Investigating Dlx and Zbed1 Protein Interactions During Cell Cycle Progression

Home , DLX3, DLX gene family, ZBED1

By Shayne McLaughlin

A Thesis presented to The University of Guelph

In partial fulfilment of requirements for the degree of Master of Science in Molecular and Cellular Biology

Guelph, Ontario, Canada © Shayne McLaughlin, March, 2019

Abstract

INVESTIGATING DLX AND ZBED1 PROTEIN INTERACTIONS DURING CELL CYCLE PROGRESSION

Michael Shayne McLaughlin Advisor: University of Guelph, 2019 Professor A. Bendall

Genes of the vertebrate distal-less homeobox (Dlx) family encode a group of homeodomain transcription factors that act as pro-differentiation factors in a variety of tissues.

Recently, we have shown that forced expression of both Dlx5 and Dlx6 decreases the rate of cellular proliferation in various cell types, at least in part by delaying entry into S-phase. In

Drosophila, this may be accomplished by a protein-protein interaction involving the homologue

Distal-less (Dll) and the DNA replication-related element binding factor (DREF) – a pro- proliferative transcription factor that positively regulates many genes associated with cell cycle progression. Here, we demonstrate that this interaction may be a conserved mechanism for Dlx- mediated antagonism of cell division since chick Dlx6 physically interacts with the chick orthologue of DREF, Zbed1. We investigated the functional significance of this physical interaction by examining aspects of cell cycle progression in several cell lines. Acknowledgements

I would first like to extend my gratitude to my advisor Dr. Andrew Bendall for his support, guidance, and patience during the course of this thesis. I am extremely thankful to have had the opportunity to work in this laboratory. I would like to extend my thanks to Dr. Mosser, who is not only a member of my committee but also supervised my undergraduate project. His suggestions have helped tremendously.

I would like to thank all of the members of the Bendall Laboratory who I have had the pleasure of working with for the past several years, especially Sarah Yull, with whom I have worked closely for the last two semesters. Furthermore, I would like to extend this thanks to the members of the neighboring laboratories (Dr. Nina Jones, Dr. Richard Mosser, and (formerly)

Dr. Terry Van Ray), but especially Lina Abou Zeid, Haider Khan, Claire Martin, and Manali

Tilak, who have been instrumental in fostering an enjoyable environment, but also explaining various techniques for which I was unfamiliar. I would like to extend my gratitude to Paul Kron, who was instrumental in teaching me to use the flow cytometer.

Lastly, I would like to thank my friends, and family, for all the love and support they have given me throughout this project. In particular, I want to take this opportunity to thank my girlfriend, who has been a pillar of love and support for me throughout the past two years.

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Declaration Of Work Performed All work contained herein was performed by Shayne McLaughlin, with the following exceptions. Sarah Yull assisted in the preparation and collection, and western blots of the selected HeLa cell populations.

Table of Contents ABSTRACT ...... ii

ACKNOWLDEGEMENTS ...... iii

DECLARATION OF WORK PERFORMED ...... iv

LIST OF FIGURES ...... iix

LIST OF TABLES ...... ix

ABBREVIATIONS ...... x

CHAPTER 1: INTRODUCTION ...... 1

THE ROLES OF PROLIFERATION AND DIFFERENTIATION IN EMBRYONIC

DEVELOPMENT ...... 1

THE G1/S RESTRICTION POINT ...... 2

DREF/ZBED1, THE “MASTER KEY” TO CELLULAR PROLIFERATION ...... 8

DLX HOMEOBOX GENES ...... 11

CONTEXT DEPENDENT EFFECTS OF DLX5 ON CELLULAR PROLIFERATION ...... 14

RATIONALE ...... 18

HYPOTHESIS ...... 19

CHAPTER 2: MATERIALS AND METHODS ...... 20

PLASMIDS ...... 20

PGEX PROTEIN INDUCTION ...... 21

GLUTATHIONE BEAD PREPARATION ...... 21

GST-PULLDOWN ...... 22

CELL LINES AND REAGENTS ...... 22

TRANSFECTION AND SELECTION ...... 22

COVERSLIP PREPARATION ...... 23

EdU INCORPORATION ASSAY ...... 23

INDIRECT IMMUNOFLUORESCENCE ...... 23

FLOW CYTOMETRY ...... 25

IMMUNOBLOTTING ...... 26

QUANTITATIVE POLYMERASE CHAIN REACTION ...... 27

STATISTICAL ANALYSIS ...... 29

CHAPTER 3: RESULTS ...... 30

3.1 THE TRANSCRIPTIONAL EFFECTS OF DLX5 EXPRESSION AT THE RESTRICTION

POINT ...... 30

3.2 INVESTIGATING ZBED1 AND DLX FAMILY PROTEIN-PROTEIN INTERACTIONS

...... 32

3.3 TRANSCRIPTIONAL EFFECTS OF ZBED1 EXPRESSION AT THE RESTRICTION

POINT ...... 34

3.4 THE FUNCTIONAL EFFECTS OF ZBED1 AND DLX5 EXPRESSION ON CELL CYCLE

PROGRESSION ...... 36

3.5 FLOW CYTOMETRIC ANALYSES ...... 40

CHAPTER 4: DISCUSSION ...... 43

4.1: TRANSCRIPTIONAL REGULATION OF THE RESTRICTION POINT ...... 44

4.2: FUNCTIONAL EFFECTS OF DLX5 REGULATION ON CELLULAR PROLIFERATION

...... 46

4.3: ZBED1-DLX PROTEIN-PROTEIN INTERACTIONS – A CONSERVED MECHANISM

OF REGULATORY CONTROL ...... 48

4.4 CONCLUSION ...... 52

BIBLIOGRAPHY ...... 53

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List of Figures

Figure 1.1 Overview of the cell cycle and the three cell cycle checkpoints ...... 2

Figure 1.2 The Rb-E2F Pathway ...... 4

Figure 1.3 The c-Myc Pathway ...... 6

Figure 1.4 Rapid and prolonged DNA damage response pathways ...... 7

Figure 1.5 The organization of the human Dlx family ...... 12

Figure 3.1 No effect of Dlx5 expression at the restriction point in HEK293 cells ...... 31

Figure 3.2 Dlx6 and Zbed1 interact in vitro ...... 33

Figure 3.3 Neither Dlx5 nor Zbed1 alter transcription of a set of genes that regulate the restriction point in HeLa cells ...... 35

Figure 3.4 Dlx5 reduces EdU incorporation in DF-1 cells ...... 37

Figure 3.5 Forced Zbed1 expression does not prevent Dlx5’s anti-proliferative effects ...... 39

Figure 3.6 Dlx5 and Zbed1 do not significantly alter cell cycle profiles in HeLa cells ...... 42

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List of Tables

TABLE 2.1 PLASMIDS USED IN THIS STUDY ...... 20

TABLE 2.2 IMMUNOFLUORESCENCE DETECTION ...... 25

TABLE 2.3 PRIMERS FOR QUANTITATIVE PCR ANALYSIS ...... 28

ABBREVIATIONS

AER Apical ectodermal Ridge AKT Protein Kinase B Amp Ampicillin AP Alkaline phosphatase ATM ataxia-telangiectasia mutant ATR ataxia-telangiectasia BCA Bicinchoninic acid bp Base pair BMP2 Bone morphogenic protein 2 BRM Brahma C terminal Carboxy terminal CAMKII Calmodulin dependent kinase II CDC Cell division cycle CDK Cyclin dependent kinase Chk1 Checkpoint kinase 1 Chk2 Checkpoint kinase 2 CKI Cyclin dependent kinase inhibitors DEPC Diethyl pyrocarbonate DMEM Dulbecco’s modified Eagle’s medium dNTP Nucleouside triphosphate DPBS Dulbecco's phosphate-buffered saline DRE Drosophila DNA replication-related element DREF Drosophila DNA replication-related element binding factor Dll Distal-less Dlx Vertebrate Distal-less homeobox DTT Dithiothreitol EDTA Ethylenediaminetetraacetic acid

EdU 5-ethynyl-2’-deoxyuridine Exd Extradenticle FBS Fetal bovine serum FITC Fluorescein isothiocyanate G1 Gap 1 phase of cell cycle G2 Gap 2 phase of cell cycle G418 Geneticin GAPDH Glyceraldehyde 3-phosphate dehydrogenase GFP Green fluorescent protein GST Glutathione S-transferase HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid Hth Homothorax IRS-2 Insulin receptor substrate-2 kDa Kilodalton LB Lysogeny Broth M Mitosis phase of cell cycle MgCl2 Magnesium Chloride Msx Msh homeobox N terminal Amino terminal NaCl Sodium chloride NaF Sodium Fluoride NaPPi Sodium Phosphate Iodine ORC Origin recognition complex Osc Osteocalcin Osx Osterix PBS Phosphate buffered saline PBS-T Phosphate buffered saline with Tween 20 PCNA Proliferating cell nuclear antigen PEI Polyethylenemine

PI Propidium iodide PKA Protein Kinase A PKB Protein Kinase B PLC Phospholipase C PVDF Polyvinylidene fluoride Rb Retinoblastoma protein RT-qPCR Reverse-Transcriptase Quantitative polymerase chain reaction SDS-PAGE Sodium dodecyl sulfate–polyacrylamide gel electrophoresis SHFM Split-hand/split-foot malformation Zbed1 Zinc Finger BED-Type Containing 1 Zerknüllt Zen

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Chapter 1: Introduction

The Roles of Proliferation and Differentiation in Embryonic Development

Embryonic development is a highly-regulated process involving thousands of genes that control a multitude of cellular processes, including cellular growth, proliferation, and differentiation, cellular death and senescence (1-5). While rapid proliferation is a hallmark of early embryonic development, and the determination of specific cellular fates through differentiation is critical for this process, it is obvious that these two processes are incompatible with one another. In support of this conclusion, differentiating cell cultures are primarily characterized by a marked decrease in their proliferative potential, a phenomenon which has been demonstrated to be correlated with a lengthened cell cycle (6). Furthermore, many cell types (ranging from neuronal to epithelial cells) undergo terminal differentiation (7, 8). This process is associated with a permanent exit from the cell cycle, and therefore a complete cessation of cellular proliferation (9).

