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Role of DNA methyltransferases 3A and 3B in inheritance of DNA methylation patterns
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Role of DNA methyltransferases 3A and 3B in inheritance of DNA methylation patterns
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Content
ROLE OF DNA METHYLTRANSFERASES 3A AND 3B IN
INHERITANCE OF DNA METHYLATION PATTERNS
by
Shikhar Sharma
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETIC, MOLECULAR AND CELLULAR BIOLOGY)
December 2010
Copyright 2010 Shikhar Sharma
ii
EPIGRAPH
“Research is to see what everybody else has seen, and to think what nobody else has
thought.”
-- Albert Szent-Györgi (1893-1986)
iii
DEDICATION
I dedicate this work to my mother whose hard work and perseverance has enabled me to
pursue a career in science, my father who has taught me to value principles and integrity
and my brother who has not only been my biggest support but has also inspired me to aim
for excellence and to give my best at everything I do.
iv
ACKNOWLEDGEMENTS
I would like to thank the following people who made the work in this thesis possible:
Dr. Peter A. Jones, my mentor, for his invaluable guidance and unfailing support
throughout the course of this work. You have not only taught me how to work and think
like a true scientist but also how to face setbacks and overcome them with a never-to-die
spirit. Your passion for science and your incredible work management skills have always
inspired me to push myself beyond my limits. Your words of wisdom will stay with me
for the rest of my career.
Dr. Gangning Liang, who is the lifeline of Jones lab and without whom none of
this work would have been possible. Your scientific acumen and practical approach along
with your patient and ‘never give up’ attitude has truly been an inspiration. I have
enjoyed every single day working with you in the Jones lab and would like to thank you
for the tremendous support you provide to the entire lab which really makes research easy
and fun.
Dr. Shinwu Jeong, for his mentorship during my initial days in the lab. You set an
example for me through your meticulous way of doing research along with your amazing
ability to always maintain a positive attitude.
Dr. Terry Kelly, for always being there when I needed her support and advice and
for always entertaining my weird scientific questions! I really enjoyed the experience of
working with you and have learnt a lot from you.
v
Dr. Daniel De Carvalho, for all his support and help during this work. It was a
pleasure working with you.
Dr. Yvonne Tsai, for her efficient lab management which makes an invaluable,
though often unnoticed, contribution to the science done in Jones lab.
My fellow graduate students Dr. Erika Wolff, Dr. Connie Cortez, Dr. Joy Lin, Dr.
Jeffrey Friedman, Flora Han and Sheng-Fang Su, who were all so much more than
colleagues. Postdocs Dr. Phillippa Oakford, Dr. Jueng Soo You, Dr. Claudia Andreu-
Vieyra, Dr. Gerda Egger, Dr. Tina Miranda and Dr. Xiaojing Yang, for all your help and
for making the lab a truly amazing place to work everyday. My past and present
committee members, Drs. Michael Stallcup, Gerhard Coetzee, Peter Laird, and Woojin
An, for their support and feedback. My scientific collaborators, Dr. Daniel Weisenberger,
Dr. Allen Yang, Dr. Xiaojiang Chen, Dr. Siho Choi and Daniel Gerke, for all their
assistance.
My friends and family including Mom, Dad, Abhishek, Divya and Tumul, for
their unyielding support and encouragement throughout this process.
vi
TABLE OF CONTENTS
Epigraph ii
Dedication iii
Acknowledgements iv
List of Figures ix
Abstract xii
Chapter 1: Epigenetic Mechanisms in Normal Development and Disease 1
Introduction 1
Epigenetic mechanisms in normal cells 3
DNA methylation 3
Table 1.1 Epigenetic mechanisms involved in regulating gene
expression and chromatin structure in normal mammalian cells 4
Covalent histone modifications 7
Interplay of DNA methylation and histone modifications 10
Nucleosome positioning & histone variants 11
miRNAs 13
Aberrant reprogramming of the epigenome in cancer 14
DNA methylation aberrations in cancer 15
Changes in histone modifications in cancer 19
Epigenetic switching in cancer 22
Role of nucleosome positioning in cancer 23
Deregulation of miRNAs in cancer 25
The cancer stem cell model 27
Epigenetic therapy of cancer 30
Future prospects and challenges 33
Overview of thesis research 35
Chapter 2: Strong Anchoring of DNMT3A/3B to Nucleosomes containing
Methylated DNA 40
Introduction 40
Materials and Methods 45
Results 51
DNMT enzymes associate with chromatin with different
affinities 51
DNMT3A/3B, but not DNMT1, strongly associate with
nucleosomes 53
vii
DNMT1 interacts with the linker DNA whereas DNMT3A/3B
physically associate with mononucleosomes 56
The cancer-specific ΔDNMT3B variants weakly associate with
nucleosomes 59
DNMT3B requires intact protein structure for anchoring to
nucleosomes 60
DNMT3A PWWP binds strongly to native nucleosomes through
interaction with H3K36me3 mark 63
DNMT3A/3B are enriched in methylated CpG islands and
repetitive DNA elements 67
DNMT3A/3B associate with di- and tetra-nucleosomal Alu and
LINE structures respectively 70
Discussion 73
Chapter 3: Role of Auxiliary Proteins in Anchoring of DNMT3A/3B
to Nucleosomes 79
Introduction 79
Materials and Methods 82
Results 84
Binding pattern of different DNMT3A/3B associated proteins to
nucleosomes 84
G9a and SUV39h1 strongly associate with nucleosomes similar
to DNMT3A/3B 87
G9a is not essential for maintenance of DNA methylation in
somatic cells 90
DNMT3A/3B do not require G9a for anchoring to nucleosomes 91
Discussion 96
Chapter 4: Role of DNA Methylation in Association of DNMT3A/3B
with Nucleosomes 99
Introduction 99
Materials and Methods 102
Results 108
DNMT3A protein level decreases on depletion of global DNA
methylation 108
Residual DNMT3A protein remains tightly bound to chromatin
in the DKO cells 113
Decreased protein stability of DNMT3A in hypomethylated
DKO cells 115
Restoration of global DNA methylation rescues DNMT3A
protein level 115
DNA methylation induced DNMT3A increase is mediated by
strong anchoring to nucleosomes 120
viii
Reduced nucleosome binding and degradation of unbound
DNMT3B upon depletion of DNA methylation 125
Synergistic activity of DNMT3A/3B is mediated by their
anchoring to nucleosomes 128
Discussion 134
Chapter 5: Summary and Conclusions 139
References 146
Appendix A: DNA Methylation Analysis by Digital Bisulfite Genomic
Sequencing and Digital MethyLight 168
ix
LIST OF FIGURES
Figure 1.1 Epigenetic gene silencing mechanisms in mammals 9
Figure 1.2 DNA methylation changes in cancer 16
Figure 1.3 Reprogramming of the epigenome during development and
tumorigenesis 20
Figure 1.4 Current model for inheritance of DNA methylation 36
Figure 2.1 Chromatin binding affinities of DNMTs and various other
chromatin-associated proteins 52
Figure 2.2 DNMT3A/3B, but not DNMT1, associate strongly with
polynucleosomes 55
Figure 2.3 DNMT3A/3B physically associate with mononucleosomes 57
Figure 2.4 Cancer-specific truncated delta DNMT3B variants associate
weakly with nucleosomes 61
Figure 2.5 Truncated DNMT3B proteins display weak binding to both
mono- and poly-nucleosomes 62
Figure 2.6 DNMT3A-PWWP and DNMT3B-PWWP bind to native
nucleosomes 64
Figure 2.7 DNMT3A-PWWP binds more strongly to nucleosomes than
DNMT3B-PWWP, through association with H3K36me3 mark 66
Figure 2.8 DNMT3A/3B preferentially bind to methylated repeats and
CpG islands 68
Figure 2.9 Association of DNMT3A/3B with di- and tetra-nucleosomal
structures present at Alu and LINE1 sequences respectively 71
Figure 2.10 Proposed model for inheritance of DNA methylation patterns
by DNMT1 and DNMT3A/3B 77
Figure 3.1 DNMT3A/3B along with G9a bind to intact nucleosomal
structures 85
x
Figure 3.2 G9a and SUV39h1 associate strongly with polynucleosomes
similar to DNMT3A/3B 88
Figure 3.3 G9a and SUV39h1 associate with mononucleosomes 89
Figure 3.4 Depletion of G9a does not impair maintenance of DNA
methylation in somatic cells 92
Figure 3.5 DNMT3A/3B and G9a do not require each other for
anchoring to nucleosomes in somatic cells 93
Figure 3.6 G9a cannot anchor ∆DNMT3B truncated variants to
nucleosomes 95
Figure 4.1 Transcription-independent decrease in DNMT3A protein level
in hypomethylated DKO cells 109
Figure 4.2 Residual DNMT3A protein remains localized within nuclei in
DKO cells 111
Figure 4.3 Decrease in DNMT3A level in DKO cells is not due to protein
mislocalization 112
Figure 4.4 Residual DNMT3A protein remains strongly anchored to
chromatin in hypomethylated DKO1 cells 114
Figure 4.5 Removal of DNA methylation results in decreased stability of
DNMT3A protein in DKO cells 116
Figure 4.6 Increase in DNA methylation upon expression of exogenous
DNMTs in DKO cells 118
Figure 4.7 Pre-existing methylation at genomic loci guides and stimulates
DNA methylation by DNMTs in somatic cells 119
Figure 4.8 Transcription-independent increase in DNMT3A protein level
upon increase in DNA methylation 121
Figure 4.9 The increased level of DNMT3A protein in infected DKO cells
remains tightly bound to nucleosomes 122
Figure 4.10 Recombinant DNMT3A can remain associated with nucleosomes
containing methylated DNA 124
xi
Figure 4.11 mRNA and protein expression analysis of exogenous Myc-tagged
DNMT3B1 in HCT116 derivative cell lines 126
Figure 4.12 Weak nucleosome binding and selective degradation of unbound
DNMT3B in the absence of elevated DNA methylation levels 127
Figure 4.13 DNMT3B catalytically-inactive mutant stimulates DNA
methylation by DNMT3A 130
Figure 4.14 DNMT3B catalytically-inactive mutant interacts with
DNMT3A similar to wild-type DNMT3B1 and DNMT3L 131
Figure 4.15 DNMT3B mutant stimulates DNA methylation by increasing
DNMT3A’s association with nucleosomes 133
Figure 4.16 Model for selective stabilization of DNMT3A/3B through
anchoring to nucleosomes containing methylated DNA 136
xii
ABSTRACT
Proper propagation of epigenetic information during somatic cell divisions is
critical for preserving gene expression patterns and cellular identity. However, the
molecular mechanisms responsible for faithful inheritance of epigenetic marks are still
poorly understood. In this thesis work, I have studied the inheritance of DNA methylation
patterns through somatic divisions, focusing on the role of DNA methyltransferases 3A
and 3B in this process.
DNA methylation patterns are established during development and then
maintained through multiple somatic cell divisions by co-operative activity of the de novo
and maintenance DNA methyltransferases - DNMT3A/3B and DNMT1, respectively. A
key question that remains unresolved is how the de novo DNMT3A/3B enzymes assist in
faithful inheritance of methylation patterns in somatic cells while guarding against
aberrant de novo DNA methylation. Using sucrose density gradient analyses of
fractionated chromatin, I have shown that almost all of the cellular contents of
DNMT3A/3B, but not DNMT1, are strongly anchored to nucleosomes containing
methylated DNA, allowing little free DNMT3A/3B to exist in the nucleus. This binding
of DNMT3A/3B to nucleosomes does not require the presence of other known
chromatin-associated proteins such as PCNA, HP1, MeCP2, EZH2, HDAC1, UHRF1
and G9a, but does require synergistic interactions of the conserved domains of
DNMT3A/3B with the intact nucleosomes. The PWWP domain of DNMT3A assists such
nucleosome binding through interaction with the H3K36me3 mark. Further, tight binding
xiii
of DNMT3A/3B to nucleosomes in the presence of DNA methylation stabilizes these
proteins. Drastic reduction of cellular DNA methylation levels results in a dramatic
transcription-independent decrease of DNMT3A/3B proteins due to reduced nucleosome
binding and subsequent degradation of the unstable free protein. Stabilization of
DNMT3A/3B on nucleosomes in methylated regions promotes propagation of DNA
methylation with DNMT3A/3B working synergistically in this maintenance process.
Taken together, this thesis work presents an unexpected self-regulatory inheritance
mechanism which not only ensures somatic propagation of methylated states by DNMT1
and DNMT3A/3B enzymes but also prevents aberrant de novo methylation by causing
degradation of free DNMT3A/3B enzymes.
1
CHAPTER 1
EPIGENETIC MECHANISMS IN NORMAL DEVELOPMENT AND DISEASE
INTRODUCTION
Chromatin structure defines the state in which genetic information in the form of
DNA is organized within a cell. This organization of the genome into a precise compact
structure greatly influences the abilities of genes to be activated or silenced. Epigenetics,
originally defined by C. H. Waddington as “the causal interactions between genes and
their products, which bring the phenotype into being” (Waddington, 1942), involves
understanding chromatin structure and its impact on gene function. Waddington’s
definition initially referred to the role of epigenetics in embryonic development, however,
the definition of epigenetics has evolved over time as it is implicated in a wide variety of
biological processes. The current definition of epigenetics is “the study of heritable
changes in gene expression that occur independent of changes in the primary DNA
sequence.” Most of these heritable changes are established during differentiation and are
stably maintained through multiple cycles of cell division, enabling cells to have distinct
identities while containing the same genetic information. This heritability of gene
expression patterns is mediated by epigenetic modifications, which include methylation
of cytosine bases in DNA, post-translational modifications of histone proteins as well as
the positioning of nucleosomes along the DNA. The complement of these modifications,
collectively referred to as the epigenome, provides a mechanism for cellular diversity by
2
regulating what genetic information can be accessed by cellular machinery. Failure of the
proper maintenance of heritable epigenetic marks can result in inappropriate activation or
inhibition of various signaling pathways and lead to disease states such as cancer (Egger
et al., 2004; Jones and Baylin, 2002).
Recent advances in the field of epigenetics have shown that human cancer cells
harbor global epigenetic abnormalities, in addition to numerous genetic alterations (Jones
and Baylin, 2002, 2007). These genetic and epigenetic alterations interact at all stages of
cancer development, working together to promote cancer progression (Jones and Laird,
1999). The genetic origin of cancer is widely accepted, however recent studies suggest
that epigenetic alterations may be the key initiating events in some forms of cancer
(Feinberg et al., 2006). These findings have led to a global initiative to understand the
role of epigenetics in the initiation and propagation of cancer (Jones and Martienssen,
2005). The fact that epigenetic aberrations, unlike genetic mutations, are potentially
reversible and can be restored to their normal state by epigenetic therapy makes such
initiatives promising and therapeutically relevant (Yoo and Jones, 2006).
This introductory chapter presents a comprehensive look at the current
understanding of the epigenetic mechanisms at work in normal mammalian cells and their
comparative aberrations that occur during carcinogenesis. I also discuss the idea of
cancer stem cells as the originators of cancer and the prospect of epigenetic therapy in
designing efficient strategies for cancer treatment.
3
EPIGENETIC MECHANISMS IN NORMAL CELLS
Chromatin is made of repeating units of nucleosomes, which consist of ~146 base
pairs of DNA wrapped around an octamer of four core histone proteins (H3, H4, H2A
and H2B) (Luger et al., 1997). Epigenetic mechanisms that modify chromatin structure
can be divided into four main categories: DNA methylation, covalent histone
modifications, non-covalent mechanisms such as incorporation of histone variants and
nucleosome remodeling and non-coding RNAs including microRNAs. These
modifications work together to regulate the functioning of the genome by altering the
local structural dynamics of chromatin, primarily regulating its accessibility and
compactness. The interplay of these modifications creates an “epigenetic landscape”
which regulates the way the mammalian genome manifests itself in different cell types,
developmental stages and disease states, including cancer (Bernstein et al., 2007; Jiang
and Pugh, 2009; Jones and Baylin, 2007; Kouzarides, 2007; Suzuki and Bird, 2008;
Zhang et al., 2007). The distinct patterns of these modifications present in different
cellular states serve as a guardian of cellular identity (Table 1.1). Here we will discuss the
important aspects of the key epigenetic mechanisms present in normal cells.
DNA methylation
DNA methylation is perhaps the most extensively studied epigenetic modification
in mammals. It provides a stable gene silencing mechanism which plays an important role
in regulating gene expression and chromatin architecture, in association with histone
modifications and other chromatin associated proteins. In mammals, DNA methylation
4
DNA
Methylation
All Cell Types
- Stable Heritable Modification
- Gene Silencing
- Chromatin Organization
- Imprinting, X-Chromosome Inactivation, Silencing of
Repetitive Elements
- Mediated by DNMTs
ES Cells
- Bimodal Distribution Pattern
Global CpG Methylation
CpG Islands Unmethylated
- Pluripotency Gene Promoters Unmethylated
Somatic Cells
- Tissue Specific Methylation of some CpG Islands and most
non-CpG Island Promoters
- Pluripotency Gene Promoters Methylated
Covalent
Histone
Modifications
All Cell Types
- Labile Heritable Modification
- Both Gene Silencing (H3K9me, H3K27me etc.) & Gene
Activation (H3K4me, Acetylation etc.)
- Specific Distribution Patterns of Histone Marks contribute to
Chromatin Organization
- Mediated by HMTs, HDMs, HATs & HDACs etc.
ES Cells
- Bivalent Domains - Coexistence of Active and Repressive
Marks (H3K4me & H3K27me) at promoters of
developmentally important genes
- Plastic Epigenome
Somatic Cells
- Loss of Bivalency and Restricted Epigenome
- Establishment of Tissue Specific Monovalent H3K27me and
H3K4me Domains
- Presence of Large Organized Chromatin K9 Modifications
(LOCKs)
Nucleosome
Positioning
& Histone
Variants
All Cell Types
- Labile Epigenetic Regulatory Mechanism
- Both Gene Silencing and Gene Activation by modulating
chromatin accessibility
- Mediated by ATP dependent Chromatin Remodeling
Complexes
- Both sliding of existing and incorporation of new
nucleosomes
- H2A.Z and H3.3 preferentially localized to gene promoters
which are active or poised for activation
- Acetylated H2A.Z associates with Euchromatin and
ubiquitylated H2A.Z with Facultative Heterochromatin
microRNAs
All Cell Types
- Labile Epigenetic Regulatory Mechanism
- Gene Silencing
- Tissue-specific expression
- Can be Epigenetically Regulated
Table 1.1 Epigenetic mechanisms involved in regulating gene expression and
chromatin structure in normal mammalian cells. Epigenetic mechanisms including
DNA methylation, covalent histone modifications, nucleososme positioning and
microRNAs are essential for normal mammalian development and regulation of gene
expression. The distinct combinatorial patterns of these modifications, collectively
termed the epigenome, are key determinants of cell fate and gene activity.
5
primarily occurs by the covalent modification of cytosine residues in CpG dinucleotides.
CpG dinucleotides are not evenly distributed across the human genome but are instead
concentrated in short CpG-rich DNA stretches called “CpG islands” and regions of large
repetitive sequences (e.g. centromeric repeats, retrotransposon elements, rDNA etc.)
(Bird, 2002; Takai and Jones, 2002). CpG islands are preferentially located at the 5’ end
of genes and occupy ~60% of human gene promoters (Wang and Leung, 2004). While
most of the CpG sites in the genome are methylated, the majority of CpG islands usually
remain unmethylated during development and in differentiated tissues (Suzuki and Bird,
2008). However, some CpG island promoters become methylated during development,
which results in long-term transcriptional silencing. X-chromosome inactivation and
imprinted genes are classic examples of such naturally occurring CpG island methylation
during development (Bird, 2002). Some tissue-specific CpG island methylation has also
been reported to occur in a variety of somatic tissues, primarily at developmentally
important genes (Eckhardt et al., 2006; Illingworth et al., 2008). In contrast, the repetitive
genomic sequences that are scattered all over the human genome are heavily methylated
which prevents chromosomal instability by silencing non-coding DNA and transposable
DNA elements (Suzuki and Bird, 2008). DNA methylation can lead to gene silencing by
either preventing or promoting the recruitment of regulatory proteins to DNA. For
example, It can inhibit transcriptional activation by blocking transcription factors from
accessing target binding sites e.g. c-myc and MLTF (Prendergast and Ziff, 1991; Watt
and Molloy, 1988). Alternatively, it can provide binding sites for methyl-binding domain
(MBD) proteins, which can mediate gene repression through interactions with histone
6
deacetylases (HDACs) (Jones et al., 1998; Nan et al., 1998). Thus, DNA methylation
uses a variety of mechanisms to heritably silence genes and non-coding genomic regions.
The precise DNA methylation patterns found in the mammalian genome are
generated by the de novo methyltransferases - DNMT3A and DNMT3B, which act
independent of replication and show equal preference for both unmethylated and
hemimethylated DNA. The methylation patterns are then heritably maintained through
multiple somatic cell divisions by the maintenance DNA methyltransferases - DNMT1,
which acts during replication preferentially methylating hemimethylated DNA (Kim et
al., 2002; Okano et al., 1999). However, recent studies suggest that DNMT1 alone cannot
faithfully propagate DNA methylation patterns and requires cooperative activity of the de
novo DNMT3A/3B enzymes in this process (Chen et al., 2003; Liang et al., 2002; Rhee
et al., 2002; Riggs and Xiong, 2004).
While the role of CpG island promoter methylation in gene silencing is well
established, much less is known about the role of methylation of non-CpG island
promoters. Recent studies have shown that DNA methylation is also important for the
regulation of non-CpG island promoters. For example, tissue-specific expression of
MASPIN, which does not contain a CpG island within its promoter, is regulated by DNA
methylation (Futscher et al., 2002). Similarly, methylation of the non-CpG island Oct-4
promoter, strongly influences its expression level (Hattori et al., 2004). Since CpG
islands occupy only ~60% of human gene promoters, it is essential to elucidate the role of
non-CpG island methylation in order to fully understand the global role of DNA
methylation in normal tissue (Wang and Leung, 2004).
7
Covalent histone modifications
Histone proteins, which comprise the nucleosome core, contain a globular C-
terminal domain and an unstructured N-terminal tail (Luger et al., 1997). The amino
terminal tails of nucleosomes can undergo a variety of post-translational covalent
modifications including methylation, acetylation, ubiquitylation, sumoylation, and
phosphorylation on specific residues (Kouzarides, 2007). These modifications regulate
key cellular processes such as transcription, replication and repair (Kouzarides, 2007).
The complement of modifications is proposed to store the epigenetic memory inside a
cell in the form of a “histone code” which determines the structure and activity of
different chromatin regions (Jenuwein and Allis, 2001). Histone modifications work by
either changing the accessibility of chromatin or by recruiting and/or occluding non-
histone effector proteins, which decode the message encoded by the modification
patterns. The mechanism of inheritance of this “histone code” however, is still not fully
understood.
Unlike DNA methylation, histone modifications can lead to either activation or
repression depending upon which residues are modified and the type of modifications
present. For example, lysine acetylation correlates with transcriptional activation (Hebbes
et al., 1988; Kouzarides, 2007) while lysine methylation leads to transcriptional
activation or repression depending upon which residue is modified and the degree of
methylation. For example, trimethylation of lysine 4 on histone H3 (H3K4me3) is
enriched at transcriptionally active gene promoters (Liang et al., 2004) while
trimethylation of H3K9 (H3K9me3) and H3K27 (H3K27me3) is present at gene
8
promoters that are transcriptionally repressed (Kouzarides, 2007). These two
modifications together constitute the two main silencing mechanisms in mammalian
cells, H3K9me3 working in concert with DNA methylation and H3K27me3 largely
working exclusive of DNA methylation (Figure 1.1). A vast array of active and
repressive histone modifications have been identified, which constitute a complex gene
regulatory network essential for the physiologic activities of cells (Bernstein et al., 2007;
Kouzarides, 2007). Genome-wide studies showing distinct localization and combinatorial
patterns of these histone marks in the genome have significantly increased our
understanding of how these diverse modifications act in a cooperative manner to regulate
global gene expression patterns (Barski et al., 2007; Wang et al., 2008).
Specific patterns of histone modifications are present within distinct cell types
and are proposed to play a key role in determining cellular identity (Mikkelsen et al.,
2007; Ringrose and Paro, 2007). For example, embryonic stem (ES) cells possess
“bivalent domains” which contain coexisting active (H3K4me3) and repressive
(H3K27me3) marks at promoters of developmentally important genes (Azuara et al.,
2006; Bernstein et al., 2006). Such bivalent domains are established by the activity of two
critical regulators of development in mammals: the polycomb group (PcG) which
catalyzes the repressive H3K27 trimethylation mark and is essential for maintaining ES
cell pluripotency through silencing cell-fate specific genes; and potentially the trithorax
group (trxG) which catalyzes the activating H3K4 trimethylation mark and is required for
maintaining active chromatin states during development (Ringrose and Paro, 2007). This
bivalency is hypothesized to add to phenotypic plasticity, enabling ES cells to tightly
9
Figure 1.1 Epigenetic gene silencing mechanisms in mammals. (A) An active gene
shows an open chromatin structure consisting of an unmethylated promoter region (small
white circles on DNA strands), with no nucleosome upstream of the transcription start
site (TSS, thick black arrow), an enrichment of active histone marks such as acetylation
(green triangle, Ac) and H3K4 methylation (green circles, 4) and high levels of H2A.Z on
nucleosomes (orange) surrounding the TSS. The open chromatin structure is permissible
for binding of transcription factors and RNA Pol-II, which mediate active transcription
on such promoters. Repression of such active genes (indicated by red arrows) can be
achieved in normal cells by two main mechanisms: (B) Gene repression by the action of
Polycomb Repressive Group (PRC1 and PRC2) which mediate the repressive H3K27
methylation (red circles, 27), is accompanied by the removal of acetylation by histone
deacetylases, loss of H3K4 methylation, chromatin compaction and nucleosome
occupancy in the NFR and ubiquitylation of H2A.Z; (C) Long-term silencing through
DNA methylation is performed by DNA methyltransferases. DNA methylation (small red
circles on DNA strands) is often accompanied by the repressive H3K9 methylation (red
circles, 9), on promoters, which leads to chromatin compaction by recruitment of HP1.
