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Targeting chromatin modification in human cancer: SMYD3 mediated ERα transcription regulation. Cooperative role between H3.3 and HP1ϒ

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Targeting chromatin modification in human cancer: SMYD3 mediated ERα transcription regulation. Cooperative role between H3.3 and HP1ϒ

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Content TARGETING CHROMATIN MODIFICATION IN HUMAN CANCER:

SMYD3 MEDIATED ERα TRANSCRIPTION REGULATION.

COOPERATIVE ROLE BETWEEN H3.3 AND HP1γ.

by

Hyunjung Kim

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETIC, MOLECULAR, AND CELLULAR BIOLOGY)

May 2011

Copyright 2011 Hyunjung Kim

ii

ACKNOWLEDGEMENTS

It would not have been possible to come to this finish line without the guidance
and the support of several individuals who provided their valuable input in the
preparation and completion of the study. I am heartily thankful to my advisor, Dr.
Woojin An, who has supported my thesis with his patience and steadfast
encouragement. I am grateful for his support from the initial to the final level
enabled me to develop an understanding of the subject.

I would like to express my sincere gratitude to Dr. Michael Stallcup, Chair of the
Department of Biochemistry, who had kind concern and consideration regarding
my academic requirements and unfalling support as my dissertation committee
adviser. Dr. Nouri Neamati, for the advice and support to complete this study as
dissertation committee.

I would like to thank my colleague, Dr. Kyu Heo, who as a good friend, always
willing to help and give his best suggestions. It would have been a lonely lab
without him. Many thanks to Dr. Jongkyu Choi, Dr. Kyunghwan Kim and other
workers in the laboratory of Dr. An for helping me to develop the study in one
way or another. I would also like to thank my dear friends, Helty, Vivian, Bernice,
and Jennifer, who have been with me from the beginning of the doctoral

iii

program, and Seongken Jeon, for cheering me up and stood by me through the
good times and bad. Dr. Kwangho Lee, Dr. Siho Choi, Heejin Kim for reagents
and priceless friendship. Korean journal club members for encouraging me with
their best wishes.

Last but not the least, my family for supporting me throughout all my studies with
their undying love and the one above all of us, the almighty God, for answering
my prayers and giving me the strength to walk to the finish line despite wanting to
throw in the towel and leave, thank you, Lord.

iv

TABLE OF CONTENTS

ACKNOWLEDGEMENTS……………………………………………................ ii

LIST OF FIGURES………………………………………………………………. v

ABSTRACT………………………………………………………………………. vii

CHAPTER 1: INTRODUCTION………………………………………………… 1

CHAPTER 2: Requirement of Histone Methyltransferase, SMYD3
for Estrogen Receptor mediated transcription……………….. 15

CHAPTER 3: Histone variant H3.3 stimulates HSP70 transcription through
cooperation with HP1γ………………………………………….. 50
Table 3-1…………………………………………………………. 90

CHAPTER 4: CONCLUDING REMARKS…...……………...………………… 91

BIBLIOGRAPHY…………………………………………………………………. 95

v

LIST OF FIGURES

Fig. 1-1. Hierarchical structure of Chromatin……………………………. 2
Fig. 1-2. Structure of nucleosome………………………………………… 5
Fig. 1-3. The domains of the core histone proteins…………………….. 6
Fig. 1-4. H3 variants in different organisms……………………………… 8
Fig. 1-5. Sequence alignment of H3. proteins…………………………… 11
Fig. 1-6. Alignment of HP1 amino acid sequences……………………... 14
Fig. 2-1. Direct interaction between SMYD3 and ER…………………… 25
Fig. 2-2. Requirement of NHSC and EEL motifs for SMYD3 HMT
activity……………………………………………………………... 28

Fig. 2-3. Coactivator function of SMYD3 in ER transcription………….. 32
Fig. 2-4. Estrogen-induced accumulation of SMYD3 and H3-K4
methylation at ER target genes………………………………… 36

Fig. 2-5. Requirement of SMYD3 for ER transcription…………………. 41
Fig. 2-6. Reduction of H3-K4 methylation upon SMYD3 depletion……. 44
Fig. S3-1. Preparation of H3/H3.3 nucleosomes…………………………. 58

Fig. S3-2. Immunostaining of HP1 isoforms………………………………. 59

Fig. S3-3. Co-localization of ectopic H3.3 and HP1γ at HSP70
promoters..………………………………………………………... 60

Fig. 3-1. Selective interaction of HP1γ with H3.3 nucleosomes……….. 63
Fig. 3-2. Requirements for H3.3 and HP1γ in HSP70 transcription…… 66

vi

Fig. S3-4. Interdependent promoter occupancy of ectopic H3.3 and
HP1γ………………………………………………………………. 69

Fig. 3-3. Interdependent localization of H3.3 and HP1γ at HSP70
promoters…………………………………………………………. 70

Fig. 3-4. Enrichment of active modifications in H3.3 nucleosomes…… 73
Fig. 3-5. HP1γ interacts preferentially with H3 tails carrying active
histone marks…………………………………………………….. 77

Fig. S3-5 Differential expression patterns of the HP1 proteins in cancer
cells………………………………………………………………... 79

Fig. 3-6. Affects of H3.3/HP1γ knockdown on HSP70 transcription…... 81
Fig. 3-7. H3.3/HP1γ knockdown-induced alterations in cancer cell
growth……………………………………………………………... 83

vii

ABSTRACT

The genome of eukaryotic cells is composed in nucleoprotein structure called
‘chromatin’. The basic repeating unit of chromatin is nucleosome, which consists
of two pairs of H2A-H2B dimmers, and one H3-H4 tetramer wrapped around 147
bp of double stranded DNA (dsDNA). Although, the structure of chromatin is
necessary for hierarchical compaction of the entire genome, it is also an obstacle
for some of the most important cellular processes like transcription, DNA repair,
DNA replication. In order to alter the structure between histones and DNA,
eukaryotic cells imply three basic mechanisms: chromatin remodeling,
incorporation of histone variants, and post-translational modification of histones.
Post-translational modification is considered as epigenetic modification, which
plays a vital role in gene regulation as well as maintaining genome stability.

Although, many different post-translational modifications on histones are
identified, histone acetylation mediated by histone acetyltransferases (HATs) and
histone methylation mediated by histone methyltransferases (HMTs) are two
major modifications. The detailed mechanistic processes on how these covalent
modifications are introduced to the target amino acid is not clear, different
modifications and combinations of modifications, the so-called ‘histone code’,
produce entirely different cellular processes such as transcription activation,

viii

repression, DNA repair, DNA replication, and so on. Therefore, studying histone
modifying enzymes can bring detailed understanding on the functions of different
covalent modifications at the cellular level. SMYD3 is one of the histone
methyltransferases that methylates histone H3-K4. Recent studies showed that
SMYD3 is frequently overexpressed in different types of cancer cells, but how
SMYD3 regulates the development and progression of these malignancies
remains unknown. I report here the previously unrecognized role of SMYD3 in
estrogen receptor (ER)-mediated transcription via its histone methyltransferase
activity. I demonstrate that SMYD3 functions as a coactivator of ER and
potentiates ER activity in response to ligand. SMYD3 directly interacts with the
ligand binding domain (LBD) of ER and is recruited to the proximal promoter
regions of ER target genes upon gene induction. Importantly, the chromatin
immunoprecipitation analyses provided compelling evidence that SMYD3 is
responsible for the accumulation of di- and trimethylation of H3–K4 at the
induced ER target genes. Furthermore, RNA interference-directed down-
regulation of SMYD3 reveals that SMYD3 is required for ER-regulated gene
transcription in estrogen signaling pathway. Thus, these results identify SMYD3
as a new coactivator for ER-mediated transcription, providing a possible link
between SMYD3 overexpression and breast cancer.

The exchange of histone variants is another mechanism, which can regulate
transcription and other cellular processes. Unlike canonical histones, histone

ix

variants can be incorporated into the chromatin independent from DNA
replication. Histone H2A and H3 variants have been introduced to the field of
epigenetics for decades and numbers of different studies have shown the
evidence of the histone variants involved in specific cellular processes causing
different chromatin states.

Histone H3.3 is one of H3 ‘replacement’ variants. Although, there are only 5
amino-acid sequence differences between human H3 and H3.3, the
incorporation, localization, and the specification of H3.3 into the chromatin seems
very distinct from the histone, H3. For instance, H3.3 is significantly recognized in
transcriptionally active loci. However, it is not clear how it exerts its function such
as whether H3.3 recognizes the active chromatin states or other proteins aid the
incorporation of this variant.
A number of studies suggest that H3.3 plays a role in transcriptionally active
chromatin and may be involved in the epigenetic maintenance of chromatin
status.

It has been shown that H3.3 incorporation was triggered into HSP70 gene loci
upon transcriptional activation in Drosophilla. However, the mechanistic details
on how H3.3 is incorporated into the chromosome were not discussed such as
whether H3.3 incorporation depends on other protein binding or how it targets to
the active loci. In this study, we purified H3 and H3.3 specific mononucleosomes

x

from a human cell line to search for proteins specifically preferential for H3.3
containing mononucleosomes. Interestingly, we observed preferential binding of
one of the HP1 proteins, HP1ϒ, to H3.3 containing mononucleosome and we
further investigated the function of HP1ϒ and H3.3 in regulating HSP70 genes
which affects the growth of breast cancer cell.

1

CHAPTER 1: INTRODUCTION

Chromatin structure and gene regulation

Eukaryotic cells carry their genetic information in a genetic blueprint called ‘DNA’.
Each of the cells in human body contains DNA, which is far greater than the size
of the nucleus in which it is contained. In order to fit the DNA without losing any
genetic information, the DNA has to be compacted in hierarchical structure called
‘chromatin’. However, DNA is not packaged directly into final structure of
chromatin. Instead, it contains several hierarchies of organization (Fig. 1-1). The
fundamental structure of chromatin is achieved by the winding of double stranded
DNA (dsDNA) around a protein core to produce a “bead-like” structure (Fig. 1-2)
called a nucleosome (McGhee et al., 1980). According to the crystal structure of
nucleosome solved by Luger et al., a mononucleosome is comprised of histone
octamers made with two copies of histone H2A, H2B, H3, and H4 wrapped by
147 base pairs (bp) dsDNA. This structure introduces 14 contact points between
histones and DNA which allows nucleosome to be highly stable protein-DNA
complex.

For decades, the assumed role of chromatin was only limited to DNA compaction
and subsequent gene repression. However, there has been tremendous

2

Figure. 1-1. Hierarchical structure of Chromatin. Schematic illustration of
chromatin fiber condensation. Shown are the steps involved in the folding of
extended nucleosomal arrays into maximally folded chromatin fibers (Hansen,
2002).

Annual Review of Biophysics and Biomolecular Structure Vol. 31: 361-392
(2002).

3

progress in this field over the last 10 years, which elevated chromatin in a key
position in gene regulation. A large molecular machine with dedicated functions
to disrupt chromatin structure and facilitate transcription, the high-resolution
structural description of the histone octamer, the architectural motif, and the fold
of all core histones indicate specialized functions of individual chromosomal
domains. In addition, the existence of histone variants contributes to a unique
nucleosomal structure to regulate a wide range of nuclear functions. Another
important evidence that showed the importance of chromatin structure in gene
regulation was histone modifications on N-termini.

Histone modifications

The core histone protein consists of the globular domains that form the spool
onto which the nucleosome DNA is wrapped. These domains are structurally
similar three-helix “handshake” motifs, which interlock to form heterodimers of
H2A/H2B and H3/H4 (Arents and Moudrianakis, 1993). The remaining of the core
histone protein comprises the structurally undefined “tail” domains (Fig. 1-3).
These N-terminal tails of the core histone protein are relatively exposed to the
cellular environment and are readily accessible to histone modifying enzymes
such as acetyltransferases, methyltransferases, kinases (Grunstein, 1997; Zhang
and Reinberg, 2001), ubiquitinligases, and other chemical modifications (Hill and
Thomas, 1990; Lambert and Thomas, 1986). In addition, these tail domains

4

mediate internucleosomal interactions within condensed chromatin structures.
Since the tail domains can adopt multiple conformations depending on the
conditions of the media, the exact tail conformation and interactions are still in
unclear state.

Numerous kinds of posttranslational modifications occur on the N-terminal tails of
the core histone protein that modulate the state of chromatin. Therefore, the
histone tails are significant regulatory end points for signal transduction within the
eukaryotic nucleus. However, how the tail domains mediate structural and
functional states of chromatin and how different posttranslational modifications
within the tail domains mediate transcriptional regulation is unclear.

Histone Variant, H3.3

In addition to covalent histone-tail modifictions, histone variants localized at
specific chromatin loci also play important roles in the formation and
maintenance of transcriptionally active and inactive chromatin as well as in
maintenance of epigenetic memory. There are four isoforms of histone H3
(CENP-A, H3.1, H3.2, and H3.3) identified in mammals. Histone H3.1 is the
canonical histone H3 (Fig. 1-4), which is strictly incorporated into the chromatin
during S Phase. Histone H3.2 is closely related to H3.1 and belongs to the family

5

Figure. 1-2. Structure of nucleosome image of the elctron microscope. The
image of electron microscope shows the “beads-on-a-string” form of the native
chromatin after treatments that unpack. A) The native chromatin structure in 30-
nm fiber form. B) The decondensed, “beads-on-a-string” form of chromatin
(Alberts et al. 2002).

