The Effects of Dlx5 and Dlx6 on Differentiation of Atdc5 Cells

THE EFFECTS OF DLX5 AND DLX6 ON DIFFERENTIATION OF ATDC5 CELLS

A Thesis

Presented to

The Faculty of Graduate Studies

The University of Guelph

DANIELLE ABERNETHY

In partial fulfillment of requirements

for the degree of

Master of Science

February 2008

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While these forms may be included Bien que ces formulaires in the document page count, aient inclus dans la pagination, their removal does not represent il n'y aura aucun contenu manquant. any loss of content from the thesis. Canada ABSTRACT

THE EFFECTS OF DLX5 AND DLX6 ON DIFFERENTIATION OF ATDC5 CELLS

Danielle Abernethy Advisor: University of Guelph, 2008 Professor A.J. Bendall

The mammalian Dlx gene family consists of six members which encode homeobox-containing proteins. These proteins are widely expressed during development; the major sites of expression include the forebrain, branchial arches, axial and appendicular skeleton, epidermis and placenta. Both Dlx5 and Dlx6 are regulators of chondrocyte differentiation during skeletogenesis, however, the mechanisms underlying this function are not known. To provide further insight into the role(s) of Dlx5 and Dlx6 in chondrocyte differentiation, a cell line that differentiates in a similar manner to in vivo chondrocytes was used. Microscopic analysis of these cells following overexpression of either Dlx5 or Dlx6 revealed a lower cell number compared to controls. Condensations were not seen and no chondrogenic nodules were formed, in contrast to control cultures.

Real-time PCR analysis indicated that Dlx6 overexpression significantly reduced the activation of Col2al and Sox9 expression but upregulated osteocalcin expression. These results indicate that Dlx6 may induce an alternate differentiation pathway of ATDC5 cells. ACKNOWLEDGEMENTS

First and foremost, I would like to thank my advisor, Dr. Andrew Bendall, for your guidance and for providing me with the opportunity to leam, explore and appreciate the field of molecular biology and genetics. Special thanks to members of the Bendall lab, notably Melissa Coubrough and Claire Hsu for making various plasmid constructs. I gratefully acknowledge Dr. Colasanti, Dr. Jones, Dr. Mosser and members of the

Department of Molecular and Cellular Biology.

To my family, thank you for all your unwavering support and love. To my fiance

Tim, thank you for helping me and being there for me over the years, in ways too numerous to count.

1 TABLE OF CONTENTS

ACKNOWLEDGEMENTS i TABLE OF CONTENTS ii LIST OF TABLES v LIST OF FIGURES vi LIST OF ABBREVIATIONS vii Chapter 1: Introduction 1 1. The Dlx Family 1 1.1 Genomic structure and evolution of Dlx genes 1 1.2 Expression of Dlx genes 6 1.2.1 Regulators pf Dlx gene expression 8 1.2.2 Dlx gene expression during early development 10 1.2.3 Dlx genes are expressed in the developing forebrain 11 1.2.3.1 Telencephalon 12 1.2.3.2 Diencephalon 12 1.2.4 Dlx genes are expressed in the branchial arches 13 1.2.5 Dlx genes are expressed in the teeth 14 1.2.6 Dlx is expressed in the sensory organs 16 1.2.9 Dlx genes are expressed in the epithelium 18 1.2.10 Dlx genes are expressed in the thymus 18 1.2.11 Dlx genes are expressed in the placenta 18 1.2.12 Dlx gene expression in cancer cells and possible role in apoptosis 19 1.2.13 Dlx gene expression in the tetrapod appendage and skeleton 19 1.3 Dlx gene function 21 1.3.1 The function of Dlx genes can be inferred by studies of Dlx knockouts 21 1.3.2 Function of Dlx genes determined by misexpression studies and identification of direct gene targets 24 1.3.2.1 Forebrain/Branchial arches 24 1.3.2.2 Epidermis 25 1.3.2.3 Placenta 25 1.3.2.4 Hematopoietic cells 26 1.3.3 The function of Dlx proteins can be mediated by protein binding partners .26 2. Endochondral Ossification 30 2.1 Physiological aspects of skeletal development 30 2.2 Growth factors regulate endochondral ossification 36 2.2.1 Bone Morphogenetic Proteins : 36 2.2.2 Indian Hedgehog/Parathyroid Hormone-related Protein 36 2.2.3 Other growth factors 37 2.3 Transcription factors regulate cell identity and differentiation during endochondral ossification 38 2.3.1 Sox genes 38 2.3.2 Runx2 39 2.3.3 Osterix .'....' 40 2.3.4 Dlx proteins 40

n Chapter 2: The role of Dlx5 and Dlx6 in chondrogenesis in ATDC5 cells 43 1. Introduction 43 1.1 Objectives 43 1.2 A chondroprogenitor cell line 44 1.3 Chondrogenesis in the ATDC5 cell line 45 2. Materials and Methods 46 2.1 Common laboratory reagents 46 2.2Plasmids ....46 2.3 Cell culture 47 2.4 RNA extraction, cDNA synthesis and RT-PCR 48 2.5 Real-Time PCR ...48 2.6 Cell staining and microscopy 50 2.7 Western blotting 50 3. Results 53 3.1 Selection of the ATDC5 cell line as a chondrogenic model system 53 3.2 Overexpression of Dlx5 andDlx6 54 3.3 Dlx5 and Dlx6 overexpression resulted in decreased cell number and suppressed chondrogenic nodule formation 54 3.4 Real-time PCR analysis of gene expression in ATDC5 cells overexpressing Dlx5 andDIx6 61 3.4.1 Induction profile of ZZ/JS-infected cultures 66 3.4.2 Dlx6 suppresses Col2al activation 73 3.4.3 CollOal expression is not significantly changed by the expression of Dlx5 mdDlx6 73 3.4.4 Both Dlx5 and Dlx6 upregulate expression of osteocalcin 73 3.4.5 Osteopontin expression is slightly downregulated by Dlx6 overexpression.74 3.4.6 Sox9 expression is significantly downregulated by both Dlx5 and Dlx6 74 3.4.7 Overexpression of Dlx5 and Dlx6 has no effect on Runx2 expression 74 3.4.8 Dlx5 and Dlx6 overexpression results in increased induction of Ihh expression 74 3.4.9 Dlx3 expression is differentially controlled by Dlx5 and Dlx6 75 3.4.10 Dlx6 has no effect on the expression of Dlx5 75 3.4.11 Dlx5 has no effect on the expression of Dlx6 75 4. Discussion 77 4.1 Stage of differentiation of ATDC5 cells 77 4.3 Dlx5 and Dlx6 show some opposing roles in chondrocyte differentiaton 79 4.4 Dlx6 overexpression blocks early chondrocyte differentiation and promotes late differentiation in ATDC5 cells 80 5. Future directions 82 Appendix 1: Measuring the transcriptional activity of Dlx proteins 83 Al.l Introduction.. 83 A1.2 Materials and Methods 83 Plasmids .• 83 Cell culture and transfection 84 Cell harvesting and reporter assays 84 A1.3 Results and Discussion 85

iii Dlx proteins can activate transcription from the SV40 promoter 85 Appendix 2: Mapping the transactivation domains in Dlx5 and Dlx6 88 A2.1 Introduction 88 A2.2 Materials and Methods 88 Plasmids 88 A2.3 Results and Discussion.... 91 Characterizing transactivation domains of chick Dlx5 and Dlx6 91 REFERENCES :95

iv LIST OF TABLES

Table 1: Known Dlx genes from several vertebrate species arranged according to their similarity to the murine Dlx genes

Table 2: Primers used for real-time PCR

Table 3: Summary of the effects ofDlx5 and Dlx6 overexpression on the induction of chondrogenic marker genes

v LIST OF FIGURES

Figure 1: The structure of the human DLX genes

Figure 2: Regulation of chondrocyte differentiation

Figure 3: Melting curve analysis for assayed genes: Actin (A), Col2al (B), CollOal (C), Osc (D), Osp (E), Sox9 (F), Runx2 (G), Ihh (H), Dlx3 (I), DZx5 (J) and Dlx6 (K)

Figure 4: Changes in actin expression levels over the 17 day differentiation period relative to day 0

Figure 5: Expression of chondrogenic genes in undifferentiated ATDC5 cells

Figure 6: Overexpression of Dlx5 and Dlx6 in ATDC5 cells

Figure 7: Microscope photographs of differentiating ATDC5 cells

Figure 8: Microscope photographs of differentiating ATDC5 cells stained with Alcian blue

Figure 9: Real-time analysis of chondrogenic marker genes (A) Col2al, (B) CollOal, Osc and Osp and (C) Runx2, Ihh, Sox9, Dlx3, Dlx5 and Dlx6 in ATDC5 cells infected with the LZRS control plasmid

Figure 10: Real-time analysis of chondrogenic structural genes, Col2al (A), CollOal (B), Osc (C) and Osp (D), in differentiating ATDC5 cells

Figure 11: Real-time analysis of chondrogenic transcription and growth factors, Sox9 (A), Runx2 (B), Ihh (C), Dlx3 (D), Dlx5 (E) and Dlx6 (F), in differentiating ATDC5 cells

Figure 12: Fold activation of murine Dlx proteins on the SV40 promoter

Figure 13: Chicken Dlx5 and Dlx6 polypeptides

Figure 14: Dlx5 and Dlx6 are transcriptional activators

VI LIST OF ABBREVIATIONS

3P-HSD 3 p-hydroxysteroid dehydrogenase/isomerase

AER apical ectodermal ridge

AP-2y activator protein-2y

Arx Aristaless-like

BMP bone morphogenetic protein

BP-1 p protein-1

Bsp bone sialoprotein

C/EBP|3 CCAAT/enhancer-binding protein |3

CD campomelic dysplasia

CMV cytomegalovirus

CNC cranial neural crest

CNS central nervous system

CTD C-terminal domain

Dlx Distal-less-like

DPvEF DNA replication-related element-binding factor

E Engrailed

ECM extra-cellular matrix

Ex embryonic day

FBS fetal bovine serum fez forebrain embryonic zinc finger

FGF fibroblast growth factor

vii FGFR1 FGF receptor 1

GABA y-aminobutyric acid

GAD glutamic acid decarboxylase

Gla y-carboxyglutamic acid

GnRH gonadotropin-releasing hormone

Grip lb glutamate receptor interacting protein lb

HD homeodomain

HEK human embryonic kidney

HH Hamburger-Hamilton stage

HMG high mobility group

HOM homeotic

Hox homeobox

HSV-1 herpes simplex virus-1

ICAM intercellular adhesion molecule

IGF1 insulin growth factor 1

IGF1R IGF1 receptor

Ihh Indian Hedgehog

IR insulin receptor

IRD interspersed repeat domain

LGE lateral ganglionic eminence

LTR long terminal repeats

MAGE melanoma antigen

MASH-1 mammalian achaete-schute homologue-1

Vlll MGE medial ganglionic eminence

MHD MAGE homology domain

Msh muscle segment homeobox gene

Msx Msh-like

N-cadherin neural cadherin

NCAM neural cell adhesion molecule

NF-Y nuclear factor-Y

NLS nuclear localization signal

NRP-2 neurophilin-2

NTD N-terminal domain

Osc osteocalcin

Osp osteopontin

Osx Osterix

P75NTR p75-neurotrophin receptor

PAGE polyacrylamide gel electrophoresis

PCR polymerase chain reaction

PDZ post-synaptic density protein

PEI polyethylenimine

PKC protein kinase C

PTHrP parathyroid hormone-related peptide

RNAi RNA interference

RT reverse transcription

SDS sodium dodecyl sulphate

IX SHFM split hand/split foot malformation

Sox9 SRY-type HMG box 9

SP-1 specificity protein-1

SV40 simian virus 40

TBP TATA-binding protein

TDO tricho-dento-osseus syndrome

TGF-p1 transforming growth factor-(3

V VP16

VEGF vascular endothelial growth factor

VP16 virion protein 16

ZLI zona limitans intrathalamica

ZPA zone of polarizing activity

X Chapter 1: Introduction

1. The Dlx Family

1.1 Genomic structure and evolution of Dlx genes

Homeobox genes encode a large group of homeodomain-containing transcription factors that regulate axial patterning, segment or cell identity and proliferation. The 180 bp homeobox-encoding motif was originally discovered in the genes of the Drosophila homeotic (HOM-C) complexes (Krumlauf, 1994). It encodes a highly conserved 60 amino acid DNA-binding domain that forms a helix-turn-helix motif. Three helices form a globular structure having an N-terminal extension: helix 3 is referred to as the recognition helix (reviewed in (Banerjee-Basu and Baxevanis, 2001). All homeodomains bind a core recognition sequence, TAAT. Two separate regions of the homeodomain contact this sequence: the N-terminal arm makes contacts in the minor groove of DNA at the 5' end of the TAAT, and the recognition helix binds in the major groove at the 3' end.

(Banerjee-Basu and Baxevanis, 2001; Kissinger et al., 1990). In vertebrates, there are several classes of homeobox genes, classified by conserved variation in their homeoboxes

(Banerjee-Basu and Baxevanis, 2001). The most well known is the HOX class, homologues of the Drosophila HOM clusters. In humans, there are 39 Hox genes, arranged in four clusters: HoxA, HoxB, HoxC and HoxD. Mutations within these genes cause the transformation of one body structure into another. It is believed that this class originated from tandem duplications of a common ancestral HOM/Hox gene. Other classes include Paired, LIM, POU and extended Hox. The class most closely related to

Hox is the NK class, which includes CSX and NKX genes, and also consists of a subclass

1 called DL (Distal-less-like). The DL class includes six Dlx and three Msx members

(Banerjee-Basu and Baxevanis, 2001).

The Dlx (Distal-less-like) genes were originally identified through their similarity

to the Drosophila gene Distal-less. This protein is expressed in the head and distally in

appendages of the fly and is required for distal limb development (Cohen et al., 1989). In

mammals, the Dlx genes exist as three pairs of closely linked, convergently transcribed

genes: Dlxl/Dlx2, Dlx3/Dlx4 and Dlx5/Dlx6 (Figure 1). Mammalian Dlx genes are all

composed of three exons and two introns, with the homeodomain spanning exons two and

three (Sumiyama et al., 2002). The location of the introns has been shown to be conserved

within all vertebrates (Stock, 2005). Each of the bigene clusters is distantly linked to one

of the Hox clusters: Dlxl/Dlx2 is linked to the HoxD cluster on chromosome 2 (in

humans), Dlx3/Dlx4 is linked to the HoxB cluster on chromosome 17 and Dlx5/Dlx6 is

linked to the HoxA cluster on chromosome 7 (Nakamura et al., 1996; Simeone et al.,

1994; Sumiyama et al., 2003). The HoxC cluster may have once had a Dlx pair linked to

it; if so, it was subsequently lost during vertebrate evolution (Stock et al., 1996). Between

orthologues and paralogues, the Dlx homeodomain is highly conserved, whereas the rest of the sequence is relatively divergent (Bendall and Abate-Shen, 2000; Sumiyama et al.,

2003). For example, Dlx5 and Dlx6 share 91.7% amino acid similarity within the homeodomain, but their N-terminal and C-terminal domains show only 34.6% and 38.8% similarity, respectively (Hsu et al., 2006). This is especially true of Dlx4 which has been shown to be more divergent than the rest of the family and is evolving about ten times more rapidly than other Dlx members (Nakamura et al., 1996; Sumiyama et al., 2002).

2 Figure 1: The structure of the human DLX genes. The sizes of intergenic regions are shown. Boxes indicate exons. The arrows show the direction of transcription. Dlxl Dfx2

I2.3kb -H— V

Dix6 DIxS on 10.7kb VX7 -V- II

DIx4 Dlx3 n n 17.6kb n -//- V

4 This is likely due to reduced selection pressure on Dlx4, especially within eutherian

mammals, resulting in reduced functionality (Coubrough and Bendall, 2006). The Dlx

genes can be further divided into two clades due to sequence similarity in amino- and

carboxy-terminal domains: Dlxl/Dlx4/Dlx6 and Dlx2/Dlx3/Dlx5 (Stock et al., 1996), with

one member from each bigene pair in each clade.

