy (H,H’ arrows). (I) 1615ΔA causes shorter and wider cartilages to develop, with a pinching phenotype in the perichondrium (I’) and showed delay in COL10A1 staining (J,J’ arrows). Key: h – humerus, r – radius, u – ulna.    99  Figure 4.3. Near-adjacent sections of forelimbs injected with DVL1 viruses. Embryos were injected into the presumptive limb field at stage 15 and fixed at stage 34. Every 10th section is shown. Viral (GAG) expression is shown in Figure 4.2. Sections are cut laterally through the DV axis of the limb and show that the mutant viruses are creating a pinching phenotype (arrows). The connections between the different regions of cartilage are present in the sections thus proving that no separate, ectopic condensations are produced (arrowheads). Scale bar=500 µm. Key: h – humerus, r – radius, u – ulna.    100  Table 4.2. Qualitative analysis of DVL1 forelimb phenotype at stage HH34. Biological replicates and detailed phenotype analysis and COL10A1 expression of embryos injected with viruses into prospective limb region. Limbs can be short, or both short and dysmorphic. Dysmorphic phenotypes were pinching phenotype of the perichondrium.  COL10A1 Phenotype  No expression Weak expression Strong expression Normal Slightly dysmorphic Very dysmorphic Short RCAS::GFP (n=4) 0 0 4 100% 4 100% 0 0 0 RCAS::DVL1 (n=6) 5 83.33% 1 16.66% 0 1 16.66% 0 0 5 83.33% RCAS::DVL1 1519∆T (n=6) 1 16.66% 3 50% 0 0 1 16.66% 3 50% 2 33.33% RCAS::DVL1 1529∆G (n=3) 0 3 100% 0 0 0 3 100% 0 RCAS::DVL1 1615∆A (n=6) 3 50% 3 50% 0 0 1 16.66% 5 83.33% 0    101  Figure 4.4. Mutant DVL1 injected forelimbs are unable to form intact cartilage elements. Embryos were injected into the presumptive limb field at stage 15 and fixed at stage 29 with RCAS viruses. (A-E) Sagittal sections of injected forelimbs were stained with Alcian blue and Picrosirius red. Irregular boarders forming the developing cartilage (arrows). Viral (GAG) expression in green (insets). (A’-E’) Limbs injected with mutant DVL1 viruses are unable to form intact cartilage elements. Inset shows GAG expression in box region in adjacent sections. (A’’-E’’) increased magnification of boxed region in A’-E’. Islands of cartilage and mesenchyme seen in the mutant injected limbs (arrowheads). Scale bar=200 µm A-E and including insets; 200 µm A’-E’ and 50 µm insets; 50 µm A”-E”. Key: h – humerus, r – radius, u – ulna.         102 Table 4.3. Qualitative analysis of DVL1 forelimb phenotype at stage HH29. Biological replicates and detailed phenotype analysis of embryos injected with viruses into prospective limb region. Limbs can be short, or both short and dysmorphic. Dysmorphic phenotypes were disrupted perichondrium and/or pockets of SOX9-negative tissue within the cartilage as seen in Figure 4.4.   Normal Slightly dysmorphic Very dysmorphic short RCAS::GFP (n=5) 5 100% 0 0 0 RCAS::DVL1 (n=6) 2 33.33% 1 16.66% 0 3 50% RCAS::DVL1 1519∆T (n=6) 2 33.33% 1 16.66% 0 3 50% RCAS::DVL1 1529∆G (n=6) 1 16.66% 2 33.33% 3 50% 0 RCAS::DVL1 1615∆A (n=6) 1 16.66% 2 33.33% 2 33.33% 1 16.66%  4.2.3 DVL1 variants do not affect cell proliferation During normal limb development, there are distinct regional differences in proliferation in endochondral bones. Initially the condensation has no polarity and cartilage is even throughout. By stage HH29, the epiphyses have qualitatively higher proliferation indices than the presumptive diaphysis. By stage HH30 (section 3.2.6), the proliferation in the diaphysis drops to a very low level setting the stage for differentiation. In DVL1 virus injected limbs, there were no significant differences in the total percentage of labeled cells in the cartilage at stage HH29 (Figure 4.5). There were very few BrdU positive cells in the centre of the ulna or the future diaphysis in GFP injected forelimbs (Figure 4.5 A’). However, in the wtDVL1 injected limbs, proliferation appeared to be even across the ulna (Figure 4.5 B’). DVL1 variants show irregular borders around the developing ulna (Figure 4.5 C’-E’) and thus we could not assess regional differences in these limbs.      103     104 Figure 4.5. Wild-type DVL1 and DVL1 variants injected forelimbs do not affect cell proliferation at stage HH29 in the developing ulna. Embryos were injected with viruses into the limb bud at stage HH15 and fixed four days post-injection. (A-E) Sagittal sections of the limbs show sections probed for anti-BrdU labeling (green) and viral (anti-GAG) expression in insets. (A’-E’) show increased magnification of BrdU labeled cells in the developing ulna, with hatched marks outlining the ulna. (A,A’) BrdU label was noticeably lacking in the diaphysis in GFP controls (arrow in A’). (B,B’) wtDVL1 prevented proliferation from decreasing in the diaphysis so regional differences were eliminated. (C-E’) Most of the mutant limbs had such irregular morphology that it was difficult to see regional differences. (F) BrdU positive cells in the entire ulna were quantified. No overall difference in cell proliferation was detected. One-way ANOVA, Tukey’s post hoc test, n=3. Scale bar=500 µm (A-E), 100 µm (A’-E’). Key: r – radius, u – ulna.  4.2.4 DVL1 variant 1529ΔG randomizes cell polarity We used Golgi positioning to mark cell polarity and we found that in control injected limbs (RCAS::GFP) Golgi was located closer to 90° to the long axis of the limb (Figure 4.6 A,A’,F; 65.37 ± 1.22° SD). In wtDVL1 limbs the Golgi location was a bit more variable; however, in the RCAS::DVL11529ΔG injected limbs there was a significantly greater randomization compared to the control embryos (45° is complete randomization, mean = 44.98 ± 5.62° SD, p<0.005).  Additionally, we wanted to find out if specific proteins involved in PCP were disorganized so we examined Prickle expression. In the chondrocytes, Prickle1 is typically located at opposite ends of the flattened chondrocytes of the mouse embryo (Kuss et al., 2014). We found that in our control injected limbs, pan-Prickle was more concentrated at the ends of the chondrocytes similar to the mouse; the wtDVL1 also had similar organization; whereas mutant DVL1 chondrocytes appeared to have more disorganized Prickle orientation (Figure 4.7 A-E’). We noticed that GFP control chondrocytes were flattened and elongated and could form the appearance of columns (flattened chondrocytes stacking on top of one another). Wild-type DVL1 chondrocytes appeared more compact with slightly random prickle orientation but still appeared to form columns. Mutant DVL1 injected limbs do not form proper flattening and elongation of the chondrocytes. There are regional differences seen within the cartilage (Figure 4.7 C’’’-E’’’). However, cells located near the perichondrium in control and wild-type limbs do not show as much stacking, and the mutant DVL1 chondrocytes have no order near the perichondrium, (Figure 4.7 C’’-E’’). DVL11529ΔG chondrocytes had no polarity and lacked the elongated appearance. Chondrocytes were very disorganized, so we decided to base our reporter assays in section 4.2.6-4.2.8 on studies using    105 1529ΔG since this variant had the most penetrant phenotypes in the skeletal studies and the cell orientation studies.     