University of Iowa University of Iowa

Iowa Research Online Iowa Research Online

Theses and Dissertations

Summer 2014

Intrinsic and extrinsic regulation of DNA methylation during Intrinsic and extrinsic regulation of DNA methylation during

malignant transformation malignant transformation

Bo-Kuan Wu University of Iowa

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Part of the Cell Biology Commons

Copyright © 2014 Bo-Kuan Wu

This dissertation is available at Iowa Research Online: https://ir.uiowa.edu/etd/1419

Recommended Citation Recommended Citation Wu, Bo-Kuan. "Intrinsic and extrinsic regulation of DNA methylation during malignant transformation." PhD (Doctor of Philosophy) thesis, University of Iowa, 2014. https://doi.org/10.17077/etd.iy8vus1s

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Part of the Cell Biology Commons

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INTRINSIC AND EXTRINSIC REGULATION OF DNA

METHYLATION DURING MALIGNANT TRANSFORMATION

by

Bo-Kuan Wu

A thesis submitted in partial fulfillment of the requirements for the Doctor of

Philosophy degree in Molecular and Cellular Biology in the Graduate College of

The University of Iowa

August 2014

Thesis Supervisor: Professor Charles Brenner

Graduate College The University of Iowa

Iowa City, Iowa

CERTIFICATE OF APPROVAL

_______________________

PH.D. THESIS

_______________

This is to certify that the Ph.D. thesis of

Bo-Kuan Wu

has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Molecular and Cellular Biology at the August 2014 graduation.

Thesis Committee: ___________________________________ Charles Brenner, Thesis Supervisor

___________________________________ Frederick Domann

___________________________________ Adam Dupuy

___________________________________ Dawn Quelle

___________________________________ Michael Wright

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ACKNOWLEDGMENTS

First, I would like to thank my mentor, Dr. Charles Brenner. He provided me a

great opportunity to get excellent training in his lab with ample freedom and full support.

He introduced me to the interesting DNA methylation field, which I plan to keep

focusing on in the future.

I am very grateful to members of my thesis committees: Dr. Frederick Domann,

Dr. Dawn Quelle, Dr. Adam Dupuy and Dr. Michael Wright. Thank you for your

invaluable advice for my research in every seminar and progress report.

I would like to thank all members of Dr. Brenner’s lab, particularly Dr. Rebecca

Fagan and Mr. Samuel Trammell. I greatly appreciate your help.

Finally, I would like to thank my family for their patience and encouragement.

I appreciate everything throughout these six years. Now, I am confident to face

my next challenge.

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ABSTRACT

Cytosine methylation of CpG dinucleotides is an epigenetic modification that

cells use to regulate gene expression, largely to promote transcriptional silencing. Focal

hypermethylation of tumor suppressor genes (TSGs) accompanied by genomic

hypomethylation are epigenetic hallmarks of malignancy. DNA methyltransferase 1

(DNMT1) is the principle vertebrate enzyme responsible for maintenance of DNA

methylation and its dysregulation has been found to lead to aberrant methylation in

cancer. In addition, recent findings demonstrated that the ten-eleven translocation 1

(TET1) protein functions as a 5-methylcytosine dioxygenase that converts 5-

methylcytosine (5mC) bases to 5-hydroxymethylcytosine (5hmC) to mediate active DNA

demethylation. Emerging evidence suggests that TET1 might function as a TSG. To

understand the dynamic regulation of DNA methylation during cellular transformation,

my work focused on intrinsic regulation of DNMT1 and how TET1 regulates DNA

demethylation in generating a cancer methylome.

The replication foci targeting sequence (RFTS) is an N-terminal domain of

DNMT1 that inhibits DNA-binding and catalytic activity, suggesting that RFTS deletion

would result in gain of DNMT1 function. However, other data suggested that RFTS may

be a positively acting domain. To test biochemical and structural predictions that the

RFTS domain of DNMT1 is inhibitory, we established cellular systems to evaluate the

function of DNMT1 alleles. The data indicate that deletion of RFTS is necessary and

sufficient to promote cellular transformation, focal hypermethylation of specific TSGs,

and global hypomethylation. These data and human mutation data suggest that RFTS

domain is a target of tumor-specific dysregulation.

RAS mutations are frequently observed in multiple malignancies. Methylation-

associated silencing of TSGs is a hallmark of RAS-driven-tumorigenesis. I discovered

that suppression of TET1 by the ERK signaling cascade is responsible for promoter

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hypermethylation and the malignant phenotype in KRAS-transformed cells. Restoration

of TET1 expression reactivates silenced TSGs and reduces colony formation. Moreover,

TET1 knockdown in a cell depleted for KRAS is sufficient to rescue the inhibition of

colony formation by KRAS knockdown. My findings suggest that impaired TET1-

mediated DNA demethylation is a target responsible for epigenetic changes in cancers

with KRAS activation.

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TABLE OF CONTENTS

LIST OF TABLES ............................................................................................................ vii

LIST OF FIGURES ......................................................................................................... viii

LIST OF ABBREVIATIONS………………………………………………......................x

CHAPTER

I. INTRODUCTION ............................................................................................1

II. RFTS-DELETED DNMT1 ENHANCES TUMORIGENICITY WITH FOCAL HYPERMETHYLATION AND GLOBAL HYPOMETHYLATION ..................................................................................7

2.1 Abstract ..............................................................................................7 2.2 Introduction ........................................................................................8 2.3 Materials and Methods ....................................................................11

2.3.1 Cell culture………. ...........................................................11 2.3.2 Establishment of stable cell lines ......................................11 2.3.3 Proliferation and invasion assay ........................................11 2.3.4 RT-qPCR ...........................................................................12 2.3.5 Immunoblotting .................................................................12 2.3.6 Adherent and soft-agar colony formation ..........................12 2.3.7 Methylation assay ..............................................................13 2.3.8 ChIP ...................................................................................13 2.3.9 Nuclease-protection assay .................................................14 2.3.10 HELP assay and data analysis ...........................................14 2.3.11 Stastical analysis ................................................................14

2.4 Results ..............................................................................................14 2.4.1 Deletion of RFTS enhances the oncogenic activity

of DNMT1 .......................................................................14 2.4.2 Promoter hypermethylation and transcriptional

silencing of DAPK and DUOX1 is driven by DNMT1 ............................................................................16

2.4.3 Strong alleles of DNMT1 condense chromatin structure at the DAPK and DUOX1 promoters ................17

2.4.4 DNA demethylating agent 5-aza-deoxycytidine (5-aza-dC) reverses gene silencing and diminishes the transformation ability of strong DNMT1 alleles. .............18

2.4.5 Genome-wide promoter methylation analysis reveals that DNMT1-ΔRFTS cells produce a methylation pattern similar to DNMT1 cells, though more intense ...............................................................................18

2.4.6 DNMT1-ΔRFTS cells exhibit genomic hypomethylation ...............................................................20

2.4.7 DNMT1-ΔRFTS expression has similar effects in H358 lung cancer cells .....................................................21

2.5 Discussion ........................................................................................22

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III. SUPPRESSION OF TET1-DEPENDENT DNA DEMETHYLATION IS ESSENTIAL FOR KRAS-MEDIATED TRANSFORMATION ..............43 3.1 Abstract ............................................................................................43 3.2 Introduction ......................................................................................44 3.3 Materials and Methods ....................................................................47

3.3.1 Cell culture………. ...........................................................47 3.3.2 Establishment of stable cell lines ......................................47 3.3.3 RT-qPCR ...........................................................................48 3.3.4 Immunoblotting .................................................................48 3.3.5 Proliferation assay .............................................................49 3.3.6 Adherent and soft-agar colony formation ..........................49 3.3.7 DNA dot blot assay ...........................................................49 3.3.8 MeDIP and hMeDIP ..........................................................50 3.3.9 Bisulfite sequencing ..........................................................50 3.3.10 ChIP ...................................................................................50 3.3.11 siRNA transfection ............................................................51 3.3.11 Stastical analysis ................................................................51

3.4 Results ..............................................................................................51 3.4.1 Oncogenic KRAS expression is sufficient to

transform non-malignant HBEC3 cells ............................51 3.4.2 Oncogenic KRAS expression causes

hypermethylation-mediated silencing of TSGs and loss of imprinting ..............................................................52

3.4.3 KRAS negatively regulates TET1 expression through the ERK signaling pathway ................................54

3.4.4 Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation. ....................................................55

3.4.5 Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 Cells ...............................................56

3.4.6 KRAS-mediated suppression of TET1 is required for maintenance of the malignant phenotype in H1299 cancer cells .......................................................................58

3.5 Discussion ........................................................................................59 IV. CONCLUSION AND FUTURE DIRECTION ..............................................86

4.1 Implication of DNMT1 RFTS domain mutant and RFTS domain association protein (RAP) in cancer ...................................86

4.2 Implication of suppression of TET1 in KRAS-dependent transformation .................................................................................89

REFERENCES ..................................................................................................................91

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LIST OF TABLES

Table

2.1 Target list of TSGs have been found with hypermethylation-mediated gene silencing in lung cancers ...........................................................................................38

2.2 Summary of the changes of promoter methylation and gene expression in DNMT1-expressing cell lines ...................................................................................39

2.3 KEGG pathway enrichment analysis ........................................................................40

2.4 DNMT1 RFTS domain mutations were found in cancer (COSMIC database) ........41

2.5 Primer list. .................................................................................................................42

3.1 Target list of hypermethylated and silenced lung cancer TSGs ...............................81

3.2 Summary of the changes of promoter methylation and gene expression in KRAS-expressing cell lines ......................................................................................82

3.3 Human primers .........................................................................................................83

3.4 Mouse primers ..........................................................................................................85

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LIST OF FIGURES

Figure

2.1 Deletion of RFTS enhances the oncogenic activity of DNMT1 ...............................25

2.2 Ectopic expression of DNMT1-ΔRFTS enhances invasion activity without proliferation ..............................................................................................................26

2.3 DNMT1-ΔRFTS promotes increased methylation and silencing of the DAPK and DUOX1 genes ....................................................................................................27

2.4 DNMT1-ΔRFTS decreases chromatin accessibility at DAPK and DUOX1 promoters ..................................................................................................................29

2.5 5-aza-dC treatment reactivates TSG expression and suppresses DNMT1-dependent transformation .........................................................................................30

2.6 Ectopic expression of DNMT1 alleles does not radically alter global methylation intensities ..............................................................................................31

2.7 DNMT1-ΔRFTS expression enhances global DNMT1 methylation changes ..........32

2.8 Genomic hypomethylation is found in DNMT1-ΔRFTS cells .................................33

2.9 The methylation levels of LINE1 were not changed in DNMT1 or DNMT1-ΔRFTS cells ..............................................................................................................34

2.10 Ectopic expression of DNMT1-ΔRFTS in H358 cells is sufficient to enhance proliferation, invasion and soft-agar colony formation ............................................35

2.11 Ectopic expression of DNMT1-ΔRFTS in H358 cells caused gene silencing of DAPK and DUOX1 and demethylation of SAT2 ................................................36

2.12 Dual roles for RFTS domain in DNMT1-dependent DNA methylation ..................37

3.1 Oncogenic KRAS expression is sufficient to transform non-malignant HBEC3 cells .............................................................................................................63

3.2 Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs .........................................................................................................................64

3.3 Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs and loss of imprinting .....................................................................................66

3.4 KRAS negatively regulates TET1 expression through the ERK signaling pathway .....................................................................................................................67

3.5 ERK pathway inhibition reactivates silenced H19 expression in KRAS cells .........68

3.6 Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation ..........................................................69

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3.7 Reduction of 5hmC and TET1-association are responsible for KRAS-mediated DNA hypermethylation .............................................................................71

3.8 Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 cells ......................................................72

3.9 Kras-mediated suppression of Tet1 is associated with decreased 5hmC and increased 5mC levels ................................................................................................74

3.10 Kras promotes transformation by inhibiting Tet1 expression ..................................75

3.11 Erk pathway inhibition increases Tet1 expression in Kras-transformed NIH3T3 cells, while Akt pathway inhibition shows no effect .................................77

3.12 KRAS-mediated suppression of TET1 is required for maintaining malignant phenotype in H1299 cancer cells ..............................................................................78

3.13 KRAS-mediated suppression of TET1 is found in HepG2 hepatoma cancer cells ...........................................................................................................................80

4.1 Multiple sequence alignment analysis of the RFTS domain of DNMT1 showed mutations found in cancer patients were occurred in conserved loci. .........90

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LIST OF ABBREVIATIONS

5-aza-dC 5-aza-deoxycytidine

5caC 5-carboxylcytosine

5fC 5-formylcytosine

5hmC 5-hydroxymethylcytosine

5mC 5-methylcytosine

AP1 activator protein 1

BAH bromo-adjacent homolog

ChIP chromatin immunoprecipitation

COSMIC catalogue of somatic mutation in cancer

DMAP1 DNMT1 associated protein 1

DNMT1 DNA methyltransferase 1

DNMT3A DNA methyltransferase 3A

DNMT3B DNA methyltransferase 3B

EGF epidermal growth factor

ERK extracellular signaling-regulated kinase

ESC embryonic stem cells

HBEC3 human bronchial epithelial cells

HELP HpaII tiny fragment enrichment by ligation-mediated PCR

hMeDIP 5-hydroxymethylcytosine DNA immunoprecipitation

HSAN1E hereditary sensory and autonomic neuropathy

ICR imprinting control region

KEGG Kyoto Encyclopedia of Genes and Genomes

MeDIP methylated DNA immunoprecipitation

NAA10 N-_-acetyltransferase 10 NatA catalytic subunit

NLS nuclear localization signal

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ns no significant difference

NSCLC nonsmall cell lung cancer

pAKT phospho-AKT

PCNA proliferating cell nuclear antigen

pERK phospho-ERK

RAPs RFTS-targeted DNMT1 associated proteins

RFTS Replication foci targeting sequence

SAT2 Satellite 2 repeat sequences

siRNA small interfering RNA

Sp1 specificity protein 1

TAB-seq Tet-assisted bisulfite sequencing

tAKT total-AKT

TCF T-cell-factor

tERK total-ERK

TET Ten-eleven translocation

TSGs tumor suppressor genes

UHRF1 ubiquitin-like containing PHD and RING finger domain protein 1

USP7 ubiquitin-specific-processing protease 7

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CHAPTER I

INTRODUCTION

DNA methylation is an epigenetic modification involved in development,

transcription, imprinting, X chromosome inactivation and genomic structure (Baylin

2005). In mammals, DNA methylation typically occurs at the cytosine base of CpG

dinucleotides (Bernstein, Meissner, and Lander 2007; Laird and Jaenisch 1996). Most

CpG dinucleotides fall in repetitive sequences and are heavily methylated to form

heterochromatin to maintain genomic stability (Jones and Baylin 2007). In addition, the

genomic distribution of CpG dinucleotides is uneven. Apart from heterochromatin, they

are usually clustered in gene promoter regions term CpG islands (M. M. Suzuki and

Bird 2008). Half of the genes in mice and humans contain CpG islands (M. M. Suzuki

and Bird 2008; Singal and Ginder 1999). Promoter CpG islands are tend to become

methylated to repress expression of downstream genes (Jones and Baylin 2007). Since

DNA methylation plays an important role in regulating many cellular processes,

abnormal DNA methylation is associated with diseases, including cancer (Robertson

2005).

