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Identification of two negative regulators of p53: H1.2 complex and VprBP
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Identification of two negative regulators of p53: H1.2 complex and VprBP
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IDENTIFICATION OF TWO NEGATIVE REGULATORS OF p53:
H1.2 COMPLEX AND VprBP
by
Kyunghwan 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
(BIOCHEMISTY AND MOLECULAR BIOLOGY)
May 2010
Copyright 2010 Kyunghwan Kim
ii
ACKNOWLEDGEMENTS
During the process of completing these projects, it became quite clear to me that I cannot
complete a Ph.D. course for myself. Even though the list of individuals I would like to
thank extends beyond the limits of this format, I wish to thank the following persons for
their dedication and support.
My mentor, Dr. Woojin An, has been a significant presence in my life. First of all, he not
only provided me an opportunity to study in the new field, but also taught me how to
progress my project and overcome problems. I will always be thankful for his support,
patience, and deep concern not only for me, but also for my family.
I wish to thank Dr. Michael Stallcup and Dr. Clay Wang for gratefully serving as
committee members and their advice to be better scientist. I am thankful my lab members
Dr. Jongkyu Choi, Dr. Kyu Heo, Hyunjung Kim and Bomi Lee for their assistance. Just
like family, we have been spending our time in the lab.
I would like to thank my friends Dr. Kwangwon Jeong, Seokwoo Hong, Yongseok Kim
and Dongjin Youn for their endless support. Without them, I cannot move on to the next
stage of my life.
I really thank my extended family for their understanding and support during these years
that I have persued my doctorate.
To my wife Sunju Yi, and our children Bomi Kim, Taeyeop Kim, all I can say is “ I love
you”. Your love, commitment and encouragement have supported me, especially in those
many days when I spent more time with my pipet than with you
iii
TABLE OF CONTENTS
Acknowledgements................................................................................………….……….ii
List of Figures…………………………………………………………………………….iv
Abstract……………………………………………………………………………….......vi
Chapter 1: Introduction………………………………………………………………..1
Chapter 2: Isolation and Characterization of a Novel H1.2 Complex
That Acts as a Repressor of p53-Mediated Transcription………………...8
Chapter 3: VprBP Antagonizes p53-Mediated Transactivation via
Inhibition of H3 Acetylation……………………………………………..49
Chapter 4: Concluding Remarks…………………………………………………......94
Bibliography………………………………………………………………………….…..98
iv
LIST OF FIGURES
Fig. 1. Posttranslational modification sites identified
in human H1 isoforms…………………………………………….5
Fig. 2-1. Preparation of H1.2 complex and recombinant H1.2……………21
Fig. 2-2. Interaction of H1.2 with its associated factors…………………..26
Fig. 2-3. Repressive effects of H1.2 complex in
chromatin acetylation and transcription…………………………30
Fig. 2-4. Direct interaction of H1.2 with p53……………………………..35
Fig. 2-5. Functional characterization of H1.2, PUR α and YB1……...........40
Fig. 2-6. Model for the promoter-selective inhibition of
p53-dependent transcription by H1 complex…………………….45
Fig. 3-1. Effects of VprBP overexpression and knockout
on cell viability…………………………………………………..62
Fig. 3-2. VprBP-mediated repression of chromatin transcription
and H3 acetylation……………………………………………….66
Fig. 3-3. Selective interaction of VprBP with unmodified H3 tail………...70
Fig. 3-4. Requirement of LisH domain for VprBP action…………………73
Fig. 3-5. Dynamics of promoter occupancy of VprBP, p53
and HDAC1……………………………………………………...78
Fig. 3-6. Physical association of VprBP with p53 and HDAC1…………...83
Fig. 3-7. Stimulation of p53 transcription and apoptosis
by VprBP knockdown……………………………………………86
Fig. 3-8. Model for the role of VprBP in p53 transrepression……………..93
v
ABSTRACT
p53 is a transcription factor which regulates the cell cycle, DNA repair and apoptosis in
response to DNA damage. Even though the mechanisms of p53 accumulation and
activation have been extensively studied, little is known about how p53 function is
negatively regulated. Here we described H1.2 complex and VprBP as novel negative
regulators of p53-mediated transcription.
Linker histone H1 has been generally viewed as a global repressor of transcription by
preventing the access of transcription factors to sites in chromatin. However, recent
studies suggest that H1 can interact with other regulatory factors for its action as a
negative modulator of specific genes. To investigate these aspects, we established a
human cell line expressing H1.2, one of the H1 subtypes, for the purification of H1-
interacting proteins. Our results showed that H1.2 can stably associate with sets of
cofactors and ribosomal proteins which can significantly repress p53-dependent, p300-
mediated chromatin transcription. This repressive action of H1.2 complex involves direct
interaction of H1.2 with p53, which in turn blocks p300-mediated acetylation of
chromatin. YB1 and PUR α, two factors present in the H1.2 complex, together with H1.2
can closely recapitulate the repressive action of the entire H1.2 complex in transcription.
ChIP and RNAi analyses further confirmed that the recruitment of YB1, PUR α and H1.2
to the p53 target gene Bax is required for repression of p53-induced transcription.
Therefore, these results reveal a previously unrecognized function of H1 as a
vi
transcriptional repressor, as well as the underlying mechanism involving specific sets of
factors in this repression process.
HIV-1 Vpr binding protein (VprBP) has been implicated in DNA replication and cell
cycle regulation, but its precise role remains unclear. Here we report that VprBP
epigenetically regulates p53-induced transcription and apoptotic pathway. VprBP is
recruited by p53 to occupy p53-responsive promoters and suppress p53 transactivation in
the absence of stress stimuli. To maintain target promoters in an inactive state, VprBP
stably binds to nucleosomes by recognizing unacetylated histone H3
tails after the initial
recruitment. Promoter-localized deacetylation of H3 tails by HDAC1 is prerequisite for
VprBP to tether and act as a bona fide inhibitor at p53 target genes. Consistent with these
results, VprBP knockdown leads to activation of p53 target genes that initiate cell cycle
arrest and apoptosis upon DNA damage, and causes an increase in DNA damage-induced
cell death. Significantly, VprBP is overexpressed in human cancer cells, and this
increased level of VprBP is tightly correlated with reduced apoptosis. Our results thus
reveal a new role for VprBP in regulation of p53 signaling pathway, as well as molecular
mechanisms of cancer development related to VprBP overexpression.
1
CHAPTER 1: Introduction
p53 function and Regulation
p53 is an important tumor suppressor that plays a pivotal role in regulating cell cycle
progresssison, DNA repair and apoptosis in response to genotoxic stresses (Menendez et
al., 2009; Riley et al., 2008; Vousden and Prives, 2009). p53 missense mutations are
observed in about 50% of human cancers, and the misregulation of p53 activity by a
group of regulators also contribute to cancer development. p53 inhibits tumor formation
by acting as a sequence-specific DNA binding factor that activates transcription of
various target genes, including p21, GADD45, Bax, Noxa and Mdm2.
The tumor suppressor activity of p53 is tightly controlled by proteosomal degradation
and post-translational modifications (Appella and Anderson, 2001). In unstressed cells,
p53 is maintained at low levels by Mdm2-mediated proteolytic degradation. Upon DNA
damage, p53 undergoes posttranslational modifications including phosphorylation,
acetylation and methylation that contribute to its accumulation and activation.
Structurally and functionally, p53 protein can be divided into four major domains: the N-
terminal transactivation domain (residues 1-83), the central DNA binding domain
(residues 102-292), the tetramerization domain (residues 323-355) and the C-terminal
regulatory domain (residues 364-393). The DNA-binding domain specifically recognizes
2
DNA response elements at the promoter regions of target genes, and C-terminal
regulatory domain helps to serach for target response elements.
Although these domains themselves play a distinct role in p53 function, the tumor
suppressor activity of p53 is also tightly controlled by post-translational modifications
(Aylon and Oren, 2007). In the event of DNA damage, p53 is phosphorylated at a number
of residues within N-terminal activation domain, which dissociates p53 from the negative
regulator Mdm2 and stabilizes it. p53 also undergoes acetylation at the C-terminal region
upon DNA damage. This modification promotes cofactor recruitment to the target gene
promoters, resulting in transcriptional activation.
Chromatin and transcription
Since the discovery of the nucleosome as an element of chromatin in 1974 (Kornberg
and Thomas, 1974), it has been elucidated that chromatin is untimately involved in the
regulation of all DNA dependent cellular processes including gene transcription. The
nucleosome is the fundamental unit of chromatin and consists of 146 bp of DNA
wrapped around an octamer of four core histones (H2A, H2B, H3 and H4) (Kornberg and
Lorch, 1999; Luger and Richmond, 1998; van Holde, 1988; Workman and Kingston,
1998). The N-terminal tails of core histones are unstructured and undergo
posttranslational modifications including acetylation, methylation, phosphorylation and
ubiquitylation. These modifications allow tightly packed chromatin to possess dynamic
3
properties for proper regulation of gene transcription (Kouzarides, 2007). So far, two
well-characterized mechanisms of action of histone modifications in chromatin
transcription have been accepted. One is the dissociaton of nucleosomal contacts to create
a loose chromatin structure, and the other is the recruitment of nonhistone proteins to
specific chromatin domains. Acetylation of lysine residues at the N-terminal histone tails
is one of the most studied modifications. Histone acetylation reorganizes chromatin by
neutralizing the positive charge of the lysine residue and allowing access of regulatory
proteins to the promoter. Histones also can be methylated at lysine and arginine residues
by histone methyltransferases. Unlike acetylation, methylation does not alter the overall
charge of the lysine and argine residues and therefore is thought to allow the recruitment
of regulatory proteins with methyl group recognizing domains (chromo, tudor, MBT and
PHD domains). Depending on the metylation site, gene transcriptions are differentially
regulated. For example, methylations of H3K4 and H3K36 are involved in gene
activation, whereas methylations of H3K9 and H3K27 are implicated in gene repression.
Although the relative contributions of histone modificaions to chromatin transcription are
well characterized, the mechanism by which these modifications regulate gene
transcription remains elusive.
Linker histone
In eukaryotes, DNA is packaged into a chain of nucleosomes that is composed of a two
copies of four core histones and DNA. Another major class of chromatin protein called
4
the linker histone H1 binds to the linker DNA entering and exiting the nucleosome and
contributes to the formation of the stable nucleosome and 30nm chromatin fiber (Bustin
et al., 2005; Happel and Doenecke, 2009; Woodcock et al., 2006). The linker histone has
a tripartite structure. It consists of a short-flexible N-terminal domain, a highly conserved
globular domain and a long, extended C-terminal domain. N-terminal and C-terminal
domains are thought to be involved in interaction with regulatory factors, and globular
domain is responsible for linker DNA binding.
Linker histone H1 is the most divergent group of histone proteins. In mammals, there are
at least eight H1 subtypes that display distinct patterns of expression during development
and differentiation. H1 subtypes can be subdivided based on their temporal and spatial
expression. In terms of temporal expression, the replication dependent subtypes H1.1,
H1.2, H1.3, H1.4 and H1.5 and the testis specific subtype H1t are mainly expressed in S-
phase, whereas the replacement histone H1.0 is expressed in a replication independent
manner. Spatially, H1.2, H1.3, H1.4 and H1.5 are present in most somatic cells, and
H1.0, H1.1, H1t and H1oo are expressed in the specific cell types (Happel and Doenecke,
2009; Izzo et al., 2008; Parseghian and Hamkalo, 2001).
As for the four core histones, H1 proteins undergo various posttranslational modification
including phosphorylation, acetylation, methylation and ubiquitylation (Fig. 1). Although
H1 phosphorylation is generally associated with chromatin decondensation, the precise
contribution of H1 phosphorylation to transcription still remains unclear.
5
Fig. 1. Posttranslational modifications of human H1 isoforms. Blue residue:
phosphorylation; red residue: methylation; green residue: formylation; yellow
highlighted: acetylation; grey highlighted: αN-terminal acetylation; underlined:
ubiquitination. The sequence stretch representing the winged helix motif is marked with a
black bar. Site of acetylation/methylation has not been assigned exactly: either
acetylation of K168 and methylation of K169 or acetylation and methylation of K168 in
H1.3 and H1.4 and of K167 and of K168 in H1.5, respectively. (Happel N et al., Gene
2009)
6
Very recently, one of the H1 isoforms, H1.4, was found to be methylated by PRC4
complex, resulting in transcriptional repression (Kuzmichev et al., 2004), but again the
function of this modification is not well defined. The heterogeneity of primary sequence,
expression patterns and posttranslational modifications suggests that H1 histones
differentially contribute to the gene regulation.
VprBP
VprBP, originally identified as HIV-1 auxiliary regulatory protein Vpr binding protein, is
present in higher eukaryotes including fly, worm, mouse and human (Zhang et al., 2001).
VprBP is ubiquitiously expressed in human and mouse tissue, but shows preferential
expression during the formation of testis tissue, suggesting that VprBP is important for
testis development. Very recently, it has been shown that VprBP acts as a component of
Cul4-DDB1 E3 ligase complex (Huang and Chen, 2008; Le Rouzic et al., 2007; McCall
et al., 2008; Tan et al., 2007; Wen et al., 2007; Zhang et al., 2008). VprBP knockdown
increased the expression level of p21 cyclin-dependent kinase inhibitor, and VprBP
ablation in mouse embryonic fibroblasts blocks the cell cycle progression and facilitates
subsequent apoptosis (Hrecka et al., 2007; McCall et al., 2008). All these informations
suggest that VprBP might be involved in cell cycle progression and apoptosis.
The goal of this thesis is 1) to purify, identify and characterize H1.2 interacting proteins,
2) to study the function of H1.2 complex as a repressor of p53 transactivation and 3) to
7
elucidate the mechanism of repressive action of VprBP in p53 tranactivation. We identify
H1.2 complex and VprBP as negative regulators of p53-mediated transcription. These
results may contribute to the new paradigm for understanding epigenetic regulation of
p53 signaling pathway.
8
CHAPTER 2: Isolation and Characterization of a Novel H1.2 Complex
That Acts as a Repressor of p53-Mediated Transcription
INTRODUCTION
Histones are the major protein components to compact genomic DNA into the limited
volume of the nucleus as a highly-organized chromatin structure. The basic element of
chromatin is the nucleosome, which consists of 146 base pairs of DNA wrapped around
an octameric core of histones containing two molecules each of H2A, H2B, H3 and H4
(Kornberg and Lorch, 1999; Luger and Richmond, 1998; van Holde, 1988; Workman and
Kingston, 1998). This repeating unit of chromatin is associated with another type of
histone called linker histone H1 to achieve an additional level of compaction, making
genes inaccessible to transcription factors and preventing their expression (Brown, 2003;
Bustin et al., 2005; Georgel and Hansen, 2001; Parseghian and Hamkalo, 2001;
Woodcock et al., 2006). Mammalian cells have at least eight histone H1 subtypes
including H1.1 through H1.5 and somatic cell specific H1o as well as germ cell-specific
H1t and H1oo, all consisting of a highly conserved globular domain and less conserved
N- and C-terminal domains (Bustin et al., 2005; Parseghian and Hamkalo, 2001;
Thiagalingam et al., 2003). The existence of multiple H1 subtypes and the diversity of
their amino acid sequences raise the possibility that individual subtypes have
nonredundant functions in various cellular processes. In addition, the expression of each
H1 subtype depends on the tissue, phase of the cell cycle, and developmental stage,
9
further suggesting the specific contribution of linker histone subtypes for regulation of
various cellular processes (Bustin et al., 2005; Khochbin, 2001; Parseghian and Hamkalo,
2001).
Although most studies have focused on the contribution of H1 as a structural component
of the nucleosome, it is becoming apparent that H1 also acts as a repressor for specific
gene transcription (Alami et al., 2003; Brown et al., 1996; Fan et al., 2005; Shen and
Gorovsky, 1996). This repressive capacity of H1 on transcription appears to be
accomplished by its localization at particular chromosomal domains with specific
transcription regulators. Msx1 recruits a linker histone H1 to the MyoD gene, and this
selective localization correlates with a repressive chromatin state and gene repression
(Lee et al., 2004). Simultaneous inactivation of three H1 subtype genes (H1.2, H1.3 and
H1.4) in mouse embryonic cells significantly affects the expression of a subset of genes,
supporting rather specific action of H1 in gene regulation (Peitz et al., 2002). A recent
study demonstrating that specific sets of ribosomal proteins interact with H1 to suppress
transcription also provides support for a rather complex mechanism for the effect of H1
in gene regulation (Ni et al., 2006). It thus appears that the linker histone H1 requires
other regulatory factors to retain its optimal capacity and specificity for epigenetic gene
regulation.
