Two Alleles of Med31 Provide a Model to Study Delayed Fetal Growth, Proliferation and Placental Development

Two alleles of Med31 provide a model to study delayed fetal growth, proliferation and placental development.

A thesis submitted to The University of Manchester for the degree of Doctor of Philosophy in the Faculty of Life Sciences

2016

Kathryn Wolton

FACULTY OF LIFE SCIENCES

Table of Contents

TABLE OF CONTENTS 2 LIST OF FIGURES 5 LIST OF TABLES 8 ABBREVIATIONS 9 DECLARATION 13 COPYRIGHT 13 ACKNOWLEDGEMENTS 14 ABSTRACT 15 CHAPTER ONE: INTRODUCTION 16 1.1 Introduction 16 1.2 Mouse Development Overview 17 1.3 Placental Development 18 1.3.1 Early development of the placenta 18 1.3.2 The definitive placenta 20 1.3.3 Gas/nutrient exchange 22 1.3.4 Placental development and FGR 23 1.4 Skeletal Development 23 1.4.1 Endochondral ossification 23 1.4.2 Early limb development 24 1.4.3 Condensation of pre-chondrogenic mesenchyme 25 1.4.4 Chondrogenesis 26 1.4.5 Molecular mechanisms of growth plate outgrowth 27 1.5 Cellular Proliferation 31 1.5.1 The cell cycle 31 1.5.2 Checkpoint regulation 32 1.6 Genetic Approaches and Mutagenic Screening 34 1.6.1 Balancer chromosome 11 mutagenesis screen 35 1.7 The Mediator Complex 39 1.7.1 Structure and organization 39 1.7.2 Mediator function 45 1.7.3 Mediator mouse mutants 48 1.7.4 Mediator and development 52

1.8 Mouse Lines Used In This Project 53 1.8.1 The Med31 Null line 54 1.8.2 The Med31 Y57C line 54 1.9 Summary and Rationale For Study 55 1.10 Aims and Objectives 56 CHAPTER TWO: MATERIALS AND METHODS 58 2.1 Med31 Null and Med31 Y57C Mice 58 2.2 Genotyping and Sequencing Med31 59 2.3 Med31/Med7N Structural Modelling 61 2.4 SDS-PAGE and Western Blotting 61 2.5 Embryo Dissection and Analysis 62 2.6 Skeletal Analysis 62 2.7 Histology 63 2.8 Morphological Assessments of the Placenta 65 2.9 Immunohistochemistry 65 2.10 HEK293 Culture and Immunocytochemistry 67 2.11 Mouse Embryonic Fibroblast Culture 68 2.12 Flow Cytometry 68 2.13 cDNA Preparation 68 2.14 Quantitative PCR (qPCR) 69 2.15 Statistical Analysis 70 2.16 Microscopy 70 2.17 Mouse Lines 71 CHAPTER THREE: IDENTIFICATION OF A NEW MUTANT ALLELE 73 OF MED31 3.1 Introduction 73 3.2 Results 75 3.2.1 Confirmation of the l11Jus8 genotype 75 3.2.2 Isolation of the Med31 Y57C mutation 75 3.2.3 Med31 Y57C protein function 80 3.2.4 Med31/Med7N interaction 84 3.2.5 The Med31 Y57C allele affects embryonic growth 84 3.3 Discussion 90 3.4 Summary 92

CHAPTER FOUR: THE ROLE OF MED31 IN EMBRYONIC GROWTH 93 AND CELL PROLIFERATION 4.1 Introduction 93 4.2 Results 94 4.2.1 Med31 regulates embryonic growth 94 4.2.2 Endochondral ossification 100 4.2.3 Gene expression analysis of the endochondral growth plate 104 4.2.4 Med31 regulates cell proliferation 108 4.2.5 Gene expression analysis of key cell cycle genes 113 4.3 Discussion 114 4.4 Summary 119 CHAPTER FIVE: THE ROLE OF MED31 IN PLACENTAL DEVELOPMENT 120 5.1 Introduction 120 5.2 Results 121 5.2.1 Morphological defects of the Med31 Null placenta 121 5.2.2 Gene expression analysis of the Med31 Null placenta 127 5.3 Discussion 130 5.4 Summary 133 CHAPTER SIX: DISCUSSION, LIMITATIONS AND FUTURE 135 DIRECTIONS 6.1 Final Discussion 135 6.1.1 Growth, endochondral ossification and proliferation 135 6.1.2 Placental development 137 6.1.3 Med31 function during development 139 6.2 Future Directions 143 REFERENCES 145

Word Count: 37,300.

List of Figures

Chapter One Fig. 1.1. The early and definitive mouse placenta. 19 Fig. 1.2. The haemotrichorial mouse placenta. 20

Fig. 1.3. The endochondral growth plate. 27 Fig. 1.4. Proliferation zone feedback loop in bone. 29

Fig. 1.5. The cell cycle and checkpoint stages. 34

Fig. 1.6. The chromosome 11 balancer region. 36 Fig. 1.7. Heterozygous balancer mice are identifiable by eye. 37

Fig. 1.8. Chromosome 11 ENU balancer screen. 38 Fig. 1.9. Mediator organization. 40

Fig. 1.10. Schematic representation of electron micrograph cMED 43 interaction with Pol II. Fig. 1.11. Structural overview of the Med7N/Med31 submodule. 44 Fig. 1.12. Initiation of transcription. 47

Chapter Two Fig. 2.1. STS marker genotyping of animals. 60

Fig. 2.2. Landmarks for skeletal analysis. 63 Chapter Three Fig. 3.1. The balancer 11 mutagenesis screen complementation test. 74

Fig. 3.2. l11Jus8 false positive confirmation. 76

Fig. 3.3. STS genotyping gel. 77 Fig. 3.4. Generation of recombinant animals and isolation of the Med31 79 Y57C mutation. Fig. 3.5. Confirmation of the Med31 Y57C mutation. 80 Fig. 3.6. The Y57C residue is conserved and the mutation is damaging. 81

Fig. 3.7. The Med31 Y57C protein. 82

Fig. 3.8. The Med31 Y57C embryos. 83 Fig. 3.9. FLAG tagged Med31 and HALO tagged Med7 were co-transfected 85 into HEK293T cells.

Fig. 3.10. Transfection experiments conducted by the protein expression 86 facility, University of Manchester.

Fig. 3.11. Crossing the Med31 Null line with the l11Jus8 line does not rescue 88 the Med31 Null homozygous growth phenotype.

Fig. 3.12. Med31 Null/l11Jus8 embryos have limb growth defects. 89 Chapter Four Fig. 4.1. Med31 Y57C post natal measurements at 3 weeks. 94 Fig. 4.2. Med31 Y57C homozygous embryos display developmental growth 96 defects. Fig. 4.3. E17.5 skeletal preparations. 97 Fig. 4.4. E17.5 Med31 Null limb preparations. 98 Fig. 4.5. E17.5 Med31 Y57C limb preparations. 99 Fig. 4.6. The E15.5 radial endochondral growth plate. 102 Fig. 4.7. The chondrocytes of the growth plate. 103 Fig. 4.8. Hypertrophic cell sizes. 105 Fig. 4.9. mRNA levels of key endochondral genes. 107 Fig. 4.10. Med31 Y57C homozygous embryos have proliferation defects. 108 Fig. 4.11. Med31 Y57C MEF growth curves. 109 Fig. 4.12. Ki67 staining of E16.5 growth plates. 111 Fig. 4.13. MEF cell cycle stage comparison. 112 Fig. 4.14. mRNA levels of cell cycle genes. 113

Chapter Five Fig. 5.1. Two Med31 alleles show different placental morphology at E17.5. 122

Fig. 5.2. Fetal and placental weights. 123

Fig. 5.3. Med31 Null homozygous placentas exhibit growth defects. 125 Fig. 5.4. High magnification of the placenta reveals morphological differences. 126 Fig. 5.5. PAS staining of placental glycogen. 127 Fig. 5.6. mRNA expression of key placental genes. 129 Chapter Six Fig. 6.1. Med31 allelic series in the mouse. 139 Fig. 6.2. Two alleles of Med31 provide a model to study its function during 142 development.

List of Tables

Chapter One Table 1.1. Comparison of Mediator subunits between species. 42 Table 1.2. Summary of Mediator complex developmental 49 phenotypes. Chapter Two Table 2.1. Embryonic genotyping primers. 58 Table 2.2. PCR primers. 61 Table 2.3. Microscopy and magnifications. 70 Table 2.4. Mouse lines used. 71 Table 2.5. Primer sequences used for qPCR analysis. 72 Chapter Three Table 3.1. l11Jus8 false positive identification. 76 Table 3.2. Example recombinant genotypes. 78

List of Abbreviations

129S5- 129S5/SvEvBrd

A/P- Anterior/Posterior

AER- Apical-ectodermal ridge

BL6-C57BL/6

BMP- Bone morphogenic protein

BSA- Bovine serum albumin

Ccnb1- Gene encoding cyclin B

CDK- Cyclin-dependent kinase cDNA- Complementary DNA

ChIP- Chromatin immunoprecipitation cMED- Core Mediator

Co-IP- Co-immunoprecipitation

Col II- Collagen type II

Col X- Collagen type X

Col1a1- Gene encoding collagen type I

Col2a1- Gene encoding collagen type II

CT- Cycle threshold

CTD- Carboxy-terminal domain

DAB- 3,3-diaminobenzedine

DAPI- 4,6-diamidino-2-phenylindole

DEPC- Diethylpyrocarbonate

DGS- DiGeorge syndrome dH2O- Distilled water

DMEM- Dulbeccos’ modified eagles medium

DNA- Deoxyribonucleic acid

DPE- Downstream promoter element

E- Embryonic day

ECL- Enhanced chemiluminescence

ECM- Extracellular matrix

ENU- N-ethyl-N-Nitrosurea

EPC- Ectoplacental cone

ExE- Extraembryonic ectoderm

F: P- Fetal: placental weight ratio

FBS- Fetal bovine serum

FGF- Fibroblast growth factor

FGR- Fetal growth restriction

FITC- Fluorescein isothiocyanate

G- Generation

Gapdh- Gene encoding Glyceraldehyde 3-phosphate dehydrogenase

GATA-1- Globin transcription factor 1

Gcm1- Glial cells missing homologue 1

GLUT- Glucose transporters

GTFs- General transcription factors

HEK- Human embryonic kidney

HRP- Horseradish peroxidase

ICC- Immunocytochemistry

ICM- Inner cell mass

Igf2- Insulin-like growth factor 2

IHC- Immunohistochemistry

Ihh- Indian hedgehog

JZ- Junctional zone

LZ- Labyrinth zone

M- Mitosis

MEFs- Mouse embryonic fibroblasts miR- Micro RNA

MMPs- Matrix Metalloproteinases mRNA- Messenger RNA mTOR- Mechanistic target of rapamycin

NGS- Next generation sequencing

OS- Human osteosarcoma

P/S- Penicillin/Streptomycin

PAS- Periodic-acid Schiff

PBS- Phosphate buffered saline

PBS-T- Phosphate buffered saline with Tween 80

PFA- Paraformaldehyde

PHH3- Phospho histone H3

PIC- Pre initiation complex

Pol II (G)- Gdown1 polypeptide

Pol II- RNA polymerase II

PPARƴ- Peroxisome proliferator-activated receptor

PTCH- Patched

Pthlh- Gene encoding PTHrP

PTHrP- Parathyroid hormone-related protein

PVDF- Polyvinylidene fluoride qPCR- Quantitative polymerase chain reaction

Rec- Recombinant

RNA- Ribonucleic acid

Runx2- Runt related transcription factor 2

SDS- Sodium dodecyl sulphate

SE- Standard error

Shh- Sonic hedgehog

Slc2a3- Gene encoding GLUT3

SMC- Smooth muscle cell

Sox9- Sex determining regionY box 9

Sp7- Gene encoding Osterix

SpA-TGCs- Spiral trophoblast giant cells

Spp1- Gene encoding Osteopontin

SPR- Surface plasmon resonance

SRB- Suppressor of RNA polymerase B

SSLP- Short sequence length polymorphism

S-TGCs- Sinusoidal trophoblast giant cells

STS- Sequence tagged site

Syntb- Syncytiotrophoblast

TBS- Tris buffered saline

TF- Transcription factor

TGCs- Trophoblast giant cells

Tpbpa- Trophoblast specific protein alpha

Wnt- Wingless-related integration site

Declaration

No portion of the work referred to in the thesis has been submitted in support of an application for another degree or qualification of this or any other university or other institute of learning.

I. The author of this thesis (including any appendices and /or schedules to this thesis) owns certain copyright or related rights in it (the “Copyright”) and she has given The University of Manchester certain rights to use such Copyright, including for administrative purposes. II. Copies of this thesis, either in full or in extracts and whether in hard or electronic copy, may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as amended) and regulations issued under it or, where appropriate, in accordance with licensing agreements which the University has from time to time. The page must from part of any such copies made. III. The ownership of certain Copyright, patents, designs, trade marks and other intellectual property (the “Intellectual Property”) and any reproductions of which may be described in this thesis, may not be owned by the author and may be owned by third parties. Such Intellectual Property and Reproductions cannot and must not be made available for use without prior permission of the owner(s) of the relevant Intellectual property and Reproductions.

IV. Further information on the conditions under which disclosure, publication and commercialization of this thesis, the Copyright and any Intellectual Property and/or Reproductions described in it may take place is available in the University IP Policy (see http://www.campus.manchester.ac.uk/medialibrary/policies/intellectual- property.pdf), in any relevant Thesis restriction declarations deposited in the University Library’s regulations (see http://www.manchester.ac.uk/library/aboutus/regulations) and in The Univeristy’s policy on presentation of Thesis.

Acknowledgements

Firstly I must take the opportunity to thank my supervisor Dr Kathryn Hentges for giving me this opportunity to develop my scientific skills and allowing me to complete a qualification of this level. I was inspired by the hard work and dedication of Dr Hentges, and would like to say thank you for the continued support and guidance I received. I would also like to thank Professor Ray Boot-Handford for his role during my PhD as academic advisor, his extensive knowledge on the molecular mechanisms underpinning limb development was invaluable for the progression of this project. I would also like to thank all the past and present member of the Hentges lab group, who I found to be extremely supportive and helpful, with special thanks to Michael Boylan and Gennadiy Tenin.

On a personal level I would not have been able to complete this PhD without the love and support of my family: my parents Mark and Anne, my brother Ed and also my Grandma for always having words of encouragement when they have been most needed. I also want to thank my friends who have been with me and supported me throughout my eight years at The University of Manchester.

Finally I would like to acknowledge all the help and support my industrial sponsor Syngenta provided during my PhD. I feel extremely lucky to now be part of the team! Special thanks go to Dr Emma Barnes and Dr Jayne Wright for all their help and support.

Abstract

Fetal growth restriction (FGR) is the failure of a fetus to reach its pre-determined genetic growth potential during development. FGR is associated both with poor outcome in the neonatal period, and the onset of major adult diseases such as diabetes, hypertension and obesity. Therefore understanding what causes restricted fetal growth is important both for improving neonatal health, and for the minimization of major worldwide healthcare burdens. Described here are two mutant mouse lines, each with a distinct mutation in the Mediator complex gene Med31. These mutations result in reduced fetal growth, allowing for the investigation of the role of Med31 in the proper control of growth during development.

The first mutant mouse line (Med31 Null) carries a C/T point mutation in exon 4 of Med31. Homozygous mutant embryos display reduced growth during development, characterized by their reduced size and smaller forelimbs compared to their heterozygous littermate controls. The second mutant mouse line (Med31 Y57C) carries a T/C point mutation in exon 3 of Med31. Similarly, homozygous mutant embryos display reduced fetal growth with reductions in forelimb length compared to their heterozygous littermate controls. In both mutant lines whole embryo growth and endochondral ossification within the limbs is perturbed. This is due to defects in cellular proliferation and the misexpression of the cell cycle genes Ccnb1 and Mtor within the mutant embryos. Additionally, the Med31 Null line is embryonic lethal by E18.5 and displays morphological defects of the placenta compared to heterozygous littermate controls. These morphological differences are suggestive of defects in the function of the placenta, and are proposed as the cause of embryonic lethality. In support of this the Med31 Y57C line is viable with no defects in placental development.

New roles for Med31 in embryonic growth, cellular proliferation and placental development are identified. Moreover the two mutant lines constitue an allelic series of Med31, and the two mutations provide insights into the various ways Med31 is able to regulate transcription during development.

Chapter One: Introduction

1.1 Introduction Fetal growth restriction (FGR) is characterized by a fetus’ inability to reach its full genetic growth potential. It represents a significant human health concern, with an incidence rate of 5-10% in developed countries (Saleem et al., 2011). FGR is associated with poor outcome in the neonatal period. It is also associated with many adult diseases which represent major health burdens in the developed world, such as: cardiovascular disease, type II diabetes, hypertension and obesity (Barker, 2006).

The mouse is an ideal model for the study of fetal growth, as fundamental developmental processes such as implantation, organogenesis and growth are broadly comparable to those in human embryos. Also the gross comparative stages are well mapped and defined (Xue et al., 2011). Furthermore the mouse is a genetically tractable organism, and multiple models have been used to support clinical data on FGR.

Normal fetal growth is dependent on a number of cellular processes such as proliferation, hypertrophy and differentiation. These, and the complex control mechanisms which regulate them, can be studied using mouse models. The mouse is considered a very suitable organism for modelling human fetal development, as organogenesis and growth very closely mirror human development but on a greatly reduced timescale. This is illustrated by the mapping of Theiler stages of mouse development onto Carnegie stages of human development (Xue et al., 2011). The mouse genome aligns directly with approximately 40% of the human genome, with around 80% of human genes known to have a corresponding gene in the mouse.

One way to study the function of these genes in development is to examine the consequence of targeted mutations on the phenotype of an organism. The availability of a fully sequenced mouse genome allows the generation of transgenic animals with knock in/knock out mutations of specific genes. These methods have proved useful in linking certain genes with specific processes in development. However one limitation to this method is that prior knowledge must exist linking a particular gene to a particular process. Therefore novel gene regulatory mechanisms may be missed.

A forward genetics approach addresses these limitations. Forward genetics uses random mutagenesis so that no prior assumptions are made regarding the genes associated with particular phenotypes and mutations are created randomly, without bias. In this way unexpected links can be made between genes and phenotypes.

The Mediator complex has been identified as a major regulator of transcription (Kornberg, 2005), and is required for the correct regulation of many fundamental developmental processes (Hentges, 2011). Investigated in this work are two developmental mouse mutants of the Mediator complex gene Med31. Both these muant lines display a slow growth phenotype, though it should be noted that these phenotypes differ from each other in severity. As such they provide further insight into the role of Med31 during embryonic growth and development. Understanding the mechanisms resulting in the slow growth phenotypes may provide insights into the causes of FGR in humans. Both lines were generated in a forward genetics screen, and each has a distinct phenotype in the areas of: a) Placental development b) Skeletal development c) Cellular proliferation These areas of developmental biology are discussed in more detail in the course of this introduction.

1.2 Mouse development overview Gestation length varies between different mouse strains. The average length is considered to be twenty to twenty one days (Murray et al., 2010). Identification of a copulation plug and therefore a potential pregnancy occurs at embryonic day (E) 0.5, estimating that fertilization took place at approximately midnight (and therefore when the animals are most active), and is confirmed some hours later by the presence of a vaginal mucus plug. The preimplantation period lasts until E4.5, during which embryos progress from the morula to the blastocyst stage (Snow, 1981). At the blastula stage, due to cell polarization events, the embryo consists of an inner cell mass (ICM) and an outer layer of trophectoderm cells. By E4.5 the embryo exists as a hollow cylinder containing only a few cell types, and has begun the implantation

17 process within the uterine wall. It is now that the embryo begins the intricate series of morphological and molecular changes which are necessary to establish the body plan. Throughout gastrulation various germ layers emerge, for example the mesoderm and the definitive endoderm, together with the specification of specialized tissue (Mitiku and Baker, 2007).

Cardiomyocytes begin to differentiate from precursor cells in the primitive streak at around E7.5. This leads to the development of a heart tube by E8, with contracting cardiomyocytes which have an established pacemaker by E9. The linear heart tube begins to loop at E9, and differentiate by E10 into the four-chambered structure necessary to support the embryo. It continues to develop structurally until around E14.5. Other major events in early organogenesis include embryo turning at E8, followed by neural tube closure at E8.5 and the formation of the three primitive brain vesicles (Leatherbury and Waldo, 1995; Kaufman, 1992). Completion of organogenesis at E14.5 in the mouse embryo corresponds in humans to the final Carnegie stage of embryonic development in week 8 (O'Rahilly, 1979).

1.3 Placental Development 1.3.1 Early development of the placenta The placenta is the first organ to form during mammalian embryogenesis. In the mouse it starts to develop at E3.5 from a specialized trophectoderm layer which forms the outer epithelial layer of the developing blastocyst, surrounding the blastocoel and ICM. Around the implantation window (E4.5) the mural trophectoderm, which is not in direct contact with the ICM, begins to differentiate into primary trophoblast giant cells (TGCs), which are invasive terminally differentiated cells, allowing implantation of the blastocyst to the uterine wall (Sutherland, 2003). The trophectoderm in contact with the ICM (polar trophectoderm) receives proliferative signals from the ICM and forms the extraembryonic ectoderm (ExE) and the ectoplacental cone (EPC) (Fig. 1.1).

The next major process in mouse placental development occurs around E8.5 to initiate development of the mature placenta. Chorioallantoic attachment sees the embryonic allantois fusing with the chorion, which is derived from the expanding ExE. Expression of the transcription factor glial cells missing homologue 1 (Gcm1) marks

18 the basal surface of the chorion (Anson-Cartwright et al., 2000). It also marks the initial points of folding and creation of villi, necessary for the formation of the labyrinth zone (LZ), and the invasion of fetal blood vessels (Cross et al., 2006).

Fig. 1.1. The early and definitive mouse placenta. (A) The origins of the extraembryonic lineages begin at E3.5 with the formation of the blastocyst. (B) At E8 chorioallantoic attachment occurs, and the cell types which comprise the three placental layers begin to differentiate. (C) The mature placenta consists of three layers: the labyrinth (Lab), the spongiotrophoblast (SpT/JZ), and the maternal decidua (Dec). Abbreviations: spiral, parietal, canal and sinusoidal trophoblast giant cells (SpA-TGC, P-TGC, C-TGC and S-TGC respectively). Adapted from Cross et al. (2006).

The developing embryo initially forms under low concentrations of oxygen, before the uteroplacental circulation is established, and many placental functions are regulated by oxygen concentration. Therefore the regulation of maternal blood flow into the placenta is a tightly regulated process (Tuuli et al., 2011). In both humans and mice the placenta is haemochorial, however the circulatory exchange of the murine LZ is organized differently to human placentas. The mouse placenta is termed haemotrichorial as three distinct layers of trophoblast cells separate the fetal and maternal blood (Fig. 1.2). These consist of a mononuclear sinusoidal TGCs (S-TGCs) layer (shown in Fig. 1.2 as mononuclear trophoblast cells) and two fetal layers of

19 multinucleated syncytiotrophoblast (Syntb) cells which form a contiguous layer allowing for efficient haemochorial exchange between mother and embryo (Coan et al., 2005). In humans there is only one layer of Syntb cells and therefore the placenta is termed haemomonochorial. During labyrinth development the spongiotrophoblast layer (or junctional zone; (JZ)) structurally supports the LZ. It is composed of non- syncytial cells, thought to be derived from the EPC.

Fig. 1.2. The haemotrichorial mouse placenta. A layer of S-TGCs (I) shown as mononuclear trophoblast cells and a bilayer of syncytiotrophoblast (II and III) separate maternal and fetal circulations. Taken from Watson and Cross (2005).

1.3.2 The definitive placenta The murine definitive placenta is established by E10.5 and continues to grow in size in order to meet the increasing demands of the growing fetus. Haemochorial exchange commences from E11.5 (Georgiades et al., 2002). The mature placenta has a clearly defined anatomical and functional structure composed of two fetally derived zones and a maternally derived zone. The LZ (derived from the ExE) and the JZ (derived from the EPC) associate with the maternally derived decidua basalis. The decidua contains maternal vessels which carry blood to and from the placenta in order to keep

20 it oxygenated. Placental function is determined by the two fetal zones and therefore embryonic mutations in either the LZ or JZ can impact placental function.

The JZ lies adjacent to the decidua, and its exact function still remains somewhat unclear. However it is known to be essential for embryonic growth and survival (Tanaka et al., 1997; Hitz et al., 2005; Oh-McGinnis et al., 2011). Early in development the JZ provides structural support in order to aid labyrinth growth and development, and this would appear to continue as the placenta grows. The JZ is composed of two trophoblast subtypes, spongiotrophoblast cells, which provide the structural rigidity and support and later, glycogen cells. Both cell types are identifiable by their expression of trophoblast specific protein alpha (Tpbpa) (Lescisin et al., 1998) and are capable of secreting lactogenic and growth hormones necessary for maternal adaptations to pregnancy, and of regulating endocrine signalling during development (Coan et al., 2006; Tunster et al., 2013).

Glycogen cells of the JZ also provide a source of late gestation energy for the developing fetus. It is thought that as the embryo continues growing into late gestation the placenta must provide an extra source of energy. This is particularly so as the placenta stops gaining mass around E16.5, in contrast to the embryo which continues growth until the end of gestation ~ (E21). Moreover parturition is particularly demanding of energy. The catabolism of glycogen, which has been accumulating within the glycogen cells of the JZ from E12.5, into glucose is thought to constitute this extraembryonic late energy source (Coan et al., 2006; Bouillot et al., 2006).

Beneath the JZ lies the LZ which is the main site of nutrient, gas and waste exchange between fetus and mother. It is composed of a meshwork of highly branched villous structures in order to provide efficient exchange. Defects in this branching process are associated with embryonic lethality and FGR (Cross et al., 2003). The placenta receives oxygenated maternal blood via spiral arteries, which penetrate through the decidua and are lined with spiral TGCs (SpA-TGCs) (Fig. 1.1C). These spiral arteries are able progressively to dilate during development as they lose smooth muscle lining. This allows increasingly large volumes of maternal blood to enter the placenta as the fetus’ demands increase (Adamson et al., 2002).

Blood flow from the decidua into the JZ and then the LZ is via arterial canals. These carry blood to the embryonic base of the placenta, at which point the maternal blood

21 percolates through the sinusoid spaces within the labyrinth in a countercurrent direction to that of fetal capillary blood flow. As the blood percolates back to the apical side of the placenta the sinusoids coalesce into larger channels in the JZ, and finally into large venous sinuses within the decidua to remove waste materials (Adamson et al., 2002).

1.3.3 Gas/nutrient exchange The double layer of Syntb cells in the murine placenta (Fig. 1.2) represents the physical barrier for solutes between the maternal and fetal circulations (Coan et al., 2005).They adhere via desmosomes and gap junctions, allowing cytoplasmic contact between the two layers. Gas exchange is primarily by rapid diffusion between these surface areas, as the lipophilic molecules are able to easily penetrate the plasma membranes (Sibley et al., 1997). Other solutes are able to passively diffuse, with the rate depending on various factors including concentration gradients and the thickness of the exchange barrier (Sibley and Boyd, 1988).

In the case of hydrophilic molecules such as glucose (the main fetal energy source during development) uptake is mediated by various different transporters, as passive diffusion through plasma membranes is not possible (Dilworth and Sibley, 2012). Glucose transporters (GLUT) allow the uptake of maternal glucose via facilitated diffusion (Uldrey and Thorens, 2004). The main two isoforms in the murine placenta are GLUT1 (localized everywhere) and GLUT3 (localized within the labyrinth) (Zhou and Bondy, 1993; Shin et al., 1997). GLUT3 is thought to be the isoform responsible for meeting the nutrient demands of the growing fetus during late gestation. However in cases of compromised fetal growth, upregulation of GLUT3 alone is not sufficient to rescue a poor growth phenotype (Constância et al., 2005).

Correct uptake of amino acids is also vital for normal fetal development, as altered supply can lead to FGR (McCormick, 1985). In some cases of FGR associated reductions of a range of amino acid transport systems, including system A, , L and y+L within the syncytiotrophoblast cells of the placenta have been observed (Mahendran et al., 1993; Cetin et al., 1995; Glazier et al., 1997; Norberg et al., 1998; and reviewed in Gaccioli et al., 2013).

1.3.4 Placental development and FGR FGR is characterized by the fetus’ inability to reach its pre- determined genetic growth potential, most commonly due to placental insufficiency (Miller et al., 2008). Placental insufficiency ultimately results in the failure of a fetus to acquire sufficient amounts of nutrients and oxygen for adequate growth. This can be due to morphological placental defects (e.g. increased thickness of the exchange barrier) (Mayhew et al. 2007), decreases in nutrient transport (Sibley et al., 2005) or altered fetal/maternal blood flow (Mills et al., 2005).

In mice, late stage embryonic lethality has been linked to mutations in genes essential for placental development and function (Hemberger and Cross, 2001). This can result in placental insufficiency and therefore FGR. Currently, genetic models of FGR in mice are few in number (Swanson and David, 2015). However the role of Insulin-like growth factor 2 (Igf2) has been studied extensively and provides evidence of links between the development of small placentas and FGR (Constância et al., 2002 and 2005).

1.4 Skeletal Development 1.4.1 Endochondral ossification Skeletal bones are formed in two distinct ways during development. The flat bones of the skull, mandible and clavicle are formed directly from the differentiation of mesenchymal cells into osteoblasts, in a process called intramembranous ossification. The components of the axial and appendicular skeleton are formed by the differentiation of mesenchymal cells into a cartilage template, which is then converted to bone in a process called endochondral ossification. The endochondral cartilage template laid down creates an overall structure of two epiphyseal ends separated by the diaphysis, which becomes the primary ossification centre. This structure is formed by the development of the growth plate, which is solely responsible for the longitudinal outgrowth of the bones. The growth plate is the most fundamental structure in the development of the endochondral skeleton, as demonstrated by various mutations affecting this structure, which are discussed later.

The growth plate can be divided into zones based on both the function and morphology of the chondrocytes of which it consists. At the outer edge of each

23 epiphysis lie the stem-like resting chondrocytes (resting zone) identifiable by their round morphology (Abad et al., 2002). Below this lies the proliferative zone, where chondrocytes are constantly proliferating. These cells have a unique and easily identifiable alignment in the growth plate. They divide so as to make organized stacks of cells running down the long axis of the bone. As each cell divides in this way the spatial orientation of the column forces the longitudinal growth of the growth plate and ultimately determines the overall length of the bone. The next zone is termed the hypertrophic zone, and it is here that the chondrocytes have been signalled to stop dividing, and start accruing cellular mass. These cells also begin to produce a collagen type X (Col X) matrix which is unique to endochondral hypertrophic cells. It is this matrix which becomes ossified, the hypertrophic cells apoptose and gaps left behind are invaded by blood vessels and osteoblasts (Liu et al., 2010).

1.4.2 Early limb development Following gastrulation the mesoderm is divided spatially into three segments, paraxial, intermediate and lateral, based upon proximity to the closing neural tube (Zhang et al., 1998). The early formation of the vertebrate limb bud results from the migration of cells from the lateral plate mesoderm (Wyngaarden et al., 2010). These cells migrate to pre-determined limb fields by means of epithelial- mesenchymal interactions (Martin and Harland, 2001). They then proliferate, initiating the formation of a limb bud.

Limb buds are surrounded by a layer of ectoderm, which forms a pocket like structure over the proliferating mesenchyme. This creates a structure known as the apical- ectodermal ridge (AER), a specialized epithelium which mediates a variety of signalling processes and runs along the distal limb bud tip (Zeller et al., 2009). Removal of the AER from the wing buds of chicken embryos showed that the AER produces signals that promote outgrowth of limbs (Summerbell et al., 1973). These signalling molecules were identified as being from the fibroblast growth factor (FGF) family, and several experiments have shown they can be substituted for the AER (Fallon et al., 1994; Crossley et al., 1996).

1.4.3 Condensation of pre-chondrogenic mesenchyme Subsequent to these migratory events, and in conjunction with the formation of the limb field, is the sequestration of groups of mesenchymal cells. These firstly

24 aggregate and then differentiate into chondrocytes. Described as a ‘membranous skeleton’ by Grüneburg (1963), these skeletal condensations are the primary resource from which the skeleton is built and modified (Hall and Miyake, 2000). Pre- chondrogenic condensations occur over a 12 hour period between E9.5-E10.5 (Hall and Miyake, 2000; Karsenty et al., 2009). These condensations aggregate previously dispersed mesenchyme in order to initialise the chondrogenic differentiation process. Subsequent chondrogenic condensations act to form skeletal elements, more than one bone or cartilage anlagen can arise from a single condensation.