It has been demonstrated that a lengthened cell cycle – particularly during the G1 phase – is causally related to the onset of differentiation, and therefore differentiation may be dependent upon factors which inhibit cell cycle progression or promote cell cycle exit (10). Consequently, pro-differentiation signalling is associated with decreased cellular proliferation, while pro- proliferative signalling has also been associated with a decrease in differentiation (11, 12).

Lastly, it has been demonstrated that regulation of the proliferation/differentiation dichotomy is critical to the process of embryonic development, as the dysregulation of these processes often leads to lethal developmental defects and is associated with the onset of a multitude of pathological conditions, including cancer (13-15).

The G1/S Restriction Point

To proceed through the cell cycle and successfully divide, a eukaryotic cell must first overcome a series of checkpoints which occur between successive stages of the cell cycle. These checkpoints were first observed in Escherichia coli as a means to facilitate DNA damage repair

(16). Indeed, as vital mechanisms governing eukaryotic replication, it has been noted that components of these cell cycle checkpoints are highly conserved across species (17-22). Known as the G1/S phase checkpoint (or the restriction point in animals), the G2/M checkpoint, and the metaphase checkpoint, they serve as mechanisms by which a cell may respond to defects during the cell cycle (Figure 1.1) (23-31). Due to the causal relationship of an increase in the length of the G1 phase and the onset of differentiation, this review will focus primarily on the restriction point (6,10).

Figure 1.1 Overview of the cell cycle and the three cell cycle checkpoints. To divide, cells must overcome three cell cycle checkpoints. These occur at the boundary of the G1 and S phases (the restriction point), the boundary of the G2 and M phases (G2/M or G2 checkpoint) and during the M phase respectively (M phase or spindle checkpoint).

Throughout the G1 phase, members of the RB (retinoblastoma) pocket protein family associate with E2Fs, transcription factors associated with the regulation of a variety of pro- proliferative genes (32, 33, 34, 35). Rb itself binds to activator E2Fs which function to upregulate the transcriptional activity of target genes (32). This association acts to repress E2F activity and thus inhibit the transcription of E2F targets. Likewise, other members of the pocket protein family (p107, p130) associate with repressor E2Fs, effectively functioning as their nuclear localization signal, and therefore, allowing these proteins to repress pro-proliferative targets (usually the same targets regulated by activator E2Fs) (Figure 1.2A) (32, 33, 36).

As a response to upstream pro-proliferative factors, cyclin D-associated cyclin-dependent kinases 4 and 6 (cdk4/6), which are normally inhibited by the presence of the CDK inhibitor

(Cdki) p16 family members, phosphorylate these pocket proteins (37-39). This phosphorylation of Rb, p107 and p130 pocket proteins is associated with the shuttling of their associated repressor E2Fs into the cytoplasm (36). As the association between pRB and activator E2Fs is disrupted, these transcription factors can now bind pro-proliferative target genes (31-39). E2F activity leads to the upregulation of cyclin transcription and thus the upregulation of cyclin- dependent kinase 2 (Cdk2) activity, leading to increased phosphorylation of pocket proteins and the establishment of a positive feedback loop (Figure 1.2B) (40). Furthermore, activated cyclin

E-Cdk2 can stimulate its own activity by phosphorylating its inhibitor, the Cdki p27, leading to the degradation of this protein (41). Cyclin E-Cdk2 is thought to ultimately drive DNA replication and thus S phase entry through the phosphorylation and activation of Cdc45, leading to the formation of the preinitiation complex by Cdc45 loading onto chromatin (42). It has been demonstrated that Cdk2 activity is critical for cell cycle progression, as inhibition of Cdk2 by

Cdki proteins p18, p27 and p19 prevents S phase progression (42).

Figure 1.2 The Rb-E2F pathway. Green arrows denote mechanisms which upregulate transcription or activity while grey arrows or black arrows indicate repression of transcription or protein activity. Prohibition signs indicate repression of these events or the activity of a given protein. A) The inactive Rb-E2F pathway. CDK activity is repressed by associated CKIs. B) The activated Rb-E2F pathway. Pro-proliferative factors lead to the inactivation of p16, the buildup of cyclin and the upregulation of CDK activity, establishing a positive feedback loop with the activation of CDK2. The activation of CDK2 ultimately drives S-phase entry.

The c-Myc proto-oncogene can also drive S phase entry as a response to stimulation by mitogens (Figures 1.3A, 1.3B). c-Myc serves as a transcription factor that directly regulates transcription of cyclins, E2Fs, cyclin-dependent kinases, cyclin-dependent kinase inhibitors and

Cdc25a (43-45). By direct upregulation of Cyclin D2, c-Myc indirectly sequesters Cdk2 inhibitor p27 via Cyclin D-cdk4/6, stimulating cyclin E-Cdk2 activity and generating a positive feedback loop via pocket protein phosphorylation and p27 degradation (46).

The cell has several mechanisms to prevent S phase entry as a response to DNA damage

(Figure 1.4). These are known as DNA damage response checkpoint pathways which operate during the restriction point and S phase itself, which, depending on the DNA damage detected, can induce either a rapid and transient response or a sustained response to block cell cycle progression (47, 48, 49-52). In these pathways, DNA damage is detected by ataxia-telangiectasia

(ATR) and ataxia-telangiectasia mutant (ATM) kinases, which subsequently phosphorylate and activate downstream kinases, checkpoint kinases 1 and 2 (chk1 and chk2). In the rapid response pathway, chk1 and chk2 target cdc25a for ubiquitination and subsequent degradation (49-52).

Loss of cdc25a leads to the sustained phosphorylation of Cdk2, thus inhibiting its activity and preventing cell cycle entry (49-52). A more prolonged response is achieved by chk1- and chk2- mediated stabilization of p53, thereby preventing its degradation by mdm2 (53). The subsequent accumulation of p53 results in the upregulation of p21 transcription, and this protein subsequently accumulates and acts to inhibit cyclin E-Cdk2, thus preventing S-phase progression while the cell either commits to repairing the DNA damage or undergoes apoptosis (47, 48).

It is clear that the restriction point is a complex regulatory mechanism that controls cell cycle progression through various signalling pathways. Thus, it seems evident that the

5 checkpoint, and the competing pro-proliferative and pro-differentiative genes involved in its regulation, is an ideal locus for the control of the proliferation/differentiation dichotomy.

Figure 1.3 The c-Myc Pathway. Green arrows denote mechanisms which upregulate transcription or protein activity while grey arrows indicate repression of transcription or activity. Prohibition signs indicate that these activities are being prevented. A) The inactive c-Myc Pathway. While c-Myc transcription is inhibited, Cdk2 remains in its inactive state, and thus cell cycle progression cannot occur. B) The activated c-Myc pathway. In response to mitogen signalling, c-Myc is transcriptionally upregulated and in turn acts to upregulate various pro- proliferative targets at the restriction point, culminating in the activation of CDK2, the establishment of the positive feedback loop and S-phase entry.

Figure 1.4 Rapid and prolonged DNA damage response pathways. Green arrows indicate the upregulation of transcription or stability of a protein while red arrows indicate a repressive effect on target expression, protein stability, or protein activity. Red X’s denote the degradation of a protein. The detection of DNA damage leads to the activation of ATM or ATR, beginning a signalling cascade which culminates in the repression of CDK2 and prevention of S phase entry.

DREF/Zbed1, The “Master Key” To Cellular Proliferation

The DNA-replicated related element factor (DREF), was first discovered in Drosophila as a protein responsible for binding an 8 base pair 5'-TATCGATA consensus sequence dubbed the Drosophila DNA replication-related element (DRE) (54). DRE was first identified as a sequence necessary for the upregulation of genes encoding for proteins required for cellular proliferation, specifically, DNA Polymerase alpha and the Proliferating Cell Nuclear Antigen

(PCNA) and, as such, disruption of the DRE sequence led to drastic decreases in their expression

(54). Soon, over 153 copies of DRE were identified, upstream of over 61 genes related to cellular proliferation, leading researchers to postulate that DRE was an element commonly associated with the regulation of these genes, and therefore more broadly associated with the regulation of cellular proliferation (55).

Isolation of DREF cDNA revealed that the gene encodes for a transcription factor consisting of 701 amino acid residues and with a molecular weight of approximately 80 kDa, and subsequent experiments identified a DNA binding domain situated in the N-terminal domain

(amino acids 16-115 specifically) (56). DRE binding necessitates the formation of a homodimer via this N-terminal domain (57).

As previous research involving the DRE sequence suggested, it would later be demonstrated that DREF was responsible for the regulation of a plethora of genes required for cellular proliferation. In addition to DNA polymerase alpha, and PCNA, DREF has been demonstrated to play a key role in the regulation of the origin recognition complex (ORC), being directly implicated in the transcriptional control of two of the complex’s six subunits orc2 and orc5, thereby implicating it in the control of DNA replication initiation (58, 59). Additional studies revealing the transcriptional regulation of DNA primase, and RCF1 by DREF lend

8 further credence to this assertion (60, 61). In addition to its regulation of DNA replication initiation, DREF also plays a critical role in the broader regulation of transcription initiation. For instance, DREF is a transcriptional regulator of the TATA-box Binding Protein (TBP), which in yeast has been shown to be required for the function of all three RNA polymerases (62, 63).

A multitude of genes that DREF has been suggested to regulate are involved in cell cycle progression. Indeed, DREF has been implicated in the transcriptional regulation of Cyclin A and

E2F transcription factors, both of which are critical to overcoming the restriction point (64, 65).

Furthermore, DREF has been shown to indirectly regulate the expression of cdc25 through its transcriptional regulation of the osa and moira genes, that encode subunits of the BRM complex, which is involved in the remodelling of chromatin (66, 67, 68). Cdc25 is a critical phosphatase that is responsible for the activation of Cdks, and is thereby involved in overcoming cell cycle checkpoints and promoting cell cycle progression (69). Furthermore, DREF is necessary for the transcriptional activation of p53, which, when it accumulates, works to prevent cell cycle progression in order to stimulate DNA repair (70, 47,48). Taken together, these studies indicate that DREF plays a significant role in the control of cell cycle progression. Indeed, knockdown of

DREF indicates that it is required for cells to progress through the late G1 and S phases of the cell cycle (71).