Molecular Biology of the Cell. 4
th
edition.
Alberts B, Johnson A, Leis J, et al. New York, Garland Science 2002

6

Figure. 1-3. The domains of the core histone proteins. Tail sequences for the
main human histone variants are shown. The gray box indicates the histone fold
domains. The red T’s indicates the peptide bond closest to the histone fold
domain susceptible to trypsin proteolysis in the nucleosome core. The vertical
blue arrows indicate the approximate point where the tails exit either through
(H2B and H3) or over/under (H2A/H4) the dsDNA wrapping the nucleosome
(Zheng C, Hayes JJ, 2003).

Biopolymers. 2003 Apr;68(4):539-46.

7

of S-phase subtypes (Franklin and Zweidler, 1977). However, H3.3 is expressed
in cells during all stages of the cell cycle as well as in quiescent cells (Wu et al.,
1982). In fact, H3.3 can be incorporated into nucleosomes independent of DNA
synthesis. Given these properties, it has been indicated that H3.3-H4 replaces
H3-H4 in nucleosomes, particularly during transcriptional activation (Ahmad and
H3nikoff 2002; H3nikoff et al. 2004). It is clear that the incorporation of H3.3
contributes a significant role in cell fate, albeit little is known about this variant.

H3.3 is a highly conserved protein, which is present in all eukaryotes (Malik and
Henikoff, 2003). H3.3 differs from canonical H3 or H3.2 by only four amino acids
at positions 31, 87, 89, and 90 (Fig 1-5). Interestingly however, these minor
differences in amino acid residues are responsible for intrinsic stability between
H3 and H3.3. The main features of H3.3 are its involvement in transcription
activation and its potential role in the epigenetic transmission of active chromatin
states (Hake and Allis, 2006).

Transcription coupled deposition of H3.3 has been directly observed in vivo on
Drosophila polytene chromosomes (Schwartz and Ahmad, 2005). Other studies
have shown the results with similar conclusion by analyzing the distribution of
H3.3 nucleosomes at high resolution by chromatin immunoprecipitation (ChIP)
(Mito et al., 2005; Wirbelauer et al., 2005). These studies have revealed that

8

Figure. 1-4. H3 variants in different organisms. (A) Schematic of evolutionary
appearance of histone H3 variants. All organisms express a centromerespecific
H3 variant (CENP-A, filled blue circle). In addition to the centromeric
H3 variant, the following H3 variants are expressed in these organisms: S.
cerevisiae contains only H3.3 (blue gradient circle); S. pombe expresses a
hybrid H3 protein that contains amino acids characteristic for H3.3 and H3.2;
Arabidopsis thaliana, Xenopus laevis, and Drosophila melanogaster (for
example) express H3.3 and H3.2 (blue circle with white dots); mammals such as
Mus musculus and H. sapiens express H3.3, H3.2, H3.1 (white circle with blue
dots), and a testis-specific H3.1t (white circle with blue stripes) variant of
unknown function. H3.3 has been associated with euchromatin and
transcriptional activation. H3.2 and H3.1 might localize to heterochromatin and
are involved in transcriptional silencing. (B) Alignment of human noncentromeric
histone H3 variants. Differences in amino acid sequence among human H3.3,
H3.2, H3.1, and H3.1t are shown in white boxes. Cysteine residues are
highlighted in red (Cys 96 in dark red and Cys 110 in pink). Identical amino acids
are shown in gray. TS, tissue-specific. The region where most amino acid
differences between the variants are found is underlined as a potential
chaperone recognition domain (see text for details), and the chaperones binding
to H3 variants are depicted below (Hake et al., 2005).

9

Figure. 1-4: Continued.

Proc Natl Acad Sci U S A. 2006 Apr 25;103(17):6428-35. Epub 2006 Mar 29.
Review(Orsi et al., 2009)

10

H3.3 is highly enriched at the promoters of active genes, suggesting that
chromatin remodeling associated with transcriptional initiation is also responsible
for H3.3 deposition (Chow et al., 2005). Recent studies, though, found the
enrichment of H3.3 at regulatory sites of active as well as at silent genes (Jin and
Felsenfeld, 2006; Mito et al, 2007; Nakayama et al., 2007). These observations
suggest two distinct roles of H3.3 involved in transcription regulation.

In addition to the incorporation of H3.3 at sites of active chromatin, H3.3 is
enriched with posttranslational modifications associated with transcription
activation, such as histone H3 lysine 4 (Hake et al., 2006; McKittrick et al., 2004;
Mito et al., 2005). How these posttranslational modifications are established on
H3.3 and their importance in gene activity to this variant are crucial questions.

Heterochromatin binding Proteins (HP1)

HP1 is a conserved chromatin binding protein originally identified in Drosophila
as a non-histone chromosomal protein functioning in heterochromatin-mediated
gene silencing (Eissenberg et al., 1990). HP1 proteins contain two conserved
domains; the chromodomain (CD) at the N-terminus, and the chromo shadow
domain (CSD) at the C-terminus (Fig. 1-6). The CD was shown to bind to
chromatin, while the CSD is implicated in various protein-protein interactions and

11

Figure. 1-5. Sequence alignment of H3.3 proteins. A) CLUSTAL alignment of
H3 and H3.3 histones. Significantly conserved amino-acid residues are shaded in
gray. In positions 31, 87, 89, and 90, amino acids from canonical H3s are shaded
in red, those from H3.3 family are shaded in green and those are fitting these
categories are shaded in purple. The single residue that differentiates mouse
H3.1 and H3.2 and differs among the H3.3 family is shaded in cyan. B) Residues
31, 87, 89, and 90 are positioned on a schematic representation of nucleosomal
H3 protein (Orsi et al., 2009).

Int. J. Dev. Biol. 53: 213-243 (2009)

12

dimerization.

There are three different HP1 variants in mammalian cells; HP1a, HP1b, and
HP1g (Fig. 1-6). In mammalian cells, the cytological distributions of the three
HP1 variants was reported to be distinct. For instance, HP1a and HP1b are
found at heterochromatin, while HP1g is found in both heterochromatic and
euchromatic regions (Minc et al., 1999; Minc et al., 2000). Consistent with the
cytological observation, HP1g was found to associate with actively transcribed
gene regions and plays a role in efficient transcriptional elongation(Lomberk et al.,
2006). Interestingly, depletion of each HP1 subtype in knockdown studies have
shown the functional specificities of HP1 variants in the cell(Aucott et al., 2008;
Serrano et al., 2009).

Numerous studies have shown that HP1 plays a role in heterochromatin
formation and gene silencing in many organisms in the past. However, recent
studies has shown the involvement of HP1 in different functions regarding
chromatin configuration such as the interaction of HP1 with nuclear membrane,
metaphse chromatid cohesion, centromere organization, telomere capping and
silencing and positive control of gene expression. However, it is unknown what
determines the specific functions among different HP1 subtypes, and how HP1
proteins can perform different functional roles and still remain to be answered. It
could be that each HP1 protein has different partners in different contexts or

13

posttranslational modifications that allow different interactions in different
contexts.

14

Figure. 1-6. Alignment of HP1 amino acid sequences. A) Alignment of
mammalian HP1 amino acid sequences. The sequences of mammalian HP1b
and Hp1g are compared with HP1a. The percentage of homology is based on
comparison to HP1a (Indicated in white/gray). B) Alignment of Hp1 amino acid
sequences between different species. The Hp1 amino acid sequences from
several non-mammalian species are compared to the corresponding mammalian
HP1 ariant and their homology with mammalian Hp1 is indicated in the color
diagram (Zeng et al.).

Epigentics 16;5(4): 287-292 (2010)

15

CHAPTER 2: Requirement of Histone Methyltransferase SMYD3
for Estrogen Receptor – mediated Transcription*

CHAPTER 2 INTRODUCTION

Estrogen receptor (ER) α is a member of the nuclear receptor

superfamily and
the primary biosensor for estrogen (Mangelsdorf et al., 1995; McKenna et al.,
1999). Upon activation by estrogen, ER binds to specific DNA sequences called

estrogen response elements (EREs) to induce expression of a number of target

genes in specific organs, including

the female reproductive organs, the central
nervous system,

and bone (Leng et al., 1994; Mangelsdorf et al., 1995; Metivier
et al., 2003). ER is comprised of several structural domains that are highly
conserved in the various nuclear receptors: the N-terminal transcription activation
domain, the DNA binding domain, the hinge region and the C-terminal conserved
ligand binding domain (Enmark and Gustafsson, 1999; Godowski et al., 1988;
Yamamoto et al., 1988). Like other nuclear receptors, the ER collaborates with a
number

of transcriptional cofactors to effectively modulate transcription

of its
target genes (McDonnell et al., 2002; Metivier et al., 2003; Schotta et al., 2002;
Shang et al., 2000). These cofactors appear to regulate the chromatin
configuration

in a highly specific manner by controlling nucleosomal
rearrangement

and histone modifications at the promoter (Daujat et al., 2002;

16

Imhof and Wolffe, 1998; McKenna and O'Malley, 2002). This targeted alteration
of

chromatin structure allows the transcriptional machinery to access the
chromatin DNA and form functional preinitiation complexes, thereby facilitating
transcription initiation (Becker, 2002; Kim et al., 2001; Thomas et al., 2006).

Two major types of chromatin remodeling have been widely investigated for ER
transcription. One such remodeling activity includes ATP-dependent chromatin
remodeling factors which alter structure and position of nucleosomes at the
promoters of ER target genes. These include proteins such as brahma-related
BRG1 (also known as hBRG1 or hSNF2) and BRM, both of which are subunits of
the mammalian homologue of the yeast SWI/SNF complex (Chen et al., 2003;
Yoshinaga et al., 1992). The second class of remodeling factors includes a
diverse group of single/multisubunit factors that effect posttranslational
modifications of the histone tails protruding from the surface of the nucleosome
(Huang et al., 2003; Kobayashi et al., 2000; Yoshinaga et al., 1992). Among the
well-known histone modifying factors acting in ER-mediated transcription are
histone acetyltransferases including p300/CBP and GCN5/PCAF and histone
methyltransferases including the arginine methyltransferases CARM1 and
PRMT1, as well as SET-domain lysine methyltransferases such as G9a, RIZ1,
NSD1 and MLL2 (Carling et al., 2004; Kim et al., 2001; Lee et al., 2006;
Rayasam et al., 2003; Schotta et al., 2002). These remodeling factors are
recruited to the promoter proximal region of some of the ER target genes

17

(DiRenzo et al., 2000; Metivier et al., 2003) and facilitate either remodeling or
removal of the underlying nucleosome, thereby increasing the accessibility of
promoter regions to the transcription machinery.

Recent studies identified that SMYD3 possesses histone methyltransferase
activity responsible for catalyzing methylation of histone H3 at K4 (Hamamoto et
al., 2004). SMYD3 contains a SET domain, which is crucial for HMT activity, and
a MYND-type zinc-finger domain (zf-MYND) domain, which is common to
developmental proteins (Ansieau and Leutz, 2002; Lutterbach et al., 1998).
Interestingly, misregulation of H3 methylation events upon over-expression of
SMYD3 has been shown to correlate with the development and progression of
colorectal and hepatocellular carcinoma (Hamamoto et al., 2004). In addition

to
its role in growth of cancer cells, a possible role of SMYD3 in transcription has
been supported by its interaction with RNA polymerase II to form transcriptional
complexes (Hamamoto et al., 2004). In fact, a microarray analysis of SMYD3-
transfected cells has revealed that a large number of genes were up-regulated
more than 3-fold in the SMYD3-overexpressing cells compared with those in the
normal cells (Hamamoto et al., 2004). Of special relevance to the present study
is that the over-expressed levels of SMYD3 have been observed in breast cancer
tissues as well as breast cancer cell lines with associated effects on cancer
growth (Hamamoto et al., 2006). ER serves as a sequence specific transcription
factor to regulate a cascade of gene targets whose products mediate the

18

initiation, development, and metastasis of breast cancers. Thus, these results
support the idea that SMYD3 might play a functional role in the transactivation of
ER-mediated gene transcription in breast cancer cells.

As a starting point for study of transcriptional processes

regulated by SMYD3, we
checked a possible role of SMYD3 in the ER signaling process. From molecular
and cellular studies, we have

obtained evidence indicating that SMYD3 is
critically involved in ligand-activated, ER-mediated transcription, by methylating
histone H3-K4 at the ERE in the promoter regions of target genes. The function
of SMYD3 in ER-mediated transcription requires its direct interaction with ER,
which in turn allows its recruitment to promoter regions of ER target genes.
Down-regulation of SMYD3 expression and concomitant reduction of H3-K4
methylation substantially repressed expression of ER target genes, revealing a
major role for SMYD3 as regulator of ER-mediated target gene transcription.