Because of the obvious link between DNA hypermethylation and transcriptional

repression, tumor suppressor genes (TSGs) have been considered to the most highly

regulated sites for methylation alteration during tumorigenesis (Hughes et al. 2013; Issa

2004). Methylation-associated silencing of TSG induces cancer formation and

progression. Thus, chemotherapy that aims to effect DNA demethylation has become a

promising anti-cancer approach that might reactivate TSG expression (Strathdee and

Brown 2002; Szyf 2005). However, in addition to promoter hypermethylation, global

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genomic hypomethylation has been found in cancer (Jackson et al. 2004). Genomic

hypomethylation could cause transcription activation of oncogenes, chromosome

rearrangement and genomic instability (Ehrlich 2002; Ehrlich 2009). The relationship

between regional hypermethylation and global hypomethylation remains unclear.

In mammals, there is a group of enzymes are responsible for establishing and

maintaining DNA methylation pattern (Chen and Riggs 2011; Rountree et al. 2001).

DNA methyltransferase 3A (DNMT3A) and DNA methyltransferase 3B (DNMT3B)

mediate de novo methylation to deposit methyl groups to naked DNA (Chen and Riggs

2011; Rountree et al. 2001). After DNA replication, DNA methyltransferase 1

(DNMT1) is the principal enzyme responsible for maintenance of cytosine methylation

at CpG dinucleotides (Law and Jacobsen 2010). DNMT1 copies the present methylation

patterns from the parental DNA strand to the newly synthesized strand (Chen and Riggs

2011; Rountree et al. 2001). However, impaired maintenance DNA methylation activity

could cause passive DNA demethylation after DNA replication (Law and Jacobsen

2010). Thus, DNMT1 bears responsibility to increase or decrease the degree of DNA

methylation. Indeed, dysregulation of DNMT1 is associated with either promoter

hypermethylation (J. Wu et al. 1993; Bakin and Curran 1999; Biniszkiewicz et al. 2002)

or genomic hypomethylation (E. Li, Bestor, and Jaenisch 1992; Gaudet 2003). Thus,

changing the expression of DNMT1 alone is not sufficient to recapitulate the regionally

increased and globally decreased DNA methylation changes observed in cancer.

DNMT1 harbors N-terminal regulatory domain and C-terminal catalytic domain.

The long N-terminal regulatory domain is composed of DNMT1 associated protein 1

(DMAP1), proliferating cell nuclear antigen (PCNA)-binding domain (PBD), nuclear

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localization signal (NLS), Replication foci targeting sequence (RFTS), CXXC, and

bromo-adjacent homolog (BAH) (Qin, Leonhardt, and Pichler 2011). These regulatory

domains have been described as essential for catalytic activity, protein association and

target specificity (Qin, Leonhardt, and Pichler 2011). Previous papers showed the RFTS

domain (Leonhardt et al. 1992) and the CXXC domain (M. Pradhan et al. 2008) are

positively acting domains required for DNMT1 activity. However, recent papers

showed these two domains could be endogenous inhibitors of DNMT1 (Syeda et al.

2011; Song et al. 2011). With a sensitive fluorescence-based assay, Syeda et al.

demonstrated that deletion of the RFTS domain lead to a 640-fold increase in DNMT1

methylation activity. Thus, the regulatory domain of DNMT1 could regulate DNMT1

enzyme activity intrinsically. However, the effect on reprogramming DNA methylation

during tumorigenesis is still obscure.

In normal cell, the level of DNMT1 is tightly regulated during cell cycle (Szyf et

al. 1985; Szyf, Bozovic, and Tanigawa 1991). However, there is substantial evidence

that DNMT1 is up-regulated and hyperactive in some cancer types, including lung (Lin

et al. 2007; Xing et al. 2008; Kwon et al. 2006), colorectal (el-Deiry et al. 1991; De

Marzo et al. 1999), liver (Park, Yu, and Shim 2006; Saito et al. 2003) and gastric (Etoh

et al. 2004) cancers, indicating extrinsic mechanisms to enhance DNMT1 expression

and activity. Several oncogenic pathways could affect DNMT1 function by

transcriptional, post-transcriptional and post-translational mechanisms. At the

transcription level, activators, including AP1 (Rouleau and MacLeod 1995; MacLeod,

Rouleau, and Szyf 1995), E2F1 (McCabe, Davis, and Day 2005; H. Kimura 2003),

specificity protein 1 (Sp1) (Lin, Wu, et al. 2010), T-cell-factor (TCF) (Campbell and

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Szyf 2003) and STAT3 (Q. Zhang 2006), increase DNMT1 transcription. On the other

hand, pRb (H. Kimura 2003) and p53 (Peterson, Bögler, and Taylor 2003) are negative

regulators. These transcription factors bind to cognate sites on the DNMT1 promoter

region to repress transcription. Utilizing post-transcriptional regulation, AUF1

(Torrisani et al. 2006) and microRNA-152 (J. Huang et al. 2010) could specifically

target the 3’-UTR of DNMT1 mRNA (Detich 2001) to affect mRNA stability. Finally,

post-translational modification of DNMT1 via phosphorylation (L. Sun et al. 2007; Lin,

Hsieh, et al. 2010) or methylation (Estève et al. 2009; Wang et al. 2008) affects its

protein degradation, which is dependent on the ubiquitin-proteasome pathway (L. Sun

et al. 2007; Lin, Hsieh, et al. 2010; Estève et al. 2009; Wang et al. 2008). Thus,

oncoproteins and tumor suppressor proteins play roles in regulating Dnmt1 activity

These frequent cancer-specific alterations produce cells with significantly

increased DNMT1 activity. The most notable case is the oncogenic RAS pathway.

There is a growing bodies of evidence showed that DNMT1 is regulated by the

RAS/AP1 pathway (Rouleau and MacLeod 1995; MacLeod and Szyf 1995; Bigey et al.

2000) and DNMT1 is essential for RAS-induced DNA hypermethylation phenotype

(Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007a). RAS is an onco-protein

belongs to the low molecular GTP-binding protein (Downward 2003). RAS signaling

regulates cell growth, cell differentiation and survival. Activated mutant RAS has been

found in about 20-25% human tumors and up to 90% in some cancer type (Bos 1989).

In response to extracellular signals, RAS switches from an inactive GDP-bound form

into an active GTP-bound form and subsequently transmits the signal to a cascade of

downstream serine/threonine kinases, consisting of RAF, MEK1 and extracellular

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signaling-regulated kinase (ERK) (Downward 2003). After phosphorylation and

activation, ERK translocates from the cytoplasm to nucleus to increase the expression

of FOS. ERK also phosphorylates and activates c-Jun, the transcriptional activator. c-

Jun and c-fos further form a transcription factor, activator protein 1 (AP1), to activate

transcription by targeting AP1 regulatory element (Shaulian and Karin 2002).

Early experiments established that the RAS/AP1 pathway can directly regulate

DNMT1 transcription (Rouleau and MacLeod 1995; MacLeod and Szyf 1995; Bigey et

al. 2000). In addition, forced expression of H-ras in several cell lines increases DNMT1

transcription (Rouleau and MacLeod 1995; Pakneshan, Szyf, and Rabbani 2005),

whereas inhibition of RAS , MEK or c-jun decreases the DNMT1 mRNA level

(MacLeod, Rouleau, and Szyf 1995; Lu et al. 2007). Furthermore, Gazin et al.

identified that Kras activation leads to a cascade in which Dnmt1 is recruited to a

specific set of genes to be silenced including Fas, Sfrp1, Par4, Plagl1, H2-K1 and Lox

(Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007a). They performed a genome-

wide shRNA screen to identify the essential genes required for Ras-mediated epigenetic

silencing of Fas and obtained 28 genes, including Dnmt1. Knockdown any of these 28

genes resulted in promoter demethylation and reexpression of Fas in NIH3T3 cells that

had been transformed by Kras. These data indicated that Dnmt1 plays an essential and

common role in Ras-mediated epigenetic silencing. Sufficient Dnmt1 level and

subsequently epigenetic silencing of some TSGs are the key steps for the initiation and

maintenance stage of Ras-mediated transformation.

Since DNMT1-dependent epigenetic silencing of TSGs contributes to initiate

and maintain cancer, it is reasonable to test whether TSG reactivation could reverse the

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transformation phenotype. In cell culture, knocking down DNMT1 by siRNA as well as

5-aza-dC treatment reversed the epigenetic silencing of TSGs and transformation

(MacLeod and Szyf 1995; Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007a;

Bakin and Curran 1999). In mice, genetic reduction of Dnmt1 mice can effectively

reduce tumor formation by mutant ApcMin (Laird et al. 1995; Eads, Nickel, and Laird

2002) or NNK treatment (Belinsky et al. 2003) as well as 5-aza-dC treatment. These

observations indicate that sufficient DNMT1 level and subsequent epigenetic silencing

of specific TSGs are key steps in tumorigenesis, which validates DNMT1 as an

important target for cancer therapy.

In this thesis, we aimed to dissect the intrinsic and extrinsic regulation of DNA

methylation by DNMT1 during malignant transformation. By understanding these

mechanisms individually or collectively, it might provide us a new way to specifically

target DNMT1 for cancer therapy.

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CHAPTER II

RFTS-DELETED DNMT1 ENHANCES

TUMORIGENICITY WITH FOCAL

HYPERMETHYLATION AND GLOBAL

HYPOMETHYLATION

2.1 Abstract

Site-specific hypermethylation of tumor suppressor genes accompanied by

genome-wide hypomethylation are epigenetic hallmarks of malignancy. However,

molecular mechanisms that drive these linked changes in DNA methylation remain

obscure. DNA methyltransferase 1 (DNMT1), the principle enzyme responsible for

maintaining methylation patterns is commonly dysregulated in tumors. Replication foci

targeting sequence (RFTS) is an N-terminal domain of DNMT1 that inhibits DNA-

binding and catalytic activity, suggesting that RFTS deletion would result in gain of

DNMT1 function. However, a substantial body of data suggested that RFTS is required

for DNMT1 activity. Here, we demonstrate that deletion of RFTS alters DNMT1-

dependent DNA methylation during malignant transformation. Compared to full-length

DNMT1, ectopic expression of hyperactive DNMT1-ΔRFTS caused greater malignant

transformation and enhanced promoter methylation with condensed chromatin structure

that silenced DAPK and DUOX1 expressions. Simultaneously, deletion of RFTS

impaired DNMT1 chromatin association with pericentromeric Satellite 2 (SAT2) repeat

sequences and produced DNA demethylation at SAT2 repeats and globally. To our

knowledge, RFTS-deleted DNMT1 is the first single factor that can reprogram focal

hypermethylation and global hypomethylation in parallel during malignant

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transformation. Our evidence suggests that the RFTS domain of DNMT1 is a target

responsible for epigenetic changes in cancer.

2.2 Introduction

Methylation of cytosine bases in CpG dinucleotides is an epigenetic

modification in mammals involved in development, imprinting and X chromosome

inactivation, which becomes dysregulated in carcinogenesis (Robertson 2005).

Increased DNA methylation in promoter regions is highly associated with

transcriptional silencing and hypermethylation-mediated silencing of tumor suppressor

genes (TSGs) is a critical epigenetic driver of cancer (Issa 2004; Belinsky 2004).

However, in addition to site-specific hypermethylation, a reduction of total cytosine

methylation of the genome, targeted in part to repetitive elements, is also common in

malignancies (Jones and Baylin 2002; Feinberg et al. 1988; Eden et al. 2003). Global

hypomethylation may be a driver of cancer by promoting genomic instability and

oncogene activation. Although both hypermethylation and hypomethylation are found

in cancer, the molecular mechanisms that link these alterations are not clear (Ehrlich

2002).

DNA methyltransferase 1 (DNMT1) is the enzyme most responsible for the

maintenance of DNA methylation patterns during DNA replication. In addition to

maintenance methylation activity, DNMT1 can perform de novo DNA methylation, like

DNMT3A and DNMT3B (Christman et al. 1995; S. Pradhan et al. 1997; Jair et al.

2006). Dysregulated DNMT1 is capable of promoting hypermethylation or

hypomethylation and, indeed, Dnmt1 has been reported to be up-regulated or down-

regulated in different types of cancer (el-Deiry et al. 1991; Belinsky et al. 1996; Lin et

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al. 2007; F. Kimura et al. 2003). Methyltransferase activity of DNMT1 has been

considered the critical effector in reprogramming cancer methylation patterns.

However, enforced overexpression of DNMT1 produces hypermethylation of TSGs and

increases methylation of the whole genome (J. Wu et al. 1993; Bakin and Curran 1999;

Biniszkiewicz et al. 2002). Conversely, deficient DNMT1 causes global

hypomethylation without site-specific hypermethylation (E. Li, Bestor, and Jaenisch

1992; Gaudet 2003). Thus, changing the expression of DNMT1 alone is insufficient to

recapitulate the linked and vectorially opposite epigenetic alterations observed in cancer.

DNMT1 consists of a series of globular domains N-terminal to the catalytic

domain, which are implicated in functions essential for catalytic activity, protein

association and target specificity. By interacting with different proteins, DNMT1 may

be enriched in or dissociated from specific genomic loci (Di Croce et al. 2002; Viré et

al. 2006; Esteve et al. 2006; Smallwood et al. 2007; Chuang 1997; Hervouet et al. 2010;

S. Pradhan and Kim 2002) with the distribution of DNMT1 determining the methylation

status of specific target sites. For example, the oncogenic PML-RAR fusion protein

recruits DNMT1 to the RARβ2 promoter to stimulate methylation (Di Croce et al. 2002),

whereas disruption of the DNMT1-proliferating cell nuclear antigen (PCNA) interaction

results in hypomethylation of cellular DNA (Chuang 1997; Hervouet et al. 2010).

Seemingly, both enzyme activity and chromatin occupancy of DNMT1 are important

for DNMT1-dependent changes in DNA methylation observed during tumorigenesis.

The replication foci targeting sequence (RFTS) was defined as an N-terminal

domain required for associating DNMT1 with replication foci (Leonhardt et al. 1992).

Multiple lines of evidence suggested that RFTS is a positively acting domain required

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for DNMT1 function (Margot et al. 2000; Leonhardt et al. 1992; Bostick et al. 2007;

Sharif et al. 2007). Surprisingly, recombinant forms of DNMT1 with the RFTS domain

bind DNA poorly, while specific deletion of RFTS activates DNA binding (Syeda et al.

2011). Moreover, using a fluorigenic methylation assay, we showed that deletion of

RFTS leads to a 640-fold increase in methylation activity due to relief of DNA-

competitive inhibition by RFTS. Further biochemical and structural experiments

support RFTS as a DNA-competitive endogenous inhibitor of DNMT1 that must be

removed from the DNA active site for DNMT1 activity (Takeshita et al. 2011;

Hashimoto et al. 2012; Berkyurek et al. 2014; Bashtrykov, Jankevicius, et al. 2014;

Bashtrykov, Rajavelu, et al. 2014). In contrast to our biochemical insights into negative

regulation by RFTS, CXXC was proposed as a nonmethylated DNA-binding inhibitor

of DNMT1 activity (Song et al. 2011). Human genetics has the potential to help resolve

DNMT1 domain function because an autosomal dominant hereditary sensory and

autonomic neuropathy (HSAN1E) maps to the RFTS domain (Klein et al. 2011). This

mutation is associated with genomic hypomethylation and promoter hypermethylation

(Klein et al. 2011).