Linking proliferation to differentiation, there is evidence to suggest that homeodomain proteins might achieve differentiation in part through downregulation of DREF or through the regulation of its downstream targets, leading researchers to suggest that DREF may be the

“master key” to cellular proliferation (72). For instance, the expression of the homeodomain protein Zerknüllt (Zen) downregulates the PCNA promoter (57). Down-regulation was demonstrated to be due to the decreased expression of DREF, a transcriptional target of Zen (57,

73). Similarly, researchers have observed that homeodomain proteins may also regulate pro- proliferative DREF targets directly, for instance, the downregulation of PCNA by the homeodomain protein, Cut, which, upon induction of differentiation, localizes to the PCNA promoter, where it functions as a repressor (74). Another homeodomain protein, Dll, has been shown to physically interact with DREF – (75). Through GST-pulldown assays using full length proteins and deletion derivatives, researchers determined that, Dll, via its N-terminal (amino acids 30-124) and C-terminal domains (amino acids 190 to 327) binds to the N-terminal DNA binding domain of DREF via its own N-terminus (amino acids 16 to 125) in vitro (75) This protein-protein interaction prevented DREF binding to the promoters of its pro-proliferative targets (75).

DREF orthologues, known as Zinc Finger BED-Type Containing 1 (Zbed1) have been identified in most vertebrate species and, although to date they have been poorly characterized, these genes and the proteins that they encode retain similarities with DREF that would suggest they may retain similar functions (76). However, these proteins also have a modest level of overall identity with DREF, with only 22% of the protein conserved between Drosophila DREF and human Zbed1 (77). Nevertheless, the human Zbed1 retains a DNA binding domain in its N- terminus, as well as dimerization activity owing to an ATC subdomain in its C-terminal region, which is necessary for DNA binding (78, 79). There are also DRE related sequences (5’-

TGTCG(C/T)GA(C/T)A) throughout the genome to facilitate transcription factor binding (80).

Indeed, research suggests that Zbed1 expression is induced in the G1-S phase of the cell cycle

(80).

Recent evidence has suggested that Zbed1 may not play such a vital role in cell cycle progression in vertebrates as its Drosophila homologue, as Zbed1 expression did not necessarily

10 overlap with that of Ki-67, a marker of cellular proliferation (76). In some tissues, Zbed1 expression extended beyond the expression of Ki-67, while in others, it appeared to be more restricted (76). While the researchers concluded that this indicated that Zbed1 was ‘not’ involved in cell proliferation, they indicated the possibility that the protein may act as a repressor of cellular differentiation (76). Therefore, despite the most recent information which puts Zbed1’s ability to regulate cell cycle progression into question, it is important to consider these vertebrate orthologues of DREF as possible binding partners for homeodomain proteins, such as those of the Dlx family.

Dlx Homeobox Genes

Homeobox genes were originally discovered in Drosophila, as researchers noted sequences of conserved DNA within the Antennapedia and Bithorax homeotic gene complexes

(81). These homeobox genes contained a conserved 180 bp region and encoded for a family of transcription factors characterized by their homeodomain – a domain of 60 amino acids that serves as the protein’s DNA binding domain. The homeobox was later demonstrated to be evolutionarily conserved across angiosperm, fungi, and metazoan species (81-83).

The vertebrate Distal-less homeobox (Dlx) genes represent a family of regulatory proteins which have critical functions throughout embryonic development. A total of six Dlx gene paralogues have been discovered in mammals, fish, birds and amphibians (84-91). First order cis-paralogous pairs reside on three different chromosomes, however there is little similarity between the encoded proteins, outside of the homeodomain (90, 92). The Dlx gene family has two identifiable clades across bi-gene clusters, with Dlx2, Dlx3 and Dlx5 forming a single clade and Dlx1, Dlx4 and Dlx6 forming another (90, 92) (Figure 1.5). Dlx genes are arranged in specific paralogous bi-gene clusters (specifically Dlx1-Dlx2, Dlx3-Dlx4 and Dlx5-

Dlx6), in which clustered pairs are convergently transcribed and display a level of functional redundancy (84, 89, 93-95).

Figure 1.5 The organization of the Dlx family in Homo sapiens. The Dlx family forms 3 paired bi-gene clusters (specifically Dlx1-Dlx2, Dlx3-Dlx4 and Dlx5-Dlx6) across chromosomes in which clustered pairs are convergently transcribed. The genes can be further organized into two distinct clades or subfamilies; Dlx2, Dlx3 and Dlx5 form one clade while Dlx1, Dlx4 and Dlx6 form another.

Throughout embryonic development, Dlx proteins are expressed in various tissues, including the forebrain and craniofacial primordia as well as in the limb buds, and Dlx expression is thought to be critical to the development of these tissues (95-99). Dlx1-/- and Dlx2-/- mice are characterized by abnormal morphogenesis of the first and second branchial arches, though this phenotype is more severe in Dlx1/2-/- double knockouts, suggesting the expression of either Dlx1 or Dlx2 may compensate for the loss of their paired gene (100). Targeted deletions of

Dlx3 in the neural crest cells of mice is associated with the impaired differentiation of odontoblasts and the formation of brittle teeth (101). While, Dlx5-/- murine embryos are characterized by impaired craniofacial development, owing to altered specification of the mandibular arch, endogenous Dlx5 expression also appears to be an important regulator of maxilla, mandible, and tooth specification in Homo sapiens (92, 102, 103).

Targeted knockdown of Dlx5 and Dlx6 expression has been shown to result in ossification defects in murine models, owing to the critical roles of these proteins in chondrocyte maturation and differentiation of osteoblasts (104). However, whereas Dlx5/6-/- mice embryos demonstrated reduced chondrocyte differentiation, this defect could be rescued by the overexpression of Dlx5 (105). Additionally, while Dlx5/6-/- mice embryos exhibited defects akin to split-hand/split-foot malformation (SHFM), this effect could be rescued by Dlx5 overexpression in the apical ectodermal ridge (AER) (99, 106). Taken together, these studies support the conclusion that Dlx genes exhibit functional redundancy across bi-gene clusters.

It is clear that Dlx homeodomain proteins play a vital role throughout the process of embryonic development and that their dysregulation is associated with developmental defects.

Therefore, investigating the means by which the Dlx family regulates differentiation is critical to understanding the mechanisms driving embryonic development.

Context Dependent Effects of Dlx5 on Cellular Proliferation

Dlx5 is expressed in cells and tissues that are differentiating (106-108). During development of the long bones of the limb, Dlx5 is expressed in both the hypertrophic region as well as in chondrocytes poised to differentiate, while its expression is absent in the proliferative zone (106-108). Furthermore, overexpression of Dlx5 is associated with an increase in the hypertrophic region of chondrocytes, resulting in decreased proliferative potential and the upregulation of chondrocyte differentiation in both murine and chick models, suggesting that

Dlx5 drives chondrocyte differentiation at the expense of their proliferative potential (106, 107,

109). In support of this suggestion, an increased rate of differentiation and a decrease in proliferative potential was demonstrated in vitro in chick fibroblasts with Dlx5 overexpression

(110). Furthermore, cultured Dlx5-/- osteoblasts were unable to maintain the same rates of proliferation and differentiation observed in Dlx5-expressing controls (110). Finally, dysregulation of chondrocyte maturation by Dlx5 is associated with developmental defects, as reduced ossification in long bones is seen in murine Dlx5-/- models, while Dlx5 overexpression in murine limbs results in shortened skeletal elements (107, 109).

Conversely, Dlx5 expression has also been associated with the upregulation of pro- proliferative targets, and high levels of endogenous Dlx5 expression is a common characteristic of cancerous human cell lines (111). To this effect, knockdown of Dlx5 expression in an ovarian cancer cell line resulted in attenuated IRS-2-AKT signalling, leading to decreased expression of cyclins A, B1, D1, D2, and E, and ultimately resulting in a marked decrease in proliferative potential (111). Furthermore, it has been demonstrated that Dlx5 is capable of binding to the promoter of the pro-proliferative c-Myc proto-oncogene, and knockdown of Dlx5 expression was correlated with a decrease in the expression of c-Myc and a decrease in proliferative potential

(112). Interestingly, endogenous expression of Dlx5 appears to have a positive role in cell proliferation in several developmental contexts, including during the development of the chick limb bud, in which Dlx5 expression in the AER specifies the continued proliferation of the underlying mesenchymal cells (113). Similarly, in murine models in which both Dlx5 and Dlx6 expression has been disrupted, inner ear defects arose that were subsequently linked to a decrease in cellular proliferation (114). Thus, it is apparent that Dlx5 is capable of both upregulation and downregulation of pro-proliferative targets, depending on cellular context, although the former scenario may be through indirect means.

The notion that Dlx5 activity may be dependent upon the context in which this protein is expressed is not without precedent. Context-dependent effects of Dlx5 expression have been demonstrated in its regulation of the Osteocalcin (Osc) gene, in which Dlx5 acts to either transcriptionally repress or stimulate Osc. In cancerous tissues, such as osteosarcoma cells, Dlx5 overexpression is associated with a reduction in the activity of the Osc promoter (115).

Conversely, in a murine osteoblast cell line, Dlx5 overexpression has been demonstrated to stimulate the transcription of Osc (110, 116). Thus, it is clear that the effects of Dlx5 may be context-dependent, and that activation or repression of Dlx5 target genes may be dependent upon post-translational modifications to this transcription factor, or alternatively the presence or absence of co-factors or protein partners (117).

Indeed, post-translational modifications have been shown to play a vital role in the transcriptional activity of Dlx5. In C2C12 cells, it was demonstrated that Dlx5 is a substrate of p38, which acts to phosphorylate Ser-34 and Ser-217 residues on Dlx5 (118). These modifications resulted in the increased expression of Osx, indicating that Ser-34 and Ser-217 phosphorylation increased the transactivation potential of Dlx5 (118). Furthermore, Dlx5

15 phosphorylation by Calmodulin dependent kinase II (CAMKII), Protein Kinase A (PKA) and

Protein Kinase B (PKB) is also correlated with an increase in transactivation potential of Dlx5 during osteoblast differentiation (119-121). Thus, it has been proposed that Dlx5 may be naturally unstable, and the modulation of the stability of this protein by protein kinases may play a vital role in its activity (121). However, post-translational modifications associated with the repression of pro-proliferative genes by Dlx5 have yet to be identified.

