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University of Iowa University of Iowa
Iowa Research Online Iowa Research Online
Theses and Dissertations
Summer 2014
Intrinsic and extrinsic regulation of DNA methylation during Intrinsic and extrinsic regulation of DNA methylation during
malignant transformation malignant transformation
Bo-Kuan Wu University of Iowa
Follow this and additional works at: https://ir.uiowa.edu/etd
This dissertation is available at Iowa Research Online: https://ir.uiowa.edu/etd/1419
Recommended Citation Recommended Citation Wu, Bo-Kuan. "Intrinsic and extrinsic regulation of DNA methylation during malignant transformation." PhD (Doctor of Philosophy) thesis, University of Iowa, 2014. https://doi.org/10.17077/etd.iy8vus1s
Follow this and additional works at: https://ir.uiowa.edu/etd
A thesis submitted in partial fulfillment of the requirements for the Doctor of
Philosophy degree in Molecular and Cellular Biology in the Graduate College of
The University of Iowa
August 2014
Thesis Supervisor: Professor Charles Brenner
Graduate College The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Bo-Kuan Wu
has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Molecular and Cellular Biology at the August 2014 graduation.
Thesis Committee: ___________________________________ Charles Brenner, Thesis Supervisor
___________________________________ Frederick Domann
___________________________________ Adam Dupuy
___________________________________ Dawn Quelle
___________________________________ Michael Wright
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ACKNOWLEDGMENTS
First, I would like to thank my mentor, Dr. Charles Brenner. He provided me a
great opportunity to get excellent training in his lab with ample freedom and full support.
He introduced me to the interesting DNA methylation field, which I plan to keep
focusing on in the future.
I am very grateful to members of my thesis committees: Dr. Frederick Domann,
Dr. Dawn Quelle, Dr. Adam Dupuy and Dr. Michael Wright. Thank you for your
invaluable advice for my research in every seminar and progress report.
I would like to thank all members of Dr. Brenner’s lab, particularly Dr. Rebecca
Fagan and Mr. Samuel Trammell. I greatly appreciate your help.
Finally, I would like to thank my family for their patience and encouragement.
I appreciate everything throughout these six years. Now, I am confident to face
my next challenge.
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ABSTRACT
Cytosine methylation of CpG dinucleotides is an epigenetic modification that
cells use to regulate gene expression, largely to promote transcriptional silencing. Focal
hypermethylation of tumor suppressor genes (TSGs) accompanied by genomic
hypomethylation are epigenetic hallmarks of malignancy. DNA methyltransferase 1
(DNMT1) is the principle vertebrate enzyme responsible for maintenance of DNA
methylation and its dysregulation has been found to lead to aberrant methylation in
cancer. In addition, recent findings demonstrated that the ten-eleven translocation 1
(TET1) protein functions as a 5-methylcytosine dioxygenase that converts 5-
methylcytosine (5mC) bases to 5-hydroxymethylcytosine (5hmC) to mediate active DNA
demethylation. Emerging evidence suggests that TET1 might function as a TSG. To
understand the dynamic regulation of DNA methylation during cellular transformation,
my work focused on intrinsic regulation of DNMT1 and how TET1 regulates DNA
demethylation in generating a cancer methylome.
The replication foci targeting sequence (RFTS) is an N-terminal domain of
DNMT1 that inhibits DNA-binding and catalytic activity, suggesting that RFTS deletion
would result in gain of DNMT1 function. However, other data suggested that RFTS may
be a positively acting domain. To test biochemical and structural predictions that the
RFTS domain of DNMT1 is inhibitory, we established cellular systems to evaluate the
function of DNMT1 alleles. The data indicate that deletion of RFTS is necessary and
sufficient to promote cellular transformation, focal hypermethylation of specific TSGs,
and global hypomethylation. These data and human mutation data suggest that RFTS
domain is a target of tumor-specific dysregulation.
RAS mutations are frequently observed in multiple malignancies. Methylation-
associated silencing of TSGs is a hallmark of RAS-driven-tumorigenesis. I discovered
that suppression of TET1 by the ERK signaling cascade is responsible for promoter
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hypermethylation and the malignant phenotype in KRAS-transformed cells. Restoration
of TET1 expression reactivates silenced TSGs and reduces colony formation. Moreover,
TET1 knockdown in a cell depleted for KRAS is sufficient to rescue the inhibition of
colony formation by KRAS knockdown. My findings suggest that impaired TET1-
mediated DNA demethylation is a target responsible for epigenetic changes in cancers
with KRAS activation.
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TABLE OF CONTENTS
LIST OF TABLES ............................................................................................................ vii
LIST OF FIGURES ......................................................................................................... viii
LIST OF ABBREVIATIONS………………………………………………......................x
CHAPTER
I. INTRODUCTION ............................................................................................1
II. RFTS-DELETED DNMT1 ENHANCES TUMORIGENICITY WITH FOCAL HYPERMETHYLATION AND GLOBAL HYPOMETHYLATION ..................................................................................7
2.3.1 Cell culture………. ...........................................................11 2.3.2 Establishment of stable cell lines ......................................11 2.3.3 Proliferation and invasion assay ........................................11 2.3.4 RT-qPCR ...........................................................................12 2.3.5 Immunoblotting .................................................................12 2.3.6 Adherent and soft-agar colony formation ..........................12 2.3.7 Methylation assay ..............................................................13 2.3.8 ChIP ...................................................................................13 2.3.9 Nuclease-protection assay .................................................14 2.3.10 HELP assay and data analysis ...........................................14 2.3.11 Stastical analysis ................................................................14
2.4 Results ..............................................................................................14 2.4.1 Deletion of RFTS enhances the oncogenic activity
of DNMT1 .......................................................................14 2.4.2 Promoter hypermethylation and transcriptional
silencing of DAPK and DUOX1 is driven by DNMT1 ............................................................................16
2.4.3 Strong alleles of DNMT1 condense chromatin structure at the DAPK and DUOX1 promoters ................17
2.4.4 DNA demethylating agent 5-aza-deoxycytidine (5-aza-dC) reverses gene silencing and diminishes the transformation ability of strong DNMT1 alleles. .............18
2.4.5 Genome-wide promoter methylation analysis reveals that DNMT1-ΔRFTS cells produce a methylation pattern similar to DNMT1 cells, though more intense ...............................................................................18
III. SUPPRESSION OF TET1-DEPENDENT DNA DEMETHYLATION IS ESSENTIAL FOR KRAS-MEDIATED TRANSFORMATION ..............43 3.1 Abstract ............................................................................................43 3.2 Introduction ......................................................................................44 3.3 Materials and Methods ....................................................................47
3.4 Results ..............................................................................................51 3.4.1 Oncogenic KRAS expression is sufficient to
