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CHAF1A Regulates PRC2-mediated Epigenetic Memory
by
Mina Rafiei
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy
Department of Laboratory Medicine and Pathobiology University of Toronto
© Copyright by Mina Rafiei 2014
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CHAF1A Regulates PRC2-mediated Epigenetic Memory
Mina Rafiei
Doctor of Philosophy
Department of Laboratory Medicine and Pathobiology University of Toronto
2014
Abstract
During DNA replication both genetic and epigenetic information on DNA and nucleosomes
needs to be faithfully reproduced. Although DNA replication is well described, the mechanism
of nucleosome and epigenetic replication is not well understood. We studied this issue in the
context of the IFN!-responsive genes (ISGs). Previous work in our lab showed that in the
BRG1-deficient SW13 adenocarcinoma cells, the Polycomb Repressive Complex 2 (PRC2)
blocks IFN!-induction of CIITA and a subset of other ISGs. We detected the repressive PRC2–
mediated epigenetic mark, H3K27me3, across the CIITA locus and at the promoters of other
PRC2-regulated ISGs. We hypothesized that other unknown factors collaborate with PRC2 in
repression of ISGs. To identify the components of this repressive apparatus, we performed an
siRNA screen using a BAC-CIITA vector containing all the remote elements crucial for CIITA
regulation. The screen yielded an unexpected novel link between the histone chaperone,
CHAF1A, and PRC2. We show that siCHAF1A rescues PRC2-repressed genes in a cell cycle
dependent manner. Interestingly, siCHAF1A does not lead to a loss but a redistribution of
H3K27me3. Therefore, we believe that CHAF1A coordinates repressive epigenetic memory by
regulating the genomic positions at which PRC2 methylates histone H3K27 during DNA
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synthesis. Our study thus, for the first time provides important clues regarding how the
epigenetic memory of this important PRC2-mediated mark is maintained in mammalian cells.
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Acknowledgments
I would like to sincerely thank to Dr. Rod Bremner and the department of Laboratory Medicine
and Pathobiology, at the University of Toronto for giving me the opportunity to study and learn
science. I am grateful for the financial support provided by CIHR and other sources at Dr. Rod
Bremner’s lab. I am indebted to my PhD committee, Drs. Jeff Wrana, Andrew Emili and Gerold
Schmitt-Ulms for their invaluable advice and support. I appreciate all the help from Dr. Marek
Pacal and my fellow students and co-workers at the lab of Dr. Rod Bremner. I am thankful for all
the help from Dr. Kevin Brown for analyzing the screen data and from Alexandro Datti, Thomas
Sun and Frederick Sagayaraj Vizeacoumar at S.M.A.R.T., the Mount Sinai robotic facility. Last
but not least, I am grateful for the all understanding and support I received from my family.
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Table of Contents
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Acknowledgments.......................................................................................................................... iv!
Table of Contents ............................................................................................................................ v!
List of Figures ................................................................................................................................. x!
List of Appendices ......................................................................................................................... xi!
List of Abbreviations .................................................................................................................... xii!
Chapter 1 ......................................................................................................................................... 1!
1! General Introduction .................................................................................................................. 1!
1.1! Gene regulation ................................................................................................................... 2!
1.1.1! Gene regulation at the transcriptional level ............................................................ 2!
1.1.1.3! Chromatin and epigenetics...................................................................................... 4!
1.2! Gene regulation by PRC2 and TrxG................................................................................... 6!
1.2.1! PcG complexes........................................................................................................ 7!
1.2.2! TrxG........................................................................................................................ 9!
1.3! IFN! .................................................................................................................................. 11!
1.4! CIITA regulation ............................................................................................................... 11!
1.5! Identifying new factors in CIITA regulation ..................................................................... 12!
1.5.1! Replication Fork proteins, epigenetic memory and CIITA regulation ................. 13!
1.5.2! Histone Chaperones, epigenetics and cancer ........................................................ 15!
1.5.3! Nucleosome replication: Conservative vs. Semiconservative .............................. 15!
1.5.4! Chaperones regulate transport, modification, and deposition of new histones..... 17!
1.5.5! Chaperones recycle old histones during replication ............................................. 20!
1.5.6! Chaperones and Epigenetic Memory: The Example of CAF1 ............................. 24!
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1.5.7! Other potential roles for CAF1 in maintaining repressive chromatin states......... 29!
1.5.8! CAF1 and Cancer.................................................................................................. 29!
1.5.9! ASF1, HIRA and cancer ....................................................................................... 30!
1.5.10! DEK, DAXX, H3.3 and Cancer............................................................................ 31!
1.5.11! Future directions ................................................................................................... 34!
1.6! Hypothesis......................................................................................................................... 35!
Chapter 2 ....................................................................................................................................... 36!
2! RNAi Screens to Identify new components regulating IFN! responsive genes (ISGs)........... 36!
2.1! Abstract ............................................................................................................................. 37!
2.2! Introduction....................................................................................................................... 38!
2.3! Material and Methods ....................................................................................................... 42!
2.3.1! Cell culture and adenovirus .................................................................................. 42!
2.3.2! HTP siRNA screening........................................................................................... 42!
2.3.3! BAC construction and reporter assays .................................................................. 43!
2.3.4! Statistical Methods for Hit Selection .................................................................... 43!
2.4! Results............................................................................................................................... 44!
2.4.1! The HTP siRNA screen optimization ................................................................... 44!
2.4.2! The primary WT and Suppressor screens ............................................................. 52!
2.4.3! The secondary WT and Suppressor screens for hits validation ............................ 55!
2.4.4! Tertiary Validation on Endogenous CIITA ........................................................... 57!
2.5! Discussion ......................................................................................................................... 58!
Chapter 3 ....................................................................................................................................... 60!
3! CHAF1A Regulates the Distribution of PRC2-mediated epigenetic mark, H3K27me3, During Replication ................................................................................................................... 60!
3.1! Abstract ............................................................................................................................. 61!
3.2! Introduction....................................................................................................................... 62!
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3.3! Material and methods........................................................................................................ 64!
3.3.1! Cell culture and siRNA transfection ..................................................................... 64!
3.3.2! RNA extraction and reverse transcription (RT).................................................... 64!
3.3.3! Western Blotting ................................................................................................... 64!
3.3.4! Chromatin immunoprecipitation (ChIP). .............................................................. 65!
3.3.5! Quantitative Real-Time PCR (qPCR)................................................................... 65!
3.3.6! Plasmid construction............................................................................................. 66!
3.3.7! Immunostaining .................................................................................................... 66!
3.3.8! Cell cycle block..................................................................................................... 66!
3.4! Results............................................................................................................................... 67!
3.4.1! CHAF1A and PCNA repress CIITA induction ..................................................... 67!
3.4.2! CIITA rescue by CHAF1A KD is BRM independent ........................................... 71!
3.4.3! The effect of siCHAF1A on known regulators of IFN! signaling........................ 73!
3.4.4! Effects of siCHAF1A on cell survival and DNA damage .................................... 75!
3.4.5! Depleting other components linked to histone deposition, DNA replication or gene silencing does not rescue CIITA ................................................................... 78!
3.4.6! CHAF1A represses multiple PRC2-regulated ISGs ............................................. 81!
3.4.7! CHAF1A is required for PRC2-mediated chromatin modification ...................... 83!
3.4.8! General role for CHAF1A in promoting PRC2-mediated gene silencing ............ 85!
3.4.9! CHAF1A-mediated repression requires S-phase .................................................. 87!
3.4.10! SiCHAF1A causes a dramatic redistribution of H3K27me3 without changing PRC2 localization ................................................................................................. 89!
3.5! Discussion ......................................................................................................................... 92!
3.5.1! A novel direct link between CHAF1 and PRC2-mediated gene repression during replication .................................................................................................. 92!
3.5.2! CHAF1A is required for printing the H3K27me3 mark at the proper targets ...... 93!
4! Discussion and Future Directions ............................................................................................ 97!
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4.1! Discussion ......................................................................................................................... 98!
4.1.1! CHAF1A maintains H3K27me3 during replication ............................................. 98!
4.1.2! CHAF1A may guide the H3K27me3 deposition indirectly.................................. 99!
4.1.3! CHAF1A may activate PRC2 ............................................................................. 101!
4.2! Future directions ............................................................................................................. 102!
4.2.1! The global effect of depleting CHAF1A on PRC2 regulated genes and redistribution of H3K27me3 ............................................................................... 102!
4.2.2! DNA methylation and histone modification ....................................................... 102!
4.2.3! Identifying the components involved in the CHAF1A-dependent maintenance of H3K27me3...................................................................................................... 103!
4.2.4! Structure-Function analyses of CHAF1A........................................................... 104!
4.2.5! Using aniPOND to screen for additional factors co-operating with CHAF1A at the replication forks ............................................................................................ 105!