To understand the molecular mechanisms by which transcription is down-regulated by
H1, we purified factors stably associated with H1.2, one of the human H1 subtypes, by
10
employing an epitope-tagging and stable cell line approach. Our functional analysis
demonstrated that the purified H1.2 complex represses p53-dependent, p300-mediated
chromatin transcription by blocking chromatin acetylation. The result that H1.2 alone is
defective in repression underscores the significance of factors associated with H1.2 for
repressive action of H1.2 in transcription. Furthermore, we found that the physical
interaction of the H1.2 complex with p53, most likely through H1.2 present in the
complex, provides a novel mechanism for the transcriptional repression by the H1.2
complex. Therefore, apart from a role of H1 in maintaining higher order chromatin
structure, our results provide new insights into the molecular mechanism of action of
linker histone H1 in specific transcription events.
EXPERIMENTAL PROCEDURES
Generation of H1.2 cell line. To construct pIRES-FHH1.2 vector used for mammalian
expression of H1.2, human H1.2 gene was amplified by PCR and inserted into NotI and
EcoRI sites of pIRESneo containing FLAG and HA tags. HeLa-S cells (1x10
6
) were
transfected with 3 µg of pIRES-FHH1.2 using Lipofectamine (Invitrogen), and positive
clones were selected with G418 (500 µg/ml, Invitrogen) for three weeks. H1.2 expression
within the isolated clones was confirmed by 4-20% gradient sodium
dodecyl sulfate-
polyacrylamide gel electrophoresis (SDS-PAGE)
and Western blotting with anti-FLAG
(Sigma) and anti-HA antibodies (Santa Cruz Biotechnology). The selected H1.2 cell line
11
was grown in 8 liter spinner culture in DME-PO
4
medium (Irvine Scientific), and nuclear
extract (0.5 g) was prepared as recently described (Heo et al., 2007).
Purification and identification of H1.2 complex. To purify the H1.2 complex, the
nuclear extract prepared from the H1.2 cell line was first loaded onto a Phosphocellulose
P11 column (Whatman) equilibrated with BC150
buffer (20 mM HEPES-KOH [pH 7.9],
0.5 mM EDTA, 0.05% NP-40,
10% glycerol, 1 mM dithiothreitol [DTT], protease
inhibitors,
150 mM KCl). The bound proteins were step-eluted with BC300, BC500,
BC850, BC1200 and BC2000. The BC1200 fraction containing the H1.2 complex was
dialyzed in BC300 buffer, and further purified by using M2 agarose affinity
chromatography (Sigma) (Heo et al., 2007). The preparation was confirmed by SDS-
PAGE with 4-20% linear gradient and immunoblotting using either anti-FLAG or anti-
HA antibody. A portion of the H1.2 complex isolated after M2 agarose affinity
chromatography (0.2 ml) was applied to a 5 ml 15-40%
glycerol gradient in BC250
containing 0.1% Nonidet
P-40. After centrifugation at 150,000 g for 20 h in
SW
55Ti rotor at 4 °C, fractions (150 µl) were collected from
the top of the tube. The
distribution of the H1.2 complex was
determined by
silver staining and Western blotting
of 4-20% SDS-PAGE gels. For mass spectrometry analysis, purified factors were
resolved in 4-20% gradient SDS-PAGE, and proteins were visualized by Coomassie blue
staining. Bands were excised from the gel and submitted to the protein sequencing
facility at the USC core mass spectrometry facility for in-gel trypsin digestion, followed
by peptide sequencing according to facility protocols. The presence of the identified
12
proteins within the purified H1.2 complex was further confirmed by Western blot
analysis. Antibodies used for Western blot analysis were as follows: anti-Nucleolin and
anti-lamin A/C from Santa Cruz Biotechnology; anti-CAPER α from Bethyl Laboratories;
anti-WDR5 from Abcam; anti-tubulin from Calbiochem; anti-H2B from Upstate Biotech;
anti-DNA-PK from Dr. Lieber (Ma et al., 2005), anti-PARP1 from Dr. Comai (Li et al.,
2004), anti-FIR from Dr. Levens (Liu et al., 2000), anti-YB1 from Dr. Kohno (Okamoto
et al., 2000) and anti-PUR α from Dr. Johnson (Daniel et al., 2004). Anti-ASXL1 was
from Dr. Brock.
Construction and expression of recombinant H1.2. Wild type and C-terminal deleted
H1.2 constructs were generated by subcloning H1.2 gene fragments encoding amino
acids 1-213 and amino acids 1-109. E. coli Rosetta 2 (DE3) pLysS cells (Novagen) were
transformed with the resulting pET-H1.2 construct, grown in 1 L of Luria Broth (LB) at
37 °C. Harvested bacteria were lyzed in 25 ml of lysis buffer (20 mM Tris, pH 8.0, 10%
glycerol, and 3 mM DTT, 0.5 M KCl) by sonication, and the cleared lysate (125 mg) was
bound to Ni-NTA-affinity resin (Novagen) in batch by rocking at 4 °C for 1 h. After
binding, the resin was washed five times with lysis buffer and eluted with BC300 buffer
containing 0.25 M imidazole. The eluted protein was loaded onto a 1 ml CM-Sephadex
C-25 column (Amersham Biosciences) pre-equilibrated with BC300. After extensive
washing with BC300, the bound proteins were step-eluted with BC300, BC500, BC800
and BC1200. The combined BC800 fractions containing H1.2 were dialyzed against
BC300 buffer and applied to SP-Sepharose column (Amersham Biosciences) pre-
13
equilibrated with BC300. Elution was again carried out by using a four-phase salt
gradient (0.3, 0.5, 0.8 and 1.2 M) in BC buffer. H1.2-containing 0.8 M fractions were
collected, and dialyzed against BC100. Protein concentrations were determined by BCA
protein assay (Pierce) using BSA as a protein standard.
GST-pull down assays. To generate H1.2 fused to glutathione S-transferase (GST), H1.2
DNA sequence was subcloned into the EcoRI and BamHI sites of pGEX-2T (Amersham
Pharmacia Biotech). Recombinant GST and GST-H1.2 proteins were expressed in
Escherichia coli Rosetta 2 (DE3) pLysS and purified by affinity chromatography on
glutathione-Sepharose 4B beads (GE Healthcare) according
to the manufacturer's
protocol. Vectors encoding ASXL1, β-catenin, TGase7, CAPER α, YB1 and WDR5 were
prepared by subcloning their cDNAs into pcDNA3.1/His vector (Invitrogen). Vectors
encoding GST-PUR α and HA-FIR were as recently described (Johnson et al., 1995; Liu
et al., 2000). The ASXL1, β-catenin, TGase7, CAPER α, YB1 and WDR5 were
synthesized by in vitro translation using the TNT coupled transcription-translation
system,
with conditions as described by the manufacturer (Promega). FLAG-tagged PARP1 was
expressed in insect Sf9 cells and purified on M2 agarose according to standard protocol.
HA-tagged FIR and GST-fused PUR α were purified as recently described (Johnson et al.,
1995; Liu et al., 2000). The GST-tag was removed from GST-PUR α by thrombin
cleavage kit (Novagen). In vitro binding experiments
were carried out using purified
recombinant GST or GST-H1.2
(2 µg) proteins bound to
glutathione-Sepharose 4B beads
and one of the prepared components of the H1.2 complex in 0.5 ml of binding buffer
(25
14
mM HEPES [pH 7.8], 0.2 mM EDTA, 20% glycerol, 150 mM KCl,
and 0.1% Nonidet P-
40). Glutathione beads were washed four times in binding buffer and boiled in SDS
sample buffer to elute bound proteins, which were then analyzed by SDS-PAGE and
Western blot analysis with antibodies as described in Fig. 2-2B. For identification of p53
interaction domain of H1.2, expression
vectors for H1.2 deletion mutants, i.e. H1.2-NT,
H1.2-GD, H1.2-CT, were also generated
by inserting PCR-amplified cDNA fragments
encoding amino acids
1-34 (for NT), 35-109 (for GD) and 110-213 (for CT) of H1.2 into
the pGEX4T-1 (Amersham Biosciences). The GST-p53 fusion proteins comprising
full-
length p53, its first 83 residues (for NT), residues 120-290 (for DBD) and its final 104
residues (for CT) were also prepared as described (Le Rouzic et al., 2007).
Immunoprecipitation assays. The genes encoding H1.2 and PUR α were prepared by
PCR amplification and subcloned into pcDNA3.1/His. Construction of pCMV-Taq2
PARP1 vector was previously described (Huang et al., 2006). All other expression
vectors were identical to those used in the TNT coupled transcription-translation
system.
293T cells (3x10
6
) were transiently transfected with 3 µg of an expression vector for
FLAG-PARP1, Xpress- β-catenin, Xpress-TGase7, HA-FIR, Xpress-CAPER α, Xpress-
YB1, Xpress-PUR α or Xpress-WDR5 along with 3 µg of an expression vector for FLAG-
or Xpress-H1.2. Total amounts of the expression vectors were kept constant by adding
empty vectors. Two days after transfection, cells were harvested and total cell extracts
were clarified by centrifugation. Immunoprecipitation was performed with anti-FLAG,
anti-HA and anti-Xpress antibodies (Invitrogen) as previously described (An et al., 2004).
15
Co-precipitated proteins were detected by Western blot analysis with anti-FLAG, anti-
HA, and anti-Xpress antibodies. To generate H1.2 deletion mutants, human H1.2 gene
fragments encoding amino acids 35-213 ( ∆NT), amino acids 1-109 ( ∆CT) and amino
acids 110-213 (CT) were subcloned into pcDNA3.1/His. A construct expressing amino
acids 290-393 of p53 was generated by inserting a corresponding gene fragment into
pIRESneo with the FLAG and HA tags. Other p53 expressing constructs have been
described recently (An et al., 2004). Transfection and immunoprecipitation assays with
mutant p53 and H1.2 were performed as described above. For coimmunoprecipitation of
endogenous proteins, 293T cell lysates were immunoprecipitated
with anti-p53
monoclonal antibody (DO-1, Santa Cruz) followed by immunoblotting with antibodies
against H1.2 (Abcam) and p53.
Immunofluorescence microscopy analysis. The vector pEGFP-H1.2 was generated by
subcloning H1.2 gene into the EcoRI and BamHI sites of pEGFP-C1, and other
expression plasmids were prepared as described in immunoprecipitation assays. HeLa
cells were grown on 18 mm glass coverslip to 40% confluency with DME-PO
4
medium
supplemented with 10% fetal bovine serum and transfected with 0.3 µg of mammalian
expression vectors for EGFP-H1.2 along with 0.3 µg of an expression vector for Xpress-
ASXL1, FLAG-PARP1, HA-FIR, Xpress-CAPER α, Xpress-YB1, Xpress-PUR α and
Xpress-WDR5 as indicated in Fig. 2D. Two days after transfection, cells were briefly
washed with PBS, fixed with 4% para-formaldehyde in PBS for 15 min and
permeabilized with 0.3% Triton X-100 in PBS for 15 min at room temperature. For
16
localization of other proteins, fixed cells were blocked in 3% BSA and incubated with the
α-FLAG, α-Xpress or α-HA antibody,
diluted in PBS containing 3% bovine serum
albumin, and subsequently with
the Cy3-conjugated secondary antibodies (Jackson
Laboratory). All incubations were at room temperature for 2 h. Confocal laser
microscopy was performed with a Zeiss LSM 510 dual-photon confocal microscope at
63X magnification, and digital images were analyzed with the Adobe Photoshop
software.
Transcription and histone acetyltransferase assays. The assembly of chromatin
templates with recombinant ACF, recombinant NAP1, recombinant core histones and
p53RE/G-less plasmid
DNA was performed as recently described (Heo et al., 2007).
FLAG-tagged human p300 and p53 proteins were expressed and purified on M2-agarose
(Sigma) according to standard
procedures and previously described (An and Roeder,
2004). For construction of the YB1 expression vector, coding sequence of YB1 was
PCR-amplified and inserted into the NdeI and BamHI sites of the pET-15b. The
recombinant YB1 protein was expressed in E. coli Rosetta 2 (DE3) pLysS cells and
purified with Ni-NTA-affinity resin (Novagen). The recombinant PUR α was prepared as
described above. Transcription assays were performed with 40 ng of chromatin or free
DNA as recently described (Heo et al., 2007), except that H1.2 or H1.2 complex was
added together with p300 and acetyl-CoA. When H1.2 (40 ng and 80 ng) was used in
transcription, BSA (160 ng and 320 ng) was also included to adjust a final concentration
similar to H1.2 complex (200 ng and 400 ng). For HAT assays, chromatin template (100
ng) was incubated with H1.2 (80 ng with 320 ng of BSA) or H1.2 complex (400 ng) in
17
the presence of p53 (30 ng), p300 (40 ng) and 2.5 µM [
3
H]-acetyl-CoA. Transcription
assays with wild type/C-terminal deleted H1.2, YB1 and PUR α were as described above
except that H1.2 (50 ng), YB1 (100 ng) and PUR α (100 ng) were added together with
p300. For reporter gene assays with wild type/C-terminal deleted H1.2, YB1 and PUR α,
H1299 cells were grown to 50% confluency (1x10
5
) on 12-well plates in DMEM with
10% fetal bovine serum. Transfection assays were performed, as indicated, with reporter
plasmid (200 ng) bearing p53 response element, p53 expression vector (100 ng), and
expression vector (200 ng) of H1.2, YB1 or PUR α. Total amount of plasmid DNA was
adjusted to 1 μg by adding empty vector. Cells were harvested at 48 h and analyzed for
luciferase activity as described previously (An et al., 2004).
RT-PCR and Chromatin immunoprecipitation (ChIP) assays. ChIP assays were
performed essentially as described (An et al., 2004; Ivanov et al., 2007) by using H1299
cells after transfection either with the plasmids expressing p53, Xpress-PUR α, FLAG-
YB1 and Gal4-H1.2 or with empty control vector. The following primers were used for
PCR amplification: Bax, 5'-TATCTCTTGGGCTCACAAG-3' and 5'-
ACTGTCCAATGAGCATCTCC-3'; GAPDH, 5’-CAGCACAGCCCACAGGTTTCC-3’
and 5’-CCTGGCTCCTGGCATCTCTGG-3’. Anti-p53 (FL393, Santa Cruz
Biotechnology), anti-GAL4-DBD (Santa Cruz Biotechnology), anti-FLAG and anti-
Xpress antibodies were used to immunoprecipitate DNA. Total RNA was also isolated
with RNeasy Mini Kit (Qiagen) and subjected to RT-PCR as described previously (An et
al., 2004). The following primers were used for RT-PCR: Bax: 5’-
18
CGTCCACCAAGAAGCTGAGCG-3’ and 5’-AGCACTCCCGCCACAA AGATG-3’;
Actin: 5’-GTGGGGCGCCCCAGGCACCA-3’ and 5’-CTCCTTAATGTCACGCACGA
TTTC-3’. The PCR products were resolved on a 1.5% agarose gel containing ethidium
bromide.
Construction of H1.2, YB1 and PUR α shRNA plasmids and stable transfection. The
design and construction of the shRNA clones against H1.2, YB1 and PUR α was
performed according to the manufacturer’s protocol (Ambion). shH1.2:
gatccAGAGCGTAGCGGAGTTTCTttcaagagaAGAAACTCCGCTACGCTCTTTttttggaa
a, agcttttccaaaaAAAGAGCGTAGCGGAGTTTCTtctcttgaaAGAAACTCCGCTACGCT
CTg, shYB1: gatccGAAGGTCATCGCAACGAAGttcaagagaCTTCGTTGCGATGAC
CTTCTTttttggaaa, agcttttccaaaaAAGAAGGTCATCGCAACGAAGtctcttgaaCTTCGT
TGCGATGACCTTCg, shPUR α: gatccgCCGCAAGTACTACATGGATttcaagagaAT
CCATGTAGTACTTGCGGTTttttggaaa, agcttttccaaaaAACCGCAAGTACTACATGG
ATtctcttgaaATCCATGTAGTACTTGTGGcg (target sequences were capitalized) were
subcloned into pSilencer 2.1-U6 neo plasmid (Ambion) and used for generation of stable
cell lines. RNA was extracted from U2OS cells stably transfected with shRNA of H1.2,
YB1 or PUR α with or without Adriamycin (0.5mg/ml for 8 h, Fluka) treatment by using
RNeasy Mini Kit. The cDNA was synthesized from purifed RNA with SuperScript III
First-Strand Kit (Invitrogen), and relative changes in expression of Bax gene were
assessed by real time PCR.