hypermethylation-mediated silencing of TSGs and loss of imprinting ..............................................................52
3.4.3 KRAS negatively regulates TET1 expression through the ERK signaling pathway ................................54
3.4.4 Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation. ....................................................55
3.4.5 Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 Cells ...............................................56
3.4.6 KRAS-mediated suppression of TET1 is required for maintenance of the malignant phenotype in H1299 cancer cells .......................................................................58
3.5 Discussion ........................................................................................59 IV. CONCLUSION AND FUTURE DIRECTION ..............................................86
4.1 Implication of DNMT1 RFTS domain mutant and RFTS domain association protein (RAP) in cancer ...................................86
4.2 Implication of suppression of TET1 in KRAS-dependent transformation .................................................................................89
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
2.4 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
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
2.6 Ectopic expression of DNMT1 alleles does not radically alter global methylation intensities ..............................................................................................31
2.7 DNMT1-ΔRFTS expression enhances global DNMT1 methylation changes ..........32
2.8 Genomic hypomethylation is found in DNMT1-ΔRFTS cells .................................33
2.9 The methylation levels of LINE1 were not changed in DNMT1 or DNMT1-ΔRFTS cells ..............................................................................................................34
2.10 Ectopic expression of DNMT1-ΔRFTS in H358 cells is sufficient to enhance proliferation, invasion and soft-agar colony formation ............................................35
2.11 Ectopic expression of DNMT1-ΔRFTS in H358 cells caused gene silencing of DAPK and DUOX1 and demethylation of SAT2 ................................................36
2.12 Dual roles for RFTS domain in DNMT1-dependent DNA methylation ..................37
3.1 Oncogenic KRAS expression is sufficient to transform non-malignant HBEC3 cells .............................................................................................................63
3.2 Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs .........................................................................................................................64
3.3 Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs and loss of imprinting .....................................................................................66
3.4 KRAS negatively regulates TET1 expression through the ERK signaling pathway .....................................................................................................................67
3.6 Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation ..........................................................69
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3.7 Reduction of 5hmC and TET1-association are responsible for KRAS-mediated DNA hypermethylation .............................................................................71
3.8 Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 cells ......................................................72
3.9 Kras-mediated suppression of Tet1 is associated with decreased 5hmC and increased 5mC levels ................................................................................................74
3.10 Kras promotes transformation by inhibiting Tet1 expression ..................................75
3.11 Erk pathway inhibition increases Tet1 expression in Kras-transformed NIH3T3 cells, while Akt pathway inhibition shows no effect .................................77
3.12 KRAS-mediated suppression of TET1 is required for maintaining malignant phenotype in H1299 cancer cells ..............................................................................78
3.13 KRAS-mediated suppression of TET1 is found in HepG2 hepatoma cancer cells ...........................................................................................................................80
4.1 Multiple sequence alignment analysis of the RFTS domain of DNMT1 showed mutations found in cancer patients were occurred in conserved loci. .........90
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LIST OF ABBREVIATIONS
5-aza-dC 5-aza-deoxycytidine
5caC 5-carboxylcytosine
5fC 5-formylcytosine
5hmC 5-hydroxymethylcytosine
5mC 5-methylcytosine
AP1 activator protein 1
BAH bromo-adjacent homolog
ChIP chromatin immunoprecipitation
COSMIC catalogue of somatic mutation in cancer
DMAP1 DNMT1 associated protein 1
DNMT1 DNA methyltransferase 1
DNMT3A DNA methyltransferase 3A
DNMT3B DNA methyltransferase 3B
EGF epidermal growth factor
ERK extracellular signaling-regulated kinase
ESC embryonic stem cells
HBEC3 human bronchial epithelial cells
HELP HpaII tiny fragment enrichment by ligation-mediated PCR
hMeDIP 5-hydroxymethylcytosine DNA immunoprecipitation
HSAN1E hereditary sensory and autonomic neuropathy
ICR imprinting control region
KEGG Kyoto Encyclopedia of Genes and Genomes
MeDIP methylated DNA immunoprecipitation
NAA10 N-_-acetyltransferase 10 NatA catalytic subunit
NLS nuclear localization signal
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ns no significant difference
NSCLC nonsmall cell lung cancer
pAKT phospho-AKT
PCNA proliferating cell nuclear antigen
pERK phospho-ERK
RAPs RFTS-targeted DNMT1 associated proteins
RFTS Replication foci targeting sequence
SAT2 Satellite 2 repeat sequences
siRNA small interfering RNA
Sp1 specificity protein 1
TAB-seq Tet-assisted bisulfite sequencing
tAKT total-AKT
TCF T-cell-factor
tERK total-ERK
TET Ten-eleven translocation
TSGs tumor suppressor genes
UHRF1 ubiquitin-like containing PHD and RING finger domain protein 1
USP7 ubiquitin-specific-processing protease 7
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CHAPTER I
INTRODUCTION
DNA methylation is an epigenetic modification involved in development,
transcription, imprinting, X chromosome inactivation and genomic structure (Baylin
2005). In mammals, DNA methylation typically occurs at the cytosine base of CpG
dinucleotides (Bernstein, Meissner, and Lander 2007; Laird and Jaenisch 1996). Most
CpG dinucleotides fall in repetitive sequences and are heavily methylated to form
heterochromatin to maintain genomic stability (Jones and Baylin 2007). In addition, the
genomic distribution of CpG dinucleotides is uneven. Apart from heterochromatin, they
are usually clustered in gene promoter regions term CpG islands (M. M. Suzuki and
Bird 2008). Half of the genes in mice and humans contain CpG islands (M. M. Suzuki
and Bird 2008; Singal and Ginder 1999). Promoter CpG islands are tend to become
methylated to repress expression of downstream genes (Jones and Baylin 2007). Since
DNA methylation plays an important role in regulating many cellular processes,
abnormal DNA methylation is associated with diseases, including cancer (Robertson
2005).
Because of the obvious link between DNA hypermethylation and transcriptional
repression, tumor suppressor genes (TSGs) have been considered to the most highly
regulated sites for methylation alteration during tumorigenesis (Hughes et al. 2013; Issa
2004). Methylation-associated silencing of TSG induces cancer formation and
progression. Thus, chemotherapy that aims to effect DNA demethylation has become a
promising anti-cancer approach that might reactivate TSG expression (Strathdee and
Brown 2002; Szyf 2005). However, in addition to promoter hypermethylation, global
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genomic hypomethylation has been found in cancer (Jackson et al. 2004). Genomic
hypomethylation could cause transcription activation of oncogenes, chromosome
rearrangement and genomic instability (Ehrlich 2002; Ehrlich 2009). The relationship
between regional hypermethylation and global hypomethylation remains unclear.