4.3! Concluding remarks ........................................................................................................ 105!
References................................................................................................................................... 107!
Appendices.................................................................................................................................. 131!
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List of Figures
Fig 1.1 Histone chaperone functions during DNA replication…………………………………23
Fig 1.2 Multiple roles for CAF1 at heterochromatin…………………………………………...28
Fig 2.4.1 Optimizing siRNA transfection and viral transduction conditions…………………..47
Fig 2.4.2 Optimizing the timing and order of the screen steps………………………………....48
Fig 2.4.3 Automated HTP version of the optimized protocol…………………………………..49
Fig 2.4.4 Testing the “B score” method of hit analysis in the optimized protocol …………….50
Fig 2.4.5 A. Plot of B scores from the pilot Wt screen………………………………………….51
Fig 2.4.6 Schematic diagram of the primary siRNA screen …………………………………...53
Fig 2.4.7 Schematic diagram of secondary validation screen…………………………………..56
Fig 3.4.1 CHAF1A and PCNA repress endogenous CIITA induction………………………….69
Fig 3.4.2 CIITA rescue by siCHAF1A is BRM-independent……………………………………72
Fig 3.4.3 The effect of siCHAF1A on the level of IRF1, p-STAT1 or STAT1 proteins………..74
Fig 3.4.4 Effects of siCHAF1A on cell growth, apoptosis and DNA damage. …………………76
Fig 3.4.5 The role of other known CHAF1A interacting proteins in CIITA silencing…………..80
Fig 3.4.6 The effect of CHAF1A KD on PRC2/BRG1dependent ISGs…………………………82
Fig 3.4.7 The effect of CHAF1A KD on H3K27me3 at PRC2/BRG1 regulated ISGs………….84
Fig 3.4.8 General effect of CHAF1A in PRC2-mediated gene silencing ……………………….86
Fig 3.4.9 The role of CHAF1A in gene silencing is cell cycle dependent………………………88
Fig 3.4.10 CHAF1A KD alters H3K27me3 distribution………………………………………...90
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List of Appendices
Apx1. List of primers for RT-PCR……………………………………………………………..128
Apx 2. List of SiRNAs………………………………………………………………………….130
Apx 3. List of Abs………………………………………………………………………………131
Apx 4. List of activator and inhibitor hits form primary and secondary screens…………….....132
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List of Abbreviations
Ago: argonaut
aniPOND: accelerated native iPOND
BAF: BRG associated factor
BAC: bacterial artificial chromosome
BRG1: Brahma related gene1
BRM: Brahma
Bromodomain: binds acetylated histones
CHD: chromodomain and helicase-like domain
CIITA: MHC ClassII transactivator
ChIP: chromatin immunoprecipitation
ChIP-chip: immunoprecipitation coupled with DNA tiling array
Chromodomain: binds methylated histones
CpG: cytosine Guanine nucleotide
CBX: chromobox homolog
CHAF1A: chromatin assembly factor1
COSMIC: somatic mutation in cancer
CSE1l: CSE1 chromosome segregation 1-like
DNMT: DNA (cytosine-5-)-methyltransferase
ETM: epithelial-mesenchymal transition
EZH: zeste homolog 2
EED: embryonic ectoderm development
EHMT2: euchromatic histone-lysine N-methyltransferase 2
FXYD1: FXYD domain containing ion transport regulator 1
GBP1: guanylate-binding protein1
H3K4ac: acetylated lysine4 at histone3
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H3K9me3: tri-methylated lysine 9 at histone H3
H3K27me3: tri-methylated lysine 27 at histone 27
HIRA: HIR histone cell cycle regulation defective homolog A
HDAC: histone deacetylase
HP1: heterochromatin protein1
HOX: homeobox
HOTAIR: HOX transcript antisense RNA
HTTP: high throughput
IFI2: interferon, gamma-inducible protein
IFITM3: interferon-induced transmembrane protein 3
IFN: interferon
IRF1: interferon regulatory factor1
IP: immunoprecipitation
iPOND: isolation of proteins on nascent DNA
ISWI: imitation SWI
IFNGR1: interferon gamma receptor 1
JAK: Janus family protein tyrosine kinase
JARID: Jumonji AT-rich interactive domain
K: lysine
LSD1: lysine specific demethylase 1
Luc: luciferase
LincRNA: intergenic non-coding RNA
MLL: Mixed lineage leukemia
MAPK13: mitogen-activated protein kinase 13
MHC: major histocompatibility complex
MBD: methyl-CpG binding domain protein 1
MeCP2: methyl CpG binding protein 2
MMS: methyl methane-sulfonate
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ncRNA: non-coding RNA
PIV: promoter IV of CIITA
PLRG1: pleiotropic regulator 1
PITX2: paired-like homeodomain transcription factor2
PcG: Polycomb protein group
PCNA: proliferative cell nuclear antigen
POLE: polymerase (DNA directed), epsilon
POLD: polymerase (DNA-directed), delta
PRC: Polycomb Repressive Complex
PRPF8: PRP8 pre-mRNA processing factor 8 homolog
qPCR: real-time quantitative PCR
RT: reverse transcription
R: arginine
RING: ring finger protein
ROR1: receptor tyrosine kinase-like orphan receptor 1
RBBP: retinoblastoma associated protein
RISC: RNA-induced silencing
SD: standard deviation
SET: SuVar39, Enhancer of Zeste, and Trithorax
SLIDE: SANT-Like ISWI domain
SOCS: suppressor of cytokine signaling
SNAIL: snail zinc finger
STAT1: signal transducer and activator of transcription
SUV39H: suppressor of variegation 3-9 homolog
SUZ12: suppressor of zeste
SWI/SNF: mating type switching and sucrose non-fermenting
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SUMO: sumoylation
TET: ten-eleven translocations
TrxG: trithorax group
Ub1: mono-ubiquitylation
Wt: wild type
XCI: X chromosome inactivation
1
Chapter 1
1 General Introduction
2
1.1 Gene regulation
Gene regulation in eukaryotes is a complex and sophisticated process of fundamental importance
for all biological events. The expression of genes in cells can be regulated on several levels in
response to internal and external stimuli. First, the DNA regulatory elements and promoters and
proteins that bind to these elements can control the rate of transcription. Second, mRNA splicing
and stability affect the amount of mRNA available. Finally, if relevant, the dynamics of mRNA
translation and post-translational protein modifications will crucially influence the final amount
of gene product generated [1, 2]. Here we focus on transcriptional level of gene regulation.
1.1.1 Gene regulation at the transcriptional level
The control of gene expression at the transcriptional level is the result of three critical
components. (1) DNA binding of transcriptional factors (TFs) that either increase (activators) or
decrease (repressors) the interaction and stabilization of RNA polymerase at the promoter of the
genes. (2) The presence of co-regulators which collaborate with TFs and can induce (co-
activators) or reduce (co-repressors) the transcriptional rate of genes. (3) The activity of factors
which affect chromatin structure by altering the accessibility of the DNA to the transcriptional
machinery. These latter factors can be histone modifying factors or chromatin remodeling
complexes (see 1.2.2.1 for a detailed discussion) [3-8].
1.1.1.1 Transcription Factors
TFs are proteins that bind to regulatory DNA sequences and either enhance or inhibit
transcription [9]. TFs are a large family of proteins that can be classified based on their: 1)
mechanism of action, 2) spatial and temporal regulatory function and 3) DNA binding domain
structure.
There are two major classes of TFs based on their mechanism of action. The first class
includes the so called general TFs such as TFIIA, TFIIB, TFIID, TFIIE, TFIIF and TFIIH that
are required for the basal level of transcription. General TFs form pre-initiation complexes at
gene promoters which are essential for the assembly of RNA polymerase II machinery and
initiation of transcription [10-14]. The second class includes TFs (e.g., c-MYC, OCT-1, SP-1)
that bind upstream of the initiation sites to induce or repress transcription.
3
TFs can also be classified based on the spatial and temporal regulatory function. For
example, some of the transcription factors are constitutively expressed in all cell types (e.g.,
general TFs). However a large family of transcription factors is activated in specific cell types at
different developmental stages or in response to internal or external stimuli. During embryonic
development, the expression of cell specific transcriptional factors is tightly controlled to switch
on or off cell type specific genes. For example, homeobox (Hox) genes control the body plan of
the embryo and determine the type of segment structures (e.g. legs, antennae, and wings in fruit
flies or the different vertebrate ribs in humans) that will form on a given segment [15]. TFs also
respond to intercellular (e.g., TGF", interferons) and environmental (e.g., heat shock, hypoxia)
signals. For example, in the TGF" signaling pathway, the binding of ligand to a receptor leads to
phosphorylation of receptor-regulated SMADs (R-SMADs) which then bind the coSMAD
SMAD4. R-SMAD/coSMAD complexes translocate to the nucleus where they act as TFs
regulating target gene expression [16-18].