CHAPTER 2 MATERIALS AND METHODS

Plasmid construction. For mammalian expression of SMYD3, SMYD3 cDNA
was PCR-amplified from the pool of MCF-7 cDNA using a 5’ primer (5’-
AAGGAAAAAAGCGGCCGCATGCGATGCTCTCAGTGCCGC) and a 3’ primer
(5’CGCGGATCCTTAGGATGCTCTGATGTTGGCGT), which introduced BamHI

19

and NotI sites at the 5’ and 3’ ends, respectively. The PCR products were
digested with NotI-BamHI and inserted into NotI and BamHI sites of pIRES
containing Flag and HA tags to generate the plasmid for mammalian expression.
For bacterial expression, SMYD3 cDNA was PCR-amplified with a forward primer
(5’-GGAATTCCATATGCGATGCTCTCAGTGCCGCGT-3’) and a reverse primer
(5’-CGCGGATCCTTAGGATGCTCTGATGTTGGCGT-3’), introducing NdeI and
BamHI sites at the 5’ and 3’ ends, respectively. The corresponding products
were digested with NdeI-BamHI and inserted into NotI and BamHI sites of pET11
containing Flag tag. The same procedure was followed to construct the bacterial
expression plasmids encoding the mutant SMYD3 (∆NHSC, ∆EEL, and ∆NHSC
& EEL) except that the original SMYD3 cDNA was first deleted at NHSC and/or
EEL motifs by using the QuikChange mutagenesis kit (Stratagene). All mutations
were confirmed by DNA sequencing.

Pulldown and immunoprecipitation assays. For in vitro interaction assays with
SMYD3, Flag epitope-tagged SMYD3 was synthesized in vitro by using TNT-
Quick coupled transcription/tranlsation system (Promega) and incubated with
GST-ER coupled to Glutathione Sepharose beads (Pharmacia) at 4°C in 1 ml of
binding buffer (20 mM Tris-HCl, pH 7.3, 0.2 M KCl, 0.2 mM EDTA, 20% Glycerol,
and 0.01% NP-40) for overnight. After washing three times with 500 µl of binding
buffer, the beads were subjected to 10% SDS-PAGE and Western blot analysis
using anti-Flag antibody. For interaction assays with AF-I and AF-II domains of

20

ERα, the domains were synthesized by using TNT-Quick coupled
transcription/translation system (Promega) and incubated with recombinant Flag-
tagged SMYD3 prepared from E. Coli along with Flag-M2 agarose beads
(Sigma). For coimmunoprecipitation assays, Flag tagged and untagged SMYD3
proteins were expressed in 293T and MCF-7 cells, and whole cell lysates were
prepared from the cells with lysis buffer (20 mM HEPES, pH 7.8, 150 mM NaCl,
10 mM EDTA, 2 mM EGTA, and 2 mM DTT, and 0.1% NP-40). The cell lysates
were mixed with Sepharose beads conjugated with anti-Flag antibody (Sigma)
and rotated at 4°C for overnight before removal of the supernatant. The resulting
samples were analyzed by Western blot analysis using anti-Flag and anti-ER
antibodies (Santa Cruz Biotechnology). To coimmunoprecipitate endogenous
SMYD3 and ER, 293T cell lysates

were reacted with anti-ER antibody or normal

rabbit IgG (Santa Cruz Biotechnology) for overnight. After centrifugation,
immunocomplexes

in the supernatants were precipitated with Protein A/G
sepharose

(Millipore) and separated

on 15% SDS-PAGE. Immunoblot analyses
were performed with

anti-ER and anti-SMYD3 antibodies.

Histone methyltransferase assay. HMT assays were performed as described
previously (Nishioka et al., 2002). 293T cells were transfected with empty
expression plasmid (negative control) or plasmids expressing Flag-HA-tagged
wild-type SMYD3 (Flag-HA-pIRES-SMYD3), mutant SMYD3 (Flag-HA-pIRES-
∆EEL, Flag-HA-pIRES-∆NHSC, and Flag-HA-pIRES-∆EEL&∆NHSC) and SET7/9

21

protein (positive control), or pFLAG-HA-pIRES (negative control), and the
proteins were purified by immunoprecipitation with anti-Flag antibody.
Recombinant histone octamers (1 µg) were incubated with SMYD3 for 1 hour at
30ºC in HMT reaction buffer (100 mM HEPES, pH7.8, 300 mM KCl, 2.5 mM
EDTA, 25 mM dithiothreitol, 50 mM sodium butyrate) in the presence of 2.3 µM
[
3
H]AdoMet or 50 µM cold AdoMet. Proteins were resolved on 15% SDS-PAGE
gel and visualized by fluorography. The antibodies used for detection of
mono/di/tri-methyl H3-K4 were from Abcam.

Luciferase Assay. 293T and MCF-7 cells were plated into 12-well plates at a
density of 1X10
5
cells. Cells were transfected using Lipofectamine (Invitrogen)
with 200 ng MMTV-luciferase reporter vector and wild-type and mutant SMYD3
expressioon vectors in dose dependent manner. Luciferase acitivy was assayed
with a following the protocol provided by Promega. The cells were lysed directly
in the plates by 200 ul of lysis buffer. Luciferase activity was measured on 100 ul
of lysate aliquots after injection of 100 ul of luciferase detection solution. Three
independent assays were done in duplicate.

Chromatin immunoprecipitation. ChIP experiments were performed with MCF-
7 cells according to the procedure described by Kim et al after estrogen
treatment (100 nM) (Kim et al., 2001). The immunoprecipitated DNA was
amplified by qPCR using the following primers: 5’-

22

GGCCTCCTTAGGCAAATGTT-3’ (pS2 forward), 5’-
CCTCCTCTCTGCTCCAAAGG-3’ (pS2 reverse), 5’-
GCCACAGGCAGCTTTAGTTC-3’ (CTSD forward), 5’-
CATTCACAGCCTCCACCTTT-3’ (CTSD reverse), 5’-
TGTGCTCAGTGACCCTTGTG-3’ (GREB1 forward) and 5’-
CTGCCCCAACAACTGAAAGA-3’ (GREB1 reverse). 5’-
CCATCATGCTGAAGTCAGTG-3’ (PS2 Upstream forward) 5’-
GTGAGTATCTTTCAGAAGATG-3’ (PS2 Upstream reverse) 5’-
CCTCACAGGTGCGTATCTCA-3’ (CTSD Upstream forward) 5’-
AGCAAGGGGTGAAAGATGGT-3’ (CTSD UPstream reverse) 5’-
TATTCCAGTGGCTGTCTTTGC-3’ (GREB1 Upstream forward) 5’-
AGGGGTCCACAGGACATGA-3’ (GREB1 Upstream reverse) An antibody for ER
was from Santa Cruz Biotechnology and antibody for SMYD3 was from Abcam.
SMYD3 shRNAs – For RNAi depletion of SMYD3, MCF-7 cells were transfected
with 3 µg of shSMYD3 by using Lipofectamine (Invitrogen). 48 h post-
transfection, mRNA was extracted using Trizol (Invitrogen), and changes in gene
expression were assessed by real time PCR. The sequences for shRNAs used
in the assays are as follows: 5’-
GATCCGCATCTACCAGCTGAAGGTGTTCAAGAGACACCTTCAGCTGGTAGA
TGTTTTTTGGAAA-3’and 5’-AGCTTTTCCAAAAAACATCTACCAGCTGAAGGT
GTCTCTTGAACACCTTCAG CTGGTAGATGCG-3’

23

CHAPTER 2 RESULTS

2.1 SMYD3 interacts with ER via distinct domains.

As a first step toward exploring a potential role of SMYD3 in ER transcription, we
analyzed the ability of SMYD3 to interact with ER N-terminal and DNA binding
domains (NTD+DBD) and ER ligand binding domain (LBD) (Fig. 2-1A). Flag-
tagged SMYD3 was incubated with equimolar amounts of GST-fused NTD+DBD
and GST-fused LBD that were immobilized on Glutathione-Sepharose beads.
After extensive washing of the beads, SMYD3 binding was analyzed by Western
blot analysis with anti-SMYD3 antibody. As shown in Figure 2-1C, SMYD3 was
able to bind to the ER LBD (lane 8), but not to the ER NTD+DBD (lane 7). The
lack of SMYD3 interaction with GST alone (lane 6) further confirmed the
specificity of the binding reactions. Reverse binding experiments using Flag-
SMYD3 immobilized on M2 affinity beads also showed the identical interaction of
ER LBD, further confirming the specific interaction between SMYD3 and ER (Fig.
2-1B). To determine whether SMYD3 is able to interact with ER in cellular
environments, immunoprecipitation was performed after transiently expressing
untagged ER and Flag-tagged SMYD3 in 293T cells (Fig. 2-1D). Cell lysates
were prepared and subjected to immunoprecipitation of ectopic SMYD3 with anti-
Flag M2 agarose beads, and the stable association of ER was analyzed by

24

Western blot analysis using anti-ER antibody. Consistent with our in vitro
interaction results, immunoprecipitation of ectopic SMYD3 resulted in co-
precipitation of ER (lane 8). To further verify cellular interaction between SMYD3
and ER in physiological conditions, we immunoprecipitated MCF-7 cell lysates
with anti-ER antibody and checked the coimmunoprecipitation of endogenous
SMYD3. As shown in Figure 1E, endogenous SMYD3 was readily detected by
Western blot analysis of anti-ER immunoprecipitates from MCF-7 cell extracts
(lane 3), but not immunoprecipitates obtained with a control IgG (lane 2). The
lack of interaction of ER with H1 further confirmed the specificity of the interaction
between ER and SMYD3 (lane 3). Collectively, these experiments demonstrate
the direct interaction of SMYD3 with ER in vitro and in vivo, which is dependent
upon ER LBD.
2.2 SMYD3 requires both NHSC and EEL motifs for its HMT activity.

It has been shown that SMYD3 can methylate histone H3 at K4 by means of its
set domain that contains NHSC and EEL motifs (Fig. 2-2A) (Hamamoto et al.,
2004; Huang et al., 2006b). In this case, however, only histone H3 was used as
a substrate to analyze the HMT activity of SMYD3, and the possible ability of
SMYD3 to methylate other core histones was not analyzed. Thus, we examined
SMYD3 HMT activity using recombinant histone octamers reconstituted with
bacterially-expressed four core histones. Recent studies showed that

25

Figure. 2-1. Direct interaction between SMYD3 and ER. A. Schematic
diagrams of the full length ERα and SMYD3. B. In vitro interaction of SMYD3
with ERα. Recombinant ER and SMYD3 and in vitro-translated LBD+DBD and
LBD were immunoprecipitated with anti-Flag M2 agarose beads and analyzed by
Western blot analysis with indicated antibodies. Asterisks (*) indicate the purified
protein. C. Specific interaction of SMYD3 with ER LBD. In vitro translated
SMYD3 was incubated with GST, GST-NTD+DBD or GST-LBD immobilized on
glutathione beads and SMYD3 interaction was analyzed by Western blot analysis
using anti-SMYD3 antibody. D. Cellular interaction of SMYD3 with ER. Flag-
tagged SMYD3 and ER were transiently expressed in 293T cells, and cell
extracts were immunoprecipitated with anti-Flag antiboby.
Coimmunoprecipitation of ER was analyzed by Western blot using ER antibody.
E. Interaction between endogenous SMYD3 and ER. Cell lysates were prepared
from MCF-7 cells and subjected to immunoprecipitaion using anti-ERα antibody.
Coimmunoprecipitation of endogenous SMYD3 was determined by Western blot
using anti-SMYD3 antibody. H1 is also included as a negative control.

26

Figure. 2-1: Continued

27

recombinant SMYD3 methylates H3 to a very limited extent, whereas cellular
SMYD3 has a strong HMT activity (Hamamoto et al., 2004). These results imply
that a post-translational activation might be required to potentiate SMYD3,
possibly through a conformational change of the protein. For this reason, we
employed the Flag-tagged SMYD3 immunoprecipitated from transfected cells
and [
3
H]-labelled SAM for our HMT assays (Fig. 2-2B). In agreement with
previous reports (Hamamoto et al., 2004; Hamamoto et al., 2006), our HMT
assays showed that histone H3 is specifically methylated by SMYD3 in
reconstituted histone octamers (Fig. 2-2B, lane 4). Since the NHSC and EEL
motifs are highly conserved among HMT family members (Hamamoto et al.,
2004; Huang et al., 2006b), we next checked the possible requirement of these
two motifs for SMYD3 HMT activity. Deletion of the NHSC or EEL motif
(SMYD3-ΔNHSC and SMYD3-ΔEEL) abolished the ability of SMYD3 to
methylate histone H3 (lanes 5 and 6), exactly mirroring those reported with the
similar SMYD3 mutants in 293T cells (Hamamoto et al., 2004). Consistent with
these observations, concomitant deletions of NHSC and EEL motifs (SMYD3-
ΔNHSC&EEL) also showed no detectable methylation of histone H3 (lane 7).
To further characterize SMYD3 activity, HMT reactions were analyzed by
Western blot analysis using antibodies specifically recognizing mono-, di-, or tri-
methylation of H3-K4. Histone octamers were methylated by SMYD3 in the
presence of nonradioactive SAM, and the methylation state of histone H3 was

28

Figure. 2-2. Requirement of NHSC and EEL motifs for SMYD3 HMT activity.
A. Conserved amino acid sequences within SET domains. B. H3 targeted
activity of SMYD3. HMT assays were performed with full-length SMYD3 or
NHSC/EEL-deleted SMYD3 proteins expressed in 293T cells using ³H-SAM and
recombinant histone octamers. Recombinant SET7 was included as a control. C.
Di-/trimethylation of H3-K4 by SMYD3. HMT assays were performed as in Fig.
2B, but using cold SAM. H3-K4 methylation was determined by Western blot
with antibodies recognizing mono-/di-/trimethylation of H3-K4.