DNA Methylated silenced promoters show a depletion of H2A.Z, loss of H3K4
methylation and histone de-acetylation. Ac: acetylation, Ub: ubiquitination, K4-HMT:
Histone H3 lysine 4 histone methyltransferase, K9-HMT: Histone H3 lysine 9 histone
methyltransferase, HAT: histone acetyltransferase, HDAC: histone deacetylase, PRC1
and PRC2: polycomb repressive complex 1 and 2, EZH2: enhancer of zeste homologue 2,
HP1: heterochromatin protein 1, Pol-II: RNA polymerase II.
10
regulate gene expression during different developmental processes. Differentiated cells
lose this bivalency and acquire a more rigid chromatin structure, which may be important
for maintaining cell fate during cellular expansion (Mikkelsen et al., 2007). This
hypothesis is supported by the recent discovery of large condensed chromatin regions
containing the repressive H3K9me2 mark, termed “LOCKs” (large organized chromatin
K9 modifications), in differentiated ES cells which can maintain silencing of large
genomic regions in differentiated tissues (Wen et al., 2009).
Histone modification patterns are dynamically regulated by enzymes that add and
remove covalent modifications to histone proteins. Histone acetyltransferases (HATs)
and histone methyltransferases (HMTs), add acetyl and methyl groups, respectively while
histone deacetylases (HDACs) and histone demethylases (HDMs) remove acetyl and
methyl groups, respectively (Haberland et al., 2009; Shi, 2007). A number of histone
modifying enzymes including various HATs, HMTs, HDACs and HDMs have been
identified in the past decade (Kouzarides, 2007). These histone modifying enzymes
interact with each other as well as other DNA regulatory mechanisms to tightly link
chromatin state and transcription.
Interplay of DNA methylation and histone modifications
In addition to performing their individual roles, histone modifications and DNA
methylation interact with each other at multiple levels, to determine gene expression
status, chromatin organization and cellular identity (Cedar and Bergman, 2009). Several
histone methyltransferases including G9a, SUV39H1 and PRMT5, can direct DNA
11
methylation to specific genomic targets by directly recruiting DNMTs to stably silenced
genes (Lehnertz et al., 2003; Tachibana et al., 2008; Zhao et al., 2009). In addition to the
direct recruitment of DNMTs, histone methyltransferases and demethylases also
influence DNA methylation levels by regulating the stability of DNMT proteins (Esteve
et al., 2009; Wang et al., 2009). DNMTs can in turn recruit HDACs and methyl binding
proteins (MBPs) to achieve gene silencing and chromatin condensation (Jones et al.,
1998; Nan et al., 1998). DNA methylation can also direct H3K9 methylation through
effector proteins, such as MeCP2, thereby establishing a repressive chromatin state (Fuks
et al., 2003b). The interactions between DNA methylation machinery and histone
modifying enzymes further enhance the complexity of epigenetic regulation of gene
expression, which determines and maintains cellular identity and function.
Nucleosome positioning & histone variants
Non-covalent mechanisms, such as nucleosome remodeling and replacement of
canonical histone proteins with specialized histone variants, also play an important role in
how chromatin structure regulates gene activity. In addition to serving as the basic
modules for DNA packaging within a cell, nucleosomes regulate gene expression by
altering the accessibility of regulatory DNA sequences to transcription factors (Jiang and
Pugh, 2009). Genome-wide nucleosome mapping data for various eukaryotic organisms
reveal a common organizational theme with precise positioning of nucleosomes around
gene promoters, compared to the relatively random pattern found in gene bodies
(Mavrich et al., 2008). Nucleosome Free Regions (NFRs) present at the 5’ and 3’ ends of
12
genes are thought to provide the sites for assembly and disassembly of the transcription
machinery (Yuan et al., 2005). The loss of a nucleosome directly upstream of the TSS is
tightly correlated with gene activation (Lin et al., 2007; Shivaswamy et al., 2008).
Furthermore, the presence of an NFR at gene promoters with basal level of transcription
correlates with their ability for rapid activation upon stimulation (Gal-Yam, 2006). In
contrast, occlusion of the transcription start site (TSS) within the NFR by a nucleosome is
associated with gene repression (Schones et al., 2008). Modulation of the NFRs is
orchestrated by ATP-dependent chromatin remodeling complexes, which modify the
accessibility of DNA regulatory sites through both sliding and ejection of nucleosomes
(Smith and Peterson, 2005). The interaction of nucleosome remodeling machinery with
DNA methylation and histone modifications plays a pivotal role in establishing global
gene expression patterns and chromatin architecture (Figure 1.1) (Harikrishnan et al.,
2005; Wysocka et al., 2006).
In addition to physical alterations in nucleosomal positioning via nucleosome
remodelers, the incorporation of histone variants e.g. H3.3 and H2A.Z, into nucleosomes
also influences nucleosome occupancy and thus gene activity (Santenard and Torres-
Padilla, 2009; Sarma and Reinberg, 2005). Unlike the major histone subtypes whose
synthesis and incorporation is coupled to DNA replication in S phase, these variants are
synthesized and incorporated into chromatin throughout the cell cycle (Santenard and
Torres-Padilla, 2009). H3.3 and H2A.Z are preferentially enriched at promoters of active
genes or genes poised for activation and can mediate gene activation by altering the
stability of nucleosomes (Jin and Felsenfeld, 2007). H2A.Z incorporation may also
13
contribute to gene activation by protecting genes against DNA methylation (Zilberman et
al., 2008). In ES cells, H2A.Z co-localizes with bivalent domains where it may assist in
maintaining key developmental genes in a poised state (Creyghton et al., 2008). Like
canonical histones, histone variants undergo various post-translational modifications,
which determine their nuclear localization and function. For example, acetylated H2A.Z
primarily associates with active genes in euchromatin while ubiquitylated H2A.Z
associates with facultative heterochromatin (Svotelis et al., 2009; Zlatanova and Thakar,
2008). Taken together, the inclusion of histone variants within nucleosomes provides
anadditional epigenetic mechanism utilized by cells to modify chromatin structure
according to the needs of diverse cellular processes.
miRNAs
MicroRNAs (miRNAs) are small, ~22 nucleotide, non-coding RNAs that regulate
gene expression through post-transcriptional silencing of target genes. Sequence-specific
base pairing of miRNAs with 3’ untranslated regions of target mRNA within the RISC
complex (RNA-induced silencing complex) results in target mRNA degradation or
inhibition of translation (He and Hannon, 2004). miRNAs are expressed in a tissue
specific manner and control a wide array of biological processes including cell
proliferation, apoptosis and differentiation. The list of miRNAs identified in the human
genome and their potential target genes is growing rapidly, demonstrating their extensive
role in maintaining global gene expression patterns (Zhang et al., 2007). Like normal
genes, the expression of miRNAs can be regulated by epigenetic mechanisms (Saito and
14
Jones, 2006). In addition, miRNAs can also modulate epigenetic regulatory mechanisms
inside a cell by targeting enzymes responsible for DNA methylation (DNMT3A and
DNMT3B) and histone modifications (EZH2) (Fabbri et al., 2007; Friedman et al., 2009).
Such interaction among the various components of the epigenetic machinery re-
emphasizes the integrated nature of epigenetic mechanisms involved in the maintenance
of global gene expression patterns.
ABERRANT REPROGRAMMING OF THE EPIGENOME IN CANCER
The precise epigenomic landscape present in normal cells undergoes extensive
distortion in cancer (Jones and Baylin, 2007). These epimutations, along with
widespread genetic alterations, play an important role in cancer initiation and progression
(Jones and Baylin, 2002). The cancer epigenome is characterized by global changes in
DNA methylation and histone modification patterns as well as altered expression profiles
of chromatin modifying enzymes. These epigenetic changes result in global dysregulation
of gene expression profiles leading to the development and progression of disease states
(Egger et al., 2004). Epimutations can lead to silencing of tumor suppressor genes
independently and also in conjunction with deleterious genetic mutations or deletions,
thus serving as the second hit required for cancer initiation according to the “two-hit”
model proposed by Alfred Knudson (Jones and Laird, 1999). In addition to inactivating
tumor suppressors, epimutations can also promote tumorigenesis by activating
oncogenes. The events that lead to initiation of these epigenetic abnormalities are still not
fully understood. Nevertheless, since epigenetic alterations, like genetic mutations, are
15
mitotically heritable, they are selected for in a rapidly growing cancer cell population and
confer a growth advantage to tumor cells resulting in their uncontrolled growth.
DNA methylation aberrations in cancer
Cancer initiation and progression are accompanied by profound changes in DNA
methylation which were the first epigenetic alterations identified in cancer (Feinberg and
Vogelstein, 1983). A cancer epigenome is marked by genome-wide hypomethylation and
site-specific CpG island promoter hypermethylation (Figure 1.2) (Jones and Baylin,
2002). While the underlying mechanisms which initiate these global changes are still
under investigation, recent studies indicate that some changes occur very early in cancer
development and may contribute to cancer initiation (Feinberg et al., 2006).
Global DNA hypomethylation plays a significant role in tumorigenesis and occurs
at various genomic sequences including repetitive elements, retrotransposons, CpG poor
promoters, introns and gene deserts (Riggs and Jones, 1983; Rodriguez et al., 2006).
DNA hypomethylation at repeat sequences leads to increased genomic instability by
promoting chromosomal rearrangements (Eden et al., 2003; Jones and Baylin, 2002).
Hypomethylation of retrotransposons can result in their activation and translocation to
other genomic regions, thus increasing genomic instability (Howard et al., 2008).
Induction of genomic instability by hypomethylation is best exemplified in patients with
the ICF (immunodeficiency, centromeric region instability and facial anomalies)
syndrome, which have a germ-line mutation in the DNMT3B enzyme resulting in
hypomethylation and subsequent chromosomal instability (Ehrlich, 2003). Similar loss of
16
Figure 1.2 DNA methylation changes in cancer. In normal cells, CpG island promoters
are generally unmethylated and when active, as in the case of tumor suppressor genes, are
accompanied by active histone marks such as acetylation and H3K4 methylation (green
circles, 4) allowing for a transcriptionally active open chromatin structure. However,
repetitive regions, transposons, CpG poor intergenic regions and imprinted gene
promoters are heavily methylated and accompanied by repressive histone marks such as
H3K9 methylation (red circles, 9) which together form a silent chromatin state. During
tumorigenesis, tumor suppressor gene promoters with CpG islands become methylated,
resulting in the formation of silent chromatin structure and aberrant silencing (indicated
by the red arrow). In contrast, the repetitive sequences, transposons and imprinted gene
promoters become hypomethylated resulting in aberrant activation (indicated by the
green arrow).
17
DNA methylation and genomic instability is implicated in a variety of human cancers
(Howard et al., 2008). In addition, DNA hypomethylation can lead to the activation of
growth-promoting genes, such as R-Ras and MAPSIN in gastric cancer, S-100 in colon
cancer and MAGE (melanoma-associated antigen) in melanoma (Wilson et al., 2007) and
a loss of imprinting (LOI) in tumors (Rainier et al., 1993). In Wilms’ tumor,
hypomethylation induced LOI of IGF2, an important autocrine growth factor, results in
its pathological biallelic expression (Ogawa et al., 1993). LOI of IGF2 has also been
linked with an increased risk of colorectal cancer (Cui et al., 2003). Thus, DNA
hypomethylation leads to aberrant activation of genes and non-coding regions through a
variety of mechanisms which contributes to cancer development and progression.
In contrast to hypomethylation, which increases genomic instability and activates
proto-oncogenes, site-specific hypermethylation contributes to tumorigenesis by silencing
tumor suppressor genes. Since the initial discovery of CpG island hypermethylation of
the Rb promoter (a tumor suppressor gene associated with retinoblastoma) (Greger et al.,
1989), various other tumor suppressor genes including p16, MLH1 and BRCA1, have also
been shown to undergo tumor-specific silencing by hypermethylation (Baylin, 2005;
Jones and Baylin, 2002, 2007). These genes are involved in cellular processes, which are
integral to cancer development and progression, including DNA repair, cell cycle, cell
adhesion, apoptosis and angiogenesis. Epigenetic silencing of such tumor suppressor
genes can also lead to tumor initiation by serving as the second hit in the Knudson’s
“two-hit” model (Jones and Laird, 1999). In addition to direct inactivation of tumor
suppressor genes, DNA hypermethylation can also indirectly silence additional classes of
18
genes by silencing transcription factors and DNA-repair genes. Promoter
hypermethylation induced silencing of transcription factors such as RUNX3 in esophageal
cancer (Long et al., 2007), and GATA-4 and GATA-5 in colorectal and gastric cancers
(Akiyama et al., 2003), leads to inactivation of their downstream targets. Silencing of
DNA-repair genes (e.g. MLH1, BRCA1 etc.) enables cells to accumulate further genetic
lesions leading to the rapid progression of cancer.
While the ability of DNA hypermethylation to silence tumor suppressor genes in
cancer is well established, how genes are targeted for this aberrant DNA methylation is
still unclear. One possibility is that silencing specific genes by hypermethylation provides
a growth advantage to cells resulting in their clonal selection and proliferation. Tumor-
specific CpG island methylation can occur through a sequence specific instructive
mechanism by which DNMTs are targeted to specific genes by their association with
oncogenic transcription factors. Aberrant hypermethylation and silencing of specific
target gene promoters by the PML-RAR fusion protein in acute promyelocytic leukemia
is an example of such a mechanism (Di Croce et al., 2002). Large stretches of DNA can
become abnormally methylated in cancer (Frigola et al., 2006) causing some CpG islands
to be hypermethylated as a result of their location inside such genomic regions which
have undergone large-scale epigenetic reprogramming. Another interesting mechanism
proposes a role of histone marks in the tumor-specific targeting of de novo methylation
and will be discussed in detail in the next section. Interestingly, regions that are
hypermethylated in cancer are often pre-marked with H3K27me3 polycomb mark in
embryonic stem (ES) cells suggesting a link between the regulation of development and
19
tumorigenesis (Figure 1.3) (Ohm et al., 2007; Schlesinger et al., 2007; Widschwendter et
al., 2007). This observation also partially explains the theory of “CpG island methylator
phenotype” or CIMP which hypothesizes that there is coordinated methylation of a subset
of CpG islands in tumors since many of these CIMP loci are known polycomb targets
(Weisenberger et al., 2006; Widschwendter et al., 2007). Further understanding of how
specific genomic regions are targeted for DNA hypermethylation in cancer will
potentially lead to additional therapeutic targets.
Changes in histone modifications in cancer
Recent advances in high throughput sequencing have enabled genome wide
mapping of chromatin changes occurring during tumorigenesis. These studies have
revealed a global loss of acetylated H4-lysine 16 (H4K16ac) and H4-lysine 20
trimethylation (H4K20me3) (Fraga et al., 2005). Such loss of histone acetylation, which
is mediated by histone deacetylases (HDACs) results in gene repression. HDACs are
often found overexpressed in various types of cancer (Halkidou et al., 2004; Song et al.,
2005) and thus, have become a major target for epigenetic therapy. Histone
acetyltransferases (HATs), which work in concert with HDACs to maintain histone
acetylation levels, can also be altered in cancer. Aberrant formation of fusion proteins
through chromosomal translocations of HAT and HAT-related genes (e.g. MOZ, MORF,
CBP and p300) occurs in leukemia (Yang, 2004). Mistargeting of such deleterious fusion
proteins contributes to global alterations in histone acetylation patterns in cancer.
20
Figure 1.3 Reprogramming of the epigenome during development and
tumorigenesis. (A) In embryonic stem cells, developmentally important genes are
marked by a unique “bivalent domain” structure, consisting of the active H3K4
methylation (green circles, 4) and repressive H3K27 methylation (red circles, 27) marks
together with H2A.Z. Such bivalent domains are important for maintaining epigenomic
plasticity that is required during development. During differentiation, the “bivalent
domains” are lost, giving way to the establishment of a more rigid “monovalent domain”
structure which is either active (indicated by the green arrow) or repressive (indicated
by the red arrow) depending upon which mark is maintained. (B) In cancer, cells undergo
aberrant somatic reprogramming which results in gene silencing through formation of a
compact chromatin structure. Silencing can occur through PRC (Polycomb Repressive
Complex) reprogramming – silencing of active genes by the polycomb group; DNA
methylation reprogramming – silencing through de novo hypermethylation (small red
circles on DNA strands) accompanied by H3K9 methylation (red circles, 9); or epigenetic
switching – replacement of gene repression by the polycomb mark with long-term
silencing through DNA methylation. Ub: ubiquitylation.
21
In addition to changes in histone acetylation, cancer cells also display widespread
changes in histone methylation patterns. Alterations in H3K9 and H3K27 methylation
patterns are associated with aberrant gene silencing in various forms of cancer (Nguyen
et al., 2002; Valk-Lingbeek et al., 2004). Dysregulation of histone methyltransferases
responsible for repressive marks results in altered distribution of these marks in cancer
and leads to aberrant silencing of tumor suppressor genes. For example, EZH2, which is
the H3K27 histone methyltransferase, is overexpressed in breast and prostate cancer
(Valk-Lingbeek et al., 2004). Increased levels of G9a, the H3K9 histone
methyltransferase, has been found in liver cancer, and is implicated in perpetuating
malignant phenotype possibly through modulation of chromatin structure (Kondo et al.,
2008; Kondo et al., 2007) . Chromosomal translocations of MLL, the H3K4 histone
methyltransferase, leads to ectopic expression of various homeotic (Hox) genes and plays
a key role in leukemic progression (Krivtsov and Armstrong, 2007).
In addition to histone methyltransferases, lysine specific-demethylases which
work in coordination with HMTs to maintain global histone methylation patterns, are also
implicated in cancer progression (Cloos et al., 2008). LSD1, the first identified lysine-
demethylase, can effectively remove both activating and repressing marks (H3K4 and
H3K9 methylation, respectively) depending on its specific binding partners (Metzger et
al., 2005; Shi et al., 2004), thus, acting as either a co-repressor or a co-activator. After
LSD1, several other histone lysine demethylases have been discovered including Jumonji
C domain (JmjC) proteins. Several of these histone demethylases are upregulated in
prostate cancer, thus making them potential therapeutic targets (Shi, 2007). However,
22
since histone demethylases, like LSD1, can perform both activating and repressive
functions, it is essential to first understand their precise context-dependent roles before
their therapeutic inhibition can be used as an effective cancer treatment strategy. Despite
these challenges, targeting histone demethylases is a promising treatment option for the
future as revealed by a recent study which showed that inhibition of LSD1 in
neuroblastoma, causes decreased proliferation in vitro and inhibition of xenograft growth
(Schulte et al., 2009).
Epigenetic switching in cancer
As mentioned previously, DNA methylation and histone modifications work
independently and in concert to alter gene expression during tumorigenesis. A key facet
of such silencing mechanisms is the formation of a rigid repressive chromatin state which
results in reduced cellular plasticity. The recent discovery of tumor-specific de novo
methylation of polycomb target genes, which are silenced by H3K27me3 in normal cells,
is another example of this phenomenon (Ohm et al., 2007; Schlesinger et al., 2007;
Widschwendter et al., 2007). In ES cells, developmentally important genes are reversibly
silenced by polycomb proteins through the establishment of the repressive H3K27me3
mark. After differentiation, these genes continue to be repressed through the maintenance
of the polycomb mark on unmethylated promoters by EZH2. In cancer, the polycomb
mark is replaced by de novo DNA methylation possibly through the recruitment of
DNMTs via the polycomb complex (Vire et al., 2006). This tumor-specific “epigenetic
switching” of the plastic polycomb mark with more stable DNA methylation results in
23
the permanent silencing of key regulatory genes which may contribute to cell
proliferation and tumorigenesis (Figure 1.3) (Gal-Yam et al., 2008). However, which
transformation-associated factors trigger this switch is still unclear.
Role of nucleosome positioning in cancer
The roles of DNA methylation and histone modifications in cancer initiation and
progression are well established, however the changes in chromatin structure that
accompany DNA methylation and histone modification changes are less well understood.
Emerging data has revealed that nucleosome remodeling works in concert with DNA
methylation and histone modifications, and plays a central role in tumor-specific gene
silencing. DNA methylation induced silencing of tumor suppressor genes in cancer
involves distinct changes in nucleosome positioning resulting in nucleosome occupancy
at TSS (Figure 1.2). Reactivation of such silenced genes using DNMT inhibitors is
accompanied by a loss of nucleosomes from the promoter region (Lin et al., 2007). In
addition, nucleosome remodeling can lead to aberrant gene silencing via the transmission
of repressive epigenetic marks to tumor suppressor gene promoters. Recent work by
Morey et al. (2008) demonstrated that nucleosome remodeling and deacetylase
corepressor complex (NuRD) plays a central role in aberrant gene silencing in leukemia
via the oncogenic transcription factor, PML-RARa. The NuRD complex facilitates
recruitment of the polycomb repressive complex 2 (PRC2) and DNMT3A to PML-RARa
target gene promoters leading to their permanent silencing by establishing a repressive
chromatin state (Morey et al., 2008). NuRD can also be recruited to methylated
24
promoters through its interaction with the MBD2 protein (Feng and Zhang, 2001).
Sustained binding of NuRD to such promoters may assist in preserving their repressive
state through maintenance of DNA methylation.
Alterations in the SWI/SNF complex, an ATP-dependent chromatin remodeling
complex, are also associated with cancer development (Reisman et al., 2009). Abrogation
of SWI/SNF function through alterations in its various subunits can result in malignant
transformation. The BAF47 (hSNF5) subunit of the SWI/SNF complex is a bona fide
tumor suppressor and its inhibition in rhabdoid tumors causes inactivation of the p21 and
p16 pathways leading to oncogenic transformation (Chai et al., 2005). Furthermore,
BRG1 and BRM, the catalytic subunits of SWI/SNF, are silenced in about 15-20% of
primary non-small-cell lung cancers (Reisman et al., 2003). Treatment of BRM null cell
lines with HDAC inhibitors has been shown to restore its expression thus making it a
promising target for epigenetic therapy. However, such treatment also resulted in
acetylation of BRM protein which abrogated its function (Reisman et al., 2009).
Development of specific HDAC inhibitors, which can circumvent BRM acetylation, is
essential for successful induction of functional BRM in tumors, which can be used as a
prospective therapeutic target in the future.
Interestingly, a context dependent oncogenic role of BRG1 has also been
proposed. Work by Naidu et al. (2009) reveals that BRG1 contributes to cancer
development by constraining p53 activity through the destabilization of the p53 protein.
Opposing roles of SWI/SNF subunits highlights the requirement for a deeper
25
understanding of the role of nucleosome remodeling in cancer development in order to
develop effective tumor-specific therapies (Naidu et al., 2009).
In addition to remodeling complexes, the histone variant H2A.Z has also been
implicated in tumorigenesis. H2A.Z is overexpressed in several types of cancer and has
been associated with the promotion of cell cycle progression (Svotelis et al., 2009).
Interestingly, loss of H2A.Z has also been implicated in tumor progression through
possible destabilization of chromosomal boundaries resulting in spreading of repressive
chromatin domains and de novo hypermethylation of tumor-suppressor gene promoters
(Witcher and Emerson, 2009).
Deregulation of miRNAs in cancer
Accumulating evidence from studies comparing miRNA expression profiles in
tumors and corresponding normal tissues indicate widespread changes in miRNA
expression during tumorigenesis (Lu et al., 2005). Since miRNAs regulate genes involved
in transcriptional regulation, cell proliferation and apoptosis (the most common processes
deregulated in cancer), alteration in their expression can promote tumorigenesis. miRNAs
can function as either tumor suppressors or oncogenes depending upon their target genes.
Many tumor suppressor miRNAs that target growth promoting genes are repressed in
cancer. For example, miR-15 and 16 which target BCL2, an anti-apoptotic gene, are
downregulated in chronic lymphocytic leukemia, while let-7 which targets the oncogene,
RAS, is downregulated in lung cancer (Ventura and Jacks, 2009; Zhang et al., 2007).
Furthermore, miR-127, which targets BCL6, is significantly downregulated in prostate
26
and bladder tumors (Saito et al., 2006) and mir-101, which targets polycomb group
protein EZH2, is downregulated in bladder transitional cell carcinoma (Friedman et al.,
2009). In contrast to tumor suppressor miRNAs, oncogenic miRNAs, which target
growth inhibitory pathways, are often upregulated in cancer. For example, miR-21, which
targets PTEN, is upregulated in human glioblastoma (Chan et al., 2005). miRNA-155 is
upregulated in breast, lung and several hematopoietic malignancies (Kluiver et al., 2006).