In Drosophila and other protostomes, such as Caenorhabditis elegans (ceh-43), there is only one Distal-less related gene; however, in vertebrates such as humans and zebrafish, there are between 6-8 Dlx genes. It is believed that an ancestor of vertebrates had a tandem duplication of the original Distal-less gene, followed by several large chromosomal duplications, leading to the full complement seen in vertebrates today. The bigene arrangement found in all species with more than one Distal-less gene supports the tandem duplication theory. The earliest example of this duplication is in the ascidian

Ciona intestinalis of the urochordate subphylum. It was found to have a single Dlx bigene cluster, which would make the tandem duplication of Dlx before the common ancestor of all the extant chordates (Caracciolo et al., 2000). However, of the euchordata, amphioxus was shown to only have one Dlx gene, likely haying lost the second one in that lineage

(Sumiyama et al., 2003). The lamprey, a jawless vertebrate not belonging to the gnafhostome lineage, was found to have four Dlx genes, two of which are in a bigene arrangement, and two others as unpaired genes (Neidert et al., 2001). It is believed that the unpaired Dlx genes were once part of bigene clusters in an ancestral jawless fish

(Neidert et al., 2001; Sumiyama et al., 2003). Finally, in a gnathostome ancestor there existed three bigene clusters of Dlx genes, each linked to a Hox cluster. The Dlx bigene clusters and the Hox clusters likely underwent duplication in the same genomic events

5 (Stock et al., 1996). Zebrafish and other teleost fish have additional Dlx genes, likely due to a genome duplication event within that lineage (Sumiyama et al., 2003).

Currently, all vertebrates, and many invertebrates studied, have been found to have some complement of Distal-less related genes, including the mouse (Nakamura et al., 1996; Porteus et al., 1991; Price et al., 1991; Robinson and Mahon, 1994; Simeone et al., 1994), human (Nakamura et al., 1996; Selski et al., 1993; Simeone et al., 1994), rat

(Shirasawa et al., 1994; Zhao et al., 1994), newt (Beauchemin and Savard, 1992),

Xenopus (Asano et al., 1992; Dirksen et al., 1993; Papalopulu and Kintner, 1993),

Eleutherodactylus coqui (Fang and Elinson, 1996), zebrafish (Akimenko et al., 1994), lamprey (Neidert et al., 2001) and also invertebrates such as ascidians (Caracciolo et al.,

2000). With so many Dlx genes discovered in such a short period of time, the nomenclature of the family is complex. Table 1 includes a list of the Dlx genes from several species arranged according to their similarity to the mouse Dlx genes.

1.2 Expression of Dlx genes

Expression analyses of Dlx genes are complicated by the presence of multiple transcripts of some members. Dlxl has six different transcripts, including one anti-sense

(McGuinness et al., 1996), Dlx5 has three different transcripts, only one of which encodes a homeodomain, Dlx6 has one sense and one anti-sense RNA (Liu et al., 1997; Yang et al., 1998) and Dlx4 has three alternately spliced transcripts (often referred to by different names: Dlx4, Dlx7 and P Protein-1 (BP-1) (Nakamura et al., 1996). These alternate transcripts are thought to play a role in the control of Dlx gene expression or modulation of protein function.

6 Table 1: Dlx genes from several vertebrate species arranged according to their similarity to the murine Dlx genes. Earlier names are shown in parentheses.

Mouse Dlxl Dlx2 Dlx3 Dlx4 DlxS Dlx6 (Tes-1) (Dlx7, BP-1) Human DLX1 DLX2 DLX3 DLX4 DLX5 DLX6

(DLX7,

DLX8)

Chicken Dlxl Dlxl Dlx3 Dlx5 Dlx6

Rat rDlx

Zebrafish dlxl a dlx2a dlx3b dlx4b dlx5a dlx6a

(dlxl) (dlx2) (dlx3) (dlx7) (dlx4) (dlx6)

dlx2b dlx4a

(dlx5) (dlx8)

Xenopus X-DLL1 X-dll2 X-dll3 X-dll

X-dll4

E. coqui Dlxl Dlx2 Dlx3 Dlx4

Newt NvHBox4 NvHBox5

7 1.2.1 Regulators of Dlx gene expression

Dlx gene expression is controlled by intergenic enhancers that lie between the

bigene clusters. There is a high level of sequence conservation in orthologous regions, but

not in paralogues, even though there is some overlap in expression. In the Dlxl/Dlx2,

Dlx3/Dlx4 and Dlx5/Dlx6 intergenic regions, several conserved enhancers have been

found which direct tissue specific expression. Between Dlxl and Dlx2, one enhancer was

shown to direct forebrain expression (Ghanem et al., 2003), while two others were

responsible for expression in the branchial arches (Ghanem et al., 2003; Park et al., 2004).

The forebrain specific enhancer is bound by MASH-1 (mammalian achaete-schute

homologue-1), which regulates expression along with numerous other transcription

factors (Poitras et al., 2007). There is one region in the Dlx3/Dlx4 intergenic sequence

responsible for branchial arch expression (Sumiyama et al, 2003) and in the Dlx5/Dlx6

intergenic region, two enhancers have been found to direct forebrain expression and one

for the branchial arches (Ghanem et al., 2003; Park et al., 2004).

One of the primary regulatory mechanisms of Dlx gene expression is auto- and

cross-regulation by other Dlx family members. There are Dlx-binding sites in all three

intergenic regions (Poitras et al., 2007; Sumiyama and Ruddle, 2003; Zerucha et al.,

2000): for example, Dlxl, Dlx2 and Dlx5 can all induce transcription from the Dlx5/Dlx6

intergenic enhancer (Stuhmer et al., 2002) while Dlx2 was shown to specifically bind at the sequence 5'(A/C/G)TAATT(G/A)(C/G)3' within this enhancer (Zerucha et al., 2000).

Dlxl and Dlx2 can bind the Dlx5/Dlx6 enhancer in the embryonic striatum and Dlx2 can bind in the newborn retina, showing cell-specific transcription (Zhou et al., 2004). It was

also shown that a complex of Dlx2 and an RNA molecule, Evf-2, transcribed from the

8 Dlx5/Dlx6 intergenic region, can bind to the Dlx5/Dlx6 enhancer to increase transcription

(Feng et al., 2006).

Study of the upstream effectors of the Dlx genes has identified numerous growth factors as regulators, notably, the bone morphogenetic proteins (BMPs) of the transforming growth factor-P (TGF-(3) superfamily. Epithelial expression of Dlx2 in the first branchial arch is controlled by BMP4 and mesenchymal expression is regulated by fibroblast growth factor 8 (FGF8) (Thomas et al., 2000). In vivo, Dlx5 is regulated by

BMP2 in chick calvarial cells (Holleville et al., 2007), BMP4 and BMP7 in the chick skull (Holleville et al., 2003), and FGF2 in the ectoderm and mesoderm of ectopic limb buds (Ferrari et al., 1999). In cell lines, Dlx5 is responsive to both BMP2 and BMP7 in

CHIOP/2 cells (Shea et al., 2003), BMP2 and BMP4 in C2C12 myoblasts (Lee et al.,

2003) and BMP4 in osteoblasts (Miyama et al., 1999); Dlx5 and Dlx6 are activated by

FGF8 and FGF9 (Park et al., 2004) and Dlx3 is induced by BMP2 in osteoblasts (Hassan et al., 2004; Hassan et al., 2006). Dlx genes can also be repressed by BMP antagonists such as Noggin (Holleville et al., 2003) and chordin (Luo et al., 2001a).

Other methods of control are mediated by proteins such as fez (forebrain embryonic zinc finger) in zebrafish, which enhances expression of dlx2 and dlx6 (Yang et al., 2001). The Dlx3 promoter has several cell-specific binding elements. In keratinocytes, the protein nuclear factor Y (NF-Y) binds to a CCAAT element and another protein, specificity protein 1,(SP-1), binds a specific sequence to activate Dlx3 expression (Park and Morasso, 1999). In the placenta, CCAAT/enhancer-binding protein p (C/EBPP) binds to the CCAAT element to drive basal expression (Holland et al., 2004). In epidermal cells, Dlx3 was shown to be activated by increasing extracellular Ca + levels through a

9 protein kinase C (PKC) pathway (Park et al., 2001). PKC phosphorylates Dlx3 and

inhibits its DNA-binding ability (Park et al., 2001). This indicates a level of post-

translational control for at least some of the Dlx proteins.

1.2.2 Dlx gene expression during early development

In many species, Dlx genes are expressed at very early stages of development. In

Xenopus, X-dll is expressed in oocytes and the early embryo (Asano et al., 1992). At the

blastula stage, X-dll2 from Xenopus is expressed in the ectoderm (Woda et al., 2003), and

in chick Dlx5 is expressed in the area pellucida (Pera et al., 1999). During gastrulation, X-

dll2 and X-dll3 are expressed in the ectoderm of Xenopus (Dirksen et al., 1994; Luo et al.,

2001b), specifically the dorsal ectoderm (Woda et al., 2003), dlx3 is expressed in the

ectoderm of zebrafish (Akimenko et al., 1994) and in chick Dlx3 is expressed at the distal

tip of the primitive streak (Pera et al., 1999). In the neurula, X-dll is expressed in the head

region and X-dll2 in the non-neural ectoderm of Xenopus (Woda et al., 2003). The neural

plate, which eventually forms the central nervous system (CNS), expresses several Dlx

genes: Dlx5 is expressed in mouse around E7.25 (embryonic day 7.25); (Yang et al.,

1998), X-dlU fxora.Xenopus (Luo et al., 2001b) and Dlx4 from the direct developing frog

Eleutherodactylus coqui (Fang and Elinson, 1996) are also expressed within the neural

plate. In chick, Dlx5 is expressed in ectodermal cells that border the neural plate (Pera et

al., 1999). As development continues, expression of Dlx5 within the mouse moves to the

rostral ectoderm in the interior neural ridge (E7.5), to the lateral margins of the neural

plate and the pre-migratory neural crest (E7.75) (Yang et al., 1998). In chick, at HH9

(Hamburger-Hamilton stage 9), Dlx3 is expressed in the anterior neural fold and Dlx5 is

expressed in the prospective neural crest at HH10 (Pera et al., 1999). Presumptive neural

10 crest cells in zebrafish express dlx2 at 12h (Akimenko et al., 1994) and in E. coqui Dlx2 is expressed in the same region (Fang and Elinson, 1996). The cranial neural crest (CNC) cells, which eventually migrate to form the facial structures and the branchial arches, begin expressing Dlx5 in mouse at E8.5 (Yang et al., 1998) and express X-dll2 mXenopus

(Luo et al., 2001b).

1.2.3 Dlx genes are expressed in the developing forebrain

In all species studied, four Dlx genes (Dlxl, Dlx2, Dlx5 and Dlx6) are expressed in the developing forebrain, with no expression seen in the midbrain or hindbrain. Neither

Dlx3 nor Dlx4 are expressed in the CNS of mammals, however, Dlx3 is expressed in the forebrain of the chick embryo (Zhu and Bendall, 2006). In zebrafish, expression of dlx genes starts as early as 13h for dlx2, dlx4 (Akimenko et al., 1994), and dlxl and dlx5

(Ellies et al., 1997). In Xenopus, X-DLL1 is found in the forebrain around stage 24

(Dirksen et al., 1993), and in mouse Dlxl, Dlx2, Dlx5 and Dlx6 are all expressed in the forebrain around E9.0 (Andrews et al., 2003; Simeone et al., 1994). The forebrain is divided into the telencephalon (eventually forming the cerebral cortex, basal ganglia and hippocampus) and the diencephalon (forms the thalamus, epithalamus, pretectum and possibly the hypothalamus). In both the telencephalon and diencephalon there are three zones: the ventricular zone (containing mainly undifferentiated, proliferating cells), the subventricular zone (with proliferating cells), and the mantle zone (consisting of post mitotic, differentiating cells). In the mouse, Dlxl and Dlx2 are found mainly in the ventricular and subventricular zones, whereas, Dlx5 is expressed in both the subventricular and mantle zones; Dlx6 is found only in the mantle (Liu et al., 1997).

11 1.2.3.1 Telencephalon

In the mouse telencephalon, the first Dlx gene expressed is Dlx2 in the basal region at E9.0-E9.75 (Bulfone et al., 1993a). This is quickly followed by Dlx5 in the ventral subcortical areas at E9.5 (Eisenstat et al., 1999). Expression of Dlx5 is then expanded to the ganglionic eminences, both the lateral ganglionic eminence (LGE) and medial ganglionic eminence (MGE). Cells from these areas will eventually migrate to the cortex and become y-aminobutyric acid (GABA)ergic neurons. Expression of both Dlxl and Dlxl is found in this region by E12.5 (Bulfone et al., 1993b) and Dlx5 is expressed by

E10 and continues to E12.5 (Simeone et al., 1994). Around the same time, Dlxl and Dlx2 are also found in the basal ganglia and the septum (Fernandez et al., 1998). As differentiation proceeds (from E18.5-P7) both Dlxl and Dlx2 are expressed in the cerebral cortex and the hippocampus (Porteus et al., 1991; Saino-Saito et al., 2003).

During early differentiation, Dlx2 expression is stronger than Dlxl. After birth the opposite is seen, with much more Dlxl transcription compared to Dlx2 (Saino-Saito et al.,

2003).

1.2.3.2 Diencephalon

Expression in the diencephalon begins with Dlxl and Dlx2 in the primordial ventral thalamus from E9.0-E9.75 (Andrews et al., 2003; Bulfone et al., 1993a). Also,

Dlx5 and Dlx6 are first expressed in both the ventral and rostral regions around E9.5-

E10.5 (Eisenstat et al., 1999; Simeone et al., 1994; Yang et al., 1998). By E12.5, Dlx5 and

Dlx6 expression is also found in the ventral thalamus (Simeone et al., 1994). Dlxl, Dlx2 and Dlx5 are expressed in the hypothalamus around E12.5 (Eisenstat et al., 1999; Saino-

Saito et al., 2003). Expression of Dlxl and Dlx2 is found in the ganglionic eminences

12 around El 1.5 (Yang et al., 1998). While no Dlx genes have been found within the zona

limitans intrathalamica (ZLI) around El2-13 both Dlxl and Dlx2 expression is detected

ventral to it (Price et al., 1991). After birth (PO), all four genes are still expressed in the

ventral thalamus, and Dlxl and Dlx2 are expressed within the ganglionic eminences

(Jones and Rubenstein, 2004). In the adult mouse, Dlxl (but not Dlx2) and Dlx5 are

expressed in the ventral thalamus and Dlx5 is also expressed in the epithalamus (Jones

and Rubenstein, 2004).

1.2.4 Dlx genes are expressed in the branchial arches

The branchial arches (or pharyngeal arches) are embryonic structures that develop

into the structures of the face, ear and neck. The cells within these arches originate from

migrating CNC cells. In mammals, there are six branchial arches, however, since the fifth

branchial arch does not develop into any structures they are numbered I, II, III, IV, and VI

(Graham, 2003; Graham et al., 2005). The first branchial arch is divided into two regions:

the maxillary and mandibular processes. This arch forms several structures: the maxilla,

the mandible, the auditory tube, jaw muscles, Meckel's cartilage and the trigeminal nerve.

The second arch (also called the hyoid arch) develops into the hyoid and facial muscles.

The third arch forms part of the hyoid and the thymus and the fourth and sixth arches

form the thyroid, the larynx and neck muscles (Graham, 2003; Graham et al., 2005).

All six Dlx genes from mice are expressed in the branchial arches. They form a

nested expression pattern in both the mediolateral axis and the proximodistal axis. In the mediolateral axis, Dlx5 and Dlx6 are expressed most medially, then Dlxl and Dlx2 with

Dlx3 and Dlx4 most laterally (Qiu et al., 1997). In the proximodistal axis, Dlx3 and Dlx4 expression is the most distally restricted, with Dlx5 and Dlx6 more proximal and Dlxl

13 and Dlx2 expressed along the entire axis (Qiu et al., 1997). Other species also show similar branchial arch expression: in Xenopus, expression of X-dlU, X-dll4 (Papalopulu and Kintner, 1993), X-dll2 (Dirksen et al., 1994) and X-DLL1 (Dirksen et al., 1993) have all been detected; in zebrafish, dlx2, dlx3 and dlx4 are all expressed (Akimenko et al.,

1994), in E. coqui Dlx2 and Dlx4 are transcribed in the branchial arches (Fang and

Elinson, 1996) and in chick all five known Dlx genes are expressed in the branchial arch by HH13 (Brown et al., 2005; Pera and Kessel, 1999; Pera et al., 1999).

In mouse, Dlx5 and Dlx6 are expressed first at E8.5 in the mesenchyme of the first arch, continuing through E9.0 with weaker expression in the other arches (Simeone et al.,

1994; Yang et al., 1998). Dlx2 expression can be detected at E8.75 in the first and second arch mesenchyme, with the expression domain widening to all four arches from E9.0-9.75

(Bulfone et al., 1993a; Clouthier et al., 2000; Thomas et al., 2000). Dlxl is expressed slightly later than Dlx2 (E9.5), but in the same regions (Qiu et al., 1997). Dlx3 is expressed around E9.5 in the distal part of the branchial arches, specifically in the mandibular part of the first arch (Clouthier et al., 2000; Robinson and Mahon, 1994). By

El 0.5, Dlxl and Dlx2 are expressed in both the mandibular and maxillary processes of the first arch; all other Dlx genes are expressed only in the mandibular process (Qiu et al.,

1997; Thomas etal., 2000).