106   Figure 4.6. DVL11529ΔG virus randomizes chondrocyte polarity in the forelimb. (A-E) Sagittal slices of injected forelimbs fixed 4 days post-injection (stage HH29) and stained with anti-Golgi (GM130, red). The angle between the Golgi-nucleus axis and the long axis of the bones in the ulna was measured (inset in A’). (F-J) Graphical representation of angular data for each virus type (75 cm2). Blue line is the average of each specimen, red line is the overall mean for the group and green lines represent the standard deviation. The cells were significantly more randomly oriented with DVL11529ΔG compared to GFP control virus (One-way ANOVA and Tukey’s post hoc test, p<0.005). GFP, DVL11519ΔT, DVL11615ΔA n=3; wtDVL1, DVL11529ΔG n=4. Scale bar=200 µm (A-E), 20 µm (A’-E’). Key: Pr – proximal, Di – distal    107       108 Figure 4.7. DVL1 variants randomize chondrocyte PCP molecule, Prickle in the forelimb. (A-E) Sagittal sections of injected forelimbs fixed 4 days post-injection (stage HH29) show staining for pan-Prickle and counterstained with Hoechst for nuclei; viral expression (GAG) shown in the inset. (A’-E’) Increased magnification of box region (A-E) in the centre of the cartilage in the zeugopod of the forelimb.  Double arrows indicate Prickle distribution at the ends of the elongated chondrocytes in the control forelimbs. (C’-E) Variant DVL1 limbs show disorganized prickle orientation. (A’’-E’’) Increased magnification of box region in (A-E) near the perichondrium. The dashed line is the boundary with the perichondrium. (A’’’-E’’’) Increased magnification of box region in (A-E) showing cartilage organization. (A’’’) Dashed lines show possible column formation. Asterisk is labeling tissue fold as seen in (A). (B’’’) Chondrocytes are more compact than in A’’’, but column formation is present (dashed lines). (C’’’) Regions within the cartilage have different organization of chondrocytes. Some chondrocytes appear organized with elongated cells (chondrocytes within dashed circle in (c)) indicated by double headed arrows. Chondrocytes also have disorganized prickle orientation and are not elongated (arrows in (c’). Asterisk is labeling undifferentiated mesenchyme. (D’’’) Two regions of compact chondrocytes that lack orientation as in (d) and (d’). Asterisks are labeling undifferentiated mesenchyme. (E’’’) slightly disorganized cells in (e) that appear less compact as in (e’) n=4 for each virus type. Scale bars as indicated. Key: Di – distal, Pr – proximal.   109 4.2.5 Transient transfections of DVL1 plasmids show functional variants  We wanted to determine whether there was a difference in the intracellular distribution of wtDVL1 versus mutant DVL1 proteins. The plasmid constructs used had N-terminal Flag tags, so protein localization was accomplished with anti-Flag antibodies (Figure 4.8). At 48h post-transfection wild-type and mutant DVL1 show multiple discrete puncta in the cytoplasm. There were several cells with intense staining throughout the cytoplasm which could indicate that in these cells expression levels were very high and protein was retained in the ER. Although the antibody recognizes the Flag-tag only, the distribution of the signal is consistent with that published for antibodies to DVL2 (Smalley et al., 2005). There are no antibodies that work well for DVL1.   Figure 4.8. DVL1 labeling of transfected cells in HEK293 cells. Cells were transfected using pcDNA3.2 plasmids and fixed 48h post-transfection. (A-D) Transfected Flag-tag DVL1 cells show puncta (pink). Confocal Z stack was taken over 7 microns to include the entire cell membrane and these images are maximum intensity projections.    110 4.2.6 DVL11529ΔG overactivates JNK-PCP activity In non-canonical Wnt signaling Dvl is said to be inactive when the C-terminus is experimentally bound to its own PDZ domain forming a closed looping conformation (Lee et al., 2015). When Dvl becomes active it opens the loop and activates the WNT signaling pathways. We tested whether the DVL1 C-terminal mutations would cause sustained activation of non-canonical WNT signaling using the ATF2 luciferase reporter. We found that wtDVL1 slightly activated ATF2 compared to empty plasmid (p<0.05), whereas DVL11529ΔG showed significantly greater activation (p<0.001). The addition of WNT5A protein further propagated the response for both wild-type (p<0.001) and variant DVL1 (p<0.001) (Figure 4.9).    Figure 4.9. DVL11529ΔG overactivates non-canonical PCP signaling. Luciferase assays conducted on HEK293 cells. Plasmids were transiently transfected 48h prior to lysing the cells. WNT5A protein were added 24h prior to lysing. (A) WNT5A protein activates ATF2 luciferase reporter on its own. wtDVL1 shows greater increase activation of ATF2 compared to empty plasmid, but not significantly different with the addition of WNT5A protein. DVL11529ΔG significantly activates ATF2. (B) Greater activation of JNK-PCP pathway represented by thicker lines in the signaling pathway. One-way ANOVA, Tukey’s post hoc. Key: enWNT – endogenous WNT, mutDVL1 – DVL11529ΔG, n.s. – not significant, letters (a,b,c,d,e,f) indicate treatment on bar graph. P values compared to empty plasmid: b<0.001, c<0.05, d<0.001, e<0.001, f<0.001.    111  4.2.7 DVL11529ΔG does not activate WNT calcium signaling  One of the non-canonical WNT pathways involves calcium signaling and we tested whether the DVL1 constructs would activate this pathway with an NFAT-luciferase reporter. The wild-type DVL1 and mutant DVL11529ΔG equally activate the calcium signaling pathway (p<0.001) (Figure 4.10). In order to see detectable differences between the two plasmids we propagated the signal using a plasmid expressing constitutively active NFAT (caNFAT). Under these circumstances, wtDVL1 showed greater activation compared to mutant DVL1 (1529ΔG) (wtDVL1 p<0.01, DVL11529ΔG p=0.3). Thus, it is possible that the mutations in DVL1 prevent the protein from propagating calcium signaling when a stimulus is present.       