While the exact mechanism of action of miR-155 is still unclear, there are suggestions
that it may play a role in class switch recombination process by targeting AID
(activation-induced cytidine deaminase) (Dorsett et al., 2008). The oncogenic miR-17~92
cluster, which targets pro-apoptotic gene Bim, is found overexpressed in several kinds of
cancer (Mendell, 2008).
Changes in miRNA expression can be achieved through various mechanisms
including chromosomal abnormalities, transcription factor binding and epigenetic
alterations (Deng et al., 2008). The initial report by Saito et al. (2006), which
demonstrated that miR-127, a tumor suppressor miRNA embedded in a CpG island, was
silenced in cancer by DNA methylation, has led to subsequent discovery of several other
miRNAs which are also silenced by epigenetic mechanisms in cancer (Lujambio et al.,
2008, Toyota, 2008 #301; Toyota et al., 2008). Since such epigenetic repression of tumor
suppressor miRNAs can be potentially reversed by treatment with chromatin modifying
drugs, they can serve as promising targets for epigenetic therapy. Saito et al. successfully
demonstrated re-activation of miR-127 in T24 bladder cancer cells following treatment
with chromatin modifying drugs including DNA methylation and HDAC inhibitors
27
(Egger et al., 2006; Saito et al., 2006). Such drug induced activation of tumor suppressor
miRNAs holds great promise for the future of cancer therapeutics.
The cancer stem cell model
Recent work suggests that the global epigenetic changes in cancer, may involve
the dysregulation of hundreds of genes during tumorigenesis. The mechanism by which a
tumor cell accumulates such widespread epigenetic abnormalities during cancer
development is still not fully understood. The selective advantage of these epimutations
during tumor progression is possible, but it is unlikely that the multitude of epigenetic
alterations that reside in a cancer epigenome occur in a random fashion and then
accumulate inside the tumor due to clonal selection. A more plausible explanation would
be that the accumulation of such global epigenomic abnormalities arises from initial
alterations in the central epigenetic control machinery, which occur at a very early stage
of neoplastic evolution. Such initiating events can predispose tumor cells to gain further
epimutations during tumor progression in a fashion similar to accumulation of the genetic
alterations that occurs following defects in DNA repair machinery in cancer. The “cancer
stem cell” model suggests that the epigenetic changes, which occur in normal stem or
progenitor cells, are the earliest events in cancer initiation (Feinberg et al., 2006). The
idea that these initial events occur in stem cell populations is supported by the common
finding that epigenetic aberrations are some of the earliest events that occur in various
types of cancer and also by the discovery that normal tissues have altered progenitor cells
in cancer patients (Cui et al., 2003; Matsubayashi et al., 2003; Peters et al., 2007). This
28
stem cell based cancer initiation model is consistent with the observation that tumors
contain a heterogeneous population of cells with diverse tumorigenic properties (Al-Hajj
et al., 2003). Since epigenetic mechanisms are central to maintenance of stem cell
identity (Mikkelsen et al., 2007; Surani et al., 2007), it is reasonable to speculate that
their disruption may give rise to a high risk aberrant progenitor cell population which can
undergo transformation on gain of subsequent genetic gatekeeper mutations. This
epigenetic disruption can lead to an overall increase in number of progenitor cells along
with an increase in their ability to maintain their stem cell state, forming a high risk
substrate population which can readily become neoplastic on gain of additional genetic
mutations (Jones and Baylin, 2007).
Several findings have recently emerged in support of the “cancer stem cell”
model. Mice with a loss of imprinting (LOI) at the IGF2 locus and an Apc mutation show
an expansion in the progenitor cell population of the intestinal epithelium, with the
epithelial cells showing higher expression of progenitor cell markers and shifting towards
a less-differentiated state (Sakatani et al., 2005). These mice were also at a higher risk for
intestinal tumors relative to control mice (Sakatani et al., 2005). Interestingly, humans
with LOI of IGF2 also show a similar de-differentiation of normal colonic mucosa cells
along with a higher risk for colorectal cancer (Cui et al., 2003). Also, stem cell-like
characteristics of tumor cells were displayed through successful cloning of mouse
melanoma and medulloblastoma nuclei to form blastocysts and chimeric mice
(Hochedlinger et al., 2004).
29
DNA methylation induced silencing of genes involved in the regulation of stem/
precursor cells’ self renewal capacity, such as p16, APC, SFRPs etc., is commonly
observed in the early stages of colon and other cancers (Jones and Baylin, 2007).
Aberrant silencing of these so called “epigenetic gatekeeper” genes in conditions of
chronic stress, such as inflammation, enables stem/precursor cells to gain infinite renewal
capacity thereby becoming immortal. These pre-invasive immortal stem cells are selected
for and then form a pool of abnormal precursor cells that can undergo further genetic
mutations leading to tumorigenesis (Baylin and Ohm, 2006; Jones and Baylin, 2007) .
Human ES cells with cancer-cell characteristics including higher frequency of teratoma-
initiating cells (TICs), growth factor and niche independence have also been found
(Werbowetski-Ogilvie et al., 2009). These partially transformed stem cells display a
higher expression of pluripotency markers suggesting their enhanced “stemness” along
with high proliferative capacity (Werbowetski-Ogilvie et al., 2009).
Polycomb proteins, which control the silencing of developmental regulators in ES
cells, provide another link between stem cell biology and cancer initiation. Polycomb
proteins are commonly upregulated in various forms of cancer (Valk-Lingbeek et al.,
2004). In addition, genes that are marked by polycomb repressive mark H3K27me3 in ES
cells are often methylated in cancer suggesting the presence of a shared regulatory
framework, which connects cancer cells with stem/progenitor cell populations. Such
findings support the hypothesis of epigenetics playing a central role in early neoplasia
and cancer stem cells being the key perpetuators of cancer (Schlesinger et al., 2007;
Widschwendter et al., 2007).
30
EPIGENETIC THERAPY OF CANCER
The reversible nature of the profound epigenetic changes that occur in cancer has
led to the possibility of “epigenetic therapy” as a treatment option. The aim of epigenetic
therapy is to reverse the causal epigenetic aberrations that occur in cancer, leading to the
restoration of a “normal epigenome”. Many epigenetic drugs have been discovered in the
recent past that can effectively reverse DNA methylation and histone modification
aberrations that occur in cancer (Yoo and Jones, 2006). DNA methylation inhibitors were
among the first epigenetic drugs proposed for use as cancer therapeutics. The remarkable
discovery that treatment with cytotoxic agents, 5-azacytidine (5-aza-CR) and 5-aza-2’-
deoxycytidine (5-aza-CdR) lead to the inhibition of DNA methylation which induced
gene expression and caused differentiation in cultured cells led to the realization of the
potential use of these drugs in cancer therapy (Constantinides et al., 1977). These
nucleoside analogs get incorporated into the DNA of rapidly growing tumor cells during
replication and inhibit DNA methylation by trapping DNA methyltransferases onto the
DNA, leading to their depletion inside the cell (Egger et al., 2004). This drug induced
reduction of DNA methylation causes growth inhibition in cancer cells by activating
tumor suppressor genes aberrantly silenced in cancer (Yoo and Jones, 2006). 5-aza-CR
(azacitidine) and 5-aza-CdR (decitabine) have now been FDA approved for use in the
treatment of myelodysplastic syndromes (MDS) and promising results have also emerged
from the treatment of other hematological malignancies such as acute myeloid leukemia
(AML) and chronic myeloid leukemia (CML) using these drugs (Plimack et al., 2007).
The possible clinical use of other improved DNA methylation inhibitors such as
31
zebularine, which can be orally administered, is currently under investigation (Cheng et
al., 2004).
The ability of these drugs to be incorporated into DNA raises concerns regarding
their potential toxic effect on normal cells. However, since these drugs only act on
dividing cells, one can argue that treatment with these drugs should mainly target rapidly
dividing tumor cells and should have minimal effects on slowly dividing normal cells.
This argument has been supported by studies demonstrating minimal side effects of long-
term treatment with DNA methylation inhibitors (Yang et al., 2003). Nevertheless, an
alternative approach involving the development of non-nucleoside compounds, which can
effectively inhibit DNA methylation without being incorporated into DNA, is also being
actively pursued. Development of several small molecule inhibitors such as SGI-1027,
RG108 and MG98 is a step in that direction (Cortez and Jones, 2008; Datta et al., 2009).
These molecules can achieve their inhibitory effects by either blocking catalytic/cofactor
binding sites of DNMTs or by targeting their regulatory mRNA sequences; however, the
weak inhibitory potential of these drugs indicates a need for the development of more
potent inhibitory compounds in future.
Aberrant gene silencing in cancer is also associated with a concomitant loss of
histone acetylation. Re-establishing normal histone acetylation patterns through treatment
with HDAC inhibitors has been shown to have anti-tumorigenic effects including growth
arrest, apoptosis and the induction of differentiation. These anti-proliferative effects of
HDAC inhibitors are mediated by their ability to reactivate silenced tumor suppressor
genes (Carew et al., 2008). SAHA (suberoylanilide hydroxamic acid), which is an HDAC
32
inhibitor, has now been approved for use in clinic for treatment of T cell cutaneous
lymphoma. Several other HDAC inhibitors such as depsipeptide and phenylbutyrate are
currently under clinical trials (Cortez and Jones, 2008).
The interaction between different components of the epigenetic machinery has led
to the exploration of effective combinatorial cancer treatment strategies, which involve
use of both DNA methylation and HDAC inhibitors together. Such combination
treatment strategies have been found to be more effective than individual treatment
approaches. For example, the de-repression of certain putative tumor suppressor genes
was only seen when 5-Aza-CdR and trichostatin A were combined (Cameron et al.,
1999). Anti-tumorigenic effects of depsipeptide were enhanced when leukemic cells were
simultaneously treated with 5-Aza-CdR (Klisovic et al., 2003). Synergistic activities of
DNA methylation and HDAC inhibitors were also demonstrated in a study showing
greater reduction of lung tumor formation in mice when treated with phenylbutyrate and
5-Aza-CdR together (Belinsky et al., 2003).
Apart from DNA methylation and HDAC inhibitors, histone methyltransferase
inhibitors have also been actively explored recently. One such inhibitor compound,
DZNep, was shown to successfully induce apoptosis in cancer cells by selectively
targeting polycomb repressive complex 2 (PRC2) proteins, which are generally
overexpressed in cancer (Tan et al., 2007). While the specificity of DZNep was
challenged in a subsequent study (Miranda et al., 2009), these findings reinforce the
potential of HMT inhibitors and the need for further development of specific histone
methylation inhibitors.
33
miRNAs also represent promising targets for epigenetic therapy. The finding by
Saito et al. (2006) that downregulation of the oncogene BCL6 via re-activation of miR-
127 following treatment with 5-Aza-CdR and 4-phenylbutyric acid strongly advocates in
favor of the potential of a miRNA based treatment strategy (Saito et al., 2006). In
addition, the introduction of synthetic miRNAs, which mimic tumor suppressor miRNAs,
can be used to selectively repress oncogenes in tumors. miRNAs, such as miR-101 which
targets EZH2 (Friedman et al., 2009), can be used to regulate the aberrant epigenetic
machinery in cancer which may assist in restoring of the normal epigenome. However,
the lack of efficient delivery methods is a major hurdle in the effective use of this
strategy. Development of efficient vehicle molecules for targeted delivery of synthetic
miRNAs to tumor cells is of prime importance in future.
FUTURE PROSPECTS AND CHALLENGES
The epigenetic revolution that has come about in the field of biology during the
last few decades has challenged the long-held traditional view of the genetic code being
the key determinant of cellular gene function and its alteration being the major cause of
human diseases. Advances made in the field of cancer epigenetics have led to the
realization that the packaging of the genome is potentially as important as the genome
itself, in regulating the essential cellular processes required for preserving cellular
identity and also in giving rise to disease states like cancer. Deeper understandings of the
global patterns of these epigenetic modifications and their corresponding changes in
cancer have enabled the design of better treatment strategies. A combinatorial approach
34
utilizing different epigenetic therapeutic approaches along with standard chemotherapy
holds significant promise for successful treatment of cancer in future. Such approaches
might also help in sensitizing cancer cells, especially cancer stem cells, which are
refractory to standard chemotherapy. Further understanding of the mechanisms required
for the maintenance of normal epigenome and their alterations which give rise to the
malignant phenotype, along with development of more specific epigenetic drugs may
hold the key to our ability to successfully reset the abnormal cancer epigenome.
35
OVERVIEW OF THESIS RESEARCH
As summarized already, the epigenome consists of several layers of heritable
transcriptional regulations imposed upon the genome, including DNA methylation,
histone modifications and nucleosome positioning. These epigenetic mechanisms are
essential for normal development and maintenance of tissue specific gene expression
patterns (Bird, 2002; Suzuki and Bird, 2008). Proper inheritance of epigenetic
modifications through cell divisions is critical for preserving cellular identity and
preventing malignant cellular transformation (Jones and Baylin, 2007; Sharma et al.,
2010). Here I have studied the process of inheritance of DNA methylation patterns
through somatic divisions, focusing on the role of the de novo DNA methyltransferases
3A and 3B (DNMT3A/3B). According to the existing model, DNA methylation patterns
are established during development by the de novo DNMT3A/3B enzymes and then
stably propagated through somatic divisions by the maintenance activity of DNMT1
(Figure 1.4). However, recent studies indicate that DNMT1 alone cannot faithfully
maintain methylation patterns and requires co-operative activity of DNMT3A/3B in this
process, particularly for maintenance of methylation at repetitive elements (Chen et al.,
2003; Liang et al., 2002; Rhee et al., 2002; Riggs and Xiong, 2004). The mechanisms
through which DNMT3A/3B may perform this maintenance role remain unclear.
Genomic DNA exists in the form of chromatin within nuclei. Thus, to ascertain
the roles of DNMTs in inheritance of DNA methylation, it is important to first understand
their interactions with chromatin. I started my work on this project by examining the
interaction of DNMT3A/3B and DNMT1 enzymes with chromatin as described in
36
Figure 1.4 Current model for inheritance of DNA methylation. The current model
focuses primarily on the role of DNMT1, the maintenance methyltransferase, in
propagation of methylation patterns through cell divisions. During replication, DNMT1
(in purple) is proposed to read the methylation patterns on the template strand (in the
form of hemimethylated DNA) and copies it to the daughter strand through its association
with the replication fork by proliferating cell nuclear antigen (PCNA, in blue) and
possibly by ubiquitin-like plant homeodomain and ring finger domain containing protein
1 (UHRF1, in yellow).
Old DNA strand
New DNA strand
Replication
Nucleosome
Methylated/unmethylated
CpG site
Repeat region
DNMT1 DNMT1
UHRF1 UHRF1
PCNA PCNA
37
Chapter 2. We used sucrose density gradients of nucleosomes prepared by partial and
limit micrococcal nuclease digestion, coupled with Western Blot analysis for studying
interactions of chromatin-associated proteins with native nucleosomes. Our initial work
showed that little free DNMT3A/3B exist in the nucleus and that almost all of the cellular
contents of DNMT3A/3B, but not DNMT1, are strongly anchored to a subset of
nucleosomes. Strong binding of DNMT3B protein to nucleosomes requires a full-length
protein. The truncated ΔDNMT3B isoforms, which are expressed specifically in cancer,
weakly associate with nucleosomes which may play a role in the aberrant DNA
methylation associated with these isoforms. Analysis of individual DNMT3A/3B
domains revealed that the PWWP domain of DNMT3A interacts strongly with native
nucleosomes and may assist in the association of DNMT3A with nucleosomes through
interaction with H3K36me3 mark. Further, our data show that DNMT3A/3B bind
specifically to nucleosomes containing methylated repetitive elements and methylated
CpG islands. These data suggest specific compartmentalization of these enzymes to their
target chromatin domains.
Targeting of DNMT3A/3B enzymes to specific chromatin domains may be
mediated by recruitment through other chromatin-associated proteins in addition to direct
interaction of DNMT3A/3B enzymes with the nucleosomes. In Chapter 3, I investigated
the role of accessory proteins, known to interact with DNMT3A/3B, in their strong
anchoring to nucleosomes. Our data shows that binding of DNMT3A/3B does not require
the presence of other well known chromatin modifying enzymes or proteins such as
PCNA, HP1, MeCP2, EZH2, HDAC1 and UHRF1, but does require an intact
38
nucleosomal structure. The H3K9 methyltransferases, G9a and SUV39h1, bind strongly
to both mononucleosomes and polynucleosomes similar to that observed for
DNMT3A/3B. However, knockdown of G9a revealed that G9a is not required for
DNMT3A/3B binding to the nucleosomes and for maintenance of DNA methylation in
somatic cells, suggesting existence of other mechanisms involved in this process.
Recently, a common theme for inheritance of histone marks has emerged where
the mark recruits and retains its own modifying enzyme and triggers renewal by
stimulating that enzyme through possible allosteric activation mechanisms (Collins et al.,
2008; Felsenfeld and Groudine, 2003; Hansen et al., 2008). Our previous data showed
that the majority of DNMT3A/3B within a somatic cell remain associated with their
products through anchoring to nucleosomes containing methylated DNA. This prompted
me to investigate the existence of a self-renewal mechanism in inheritance of DNA
methylation by DNMT3A/3B similar to that observed for histone marks. In Chapter 4, I
have shown that the presence of DNA methylated regions is essential for DNMT3A/3B’s
association with chromatin and for maintaining their cellular levels. Anchoring to
nucleosomes containing methylated DNA stabilizes DNMT3A/3B proteins. Reduction in
DNA methylation levels results in reduced DNMT3A/3B binding to nucleosomes
accompanied by selective degradation of the free enzymes by the cellular machinery.
Further, pre-existing methylation stimulates DNA methylation propagation by stabilizing
DNMT3A/3B on nucleosomes and DNMT3A/3B work synergistically to propagate DNA
methylation patterns in somatic cells.
39
Taken together, my work has revealed an unanticipated mechanism for
inheritance of DNA methylation patterns where the epigenetic mark not only recruits the
catalyzing enzyme but also regulates the protein level, i.e. the enzymatic product (5-
methylcytosine) determines the level of the methylase, thus forming a novel homeostatic
inheritance system. Such a mechanism not only ensures faithful somatic propagation of
methylated states by DNMT1 and DNMT3A/3B enzymes but also prevents aberrant de
novo methylation by causing degradation of free DNMT3A/3B enzymes.
40
CHAPTER 2
STRONG ANCHORING OF DNMT3A/3B TO
NUCLEOSOMES CONTAINING METHYLATED DNA
INTRODUCTION
DNA methylation is a stable gene silencing mechanism required for key
biological processes including embryogenesis, genomic imprinting, X-chromosome
inactivation, repression of transposons and maintenance of tissue specific gene
expression patterns (Bird, 2002; Suzuki and Bird, 2008). Aberrant methylation
contributes to tumorigenesis and other diseases (Jeong et al.; Jones and Baylin, 2007).
Thus, proper maintenance of DNA methylation patterns is essential for preserving
cellular identity and preventing malignant cellular transformation.
In mammals, DNA methyltransferases (DNMTs), DNMT3A, DNMT3B and
DNMT1, primarily establish and maintain global DNA methylation patterns (Li et al.,
1992; Okano et al., 1999). A model explaining inheritance of DNA methylation patterns
through somatic divisions was proposed more than three decades ago by Riggs (Riggs,
1975) and Holiday and Pugh (Holliday and Pugh, 1975); however, the exact molecular
mechanisms responsible for the faithful propagation of methylation patterns are still
poorly understood. It has been proposed that DNMT1 acts mainly as a “maintenance
methyltransferase” during DNA synthesis, and DNMT3A and DNMT3B act, as “de
novo” enzymes. In accordance with this hypothesis, studies have shown that DNMT1
41
preferentially methylates hemimethylated DNA in vitro (Bestor and Ingram, 1983), and is
tethered to replication foci during S-phase (Leonhardt et al., 1992). In contrast, DNMT3A
and DNMT3B (DNMT3A/3B) have no preference for hemimethylated DNA (Okano et
al., 1998), and are required for de novo methylation of genomic DNA (Okano et al.,
1999). However, more recent studies indicate that DNMT1 may also be required for de
novo methylation of genomic DNA (Egger et al., 2006; Jair et al., 2006), and Dnmt3a/3b
are also required for maintenance functions (Chen et al., 2003; Liang et al., 2002; Riggs
and Xiong, 2004). Furthermore, the different DNMTs cooperate in maintaining the
methylation of certain regions of the genome, particularly repetitive elements (Liang et
al., 2002; Rhee et al., 2002). Though the maintenance role of DNMT1 in propagation of
methylation patterns has been well elucidated; the mechanisms through which the de
novo DNMT3A/3B enzymes assist in the maintenance process are still unclear.
In somatic cells, DNMT1 is found diffusely localized throughout nuclei in non-S-
phase cells (Leonhardt et al., 1992) and has been suggested to perform its maintenance
activity during replication by associating with the replication foci through interactions
with proliferating cell nuclear antigen (PCNA) (Chuang et al., 1997) and ubiquitin-like,
containing PHD and RING finger domains 1 (UHRF1) that binds to hemimethylated
DNA (Arita et al., 2008; Avvakumov et al., 2008; Bostick et al., 2007; Hashimoto et al.,
2008; Sharif et al., 2007). In contrast to the dynamic interactions of DNMT1 with the
chromatin, DNMT3A/3B enzymes are usually found tightly associated with
heterochromatin regions in most transient expression assays (Bachman et al., 2001; Chen
et al., 2002) suggesting a different mechanistic basis for their maintenance activity. As
42
genomic DNA in chromatin is packaged into nucleosomes, which limit the accessibility
of target sites to the enzymes, the nature of DNMTs interactions with nucleosomes would
be critical for their ability to faithfully maintain genomic methylation.
Genetic and biochemical studies have provided insights into the distinct and
cooperative functions of the DNMT enzymes, however few of these studies have
addressed how they interact with chromatin in vivo. Targeting of DNMT3A/3B enzymes
to specific chromatin regions may involve direct interaction of their unique domains with
certain chromatin marks. It may also be mediated by interaction with accessory proteins
which can recruit DNMT3A/3B to chromatin domains in vivo (discussed later in Chapter
3). Recombinant DNMT3A/3B enzymes can indeed methylate the CpG sites on
nucleosomes assembled in vitro , suggesting ability of these enzymes to associate directly
with the chromatin through their domains (Gowher et al., 2005b; Okuwaki and Verreault,
2004; Robertson et al., 2004; Takeshima et al., 2006). DNMT3A/3B enzymes contain an
N-terminal regulatory region having a PWWP domain and a PHD-like zinc finger domain
also known as ADD (ATRX-DNMT3-DNMT3L) domain (Goll and Bestor, 2005;
Hermann et al., 2004). The PWWP domain of Dnmt3a and 3b has been shown to be
essential for targeting of these enzymes to heterochromatin. The PWWP domain of
Dnmt3b has also been reported to interact non-specifically with DNA, suggesting a role
in interaction of the enzyme with the chromatin (Chen et al., 2004; Ge et al., 2004).
The cysteine-rich ADD domain found in DNMT3A/3B has been shown to interact
with histone deacetylase1 (Fuks et al., 2001), the transcriptional repressor RP58, the
silencer of heterochromatin HP1 and the histone methyltransferase SUV39h1 (Goll and
43
Bestor, 2005) but whether it plays a role in targeting of DNMT3A/3B to specific
nucleosomes remains an open question. Recently, the ADD domain of DNMT3L, a
regulatory factor which stimulates DNMT3A/3B activity in ES cells, has been found to
specifically interact with unmethylated H3K4 residues indicating that ADD domain can
actually mediate targeting of these enzymes (Ooi et al., 2007; Otani et al., 2009; Zhang et
al.). Thus, to further understand the mechanisms guiding DNMT3A/3B maintenance
activity in somatic cells, it is essential to study the interaction of these enzymes with the
chromatin in vivo and to determine the protein regions guiding such interactions.
In this study, we first investigated how different DNMT enzymes interact with
chromatin at the nucleosomal level in somatic cell lines. Micrococcal nuclease (MNase)
treatment of nuclei in a low ionic strength buffer digests nucleosomal linker DNA
regions, thereby minimizing the disruption of protein complexes on the nucleosomes. We
prepared nucleosomes from partial or limit MNase-digested nuclei and resolved them on
sucrose density gradients to analyze their interactions with chromatin proteins. This
method allows for the study of in vivo interactions between the chromatin modification
enzymes and their actual nucleosomal substrates in the native state. The results indicate
that while DNMT1 interacts primarily with linker DNA, DNMT3A/3B enzymes interact
strongly with nucleosomes. Such strong binding to nucleosomes requires synergistic
interactions of individual conserved domains of DNMT3A/3B with the chromatin.