In this study, we investigate the functional impact of two DNMT1 regulatory

domains, RFTS and CXXC. Here we show that two-fold overexpression of full-length

and various deletion mutants resulted in degrees of promoter hypermethylation,

chromatin condensation, and transcriptional repression. Ectopic expression of DNMT1-

ΔRFTS led to the most malignant phenotype, which required the presence of the CXXC

domain for oncogenic ability. Surprisingly, DNMT1-ΔRFTS cells further showed a

cancer-like global hypomethylation phenotype. These data suggest that tumor-specific

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targeting of RFTS function may mediate the commonly observed linkage of focal

hypermethylation and global hypomethylation.

2.3 Materials and Methods

2.3.1 Cell culture

HBEC3 cells and stable cell lines were grown in KSFM media supplemented

with bovine pituitary extract and recombinant human EGF. H358 cells (CRL-5807,

ATCC) and stable cell lines were grown in RPMI1640 media with 10% serum.

2.3.2 Establishment of stable cell lines

To establish full length or deletion mutant human DNMT1 overexpression stable

lines, a full-length DNMT1 cDNA clone (SC325419, Origene) was used as template to

amplify full-length or deletion mutant DNMT1 fragments. PCR fragments were first

T/A cloned into pGEM-T easy vector (A1360, Promega) and then subcloned into

pLenti6/V5 vector (K4950-00, Invitrogen). Viral production and transduction was

performed using the ViralPower Bsd Lentiviral Support Kit (K4950-00, Invitrogen).

Primer pairs used for plasmid constructions are provided in Table 2.5.

2.3.3 Proliferation and invasion assay

To analyze proliferation, 1,000 (HBEC3) or 5,000 (H358) cells were seeded in

replicates of 6 in KSFM media with or without EGF supplementation in 96-well plates.

Relative cell numbers were analyzed using Resazurin (R7017, Sigma) 72 hrs after

seeding. Data were collected from 4 independent experiments. Cell invasion was

analyzed using the CultreCoat® 24 Well Low BME Invasion Assay kit (3481-024-01,

Trevigen).

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2.3.4 RT-qPCR

Total RNA was extracted using the RNeasy Mini Kit (74106, Qiagen) with

DNase treatment (79254, Qiagen) to eliminate DNA contamination. Equal amounts of

RNA were reverse transcribed to generate cDNA using the QuantiTect Reverse

Transcription Kit (205314, Qiagen). Specific primer pairs were then used to amplify

target genes (Table 2.5). qPCR reactions were conducted with iQ SYBR Green

Supermix (170-8884, Bio-Rad). All data were collected from 3 or 4 independent

experiments.

2.3.5 Immunoblotting

Protein extracts from each stable cell lines were prepared in RIPA buffer

(89900, Thermo Scientific) according to the manufacturer's instructions. Equal amounts

of protein were separated using NuPAGE® Novex® 3–8% Tris-Acetate Gel and

transferred to 0.2 µm nitrocellulose membrane at 4 °C overnight. DNMT1

(WH0001786M1, Sigma) and actin (ab3280, Abcam) were detected using specific

antibodies and visualized by SuperSignal West Femto Substrate (34096, Thermo

Scientific).

2.3.6 Adherent and soft-agar colony formation

For adherent colony formation, 200 cells were seeded on 10-cm culture dishes in

KSFM media without EGF, and allowed to grow for 12 days, followed by 4%

methylene blue (M9140, Sigma) staining. Colonies > 3 mm were counted. For soft-agar

colony formation, 10,000 (HBEC3) or 5,000 (H358) cells were resuspended in media

with 0.4% agarose and plated over a layer of 0.6 % agarose. Cells were incubated at 37

°C for 6 (HBEC3) or 4 (H358) weeks and then colonies were stained with MTT

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(M5655, Sigma). Colony images were acquired with ChemiDoc XRS (Bio-Rad) and

quantified using Quantity One software (Bio-Rad). All data were collected from 2 or 3

independent experiments, each performed in triplicate.

2.3.7 Methylation assay

Genomic DNA was extracted using DNeasy Blood & Tissue Kit (69506,

Qiagen). Promoter methylation analysis was performed using MethylMiner Methylated

DNA Enrichment Kit (ME10025, Invitrogen) or bisulfite sequencing. For MethylMiner

experiments, genomic DNA was first fragmented by sonication to an average size of

400 bps. Methylated DNA was captured following the manufacturer protocol. The

methylation level was analyzed using specific primer sets with qPCR (Table 2.5). 10%

of input DNA was used as a control. Data were collected from 3 independent

experiments. For bisulfite sequencing, genomic DNA was treated with bisulfite using

EpiTect Bisulfite kit (59124, Qiagen). Bisulfite treated DNA was then used as a

template and PCR was performed using specific primer pairs (Table 2.5). Final PCR

products were gel purified and cloned into the pGEM-T easy vector (A1360, Promega).

Independent clones were subjected to sequencing. Global methylation was analyzed by

HPLC. Genomic DNA was digested to nucleosides using DNA Degradase Plus Kit

(E2021, Zymo Research). 1 µg digested DNA was separated on a 25 cm x 4.6 µm C18

Supelco column using 7.5 mM ammonium phosphate and methanol. Retention time of

each nucleoside was determined by nucleotide standards. Data were collected from 3

independent experiments, each performed in duplicate.

2.3.8 ChIP

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ChIP was performed with Magna ChIP HiSens chromatin immunoprecipitation

kit (17-10461, Millipore) and analyzed using qPCR. 10% of input DNA was used as a

control. All data were collected from 3 independent experiments. DNMT1 antibody

(IMG-261A, Imgenex) was used.

2.3.9 Nuclease-protection assay

Chromatin accessibility was assessed using EpiQ™ Chromatin Preparation Kit

(172-5402, Bio-Rad). Primers used are listed in Table 2.5. All data were collected from

3 independent experiments, each performed in triplicate.

2.3.10 HELP assay and data analysis

Genomic DNA was extracted using the DNeasy Blood & Tissue Kit (69506,

Qiagen). The HELP assay was conducted in the Epigenomics Core Facility of Weill

Cornell Medical College. Samples were hybridized to a custom oligonucleotide array

(Roche NimbleGen, Madision, WI; design name:

100128_HG19_MKF_HELP_ChIP_HX3). HELP data were processed using the HELP

package in R from Bioconductor. Primary data are available from the NCBI GEO

public database (accession number: GSE57829).

2.3.11 Statistical analysis

All data were presented as mean ± SD. One-way ANOVA was used to calculate

P-value and determine significance. P-value lower than 0.05 was considered

statistically significant.

2.4 Results

1.4.1 Deletion of RFTS enhances the oncogenic activity of

DNMT1

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To dissect the functional role of two regulatory domains in human DNMT1, we

established stable cell lines in non-malignant human bronchial epithelial cells (HBEC3)

cells (Sato et al. 2006) that expressed full-length DNMT1 (DNMT1 cells), RFTS-

deleted DNMT1 (DNMT1-ΔRFTS cells), CXXC-deleted DNMT1 (DNMT1-ΔCXXC

cells), RFTS/CXXC double-deleted DNMT1 (DNMT1-ΔR/C cells), or vector alone

(vector cells). mRNA and protein validation showed that each DNMT1 construct was

expressed at levels equivalent to that of endogenous DNMT1 in HBEC3 cells (Fig.

2.1A). Because DNMT1 has protooncogenic properties (J. Wu et al. 1993; Bakin and

Curran 1999; Damiani et al. 2008), we evaluated transformation markers of the

resulting cell lines. There was no significant proliferation difference between vector

cells and DNMT1-expressing cells, with or without supplementation of epidermal

growth factor (EGF) (Fig. 2.2A). We then examined whether DNMT1 overexpression

could alter invasive activity, a hallmark of cancer metastasis (Fig. 2.2B). Although

invasion was slightly enhanced in all DNMT1-expressing cells, only DNMT1-ΔRFTS

and DNMT1-ΔR/C cells showed a significant difference compared to vector cells,

suggesting that deletion of the RFTS domain provides an invasive advantage.

To further analyze the oncogenic properties of these cells, we examined

anchorage-dependent growth (Fig. 2.1B). Overexpression of all forms of DNMT1

increased the number of colonies observed. However, whereas DNMT1 cells produced

two-fold more colonies, DNMT1-ΔRFTS cells produced three-fold more colonies. We

further tested the anchorage-independent colony-forming ability of these cells by soft-

agar colony formation (Fig. 2.1C). Because HBEC3 is a non-malignant cell line, vector

cells produced fewer than 5 colonies per dish. However, DNMT1 cells produced 30

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colonies and DNMT1-ΔRFTS cells produced 130 colonies in soft agar. Interestingly,

though DNMT1-ΔRFTS cells elevated soft-agar colony-forming potential, removing the

CXXC domain in DNMT1-ΔR/C cells completely abolished the effect. These data

indicate that the CXXC domain is essential for the oncogenic activity of DNMT1-

ΔRFTS. Thus, DNMT1 acts as weak oncoprotein, whose transforming activity is

enhanced by deletion of the inhibitory RFTS domain.

2.4.2 Promoter hypermethylation and transcriptional

silencing of DAPK and DUOX1 is driven by DNMT1.

Several TSGs are methylation-associated silencing in lung cancer (Table 2.1).

To test whether introduction of ectopic DNMT1 alleles is sufficient to alter DNA

methylation of these genes, we examined the methylation status of promoter-associated

CpG islands and relevant transcript levels for each of 24 TSGs (Table 2.2). Of these

genes, DAPK (D. H. Kim et al. 2001; Pulling et al. 2004) and DUOX1 (Luxen,

Belinsky, and Knaus 2008) were consistently hypermethylated and silenced by all

DNMT1 alleles. By performing methylated DNA immunoprecipitation (MeDIP), we

found a ~2-fold increase in methylation of the DAPK promoter in all DNMT1-

expressing cells (Fig. 2.3A). Bisulfite sequencing indicated that there were few

methylated CpGs in vector cells (1.6%) (Fig. 2.3B). DNMT1 and DNMT1-ΔRFTS

overexpression up-regulated methylation more than ten-fold to 17.6% and 24.6%,

respectively. Although there was no apparent difference between DNMT1 and

DNMT1-ΔRFTS from MeDIP analysis, bisulfite sequencing showed that DNMT1-

ΔRFTS expression drove greater methylation of the DAPK promoter than did wild-type

DNMT1. Furthermore, we analyzed the association between DNMT1 and the DAPK

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promoter region by DNMT1 chromatin immunoprecipitation (ChIP) (Fig. 2.3C). We

found increased chromatin occupancy of DNMT1 on the DAPK promoter region in all

DNMT1-expressing cells.

The transcript levels of DAPK were consistent with its increased promoter

methylation. Compared to the vector control, expression of DAPK was decreased in all

DNMT1-expressing cells, among which DNMT1-ΔRFTS cells showed the greatest

repressive effect (Fig. 2.3D). A similar correlation in DNA methylation, DNMT1

chromatin occupancy and mRNA expression were also found in DUOX1 in all

DNMT1-expressing cells (Fig. 2.3). In agreement with the transformation result,

DNMT1-ΔRFTS overexpression resulted in the greatest effect on hypermethylation-

mediated silencing of DAPK and DUOX1, indicating that RFTS-deleted DNMT1 is a

gain of function mutant. Deletion of the two different regulatory domains of DNMT1

did not change target preference and chromatin occupancy of DNMT1. In DNMT1-

expressing cells, the same target genes were affected with different degree of

methylation, which could be attributed to differences in DNA methyltransferase activity

(Syeda et al. 2011).

2.4.3 Strong alleles of DNMT1 condense chromatin

structure at the DAPK and DUOX1 promoters.

To test whether RFTS and CXXC domains alter the ability of DNMT1 to

condense chromatin structure, we analyzed the sensitivity of the DAPK and DUOX1

promoter regions to DNase treatment (Fig. 2.4). Our data indicate that forced expression

of DNMT1 or DNMT1-ΔRFTS reduced chromatin accessibility at both promoters

compared to vector, with DNMT1-ΔRFTS producing the greatest reduction of DNase

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sensitivity. Thus, silencing of DAPK and DUOX1 is due to DNMT1-dependent

alteration of chromatin, and can be increased by deletion of the inhibitory RFTS

domain.

2.4.4 DNA demethylating agent 5-aza-deoxycytidine (5-

aza-dC) reverses gene silencing and diminishes the

transformation ability of strong DNMT1 alleles.

We have demonstrated that the expressions of DAPK and DUOX1 are inhibited

concurrent with an increase in promoter methylation. We tested whether this

methylation-mediated repression is reversible by treating cells with the DNA

demethylating agent 5-aza-dC and analyzing expressions of DAPK and DUOX1. As

expected, 5-aza-dC treatment significantly increased DAPK and DUOX1 levels in

DNMT1-expressing cells (Fig. 2.5A). Moreover, 5-aza-dC-treated DNMT1 and

DNMT1-ΔRFTS cells completely lost activity in anchorage-independent growth (Fig.

2.5B), indicating that increased methylation-dependent gene regulation is responsible

for the oncogenic properties of DNMT1 and DNMT1-ΔRFTS and it is reversible.

2.4.5 Genome-wide promoter methylation analysis reveals

that DNMT1-ΔRFTS cells produce a methylation pattern

similar to DNMT1 cells, though more intense.

Targeted screening of TSGs indicated that DAPK and DUOX1 promoters are

hypermethylated in all DNMT1-expressing cells. DNA from vector, DNMT1 and

DNMT1-ΔRFTS cells were used in the high-resolution HpaII tiny fragment enrichment

by ligation-mediated PCR (HELP) assay (Khulan 2006). This genomic methylation

array of 720,001 probes was designed to focus on CpG islands near transcription start

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sites. The Pearson Correlation test (Di Croce et al. 2002; Thompson et al. 2008) showed

that the majority of methylation intensity signals was similar in all cell lines (Fig. 2.6).

We then analyzed DNA methylation profiles by pairwise comparisons of DNMT1,

DNMT1-ΔRFTS and vector control cells. As seen in the volcano plots (Fig. 2.7A), there

were hyper- and hypomethylation sequences in both DNMT1-expressing cells, with

DNMT1-ΔRFTS cells exhibiting greater changes. To systematically analyze

methylation targets, we performed Kyoto Encyclopedia of Genes and Genomes

(KEGG) pathway enrichment analysis to examine the top 1,000 hypermethylated or

hypomethylated genes in DNMT1-ΔRFTS cells compared to vector cells (Table 2.3).

Pathway analysis indicated that DNMT1-ΔRFTS drove increased methylation of

specific genes involved in cell adhesion, migration and signaling pathways, which are

highly related to carcinogenesis. In addition, hypomethylated gene targets in DNMT1-

ΔRFTS cells were enriched in lysosomal functions and enzymes in phenylalanine and

tyrosine metabolism.

We generated heat maps to compare methylation differences in all three sample

sets by selecting fragments with more than 4-fold signal changes between DNMT1-

ΔRFTS and vector cells (Fig. 2.7B). This big-data visualization analysis showed that

DNMT1 and DNMT1-ΔRFTS cells share similar methylation targets and that DNMT1-

ΔRFTS cells produces a higher degree of methylation. The DNMT1 and DNMT1-

ΔRFTS cells clustered together, suggesting that the RFTS domain does not control

locus specificity but limits the degree of DNA modification, just as it limits in vitro

activity (Syeda et al. 2011).

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2.4.6 DNMT1-ΔRFTS cells exhibit genomic

hypomethylation.