1.2.5 Dlx genes are expressed in the teeth

The developing tooth starts as an aggregation of cells derived from the ectoderm of the first branchial arch and the ectomesenchyme of neural crest cells, called a tooth bud. The earliest cell aggregates are called the dental lamina which connects the developing tooth bud to the epithelial layer of the mouth. The tooth bud is divided into

14 three parts: the enamel organ, the dental papilla and the dental follicle. The enamel organ consists of the inner and outer epithelium which will eventually form the ameloblasts, which make the enamel of the tooth. The dental papilla produces odontoblasts, which are dentin-forming cells, and mesenchymal cells within the papilla eventually form the tooth pulp. The dental follicle forms the cementoblasts, osteoblasts and fibroblasts.

Dlx genes are expressed in all of the regions of the developing teeth. Expression of Dlx genes begins around El2.5 when Dlxl and Dlx2 transcripts are found in the region where teeth will develop (Bulfone et al., 1993b; Porteus et al., 1991; Thomas et al., 2000).

At the same time, Dlx2 and Dlx3 are found in the dental lamina of both the molar and incisor primordia (Zhao et al., 2000). Both Dlx3 and Dlx5 are expressed in the epithelial and mesenchymal cells of the developing teeth (Davideau et al., 1999; Ghoul-Mazgar et al., 2005). By E14.5-E15.5 Dlxl and Dlx6 are expressed in the dental follicle, Dlx3, Dlx5 and Dlx4 transcription can be found in the dental papilla and Dlx5 expression is also found in the cervical loop (Zhao et al., 2000). By E16.5-E17.5 all six Dlx genes are expressed within the molars, Dlxl is still expressed in the dental follicle, Dlx3 and Dlx7 are now expressed within the cervical loop, Dlx5 and Dlx6 are expressed in the dental papilla and Dlx2 is expressed in the enamel epithelia (Zhao et al., 2000). Later in differentiation, Dlxl is expressed in the ameloblast layer (Robinson and Mahon, 1994),

Dlx3 is expressed in both odontoblasts and ameloblasts (Ghoul-Mazgar et al., 2005;

Robinson and Mahon, 1994) and Dlx5 is expressed within the osteogenic mesenchyme around the tooth (Levi et al., 2006).

15 1.2.6 Dlx is expressed in the sensory organs

Dlx genes are expressed in all of the sensory placodes of the developing sensory organs, including the eye, olfactory organs and the inner ear.

Only a few Dlx genes have been shown to be expressed in the developing eye. At

E10, mouse Dlxl is expressed in the optic chiasma, the region where the optic nerves cross in the brain (Price et al., 1991). In Xenopus, X-DLL1 andX-dll4 are expressed in the ganglion layer of the developing retina and X-dll3 andX-dll4 are both expressed in the optic chiasma ridge (Dirksen et al., 1993; Papalopulu and Kintner, 1993).

The inner ear is formed by an area of thickening in the embryonic epithelial layer, called the otic (or auditory) placode, which invaginates, forming the otic pit. This pit becomes enclosed to form the otic vesicle (or otocyst), which eventually develops into the vestibular organs required for both hearing and equilibrium. Dlx genes are expressed in the developing ear in all vertebrate species. In Xenopus, X-DLL1, X-dll2 and X-dllS are all expressed in the otic vesicles (Dirksen et al., 1993; Dirksen et al., 1994; Papalopulu and

Kintner, 1993). In zebrafish, dlx3 and dlxl are expressed in the otic placodes (Akimenko et al., 1994; Ekker et al., 1992; Ellies et al., 1997) and then later dlx3, dlx4 and dlx6 expression is found in the otic vesicle (Ekker et al., 1992; Ellies et al., 1997). In chick,

Dlx3 expression is found first in the otic placode, followed by that of Dlx5 (Brown et al.,

2005; Pera and Kessel, 1999; Pera et al., 1999). This expression continues into the otic pit where Dlxl, Dlx2 and Dlx6 are also expressed (Brown et al., 2005). Dlx3, Dlx5 and Dlx6 are all expressed in the otic vesicle, while, Dlxl, Dlx2 and Dlx5 are expressed in the cochlear and vestibular nerves of the inner ear (Brown et al., 2005). E. coqui also shows

Dlx expression during ear development as Dlx2 is expressed in the otic vesicle (Fang and

16 Elinson, 1996). In the mouse, Dlx2 is first expressed in the ectoderm of the otic pit at

E8.75 and continues into the otic vesicle by E9.75 (Bulfone et al, 1993b). Around the same time, Dlx5 and Dlx6 are being expressed in the otic vesicle from E8.5-E9.0 and become restricted to the vestibular region by El2.5 (Robinson and Mahon, 1994).

In the developing olfactory organ, a thickening of the epithelium forms the olfactory (or nasal) placode which eventually becomes the vomeronasal organ in the palate and the olfactory epithelium of the nose. This is a specialized epithelial tissue, which is covered in olfactory receptors which mediate the sense of smell. Sensory input is sent through olfactory receptor neurons to the olfactory bulb, a structure in the brain that filters these inputs. In Xenopus, X-dll2 and X-dll3 are expressed in the olfactory placodes

(Dirksen et al., 1994; Papalopulu and Kintner, 1993). In zebrafish, dlx3, dlx7, dlx4 and dlx6 are all expressed in the olfactory placodes (Akimenko et al., 1994; Ellies et al.,

1997), as is Dlx4 in E. coqui (Fang and Elinson, 1996). In chick, Dlx3 expression is first detected in the olfactory placodes, which then expands to the olfactory pits, while Dlx5 is only expressed in the olfactory placodes (Pera and Kessel, 1999; Pera et al., 1999). In the mouse, Dlx5 and Dlx6 are expressed in the olfactory placodes from E8.0-E9.0 and from

E10.5-E13.5 are then expressed in the olfactory epithelium of the nasal cavity (Levi et al.,

2006; Simeone et al., 1994; Yang et al., 1998). Dlx2 is expressed in the olfactory bulb by

E17.5 with expression of both Dlxl and Dlx2 continuing in this region until adulthood

(Porteus et al., 1991; Saino-Saito et al., 2003). Both Dlx2 and Dlx5 are expressed in the developing vomeronasal organ which is responsible for pheromone sensing (Levi et al.,

2006; Porteus et al., 1991).

17 1.2.9 Dlx genes are expressed in the epithelium

Many Dlx genes are expressed within the developing epidermis, feathers and hair.

In both Xenopus and newt, expression occurs in the epidermis (for X-dll2 and NvHBox4 and NvHBox5 respectively) (Beauchemin and Savard, 1992; Dirksen et al., 1994;

Morasso et al., 1994). In the mouse, Dlx3 is expressed in the matrix cells of whisker follicles and suprabasal cells in interfollicular epidermis (Robinson and Mahon, 1994).

Dlx3 is also expressed in terminally differentiated keratinocytes (Morasso et al., 1996). In the chick, Dlx2 and Dlx3 are expressed in the epidermal placodes and the epithelium of the feather bud, and Dlx5 is expressed in the bud epidermis (Rouzankina et al., 2004).

Dlx5 from chick is also expressed in the prospective non-neural epidermis (Pera et al.,

1999).

1.2.10 Dlx genes are expressed in the thymus

Dlxl, Dlx2, Dlx3, Dlx5 and Dlx6 are all expressed in the thymus during development (Woodside et al., 2004). Dlxl and Dlxl are found in the thymocytes and

Dlx3 and Dlx5 are expressed in stromal epithelial cells of the thymus (Woodside et al.,

2004).

1.2.11 Dlx genes are expressed in the placenta

Murine Dlx3 is expressed in several areas of the placenta: the ectoplacental cone and the chorionic plate, and later in the labyrinthine trophoblasts of the chorioallantoic placenta (Morasso et al., 1999). Human DLX4 has also been shown to be expressed in the placenta (Quinn et al., 1997).

18 1.2.12 Dlx gene expression in cancer cells and possible role in apoptosis

Dlx4 has been shown to be up-regulated in several cancers and cancerous cell lines, including: breast cancer (Man et al., 2005), myeloid and lymphoid leukemias (Haga et al., 2000), and lymphoma (Haga et al., 2000), and in erythromyeloid (Haga et al.,

2000), hematopoietic, leukemia and choriocarcinoma cell lines (Shimamoto et al., 1997;

Sun et al., 2006). DLX1 was also expressed in human non-lymphoid hematopoietic cell lines (Chiba et al., 2003). DLX4 overexpression in a B-cell line was able to inhibit apoptosis (Shimamoto et al., 2000). It also up-regulated intercellular adhesion molecule-1

(ICAM-1) andICAM-2, leading to increased cell adhesion properties (Shimamoto et al.,

2000). However, it was also found that Dlxl, Dlx2, Dlx3 and Dlx4 were expressed at higher levels in cell lines that are more sensitive to apoptotic induction (Ferrari et al.,

2003). The role of various Dlx genes in cell growth and apoptosis needs to be studied further.

1.2.13 Dlx gene expression in the tetrapod appendage and skeleton

Within the apical ectodermal ridge (AER), a major signaling center during tetrapod appendage development, expression of several dlx members from zebrafish are found: dlxl, dlx3 and dlx4 in the anterior region and dlxl in the posterior region

(Akimenko et al., 1994; Ellies et al., 1997). In E. coqui, Dlx4 expression is found within the AER and in chick both Dlx2 and Dlx5 are expressed (Fang and Elinson, 1996; Ferrari et al., 1995; Pera and Kessel, 1999). Dlx expression is also found throughout development of the appendage bud itself. In zebrafish, dlxl is expressed in the presumptive fin bud, dlx2, dlx5, dlx4 and dlx6 are expressed in the pectoral fin bud, and dlx3 and dlxl are expressed in the median fin fold (Akimenko et al., 1994; Ellies et al., 1997). In E. coqui,

19 both Dlx2 and Dlx4 are expressed within the limb bud (Fang and Elinson, 1996; Pera and

Kessel, 1999). In chick, Dlx3, Dlx5 and Dlx6 are all expressed within the limb bud

(Bendall et al., 2003; Hsu et al., 2006; Pera and Kessel, 1999). Dlx5 gets progressively more restricted in expression, starting in the surface ectoderm of the trunk, then in the ectoderm overlying the prospective wing and leg-forming regions, then the distal tip of the ventral ectoderm, then it is finally restricted to limb mesenchyme (Ferrari et al.,

1999). Later, it is found in the distal limb bud, then the mesenchyme of the anterior limb bud, then the posterior mesenchyme and lastly in the distal mesenchyme (Ferrari et al.,

1995).

Dlx genes are expressed during many stages of intramembranous and endochondral ossification. Dlx genes are expressed within developing chondrocytes. Dlx3 and Dlx5 are expressed in mesenchymal progenitors and within proliferating chondroprogenitors (Bendall et al., 2003; Ferrari and Kosher, 2002; Ghoul-Mazgar et al.,

2005; Hassan et al., 2004). Dlx3 and Dlx5 are expressed in pre-hypertrophic and hypertrophic chondrocytes and Dlx3 is expressed later during the mineralization stage

(Bendall et al., 2003; Ferrari and Kosher, 2002; Ferrari et al., 1995; Hassan et al., 2004).

The perichondrium is a specialized structure surrounding the differentiating chondrocytes, which eventually ossifies and forms the bony collar. Dlx3, Dlx5 and Dlx6 are all found within the developing perichondrium and Dlx5 expression persists in the periosteum

(Bendall et al., 2003; Ferrari and Kosher, 2002; Ferrari et al., 1995; Hassan et al., 2004;

Simeone et al., 1994).

Dlx5 is expressed in osteogenic precursors before differentiation and in mature osteoblasts (Holleville et al., 2007; Holleville et al., 2003). Dlx3 is also found in

20 osteoblasts involved in both intramembranous and endochondral ossification, although its

expression is higher in endochondral bone (Ghoul-Mazgar et al., 2005; Hassan et al.,

2004).

1.3 Dlx gene function

1.3.1 The function of Dlx genes can be inferred by studies of Dlx knockouts

The function of Dlx genes can be determined by studying the phenotype of knock

out mice. Knock-outs of Dlx genes have been done by targeting either one gene, or both

genes of a linked pair. Double Dlx gene knock-outs usually show a more severe phenotype than single ones reflecting the overlapping expression and overlapping

functions of linked gene pairs.

Dlxl null mutants are viable at birth, with few gross abnormalities, however, they

die within a month (Qiu et al., 1997). By contrast, a Dlxl null mutation causes death within a few hours, with obvious abdominal distention (Qiu et al., 1995). The Dlxl mutant also shows numerous defects in skeletal elements derived from the branchial

arches and in the dermal bones of the lateral skull wall (Qiu et al., 1995). These are likely

functions that are specific to Dlxl. A double knock-out of the two genes produces a much more drastic phenotype. Newborns die within a few hours, similar to the Dlxl null mice.

They have no maxillary molars, whereas both Dlxl and Dlxl null mice had normal teeth, making both genes essential for maxillary molar formation (Qiu et al., 1997; Thomas et

al., 1997). They also have defects in CNC-derived skeletal components of the proximal branchial arches (Qiu et al., 1997). Defects in the forebrain have been shown, including the basal ganglia and the striatum (Anderson et al., 1997; Qiu et al., 1997). This shows a

21 role for Dlxl and Dlx2 in developing teeth, in the forebrain and in patterning and

ossification of the branchial arches and the skull.

A deletion mutant of Dlx3 exists, however this deletion results in embryonic death

around day 9.5-1.0 due to placental defects (Morasso et al, 1999). Clearly, Dlx3 is

essential for placental development. However, this makes it difficult to determine later

functions of Dlx3. There have been a few studies of various natural defects. A zebrafish

mutant of both dlx3 and dlxl lacks sensory (both otic and olfactory) placodes (Solomon

and Fritz, 2002). Also, a human disease, tricho-dento-osseus syndrome (TDO) has been

mapped to the DLX3 locus (Price et al., 1998). These individuals are born with curly and

kinked hair, taurodontism (elongation of the dental pulp chamber, causing enamel

defects), cranial thickening and some other variable defects in hair, teeth and bone (Price

et al., 1998). They also have a general increase in bone mineral density caused by an

alteration in endochondral and intramembranous bone formation (Haldeman et al., 2004).

The Dlx4 knockout phenotype is not known and there have been no reports

addressing its normal function. Dlx4 has been reported as being upregulated in several cancer cell lines. Also, when Dlx4 transcripts were knocked down using RNA interference (RNAi) in two different cell lines (choriocarcinoma and erythroleukemia), its inhibition caused an increase in apoptosis, indicating that Dlx4 can suppress apoptosis

(Shimamoto et al., 2000; Sun et al., 2006).

Dlx5 knock-out mice display numerous defects. These mice are bora slightly smaller than normal and die shortly after birth with a distended abdomen, similar to Dlxl and Dlxl/Dlx2 mutants (Acampora et al., 1999). They have cranial defects due to a delayed ossification of the roof of the skull (Acampora et al., 1999; Depew et al., 1999)

22 and there are craniofacial abnormalities in regions derived from the first four branchial

arches, such as a shortening of Meckel's cartilage (Depew et al., 1999). Several sensory

organs are malformed. The olfactory and otic placodes show defects (Depew et al., 1999)

and the vestibular organ (inner ear) shows severe malformation, including an absence of

semicircular canals (Merlo et al., 2002). The simultaneous deletion of Dlx5 and Dlx6

shows similar, but more severe, abnormalities. Dlx5/6'/' mice die soon after birth. They

have no calvaria, producing exencephaly, a likely cause of their early death (Robledo et

al., 2002). They have limb defects causing missing central digits, notably in the hindlimb.

In most bones there is an absence or delay of endochondral ossification and in both

chondrocyte and osteoblast differentiation (Robledo et al., 2002). In the vestibular organ,

several structures fail to form, resulting in a complete loss of function (Robledo and

Lufkin, 2006). These observations suggest that Dlx5 and Dlx6 are necessary for limb

patterning and formation, skeletal ossification and sensory organ formation. Dlx6 seems

to be able to compensate for the lack of Dlx5 during endochondral ossification and limb

formation, but Dlx5 mutants still show defects in sensory organ and craniofacial

structures (Acampora et al., 1999). Dlx5/6'~ mice display a partial phenocopy of the human disease split hand/split foot malformation (SHFM), an autosomal dominant disease mapped to the Dlx5/Dlx6 locus characterized by missing central digits, fusion of the remaining digits, sensorineural deafness and vestibular malformations (Robledo et al.,

2002). The single Dlx6 null phenotype has not yet been published.

23 1.3.2 Function of Dlx genes determined by misexpression studies and identification of direct gene targets

Dlx genes have been shown to be important in the development of several organs: the skeleton, the forebrain, the branchial arches, the epidermis, the placenta and hematopoietic cells. Overall, Dlx is important in controlling cell migration and differentiation and in regulating epithelial:mesenchymal interactions.