Deletion of protein domains, as observed with cancer-specific truncated delta variants of
DNMT3B, diminishes binding of these enzymes to nucleosomes which might contribute
to the spurious methylation associated with these variants in cancer. Investigation of the
44
role of individual DNMT3A/3B domains revealed that the PWWP domain of DNMT3A
binds strongly to native nucleosomes through interaction with H3K36me3 mark, which
may assist in association of DNMT3A with the chromatin. Further, our data shows that
DNMT3A/3B bind specifically to nucleosomes containing methylated repetitive elements
and methylated CpG islands. Taken together, these data suggest that inheritance of DNA
methylation requires cues from chromatin component in addition to hemimethylation.
45
MATERIALS AND METHODS
Cell culture
HCT116, a human colon cancer cell line, and 293T cells, were maintained in
McCoy’s 5A and DMEM, respectively, containing 10 % inactivated fetal bovine serum,
100 units/ml penicillin, and 100 μg/ml streptomycin. Puromycin was included in the
culture medium at 3 μg/ml to maintain transfected 293T cells.
Expression vector construction
A modified version of the pIRESpuro3 vector (Clontech), pIRESpuro/Myc, was
constructed by ligating Myc tag DNA sequence into NheI-EcoR1 site of pIRESpuro3
vector. Human 3A1, 3B1, and 3B2 cDNA were kindly provided by A. Riggs (the City of
Hope). DNMT3A1, -3B1, and -3B2 cDNA were cloned into EcoR1-NotI site of
pIRESpuro/Myc vector and used for expressing N-terminal Myc-tagged DNMTs in
mammalian cells. Human ΔDNMT3B2 cDNA was amplified from pcDNA3/Myc-DNMT
3B1 using polymerase chain reaction (PCR), and ligated into EcoRI and BstX1 site of
pIRESpuro/Myc-DNMT3B2. To generate ΔDNMT3B4 cDNA, the corresponding N-
terminal sequence of ΔDNMT3B4 was amplified from HCT 116 cDNA, which expresses
ΔDNMT3B4, using RT-PCR and ligated into EcoRI and MscI site of IRESpuro/Myc
respectively. The constructs for expression of Myc-tagged truncated DNMT3B proteins
were prepared by Ms. Flora Han. Sequences coding for specific DNMT3B regions
[DNMT3B#7 (aa. 1-533); DNMT3B#9 (aa. 1-350); DNMT3B#5 (aa. 151-350);
DNMT3B#11 (aa. 1-216)] were amplified from Myc-DNMT3B1 construct and cloned
46
into EcoR1-NotI site of pIRESpuro/Myc vector All expression vector constructs were
transfected into 293T cells using lipofectamine 2000 (Invitrogen) and the cells stably
expressing DNMTs were selected in the presence of 3 μg/ml puromycin for three weeks.
Nuclei preparation
Nuclei were prepared according to the procedure described previously (Gal-Yam,
2006). Briefly, cells were trypsinized and washed once with PBS. The cells were then
resuspended in ice-cold RSB buffer (10 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM
MgCl
2
) containing protease inhibitors and kept on ice for 10 min before Dounce
homogenization in the presence of 0.5-1 % NP-40 to break up cell membranes. Nuclei
were washed twice with RSB plus the protease inhibitors (Roche) without the detergent.
Salt extraction of nuclei
Nuclei from 5x10
6
cells were resuspended in 500 μl of ice-cold RSB containing
0.25 M sucrose and protease inhibitors and various concentrations of NaCl, and kept at
4°C for 5 min. Nuclei were then harvested by microcentrifugation at 5000 rpm. The
nuclear pellet was dissolved in a SDS loading buffer and subjected to Western blotting.
MNase digestion and sucrose density gradient centrifugation
Purified nuclei (1x10
8
) resuspended in 1 ml of RSB containing 0.25 M sucrose, 3
mM CaCl
2
, and 100 μM PMSF, were digested with 5 units, for partial digestion, or 500
units, for the limit digestion, of MNase (Worthington) for 15 min at 37°C, and then the
47
reaction was stopped with EDTA/EGTA (up to 10 mM). After microcentrifugation at
5,000 rpm for 5 min, the nuclear pellet was resuspended in 0.3 ml of the buffer (10 mM
Tris-HCl, pH 7.4, 10 mM NaCl) containing 5 mM EDTA/EGTA, gently rocked for 1h at
4°C, and followed by microcentrifugation to obtain soluble nucleosomes, which were
then fractionated through a sucrose density gradient solution (5-25% sucrose, 10 mM
Tris-HCl, pH 7.4, 0.25 mM EDTA) containing NaCl of indicated concentrations, at
30,000 rpm for 16 h at 4°C. For the control experiment, purified nuclei (1x10
8
) were
incubated in 650 μl of RSB buffer containing 300mM NaCl for 5 minutes at 4°C.
Chromatin-free nuclear extract was prepared by centrifuging the incubated nuclei at
13,000 rpm for 10 min and collecting the supernatant. 550μl of the extract was then
loaded onto the sucrose gradients. Nucleosomes for a native ChIP assay were prepared in
the digestion buffer, 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 8 mM MgCl
2
, 3 mM
CaCl
2
, for 15 min at 37°C. The reaction was stopped with EDTA up to 10 mM, and left at
RT for 20 min, before collecting the soluble fraction of nucleosomes. Fractions were
taken from the top of the centrifuge tube to 16 aliquots.
Western blot analysis
Proteins from the same volume of each fraction (150-250 μl) were concentrated
by TCA precipitation, dissolved in SDS/β-mercaptoethanol loading buffer, and resolved
on a 4-15 % gradient SDS/PAGE gel (Bio-Rad, Hercules, CA). Antibodies against H3
(ab1791), histone H1 (ab7789), H3K9me3 (ab1773), H3K36me3 (ab9050), H4K20me3
(ab9053), DNMT3A (ab2850), HDAC1 (ab7028), and HP1alpha (ab9057) were
48
purchased from Abcam Inc.(Cambridge, UK); EZH2 (ac22) from Cell Signaling
Technology, Inc.(Danvers, MA); MeCP2 (07-013) from Upstate, Inc. (Charlottesville,
VA); DNMT1 (sc-20701), DNMT3B (sc-10235), and PCNA (sc-56), from Santa Cruz
Biotech. (Santa Cruz, CA); H3K27me3 (07–449), Myc epitope tag (05-724) from Upstate
(now Millipore, Billerica, MA). Image of individual proteins was visualized using ECL
detection system (Thermo Scientific, Waltham, MA and Millipore, Billerica, MA).
Construction and expression of PWWP and ADD (PHD-like) domains
PWWP and ADD domains of DNMT 3A, DNMT3B and DNMT3L [3A-PWWP
(aa 283-424), 3A-PHD-like (aa 532-592), 3B-PWWP (aa 216-350), 3B-PHD-like (aa
473-533), 3L-PHD-like aa 91-151] were cloned in EcoR1-BamHI site of pGEX-6P-1
(Amersham). The recombinant GST-fusion proteins were then expressed in E.coli and
bacterial lysates containing GST-fusion proteins were prepared. For checking proper
folding of the domain proteins, fusion proteins were purified using a glutathione resin
column. The recombinant peptides were eluted from the column using PreScission
protease. The eluted proteins were analyzed on a gel and further purified using a S75 gel
filtration column. I performed these experiments in Dr. Xiaojiang Chen’s lab.
Nucleosome binding assay
Native mononucleosomes isolated from HeLa cells were incubated with bacterial
lysates containing GST-PHD fusion proteins overnight at 4°C in binding buffer
containing 50 mM Tris-HCl pH 8.0, 100 mM NaCl, 0.1% NP-40, 10 μM ZnCl
2
, 1 mM
49
DTT and protease inhibitors. The samples were then incubated with Glutathione
Sepharose beads (GE Healthcare) at 4°C for 2 hrs. Beads were washed five times with the
binding buffer. The bound material was then used for DNA and protein extraction. Bound
DNA was resolved on a 1.5% agarose gel. The bound proteins were analyzed by
immunoblot analysis using antibodies specific for different histone marks.
Quantification of DNA methylation levels
Ms-SNuPE assay was performed as described previously (Yoo et al., 2007).
Genomic DNA was prepared from 293T cells transfected with myc-DNMT3A/B
expression vectors. Primers used for this assay are described in the manuscript (Jeong et
al., 2009).
Chromatin immunoprecipitation assay
ChIP assays were performed as described previously (Lin et al., 2007). The
following antibodies were used: 10 μg of anti-Myc antibody (05-724 from Upstate), anti-
DNMT3A (ab2850 from Abcam), and anti-CD8 (sc-32821 from Santa Cruz Biotech) as a
non-specific control antibody. Primers and probes used are described in the manuscript
(Jeong et al., 2009). For native ChIP, we used DNMT3A-specific (Ab2850) and
DNMT3B-specific (ab2851) antibodies. Nucleosomes were incubated with antibodies
over night at cold room in the MNase digestion buffer containing 10 mM EDTA, 0.2 mM
PMSF, and 0.1 % NP-40, and then incubated with protein-A agarose beads for 2 hr,
50
before washing the beads, two times for each, with 150 mM NaCl-, and 300 mM NaCl-
containing buffer containing 20 mM Tris-HCl, pH 7.4, 1 mM EDTA, and 0.1 % NP-40.
AP-PCR assay
Sixty nanograms of DNA of the input and unbound, and two microliters of
antibody-bound DNA were used for each AP-PCR reaction, to make sure that DNA
amounts of the input and unbound were much more than the bound DNA. Four random
primers (GC-rich) were added into each PCR reaction. Detailed procedure of
methylation-sensitive AP-PCR was previously described (Liang et al, 2002, Liang et al,
2004).
Southern blot
Southern blot analysis was performed as described previously (Weisenberger et
al., 2004). Equivalent amounts of DNA from each gradient fraction were ran on a 1%
agarose gel, and transferred to a nylon membrane (GeneScreen; Perkin Elmer Corp.,
Boston, MA). The radiolabeled probes were prepared using High Prime (Roche
Diagnostics, Germany) according to manufacturer’s protocol. For preparing genomic
DNA probe, genomic DNA from HCT116 cells was digested with EcoRI followed by
radiolabelling with High Prime. Consensus sequences of Alu and LINE probes used are
described in the related manuscripts (Jeong et al., 2009; Wolff et al., 2010).
51
RESULTS
DNMT enzymes associate with chromatin with different affinities
The DNA methyltransfreases, DNMT1, DNMT3A and DNMT3B, have been
previously shown to localize to distinct regions of chromatin, with DNMT1 showing a
diffused distribution throughout the nuclei while DNMT3A/3B showing a highly
punctuate distribution. (Bachman et al., 2001; Chen et al., 2002; Leonhardt et al., 1992).
To determine whether the difference in distribution patterns of DNMTs is due to a
difference in their chromatin binding affinities, we performed a salt extraction
experiment. This experiment was conducted by Dr. Shinwu Jeong in our lab.
Purified nuclei from HCT116 human colon cancer cells were incubated in buffers
with increasing concentrations of NaCl from 10mM to 400mM, in the presence of
protease inhibitors. Amount of various chromatin-associated proteins remaining inside
the nuclei following washes with increasing salt concentrations were then measured using
western blot analysis (Figure 2.1). As expected, under all salt concentrations the histone
octamer stayed intact inside the nuclei indicated by similar amounts of histone H3
remaining in the nuclei. DNMT1 levels within the nuclei were reduced to about half at
100 mM salt, and the majority of it disappeared at 200 mM salt indicating a weak
association of DNMT1 with chromatin. Strikingly, both DNMT3A/3B protein levels
remained almost constant within the nuclei up to 400 mM NaCl, displaying their strong
affinities for chromatin. The two bands for DNMT3B represent its two isoforms
expressed in HCT116 cells, DNMT3B2 being the upper band, and catalytically inactive
DNMT3B3, the lower (Aoki et al., 2001; Chen et al., 2005; Weisenberger et al., 2004).
52
Figure 2.1 Chromatin binding affinities of DNMTs and various other chromatin-
associated proteins. Nuclei purified from WT HCT116 cells were incubated in
nondenaturing extraction buffers containing 10 to 400 mM NaCl for 5 min. Equivalent
volumes of pellet fractions were subjected to western blot analysis using specific
antibodies. Ponceau S staining shows core histones transferred onto the membrane from
the SDS/PAGE gel. Experiment conducted by Dr. Shinwu Jeong.
53
We also tested various other chromatin-associating proteins which are known to interact
with DNMTs such as proliferating cell nuclear antigen (PCNA) (Chuang et al., 1997),
histone deacetylase 1 (HDAC1) (Fuks et al., 2000; Fuks et al., 2001; Robertson et al.,
2000), Methyl-CpG binding protein 2 (MeCP2) (Kimura and Shiota, 2003), Enhancer of
Zeste homolog 2 (EZH2) (Vire et al., 2006), and heterochromatin protein 1 alpha
(HP1alpha) (Fuks et al., 2003a; Lehnertz et al., 2003; Smallwood et al., 2007). However,
unlike DNMT3A/3B, all these proteins showed relatively weaker chromatin binding
affinities. The levels of PCNA and HDAC1, retained inside the nuclei decreased with
increasing salt concentrations, leaving some residual protein at the higher salt
concentrations. The amounts of MeCP2, EZH2 and HP1alpha showed constant levels of
the protein up to 200 mM salt, but decreased at higher salt concentrations. These data
suggest that DNMT3A/3B’s strong binding to chromatin is independent of these auxiliary
proteins (discussed in detail in Chapter 3).
DNMT3A/3B, but not DNMT1, strongly associate with nucleosomes
To further assess the differential interactions of DNMTs with chromatin, we
analyzed association of these enzymes with nucleosomes using sucrose density
centrifugation analysis which allows for the study of in vivo interactions between the
chromatin modification enzymes and their actual nucleosomal substrates in the native
state. This experiment was performed in collaboration with Dr. Shinwu Jeong.
Purified nuclei of HCT116 cells were subjected to partial digestion with a low
concentration of MNase, which cuts the linker DNA regions to generate nucleosomal
54
fragments of various sizes, under low salt conditions so that no artificial rearrangements
of nuclear proteins would occur. The digested nuclei were incubated in low ionic buffer
to collect diffused nucleosomes, which were loaded onto sucrose gradients and separated
by ultracentrifugation. Since DNMT3 enzymes remained associated with chromatin even
at 300 mM salt while other proteins started dissociating at such salt concentrations
(Figure 2.1), we used sucrose gradients containing 300 mM NaCl to determine the key
factors required for DNMTs binding to the nucleosomes (Figure 2.2). The relative
amounts and sizes of DNA in consecutive gradient fractions increased from fractions 6 to
16. Mononucleosomal DNA fragments were enriched largely in fractions 6 and 7,
dinucleosomes in fraction 9-10, and trinucleosomes, in 11-12, and so forth. The
distribution of histones H3 and H1 matched that of the nucleosomal DNA fragments and
the bulk histone proteins stained with Ponceau S solution indicating the intact structure of
nucleosomes sedimenting in each fraction.
DNMT1 sedimented in fractions 4 to 7 only, separate from the fractions
containing nucleosomes, showing that DNMT1 was not able to bind to polynucleosomes
at high salt concentrations. In contrast, the DNMT3A/3B enzymes remained associated
with the nucleosomal fractions. The majority of the DNMT3A/3B were detected in
polynucleosomal fractions 8-16 and not in the mononucleosomal fractions 6-7, consistent
with the relative abundance of polynucleosomes compared to mononucleosomes in a
partial digest (Figure 2.2). Overexposure of the immunoblot showed low amounts of
DNMT3A/3B sedimenting in fraction 6-7 also (data not shown). However, the patterns of
DNMT3A and DNMT3B distribution did not completely mirror each other indicating
55
Figure 2.2 DNMT3A/3B, but not DNMT1, associate strongly with polynucleosomes.
Purified nuclei from HCT116 cells were subjected to partial digestion with MNase.
Nucleosomes released from the nuclei were then resolved by ultracentrifugation on a
sucrose density gradient (5%-25%) containing 300 mM NaCl. Gradients were fractioned
into 16 aliquots numbered 1-16 starting from the top of the centrifuge tube. Absorbance
of each fraction was read at 260 nm. DNA purified from each fraction was resolved by
agarose gel electrophoresis and stained with EtBr. To probe the distribution of proteins in
each fraction, Western blotting was performed with various antibodies.
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 C
56
presence of some unique DNMT3A and DNMT3B binding regions. On the other hand,
EZH2 and HDAC1 showed main peaks in fractions 6-8, separate from the
polynucleosomes. These two proteins or complexes of them, unlike DNMT3A/3B, were
therefore not able to bind to polynucleosomes at high salt concentrations. These data
suggest that DNMT3A/3B, but not DNMT1, strongly associate with nucleosomes even at
a high salt concentration.
DNMT1 interacts with the linker DNA whereas DNMT3A/3B physically associate
with mononucleosomes
To further confirm that DNMT3A/3B’s cosedimentation with polynucleosomes is
mediated by their physical association with nucleosomes and also to determine whether
their strong interaction with chromatin requires polynucleosomal structures or whether
they could also bind to mononucleosomes, we next prepared mononucleosomes
consisting of approximately 150 bp of DNA wrapped around an octamer of histones by
more extensive MNase digestion of purified HCT116 nuclei and analyzed them on
sucrose gradients at 300 mM NaCl. Mononucleosomes containing 150 bp DNA
fragments and core histones, localized in a peak at fraction 6 (Figure 2.3A). DNMT1 was
mostly present in fraction 4, and UHRF1 formed a peak at fraction 3, both separated from
the nucleosome containing fractions. PCNA was also localized in nucleosome-free
fractions. Our data from 100 mM NaCl-containing gradients showed that DNMT1 could
interact with polynucleosomes on 100 mM NaCl-containing gradients, however, it
57
Figure 2.3 DNMT3A/3B physically associate with mononucleosomes. (A) Purified
nuclei from HCT116 cells were subjected to limit digestion with MNase at low ionic
strength. Nucleosomes released from the nuclei were then resolved by ultracentrifugation
on a sucrose density gradient (5%-25%) containing 300 mM NaCl. Gradients were
fractioned into 16 aliquots numbered 1-16 starting from the top of the centrifuge tube.
Absorbance of each fraction was read at 260 nm. To probe the distribution of proteins in
each fraction, Western blotting was performed with various antibodies. (B) Native
proteins released from nuclei at high ionic strength (300mM) were resolved by
ultracentrifugation on a sucrose density gradient (5%-25%) containing 300 mM NaCl.
Gradients were fractioned into 16 aliquots numbered 1-16 starting from the top of the
centrifuge tube. To probe the distribution of proteins in each fraction in a chromatin free
context, western blotting was performed with various antibodies. (C) Sucrose gradient for
mononucleosomal digest from DKO8 cells.
Core Histones
DNMT3A
A
B
C
58
dissociated from mononucleosomes even at low ionic strength, suggesting that DNMT1
may weakly interact with linker DNA (data not shown).
EZH2 and HDAC1 showed similar sedimentation profiles on the 300 mM NaCl-
containing gradients with mononucleosomes as previously observed on gradients with
polynucleosomes (compare Figure 2.2 to 2.3A), strongly suggesting that these proteins
were not bound to nucleosomes at a high salt concentration. In contrast, both
DNMT3A/3B localized to peaks in fraction 7, indicating that DNMT3A/3B must be
bound to mononuclesomes rather than the linker DNA, and that their presence altered the
sedimentation of bound nucleosomes by one fraction relative to bulk mononucleosomes.
Also, the sedimentation profiles of DNMT3A/3B show a marked change when the extent
of MNase digestion was altered from partial to limit (compare Figure 2.2 to 2.3A).
MNase is a DNase which specifically targets linker DNA for digestion and has no effect
on proteins, so that it would not be expected to alter the distribution of DNMT3A/3B
unless the enzymes were physically associated with chromatin.
To confirm that EZH2 and HDAC1 which cosedimented with nucleosomes in
fractions 6-8 at 300 mM NaCl were not physically bound to chromatin, we prepared
nuclear extracts devoid of chromatin and containing only native proteins and protein
complexes eluting out from nuclei at high salt concentrations, and analyzed them on
sucrose gradients containing 300 mM NaCl (Figure 2.3B). Even in a chromatin free
environment, EZH2 and HDAC1 still sedimented in the same fractions showing main
peaks in fraction 6-8 similar to their sedimentation profiles in the presence of
nucleosomes. This control experiment thus proves that EZH2 and HDAC1 do not bind to
59
nucleosomes at high salt concentrations even though they sediment in the same fractions
possibly as a part of large multiprotein complexes. Likewise, DNMT1 and PCNA
showed similar sedimentation profiles to those in the presence of polynucleosomes at 300
mM NaCl. DNMT3A could not be detected in any of the fractions in the chromatin-free
gradient. These data indicate that DNMT3A/3B are physically bound to nucleosomes and
these non-histone proteins are not involved in the stable complex between DNMT3A/3B
and nucleosomes.
Since DNMT3A and DNMT3B can form heterocomplexes (Kim et al., 2002), we
also tested whether the nucleosomal binding of one enzyme requires the presence of the
other. We performed the same experiment using a DNMT3B-deficient HCT116 cell line,
DKO8 (DNMT1
ΔE2-5
, DNMT3B
−/−
) (Rhee et al., 2002), and found that DNMT3A
cosedimented with mononucleosomes in the absence of DNMT3B, indicating that the
interaction between the DNMT3A and 3B are not required for DNMT3A’s association
with nucleosomes (Figure 2.3C).
The cancer-specific ΔDNMT3B variants weakly associate with nucleosomes
Various isoforms of DNMT3A/3B are expressed in mammalian cells (Goll and
Bestor, 2005). Recently a family of DNMT3B isoforms, the delta variant ΔDNMT3B
isoforms, has been associated with cancer-specific aberrant methylation (Wang et al.,
2006a; Wang et al., 2006b). We next wanted to investigate whether such aberrant
methylation patterns resulted from impaired association of these isoforms with the
chromatin. To examine the interaction pattern of various DNMT3B isoforms with
60
nucleosomes, we expressed various Myc-tagged fusion proteins of DNMT3A/3B in
human 293T cells (Figure 2.4A) and tested their distribution in mononucleosomal digests
on sucrose gradients at a high salt concentration (Figure 2.4B). The endogenous DNMT1
and DNMT3A/3B showed distribution patterns similar to those of HCT116 cells
indicating that the strong nucleosomal association of DNMT3A/3B takes place in both
cell types and is not due to potential cell type specific interactions. Full length Myc-
tagged DNMT3A/3B also showed a strong association with mononucleosomes. However,
neither ΔDNMT3B2, which lacks the initial 199 aa of the N-terminal region (N-terminal
domain) nor ΔDNMT3B4 (which lacks the N-terminal domain and a part of the PWWP
domain, see Figure 2.4A) was strongly anchored to mononucleosomes. Even at 100 mM
NaCl concentration, majority of ΔDNMT3B2 and ΔDNMT3B4 proteins dissociated from
nucleosomes (Figure 2.4C). These data indicate that the N-terminal region of DNMT3B
plays an essential role in its strong nucleosomal binding.
DNMT3B requires intact protein structure for anchoring to nucleosomes
To further dissect the regions of DNMT3B protein which play a role in its strong
anchoring to nucleosomes, we expressed Myc-tagged truncated DNMT3B proteins in
human 293T cells (Figure 2.5A) and first tested their distribution in mononucleosomal
digests in 300 mM NaCl sucrose gradients. Unlike full length Myc-tagged DNMT3B
protein which showed strong association with the mononucleosomes (Figure 2.5B), all of
the truncated proteins showed very weak affinity for binding to the mononucleosomes
and sedimented in the nucleosome-free fractions 1-4. To ascertain if the truncated
61
Figure 2.4 Cancer-specific truncated delta DNMT3B variants associate weakly with
nucleosomes. (A) Map of DNMT3A/3B isoforms showing the PWWP and PHD-like
domains located in the N-terminal regions, and the catalytic methylase domains in the C-
terminal region. (B) Mononucleosomes from 293T cells transfected with expression
vectors of various Myc-DNMT3A/3B deletion proteins were subject to sucrose gradients
containing 300 mM NaCl. Endogenous and exogenous enzymes on the gradient were
detected by Western blot analysis using specific antibodies against endogenous protein
and anti-Myc Ab. (C) Mononucleosomes from 293T cells transfected with expression
vectors of Myc-∆DNMT3B2 or Myc-∆DNMT3B4 deletion proteins were subject to
sucrose gradients containing 100 mM NaCl. Exogenous enzymes on the gradient were
detected by western blotting using anti-Myc Ab.
A
B
C
62
Figure 2.5 Truncated DNMT3B proteins display weak binding to both mono- and
poly-nucleosomes. (A) Map of various DNMT3B truncated constructs prepared for
transfection in 293T cells. (B) Mononucleosomes from 293T cells transfected with
expression vectors of various Myc-DNMT3B deletion proteins were subject to sucrose
gradients containing 300 mM NaCl. (C) Polynucleosomes from 293T cells transfected
with expression vectors of various Myc-DNMT3B deletion proteins were subject to
sucrose gradients containing 300 mM NaCl. Exogenous enzymes on the gradient were
detected by Western blot analysis using anti-Myc Ab.
DNMT3B #7
DNMT3B #5
DNMT3B #9
DNMT3B #11
DNMT3B1
A
B
C
63
DNMT3B proteins may anchor to nucleosomes in the presence of linker DNA, we
prepared polynucleosomes by subjecting the transfected nuclei to partial MNase digestion
and then resolved them on a 300 mM NaCl-containing sucrose gradient. Even with
polynucleosomes, the truncated DNMT3B proteins showed very weak binding and the
majority of the protein sedimented in the nucleosome-free fractions (Figure 2.5C).
