UNIVERSITY OF CINCINNATI
Date:______
I, ______, hereby submit this work as part of the requirements for the degree of: in:
It is entitled:
This work and its defense approved by:
Chair: ______
RB-mediated Regulation of Transcription and Epigenetic Modifications
A dissertation submitted to the
Division of Research and Advanced Studies of the University of Cincinnati
in partial fulfillment of the requirements for the degree of
DOCTORATE OF PHILOSOPHY (Ph.D.)
In the department of Cell Biology, Neurobiology, and Anatomy
of the College of Medicine
2006
By
Hasan Siddiqui
B.S., University of South Alabama, 2000
Committee Chairman: Erik Knudsen, Ph.D
ABSTRACT
The retinoblastoma tumor suppressor protein (RB) is the ‘master regulator’ of
cellular proliferation and is targeted for inactivation in the majority of human
tumors. RB inhibits proliferation by repressing the transcription of genes essential
for cell cycle progression. Although RB interacts with numerous chromatin
modifying (e.g., histone deacetylases or HDACs, histone methyltransferases or
HMTases) and remodeling (e.g., SWI/SNF chromatin remodeling complex) enzymes, the functional significance of these interactions is not apparent. Here we investigated the role of chromatin modifying and remodeling enzymes in RB- mediated transcriptional repression and epigenetic modifications. We find that active RB mediates histone deacetylation on endogenous RB target gene promoters. We also demonstrate that this deacetylation is HDAC dependent, since the HDAC inhibitor trichostatin A (TSA) prevented histone deacetylation at each promoter. However, TSA treatment blocked RB repression of only a specific subset of genes, demonstrating that the requirement of HDACs for RB-mediated transcriptional repression is promoter specific. Furthermore, we find that cell cycle inhibitory action of RB is not intrinsically dependent on the ability to recruit
HDAC activity. The HDAC-independent repression was not associated with DNA methylation or gene silencing since it was readily reversible. We show that this form of repression resulted in altered chromatin structure and was dependent on
SWI/SNF chromatin remodeling activity. Importantly, we demonstrate that
SWI/SNF is required for histone deacetylation of RB target promoters.
Furthermore, we show that loss of RB disrupts a specific epigenetic modification
and heterochromatin protein 1 (HP1) dynamics in the chromatin. These studies highlight the influence of RB on transcriptional repression and epigenetic modifications.
DEDICATION
In Loving Memory of My Father
S.B. Siddiqui, PhD
(1942-2004)
Acknowledgments
I am greatly indebted to my advisor Dr. Erik Knudsen for his guidance and
support throughout my graduate career. It was an exciting and memorable experience to have the opportunity to work under his supervision. Needless to say, I will always cherish my time in his laboratory. I am grateful to members of my thesis committee: Drs Robert Brackenbury, Sohaib Khan, Karen Knudsen and Bernard Weissman for their sincere suggestions, advice and help in both academic and personal matters. I want to thank all members of the Knudsens’ laboratories, past and present, for their help and suggestions during the course of my study. This work would not have been possible without the unconditional assistance from Ranjaka Gunawardena, Sejal Ranmal-Fox and David Solomon. I truly appreciate their help. I would also like to thank Emily Bosco, a great lab- colleague, who sacrificed valuable time on proof reading my manuscripts.
I wish to express my heartfelt appreciations to my friends Moinul Islam,
Andromeda Nauli, Saydur Rahman and Kazi Sayeed for their friendship, support, and humorous discussions during my stay in Cincinnati.
I am immensely grateful to my wonderful parents S.B. Siddiqui and Firoza
Begum for their endless love, support and encouragements, without which I would not have come this far. I would also like to thank my brother Raihan for his support. Finally, I wish to acknowledge my wife Rumana for her patience, love, support, understanding and companionship. For all that, I am truly grateful to her.
TABLE OF CONTENTS
Page
Table of Contents………………………………………………………..1
List of Tables and Figures……………………………………………...3
Chapter I: Introduction……………………………………………….....5
A. Cancer and Cell Cycle…………………………………………5 B. Retinoblastoma Tumor Suppressor Protein (RB)………...8 C. Growth Suppression by RB…………………………………10 D. The E2F Family of Transcription Factors: Cellular target of RB……………………………………………………11 E. Chromatin Modifying and Remodeling Enzymes………..14 F. RB and Chromatin…………………………………………….18 G. Reference List…………………………………………………..21
Chapter II: Histone Deacetylation of RB-responsive Promoters: Requisite for Specific Gene Repression but Dispensible for Cell Cycle Arrest…………………………………………………………………....32
A. Abstract………………………………………………………...32 B. Introduction……………………………………………………34 C. Materials and Methods……………………………………....37 D. Results………………………………………………………….43 E. Discussion……………………………………………………..53 F. Reference List………………………………………………....59
Chapter III: Hierarchical Requirement of SWI/SNF in RB-mediated Repression of Plk1……………………………………………………..89
A. Abstract……………………………………………………...... 89 B. Introduction……………………………………………………91
1 C. Materials and Methods…………………………………...... 94 D. Results………………………………………………………….96 E. Discussion………………………………………………….…105 F. Reference List…………………………………………….…..109
Chapter IV: Loss of RB Compromises Specific Heterochromatin Modifications and Modulates HP1α Dynamics………………………..132
A. Abstract……………………………………………………...132 B. Introduction……………………………………………...….134 C. Materials and Methods………………………………...... 137 D. Results………………………………………………...……..140 E. Discussion…………………………………………………...145 F. Reference List……………………………………………....150
Chapter V: Summary and Conclusions………………………………....167
A. HDAC-dependent gene repression by RB……………168 B. Requirement of HDAC activity for RB-mediated cell cycle arrest…………………………………………….171 C. HDAC-independent gene repression by RB………....172 D. Cross talk between SWI/SNF activity and histone deacetylation…………………………………..…174 E. Regulation of Epigenetic Modification by RB………..175 F. Reference List……………………………………………...178
2 List of Tables and Figures
Figure Page
I-1 Functional inactivation of the RB pathway of cell cycle regulation………..7
I-2 Models of RB-mediated inhibition of E2F-regulated gene transcription....13
II-1 PSM-RB (active RB) represses a set of cell cycle genes and induces cell
cycle arrest (A-B)………………………………………………………………72
II-1 PSM-RB (active RB) represses a set of cell cycle genes and induces cell
cycle arrest (C-E)………………………………………………………………73
II-2 Active RB induces histone deacetylation at promoters of specific cell cycle
genes……………………………………………………………………………75
II-3 RB induced deacetylation at specific promoters is mediated through
histone deacetylases (HDACs)………………………………………………77
II-4 HDAC activity is dispensable for RB-mediated cell cycle arrest………….79
II-5 HDAC-dependent deacetylation is required by RB to repress specific
promoters (A-C)………………………………………………………………..81
II-5 HDAC-dependent deacetylation is required by RB to repress specific
promoters (D)…………………………………………………………………..83
II-6 The cyclin A promoter is not subjected to stable gene silencing (A-B)….84
II-6 The cyclin A promoter is not subjected to stable gene silencing (C)……86
II-7 RB-mediated repression of the cyclin A promoter involves chromatin
remodeling (A)………………………………………………………………….87
II-7 RB-mediated repression of the cyclin A promoter involves chromatin
remodeling (B)………………………………………………………………….88
3 III-1 RB pathway represses Plk1 expression (A-C)……………………………119
III-1 RB pathway represses Plk1 expression (D-E)……………………………120
III-2 SWI/SNF is required for RB-mediated repression of Plk1 (A-B)………..121
III-2 SWI/SNF is required for RB-mediated repression of Plk1 (C-E)………..122
III-3 SWI/SNF is dispensable for E2F chromatin/promoter association(A-B).124
III-3 SWI/SNF is dispensable for E2F chromatin/promoter association(C-E).125
III-4 SWI/SNF is not required for pocket protein association with the Plk1
promoter ……………………………………………………………………...127
III-5 SWI/SNF is required for histone deacetylase-mediated repression of the
Plk1 promoter (A)…………………………………………………………….128
III-5 SWI/SNF is required for histone deacetylase-mediated repression of the
Plk1 promoter (B-D)………………………………………………………….129
III-6 Schematic diagram of transcriptional regulation of Plk1…………………131
IV-1 Conditional loss of RB leads to target gene deregulation……………….159
IV-2 Acute RB loss results in specific loss of heterochromatin associated
histone modifications (A-C)…………………………………………………160
IV-2 Acute RB loss results in specific loss of heterochromatin associated
histone modifications (D-F)…………………………………………………161
IV-3 Acute RB loss does not relocate euchromatin markers into
heterochromatin domains…………………………………………………..162
IV-4 Acute RB loss modulates HP1α dynamics (A)…………………………...164
IV-4 Acute RB loss modulates HP1α dynamics (B)…………………………...165
IV-4 Acute RB loss modulates HP1α dynamics (C)…………………………...166
4 Chapter I:
Introduction
Cancer and Cell Cycle
Cancer refers to approximately 100 different diseases that arise in various tissues of the human body through distinct mechanisms. Each cancer has unique features yet the basic processes that lead to tumor formation appear to be similar. Tumor cells acquire the ability to proliferate without growth signals, invade surrounding tissues, migrate to distal tissues (metastasis), bypass programmed cell death (apoptosis) and promote blood vessel formation
(angiogenesis) (24). In normal healthy individuals, cells undergo division only when surrounding cells emit signals. This ensures that the daughter cells have sufficient nutrients available for its survival. However, tumor cells divide without any external growth promoting signals. As tumor cells continue to divide, bypassing growth inhibitory signals, they invade nearby tissues creating havoc in normal healthy tissues. It is therefore essential that proliferation is carefully regulated for homeostasis.
In normal cells, cell division is a carefully choreographed process controlled by a host of cellular proteins. This process, called the cell cycle, is composed of four stages. In G1 (gap 1), the cell prepares to copy its DNA. The actual replication of DNA is accomplished in the next stage, called S (synthesis) phase, tightly controlled by various proteins such that every base pair in the host DNA is faithfully copied in the daughter DNA. Upon successful completion of S phase,
5 cells go through G2 (gap 2) essentially preparing for the next phase. In M
(mitosis) phase, the cell finally divides its DNA content into two daughter cells ensuring that each new progeny is endowed with the exact set of chromosomes as the parental cell. The progeny cells immediately enter G1 and may go through the cell cycle again or may enter G0 (quiescence) until appropriate signals stimulate them to enter the cell cycle again. The coordinated efforts of various cell cycle regulatory proteins ensure that cell division occurs only when the physiological environment has the resources to sustain new cells. Although the process is carefully regulated, it sometimes goes awry, leading to malignant transformation, as happens during tumorigenesis. Thus, cell machinery has devised pathways to inhibit proliferation when the physiological environment is unfavorable for cell growth. One critical pathway for growth inhibition centers on the retinoblastoma tumor suppressor protein (RB).
The RB pathway of cell cycle regulation has been termed the ‘master regulator’ of cell cycle progression (28). This term aptly describes the role of this pathway in regulating cell cycle, as virtually all growth inhibitory signals converge on components of this pathway to efficiently halt cell proliferation. Thus for tumor progression, inactivation of this pathway is central to the tumor cell’s ability to proliferate without a license. In fact, virtually all forms of cancer have defects in the RB pathway, emphasizing its critical role for tumor inhibition.
6 Amplification Overexpression
Cyclin D
p16ink4a + RB E2F
CDK4/6
Deletion Deletion Point Mutation Mutation Point Mutation Methylation Disruption of p16 binding Viral Oncoprotein Binding
Figure I-1: Functional inactivation of the RB pathway of cell cycle
regulation. Upstream positive (p16ink4a) and negative (Cyclin
D/CDK4, 6) regulators of RB are found to be mutated in many forms
of cancer. Mutations in p16ink4a can inactivate it leading to
deregulated cyclin D/CDK-6 activity. Mutations can also affect
cyclin D and CDK4,6 resulting in increased activity of this complex.
Either of these two processes is sufficient to inactivate RB by
hyperphosphorylation. RB itself can be inactivated by deletion,
point mutation and binding to viral oncoproteins (e.g. E7 of HPV16).
Adapted from Mulligan and Jacks, 1998.
Functional inactivation of RB can occur through several disparate mechanisms including biallelic deletion of the Rb gene, inactivation by viral oncoproteins such as SV40 large T antigen, adenoviral E1A and human papillomavirus E7 or mutation of upstream regulators (p16ink4a and Cyclin
7 D/CDK4) (2, 3, 35, 54, 57, 67). Although the mode of inactivation of the RB
pathway differs in different cancers, the end result is loss of restraint on cell cycle
machinery, which allows cell division to occur without any barrier. The
importance of the RB pathway in growth suppression is underscored by the
observation that approximately 90% of human tumors show defects in the RB
pathway.
Retinoblastoma Tumor Suppressor Protein (RB)
Rb was the first tumor-suppressor gene to be identified and was
subsequently mapped to the long arm of chromosome 13 (13q14). The human
Rb gene consists of 27 exons encompassing 180 kb of genomic DNA. It is
transcribed into a 4.8 kb mRNA with no convincing evidence of alternate splice
forms. The open reading frame of the Rb gene is approximately 2.7 kb within 4.8 kb Rb mRNA. The protein product of the Rb gene is 928 residues long (921 in mouse) and has a molecular weight of 110 KDa. The promoter region of the Rb gene contains binding motifs for SP1 and ATF, but does not contain any TATA or
CAAT elements. The 5’ end of the Rb gene contains CpG islands which are normally unmethylated but are observed to be hypermethylated in certain tumors.
RB is a ubiquitously expressed nuclear phosphoprotein. Biochemical analyses have revealed four discrete structural domains of RB (26). Three of these domains, the N terminus, R motif and A/B pocket, are resistant to partial proteolysis while the fourth domain, the C-terminus, is susceptible to proteolysis.
Although the N and C termini of RB are structurally poorly defined, they are
8 important for growth suppression. The C terminal region of RB contains a nuclear
localization signal and cyclin binding motifs ([R/K]xL) that are important for its
interaction with D type cyclins. It interacts with the A/B pocket domain and this
interaction is strengthened by phosphorylation of C terminal phosphorylation
residues. In addition, the C terminus also binds the oncoproteins c-Abl and
MDM2 (70). Deletion of exons 24 and 25 compromises many C terminal
functions and results in low penetrance retinoblastoma, underscoring the
importance of the C terminus for biological activity of RB (7). The N terminus also binds several proteins and appears to be important for RB function, since deletion mutants have been identified in human tumors (15, 30). The A/B pocket domain is the most well defined domain of the RB protein. Structural analyses using X-ray crystallography has shown that these two domains essentially form a protein binding pocket (38). Initially this pocket domain was identified through its ability to bind viral oncoproteins E1A and SV40 large T antigen (17).
Subsequently, several studies revealed that this region binds many cellular proteins including the E2F family of transcription factors (29, 69). Most naturally occurring mutations of RB disrupt the integrity of the A/B pocket, highlighting its
importance for RB function (23, 74). Genetic and biochemical analysis have
shown that the A/B pocket domain of RB is necessary but not sufficient for the
growth suppressive property of RB (29, 50). These studies reveal that A/B pocket
as well as the C terminal amino acid sequences are required for cell cycle
inhibition by RB. Homology analysis of the pocket domain revealed that two other
cellular proteins, p107 and p130, have similar domains. Collectively, these
9 proteins (RB, p107 and p130) are termed pocket proteins. Although they share
domain similarity and have similar properties in cell culture models, only RB is
observed to be mutated in human cancer emphasizing its unique role as a tumor
suppressor.
Growth Suppression by RB
RB is a multifunctional protein and has been implicated in a wide range of
cellular processes, including regulation of cell cycle, differentiation, apoptosis,
DNA replication, DNA damage checkpoint response. However, most functional
studies of RB have focused on its ability to inhibit proliferation. Overexpression studies have demonstrated that RB can arrest cells in G1 and cells deficient in
RB undergo accelerated G1 transition (22, 27, 32, 50). Together, these studies revealed that RB is a potent negative regulator of cellular proliferation. It was also demonstrated that during cell cycle arrest by RB, critical cell cycle regulators such as dihydrofolate reductase (DHFR) and cyclin E were transcriptionally repressed. Since RB lacks DNA binding motifs, questions were raised as to how it may influence transcription. It was postulated that RB mediates its biological action through various other proteins.
RB interacts with a myriad of cellular proteins. The repertoire of RB binding proteins includes transcription factors, chromatin modifying and remodeling enzymes, and viral oncoproteins. Early studies demonstrated that proteins from DNA tumor viruses, including adenoviral E1A, SV40 large T
antigen and E7 of human papilloma virus type 16 (HPV16) can bind RB and drive
10 quiescent cells into S phase (11, 18, 47, 71). Subsequent studies aimed at
identifying cellular substrates of RB led to the discovery of E2 factor (E2F). It is
now known that the E2F family of transcription factors is the critical target of the
RB family of pocket proteins.
The E2F Family of Transcription Factors: Cellular Target of RB
E2F was first identified as cellular factors that are required for the activation of the E2 viral promoter. In vivo, E2Fs exist as heterodimers with differentiation regulated transcription factor (DRTF) proteins (DP). To date, eight
E2Fs (E2F1-E2F8) and three DPs (DP1-DP3) have been discovered in mammalian cells although fewer members have been identified in lower eukaryotes (13). The classical E2Fs (E2F1 through E2F5) can be divided into two subgroups based on their interaction with pocket proteins and their role in transcription. Activating E2Fs consist of E2F1, E2F2 and E2F3a and primarily interact with RB. E2F4 and E2F5 are repressive E2Fs and generally interact with p107 and p130 respectively. Recently identified E2F family members (E2F3b,
E2F6, E2F7 and E2F8) are less well characterized and likely function as repressors of E2F-dependent transcription (13). Among the classical E2Fs, activating E2Fs drive cell proliferation whereas repressor E2Fs are likely required for cell cycle exit and differentiation (66). To regulate proliferation, E2Fs bind to the promoter region of target genes via the consensus 5’-TTTCGCGC-3’ sequence to initiate or repress transcription.
11 E2F regulated genes are essential for cell cycle progression because they
include cell cycle regulators (e.g., cyclin E, cyclin A, cdc2, cdk2), de novo
nucleotide synthesis enzymes (dihydrofolate reductase, thymidylate synthase, thymidine kinase) and replication proteins (PCNA, DNA polα and δ, RPA1,2,3,
CDC6, MCM2,3,4,5,6,7, ORC1) . Although the number of E2F regulated genes is steadily increasing, it has become clear that proliferation control is central to the role of E2Fs. Overexpression of E2F1, E2F2 and E2F3 drives quiescent cells to re-enter the cell cycle (34, 40, 51). This is dependent on DNA binding and transcriptional activity of the proteins (34). The activating E2Fs can override various anti-proliferative signals including TGF-β and Cdk inhibitors (p16, p21 and p27) (12, 43, 56). Conversely, inhibition of E2F activity by dominant-
negative mutants of E2F1, DP1 and DP2 or by competitive RNA molecules
increases cells in G1 phase, thereby reducing portions of S phase cells (14, 19,
33, 72). The critical role of activating E2Fs in driving S-phase progression has
been validated by examining endogenous E2Fs through a variety of different
methods. For instance, selective inactivation of E2F3 by microinjection of anti-
E2F3 antibody dramatically arrests proliferating cells (39). Combined inactivation
of activating E2Fs results in elevated p21Cip1 protein with concomitant decrease
in CDK activity and RB phosphorylation (73). Collectively, these findings
demonstrate that activating E2Fs are critical for cellular proliferation. Thus,
whereas activating E2Fs promote proliferation, RB acts to inhibit it suggesting
that the two classes of proteins antagonize each other’s activity.
12 A RB
E2F DP
B Co-repressors RB
E2F DP
Figure I-2: Models of RB-mediated inhibition of E2F-regulated
gene transcription. A) RB binds to transactivation domain of E2F
and blocks its activity (inhibition of activation). B) RB recruits
additional corepressors and actively modifies/remodels the
chromatin to repress transcription (active repression)
Genetic and biochemical studies have revealed that RB opposes activating
E2Fs (25, 59). Furthermore, microarray analysis shows that RB efficiently represses a large number of E2F-regulated genes that are required for cell cycle progression (44). However, the mechanism underlying this repression is poorly understood. Two models have been proposed to describe how RB might inhibit
E2F-mediated transcription (Fig I-2). First, RB binds to activating E2Fs and blocks their transactivation (20, 25). Second, RB recruits multiprotein complexes
13 to E2F-regulated gene promoters that actively repress transcription (6, 58, 68).
This active repression presumably involves altering DNA topology such that transcription factors are unable to bind to their target sequences. This would require that RB assembles chromatin modifying and remodeling enzymes to
E2F-regulated gene promoters. Indeed, RB has been shown to interact with enzymes that are involved in covalent histone modification and ATP-dependent chromatin remodeling.
In addition to E2Fs, RB interacts with multiple proteins that have known corepressor function. Often these interactions occur between and LXCXE motif in the corepressor molecule and the A/B pocket domain of RB. For instance, RB interacts with histone deacetylases (HDACs) (5, 10, 36, 41, 42), histone methyltranferases (HMTases) (49) and components of the ATP-dependent chromatin remodeling enzymes SWI/SNF (BRG1 and BRM) (16, 62), which all contain LXCXE motifs. These corepressors are believed to play important roles
in RB-mediated transcriptional repression and cell cycle arrest. Consistent with
this hypothesis, combined loss of BRG1 and BRM, the core ATPase of the
SWI/SNF chromatin remodeling complex, compromises RB-mediated
transcriptional repression and cell cycle arrest (64). However, the role of HDACs
and HMTases for RB function is not clear.