When the mesenchymal cells are stimulated to migrate, they remain clustered as a group due to increased adherence (Olsen et al., 2000; Hall and Miyake, 1995). This process involves fibronectin, transforming growth factor beta and neural cell adhesion molecules.

Bone morphogenic protein (BMP) signalling has also been implicated in the initiation of condensation in micromass cultures of limb mesenchyme (Barner and Niswander, 2007). The importance of BMP signalling in the initiation of condensation has further been demonstrated by deletion of the genes Bmp2 and Bmp4 from pre-chondrogenic mesenchyme, which results in the loss of radial and ulnar skeletal elements in mice (Bandyopadhyay et al., 2006).

Hoxa2 control of pre-chondrogenic condensations was first shown in Hoxa2 null mice, which displayed ectopic growth of cartilage and duplicated skeletal elements (Gendron-Maguire et al., 1993). Additionally, over expression of Hoxa2 throughout the entire developing endochondral skeleton led to spatially restricted failure of bone formation (Tavella and Bobola, 2010). These observations suggested an inhibitory role for Hoxa2 in skeletal development. However, as there was disruption in the patterning of the condensations, and a difference in the distribution of Hoxa2 positive cells when compared to wild type, it is likely that Hoxa2 functions to regulate the spatial distribution of chondrocytes, without altering function.

The transcription factor sex determining regionY box 9 (Sox9) plays a pivotal role in all aspects of endochondral ossification, including condensation. Deletion of Sox9 from limb mesenchyme prior to the onset of condensation results in the complete

25 absence of cartilage and bone, and consequently a total failure of limb development (Akiyama et al., 2002). Sox9 is expressed in all chondroprogenitors (except hypertrophic cells) during chondrogenesis (Zhao et al., 1997). Immunohistochemical experiments have shown that Sox9 co-localises with collagen type II (Col II) (Ng et al., 1997) which is the main extracellular matrix (ECM) component of cartilage (Grande et al., 1997). Expression of Col II during chondrogenesis is driven by Sox9 activity, as the chondrocyte speciﬁc enhancer element of Col2a1 is a direct target for Sox9 (Lefebvre et al., 1997) and Sox9 is known to directly bind to the first intron of Col2a1 (Bell et al., 1997). This intron contains the regulatory sequences required for the expression of Col2a1 (Zhou et al., 1995).Taken together these experiments confirm the role of Sox9 in the transcription of Col II in chondroprogenitor cells.

1.4.4 Chondrogenesis Following initial aggregation, cells located centrally within the condensation soon commit to a chondrogenic fate (Shimizu et al., 2007). Once chondrocytes are formed, the cells go through a differentiation process leading to the development of osseous tissue. A few layers of peripheral cells however do not go through this process, and instead form the perichondrium (Caplan and Pechak, 1987). The morphogens sonic hedgehog (Shh) and wingless-related integration site (Wnt) and the transcription factors Sox4, Sox11 and Sox12 are important for the maintenance of this skeletogenic mesenchymal cell population (reviewed by Lefebvre and Smits, 2005). The perichondrium, a sheath like structure which forms around the growth plate, has been suggested to have important signalling functions in the control of growth plate chondrocyte maturation (Vortkamp et al., 1996; Karsenty et al., 2009).

Induction of chondrogenesis is controlled by the transcription factors Sox5, Sox6 and Sox9. This trio alone is sufficient for the generation of chondrocytes from mesenchymal progenitors (Ikeda et al., 2004). Sox5 and Sox6 are downstream of Sox9 (Akiyama et al., 2002), and Sox9 is able to control its own expression (Kumar and Lassar, 2009).

To establish the growth plate, chondrocytes begin proliferating rapidly. These cells secrete cartilage specific ECM molecules, such as Col II, collagen type IX and collagen type XI (Shimizu et al., 2007; Aszodi et al., 2000). They become flattened to

26 form ordered structural columns of cells. This initial growth, later hypertrophic expansion, and their subsequent lateral outgrowth create the morphological template needed for formation of the long bones found in limbs. This results in globular ends, epiphyses, flanking a long middle shaft, diaphysis. Once the diaphysis structure is formed, the longitudinal growth of the structure is dependent on the differentiation pathway of the growth plate (Fig. 1.3). Chondrocytes located at each epiphysial end do not undergo this maturation process, and instead remain undifferentiated and thus able to form the articular cartilage needed at joints (Lutfi, 1974).

Fig. 1.3. The endochondral growth plate. The chondrocytes of the growth plate are organized into several zones. Resting chondrocytes become proliferative and eventually mature to the large hypertrophic cells. The matrix deposited by these large cells becomes calcified as blood vessels and osteoblasts invade the area. Taken from Wallis (1996).

1.4.5 Molecular mechanisms of growth plate outgrowth The primary mechanism governing the switch of chondrocytes to proliferation, and then to hypertrophy is controlled largely by the feedback loop outlined in Fig. 1.4. The morphogen indian hedgehog (Ihh) is an essential regulator of limb development. It is required for the normal regulation of proliferation and maturation of chondrocytes (Kronenberg, 2003).

Ihh is produced by pre-hypertrophic cells and stimulates a pathway localised to the perichondrium by signalling via the transmembrane receptor patched (PTCH), which results in the activation of the transcription factor Gli in these cells. PTCH and Gli are

27 both strongly expressed in the perichondrium and excluded from cartilage. This demonstrates that the perichondrium is the target for Ihh signalling in limb development (Vortkamp et al., 1996).

In the absence of Ihh signal chondrocytes leave the proliferative zone prematurely, contributing to a reduction in limb size and an increase in the hypertrophic fraction (Kronenberg, 2003). This is due to a loss of parathyroid hormone-related protein (PTHrP), which is secreted by both the perichondrium and early proliferative chondrocytes (Lanske et al., 1996). As the receptor for PTHrP is located on proliferating chondrocytes the following model was proposed. PTHrP is secreted in a spatially defined area, which limits its activity to cells near the perichondrium and early proliferative zone. The effect of PTHrP on these chondrocytes is to ensure they remain proliferative. As the cells move away from the perichondrium the signal becomes dampened. Cells are no longer stimulated to remain proliferative and start to mature, forming a zone of pre-hypertrophic cells which start to secrete Ihh. Ihh signals to the perichondrium, via PTCH, activating Gli which induces expression of Pthlh (the gene encoding PTHrP) in the perichondrium, starting the process again. Thus the interaction of these two signalling molecules during chondrogenesis creates a negative feedback loop, which controls the maintenance of the proliferative zone, and the initiation of hypertrophy (Fig. 1.4).

In this way both Ihh and PTHrP can be said to positively regulate the proliferative zone of the growth plate. The deletion of either Ihh or Pthlh results in a decrease in overall limb length, attributed to a failure of proper chondrocyte proliferation and subsequent acceleration into hypertrophy (Karaplis et al., 1994). However in both Ihh and Pthlh null mice, chondrocytes are able to transition into hypertrophy. This indicates the involvement of as yet unidentified additional mechanisms (Amizuka et al., 1994; Karp et al., 2000).

Fig. 1.4. Proliferation zone feedback loop in bone. (1) PTHrP is secreted from perichondrial cells and chondrocytes at the ends of long bones. PTHrP acts on receptors located on proliferating chondrocytes to keep the chondrocytes proliferating and, thereby, to delay the production of Ihh. When the source of PTHrP production is sufficiently distant, then Ihh is produced. (2) Ihh acts on its receptor located on chondrocytes to increase the rate of proliferation. (3) Through a poorly understood mechanism, it stimulates the production of PTHrP at the ends of bones. (4) Ihh also acts on perichondrial cells to convert these cells into the osteoblasts of the bone collar. Taken from Kronenberg (2003).

Sox9 has also been linked to the regulation of the proliferative zone. It is expressed in proliferating chondrocytes, as well as in sites of condensations. Of the Sox trio, only Sox9 is required for expression of Ihh and Pthlh in chondrocytes (Akiyama et al., 2002; Smits et al., 2001).

Hypertrophy is the means, other than proliferation, of increasing the size of a given organ or appendage. It is the process by which cells themselves increase in size, without replication of the genome. Hypertrophy is typically defined as a cell cycle G0/G1 phase arrest, in which there is increased ribonucleic acid (RNA) and protein

29 synthesis without division. This illustrates why in the growth plate, hypertrophy and proliferation are mutually exclusive processes (Shankland and Wolf, 2000).

In chondrocyte hypertrophy the cells increase in size and express stage specific markers, such as Col X. This precludes differentiation of osteoblast cells, matrix remodelling, vascular invasion and mineralization. These processes constitute the end point of endochondral ossification, in which cartilage is replaced by bone (Kim et al., 1999). This part of skeletogenesis is arguably less well understood than others (Karsenty et al., 2009). Currently the transcription factor runt related transcription factor 2 (Runx2) is posited as the master regulator of hypertrophy. What governs Runx2 expression is less well known. Moreover Runx2 has differing functions in the regulation of hypertrophy, depending on the time and location of its expression. For example it is well understood that Runx2 expression in the prehypertrophic zone is necessary for the proper maturation of chondrocytes. However there is some evidence which suggests a role for Runx2 in the inhibition of maturation, by interaction with FGF18 in the perichondrium (Hinoi et al., 2006).

Runx2 knockout mice show a complete absence of hypertrophic cells in the E17.5 humerus, and a reduction of the hypertrophic zone in both the ulna and radius (Kim et al., 1999). Ultimately loss of Runx2 leads to a delay in hypertrophy and therefore ossification. Clearly therefore correct Runx2 expression is important for the timely development of the growth plate, through regulation of hypertrophy.

As well as contributing to an increase in the length of the limb, the hypertrophic chondrocytes secrete factors that prepare the cartilage matrix for vascular invasion and bone deposition (White and Wallis, 2001). To facilitate these two processes the matrix must first be remodelled. Secretion of Col X is the earliest marker of this process and considered the main molecular marker of hypertrophy. One property of Col X is its unique shape, which is thought to form into a hexagonal like structure around the expanding chondrocytes (White and Wallis, 2001). Matrix metalloproteinases (MMPs) are secreted to help degrade the main components of the previously formed cartilage matrix; collagens by MMP13 and MMP2, and proteoglycans by MMP10. Other proteins secreted include angiogenic factors such as vascular endothelial growth factor 1. Collectively these processes create an environment which enables the differentiation of osteoblast precursor cells, a process

30 also regulated by Runx2, and the transcription factors Osterix and activating transcription factor 4 (Lefebvre and Bhattaram, 2010). Once osteoblasts are formed they start depositing the molecules necessary for ossification of the ECM.

There is a second differentiation pathway of early mesenchymal progenitor cells, in which cells are stimulated towards osteoblast differentiation rather than chondrocyte differentiation. Sox9 expression represses Runx2 expression in chondrocyte cells. Cells committed to osteoblast differentiation do not express Sox9, which allows Runx2 to remain expressed in the cells and activate its target genes, including osteoblast specific Osterix (Sp7), Collagen type I (Col1a1) and Osteopontin (Spp1). Mature osteoblasts completely lack Sox9 expression, and the expression of Sp7 is considered a marker of a fully committed osteoblast cell.

1.5 Cellular proliferation 1.5.1 The cell cycle The overall size of an animal, organ or even appendage depends on the number and size of the cells it contains (Conlon and Raff, 1999). Therefore regulation of cell proliferation, cell size, and cell survival determines the overall growth capabilities of an embryo during development.

Cellular proliferation is regulated by a cycling process in which cells undergoing division must pass through several stages. These stages have rigorous checkpoints which must be ‘cleared’ for progression into the next stage of the cell cycle. These are known as G1, S, G2 (collectively interphase) and M (mitosis) (Nurse, 2000). The mammalian cell cycle takes approximately twenty four hours. Mitosis takes approximately thirty minutes during each cycle and is the focus of much research, as it is within mitosis that any errors in the mechanical phase of cell division will occur (Alberts et al., 2002).

The time each cell spends in each stage is dependent on the completion of certain processes that must occur. In G1 for example, cells must grow to an appropriate size and synthesize the hundreds of different proteins needed for continuation of the cycle. The length of gap phases can vary greatly depending on external conditions and extracellular signals. If these conditions are not met cells can enter a state called G0,

31 where cells remain dormant until the required conditions are met. When conditions are met the cell can move into the S phase of the cycle, in which the chromosomes are replicated. A fundamental function of the cell cycle is to ensure accurate replication of the genome, via semi conservative replication. This process involves the generation of replication complexes generated by the co-operation of numerous enzymes that have been produced in G1. Following this, cells must then pass through a secondary gap phase (G2) in order to advance into M (Alberts et al., 2002).

M is composed of several stages. The first stage is prophase, during which the chromosomes condense to form sister chromatids, the nuclear envelope breaks down and microtubules form spindle poles which physically control the mechanical aspects of cell division (Drouin et al., 1991; Georgatos et al, 1997). Sister chromatids align in the centre of the cell during metaphase (Rieder and Salmon, 1998), and separate to opposite ends of the cell during anaphase (Nasmyth, 1999). The cell is then able to separate identical pockets of genetic material into different cytosolic compartments (telophase and cytokinesis) thereby creating a daughter cell.

Control of these phases must be tightly regulated. The sequence of events must be preserved and the directionality of the cycle maintained (Nurse, 2000). Furthermore the timing of the events must be coupled to cell growth to ensure that a cell does not divide when it is too small, or too large (Minet et al., 1979). This is particularly important during development, in which the overall growth of the organism relies on continuous accurate cellular proliferation and therefore appropriate cell cycle regulation.

1.5.2 Checkpoint regulation Checkpoint regulation is controlled by small heterodimeric protein kinases. They act as master regulators at three distinct checkpoints of the cycle, G1/S, S/G2 and G2/M. They are composed of a regulatory subunit, cyclin and a catalytic subunit, cyclin dependent kinase (CDK) (Dunphy et al, 1988; Gong and Ferrell, 2010). The CDK subunit phosphorylates numerous proteins during cell cycle to control their function. The ability of CDK to regulate progression from one stage of the cycle to the next is largely dependent on the fluctuating concentrations of cyclin at different times within the cell cycle (Tyson and Murray, 1989). 32

G1/S cyclin-CDK complexes (cyclin D1,2,3- CDK4/CDK6 and cyclin E -CDK2) are expressed as soon as cells are stimulated to replicate (Sherr and Roberts, 2004). An increase in their activity commits the cell to the cycle (Moffat and Andrews, 2004). These complexes promote progression from G1 to S phase in various ways. They phosphorylate and inactivate the ubiquitin ligase complex Cdh1 that degrades mitotic cyclins. They activate the transcription of genes which are involved in producing enzymes needed for S phase, and genes coding S phase cyclin-CDK protein components (Harper and Adams, 2001). They also phosphorylate inhibitors of S phase cyclin-CDKs, such as p27, resulting in their subsequent degradation. This liberates the S cyclin-CDKs that the inhibitors were bound to (Vlach et al, 1997; Tomoda et al, 1999).

The S Phase cyclin-CDK complex (cyclin A and CDK2) phosphorylates components of DNA pre-replication complexes which are assembled on replication origins during G1 (Tanaka et al., 2007). This initiates genomic replication and inhibits formation of new complexes, ensuring each chromosome replicates only once. Following the completion of DNA synthesis, inhibitory phosphates, which have been placed on G2/M cyclin-CDK, are removed (Dasso and Newport, 1990; Michael and Newport, 1998).

G2/M cyclin-CDK (cyclin B and CDK1) phosphorylate numerous proteins involved in multiple phases of mitosis. These phases include chromosome condensation, nuclear envelope breakdown, spindle assembly and all the initial actions needed for a cell to pass from G2 to M (Ohi and Gould, 1999). A cell becomes committed to mitosis upon the accumulation of cyclin B and CDK1 within the nucleus (Pines and Rieder, 2001). Two independent checkpoint pathways exist to prevent G2/M progression. In the scenario of DNA damage, the kinase Chk1 and Cdc25 phosphatase can rapidly prevent entry into M by depressing the activity of cyclin B/CDK1 (Chen et al., 2003). In the case of environmental insult, the p38 kinase pathway will also depress cyclin B/CDK1 activity (Mikhailov et al., 2005). Without correct expression and activation of either cyclin B or CDK1 a cell will not progress through M phase, critically limiting the proliferative ability of the cell.

Fig. 1.5. The cell cycle and checkpoint stages. To ensure cells proliferate appropriately the cell has an intrinsic system which manages the progression through cell growth, DNA synthesis and division. The cycle can be split up into G1,S, G2 and M, and between each stage exists a checkpoint which allows the progression of the cycle. For example, the ability of a cell to become mitotic and physically begin division is dependent on the correct expression of cyclin B and subsequent activation of the kinase CDK1. Taken from Alberts et al. (2002).

1.6 Genetic approaches and mutagenic screening Despite intensive research understanding of the genetic controls necessary for the co- ordination of embryonic development is still quite minimal. This is indicative of the vastly complex systems which need to be understood. Various genes need to be differentially expressed throughout development in order to generate diverse cell types, which enables the generation of distinct tissues and organs.

Identifying roles for the predicted 25,000 genes within the human genome was significantly advanced by the complete sequencing of the human genome in 2003 (Chial, 2008). The mouse genome was sequenced at the same time and provided further support for using the mouse as a model system of human development (Mouse Genome Sequencing Consortium, 2002).

A powerful and efficient way to determine gene function in model organisms is to employ high throughput mutagenesis screens. The chemical mutagen N-ethyl-N- nitrosurea (ENU) is an alkylating agent, and acts by transferring the ethyl group of ENU to nucleobases (usually thymine) in nucleic acids (Russell et al., 1979). ENU targets spermatogonial cells and induces point mutations randomly throughout the

34 genome, at the rate of about 1 base change every 700 loci (Davis and Justice, 1998; Hentges and Justice, 2004). A further advantage of this technique is that ENU mutations have particularly high frequencies of missense and splice site mutations, which often give rise to an identifiable phenotype (Boles et al., 2009; Hentges and Justice, 2004; Hentges et al., 2006).

Mutagenesis screens are highly effective at rapidly generating new phenotypes, but identification of causative mutations within genes can be a laborious process. The reduction of possible mutations to a known targeted area within the genome is therefore extremely helpful for identifying candidate genes. This can be achieved by the use of a balancer chromosome, which is an engineered inversion within a defined region on a chromosome. The region of interest can be physically inverted by incorporating loxP sites at either end, followed by Cre-mediated recombination between the two loxP sites leading to the desired rearrangement (Zheng et al., 1999). This inversion then prohibits homologous recombination between two homologous chromosome pairs during meiosis, and thereby maintains any ENU induced mutations within the balancer region. The origins of this thesis research lie in an ENU balancer mutagenesis screen created to identify novel mutations that were homozygous lethal during embryonic development (Kile et al., 2003).

1.6.1 Balancer chromosome 11 mutagenesis screen It was first observed in D. melanogaster that two lethal mutations could be maintained in a balanced stock if they were in-trans in the same region of the chromosome and if recombination was suppressed. Since then balancer chromosomes have commonly been used in stock maintenance, as well as in mutagenesis screens in the study of drosophila (Ashburner, 1989).

The first mouse balancer chromosome was engineered by inverting a 24cM region between the Trp53 and Wnt3 genes on chromosome 11 (Zheng et al., 1999). LoxP sites were introduced at either end of the desired region, and recombination between these two sites generated an inversion of this area in the presence of Cre recombinase. This would therefore fix any mutation within this region in-trans on the corresponding chromosome, as recombination between the pair would be prevented. This particular region of chromosome 11 was chosen for a number of reasons. Firstly, the region is 35 gene dense and highly syntenic with a region on human chromosome 17, where various diseases are known to map. Secondly by engineering one of the loxP sites to interrupt Wnt3, any mice homozygous for the balancer chromosome are early embryonic lethal, since previous data has shown that Wnt3 is necessary for embryonic survival beyond gastrulation (Liu et al., 1999) (Fig. 1.6).

Fig. 1.6. The chromosome 11 balancer region. The chromosome 11 balancer is an inversion (Inv) on chromosome 11 created by Zheng et al. (1999). This was used in an ENU mutagenesis screen to keep mutations in-trans on the opposite chromosome (Kile et al., 2003). Male and female mice were created from the screen which were heterozygous for the balancer region (Inv) and carried the ENU recessive lethal mutation (see also Fig. 1.8). Heterozygous mice for the balancer and mutation are considered wild type. When bred together they produce offspring which are also heterozygous (control littermates) and offspring which are homozygous for the mutation (mutant embryos). Typically the mutations are recessive lethal. Offspring homozygous for the balancer chromosome are early embryonic lethal due to disruption of the Wnt3 gene. Taken from Zheng et al. (1999).

A further useful genetic modification was the targeting of Wnt3 with a vector containing the dominant Agouti K14 promoter transgene, so that mice carrying the balancer chromosome are easily identifiable by eye as a result of the agouti pigment in their skin, colouring ears and tail yellow (Fig. 1.7). This allows identification by eye of mice which are heterozygous for the balancer chromosome.

A balancer mutagenesis screen was used to generate the recessive mutant mouse lines used in this thesis research (Kile et al., 2003). These lines were selected for further study because the homozygous mutant embryos displayed interesting developmental phenotypes. These lines were generated by breeding female mice which carried the balancer chromosome 11, with male mice which had been mutagenized by ENU. This occurred several years before the start of this thesis research.

Fig. 1.7. Heterozygous balancer mice are identifiable by eye. Heterozygous offspring generated by the cross described in Fig.1.6 are identifiable by eye due to the presence of the K14-Agouti promoter transgene within the balancer region. Mice heterozygous for the balancer chromosome have agouti pigment present in their skin, which colours ears and, most noticeably, the tail yellow (starred). Adapted from Hentges and Justice (2004).

In the original mutageneis screen the first generation (G) matings (G0) produced offspring which were heterozygous for both the balancer chromosome and the ENU induced mutation, or heterozygous for the balancer chromosome alone (Fig. 1.8). Next, G1 animals were bred to produce G2 offspring which were all heterozygous for both the balancer chromosome and the ENU induced mutation. G2 animals are the stock animals which maintain the line. These were used during this research to set up timed matings. G2 are bred to produce G3 offspring which are heterozygous for the balancer chromosome and the mutation, or homozygous for the mutation only (Fig. 1.8). These G3 offspring were examined during development as part of the original screen, and displayed interesting developmental phenotypes. As a result of the 2003 screen, fifty five recessive mouse lines with homozygous developmental phenotypes were generated (Kile et al., 2003). Two of these were used for study during this research. The causative mutation in one of these lines was known prior to this

37 research. This was a C/T point mutation in exon 4 of the Mediator complex gene Med31 (Risley et al., 2010). The second, discovered during this research, is a T/C point mutation in exon 3 of the same gene, Med31.

Fig. 1.8. Chromosome 11 ENU balancer screen. In the original screen (Kile et al., 2003) male mice of the strain C57BL/6J (red bars) were mutagenized (star symbol) with the DNA alkylating agent ENU and crossed to females of the strain 129S5 (blue bars) which were heterozygous for the balancer chromosome 11 (blue arrowhead bar). This generated offspring in G1 which were heterozygous for the C57BL/6J mutation and the 129S5 balancer chromosome (G1 right). Additionally there were offspring which were heterozygous for the balancer chromosome and C57BL/6J chromosome without the mutation (G1 left). These animals were crossed together to generate more animals carrying both the mutation and the balancer chromosome (G2). The G2 generation (bracketed) are the animals used to set up timed matings to generate the G3 offspring described in this research. G2 animals are maintained as a heterozygous stock and are set up in timed matings to produce G3 homozygous mutant C57BL/6J embryos (left) and control littermates which are heterozygous mutant C57BL/6J and balancer 129S5 (right). Adapted from Hentges and Justice (2004). 38

1.7 The Mediator Complex 1.7.1 Structure and organization RNA polymerase II (Pol II) is responsible for the expression of all protein coding (and the majority of non-coding RNA genes), and synthesis of all messenger RNA (mRNA) molecules in eukaryotes. The intricate regulation of Pol II transcription underlies all events necessary for normal cell differentiation and development.

Pol II is not sufficient to initiate transcription (Roeder, 1996). Instead Pol II functions and is regulated through a multiprotein complex, termed the pre initiation complex (PIC). General transcription factors (GTFs) including TFIIA, TFIIB, TFIID, TFIIE, TFIIF and TFIIH are necessary components of the PIC, and they interact both with Pol II and activator/repressor proteins to regulate the presence of the PIC at promoter regions (Orphanides et al., 1996). In 1990, using yeast biochemistry, Kornberg and colleagues identified that extra factors interacted with Pol II. These factors were thought to mediate the transcriptional activation process, be necessary for response to transcriptional regulators, and regulate activator-dependent transcription in vivo (Kelleher et al., 1990).

These extra factors were identified following a genetic screen in S.cerevisiae for mutations which could suppress the conditional phenotypes of Pol II carboxy-terminal domain (CTD) truncation. This led to the discovery that several suppressor of RNA polymerase B (SRB) proteins create a multisubunit structure which binds tightly to Pol II (Thompson et al., 1993). This was the first discovery of what was later named the Mediator complex, however many more studies in yeast were performed to isolate all components of this multi subunit structure. The complexity of the structure and the different genetic screens used to fully identify all the components gave rise to a plethora of terminologies to describe the subunits in S.cerevisiae (SRB, SOH, PGD, CSE, NUT, SSN) (Biddick and Young, 2005). Furthermore multiple laboratories then used a variety of procedures to isolate mammalian complexes, which were also given differing names (TRAP/SMCC, NAT, ARC, DRIP, Srb/MED) (Jiang et al., 1998; Sun et al., 1998; Ito et al., 1999; Näär et al., 1999; Malik et al., 2000). In 2004 a unified nomenclature for Mediator was established which consisted of Med1 through Med31, together with the CDK8 subunit (Guglielmi et al., 2004).

Mediator is described as a molecular adaptor. It links gene specific transcription factors, located at upstream promoter and enhancer elements, with Pol II and the GTFs at the main promoter site (Koleske and Young, 1995; Conaway et al., 2005). Mediator functions as a ‘modular interface’, and is unable to act directly on transcriptional enhancer/silencer elements (Poss et al., 2013). However, since the complex is considered a ubiquitous regulator of gene expression, it is often also described as a GTF (Ansari et al., 2009).

The specification and maintenance of cell fate in multicellular organisms is critically dependent upon the precise spatio-temporal control of Pol II transcription in response to a determinative set of signals, both intrinsic and extrinsic to the cell. Because of its central importance in organismal biology, metazoans have evolved an elaborate protein machinery to ensure proper control of transcription. This is discussed below.

Mediator is a dynamic, multi-module, multi-protein complex. It consists of four main modules, termed ‘head’ ‘middle’ ‘tail’ and ‘kinase’ which have variable subunit composition and an overall structural flexibility (Larivière et al., 2013). Each submodule is composed of a number of Mediator subunits (Med proteins), as illustrated by Fig. 1.9.

Fig. 1.9. Mediator organization. Based on collective studies described in section 1.7.1. A proposed schematic of modular organization and subunit composition of Mediator. Blue subunits indicate placement within the tail module, green within the middle module, purple within the head module, and red units are the variable kinase domain. Med14 (in grey) is proposed to form part of an architectural backbone. 40

Table 1.1 outlines the inter-species subunit sequence and residue conservation in H.sapiens, S.cerevisiae, D. melanogaster and M.musculus. As the focus of this research is the subunit Med31 highlighted in yellow are comparisons between the M.musculus and H. sapiens subunits. In blue, the same comparison for a second subunit (Med7) is shown. Med7 is located within the middle module of Mediator and is known to directly interact with Med31 (Koschubs et al., 2009).

A fundamentally important feature of Mediator is its ability to change subunit composition (Tsai et al., 2014). Subunits are known to be substituted in order to regulate specific biological functions (reviewed in Allen and Taatjes, 2015), but little is known about how subunit exchange is regulated. In addition, Mediator structure is able to change dramatically upon binding to other proteins or protein complexes. In each case structural changes in Mediator are associated with changes in function. Structural shifts that occur upon transcription factor (TF) binding, which seem to propagate throughout the entire Mediator complex, correlate with the activation of Pol II transcription (Taatjes et al., 2002). Large scale structural shifts also occur upon the binding of Mediator to Pol II, or the inhibitory CDK8 module (Tsai et al., 2013). These structural changes may ensure that the binding of Pol II and the CDK8 module to Mediator are mutually exclusive, in order to allow both for activation and inhibitory effects on the complex via different mechanisms.

The overall structure of the complex, both in yeast and mammals, is modular. The subunits are organized into head, middle, tail and kinase modules (Asturias and Kornberg, 1999). The subunits composing the head and middle modules are tightly associated with each other, and it is generally accepted that these comprise a core Mediator (cMED). This has been described in both S.cerevisiae (Plaschka et al., 2015; Liu et al., 2001) and in H. sapiens (Cevher et al., 2014). The term cMED refers to the ability of the submodules to associate stably with one another, and activate transcription in vitro. Both the head and middle modules are known to physically make contact with Pol II (Davis et al., 2002). This is illustrated in Fig. 1.10 which shows a reconstructed electron micrograph image of cMED binding to Pol II.

In cMED, both S.cerevisiae and H. sapiens require the additional presence of the

Table 1.1. Comparison of Mediator subunits between species. Basic comparison of Mediator subunits in H. sapiens (Hs) S.cervisiae (Sc), D. melanogaster (Dm), and M. musculus (Mm) by percent identity, percent similarity. Highlighted in blue Mm Med7 has 98% protein sequence similarity to Hs Med7, and in yellow Mm Med31 has 100% similarity to Hs Med31. Adapted from Poss et al. (2013).

Med14 subunit, which has traditionally been considered a component of the tail module. Current opinion of Mediator structure however places Med14 as an architectural backbone, with the ability to link all four modules together (Cevher et al., 2014).

Fig. 1.10. Schematic representation of electron micrograph cMED interaction with Pol II. Pol II (grey) interacts directly with the head (blue) and middle (purple) modules of Mediator forming a core Mediator structure necessary for transcription (cMED). Taken from Plaschska et al. (2015).

The mammalian Mediator head module is known to contain the subunits Med6, Med8, Med11, Med17, Med18, Med20 and Med22 (Larivière et al., 2012). The middle module contains the subunits Med1, Med4, Med7, Med9, Med10, Med21, and Med31 (Guglielmi et al, 2004; Béve et al, 2005). Interestingly, whilst attempting to fully elucidate the structure of the middle domain a separate submodule was identified, the Med7N/Med31 submodule (Fig. 1.11). Using a combination of biochemical, X ray crystallography, phenotypic and transcriptome techniques in S.cerevisiae the Med7N/Med31 submodule was established as structurally and functionally distinct, and required for activated transcription of certain sets of genes (Koschubs et al., 2009). To date this is the first identification of such a submodule within cMED having independent transcriptional activity. Such a submodule has not yet been described in mammalian Mediator.

Fig. 1.11. Structural overview of the Med7N/Med31 submodule. Ribbon model representation of S.cervisiae Med7N/Med31 crystal structure. Taken from Koschubs et al. (2009).

There still remains some debate over the exact subunit composition of the tail module. It is generally accepted that the tail seems mostly to be involved in regulating specific signal transduction events, through interaction with gene specific transcription factors (Malik and Roeder, 2010; Conaway and Conaway, 2011). However it should also be noted that this behaviour is not exclusive to the tail module, as examples of transcription factor interactions have been shown to occur within cMED. These will be discussed later.

Mediator can also incorporate a variable kinase module which can reversibly associate (Knuesel et al., 2009). The module (CDK8) is often described as comprising CDK8, cyclin C, Med12 and Med13. However Sato et al. (2004) identified some subsets of Mediator with different isoforms of these kinase module subunits (CDK19, Med12L and Med13L) which associate in a mutually exclusive manner to their counterpart. It is believed that these different subunit isoforms are another mechanism for providing additional complexity in the regulatory control of transcription (Conaway and Conaway, 2011). It could be that these different isoforms function at different promoters, or different cell types to regulate distinct spatio-temporal transcription events (Conaway et al., 2005).

CDK8 is thought to negatively regulate transcription. It is thought to impart a control feedback on Mediator function. The CDK8 subunit can sterically block Mediator from interacting with Pol II and thus inhibit transcription (Elmlund et al., 2006). It does this 44 by inducing a conformational change in Mediator structure (Knuesel et al., 2009). However the module has also been implicated in various roles for gene activation (Galbraith et al., 2010). Currently, the regulation controlling CDK8 binding to Mediator is unclear, however recent studies have suggested that turnover of the Med13 subunit by means of phosphorylation and ubiquitylation are involved (Davis et al., 2013).

1.7.2 Mediator function One of the first events to take place during transcription is the formation of the PIC on the core promoter to specify the transcription start site. Although Poll II can unwind DNA, and synthesize RNA from it, it cannot alone recognize promoter sites along the DNA at which to do this. The PIC is a multi-subunit 4 MDa assembly of proteins which enables Pol II to identify which genes need to be expressed, at what time and where (Kornberg, 2005).