112 Figure 4.10. DVL11529ΔG partially activates calcium WNT signaling. NFAT luciferase reporter was used to assess non-canonical WNT Ca2+ signaling activity. HEK293 cells were transiently transfected (48h) with DVL1 plasmids (wild-type and 1529ΔG). (A,B) DVL1 plasmids showed a significant increase in NFAT reporter activity alone (p<0.001), left side of graph. (A,C) Concomitantly with caNFAT, there was no detectable difference between empty plasmid and 1529ΔG (p=0.3); whereas wtDVL1+caNFAT was significantly different than DVL11529ΔG+caNFAT (p<0.01), right side of graph.. One-way ANOVA, Tukey’s post hoc. Key: enWNT – endogenous WNT, mutDVL1 – DVL11529ΔG.  4.2.8 DVL11529ΔG does not activate endogenous canonical WNT signaling  DVL proteins are required for signaling in the canonical WNT pathway therefore we tested the effect of the C-terminal mutation using SuperTopflash luciferase assays. Wild-type DVL1 was able to activate the canonical pathway, whereas mutant DVL11529ΔG was unable to do so (Figure 4.11 A,B). We found greater increase activation of wtDVL1 after the addition of WNT3A protein; however, DVL11529ΔG still failed to activate the reporter (Figure 4.11 C,D). Activation levels of 1529ΔG + WNT3A were much lower than WNT3A protein on its own (concomitantly with the empty plasmid). This indicates that the 1529ΔG variant is dominantly interfering with the activation of the canonical WNT signaling pathway. Next, we wanted to antagonize the β-catenin destruction complex directly by using LiCl. With the addition of LiCl, we saw significantly increased activation with wtDVL1 and DVL11529ΔG (Figure 4.11 E,F). This synergistic  activation of STF reporter with LiCl indicates that LiCl is acting downstream of DVL. In addition, the enhanced activation of the STF reporter in the presence of LiCl and DVL1 suggests that the extra DVL molecules are able to move to the receptors, causing further disruption of the CTNNB1 destruction complex. LiCl on its own is unable to completely block formation of all destruction complexes.     113     114 Figure 4.11. DVL11529ΔG does not activate STF luciferase reporter. Luciferase assays conducted on HEK293 cells. Plasmids were transiently transfected 48h prior to lysing the cells. LiCl or WNT3A protein were added 24h prior to lysing. (A,B) Wild-type DVL1 plasmid significantly activates canonical WNT signaling whereas mutant DVL1 shows no activation of the reporter. (C,D) WNT3A protein increases activation alone or in combination with wtDVL1. Mutant DVL1 antagonizes the effects of WNT3A. (E,F) LiCl activates the STF reporter alone. In combination with wtDVL1 or mutant DVL1 there is a synergistic increase in pathway activation. One-way ANOVA, Tukey’s post hoc ***p<0.001. Key: enWNT – endogenous WNT, mutDVL1 – DVL11529ΔG.      115 4.3 Discussion Here, we studied the effects of wild-type DVL1 and DVL1 variants on the development of the chicken forelimb. We analyzed whole embryo morphogenesis, cell polarity, and transcriptional activation of WNT signaling pathways. The DVL1 variants cause a frameshift which leads to a nonsense basic peptide being added on after the DEP domain. The read-through continues and shares 109 amino acid sequence for all variants until a stop codon is reached (Mansour et al., 2018; White et al., 2015). White et al. (2015) and Bunn et al. (2015) suggested that the DVL1 mutant RNA escapes nonsense mediated decay and tested this by performing qPCR from two subjects that were compared to wild-type controls. They found expression of both wild-type and mutant transcripts in affected individuals (Bunn et al., 2015; White et al., 2015). The transcripts continued past the frameshift. Bunn et al. (2015) performed western blot analysis on DVL1 1519∆T variant and found similar protein in the lysate compared to wtDVL1. Furthermore, in a recent study on pug-nosed dogs, the authors identified a similar frame shift mutation in DVL2 (Mansour et al., 2018). They used a stable expression system involving N-terminal tagged constructs cloned into lentiviruses and expressed in NIH/3T3 cells. C-terminal tags cannot be added to DVL since they interfere with function (Lee et al., 2015; Qi et al., 2017). Mansour et al. (2018) performed westerns and found that mutant and wild-type DVL2 proteins were expressed at similar levels. It is very likely that the same would be true for all DVL1 variants. We looked at transfected HEK293 cells with DVL1 variants containing an N-terminal Flag-tag and saw expression within the cells forming discrete puncta. Others have shown that DVL is distributed as intracellular puncta (Bunn et al., 2015). Thus, our data is consistent with the idea that variant protein is generated, complete with the basic non-sense peptide at the C-terminal. 4.3.1 Defects during chondrogenesis combine to give limb phenotype in RS  The current study supports the hypothesis that dominant RS mutations in DVL1 affect early growth of the cartilage condensations. We studied three out of the thirteen currently discovered mutations in DVL1 in dominant RS (Bunn et al., 2015; White et al., 2015; White et al., 2018; White et al., 2016a) using retroviruses to misexpress the mutant gene overtop of the endogenous Gallus gene. Results were compared to GFP control and wild-type DVL1 viruses. Disrupted cartilage condensations seen with DVL1 variants is clearly illustrated by the phenotype of the limb when the cartilage shape is first present (stage HH29), and the cartilage elements are unable to form    116 intact condensations. Furthermore, at later stages of development before skeletogenesis has taken place (stage HH34), we demonstrate that the DVL1 variants delay hypertrophy with reduced COL10A1 expression. This provides evidence that disrupted condensing chondrocytes during the initial stages of development with delayed hypertrophy impedes long bone elongation. Furthermore, cartilage elements in DVL1 variants are severely disorganized, containing SOX9-positive and -negative regions, and display polarity defects with randomized Golgi bodies and Prickle orientation. Conversely, wtDVL1 limbs comprise of a different phenotype, with wider and shorter skeletal elements, an intact perichondrium and mild polarity changes.  During endochondral ossification, mesenchymal cells condense and differentiate into chondrocytes that shape a template for bone formation (Kronenberg, 2003). We observed a major disruption to the mesenchymal compaction that is necessary to form the bone template by all three DVL1 variants. The retrovirus is turned on gradually from about 24h after injection into the limb field. Progenitor cells for the cartilage are already present in the stage E.10-10.5 mouse embryos, comparative to chicken stage HH22-24 (Akiyama et al., 2005). Therefore, we have hit the window when chondrocytes are being specified. It appears that chondrocytes are still specified from the mesenchyme, but it is the organization of the cells into a compact mass that fails to occur. From the Alcian blue staining and SOX9 expression we could identify that cartilage matrix was being made, albeit regions within the cartilage were SOX9-negative. Slightly later timepoints found that COL10A1 was expressed in pockets within the cartilage. Thus, chondrocytes are forming. The complex disorganization is more likely due to intracellular mislocalization of PCP markers in chondrocytes, as shown by the diffuse Prickle staining and randomized cell orientation in stage HH29 cartilage. The fact that DVL1 is a cytoplasmic protein means that if a region of cells is dominantly affected by the variant, then the peripheral cells even if they are more normal, could still be mixed into a poorly organized condensation. In contrast, the condensations are smooth and regular in the wtDVL1-infected limbs which is consistent with more normal PCP (normal Prickle expression, cell axes perpendicular to the long axis of the bone). Thus, our molecular and histology data suggests that variant DVL1 protein dominantly interferes with chondrocyte organization at earlier stages.  