The epigenetic signature of cancer is regional hypermethylation with global

hypomethylation. We thus analyzed fractional methyl cytosine content by HPLC (Fig.

2.8A) (Leonhardt et al. 1992; Magaña et al. 2008). Previous studies have shown that

highly overexpressed DNMT1 causes genomic hypermethylation (J. Wu et al. 1993;

Bakin and Curran 1999; Biniszkiewicz et al. 2002). However, in our DNMT1 cells in

which ectopic expression is at endogenous levels, there was not a significant increase in

genomic DNA methylation. In contrast, DNMT1-ΔRFTS cells, which displayed the

highest levels of focal hypermethylation, had reduced overall methyl cytosine content in

comparison to vector or other DNMT1-expressing cells. We considered that the main

site of demethylation might occur in Satellite 2 repeat sequences (SAT2) for three

reasons. First, the RFTS domain mediates association of DNMT1 to pericentromeric

heterochromatin to maintain dense methylation (Easwaran et al. 2004; Schneider et al.

2013) and SAT2 is the most abundant repeat in the region (Ting et al. 2011). Second,

SAT2-specific hypomethylation has been found in DNMT1-null cells(Rhee et al. 2000;

Espada 2004) and in patients with RFTS-mutated DNMT1 (Klein et al. 2011). Third,

DNA hypomethylation and RNA up-regulation of SAT2 are highly associated with

various cancers and contributes to genomic instability (Ting et al. 2011). To test

whether SAT2 is hypomethylated in our cell lines, we performed bisulfite sequencing

(Fig. 2.8B and C). SAT2 methylation was significant reduced from 66% (vector) and

63% (DNMT1) to 48% in DNMT1-ΔRFTS cells. Further, we analyzed chromatin

occupancy of DNMT1 on SAT2 loci by DNMT1 ChIP (Fig. 2.8D). The data indicate

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that DNMT1-ΔRFTS expression reduced association between DNMT1 and SAT2 loci.

Unlike what was observed for SAT2, there was no significant methylation change in

LINE1 DNA repeat (Fig. 2.9). Therefore, specific demethylation of SAT2 might be due

to the impaired association between DNMT1-ΔRFTS and the pericentromeric region.

Our results suggest that demethylation of SAT2 and promoter hypomethylation detected

in the HELP assay might both contribute to reduced genomic methylation observed in

DNMT1-ΔRFTS cells. Moreover, we investigated whether SAT2 DNA

hypomethylation is associated with transcription. Indeed, expression of SAT2 non-

coding RNA was increased in DNMT1-ΔRFTS cells, but was not altered in cell lines

with alleles of DNMT1 that are weaker (Fig. 2.8E).

2.4.7 DNMT1-ΔRFTS expression has similar effects in

H358 lung cancer cells.

To rule out a cell-specific effect, we established stable cell lines in H358 cells to

determine the effect of DNMT1-ΔRFTS expression in malignant cells (Fig. 2.10A).

DNMT1-ΔRFTS expression enhanced the proliferation, invasion and soft-agar colony

growth of the cells, while full-length DNMT1 expression behaved similarly to vector

control cells (Fig. 2.10B-D). Moreover, ectopic expression of DNMT1-ΔRFTS slightly

inhibited expressions of DAPK and DUOX1 (Fig. 2.11A). Although DNMT1-ΔRFTS

cells did not express significantly more SAT2 RNA transcripts (Fig. 2.11A), there was a

notable methylation reduction of SAT2 in DNMT1-ΔRFTS cells (Fig. 2.11B and C).

These data suggest that the biological function of DNMT1-ΔRFTS is not cell-specific.

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2.5 Discussion

We proposed that the RFTS domain is DNA-competitive and inhibitory (Syeda

et al. 2011), while other data suggested that the CXXC domain is DNA-competitive and

inhibitory (Song et al. 2011). Here, we hypothesized that expression of hyperactive

DNMT1 lacking an autoinhibitory domain could enhance transformation by altering

DNA methylation. Thus, we modestly expressed full-length and deletion forms of

DNMT1 in immortalized HBEC3 cells to examine their oncogenic potential and

alteration in DNA methylation. Full-length DNMT1 expression triggered cellular

transformation while DNMT1-ΔRFTS functioned as a stronger oncoprotein. The

oncogenic effects of DNMT1-ΔRFTS depended on the presence of the CXXC domain,

which is apparently a positive factor. Deletion of either regulatory domain resulted in

the same apparent target preference; expression of all DNMT1 alleles increased

methylation of DAPK and DUOX1 promoters. DNMT1 and DNMT1-ΔRFTS cells share

similar hypermethylated targets in genome-wide analysis as well, though deletion of

RFTS increased the degree of DNA methylation.

Given previous findings that overexpressed or activated DNMT1 caused non-

specific genomic hypermethylation (J. Wu et al. 1993; Bakin and Curran 1999;

Biniszkiewicz et al. 2002), we surprisingly discovered that DNMT1-ΔRFTS expression

at endogenous levels led to demethylation in SAT2 and in the genome. This finding is

consistent with a previous study on RFTS-mutated DNMT1 (Klein et al. 2011), in

which point mutations in the RFTS domain caused SAT2 and genomic hypomethylation.

The study also showed that mutations in the RFTS domain of DNMT1 impaired binding

with heterochromatin (Klein et al. 2011). We confirmed the impaired association

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between DNMT1 and SAT2 loci in DNMT1-ΔRFTS cells by DNMT1 ChIP. Because

DNMT1 may function as an oligomeric complex (Fellinger et al. 2009), RFTS-deleted

DNMT1 and endogenous DNMT1 may form a heterooligomer that is impaired in

association with pericentromeric heterochromatin. Thus, we suggest a model in which

deletion of the RFTS domain activates DNMT1 for euchromatic DNA-binding, but

decreases chromatin occupancy of DNMT1 to heterochromatic SAT2 loci by virtue of a

missing protein interaction, leading to passive DNA demethylation (Fig. 2.12).

Searching the catalogue of somatic mutation in cancer (COSMIC) database, 26

mutation sites within the RFTS domain of DNMT1 were found (Table 2.4). These

DNMT1 mutants could promote malignancy by increasing DNA methyltransferase

binding and activity on euchromatic DNA while being disadvantaged in pericentromeric

SAT2 association and methylation.

RFTS-targeted DNMT1 associated proteins (RAPs) are likely to participate in

these mechanisms. To our knowledge, there are at least three known RAPs including

ubiquitin-like containing PHD and RING finger domain protein 1 (UHRF1) (Bostick et

al. 2007; Sharif et al. 2007; Bashtrykov, Jankevicius, et al. 2014), ubiquitin-specific-

processing protease 7 (USP7) (Felle et al. 2011) and N-α-acetyltransferase 10 NatA

catalytic subunit (NAA10) (Lee et al. 2010). These proteins have been shown to recruit

DNMT1 to specific loci and stimulate its methylation activity, causing site-specific

hypermethylation. Moreover, these proteins were found up-regulated in lung cancers

(Unoki et al. 2010; Daskalos et al. 2011; Lee et al. 2010), indicating that release of the

RFTS domain inhibition might drive cancer formation via hypermethylayion. If this is

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the case, targeting of these binding partners could be a promising therapeutic strategy to

limit DNMT1-dependent hypermethylation in cancer.

Because of the obvious link between hyperactive DNMT1 and transcriptional

repression of TSGs, DNMT1-mediated DNA hypermethylation is emerging as a crucial

therapeutic target (Laird et al. 1995; M. Suzuki et al. 2004). The current approach is to

inhibit expression or hyperactivity of DNMT1 (Ramchandani et al. 1997; McCabe et al.

2006; Datta et al. 2009). However, demethylating agents lead to unavoidable non-

specific genomic demethylation causing genomic instability or oncogene reactivation

and cause selective opportunities for cancer progression (Szyf 2003; Loriot 2006;

Yaqinuddin et al. 2009; Morey Kinney et al. 2010). Because the RFTS domain

functions as a key regulator of DNMT1 function, targeting RFTS interactions may

revert euchromatin-associated DNMT1 activation while also normalizing

pericentromeric DNA methylation.

In conclusion, our study reveals the functional roles of the RFTS domain of

DNMT1 in maintenance of a nontransformed epigenome. We have demonstrated that

deletion of RFTS enhanced the oncogenic potential of DNMT1 by increasing promoter

methylation of TSGs such as DAPK and DUOX1. However, DNMT1-ΔRFTS also

decreased association of DNMT1 with the pericentromeric region, causing SAT2

demethylation. Because DNMT1-ΔRFTS was able to reprogram the overall methylation

pattern of epithelial cells in a manner that is common in cancer, the data suggest that

RFTS may be a target of tumor-specific dysregulation.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001; ns, indicates no significant difference in comparison to vector cells.

Figure 2.1. Deletion of RFTS enhances the oncogenic activity of DNMT1. (A) HBEC3 stable cell lines were established to express full-length and DNMT1 deletion forms near endogenous DNMT1 levels. The levels of DNMT1 were determined by RT-qPCR (left) and western blotting (right). Data were normalized to vector cells. (B) Adherent colony formation. (C) Soft-agar colony formation.

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*, P < 0.05 in comparison to vector cells. Figure 2.2. Ectopic expression of DNMT1-ΔRFTS enhances invasion activity without proliferation. (A) Neither full-length nor mutant DNMT1 overexpression changed proliferation rates in the presence or absence of EGF. Data were normalized to vector cells cultured with EGF. (B) DNMT1-ΔRFTS and DNMT1-ΔR/C cells showed slightly enhanced invasion. Invasion ability was quantified by the CultureCoat 24 Well Low BME Cell Invasion Assay. Data were normalized to vector cells with n = 4.

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Figure 2.3. DNMT1-ΔRFTS promotes increased methylation and silencing of the DAPK and DUOX1 genes. (A) The methylation levels of DAPK (left) and DUOX1 (right) promoter-associated CpG islands were analyzed by qPCR. Methylated DNA was analyzed using the MethylMiner kit and amplified with specific primers. (B) Bisulfite sequencing results for DAPK (left) and DUOX1 (right) promoters. White squares represent unmethylated cytosines and black squares represent methylated cytosines in CpG sites. The percentage of methylated CpG dinucleotides from 8 independent clones is indicated. (C) DNMT1 chromatin occupancy was analyzed using DNMT1 ChIP and qPCR. (D) mRNA levels of DAPK (left) and DUOX1 (right) were analyzed by RT-qPCR and normalized to vector cells

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison

to vector cells.

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*, p < 0.05; ***, p < 0.001 in comparison to vector cells.

Figure 2.4. DNMT1-ΔRFTS decreases chromatin accessibility at DAPK and DUOX1 promoters. Cells were treated with or without DNA nuclease for 1 hr, prior to detection of promoter DNA by qPCR. The index of chromatin accessibility = 2 ((Ct DNase

treated)-(Ct Untreated)).

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***, p < 0.001 in comparison to the DMSO treated control. Figure 2.5. 5-aza-dC treatment reactivates TSG expression and suppresses DNMT1-dependent transformation. (A) mRNA levels of DAPK (left) and DUOX1 (right) were analyzed by RT-qPCR after 100nM 5-aza-dC treatment for 5 days and normalized to vector cells treated with DMSO. (B) Soft-agar colony formation after 5-aza-dC treatment.

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Figure 2.6. Ectopic expression of DNMT1 alleles does not radically alter global methylation intensities. Pairwise unsupervised clustering followed by Pearson correlations of normalized ratios from HELP assay indicated that the majority of methylation intensities are little changed in DNMT1-expressing cells in comparison to vector cells. DNMT1 and DNMT1-ΔRFTS cells shared more similarity than vector cells.

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Figure 2.7. DNMT1-ΔRFTS expression enhances global DNMT1 methylation changes. (A) Genome-wide promoter DNA methylation profiles were obtained using the HELP assay. Volcano plots are the x-axis scores probe-specific methylation ratios and the y-axis scores p-values for the confidence of measurements. The plots allow visualization of methylation differences between vector and DNMT1 cells as well as the differences between vector and DNMT1-ΔRFTS cells. Probes sets that showed significant hyper- or hypomethylation (p < 0.05 for methylation changes (log2(HpaII/MspI)) > 2) are shown in cyan. All other probes are shown in red. (B) Heat map illustration of HpaII-enrichment fragments with methylation changes (log2(HapII/MspI)) > 2 between vector and DNMT1-ΔRFTS cells.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to vector cells.

Figure 2.8. Genomic hypomethylation is found in DNMT1-ΔRFTS cells. (A) 5-methylcytosine (mC) content of the total cytosine pool was determined by HPLC. (B) Bisulfite sequencing of SAT2. White squares represent unmethylated CpGs, black squares represent methylated CpGs, and grey squares represent undetermined sites. Each row is an independent sequencing result. (C) Quantitation of SAT2 bisulfite sequencing. (D) DNMT1 chromatin occupancy was analyzed using DNMT1 ChIP and qPCR. (E) Expression of SAT2 non-coding RNA was analyzed by RT-qPCR and normalized to vector cells.

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Figure 2.9. The methylation levels of LINE1 were not changed in DNMT1 or DNMT1-ΔRFTS cells. (A) Bisulfite sequencing of LINE1. (B) Quantitation of LINE1 bisulfite sequencing. n = 20. ns, indicates no significant difference in comparison to vector cells.

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*, P < 0.05; ***, p < 0.001 in comparison to vector cells.

Figure 2.10. Ectopic expression of DNMT1-ΔRFTS in H358 cells is sufficient to enhance proliferation, invasion and soft-agar colony formation. (A) H358 stable cell lines were established to express full-length DNMT1 or DNMT1-ΔRFTS near the endogenous DNMT1 levels. The levels of DNMT1 were determined by western blotting. (B) Both DNMT1 and DNMT1-ΔRFTS increased the proliferation rate in H358 cells. Data were normalized to vector cells. (C) DNMT1-ΔRFTS showed slightly enhanced invasion. Invasion ability was quantified by the CultureCoat 24 Well Low BME Cell Invasion Assay. Data were normalized to vector cells. n = 4. (D) DNMT1-ΔRFTS cells showed the greatest colony formation in soft agar.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to vector cells.

Figure 2.11. Ectopic expression of DNMT1-ΔRFTS in H358 cells caused gene silencing of DAPK and DUOX1 and demethylation of SAT2. (A) Expressions of DAPK, DUOX1 and SAT2 were analyzed using RT-qPCR. (B) Bisulfite sequencing of SAT2. (C) Quantitation of SAT2 bisulfite sequencing. n = 20.

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Figure 2.12. Dual roles for RFTS domain in DNMT1-dependent DNA methylation. (A) RFTS-targeted DNMT1 associated proteins (RAP) are proposed to relieve inhibition of DNMT1 for access to euchromatin. (B) The RFTS domain mediates association between DNMT1 and pericentromeric heterochromatin. (C) In cancer, overexpression of RAPs or mutation of RFTS is proposed to relieve DNMT1 inhibition, thereby increasing methylation and silencing of TSGs. However, because the RFTS domain is required for association with heterochromatic SAT2 sequences, DNMT1 with mutant RFTS may be less associated with such sequences, accounting for global hypomethylation.

