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Transcriptional regulation by epigenetic mechanisms
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Transcriptional regulation by epigenetic mechanisms
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Content
TRANSCRIPTIONAL REGULATION BY EPIGENETIC MECHANISMS
by
Joy Chieh-Yu Lin
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(BIOCHEMISTRY AND MOLECULAR BIOLOGY)
August 2007
Copyright 2007 Joy Chieh-Yu Lin
ii
DEDICATION
This thesis is dedicated to my parents who supported me throughout my entire
graduate study and in fond memory of my beloved brother, Wells Lin.
iii
ACKNOWLEDGEMENTS
I would like to express my deepest gratitude to the following people who made the
work in this thesis possible:
Dr. Peter A. Jones, my thesis advisor, for his unconditional support and guidance.
Drs. Louis Dubeau, Peter W. Laird, Robert Stellwagen, and Darryl Shibata for their
insights that challenged my thinking.
Drs. Shinwu Jeong and Gangning Liang for their enlightening scientific discussions,
technical assistance and instructive comments and timely evaluation of this thesis.
Drs. Einav Nili Gal-Yam and Gerda Egger for their scientific insights and advice.
Past and present members of the Jones lab: Ana Aparicio, Jonathan Cheng, Jody
Chuang, Connie Cortez, Sonia Escobar, Merhnaz Fatemi, Jeff Friedman, Tony Li,
Kaylene Lin, Tina Miranda, Anna Van Rietschoten, Yoshimasa Saito, Daiya Takai,
Yvonne C. Tsai, Dan Weisenberger, Erika Wolff and Christine Yoo for providing a
nurturing lab environment and making lab work enjoyable.
Jones lab meeting participants, in particular, Drs. Gerry Coetzee and Judd Rice for
their constructive criticisms and comments.
iv
TABLE OF CONTENTS
DEDICATION ii
ACKNOWLEDGMENTS iii
LIST OF FIGURES vii
ABSTRACT ix
CHAPTER 1: EPIGENETIC MECHANISMS 1
INTRODUCTION 1
DNA METHYLATION 3
The heritability of DNA methylation 3
CpG island 4
DNA methylation in transcriptional repression 4
DNA methylation in germline- and tissue-specific genes 5
Promoter hypermethylation in cancer 6
Hypomethylation in cancer 8
HISTONE MODIFICATIONS 10
Histone modification and transcription 10
Histone methylation 11
Histone acetylation 13
Histone ubiquitylation 14
Roles of histone modifications 14
Histone modifications in cancer 16
NUCLEOSOME OCCUPANCY 17
Regulation of nucleosomes 18
Histone variants 19
Transcription factor binding to nucleosomal DNA 19
Nucleosome occupancy in transcription initiation 21
CpG island is inherently nucleosome-free 21
Nucleosome remodeling and links to cancer 22
DNA METHYLATION, HISTONE MODIFICATIONS AND
NUCLEOSOME REMODELING ARE INTERRELATED 23
Epigenetic therapy 25
CONCLUSIONS 27
OVERVIEW OF THESIS RESEARCH 28
v
CHAPTER 2: DISTINCT LOCALIZATION OF HISTONE H3
ACETYLATION AND H3-K4 METHYLATION TO THE
TRANSCRIPTION START SITES IN THE HUMAN
GENOME 32
INTRODUCTION 32
MATERIALS AND METHODS 34
Cell lines 34
ChAP assay 34
PCR analysis of immunoprecipitated DNA 35
Real-Time PCR amplification of immunoprecipitated DNA 35
RESULTS 36
Validation of ChAP assay by conventional ChIP assay 36
The markers of active chromatin were preferentially located near
transcription start sites of genes 38
Preferential localization of H3-K9/14 acetylation and H3-K4
methylation at the transcription start site in the p16 gene 42
DISCUSSION 45
CHAPTER 3: THE BIDIRECTIONAL MLH1 PROMOTER IS
SILENCED IN BOTH DIRECTIONS BY DNA
METHYLATION 48
INTRODUCTION 48
MATERIALS AND METHODS 50
Cell culture 50
5-aza-2’-deoxycytidine (5-aza-CdR) treatment 50
Plasmid construction 51
Cell transfection and dual luciferase reporter assay 52
Methylation-sensitive single nucleotide primer extension 52
(Ms-SNuPE)
RT-PCR 53
Bisulfite genomic sequencing 54
Rapid amplification of cDNA ends (5’-RACE) 55
RESULTS 55
Organization and expression analysis of the EPM2AIP1 and
MLH1 genes 55
The promoter of EPM2AIP1 and MLH1 is bidirectional 56
Correlation of methylation status of the bidirectional promoter
and the expression patterns of EPM2AIP1 and MLH1 58
Kinetics of EPM2AIP1 and MLH1 mRNA induction and
demethylation of the bidirectional promoter 59
DISCUSSION 64
CHAPTER 4: ROLE OF NUCLEOSOMAL OCCUPANCY IN THE
EPIGENETIC SILENCING OF THE MLH1 CpG ISLAND 67
vi
INTRODUCTION 67
MATERIALS AND METHODS 70
Cell culture 70
5-aza-2’-deoxycytidine (5-aza-CdR) treatment 70
RT-PCR 70
Bisulfite genomic sequencing 71
M.SssI treatment 72
DNaseI footprinting 73
Mononucleosomal DNA preparation and analysis 74
ChIP assays 75
Real-Time PCR amplification of immunoprecipitated DNA 75
RESULTS 75
The MLH1 (1a+1b) promoter has only one highly positioned
nucleosome in expressing cells while it is occupied by
nucleosomes in nonexpressing cells 75
Distinct chromatin structures at the EPM2AIP1/MLH1 promoter
in expressing and nonexpressing cells 78
Nucleosome eviction after 5-aza-CdR treatment 85
DISCUSSION 93
CHAPTER 5: SUMMARY AND CONCLUSIONS 102
REFERENCES 107
vii
LIST OF FIGURES
Figure 1.1 Three major epigenetic mechanisms: DNA methylation,
histone modification and nucleosome occupancy 2
Figure 1.2 Gene silencing by promoter hypermethylation in cancer 7
Figure 1.3 DNA methylation, histone modifications and nucleosome
remodeling are interrelated 26
Figure 2.1 Validation of ChAP assay by conventional ChIP assay 37
Figure 2.2 The markers of active chromatin were preferentially located
near transcription start sites of genes 41
Figure 2.3 Comparison of the levels of acetylated H3 and dimethylated
H3-K4 in the 5’ regions versus the body of genes HTATIP2
and MAN1 43
Figure 2.4 Preferential localization of H3-K9/14 acetylation and H3-K4
methylation at the transcription start site in the p16 gene 44
Figure 3.1 Organization and promoter analysis of the EPM2AIP1 and
MLH1 genes 57
Figure 3.2 Correlation of methylation status of the bidirectional promoter
and the expression patterns of EPM2AIP1 and MLH1 in
various cell lines 60
Figure 3.3 Correlation of methylation status of the bidirectional promoter
and the expression patterns of EPM2AIP1 and MLH1 in two
cell lines by bisulfite sequencing 61
Figure 3.4 Kinetics of EPM2AIP1 and MLH1 mRNA induction and
demethylation of the bidirectional promoter 63
Figure 4.1 Detection of hypersensitive sites by DNaseI digestion and
nucleosomal depletion by mononucleosomal DNA analysis
and ChIP 77
Figure 4.2 Detection of nucleosome depletion by mononucleosomal
DNA analysis 80
viii
Figure 4.3 Distinct chromatin structures at the EPM2AIP1/MLH1
promoter in expressing LD419 and nonexpressing RKO
cells 82
Figure 4.4 Accessibility of native chromatin to M.SssI at the EPM2AIP1/
MLH1 promoter region in expressing LD419 cells 83
Figure 4.5 Gene reactivation after 5-aza-CdR treatment by quantitative
RT-PCR 87
Figure 4.6 Chromatin structural changes upon gene reactivation by
5-aza-CdR 88
Figure 4.7 Methylase-based single promoter analysis (MSPA) of
RKO cells following 5-aza-CdR treatment 89
Figure 4.8 Eviction of nucleosomes by 5-aza-CdR treatment 90
Figure 4.9 Correlation between expression level and % of nucleosome-
depleted molecules 94
Figure 4.10 A simplified model for the epigenetic silencing of the
MLH1 gene 99
ix
ABSTRACT
Transcriptional regulation can be controlled by epigenetic mechanisms
including DNA methylation, histone modifications and nucleosome occupancy. We
first mapped histone H3 acetylation and H3-K4 methylation patterns globally across
the human genome using our novel genome scanning method called ChAP and
showed that they are located preferentially to the transcriptional start sites in the
human genome. The strong concordance of these modifications at the 5’ regions of
genes indicate that these modifications may serve as critical signals for defining the
start of the transcribed domain. We then examined these epigenetic parameters in
detail using the bidirectional CpG island promoter, MLH1 gene as an example.
MLH1, which has two transcriptional start sites and the other gene EPM2AIP1, are
coordinately expressed in cell lines with low promoter methylation and both are
silenced together in cancer cell lines with promoter hypermethylation, suggesting the
promoter acts as a unit. In addition to DNA methylation, silencing of the three
transcription start sites in the bidirectional MLH1 promoter in cancer cells also
involves distinct changes in nucleosomal occupancy. Three nucleosomes, almost
completely absent from the start sites in normal cells, are present on the methylated
and silenced promoter, suggesting that epigenetic silencing may be accomplished by
the stable placement of nucleosomes into previously vacant positions. Indeed,
activation of the promoter by demethylation using 5-aza-2’deoxycytidine involves
nucleosome eviction.
1
CHAPTER 1
EPIGENETIC MECHANISMS
INTRODUCTION
Transcriptional activation is a multi-step process that involves modulation of
chromatin structure such as the accessibility of DNA, interaction of RNA
polymerase and transcription factors, as well as direct regulation of polymerase
during different stages of transcription. In addition to genetic factors, there are other
factors, collectively termed “epigenetics” which is defined as a heritable change in
the pattern of gene expression without any changes in the primary nucleotide
sequence in DNA (Bird, 2002). Biochemical analyses of transcriptional initiation
and elongation have provided information on the molecular interactions underlying
transcriptional control. Additionally, the discovery and subsequent characterization
of protein complexes and enzymatic activities that function as chromatin remodeling
machines, co-activators, co-repressors, or mediators of transcription factors have
unraveled many of the sophisticated regulatory mechanisms that integrate
physiological signals within a cell. The focus of this chapter is to examine in detail
three major epigenetic mechanisms: DNA methylation, histone modifications and
nucleosome occupancy and characterize the changes in each process in the context of
tumorigenesis (Fig. 1.1).
2
Fig. 1.1. Three major epigenetic mechanisms: DNA methylation, histone
modification and nucleosome occupancy. Red circles show CpG dinucleotides. Ac
refers to acetyl groups. Nucleosomes, structural units of chromatin, consist of 146
bp DNA wound around a core of histone proteins.
3
DNA METHYLATION
DNA methylation is catalyzed by DNA methyltransferases (DNMTs) which
add a methyl group to the 5-carbon (C5) of the cytosine ring in DNA to form 5-
methylcytosine (5mC). This covalent modification of DNA, mostly in the context of
the CpG dinucleotide, occurs postreplicatively. Due to the high rate of the
deamination of 5-methylcytosine to form thymine, which is not readily repaired, the
CpGs are depleted progressively over evolutionary time. Thus, the observed
frequency of the CpGs in the human genome is considerably less than predicted
(Takai and Jones, 2002).
The heritability of DNA methylation
The 5-methylation of cytosine is replicated as cytosine. Due to the symmetry
of the CpG dinucleotide pair, each daughter strand has a hemimethylated template
which is subsequently copied on the newly replicated strand, propagating the original
methylation pattern (Holliday and Pugh, 1975; Riggs, 1975). DNMT1 is believed to
be responsible for maintenance DNA methylation while DNMT3A and DNMT3B
mostly catalyze the de novo CpG methylation (Robertson, 2002). Thus, methylation
patterns can be propagated through cell division and be clonally maintained in a
population of cells (Wigler et al., 1981). This is a classic example of epigenetic
inheritance (Bird, 2002).
4
CpG islands
One important feature of DNA methylation patterns is the presence of CpG
islands, areas highly rich in unmethylated CpGs, which are normally unmethylated in
the germline and are often found at the 5’ends of genes or the promoter regions (Bird,
1986; Gardiner-Garden and Frommer, 1987; Takai and Jones, 2002). About half of
human genes are associated with CpG islands in their promoter regions (Antequera
and Bird, 1993). CpG islands are normally kept unmethylated in somatic cells,
allowing CpG-island-containing genes to be transcribed in the presence of the
necessary transcriptional activators while methylated CpG islands are often
associated with transcriptional repression. Examples include transcriptionally silent
genes on the inactive X chromosome in females and some imprinted genes, which
are programmed to allow the expression of only one allele of the gene in normal
tissues (Bird, 2002). Interestingly, methylation was recently found to be
concentrated at gene bodies on the active X chromosome, affecting multiple
neighboring CpGs (Hellman and Chess, 2007).
DNA methylatioin in transcriptional repression
DNA methylation is a characteristic of heterochromatin structure and it
contributes to the formation of a repressive environment along with other
characteristics such as the presence of heterochromatin protein 1, H3-K9 histone
methylation, low histone acetylation and nucleosomes (Pennings et al., 2005). Thus,
DNA methylation may be used to ensure late replication of the bulk of the genome,
5
thereby facilitating transcriptional silencing of these regions via repressive chromatin
formation (Bird and Wolffe, 1999). Hence, DNA methylation may in turn help
maintain regions rich in repetitive elements, viral insertions and transposons in a
transcriptionally inactive configuration (Bestor, 1990).
DNA methylation can function to promote or to prevent binding of regulatory
proteins. One mechanism involves the recruitment of a family of methyl-CpG
binding proteins to the methylated CpGs, repressing transcription through
interactions with histone deacetylases (HDACs), thereby compacting the chromatin
locally (Jones and Laird, 1999). Transcriptional repressors such as Kaiso have been
shown to bind methylated DNA via a distinct type of zinc finger meCpG-binding
domain, resulting in methylation-dependent repression (Prokhortchouk et al., 2001).
Alternatively, methylated CpGs can prevent DNA binding proteins from binding to
their target sites directly by steric hindrance as seen for CTCF whose binding is
abolished by DNA methylation at the target sites in the H19 locus (Hark et al., 2000).
DNA methylation in germline- and tissue-specific genes
Many germline- and tissue-specific genes have been proposed to be
regulated by DNA methylation (Costello and Vertino, 2002). Genomic imprinting is
established by DNA methylation on one of the two parental alleles of a gene to
ensure monoallelic expression of the gene and the inactivation of one of the two X
chromosomes in females is set up in the same fashion (Heard and Disteche, 2006).
On the other hand, loss of imprinting of the insulin-like growth factor-II (IGF2) gene
6
has been observed in Wilms tumor (Ravenel et al., 2001). Similarly, the expression
of the MAGE family of germline-specific genes is also controlled by DNA
methylation (Bodey et al., 2002). Specifically, MAGE-1 expression has been shown
to correlate with the demethylated form of the Ets binding sites of its promoter
(Janssen et al., 1999). DNA methylation has also been postulated as a mechanism
for tissue-specific expression such as the maspin (SERPINB5) gene in which the
promoter is unmethylated in normal cells expressing the gene and is methylated in
those that do not express the gene (Walsh and Bestor, 1999). These findings indicate
that DNA methylation is essential in the establishment and maintenance of imprinted
gene and tissue-specific gene expression.
Promoter hypermethylation in cancer
Methylation of gene promoter CpG islands is associated with abnormal
silencing of transcription (Fig. 1.2) which is one mechanism for inactivation of
tumor suppressor genes (Herman and Baylin, 2003; Jones and Laird, 1999).
According to Knudson’s hypothesis, two hits are required for a complete loss of
function of both copies of a tumor suppressor gene (Jones and Laird, 1999; Knudson,
2001). With respect to genetic changes, one copy can be inactivated by germline or
somatic mutations, leading to the first hit of the two-hit inactivation while the second
hit generally involves loss of heterozygosity. Epigenetically speaking, aberrant
promoter hypermethylation can have similar functional effect as mutations, also
resulting in the loss of one copy, and biallelic methylation can knock out both copies
7
Fig. 1.2. Tumor suppresor gene silencing by promoter hypermethylation in
cancer. 5’ CpG islands are usually unmethylated in normal cells. Cancer cells are
often found to be associated with hypermethylation of CpG islands of tumor
suppressor genes, leading to gene silencing. Open circles represent unmethylated
CpG dinucleotides; solid circles, methylated CpG dinucleotides.
8
of the gene. There are many examples of tumor suppressor genes that have been
shown to be inactivated by promoter hypermethylation including the cell-cycle-
control p16(INK4A) gene in many cancers, the mismatch repair gene MLH1 and the
APC gene in colorectal cancer and the VHL gene in renal cancer (Jones and Laird,
1999).
Hypomethylation in cancer
Global genomic hypomethylation has been documented in many types of
cancer such as prostate metastatic tumors, B-cell chronic lymphocytic leukemia and
hepatocellular carcinomas (Bedford and van Helden, 1987; Lin et al., 2001;
Wahlfors et al., 1992). Hypomethylation could potentially lead to chromosomal
instability, activation of retrotransposons and oncogenes. One possible outcome of
hypomethylation may affect the integrity of chromosomes in cancer. Normally, the
stability of pericentromeric regions of the chromosome is maintained by appropriate
amounts of DNA methylation. For example, structural alterations of the
pericentromeric regions of chromosomes have been associated with global genomic
hypomethylation in breast adenocarcinomas (Narayan et al., 1998). Indeed, tumors
can be induced in mice carrying a hypomorphic DNA methyltransferase 1 (Dnmt1)
allele, indicating a possible cause and effect relationship (Gaudet et al., 2003).
Additionally, regions of the genome that are normally repressed by DNA
methylation could be reactivated due to a weakened repression brought upon by loss
of methylation. This might lead to undesirable expression of potentially harmful
9
elements such as retrotransposons or repetitive elements or imprinted genes such as
those on the inactive X chromosome (Walsh and Bestor, 1999). Long interspersed
nuclear elements (LINEs) are the most abundant retrotransposons in the human
genome (Costello and Plass, 2001; Kazazian and Moran, 1998). Full length LINE-1s
have two alternative open reading frames, one encoding a nucleic acid binding
protein while the other encoding a protein with endonuclease and reverse
transcriptase activities, enabling their migration to other regions via an RNA
intermediate (Costello and Plass, 2001). Thus, they can potentially become
integrated and disrupt essential growth regulating genes (Costello and Plass, 2001).
While many LINEs are methylated and transcriptionally repressed, loss of promoter
methylation and transcriptional activation of LINEs have been associated with some
sporadic cancer types such as the APC gene in colon cancer (Costello and Plass,
2001). Loss of imprinting in the IGF2 gene leading to biallelic expression of the
gene has been associated with higher colorectal cancer risk (Cui et al., 2003).