Surprisingly, we did not see any changes with proliferation in our data in the entire cartilage element. However, GFP controls lacked proliferation in the diaphysis region compared to wild-type and variant DVL1 injected limbs, which displayed a more even appearance of proliferation    117 throughout the cartilage. We were unable to quantify proliferating cells by region (epiphysis or diaphysis) as it was too difficult to mark the boundaries due to the irregularity of the cartilage elements in the mutant DVL1 forelimbs. As well, in GFP control or wtDVL1 injected forelimbs the cartilage elements contained flattened, elongated chondrocytes. In contrast, DVL1 variants showed underdeveloped skeletal elements. Despite the initial failure to form intact cartilage elements after skeletogenesis has begun (stage HH38), the limbs with mutant forms of DVL1 are able to form bone. All the elements are present, albeit some digits have delayed ossification. Thus, in humans, it is possible that growth plates still form later on and limbs are able to undergo close-to-normal morphogenesis. There are not many clinical details available for the limbs of patients with DVL1 mutations, so it is not clear whether bones are dysmorphic as well as shortened.   4.3.2 Genotype-phenotype correlations in the chicken model for DVL1 variants The data supports potential genotype-phenotype correlations. The chicken embryo allows in vivo study of the pathogenic mechanism  by which DVL1 mutations cause dominant RS phenotypes. While  CRISPR/Cas9 could be used to knock-in specific single nucleotide deletions in the mouse but single base pair changes are more challenging than engineering a larger loss-of-function deletion. The chicken embryo allows us to study misexpression of the exact frameshift mutations in human DVL1 by use of retroviral-mediated gene delivery on top of the endogenous wild-type chicken homologue. We found that all three variants produced similar phenotypes, all reminiscent of RS; however, the variant 1529ΔG was slightly more severe with significant cell polarity disruptions compared to the two other variants studied (1519ΔT, 1615ΔA). All three patients with DVL1 variants (1519ΔT, 1529ΔG, 1615ΔA) have skeletal defects affecting the limbs with mesomelia and brachydactyly; although, the severity of each patient is not described (White et al., 2015). Only the patient with 1529ΔG mutation had the arm span and height noted in the case report, with an arm span/height ratio slightly less than average at 0.92 (average: 1.00-1.02 for females) (Hepper et al., 1965). Radiographs would be necessary to fully analyze the bone phenotypes.  4.3.3 Planar cell polarity signaling is disrupted in the chicken model for RS During development, chondrocytes acquire an ellipsoidal shape and form distinct columns that essentially elongate the developing limb along the PD axis (Kuss et al., 2014; Li et al., 2017). PCP signaling has been suggested to coordinate column formation (Gao et al., 2011; Li and Dudley,    118 2009). The ultimate result of active JNK-PCP signaling in vertebrates is to produce asymmetric distribution of core PCP molecules FZD, DVL, Prickle, Vangl2, which is then translated into morphogenetic changes such as convergent extension (Devenport, 2016; Keller et al., 2000). Convergent extension is also thought to occur in growing cartilage (Li et al., 2017; Li et al., 2015; Romereim and Dudley, 2011). When some of the PCP core molecules such as Vangl2 and Prickle are deleted, one of the main phenotypes is abnormal skeletal elongation (Gao et al., 2011; Liu et al., 2014; Randall et al., 2012; Wang et al., 2011; Yang et al., 2013; Yin et al., 2012). We examined Prickle expression and found qualitative differences that fit with the hypothesis that the core PCP molecules are abnormally distributed in the presence of mutant DVL1. Attempts to localize Vangl2 staining was unsuccessful due to lack of cross-reactivity with the chicken protein. Nevertheless, we were able to measure the levels of activity in the JNK-PCP pathway with biochemical assays. We saw a striking increase in activation of the non-canonical ATF2 luciferase reporter with DVL11529ΔG compared to wtDVL1. These data suggest the alterations in DVL in the chondrocytes of patients with RS results in a gain-of-function that dominates over the wild-type protein. 4.3.4 Biochemical defects caused by DVL1 variants are a complex mixture of gain-of-function in the JNK-PCP pathway concomitant with a loss-of-function in the canonical WNT signaling pathway and no change in the Calcium signaling pathway It has been shown that DVL in its active conformation (open) activates both canonical and non-canonical WNT signaling (Lee et al., 2015; Qi et al., 2017). We proposed that the DVL1 frameshift mutations would lead to an open conformation which favors the non-canonical PCP signaling pathway. Indeed, we have proven that the substitution of an abnormal peptide plus the loss of the extreme C-terminus significantly enhanced activation of the non-canonical ATF2 luciferase reporter that was not present with the wtDVL1. The activation of JNK-PCP by variant DVL1 occurs in the presence or absence of exogenous WNT5A. It is possible that the strong effect is not only due to loss of the C-terminal but also the novel/abnormal peptide. The mutation could lead to loss of binding of a C-terminal interacting protein that normally negatively regulates DVL1. One such inhibitor that is directly related to RS is NXN (Nucleoredoxin). NXN was found to be mutated in a single case of recessive RS (White et al., 2018). Studies carried out in cell lines and in Xenopus embryos showed that NXN (also known as NRX or Nucleoredoxin) can bind to DVL proteins and    119 inhibit non-canonical WNT signaling.  Knockdown of NXN with Morpholinos prevented elongation of Xenopus embryos which is driven by convergent extension (Funato et al., 2008). Furthermore, NXN expression blocked the increase in phosphorylation of JNK produced by exogenous DVL. It is interesting that NXN is inhibitory for both the canonical and non-canonical WNT signaling pathways (Funato et al., 2006; Funato et al., 2010). The prediction would be that the loss of NXN function in RS would result in gain of activity in both the JNK-PCP and canonical signaling pathways. However, as we will discuss, the DVL1 variant, unexpectedly fails to activate the canonical pathway.  