Chromatin Modifying and Remodeling Enzymes:
In eukaryotes, the nucleosome serves as a barrier to transcription that
must be overcome to initiate gene transcription. Transcriptional activators recruit
14 histone acetyltransferases (HATs), which add acetyl groups to the lysine tails of
core histones. Addition of the acetyl group neutralizes the positive charge on the histone tails, weakening the histone-DNA interaction, resulting in loosening of the chromatin structure. This disruption is believed to unfold the DNA thereby allowing transcription factors and basal transcriptional machinery to gain access to the promoter regions of genes to be activated. Conversely, transcriptional repressors utilize the activity of histone deacetylases to inhibit transcription.
HDACs remove the acetyl groups from the lysine residues of histone tails, which results in a positively charged histone tail that enhances the histone-DNA interaction. This enhanced histone-DNA interaction is believed to facilitate nucleosome formation and results in a compact chromatin structure that is inaccessible to the transcriptional machinery. Thus transcriptional repressors recruit HDACs to form a barrier between transcriptional activators and their cognate DNA sequences.
HDACs belong to a large family consisting of >10 different enzymes, which are divided into three classes. Of these, class I enzymes, which include
HDAC1, HDAC2 and HDAC3, have been shown to interact with RB (5, 9, 10, 36,
37, 41, 42). Initial studies have shown that RB physically interacts with HDAC1 and could mediate histone deacetylation at a synthetic promoter. Subsequent studies revealed that cell cycle dependent histone acetylation changes occurred in several endogenous RB/E2F-regulated promoters (46, 52). However, a direct demonstration of the role of HDACs during RB-mediated transcriptional repression and cell cycle arrest has been lacking.
15 Histone methylation as a mechanism of transcriptional regulation has
gained increased attention in the past few years. Unlike acetylation of histones
which decreases the positive charge, methylation does not alter the overall
charge of histone tails. Histone lysines can be mono, di and trimethylated and
each additional methyl group enhances basicity and hydrophobicity of the
histones. Moreover, the affinity of histones for DNA increases as methyl groups
are added (4, 8). Thus, methylation of core histones likely alters histone-DNA
interactions thereby influencing nucleosome structure and function. Methylation of histones is accomplished by histone methyltransferases (HMTase) using the cofactor S-adenosylmethionine (SAM). Early studies suggested that histone methylation is relatively stable, aid in establishment specific genomic domains.
However, the recent discovery of lysine specific demethylases (LSDs) suggests
that histone methylation is likely to be a dynamic process (45, 60).
The eukaryotic genome can be divided into transcriptionally active
(euchromatin) and inactive (heterochromatin) domains. Thus, euchromatin
consists of gene-rich regions of the chromatin whereas heterochromatin is
generally gene-poor, containing large blocks of repetitive DNA elements.
Heterochromatin assembly requires methylation of specific lysines on histone
tails and heterochromatin protein 1 (HP1) binding to the chromatin. A number of
HMTases have been discovered and shown to contain specificity for histone
substrate. For instance, Suv39h1/h2 specifically trimethylates histone H3 lysine
9, whereas Suv4-20h1/h2 preferentially methylates histone H4 lysine 20 (55).
Primary mouse embryonic fibroblasts from Suv39h1/h2 mice show enhanced
16 mitotic defects and poorly characterized heterochromatin domains (53).
Furthermore, overexpression of SUV39H1 can induce ectopic heterochromatin
formation (53). Collectively, these findings suggest a critical role for HMTases for
heterochromatin assembly and maintenance.
Unlike histone modification enzymes which covalently modify the histones, the SWI2/SNF2 superfamily of chromatin remodeling enzymes alter chromatin structure through a process that requires energy in the form of ATP. The chromatin remodeling superfamily is classified into three subfamilies each containing multiprotein complexes: 1) the SWI2-SNF2 subfamily, 2) the ISWI subfamily and 3) CHD subfamily. Although chromatin remodeling complexes are
composed of multiple subunits, the ATPase subunit is critical for its function. In
the SWI/SNF complex, which is a member of SWI/SNF2 subfamily of chromatin
remodelers, the core subunit is either Brahma (BRM) or Brahma related gene 1
(BRG1). The SWI/SNF complex consists of at least 11 subunits: the core
ATPase (BRG1 or BRM) and several BRG1-associated factors (BAFs). The
complex binds both DNA and nucleosomes with high affinity and this interaction
is believed to occur at the minor groove of the DNA helix. Initially SWI/SNF was
believed to promote transcriptional activation since it was shown to increase
accessibility of transcriptional activators and restriction enzymes to the
nucleosomal DNA. However in microarray studies, many genes were
upregulated when SWI/SNF was inactivated indicating a role for SWI/SNF in
transcriptional repression (31, 65). Moreover, BRG1 and BRM have been
isolated from complexes that also contain repressor molecules including Sin3,
17 HDAC1 and HDAC2 (61). Collectively, these studies support a role for SWI/SNF in both transcriptional activation and repression.
RB and Chromatin
It is well established that RB is a potent growth inhibitor and transcriptional repressor. However, the mechanism of its transcriptional repressor function has not been investigated in depth. Although numerous histone modifying and chromatin remodeling enzymes have been shown to interact with RB, the functional significance of most of these interactions remain unexplored. The first chromatin remodeling enzyme shown to interact with RB was the BRG1 subunit of the SWI/SNF chromatin remodeling complex. Additional studies have shown that cell lines deficient in BRG1 or BRM are resistant to RB mediated cell cycle arrest, suggesting that SWI/SNF is a requisite for the biological action of RB (63,
75). However, the mechanism underlying this phenomenon has not been investigated.
Several independent groups have shown that class I HDACs (HDAC1,
HDAC2 and HDAC3) interact with RB (5, 41, 42). Furthermore, promoter activity of a synthetic reporter construct was shown to be repressed by RB in a HDAC dependent manner (41). However, since a synthetic promoter may not behave in a manner analogous to an endogenous promoter, these studies do not necessarily reveal the true requirement of HDACs for RB-mediated transcriptional repression. Furthermore, as HDACs modify histones on the chromatin, the use of a plasmid based reporter construct to analyze histone
18 acetylation/deacetylation may not accurately reflect the biological functions of these enzymes. Thus, a more direct approach using endogenous RB target genes and their promoters in the context of the chromatin will help delineate the physiological requirement of HDACs for RB-mediated transcriptional repression and cell cycle arrest.
RB also interacts with several histone methyltransferases (21, 49). Nielsen et. al. first demonstrated that RB physically interacts with SUV39H1 and that
SUV39H1 directed methylation of H3K9 was critical for RB-mediated transcriptional repression of the cyclin E gene (49). However, subsequent studies did not reveal any other RB-regulated genes that required SUV39H1 activity for repression in cells progressing through the cell cycle. Thus, it appears that RB- mediated repression of target genes in cycling cells may not require H3K9 methylation by SUV39H1 although this activity has been shown to be indispensable for repression of some RB target genes in senescent and differentiated cells (1, 48). The potential involvement of the recently identified
HMTase Suv4-20h1 and Suv4-20h2 for RB-mediated transcriptional repression remains unknown (21, 55). However, since both the SUV39H and Suv4-20
HMTases are involved in heterochromatin assembly and maintenance pathway, it is possible that RB plays a role in regulating the chromatin structure by directing these enzymes to sites of heterochromatin assembly and regulating their activity.
19 Here we investigated the role of RB in modulating transcription and chromatin structure. Specifically, we investigated i) the role of HDACs in RB- mediated transcriptional repression and cell cycle arrest, ii) the interplay between SWI/SNF and histone deacetylation for RB-mediated transcriptional repression and iii) the role of RB in regulating chromatin structure. Our findings reveal that RB-mediated repression of target genes occur in concert with histone deacetylation of target gene promoters. However, this deacetylation is not required for RB-mediated cell cycle arrest since blockade of HDAC activity did not rescue cells from RB-induced cell cycle arrest. Our findings also demonstrate that RB utilizes HDAC activity to repress only a subset of target genes. Critical
RB targets, including cyclin A, which is required for S-phase progression, were
HDAC-independent but SWI/SNF dependent for repression. We also demonstrate that histone deacetylation of RB-responsive genes was dependent on SWI/SNF activity. Finally, we also demonstrate a more global role of RB in regulating chromatin structure by influencing components of the heterochromatin assembly pathway.
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31 Chapter II:
Histone Deacetylation of RB-responsive Promoters: Requisite for Specific Gene Repression but Dispensable for Cell Cycle Arrest
ABSTRACT
The retinoblastoma tumor suppressor protein (RB) is targeted for inactivation in the majority of human tumors, underscoring its critical role in attenuating cellular proliferation. RB inhibits proliferation by repressing the transcription of genes essential for cell cycle progression. To repress transcription, RB assembles multiprotein complexes containing chromatin modifying enzymes including histone deacetylases (HDACs). However, the extent to which HDACs participate in transcriptional repression and are required for RB-mediated repression has not been established. Here we investigated the role of HDACs in RB dependent cell cycle inhibition and transcriptional repression. We find that active RB mediates histone deacetylation on cyclin A, cdc2, topoisomerase IIα, and thymidylate synthase promoters. We also demonstrate that this deacetylation is HDAC dependent, since the HDAC inhibitor trichostatin A (TSA) prevented histone deacetylation at each promoter.
However, TSA treatment blocked RB repression of only a specific subset of genes, demonstrating that the requirement of HDACs for RB-mediated
32 transcriptional repression is promoter specific. The HDAC-independent repression was not associated with DNA methylation or gene silencing, but was readily reversible. We show that this form of repression resulted in altered chromatin structure and was dependent on SWI/SNF chromatin remodeling activity. Importantly, we find that cell cycle inhibitory action of RB is not intrinsically dependent on the ability to recruit HDAC activity. Thus, while HDACs do play a major role in RB-mediated repression, they are dispensable for repression of critical targets leading to cell cycle arrest.
33 INTRODUCTION
The retinoblastoma tumor suppressor, RB, functions as a negative
regulator of cell cycle progression that is frequently inactivated in human cancers
(10,22,75). In G0 and early G1 cells, RB is hypophosphorylated and inhibits the
transition into S-phase of the cell cycle. Mitogenic signaling cascades activate
CDK4/cyclin D1 complexes that initiate the phosphorylation of RB on a sub-set of serine and threonine residues (65). Subsequent phosphorylation catalyzed by
CDK2/cyclin E leads to RB hyperphosphorylation (23). These combined events serve to functionally inactivate RB and thereby facilitate progression through S- phase (2,23). In contrast with mitogenic signaling pathways, anti-mitogens (e.g.
TGF-β or DNA-damage) serve to inhibit RB phosphorylation and prevent
progression through the cell cycle (28). Thus, RB integrates multiple signaling cascades to modify proliferation. In cancer, RB is inactivated through several
disparate mechanisms. These modes of inactivation include the bialleleic
inactivation of the RB gene, binding by oncoproteins of DNA tumor viruses, and
aberrant phosphorylation (2,3,30,59,62,76). Through these distinct mechanisms
of RB inactivation, tumors evade cell cycle regulation and proliferate
uncontrollably.
RB functions to inhibit cellular proliferation by assembling complexes
involved in transcriptional repression. Biochemical analyses have shown that RB
interacts with a plethora (>100) of different cellular proteins (46). The significance
of most of these interactions remains elusive. However, the E2F family of
transcription factors represent critical targets of RB (8,16,49). E2F complexes
34 exist in vivo as heterodimers composed of subunits from E2F (E2F1-6) and DP
(DP1 and DP2) gene families. E2F-DP heterodimers bind to specific DNA sequences and function as transcriptional activators. E2F responsive genes include cell cycle regulators, such as cyclin E, cyclin A, cdc2, and cdk2
(5,12,18,20,29,54,61,73) and factors important for DNA synthesis, including DNA
polymerase α, thymidine kinase, and dihydrofolate reductase (31,53,57,69).
Recently, E2F proteins have been shown to directly interact with the promoters of
many of these genes (72,78).
Genetic and biochemical analyses have shown RB to functionally
antagonize E2F activity (27,64). Additionally, we have recently shown that RB
potently represses a significant number of E2F-regulated genes that are requisite
for cell cycle progression (42). Currently, there are two models describing how
RB impinges upon E2F directed transcription: 1) RB binds to the E2F family of
transcription factors, thus blocking their transactivation capacity (17,27); and 2)
RB assembles large multi-protein complexes at E2F-regulated promoters that
actively repress transcription (7,63,77). A number of functional studies
demonstrate that E2F-dependent repression is required for RB to inhibit
proliferation. For example, E2F alleles which displace E2F/RB complexes from
DNA abrogate RB-dependent cell cycle control (82).
To facilitate transcriptional repression, RB not only interacts with E2F, but
also multiple co-repressor molecules. Many of these interactions are mediated by
LXCXE-motifs present in the co-repressor interacting with the A/B pocket domain
of RB (6,15,68). For example, RB interacts with histone deacetylases (HDAC)
35 (6,11,36,39,40), ATP-dependent chromatin modifying enzymes BRG-1/BRM
(15,68), and histone methyltransferases (52) which contain LXCXE-motifs. It is believed that these enzymes play critical roles in RB-mediated transcriptional repression. Such an idea is supported by the observation that the combined loss of BRG-1 and BRM compromise RB-mediated transcriptional repression and cell cycle inhibition (71). In contrast, the discrete role of HDACs in cell cycle control is less clear.
Acetylation and deacetylation of core histones is a key mechanism by which transcription can be altered (21). In general, transcriptional activators recruit histone acetyltransferases (HATs), which add acetyl groups to the lysine residues of histone tails. This acetylation neutralizes the positive charge on lysine, resulting in loosening of chromatin structure. Such disruption is believed to unfold DNA allowing basal transcription machinery to gain access to the promoter regions of genes to be activated. In contrast, histone deacetylases
(HDACs) remove acetyl groups from lysine residues of core histones, thereby preventing basal transcription machinery access to the promoter. Indeed, transcriptional repressors recruit HDACs and utilize their function to suppress transcription (21,24-26,35,51). HDACs belong to a family of enzymes divided into three different classes, encompassing greater than 10 distinct proteins (43).
Three of these HDACs (HDAC1-3) have been shown to interact with RB
(6,9,11,36,37,39,40). It was initially demonstrated that RB could recruit HDAC1 and mediate histone deacetylation at a synthetic promoter (39). Further studies have shown that several endogenous RB/E2F-regulated promoters exhibit
36 changes in promoter histone acetylation as a function of cell cycle position
(47,58) . However, direct demonstration of a role for HDACs in cell cycle
inhibition and transcriptional repression mediated by RB has not been extensively elucidated.
Here we focused on delineating the role of HDACs in RB function. We find that RB-mediated repression occurs in concert with the deacetylation of histones.
These deacetylation events are catalyzed through the action of HDACs, as addition of the HDAC inhibitor TSA prevents RB-mediated histone deacetylation.
The functional requirement of HDACs exhibited promoter specificity, as only a sub-set of RB targets remained repressed when HDAC activity is inhibited. The
HDAC-independent repression was shown to be readily reversible, independent of DNA methylation, and required functional SWI/SNF activity. Importantly, inhibition of HDAC activity with TSA was not sufficient to overcome RB-mediated cell cycle arrest. Together, these data provide critical insights into the action of
RB in transcriptional repression and the relative role of HDACs in cell cycle control.
MATERIALS AND METHODS
Cell culture: A5-1 cell line, harboring conditional expression of a phosphorylation site mutant RB (PSM-RB) (1), was cultured in Dulbecco’s
modified Eagle’s medium (DMEM) supplemented with 10% heat-denatured fetal
bovine serum (FBS), glutamine, penicillin/streptomycin, G418 (400 µg/ml),
37 hygromycin B (200 µg/ml), and doxycycline (1 µg/ml). To induce expression of
PSM-RB, cells were washed with PBS and maintained in media lacking doxycycline for times as indicated. Trichostatin A (Sigma, St. Louis, MO) was
added to the culture media in concentrations as described. Freshly prepared
media containing 5 µM 5-aza-2-deoxycytidine (Sigma, St. Louis, MO) was added every 24 hrs for 96 hrs, and cells were subsequently maintained in the presence or absence of Dox for another 18 hrs.
Western blotting: Immunoblotting was performed following standard biochemical techniques. To detect histone H4 and acetylated histone H4, proteins were isolated by acid-extraction. Antibodies used were RB 851 (gift from
Dr. Jean Wang), Cyclin A (sc-751; Santa Cruz), Cdc2 (sc-747; Santa Cruz ),
topoisomerase IIα (topoIIα: TopoGen, Inc.), thymidylate synthase (gift from Dr
Masakazu Fukushima), RNRII (sc-10848, Santa Cruz), BRG-1 (sc-17796, Santa
Cruz), β-tubulin (sc-5274, Santa Cruz), FLAG M2 (F-3165; Sigma), histone H4
(07-108; Upstate Biotechnology), acetylated histone H4 (06-866; Upstate
Biotechnology), HDAC1 (sc-7872; Santa Cruz ) and HDAC3 (sc-11417; Santa
Cruz).
Plasmids: Primers were used to amplify from rat genomic DNA the following gene promoter regions: Topoisomerase IIα, bases -217 to -19 (79);
Thymidylate synthase, bases -124 to -5 (38); Cdc2, bases -193 to -1 (66); Cyclin
A, bases -135 to +33 (67); Ribonucleotide reductase II, bases –310 to –26.
These promoter regions were cloned into the firefly luciferase expression vector
38 pGL2-Basic (Promega, Madison, WI) and then confirmed by DNA sequencing.
The pTS-dnBRG-1 construct has been previously described (13,14).
Generation of stable cell lines and transcriptional reporter assays:
The reporter constructs along with the empty vector pGL2-B were integrated into
the A5-1 cell line, and three clones were isolated for each construct. Each clone
was cultured in the presence or absence of Dox for 18 hours. Cells were then
harvested, and luciferase activity was quantified using the Promega luciferase
assay kit. Luciferase activity was normalized to total protein concentration by DC protein assay (Bio-Rad, Hercules, CA). Reporter assays were also performed on cells treated with 100 nM of the HDAC inhibitor trichostatin A (Sigma, St. Louis,
MO) for 18 hours in the presence or absence of Dox. The pTS-dnBRG-1 plasmid was transfected into A5-1 cells and selected for the stable inducible expression of dnBRG-1 by FLAG immunoblot.
RT-PCR: A5-1 cells were maintained in the presence or absence of Dox for 18 hours prior to RNA extraction using TRIzol (Invitrogen, Carlsbad, CA).
Reverse transcription of purified RNA was performed using oligo-dT priming and
SuperscriptII RT (Invitrogen). cDNA was then amplified for 24 to 28 cycles using the following primer pairs: Topoisomerase IIα, 5’-
TGCCCAGTTAGCTGGGTCAGTG-3’, 5’-TGAGCATTGTAAAGATGTACCT-3’
(200 bp); Thymidylate synthase, 5’-TTTTATGTGGTGAATGGGGAGC-3’, 5’-
TGGGAAAGGTCTTGGTTCTCGC-3’ (231 bp); Cdc2, 5’-
GGATTGTGTTTTGTCACTCCCG-3’, 5’-CCTATGCTCCAGATGTCAACCG-3’
(229 bp); Cyclin A, 5’-GAGAATGTCAACCCCGAAAAAG-3’, 5’-
39 TGGTGAAGGCAGGCTGTTTAC-3’ (205 bp); RNRII, 5’-
CTTCAACGCCATTGAGACAA-3’, 5’-TCACAGTGCAGACCCTCATC-3’ (234 bp);
β-actin, 5’-ATGGATGACGATATCGCTGC-3’, 5’-CTTCTGACCCATACCCACCA-
3’ (150 bp); and BRG1-FLAG, 5’-GCCCGTGGACTTCAAG-3’, 5’-
CGTCGTCCTTGTAGTCG-3’ (450 bp). PCR products were resolved by agarose gel electrophoresis and visualized by ethidium bromide staining. RT-PCR was also performed on RNA extracted from A5-1 cells treated with 100 nM TSA for 18 hours in the presence or absence of Dox.
BrdU incorporation and flow cytometry: A5-1 cells were cultured in the presence or absence of Dox and 100 nM TSA for 16 hrs. To detect progression through S-phase, cells were pulse-labeled with bromodeoxyuridine (BrdU:
Amersham). Following 8 hrs of labeling, cells were fixed and BrdU incorporation was detected by indirect immunofluorescence (anti-BrdU, Accurate Scientific).
Results are representative of three independent experiments. Flow cytometry was performed as previously described (32).
Chromatin immunoprecipitation (ChIP) assays: ChIP assays were performed as previously described (78) with a few modifications. A5-1 cells were cultured in 15 cm culture plates with or without Dox and TSA for 24 hours.