Hypomethylation can also potentially lead to abnormal activation of
protooncogenes, thereby providing a selective growth advantage for the tumor cells
which then dominate the cell population through clonal expansion (Holliday and
Pugh, 1975). Hypomethylation within the body of genes has been associated with
some primary cancers such as in the cases of the CMYC gene in bladder cancer and
the HRAS gene in non-small cell lung cancer (Del Senno et al., 1989; Vachtenheim
et al., 1994).
10
HISTONE MODIFICATIONS
The nucleosome, the fundamental unit of chromatin, is composed of an
octamer of the four core histones (H3, H4, H2A, H2B) around which DNA is
wrapped. The core histones are largely globular while their N-terminal tails are
unstructured. These highly basic tails interact with the negatively charged DNA
backbone or with other proteins. Specific residues within these histone tails are
subject to diverse post-translational covalent modifications, including acetylation,
methylation, phosphorylation, ubiquitination and so on. These modifications
influence the interaction of the histone tails with DNA or with other proteins that are
required for specific downstream cellular processes such as transcription, replication,
repair and recombination. Another layer of regulation comes from the fact that not
all modifications are required to be on the same histone simultaneously. Thus, the
timing of the formation of a modification may be fine-tuned according to the present
needs of the cell. In addition, modifications are dynamic and they can appear and
disappear within minutes in response to cellular stimuli (Kouzarides, 2007).
Histone modification and transcription
Chromatin is generally classified into two states: euchromatin, which is
associated with active transcription, and heterochromatin, which is associated with
inactive transcription. These distinct types of chromatins are established by distinct
sets of modifications. Modifications that are often associated with euchromatin
11
include H3 and H4 acetylation, di- or trimethylation of H3-K4 whereas H3-K9 and
H3-K27 and also H4-K20 methylation are linked with heterochromatin (Schotta et
al., 2004). However, a given modification can be activating or repressing depending
on a given cue. For example, H3-K9 methylation acts as an inactive mark in the
promoter region and as an active mark in the coding region. This modification has
been shown to be associated with transcription elongation in mammalian chromatin
(Vakoc et al., 2005), showing that the location of a modification has different effects
on transcription. In other words, it can activate transcription at one location while at
another location, it represses the process. This is also exemplified in Set2-mediated
methylation of histone H3-K36 which is normally within the ORF of actively
transcribed genes, but it becomes repressive at the promoter region (Landry et al.,
2003; Strahl et al., 2002).
Histone methylation
Methylation of lysines and arginines can have different forms: mono-, di-, or
trimethyl for lysines and mono- or di- (asymmetric or symmetric) for arginines
(Kouzarides, 2007). Lysine methyltransferases are highly specific, modifying one
single lysine on a particular histone. Lysine methylation can be either activating or
repressing. H3-K4 and H3-K36 methylations are generally activating whereas H3-
K9 and H3-K27 methylations are often associated with repression.
H3-K4 residue is methylated by the Set1 complex in yeast. A gradient of this
modification is established across the entire ORF in which methylation is localized
12
to the 5’ end, dimethylation is enriched in the middle and monomethylation is
enriched at the 3’ end (Guo et al., 2006). One plausible explanation is that H3-K4
monomethylation is maintained at a basal level. Set1, which is associated with
elongating Pol II at the 5’ end, converts monomethylation into dimethylation and
dimethylation into trimethylation. As a result, the stepwise addition of methyl
groups may in turn establish a gradient of methylation across the ORF (Li et al.,
2007). In humans, histone H3-K4 methylation is catalyzed by MLL/WRD5 complex
(Wysocka et al., 2005). The extent of modification, i.e. mono-, di- and
trimethylation can influence the rate of transcription (Li et al., 2007). The role of
H3-K4 methylation has also been implicated in signaling. It can indirectly activate
and repress through the interactions with chromatin-remodeling factors and histone-
modifying complexes via PHD domains, thereby recruiting downstream complexes
(Mellor, 2006b).
H3-K36 methylation is modified by Set2 and is enriched at the 3’ end of
active genes. This modification has been found to be associated with an elongating
RNA Pol II. H3-K36 methylation recruits EAF3 through the chromodomain, which
then recruits Rpd35 deacetylase complex to the coding region, hypoacetylating the
region (Morris et al., 2005). Deacetylation immediately after the passage of RNA
Pol II closes up chromatin, thereby suppressing inappropriate initiation from internal
start sites (Morris et al., 2005).
H3-K9 methylation represses transcription by recruiting methylating enzymes
and HP1 to the promoter region of genes (Lachner et al., 2001). MeCP2 has been
13
shown to be associated with H3-K9 methylation in establishing the repressed state
(Fuks et al., 2003b). As mentioned previously, in addition to its role in
transcriptional repression, H3-K9 methylation has also been associated with
transcription elongation in mammalian chromatin (Vakoc et al., 2005). In addition to
H3-K9 methylation, H4-K20 trimethylation is found to be enriched at pericentric
heterochromatin and it is modified by two SET domain histone methyltransferases,
Suv4-20h1 and Suv4-20h2 which interact with HP1 isoforms (Schotta et al., 2004).
H4-K20 has been implicated in gene silencing (Schotta et al., 2004). H3-K27
methylation has been shown to silence HOX gene expression although the precise
mechanism remains to be deduced (Kouzarides, 2007).
Arginine methylation is mediated by protein arginine methyltransferases
( PRMTs) which are recruited to the promoter region of genes by transcription factor
binding (Lee et al., 2005). Like lysine methylation, arginine methylation can either
activate or silence transcription (Lee et al., 2005).
Histone acetylation
Histone acetylation is catalyzed by a variety of histone acetyltransferases
(HATs) consisting of three main families, GNAT, MYST and CBP/p300
(Kouzarides, 2007). Histone acetylation can occur at multiple lysine residues, and
different combinations of residues involved have distinct effects on transcription.
While most acetylation has been found within the N-terminal tail of histones, it has
been found on K56 within the core domain of H3 (Kouzarides, 2007). Acetylation is
14
unique because not only the location or the site is crucial, but the number of the
residues involved can influence the outcome of the chromatin state as well
(Kurdistani et al., 2004). This modification has been implicated almost solely in
transcription activation. The collaborative effort between histone acetyltransferases
and deacetylases establish the extent and the turnover of acetylation, usually within
minutes (Katan-Khaykovich and Struhl, 2002).
Histone ubiquitylation
This modification is relatively bulky. However, it has been shown to have
minimal effect on nucleosome structure (Jason et al., 2002). Ubiquitylation
of
histone H2B leads to methylation of histone H3 on specific
lysine residues.
Although the sequence of events seems to be
highly conserved among eukaryotes, its
mechanistic function
in transcription remains largely unknown.
Roles of histone modifications
Most obviously, histone modifications other than methylations can alter the
overall net charge of nucleosomes. A change in charge can potentially disrupt and
loosen the interactions between nucleosomal DNA and histone proteins, leading to
the desired chromatin state (Workman and Kingston, 1998). Histone modifications
also act as signals which can be interpreted by other effector proteins, thereby
recruiting the appropriate players to the site of action. Thus, histone modifications
15
can influence chromatin dynamics and structures directly by disrupting the contacts
between DNA and histone proteins or indirectly by recruiting other proteins.
Histone modifications can also affect higher-order chromatin structure by
altering the interactions between histones in neighboring nucleosomes. Acetylation
can loosen chromatin through neutralization of the basic charge of the lysine. Any
change in histone charge will lead to structural changes in chromatin because
internucleosomal contacts are essential for stabilizing higher-order chromatin
structure. H4-K16 acetylation has been shown to prevent the compaction of 30 nm
fibers (Shogren-Knaak et al., 2006). An interesting fact to note is that a single
modification can induce multiple effects since H4-K16 also has been found to
impede ATP-dependent chromatin assembly (Shogren-Knaak et al., 2006).
Nonhistone proteins are recruited to histone marks by binding via specific
domains. Acetylation is recognized by bromodomains, methylation is recognized by
chromo-like and PHD domains and phosphorylation is recognized by a domain
within 14-3-3 proteins (Kouzarides, 2007).
Apart from inter- and intra-nucleosomal interactions, the modifications also
interact with each other at various levels. Such communications or “crosstalk”
between modifications leads to important consequences. Neighboring modifications
can affect the binding of an effector protein of an adjacent modification either
positively or negatively. For example, the binding of the protein could be
strengthened in the presence of a second modification. On the other hand, binding
can also be perturbed by an adjacent modification as it has been shown that H3-S10
16
phosphorylation disrupts HP1 binding to H3-K9 methylation (Dormann et al., 2006).
To add more complexity, the catalytic activity of an enzyme could be greatly
reduced by modification of its cognate substrate such as the methylation of H3-K36
by Set2, which is affected by H3-P38 isomerization (Nelson et al., 2006).
Interestingly, crosstalk can also involve modifications on different histones. The
dynamic nature of histone modifications establishes the stability of a given
chromatin structure.
Histone modifications in cancer
Histone modifications play a pivotal role in establishing the nuclear
framework and in regulating transcription. Thus, global alterations of histone
modification patterns can impact the architecture and integrity of the genome,
disrupting normal gene expression patterns which could lead to cancer. Generally,
histones are maintained in an acetylated configuration in active genes and in
deacetylated state in repressed genes. Thus, gene silencing in cancer may involve
the cooperation between DNA methylation and histone deacetylation in which
histone deacetylases (HDACs) are recruited by methyl cytosine-binding proteins that
bind to methylated DNA (Jones and Laird, 1999). However, histone hypoacetylation
can occur in the absence of DNA hypermethylation as in the transcriptional silencing
of the p21(WAF1) gene (Richon et al., 2000). Histone hypoacetylation is therefore a
prominent mark in cancer cells. In addition, loss of H3-K4 trimethylation and gain
17
of H3-K9 methylation and H3-K27 trimethylation have been associated with
abnormally silenced genes in cancer (Fahrner et al., 2002; McGarvey et al., 2006).
Recently, genome-wide studies examining alterations in histone
modifications have provided more insight into the underlying mechanisms of cancer.
A study of global post-translational modifications of histone H4 in a panel of normal
tissues, cancer cell lines and primary tumors has characterized losses of H4
monoacetylation and trimethylation (Fraga et al., 2005). Also, these changes seemed
to appear early and accumulated during tumorigenesis, suggesting that they may
contribute directly to tumor progression (Fraga et al., 2005). Similarly, another
study has also shown that breast cancer cells are characterized by decreased histone
H4 trimethylation and monoacetylation (Tryndyak et al., 2006). These findings
indicate that global loss of H4 monoacetylation and trimethylation may be another
important feature of cancer cells (Esteller, 2007).
NUCLEOSOME OCCUPANCY
Nucleosomes are the repeating structural units of chromatin, each consisting
of 146 bp of DNA wound around a protein core composed of the histones H2A, H2B,
H3 and H4. The nucleosomes are packed into higher orders of structure to confine
the entire length of the eukaryotic chromosomes into the nucleus. This packaging
inevitably imposes a barrier for many molecular processes including transcription,
replication, recombination, and DNA repair among others that require access to the
18
DNA. Therefore, an understanding of the structure of chromatin and how it is
regulated by various parameters is invaluable.
Regulation of nucleosomes
Nucleosomes have a dynamic nature. In temporal and spatial fashion, histone
and chromatin modifying enzymes and transcription regulatory proteins interact to
allow for the inter-conversion between permissive and non-permissive chromatin
configurations (Mellor, 2006a). Chromatin modifiers or chromatin remodeling
enzymes can alter the chromatin structures by breaking up the contacts between
histones and nucleosomal DNA using ATP hydrolysis as a mechanism to increase
the accessibility of nucleosomal DNA to protein binding such as transcription factors
(Saha et al., 2005; Varga-Weisz and Becker, 2006). As a consequence, DNA can be
unwound from the histone proteins or alternatively, nucleosomes can be mobilized to
different locations (sliding). Both processes can lead to exposure of DNA to the
relevant proteins. It is debatable whether nucleosomes could be mobilized only by
sliding or could actually be displaced completely. However, recent evidence has
suggested that complete displacement does occur in vivo (Li et al., 2007; Workman,
2006). Compared with their counterparts, histones H2A and H2B appear to be more
displaceable than histones H3 and H4 (Li et al., 2007; Workman, 2006).
Histones can be displaced or “evicted” by various means. Displacement or
eviction of histones can be achieved by chromatin remodeling enzymes such as the
Swi/Snf (Guyon et al., 1999; Phelan et al., 1999; Whitehouse et al., 1999). In
19
addition, displacement can be mediated by transcription factor binding, forming an
unstable complex of the protein and the nucleosome which in turn contributes to
nucleosome displacement (Steger et al., 1998). Alternatively, eviction can be
mediated by RNA Pol II through its passage along the DNA (Adelman and Lis,
2002). Following eviction, however, the displaced nucleosomes can still rebind to
the same DNA, thus protein chaperones such as nucleophosmin are required to
prevent the rebinding of the nucleosomes (Okuwaki et al., 2001).
Histone variants
Histone variants are expressed outside the S phase and they are incorporated
into the chromatin in a replication-independent fashion (Kornberg and Lorch, 1999).
Histone variants share many similarities with the canonical histones. The amino acid
sequence between them is conserved with minor differences, entailing similar
functions between them. Like their counterparts, histone variants can also be
mobilized by chromatin remodeling complexes through ATP hydrolysis or by
chaperones (Wu et al., 2005).
Transcription factor binding to nucleosomal DNA
In order for transcription to occur, transcription factors are required to bind to
the appropriate sites on DNA. How do the transcription factors obtain access to the
binding sites which can be buried in chromatin? First of all, there is evidence
showing that nucleosomal DNA may not be completely inaccessible. It has been
20
shown that rapid repair on nucleosomal DNA does occur at both inactive and active
genes (Thoma, 2005). Also, since histone dissociation is too slow for the rapid
repair observed, the unwrapping of nucleosomes seems much more likely to be
responsible for increasing the accessibility of nucleosomal DNA (Thoma, 2005).
On the other hand, transcription factor binding sites have been found at sites
on DNA that are intrinsically free of nucleosomes. This observation is consistent
with recent genome-wide studies which have observed depletion of nucleosomes just
upstream of transcriptional start sites in yeast, flies and humans (Heintzman et al.,
2007; Lee et al., 2004; Mito et al., 2005; Ozsolak et al., 2007). These studies
indicate that lack of nucleosomes is needed to allow access of the transcriptional
machinery to DNA. Remarkably, it has been suggested that nucleosomal positioning
is determined merely by the DNA sequence (Cohanim et al., 2006). Therefore, it
appears that eukaryotic cells deliberately place transcription binding sites in
accessible regions in DNA (Li et al., 2007). In addition, transcription factor binding
can also be regulated by histone modifications which are selective in the sense that
they decide which proteins are to be recruited (Yan and Boyd, 2006).
Nucleosomes pose a strong barrier to RNA Pol II which pauses and
backtracks at intrinsic pause sites on the nucleosomal DNA as it transcribes along
(Kireeva et al., 2005). Transcript cleavage factor TFIIS then reactivates the
backtracked Pol II and promotes transcription through the nucleosome (Kireeva et al.,
2005).
21
Nucleosome occupancy in transcription initiation
To initiate transcription by Pol II, activators bind to the core promoter,
leading to the recruitment of adaptor complexes and transcription factor binding. Pol
II and other transcription factors TFIID, TFIIA and TFIIB form the preinititiation
complex (PIC). In order to fit the PIC, increased histone acetylation is required to
open up the chromatin (Li et al., 2007) which is consistent with the observation that
increased histone acetylation at the promoter region is usually coupled with active
transcription (Ito et al., 2000). In addition, some nucleosomes must be evicted to
accommodate PIC formation (Li et al., 2007). The histone variant H2A.Z (Htz1) is
preferentially enriched at the promoter region because it is relatively easier to evict
(Zhang et al., 2005). Two models of chromatin regulation during transcription
initiation have been proposed. In one model, partial PIC can form without histone
loss and histones are displaced by transcription factor binding while in another
model, histones must be evicted prior to PIC formation (Li et al., 2007).
CpG island is inherently nucleosome-free
CpG islands have an alternative structure to bulk chromatin in normal cells
(Tazi and Bird, 1990). They are characterized by a lack of cytosine methylation, low
levels of histone H1, high levels of histone acetylation and hypersensitivity to
DNaseI which has been equated with nucleosome-free regions (Mucha et al., 2000;
Pfeifer and Riggs, 1991; Tazi and Bird, 1990). Genome wide analysis has shown
that DNaseI sensitive sites are often present in both the expressing and
22
nonexpressing state on many autosomal genes (Mito et al., 2007; Sabo et al., 2004).
In a more specific case, the start site of the maternally imprinted and repressed Igf2
gene is unmethylated and DNaseI hypersensitive to the same extent as the paternal
gene which is strongly expressed (Sasaki et al., 1992a). These and other data suggest
that the “open” or permissive state of CpG islands for potential transcription is
mitotically heritable (Davey et al., 2004). A promoter permissive for transcription
may be therefore kept nucleosome-free, allowing it to become activated by binding
of appropriate transcriptional activators (Li et al., 2007). A central goal of tumor
biology is to understand the mechanisms by which this permissive chromatin
configuration is converted into a state which is permanently repressed and non-
permissive for expression.
Nucleosome remodeling and links to cancer
Recent evidence has suggested a role of the SWI/SNF complex, which
mobilize nucleosomes using ATP hydrolysis in tumorigenesis (Roberts and Orkin,
2004). Many subunits of the SWI/SNF complex have been found to possess intrinsic
tumor suppressor activities such as Snf5, a potent tumor suppressor that is
specifically inactivated in lethal childhood cancers (Sansam and Roberts, 2006).
Abnormal expression of BRG1/BRM, catalytic subunits of the SWI/SNF complex, is
associated with tumor development and increased invasiveness in prostate cancers
(Sun et al., 2007). Also, SWI/SNF has been found to interact with the tumor
suppressor protein RB, thereby mediating RB regulation of the cell cycle (Giacinti
23
and Giordano, 2006). Collectively, these findings suggest that nucleosomal
remodeling may also play a pivotal role in the epigenetic silencing in cancer.
DNA METHYLATION, HISTONE MODIFICATIONS AND NUCLEOSOME
REMODELING ARE INTERRELATED
DNA methylation and histone modifications have been suggested to be
linked by experiments showing that MeCP2, a methyl-CpG binding protein, recruits
chromatin modifying enzymes such as HDACs to sites of DNA methylation (Nan et
al., 1998). The most striking piece of evidence comes from studies in Neurospora in
which mutations in histone lysine 9 methyltransferase result in loss of cytosine
methylation (Tamaru and Selker, 2001). In mouse, the Suv39h-mediated histone
H3-K9 methylation is required to direct Dnmt3b-dependent DNA methylation at
pericentric repeats, showing a functional link between histone modification and
DNA methylation (Lehnertz et al., 2003). In humans, the DNA methyltransferases
are found to be associated with HP1 and the SUV39H1 histone methyltransferase,
chromatin proteins responsible for genes repression and heterochromatin (Fuks et al.,
2003a). Loss of K16 acetylation and K20 trimethylation of histone H4 has been
shown to be associated with global DNA hypomethylation, linking DNA methylation
and histone modifications in determining chromatin structures. A functional link
between the Polycomb group and DNA methyltransferases has been reported in
which DNMTs are recruited directly to regulatory regions of PcG-repressed genes by
24
the H3-K27 methyltransferase EZH2, leading to local DNA methylation (Quina et al.,
2006; Vire et al., 2006). This suggests that heritable patterns of gene silencing may
be established and propagated by a complex interplay between different silencing
pathways.