In mammals, there is a group of enzymes are responsible for establishing and
maintaining DNA methylation pattern (Chen and Riggs 2011; Rountree et al. 2001).
DNA methyltransferase 3A (DNMT3A) and DNA methyltransferase 3B (DNMT3B)
mediate de novo methylation to deposit methyl groups to naked DNA (Chen and Riggs
2011; Rountree et al. 2001). After DNA replication, DNA methyltransferase 1
(DNMT1) is the principal enzyme responsible for maintenance of cytosine methylation
at CpG dinucleotides (Law and Jacobsen 2010). DNMT1 copies the present methylation
patterns from the parental DNA strand to the newly synthesized strand (Chen and Riggs
2011; Rountree et al. 2001). However, impaired maintenance DNA methylation activity
could cause passive DNA demethylation after DNA replication (Law and Jacobsen
2010). Thus, DNMT1 bears responsibility to increase or decrease the degree of DNA
methylation. Indeed, dysregulation of DNMT1 is associated with either promoter
hypermethylation (J. Wu et al. 1993; Bakin and Curran 1999; Biniszkiewicz et al. 2002)
or genomic hypomethylation (E. Li, Bestor, and Jaenisch 1992; Gaudet 2003). Thus,
changing the expression of DNMT1 alone is not sufficient to recapitulate the regionally
increased and globally decreased DNA methylation changes observed in cancer.
DNMT1 harbors N-terminal regulatory domain and C-terminal catalytic domain.
The long N-terminal regulatory domain is composed of DNMT1 associated protein 1
Bakin and Curran 1999; Biniszkiewicz et al. 2002). However, in our DNMT1 cells in
which ectopic expression is at endogenous levels, there was not a significant increase in
genomic DNA methylation. In contrast, DNMT1-ΔRFTS cells, which displayed the
highest levels of focal hypermethylation, had reduced overall methyl cytosine content in
comparison to vector or other DNMT1-expressing cells. We considered that the main
site of demethylation might occur in Satellite 2 repeat sequences (SAT2) for three
reasons. First, the RFTS domain mediates association of DNMT1 to pericentromeric
heterochromatin to maintain dense methylation (Easwaran et al. 2004; Schneider et al.
2013) and SAT2 is the most abundant repeat in the region (Ting et al. 2011). Second,
SAT2-specific hypomethylation has been found in DNMT1-null cells(Rhee et al. 2000;
Espada 2004) and in patients with RFTS-mutated DNMT1 (Klein et al. 2011). Third,
DNA hypomethylation and RNA up-regulation of SAT2 are highly associated with
various cancers and contributes to genomic instability (Ting et al. 2011). To test
whether SAT2 is hypomethylated in our cell lines, we performed bisulfite sequencing
(Fig. 2.8B and C). SAT2 methylation was significant reduced from 66% (vector) and
63% (DNMT1) to 48% in DNMT1-ΔRFTS cells. Further, we analyzed chromatin
occupancy of DNMT1 on SAT2 loci by DNMT1 ChIP (Fig. 2.8D). The data indicate
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that DNMT1-ΔRFTS expression reduced association between DNMT1 and SAT2 loci.
Unlike what was observed for SAT2, there was no significant methylation change in
LINE1 DNA repeat (Fig. 2.9). Therefore, specific demethylation of SAT2 might be due
to the impaired association between DNMT1-ΔRFTS and the pericentromeric region.
Our results suggest that demethylation of SAT2 and promoter hypomethylation detected
in the HELP assay might both contribute to reduced genomic methylation observed in
DNMT1-ΔRFTS cells. Moreover, we investigated whether SAT2 DNA
hypomethylation is associated with transcription. Indeed, expression of SAT2 non-
coding RNA was increased in DNMT1-ΔRFTS cells, but was not altered in cell lines
with alleles of DNMT1 that are weaker (Fig. 2.8E).
2.4.7 DNMT1-ΔRFTS expression has similar effects in
H358 lung cancer cells.
To rule out a cell-specific effect, we established stable cell lines in H358 cells to
determine the effect of DNMT1-ΔRFTS expression in malignant cells (Fig. 2.10A).
DNMT1-ΔRFTS expression enhanced the proliferation, invasion and soft-agar colony
growth of the cells, while full-length DNMT1 expression behaved similarly to vector
control cells (Fig. 2.10B-D). Moreover, ectopic expression of DNMT1-ΔRFTS slightly
inhibited expressions of DAPK and DUOX1 (Fig. 2.11A). Although DNMT1-ΔRFTS
cells did not express significantly more SAT2 RNA transcripts (Fig. 2.11A), there was a
notable methylation reduction of SAT2 in DNMT1-ΔRFTS cells (Fig. 2.11B and C).
These data suggest that the biological function of DNMT1-ΔRFTS is not cell-specific.
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2.5 Discussion
We proposed that the RFTS domain is DNA-competitive and inhibitory (Syeda
et al. 2011), while other data suggested that the CXXC domain is DNA-competitive and
inhibitory (Song et al. 2011). Here, we hypothesized that expression of hyperactive
DNMT1 lacking an autoinhibitory domain could enhance transformation by altering
DNA methylation. Thus, we modestly expressed full-length and deletion forms of
DNMT1 in immortalized HBEC3 cells to examine their oncogenic potential and
alteration in DNA methylation. Full-length DNMT1 expression triggered cellular
transformation while DNMT1-ΔRFTS functioned as a stronger oncoprotein. The
oncogenic effects of DNMT1-ΔRFTS depended on the presence of the CXXC domain,
which is apparently a positive factor. Deletion of either regulatory domain resulted in
the same apparent target preference; expression of all DNMT1 alleles increased
methylation of DAPK and DUOX1 promoters. DNMT1 and DNMT1-ΔRFTS cells share
similar hypermethylated targets in genome-wide analysis as well, though deletion of
RFTS increased the degree of DNA methylation.
Given previous findings that overexpressed or activated DNMT1 caused non-
specific genomic hypermethylation (J. Wu et al. 1993; Bakin and Curran 1999;
Biniszkiewicz et al. 2002), we surprisingly discovered that DNMT1-ΔRFTS expression
at endogenous levels led to demethylation in SAT2 and in the genome. This finding is
consistent with a previous study on RFTS-mutated DNMT1 (Klein et al. 2011), in
which point mutations in the RFTS domain caused SAT2 and genomic hypomethylation.