Protein-protein interactions may play an important role in the determination of Dlx5 activation or repression of target genes. Due to their wide range of targets, it has long been suggested that protein-protein interactions may regulate the transcriptional activity of homeodomain proteins. Indeed, a subset of homeodomain proteins, Hox, utilize protein partners such as Homothorax (Hth) and Extradenticle (Exd) to further regulate their binding specificity

(122). Few interactions with Dlx5 have been identified. However, one such interaction – the association of Dlx5 with the homeodomain protein, Msx – results in inhibition of Dlx5 DNA binding activity, and thus, transcriptional activity, for both transcription factors (123).

Furthermore, the Drosophila Dlx homologue, Distal-less (Dll), has been shown to associate with both Hth and Exd, resulting in the alteration of its transcriptional activity (124).

Researchers have attempted to characterize the mechanisms by which Dlx5 modulates proliferative potential in several murine, chick and human cell types. Transfection of proliferating C2C12 cells with vectors encoding Dlx5 or its cis-paralogue, Dlx6, resulted in a marked decrease in proliferative potential compared to vector controls (117). The same reduction in proliferative potential upon Dlx5 or Dlx6 transfection and subsequent overexpression was seen in the chick DF-1 cell line, in which homeodomain mutants unable to bind DNA were incapable of replicating this effect (117). Thus, the DNA binding activity of Dlx5 is necessary for

16 reduction of proliferative potential in these cell lines. However, introduction of the Dlx5 overexpressing vector to human embryonic kidney cells expressing the SV40 large T-antigen

(HEK293T) demonstrated no change in proliferative potential (117). This suggests that a Dlx5- mediated reduction in proliferation may depend on its effects on known SV40 large T-antigen targets, specifically the p53 tumour suppressor protein and Retinoblastoma protein (Rb), which are proteins associated with inhibition of cell cycle progression at the restriction point (125-131).

There is additional evidence to suggest that Dlx5 can modulate proliferation and differentiation by regulating the transcriptional activity of genes involved in the restriction point.

Flow cytometry assays conducted on C2C12 cells transfected and subsequently selected for overexpression of Dlx5 demonstrated a statistically significant increase in the proportion of cells remaining in the G1 phase of the cell cycle, thus implicating Dlx5 in the regulation of the restriction point (117). This data is consistent with previous studies that linked an increase in the length of the G1 phase with the onset of differentiation (6, 10). Interestingly, while previous studies had implicated overexpression of Dlx5 and its binding to the promoter of the c-Myc proto-oncogene with the upregulation of its transcriptional activity in Jurkat cells, and activation of a luciferase reporter in C2C12 and DF-1 cell lines, the same assay demonstrated transcriptional repression of the c-Myc promoter as a response to Dlx5 binding in HEK293T cells (112, 117). Taken together, these data strongly support the hypothesis that Dlx5 activity is context-dependent, and thus, the effects of this transcription factor must continue to be investigated in both pro-proliferative or differentiating cell lines to determine the mechanism by which Dlx5 can differentially regulate a diverse range of genes.

As Dlx5 is both a potential binding partner for Zbed1, and has previously been shown to regulate the transcriptional activity of several components of the restriction point, such as the c-

Myc proto-oncogene, investigating the effects of Dlx5 expression on the progression through this checkpoint is a promising endeavour which may help elucidate the mechanisms by which Dlx5 activity remains context-dependent (see figures 1.3A, 1.3B) (112, 117).

Rationale

The regulation of cell proliferation and differentiation is of critical importance throughout development and involves both common and lineage-specific regulations. While it is understood that homeobox genes play a very important role in the specification of tissue differentiation, in part through decreasing proliferative potential, the exact mechanisms by which they achieve this function remain largely unknown. To better understand these processes would give insight into how their aberrant regulation may lead to developmental defects, and pathological disease, and also, just as importantly, will expand our understanding of basic developmental processes. While the Drosophila homologue of Dlx proteins, Dll, and its effects on cell cycle progression are better understood, it is important to investigate whether or not these mechanisms are conserved, and how these processes might differ in more complex organisms. Here, we investigate the effects of Dlx5 on the restriction point, and examine whether Dlx binding to Zbed1 is a conserved mechanism of regulatory control of cell cycle progression. Our goal is to elucidate the exact mechanisms by which Dlx5 and Dlx6 regulate cell cycle progression.

Hypothesis

Dlx5 and Dlx6 mediate cell cycle progression through direct and indirect regulation of the restriction point, and that Zbed1-Dlx protein binding is a conserved mechanism for regulatory control of this important cell cycle checkpoint.

Chapter 2: Methods

Plasmids

Table 2.1 Plasmids used in this study

Descriptor Vector Experiments Source eGFP pCAGGS EdU Incorporation assay (132) c-Myc-tagged Dlx5 pcDNA3 EdU Incorporation assay (107)

(chicken) c-Myc-tagged Dlx5 pcDNA3 qPCR experiments, Flow (133)

(mouse) Cytometry

Vector pcDNA3 qPCR experiments, Flow (Invitrogen)

Cytometry

Flag-tagged Zbed1 pcFLAG GST-Pulldown assays, EdU This study

(chicken) incorporation assays, Flow

Cytometry, qPCR

Flag-tagged Zbed1 pcFLAG GST-Pulldown This study

(N-term) (chicken)

GST pGEX-2T GST-Pulldown (Pharmacia)

GST-tagged Dlx6 pGEX-2T GST-Pulldown C. Paradiso & A. Bendall, unpublished.

(mouse)

The chicken Zbed1 coding sequence was amplified from cDNA synthesized from whole chicken embryos at Hamburger and Hamilton stage 27-28 and cloned into pcFLAG as an Xba1-

BamH1 fragment in order to place a FLAG epitope tag at the amino-terminus of the encoded

20 protein. Two PCR artefacts that led to missense codons were subsequently corrected by site- directed mutagenesis. In the course of correcting those errors a clone was derived with a frame shift and premature stop codon downsream of codon 122. This clone was used to express the amino-terminal (DNA-binding) domain of Zbed1. pGEX2T-Dlx6 contained the murine Dlx6 coding sequence as an in-frame BamHI-SmaI fragment downstream of GST. pGEX-Protein Induction

E. coli BL21 (DE3) cells were transformed with the parental pGEX plasmid or pGEX-

Dlx6 and plated on 100 µg/mL ampicillin (Amp) LB agar plates. Following overnight incubation at 37°C, colonies were grown overnight in a shaking incubator in a 5 mL culture with 100 µg/mL ampicillin. The following morning, 45 mL of LB amp was added to the culture. Cultures were grown in a shaking incubator at 37°C until log phase (OD600=0.5~0.8). Once the density was within this range, IPTG was added to a concentration of 1 mM to induce protein expression and cells were incubated for a further 3 hours at 37°C. Following incubation, cells were pelleted at

6,000 rpm and resuspended in 1 mL of PLC+ buffer (10% glycerol, 50mM Hepes pH 7.5, 150 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA, 10 mM NaPPi, 100 mM NaF, 1% Triton X-100, 10 ug/ml Aproptinin, 10 ug/ml Leupopeptin, 0.1 mM Ortho Vanadate, 0.1 mM PMSF). Cells were then sonicated with three thirty-second pulses on ice and centrifuged at 13,000 rpm for 10 minutes at 4°C.

Glutathione Bead Preparation

250 µL of Glutathione Sepharose 4B beads (GE Healthcare) were aliquoted and centrifuged at 3,000 rpm for 30 seconds. After removal of supernatant via aspiration, beads were washed 2x in 500 uL PBS and 1x in PLC+ lysis buffer. Following dilution to 1 mL total volume in PLC+ buffer, beads were resuspended by vortexing and incubated on a nutator with extracts

21 from the bacteria induced to express Dlx6-GST at 4°C for 3 hours. Following incubation, the sepharose beads were pelleted at 3,000 rpm at 4°C for 1 minute and washed 3x in 1 mL of PLC buffer (10% glycerol, 50 mM HEPES pH 7.5, 150 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA, 10 mM NaPPi, 100 mM NaF, 1% Triton X-100). Washed beads were resuspended in 100 µl of PLC buffer + 1 mM DTT. To confirm protein expression, 20 µL of bead slurry was added to 20 µL of

2x SDS, boiled for 2 minutes, and pelleted at 3,000 rpm for 1 minute. Samples were loaded onto an SDS-PAGE gel, which was then stained with Coomassie blue stain (0.5 g Coomassie, 500 ml methanol, 100 mL acetic acid, 400 mL H2O) for 1 hour, and washed with destaining solution

(7% acetic acid, 5% methanol, in H2O) overnight to visualize the separated proteins.

GST-Pulldown

Twenty-four hours post transfection DF-1 cells were collected and lysed in PLC+ buffer, and sonicated 3x. Cell lysates were then mixed with bead slurries and incubated on a nutator overnight at 4°C. Samples were then washed in PBS buffer 3x with vortexing before being resuspended in 100 µl of PLC+ buffer. To elute bound proteins, 20 µL of bead slurry was added to 20 µL of 2x SDS, boiled for 2 minutes, and pelleted at 3,000 rpm for 1 minute. 20 µL of sample supernatant per pulldown was then separated by SDS-PAGE.

Cell Lines and Reagents

Human Embryonic Kidney (HEK293), HeLa (originally derived from human cervical carcinoma) and chicken fibroblast (DF-1) cell lines were cultured in Dulbecco’s modified

Eagle’s medium (DMEM; Thermo Scientific), supplemented with 10% fetal bovine serum (FBS;

VWR), 100 U/mL penicillin, 100 μg/mL streptomycin, and 2 mM L-glutamine (Thermo

Scientific). Cell cultures were maintained at 37°C and at 5% CO2.

Transfection and Selection

Cells were seeded 24 hours prior to transfection, at which point they were transfected at

60-70% confluency using polyethylenimine (PEI) at a ratio of 4.5 μg PEI per μg of plasmid

DNA. To prevent cytotoxic stress, transfected DF-1 and HeLa populations were treated with new media 4 and 6 hours post-transfection respectively. For HeLa cells transfected with either pcDNA3 or pcFLAG, selection was achieved via incubation with 500 µg/mL Geneticin (G418)

(Bioshop) beginning at 1 day after transfection for a period of 4 days alongside an un-transfected control dish to ensure lethality in cells lacking the vector.