DNA methylation and nucleosome remodeling have also been shown to be
interrelated (Jones and Baylin, 2007). Methyl-CpG-binding domain-containing
protein, MBD3, has been reported to interact with the nucleosome remodeling NuRD
complex, thereby directing the complex to methylated DNA (Jones and Baylin, 2007;
Zhang et al., 1999). Gene silencing by DNA methylation may be accomplished
through the cooperation between the NuRD complex and MBD3 (Zhang et al., 1999).
Additionally, it has been shown in vivo that MeCP2 recruits Brahma (Brm), a
catalytic component of the SWI/SNF-related chromatin remodeling complex in
transcriptional silencing (Harikrishnan et al., 2005). Furthermore, it has been shown
that the NURF remodeling complex is stabilized through interaction with H3 tails
trimethylated at K4, coupling histone modifications with chromatin remodeling
(Wysocka et al., 2006). These finding reinforce the notion that DNA methylation,
histone modifications and nucleosome remodeling are interrelated (Fig. 1.3) and
these processes may work in concert in gene silencing leading to a heritable
repressive state (Jones and Baylin, 2007).
25
Epigenetic therapy
Epigenetic alterations in cancer are potentially reversible and the reactivation
of epigenetically silenced genes could have a profound antitumor effect (Egger et al.,
2004). In experimental settings, it has been shown that re-expression of silenced
genes can be maintained for years by DNA demethylating agents (Mack, 2006). In
clinical settings, these drugs have been shown to exhibit antitumor activity in
patients with regimen at low doses (Mack, 2006). Indeed, two such drugs, 5-
azacytidine and 5-aza-2’-deoxycytidine have been approved by US food and Drug
Administration (FDA) to treat myelodisplastic syndrome (Jones and Taylor, 1980;
Mack, 2006).
Another promising alternative for cancer treatment is HDAC inhibitors
(Esteller, 2007). One mechanism of action of these drugs lies in their ability to
reactivate tumor-suppressor genes, such as p21(WAF1) through enhanced histone
acetylation around the p21(WAF1) promoter (Ocker and Schneider-Stock, 2007).
HDAC inhibitors have been shown to be well tolerated in patients, and the first drug
of this category, suberoylanilide hydroxamic acid (SAHA), has been approved by the
FDA for the treatment of cutaneous T-cell lymphoma (Esteller, 2007; Mack, 2006).
In addition, DNA methylation seems to dominate histone deacetylation in gene
silencing because there are reports showing that HDAC inhibitors fail to reactivate
cancer genes when they are used alone, but they do demonstrate synergistic effects if
demethylating agents such as 5-aza-CdR are used first (Cameron et al., 1999; Saito et
al., 2006; Suzuki et al., 2002). Indeed, since many epigenetic processes involved in
26
Fig. 1.3. DNA methylation, histone modifications and nucleosome remodeling
are interrelated and thought to all contribute to gene silencing. (Jones P.A., and
Baylin S.B. 2007. The epigenomics of cancer. Cell 128, 683-692)
27
in gene silencing appear to be interrelated, combination therapies may be an effective
therapeutic strategy (Jones and Baylin, 2007).
CONCLUSIONS
The dynamics of chromatin structure are regulated in a very sophisticated
way through multiple layers of epigenetic controls including DNA and histone
modifications, chromatin remodeling, histone variants and nucleosome eviction.
Global alterations of epigenetic modifications can potentially influence the structure
and integrity of the genome and perturb normal patterns of gene expression. Indeed,
numerous studies have indicated that an abnormal pattern of epigenetic
modifications is a hallmark of cancer. Until recently, studies of epigenetic silencing
have predominantly focused on local changes by a single parameter at specific loci.
With increasing realization of the interplay of these epigenetic processes, studies on
the overall pattern of epigenetic marks in cancer cells will undoubtedly shed light on
the mechanisms of epigenetic defects in tumorigenesis. Additionally, a human
epigenome project has been proposed which will enable us to compare epigenetic
profiles in normal cells versus cancer cells. Thus, unbiased, genome-wide
approaches using technologies such as ChIP-on-chip and tiling microarrays have
been invaluable.
Although more and more genome-wide studies are beginning to emerge,
profiles of various epigenetic parameters such as global occupancy of chromatin
28
modifiers are still needed. Epigenetic changes are potentially reversible, making
them desirable targets for antitumor therapeutics. However, one major drawback is
the issue on specificity. Global profiling of a single parameter may potentially
improve the specificity of drug design, such as of an inhibitor that specifically
inhibits a given histone modification at a specific residue.
OVERVIEW OF THESIS RESEARCH
In eukaryotic cells, DNA molecules are packaged into chromatin, forming
nucleoprotein complexes. The nucleosome, basic unit of the chromatin, comprises
146 bp of DNA wrapped around an octamer of core histones: H2A, H2B, H3 and
H4. The modulation of chromatin structure is essential to transcriptional regulation
since it affects the accessibility and the recruitment of other factors to the DNA. In
addition to genetic determinants, transcription can be regulated by epigenetic
mechanisms including DNA methylation, histone modifications and nucleosome
occupancy which cooperate to alter chromatin structure. This thesis describes the
control of transcription by each mechanism in depth and discusses the changes in
each in the context of tumorigenesis.
In Chapter 2, we sought to develop a novel assay capable of scanning global
histone modification patterns in cells. At that time, most studies in the field of
chromatin structure changes focused mainly on examining histone modifications at
individual loci, one locus at a time, and it was believed that histone acetylation and
29
K4 methylation were widespread throughout transcribed units. Thus, we developed
the ChAP assay which combines chromatin immunoprecipitation (ChIP) and
arbitrarily primed polymerase chain reaction (AP-PCR), allowing for the rapid and
unbiased comparison of histone modification patterns. Using this approach, we were
able to show that promoter regions of active genes in humans are highly acetylated at
histone H3 and methylated at H3-K4 while the transcribed regions previously
thought to have the same modifications are actually maintained in a deacetylated H3
and demethylated H3-K4 configuration. This study was one of the pioneering
examples focusing on global histone modifications and many other genome-wide
studies soon followed.
While global studies are important, it is also interesting to examine the
organization of specific loci at high resolution to understand fully the individual
factors involved in transcription regulation. Thus, in later chapters, we focused on
the CpG island bidirectional promoter of MLH1 and EPM2AIP1 genes and
determined how it is regulated by these three mechanisms. Chapter 3, we
investigated the role of DNA methylation for the transcriptional control of this
promoter. First of all, 5’-rapid amplification of cDNA ends (5’-RACE) analysis
showed that the two start sites were located 321 bases from each other, which could
potentially accommodate two nucleosomes. Promoter deletion analysis showed that
the 321-bp intergenic region indeed has bidirectional promoter activity. The genes
are coordinately expressed in cell lines with low promoter methylation and both
genes are silenced together in a series of cancer cell lines in a process associated with
30
dense CpG island methylation. Furthermore, cell lines displaying promoter
hypermethylation were treated with 5-aza-2’-deoxycytidine (5-aza-CdR), leading to
the concurrent reactivation of both genes after drug treatment. Therefore epigenetic
silencing results in the suppression of both genes, showing the promoter acts as a
unit. Our results show that epigenetic silencing can result in the simultaneous
inactivation of two genes.
Chapter 4 describes the role of chromatin structures, mainly by histone
modifications and nucleosome occupancy, in the epigenetic silencing of the MLH1
CpG island. Our results indicate that silencing of the three transcription start sites in
the bidirectional MLH1 promoter CpG island in cancer cells involves distinct
changes in nucleosomal occupancy. Three nucleosomes, almost completely absent
from the start sites in normal cells, are present on the methylated and silenced
promoter, suggesting that epigenetic silencing may be accomplished by the stable
placement of nucleosomes into previously vacant positions. Activation of the
promoter by demethylation using 5-aza-2’-deoxycytidine involves nucleosome
eviction. Thus, epigenetic silencing of tumor suppressor genes may involve heritable
changes in nucleosome occupancy enabled by cytosine methylation.
The studies in this thesis have important biological and clinical implications
because numerous studies have shown that an abnormal pattern of epigenetic
modifications is a hallmark of cancer. Understanding of the global pattern of
epigenetic marks in both normal and cancer cells will indeed help elucidate the
defects caused by epigenetic factors in tumorigenesis. Furthermore, since epigenetic
31
alterations in cancer are potentially reversible, our results indicate that regulatory
proteins involved in nucleosome remodeling may be alternative therapeutic targets in
addition to DNA demethylating agents and HDAC inhibitors.
32
CHAPTER 2
DISTINCT LOCALIZATION OF HISTONE H3 ACETYLATION AND H3-
K4 METHYLATION NEAR THE TRANSCRIPTION START SITES IN THE
HUMAN GENOME
INTRODUCTION
Eukaryotic genomes are organized into chromatin, a higher-order structure
made up of nucleosomes, which consists of 146 bp of DNA wrapped around an
octamer of histone proteins. The tails of histones are subject to a diversity of
posttranslational modifications such acetylation, methylation and phosphorylation,
regulating chromatin structure and function in various cellular processes such as
transcription, replication and repair. Recently, there is increasing evidence from
yeast studies showing that histone modifications play an essential role in
transcription regulation (Hartzog, 2003; Kurdistani and Grunstein, 2003). On the
other hand, the data for the mammalian system is scarce and most studies have
predominantly focused on histone modifications at specific loci. In yeast,
transcription initiation is linked with histone acetylation and methylation while
elongation is associated with not only these modifications but also the recruitment of
elongation factors. Specifically, histone H3-K9/14 acetylation and H3-K4
methylation are associated not only with the promoter regions but also with coding
regions, indicating the role of these modifications in elongation in yeast (Bernstein et
al., 2002; Ng et al., 2003; Santos-Rosa et al., 2002; Xiao et al., 2003).
33
While the same histone modifications have been linked with transcriptional
initiation at the 5’ regions of some human genes (Litt et al., 2001), the general
applicability of the yeast genome is in doubt due to fundamental differences between
genomes. The average gene size in Saccharomyces cerevisiae is 2 kb while it is
about 27 kb in the human genome. However, the yeast genome is much more
compact than the human genome, mainly due to the presence of large introns
including interspersed repeats (Goffeau et al., 1996; Venter et al., 2001). Thus, we
attempted to determine whether the patterns of histone modifications at the start sites
are the same in the human genome, as some studies have indicated significant
differences in H3-K4 methylation patterns at some loci between chicken and
yeast(Myers et al., 2001; Schneider et al., 2004)
To address this question, we developed a novel genome scanning technique
termed ChAP, coupling the chromatin immunoprecipitation (ChIP) assay with the
scanning capabilities of arbitrarily primed PCR (AP-PCR) to determine patterns of
histone modifications genome wide in native chromatin environment in a
nondiscriminatory fashion. This assay screens a random sample across the entire
mammalian genome, differing from the “ChIP-on-chip” assays which are limited by
the sequences applied to the microarray. Our results indicate that methylated lysine
4 (K4) and acetylated K9/14 of histone H3 are distinctly localized to the 5’ regions
of transcriptionally active human genes, showing similar chromatin structures to
those in yeast while the large transcribed regions previously thought to have the
34
same modifications are, in fact, are maintained in a deacetylated conformation in
regions read by elongating polymerase.
MATERIALS AND METHODS
Cell culture
A bladder cancer cell line, T24 and normal LD419 human bladder fibroblasts cell
line were cultured and maintained as described previously (Liang et al., 2002).
ChAP assay
ChAP assay couples the ChIP assay and AP-PCR genomic screening abilities. The
protocol for the ChIP assay was described previously (Nguyen et al., 2001).
Antibodies used were: 15 μg of anti-methyl CpG binding protein 2 (MeCP2), 10 µg
of either anti-acetylated H3-K9/14 (Upstate), anti-dimethylated H3-K4 (Upstate),
anti-trimethylated H3-K4 (Abcam) or normal mouse IgG (Santa Cruz) as nonspecific
antibody control. The AP-PCR assays were performed as described previously
(Liang et al., 2002). Five microliters of each ChIP DNA was amplified by AP-PCR
using a combination of three or four random primers, either GC-rich or GC-poor.
Fragments of interest were isolated, cloned and sequenced and the resulting
nucleotide sequences were compared with the GenBank sequences using BLAST
(www.ncbi.nlm.nih.gov/blast), the University of California at Santa Cruz Human
35
Genome Browser (http://genome.cse.ucsc.edu) and the CpG island searcher program
(www.uscnorris.com/cpgislands) (Takai and Jones, 2003).
PCR analysis of immunoprecipitated DNA
PCR amplification was performed using Expand DNA polymerase (Roche) with 5 µl
of immunoprecipitated DNA, 5 µl of nonspecific antibody negative control (NAC),
or 1 µl of input chromatin (Nguyen et al., 2001). PCR conditions and primers were
described as previously (Liang et al., 2004).
Real-Time PCR amplification of immunoprecipitated DNA
Quantitative PCR was performed using AmpliTaq Gold DNA polymerase (Applied
Biosystems) and TaqMan probes (Biosearch) with DNA Engine Opticon System (MJ
Research). Five µl of immunoprecipitated DNA, 5 µl of nonspecific antibody
negative control (NAC), or 1 µl of input sample (0.2%) were used. Primers were
described as previously (Liang et al., 2004). The following PCR conditions were
used: 95ºC for 10 min, and 45 cycles of 95ºC for 15 s and 59ºC for 1 min. For each
PCR, a set of known amounts of DNA was included as quantitation standard and
immunoprecipitated samples with nonspecific antibody were also included. The
fraction of immunoprecipitated DNA was calculated as (amount of
immunoprecipitated sample with antibody – amount of NAC) / (amount of input
DNA (1%) – amount of NAC).
36
RESULTS
Validation of ChAP assay by conventional ChIP assay
Standard ChIP analyzes were performed on the human bladder cancer cell
line, T24, with antibodies against MeCP2, H3-K9/14 acetylation and H3-K4
dimethylation. Next, immunoprecipitated DNA samples were amplified by AP-PCR
using random GC-rich or non-GC-rich 10-mer primer sets and then the radioactive
products were run on polyacrylamide gels (Fig. 2.1a). The MeCP2 antibody was
used as a marker for inactive chromatin as a control (Nguyen et al., 2001; Nguyen et
al., 2002). Normal mouse IgG was used as a nonspecific antibody negative control
(NAC). Bands present in the lanes from immunoprecipitates did not show up in the
NAC lane but these bands were present in the lane for input DNA which consisted of
a diluted sample of genomic DNA (Fig. 2.1a). Bands present in the
immunoprecipitated lanes and not present in the NAC lane were thought to be
informative. Bands from the active markers (acetylated H3-K9/14 and dimethylated
H3-K4) were coupled but were absent from the inactive marker, MeCP2, indicating
that the active and inactive chromatin markers were mutually exclusive on the AP-
PCR gels (Fig. 2.1a). This result shows the application and validity of the ChAP
assay. Different combinations of three to four random primers (GC-rich or GC-poor)
were used for each AP-PCR and 20 AP-PCRs (10 each for GC-rich and GC-poor
primers) were analyzed and 288 informative bands were obtained. In these
experiments, about 85% of the bands from the lanes for acetylated H3-K9/14 and
37
Fig. 2.1. Validation of ChAP assay by conventional ChIP assay. ChIP assays
were performed on T24 cells with antibodies specific for MeCP2 as a control
(inactive marker), acetylated H3-K9/14 and dimethylated H3-K4. The
immunoprecipiated DNA was amplified by AP-PCR and conventional ChIP-PCR. (a)
An example of ChAP assay gel generated by using radioactively labeled AP-PCR
products. Arrows indicate potentially informative bands (bands present in the
immunoprecipitated lanes and absent from the nonspecific antibody control or NAC
lane), which were excised, cloned and sequenced. Solid arrows indicate bands
precipitated by active markers (acetylated H3-K9/14 and dimethylated H3-K4).
Open arrows show bands precipitated by the inactive marker, MeCP2. (b) Sixteen
randomly selected bands from the ChAP assay were validated with standard ChIP
assay. For all 16 bands, ChIP-PCR of the immunoprecipitated DNA showed the
concordance of H3-K9/14 acetylation and H3-K4 dimethylation. Input DNA
represents total input chromatin (0.2%).
38
dimethylated H3-K4 were coupled while these DNA samples were either weakly or
not precipitated by MeCP2 (data not shown). Since a strong concordance between
H3-K9/14 acetylation and H3-K4 dimethylation was observed, these modifications
were very likely to be present at the same DNA sequences.
Next, 57 of the 288 informative bands were randomly excised, cloned and
sequenced. To verify the concordance of both the active markers on the isolated
chromatin fragments, conventional ChIP analyses were performed on 16 randomly
chosen fragments (Fig. 2.1b). RT-PCR analyses were performed to show that all 16
genes were expressed (data not shown). For all 16 bands, ChIP-PCR of the
immunoprecipitated DNA showed that H3-K9/14 acetylation and H3-K4
dimethylation were coupled, showing the reliability of the ChAP assay (Fig. 2.1b).
The markers of active chromatin were preferentially located near transcription
start sites of genes
Nearly all of these 57 bands immunoprecipitated by the active markers were
associated with genes and about 14% (8 of 57) were in nongene regions, consisting
mainly of repetitive elements (Table 2.1). About 58% (33 of 57) of the fragments
were located within 500 bp of either side of the transcriptional start site of a known
gene or an EST in the database while 28% (16 of 57) were located within the body of
a gene, defined as any region 500 bp or farther downstream of the transcription start
site (Fig. 2.2a and b). GC contents of the primers did not seem to create any bias on
the distribution of the bands since the bands were highly localized to the 5’ region of
39
Table 2.1. Description of 57 fragments identified from the ChAP assay. The 5’
region is defined within 500 bp of a transcriptional start site or the 5’ end of an EST.
Body of genes refers to regions 500 bp or farther downstream of the transcriptional
start site. Other, nongene regions. P, fragments amplified by GC-poor primers; G,
GC-rich primers. CGI, CpG island; N, non-CpG island; Y, CpG island; Bidir.,
bidirectional promoter.
FRAGMENTS Size (bp) CGI
P6-2 330 N
W6-1 158 N
G6-1 250 N
G6-7 451 N
P1-1 275 N
P4-3 413 N
P6-1 315 N
P9-2 440 N
P10-2 280 N
G4-1 238 N
G5-4 220 N
G8-1 437 N
G6-10 750 N
P10-3 271 N
G10-3 612 N
G3-2 398 Y
G10-4 362 Y
G4-3 244 Y
G6-8 165 Y
G7-18 130 Y
G9-1 576 Y
G7-7 389 Y
G7-10 175 Y
G7-15 185 Y
G8-5 245 Y
P1-3 166 Y
G2-4 191 Y
G3-3 188 Y
G6-3 469 Y
G7-3 476 Y
G7-11 207 Y
G8-3 284 Y
G9-2 285 Y
G10-1 193 Y
G10-6 299 Y
G10-9 262 Y
P4-4 268 N
P4-5 245 N
P5-1 254 N
G5-5 159 N
G6-6 294 N
P1-2 350 N
P7-1 264 N
P10-1 273 N
P10-4 289 N
P10-5 187 N
G8-4 202 N
G2-5 381 N
G5-3 227 N
G6-4 434 N
G2-1 176 N
G10-8 299 Y
G2-3 209 Y
G3-1 451 Y
G7-14 317 Y
G7-9 262 Y
IDENTIFICATION
5' of EST (AW673469)
5' of BDNF gene
5' of EST (AL050022 andBI459794) (Bidir.)