The study also showed that mutations in the RFTS domain of DNMT1 impaired binding
with heterochromatin (Klein et al. 2011). We confirmed the impaired association
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between DNMT1 and SAT2 loci in DNMT1-ΔRFTS cells by DNMT1 ChIP. Because
DNMT1 may function as an oligomeric complex (Fellinger et al. 2009), RFTS-deleted
DNMT1 and endogenous DNMT1 may form a heterooligomer that is impaired in
association with pericentromeric heterochromatin. Thus, we suggest a model in which
deletion of the RFTS domain activates DNMT1 for euchromatic DNA-binding, but
decreases chromatin occupancy of DNMT1 to heterochromatic SAT2 loci by virtue of a
missing protein interaction, leading to passive DNA demethylation (Fig. 2.12).
Searching the catalogue of somatic mutation in cancer (COSMIC) database, 26
mutation sites within the RFTS domain of DNMT1 were found (Table 2.4). These
DNMT1 mutants could promote malignancy by increasing DNA methyltransferase
binding and activity on euchromatic DNA while being disadvantaged in pericentromeric
SAT2 association and methylation.
RFTS-targeted DNMT1 associated proteins (RAPs) are likely to participate in
these mechanisms. To our knowledge, there are at least three known RAPs including
ubiquitin-like containing PHD and RING finger domain protein 1 (UHRF1) (Bostick et
al. 2007; Sharif et al. 2007; Bashtrykov, Jankevicius, et al. 2014), ubiquitin-specific-
processing protease 7 (USP7) (Felle et al. 2011) and N-α-acetyltransferase 10 NatA
catalytic subunit (NAA10) (Lee et al. 2010). These proteins have been shown to recruit
DNMT1 to specific loci and stimulate its methylation activity, causing site-specific
hypermethylation. Moreover, these proteins were found up-regulated in lung cancers
(Unoki et al. 2010; Daskalos et al. 2011; Lee et al. 2010), indicating that release of the
RFTS domain inhibition might drive cancer formation via hypermethylayion. If this is
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the case, targeting of these binding partners could be a promising therapeutic strategy to
limit DNMT1-dependent hypermethylation in cancer.
Because of the obvious link between hyperactive DNMT1 and transcriptional
repression of TSGs, DNMT1-mediated DNA hypermethylation is emerging as a crucial
therapeutic target (Laird et al. 1995; M. Suzuki et al. 2004). The current approach is to
inhibit expression or hyperactivity of DNMT1 (Ramchandani et al. 1997; McCabe et al.
2006; Datta et al. 2009). However, demethylating agents lead to unavoidable non-
specific genomic demethylation causing genomic instability or oncogene reactivation
and cause selective opportunities for cancer progression (Szyf 2003; Loriot 2006;
Yaqinuddin et al. 2009; Morey Kinney et al. 2010). Because the RFTS domain
functions as a key regulator of DNMT1 function, targeting RFTS interactions may
revert euchromatin-associated DNMT1 activation while also normalizing
pericentromeric DNA methylation.
In conclusion, our study reveals the functional roles of the RFTS domain of
DNMT1 in maintenance of a nontransformed epigenome. We have demonstrated that
deletion of RFTS enhanced the oncogenic potential of DNMT1 by increasing promoter
methylation of TSGs such as DAPK and DUOX1. However, DNMT1-ΔRFTS also
decreased association of DNMT1 with the pericentromeric region, causing SAT2
demethylation. Because DNMT1-ΔRFTS was able to reprogram the overall methylation
pattern of epithelial cells in a manner that is common in cancer, the data suggest that
RFTS may be a target of tumor-specific dysregulation.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001; ns, indicates no significant difference in comparison to vector cells.
Figure 2.1. Deletion of RFTS enhances the oncogenic activity of DNMT1. (A) HBEC3 stable cell lines were established to express full-length and DNMT1 deletion forms near endogenous DNMT1 levels. The levels of DNMT1 were determined by RT-qPCR (left) and western blotting (right). Data were normalized to vector cells. (B) Adherent colony formation. (C) Soft-agar colony formation.
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*, P < 0.05 in comparison to vector cells. Figure 2.2. Ectopic expression of DNMT1-ΔRFTS enhances invasion activity without proliferation. (A) Neither full-length nor mutant DNMT1 overexpression changed proliferation rates in the presence or absence of EGF. Data were normalized to vector cells cultured with EGF. (B) DNMT1-ΔRFTS and DNMT1-ΔR/C cells showed slightly enhanced invasion. Invasion ability was quantified by the CultureCoat 24 Well Low BME Cell Invasion Assay. Data were normalized to vector cells with n = 4.
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Figure 2.3. DNMT1-ΔRFTS promotes increased methylation and silencing of the DAPK and DUOX1 genes. (A) The methylation levels of DAPK (left) and DUOX1 (right) promoter-associated CpG islands were analyzed by qPCR. Methylated DNA was analyzed using the MethylMiner kit and amplified with specific primers. (B) Bisulfite sequencing results for DAPK (left) and DUOX1 (right) promoters. White squares represent unmethylated cytosines and black squares represent methylated cytosines in CpG sites. The percentage of methylated CpG dinucleotides from 8 independent clones is indicated. (C) DNMT1 chromatin occupancy was analyzed using DNMT1 ChIP and qPCR. (D) mRNA levels of DAPK (left) and DUOX1 (right) were analyzed by RT-qPCR and normalized to vector cells
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison
to vector cells.
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*, p < 0.05; ***, p < 0.001 in comparison to vector cells.
Figure 2.4. DNMT1-ΔRFTS decreases chromatin accessibility at DAPK and DUOX1 promoters. Cells were treated with or without DNA nuclease for 1 hr, prior to detection of promoter DNA by qPCR. The index of chromatin accessibility = 2 ((Ct DNase
treated)-(Ct Untreated)).
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***, p < 0.001 in comparison to the DMSO treated control. Figure 2.5. 5-aza-dC treatment reactivates TSG expression and suppresses DNMT1-dependent transformation. (A) mRNA levels of DAPK (left) and DUOX1 (right) were analyzed by RT-qPCR after 100nM 5-aza-dC treatment for 5 days and normalized to vector cells treated with DMSO. (B) Soft-agar colony formation after 5-aza-dC treatment.