1.3.2.1 Forebrain/Branchial arches

Ectopic expression of both Dlx2 and Dlx5 in the neural tube inhibited migration of cells to the branchial arches and induced aggregation of neural crest cells (McKeown et al., 2005). Also, ectopic expression of Dlx2 in the branchial arch mesenchyme induced cell adhesion molecules such as neural cadherin (N-cadherin) and neural cell adhesion molecule (NCAM), causing increased mesenchymal condensation (McKeown et al.,

2005). However, Dlxl and Dlx2 were shown to repress neurophilin-2 (NRP-2) by binding to its promoter (Le et al., 2007). NRP-2 is a receptor for class III semaphorins which inhibit neuronal migration. Down-regulating NRP-2 thus facilitates interneuron migration

(Le et al., 2007). These results indicate that Dlx can regulate cell migration through the control of cell signaling molecules. Dlx proteins also regulate many other forebrain- specific genes. The gene for gonadotropin-releasing hormone (GnRH), which is expressed in cells migrating from the nasal region to the hypothalamus, contains four conserved homeodomain-binding sites. Msxl was shown to bind and repress this gene, whereas Dlxl, Dlx2 and Dlx5 can bind these elements and increase transcription of

GnRH(Givens et al., 2005). Aristaless (Arx), is a transcription factor expressed in the ventral thalamus, hypothalamus, ganglionic eminences, subpallial telencephalon and

24 cortical GABAergic neurons (Cobos et al., 2005). Ectopic expression of Dlxl, Dlx2 and

Dlx5 was shown to up-regulate expression of Arx in the forebrain (Cobos et al., 2005).

There was also a reduction of Arx in Dlxl/Dlx2 null mice (Cobos et al., 2005). Ectopic

expression of Dlx2 and Dlx5 in the cerebral cortex induces glutamic acid decarboxylases

(GADs) which synthesize GABA (Stuhmer et al., 2002). The Wnt-1 enhancer also has a homeodomain-binding site which is required for forebrain expression. This element may be bound by either Dlx2 or Emx2 in vivo (Her et al., 1995).

1.3.2.2 Epidermis

Dlx2 misexpression in chick feather buds causes feather loss and feather bud fusions and also up-regulates expression of NCAM and tenascin (Rouzankina et al.,

2004). This indicates both a positive role (feather bud fusions indicate a promotion of feather bud development) and a negative role (lack of feather bud initiation) for Dlx2 in feather development. When Dlx3 is ectopically expressed in the basal cell layer it results in an abnormal epidermal phenotype and perinatal lethality (Morasso et al., 1996). It causes expression of profilaggrin and loricirin, which are normally only expressed in late stages of epidermal differentiation (Morasso et al., 1996). Dlx3 can therefore transform basal cells into more differentiated keratinocytes.

1.3.2.3 Placenta

Dlx3 is responsible for expression of two important placental hormone-related genes. Chorionic gonadotropin is a glycoprotein hormone consisting of an a and P subunit. The a subunit has a promoter sequence responsible for placental-specific expression. Dlx3 was found to bind this element and overexpression of Dlx3 triggered chorionic gonadotropin expression (Roberson et al., 2001). 3p-hydroxysteroid

25 dehydrogenase/isomerase (3P-HSD) is responsible for biosynthesis of steroid hormones.

In the placenta, one isoform is responsible for placental progesterone production, which is

necessary for maintenance of pregnancy. There is a trophoblast-specific enhancer with

three transcription factor binding sites. The transcription factor activator protein-2y (AP-

2y) and Dlx3 bind one site each and are both required for maximal expression (Peng and

Payne, 2002). However, whether this is biologically relevant is not clear as Dlx3 and 3/?-

HSD were later shown to not be expressed in the same placental cell types (Berghorn et

al., 2005).

1.3.2.4 Hematopoietic cells

Different isoforms of Dlx4 show different effects on the P-globin promoter, a

gene expressed strongly in the placenta and kidney. One isoform, called BP-1 (P protein

ic repressed the fi-globin promoter through two silencer elements (Chase et al., 2002; Fu

et al., 2001), while Dlx7 showed no repression (Fu et al., 2001). This indicates that

different isoforms of Dlx genes may have different properties and functions.

Together, these studies consistently emphasize that Dlx proteins can control

differentiation by inducing expression of cell-specific proteins.

1.3.3 The function of Dlx proteins can be mediated by protein binding partners

The Dlx family regulates gene expression through their ability to bind the core

DNA sequence TAATT. Extended sequences have been identified for both Dlx3 and

Dlx5. Dlx3 binds the DNA consensus sequence: 5' (A/C/G)TAATT(G/A)(C/G) 3'

(Feledy et al., 1999) and Dlx5 binds a DNA element on the Bspl promoter, found to be:

5'TTAATTGA 3' (Benson et al, 2000). However, the low sequence specificity of

26 homeodomains compared with other DNA-binding motifs (most homeodomains will bind

the core sequence TAAT) is not sufficient to account for their specificity of action

(Hayashi and Scott, 1990). It has thus been proposed that increased specificity is

conferred by protein-protein interactions (Hayashi and Scott, 1990).

Several proteins have been found to interact with the Dlx family. The most

characterized interaction has been between the Dlx family and another closely related

family, Msx. The Msx (Ms7z-like) family contains homeobox genes related to the

Drosophila Msh (muscle segment homeobox) gene. In mammals there are three members:

Msx 1-3 (Davidson, 1995). All members of the family have been found to be repressors of transcription. They are expressed in many tissue types during development, including the dermis and epidermis of both fetal and adult skin (Stelnicki et al., 1997), the epithelial and mesenchymal cells that form the placenta (Quinn et al., 2000), migrating muscle precursor cells (Bendall et al., 1999), limb buds, branchial arches, and the dorsal neural tube (Zhang et al., 1997). It was originally found that DNA-binding is not required for the repressor activity of the Msx proteins (Catron et al., 1995) and was later discovered that

Msx can directly interact with the TATA-binding protein (TBP), a component of the basal transcription machinery, which may be a possible mechanism of direct transcriptional repression (Zhang et al., 1996). There are several functions attributed to the Msx family.

Both Msxl and Msx2 have been shown to inhibit differentiation of myoblasts, adipocytes, fibroblasts, osteoblasts and chondrocytes, making them general inhibitors of mesenchymal cell differentiation (Hu et al., 2001; Takahashi et al., 2001). One of the mechanisms by which this is accomplished is their ability to up-regulate cyclin Dl expression and increase Cdk4 activity, likely through an indirect interaction since they are

27 generally repressors (Hu et al., 2001). Therefore, Msx proteins generally maintain cell

proliferation and inhibit differentiation. It has become clear that the repressor function of

Msx proteins is likely due to various protein-protein interactions with the Msx

homeodomain that preclude DNA-binding. In this way, DNA binding by Pax3 (Bendall et

al., 1999), Lhx2 (Bendall et al., 1998) and Dlx proteins is inhibited by interaction with

Msxl. The Msx and Dlx families are expressed in many of the same tissues, including the

branchial arches and the limb buds (Zhang et al., 1997). The two families seem to be

antagonists of each other. Examples of known interactions include Msx2 and Dlx4 in the

regulation of epithelial-mesenchymal interactions in the placenta (Quinn et al., 2000) and

Msxl and Dlx3 interactions in murine epidermal tissues (Bryan and Morasso, 2000).

There are also several interactions during skeletal development, as discussed later. A

double knock-out of Msxl and Dlx5 was created, which demonstrated some independent

functions of each gene (Dlx5 in the mandible and Msxl in the middle ear), some

synergistic functions (deposition of bone tissue) and some overlapping functions (in palate growth and closure) (Levi et al., 2006).

The melanoma antigen (MAGE) family of proteins all contain a MAGE homology domain (MHD); MAGE-D1 (also known as Dlxin-1 and NRAGE) belongs to a subfamily which also contain a second homology domain named MHD2 which is separated from the first MHD by an interspersed repeat domain (IRD) consisting of 25 tandem repeats of the hexapeptide WQXPXX (where X represents any amino acid) (Barker and Salehi, 2002).

MAGE-D1 was recently implicated as a mediator of apoptosis by its binding to the p75 neurotrophin receptor (p75NTR). This binding inhibits the cell cycle and eventually facilitates p75NTR-mediated apoptosis in neuronal cells (Salehi et al., 2000). MAGE-D1

28 is broadly expressed during development in the brain, skeletal elements and osteoblasts

(Masuda et al., 2001). MAGE-D1 was originally found as a binding partner for Dlx5, although it was subsequently also shown to interact with Dlx4 and Msx2 (Masuda et al.,

2001). This interaction is mediated through the IRD of MAGE-D1 and the N-terminus of

Dlx5 and has been shown to increase the transactivation potential of Dlx5 (Masuda et al.,

2001). It also seems to act as an adaptor protein for the Dlx family. For example, MAGE-

Dl interacts with Prajal, a RING-finger protein which acts as a ubiquitin ligase leading to the degradation of MAGE-D1, which in turn down-regulates Dlx5-dependent transcription (Sasaki et al., 2002). Another protein, necdin, was also found to be associated with Dlx proteins through its interaction with MAGE-D1 (Kuwajima et al.,

2006). Necdin, a member of the MAGE family, is a post-mitotic neuron-specific nuclear protein that negatively regulates cell cycle progression. Necdin interacts with MAGE-D1 through their respective MHDs while MAGE-D1 interacts with Dlx through its IHD.

Necdin is co-expressed with Dlx proteins in the forebrain. There, it was found to enhance

Dlx2-mediated transcription of the Wntl promoter and promote GABAergic neuron differentiation (Kuwajima et al., 2006). It can also associate with Msx2 through MAGE-

Dl and de-repressed Msx2-induced repression of myogenic differentiation (Kuwajima et al., 2004). Thus, MAGE-D1, in association with necdin, seems to arrest the cell cycle, promoting differentiation.

A few members of the Dlx family have been implicated in Smad signaling, part of the TGF-p/BMP signaling pathway, by their ability to bind different Smads. The Dlxl homeodomain can interact with Smad4 in hematopoietic cells and repress the transactivation of Activin A (activin A-induced erythroid differentiation) (Chiba et al.,

29 2003). Dlx3 can also bind Smad6 through its homeodomain in differentiated placental

trophoblasts (Berghorn et al., 2006).

Runx2 (formerly Cbfal), a runt domain transcription factor, has been found to be

a direct inducer of the osteocalcin gene and to be necessary for osteoblast differentiation

(Ducy et al., 1997; Otto et al., 1997). It was found that Runx2 can bind Msx2, repressing

its transcriptional activity; Dlx5 then relieves this repression by binding Msx2 (Shirakabe

et al., 2001). Also, Dlx3 was found to be able to activate the osteocalcin promoter, but it

can also bind to Runx2 and reduce Runx2-mediated transcription (Hassan et al., 2004).

Another protein known to interact with the Dlx family of proteins is GRIP lb

(glutamate receptor interacting protein lb) although the significance of this interaction is

unclear (Yu et al. 2001). Griplb, a smaller form of Grip, shows a restricted pattern of

expression that overlaps with Dlx2 and Dlx5, where they have been shown to bind to its

fourth PDZ (post-synaptic density protein) domain through their N-termini (Yu et al.

2001).

In addition to proteins, Evf-2, a non-coding RNA, encoded by the Dlx5/6 ultra-

conserved intergenic enhancer region, can form a complex with Dlx2. This complex then

specifically activates the Dlx5/6 forebrain enhancer (Feng et al., 2006).

2. Endochondral Ossification

2.1 Physiological aspects of skeletal development

The vertebrate embryo makes bones by two different mechanisms:

intramembranous ossification, which occurs in the flat bones of the skull and the clavicle,

and endochondral ossification, the process used in all other bones. Endochondral

N ossification makes use of a cartilage intermediate, whereas intramembranous ossification

30 involves the differentiation of mesenchymal cells directly into osteoblasts. Endochondral ossification begins with the condensation of mesenchymal cells and their differentiation into proliferating chondroblasts. These cells then differentiate to form pre-hypertrophic chondrocytes; after further differentiation, these cells undergo hypertrophy, and die.

Vascularization of this cartilage tissue brings in osteoblasts, which begin laying a matrix of trabecular bone, replacing the original cartilage matrix. A diagram of the different steps of chondrocyte differentiation is shown in Figure 2.

The mesenchymal cells which will eventually form bone come from three locations: neural crest cells of the neural ectoderm will form the craniofacial bones, the sclerotome of the paraxial mesoderm (somites) will form the axial skeleton and the somatopleure of the lateral plate mesoderm will eventually become the appendicular skeleton (Goldring et al., 2006). These cells are pluripotent and give rise to many different cell types: osteoblasts, chondrocytes, adipocytes, myoblasts and fibroblasts

(Komori, 2006). Once they have migrated to their eventual locations, mesenchymal cells destined to become chondrocytes begin condensing. This condensation occurs by the increased expression of cell adhesion molecules such as N-cadherin and NCAM (Goldring et al., 2006). The shape of these cells is dictated by interactions of the mesenchyme with the overlying epithelium.

Proliferating chondroblasts have a highly developed ECM consisting of Type II collagen, Type IX collagen and aggrecan. This ECM is essential to the growth of the cells; a defective ECM has been shown to affect chondrocyte differentiation and alter the expression of differentiation markers (Barbieri et al., 2003). Around the chondroprogenitor cells is a thin layer of mesenchymal cells called the perichondrium,

31 Figure 2: Regulation of chondrocyte differentiation. The cell types at stages of endochondral ossification are shown. Factors important for the transition between different cell types are indicated in the middle. The expression of several marker genes at different stages are indicated at the bottom of the figure.

32 Hypertrophic Proliferating Pre-hypertrophic chondrocyte Mesenchymal Chondroprogenitor chondrocyte precursor chondrocyte BONE

Sox9 Sox9 Sox5 Ihh Sox6 Runx2 Dlx5 Dlx6

Sox9 Sox9 Type II Ihh Osc Runx2 D!x3 collagen Runx2 Osp Dlx5 Sox9 Type II TypeX collagen collagen Dlx3 Dlx3 Dlx5 Dlx5 which supplies many signaling molecules to the dividing chondrocytes. Differentiation of

chondrocytes begins at the centre of the bone (diaphysis), with reserve chondrocytes at

the ends (epiphyses) (Lai and Mitchell, 2005). Differentiation begins when these cells

become pre-hypertrophic chondrocytes (Yoon et al., 2006). After further differentiation, the chondrocytes begin to hypertrophy, increasing their cellular fluid volume by as much

as 20-fold (Goldring et al., 2006). They begin producing a different ECM conducive to mineralization and blood vessel invasion consisting of Type X collagen and alkaline phosphatase (Yoon et al., 2006). They also express both osteopontin and osteocalcin

(Barak-Shalom et al., 1995; Sims et al., 1997). This ECM eventually becomes mineralized (Shum and Nuckolls, 2002). At the same time, the perichondrium around the hypertrophic chondrocytes differentiates directly into osteoblasts, becoming first the periosteum, then the fully mineralized bony collar. Angiogenesis begins, and the chondrocytes undergo apoptosis (Goldring et al., 2006). Vascularization brings in osteoblasts derived from mesenchymal cells that begin removing the cartilage matrix and replacing it with a bone matrix, consisting of Type I collagen, osteocalcin and osteopontin

(Komori, 2006; Wagner and Karsenty, 2001).

A pool of proliferating chondrocytes remains at each end of the bone and longitudinal bone growth is driven by proliferation and enlargement of these cells (Yoon et al., 2006). These reserve cells eventually disappear at the end of puberty, when bone growth has finished (Karsenty and Wagner, 2002).

The contents of the ECM at each stage is essential for proper progression of ossification. The collagens are the most abundant members of the ECM and the major structural element of all connective tissue. Vertebrates have around 27 different collagens

34 and another 20 different proteins with "collagen-like" domains (Myllyharju and

Kivirikko, 2004). Collagens form a typical, right-handed triple helix, with three polypeptide chains, either homotrimers, with three identical chains, or heterotrimers, containing different chain combinations (Gelse et al., 2003). The collagen domain consists of repeats of Gly-X-Y, with glycine every third position to allow for tight packing of collagen a-helices (glycine is the smallest amino acid) and where X and Y are frequently occupied by proline and 4-hydroxyproline. There are several groups of collagens. The fibril-forming collagens are the most common and consist of Type I and

Type II collagen (expressed from the Collal and Col2al genes, respectively), among others. They assemble into highly oriented, supramolecular aggregates. The fibril-forming group are synthesized as procollagens and have to have an N- and C-terminal domain cleaved to form into their fibrils (Kadler et al., 2007). They also undergo extensive post- translational modifications, including adding hydroxyproline residues and glucosyl and galactosyl groups, which mediate interactions with proteoglycans (Gelse et al., 2003).

Type X collagen (expressed from the CollOal gene) is one of the hexagonal network- forming collagens with special N- and C-terminal domains which allow its assembly into networks.

Both osteocalcin and osteopontin are secreted by osteoblasts into bone ECM.