5' of BDNF and EST (BC029795)
5' of VPS33B and EST (AK027754) (Bidir.), body of PRC1
5' of HNRU gene
5' of EST(L02867)
5' of DBL (oncogene)
5' of NOTCH2 gene
5' of GEMIN6 gene
5' of EST (AK056022) or body of UXS1
5' of SIPA1 gene
5' of FLJ22402 gene
5' of EST (L33810)
5' of EST (AI949649) and body of BCL10 gene
5' of EST (AF338242)
5' of L02867 gene
5' of TCEB2 gene
5' of MCM2 gene
5' of AXOT and EST (T86526) (Bidir.)
5' of GFI1 or EST ( BM920225)
5' of MGC23445
5' of FLJ20534 gene
5' of PTPRU gene
5' of ESTs (AI633959 and BE669903) (Bidir.)
5' of LAMB3 gene
5' of EST (X13294)
5' of genes ( BC010012)
5' of EST (AB033030)
5' of HTATIP2 gene
5' of EST (BG527537)
5' of CPSF1 gene
5' of DKFZp761B1514 gene
5' of MAN1 gene
5' of ARL7 and body of EST (BG990421)
Body of EST(AW236867) and containing LTR
Body of C11orf17 gene
Body of gene (CAPN11)
Body of EST (AB046797)
Body of BDH gene
Body of EST (BC027456)
Chromosome 10
Chromosome 6
Chr.18 (Sine and Line)
Body of genes (HOXC4, KIAA1536, HOXC6)
Body fo ACPP gene
Body of EST (AB011106) (with LTR)
Body of MAP2K5 gene
body of GRAF gene
Chromsome 5
5' of ESTs (AA158879 and BG830482) (Birdir.)
5' Region Body of Genes and Others
Body of HOXC4 gene
Body of EST (XM_099125)
no match
no match
no match
}
}
{
{
FRAGMENTS Size (bp) CGI
P6-2 330 N
W6-1 158 N
G6-1 250 N
G6-7 451 N
P1-1 275 N
P4-3 413 N
P6-1 315 N
P9-2 440 N
P10-2 280 N
G4-1 238 N
G5-4 220 N
G8-1 437 N
G6-10 750 N
P10-3 271 N
G10-3 612 N
G3-2 398 Y
G10-4 362 Y
G4-3 244 Y
G6-8 165 Y
G7-18 130 Y
G9-1 576 Y
G7-7 389 Y
G7-10 175 Y
G7-15 185 Y
G8-5 245 Y
P1-3 166 Y
G2-4 191 Y
G3-3 188 Y
G6-3 469 Y
G7-3 476 Y
G7-11 207 Y
G8-3 284 Y
G9-2 285 Y
G10-1 193 Y
G10-6 299 Y
G10-9 262 Y
P4-4 268 N
P4-5 245 N
P5-1 254 N
G5-5 159 N
G6-6 294 N
P1-2 350 N
P7-1 264 N
P10-1 273 N
P10-4 289 N
P10-5 187 N
G8-4 202 N
G2-5 381 N
G5-3 227 N
G6-4 434 N
G2-1 176 N
G10-8 299 Y
G2-3 209 Y
G3-1 451 Y
G7-14 317 Y
G7-9 262 Y
IDENTIFICATION
5' of EST (AW673469)
5' of BDNF gene
5' of EST (AL050022 andBI459794) (Bidir.)
5' of BDNF and EST (BC029795)
5' of VPS33B and EST (AK027754) (Bidir.), body of PRC1
5' of HNRU gene
5' of EST(L02867)
5' of DBL (oncogene)
5' of NOTCH2 gene
5' of GEMIN6 gene
5' of EST (AK056022) or body of UXS1
5' of SIPA1 gene
5' of FLJ22402 gene
5' of EST (L33810)
5' of EST (AI949649) and body of BCL10 gene
5' of EST (AF338242)
5' of L02867 gene
5' of TCEB2 gene
5' of MCM2 gene
5' of AXOT and EST (T86526) (Bidir.)
5' of GFI1 or EST ( BM920225)
5' of MGC23445
5' of FLJ20534 gene
5' of PTPRU gene
5' of ESTs (AI633959 and BE669903) (Bidir.)
5' of LAMB3 gene
5' of EST (X13294)
5' of genes ( BC010012)
5' of EST (AB033030)
5' of HTATIP2 gene
5' of EST (BG527537)
5' of CPSF1 gene
5' of DKFZp761B1514 gene
5' of MAN1 gene
5' of ARL7 and body of EST (BG990421)
Body of EST(AW236867) and containing LTR
Body of C11orf17 gene
Body of gene (CAPN11)
Body of EST (AB046797)
Body of BDH gene
Body of EST (BC027456)
Chromosome 10
Chromosome 6
Chr.18 (Sine and Line)
Body of genes (HOXC4, KIAA1536, HOXC6)
Body fo ACPP gene
Body of EST (AB011106) (with LTR)
Body of MAP2K5 gene
body of GRAF gene
Chromsome 5
5' of ESTs (AA158879 and BG830482) (Birdir.)
5' Region Body of Genes and Others
Body of HOXC4 gene
Body of EST (XM_099125)
no match
no match
no match
}
}
{
{
40
genes, in spite of the GC contents. Indeed, 23 of 40 of the fragments for GC-rich
primers (G series) and 10 of 17 of the fragments for GC-poor primers (P series) were
localized to 5’ regions. In addition, 20 of the 33 fragments located in the 5’ regions
of genes were predicted to harbor CpG islands as expected for human 5’ regions
(Takai and Jones, 2003; Venter et al., 2001).
We next compared the distribution of chromatin fragments to the expected
frequency in the human genome using the completed sequences (Venter et al., 2001).
Only 1-2% of the human genome are predicted to be localized at the 5’ regions of
genes assuming an average of two transcription start sites per gene whereas about
58% (33 of 57) of fragments immunoprecipitated by the active markers, H3-K9/14
acetylation and H3-K9 dimethylation, were represented at least 30-fold more
frequently than predicted on a random basis (Fig. 2.2b, P < 0.0001). This shows that
33 of the 57 bands are localized preferentially near the 5’ start site for each gene as
depicted graphically (Fig. 2.2c, gray box).
Since the ChAP assay showed that the active markers were preferentially
localized near transcription start sites, we next performed real-time PCR to
determine the levels of H3-K9/14 acetylation and H3-K4 dimethylation at the
transcription start sites to validate the results. Two examples of genes isolated from
AP-PCR gels, HIV-1 Tat interactive protein 2 gene (HTATIP2) and integral inner
nuclear membrane protein gene (MAN1) were included in the analysis. Also, LD419
normal bladder fibroblasts in addition to T24 bladder cancer cells were analyzed.
H3-K4 trimethylation was also measured because this modification was recently
41
Fig. 2.2. The markers of active chromatin were preferentially located near
transcription start sites of genes. (a) Map of the 5’ region and the body of a gene.
(b) Comparison of the 5’ region and body of genes in bands targeted by acetylated
H3-K9/14 and dimethylated H3-K4 with respect to the expected distribution of these
regions in the entire human genome. Others, nongene regions. (c) Sizes and
locations of the sequences with respect to the 5’ start sites of 38 regions targeted by
acetylated H3-K9/14 and dimethylated H3-K4 antibodies within 1 kb of the nearest
the transcription start site. Each horizontal bar indicates an individual band excised
from the AP-PCR gels, cloned and sequenced. All bands are positioned with respect
to the start sites of their respective transcripts in the GenBank database. Vertical
bars show the number of sequences in 200-bp intervals at the indicated position
relative to the start site. The region within 500 bp of either side of a transcriptional
start site is shown in the gray box.
42
shown to be enriched exclusively in active genes in yeast (Ng et al., 2003; Santos-
Rosa et al., 2002). The results confirmed the presence of all three active markers
near the transcription start sites of both genes and these markers were 6 to 122 times
more enriched at the start sites relative to downstream regions (Fig. 2.3).
Preferential localization of H3-K9/14 acetylation and H3-K4 methylation near
the transcription start site in the p16 gene
To further validate our data, we extended to include a detailed study of
histone modifications in a 7-kb region of the transcriptional unit of the p16(INK4A)
gene in both cell types using antibodies against H3-K9/14 acetylation, H3-K4 di- and
trimethylation (Fig. 2.4). We chose this gene because it was not isolated from the
ChAP analysis and its promoter region was characterized in detail previously
(Nguyen et al., 2001; Nguyen et al., 2002). The endogenous p16(INK4A) promoter
was almost completely methylated in T24 cells and this methylation was associated
with histone deacetylation and H3-K4 hypomethylation (Nguyen et al., 2001;
Nguyen et al., 2002). Quantitative PCR analyzes showed that all three markers were
about 40- to 51-fold enriched around the transcription start site (Fig. 2.4, regions 4-7)
in expressing LD419 cells compared with nonexpressing T24 cells. Also, these
modifications decreased dramatically in regions 3 and 8 in LD419 cells (Fig. 2.4).
These results were consistent with the data from HTATIP2I and MAN1, which were
identified by the ChAP analysis (Fig. 2.3). The distance between regions 3 and 8 is
about 2 kb which is approximately the size of an average yeast gene. This ma
43
Fig. 2.3. Comparison of the levels of acetylated H3 and dimethylated H3-K4 in
the 5’ regions versus the body of genes HTATIP2 (a) and MAN1 (b). ChIP assays
were performed with antibodies against acetylated H3-K9/14 and di- and
trimethylated H3-K4. Data reflect three different real-time PCRs of three
independent ChIP assays. Solid boxes, exons; arrows, predicted transcription start
sites. Regions analyzed by real-time PCR are shown as triangles (filled, 5’ region;
open, body of gene). The numbers reflect the fraction of immunoprecipitated DNA
calculated as a percentage of input DNA. For each PCR, a set of known amounts of
DNA were included as quantitation standard. Error bars show the standard deviation
from three independent experiments.
44
Fig. 2.4. Preferential localization of H3-K9/14 acetylation and H3-K4
methylation near the transcription start site in the p16 gene. (a) Region of the
p16 gene analyzed. The arrow indicates the transcription start site. CpG islands are
shown as gray boxes and repetitive elements are shown as hatched boxes.
Horizontal bars below the map show the regions amplified by real-time PCR. (b)
Levels of histone modifications along the p16 gene (regions 1-10) in expressing
LD419 cells (filled bars) and the nonexpressing T24 cells (open bars). Error bars
reflect the standard deviation from three to five independent experiments.
45
explain why histone H3 acetylation and H3-K4 methylation have not been shown to
be substantially decreased in the compact coding regions of yeast. Also, these data
indicate that these modifications, potentially involved in transcription initiation
and/or the transition between initiation and elongation, may be confined to promoter
regions, similar to yeast (Hartzog, 2003).
DISCUSSION
Our results, obtained by ChAP analysis and confirmed by real-time PCR,
have shown the preferential localization of H3-K9/14 acetylation and H3-K4
methylation near the transcription start sites of human genes (500 bp on either side of
the start sites as we defined here). Although we could not discern whether these
modifications were located upstream or downstream of the transcription start sites
due to the limited resolution of sonication in the ChIP assays. These data suggest
that these modifications may be required for transcription initiation and possibly the
transition stage between initiation and elongation but not throughout regions
transcribed by RNA polymerase II (Pol II). Indeed, we have previously shown that
regions consisting of heavy CpG methylation, MeCP2 binding and nuclease
inaccessibility do not block transcriptional elongation (Nguyen et al., 2001; Nguyen
et al., 2002). On the other hand, H3-K9/14 may become unmodified immediately
after the passage of Pol II as Hos2, a histone deacetylase has been shown to be
46
associated with the coding region of active genes and deacetylated histones H3 and
H4 in yeast (Kurdistani and Grunstein, 2003).
In contrast of our results, H3-K4 methylation has not been shown to be
substantially decreased in the coding regions in yeast. Instead, a gradient of this
modification is established across the entire ORF in which trimethylation is enriched
to the 5’ end, dimethylation is enriched in the middle and monomethylation is
enriched at the 3’ end (Guo et al., 2006). One plausible explanation is that H3-K4
monomethylation is maintained at a basal level. Set 1, which is associated with
elongating Pol II at the 5’ end, converts monomethylation into dimethylation and
subsequently into trimethylation. Thus, the stepwise addition of methyl groups may
in turn create a gradient of methylation across the ORF (Li et al., 2007). In contrast
to acetylation, histone methylation is not known to be readily reversible and it
appears that nucleosomes without di- and trimethylated H3-K4 are traversed by
human Pol II. Indeed, it has been shown that H3-K4 methylation does not influence
transcriptional elongation per se (Pavri et al., 2006).
In humans, the transcribed regions may be maintained in a deacetylated
configuration to prevent inappropriate transcript initiation from cryptic promoters.
In yeast, it has been shown that histones evicted in front of elongating Pol II seem to
be deposited onto DNA soon after the passage of Pol II in order to maintain
chromatin stability during transcription and to prevent cryptic promoters from being
accessible to transcription factors (Li et al., 2007). In addition, since the antibodies
against di- and trimethylated H3-K4 and acetylated H3-K9/14 used bind not only H3
47
but also H3.3, our results also suggest that the 5’ regions of active human genes may
have higher levels of histone H3.3 than transcribed regions. This is consistent with
the observation that histone H3.3 deposition is enriched on active chromatin (Tagami
et al., 2004).
In summary, the ChAP assay provides an unbiased, rapid and robust
comparative approach to assess the pattern and changes of histone marks at a global
scale. Most importantly, compared to standard “chip-on-chip” assays, it is not
limited by the availability of coding sequences that comprise only a small proportion
of the entire human genome. The power of this assay can be strengthened by the
ever increasing availability of specific antibodies to modified histones and other
chromatin-associated proteins. The strong concordance of H3-K9/14 acetylation and
H3-K4 methylation near the transcription start sites of genes indicate that these
modifications may serve as critical signals for defining the start of the transcribed
domain.
48
CHAPTER 3
THE BIDIRECTIONAL MLH1 PROMOTER IS SILENCED IN BOTH
DIRECTIONS BY DNA METHYLATION
INTRODUCTION
Bidirectional gene pairs that are transcribed head-to-head on opposite strands
have been described and comprise about 10% of human genes (Adachi and Lieber,
2002; Lavia et al., 1987; Takai and Jones, 2004; Trinklein et al., 2004). However,
the transcriptional control mechanism of most such genes is not known. In the
present study, we characterized the bidirectional CpG island promoter between the
human genes encoding MLH1 and EPM2AIP1. MLH1 is a major component of the
DNA mismatch repair (MMR) system which corrects replication errors (Modrich
and Lahue, 1996). Germline mutations in MLH1 and other MMR genes have been
shown to confer susceptibility to hereditary nonpolyposis colorectal cancer (HNPCC)
(Peltomaki, 1997). Inactivation of MLH1 is also mediated by epigenetic
mechanisms where promoter hypermethylation of MLH1 has been documented as
causing silencing of expression in sporadic colorectal cancer with microsatellite
instability (Cunningham et al., 1998; Eads et al., 1999; Herman et al., 1998; Kane et
al., 1997; Veigl et al., 1998; Xiong et al., 2001).
The gene EPM2AIP1, which consists of a single exon, is divergently
transcribed antisense to MLH1. EPM2AIP1 has been identified as interacting with
49
laforin protein in Lafora disease by a yeast two-hybrid system (Ianzano et al., 2003).
Lafora’s disease, an autosomal-recessive disorder, is an adolescent progressive
myoclonus epilepsy with less than 0.01% prevalence in humans (Minassian et al.,
2000). It is characterized by prominent myoclonic seizures, diminished mental and
neurologic functions, resulting inevitably in death within 10 years of onset
(Minassian et al., 2000). Laforin, a protein encoded by the gene EPM2A, is
frequently mutated in Lafora disease and it acts as a dual-specificity phosphatase
(Ianzano et al., 2003). However, the function of EPM2AIP1 is unclear. Therefore,
investigating the regulation of the bidirectional promoter may not only expand the
understanding of DNA repair, but also shed light on the underlying mechanism of
Lafora disease.
The start sites of the genes are located within a CpG island of 1.6 kb in length
and we determined that they are 321 bp apart by 5’-RACE. Since promoter
hypermethylation inactivates MLH1 in HNPCC, and given that the two genes share a
common CpG island with transcriptional start sites spaced only 321 bp apart, we
hypothesized that both genes may be regulated through the common CpG island by
epigenetic mechanisms such as DNA methylation. The objective of this study was to
analyze the methylation status on a bidirectional promoter of an important cancer
susceptibility gene and to delineate how the expression of two genes sharing one
CpG island is regulated. We performed dual luciferase reporter assay to show that
the promoter is active in both directions. Also, to determine the relationship between
expression and DNA methylation, we examined methylation levels by methylation-
50
sensitive single nucleotide primer extension (Ms-SNuPE) and bisulfite sequencing
and gene expression by RT-PCR in various cell lines. The results showed a strong
correlation between promoter hypermethylation and the lack of expression of both
genes. In addition, we treated cell lines with promoter hypermethylation with 5-aza-
2’-deoxycytidine (5-aza-CdR) and both genes were reactivated simultaneously after
drug treatment, suggesting similar mRNA induction kinetics. Our findings strongly
suggest that both genes may be coregulated and that the hypermethylation of the
shared CpG island leads to the concurrent silencing of both genes.
MATERIALS AND METHODS
Cell culture
Colorectal cancer cell lines (LoVo, HCT116, LS174T, LS123, HT29, SW620, SW48,
RKO), bladder cancer cell lines (T24, J82), and a cervical cancer cell line (Phelan et
al.) were obtained from the American Type Culture Collection (ATCC). All cells
were cultured as recommended by ATCC. Normal LD419 human bladder
fibroblasts cell line was generated in our laboratory and cultured in McCoy’s 5A
supplemented with 20% FBS.
5-aza-2’-deoxycytidine (5-aza-CdR) treatment
Cells were plated (4 x 10
5
cells/100 mm dish or 2 x 10
6
cells/150 mm dish) and 24 h
later, treated with 10
-5
M 5-aza-CdR (Sigma) for 24 h. The culture was then
51
replenished with fresh medium without the drug for two more days and then nuclei,
DNA and RNA were isolated from the drug-treated culture.