Finally, based on DNA binding domains, TFs can be classified to 5 major super classes:
1) Basic domain, 2) Zinc-coordinating DNA binding domains, 3) Helix-turn-helix, 4) beta-
scaffold factors with minor groove 5) Others. Each super class contains different sub classes and
families. Not surprisingly, altogether this is a large number of proteins. Indeed, TFs make up at
least 10% of genes in the genome, which makes this family the single largest family of human
proteins [19].
1.1.1.2 Transcriptional co-regulators
The regulatory functions of TFs as activators or repressors are achieved in collaboration with co-
regulators (co-activators or co repressors). Co-regulators do not typically have DNA binding
domains and are recruited to the DNA sequences by TFs. There are different classes of co-
regulators. For example repression of E-cadherin, which is an epithelial marker and is observed
in epithelial-mesenchymal transition (EMT), is achieved by collaboration of TFs activated by
different signaling pathways such as TGF", Notch, Wnt, as well as hypoxia and co-regulators
including Snail, Slug, Twist, ZEB1 and so on [20-22]. Co-regulators can be histone modifying
enzymes such as histone acetyltransferases (HATs) that act as co-activators or histone
deacetylases and histone methylases of lysine K9 and K27 which are co-repressors [13, 23-29].
4
A special class of co-regulators includes ATP-dependent chromatin remodeling
complexes such as SWI/SNF that can be either co-activators by making the chromatin more
accessible to transcription machinery or repressors by making the chromatin more condensed and
not accessible for transcription [30-35]. These will be discussed in detail in the section 1.2.2.1.
1.1.1.3 Chromatin and epigenetics
DNA is packed into a compact structure called chromatin so that it can fit into the cell nucleus
[36]. Chromatin is a dynamic and highly organized system made of repeating units referred to as
nucleosomes [36-40]. In eukaryotic cells about 147 base pairs (bps) of DNA are wrapped around
a histone octamer (two of H3/H4 and two of H2A/H2B) to form a nucleosome [39-41]. The H1
histone, which is called the linker histone, binds the DNA segments between nucleosomes and
helps wrap nuleosomes into a higher conformation [39, 40]. Chromatin is classified into two
forms, based on its structure. The first, euchromatin, is less condensed, gene-rich and replicates
early. The second, heterochromatin, is highly condensed, mostly not transcribed and replicates
late in the S phase of the cell cycle [39, 40, 42-44].
Chromatin structure functions as a barrier for transcription and needs to be rendered
accessible for transcription factors and transcriptional machinery. This can be done, in
eukaryotic cells in two ways. The first, carried out by ATPase chromatin remodeling complexes
which alter nucleosome structure using energy derived from ATP hydrolysis [45-47]. The
second, performed by histone modifying enzymes, relaxes histone-histone and histone-DNA
interactions through covalent histone modifications.
There are different types of histone modifications such as acetylation, phosphorylation,
methylation, ubiquitination and sumoylation that are context-dependent and can either repress or
activate gene transcription. Two important and well known histone modifications associated
with gene repression are the methylated lysine 9 and 27 of histone H3 (H3K9 and H3K27). The
former is mediated by several different histone methyltransferases (see below) and the latter is
mediated by Polycomb Group (PcG) protein complexes [48].
5
In addition to histone modifications, DNA can also be methylated which usually
correlates with gene repression. The DNA and histone modifications together are referred to as
“epigenetic code” (epi = over, beyond) which defines the status of gene expression as either
silent or active without changes in DNA sequence [49]. A strict definition of epigenetics only
includes chromatin events that are inherited from one cell or organism generation to the next.
However, the term is also used more broadly to refer to chromatin modifications in any cell, even
post-mitotic cells.
1.1.1.3.1 DNA methylation
DNA methylation is an epigenetic modification carried out by DNA methyltransferases that
covalently add methyl groups to the cytosine nucleotides of DNA. It occurs in those parts of
DNA sequences where a cytosine nucleotide sits next to a guanine nucleotide; such sequence is
referred to as CpG [50]. Unmethylated CpG clusters are called CpG islands, which are usually
associated with gene promoters in normal cells [51-53]. Abnormal hypermethylation of CpG
islands is often seen at the promoters of tumor suppressors, leading to their inactivation, in
different types of cancers such as colon cancer, glioma and leukemia [54, 55]. DNA methylation
attracts methyl-CpG-binding proteins such as MBD1 and MeCP2, which in turn promote binding
of histone modifying enzymes such as histone deacetylases [56, 57]. The combination of DNA
and histone modification leads to chromatin compaction and gene repression [58].
DNA methylation/demethylation is mediated by different classes of DNA methyl
enzymes. One class of DNA methyltransferases is the DNMT, DNA (cytosine-5-)-
methyltransferase family. One member of DNMT family is DNMT1 which maintains the DNA
methylation pattern during replication [59]. DNMT1 recognizes methylated CpGs at parental
DNA strands and transfers methyl groups to the correct CpG sites on the daughter DNA [60].
DNMT1 also interacts with Proliferative Cell Nuclear Antigen (PCNA) at the replication forks,
which may be important in the maintenance of DNA methylation during replication [58, 61].
Other DNMT family members such as DNMT3a and DNMT3b are involved in de novo
methylation of DNA during development [59]. Recent studies indicate that Ten-Eleven
Translocation (TETs) DNA methylase enzymes, which oxidize 5-methylcytosine (5mC) to 5-
hydroxymethylcytosine (5hmC), serve to demethylate DNA [62]. DNA
methylation/demethylation is a dynamic event and has crucial impact on gene regulation. The
6
cross talk between DNA and histone epigenetic modifications leads to gene activation or
repression.
1.1.1.3.2 Histone methylation
Both histones H3 and H4 can be methylated and these marks can be either activating or
repressive. Methylation of H3 is well studied, and both lysine (K) and arginine (R) residues can
be methylated to a different degree: mono (me1), di (me2) or tri-methylation (me3). The
methylation of K 4 or 36 of histone H3 is linked to gene activation while the methylation of K 9,
27, or 79 of this histone lead to gene repression [48, 63]. SET1 and Mixed lineage leukemia
(MLL) family of histone methyl transferases are involved in H3K4 methylation [64]. Histone
methylation is reversible and can be erased by specific histone demethylases. For example
Lysine-specific histone demethylase1 (LSD1) and Jumonji AT-rich interactive domain 1
(JARID1) are responsible for erasing methyl groups from histone H3K4 [65].
The trimethylation of K 9 of histone H3 (H3K9me3) and of H3 lysine 27 (H3K27me3)
are two major and well-studied repressive marks that have been conserved throughout eukaryotic
evolution [66]. These latter histone modifications are mediated by several different types of
histone methyltrasferases such as Suv39 family containing SET domain-containing enzymes [48,
67, 68]. The methylation of H3K9 provides docking sites for proteins such as heterochromatin
protein 1(HP1) which induces chromatin condensation and maintains gene silencing [69-73].
The H3K27me3 modification is mediated by PcG proteins. Components of PcG complex
contain chromodomains which bind H3K27me3 and mediate gene silencing. We will discuss
these in more detail below.
1.2 Gene regulation by PRC2 and TrxG
The DNA sequences and genetic information are shared by all cells but what distinguishes their
identity and fate is their different gene expression patterns. These patterns (i.e., active or silent)
are established during development. Next, the memory of the transcriptional state needs to be
maintained during cell division and DNA replication [74]. Trithorax group (TrxG) and PcG
complexes are known to maintain the memory of the gene status as active and repressed,
respectively [75, 76].
7
PcG proteins were identified more than 50 years ago in Drosophila as negative HOX
genes regulators, revealed by mutations of HOX gene activators [74, 77]. Later, TrxG proteins
were identified via mutations that rescued the phenotype observed in PcG protein mutations [75,
76]. Therefore, PcG and TrxG show antagonistic effects on gene regulation [78, 79]. The
function of PcG and TrxG are evolutionarily conserved. In vertebrates, TrxG as PcG are also
crucial gene regulators but have been also implicated in other biological events such as cell fate,
differentiation and proliferation, X-chromosome inactivation, stem cell identity and cancer [79,
80]. It has been shown that the PcG and TrxG complexes are bound to the same DNA sequences
which allows for switchable regulatory platforms [76].
1.2.1 PcG complexes
PcGs are essential in maintaining the expression status of genes imprinted during early
development and their dysfunction causes homeotic transformations[81]. There are five different
PcG complexes of which PRC1 and PRC2 are the best characterized [23].