Osteopontin is a phosphorylated glycoprotein that has a tripeptide sequence RGD (Arg-

Gly-Asp) recognized by a receptor on osteoclasts. It is believed that osteopontin binds osteoclasts and facilitates local bone resorption (Kavukcuoglu et al., 2007). Osteocalcin is a small Ca -binding protein containing three Gla (y-carboxyglutamic acid) residues,

35 which facilitate the protein's adsorption to hydroxyapatite (the main mineral in bone). It also participates in regulation of mineralization and bone turnover (Dowd et al., 2003).

2.2 Growth factors regulate endochondral ossification

2.2.1 Bone Morphogenetic Proteins

One of the most widely studied and important growth factors during endochondral ossification is the BMP family. BMPs are members of the TGF-p superfamily and they signal through serine/threonine kinase transmembrane receptors. Their signal is then transduced within the cell by Smad proteins (Ryoo et al., 2006). Most of the BMPs expressed during chondrogenesis are found within the perichondrium (BMP2, BMP4 and

BMP7), however BMP7 has been shown to be expressed in proliferating chondrocytes

(Goldring et al., 2006) and BMP6 was found in both pre-hypertrophic and hypertrophic chondrocytes (Lai and Mitchell, 2005). BMPs have numerous roles throughout endochondral ossification. BMP expression was found to be upstream of Sox9, which likely makes it one of the first factors involved in chondrogenesis (Healy et al., 1999).

BMPs were also shown to up-regulate Indian hedgehog (Ihh) expression (Lai and

Mitchell, 2005; Yoon et al., 2006) and BMP2, BMP4 and BMP7 were all shown to increase Runx2 expression (Shum and Nuckolls, 2002). However, BMPs can also have an inhibitory effect on chondrogenesis. BMP4 was shown to induce expression of Msx2, which is a negative regulator of Sox9 and chondrogenesis (Shum and Nuckolls, 2002).

Overall, BMP signaling positively regulates both chondrogenesis and osteogenesis

(Wagner and Karsenty, 2001).

36 2.2.2 Indian Hedgehog/Parathyroid Hormone-related Protein

Ihh belongs to the hedgehog family which encode secreted growth and patterning

molecules (Lai and Mitchell, 2005). Ihh protein is expressed by pre-hypertrophic

chondrocytes and signals to both chondrocytes and the perichondrium (Colnot, 2005;

Goldring et al., 2006). Ihh functions in a feedback loop with another signaling molecule,

parathyroid hormone-related peptide (PTHrP). PTHrP is found in the periarticular

perichondrium (at the epiphyses). Ihh up-regulates PTHrP, which stimulates cell

proliferation and inhibits chondrocyte hypertrophy thus regulating the size of the pre-

hypertrophic zone (Goldring et al, 2006; Lai and Mitchell, 2005; Shum and Nuckolls,

2002; Wagner and Karsenty, 2001). Ihh also has roles that are independent of PTHrP. It

was shown to induce proliferation of chondrocytes and it is also an important regulator of perichondrial development (Lai and Mitchell, 2005), perhaps because of its ability to

induce Runx2 in the perichondrium (Komori, 2006).

2.2.3 Other growth factors

FGFs regulate limb initiation and limb bud outgrowth (Goldring et al., 2006).

They have also been shown to negatively regulate proliferation and differentiation of chondrocytes, in different cell types and at different stages of differentiation, showing opposing actions to BMP in the growth plate (Yoon et al., 2006). In fact, BMP signaling inhibits FGF receptor 1 (FGFR1) expression (Yoon et al, 2006). Insulin growth factor 1

(IGF1) has been shown to stimulate proliferation and differentiation of chondrocytes

(Phornphutkul et al., 2006). Vascular endothelial growth factor (VEGF) is produced in cartilage and the perichondrium and is essential for angiogenesis and invasion of osteoblasts to make the bone matrix (Colnot, 2005; Goldring et al., 2006).

37 2.3 Transcription factors regulate cell identity and differentiation during endochondral ossification

Osteochondroprogenitor mesenchymal cells express both Sox9 and Runx2

(Akiyama et al., 2005; Ng et al., 1997; Zhou et al., 2006). Whichever transcription factor is expressed at the highest level determines whether the cells will become chondrocytes or osteoblasts (Kawakami et al., 2006). Sox9 also seems to be dominant over Runx2 since it can interact with Runx2 through their respective DNA-binding domains and thus repress Runx2 transcriptional activity (Zhou et al., 2006).

2.3.1 Sox genes

Sox9 (SRY-type HMG box 9) belongs to the Sox gene family, related to the testis determination factor Sry (Wright et al., 1993). The DNA-binding domain of Sox9 is an

Sry-like high mobility group (HMG) box (Healy et al., 1999). Sox9 mutants are not viable, so conditional and heterozygous mutants are usually studied. Inactivation of Sox9 in the limb buds before mesenchymal condensation leads to a complete absence of cartilage and bone (Akiyama et al., 2002). Inactivation after mesenchymal condensation leads to severe chondrodysplasia, with chondrocyte proliferation inhibited (Akiyama et al., 2002). Sox9+/~ mice die perinatally with a cleft palate and bending of skeletal structures derived from cartilage (Bi et al., 2001). Pre-cartilaginous condensations are delayed and there is a larger hypertrophic zone (Bi et al., 2001). These heterozygous mutant mice are phenocopies of the human disease campomelic dysplasia (CD).

Symptoms of this disorder include congenital bowing and angulation of the long bones, defects in cartilage formation and XY sex reversal and affected individuals usually die within a few months due to respiratory failure (Foster et al., 1994). This disease was

38 mapped to the SOX9 locus and was found to be an autosomal dominant disease caused by

a mutation in one of the SOX9 alleles (Foster et al., 1994; Giordano et al., 2001; Wagner

et al., 1994). This information indicates that Sox9 is essential for cartilage and bone

formation by positively regulating chondrocyte proliferation and negatively regulating hypertrophy. Sox9 regulates several early chondrogenic differentiation markers. The chondrocyte-specific enhancer in intron 1 of CoUal, has three HMG-binding domains, one of which is bound by Sox9, resulting in increased transcription (Goldring et al., 2006;

Ng et al., 1997; Shum and Nuckolls, 2002; Zhou et al., 1998). Sox9 can also bind the

Colllal gene enhancer and increase transcription (Shum and Nuckolls, 2002).

Sox9 is necessary for expression of two other Sox genes, L-Sox5 and Sox6; neither

L-Sox5 nor Sox6 expression is detected in Sox9 null mutants (Akiyama et al., 2002). Since

L-Sox5 and Sox6 have no transactivation domains, it is believed that they increase the transcriptional activity of Sox9 by forming a complex of all three proteins (Karsenty and

Wagner, 2002). L-Sox5 and Sox6 are also essential for bone formation. A single null mutant of either gene results in mild skeletal abnormalities, however, a double knock-out,

SoxS/6'^, causes severe chondrodysplasia, similar to Sox9+/~ mutants (Smits et al., 2001).

2.3.2 Runx2

Runx2 (formerly Cbfal and PEBP2), one of three mammalian Runx genes, is related to the Drosophila pair-rule gene runt (Ogawa et al., 1993). Runx proteins recognize the DNA consensus sequence PyGPyGGTPy (Komori, 2006) and Runx2 has a strong Glu and Ala-rich transactivation domain (Akiyama et al., 2005). Runx2 null mutants die just after birth due to respiratory failure. These mice show a complete lack of ossification due to a lack of osteoblast differentiation (Komori et al., 1997; Otto et al.,

39 1997). In a culture of i?«wx2-deficient calvarial cells, an osteoblast phenotype was not seen, however, adipocytes and chondrocyte differentiation of the mesenchymal cells did occur (Kobayashi et al., 2000). This indicates that Runx2 is necessary for osteogenesis and inhibits adipocyte and chondrocyte differentiation of mesenchymal cells. Runx2 likely induces osteoblast differentiation by its regulation of osteoblast-specific genes like

Collal, Osteopontin, Bsp and Osteocalcin (Komori, 2006). It may also up-regulate the expression of VEGF, thus promoting angiogenesis (Shum and Nuckolls, 2002).

During chondrogenesis, Runx2 is also expressed in the perichondrium and pre- hypertrophic chondrocytes (Goldring et al., 2006; Shum and Nuckolls, 2002). Mutations in Runx2 lead to a lack of chondrocyte hypertrophy (Kim et al., 1999). It also induces the expression of Ihh (Komori, 2006; Lai and Mitchell, 2005). This indicates a role for Runx2 in endochondral ossification in both chondrogenesis and osteogenesis.

2.3.3 Osterix

Another transcription factor essential for osteoblast differentiation is the zinc finger, SP family member Osterix (Osx) (Komori, 2006). Null mutations of Osx result in a lack of bone formation due to the absence of mature osteoblasts (Nakashima et al.,

2002). This gene is activated downstream of Runx2 and can activate the osteocalcin and

Collal genes (Akiyama et al., 2005).

2.3.4 Dlx proteins

Dlx proteins are expressed during bone formation in both chondrocytes and osteoblasts. Both Dlx5 and Dlx6 were shown to increase chondrogenic nodule formation, downstream of mesenchymal cell aggregation with the homeodomain being essential for

40 this effect (Hsu et al., 2006). In the limbs, misexpression of Dlx5 leads to shortened

skeletal elements, likely due to reduced chondrocyte proliferation and enhanced

chondrocyte hypertrophy (Bendall et al., 2003; Ferrari and Kosher, 2002). Dlx5 induces

chondrocyte differentiation leading to a greater number of hypertrophic chondrocytes and

inducing Type X collagen, osteopontin and an expanded mineralized cartilage matrix

(Ferrari and Kosher, 2002). This indicates that Dlx5, and possibly Dlx6, are inducers of both early and late chondrocyte differentiation (Bendall et al., 2003).

In osteoblasts, overexpression of Dlx5 induced osteogenic markers, such as

Collal, osteopontin, alkaline phosphatase, osteocalcin and Runx2 and accelerated mineralization of the extra-cellular matrix (ECM) (Holleville et al., 2007; Miyama et al.,

1999; Tadic et al., 2002). It also enhanced the formation of periosteal bone (Ferrari and

Kosher, 2002). Overexpression ofDlx3 has shown a similar effect, by inducing expression of Runx2, osteocalcin and alkaline phosphatase (Hassan et al., 2006). Several osteoblast-specific markers are directly up-regulated by Dlx proteins. Transcription of

Runx2 is partially mediated by Dlx proteins in osteoblasts. There are three Dlx-responsive elements in the promoter region ofRunx2. Both Dlx3 and Dlx5 have been shown to enhance expression of Runx2 (Hassan et al., 2006; Lee et al., 2005). Runx2 was also repressed by Msx2 (Hassan et al., 2006; Lee et al., 2005). Msx2 was able to inhibit the transcriptional activity of Runx2 by binding to its DNA-binding domain; Dlx5 relieved this repression by binding to Msx2 (Shirakabe et al., 2001). Conflicting data has been presented on the role of Dlx3 and Dlx5 during osteocalcin expression. Dlx5 was originally shown to repress the osteocalcin promoter, with anti-sense inhibition of Dlx5 causing increased osteocalcin transcription (Ryoo et al., 1997). However, Dlx5 was also

41 shown to mildly activate the basal promoter, and reverse Msx2 repression, an effect

independent of the DNA-binding activity of Dlx5 (Newberry et al., 1998). Also, the

osteocalcin gene was shown to be repressed by Msx2 in proliferating osteoblasts with

Dlx3, Dlx5 and Runx2 being recruited post-proliferatively to initiate transcription

(Hassan et al., 2004). Dlx3 alone was shown to stimulate transcription of osteocalcin, however a Dlx3/Runx2 interaction reduced Runx2-mediated transcription (Hassan et al.,

2004). Control of osteocalcin expression therefore seems to require a complex interaction between several transcription factors, including Msx2, Dlx3, Dlx5 and Runx2. Lastly, the bone sialoprotein (Bsp) promoter contains two Runx2-binding sites and one homeodomain-binding site, with all three sites essential for maximal activity of Bsp in osteoblasts (Roca et al., 2005). Dlx5 has been shown to bind the Bsp promoter either independently or in a complex with Runx2 to stimulate activity (Benson et al., 2000;

Roca et al., 2005).

The role of Dlx5 and Dlx6 in endochondral ossification requires further study.

The main focus of this thesis was to characterize the response of a chondrogenic cell model in vitro to misexpression of either Dlx5 or Dlx6. The longer term goal of this study was to establish an in vitro system that would complement ongoing in vivo studies that seek to characterize the mechanism of action of Dlx transcription factors during chondrocyte differentiation.

42 Chapter 2: The role of Dlx5 and Dlx6 in chondrogenesis in

ATDC5 cells

1. Introduction

1.1 Objectives

Dlx5 and Dlx6 are both expressed in mesenchymal chondroprogenitors, pre- hypertrophic and hypertrophic chondrocytes (Bendall et al., 2003; Ferrari and Kosher,

2002; Ferrari et al., 1995; Ghoul-Mazgar et al., 2005; Hsu et al., 2006). They also have been shown to induce both early and late chondrogenic differentiation (Bendall et al.,

2003; Chin et al., 2007; Ferrari and Kosher, 2002), but how those effects are mediated is not known. No direct downstream gene targets of Dlx5 and Dlx6 in chondrocytes have been identified to date. The domain requirements for Dlx5 and Dlx6-mediated chondrogenesis is somewhat different (Hsu et al., 2006) and this is reflected in the transcriptional activities of their amino- and carboxy-terminal domain (Appendix 2).

Since their transactivation domains are quite divergent, this may explain their functional specificity in regions of co-expression.

To provide further insight into the role of Dlx5 and Dlx6 in chondrogenesis, an in vitro cell line that differentiates in a similar pattern to in vivo chondrocytes was used.

Dlx5 and Dlx6 were overexpressed in ATDC5 cells and several genes analyzed by quantitative PCR to determine the effect of Dlx5 or Dlx6 on chondrogenic differentiation.

Cell morphology and chondrogenic nodule formation was also monitored

43 1.2 A chondroprogenitor cell line

The ATDC5 cell line was derived from a murine teratocarcinoma. These multipotent fibroblast cells were found to undergo differentiation in vitro in a similar pattern to chondrocytes in vivo, making them an excellent model of chondrogenesis

(Atsumi et al., 1990). At sub-confluence, these cells exhibit a fibroblastic morphology and they stop proliferating at confluence, however, with the addition of 10 u.g/mL of insulin they begin proliferating again 2-3 days later, aggregate and form chondrogenic nodules that stain with Alcian blue (Atsumi et al., 1990).

Several factors are able to induce chondrogenic differentiation in ATDC5 cells.

The most commonly used is insulin. High levels of insulin (10 u.g/mL) were originally believed to exert an effect on the IGF1 receptor (IGF1R) rather than the insulin receptor

(IR), since they are known to bind each others receptors, albeit with lower affinity

(Phornphutkul et al., 2006) and IGF1 has a known role in promoting chondrogenic differentiation in vivo (Goldring et al., 2006). However, it has been shown that insulin is able to induce chondrogenesis through the IR, as well as the IGF1R, implicating insulin as having a role in chondrogenesis (Phornphutkul et al., 2006).

The BMP proteins have also been shown to have a positive effect on differentiation of ATDC5 cells. Both BMP2 and BMP4 can induce chondrogenic differentiation without the requirement for a condensation phase (Shukunami et al., 1998;

Wahl et al., 2004). BMP proteins up-regulate Type II collagen and, later, Type Xcollagen, alkaline phosphatase and osteopontin (Shukunami et al., 1998). Cellular hypertrophy and mineralization can be enhanced by adding ascorbic acid to the media and growing the cells at 3% CO2 (Akiyama et al., 2000). Adding ascorbate 2-phosphate, along with

44 insulin, was found to significantly reduce the pre-chondrogenic proliferation phase from

21 to 7 days, with greater production of larger chondrogenic nodules and induction of hypertrophy around 7-10 days (Altaf et al., 2006).

Conversely, some agents have been identified which restrict differentiation of

ATDC5 cells. FGF2 is a potent mitogen for these cells and its addition blocks differentiation (Shukunami et al., 1998). Dexamethasone inhibits insulin-induced condensation and nodule formation and also down-regulates important chondrogenic markers such as Type II collagen and Runx2 (Fujita et al., 2004). Hypoxia (1% O2 - the in vivo physiological condition during cartilage formation) was also found to be sufficient to induce chondrogenesis. However, when insulin treatment and hypoxia were combined, it delayed and suppressed insulin-mediated early chondrogenesis and nearly completely blocked hypertrophic differentiation (Chen et al., 2006).

1.3 Chondrogenesis in the ATDC5 cell line

While chondrogenic differentiation of ATDC5 cells follows a predictable pattern of marker gene induction, the exact time course is seen to vary from study to study.