The lack of activation of canonical WNT signaling by mutant DVL11529ΔG also does not agree with the data from Qi et al. (2017) who suggest that loss of the C-terminus would increase signaling. The differences are likely due to the basic abnormal peptide that has been substituted for the C-terminus in the DVL1 mutations. This implies that the peptide is not passive but is actively interfering with canonical WNT signaling. Indeed, adding exogenous DVL1 variant plasmid in combination with WNT3A significantly inhibits activation of the STF reporter. It will be interesting to see whether the variant protein can still associate with the FZD receptor.  Finally, we have also explored an alternative, non-canonical pathway involving Calcium signaling. Although mechanisms by which DVLs transduce WNT/Ca2+ signaling remains unclear, it has been shown that deleting the PDZ domain in Xenopus DVL removes the ability to activate protein kinase C in the Ca2+ pathway (Sheldahl et al., 2003). Based on these data, we expected to see no increase in Ca2+ signaling activity by the DVL11529ΔG variant, similar to empty plasmid controls. However, we showed that wild-type DVL1 and DVL11529ΔG were equally able to activate the NFAT luciferase reporter. Thus, the mutation did not impair normal levels of signaling in response to endogenous ligands. We have localized the defects in signaling to the canonical and JNK-PCP pathways and uncovered some possible functional effects caused by the abnormal peptide (Table 4.4).        120 Table 4.4. Summary of the dominant effects of DVL1 mutations on biological function.  Biological function Experimental Readout Loss of function  mutDVL1 similar to control or lower levels than wtDVL1 Gain of function mutDVL1 similar to wtDVL1 Neomorphic function  mutDVL1 different than wtDVL1 Cell cycle Proliferation 1.  No increase in proliferation at stage 29 compared to GFP 1.   Even distribution of proliferating cells across the cartilage  Signaling JNK-PCP   1.  Increase in ATF2 in HEK293 cells 2.  Increase in ATF2 in HEK293 cells when WNT5A was added  Canonical  β-catenin 2.  Complete inability to induce canonical signaling  3.  Dominantly inhibits canonical signaling induced by WNT3A, opposite to wtDVL1 Calcium pathway 3.  Unable to activate high levels of NFAT with addition of a stimulus, similar to empty plasmid 2.   Able to activate Ca+2 reporter  Translation Protein synthesis   3.   Similar expression pattern in ICC  Morphogenesis Skeletogenesis  4.   Delayed hypertrophy 5.   Shortened limbs  4.  Irregular cartilage condensations 5.  Pockets of hypertrophic chondrocytes Planar Cell Polarity PCP phenotypes    6.  Randomized orientation of chondrocytes 7.  Prickle expression diffuse      121 5. General Discussion The chicken embryo allowed us to study five variants in dominant RS, two variants in WNT5A and three in DVL1, as well as comparing the data to wild-type genes and a control virus (GFP or AlkPO4). The chicken embryo is a tractable animal model and these experiments elucidated underlying mechanisms of dominant RS that could not be predicted using bioinformatics tools. The combination of in vivo and in vitro experiments has also aided in the understanding of non-canonical WNT signaling. The RCAS system provides a formidable tool for regional misexpression of exogenous genes during embryogenesis. One strategy that we took in this work was to not stop at wholemount skeletal stains to show the phenotype. Instead we used histomorphometry of sections so that we could map the phenotypes on top of the expression of the RCAS retroviruses. This approach was not used in previous studies from our lab (Bond et al., 2016).   5.1 Strengths and weaknesses of the chicken model system as compared to the mouse One weakness of the chicken embryo is that it is not a genetic model. Therefore, we are unable to knockout genes or express them at physiological levels. It is known that the RCAS system has a strong promoter that drives levels of expression higher than physiological levels.  Indeed in other studies from our lab we have shown >10-fold increase at 48h post-injection of RCAS::WNT5A going up to >100−200-fold increase at 96h post-injection (see results for 48h post-injection into the mandible (Hosseini-Farahabadi et al., 2017). Therefore, the phenotypes could be due to stress on the cells. However, in many studies conducted by our lab (Geetha-Loganathan et al., 2014; Higashihori et al., 2010; Hosseini-Farahabadi et al., 2017; Nimmagadda et al., 2015) and others (Fuchs et al., 2010; Tiecke et al., 2006), there is little evidence of apoptosis, a consequence of too much protein in the cell. I have shown the lack of increased apoptosis in the mandible and limb for RCAS::WNT5AC83S (Chapter 3) and Hosseini-Farahabadi et al. (2017). It is important to note that neither the mutant nor wtWNT5A viruses are affecting the level of the endogenous Gallus WNT5A gene which we could detect with chicken-specific primers (Hosseini-Farahabadi et al., 2017). In unpublished data from our lab, DVL1 viruses were injected into the face and levels measured with qRT-PCR. The mutant and wild-type DVL1 RNA levels were about 150-fold increase relative to GFP virus; however, there was no change in Gallus DVL1. Thus, there are no    122 abnormal feedback loops induced by the viruses. It is important also to consider that the RNA levels were similar for mutant infected and wild-type genes so any differences in phenotype can be attributed to the mutation.  The experiments conducted here have viral expression in the background of the normal Gallus genome, which is an important advantage of using the chicken embryo. Expression of traits in humans is dependent on the modifier genes present in the rest of the genome. Dominant RS is heterozygous condition in which 50% of the protein is normal. The phenotypes are penetrant even with the normal allele but there is some variability in the expressivity of the phenotype (White et al., 2018). Therefore, we need a system in which modifier genes play less of a role. The misexpression of mutant gene overtop of the Gallus gene at high levels achieves this result. In this study I was successful in finding differences between wild-type and variant genes which can be related to the RS phenotypes.  