PRC1 contains catalytic subunits RING1A and RING1B, which catalyze lysine 119
mono-ubiquitylation (K119) of histone H2A (H2AUb1). The PRC1 core complex contains the
chromodomain-containing protein Chromo Box Homolog 4 (CBX4), which aids SUMO E3
ligase to sumoylate many other proteins including DNA methyltransferases, such as DNMT3a
[82]. However the function of sumoylation in PcG-mediated gene silencing is not completely
clear.
The catalytic subunit of PRC2 that mediates di-methylation and tri-methylation of
H3K27, is formed by Enhancer of Zeste homolog 2 (EZH1/2), which binds to Suppressor of
zeste 12 (SUZ12) and one of Embryonic ectoderm development isoforms (EED 1-4) for full
activity [83]. The core of the PRC2 complex also contains cofactors such as Retinoblastoma-
associated protein, RbAp46/48 (also known as RBBP7/4), and JARIDs. Finally, histone
deacetylases, HDAC1 and HDAC2, transiently bind the core complex and are required for its
recruitment to target loci [83-87].
EZH2 mediates di-methylation and tri-methylation of H3K27, a modification recognized
by the chromodomain subunit of PRC1 complex, which then leads to chromatin structure
alteration and transcriptional repression [88, 89]. It has been shown that RNA polymerase II
8
(RNApol II) pauses at some PRC2-rich regions and it is believed that short transcripts that are
bound to such paused RNApol II also recruit PRC2 [90, 91]. Recent studies have shown that the
function of PRC2 is not limited to gene regulation and it is also involved in other biological
process of cells such as cell cycle control and DNA damage [92-94].
PRC2 regulates thousands of genes including many cell type dependent transcriptional
factors [78, 95]. PRC2 components such as EZH2 are upregulated and act as oncogenes in
several types of cancers [96, 97]. For example, EZH2 has been used as a biomarker for breast
and prostate metastasis. Also, over expression of PRC2 induces silencing at the promoter of
tumor suppressor genes [97, 98]. Recent studies have identified mutation of EZH2 in SET-
domain of the protein which is associated with gain of function and induction in H3K27me3
repressive mark [99, 100]. Therefore, it is crucial, for the sake of improved understanding and
treatments of cancer, to decipher how PRC2 is regulated and distributed in cancer cells.
1.2.1.1 Mechanism of PRC2 recruitment
PRC2 is recruited to the specific DNA sequences, Polycomb Responsive Elements (PREs), in
Drosophila but only recently DNA elements have been described that recruit PRC2 to their
targets in vertebrates[101]. Importantly, PRC2 recruitment to target genes can be assisted by a
variety of other factors [86, 87]. For example, recently, non-coding RNAs (ncRNAs) have been
described as important PRC2 recruiters in vertebrates [85]. The involvement of ncRNAs was
first reported in the context of HOXD cluster repression. Here, the recruitment of PRC2 to its
regulatory elements is mediated by long ncRNA (lincRNA) HOTAIR which is expressed from
the HOXC cluster [102]. Moreover, ncRNAs are also involved in the X chromosome
inactivation (XCI) by recruiting EZH2 to initiate and spread silencing [102]. Recent studies
have been showing more evidence that lincRNAs are associated with PRC2 at the targeted genes
and knocking down these lincRNAs activates PRC2-repressed genes [103, 104].
In addition to ncRNAs and lincRNAs, there are several protein factors which are
involved in PRC2 recruitment to the target genes. These proteins include, the RNAi machinery
factors such as Argonauts, the transcription factor Snail family zinc finger 1 (SNAIL1) or the
Jumonji protein JARID2 [13, 105].
9
Recent studies show that PRC2 is recruited to hypomethylated CpG rich regions [106-
108]. However the mechanism of the recruitment and to which extent DNA methylation repels
PRC2 complex recruitment still needs to be addressed.
1.2.1.2 PRC2-dependent epigenetic inheritance
Another important question concerning PRC2 activity is how PRC2-mediated signatures are
inherited during DNA replication when the histone structure and epigenetic marks have been
challenged. During S-phase, nucleosomes are disrupted ahead of the replication fork and then
reassembled to the newly synthesized DNA. Therefore a specific mechanism must exist to
preserve the parental PRC2-mediated silencing marks.
To date, two models have been proposed to explain the inheritance of the PRC2-mediated
epigenetic mark, H3K27me3. The first one is a “positive feedback” model in which the
preexisting H3K27me3 marks stimulate PRC2, thus methylating newly deposited histones [109,
110].
Recently, a Drosophila study has proposed another model. This model posits that
H3K27me3 inheritance does not rely on the actual mark but on the presence of the PRC2
complex [111]. Here, PRC2 remains bound to DNA during replication and the pre-existing
marks are in fact stripped off histones during replication. PRC2 components modify both
parental and daughter histones at the same place again in the G2 phase after DNA replication
[111].
1.2.2 TrxG
TrxG proteins are categorized into three classes based on their function. In vertebrates, the first
class includes the SET domain containing complexes that methylate Lys 4 of histone H3
(H3K4me3), a gene activating mark [79]. This class also includes Myeloid/Lymphoid Or
Mixed-Lineage Leukemia, MLL histone methyltransferases which mediate global gene
activation [76, 112].
In the second class of TrxG proteins are the sequence-specific DNA binding proteins that
contain some histone modifying enzymes and chromatin remodeling complexes. DNA binding
proteins localize to TrxG responsive elements (TREs), found in Drosophila, which often overlap
10
with PREs [76, 78]. It has been suggested that CpG-rich sequences may perform this role and
recruit PcGs and TrxGs to DNA [106].
The third class contains the ATP-dependent chromatin remodeling complexes which
activate genes by inducing nucleosome eviction or nucleosome sliding or changing the chromatin
architecture (e.g., creating loops) all of which renders DNA target motifs more or less accessible
for transcription activators or repressors and RNA polymerase machinery [113]. There are
several types of ATP-dependent chromatin remodeling complexes and they regulate a large
number of genes in different cell types but they are also involved in other cellular process such
as cell cycle or cell signaling [33, 114, 115]. We will discuss these remodeling complexes in
more detail below.
1.2.2.1 ATP-dependent chromatin remodeling complexes
Cells use the ATP-dependent chromatin remolding complexes to adjust the DNA surrounding
histones to make it more accessible to transcriptional factors and RNA polymerase or providing
higher level of chromatin structure by looping which are associated with gene activation or
repression respectively. They are protein complexes conserved from yeast to mammals and the
complexes can be categorized into several subfamilies based on the presence of conserved
domains [46, 116].
One subfamily of ATPase chromatin remodelers includes the imitation switch (ISWI)
complexes, which contain different histone binding domains such as SANT and SLIDE (SANT-
Like ISWI) domains [117]. Next, the chromodomain helicase DNA-binding (CHD) subfamily
binds to acetylated histones [118]. The SWI/SNF chromatin remodeling complex is another
important subclass which was first identified in yeast in a screen aimed to identify genes that
regulate mating-type switching (SWI) and sucrose non-fermenting (SNF) phenotypes [119-121].
In mammals, there are two major SWI/SNF complexes: BAF (hSWI/SNFA) or PBAF
(hSWI/SNFB). These two complexes have 10 subunits in common, however, the catalytic
subunit of PBAF is Brahma-Related gene1 (BRG1) whereas BAF can associate with either
BRG1 or Brahma (BRM) [122]. Human BRG1 and BRM show approximately 74% identity on
the amino-acid level and although they both act as engines of the SWI/SNF complexes, they
seem to play different roles during differentiation and proliferation and cell signaling [123-125].
11
BRG1 is essential for tumor suppression and plays important roles in differentiation, gene
regulation and cell cycle control [126]. For example, BRG1 is mutated, deleted or silenced in
several types of cancers such as non-small cell lung carcinoma, prostate cancer, breast cancer
and adrenal carcinoma [127]. BRG1 heterozygous mice are prone to transformation [123].
Importantly, our lab showed that BRG1 is required for the induction of a subset of Interferon !
(IFN!)- responsive genes [33, 115, 128]. Another pathway regulated by both BRG1 and BRM is
the Notch signaling pathway. Here, their activity is antagonized by PcG complexes [125, 129].
This is very interesting because, as described in Chapter 3, we observed the co-operation of
BRG1 and PRC2 in the regulation of genes downstream of the IFN! signaling pathway.
1.3 IFN!
Interferons are soluble proteins, which were initially described as factors secreted in response to
viral infection, designated to inhibit cell proliferation and viral replication [130]. There are two
types of IFNs, IFN #/" (type I) and IFN! (type II). Both types have antiviral and
antiproliferation activities but bind to different cell surface receptors and have different
biological effects [131, 132].