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Figure 2.6. Ectopic expression of DNMT1 alleles does not radically alter global methylation intensities. Pairwise unsupervised clustering followed by Pearson correlations of normalized ratios from HELP assay indicated that the majority of methylation intensities are little changed in DNMT1-expressing cells in comparison to vector cells. DNMT1 and DNMT1-ΔRFTS cells shared more similarity than vector cells.
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Figure 2.7. DNMT1-ΔRFTS expression enhances global DNMT1 methylation changes. (A) Genome-wide promoter DNA methylation profiles were obtained using the HELP assay. Volcano plots are the x-axis scores probe-specific methylation ratios and the y-axis scores p-values for the confidence of measurements. The plots allow visualization of methylation differences between vector and DNMT1 cells as well as the differences between vector and DNMT1-ΔRFTS cells. Probes sets that showed significant hyper- or hypomethylation (p < 0.05 for methylation changes (log2(HpaII/MspI)) > 2) are shown in cyan. All other probes are shown in red. (B) Heat map illustration of HpaII-enrichment fragments with methylation changes (log2(HapII/MspI)) > 2 between vector and DNMT1-ΔRFTS cells.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to vector cells.
Figure 2.8. Genomic hypomethylation is found in DNMT1-ΔRFTS cells. (A) 5-methylcytosine (mC) content of the total cytosine pool was determined by HPLC. (B) Bisulfite sequencing of SAT2. White squares represent unmethylated CpGs, black squares represent methylated CpGs, and grey squares represent undetermined sites. Each row is an independent sequencing result. (C) Quantitation of SAT2 bisulfite sequencing. (D) DNMT1 chromatin occupancy was analyzed using DNMT1 ChIP and qPCR. (E) Expression of SAT2 non-coding RNA was analyzed by RT-qPCR and normalized to vector cells.
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Figure 2.9. The methylation levels of LINE1 were not changed in DNMT1 or DNMT1-ΔRFTS cells. (A) Bisulfite sequencing of LINE1. (B) Quantitation of LINE1 bisulfite sequencing. n = 20. ns, indicates no significant difference in comparison to vector cells.
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*, P < 0.05; ***, p < 0.001 in comparison to vector cells.
Figure 2.10. Ectopic expression of DNMT1-ΔRFTS in H358 cells is sufficient to enhance proliferation, invasion and soft-agar colony formation. (A) H358 stable cell lines were established to express full-length DNMT1 or DNMT1-ΔRFTS near the endogenous DNMT1 levels. The levels of DNMT1 were determined by western blotting. (B) Both DNMT1 and DNMT1-ΔRFTS increased the proliferation rate in H358 cells. Data were normalized to vector cells. (C) DNMT1-ΔRFTS showed slightly enhanced invasion. Invasion ability was quantified by the CultureCoat 24 Well Low BME Cell Invasion Assay. Data were normalized to vector cells. n = 4. (D) DNMT1-ΔRFTS cells showed the greatest colony formation in soft agar.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to vector cells.
Figure 2.11. Ectopic expression of DNMT1-ΔRFTS in H358 cells caused gene silencing of DAPK and DUOX1 and demethylation of SAT2. (A) Expressions of DAPK, DUOX1 and SAT2 were analyzed using RT-qPCR. (B) Bisulfite sequencing of SAT2. (C) Quantitation of SAT2 bisulfite sequencing. n = 20.
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Figure 2.12. Dual roles for RFTS domain in DNMT1-dependent DNA methylation. (A) RFTS-targeted DNMT1 associated proteins (RAP) are proposed to relieve inhibition of DNMT1 for access to euchromatin. (B) The RFTS domain mediates association between DNMT1 and pericentromeric heterochromatin. (C) In cancer, overexpression of RAPs or mutation of RFTS is proposed to relieve DNMT1 inhibition, thereby increasing methylation and silencing of TSGs. However, because the RFTS domain is required for association with heterochromatic SAT2 sequences, DNMT1 with mutant RFTS may be less associated with such sequences, accounting for global hypomethylation.
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Table 2.1. Target list of TSGs have been found with hypermethylation-mediated gene silencing in lung cancers Function Gene Function Gene
Cell cycle P16
Cell adhesion
CDH1
CDH13
Growth/ Differentiation
APC TIMP3
RARß TSLC1
DUOX1 LAMA3
DUOX2 RECK
IGFBP3
GATA4
Apoptosis
DAPK
WWOX RASSF1A
MTHFR FHIT
FAS
DNA repair MGMT NORE1A
BCL2
Detoxification GSTP1 SEMA3B
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Table 2.2. Summary of the changes of promoter methylation and gene expression in DNMT1-expressing cell lines. Cell lines DNMT1 DNMT1-
failed to have this effect (Figure 3.12B). Moreover, KRAS knockdown inhibited colony-
forming activities (Figure 3.12C), indicating that H1299 cells are addicted to KRAS
expression. To determine whether TET1 is functionally important in KRAS knockdown
cells, we treated cells with KRAS siRNA, TET1 siRNA or combined KRAS and TET1
siRNAs. We confirmed that TET1 knockdown was sufficient to prevent TET1 induction
in KRAS/TET1 double knockdown cells (Figure 3.12D). Colony-forming assays
performed with siRNA-treated cells indicated that TET1 knockdown in a cell depleted
for KRAS is sufficient to rescue the inhibition of colony formation by KRAS
knockdown (Figure 3.12E). Thus, despite the many targets downstream of PI3K-AKT
and RAF-MEK-ERK cascades and the complexity of RAS-driven oncogenesis, TET1
suppression is sufficient to restore H1299 malignancy.
3.5 Discussion
Cancers with RAS activation exhibit aberrant promoter hypermethylation and
transcriptional silencing of TSGs. Sustained epigenetic repression of TSGs not only
promotes tumor initiation, but also maintains their survival and malignant properties.
Based on the fact that DNMT isozymes convert cytosine bases to 5mC, DNMT
enzymes, especially DNMT1 (Gazin, Wajapeyee, Gobeil, Virbasius, and Green 2007b),
have been considered the main effectors that drive DNA hypermethylation during RAS-
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induced tumorigenesis. This work reveals that suppression of TET1 expression is
essential for KRAS-induced DNA hypermethylation in cancer cells (Figure 3.12F).