However, some truncated protein showed very weak interaction with polynucleosomes,
possibly though non-specific interactions with the linker DNA. These experiments were
further performed using sucrose gradients with 150 mM salt concentration to test if the
truncated proteins may possibly be able to bind to nucleosomes at a lower physiological
salt concentration. Even at 150 mM salt concentration, the truncated DNMT3B proteins
did not show any binding to nucleosomes (data not shown). These data suggest that an
intact structure of DNMT3B protein is required for strong anchoring to nucleosomes
where different domains may work synergistically to enable tight binding of the protein
to the nucleosomes.
DNMT3A PWWP binds strongly to native nucleosomes through interaction with
H3K36me3 mark
To examine the role of the conserved domains (PWWP and ADD domains) found
in the N-terminal regulatory region of DNMT3A/3B in the strong anchoring of
DNMT3A/3B enzymes to nucleosomes, GST-tagged PWWP and ADD domains of
DNMT3A/3B and DNMT3L were expressed in E.coli (Figure 2.6A). A nucleosome
binding assay was performed using native nucleosomes extracted from HeLa cells and
64
Figure 2.6 DNMT3A-PWWP and DNMT3B-PWWP bind to native nucleosomes. (A)
Ponceau stain showing protein expression of various GST-tagged DNMT3A/3B and
DNMT3L domains expressed in E.coli. (B) Agarose gel showing genomic DNA obtained
after digestion of HeLa nuclei with varying concentrations of MNase. (C) Nucleosome
Binding Assay: Bacterial lysates containing individual GST-tagged DNMT3 domains or
GST were incubated with native HeLa nucleosomes (containing primarily
mononucleosomes) followed by pull down using GST resin. Bound material was
analyzed by immunoblotting with H3 antibody.
A
B
C
65
the recombinant GST-tagged DNMT3 PWWP and ADD domains. Native
mononucleosomes, prepared through MNase digestion of purified HeLa nuclei (Figure
2.6B), were incubated with GST-tagged DNMT3 domains and their interactions were
assessed by GST pull-down experiments. Analysis of bound material by immunoblotting
with H3 antibody revealed that both DNMT3A-PWWP and DNMT3B-PWWP could
bind to native nucleosomes (Figure 2.6C). The ADD domains however showed very
weak binding to the native nucleosomes in our assay which can be due to improper
folding of the small ADD domain proteins.
We next examined the strength of nucleosome binding of DNMT3A-PWWP and
DNMT3B-PWWP domains through a salt extraction assay. The binding affinity of the
PWWP domains decreased significantly as the salt concentration of the binding buffer
was increased from 100 to 300 mM (Figure 2.7A). Interestingly, the affinity of
DNMT3A-PWWP for binding to native nucleosomes was found to be much higher than
that of DNMT3B-PWWP. This finding is opposite to the previous findings regarding
DNA binding properties of these domains which showed that Dnmt3B-PWWP binds
stronger to naked DNA than Dnmt3A-PWWP (Qiu et al., 2002). This led us to
investigate whether DNMT3A-PWWP could bind to histones, the other key component
of a nucleosome apart from DNA. We separated the bound proteins pulled down by the
GST-DNMT3A-PWWP domain in the native nucleosome binding assay and
immunoblotted them with antibodies for various histone marks. We could observe
enrichment of the H3K36me3 mark in comparison to H3K9me3, H3K27me3 and
H4K20me3, even though the preferential binding appeared to be weak (Figure 2.7B).
66
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
H3-K36me3 H3-K9me3 H3-K27me3 H4-K20me3
Pull Down / Input ( Mark/H3)
Figure 2.7 DNMT3A-PWWP binds more strongly to nucleosomes than DNMT3B-
PWWP, through association with H3K36me3 mark. (A) Nucleosome Binding Assay
was performed under different salt conditions (100, 150 and 300 mM) to determine the
strength of binding. Bound material was analyzed by immunoblotting with H3 antibody.
(B) Signals for individual marks were normalized with H3 signal and the enrichment in
pull down sample was estimated in comparison to the input. Data presented in this figure
is representative of two biological replicate experiments.
A
B
67
Recently, similar findings have been reported by Dhayalan et al. (Dhayalan et al., 2010),
who have also shown specific binding of Dnmt3a-PWWP to the H3K36me3 mark using
peptide arrays and nucleosome binding assays. In agreement with our data, they have also
found that H3K36me3 mediated binding to nucleosomes was specific for Dnmt3a-
PWWP, with Dnmt3b-PWWP showing minimal peptide binding. Taken together, these
data suggest a guiding mechanism where H3K36me3 assists recruitment of DNMT3A to
nucleosomes through its PWWP domain, in agreement with previous studies indicating
an essential role of the PWWP domain in localization of Dnmt3a in the nuclei (Qiu et al.,
2002).
DNMT3A/3B are enriched in methylated CpG islands and repetitive DNA elements
To ascertain the specificity of DNMT3A/3B enzymes for distinct chromatin
regions, we examined the genomic regions bound by these enzymes. Previous studies
have shown that DNMT3A/3B are enriched on methylated CpG islands (CGIs)
(Schlesinger et al., 2007) and the maintenance of methylated state of such CGIs is
relatively insensitive to DNMT1 depletion which suggests a role for DNMT3A/3B in
maintaining CGI methylation (Egger et al., 2006). Based on these observations we
hypothesized that DNMT3A/3B might preferentially bind to nucleosomes associated with
methylated CGIs. To test this hypothesis, we used the 293T cells transfected with myc-
DNMT3A/3B proteins. We first determined the DNA methylation levels in various CGI
regions in those cells and then selected six highly methylated regions, including D4Z4
repeats, and three unmethylated regions for our analysis (Figure 2.8A). We next
68
Figure 2.8 DNMT3A/3B preferentially bind to methylated repeats and CpG islands.
(A) The levels of DNA methylation in nine different CpG islands in 293T cells
transfected with myc-DNMT3A1 or DNMT3B1 constructs were determined by Ms-
SNuPE. (B) ChIP assays were performed with anti-DNMT3A antibodies for the cells
with myc-DNMT3A1 or anti-Myc antibody for the cells with myc-DNMT3B1. Values
are the averages of at least triplicate determinations with standard errors as indicated.
These experiments were performed by Dr. Joy Lin and Dr. Gangning Liang.
69
performed ChIP experiments with anti-myc antibodies and found that DNMT3A was
enriched at the methylated ABCD1, MLH3, and IRF5 regions, however, it was less
enriched in the other three methylated CDKN2A, EPM2AIP1, and D4Z4 regions. In
contrast, DNMT3B was enriched in all of the six highly methylated regions (Ehrlich,
2003; Weisenberger et al., 2004). Both DNMT3A and DNMT3B did not show
enrichment in the three unmethylated regions AHR, HOXB13, MLLT3 (Figure 2.8B).
These results indicate that DNMT3A/3B preferentially bind to methylated CGIs, with
DNMT3A occupying a subset of methylated CGIs shared by DNMT3B.
To gain a more global perspective, we next performed a native ChIP assay
followed by direct sequencing of the immunoprecipitated DNA. Nucleosomes isolated
from MNase-digested nuclei were immunoprecipitated using DNMT3A/3B-specific
antibodies and the DNA purified used for whole genome amplification. Sequence
analysis of 24 DNA clones obtained from this amplified DNA sample showed that 83%
of the clones contained SINE and LINE repeats of which 4% contained CGIs (Figure
2.8C). The human genome comprises of approximately 35% repetitive DNA and 65%
single copy DNA sequences (Smit and Riggs, 1995; Venter et al., 2001). Thus, the
proportion of repeat sequences found in the amplified DNA sample is more than 2-fold
higher than expected. This suggests that DNMT3A/3B enzymes preferentially associate
with repetitive DNA elements.
To confirm that DNMT3A/3B may also specifically bind to CGI regions, we next
characterized DNA fragments enriched in DNMT3A/3B-bound fraction using arbitrary
primed PCR (AP-PCR), which preferentially amplifies GC-rich regions by using GC rich
70
random primers (Liang et al., 2004). Sequence analysis of 36 DNA fragments obtained
from the AP-PCR amplified DNA sample showed that 71% of the sequenced fragments
were repeats and 36% were CGIs (Figure 2.8C). Fourteen fragments were selected out of
these 36 fragments, and all of these confirmed to be methylated at internal HpaII (CCGG)
sites using methyl sensitive AP-PCR (Liang et al., 2002) (data not shown). These data
together indicate that DNMT3A/3B preferentially binds to methylated repeat sequences
and CGIs, which is consistent with their essential role in maintaining DNA methylation at
repeat elements (Liang et al., 2002; Rhee et al., 2002)
DNMT3A/3B associate with di- and tetra-nucleosomal Alu and LINE structures
respectively
To further explore DNMT3A/3B’s association with the repeat elements, we next
examined distribution of the Alu and LINE repeat sequences, which associate with
DNMT3A/3B, on sucrose gradients. Purified nuclei from HCT116 cells were subjected to
partial digestion using MNase and the chromatin was then fractionated on a 300mM
sucrose gradient. Proteins enriched in each fraction were analysed using Western blot
analysis as described previously (Figure 2.9A). DNA extracted from equivalent volume
of each fraction was then run on an agarose gel. We first performed a genomic Southern
using radioactively labeled input DNA which showed the majority of DNA fragments
being present as mononucleosomes and dinucleosomes (Figure 2.9B). Interestingly,
probing the blot with the Alu repeat sequence, found enriched in DNMT3A/3B
71
Figure 2.9 Association of DNMT3A/3B with di- and tetra-nucleosomal structures
present at Alu and LINE1 sequences respectively. (A) Nucleosomes obtained from
partial MNase digestion of HCT116 nuclei were resolved on a sucrose density gradient
containing 300mM NaCl. Proteins enriched in each fraction were analyzed by
immunoblotting. (B) Southern experiments were performed for genomic DNA, Alu and
LINE1 to examine their enrichment in the DNA in each fraction. Genomic DNA was
enriched in the mono- and dinucleosome fractions. However Alu showed preferential
enrichment in di-nucleosomes. LINE1 elements showed enrichment in the di- and
tetranucleosome fractions. According to our model, the L1 promoters with a
tetranucleosomal structure should be inactive and methylated.
72
immunoprecipitates (conducted by Dr. Shinwu Jeong), showed preferential enrichment in
dinucleosomal fragments.
Since LINE repeat sequences were also found enriched in DNMT3A/3B
immunoprecipitation experiment, we next probed the blot with a LINE sequence, L1
(LINE1) promoter sequence. L1 promoter sequence showed enrichment in both the
dinucleosome and tetranucleosome fractions (Figure 2.9B). Later, a more detail study of
one such individual LINE1 element present within the MET gene, MET-LINE1, done by
Dr. Erika Wolff in our lab revealed that an active LINE1 element undergoes a switch
from a di- to tetra-nucleosomal state when it is silenced through DNA methylation (Wolff
et al., 2010). Furthermore, this switch is reversed when the cells are depleted of DNMTs
and undergo severe hypomethylation as observed in DNMT-deficient HCT116 DKO1
(DNMT1
ΔE2-5
, DNMT3B
−/−
) cells, indicating that DNA methylation and possibly binding
of DNMT3A/3B is essential for maintaining the LINE1 repeats in an inactive tetra-
nucleosomal state (Wolff et al., 2010). Taken together, these data suggest that
DNMT3A/3B associate with unique di- and tetra-nucleosomal chromatin structures found
at repeat elements, which may enable them to faithfully maintain their methylated states.
73
DISCUSSION
Interaction of DNMT enzymes with their substrate DNA in a nucleosomal context
is essential for proper maintenance of DNA methylation patterns. In this study, we
examined the association of different DNMT enzymes with the nucleosomes through
fractionation of MNase digested chromatin upon sucrose density gradients followed by
analysis of chromatin-associated protein using Western blotting. Our data show that
DNMT1 associates weakly with chromatin, mostly at linker or nucleosome-free DNA
regions possibly assisted by factors such as PCNA and/or UHRF1 (Bostick et al., 2007;
Chuang et al., 1997; Ooi and Bestor, 2008). The “loose” association of DNMT1 with
chromatin observed in our experiments is consistent with a highly dynamic, transient
nature of the interaction of Dnmt1 with the replication machinery (Schermelleh et al.,
2007; Schermelleh et al., 2005) and its continuous loading onto constitutive
heterochromatin during G2/M phase (Easwaran et al., 2004). In addition to its
replication-coupled maintenance activity, DNMT1 may further be able to methylate
hemimethylated sites on the linker DNA missed during the replication process in
association with UHRF1 (Arita et al., 2008; Avvakumov et al., 2008; Hashimoto et al.,
2008; Sharif et al., 2007).
In contrast to the dynamic nature of the interaction of DNMT1 with the
chromatin, DNMT3A/3B enzymes exhibit strong association with nucleosomes
suggesting a highly compartmentalized activity of these enzymes mediated by their tight
association with target chromatin regions. This data is consistent with the highly
localized distribution of DNMT3A/3B enzymes observed within nuclei (Bachman et al.,
74
2001; Chen et al., 2002). Since the sedimentation properties of DNMT3A/3B were
strongly influenced by the extent of MNase digestion, which as a DNase, would not
directly alter the sedimentation behavior of a protein, our data suggests a strong physical
interaction between DNMT3A/3B and nucleosomes. Our data further shows that
DNMT3B requires an intact protein structure for stable binding to nucleosomes, since all
of the ΔDNMT3B variants and other truncated versions of DNMT3B protein examined
easily dissociated from mononucleosomes. Since these variants possess active de novo
methylation activity, this lack of tethering might play a role in the generation of spurious
methylation patterns such as those found in lung cancer cells which express high levels of
ΔDNMT3B variants (Wang et al., 2006a; Wang et al., 2006b).
Individual domains of DNMT3A/3B proteins have previously been suggested to
target these enzymes to silent chromatin regions. The PWWP domain of DNMT3A/3B
has been shown to be required for targeting to heterochromatin (Chen et al., 2004; Ge et
al., 2004). The DNMT3B-PWWP domain has also been shown to interact with DNA
non-specifically which may stabilize interaction of the protein with the nucleosomes (Qiu
et al., 2002). Our data shows that unlike DNMT3B-PWWP, DNMT3A-PWWP interacts
more strongly with native nucleosomes through its association with the H3K36me3 mark
suggesting a role of H3K36me3 in guiding DNMT3A activity to specific chromatin
domains. Similar distribution of DNA methylation and H3K36me3 observed in genome-
wide studies supports presence of such a mechanism (Meissner et al., 2008). Recently,
ADD domains of DNMT3A and DNMT3L have also been implicated in associating these
enzymes with the chromatin through its interaction with unmethylated H3K4 residues
75
(Ooi et al., 2007; Zhang et al., 2010). However, it is interesting to note that while these
domains can interact individually with native nucleosomes and have been proposed to
target DNMT3 enzymes, they are alone insufficient to anchor DNMT3 enzymes to the
nucleosomes. Our data shows that an intact DNMT3B protein structure is necessary for
its strong binding to nucleosomes suggesting a requirement of synergistic activity of
different domains and structural motifs for achieving DNMT3A/3B’s strong association
with the nucleosomes.
We have further shown that DNMT3A/3B are bound to regions containing
methylated repeats and CpG islands. Also, DNMT3A/3B possibly recognize unique di-
and tetra-nucleosomal structures present at Alu and LINE1 repeat sequences respectively.
Alu and LINE repeats are kept methylated in somatic cells preferentially by the activities
of DNMT3A/3B enzymes. Alu sequences are known to influence nucleosome positioning
in chromatin and to exist predominantly as dinucleosomes in nuclei (Englander and
Howard, 1995; Kato et al., 2003; Salih et al., 2008; Tanaka et al.). It will be interesting to
see whether DNA methylation of Alu sequences observed in mammalian genomes is
required for formation of such dinucleosomal structures. Recent observation of increased
nucleosome occupancy and rigidity upon DNA methylation supports such a possibility
(Choy et al. 2010). In addition, formation of inactive tetra-nucleosomal structures at
LINE1 elements has also been shown to accompany DNA methylation by our lab,
suggesting a role of methylation in stabilization of such nucleosomal structures (Wolff et
al., 2010). How such nucleosomal structures may be recognized by DNMT3A/3B is still
76
an open question. It may involve oligomerization of DNMT3A/3B proteins similar to that
recently observed with DNMT3A-DNMT3L complex (Jia et al., 2007).
The preferential association of DNMT3A/3B with nucleosomes containing
methylated repeats and CpG islands is interesting since it may provide some insights into
the manner in which these de novo enzymes may assist DNMT1 in propagation of
methylation patterns. The original model proposed for inheritance of DNA methylation
suggests that patterns are established de novo during embryogenesis and then faithfully
copied by a “maintenance” enzyme (Holliday and Pugh, 1975; Riggs, 1975). DNMT1
clearly can serve such a maintenance function through its replication-coupled activity
(Bestor and Ingram, 1983; Bostick et al., 2007; Chuang et al., 1997; Leonhardt et al.,
1992). However, the maintenance of global DNA methylation seems to require
DNMT3A and/or 3B in cooperation with DNMT1 since knock out of these enzymes
leads to a gradual loss of the majority of CpG methylation (Chen et al., 2003; Liang et al.,
2002; Rhee et al., 2002). Our data may help explain how CpG rich regions, such as
repeats and CGIs are kept methylated, since they suggest that DNMT3A/3B are highly
localized to a subset of nucleosomes containing methylated CpG sites. Based on our
current findings, we propose that DNMT3A/3B associated with nucleosomes containing
methylated DNA may have roles in maintaining methylated states of such regions (Figure
2.10). Since DNMT1 is a processive enzyme, it may not be able to faithfully copy
methylation patterns during the rapid replication process especially in regions containing
a large number of CpG sites such as at CpG islands and some repeats (Schermelleh et al.,
2007), leaving behind some unmethylated CpG sites. DNMT3A/3B, which remain
77
Figure 2.10 Proposed model for inheritance of DNA methylation patterns by
DNMT1 and DNMT3A/3B. The de novo DNMT3A and DNMT3B enzymes
(represented by DNMT3, in orange) remain bound to chromatin in somatic cells,
preferentially binding to nucleosomes that contain methylated DNA. During replication,
DNMT1 (in purple), which is the predominant DNA methylase in the somatic cells, reads
the methylation patterns on the template strand and copies it to the daughter strand
through its association with the replication fork by proliferating cell nuclear antigen
(PCNA, in blue) and possibly by ubiquitin-like plant homeodomain and ring finger
domain containing protein 1 (UHRF1, in yellow). We propose that soon after DNA
replication, DNMT3A and DNMT3B complete the methylation process and correct errors
that are left by the DNMT1 enzyme. Because these enzymes are compartmentalized to
the chromatin region containing methylated DNA, they do not ‘read’ the parental strand
for DNA methylation patterns but rather methylate newly replicated cpG sites that are
unmethylated. In cancer, the cancer-specific ∆DNMT3B isoforms (in pink), which
weakly associate with the nucleosomes, may cause spurious de novo methylation due to
their aberrant targeting. Such aberrant methylation may later get inherited through
maintenance activity of DNMT1 and DNMT3A/3B.
78
associated with such regions, may then methylate the remaining unmethylated CpG sites
in those regions, resulting in faithful inheritance of their methylated states. On the other
hand, ΔDNMT3B variants may be aberrantly targeted, contributing to abnormal
methylation patterns observed in tumorigenesis (Wang et al., 2006a; Wang et al., 2006b).
79
CHAPTER 3
ROLE OF AUXILIARY PROTEINS IN
ANCHORING OF DNMT3A/3B TO NUCLEOSOMES
INTRODUCTION
DNMT3A/3B, but not DNMT1, show a strong association with the nucleosomes,
suggesting a difference in the maintenance mechanisms guiding the activities of these
enzymes in somatic cells (Jeong et al., 2009). Unlike the transient interactions of DNMT1
with chromatin, DNMT3A/3B remain preferentially bound to chromatin regions
containing methylated repeats and CpG islands. In addition to the direct interactions of
DNMT3A/3B domains with nucleosomes (as described in Chapter 2), targeting of these
de novo enzymes to specific chromatin regions in vivo may also involve interaction with
auxiliary factors (Hermann et al., 2004; Klose and Bird, 2006). Recently Dnmt3L has
been found to connect Dnmt3a2 to nucleosomes in ES cells (Ooi et al., 2007). However,
Dnmt3L is expressed only during gametogenesis and embryonic stages (Aapola et al.,
2000; Bourc'his et al., 2001), suggesting that other mechanisms might be necessary for
directing DNMT3A/3B enzymes to specific chromatin domains in somatic cells.
Previous studies have revealed interaction of DNMT3A/3B proteins with various
chromatin-associated proteins including hetrochromatin protein 1 (HP1), histone
deacetylase 1 (HDAC1), UHRF1 and histone methyltransferases such as EZH2 &
SUV39h1, suggested to play a role in recruitment of DNMT3A/3B to specific chromatin
80
regions for de novo methylation in embryonic stem (ES) cells (Klose and Bird, 2006;
Law and Jacobsen, 2010). Recently, G9a, another H3K9 methyltransferase, has been
suggested to recruit DNMT3A/3B to euchromatic H3K9 methylated regions for de novo
methylation in ES cells (Tachibana et al., 2008). While the roles of DNMT3A/3B’s
interactions with chromatin-associated auxiliary proteins in de novo methylation in ES
cells has been well studied, their roles in targeting of DNMT3A/3B enzymes to specific
chromatin regions for maintenance of methylation patterns in somatic cells remains
unclear.
G9a, along with its partner protein GLP, is crucial for H3K9 (mainly H3K9me
and H3K9me2) methylation of euchromatin and is involved in transcriptional silencing
(Tachibana et al., 2002; Tachibana et al., 2005). G9a also binds to its own product,
H3K9me and H3K9me2 residues, through its ankyrin domain, a mechanism suggested to
play a role in propagation of H3K9 methylation through cell divisions (Collins and
Cheng, 2010; Collins et al., 2008). G9a physically interacts with DNMT3A/3B and
promotes de novo methylation in ES cells through their recruitment to G9a-target genes
(Epsztejn-Litman et al., 2008). SUV39h1 is responsible for heterochromatic H3K9
methylation and assists in establishment of heterochromatin in association with HP1
(Peters et al., 2001). HP1 binds to H3K9me3 residues and is thought to work downstream
of SUV39 HMTases to stabilize heterochromatic domains (Bannister et al., 2001;
Nakayama et al., 2001). HP1 also interacts with SUV39h1 and targets it to H3K9me3
regions, which is thought to be critical for inheritance of H3K9 methylation in
heterochromatic regions. SUV39h1 has been proposed to direct DNA methylation to
81
major satellite repeats via recruitment of DNMT3A/3B by HP1 (Felsenfeld and
Groudine, 2003; Lehnertz et al., 2003; Peters et al., 2001). Whether these proteins play a
similar role in associating DNMT3A/3B with methylated chromatin regions in somatic
cells is unknown.
Here we show that binding of DNMT3A/3B to nucleosomes in somatic cells does
not require auxiliary proteins such as HP1, MeCP2, EZH2, HDAC1, UHRF1 and G9a,
but does require an intact nucleosomal structure. We used sucrose density gradients of
nucleosomes prepared by partial and limit micrococcal nuclease digestion, coupled with
Western Blot analysis to probe for the interactions of DNMTs and other chromatin
associated proteins with the native nucleosomes. While DNMT3A/3B remained strongly
anchored to nucleosomes at high salt concentrations, auxiliary proteins such as HP1,
MeCP2, EZH2, HDAC1 and UHRF1 dissociated from the nucleosomes. The H3K9
methyltransferases, G9a and SUV39h1, displayed strong binding to both
mononucleosomes and polynucleosomes, similar to that observed for DNMT3A/3B.
However, knockdown of G9a did not inhibit binding of DNMT3A/3B to nucleosomes
and DNA methylation levels of several G9a-associated genes remained unaltered in G9a
knockdown cells, suggesting no role of G9a in maintenance of DNA methylation through
DNMT3A/3B. In addition, association of G9a with nucleosomes was also found to be
independent of DNA methylation.
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MATERIALS AND METHODS
Cell culture
HCT116 and 293T cells, were maintained in McCoy’s 5A and DMEM,
respectively, containing 10 % inactivated fetal bovine serum, 100 units/ml penicillin, and
100 μg/ml streptomycin. Puromycin was included in the culture medium at 3 μg/ml to
maintain transfected cells.
MNase digestion and sucrose density gradient centrifugation
Sucrose density gradient experiments were performed as described previously in
Chapter 2. The EtBr treatment to the mononucleosome samples was done by adding 20
mg/ml EtBr to the samples (300 μg/ml at final), followed by incubation at room
temperature for 10 min before loading onto the gradient.