IFN! signaling is initiated by binding of the ligand to IFN! receptors (IFNGR1-
IFNGR2), which results in their oligomerization. This in turns leads to the phosphorylation of
receptor-associated Janus kinases, JAK1 and JAK2 [131, 132]. These kinases phosphorylate
Signal transducer and activator of transcription-1 (STAT1) [131, 133, 134]. Dimerized p-
STAT1 translocates to the nucleus and binds to regulatory elements of IFN! stimulated genes
(ISGs) [131, 133, 134]. ISGs respond to IFN! in most cell types [135, 136]. Unfortunately, very
little is known about the regulation of ISGs in cancer. As discussed below and in more detail in
Chapters 2 and 3, using the ISG Class II transactivator (CIITA), the master regulator of MHC
class II, as a model, our lab has uncovered several novel factors in ISG regulation in cancer cells
[137].
1.4 CIITA regulation
CIITA is the master regulator and a co-activator of MHC class II (MHC) induction [138].
CIITA is regulated by four alternative promotes (I-IV). Promoters (I-III) are used for the
constitutive form of CIITA induction which is observed in antigen presenting cells such as B
12
lymphocytes and dendritic cells [138]. Promoter IV is used in the inducible form of CIITA in
other cell types and drives the IFN! response [139].
Immune cells target not only infected cells but can also detect transformed cells and
destroy them [131]. This process is called immunosurveillance [131]. CIITA is silent and not
responsive to IFN!, in tumors such as Uveal melanoma, adrenal carcinoma and colorectal
cancers [140, 141]. It is likely that silencing of CIITA is a strategy that cancer cells use to switch
off MHCII and escape immunosurveillance. Therefore, understanding the mechanism of CIITA
regulation should help improve drug design to treat cancers where MHCII is inactivated.
Our lab has used CIITA as a model to understand the regulation of ISGs but also to study
gene regulation in general and has made significant contributions in this field. Our previous
work linked SWI/SNF to regulation of ISGs in cancer cells and found that BRG1 is required for
CIITA to respond to INF! [33, 115, 137]. Our lab identified multiple STAT1/IRF1 enhancers
across the CIITA locus, spanning >100 kb far from transcription start site, which are critical for
CIITA induction in response to IFN! [137]. In addition, we showed that in the BRG1-deficient
adrenal carcinoma SW13 cells CIITA and a subset of ISGs are regulated by PRC2 repressive
complexes. We detected the PRC2-signature chromatin mark, H3K27me3 as well as the PRC2
components SUZ12 and EZH2 at multiple sites across the CIITA locus (unpublished data,
Chapter 3). We showed that re-introducing BRG1 to SW13 cells restored the IFN!
responsiveness of CIITA [33, 115]. Moreover, we showed that knocking down PRC2
components (EZH2 and SUZ12) rescued CIITA induction and reduced H3K27me3 across the
CIITA locus (unpublished data, Chapter 3).
Together, these findings raise the possibility that inactivation of SWI/SNF and/or
activation of PRC2-mediated repression might be mechanisms cancer cells use to silence MHCII
and escape immunosurveillance.
1.5 Identifying new factors in CIITA regulation
Using CIITA as a model of ISG regulation has several advantages. First, CIITA is a very
important ISG and a master regulator of MHC class II, and thus the knowledge of CIITA
regulation has broad clinical relevance. Further, as we have found, this gene is regulated by both
PRC2 and BRG1, members of PcG and TrxG family of proteins, respectively. Since multiple
13
genes are regulated by these two factors, uncovering how they work in the context of CIITA
could likely be extended to many other PcG /TrxG regulated genes.
Z. Ni in our lab built a 194 kb BAC-CIITA luciferase (Luc) reporter containing all the
newly identified remote enhancers, and showed that the construct is, like the endogenous CIITA
locus, also BRG1-dependent. As described in Chapter 2, we took advantage of this reagent and
performed high throughput (HTP) siRNA screens to determine: A. whether we could rescue
CIITA responsiveness in BRG1-deficient cells by suppressing factors that repress this target
(Suppressor screen) and B. which additional regulators (activators or inhibitors) affect CIITA
regulation in the presence of BRG1 (Wild type screen). In Chapter 2, the optimization
procedures for the different steps of the siRNA screens are discussed and the identified hits are
reported. In Chapter 3 we focus on two hits identified by the Suppressor screen, both replication
fork proteins, and dissect the mechanism of these factors in the CIITA regulation.
1.5.1 Replication Fork proteins, epigenetic memory and CIITA regulation
As we will describe later, our screen for repressors of CIITA in BRG1-deficient cells uncovered
two candidates, CHAF1A (P150), the large subunit of CAF-1 histone chaperone complex and
PCNA, Proliferating Cell Nuclear Antigen, which is a cofactor of DNA polymerase delta. Both
identified hits are required for DNA replication and are also involved in replication-coupled gene
silencing.
The main function of CAF-1 is to assemble the histones H3/H4 into DNA at the
replication fork and the complex includes CHAF1A, CHAF1B and RbAp46/48 (RBBP4) [142].
Interestingly, RbAp46/48 is a component of several repressor complexes including PRC2 [84].
Recent studies have shown that CHAF1A is required for maintaining H3K9me3 epigenetic
modifications in heterochromatin regions after DNA replication (epigenetic memory) and was
identified in an siRNA screen aimed to identify factors required for silencing of a randomly
integrated vector in HeLa cells [143-145]. PCNA is a DNA polymerase accessory protein which
interacts with CHAF1A at the replication fork. There is growing evidence implicating both
PCNA and CHAF1A in gene silencing [146, 147]. We found that knock down of both of these
14
factors enhanced responsiveness of CIITA and a subset of other PRC2-regulated ISGs as well as
the basal levels of non-ISGs in SW13 cells. We have now evidence indicating that CHAF1A
controls CIITA expression by proper transferring of PRC2- dependent repression mark,
H3K27me3, during S-phase which is discussed in Chapter3.
In the next section, published as a chapter in “SYSTEMS ANALYSIS OF CHROMATIN-
RELATED PROTEIN COMPLEXES IN CANCER”, we review in detail the current knowledge of
histone chaperones and epigenetic memory [148]. This section was written together with my
supervisor, Rod Bremner.
15
1.5.2 Histone Chaperones, epigenetics and cancer
The first histone chaperone was isolated as a nuclear protein that prevents histone precipitation
and facilitates nucleosome formation [149]. Chaperones are typically acidic, ideal for binding to
basic, positively charged histones [150]. They bind histones in the cytoplasm, carry them into
the nucleus and facilitate nucleosome formation (Fig 1.1) [151, 152]. Chaperones were
originally thought of as simple carriers that prevent nonspecific DNA-histone interactions, but
are now known to regulate histone disassembly and reassembly, covalent histone and DNA
modifications, and the maintenance of epigenetic and chromatin states during replication,
transcription and DNA repair.
Classically, epigenetics is “the inheritance of variation (-genetics) beyond (epi-) changes
in the DNA sequence” [153], but the term is now used commonly to describe any modification
of a nucleosome, even in post-mitotic cells where “inheritance” is by definition a misnomer.
When DNA is replicated or repaired, cells “remember” which genes were silent or active and
nucleosomal positioning and covalent histone modifications are crucial for this process.
Chaperones redistribute parental histones and deposit new histones on replicated DNA and thus
they also have a profound role in epigenetic memory. Understanding this process is important
because epigenetic memory is perturbed in multiple human diseases, including cancer as we
review here for histone chaperones.
1.5.3 Nucleosome replication: Conservative vs. Semiconservative
Nucleosomes consist of a core (H3-H4)2 tetramer and two H2A-H2B dimers attached on
either side, with 146 bp of DNA wrapped around the entire octamer. Histones as well as DNA
are replicated, and must be incorporated into new nucleosomes and covalently modified to
maintain the chromatin state. To understand how histone chaperones operate it is important to
first understand nucleosome replication. For simplicity, we will limit our discussion on
replication of the (H3-H4)2 tetramer to the three non-centromeric mammalian somatic H3
16
variants, H3.1, H3.2 and H3.3 that combine with an invariant H4 [154]. H3.1 and 3.2
show similar properties and we will refer predominantly to the more highly expressed H3.1
isoform.