Coverslip preparation

Glass coverslips were sterilized by dipping in 70% ethanol and passing through a flame.

Following two washes with phosphate buffered saline PBS, slides were coated with 0.1 M poly-

D-lysine for 30 minutes, washed twice with sterile deionized water and allowed to dry in a biosafety cell culture hood for two hours. Once dry, slides were washed once more with DMEM before cells were seeded onto the coverslips.

EdU Incorporation Assay

DF-1 cells were seeded directly onto poly-D-lysine coated coverslips and allowed to settle for 24 hours before transfection. Twenty-four hours post-transfection, cells were incubated for 4 hours with 10 µM EdU in DMEM at 37°C and 5% CO2 and fixed with 3.7% formaldehyde

PBS for 15 minutes at room temperature. Fixed cells were then permeabilized using 0.5% Triton-

X-100 in PBS for 20 minutes at room temperature. The Click-iT® EdU Alexa Fluor® 594

Imaging Kit (Life Technologies) was used to detect EdU incorporation as per the manufacturer’s instructions.

Indirect Immunofluorescence

Following the Click-iT® reaction, coverslips were washed twice with PBS supplemented with 10 mM glycine (PBS-G). Mouse primary antibodies were diluted to 1:50 with 3% bovine serum albumin (BSA) in PBS-G. c-Myc-tagged Dlx proteins were detected with a 9E10 α-MYC antibody (Developmental Hybridoma Bank), while FLAG-tagged Zbed1 proteins were detected with a monoclonal anti-FLAG M2 antibody (Sigma Aldritch) or a rabbit polyclonal anti-

DYKDDDDK antibody (Invitrogen)) diluted 1:50 or 1:1,000 in 3% BSA in PBS-G respectively.

Coverslips were treated for 2 hours at room temperature in the dark. Following primary antibody incubation, coverslips were washed 3x with PBS-G and incubated for 45 minutes with secondary

FITC-conjugated goat anti-mouse antibody (Genetex) diluted 1:100 with 3% BSA in PBS-G, or secondary AlexaFluor 647 goat anti-rabbit antibody (Invitrogen) diluted 1:500 with 3% BSA in

PBS-G. Following 3x washes with PBS-G, coverslips were incubated in 5 µg/mL Hoescht 33342 in PBS for 30 minutes in the dark at room temperature. Coverslips were washed an additional 2x with PBS and 1x with sterile deionized water and allowed to dry before mounting in a sterile

50% glycerol solution. Slides were imaged with a Leica DM IRE2 inverted fluorescence microscope. Untransfected DF-1 cells were used as a control for exposure levels for each experiment, to exclude the possibility of false positives. To ensure there was no bleed-through between different targets, single treatments for DAPI, Alexafluor-594, FITC or Alexafluor-647 were imaged at all four wavelengths. Nevertheless, as the same cube was used to detect

Alexafluor-594 and Alexafluor-647, the extra precaution of examining specimens positive for both on the confocal microscope was done at the Molecular and Cellular Imaging Facility at the

University of Guelph.

Table 2.2 Immunofluorescence Detection

Target Detection Detection Microscope Cubes

Controls

GFP Untransfected N/A Leica DM IRE2 GFP (470/40 nm)

EdU No EdU Alexafluor-594 Leica DM IRE2 Y5 (620/60 nm)

DAPI Untreated Hoescht Leica DM IRE2 A4 (360/40 nm)

Dlx5 Untransfected FITC Leica DM IRE2 GFP (470/40 nm)

Zbed1 Untransfected FITC Leica DM IRE2 GFP (470/40 nm)

Dlx5+Zbed1 Untransfected FITC, Leica DM IRE2 IRE2: GFP, Y5

(EdU +/-) Alexafluor-647 Confocal: Leica DM

6000B

Flow Cytometry

Selected HeLa cell populations were seeded at equal densities in 6 cm dishes. Twenty- four hours post-seeding, cell populations were synchronized by treatment with 100 ng/ml nocodazole (Sigma-Aldritch) in DMEM for 24 hours. To release the cell synchronization, cells were washed thrice with PBS and re-plated in a new 6 cm dish. One dish of each synchronized population was collected at this release time (0 hours) and further dishes were collected at 12 hours, 18 hours, and 24 hours post-release. Transfected cells used as asynchronous controls were also collected at the 0 hour time-point for flow cytometry. Cells were centrifuged at 1,000 xg and resuspended at a concentration of 1x106 cells/mL in DPBS (Dulbecco's phosphate-buffered saline). Cells were centrifuged again at 1,000 xg, resuspended in PBS and fixed by dropwise

26 addition of chilled 70% ethanol with gentle vortexing. Cells were then incubated at -20°C for at least 24 hours and then pelleted at 3,000 xg. Pellets were washed 3x in PBS and then resuspended in 50 μL of 100 μg/mL RNase A (Sigma Aldritch) per 1x106 cells prior to 10 minutes of incubation at room temperature and the addition of 400 µL of 50 µg/mL propidium iodide (PI) (Bioshop Canada Inc) in PBS (per 1x106 cells), and an additional 30 minute incubation in a water bath, in the dark, at 37°C. DNA content was determined using a Becton

Dickinson FACSCalibur flow cytometer at excitation 370 nm and emission of 488nm. Fl2 histograms were analyzed using Flowploidy package in R software (134).

Immunoblotting

Cells were removed from their culture dish via cell scraping and resuspended in chilled

PBS. Samples were pelleted and resuspended in 1x Laemmli buffer (10% glycerol, 2% SDS,

0.05 M Tris-HCl pH 6.8) containing protease inhibitors (Complete Mini Protease Inhibitor;

Roche) before sonication. Samples were boiled at 100°C for 5 minutes in a heat block and protein concentrations were quantified with a BCA protein assay kit (Pierce). Following equalization of protein concentration to 4 µg/ul, 40 µg of each protein sample was separated by

SDS-PAGE on a 13% poly-acrylamide gel. Proteins were then transferred onto PVDF membrane

(pre-wet in methanol) at 300 mV, for 1 hour at 4°C in circulating transfer buffer (25mM Tris,

192mM glycine, 20% methanol in H20). Membranes were subsequently blocked in 5% skim milk powder in PBS-T (PBS containing 0.1% Tween-20). Membranes were incubated with primary antibodies in blocking solution plus 0.05% sodium azide overnight; to detect Flag- tagged proteins, a primary monoclonal anti-flag M2 mouse antibody was used at a dilution of

1:7,000, while to detect c-Myc-tagged proteins, a primary mouse α-MYC antibody (9E10) was used at a dilution of 1:1,000.

The next morning, following one 15 minute and 3x 5 minute washes in PBS-T, membranes were incubated with 1:10,000 mouse horseradish peroxidase conjugated secondary antibody in blocking solution for a period of 1 hour. Following one 15 minute and 3x 5 minute washes in PBS-T, membranes were imaged with a Bio-Rad molecular Imager Chemi Doc XRS+ and ImageLab software (Bio-Rad), using Western Lighting Chemiluminescence Reagent Plus

(Perkin Elmer Life Sciences) in order to detect protein bands.

Quantitative Polymerase Chain Reaction

RNA Isolation: Transiently transfected HEK293 cells were collected 24 or 48 hours post- transfection using a cell scraper and resuspended in 1 mL of chilled PBS. G418-selected populations of HeLa cells were seeded at one million cells per 6 cm dish and similarly collected

24 hours later. Following pelleting at 9,000 xg for 10 minutes, samples were solubilized in 3 mL

TRizol for 10 minutes at room temperature before adding 0.6 mL of chloroform and vortexed, followed by incubation for 5 minutes at room temperature. Cells were then pelleted at 4°C for

9,000 rpm for 15 minutes and the upper phase was transferred to a new tube. Isopropanol (1.5 mL) was added, mixed by inversion and incubated for 10 minutes before centrifugation at 9,000 rpm for 15 minutes at 4°C. After removal of the supernatant, pellets were dissolved in 20 µL of

DEPC-treated H2O. Genomic DNA was removed from each sample using the RapidOut DNA

Removal Kit (Thermo Fischer) as per the manufacturers’ instructions. For reverse transcription of the RNA, 1 µL of 500 µg/mL Oligo(dT) and 1 µL 10 mM dNTP mix was added to 1 µg of

RNA plus sterile distilled water to a final volume of 12 µL. The mixture was heated to 65°C in a water bath and then quickly chilled on ice before the addition of 4 µL 5x First-Strand Buffer, and

2 µL 0.1M DTT. The solution was incubated at 42°C for 2 minutes and then 1 µL (200 units) of

Superscript II reverse transcriptase was added to the mixture. Following 50 minutes of

incubation at 42°C, the enzyme was inactivated at 72°C for 15 minutes. qPCR experiments were

performed using the StepOnePlus real time PCR-instrument at the University of Guelph

Advanced Analysis Centre. The expression of the housekeeping gene, GAPDH, was used to

standardize Ct values across samples. A full list of primers used is shown in Table 2.3.

Table 2.3 Primers for quantitative PCR analysis

Target Forward Primer (5’ ---- 3’) Product Length Used previously Reverse Primer (5’ ---- 3’)

MYC Fw: AATGAAAAGGCCCCCAAGGTAGTTATCC 122bp (135) Rv: GTCGTTTCCGCAACAAGTCCTCTTC

CCND1 Fw: ACAAACAGATCATCCGCAAACAC 144 bp (136) (Cyclin D1) Rv: TGTTGGGGCTCCTCAGGTTC

CCND2 Fw: CTGTGTGCCACCGACTTTAAGTT 75 bp (137) (Cyclin D2) Rv: GATGGCTGCTCCCACACTTC

CCNE1 Fw: AGCCAGCCTTGGGACAATAAT 134 bp (138) (Cyclin E1) Rv: GAGCCTCTGGATGGTGCAAT

EBF1 Fw: CAGCCCGCCTTGATCTTCTA 200 bp No Rv: CTTCCACTCCGTTGGATGCT Zbed1 Fw: GAAGACCCCCTCAAGTGGTG 119 bp No Rv: GATCCGAAGAGACGCTCAGG p21 Fw: GCAGACCAGCATGACAGATTT 70 bp (139)

Rv: GGATTAGGGCTTCCTCTTGGA p27 Fw: CAAATGCCGGTTCTGTGGAG 177 bp (140) Rv: TCCATTCCATGAAGTCAGCGATA p53 Fw: TAACAGTTCCTGCATGGGCGGC 129 bp (141)

Rv: GATCCGAAGAGACGCTCAGG

GAPDH Fw: AATCCCATCACCATCTTCCA 82 bp (142)

Rv: TGGACTCCACGACGTACTCA

Statistical Analysis For EdU incorporation assays, labeled and unlabeled cells were counted manually and recorded in MS Excel. Fold changes for the RT-qPCR results were calculated from the delta delta CT values. Statistical analysis for both experiments was conducted by determining the standard deviation and performing a one tailed T-test. For flow cytometric data, the R package

(134), Flowploidy, was used to analyze cell cycle profiles, with standard deviation and two tailed

T-tests used to determine significance. A value of P<0.05 was considered significant.