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Table 2.1. Target list of TSGs have been found with hypermethylation-mediated gene silencing in lung cancers Function Gene Function Gene

Cell cycle P16

Cell adhesion

CDH1

CDH13

Growth/ Differentiation

APC TIMP3

RARß TSLC1

DUOX1 LAMA3

DUOX2 RECK

IGFBP3

GATA4

Apoptosis

DAPK

WWOX RASSF1A

MTHFR FHIT

FAS

DNA repair MGMT NORE1A

BCL2

Detoxification GSTP1 SEMA3B

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Table 2.2. Summary of the changes of promoter methylation and gene expression in DNMT1-expressing cell lines. Cell lines DNMT1 DNMT1-

ΔRFTS DNMT1-ΔCXXC

DNMT1-ΔR/C

Promoter hypermethylation with gene silencing

DAPK DAPK DAPK DAPK DUOX1 DUOX1 DUOX1 DUOX1

Promoter hypermethylation without gene silencing

RASSF1A RASSF1A RASSF1A RASSF1A DUOX2 DUOX2 DUOX2 DUOX2 GATA4 GATA4 GATA4

Gene silencing without promoter hypermethylation

FHIT FHIT FHIT FHIT MTHFR MTHFR MTHFR MTHFR

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Table 2.3. KEGG pathway enrichment analysis

Pathway Gene Counts p value Hyper-

Focal Adhesion 31 9.12E-07 Fructose and mannose metabolism 11 1.28E-05 Leukocyte transendothelial migration 19 1.13E-04 Tight junction 19 5.73E-04 ECM-receptor interaction 11 0.021146166 Neurotrophin signaling pathway 14 0.024730832 Pathways in cancer 28 0.034601812

Hypo- Lysosome 13 0.024611161 Phenylalanine metabolism 5 0.028973119 Tyrosine metabolism 7 0.030538571

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Table 2.4. DNMT1 RFTS domain mutations were found in cancer (COSMIC database)

Cancer Type Mutation Position Conservation

Melanoma P351L P

Stomach K352E K (Q in Rat)

Uterine Q358R Q

Melanoma L365F L

Melanoma D373N D

Lung squamous T382R T

Bladder I388M I (V in Rat)

Stomach H416Y R in Rat, Mouse and Sheep

Melanoma P421L P

HEC251 (endometrial) D423N D

Stomach E428K E

Colorectal E432K E

22RV1 (Prostate Cancer) L433F L

Bladder D470N D

Uterine F479L F

Uterine/ Colorectal E485K E

Melanoma E504K E

Colorectal Q517P Q

Ovary T523A T (V in Mouse and Rat)

Melanoma I531R I

Esophagus T534M T

HS994T (Skin Cancer) S549F S

Colorectal A554T/P A

Head & Neck E559Q E (S in Mouse and Rat)

Esophagus E566K E

Prostate D583N D (A in Mouse S in Rat)

SW620 (Large Intestine) L587M L

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Table 2.5. Primer list Forward Reverse Plasmid construct TA cloning CACCATGCATCATCATCATCATCATCCGGC

GCGTACCGCCCCA GTCGACCTAGTCCTTAGCAGCTTCCT

RFTS deletion ACCAAGCTGGTCTACCAGATC AGGGTGGGTCTTGGAGTTCATG

CXXC deletion TCACCCAAAAAAATGCACCAG GTTCTCCTTGTCTTCTCTGTC

R/C deletion TCACCCAAAAAAATGCACCAG AGGGTGGGTCTTGGAGTTCATG

RNA DNMT1 GAGCTACCACGCAGACATCA CGAGGAAGTAGAAGCGGTTG

DAPK CTCCCCATTTCTTGGAGACA CCAGGGATGCTGCAAACTAT

DUOX1 ATGTGCCAGATACCCAAAGC CAGCTGACGGATGACTTGAA

SAT2 CATCGAATGGAAATGAAAGGAGTC ACCATTGGATGATTGCAGTCAA

GAPDH GAGTCAACGGATTTGGTCGT GACAAGCTTCCCGTTCTCAG

MeDIP/ ChIP DAPK GCTTTTGCTTTCCCAGCCAGGGC ATCGCACTTCTCCCCGAAGCCAA

DUOX1 CCATGGGACTTGTGAAGGCGGAC CTCCCGGGGCGCAGGTAGAG

SAT2 CATCGAATGGAAATGAAAGGAGTC ACCATTGGATGATTGCAGTCAA

Bisulfite sequencing DAPK 1st PCR TTTTATTTATTTTTTAGTTGTGTTTT TAAAAACAATCTCTCTCCAACCTAC

DAPK 2nd PCR TTAGTTTTTGTTTTTTTAGTTAGGG AACAATCCCCAAAACCACAT

DUOX1 1st PCR GGTTTTGGATTTGGAGTTTAGATT AAAAAACTAACATTCCCCTTTCTTC

DUOX1 2nd PCR GTTTTATGGGATTTGTGAAGG CTACCCTTAAAACTCCCTCCC

SAT2 1st PCR TGGAATTATTATTAAATGGTAATTTAATGG AAATAATTACAATCAATTCATTC

SAT2 2nd PCR TTAATGGAAAGGAATGGAAT TTCCATTAAATAATAACTCC

LINE1 TTATTAGGGAGTGTTAGATAGTGGG TACCTAAACAAACCTAAACAATAAC

Chromatin accessibility DAPK GTGGGTGTGGGGCGAGTGGGTGT CTCTCGGCTCCTTGCCGCCTTTT

DUOX1 CGCCTCCCACCCTCTCCCCAGCC GGCCCGCGGAGCCCTCTCTC

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CHAPTER III

SUPPRESSION OF TET1-DEPENDENT DNA

DEMETHYLATION IS ESSENTIAL FOR KRAS-

MEDIATED TRANSFORMATION

3.1 Abstract

Hypermethylation-mediated tumor suppressor gene (TSG) silencing is a central

epigenetic alteration in RAS-dependent tumorigenesis. Ten-eleven translocation (TET)

enzymes can depress DNA methylation by hydroxylation of 5-methylcytosine (5mC)

bases to 5-hydroxymethylcytosine (5hmC). Here we report that suppression of TET1 is

responsible for KRAS-induced DNA hypermethylation and cellular transformation. In

two non-malignant cell lines, HBEC3 and NIH3T3, oncogenic KRAS promotes

transformation by inhibiting TET1 expression via the ERK signaling pathway. This

reduces chromatin occupancy of TET1 at TSG promoters, reduces levels of 5hmC, and

increases levels of 5mC and transcriptional silencing. Restoration of TET1 expression

by ERK pathway inhibition or ectopic TET1 reintroduction in KRAS-transformed cells

reactivates TSGs and inhibits colony formation. Additionally, H1299 cancer cells

require persistent TET1 suppression by KRAS to maintain malignancy. KRAS

knockdown increases TET1 expression and diminishes colony-forming ability, while

KRAS/TET1 double knockdown bypasses KRAS dependency. Thus, suppression of

TET1-dependent DNA demethylation is critical for KRAS-mediated transformation.

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3.2 Introduction

RAS proteins are a family of 21 kDa proteins that accomplish signal

transduction by coupling receptor engagement to downstream pathway activation

(Downward 2003; Pylayeva-Gupta, Grabocka, and Bar-Sagi 2011). RAS proteins,

which include KRAS, HRAS and NRAS, share similar functions in regulating cell

proliferation, differentiation and survival. Gain-of-function mutations in RAS genes are

found frequently in malignancies (Jones and Baylin 2002; Pylayeva-Gupta, Grabocka,

and Bar-Sagi 2011; Belinsky 2004; D’Arcangelo and Cappuzzo 2012), and multiple

malignancies depend on RAS mutations to maintain malignant phenotypes (Bestor

2000; Chin et al. 1999). Hyperactive RAS drives constitutive signaling through the

RAF-MEK-ERK and PI3K-AKT cascades (J. Wu et al. 1993; Schubbert, Shannon, and

Bollag 2007; Belinsky et al. 1996) driving cellular transformation (Laird et al. 1995;

Greig et al. 1985; M. Suzuki et al. 2004). Accordingly, targeting RAS-related signaling

pathways is a central goal of molecular oncology (Pylayeva-Gupta, Grabocka, and Bar-

Sagi 2011; Downward 2003).

Cytosine methylation of CpG dinucleotides is an epigenetic modification that

cells use to regulate gene expression, largely to promote transcriptional silencing. Focal

hypermethylation of tumor suppressor genes (TSGs) accompanied by genomic

hypomethylation are epigenetic hallmarks of malignancy (Pylayeva-Gupta, Grabocka,

and Bar-Sagi 2011; Jones and Baylin 2002; D’Arcangelo and Cappuzzo 2012; Belinsky

2004). Three DNA methyltransferases (DNMTs), the de novo enzymes DNMT3A and

DNMT3B and the maintenance enzyme DNMT1, are responsible for establishment and

maintenance of DNA methylation patterns (Chin et al. 1999; Bestor 2000). Aberrant

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overexpression of DNMTs contributes to cancer-associated DNA hypermethylation

(Schubbert, Shannon, and Bollag 2007; J. Wu et al. 1993; Belinsky et al. 1996).

Inhibition of DNMTs in cancers can revert DNA hypermethylation, reactivate silenced

TSGs and diminish tumorigenicity (Greig et al. 1985; Laird et al. 1995; M. Suzuki et al.

2004), indicating that DNA methylation is reversible by modulating DNMT activities.

Previous studies showed that RAS-driven transformation drives methylation-

associated silencing of TSGs to inhibit apoptosis and promote cell proliferation

(Borrello et al. 1987; Patra 2008; Gazin, Wajapeyee, Gobeil, Virbasius, and Green

2007b; Serra et al. 2014). RAS activation was shown to trigger DNA hypermethylation

through elevated DNMT transcription (Pruitt et al. 2005; Chang, Cho, and Hung 2006;

Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007b; Bakin and Curran 1999) and

inhibition of DNMT expression has been shown to be sufficient to reverse RAS-induced

hypermethylation and transformation (MacLeod and Szyf 1995; Ramchandani et al.

1997). Thus, DNMT enzymes have been considered the principal mediators of DNA

methylation driven by RAS activation and have been targeted by early stage drug

discovery efforts (Fagan, Wu, et al. 2013; Fagan, Cryderman, et al. 2013; J. Huang et al.

2013).

Recent findings demonstrated that the ten-eleven translocation (TET) family

proteins, including TET1, TET2 and TET3, function as iron and α-ketoglutarate-

dependent 5-methylcytosine dioxygenases that convert 5-methylcytosine (5mC) bases

to 5-hydroxymethylcytosine (5hmC) bases (Tahiliani et al. 2009; Ito et al. 2010). 5hmC

is proposed as an intermediate in passive and active DNA demethylation (Kohli and

Zhang 2013; Pastor, Aravind, and Rao 2013; S. C. Wu and Zhang 2010; H. Wu and

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Zhang 2014), suggesting novel mechanisms to regulate methylation dynamics and gene

reactivation. Presence of 5hmC in genomic DNA impairs maintenance methylation by

preventing DNMT1 recognition (Valinluck and Sowers 2007; Hashimoto et al. 2012),

thereby facilitating passive demethylation linked to DNA replication. In addition, 5hmC

can be further converted by TET proteins to 5-formylcytosine (5fC) and 5-

carboxycytosine (5caC) (Ito et al. 2011), which are replaced by cytosine through DNA

repair processes (Cortellino et al. 2011; He et al. 2011). TET-mediated active

demethylation is independent of DNA replication (Pastor, Aravind, and Rao 2013; S. C.

Wu and Zhang 2010). TET proteins and 5hmC modifications are abundant in mouse

embryonic stem cells (ESC) (Ito et al. 2010; Ficz et al. 2012; Koh et al. 2011) and in the

brain (Kriaucionis and Heintz 2009; J. U. Guo et al. 2011; Kaas et al. 2013). In addition

to the roles of TET-driven DNA modification in ESC and neuronal systems, emerging

evidence suggests that TET-dependent DNA demethylation plays a role in

tumorigenesis. In solid tumors, expression of TET genes is dramatically reduced and is

highly associated with reduced 5hmC (Lian et al. 2012; Yang et al. 2013; Ko et al.

2010) and hypermethylation-mediated silencing of TSGs (Hsu et al. 2012; M. Sun et al.

2013). These data suggest that TET genes themselves may have TSG activity. However,

whether TET-mediated DNA demethylation plays a role in RAS-induced DNA

hypermethylation and malignant transformation remains unclear.

In this study, we used two non-malignant cell lines to dissect KRAS-driven

transformation and the establishment of cancer-associated DNA hypermethylation.

Unexpectedly, instead of an increase of DNMT expression, we discovered that TET1 is

transcriptional suppressed via the RAS-ERK signaling pathway. Regional decreases in

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5hmC were accompanied by TSG promoter hypermethylation and gene silencing.

Forced TET1 reintroduction not only reactivated silenced TSGs but also abolished

KRAS-induced colony-forming ability. Moreover, KRAS depletion by small interfering

RNA (siRNA) up-regulated TET1 expression in cancer cells. Strikingly, knocking down

TET1 restores colony-forming ability to KRAS depleted cells, indicating that persistent

TET1 suppression by KRAS is required to maintain the malignant phenotype and that

TET1 suppression is sufficient to carry our KRAS transformation several steps

downstream from KRAS. These data establish that impaired TET1-mediated DNA

demethylation is a critical mediator of tumor initiation and maintenance in KRAS-

transformed cells.

3.3 Material and Methods

3.3.1 Cell culture

HBEC3 cells and stable cell lines were grown in KSFM media supplemented

with bovine pituitary extract and recombinant human EGF unless specific indicated.

NIH3T3 cells (CRL-1658, ATCC), Kras-transformed NIH3T3 cells (CRL-6361,

ATCC) and HepG2 cells were grown in DMEM media with 10% FBS. H1299 cells

were grown in RPMI-1640 media with 10% FBS.

3.3.2 Establishment of stable cell lines

To establish oncogenic KRAS-expressing stable lines in HBEC3 cells, a full-length

human KRAS-G12V cDNA clone (gift of Dr. John Minna) was used as template to

generate a KRAS-G12V construct with an N-terminal myc-tag. For transient TET1

reintroduction, a catalytic domain of human TET1 cDNA clone (plasmid 39454,

Addgene) (J. U. Guo et al. 2011) was used as template to generate a TET1 construct

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with an N-terminal myc-tag. PCR fragments were first T/A cloned into pGEM-Teasy

vector (Promega) and then subcloned into pLenti6/V5 vector (Invitrogen). Viral

production and transduction was performed using ViralPower Bsd Lentiviral Support

Kit (Invitrogen). Monoclonal cell lines were selected by serial dilution in 96-well plates

with 5 µg/ml Blasticidin (Invitrogen). Primer pairs used for plasmid construction are

provided in Table 3.3

3.3.3 RT-qPCR

Total RNA was extracted using the RNeasy Mini Kit (Qiagen) with DNase treatment

(Qiagen) to eliminate DNA contamination. Equal amounts of RNA were reverse

transcribed to generate cDNA using an iScript cDNA Synthesis Kit (Bio-Rad). Specific

primer pairs were then used to amplify target genes (Table 3.3 and 3.4). qPCR reactions

were conducted with iQ SYBR Green Supermix (Bio-Rad). All data were collected

from 3 or 4 independent experiments.

3.3.4 Immunoblotting

Protein extracts from each stable cell lines were prepared in RIPA buffer (Thermo

Scientific) according to the manufacturer's instructions. Equal amounts of protein were

separated using NuPAGE® Novex® 4-12% Bis-Tris Gel and transferred to 0.2 µm

nitrocellulose membrane at 4 °C overnight. Proteins were detected using specific

antibodies and visualized by SuperSignal West Femto Substrate (Thermo Scientific).