29

Figure. 2.2: Continued

30

analyzed by probing with H3-K4 mono-, di-, and trimethyl-specific antibodies. Our
Western blotting of the HMT reaction detected a high signal for di- and tri-
methylation of H3-K4 (Fig. 2-2C, lane 4). Incontrast, the reaction product
showed a very weak signal for monomethylation (lane 4). Collectively, these
data confirm the intrinsic preference of SMYD3 for di- and trimethylation of
histone H3-K4 as well as the critical requirement of NHSC and EEL motifs for
SMYD3 HMT activity.

2-3. SMYD3 functions as a co-activator for ER-mediated transcription.

Since recent microarray analyses indicate that SMYD3 is associated with the
expression of several genes (Hamamoto et al., 2004), we next evaluated its
possible role as a coactivator in ER-mediated transcription. The human breast
cancer MCF-7 cells were transfected with luciferase reporter plasmid and ER
expression vector along with SMYD3 expression vector, and luciferase activity
was measured 48 h after transfection. As expected, ER activated the expression
of a transiently transfected reporter gene in an estrogen-dependent manner (Fig.
2-3A). Remarkably, when increasing amounts of wild type SMYD3 were
expressed together with ER, the ligand-dependent activation of reporter gene
transcription was significantly enhanced (Fig. 2-3A). We also observed a modest
enhancement of ligand-independent, ER-mediated activation of reporter gene
expression induced by SMYD3 (Fig. 2-3A). Next, to check whether the NHSC

31

and EEL motifs are important for coactivator function of SMYD3, the reporter
gene assays were repeated with SMYD3 mutant constructs lacking NHSC and/or
EEL motifs. The coactivator function of SMYD3 was significantly impaired by
independent or simultaneous deletion of NHSC and EEL motifs, confirming the
requirement of these two motifs for the coactivator function of SMYD3 (Fig. 2-
3B). To confirm the above results, we also repeated the experiments using
human epithelial 293T cells; similar results were obtained from these parallel
experiments (Fig. 2-3C and 2-3D). Our studies proved the critical
requirement of NHSC and EEL motifs in SMYD3 HMT activity. A possible
explanation for the above data is that changes in the HMT activity are likely to be
responsible for the repressed transcription caused by the deletion of the NHSC
and EEL motifs. However, another possibility is that NHSC and EEL motifs of
SMYD3 are critical for the physical interaction between SMYD3 and ER; thus the
lack of these motifs might modulate downstream transcription activities. To
check this possibility, we analyzed the binding of wild type and deletion mutant
SMYD3 proteins to a fixed concentration of the ligand binding domain (LBD) and
the N-terminal and DNA binding domains (NTD+DBD) of ER. As expected from
our initial binding data (Fig. 2-1B and 2-1C), full length SMYD3 was able to bind
to GST-LBD fusion (Fig. 2-3E, lane 4), but not to GST-NTD+DBD fusion (Fig. 2-
3E, lane 3). Furthermore, identical binding assays with the deleted forms of
SMYD3 showed a comparable binding capacity of these mutant SMYD proteins
to ER (lanes 7, 10 and 13). The results from these binding experiments, together

32

Figure. 2-3. Coactivator function of SMYD3 in ER transcription. A and C.
Effect of SMYD3 on ER-mediated transcription in vivo. MCF-7 (A) and 293T
cells (C) in 12-well plates were transiently transfected with MMTV-ERE reporter
gene together with ER expression vector and various amounts of SMYD3
expression vectors in uninduced and estrogen induced conditions as indicated.
The representative data from three independent experiments are shown, and the
error bars indicate as the means ±S.E. B and D. Requirement of HMT activity for
SMYD function. Reporter gene assays were as in Fig. 2-3A and 2-3C, but with
mutant SMYD3 expression vectors. E. In vitro interaction of ER with SMYD3
lacking NHSC/EEL motifs. ER NTD+DBD and LBD fused to GST were incubated
with SMYD3 lacking NHSC/EEL motifs, and SMYD3 binding was analyzed by
immunoblot with anti-SMYD3 antibody.

33

Figure. 2-3: Continued

34

with the results from our HMT assays, strongly suggest that inactivation of SMYD3 HMT
activity, not changes in SMYD3-ER interaction, and are the major cause of impaired
transcription caused by the deletion of NHSC and EEL motifs.

2-4. SMYD3 augments ER target gene expression via histone H3-K4
methylation.

Having shown the ability of SMYD3 to coactivate ER-mediated transcription, we
sought to determine the participation of SMYD3 in transcription of three
endogenous ER target genes. Thus, MCF-7 cells were treated with 100 nM of
estrogen for 0, 30, 60, 90, and 120 min, and the recruitment of ER and SMYD3 to
the promoters of three target genes (pS2, CTSD and GREB1) were examined by
chromatin immunoprecipitation (ChIP) (Fig. 2-4). The presence of the promoter
regions in the chromatin immunoprecipitates was analyzed by quantitative real-
time PCR using specific pairs of primers spanning the estrogen responsive
regions in the promoters. Immunoprecipitation with the ER-specific antibody
showed an obvious increase with respect to ER occupancy of the target gene
promoters within 30 min of estrogen treatment (Fig. 2-4, ER). ER promoter
occupancy was significantly declined and returned to baseline after 60 min of
estrogen treatment. The second phase of ER recruitment to the promoter was
evident after 90 min of estrogen treatment. These results are consistent with

35

previous indications that ER acts in a transient but repeated fashion on the same
target gene promoter following estrogen stimulation (Shang et al., 2000).

We next investigated the promoter recruitment of SMYD3 and the relationship of
this process to the ER occupancy and H3-K4 methylation state at the three
promoters following estrogen treatment. SMYD3 showed its accumulation on the
promoters to a very high level at the 30 min point, but showed an apparent
dissociation from the promoters at the 60 min point (Fig. 2-4, SMYD3). The
second cycle of promoter occupancy began at 90 min point and returned to the
baseline level at 120 min point. Parallel analysis over the same time period
showed that the level of di- and tri-methyl K4 of histone H3 peaked at 30 min and
90 min time points (Fig. 2-4, H3K4me2 and H3K4me3). In contrast, we did not
see any apparent change in mono-methylation of H3-K4 after estrogen treatment
(H3K4me1). In addition, SMYD3 recruitment and H3-K4 methylation were
targeted by ER to the ERE region, because their localizations were
minimal/undetectable in a region ∼2 kb uptream of the ERE site (Fig. 2-4,
Upstream). The coincidental appearance and decrease of SMYD3 and di-/tri-
methyl H3-K4 strongly suggest that the accumulation of di-/tri-methyl H3-K4 is
due to the enhanced recruitment of SMYD3 following the onset of estrogen
treatment. Moreover, a very similar timing of SMYD3 occupancy and the
corresponding H3-K4 methylation in all three target genes strongly supports that

36

Figure. 2-4. Estrogen-induced accumulation of SMYD3 and H3-K4
methylation at ER target genes. MCF-7 cells were cultured under estrogen-
deprived conditions for 3 days and subjected to ChIP analysis after treating with
estrogen for 0, 30, and 60, 90 and 120 min. ChIP assays were performed using
antibodies specifically recognizing ER, SMYD3, and H3-K4 mono-/di-/tri-
methylation. Input DNA and immunoprecipitated DNA were quantified by real-
time PCR using specific primer sets as indicated. The results are shown as
percentage of input. The error bar indicates as the means ±S.E.

36

37

Figure. 2-4: Continued

38

Figure. 2-4: Continued

39

Figure. 2-4: Continued

39

40

ER employs similar dynamics to recruit SMYD3 for H3-K4 methylation. These
studies thus confirm that ER can recruit SMYD3 to its natural target genes and
mediate de novo methylation of H3-K4 in the process of gene transcription.

2-5. The functional role of SMYD3 in ER-mediated transcription.

To assess the requirement of SMYD3

in ER transcription activity in vivo, we
examined the effect of manipulating SMYD3 levels on ER target gene expression
in MCF-7 cells. We generated a short hairpin RNA (shRNA) construct for
SMYD3 and tested its knockdown effects on SMYD3 expression. The RT-PCR
confirmed that cell transfection with SMYD3

shRNA

decreased the SMYD3
mRNA level by 75% in the absence of E2 and 50% in the presence of E2 (Fig. 2-
5B, SMYD3). The reduced protein level was also confirmed by Western blot
analysis of endogenous SMYD3 using anti-SMYD3 antibody (Fig. 2-5A). This
shRNA-induced silencing of SMYD3 gene significantly reduced the ligand-
induced expression of pS2, CTSD and GREB1 genes compared with negative
control (non-target shRNA) cells (Fig. 2-5B). Albeit the expression

of ER target
genes was much weaker without estrogen treatment, we also could detect a
moderate reduction in ER transcription activity after SMYD3 shRNA transfection
in this uninduced condition (Fig. 2-5B, pS2, CTSD, GREB1).
Figure. 2-5. Requirement of SMYD3 for ER transcription. A. Validation of
SMYD3 knockdown. MCF-7 cCells were transfected with SMYD3 shRNA or
control shRNA, and expression of SMYD3 protein was checked by Western
blotting of whole-cell lysates using anti-SMYD3 antibody. β-actin was used as an
internal control (Lower panel). B. Repression of ER transcription by SMYD3
knockdown. CMCF-7 cells were transfected with SMYD3 shRNA, and treated
with 100 nM estrogen 24 h post-transfection. mRNA levels were analyzed by
real time PCR . Error bars indicates as the means ± S.E. from the experiments
performed in duplicate, and the experiments were repeated three times.

41

42

Figure 2-5.: Continued

43

2-6. SMYD3 is responsible for the level of H3-K4 mehylation in ER-target
genes.

To furher investigate the effect of SMYD3 knockdown on promoter occupancy of
H3-K4 methylation, we performed ChIP assays on PS2 gene with SMYD3-
depleted and control MCF-7 cells following E2 treatment (Fig. 2-6A). As
expected, SMYD3 depletion resulted in a dramatic reduction in
immunoprecipitation of the ERE region using SMYD3 antibody (Fig. 2-6B,
SMYD3). However, the depletion had no obvious effect on the cyclic promoter
occupancy of ER after the addition of E2 (Fig. 2-6B, ER). Importantly, similar
ChIP analysis showed that RNAi-mediated depletion of SMYD3 distinctly
diminished promoter-targeted accumulation of di-/tri-methylation of H3-K4 (Fig. 2-
6B, H3K4me2 and H3K4me3). In contrast, the depletion had minimal effect on
the level of mono-methylation of H3-K4 at pS2 promoter upon E2 treatment (Fig.
2-6B, H3K4me1), indicating that SMYD3 is not required to establish H3-K4
mono-methylation. Taken together, these data indicate that endogenous SMYD3
plays an important role in the augmentation

of ER-mediated transcription in the
ligand-treated environment.

44

Figure. 2-6. Reduction of H3-K4 methylation upon SMYD3 depletion. A,
Western blot analysis of SMYD3 knockdown. MCF7 cells were transfected with
SMYD3 shRNA for 24 h and subjected to Western blot analysis after E2
treatment for 0, 30, 60, 90, and 120 min. β-actin was used as a loading control
(Lower panel). B. Inhibition of H3-K4 methylation at pS2 gene upon SMYD3
knockdown. ChIP assays were performed using antibodies against ER, SMYD3
and mono-/di-/tri-methylated H3-K4.as in Fig. 2-4, but after SMYD3 knockdown.