19
RESULTS
2.1 Linker histone H1.2 stably associates with multiple regulatory factors in living
cells.
As a first step in exploring the repressive roles of H1 in transcription, we generated a
HeLa-derived cell line that constitutively expresses FLAG and HA-tagged H1.2 for the
purification of the H1.2 complex (see EXPERIMENTAL PROCEDURES for details).
Our Western blot analysis with FLAG antibody confirmed that the major fraction of
expressed H1.2 was present in the nucleus (Fig. 2-1B). Similar Western analysis of the
cell line nuclear extracts with H1.2 antibody also confirmed comparable levels of ectopic
H1.2 versus endogenous H1.2 (Fig. 2-1C), thus minimizing the possibility that ectopic
H1.2 nonspecifically interacts with other factors due to its non-physiological
concentration. To enhance the purity of the H1.2 complex in our purification, the nuclear
extract prepared from the cultured H1.2 cell line was initially fractionated on a P11
phosphocellulose column with increasing salt concentrations (Fig. 2-1A). The 1.2 M
fraction containing ectopic H1.2 was further purified by immunoaffinity chromatography
using anti-FLAG antibody. An SDS-PAGE analysis of the purified H1.2 complex
consistently identified 16 bands that are not observed with the control purification,
similarly conducted with HeLa nuclear extracts (Fig. 2-1E). In an effort to confirm the
stability of the H1.2 complex, we further purified it by ultracentrifugation in a 15–40%
glycerol gradient under stringent condition (250 mM KCl, 0.1% NP40). As shown in Fig.
20
2-1G, the complex sedimented as a single discrete peak in the glycerol gradient (fractions
21-25), suggesting that H1.2 forms a single complex. In order to define the functional
role of H1 as a single protein, His-tagged H1.2 was also expressed in bacteria and
purified by three consecutive chromatographies using Ni-NTA, CM Sephadex C-25, and
SP-HP as described in EXPERIMENTAL PROCEDURES (Fig. 2-1D). The overall
procedures for purification of the H1.2 complex and the recombinant H1.2 are
summarized in Fig. 2-1A and 2-1D.
To identify the factors present in the H1.2 complex, the major protein bands were excised
from the gel and subjected to mass spectrometry analysis. The most prominent proteins
identified in our analysis were four endogenous ribosomal proteins (L13a, L7a, L22 and
S3) among the cluster of abundant low-molecular weight proteins. These results are
consistent with recent results from Drosophila, indicating that H1 interacts with multiple
nuclear ribosomal proteins for more efficient repression of transcription (Ni et al., 2006).
In further support of a repressive role for H1 in transcription, four of the proteins (YB1,
FIR, PARP1 and PUR α) present in the purified complex also belong to corepressor
family of proteins (An et al., 2004; Knapp et al., 2006; Lasham et al., 2000; Liu et al.,
2000). Somewhat surprisingly, however, we also found among the pull-down factors
ASXL1, nucleolin, β-catenin, and CAPER α which were originally identified as
coactivators in gene activation (Cho et al., 2006; Clevers, 2006; Dowhan et al., 2005;
Doyen et al., 2006). In addition to transcription-related factors, mass spectrometry
analysis also identified proteins that have a role in other cellular processes such as
21
Fig. 2-1. Preparation of H1.2 complex and recombinant H1.2. A, Schematic diagram
for the purification of H1.2 complex from the stable cell line. Numbers indicate KCl
concentration used to purify the individual fractions. Each elution was separated by 4-
20% gradient SDS-PAGE and probed with HA and FLAG antibodies as indicated (lane
2). The control preparation with normal HeLa nuclear extract is also included (lane 1). B,
Subcellular localization of ectopic H1.2. Cytoplasmic and nuclear extracts were prepared
as recently described (Heo et al., 2007) and analyzed by immunoblots with anti-FLAG,
anti-lamin A/C and anti-tubulin antibodies. C, Relative levels of ectopic H1.2 versus
endogenous H1.2. Nuclear extracts were prepared from control cells (lane 1) and H1.2
expressing cells (lane 2), and Western blot analysis were performed with anti-H1.2
antibody. D, Purification of recombinant H1.2. Recombinant H1.2 (rH1.2) was purified
as described in EXPERIMENTAL PROCEDURES. The purity of the purified H1.2 was
confirmed by SDS-PAGE and Coomassie staining analysis. E, Mass spectrometric
identification of H1.2 complex. After H1.2 complex was fractionated by 4-20% gradient
SDS-PAGE, bands were excised and subjected to mass spectrometry analysis as
described in EXPERIMENTAL PROCEDURES. Identified components of H1.2 complex
are indicated on the right. Molecular mass markers are indicated on the left. Lane 1,
mock-purified control; lane 2, H1.2 complex. F, Immunoblot confirmation of identified
factors. Purified H1.2 complex was separated by 4-20% gradient SDS-PAGE, and
presence of selected factors was analyzed with indicated antibodies. Lane 1, HeLa
nuclear extract input; lane 2, mock-purified control; lane 3, H1.2 complex. G, Glycerol
gradient centrifugation of H1.2 complex. H1.2 complex, purified on Phosphocellulose
P11 and anti-FLAG antibody affinity columns, was separated by 15-40% glycerol
gradient centrifugation as described in EXPERIMENTAL PROCEDURES. Fractions
were loaded onto 4-20% SDS-polyacrylamide gel, and proteins were detected by silver
staining (one upper panel) or Western blot (six lower panels).
22
Fig. 2-1. Continued.
23
Fig. 2-1. Continued.
24
cellular protein shuttling (Importin7/90), chromatin signaling (WDR5), protein
metabolism (TGase7), apoptosis (hnRNP K) and nucleosome formation (H2A/H2B)
(Bauerle et al., 2002; Grenard et al., 2001; Kouzarides, 2007; Luger, 2006; Moumen et al.,
2005). Another interesting finding is the presence of DNA-PK and PP1 in the H1.2
complex, which are known to phosphorylate and dephosphorylate H1, respectively (Ma
et al., 2005; Paulson et al., 1996). Thus our observations bear an important implication on
a possible competitive action of these two activities to regulate H1.2-dependent processes.
Our mass spectrometry results were further validated by immunoblot analysis using
available antibodies (Fig. 2-1F).
2.2 H1.2 forms a stable complex with its associated factors via direct or indirect
interactions.
In order to obtain a detailed interaction map of H1.2 with its associated factors, we next
analyzed the ability of H1.2 to interact in vitro with individual factors. GST-H1.2 fusion
proteins were pre-bound to glutathione-Sepharose beads and incubated with an equimolar
amount of each of nine selected factors which were prepared as recombinant (PARP1,
FIR and PURα) or in vitro translated (ASXL1, TGase7, CAPER α, β-catenin, YB1 and
WDR5) proteins containing FLAG, Xpress or HA epitope tags at their amino termini (see
EXPERIMENTAL PROCEDURES for details). After extensive washing of the beads,
bound proteins were analyzed by Western blot analysis with anti-FLAG, anti-Xpress or
anti-HA antibody. As shown in Fig. 2-2B, H1.2 was able to directly interact with ASXL1,
25
PARP1, FIR, CAPER α, YB1, PUR α and WDR5, but similar experiments with TGase7
and β-catenin did not show any binding to H1.2. The lack of interactions of any of the
factors with GST alone further confirmed the specificity of their interactions.
To determine if H1.2 and its associated factors are also capable of similar interactions in
vivo, immunoprecipitation was performed with 293T cells transiently expressing FLAG-
or Xpress-H1.2 and one of FLAG-PARP1, Xpress- β-catenin, Xpress-TGase7, HA-FIR,
Xpress-CAPER α, Xpress-YB1, Xpress-PUR α and Xpress-WDR5 (Fig. 2-2C). The
second day after transfection, cell lysates were prepared and subjected to
immunoprecipitation of H1.2 with anti-FLAG or anti-Xpress antibody, and interactions
of co-expressed factors were further analyzed by immunoblotting with antibodies specific
to epitopes within the factors. As shown in Fig. 2-2C, immunoprecipitation of H1.2
resulted in the co-precipitation of PARP1, FIR, CAPER α, YB1, PUR α and WDR5 (lanes
1-3 and 7-9), thus confirming their physical interaction with H1.2 in cellular conditions.
However, consistent with our in vitro results, Western blot analysis showed no detectible
interaction of H1.2 with TGase7 and β-catenin. Reverse immunoprecipitation analysis
using antibodies specific for associated factors (lanes 4-6 and 10-12) also showed the
same interaction of H1.2 with the factors, further confirming the specificity of their
interactions with H1.2.
To further support interaction between H1.2 and its associated factors, we next performed
cellular co-localization analysis. Plasmids encoding EGFP-H1.2 and epitope-tagged
factors (Xpress-ASXL1, FLAG-PARP1, HA-FIR, Xpress-CAPER α, Xpress-YB1,
26
Fig. 2-2. Interaction of H1.2 with its associated factors. A, Purification of GST-H1.2
fusion protein. GST or GST-H1.2 was purified as described in EXPERIMENTAL
PROCEDURES. The purity of the proteins was analyzed by 15% SDS-PAGE and
Coomassie staining analysis. Lane 1, GST; lane 2, GST-H1.2. B, In vitro binding assay
of H1.2 and its associated factors. In vitro translated proteins (Xpress-ASXL1, Xpress- β-
catenin, Xpress-TG7, Xpress-CAPER α, Xpress-YB1 and Xpress-WDR5) and
recombinant proteins (FLAG-PARP1, HA-FIR and PUR α) were incubated with GST or
GST-H1.2 as described in EXPERIMENTAL PROCEDURES. After extensive washing,
bound proteins were analyzed by immunoblot with anti-Xpress (for ASXL1, β-catenin,
TG7, CAPER α, YB1 and WDR5), anti-HA (for FIR), anti-PARP1 and anti-PUR α
antibodies. Lane 1, 10% input; lane 2, GST alone; lane 3, GST-H1.2. C, In vivo binding
assay of H1.2 and its associated factors. FLAG or Xpress-tagged H1.2 was transiently
transfected with its associated factors (FLAG-, HA- or Xpress-tagged), and
immunoprecipitation was performed as described in EXPERIMENTAL PROCEDURES.
Lanes 1, 4, 7 and 10, factor only expression; lanes 2 ,5 ,8 and 11, H1.2 only expression;
lanes 3 ,6 ,9 and 12, H1.2 and factor co-expression. Asterisks indicate non-specific bands.
D, Cellular co-localization of H1.2 with its associated factors. HeLa cells were
transfected with expression vectors encoding EGFP-H1.2 and Xpress-ASXL1, FLAG-
PARP1, HA-FIR, Xpress-CAPER α, Xpress-YB1, Xpress-PUR α or Xpress-WDR5 as
indicated. Cells were fixed with paraformaldehyde and immunostained with anti-FLAG,
anti-Xpress or anti-HA antibody, followed by the Cy3-conjugated secondary antibodies.
After mounted on glass slides with VECTASHIELD with DAPI (Vector Laboratories),
confocal microscopy was performed as detailed in EXPERIMENTAL PROCEDURES.
H1.2 is stained green and its associated factor is stained red. Nucleus is stained blue and
co-localizations of H1.2 and its associated factors are shown in the Merge. The scale bar
represents 5 μm.
27
Fig. 2-2. Continued.
28
Xpress-PUR α and Xpress-WDR5) were constructed and co-transfected into HeLa cells,
and their cellular localizations were analyzed by fluorescence confocal
microscopy.
Consistent with results from the H1.2 stable cell line (Fig. 2-1), our fluorescence
microscopy specifically localized H1.2 to the cell nucleus as bright green spots in all
cases. Indirect immunofluorescence studies also produced the positive red staining for
ASXL1, PARP1, CAPER- α, YB1 and WDR5 mostly in the nucleus, while similar studies
detected FIR in both the cytoplasm and nucleus. In contrast, minimal localization
of
PUR α in nucleus was detected (Fig. 2-2D), possibly due to the lack of its phosphorylation
which is known to govern its
nuclear localization (Barr and Johnson, 2001). Next, co-
localization of H1.2 with expressed factors was examined by superimposing the green
and red optical channels produced by H1.2 and its associated factors. H1.2 displayed
localization patterns
similar to those of ASXL1, PARP1, FIR, CAPER α, YB1, and
WDR5 within the nucleus. Although PUR α was visualized primarily in the cytoplasm,
there was also considerable yellow staining in the nucleus, indicating that at least nuclear
PUR α can be co-localized with H1.2. These results suggest that H1.2 proteins can be
associated with its associated proteins in vivo.
2-3. H1.2 complex represses p53-dependent chromatin transcription.
The finding that specific regulatory factors are associated with H1.2 prompted us to
determine whether these factors have any effect on transcription. To investigate the effect
on DNA and chromatin transcription at the same time, two different transcription
29
templates whose transcription is dependent on p53 were prepared; p53ML-S producing
200 nucleotide long transcript and p53ML-L producing 280 nucleotide long transcript
(Fig. 2-3A). Chromatin was assembled only on p53ML-S DNA with recombinant core
histones using recombinant ACF and NAP1. The transcription reaction contained equal
amounts of p53ML-S chromatin (40 ng) and p53ML-L DNA (40 ng) templates to
simultaneously monitor alteration of chromatin and DNA transcription. Transcription
assays were carried out with p53, p300 and acetyl-CoA as recently described (An et al.,
2004), except that the H1.2 complex or H1.2 was added together with p300 and acetyl-
CoA, as summarized in Fig. 2-3B. Transcription from chromatin template was
completely
dependent upon p53, p300, and
acetyl-CoA, whereas transcription from a histone-free
DNA template was activated only by p53, independent of p300 and acetyl-CoA (Fig. 2-
3C, lanes 1-4). As shown in Fig. 2-3C, H1.2 alone showed only a slight inhibitory effect
on transcription of both chromatin and DNA templates
at the highest concentrations
tested (lanes 5-8). In contrast, when we extended our assays to the H1.2 complex, a
significant
inhibitory effect on chromatin transcription was observed (lanes 9-12). Similar
experiments with DNA templates
failed to reveal any distinct effects of the H1.2 complex
on DNA transcription (lanes 9-12). Addition of the H1.2 complex prior to p53 and/or
p300 also reduced transcription to a level comparable to that observed following
simultaneous addition of the H1.2 complex and p300 (data not shown).
Recent studies proved the contribution of histone acetylation per se in p300-mediated
chromatin transcription (An et al., 2002). Thus, a possible interpretation of our
transcription results is that the H1.2 complex represses
p300-mediated acetylation at the
30
Fig. 2-3. Repressive effects of H1.2 complex in chromatin acetylation and
transcription. A, Schematic representation of transcription templates. Arrows implicate
length of DNA to be transcribed. p53 RE, p53 response element. B, Schematic summary
of transcription and chromatin HAT assays. NTPs and PIC indicate nucleotide
triphosphates and pre-initiation complex, respectively. C, Transcription assays with
recombinant H1.2 and H1.2 complex. p53ML-S chromatin template (40 ng) and p53ML-
L DNA (40 ng) were transcribed with p53 (15 ng), p300 (20 ng) and/or acetyl-CoA (10
μM) as summarized in Fig. 2-3B and as recently described (An and Roeder, 2004). H1.2
complex (200 ng and 400 ng) and recombinant H1.2 (40 ng and 80 ng of H1.2 mixed
with 160 ng and 320 ng of BSA) were used in transcription. Note that results were
obtained from three seperate transcription experiments as indicated by boxes. Data were
quantitated by phosphorimager and normalized to reactions with p53ML-L DNA and p53
alone (100%). D, Chromatin HAT assays with recombinant H1.2 and H1.2 complex.
Chromatin template (100 ng) was incubated with recombinant H1.2 (80 ng of H1.2 mixed
with 320 ng of BSA) or H1.2 complex (400 ng) in the presence of p53 (30 ng), p300 (40
ng) and 2.5 μM [
3
H]-acetyl-CoA. Txn, relative transcription levels; ND, nondetectable.
31
32
promoter region after its recruitment by p53. This possibility was investigated by
checking whether the H1.2 complex is an efficient repressor
of p300-mediated acetylation
of chromatin. Consistent with recent results, p300-mediated acetylation was completely
dependent on p53 that is known to recruit p300 for promoter-targeted acetylation (Fig. 2-
3D, lane 3) (An et al., 2002). In further analysis with the H1.2 complex, we observed a
significant inhibition of p300-mediated acetylation of chromatin templates (lane 5).
When the same concentration of recombinant H1.2 was examined, only a slight inhibitory
effect has been detected (lane 4). Collectively, these results demonstrate that the H1.2
complex can repress p53-dependent transcription from chromatin by down-regulating
p300-mediated acetylation of chromatin.