Formaldehyde (Fisher Scientific) was added directly into the culture medium to a final concentration of 1% and fixed for 15 min at room temperature with mild shaking. To stop the fixation reaction, glycine was added to a final concentration of 0.125 M. Cells were then washed with ice-cold PBS and harvested by trypsinization (20% in PBS). Cells were lysed in cell lysis buffer (5 mM PIPES
40 [pH 8.0], 85 mM KCl, 0.5% NP-40, 0.5 mM phenylmethylsulfonyl fluoride, 100 ng
leupeptin and aprotinin/ml) and then incubated on ice for 10 min. Nuclei were
collected by microcentrifugation (3,000Xg for 5 min) and resuspended in nucleus lysis buffer (50 mM Tris-Cl pH 8.1, 10 mM EDTA, 1% SDS, 0.5 mM phenylmethylsulfonyl fluoride, 100 ng/ml leupeptin and aprotinin). After incubation on ice for 10 min, nuclei were sonicated 7 times with 10 sec pulses and then centrifuged. Chromatin solution was precleared with Staphylococcus aureus protein A-positive cells for 15 min at 40C. Prior to use, Staph A cells were blocked with sheared herring sperm DNA and bovine serum albumin (BSA) for at least 3 hrs at 40C. Precleared chromatin from approximately 107 cells was incubated with 1 µg of the indicated antibodies and without an antibody for at least 3 hrs at 40C. Mock IP contained nuclear lysis buffer only. Antibodies used
were: acetylated histone H4 (06-866; Upstate Biotechnology), dimethyl histone
H3 K9 (07-212, Upstate Biotechnology), E2F4 (sc-1082X; Santa Cruz), Dbf-4
(sc-11354; Santa Cruz), and HDAC1 (sc-7872; Santa Cruz). Staph A cells were
then added and incubated at room temperature for 15 min. Staph A-Immune complexes were washed twice in dialysis buffer (2 mM EDTA, 50 mM Tris-Cl [pH
8.0], 0.2% N-lauryl sarcosine), 5 times in IP wash buffer (100 mM Tris-Cl [pH
9.0], 500 mM LiCl, 1% NP-40, 1% deoxycholic acid) and twice in TE (10 mM Tris
[pH 8.0], 1 mM EDTA). After washing, immune complexes were eluted by adding elution buffer (50 mM NaHCO3, 1% SDS). Inputs were processed from 1% of the total chromatin used in immunoprecipitations. Cross-links were reversed by the addition of NaCl to a final concentration of 300 mM, and RNA was removed by
41 addition of 10 µg of RNase per sample followed by incubation at 650C for 6 h.
DNA was purified using QIAquick PCR Purification Kit by following the manufacturer’s protocol (QIAGEN, Valencia, CA). Promoter regions for the indicated genes were then amplified using the following primer pairs: Cyclin A 5’-
CGACCGGCGCTCCTGGTGACGTC-3’, 5’-TGGCGGCCGACGCACGGAGCA-
3’; Cdc2 5’-TGAGCTCAAGAGTCAGTTGGCGCC-3’, 5’-
CGGCACAGCAGTTTCAAACTCAC-3’; TopoIIα 5’-
GACCGTCTGCGATTGATTGC-3’, 5’-TGACCGTCCTGAAGGGGCTC-3’;
Thymidylate synthase 5’-GGGTCTGTCAATTTCGG-3’, 5’-
GAGCAGTCTGGTGGCAGTGTAGTC-3’; Myogenin 5’-
AGAGGGAAGGGGAATCACAT-3’, 5’-TCCATCAGGTCGGAAAAGAC-3’, HPRT
5’-CAGGCCCAACTTGTCAGAAC-3’, 5’-TGCACAACACCTCAGAGACG-3’. PCR was performed in a 50 µl volume containing 3 µl of purified DNA, 50 ng of each primer set, 0.25 units of Taq DNA polymerase (Promega) and 5 µCi of [α-32P] dCTP. PCR parameters were 940 C for 4 min; 27 cycles of 940 C for 30 sec, annealing for 30 sec, 720 C for 30 sec, and a final extension at 720 C for 10 min.
PCR products were separated on a 6% polyacrylamide gel and visualized with a phosphorimager (Molecular Dynamics, CA).
Restriction enzyme accessibility assay: The restriction enzyme accessibility assay was used to investigate nucleosome position at the cyclin A promoter. This assay was performed as previously described (4,56). Nuclei were harvested from A5-1 cells grown in the presence and absence of Dox for 18 hrs.
Nuclei permeabilized in 0.5% NP-40 were digested with EagI for 2 hrs at 37oC
42 followed by digestion in 1 mg/ml proteinase K (Sigma) overnight. Protein-free
genomic DNA was extracted and subsequently digested overnight with KpnI and
EcoRI. Digested DNA was then resolved by agarose gel electrophoresis and
transferred onto an Immobilon-Ny nylon membrane (Millipore). Enzyme
accessibility was visualized by radioactive Southern blot with a probe generated
by PCR against bases -135 to +33 of the cyclin A promoter.
RESULTS
Active RB represses cell cycle genes: Studying the function of RB in
mediating transcriptional repression is hampered, as wild type RB is readily
phosphorylated/inactivated by CDK/cyclin complexes in most cell types. To
circumvent this problem, we utilized a phosphorylation site mutant of RB (PSM-
RB) that is refractory to phosphorylation and is therefore constitutively active. In
this study, a Rat-1 derived cell line with tetracycline-regulated PSM-RB
expression (A5-1) was utilized to specifically analyze the downstream action of
RB. In order to confirm the cell cycle inhibitory potential of PSM-RB in this
setting, we examined bromodeoxyuridine (BrdU) incorporation of A5-1 cells
cultured in the presence or absence of the tetracycline analogue doxycycline
(Dox). Consistent with previously reported data, we found that the induction of
PSM-RB upon removal of Dox dramatically reduced the number of BrdU-positive cells (Fig. II-1A). Therefore, A5-1 cells represent an effective model to study RB activity.
43 We have recently identified and validated a large number of RB repressed
target genes using microarray analysis (42). To initially investigate the promoter
activity of representative genes, we cloned regions of the cyclin A, cdc2,
topoisomerase IIα (topo IIα), and thymidylate synthase (TS) promoters into the
luciferase reporter vector pGL2-B (Fig. II-1B). These cloned promoter fragments
represent less than 2 nucleosomes of DNA and are thus useful for investigating
the regulatory action of RB on a relatively small region of chromatin. The
reporter constructs and the vector (pGL2-B) were then integrated into A5-1 cells
to provide an effective means for studying promoter activity in the context of
chromatin. Three independent clones for each construct were selected and
maintained in medium with or without Dox for 18 hours. We found that the activity
of cyclin A, cdc2, topoIIα, and TS promoters was reduced 3 to 5 fold upon induction of PSM-RB expression (Fig. II-1C, compare +Dox to –Dox). In contrast, PSM-RB had no effect on the basal transcription of pGL2-B. These data demonstrate that active RB represses the promoter activity of these genes in chromatin.
Next we investigated the correlation between promoter activity and endogenous RNA levels (Fig. II-1D). We utilized RT-PCR to determine the levels of endogenous RNA in A5-1 cells that were cultured in the presence or absence of Dox. Induction of active RB attenuated the expression of cyclin A, cdc2, topoIIα and TS RNA (Fig. II-1D, compare +Dox to –Dox). In contrast, induction of
PSM-RB had no effect on the expression of the β-actin gene. Consistent with these observations, endogenous protein levels of cyclin A, cdc2, topoIIα, and TS
44 were also attenuated when Dox was removed from the mediium (Fig. II-1E,
compare +Dox to -Dox). The attenuations are due to the presence of PSM-RB and not merely to differences in loading, as revealed by immunoblotting for β- tubulin. Immunoblotting with a PSM-RB specific antibody clearly demonstrated that PSM-RB is only expressed in the absence of Dox (Fig. II-1E). Taken together, these results confirm that active RB inhibits the expression of cyclin A, cdc2, topoIIα, and TS genes by repressing promoter activity.
Active RB induces histone deacetylation at promoters of specific cell cycle genes: RB has been shown to recruit HDACs to the promoters of target genes (81). This event is believed to be responsible for transcriptional repression
and the subsequent anti-proliferative action of RB. However, the extent to which
HDACs are required for RB to act as a transcriptional repressor and tumor
suppressor is unclear. To assess the action of HDAC enzymatic activity in RB-
mediated transcriptional repression, we utilized in vivo formaldehyde cross-
linking of DNA-protein complexes followed by chromatin immunoprecipitation
(ChIP). Initially, we validated the linearity of PCR amplification over a wide range
of template concentrations. Increasing amounts of input chromatin were amplified
with primers specific for the cdc2 promoter in the presence of [α- 32P] dCTP for quantitation. As shown in figure II-2A, the PCR reaction was linear throughout the titration, indicating that changes in chromatin levels could be accurately observed by radioactive PCR. Using this approach, we then investigated the promoters of the cyclin A, cdc2, topoIIα and TS genes for changes in histone H4 acetylation, as these targets were repressed by PSM-RB (Fig. II-2B).
45 Specifically, A5-1 cells were cultured in the presence or absence of Dox for 24
hours, were formaldehyde cross-linked, and ChIP assays were performed
utilizing acetylated histone H4 antibody. For each target promoter, input lanes
(non-immunoprecipitated DNA) confirmed that equal amounts of chromatin were used in all ChIP assays (Fig. II-2B, lanes 1 and 2). The Dbf-4 antibody was
utilized as a negative control, since the replication factor Dbf-4 is not expected to
occupy these promoters (Fig. II-2A, lanes 5 and 6). We found that, in presence
of PSM-RB, acetylated histone H4 association with the promoters of cyclin A, cdc2, topoIIα, and TS was nearly abolished (Fig. II-2B, compare lanes 3 and 4).
This effect is specific to RB-repressed genes, as no change in histone acetylation
was observed on the hypoxanthine-guanine phosphoribosyltransferase (HPRT)
promoter (Fig. II-2B). In contrast with acetylated histone H4 occupancy, which
was diminished in the presence of PSM-RB, we found that E2F4 occupancy was
unaffected by the induction of active RB (Fig. II-2C, compare lanes 3 and 4).
Moreover, we detected HDAC1 bound to cdc2 promoter preferentially in
presence of PSM-RB (Fig. II-2C, compare lanes 5 and 6). Therefore, the
decrease in acetylated histone H4 on these promoters is a consequence of PSM-
RB induction and not all protein-promoter interactions were similarly affected.
The reduction in acetylated histone H4 at these promoters strongly suggests that
these promoters are actively deacetylayed. This observation is consistent with
RB/HDAC complexes acting on these promoters.
RB-induced histone deacetylation at specific promoters is mediated
by HDACs: Histone deacetylation at the promoters of cyclin A, cdc2, topoIIα,
46 and TS in the presence of active RB (shown in Fig. II-2) supports a critical role
for HDAC in transcriptional repression. To determine the explicit requirement of
HDACs for this transcriptional repression, we used trichostatin A (TSA), a
pharmacological inhibitor of HDAC activity. Initially we determined the effect of
TSA on bulk histone acetylation (Fig. II-3A). A5-1 cells were cultured in the
presence and absence of Dox and TSA for 24 hours. Cells were then harvested,
and the levels of acetylated histone H4 and total histone H4 were evaluated by
immunoblotting (Fig. II-3A). We found that TSA treatment resulted in the marked
accumulation of acetylated histone H4 in a dose-dependent manner (Fig. II-3A,
top panel compare lanes 1, 2, 3 and 4). Importantly, analysis of total histone H4
levels showed that TSA inhibited deacetylation and did not merely augment the expression of histone H4. Furthermore, TSA did not affect the expression of
PSM-RB, HDAC1, or HDAC3. While performing these studies, we found that
100 nM TSA had a minimal effect on cell viability (data not shown). Therefore, we
employed 100 nM TSA to study the requirement of HDAC activity for RB-
mediated transcriptional repression.
To investigate the requirement of HDAC activity for RB-mediated histone
deacetylation, we used TSA to inhibit HDAC activity and then monitored
promoter histone acetylation. For this analysis, A5-1 cells were treated with or
without Dox and TSA for 24 hours. These cells were then used as substrates for
ChIP assays with antibodies specific for acetylated histone H4 (Fig. II-3B). Non- immunoprecipitated chromatin (Inputs) showed relative amount of chromatin used in each immunoprecipitation (Fig. II-3B, lanes 2-5). Immunoprecipitation
47 with a non-specific antibody Dbf4 (Fig. II-3B, lanes 7-10), without an antibody
(Fig. II-3B, lanes 11-12), and without chromatin (Fig. II-3B, lane 6) did not result
in appreciable PCR product and thus provided evidence of specificity for the
ChIP assay. Consistent with data described above, we observed histone H4 deacetylation on the promoters analyzed (Fig. II-3B, compare lanes 13 and 14).
Surprisingly, TSA by itself did not augment the histone H4 acetylation observed
on any promoter in the presence of Dox (Fig. II-3B, compare lanes 13 and 15),
indicating that HDAC activity was likely absent from the promoter in normal
asynchronously proliferating cells. However, inhibition of HDAC activity by TSA
resulted in marked acetylation of histone H4 on the promoters of topoIIα and TS,
in the presence of PSM-RB (Fig. II-3B, compare lane 14 to 16). Similar results
were obtained for cyclin A and cdc2 promoters. The use of [α 32P] dCTP in PCR
enabled the quantitative analysis of product signal intensity. PCR product bands
from acetylated histone H4 immunoprecipitations were normalized to their
corresponding inputs to account for variations in chromatin used in the ChIP
assay. Quantitation of PCR amplified products using phosphorimager clearly
showed that TSA results in re-acetylation of these promoters (Fig. II-3C).
Collectively, these data demonstrate that RB-induced deacetylation of cyclin A,
cdc2, topoIIα, and TS promoters require HDAC activity. Since histone deacetylation is a critical mechanism of transcriptional repression, these results suggest that TSA may reverse RB-mediated cell cycle inhibition.
HDAC activity is dispensable for RB-mediated cell cycle arrest: To monitor the influence of HDAC inhibition on cell cycle progression, A5-1 cells
48 were cultured in the presence or absence of Dox and TSA for 16 hours before
being pulsed with BrdU for 8 hours. Surprisingly, cells cultured in the absence of
Dox remained inhibited for BrdU incorporation in the presence of TSA (Fig. II-4A and B). To examine cell cycle distribution, we performed flow cytometric analysis of cells treated with or without Dox and TSA. As shown in Fig. II-4C, we found that treatment with or without TSA, concurrent with PSM-RB (-Dox) expression, did not significantly alter cell cycle position. Taken together, these data demonstrate that RB is able to maintain cell cycle arrest even in the absence of
HDAC activity.
HDAC requirement for repression is promoter specific: The failure of
TSA to rescue cells from RB-mediated cell cycle arrest suggested that inhibition of HDAC activity was not sufficient to alleviate transcriptional repression. Having established that histone deacetylation at cyclin A, cdc2, topoIIα, and TS promoters by RB is the result of HDAC activity (Fig II-3B), we sought to elucidate the functional significance of this event (Fig. II-5). A5-1 cells with integrated luciferase reporters of cyclin A, cdc2, topoIIα, TS, and vector pGL2-B were cultured in the presence or absence of Dox and TSA for 18 hours. Consistent with the failure of TSA treatment to augment promoter histone acetylation in the absence of PSM-RB (+Dox), TSA did not significantly stimulate basal promoter activity (not shown). Analysis of RB-mediated repression showed that TSA significantly alleviated RB-mediated promoter repression of cdc2, topoIIα, and TS
(Fig. II-5A). However, the promoter activity of cyclin A was not recovered in the presence of TSA (Fig. II-5A).
49 The expression of these targets was also monitored through investigation
of endogenous RNA levels (Fig. II-5B). Consistent with the reporter assays, we found that TSA ameliorated the attenuation of cdc2, topoIIα, and TS RNA levels that is mediated by PSM-RB (Fig. II-5B). Again at the RNA level, TSA failed to
relieve the inhibition of cyclin A by PSM-RB. As expected, β-actin RNA levels
were unaffected upon treatment with either Dox or TSA (Fig. II-5B).
To determine whether the changes in RNA levels led to meaningful
changes in protein levels, immunoblot analysis was performed. Analysis of cyclin
A, cdc2, topoIIα, and TS protein levels demonstrated PSM-RB-mediated
attenuation (Fig. II-5C, compare lanes 1 and 2). Consistent with the promoter and
RNA analyses, treatment with TSA reversed the RB-mediated attenuation of
cdc2, topoIIα, and TS protein levels (Fig. II- 5C, compare lanes 3 and 4).
However, TSA did not recover RB-mediated inhibition of cyclin A protein levels
(Fig. II- 5C, compare lanes 3 and 4).
In addition to cyclin A, we analyzed the promoter activity of other genes
associated with the RB/E2F-signaling axis and found that ribonucleotide
reductase subunit II (RNRII) promoter activity was not recovered with TSA
treatment (Fig. II-5D). Specifically, an integrated RNRII reporter was repressed
by PSM-RB and this repression was not alleviated with TSA (Fig II-5D, left
panel). Moreover, RNRII RNA (Fig. II-5D, middle panel) and protein (Fig. II-5D,
right panel) levels were similarly reduced by RB action in the absence and
presence of TSA. Thus, RNRII, like cyclin A, is also an HDAC-independent
target of RB-mediated repression.
50 HDAC-independent mechanism of cyclin A repression: It is known that transcriptional repression elicited by RB can occur through the action of co- repressors in addition to HDACs. Specifically, it has recently been demonstrated that the cyclin A promoter is irreversibly silenced during induced senescence
(48). This pathway likely involves histone H3 lysine 9 methylation, heterochromatin protein 1 (HP1) recruitment, and DNA methylation of the promoter (34,48,52). Therefore, we initially analyzed the histone H3 lysine 9 methylation of the cyclin A promoter during RB-mediated arrest (Fig. II-6A). Cells were cultured in the presence or absence of Dox to induce PSM-RB and isolated chromatin was subjected to ChIP analysis using antibodies specific for histone
H3 methylated on lysine 9. We observed histone H3 lysine 9 methylation on the myogenin promoter that is silenced in fibroblastic cells (Fig. II-6A). In contrast, we failed to detect histone H3 lysine 9 methylation on the cyclin A promoter above background, suggesting that this silencing mechanism is not responsible for cyclin A repression. Consistent with this observation, culture in 5-Aza-2- deoxcytidine which blocks DNA methylation and reverses silencing
(41,45,50,80), failed to augment cyclin A promoter activity or protein levels in the presence of PSM-RB (Fig. II-6B). Lastly, we determined whether the RB- mediated repression of the cyclin A promoter was reversible. To do so, cells were cultured in the absence of Dox to induce PSM-RB, and cyclin A promoter activity was repressed (Fig. II-6C, left panel). Re-addition of Dox to the media resulted in the restoration of cyclin A promoter activity. Additionally, cyclin A protein levels were restored following the re-addition of Dox (Fig. II-6C, right
51 panel). Together, these results indicate that stable epigenetic silencing
mechanisms are not responsible for the observed RB-mediated repression of the cyclin A promoter.
In addition to HDACs and silencing mechanisms, we and others have demonstrated that SWI/SNF activity plays a requisite role in the repression of the cyclin A promoter (70,81). Consistent with these studies, we observed that unlike 5-Aza-2-deoxcytidine or TSA, ectopic expression of dominant negative
BRG-1 (dnBRG-1) that inhibits SWI/SNF activity augments the expression of cyclin A RNA (Fig. II- 7A, left panel) and protein (Fig. II-7A, right panel) levels in the presence of PSM-RB (Fig. II-7A). These results indicate that chromatin remodeling represents a critical means through which cyclin A repression occurs.
One possible explanation is that SWI/SNF functions in concert with RB to position nucleosomes near the transcription start site to inhibit transcription of the cyclin A promoter. Analysis of promoter structure was carried out in the presence or absence of PSM-RB using restriction enzyme accessibility assay in the proximity of the transcription start site. We observed that chromatin from cells cultured in the absence of PSM-RB (+Dox) was readily digestible (Fig. II-
7B, lane 3), whereas chromatin from cells expressing PSM-RB (-Dox) was resistant to enzyme cleavage (Fig. II-7B, lane 4). This change in chromatin structure was largely dependent upon SWI/SNF as dnBRG-1 retarded the formation of the nuclease resistant chromatin structure (Fig. II-7B, lane 6).
These results indicate that chromatin remodeling occurs on the cyclin A promoter to mediate transcriptional repression.
52 DISCUSSION
RB represses the expression of multiple genes involved in cell cycle transitions. This transcriptional control has been attributed to multiple corepressors recruited by RB. Here we specifically focused on elucidating the role of HDACs in RB-mediated transcriptional repression of four critical targets
(cyclin A, cdc2, topoIIα, and TS). We demonstrate that active RB leads to histone deacetylation on the promoters of these four genes, and the observed deacetylation is dependent on HDAC activity. This action of HDACs was required for the transcriptional repression of cdc2, topoIIα, and TS genes, as TSA reversed RB-mediated repression. However, this action was promoter specific as TSA failed to recover cyclin A or RNRII levels. Analysis of the HDAC- independent repression of cyclin A indicated that it was not due to epigenetic silencing mechanisms, but was reversible and involved chromatin remodeling.
Importantly, we demonstrate that cell cycle inhibitory action of RB is independent of HDAC enzymatic activity. Together, these results demonstrate the intricate interplay between RB and HDACs in transcriptional regulation and cell cycle control.
Role of HDACs in RB-mediated transcriptional repression: HDACs represent co-repressors identified to interact with RB (6,39,40).However, relatively few studies have analyzed their role in RB-mediated transcriptional repression and cell cycle control. Originally, it was demonstrated that HDAC activity is associated with RB, but only in the context of a synthetic promoter was
53 histone deacetylation observed (39). Subsequent studies have shown that
specific RB/E2F target genes in fact undergo changes in promoter histone
acetylation during the cell cycle (47,58). Our results shown here demonstrate
that RB-mediated repression leads to histone deacetylation at all promoters analyzed. In principle, such an effect could be due to either recruitment of HDAC or the inhibition of HAT activity. Inhibition of HAT as a mechanism for promoter
deacetylation is not without merit, as E2F proteins likely employ HAT-dependent
mechanisms for gene activation (74). By using the HDAC inhibitor TSA we could
specifically demonstrate that the deacetylation of these promoters is dependent on the enzymatic activity associated with class I HDAC molecules (43). The effect of RB on histone acetylation is not due to a bulk-deacetylation phenomenon, as total cellular levels of acetylated histones are not changed by the expression of PSM-RB. Interestingly, while TSA does lead to bulk histone hyperacetylation, it does not lead intrinsically to the hyperacetylation of histones
at the E2F/RB regulated promoters studied here. This could be because the
histones at these promoters are already hyperacetylated or because HDAC is not
associated with the promoter in asynchronously proliferating cells. Such a
hypothesis is supported by the finding that we observed HDAC1 specifically
associated with the cdc2 promoter when PSM-RB was expressed. Clearly in the
context of RB-mediated repression, the mechanism of histone deacetylation is
HDAC-dependent and can be reversed by TSA on all promoters analyzed. The inhibition of histone deacetylation by TSA had a functional consequence in the context of cdc2, topoIIα and TS gene expression. Interestingly, even in cases
54 where TSA only partially reversed RB-mediated histone deacetylation, TSA was
capable of fully restoring promoter activity (i.e., the cdc2 promoter). Such a finding, for which there is precedent in the literature (60, 83), suggests that only a moderate level of histone acetylation is required for transcriptional activity on specific promoters. In fact, TSA fully restored the promoter activity, endogenous
RNA, and protein levels of cdc2, topoIIα, and TS in the presence of PSM-RB.