Before differentiation, ATDC5 cells behave as undetermined mesenchymal cells expressing Collal, but not Col2al (Shukunami et al., 1996). Upon addition of insulin they enter the proliferating chondroprogenitor phase and exit the cell cycle by day 21 -24 with the hallmarks of hypertrophic chondrocytes. (Shukunami et al., 1997; Shukunami et al., 1996). They begin forming chondrogenic nodules around day 8-10 (Shukunami et al.,

1996). By day 33 the cultures start depositing a calcified matrix (Shukunami et al., 1997).

ATDC5 cell differentiation is accompanied by the same suite of gene expression as is seen in vivo: Type II collagen is up-regulated as early as day 8 following the addition of

45 insulin to confluent cultures and persists until day 30 (Akiyama et al., 1997; Shukunami et al., 1997). PTH/PTHrP is expressed from day 10-30 (Shukunami et al., 1997), Ihh is present from day 12-45 (Akiyama et al., 1997) and Type Xcollagen is expressed later, from day 24-45 (Akiyama et al., 1997; Shukunami et al., 1997). Several BMP family members have also been found to be expressed during ATDC5 differentiation. BMP4 was produced during the entire process, BMP6 was found during cartilage nodule formation and BMP7 was only expressed after the cells had begun to calcify (Akiyama et al., 2000).

2. Materials and Methods

2.1 Common laboratory reagents

Unless otherwise stated, common laboratory chemicals were purchased from

Fisher Scientific (Whitby, Ontario). Modifying and restriction enzymes utilized were purchased from Invitrogen (Burlington, Ontario) and oligonucleotides were purchased from IDT (Toronto, Ontario). Cell culture reagents were purchased from Invitrogen, unless noted otherwise.

2.2 Plasmids

Mammalian retroviral plasmids were based onpBMN-LZRS (Yang et al., 1999).

LZRS-myc-Dlx5 and LZRS-myc-Dlx6 were constructed previously (C. Hsu, A. Bendall, unpublished). 2x myc-tagged Dlx coding sequences were subcloned as EcoRI-Hindlll fragments into LZRSA (Hu et al., 2001). A non-insert control LZRS plasmid was constructed by removing the Dlx5 coding sequence from LZRS-myc-Dlx5 using HindlH and EcoRI, end filling the 5' overhang with Klenow (New England Biolabs) and recircularizing the plasmid with T4 DNA ligase.

46 2.3 Cell culture

Phoenix Ecotropic cells (American Type Culture Collection, ATCC) were maintained in DMEM supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, 100 ug/ml streptomycin, and 2 mM L-glutamine in a humidified incubator at

6 37°C, 5%C02. lxlO cells were seeded into a 6 cm plate 24 hours prior to transfection.

Cells were transfected at 50-70% confluence with 6 ug of LZRS retroviral expression plasmids, using 2.5 ul of Lipofectamine 2000 (Invitrogen) per |ug of DNA, according to the manufacturer's protocol. 48 hours post-transfection, 4 ug/ml puromycin (Sigma) was added to the media to select for transfected cells. Cells were grown in puromycin- containing medium for 24 hours, then split into 15 cm plates. Virus was harvested 24 hours after cells became confluent, as follows. Medium was removed from the transfected

Phoenix plates, then passed through a 0.45 uM filter to remove cell debris. 4 ug/ml of polybrene (Sigma) was added to the virus-containing medium, which was then added directly to target cells as indicated below.

ATDC5 cells (ATCC) were maintained in DMEM/Ham's F12 (1:1) with 5% FBS,

100 U/ml penicillin, 100 ug/ml streptomycin, 2 mM L-glutamine, 3 x 10"8 M sodium selenite (Sigma) and 10 |J.g/ml apo-transferrin (Sigma) in a humidified incubator at 37°C,

5% CO2. For PvNA extraction, 4.5 x 104 cells were seeded into 12-well plates. For staining and microscopy, 1 x 105 cells were seeded into 3.5 cm dishes. Cells were grown to 90-95% confluence for infection. For infection, the medium on the cells was replaced with a 1:1 mixture of complete medium and virus-containing medium collected from

Phoenix Ecotropic cells; cells were infected on day 0, with a second infection of a 1:1 mixture 24 hours later.

47 Phoenix Ecotropic cells; cells were infected on day 0, with a second infection of a 1:1 mixture 24 hours later.

2.4 RNA extraction, cDNA synthesis and RT-PCR

Duplicate infected ATDC5 cell cultures were harvested at the following intervals: day 0,1, 2, 4, 6, 8, 10, 12, 14, and 17. Total RNA was extracted using 500 uL TRIzol

(Invitrogen) per well, according to the manufacturer's protocol and quantitated by measuring the optical density at 260nm. 1 ug of RNA was reverse transcribed into cDNA using oligo dTis (IDT) and Superscript II (Invitrogen) according to the manufacturer's protocol. Semi-quantitative reverse transcription polymerase chain reaction (RT-PCR) using cDNA from day 0 was performed to test real-time PCR primers (Table 2). The reaction consisted of 50 ng cDNA, 20 pmol each primer and 10 U Taq polymerase (UBI).

Amplification was done in an Eppendorf mastercycler as follows: an initial denaturation at 94°C for 5 minutes, 35 cycles of (94 °C for 60 seconds, 56°C for 30 seconds and 72 °C for 90 seconds), with a final extension at 72°C for 10 minutes. PCR products were run on a 1% agarose gel in TAE containing 0.01% ethidium bromide and visualized on a

Typhoon 9400 Variable Mode Imager (Amersham Biosciences) with ImageQuant 5.2 software (Molecular Dynamics)

2.5 Real-Time PCR

The primers for all assayed genes were designed using Primer Express software

(Applied Biosystems). The genes, primer sequences, and amplicon length are listed in

Table 2. For a single real-time PCR reaction in 25 ul, 50 ng of cDNA and 400 nM each primer, with lx SYBR-Green Mastermix (SYBR Green I dye, AmpliTaq Gold® DNA

48 Table 2: Primers used for Real-Time PCR.

Gene Forward primer Reverse primer Amplicon name length p-Actin 5 '-CCTGAACCCTAAGGCCAACC-3' 5 '-CACAGCCTGGATGGCTACG-3' 91 bp Col2al 5'CCAGGGCTCCAATGATGTAGA3' 5 'TGTTTCGTGCAGCCATCCT3' 84 bp CollOal 5 '-AGCTGCGCCACGCATCT-3' 5'- 81 bp TACAAATGAACCTGTCTGGGTACCT- 3' Dlx3 5 '-GCATCTGCACAACGCGG-3' 5 '-GTGTTCCAGCGGCACCTC-3' 81 bp Dlx5 5'- 5 '-GGCATCTCCCCGTTTTTCAT-3' 82 bp GACTGACGCAAACACAGGTGA-3' Dlx6 5 '-TCTGCCAAGGGCGTCAGTAT-3' 5 '-GGTGTCCTGGTGTGGTGAGG-3' 81 bp Ihh 5'-GGACGAGGAGAACACGGGT-3' 5'-TCATGACAGAGATGGCCAGTG-3' 83 bp Osc 5'-GCCTTCATGTCCAAGCAGGA-3' 5'-GCTAAGGGCTCTGGCCACT-3' 81 bp Osp 5'-CAAGCAATTCCAATGAAAGCC- 5'-CCTCGCTCTCTGCATGGTCT-3' 84 bp 3' Runx2 5' - ACGCCCTGTCTTCC AC A-3' 5'- 81 bp GGCAGTGTCATCATCTGAAATACG- 3' Sox9 5'-GCATCTGCACAACGCGG-3' 5' -CCTCC ACG A AGGGTCTCTTCT-3' 86 bp polymerase, dNTPs with dUTP, ROX, and optimized buffer components) (Applied

Biosystems) was used. A three temperature cycling profile (95°C for 10 sec, 60°C for 15 sec and 72°C for 20 sec) was run in a Rotor-Gene 6000 Series machine (Corbett). The

specificity of each PCR reaction was assessed by performing a melting curve analysis after each reaction (Figure 3). To normalize for cDNA between samples, fi-actin was used as an internal control. Data was analyzed using Rotor-Gene 6000 Series software. To analyze relative induction over a basal level (day 0) the delta-delta CT relative quantification method was used (Livak and Schmittgen, 2001). Samples at each timepoint were performed in duplicate and averaged. The experiments were done three times.

2.6 Cell staining and microscopy

ATDC5 cells in 3.5 cm dishes were rinsed in 1 x PBS, then fixed with 100% methanol for 5 minutes. They were then stained in Alcian blue (0.01% Alcian blue, 20% acetic acid in ethanol) overnight. Chondrogenic nodules were examined using a Leica MZ

125 microscope at lOx magnification. Cells were examined using a Leica DM IRE2 microscope at 20x magnification. Images were processed in Openlab version 4.0.1

(Impro vision).

2.7 Western blotting

Whole cell extracts of infected ATDC5 cells (day 4) were made using 2x sodium dodecyl sulphate (SDS) sample buffer (0.1 M Tris pH 6.8, 20% glycerol, 4% SDS, 200 mM DTT, 0.001% bromophenol blue). Proteins were separated on a 13.5% SDS- polyacrylamide gel and transferred to Westran clear signal nitrocellulose membrane

(Mandel Scientific Company). The membrane was probed using 1:1000 anti-c-myc

50 Figure 3: Melting curve analysis for assayed genes: Actin (A), Col2al (B), CollOal (C), Osc (D), Osp (E), Sox9 (F), Runx2 (G), Ihh (H), D/xJ (I), Dlx5 (J) and £>/xtf (K). On the x-axis is the temperature in °C from 70°C to 95°C and the y-axis indicates the negative derivative of fluorescence over temperature (-dF/dT).

51 V "'"*%*X

*•***. If -V_^ 90 9S 75 B 85 90 75 90 65 90

52 antibody (9E10) and 1:5000 sheep anti-mouse secondary antibody, conjugated to

horseradish peroxidase (BioCan Scientific). Detection was accomplished using ECL-Plus

Western Blotting Detection System (GE Healthcare), according to the manufacturer's

protocol. Bands were visualized with a Typhoon 9400 Variable Mode Imager (Amersham

Biosciences) and ImageQuant Version 5.2 software (Molecular Dynamics).

3. Results

3.1 Selection of the ATDC5 cell line as a chondrogenic model system

In order to determine the effect of Dlx5 and Dlx6 on chondrogenesis, we used a cell line that differentiates in a multi-step process similar to chondrocytes in vivo. ATDC5 cells can be induced to differentiate by the addition of insulin (Atsumi et al., 1990).

However, we note that ATDC5 cells spontaneously differentiate upon reaching confluence, without adding insulin, albeit more slowly. Therefore, misexpressing Dlx5 or

Dlx6 in ATDC5 cells in the absence of insulin, allows the investigation of Dlx5 and Dlx6-

specific effects.

In order to determine the effects of Dlx5 and Dlx6 on the differentiation of

ATDC5 cells, real-time PCR using gene-specific primers was used. Primers were designed for several genes of interest, spanning introns so that cDNA-specific fragments were amplified. Sox9 was chosen as a marker of early chondrogenesis, since it is expressed in mesenchymal condensations and proliferating chondrocytes. Col2al is a marker of proliferating chondrocytes and pre-hypertrophic chondrocytes, whereas

CollOal is expressed in hypertrophic chondrocytes. Ihh is expressed in pre-hypertrophic chondrocytes and Runx2 is expressed in both pre-hypertrophic and hypertrophic chondrocytes. Both osteopontin and osteocalcin are expressed late in chondrocyte

53 differentiation, as well as in osteoblasts. All of these markers can be used to determine the stage of differentiation of ATDC5 cells. Actin was chosen as an internal control, since it was found that its levels did not change significantly during chondrogenesis in ATDC5 cells (Figure 4).

Expression of these markers was examined using semi-quantitative RT-PCR of

ATDC5 RNA at confluence, but before differentiation. All of the genes of interest were expressed in these cells, including several Dlx genes (Figure 5). This validates the specificity of the primers and indicates that all of these genes are expressed in this cell line at low but detectable levels.

3.2 Overexpression of Dlx5 and Dlx6

The level of expression of both Dlx5 and Dlx6 were monitored using real-time

PCR for each infection and misexpression was also confirmed using Western blotting.

Figure 6A and 6B show the average fold induction of Dlx5 and Dlx6 expression from three independent experiments. The level of induction is relatively consistent, achieving a

100-200 fold increase above the endogenous levels measured at day 0. Figure 6C shows a

Western blot of LZRS, LZRS-myc-Dlx5 and LZRS-myc-Dlx6-infected cells. There is no expression of myc-tagged protein in the LZRS control lane (lane 1), but both Dlx5 and

Dlx6 protein are visible in lanes 2 and 3 respectively. Actin was used as a loading control.

3.3 Dlx5 and Dlx6 overexpression resulted in decreased cell number and suppressed chondrogenic nodule formation

Microscopy and Alcian blue staining were used to compare cell morphology and chondrogenic nodule formation following infection of ATDC5 cells with Dlx5 and Dlx6-

54 Figure 4: Changes in actin expression levels over the 17 day differentiation period relative to day 0. ATDC5 cells were harvested at various time points and RNA was extracted using TRIzol. 1 ug of RNA was reverse transcribed into cDNA then used for real-time PCR analysis using primers specific to actin. Results shown are representative data from one experiment using duplicate samples.

55 m i. ___ J i S — L_ n e in actin expressio Chang t -» -»

Days post infection Figure 5: Expression of chondrogenic genes in undifferentiated ATDC5 cells. Confluent ATDC5 cells were harvested and RNA was extracted using TRIzol. 1 ug of RNA was reverse transcribed into cDNA using Superscript II. PCR was performed using real-time primers for several genes and Taq polymerase. PCR products were run on a 1% agarose gel. Amplicon length is between 80 and 90 bp. * indicates a non-specific product.

57 i'

;7 «-

i 1 1 0<*

c 0"b 3 •f- ^ Co' ..v* «. x Co,\a*1

<\<\ 1 ^

Q. Q_

O£ °O

58 Figure 6: Overexpression of Dlx5 and Dlx6 in ATDC5 cells. (A) Real-time PCR of Dlx5 overexpression. ATDC5 cells were infected with control retrovirus or encoding Dlx5. Cells were harvested at the times indicated and RNA was extracted using TRIzol. 1 ug of RNA was reverse transcribed into cDNA, then used for real-time analysis using primers for Dlx5 (B) Real-time PCR of Dlx6 overexpression. ATDC5 cells were infected with control or retrovirus encoding Dlx6. Cells were harvested at the times indicated and RNA was extracted using TRIzol. 1 ug of RNA was reverse transcribed into cDNA, then used for real-time analysis using primers for Dlx6 (C) Western blot of LZRS, LZRS-Dlx5 and LZRS-Dlx6-inkcted cells. Cells were harvested 4 days post-infection then resuspended in 2x sample buffer. The whole cell lysate was run on a 13.5% polyacrylamide gel with a pre-stained protein ladder. Anti-myc primary antibody was used to detect myc-tagged Dlx proteins. The membrane was re-probed with a (3-actin antibody (bottom panel).

59 Relative fold induction of Dlx5 and Dlx6

OIOOIOOIOOIOOIO ooooooooooo

• n

1 1 1 o ^1> o

-V s CAG

> n n encoding retroviruses. Control LZRS-infected samples quickly became very confluent

(Figure 7). By day 6 the cells started to condense, and by day 14 chondrogenic nodules were obvious. The Z,Z/?5-Z)/x5-infected cells did not change greatly over the differentiaton period. By day 14 they were moderately confluent, but there were obviously fewer cells than the LZRS control and cells were not condensed. The LZRS-

D/jc(5-treated cells also had fewer cells than the ZZftS-treated cells. However, unlike the

Z,Z/?S'-.D/;c5-infected cells, by day 12 the cells were starting to grow larger and by day 14 the cells appeared considerably larger. By day 17, there was a significant amount of cell death and the remaining attached cells were dying.

Alcian blue staining of these cultures revealed that chondrogenic nodules formed in the control cultures by day 14 (seen as dark areas in Figure 8). While large chondrogenic nodules are visible in the LZRS control cells, no nodule formation is present in either the LZRS-Dlx5 or LZRS-Dlx6-infected cells.

3.4 Real-time PCR analysis of gene expression in ATDC5 cells overexpressing

Dlx5 and Dlx6

We next examined the expression of various chondrogenic and osteogenic marker genes to investigate the basis for the differences in cell behaviour following misexpression of Dlx5 or Dlx6. Real-time PCR of several chondrogenic genes was used to determine the effect of Dlx5 and Dlx6 on chondrogenesis in ATDC5 cells. ATDC5 cells were grown to confluence (day 0). They were infected with LZRS or a LZRS encoding Dlx5 or Dlx6 for two consecutive days, then cultured with the media changed every second day thereafter. RNA was extracted from the cells at day 0, 1, 2, 4, 6, 8, 10,

61 Figure 7: Microscope photographs of differentiating ATDC5 cells. ATDC5 cells were transduced with a retrovirus encoding either Dlx5, Dlx6 or a control virus (LZRS). Cells were fixed in 100% methanol between day 0 (reaching confluence) and day 17 and stained in Alcian blue overnight. Cells were examined using a Leica DM IRE2 microscope at 20x magnification and visualized in Openlab. Day 0, 4, 8,12,14 and 17 are shown with LZRS-, LZRS-Dlx5- and LZRS-Dlx6-'mfected cells.