2-4. H1.2 directly interacts with p53 via its C-terminal domain.
To investigate if the H1.2-complex inhibits p53-mediated transcription by a possible
interaction between H1.2 and p53, we also performed
a series of in vitro protein-protein
interaction assays.
In initial experiments, purified FLAG-p53 protein was incubated
with
GST-H1.2 full length, GST-H1.2 N-terminal domain, GST-H1.2 globular domain or
GST-H1.2 C-terminal domain which was immobilized on glutathione-Sepharose beads.
After rigorous washing, p53 binding was analyzed by Western blot analysis using anti-
FLAG
antibody. As shown in Fig. 2-4A, p53 can bind to the full-length and C-terminal
domains (lanes 10 and 13), but not to the N-terminal and globular domains (lanes 11 and
12), of H1.2. In mapping the region of p53 required for H1.2 binding, we also found that
33
H1.2 interacts with the p53 C-terminal domain (lane 18), but not with the p53 N-terminal
and DNA binding domains (lanes 16 and 17).
On the basis of ability of p53 to directly
interact with H1.2, we also checked whether p53 can interact with the entire H1.2
complex via its recognition of H1.2 present in the complex. Thus immobilized GST-p53
was incubated with the purified H1.2 complex, and factors bound to p53 were identified
by Western blot analysis. As expected, we found that the entire H1.2 complex is indeed
able to bind to GST-p53 but not to GST alone (Fig. 2-4B).
To confirm these in vitro results in vivo, we transiently expressed Xpress-H1.2 and
FLAG-p53 in 293T cells for immunoprecipitation. As shown in Fig. 2-4C, FLAG-p53
was co-immunoprecipitated from
cells in an Xpress-H1.2 dependent manner (lane 3), but
not from control
cells that received the control empty plasmid (pcDNA3.1/His) (lane 1).
These results were further confirmed by an inverse experiment, in which the cell lysate
was
subjected to immunoprecipitation with an anti-FLAG antibody to precipitate p53
(lane 6). To define the region of H1.2 necessary for p53 binding, several H1.2 deletion
mutants were also analyzed for their ability
to interact with p53. Consistent with the in
vitro binding data, H1.2 mutant in which N-terminal region (amino acids 1-34)
was
deleted still retained the ability to bind
to p53 (lane 13). However, when the C-terminal
region
(amino acids 110-213)
of H1.2 was deleted (H1.2 ∆CT), no binding of H1.2 to p53
was
observed (lane 14), indicating that the C-terminal domain (amino acids 110-213)
of
H1.2 is required for p53 association. Indeed, an H1.2 mutant without both N-terminal and
globular domains of H1.2 showed p53 binding comparable to that observed with full
34
length H1.2 (lane 15).
To determine the H1.2-binding
region of p53, we also analyzed one
p53 mutant containing only C-terminal domain and two mutants lacking N- and C-
terminal domains. Consistent with in vitro results (Fig. 2-4A), a p53 C-terminal deletion
mutant (amino acids 1-300) showed no interaction with H1.2 (lane 20), whereas an N-
terminal deletion mutant (amino acids 81-393) showed a wild type level of H1.2 binding
(lane 19). p53 C-terminal domain also showed a strong binding to H1.2, similar to that of
full length p53 (lane 21), arguing that p53 C-terminal domain specifically interacts with
H1.2 C-terminal domain. To further verify cellular interaction between H1.2 and p53 in
physiological conditions, we immunoprecipitated 293T cell lysates with anti-p53
antibody and examined the coimunopreciptation of endogenous H1.2. In addition to p53,
we also could confirm the presence of H1.2 in our immunoprecipitates (Fig. 2-4D).
We next checked whether p53 can interact with the entire H1.2 complex. FLAG-p53 was
co-expressed with Xpress-H1.2 in 293 cells, and cell extracts were prepared and
subjected to immunoprecipitation with FLAG antibody. We checked bound proteins by
Western blot analysis. As shown in Fig. 2-4E, FLAG-p53
was co-immunoprecipitated
with nucleolin, FIR, YB1, PUR α and WDR5 in Xpress-H1.2-expressed cells (lane 3), but
none of these proteins could be found in the control precipitation with mouse IgG (lane 2)
confirming specificity of the precipitation. Together, these experiments demonstrate the
interaction of the H1.2 complex with p53 in vivo and in vitro, which seems to be
mediated through a direct interaction between p53 C-terminal domain and H1.2 C-
terminal domain.
35
Fig. 2-4. Direct interaction of H1.2 with p53. A, p53 interaction with H1.2 in vitro.
GST-H1.2 mutants (lanes 1-4) or GST-p53 mutants (lanes 5-7) were analyzed by SDS-
PAGE and Coomassie staining analysis. For interaction studies, GST-H1.2 mutants and
GST-p53 mutants were incubated with FLAG-tagged p53 and His-tagged H1.2,
respectively. After washing, binding of p53 and H1.2 was analyzed by Western blot
analysis with anti-FLAG or anti-His antibody. Lane 1, GST-H1.2 FL (full length, amino
acids 1-213); lane 2, GST-H1.2 NT (amino acids 1-34); lane 3, GST-H1.2 GD (amino
acids 35-109); lane 4, GST-H1.2 CT (amino acids 110-213); lane 5, GST-p53 NT (amino
acids 1-83); lane 6, GST-p53 DBD (amino acids 120-290); lane 7, GST-p53 CT (amino
acids 290-393); lanes 8 and 14, 10% input of FLAG-p53 and His-H1.2; lanes 9 and 15,
GST control; lanes 10-13, p53 bound to GST-H1.2 mutants; lanes 16-18, H1.2 bound to
GST-p53 mutants. B, p53 interaction with H1.2 complex in vitro. GST (lane 1) or GST-
p53 full length (lane 2) was incubated with H1.2 complex, and pull-down fractions were
analyzed by Western blot analysis using indicated antibodies. C, p53 interaction with
H1.2 in vivo. H1.2 and p53 were expressed in 293T cells and immunoprecipitated using
anti-FLAG and anti-Xpress antibodies as indicated (lanes 1-6). Similar experiments were
also performed after expression of p53 and H1.2 deletion mutants as described in
EXPERIMENTAL PROCEDURES (lanes 10-21). Lanes 1 and 4, p53 only expression;
lanes 2 and 5, H1.2 only expression; lanes 3 and 6, p53 and H1.2 co-expression; lanes 7-
9, expressed H1.2 mutants in whole cell lysates; lanes 10-12, H1.2 only controls; lanes
16-18, p53 only controls; lanes 13-15 and 19-21, H1.2 mutants and p53 mutants co-
expressions. The asterisk indicates non-specific band containing IgG light chain. D,
Mutual interaction of endogenous p53 and H1.2. Whole cell extracts from 293T cells
were immunoprecipiated with anti-p53 antibody (DO-1) and analyzed by Western
blotting with anti-H1.2 and anti-p53 antibodies as indicated. Lane 1, whole cell lysate;
lane 2, control IgG; lane 3, anti-p53 precipitates. E, p53 interaction with H1.2 complex in
vivo. 293T cells were transfected with FLAG-tagged p53 and Xpress-tagged H1.2
expressing plasmids, and cell lysates were prepared two days after transfection. Lysates
were immunoprecipitated with anti-FLAG and analyzed by Western blot analysis using
indicated antibodies. Lane 1, whole cell lysate; lane 2, control IgG; lane 3, anti-FLAG
precipitates.
36
Fig. 2-4. Continued.
37
2.5 Repression of p53-dependent transcription by H1.2 requires YB1 and PUR α
Although the H1.2 complex could repress p53-dependent, p300-mediated chromatin
transcription, it is unclear which factors are mainly involved in this repression. Recent
studies indicated that YB1 can down-regulate p53-induced transactivation of genes
involved in apoptotic process by its interaction with p53 (Homer et al., 2005). Results
from coimmunoprecipitation analysis also suggested that YB1 can directly interact with
PUR α for their functional synergy (Lasham et al., 2000; Safak et al., 1999). Thus, having
found that H1.2 can also interact with both YB1 and PUR α in our studies (above), we
asked whether YB1 and PUR α together with H1.2 can repress p53-dependent, p300-
mediated chromatin acetylation and transcription as observed with the entire H1.2
complex. We first checked the effects of H1.2, YB1 and/or PUR α on chromatin
transcription by using recombinant H1.2, YB1 and PUR α (Fig. 2-1C and 2-5A). As
shown in Fig. 2-5B, p300 stimulatory effect on p53-induced transcription was unaffected
by YB1, PUR α or H1.2 (lanes 4-6). Similar experiments with pair-wise combinations of
H1.2, YB1 and PUR α also showed no detectable change in transcription (lanes 7-9).
However, simultaneous addition of H1.2, YB1 and PUR α resulted in a significant
repression of transcription, supporting functional cooperativity of H1.2, YB1 and PUR α
for transcription repression (lane 10). To test the possibility that transcription repression
by H1.2, YB1 and PUR α might reflect their repressive action on chromatin acetylation,
we also assessed their effect on p53-dependent acetylation of chromatin by p300.
Chromatin acetylation was significantly repressed when H1.2, YB1 and PUR α were
38
added together (lane 20), but not when the three proteins were added individually or in
pairs (lanes 14-19). Since the C-terminal domain of H1.2 was required for p53 interaction
(Fig. 2-4), we next tested the effect of H1.2 C-terminal deletion on chromatin
transcription and acetylation. As expected, deletion of H1.2 C-teminal domain
significantly compromised the repressive effects of H1.2, YB1 and PUR α on p53-
dependent, p300-mediated chromatin acetylation and transcription (Fig. 2-5C).
To validate the conclusions from the in vitro studies described above, p53-deficient
H1299 cells were transfected with p53 expression vector and luciferase reporter construct
(derived from Bax gene) along with plasmids expressing H1.2, YB1 and PUR α, and
luciferase reporter assays were carried out at 48 h after transfection. In agreement with
our in vitro results, expression of H1.2, YB1 and PUR α showed a severe repression of
Bax reporter gene transcription mediated by p53 (Fig. 2-5D, lane 9). However, individual
or pair-wise expression of the three proteins minimally disrupted p53-induced
transcription in all cases (lanes 3-8), again indicating their cooperative action in this
inhibitory process. Furthermore, no significant repression of p53-induced transcription
was observed with C-terminal deleted H1.2, YB1 and PUR α (lane 11). These results are
consistent with the results of the in vitro analyses (Fig. 2-5B and 2-5C) and suggest that
H1.2 C-terminal domain is critical for optimal activities of H1.2, YB1 and PUR α.
In view of the significant effects of H1.2, YB1 and PUR α on p53 transcription, ChIP
analysis was also performed in p53-deficient H1299 cells which were transfected with
p53. Due to the unavailability of ChIP grade antibodies against H1.2, YB1 and PUR α, we
39
expressed Gal4-H1.2, FLAG-YB1 and Xpress-PUR α. We checked the recruitment of
expressed proteins to the Bax p53 response element and GAPDH minimal promoter. As
shown in Fig. 2-5E, our results show the significant level of p53 as well as recruitment of
H1.2, YB1 and PUR α at the response element region of the Bax gene. In contrast, similar
ChIP analyses on GAPDH promoter did not detect YB1 and PUR α, but showed the level
of H1.2 comparable to that observed in the Bax gene, most likely due to its global
localization to alter chromatin structure. These results were reliable, since cells
transfected with an empty vector did not show any detectible precipitation of DNA
fragments. Consistent with ChIP results, RT-PCR analysis also confirmed that H1.2, YB1
and PUR α are capable of significantly repressing p53-induced transcription of the Bax
gene (Fig. 2-5F, lane 4). By contrast, YB1 and PUR α in the absence of H1.2 showed a
modest repressive effect on transcription (lane 3), probably resulted from the minimal
action of endogenous H1.2.
To further confirm the repressive role of YB1, PUR α and H1.2 on transcription of Bax
gene, we knockdowned expression of H1.2, YB1 and PUR α in human U2OS
osteosacoma cells by stably transfecting shRNAs targeting YB1, PUR α or H1.2 (Fig. 2-
5G). Cell strain expressing a vector without a shRNA molecule was used as a control
(lane 1). Our Western analysis confirmed that the cell strain expressing H1.2 shRNA
molecule expressed the much lower level of H1.2, compared to the level in the control
cell strain (lane 2). Interestingly, YB1 and PUR α shRNA repressed expression of YB1
and PUR α, but they also had effect on the level of H1.2 (lanes 3 and 4), perhaps because
40
Fig. 2-5. Functional characterization of H1.2, PUR α and YB1. A, Analysis of
recombinant proteins. Recombinant C-terminal tailless H1.2, PUR α and YB1 were
purified as described in EXPERIMENTAL PROCEDURES. The purity of the purified
proteins was confirmed by SDS-PAGE and Coomassie staining analysis. B, Repressive
action of H1.2, PUR α and YB1 in p300-mediated, p53-dependent transcription.
Chromatin template was transcribed with recombinant H1.2 (50 ng), PUR α (100 ng)
and/or YB1 (100 ng) as indicated. Final protein concentrations in all reactions were
adjusted to be identical by including BSA. C, Requirement of H1.2 C-terminus for
transcriptional repression. Transcription assays were performed as in Fig. 2-5B but with
mutant H1.2 in which the C-terminal tail has been deleted. D, Effect of H1.2, PUR α and
YB1 on p53-dependent transcription in vivo. H1299 cells were transiently transfected
with Bax reporter gene together with vectors that express p53, PUR α, YB1 and/or wild
type/C-terminal tailless H1.2 as indicated. Data are represented as the means ± SEM of
three independent experiments. E, ChIP analysis of Bax promoter. H1299 cells were
mock-transfected (lane 2) or transfected with p53, H1.2, PUR α and YB1 (lane 3), and
subjected to ChIP analysis with antibodies specific for the indicated proteins. A sample
containing 5% total input chromatin was also included for each ChIP assay (lane 1).
Similar ChIP experiments on the GAPDH minimal promoter region were also included as
a control (lanes 4-6). F, RT-PCR analysis of Bax RNA. H1299 cells were transfected
with p53, H1.2, PUR α and/or YB1 as indicated. RT-PCR was performed on total RNA
isolated from transfected or mock-transfected cells. RT-PCR of Actin RNA was used as a
loading control. G, Validation of H1.2, PUR α and YB1 knockdown. Cells were stably
transfected with shRNA of H1.2, YB1 or PUR α, and expression of targeted proteins was
checked by Western blot analysis of cell lysates with anti-H1.2, anti-YB1 and anti-PUR α
antibodies (lanes 2-4). As a control, cells were also transfected with a mock shRNA
vector (lane1). H, Up-regulation of Bax gene transcription upon H1.2, PUR α and YB1
knockdown. RNA was extracted from cells transfected with shRNA of H1.2, YB1 or
PUR α, and relative changes in expression of Bax gene were assessed by Real time PCR.
41
Fig. 2-5. Continued.
42
Fig. 2-5. Continued.
43
YB1 and PUR α could positively regulate H1.2 expression by binding to the CCAAT box
present in the H1.2 gene promoter (lanes 3 and 4) (Meergans et al., 1998; Samuel et al.,
2005). We then checked the effect of depletion of H1.2, YB1 or PUR α on transcription of
Bax gene with or without DNA damage induced by adriamycin treatment. Albeit to a
different extent, three cell strains showed upregulation of Bax gene transcription in all
cases (Fig. 2-5H), strongly suggesting that YB1, PUR α and H1.2 are necessary for
optimal repression of Bax gene.
DISCUSSION
Previous studies addressing the role of H1 in transcription have focused on identifying its
properties in the formation and maintenance of condensed chromatin structure which
could globally inhibit transcription initiation (Bednar et al., 1998; Brown et al., 2006;
Georgel and Hansen, 2001). However, there have been an increasing number of examples
in which H1 plays a more specific role in transcription by differentially acting at the level
of individual genes. The original model for this H1 specificity is based on the results
obtained from gene knockout experiments in Tetrahymena, fungus and yeast (Barra et al.,
2000; Shen and Gorovsky, 1996; Ushinsky et al., 1997). These studies showed that
deletion of H1 gene keeps the organism alive, but specific subsets of genes are
differentially regulated. Other studies with higher eukaryotes also showed that linker
histones have gene selectivity in their repressive actions as shown in the expression of the
MyoD, Xbra, and Bmp-4 genes (Lee et al., 2004; Steinbach et al., 1997). Moreover, a
44
similar specificity of H1 has been shown in a recent study which revealed that
simultaneous inactivation of three out of six H1 subtype genes does not influence global
transcription but primarily affects the activity of specific genes (Fan et al., 2005). These
gene specific effects of H1 might result from its interaction with sequence-specific DNA
binding proteins or specific regulatory factors, as has been shown with Msx1, BAF, SirT1,
HP1 and DFF40 (Daujat et al., 2005; Lee et al., 2004; Montes de Oca et al., 2005;
Vaquero et al., 2004; Widlak et al., 2005).