Such a finding is critical, as it demonstrates that HDAC activity represents the sole means through which RB mediates repression of these targets. In contrast, we failed to observe recovery of cyclin A expression when HDAC activity was inhibited in presence of RB, even though promoter histones were acetylated
following TSA treatment.
HDAC-independent mechanisms of transcriptional repression: The findings we observe with cyclin A gene regulation indicate that RB utilizes mechanisms in addition to histone deacteylation to mediate repression. Such a conclusion is not without precedent (44,81). However, to definitively make such a conclusion it is critical to determine that inhibition of HDAC activity actually reversed the histone deacetylation on the promoter. We clearly observed that
TSA was sufficient to efficiently reverse RB-mediated histone deacetylation on the cyclin A promoter. Thus, the failure of TSA to reverse RB-mediated repression of the cyclin A promoter is due to a mechanism that is clearly distinct from HDAC activity. Additionally, while we did not explicitly evaluate the RNRII promoter, it behaved in a manner similar to that of cyclin A in that its repression
55 was independent of HDAC activity. Therefore, it seems likely that repression of
cyclin A and RNRII by RB is dependent on other chromatin modifying factors.
RB has been shown to associate with other chromatin modifying enzymes
to mediate transcriptional repression. Specifically, recent studies indicate that
RB target genes (including cyclin A) are silenced through a mechanism involving
histone methylation and HP1 chromatin association (48,52). Such silencing
which is observed in senescent cells is irreversible (48,52). Here, we find that
the repression of cyclin A by PSM-RB does not involve irreversible silencing
mechanisms, as we failed to detect an influence of histone or DNA methylation
on RB-mediated repression of the cyclin A promoter. Additionally, the repression
of the cyclin A promoter was readily reversible. Such a result is consistent with
the ability of quiescent cells with the cyclin A promoter repressed to re-enter the cell cycle (48) and with the failure to detect histone methylation or HP1 chromatin association with the cyclin A promoter in quiescent cells (48).
In addition to silencing mechanisms and HDACs, RB is known to associate with components of the SWI/SNF chromatin remodeling complex and the activity of SWI/SNF is required for the repression of cyclin A. For example, expression of PSM-RB or p16ink4a to activate endogenous RB in BRG-1/BRM deficient cell lines does not attenuate cyclin A and RNRII levels or lead to cell cycle arrest (70,81, Gunawardena, unpublished data). One means through which SWI/SNF acts is by facilitating histone acetylation and deacetylation. This action of SWI/SNF is not sufficient for cyclin A repression as the promoter remained repressed even with histones acetylated. Rather, our results suggest
56 that SWI/SNF is functioning to modify chromatin structure to inhibit transcription.
Consistent with this model, we find that the nuclease accessibility in the region of the cyclin A transcription start site is inhibited during RB-mediated repression.
Thus, chromatin topology, not histone modification, is likely sufficient for RB- mediated repression events on the cyclin A promoter.
HDAC activity is not required for RB-mediated cell cycle inhibition:
Another finding from our studies is that inhibition of HDAC activity by TSA does not rescue cells from arrest imposed by RB. This result is quite surprising, as
TSA exerts pronounced effects on gene expression, but has no detectable effect on RB-mediated cell cycle inhibition. Most likely failure of TSA to block RB- mediated repression of cyclin A and similarly regulated genes (e.g. RNRII) is responsible for this phenomenon. Cyclin A is required for traversing of the cell cycle (19,55), and thus lack of cyclin A expression alone might well explain the failure of cells to proliferate when HDAC activity is inhibited. An important extension of our work is based on the current use of HDAC inhibitors in clinical trials for treatment of several forms of cancer (33). In this context, HDAC inhibitors are believed to reactivate genes that have been inappropriately silenced during tumor progression and thus inhibit tumor growth. Based on our findings that TSA does not reverse RB-mediated cell cycle arrest, we conclude that the treatment of tumors wih HDAC inhibitors will not have the undesired effect of inactivating the RB pathway of cell cycle inhibition.
57 In summary, transcriptional repression of cell cycle genes by RB is a
complex process involving multiple chromatin modifying factors. Here we find
one class of factors, HDACs, play a critical role in the transcriptional repression programs elicited by RB. This action of HDACs is promoter specific and as such is not required for RB-mediated cell cycle inhibition.
ACKNOWLEDGMENTS
We are grateful to Dr. Karen Knudsen, Dr. Christopher Mayhew, Steven
Angus, Craig Burd, Michael Markey, and Christin Petre for critical reading of the manuscript. We thank Shelly Barton, Anthony Imbalzano, Bernard Weissman
and members of the Knudsens’ laboratories for helpful discussions. Sandy
Schwemberger provided technical assistance with flow cytometry analysis.
This work was supported by grants to ESK from the National Cancer
Institute (CA82525).
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71 A
100
80
60
40
20
Relative % BrdU Postive BrdU % Relative 0 +Dox -Dox
B
rat topoIIα promoter -217 GACCGTCTGCGATTGATTGCTGTAAACAGAGCAAATGAGAGCGAGTCGGTGATTGGTTCCTCTGGATTTGGGCGGGCT C/EBP ICB4 E2F -137 ATGAATGTTCAAATGAAACTAGGGAAGCTCTCCTAACCGACGCAACGTCTGTGGAGAAGCGTCCAAGTCGCCGGTCGC B-myb -57 AGCGTTTCTCACTGATTTTGTCGCTTTTAGGATTCTCGAG rat cyclin A promoter -135 CGACCGGCGCTCCTGGTGACGTCACTGGCCCCGAACCGCCCTCATTAGGTCCATTTCAATAGTCGCGGGCTACTTGAA ATF Sp1 E2F -55 ACAAGAACAGCCGGCGGGCCGCCGGCTCGCGGCACTTGGGCTCTGCGCTCCTCCCTCCCGCAGCGCCTGCTCCGTGCG E2F +26 GGCCGCA rat TS promoter -124 GGTCTGTCAATTTCGGCGGCCAGAAGCTTCCAGCAGGAAGAGGCGGGCTCGAGAGGAGGAAAAGAGCAACCGGAAGGT E2F Ets Sp1 Ets Ets -44 GGGTTTTGGCGCCGACTACACTGCCACCAGACTGCTC E2F rat cdc2 promoter -193 TGAGCTCAAGAGTCAGTTGGCGCCCGCCCTCCTGTAATTCCTCCGGCCGCGGTTTCCGCTCCCTTTCGCGCTCTGCAC E box Sp1 E2F -113 CCAGGCAGCCCGGGCCGTAGCTTAGCTCGGCTCTGATTGGCTCCTTTGAACGTCTACGTGCAATCGGATTGGCGGACC
-33 GGGAGCTTTACCGCGGTGAGTTTGAAACTGCTG
Figure II-1: PSM-RB (active RB) represses a set of cell cycle genes and induces cell cycle arrest. A) A5-1 cells were cultured in the presence or absence of Dox for 16 hrs prior to BrdU labeling for an additional 8 hrs.
Percentage of BrdU positive cells was determined from three independent experiments. B) The cloned rat regulatory regions of different RB target genes analyzed in present study.
72 C
100
80
60 +Dox 40 -Dox
20 Relative luciferase activity (%) 0 TopoIIα TS Cyclin A Cdc2 pGL2-B D TopoIIα TS Cyclin A Cdc2 β-actin Dox : ++++--+ - - - +
12 34678910 5 E + - + - : Dox PS M- RB Cdc 2
Tubulin TopoIIα
Cyclin A TS 12 12
Figure II-1: PSM-RB (active RB) represses a set of cell cycle genes and induces cell cycle arrest. C) A5-1 parental cell line was stably transfected with rat promoter constructs for the indicated genes or empty vector (pGL2-B) driving expression of the firefly luciferase gene. Three clones were selected for each construct, maintained in the presence or absence of Dox for 18 hrs, and then assayed for luciferase activity. D) A5-1 cells were cultured in the presence or absence of Dox for 18 hrs. Total RNA was then extracted and reverse transcribed into cDNA. This cDNA was subjected to linear PCR amplification with
73 specific primers for the indicated genes. E) A5-1 cells were cultured in the presence (lane 1) or absence (lane 2) of Dox for 24 hrs. Total protein was isolated, resolved by SDS-PAGE, and immunoblotted with antibodies as indicated.
74 A Cdc2 Input
l µ l 5 µ l O 2 5 µ l l l 2 .1 .2 .5 µ µ µ H 0 0 0 1 2 4
1234567
R2 = 0.9817
012345 Input (uL)
B Input Ac-H4 Dbf-4 +++- --: Dox
Cyclin A
Cdc2
TopoIIα
TS
HPRT
12 346 5
C Dbf-4 E2F4 HDAC1 k c o + - + - + - M : Dox Cdc2
12 3 456 7
Figure II-2: Active RB induces histone deacetylation at promoters of specific cell cycle genes: A) Total chromatin was isolated from A5-1 cells cultured in the presence of Dox and increasing amounts of chromatin (0 to 4 ul) were subjected to PCR in the presence of [α-32P] dCTP and primers specific for the cdc2 promoter. Production of PCR product was quantified using a phosphor
75 imager. B) A5-1 cells were cultured in the presence (lanes 1, 3, 5) or absence
(lanes 2, 4, 6) of Dox for 24 hrs, cross-linked with formaldehyde, and ChIP
assays were performed as described. Residency of acetylated histone H4 at the
indicated gene promoters was determined by carrying out the ChIP with
antibodies specific to acetylated histone H4 (lanes 3 and 4). Input (lanes 1 and
2) refers to PCR containing 1% of total chromatin used in immunoprecipitation.
Immunoprecipitation with Dbf-4 (lanes 5 and 6) is a negative control. PCR
products were detected by autoradiography. C) As in B) except immunoprecipitation was performed with antibodies specific for Dbf-4 (lanes 1 and 2), E2F4 (lanes 3 and 4), and HDAC1 (lanes 5 and 6). The mock represents a ChIP assay without the inclusion of chromatin substrate. PCR products were detected by autoradiography.
76 A ++++ -- - -: Dox 0 501002000 50 100 200 : TSA (nM) Acetylated Histone H4
Histone H4
PSM-RB
HDAC1
HDAC3 123478 56
B Inputs Dbf-4No Ab Ac-H4
+ - + - k + - + - + - + --+ : Dox c O o 2 H - - + ++M --+ + + --+ + : TSA TopoIIα
TS
1234 5678 910111213141516
C Cyclin A Cdc2 TS TopoIIα 100%% 100% 100%% 100%
75%% 75% 75%% 75%
50%% 50% 50%% 50%
25%% 25% 25%% 25% Relative binding
0%% 0% 0% 0% + - + - + - + - + - + - + - + - + - + - + - + - : Dox ---- ++ ---- ++ ---- ++ ---- ++: TSA Dbf4 Ac H4 Dbf4 Ac H4 Dbf4 Ac H4 Dbf4 Ac H4
Figure II-3: RB induced deacetylation at specific promoters is mediated
through histone deacetylases (HDACs): A) A5-1 cells were maintained in
medium with (lanes 1-4) or without Dox (lanes 5-8) and the HDAC inhibitor TSA
(0, 50, 100, 200 nM) for 24 hrs. Proteins were resolved by SDS-PAGE and
immunoblotted with antibodies as indicated. B) A5-1 cells were cultured in the
presence or absence of Dox and TSA (100 nM). Cells were cross-linked after 24
77 hrs, and ChIP assay was performed with an acetylated histone H4 specific
antibody (lanes 13-16). Inputs (lanes 2-5) represent PCR with 1% of chromatin
utilized in immunoprecipitations. Controls include: H2O (PCR control, lane 1),
Mock (IP without chromatin, lane 6), Dbf-4 (non-specific antibody, lanes 7-10),
and No Ab (IP without antibody, lanes 11-12) C) PCR products from Fig. 3B were quantified using phosphor imager and data were normalized to account for variation in inputs.
78 A +Dox -Dox HoechstBrdU Hoechst BrdU
-TSA
+TSA
B 100
80
60
40
20
0 RelativeBrdU % Positve +Dox -Dox +Dox -Dox -TSA +TSA C
G0-G1=67.44% G0-G1=90.33% G0-G1=59.40% 1200 G0-G1=86.33% 1500 1200 S=24.14% 1200 S=9.39% S=30.09% S=8.55% G2-M=8.42% G2-M=0.28% G2-M=10.51% 900 G2-M=5.11% 900 900 1000 600 600 600 # of cells # of cells # of cells 500 # of cells 300 300 300
0 0 0 0 0 200 400 600 0 200 400 600 0 200 400 600 0 200 400 600 PI PI PI PI +Dox, -TSA -Dox, -TSA +Dox, +TSA -Dox, +TSA
Figure II-4: HDAC activity is dispensable for RB-mediated cell cycle arrest:
A) A5-1 cells were cultured in the presence or absence of Dox and TSA for 16
hrs prior to BrdU labeling for an additional 8 hrs. Cells were fixed and stained
with anti-BrdU antibody and Hoechst to identify proliferating cells and nuclei,
79 respectively. B) Percentage of BrdU positive cells was determined from three independent experiments. C) A5-1 cells were cultured in the presence or absence of Dox and TSA for 24 hrs. Cells were harvested, fixed with ethanol,
and stained with propidium iodide (PI). Cell cycle distribution was then
determined by flow cytometry. DNA content (PI intensity) is plotted against cell
number. Percentage of cells in G0-G1, S, and G2-M was determined by ModFit
software.
80 A 120%
100%
80%
60% +Dox -Dox 40% Relative Luciferase Activity 20%
0% - + - + - + - + - + : TSA TopoIIα TS Cyclin A Cdc2 pGL2-B
BC + --+ : Dox - - + + : TSA TopoIIα TS Cyclin A Cdc2 β-actin Cyclin A Dox : +----- + ++ + TopoIIα + TSA TS 1234 5 6 78910 Cdc2
PSM-RB
Tubulin
1234
Figure II-5: HDAC-dependent deacetylation is required by RB to repress
specific promoters: A) A5-1 integrated reporter cell lines from Fig. 1C were
cultured in the presence or absence of Dox and 100 nM TSA for 18 hrs. Cells
were harvested and assayed for luciferase activity. B) A5-1 cells were cultured in 100 nM TSA in the presence or absence of Dox for 18 hrs. Total RNA was then extracted and reverse transcribed into cDNA. This cDNA was subjected to linear PCR amplification with specific primers for the indicated genes. C) A5-1
81 cells were cultured in the presence (lanes 1 and 3) or absence (lanes 2 and 4) of
Dox in the absence (lanes 1 and 2) or presence (lanes 3 and 4) of 100 nM TSA for 24 hrs. Total protein was isolated, resolved by SDS-PAGE, and immunoblotted with antibodies as indicated.
82 D
rat RNRII promoter -262 CCCCTTGAGCTCTCCAATCCCCGAGCCTCTAGCAATCCAAAACGGTTCCGGCCCCTCGAGCGCCCGCGCGCACCAGGGCG CCAAT -182 GCGCAGACTCCTTGTAGGTCTTTGTGCGGAAGGCCTACGGCGCAACTCAAATCTCCCGCGCTGAGCGGCGAGCGGGGTCA E2F -92 GCGGCGTCCAATCGCAGCCGAGGACACGCCCACCTCGGACCGCGCGATTCGAACGCCCTTTAAAGGGTGCGGACGCCGGC CCAAT TATA -12 AGCTGCCGGTGCACCGGATTCCAGCTGTTCCCTCTTCTCCTCGTC
100%
75% Dox : + - + - Dox : + + - - TSA : -- + + TSA : --+ + 50%
25% 12 34 12 34
Relative Luciferase Activity 0% ++- - : Dox - - + + : TSA
Figure II-5: HDAC-dependent deacetylation is required by RB to repress
specific promoters: D) Top panel: The cloned regulatory region of the rat
RNRII promoter. Left panel: A5-1 parental cell line was stably transfected with a rat RNRII promoter reporter driving expression of the firefly luciferase
gene. Cells were cultured in the presence or absence of Dox with or without 100
nM TSA as indicated and relative luciferase activity was determined. Middle
panel: A5-1 cells were cultured in the presence (lanes 1 and 3) or absence
(lanes 2 and 4) of Dox and the absence (lanes 1 and 2) or presence (lanes 3 and
4) of TSA for 18 hours. Cells were harvested and RNRII RNA levels were determined by RT-PCR. Right panel: A5-1 cells were cultured in the presence
(lanes 1 and 2) or absence (lanes 3 and 4) of Dox and the absence (lanes 1 and
3) or presence (lanes 2 and 4) of TSA for 24 hours. Cells were harvested and
RNRII protein levels were determined by immunoblotting.
83 A Inputs Dbf-4 CH3-K9-H3 +++---:Dox Cyclin A
Myogenin
12 3456
B Cyclin A-Luc 100%
+ - - : Dox 75% - - + : 5-Aza-2-dC
Cyclin A 50%
Tubulin 25% 12 3 Relative Luciferase Activity 0% + - --: Dox --+ - : 5-Aza-2-dC -- -+ : TSA
Figure II-6: The cyclin A promoter is not subjected to stable gene silencing:
A) A5-1 cells were cultured in the presence or absence of Dox as indicated.
Chromatin was isolated and utilized in ChIP assays with antibodies specific for
di-methylated K9 histone H3 (lanes 5 and 6). Input (lanes 1 and 2) and Dbf-4
(lanes 3 and 4) controls are shown. Chromatin was amplified with primers
specific for the cyclin A and myogenin promoters, and products were detected by
autoradiography. B) A5-1 cells harboring the integrated cyclin A reporter were
cultured in the presence of 5-Aza-2-dC as described in the materials and
methods and then cultured in the absence of Dox for 24 hours. Relative
84 luciferase activity was determined by reporter assay (left panel) and endogenous protein levels were determined by immunoblotting (right panel).
85 C Cyclin A-Luc , 150% rs h s rs r h 4 h 2 4 4 x 2 125% 2 o x x x o o -D o D D + -D + 100% Cyclin A 75%
50% PSM-RB
25% Tubulin Relative Luciferase Activity Luciferase Relative
0% 123 x rs s, o h hr D 4 rs + 2 24 h x 4 o ox 2 -D -D ox +D
Figure II-6: The cyclin A promoter is not subjected to stable gene
silencing: C) A5-1 cells harboring the integrated cyclin A reporter were cultured in the presence or absence of Dox for 24 hours. To attenuate PSM-RB, Dox was re-administered to indicated cultures. Relative luciferase activity was determined by reporter assay (left panel) and endogenous protein levels were determined by immunoblotting (right panel).
86 A
A5-1 A5-1 A5-1 A5-1 pTS-dnBRG-1 pTS-dnBRG-1 Dox : ++-- Dox : ++-- Cyclin A Cyclin A BRG1-FLAG BRG1-FLAG PSM-RB
β-actin Tubulin
1234 12 3 4
Figure II-7: RB-mediated repression of the cyclin A promoter involves chromatin remodeling. A) Parental A5-1 cells (lanes 1 and 2) or A5-1 cells engineered to inducibly co-express dnBRG-1 and PSM-RB (lanes 3 and 4) were cultured in the presence (lanes 1 and 3) or absence (lanes 2 and 4) of Dox for 24 hours. Cells were harvested and protein and RNA levels were determined by RT-
PCR (left panel) or immunoblotting (right panel) as indicated.
87 naked A5-1 B A5-1 rat cyclin A locus genomic DNA pTS-dnBRG-1 EagI : Dox : l + - ++-- na io t e rip sit I I sc t I n r oR EagI uncut pn a ta ag c K tr s E E -713 +28 +914
-135 +33 EagI cut probe 12 3 456
Figure II-7: RB-mediated repression of the cyclin A promoter involves chromatin remodeling. B) Left panel: Genomic structure of the rat cyclin A locus.
Right panel: Southern blot analysis of genomic DNA isolated from A5-1 cells (lanes
1 and 2). Parental A5-1 cells (lanes 3 and 4) or A5-1 cells engineered to inducibly
express dnBRG-1 (lanes 5 and 6) were cultured in the presence or absence of Dox
as indicated. Permeabilized nuclei were subjected to digestion with Eag1, and then
isolated genomic DNA was subjected to cleavage with Kpn1 and EcoR1.
Restriction fragments were detected by radioactive Southern blot.
88
Chapter III:
Hierarchical Requirement of SWI/SNF in RB-mediated Repression of Plk1
ABSTRACT
Polo like kinase 1 (Plk1) is a critical regulator of cell cycle progression that harbors oncogenic activity and exhibits aberrant expression in multiple tumors.