The advantages of the mouse are that loss-of-function can be studied and heterozygous genotypes can be engineered. It is important to use an animal model that undergoes endochondral ossification, similar to humans, rather than zebrafish as most of their skeleton develops through intramembranous ossification and they lack an appendicular skeleton (Weigele and Franz-Odendaal, 2016). The CRISPR/Cas9 system can be used to knock-in specific gene variants reported in humans with RS. The adult phenotypes can be traced and the effects of the mutation in specific tissue can be studied with conditional expression. The downside is that there are numerous gene variants. It takes several rounds of breeding to generate the heterozygous animals. There is also risk that the mutant allele may not produce a phenotype in the heterozygous state and that breeding to the homozygous state or crossing into other backgrounds may be necessary. It is useful to carry out experiments on less expensive animal models and the chicken fills the gap. 5.2 Comparison of the effects of the WNT5A and DVL1 mutations and how these inform us about the pathogenesis of RS Apart from the general limb shortening there are a number of other features consistently shared by the WNT5A and DVL1 variants studied here. These similarities could explain why patients with missense mutations in WNT5A have similar phenotypes to those with a C-terminal change in DVL1 or DVL3. At the biochemical level, both mutant WNT5A and DVL1 activate the non-canonical JNK-PCP signaling pathway. This suggests that part of the phenotype is due to a gain in activity. The WNT5AC182R variant is more active in the JNK-PCP pathway than C83S. Moreover, both    123 variants (DVL11529ΔG, WNT5AC83S) do not activate canonical WNT signaling above the levels of empty (pcDNA3.2) plasmid (expected for WNT5AC83S, unexpected for DVL11529ΔG). At the cellular level, gene variants in WNT5A and DVL1 randomize chondrocyte polarity which is correlated with a lack of organization in the hypertrophic zone. Both types of variants delay hypertrophy and ultimately lead to shortened limbs. We noticed that GFP control injected limbs have a lower proliferation in the diaphysis relative to the epiphysis, whereas both gene variants remove this differential proliferation. Taken together, the limb defects in dominant RS appear to be mainly due to the effects on chondrocyte polarity as well as the gain in JNK-PCP signaling activity. 5.3 Differences between WNT5A and DVL1 mutations suggest other mediators may be involved The detailed examination of the gene variants in WNT5A and DVL1 uncovered some unanticipated differences. First, the shape of the cartilage condensations was far more irregular with the DVL1 variants compared to the WNT5A variants. As previously mentioned, this phenotypic difference is likely due to a cell autonomous effect of DVL1 creating pockets of infected cells that are disrupting morphogenesis. In contrast WNT5A codes for a secreted protein so the effects are likely to be more even throughout the cartilage condensation.  One of the most interesting differences was in the biochemical studies. On initial analysis, both variants do not activate the STF luciferase reporter. WNT5A is not supposed to activate this reporter (Mikels and Nusse, 2006), but the surprise came from the DVL1 variant. Here, the wtDVL1 activated STF luciferase but the 1529ΔG variant was completely unable to do so. Thus, the decreased ability to activate canonical signaling by DVL11529ΔG could be contributing to the phenotype. Next, we looked at the ability of WNT5A to antagonize canonical signaling. Indeed, wtWNT5A was an effective antagonist but WNT5AC83S was completely inactive in this assay. In contrast, DVL11529ΔG was a strong inhibitor of WNT3A stimulated signaling. Therefore, it appears that the DVL1 and WNT5A variants have opposite effects on the canonical pathway. However, there was one condition under which WNT5AC83S resembled DVL11529ΔG and that was when Ror2 receptor was supplied. In the presence of Ror2, WNT5AC83S was able to antagonize WNT3A. This was the first hint that Ror2 could be involved in mediating the RS phenotypes.  Contrasting behaviours between mutant WNT5A and DVL1 were observed in the LiCl experiments in which the canonical pathway was activated. WNT5A variants were similar to the wtWNT5A and inhibited the activity stimulated by 6 mM LiCl. We suspect that WNT5A variants    124 can still bind to Ror2 and thus inhibit canonical signaling downstream of GSK3β (Mikels et al., 2009; Mikels and Nusse, 2006). In contrast, both wtDVL1 and DVL11529ΔG synergized with LiCl. The variant DVL1 was likely able to complex with receptors, increasing STF activation by 2.6 fold above empty plasmid. The wtDVL1 also dramatically increased activation of STF (3.0 fold). We had expected the higher LiCl concentration (24 mM) would have completely blocked all CTNNB1 destruction complexes. Evidently, there were residual CTNNB1 complexes that were disrupted by the addition of DVL1.     125     126 Figure 5.1. Summary of the dominant effects of WNT5A and DVL1 mutations on skeletal morphogenesis. The forelimb and mandible field are injected at stage 15 prior to budding. ① Controls (empty plasmid, GFP or AlkPO4 virus) show normal cell polarity and flattened chondrocyte shape in the limb and mandible. ② Controls injected in the forelimbs show hypertrophy, low proliferation in the diaphysis and elongation of the cartilage condensations in the proximo-distal axis. ③ Control injected mandibles show no change in cell proliferation. ④ Wild-type viruses shows intermediate changes in chondrocyte polarity and altered flattening of chondrocytes in the limb and face. ⑤ wtDVL1 forelimbs have delayed hypertrophy and increased proliferation in the diaphysis. wtDVL1 shows increase in canonical signaling and moderate increase in JNK-PCP signaling. ⑥ wtWNT5A forelimbs have increased proliferation in the diaphysis and similar hypertrophy compared to controls. ⑦ wtWNT5A mandibles have no change in cell proliferation. ⑧ wtWNT5A blocks canonical signaling and has a moderate increase in JNK-PCP signaling. ⑨ Mutant viruses show randomized cell polarity and rounder chondrocyte shape. ⑩ mutDVL1 forelimbs show pockets of hypertrophy and increased cell proliferation in the diaphysis. mutDVL1 failed to activate canonical signaling and over-activates JNK-PCP signaling. ⑪ mutWNT5A forelimbs caused a lack of elongation with increased proliferation in the diaphysis and blocked hypertrophy leading to distortion of growth. ⑫ mutWNT5A mandibles showed no change in cell proliferation. ⑬ mutWNT5A is unable to antagonize the canonical pathway unless Ror2 is present and like DVL1 variants causes increased JNK-PCP signaling. Key: Di – distal, FL – forelimb, mc – Meckel’s cartilage, Md – mandible, Pr – proximal, r – radius, u – ulna.  5.4 Debunking the myth that Dominant Robinow Syndrome, is caused by hypomorphic effects of the WNT5A gene mutations  It has been reported that mutations in WNT5A are acting as a partial loss-of-function (hypomorphic alleles) (Person et al., 2010); whereas those in DVL1 are predicted to result in dominant negative