As a major step toward investigating the effect of H1 on a specific transcription pathway,
we sought to determine if H1 can stably interact with any other proteins with activities
potentially critical for its repressive action on transcription. The significant feature of the
present study is the purification, identification and characterization of the H1.2 complex,
acting as a repressor of p53-dependent, p300-mediated transcription from chromatin. Our
discovery that p300-mediated chromatin acetylation is significantly repressed by the H1.2
complex raises the possibility that it prevents p300 from being recruited to the promoter
region by p53. Another possibility is that the H1.2 complex does not affect recruitment of
p300, but diminish accessibility of core histone tails by inducing localized compaction of
chromatin. However, given the demonstration of a direct interaction of H1.2 with p53, it
is likely that H1.2 acts as an anchoring protein for other regulatory factors which prevent
p53-dependent recruitment of p300 (Fig. 2-6). In all cases, H1.2 itself showed minimal
effects on chromatin acetylation and transcription, further confirming that factors co-
purified with H1.2 play a key role for repressive action of H1.2. This finding is somewhat
45
Fig. 2-6. Model for the promoter-selective inhibition of p53-dependent transcription
by H1 complex. We propose that the H1.2 complex binds to p53 and disrupts p53-
mediated recruitment of chromatin regulating factors to the promoter of target genes.
This repressive chromatin state will in turn interfere with the formation of functional
preinitiation complexes at the promoter to block gene transcription (see DISCUSSION
for details). Ac; acetylation.
46
surprising because previous in vitro studies showed that H1 itself is capable of repressing
chromatin remodeling and transcription (Georgel and Hansen, 2001; Woodcock et al.,
2006). This may reflect the use of higher concentration of H1 in reactions which will
result in reorganization of overall chromatin structure. In fact, we also could detect partial
repression of transcription at a molar ratio higher than two H1 molecules per nucleosome
(data not shown). In this regard, some distinctions need to be made between the
repressive effect of the H1.2 complex that we have purified and the previously reported
effect of H1 as a single structural protein.
Having found that H1.2 engages multiple factors for its repressive action on p53-
mediated transcription, our next question was whether H1.2 requires any specific factors
to elicit its repressive activity. To investigate this potentially important aspect, we have
selected PUR α and YB1 for functional reconstitution of the entire H1.2 complex. Since
YB1 and H1.2 can interact with p53 (Homer et al., 2005, Fig. 2-4A and 2-4C) and PUR α
can stably associate with p53 (Fig. 2-4E), it is possible that PUR α and YB1 can
coordinate the repressive action of H1.2 in transcription. Indeed, our analysis revealed
that H1.2, PUR α and YB1 can closely recapitulate repressive effects of the entire H1.2
complex by blocking p300-mediated chromatin acetylation. These data suggest that
repressive action of the H1.2 complex may be mediated by a subset of factors, at least
one of which is H1.2.
47
Given that several other factors associated with H1.2 are also known as a repressor of
other activators and genes, it will be interesting to sort out key factors involved in various
repressive processes in our future study. For example, our finding that PARP1 is a
component of the H1.2 complex implies that PARP1 may participate as a key factor in
H1-induced chromatin repression. Since PARP1 can physically interact with H1.2 (Fig.
2-2) and p53 (Vaziri et al., 1997; Wesierska-Gadek and Schmid, 2000), it probably can
function as a repressor by facilitating H1.2 interaction with p53. Our results appear to be
contrary to the recent report indicating that H1 and PARP1 exclusively reside in distinct
chromatin domains (An et al., 2004). However, this difference could simply be explained
by the fact that the previous study was conducted with the mixture of all H1 subtypes,
while the present study was undertaken with one of the subtypes, H1.2. Thus our results
can be interpreted as a consequence of specific interaction of PARP1 with H1.2 among
all subtypes. It is also interesting to note that the H1.2 complex contains several cofactors
(e.g., CAPER α and nucleolin) which are known to activate transcription, but transcription
is still significantly repressed by the H1.2 complex. These results may be due to the use
of specific transcription reactions in our studies. For example, previous studies used
estrogen receptor (ER) as an activator to show coactivator function of CAPER α (Dowhan
et al., 2005) whereas our studies used p53 to study the effect of the factors. We assume
that CAPER α in the H1.2 complex minimally contributes to H1.2 action onto p53-
dependent transcription, allowing other factors to retain their repressive action in
transcription. Therefore it will be interesting to test functional contribution of the H1.2
complex in transcription induced by various activators in future studies.
48
Another interesting finding is the purification of free H2A and H2B as components of the
H1.2 complex. Although we do not have a clear explanation, the crystallographic
structure of the nucleosome indicates that the C-terminal domain of H2A is localized in
close proximity to linker DNA where H1 proteins are preferentially localized (Luger,
2006). Thus our results bear an important implication on possible ability of the C-
terminal domain of H2A to interact with H1.2, which will affect the binding of H1.2 to
the nucleosome as a structural component. Furthermore, in view of the diversity in amino
acid sequence and regulation of the synthesis of H1 subtypes (Thiagalingam et al., 2003)
as well as the difference in their distribution with respect to particular genes (Khochbin,
2001; Parseghian and Hamkalo, 2001), it will be important to check if individual
subtypes may have nonredundunt functions in the regulation of specific genes by
associating with distinct factors. The ability to purify factors associated with different H1
subtypes and to analyze their function will be most useful to address these questions.
49
CHAPTER 3: VprBP Antagonizes p53-Mediated Transactivation via
Inhibition of H3 Acetylation.
INTRODUCTION
VprBP was first identified as a protein that has an ability to interact with HIV-1 viral
protein R (Vpr) by coimmunoprecipitation assays (Zhang et al., 2001). VprBP is a 1507-
amino-acid protein that shares the conserved domains including YXXY repeats, Lis
homology domain and WD40 domain. Despite the lack of molecular characterization of
VprBP, recent studies suggest that VprBP can specifically associate with DDB1 to act as
a substrate-recognition subunit of the CUL4-DDB1 ubiquitin E3 ligase complex (Huang
and Chen, 2008; Le Rouzic et al., 2007; McCall et al., 2008; Tan et al., 2007; Wen et al.,
2007; Zhang et al., 2008). Through binding to Vpr, VprBP allows Vpr to modulate the
intrinsic catalytic activity of CUL4-DDB1 complex, which in turn leads to the induction
of G2 phase cell cycle arrest in the virus-infected cells. In addition, VprBP-depleted cells
were shown to activate DNA damage check points and increase the cellular level of CDK
inhibitor p21, linking VprBP to the control of cell cycle arrest and apoptosis (Hrecka et
al., 2007).
p53 is an important tumor suppressor which induces either cell cycle arrest or apoptosis
in response to DNA damage (Menendez et al., 2009; Riley et al., 2008; Vousden and
Prives, 2009). p53 regulates these processes mainly by acting as a sequence-specific
50
transcription factor that regulates expression of a number of target genes. This
transcription reaction is to a large extent regulated at the level of chromatin, which
establishes a physical barrier for the binding of regulatory factors to the promoter region
of a target gene. The fundamental building block of chromatin is the nucleosome core
particle, which is comprised of 147 bp of DNA spooled around a hetero-octamer of
histones H2A, H2B, H3, and H4 (Kornberg and Lorch, 1999; Luger et al., 1997; van
Holde, 1988; Workman and Kingston, 1998). The most dynamic parts of the nucleosome
are amino-terminal domains (called histone “tails”) of core histones, which protrude
away from the DNA. The major contributions of individual histone tails in gene
transcription are made through their posttranslational modifications including acetylation,
methylation, phosphorylation and ubiquitination (Berger, 2007; Kouzarides, 2007; Li et
al., 2007; Peterson and Laniel, 2004). Among these epigenetic processes, histone
acetylation has been implicated as a critical epigenetic mark for activation of p53 target
genes upon DNA damage (An et al., 2004; Donner et al., 2007; Doyon et al., 2004;
Espinosa and Emerson, 2001; Ivanov et al., 2007). While acetylations of all four histone
tails have been linked to active transcription, there is an emerging body of evidence to
support that acetylations of H3 and H4 tails are particularly important for transcription of
p53 target genes (An et al., 2004; Donner et al., 2007; Doyon et al., 2004; Espinosa and
Emerson, 2001; Ivanov et al., 2007; Liu et al., 2003). When cells are exposed to stress
conditions, p53 recruits histone acetyltransferases (HATs) to establish distinct histone
acetylation at its target genes, which will in turn allow the transcriptional machinery to
access the promoters and initiate the high level of transcription. Because histone
51
acetylation is actively regulated by a competitive action of HAT and histone deacetylase
(HDAC) (Katan-Khaykovich and Struhl, 2002; MacDonald and Howe, 2009; Roth et al.,
2001; Shahbazian and Grunstein, 2007), the deregulation of this epigenetic process can
lead to aberrant repression of p53 target genes, ultimately resulting in malignant
transformation and progression of a variety of cancers. Given this reversible nature of
histone acetylation and deacetylation, it would be necessary that cells employ additional
factors which can recognize and lock in a distinct epigenetic status of gene promoter
regions. In relation to the present study, the cellular depletion of VprBP leads to the
increased level of the p53 target gene p21 (Hrecka et al., 2007). These results raise
questions about whether VprBP is able to down-regulate p53-mediated transcription and,
if so, how this would affect cellular responses to DNA damage.
In this study, we characterize the role of VprBP in the p53 signal transduction pathway
and define the mechanisms responsible for the observed phenomenon. We show that
VprBP is recruited by p53 and attenuates p53 transactivation through interaction with
histone H3
tails and inhibition of their acetylation at promoter regions. HDAC1-mediated
deacetylation of H3 tails under non-stressed conditions contributes to the stable
localization of VprBP at p53 target genes. Furthermore, VprBP is over-expressed in
cancer cells, and RNA interference (RNAi)-mediated depletion of VprBP distinctly
augments DNA damage-induced apoptotic cell death. Together, these results reveal a
hitherto unknown function of VprBP in the control of p53-dependent transcription and
DNA damage-induced apoptotic response.
52
EXPERIMENTAL PROCEDURES
Cell culture. 293T, HeLa, LD611 and MCF7 cells were cultured in Dulbecco's modified
Eagle's medium (DMEM) supplemented with 10% FBS. MCF10-2A cells were grown in
1:1 mixture of DMEM and DMEM-F12 supplemented with 20 ng/ml epidermal growth
factor, 100 ng/ml cholera toxin, 0.01 mg/ml insulin, 500 ng/ml hydrocortisone and 5%
horse serum. Urotsa cells were grown in DMEM Low glucose containing 10% FBS.
MLC cells were grown in T medium containing 10% FBS. LNCaP cells were grown in
RPMI-1640 with 10% FBS. Wild type and VprBP
flox/-
MEF cells were propagated in
DMEM supplemented with 10% FBS as previously described (McCall et al., 2008).
Plasmids and expression vectors. p53ML601-14 plasmid was constructed as follows:
The DNA fragment (p53ML) bearing p53 response elements, adenovirus major late
promoter and G-less cassette was isolated from EcoRI digestion of p5GAML array
plasmid (An et al., 2004). To construct p53ML601-7, the p53ML fragment was ligated
into EcoRI sites of p601-7 containing seven direct repeats of the 207 bp 601 nucleosome
positioning sequence. The EcoRI site at the 3’ end of p53ML601-7 was removed by site-
directed mutagenesis. Finally, the 601-7 fragment generated by PCR amplification from
p601-7 was cut with EcoRI and ApoI sites and subcloned into the EcoRI sites at the 5’
end of p53ML601-7 to create p53ML601-14 as illustrated in Fig. 3-2A.
53
Bacterial expression vectors for core histones, Flag-p53 and GST-histone tail fusion
proteins were as described (An et al., 2004; An and Roeder, 2003; Luger et al., 1997).
For construction of GST-fusions of p53 and HDAC1, PCR products were prepared with
primers spanning the corresponding genes and inserted into the BamHI and EcoRI sites
of pGEX-4T1. Bacterial expression constructs of N-terminal domain (NT, amino acids
1-750), LisH and C-terminal domains (LisH+CT, amino acids 751-1507) and C-terminal
domain (CT, amino acids 910-1507) of VprBP were generated by PCR tagging of the
corresponding cDNA and subcloning them into pET-15b or pET-11d vector in frame
with 5’ hexa-His or Flag sequences, respectively. For mammalian expression of HDAC1
and p53, corresponding cDNA fragments were amplified from the appropriate plasmids
by PCR using specific primers and ligated into the correct reading frame of pIRES
containing 5’ Flag or pCDNA3.1/His vector. Further details of plasmid constructions are
available upon request.
Preparation of recombinant proteins and H3 tail peptides. Recombinant histones
were expressed in Escherichia coli Rosetta 2 (DE3) pLysS cells (Novagen) and purified
as described previously (An et al., 2004; Dyer et al., 2004). GST-fused proteins were
expressed in E.coli Rosetta 2 (DE3) pLysS cells and purified on glutathione-Sepharose
4B beads as described (Kim et al., 2008). Bacterially expressed His-tagged and Flag-
tagged VpBP fragments were initially purified with Ni-NTA and M2 agarose,
respectively, and further purified with Q Sepharose and SP-HP column according to
standard procedures. VprBP and p300 proteins were expressed as His-tagged proteins in
54
insect (Sf9) cells using a baculovirus vector and purified by standard procedures with Ni-
NTA affinity chromatography. The peptides corresponding to the N-terminal tail of H3
(amino acids 1-28) were synthesized in Genemed Synthesis Inc (South San Francisco,
CA) by solid-phase Fmoc/tBu chemistry using an automated peptide synthesizer. The
synthesized peptides were purified by RP-HPLC on a Zorbax SB300-C8 column (9.4 mm
× 25 cm) using a water (0.1% TFA) to acetonitrile (0.1% TFA) gradient, and peptide
purity was confirmed using ES-MS and amino acid analysis.
Reconstitution of nucleosome arrays and mononucleosomes. For nucleosome array
reconstitution, the p53ML601-14 plasmid was digested with EcoRI and HindIII, and the
3.4 kb p53ML-601 array DNA fragment was gel purified. For mononucleosome
reconstitution, the 207 bp p53RE DNA fragment containing p53 response elements was
PCR amplified from the p53ML601-14 plasmid using a pair of 5’ biotinylated primers.
Nucleosome array and mononucleosome were reconstituted onto purified DNA
fragments by salt gradient dialysis (Hamiche et al., 1999; Ito et al., 1997). The
reconstituted nucleosome arrays and mononucleosomes were purified from free DNA and
core histones by sedimentation in a 5-30% (vol/vol) glycerol gradient as previously
described (Jaskelioff et al., 2000). Micrococcal nuclease (MNase) digestion of
nucleosome arrays (2 µg) was performed at room temperature for 5 min in 100 µl
reaction containing 2 mM CaCl
2
and 4 mU MNase (Sigma). Reconstituted nucleosome
arrays and mononucleosomes were analyzed on 1% agarose
nucleoprotein gels stained
with ethidium bromide.
55
In vitro transcription and HAT assays. In vitro transcription assays were as described
(Kim et al., 2008) except that 100 ng of p53ML-601 nucleosome arrays or an equimolar
amount of DNA templates was used for each reaction. VprBP (25 or 50 ng) was added
before, after or together with p300 (20 ng) and acetyl-CoA (10 µM). The RNA products
from the reactions were analyzed by gel electrophoresis and autoradiography. For
chromatin HAT assay, p53ML-601 nucleosome arrays (200 ng) were preincubated with
p53 (15 ng) for 20 min and then with p300 (20 ng), acetyl-CoA (10 μM) and/or VprBP
(25 or 50 ng) for another 60 min. HAT reactions were stopped by boiling in SDS-PAGE
loading buffer and analyzed by Western blotting with H3, acetyl-H3, H4 and acetyl-H4
antibodies (Abcam).