The PIC is assembled upon binding of the promoter region by the GTF TFIID that recognizes the TATA box sequence, which forms one of the core elements of eukaryotic promoters and enables PIC assembly. The TATA box contains a consensus sequence (TATA(A/T)(A/T)(A/G) which is recognized by the TATA binding protein subunit of TFIID. Other promoter core elements exist. These include Initiator (Chalkley and Verrijzer, 1999), and the downstream promoter element (DPE) (Burke and Kadonaga, 1997) which are recognized by another GTF (TFIIB). The common purpose of these sites is to enable the initial binding of a GTF, initiating a cascade of events leading to the recruitment of Pol II at the promoter site.

Finally TFIIH (which contains two DNA helicases; XPB, XPD) promotes adenosine triphosphate dependent unwinding of a short stretch of promoter DNA (Lu et al., 1991; Holstege et al., 1996). This enables Pol II to locate and recognize the transcription start site. It is important to note that nearly all of these studies have examined TATA promoters, which are the most highly regulated (Grünberg and Hahn, 2013).

Mediator is necessary for assembly of the PIC, and displays a functional synergy with TFIID in this process (Baek et al., 2002) as well as interacting with TFIIH to recruit it

45 to the PIC and stimulate its kinase activity (Esnault et al., 2008). TFIIH phosphorylation of Pol II is necessary in order to initiate elongation which is the next process in transcription (Akoulitchev et al., 1995). An additional role for Mediator in PIC assembly seems to involve Pol II complexes which contain the additional Gdown1 polypeptide (Pol II (G)). Gdown1 competes with TFIIF for binding to Pol II, thereby inhibiting the assembly of the PIC and repressing transcription (Jishage et al., 2012). However in the presence of Mediator, TFIIF can bind to Pol II (G) complexes and this in turn may promote the formation of active PICs.

Collectively these events all lead to the correct assembly of the PIC at the promoter region of a particular gene, but not the actual initiation of transcription. For this, the transcriptional machinery requires further input from Mediator (Kornberg, 2007). Activator proteins located at their relevant enhancer are able to physically interact with Mediator, which subsequently binds to Pol II via its CTD (Thompson et al., 1993; Kim et al., 1994; Asturias and Kornberg, 1999) to stimulate the initiation of transcription (Kornberg, 2005). This process is illustrated in Fig. 1.12. An outstanding question was how Mediator could interact with activator proteins located at up/downstream locations to the PIC. It was found that Mediator can directly interact with the cohesin loading factor Nipbl and cohesin (Kagey et al., 2010). Thus, a proposed mechanism for bringing enhancer/activator regions in proximity to promoter regions, and therefore the PIC, includes chromatin looping. This looping is stabilized by the presence of cohesin and Mediator, which have been shown to co-localize at both regions (Kagey et al., 2010).

In addition to having a vital role in the initiation of transcription, Mediator is important for elongation, which is the next discrete stage of transcription. To date studies have outlined the involvement of Mediator in several aspects of elongation. It plays a role in: overcoming the negative regulation in elongation (Malik et al., 2007), recruiting elongation and pre-mRNA processing factors (Takahashi et al., 2011) and controlling phosphorylation of the heptapeptide repeats in the Pol II CTD (Boeing et al., 2010).

Fig. 1.12. Initiation of transcription. Mediator acts as a bridge between Pol II, the PIC containing the general transcription factors, DNA regulatory elements, and gene specific transcription factors in order to co-ordinate transcription within the cell. It is also thought to be involved with chromatin looping. Taken from Alberts et al. (2002).

The head module of Mediator binds directly to Pol II via the CTD, through four heptarepeats (Robinson et al., 2012). Phosphorylation of the CTD precludes Mediator and Pol II from interacting (Søgaard and Svejstrup, 2007). It is thought that during transcription Mediator acts to control this phosphorylation of the CTD by recruiting TFIIH to the PIC and inducing its kinase activity (Esnault et al., 2008). This would then result in Mediator dissociation from Pol II, thereby permitting rapid release of promoter-proximally paused Pol II into elongation (Wong et al., 2014).

Due to these multiple roles Mediator is considered to be as necessary for transcription as Pol II. Mediator is able to assemble the general transcriptional machinery for basal transcription, modify chromatin structure for initiation, and help regulate elongation. In mammals it also co-ordinates activated transcription, by linking gene-specific transcription factors to its own individual Med subunits to regulate distinct transcriptional events. Conversely, in yeast its interactions with transcription factors predominantly regulate repressor functions in transcription.

1.7.3 Mediator mouse mutants In this thesis, the mammalian Mediator complex will be the focus, particularly the idea that individual subunits of Mediator are able to regulate gene-activated transcription during development, through interaction with specific transcription factor partners. Development is a complex and highly regulated process relying on the differential expression of particular genes at specific times in order to generate specific cell types, tissues, organs and systems. This regulation is governed by a variety of TFs, which are stimulated to modulate gene expression by means of repressing or activating transcription. The critical role of this regulation during normal development has been confirmed by various genetic studies using mouse mutants (reviewed in Hentges, 2011). How various transcriptional events, necessary for normal development, are ultimately controlled can be further elucidated through the use of animal models with disruptions in these processes. As it is known that the overall function of the Mediator complex is to enable both basal and activated transcription, and that depletion of Mediator from human nuclear extracts prevents transcription completely (Mittler et al., 2001), the use of Mediator mouse mutants provides a way to investigate the control of transcription during development.

Mutation studies on individual Mediator proteins have shown that the separate protein components of Mediator are involved in the transcriptional regulation of distinct sets of developmental genes, and that this control is crucial for the regulation of specific developmental functions, as outlined in Table 1.2. Given the essential function Mediator and its components collectively perform, it is perhaps unsurprising that mutations in these genes disrupt development. However, as discussed below the phenotypes of individual Mediator subunit mutations are distinct. This demonstrates the requirement for specific Mediator genes in specific areas of development. To date various Mediator subunit specific phenotypes have been identified in mice and mammalian cells. These include examples from each of the four Mediator modules, ‘head’ ‘middle’ ‘tail’ and ‘kinase’. Although all Mediator subunit knock outs are embryonic lethal in mice, death occurs at different developmental time points. The different knock outs result in a huge variety of phenotypes, also outlined in Table 1.2. These are discussed in more detail within the text, which also details the location within Mediator of each mutated subunit.

Mediator gene Organism Mutant phenotype Med1 M. musculus Cardiac hypoplaysia, placental insufficiency, liver haemorrhage and necrosis, growth delay; Erythropoiesis defects. Med6 D. melanogaster Third-instar larval lethality, absence of imaginal discs Med12 C. elegans Ectopic vulva cells; Defects in asymmetric cell division Med12 D. melanogaster Aberrant photoreceptor differentiation; Disrupted wing disc cell affinity; Deficient expression of Wg target genes; Impaired crystal cell development Med12 D. rerio Defects in brain, neural crest, and kidney development; Deficits in differentiation in neuronal subtypes and cardiovascular defects; Abnormal cranial neural crest, cartilage, and ear development; Endodermal differentiation defects; Reduced hindbrain cell proliferation and aberrant rhombomere boundaries Med12 M. musculus Dysregulation of Nanog target genes in ES cells; Abnormal neural tube closure, axis elongation, somitiogenesis, and cardiac development Med12 Arabidopsis Delayed expression of embryonic patterning genes, uncoupled cell division, pattern formation and morphogenesis Med13 C. elegans Embryonic lethality, vulva defects Med13 D. melanogaster Aberrant photoreceptor differentiation; Disrupted wing disc cell affinity; Deficient expression of Wg target genes; Impaired crystal cell development Med13 Arabidopsis Delayed expression of embryonic patterning genes, uncoupled cell division, pattern formation and morphogenesis Med14 D. rerio Slight reduction in retinal amacrine cell number, rod cell formation unaffected Med15 D. melanogaster Loss of wing veins, reduction in wing size Med21 M. musculus Arrest at blastocyst stage

Med23 C. elegans Larval lethality, vulva and gonad defects Med23 M. musculus Systemic circulatory failure Med24 D. rerio Lacks retinal dopanergic amacrine cells, decreased rhodopsin expressing cells; Reduction in enteric nervous system neurons Med24 M. musculus Cardiac hypoplasia, poor vascular development, thin neural tube, intrinsic cell growth defects Med25 D. rerio, M. Palatal malformations musculus Med27 D. rerio Reduction in retinal amacrine cell layer, increase in rhodopsin expressing cells Med28 NIH3T3 cells, Smooth muscle gene expression defects C2C12 cells Med31 D. melanogaster Cell fate and A/P axis defects

Table 1.2. Summary of Mediator complex developmental phenotypes. Mutations in various Mediator subunits cause a variety of developmental phenotypes revealing complex and diverse roles for Mediator during development. Adapted from Hentges (2011).

Head Module Knockdown of Med28 in NIH3T3 cells results in the over expression of specific smooth muscle cell (SMC) genes including TagIn, Actg2, and Cnn2, whilst suppression of Med28 in the myoblast cell line C2C12 results in the transdifferentiation of these cells to SMCs (Beyer et al., 2007). Therefore Med28 is thought to act as a repressor of SMC differentiation. Although this study did not use a mouse mutant, it is accepted that abnormalities in SMC growth and differentiation during development lead to severe abnormalities in the embryo.

Middle Module Med1 knock out embryos are early embryonic lethal (E11.5) and display a range of developmental phenotypes, including cardiac hypoplasia, neural tube abnormalities and cell cycle defects (Ito et al., 2000). Hypomorphic conditional knock out Med1 mice survive further into development, and thereby other phenotypes, such as hepatic necrosis, become apparent. These mice are viable to E13.5 whereupon they start to die

50 due to haematopoietic placental insufficiency (Landles et al., 2003). It was next shown that Med1 acts as a co-activator for globin transcription factor 1 (GATA-1) mediated transcription during development (Stumpf et al., 2006), in which erythropoeisis is controlled through its interaction with another Med subunit (Med27).

Med1 conditional knock out mice exhibit a specific block in erythroid development, and die relatively early in development (Stumpf et al., 2010). Furthermore T Cell conditional knock out of Med1 specifically blocks invariant natural killer T cell development, a unique lineage of T lymphocytes (Yue et al., 2011).

Knockdown of Med14 in 3T3-L1 cells impairs adipogenesis through a loss of peroxisome proliferator- activated receptor (PPARƴ) transcriptional activity (Grøntved et al., 2010). Additionally Med14 is necessary for normal retinal development in D. rerio (zebrafish) (Dürr et al., 2006). Currently no mouse model exists to study the effects of Med14 in vivo. Med21 knock out embryos fail to survive past the blastocyst stage of development (Tudor et al., 1999) whilst short interference RNA knock down affects keratinocyte differentiation (Oda et al., 2010).

Tail Module Mice lacking Med23 are embryonic lethal by E10.5, and homozygous mutant embryos appear smaller than their wild-type littermates (Balamotis et al., 2009). Besides the apparent anomalies in blood vessel formation, E9.5 null embryos exhibited a severely disorganized nervous system and defective neural tube closure. Med23 is thought to regulate neural cell differentiation by modulating the expression of Bmp4 (Zhu et al., 2015).

Med24 null mice are early embryonic lethal with a wide variety of developmental malformations, including abnormal central nervous system development, open neural tube, severe cardiac hypoplasia, poor vasculature and anaemia. Cultured fibroblasts from these mice showed defective activator driven transcriptional activation, as well as an attenuation of basal transcription (Ito et al., 2002).

Kinase Module Hypomorphic Med12 mutants fail to develop past E9.5-10.5. They are thought to be incapable of completing gastrulation due to perturbations in the Wnt/ß Catenin signaling pathway. As a consequence embryos display severe neural tube closure

51 defects, severe defects in somitogenesis and a failure of correct cardiac formation (Rocha et al., 2010). Like the Med1 subunit, Med12 is known to have physical and functional interactions with transcription factors such as Nanog (Tutter et al., 2009) and Sox10 (Vogl et al., 2013).

CDK8 null embryos are lethal by E2.5, have fragmented blastomeres and do not proceed to compaction (Westerling et al., 2007), indicating an essential role for CDK8 in cell viability and early development.

1.7.4 Mediator and development There are an ever increasing number of links emerging between Mediator subunit mutations and various human diseases, including developmental disorders and cancers. Not only is this medically relevant but it further enhances our growing understanding of the specific cellular functions of each subunit.

Developmental disorders Missense mutations in the Med12 subunit are associated with several developmental X-linked intellectual disability disorders (reviewed in Graham and Schwartz, 2013). Currently no mechanistic links have been identified, however the disorders commonly cause micrognathia, open mouth and craniosynostosis.

Hemizygous deletions of the chromosomal region encoding the Med15 subunit lead to the developmental disorders collectively referred to as DiGeorge syndrome (DGS) (Berti et al., 2001). Loss of the Med15 subunit may be involved in the aberrant development of some of the structures affected in DGS. These include tetralogy of Fallot, cleft palate and lip, and craniosynostosis.

A Med25 missense mutation has been associated with a group of neurological disorders called Charcot-Marie-Tooth disease, which results from motor and sensory neuropathy. The proposed mechanistic link is that the amino acid substitution (A335V) results in the Med25 protein being able to interact with an extended range of src Homology-3 protein binding domains, possibly affecting the specificity and degree of activation of downstream target genes of Med25 in the peripheral nervous system (Leal et al., 2009).

Congenital heart disease occurs in approximately 0.7–1.0% of live births, and thus represents the most common severe birth defect in humans (Samánek, 2000). Transposition of the great arteries accounts for 5–7% of all congenital heart disease and is marked by ventriculoarterial discordance, in which the aorta and pulmonary arteries arise from the right and left ventricles, respectively, in a reversal of the normal cardiac outflow. It requires major corrective surgery. Relatively little is known about the pathogenesis of this disease, however Med13L has been implicated in the aetiology, although there is currently no mechanistic explanation for this (Muncke et al., 2003).

In summary, the role of Mediator during development is fundamental, wide reaching and to date not fully understood. By investigating further Mediator mutant lines a more complete understanding of the requirement for Mediator in developmental processes can be gained.

1.8 Mouse Lines Used In This Project Two recessive mutant mouse lines from the mutagenesis screen outlined in section 1.6 were selected for study during this research. The causative mutation in one of these lines was known prior to this study. This was a C/T point mutation in exon 4 of the Mediator complex gene Med31. It results in embryonic lethality in homozygous embryos by E18.5 (Risley et al., 2010). This mouse line is named ‘Med31 Null’. Med31 Null homozygous embryos are considered mutant. They genotype as homozygous C57BL/6J (BL6). Med31 Null heterozygous embryos are the control littermates. They genotype as heterozygous BL6/129S5.

The second mouse line was originally named l11Jus8 by Kile et al., (2003), to identifiy it from the other lines created in the mutagenesis screen. During the course of this research three mutations in l11Jus8 were identified (Tenin et al., 2014). One of these was a T/C point mutation in exon 3 of Med31. This mutation was subsequently isolated into a distinct line during this study for phenotypic analysis of the homozygous embryos. This mouse line is named ‘Med31 Y57C’. Unlike the Med31 Null line, homozygous Med31 Y57C embryos are not embryonic lethal. Med31 Y57C homozygous embryos are considered mutant.They genotype as homozygous BL6. Med31 Y57C heterozygous embryos are the control littermates. They genotype as heterozygous BL6/129S5. 53

Both these lines were maintained as heterozygous for their mutation and the balancer region (BL6/129S5). Homozygous mutant embryos and heterozygous control embryos were generated following timed matings of heterozygous parent stock (Fig. 1.8).

1.8.1 The Med31 Null line In 2010 the causative mutation was identified following meiotic mapping of the mutant phenotype, within the balancer interval. Meiotic mapping focused the area of interest to approximately 40 genes within the balancer region. These genes were sequenced to identify the causative mutation. This was found to be a point mutation (C/T) in the fourth exon of Med31, at the 72 Mb region on chromosome 11 (Risley et al, 2010).

Med31 encodes the Med31 protein, which forms part of the multi-protein Mediator complex. Sequence analysis suggested that the point mutation should result in the insertion of an early stop codon, 26 amino acids short of the normal transcript length, producing a truncated protein. Analysis of the region excluded from the final transcript showed that no annotated functional domains would be missing from the mutant Med31 protein. qPCR analysis showed that mutants express Med31 transcripts at levels equivalent to wild type (Risley et al., 2010). However as a consequence of the truncation, the Med31 protein is rapidly degraded by the proteasome and there is no evidence of protein within homozygous mutant embryos. Therefore this mutation was considered to result in a null allele of Med31 (Risley et al., 2010). Med31 Null homozygous embryos were previously described as showing developmental delay and reduced embryonic growth.

1.8.2 The Med31 Y57C line The second mouse line selected for study from the ENU screen was named l11Jus8 by the original investigators, and homozygous embryos were embryonic lethal (Kile et al., 2003). To identify the causative mutation, next generation sequencing was performed on a single E12.5 l11Jus8 homozygous embryo. Sequence variants were confirmed by exome sequencing and additionally by Sanger sequencing on a further three l11Jus8 homozygous embryos. Three coding changes were found; Med31 (T/C), Nf1 (T/A), and Erbb2 (T/G) (Tenin et al., 2014). This point mutation in Med31 results in a Y57C missense change within the Med31 protein. This mutation was isolated into

54 a distinct mouse line for further study during the course of this research. This line is named Med31 Y57C.

1.9 Summary and Rationale for Study FGR is a common condition worldwide, and many possible causes have been identified. The precise mechanisms of these causes are still being unravelled, and further advanced by progress in the field of developmental biology. Mouse mutants with specific defects in development, which affect the growth of the fetus, are helping to refine understanding of these mechanisms.

Mediator is known to be of critical importance in numerous roles during development, as outlined in section 1.7. This ranges from fundamental roles for the whole complex during transcription, to specific roles for each of the subunits, as illustrated in Table 1.2. This work focuses on two mouse lines with mutations in the Mediator complex gene Med31. These mutant lines were previously generated during a forward mutagenesis screen. This screen introduced mutations into the genome which resulted in specific developmental defects (Kile et al., 2003).

Prior to this research, the first mouse line was known to carry a mutation in Med31 and is called the Med31 Null line. The Med31 Null line was known to have a delay in endochondral ossification, slow growth during development and defects in cell proliferation (Risley et al., 2010). During the course of this research a different mutation in Med31 was discovered, in a second mouse line (the l11Jus8 line), which had also been generated by the mutagenesis screen (outlined in section 1.6.1).

The Med31 mutation discovered in the l11Jus8 line was isolated into a distinct mouse line during this work (named the Med31 Y57C line). This line was investigated for the same phenotypes which had previously been identified in the Med31 Null line (defects in endochondral ossification and cell proliferation). The original analysis of these phenotypes, performed in the Med31 Null line by Risley et al. (2010), was also expanded.

Homozygous Med31 Null embryos are lethal by E18.5, whereas homozygous Med31 Y57C embryos are not embryonic lethal. To investigate this difference in viability placental development was examined. Given that placental function is determined by the fetally derived LZ and JZ, embryonic mutations can impact placental function (as 55 discussed in section 1.3.2). Furthermore, in mice, late stage embryonic lethality has been linked to mutations in genes which are essential for placental development and function (Hemberger and Cross, 2001). These mutations can result in placental insufficiency, which is implicated in FGR. Moreover both Med31 mutant lines display a slow growth phenotype during development, which could be a consequence of placental insufficiency. Taken together these facts provided a compelling justification to investigate placental development.

Altogether this necessitated a comparative analysis of the consequences of the two different mutant alleles of Med31 in three specific areas; placental development, skeletal development, and cell proliferation.

1.10 Aims and Objectives Loss of function mutants, in individual Mediator complex genes, have very specific developmental defects (Table 1.2). The requirement for individual subunits of Mediator during development is illustrated by the fact that most Mediator subunit knock outs in mice are early embryonic lethal. This suggests a general requirement for Mediator in multiple aspects of embryonic development, however early developmental death prevents further investigation of the requirement for each subunit throughout the whole course of development.

In contrast the Med31 Null line survives to late gestation (E18.5) and therefore the requirement for this subunit throughout development can be investigated, through study of its phenotypes. Similarly, because the missense mutation in the Med31 Y57C line is not embryonic lethal, the specific requirement for the 57th tyrosine in the normal function of Med31 throughout development can be investigated. As some of the Med31 Y57C phenotype is distinct from the Med31 Null phenotype this provides an opportunity to investigate the more subtle requirements for Med31 during development, as opposed to examining only presence/absence of protein.

The aim of investigating two different mouse lines, with two different alleles of Med31, was to reveal more about how this particular subunit of Mediator functions during development. The particular objectives of this thesis research therefore were:

Chapter Three

 Confirm that the l11Jus8 mouse line originating from the mutagenesis screen (Kile et al., 2003) carried a novel point mutation in Med31 (Tenin et al., 2014).  Isolate this mutation into a distinct mouse line (Med31 Y57C line).  Examine the consequences of the Y57C mutation on Med31 protein function.  Investigate the ability of the Med31 Y57C protein to interact with the Med7N subunit of Mediator.  Determine whether the Med31 Y57C allele when homozygous results in a slow growth phenotype.

Chapter Four- in both Med31 mutant lines

 Investigate embryonic growth rates.  Investigate the development of the endochondral growth plate using histology and immunohistochemistry (IHC).  Identify expression defects within the endochondral growth plate by quantitative polymerase chain reaction (qPCR).  Investigate intrinsic cell proliferation defects using cell growth curves and flow cytometry.  Investigate cell proliferation defects within the endochondral growth plate.  Identify expression defects in cell cycle genes by qPCR (in the Med31 Y57C line only).

Chapter Five – in the Med31 Null line

 Quantitative and qualitative histological analysis of the placenta.  Identify expression defects in key placental genes by qPCR.

The overall aim of this thesis research is therefore to describe an allelic series of Med31 in order to better understand the way this particular subunit functions during development.

Chapter Two: Materials and Methods

2.1 Med31 Null and Med31 Y57C Mice Generation of the Med31 Null line (Risley et al., 2010) and identification of the Med31 Y57C mutation (Tenin et al., 2014) have been previously described. All mice were maintained in the Biological Service Facility of the University of Manchester, UK, with local ethical approval and according to UK Home Office requirements (Home Office project PPL 40/2912). Mice were out-crossed a minimum of 6 times to 129S5/SvEvBrd (129S5) animals, although the use of the chromsome 11 balancer ensured that the candidate region on chromosome 11 remained heterozygous C57BL/6 (BL6) and 129S5. Such mice are described throughout as Med31 Null or Med31 Y57C heterozygous animals/embryos. The K14-agouti transgene integrated on the balancer chromosome allowed the identification of these adult animals without molecular genotyping. However embryo genotyping to distinguish between the 129S5 and BL6 chromosomes was performed by PCR analysis of genomic DNA using sequence tagged sites (STS) markers D11Mit4, D11Mit35, D11Mit99 and D11Mit327.

5’-3’ Sequence STS marker Forward Reverse D11Mit4 CAGTGGGTCATCAGTACAGCA AAGCCAGCCCAGTCTTCATA D11Mit35 AGTAACATGGAACATCGACGG TGCTCAGCTCTGGAGTGCTA D11Mit99 CTGTAGGTAAAATACACTTGCCG GGTGGACAGACCCTTCTGAA D11Mit327 ATTACAGTTGACTGATACCAATCAGC TCAGGCTCCACTGTGAAATG

Table 2.1. Embryonic genotyping primers. Primer sequences for various sequence tagged sites (STS) along chromosome 11 used to differentiate between the 129S5 and BL6 chromosomes. These primer sequences have been described previously (Kile et al., 2003)

For both the Med31 Null and Med31 Y57C lines heterozygous animals (which genotype as BL6 and 129S5) were bred together to generate homozygous mutant embryos which genotyped only as BL6. Embryos homozygous for the balancer chromosome are embryonic lethal and so no animals would genotype only as 129S5 58 following this cross (see section 1.6.1).

2.2 Genotyping and sequencing Med31 To isolate the Med31 Y57C mutation, genomic DNA obtained from ear punches of adult mice or yolk sacs of embryos was extracted using 50mM NaOH, and heated for 30 minutes at 95C. The genomic DNA was subjected to PCR to differentiate between BL6 (mutagenized) and 129S5 (balancer) chromosomes using the primer sequences outlined in Table 2.1. For the isolation of the Med31 Y57C line and some experimental crosses, heterozygous mice were bred to the wild type 129S5 strain to remove the balancer chromosome. Mice without the balancer chromosome were mated and their offspring genotyped to determine whether they were homozygous BL6, homozygous 129S5, or heterozygous in the balancer region. Adult mice with a chromosome that was partially of BL6 and partially of 129S5 background in the balancer region were identified as ‘recombinant’. The PCR reaction mix consisted of a total 10µL containing 5µL dH2O, 3µL redtaq PCR mix (Bioline), 1µL DNA extraction, 1µL primer mix (0.75mM each primer). PCR conditions were as follows:

95C x 5 minutes 95C x 1 minute 60C x 1 minute x35 cycles 72C x 1 minute 72C x 5 minutes

PCR products were separated on a 5% (w/v) agarose gel containing 5μL/100mL ethidium bromide by electrophoresis. Gels were visualized using a gene flash bioimaging system (Syngene). Animals were genotyped as: wild type (homozygous 129S5), Med31 Null homozygous, Med31 Y57C homozygous (both BL6 homozygous), Med31 Null heterozygous or Med31 Y57C heterozygous (both 129S5/BL6). This was determined by the presence of different sized bands on the gel which were due to differences in the lengths of the STS markers on the 129S5 and BL6 chromosomes (Fig. 2.1).

Fig. 2.1. STS marker genotyping of animals. Agarose gel showing the difference in PCR product size when animals are genotyped with an STS marker. Each marker is a different length on the two chromosomes (BL6 and 129S5). Therefore the size of the products on the gel identifies the chromosome the STS comes from. Example shown is using the marker Mit327. Mit327 is shorter in length on the BL6 chromosome than on the 129S5 chromosome and so runs further on the gel. Reading from left to right the first animal is homozygous BL6, and the second animal is heterozygous 129S5/BL6. Samples were run alongside a sample of known genotype for comparison. On this gel the comparison was with a wild type sample which genotyped as homozygous 129S5. Actual sizes were not recorded.

To confirm that the Y57C mutation had been successfully isolated, amplification of Med31 was performed using the primers listed in Table 2.2 following the same PCR conditions listed above. Products were run on a 1% (w/v) agarose gel containing 5μL/100mL safe view (NBS Biologicals) and purified from the gel by spinning over glass wool at 4193 x g for 5 minutes. An equal volume of isopropanol and 1/10 flow through volume of 3M NaOAc was added to the elution and frozen at -20C for at least an hour. DNA was pelleted following a 10 minute spin at 28341 x g and resuspended in 20L dH2O.

The next step (cycle sequencing) required 100ng eluted DNA alongside, 3L sequence buffer, 2L BigDye v3.1, 2L forward primer mixed up to 20L total volume with dH2O. PCR conditions for cycle sequencing were as follows: 96C x 1 minute 96C x 30 seconds 50C x 15 seconds x25 cycles 60C x 4 minutes

Following cycle sequencing DNA was precipitated by adding 16L dH2O, and 64L

95% ethanol, before being vortexed and left at room temperature for 15 minutes. DNA was pelleted following a 20 minute centrifugation at 28341 x g, the supernatant aspirated and the pellet washed using 70% ethanol. Following a final 10 minute centrifugation, the supernatant was aspirated and the pellet heated at 90C for 1 minute. Following precipitation, samples were sent to the Core Sequencing Facility (Faculty of Life Sciences, Manchester) for Sanger sequencing. The same protocol was used to amplify and sequence the genes outlined in section 3.2.2 within the l11Jus8 line.

5’-3’ Sequence

Forward Reverse

Med31 Exon 3 AGTAGTCCAGCGGTTGGTTG TCATTTTCCATTCCCACACA Med31 Exon 4 ATGGCAGGCTTGAGATCATT CCACTGTATGTCACCCCTATGTT Dlg4 CCCTGGGAAGTAGAAGACGG ACAGGGGTGTGGGTTAAAGT Olfr1 ATAAAGGGGGCTCTGGAAAG GGTGACTCAGGTCCCTCAAA Gemin4 TGACCAATCTTGGCACACAT GCAGGACAAACTCCTTGAGC

Synrg AGACTGCCCCACATCATCTC CTGGTGGCTGTCAATGGTTC Tbx2 CCTCCCTCTCTCCCCGTAG GGTTCCTTGCTGCACCTCT Cdk12 AAACAATAGCCCAGCACCAC GGTCAGTTTGGCTCTGAAGC

Table 2.2. PCR primers. Each primer set was used to amplify each exon/gene during PCR from genomic DNA. Using this PCR template only, the forward primer is used during the cycle sequencing reaction. Med31 primers have been described previously (Exon 4 Risley et al., 2010) (Exon 3 Tenin et al., 2014). Other primers were designed using the Primer 3 tool and sequence specificity checked using the UCSC genome browser.

al.,2.3 2010) Med31/Med7N (Tenin et al., 2014). structural modelling Modelling was performed by Professor Simon Lovell, at The University of Manchester using MAGE/PROBE (Word et al., 2000).

2.4 SDS-PAGE and Western blotting To examine Med31 Y57C homozygous embryos for the presence of Med31 protein, concentration of cell lysate was measured using the Bio-Rad protein assay (cat no. 500-0006). E14.5 embryonic cell lysate was prepared using RIPA buffer according to manufactuer instructions (Sigma Aldrich). For SDS-PAGE, 50 μg protein lysate was loaded onto a 15% polyacrylamide gel, and run at 100 Volts for 1 hour in a running

61 buffer (10x stock solution: 25mM tris, 190mM glycine, 10g SDS up to 1L with dH2O). Transfer was performed on ice for 1hour 30 minutes at 0.2 Amps using a transfer buffer (10x stock solution: 25mM tris, 190mM glycine and 200mL methanol in dH2O up to 1L). PVDF membranes were blocked overnight in 5% milk in TBS at 4 °C, incubated in primary antibody: anti-Med31 (ag9008, Proteintech), anti-β-actin (sp124, Sigma), for 1 hour at room temperature (anti-Med31 at 1:1000 dilution) (anti- β-actin at 1:250) washed and incubated in secondary antibody (anti-rabbit HRP, Sigma) at 1:20,000 dilution for 1 hour at room temperature. ECL solution (Amersham) was used to detect signal from blots which were developed for five minutes.

2.5 Embryo dissection and analysis Med31 Null and Med31 Y57C heterozygous mice were set up for timed matings to generate homozygous mutant embryos for examination. The day of the vaginal mucus plug was considered as E0.5. At the desired time point, embryos and/or placentas were dissected out of the uterus and placed on ice for 30 minutes. Prior to fixation, embryos were imaged while submerged in PBS using a Leica MZ6 microscope and DFC420 camera.

2.6 Skeletal analysis E18.5 and E17.5 embryos were dissected out of the uterus and incubated in ice cold PBS for 2 hours. They were then eviscerated by opening the abdominal wall, and skinned, with special care taken with the limbs and tail. Embryos were then placed in 100% ethanol, which was changed after 20 minutes, and left to fix for two days at room temperature. Cartilages were then stained blue by preparing a mixture of 150mg/L alcian blue in 80% ethanol/20% acetic acid and leaving to incubate overnight at room temperature. To rinse and postfix the embryos, the alcian blue was replaced with 100% ethanol and embryos were left to incubate in this overnight. To digest any soft tissues remaining on the embryo at this point a 2% KOH solution was added to the embryos which were left overnight. The embryos were then stained with alizarin red (50mg/L alizarin red in 2% KOH) for 3 hours at room temperature, to stain areas of bone. Embryos were then rinsed and cleared in 1.5% KOH overnight to ensure skeletal elements were completely visible. 1.5% KOH solution was replaced

62 with 25% glycerol (in dH2O) for storage. Embryos were transferred to 50% glycerol, 50% PBS for imaging using a Leica MZ6 microscope and DFC420 camera. Measurements were conducted using the ImageJ software measuring tool, after reproducible parameters were decided upon to act as measuring ‘landmarks’ on the skeleton as shown in Fig. 2.2.

Fig. 2.2. Landmarks for skeletal analysis. In order to perform accurate measurements of embryos and limbs, specific anatomical landmarks were chosen, which would allow reproducibility of measurements between embryos. (A) Whole embryo length measurements started from the bottom end of the parietal skull bone down to the last ossified vertebra in the embryo. (B) For whole limb measurements the starting point was the tip of the middle digit, to the end of the humerus. Individual bone lengths were measured as illustrated. Measurements taken shown by yellow points.