or gain-of-function proteins (White et al., 2015; White et al., 2018). Our data suggests that the mutations in dominant RS have neomorphic effects that interfere with the function of the wild-type proteins. Person et al. (2010) originally ruled out the possibility of WNT5A variants acting as a dominant negative mutation and instead suggested that the mutations are having a hypomorphic affect based on functional expression experiments in zebrafish and Xenopus embryos. Mutant WNT5A phenotypes were less extreme compared to either wild-type- or dominant negative-WNT5A zebrafish embryos as the mutants produced phenotypes similar to Wnt5 loss-of-function embryos (Person et al., 2010). Rather, the group suggests that the mutations are hypomorphic since WNT5A variants have reduced function on animal cap development in Xenopus embryos (Person et al., 2010). However, their phenotypes could be the result of over-activation of the non-canonical pathway involved in convergent extension, rather than the effect of a less active form of WNT5A. Over-activation of the non-canonical pathway impacts    127 convergent extension indirectly, as seen with overexpression data from others (Lee et al., 2015; Qi et al., 2017).  There are two pieces of our data that suggest WNT5A variants lead to partial loss-of-function. The first is that variant WNT5A cannot antagonize canonical signaling. In humans with RS, there is still a normal copy of the gene, so inhibition of the canonical pathway could still occur. The second piece of data that suggests the WNT5A variants cause a loss-of-function is that the level of WNT5AC83S protein in conditioned media was significantly lower than wtWNT5A. Lower secretion would amount to about 70% of the wild-type levels in dominant RS (50% from the normal allele and 20% from the mutant allele). It is unlikely that this small difference can explain the strong effects on cartilage development. We have found several instances where WNT5A variants can dominantly override the actions of the wild-type gene, particularly in regulating chondrocyte polarity. These morphology changes occur despite the lower levels of protein expressed from the viral constructs (same viruses were used in both the in vitro and in vivo work).  5.5 Overall significance WNT signaling is a highly recognized group of signal transduction pathways. A better understanding of the less-studied non-canonical WNT signaling pathway is imperative as it is relevant to a large number of developmental mechanisms including planar cell polarity, cytoskeletal dynamics, and convergent extension (Gomez-Orte et al., 2013; Yamanaka et al., 2002). Furthermore, the developing limb is a highly recognized model to study pattern formation and skeletogenesis (Davey et al., 2018b). Also, understanding the development of Meckel’s cartilage is important as it primarily controls the size of the mandible (Mori-Akiyama et al., 2003). In this report I describe work conducted on mutations in dominant RS that disrupt morphogenesis of the forelimbs or mandible. This project can aid in understanding how non-canonical WNT signaling is involved in limb and face morphogenesis. 5.6 Future directions The ability to misexpress genes in the chicken embryo has greatly facilitated the analysis in studying the mechanism of dominant RS; however, wild-type or variant genes are not endogenously expressed in the entire mesenchyme of the human limb including the muscles. Our results using GAG antibody have demonstrated that the virus can infect the entire mesenchyme and could preclude development of the cartilage. We did not see any inhibition of myogenesis in    128 our studies, so the responding cell type appears to be mainly chondrocytes. Future experiments could incorporate a chondrogenic tissue-specific promoter to target cells undergoing chondrogenesis. RCAS vectors express inserted sequences from the retroviral (universal) promoter located within the long terminal repeat. Contrastingly, RCAN vectors lack the src splice acceptor site, and can express an inserted gene from an internal promoter (Sato et al., 2002). It has been shown that retroviral vectors are able to target the expression of a DNA insert to specific cell types in vivo via RCAN to contain predicted promoter sequences of a chondrogenic gene (Lambeth et al., 2014; Sato et al., 2002). COL2A1 could be used as the tissue-specific promoter to target chondrogenesis in the chicken embryo. Conversely, WNT5A or DVL1 could also be used as promoters to express the mutant genes where endogenous genes in dominant RS are expressed. This technique could reduce viral load and drive expression in a temporal and spatially restricted manner.   Growth of the limb occurs in the PD axis with the main contributors elongating the limbs being ECM deposition and cell volume enlargement starting at the primary ossification centre (Li et al., 2015). The growth plate is a secondary ossification center located near the epiphysis and plays a role in the ossification and elongation of long bones (Marchini and Rolian, 2018), and disturbances to the growth plate can lead to dwarfism (Mundlos and Olsen, 1997). Dominant RS variants could cause precocious maturation of the growth plate by disrupting chondrocyte alignment. Our results show a delay in progressing to hypertrophy, which might affect bone length, and may be exacerbated by a poorly functioning growth plate. However, we are limited in that the chicken embryos must be euthanized prior to hatching. Nevertheless, future work could observe the growth plate histology of the chicken embryo at later stages, just prior to hatching. Furthermore, the perichondrium is another factor contributing to the elongation of the limb (Rooney and Archer, 1992). WNT5A is expressed in the perichondrium and therefore could be disrupting its function. Wnt5a-/- mice show delayed PTHrP, which is located in the perichondrium (Hartmann and Tabin, 2000; Vortkamp et al., 1996; Yang et al., 2003). As well, IHH regulates PTHrP which is also downstream of WNT5A signaling and is required for cartilage differentiation (St-Jacques et al., 1999). Additionally, Stricker et al. (2006) studied truncated forms of ROR2 that cause recessive RS and used RCAS retroviruses to deliver the mutant genes to the chicken embryo. The group found that mutant ROR2 truncates the limb and the skeletal elements lack IHH expression (Stricker    129 et al., 2006). The variants in dominant RS could disrupt perichondrium development, therefore signaling pathways including IHH should be observed via gene expression.  