Primary antibodies include TET1 (09-872, Millipore); DNMT1 (WH0001786M1,

Sigma); RAS (05-1072, Millipore); myc Tag (05-724, Millipore); actin (ab3280,

Abcam); Phospho-AKT (4060, Cell Signaling); Total-AKT (9272, Cell Signaling);

Phospho-ERK (9101, Cell Signaling) and Total-ERK (9102, Cell Signaling). Secondary

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antibodies are goat anti-rabbit IgG (Thermo Scientific) and goat anti-mouse IgG

(Thermo Scientific).

3.3.5 Proliferation assay

1,000 cells were seeded in replicates of 6 in KSFM media with or without EGF

supplementation in 96-well plates. Relative cell numbers were analyzed using

Resazurin (Sigma) 72 hrs after seeding. All data were collected from 4 independent

experiments.

3.3.6 Adherent and soft-agar colony formation

For adherent colony formation, 50 (HBEC3) or 200 (NIH3T3 and H1299) cells were

seeded on 6-well plates, allowed to grow for 9 (NIH3T3), 10 (H1299) or 12 (HBEC3)

days, followed by 4% methylene blue (Sigma) staining. Colony size > 2 mm were

counted. For soft-agar colony formation, 10,000 cells were resuspended in media with

0.4% agarose and plated over a layer of 0.6% agarose. Cells were incubated at 37 °C for

3 (NIH3T3 and H1299) or 4 (HBEC3) weeks and colonies were stained with MTT

(Sigma). Colony images were acquired with ChemiDoc XRS (Bio-Rad) and quantified

using Quantity One software (Bio-Rad). All data were collected from 2 or 3

independent experiments, each in triplicate.

3.3.7 DNA dot blot assays

For global 5mC and 5hmC levels, DNA dot blots were performed with a 96-well

manifold. Genomic DNA was extracted using DNeasy Blood & Tissue Kit (Qiagen). 1

µg genomic DNA and serial 2-fold dilutions were mixed with 0.4 M NaOH, 10 mM

EDTA and denatured at 100°C for 10 min. Samples were then chilled on ice and

neutralized with an equal volume of 2 M ammonium acetate pH 7.0 and loaded onto a

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20X SSC rinsed Hybond-ECL nitrocellulose membrane. 5mC and 5hmC were detected

using specific antibodies (5mC, 39769, Active motif; 5hmC, BI-MECY, Eurogentec)

and visualized by SuperSignal West Femto Substrate (Thermo Scientific).

3.3.8 MeDIP and hMeDIP

Promoter methylation analysis was performed using MethylMiner Methylated DNA

Enrichment Kit (Invitrogen) and promoter hydroxymethylation analysis was performed

using HydroxyMethyl Collector (Active Motif). Genomic DNA was first fragmented by

sonication to an average size of 400 bp. Methylated DNA or hydroxymethylated DNA

was captured and eluated following the manufacturers’ protocols. 5mC and 5hmC levels

were analyzed using specific primer sets with qPCR (Table 3.3 and 3.4). 10% of input

DNA was used as a control. All data were collected from 3 independent experiments.

3.3.9 Bisulfite sequencing

For 5mC detection, genomic DNA was treated with bisulfite using EpiTect Bisulfite kit

(Qiagen). Bisulfite treated DNA was then used as a template and PCR was performed

using specific primer pairs (Table 3.3 and 3.4). Final PCR products were gel purified

and cloned into the pGEM-T easy vector. Independent clones were subjected to

sequencing. For 5hmC detection, genomic DNA was applied to 5hmC TAB-Seq Kit

(WiseGene) following the manufacturer protocol prior to bisulfite coversion.

3.3.10 ChIP

ChIP was performed with Magna ChIP HiSens chromatin immunoprecipitation kit

(Millipore), TET1 antibody (09-872, Millipore), and analyzed using qPCR (Table 3.3

and 3.4). 10% of input DNA was used as a control. All data were collected from 3

independent experiments.

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3.3.11 siRNA transfection

Cells were transfected with 10 nM siRNA using the Lipofectamine RNAiMAX Reagent

(Invitrogen). siRNAs were purchased from PreDesigned Oligo Sets (Integrated DNA

Technologies), including siControl (DS NC1); siKRAS-1 (N004985.12.3); siKRAS-2

(N004985.12.5); siTET1-1 (N030625.12.1) and siTET1-2 (N030625.12.2).

3.3.12 Statistical analysis

All data were presented as mean ± SD. Paired Student’s t tests or one-way ANOVA was

used to calculate P-value and determine significance. P-values below 0.05 were

considered statistically significant.

3.4 Results

3.4.1 Oncogenic KRAS expression is sufficient to

transform non-malignant HBEC3 cells

Expression of KRAS-G12V has the ability to transform a broad spectrum of

non-malignant cells (Pylayeva-Gupta, Grabocka, and Bar-Sagi 2011; Patra 2008).

However, a previous report showed that overexpression of KRAS-G12V was

insufficient to transform immortalized human bronchial epithelial cells (HBEC3),

apparently due to lack of induction of downstream signals (Sato et al. 2006). To probe

the biological effect of oncogenic KRAS in HBEC3 cells, we established stable cell

lines with KRAS-G12V marked by an N-terminal myc-tag. After serial dilution to

select monoclonal cell lines, three KRAS clones (R1, R2 and R3) and two vector

control clones (V1 and V2) were selected and examined by western blot (Figure 3.1A).

In R1, R2 and R3, expression of myc-KRAS was about 30% of the level of endogenous

RAS proteins. However, as shown in Figure 3.1A, expression of KRAS-G12V was

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associated with activation of AKT and ERK as evidenced by a 2-fold induction of

phospho-AKT and 6-fold induction of phospho-ERK.

We found a 23% increase in cell proliferation in KRAS cells (Figure 3.1B).

Additionally, because KRAS is an effector of epidermal growth factor (EGF) receptor

signaling (Yarden and Sliwkowski 2001; Sharma et al. 2007), we considered whether

expression of hyperactive KRAS could enable bypass of EGF-dependent growth of

HBEC3 cells (Sato et al. 2006) (Figure 3.1B). Without EGF supplementation, vector

cells lost half their proliferation ability. However, KRAS cell lines without EGF

supplementation showed the same extent of proliferation as vector cells with EGF,

indicating a KRAS-mediated bypass. To further evaluate the oncogenic properties of

KRAS cells, adherent and soft-agar colony formation was assessed. As shown in Figure

3.1C, adherent colony formation was increased 6-fold in KRAS cells while soft-agar

colony formation in the presence of EGF was increased to more than 100-fold. Without

EGF supplementation, KRAS cells produced more than 10 colonies while vector cells

produced none. In summary, HBEC3 cells can be used to dissect hyperproliferation,

EGF-independence and colony formation that are driven by KRAS mutation.

3.4.2 Oncogenic KRAS expression causes

hypermethylation-mediated silencing of TSGs and loss of

imprinting

Aberrant DNA methylation is a hallmark of cancer and RAS activation has been

shown to drive DNA hypermethylation during tumorigenesis (Pruitt et al. 2005; Chang,

Cho, and Hung 2006; Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007b; Bakin

and Curran 1999). Although there was no increase in 5mC content in KRAS-

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transformed cells (Figure 3.2A), we surveyed 24 TSGs reported to be silenced by

promoter hypermethylation in lung cancers (Belinsky 2004) (Table 3.1 and 3.2) by

quantitative methylated DNA immunoprecipitation (MeDIP). An increase in promoter

methylation was found in five of the 24 TSGs in KRAS cells, including DAPK (D. H.

Kim et al. 2001), MGMT (Pulling et al. 2003), DUOX1 (Luxen, Belinsky, and Knaus

2008), TIMP3 (Bachman et al. 1999) and GATA4 (M. Guo et al. 2004) (Figure 3.2B and

3.3A). Bisulfite sequencing indicated 2 to 20-fold methylation increases in the

promoters of DAPK, MGMT and DUOX1 in R2 cells in comparison to V1 cells (Figure

3.2C), demonstrating that KRAS activation caused DNA hypermethylation of specific

TSGs. Because promoter hypermethylation is highly associated with transcriptional

silencing, we analyzed expression of the five target genes. As shown in Figure 3.2D and

3.3B, the mRNA level of all five genes was markedly decreased in KRAS cells.

In addition to hypermethylation of TSGs, loss of imprinting is an additional type

of dysregulated methylation in malignancies. We focused on the well-studied H19

imprinting control region (H19 ICR) (Steenman et al. 1994) to examine the methylation

change associated with KRAS activation. Bisulfite sequencing indicated that the

methylation level of H19 ICR was increased from 40.7% in V1 cells to 65.9% in R2

cells (Figure 3.3C). Hypermethylation of H19 ICR was accompanied by silenced H19

and activated IGF2 expression (Figure 3.3D).

To test whether promoter hypermethylation was sufficient to suppress gene

expression and whether methylation-associated gene silencing was reversible, we

treated cells with the demethylating reagent, 5-aza-deoxycytidine (5-aza-dC) (Jones et

al. 1982). As shown in Figure 3.2E and 3.3E, 5-aza-dC reactivated expression of all five

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TSGs and reverted expression of H19 and IGF2, indicating that transcriptional silencing

is driven by promoter hypermethylation and is reversible. In addition, 5-aza-dC

treatment decimated colony formation in KRAS-transformed cells compared to DMSO

treatment (Figure 3.2F). Thus, HBEC3 cellular transformation depends upon an altered

methylation status that is commonly found in human cancers.

3.4.3 KRAS negatively regulates TET1 expression through

the ERK signaling pathway

DNMT enzymes, especially DNMT1, have been considered to be the major

effectors of RAS-induced hypermethylation in various cells (Patra 2008; Gazin,

Wajapeyee, Gobeil, Virbasius, and Green 2007b). Thus, we tested whether levels of

DNMT1 were increased in KRAS cells. However, we did not observe any difference of

DNMT1 expression between vector and KRAS cells at the mRNA or protein levels

(Figure 3.4A). We further examined the other two DNA methyltransferases, DNMT3A

and DNMT3B (Figure 3.5A). Although there was a slight decrease in DNMT3B

expression in KRAS cells, this cannot be linked to promoter hypermethylation. In

addition to activation of DNA methyltransferase, another possible mechanism to cause

hypermethylation is suppression of enzymes that act on 5mC substrates, such as TET1,

TET2 and TET3. As shown in Figure 3.4A and 3.5A, KRAS activation nearly

extinguished expression of TET1 at the mRNA and protein levels. No change was

observed in TET2 and TET3 expression.

RAS activation drives two major protein kinase cascades, namely the

PI3K/AKT and RAF/MEK/ERK cascades. To dissect the mediator of TET1

extinguishment by KRAS, we used specific inhibitors of PI3K and MEK. Because these

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signals are essential for cell survival, we used low doses, i.e., 2 µM PI3K inhibitor

LY294002 (Vlahos et al. 1994) or 30 µM MEK inhibitor PD98059 (Dudley et al. 1995)

to titrate KRAS signaling without reducing cell viability. As shown in Figure 3.4 B,

TET1 expression in KRAS cells treated with the MEK inhibitor was restored to the

same level as in vector cells. However, no effect was observed after partial inhibition of

PI3K. Moreover, DNMT1 expression was not affected by either inhibitor (Figure 3.4B).

Remarkably, ERK pathway inhibition caused up to 3-fold transcriptional increases of

DAPK, MGMT, DUOX1 and H19 in KRAS cells (Figure 3.4C and 3.5B). Because

epigenetic silencing of TSGs is essential for KRAS-mediated transformation in HBEC3

cells, we tested whether KRAS-mediated transformation was also regulated by one or

the other kinase cascade. KRAS cells treated with PD98059 or LY294002 for 6 days

were subjected to adherent and soft-agar colony-forming assays. As shown in Figure

3.4D, ERK pathway inhibition significantly reduced colony-forming abilities of KRAS

cells, while AKT pathway inhibition had no effect. Together, our data indicate that

KRAS decreases TET1 transcription and promotes cellular transformation through the

ERK pathway.

3.4.4 Reduction of TET1 and 5hmC are responsible for

KRAS-mediated DNA hypermethylation and cellular

transformation

To clarify the consequence of TET1 reduction in KRAS cells, we examined 5hmC

levels in the genome. Though there was no dramatic change in genomic 5hmC in vector

and KRAS cells (Figure 3.6A), we used 5-hydroxymethylcytosine DNA

immunoprecipitation (hMeDIP) to discover a 2 to 4-fold decrease in 5hmC in promoter

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regions of the five TSGs and H19 ICR that are hypermethylated by mutant KRAS

expression (Figure 3.6B and 3.7A). Because traditional bisulfite sequencing cannot

distinguish 5mC and 5hmC (Y. Huang et al. 2010), we used Tet-assisted bisulfite

sequencing (TAB-seq) (M. Yu et al. 2012) to identify specific 5hmC modifications in

V1 and R2 cells. As shown in Figure 3.6C, 5hmC modifications were decreased from

8.1% (V1) to 4.5 % (R2) in the DAPK promoter, 9.8% (V1) to 3.9% (R2) in the MGMT

promoter and 9.2% (V1) to 4.1% (R2) in the DUOX1 promoter, respectively.

Given the finding that KRAS activation inhibits TET1 expression, the decrease

of 5hmC in targeted genes might be due to reduced chromatin association with TET1.

By TET1 chromatin immunoprecipitation (ChIP), we found that TET1 chromatin

occupancy was reduced at the examined promoters in all KRAS cell lines (Figure 3.6D

and 3.7B). To test whether loss of TET1 was responsible for gene silencing and cellular

transformation observed in KRAS cells, we reintroduced TET1 expression in KRAS

cell lines. As shown in Figure 3.6E and 3.7C, ectopic expression of the catalytic domain

of human TET1 (aa 1418-2136) (J. U. Guo et al. 2011) was sufficient to reactivate

KRAS-mediated gene silencing. Moreover, as shown in Figure 3.6F, restoration of

TET1 expression also suppressed KRAS-mediated transformation. Thus, TET1

suppression is required to maintain TSG silencing and transformation in KRAS cells.

3.4.5 Loss of Tet1 expression is associated with decreased

5hmC and increased 5mC content in Kras-transformed

NIH3T3 Cells

Previous work showed that oncogenic Kras expression caused methylation-

mediated silencing of TSGs in NIH3T3 mouse fibroblast cells in a manner that depend

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on Dnmt1, Erk and other positively acting factors (Ramchandani et al. 1997; McCabe et

al. 2006; Datta et al. 2009). We hypothesized that suppression of Tet1-mediated DNA

modifications might underlie Kras-driven hypermethylation in this system. As shown in

Figure 3.8A and 3.9A, Dnmt1 was increased 2-fold in oncogenic Kras-transformed

NIH3T3 (Kras) cells. In contrast, Tet1 was decreased 2-fold while Tet2 and Tet3 were

also modestly down-regulated in Kras cells. At the genome level, Kras activation

resulted in a nearly 2-fold increase in 5mC accompanied by a 30% decrease of 5hmC

levels (Figure 3.8B). Kras-dependent hypermethylation and silencing in NIH3T3 cells

includes Fas, Sfrp1 and Lox (Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007b).