45

Figure. 2-6: Continued

46

CHAPTER 2 DISCUSSION

In this

study, we investigated a possible role of SMYD3 histone methyltransferase
in activating ER target genes. The present data demonstrate that (i) SMYD3
physically interacts with ER both in vitro and in vivo, (ii) SMYD3 acts as a
transcriptional coactivator of ER that enhances ER-mediated transcription, (iii)
SMYD3-imparted transactivation correlates with SMYD3 recruitment and H3-K4
methylation at ER target genes, and (iv) SMYD3 knockdown significantly reduces
the ligand-induced expression of ER target genes. These results reveal an
essential role of SMYD3 in modulating ER-mediated transcription and provide an
example of epigenetic regulation of ER function. The ability of ER to activate
transcription requires the repeated cycling of various coregulators onto its target
gene promoters in the presence of continuous
stimulation by estrogen (Metivier et al., 2003; Shang et al., 2000). Currently, at
least two mechanistic models have been proposed

to describe the function of
these coregulators. First, they

transmit the signal of ligand-induced ER
conformational change to the basal transcription machinery (Lemon and
Freedman, 1999). Second, they are associated with targeted chromatin
remodeling by ER (Belandia and Parker, 2003). Recent biochemical

and genetic
studies support that methylation of histone H3 at K4 is characteristic to gene
activation, and removal of this modification is involved in transcriptional

47

repression (Lachner and Jenuwein, 2002; Lachner et al., 2001; Rea et al., 2000).
The data presented here demonstrate that SMYD3, through its HMT activity,
plays a significant role in dictating the transcriptional activity of ER. That the
effects of SMYD3 were found to be dependent upon the ability to interact with ER
LBD implies that SMYD3 is a functionally important component of estrogen-
stimulated ER transcription. Combined with the observation that shRNA-induced
silencing of SMYD gene inhibits ER target gene expression, these results argue
strongly in favor of SMYD3 as an integral component of the ER response. While
our analyses have been restricted to ER-dependent function of SMYD3, previous
studies indicated that activation of other nuclear receptors also involves H3-K4
methylation (Dreijerink et al., 2006; Guccione et al., 2007). As such, elucidation
of a possible role of SMYD3 in the promoter-localized H3-K4 methylation and the
consequent activation of transcription at other nuclear receptor target genes is an
important issue that warrants further investigation.
Many SET domain-containing

proteins with HMT activity harbor two conserved
amino acid sequence called NHSC and EEL motifs (Hamamoto et al., 2004;
Huang et al., 2006b). SMYD3 also possesses

these motifs within its SET
domain, and they have been shown to be critical for SMYD3 enzymatic

activity in
HMT reaction (Hamamoto et al., 2004). Our studies performed with SMYD3
mutants demonstrated that, although the NHSC and EEL motifs within set
domain are essential for SMYD3 HMT activity, these motifs are dispensable for
SMYD3 binding to ER (Fig. 2-3E). Importantly, when these deletion mutants

48

were checked in ER luciferase

reporter assays, they failed to show coactivator
function (Fig. 2-3A-D). A simple interpretation of these results is that HMT
activity of SMYD3 is required for

its action on the transactivation of reporter gene
following estrogen stimulation. These characteristics ascribed to NHSC and EEL
motifs fit very well with the generic properties previously assigned to the
NHSC/EEL motif-containing set domains in regulating and mediating the
enzymatic activity of HMT proteins (Hamamoto et al., 2004; Huang et al., 2006a).
These results also suggest that NHSC and EEL motifs of SMYD3 could provide
the molecular

target for regulation of H3-K4 methylation-dependent
transcriptional

responses by ER. More thorough domain mapping and
mutagenesis experiments

will be required to provide further insights into the ER-
SMYD3

interactions, which facilitate SMYD3 recruitment and H3-K4 methylation
in ER-mediated transcription.

We have shown that ER promoter occupancy upon E2 treatment coincides with
promoter recruitment of SMYD3 and appearance of di-/tri-methyl H3-K4 (Fig. 2-
4A-C). This similar timing of ER occupancy and SMYD3 recruitment strongly
supporting that H3-K4 methylation per se endows coactivator properties of
SMYD3 in regulating ER-mediated transcription. In further support of a
regulatory role of SMYD3-mediated H3-K4 methylation, SMYD3 depletion
showed a significant effect on the level of di-/tri-methylation, but not mono-
methylation, of H3-K4 at the promoter of pS2 gene upon E2 treatment (Fig. 2-6).

49

However, it is also possible that SMYD3 augments the activities of other
transcription components at the initial stage of gene induction. In fact, a recent
study demonstrated that SMYD3 interacts with an RNA helicase to form a
complex with RNA polymerase II (Hamamoto et al., 2004). Hence, SMYD3 could
act as a "bridge" protein that mediates functional interaction between ER and
RNA Pol II to coordinate the tightly integrated

processes of chromatin remodeling
and transcription in ER-driven

transcription (Rachez et al., 1999; Torchia et al.,
1998). Furthermore, increasing evidence suggests that coactivator proteins such
as p300, PRMT1 and CARM1 act together as potential regulators of ER-induced
transcription (Chen et al., 2000; Daujat et al., 2002; Huang et al., 2003; McKenna
et al., 1999; Spencer et al., 1997). Thus, it is possible that a distinct group of
coactivators play a crucial role in orchestrating ER-mediated transcription. In this
respect, it will be interesting to determine the relative contribution of SMYD3,
together with other coactivators that associate with ER target gene expression.

In conclusion, the analysis of SMYD3 described here establishes its role in ER-
mediated transcription as a coactivator. H3-K4 methylation appears to be critical
for SMYD3 function in ER transcription. Identifying cofactors that influence ER
has been important in the development of effective

therapies and prevention of
breast cancer. Since our studies provide clues as to the requirement of SMYD3
HMT activity for its function, the development of HMT inhibitors may be of
therapeutic

benefit in modulating SMYD3 action in ER transcription.

50

CHAPTER 3: Histone variant H3.3 stimulates HSP70
transcription through cooperation with HP1γ.

CHAPTER 3 INTRODUCTION

The dynamic nature of chromatin is functionally important for the regulation of
diverse DNA-dependent processes in the nucleus including transcription,
replication, and DNA repair. While the mechanisms are not well understood,
nucleosome remodeling activities modulate the functional state of chromatin. In
addition to ATP-dependent nucleosome remodeling and covalent histone
modifications, deposition of histone variants into the nucleosome is thought to be
critical in transcriptional regulation (Henikoff, 2008; Kouzarides, 2007). H3.3 is
the predominant form of histone H3 variants, which differs by four amino acids
from the replication dependent histones H3.1 and H3.2, (generally referred

to as
H3) (Albig et al., 1995; Wells et al., 1987). The H3.3 variant is expressed
throughout the cell cycle and incorporated into chromatin in DNA replication-
independent manner whereas histone H3 is primarily expressed and
incorporated during S phase (Tagami et al., 2004). In human cells, replication-
independent deposition of H3.3 is mediated by the HIRA chaperone and the
ATRX chromatin remodeler through a mechanism distinct from that of replication-
coupled deposition of H3 by the CAF-1 (Drane et al.; Goldberg et al.). H3.3 is

51

preferentially distributed over the promoter regions and its accumulation
coincides with higher levels of H3-K4 methylation and bound RNA polymerase II
(Chow et al., 2005; Jin et al., 2009; McKittrick et al., 2004; Mito et al., 2005).
Further support for such a transcription-coupled H3.3 deposition is provided by
studies in Drosophila demonstrating that the heat shock-induced transcription of
HSP70 genes coincides with the replacement of H3 with H3.3 (Sakai et al., 2009;
Schwartz and Ahmad, 2005). In addition to its enrichment over active genes, a
genome-wide analysis of H3.3 distribution indicated that a fraction of H3.3 also
localizes to constitutive heterochromatin regions at telomeres (Goldberg et al.;
Wong et al.).

Another group of proteins specifically marking chromatin state consists

of
heterochromatin

protein 1 (HP1). Mammalian cells possess three closely related
isoforms of HP1 based on their size and amino acid sequence similarity (Singh et
al., 1991). HP1α primarily localizes to pericentric heterochromatin, HP1β binds to
both heterochromatin and euchromatin, and HP1γ exclusively targets
euchromatin (Dialynas et al., 2007; Inoue et al., 2008; Serrano et al., 2009). All
these isoforms have been originally characterized as regulatory proteins that
establish an inactive state of chromatin, but recent studies have challenged this
view (Hediger and Gasser, 2006; Hiragami and Festenstein, 2005). One of the
first clear indications that HP1 acts as a positive regulator came from a previous
work demonstrating that H3K9me and HP1γ are enriched at the coding regions

52

of actively transcribed genes (Vakoc et al., 2005). An activating role of HP1 in
transcription was further supported by recent studies showing that HP1γ
associates with the Survivin gene in its active state (Smallwood et al., 2007).
Another notable finding is that Drosophila HP1c (HP1γ homolog) stimulates
transcription elongation by bridging the interaction between histone chaperone
FACT and phosphorylated RNA polymerase II (Kwon et al.). All these results
suggest that, besides its most commonly cited role in heterochromatin formation,
HP1 can trigger an active chromatin structure once it is targeted to specific genes
within euchromatin.

Of special relevance to the present study, the rapid incorporation of H3.3 and the
recruitment of HP1c have been shown to play a role in controlling transcription of
the HSP70 genes under heat shock condition in Drosophila (Schwartz and
Ahmad, 2005). These findings raise a question about whether H3.3 and HP1γ
are also required for higher rates of transcription at induced HSP70 genes in
human cells and if so, how their effects are generated. In this study, we
addressed these questions under relevant conditions by monitoring independent
and cooperative actions of H3.3 and HP1γ in HSP70 gene transcription. We
found that H3.3 and HP1γ are co-localized at HSP70 promoters and establish
transcriptional competence in response to heat shock. Detailed investigation of
the underlying mechanism revealed a selective connection between HP1γ
localization and active histone modifications enriched in H3.3 nucleosomes.

53

Furthermore, HP1γ and H3.3 are functionally interdependent as RNA
interference (RNAi)-mediated depletion of either HP1γ or H3.3 inhibits HSP70
transcription and cell proliferation.

CHAPTER 3 MATERIALS AND METHODS

Cell culture, Antibodies and Constructs. HeLa and MCF-7 cells were grown in
Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal
bovine serum (FBS) and 1% penicillin/streptomycin.

Antibodies used in this study
are as follows: H3ac, H3K9me3 and H3.3 antibodies from Abcam, IgG antibody
from Santa Cruz Biotechnology, Flag antibody from Sigma, HP1α, HP1β, HP1γ
antibodies from Millipore, H3K4me3 antibody from Active Motif and H3K27me3
antibody from Dr. J. Rice. For mammalian expression of H3 and H3.3, their
cDNAs were PCR-amplified and ligated into pIRES mammalian vector in frame
with 5’ Flag and HA sequences.

Nucleosome purification. HeLa cells were transfected with expression vectors
for Flag- and HA-tagged versions of human H3 and H3.3. 48 h post-transfection,
cells were harvested and lysed with buffer A (20 mM HEPES, pH 7.4, 10 mM
KCl, 1.5 mM MgCl
2
, 0.34 M sucrose, 10% glycerol, 1 mM dithiothritol, and
protease inhibitor cocktail) containing 0.2% Triton X-100. Nuclei were pelleted by

54

centrifugation at 1,000 g, resuspended in buffer A containing 2 mM CaCl
2
and
digested with 0.6 U microccocal nuclease (MNase, Sigma) at 37°C for 28 min.
Digested nuclei were collected and incubated in nuclear extraction buffer (20 mM
HEPES, pH 7.4, 420 mM NaCl, 1.5 mM MgCl
2
, 0.2 mM EGTA, and protease
inhibitor cocktail) for 1 h, and centrifuged to remove nuclear debris. After
adjusting the salt concentration of the extract to 150 mM NaCl, ectopic H3/H3.3-
containing nucleosomes were isolated by two consecutive immunoprecipitation
using anti-Flag and anti-HA antibodies (Sigma) in washing buffer (20 mM
HEPES, pH 7.8, 300 mM NaCl, 1.5 mM MgCl
2
, 0.2 mM EGTA, 10% Glycerol,
0.2% Triton X-100, and protease inhibitor cocktail). Bead-bound nucleosomes
and proteins were analyzed by Western blot analysis.

Immunofluorescence. Heat shock-treated HeLa cells were fixed in freshly
prepared 4% formaldehyde solution at room temperature. Cells on cover slips
were permeabilized in Solution P (1% BSA and 0.2% Triton X-100 in PBS) for 10
min, washed with PBS, and blocked with 10% BSA in PBS. Cells were incubated
in stepwise with a primary antibody overnight and a secondary fluorescent
antibody for 1 h at 25°C. Images were captured on a Zeiss ApoTome Axiocam
MRc microscope equipped with a 60X oil-immersion lens. Image processing was
performed with Photoshop.

55

qRT-PCR, Chromatin Immunoprecipitation (ChIP), and shRNA. For
quantitative reverse transcription PCR (qRT-PCR) analysis of HSP70 gene
expression, total RNA was isolated using the Trizol reagent (Invitrogen) following
the manufacturer’s instructions. RNA was converted to cDNA using iScript cDNA
Synthesis Kit (Bio-Rad), and gene expression was assessed by using the IQ
SYBR Green Supermix (Bio-Rad) and the MYiQ2 real time cycler (Bio-Rad).
Assays were normalized to glyceraldehyde-3-phosphate dehydrogenase
(GAPDH) mRNA levels. The primers used quantitative real time PCR (qPCR) are
listed in the Supplementary data (Table S1). ChIP assays were performed
essentially as described (Kim et al., 2009). DNA was recovered from heat/mock-
treated HeLa cells by PCR purification column kit. Input DNA (1%) was used for
normalization. Primers used in this study are listed in Supplementary data (Table
S2). The sequences encoding the short hairpin RNAs (shRNA) for this study are
listed in Supplementary Data (Table S3). The pLKO vectors containing specific
shRNA were constructed according to the manufacturer's instructions (Open
Biosystems). All the constructs were verified by DNA sequencing. For
knockdown experiments, cells (1X10
5
) were transfected with shRNA expression
constructs using Lipofectamine reagent (Invitrogen) and selected with puromycin
(5µg/ml) for 2 weeks.