Lentiviral knockdown
The lentivirus particles containing N/S, shG9a5 and shG9a7 shRNA sequences
were prepared by Mr. Daniel Gerke from Dr. Michael Stallcup’s lab. For lentivirus
production, the vesicular stomatitis virus envelope protein G expression construct
pMD.G1, the packaging vector pCMV ΔR8.91 and the transfer vector
pHRCMVpuroSin8 were used as described previously (Ou et al., 2009). Short hairpin
RNA sequences encoding non-specific and sequences targeting G9a were as follows: N/S
5’- AAAACTGCAGAAAAAGGGTAGGTTCGACTAGCAGGACTCTTCTCTTGAA
83
AGAGTCTTGCTAGTTGAACCTACCCGGTGTTTCGTCCTTTCCACAAG-3’;
shG9a5 5’- AAAACTGCAGAAAAAGACAGCAAGTCTGAAGTTGAAGCTCTCTCT
TGAAGAGCTTTAACTTCATACTTGCTGTCGGTGTTTCGTCCTTTCCACAAG -3’
and shG9a7 5’-AAAACTGCAGAAAAAGGATGAATCTGAGAATCTTGAGGG
ATCTCTTGAATCCCTCCAGATTCTTAGATTCATCCGGTGTTTCGTCCTTTCCA
CAAG -3’. Infected HCT116 cells were selected in the presence of 3 μg/ml puromycin
for two weeks.
Western blot analysis
Western blot analysis was performed as described previously in Chapter-2.
Antibodies used against SUV39h1 (ab12405) were purchased from Abcam
Inc.(Cambridge, UK); G9a (G 6919) from Sigma (Saint Louis, MO); anti-UHRF1
antibody (#612264) from BD Biosciences.
Quantification of DNA methylation levels
Ms-SNuPE assay was performed as described previously (Yoo et al., 2007).
Genomic DNA was prepared from HCT116 cells infected with either N/S, shG9a5 or
shG9a7 constructs 14 days after lentiviral infection.
84
RESULTS
Binding pattern of different DNMT3A/3B associated proteins to nucleosomes
To test the role of auxiliary proteins in the strong nucleosome anchoring
manifested by DNMT3A/3B, we examined their nucleosomal binding patterns using
sucrose density gradient analysis. Mononucleosomal MNase digests from HCT116 cells
were analyzed on sucrose gradients containing 300 mM NaCl. As observed previously,
mononucleosomes containing ~146 bp DNA fragments and core histones, localized in a
peak at fraction 6 (Figure 3.1A). DNMT1 was found dissociated from nucleosomes
forming a peak at fraction 4 while DNMT3A/3B showed strong nucleosome binding,
sedimenting in nucleosome containing fractions and forming a peak at fraction 7.
However various DNMT3A/3B associated proteins like HP1alpha, MeCP2 and UHRF1,
previously suggested to recruit DNMT3A/3B to the chromatin, sedimented separately
from both mononucleosomes and DNMT3A/3B. EZH2 and HDAC1 showed main peaks
in fractions 6-8 as observed previously, independent of the sedimentation profile of the
nucleosomes. These results show that these non-histone proteins are not required for
association of DNMT3A/3B with nucleosomes. In contrast to the auxiliary proteins
described above, G9a, the H3K9 histone methyltransferase, remained tightly anchored to
the nucleosomes similar to DNMT3A/3B enzymes.
To further explore whether these enzymes require the intact nucleosomal
structures for their association with chromatin, we performed the sucrose gradient
chromatin fractionation analysis on the mononucleosomes treated prior with ethidium
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Figure 3.1 DNMT3A/3B along with G9a bind to intact nucleosomal structures.
Mononucleosomes released by extensive digestion with MNase were resolved on a
sucrose density gradient (5%-25%) containing 300 mM NaCl. Mononucleosomes were
incubated in the absence (A) or presence (B) of 300 μg/ml EtBr for 10 min at room
temperature, before loading onto the gradients. Gradients were fractionated and analysed
as described previously. EtBr treatment and chromatin fractionation were performed by
Dr. Shinwu Jeong.
86
bromide (EtBr). The EtBr treatment and chromatin fractionation was performed by Dr.
Shinwu Jeong. EtBr disrupts the nucleosomal structure by intercalating into DNA within
the nucleosomes (McMurray et al., 1991; McMurray and van Holde, 1986), without
interfering with the protein-protein interactions (Lehnertz et al., 2003; Nielsen et al.,
2001). Mononucleosomes were incubated with EtBr prior to loading onto sucrose
gradients containing 300 mM NaCl (Figure 3.1B). DNA fragments of 150 bp were found
mainly in fractions 3 and 4 whereas the histone proteins were detected in fractions 7-16,
possibly due to aggregation (McMurray and van Holde, 1986). The distributions of
DNMT1, HP1-alpha, EZH2, or HDAC1 were not affected by EtBr, again suggesting that
they were not strongly bound to nucleosomes and were freely sedimenting, possibly as
complexes with other proteins in the gradients. However, the distributions of
DNMT3A/3B changed dramatically upon disruption of nucleosomal structure by EtBr,
with the enzymes now sedimenting mainly in factions 3-5, which contained the majority
of the DNA but not the histone components of the nucleosome. These data show that
DNMT3A/3B enzymes require intact nucleosomal structure for their association with
chromatin. G9a also displayed a similar dissociation from nucleosomes upon EtBr
treatment. These data again demonstrate that these enzymes are physically bound to the
nucleosomes and are not directly binding to H3 or simply cosedimenting with them on
the gradients.
87
G9a and SUV39h1 strongly associate with nucleosomes similar to DNMT3A/3B
To understand if G9a plays a role in DNMT3A/3B binding to nucleosomes, we
subjected purified HCT116 nuclei to partial digestion with MNase, yielding a mixture of
mono- and poly-nucleosomes. The nucleosomal digests were then fractionated on sucrose
gradients containing 300 mM NaCl and the distribution of chromatin-associated proteins
was analyzed through immunoblotting (Figure 3.2A). G9a associated strongly with
polynucleosomes with substantial amounts of G9a protein sedimenting in nucleosome-
containing fractions, possibly in association with its own enzymatic products, H3K9me
and H3K9me2 (Collins et al., 2008). Strikingly, the sedimentation profile of G9a was
very similar to that of DNMT3A/3B (Figure 3.2B). Interestingly, SUV39h1 was also
found associated with the polynucleosomes in our gradients. The sedimentation profile of
SUV39h1 was found to be slightly shifted towards the bottom fractions of the gradient,
which contained the larger chromatin fragments indicating association with the condense
heterochromatin (Figure 3.2A,B). However, unlike DNMT3A/3B, we did observe
nucleosome-free G9a and SUV39h1 proteins also in our gradients (data not shown).
We next asked whether G9a and SUV39h1 could bind to mononucleosomes.
Analysis of mononucleosomal digests from HCT116 cells on 300 mM sucrose gradients
showed that G9a and SUV39h1 could bind to mononucleosomes (Figure 3.3A).
However, under such extensive digestion of chromatin, a substantial portion of cellular
SUV39h1 protein dissociated from the nucleosomes, possibly due to disruption of
condensed heterochromatin structure. The sedimentation profiles of G9a and SUV39h1
showed a marked change when the extent of MNase digestion was altered from partial to
88
Figure 3.2 G9a and SUV39h1 associate strongly with polynucleosomes similar to
DNMT3A/3B. (A) Nucleosomes released from nuclei partially digested with MNase at
low ionic strength were resolved by ultracentrifugation on a sucrose density gradient
(5%-25%) containing 300 mM NaCl. Gradients were fractionated and analysed as
described previously. (B) Quantitation of protein bands obtained from the western blot
was done using Quantity One software (Bio-Rad). Plotting of levels of individual
proteins in each fraction shows co-sedimentation of G9a and DNMT3A/3B while
SUV39h1 shows a sedimentation profile shifted towards bottom of the gradient
indicating association with heavier condensed heterochromatin fragments.
B
A
89
Figure 3.3 G9a and SUV39h1 associate with mononucleosomes. Mononucleosomal
digests prepared by extensive MNase digestion of nuclei from (A) HCT116 cells and (B)
293T cells were resolved by ultracentrifugation on a sucrose density gradient (5% to
25%) containing 300 mM NaCl. Gradients were fractionated and analysed as described
previously.
A
B
Core Histones
G9a
HCT116, 300mM, Limit Digestion
C 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Core Histones
G9a
C 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
293T cells, 300mM, Limit Digestion
SUV39h1
SUV39h1
90
extensive, similar to that observed for DNMT3A/3B previously, indicating physical
association of these proteins with nucleosomes. Similar binding of G9a and SUV39h1 to
mononucleosomes was also observed in 293T cells indicating that the strong nucleosomal
association of these proteins takes place in both cell types and is not due to potential cell
type specific interactions (Figure 3.3B). SUV39h1 has previously been suggested to be
recruited to heterochromatin via interaction with HP1alpha which recognizes H3K9me3,
thereby forming a positive-feedback loop required for faithful propagation of H3K9me3
mark (Margueron and Reinberg, 2010). Interestingly, even when HP1alpha dissociated
from nucleosomes in our 300mM NaCl containing sucrose gradients, a fraction of
cellular SUV39h1 protein remained tightly associated with the nucleosomes indicating
presence of an HP1alpha independent mechanism for binding of SUV39h1 to the
nucleosomes, as suggested by one previous study (Krouwels et al., 2005). However, since
HP1alpha is responsible for recruiting DNMT3A/3B to SUV39h1 methylated H3K9me3
regions, anchoring of DNMT3A/3B to nucleosomes through such an HP1alpha
independent chromatin binding of SUV39h1 seems unlikely.
G9a is not essential for maintenance of DNA methylation in somatic cells
Recently, G9a was shown to direct DNA methylation to retrotransposons, major
satellite repeats and densely methylated CpG-rich promoters in ES cells through
recruitment of DNMT3A/3B proteins (Epsztejn-Litman et al., 2008). To ascertain
whether a similar role of G9a exists in maintenance of DNA methylation in somatic cells,
we knocked-down G9a in HCT116 cells. G9a protein level was severely reduced in G9a
91
shRNA (sh-G9a5, sh-G9a7) infected cells compared to the non-specific (N/S) shRNA
control infected cells (Figure 3.4A). DNMT3A protein levels did not show any difference
among the infected cell lines. Next we examined DNA methylation levels at various CpG
poor and CpG island promoter regions and repeats in G9a-knockdown (G9a-kd) and
control infected HCT116 cells. We selected six highly methylated regions including some
G9a-target regions (CpG poor promoters: RUNX3P1, MAGE-A1, SPANXA1; CpG island
promoters: ATBF1, XAGE1; Repeats: LINE1) and one unmethylated region (CpG island:
RUNX3P2) for our analysis. We did not observe any change in DNA methylation levels
at the analyzed regions in the G9a-kd cells compared to control infected HCT116 cells
(Figure 3.4B). This data suggests that unlike in ES cells, G9a is not essential for
maintenance of DNA methylation at these loci in somatic cells, in agreement with some
other recent studies (Kondo et al., 2008; Link et al., 2009).
DNMT3A/3B do not require G9a for anchoring to nucleosomes
The finding that DNA methylation is maintained in the absence of G9a in somatic
cells prompted us to examine whether DNMT3A/3B, which are recuited by G9a for DNA
methylation in ES cells, are still strongly anchored to nucleosomes in the G9a-kd somatic
cells. We analyzed mononucleosomal digests from HCT116 cells infected with either
G9a shRNA or non-specific control on 300 mM sucrose density gradients. DNMT3A/3B
along with G9a were tightly associated with nucleosomes in the control cells as expected
(Figure 3.5A). Interestingly, even in G9a- knockdown cells, DNMT3A/3B remained
strongly bound to the nucleosomes, indicating that strong association of DNMT3A/3B
92
CpG Poor CpG Island Repeat
Figure 3.4 Depletion of G9a does not impair maintenance of DNA methylation in
somatic cells. (A) Western blot of nuclear extracts from HCT116 cells infected with
either control (N/S) shRNA or shRNAs against G9a (sh-G9a5 or shG9a7) prepared 14
days after infection. G9a knochdownat protein level was confirmed using antibody
against G9a. (B) The levels of DNA methylation at different loci in HCT116 cells
infected with either control (N/S) shRNA or shRNAs against G9a (sh-G9a5 or shG9a7)
were measured through Ms-SNuPE 14 days after infection.
A
B
93
Figure 3.5 DNMT3A/3B and G9a do not require each other for anchoring to
nucleosomes in somatic cells. Mononucleosomal digests prepared by extensive MNase
digestion of nuclei from (A) HCT116 cells infected with either control N/S shRNA or
shG9a5 shRNA construct and (B) hypomethylated DKO1 cells, were resolved by
ultracentrifugation on a sucrose density gradient (5% to 25%) containing 300 mM NaCl.
Gradients were fractionated and analysed as described previously.
Core Histones
G9a
C 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
DKO1
A
B
94
with methylated chromatin regions observed in somatic cells is not mediated by G9a. We
also asked whether impaired DNA methylation could feed back on H3K9 methylation
and disrupt binding of G9a to H3K9 methylated regions. Mononucleosomal digests from
severely hypomethylated DNMT-deficient DKO1 (DNMT1
ΔE2-5
, DNMT3B
−/−
) cells
were analyzed on sucrose gradients containing 300 mM NaCl. The majority of G9a was
found tightly anchored to nucleosomes even in DKO1 cells indicating that a loss of DNA
methylation and absence of DNMTs does not affect G9a’s association with the chromatin
(Figure 3.5B).
Next, we asked whether G9a could target the truncated delta DNMT3B variants,
which show weak chromatin binding affinities, to nucleosomes (Jeong et al., 2009). G9a
binds to DNMT3A/3B through the interaction of its ankyrin domain with the catalytic
domains of DNMT3A/3B, therefore, it should be able to physically interact with the
catalytically active delta DNMT3B variants (Figure 3.6A). Mononucleosomal digests
from 293T cells expressing Myc-tagged delta DNMT3B variants were analyzed on
sucrose gradients containing 300 mM salt. While G9a remained tightly bound to the
nucleosomes, the delta DNMT3B variants completely dissociated from the nucleosomes
(Figure 3.6B). Taken together, these data suggest that unlike ES cells, where G9a is
essential for DNA methylation through recruitment of DNMT3A/3B enzymes to the
H3K9me3 containing chromatin regions, G9a is not essential for anchoring of
DNMT3A/3B to the nucleosomes and for propagation of DNA methylation in somatic
cells.
95
Figure 3.6 G9a cannot anchor ∆DNMT3B truncated variants to the nucleosomes.
(A) Map of delta DNMT3B isoforms showing the PWWP and PHD-like domains located
in the N-terminal regions, and the catalytic methylase domains in the C-terminal region.
(B) Mononucleosomal digests prepared by extensive MNase digestion of nuclei from
293T cells expressing either ∆DNMT3B2 or ∆DNMT3B4, were resolved by
ultracentrifugation on a sucrose density gradient (5% to 25%) containing 300 mM NaCl.
Gradients were fractionated and analysed through immunoblotting as described
previously.
A
B
96
DISCUSSION
The strong association of DNMT3A/3B with nucleosomes in somatic cells is very
striking. Such anchoring may be critical for maintenance of DNA methylated states
through somatic divisions. Similar mechanisms of epigenetic inheritance have recently
been proposed for propagation of histone marks where the enzymes responsible for
establishment of the respective histone marks remain associated with the chromatin
domains containing their own mark enabling faithful maintenance (Collins et al., 2008;
Felsenfeld and Groudine, 2003; Hansen et al., 2008). Strong binding of H3K9
methyltransferases, G9a and SUV39h1, to nucleosomes observed in our sucrose density
gradient experiments supports such hypotheses. Since histone marks are dynamically
regulated by the combined action of histone methylases and demethylases, continued
association of such HMTases with chromatin regions containing their enzymatic product
would be critical for faithful propagation of histone marks in somatic cells. Our data also
reveals the existence of an HP1 independent targeting mechanism for association of
SUV39h1 with the chromatin. Since HP1 displays a highly transient interaction with
heterochromatin (Krouwels et al., 2005), the presence of additional mechanisms which
could target SUV39h1 to the heterochromatic regions would be critical for proper
maintenance of heterochromatin.
Recruitment of DNMT3A/3B to specific chromatin domains through interactions
with various chromatin associated proteins has previously been shown to be essential for
de novo methylation in ES cells (Law and Jacobsen, 2010). In this study, we examined
whether the interactions with such auxiliary proteins are essential for the strong
97
anchoring of DNMT3A/3B to nucleosomes in somatic cells. Our data shows that most of
the DNMT3A/3B interacting auxiliary proteins such as HP1, MeCP2, EZH2, HDAC1
and UHRF1, shown to be critical for DNA methylation by DNMT3A/3B in ES cells,
associate weakly with the chromatin and are not required for the tight binding of
DNMT3A/3B to the nucleosomes as observed in somatic cells. Rather, an intact
nucleosomal structure is required for DNMT3A/3B association with nucleosomes,
suggesting that their interactions with both the DNA and the histone components of the
nucleosome are essential for such binding. Such interactions, possibly through their
specific domains such as PWWP and ADD domain (Dhayalan et al., 2010; Ooi et al.,
2007; Qiu et al., 2002; Zhang et al., 2010), may work synergistically to anchor these
proteins to the nucleosomes as also suggested by our data in Chapter 2.
H3K9 methylation and DNA methylation patterns are found to be highly
coincident in mammalian cells. In plants, H3K9 methylation has previously been shown
to direct DNA methylation (Margueron and Reinberg, 2010). Recently, several studies
have suggested existence of a similar link between H3K9 methylation and DNA
methylation in mammalian cells. Knockdown of H3K9 HMTases, SUV39h1, SUV39H2
or G9a, has been shown to result in reduced DNA methylation at certain loci in ES cells
(Law and Jacobsen, 2010). Recruitment of DNMT3A/3B to H3K9 methylated regions
through interactions with proteins such as HP1, SUV39h1 and G9a has been proposed to
be critical for DNA methylation of such loci in ES cells. Here we show that unlike ES
cells, these proteins are not essential for targeting of DNMT3A/3B to nucleosomes in
somatic cells. Depletion of G9a in somatic cells does not impair maintenance of DNA
98
methylation at G9a-target loci, suggesting the presence of other mechanisms involved in
faithful propagation of DNA methylation patterns in somatic cells. This hypothesis is
further supported by our finding that DNMT3A/3B remain bound to their target regions
even in the absence of G9a in somatic cells indicating a G9a-independent targeting
mechanism that enables proper compartmentalization of these enzymes. Interestingly, we
also found that the presence/absence of DNA methylation does not affect association of
G9a, which binds to H3K9me and H3K9me2 residues, with nucleosomes, suggesting that
impaired DNA methylation does not feedback on H3K9 methylation in somatic cells.
Taken together, these data indicate that anchoring of DNMT3A/3B to nucleosomes in
somatic cells does not require the known DNMT3A/3B interacting auxiliary proteins and
may involve direct interaction with the nucleosomes itself. Proteins involved in H3K9
methylation such as G9a, HP1 or SUV39h1 may play a role in de novo methylation in ES
cells through recruitment of DNMT3A/3B, but are not essential for maintenance of DNA
methylation in differentiated somatic cells suggesting presence of some other mechanism
involved in this maintenance process.
99
CHAPTER 4
ROLE OF DNA METHYLATION IN ASSOCIATION OF
DNMT3A/3B WITH NUCLEOSOMES
INTRODUCTION
In mammals, DNA methylation patterns are generally thought to be established
during embryonic development by de novo DNA methyltransferases 3A and 3B (Okano
et al., 1999) and then stably maintained through multiple somatic divisions by the
‘maintenance activity’ of DNMT1 both during and after replication (Law and Jacobsen,
2010). However, recent studies suggest that DNMT1 requires co-operative activity of the
de novo DNMT3A/3B enzymes for proper maintenance of methylation patterns (Chen et
al., 2003; Liang et al., 2002; Riggs and Xiong, 2004), which are ubiquitously expressed
in somatic cells. Based on our previous data, we proposed a revised model of inheritance
assigning DNMT3A/3B to a maintenance role in somatic cells (Jones and Liang, 2009);
however, questions still remain regarding the molecular mechanisms guiding the
maintenance activity of these de novo enzymes.
In embryonic stem (ES) cells, DNMT3A/3B establish methylation patterns in
association with DNMT3L, a regulatory factor which stimulates DNMT3A/3B de novo
activity (Gowher et al., 2005a) and targets them to nucleosomes containing unmethylated
H3K4 residues (Ooi et al., 2007). Methylated H3K4 containing chromatin regions remain
refractory to such DNA methylation (Okitsu and Hsieh, 2007; Weber et al., 2007).
100
Further, heterochromatin protein 1 (HP1) recruits DNMT3A/3B to H3K9me3 residues,
established by histone methyltransferase (HMTase) Suv39h1/2, enabling de novo DNA
methylation in pericentric heterochromatin (Lehnertz et al., 2003). In euchromatic
regions, G9a, another H3K9 HMTase, recruits DNMT3A/3B for de novo methylation of
early embryonic gene promoters (Epsztejn-Litman et al., 2008). UHRF1, which assists
DNMT1 in locating to hemimethylated sites (Sharif et al., 2007), also targets
DNMT3A/3B for de novo methylation in ES cells (Meilinger et al., 2009). However,
DNMT3L is expressed only during gametogenesis and embryonic stages and not in
somatic tissues (Aapola et al., 2000; Bourc'his et al., 2001). Further, we and others have
recently shown that HP1 and UHRF1 are not required for DNMT3A/3B’s association
with nucleosomes (Jeong et al., 2009) and G9a does not affect maintenance of DNA
methylation in somatic cells (Kondo et al., 2008; Link et al., 2009). Thus, other
mechanisms must exist to ensure proper localization of these enzymes to silent chromatin
regions in somatic cells (Bachman et al., 2001), enabling faithful maintenance of
methylated states.
We and others have previously shown that the majority of DNMT3A/3B within a
somatic cell are strongly anchored to nucleosomes containing methylated DNA with little
free DNMT3A/3B proteins existing (Jeong et al., 2009; Schlesinger et al., 2007). Here we
show that the presence of such methylated regions is essential for DNMT3A/3B’s
association with chromatin and quite unexpectedly, also for maintaining the cellular
levels of these enzymes. Reduction in DNA methylation levels results in reduced
DNMT3A/3B binding to nucleosomes accompanied by selective degradation of the free
101
enzymes through the proteosomal pathway. Restoration of DNA methylation increases
DNMT3A/3B protein levels through their stabilization on nucleosomes. Further, pre-
existing methylation stimulates propagation of DNA methylation in vivo by stably
anchoring DNMT3A/3B to nucleosomes. DNMT3A/3B work synergistically to
propagate methylation patterns with DNMT3B stimulating DNMT3A activity by
promoting its association with nucleosomes, similar to DNMT3L. Taken together, these
data suggest an inheritance model where DNMT3A/3B remain localized to silent
methylated domains by binding to nucleosomes containing methylated DNA, enabling
faithful maintenance of methylated states in cooperation with DNMT1; while non-
anchored DNMT3A/3B enzymes get selectively degraded preventing spurious de novo
methylation.
102
MATERIALS AND METHODS
Cell culture and Drug treatment
HCT116 derivative cell lines were maintained in McCoy’s 5A medium containing
10 % inactivated fetal bovine serum, 100 units/ml penicillin and 100 μg/ml streptomycin.
Puromycin was included in the culture medium at 3 μg/ml to maintain infected HCT116
derivative cell lines. When indicated, cycloheximide (Sigma) was added to a final
concentration of 50 μg/ml. The proteosome inhibitor MG132 (Calbiochem) was used at
10 μM for 2 h prior to CHX treatment.
RNA Isolation and RT-PCR
Total RNA was isolated by using TRIzol reagent (Invitrogen, Carlsbad, CA). For
HCT116 derivative cells infected with different DNMTs, RNA was extracted 8 weeks
after infection. Total RNA (4 μg) was transcribed with M-MLV reverse transcriptase
(Invitrogen). Real-time quantitative PCR was performed on the Opticon Real-time PCR
system (Bio-Rad, Hercules, CA). Minus-RT reactions generated were used as negative
controls. For quantitation, standard dilution curves were included on each plate. All
samples were analyzed in triplicates. GAPDH and PCNA were used as endogenous
controls. Individual data were converted to relative values based on the standard curve
and were normalized to GAPDH or PCNA values of the same sample. For primer and
probe sequences, refer to the manuscript.
103
Expression vector construction
Human 3B1, ΔDNMT3B2 and DNMT3L cDNA sequences having the Myc tag
DNA sequence ligated to their 5’ ends were amplified from the pIRESpuro/Myc
constructs (Jeong et al., 2009) (a modified version of the pIRESpuro3 vector, Clontech),
a generous gift from Allen Yang (USC), using polymerase chain reaction (PCR). Myc-
tagged catalytically-inactive mutant of DNMT3B1, having a cysteine to serine alteration
in the catalytic domain corresponding to position 657 of DNMT3B1 protein, was
prepared using a site-directed mutagenesis kit (Stratagene). The mutation was confirmed
by sequencing both strands of the construct. For preparation of the constructs, the
lentivirus vector pLJM1 was linearized using AgeI and EcoRI restriction enzymes and the
Myc tagged DNMT cDNAs were cloned in it using In-fusion advantage PCR cloning kit
(Clontech) following manufacturer’s protocol. Lentiviral constructs were prepared by Dr.