As shown in Figure 3.8C and 3.9A, the mRNA expression of these genes was nearly

extinguished by Kras activation. To gain further insight into the dynamics of 5mC and

5hmC, we compared 5mC and 5hmC content in promoter regions in parallel. Our data

showed intense methylation increases from 0% 5mC to 80% 5mC concomitant with a 4-

fold 5hmC decrease in Kras cells compared to NIH3T3 cells (Figure 3.8D, 3.9B and

3.9C). As shown in Figure 3.8E and 3.9D, bisulfite sequencing indicated that there were

few or no 5mC modifications in NIH3T3 cells while Ras activation up-regulated

methylation to greater than 70% in the examined promoters. Increases in 5mC were

accompanied by up to 3-fold reduction in 5hmC in Kras cells. Strikingly, all 7

interrogated CpG sties in the Fas promoter were 95-100% in the 5hmC state in NIH3T3

cells. Upon Kras transformation, these CpG sites were converted to 50-100% 5mC.

These data indicate that NIH3T3 cells employ a strong Tet-dependent DNA

modification activity to maintain TSG promoters at low methylation status. Consistent

with this interpretation, Tet1 is highly associated with Fas, Sfrp1 and Lox promoters in

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NIH3T3 cells and is largely evacuated from them in Kras-transformed NIH3T3 cells

(Figure 3.8F and 3.9E).

As shown in Figure 3.10A and 3.11A, Erk pathway activity is required for

down-regulation of Tet1 in Kras-transformed NIH3T3 cells. Erk inhibition reactivated

silenced TSGs (Figure 3.10B) and reduced colony formation (Figure 3.11B and 3.10C),

while Akt inhibition showed no significant changes (Figure 3.10B and 3.10C).

Reintroduction of TET1 expression was also sufficient to increase expression of Fas,

Sfrp1 and Lox nearly 3-fold (Figure 3.10D). By reintroducing TET1 expression to Kras-

transformed NIH3T3 cells, we greatly reduced colony-forming ability (Figure 3.10E).

Thus, in NIH3T3 and HBEC3 cells, KRAS activation suppresses TET1

transcription through the ERK signaling pathway. Reduction of TET1 led to decreased

5hmC, increased 5mC levels, and silencing of TSG promoter regions associated with

reduced TET1 chromatin occupancy. Restoration of TET1 by ERK pathway inhibition

or reintroducing ectopic TET1 gene expression reactivated silenced TSGs and reduced

colony formation. These data identify TET1 in an essential axis of KRAS-ERK TSG

methylation in the transition from an immortalized cell to a malignant cell.

3.4.6 KRAS-mediated suppression of TET1 is required for

maintenance of the malignant phenotype in H1299 cancer

cells

To dissect the connection between KRAS and TET1 in fully malignant cells, we

used siRNA treatment to determine TET1 expression after KRAS depletion in H1299

lung cancer cells. After treating with KRAS siRNA for 2 days, TET1 mRNA and protein

increased nearly 2-fold compared to mock-transfected cells or control siRNA, while

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DNMT1 expression stayed the same (Figure 3.12A and 3.12B). As shown in Figure

3.13, KRAS-mediated suppression of TET1 was also observed in HepG2 hepatoma

cancer cells, indicating that negative regulation by KRAS of TET1 is not cell type-

specific. In agreement with our findings in HBEC3 and NIH3T3 cells, inhibition of the

ERK signaling pathway reactivated TET1 expression, whereas AKT pathway inhibition

failed to have this effect (Figure 3.12B). Moreover, KRAS knockdown inhibited colony-

forming activities (Figure 3.12C), indicating that H1299 cells are addicted to KRAS

expression. To determine whether TET1 is functionally important in KRAS knockdown

cells, we treated cells with KRAS siRNA, TET1 siRNA or combined KRAS and TET1

siRNAs. We confirmed that TET1 knockdown was sufficient to prevent TET1 induction

in KRAS/TET1 double knockdown cells (Figure 3.12D). Colony-forming assays

performed with siRNA-treated cells indicated that TET1 knockdown in a cell depleted

for KRAS is sufficient to rescue the inhibition of colony formation by KRAS

knockdown (Figure 3.12E). Thus, despite the many targets downstream of PI3K-AKT

and RAF-MEK-ERK cascades and the complexity of RAS-driven oncogenesis, TET1

suppression is sufficient to restore H1299 malignancy.

3.5 Discussion

Cancers with RAS activation exhibit aberrant promoter hypermethylation and

transcriptional silencing of TSGs. Sustained epigenetic repression of TSGs not only

promotes tumor initiation, but also maintains their survival and malignant properties.

Based on the fact that DNMT isozymes convert cytosine bases to 5mC, DNMT

enzymes, especially DNMT1 (Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007b),

have been considered the main effectors that drive DNA hypermethylation during RAS-

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induced tumorigenesis. This work reveals that suppression of TET1 expression is

essential for KRAS-induced DNA hypermethylation in cancer cells (Figure 3.12F).

In the Kras-transformed NIH3T3 system, when PI3K and MEK are inhibited,

the Fas and Sfrp1 promoters are rapidly demethylated even when an inhibitor of DNA

replication is applied (Wajapeyee et al. 2013). These data implied a mechanism for

active DNA demethylation, which had not been identified. Moreover, forced expression

of oncogenic BRAF kinase, which functions between RAS and ERK, is sufficient to

transform NIH3T3 cells in a manner that reduced expression of Tet genes and genomic

5hmC levels (Kudo et al. 2012). As shown in Figure 3.8E, the ability of NIH3T3 cells

to be self-limiting by virtue of Fas expression is so important that the Fas promoter is

apparently kept in a 5hmC modified state by Tet1 so that it cannot be silenced by

methylation. Kras transformation depletes Tet1 and allows Dnmt enzymes to convert

nonmodified CpG dinucleotides to 5mCpG.

Although similar KRAS-mediated TET1 suppression was found in HBEC3 and

NIH3T3 cells, there are two important differences. First, decreased Tet1 was

accompanied by increased Dnmt1 in Kras-transformed NIH3T3 cells, while TET1 was

reduced without DNMT1 alteration in KRAS-transformed HBEC3 cells. These cell-

type specific effects indicate that KRAS can regulate dynamic DNA methylation by

inhibiting TET1 expression alone or by further coupling with increased DNMT1.

Further studies should reveal whether TET1 reduction and DNMT1 induction by KRAS

activation work collaboratively or independently on targeted genes to cause promoter

hypermethylation during tumorigenesis. Second, a significant reduction in genomic

5hmC was observed in Kras-transformed NIH3T3 cells but not in HBEC3 cells,

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suggesting that extinguishing TET1 expression may be insufficient to reduce global

5hmC. This may be the case because TET proteins regulate 5mC conversion to 5hmC at

distinct genomic loci. TET1 localizes to CpG-rich promoters via its CXXC domain (Y.

Huang et al. 2014; Xu et al. 2011). However, TET2, which lacks the CXXC domain,

associates primarily with gene bodies (Y. Huang et al. 2014). Indeed, in ESC, Tet2

knockdown causes a greater reduction in genomic 5hmC levels than Tet1 knockdown

(Y. Huang et al. 2014). In addition, TET family proteins may be partially redundant

with the potential for TET2 and TET3 to maintain genomic 5hmC levels when TET1 is

not expressed. Consistent with this hypothesis, double depletion of Tet1 and Tet2 more

significantly reduces 5hmC levels than individual depletion (Koh et al. 2011; Dawlaty

et al. 2013).

Our finding that the RAS-ERK signaling pathway suppresses TET1 expression

during and after malignant transformation has implications for regulation of Tet1

expression in ESC. Tet1 transcripts stay at high levels in the pluripotent state, but drop

rapidly in differentiation in concert with the pluripotency transcription factor Oct4 (Koh

et al. 2011). However, the connection between Oct4 and Tet1 remains unclear.

Evidence has been shown that Oct4 maintains undifferentiated ESC status by inhibiting

the Erk pathway (L. Li et al. 2010). We suggest that Oct4 inhibition of the Erk pathway

maintains Tet1 expression, such that loss of Oct4 results in Tet1 suppression.

Though it is possible for oncogenes to be dispensable after establishment of

neoplastic transformation, oncogene addiction is common (Weinstein 2002), is well

documented in RAS-dependent malignancies (Chin et al. 1999; Singh et al. 2009), and

depends on the RAS-driven DNA hypermethylation phenotype (Wajapeyee et al. 2013).

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In our study, because TET1 reexpression blocks transformation and because TET1

knockdown can allow KRAS knockdown cells to retain a malignant phenotype, we

identified TET1 repression as a critical component of the RAS program. Though

functional TET1 reintroduction is facile in the laboratory setting, there is little optimism

that 100% of a patient’s solid tumor cells could be made to re-express a tumor

suppressing activity. On the other hand, several inhibitors of the EGFR-RAS-RAF-

MEK-ERK axis are under development (Pao and Chmielecki 2010; Downward 2003;

Engelman et al. 2008; Karapetis et al. 2008). Because these drugs may depend on re-

activating TET1 expression for efficacy, TET1 re-repression or increased 5hmC may

serve as biomarkers of functional reversion of RAS transformation.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001; ns, no significant difference in comparison to V1 cells. ###, p < 0.001 in comparison to V1 cells without EGF. Figure 3.1. Oncogenic KRAS expression is sufficient to transform non-malignant HBEC3 cells. (A) HBEC3 stable clones were established to express oncogenic KRAS. Protein levels of RAS, phospho-AKT (pAKT), total-AKT (tAKT), phospho-ERK (pERK) and total-ERK (tERK) were determined by western blotting. (B) KRAS cell lines without EGF proliferate as well as vector cell lines with EGF. Data were normalized to V1 cells with EGF. (C) Adherent and soft-agar colony formation indicate that KRAS transforms HBEC3 cells. All data are presented as mean ± SD.

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Figure 3.2. Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs. (A) Genomic 5mC levels in HBEC3-derived stable cell lines were measured by DNA dot blot assay. (B) Methylation levels of promoter-associated CpG islands were analyzed by qPCR. (C) 5mC bisulfite sequencing of DAPK, MGMT and DUOX1 promoters. White squares represent non-methylated cytosines and black squares represent methylated cytosines in CpG sites. The percentages of methylated CpG from 6 independent clones are indicated. (D) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (E) After 100 nM 5-aza-dC treatment for 5 days, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO treated control. (F) Adherent and soft-agar colony formation after 5-aza-dC treatment indicate that KRAS transformation depends on the hypermethylation phenotype. All data are presented as mean ± SD.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or DMSO treated control.

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**, p < 0.01; ***, p < 0.001 in comparison to V1 cells or the DMSO treated control.

Figure 3.3. Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs and loss of imprinting. (A) Methylation levels of promoter-associated CpG islands were analyzed by qPCR. (B) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (C) 5mC bisulfite sequencing of H19 ICR. White squares represent unmethylated cytosines and black squares represent methylated cytosines in CpG sites. The percentages of methylated CpG from 20 independent clones are indicated. (D) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (E) After 5-aza-dC treatment, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO treated control. All data are presented as mean ± SD.

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**, p < 0.01; ***, p < 0.001 in comparison to V1 cells or DMSO treated control. Figure 3.4. KRAS negatively regulates TET1 expression through the ERK signaling pathway. (A) In HBEC3 cell lines, mRNA levels of DNMT1 and TET1 were determined by RT-qPCR and normalized to V1 cells. Protein levels were determined by western blotting. (B) After 30 µM ERK pathway inhibitor PD98059 or 2 µM AKT pathway inhibitor LY294002 treatment for 6 days, protein levels of DNMT1 and TET1 were determined by western blotting. (C) After ERK pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to DMSO control. (D) Adherent and soft-agar colony formation after ERK pathway or AKT pathway inhibition indicate that cellular transformation is mediated by the ERK pathway. All data are presented as mean ± SD.

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***, p < 0.001 in comparison to V1 cells or the DMSO treated control. Figure 3.5. ERK pathway inhibition reactivates silenced H19 expression in KRAS cells. (A) mRNA levels were determined by RT-qPCR and normalized to V1 cells. (B) After ERK pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO control. All data are presented as mean ± SD.

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Figure 3.6. Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation. (A) Genomic 5hmC levels in HBEC3-drived cell lines were measured by DNA dot blot assay. (B) Hydroxymethylation levels of promoter-associated CpG islands were analyzed by qPCR. (C) TAB-seq 5hmC of DAPK, MGMT and DUOX1 promoters. White circles represent cytosines or 5mC, black circles represent 5hmC in CpG sites, and Xs represent undetermined sites. The percentages of 5hmC from 20 independent clones are indicated. (D) TET1 chromatin occupancy was analyzed using TET1 ChIP and qPCR. (E) After TET1 viral transduction for 6 days, mRNA levels were analyzed by RT-qPCR and normalized to vector viral transduction control. (F) Adherent and soft-agar colony formation after TET1 viral transduction indicate that TET1 re-expression reverts the transformed phenotype. All data are presented as mean ± SD.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or vector virus control.

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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or the vector virus control.

Figure 3.7. Reduction of 5hmC and TET1-association are responsible for KRAS-mediated DNA hypermethylation. (A) Hydroxymethylation levels of promoter-associated CpG islands were analyzed by qPCR. (B) TET1 chromatin occupancy was analyzed using TET1 ChIP and qPCR. (C) After TET1 viral transduction, mRNA levels were analyzed by RT-qPCR and normalized to the vector viral transduction control. All data are presented as mean ± SD.

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Figure 3.8. Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 cells. (A) mRNA levels were determined by RT-qPCR and normalized to NIH3T3 cells. Protein levels were determined by western blotting. (B) Genomic 5mC and 5hmC levels were measured by DNA dot blot assay. (C) Fas expression was determined by RT-qPCR and normalized to that of NIH3T3 cells. (D) Methylation and hydroxymethylation levels of Fas promoter were analyzed by qPCR. (E) Bisulfite sequencing for 5mC and 5hmC. The percentages of 5mC or 5hmC were indicated. (F) Tet1 chromatin occupancy was analyzed using Tet1 ChIP and qPCR. The data indicate that Kras transformation depresses Fas expression by converting the promoter from a 5hmC state to a 5mC state due to depletion of Tet1. All data are presented as mean ± SD.

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**, p < 0.01; ***, p < 0.001 in comparison to NIH3T3 cells.

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**, p < 0.01; ***, p < 0.001 in comparison to NIH3T3 cells.

Figure 3.9. Kras-mediated suppression of Tet1 is associated with decreased 5hmC and increased 5mC levels. (A) mRNA levels were determined by RT-qPCR and normalized to NIH3T3 cells. (B) Methylation and (C) hydroxymethylation levels of Sfrp1 and Lox promoters were analyzed by qPCR. (D) Bisulfite sequencing for 5mC and 5hmC. The percentages of 5mC or 5hmC are indicated at each promoter without and with Kras transformation. (E) Tet1 chromatin occupancy was analyzed using Tet1 ChIP and qPCR. All data are presented as mean ± SD.

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Figure 3.10. Kras promotes transformation by inhibiting Tet1 expression. (A) After 25 µM PD98059 or 2.5 µM LY294002 treatment for 4 days, protein levels of Dnmt1 and Tet1 were determined by western blotting. (B) After Erk pathway or Akt pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to DMSO control. (C) Adherent and soft-agar colony formation after Erk pathway or Akt pathway inhibition indicate that cellular transformation is mediated by the ERK pathway in KRAS-transformed NIH3T3 cells. (D) After TET1 viral transduction for 6 days, mRNA levels were analyzed by RT-qPCR and normalized to vector viral transduction control. (E) Adherent and soft-agar colony formation after TET1 viral transduction indicate that TET1 re-expression reverts Kras-mediated malignancy. All data are presented as mean ± SD.