In vitro binding assay. Flag-tagged HP1 proteins (2 µg) were immobilized on
M2 agarose beads and incubated with unmodified/modified H3/H4 tail peptides

56

(5 µg) in binding buffer (10 mM Tris-HCl, pH 7.8, 1 mM EDTA, 10% glycerol, 150
mM KCl, and 0.02% Nonidet P-40) for 6 h at 4°C. After washing beads three
times with binding buffer, bound tail peptides were resolved on 4-20% gradient
SDS-PAGE and detected using Silver Stain Plus (Bio-Rad).

MTT and colony formation assays. Cell proliferation was assessed by the 3-
(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay
according to the manufacturer’s instructions (Calbiochem). Briefly, HP1γ/H3.3-
depleted MCF7 cells were incubated with 1 ml of MTT (5 mg/ml) for 3 h at 37°C.
The MTT formazan precipitate was dissolved with 1 ml of MTT solvent and
cellular proliferation was

determined from the conversion of MTT to formazan
using a Microplate Reader Model 680 (Bio-Rad) at a wavelength of 570 nm with
background subtraction at 650 nm. Experiments were done in triplicate. For soft
agar colony formation assay, MCF-7 cells (1 X 10
5
) were suspended in semisolid
medium (DMEM 10% FBS plus 0.33% ultra pure noble agar). The mixture was
added over a layer of 0.8% agar in DMEM on 60 mm plate and incubated at 37°C
in a humidified 5% CO
2
atmosphere. Colonies were counted after three weeks.
All assays were run in triplicate, and results presented are the average of three
individual experiments.

57

CHAPTER 3 RESULTS
3-1. H3.3 and HP1γ are co-enriched at HSP70 promoters

To study the interaction of HP1 isoforms with nucleosomes containing H3.3 or
H3, we transiently expressed Flag-HA tagged versions (f-h) of H3.3 and H3 in
HeLa cells (Supplementary Figure 3-S1B). Soluble chromatin fragments were
prepared from cell nuclei of the transfected cells and digested with micrococcal
nuclease (MNase) to yield mainly mononucleosomes, as confirmed by gel
electrophoresis of nucleosomal DNA (Figure 3-1A, lanes 3 and 4). The
mononucleosomes were subjected to double immunoprecipitations with anti-Flag
and anti-HA antibodies to selectively purify mononucleosomes containing f-h-
H3.3 or f-h-H3 (Supplementary Figure 3-S1A). As expected, almost equimolar
levels of H2A, H2B, H4, and endogenous/ectopic H3.3/H3 were found in purified
nucleosomes (Figure 3-1A, lanes 6 and 7). Western analyses with H3 antibody
recognizing both H3 and H3.3 confirmed that similar levels of ectopic H3 and
H3.3 are present in purified nucleosomes (Figure 3-1B, α-H3). In addition, our
analyses using H3.3 specific antibody showed only trace amounts of
endogenous untagged H3.3 in H3.3-containing nucleosomes (α-H3.3, lane 3),
indicating that the lower band detected by Western blotting with H3 antibody (α-
H3, lane 3) mainly contains endogenous H3. Next, we examined whether any
HP1 isoforms are associated with H3.3 nucleosomes by Western blotting using

58

Figure. S3-1. Preparation of H3/H3.3 nucleosomes. (A) HeLa cells were
transfected with Flag-HA-H3 (f-h-H3) or -H3.3 (f-h-H3.3) constructs for 48 h. After
shearing the nuclei to solubilize chromatin, mononucleosomes were prepared by
micrococcal nuclease digestion of the soluble chromatin fractions. Ectopic
H3/H3.3-containing mononucleosomes and their interacting proteins were
sequentially immunoprecipitated with anti-Flag and anti-HA antibodies. (B)
Expression levels of ectopic H3 and H3.3 were assessed by Western blots of cell
lysates using anti-Flag antibody.

59

Figure. S3-2. Immunostaining of HP1 isoforms. Cells were immunostained
with HP1α and HP1β antibodies (green channel) and H3.3 antibody (red
channel). The nuclei were also stained with DAPI (blue).

60

Figure. S3-3. Co-localization of ectopic H3.3 and HP1γ at HSP70 promoters.
(A) Cells were transfected with Flag-H3.3 (f-H3.3) for 48h and treated with heat
shock at 42°C for 30 min and then, analyzed by ChIP analysis on HSPA1 and
HSPA6 promoters as described in Figure 1D. (B) FLAG-tagged versions of HP1
isoforms were expressed in cells, and Chip assays were performed as in Figure
1D.

61

antibodies specific to three HP1 isoforms. Our results indicated a stable
association of HP1γ, but weak or undetectable associations of HP1α and HP1β
with H3.3 nucleosomes (α-HP1α/HP1β/HP1γ, lane 3). The minimal association of
all HP1 isotypes with H3 nucleosomes also confirmed the specificity of the HP1γ-
and H3.3 nucleosome interaction (α-HP1α/HP1β/HP1γ, lane 2). To further
support the observed interaction, we

conducted immunofluorescence imaging of
endogenous H3.3 and HP1γ proteins in HeLa cells. As depicted in Figure 3-1C
and Supplementary Figure 3-S2, double immunostaining of H3.3 and HP1γ
proteins

showed a significant colocalization of H3.3 with HP1γ, but not with HP1α
and HP1β in the cell nucleus.

Having shown the ability of HP1γ to selectively bind to H3.3 nucleosomes, we
sought to determine whether this interaction is related to their cellular functions.
Recent studies demonstrated that HP1γ and H3.3 are associated with heat
shock-induced transcription of HSP70 genes in Drosophila (Sakai et al., 2009;
Schwartz and Ahmad, 2005). Thus, we first examined the localization of H3.3
and HP1γ at two HSP70 genes, HSPA1 and HSPA6, in HeLa cells after heat
shock by chromatin immunoprecipitation (ChIP). Two sets of primers were
employed to detect crosslinking of H3.3 and HP1 proteins in the promoter and
transcribed regions by qPCR. As shown in Figure 3-1D, heat shock treatment
resulted in a rapid accumulation of H3.3 protein in the promoter regions of
HSPA1 and HSPA6 genes (α-H3.3). On the contrary, H3.3 was minimally

62

localized in the coding region and showed little change in its level after heat
shock treatment (α-H3.3). In exploring the alteration of HP1γ occupancy, we also
found that HP1γ level was significantly increased in the promoter region (~10-
fold) but only modestly in the coding region during the activation process (α-
HP1γ). Remarkably, however, parallel ChIP assays with anti-HP1α and anti-
HP1β antibodies repeatedly demonstrated no detectable accumulation of HP1α
and HP1β in the promoter and coding regions in response to heat shock (α-HP1α
and α-HP1β). To further confirm the above results, the ChIP assays were
repeated using cells transiently transfected with Flag-tagged versions of H3.3,
HP1α, HP1β, and HP1γ; similar results were obtained with these ectopic proteins
(Supplementary Figure 3-S3). These results argue strongly against the possibility
that H3.3 and HP1γ antibodies cross-react with other proteins, and indicate that
the distribution of H3.3 and HP1γ proteins in the cell is indeed accurately
established.

3-2. Both H3.3 and HP1γ are required for HSP70 transcription.

Heat shock-induced enrichments of H3.3 and HP1γ described above raised the
possibility that they are required for transcription of HSP70 genes. To check this
possibility, we first depleted H3.3 and HP1γ individually by a vector-based RNA
interference (RNAi) technique. Western blot and qRT-PCR analyses confirmed

63

Figure. 3-1. Selective interaction of HP1γ with H3.3 nucleosomes. (A) HeLa
cells were transfected with control (lane 2), histone H3 (lane 3), and H3.3 (lane 4)
expression vectors for 48 h, and mononucleosomes were prepared as
summarized in Supplementary Figure 3-S1A. Total nucleosomal DNA was
subjected to 2% agarose gel electrophoresis and visualized with ethidium
bromide (lanes 2-4). Mononucleosomes containing H3 or H3.3 were purified by
sequential immunoprecipitations using anti-Flag and anti-HA antibodies, and
histone compositions of the purified nucleosomes were analyzed by Coomassie
staining following 15% SDS-PAGE (lanes 6 and 7). Lane 1, 0.1-12 kb DNA
ladder; lane 5, control mock-purified material. (B) Proteins co-purified with H3
and H3.3 nucleosomes were separated by 15% SDS-PAGE, and the presence of
indicated proteins was analyzed by Western blotting. Lane 1, mock-purified
material; lane 2, H3 mononucleosomes; lane 3, H3.3 mononucleosomes. (C)
HeLa cells were immunostained for HP1γ (green channel), H3.3 (red channel),
and DAPI (blue channel). Image overlays show colocalization between HP1γ and
H3.3. (D) HeLa cells were mock-treated (-) or heat-treated (+) for 30 min, and
quantitative chromatin immunoprecipitation (ChIP) assays of promoter (■) and
coding ( ) regions of HSPA1 and HSPA6 genes were performed using antibodies
specifically recognizing H3.3, HP1α, HP1β, and HP1γ. IgG antibody was used as
a negative control. Input DNA and immunoprecipitated DNA were analyzed by
qPCR analyses using primer sets depicted in the top panel. The results are
shown as percentage of input. The error bar indicates the means ± S.D.

64

Figure. 3-1: Continued

65

that transfection of HeLa cells with shRNAs against H3.3 and HP1γ efficiently
depleted their responsive target proteins (Figures 3-2A and 3-2C). When qRT-
PCR was performed, we detected a distinct reduction in transcription of HSPA1
and HSPA6 genes upon shRNA-mediated silencing of HP1γ (Figure 3-2B).
Similarly, upon the knockdown of H3.3, there was a substantial decrease in
transcription (Figure 3-2D). On the other hand, the knockdown of H3.3 or HP1γ
did not change transcription of a control gene, Actin (Figures 3-2B and 3-2D).
Considering the possibility that H3.3 and HP1γ could act together to regulate
HSP70 genes, we also checked whether double knockdown of H3.3 and HP1γ
exhibits more severe defect in transcription (Figure 3-2E). Interestingly, however,
the simultaneous knockdown of H3.3 and HP1γ attenuated HSP70 transcription
to a similar level as that observed in individual knockdown of H3.3 or HP1γ
(Figure 3-2F). These data constitute a powerful argument that both H3.3 and
HP1γ are indeed essential for the transcriptional activation of HSP70 genes
upon heat shock.

3-3. H3.3 and HP1γ are interdependent for their localization at HSP70
promoters.

To gain support for the knockdown results above, we next checked whether the
depletion of H3.3 or HP1γ is directly attributable to their promoter occupancy. As

66

Figure. 3-2. . Requirements for H3.3 and HP1γ in HSP70 transcription. (A)
HeLa cells were stably transfected with shRNAs targeting to nonspecific control
or HP1γ. The efficiency and selectivity of knockdown were determined by
Western blotting (left panel) and qRT-PCR (right panel). (B) Mock-depleted (NC
shRNA) or HP1γ-depleted (HP1γ shRNA) HeLa cells were heat shocked as in
Figure 3-1D. Total mRNA was purified and subjected to qRT-PCR to examine
transcription levels of HSPA1 and HSPA6 genes. All transcription levels were
quantified relative to uninduced conditions and normalized to that of GAPDH.
Average and standard deviation of three independent experiments are shown. (C
and D) Cells were transfected with shRNA targeting H3.3 or irrelevant control,
and Western blotting and qRT-PCR analyses were performed as described in
Figure 3-2B. Note that H3.3 protein is encoded by two different genes, H3.3A
and H3.3B. (E and F) H3.3 and HP1γ were simultaneously depleted, and
Western blotting and qRT-PCR analyses were essentially as in Figure 3-2B.

67

Figure. 3-2: Continued

68

expected, the mock-depleted cells showed an apparent increase of HP1γ and
H3.3 at HSPA1 and HSPA6 promoters after heat shock treatment (Figures 3-3A
and 3-3B, NC shRNA). When ChIP experiments were performed using HP1γ-
depleted cells, near complete loss of HP1γ was detected at both promoters
(Figure 3-3A, upper panel). Unexpectedly, however, HP1γ-depleted cells also
showed a significant reduction in promoter occupancy of H3.3 (α-HP1γ, H3.3
shRNA) (Figure 3-3A, lower panel). This observation indicates that the heat
shock-stimulated incorporation of H3.3 into the promoter nucleosomes is
dependent on the presence of HP1γ. Analogously, when ChIP experiments were
repeated using H3.3-depleted cells, a decreased level of H3.3 in the promoter
nucleosomes coincided with a base line level of promoter-bound HP1γ (Figure 3-
3B), suggesting that H3.3 is necessary component for the promoter localization
of HP1γ. Because HSPA1 and HSPA6 promoters are minimally occupied by
H3.3 and HP1γ in the absence of heat shock, knockdown of H3.3 or HP1γ failed
to show any apparent alteration and interdependency in their promoter
occupancy under this normal condition. We also repeated the entire experiments
in cells transiently transfected with Flag-tagged H3.3 and HP1γ; similar results
were obtained from these experiments (Supplementary Figure 3-S4).