Plasmid construction
Genomic DNA was used for PCR to prepare various lengths of insert covering the
intergenic region. In the MLH1 direction, the amplified fragments were from
nucleotide -324 to -324 to +10, -286 to +10, -261 to +10, -236 to +10 and -211 to
+10 (relative to MLH1 1a): forward (Fwd) primers containing an XhoI site
(underlined): 5’-AACTCGAGCACAAGCCCGGTTCCGGCAT-3’ (-324 to +10),
5’-AACTCGAGATATTTCGTATTCCCCGAG-3’ (-286 to +10), 5’-
AACTCGAGAAAAACGAACCAATAGGAAGAGCG-3’ (-261 to +10), 5’-
AACTCGAGACAGCGATCTCTAACGC-3’ (-236 to +10) and 5’-
AACTCGAGATATCCTTCTAGGTAGCGGGCA-3’ (-211 to +10); the reverse
(Venter et al.) primer containing a HindIII site (underlined), 5’-ATTAAGCTTG
AAACGTCTAGATGCTCAACGG-3’. In the EPM2AIP1 direction, the amplified
fragments were from nucleotide -324 to +10, -324 to -183, -324 to -208, -324 to -232
and -324 to -258 (relative to MLH1 1a): Fwd primers containing an XhoI site
(underlined), 5’-AACTCGAGAAACGTCTAGATGCTCAACGG-3’ (-324 to +10),
5’-AACTCGAGTACTGCCCGCTACCTAGAAGGAT-3’ (-324 to -183), 5’-
AACTCGAGCGCTTGCGCGTTAGAGATCGCT-3’ (-324 to -208), 5’-
AACTCGAGCCGCTCTTCCTATTGGTTCGTTTTTA-3’ (-324 to -232) and 5’-
AACTCGAGAGGAGCTCGGGGAATACGAAAT-3’ (-324 to -258); the Rev
52
primer containing a HindIII site (underlined), 5’-ATTAAGCTTCACAAG
CCCGGTTCCGGCAT-3’. PCR products were double digested with XhoI and
HindIII, gel purified and ligated in the XhoI and HindIII sites of pGL3-Basic
(Promega) using T4 DNA ligase. Plasmids were purified with a miniprep kit
(Promega) and sequenced by an automated DNA sequencer at Laragen (Los
Angeles).
Cell transfection and dual luciferase reporter assay
HeLa cells were seeded (1.2 x 10
4
cells per well, 96-well dish) 24 h before
transfection. Transient transfections were performed by Lipofectamine Plus
(Invitrogen). MLH1 and EPM2AIP1 promoter/firefly luciferase fusion vector (1 μg
per well) and pRL-SV40 vector (10 ng per well, Promega) were cotransfected into
HeLa cells. Cells were incubated until 24 h post-transfection and Luciferase assay
was performed (Dual-Luciferase Reporter Assay System, Promega). The firefly
luciferase activity of each test plasmid was normalized with the Renilla luciferase
activity, which was used as an internal control to correct for the variation of
transfection efficiency.
Methylation-sensitive single nucleotide primer extension (Ms-SNuPE)
Genomic DNA was treated with sodium bisulfite as previously described(Frommer
et al., 1992). The mean cytosine methylation levels of CpG sites were determined by
Ms-SNuPE assay(Nguyen et al., 2001). Briefly, PCR primers were designed to
53
amplify specifically the bisulfite-converted DNA: Fwd: 5’-TAAGGGGAGAG
GAGGAGTT-3’; Rev: 5’-CAATCATCTCTTTAATAACATTAACTAACC3’. The
PCR conditions were 95°C for 2 min, 40 cycles of 95°C for 1 min, 54°C for 50 s,
72°C for 1 min, and 72°C for 10 min. PCR products were gel purified by the
Qiaquick Gel Extraction Kit (Qiagen), and the template was resuspended in 30 μL of
H
2
O. The 5’ to 3’ sequences of Ms-SNuPE primers were as follows: S1:
TTTTAGTAGAGGTATATAAGTT; S2: GCGTATATTTTTTTAGGTAG; S3:
AGG GTGGGGTTGGATGG. Ms-SNuPE reactions were performed as previously
described(Gonzalgo and Jones, 1997). Primer extension conditions were 95°C for 1
min, 49°C (59°C for S3) for 30 s, and 72°C for 20 s. Reactions were terminated by 3
μL of stop solution, denatured at 95°C for 5 min and resolved on a 15% denaturing
polyacrylamide gel (7 M urea). The methylation levels were quantitated by a
Molecular Dynamics PhosphorImager.
RT-PCR
Total RNA was isolated from cells by the Trizol reagent (Invitrogen). Reverse-
transcription was performed with random primers. The 5’ to 3’ sequences of the
primers used in the PCR are: β-ACTIN Fwd: TTTGAGACCTTCAACACCCCAG;
β-ACTIN Rev: TTTCGTGGATGCCACAGGA; MLH1 Fwd: CAGCTAA
TGCTATCAAAGAGATGATTG (1a) and GAGACCTTTTAAGGGTT GTTTGG
(1b); MLH1 Rev: GTTGTAAGAGTAACATGAGCCACATG; EPM2AIP1 Fwd:
TTTGTGGCCTATGAGAACTACC; EPM2AIP1 Rev: GCTCTGATTCAGATTC
54
CGTTAG. The PCR conditions were 95°C for 9 min, 30 cycles (25 cycles for β-
ACTIN) of 95°C for 30 s, 62°C for 1 min, 72°C for 1 min, and 72°C for 10 min.
PCR products were resolved on 2% agarose gels. Quantitative RT-PCR was
performed using an Opticon light cycler with SYBR green I (Sigma), using the
above primers. All values were normalized to β-ACTIN expression ratios and a set
of known amounts of standards was used for quantitation.
Bisulfite genomic sequencing
DNA extracted from cells were treated with bisulfite, PCR amplified using primers
specific to the bisulfite-converted DNA, and then ligated into the pGEM-T easy
vector (Promega). For MLH1 1a, two PCR amplifications of DNA sizes of 536 bp
and 708 bp were produced. The 5’ to 3’ sequences of the primers were: 536 bp, Fwd:
TAAGGGGAGAGGAGGAGTT; Rev: CAATCATCTCTTTAATAACATTAACT
AACC; 708 bp DNA: Fwd: ATAGATTAGGTATAGGGTTTT AT; Rev:
CAATCATCTCTTTAATAACATTAACTAACC. The PCR conditions for the 536
bp DNA were 95°C for 2 min, 40 cycles of 95°C for 1 min, 54°C for 50 s and 72°C
for 1 min, followed by 72°C for 10 min. The PCR conditions for the 708 bp
fragment were 95°C for 2 min, 40 cycles of 95°C for 1 min, 52°C for 50 s and 72°C
for 1 min, followed by 72°C for 10 min. Individual plasmid molecules were
sequenced by an automated DNA sequencer at Laragen (LA) and at the
microchemical core laboratory at the University of Southern California (USC).
55
Rapid amplification of cDNA ends (5’-RACE)
Total RNA was extracted from LoVo cells as described above and the 5’ ends of
mRNA were determined using the RLM-RACE Kit (Ambion) according to the
manufacturer’s instructions. The 5’ to 3’ sequences of gene-specific primers were:
EPM2AIP1 outer: GGCTCCACCACCAAATAACGCT; EPM2AIP1 inner:
GCCGGAAAACTAGAGCCTCGTCGACT; MLH1 outer: CAATATCCAGA
TCTTCTTTCCTGATCCC; MLH1 inner: CCTCCCTCTTTAACAATCACTT
GAATAC. The universal outer and inner primers were provided with the kit. 5’-
RACE reaction products were cloned into a pGEM-T easy vector (Promega) and
sequenced.
RESULTS
Organization of the EPM2AIP1 and MLH1 genes
The bidirectional genes EPM2AIP1 and MLH1 on human chromosome
3p22.1 are located within a CpG island of 1.6 kb in length, which has a GC content
of 57% and a CpG observed/expected ratio of 0.80 (Takai and Jones, 2003), (Fig.
3.1a). The transcriptional start sites of the two genes were suggested to be 470 bp
apart on the GenBank website. However, 5’-RACE analysis using mRNA isolated
from LoVo colon cancer cells (data not shown) established that MLH1 has two
transcriptional start sites (1a and 1b) spaced 309 bp apart which could accommodate
almost two nucleosomes, while EPM2AIP1 initiates transcription at only one site
56
321 bp upstream of the first initiation site 1a of MLH1 (Fig. 3.1a). The start site of
EPM2AIP1 was found to be about 150 bp closer to MLH1 (1a) than indicated by the
GenBank website and, therefore, the divergent transcription start sites are actually
321 bp apart, which could also accommodate approximately two nucleosomes.
The promoter of EPM2AIP1 and MLH1 is bidirectional
To confirm that the promoter of EPM2AIP1 and MLH1 has activity in both
directions, a dual luciferase reporter assay with a series of promoter deletion mutants
was performed (Fig. 3.1b). Different lengths of DNA fragments spanning the
intergenic region were inserted into pGL3-Basic and transiently transfected into
HeLa cells, which express the two endogenous genes. In the MLH1 direction, the
promoter activity decreased by more than 50% (relative to construct #1) when the
region spanning –324 to –286 bp (relative to MLH1 1a) was deleted, indicating this
region might be important for MLH1 transcription. The promoter activity decreased
even more with further deletion to -261 but it increased when the –261 to –236 bp
region was deleted, suggesting the presence of a repressor in this region and further
deletion of the -236 to -211 region again caused additional significant decrease of
promoter activity. It is of note that a putative protein binding site was reported in the
region of –236 to -211 bp by an in vivo methylation footprinting assay (Arita et al.,
2003); however, there is no known transcription factor consensus sequence in this
particular region. Promoter activity in the EPM2AIP1 direction decreased greatly
with the deletion of base pairs +10 to -183 (relative to MLH1 1a) and even more so
57
Figure 3.1. Organization and promoter analysis of the EPM2AIP1 and MLH1
genes. (a) The bidirectional promoter and the CpG island. Horizontal arrows show
the transcriptional start sites as established by 5’-RACE analysis and black boxes
show the respective first exons. Black tick marks indicate CpG dinucleotides.
Horizontal bar underneath the tick marks represents the CpG island. (b) Promoter
analysis. Top is the map of the region analyzed. Promoter DNA fragments inserted
upstream of the firefly luciferase gene in the promoterless pGL3- basic vector
(pGL3-B) are shown on the left as gray (MLH1) and hatched (EPM2AIP1) areas.
The orientation of transcription in each construct is depicted as an arrow. On the
right are luciferase activities of corresponding constructs. The mean activity of
construct #1 was set at 100% and the relative activity of each construct was
calculated. Data shown on the right are the means + SD (n=12).
58
with further deletion from –183 to –208 bp. The promoter activity was comparable
to the promoter-less control with further deletion from -208 to -258 bp. Therefore,
the region spanning +10 to –208 bp was essential for the maximal expression of
EPM2AIP1. In essence, the intergenic region acts as a promoter shared by the two
genes and indeed functions in both directions with MLH1 promoter activity
approximately two times stronger than EPM2AIP1 promoter activity.
Correlation of endogenous methylation status of the bidirectional promoter and
the expression patterns of EPM2AIP1 and MLH1
To determine the relationship between the methylation status of the
bidirectional promoter and the expression of the two genes, we used the quantitative
methylation-sensitive single nucleotide primer extension (Ms-SNuPE) assay (Bender
et al., 1999) and RT-PCR in normal human fibroblast cells and various colon and
bladder cancer cell lines. The mean cytosine methylation levels of several CpG sites
in the MLH1 promoter were determined based on bisulfite treatment followed by
single nucleotide primer extension, eliminating the use of restriction enzymes
common in other methods for methylation analysis. Normal human fibroblast
LD419 cells and cell lines with low methylation levels (LoVo, HCT116, T24, J82,
LS174T, LS123, HT29 and SW620), expressed EPM2AIP1 and both transcripts of
MLH1 (Fig. 3.2). Conversely, none of the transcripts were produced in the colon cell
lines SW48 and RKO, displaying promoter hypermethylation (93% and 95%,
respectively, Fig. 3.2). Treatment of both cell lines (SW48 and RKO) with the
59
demethylating agent 5-aza-CdR for 24 h (Jones and Taylor, 1980), caused a decrease
in the level of CpG methylation with a concordant reactivation of all three transcripts
(Fig. 3.2). These results indicate that both genes may be controlled by the common
CpG island.
The PCR products of bisulfite-treated genomic DNA from two cell lines with
low and high methylation levels were cloned and sequenced to investigate the
detailed patterns of methylation. Almost every CpG sites in the region was
unmethylated in HT29 cells (Fig. 3.3) which was consistent with the methylation
level determined by Ms-SNuPE (Fig. 3.2). For RKO cells, virtually all the CpG sites
were methylated as expected (Fig. 3.3). In all the cell lines, the expression of
EPM2AIP1 and both of the MLH1 transcripts appeared to be coupled, suggesting
coregulation of transcription. Also, both genes were reexpressed after 5-aza-CdR
treatment, indicating that they might be regulated by the common CpG island. Thus,
the endogenous methylation status of the bidirectional promoter correlates quite well
with the expression patterns of both genes, as was also shown by others (Shu et al.,
2006).
Kinetics of EPM2AIP1 and MLH1 mRNA induction and demethylation of the
bidirectional promoter
We next asked whether reactivation of both genes by 5-aza-CdR occurred
with the same kinetics. The hypermethylated RKO cells were exposed to 10 μM 5-
aza-CdR for 24 h and the expression patterns of both genes monitored by RT-PCR
60
Figure 3.2. Correlation of methylation status of the bidirectional promoter and
the expression patterns of EPM2AIP1 and MLH1 in various cell lines. The
average methylation levels of 3 CpGs in the EPM2AIP1/MLH1 promoter region
were analyzed by Ms-SNuPE and expressed as % methylation as quantitated by a
PhosphorImager. The expression of both genes was determined by RT-PCR in
normal LD419 human bladder fibroblast cell line and various human cancer cell
lines as shown in the bottom. RT-PCR products were run on an agarose gel and
stained with EtBr. β-ACTIN expression served as a control for the input amount of
cDNA. RT- served as negative control for the intronless gene, EPM2AIP1.
61
Figure 3.3. Correlation of methylation status of the bidirectional promoter and
the expression patterns of EPM2AIP1 and MLH1 in expressing HT29 and
nonexpressing RKO cells by bisulfite sequencing. Expression status of each cell
line was determined by RT-PCR as shown in Fig. 3.2. Solid circles, methylated CpG
dinucleotides; open circles, unmethylated CpG dinucleotides. Horizontal rows of
circles indicate individual molecules that were sequenced after PCR amplification
and cloning of bisulfite-treated DNA.
62
over 93 days. EPM2AIP1 and the first transcript (1a) of MLH1 (immediately
adjacent to EPM2AIP1) were induced slightly 24 h after drug treatment while the
expression of the other MLH1 transcript showed a delayed induction at 48 hrs after
drug treatment (Fig. 3.4a). From this point on, the expression levels of EPM2AIP1
and MLH1 transcripts increased substantially and showed the strongest expression
level 72 hrs after treatment. Expression was sustained up to day 51 and began to
decrease thereafter (Fig. 3.4a), as we have found for other reactivated genes such as
p16 (Bender et al., 1998). Once again, the expression of both genes showed
concordant behavior.
Bisulfite-genomic sequencing was next performed to follow the kinetics of
demethylation and remethylation of the promoter region. The untreated RKO cells
show extensive methylation (Fig. 3.4b) which was consistent with the previous result
(Fig. 3.3b), and substantial demethylation was observed 72 h after treatment. A
decrease of about 50 % of the number of methylated CpG sites was observed which
corresponded to the theoretical maximal demethylation of 50% after one cycle of
replication in the presence of the aza nucleotide. Furthermore, extensive
remethylation was observed 72 days after treatment; however, the remethylation did
not return to the original methylation level as seen in untreated RKO cells (Fig. 3.4b).
Also, 72 h after treatment, about 50% (6 of 12 DNA strands) of the molecules were
substantially demethylated and 72 days after treatment, about 10% (1 in 9 DNA
strands) of the molecules remained substantially unmethylated. If it is assured that
complete demethylation is necessary for expression, then the mRNA level at day 72
63
Figure 3.4. Kinetics of EPM2AIP1 and MLH1 mRNA induction and
demethylation of the bidirectional promoter. RKO cells were treated with 5-aza-
CdR (10 μM) for 24 h. DNA and RNA were isolated at different time points after
drug treatment. (a) Expression patterns of EPM2AIP1 and MLH1 genes were
analyzed by RT-PCR. (b) The methylation status of individual molecules of the
region at different times after drug addition was analyzed by bisulfite sequencing.
Solid circles, methylated CpG dinucleotides; open circles, unmethylated CpG
dinucleotides. Horizontal rows of circles indicate individual molecules that were
sequenced after PCR amplification and cloning of bisulfite-treated DNA.
64
would be expected to be about 1/5 or 20% of that at 72 hr after treatment, which was
as observed (Fig. 3.4a).
DISCUSSION
The transcriptional mechanism of bidirectional CpG island promoters has not
been thoroughly investigated. In this study, we focused on the bidirectional
promoter of MLH1 and EPM2AIP1 and tested the hypothesis that both genes might
be concordantly silenced by DNA methylation through their common CpG island.
First of all, deletion mutant analysis showed that the 321-bp intergenic region was
indeed sufficient for the expression of both genes. The methylation status of the
CpG island between the pair of genes was correlated with gene expression in various
cell lines to identify the effects of methylation on transcription of the pair. Promoter
hypermethylation correlated with lack of expression of both genes. In addition, cell
lines with promoter hypermethylation were treated with 5-aza-CdR, which led to the
concurrent reactivation of both genes. Due to a gradual resilencing effect, the
expression of both genes decreased coordinately after growth in the absence of 5-
aza-CdR, again suggesting a common silencing mechanism. The fact that two genes
can be simultaneously silenced by de novo methylation suggests that caution should
be exercised when ascribing a phenotype to a given gene such as MLH1.
In regard to MLH1, two transcripts were always produced, with one encoding
a full-length protein of 756 amino acids and another encoding a truncated form
65
missing the N-terminal ATPase domain. The N-terminal ATPase domain has been
suggested to confer the activation of the DNA repair machinery (Guarne et al., 2004)
and thus, missing this domain will render the repair inactive. The C-terminal domain
of MLH1 is thought to mediate dimerization with hPM1 and hPMS2 (Guerrette et al.,
1999). The outcome of producing a truncated protein leading possibily to inactive
repair, along with a full-length protein is not apparent. MMR functions to repair
subtypes of DNA lesions including base-base mismatches and insertion-deletion
loops (IDLs) (Kondo et al., 2001). MLH1 can form different heterodimers with
hPMS, hPMS2 and hMLH3 and all three proteins compete for the binding to MLH1
(Kondo et al., 2001). Each dimerization appears to repair a specific subtype of
replication errors. Therefore, it is possible that a truncated form of MLH1 protein
with only the intact C-terminal domain (dimerization domain) is produced to bind to
those dimerization partners not needed to ensure the activation of only one MMR
pathway specific for the subset of DNA errors. In other words, the truncated form
missing the N-terminal ATPase domain is adapted to compete out unnecessary
dimerizations, preventing the activation of other nonspecific MMR pathways.
Interaction assays involving different MLH1 heterodimers may clarify these
possibilities.