2.7 Histology To perform histological analysis, embryos, whole limbs and placentas were fixed in 4% PFA overnight at 4C and subsequently rehydrated with 2x washes of 50% methanol and 2x washes of 100% methanol. Samples were stored in 100% methanol

63 and frozen at least overnight at -20C. To process the tissue for embedding, the 100% methanol was replaced with 50% methanol/50% histoclear (Fisher Scientific) for at least 1 hour, this was then changed for 100% histoclear for 2 hours- in both instances samples were placed on a shaker at room temperature. Samples were then moved to a hybridiser oven (65C) in 50% paraffin wax (Invitrogen) 50% histoclear (for 1 hour) before finally being placed in 100% paraffin wax for at least an hour before embedding into plastic casettes. Wax was allowed to set at 4C for 24 hours before sectioning. For limb sections, the forelimbs were dissected from the embryo above the shoulder and the limbs orientated for sagittal sectioning. For placental sections, the placenta was orientated for saggital sectioning.

Sections of each sample were cut at 5 or 8 microns on a Finesse E microtome (Thermo Shandon) straightened thermostatically using a 42C water bath, and mounted onto polysine coated microscope slides (Thermo Scientific). Samples were placed on a 60C hot plate for 5-10 minutes and left to dry before staining.

Hamatoxylin and Eosin: Limb and placental sections were dewaxed by placing them in three changes of xylene (Genta Environmental Limited) for 5 minute intervals and subsequently rehydrated in descending alcohol series for 30 second intervals. All descending alcohol series for section 2.7 and 2.9 follow: 100%, 75%, 50% and 10% ethanol then dH2O. Slides were placed in pre filtered Harris’ haematoxylin (Thermo

Scientific) for 8 minutes, and immediately rinsed in running H2O for a further 5 minutes. Samples were then immersed in 0.5% eosin y (Thermo Scientific) for 30 seconds and washed again in H2O for 30 seconds. They were then further passed through an ascending alcohol series to dehydrate. All ascending alcohol series for section 2.7 follow: dH20, 10%, 50%, 75% and 100% ethanol. Slides were then placed in two changes of xylene and mounted with coverslips using Dpx mounting media (Fisher Scientific).

Alcian Blue and Nuclear Fast Red: E15.5 limb sections were dewaxed by placing them in three changes of xylene for 5 minute intervals and subsequently rehydrated in descending alcohol series for 30 second intervals. Slides were placed in 1% alcian blue for 5-8 minutes, and washed in running tap water for 30 seconds. Sections were then counterstained using nuclear fast red (0.1%) for 4 minutes. Slides were passed through ascending alcohol series to dehydrate, placed in two changes of xylene and

64 mounted with coverslips using Dpx mounting media.

PAS staining: To examine glyocgen content within the placenta, saggital sections were dewaxed by placing them in three changes of xylene for 5 minute intervals, and subsequently rehydrated in descending alcohol series for 30 second intervals. Slides were then stained for the presence of glycogen using methods described in the PAS kit (Sigma Aldrich).

2.8 Morphological Assessments of the Placenta To investigate placental morphology and size differences in the Med31 Null line, E14.5 saggital sections were measured in Image J using the measuring tool at the lowest magnification possible, in one mid section for 3 placentas/genotype. Whole placental area was taken as a measurement, then areas of each specific zone. The areas were marked out manually, using the measuring tool described in section 2.6 which was also used for embryo/limb measurements, and total area of each marked out zone was calculated in Image J. The total area of both the LZ and JZ in each genotype was reported independently in Fig. 5.3. However for more accurate analysis, the areas of each zone were then calculated as a percentage of the total placental area. To investigate the sinusoid dilation which was observed in E14.5 high magnification sections (x20), the circumference of the vessels were measured using the measuring tool in Image J. 120 sinusoids from 3 different placentas were measured for each genotype.

2.9 Immunohistochemistry Ki67: To examine mitotic activity, E16.5 limb sections were rehydrated in a descending alcohol series followed by 3x 5 minute PBS washes. Slides were then divided to include a control section in which the primary antibody was substituted for blocking solution only. Antigen retrieval was by boiling in sodium citrate (10mM pH6.0). Slides were boiled for 20 minutes at 20% power using a standard microwave. Slides were left to cool for 20 minutes and subsequently washed 3x with PBS.

Sections were incubated with 3% H2O2 in PBS for 10 minutes at room temperature, and rinsed with PBS. A blocking solution of 10% goat serum in PBS was applied for 1 hour at room temperature, followed by incubation with the primary antibody (drm004 Ki67, Acris) at a dilution of 1:200 (in blocking solution) overnight at 4C. Slides

65 were then washed for 5 minutes 3x with PBS-T and the secondary antibody applied (anti-rabbit biotinylated, Vector Labs) at a dilution of 1:200 in 5% goat serum for 1 hour 45 minutes at room temperature, and then washed for 5 minutes 3x with PBS-T. ABC developer kit (Vector Labs) was applied for 30 minutes, followed by 3x 5 minute wash with PBS-T. DAB substrate (Vector Labs) was then applied to the samples and left to develop until there was a colour change visible (<5 minutes).

Slides were then washed in running H2O for 10 minutes.

Col X: To examine Col X protein expression, E16.5 limb sections were rehydrated in a descending alcohol series, followed by 3x 5 minute PBS washes. Slides were then divided to include a control section in which the primary antibody was substituted for blocking solution only. Antigen retrieval was 0.5mg/mL trypsin in PBS for 12 minutes at room temperature, followed by 3x 5 minute PBS washes. Sections were incubated with 3% H2O2 in PBS for 10 minutes at room temperature, and then rinsed with PBS. Samples were blocked using 2% goat serum/PBS for 1 hour at room temperature. Primary antibody (Col X, kind gift from Professor Ray Boot-Handford) was applied at 1:500 in blocking solution overnight at 4C. Slides were then washed for 3x 5 minutes with PBS-T and the secondary antibody applied (anti-rabbit biotinylated, Vector Labs) at a dilution of 1:500 in 2% goat serum for 2 hours at room temperature, and then washed for 3x 5minutes with PBS-T. ABC developer kit (Vector Labs) was applied for 30 minutes, followed by 3x 5 minutes with PBS-T. Vector VIP substrate (Vector Labs) was then applied to the samples and left to develop until there was a colour change visible (<5minutes). Slides were then washed in running H2O for 10 minutes.

Wholemount phospho-histone H3 (PHH3): To examine mitotic activity PHH3 staining was employed. Whole E10.5 embryos were rehydrated in a descending alcohol series, followed by 3x 5 minute PBS washes. They were then incubated in 3%

H2O2 for 15 minutes at room temperature, and rinsed 3x with PBS. Embryos were then incubated in PBS-T (0.2%) for 10 minutes, followed by incubation in blocking solution (50L goat serum in 3.3mL PBS plus 1% BSA) for 30 minutes at room temperature. This was followed by incubation in the primary antibody for 1 hour at room temperature (06-570 PHH3, Millipore 1:100) in 1% BSA/PBS. Embryos were washed 3x 10 minutes in PBS then incubated in the secondary antibody for 1 hour at

66 room temperature (anti rabbit biotinylated, Vector Labs 1:250). They were then washed 3x 10 minutes in PBS and incubated in ABC substrate (Vector Labs) for 30 minutes at room temperature. Embryos were then given a final wash 3x 10 minutes in PBS and incubated in DAB substrate until there was a colour change visible (<5minutes).

2.10 HEK293 Culture and Immunocytochemistry HEK293T (HEK) cells were cultured in DMEM media supplemented with 10% FBS and 1% Penicillin/Streptomycin (P/S) mix (all Sigma Aldrich), at 37°C in a 5% CO2 humidified incubator. Under normal culture conditions cells were allowed to reach 80% confluency, rinsed with PBS, then trypsinized and split 1:7 as appropriate. Transfection plates (24 well plate, Corning) were prepared following trypsinization of a confluent flask of HEK cells. Cells were then counted manually using a heamocytometer, and resuspended at a volume of 1.67x 105 cells/well in 500µl of complete culture medium, and incubated overnight as per normal culture conditions. 1µg mammalian expression construct (Med31 FLAG- Risley et al., 2010) and (custom Med7 HALO-Promega FHC 03184) was transfected into cells using 1µl X- tremeGENE HP DNA transfection reagent (Roche) diluted into 100µl serum-free media. Plates were left at least 24 hours before examining protein expression. To detect protein expression by immunocytochemistry the culture media was removed at least 24 hours post-transfection and the cells were rinsed with PBS, then fixed using 2% PFA for 20 minutes. Cells were rinsed again and permeabilized using 0.2% Triton-X in PBS for 10 minutes, and rinsed again. Blocking was 5% goat serum in PBS for 40 minutes, followed by incubation with primary antibody at 1:100 (anti FLAG, sigma Aldrich) (anti HALO, Promega) in blocking solution for 1 hour 30 minutes at room temperature. Control wells received blocking solution only. Cells were then washed thoroughly using 3x 10 min PBS washes, before being incubated with the secondary antibody (anti rabbit biotinylated, Vector Labs 1:500) for 1 hour with fluorescent detection for the HALO construct by a further 30 minute incubation with Cy3-streptavidin (Sigma 1:500) and for the FLAG construct by using a secondary antibody directly conjugated to FITC (Sigma 1:500). Cells were then mounted onto slides using Dpx medium containing DAPI for visualization of the nucleus.

2.11 Mouse Embryonic Fibroblast Culture E14.5 embryos were harvested as previously described. Then the head, limbs, and tail were removed and the embryo eviscerated. This left a torso flank of the embryo which was placed in a 10cm plate and digested using 1mL trypsin, followed by mincing of the tissue with a sterile razor blade. Samples were then placed in the CO2 incubator at 37 C for at least 45 minutes. After digestion, 4mL DMEM (10% FBS, 1% P/S) was added to the plate, and pipetted vigorously so as to disperse the tissue and separate cells. Cells were transferred to a new plate, where a further 6-15mL culture media was added. Cells were allowed to attach for 24 hours before the media was changed to remove any debris, and then subsequently grown to confluency. Mouse embryonic fibroblast (MEF) identity was confirmed visually, as this is a common method used to prepare MEF cells (Xu, 2005; Risley et al., 2010). MEFs were trypsinized and resuspended for counting. Cell number within a 20µl sample of this suspension was counted using a haemocytometer and scaled accordingly to the final suspension volume. MEF growth curves were constructed as described previously (Risley et al., 2010).

2.12 Flow Cytometry MEF cells were cultured as normal, grown to 80% confluency and harvested following trypsinization. Cells were fixed immediately in 70% ice cold ethanol with brief vortexing, then left on ice for 30 minutes. Ice cold PBS was added, and cells were centrifuged for 5 minutes at 4193 x g to pellet. Cells were then treated with 50µL ribonuclease (100µg/mL stock, Promega) and 200µL propidium iodide was added (50µg/mL stock) which was provided by The Univeristy of Manchester flow cytometry facility, which performed the analysis. Both forward scatter and side scatter analysis was used to identify single cells.

2.13 cDNA Preparation Embryos were dissected out and genotyped as described using STS markers. Individual E14.5 limbs and E14.5 placentas were placed in RNA later solution (Life Technologies) for storage, and labelled with the appropriate genotype. To extract RNA from the tissues, 1mL TRIzol (Life Technologies) per 50-100mg tissue was

68 added and samples were homogenized in the solution. To ensure complete dissociation of nucleoprotein complexes, samples were left to stand for 5 minutes at room temperature. 0.2mL chloroform (per 1mL TRIzol) was added to the sample, allowed to stand again at room temperature for 2-15 minutes, and centrifuged at 42931 xg for 15 minutes at 4C. The upper aqueous phase was aspirated to a new tube and 0.5mL isopropanol (per 1mL TRIzol) added. The sample was left to stand for 5- 10 minutes at room temperature before centrifuging again at 42931 xg for 10 minutes at 4C. The RNA pellet was then washed with 1mL 75% ethanol by vortexing and centrifuging once more at 8217 xg for 5 minutes at 4C. The pellet was briefly air dried, then re-suspended in 40L DEPC dH2O. Complementary DNA (cDNA) was synthesized from 5µg RNA in all cases by reverse transcription using Tetro cDNA synthesis kit (Bioline) according to the manufacturer’s instructions.

2.14 Quantitative PCR (qPCR) Mouse specific primers were designed using Primer 3 and UCSC Genome Bioinformatics (http://genome.ucsc.edu). UCSC was used to confirm genomic location and sequence specificity of previously published qPCR primer sequences. Primers were synthesized by Eurogentec. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an internal control (primer sequences listed in Risley et al., 2010).

Reactions were set up using Promega qPCR Core kit for SYBR Green (GoTaq qPCR Master Mix, no A6001 & A6002, Promega). All reactions were performed with three biological and three technical triplicates. The thermal cycling parameters used using ABI 7000 SDS were: 95 °C for 10 minutes 95 °C for 15 seconds 40 cycles 60 °C for 1 minute

Each biological run produced three CT values for each gene. mRNA fold changes for each gene were calculated in the following way. Within each biological run ΔCT values within each genotype were calculated and normalised to GAPDH as a reference gene as the following: 2^ (average CT of GAPDH/ average CT of gene of

69 interest).This generated three ΔCT values from nine raw CT values. The three ΔCT values for each gene were then averaged. The fold change of mRNA expression for each gene within each genotype was then calculated using the mRNA expression for that gene in the wild type control sample. This was achieved by setting the wild type sample to 1 in the following way: average ΔCT for wild type genotype/ average ΔCT for wild type genotype. Fold change compared to wild type for other genotypes was then calculated using the following: average ΔCT for gene of interest/ average ΔCT for wild type genotype.

Reactions were performed on an ABI 7000 sequence Detection System (Applied Biosystems). All primers were validated by checking the melting curves generated from the run and by running qPCR products on an agarose gel to ensure single products of the expected size were produced.

2.15 Statistical Analysis Data were presented graphically as mean with range. Data were analysed statistically using GraphPad Prism software. Not all data were normally distributed following a Shapiro-Wilks test, and so Gaussian distribution was not assumed. Therefore to compare two data sets the non parametric test of choice was Mann Whitney, to compare three data sets Kruskall Wallis was used. Significance was at the 95% confidence level with P<0.05 and reported to 4 significant digits. For Figs. 4.10, 4.11 and 4.13 specifically the data warranted analysis by two-way ANOVA with Sidaks multiple comparisons.

2.16 Microscopy

Leica MZ6 stereomicroscope Leica DMIL fluorescence microscope (x0.63-2.0) (x2.5-40)

Fig. 3.8 (x0.63) Fig. 3.9 (x40) Fig. 3.11 (x0.63) Fig. 4.6 (x20) Fig. 3.12 (x1.0) Fig 4.7 (x20 and x40) Fig. 4.2 (x0.63 and x0.8) Fig. 4.12 (x40)

Fig. 4.3 (x0.63) Fig. 5.1 (x2.5) Fig. 4.4 (x1.0) Fig.5.3 (x5) Fig. 4.5 (x1.0) Fig. 5.4 (x20) Fig. 4.10 (x2.0) Fig. 5.5 (x20)

Table 2.3. Microscopy and magnifications. The microscope and objective used is detailed for each relevant figure.

2.17 Mouse Lines

Mouse Line Genetic background Other notes (allele1/allele2) Wild Type 129S5/129S5 Without balancer region

Med31 Null 129S5/BL6 Taken from mutagenesis screen heterozygous

Med31 Null BL6/BL6 Taken from mutagenesis screen homozygous l11Jus8 129S5/BL6 Taken from mutagenesis screen

Med31 Y57C 129S5/BL6 Isolated from l11Jus8 by KW heterozygous

Med31 Y57C BL6/BL6 Isolated from l11Jus8 by KW homozygous

Med31 Null/ l11Jus8 BL6/BL6 Offspring of Med31 Null heterozygous x l11Jus8

Med31 Null/l11Jus8 129S5/BL6 Either Med31 Null heterozygous or heterozygous controls l11Jus8

Table 2.4. Mouse lines used. Mouse lines refererred to in this thesis, the genetic background of each line and where they were obtained from. 129S5 contains the balancer chromosome (apart from in the wild type line), BL6 contains the mutagenized region. Refer to Fig. 1.6 and 1.8 for mutagenesis screen details.

al., 2010) (Tenin et al., 2014).

Gene Forward 5’-3’ Reverse 5’-3’ Amplicon size

(Base Pairs) Pthlh GTTTCTTCCTCCACCATCTG ATCTGCCCTCATCGTCTG 103

Igf2 CAAGTGGATTAATTATACGCTTTCTG AGAGGCGGGTAGGCTCAC 71 Tpbpa TCTGTAAAGTGATCCCTCCTTAATC ATTTGGGAGAGAGAAAGAAGAGGTA 202

Gcm1 AGGCAAGAAGAGCCATGAAG CCTCCCCTTCATCCGTAAG 68 Slc2a3 ATGGGGACAACGAAGGTGAC GTCTCAGGTGCATTGATGACTC 87 Mtor ACGTCACCATGGAGCTTCGA CAAATCTGCCAATTCTGGTGG 89 Ccnb1 CAGCCTCTGTGAAACCAGTGC CTGTCCTCGTTATCTATGTCCTCG 124 Sox5 CCCCTGATCCAGAGCACTTAC CCGCAATGTGGTTTTCGCT 155 Sox6 AATGCACAACAAACCTCACTCT AGGTAGACGTATTTCGGAAGGA 165 Sox9 GACAAGCGGAGGCCGAA CCAGCTTGCACGTCGGTT 187 Col2a1 CCTCCGTCTACTGTCCACTGA ATTGGAGCCCTGGATGAGCA 89 Ihh CTCTTGCCTACAAGCAGTTCA CCGTGTTCTCCTCGTCCTT 106 Runx2 AGAGTCAGATTACAGATCCCAGC TGGCTCTTCTTACTGAGAGAGG 210 Sp7 TCTTCCGGGAACAGATACAGG TGGTGTCCAATAGTCTGGTCA 126

Table 2.5. Primer sequences used for qPCR analysis. Primer sequences for Pthlh were a kind gift from Chloe Duval (University of Manchester). Primer sequences for Mtor and Ccnb1 have previously been described (Risley et al., 2010). Additional primer sequences were designed using Primer 3 software and checked for sequence specificity using the UCSC genome browser. All primers designed to work at 60°C. qPCR products were run on a 1% agarose gel to check product size. Chapter Three

Identification of a new mutant allele of Med31

3.1 Introduction The Med31 Null line was generated prior to this study (outlined in section 1.8.1). As part of this work the Med31 Y57C mouse line was established from a second mouse line (l11Jus8) generated by the ENU mutatgenesis screen (outlined in section 1.8.2). Kile et al. (2003) performed an ENU mutagenesis screen on mouse chromosome 11 to generate mouse lines with recessive developmental phenotypes. The screen produced heterozygous mice which carried a point mutation within the balancer chromosome region (shown in Fig. 1.8). These heterozygous mice were then crossed to generate mutant embryos homozygous for the point mutation (Fig. 1.8). In total fifty five different lines were generated in this way.

Thirty three of these lines had recessive developmental phenotypes which resulted in embryonic lethality. In theory for each of these lines the causative point mutation could be located within a different gene. In order to determine if this was true Kile et al. (2003) performed a complementation test (Fig. 3.1).

A complementation test is a useful genetic tool which identifies whether two mutations, which result in the same phenotype, are present within the same gene. In these tests two different recessive lines which have the same phenotype are crossed. The phenotype of the offspring is compared to the phenotype of the homozygous recessive mutants of each line. If the phenotype is absent in the offspring, this indicates that the recessive mutations carried in each parent are located within different genes. If however following a cross the phenotype remains, this indicates that the same gene is mutated in both lines.

In this instance, as each of the thirty three mutant lines was embryonic lethal, if any of the crosses produced viable offspring this would indicate that the mutations were located on different genes. As part of this test the Med31 Null line and the l11Jus8 line were crossed together, and lethality in the offspring was not rescued. This indicated that both lines carried mutations in the same gene.

Fig. 3.1. The balancer 11 mutagenesis screen complementation test. The lines generated in the screen were crossed together to check for complementation. This shows if the mutations in each line are in different genes. The Med31 Null line (red arrow) was crossed with the l11Jus8 line (blue arrow). The black box outlined in blue shows that complementation did not occur. This indicated that the mutations in the Med31 Null line and l11Jus8 line are carried in the same gene. Adapted from Kile et al. (2003).

This indicated that the l11Jus8 line carried a point mutation in Med31.To confirm this, next generation sequencing (NGS) was performed on a single homozygous l11Jus8 embryo, which identified that three coding mutations were carried in the l11Jus8 line. The mutations were: T72,213,810C in Med31 (T/C) T79,447,501A in Nf1 (T/A) T98,433,986G in Erbb2 (T/G) These are detailed in Tenin et al. (2014).The coding changes were subsequently confirmed by Sanger sequencing on three homozygous l11Jus8 embryos.

As part of this thesis research the l11Jus8 line was examined to establish whether other mutations in non-coding genes were present. Furthermore following confirmation of the l11Jus8 genotype, the T/C point mutation in Med31 was isolated to a distinct mouse line for further investigation of the effects of this mutant allele.

3.2 Results 3.2.1 Confirmation of the l11Jus8 genotype NGS identified the presence of three coding mutations in the l11Jus8 line. One consequence of using this technique is the generation of false positive data. Point mutations which don’t exist within the genome can be mistakenly identified as being present due to replication errors. To definitively establish the genotype of the l11Jus8 mouse line, several genes were checked for sequence variations which had been identified during NGS. However none of the predicted base changes were confirmed by Sanger sequencing in l11Jus8 embryos (Table 3.1) (Fig. 3.2) (Tenin et al., 2014).

3.2.2 Isolation of the Med31 Y57C mutation The l11Jus8 line carried three coding mutations located on chromosome 11 (Tenin et al., 2014). In order to study the effects of the novel Med31 mutation during development, it was isolated into a distinct mouse line (the Med31 Y57C line). Thus the Med31 Y57C line carries only the T/C point mutation in Med31.

base position annotation status change 70023993 T/C Dlg4 false positive

73395083 T/G Olfr1 false positive

76211485 C/T Gemin4 false positive

83985389 T/A Synrg false positive

85837942 G/T Tbx2 false positive

98230132 G/A Cdk12 false positive

Table 3.1. l11Jus8 false positive identification. Several coding genes were identified as false positives during NGS of the l11Jus8 line (Tenin et al., 2014).

Fig. 3.2. l11Jus8 false positive confirmation. Each residue (highlighted in yellow) was confirmed to be a false positive by visual examination of the chromatogram and corresponding sequence.

To achieve this, the three mutations carried on chromosome 11 had to be separated. This was done by outcrossing the l11Jus8 line to 129S5 animals. Offspring generated from this cross each carried different chromosomal rearrangements. These rearrangements may/may not include regions of the chromosome carrying the mutated genes, this is called recombination. It can be tracked in animals by genotyping for known markers along the chromosome.

Recombinant offspring can be identified using STS markers along the chromosome. The length of these markers differs on the chromosomes of different mouse strains. The markers can therefore be used to differentiate between two different strains. All 3 mutations within the l11Jus8 line were located on a chromosome of BL6 origin, because the original ENU mutagenesis screen mutagenized mice of this strain (Fig. 1.8). Genotyping an animal for a marker reveals the markers length. The length indicates whether the part of the chromosome on which the marker is located is from one strain or another (i.e. BL6 or 129S5). This is because the length of each of the markers on a BL6 chromosome is different to the length of the markers on a chromosome from a 129S5 animal (illustrated previously in Fig. 2.1 and next in Fig. 3.3). In the same way genotyping an animal for two separate markers reveals which strain the region of chromosome between the markers comes from.

Fig. 3.3. STS genotyping gel. Using the STS markers Mit4, Mit35 or Mit99 animal 1 genotyped as homozygous for the BL6 chromosome (red arrow) and animal 2 genotyped as heterozygous for the BL6 chromosome (red arrow) and the 129S5 chromosome (green arrow). This is because the Mit4, Mit35 and Mit99 marker sequences are shorter in length on the BL6 chromosome than on the 129S5 chromosome. Actual band sizes were not used to identify genotypes, however samples were run next to ones with an established genotype for comparison. Bracketed region indicates excess primers not important for genotyping.

Following the outcross, three markers (D11Mit4, D11Mit35 and D11Mit99) were used to identify offspring which carried both the Med31 and Nf1 mutations but did not carry the Erbb2 mutation. These offspring carried a ~10Mb fragment of the BL6 chromosome (inherited from the mutagenized parent) between the markers Mit4 and Mit35, but genotyped as 129S5 along the rest of the region, thereby excluding Erbb2 (Fig. 3.4A). Of the 46 animals genotyped for these three markers, 4 animals had this genotype. These were called recombinant one (Rec1) (examples shown in Table 3.2).

Mouse Mit4 Mit35 Mit99 806 Het Het Het 807 Het 129 129 808 129 129 129

Table 3.2. Example recombinant genotypes. Following the outcross of the l11Jus8 line to 129S5 animals recombinant offspring were genotyped. Those which carried the Med31 and Nf1 mutations would genotype as heterozygous BL6/129 at Mit4. To be sure the Erbb2 mutation was not included these animals also had to genotype as homozygous 129 with the markers Mit35 and Mit99. Animal 807 (highlighted in yellow) is an example of the correct genotype to be classed a Rec1 animal. (Total 4/46 animals). See also Fig. 3.4.

As no markers exist between Med31 and Nf1 these 4 animals were sequenced to identify if recombination had separated these genes (Fig. 3.4B). Of these 4 animals, 2 carried the Nf1 mutation, and 2 did not. All 4 were heterozygous for the Med31 mutation. The 2 animals which carried only the Med31 mutation were classified Rec2. These mice were bred as a heterozygous cross to establish the Med31 Y57C line.

Heterozygous and homozygous animals were differentiated thereafter by genotyping only for the Mit4 marker. The Med31 coding change was also subsequently confirmed by Sanger sequencing on numerous Med31 Y57C heterozygous and homozygous embryos (Fig. 3.5).

Fig. 3.4. Generation of recombinant animals and isolation of the Med31 Y57C mutation. (A) The l11Jus8 line carried three coding mutations in a 35Mb region of chromosome 11 on a C57BL/6 chromosome. This line was outcrossed to a wildtype 129S5 line to generate offspring which had natural recombination events between the two chromosomes. Animals identified as Rec1 genotyped as BL6/129 with the marker Mit4 (and therefore carried both the Med31 and Nf1 mutations), and genotyped as homozygous 129S5 along the rest of the region (and therefore did not carry the Erbb2 mutation). 4/46 animals were identified as Rec1. (B) As no marker seperates the Med31 and Nf1 genes, Rec1 animals had to be Sanger sequenced to identify recombination between these regions. Animals were identified as Rec2 if they carried the Med31 point mutation, but not the Nf1 mutation (2/4 animals). 79

Fig. 3.5. Confirmation of the Med31 Y57C mutation. Following isolation of the Med31 Y57C line by crossing of Rec 2 animals, several animals were subsequently sequenced to confirm isolation of the mutation. Included are example chromatograms showing both Med31 Y57C heterozygous (A/G) and homozygous animals (G/G). Once the line was firmly established, embryos could be identified using the STS genotyping previously described (Figs. 2.1 and 3.3).

3.2.3 Med31 Y57C protein function The T/C Med31point mutation results in a Y57C substitution within the Med31protein. This tyrosine residue is fully conserved through multiple species (Fig. 3.6A) and is located within the region of Med31which directly contacts Med7N in yeast. This region is called the binding interface. Bioinformatic analysis using the online tools Polyphen 2.0 and PROVEAN predict that the substitution is damaging and deleterious to Med31 protein function respectively (Fig. 3.6B).

Western blot analysis shows that the Med31 Y57C mutation has no effect on the expression of the Med31 protein in homozygous embryos, compared to heterozygous controls (Fig. 3.7A). This result provides support to the hypothesis that, unlike the previous Med31 null mutation, the Med31 Y57C mutation does not produce a null allele of the gene.

The 57th tyrosine in Med31 is conserved in S.cerevisiae. Structural modelling, which utilized the S.cerevisiae crystal structure of Med31, illustrates that the Y57C amino acid change within Med31 disrupts van der Waals interactions between the protein and

Med7N (Fig. 3.7B). This likely affects the ability of these two proteins to bind normally.

The Med31 Y57C allele is viable when homozygous, and there is no significant difference in the number of homozygous animals genotyped, compared to control animals, at ages three weeks and over (Fig. 3.8A). In support of this the Med31 Y57C homozygous embryos show no gross morphological phenotype at late gestation (E17.5). However these embryos do appear smaller than littermates (Fig. 3.8B).

Fig. 3.6. The Y57C residue is conserved and the mutation is damaging. (A) The Med31 Y57C mutation results in the 57th tyrosine in Med31 (highlighted in red, and conserved across multiple species) being substitured for a cysteine. (B) PolyPhen 2.0 predicts this substitution to be ‘probably damaging’ with the highest level of specificity given (Adzhubei et al., 2010). (C) PROVEAN predicts that the mutation will be deleterious to the biological function of the Med31 protein. The PROVEAN score indicates the degree of functional impact, the lowest threshold for a ‘deleterious’score is -2.5, the Y57C mutation scored -8.5 (Choi et al., 2012).

Fig. 3.7. The Med31 Y57C protein. (A) To investigate whether the mutation in the Med31 Y57C line affects the expression of Med31 protein, Med31 Y57C homozygous embryos (n=3) had protein extracted which was subjected to western blotting. This was also done in Med31 Y57C heterozygous controls (n=3). Blots were also probed with ß-actin to ensure equal loading. Med31 Y57C homozygous embryos have Med31 protein during development. (B) Med31 interacts with the Med7N subunit of Mediator. Direct van der Waals contacts (left image; white arrow) are made in the interface between the 57th Tyrosine of Med31 (red and yellow) and a residue on Med7N (green and grey). In the Y57C homozygous embryos the Y57C mutation results in a loss of van der Waals contacts between Med31 and Med7N within the binding interface (right image; white arrow).

Fig. 3.8. The Med31 Y57C embryos. (A) 40 animals were genotyped at least 3 weeks post-natally to determine viability of the Y57C allele. There was no significant deviation from expected Mendelian ratios between the genotypes (Chi Squared, P= 0.3329). (B) Late gestation Med31 Y57C homozygous embryos (E17.5) show no gross morphological abnormalities. However they are consitently smaller (right image) than heterozygous control littermates (left image). x0.63 magnifcation. This was the smallest objective available and at this magnification it was not possible to fit the Med31 Y57C E17.5 heterozygous embryos completely into the field of view (n=9 heterozygous, n=5 homozygous).

3.2.4 Med31/Med7N Interaction The next stage was to construct a mutant form of a Med31 FLAG tagged mammalian expression construct, which carried the Med31 Y57C mutation, achieved by using site directed mutagenesis. This was to determine whether the Med31 Y57C mutation affected the ability of Med31 and Med7N to physically interact with one another, as predicted by the structural modelling.

Next tagged versions of Med31 and Med7 were over-expressed in HEK293T cells (Med31-FLAG and Med7-HALO) (Fig. 3.9).This was in order to perform co- immunoprecipitation (co-IP) using anti-FLAG and anti-HALO antibodies which would reveal if the binding of Med31 and Med7N was affected by the Med31 Y57C mutation, compared to wild type. However it was observed that there were unexpected localization differences between the two proteins within HEK293T cells, possibly due to different levels of expression. Most surprisingly there was a lack of strong expression within the nucleus. This contrasts with endogenous Med31, which has high nuclear expression in HEK293T cells (Fig. 3.9E).

As the localization differences between the two tagged proteins could not be explained, and as it was necessary for them to be overexpressed in the same cellular compartments in order to perform the co-IP experiments, it was decided to collaborate with the protein expression facility (Faculty of Life Sciences) to purify soluble forms of the Med31 wild type and Med 31 Y57C proteins. The intention was to perform surface plasmon resonance experiments (SPR, Biacore) in which the soluble forms of Med31 would be passed over a chip displaying immobilized Med7. This would allow comparison of a variety of binding strengths and binding kinetics between Med31 (in both wild type and Y57C lines) and Med7. Unfortunately the protein expression facility was unable to purify high enough yields of either Med31 wild type or Y57C protein necessary for the SPR experiments following transient chemical transfection of the Med31 FLAG plasmids into HEK293 cells (Fig. 3.10). This is consistent with a recent publication, which has determined that Med31 is insoluble (Schneider et al., 2015).

3.2.5 The Med31 Y57C allele affects embryonic growth The complementation test, outlined in section 3.1, revealed that when the Med31 Null line and the l11Jus8 mouse line were crossed, lethality in the offspring was not rescued.

Fig. 3.9. FLAG tagged Med31 and HALO tagged Med7 were co-transfected into HEK293T cells. (A) Shows the expression of the HALO tag within transfected cells. (B) Shows the expression of the FLAG tag within the same cells following co-transfection. (C) A merged image of both stains. (D) Shows a no antibody control with only DAPI nuclear stain. Images representative of three co- transfections run alongside three no primary antibody control experiments. (E) Shows that endogenous Med31 (red) is strongly colocalized with DAPI in the nucleus (blue). All images x40 magnification.