In order to fully characterize the mediators of RS it will be necessary to profile gene expression using RNAseq (Kolodziejczyk et al., 2015; Wang et al., 2009). The WNT pathway is capable of transactivating other signaling pathways during development (Attisano and Labbe, 2004; Song et al., 2015) but in reality, we cannot predict the target genes that might be affected by WNT5A or DVL1 mutations. Although the WNT5A ligand and DVL1 adaptor protein will not directly change gene expression, the indirect targets gleaned from RNAseq data will still be highly informative. Pathway analysis will identify whether a particular pathway is differentially expressed and therefore worth following up. Currently, our lab has generated RNAseq data for DVL11519ΔT injected into the upper face. We plan on extending these studies to the limb in order to explore in an unbiased manner the possible downstream pathways affected by the abnormal non-canonical WNT signaling.  Furthermore, if we had access to fibroblast from patients and could generate induced pluripotent stem cells (iPSC), we could detect downstream targets by reprogramming them into iPSC and then differentiating them towards the osteogenic lineage (Csobonyeiova et al., 2017; Wu et al., 2017). This would enable us to perform gene expression or biochemical analysis which would be less artificial than using expression plasmids. The use of iPSC-osteogenic cells creates an advantage for a model to study RS. Many genetic bone disorders have finite treatment possibilities due to the absence of appropriate animal models; thus, iPSC-derived diseased models could enable us to better understand the origin and pathology of RS. With the use of iPSC-osteogenic cells, we could create a “disease in a dish” model to investigate differences in the chondrogenic profile of endochondral ossification between RS-iPSC-chondrocytes and control-iPSC-chondrocytes.   The caveat is that 3D morphogenesis would not occur and polarity defects could not be studied.  We did not explore in depth the effect of the variants on protein localization inside the cell, how the variants affect receptor affinity or binding to intracellular proteins. Our chondrocyte data suggests there are abnormalities in the distribution of core PCP components and these could lead to the disease phenotypes.   Finally, we are particularly fascinated by the effects of the DVL1 variants on cartilage condensation. We would like to extend these studies by following the typical markers of chondroprogenitor cells, markers of increased adhesion and ECM to determine exactly which steps    130 are affected by DVL1 variants (Tavella et al., 1994; Yoon et al., 2005). To isolate the effects on condensing cartilage with DVL1 variants we can turn to micromass culture which recapitulate the early steps of chondrogenesis in the limb and face (Bobick et al., 2007; Hosseini-Farahabadi et al., 2013; Langille, 1994; Ralphs, 1992; Underhill et al., 2014). Using this in vitro system, we can quantify the number, size and shape of nodules which vary according to levels of WNT signaling (Hosseini-Farahabadi et al., 2013).  Elongation of the limbs and mandible is driven by proliferation, chondrocyte hypertrophy and the ability for chondrocytes to pivot around each other to form columns (Le Pabic et al., 2014; Li and Dudley, 2009). Daughter cells are displaced laterally and intercalate into stacked columns. Disruption of the plane of cell division can lead to altered stacking and modify the dimension of the bones (Li and Dudley, 2009). We attempted to look at cell mitoses in the limb using phospho-histone 3 antibody (Hans and Dimitrov, 2001); however, the results were inconclusive as there were not enough proliferative cells in the cartilage (data not shown). We can use another approach to see whether RS mutations interfere with intercalated cell division. Others have used replication incompetent avian retroviruses (RIA viruses) (Li et al., 2017). RIA viruses encode distinct fluorescent proteins so that individual clones are marked by distinct colors. Using different retroviral envelopes, this system could allow us to perform lineage tracing in the cartilage of infected limbs with the mutant variants in dominant RS to determine cell intercalation and chondrocyte stacking in the cartilage.  A simpler approach is to use low titre RCAS retroviruses with the DVL1 variants to observe mosaic cartilage infection. This experiment can be a perfect follow-up assay to detect the distorted condensation seen with the mutant DVL1 limbs. Low titre infection will create islands of infected cells and will help pin down the relationship of infected chondrocytes to non-infected chondrocytes. This experiment can tell us how the mutant virus is affecting condensation shape and will enable us to detect whether the mutant cells are pivoting towards each other. Although, a limitation to this experiment is that only a subset of embryos will be informative, depending of the extent of viral spread. In addition to studying expression of WNT signaling with the WNT readouts using luciferase assays, other luciferase reporter systems can be used to study transcription expression of genes involved in cartilage formation such as SOX-responsive reporter (Weston et al., 2002) or SBE-4 reporter for BMP signaling (Zawel et al., 1998). These experiments would entail using micromass    131 cultures versus HEK293 cells for cartilage formation. Another assay regarding luciferase reporter would be to replicate an experiment conducted by Bunn et al. (2015) by adding both wild-type constructs and mutant constructs in equal amounts to mimic the heterozygous mutation.  5.7 Concluding remarks To date there are currently four genes involved in dominant RS that appear to affect non-canonical WNT signaling (ROR2, WNT5A, FZD2, DVL1, DVL3, RAC3, GPC4, NXN) (Bunn et al., 2015; Person et al., 2010; Roifman et al., 2015; White et al., 2015; White et al., 2018; White et al., 2016b). This study has shown that WNT5A and DVL1 variants activate PCP which is correlated with randomization of polarity in the developing cartilage. Regions of the face and limb both require growth of cartilages during prenatal development are affected by RS. RS is a rare disease that affects 1:500,000 live births but is so far one of the few genetic diseases that specifically affects non-canonical WNT signaling. With access to cheaper exome sequencing more patients with RS-like features will receive a genetic diagnosis. Thus, the prevalence of RS which has relatively low morbidity and can be transmitted through families may increase. In the bigger context of human biology, the study of rare diseases is the best way to understand gene function in humans. Understanding the effect of the mutation on many aspects of gene function can lead to therapies for a wide variety of diseases. Consequently, there are many large international efforts devoted to modeling rare diseases in a variety of animal models (Foley, 2015; Groft and Posada de la Paz, 2017; Hieter and Boycott, 2014; Wangler et al., 2017). 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