In short, we have redefined the start sites of MLH1, shown the presence of a
truncated transcript, and demonstrated that both MLH1 transcripts are coordinately
expressed with the EPM2AIP1 gene. The data also show that CpG island cytosine
methylation, which is often found in colon cancer (Cunningham et al., 1998; Eads et
66
al., 1999; Herman et al., 1998; Kane et al., 1997; Veigl et al., 1998; Xiong et al.,
2001), is associated with the silencing of two genes at one time. This means that
caution should be applied before ascribing a phenotype to the inactivation of one
gene in such a pair.
67
CHAPTER 4
ROLE OF NUCLEOSOMAL OCCUPANCY IN THE EPIGENETIC
SILENCING OF THE MLH1 CpG ISLAND
INTRODUCTION
Abnormal promoter CpG island methylation is the most well-established
mechanism for epigenetic silencing of tumor suppressor genes in tumorigenesis
(Baylin and Ohm, 2006; Herman and Baylin, 2003; Jones and Laird, 1999). Much
effort has been spent on investigating other epigenetic changes such as covalent
histone modifications responsible for aberrant gene regulation in cancer. Inactive
hypermethylated promoters are associated with a closed or repressive chromatin
configuration, characterized by deacetylated histones and a variety of inactive
histone marks. For examples, in colon and breast cancer cells, the silencing of
MLH1 (Fahrner et al., 2002; McGarvey et al., 2006) and RASSF1A (Strunnikova et
al., 2005) is associated with deacetylated histone H3 and increased H3-K9
methylation. Furthermore, it has been shown that H3-K9 methylation can occur
prior to DNA methylation in the silencing of p16(INK4a) (Bachman et al., 2003),
highlighting the importance of histone modifications. Since previous studies have
focused primarily on promoter hypermethylation and covalent histone modifications,
the objective of this study was to explore additional determinants, such as the role of
nucleosomal occupancy, in gene silencing in cancer since this has been implicated in
transcriptional control in yeast, flies and humans (Bernstein et al., 2004; Heintzman
68
et al., 2007; Lee et al., 2004; Lieb and Clarke, 2005; Mito et al., 2005; Ozsolak et al.,
2007; Yuan et al., 2005).
We focused on MLH1, a key player in the DNA mismatch repair system
(Modrich and Lahue, 1996), which is frequently silenced by promoter
hypermethylation in various cancers (Herman et al., 1998; Kanaya et al., 2003; Kane
et al., 1997; Murata et al., 2002; Xiong et al., 2001). MLH1, which we determined to
have two transcripts 1a and 1b, is a member of a class of bidirectional gene pairs that
are transcribed head-to-head on opposite strands, comprising about 10% of human
genes (Adachi and Lieber, 2002; Lavia et al., 1987; Takai and Jones, 2004; Trinklein
et al., 2004). The divergently transcribed gene to MLH1 is EPM2AIP1, whose
function is unclear, but which has been identified as interacting with laforin protein
in Lafora disease, a disorder characterized by prominent myoclonic seizures,
diminished neurologic functions and death occurring within 10 years of onset
(Ianzano et al., 2003).
Until recently, methods for examining nucleosome positioning have relied on
low-resolution nuclease digestions which reveal only the average state of a given
promoter on all the molecules in a cell population. However, we were able to
analyze the scenario on individual molecules using our new high-resolution
methylase-based single promoter analysis assay (MSPA), providing insights into the
dynamics of chromatin remodeling (Fatemi et al., 2005; Gal-Yam et al., 2006). Our
results, confirmed by traditional methods such as DNaseI digestion,
mononucleosomal DNA analysis and chromatin immunoprecipitation, show
69
remarkable nucleosome depletion just upstream of each start site on the active
promoter while the inactive promoter is associated with nucleosome occupancy in a
mitotically heritable fashion. These findings are consistent with emerging evidence,
indicating that gene activation is accompanied by nucleosome disassembly at the
transcriptional start site (Boeger et al., 2003; Mellor, 2005; Reinke and Horz, 2003;
Workman, 2006). Specific examples show a decrease in histone content on the
inducible IL2 gene upon T-cell activation (Chen et al., 2005) and the constitutive
depletion of nucleosomes from the stress-inducible GRP78 promoter, regardless of
its induction state (Gal-Yam et al., 2006). Depletion of nucleosomes just upstream
of transcriptional start site has been observed in genome-wide screens of yeast, flies
and humans (Bernstein et al., 2004; Heintzman et al., 2007; Lee et al., 2004; Mito et
al., 2005; Ozsolak et al., 2007; Yuan et al., 2005). These data mostly pertain to
immediate changes associated with remodeling in response to an activating signal.
Epigenetic silencing, however, is characterized by a mitotically heritable state which
is not generally reversible unless DNA methylation or histone marks are reversed.
An early study by Tazi et al (Tazi and Bird, 1990), compared the properties
of active CpG island chromatin and bulk inactive chromatin globally, showing that
the former is H1-deficient, H3 and H4-hyperacetylated and nucleosome-depleted.
However, the transition between the two states was not examined. Here we show
that nucleosomes are re-depleted from promoter molecules upon gene reactivation by
demethylation using 5-aza-2’-deoxycytidine (5-aza-CdR), suggesting that epigenetic
silencing may be associated with nucleosome reassembly at previously deplted
70
regions. Thus, our results provide strong evidence that epigenetic silencing of a
tumor suppressor gene involves not only DNA methylation and histone
modifications, but also heritable nucleosome occupancy changes in epigenetic
silencing of a tumor suppressor gene.
MATERIALS AND METHODS
Cell culture
A colorectal cancer cell line, RKO was obtained from the American Type Culture
Collection (ATCC) and cultured as recommended by ATCC. Normal LD419 human
bladder fibroblasts cell line was generated in our laboratory and cultured in McCoy’s
5A supplemented with 20% FBS.
5-aza-2’-deoxycytidine (5-aza-CdR) treatment
Cells were plated (4 x 10
5
cells/100 mm dish or 2 x 10
6
cells/150 mm dish) and 24 h
later, treated with 10
-5
M 5-aza-CdR (Sigma) for 24 h. The culture was then
replenished with fresh medium without the drug for two more days and then nuclei,
DNA and RNA were isolated from the drug-treated culture.
RT-PCR
Total RNA was isolated from cells by the Trizol reagent (Invitrogen). Reverse-
transcription was performed with random primers. The 5’ to 3’ sequences of the
71
primers used in the PCR are: β-ACTIN Fwd: TTTGAGACCTTCAACACCCCAG;
β-ACTIN Rev: TTTCGTGGATGCCACAGGA; MLH1 Fwd: CAGCTAAT
GCTATCAAAGAGATGATTG (1a) and GAGACCTTTTAAGGGTTGTTTGG
(1b); MLH1 Rev: GTTGTAAGAGTAACATGAGCCACATG; EPM2AIP1 Fwd:
TTTGTGGCCTATGAGAACTACC; EPM2AIP1 Rev: GCTCTGATTCAGATTCC
GTTAG. Quantitative RT-PCR was performed using an Opticon light cycler with
SYBR green I (Sigma), using the above primers. All values were normalized to β-
ACTIN expression ratios and a set of known amounts of standards was used for
quantitation.
Bisulfite genomic sequencing
DNA extracted from cells were treated with bisulfite, PCR amplified using primers
specific to the bisulfite-converted DNA, and then ligated into the pGEM-T easy
vector (Promega). For MLH1 1a, three PCR amplifications of DNA sizes of 250,
536 and 708 bp were produced. The 5’ to 3’ sequences of the primers were: 250 bp
DNA: Fwd: ATAGATTAGGTATAGGGTTTTAT; Rev: AACTTATATACC
TCTACTAAAATA AT; 536 bp DNA: Fwd: TAAGGGGAGAGGAGGAGTT;
Rev: CAATCATCTCTTTAATAACATTAA CTAACC; 708 bp DNA: Fwd:
ATAGATTAGGTATAGGGTTTTAT; Rev: CAATCATCTCTTTAATAACATTA
ACTAACC. The PCR conditions for the 250 bp DNA were 95°C for 2 min, 40
cycles of 95°C for 1 min, 51°C for 50 s and 72°C for 1 min, followed by 72°C for 10
min. The PCR conditions for the 536 bp DNA were 95°C for 2 min, 40 cycles of
72
95°C for 1 min, 54°C for 50 s and 72°C for 1 min, followed by 72°C for 10 min.
The PCR conditions for the 708 bp fragment were 95°C for 2 min, 40 cycles of 95°C
for 1 min, 52°C for 50 s and 72°C for 1 min, followed by 72°C for 10 min. For
MLH1 1b, the 5’ to 3’ sequences of the primers were: Fwd: TGA
GGTGATTGGTTGAAGGTAT; Rev: ACTTACACTCCAAACAACCCTTA. In
addition, the selective amplification in the MSPA assay on RKO cells was performed
with the reverse primer only annealing to the unmethylated molecules to eliminate
the amplification of extensively methylated clones which could not be analyzed (see
the main text for rationale). To increase the specificity of the primer to unmethylated
DNA, a mismatch was incorporated at the 3’ end of the reverse primer which will
destabilize unspecific duplex formation. For selective amplification, the 5’ to 3’
sequences of the primers were: Fwd: TGGGTTGGAAAATTAGAGTTTTGTT;
Rev: ACCAAATAACCCCTACCACAAATA. The PCR conditions were 95°C for
2 min, 44 cycles of 95°C for 1 min, 55°C for 50 s and 72°C for 1 min, followed by
72°C for 10 min. Two independent bisulfite-sequencing reactions were done to
avoid introducing a bias in the analyses. Individual plasmid molecules were
sequenced by an automated DNA sequencer at Laragen (LA) and at the
microchemical core laboratory at the University of Southern California (USC).
M.SssI treatment
Nuclei preparation and M.SssI reactions were performed as described
previously(Fatemi et al., 2005). Briefly, purified genomic DNA and freshly
73
extracted nuclei were treated with M.SssI for 15 min at 37°C. Reactions were
stopped by the addition of an equal volume of stop solution (20 nM Tris-HCl, pH 7.9,
600 mM NaCl, 1% SDS, 10 mM EDTA, 400 μg/mL proteinase K), incubated at
55°C for overnight and DNA was purified by phenol/chloroform extraction and
ethanol precipitation.
DNaseI footprinting
Nuclei from LD419 cells were extracted as described above. Nuclei were
resuspended in RSB buffer (10 mM Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2)
plus 0.25 M sucrose, incubated with various concentrations of DNaseI (Worthington)
at 37°C for 15 min to obtain suitable range of digestion of genomic DNA revealed by
EtBr staining. Digested DNA, purified as described above, was cut again by DraI
restriction enzyme to be resolved on a 1.5% agarose gel, which was then Southern
blotted. Blot was hybridized with a 163 bp PCR amplified DraI-probe spanning -29
to 134 bp relative to MLH1 1b. The probe was labeled with [α-
32
P]dCTP using the
High Prime (Roche) and hybridized by ExpressHyb Hybridization Solution (BD
Biosciences). The 5’ to 3’ PCR primer sequences for the probe amplification were:
Fwd: GTTCCCTGACGTGCCAGTCA; Rev: AAATTAAGTGGCTTCCTTACTTA
GTTAACG. The blot was visualized by Molecular Dynamics PhosphorImager.
74
Mononucleosomal DNA preparation and analysis
Detailed protocols were published previously(Gal-Yam et al., 2006). Quantitative
PCR was performed using AmpliTaq Gold DNA polymerase (Applied Biosystems)
and TaqMan probes (Biosearch) with DNA Engine Opticon System (MJ Research,
Cambridge, MA). The 5’ to 3’ sequences for the primers and probes used were:
region A: Fwd: CTACGATGAGGCGGCGAC; Rev: GACCCAGCGTT
ATTTGGTGGT; probe: CCAGGCACAGGGC; region B: Fwd: CCTCAGC
AGAGGCACACAAG; Rev: AATACGAAATATCCATCCAATAGGA; probe:
CGGTTCCGGCATCT; region C: Fwd: CAGCAACCCACAGAGTTGAGAA; Rev:
GCGGCAGCTATTGATTGGA; probe: TTGACTGGCATTCAA; region D: Fwd:
CGTAAGCTACAGCTGAAGGAAGAA; Rev: CGTCTAGATGCTCAACGGAAG
TG; probe: CACGAGGCACTGAGG; region E: Fwd: GGCTGGACGA
GACAGTGGTG; Rev: CAGTTCTCAATCATCTCTTTGATAGCA; probe: AAG
TTATCCAGCGGCCAG; region F: Fwd: TCACTCAATGGCGCGGAC; Rev:
TGGCACGTCAGGGAACCC; probe: CAGCGCATGCCCACA; region G: Fwd:
TGTAAGTGGAGGAATATACGTAGTGTTGT; Rev: TCGCATGTTCTGCATA
CATAATTTT; probe: AATGGTACCGTTAACTAAG; region H: Fwd: CTGTGT
TACTGTTTTCTTGCTTTTCAT; Rev: GAATTGTGCCTTTGGATGTGAAC;
probe: CATTCCAGAAATCATC. PCR conditions were used: 95°C for 9 min, and
45 cycles of 95°C for 30 s, 62°C for 1 min and
72°C for 1 min, followed by 72°C for
10 min.
75
ChIP assays
ChIP analyses were performed as described previously(Nguyen et al., 2001).
Antibodies used were: 10 μg of either anti-Histone H3 (Abcam) or anti-acetylated
Histone H3 (Upstate) and 1 μg of rabbit IgG (Upstate) as nonspecific antibody
control.
Real-Time PCR amplification of immunoprecipitated DNA
Quantitative PCR was performed using AmpliTaq Gold DNA polymerase (Applied
Biosystems) and TaqMan probes (Biosearch) with DNA Engine Opticon System (MJ
Research, Cambridge, MA). Same primers and probe sequences from the
mononucleosomal DNA analysis were used. The following PCR conditions were
used: 95°C for 10 min, and 45 cycles of 95°C for 15 s and
59°C for 1 min. For each
PCR, a set of known amounts of DNA were included as quantitation standard and
immunoprecipitated samples with nonspecific antibody were also included. The
fraction of immunoprecipitated DNA was calculated as percentage of input DNA.
RESULTS
The MLH1 (1a+1b) promoter has only one highly positioned nucleosome in
expressing cells while it is occupied by nucleosomes in nonexpressing cells
As nucleosomal positioning and histone tail modifications play essential roles
in transcriptional regulations (Bernstein and Allis, 2005; Rice and Allis, 2001), we
76
investigated chromatin structure at all three transcriptional start sites. We first used
DNaseI hypersensitivity assay, a conventional approach to map chromatin structure.
Nuclei from LD419 and RKO cells were treated with increasing concentrations of
DNaseI to obtain suitable levels of digestion (Fig. 4.1a).
Examination of the Southern blot images revealed a region of
hypersensitivity in the lanes of minimally digested LD419 chromatin samples (Fig.
4.1b, lanes 2-4), which was mapped to the promoter region shared by the two genes,
between MLH1 1a and EPM2AIP1. Interestingly, another hypersensitive region
corresponding to the region upstream of MLH1 1b was also detected at higher
enzyme concentrations (Fig. 4.1b, lane 4). These patterns were not detected in
control naked DNA, suggesting the lack of sequence preference of DNaseI for that
region. At the genomic level, RKO nuclei were digested to a greater extent with
higher concentrations of DNaseI, compared to LD419 (Fig. 4.1a, lanes 11-14).
However, even under such digestion conditions, at the specific loci examined, the
1399 bp DraI fragment persisted up to lane 14 (Fig.4.1b) without showing any
discrete DNaseI hypersensitivity. The region examined in RKO cells is therefore
more compact and inaccessible to DNaseI. Thus, the promoter regions just upstream
of all three start sites are highly accessible in expressing LD419 cells, but not in
nonexpressing RKO cells. The hypersensitive sites in LD419 cells may suggest the
lack of nucleosomes, as a result of changes in chromatin structure correlated with
transcriptional activity of the genes. To confirm the nucleosome depletion suggested
by DNaseI footprinting, we examined the presence of nucleosomes in the promoter
77
Figure 4.1. Detection of hypersensitive sites by DNaseI digestion and
nucleosomal depletion by mononucleosomal DNA analysis and ChIP. Genomic
naked DNA and nuclei from LD419 and RKO cells were treated with increasing
concentrations of DNaseI and DNA from each sample was then purified and treated
with DraI, and the digestion products analyzed by Southern blotting. Naked DNA,
as a control, was used to confirm the lack of sequence specificity of the enzyme. (a)
Digested samples prior to DraI treatment were resolved by gel electrophoresis as
shown. (b) Southern blot analysis revealed DNaseI hypersensitivity in the
EPM2AIP1/MLH1 promoter. On the left, drawn to scale, the 1399 bp DNA
fragment generated by DraI digestion, transcription start sites (arrows) and probe
fragment (black box) are indicated. Numbers show the fragment size in bp.
DNaseI:
LD419 RKO
Naked
DNA
a
1 2 3 4 5 6 7 8 9 10 1112131415
DNaseI:
EPM2AIP1 MLH1 1a
LD419 RKO
Naked
DNA
b
1399
321 309 636
DraI
DraI
MLH1 1b
1 2 3 4 5 6 7 8 9 101112131415
78
regions at the mononucleosomal level by two other traditional methods. Nuclei from
LD419 and RKO cells were partially digested with MNase, yielding molecules of
various nucleosomal repeats. Nucleosomes from the digested nuclei were then run
on a sucrose gradient to isolate fractions enriched in mononucleosomes and the DNA,
derived from the mononucleosomes, was analyzed by quantitative PCR using 8
primer sets positioned across the region (Fig. 4.2). Consistent with the DNaseI
results, in expressing LD419 cells, the region between the EPM2AIP1 and MLH1 1a
transcription initiation sites, TISs (Fig. 4.2, B-D) and the region of about 150 bp just
upstream of MLH1 1b TIS (Fig. 4.2, F) showed virtually no signals after PCR while
the other regions analyzed showed relatively higher enrichment. Interestingly, there
is one, and only one, highly positioned nucleosome in the MLH1 promoter (1a+1b),
as shown by the dramatically increased enrichment just downstream of MLH1 1a TIS
(Fig. 4.2, E), corresponding to the region between two hypersensitive sites (Fig. 4.1b,
lane 4). On the other hand, no analogous nucleosomal depletion was observed in
nonexpressing RKO cells (Fig. 4.2). Taken together, these results argue that the
promoter regions just upstream of all three start sites are depleted of nucleosomes in
expressing LD419 cells, but are occupied by nucleosomes in nonexpressing RKO
cells.
Distinct chromatin structures at the EPM2AIP1/MLH1 promoter
Next, we used the chromatin immunoprecipitation assay (ChIP) to validate
the above results and to compare the chromatin structures at the bidirectional
79
promoter in expressing and nonexpressing cells. Since we wanted as high a
resolution map as possible over the relatively small region between the start sites,
particular attention was paid to ensuring that the fragments generated by chromatin
sonication were of the order of 200 bp.
Consistent with the DNaseI and the mononucleosomal DNA analyses, H3 occupancy
was clearly lower in the region between the EPM2AIP1/MLH1 1a start site in
expressing LD419 cells and much higher in nonexpressing RKO cells (Fig. 4.3, B-D).