Fig. 3.10. Transfection experiments conducted by the protein expression facility, University of Manchester. (A) Initially the efficiency of Med31 FLAG transient transfections was assessed and shown to be variable in two repeats. (B) shows transfection method scale up (~10 fold) after which cells were lysed and soluble material was left to bind to FLAG resin/beads overnight. Bound material was eluted to release the Med31 FLAG-fusion protein. However there was evidence of other bands present after the purification process. Molecular weight ladder (M), Solute (Sol), Flow through (FT) Wash (W).

Before the Med31 Y57C mutation was isolated from the l11Jus8 line it was important to determine whether the Med31 Y57C mutation could recover the phenotype (delay in embryonic growth) seen in the Med31 Null line. If the phenotype was not rescued it would indicate that the mutations (including Med31 Y57C) carried in the l11Jus8 line had detrimental effects on embryonic growth. Timed matings between Med31 Null heterozygous animals and l11Jus8 heterozygous animals were set up, in order to generate embryos which were of the genotype Med31 Null/111Jus8. Embryos were taken at both E17.5 and E18.5 for analysis of both late stage lethality and embryonic growth.

Animal one: Med31 Null/129S5 X Animal two: 11Jus8/129S5

Med31 Null/l11Jus8

No embryos from the two litters taken at E18.5 (n=11) genotyped as Med31 Null/l11Jus8. This suggests that lethality occurs before E18.5 in these animals. This is supported by the data from the complementation test in section 3.1.

E17.5 Med31 Null/l11Jus8 embryos have a slow growth phenotype. They are significantly smaller than either Med31 Null/129S5 or l11Jus8/129S5 control embryos (p =<0.0001) (Fig. 3.11). This suggests that the Med31 Y57C allele could be cumulatively contributing to the delay in embryonic growth in the Med31 Null/l11Jus8embryos. Data from the Med31 Y57C homozygous embryos (Fig. 3.8B) supports this hypothesis.

E17.5 Med31 Null/l11Jus8 embryos also display significant growth defects in the limbs (Fig. 3.12). The overall limb length, and length of the forelimb bones (humerus, radius and ulna) were significantly reduced compared to Med31 Null/129S5 or l11Jus8/129S5 control embryos (p =<0.0001).

Fig. 3.11. Crossing the Med31 Null line with the l11Jus8 line does not rescue the Med31 Null homozygous growth phenotype. E17.5 Med31 Null/l11Jus8 embryos (n=7) are significantly smaller than heterozygous littermates (Med31 Null/129S5 or l11Jus8/129S5 n=12). Data presented as mean with range. ****p <0.0001 (Mann-Whitney test). x0.63 magnification.

Fig. 3.12. Med31 Null/l11Jus8 embryos have limb growth defects. E17.5 Med31 Null/l11Jus8 embryos (n=7) have significantly shorter limb length and individual forelimb bone length than heterozygous littermates (Med31 Null/129S5 or l11Jus8/129S5 n=12). Data presented as mean with range. ****p <0.0001 (Mann-Whitney test). Magnification x1.0.

3.3 Discussion The overall aim of this chapter was to outline and describe a newly identified mutant mouse allele of the Mediator complex gene Med31. This is only the second mammalian allele of Med31 to be described to date, and provides an opportunity to investigate a hypomorphic allele of the gene in mammals.

Previous studies on Med31 include work in D.melanogaster, in which dMED31 was found to be a novel maternal-effect gene required for embryonic viability, and establishment of the anterior-posterior (A/P) axis (Bosveld et al., 2008). Interestingly the embryonic lethality relates to the maternal rather than the embryonic genotype, and only females which are homozygous for the hypomorphic dMED31 allele produce non- viable embryos. Additionally, a small proportion of embryos are viable, yet have defects in their abdominal segmentation patterning. This demonstrates that maternal dMED31 is required in the early drosophila embryo. The mechanisms involved include regulation of zygotic gap gene expression, which in turn drives the expression of other sets of genes (pair-rule genes) needed for segmentation of the embryo. Although these developmental processes are specific to D.melanogaster, as Med31 is one of the most highly conserved Mediator subunits, these experiments provided some of the first indications as to the potential importance of this subunit in all higher eukaryotes. The newly isolated Y57C allele of Med31 in mouse, presented here for the first time, provides new and further opportunity to study the requirement for this gene in development in higher organisms.

The Med31 Y57C allele does not alter Med31 protein expression compared to control animals, and it is therefore reasonable to conclude that the Y57C mutation does not produce a null allele, as protein is present within homozygous embryos (Fig. 3.7A). The missense mutation however does replace the 57th tyrosine in Med31 with a cysteine. This particular tyrosine residue displays extremely high conservation between multiple species (Fig. 3.6A) which indicates its central importance to the correct functioning of the protein. Analysis of human sequence variations from the 1000 genomes project reveals that this particular variant has not yet been identified in the human population. However, bioinformatic analysis using the online software PolyPhen 2.0 predicts that the Y57C substitution within Med31 would be ‘probably damaging’ to

90 protein function. This is predicted with the highest level of specificity given by this software (Adzhubei et al., 2010). Analysis with PROVEAN predicts that the Y57C mutation would be ‘deleterious’ to the biological function of the Med31 protein. The PROVEAN score indicates the degree of functional impact, the lower the score the more severe the effect. The lowest threshold for a ‘deleterious’score is -2.5, the Y57C mutation scored -8.5 (Choi et al., 2012). Together these strongly suggest that the Y57C mutation results in a hypomorphic allele of Med31 because of the loss of normal Med31 protein function.

Previously, work investigating the interaction between Med31and Med7N has been conducted in S.cerevisiae, in which the transcriptionally active submodule was first identified (Koschubs et al., 2009). The results presented in Koschubs et al. (2009) provided evidence to demonstrate that the 57th tyrosine in Med31 is located within the interface contact between the Med31 and Med7N protein. These findings support the modelling data presented in this thesis (Fig. 3.7B) in which loss of the 57th tyrosine affects the interaction of these two subunits. Currently no other experimental data provides evidence of a mutation affecting a residue located in the Med31/Med7N binding interface. Unfortunately it was impossible to obtain a solubilized form of Med31 in order to test this experimentally (Fig. 3.10). Therefore all conclusions linking the Med31 Y57C phenotype to a disruption in Med31/Med7N binding rely on a combination of the genetic analysis and modelling data presented here. The conservation of the tyrosine residue, and high total conservation of the Med31 sequence between M.musculus and S.cerevisiae (Table 1.1), means that the modelling data would prove useful for predicting the effects of this mutation across species.

Analysis of skeletal preparations from the Med31 Null/l11Jus8 embryos indicated that the Med31 Y57C allele has a detrimental effect on embryonic growth as these embryos are significantly smaller than Med31 Null heterozygous embryos, which do not carry the Med31 Y57C allele and are known to be comparable in size to wild type embryos (Fig. 3.11). Furthermore the Med31 Null/l11Jus8 embryos display limb growth defects whereas the Med31 Null heterozygous embryos do not (Fig. 3.12). Like the Med31 Null allele the Med31 Y57C allele appears to have no effect on embryonic growth when the embryos are heterozygous for the Y57C mutation. The negative effect of the Y57C allele on embryonic growth was later confirmed by analysis of Med31 Y57C

91 homozygous embryos, which appear smaller than littermates (Fig. 3.8). Interestingly the growth phenotype observed in Med31 Y57C homozygous embryos appears less severe than in Med31 Null/l11Jus8 embryos. Together these preliminary results indicated that the role of the Y57C mutation in growth and skeletal development be examined further.

The combination of the Med31 Null and Med31 Y57C alleles also results in embryonic lethality. This result was first shown in the complementation test performed by Kile et al. (2003). It is further supported by the failure to recover any Med31 Null/l11Jus8 embryos at E18.5 (n=11). Although this result was not considered significant for the number of samples in this preliminary test, it does provide further support for this conclusion as it constitutes a marked deviation from expected Mendelian ratios.

Furthermore as the Med31 Y57C allele is viable when homozygous, it provides the first opportunity for long term study for the requirement of Med31 function in adult mice. This hypomorphic allele may reveal as yet previously unidentified requirements for Med31 function in numerous adulthood processes, which would not otherwise be observable as Med31 Null heterozygous mice are comparable to wild type and Med31 Null homozygous embryos are lethal. Additionally the Med31 Y57C line could provide a model to study the consequences of delayed fetal growth in the health outcomes of adult mice. It is well known that a common feature of nonlethal fetal growth restriction is a period of post natal growth, termed ‘catch up growth’ (Ozanne and Hales, 2004). Additionally it is well understood in humans that FGR and the subsequent catch up growth are implicated in the etiology of numerous adulthood diseases, many of which are metabolic in origin, through a process termed fetal programming (de Boo and Harding, 2006). FGR has been induced in animal models using various mechanisms, including maternal calorie restriction and induced fetal hypoxia. However genetic models are few in number (Swanson and David, 2015).

3.4 Summary Described above is the identification and isolation of a new mutant allele of Med31 in mice (Med31 Y57C). A previous mutant mouse line of Med31 (Med31 Null) showed delayed development, and defects in endochondral ossification and cell proliferation (Risley et al., 2010). The next research aims were to investigate whether the Med31 Y57C mouse line showed similar phenotypes to the Med31 Null line, and investigate the mechanisms behind the phenotypes in both lines in more detail.

Chapter Four The role of Med31 in embryonic growth and cell proliferation

4.1 Introduction Both the Med31 Null and Med31 Y57C mouse lines display a slow growth phenotype in their homozygous embryos during development. This has been discussed previously in the Med31 Null line (Risley et al., 2010), and was highlighted in chapter 3 for the Med31 Y57C line following isolation of the mutation into a distinct mouse line. It was discovered that Med31 Y57C homozygous embryos appear smaller in late gestation (E17.5) than control littermates (Fig. 3.8B), and the data in chapter 3 provided evidence to support a role for this allele in embryonic growth. This is investigated further within this chapter by looking at the growth of the Med31 Y57C homozygous embryos at several points during development.

The growth defects shown by both the Med31 Null and Med31 Y57C embryos are investigated here in further detail. The growth of the limb and the rate of ossification during development are often used as a marker to identify subtle growth defects in the developing embryo (Brooke et al., 1984). It is widely acknowledged for example that maternal toxicity during development can result in reduced fetal growth, with consequent delays in ossification (Moore et al., 2013). These delays in ossification manifest on a gross level as a reduction in overall limb length. Therefore a decrease in embryonic limb length can indicate problems with fetal growth.

Presented in this chapter are data which show that both Med31 mutant lines display defects in the growth of the limbs during development. This is investigated further by morphological examination of the endochondral growth plate. Futhermore, as Mediator proteins are known to regulate distinct transcriptional events during development (outlined in section 1.7.3 and Table 1.2) expression of key genes which regulate growth plate development is examined.

Following examination of the growth plate it became clear that it was important to consider the role of cellular proliferation in the development of the Med31 Null and Med31 Y57C growth plate phenotype, particularly as the Med31 Null line was known to have proliferation defects (Risley et al., 2010).

4.2 Results 4.2.1 Med31 regulates embryonic growth Slow growth during development and subsequent FGR significantly increases the risk of complication in the neonatal period, particularly in premature births (Bernstein et al., 2000). Furthermore, the post natal ‘catch up growth’ associated with FGR is linked to the development of major adult diseases such as diabetes, hypertension, obesity and heart disease in humans (reviewed in Galjaard et al., 2013). This is particularly important when considering the phenotype of the Med31 Y57C homozygous embryos. Unlike the Med31 Null homozygous embryos, which are embryonic lethal by E18.5, these embryos are viable and develop into fertile animals (Fig. 3.8A). Preliminary examination of adult animals has shown no obvious differences in the size of Med31 Y57C homozygous mice compared to controls when X rayed 3 weeks post natally (Fig. 4.1).

Fig. 4.1. Med31 Y57C post natal measurements at 3 weeks. There are no significant differences post natally at three weeks in either body length or limb length of Med31 Y57C homozygous animals compared to Med31 Y57C heterozygous animals (n=5/genotype) (Mann-Whitney test). Animals were X-rayed. X-rays were measured manually. Data presented as mean with range.

Med31 Null homozygous embryos display a slow growth phenotype with a consequent delay in development (Risley et al., 2010). Thus it was important to determine if the Med31 Y57C homozygous embryos also showed a slow growth phenotype throughout development, with resulting developmental delay. The development of the Med31

Y57C homozygous embryo was examined at varying time points during gestation to determine if a slow growth phenotype was present in early development, and if so whether it persisted through to late gestation. At all stages the Med31 Y57C homozygous embryos were smaller in size than heterozygous littermates. However the difference in size between homozygous and heterozygous embryos in late gestation (E17.5) was attenuated compared to earlier in development, and was more variable between embryos (Fig. 4.2).

To quantify these differences in embryo size in both Med31 mutant lines, E17.5 skeletal preparations were made (Fig. 4.3). This gestation time point was chosen because it is the latest point at which recovery of Med31 Null embryos is still achievable. These skeletal preparations were then subjected to detailed measurement. Whole embryo length was chosen as a standard marker to quantify growth. However, more subtle growth changes are often most apparent in embryonic limbs, due to the rapid growth and proliferation seen within this area relative to the rest of the embryo. Therefore limb length and the lengths of the forelimb bones were also measured.

Med31 Null homozygous embryo length is significantly reduced at E17.5 compared to heterozygous controls (p=0.001) (Fig. 4.3). Med31 Y57C homozygous embryo length is not significantly different to heterozygous controls at E17.5 (Fig. 4.3). However it was observed that there was a wider range of embryo lengths within the Med31 Y57C homozygous genotype than any other genotype. It was also observed from the skeletal preparations that in both Med31 mutant lines the limb appeared to show the most marked decrease in growth relative to the rest of the embryo. Therefore the length of the whole forelimb, from the proximal end of the humerus to the distal end of the middle digit, was measured in each line. Additionally the lengths of the individual bones within the forelimb (the humerus, radius and ulna) were measured.

The Med31 Null homozygous forelimb was significantly reduced in length compared to littermate controls, as were the individual bone lengths (p=0.001) (Fig. 4.4). This was also true in the Med31 Y57C homozygous mice compared to controls, although the difference in length was less significant in each case (p=0.0045-0.007) (Fig. 4.5).

Fig. 4.2. Med31 Y57C homozygous embryos display developmental growth defects. Throughout a 7 day course of development the Med31 Y57C homozygous embryos have developed to a comparable stage as heterozygous controls. However they are visibly smaller at each developmental stage. As development progresses the difference in size becomes more variable, with some later gestation embryos appearing nearly comparable in overall size. E10.5 and E15.5 taken at x0.8 magnification. E17.5 taken at x0.63 magnification (lowest available; image not cropped). n=3/timepoint.

Fig. 4.3. E17.5 skeletal preparations. Embryos were measured using the measuring tool in Image J from the top of the parietal skull bone to the last ossified vertebra. Med31 Null homozygous (n=5) embryos are smaller in length than heterozygous controls (n=9) ***p=0.001. Med31 Y57C homozygous embryos (n=5) are not significantly (ns) different in length to heterozygous controls (n=9). Mann-Whitney test. Magnification x0.63. Data presented as mean with range.

Fig. 4.4. E17.5 Med31 Null limb preparations. Limbs were measured using the measuring tool in Image J from the proximal end of the humerus to the distal end of the middle digit. Individual bone measurements are illustrated in section 2.6. Med31 Null homozygous (n=5) forelimb, humerus, and radial lengths were significantly reduced compared to controls (n=9) (***p= 0.001) as was ulna length (***p= 0.0005). Mann-Whitney test. Magnification x1.0. Data presented as mean with range.

Fig. 4.5. E17.5 Med31 Y57C limb preparations. Limbs were measured using the measuring tool in Image J from the proximal end of the humerus to the distal end of the middle digit. Individual bone measurements are illustrated in section 2.6. Med31 Y57C homozygous (n=5) forelimb (** p= 0.007), humerus (**p= 0.006), radius (**p= 0.0065) and ulna length (**p= 0.0045) were all significantly reduced compared to heterozygous controls (n=9). Mann Whitney test. Magnification x1.0. Data presented as mean with range.

As the most significantly affected area of growth in both the Med31 mutant lines is the long bones of the limbs, it was important to determine whether this was due to a specific defect in endochondral ossification, or was a marker of a general growth delay. During mouse development the forelimb begins to develop as a mesenchymal bud at E10.5 and by E14.5 a full cartilage anlagen has been deposited. This is ossified from E15.5 onwards, as outlined in section 1.4.

4.2.2 Endochondral ossification To determine whether in both Med31 mutant lines there was a specific defect in endochondral ossification, embryonic forelimbs were sectioned and stained. Histological analysis indicated that the cells of the growth plate had properly differentiated into proliferating and hypertrophic chondrocytes, and that the endochondral growth plate appeared correctly specified in terms of its gross structure by E15.5 (Fig. 4.6). However in both Med31 mutant lines there were differences in the size and extent of growth plate development compared to controls.

E15.5 Med31 Null heterozygous animals have a clear primary ossification centre present within the radius with distinct zones of flanking hypertrophic cells and evidence of columnar/stacked proliferating chondrocytes, visible only in the periphery of images. In contrast the Med31 Null homozygous animals display no primary ossification centre, and instead the growth plate is almost entirely composed of hypertrophic cells, with extensive proliferation zones still visible (Fig 4.6). Ossification in the Med31 Null homozygous growth plates does however take place by E17.5 (Fig. 4.4). Thus it would appear that the process of endochondral ossification is occurring at a delayed rate in the Med31 Null homozygous growth plates, compared to Med31 Null heterozygous growth plates.

E17.5 Med31 Y57C homozygous embryos have a decrease in forelimb and individual bone lengths compared to Med31 Y57C heterozygous controls (Fig. 4.5). E15.5 Med31 Y57C heterozygous embryos displayed a primary ossification centre within the radius (Fig. 4.6). The overall structure and proportions of the growth plate appeared similar to the Med31 Null heterozygous growth plate. In contrast to the Med31 Null homozygous embryos, the Med31 Y57C homozygous embryos have also formed an ossification centre (Fig. 4.6). However both the hypertrophic zone and primary ossification centers

100 appear noticeably smaller in the Med31 Y57C homozygous growth plate than in the Med31 Y57C heterozygous controls. This is perhaps indicative of a delay in the development of the growth plate. However this delay is not as severe as that observed in the Med31 Null homozygous embryos.

E14.5 high magnification images of the growth plate cells reveal no overt differences in the morphology in either Med31 mutant line (Fig. 4.7). The size and appearance of both the resting chondrocytes and the proliferative chondrocytes is unremarkable in either mutant line. Importantly there is clear organization of columnar stacks of cells within the proliferative zone. The organization and proliferation of the cells ultimately drives the outgrowth of the bone. At E14.5 cells have differentiated into hypertrophy but no ossification center is present.

Although by E14.5 the hypertrophic cells have differentiated in both Med31 mutant lines, they appeared smaller in size compared to littermate controls. This is significant as it indicates that either, there is a delay in the transition to hypertrophy and therefore the cells have had less time to accrue mass or, the cells are defective in becoming hypertrophic and are somehow specified incorrectly.

To test if the latter was true the expression of Col X was examined. Col X is only expressed by hypertrophic chondrocytes of the growth plate. The Col X matrix which is deposited provides the initial framework for calcification within the primary ossification centre. Therefore if Col X expression was absent or defective in any way, this could be expected to have a detrimental effect on the process of ossification. However, although at E16.5 the hypertrophic cells remain smaller than controls, both Med31 mutant lines showed normal expression of Col X around the hypertropic cells (Fig. 4.7).

In order to determine whether there was a delay in the differentiation of hypertrophic cells in either of the Med31 mutant lines, hypertrophic cell size measurements were taken at different developmental stages and compared (Fig. 4.8). At E15.5 there was a significant decrease in the size of Med31 Null homozygous hypertrophic cells compared to heterozygous controls (p= <0.0001). This was also true in the Med31 Y57C homozygous mutants, which also showed significantly smaller hypertrophic cells than controls (p=0.007) (Fig. 4.8).

101

Fig. 4.6. The E15.5 radial endochondral growth plate. Whole E15.5 limbs were sectioned and stained with alcian blue and nuclear fast red. Med31 Null homozygous embryos (n=5) do not have a fully specified radial growth plate compared to littermate controls. The Med31 Null homozygous growth plate lacks an ossification centre (B) and instead is still mainly comprised of hypertrophic cells (H).There is some evidence that ossification is starting to occur around this point (arrowhead). P marks proliferative zones. In the Med31 Y57C homozygous radial growth plate (n=4) ossification is evident (B), however the size of each of the zones is reduced compared to heterozygous littermate controls. x20 magnification.

102

Fig. 4.7. The chondrocytes of the growth plate. H&E staining of the E14.5 growth plate reveals that both Med31 mutant lines have all types of chondrocyte cell specified correctly in the growth plate, with no overt morphological abnormalities (n=3/genotype). Immunohistochemistry on the E16.5 growth plate revelas that the hypertrophic cells express the protein Col X in all genotypes (n=3/genotype). Shown alongside is a no primary antibody (Col X) control section (n=1/genotype).

103

From E15.5 to E16.5 there was an increase in hypertrophic cell size in both Med31 Null heterozygous and Med31 Null homozygous animals (p=0.0236 and p=0.0199 respectively) (Fig. 4.8). This indicates that the Med31 Null homozygous hypertrophic cells are capable of gaining mass during development. However, when the size of E15.5 Med31 Null heterozygous cells and E16.5 Med31 Null homozygous cells was compared there was no significant difference in size (Fig. 4.8). This suggests that the mutant cells are delayed in hypertrophy by approximately one embryonic day.

Similarly from E15.5 to E16.5 there was an increase in hypertrophic cell size in both Med31 Y57C heterozygous and Med31 Y57C homozygous embryos (p= 0.001 and p=0.0103 respectively) (Fig. 4.8). This indicates that the Med31 Y57C homozygous hypertrophic cells are capable of gaining mass during development. However, unlike in the Med31 Null line, E16.5 Med31 Y57C homozygous cells are significantly larger than E15.5 Med31 Y57C heterozygous cells (p=0.0122) (Fig. 4.8). This suggests that a less severe delay than is present in the Med31 Null line, in which E16.5 homozygous cells are comparable in size to E15.5 heterozygous cells.

4.2.3 Gene expression analysis of the endochondral growth plate The next stage was to investigate whether the morphological differences, seen in both Med31 mutant lines compared to controls, were due to gene expression defects within the growth plate. Both Med31 phenotypes are distinct from other Mediator mutants. Mediator proteins are known to regulate activated transcription. Thus it was hypothesized that Med31 may regulate the expression of particular sets of genes during development. Therefore it was important to determine whether there were expression defects in any key genes which regulate growth plate development (outlined in section 1.4).

The morphological investigations of growth plate development in both Med31 mutant lines informed the choice of genes selected for qPCR analysis. As growth plate development appeared to be delayed in both lines (albeit to differing degrees) it was decided to focus on genes which control the development of the growth plate. Expression defects could explain a deviation from the normal developmental progression, which in turn would provide a mechanistic explanation for the overall phenotype.

104

Fig. 4.8. Hypertrophic cell sizes. The circumferences of hypertrophic cells within the growth plate were measured in Image J using the measuring tool. Each data point is the mean of three values. 10 cells were counted in three consecutive sections (n=3/genotype). Measurements were taken at both E15.5 and E16.5. P values corresponding to *, **, **** annotations are given within the text. Mann Whitney test. Data presented as mean with range.

105

The genes selected for expression analysis were known regulators of distinct phases of growth plate development (as reviewed in section 1.4). Sox9 and Col2a1 expression defects had previously been identified in E10.5 Med31 Null homozygous embryos (Risley et al., 2010). To investigate whether these expression defects persisted into later development, their expression was analysed in E14.5 growth plates. Med31 Y57C homozygous embryos were also examined for comparative analysis of the two alleles (Fig. 4.9). To understand fully the details of Sox9 function in the mutant growth plates the associated downstream genes Sox5 and Sox6 were also included for expression analysis in both lines (Fig. 4.9).

Both Med31 mutant lines have smaller hypertrophic cells than controls (Fig. 4.8). This could indicate a problem with the switch from proliferation to hypertrophy within the growth plate. Together PTHrP and Ihh are responsible for the maintainance of this switch.Therefore expression defects in the genes which produce these proteins may affect hypertrophy within the growth plate. The expression of Pthlh and Ihh were therefore examined (Fig. 4.9). Furthermore the transcription factor Runx2 is known to be a master regulator of the switch to hypertrophy. Defects in the expression of this protein might therefore be expected to delay the progression of hypertrophy. As such the expression of Runx2 was also investigated (Fig. 4.9).

Both the Med31 mutant lines have shorter limb bones than controls (Fig. 4.4 and 4.5). This could be due to a problem with the differentiation of osteoblast cells (discussed in section 1.4).The transcription factor Osterix is the key regulator of osteoblast differentiation. Defects in the expression of this protein might delay osteoblast differentiation, with a consequent effect on the ossification of the growth plate and therefore the length of the limb bones. As such the expression of Sp7 was also investigated (Fig. 4.9).

At E14.5 no expression defects were found in any of the genes examined in either Med31 mutant line. This suggests that the differences in growth plate development and hypertrophic cell maturation in both lines may not be due to specific defects in the expression of endochondral genes. Instead it may be indicative of a more general delay in growth.

106

Fig. 4.9. mRNA levels of key endochondral genes. Relative expression levels of each gene were calculated using cDNA prepared from E14.5 limbs (n=3/genotype). The internal reference gene was Gapdh which was similarly expressed across all genotypes. No expression defects were detected in either Med31 mutant line. Data presented as mean with SE.

107

4.2.4 Med31 regulates cell proliferation A more general growth defect implies that the embryo is following a normal growth pattern but at a reduced rate of growth. One result of this could be that the embryo cannot reach its full genetic growth potential during development, which is a feature of FGR. As previously reported E10.5 Med31 Null homozygous embryos have proliferation defects. These were noticed globally in the embryo, but were particularly noticeable in the forelimb buds (Risley et al., 2010).

In order to determine whether these cell proliferation defects were also present in the Med31 Y57C homozygous embryos, the mitotic activity of cells at E10.5 in the embryo was measured using phospho-histone H3 (PHH3) staining. This technique only marks cells in the mitotic phase of the cell cycle. There was a decrease in the mitotic activity of cells in the Med31 Y57C homozygous embryos, as compared to controls (Fig. 4.10). This was measured as the number of stained cells per unit of tissue area. Similarly to the Med31 Null homozygous embryos the reduction in the number of mitotic cells compared to controls was significant in the embryo trunk (p=0.029), but was most noticeable within the limb bud (p= 0.017) (Fig. 4.10).

Fig. 4.10. Med31 Y57C homozygous embryos have proliferation defects. Med31 Y57C homozygous limb buds have fewer proliferating cells (per mm2).There is also a decrease in mitotic activity within the trunk, but this was not significant in the embryo head (n=3) (trunk adjusted *p=0.029) (limb bud adjusted *p=0.017). Data analysed using 2-Way ANOVA with Sidaks multiple comparisons. Data presented as mean with SE. x2.0 magnification. 108

The next step was to determine whether the Med31 Null homozygous and Med31 Y57C homozygous embryos had a cell intrinsic proliferation defect. These experiments would establish if the mitotic defects observed in the mutant embryos were independent of any extrinsic factors such as environmental insult to the embryo, or nutritional deficiencies. Mouse embryonic fibroblasts (MEFs) were extracted from E14.5 embryos and cultured. The rate of growth was measured for 7 days.

A MEF growth curve had previously been generated for the Med31 Null homozygous embryos.This showed a delay in the growth of MEFs over the 7 day period compared to controls (Risley et al., 2010). A MEF growth curve was generated for the Med31 Y57C homozygous embryos. This too showed a delay in the growth of MEFs over a seven day period, when compared to MEFs extracted from control littermates (Fig. 4.11).

Fig. 4.11. Med31 Y57C MEF growth curves. One week growth curves were generated for E14.5 Med31 Y57C MEFs (n=3/genotype). There is a significant decrease at both day 5 and day 7 in the number of cells per well in the Med31 Y57C homozygous genotype compared to the Med31 Y57C heterozygous genotype (day 5 adjusted *p=0.023 day 7 adjusted ****p<0.0001). 2-Way ANOVA with Sidaks multiple comparisons. Triplicate experiments/n/genotype. Data presented as mean with SEM.

109

The MEF growth curves therefore confirmed that both Med31 mutant lines show defects in proliferation. The mutant embryonic cells divide more slowly compared to control cells. This was most noticeable within the developing limb bud in both lines. Furthermore this appears to be a mitotic-specific defect, as evidenced by the reduction in M phase specific PHH3 staining. This proliferation defect persists into later development, and is still present in both Med31 mutant lines at E14.5, as evidenced by the MEF growth curves. Furthermore this is an intrinsic defect, as culturing embryonic cells in vitro removed any potential confounding environmental/nutritional factors.

It is the proliferative capacity of chondrocytes in the proliferative zone of the growth plate which drives bone outgrowth (section 1.4). Therefore if the proliferation defects seen in the early embryo and E14.5 MEF cultures were present in the endochondral growth plate this could significantly affect outgrowth of the bones, resulting in a reduction in overall limb length. This was examined in both Med31 mutant lines.

E16.5 sections of growth plate from both lines were stained with the mitotic marker Ki67. There was a reduction in the number of positively stained cells in both Med31 Null homozygous and Med31 Y57C homozygous proliferative zones (Fig. 4.12). This demonstrates a reduction in the number of proliferating cells in each mutant growth plate.

As the data generated evidence of persistent proliferation defects within both Med31 lines, it was important to examine whether the cell cycle was progressing correctly, compared to control cells. Therefore MEFs from both lines were cultured and subjected to flow cytometry analysis. Flow cytometry can determine the proportion of cells within each phase of the cell cycle by measurement of the dye propidium iodide. This is incorporated at a known rate into DNA, the amount of which varies within the cell, between stages of the cell cycle. The amount of dye reflects the amount of nuclear material within the cell, and therefore cell cycle stage. There was a significant reduction in the percentage of Med31 Null homozygous cells in G1 compared to controls. The difference appeared to be distributed equally between S and G2/M (Fig. 4.13A). There was no significant difference across any of the phases between Med31 Y57C homozygous cells and controls (Fig. 4.13B).

110

Fig. 4.12. Ki67 staining of E16.5 growth plates. E16.5 growth plate proliferative zones were stained with Ki67 to examine mitotic activity. Note the dramatic decrease in the number of positively stained cells in the Med31 Null homozygous growth plate compared to Med31 Null heterozygous controls. There was also a noticable reduction in the number of positively stained cells in the Med31 Y57C homozygous growth plate compared to Med31 Y57C heterozygous controls, although less dramatic than that observed in the Med31 Null homozygous embryos (n=3/genotype). x40 magnification.

111

Fig. 4.13. MEF cell cycle stage comparison. E14.5 MEFs were cultured as normal and fixed for analysis by flow cytometry at 80% confluency (n=3/genotype).There was a significant decrease in the percentage of Med31 Null homozygous cells in G1 compared to heterozygous controls (adjusted *p=0.02) but no other significant changes were observed. Data analysed using 2-Way ANOVA with Sidaks multiple comparisons. Data presented as mean with SE.

112

4.2.5 Gene expression analysis of key cell cycle genes Mediator complex proteins regulate gene expression. Therefore it was decided to investigate whether there were expression defects in key cell cycle genes. As outlined in section 1.5, each phase of the cell cycle is tightly regulated by the activity of cyclin-CDK complexes. The proliferation defects observed in the Med31 mutant embryos and MEFS are mitotic. The cyclin-CDK complex responsible for the regulation of progression from G2 to M phase is cyclin B and CDK2. It had previously been reported that the Med31 Null homozygous embryos have mRNA expression defects in Ccnb1 (cyclin B) and protein expression defects in CDK2 (Risley et al., 2010). Furthermore Med31 Null homozygous embryos have reduced mRNA expression of Mtor (mTOR). mTOR is a key regulator of cell cycle and acts to couple cell growth with cell proliferation (Schmelzle and Hall, 2000). Similarly the Med31 Y57C homozygous embryos have reduced mRNA expression of both Ccnb1 and Mtor (Fig. 4.14). Defects in the expression of either of these genes might negatively impact the progression of a cell into mitosis, and reduce the proliferative capacity of the cell.

Fig. 4.14. mRNA levels of cell cycle genes. Relative mRNA expression for each gene was calculated using cDNA prepared from E14.5 embryos (n=3/genotype). The internal reference gene was Gapdh which was similarly expressed across all genotypes.There was a significant decrease in both Mtor and Ccnb1 mRNA expression within the Med31 Y57C homozygous embryos compared to heterozygous controls (*p= 0.01) Mann-Whitney test. Data presented as mean with SE.

