University of Iowa
Iowa Research Online
Theses and Dissertations
Summer 2014
Intrinsic and extrinsic regulation of DNA
methylation during malignant transformation
Bo-Kuan Wu
University of Iowa
Copyright 2014 Bo-Kuan Wu
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/1419
Recommended Citation
Wu, Bo-Kuan. "Intrinsic and extrinsic regulation of DNA methylation during malignant transformation." PhD (Doctor of Philosophy)
thesis, University of Iowa, 2014.
http://ir.uiowa.edu/etd/1419.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Cell Biology Commons
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
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.
ii
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 5methylcytosine (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. Methylationassociated 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
iii
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 TET1mediated DNA demethylation is a target responsible for epigenetic changes in cancers
with KRAS activation.
iv
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
2.2
2.3
2.4
2.5
Abstract ..............................................................................................7
Introduction........................................................................................8
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
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 (5aza-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
Discussion ........................................................................................22
v
III.
SUPPRESSION OF TET1-DEPENDENT DNA DEMETHYLATION
IS ESSENTIAL FOR KRAS-MEDIATED TRANSFORMATION ..............43
3.1
3.2
3.3
3.4
3.5
IV.
Abstract ............................................................................................43
Introduction......................................................................................44
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
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 Krastransformed NIH3T3 Cells ...............................................56
3.4.6
KRAS-mediated suppression of TET1 is required for
maintenance of the malignant phenotype in H1299
cancer cells .......................................................................58
Discussion ........................................................................................59
CONCLUSION AND FUTURE DIRECTION ..............................................86
4.1
4.2
Implication of DNMT1 RFTS domain mutant and RFTS
domain association protein (RAP) in cancer ...................................86
Implication of suppression of TET1 in KRAS-dependent
transformation .................................................................................89
REFERENCES ..................................................................................................................91
vi
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
vii
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 DNMT1dependent 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 DNMT1ΔRFTS 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
viii
3.7
Reduction of 5hmC and TET1-association are responsible for KRASmediated 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
ix
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
x
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
xi
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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. cJun 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 genomewide 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 DNAbinding 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 DNMT1dependent 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 downregulated 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 DNAcompetitive 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 DNMT1ΔRFTS 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), RFTSdeleted 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 softagar 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 DNMT1ΔRFTS. 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 DNMT1expressing 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 DNMT1ΔRFTS 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 hypermethylationmediated 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 DNMT1expressing 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 (5aza-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 DNMT1ΔRFTS 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 DNMT1ΔRFTS and vector cells (Fig. 2.7B). This big-data visualization analysis showed that
DNMT1 and DNMT1-ΔRFTS cells share similar methylation targets and that DNMT1ΔRFTS cells produces a higher degree of methylation. The DNMT1 and DNMT1ΔRFTS 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 noncoding 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 nonspecific 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-specificprocessing 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 nonspecific 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 RTqPCR (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 RTqPCR 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 5aza-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) 5methylcytosine (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.
!
!
34!
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.
!
!
35!
*, 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) DNMT1ΔRFTS cells showed the greatest colony formation in soft agar.
!
!
36!
*, 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.
!
!
37!
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.
!
!
38!
Table 2.1. Target list of TSGs have been found with hypermethylation-mediated
gene silencing in lung cancers
Function
Gene
Cell cycle
P16
Function
Gene
CDH1
CDH13
APC
TIMP3
Cell adhesion
RARß
TSLC1
DUOX1
LAMA3
DUOX2
RECK
Growth/ Differentiation
IGFBP3
GATA4
DAPK
WWOX
RASSF1A
MTHFR
FHIT
Apoptosis
DNA repair
MGMT
FAS
NORE1A
BCL2
Detoxification
!
GSTP1
SEMA3B
!
39!
Table 2.2. Summary of the changes of promoter methylation and gene expression
in DNMT1-expressing cell lines.
Cell lines
DNMT1
Promoter hypermethylation with
gene silencing
Promoter hypermethylation
without gene silencing
DAPK
DUOX1
RASSF1A
DUOX2
GATA4
FHIT
MTHFR
Gene silencing without
promoter hypermethylation
!
DNMT1ΔRFTS
DAPK
DUOX1
RASSF1A
DUOX2
GATA4
FHIT
MTHFR
DNMT1ΔCXXC
DAPK
DUOX1
RASSF1A
DUOX2
GATA4
FHIT
MTHFR
DNMT1ΔR/C
DAPK
DUOX1
RASSF1A
DUOX2
FHIT
MTHFR
!