However, the mechanism through which Plk1 expression is regulated has not been extensively studied. Here we demonstrate that Plk1 is a target of the retinoblastoma tumor suppressor (RB) pathway. Activation of RB and related pocket proteins p107/p130 mediate attenuation of Plk1. Conversely, RB loss deregulates the control of Plk1 expression. RB-pathway activation resulted in the repression of Plk1 promoter activity, and this action was dependent on the
SWI/SNF chromatin remodeling complex. Although SWI/SNF subunits are lost during tumorigenesis and cooperate with RB for transcriptional repression, the mechanism through which SWI/SNF impinges on RB action is unresolved.
Therefore, we delineated the requirement of SWI/SNF for three critical facets of
Plk1 promoter regulation: transcription factor binding, co-repressor binding, and histone modification. We find that E2F4 and pocket protein association with the
Plk1 promoter is independent of SWI/SNF. However, these analyses revealed that SWI/SNF is required for histone deacetylation of the Plk1 promoter. The importance of SWI/SNF-dependent histone deacetylation of the Plk1 promoter
89 was evident, as blockade of this event restored Plk1 expression in the presence of active RB. In summary, these data demonstrate that Plk1 is a target of the RB pathway. Moreover, these findings demonstrate a hierarchical role for SWI/SNF in the control of promoter activity through histone modification.
90 INTRODUCTION
Progression through the cell cycle is a carefully choreographed process that
is often deregulated in cancer cells (33-35). It is believed that deregulation of
proliferation control serves to fuel tumor development and progression.
Interplay between cell cycle regulatory proteins is increasingly relevant for
understanding the underlying basis of appropriate cell cycle control and the
development of aberrant proliferation in cancer. Here we demonstrate a novel
regulation of the mitotic polo like kinase 1 (Plk1) by the retinoblastoma tumor
suppressor (RB) pathway.
Initially identified through homology with the Drosophila polo, Plk1 governs multiple events associated with G2/M progression (7, 8, 19, 28). For example,
Plk1 is a determinant of mitosis promoting factor, CDK1/cyclin B, which
stimulates entry into mitosis (19, 45). Additionally, centrosome duplication and
assembly of the mitotic spindle apparatus are regulated through the action of
Plk1 (17, 19). Recently, a number of studies have implicated a role for Plk1 in
cancer. For example, numerous tumors types (e.g. colorectal cancer, squamous
cell carcinomas of the head and neck and melanoma) aberrantly express Plk1
(13, 15, 16, 43). Consistent with a causative role in tumorigenesis, ectopic expression of Plk1 can transform cells in culture (37). Conversely, Plk1 ablation in tumor cells results in mitotic failure and cell death (5, 17, 20, 38).
Plk1 expression is repressed in resting cells and induced only as cells
progress through G1/S (18, 50). The induction of Plk1 protein levels is largely dependent on the regulation Plk1 promoter activity, and cis-acting elements that
91 modulate cell cycle dependence have been defined (18, 47). Particularly, the
Plk1 promoter is subject to transcriptional repression through cell cycle-
dependent element (CDE) / cell cycle-gene homology region (CHR) elements
that are required for the cell cycle dependence of Plk1 promoter activity (47, 55).
However, the mechanism through which silencing of Plk1 expression occurs as cells exit the cell cycle and the source of the deregulated expression in tumor
cells is unknown.
A critical regulator of G1/S-dependent gene expression is the RB-pathway.
RB is functionally inactivated in the majority of tumors via a number of discrete
mechanisms (26, 33-35). It is believed that RB, in concert with the related pocket
proteins p107 and p130, functions as a tumor suppressor through its capacity to
repress the transcription of critical targets in a cell cycle-dependent manner and
thus prevent proliferation (4, 26). In G0 or early G1 cells, RB and related
proteins are hypophosphorylated and form complexes with the E2F family of
transcription factors (11, 12, 27). The E2F family of transcription factors is
involved in the regulation of numerous genes required for cell cycle progression.
RB, p107 and p130 mediate transcriptional repression and subsequent
attenuation of E2F-regulated genes by recruiting additional co-repressors (e.g.
histone deacetylases) that modify chromatin structure (11, 12, 24). Repression is
alleviated when RB-family members are phosphorylated by CDK-complexes in
mid G1, thus enabling progression through the cell cycle (23). Transcriptional
repression is viewed as requisite for RB function in tumor suppression based on
genetic and biochemical data (12, 54). Additionally, we and others have found
92 that loss of SWI/SNF chromatin remodeling factors compromises RB-mediated transcriptional repression (31, 41, 42, 53).
SWI/SNF is a heterogeneous multi-subunit chromatin remodeling complex
(25, 29). This complex utilizes the energy of ATP to remodel chromatin structure and contains either BRG1 or BRM as the central ATPase (25). The activity of the core ATPase subunit is required by the SWI/SNF complex to regulate gene transcription (6, 14, 29). Prior studies have demonstrated that the combined loss of BRG1 and BRM result in resistance to activation of the RB-pathway and aberrant cell cycle progression (31, 42). Additionally, loss of SWI/SNF activity is associated with a failure of RB to elicit transcriptional repression of specific targets (e.g. cyclin A). Based on studies in yeast, the loss of SWI/SNF could disrupt virtually any step associated with transcriptional repression (6, 29). For example, SWI/SNF could be required for the assembly of E2F proteins at promoter; for the retention of RB or related proteins at the promoter; or for subsequent modifications of the promoter. However, the mechanism through which SWI/SNF cooperates with RB for transcriptional repression is not understood.
In this study, we specifically focused on elucidating the regulation of Plk1 expression. We show that Plk1 is repressed via activation of the RB-pathway and that Plk1 expression is deregulated through targeted RB loss. Moreover, we demonstrate that the repression of Plk1 by RB is dependent on SWI/SNF activity.
Analysis of SWI/SNF function demonstrates that chromatin remodeling is not required for the association of E2F or RB family members at the Plk1 promoter.
93 In contrast, histone deacetylation of the Plk1 promoter was dependent on
SWI/SNF and critical for the observed transcriptional repression. Thus, this study
provides critical insight into the mechanism through which Plk1 transcription is
regulated and demonstrates the intricate relationship between SWI/SNF and
histone deacetylases during RB-mediated transcriptional repression.
MATERIALS AND METHODS
Cell Culture, Plasmids, Infections and Transfections: SW13, TSUPr-1,
U2OS and A5-1 cells were maintained in Dulbecco’s Modified Eagle’s Media
(DMEM), supplemented with 10% fetal bovine serum (FBS), 100 units of
0 pencillin-streptomycin, 2 mM L-glutamine at 37 C in 5% CO2. In addition, A5-1 cells were maintained in G418 (200 µg/ml), hygromycin B (200 µg/ml) and doxycycline (Dox, 1 µg/ml). Primary RbloxP/loxP mouse adult fibroblasts (MAFs) were isolated from RbloxP/loxP mice (48). Cells were propagated by routine sub-
culturing in DMEM containing 10% fetal calf serum (FCS) supplemented with
100 units/ml pencillin-streptomycin and 2 mM L-glutamine. Primary cultures were between passage 2 to 6. Plasmids encoding β-gal, p16ink4a, PSM-RBP,
and BRG1 have been previously described (40, 41, 47). The reporter constructs
3xE2F-Luc and Plk1-Luc have been previously described (40, 41, 47).
Adenoviruses encoding GFP and p16ink4a have been previously described (1).
The Cre-encoding adenovirus was obtained from the Iowa University vector core.
Infections were performed at a multiplicity of infection of 50 to 100 for
94 approximately 95-100% infection efficiency after 24 hours as determined by GFP
fluorescence.
Reporter assays, immunoblotting and RT-PCR: Immunoblotting was
performed using standard techniques. The following antibodies were used: RB
purified mouse anti-human (Mat No 554136, BD), p107 (sc-318, Santa Cruz), p130 (sc-317, Santz Cruz), Plk1 (sc 17783, Santa Cruz; 06-813, Upstate
Biotechnology), BRG1 (sc-17796, Santa Cruz), p16ink4a (sc-759, Santa Cruz),
E2F1 (gift from A. Yee), E2F2 (sc-9967, Santa Cruz), E2F4 (sc-1082, Santa
Cruz), Lamin B (sc-6217, Santa Cruz), Vimentin (gift from Dr. Wallace Ip). All immunoblots were repeated at least three times with independent samples.
Reporter assays were performed as described previously (36). The reporter assays were performed in triplicate and from three independent experiments. RT-
PCR was performed as described previously (36). The following primer pairs were utilized: human Plk1, 5’-CCA GAG GGA GAA GAT GTC CA-3’ and 5’-ATA
ACT CGG TTT CGC TGC AG-3’; human GAPDH, 5’-TGG AAA TCC CAT CAC
CAT CT-3’ and 5’-TTC ACA CCC ATG ACG AAC AT-3’; rat Plk1, 5’-TTT GTG
TTC GTG GTT TTG GA-3’ and 5’-TTC TTC CGT TCC CCT TCA TA-3’. The rat
β-actin primers have been previously described (36). Experiments were performed at least three times and representative data is shown.
Chromatin Immunoprecipitation (ChIP) assay: Chromatin immunoprecipitation (ChIP) assays were performed as previously described
(36). The following antibodies were utilized for ChIP: p107 (sc-318; Santa Cruz), p130 (sc-317; Santa Cruz), E2F4 (sc-1082; Santa Cruz), acetylated histone H4
95 (06-866; Upstate Biotechnology) and Dbf4 (sc-11354; Santa Cruz). The following primers were used to amplify regions of the human Plk1 promoter: 5’-
GGTTTGGTTTCCCAGGCTAT-3’ and 5’-GCTGGGAACGTTACAAAAGC-3’.
ChIP assays were repeated at least twice with independent samples.
RESULTS
The RB-pathway regulates Plk1 expression: The Plk1 gene is subject
to deregulated expression in a variety of tumor types and has been shown to
harbor oncogenic activity(37, 52). Additionally, it has been previously shown that
Plk1 is a G1/S regulated gene (9, 47). However, the mechanisms through which
Plk1 expression is controlled or deregulated in human cancers are poorly
understood. In a micro-array screen we identified Plk1 as a target for RB-
mediated repression (22). Therefore, we initially determined the activity of the
RB pathway on Plk1 protein levels (Fig. III-1). To activate endogenous RB (and
the related proteins p107 and p130), U2OS cells were infected with adenoviruses
encoding either GFP (control) or p16ink4a which prevents their phosphorylation
by inhibiting CDK4 activity. As expected, the expression of p16ink4a resulted in
RB dephosphorylation (Fig. III-1A), indicating endogenous RB-pathway
activation. Plk1 protein levels were significantly attenuated in those cells infected
with p16ink4a encoding adenovirus, as compared to cells infected with the GFP
encoding virus (Fig. III-1A).
96 To determine if targeted RB loss, such as occurs in cancer, influences
Plk1 expression, conditional knockout of the Rb gene was employed. Mouse adult fibroblasts (MAFs) of RbloxP/loxP genotype were infected with recombinant adenoviruses encoding either GFP or Cre-recombinase. In this system, the endogenous Rb locus is subject to recombination through the expression of the
Cre-recombinase and RB protein expression is ablated (not shown). Under this condition, we find that Plk1 protein levels were elevated following RB loss (Fig.
III-1B, compare lanes 1 and 2). Similarly, by micro-array analysis we observe a
2.4-fold elevation in Plk1 RNA levels with RB loss (Markey et al., in preparation).
Thus, endogenous RB serves to maintain the appropriate levels of Plk1. To determine the subsequent action of endogenous RB on attenuating Plk1 protein levels under physiological stress, we examined the expression of Plk1 following exposure to camptothecin (CPT). CPT induces an RB-dependent checkpoint response, wherein specifically those cells deficient in RB (Cre-infected) continue cell cycle progression in the presence of CPT (Fig. III-1C). Consistent with protein analyses, MAFs deficient in RB showed up-regulation of Plk1 RNA levels compared to MAFs harboring RB (Fig. III-1D, compare lanes 1 and 3).
Treatment with CPT resulted in the repression of Plk1 RNA levels in RB proficient MAFs. (Fig. III-1D, compare lanes 1 and 2). However, Cre-mediated ablation of RB largely relieved the repression of Plk1 transcription following treatment with CPT (Fig. III-1D, compare lanes 2 and 4). These observations were further supported by analysis of Plk1 protein levels (Fig. III-1E, compare
97 lanes 2 and 4). Collectively, these results demonstrate that Plk1 is a target of the
RB-pathway, and specific RB loss results in deregulation of this critical target.
RB-mediated repression of Plk1 is compromised in SWI/SNF
deficient cells: Having established that Plk1 protein levels are modulated by the
RB-pathway, we next sought to elucidate the mechanism of this regulation.
Since SWI/SNF activity is compromised in specific cancers (30, 31) and known to be required for repression of selected RB target genes (e.g. cyclin A) (41, 53),
the action of SWI/SNF in Plk1 regulation was investigated. The SW13 cell line
does not express the BRG1 and BRM ATPases requisite for SWI/SNF activity,
whereas TSUPr-1 cells express BRM and are sensitive to RB-mediated signaling
(42). To activate the endogenous RB pathway, adenoviral transduction of
p16ink4a that maintains RB in its hypo-phosphorylated/active state was utilized.
As shown, expression of p16ink4a led to the dephosphorylation of RB and
related proteins p107 and p130 in both cell types (Fig. III-2A). To determine the
coordinate action of the RB pathway and SWI/SNF upon Plk1 expression, we
initially analyzed Plk1 promoter activity. SW13 cells and TSUPr-1 cells were co-
transfected with Plk1 reporter plasmid and either vector control or p16ink4a-
encoding plasmids (Fig. III-2B). p16ink4a expression potently repressed Plk1
promoter activity in TSUPr-1 cells. In contrast, p16ink4a failed to repress Plk1 in
the SW13 cell line. Consistent with the reporter assays, we observed attenuation
of endogenous RNA of Plk1 in TSUPr-1 cells infected with p16ink4a (Fig. III-2C,
compare lanes 3 and 4). As expected, expression of GAPDH did not change in
either of these cell lines even when infected with p16ink4a. In contrast, p16ink4a
98 failed to attenuate endogenous RNA levels of Plk1 in SW13 cells (Fig. III-2C, compare lanes 1 and 2). The SWI/SNF dependent reduction in promoter activity and RNA levels were reflected in the specific attenuation of Plk1 protein levels by p16ink4a infection in TSUPr-1 cells (Fig. III-2D). Taken together, these results indicate that SWI/SNF activity is critical for RB-pathway mediated repression of
Plk1. To confirm this observation, BRG1 expression was restored in SW13 cells in combination with p16ink4a, and Plk1 promoter activity was analyzed through reporter analysis (Fig. III-2E). Similar to our earlier results, p16ink4a alone failed to repress Plk1 promoter activity (Fig. III-2E). However, co-transfection of BRG1 and p16ink4a significantly repressed Plk1 promoter activity in the BRG1/BRM- deficient SW13 cells. Collectively, these data demonstrate that SWI/SNF activity is required for RB-mediated repression of the Plk1 promoter and attenuation of its RNA and protein levels.
SWI/SNF is dispensable for the assembly of E2F or pocket proteins at the Plk1 promoter: Failure of the RB pathway to repress Plk1 in the absence of SWI/SNF activity could be attributed to multiple functions of this complex. In yeast, it has been shown that SWI/SNF activity is required for the assembly of transcription factor complexes on chromatin (3, 6). Therefore, SWI/SNF loss could bypass RB-mediated repression through two basic mechanisms. First,
SWI/SNF could be required for E2F action. Specifically, SWI/SNF could be required for the expression of E2F family members or for E2F-chromatin interactions that are required for RB-pathway mediated repression (51). Second,
99 SWI/SNF could be required for pocket proteins to associate with E2F-factors on target promoters.
Initially, we probed the action of SWI/SNF directly on E2F proteins. The
E2F-family is broadly defined as activating (E2F1-3) or repressive (E2F4-5) E2Fs based on their predominant transcriptional role (46). Immunoblot analyses demonstrated that both SW13 and TSUPr-1 cells expressed similar levels of
E2F1 (‘activating E2F’) and E2F4 proteins (‘repressive E2F’) (Fig. III-3A). This result is consistent with microarray analyses that did not identify E2F-family members as targets of SWI/SNF (29) and suggests that limitation of E2F expression does not underlie the resistance to RB-pathway activation in SW13 cells. Therefore, several approaches were utilized to subsequently delineate the role of SWI/SNF on the functional interaction of E2F with chromatin. Initially, we determined whether E2F proteins were compromised for transcriptional activation, as the RB-pathway is compromised for transcriptional repression. To perform these analyses we used a synthetic promoter composed of multimerized
E2F sites (3xE2F-Luc), wherein promoter activity is dependent on E2F binding for activity. Endogenous E2F activity was readily detected with this reporter (Fig.
III-3B). Additionally, ectopic expression of E2F2 clearly activated the reporter (3- fold) in the absence of SWI/SNF (Fig. III-3B). These results suggest that E2F binding to a simple promoter element and stimulation of transcription is independent of SWI/SNF action.
100 Since transcription factors interact with chromatin in a dynamic fashion,
we assessed the action of SWI/SNF on E2F2 retention in living cells. In the case
of E2F-proteins, we have previously shown that the diffusion rate of these
proteins in living cells is dependent on chromatin association (2). Here we
constructed an expression vector encoding enhanced green fluorescent protein
(GFP) fused to the N-terminus of human E2F2 (GFP-E2F2). Expression of the
GFP-E2F2 fusion protein was verified by immunoblotting (Fig. III-3C, Middle
Panel). Additionally, the GFP-E2F2 construct efficiently stimulated transcription from the 3xE2F-Luc reporter (not shown). Having validated the functional activity of GFP-E2F2, it was utilized in fluorescence recovery after photobleaching
(FRAP) analysis to determine the influence of SWI/SNF on the nuclear retention of E2F2. SW13 and TSUPr-1 cells were transfected with GFP-E2F2 expression plasmids and FRAP analysis was performed 18 hours post-transfection. Under these conditions, we did not observe a significant difference in the mobility of
GFP-E2F2 between the two cells lines (data not shown). To specifically address whether SWI/SNF influenced the mobility of E2F2 in SW13 cells, we co- transfected these cells with GFP-E2F2 and either vector or BRG1. We failed to observe any difference in fluorescence recovery in the presence or absence of
BRG1 (Fig. III-3D). As expected, freely diffusible GFP rapidly recovered fluorescence after photobleaching, whereas GFP-Histone H2B failed to recover due to its tight association with chromatin. Taken together, these findings demonstrate that SWI/SNF does not globally affect E2F2 chromatin retention as assessed by live cell imaging.
101
To specifically determine if SWI/SNF is required for the association of an
E2F family member with the Plk1 promoter, chromatin immunoprecipitation
(ChIP) assays were performed. SW13 and TSUPr-1 cells were infected with p16ink4a encoding adenovirus, protein-DNA complexes were crosslinked with formaldehyde and immunoprecipitated with antibodies to E2F4 and Dbf4 (non- specific antibody control). Immunoprecipitated DNA was purified and utilized in quantitative radioactive PCR using primers flanking the E2F-binding sites present in the Plk1 promoter. Amplification of the DNA was well within the linear range of
PCR (data not shown). E2F4 occupancy was observed in both TSUPr-1 and
SW13 cells (Fig. III-3E, compare lanes 1 and 2). Together, these results demonstrate that SWI/SNF is not required for the assembly of E2F4 on promoters.
In addition to regulating transcription factor association, SWI/SNF could regulate the ability of pocket proteins to assemble at promoters. As shown in Fig.
III-2A, RB and the related proteins p107 and p130 are expressed in TSUPr-1 and
SW13 cells, and these proteins were dephosphorylated following the expression of p16ink4a. Next, we determined if RB had the capacity to functionally interact with E2F proteins in the absence of SWI/SNF. It is known that RB binding to E2F proteins will inhibit their transactivation function. As such, the 3xE2F-Luc reporter construct was utilized to determine if RB has the capacity to physically interact with E2F family members on a promoter. In SW13 cells the expression of a constitutively active allele of RB (PSM-RB) inhibited virtually all activation of the
102 reporter (Fig. III-4A). This result indicates that RB retains the capacity to
efficiently interact with E2F in the absence of SWI/SNF activity.
To determine the requirement for SWI/SNF in the assembly of pocket
proteins on the Plk1 promoter, ChIP analysis was performed (44, 49). We
observed approximately equal recruitment of p130 to promoters in the presence or absence of SWI/SNF (Fig. III-4B, lanes 1 and 2). In contrast, there was enhanced p107 recruitment in the SWI/SNF deficient cells (Fig. III-4B, lanes 1 and 2). Thus, SWI/SNF is not required for these pocket proteins to assemble on promoters. Similarly, we detected RB at the Plk1 promoter in the absence of
SWI/SNF (data not shown). However, in our hands the occupancy of RB at the
Plk1 promoter (as well as other promoters) is not consistently detectable.
Therefore, similar to the situation with E2F factors, BRG1/BRM was not required for promoter association of p107/p130.
SWI/SNF is required for Plk1 promoter histone deacetylation: It is
believed that one of the key components of RB-mediated transcriptional repression is not only its recruitment to chromatin, but also subsequent histone modifications at the promoter (10). Specifically, RB and related proteins recruit histone deacetylase activities that result in promoter hypoacetylation. We have recently shown that such modifications represent a critical means through which
RB functions to repress transcription (36). Therefore, we determined whether
SWI/SNF influences promoter histone acetylation. ChIP assays were performed using anti-acetyl histone H4 antibody to determine the acetylation status of histone H4 at the Plk1 promoter (Fig. III-5A). As shown, infection with
103 p16ink4a resulted in significant histone deacetylation of the Plk1 promoter in
TSUPr-1 cells (Fig. III-5A, upper left panel , compare lanes 3 and 4), which was evident from quantification of independent experiments (Fig. III-5A, upper right
panel). In contrast, p16ink4a infection failed to induce any histone deacetylation
of Plk1 promoter in SW13 cells, indicating that SWI/SNF is required for histone
deacetylation of the Plk1 promoter.