H3 tail- and nucleosome-binding assays. To analyze VprBP-tail interaction in vitro,
GST-histone tails (2 µg) were immobilized on glutathione-agarose
beads and incubated
with His-VprBP (2 µg) for 16 h at 4 °C in 750 µl of binding buffer (25 mM Tris-HCl, pH
7.8, 0.2 mM EDTA, 20% glycerol, 150 mM KCl, 0.1% Nonidet P-40). The beads were
washed three times with binding buffer to remove VprBP not bound to histone tails, and
bound proteins were subjected to SDS–PAGE followed by immunoblotting with anti-His
antibody (Novagen). To identify the tail binding domain of VprBP, similar pulldown
experiments were performed with 2 µg of N-terminal domain (NT, amino acids 1-750),
LisH and C-terminal domains (LisH+CT, amino acids 751-1507) or C-terminal domain
(CT, amino acids 910-1507). For H3 tail peptide binding experiments, His-VprBP (2 µg)
56
was coupled with Ni-NTA agarose beads and incubated with unmodified or acetylated
H3 peptides (2 µg) for 16 h at 4 °C in the binding buffer. The reaction mixtures were
then pelleted by centrifugation, and the beads were washed three times in washing buffer
(25 mM Tris-HCl, pH 7.8, 0.2 mM EDTA, 20% glycerol, 200 mM KCl, 0.1% Nonidet P-
40). The VprBP protein bound to the coupled peptides was resolved by 4-20% SDS-
PAGE and analyzed by silver staining. For nucleosome binding assays, 2 µg DNA
equivalents of mononucleosomes were preacetylated by p53 (150 ng) and p300 (200 ng)
supplemented with acetyl-CoA (10 µM) and immobilized on Streptavidin agarose
(Novagen). After extensive washing with washing buffer (25 mM Tris-HCl, pH 7.8, 0.2
mM EDTA, 20% glycerol, 300 mM KCl, 0.1% Nonidet P-40), His-tagged VprBP was
added to the nucleosomes and incubated in 500 µl of binding buffer at 4 °C for 16 h. The
beads were washed three times with binding buffer,
and nucleosome-bound VprBP was
detected
by Western blotting using the anti-His antibody.
Protein-protein interaction. For GST pull-down assays, His-tagged VprBP (2 µg) was
incubated with GST-fused p53 (2 µg) or HDAC1 (2 µg) immobilized on glutathione-
agarose
beads in 750 µl of binding buffer (25 mM Tris-HCl, pH 7.8, 0.2 mM EDTA, 20%
glycerol, 150 mM KCl, 0.1% Nonidet P-40) for 16 h
at 4 °C with gentle rotation. After
washing beads three times with binding buffer, bound
VprBP was resolved by 8% SDS-
PAGE and detected by Western blot
analysis using anti-His antibody. Similar in vitro
binding assays were carried out using Flag-p53 (2 µg) and GST-HDAC1 (2 µg), which
were analyzed by Western blotting with anti-Flag antibody. For in vivo interaction
57
studies, 293T cells were transiently transfected with expression vectors encoding Myc-
VprBP, Flag-p53, Flag-HDAC1 and His-HDAC1. Two days after transfection, cells
were solubilized and the cleared lysates were subjected to immunoprecipitation with anti-
Flag (Sigma), anti-Myc (Invitrogen) or anti-His antibody. The bound proteins were
eluted by boiling in SDS sample buffer and analyzed by Western blot analysis. For co-
immunoprecipitation of endogenous proteins, 293T cell lysates (1 mg) were
immunoprecipitated using anti-p53 (DO1, Santa Cruz Biotechnology, Inc.) or anti-
HDAC1 antibody (Active Motif) and immunoblotted with anti-VprBP (Proteintech
Group, Inc), anti-HDAC1 or anti-p53 (FL-393, Santa Cruz Biotechnology, Inc.) antibody.
Reverse transcription PCR. For quantitative reverse transcription PCR analysis of p21
and Noxa gene expression, total RNA was isolated using the RNeasy mini kit (Qiagen)
according to the manufacturer’s instructions. Quantitative PCR was performed using the
IQ SYBR Green Supermix (Bio-Rad) and the IQ5 real time cycler (Bio-Rad). Assays
were normalized to β-actin mRNA levels. All reactions were run in triplicate, and data
presented is the average of three individual experiments. The primers used for qPCR are
listed in the Table 3-1.
Chromatin immunoprecipitation (ChIP) and RNA interference. ChIP assays with
293T cells, either treated or not treated with etoposide, were performed using the ChIP
assay kit from Upstate/Millipore according to the manufacturer’s protocol and as recently
described (Kim et al., 2008). Antibodies specific to VprBP, H3, Acetylated H3, p53
58
59
(DO-1) and HDAC1 were used for immunoprecipitation. Primers used for quantitative
real time PCR (RT-qPCR) are listed in Table 3-1. All samples were run in triplicate and
results were averaged. The positions of the PCR primers within the promoter regions are
shown in Fig. 3-4.
For shRNA-based knockdown experiments with 293T cells, DNA oligonucleotides
encoding shRNAs specific for VprBP mRNA (5’-AATCACAGAGTATCTTAGA-3’),
p53 mRNA (5’-GACTCCAGTGGTAATCTAC-3’) and HDAC1 mRNA (5’-
GCAGATGCAGAGATTCAAC-3’) were subcloned into the H1 promoter-driven vector
pSUPER.puro (OligoEngine). 293T cells were transfected with the indicated shRNA
expression constructs for 72 h, and then subjected to ChIP analysis. For gene knockdown
experiments with LD611, LNCaP and MCF7 cells, 21 nucleotide siRNA duplexes with 3 ′
dTdT overhangs corresponding to VprBP mRNA (5’-UCACAGAGUAUCUUAGAGA-
3’) were synthesized by the DNA Core Facility at the University of Southern California.
Cells were transfected with siRNA oligonucleotides (60 µM) or negative control siRNA
(Ambion). At 48 h post-transfection, cells were treated with or without etoposide (100
µM) and harvested and subjected to qRT-PCR analysis.
Cell viability and apoptosis assay. The cytotoxicity of etoposide treatment was
estimated by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay
according to the standard protocol. Briefly, HeLa and 293T cells were transfected with
VprBP expression vector or empty vector for 24 h, and treated with or without etoposide
(100 µM). After washing cells with PBS, 1 ml of MTT (0.5 mg/ml) was added to cells
60
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. MTT assays with wild type and VprBP
flox/-
MEF cells were
performed as with HeLa and 293T cells, except that cells were first transfected with Cre
recombinase peptides to delete VprBP gene as recently described (Peitz M 2002). For
apoptosis assay, LD611, LNCaP, and MCF7 cells were cultured in 100 mm plates and
treated with or without etoposide (100 µM) for 24 h. After the treatment, the cells were
harvested
with trypsin/EDTA, resuspended in the binding buffer and then
incubated with
FITC-conjugated Annexin V and propidium iodide (FITC Annexin V apoptosis detection
Kit I; BD Pharmingen),
according to the manufacturer's protocol. The numbers of
apoptotic
cells were monitored with flow cytometry.
RESULTS
3-1. VprBP regulates cell viability and p53 target gene expression.
As an initial step toward gaining insights into the biological function of VprBP, we first
investigated its effect on cell proliferation. HeLa and 293T cells were transiently
transfected with an expression plasmid encoding VprBP or empty vector, and the cell
viability was determined by the MTT assay after etoposide or control DMSO treatment.
The expression of VprBP caused a substantial increase in the viability of both HeLa and
61
293T cells following DNA damage (Fig. 3-1A). In an attempt to further define the role
of VprBP in the control of cell viability, we have employed conditional VprBP
flox/-
MEF
cells that allow switching on and off of VprBP expression under the control of Cre
recombinase. As determined by quantitative Western blot analysis, the transfection of
wild type MEF cells with cell permeable TAT-Cre recombinase didn’t affect VprBP
expression, but the same treatment of VprBP
flox/-
MEF cells with the TAT-Cre
recombinase resulted in the almost complete ablation of VprBP expression (Fig. 3-1B,
lanes 1-4). Viability assays of wild type and VprBP
flox/-
MEF cells revealed no
significant difference in cell viability upon etoposide treatment without Tat-Cre
recombinase transfection (Fig. 3-1B, right panel, -Cre). By contrast, MTT assays
revealed that cell viability upon etoposide treatment was markedly reduced by Cre
recombinase-mediated deletion of VprBP gene in VprBP
flox/-
MEF (Fig. 3-1B, right panel,
+Cre), implicating VprBP as having a positive effect on cell survival.
To address whether the observed contribution of VprBP to cell viability is accompanied
by an alteration in p53 pathway, we next measured the level of transcription of two
known p53 target genes, p21 and Noxa. As summarized in Fig. 3-1C, HeLa and 293T
cells transfected with VprBP demonstrated a distinct decrease in DNA damage-induced
transcription of the genes, compared to control cells transfected with empty vector.
When a possible contribution of VprBP to p53-responsive gene transcription was
measured in wild type and VprBP
flox/-
MEF cells, a distinct boost in DNA-damage-
induced transcription of p21 and Noxa was detectable after Cre-mediated deletion of
62
Figure 3-1. Effects of VprBP overexpression and knockout on cell viability. A,
Increased cell viability by VprBP overexpression. HeLa and 293T cells were transiently
transfected with empty vector or VprBP expression vector for 24 h, and then treated with
etoposide (100 µM) for 24 h. Cell viability was measured by analysis of MTT
conversion. All reactions were performed in triplicate. B, Decreased cell viability by
VprBP knockout. Wild type and VprBP
flox/-
MEF cells were treated with Cre
recombinase peptides for 72 h, and VprBP depletion was confirmed by Western blot
analysis using anti-VprBP antibody (lanes 1-4). Cell viability assays were identical to
Fig. 3-1A. C, Transcriptional repression of p53-responsive genes by VprBP
overexpression. Transfection with empty or VprBP expression vector and etoposide
treatment were identical to Fig.3-1A. mRNA levels were analyzed by quantitative real-
time reverse transcription PCR. Average and standard deviation of three independent
experiments are shown. D, Transcriptional activation of p53-responsive genes by VprBP
knockout. Wild type and VprBP
flox/-
MEF cells were treated with etoposide as in Fig. 3-
1B, and total RNA was isolated and analyzed by quantitative reverse transcription PCR
analysis as in Fig. 3-1C.
63
Figure 3-1. Continued.
64
VprBP gene in VprBP
flox/-
MEF cells, but not in wild type MEF cells (Fig. 3-1D). Taken
together, these observations imply that VprBP negatively regulates p53 target gene
transcription.
3-2. VprBP represses p53-mediated transcription from chromatin
The fact that changes in cellular level of VprBP directly correlate with alterations in p21
and Noxa transcription (Fig. 3-1C and 3-1D) suggests that VprBP may be involved in the
p53 transcription pathway. To check this possibility, we next prepared the recombinant
VprBP (Fig. 3-2B) and examined its effects on p53-mediated transcription using a cell
free assay system. For reconstitution of linear nucleosome arrays, we employed p53ML-
601 array template containing p53 response elements and the adenovirus major late core
promoter and, on both sides, seven direct repeats of the 601 nucleosome positioning
sequence (Fig. 3-2A). Unmodified 17-mer nucleosomal arrays were assembled from
recombinant histone octamers (Fig. 3-2C) and p53ML-601 array DNA templates by using
a salt gradient dialysis method. After confirming the successful reconstitution by
electrophoretic mobility shift and partial micrococcal nuclease digestion (Fig. 3-2D and
3-2E), transcription assays were carried out with p53 and p300 HAT as summarized in
Fig. 3-2F. Transcription from p53ML-601 nucleosome arrays was completely
dependent
upon p53, p300 HAT, and acetyl-CoA,
whereas transcription from p53ML-601 DNA
template showed a dependence only on the activator
p53 (Fig. 3-2G, lanes 1-3). When
the effect of VprBP was examined, we found that addition of VprBP prior to p300
significantly decreased the level
of transcription from nucleosome array templates (Nuc.
array, lanes 4 and 5). However, no detectable change in transcription was observed upon
65
simultaneous addition of VprBP and p300 or sequential addition of VprBP after p300
(Nuc. array, lanes 10, 11, 16 and17). A similar transcription analysis with DNA
templates also failed to show any effect of VprBP on transcription (Fig. 3-2G, DNA),
suggesting that the observed repression of chromatin transcription is likely to be the
results from interference of chromatin modifying activities.
Because the primary role of acetylation of both H3 and H4 tails has been well-illustrated
in p53 transcription (An et al., 2004; Doyen et al., 2006; Espinosa and Emerson, 2001;
Ivanov et al., 2007), a possible interpretation of these results is that
VprBP could repress
p300-mediated acetylation of H3 and H4 at the
promoter region. This possibility
was
investigated by checking whether VprBP is also repressive for acetylation of H3 and H4
within reconstituted nucleosome arrays. Western blot of HAT reactions confirmed that
p300-mediated histone acetylation in the context of nucleosome arrays is
completely
dependent on p53, which is known to recruit p300
for promoter-targeted acetylation (Fig.
3-2H, lanes 1-3). In parallel experiments with VprBP, we observed a significant
inhibition of H3 acetylation, but not H4 acetylation, by addition of VprBP prior to p300
(lanes 4 and 5), indicating selective repressive effect of VprBP on H3 acetylation reaction.
As expected from transcription assays, there was no apparent repression of H3
acetylation, when VprBp was added after or simultaneously with p300 (lanes 10, 11, 16
and 17). When the same concentration of the heat-inactivated VprBP was
examined for
transcription and HAT assays, no inhibitory effect was detected (Fig.3-2G and 3-2H,
66
Fig. 3-2. VprBP-mediated repression of chromatin transcription and H3 acetylation.
A, Schematic depiction of the p53ML-601 nucleosome array. The p53ML-601 array
template contains the central three nucleosome length (594 bp) fragment bearing p53
response elements, adenovirus major late promoter and G-less cassettes flanked on either
side by seven repeats of 207 bp 601 nucleosome positioning sequences. B, Analysis of
purified VprBP by 8% SDS-PAGE and Coomassie blue staining. Molecular size markers
are indicated in kDa. C, Preparation of recombinant histone octamers. Reconstituted
histone octamers were analyzed by 15% SDS-PAGE and Coomassie blue staining. Lane
1, intact histone octamer; lane 2, histone octamer containing H3 mutated at the four major
acetylation sites (K9, K14, K18 and K23). D, 1% agarose gel of nucleosomal arrays
reconstituted with the 3.4 kb p53ML-601 array DNA fragment. The nucleoprotein gel
was ethidium bromide-stained, and major nucleosome array species are indicated by
arrow (lane 3). Lane 1: 1kb DNA ladder; lane 2: free p53ML-601 array DNA fragment.
E, Micrococcal nuclease (MNase) digestion of the p53ML-601 nucleosome arrays.
Nucleosome arrays were reconstituted on the 3.4 kb p53ML-601 array DNA fragment
and digested with MNase (4 mU) for 5 min to generate a ladder of partial digestion
products (lanes 2 and 3). A 123 bp DNA ladder was used as the size marker (M, lane 1).
The predicted position of nucleosomes is indicated to the right by arrows. F, Outline of
chromatin HAT and transcription assays. Abbreviations: PIC, preinitiation complex;
NTPs, nucleotide triphosphates. G, Repressive effect of VprBP on chromatin
transcription. p53ML-601 nucleosome array or histone free p53ML-601 DNA was
transcribed in the presence of p53, p300, acetyl-CoA and/or VprBP as indicated. Prior to
transcription, p300 and VprBP were added together or sequentially as indicated. All
transcriptions were conducted simultaneously under identical conditions, so that the
transcription signals are directly comparable. Heat-inactivated VprBP was used in
control transcription reactions and marked by five point star (lanes 6, 12 and 18).
Nucleosome free p53ML-601 array is labeled DNA, while p53ML-601 nucleosome array
is labeled Nuc. array. Data were quantitated by phosphoimager, and the results shown
are representative of three independent experiments. H, Repressive effect of VprBP on
H3 acetylation. p53ML-601 nucleosome array was incubated with p53, p300, acetyl-
CoA and/or VprBP as summarized in Fig. 3-2F. Acetylation of nucleosome arrays was
detected by Western blotting with anti-acetyl H3 and H4 antibodies ( α-AcH3 and α-
AcH4). Western blot analyses of H3 and H4 confirmed equal loading of histones ( α-H3
and α-H4).
67
Fig. 3-2. Continued.
68
Fig. 3-2. Continued.
69
lanes 6, 12 and 18), further supporting specificity of VprBP-induced repression of H3
acetylation and transcription of nucleosomal arrays.