40!
Table 2.3. KEGG pathway enrichment analysis
Pathway
Gene Counts
p value
Focal Adhesion
Fructose and mannose metabolism
Leukocyte transendothelial migration
Tight junction
ECM-receptor interaction
Neurotrophin signaling pathway
Pathways in cancer
31
11
19
19
11
14
28
9.12E-07
1.28E-05
1.13E-04
5.73E-04
0.021146166
0.024730832
0.034601812
Lysosome
Phenylalanine metabolism
Tyrosine metabolism
13
5
7
0.024611161
0.028973119
0.030538571
Hyper-
Hypo-
!
!
41!
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
!
!
42!
Table 2.5. Primer list
Forward
Reverse
Plasmid construct
TA cloning
RFTS deletion
CXXC deletion
R/C deletion
CACCATGCATCATCATCATCATCATCCGGC
GCGTACCGCCCCA
ACCAAGCTGGTCTACCAGATC
TCACCCAAAAAAATGCACCAG
TCACCCAAAAAAATGCACCAG
GTCGACCTAGTCCTTAGCAGCTTCCT
GAGCTACCACGCAGACATCA
CTCCCCATTTCTTGGAGACA
ATGTGCCAGATACCCAAAGC
CATCGAATGGAAATGAAAGGAGTC
GAGTCAACGGATTTGGTCGT
CGAGGAAGTAGAAGCGGTTG
CCAGGGATGCTGCAAACTAT
CAGCTGACGGATGACTTGAA
ACCATTGGATGATTGCAGTCAA
GACAAGCTTCCCGTTCTCAG
GCTTTTGCTTTCCCAGCCAGGGC
CCATGGGACTTGTGAAGGCGGAC
CATCGAATGGAAATGAAAGGAGTC
ATCGCACTTCTCCCCGAAGCCAA
CTCCCGGGGCGCAGGTAGAG
ACCATTGGATGATTGCAGTCAA
TTTTATTTATTTTTTAGTTGTGTTTT
TTAGTTTTTGTTTTTTTAGTTAGGG
GGTTTTGGATTTGGAGTTTAGATT
GTTTTATGGGATTTGTGAAGG
TGGAATTATTATTAAATGGTAATTTAATGG
TTAATGGAAAGGAATGGAAT
TTATTAGGGAGTGTTAGATAGTGGG
TAAAAACAATCTCTCTCCAACCTAC
AACAATCCCCAAAACCACAT
AAAAAACTAACATTCCCCTTTCTTC
CTACCCTTAAAACTCCCTCCC
AAATAATTACAATCAATTCATTC
TTCCATTAAATAATAACTCC
TACCTAAACAAACCTAAACAATAAC
AGGGTGGGTCTTGGAGTTCATG
GTTCTCCTTGTCTTCTCTGTC
AGGGTGGGTCTTGGAGTTCATG
RNA
DNMT1
DAPK
DUOX1
SAT2
GAPDH
MeDIP/ ChIP
DAPK
DUOX1
SAT2
Bisulfite sequencing
DAPK 1st PCR
DAPK 2nd PCR
DUOX1 1st PCR
DUOX1 2nd PCR
SAT2 1st PCR
SAT2 2nd PCR
LINE1
Chromatin accessibility
DAPK
DUOX1
!
GTGGGTGTGGGGCGAGTGGGTGT
CGCCTCCCACCCTCTCCCCAGCC
CTCTCGGCTCCTTGCCGCCTTTT
GGCCCGCGGAGCCCTCTCTC
!
43!
CHAPTER III
SUPPRESSION OF TET1-DEPENDENT DNA
DEMETHYLATION IS ESSENTIAL FOR KRASMEDIATED 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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44!
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 BarSagi 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
!
!
45!
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 methylationassociated 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-ketoglutaratedependent 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
!
!
46!
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 5carboxycytosine (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
!
!
47!
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 KRAStransformed 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
!
!
48!
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
!
!
49!
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
!
!
50!
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.
!
!
51!
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
!
!
52!
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-
!
!
53!
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
!
!
54!
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
!
!
55!
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
!
!
56!
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 methylationmediated 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 4fold 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 Krastransformed 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 typespecific. 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 colonyforming 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 celltype 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-RAFMEK-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 reactivating 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 promoterassociated 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 5aza-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.