Chapter 3: Results

3.1 The Transcriptional Effects of Dlx5 Expression at the Restriction Point

Previous experiments in our laboratory have indicated that Dlx5 and Dlx6 expression results in a lengthening of the G1 phase of the cell cycle (117). However, the exact mechanism by which Dlx5 and Dlx6 decrease the proliferative potential of the cells in which they are expressed remains largely unknown.

The misexpression of Dlx5 appears to drive proliferation in various cancerous cell lines, indicating various regulatory targets operating at the restriction point (111, 112). In ovarian cancer cells, knockdown of Dlx5 results in the decreased expression of the cyclins A, B1, D1,

D2, and E due to an attenuated IRS-2-AKT signalling pathway, while in lung cancer cells the knockdown of Dlx5 resulted in decreased expression of the c-Myc proto-oncogene (111, 112).

However, Dlx5 is known to have opposing effects in developmental and cancerous contexts, the best example of which is its regulation of Osc, where in osteoblast sarcoma cells, the overexpression of Dlx5 results in the decreased activity of the Osc promoter, whereas the gene is a positive regulator of Osc transcription in murine osteoblasts (115, 110, 116).

To determine the broader effects of Dlx5 expression on the restriction point, nine genes of interest were identified, including both positive (c-Myc, Ebf1, and the cyclins E1, D1, and D2) and negative (p21, p27, p53) regulators of this control point, as well as Zbed1. HEK293 cells were transfected with empty pcDNA3 vector or pcDNA3-Dlx5. RNA samples were collected 24 hours or 48 hours post-transfection, and the transcript levels were quantified via RT-qPCR

(Figures 3.1A, 3.1B). The overexpression of Dlx5 did not have any notable impact on the expression of any of the genes analyzed.

Figure 3.1 Effect of Dlx5 misexpression at the restriction point in transiently-transfected HEK293 cells. A) Expression of pro-proliferative (c-Myc, EBF1, Zbed1, and Cyclin E1, Cyclin D1 and Cyclin D2), and anti-proliferative (p21, p27, p53) restriction point genes in HEK293 cells collected 24 hours (A) or 48 hours (B) post-transfection with pcDNA3-Dlx5 compared to vector controls. Data are normalized against GAPDH. Exogenous Dlx5 was detected at levels averaging 600-fold higher than in vector controls (not shown).

3.2 Investigating Zbed1 and Dlx family protein-protein interactions

As the overexpression of Dlx5 did not significantly alter transcription at the restriction point in HEK293 cells, it became evident that the Dlx family may not cause reduced proliferative potential through the regulation of this cell cycle checkpoint. Therefore, we identified possible protein-protein binding partners which could explain this phenomenon.

As the Drosophila homologue of the Dlx family, Dll, has been demonstrated to interact with the pro-proliferative transcription factor DREF in vitro (75), we selected the vertebrate homologue of DREF, Zbed1, as a candidate for interactions with Dlx family proteins. In

Drosophila, Dll interacts specifically with the DREF N-terminal region, thus preventing its DNA binding, and thereby acting to negatively control the transcriptional effects of this protein (75).

As DREF has previously been demonstrated to be vital to progress through the restriction point and into the S-phase of the cell cycle, a similar interaction between Dlx proteins and the vertebrate homologue of DREF, Zbed1, could explain how Dlx proteins achieve the lengthening of the G1 phase necessary to induce differentiation (6, 10, 71, 117).

To investigate whether Zbed1 and Dlx protein interactions represented a conserved mechanism of regulation, we examined interactions between chicken Zbed1 and mouse Dlx6 via

GST-pulldown assay. GST or GST-Dlx6 was purified from E.coli and immobilized on

Glutathione-Sepharose beads and incubated with the cell lysates of transfected DF-1 cells. GST-

Dlx6 pulled down the full length Zbed1 protein, while GST did not (Figure 3.2A). We also observed a specific interaction between Dlx6-GST and the 13kDa N-terminal region of Zbed1 in one experiment (Figure 3.2B) however attempts to repeat this result were inconclusive as the N- terminal region repeatedly stuck to the GST beads as well (data not shown). Therefore, while we

33 have not yet mapped the interaction domain of Dlx6, these results do demonstrate that Dlx6 interacts with Zbed1 in vitro.

Figure 3.2 Dlx6 and Zbed1 interact in vitro. GST pulldown assay using GST-Dlx6 and the cell lysates of DF-1 cells transfected with pcFLAG-Zbed1 (A) or pcFLAG-Zbed1(N), encoding a 13 kDa Zbed1 N-terminal domain (B). GST (lane 2) or GST-Dlx6 (lane 3) were immobilized on sepharose beads and incubated with the cell lysates. 1% of the cell lysate was used to visualize the input protein.

3.3 Transcriptional effects of Zbed1 expression at the restriction point

The Drosophila homologue of Zbed1, DREF, has been demonstrated to transcriptionally regulate several genes at the restriction point, such as the E2F transcription factors and Cyclin A

(64, 65). We examined the effects of forced Zbed1 expression upon a subset of our genes of interest in HeLa cells. HeLa cells were transfected with empty pcDNA3, pcDNA3-c-Myc-Dlx5, or pcFlag-Zbed1, and selected for four days in G418 before samples were collected and quantified by RT-qPCR. Neither Zbed1 nor Dlx5 expressing populations demonstrated significant changes in transcriptional profile when compared to the vector control (Figure 3.3A).

As the primers used for Zbed1 in this experiment were designed for the human sequence, and did not recognize the exogenous chick Zbed1 transcript, protein expression was verified by western blotting (Figure 3.3B).

Figure 3.3 Neither Dlx5 nor Zbed1 alter transcription of a set of restriction point regulatory genes in drug-selected populations of HeLa cells. A) The relative expression of pro- proliferative (c-Myc, Zbed1, and Cyclins E1 and D1) and anti-proliferative (p21, p27, p53) restriction point genes in HeLa cells. HeLa cells transfected with pcDNA3, pcDNA3-Dlx5 or pcFlag-Zbed1 were selected for G418 resistance compared to empty vector controls. Data are normalized against GAPDH. B) Verification of exogenous protein expression by SDS-PAGE and western blotting. Selected cell populations were seeded into 6 cm dishes and collected alongside RNA samples for each biological replicate. Blots in the lower panel have been cropped to remove non-relevant lanes.

3.4 The functional effects of Zbed1 and Dlx5 expression on cell cycle progression

With a physical interaction identified between the Dlx6 and Zbed1, we investigated the functional consequences of such an interaction on cell cycle progression using EdU incorporation assays.

A thymine analogue, EdU can be used to identify cells which have progressed into the S- phase of the cell cycle, as cells incubated with EdU will incorporate the analogue into newly synthesized DNA. The incorporation of an alkaline-conjugated EdU analogue could be detected by immunofluorescence microscopy using an azide-conjugated AlexaFluor dye and copper catalyzed CLICK reaction (143).

DF-1 cells were seeded onto coverslips and then transfected 24 hours later. Twenty-four hours post-transfection, DF-1 cells were incubated with EdU for 4 hours. Cells positive for Dlx5 or Zbed1 were detected with anti-myc 9E10 and anti-FLAG M2 primary antibodies respectively, and visualized via a FITC-conjugated secondary antibody, while nuclei were stained using

Hoescht 33342. To ensure that the effects of these proteins on cell cycle progression were not a result of stress from the transfection process, cells transfected with GFP were used as a control.

To determine the effects of Dlx5, Zbed1 or GFP on S-phase entry, we compared the proportion of GFP-, Dlx5-, or Zbed1-expressing cells that had incorporated EdU with untransfected cells from the same slide (Figure 3.4A, 3.4B). While GFP expression was not associated with a significant alteration in the percent EdU incorporation, consistent with previous experiments in other cell lines, Dlx5 expression resulted in a significant decrease (t test < 0.05) in EdU-positive cells, decreasing from 53% in Dlx5-negative DF-1 cells to 10.6%. Consistent with recent evidence suggesting that Zbed1 does not stimulate cellular proliferation in vertebrates, no

37 significant increase in EdU incorporation was demonstrated between Zbed1-positive and Zbed1- negative DF-1 cells.

Figure 3.4 Dlx5 reduces EdU incorporation in transiently-transfected DF-1 cells. DF-1 cells seeded onto poly-D-Lysine coated coverslips and transfected 24 hours later. Twenty-four hours post-transfection cells were incubated for 4 hours in EdU. A) Representative images of transfected cells. B) Average EdU incorporation in transfected and un-transfected populations across at least 3 experiments for each expressed protein. Original magnification 10x. *p<0.05 (unpaired t-test)

Next, DF-1 cells were co-transfected as described above and incubated for 4 hours in

EdU. Zbed1 was detected using a rabbit anti-FLAG primary antibody and an AlexaFluor 647- conjugated anti-rabbit secondary antibody, while Myc-tagged Dlx5 was detected with an anti-c-

Myc 9E10 mouse monoclonal primary antibody and a FITC-conjugated anti-mouse secondary antibody. DF-1 nuclei were stained with Hoescht 33342 and the percentage of EdU positive, dual-expressing cells, were compared to the percentage of EdU-positive double-negative cells

(Figure 3.5A). Interestingly, a marked decrease in EdU incorporation was witnessed in cells co- expressing these proteins, with 37% of unlabeled cells incorporating EdU, while only 5% of labeled Dlx5/Zbed1-expressing cells incorporated EdU (Figure 3.5B, p < 0.05). There were insufficient numbers of single-transfected cells in these experiments to analyse separately. Thus, forced expression of Zbed1 did not prevent the negative effects of Dlx5 expression on cell cycle progression.