In the Kras-transformed NIH3T3 system, when PI3K and MEK are inhibited,
the Fas and Sfrp1 promoters are rapidly demethylated even when an inhibitor of DNA
replication is applied (Wajapeyee et al. 2013). These data implied a mechanism for
active DNA demethylation, which had not been identified. Moreover, forced expression
of oncogenic BRAF kinase, which functions between RAS and ERK, is sufficient to
transform NIH3T3 cells in a manner that reduced expression of Tet genes and genomic
5hmC levels (Kudo et al. 2012). As shown in Figure 3.8E, the ability of NIH3T3 cells
to be self-limiting by virtue of Fas expression is so important that the Fas promoter is
apparently kept in a 5hmC modified state by Tet1 so that it cannot be silenced by
methylation. Kras transformation depletes Tet1 and allows Dnmt enzymes to convert
nonmodified CpG dinucleotides to 5mCpG.
Although similar KRAS-mediated TET1 suppression was found in HBEC3 and
NIH3T3 cells, there are two important differences. First, decreased Tet1 was
accompanied by increased Dnmt1 in Kras-transformed NIH3T3 cells, while TET1 was
reduced without DNMT1 alteration in KRAS-transformed HBEC3 cells. These cell-
type specific effects indicate that KRAS can regulate dynamic DNA methylation by
inhibiting TET1 expression alone or by further coupling with increased DNMT1.
Further studies should reveal whether TET1 reduction and DNMT1 induction by KRAS
activation work collaboratively or independently on targeted genes to cause promoter
hypermethylation during tumorigenesis. Second, a significant reduction in genomic
5hmC was observed in Kras-transformed NIH3T3 cells but not in HBEC3 cells,
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suggesting that extinguishing TET1 expression may be insufficient to reduce global
5hmC. This may be the case because TET proteins regulate 5mC conversion to 5hmC at
distinct genomic loci. TET1 localizes to CpG-rich promoters via its CXXC domain (Y.
Huang et al. 2014; Xu et al. 2011). However, TET2, which lacks the CXXC domain,
associates primarily with gene bodies (Y. Huang et al. 2014). Indeed, in ESC, Tet2
knockdown causes a greater reduction in genomic 5hmC levels than Tet1 knockdown
(Y. Huang et al. 2014). In addition, TET family proteins may be partially redundant
with the potential for TET2 and TET3 to maintain genomic 5hmC levels when TET1 is
not expressed. Consistent with this hypothesis, double depletion of Tet1 and Tet2 more
significantly reduces 5hmC levels than individual depletion (Koh et al. 2011; Dawlaty
et al. 2013).
Our finding that the RAS-ERK signaling pathway suppresses TET1 expression
during and after malignant transformation has implications for regulation of Tet1
expression in ESC. Tet1 transcripts stay at high levels in the pluripotent state, but drop
rapidly in differentiation in concert with the pluripotency transcription factor Oct4 (Koh
et al. 2011). However, the connection between Oct4 and Tet1 remains unclear.
Evidence has been shown that Oct4 maintains undifferentiated ESC status by inhibiting
the Erk pathway (L. Li et al. 2010). We suggest that Oct4 inhibition of the Erk pathway
maintains Tet1 expression, such that loss of Oct4 results in Tet1 suppression.
Though it is possible for oncogenes to be dispensable after establishment of
neoplastic transformation, oncogene addiction is common (Weinstein 2002), is well
documented in RAS-dependent malignancies (Chin et al. 1999; Singh et al. 2009), and
depends on the RAS-driven DNA hypermethylation phenotype (Wajapeyee et al. 2013).
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In our study, because TET1 reexpression blocks transformation and because TET1
knockdown can allow KRAS knockdown cells to retain a malignant phenotype, we
identified TET1 repression as a critical component of the RAS program. Though
functional TET1 reintroduction is facile in the laboratory setting, there is little optimism
that 100% of a patient’s solid tumor cells could be made to re-express a tumor
suppressing activity. On the other hand, several inhibitors of the EGFR-RAS-RAF-
MEK-ERK axis are under development (Pao and Chmielecki 2010; Downward 2003;
Engelman et al. 2008; Karapetis et al. 2008). Because these drugs may depend on re-
activating TET1 expression for efficacy, TET1 re-repression or increased 5hmC may
serve as biomarkers of functional reversion of RAS transformation.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001; ns, no significant difference in comparison to V1 cells. ###, p < 0.001 in comparison to V1 cells without EGF. Figure 3.1. Oncogenic KRAS expression is sufficient to transform non-malignant HBEC3 cells. (A) HBEC3 stable clones were established to express oncogenic KRAS. Protein levels of RAS, phospho-AKT (pAKT), total-AKT (tAKT), phospho-ERK (pERK) and total-ERK (tERK) were determined by western blotting. (B) KRAS cell lines without EGF proliferate as well as vector cell lines with EGF. Data were normalized to V1 cells with EGF. (C) Adherent and soft-agar colony formation indicate that KRAS transforms HBEC3 cells. All data are presented as mean ± SD.
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Figure 3.2. Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs. (A) Genomic 5mC levels in HBEC3-derived stable cell lines were measured by DNA dot blot assay. (B) Methylation levels of promoter-associated CpG islands were analyzed by qPCR. (C) 5mC bisulfite sequencing of DAPK, MGMT and DUOX1 promoters. White squares represent non-methylated cytosines and black squares represent methylated cytosines in CpG sites. The percentages of methylated CpG from 6 independent clones are indicated. (D) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (E) After 100 nM 5-aza-dC treatment for 5 days, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO treated control. (F) Adherent and soft-agar colony formation after 5-aza-dC treatment indicate that KRAS transformation depends on the hypermethylation phenotype. All data are presented as mean ± SD.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or DMSO treated control.
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**, p < 0.01; ***, p < 0.001 in comparison to V1 cells or the DMSO treated control.
Figure 3.3. Oncogenic KRAS expression causes hypermethylation-mediated silencing of TSGs and loss of imprinting. (A) Methylation levels of promoter-associated CpG islands were analyzed by qPCR. (B) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (C) 5mC bisulfite sequencing of H19 ICR. White squares represent unmethylated cytosines and black squares represent methylated cytosines in CpG sites. The percentages of methylated CpG from 20 independent clones are indicated. (D) mRNA levels were analyzed by RT-qPCR and normalized to V1 cells. (E) After 5-aza-dC treatment, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO treated control. All data are presented as mean ± SD.
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**, p < 0.01; ***, p < 0.001 in comparison to V1 cells or DMSO treated control. Figure 3.4. KRAS negatively regulates TET1 expression through the ERK signaling pathway. (A) In HBEC3 cell lines, mRNA levels of DNMT1 and TET1 were determined by RT-qPCR and normalized to V1 cells. Protein levels were determined by western blotting. (B) After 30 µM ERK pathway inhibitor PD98059 or 2 µM AKT pathway inhibitor LY294002 treatment for 6 days, protein levels of DNMT1 and TET1 were determined by western blotting. (C) After ERK pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to DMSO control. (D) Adherent and soft-agar colony formation after ERK pathway or AKT pathway inhibition indicate that cellular transformation is mediated by the ERK pathway. All data are presented as mean ± SD.