To determine the effect of histone deacetylation on Plk1 expression, we
utilized an immortalized rat fibroblast cell line that expresses PSM-RB in a tet-off
inducible fashion (A5-1). In this system, removal of the tetracycline analogue doxycycline (Dox) results in the induction of PSM-RB (Fig. III-5B). Under these conditions, Plk1 protein levels were efficiently down-regulated (Fig. III-5B). These results were confirmed at the level of endogenous RNA (Fig. III-5C, lanes 1 and
2). To investigate the coordinate action of SWI/SNF and histone deacetylation in this system, two approaches were employed. First, an inducible cell line (A5-1 pTS-dnBRG1) that expresses a dominant negative mutant of BRG1 was utilized to specifically determine the requirement for SWI/SNF during RB-mediated repression in this system. Removal of Dox from the media results in the coordinate induction of both PSM-RB and mutant BRG1 as we have previously reported (36). Under these conditions RB-mediated attenuation of Plk1 RNA levels were compromised (Fig. III-5B, lanes 3 and 4). Therefore, RB-mediated repression of Plk1 is dependent on SWI/SNF activity consistent with what was observed in SW13 cells. To address whether inhibition of HDAC activity specifically has the potential to block RB-mediated attenuation of Plk1, a
104 pharmacological inhibitor of deacetylase enzymes, trichostatin A (TSA), was
utilized. As shown, TSA significantly reversed the RB-mediated attenuation of
Plk1 RNA levels (Fig. III-5C, lanes 5 and 6). To directly investigate Plk1
promoter activity, the Plk1 reporter construct was integrated into A5-1 cells. In
this system, there was a 10-fold reduction in Plk1 promoter activity in presence of
active RB (Fig. III-5D). Treatment with the histone deacetylase inhibitor TSA
partially alleviated RB-mediated repression of Plk1 (Fig. III-5D). Collectively,
these data demonstrate that SWI/SNF is required for the deacetylation of the
Plk1 promoter and that this event is critical for repression mediated by the RB- pathway. Our findings support a model wherein both SWI/SNF and histone deacetylation are required for RB-mediated repression of Plk1 (Fig. III-6).
Moreover, histone deacetyltion of Plk1 promoter requires SWI/SNF activity, thus placing SWI/SNF and histone deacetylation in a hierarchical order for the repression of Plk1 expression.
DISCUSSION
Plk1 plays critical roles in progression through the cell cycle. Specifically,
Plk1 is implicated in a variety of processes associated with mitotic progression.
These activities range from roles in centrosome duplication (which occurs at the
G1/S transition) to spindle-pole maturation required for a productive nuclear division(19, 28). As such, the regulation of Plk1 expression is tightly controlled and modification of Plk1 levels is associated with diverse effects on cell biology.
105 For example, loss of Plk1 is inconsistent with cellular viability and induces catastrophic events in mitosis (20). In contrast, ectopic expression of Plk1 is associated with cellular transformation and is derergulated in human cancers (37,
50, 52). It has been previously demonstrated that Plk1 expression is stimulated as cells progress toward the G1/S transition (9, 47). Here we show that this gene regulation is manifested through the activity of the RB-pathway. Since the RB- pathway is compromised in the majority of tumors, these analyses provide a likely mechanism for the deregulation of Plk1 expression observed in tumors.
The mechanism through which RB regulates critical down-stream target gene expression has been hypothesized to involve the recruitment of co- repressors (10, 11). Prior studies have demonstrated that RB can recruit a myriad of co-repressors to facilitate transcriptional repression (21, 41). Here we show that the activity of SWI/SNF is required for the attenuation of Plk1 with RB- pathway activation. Such a result is consistent with the requirement for SWI/SNF in the repression of several additional RB target genes including cyclin A, cdc2 and cyclin E (39, 53). These findings suggest that repression by RB may be generally dependent on SWI/SNF activity, underscoring the ability of SWI/SNF deficiency to render cells resistant to the acute cell cycle arrest elicited by activation of the RB-pathway. In the context of RB-mediated repression of the
Plk1 promoter, there are several possible requisite actions for SWI/SNF. First, the transcription factor responsible for recruiting RB and related proteins could fail to associate with its cognate response element in the absence of SWI/SNF activity. Such a phenomenon is observed in yeast, wherein the Gal4
106 transcription factor requires SWI/SNF activity to associate with chromatin in vitro
(3). Similarly, the recruitment of GCN5 containing complexes to specific promoters is dependent on SWI/SNF activity (6). In the case of the RB-pathway this transcription factor would be the E2F-family of transcription factors, and it is well known that disruption of E2F-chromatin association represents one means to bypass RB-mediated arrest (32, 54). However, we show here that SWI/SNF is not required for the basic retention of E2F proteins on chromatin, using both reporter assays and a live cell imaging approach. Additionally, we can readily detect E2F4 at the Plk1 promoter in the absence of SWI/SNF activity. Second,
RB and related proteins could fail to stably interact with E2F at the promoter in the absence of SWI/SNF. We addressed this possibility by delineating the binding of RB to E2F in a simple functional interaction study, which revealed that
RB does retain the capacity to interact with E2F proteins to inhibit transactivation.
Additionally, we could clearly detect the RB related proteins p107 and p130 at the Plk1 promoters by ChIP. Thus, assembly of potential repressor complexes at target promoters occurs independently of SWI/SNF. Lastly, SWI/SNF activity could in fact be required for chromatin modifications leading to repression. It has been previously established that RB-repressor complexes can utilize histone deacetylation as a means to facilitate transcriptional repression (36, 53).
Analyses of the Plk1 promoter clearly demonstrated histone deacetylation during repression. However, this deacetylation was dependent on SWI/SNF activity.
In the case of the RB family of proteins, histone deacetylation plays n important role in transcriptional repression of specific genes. For example, the Cyclin E,
107 TopoIIα, TS, and Cdc2 genes are repressed via the RB pathway in an HDAC- dependent manner (36, 53). Our results suggest that the requisite action of
SWI/SNF in the repression of these genes could be solely through the control of the histone deacetylation of these promoters. Such a possibility is demonstrated in the case of Plk1 where the inhibitor of histone deaceytlation, TSA, significantly reversed RB-mediated repression.
In conclusion, these studies delineate a critical mechanism through which
Plk1 is transcriptionally regulated. Specifically, the RB-pathway is responsible for repression of the Plk1 promoter. This repression is dependent on SWI/SNF functioning in a hierarchical manner to control histone deacetylation of the Plk1 promoter.
ACKNOWLEDGMENTS
The authors thank Dr. Karen Knudsen and all members of both Knudsen laboratories for thought-provoking discussion and assistance with manuscript preparation. Additionally, we were assisted by the generous contributions of reagents from: Dr. Peter Stambrook, Dr. Anthony Imbalzano, Dr. Igor Roninson and Dr. Takeshi Uchiumi. ESK is supported by grants from National Cancer
Institute (CA106471); CNM is supported by NCI training grant T32CA59268.
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118 a k4 e in r A P 6 B C F 1 P P G p F F -G -G d d ppRB A A pRB Plk1 Plk1 β-tubulin Lamin B 1 2 1 2
RbloxP/loxP MAF C 120
100
80
60
40 BrdU PositiveBrdU
(% untreated control) untreated (% 20
0 Mock CPT Mock CPT Ad-GFP Ad-GFP Cre
Figure III-1: RB pathway represses Plk1 expression (A) U2OS cells were infected with either GFP (lane 1) or p16ink4a (lane 2) encoding adenoviruses.
Cells were harvested 36 hours post-infection and total protein was resolved by
SDS-PAGE. The indicated proteins were detected by immunoblotting. (B)
RbloxP/loxP MAFs were infected with GFP or Cre encoding adenoviruses. Cells
were harvested 6-days post-infection, and immunoblotted for Plk1 and β-tubulin.
(C) Ad-GFP and Ad-Cre infected RbloxP/loxP MAFs were treated with 10 µM camptothecin for 16 hrs 6-days post-infection and subsequently pulse labeled with BrdU for 12 hrs. The cells were fixed and the percent BrdU incorporation was determined. Untreated controls were set to 100%.
119 e DEre r C C FP FP FP FP G -G -G -G - d d d d A A A A - + - + : CPT - + - + : CPT Plk1 Plk1
GAPDH Vimentin 1 2 3 4 1 2 3 4
Figure III-1: RB pathway represses Plk1 expression (D) RbloxP/loxP MAFs treated with 5 µM camptothecin for 16 hrs 6-days post Ad-GFP and Ad-Cre infection were harvested, total RNA was isolated and RT-PCR was performed.
(E) Similar as in Fig 3C, except cells were harvested prior to BrdU addition. Total protein was isolated, resolved by SDS-PAGE and immunoblotted for Plk1 and
Vimentin.
120 A B
SW 13 TSUPr-1 Plk1- Luc a a 4 4 k k n n i i 100% P 6 P 6 1 F 1 F p p G G 75% Figure 2 ppRB pRB 50% pp107 p107
Relative luciferaseRelative activity 25%
pp130 p130 0% a r r 4 o a k t 4 to in c k c 6 e in e 1 v 6 v p 1 p16ink4a p
12 3 4 SW13 TSUPr-1
Figure III-2: SWI/SNF is required for RB-mediated repression of Plk1 (A)
SW13 (lanes 1 and 2) and TSUPr-1 (lanes 3 and 4) cells were infected with GFP
(lanes 1 and 3) or p16ink4a (lanes 2 and 4) encoding adenoviruses. Cells were
harvested 36 hrs post-infection, total protein was resolved by SDS-PAGE, and
the indicated proteins were detected by immunoblotting. (B) SW13 and TSUPr-1 cells were co-transfected with CMV β-gal and Plk1-Luc reporter plasmids and either vector or p16ink4a expression plasmids. Relative luciferase activity was
normalized to β-gal activity for transfection efficiency and vector control was set
to 100.
121 C SW13 TSUPr-1 D SW13 TSUPr-1
a a a a 4 4 4 4 k k k k n n n in i i i P P P P 6 6 6 6 F F F 1 F 1 1 G p G p1 G p G p Plk1 Plk1
GAPDH Lamin B 1234 12 3 4
E SW13 Plk1-Luc 125
100
75
50
25
Relative Luciferase Activity 0
r 1 a G to 4 c k R e in B V 6 + 1 a p 4 k in 6 1 p
Figure III-2: SWI/SNF is required for RB-mediated repression of Plk1 (C) SW13
(lanes 1 and 2) and TSUPr-1 (lanes 3 and 4) cells were infected with either GFP
(lanes 1 and 3) or p16ink4a (lanes 2 and 4) encoding adenoviruses. Total RNA was extracted 24 hrs post-infection and subjected to linear RT-PCR amplification with primers specific for the indicated genes. (D) SW13 (lanes 1 and 2) and TSUPr-1
(lanes 3 and 4) cells were infected with either GFP (lanes 1 and 3) or p16ink4a (lanes
2 and 4) encoding adenoviruses. Total protein was resolved by SDS-PAGE, and the indicated proteins were detected by immunoblotting. (E) SW13 cells were co- transfected with CMV β-gal and Plk1-Luc reporter plasmids and either vector,
122 p16ink4a, or p16ink4a and BRG1 expression plasmids. Relative luciferase activity was normalized to β-gal activity for transfection efficiency and vector control was set to
100.
123 AB 3xE2F-Luc 4 SW 13 TSUPr-1
a 4 k a n 4 i 3 k 6 n i P 1 P p 6 F F 1 G G p E2F1 2
E2F4 1 Relative Luciferase Activity Relative 12 34 0 2 tor c 2F e E V
Figure III-3: SWI/SNF is dispensable for E2F chromatin/promoter association (A)
SW13 (lanes 1 and 2) and TSUPr-1 (lanes 3 and 4) cells were infected with either
GFP (lanes 1 and 3) or p16ink4a (lanes 2 and 4) encoding adenoviruses. Total protein was resolved by SDS-PAGE and the indicated proteins were detected by immunoblotting. (B) SW13 cells were transfected with CMV β-gal, the reporter construct 3XE2F-Luc, and either vector or E2F2 as indicated. Relative luciferase activity was normalized to β-gal activity for transfection efficiency and vector control was set to 1.
124 CD 1.0 P GF GFP-E2F2 0.8 anti-GFP free GFP GFP-E2F2 + vector 0.6 GFP-E2F2 + Brg1 anti-E2F2 GFP-Histone H2B 0.4 12
0.2
0.0 0123456 Plk1 Promoter Time (sec) SW13 TSUPr-1 E Inputs
DBF4
E2F4
12
Figure III-3: SWI/SNF is dispensable for E2F chromatin/promoter association (C)
U2OS cells were transiently transfected with free GFP alone or GFP-E2F2. Total protein was resolved by SDS-PAGE and GFP-E2F2 protein was detected with antibodies specific for GFP and E2F2. (D) SW13 cells on 25-mm cover slips were co- transfected with GFP, GFP-Histone H2B or GFP-E2F2 and either empty vector or wild type BRG1. Coverslips were transferred to live cell imaging chambers and nuclear
FRAP analysis was performed 18 hrs post transfection. (E) SW13 (lanes 1) and
TSUPr-1 (lanes 2) cells were infected with p16ink4a encoding adenoviruses. ChIP assays were performed with antibodies for E2F4 and Dbf4 (non specific). Input and
125 immunoprecipitated DNA was amplified by PCR with primers specific for the Plk1 promoter.
126 A B
3xE2F-Luc 1.5 Plk1 Promoter SW13 TSUPr-1 Input 1 DBF4
p107 0.5 p130 Relative LuciferaseRelative Activity
0 12 r B 2 to R F c - 2 e M E V S + P B -R M S P
Figure III-4: SWI/SNF is not required for pocket protein association with the
Plk1 promoter: (A) SW13 cells were transfected with CMV β-gal and 3XE2F-Luc reporter plasmids and either vector, PSM-RB, or PSM-RB and E2F2 expression plasmids as indicated. Relative luciferase activity was normalized to β-gal activity for transfection efficiency. (B) SW13 (lanes 1) and TSUPr-1 (lanes 2) cells were infected with p16ink4a encoding adenoviruses. ChIP assays were performed with antibodies for p107 and p130. Input and immunoprecipitated
DNA was was amplified by PCR using primers specific for the Plk1 promoter.
127 A SW13 TSUPr-1 Relative Ac H4 Binding a a 4 4 k k k in 100.0 n k c i c 6 o 6 o 1 1 p M p M 75.0 Input 50.0
DBF4 25.0 Relative % Input
0.0 k a k a c 4 c 4 o k o k M in M in Ac H4 6 6 1 1 p p 12 34 SW13 TSUPr-1
Figure III-5: SWI/SNF is required for histone deacetylase-mediated repression of the Plk1 promoter (A) Left Panel SW13 (lanes 1 and 2) and
TSUPr-1 (lanes 3 and 4) cells were either mock infected (lanes 1 and 3) or infected with p16ink4a (lanes 2 and 4) encoding adenoviruses. ChIP assays were performed with antibodies for acetylated histone H4. Input and immunoprecipitated DNA was amplified by PCR using primers specific for the
Plk1 promoter. Right Panel Quantitative analyses of histone acetylation of the
Plk1 promoter from two independent experiments.
128 BC
+ :Dox A5-1 - A5-1 pTS-dnBRG1 A5-1 + TSA PSM-RB +- +- +-:Dox
Plk1 Plk1
Tubulin β-Actin 12 12 34 56
Plk1-Luc D 100
75
50
25 Relative Luciferase Activity 0 +Dox -Dox +Dox -Dox -TSA +TSA
Figure III-5: SWI/SNF is required for histone deacetylase-mediated repression of the Plk1 promoter (B) A5-1 cells were cultured in the presence
(lane 1) or absence (lane 2) of doxycycline for 24 hours. Total protein was resolved by SDS-PAGE and the indicated proteins were detected by immunoblotting. (C) A5-1 (lanes 1,2, 5 and 6) or A5-1 pTS-dnBRG1 (lanes 3 and 4) cells were cultured in the presence (lanes 1, 3 and 5) or absence (lanes 2,
4 and 6) of Dox and the addition of 100 nM TSA (lanes 5 and 6). Total RNA was isolated 24 hours post-treatment and the indicated RNA levels were determined
129 by RT-PCR analyses. (D) The Plk1-Luc reporter construct was stably integrated into A5-1 cells and three clones were selected. Cells were cultured with or without Dox in the presence or absence of TSA. Relative luciferase activity was determined from three independent experiments with the +Dox condition set to 100.
130 Pocket Protein SW I/SNF E2F DP Independent Plk1 Promoter Ac-Histone
SW I/SNF Pocket Protein SW I/SNF E2F DP X Dependent Plk1 Promoter Histone deacetylation
Figure III-6: Schematic diagram of transcriptional regulation of Plk1. Here we show that activation of the Plk1 promoter is regulated by E2F and pocket proteins. In SWI/SNF deficient cells, E2F4 and pocket proteins are recruited to the Plk1 promoter. Despite the presence of the pocket proteins, histones remain acetylated and the promoter retains activity. By contrast, in the presence of
SWI/SNF, the E2F4 and pocket protein recruitment results in histone deacetylation at the Plk1 promoter and subsequent promoter repression. In summary, our data support a model of hierarchy between SWI/SNF and histone deacetylation of the Plk1 promoter.
131
Chapter IV:
Loss of RB Compromises Specific Heterochromatin Modifications and Modulates HP1α Dynamics
ABSTRACT
Heterochromatin domains are important for gene silencing, centromere
organization and genomic stability. These genomic domains are marked with
specific histone modifications, heterochromatin protein 1 (HP1) binding and DNA
methylation. The retinoblastoma tumor suppressor, RB mediates transcriptional repression and functionally interacts with a number of factors that are involved in heterochromatin biology including HP1, SUV39H1, DNMT1 and components of the SWI/SNF chromatin remodeling complex. To analyze the specific influence of RB loss on chromatin modification, mouse adult fibroblasts derived from
RbloxP/loxP mice were utilized to acutely knock out RB. In this setting, target genes of RB are deregulated. Additionally, changes in histone modifications were observed. Specifically, histone H4 lysine 20 trimethylation was absent from heterochromatin domains following loss of RB and there were changes in the relative levels of histone modifications between RB-proficient and deficient cells.
While RB loss significantly altered the modifications associated with heterochromatin domains, these domains were readily identified and efficiently mediated the recruitment of HP1α. Kinetic analyses of HP1α within the
heterochromatin domains present in RB-deficient cells indicated that loss of RB
132 retarded HP1α dynamics, indicating that HP1α is paradoxically more tightly associated with heterochromatin in the absence of RB function. Combined, these analyses demonstrate that loss of RB has global effects on chromatin modifications and dynamics.
133 INTRODUCTION
The retinoblastoma tumor suppressor protein (RB) belongs to a family of
proteins collectively known as pocket proteins that includes RB, p107 and p130
(8). Although all members of the pocket protein family have the potential to block
cell cycle progression, RB is specifically targeted for inactivation in cancer.
Classically, RB is believed to function as a tumor suppressor through its ability to block cell cycle progression. In G0 and G1 phases of cell cycle, RB is hypophosphorylated and inhibits the transition into S phase of the cell cycle.
Phosphorylation of RB by CDK4/cyclin D1 and CDK2/cyclin E during mitogenic signal transduction leads to its inactivation thereby allowing cells to progress through S-phase of the cell cycle (19, 42). RB inhibits cellular proliferation by antagonizing E2F family of transcription factors (20, 41). The E2F family of transcription factors are important for transcription of genes that are involved in cell cycle progression, DNA synthesis and apoptosis (5, 9, 12, 13, 22, 23, 34-36,
40, 45, 47). Additionally, RB actively represses transcription of E2F regulated genes by assembling transcriptional repressor complexes comprising SWI/SNF and HDACs that alter chromatin structure so it is less permissive to transcription factors.
Studies of the tumor suppressive function of RB have traditionally focused on aspects of cell cycle control particularly the transcriptional repressive function of RB. To mediate transcription, RB utilizes several chromatin remodeling and modifying enzymes including SWI/SNF chromatin remodeling enzymes, histone deacetylases (HDACs) and histone methyltransferases (HMTase) (6, 10, 26, 32,
134 44). It has been demonstrated that SWI/SNF is required for RB-mediated cell cycle arrest (46, 48). In addition, HDACs are important for deacetylation of RB- responsive promoters leading to repression of a subset of RB target genes.
However, HDAC activity is dispensable for RB-mediated cell cycle arrest (43).
While methyltransferase activity has been shown to be required for RB-mediated repression of cyclin E and dihydrofolate reductase (DHFR) genes, RB does not appear to employ HMTase activity for repression of other target genes in cycling cells (1, 6, 31). Thus, covalent modifications, specifically histone deacetylation play an important role during RB-mediated transcriptional repression.