62 LZRS Dlx5 Dlx6

63 Figure 8: Microscope photographs of differentiating ATDC5 cells stained with Alcian blue. ATDC5 cells were transduced with a retrovirus expressing either Dlx5, Dlx6 or a control virus (LZRS). Cells were fixed in 100% methanol between day 0 (reaching confluence) and day 17 and stained in Alcian blue overnight. Cells were examined using a Leica MZ 125 microscope at lOx magnification and imaged using Openlab (Improvision). Day 10, 12, 14 and 17 are shown with LZRS-, LZRS-Dlx5- and LZRS- Z)/x<5-infected cells. Arrows indicate chondrogenic nodules.

64 Day 10 Day 12 Day 14

in I N

• ,J^ff; ' „ '.^.iVJ"- , ';'«'':•*"" . . .. r^t;.| v^"iwwi^^.*^*HfeSl '. • • -. '••--•.;• ^^^yMrnHH .;..'.-r,.:.y -*.i\ .'".' ' V ^w^n^n^H

'**'" *•!» 'J* -*•/?"* ^ J>V ^'Mj^W iMfiSf ^ii^ . . * •*. t«A&jSa§g| ***** *****J[eli£ "*:itJ^fflHB * 'J A'/^^PQ ' •> ,< .v. BiiBWBliniiiifci' m x ' ' 7 i>~- • i ^. ~-^iif"n|"!BBH3|MJ|j , 14 --^•^SIMBSJ I '- , (* ^ ... f ,i ,>" , ! 'i-i* '^'^M" '*"*'*?sf*!«Si®^SHlI ^W^r^^^mnM Ife'v-& '•.-•' •" ^• PSB&&&K 1

65 12, 14 and 17 and reverse transcribed into cDNA, which was then used as a template for real-time PCR. Real-time PCR was analyzed by the delta-delta CT method: CT values were all normalized to actin, then presented as fold induction over day 0 levels. The resulting graphs are shown in Figures 10 and 11. An induction profile of each gene is shown for control ZZflS-infected cultures in Figure 9.

3.4.1 Induction profile of ZZftS-infected cultures

In control cultures, there is a steady increase in Col2al expression, leading to an induction above day 0 of around 500 fold by day 17. The fold induction of CollOal in

LZRS control samples shows a slight increase over the differentiation period, from around

5-fold induction, to between 10-15-fold, however, this expression never reaches the levels of induction seen with Col2al. The levels of Osc increase slowly over the differentiation period in LZRS control samples, eventually reaching around 50-fold induction over day 0.

Osp shows a large increase in expression in LZRS control samples, with up to 100-fold induction. Sox9 shows a steady increase in expression to a level of about 20-fold over day

0 in control samples. Runx2 expression in control samples shows an increase early in the differentiation period, to around 8-fold induction, which then becomes fairly stable in the later stages, with small amounts of fluctuation. Ihh expression increases slowly later in the differentiation period, to around 10-fold induction in the LZRS control samples. The level of induction of Dlx3 expression slowly increases over the experimental period, with more significant increases especially in the last few days, to around 15-fold induction.

The levels of Dlx5 show a fairly steady expression level in control samples, with around

6-fold induction. The levels of Dlx6 in control samples show little increase in induction over the differentiation period, with only as much as 4-fold induction.

66 Figure 9: Real-time PCR analysis of chondrogenic marker genes (A) Col2al, (B) CollOal, Osc and Osp and (C) Rural, Ihh, Sox9, Dlx3, Dlx5 and Dlx6 in ATDC5 cells infected with the LZRS control plasmid. ATDC5 cells were infected with a LZRS control plasmid. Cells were harvested at different time points from day 0 (reaching confluence) to day 17. RNA was extracted using TRIzol and reverse transcribed using Superscript II. cDNA was then using in real-time PCR with different primer sets.

67 (A)

600"

^•^ 500 -^—Col2a1

400 -

300 -

200 - ,-"•" 100

0 0 1 2 4 6 8 10 12 14 17

Days post-infection

-m-~ Conoal 100 Osc / 80 Osp /

y 'i 40 / '

20 -

_ -•- •"" """•--« 0 - 0 1 2 4 6 I 8 I 10 I 12 14 17

Days post-infection

68 Figure 10: Real-time analysis of chondrogenic structural genes, Col2al (A), CollOal (B), Osc (C) and Osp (D), in differentiating ATDC5 cells. ATDC5 cells were infected with a retrovirus encoding either Dlx5, Dlx6 or a control virus. Cells were harvested at different time points from day 0 (reaching confluence) to day 17. RNA was extracted using TRIzol and reverse transcribed using Superscript II. cDNA was then using in real time PCR with different primer sets. Graphs shown indicated values normalized to actin then presented as a fold induction over the levels of day 0. * indicates a statistically significant difference between Dlx6 and LZRS control (p<0.05).

69 > Relative fold induction of Col10a1 Relative fold induction of Col2a1 s ss s *= Hi

~ * -

Day s post-inf e K * * on_ to f * p s pH ,_ * fiB 1 f3

MI»! ( j£ O * IMgiPiJ ' =3 *

Relative fold induction of Osc Relative fold induction of Osp i & s s s 3 S

O in T3 Figure 11: Real-time analysis of chondrogenic transcription and growth factors, Sox9 (A), Rural (B), Ihh (C), Dlx3 (D), Dlx5 (E) and Dlx6 (F), in differentiating ATDC5 cells. ATDC5 cells were infected with a retrovirus encoding either Dlx5, Dlx6 or a control virus. Cells were harvested at different time points from day 0 (reaching confluence) to day 17. RNA was extracted using TRIzol and reverse transcribed using Superscript II. cDNA was then using in real-time PCR with different primer sets. Graphs shown indicated values normalized to actin then presented as a fold induction over the levels of day 0. *, f, % indicates a statistically significant difference (p<0.05) between Dlx6 and LZRS control, Dlx5 and the LZRS control and Dlx5 and Dlx6, respectively.

71 O Relative fold induction of Dlx5 Relative fold induction of Ihh Relative fold induction of Sox9 w a a es « a

.....!.!.™"i;Li. _+" Day s post-infectio n

Dl x O x „ un ilml liilllllllilllHill IIIIIIII ill 1 '

• • ' n _ j-— 1 (Hi—iiiiiii iiiiiiwiiiii in i

Relative fold induction of Dlx6 Relative fold induction of Dlx3 Relative fold induction of Runx2 o 3 B SS 6 S S MM ii Day s y s post-infectio n a CT> X I X ?' 3

iv) fsmmmm^t 1 * ^^^J 1 bi

=3 •-i * 3.4.2 Dlx6 suppresses CoUal activation

The expression of Col2al in samples with Dlx5 overexpression shows some variability. At earlier timepoints, from day 1 through day 10, Dlx5 slightly upregulates expression of CoUal compared to the control cultures, however, this was not statistically significant. After day 10, Dlx5 seemed to show little effect or a slight repression of

CoUal, but this is also not significant. In DZxtf-treated samples, CoUal shows a marked reduction in activation, with the highest fold induction being about 40 fold over day 0 expression. The downregulation of CoUal by Dlx6 overexpression is statistically significant from day 2 to day 17, with the exception of day 12.

3.4.3 CollOal expression is not significantly changed by the expression of Dlx5 and Dlx6

The fold induction seen in samples infected with LZRS-Dlx5 show an overall reduction in CollOal expression, however, since the expression of CollOal is so variable, these observations are not statistically significant. D/xtf-treated samples show a similar pattern to those of Z)/x5-treated samples, with a slight reduction in expression that is not significant.

3.4.4 Both Dlx5 and Dlx6 upregulate expression of osteocalcin

In samples treated with both Dlx5- and Z)/x6-encoding retrovirus, a marked increase in Osc expression was seen. Since LZRS-Dlx5-infected samples showed a high degree of variability, none of the observations are significant, even though an obvious trend of increased expression exists. However, misexpression ofDlx6 significantly increases expression of Osc from day 4 to day 14, reaching levels of induction of around

73 300-fold. There appears to be a trend of slowly increasing expression of Osc, likely as

levels of Dlx6 accumulate.

3.4.5 Osteopontin expression is slightly downregulated by Dlx6 overexpression

Dlx5 overexpression causes a slight increase in Osp expression at some timepoints, however, this effect is not significant. Overexpression of Dlx6 resulted in consistently lower levels of Osp transcription compared to the control cultures, although only day 1 shows a statistically significant difference.

3.4.6 Sox9 expression is significantly downregulated by both Dlx5 and Dlx6

Both Dlx5 and Dlx6 overexpression suppressed Sox9 induction. LZRS-Dlx5- infected samples show a consistent repression of Sox9 induction by as much as 4-fold.

This observation is only statistically significant on day 4 and day 8, although the trend continues through the entire differentiation period. Dlx6 overexpression shows a similar level of repression of Sox9 as Dlx5, with levels of Sox9 induction clearly lower throughout the experiment. LZRS-Dlx6-infected samples show a significant downregulation on both day 4 and day 17.

3.4.7 Overexpression of Dlx5 and Dlx6 has no effect on Runx2 expression

The addition of either Dlx5 or Dlx6 causes no significant changes in Runx2 expression compared to the control cultures.

3.4.8 Dlx5 and Dlx6 overexpression results in increased induction of Ihh expression

Both Dlx5 and £)/x<5-treated cells show a fairly consistent increase in Ihh induction compared to the control cultures. ZZi?5'-£)/x5-infected samples show a modest increase in

74 expression over the control, with a significant increase on day 4. Dlx6 overexpression results in a slightly higher level of Ihh induction than DZx5-treated samples and both day

2 and day 12 show significantly increased expression from the control levels.

3.4.9 Dlx3 expression is differentially controlled by Dlx5 and Dlx6

Dlx5 overexpression appears to cause an increase in induction of Dlx3 over much of the differentiation period, especially on day 1 when it is significantly increased.

Conversely, Dlx6 causes a downregulation of Dlx3 expression with significant values on day 12 and day 17. This effect of Dlx6 is only seen after day 8 however, as before that

Dlx6 appears to upregulate Dlx3 expression, albeit not significantly.

3.4.10 Dlx6 has no effect on the expression of Dlx5

The addition of Dlx6 overexpression caused a slight decrease in Dlx5 expression, but this effect was not significant.

3.4.11 Dlx5 has no effect on the expression of Dlx6

Dlx5 overexpression appears to result in little change to the expression of Dlx6.

Although a few days show slightly decreased levels of Dlx6, these values are not significantly different.

A summary of the effects of Dlx5 and Dlx6 misexpression in the real-time PCR results is shown in Table 3.

75 Table 3: Summary of the effects of Dlx5 and Dlx6 overexpression on the induction of chondrogenic marker genes

Dlx5 Dlx6 Sox9 4 4 Col2al 4 Ihh H H Runx2

CollOal ^— ^—

Osc —— II Osp fl Dlx3 tf U Dlx5 N/A

Dlx6 N/A

76 4. Discussion

4.1 Stage of differentiation of ATDC5 cells

As all of the chondrogenic marker genes were upregulated during the differentiation period, estimating the level of differentiation of the ATDC5 cells is difficult. Early studies using ATDC5 cells found no expression of most of these genes at day 0 (Shukunami et al., 1997), however, this study detected low levels of expression of all of them (Figure 4). This could be explained by the different techniques used. Earlier studies predominantly used Northern blot to detect expression, however, this study used either semi-quantitative or real-time PCR, which are more sensitive and able to detect even low expression levels. More recent studies using real-time PCR did find much earlier expression of chondrogenic marker genes, with increases in expression of Col2al by day 2 and Ihh and Runx2 by day 4 after the addition of insulin (Chen et al., 2005;

Huang et al., 2004). In this study, the levels of most of the genes examined steadily increased, but none of them had begun to decrease by the end of the study, likely due to the much slower progression of differentiation in ATDC5 cells without insulin.

The ZZ/?5-treated cells began condensing by day 6 and by day 14 chondrogenic nodules were visible, indicating that those cells were in the proliferating stage of chondrocyte differentiation. However, both the Dlx5 and DZx<5-expressing cells had fewer cells, did not form chondrogenic nodules and by day 14, the D/xtf-infected cells have a very different morphology (Figures 7 and 8). Therefore, it seems that the control LZRS- infected cells and the LZRS-Dlx5 and LZRS-Dlx6-treated cells are at different stages of differentiation.

77 4.2 Dlx5 and Dlx6 reduce ATDC5 cell number

Dlx proteins have been shown to positively regulate differentiation of numerous

cell types (Ferrari and Kosher, 2002; Holleville et al., 2007; Miyama et al., 1999;

Morasso et al., 1996; Tadic et al., 2002). In this study, both Dlx5 and Dlx6 positively

regulate ATDC5 differentiation, albeit somewhat differently. Bright field microscopy

shows a clear reduction in cell number in LZRS-DU5 and LZRS-Dlx6-'mfected cultures

compared to the LZRS control which continued to proliferate and condense over the

course of the experiment (Figures 9,10 and 11). The effect on cell number was much

stronger in ZZftS-DZxo'-infected cells compared to LZRS-Dlx5, with the Dlx5

misexpressing cells becoming more confluent near the end of the 17 day period. Whether

this effect is due to a positive regulation of differentiation by upregulating late markers of

chondrogenesis, or a direct negative regulation of cell proliferation is not yet clear. There

is some evidence for both of these possibilities. Dlx proteins activate expression of late

stage markers in several cell types: GAD in the forebrain (Stuhmer et al., 2002), Bsp in

osteoblasts (Benson et al., 2000; Roca et al., 2005), profllaggrin and loricirin in

keratinocytes (Morasso et al., 1996) and chorionic gonadotropin and 36-HSD in the placenta (Peng and Payne, 2002; Roberson et al., 2001). However, there is also some

circumstantial evidence to suggest they also suppress cell proliferation. Two Dlx-binding proteins, Msx and MAGE-D1 have been shown to regulate cell proliferation: Msx positively regulating proliferation by upregulating cyclin Dl and increasing CDK4

activity (Hu et al., 2001) and MAGE-D1 negatively regulating cell cycle progression by

associating with necdin (Kuwajima et al., 2006). Both proteins are known to interact with

Dlx and either repress its function (Msx), or increase transcriptional activation (MAGE-

78 Dl) (Masuda et al., 2001; Zhang et al., 1997). The Drosophila homologue, Distal-less, has recently been shown to bind a transcription factor, DREF, required for expression of proliferation-related genes, and inhibits its ability to bind DNA (Hayashi et al., 2006).

The reduction in cell number may also be accounted for by increased cell death, particularly in LZRS-Dlx6-infected cultures where many cells were clearly dying by day

17. In this respect, one of the Dlx family members, DLX4, has been shown to suppress apoptosis (Shimamoto et al., 2000; Sun et al., 2006) and it was also found that Dlxl,

Dlx2, Dlx3 and Dlx4 were expressed at higher levels in cell lines that are more sensitive to apoptotic induction (Ferrari et al., 2003). However, the mechanism of control of proliferation for the Dlx proteins remains to be further studied.

4.3 Dlx5 and Dlx6 show some opposing roles in chondrocyte differentiaton

Most of the linked Dlx gene pairs are co-expressed and all Dlx genes share very high homology within the homeobox sequence, however, they also clearly encode some specific functions. For example, while double knockouts of both Dlxl and Dlxl show a more severe phenotype than the respective single knockouts, indicating overlapping functions, both single knockouts demonstrate some functions that cannot be compensated for by the other gene (Qiu et al., 1997; Qiu et al., 1995; Thomas et al., 1997). Dlx5 knockouts display defects in ossification of the skull, the branchial arches and the sensory organs, for which Dlx6 cannot compensate (Acampora et al., 1999; Depew et al., 1999).

Although a Dlx6 knockout phenotype has not yet been published, it is likely that Dlx6 also has some specific functions.

Besides the altered behaviour of Dlx5 and DZrtf-misexpressing cells, Dlx5 and

Dlx6 also showed differential effects on specific gene prqducts. In this study, it was

79 shown that Dlx6 overexpression reduced levels ofDlx3 expression, while Dlx5 overexpression caused an increase in Dlx3 expression (Figure 11). Cross-regulation of

Dlx gene expression has been shown to occur, as Dlxl, Dlx2 and Dlx5 can activate transcription from the intergenic Dlx5/6 enhancer (Stuhmer et al., 2002; Zerucha et al.,

2000; Zhou et al., 2004). However, it has not been shown that a Dlx protein can repress the transcription of another. Whether Dlx5 is directly upregulating Dlx3 by binding to a

Dlx3/4 enhancer or whether Dlx6 is directly repressing Dlx3 transcription is not clear.