Nucleosomes could, in theory, follow conservative or semi-conservative modes of
replication. In the former model, old (parental) histones are transferred intact to one of the two
replicated DNA strands, and new histones are used to generate a second nucleosome at the
matching position. In semi-conservative replication, the parental histone octamers are split, and
mix with new histones to create two hybrid nucleosomes. An elegant study suggests that in
HeLa cells H2A-H2B dimers are always replicated semi-conservatively (new mixed with old),
but H3.1-H4 tetramers undergo conservative replication [155]. Inducible Flag-tagged Histone
H3.1 plus growth in normal lysine (K0) media was used to label old nucleosomes, then Flag-
H3.1 expression was extinguished, cells were arrested at G2/M, released and grown in heavy
lysine isotope (K8) to label new histones through the next S-phase. Mononucleosomes with old
Flag-tagged H3.1 were purified, and Mass Spectrometry (MS) used to analyze gel purified
histones. Subtracting background label, only 2% of H3.1 or H4 in Flag-tagged (i.e. old)
nucleosomes contained K8 (i.e. new) label, and this did not increase much even after a second S-
phases. Thus, there is almost no splitting of (H3.1-H4)2 during S phase, indicating conservative
replication. In contrast, H2A and H2B in Flag-tagged nucleosomes were ~ 50% labeled with K8
after one S-phase, indicating semi-conservative replication. The latter is consistent with the fact
that H2A-H2B dimers are not in contact in the nucleosome, but reside either side of the tightly
bound pair of H3-H4 dimers [156].
The replication/deposition of (H3.3-H4)2 tetramers is different from that of (H3.1-H4)2
tetramers. Human H3.1 or H3.2 differ at only 5 or 4 amino acid positions from H3.3,
respectively, but whereas their expression is restricted to S-phase, H3.3 is expressed throughout
the cell cycle [157]. Also, while Asf1 (anti silencing function 1) and CAF1 are the chaperones
that deposit new H3.1-H4 dimers, HIRA (histone regulator A), DAXX (death associated protein
six) or DEK [158] chaperones deposit new H3.3-H4 dimers [159-163]. Consistent with separate
deposition pathways, H3.1/3.2-H4 dimers are not found with H3.3-H4 dimers in nucleosomes
[160]. Using the labeling system discussed above, Xu et al found that ~6% and ~20% of
nucleosomes with old Flag-tagged/K0 H3.3 contained new K8-labeled H3.3 after one or two S-
17
phases, respectively [155]. This (H3.3-H4)2 tetramer splitting occurs during DNA synthesis but
not during replication-independent deposition, as it is greatly reduced when S-phase is blocked.
In summary, H3.1-H4 tetramers do not split but undergo conservative replication, some
H3.3-H4 tetramers split and undergo semi-conservative replication, and new H3.3-H4 complexes
added outside S-phase are deposited as tetramers and do not mix with H3.1-H4 dimers.
1.5.4 Chaperones regulate transport, modification, and deposition of new histones
Newly generated histones must avoid aggregation, move into the nucleus, accumulate
post-translational modifications, and form nucleosomes on DNA. Chaperones play crucial roles
in all these processes. A recent study combined protein purification, biochemical reconstitution
assays, and RNAi/genetics to deduce a comprehensive view of the early stages of histone
synthesis and transport into the mammalian or yeast nucleus (Fig 1.1) [164]. Here, we highlight
the mammalian process.
As H3.1 emerges from the ribosome its folding is assisted by HSC70 [164]. H3.1 is
monomethylated at this early stage on lysine 9 (K9me1), although the mechanism is unclear
[165]. The newly synthesized non-replicative histone H3.3 is typically dimethylated (K9me2) or
exhibits acetylation (K9ac and K14ac) [165].
From HSC70, H3 is passed to HSP90 which cooperates with the chaperone tNASP1
(“testicular Nuclear Autoantigenic Sperm Protein) to assemble H3.1-H4 dimers (Fig 1.1) [164].
Through splicing the NASP gene also encodes “somatic” NASP (sNASP) [166]. H3-H4 dimers
in the tNASP-HSP90 complex are passed onto sNASP, which also recruits the HAT1 (histone
acetyl transferase 1) holoenzyme, made up of HAT1 and the chaperone RbAp46 (RBBP7) [164].
RbAp46 and its fly homolog p55 bind histone H4 [167] and sNASP preferentially binds histone
H3 [168], providing a logical explanation as to how histone dimers are held by the sNASP-
RbAp46 dual chaperone complex. When H3-H4 dimers are passed to the sNASP-RbAp46
complex a dramatic reduction in H3K9me occurs by an unknown mechanism, and HAT1
18
catalyzes H4 acetylation on K5 and K12 (Fig 1.1). sNASP can dimerize [169], and complexes
have been observed containing one or two H3-H4 dimers, but not tetramers [164] which, as
described later, form in the nucleus (Fig 1.1). In budding yeast (S. cerevisiae) the equivalent of
sNASP is Hif1, and the Hat1p/Hat2 holoenzyme performs acetylation, much like the mammalian
HAT1-Rbap46 complex. HAT1 or Hat1p driven K5ac and K12ac modifications appear to
enhance association with nuclear transporters and thus nuclear uptake [170, 171].
NASP is essential for murine development [166], and plays a key role in adjusting the
soluble reservoir of H3-H4. Reducing NASP leads to autophagy-mediated depletion of soluble
H3-H4, whereas reducing HAT1 (and thus histone acetylation) or Asf1 does not lower cytosolic
H3-H4 [172]. Thus, NASP provides a tunable cytoplasmic source of H3-H4.
After passing from the HSC70 to the tNASP complex and then to the
sNASP/Rbap46/HAT1 complex, the final cytosolic stage involves the transfer of H3-H4 dimers
to a complex containing the conserved chaperone ASF1 and Importin-4. There are two human
ASF1 genes, A and B, but only the B protein is found at this stage, although A likely takes over
when B is artificially removed [164]. Importin-4 is a karyopherin family member that mediates
transport through the nuclear pore. ASF1 acts as a sink for H3-H4 dimers since most non-
nucleosomal nuclear histone H3-H4 is found associated with this chaperone [173].
Once in the nucleus, H3.1-H4 dimers pass from ASF1 to the CAF1 (chromatin assembly
factor 1) complex. H3.3-H4 dimers are handled by HIRA for deposition on genic regions [159,
161], or DEK for an as yet uncharacterized target [158], and DAXX for deposition on
heterochromatin [162, 163, 174]. For simplicity, we will focus on the H3.1-H4/CAF1 interaction
below, but we will also discuss DEK and DAXX in the section on chaperones and cancer.
CAF1 was originally isolated as a human complex that promotes in vitro nucleosome
assembly on replicating SV40 viral DNA templates [142]. It is conserved from yeast to humans
and consists of three subunits termed Cac1-3 in yeast [175], p180, p105 and p55 (NURF) in
Drosophila [176, 177] , or p150 (CHAF1A), p60 (CHAF1B) and p48 in humans [142]. The latter
is also called Rbap48 (RBBP4), a close relative of Rbap46 (RBBP7) mentioned above (see
sNasp/Rbap46/HAT1 complex, (Fig 1.1)). Asf1 can bind directly to the mid-sized subunit of
CAF1 (p60/CHAF1B) [178, 179], while the small subunit (Rbap48) binds both H3 and H4 [180].
19
For many years it was unclear whether newly synthesized histones H3 and H4 form
tetramers prior to deposition by CAF1, or only after deposition on DNA. Three recent studies all
suggest the former. The first found that mutations affecting neuronal fate in C. Elegans mapped
to a C-terminal region of histone H3 required for H3-H4 tetramerization [181]. C. Elegans has
24 histone H3 genes, but mutation of only one acted as a dominant negative to block
nucleosome formation. Depleting CAF1 or PCNA (required to recruit CAF1 – see below)
caused the same neuronal phenotype, whereas depleting Asf1 did not alter fate. These data
suggest that the phenotype results from an inability of CAF1 to assemble H3-H4 tetramers. The
second piece of evidence came from cross-linking studies which found that a single molecule of
yeast CAF1 binds H3-H4 tetramers [182]. And the third involved thermodynamic studies
showing that once yeast ASF1-H3-H4 binds to CAF1, ASF1 is dislodged and a second H3-H4
dimer is introduced to form the H3-H4 tetramer (Fig 1.1) [183].
The latter study also found that the affinity of CAF1 for unmodified H3-H4 dimers is ~2-
fold lower than that of ASF, raising the question of how the transfer could be thermodynamically
favorable. Notably, however, post-translational histone marks increase the affinity of histones
for downstream chaperones [184]. In the nucleus the yeast or mammalian acetyl transferases
Rtt109 or CBP/p300 catalyze H3K56 acetylation, respectively, which is stimulated considerably
by Asf1 or the yeast histone chaperone Vps75 [185-187], and this modification promotes
association of H3 with CAF1 or the yeast histone chaperone Rtt106 [188]. Yeast lacking Asf1 or
H3K56 acetylation have a reduced life span [189]. Another modification that influences binding
is PAK2-mediated phosphorylation of H4 Ser 47, which diverts H3.3-H4 dimers away from
CAF1 to HIRA [190]. Thus, chaperones facilitate histone modification and/or exploit these
events to guarantee the passage of histones along the chaperone chain in the right direction and
with the correct partners. Several other marks have been described on newly synthesized
histones, several of which affect chromatin assembly and sensitivity to DNA damaging agents,
but the underlying mechanisms are largely unclear [191].