A zone of nucleosomal depletion or “dip” in histone H3 occupancy was seen in the
EPM2AIP1/MLH1 1a promoter region of the expressing LD419 cells (Fig. 4.3, B-D).
H3 occupancy in nonexpressing RKO was high in all regions and no prominent dip
was present in the EPM2AIP1/MLH1 1a promoter region (Fig. 4.3, B-D). However,
ChIP analysis did not detect the depletion of the nucleosome upstream of the start
site of MLH1 1b in LD419 (Fig. 4.3, F), probably as a result of the limited resolution
of sonication. As expected, overall histone H3 acetylation was high in expressing
LD419 as compared to RKO (Fig. 4.3). Remarkably, the H3 acetylation pattern in
LD419 recapitulated the patterns seen in the H3 and mononucleosomal DNA
analyses, showing a prominent dip in the EPM2AIP1/MLH1 1a promoter and a lesser
dip in the region just upstream of MLH1 1b (Fig. 4.3, LD419, regions B-D and F).
While many studies have shown that promoter regions are often associated with
increased acetylation in active promoters without specific references to H3
occupancy, our results indicate that the increased acetylation is actually coupled with
H3 enrichment in regions surrounding the promoter (Fig. 4.2, A and E). H3
80
Figure 4.2. Detection of nucleosome depletion by mononucleosomal DNA
analysis Nuclei from LD419 and RKO were digested partially with MNase and the
reaction mixture was run on a sucrose gradient to isolate mononucleosomal DNA.
Enrichment of mononucleosomal DNA was analyzed by real-time PCR using
primers specific for eight regions (A to H) shown as black rectangles as depicted on
top. The fraction of mononucleosomal DNA was calculated as a percentage of input
genomic DNA.
RKO
LD419
Mononucleosomes
150 bp
EPM2AIP1 MLH1 1a MLH1 1b
0
8
0
8
a b c d e f g h a b c d e f g h
%(IP/Input) %(IP/Input)
A B C D E F G H A B C D E F G H
A B C D E F G H A B C D E F G H
81
acetylation was minimal or almost none existent, in all regions in nonexpressing
RKO cells (Fig. 4.3, A-H).
To study the region in greater detail, we used a new high-resolution methylase-based
single promoter analysis assay (MSPA) we have developed, which allows for the
footprinting of individual DNA molecules (Fatemi et al., 2005; Gal-Yam et al., 2006;
Kladde and Simpson, 1996). Nuclei from expressing, unmethylated LD419 cells
were treated with M.SssI, a cytosine-C5 CpG DNA methyltransferase, which
methylates all accessible CpG sites. While this enzyme methylates every CpG site in
purified DNA, it is unable to methylate CpG sites which are found within a
nucleosome, or bound by tight-binding transcription factors (Fatemi et al., 2005;
Gal-Yam et al., 2006; Kladde and Simpson, 1996). Following M.SssI treatment,
bisulfite conversion of extracted DNA and PCR amplification of the promoter region,
single PCR products were cloned and sequenced to display the accessibilities of
individual DNA strands to the methylase.
As expected, the regions analyzed were virtually unmethylated in untreated
LD419 nuclei (Fig. 4.4a). Control experiments showed that naked DNA extracted
from the same cells was almost completely methylated by M.SssI treatment at the
region under the same experimental conditions used for nuclei, with no preferential
sites of methylation (Fig. 4.4b). Analysis of the M.SssI treated nuclei revealed that
the 321 bp region between the two start sites (EPM2AIP1 and MLH1 1a) and also a
region of about 150 bp upstream of MLH1 1b were largely accessible to M.SssI as
shown by the extensive acquired methylation, indicating the absence of nucleosomes
82
Figure 4.3. Distinct chromatin structures at the EPM2AIP1/MLH1 promoter in
expressing LD419 and nonexpressing RKO cells. ChIP analysis performed with
antibodies against histone H3 and acetylated histone H3. Immunoprecipitated DNA
was analyzed by real-time PCR as described in Fig. 4.2. The fraction of
immunoprecipitated DNA was calculated as a percentage of input DNA. The data
reflect the average of duplicate experiments and three independent chromatin
preparations were done to confirm the reproducibility of results.
0
3
%(IP/Input)
RKO
0
5
%(IP/Input)
H3 ChIP
LD419
%(IP/Input)
RKO
Ac-H3 ChIP
0
8
16
%(IP/Input)
LD419
0
8
16
150 bp
EPM2AIP1 MLH1 1a MLH1 1b
A B C D E F G H
150 bp
EPM2AIP1 MLH1 1a MLH1 1b
A B C D E F G H A B C D E F G H
A B C D E F G H A B C D E F G H
A B C D E F G H A B C D E F G H
83
Figure 4.4. Accessibility of native chromatin to M.SssI at the EPM2AIP1/MLH1
promoter region in expressing LD419 cells. Nuclei were extracted from expressing
unmethylated LD419 cells and then were treated with M.SssI for 15 min followed by
bisulfite genomic sequencing. Two independent bisulfite-sequencing reactions were
done to avoid introducing a bias in the analyses. Four PCR products of different
sizes as indicated by the blue dotted lines, were included in the analysis. (a)
Untreated nuclei. (b) Naked DNA treated with M.SssI. (c) Nuclei treated with
M.SssI. Horizontal lines with circles indicate individual molecules that were
sequenced after PCR amplification and cloning of bisulfite-treated DNA. Solid
circles, methylated CpG dinucleotides; open circles, unmethylated CpG
dinucleotides. Pink bars indicate inaccessible areas or patches to M.SssI, suggesting
presence of nucleosomes. Patches are defined as at least two consecutively
unmethylated sites flanked on each side by least two consecutively methylated CpG
sites (Fatemi et al., 2005). Blue rectangles show the putative protein binding regions.
84
in these regions (Fig. 4.4c). No nucleosome footprint was seen in 28 out of 30
promoter replicas analyzed in the 321 bp region and none was seen in all of 27
molecules analyzed in the 150 bp upstream of MLH1 1b (Fig. 4.4c). Furthermore,
there were two clear patches of substantial inaccessibility, suggesting the presence of
nucleosomes downstream of MLH1 1a and EPM2AIP1 start sites (Fig. 4.4c, pink).
Patches are defined as at least two consecutively unmethylated sites flanked on each
side by least two consecutively methylated CpG sites (Fatemi et al., 2005). Patches
at both the 5’ and 3’ ends were also considered as nucleosomes on the basis of the
mononucleosome and ChIP experiments (Fig. 4.2 and 4.3). In fact, a well-defined
nucleosome was precisely positioned in 93% (25/27) of molecules examined in the
region downstream of MLH1 1a (Fig. 4.4c, pink).
Two inaccessible patches were also detected whose sizes were too small to
qualify as nucleosomes. One patch of about 30 bp was observed in the region
between EPM2AIP1 and MLH1 1a, corresponding to the region (-236 to -211) of a
putative protein binding site mentioned earlier (Fig. 4.4c, blue). Another patch of
about 20 bp was also detected in the region between MLH1 1a and 1b, which may be
an unidentified protein binding site important for MLH1 1b start site function (Fig.
4.4c, blue). In addition, there was another region of substantial inaccessibility
downstream of MLH1 1b which could indicate another nucleosome by comparison
with Fig. 4.1-4.3. Together these results suggest that in expressing LD419 cells, the
region between EPM2AIP1 and MLH1 1a is devoid of nucleosomes while there is
precisely one nucleosome positioned downstream of MLH1 1a, followed by another
85
nucleosome-free region upstream of the MLH1 1b start site. The nucleosome-free
regions correspond to the hypersensitive regions seen in the DNaseI assay (Fig. 4.1b,
lane 4) and also to precisely the same regions with the lowest enrichment seen in the
mononucleosomal DNA analysis (Fig. 4.3, regions B-D and F).
Nucleosome eviction after 5-aza-CdR treatment
Since the promoter in nonexpressing RKO cells was fully occupied by nucleosomes,
we were interested to see the chromatin structural changes accompanying gene
activation. First we quantified the level of expression of the silenced genes after 5-
aza-CdR treatment using quantitative RT-PCR. As found earlier (Fig. 3.2) 24 h 5-
aza-CdR treatment caused a concordant reactivation of all three transcripts as shown
by the dramatic increase in the expression level 72 h after addition of the drug (Fig.
4.5). However, the drug-induced expression level at 72 h was not as high as the level
in untreated LD419 cells, which was 4 times higher for all three transcripts (Fig. 4.5).
Expression for all three transcripts was sustained even 44 days after drug treatment
although with a 3 to 7 fold decrease in level of expression (Fig. 4.5). The decreased
expression as a function of time after treatment has been observed with other genes
activated by 5-aza-CdR treatment and is due to a gradual resilencing effect (Bender
et al., 1999).
We then analyzed the chromatin changes upon gene activation by ChIP
analysis of RKO cells 72 h and 44 days after 5-aza-CdR treatment. The H3
acetylation was minimal in untreated RKO cells and increased substantially by
86
several fold in every region examined 72h after drug treatment (Fig. 4.6, A-H),
which has been reported by others (Fahrner et al., 2002). Even 44 days after drug
treatment, the H3 acetylation was still maintained at levels intermediate between the
levels seen in untreated and 72 h in virtually all regions. Strikingly, after gene
activation, acetylation levels recapitulated the patterns seen in the mononucleosomal
DNA analysis showing a dip in the EPM2AIP1/MLH1 1a promoter and lesser dip
upstream of the MLH1 1b at both 72h and 44 days. Qualitatively, the changes in H3
acetylation levels correlate well with the changes in gene expression levels.
As ChIP analysis only yields an average result for the total cell population with
limited resolution, we applied the MSPA assay to detect chromatin structural
changes in RKO cells following gene activation upon 5-aza-CdR treatment. Almost
all of the CpG sites in the promoter were endogenously methylated in untreated RKO
cells (Fig. 4.7a). Two populations of molecules were visible 72 h after treatment
with 5-aza-CdR (Fig. 4.8a). In accordance with the expected mechanism of action of
the drug (Egger et al., 2004), about 62% (15/24) of the molecules were methylation-
free or had sporadic residual methylation (Fig. 4.8a). These molecules, not present
in the parent nuclei before 5-aza-CdR treatment, could potentially serve as substrates
for M.SssI, thus allowing us to search for new footprints using the MSPA mapping
technology.
On the other hand, 38% (9/24) of the sequenced molecules were still
substantially methylated, and were not suitable substrates for M.SssI because these
molecules would remain substantially methylated following subsequent M.SssI
87
Figure 4.5. Gene reactivation after 5-aza-CdR treatment by quantitative RT-
PCR. RKO cells were treated with 5-aza-CdR and then harvested 72 h and 44 days
after drug addition for RT-PCR and ChIP analyzes. Expression levels were
normalized with β-ACTIN which served as a control for the input cDNA. A minus-
RT control served as negative control for the intronless gene, EPM2AIP1 (data not
shown). The data represent the average of duplicate PCRs and two independent
cDNA preparations were done to confirm the reproducibility of results.
MLH1 1a
0
0.005
0.01
0.015
0.07
LD419 Unt 72 h 44 d
MLH1 1a/ -ACTIN
MLH1 1b
0
0.002
0.004
0.006
0.02
LD419 Unt 72 h 44 d
EPM2AIP1/β β β β-ACTIN
EPM2AIP1
0
0.01
0.02
0.03
0.1
LD419 Unt 72 h 44 d
MLH1 1a
0
0.005
0.01
0.015
0.07
LD419 Unt 72 h 44 d
MLH1 1a/β β β β-ACTIN
MLH1 1b
0
0.002
0.004
0.006
0.02
LD419 Unt 72 h 44 d
EPM2AIP1/ -ACTIN
EPM2AIP1
0
0.01
0.02
0.03
0.1
LD419 Unt 72 h 44 d
MLH1 1b/β β β β -ACTIN
MLH1 1a
0
0.005
0.01
0.015
0.07
LD419 Unt 72 h 44 d
MLH1 1a/ -ACTIN
MLH1 1b
0
0.002
0.004
0.006
0.02
LD419 Unt 72 h 44 d
EPM2AIP1/β β β β-ACTIN
EPM2AIP1
0
0.01
0.02
0.03
0.1
LD419 Unt 72 h 44 d
MLH1 1a
0
0.005
0.01
0.015
0.07
LD419 Unt 72 h 44 d
MLH1 1a/β β β β-ACTIN
MLH1 1b
0
0.002
0.004
0.006
0.02
LD419 Unt 72 h 44 d
EPM2AIP1/ -ACTIN
EPM2AIP1
0
0.01
0.02
0.03
0.1
LD419 Unt 72 h 44 d
MLH1 1b/β β β β -ACTIN MLH1 1b/β β β β -ACTIN
88
Figure 4.6. Chromatin structural changes upon gene reactivation by 5-aza-CdR.
RKO cells were treated with 5-aza-CdR and then harvested 72 h and 44 days after
drug addition for ChIP analysis with antibody against acetylated H3. At 72 h,
acetylation increased dramatically in all the regions analyzed. Immunoprecipitated
DNA was analyzed by real-time PCR as described (Fig. 4.4). The fraction of
immunoprecipitated DNA was calculated as a percentage of input DNA. The data
reflect the average of duplicate experiments and two independent chromatin
preparations were done to confirm the reproducibility of results.
150 bp
EPM2AIP1 MLH1 1a MLH1 1b
A B C D E F G H A B C D E F G H
0
0.5
1
1.5
2
%(IP/Input)
Unt
72 h
44 days
Unt
72 h
44 days
A B C D E F G H
RKO+5-aza-CdR: Ac-H3 ChIP
89
Figure 4.7. Methylase-based single promoter analysis (MSPA) of RKO cells
following 5-aza-CdR treatment. (a) Nuclei from untreated RKO cells were
extracted and subjected to bisulfite genomic sequencing. (b) RKO cells were treated
with 5-aza-CdR for 24 h and harvested 72 h after drug treatment started. Naked
DNA from drug-treated cells was extracted and then subjected to M.SssI treatment,
followed by bisulfite genomic sequencing. The promoter was almost fully
methylated after M.SssI treatment. Horizontal lines with circles indicate individual
molecules that were sequenced after PCR amplification and cloning of bisulfite-
treated DNA. Solid circles, methylated CpG dinucleotides; open circles,
unmethylated CpG dinucleotides
Untreated RKO nuclei
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp
RKO+5-aza-CdR: Naked DNA+M.SssI
b
a
90
Figure 4.8. Eviction of nucleosomes by 5-aza-CdR treatment. RKO cells were
treated with 5-aza-CdR for 24 h and harvested 72 h after drug treatment started.
Both DNA and nuclei from drug-treated RKO cells were extracted and then subject
to M.SssI treatment, followed by bisulfite genomic sequencing. (a, d) Demethylation
of the promoter in RKO cells 72 h (a) and 44 days (d) after addition of 5-aza-CdR.
(b,c,e,f) PCR analyzes were done using selective primers that only anneal to
unmethylated molecules to filter out extensively methylated molecules in (a) and (d)
which were not suitable for M.SssI treatment. (b, e) Selective amplifications were
done on the samples in (a) and (d) to verify the specificity of the primers. (c, f)
M.SssI treatments on nuclei from drug-treated RKO cells 72h (c) and 44 days (f)
after drug addition. Please refer to Fig. 4.4 for descriptions of molecules. Pink bars
indicate inaccessible areas or patches to M.SssI, suggesting presence of nucleosomes.
Patches are defined as at least two consecutively unmethylated sites flanked on each
side by at least two consecutively methylated CpG sites. The DNA molecules with
nucleosome depletion in the promoter region were boxed in blue.
Demethylated
by 5-aza-CdR
RKO+5-aza-CdR (72h)
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp
Demethylated
by 5-aza-CdR
RKO+5-aza-CdR (72h)
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp
Nucleosomes
present
Nucleosomes
absent
Nuclei+M.SssI (44 days) Nuclei+M.SssI (72h)
Nucleosomes
present
Nucleosomes
absent
Nuclei+M.SssI (44 days) Nuclei+M.SssI (72h)
RKO+5-aza-CdR (44 days)
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp RKO+5-aza-CdR (44 days)
EPM2AIP1 MLH1 1a
1 0 0 2 0 0 3 0 0 4 0 0 5 0 0 6 0 0 7 0 0
150 bp
RKO+5-aza-CdR (72h)
Selective
Amplification
RKO+5-aza-CdR (44 days) RKO+5-aza-CdR (72h)
Selective
Amplification
RKO+5-aza-CdR (44 days)
a
b
c
d
e
f
91
treatment (Fig. 4.8a). Thus, to filter out such non-informative molecules, selective
primers were designed based on the patterns seen in LD419 cells after M.SssI
treatment (Fig. 4.4c). The primers specifically annealed to unmethylated regions
such as those seen in LD419 cells (Fig. 4.4c), amplifying the region containing the
bidirectional EPM2AIP1/MLH1 1a promoter. As controls, no PCR products were
produced from the amplification of the DNA from untreated RKO cells (data not
shown) while after 5-aza-CdR treatment, only substantially unmethylated molecules
were amplified and cloned, confirming the specificity of the primers (Fig. 4.8b).
We first verified that naked DNA extracted from 5-aza-CdR treated cells,
could be fully converted to methylated molecules by M.SssI. This was indeed the
case, since 95% (20/21) of the sequenced molecules were extensively methylated
after M.SssI treatment (Fig. 4.7b). Therefore, residual 5-aza-cytosine in DNA did
not complicate the analysis by generation of spurious patterns.
Nuclei prepared from 5-aza-CdR treated cells 72 h after drug addition, were
then exposed to M.SssI and promoter molecules were sequenced and analyzed by
selective amplification (Fig. 4.8c). About 54% (20 of 37 molecules) had patches of
protected areas large enough to accommodate nucleosomes (as previously defined),
many of which had the diagnostic footprints of a nucleosome (Fig. 4.8c) which are
∼150 or multiples of 150 bp in size (Fatemi et al., 2005). These represent
demethylated molecules which still harbored nucleosomes in the MLH1 promoter.
In contrast, a second subset of promoter had high levels of M.SssI accessibility
between the transcription start sites, flanked by inaccessible areas, indicative of a
92
nucleosome-depleted region reminiscent of the patterns seen in the expressing cells
(Fig. 4.8c, blue box, compared to Fig. 4.4c). Thus, the data strongly suggests that at
least in a subset of promoters the nucleosomes are evicted when the genes become
reactivated after drug treatment. As for the demethylated promoters which seemed
to be occupied by nucleosomes, these could possibly reflect hemimethylated
promoter molecules which are still present 72 h after drug addition according to the
drug mechanism (Bender et al., 1999; Egger et al., 2004), and thus nucleosomes
might be trapped on the hemimethylated DNA (Fig. 4.8c).
To eliminate this variable, identical analyses were done on RKO cells treated
with 5-aza-CdR, followed by 44 days of culture without the drug, at which time no
or very little hemimethylated DNA would be expected to be present. Bisulfite
genomic sequencing revealed that about 7% (2/29) of the sequenced molecules were
still completely unmethylated while the rest were extensively methylated, probably
due to remethylation after removal of the drug (Bender et al., 1999). In other words,
the proportion of demethylated molecules was reduced by about 9-fold from 62.5%
(15/24) at 72 h to 7% (2/29) at 44 days (Fig. 4.8a, d). Interestingly, the expression
level was also reduced by 7, 3 and 7-fold for the MLH1 1a, 1b and EPM2AIP1
transcript, respectively (Fig. 4.5). Thus, these results again strongly show that
promoter methylation status correlates quite well with gene expression as shown
earlier (Fig. 3.2).