3-3. VprBP binds to unmodified H3 tail in the context of nucleosome
A possible mechanism underlying the inhibitory action of VprBP in transcription is that
VprBP interacts with histone tails, especially H3 tails, to repress their acetylation. To test
this possibility, we examined interactions of VprBP with a fixed concentration of GST-
histone tail fusion proteins pre-bound to glutathione-Sepharose beads. After extensive
washing of the beads, VprBP binding was determined by Western blotting with anti-His
antibody. As shown in Fig. 3-3A, we found that VprBP binding was highly selective for
H3 tail relative to other histone tails. Because VprBP repressive effect was lost when
nucleosomal arrays were preacetylated by p300 (Fig. 3-2G), we further examined the
effect of tail acetylation on tail-VprBP interaction using synthetic H3 tail peptides. When
assayed with immobilized His-VprBP, unmodified H3 tail peptides showed a distinct
interaction with VprBP, as confirmed by silver staining after SDS-PAGE of the reactions
(Fig. 3-3B, lane 4). By contrast, similar binding experiment with H3 tail peptides bearing
lysine acetylations at K9, K14, K18 and K23 showed no detectable interaction of VprBP
(lane 5). Additionally, parallel binding experiment with mononucleosomes reconstituted
on a 207 bp p53RE DNA fragment (Fig. 3-3C and 3-3D) showed VprBP binding both to
nucleosomes containing wild type H3 and nucleosomes containing lysine-mutated H3
(Fig. 3-3E, lanes 3 and 5). Notably, however, the observed VprBP-nucleosome
interaction was distinctly inhibited by p300-mediated H3 acetylation, as reflected by an
70
Fig. 3-3. Selective interaction of VprBP with unmodified H3 tail. A, Selective
interaction of VprBP with H3 N-terminal tail. His-tagged VprBP was tested for binding
to GST (lane 2) or GST-histone tail fusion (lanes 3-6) proteins. VprBP binding to histone
tails was determined by Western blot analysis using anti-His antibody. Lane 1 represents
10% of VprBP used in the binding reactions. B, Selective interaction of VprBP with
unmodified H3 tail. Unmodified (H3) and acetylated (AcH3) H3 peptides (amino acids
1-28) were synthesized and incubated with His-tagged VprBP immobilized onto Ni-NTA
agarose beads. After extensive washing, bound H3 peptides were resolved in a 4-20%
SDS-PAGE and silver-stained. Input lanes 1 and 2 represent 10% of proteins used in the
binding reactions. C, Analysis of reconstituted p53RE nucleosome. Mononucleosomes
containing wild type (lane 3) or acetylation site-mutated (lane 4) H3 were reconstituted
with p53RE DNA fragment, purified by glycerol gradient and analyzed by 1 % agarose
nucleoprotein gels. Lane 1: 100 bp DNA ladder; lane 2: free p53RE DNA fragment. D,
HAT assays with p53 RE nucleosomes. p53RE nucleosomes (200 ng) containing wild
type (lanes 1-3) or acetylation site-mutated (lanes 4-6) H3 were incubated with p53 (15
ng), p300 (10 ng) and acetyl-CoA (10 µM) as described in Materials and Methods. H3
acetylation was analyzed by Western blot analysis (higher panel). Histone H3 was used
as loading control (lower panel). E, Preferential binding of VprBP to unmodified
nucleosome. Nucleosomes containing wild type or mutant H3 were reconstituted on
biotinylated 207 bp p53 RE and incubated with p300, p53 and/or acetyl-CoA. After
immobilizing reconstituted nucleosomes and free DNA on Streptavidin agarose beads,
the interaction assays were performed with VprBP. The presence of VprBP in the beads
was analyzed by Western blotting with anti-His antibody.
71
Fig. 3-3. Continued.
72
apparent decrease in binding of VprBP to wild type H3 nucleosome, but not to H3
mutated nucleosome (Fig. 3-3E, lane 4 versus lane 6).
3-4. The LisH domain of VprBP is required for its repressive action
VprBP contains an evolutionarily conserved Lis homology (LisH) domain in the central
region and several WD40 repeats in the carboxyl terminus (Fig. 3-4A). Since these
motifs are often critical for known functions of full length proteins, we were interested in
determining which part of VprBP is required for its repressive action. To this end, three
different VprBP deletion mutants were prepared from bacteria (Fig. 3-4B) and tested in
transcription and HAT assays. N-terminal region (residues 1-750, NT) and C-terminal
region containing WD40 repeats (residues 910-1507, CT) showed no detectable change
in p53-dependent, p300-mediated transcription of nucleosome arrays (Fig. 3-3C, Nuc.
array, lanes 4, 5, 10 and 11). In striking contrast, C-terminal fragment fused
evolutionarily conserved LisH domain (residues 751- 1507, LisH+CT) repressed the
nucleosome array transcription to a level comparable to that observed with full-length
VprBP (Nuc. array, lanes 7 and 8). As expected, DNA transcription was not altered by
these VprBP deletion mutants in all cases (Fig. 3-4C, DNA). In parallel HAT assays with
nucleosome arrays, we also observed a significant inhibition of p300-mediated
acetylation of H3 by LisH+CT fragment, but not by NT or CT fragment (Fig. 3-4D, α-
AcH3). Because
VprBP binds to unmodified H3 tails
to act as a transcription repressor
(Fig. 3-3A and 3-3B), the above results argue that LisH domain might be important for
the interaction between VprBP and H3 tails. In fact, GST-pull-down assays with
73
Fig. 3-4. Requirement of LisH domain for VprBP action. A, Schematic illustration of
VprBP deletion mutants. Numbers indicate amino acid residues. The Lis homology
motif (LisH) and WD40 repeat motif (WD40) are indicated. NT: N-terminal domain;
LisH+CT: Lis homology and C-terminal domain; CT: C-terminal domain. B, Preparation
of VprBP deletion mutants. His-tagged VprBP proteins were prepared as described in
Materials and Methods and analyzed on 8% SDS-PAGE stained with Coomassie blue.
Lane 1: N-terminal domain; lane 2: Lis homology and C-terminal domain; lane 3: C-
terminal domain. C, Differential effects of VprBP deletion mutants on chromatin
transcription. Transcription reactions were essentially as described in Fig. 3-2G, but
contained VprBP deletion mutants which were added prior to p300. The asterisk
indicates the usage of heat-inactivated VprBP. D, Differential effects of VprBP deletion
mutants on H3 acetylation. HAT assays were performed as in Fig. 3-2H, but with the
indicated VprBP deletion mutants. The heat-inactivated VprBP (marked with asterisk)
was also included. E, Preferential binding of VprBP LisH domain to H3 tail. GST-pull
down assays were performed with GST-H3 tail and different deletion VprBP constructs
as indicated. Input (lane 1) represents 10% of mutant proteins used in the binding
experiments. F, Preferential binding of VprBP LisH domain to unmodified H3 tail.
Binding assays were performed using synthetic H3 tail peptides as in Fig. 3-3B, but with
the indicated VprBP deletion mutants. The asterisks indicate the positions of unmodified
and acetylated H3 tail peptides. Lanes 1 and 2 show 10% of tail peptides used in the
binding reactions.
74
75
deletion mutants of VprBP revealed that H3 tails can interact directly with LisH+CT
fragment, but not with NT and CT fragments (Fig. 3-4E, lane 3), indicating that LisH
domain is responsible for the interaction of VprBP with H3 tails. Similar binding
experiments with unmodified or acetylated H3 tail peptides also showed that, in contrast
to NT and CT fragments, LisH+CT fragment of VprBP can bind to these peptides (Fig. 3-
4F, lanes 4 and 5). However, LisH+CT fragment showed a marked preference for
unmodified H3 peptides over acetylated H3 peptides (compare the asterisk-marked
bands), underscoring the specificity of the interaction and verifying the importance of
LisH domain in VprBP interaction.
3-5. VprBP down-regulates p53 target genes by modulating histone acetylation.
To assess the in vivo relevance of our in vitro results, we checked whether VprBP can
antagonize the transcription of p53 target gene in response to DNA damage by RNAi-
complemented ChIP assays. As first confirmed by Western blot analysis, the transfection
of 293T cells with shRNA targeting VprBP efficiently depleted VprBP (Fig. 3-5A, lanes
1 and 2). In checking the transcription of two p53-responsive genes, p21 and Noxa, by
quantitative reverse transcription PCR, we found that VprBP knockdown significantly
enhanced DNA-damage-induced activation of p21 and Noxa to a comparable extent (Fig.
3-5B). Similar experiments without DNA damage also showed, although to a lower level
than that obtained with DNA damage, a distinct boost in transcription of p21 and Noxa
genes after VprBP knockdown (Fig. 3-5B).
76
When ChIP experiments were performed at the p21 promoter region in VprBP-depleted
293T cells under normal conditions, we detected a dramatic reduction in the promoter
occupancy of VprBP, but a minimal change in the level of p53 (Fig. 3-5C, top panel).
Given the dependence of unacetylated state of H3 tails for VprBP-H3 tail interaction (Fig.
3-3B and 3-3E), we further examined the effect of VprBP depletion on promoter
localization of HDAC1, which acts as a key epigenetic repressor of p21 transcription by
blocking histone acetylation (Duan et al., 2005; Lagger et al., 2003). Significantly, the
VprBP depletion led to the decrease of HDAC1 occupancy and the concomitant
accumulation of acetylated H3 at the promoter region (top panel). In parallel ChIP
experiments using p53-depleted cells, we also detected near complete loss of VprBP as
well as a dissociation of HDAC1, and a significant reduction of H3 acetylation (middle
panel). This observation strongly suggests that the initial recruitment of VprBP and
HDAC1 to the p21 promoter is mediated by p53. One anomaly in this result is that the
dissociation of HDAC1 from the p21 promoter coincides with the reduction of H3
acetylation (middle panel, AcH3). The decline of H3 acetylation may be due to the
failure of p53-mediated recruitment of a HAT, which would initially acetylate H3. Also
consistent with our results indicating a stable association of VprBP with unmodified H3
tail, our ChIP assays after HDAC1 knockdown showed a significant decrease in promoter
localization of VprBP and HDAC1 but no change in p53 occupancy (bottom panel). To
explore the similar effect of VprBP on other p53 target genes, we also carried out the
same analysis on Noxa gene. Consistent with the results from p21 gene, shRNA-
77
mediated depletion of VprBP at the Noxa promoter region was found to coincide with
dissociation of HDAC1 and apparent increase in H3 acetylation (Fig. 3-5D, top panel).
Depletion of p53 and HDAC1 also resulted in a dramatic reduction in
immunoprecipitation of the promoter region using VprBP antibody (middle and bottom
panels), strongly supporting that the VprBP plays an inhibitory role in other p53 target
genes as well.
To further investigate the repressive role of VprBP, we performed ChIP analyses with
293T cells that were either untreated or treated with DNA damaging agent etoposide (Fig.
3-5E). Predictably, etoposide-induced DNA damage resulted in an accumulation of p53
and a large increase in H3 acetylation at the p21 promoter region (upper panel). In
correlation with the increased level of H3 acetylation, the release of VprBP from the
promoter region was observed (upper panel). ChIP assays also showed that the release of
VprBP from the p21 promoter region coincides with decreased occupancy of HDAC1 at
the promoter region (upper panel), suggesting that HDAC1 binding and H3 deacetylation
at sites surrounding p21 promoter are dependent on VprBP. In contrast, a minimal
alteration in the level of p53, VprBP, H3 acetylation and HDAC1 at the distal region was
observed upon DNA damage in all cases. We also repeated the entire experiments on
Noxa gene; again similar results were obtained from these parallel experiments (Fig. 3-
5E, lower panel), and these results again argue for the possibility that the repressive
action of VprBP pertains to other p53 response genes.
78
Fig. 3-5. Dynamics of promoter occupancy of VprBP, p53 and HDAC1. A, Depletion
of VprBP, p53 and HDAC1. 293T cells were transfected with VprBP shRNA (lane 2),
p53 shRNA (lane 4), HDAC1 shRNA (lane 6), or control shRNA (lanes 1, 3 and 5) and
Western blot analyses were performed at day 3 post-transfection with shRNA. Actin was
used as an internal control (lower panel). B, p53 target gene activation by VprBP
knockdown. VprBP-depleted and control 293T cells were treated with 100 µM etoposide
for 8 h, and mRNA levels were analyzed by quantitative real-time reverse transcription
PCR. Average and standard deviation of three independent experiments are shown. C,
VprBP depletion-promoted H3 acetylation at p21 promoter. VprBP, p53 and HDAC1
were depleted as in Fig. 3-5A, and ChIP assays of promoter and distal regions were
performed using antibodies specifically recognizing VprBP, p53, HDAC1, H3 and
acetylated H3. Input DNA and immunoprecipitated DNA were quantified by quantitative
real-time PCR (qPCR) analyses using distal and proximal primer sets. The results are
shown as percentage of input, and the error bar indicates the means ± S.E. Shown on the
top is schematic diagram of p21 promoter and distal regions subjected to ChIP analysis.
D, VprBP depletion-promoted H3 acetylation at Noxa promoter. ChIP analyses were
essentially as described in Fig. 3-5C, but over the Noxa gene promoter. E, DNA damage-
induced dissociation of VprBP. 293T cells were treated with or without etoposide (100
µM) for 4 h, and then analyzed by ChIP analysis of p21 and Noxa promoters as described
in Fig. 3-5C and 3-5D.
79
80
81
82
3-6. VprBP physically interacts with p53 and HDAC1
To determine if the recruitment and repressive action of VprBP at p53 target genes reflect
its direct interaction with p53 and HDAC1, we performed a series of interaction studies.
We initially determined whether VprBP and HDAC1 can directly interact with p53 in
vitro by GST-pull-down assays. In these experiments, GST-p53 and GST-HDAC1
immobilized on Sepharose beads were incubated with His-VprBP and Flag-p53, and the
immobilized proteins were then analyzed by immunoblot after extensive washing. As
shown in Fig. 3-6A, His-VprBP is specifically precipitated from the reaction by both
GST-p53 and GST-HDAC1 (lanes 3 and 6). In contrast, pull-down experiments in which
GST-HDAC1 was used under the same conditions failed to show any detectable binding
to Flag-p53. To validate these in vitro results in vivo, extracts from 293T cells
transiently transfected with expression plasmids for Myc-VprBP, Flag-p53 and Flag-/His-
HDAC1 were immunoprecipitated using anti-Flag antibody. Consistent with in vitro
binding data, our Western blot analysis showed a strong interaction of VprBP with p53
and HDAC1 (Fig. 3-6B, lanes 3 and 6), but no detectable interaction between p53 and
HDAC1 was observed in our analysis (lane 9). The interaction of VprBP with p53 and
HDAC1 in physiological conditions was further tested by western blot analysis of α-p53
or α-HDAC1 immunoprecipitates from extracts of 293T cells. Again, both p53 and
HDAC1 were coimmunoprecipitated with VprBP, but HDAC1 failed to show
coimmunoprecipitation of p53 (Fig. 3-6C). As indicated in the control reactions, these
results cannot be attributed to nonspecific interactions.
83
Fig. 3-6. Physical association of VprBP with p53 and HDAC1. A, VprBP interaction
with p53 and HDAC1 in vitro. His-tagged VprBP or Flag-tagged p53 was incubated with
GST or GST-fused p53 or HDAC1, and bound proteins were analyzed by Western
blotting using anti-His and anti-Flag antobodies as indicated. Input corresponds to 10%
of the material used in the binding reactions. B, VprBP interaction with p53 and HDAC1
in vivo. 293T cells were transfected for 48 h with epitope tagged forms of VprBP, p53
and/or HDAC1. Whole cell lysates were prepared and subjected to immunoprecipitation
with anti-Flag antibody followed by Western blotting with indicated antibodies (IP: α-
Flag). Cell extracts were also analyzed by Western blotting to confirm that equivalent
amounts of proteins from each lysate were used for immunoprecipitation (Total lysate). C,
Interaction of endogenous VprBP with p53 and HDAC1. Whole cell extracts were
prepared from 293T cells and immunoprecipitated with anti-p53 ( α-DO1) or anti-HDAC1
( α-HDAC1) antibody. The precipitates were analyzed by Western blotting with
antibodies against VprBP, HDAC1 and p53 as indicated. The asterisk in the lane 3
indicates a nonspecific band containing IgG heavy chain.
84
Fig. 3-6. Continued.
85
3-7. Proapoptotic effect of VprBP knockdown in cancer cells
Given the demonstrated action of VprBP as a negative regulator of p53, we next assessed
the expression level of VprBP in three different cancer cell lines expressing wild type
p53; bladder cancer cell line (LD611), prostate cancer cell line (LNCaP) and breast
cancer cell line (MCF7). From our Western blot analysis of these cancer cells, a much
higher level of VprBP expression was evident in comparison to the corresponding normal
cells (Urotsa, MLC and MCF10-2A) (Fig. 3-7A, α-VprBP). Prostate (LNCaP) and breast
(MCF7) cancer cells also showed a higher level of HDAC1, compared to the normal cells,
( α-HDAC1, lanes 4 and 6 versus lanes 3 and 5) arguing for the functional connection
between VprBP and HDAC1 as observed at the p53 target genes. However, we detected
no significant difference in the steady-state level of HDAC1 between bladder cancer
(LD611) and normal (Urotsa) cells ( α-HDAC1, lane 1 versus lane 2). It is tempting to
speculate that overexpressed VprBP acts as a major platform upon which HDAC1 exerts
effects on transcription events in the LD611 cancer cells.