Covalent modifications that occur at histones have important implications not just for transcription but other cellular processes linked to tumorigenesis including gene silencing, senescence, differentiation, and DNA damage response. Specific histone modifications mark structural and functional DNA domains that are important for maintaining genomic stability. The eukaryotic genome is organized into two types of domains: euchromatin and heterochomatin. In general, euchromatic domains are gene-rich and accessible to DNA binding transcription factors. In contrast, heterochromatic domains are inaccessible to transcription factors and are composed of repetitive elements that normally do not contain genes. While large heterochromatic domains are found in the centromeric and telomeric regions of the chromosome, smaller domains are distributed throughout the genome. Originally thought to be ‘junk DNA’, heterochromatin is now known to possess important functions. Heterochromatin domains that surround the centromeric region are important for its function in
135 proper chromatin segregation (3, 18, 33, 37). Furthermore, heterochromatin inhibits recombination between homologous repeats thereby stabilizing repetitive
DNA sequences in the genome (15, 17). Although, the mechanisms underlying formation and propagation of heterochromatin are not fully understood, trimethylation of histone H3 lysine 9 (H3K9), trimethylation of histone H4 lysine
20 (H4K20) and recruitment of heterochromatin protein 1 (HP1) appear to be intimately involved (16, 39). Trimethylation of heterochromatic histone H3K9 is accomplished by Suv3-9h1 and Suv3-9h2 (38), while Suv4-20h1, Suv4-20h2 trimethylates histone H4K20 (39). RB physically interacts with both these methyltransferases although the biological significance of these interactions for
RB function remain unclear (14, 32).
Aberrations in chromatin structure and epigenetic programs are observed in a variety of human cancers. Traditionally, these influences on chromatin are viewed as promoter specific events affecting gene expression programs and the silencing of tumor suppressor genes. However, large-scale changes in chromatin structure/modification have recently been observed in cancer. For example, in a screen of primary tumors and tumor cell lines, it was reported that loss of acetylation of histone H4 lysine 16 and trimethylation of histone H4 lysine 20 is a salient feature of cancer (11). Critical mediators of chromatin structure can function as tumor suppressors in mouse models, and since RB can directly interact with such factors (SWI/SNF, HDACs, HMTases), we investigated the action of RB loss on chromatin structure. We find that loss of RB did not have discernable effect on heterochromatin structures as these structures were readily
136 identified following staining with DAPI. Similarly, RB loss did not impact
heterochromatin trimethylation of H3K9 and recruitment of heterochromatin protein 1α (HP1α) to heterochromatin domains. However, RB loss led to specific loss of trimethylated H4K20 from heterochromatin domains. Furthermore, kinetic studies revealed decreased HP1α mobility with RB loss. Our findings demonstrate a novel role of RB in regulating HP1α dynamics and H4K20 trimethylation in heterochromatin domains.
MATERIALS & METHODS
Cell culture, plasmids and trasfections: Primary RbloxP/loxP mouse adult fibroblasts (MAFs) were cultured in Dulbecco’s Modified Eagle’s Medium
(DMEM) supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin-
0 streptomycin and 2 mM L-glutamine at 37 C and 5% CO2. MAFs were obtained from RbloxP/loxP genotype mice as previously described (29). MAFs from passages
3 to 5 were used in this study. Replication-defective recombinant adenovirus
expressing green fluorescent protein alone (Ad-GFP) or GFP and Cre recombinase (Ad-GFP-Cre) were generous gifts from G Leone (Ohio State
University). Conditional knockout of RB in Rb-floxed primary MAFs was achieved by infecting the cells with adenoviral GFP-Cre at an MOI of approximately 20.
Infection efficiency was determined to be 90-95% through GFP fluorescence.
Cells were cultured at least 6 days post-infection before use in this study.
Plasmids expressing recombinant GFP-H10, H2B-GFP and GFP-HP1α were generous gifts from David Brown, National Cancer Institute; Geoff Wahl,
137 Salk Institute and D Kirschmann, University of Iowa respectively. Thomas
Jenuwein (Institute of Molecular Pathology, Vienna, Austria) and Susanne Wells
(Cincinnati Children’s Hospital Medical Center) kindly provided Suv39h double
null and TKO cells respectively.
Immunofluorescence: MAFs previously infected with GFP or GFP-Cre
were seeded on coverslips in 6-cm dishes. Following attachment to the
coverslips, cells were fixed with methanol for 15 min at room temperature. The
following primary antibodies were used: histone H3 (05-499; Upstate), acetyl histone H3K9 (06-942; Upstate), trimethyl histone H3K9 (ab8898-100; Abcam), trimethyl histone H4K20 (ab9053; Abcam) and acetyl histone H4 (06-866;
Upstate).
Fluorescence Recovery After Photo-bleaching (FRAP): FRAP experiments were performed as previously described with few modifications (2).
For FRAP analysis, MAFs were seeded onto 25mm cover glass (Fisher
Scientific) and transfections were performed using FuGENE (Roche). FRAP experiments were performed 48 hour post transfection by transferring the cover glass into a live-cell imaging chamber (Atto) set in a water-jacketed stage incubator at 370C. FRAP was performed on a Zeiss LSM510 laser scanning
confocal unit attached to a Zeiss Axiovert inverted microscope equipped with a
C-Apochromat 63X 1.4 NA objective. Photo-bleaching was achieved by 100%
transmission of 488-nm light from an argon laser running at 6.3 mW. Fluorescent
intensity of the bleached area and a distal unbleached area of the nucleus of
equal size were measured every 100 ms for indicated lengths of time after photo-
138 bleaching. These values were compared to each other to produce a relative
fluorescent intensity, which was then normalized to pre-bleached intensity
values. Data represented here were collected from at least 12 nuclei per
condition from multiple independent experiments.
Immunoblotting: Immunoblotting was performed as previously described
(43). Antibodies against the following proteins were used: cyclin A (sc-751; Santa
Cruz), MCM7 (sc-9966; Santa Cruz), PCNA (sc-56; Santa Cruz), Plk1 (sc-17783;
Santa Cruz), thymidylate synthase (TS; gift from Masakazu Fukushima) and
vimentin (gift from Wally Ip).
Polymerase Chain Reaction (PCR): Cre-mediated recombination of
RbLoxP/LoxP MAFs was verified by PCR. Genomic DNA was isolated using DNeasy
Tissue Kit (Qiagen, Valencia, CA) following manufacturer’s protocol. The following PCR primers were used Rb212 (5’-
GAAAGGAAAGTCAGGGACATTGGG-3’) and Rb18 (5’-
GGCGTGTGCCATCAATG-3’) (28).Thermal cycling conditions consisted of 30 cycles of 30 sec at 950C, 30 sec at 550C and 30 sec at 720C.
Chromatin Immunoprecipitation (ChIP): ChIP assays were performed as previously described (43). The following primer pairs were used: topoIIα, 5’-
CTGCAAACAGAGCCAATGAG-3’ and 5’-ATGGTGACGGTCCTGAAGAG-3’;
RNRII, 5’-CGCAGGCTCCTTAAAGGTC-3’ and 5’-AGGCGAAAACAGCTGGAAT-
3’.
139 RESULTS
Conditional loss of RB results in downstream target deregulation: To
investigate the role of RB in regulating chromatin structure, a conditional
knockout of RB using the Cre-Lox system was utilized. Mice of RbloxP/loxP genotype have been previously described (28). Mouse adult fibroblasts (MAFs) were isolated from RbloxP/loxP mice and cells were transduced with adenoviruses
encoding either GFP-tagged Cre-recombinase (Ad-GFP-Cre) or GFP (Ad-GFP) as a control. Initially, the efficiency of recombination was verified by genomic
PCR. As shown in figure IV-1A, cells infected with Ad-GFP yield a 746 bp PCR product, while cells infected with Ad-GFP-Cre reveal a recombined product of
260 bp. Under these conditions the RB protein is absent and the cells are
functionally deficient for RB based on a variety of functional assays (4, 29). To
determine the influence of RB loss on transcriptional targets, we analyzed the
protein levels of cyclin A, minichromosome maintenance protein 7 (MCM 7),
proliferating cell nuclear antigen (PCNA), polo-like kinase 1 (Plk1) and
thymidylate synthase (TS) (Fig. IV-1B). The levels of each of these proteins
were enhanced with the loss of RB, indicating that RB-mediated transcriptional
repression has been relieved.
Acute RB loss results in specific loss of heterochromatin associated
histone modifications: RB is known to functionally interact with many factors that are important for heterochromatin assembly. Therefore, the influence of RB loss on heterochromatin structure was investigated. In murine fibroblasts, distinct peri-centromeric heterochromatin is readily identified following non-specific DNA
140 staining. As shown in figure IV-2A, loss of RB did not have a measurable effect on the size and appearance of heterochromatin domains. To specifically investigate the molecular effects of RB loss on heterochromatin domains, immunfluorescence microscopy was performed using antibodies that detect specific histone modifications. Initially, modifications that are enriched within heterochromatin domains were investigated. Analyses of H3K9 trimethylation demonstrated that, irrespective of the presence of RB, the majority of H3K9 trimethylation was confined within the heterochromatin domains (Fig. IV-2A).
However, from the analyses of multiple samples, there was an overall trend toward reduced intensity for H3K9 trimethyl staining throughout the nuclei of RB deficient cells. This reduction in trimethyl H3K9 intensity with RB loss was evident from quantitative analysis using MetaMorph® image analysis software
(Fig. IV- 2B).
To further investigate the regulation of this modification, cells deficient in the predominant mediators of H3K9 trimethylation, Suv39h1 and Suv39h2, were analyzed (Fig. IV-2C). In contrast with the RB-deficient cells, the Suv39h-/- cells revealed virtually no detectable trimethyl H3K9 staining. Thus, although RB
physically interacts with Suv39h1 and Suv39h2, the loss of RB is not
synonymous with the loss of the Suv39h1 and Suv39h2 histone methyl
transferases (HMTases) (Fig. IV-2C). Since the RB-related proteins, p107 and
p130 can interact with many of the same chromatin modifying enzymes that are
engaged by RB, the net effect of deleting all pocket proteins was assessed using
fibroblasts harboring the combined knockout of RB, p107 and p130 (TKO cells).
141 In these cells, like those deficient for RB above, H3K9 trimethylation was readily
observed in heterochromatin domains (Fig. IV-2C). Thus, both the complete loss
of pocket proteins and loss of RB exert similar effects on H3K9 methylation.
Next, the localization and pattern of histone H4K20 trimethylation was
analyzed. Focal enrichment of this modification was readily observed in regions
of heterochromatin in MAFs infected with adenoviral GFP (Fig. IV-2D). In
contrast, cells infected with Ad-GFP-Cre showed a deregulated pattern of histone
H4K20 trimethylation, such that the heterochromatin domains were largely devoid of this modification (Fig. IV-2D). A quantitative analysis of histone H4K20
trimethylation pattern showed that the GFP-Cre infected cells exhibited a general
reduction in H4K20 trimethylation compared to cells infected with GFP alone
(Fig. IV-2E). Together, these data demonstrate that acute loss of RB alone is
sufficient to destabilize the trimethylation pattern of histone H4K20 in constitutive
heterochromatin. Analyses of Suv39h-/- cells and TKO cells demonstrated a
striking lack of H4K20 reactivity within heterochromatin domains (Fig. IV-2F).
Thus, there is a significant distinction between the relative contribution of RB and
other pocket proteins or the Suv39h1 and Suv39h2 HMTases in regard to H4K20
trimethylation.
Acute loss of RB does not relocate markers of euchromatin into
heterochromatin domains: These combined findings led us to question whether
the diminution of markers of heterochromatin (i.e. H4K20 trimethylation) would be
associated with increased histone acetylation within the heterochromatin
domains present in RB-deficient cells. We found that acetylation of H3K9 was
142 excluded from heterochromatin regions in GFP infected cells, and this
localization was similarly maintained within the context of GFP-Cre infected cells
(Fig. IV-3A). Thus, the acute loss of RB does not facilitate the extensive histone
acetylation of heterochromatin domains. A quantitative analysis revealed an
increase in intensity of acetylated H3K9 in GFP-Cre infected cells (Fig IV-3B).
Similar results on global acetylation with histone H4 were also observed (Fig. IV-
3C and 3D). These findings are consistent with the increased transcriptional
activity of these cells as a result of RB loss, since acetylation of histones is a
marker for active transcription. Consistent with this idea, analyses of specific
promoters of genes that are upregulated with the loss of RB by ChIP analyses
demonstrated an increase in histone H4 acetylation (Fig. IV-3E). Therefore, RB
loss resulted in increased histone acetylation in euchromatic regions of the
nucleus and in the promoters of specific RB regulated genes but did not result in
global redistribution of euchromatic markers into heterochromatin domains.
Acute RB loss alters HP1α dynamics: In replicating cells, the nucleosome is assembled by first wrapping the DNA around H3-H4 tetramer and
then the addition of a H2A-H2B dimer. Although quite stable once assembled,
histone-DNA interactions within the nucleosome are continually altered in living
cells. In response to a variety of cellular processes, histones are exchanged
within the nuclesomes (24). Since histone modifications have the potential to
either compact or loosen the chromatin, we examined how RB-mediated histone modifications influenced histone and non-histone protein interaction with DNA.
Initially, the ability of heterochromatin protein 1α (HP1α) to localize to
143 heterochromatin domains was assessed in living cells. In these experiments, RB
proficient or deficient MAFs were transfected with GFP-HP1α expression plasmids. Initially, we monitored the localization of GFP-HP1α using high resolution confocal microscopy. Under these conditions, the GFP-HP1α was readily observed in heterochromatin domains of both RB proficient and RB
deficient cells (Fig. IV-4A). Therefore, the presence of RB does not qualitatively modulate the localization of HP1α. Similar localization was observed in the TKO cells. However, the Suv39h-/- cells failed to localize HP1α to heterochromatin
(Fig IV-4A). Thus, the recruitment of HP1α followed the tri-methylation pattern of
H3K9 and was not altered by pocket proteins (Fig IV-2).
The dynamic properties of chromatin can be monitored in live cells by investigating the dynamics of GFP-HP1α in chromatin domains. To achieve this, transfected cells were subjected to fluorescence recovery after photobleaching
(FRAP) analyses. In this approach, a discrete heterochromatin domain is bleached and the dynamic recovery of fluorescence into this bleached region is an indication of the stability of the interaction with the given chromatin associated factor. In the case of HP1α, exchange occurs more slowly when engaged in heterochromatin domains and this is a measure of the extent to which it is actively associated with chromatin. In RB-proficient MAFs, HP1α recovered at a t1/2 of ~2.5 s, consistent with previous findings (7). Surprisingly, the loss of RB
had a net effect of decreasing the mobility of HP1α at heterochromatin domains.
Thus, the loss of RB reduces the dynamic exchange of HP1α (Fig. IV-4B). A
possible explanation for this could be an overall change in the behavior of
144 nucleosome dynamics. To address this, the dynamics of histones H10 and H2B was determined. As shown in figure IV-4C, histone H2B readily associated with heterochromatin domains. However, in contrast with the behavior of HP1α, there
was no discernible effect of RB loss on the mobility of histone H2B (Fig IV-4C).
Similar results were also observed with H10 (data not shown). Thus, the loss of
RB has a specific effect on HP1α dynamics within heterochromatin domains, yet does not have a global influence on histone dynamics.
DISCUSSION
The proximal influence of RB on cell cycle is believed to be manifest through the control of gene expression. RB is known to interact with a variety of transcription factors and chromatin modifying proteins, which contribute to both heterochromatin and gene regulation. Here we utilized a conditional knockout model to assess the action of acute RB loss, as this approach alleviates the compensation that can be manifest by other pocket proteins (ie. p107 and p130).
As expected, the acute loss of RB resulted in increased levels of E2F-regulated genes. This is confirmed by microarray analyses which identified approximately
300 genes classically associated with the E2F pathway that are deregulated as a consequence of RB loss (in preparation). Additionally, the loss of RB in this model deregulates cell cycle control processes (4, 29). Thus, this model provides a means to delineate the effect of RB loss on chromatin. The principal effects of
RB on chromatin have been investigated in the context of gene promoters. In this
145 setting, RB function has the effect of inducing histone deacetylation and histone methylation. Correspondingly, loss of RB is associated with enhanced histone acetylation of target promoters, which we also observe. Therefore, RB loss lead to discrete chromatin modifications at specific promoter regions associated with the deregulation of gene expression.
Prior studies have suggested that RB could function not only in the context of promoters, but on global chromatin structure. This concept was first supported by data from Herrera et al., who found that loss of RB led to increased histone
H1 phosphorylation and relaxed chromatin structure (21). Subsequent studies
have indicated that RB plays an active role in senescence induced
heterochromatin foci in human cells (30). Therefore, we specifically probed the
influence of RB loss on the well-defined heterochromatin domains present in the
nuclei of murine fibroblasts. These domains are well characterized as largely
hypo-acetylated and enriched for trimethylated H3K9 and H4K20. Loss of RB
had little effect on multiple facets of these heterochromatin domains. First, H3K9
trimethylation was retained within heterochromatin domains with loss of RB.
Second, histone acetylation was not observed within these domains following
loss of RB. Third, the influence of RB loss on heterochromatin domains is not
absolute as heterochromatin structures are readily apparent by DAPI staining.
Moreover, HP1α recruitment to heterochromatin domains occurs even in the
absence of RB. In this regard, loss of RB is quite distinct from the loss of
Suv39h1 and Suv39h2 that preclude H3K9 trimethylation and the recruitment of
HP1 to heterochromatin domains. However, RB loss did influence the decoration
146 of the heterochromatin domains with H4K20 trimethylation. This effect was relatively specific for the heterochromatin domains as peripheral H4K20 trimethylation could be observed in the nucleoplasm of RB-deficient cells. This specific effect of RB loss contrasted both with the loss of Suv39h1, Suv39h2 and
RB/p107/p130, wherein virtually no H4K20 was detectable. Thus, the role of RB in this setting is specific for heterochromatin and suggests that p107 or p130 can compensate for some function of RB in directing H4K20 methylation, but not to heterochromatin domains. These data are consistent with a recently published report using TKO MEFs (14). However, while Gonzalo et. al., find loss of histone trimethylation of H4K20 in TKO MEFs, RB null MEFs have no apparent influence on heterochromatin. This difference is likely due to the nature of the knock-out
(chronic vs. acute), since compensation by RB family members could mask the loss of histone H4K20 trimethylation in RB null MEFs. However, in acute knockout RB MAFs H4K20 trimethylation disruption is detected due to a lack of compensation. Therefore, loss of H4K20 trimethylation and focal enrichment of this mark on heterochromatin domains is detected in acute RB knockout MAFs.
The mechanism through which RB mediates this change in H4K20 methylation is not apparent. Previous studies have demonstrated loss of H4K20 trimethylation in TKO MEFs through a mechanism discrete from E2F which mediates much of the RB-dependent gene regulation (14). In the case of RB- mediated transcriptional repression, RB recruits various corepressors including
HDACs, SWI/SNF and SUV3-9H1 to promoters of target genes (6, 10, 25, 26,
32). Thus, RB could be required to recruit Suv4-20h HMTases to sites of
147 heterochromatin assembly. Such a possibility is supported by the finding that
HP1α is appropriately enriched in heterochromatin domains, yet the dynamics of
HP1α are altered with RB loss. Prior studies have shown that HP1 is required for the loading of Suv4-20h2 HMTase on chromatin (39). Thus, the effect of RB on
HP1α could be responsible for the failure to mediate H4K20 trimethylation. This mechanism is appealing since it would suggest a specific effect of RB on HP1α function in heterochromatin, as opposed to a more ubiquitous role in regulating
Suv4-20h1/h2 function throughout the nucleus.
The functional effect of RB loss on heterochromatin domains is not intrinsically apparent. There are non coding RNA transcripts produced by peri- centromeric chromatin, which are believed to be important for targeting and propagating heterochromatin formation through pathways that involve the RNAi machinery (27). We did not observe alteration in the transcript levels of major or minor satellite repeats with acute RB loss (data not shown). Thus, the change in
H4K20 methylation is apparently not sufficient to mediate changes in transcription within heterochromatin. Since H4K20 trimethylation is associated with heterochromatin architecture, we postulated that loss of RB may influence the dynamics of proteins within heterochromatin domains. Therefore, we utilized
FRAP to monitor chromatin dynamics. Loss of RB did not have a profound
effect on histone dynamics. However, we observed reduced mobility of HP1α
with the loss of RB. This effect of RB is directly counter to the effect of HDAC
inhibitors or loss of Suv39h1 and Suv39h2, which have the net effect of
destabilizing HP1α with heterochromatin domains. How these changes in HP1α
148 and H4K20 methylation influence chromatin biology remain a subject of ongoing
study. Cells deficient in RB are prone to non-dysjunction effects which influence
ploidy in both cell culture and animal models. Thus, these modifications of
chromatin could compromise the efficiency of centromere function.
In summary, our findings demonstrate that RB regulates trimethylation of
H4K20 and HP1α-chromatin association in heterochromatin domains. Thus, although both Suv39h and Suv4-20h interact with RB, only Suv4-20h directed trimethylation of H4K20 is affected by RB loss. Similarly, RB loss does not impact
HP1α localization at heterochromatin domains but influences its dynamic association with heterochromatin. Thus, our data identify specific aspects of heterochromatin biology regulated by RB.
ACKNOWLEDGMENTS
We thank Emily Bosco, Christopher Mayhew and William Zagorski for critical reading of the manuscript. We are grateful to Nancy Kleene for technical assistance with confocal microscopy. We thank all members of the Knudsens’ and Reed laboratories for helpful discussions. This work is supported by grants to ESK from the National Cancer Institute (CA 104213-02). HS is supported by the Albert J. Ryan Foundation.
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158 AB e r e e C r r - C C P P - - F P P F P P F G G F F F - - G G G O d d -G - - - 2 d A A d d d H A A A A Cyclin A Plk-1 746 bp MCM7 TS 260 bp
PCNA Vimentin
Figure IV-1: Conditional loss of RB leads to target gene deregulation. A)
RbLoxP/LoxP mice were sacrificed, peritoneal fascia was isolated to culture
fibroblasts. Mouse adult fibroblasts (MAFs) were infected with adenovirus
expressing either GFP (Ad-GFP) or GFP-Cre (Ad-GFP-Cre). Cells were
harvested 6 days post-infection and DNA was isolated from these cells. Isolated
DNA was subjected to PCR amplification using primers specific for flanking
regions of exon 3 of Rb gene. B) MAFs infected with either Ad-GFP or Ad-GFP-
Cre were harvested 6 days post-infection. Total protein was isolated from these cells, resolved in SDS-PAGE and transferred into ImmobolinP membrane.