Again, only substantially unmethylated molecules were detected by selective
amplification from 44 days after 5-aza-CdR treatment (Fig. 4.8e). Among them,
93
about 92% (12/13) had extensive accessibility to M.SssI between the transcription
start sites, indicative of a nucleosome depleted region, recapitulating the patterns
seen in expressing cells (Fig. 4.8f, blue box). These results show that nucleosomes
are evicted from a subset of promoters upon gene reactivation by drug-induced
methylation. Not only that, data from 44 days indicate that nearly all the
demethylated promoter molecules lack nucleosomes, establishing the heritability of
nucleosomal eviction on the demethylated, active promoter.
By combining the data from quantitative RT-PCR and the MSPA analyses
(Fig. 4.4-4.8), we found a remarkable correlation between the expression level and
the percentage of nucleosome-depleted molecules for MLH1 1a and EPM2AIP1
where the decrease in expression is associated with a reduced nucleosome-depleted
population (Fig. 4.9). Thus, it seems very likely that reactivation of such an
epigenetically silenced gene requires nucleosome depletion in addition to
demethylation of DNA and the application of positive histone marks.
DISCUSSION
Depletion of nucleosomes just upstream of transcriptional start sites has
recently been observed in genome-wide screens of yeast, flies and humans
(Heintzman et al., 2007; Lee et al., 2004; Mito et al., 2005; Ozsolak et al., 2007).
This strongly suggests that lack of nucleosomes is needed to allow access of the
transcriptional machinery to the promoter. Indeed nucleosomal remodeling factors
94
Figure 4.9. Correlation between expression level and % of nucleosome-depleted
molecules. Data were graphed based on the results from Fig. 4.5-4.8. A decrease in
expression is associated with a reduced nucleosome-depleted population.
MLH1 1a
EPM2AIP1
95
have been shown to be causally linked to transcription activation. For example,
nucleosomes in the PHO5 promoter are completely disassembled upon
transcriptional activation in yeast (Boeger et al., 2004). It therefore seems likely that
removal of at least one nucleosome is necessary for gene transcription.
These results, obtained with a complex human promoter, build on these
studies in several important ways. Firstly, the high resolution MSPA method,
validated by traditional approaches, shows that there is a constitutive complete
absence (i.e. total depletion) of a single nucleosome upstream of each start. This is
similar to the GRP78 promoter in which no trace of a nucleosome was seen in 356
promoter replicas we examined (Gal-Yam et al., 2006). While it is possible that this
might be due to a rapid and reversible nucleosomal occupancy, which is unlikely
because fixation with formaldehyde before ChIP analysis confirmed very low
occupancy in the regions examined. Also, we have shown in Chapter 2 the
preferential localization of H3-K9/14 acetylation and H3-K4 methylation near the
transcription start sites of human genes although we could not discern whether these
modifications were found upstream or downstream of the start sites at the time due to
the limited resolution of ChIP assays. Indeed, with the high resolution MSPA assays
and ChIP assays have confirmed that H3 acetylation is highly localized just
downstream of the transcription start site which could not be verified with just ChIP
assay alone.
Secondly, it is remarkable that the entire 630 base pair promoter, which
generates three transcripts, acts as a unit in the sense that either all transcripts were
96
present or none were present, so that the start sites seem to be coordinately controlled.
Equally remarkable is the presence of a highly positioned, acetylated nucleosome in
almost all the promoter replicas in expressing cells. This finding might help explain
the existence of multiple transcription start sites in many mammalian CpG island
promoters. CpG islands often do not have TATA boxes and so initiate transcription
in quite a heterogeneous fashion. Perhaps the nucleosome occupancy near the start
sites is responsible for defining the transcripts generated in a given CpG island.
Support for this idea comes from those observations of a high level of positional
variation and heterogeneous start sites in the p16 gene (Fatemi et al., 2005) in
contrast to the three, “sharply defined” sites observed by 5’-RACE in MLH1.
Further work will clearly be necessary to resolve these issues.
The third implication of this work relates to the role of occupancy in
epigenetic silencing which is, by definition, mitotically heritable. Most work to date
has focused on constitutively active genes or on the chromatin changes associated
with gene induction. As an extension of a previous study on p16 showing that DNA
methylation may serve to lock in the repressed state after H3-K9 methylation
(Bachman et al., 2003), those data suggest that heritable DNA methylation patterns
may maintain silencing, not only by “compacting” chromatin and by the erasure and
application of appropriate histone marks, but also by positioning nucleosomes at the
start sites. The almost “digital” quality of this process was seen in our experiments
with 5-aza-CdR. Here, the erasure of DNA methylation by drug treatment led to the
reactivation of all three start sites, application of the covalent activating acetylated
97
marks and removal of the nucleosomes from the region examined. Interestingly,
even though the gene was re-silenced as a function of time after treatment, the 7% of
promoters that remained unmethylated in the mass population were also nucleosome
free. Indeed, the level of expression of all three transcripts in reactivated RKO cells
was highly correlated with the level of nucleosome deficiency. ChIP analyses were
also used to verify the data and while the application of the acetylated marks was
observed after drug induction (Fig. 4.6), the removal of the nucleosomes was less
well portrayed (data not shown). Since the H3 acetylation in untreated RKO cells is
at baseline, any incremental changes from the baseline will be easily detected
whereas the H3 occupancy is already saturated.
Also, we have shown in Chapter 2 the preferential localization of H3-K9/14
acetylation and H3-K4 methylation near the transcription start sites of human genes
using the ChAP assay which couples ChIP assay and AP-PCR. At the time, we
could not discern whether these modifications were found upstream or downstream
of the start sites at the time due to the limited resolution of ChIP assays. Indeed, here
the high resolution MSPA assays together with the ChIP assays have confirmed that
H3 acetylation is highly localized just downstream of the transcription start site
which could not be verified with just ChIP assay alone as shown previously.
These results thus suggest a model of the epigenetic silencing of the MLH1
gene in which the unmethylated and active promoters of EPM2AIP1 and MLH1 (1a
and 1b) are depleted of one nucleosome just upstream of each of the transcription
start sites (Fig. 4.10). In the silenced state, the methylated and inactive promoters
98
are occupied by nucleosomes and treatment with 5-aza-CdR causes considerable
DNA demethylation of the promoter region although some molecules remain
methylated. Remarkably, while some hemimethylated promoter molecules are
occupied by nucleosomes, the nucleosome-free zone is established in a substantial
portion of the promoter molecules. Since these promoter molecules show strikingly
similar patterns to those seen in expressing cells (Fig. 4.4c), they are probably
associated with the reexpression of genes (Fig. 4.10). Conversely, the genes are
most likely silenced in the hemimethylated promoter molecules with nucleosomes in
the promoters since hemimethylation has been shown to block transcription by
inhibiting transcription factor binding (Sasaki et al., 1992b).
The involvement of nucleosome occupancy in transcriptional regulation is best
studied at the yeast PHO5 gene. Upon gene induction, histones are first
hyperacetylated and then are evicted from the promoter in trans (Reinke and Horz,
2003) and nucleosome reassembly is obligatory for rerepression (Schermer et al.,
2005).
However, when nucleosome reassembly is blocked, transcription can occur
even in the absence of the transcriptional activators under repressing conditions,
indicating the role of nucleosomes as the final “executor” for gene silencing (Adkins
and Tyler, 2006). A similar mechanism can be envisaged for the mammalian system
in which histones are hyperacetylated, followed by eviction upon gene activation and
nucleosomes reassembly at previously depleted regions upon resilencing. DNA
methylation adds another layer of repression by maintaining the silent state,
99
Figure 4.10. A simplified model for the epigenetic silencing of the MLH1 gene.
The active promoters of 1a and 1b are depleted of at least one nucleosome just
upstream of each of the transcription start sites. In the silenced state, the inactive
promoters of 1a and 1b are occupied by nucleosomes. Treatment with 5-aza-CdR
causes substantial DNA demethylation of the promoter region although some
molecules remain methylated. While some hemimethylated promoter molecules
may be still occupied by nucleosomes, the nucleosome-free zone is established in
some of the promoter molecules. The activation of genes is probably derived from
these molecules with a nucleosome-free zone in the promoter although we could not
determine definitively. Green represents nucleosomes bearing active marks. Red
represents repressive marks. Grey represents nucleosomes occupying the region of
hemimethylation. Open circles indicate unmethylated CpG dinucleotides and filled
circles, methylated CpG dinucleotides.
Ac Ac Ac Ac Ac Ac Ac Ac
5-aza-CdR
EPM2AIP1 MLH1 1a
ACTIVE
MLH1 1a EPM2AIP1
MLH1 1b
MLH1 1b
Ac Ac Ac Ac Ac Ac Ac Ac
EPM2AIP1 MLH1 1a MLH1 1b
MLH1 1a EPM2AIP1 MLH1 1b
MLH1 1a
EPM2AIP1
MLH1 1b
SILENCED
100
contributing to its heritability. It has been shown that HDAC inhibitors such as TSA
alone cannot reactivate hypermethylated promoters including MLH1 while the very
same genes can be reactivated with 5-aza-CdR (Cameron et al., 1999; Fahrner et al.,
2002). This strongly suggests that DNA methylation may serve to lock in the
repressed state, perpetuating the silencing signal and only when the lock is removed,
can reactivation take place.
In a nutshell, these data unravel a previously unrecognized mechanism for
epigenetic silencing of tumor suppressor genes. Tazi et al previously showed that
CpG island promoters are more accessible to nucleases than bulk genomic DNA
(Tazi and Bird, 1990). Consistently, these data show with high resolution that
constitutively active CpG island promoter is depleted of precisely one nucleosome
upstream of each start site while the silenced promoter is occupied by nucleosomes.
These results also showed that treatment with 5-aza-CdR leads to not only the
erasure of DNA methylation and the acetylation of histones but also the eviction of
nucleosomes, providing a link between active and epigenetically silenced states that
was missing from previous studies.
Clearly, methyl-binding proteins such as MBD1, MBD2 or MeCP2 are
involved in silencing of methylated promoters (Klose and Bird, 2006), however, the
data suggest that the core mechanism responsible for silencing may be the insertion
of a nucleosome into a previously unoccupied site. It has been shown in vivo that
MeCP2 recruits Brahma (Brm), a catalytic component of the SWI/SNF-related
chromatin remodeling complex in transcriptional silencing (Harikrishnan et al.,
101
2005). Although the exact chromatin remodeling complex regulating the eviction of
nucleosomes observed has not been identified, nucleosomal occupancy might be the
essential outcome of a chromatin remodeling process involving covalent
modification of histones and other chromosomal proteins.
102
CHAPTER 5
SUMMARY AND CONCLUSIONS
Epigenetic mechanisms such as DNA methylation, histone modifications and
nucleosome occupancy regulate eukaryotic transcription without altering the primary
nucleotide sequence in DNA and these processes coordinately contribute to the
dynamic nature of chromatin. DNA methylation is used by the cell to ensure the
correct establishment and maintenance of patterns of gene expression, and the stable
silencing of a gene is linked with hypermethylation in its promoter region.
Chromatin structure is also controlled by covalent modifications of histone tails
which interact with the underlying DNA or with other regulatory proteins that are
required for specific downstream cellular processes. Chromatin is composed of a
repeating unit of nucleosomes in which DNA is wrapped around an octamer of
histones. Nucleosomes interact with chromatin modifying enzymes, along with
other effector proteins to allow for the inter-conversion between permissive and non-
permissive states of chromatin both temporally and spatially. The research described
in this thesis characterizes the role of each epigenetic factor in transcriptional
regulation and its contribution to a diseased state such as cancer.
In Chapter 2, my studies focused on global histone modification patterns in
cells because a detailed map of chromatin features across the entire human genome
would enrich our understanding of transcriptional regulation. At that time, most
work in the field focused on studying histone modifications at specific loci, one at a
103
time and also, it was believed that histone acetylation and K4 methylation were
enriched throughout transcribed regions. In order to assess global histone
modification patterns in cells, we developed a chromatin scanning technique called
ChAP, combining the chromatin immunoprecipitation assay (ChIP) and arbitrarily
primed PCR (AP-PCR), which allows for the rapid and unbiased comparison of
histone modification patterns within the eukaryotic nucleus. This assay differs from
the commonly used “Chip on Chip” assays in that it is not limited by the sequences
applied to the microarray. Dr. Gangning Liang performed all the ChAP assays while
I sequenced the majority of the fragments isolated from the ChAP assays. We found
that methylated K4 and acetylated K9/14 of histone H3 were both highly localized
near the transcription start sites of transcriptionally active human genes. These data
indicate that the large transcribed regions of human genes are, in fact, maintained in
a deacetylated configuration. The strong concordance of H3-K9/14 acetylation and
H3-K4 methylation near the transcription start sites of genes indicate that these
modifications may serve as critical signals for defining the start of the transcribed
domain. However, we could not discern completely whether these modifications
were found upstream or downstream of the transcription start sites due to the limited
resolution of the ChIP assays.
Motivated by our findings in Chapter 2 which show the preferential
localization of histone H3 acetylation and H3-K4 methylation to the transcription
start sites in the human genome, in Chapter 3, I shifted our attention to examine the
role of the three major epigenetic mechanisms in the transcriptional regulation in the
104
MLH1 gene, a CpG island bidirectional promoter. While it is important to study the
global pattern of histone marks, it is equally valuable to study in detail these
epigenetic processes at a specific locus. At that time, the control of CpG island
bidirectional promoter by epigenetic parameters was unclear. To address this
question, I first established by 5’-RACE analysis that MLH1 has two transcriptional
start sites (1a and 1b) spaced 309 bp apart while the other gene, EPM2AIP1 initiates
transcription at only one site, 321 bp upstream of MLH1 1a. Promoter deletion
analysis showed that the 321-bp intergenic region indeed has bidirectional promoter
activity. The genes are coordinately expressed in cell lines with low promoter
methylation while both genes are silenced together in a series of cancer cell lines in a
process associated with dense CpG island methylation. Furthermore, cell lines
displaying promoter hypermethylation were treated with 5’-aza-2’-deoxycytidine (5-
aza-CdR), leading to the concordant reactivation of both genes after drug treatment.
Thus epigenetic silencing results in the suppresson of both genes showing the
promoter acts as a unit. These results show that epigenetic silencing can result in the
simultaneous inactivation of two genes.
In Chapter 4, I sought to investigate the chromatin structure at all three
transcriptional start sites of the promoter. The high resolution MSPA method shows
that there is a constitutive complete absence of a single nucleosome just upstream of
each start site. These results were validated by traditional approaches including
DNaseI footprinting, mononucleosomal DNA analysis and chromatin
immunoprecipitations. ChIP assays also showed that histone H3 acetylation was
105
high in all regions analyzed in expressing LD419 cells as compared to nonexpressing
RKO cells. While many studies have shown that promoter regions are often
associated with increased acetylation in active promoters without specific references
to H3 occupancy, our data indicate that the increased acetylation is actually coupled
with H3 enrichment in regions surrounding the promoter.
Also, we showed previously in Chapter 2 the preferential localization of H3-
K9/14 acetylation and H3-K4 methylation near the transcription start sites of human
genes using the ChAP assay which couples ChIP assay and AP-PCR. As mentioned
above, we could not discern completely whether these modifications were found
upstream or downstream of the start sites at the time due to the limited resolution of
ChIP assays. Indeed, here the high resolution MSPA assays together with the ChIP
assays confirmed that H3 acetylation is highly localized just downstream of the
transcription start site which could not be verified with just ChIP assay alone as
shown previously.
In addition, three distinct nucleosomes almost completely absent from the
start sites in normal cells are, in fact, present on the silenced promoter. Thus,
epigenetic silencing may be characterized by the placement of nucleosomes into
previously vacant positions. The MSPA assay was also applied to detect chromatin
structural changes in nonexpressing RKO cells following gene activation upon 5-
aza-CdR treatment. Strikingly, activation of the silenced promoter by 5-aza-CdR
seems to involve nucleosome eviction, showing that epigenetic silencing of tumor
suppressor genes may involve heritable changes in nucleosome occupancy enabled
106
by cytosine methylation. Silencing in cancer involves DNA methylation and
chromatin covalent and structural changes which are somatically heritable and
contribute to the cancer phenotype. Thus, these data show that the switch from
permissive chromatin configuration to a state which is permanently repressed and
non-permissive may involve the complete absence or presence, respectively, of
nucleosomes at the three start sites of the bidirectional promoter.
My studies described in this thesis characterized the three major epigenetic
mechanisms in transcriptional regulation and how these processes contribute to
tumorigenesis. Indeed, abnormal alterations in the patterns of these epigenetic
parameters have been equated with several human diseases, in particular, cancer,
making them attractive targets for therapeutics. Profiles of normal and abnormal
epigenetic patterns will enable us to elucidate how these important parameters may
contribute to the cancer phenotype and shed light on the search for novel therapeutic
agents.
107
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Abstract (if available)
Abstract
Transcriptional regulation can be controlled by epigenetic mechanisms including DNA methylation, histone modifications and nucleosome occupancy. We first mapped histone H3 acetylation and H3-K4 methylation patterns globally across the human genome using our novel genome scanning method called ChAP and showed that they are located preferentially to the transcriptional start sites in the human genome. The strong concordance of these modifications at the 5' regions of genes indicate that these modifications may serve as critical signals for defining the start of the transcribed domain. We then examined these epigenetic parameters in detail using the bidirectional CpG island promoter, MLH1 gene as an example. MLH1, which has two transcriptional start sites and the other gene EPM2AIP1, are coordinately expressed in cell lines with low promoter methylation and both are silenced together in cancer cell lines with promoter hypermethylation, suggesting the promoter acts as a unit. In addition to DNA methylation, silencing of the three transcription start sites in the bidirectional MLH1 promoter in cancer cells also involves distinct changes in nucleosomal occupancy. Three nucleosomes, almost completely absent from the start sites in normal cells, are present on the methylated and silenced promoter, suggesting that epigenetic silencing may be accomplished by the stable placement of nucleosomes into previously vacant positions. Indeed, activation of the promoter by demethylation using 5-aza-2'deoxycytidine involves nucleosome eviction.
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Creator
Lin, Joy Chieh-Yu
(author)
Core Title
Transcriptional regulation by epigenetic mechanisms
School
Keck School of Medicine
Degree
Doctor of Philosophy
Degree Program
Biochemistry and Molecular Biology
Degree Conferral Date
2007-08
Publication Date
06/18/2009
Defense Date
05/16/2007
Publisher
University of Southern California
(original),
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Tag
bidirectional promoters,chromatin structures,OAI-PMH Harvest
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English
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Jones, Peter A. (
committee chair
), Dubeau, Louis (
committee member
), Stellwagen, Robert H. (
committee member
)
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Lin, Joy Chieh-Yu
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bidirectional promoters
chromatin structures