Figure 3.5 Forced Zbed1 expression does not prevent the anti-proliferative effects of Dlx5. DF-1 cells were seeded onto poly-D-Lysine coated coverslips and transfected 24h later. Twenty- four hours post-transfection cells were incubated for 4 hours in EdU. A) A representative image of co-transfected cells taken at an original magnification of 20x. B) Average EdU incorporation in co-expressing and double-negative populations. *p<0.05 (unpaired t-test).

3.5 Flow Cytometric Analyses

To better understand the effects of Zbed1 and Dlx5 expression on cell cycle progression, the cell cycle profiles of asynchronous and synchronous populations of HeLa cells expressing these proteins was examined by flow cytometry. 24 hours after transfecting HeLa cell populations with either pcDNA3, pcDNA3-Dlx5 or pcFlag-Zbed1, cells were selected with G418 over the course of four days. Prior to selection, cell populations were seeded at equal densities in seven 6 cm plates. Synchronization of cell populations at the G2-M checkpoint was achieved by treatment with the mitotic-spindle poison, nocodazole, for a period of 24 hours (144). Along with an asynchronous control, one dish of nocodazole-treated cells was collected at the release point, and every 6 hours thereafter up to 24 hours post-release, in order to track the progression of these cell populations through the cell cycle. Samples were fixed in ethanol, treated with RNase, and incubated with DNA intercalating dye, propidium iodide. Following incubation, the cell cycle profiles of each sample were investigated using a Becton Dickinson FACSCalibur flow cytometer. Transfected and selected cells were also collected at the 0 hour time-point for verification of exogenous protein expression.

In the asynchronous samples, no significant differences were detected in the proportion of cells in the G1, S, or G2 phases of the cell cycle in either Zbed1 or Dlx5 transfected populations when compared to the control cells (Figure 3.6A). Furthermore, while nocodazole treatment successfully inhibited passage through the G2-M checkpoint, thus synchronizing cell populations in the G2 phase, no significant difference was observed between synchronized populations at release, indicating that these populations did not react differently to the application of the drug (Figure 3.6B). However, post-release, sample collection became markedly more difficult, culminating in a failure to collect adequate cell numbers at the 6 hour

41 timepoint. Like the asynchronous controls, there were no significant differences in Zbed1- expressing, Dlx5-expressing, or empty vector transfected samples at 12, 18, or 24 hours post- nocodazole release (Figures 3.6C-E). It should be noted that, between replicates, there was a large degree of variation, particularly in the 12 and 24 hour timepoints, which could potentially have masked any protein-induced effects upon the cell cycle, however, as HeLa cells progress through the cell cycle in roughly 18-24 hours (145) and most cells had already passed through

G1 at the 12 hour timepoint, this seems unlikely. To ensure that the relevant proteins were expressed, one plate of unsynchronized cells was collected at the release time and used to verify protein expression via SDS-PAGE (3.6F).

Figure 3.6 Dlx5 and Zbed1 do not significantly alter cell cycle profiles in HeLa cells. HeLa cells were transfected with pcDNA3 vector, pcDNA3-Dlx5 or pcFlag-Zbed1 and selected in G418 for four days. Following selection, HeLa cells were seeded at equal densities in seven 6 cm plates. 24 hours post seeding, five plates were treated with 100 ng/mL nocodazole for 24 hours. HeLa cells were released from nocodazole by re-plating after 3x PBS wash. At release, both control (A, n=4) and nocodozole-treated (B, n=3) samples were collected. Following release, further samples were collected at 12 hours (C, n=3), 18 hours (D, n=2) or 24 hours (E, n=2). Cell populations were examined after propidium iodide staining using a Becton Dickinson FACSCalibur flow cytometer. Histograms were analyzed using the R Polyploidy package. (F) Verification of exogenous protein expression by SDS-PAGE and western blotting. Selected asynchronous cell populations were seeded into 6 cm dishes and collected alongside control samples.

Chapter 4: Discussion

Members of the Dlx family of homeobox genes encode for transcription factors critical throughout the process of embryonic development. During this time, Dlx family members are expressed in the forebrain, craniofacial primordia, and the limb buds of the developing embryo, and are vital for the specification and development of these tissues (95-99). Arranged into three separate bi-gene clusters, the Dlx gene pairs are transcribed convergently, and show functional redundancy within linked gene pairs, despite sharing little amino acid sequence conservation outside of the homeodomain (84, 89-90, 92-95).

Recently, it has been demonstrated that the expression of Dlx5 and Dlx6 results in a corresponding reduction in proliferative potential of various cell lines, including C2C12, DF-1,

HEK293 and HeLa cells, indicating that Dlx family members may play a more direct role in antagonizing proliferation as well as up-regulating differentiation markers (117). Reduced proliferative capacity in these cells was subsequently demonstrated to occur as a result of a lengthening of the G1 phase (117). It has long been demonstrated that the onset of differentiation is causally associated with an increased G1 phase and corresponding decrease in proliferative potential (10). Therefore, it seems apparent that the pro-differentiation effects of the Dlx family members involves their ability to reduce proliferative potential.

The dysregulation of the proliferation/differentiation balance is a causal factor in various forms of disease. Many cancers are associated with high rates of cellular proliferation achieved via the ablation or suppression of normal differentiation pathways – for example, the development of osteosarcoma is associated with underlying defects in osteogenic differentiation

(14, 15). However, modulation of this dichotomy can also serve to combat these diseases –

44 indeed, “differentiative” therapy has long been used as a powerful tool in treating cancerous tissues (146). For instance, upon treatment with the proliferation activated receptor agonist, troglitazone, it has been demonstrated that various osteosarcoma cell lines begin to undergo differentiation, coupled with a marked decrease in the proliferative capacity of these cells, as well as cell viability (146). Thus, a greater understanding of the regulatory control of proliferation and differentiation may provide novel therapeutic targets.

In this thesis, we attempted to determine by what mechanisms Dlx5 regulates cell cycle progression. Furthermore, we examined whether a previously observed protein-protein interaction between DREF and Dll in Drosophila was a conserved mechanism for the regulatory control of this process in vertebrates.

Transcriptional Regulation of the Restriction Point

Previous studies in our laboratory have demonstrated decreased proliferative potential in

HEK293 and HeLa cells transfected with Dlx5- or Dlx6-encoding plasmids when compared to vector controls (117). Furthermore, flow cytometric analyses of asymmetric populations of

C2C12 cells indicated that Dlx5 or Dlx6 expression culminated in a lengthening of the G1 phase of the cell cycle (117). As a lengthened G1 is associated with the onset of differentiation, it is possible that the antagonism of cell cycle progression by Dlx family members is important for their differentiation function.

In this thesis we attempted to characterize how Dlx5 might achieve the lengthening of the

G1 phase seen in these cell types. In cancerous tissues, the dysregulation of Dlx5 has been associated with the dysregulation of several pro-proliferative factors operating at the restriction point, namely the c-Myc proto-oncogene and the cyclins A, B, D1, D2, and E (111, 112). Due to

45 the context-dependent nature of Dlx5 regulatory functions, these studies in cancerous tissues provide insight into possible targets in other cell lines – indeed, Dlx5 appears to have opposing effects in cancerous tissues when compared to their wild-type counterparts, as is evident by the opposing regulatory effects upon Osc gene expression in osteosarcoma and mouse osteoblast cell lines (115, 110, 116).

As additional evidence to suggest that Dlx5 may regulate the restriction point, the negative effects of Dlx5 expression on proliferation in HEK293 and HeLa cells were ablated upon the expression of the SV40 large T-antigen, which impacts the progression of the restriction point via its interactions with pRB and p53 (117, 125-131).

Therefore, to examine the effects of Dlx5 expression on this cell cycle checkpoint, we conducted RT-qPCR in HEK293 cells transfected with Dlx5. However, none of the genes of interest selected for this experiment demonstrated dysregulation at 24 hours or 48 hours post transfection. As we were using unselected populations, it is possible that this is simply a result of transfection efficiency, or simply that due to the timepoint-specific nature of gene expression for several genes – namely the cyclins – that the effect of Dlx5 expression was simply too low to register on an asynchronous population. To remove transfection efficiency as a variable, we repeated RT-qPCR analyses using drug-selected HeLa cell populations. However, once again, no significant changes in gene expression were observed.

While HEK293 and HeLa cells did demonstrate a reduction in proliferative potential while expressing Dlx5, it should be noted that this effect was more pronounced in DF-1 cells

(117). Therefore, these cell lines may not have been the ideal choice for this experiment.

Functional effects of Dlx5 regulation on cellular proliferation

Corresponding with the evidence that Dlx5 expression has a repressive effect on the rate of cell proliferation in a number of cell types, the EdU labeling index of Dlx5 expressing DF-1 cells was markedly decreased in comparison to controls. However, while a decrease in S-phase entry was observed in DF-1 cells, no corresponding lengthening of the G1 phase of the cell cycle could be confirmed in flow cytometry experiments in HeLa cells selected for Dlx5 expression.

Several possibilities exist that could explain the inconsistencies in the flow cytometric analyses and the EdU incorporation assays of Dlx5-expressing populations of HeLa cells and

DF-1 cells respectively. One unlikely scenario is that the overexpression of Dlx5 prevents the synthesis of new DNA in S-phase. While intended to detect S-phase entry, EdU incorporation assays are limited to the detection of any newly synthesized DNA in which the analogue is incorporated, and thus, EdU labeling will still be observed in cells which had already entered S- phase at the onset of the EdU incubation, and EdU would not be detected in cells that have entered S-phase but have been prevented from replicating their DNA. As our data does not suggest that Dlx5 is transcriptionally regulating the restriction point, it appears unlikely, but still a possibility, that Dlx5 overexpression is preventing DNA-synthesis in cells which have already passed the checkpoint and are in S-phase. Such an occurrence could possibly be explained by interactions with the Zbed1 protein, as DREF is a vital regulator of DNA synthesis and, consequently, S-phase progression in Drosophila, and vertebrate Zbed1 expression increases through the late G1 and peaks in S-phase (80).