20
The final step in the journey of a new H3-H4 tetramer is deposition onto its highest
affinity partner, DNA. For the CAF1-(H3-4)2 complex, this is facilitated by interaction with
PCNA (proliferating cell nuclear antigen). PCNA forms a trimeric clamp that completely
encircles replicating DNA strands [192]. Its interaction with DNA involves water molecules,
allowing it to glide along the template and improve DNA polymerase processivity (the number
of nucleotides replicated without polymerase dissociation). The large subunit of CAF1
(p150/CHAF1A in humans) binds directly to PCNA [146]. As discussed below, CAF-1 is critical
for the inheritance of heterochromatin, and mutations in yeast PCNA that affect CAF1
recruitment disrupt heterochromatin-mediated silencing [193], and this interaction is also critical
for chromatin assembly after DNA damage [194]. Moreover, dominant negative CAF1 mutants
that do not bind PCNA or p60/CHAF1B disrupt chromatin deposition and lead to activation of
the DNA damage checkpoint [195].
In summary, chaperones guide H3-H4 dimers from their synthesis in the cytoplasm to
their destination on DNA through a thermodynamically favorable chain of binding reactions (Fig
1.1).
1.5.5 Chaperones recycle old histones during replication
Chaperones also disassemble nucleosomes as the replication fork passes, and then reassembles
them on both strands of replicated DNA. The first step in the disassembly of parental
nucleosomes is removal of H2A/H2B. FACT (facilitates chromatin transcription) is an
H2A/H2B chaperone and while most of the work on its function focuses on transcription, it is
also involved in altering chromatin structure during DNA replication [196, 197]. FACT is
recruited to the replication fork through an interaction with MCM4, one of six proteins (MCM2-
7) that make up the helicase that unwinds DNA [198]. Another chaperone that could be involved
21
in removing H2A/H2B from DNA is Nap1 (nucleosome assembly protein 1) which operates in
collaboration with the ATP-dependent chromatin remodeling factor RSC in vitro [199]. Nap1 is
thought to be particularly important for removal of H2A/H2B during transcription [200],
reviewed in [201].
Once H2A/H2B is removed, the more stable H3/H4 can be displaced. H3/H4 dimers are
then assembled onto nascent DNA quickly, whereas histones H2A and H2B are added 2-10 min
after fork passage [202]. Asf1 plays a key role in coordinating the removal of H3/H4 dimers with
their re-deposition behind the fork [203]. Asf1 splits (H3-H4)2 histone tetramers into dimers and
an Asf1-H3/H4-MCM sandwich briefly tethers them to the helicase (Fig 1.1). Knockdown of
both Asf1 genes (Asf1a & b) prevents nucleosome removal, the helicase fails to unwind DNA
(reflected in reduced levels of single stranded (ss) DNA levels at the fork), and cells arrest in S-
phase. Inhibiting DNA polymerase causes an accumulation of parental histones on Asf1,
identifiable by covalent marks absent on newly synthesized histones (H3K9me3/ H4K16ac)
[203]. Asf1 is also required to bring new H3-H4 dimers to DNA (see above [204]), thus when
H3-H4 dimers are over-expressed, Asf1 becomes limiting, parental histones are not dislodged,
the helicase stalls, ssDNA levels drop, and cells arrest in S-phase [203]. This S-phase block can
be rescued by elevating Asf1 levels [203]. Therefore, Asf1 coordinates both recycling of parental
and introduction of new histone H3-H4 dimers.
The above data raise an interesting conceptual problem. As noted earlier, (H3.1-H4)2
tetramers exhibit conservative replication [155]. But Asf1, which plays a key role in tethering old
H3-H4 dimers to the replication fork [203], splits tetramers [150, 205]. Once old H3.1-H4
tetramers are split into dimers, how do they re-associate and remain separate from new H3.1-H4
dimers? Re-association of old dimers might be the only option given that new tetramers are
preassembled on CAF1 (see above). Whether H3.1-H4 dimers from an old nucleosome remain
closely associated to a pair of Asf1 molecules at the fork is unclear. Perhaps Asf1 traffics old
histones through CAF1 at the fork, although in vitro assays show that CAF-I cannot assemble
histones H3 and H4 purified from cellular chromatin onto DNA [142, 206]. Whether this is the
case at the replication fork in vivo, however, is unclear. Nap1 or its close relative Vps75, form
homodimers that adopt an earmuff structure and directly bind the two H3 proteins in an intact
(H3-H4)2 tetramer [207], but whether this feature is exploited during conservative replication of
22
the core nucleosome is unknown. Alternatively, covalent modifications and/or associated
proteins (chaperones?) on old and/or new dimers preclude mixing during deposition.
There are two ASF genes in mammalian cells and while knockdown of both ASF1A and
B is required for acute arrest of cultured cells, removing ASF1B, but not A, blocks growth in
colony formation assays [208]. The reason for this difference is not fully resolved, but ASF1B
has other qualities that distinguish it from 1A, such as its down-regulation in quiescent or
senescent cells, and ASF1B deficiency causes unique effects on the transcriptome, and the
appearance of mitotic defects such as micronuclei and DNA bridges [208].
23
Fig 1.1 Histone chaperone functions during DNA replication. In the cytoplasm (top) HSC70 chaperones newly translated H3, which is methylated by an unknown enzyme. Next, HSP90 and tNASP chaperone H3K9me1, and histone H4 joins the complex to generate H3-H4 dimers. H3-H4 is then transferred to a complex containing the chaperones sNASP and RBAP46, and the enzyme HAT1. H3 is demethylated a this stage by an unknown enzyme, and HAT1 acetylates H4 on K5 and K12. Next, the acetylated H3-H4 dimer is passed to ASF/importin for nuclear import, following which CBP/p300 acetylates H3 on K56. The latter facilitates transfer to the final chaperone in the chain, CAF1, which assembles H3-H4 dimers into (H3-H4) 2 tetramers. CAF1 is tethered to PCNA at the replication fork and deposits the (H3-H4)2 tetramer on DNA, following which H2A-H2B dimers are deposited, for example, by the dimeric chaperone complex FACT, which is tethered at the fork through an interaction with MCM4 in the helicase. At the fork the helicase unwinds DNA (towards the left), and behind it (to the right) polymerases (not shown for simplicity) synthesize the new leading or lagging strands of DNA. Ahead of the helicase, parental H2A-H2B dimers are displaced by FACT, and ASF1 then separates the remaining H3-H4 tetramer into dimers. The resultant H3-H4-Asf1 complex is tethered to the fork through an interaction of the histones with the helicase. H3-H4 is then redeposited on DNA, but it is unclear whether this occurs through another chaperone intermediate (e.g. CAF1 etc), or whether tetramers form directly on DNA after release from ASF1 (see text). The octamer is then completed by addition of two H2A-H2B dimers. Old and new histone H3-H4 tetramers do not mix (as shown), but old and new H2-H2B dimers can mix (here, for simplicity, no mixing is indicated).
24
1.5.6 Chaperones and Epigenetic Memory: The Example of CAF1
Maintenance of nucleosomes during replication or repair is, by definition, a key aspect of
epigenetic memory. However, in addition to passing on old and depositing new histones,
chaperones play additional epigenetic roles. The full extent to which chaperones regulate this
process and the mechanisms therein are largely obscure. Most work has been performed on
CAF1 and ASF1, particularly on maintenance of heterochromatin. Below, we summarize the
data for CAF1 [209].
Heterochromatin is typically rich in repetitive DNA, such as centromeric satellite
sequences and telomeric repeats. These dense domains are gene poor, less accessible, coated
with repressive hypoacetylated histones and histone H3 trimethylated on lysine 9 (H3K9me3),
and replicated late in S-phase. H3K9me3 tethers heterochromatin protein 1 (HP1) which self-
associates to promote condensation [210]. Moreover, HP1 recruits the H3K9 methyl transferases
SUV39H1 and SUV39H2, which propagate this chromatin mark during replication (Fig 1.2)
[211, 212]. The DNA in these dense regions is easily visible upon staining with intercalating
fluorescent dyes such as 4$,6-diamidino-2-phenylindole (DAPI). DAPI-intense heterochromatin
spots remain even during their replication, so how is such densely packed DNA and chromatin
duplicated? As discussed below, CAF1 plays a major role in this process. Indeed, CAF1 is
crucial for heterochromatin maintenance in yeast, plant, fly, frog, mouse and human cells [213-
217].
Links between CAF1 and heterochromatin arose from work in budding yeast (S.
cerevisae) where, although dispensable for survival, its subunits are essential for efficient
silencing of marker genes proximal to telomeres [175, 218, 219]. Initially, it was thought that
silencing at mating type loci did not require CAF1, but more sensitive assays revealed that CAF1
is essential to maintain silencing, but not for re-establishment (e.g. following Sir protein
disruption) [220].