In line with the repressive action of VprBP in p53 transcription activity, a possible effect
of VprBP depletion on expression of p21 and Noxa genes was examined in these cancer
cell lines. Because the shRNA-mediated knockdown of VprBP was not efficient in these
cancer cell lines (data not shown), we employed synthetic siRNA to achieve high-
efficiency knockdown, as confirmed by Western blot analysis (Fig. 3-7B). When these
VprBP-depleted cells were treated with etoposide, the DNA damage-induced expression
86
Fig. 3-7. Stimulation of p53 transcription and apoptosis by VprBP knockdown. A,
Western blot analysis of normal and cancer cells. Exponentially growing normal (Urotsa,
MLC and MCF10-2A) and cancer (LD611, LNCaP and MCF7) cells were subjected to
Western blot analysis using antibodies against VprBP and HDAC1. Actin served as a
control for equal protein loading. B, RNAi-mediated depletion of VprBP. LD611,
LNCaP and MCF7 cells were transfected with a control siRNA (lanes 1, 3 and 5) or a
siRNA directed against VprBP (lanes 2, 4 and 6) for 72 h and analyzed by Western
blotting using the indicated antibodies. All analyses were performed in parallel for each
of the three cell lines. C, Activation of p53 target genes by VprBP depletion. Cells were
first transfected with siRNA targeting VprBP or irrelevant control as in Fig. 3-7B and
treated with or without etoposide (100 µM) for 24 h. Transcription levels of p21 and
Noxa genes were analyzed by quantitative reverse transcription PCR. D, Up-regulation of
DNA damage-induced apoptosis by VprBP depletion. After transfected with either a
control siRNA or a VprBP siRNA, cells were mock-treated or etoposide-stressed as in
Fig. 3-7C. Apoptotic status of cells was analyzed by assessing Annexin V-FITC
fluorescent intensity. The average and standard deviations are shown for three
independent experiments.
87
Fig. 3-7. Continued.
88
of p21 and Noxa genes was significantly stimulated, supporting VprBP acting as a
transcription repressor of p53 (Fig. 3-7C, Eto). Although the p53 target genes exhibited
very weak transcription without etoposide treatment, a moderate enhancement in the
target gene expression was also detectable by VprBP depletion in this uninduced
condition (Fig. 3-7C, -).
Because p53-mediated transcription of p21 and Noxa genes represents the early stage of
apoptosis, we reasoned that the p53-dependent apoptosis that is compromised by VprBP
over-expression in cancer cells may be reactivated by VprBP depletion. To test this, we
depleted VprBP in the cancer cell lines and then subjected them to apoptosis analysis.
Our results showed that VprBP depletion resulted in an apparent increase in the
proportion of apoptotic cells (Fig. 3-7D, Eto), strongly indicating that endogenous VprBP
plays an important role in governing DNA-damage induced apoptosis. The effect of the
VprBP depletion on the induction of apoptosis, albeit of lower magnitude, was also
apparent without DNA damage (Fig. 3-7D, -).
DISCUSSION
Although VprBP was originally implicated in HIV1 replication and pathogenesis based
on its interaction with HIV1 protein Vpr, a recent study detected upon VprBP knockdown
a distinct increase in transcription of p21 gene, which is related to the p53-mediated DNA
damage response pathway (Hrecka et al., 2007). In this study, we explored the
89
hypothesis that VprBP could generate a repressive environment at p53 target genes and
antagonize the apoptotic pathway in response to DNA damage. Taking advantage of our
well defined in vitro transcription system, we demonstrate that VprBP can act as a
negative regulator of p53-mediated chromatin transcription when added prior to p300
HAT. No significant repression in chromatin transcription was observed by the addition
of p300 HAT prior to VprBP or the simultaneous addition of VprBP and p300 HAT to
transcription reactions. It is worth noting that the use of recombinant histones for
chromatin assembly allowed us to exclude the effect of any prior modifications that could
influence the repressive action of VprBP. In striking contrast to chromatin transcription,
the repressive properties of VprBP cannot be recapitulated on naked DNA, pointing to a
mechanism that is dependent upon chromatin architecture. Consistent with this notion,
we found that VprBP preferentially recognizes unmodified state of four major lysine
substrates (K9, K14, K18 and K23) in H3 N-terminal tails protruding from inactive
nucleosomes. Thus, apart from our demonstration of new repressor activity in regulating
p53 function, these results point to the requirement of H3 deacetylation for VprBP-
induced transrepression.
Another important observation from our study is that the central LisH domain of VprBP
plays a dominant role in repressing p53-mediated chromatin transcription. These results
can be simply explained by an intrinsic ability of LisH domain to recognize unmodified
H3 tails. In fact, our interaction studies confirmed the requirement of LisH domain for
the interaction between VprBP and unmodified H3 tails. Notably, our results contrast
90
with those of a recent study showing that LisH domain in transducin beta-like protein 1
and its receptor (TBL1 and TBLR1) binds to the hypoacetylated H4 tail for chromatin
targeting by the nuclear receptor co-repressor complex (Choi et al., 2008). This may
reflect a low degree of sequence homology (only 30%) in their LisH domains, which
could discriminate related but distinct regions of H3 and H4 tails. Thus, whether LisH
domains in different proteins recognize different histone tails is an intriguing question
that needs to be addressed in future studies.
In accord with our in vitro studies, RNAi-complemented ChIP analyses demonstrate that
VprBP is necessary for the maintenance of repressed states of p53 target genes under
normal unstressed conditions. Importantly, the ability of VprBP in establishing this
repressed environment at p53 target promoters relies on its initial recruitment by p53 as
well as its ability to recognize unmodified H3 tails. One important aspect revealed in this
regard is that HDAC1 acts as a gatekeeper to prevent H3 acetylation at p21 and Noxa
promoters, supporting its role in stimulating a stable localization and action of VprBP at
the promoter regions. Because VprBP is required for initial recruitment of HDAC1 to the
promoters, functional significance of the VprBP-HDAC1 interaction should be viewed as
one of the key regulatory processes in establishing a repressed epigenetic environment at
p53 target genes. In further support of the interplay between VprBP and HDAC1 in
constraining p53 transcription, DNA damage-induced activation of p53 target genes was
accompanied by dissociation of VprBP and HDAC1 and concomitant accumulation of
H3 acetylation. At most eukaryotic promoters, the level of histone acetylation undergoes
91
dynamic changes, in which HATs and HDACs act continuously, generating a steady-state
level of histone acetylation (Katan-Khaykovich and Struhl, 2002). This rapid kinetics of
promoter-targeted acetylation and deacetylation processes indicates that the maintenance
of a proper deacetylation state is an important aspect of keeping p53 target genes in an
inactive state under normal cellular condition. In line with this notion, our data establish
the crucial role of VprBP in continuously perturbing H3 acetylation at the promoter
regions to maintain the inactive epigenetic state of p53 response genes. Given the fact
that histone methylation and ubiquitination events are also crucial for potentiating p53-
mediated transcription (An et al., 2004; Kim et al., 2005), VprBP binding to nucleosomes
at p53 responsive genes might also be influenced by these epigenetic modifications.
However, our observation that H3 methylation at K4, K9, K27 or K36 had little or no
effect on VprBP interaction with H3 tails (data not shown) argues against this possibility
and instead indicates a major regulatory role of H3 acetylation for VprBP-H3 tail
interaction.
The functional significance of VprBP in p53 signaling pathway is further underscored by
the finding that etoposide-induced DNA damage leads to the dissociation of VprBP from
promoter nucleosomes to transactivate p53 target genes. That DNA damage-induced
dissociation of VprBP coincides with disappearance of HDAC1 and accumulation of H3
acetylation again points to the interplay of VprBP and HDAC1. Thus, what emerges
from this epigenetic process is the opportunity for H3 acetylation to be precisely
modulated through highly restricted communication between VprBP and HDAC1. More
92
intriguing in this regard is high levels of VprBP and HDAC1 observed in cancer cell lines,
which reinforces our hypothesis that misregulation of VprBP expression maintains
repressive state of p53 target genes and interferes with the apoptotic capability of cells
(Fig. 3-8). In agreement with this hypothesis, VprBP knockdown in these cell lines
results in the apparent stimulation of p53 transcription and apoptotic cell death in
response to etoposide treatment. These data constitute a powerful argument that
overexpression of VprBP in cancer cells antagonizes transcription of p53 target genes,
encoding proapoptotic factors, upon DNA damage. Therefore, further characterization of
physiological and oncogenic activities of VprBP has broad implications for the repertoire
and complexity of the regulatory mechanisms underlying p53 signaling process. Future
work also should be aimed at providing insights into how great a portion of genes are
directly regulated by VprBP and how the specificities of VprBP-mediated repression are
properly determined.
93
Fig. 3-8. Model for the role of VprBP in p53 transrepression. VprBP recognizes
unacetylated H3 at p53 target promoters, and acts as a molecular rheostat to block
transcription initiation in normal condition. For the most efficient repression, VprBP
cooperates with HDAC1 to remove and block H3 acetylation. Upon DNA damage, the
majority of VprBP is dissociated, and HAT establishes active promoter environment for
p53 to achieve the most efficient transcription of proapoptotic genes. By contrast, in
cancer cells that overexpress VprBP, DNA damage-induced activation of p53 target
genes is significantly down-regulated by the high level of endogenous VprBP. See
Discussion for further details.
94
CHAPTER 4: Concluding Remarks
The tumor suppressor p53 determines the cell fate in response to DNA damage by
regulating the expression of multiple target genes that induce cell-cycle arrest, DNA
repair and apoptosis. Failure of this process contributes to cancer development. Previous
studies regarding the regulation of p53 function have mainly focused on p53 stabilization
upon DNA damage and coregulators involved in p53-mediated transcription of target
genes. However, how p53 target genes are properly repressed in unstressed cells is
largely unknown. Recent studies revealed that p53 resides at its target gene promoters in
unstressed cells (Jackson and Pereira-Smith, 2006; Jang et al., 2009), suggesting that
negative regulators play an important role in controling p53 functions, beside MDM2-
mediated degradation. Here, we identified H1.2 complex and VprBP acting as a key
repressor of p53-mediated transactivation.
To investigate the function of H1.2 in gene regulation, we first purified H1.2 interacting
proteins. We found that H1.2 together with its associated factors, but not H1.2 alone,
significantly inhibits p53-dependent, p300-mediated chromatin transcription by blocking
histone acetylation mediated by p300. Further, we revealed that H1.2, YB1 and PUR α
can recapitulate the repressive action of entire H1.2 complex, indicating that YB1 and
PUR α are key factors among H1.2-associated factors. Consistent with transcription data,
ChIP and RNAi analyses showed that YB1, PUR α and H1.2 are colocalized at p53 target
genes and required to maintain repressed states of transcription.
95
Our results raise several questions about the repressive action of H1.2 in p53
transactivation. Is the repressive effect of H1.2 complex on p53-mediated transcription
non-redundant? Is H1.2 complex only involved in regulating p53 target gene expression?
How is this repressive H1.2 complex released from p53 target genes after genotoxic
stress? To address these questions, it is worth trying to purify and characterize other H1
subtype interaction proteins. In our unpublished data, YB1 and PUR α, the key factors in
H1.2 complex, were found to be stably associated with other H1 subtypes, suggesting that
H1.2 can be replaced by other H1 subtypes for the corepressor functions of YB1 and
PUR α in p53 pathway (data not shown). We should also consider the expression levels
and cellular localization of H1 subtypes. Although the expression levels of H1 subtypes
vary depending on cell types, H1.2 and H1.4 are the most abundant in most somatic cells.
A recent study detected distinct subcellular localizations of H1 subtypes; H1.0-H1.3 in
euchromatin regions and H1.4 and H1.5 in heterochromatin regions (Th'ng et al., 2005).
In addition, Sancho et al observed through gene expression array experiments that
different subset of genes were altered in each H1 knock-down (Sancho et al., 2008).
Especially, H1.2 depletion in T47D cells repressed expression of cell cycle-regulatory
genes and caused cell cycle arrest. H1.4 depletion led to cell death, but other H1 subtype
depletions have little effect on cell death and cell cycle arrest. Gathering all these
informations, we can speculate that histone H1 subtypes have distinct roles in the
regulation of p53 function.
96
It also should be noted that because H1.2-interacting proteins exist in multiple complexes,
these subcomplexes may regulate different gene expressions, depending on associated
cofactors. Considering that H1b with Msx1 negatively regulates MyoD gene expression,
H1.2 with some cofactors (Asxl1, CAPER α, nucleolin and WDR5) may positively
regulate specific genes.
Finally, it will be important to figure out how H1.2 with YB1 and PUR α dissociate from
p53. Interestingly, our mass spectrometry analysis identified the presence of DNA-PK
and PP1, which are involved in phosphorylation and dephosphorylation of H1. Recently,
it has been reported that in unstressed cells, Apak with HDAC1 repress p53 functions by
inhibiting p53 acetylation. During DNA damage, Apak is phosphorylated by ATM and
dissociates from p53, leading to p53 activation and apoptosis. Thus we expect that DNA-
PK facilitates H1.2 dissociation from p53 taget promoters via its kinase activity, whereas
PP1 competitively stablizes H1.2 at the promoters by its phosphotase activity.
At the second stage of my Ph.D research, I also observed that VprBP antagonizes p53
function Here we revealed that VprBP is initially recruited by p53 to the p53-responsive
promoters and keep promoters in an inactive state under normal unstressed condition by
interacting with nucleosomes through unacetylated histone H3 tails. In addition,
deacetylation of H3 tails by HDAC1 is necessary for VprBP to occupy promoter region.
The repressive actions of VprBP and HDAC1 at p53 target genes are further confirmed
97
by the finding that knockdown of VprBP and HDAC1 stimulates p53 target gene
expression in response to DNA damage and increases apoptosis.
Given that VprBP and HDAC1 dissociate from the promoter regions upon DNA damage,
it will be interesting to find the mechanism how VprBP can be released from the
nucleosomes. As is the case for linker histone and p53, we assume that PI 3-kinase family
such as DNA-PK, ATM or ATR is involved in the dissociation of VprBP from the
nucelsome. Indeed, our pilot experiments revealed that VprBP is phosphorylated by
DNA-PK (data not shown). Thus, the phosphorylation of VprBP may destabilize the
interaction between VprBP and H3 tail region through structural change, resulting in the
release of VprBP from the chromatin.
Having found that VprBP is overexpressed in several human cancer cells and the
exaggerated increase of VprBP renders the cells more resistant to apoptotic stress, further
investigation regarding the involvement of VprBP in p53 pathway and cancer
development will allow us to understand the mechanism of tumorigenesis.
In conclusion, our findings of H1.2 complex and VprBP as negative regulators of p53
function provided a better understanding of p53 regulation. Moreover, our studies yielded
important insights on a possible approach targeted to H1.2 complex and VprBP as a new
therapeutic strategy for cancer treatment.
98
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Abstract (if available)
Abstract
p53 is a transcription factor which regulates the cell cycle, DNA repair and apoptosis in response to DNA damage. Even though the mechanisms of p53 accumulation and activation have been extensively studied, little is known about how p53 function is negatively regulated. Here we described H1.2 complex and VprBP as novel negative regulators of p53-mediated transcription.
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Asset Metadata
Creator
Kim, Kyunghwan
(author)
Core Title
Identification of two negative regulators of p53: H1.2 complex and VprBP
School
Keck School of Medicine
Degree
Doctor of Philosophy
Degree Program
Biochemistry and Molecular Biology
Degree Conferral Date
2010-05
Publication Date
04/28/2010
Defense Date
03/24/2010
Publisher
University of Southern California
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Tag
acetylation,cancer,chromatin,histone,linker histone,OAI-PMH Harvest,p53,transcription,VprBP
Language
English
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An, Woojin (
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), Stallcup, Michael R. (
committee member
), Wang, Clay C. C. (
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)
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keny0917@hanmail.net,Kyunghki@usc.edu
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Kim, Kyunghwan
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University of Southern California Dissertations and Theses
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Tags
acetylation
cancer
chromatin
histone
linker histone
p53
transcription
VprBP