113

4.3 Discussion The overarching aim of this chapter was to investigate the slow growth phenotype displayed by both Med31 mutant lines. By late gestation (E17.5) the Med31 Null homozygous embryos are significantly shorter in length than heterozygous controls. The Med31 Y57C homozygous embryos are not significantly different in embryo length compared to heterozygous controls. However there was a greater range in the length of Med31 Y57C homozygous embryos than in any other genotype examined. Furthermore there was a trend towards a shorter length embryo compared to Med31 Y57C heterozygous controls.

Measurement of the limbs in both Med31 mutant lines revealed significant reductions in the lengths of forelimbs compared to controls. Therefore the next major objective was to determine whether there were any endochondral defects which could be attributed to the loss of normal Med31 function in either mutant line. Forelimb and individual bone length is reduced in the Med31 Null homozygous embryos compared to controls. This is also true of the Med31 Y57C homozygous embryos. However the difference between Med31 Null homozygous embryos and controls is consistently greater than the difference between Med31 Y57C homozygous embryos and controls. This shows that the growth defect in the limbs is more exaggerated in the Med31 Null line.

To determine whether the reduction in bone length could be due to defects in the development of the growth plate, which determines the outgrowth of the limb long bones, histological sections of embryonic bone were examined at various embryonic stages.

Mouse mutants with defects in major endochondral processes provide a histological background against which to compare the Med31 mutant growth plates. Chondrogenic condensations do not develop in Sox9 knock out embryos.This identifies Sox9 as an essential transcription factor for regulation of chondrocyte differentiation (Bi et al., 1999). Sox9 haploinsufficiency in mice results in severe bending of skeletal elements, particularly the radius and ulna. It also results in an expansion of the hypertrophic zone with consequent premature ossification within the growth plate (Bi et al., 2001). This is likely due to the fact that Sox9 represses osteoblast differentiation, and therefore a reduction in Sox9 activity is likely to result in the premature

114 differentiation of osteoblast cells. Futhermore Sox9 is responsible for the expression of several other key genes which are necessary for chondrogenic development. These include Sox5, Sox6 and Col2a1.

Loss of Sox9 expression at E10.5 had been reported in the Med31 Null line. This is around the time condensations have started to form within the limb bud. However unlike the other reported phenotypes of Sox9 mutants, the limbs of Med31 Null homozygous embryos form fully, with no gross defects. Therefore it was necessary to establish when Sox9 expression was recovered. As E14.5 Med31 Null homozygous embryos express Sox9, this indicates that the mis-expression at E10.5 previously identified by Risley et al. (2010), was not in fact a complete loss of Sox9 expression during development, but likely a delay in expression. This is possibly due to a delay in the growth of the Med31 Null homozygous embryos. Furthermore expression of Sox9 in Med31 Y57C homozygous embryos was comparable to controls at E14.5.

Sox9 is also responsible for the expression of Col2a1 during endochondral ossification. In Col2a1 knock out mice there is a complete absence of endochondral bone formation (Li et al., 1995). Mutations in Col2a1 cause a spectrum of cartilage defects, ranging from moderate to severe forms of chondrodysplasias (Prockop et al. 1997). At E10.5 Med31Null homozygous embryos show a loss of Col2a1 expression within the limb bud concurrently with the loss of Sox9 expression (Risley et al., 2010). Loss of Col2a1 expression could therefore have been due to loss of Sox9 expression at this time. By E14.5 there are no expression defects in either Sox9 or Col2a1 in either Med31 mutant line. This supports the idea that previous misexpression of Col2a1 was due to the absence of Sox9 expression, as a consequence of developmental delay.

If expression of Sox9 had been delayed sufficiently, it may also have affected the expression of the other two Sox genes involved during endochondral bone development, Sox5 and Sox6. These are expressed downstream of Sox9 and collectively the trio is necessary for the initiation of chondrogenesis. Therefore misexpression could alter the normal timing of growth plate development. However by E14.5 the expression level of both Sox5 and Sox6 is comparable to controls in both Med31 mutant lines.

115

Histologically there is evidence of developmental delay in the E14.5 growth plate of both Med31 Null homozygous and Med31 Y57C homozygous embryos. The Med31 Null homozygous growth plate has failed to progress as far in endochondral ossification as controls. Supporting this there is evidence that the final subset of cells to differentiate in the endochondral pathway (hypertrophic cells) display delayed onset of differentiation compared to controls, which could explain their smaller size. The Med31 Y57C homozygous growth plate also shows some evidence of delay, although to a lesser extent than in the Med31 Null line. The growth plate is at a more similar stage of development to controls, however the primary ossfication centre is reduced in size. This is evidence of delay in the progression of growth plate development. It is further supported by the decreased size of the hypertrophic cells compared to controls. Again, however, the difference in size is markedly less pronounced than in the Med31 Null line, suggesting that the delay is less severe.

To rule out specific gene expression defects in the chondrocyte differentiation pathway, the expression levels of Ihh, Pthlh and Runx2 were examined. Ihh and PTHrP are responsible for providing the proliferative signals necessary to maintain and increase the size of the proliferative zone. Growth plates lacking either of these genes are shortened in length due to a premature switch to hypertrophy. In such instances hypertrophic cells would have more time to accrue mass. This means that proper maintainance of the proliferative capacity of this zone is one of the major determinants of overall limb length. However there was no difference in Ihh or Pthlh mRNA in either mutant line compared to controls. Furthermore limb histology did not support this idea, as the hypertrophic cells were reduced in size, indicating a delay in their maturation.

Similarly expression defects in Runx2 can affect the ability of cells to become hypertrophic. Runx2 knock out growth plates have a reduced hypertrophic zone in the ulna and radius (Kim et al., 1999). However there were no expression defects in Runx2 mRNA in either mutant line.

A further potential cause of delay in growth plate development is a delay in the differentiation of osteoblast cells, which calcify the matrix deposited by hypertrophic chondrocytes. However the expression of Sp7, which encodes the transcription factor Osterix, was not differentially expressed either.

116

There are numerous genes known to be involved in regulating growth plate development, some of these have been well described others have not. Additionally there could be other genes which are involved and have yet to be identified. The genes selected for expression analysis here were chosen due to the phenotype(s) observed in the embryonic growth plates, as discussed in section 4.2. However this analysis did not reveal any misexpression which could explain the phenotype(s). As Med31 is yet to be associated with any transcription factor partner, it would be difficult to pinpoint which other genes would be good candidates for expression analysis without performing genome wide studies in the whole growth plate. Such studies could identify genes which are misexpressed in either/both mutant lines, and reveal a wider picture for the involvement of Med31 in controlling the expression of genes necessary for endochondral ossification.

In the absence of global expression data, and because previous data indicated that Med31 may regulate cellular proliferation, and because the proliferative zone of the growth plate directly determines overall limb length, the following questions were posed:

 Can a reduction in cellular proliferation explain the slow growth phenotype observed in both Med31 mutant lines?  Can a reduction in cellular proliferation negatively affect the length of individual bones within the forelimb?  Are there any genes associated with cell cycle which are mis-expressed in both Med31 mutant lines which could reasonably account for such defects?

As Med31 Null homozygous embryos had proliferation defects (reduced mitotic activity and reduced MEF growth rates), this was examined in the Med31 Y57C homozygous embryos. It was confirmed that the Med31 Y57C homozygous embryos also have reduced mitotic activity. This cellular proliferation defect was further quantified in vitro by culturing MEFs and measuring their ability to divide in what is considered optimal nutritional conditions. The significant reduction in cell growth by day 5, in cells taken from Med31 Y57C homozygous embryos, reveals that these cells have an intrinsic defect in proliferation which is indepdendent of environmental context.

117

Taken together it is tempting to speculate that the intrinsic proliferation defect seen in both Med31 mutant lines is representative of a wider picture in which Med31 mutant cells are unable to proliferate at a rate comparable to controls, delaying the overall growth of the embryo. The difference in the severity of this delay between the two mutant lines, could be due to their differing proliferative capacities.

To address whether the proliferation defect seen in both Med31 mutant lines could detrimentally affect limb outgrowth, the proliferation of cells within the growth plate was examined. A reduction in the number of mitotic cells within the proliferative zone of the growth plate was observed in both lines. One consequence of this in the growth plate is that the rate of proliferative zone expansion is reduced. This in turn would delay the onset of hypertrophy, which would delay the whole endochondral ossification process. This would ultimately result in reduced limb growth. In support of this, the proliferation defects within the growth plate are most severe in the Med31 Null homozygous embryos, which have the greatest reduction in limb growth.

It was previously confirmed that the reduction in mitotic activity observed in the Med31 Null homozygous embryos was associated with a concurrent reduction in the expression of Ccnb1 (Risley et al., 2010). This is also true of the Med31 Y57C homozygous embryos. As the proliferation defects observed in both Med31 mutant embryos and MEFS are mitotic, and as the regulation of progression from G2 to M phase is controlled in part by cyclin B, the relationship between Med31 and cyclin B is a logical focus. As Ccnb1 is misexpressed in both Med31 mutants it is proposed that correct Med31function is necessary for correct expression of Ccnb1. Cyclin B function is necessary for the normal mitotic progression of a cell. It is therefore proposed that in both mutant lines the consequent loss of cyclin B expression is affecting the mitotic progression of these cells. In consequence the cells proliferate more slowly which in turn delays the growth of the embryos. This becomes most apparent in the limb for two reasons, 1) the development of the growth plate is dependent on the expansion of the proliferative zone and 2) the limb grows quickly during development in a short space of time relative to any other developmental landmark.

Both mutant lines also display reduced Mtor expression. mTOR is a critical regulator of cell growth, and indirectly controls the progression of a cell through the cycle,

118 ensuring that it does not divide too early. mTOR senses the nutritional environment available to the cell and, if favourable, promotes growth and division. The Med31 mutant MEFs were cultured in optimal growth conditions but still exhibited defects in proliferation. It is therefore possible that the reduced expression of Mtor could have resulted in delayed proliferation, as the favourable environmental conditions were not being relayed correctly to the cell. Currently however there is no other experimental data to support this suggestion. Together the data on cyclin B and mTOR indicate the need to further examine the expression of cell cycle related genes in the Med31 mutant lines.

4.4 Summary Described above are two mutant mouse lines with two different mutations in the Mediator complex gene Med31. Both mutant lines exhibit delayed growth and defects in cellular proliferation. Discussed so far are the many characteristics of growth delay which are common to both the Med31 Null homozygous and Med31 Y57C homozygous embryos; growth delay during development, reduction of limb length, mitotic specific proliferation defects and expression defects in the cell cycle genes Ccnb1 and Mtor. In the next section the main phenotypic difference between the two mutant lines (embryonic lethality) will be addressed. By comparing and contrasting the two mutant lines, the aim is to understand better the requirement for Med31 during development.

119

Chapter Five

The role of Med31 in Placental Development

5.1 Introduction The most notable difference observed between the Med31 Null and Med31 Y57C lines is the difference in viability of the homozygous embryos. The Med31 Null homozygous embryos are embryonic lethal by E18.5 (Risley et al., 2010). In contrast, as described in Chapter 3, the Med31 Y57C homozygous embryos are viable. The next important step therefore was to investigate the cause of lethality in the Med31 Null mice, and determine why survival was different in the Med31 Y57C line.

Mid-late gestation lethality is relatively rare amongst developmental mouse mutants. Most problems which arise during development and which can kill an embryo in utero are due to early defects in implantation or gastrulation, or become evident when the cardiovascular system starts to become functional (Bolon, 2015). Clearly the Med31 Null homozygous embryos had survived through implantation and gastrulation, and analysis of gross morphology failed to identify any cardiovascular defects which could lead to mid-late gestation death. Additionally there is no evidence of cardiac failure due to more subtle mechanisms (i.e. defects in conductivity), which would manifest as changes in the overall appearance of the heart and embryo.

Changes in placental function may also result in mid-late gestation lethality. The placenta is a transient but vital organ during development, and without it the embryo cannot survive in utero. The placenta’s most basic function is to exchange gas, nutrients and waste products between the mother and embryo, and therefore each developing embryo has its own developing placenta. Any problem with the correct development of the placenta at any embryonic stage can lead to placental insufficiency, growth restriction of the embryo, or death (Rossant and Cross, 2001).

The precise morphology of the placenta is important for its function. The placenta is an adaptive organ, which changes its morphological characteristics in response to environmental conditions to better suit the needs of the developing embryo.

120

Differences from normal placental morphology can therefore reveal difficulties/problems during pregnancy, from either the maternal or fetal side. Additionally morphological defects, due to mutations directly affecting placental development, might adversely affect fetal development, and could even result in a non viable pregnancy. It was therefore logical to investigate whether there were differences in placental development between the Med31 Null homozygous and Med31 Y57C homozygous embryos. If specific defects were identified in the Med31 Null line, and these were not present in the Med31 Y57C line, they could be further investigated as causative factors in embryonic lethality.

Presented in this chapter are data which show that the Med31 Null line, but not the Med31 Y57C line, has defects in the development of the placenta. This is investigated by morphological examination. To understand the morphological changes observed, and as Mediator proteins are known to regulate transcription, expression of key genes which regulate placental development is examined.

5.2 Results 5.2.1 Morphological defects of the Med31 Null placenta Med31 mRNA is expressed at comparatively high levels within the placenta in relation to other embryonic tissues examined (Risley et al., 2010). This suggests that Med31 is important for normal placental function. As the JZ and LZ are fetally derived it was important to investigate whether the embryonic genotype could be disrupting normal placental function, and thereby provide an explanation for lethality in the Med31 Null homozygous embryos. At E17.5 the Med31 Null homozygous placentas are much smaller than controls (Fig. 5.1A). As this decrease in size could potentially be explained by the proliferation defects which had previously been observed in both mutant lines, the Med31 Y57C homozygous placentas were also examined. There was no difference in gross morphology or size compared to littermate controls, and these placentas were indistinguishable from one another (Fig. 5.1B). It was noted however that both the Med31 Y57C homozygous and heterozygous placentas appeared larger than the Med31 Null heterozygous placenta at E17.5.

121

To investigate further the effect of the two Med31 mutations on embryonic and placental growth, fetal and placental wet weights were recorded. Additionally the fetal: placental (F: P) weight ratio was calculated by dividing fetal weight by placental weight. The F: P ratio provides an indication of the transfer capabilities of the placenta. A reduction in the F: P ratio can therefore indicate placental dysfunction (Sibley, 2009). There were significant differences in the late stage body weights, placental wet weights and F: P weight ratios of Med31 Null homozygous animals compared to controls (Fig. 5.2.) There was a 22% reduction in mean fetal weight compared to controls (0.934g vs. 1.204g respectively). There was an 11% reduction in mean placental weight compared to controls (0.088g vs. 0.099g respectively). Furthermore despite decreases in both fetal and placental wet weights, a significant decrease in the F: P ratio within the Med31Null homozygous genotype was observed. In contrast there was no significant difference in late stage fetal weights, placental wet weights and F: P weight ratios within the Med31 Y57C homozygous genotype (Fig. 5.2). This finding is consistent with previous observations in chapter 4, in which there were no significant differences in E17.5 embryo lengths for the Med31 Y57C homozygous genotype compared to controls.

Fig. 5.1. Two Med31 alleles show different placental morphology at E17.5. H&E staining of E17.5 placental sections revealed that Med31 Null homozygous placentas are much smaller than heterozygous controls. These embryos are lethal by E18.5. Med 31 Y57C homozygous placentas are indistinguishable in size and morphology from heterozygous controls. These embryos are viable. n=3/genotype. x2.5 magnification. Images not cropped. 122

Fig. 5.2. Fetal and placental weights. At E18.5 there is a reduction in Med31 Null homozygous fetal weight (***p=0.0002), placental weight (***p=0.0004), and fetal:placental weight ratio (**p=0.0054) (n=8) compared to heterozygous (n=16) and wild type (n=8) controls. Data analysed using One-way ANOVA (Kruskall Wallis test). At E17.5 there were no significant differences in fetal weight, placental weight, or fetal:placental weight ratio between Med31 Y57C homozygous embryos (n=6) and heterozygous controls (n=11) (Mann Whitney test). Data presented as mean with range.

123

As there were no apparent defects in Med31 Y57C homozygous placentas, and as these embryos are viable this line was not investigated further. To better understand the defects seen in the Med31 Null line, the morphology of the Med31 Null homozygous placentas was examined at E14.5, using paraffin/H&E sections. At low magnification Med31 Null homozygous placentas are smaller in size than controls. This is consistent with the observations later in development (E17.5). The placentas, although reduced in size, appear correctly organized into the three expected layers; decidua basalis, JZ and LZ (Fig. 5.3). However the JZ of the Med31 Null homozygous placentas appears ‘thinner’ than controls in comparable serial sections.

To investigate this finding further, and to determine if this ‘thinning’ was specific only to the JZ, the areas of the JZ and LZ were quantified on mid saggital sections (Fig. 5.3). To put this into context, areas of each zone were then calculated as a percentage of the total placental area (minus the decidua). At E14.5 Med31 Null homozygous placental area (mean 0.046mm2) was reduced 22% compared to controls (mean 0.059mm2). The mean areas of the heterozygous LZ and JZ were 59% and 41% (respectively) of total mean placental area, whilst the mean areas of the homozygous LZ and JZ were 67% and 33% (respectively) of total mean placental area (Fig 5.3). The homozygous LZ was 8% larger in relation to total area than the heterozygous LZ, and the homozygous JZ was 8% smaller in relation to total area than the heterozygous JZ.

At higher magnification morphological differences in the Med31 Null homozygous placentas became more apparent (Fig. 5.4). Within the JZ the presence of both spongiotrophoblast and glycogen cells was observed, however in the Med31 Null homozygous placentas these cells appeared somewhat smaller and more compacted than in controls. The sinusoidal spaces (which carry maternal blood) within the Med31 Null homozygous LZ appeared larger. They were significantly distended in comparison to controls (Fig. 5.4). Furthermore the S-TGCs which line these sinusoidal spaces appeared to be smaller and, perhaps, less fully developed in the Med31 Null homozygous placentas.

Glycogen cells within the JZ are thought to play a key role in late stage gestation energy release. The JZ of the Med31 Null homozygous placentas was reduced in size. One potential reason for this is that the glycogen cells were somehow defective at

124 accumulating glycogen which could contribute both to a reduction in overall JZ size, and deprive the embryo of late gestation energy, which in turn could prove lethal. Therefore it was necessary to verify whether the glycogen cells were accumulating glycogen. Periodic-acid Schiff (PAS) staining of the JZ revealed that the Med31 Null homozygous glycogen cells do accumulate glycogen (Fig. 5.5 arrowheads).

Fig. 5.3. Med31 Null homozygous placentas exhibit growth defects. E14.5 Med31 Null homozygous placentas are smaller than control littermates. All three compartments are present in both genotypes at E14.5 (DB decidua basalis, JZ junctional zone and LZ labyrinth zone) (n=3/genotype). x5 magnification. As the JZ appeared reduced in the Null homozygous placenta the areas of both the LZ and JZ were measured in Image J using the measuring tool. The area of the LZ and JZ are significantly reduced in Med31 Null homozygous placentas (*p= 0.0199 and **p= 0.0025 respectively). Mann Whitney test. Triplicate measurements/n/genotype. Not as ratio of whole placenta area. Data presented as mean with SE. Bottom: Each placental zone was calculated as % of total area. The homozygous LZ is 8% larger than controls.

125

Fig. 5.4. High magnification of the placenta reveals morphological differences. H&E staining of E14.5 placental saggital sections reveals that both glycogen cells (green arrowhead) and spongiotrophoblast cells (yellow star) are present within the Med31 Null homozygous JZ, although they appear smaller/under developed compared to heterozygous controls. The S-TGCs within the LZ of Med31 Null homozygous placentas also appear smaller, and the sinusoidal spaces appear larger (black arrow). x20 magnification. Sinusoidal diameter was measured in Image J using the measuring tool which measured sinusoidal lumen circumference. Null homozygous sinusoids are more distended than heterozygous controls (***p=0.0006). Mann Whitney test. Data presented as mean with range.

126

Fig. 5.5. PAS staining of placental glycogen. PAS staining reveals the presence of glycogen within the JZ of both Med31 Null heterozygous and homozygous placentas (concentrated purple dots, indicated by black arrows).Three placentas from each genotype, and three different sections from each placenta were stained and analyzed. JZ (junctional zone) LZ (labyrinth zone). x20 magnification.

5.2.2 Gene expression analysis of the Med31 Null placenta The morphological differences in the Med31 Null homozygous placentas might be explained by misexpression of functional/regulatory genes in the placenta as a direct consequence of lack of Med31. Therefore the expression of five key genes necessary for normal placental development and function was examined by qPCR. The expression of these genes was also examined in the Med31 Y57C homozygous placentas, which show no abnormalities in size/morphology, as a comparator.

As there were significant differences in the sizes of both the LZ and JZ areas compared to controls, the expression of Gcm1 and Tpbpa (which are considered markers of the LZ and JZ respectively) were examined. Both genes are known to be

127 essential for the correct development of their respective zones. Therefore if the morphological changes were associated with differences in mRNA levels, this could indicate direct problems with the development of these zones. However there were no significant differences in the expression of Gcm1 and Tpbpa between any of the genotypes examined (Fig. 5.6).

As the Med31 Null homozygous phenotype is embryonic lethal it could mean that the placentas are functionally impaired, and unable to support the developing embryos due to defects in nutrient exchange. Glucose is the main fetal energy source during development, and the main murine placental glucose transporter expressed in the placenta is GLUT3. If GLUT3 was absent from the placenta, glucose transport would be disrupted and in turn this may affect the survival of the embryo in utero. Therefore the expression of Slc2a3 (which encodes GLUT3) within the Med31 Null homozygous placenta was examined. No difference in Slc2a3 mRNA expression in any of the genotypes was observed (Fig. 5.6).

Med31 forms part of Mediator and therefore it has the potential to regulate the expression of any gene within the placenta. Furthermore, there is no evidence within the literature that Med31 interacts with a placental-specific transcription factor, which would help focus expression analysis. Therefore mouse mutants with similar phenotypes were next considered. The most well studied mouse mutants of placental development involve the gene Igf2. Both total embryonic knock out of Igf2 (Igf2 Null) and placental specific Igf2 knockout (Igf2 P0) mouse models display placental growth restriction. In Igf2 P0 placentas the reduction in placental growth is associated with a decrease in the passive permeability of the placenta, and placental growth restriction precedes FGR (Sibley et al., 2004). In contrast global knockout of Igf2 results in concurrent fetal and placental growth restriction (Constância et al., 2002). This phenotype is not dissimilar to the Med31 Null homozygous embryos, and therefore Igf2 was a good candidate for expression analysis.The expression of Pthlh was also investigated, as knock out mice are smaller than littermates due to reduced passive permeability within the placenta (Bond et al., 2008). However there were no expression defects detected for either gene in any of the genotypes (Fig. 5.6).

128

Fig. 5.6. mRNA expression of key placental genes. Relative mRNA expression levels for each gene were calculated using cDNA prepared from the E14.5 placenta (n=3/genotype). Internal reference gene was Gapdh which was similarly expressed across all genotypes. No expression defects were detected in either Med31 mutant line. Data shown as mean with SE.

129

5.3 Discussion The overall aim of this chapter was to investigate the difference in embryonic viability between the two mutant lines. The Med31 Null homozygous embryos are lethal by E18.5, whereas the Med31 Y57C homozygous embryos are viable. To address this point the development of the placenta was examined, as problems with placental function at any stage of development can result in embryonic lethality.

Additionally by comparing and contrasting the requirement for Med31 in embryo survival and placental development, between both mutant lines, the neccesity for different Med31 functions in these processes can be better understood. There were no morphological abnormalities in the Med31 Y57C homozygous placentas. However the Med31 Null homozygous placentas did show an interesting phenotype. Initial observations revealed that the Med31 Null homozygous placentas were significantly smaller than controls at both E14.5 and E17.5. However, this is perhaps unsurprising given that the Med31 Null homozygous embryos are themselves smaller. In contrast, whereas the Med31 Y57C homozygous embryos also display delayed growth during development there is no difference in placental size in these embryos compared to controls. The delayed growth is less apparent in the Med31 Y57C homozygous embryos than in the Med31 Null homozygous embryos. However if placental size was reduced in the Med31 Null homozygous embryos simply because the embryos themselves were smaller, then it would be reasonable also to expect some reduction in placental size in the Med31 Y57C homozygous embryos. Such a reduction was not observed.

Although the Med31 Null homozygous embryos and placentas are smaller than controls, the embryos are disproportionatly smaller than the placentas. This is illustrated by the F: P ratio of this genotype. The F: P ratio is derived by dividing the fetal weight by the placental weight, and provides an indication of the transfer capabilities of the placenta (i.e. how efficient the placenta is at providing the necessary support to the embryo) (Sibley, 2009).The decrease in this value compared to controls indicates that the Med31 Null homozygous placentas are dysfunctional in their transfer capabilities.

130

To investigate this dysfunction further the placentas were subjected to histological examination, as defects in morphology may reveal the cause of this dysfunction. One of the most apparent differences is that the JZ of the Med31 Null homozygous placentas are notably thinner than those of control littermates. In addition the two cell types which comprise the JZ (glycogen cells and spongiotrophoblast cells) looked underdeveloped compared to controls. This could be because these cell types are not developing correctly during JZ development, which in turn could lead to a reduced JZ area. In this instance the lack of Med31 during placental development would lead to cell type specific defects, which disrupt the normal morphology of the placenta, and thereby impair its overall function. Alternatively this change in JZ morphology could be the consequence of a more generalized Med31 Null homozygous placental insufficiency. In this instance the placenta has failed to develop appropriately to meet the required fetal demands, and has therefore adapted in morphology in an attempt to do so.

There is also a reduction in total LZ area in Med31 Null homozygous placentas when compared directly to controls. However the LZ of the Med31 Null homozygous placentas are 8% larger than controls in relation to their respective total placental areas. This indicates that the growth of the LZ is preserved preferentially over that of the JZ within the Med31 Null homozygous placentas. This favouring of growth of the LZ has previously been reported. It is thought to be an adaptive response, in which JZ growth is sacrificed in order to ensure that the LZ grows sufficiently in order to meet the energy demands of the developing fetus (Coan et al., 2008, 2010, 2011; Tunster et al., 2010). This fnding supports the idea that the placenta has failed to develop adequately to meet fetal demands and has therefore undergone morphological adaptation.

There was a significant increase in the distention of the maternal sinusoidal spaces within the LZ of the Med31 Null homozygous placentas. Changes in either/both fetal or maternal blood flow are implicated in placental insufficiency (Jansson and Powell, 2007).When the area of the sinusoidal spaces within the LZ is increased it reduces the surface area within the LZ for nutrient transport. This in turn can lead to placental insufficiency (Rodriguez et al., 2004). However it is also known that LZ sinusoidal spaces decrease in size as development progresses (Adamson et al., 2002). Therefore the increased sinusoidal space observed at E14.5 in Med31 Null homozygous

131 placentas, could be a direct result of a more generalized developmental delay. There is perhaps some evidence to support a delay in the development of the placenta when the appearance of the cell types of the JZ is considered. These cells appear smaller and underdeveloped compared to controls. Although the JZ itself is smaller, it should be comprised of fully developed cells if development had progressed normally. To determine whether the morphological abnormalities of the Med31 Null homozygous placentas could be explained by gene expression defects, the expression of key genes known to be involved in the normal development and function of the placenta was examined.

The reduced F: P ratio measured in the Med31 Null homozygous genotype indicates that placental transport in these animals is inefficient compared to controls. If severe enough this could easily impact embryonic development and could even result in embryonic lethality. As glucose is the main fetal energy source during development, a defect in glucose transport across the placenta could reasonably be expected to contribute to lethality. However there were no differences in the expression of Slc2a3 which encodes the main isoform of glucose transporter in the murine placenta. However there are many more placental transport functions which when disrupted could result in embryonic lethality and therefore these require further investigation.

As JZ morphology appeared abnormal in the Med31 Null homozygous placentas the expression of Tpbpa was examined. Tpbpa is expressed in the EPC and later within the JZ. Tpbpa positive cells are progenitors of many trophoblast subtypes including TGCs, spongiotrophoblasts and glycogen cells (Hu and Cross, 2011). As both the spongiotrophoblasts and glycogen cells of the JZ appeared underdeveloped in the Med31 Null homozygous placentas, the expression of Tpbpa was examined. Although there were no differences in expression levels of Tpbpa it does not rule out that JZ development is disrupted in these mutants, due to defects in the expression of other genes involved in its development. Alternatively, it could also be true that JZ development has followed a normal progression at a slower rate, which could account for the morphological differences observed.

132

The transcription factor Gcm1 is required for normal placental development. Placental specific Gcm1 knockout mice fail to form a LZ, and this is lethal by E10. Specifically its expression, in clusters of trophoblast cells at the base of the chorion, defines the initiation of branchpoints (Anson-Cartwright et al., 2000). Although Med31 Null homozygous placentas appear to have abnormal LZ morphology and distention of maternal sinusoids, this is unrelated to Gcm1 expression as there were no significant expression differences to controls. Although there were no differences in expression levels of Gcm1 it does not rule out that LZ development is disrupted in these mutants, due to defects in the expression of other genes involved in its development. Similarly to JZ development it could also be true that the LZ has developed normally at a slower rate, and this may explain some of the phenotype observed (e.g. distended sinusoids). However this does not adequately explain the overgrowth of the LZ at the expense of the JZ, which supports the F: P data, and suggests at an inherent insufficiency, resulting in an adaptive response which is independent of Med31

Other mouse mutants with developmental placental phenotypes have been described. These might provide indicators as to the details of the mechanistic explanations of the Med 31 Null homozygous phenotype. Two genes in particular were selected as good candidates to consider. Igf2 expression was measured as it is an important regulator of placental function. Specifically Igf2 null placentas show increased LZ growth at the expense of JZ growth (Sferruzzi-Perri et al., 2011) which is a major phenotype of the Med 31 Null homozygous placenta. Pthlh expression was measured as Pthlh knock out in mice results in changes to placental function. This in turn impacts on overall embryonic growth (Bond et al., 2008). Both are major phenotypes seen in Med 31 Null homozygous embryos. However there were no expression defects in Igf2 or Pthlh. Overall, further expression analysis of genes which regulate placental development and function is necessary to advance investigations.

5.4 Summary The two Med31 mutant lines share some developmental phenotypes in terms of growth during development. However they have marked differences in both embryonic viability and placental development. The Med31 Null homozygous embryos are embryonic lethal and show placental defects during development, whereas the Med31 Y57C homozygous embryos are viable and demonstrate no overt

133 placental defects. As a major morphological difference between the two Med31 lines is the difference in placental development, and as adequate placental function is necessary for life, it seems reasonable to conclude that the placental defects in the Med31 Null homozygous embryos could explain the difference in viability between these two lines. The Med31 Null homozygous placentas appear to display a morphologically adapted phenotype, although whether this is cause or effect in terms of lethality is difficult to determine based on current data. However, what is clear is that the differences in placental phenotypes do not appear to be underscored by defects in the transcription of several key genes responsible for the correct regulation of placental development.

134

Chapter Six

Discussion, Limitations, and Future Directions

6.1 Final Discussion The overall aim of this research was to examine the role of Med31 in embryonic development. Addressed specifically were the processes of growth, as marked by endochondral ossification and cellular proliferation, and placental development. By utilizing both a null and a hypomorphic allele of this gene it is possible to describe an allelic series of Med31, in which gene dosage effects, the resulting protein function and their effects on phenotype can be studied.

6.1.1 Growth, endochondral ossification and proliferation Both Med31 mutant lines exhibit delayed growth throughout development compared to their littermate controls (Risley et al., 2010) and (Fig. 4.2). In both lines the area of growth most markedly affected was forelimb growth. Limb growth during development occurs by means of endochondral ossification. Histological analysis revealed delays in several aspects of endochondral growth plate development. These delays were found to be in proportion to the overall growth restriction present in both lines. Ossification in the limb is the most sensitive marker of growth delay in a developing embryo. This is because ossification is dependent on the rapid proliferation of chondrocytes within the growth plate, and the limb grows extremely rapidly relative to the rest of the embryo. Together this explains why the Med31 Y57C homozygous limbs are significantly reduced in length compared to controls, even though overall embryo length is not. To better understand the general growth delay seen in the embryos of both lines, cellular proliferation was examined.