69

Figure. S3-4. Interdependent promoter occupancy of ectopic H3.3 and
HP1γ. H3.3-depleted and HP1γ-depleted cells were transfected with ectopic
H3.3 and HP1γ, then ChIP assays were performed using H3.3 and HP1γ
antibodies after heat-shock as in Figure 3-3.

70

Figure. 3-3. Interdependent localization of H3.3 and HP1γ at HSP70
promoters. (A) Mock-depleted (■) and H3.3-depleted ( ) cells were heat-treated
as in Figure 1D, and ChIP assays of HSP70 promoter regions were performed
using antibodies against H3.3 and HP1γ. (B) ChIP assays were essentially
identical to Figure 3A, but in H3.3-depleted cells.

71

3-4. Active histone marks are enriched at promoter-positioned
nucleosomes.

One possible explanation for the observed colocalization of HP1γ and H3.3 is
that H3.3 nucleosomes are enriched with specific histone modifications to
promote a stable binding of HP1γ. To investigate this possibility, we prepared
nucleosomes containing f-h-H3 or f-h-H3.3 as described in Figure S1A and
determined the levels of histone methylation and acetylation by Western blotting.
Because H3 and H3.3 are identical across the N-terminal domain, all antibodies
specific for H3 modifications should recognize H3.3 modifications with the same
affinity. Given that only trace amount of endogenous H3.3 was detected in
purified nucleosomes (Figure 3-1B, α-H3.3), our Western blot analysis mainly
evaluated the modifications of ectopic H3.3. In determining the degree of
methylation, we found much higher levels of di- and tri-methylation of ectopic
H3.3 lysine 4, compared to those of ectopic H3 lysine 4 (Figure 3-4A, lanes 2 and
3, α-H3K4me2/H3K4me3). This was not due to a general

preference of H3.3 for
HMTs present in the nucleus, as the

levels of K9 and K27 methylations of ectopic
H3.3 were similar to those of ectopic H3 (lanes 5, 6, 8 and 9). When we checked
the acetylation status of H3 and H3.3 in purified nucleosomes, we found that
ectopic H3.3 was acetylated to a higher extent than ectopic H3 (lanes 11 and 12,
α-H3ac). Interestingly, a higher level of H4 acetylation (H4ac) was also detected
in the H3.3 nucleosomes (lanes 11 and 12).

72

We next determined whether similar modifications are critical for the
transcriptional activation of HSP70 genes by employing the same ChIP protocol
as for examination of HP1γ and H3.3 localization. Of note, antibodies specific for
H3 tail modifications are unable to distinguish H3 modifications from H3.3
modifications in our ChIP assays. In agreement with our purification results
(Figure 3-4A), we found that the level of H3K4me3 distinctly increased at HSPA6
promoter in response to heat shock (Figure 3-4B, HSPA6, α-H3K4me3). Similar
results were observed at transcriptionally active HSPA1 promoter, although the
increase was not as high due to the fact that high signal was detected before
heat shock treatment (HSPA1, α-H3K4me3). In marked contrast, two repressive
modifications, H3K9me3 and H3K27me3, failed to show any changes after heat
shock treatment (α-H3K9me3 and α-H3K4me27). In parallel ChIPs with
antibodies against acetylated H3 (H3ac) and H4 (H4ac), it was observed that
both promoters are preloaded with basal levels ofH3ac and H4ac under non-heat
shock conditions (α-H3ac and α-H4ac, -). However, an apparent increase in
H3ac and H4ac at the promoters was seen after heat-induced stimulation of
HSP70 transcription (α-H3ac and α-H4ac, +). Taken together, these data imply
that cooperative functions of H3.3 and HP1γ in HSP70 transcription are
accompanied by the acquisition of active histone marks at the promoter
nucleosomes.

73

Figure. 3-4. Enrichment of active modifications in H3.3 nucleosomes. (A)
Mononucleosomes containing ectopic H3 or H3.3 were prepared as in Figures 3-
1A and 3-S1A, and analyzed by Western blotting using antibodies that recognize
H3-K4 methylation (α-H3K4me1/me2/me3) H3-K9 methylation (α-
H3K9me1/me2/me3), H3-K27 methylation (α-H3K27me1/me2/me3), H2A
acetylation (α-H2Aac), H2B acetylation (α-H2Bac), H3 acetylation (α-H3ac) and
H4 acetylation (α-H4ac). Lanes1, 4, 7, and 10, mock-purified materials; lanes 2,
5, 8, and 11, H3 mononucleosomes; lanes 3, 6, 9, and 12, H3.3
mononucleosomes. (B) Cells were mock-treated (-) or heat-treated (+) for 30 min,
and modification status of promoter nucleosomes was determined. ChIP assays
were essentially as described in Figure 3-3D.

74

Figure. 3-4: Continued

75

3-5. HP1γ interacts preferentially with H3 tails carrying active histone
marks.

The observed enrichments of H3K4me3, H3ac, and H4ac in H3.3 nucleosomes
of HSP70 promoters encourage the possibility that the positive action of HP1γ in
HSP70 transcription may be linked to its ability to recognize these modifications.
In an attempt to test this possibility, we synthesized a series of peptides
corresponding to the first 28 amino acid sequences of H3 and H4 and examined
their interaction with Flag-HP1 proteins pre-bound to M2 agarose beads (Figures
3-5A and 3-5B). After extensive washing of the beads, the binding of the tail
peptides to HP1 proteins was analyzed by SDS-PAGE and followed by silver
staining. As seen in Figure 5C, the tetra-acetylated H3 tail peptide bound avidly
to HP1γ (lane 10), but its unmodified counterpart bound minimally to HP1γ (lane
5). Under the same condition, both unmodified and acetylated H3 tail peptides
showed undetectable or minimal binding to HP1α and HP1β (lanes 3, 4, 8 and 9).
Interestingly, although high level of H4 acetylation was detected in HP1γ-bound
H3.3 nucleosomes (Figures 3-1B and 3-4A), acetylated H4 tail peptides did not
show any binding affinity to HP1γ (Figure 5C, lanes 18-20 and 23-25). When K4-
trimethylated H3 tail peptides were tested for the experiments, they showed an
apparent binding capacity to HP1γ, but not to HP1α and HP1β (lanes 13-15).
Thus, our results point to the importance of active histone marks for HP1γ-H3.3
nucleosome interaction and bring up several interesting questions: What is the

76

temporal order of these modifications upon heat shock? Which factors are
responsible for the observed modifications? What are the sequence and motif of
HP1γ to allow the recognition of active histone marks present in HSP70
promoters?

3-6. Cooperative roles of H3.3 and HP1γ are deregulated in cancer cells.

HSP70 has been positioned as a cancer-relevant survival protein and
misregulation of HSP70 expression is directly linked to cell proliferation and
dedifferentiation in human cancer (Barnes et al., 2001; Ciocca and Calderwood,
2005; Galluzzi et al., 2009; Rohde et al., 2005). Intrigued by the demonstrated
action of H3.3 and HP1γ in HSP70 transcription, we next assessed the
expression levels of H3.3 and HP1γ in three different human cancer cell lines;
breast cancer cell line (MCF-7), prostate cancer cell line (LNCaP) and bladder
cancer cell line (LD611). From our Western blot and qRT-PCR analyses of these
cancer cells, a much higher level of HP1γ expression was evident in comparison
with their normal counterparts (MCF-10-2A, MLC and Urotsa) (Figure 3-6A and
Supplementary Figure 3-S5). Additionally, similar Western blot analyses detected
a slightly lower level of HP1α in prostate (LNCaP) and bladder (LD611)
carcinoma cells (Supplementary Figure 3-S5), but no difference in the levels of
HP1α, HP1β and H3.3 in MCF-10-2A and MCF-7 (Figure 3-6A). Furthermore, the
expression levels of HSP70 as well as HSPA1 and HSPA6 genes were much

77

Figure. 3-5. Preferential binding of HP1γ to acetylated H3 peptides. (A)
Schematic diagrams of unmodified and modified histone tail peptides used in the
binding assays. Peptides contain the first 28 amino acids of the N-terminal
domain of the human H3 and H4. (B) Recombinant HP1α, HP1β, and HP1γ were
resolved by 15% SDS-PAGE and visualized by Coomassie staining (lanes 2-3).
Synthesized H3 and H4 N-terminus tail peptides were analyzed by 4-20%
gradient SDS-PAGE and visualized by silver-staining (lanes 6-10). (C)
Synthesized H3 and H4 tail peptides were incubated with Flag-HP1α, Flag-HP1β,
or Flag-HP1γ immobilized on M2 agarose beads. After extensive washing, bound
peptides were resolved on 20% SDS-PAGE and visualized by silver-staining.
Input lanes represent 10% of tail peptides used in the binding reactions.

78

Fig. 3-5: Continued

79

Figure. S3-5. Differential expression patterns of the HP1 proteins in cancer
cells. Levels of three HP1 isotypes in untransformed bladder (Urotsa) and
prostate (MLC) cells were compared with those in bladder cancer (LD611) and
prostate cancer (LNCaP) cells by Western blotting and qRT-PCR as in Figure
6A.

80

higher in MCF-7 breast cancer cell lines compared to the normal cells (Figure 3-
6B). In line with the cooperative action of H3.3 and HP1γ during HSP70 gene
induction, their requirement on HSPA1 and HSPA6 transcription was examined
in MCF7 breast cancer cells. Accordingly, we depleted H3.3 and HP1γ either
individually or simultaneously, as confirmed by qRT-PCR and Western blot
analyses (Figure 3-6C). Congruent with our results from HeLa cells, the selective
knockdown of HP1γ resulted in a distinct reduction in HSPA1 and HSPA6 mRNA
levels (Figure 3-6D). Interestingly, although a similar level of H3.3 expression
was found in normal MCF-10-2A and cancer MCF7 cells (Figure 3-6A), H3.3
depletion also repressed heat shock-induced transcription of HSPA1 and HSPA6
genes (Figure 3-6D). Importantly, when both H3.3 and HP1γ were depleted,
HSPA1 and HSPA6 transcription was repressed, but the level of repression was
similar to that detected in single knockdown of H3.3 or HP1γ (Figure 3-6D), again
supporting the idea of the dual requirement of H3.3 and HP1γ for HSP70
regulation.

3-7. The effect of H3.3 and HP1γ in the growth of MCF-7 breast cancer cells.

We next examined if knockdowns of H3.3, HP1γ, or both proteins in MCF7 cells
have effects on cell proliferation by employing MTT and clonogenic assays. As
shown in Figure 3-7A, MCF7 cells depleted of HP1γ or H3.3 displayed a
significantly reduced cell growth, showing about 60% lower than mock depleted

81

Figure. 3-6. Affects of H3.3/HP1γ knockdown on HSP70 transcription. (A)
MCF-7 breast cancer cells and untransformed MCF-10-2A breast epithelial cells
were subjected to Western blot analysis using antibodies specific for HP1α,
HP1β, HP1γ, and H3.3 (left panel). Actin served as a control for equal protein
loading. qRT-PCR was performed as in Figure 2A (right panel). (B) HSP70
expression was assessed by Western blotting with anti-HSP70 antibody (left
panel). Transcription of HSPA1 and HSPA6 genes in MCF-10-2A and MCF-7
cells was quantified by qRT-PCR and corrected for expression of the control
gene (GAPDH) (right panel). qRT-PCR was also performed to measure β-actin
mRNA expression (ACTIN). (C) MCF-7 cells were transfected with shRNAs
targeting H3.3 and/or HP1γ, and individual and simultaneous depletions of H3.3
and HP1γ were confirmed by Western blot analysis (right panel) and qRT-PCR
(center and right panels). (D) Transcription levels of HSPA1 and HSPA6 genes of
H3.3/HP1γ-depleted MCF-7 cells were analyzed as in Figure 3-6B.

82

Figure. 3-6: Continued

83

Figure. 3-7. H3.3/HP1γ knockdown-induced alterations in cancer cell
growth. (A) MCF-7 cells were depleted of H3.3 and/or HP1γ as in Figure 3-6C
and cell proliferation was measured by MTT assay. Results represent the mean ±
S.D. of three experiments performed in triplicate. (B) H3.3/HP1γ-depleted MCF-7
cells were subjected to soft agar colony formation assay. The graph illustrates
the total number of colonies present on the plate after three weeks of culture.
Error bars on the graph indicate the standard deviation from triplicate
experiments. (C) Models showing the combinatorial role of H3.3 and HP1γ in
heat shock-induced HSP70 transcription. Upon heat shock, H3.3 is incorporated
at HSP70 promoters and contributes to rapid changes in active histone
modifications to stimulate the stable localization of HP1γ at the promoter
nucleosomes. The co-enrichment of H3.3 and HP1γ converts HSP70 gene from
a repressor state to an active state, leading to a great increase in transcription of
HSP70 genes. Lack of H3.3 nucleosomes disrupts the recruitment of HP1γ,
which drives the equilibrium further toward H3.3 dissociation, resulting in
blockage of transcription initiation. See the Discussion for more details.