However, since Dlx3 is expressed at late stage chondrocyte differentiation (Hassan et al.,

2004), it is unlikely that the effect of Dlx6 on Dlx3 expression is due to the cells being in a more differentiated state. This effect must be studied further.

4.4 Dlx6 overexpression blocks early chondrocyte differentiation and promotes late differentiation in ATDC5 cells

The earliest markers of chondrocyte differentiation examined in this study were

Sox9 and Col2al, both expressed in proliferating chondrocytes. Dlx6 overexpression was found to significantly decrease the fold-activation of both Sox9 and Col2al compared to both the control- and LZRS-Dlx5-infected cells (Figures 10 and 11). Chondrocyte-specific expression of Col2al has been shown to be driven by a 48 bp region in the first intron of the gene (Zhou et al., 1998). There are three HMG binding sites within this region, of which Sox9 binds one and increases transcription of Col2al (Zhou et al., 1998). The reduction in Col2al could then be explained by the decreased Sox9 expression. Sox9 expression is controlled by BMP signaling which is mediated within the cell by activated

Smad proteins (Goldring et al., 2006; Healy et al., 1999).

80 It is unlikely that Dlx6 is directly repressing the transcription of Sox9 and Col2al:

Dlx proteins usually act as transcriptional activators and Dlx5 and Dlx6 are both expressed around the time that Sox9 and CoUal are highly expressed (Bendall et al.,

2003; Ferrari and Kosher, 2002; Ghoul-Mazgar et al., 2005; Hassan et al., 2004). The most likely method by which Dlx6 is downregulating Sox9 and CoUal is indirectly by causing the cells to mature into more differentiated chondrocytes, thereby turning off

Sox9 and Col2al expression. The cell morphology of the Z)/x5-infected cells also strongly supports this theory. By day 14, the cells have become very large and appear to be hypertrophic chondrocytes (Figure 7). Within the next three days many of the cells died, appearing to have reached terminal differentiation and undergone apoptosis.

The upregulation of Osc, a late marker of chondrogenesis and osteogenesis, by

Dlx6 would also support the hypothesis that Dlx6 overexpression causes the cells to shift to a later stage of differentiation. However, this may also be a direct effect. Dlx5 has been shown to mildly activate the basal Osc promoter and relieve Msx2 repression (Newberry et al., 1998). Also, it was shown in osteoblasts that a complex of Dlx3, Dlx5 and Runx2 can activate the Osc promoter (Hassan et al., 2004). Therefore it is possible that Dlx6 binds and activates transcription of the Osc promoter in ATDC5 cells, more strongly than

Dlx5. One problem with the idea that Dlx6 induces precocious hypertrophy is the absence of an upregulation of CollOal in D/xtf-infected cells. CollOal is a marker of hypertrophy so if these cells are hypertrophic, they should be expressing this gene. This gene showed a great deal of variation in the real-time PCRs, which could be due to inefficient or non specific primers. Repeating the real-time PCRs with a different pair of primers is needed to clarify the CollOal expression of ZZfliS-D/jctf-infected cultures.

81 5. Future directions

In order to further define the role of Dlx5 and Dlx6 in chondrogenesis, more

studies should be conducted. A study of their role in proliferation of chondrocytes is

necessary. This could be accomplished using a cell proliferation assay and using real-time

PCR to determine if various cell cycle-related factors are upregulated or downregulated in

response to Dlx overexpression. A good target may be the transcription factor DREF

(DNA replication-related element-binding factor), required for expression of many

proliferation-related genes, whose Drosophila homologue was shown to be negatively

regulated by Distal-less (Hayashi et al., 2006). It must also be determined whether the

effect of Dlx5 and Dlx6 on Dlx3 expression is direct or indirect. Repression of one Dlx

gene by another has not been previously shown, so this would be interesting.

Finally, unraveling the mechanism by which Dlx6 promotes hypertrophy in

ATDC5 cells is of utmost importance. A first step may be to determine the effect of Dlx6

on various gene promoters, including CoUal, Sox9 and Osc, to determine if these genes

are direct targets of Dlx6. It may also be necessary to examine more genes by real-time

PCR, especially late markers of chondrogenesis, such as alkaline phosphatase, VEGF,

Wnt4 and MMP-13 (Church et al., 2002; D'Angelo et al., 2000; Gerber et al., 1999;

Ishikawa et al., 1987), and further clarify the effect on CollOal.

82 Appendix 1: Measuring the transcriptional activity of Dlx proteins

Al.l Introduction

Several lines of evidence indicate that Dlx proteins are activating transcription factors. However, little is known about how transcription activation is mediated by the

Dlx proteins. Homology searches reveal no obvious activation domains in the sequences of Dlx5 and Dlx6, however, some transcriptional activation domains have been mapped for other Dlx proteins. Dlx3 was found to have an activation domain on each side of the homeodomain, one in its N-terminal domain (NTD) and another in its C-terminal domain

(CTD), which were slightly acidic (Feledy et al'., 1999). Other domains that have been located are a polyhistidine tract and glycine-rich domain in Dlx2 (McGuinness et al.,

1996) and the NTD of Dlx2, Dlx3 and Dlx5 all contain conserved tyrosine residues (Liu et al., 1997). However, more work is needed to reveal the mechanisms by which Dlx proteins activate transcription. As a first step towards this goal, the transcriptional activity of all six mammalian Dlx proteins were compared in a standardized assay.

A1.2 Materials and Methods

Plasmids

Myc-tagged, mouse Dlx open reading frames were cloned mpcDNA3 (Coubrough and Bendall, 2006). CMV-pGal contains the fi-galactosidase gene downstream of the human cytomegalovirus (CMV) immediate-early promoter, and was used to normalize reporter gene transcription with respect to transfection efficiency. pGL3-promoter and

83 pGL3-control (Promega) contain the SV40 (Simian virus 40) immediate early promoter alone or with the SV40 enhancer, respectively, upstream of the firefly (Photinus pyralis) luciferase gene.

Cell culture and transfection

Human embryonic kidney (HEK) 293 cells were maintained in DMEM supplemented with 10% FBS, 100 U/mL penicillin, 100 ug/mL streptomycin and 2 raM L-glutamine in a humidified incubator at 37°C, 5% CO2. 6.5 x 104 cells were seeded per well in a 24-well plate, 24 hours prior to transfection. Cells were transfected at 50-70% confluence with 2 jagpcDNA3-Dlx expression plasmids, 1 \xgpGL3-promoter and 0.5 (ag CMV-j3Gal using

2 (ig of polyethylenimine (PEI) (Sigma) per jug of DNA in serum-free DMEM. Each transfection was performed in duplicate or triplicate.

Cell harvesting and reporter assays

Cell extracts were collected 48 hours post-transfection using lx reporter lysis buffer

(Promega). Luciferase expression levels were measured according to the manufacturer's protocol (Promega) using a Turner TD-20e Luminometer. Values were standardized by measuring P-galactosidase activity using 2x p-gal buffer (120 raM Na2HP04, 80 mM

NaH2P04, 2 mM MgCl2, 100 mM P-mercaptoethanol and 1.33 mg/mL 0-nitrophenyl P-D galactopyranoside) and a standard protocol (Sambrook, 1989). Briefly, 10 uL of cell extract was mixed with 150 uL of P-gal buffer, then incubated at 37°C for 30 minutes.

The reaction was stopped by the addition of 500 uL of 1M Na2Co3; the optical density of the mixture was measured at 420 nm.

84 A1.3 Results and Discussion

Dlx proteins can activate transcription from the SV40 promoter

The SV40 promoter, which drives the luciferase gene in pGL3-promoter vectors,

contains two Dlx-consensus sequences (TAATT), located 89 bp apart, in an inverted

orientation (Figure 12A). WhenpcDNA3-Dlx5 was co-transfected with the pGL3- promoter reporter plasmid, this promoter responded to Dlx5 in a dose-dependent manner

(Figure 12B). We therefore used this reporter to compare the transcriptional activites of

all six mammalian Dlx proteins. Dlx expression plasmids were co-transfected with/?GZJ- promoter and CMV-fiGal into HEK 293 cells and after 48 hours a luciferase reporter

assay was performed. Luciferase values were normalized to |3-galactosidase produced

from the CMV-PGal vector as a control. The Z)/x-transfected samples were then

normalized to the vector-transfected (pcDNA3) samples for the reporter and values shown

are fold stimulation above the basal level. All of the mouse Dlx proteins were shown to

activate the SV40 promoter, although with differing strengths (Figure 12C). Dlxl, Dlx2

and Dlx3 all showed the strongest activation potential, with Dlx4, Dlx5 and Dlx6

showing lower levels. These differences may be due to reduced DNA-binding affinity to

this particular promoter or the actual strength of each of the proteins transactivation

domains. However, since the Dlx proteins have nearly identical homeodomains (Bendall

and Abate-Shen, 2000), and likely bind DNA with equal affinity, these differences are

more likely due to actual differences in activation potential between the mammalian Dlx

proteins.

85 Figure 12: Fold activation of murine Dlx proteins on the SV40 promoter. (A) Sequence of the SV40 promoter found within the plasmid pGL3-promoter: Dlx-consensus binding sites (TAATT) are highlighted in red. (B) Fold activation of murine Dlx5 on pGL3- promoter. Different levels of pcDNA3-Dlx5 were transfected into HEK293 cells: 0 ug of pcDNA3-Dlx5, 0.125 ug, 0.5 ug and 2.0 ug along with 1.0 ug of pGL3-promoter. Results shown are averages from one experiment done in triplicate. Dlx5 activation is normalized to basal luciferase values in cells transfected with 2.0 ug of pcDNA3. (C) Fold activation of murine Dlx proteins on pGL3'-promoter. 2.0 ug of each D/x-expression plasmid, pcDNA3-Dlxl-Dlx6 or pcDNA3 were transfected into HEK293 cells with 1.0 ug of pGL3- promoter. Values shown are fold stimulation of normalized luciferase values from Dlx- transfected samples over those of pcZW^J-transfected samples. Each bar represents the meansisem from 2 independent experiments, each performed in triplicate.

86 (A) ATCTCAATTA GTCAGCAACC ATAGTCCCGC CCCTAACTCC GCCCATCCCG

CCCCTAACTC CGCCCAGTTC CGCCCATTCT CCGCCCCATC GCTGACTAAT

TTTTTT'] ATGCAGAGG CCGAGGCCGC CTCGGCCTCT GAGCTATTCC

AGAAGTAGTG AGGAGGCTTT TTTGGAGGCC TAGGCTTTTG CAAAAAGCTT

(B)

35.00 -i

30.00 J [«*« g 25 00 *i 'f 20.00 - 8 15.00 - g £ 10.00 - 5.00 - •• ill i Oug 0.125ug 0.5ug l•2ug pcDNA-DlxS (ug)

160.00 -|

E 140.00 - | 120.00- B 100.00 f 2 80.00 - a 60.00 • 1 40.00 - & 20.00 - ,

o.oo- pcDNA D x1 Dx2 Dlx3 Dx4 Dlx5 Dlx6

87 Appendix 2: Mapping the transactivation domains in DIx5 and

DIx6

A2.1 Introduction

Limb bud mesenchymal cells from Dlx5/(>J~mice, showed a reduction in the potential to form chondrogenic nodules, indicating that both Dlx5 and Dlx6 are necessary for chondrogenesis. Also, at high cell densities in a micromass culture, misexpression of both Dlx5 and Dlx6 was shown to increase the nodule number. Finally, it was determined that the domains required for this function was the amino terminal domain (NTD) of Dlx5 and both the NTD and the carboxy terminal domain (CTD) of Dlx6 (Hsu et al., 2006). In order to determine if the domain required for this biological function had transcriptional activity, an in vitro assay testing the transcriptional activation potential of Dlx5 and Dlx6, and various truncations of these proteins, was performed. These results were published in

(Hsu et al., 2006).

A2.2 Materials and Methods

Plasmids

All chicken Dlx5 and Dlx6 open reading frames were subcloned into pcDNA3

(Invitrogen) from a previously constructed vector (Hsu et al., 2006). Each Dlx gene is divided into three segments: an NTD, the homeodomain (HD) (including a small portion of the NTD to conserve the nuclear localization signal (NLS)) and the CTD. All Dlx genes inpcDNA3 plasmids are preceded by two tandem c-Myc epitopes (EQKLISEEDL with a GS linker). All of the constructs used are described in Figures 13. pcDNA3-Dlx5, pcDNA3-Dlx5AC, pcDNA3-Dlx5/6 (containing the NTD and HD of Dlx5 and the

88 Figure 13: Chicken Dlx5 and Dlx6 polypeptides. Domain-specific amino acid numbers are shown. The NTD of Dlx5 is dark purple. The homeodomain is shown in yellow. The CTD is blue. The NTD of Dlx6 is light purple. The homeodomain is orange. The CTD is blue. The NTD of Dlx5 is yellow.

89 286 Dlx5 - ,; ~--y-.: :" ' :; IFt^amlgSmKsmfMKZM ,-M»T.'\;::'^T:xsr.<"iA«^s:rr.;.,.*v;r,..™!:;.,:«r':i!;.s;!i

195 Dlx5AC

123 286 Dlx5AN

123 195 Dlx5HD

267 Dlx6

200 Dlx6AC

128 267 Dlx6AN

128 200 Dlx6HD

195 201 267 Dlx5/6

90 CTD ofDlx6),pcDNA3-Dlx6, pcDNA3-Dlx6AC andpcDNA3-Dlx6ANwere all subcloned

into pcDNA3 using the restriction enzymes Apal and EcoRI. pcDNA3-Dlx6HD was

constructed by PCR using primers 5'-

TATAGGATCCGAAAACGGCGAGATCAGATTC-3' and 5' -

TATATCTAGATCACTTCAGCAGCTTCTTAAACTTGG-3'. adding the restriction

enzymes BamHl andXbal (underlined), with Slaxl3-Dlx6 (Hsu et al., 2006) as a

template. pcDNA3-Dlx5AN and pcDNA-Dlx5HD were previously constructed (Melissa

Coubrough, unpublished).

Cell culture, transfection and reporter assays were done as described in Appendix 1.

A2.3 Results and Discussion

Characterizing transactivation domains of chick Dlx5 and Dlx6

Chick Dlx expression plasmids were transfected into HEK293 cells with pGL3- promoter and CMV-flGal. Values shown are normalized to basal luciferase levels in

/?aDiV/43-transfected cells (Figure 14). Chick Dlx5 and Dlx6 show transcriptional

activation of the SV40 promoter, around 8 and 10-fold respectively. When the NTD of

Dlx5 was removed, transcriptional activation increased over that of full-length Dlx5,

showing around 40-fold activation. Removing the CTD slightly reduced activation

compared to Dlx5, while the HD had no activation potential on its own. With Dlx6, removing the NTD and the CTD resulted in similar fold-activation, slightly higher than full-length Dlx6. Once again, the homeodomain alone had no transactivation potential. A chimeric protein, containing the NTD and HD of Dlx5 and the CTD of Dlx6 showed higher activation than either Dlx5 or Dlx6.

91 Figure 14: Dlx5 and Dlx6 are transcriptional activators. 0.33 |j.g of chickpcDNA-Dlx expression plasmids orpcDNA3 were transfected into HEK293 cells with 0.5 y.g of pGL3-promoter and assayed for luciferase reporter activity after 48h. Values shown are fold stimulation of normalized luciferase activities from D/x-transfected samples over those of/?cZWv4J-transfected samples for each construct. Each bar represents the means±sem from between 3 and 12 independent experiments, each done in triplicate (Dlx5, n=12; Dlx5AC, n=7; Dlx5AN, n=8; Dlx5HD, n=4; Dlx6, n=l 1; Dlx6AN, n=7; Dlx6AC, n=8; Dlx6HD, n=3; Dlx5/6, n=5).

92 'SO

» so 4 40 4 m 4 20

l"^"~' yiiiuimmmsffiMjm^wwiMMia&miii y *SSS+SSSSilLl

93 Both Dlx5 and Dlx6 show a transactivation function on the SV40 promoter which matches their general role as transcriptional activators. However, different domains of

Dlx5 and Dlx6 seem to be important for this activity. In Dlx5, the NTD contains a potent transcriptional activation domain, which appears to be partially masked in the full-length protein since removing the CTD significantly increased its activity. In contrast, both the

NTD and CTD of Dlx6 have similarly active transcription activity. These results correspond well with the domain requirements for chondrogenic nodule formation (Hsu et al., 2006). This indicates that the increase in chondrogenesis seen when Dlx5 and Dlx6 are over-expressed is due to their action as transcriptional activators.

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