Alongside delayed growth in both lines there is an accompanying reduction in the rate of mitotic activity observed, in the whole embryo (Fig. 4.10), the limb bud (Fig. 4.10) the endochondral growth plate (Fig. 4.12), and in embryonic fibroblasts cultured in vitro (Fig. 4.11). It had previously been established that the reduction in mitotic acitivty observed in the Med31 Null homozygous embryos was associated with a concurrent reduction in the expression of Ccnb1 (Risley et al., 2010). This is also true in the Med31 Y57C homozygous embryos. Cyclin B is the main mitotic promoting

135 cyclin subunit. It associates with the kinase subunit (CDK1) in order to promote a cell from G2 to M. It is possible that a reduction in cyclin B was preventing the Med31 Null homozygous and Med31 Y57C homozygous cells from progressing into M at the same rate as control cells. Flow cytometry can be used to identify the percentage of cells within each cell cycle phase in a given population. If Med31 mutant cells were failing to progress into M correctly, there might be a subsequent increase in the percentage of cells in other stages of the cycle, alongside a reduction in the percentage of cells in M phase compared to controls. However there were some limitations with the technique employed to investigate this. Firstly it was not possible to differentiate between the phases of G2 and M, and therefore not possible to establish if there was a decrease in the numbers of mutant cells progressing from G2 to M using this method. Furthermore due to the fixation process this technique limits observations to a snapshot in time and thus is unable to compare rate of progression through the cycle in real time. Therefore, although there were no changes in the percentage of cells in each cell cycle stage compared to controls in either line (Fig. 4.13), this does not necessarily rule out the original hypothesis that mutant cells are failing to progress into M at the same rate as controls.

This research provides further evidence supporting the importance of Med31 in the regulation of cell cycle and proliferation. Med31 deletion in S.pombe causes down regulation of 8 genes associated with cell cycle progression (out of a total 238 downregulated) (Miklos et al., 2008). In C.albicans, transcriptome analysis of Med31 null mutants using gene set enrichment analysis showed an upregulation of genes expressed at both the G1/S and S/G2 phases of the cell cycle (Uwamahoro et al., 2012). Other approaches have looked at Mediator subunit expression within cancer cell lines. Interestingly Med31 is one of few Mediator subunits which are highly over expressed in multiple human osteosarcoma (OS) cell lines (Schiano et al., 2013). Furthermore it has been shown that microRNA 1 is capable of downregulating Med31 expression within OS cell lines, which subsequently suppressed the proliferation of these OS cells (Jiang et al., 2014). Taken together these data suggest a role for Med31 in the pathogenesis of osteosarcoma, through its positive influence on mitotic progression. This supports the data presented here which shows that mutations which inhibit the function(s) of Med31 result in mitotic defects.

136

Altogether, it would appear that in normal contexts Med31 may regulate the progression of cell cycle through G2/M. This in turn promotes normal proliferation rates throughout the embryo, resulting in normal growth.

6.1.2 Placental development A reduction in embryo growth alone is not sufficient to explain the lethality observed in the Med31 Null homozygous embryos. Furthermore, although the Med31 Y57C homozygous embryos have mitotic defects, these survive to parturition and show normal post natal development compared to control animals. Therefore it was important to determine the cause of lethality, and why this was only present within the Med31 Null homozygous embryos.

Placental development in the absence of Med31 was examined. It was observed that the definitive placenta was able to form (Fig. 5.1), which had both JZ and LZ within it. This indicates that Med31 is not necessary for the specification of these zones of the placenta. However the morpholological abnormalities and lethality seen in the Med31 Null homozygous placenta and embryo respectively suggest that Med31 function is required for normal placental function. Furthermore the late gestation lethality exhibited by the Med31 Null homozygous embryos indicates that haemochorial exchange is functional and able to support embryonic development into late gestation, though to what extent currently remains unknown, but gene expression for the glucose transporter GLUT3 was unaffected

One of the major morphological abnormalities observed in the Med31 Null homozygous placenta was a reduction in JZ area in relation to total placental size, whilst the growth of the LZ appeared to be spared. This had previously been reported in several studies looking at the effects of maternal undernutrition on placental morphology (Coan et al., 2010). Sferruzzi-Perri et al. (2011) showed that maternal malnutrition affects wild type placentas by reducing both JZ volume and the abundance of glycogen cells. Furthermore they observed the same findings in Igf2 null placentas, even when mothers were fed ad-libitum.

These results provide further evidence for the role of Igf2 as an important regulator of placental function, alongside previous data from studies showing its upregulation in response to placental growth restriction and fetal demand (Constância et al., 2005;

137

Dilworth et al., 2010). There were no changes in Igf2 expression in the Med31 Null homozygous placentas. This indicates that other, as yet unknown, mechanisms exist which act independently of Igf2 to regulate the adaptations of the growth restricted placenta.

Adaptation by the mouse placenta to support fetal growth has been demonstrated in several studies in which preservation and growth of the LZ is promoted at the expense of the JZ (Coan et al., 2008, 2010, 2011). In these situations, when fetal growth is restricted by placental insufficiency, this adaptation allows the embryo to recover its growth towards a normal trajectory. From data presented in this study it could be argued that the morphological changes seen in the Med31 Null homozygous placentas are not just an adaptive response. Instead they could be a primary pathological alteration in normal placental development due to the absence of Med31. This is because, although there are morphological features in the placenta in common with adapative response, there is no embryonic growth recovery by E18.5. This suggests either a failure of placental adaptation, or a prevention of embryonic recovery due to some other unknown cause.

One potential explanation for this could be that the Med31Null homozygous placenta adapts morphologically to compensate for reduced fetal growth but fails to adapt functionally. This functional failure could also, together with the intrinsic mitotic defects, be responsible for the reduction in fetal growth. One observation made from study of the Med31Null homozygous LZ was that the sinusoidal area was increased compared to controls (Fig. 5.4). This may indicate a problem with the continued branching morphogenesis required for optimal LZ function and haemochorial exchange. Such an increase in lumen area would decrease the surface area available for exchange, resulting in impaired fetal growth.

Taken together these results suggest that Med31 regulates the expression of key genes necessary for normal placental function. This is because the morphological changes observed in the Med31 Null homozygous placentas are likely to be adapative responses to a functional failure resulting from the loss of Med31 function. This impacts not only placental morphology but also the overall growth and survival of the Med31 Null homozygous embryo.

138

6.1.3 Med31 function during development Collectively the data presented here allow certain conclusions to be made about the requirements for Med31 function during development. By looking at the genotypes and phenotypes described it is possible to propose an allelic series which ranges from 100% protein function in wild type embryos, to 0% protein function in Med31 Null homozygous embryos, with a range of proposed values in between (Fig. 6.1).

Fig. 6.1. Med31 allelic series in the mouse. A proposed model of Med31 protein function in the various genotypes discussed in this thesis research. This model is based on the phenotypic and genetic data presented throughout all three results chapters and is discussed in detail within section 6.1.3. Above 50% normal Med31 function there is no evidence of growth defects within the embryos. Less than 25% normal protein function results in embryonic lethality.

Med31 Null heterozygous embryos have 50% of the normal Med31 protein function found in wild type embryos. However this amount of protein function is sufficient for embryonic viability, with the absence of any growth or placental phenotype. This informs somewhat about the amount of protein function present within the Med31 Y57C homozygous embryos. If Med31 Null heterozygous embryos are phenotypically normal, it is reasonable to assume that the Med31 Y57C homozygous embryo has a reduction of greater than 50% normal Med31 function. This is because otherwise there would be no growth phenotype. In such a case the Med31 Y57C homozygous embryos would resemble the Med31 Null heterozygous embryos in phenotype. This in turn provides more genetic explanation for the differences

139 observed between phenotypes, even within the same line. The differences seen between Med31 Y57C homozygous embryos and controls are often less significant than the differences observed between Med31 Null homozygous embryos and their controls. When looking at the proposed schematic for protein function in each of the genotypes, the potential difference in protein function between the Med31 Null homozygous embryo and its control is potentially far greater than that between the Med31 Y57C homozygous embryo and its control. Further support for this proposed model of protein function comes from the data generated by examining embryos with the genotype Med31 Null/Y57C. These embryos were generated as described in section 3.2.5. In these embryos it is known that the contribution of one allele (Med31 Null) is ‘0% protein function, and that the other allele (Med31 Y57C) must only contribute up to 25% protein function. As the Null/Y57C genotype is embryonic lethal it can therefore be concluded that the minimal amount of Med31 protein function for embryonic viability must be greater than 25%.

What has also become clear from this research is that Med31 control of transcription may be described in at least two different ways in mammalian development. Firstly Med31 is thought to behave like other Mediator complex proteins (Fig. 6.2A red diamond). It will bind directly to gene specific TFs in order to regulate distinct transcriptional events during development (Borggrefe and Yue, 2011). As of yet no TF which binds directly to Med31 has been identified. However studies in yeast and fungi have demonstrated that Med31 is required for the expression of Ace2 dependent genes (Miklos et al., 2008 and Uwamahoro et al., 2012). Furthermore other Mediator complex proteins have been described as behaving in this way. Within the middle module Med1 is known to interact directly with PPARƴ to regulate adipocyte differentiation. It also interacts with the TF GATA1 to regulate numerous processes including erythropoesis, cardiac and megokaryocyte development, amongst many others (reviewed in Yin and Wang, 2014). Secondly evidence presented in this research supports a role for the Med31/Med7N submodule in mammalian transcription (Fig. 6.2A yellow diamond). This had previously only been described in yeast.

To integrate these two ideas and propose a framework for the two Med31 mutant lines the following is proposed. In wild type context Med31 acts to regulate transcription

140

via the two mechanisms described above. Firstly through direct binding with a TF partner(s), and secondly in conjunction with Med7N to interact with a further distinct set of TF(s). Presumably both mechanisms exist in order to provide increased levels of transcriptional diversity depending on the spatial and temporal contexts within development, as well as providing for potential differences relating to tissue specificity.

In the two Med31 mutant lines, each has its own unique allele of Med31 which affects overall Med31 function in a distinct way. Ultimately this results in some phenotypes which are similar to and some which are distinct from each other. The Med31 Null homozygous embryo has an absence of Med31 during development, and consequently will be unable to regulate transcription in either of the two ways outlined above (Fig. 6.2B). In contrast the Med31 Y57C homozygous embryo does have Med31 present during development. It is hypothesized that the Y57C mutation significantly disrupts the interaction between Med31 and Med7N. This thereby affects the transcriptional activity of this submodule. However some Med31 function is thought to be retained. It is proposed that Med31 is still able to regulate some transcription, through direct interaction with TFs independently of Med7N (Fig. 6.2B). This provides an explanation for the differences in placental phenotype, and embryonic viability within these embryos compared to the Med31 Null homozygous embryos.

The phenotypes observed in the two Med31 mutant lines lead to the interesting proposal that the Med31/Med7N subunit regulates the transcription of genes which control cellular proliferation in the developing embryo. Identified genes so far are Mtor and Ccnb1. This explains why there are proliferation defects in both Med31 lines (Fig. 6.2). Additionally, in this model Med31 directly interacts with its own distinct set of TF(s) (Fig. 6.2 yellow diamond) enabling it to regulate the transcription of genes necessary for placental development independently of the Med31/Med7N submodule. This would explain why there is a placental phenotype in the Med31 Null homozygous embryos but not in the Med31 Y57C homozygous embryos (Fig. 6.2).

As yet it has not been possible to identify any genes which are misexpressed in the Med31 Null homozygous placenta. However, only a handful were selected for

141

Fig. 6.2. Two alleles of Med31 provide a model to study its function during development. (A) It is proposed that like other Mediator complex proteins Med31 is able to interact with gene specific TFs (red diamond) in order to regulate distinct transcriptional events. Secondly it is proposed that the Med31/Med7N submodule is conserved from yeast and is transcriptionally active in mammals, interacting with its own subset of TF partners (yellow diamond). (B) Med31 Null homozygous embryos completely lack Med31. This disrupts transcription through both pathways shown in (A). Med31 Y57C homozygous embryos have a mutated form of Med31. The Y57C mutation is thought to disrupt the ability of Med31 and Med7N to interact. This disrupts transcription via this submodule (yellow diamond). However it is thought direct Med31 targets (red diamond) are normally expressed. This leads to some phenotypes which are similar in both Med31 mutant lines, and some phenotypes which are unique to the Med31 Null homozygous embryos.

142 analysis in this work, and therefore further gene expression study is required. Furthermore, to understand fully whether the placental phenotype is contributing to lethality, functional studies on placental transport should be conducted, which may in turn inform gene expression analysis.

Overall, this research has contributed to a better appreciation of the value of utilizing multiple alleles for understanding gene function in mouse embryonic development. Furthermore there is clearly added value in studying not only null alleles or gene knock outs, but also missense mutations which may reveal more subtle changes in gene/protein function.

To date the majority of work on Mediator has been conducted in yeast or in human cell lines. This is particularly true for the Med31 subunit. First identified as part of the yeast Mediator complex as the subunit Soh1 (Linder and Gustafsson, 2004), Med31 is a non-essential gene in yeast (Szilagyi et al., 2002). However Med31 knock down in C. elegans results in larval lethality (Clayton et al., 2008). Transcriptome analysis from Med31 null yeast strains first introduced the possibility that Med31 is not required for the expression of all genes transcribed by Pol II, but rather the regulation of large subsets of genes (Miklos et al., 2008). This was later backed up by expression analysis in C.albicans which showed that 7% of the genome is misexpressed following deletion of Med31. This included genes responsible for regulating cell cycle progression (Uwamahoro et al., 2012). However, gene expression studies and phenotypic analysis in yeast and fungi have not previously allowed for the appreciation of subtle differences in Med31 function(s). As a consequence of this research it is now possible to start to understand the specific requirements for Med31 function in various developing tissues.

6.2 Future Directions There is still much work to be done to fully appreciate the complexity of Mediator function, not only in development but in disease and normal human health. Currently many studies of Mediator have focused on individual subunits, and the ways in which these subunits directly interact with TF partners. However, much more work is necessary to reveal the molecular basis of the cooperation between intracellular signalling and transcriptional regulation by Mediator. In addition, in light of evidence that Mediator is likely to be present in cells in multiple functionally distinct forms,

143 one major aim of future studies will be to identify the repertoire of Mediator assemblies and subassemblies, and to elucidate their functions.

To continue this thesis research there are some major areas to be investigated:

1. Investigate protein interactions between Med31 Y57C and Med7N, and determine experimentally if the the Y57C mutation disrupts the binding of these proteins.

2. Global analysis of gene expression in Med31 Null homozygous and Med31 Y57C homozygous embryonic tissues and placentas.

3. ChIP experiments to determine the transcriptional targets of Med31 in both Med31 wild type and Med31 Y57C homozygous embryos. It is possible some differences would be observed if the Med Y57C protein is still able to associate with DNA.

4. An investigation of overall Mediator assembly in Med31 Null homozygous and Med31 Y57C homozygous embryos. Perhaps the association of other Mediator proteins depends on the presence of Med31 within Mediator.

5. To determine the cause of Med31 Null homozygous embryonic lethality several placental function experiments should be undertaken. Primarily these should concern the efficiency of haemochorial exchange. Firstly this should be by quantification of glucose and amino acid transport to determine the efficiency of placental function. Further functional studies may be indicated following genome wide expression analysis of the placenta.

144

References

Abad V, Meyers JL, Weise M, Gafni RI, Barnes KM, Nilsson O, Bacher JD, Baron J (2002) The role of the resting zone in growth plate chondrogenesis. Endocrinology. 143(5):1851-7. Adamson SL, Lu Y, Whiteley KJ, Holmyard D, Hemberger M, Pfarrer C, Cross JC (2002) Interactions between trophoblast cells and the maternal and fetal circulation in the mouse placenta. Dev Biol. 250(2):358-73. Adzhubei IA, Schmidt S, Peshkin L, Ramensky VE, Gerasimova A, Bork P, Kondrashov AS, Sunyaev SR (2010) A method and server for predicting damaging missense mutations. Nat Methods. 7(4): 248-9. Akiyama H, Chaboissier MC, Martin JF, Schedl A, de Crombrugghe B (2002) The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev. 16(21): 2813-28. Akoulitchev S, Mäkelä TP, Weinberg RA, Reinberg D (1995) Requirement for TFIIH kinase activity in transcription by RNA polymerase II. Nature. 377(6549):557-60. Alberts B, Johnson A, Lewis J (2002) Molecular Biology of the Cell. 4th edition. New York: Garland Science. Allen BL, Taatjes DJ (2015) The Mediator complex: a central integrator of transcription. Nat Rev Mol Cell Biol. 16(3):155-66. Amizuka N, Warshawsky H, Henderson JE, Goltzman D, Karaplis AC (1994) Parathyroid hormone-related peptide-depleted mice show abnormal epiphyseal cartilage development and altered endochondral bone formation. J Cell Biol. 126(6):1611-23. Ansari SA, He Q, Morse RH (2009) Mediator complex association with constitutively transcribed genes in yeast. Proc Natl Acad Sci U S A. 106(39):16734-9. Anson-Cartwright L, Dawson K, Holmyard D, Fisher SJ, Lazzarini RA, Cross JC (2000) The glial cells missing-1 protein is essential for branching morphogenesis in the chorioallantoic placenta. Nat Genet. 25:311 – 314. Ashburner M (1989). Drosophila: A Laboratory Handbook and Manual. Cold Spring Harbor Laboratory. Asturias FJ, Kornberg RD (1999) Protein crystallization on lipid layers and structure determination of the RNA polymerase II transcription initiation complex. J Biol Chem. 274(11):6813-6. Aszodi A, Bateman JF, Gustafsson E, Boot-Handford R, Fässler R (2000) Mammalian skeletogenesis and extracellular matrix: what can we learn from knockout mice? Cell Struct Funct. 25(2):73-84.

145

Baek HJ, Malik S, Qin J, Roeder RG (2002) Requirement of TRAP/mediator for both activator-independent and activator-dependent transcription in conjunction with TFIID-associated TAF(II)s. Mol Cell Biol. 22(8):2842-52. Balamotis MA, Pennella MA, Stevens JL, Wasylyk B, Belmont AS, Berk AJ (2009) Complexity in transcription control at the activation domain-mediator interface. Sci Signal. 2(69):169-177. Bandyopadhyay A, Tsuji K, Cox K, Harfe BD, Rosen V, Tabin CJ (2006) Genetic analysis of the roles of BMP2, BMP4, and BMP7 in limb patterning and skeletogenesis. PLoS Genet. 2(12):216-226. Barker DJ (2006) Adult consequences of fetal growth restriction. Clin Obstet Gynecol. 49(2):270-83. Barner M, Niswander L (2007) Visualization of cartilage formation: insight into cellular properties of skeletal progenitors and chondrodysplasia syndromes. Dev Cell.12(6):931-41. Bell DM, Leung KK, Wheatley SC, Ng LJ, Zhou S, Ling KW, Sham MH, Koopman P, Tam PP, Cheah KS (1997) SOX9 directly regulates the type-II collagen gene. Nat Genet. 16(2):174-8. Bernstein IM, Horbar JD, Badger GJ, Ohlsson A, Golan A (2000) Morbidity and mortality among very-low-birth-weight neonates with intrauterine growth restriction. Am J Obstet Gynecol. 182(1 Pt 1):198-206. Berti L, Mittler G, Przemeck GK, Stelzer G, Günzler B, Amati F, Conti E, Dallapiccola B, Hrabé de Angelis M, Novelli G, Meisterernst M (2001) Isolation and characterization of a novel gene from the DiGeorge chromosomal region that encodes for a mediator subunit. Genomics. 74(3):320-32. Béve J, Hu GZ, Myers LC, Balciunas D, Werngren O, Hultenby K, Wibom R, Ronne H, Gustafsson CM (2005) The structural and functional role of Med5 in the yeast Mediator tail module. J Biol Chem. 280(50):41366-72. Beyer KS, Beauchamp RL, Lee MF, Gusella JF, Näär AM, Ramesh V (2007) Mediator subunit MED28 (Magicin) is a repressor of smooth muscle cell differentiation. J Biol Chem. 282(44):32152-7. Bi W, Deng JM, Zhang Z, Behringer RR, de Crombrugghe B (1999) Sox9 is required for cartilage formation. Nat Genet. 22(1):85-9. Bi W, Huang W, Whitworth DJ, Deng JM, Zhang Z, Behringer RR, de Crombrugghe B (2001) Haploinsufficiency of Sox9 results in defective cartilage primordia and premature skeletal mineralization. Proc Natl Acad Sci U S A. 98(12):6698-703. Biddick R, Young ET (2005) Yeast mediator and its role in transcriptional regulation. C R Biol. 328(9):773-82. Boeing S, Rigault C, Heidemann M, Eick D, Meisterernst M (2010) RNA polymerase II C-terminal heptarepeat domain Ser-7 phosphorylation is established in a mediator- dependent fashion. J Biol Chem. 285(1):188-96.

146

Boles MK, Wilkinson BM, Wilming LG, Liu B, Probst FJ, Harrow J, Grafham D, Hentges KE, Woodward LP, Maxwell A, Mitchell K, Risley MD, Johnson R, Hirschi K, Lupski JR, Funato Y, Miki H, Marin-Garcia P, Matthews L, Coffey AJ, Parker A, Hubbard TJ, Rogers J, Bradley A, Adams DJ, Justice MJ (2009) Discovery of candidate disease genes in ENU-induced mouse mutants by large-scale sequencing, including a splice-site mutation in nucleoredoxin. PLoS Genet. 5(12):e1000759. Bolon B (2015) Pathology of the Developing Mouse: A Systematic Approach. CRC Press. Bond H, Dilworth MR, Baker B, Cowley E, Requena Jimenez A, Boyd RD, Husain SM, Ward BS, Sibley CP, Glazier JD (2008) Increased maternofetal calcium flux in parathyroid hormone-related protein-null mice. J Physiol. 586(7):2015-25. Borggrefe T, Yue X (2011) Interactions between subunits of the Mediator complex with gene-specific transcription factors. Semin Cell Dev Biol. 22(7):759-68. Bosveld F, van Hoek S, Sibon OC (2008) Establishment of cell fate during early Drosophila embryogenesis requires transcriptional Mediator subunit dMED31. Dev Biol.313(2):802-13. Bouillot S, Rampon C, Tilet E, Huber P (2006) Tracing the glycogen cells with protocadherin 12 during mouse placenta development. Placenta. 27(8):882-8. Brooke OG, Wood C, Butters F (1984) The body proportions for small-for-dates infants. Early Hum Dev.10(1-2):85-94. Burke TW, Kadonaga JT (1997) The downstream core promoter element, DPE, is conserved from Drosophila to humans and is recognized by TAFII60 of Drosophila. Genes Dev.11(22):3020-31. Caplan AI, Pechak DG (1987) The cellular and molecular embryology of bone formation. Bone and mineral research. NewYork: Elsevier. Cetin I, Marconi AM, Baggiani AM, Buscaglia M, Pardi G, Fennessey PV, Battaglia FC (1995) In vivo placental transport of glycine and leucine in human pregnancies. Pediatr Res.37(5):571-5. Cevher MA, Shi Y, Li D, Chait BT, Malik S, Roeder RG (2014) Reconstitution of active human core Mediator complex reveals a critical role of the MED14 subunit. Nat Struct Mol Biol.21(12):1028-34. Chalkley GE, Verrijzer CP (1999) DNA binding site selection by RNA polymerase II TAFs: a TAF(II)250-TAF(II)150 complex recognizes the initiator. EMBO J. 18(17):4835-45. Chen MS, Ryan CE, Piwnica-Worms H (2003) Chk1 kinase negatively regulates mitotic function of Cdc25A phosphatase through 14-3-3 binding. Mol Cell Biol. 23(21):7488-97. Chial H (2008) DNA sequencing technologies key to the Human Genome Project. Nature Education. 1(1):219.

147

Choi Y, Sims GE, Murphy S, Miller JR, Chan AP (2012) Predicting the functional effect of amino acid substitutions and indels. PLoS One. 7(10):e46688. Clayton JE, van den Heuvel SJ, Saito RM (2008) Transcriptional control of cell-cycle quiescence during C. elegans development. Dev Biol. 313(2):603-13. Coan PM, Angiolini E, Sandovici I, Burton GJ, Constância M, Fowden AL (2008) Adaptations in placental nutrient transfer capacity to meet fetal growth demands depend on placental size in mice. J Physiol. 586(18):4567-76. Coan PM, Conroy N, Burton GJ, Ferguson-Smith AC (2006) Origin and characteristics of glycogen cells in the developing murine placenta. Developmental Dynamics. 235(12):3280–3294. Coan PM, Ferguson-Smith AC, Burton GJ (2005) Ultrastructural changes in the interhaemal membrane and junctional zone of the murine chorioallantoic placenta across gestation. J Anat. 207(6):783-96. Coan PM, Vaughan OR, McCarthy J, Mactier C, Burton GJ, Constância M, Fowden AL (2011) Dietary composition programmes placental phenotype in mice. J Physiol. 589(Pt 14):3659-70. Coan PM, Vaughan OR, Sekita Y, Finn SL, Burton GJ, Constância M, Fowden AL (2010) Adaptations in placental phenotype support fetal growth during undernutrition of pregnant mice. J Physiol. 588(Pt 3):527-38. Conaway RC, Conaway JW (2011) Function and regulation of the Mediator complex. Curr Opin Genet Dev. 21(2):225-30. Conaway RC, Sato S, Tomomori-Sato C, Yao T, Conaway JW (2005) The mammalian Mediator complex and its role in transcriptional regulation. Trends Biochem Sci. 2005 May;30(5):250-5. Conlon I, Raff M (1999) Size control in animal development. Cell. 96(2):235-44. Constância M, Angiolini E, Sandovici I, Smith P, Smith R, Kelsey G, Dean W, Ferguson-Smith A, Sibley CP, Reik W, Fowden A (2005) Adaptation of nutrient supply to fetal demand in the mouse involves interaction between the Igf2 gene and placental transporter systems. Proc Natl Acad Sci USA. 102(52):19219-24. Constância M, Hemberger M, Hughes J, Dean W, Ferguson-Smith A, Fundele R, Stewart F, Kelsey G, Fowden A, Sibley C, Reik W. (2002) Placental specific IGF-II is a major modulator of placental and fetal growth. Nature. 417(6892):945-8. Cross JC, Baczyk D, Dobric N, Hemberger M, Simmons DG, Yamamoto H, Kingdom JC (2003) Genes, development and evolution of the placenta. Placenta. 24(2-3):123-30. Cross JC, Nakano H, Natale DR, Simmons DG, Watson ED (2006) Branching morphogenesis during development of placental villi. Differentiation. 74(7):393-401. Crossley PH, Minowada G, MacArther CA, Martin GR (1996) Roles for FGF8 in the induction, initiation and maintenance of chick limb development. Cell. 84(1):127-36.

148

Dasso M, Newport JW (1990) Completion of DNA replication is monitored by a feedback system that controls the initiation of mitosis in vitro: studies in Xenopus. Cell. 61(5):811-23. Davis AP, Justice MJ (1998) Mouse alleles: if you've seen one, you haven't seen them all. Trends Genet. 14(11):438-41. Davis JA, Takagi Y, Kornberg RD, Asturias FA (2002) Structure of the yeast RNA polymerase II holoenzyme: Mediator conformation and polymerase interaction. Mol Cell.10(2):409-15. Davis MA, Larimore EA, Fissel BM, Swanger J, Taatjes DJ, Clurman BE (2013) The SCF-Fbw7 ubiquitin ligase degrades MED13 and MED13L and regulates CDK8 module association with Mediator. Genes Dev. 27(2):151-6. de Boo HA, Harding JE (2006) The developmental origins of adult disease (Barker) hypothesis. Aust N Z J Obstet Gynaecol. 46(1):4-14. Dilworth MR, Kusinski LC, Cowley E, Ward BS, Husain SM, Constância M, Sibley CP, Glazier JD (2010) Placental-specific Igf2 knockout mice exhibit hypocalcemia and adaptive changes in placental calcium transport. Proc Natl Acad Sci U S A. 107(8):3894-9.

Dilworth MR, Sibley CP (2012) Review: Transport across the placenta of mice and women. Placenta. 34. 34-9. Drouin R, Lemieux N, Richer CL (1991) Chromosome condensation from prophase to late metaphase: relationship to chromosome bands and their replication time. Cytogenet Cell Genet. 57(2-3):91-9. Dunphy WG, Brizuela L, Beach D, Newport J (1988) The Xenopus cdc2 protein is a component of MPF, a cytoplasmic regulator of mitosis. Cell. 54(3):423-31. Dürr K, Holzschuh J, Filippi A, Ettl AK, Ryu S, Shepherd IT, Driever W (2006) Differential roles of transcriptional mediator complex subunits Crsp34/Med27, Crsp150/Med14 and Trap100/Med24 during zebrafish retinal development. Genetics. 174(2):693-705. Elmlund H, Baraznenok V, Lindahl M, Samuelsen CO, Koeck PJ, Holmberg S, Hebert H, Gustafsson CM (2006) The cyclin-dependent kinase 8 module sterically blocks Mediator interactions with RNA polymerase II. Proc Natl Acad Sci U S A. 103(43):15788-93. Esnault C, Ghavi-Helm Y, Brun S, Soutourina J, Van Berkum N, Boschiero C, Holstege F, Werner M (2008) Mediator-dependent recruitment of TFIIH modules in preinitiation complex. Mol Cell. 31(3):337-46. Fallon JF, Lopez A, Ros MA, Savage MP, Olwin BB, Simandl BK (1994) FGF-2: apical ectodermal ridge growth signal for chick limb development .Science. 264(5155):104-7. Gaccioli F, Lager S, Powell T, Jansson T (2013) Placental Transport in Response to Altered Maternal Nutrition. J Dev Orig Health Dis. 4(2): 101–115.

149

Galbraith MD, Donner AJ, Espinosa JM (2010) CDK8: a positive regulator of transcription. Transcription. 1(1):4-12. Galjaard S, Devlieger R, Van Assche FA (2013) Fetal growth and developmental programming. J Perinat Med. 41(1):101-5. Gendron-Maguire M, Mallo M, Zhang M, Gridley T (1993) Hoxa-2 mutant mice exhibit homeotic transformation of skeletal elements derived from cranial neural crest. Cell. 75(7):1317-31. Georgatos SD, Pyrpasopoulou A, Theodoropoulos PA (1997) Nuclear envelope breakdown in mammalian cells involves stepwise lamina disassembly and microtubule-drive deformation of the nuclear membrane. J Cell Sci. 110 ( Pt 17):2129-40. Georgiades P, Ferguson-Smith AC, Burton GJ (2002) Comparative developmental anatomy of the murine and human definitive placentae. Placenta. 23(1):3-19. Glazier JD, Cetin I, Perugino G, Ronzoni S, Grey AM, Mahendran D, Marconi AM, Pardi G, Sibley CP. (1997). Association between the activity of the system A amino acid transporter in the microvillous plasma membrane of the human placenta and severity of fetal compromise in intrauterine growth restriction. Pediatr Res. 42(4):514-9. Gong D, Ferrell JE (2010) The roles of cyclin A2, B1, and B2 in early and late mitotic events. Mol Biol Cell. 21(18):3149-61. Graham JM, Schwartz CE (2013) MED12 related disorders. Am J Med Genet A. 161(11):2734-40. Grande JP, Melder DC, Zinsmeister AR (1997) Modulation of collagen gene expression by cytokines: stimulatory effect of transforming growth factor-beta1, with divergent effects of epidermal growth factor and tumor necrosis factor-alpha on collagen type I and collagen type IV. J Lab Clin Med. 130(5):476-86. Grøntved L, Madsen MS, Boergesen M, Roeder RG, Mandrup S (2010) MED14 tethers mediator to the N-terminal domain of peroxisome proliferator-activated receptor gamma and is required for full transcriptional activity and adipogenesis. Mol Cell Biol. 30(9):2155-69. Grünberg S, Hahn S (2013) Structural insights into transcription initiation by RNA polymerase II. Trends Biochem Sci. 38(12):603-11. Grüneberg H (1963). The pathology of development: a study of inherited skeletal disorders in animals. Oxford: Blackwell. Guglielmi B, van Berkum NL, Klapholz B, Bijma T, Boube M, Boschiero C, Bourbon HM, Holstege FC, Werner M (2004) A high resolution protein interaction map of the yeast Mediator complex. Nucleic Acids Res. 32(18):5379-91. Hall BK, Miyake T (1995) Divide, accumulate, differentiate: cell condensation in skeletal development revisited. Int J Dev Biol. 39(6):881-93.