Though the possibility that Dlx5 induces apoptosis has been investigated alongside EdU assays in other cell lines, this scenario was not explored in DF-1 cells in this thesis. While cell nuclei were stained for Hoescht in each experiment, and no significant differences in the

47 morphology of cell nuclei between samples were observed, no accurate measure of the rate of apoptotic activity was taken. Therefore, it may prove necessary to conduct TUNEL assays to clarify that the drop in proliferative potential in DF-1 cells was indeed entirely due to inhibited

S-phase entry and not increased apoptotic activity.

A third, and perhaps more likely, scenario is that, due to the nature of the flow cytometric analyses, changes to cell cycle length may not necessarily translate into significant changes in the cell cycle structure of the population. Indeed, in C2C12 cells, previous data in our laboratory suggests that the overall difference in cell cycle length may amount to a single hour between

Dlx5-expressing cells and non-transfected controls, with Dlx5-expressing C2C12 cells having an estimated cell cycle length of 18 hours compared to 17 hours in their counterparts (117). HeLa cells have a slightly longer cell cycle – approximately 22 hours in length and spend roughly half this time in the G1 phase of the cell cycle (144). Assuming that the lengthened cell cycle occurs entirely at the G1 phase as a response to Dlx5 expression, and therefore, G1 accounted for 12 of

23 hours of cell cycle, the G1 phase would only account for approximately 52% of the length of the cell cycle, a mere 2% increase. It seems evident that these small changes may not result in significant alterations of cell cycle profiles. While synchronization of cell populations could possibly correct for these small changes, it seems apparent that six-hour timepoints are simply too far apart to identify any changes in cell cycle progression.

Alternatively, nocodazole synchronization of transfected cell populations could be used in combination with EdU incorporation experiments over several timepoints to track the progression of these cells through G1 and into S phase and clarify differences between G1-S progression in Dlx5-expressing and non-transfected cells.

Zbed1-Dlx protein-protein interactions – a conserved mechanism of regulatory control

While the Drosophila protein DREF has been referred to as a “master key” for cell proliferation, the vertebrate homologue Zbed1 may not play as central a role during cell division.

A recent study, for example, found no absolute correlation between the presence of Ki-67 and

Zbed1 proteins across a variety of human tissues; some Zbed1+ cells were not actively cycling while other cycling populations lacked Zbed1 expression (76). This evidence may be at odds with previous experiments in which the siRNA-mediated knockdown of human Zbed1 transcripts resulted in the prevention of S-phase entry and DNA replication (80) or may indicate that Zbed1 is required for cell cycle progression in a subset of cells, rather than in all vertebrate cell types.

Therefore, as with Dlx5, the effects of Zbed1 expression may be context dependent. However, due to its high expression in differentiation-resistant tissues, Zbed1 may play a role in the repression of differentiation by a mechanism that is not yet defined (76).

Our study provides additional evidence that forced expression of Zbed1 does not stimulate S-phase entry, as Zbed1 expression did not lead to a significant increase in EdU- labelled cells when compared to control DF-1 cells. Furthermore, our qPCR results indicate no change in the mRNA levels of several pro-proliferative or anti-proliferative factors operating at the restriction point in HeLa cells expressing exogenous Zbed1. We cannot rule out that Zbed1 may transcriptionally regulate other cell cycle checkpoint genes not tested in this study though.

Finally, in both asynchronous and synchronized HeLa cell populations expressing Zbed1, no significant difference in cell cycle profile was observed when compared to the vector control.

Unfortunately, due to technical problems, we experienced difficulties examining these populations six hours post-nocodazole release and witnessed large variations at both 18 and 24 hours.

As Zbed1 expression in vertebrates peaks in the S-phase, and its ablation in HeLa cells results in defects in DNA synthesis, the transcription factor might instead be vital for DNA synthesis and S-phase progression (80). Therefore, while our data suggests that Zbed1 does not stimulate cell cycle progression through the restriction point and into S-phase, it does not preclude the possibility that Zbed1 is a critical factor for cell cycle progression in vertebrates.

Furthermore, it may simply be that beyond a minimum threshold of expression, Zbed1 does not result in any obvious changes in phenotype, and therefore, no change is apparent due to its overexpression. However, as Zbed1 expression has an imperfect correlation with Ki-67 in human tissues, it is perhaps not as ubiquitously important in vertebrate cell types as DREF appears to be in Drosophila. Moreover, like the Dlx family, Zbed1 may exhibit functional redundancy with other factors (76, 80). As control of the proliferation/differentiation dichotomy is a critical process, it seems unlikely that a single gene has over-riding control for this in vertebrates.

While our data did not support the idea that Zbed1 stimulates cell cycle progression through the restriction point in vertebrates, our study confirmed that protein-protein interactions between the Dlx family and Zbed1 is conserved; GST-Dlx6 was able to pull down the full-length

Zbed1 protein. Though we were unable to map the domains responsible for this interaction, we suspect that Dlx6 may bind to Zbed1 via its N-terminal domain, as we were able to pull down a

13kDa N-terminal mutant with GST-Dlx6. Unfortunately, due to the affinity of the 13kDa fragment for GST alone, we were unable to replicate this finding in subsequent experiments. A

Zbed1 protein lacking the N-terminal domain would help confirm that the DNA binding domain is important for the interaction with Dlx proteins.

Dlx proteins have been shown to exhibit functional redundancy within bi-gene pairs; it is therefore likely that Zbed1 also physically interacts with Dlx5. While genetically dissimilar from

50 one another apart from their DNA binding regions, it is possible that Dlx5 and Dlx6 encode for genetically distinct, yet functionally similar, terminal domains, as the structure of these Dlx proteins has not been determined. Should an interaction between Dlx5 and Zbed1 also be confirmed and mapped to a terminal domain of the Dlx proteins, it would be a strong case for structural similarities between the Dlx5 and Dlx6 proteins. Thus, it is important that potential interactions between the entire Dlx protein family and Zbed1 be investigated and mapped.

While this thesis demonstrates an interaction occurs in vitro the interaction may not necessarily occur in vivo. To demonstrate that such an interaction occurs, it may be necessary to utilize FRET (Fluorescence Resonance Energy Transfer) which would allow for the direct visualization of these protein interactions in intact cells.

While the interaction between Dlx6 and Zbed1 has been confirmed, the functional importance of the physical interaction between Dlx proteins and Zbed1 remains elusive. The data outlined in this thesis indicates that any interaction does not inhibit the effects of Dlx5 upon cell cycle progression (and indeed, it is likely that this interaction is responsible for this), as DF-1 cells co-expressing both Zbed1 and Dlx5 had an EdU labeling index that was not dissimilar to that of DF-1 cells that expressed Dlx5 alone. However, our experiment did not examine a dose responsive effect of Dlx5 versus Zbed1 expression, as fluorescent antibodies only indicated whether or not these proteins were present in a given cell, not the degree to which they were expressed. Thus, it is possible that the accumulation of Zbed1 may still represent a hurdle which pro-differentiative factors such as the Dlx proteins need to overcome in order to drive differentiation – this would be consistent with the observation that Zbed1 is highly expressed in tissues which are resistant to differentiation (76). We propose a model in which Dlx proteins need to accumulate to a certain threshold in order to overcome Zbed1, and thereby drive

51 differentiation. Indeed, a similar process explains the functional interactions between Dlx5 and

Msx2 in C2C12 cells – as a response to bone morphogenic treatment protein (BMP)-2 treatment,

Dlx5 expression results in the downstream expression of Runx2 and ultimately the upregulation of the transcription of Alkaline Phosphatase (ALP) – yet despite the fact that Dlx5 expression increases dramatically upon treatment, there is a lag between this and ALP expression (147).

However, while initially high levels of Msx2 (another homeodomain protein known to interact with Dlx5) exist in C2C12, this expression also gradually decreases as a result of BMP-2 treatment, leading researchers to propose that a threshold of Dlx5 to Msx2 protein levels had to be overcome in order to stimulate ALP expression (147). As previous data suggests that the

DNA-binding domain of the Dlx5 transcription factor is required to exert its repressive effects upon cellular proliferation (117), it seems likely that this domain will be vital for this interaction, or, possibly, that this interaction occurs on the activating domain of Dlx5, thereby preventing its transcription functions.

Due to the contradictory results indicating the importance of Zbed1 on cell cycle progression, it may be necessary to investigate the effects of Zbed1 knockdown in relation to Ki-

67 expression, particularly in tissues which are known to be resistant to differentiation. This experiment would clarify the importance of Zbed1 on cell cycle progression in various cell types, as well as whether or not its expression is correlated with a significant protective effect against differentiation. Indeed, the simultaneous over-expression of Dlx family members in these tissues with, or without Zbed1 knockdown would allow any differences in propensity to differentiate to be attributed to the presence, or absence, of Zbed1. Finally, Zbed1 could possibly be introduced to cell lines prone to differentiation to see if its presence has an inhibitory effect on this process.

In short, the mechanisms of action of Zbed1 in vertebrate cells remains to be explored.

Conclusion

Consistent with recent evidence suggesting that Zbed1 may not function as the master key to proliferation in vertebrates, the forced expression of this gene did not result in significant increases in S-phase entry in DF-1 cells, and its expression was not associated with any significant changes in the cell cycle profile or transcription of several restriction point genes in

HeLa cells selected for expression of the exogenous protein. Therefore, forced expression of

Zbed1 does not appear to stimulate proliferation in these two vertebrate cell types. However, the ablation of Zbed1 expression in HeLa cells has previously been demonstrated to lead to defects in S-phase entry and DNA-replication, owing to the transcriptional control of the histone H1 gene (80). Thus, it seems apparent that Zbed1 can function as a pro-proliferative factor in vertebrates, though its effects on the proliferation/differentiation dichotomy may be context dependent. One determining factor could be the presence of proteins which interact with Zbed1 and negatively regulate its pro-proliferative functions. In this thesis, it was demonstrated that

Zbed1 interacts with the Dlx family member, Dlx6. However, further research is required in order to determine whether this interaction extends to the rest of the Dlx family, and to determine the significance of this interaction in the regulatory control of the proliferation/differentiation switch.

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