Daniel De Carvalho. For lentivirus production, the vesicular stomatitis virus envelope
protein G expression construct pMD.G1, the packaging vector pCMV ΔR8.91 and the
transfer vector pLJM1 were used as described previously (Ou et al., 2009). Infected
HCT116 derivative cells, stably expressing various DNMTs, were selected in the
presence of 3 μg/ml puromycin for three weeks.
Nuclear and whole cell lysates preparation
Nuclei were prepared as described previously in Chapter 2. Nuclear extracts were
prepared by resuspending nuclei in RIPA buffer (50 mM Tris-HCl, pH 8.0, 150 mM
NaCl, 1% NP-40, 0.5% DOC, 0.1% SDS) followed by sonication. Whole cell lysates
104
were prepared by resuspending cells, washed twice with PBS, in RIPA buffer followed
by sonication.
Salt extraction of nuclei
Salt extraction was performed as described in Chapter 2. Nuclear pellets were
resuspended in RIPA buffer and subjected to sonication. Proteins in the supernatant were
concentrated using TCA precipitation and later resuspended in RIPA buffer. Equivalent
volumes of supernatant and pellet fractions were added to SDS loading buffer and
subjected to Western blotting.
MNase digestion and sucrose density gradient centrifugation
MNase digestion and sucrose gradients experiments were performed as described
previously in Chapter 2 (Jeong et al., 2009).
Western blot and Immunofluorescence (IF) analyses
Protein samples were dissolved in SDS/β-mercaptoethanol loading buffer, and
resolved on a 4-15 % gradient SDS/PAGE gel (Bio-Rad, Hercules, CA). Antibodies
against H3 (ab1791), DNMT3A (ab2850) were purchased from Abcam Inc. (Cambridge,
UK); EZH2 (ac22) from Cell Signaling Technology, Inc.(Danvers, MA); DNMT1 (sc-
20701), DNMT3B (sc-10235), GST (sc-138) and p53 (sc-126) from Santa Cruz Biotech.
(Santa Cruz, CA); Myc epitope tag (05-724) from Upstate (now Millipore, Billerica,
MA); G9a (G 6919) and β-Actin (A 5316) from Sigma (Saint Louis, MO). Image of
105
individual proteins was visualized using ECL detection system (Thermo Scientific,
Waltham, MA and Millipore, Billerica, MA) and bands were quantified using Quantity
One software (BioRad, Hercules, CA). Immunofluorescence experiments were performed
as described previously (Wang et al., 2009). The antibody used for DNMT3A (sc-20703)
was purchased from Santa Cruz Biotech. (Santa Cruz, CA). Detailed protocols can be
found in the related manuscript.
Native Immunoprecipitation assay
Mononucleosome extracts for the native IP experiment were prepared by
digesting nuclei with MNase in 1 ml of ice-cold digestion buffer containing 50 mM Tris-
HCl, pH 8.0, 100 mM NaCl, 8 mM MgCl2, 3 mM CaCl2, 1mM DTT and protease
inhibitors for 15 min at 37°C. The reaction was stopped with EDTA/EGTA (up to 10
mM), and left at room temperature for 20 min, before collecting the soluble fraction of
nucleosomes. Nucleosomes were incubated with antibodies overnight at 4°C in cold
room in the MNase digestion buffer and then incubated with protein-A agarose beads for
3 hr. The beads were washed three times with the digestion buffer before the bound
proteins were eluted in the elution buffer. Proteins pulled down by different antibodies
were later analyzed by Western blot analysis. Antibodies used for IP were DNMT3A
(ab2850) from Abcam (Cambridge, UK) and Myc epitope tag (05-724) from Upstate
(now Millipore, Billerica, MA). IgG (sc-2027) and CD8 (sc-32812) from Santa Cruz
Biotech. (Santa Cruz, CA) were used as the negative controls for IP.
106
Reconstitution of Mononucleosomes
Mononucleosomes were reconstituted with recombinant histones and modified
601 (M601)(Bouazoune et al., 2009) sequence by salt dialysis method(Hamiche et al.,
1999). For preparing methylated nucleosomes, M601 sequence was methylated using
M.SssI and S-adenoylmethionine (SAM) (New England Biolabs) and purified using Min
Elute Spin Columns (Qiagen) before being used for reconstitution experiments. Complete
methylation of the fragments was confirmed through HpaII-MspI digestion method. After
reconstitution, any resulting precipitates were removed using microcentrifugation.
GST-pull down assay
GST-DNMT3A (H00001788-P01) and GST-DNMT3A cat. (H00001788-Q01)
recombinant proteins were purchased from Abnova (Taipei, Taiwan). Purified GST
protein was kindly provided by Woojin An (USC). Reconstituted nucleosomes containing
either methylated or unmethylated M601 DNA were incubated with 1 μg of recombinant
GST-DNMT3A, GST-DNMT3A cat. or GST proteins over night at 4°C in a binding
buffer containing 50 mM Tris-HCl pH 8.0, 100 mM NaCl, 0.1% NP-40, 10 μM ZnCl
2
, 1
mM DTT and protease inhibitors. The samples were then incubated with GST resins (GE
Healthcare) at 4°C for 3 hrs. The resins were washed three times with the binding buffer
before elution of the bound proteins. Bound proteins were later analyzed by Western blot
analysis.
107
DNA methylation analysis
Genomic DNA (10 μg) isolated from various HCT116 derivative cell lines was
digested with methylation-sensitive restriction enzymes, HpaII or MspI (New England
Biolabs), at 37°C over night. The digested DNA was run on an agarose gel at low voltage
for 8 hrs in order to achieve good separation. The undigested DNA band in each lane was
then quantified using the ImageQuant software. Percentage of genomic methylation
present was calculated using the formula:
% methylation = ((H-M) X 100) / G ,
where H= undigested with HpaII; M= undigested with MspI and G= genomic DNA.
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RESULTS
DNMT3A protein level decreases on depletion of global DNA methylation
In somatic cells, DNMT3A/3B remain bound to nucleosomes containing
methylated DNA (Jeong et al., 2009). To investigate the role of DNA methylation in this
binding, we used a series of HCT116 colon cancer cells with homozygous deletions for
DNMT1 (DNMT1
ΔE2-5
; 1KO) (Egger et al., 2006; Rhee et al., 2000), DNMT3B
(DNMT3B
-/-
; 3BKO) or both DNMT1
and DNMT3B (DNMT1
ΔE2-5
/ DNMT3B
-/-
; double
knockout, DKO) and consequently different levels of genomic DNA methylation (Rhee
et al., 2002). For the DKO cells, which still contain residual DNMT1 activity (Egger et
al., 2006), we used two clones for our analysis, DKO1 and DKO8, having lost ~95% and
~50% DNA methylation respectively (Rhee et al., 2002). RT-PCR analysis of DNMT3A,
DNMT3B and DNMT1 transcript levels in the various HCT116 derivative cell lines
showed similar or higher levels of DNMT3A transcripts in HCT116 knockout cell lines
compared to WT HCT116; reduced levels of DNMT1
ΔE2-5
hypomorph transcripts in
1KO and the two DKO clones, with relatively higher expression in the DKO8 clone and
no detectable levels of DNMT3B transcripts in 3BKO and both DKO cell lines,
consistent with previous data (Egger et al., 2006; Rhee et al., 2002) (Figure 4.1A).
Next we examined DNMT protein levels in these cell lines through
immunoblotting of nuclear extracts. Similar to mRNA analysis, DNMT3B and DNMT1
protein levels were severely reduced in the respective knockout cell lines (Figure 4.1B).
Surprisingly, while DNMT3A mRNA levels were higher in both DKO clones, we found
dramatically reduced DNMT3A protein in them compared to WT HCT116 cells. G9a,
109
Figure 4.1 Transcription-independent decrease in DNMT3A protein level in
hypomethylated DKO cells. (A) DNMT3A, DNMT3B and DNMT1 mRNA levels in
wild-type (WT) HCT116 and different knockout cell lines were measured by RT-PCR.
Results are normalized to GAPDH mRNA levels. Data represents mean and standard
deviation of triplicate PCR reactions from a single experiment, representative of two
independent biological replicate experiments. (B) Western blot analysis of nuclear
extracts from various HCT116 derivative cell lines using different antibodies. In order to
detect the truncated DNMT1
ΔE2-5
hypomorph expressed in 1KO and DKO cells, an
antibody directed against the C-terminus of DNMT1 was used. (C) DNA methylation
analysis of WT HCT116 and knockout cell lines using methylation-sensitive restriction
enzymes. Genomic DNA isolated from the cells was digested with HpaII or MspI
enzymes and the methylation level estimated as described in the Materials and Methods
section. Data is presented as percentage of methylation retained compared to WT
HCT116 methylation levels. Data presented is from a single experiment, representative of
two biological replicate experiments.
110
another chromatin-modifying protein, did not display such large changes in protein levels
in HCT116 knockout cell lines (Figure 4.1B).
Assessment of global DNA methylation levels using methylation-sensitive
restriction enzymes revealed a direct correlation between the amount of DNMT3A
protein and level of methylation retained in the knockout cells, suggesting a possible role
of DNA methylation in maintaining cellular DNMT3A levels (Figure 4.1B, C). DKO8
cells, which had retained higher DNA methylation levels, showed higher DNMT3A
protein compared to the minimal amount present in the severely hypomethylated DKO1
cells. Immunofluorescence analyses of HCT116 and DKO cells displayed similar
reduction in DNMT3A protein levels in DKO cells as observed in western blots of their
nuclear extracts. Moreover, the residual DNMT3A protein displayed similar nuclear
distribution in DKO cells as in WT HCT116 cells, suggesting that its reduced nuclear
levels present in the DKO cells are not the result of protein mislocalization (Figure 4.2).
These findings were further confirmed through western blotting of whole cell lysates of
the HCT116 derivative cell lines (Figure 4.3). Since no such decrease in DNMT3A
protein was observed in the single DNMT1 and DNMT3B knockout cells (1KO and
3BKO respectively), which retained substantial levels of DNA methylation, maintenance
of DNMT3A levels through possible protein-protein interactions with DNMT1 and/or
DNMT3B seems unlikely.
111
HCT116
DKO8
DKO1
DAPI DNMT3A
Figure 4.2 Residual DNMT3A protein remains localized within nuclei in DKO cells.
Co-cultured HCT116, DKO8 and DKO1 cells were immunostained for DNMT3A (green)
using a rabbit polyclonal DNMT3A antibody and their nuclei (blue) were stained with
4,6-diamidino-2- phenylindole. Scale bar, 20 μm.
112
Figure 4.3 Decrease in DNMT3A level in DKO cells is not due to protein
mislocalization. Western blot analysis of whole cell extracts from various HCT116
derivative cell lines was performed using DNMT3A antibody. Actin was used as the
loading control.
DKO1
3BKO
1KO
HCT116
DKO8
DNMT3B
DNMT3A
ACTIN
DKO1
3BKO
1KO
HCT116
DKO8
DNMT3B
DNMT3A
ACTIN
113
Residual DNMT3A protein remains tightly bound to chromatin in the DKO cells
We have previously shown that DNMT3A/3B strongly associate with methylated
chromatin regions (Jeong et al., 2009). To determine whether the residual DNMT3A
protein in hypomethylated DKO cells retains similar affinity for chromatin as in WT
HCT116 cells, we performed a salt extraction experiment as described previously (Jeong
et al., 2009). Purified nuclei from HCT116 and DKO1 cells were incubated in buffers
with increasing concentrations (50mM to 400mM) of NaCl. Nuclear pellet and
supernatant fractions were independently analyzed through western blot analysis. As
expected, similar amounts of core histones remained inside the extracted nuclei under all
salt concentrations. In HCT116 cells, the DNMT3A protein level remained almost
constant within the nuclei up to 400mM NaCl indicating a strong binding affinity for
chromatin (Figure 4.4), whereas other chromatin associated proteins such as EZH2 and
G9a showed relatively weaker binding affinities with substantial amounts detected in the
supernatant at more than 200mM NaCl concentrations. Interestingly, almost all of the
DNMT3A protein present in DKO1 cells, though greatly reduced in comparison to WT
HCT116, also remained tightly associated with the chromatin up to 400mM NaCl (Figure
4.4), possibly binding to the few methylated regions remaining in the DKO1 cells.
Meanwhile, EZH2 and G9a showed weaker binding to chromatin, similar to that
observed in WT HCT116. These data suggest that binding to methylated chromatin
regions may be critical for stabilization of DNMT3A protein.
114
Figure 4.4 Residual DNMT3A protein remains strongly anchored to chromatin in
hypomethylated DKO1 cells. Nuclei purified from WT HCT116 and DKO1 cells were
incubated in nondenaturing extraction buffers containing 50 to 400 mM NaCl for 5 min.
Equivalent volumes of both supernatant and pellet fractions were subjected to western
blot analysis using specific antibodies. Ponceau S staining shows core histones
transferred onto the membrane from the SDS/PAGE gel. For detecting low levels of
DNMT3A in DKO1 cells, blots for both the supernatant and pellet fractions from DKO1
cells were overexposed for 5 fold more time duration compared to HCT116 cells, as
indicated by *.
115
Decreased protein stability of DNMT3A in hypomethylated DKO cells
To assess whether the dramatic transcription-independent decrease in steady-state
levels of DNMT3A protein observed in hypomethylated DKO cells was due to altered
protein stability, we treated WT HCT116 and DKO8 cells with the protein synthesis
inhibitor cycloheximide (CHX) (Vega et al., 2004) and measured the DNMT3A protein
remaining at different time points after treatment. DNMT3A was stable in WT HCT116
cells with 93% still remaining after 6 hrs of CHX treatment (Figure 4.5A). However, in
DKO8 cells, DNMT3A was very unstable with its level rapidly decreasing to 49% 2 hrs
after treatment. The half-life of DNMT3A protein decreased dramatically from 16 hrs in
WT HCT116 to 7 hrs in DKO8 cells (Figure 4.5B). Interestingly, after a rapid initial
decrease in DNMT3A protein level in DKO8 cells within the first 2 hrs of CHX
treatment, a fraction of DNMT3A protein remained stable thereafter till the 8 hr time
point. This fraction may possibly represent the stable DNMT3A protein bound to the
methylated chromatin regions in DKO8 cells, similar to that observed in DKO1 cells
(Figure 4.4). Taken together, these data indicate that a decrease in DNA methylation
results in destabilization of DNMT3A protein, possibly due to reduced chromatin binding
in the absence of methylated DNA regions, the main sites of DNMT3A/3B binding
(Jeong et al., 2009).
Restoration of global DNA methylation rescues DNMT3A protein level
To ascertain if depletion of DNA methylation is primarily responsible for the
decrease in DNMT3A protein, we sought to restore DNA methylation in the DKO cells.
116
Figure 4.5 Removal of DNA methylation results in decreased stability of DNMT3A
protein in DKO cells. (A) WT HCT116 and DKO8 cells were treated with
cycloheximide (CHX) and the levels of DNMT3A protein remaining at different time
points after treatment were determined by western blotting of nuclear extracts. (B)
Quantitation of protein bands was done using Quantity One software (Bio-Rad). The data
points represent DNMT3A levels, normalized to actin, at different time points presented
as the fraction of protein remaining compared to levels present before CHX treatment.
Straight lines represent linear regression adjustment of the individual time points. The
half-life of the DNMT3A protein decreased from 16 hr in WT HCT116 to 7 hr in DKO8
cells. p53 and actin were used as positive and loading controls, respectively. The data
presented is from a single experiment, which is representative of two independent
biological replicate experiments.
A
B
117
We expressed Myc-tagged DNMT3B1, ∆DNMT3B2 (Wang et al., 2006b) or DNMT3L
in DKO1 and DKO8 cells using a lentiviral system and confirmed expression of the
relevant proteins by immunoblotting (Figure 4.6A). Global DNA methylation levels were
measured 8 weeks post-infection using methylation-sensitive restriction enzymes. We
observed increased DNA methylation in both DKO cell lines infected with DNMT
constructs compared to empty vector controls (Figure 4.6B). Even though there was
equivalent mRNA expression of exogenous DNMT enzymes in the two DKO clones
(Figure 4.7A), DKO8 cells, with higher baseline methylation levels, showed a greater
increase in methylation compared to hypomethylated DKO1 cells for each individual
construct. Moreover, the increase in methylation in the infected cells was preferentially
localized to loci having low-levels of pre-existing methylation and minimal de novo
methylation of previously unmodified sites could be observed (Figure 4.7B). These
results indicate a stimulatory effect of pre-existing methylation (Kim et al., 2002) on
DNA methylation by DNMTs in vivo, possibly through stabilization of de novo
DNMT3A/3B enzymes on methylated nucleosomes as suggested by their higher protein
levels in DKO8 cells (Figure 4.1B, 4.6A). This process may further be enhanced by the
higher levels of DNMT1 hypomorph present in DKO8 cells (Vilkaitis et al., 2005)
(Figure 4.1B). Within each DKO clone, exogenous DNMT3L expressing cells showed
the most robust increase in methylation followed by DNMT3B and ∆DNMT3B2
expressing cells respectively, re-emphasizing the strong stimulatory effect of DNMT3L
on DNMT3A/3B activity observed in ES cells (Gowher et al., 2005a). These methylation
118
Figure 4.6 Increase in DNA methylation upon expression of exogenous DNMTs in
DKO cells. (A) Expression of Myc-tagged DNMT3B1, ∆DNMT3B2 and DNMT3L
proteins, infected using a lentiviral system, in DKO cells was confirmed by
immunoblotting of nuclear extracts using a Myc antibody. Lentiviral constructs prepared
by Dr. Daniel D De Carvalho. (B) DNA methylation analysis of infected DKO cells using
methylation-sensitive restriction enzymes. Genomic DNA was isolated from infected
cells eight weeks after infection and methylation level was estimated as described in
Figure 4.1. Data is presented as the percentage of methylation retained compared to WT
HCT116 methylation levels.
A
B
119
Figure 4.7 Pre-existing methylation at genomic loci guides and stimulates DNA
methylation by DNMTs in somatic cells. (A) RT-PCR analysis was performed using
primers for DNMT3B to check Myc-DNMT3B1 & Myc-∆DNMT3B2 expression in DKO
cells. The results are normalized to GAPDH mRNA levels. Data represents mean and
standard deviation of triplicate PCR reactions from a single experiment, which is
representative of two independent biological replicate experiments. (B) A supervised
cluster analysis is shown for DNA methylation at over 27000 CpG sites in HCT116 cells
and DKO cells infected with E/V or different DNMT lentiviral constructs, using the
Illumina Infinium methylation assay. Analysis was performed by Dr. Daniel D De
Carvalho.
A
B
120
data were further confirmed through Illumina Infinium analysis (Laird, 2010) for each
infected cell line (Figure 4.7B).
Interestingly, immunoblotting of nuclear extracts revealed a substantial
transcription-independent increase in DNMT3A protein level in all DNMT infected DKO
cell lines (Figure 4.8A,B). Moreover, the increase in DNMT3A correlated with the
increase in global DNA methylation levels. Considering that DNMT3A primarily
associates with methylated chromatin regions, these data suggest that presence of such
methylated regions is required for maintaining its protein level in somatic cells.
DNA methylation induced DNMT3A increase is mediated by strong anchoring to
nucleosomes
To examine whether the increase in DNMT3A protein observed upon restoration
of DNA methylation is mediated by binding to nucleosomes, we used sucrose density
gradient analysis which allows for the study of in vivo interactions between the chromatin
modification enzymes and their actual nucleosomal substrates in the native state (Jeong et
al., 2009). Mononucleosomal digests prepared by extensive micrococcal nuclease
(MNase) digestion of nuclei from infected DKO8 cells, expressing either Myc-tagged
DNMT3B1, ∆DNMT3B2 or DNMT3L, were subjected to fractionation on sucrose
gradients containing 300mM NaCl. Western blot analysis showed similar nucleosomal
profile in all gradients with mononucleosomes forming a peak at fraction 6 (Figure 4.9).
The DNMT fusion proteins displayed distinct sedimentation profiles indicating different
nucleosome binding affinities. DNMT3B1 associated strongly with nucleosomes while
121
Figure 4.8 Transcription-independent increase in DNMT3A protein level upon
increase in DNA methylation. (A) RT-PCR analysis was performed for analyzing
endogenous DNMT3A mRNA levels in the DKO8 and DKO1 cells, 8 weeks after
infection with different DNMTs. WT HCT116, 1KO and 3BKO cell lines were also
included in the analysis. Results are normalized to PCNA mRNA levels. Data represents
mean and standard deviation of triplicate PCR reactions from a single experiment, which
is representative of two independent biological replicate experiments. E/V: Empty
Vector. (B) Western blot analysis of nuclear extracts prepared from infected DKO cells
and different HCT116 derivative cell lines using a DNMT3A antibody. Histone H3 was
used as the loading control. Data presented in this figure is representative of two
biological replicate experiments. E/V: Empty Vector
A
B
122
Figure 4.9 The increased level of DNMT3A protein in infected DKO cells remains
tightly bound to nucleosomes. Mononucleosomal digests prepared by extensive MNase
digestion of infected DKO8 nuclei, were resolved by ultracentrifugation on a sucrose
density gradient (5% to 25%) containing 300 mM NaCl. Gradients were fractioned into
16 aliquots numbered 1-16 starting from the top of the centrifuge tube. To probe the
distribution of proteins in each fraction, western blotting was performed with various
antibodies after TCA precipitation of proteins from each fraction. Ponceau S staining
shows core histones transferred onto the membrane from the SDS/PAGE gel.
Mononucleosomes peaked in fraction 6 and the small proportion of higher order
oligonucleosomes remaining in the digests sedimented in later fractions. The control
lanes on the gels were loaded with unfractionated nuclear extract to monitor the quality of
the immunostaining of the membranes.
123
the truncated ∆DNMT3B2 variant showed weak association, consistent with previous
data (Jeong et al., 2009). DNMT3L showed a bimodal distribution having both
nucleosome-free and nucleosome-bound protein (fractions 1-4 and 5-16 respectively).
Strikingly, DNMT3A remained strongly associated with nucleosomes in all infected cell
lines, independent of the nucleosome binding affinities of the exogenous proteins,
suggesting a nucleosome anchorage dependent stabilization of the protein. DNMT3A
formed a peak at fraction 7 in DNMT3B1 and ∆DNMT3B2 expressing cells. In
DNMT3L expressing cells, the peak was shifted to fraction 9, indicating the formation of
heavier DNMT3A-DNMT3L tetramer encasing the nucleosome (Jia et al., 2007). We did
not observe any DNMT3A in the nucleosome-free fractions (1-4) co-sedimenting with
the unbound pool of ∆DNMT3B2 or DNMT3L fusion proteins, suggesting that increase
in DNMT3A is not due to stabilization through protein-protein interactions with the
infected proteins. Instead, DNMT3A’s stabilization is mediated by strong binding to
nucleosomes upon increase in DNA methylation, which may be required for such
binding. Our in vitro binding experiments using recombinant DNMT3A and reconstituted
nucleosomes, with either methylated or unmethylated DNA, further support this idea
showing that DNMT3A can in fact stay associated with its product (i.e. methylated DNA)
in a nucleosomal context, even in the absence of any facilitatory histone marks (Figure
4.10A,B). However, the DNMT3A catalytic domain by itself could not bind to either
methylated or unmethylated nucleosomes indicating a role for the N-terminal regulatory
region in nucleosome binding (Gowher et al., 2005b). Taken together, these data suggest
124
Figure 4.10 Recombinant DNMT3A can remain associated with nucleosomes
containing methylated DNA. (A) Reconstitution of mononucleosomes with recombinant
histones and M601 DNA was analyzed by running naked M601 DNA and reconstituted
M601 DNA, unmethylated and M.SssI methylated, on a 1.5% agarose gel. The gel was
stained with EtBr for 30 min after completion of the run. (B) Reconstituted
mononucleosomes were incubated with either GST-DNMT3A, GST-DNMT3A cat.
(catalytic domain only) or GST proteins. Proteins pulled down using GST resins were
analyzed by Western blot analysis. Input denotes reconstituted nucleosomes alone. GST
antibody was used to detect pulled down GST-tagged proteins. H3 antibody was used to
detect mononucleosomes pulled down along with the GST-tagged proteins. Un,
Unmethylated; Me, Methylated; Nuc, Nucleosomes.
A
B
125
that DNMT3A protein is stabilized by binding to nucleosomes containing its own product
(i.e. methylated DNA), which is essential for maintaining its cellular levels.
Reduced nucleosome binding and degradation of unbound DNMT3B upon depletion
of DNA methylation
DNMT3B, like DNMT3A, also compartmentalizes to methylated regions in
somatic cells via strong anchoring to nucleosomes containing methylated DNA (Jeong et
al., 2009; Schlesinger et al., 2007). To examine whether DNMT3B also binds to
nucleosomes in a DNA methylation dependent manner, we expressed Myc-tagged
DNMT3B1 in three DNMT3B-knockout HCT116 cell lines, 3BKO, DKO8 and DKO1,
which possess 92%, 26% and 5% of total genomic DNA methylation respectively (Figure
4.1C). We first tested mRNA and protein expression of the exogenous DNMT3B1 in
these cell lines. Interestingly, while DNMT3B1 mRNA levels were similar in all infected
cell lines, we found dramatically reduced DNMT3B1 protein, similar to DNMT3A, in
severely hypomethylated DKO1 cells in comparison to 3BKO and DKO8 cells (Figure
4.11A, B).