!
!
*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or DMSO treated
control.
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66!
**, 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 promoterassociated 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 softagar 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 softagar colony formation after TET1 viral transduction indicate that TET1 reexpression reverts the transformed phenotype. All data are presented as mean ± SD.
!
!
*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or vector virus
control.
!
70!
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71!
*, 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 KRASmediated DNA hypermethylation. (A) Hydroxymethylation levels of promoterassociated 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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72!
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.
!
!
**, p < 0.01; ***, p < 0.001 in comparison to NIH3T3 cells.
!
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74!
**, 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.
!
!
**, p < 0.01; ***, p < 0.001 in comparison to DMSO treated control or vector virus
control.
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!
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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 Krastransformed NIH3T3 cells, while Akt pathway inhibition shows no effect.
(A) After Erk pathway or Akt pathway inhibition, mRNA levels were analyzed by RTqPCR 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 hypermethylationmediated silencing of TSGs.
!
!
***, 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 RTqPCR 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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81!
Table 3.1. Target list of hypermethylated and silenced lung cancer TSGs.
Function
Gene
Cell Cycle
P16
Function
Cell Adhesion
Gene
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
!
82!
Table 3.2. Summary of the changes of promoter methylation and gene expression
in KRAS-expressing cell lines.
Cell lines
Promoter hypermethylation with gene
silencing
Promoter hypermethylation without
gene silencing
Gene silencing without
promoter hypermethylation
!
KRAS
DAPK/ MGMT/ DUOX1/ TIMP3/ GATA4
None
RARβ / IGFBP3/ WWOX/ LAMA3/ RECK/
NORE1A/ BCL2
!
83!
Table 3.3. Human primers.
Forward
Reverse
Plasmid Construct
KRAS TA Cloning
TET1 TA Cloning
CACCATGGAACAAAAACTTATTTCTGAAGA
AGATCTGACTGAATATAAACTTGTGG
CACCATGGAACAAAAACTTATTTCTGAAGA
AGATCTGGAACTGCCCACCTGCAGCTG
GTCGACTTACATAATTACACACTTTG
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
TTTTATTTATTTTTTAGTTGTGTTTT
TAAAAACAATCTCTCTCCAACCTAC
DAPK 2 PCR
TTAGTTTTTGTTTTTTTAGTTAGGG
AACAATCCCCAAAACCACAT
MGMT 1st PCR
GTTTTTTTGTTTTTTTTAGGTTTT
CAACATAAAAAAATAAAAAAAACCC
GTTTTTTTGTTTTTTTTAGGTTTT
CCAATCCACAATCACTACAAC
GGTTTTGGATTTGGAGTTTAGATT
AAAAAACTAACATTCCCCTTTCTTC
Bisulfite sequencing
DAPK 1st PCR
nd
nd
MGMT 2 PCR
st
DUOX1 1 PCR
!
!
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Table 3.3. Continued
DUOX1 2nd PCR
st
H19 ICR 1 PCR
nd
H19 ICR 2 PCR
!
GTTTTATGGGATTTGTGAAGG
CTACCCTTAAAACTCCCTCCC
TAGGGTTTTTGGTAGGTATAGAGTT
AAATCCCAAACCATAACACTAAAAC
ATATGGGTATTTTTGGAGGTTTTTT
AAATCCCAAACCATAACACTAAAAC
!
85!
Table 3.4. Mouse primers.
Forward
Reverse
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
mRNA
MeDIP/ hMeDIP/ ChIP
Fas (*)
GAAGTAGAAACAGAAGCTGAG
TTGCTACATCCCAACTGTAAC
Sfrp1
TTACAGCGTCCAACTCCGAC
CGGCCAGAAGGATCGGTTTA
Lox (*)
GCTGCTAGGACCTTGTGATGG
CACCCCAGATGAGAGGCCCA
Fas PCR (*)
GAAAAGAAGTAGAAATAGAAGTTGAG
CTACATCCCAACTATAACTTTACTAC
Sfrp1 1st PCR (*)
GAAAGTATTTGTTTAGTTTTTGGTTTTG
Bisulfite sequencing
nd
CAAATTAAACAACACCATTCTTATAAC
C
Sfrp1 2 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 RFTSdeleted 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 KRASdependent 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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88!
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 RASinduced 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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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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