84

Figure. 3-7: Continued

85

cells. Because simultaneous depletion of HP1γ and H3.3 failed to yield a much
greater reduction, these results constitute a powerful argument that H3.3 and
HP1γ are interdependent for their action. Similarly, MCF7 cells were depleted of
HP1γ and H3.3, and the percentage of surviving colonies was assessed three
weeks after heat shock treatment. Cell survival decreased to about 40% after
individual depletion of HP1γ and H3.3 as determined by the average percentage
of colony numbers (Figure 3-7B). Our results also showed the colonies from
MCF7 cells depleted of HP1γ or H3.3 was much smaller in size than those from
the control cells. The rate of cell proliferation and colony formation was more
obviously decreased after double knockdown of H3.3 and HP1γ (Figure 3-7B).

CHAPTER 3 DISCUSSION

It has been known for years that H3.3 incorporation and HP1γ binding to
nucleosomes are related to changes in transcriptional competency of chromatin
(9,11,12,27), but it is only recently that biochemical studies have merged to
elucidate the dynamic actions of these chromatin regulators (Font-Burgada et al.,
2008; Zhang et al., 2007). In this study we combined in vitro and in vivo
experiments to investigate the H3.3 exchange and HP1γ recruitment occurring at
human HSP70 genes induced by heat shock stimulus. Our results indicate that
H3.3 and HP1γ exert a critical role in heat shock response network through the

86

regulated transcription of HSP70, which is upstream component of the heat
shock signal transduction pathway. By using knockdown and ChIP techniques,
we were able to confirm that H3.3 exchange is necessary for transcription of two
HSP70 genes, HSPA1 and HSPA6, in response to heat shock. The heat shock-
induced enrichment of H3.3 appears to be mainly localized at the promoter, as
only modest changes in their levels were detected within the coding region. This
finding is in agreement with previous results demonstrating that H3.3-containing
nucleosomes mark promoters of transcriptionally active genes, e.g., in the MyoD
promoter (Ng and Gurdon, 2008). Somewhat surprisingly, our data show that
HP1γ is brought into HSP70 promoters when HSP70 genes are activated by heat
shock, in the same manner that heat shock stabilizes H3.3 localization at HSP70
genes. A detectable increase in H3.3 and HP1γ occupancy in coding regions was
also observed after heat shock, but the change was relatively modest. It is
therefore likely that alterations in promoter nucleosomes by H3.3 exchange and
HP1γ binding are responsible for establishing active state of HSP70 genes.

Another intriguing finding of our study is that the recruitments of H3.3 and HP1γ
to HSP70 promoters are mutually dependent, which underscores a complex
relationship between H3.3 and HP1γ in heat shock response. The mechanistic
basis for the H3.3 dependency in HP1γ recruitment is not fully elucidated in our
study. However, given that H3.3 incorporation promotes activating modifications
in the nucleosome, one obvious possibility is that these H3.3-induced

87

modifications stimulate HP1γ binding to the promoter regions (Figure 3-7C). To
support this model, our study confirmed that HP1γ overrides with HP1α and
HP1β in interacting with H3.3 nucleosomes enriched by H3/H4 acetylation and
H3-K4 methylation. Such a selective interaction could enable HP1γ to interpret a
set of combinatorial modifications and to cooperate with H3.3 at HSP70
promoters. The data presented here thus provide a glimpse into the positive
interplay of active histone modifications in HP1γ recruitment and afford a
conceptual framework on which to build similar models for other HP1γ-dependent
transcription processes. However, this model does not provide the explanation
for HP1γ-dependent function of H3.3 at the HSP70 promoter. Along with
indications that most histone variants are in dynamic equilibrium between
deposition and dissociation (Henikoff, 2008), one obvious possibility is that HP1γ
binding to H3.3 nucleosome would shift the equilibrium toward deposition. Hence,
although HP1γ is not required to facilitate the initial deposition of H3.3, we
believe that HP1γ-based decorations of H3.3 nucleosomes might be critical for
the intrinsic stability of H3.3 in the nucleosome. Due to technical limitations, this
possibility has not been examined in the current study, but should be considered
to shed light on the regulatory mechanism of HP1γ-H3.3 cooperativity. To further
support the functional significance of HP1γ and H3.3, our work demonstrates a
much higher level of HP1γ expression in cancer cells in comparison to the
corresponding normal cells. These data constitute a powerful argument that

88

overexpression of HP1γ in cancer cells results in the misregulation of the
cooperative action between H3.3 and HP1γ during HSP70 gene induction.
In fact, single or double knockdown of H3.3 and HP1γ in these cancer cell lines
leads to an apparent reduction in HSP70 transcription rate. That cell proliferation
and colony formation were obviously decreased after the same knockdown also
points to the interplay of H3.3 and HP1γ in these cancer cells. Therefore, further
characterization of cooperative activities of H3.3 and HP1γ has potential
implications in terms of cancer treatment.

It is currently unknown whether the ability of H3.3 and HP1γ to occupy other
chromatin regions is attributed by the same mechanism with which they are co-
localized in HSP70 genes. We note that a similar, but unidirectional, mechanism
has been proposed for pericentrometic heterochromatin in which H3.3 deposition
and K9 methylation contribute to the recruitment of HP1γ (Zhang et al., 2007).
These data hint at the possibility that H3.3 and HP1γ are also involved in
condensing inactive chromatin harboring high levels of the repressive mark H3-
K9 methylation. Therefore, a better understanding of cooperative actions of H3.3
and HP1γ will require further studies on how specific histone marks are initially
established and how they contribute to H3.3 exchange and HP1γ localization at
distinct chromatin regions. Also of note, Drosophila HP1c, which is human HP1γ
homolog, has been linked to transcription elongation by acting as a bridge
between FACT and elongating RNA Pol II (Kwon et al.). The observed effects of

89

HP1c are intriguing, as HP1c might perform an analogous cooperative action
with H3.3 in the post-initiation control of transcription. Hence, whether HP1γ is
capable of recognizing H3.3 nucleosomes in both the initiation and elongation
phases of transcription is an intriguing question that needs to be addressed in
future studies. Overall, our findings could be incorporated in a model that
highlights the combinatorial roles of H3.3 and HP1γ in governing signal
integration, competence, and specificity in heat-induced stimulation of HSP70
transcription (27) (Figure 3-7C). Such cooperative actions should provide
additional levels of specificity and efficiency to the signaling pathways involved in
transcriptional regulation of heat shock proteins. What emerges from this model
is a cell’s response to heat shock to be precisely regulated through highly
restricted communication between HP1γ and H3.3. The challenge for us in the
future is to determine whether other genes are also controlled by the HP1γ-H3.3
interplay and how this process is properly regulated in human cells.

90

Table. 3-1

91

CHAPTER 4: Concluding Remarks

Posttranslational modifications of histones and incorporation of histone
variants into the chromatin can affect chromatin structure and function
resulting in altered gene expression and changes in cellular processes.
Abnormal levels of gene expression and altered epigenetic patterns are
key characteristics of cancer. Changes in the level of histone acetylation
and histone methylation play important roles in the onset and progression
of cancer in numerous tumor types. Therefore, the reversal of aberrant
epigenetic changes is thought of as a potential strategy for the treatment of
cancer. In fact, there are several targeting enzymes that regulate histone
acetylation or histone methylation as epigenetic therapies, with some
demonstrating efficacy in hematological malignancies and solid tumors.
However, the exact mechanism behind these therapies still needs to be
revealed. Here, we identified SMYD3 as one of the key histone
methyltransferases required for ER-mediated transcription in breast cancer
and the cooperative role of histone variant, H3.3 and chromatin binding
protein HP1γ in regulation of HSP70, which affect the proliferation and
growth of breast cancer.

92

SMYD3 is a novel histone methylstransferase highly expressed in numerous
types of cancer, but highly overexpressed in breast cancer. To investigate the
functional role of SMYD3 in gene regulation in ERα dependent breast cancer
cells, we first checked the function of SMYD3 in ERα mediated transcription. We
found that SMYD3 can interact with ERα through LBD domain and mediate
activation of ER-target genes through methylation of H3 lysine 4 (H3K4me3).
Further, we revealed that SMYD3 and ERα co-localize on the promoters of ERα
target genes and SMYD3 mutants with inhibited methyltransferase activity
showed the loss of transactivation of MMTV promoter. Consistent with luciferase
data, ChIP and shRNA analyses showed that SMYD3 and ER are colocalized at
ER-target genes and required to enhance the transcription activity.

There were several questions raised about the transactivation of ER target genes
mediated by SMYD3; 1) Is the interaction between ER and SMYD3 specific? 2)
Does the level of H3K4 methylation change in the absence of SMYD3? In order
to address these questions, one of the histones, H1, was used to prove the
specific interaction between SMYD3 and ER. As a result, there was no
interaction between HMGB1 and ER while SMYD3 and ER specifically interacted
both in vitro and in vivo. Then, whether SMYD3 is the main methyltransferase
responsible for H3K4methylation at ER-target gene promoters, endogenous
SMYD3 was depleted using SMYD3 specific shRNA. Knockdown of SMYD3
showed decreased level of H3K4 di- and tri- methylation. Out published data

93

showed that SMYD3 is recruited to the promoters of ER-target genes, SP2,
GREB1, and CTSD, to increase the levels of H3K4 di- and tri- methylation,that
are generally known as transcription activation marker, and increase transcription
activity even higher.

Another mechanism that can control gene expression in chromatin context is
incorporation of histone variant into the nucleosome. Histone variant H3.3 plays
an important role in transcription activation. In the study of Drosophila, it was
shown that H3.3 is highly specifically incorporated in HSP70 upon heat shock.
Although, the function of HSP70 is critical in protein protection, very little is
expressed in normal cells. However, the amount of HSP70 expressed in breast
cancer is hundreds of times over that of the normal cells. Therefore, over
expressed HSP70 protects breast cancer cells from the toxic environment.

To investigate the function of histone variant in gene expression in breast cancer,
we first purified both H3 and H3.3 containing mononucleosomes from the cell.
We found that histone acetylation level of H3 and H4 as well as H3K4di- and tri-
methylation levels were enriched in H3.3 containing nucleosome compared to H3
containing nucleosome. The most interesting observation was that one subtype
of the heterochromatin binding proteins (HP1), HP1γ, was bound to H3.3
containing nucleosome.

94

Given that H3.3 is involved in HSP70 transcription activation, it will be interesting
to investigate the mechanism how H3.3 allows active state of HSP70
transcription and whether HP1γ plays a role for the incorporation of H3.3. We
observed the colocalization of H3.3 and HP1γ in the cell and found the
cooperative role between two proteins using real-time RT-PCR and ChIP. The
important functions of H3.3 and HP1γ in HSP70 gene regulation were also
shown in MTT cell proliferation assay and soft-agar colony formation assay.

In conclusion, our studies of ER-target gene regulation mediated by SMYD3 and
HSP70 gene regulation mediated by cooperative role between H3.3 and HP1γ
have provided better understandings of epigenetic regulation of gene expression
in cancer cells. Such epigenetic mechanisms, that regulate multiple aspects of
chromatin structure and function allowing transcriptionally active or repressive
state for gene expression, contributes to development of potential therapeutic
applications in the future.

95

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Abstract (if available)

Abstract The genome of eukaryotic cells is composed in nucleoprotein structure called ‘chromatin’. The basic repeating unit of chromatin is nucleosome, which consists of two pairs of H2A-H2B dimmers, and one H3-H4 tetramer wrapped around 147 bp of double stranded DNA (dsDNA). Although, the structure of chromatin is necessary for hierarchical compaction of the entire genome, it is also an obstacle for some of the most important cellular processes like transcription, DNA repair, DNA replication. In order to alter the structure between histones and DNA, eukaryotic cells imply three basic mechanisms: chromatin remodeling, incorporation of histone variants, and post-translational modification of histones.

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Creator Kim, Hyunjung (author)

Core Title Targeting chromatin modification in human cancer: SMYD3 mediated ERα transcription regulation. Cooperative role between H3.3 and HP1ϒ

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 05/05/2011

Defense Date 03/14/2011

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag chromatin,estrogen receptor,gene regulation,H3.3,histone,histone variant,HP1,HSP70,methyltransferase,OAI-PMH Harvest,post-translational modification,Transcription

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor An, Woojin (committee chair), Neamati, Nouri (committee member), Stallcup, Michael R. (committee member)

Creator Email hyunjunk.k81@gmail.com,hyunjunk@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m3917

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