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**, p < 0.01; ***, p < 0.001 in comparison to DMSO treated control or vector virus control.

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***, p < 0.001 in comparison to the DMSO treated control or NIH3T3 cells. Figure 3.11. Erk pathway inhibition increases Tet1 expression in Kras-transformed NIH3T3 cells, while Akt pathway inhibition shows no effect. (A) After Erk pathway or Akt pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO control. (B) Adherent and soft-agar colony formation. All data were presented as mean ± SD.

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Figure 3.12. KRAS-mediated suppression of TET1 is required for maintaining malignant phenotype in H1299 cancer cells.(A) After 10 µM KRAS siRNA treatment for 2 days, mRNA levels were determined by RT-qPCR and normalized to mock control without adding siRNA. Protein levels of TET1 and DNMT1 were determined by western blotting. (B) After 20 µM PD98059 or 5 µM LY294002 treatment for 2 days, protein levels were determined by western blotting. (C) Adherent and soft-agar colony formation after KRAS siRNA treatment. (D) Protein levels were determined by western blotting after siRNA treatments. (E) Adherent and soft-agar colony formation after indicated siRNA treatments. The data indicate that KRAS becomes dispensable if TET1 is knocked down. All data are presented as mean ± SD. (F) Essential role of TET1 suppression for RAS-mediated DNA hypermethylation and cellular transformation. TET1 modulates epigenetic and transcriptional regulation via hydroxylation of 5mC and subsequent DNA demethylation. TET1 targets CpG-rich promoters of TSGs to prevent DNA hypermethylation. The KRAS-ERK signaling pathway suppresses TET1 transcription. In KRAS-transformed cells, TET1 suppression decreases TET1 binding and 5hmC production at targeted promoters, resulted in hypermethylation-mediated silencing of TSGs.

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***, p < 0.001 in comparison to mock cells or siControl treated cells.

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***, p < 0.001 in comparison to the mock-transfected control. Figure 3.13. KRAS-mediated suppression of TET1 is found in HepG2 hepatoma cancer cells. After KRAS siRNA treatment, mRNA levels were determined by RT-qPCR and normalized to the mock-transfected control. Protein levels of TET1 and DNMT1 were determined by western blotting. All data are presented as mean ± SD.

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Table 3.1. Target list of hypermethylated and silenced lung cancer TSGs.

Function Gene Function Gene

Cell Cycle P16 Cell Adhesion CDH1

CDH13

Growth/ Differentiation APC TIMP3

RARß TSLC1

DUOX1 LAMA3

DUOX2 RECK

IGFBP3

GATA4 Apoptosis DAPK

WWOX RASSF1A

MTHFR FHIT

FAS

DNA repair MGMT NORE1A

BCL2

Detoxification GSTP1 SEMA3B

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Table 3.2. Summary of the changes of promoter methylation and gene expression in KRAS-expressing cell lines. Cell lines KRAS Promoter hypermethylation with gene silencing

DAPK/ MGMT/ DUOX1/ TIMP3/ GATA4

Promoter hypermethylation without gene silencing

None

Gene silencing without promoter hypermethylation

RARβ / IGFBP3/ WWOX/ LAMA3/ RECK/ NORE1A/ BCL2

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Table 3.3. Human primers.

Forward Reverse

Plasmid Construct

KRAS TA Cloning CACCATGGAACAAAAACTTATTTCTGAAGA

AGATCTGACTGAATATAAACTTGTGG GTCGACTTACATAATTACACACTTTG

TET1 TA Cloning CACCATGGAACAAAAACTTATTTCTGAAGA

AGATCTGGAACTGCCCACCTGCAGCTG GTCGACTCAGACCCAATGGTTATAGG

mRNA GAPDH GAGTCAACGGATTTGGTCGT GACAAGCTTCCCGTTCTCAG

KRAS TGTGGTAGTTGGAGCTGGTG TGACCTGCTGTGTCGAGAAT

DAPK CTCCCCATTTCTTGGAGACA CCAGGGATGCTGCAAACTAT

MGMT ACGCACCACACTGGACAGCC CCGGCACGGGGAACTCTTCG

DUOX1 ATGTGCCAGATACCCAAAGC CAGCTGACGGATGACTTGAA

TIMP3 CTGACAGGTCGCGTCTATGA AGTCACAAAGCAAGGCAGGT

GATA4 CCGGGATCTGCCGCGTTCTC GGAGTGAGGGGTCTGGGCGT

H19 CCTCCACGGAGTCGGCACAC GGCGCTGCTGTTCCGATGGT

IGF2 CGAATTGGCTGAGAAACAATTGGC TCGGATGGCCAGTTTACCCTGAAA

DNMT1 GAGCTACCACGCAGACATCA CGAGGAAGTAGAAGCGGTTG

DNMT3A CAAGCGGGACGAGTGGCTGG TCAGTGGGCTGCTGCACAGC

DNMT3B CTCAGAGGCAGTGACAGCAG TGTCTGAATTCCCGTTCTCC

TET1 ACCCCCTGTCACCTGCTGAGG GCGATGGCCACCCCACCAAT

TET2 TCACACCAGGTGCACTTCTC GGATGGTTGTGTTTGTGCTG

TET3 TCTCCCCAGTCTTACCTCCG CCAGGCTTCAGGGAACTCAG

MeDIP/ hMeDIP/ ChIP DAPK GCTTTTGCTTTCCCAGCCAGGGC ATCGCACTTCTCCCCGAAGCCAA

MGMT GAACGCTTTGCGTCCCGACG CCGAGGGAGAGCTCCGCACT

DUOX1 CCATGGGACTTGTGAAGGCGGAC CTCCCGGGGCGCAGGTAGAG

TIMP3 GGGCCGATGAGGTAATGCGGC GCCTGGGCGGCCGAGTGATA

GATA4 TGCTGGGGGAGCTTTCCGCACA TGACTGGCCTGTGGGAGTCACGTG

H19 ICR CTCACACATCACAGCCCGAG TGTGGATAATGCCCGACCTG

Bisulfite sequencing DAPK 1st PCR TTTTATTTATTTTTTAGTTGTGTTTT TAAAAACAATCTCTCTCCAACCTAC

DAPK 2nd PCR TTAGTTTTTGTTTTTTTAGTTAGGG AACAATCCCCAAAACCACAT

MGMT 1st PCR GTTTTTTTGTTTTTTTTAGGTTTT CAACATAAAAAAATAAAAAAAACCC

MGMT 2nd PCR GTTTTTTTGTTTTTTTTAGGTTTT CCAATCCACAATCACTACAAC

DUOX1 1st PCR GGTTTTGGATTTGGAGTTTAGATT AAAAAACTAACATTCCCCTTTCTTC

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Table 3.3. Continued

DUOX1 2nd PCR GTTTTATGGGATTTGTGAAGG CTACCCTTAAAACTCCCTCCC

H19 ICR 1st PCR TAGGGTTTTTGGTAGGTATAGAGTT AAATCCCAAACCATAACACTAAAAC

H19 ICR 2nd PCR ATATGGGTATTTTTGGAGGTTTTTT AAATCCCAAACCATAACACTAAAAC

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Table 3.4. Mouse primers.

Forward Reverse

mRNA β-Actin AGAGGGAAATCGTGCGTGAC CAATAGTGATGACCTGGCCGT

Dnmt1 (*) GAACCATCACCGTGCGAGAC CCAGTGGGCTCATGTCCTTG

Dnmt3A TGGTGCTTTCAAAACAGCGAG GTTTGTTAAAACCCCCTCCAGC

Dnmt3b ACTTGGTGATTGGTGGAAGC CCAGAAGAATGGACGGTTGT

Tet1 TGTCAGACATGGGGCATCAG TGTCGGGGTTTTGTCTTCCG

Tet2 TTGTTAGAAAGGAGACCCGGC TCATGTCCTGTTGACCGTGAG

Tet3 CCGGCCGAGGTGGAAATAAATG CCCTGAGGTGCTTAGCTGC

Fas (*) GATGCACACTCTGCGATGAAG CAGTGTTCACAGCCAGGAGAAT

Sfrp1 (*) CATCCATGGGGCTACAGTGA TGGCATGGTGAGTTTTCAGG

Lox (*) CTCATCTGCCTGAAAGCACAC GGGCAAAGAGGTACATCGAAG

MeDIP/ hMeDIP/ ChIP Fas (*) GAAGTAGAAACAGAAGCTGAG TTGCTACATCCCAACTGTAAC

Sfrp1 TTACAGCGTCCAACTCCGAC CGGCCAGAAGGATCGGTTTA

Lox (*) GCTGCTAGGACCTTGTGATGG CACCCCAGATGAGAGGCCCA

Bisulfite sequencing Fas PCR (*) GAAAAGAAGTAGAAATAGAAGTTGAG CTACATCCCAACTATAACTTTACTAC

Sfrp1 1st PCR (*) GAAAGTATTTGTTTAGTTTTTGGTTTTG CAAATTAAACAACACCATTCTTATAAC

C

Sfrp1 2nd PCR (*) GTTTTGTTTTTTAAGGGGTGTTGAT TTATAACACAACCTCAAATCCAC

Lox 1st PCR (*) AGGGAGGGGGTTGTTAGGATTTTG TAACAACCACCCTCTCTCCTTTCACTC

Lox 2nd PCR (*) GTTGTTAGGATTTTGTGATGGTGAGTTG CACCCCAAATAAAAAACCCATTCACTT

AC

* Gazin, C., Wajapeyee, N., Gobeil, S., Virbasius, C.-M., and Green, M.R. (2007). An elaborate pathway required for Ras-mediated epigenetic silencing. Nature 449, 1073–1077.

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CHAPTER IV

CONCLUSION AND FUTURE DIRECTION

4.1 Implication of DNMT1 RFTS domain mutant and

RFTS domain association protein (RAP) in cancer

We identified that the RFTS domain as responsible for altering DNMT1-

dependent methylation during transformation, suggesting that RFTS domain may be a

target of tumor-specific dysregulation. However, no RFTS-deleted DNMT1 has been

reported in cancers. There are two possible ways in which phenotypes similar to RFTS-

deleted DNMT1 might be produced in human cancers. First, in the COSMIC database,

we identified 26 mutation sites within the conserved RFTS domain of DNMT1 (Fig.

4.1). These DNMT1 mutants might enhance DNMT1 enzyme activity or impair

DNMT1 chromatin association. In order to identify the potential impact of those

mutations, one could generate recombinant DNMT1 by Escherichia coli expression

(Syeda et al. 2011) and test DNMT1 activity in vitro (Syeda et al. 2011). One could also

express these mutant alleles of DNMT1 in HBEC3 or H358 cells. Based on promoter

methylation assays and genomic methylation analysis, one should be able to determine

whether cancer-associated alleles of DNMT1 cause focal hypermethylation and

genomic hypomethylation as suggested by the RFTS-deleted DNMT1.

I also suggest that RAPs have the potential to affect RFTS function. To our

knowledge, UHRF1 is the most well known RAP. UHRF1 recruits DNMT1 to newly

replicated hemimethylated DNA (Bostick et al. 2007; Sharif et al. 2007). UHRF1 also

stimulates DNMT1 enzyme activity by virtue of binding the autoinhibitory RFTS

domain (Bashtrykov, Jankevicius, et al. 2014). UHRF1 was found up-regulated in

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nonsmall cell lung cancer (NSCLC) (Unoki et al. 2010; Daskalos et al. 2011). Indeed,

UHRF1 down-regulation leads to promoter hypomethylation of TSGs in A549 cells

(Daskalos et al. 2011), while UHRF1 overexpression drive genomic hypomethylation

and hepatocellular carcinoma in zebrafish (Mudbhary et al. 2014). These data indicate

that UHRF1 has the ability to regulate regional and global DNA methylation. In

addition to UHRF1, USP7 and NAA10 are RAPs (Felle et al. 2011; Lee et al. 2010). To

test the potential impact of these RAPs, one could overexpress each protein in HBEC3

or H358 cells and determine the effects on promoter and genomic DNA methylation.

Moreover, targeting of RAPs could be a promising therapeutic strategy to limit

DNMT1-dependent hypermethylation and hypomethylation in cancer. Current

approaches use demethylating agents to limit DNMT1 function and methylation levels

(Szyf 2003; Loriot 2006; Yaqinuddin et al. 2009; Morey Kinney et al. 2010). However,

incorporation of 5-aza cytosine leads to non-specific genomic demethylation, DNA and

RNA damage, and significant side effects. Directly targeting RAPs might revert

euchromatin-associated DNMT1 activation and also normalize pericentromeric DNA

methylation.

4.2 Implication of suppression of TET1 in KRAS-

dependent transformation

We found that suppression of TET1 is responsible for KRAS-induced

hypermethylation and malignant transformation. Inhibition of the ERK pathway or

reintroduction of TET1 expression is sufficient to reactivate TSG expression and inhibit

colony formation. Our data indicate that reactivation of suppressed TET1 is a means to

treat KRAS-dependent cancer. Although functional TET1 reintroduction in a patient’s

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tumor would be difficult, treatment with ERK pathway inhibitors is in clinical testing.

In cell lines we examined, colony-forming abilities of KRAS-transformed cells are

more sensitive to ERK inhibition than AKT inhibition and this is the pathway that

restores TET1 expression. Our data are consistent with xenograft KRAS-driven tumor

models which ERK inhibition is more effective than AKT inhibition (Engelman et al.

2008; Hofmann et al. 2012). Because loss of TET1-mediated hydroxymethylation is the

key mediator of KRAS-induced transformation, TET1 re-expression or increased 5hmC

level could be a biomarker to indicate functional reversion of tumors with hyperactive

KRAS.

In addition to colony-forming ability, we also found that TET1 reduction might

be the mediator of EGF-independent growth and KRAS addiction. Oncogene addiction

is a phenomenon in which cancer cells require constant activation of oncogenes or

inactivation of TSGs for survival and malignant phenotype (Weinstein 2002). Several

studies demonstrated that cancer cells with RAS activation are addicted to functional

RAS expression (Chin et al. 1999; Singh et al. 2009), which might dependent on RAS-

induced DNA hypermethylation phenotype (Wajapeyee et al. 2013). In our study,

because KRAS/TET1 double knockdown completely bypassed this KRAS dependency

by preventing TET1 induction, KRAS addiction phenomenon seems to dependent on

suppressing TET1 expression in H1299 cells. We reason that increased TET1 by KRAS

knockdown might trigger active DNA demethylation and substantly reactivate TSG

expression to diminish colony-forming abilities. In addition, TET1 could function as a

transcriptional repressor, which is independent of its catalytic activity (H. Wu et al.

2012; Williams et al. 2011). Further studies should reveal which TET1-targeted genes

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89!

are responsible for KRAS addiction in cancer cells and how increased TET1 regulates

their promoter methylation and transcription.

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* : RFTS mutations from cancer patients Figure 4.1. Multiple sequence alignment analysis of the RFTS domain of DNMT1 showed mutations found in cancer patients were occurred in conserved loci. Amino acid sequence of the RFTS region of DNMT1 from different species were analyzed by using Clustal W. RFTS mutations from cancer patients were marked by black star.

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