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***, p < 0.001 in comparison to V1 cells or the DMSO treated control. Figure 3.5. ERK pathway inhibition reactivates silenced H19 expression in KRAS cells. (A) mRNA levels were determined by RT-qPCR and normalized to V1 cells. (B) After ERK pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO control. All data are presented as mean ± SD.
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Figure 3.6. Reduction of TET1 and 5hmC are responsible for KRAS-mediated DNA hypermethylation and cellular transformation. (A) Genomic 5hmC levels in HBEC3-drived cell lines were measured by DNA dot blot assay. (B) Hydroxymethylation levels of promoter-associated CpG islands were analyzed by qPCR. (C) TAB-seq 5hmC of DAPK, MGMT and DUOX1 promoters. White circles represent cytosines or 5mC, black circles represent 5hmC in CpG sites, and Xs represent undetermined sites. The percentages of 5hmC from 20 independent clones are indicated. (D) TET1 chromatin occupancy was analyzed using TET1 ChIP and qPCR. (E) After TET1 viral transduction for 6 days, mRNA levels were analyzed by RT-qPCR and normalized to vector viral transduction control. (F) Adherent and soft-agar colony formation after TET1 viral transduction indicate that TET1 re-expression reverts the transformed phenotype. All data are presented as mean ± SD.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or vector virus control.
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*, p < 0.05; **, p < 0.01; ***, p < 0.001 in comparison to V1 cells or the vector virus control.
Figure 3.7. Reduction of 5hmC and TET1-association are responsible for KRAS-mediated DNA hypermethylation. (A) Hydroxymethylation levels of promoter-associated CpG islands were analyzed by qPCR. (B) TET1 chromatin occupancy was analyzed using TET1 ChIP and qPCR. (C) After TET1 viral transduction, mRNA levels were analyzed by RT-qPCR and normalized to the vector viral transduction control. All data are presented as mean ± SD.
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Figure 3.8. Loss of Tet1 expression is associated with decreased 5hmC and increased 5mC content in Kras-transformed NIH3T3 cells. (A) mRNA levels were determined by RT-qPCR and normalized to NIH3T3 cells. Protein levels were determined by western blotting. (B) Genomic 5mC and 5hmC levels were measured by DNA dot blot assay. (C) Fas expression was determined by RT-qPCR and normalized to that of NIH3T3 cells. (D) Methylation and hydroxymethylation levels of Fas promoter were analyzed by qPCR. (E) Bisulfite sequencing for 5mC and 5hmC. The percentages of 5mC or 5hmC were indicated. (F) Tet1 chromatin occupancy was analyzed using Tet1 ChIP and qPCR. The data indicate that Kras transformation depresses Fas expression by converting the promoter from a 5hmC state to a 5mC state due to depletion of Tet1. All data are presented as mean ± SD.
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**, p < 0.01; ***, p < 0.001 in comparison to NIH3T3 cells.
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**, p < 0.01; ***, p < 0.001 in comparison to NIH3T3 cells.
Figure 3.9. Kras-mediated suppression of Tet1 is associated with decreased 5hmC and increased 5mC levels. (A) mRNA levels were determined by RT-qPCR and normalized to NIH3T3 cells. (B) Methylation and (C) hydroxymethylation levels of Sfrp1 and Lox promoters were analyzed by qPCR. (D) Bisulfite sequencing for 5mC and 5hmC. The percentages of 5mC or 5hmC are indicated at each promoter without and with Kras transformation. (E) Tet1 chromatin occupancy was analyzed using Tet1 ChIP and qPCR. All data are presented as mean ± SD.
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Figure 3.10. Kras promotes transformation by inhibiting Tet1 expression. (A) After 25 µM PD98059 or 2.5 µM LY294002 treatment for 4 days, protein levels of Dnmt1 and Tet1 were determined by western blotting. (B) After Erk pathway or Akt pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to DMSO control. (C) Adherent and soft-agar colony formation after Erk pathway or Akt pathway inhibition indicate that cellular transformation is mediated by the ERK pathway in KRAS-transformed NIH3T3 cells. (D) After TET1 viral transduction for 6 days, mRNA levels were analyzed by RT-qPCR and normalized to vector viral transduction control. (E) Adherent and soft-agar colony formation after TET1 viral transduction indicate that TET1 re-expression reverts Kras-mediated malignancy. All data are presented as mean ± SD.
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**, p < 0.01; ***, p < 0.001 in comparison to DMSO treated control or vector virus control.
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***, p < 0.001 in comparison to the DMSO treated control or NIH3T3 cells. Figure 3.11. Erk pathway inhibition increases Tet1 expression in Kras-transformed NIH3T3 cells, while Akt pathway inhibition shows no effect. (A) After Erk pathway or Akt pathway inhibition, mRNA levels were analyzed by RT-qPCR and normalized to the DMSO control. (B) Adherent and soft-agar colony formation. All data were presented as mean ± SD.
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Figure 3.12. KRAS-mediated suppression of TET1 is required for maintaining malignant phenotype in H1299 cancer cells.(A) After 10 µM KRAS siRNA treatment for 2 days, mRNA levels were determined by RT-qPCR and normalized to mock control without adding siRNA. Protein levels of TET1 and DNMT1 were determined by western blotting. (B) After 20 µM PD98059 or 5 µM LY294002 treatment for 2 days, protein levels were determined by western blotting. (C) Adherent and soft-agar colony formation after KRAS siRNA treatment. (D) Protein levels were determined by western blotting after siRNA treatments. (E) Adherent and soft-agar colony formation after indicated siRNA treatments. The data indicate that KRAS becomes dispensable if TET1 is knocked down. All data are presented as mean ± SD. (F) Essential role of TET1 suppression for RAS-mediated DNA hypermethylation and cellular transformation. TET1 modulates epigenetic and transcriptional regulation via hydroxylation of 5mC and subsequent DNA demethylation. TET1 targets CpG-rich promoters of TSGs to prevent DNA hypermethylation. The KRAS-ERK signaling pathway suppresses TET1 transcription. In KRAS-transformed cells, TET1 suppression decreases TET1 binding and 5hmC production at targeted promoters, resulted in hypermethylation-mediated silencing of TSGs.