Immunoblotting was performed with the indicated antibodies. PCNA=proliferating
cell nuclear antigen; MCM7=mini-chromosome maintenance protein 7;
Plk1=polo-like kinase 1; TS=thymidylate synthase
159 A Trimethyl H3-K9 Hoechst Merged B Trimethyl Histone H3K9 Intens
30 Ad-GFP 25 20
15 Ad-GFP-Cre 10 Intensity (AU) 5
0 Ad-GFP Ad-GFP-Cre
C Trimethyl H3-K9 Hoechst Merged
Suv39h-/-
TKO
Figure IV-2: Acute RB loss results in specific loss of heterochromatin associated histone modifications. MAFs previously infected with either Ad-GFP or
Ad-GFP-Cre were seeded onto coverslips and fixed with methanol. Immunostaining with anti-trimethyl histone H3 lysine 9 (H3K9) and anti-trimethyl histone H4 lysine 20
(H4K20) antibodies were performed. To visualize the nuclei, cells were stained with
Hoechst. A) Ad-GFP and Ad-GFP-Cre infected MAFs immunostained with anti trimethyl histone H3K9. B) Analysis of intensity of trimethylated histone H3K9 using
MetaMorph®. Micrographs taken at identical exposure conditions were evaluated for intensity of the staining using MetaMorph®. C) Anti-trimethyl H3K9 staining of cells lacking Suv39h1/h2 (Suv39h-/-) and pocket proteins (TKO).
160 Trimethyl H4-K20 D Trimethyl H4-K20 Hoechst Merged E 120
100
Ad-GFP 80
60
40 Intensity (AU) Intensity Ad-GFP-Cre 20
0 Ad-GFP Ad-GFP-Cre F Trimethyl H4-K20 Hoechst Merged Suv39h-/-
TKO
Figure IV-2: Acute RB loss results in specific loss of heterochromatin associated histone modifications. D) Immunostaining of RbloxP/LoxP MAFs infected with Ad-GFP and Ad-GFP-Cre using anti-trimethyl histone H4 lysine 20 (H4K20) antibody. E) Quantitation of trimethyl H4K20 immunostaining in Ad-GFP and Ad-
GFP-Cre recombinase infected RbloxP/LoxP MAFs using MetaMorph® image analysis software. F) Anti-trimethyl H4K20 immunostaining of cells lacking Suv39h1/h2
(Suv39h-/-) and pocket proteins (TKO).
161 ABAcetylated H3 Hoechst Merged Acetylated H3 Int 40 35 Ad-GFP 30 25 20 15 10 Ad-GFP-Cre Intensity (AU) 5 0 Ad-GFP Ad-GFP-Cre C D Acetylated H4 Hoechst Merged Acetylated H4 100 90 Ad-GFP 80 70 60 50 40
Intensity (AU) 30 Ad-GFP-Cre 20 10 0 Ad GFP AD GFP C Ad-GFP Ad-GFP-Cre
E Ac H4 Dbf4 Input
e e e Cr Cr Cr - - - O P P P P P P 2 F F F F F F H G G G G G G TopoIIa
RNRII
12 3 4 5 6 7
Figure IV-3: Acute RB loss does not relocate euchromatin markers into heterochromatin domains. RbloxP/loxP MAFs were infected with either Ad-GFP or Ad-GFP-Cre. Cells were seeded onto cover slips, fixed with methanol, immunostained with antibodies as indicated. A) Micrographs of MAFs
162 immunostained with anti-acetyl histone H3 lysine 9 (H3K9). B) Quantitative analysis of anti-acetyl histone H3K9 staining using MetaMorph®. C) Images of
MAFs stained with anti-acetyl histone H4. D) Quantitation of immunofluorescence intensity of acetyl histone H4 using MetaMorph®. E) PCR analysis of the TopoIIα and RNRII promoters utilizing purified DNA from chromatin immunoprecipitation
(ChIP) assays performed with anti-acetyl histone H4 using formaldehyde cross- linked chromatin from Ad-GFP and Ad-GFP Cre infected RbloxP/loxP MAFs.
163 A Rb+/+ Rb-/- HP1-GFP
TKO Suv39h-/-
Figure IV-4: Acute RB loss modulates HP1α dynamics. MAFs previously infected
with either GFP or GFP-Cre were cultured such that GFP expression was lost. A)
Rb+/+, Rb-/-, pocket protein null (TKO), and Suv39h double null cells (Suv39h-/-) were
transfected with plasmid encoding GFP-HP1α. High resolution images were taken using confocal microscope.
164 B Pre-bleach Post-bleach 0''25'' 50'' GFP-HP1 Ad-GFP
0''25'' 50'' Ad-GFP-Cre
HP1 Mobility 1.2
1.0
0.8 Ad-GFP 0.6 Ad-GFP-Cre
0.4
Relative Fluorescence 0.2
0.0
0 6 6 7 7 7 7 7 .2 .0 .6 .4 .0 .8 .4 1.46 2 3 3.87 4 5 6.27 7 7 8.67 9 0.27 1 Time (s)
Figure IV-4: Acute RB loss modulates HP1α dynamics. B) FRAP analysis of
HP1α in Ad-GFP and Ad-GFP-Cre infected MAFs that have lost adenoviral expression vector as a result of prolonged culture. FRAP was performed 48 hour post-transfection with GFP-HP1α. Images were taken before and during recovery after photo-bleaching. During recovery, images were taken at times as indicated. For each time point, fluorescent emissions in the bleached region were normalized to a nearby non-bleached region. First, fluorescent emission measurement (pre- bleached) was set to 1 and subsequent measurements were expressed as a percentage of the first emission. FRAP images and quantitative analysis of FRAP experiments after photo-bleaching are shown.
165
Rb quantitation ofFRAP
Figure IV-4: loxP/loxP MAFs. C Acute RB loss modulates HP1 AcuteRBlossmodulates
Relative Fluorescence r-lahPost-bleach Pre-bleach 0.0 0.2 0.4 0.6 0.8 1.0 1.2 Ad-GFP-Cre Ad-GFP 0 8. dataforhistoneH2BinAd-G 8 8 9. 4 8 10.08
10.68
11 .28 1 1. 8 8 1
2. H2B Mobility 4 Time (s) Time 8 13.08
13 ' 50'' 50'' 0'' 0'' .68 14 .28 166 14 .88 1 5. 48 16.08
16 25'' 25'' α .68 17 dynamics. .28 17 .88 FP and Ad-GFP-Cre infected FP andAd-GFP-Creinfected
Ad-Cre Ad-GFP C) H2B-GFP FRAP imagesand Chapter V:
Summary and Conclusions
The studies reported here attempt to delineate the functional significance
of interactions between RB and various chromatin modifying and remodeling
enzymes. In addition, a hierarchical role of these enzymes for RB-mediated
transcriptional repression has also been reported. Prior to studies reported here, it was known that SWI/SNF chromatin remodeling enzymes are indispensable for
RB-mediated transcriptional repression and cell cycle arrest (28, 30). However, the significance of the RB-HDAC and RB-HMTase interactions has not been investigated in detail. Our results demonstrate the critical role of HDACs for RB- mediated repression of specific target genes (Chapter II). Importantly, studies reported here demonstrate a global role of RB in regulating chromatin architecture thereby highlighting the importance of RB-HMTase interaction in vivo
(Chapter IV). Another important finding reported here is the demonstration that
SWI/SNF is crucial for histone deacetylation of RB-responsive promoters. This finding suggests a hierarchy in the requirement of chromatin remodeling and modifying enzymes for transcriptional regulation (Chapter III). Thus our findings dissect mechanisms of RB-mediated transcriptional repression and unearth a previously unknown function of RB in epigenetic control of chromatin
modification.
167 The results reported herein have important clinical implications for several reasons. First, since HDAC inhibitors (HDACi) are currently under clinical trials as potential anti-cancer drugs, our findings indicate that HDACi do not disengage
RB pathway of cell cycle inhibition. Second, as RB pathway is inactivated in many cancers, the treatment of cancers by restoring functional RB will be dependent on active chromatin remodeling and modifying enzymes. Third, genomic instability arising in RB-deficient cells is presumably due to abnormal heterochromatin histone modifications arising from lack of HMTase activity. Thus, future therapeutics designed to restore functional RB must take into consideration the critical corepessors of RB.
HDAC-dependent gene repression by RB:
Gene regulation is a complex process involving multiple coactivators and corepressors. The formation of nucleosomes serves as a barrier that must be dealt with to allow activation of gene transcription. Early studies identified histone acetyltransferases (HATs) as chromatin modifying enzymes that function as coactivator of gene transcription by adding acetyl groups to histone tails. This acetylation of histone tails loosens the chromatin, allowing basal transcriptional machinery to access the promoter regions of genes to be activated. Subsequent studies identified histone deacetylases as enzymes capable of removing the acetyl groups thereby leading to a compact chromatin structure. Thus, as expected, transcriptional activators recruit HATs whereas repressors recruit
HDACs to regulate gene transcription. RB has been shown to function as a
168 repressor of E2F-mediated gene transcription; however the mechanism
underlying this repression was largely unknown. The discovery of HDACs as
cellular binding partners of RB suggested a model wherein RB-mediated
repression of E2F-regulated genes involved modification of chromatin structure
by HDACs to a more compact form that is inaccessible to basal transcriptional
machinery. Initial studies have demonstrated that HDAC activity was associated
with RB and histone deacetylation was shown to occur only in the context of a
synthetic promoter (12). Later studies using chromatin immunoprecipitations assays (ChIP) demonstrated that RB/E2F regulated genes undergo changes in histone acetylation during cell cycle (16, 20). However, a direct demonstration of the role of HDACs during RB-mediated transcriptional repression and cell cycle arrest was still lacking.
Our approach to dissecting the role of HDACs for RB-mediated transcriptional repression and cell cycle arrest was different from previous studies in several aspects. First, we analyzed critical endogenous RB/E2F target genes involved in S and M phase progression that had not been studied before.
Second, the promoter activity of selected target genes was investigated in the context of the chromatin by making stable reporter constructs. Third, the exclusive requirement of HDAC activity for RB-mediated cell cycle arrest was analyzed for the first time. Our results show that during RB-mediated transcriptional repression, histone deacetylation occurs at all promoters analyzed
(i.e., cdc2, cyclin A, topoIIα and TS). To specifically show that this deacetylation is a result of HDAC activity and not necessarily reduced HAT activity associated
169 with E2Fs, we utilized trichostatin A (TSA), a pharmacological inhibitor of HDAC
activity. TSA was able to inhibit promoter deacetylation of all RB target genes under study. Thus, treatment of cells with TSA allowed us to demonstrate that
promoter histone deacetylation was a result of HDAC activity. The inhibition of
HDAC activity by TSA had functional consequences in the case of cdc2, topoIIα
and TS gene expression, as TSA was able to derepress RB-mediated repression
of these genes. An interesting finding was the observation that even in cases
where TSA partially reversed histone deacetylation, promoter activity was fully
restored (e.g., cdc2). This suggests that only a moderate level of histone
acetylation is sufficient to drive promoter activity and is consistent with previous
reports (22, 31). Furthermore, TSA fully restored the promoter activity, RNA and
protein levels of cdc2, topoIIα and TS genes in presence of PSM-RB, suggesting
that recruitment of HDAC activity is the only means through which RB represses
these genes. In contrast, we did not detect derepression of cyclin A promoter
activity when HDAC activity was inhibited in presence of PSM-RB although the cyclin A promoter was acetylated following TSA treatment. Clearly, the requirement of HDAC activity for RB-mediated target gene repression was context-dependent: while some genes were HDAC-dependent, others were
HDAC-independent for repression by RB. Although our findings identify important principles of RB-mediated transcriptional repression, it does not reveal the relative contribution of these two pathways. With the introduction of promoter microarrays, it is now possible to analyze promoter occupancy of a large number
170 of genes. Current efforts are directed towards dissecting the in vivo occupancy of RB target genes using a genome-wide approach.
Requirement of HDAC activity for RB-mediated cell cycle arrest:
An important finding from our study is the demonstration that HDAC activity is dispensable for RB-mediated cell cycle arrest, since inhibition of HDAC activity by TSA does not rescue cells from arrest inflicted by RB. Although surprising, this phenomenon can be explained by the observation that TSA fails to derepress other critical RB target genes including cyclin A and RNRII.
Presumably, lack of cyclin A gene expression, even when cells are treated with
TSA, prevents cells from progressing through the cell cycle as cyclin A is critical for traversing through the S-phase. Similarly, since RNRII is responsible for dNTP metabolism, lack of RNRII expression may result in depletion the dNTP pool even though HDAC activity is inhibited by RB. Thus, repression of cyclin A and RNRII by PSM-RB might be sufficient to block cells in S phase when treated with TSA although other genes are derepressed. This finding has important therapeutic implication since HDAC inhibitors are currently under phase I and II clinical trials for treatment of several forms of cancers (11). Several of these
HDAC inhibitors including butyrates, valproic acid, SAHA, pyroxamide, depsipeptide, MS-275 and Cl-994 have been shown to reactivate genes (e.g.; p21, p16, and gelsolin) that have been inappropriately inactivated during tumorigenesis (10, 15, 21, 24, 25, 29). Thus, it is hoped that reactivation of these genes by HDAC inhibitors would contribute to cell cycle arrest, differentiation and
171 apoptosis of cancer cells. Based on our finding that TSA does not reverse RB- mediated cell cycle arrest, we conclude that treatment of tumors with HDAC inhibitors will not have the undesired effect of inactivating the RB pathway of cell cycle inhibition.
HDAC-independent gene repression by RB:
Although inhibition of HDAC activity by TSA resulted in derepression of cdc2, topoIIα and TS, it failed to derepress the cyclin A gene. ChIP analysis demonstrated that the cyclin A promoter was acetylated with TSA treatment, similar to other genes analyzed in this study. Furthermore, we also identified another gene (RNRII) that behaved in a similar manner to cyclin A. Thus, our results with cyclin A and RNRII indicate that RB utilizes additional mechanisms other than histone deacetylation to mediate transcriptional repression. Such mechanism(s) could potentially involve histone methyltransferases (HMTases),
DNA methytransferases (DNMTs) or SWI/SNF chromatin remodeling enzymes, since these have been shown to interact with RB (7, 18, 19, 23).
Recent studies have shown that RB target genes, including cyclin A, could be repressed by a mechanism that involves histone methylation and HP1 chromatin association. Such a mechanism, which has been demonstrated in senescent cells, would be irreversible (17). However, we find that repression of cyclin A by PSM-RB did not involve irreversible silencing mechanisms, as we failed to detect any influence of histone or DNA methylation during RB-mediated repression of the cyclin A promoter. Furthermore, repression of cyclin A by PSM-
172 RB was readily reversible when PSM-RB expression was inhibited. Thus, it appears that histone or DNA methylation does not play a role in repressing cyclin
A in asynchronously proliferating cells and is most likely a mechanism to repress cyclin A in senescent cells. A recent report published in EMBO journal by Ait-Si-
Ali et. al. demonstrates that Suv39h-dependent silencing of S phase genes occurs in differentiating but not cycling cells, further supporting our finding (1).
In addition to HDACs, HMTases, HP1 and DNMTs, RB has been known to associate with components of the SWI/SNF chromatin remodeling complex.
Previously work from our lab has shown that BRG1 is required for repression of cyclin A (28). For instance, expression of PSM-RB or p16ink4a to activate endogenous RB in BRG1 and Brm deficient cells does not lead to repression of cyclin A or cell cycle arrest (28, 30). A potential mechanism could be that
SWI/SNF facilitates histone acetylation or deacetylation. However, this action of
SWI/SNF is not sufficient for cyclin A repression as the promoter remained repressed even when histones were acetylated. It is likely that SWI/SNF functions to remodel the promoter structure to inhibit transcription. Consistent with this model, we observe that restriction enzyme accessibility of the cyclin A transcriptional start site is prevented by PSM-RB in SWI/SNF dependent manner.
Therefore, chromatin topology but not histone modification is likely responsible for cyclin A repression by RB. A recent report by Coisy et. al. demonstrates that cyclin A repression requires BRM-mediated chromatin remodeling, substantiating our finding (4).
173 Cross talk between SWI/SNF activity and histone deacetylation:
Although both BRG1 and HDAC1 contain LXCXE motif, BRG1 does not
require this motif to interact with RB. Thus RB can simultaneously recruit BRG1
and HDAC1 into a single complex (6, 30). Prior to our study, the significance of
RB recruiting two classes of proteins that regulate chromatin structure into a
single complex was not understood. The work reported here provides the first
evidence for a link between SWI/SNF chromatin remodeling activity and histone
deacetylation during RB-mediated transcriptional repression (Chapter III). Thus
there appears to be a hierarchy in the requirement of various corepressors for
the biological action of RB. In the case of Plk1, a RB/E2F regulated gene, both
SWI/SNF and HDAC activity was required for repression by RB. Importantly, the
activity of SWI/SNF was required for HDAC-mediated histone deacetylation of
the Plk1 promoter. Although we did not explicitly explore whether binding of
SWI/SNF occurs prior to HDAC binding, studies from yeast SWI/SNF suggest that it is a likely sequence of events.
SWI/SNF can function in both activation and repression of gene transcription. In the budding yeast Saccharomyces cerevisiae, SWI/SNF is required for the activation of HO gene. ChIP analyses to determine ordered recruitment of coactivators have found that recruitment SWI/SNF at the HO promoter occurs prior to recruitment of the histone acetyltransferases SAGA (5).
This suggests that the binding or activity of SWI/SNF is required for the binding or function of SAGA at the HO promoter. Conversely, SWI/SNF complexes may be required for histone deacetylation by HDACs, as we have demonstrated
174 (Chapter III). Our data is consistent with the previous finding that HDAC activity
in the corepressor NURD complex, containing both ATP-dependent nucleosome
remodeling and HDAC activity, is stimulated by ATP on nucleosomal templates
(3). Several possible mechanisms may explain the requirement of SWI/SNF for
histone deacetylation of the Plk1 promoter. First, nucleosome remodeling by
SWI/SNF may facilitate HDAC binding or activity at the Plk1 promoter. Second, since SWI/SNF can catalyze both the disruption and formation of nucleosomal structure in vitro (26), it may be able to promote oscillation of chromatin from repressed and remodeled structure. For complex promoters that are regulated by
several different activators, the simple removal of acetyl groups by HDACs may not be sufficient to keep them at a repressive state, unless SWI/SNF actively
remodels the nucleosome into a repressed state. Future studies will provide
insights into the mechanism of the interplay between SWI/SNF and histone
deacetylation during RB-mediated transcriptional repression.
Regulation of Epigenetic Modification by RB:
Investigators from various disciplines now agree that cancer is not only a
genetic disease but has an epigenetic component as well. Hypermethylation of
tumor suppressor genes has been detected in many forms of cancer. Additionally
a recent study by Fraga et. al. has shown that loss of H4K16 acetylation and
H4K20 trimethylation occurs in many tumor types (8). Critical modulators of
epigenetic modifications can function as tumor suppressors in mouse models
and some of these SWI/SNF, HDACs, HMTase, and HP1 have been shown to
175 interact with RB. Therefore, RB could directly play a role in regulating chromatin
structures by interacting with these factors. Our findings suggest that RB
regulates trimethylation of H4K20 at heterochromatin domains, as loss of RB
results in disruption of this modification at these structures. Additionally, HP1α
association with heterochromatin domains is influenced by RB, since RB loss
deregulates this association. Collectively, our data suggest that RB loss has a
global effect on chromatin modification and dynamics.
The mechanism through which RB mediates change in H4K20
trimethylation is not apparent. Previous studies have demonstrated loss of
H4K20 trimethylation in TKO MEFs through a mechanism discrete from E2F which mediates much of the RB-dependent gene regulation (9). In the case of
RB-mediated transcriptional repression, RB recruits various corepressors including HDACs, SWI/SNF and SUV3-9H1 to promoters of target genes (2, 7,
12, 13, 18). Thus, RB could be required to recruit Suv4-20h HMTases to sites of heterochromatin assembly. Such a possibility is supported by the finding that
HP1α is appropriately enriched in heterochromatin domains, yet the dynamics of
HP1α are altered with RB loss. Prior studies have shown that HP1 is required for the loading of Suv4-20h2 HMTase on chromatin (27). Thus, the effect of RB on
HP1α could be responsible for the failure to mediate H4K20 trimethylation. This mechanism is appealing since it would suggest a specific effect of RB on HP1α function in heterochromatin, as opposed to a more ubiquitous role in regulating
Suv4-20h1/h2 function throughout the nucleus.
176 The consequence of RB loss on heterochromatin domains is not
intrinsically apparent. Since H4K20 trimethylation is associated with
heterochromatin architecture, we postulated that loss of RB may influence the
dynamics of proteins within heterochromatin domains. Therefore, we utilized
FRAP to monitor chromatin dynamics. Loss of RB did not have a profound
effect on histone dynamics. However, we observed reduced mobility of HP1α
with the loss of RB. This effect of RB is directly counter to the effect of HDAC inhibitors or loss of Suv39h1 and Suv39h2, which have the net effect of destabilizing HP1α with heterochromatin domains. Future studies will explore the significance of the changes in HP1α dynamics and H4K20 methylation, which occur with RB loss, on chromatin biology.
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