To assess whether the decrease in DNMT3B1 resulted from a reduction in binding
affinity for nucleosomes in hypomethylated cells, we tested its distribution in
mononucleosomal digests fractionated on 300mM NaCl containing sucrose gradients. In
3BKO and DKO8 cells, the exogeneous DNMT3B1 showed strong association with
nucleosomes similar to endogeneous DNMT3A (Figure 4.12A, 4.9). However,
DNMT3B1 weakly associated with nucleosomes in severely hypomethylated DKO1 cells
126
Figure 4.11 mRNA and protein expression analysis of exogenous Myc-tagged
DNMT3B1 in HCT116 derivative cell lines. (A) RT-PCR analysis was performed using
primers for Myc-DNMT3B1 to assess its mRNA levels in infected 3BKO, DKO8 and
DKO1 cells. The results are normalized to GAPDH mRNA levels. Data represents mean
and standard deviation of triplicate PCR reactions from a single experiment,
representative of two biological replicate experiments. (B) Western blot analysis of
nuclear extracts from infected 3BKO, DKO8 and DKO1 cells. Exogenous DNMT3B1
was detected vy Myc antibody.
127
Figure 4.12 Weak nucleosome binding and selective degradation of unbound
DNMT3B in the absence of elevated DNA methylation levels. (A) Nuclei extracted
from infected cells were extensively digested with MNase and mononucleosomes
released from them were resolved by ultracentrifugation on a sucrose density gradient
(5% to 25%) containing 300 mM NaCl. The gradients were fractionated and analyzed as
described previously. (B) DKO1 cells expressing Myc-DNMT3B1 were treated with
cycloheximide (CHX) for different time points. The proteosome inhibitor, MG132 was
added 2 hr prior to CHX treatment. Nuclei extracted from each sample were then
incubated in 500 μl of ice-cold RSB containing 300 mM NaCl, 0.25 M sucrose and
protease inhibitors at 4°C for 5 min. Supernatant and nuclear fractions were separated by
centrifugation at low speed and equivalent protein amounts from each were subjected to
western blot analysis. Data is representative of two biological replicate experiments.
A
B
128
with the bulk of the overexpressed protein sedimenting in nucleosome-free fractions (2-
4), suggesting a dramatic reduction in nucleosome binding affinity upon depletion of
DNA methylation.
To further confirm this phenomenon, we subjected Myc-tagged DNMT3B1
expressing DKO1 cells to CHX treatment and analyzed protein stability of the
nucleosome-bound and -free fractions of DNMT3B1 protein. Consistent with our
previous data on endogenous DNMT3A enzyme, the overexpressed free DNMT3B1
protein underwent rapid degradation compared to the stable nucleosome-bound
DNMT3B1 protein, clearly displaying the instability of the unbound protein (Figure
4.12B). Such degradation was inhibited by treatment with the proteosome inhibitor
MG132, indicating the role of proteosomal pathway in this process. Taken together, these
data show that both DNMT3A/3B require the presence of DNA methylation for tight
binding to nucleosomes and subsequent protein stabilization. Such a mechanism would
enable faithful inheritance of methylated states through proper compartmentalization of
DNMT3A/3B while preventing spurious de novo methylation through selective
degradation of the free enzymes.
Synergistic activity of DNMT3A/3B is mediated by their anchoring to nucleosomes
In ES cells, DNMT3A/3B strongly interact and mutually stimulate each other’s
activity, thus working synergistically to establish genomic DNA methylation patterns
during development (Li et al., 2007). To ascertain whether a similar mechanism is
involved in propagation of DNA methylation in somatic cells, we expressed a Myc-
129
tagged catalytically-inactive DNMT3B1 mutant, having a cysteine to serine alteration
(position 657) which destroys catalytic activity without compromising other functions
(Hsieh, 1999), in DKO8 cells and confirmed its protein expression by immunoblotting
(Figure 4.13A). To determine whether the DNMT3B1 mutant could stimulate DNMT3A
activity, we measured the global DNA methylation level in the mutant expressing cells 8
weeks post-infection. We observed a substantial increase in methylation, demonstrating a
stimulatory effect of DNMT3B on DNMT3A activity, independent of catalytic activity
(Figure 4.13B). Immunoprecipitation experiments showed that the mutant DNMT3B1
strongly interacted with DNMT3A, similar to WT DNMT3B1, suggesting a DNMT3L-
like stimulation mechanism which occurs through physical interaction of the two proteins
(Gowher et al., 2005a) (Figure 4.14A,B). Along with an increase in DNA methylation,
we observed a substantial increase in endogenous DNMT3A protein levels in mutant
DNMT3B1 expressing cells (Figure 4.13A), similar to WT DNMT3B1 expressing cells,
suggesting DNA methylation induced stabilization of DNMT3A protein.
In ES cells, stimulation of DNMT3A/3B activity by DNMT3L partially occurs
through increased association of the enzymes with the substrate DNA, allowing these
slow acting enzymes to efficiently methylate the substrate (Yokochi and Robertson,
2002). To examine whether stimulation of DNMT3A by DNMT3B in somatic cells
occurs through a similar mechanism in a nucleosomal context, we analyzed
mononucleosomal digests from DNMT3B1 mutant expressing cells on 300 mM sucrose
density gradients. All cellular DNMT3A in infected 3BKO and DKO8 cell lines was
found tightly anchored to nucleosomes suggesting that its stimulation by DNMT3B
130
Figure 4.13 DNMT3B catalytically-inactive mutant stimulates DNA methylation by
DNMT3A. (A) Western blot analysis of nuclear extracts from DKO8 cells expressing
wild-type and catalytically-inactive Myc-tagged DNMT3B1 mutant (mut) using specific
antibodies. (B) DNA methylation analysis of infected DKO8 cells using methylation-
sensitive restriction enzymes. Genomic DNA was isolated from infected cells eight
weeks after infection and methylation level was estimated as described in Figure 4.1.
Data is presented as percentage of total genomic methylation present compared to WT
HCT116 methylation levels and is representative of two independent biological replicate
experiments. E/V: Empty Vector; mut: mutant
131
Figure 4.14 DNMT3B1 catalytically-inactive mutant interacts with DNMT3A
similar to wild-type DNMT3B1 and DNMT3L. Western blot analysis was used to
analyze proteins immunoprecipitated using Myc and DNMT3A antibodies in DKO8 cells
expressing (A) Myc-DNMT3B1 or Myc- mut (mutant) DNMT3B1 and (B) Myc-
DNMT3L. IgG was used as a negative control. Antibodies used for immunoprecipitation
(IP) are mentioned at the top and for immunoblotting on the left. Very faint bands of co-
immunoprecipitated DNMT3A were visible in IPs with Myc antibody in (A), possibly
due to very low levels of DNMT3A present in those cell lines. In (B), the upper strong
band in the Myc and CD-8 IP lanes corresponds to the mouse IgG of the IP antibody
while the lower band is for Myc-DNMT3L. Input denotes mononucleosomal digestes
prepared for immunoprecipitation experiments. mut: mutant
A
B
132
occurs through an increased binding to nucleosomes (Figure 4.15). We could not detect
DNMT3A in DKO1 cells due to its extremely low levels in this assay. The DNMT3B1
mutant displayed similar binding affinity for nucleosomes as WT DNMT3B1 suggesting
that catalytic activity has little role in nucleosome binding. Taken together, these data
show that in vivo stimulation of DNMT3A by DNMT3B occurs through an increased
binding to nucleosomes, similar to that observed with DNMT3L, enabling efficient
methylation from these slow acting de novo enzymes and their consequent stabilization
through continued association with such methylated regions.
133
Figure 4.15 DNMT3B mutant stimulates DNA methylation by increasing
DNMT3A’s association with nucleosomes. Mononucleosomes released from nuclei,
extensively digested with MNase, were resolved by ultracentrifugation on a sucrose
density gradient (5% to 25%) containing 300 mM NaCl. The gradients were fractionated
and analyzed as described previously. mut: mutant
134
DISCUSSION
Proper maintenance of epigenetic modifications within specific chromatin
domains is critical for preserving cellular identity. Recently, a common theme for
inheritance of histone marks has emerged where the mark recruits and retains its own
modifying enzyme and triggers renewal by stimulating that enzyme through possible
allosteric activation mechanisms (Collins et al., 2008; Felsenfeld and Groudine, 2003;
Hansen et al., 2008). Our work suggests involvement of a similar mechanism in
maintenance of DNA methylation patterns through DNMT3A/3B in somatic cells.
We and others have previously shown that DNMT3A/3B, but not DNMT1, are
strongly anchored to nucleosomes containing methylated DNA in somatic cells (Jeong et
al., 2009; Schlesinger et al., 2007). Our current data shows that the presence of DNA
methylation is essential for association of DNMT3A/3B with chromatin and also for
maintaining the cellular levels of the DNMT3A/3B enzymes, thereby creating a
homeostatic inheritance system. Such methylation directed binding stimulates DNA
methylation at target loci in vivo ensuring faithful maintenance of methylation patterns, a
phenomenon previously observed in inheritance of the polycomb mark (Margueron et al.,
2009). Since DNMT3A/3B are slow acting enzymes compared to DNMT1 (Yokochi and
Robertson, 2002), stable association with their target methylated regions would be key
for their ability to properly maintain methylated states. We further show that
DNMT3A/3B work synergistically in this maintenance process and DNMT3B stimulates
DNMT3A activity through increased association with nucleosomes, similar to DNMT3L.
Thus, promotion of DNA methylation by selective binding of DNMT3A/3B to
135
nucleosomes containing pre-existing methylation may serve as a critical positive feed-
back loop mechanism essential for faithful propagation of epigenetic states through
somatic cell divisions (Bachman et al., 2001; Chodavarapu et al., 2010; Margueron and
Reinberg, 2010).
Another key finding of our work is the selective degradation of free DNMT3A/3B
proteins which could not bind to chromatin in the absence of pre-existing DNA
methylation. This phenomenon may help explain how somatic cells, which still express
de novo DNMT3A/3B enzymes, prevent aberrant de novo methylation of CpG islands.
Our data suggests that once DNMT3A/3B are recruited to methylated chromatin
domains, pre-existing methylation stabilizes their binding to such regions and enables
faithful propagation of methylated states. However, in absence of DNA methylation, as
would be the case with unmethylated CpG islands, these slow acting enzymes are unable
to stably bind to the chromatin. The resulting free de novo enzymes, which could
potentially cause spurious methylation, are then selectively degraded by the proteosomal
pathway possibly through recognition of an altered conformation in the unbound state
(Figure 4.16). Histone methyltransferases, however, are not regulated in such a manner
and have been found to exist in both free and chromatin-bound forms within nuclei. This
difference can be partially explained by the fact that histone marks are far more dynamic
in nature, actively regulated by the combined action of histone methyltransferases and
demethylases (Shi and Whetstine, 2007), compared to DNA methylation which is still
believed to be a relatively stable mark in differentiated tissues (Law and Jacobsen, 2010).
136
Figure 4.16 Model for selective stabilization of DNMT3A/3B through anchoring to
nucleosomes containing methylated DNA. (A) In somatic cells, DNMT3A remains
bound to nucleosomes containing methylated DNA, enabling proper maintenance of
methylated states in co-operation with DNMT1, the maintenance enzyme, which copies
the methylation pattern during replication by associating with the proliferating cell
nuclear antigen (PCNA). (B) If DNA methylation is lowered by genetic disruption of
DNMT1 and DNMT3B in DKO cells, DNMT3A loses its ability to bind to nucleosomes
which results in destabilization and subsequent degradation of the protein. (C)
Restoration of DNA methylation in such hypomethylated cells, through expression of
DNMT3B (WT or mut) or DNMT3L, increases DNMT3A protein levels by enabling it to
bind to nucleosomes again which results in stabilization of DNMT3A protein. Exogenous
DNMT3B (WT or mut) also binds strongly to nucleosomes in the presence of DNA
methylation and synergistically increases methylation along with DNMT3A while the
excess free DNMT3B protein, which could not anchor to the nucleosomes, gets degraded
by the proteosomal machinery. mut: mutant
137
While initial recruitment of DNMT3A/3B to methylated regions may involve
other proteins, our data strongly suggests that their anchoring to chromatin primarily
depends upon pre-existing DNA methylation. However, in addition to DNA methylation,
certain histone modifications and accessory proteins may also help in selective
compartmentalization of these enzymes. For instance, unmethylated H3K4, recently
shown to bind DNMT3A (Otani et al., 2009), may assist in stable binding to silent
domains. Recruitment of DNMT3A/3B to such domains may involve UHRF1 (Meilinger
et al., 2009). On the other hand, proteins like H2A.Z, CTCF and H3K4me3 etc. which are
antagonistic to DNA methylation (Okitsu and Hsieh, 2007; Zilberman et al., 2008;
Zlatanova and Caiafa, 2009), may occlude binding of DNMT3A/3B to active/poised
regions, thus constraining their activities to silent methylated domains only. Recently,
Witcher and Emerson (Witcher and Emerson, 2009) have shown that loss of such
boundary elements indeed results in aberrant spreading of DNA methylation beyond
methylated domains. Our data suggests that these aberrations may involve DNMT3A/3B
enzymes which remain bound to methylated regions (Jeong et al., 2009). During
tumorigenesis, these de novo enzymes may progressively override the chromatin
boundaries, gradually spreading methylation beyond their specific domains to the entire
region (Graff et al., 1997) resulting in aberrant methylation of genes in clusters – a
common feature of cancer-specific hypermethylation (Coolen et al., 2010; Keshet et al.,
2006). Such a mechanism may also help explain why CpG island loci having pre-existing
methylation in a normal tissue are more susceptible to undergo de novo methylation in
cancer (Huang et al., 1999). Moreover, ectopic de novo methylation, correlated with
138
overexpression of DNMT3A/3B in several types of cancer (Jones and Baylin, 2007), may
also be maintained and propagated through continued association of DNMT3A/3B with
such regions. DNA methylation inhibitors, widely used to inhibit aberrant methylation in
cancer, target DNMTs by trapping them on DNA (Egger et al., 2004). Our data suggests
that destabilization of DNMT3A/3B upon removal of DNA methylation may provide
another mechanism for depletion of these enzymes upon treatment with hypomethylating
drugs. However, future studies are required to further understand these mechanisms,
focusing on factors determining proper compartmentalization of DNMT3A/3B to
methylated regions and mechanisms responsible for selective degradation of the unbound
protein.
In conclusion, our data suggests a model for epigenetic inheritance of DNA
methylation in somatic tissues where pre-existing methylation triggers its renewal by
recruiting and stabilizing DNMT3A/3B on methylated chromatin domains, which then
work synergistically to propagate DNA methylation in co-operation with DNMT1. Such
a mechanism not only ensures faithful maintenance of methylated states but also guards
against aberrant methylation from the de novo DNMT3A/3B enzymes.
139
CHAPTER 5
SUMMARY AND CONCLUSIONS
Epigenetic mechanisms are essential for embryonic development and cellular
differentiation. The layer of epigenetic regulation imposed upon the genome enables cells
to have distinct identities while containing the same genetic information. Epigenetic
modifications including DNA methylation, histone modifcations and nucleosome
positioning, work together to regulate functioning of the genome and enable
establishment of heritable tissue-specific gene expression patterns (Bernstein et al., 2007;
Suzuki and Bird, 2008). Failure in proper inheritance of epigenetic marks can result in
disease states such as cancer (Jones and Baylin, 2007; Sharma et al.). Thus, to preserve
distinct cellular identities after differentiation and to prevent malignant transformation, it
is essential to ensure faithful propagation of epigenetic modifications during somatic cell
divisions. However, the mechanisms responsible for maintenance of epigenetic
modifications are still poorly understood.
In this study, I have examined the inheritance process involved in faithful
propagation of DNA methylation patterns through somatic divisions. In mammals, DNA
methylation patterns are primarily established and propagated by DNA
methyltransferases, the de novo DNMT3A, DNMT3B enzymes and the maintenance
DNMT1 enzyme. At the time I started working on this project in 2007, the model
existing for inheritance of DNA methylation focused mainly on the maintenance role of
140
DNMT1 (Figure 1.4). It was proposed that DNA methylation patterns were faithfully
propagated during replication by DNMT1, which moved along with the replication fork
and copied the methylation pattern from the template strand to the newly synthesized
daughter strand (Holliday and Pugh, 1975; Riggs, 1975). However, some recent studies
had suggested that DNMT1 cannot faithfully maintain DNA methylation by itself and
required co-operative activity of DNMT3A/3B in this maintenance process, particularly
at repetitive elements (Chen et al., 2003; Liang et al., 2002; Riggs and Xiong, 2004). But
the mechanisms guiding such maintenance activity of DNMT3A/3B enzymes remained
unclear.
The primary focus of my work was to elucidate the molecular mechanisms
responsible for guiding DNMT3A/3B activity in somatic cells. I started my work by
initially studying the interaction of DNMT3A/3B proteins with chromatin. This was
important since their different nuclear distribution patterns observed in somatic cells
suggested that DNMT3A/3B and DNMT1 had distinct maintenance mechanisms. The
strong association of DNMT3A/3B, but not DNMT1, with nucleosomes observed in our
experiments is very striking. It suggests that unlike the transient interaction of DNMT1
involved in its replication-associated maintenance activity for the entire genome,
DNMT3A/3B activity is compartmentalized to certain chromatin domains where they
may assist DNMT1 in maintenance of DNA methylation. Such compartmentalization of
DNMT3A/3B proteins involves direct interaction of their conserved protein domains, the
PWWP and the ADD domain, with specific chromatin modifications such as H3K36me3
and unmethylated H3K4 residues, as shown by us and recent work from other groups.
141
However, it is interesting to note that while the individual domains may provide
specificity for interaction of DNMT3A/3B enzymes with certain chromatin regions, the
individual domains alone cannot anchor DNMT3A/3B proteins to the nucleosomes.
Furthermore, synergistic activity of different protein domains together is required to
accomplish tight nucleosome binding that is seen in vivo.
The association of DNMT3A/3B with methylated CpG islands and repetitive
elements may provide insights into the way these de novo enzymes perform their
maintenance role in somatic cells. Recently, a common model for propagation of histone
marks has emerged where the enzyme remains associated with its product enabling its
faithful maintenance through multiple somatic divisions (Collins et al., 2008; Felsenfeld
and Groudine, 2003; Hansen et al., 2008). Tight binding of DNMT3A/3B proteins to
silent methylated chromatin regions indicates existence of a similar inheritance model
involved in propagation of DNA methylation through somatic cell divisions. Through
such a mechanism, DNMT3A/3B may assist DNMT1 in maintaining the methylated
states of CpG dense and repeat regions. Since DNMT1 is a processive enzyme, it may
miss copying DNA methylation of some CpGs in CpG rich regions during the replication
process. DNMT3A/3B, which remain localized to such methylated regions, would then
restore proper methylated states by methylating the sites missed by DNMT1. Such a role
of DNMT3A/3B enzymes may serve as a proof-reading mechanism, critical for ensuring
faithful inheritance of DNA methylation patterns in somatic cells.
Recruitment of DNMT3A/3B by various other chromatin-associated proteins has
been shown to be essential for de novo methylation in ES cells. However, our data shows
142
that these auxiliary proteins are not required for continued association of DNMT3A/3B
with methylated chromatin regions in somatic cells. Unlike DNMT3A/3B, most of the
interacting proteins such as PCNA, HP1, MeCP2, EZH2, HDAC1 and UHRF1, show
weak association with nucleosomes suggesting that while they may recruit DNMT3A/3B
to specific chromatin domains, they are not essential for their strong anchoring to
nucleosomes. The H3K9 methyltransferases, G9a and SUV39h1, display strong
association with nucleosomes similar to DNMT3A/3B. Strong binding of G9a and
SUV39h1 to nucleosomes, possibly through association with their own products, would
enable them to faithfully propagate H3K9 methylation through somatic divisions.
However, tight binding of DNMT3A/3B to chromatin regions containing DNA
methylation in somatic cells does not seem to require these enzymes, suggesting the
existence of some other mechanism that directs maintenance activity of these enzymes.
Whether SUV39h1 can itself mediate targeting of DNMT3A/3B to heterochromatic
regions, independent of HP1alpha, is still an open question.
Since DNMT3A/3B enzymes possess de novo methylation activity, it is extremely
important to ensure tight regulation of their activities in differentiated somatic cells which
already possess well-established methylation patterns. Our data shows that the majority
of DNMT3A/3B proteins within nuclei in somatic cells are bound to nucleosomes
containing methylated DNA, with little free DNMT3A/3B protein existing. DNA
methylation is required for the strong association of DNMT3A/3B proteins with
chromatin indicating a product-mediated targeting mechanism, which may serve in
faithful propagation of methylated states. While the majority of DNMT3A/3B proteins
143
get properly targeted and remain tightly associated with their target chromatin regions,
any free protein not anchored to nucleosomes due to absence of stabilizing DNA
methylation gets selectively degraded by the cellular machinery. Such a mechanism
would not only ensure proper maintenance of DNA methylation at methylated loci but
would also prevent aberrant de novo methylation of unmethylated regions such as CpG
islands which could potentially result in malignant transformation. However,
overexpression of DNMT3s in cancer may result in abundant amounts of free
DNMT3A/3B enzymes which may get targeted aberrantly, leading to the spurious
methylation observed in various types of cancer. Continued association of DNMT3A/3B
enzymes with such aberrantly methylated regions may later enable propagation of
spurious methylation patterns eventually giving rise to malignant phenotype. DNA
methylation inhibitors, which have been suggested to inhibit DNMTs activity by trapping
them onto DNA (Egger et al., 2004), can successfully remove aberrant DNA methylation
from cancer cells, thereby enabling the partial resetting of the abnormal cancer
methylome. Our data suggests that destabilization and degradation of free DNMT3A/3B
upon removal of DNA methylation may serve as another mechanism for inhibition of
DNMT3A/3B by DNA methylation inhibitors. However, removal of DNA methylation
inhibitors results in rebound of DNA methylation at the aberrantly targeted loci (Yoo and
Jones, 2006), possibly reverting cells back to the malignant state. This suggests that the
reset epigenome retains cues that guide re-establishment of the cancer methylome
through DNMTs upon removal of DNA methylation inhibitors. H3K9 methylation is
found concomitant with DNA methylation and remains unaltered upon removal of DNA
144
methylation as suggested by our data showing continued association of G9a with
nucleosomes in the hypomethylated cells. Therefore, it is reasonable to hypothesize that
H3K9 methylation, which remains at the aberrantly targeted loci, may enable re-
establishment of aberrant methylation patterns at such loci upon removal of DNA
methylation inhibitors through recruitment of DNMT3A/3B by G9a or HP1/SUV39h1.
Future studies involving combinatorial epigenetic therapy, targeting both DNA
methylation and histone methylation machineries together, may enable better
understanding of such mechanisms.
In summary, this thesis work has unraveled key mechanisms involved in faithful
inheritance of DNA methylation patterns by DNMT3A/3B in co-operation with DNMT1.
We have shown that DNMT3A/3B, but not DNMT1, strongly associate with the
nucleosomes containing methylated DNA. This tight binding involves simultaneous
interactions of individual conserved DNMT3A/3B protein domains with specific
chromatin marks, together enabling stable association of the enzymes with nucleosomes.
Impaired tethering to nucleosomes, as observed with ΔDNMT3B truncated isoforms,
might lead to aberrant methylation in cancer. DNMT3A/3B do not require known
auxiliary proteins for their association with nucleosomes, but do require an intact
nucleosomal structure. Finally, this work presents a novel principle of enzyme regulation
where the levels of the catalyzing enzymes, DNMT3A/3B, are determined by the levels
of their own enzymatic products i.e. 5-methylcytosine itself. This novel mechanism not
only ensures faithful propagation of DNA methylation patterns but also prevents spurious
de novo methylation. Further understanding of the epigenetic inheritance mechanisms
145
may hold the key to our ability to elucidate the molecular events which lead to
tumorigenesis and to design better treatment strategies for successfully resetting the
cancer epigenome.
146
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168
APPENDIX A
DNA METHYLATION ANALYSIS BY DIGITAL BISULFITE
GENOMIC SEQUENCING AND DIGITAL METHYLIGHT
In addition to the work described in this thesis, I also worked on the development
of a highly-sensitive sequencing technique, ‘Digital Bisulfite Genomic Sequencing’, used
for DNA methylation analysis of tumor and normal tissue samples. I started working on
this project during my rotation in the Jones lab, in collaboration with Dr. Peter Laird’s
group, and continued working on it after joining the lab till the time it was published in
2008 (Weisenberger et al., 2008). I have attached the related manuscript that resulted
from this collaboration.
169
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175
176
177
178
Abstract (if available)
Abstract
Proper propagation of epigenetic information during somatic cell divisions is critical for preserving gene expression patterns and cellular identity. However, the molecular mechanisms responsible for faithful inheritance of epigenetic marks are still poorly understood. In this thesis work, I have studied the inheritance of DNA methylation patterns through somatic divisions, focusing on the role of DNA methyltransferases 3A and 3B in this process.
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Sharma, Shikhar
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Role of DNA methyltransferases 3A and 3B in inheritance of DNA methylation patterns
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Keck School of Medicine
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Doctor of Philosophy
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Genetic, Molecular and Cellular Biology
Publication Date
11/18/2010
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epigenetics
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