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***, p < 0.001 in comparison to mock cells or siControl treated cells.
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***, p < 0.001 in comparison to the mock-transfected control. Figure 3.13. KRAS-mediated suppression of TET1 is found in HepG2 hepatoma cancer cells. After KRAS siRNA treatment, mRNA levels were determined by RT-qPCR and normalized to the mock-transfected control. Protein levels of TET1 and DNMT1 were determined by western blotting. All data are presented as mean ± SD.
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Table 3.1. Target list of hypermethylated and silenced lung cancer TSGs.
Function Gene Function Gene
Cell Cycle P16 Cell Adhesion CDH1
CDH13
Growth/ Differentiation APC TIMP3
RARß TSLC1
DUOX1 LAMA3
DUOX2 RECK
IGFBP3
GATA4 Apoptosis DAPK
WWOX RASSF1A
MTHFR FHIT
FAS
DNA repair MGMT NORE1A
BCL2
Detoxification GSTP1 SEMA3B
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Table 3.2. Summary of the changes of promoter methylation and gene expression in KRAS-expressing cell lines. Cell lines KRAS Promoter hypermethylation with gene silencing
* Gazin, C., Wajapeyee, N., Gobeil, S., Virbasius, C.-M., and Green, M.R. (2007). An elaborate pathway required for Ras-mediated epigenetic silencing. Nature 449, 1073–1077.
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CHAPTER IV
CONCLUSION AND FUTURE DIRECTION
4.1 Implication of DNMT1 RFTS domain mutant and
RFTS domain association protein (RAP) in cancer
We identified that the RFTS domain as responsible for altering DNMT1-
dependent methylation during transformation, suggesting that RFTS domain may be a
target of tumor-specific dysregulation. However, no RFTS-deleted DNMT1 has been
reported in cancers. There are two possible ways in which phenotypes similar to RFTS-
deleted DNMT1 might be produced in human cancers. First, in the COSMIC database,
we identified 26 mutation sites within the conserved RFTS domain of DNMT1 (Fig.
4.1). These DNMT1 mutants might enhance DNMT1 enzyme activity or impair
DNMT1 chromatin association. In order to identify the potential impact of those
mutations, one could generate recombinant DNMT1 by Escherichia coli expression
(Syeda et al. 2011) and test DNMT1 activity in vitro (Syeda et al. 2011). One could also
express these mutant alleles of DNMT1 in HBEC3 or H358 cells. Based on promoter
methylation assays and genomic methylation analysis, one should be able to determine
whether cancer-associated alleles of DNMT1 cause focal hypermethylation and
genomic hypomethylation as suggested by the RFTS-deleted DNMT1.
I also suggest that RAPs have the potential to affect RFTS function. To our
knowledge, UHRF1 is the most well known RAP. UHRF1 recruits DNMT1 to newly
replicated hemimethylated DNA (Bostick et al. 2007; Sharif et al. 2007). UHRF1 also
stimulates DNMT1 enzyme activity by virtue of binding the autoinhibitory RFTS
domain (Bashtrykov, Jankevicius, et al. 2014). UHRF1 was found up-regulated in
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nonsmall cell lung cancer (NSCLC) (Unoki et al. 2010; Daskalos et al. 2011). Indeed,
UHRF1 down-regulation leads to promoter hypomethylation of TSGs in A549 cells
(Daskalos et al. 2011), while UHRF1 overexpression drive genomic hypomethylation
and hepatocellular carcinoma in zebrafish (Mudbhary et al. 2014). These data indicate
that UHRF1 has the ability to regulate regional and global DNA methylation. In
addition to UHRF1, USP7 and NAA10 are RAPs (Felle et al. 2011; Lee et al. 2010). To
test the potential impact of these RAPs, one could overexpress each protein in HBEC3
or H358 cells and determine the effects on promoter and genomic DNA methylation.
Moreover, targeting of RAPs could be a promising therapeutic strategy to limit
DNMT1-dependent hypermethylation and hypomethylation in cancer. Current
approaches use demethylating agents to limit DNMT1 function and methylation levels
(Szyf 2003; Loriot 2006; Yaqinuddin et al. 2009; Morey Kinney et al. 2010). However,
incorporation of 5-aza cytosine leads to non-specific genomic demethylation, DNA and
RNA damage, and significant side effects. Directly targeting RAPs might revert
euchromatin-associated DNMT1 activation and also normalize pericentromeric DNA
methylation.
4.2 Implication of suppression of TET1 in KRAS-
dependent transformation
We found that suppression of TET1 is responsible for KRAS-induced
hypermethylation and malignant transformation. Inhibition of the ERK pathway or
reintroduction of TET1 expression is sufficient to reactivate TSG expression and inhibit
colony formation. Our data indicate that reactivation of suppressed TET1 is a means to
treat KRAS-dependent cancer. Although functional TET1 reintroduction in a patient’s
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tumor would be difficult, treatment with ERK pathway inhibitors is in clinical testing.
In cell lines we examined, colony-forming abilities of KRAS-transformed cells are
more sensitive to ERK inhibition than AKT inhibition and this is the pathway that
restores TET1 expression. Our data are consistent with xenograft KRAS-driven tumor
models which ERK inhibition is more effective than AKT inhibition (Engelman et al.
2008; Hofmann et al. 2012). Because loss of TET1-mediated hydroxymethylation is the
key mediator of KRAS-induced transformation, TET1 re-expression or increased 5hmC
level could be a biomarker to indicate functional reversion of tumors with hyperactive
KRAS.
In addition to colony-forming ability, we also found that TET1 reduction might
be the mediator of EGF-independent growth and KRAS addiction. Oncogene addiction
is a phenomenon in which cancer cells require constant activation of oncogenes or
inactivation of TSGs for survival and malignant phenotype (Weinstein 2002). Several
studies demonstrated that cancer cells with RAS activation are addicted to functional
RAS expression (Chin et al. 1999; Singh et al. 2009), which might dependent on RAS-
induced DNA hypermethylation phenotype (Wajapeyee et al. 2013). In our study,
because KRAS/TET1 double knockdown completely bypassed this KRAS dependency
by preventing TET1 induction, KRAS addiction phenomenon seems to dependent on
suppressing TET1 expression in H1299 cells. We reason that increased TET1 by KRAS
knockdown might trigger active DNA demethylation and substantly reactivate TSG
expression to diminish colony-forming abilities. In addition, TET1 could function as a
transcriptional repressor, which is independent of its catalytic activity (H. Wu et al.
2012; Williams et al. 2011). Further studies should reveal which TET1-targeted genes
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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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