150

Hall BK, Miyake T (2000) All for one and one for all: condensations and the initiation of skeletal development. BioEssays. 22(2):138-47. Harper JW, Adams PD (2001) Cyclin-dependent kinases. Chem Rev. 101(8):2511-26. Hemberger M, Cross, JC (2001). Genes governing placental development. Trends in Endocrinology and Metabolism. 12(4):162-8. Hentges KE (2011) Mediator complex proteins are required for diverse developmental processes. Semin Cell Dev Biol. 22(7):769-75. Hentges KE (2011) The Mediator complex: crucial functions in transcription with links to development and disease. Semin Cell Dev Biol. 22(7):728. Hentges KE, Justice MJ (2004) Checks and balancers: balancer chromosomes to facilitate genome annotation. Trends Genet. 20(6):252-9. Hentges KE, Nakamura H, Furuta Y, Yu Y, Thompson DM, O'Brien W, Bradley A, Justice MJ (2006) Novel lethal mouse mutants produced in balancer chromosome screens. Gene Expr Patterns. 6(6):653-65. Hinoi E, Bialek P, Chen YT, Rached MT, Groner Y, Behringer RR, Ornitz DM, Karsenty G (2006) Runx2 inhibits chondrocyte proliferation and hypertrophy through its expression in the perichondrium. Genes Dev. 20(21):2937-42. Hitz C, Vogt-Weisenhorn D, Ruiz P, Wurst W, Floss T (2005) Progressive loss of the spongiotrophoblast layer of Birc6/Bruce mutants results in embryonic lethality. Genesis. 42(2):91-103. Holstege FC, van der Vliet PC, Timmers HT (1996) Opening of an RNA polymerase II promoter occurs in two distinct steps and requires the basal transcription factors IIE and IIH. EMBO J. 15(7):1666-77. Hu D, Cross JC (2011) Ablation of Tpbpa-positive trophoblast precursors leads to defects in maternal spiral artery remodeling in the mouse placenta. Dev Biol. 358(1):231-9. Ikeda T, Kamekura S, Mabuchi A, Kou I, Seki S, Takato T, Nakamura K, Kawaguchi H, Ikegawa S, Chung UI (2004) The combination of SOX5, SOX6, and SOX9 (the SOX trio) provides signals sufficient for induction of permanent cartilage. Arthritis Rheum. 50(11):3561-73. Ito M, Okano HJ, Darnell RB, Roeder RG (2002) The TRAP100 component of the TRAP/Mediator complex is essential in broad transcriptional events and development. EMBO J. 21(13):3464-75. Ito M, Yuan CX, Malik S, Gu W, Fondell JD, Yamamura S, Fu ZY, Zhang X, Qin J, Roeder RG (1999) Identity between TRAP and SMCC complexes indicates novel pathways for the function of nuclear receptors and diverse mammalian activators. Mol Cell. 3(3):361-70. Ito M, Yuan CX, Okano HJ, Darnell RB, Roeder RG (2000) Involvement of the TRAP220 component of the TRAP/SMCC coactivator complex in embryonic development and thyroid hormone action. Mol Cell. 5(4):683-93.

151

Jansson T, Powell TL (2007) Role of the placenta in fetal programming: underlying mechanisms and potential interventional approaches. Clin Sci (Lond).113(1):1-13. Jiang C, Chen H, Shao L, Wang Q (2014) MicroRNA-1 functions as a potential tumor suppressor in osteosarcoma by targeting Med1 and Med31. Oncol Rep. 32(3):1249- 56. Jiang YW, Veschambre P, Erdjument-Bromage H, Tempst P, Conaway JW, Conaway RC, Kornberg RD (1998) Mammalian mediator of transcriptional regulation and its possible role as an end-point of signal transduction pathways. Proc Natl Acad Sci U S A. 95(15):8538-43. Jishage M, Malik S, Wagner U, Uberheide B, Ishihama Y, Hu X, Chait BT, Gnatt A, Ren B, Roeder RG (2012) Transcriptional regulation by Pol II(G) involving mediator and competitive interactions of Gdown1 and TFIIF with Pol II. Mol Cell. 45(1):51-63. Kagey MH, Newman JJ, Bilodeau S, Zhan Y, Orlando DA, van Berkum NL, Ebmeier CC, Goossens J, Rahl PB, Levine SS, Taatjes DJ, Dekker J, Young RA (2010) Mediator and cohesin connect gene expression and chromatin architecture. Nature. 467(7314):430-5. Karaplis AC, Luz A, Glowacki J, Bronson RT, Tybulewicz VL, Kronenberg HM, Mulligan RC (1994) Lethal skeletal dysplasia from targeted disruption of the parathyroid hormone-related peptide gene. Genes Dev. 8(3):277-89. Karp SJ, Schipani E, St-Jacques B, Hunzelman J, Kronenberg H, McMahon AP (2000) Indian hedgehog coordinates endochondral bone growth and morphogenesis via parathyroid hormone related-protein-dependent and -independent pathways. Development. 127(3):543-8. Karsenty, G. Kronenberg, H.M. Settembre, C (2009) Genetic control of bone formation. Annu. Rev. Cell Dev. Biol. 25:629-48. Kaufman MH (1992) The atlas of mouse development. San Diego: Academic Press. Kelleher R, Flanagan PM, Kornberg RD (1990) A novel mediator between activator proteins and the RNA polymerase II transcription apparatus. Cell. 61(7):1209-15. Kile BT, Hentges KE, Clark AT, Nakamura H, Salinger AP, Liu B, Box N, Stockton DW, Johnson RL, Behringer RR, Bradley A, Justice MJ (2003). Functional genetic analysis of mouse chromosome 11. Nature. 425(6953):81-6. Kim IS, Otto F, Zabel B, Mundlos S (1999) Regulation of chondrocyte differentiation by Cbfa1. Mech Dev. 80(2):159-70. Kim YJ, Björklund S, Li Y, Sayre MH, Kornberg RD (1994) A multiprotein mediator of transcriptional activation and its interaction with the C-terminal repeat domain of RNA polymerase II. Cell. 77(4):599-608. Knuesel MT, Meyer KD, Bernecky C, Taatjes DJ (2009) The human CDK8 subcomplex is a molecular switch that controls Mediator coactivator function. Genes Dev. 23(4):439-51.

152

Koleske AJ, Young RA (1995) The RNA polymerase II holoenzyme and its implications for gene regulation. Trends Biochem Sci. 20(3):113-6. Kornberg RD (2005) Mediator and the mechanism of transcriptional activation.Trends Biochem Sci. 30(5):235-9. Kornberg RD (2007) The molecular basis of eukaryotic transcription. Cell Death Differ. 14(12):1989-97. Koschubs T, Seizl M, Larivière L, Kurth F, Baumli S, Martin DE, Cramer P (2009) Identification, structure, and functional requirement of the Mediator submodule Med7N/31. EMBO J. 28(1):69-80. Kronenberg HM (2003) Developmental regulation of the growth plate. Nature. 423(6937):332-6. Kumar D, Lassar AB (2009) The transcriptional activity of Sox9 in chondrocytes is regulated by RhoA signaling and actin polymerization. Mol Cell Biol. 29(15):4262- 73. Landles C, Chalk S, Steel JH, Rosewell I, Spencer-Dene B, Lalani el-N, Parker MG (2003) The thyroid hormone receptor-associated protein TRAP220 is required at distinct embryonic stages in placental, cardiac, and hepatic development. Mol Endocrinol. 17(12):2418-35. Lanske B, Karaplis AC, Lee K, Luz A, Vortkamp A, Pirro A, Karperien M, Defize LH, Ho C, Mulligan RC, Abou-Samra AB, Jüppner H, Segre GV, Kronenberg HM (1996) PTH/PTHrP receptor in early development and Indian hedgehog-regulated bone growth. Science. 273(5275):663-6. Larivière L, Plaschka C, Seizl M, Petrotchenko EV, Wenzeck L, Borchers CH, Cramer P (2013) Model of the Mediator middle module based on protein cross- linking. Nucleic Acids Res. 41(20):9266-73. Larivière L, Plaschka C, Seizl M, Wenzeck L, Kurth F, Cramer P (2012) Structure of the Mediator head module. Nature. 492(7429):448-51. Leal A, Huehne K, Bauer F, Sticht H, Berger P, Suter U, Morera B, Del Valle G, Lupski JR, Ekici A, Pasutto F, Endele S, Barrantes R, Berghoff C, Berghoff M, Neundörfer B, Heuss D, Dorn T, Young P, Santolin L, Uhlmann T, Meisterernst M, Sereda MW, Stassart RM, Zu Horste GM, Nave KA, Reis A, Rautenstrauss B (2009) Identification of the variant Ala335Val of MED25 as responsible for CMT2B2: molecular data, functional studies of the SH3 recognition motif and correlation between wild-type MED25 and PMP22 RNA levels in CMT1A animal models. Neurogenetics. 10(4):375-376. Leatherbury L, Waldo K (1995) Visual understanding of cardiac development: the neural crest's contribution. Cell Mol Biol Res. 41(4):279-91. Lefebvre V, Bhattaram P (2010) Vertebrate skeletogenesis. Curr Top Dev Biol. 90:291-317.

153

Lefebvre V, Huang W, Harley VR, Goodfellow PN, de Crombrugghe B (1997) SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro alpha1(II) collagen gene. Mol Cell Biol. 17(4):2336-46. Lefebvre V, Smits P (2005) Transcriptional control of chondrocyte fate and differentiation. Birth Defects Res C Embryo Today. 75(3):200-12. Lescisin K R, Varmuza S, Rossant J (1998) The expression of placental lactogen (PL) is regulated during trophoblast giant cell. Genes Dev. 12(8):1315–1326 Li SW, Prockop DJ, Helminen H, Fässler R, Lapveteläinen T, Kiraly K, Peltarri A, Arokoski J, Lui H, Arita M (1995) Transgenic mice with targeted inactivation of the Col2 alpha 1 gene for collagen II develop a skeleton with membranous and periosteal bone but no endochondral bone. Genes Dev. 9(22):2821-30. Linder T, Gustafsson CM (2004) The Soh1/MED31 protein is an ancient component of Schizosaccharomyces pombe and Saccharomyces cerevisiae Mediator. J Biol Chem. 279(47):49455-9. Liu G, Iwata K, Ogasawara T, Watanabe J, Fukazawa K, Ishihara K, Asawa Y, Fujihara Y, Chung UL, Moro T, Takatori Y, Takato T, Nakamura K, Kawaguchi H, Hoshi K (2010) Selection of highly osteogenic and chondrogenic cells from bone marrow stromal cells in biocompatible polymer-coated plates. J Biomed Mater Res A. 92(4):1273-82. Liu P, Wakamiya M, Shea MJ, Albrecht U, Behringer RR, Bradley A (1999) Requirement for Wnt3 in vertebrate axis formation. Nat Genet. 22(4):361-5. Liu Y, Ranish JA, Aebersold R, Hahn S (2001) Yeast nuclear extract contains two major forms of RNA polymerase II mediator complexes. J Biol Chem. 276(10):7169- 75. Lu H, Flores O, Weinmann R, Reinberg D (1991) The nonphosphorylated form of RNA polymerase II preferentially associates with the preinitiation complex. Proc Natl Acad Sci U S A. 88(22):10004-8. Lutfi AM (1974) The role of cartilage in long bone growth: a reappraisal. J Anat. 117(Pt 2):413-7. Mahendran D, Donnai P, Glazier JD, D’Souza SW, Boyd RD, Sibley CP (1993) Amino acid (system A) transporter activity in microvillous membrane vesicles from the placentas of appropriate and small for gestational age babies. Pediatr Res.34(5):661-5. Malik S, Barrero MJ, Jones T (2007) Identification of a regulator of transcription elongation as an accessory factor for the human Mediator coactivator. Proc Natl Acad Sci U S A.104(15):6182-7. Malik S, Gu W, Wu W, Qin J, Roeder RG (2000) The USA-derived transcriptional coactivator PC2 is a submodule of TRAP/SMCC and acts synergistically with other PCs. Mol Cell. 5(4):753-60. Malik S, Roeder RG (2010) The metazoan Mediator co-activator complex as an integrative hub for transcriptional regulation. Nat Rev Genet. 11(11):761-72.

154

Martin BL, Harland RM (2001) Hypaxial muscle migration during primary myogenesis in Xenopus laevis. Dev Biol. 239(2):270-80. Mayhew TM, Manwani R, Ohadike C, Wijesekara J, Baker PN (2007) The placenta in pre-eclampsia and intrauterine growth restriction: studies on exchange surface areas, diffusion distances and villous membrane diffusive conductances. Placenta. 28(2-3):233-8. McCormick MC (1985) The contribution of low birth weight to infant mortality and childhood morbidity. N Engl J Med. 312(2):82-90. Michael WM, Newport J (1998) Coupling of mitosis to the completion of S phase through Cdc34-mediated degradation of Wee1. Science. 282(5395):1886-9. Mikhailov A, Shinohara M, Rieder CL (2005) The p38-mediated stress-activated checkpoint. A rapid response system for delaying progression through antephase and entry into mitosis. Cell Cycle. 4(1):57-62. Miklos I, Szilagyi Z, Watt S, Zilahi E, Batta G, Antunovics Z, Enczi K, Bähler J, Sipiczki M (2008) Genomic expression patterns in cell separation mutants of Schizosaccharomyces pombe defective in the genes sep10 ( + ) and sep15 ( + ) coding for the Mediator subunits Med31 and Med8. Mol Genet Genomics. 279(3):225-38. Miller J, Turan S, Baschat AA (2008) Fetal growth restriction. Semin Perinatol. 32(4):274-80. Mills TA, Wareing M, Bugg GJ, Greenwood SL, Baker PN. (2005). Chorionic plate artery function and Doppler indices in normal pregnancy and intrauterine growth restriction. Eur J Clin Invest.35(12):758-64. Minet M, Nurse P, Thuriaux P, Mitchison JM (1979) Uncontrolled septation in a cell division cycle mutant of the fission yeast Schizosaccharomyces pombe. J Bacteriol. 137(1):440-6. Mitiku N, Baker JC (2007) Genomic analysis of gastrulation and organogenesis in the mouse. Dev Cell. 13(6):897-907. Mittler G, Kremmer E, Timmers HT, Meisterernst M (2001) Novel critical role of a human Mediator complex for basal RNA polymerase II transcription. EMBO Rep. 2(9):808-13. Moffat J, Andrews B (2004) Late-G1 cyclin-CDK activity is essential for control of cell morphogenesis in budding yeast. Nat Cell Biol. 6(1):59-66. Moore NP, Boogaard PJ, Bremer S, Buesen R, Edwards J, Fraysse B, Hallmark N, Hemming H, Langrand-Lerche C, McKee RH, Meisters ML, Parsons P, Politano V, Reader S, Ridgway P, Hennes C (2013) Guidance on classification for reproductive toxicity under the globally harmonized system of classification and labelling of chemicals (GHS). Crit Rev Toxicol. 43(10):850-91. Mouse Genome Sequencing Consortium (2002) Initial sequencing and comparative analysis of the mouse genome. Nature. 420: 520-562.

155

Muncke N, Jung C, Rüdiger H, Ulmer H, Roeth R, Hubert A, Goldmuntz E, Driscoll D, Goodship J, Schön K, Rappold G (2003) Missense mutations and gene interruption in PROSIT240, a novel TRAP240-like gene, in patients with congenital heart defect (transposition of the great arteries) Circulation. 108(23):2843-50. Murray SA, Morgan JL, Kane C, Sharma Y, Heffner CS, Lake J, Donahue LR (2010) Mouse gestation length is genetically determined. PLoS One. 5(8):e12418. Näär AM, Beaurang PA, Zhou S, Abraham S, Solomon W, Tjian R (1999) Composite co-activator ARC mediates chromatin-directed transcriptional activation. Nature. 398(6730):828-32. Nasmyth K (1999) Separating sister chromatids. Trends Biochem Sci. 24(3):98-104. Ng LJ, Wheatley S, Muscat GE, Conway-Campbell J, Bowles J, Wright E, Bell DM, Tam PP, Cheah KS, Koopman P (1997) SOX9 binds DNA, activates transcription, and coexpresses with type II collagen during chondrogenesis in the mouse. Dev Biol. 183(1):108-21. Norberg S, Powell T, Jansson T (1998) Intrauterine growth restriction is associated with a reduced activity of placental taurine transporters. Pediatr Res. 44(2):233-8. Nurse P (2000) A long twentieth century of the cell cycle and beyond. Cell. 100(1):71-8. Oda Y, Chalkley RJ, Burlingame AL, Bikle DD (2010) The transcriptional coactivator DRIP/mediator complex is involved in vitamin D receptor function and regulates keratinocyte proliferation and differentiation. J Invest Dermatol. 130(10):2377-88. Ohi R, Gould KL (1999) Regulating the onset of mitosis. Curr Opin Cell Biol. 11(2):267-73. Oh-McGinnis R, Bogutz AB, Lefebvre L (2011) Partial loss of Ascl2 function affects all three layers of the mature placenta and causes intrauterine growth restriction. Dev Biol. 351(2):277-86. Olsen BR, Reginato AM, Wang W (2000) Bone Development. Annu. Rev. Cell Dev. Biol. 16:191-220 O'Rahilly R (1979) Early human development and the chief sources of information on staged human embryos. Eur. J. Obstet. Gynecol. Reprod. Biol. 9(4);273-80. Orphanides G, Lagrange T, Reinberg D (1996) The general transcription factors of RNA polymerase II. Genes Dev. 10(21):2657-83. Ozanne SE, Hales CN (2004) Lifespan: catch-up growth and obesity in male mice. Nature. 427(6973):411-2. Pines J, Rieder CL (2001) Re-staging mitosis: a contemporary view of mitotic progression. Nat Cell Biol. 3(1):3-6.

156

Plaschka C, Larivière L, Wenzeck L, Seizl M, Hemann M, Tegunov D, Petrotchenko EV, Borchers CH, Baumeister W, Herzog F, Villa E, Cramer P (2015) Architecture of the RNA polymerase II-Mediator core initiation complex. Nature. 518(7539):376-80. Poss ZC, Ebmeier CC, Taatjes DJ (2013) The Mediator complex and transcription regulation. Crit Rev Biochem Mol Biol. 48(6):575-608. Prockop DJ, Ala-Kokko L, McLain DA, Williams C (1997) Can mutated genes cause common osteoarthritis? Br J Rheumatol. 36(8):827-9. Rieder CL, Salmon ED (1998) The vertebrate cell kinetochore and its roles during mitosis. Trends Cell Biol. 8(8):310-8. Risley MD, Clowes C, Yu M, Mitchell K, Hentges KE (2010) The Mediator complex protein Med31 is required for embryonic growth and cell proliferation during mammalian development. Dev Biol. 342(2):146-56. Robinson PJ, Bushnell DA, Trnka MJ, Burlingame AL, Kornberg RD (2012) Structure of the mediator head module bound to the carboxy-terminal domain of RNA polymerase II. Proc Natl Acad Sci U S A. 109(44):17931-5. Rocha PP, Scholze M, Bleiss W, Schrewe H (2010) Med12 is essential for early mouse development and for canonical Wnt and Wnt/PCP signaling. Development. 137(16):2723-31. Rodriguez TA, Sparrow DB, Scott AN, Withington SL, Preis JI, Michalicek J, Clements M, Tsang TE, Shioda T, Beddington RS, Dunwoodie SL (2004) Cited1 is required in trophoblasts for placental development and for embryo growth and survival. Mol Cell Biol. 24(1):228-44. Roeder RG (1996) The role of general initiation factors in transcription by RNA polymerase II. Trends Biochem Sci. 21(9):327-35. Rossant J, Cross JC (2001) Placental development: lessons from mouse mutants. Nat Rev Genet. 2(7):538-48. Russell WL, Kelly EM, Hunsicker PR, Bangham JW, Maddux SC, Phipps EL (1979) Specific-locus test shows ethylnitrosourea to be the most potent mutagen in the mouse. Proc Natl Acad Sci U S A. 76(11):5818-9. Saleem T, Sajjad N, Fatima S, Habib N, Ali S, Qadir M (2011) Intrauterine growth retardation - small events, big consequences. Ital J Pediatr. 37-41. Samánek M (2000) Congenital heart malformations: prevalence, severity, survival, and quality of life. Cardiol Young. 10(3):179-85. Sato S, Tomomori-Sato C, Parmely TJ, Florens L, Zybailov B, Swanson SK, Banks CA, Jin J, Cai Y, Washburn MP, Conaway JW, Conaway RC (2004) A set of consensus mammalian mediator subunits identified by multidimensional protein identification technology. Mol Cell. 14(5):685-91. Schiano C, Rienzo M, Casamassimi A, Napoli C (2013) Gene expression profile of the whole Mediator complex in human osteosarcoma and normal osteoblasts. Med Oncol. 30(4):739.

157

Schmelzle T, Hall MN (2000) TOR, a central controller of cell growth. Cell.103(2):253-62. Schneider M, Hellerschmied D, Schubert T, Amlacher S, Vinayachandran V, Reja R, Pugh BF, Clausen T, Köhler A (2015) The Nuclear Pore-Associated TREX-2 Complex Employs Mediator to Regulate Gene Expression. Cell. 162(5):1016-28. Sferruzzi-Perri AN, Vaughan OR, Coan PM, Suciu MC, Darbyshire R, Constância M, Burton GJ, Fowden AL (2011) Placental-specific Igf2 deficiency alters developmental adaptations to undernutrition in mice. Endocrinology. 152(8):3202-12. Shankland SJ, Wolf G (2000) Cell cycle regulatory proteins in renal disease: role in hypertrophy, proliferation, and apoptosis. Am J Physiol Renal Physiol. 278(4):F515- 29. Sherr CJ, Roberts JM (2004) Living with or without cyclins and cyclin-dependent kinases. Genes Dev. 18(22):2699-711. Shimizu H, Yokoyama S, Asahara H (1997) Growth and differentiation of the developing limb bud from the perspective of chondrogenesis. Dev Growth Differ. 2007 49(6):449-54. Shimizu T, Nakano K, Tsujigiwa H, Nagatsuka H, Watanabe T, Okafuji N, Kurihara S, Hasegawa H, Nagai N, Kawakami T (2007) Notch signaling in mandibular condylar cartilage development. Eur J Med Res.12(10):515-9. Shin BC, Fujikura K, Suzuki T, Tanaka S, Takata K (1997). Glucose transporter GLUT3 in the rat placental barrier: a possible machinery for the transplacental transfer of glucose. Endocrinology. 138(9):3997-4004. Sibley CP (2009) Understanding placental nutrient transfer--why bother? New biomarkers of fetal growth. J Physiol. 587(Pt 14):3431-40. Sibley CP, Boyd RD (1988) Control of transfer across the mature placenta. Oxf Rev Reprod Biol. 10:382-435. Sibley CP, Coan PM, Ferguson-Smith AC, Dean W, Hughes J, Smith P, Reik W, Burton GJ, Fowden AL, Constância M (2004) Placental-specific insulin-like growth factor 2 (Igf2) regulates the diffusional exchange characteristics of the mouse placenta. Proc Natl Acad Sci U S A. 101(21):8204-8. Sibley CP, Glazier J, D’ Souza S (1997) Placental transporter activity and expression in relation to fetal growth. Exp Physiol. 82(2):389-402. Sibley CP, Turner MA, Cetin I, Ayuk P, Boyd CA, D'Souza SW, Glazier JD, Greenwood SL, Jansson T, Powell T (2005) Placental phenotypes of intrauterine growth. Pediatr Res. 58(5):827-32. Smits P, Li P, Mandel J, Zhang Z, Deng JM, Behringer RR, de Crombrugghe B, Lefebvre V (2001) The transcription factors L-Sox5 and Sox6 are essential for cartilage formation. Dev Cell. 1(2):277-90. Snow MHL, Tam PPL, McLaren A (1981) The control and regulation of size and morphogenesis in mammalian embryos. Proc. Soc. Devl.Biol. 201-217.

158

Søgaard TM, Svejstrup JQ (2007) Hyperphosphorylation of the C-terminal repeat domain of RNA polymerase II facilitates dissociation of its complex with mediator. J Biol Chem. 282(19):14113-20. Stumpf M, Waskow C, Krötschel M, van Essen D, Rodriguez P, Zhang X, Guyot B, Roeder RG, Borggrefe T (2006) The mediator complex functions as a coactivator for GATA-1 in erythropoiesis via subunit Med1/TRAP220. Proc Natl Acad Sci U S A.103(49):18504-9. Stumpf M, Yue X, Schmitz S, Luche H, Reddy JK, Borggrefe T (2010) Specific erythroid-lineage defect in mice conditionally deficient for Mediator subunit Med1. Proc Natl Acad Sci U S A. 107(50):21541-6. Summerbell D, Lewis JH, Wolpert L (1973) Positional information in chick limb morphogenesis. Nature. 244(5417):492-6. Sun X, Zhang Y, Cho H, Rickert P, Lees E, Lane W, Reinberg D (1998) NAT, a human complex containing Srb polypeptides that functions as a negative regulator of activated transcription. Mol Cell. 2(2):213-22. Sutherland A (2003) Mechanisms of implantation in the mouse: differentiation and functional importance of trophoblast giant cell behaviour. Dev Biol. 258 (2):241–251. Swanson AM, David AL (2015) Animal models of fetal growth restriction: Considerations for translational medicine. Placenta. 36(6):623-30. Szilagyi Z, Grallert A, Nemeth N, Sipiczki M (2002) The Schizosaccharomyces pombe genes sep10 and sep11 encode putative general transcriptional regulators involved in multiple cellular processes. Mol Genet Genomics. 268(4):553-62. Taatjes DJ, Näär AM, Andel F, Nogales E, Tjian R (2002) Structure, function, and activator-induced conformations of the CRSP coactivator. Science. 295(5557):1058- 62. Takahashi H, Parmely TJ, Sato S, Tomomori-Sato C, Banks CA, Kong SE, Szutorisz H, Swanson SK, Martin-Brown S, Washburn MP, Florens L, Seidel CW, Lin C, Smith ER, Shilatifard A, Conaway RC, Conaway JW (2011) Human mediator subunit MED26 functions as a docking site for transcription elongation factors. Cell. 146(1):92-104. Tanaka M, Gertsenstein M, Rossant J, Nagy A (1997) Mash2 acts cell autonomously in mouse spongiotrophoblast development. Dev Biol. 190(1):55-65. Tanaka T, Halicka HD, Traganos F, Seiter K, Darzynkiewicz Z (2007) Induction of ATM activation, histone H2AX phosphorylation and apoptosis by etoposide: relation to cell cycle phase. Cell Cycle. 6(3):371-6. Tavella S, Bobola N (2010) Expressing Hoxa2 across the entire endochondral skeleton alters the shape of the skeletal template in a spatially restricted fashion. Differentiation. 79(3):194-202. Tenin G, Clowes C, Wolton K, Krejci E, Wright JA, Lovell SC, Sedmera D, Hentges KE (2014) Erbb2 is required for cardiac atrial electrical activity during development. PLoS One. 9(9):e107041.

159

Thompson CM, Koleske AJ, Chao DM, Young RA (1993) A multisubunit complex associated with the RNA polymerase II CTD and TATA-binding protein in yeast. Cell. 73(7):1361-75. Tomoda K, Kubota Y, Kato J (1999) Degradation of the cyclin-dependent-kinase inhibitor p27Kip1 is instigated by Jab1. Nature. 398(6723):160-5. Tsai KL, Sato S, Tomomori-Sato C, Conaway RC, Conaway JW, Asturias FJ (2013) A conserved Mediator-CDK8 kinase module association regulates Mediator-RNA polymerase II interaction. Nat Struct Mol Biol. 20(5):611-9. Tsai KL, Tomomori-Sato C, Sato S, Conaway RC, Conaway JW, Asturias FJ (2014) Subunit architecture and functional modular rearrangements of the transcriptional mediator complex. Cell. 157(6):1430-44. Tudor M, Murray PJ, Onufryk C, Jaenisch R, Young RA (1999) Ubiquitous expression and embryonic requirement for RNA polymerase II coactivator subunit Srb7 in mice. Genes Dev. 13(18):2365-8. Tunster SJ, Jensen AB, John RM (2013) Imprinted genes in mouse placental development and the regulation of fetal energy stores. Reproduction. 145(5):R117-37. Tunster SJ, Tycko B, John RM (2010) The imprinted Phlda2 gene regulates extraembryonic energy stores. Mol Cell Biol. 30(1):295-306. Tutter AV, Kowalski MP, Baltus GA, Iourgenko V, Labow M, Li E, Kadam S (2009) Role for Med12 in regulation of Nanog and Nanog target genes. J Biol Chem. 284(6):3709-18. Tuuli MG, Longtine MS, Nelson DM. (2011) Oxygen and trophoblast biology- a source of controversy. Placenta. 32 Suppl 2:S109-18. Tyson JJ, Murray JD (1989) Cyclic AMP waves during aggregation of Dictyostelium amoebae. Development. 106(3):421-6. Uldrey M, Thorens B (2004) The SLC2 family of facilitated hexose and polyol transporters. Pflugers Arch. 447(5):480-9. Uwamahoro N, Qu Y, Jelicic B, Lo TL, Beaurepaire C, Bantun F, Quenault T, Boag PR, Ramm G, Callaghan J, Beilharz TH, Nantel A, Peleg AY, Traven A (2012) The functions of Mediator in Candida albicans support a role in shaping species-specific gene expression. PLoS Genet. 8(4):e1002613. Vlach J, Hennecke S, Amati B (1997) Phosphorylation-dependent degradation of the cyclin-dependent kinase inhibitor p27. EMBO J. 16(17):5334-44. Vogl MR, Reiprich S, Küspert M, Kosian T, Schrewe H, Nave KA, Wegner M (2013) Sox10 cooperates with the mediator subunit 12 during terminal differentiation of myelinating glia. J Neurosci. 33(15):6679-90. Vortkamp A, Lee K, Lanske B, Serge GV, Kroneberg HM, Tabint CJ (1996) Regulation of Rate of Cartilage Differentiation by Indian Hedgehog and PTH-Related Protein. Science. 273(5275):613-22.

160

Wallis GA (1996) Bone growth: coordinating chondrocyte differentiation. Curr Biol.6(12):1577-80. Watson ED, Cross JC (2005) Development of structures and transport functions in the mouse placenta. Physiology. 20:180-93. Westerling T, Kuuluvainen E, Mäkelä TP (2007) Cdk8 is essential for preimplantation mouse development. Mol Cell Biol. 27(17):6177-82. White A, Wallis G (2001) Endochondral ossification: a delicate balance between growth and mineralisation. Curr Biol. 11(15):R589-91. Wong KH, Jin Y, Struhl K (2014) TFIIH phosphorylation of the Pol II CTD stimulates mediator dissociation from the preinitiation complex and promoter escape. Mol Cell. 54(4):601-12. Word JM, Bateman RC, Presley BK, Lovell SC, Richardson DC (2000) Exploring steric constraints on protein mutations using MAGE/PROBE. Protein Sci. 9(11):2251-9. Wyngaarden LA, Vogeli KM, Ciruna BG, Wells M, Hadjantonakis AK, Hopyan S (2010) Oriented cell motility and division underlie early limb bud morphogenesis. Development. 137(15): 2551–2558.

Xu J (2005) Preparation, culture, and immortalization of mouse embryonic fibroblasts. Curr Protoc Mol Biol. 28(1):1-28. Xue L, Yi H, Huang Z, Shi YB (2011) Global Gene Expression during the Human Organogenesis: From Transcription Profiles to Function Predictions. Int J Biol Sci; 7(7):1068-1076. Yin JW, Wang G (2014) The Mediator complex: a master coordinator of transcription and cell lineage development. Development. 141(5):977-87. Yue X, Izcue A, Borggrefe T (2011) Essential role of Mediator subunit Med1 in invariant natural killer T-cell development. Proc Natl Acad Sci U S A. 108(41):17105- 10. Zeller R, Lopez-Rios J, Zuniga A (2009) Vertebrate limb bud development: moving towards integrative analysis of organogenesis. Nature Reviews Genetics. 10(12):845- 58. Zhang J, Houston DW, King ML, Payne C, Wylie C, Heasman J (1998) The role of maternal VegT in establishing the primary germ layers in Xenopus embryos. Cell. 94(4):515-24. Zhao Q, Eberspaecher H, Lefebvre V, de Crobrugghe B (1997) Parallel expression of Sox9 and Col2a1 in cells undergoing chondrogenesis. Developmental Dynamics. 209(4):377-86. Zheng B, Sage M, Cai WW, Thompson DM, Tavsanli BC, Cheah YC, Bradley A (1999) Engineering a mouse balancer chromosome. Nat Genet. 22(4):375-8.

161

Zhou G, Garofalo S, Mukhopadhyay K, Lefebvre V, Smith CN, Eberspaecher H, de Crombrugghe B (1995) A 182 bp fragment of the mouse pro alpha 1(II) collagen gene is sufficient to direct chondrocyte expression in transgenic mice. J Cell Sci.108 ( Pt 12):3677-84. Zhou J, Bondy CA (1993) Placental glucose transporter gene expression and metabolism in the rat. J Clin Invest. 91(3):845-52. Zhu W, Yao X, Liang Y, Liang D, Song L, Jing N, Li J, Wang G (2015) Mediator Med23 deficiency enhances neural differentiation of murine embryonic stem cells through modulating BMP signaling. Development. 142(3):465-76.

162