Heterochromatin in S. cerevisae relies on proteins like Rap2 and the Sir family of histone
deacetylases, but different factors are utilized in higher eukaryotes, such as the HP1 family.
25
Remarkably, however, subsequent work provided that a direct link also exists between
heterochromatin maintenance and CAF1 in higher eukaryotes.
This connection came from the discovery that the N-terminus of murine Chaf1A/p150
binds directly to HP1 [219]. In human or murine cell lines, CAF1 colocalizes with HP1 in late
S-phase at sites of pericentric heterochromatin [221]. Labeling with thymidine analogues, such
as BrdU, and 3D imaging showed that heterochromatin is replicated at the surface of DAPI-
dense spots, and is then buried inside the domain after replication [222]. In addition to
trimethylation by Suv39h1/2, HP1 binding to the core of DAPI-rich heterochromatin also
requires an RNA component, but in contrast Chaf1A/p150 tethers an RNase and an Suv39h1/h2-
null resistant HP1 fraction to replicating DNA on the surface of heterochromatin domains in
mouse fibroblasts [222]. These data suggest that when DNA emerges at the surface of a
heterochromatic domain, HP1 is displaced and tethered at the replication fork by Chaf1a. Indeed,
while Chaf1a/p150 or Chaf1b/p60 knockdown reduces nucleosome deposition in S-phase DNA,
only the former causes cell cycle arrest in mid-S-phase, which is not associated with a DDR, but
rather an inability to replicate pericentric heterochromatin [222]. This RNAi-induced defect is
complemented by wild type Chaf1a, but not by mutants that do not bind HP1. Moreover, Chaf1a
knockdown does not arrest Suv39h1/h2 double null fibroblasts where HP1 is absent from DAPI-
rich heterochromatin [145]. Altogether, these data suggest that CAF1 displaces HP1 during
heterochromatin replication, and holds it at the replication fork ready for re-deposition (Fig 1.2).
CAF1-HP1 [222] exists separately from CAF1-H3.1-H4, thus a single CAF1 complex
deposit newly synthesized histones or handles old/new HP1 separately. CAF1-HP1 binds other
proteins critical for heterochromatin maintenance, including Methyl Binding Domain 1 (MBD1)
[223], which recruits the H3K9 methyltransferase SETDB [224]. While bound to CAF1,
SETDB1 stimulates mono-, but not di- or tri-, methylation of H3K9 [225]. Monomethylated K9
is an excellent substrate for di and tri methylation by Suv39h1 (Fig 1.2).
26
In summary, CAF1 performs many functions relevant to the epigenetic inheritance of
heterochromatin (Fig 1.2): i. As at other loci it loads new histone H3.1-H4 tetramers onto
replicated DNA; ii. It displaces and re-deposits old parental HP1 at the surface of
heterochromatin domains, and brings in new HP1 to maintain heterochromatin on replicated
DNA; iii. CAF1-HP1-MDB1-SETDB1 mono-methylates H3K9 on new nucleosomes, allowing
Suv39h1/h2 - tethered to old nucleosomes – to convert H3K9me to H3K9me3, and the latter can
now receive old or new HP1 from CAF1.
As well as maintenance, CAF1 is also important for the de novo formation of murine
heterochromatin as Chaf1a null mouse embryos arrest at the 16 cell stage and fail to form DAPI-
rich spots [226]. In ES cells, where chromatin is more plastic than in differentiated cell types,
Chaf1a knockdown does not cause arrest, perhaps because HP1 is easier to displace in these cells
than fibroblasts. However, these depleted ES cells die after 4 days of knockdown [226], perhaps
because the failure to form heterochromatin disrupts chromosome segregation, although the latter
was not tested. Whether this survival function for CAF1 depends on interaction with HP1 has
not been tested.
The above model of Chaf1-HP1 mediated regulation of heterochromatin was established
primarily in mouse cells. The picture is less clear in other animal species/cell types. In human,
mouse, and chick cells removing Chaf1a/p150 or Chaf1b/p60 impairs nucleosome deposition in
S-phase [145, 195, 227-229]. In chick DT40 cells, like mouse fibroblasts, there is no DDR when
CAF1 is disrupted (indeed the response to UV or HU is dampened) [229]. However, whereas
only Chaf1a knockdown perturbs S-phase progression in mouse cells, knockout of Chaf1a or
Chaf1b delays S-phase in chick cells, and also causes extensive cell death by 48 hrs [145, 229].
Binding of chick Chaf1a to PCNA and Chaf1b is required for survival, but the interaction with
27
HP1 is dispensable [229]. Whether Chaf1a HP1-binding mutants affect heterochromatin
replication in chicks is unknown. Unlike chick/mouse cells, CHAF1A small interfering RNA
(siCHAF1A) causes a DDR in the human cancer cell lines RTK, HeLa, and U2OS cells [195,
227], as does siCHAF1B in HeLa cells [228]. Viability was compromised in both the
U2OS/siCHAF1a and HeLa/siCHAF1B assays. Intact heterochromatin is important for proper
nucleation of spindles at mitosis, thus cell death in some of these scenarios might follow
disruption of this key process, but may also be the result of an S-phase DDR in some cases.
However, neither scenario applies in chicks as there is no DDR, and HP1 binding is dispensable
for survival. The extent to which the above variable responses to CAF subunit disruption reflect
differing chaperone redundancy, species/tissue specificity, and/or degree of neoplastic
transformation is unclear. Notably, an RNAi screen in HeLa cells identified CHAF1A as critical
to maintain silencing of an integrated GFP reporter gene [230]. Other heterochromatin
regulators were also identified, such as HP1 and SETDB1, consistent with the model discussed
above.
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Fig 1.2 Multiple roles for CAF1 at heterochromatin. Solid arrows indicate protein movements, dotted arrows indicate methylation events. Old and new nucleosomes, PCNA and DNA are shown as in Fig 1.1 i. CAF1 tethered to PCNA deposits new H3-H4 tetramers onto DNA (see Fig 1.1 for details). ii. A separate CAF1 complex tethers SETDB1 to the fork through an interaction with MBD1, and this enzyme monomethylates H3K9. This CAF1 complex also contains new HP1 to deposit on trimethylated H3K9. iii. SUV39H1, tethered to HP1, di and trimethylates H3K9. iv. New HP1, tethered to CAF1, is deposited on H3K9me3. v. SUV39H1 is transferred to newly deposited HP1. vii. Old (parental) HP1 is transferred to a third CAF1 complex at the fork. vii. Old HP1 is transferred (in this example) onto old, already trimethylated H3K9 (see Fig 1.1 for details on how old nucleosomes are transferred at the fork). viii. SUV39H1 is transferred to old HP1. It is likely that old and new HP1 is distributed randomly among old and new nucleosomes, although this has not been tested. The top right illustrates two complexes of CAF1 bound to one PCNA trimer, which is theoretically feasible, but whether this occurs, or whether CAF1-(H3-H4)2 and CAF1-MBD1-SETDB1-HP1 complexes bind sequentially (i.e. separately) is unknown. New and old SUV39H1/2 is not differentiated, but new enzyme is likely drawn from the nucleoplasm.
29
1.5.7 Other potential roles for CAF1 in maintaining repressive chromatin states
Beyond its well established role in regulating HP1 bound heterochromatin, CAF1 may regulate
other repressive mechanisms. The Polycomb group (PcG) of proteins represses transcription at
multiple loci, including homeobox transcription factors required for specific developmental fates
[23]. Heterozygous mutations in the large Drosophila CAF1 subunit (p180, equivalent to
p150/CHAF1A in humans) enhance the effect of heterozygous mutations affecting the PcG
protein, Pc [216]. This genetic evidence suggests that CAF1 may facilitate the epigenetic effects
of PcG complexes. But how this operates, and the specific PcG complexes affected are unknown.
The CAF-1 subunit, RBBP4 (Rbap48) or p55 (Nurf55) in Drosophila, is also a subunit of
Polycomb Repressive Complex 2 (PRC2) *, but these functions are separate because structural
studies show that Nurf55 uses the same region to interact with histones H3 and H4 (as part of its
role in CAF1) and with SUZ12 (a component of PRC2) [180]. There is no biochemical evidence
that CAF-1 and PcG interact directly, but conceivably CAF-1 might recruit other proteins to the
fork that influence PcG activity. Apart from well known interactions with PCNA, ASF1,
histones, and HP1-MBD-SETDB1 (see above), CAF1 binds several other proteins [231], but
their role in epigenetic inheritance is unclear.
1.5.8 CAF1 and Cancer
In view of their central role in regulating nucleosome density, histone modifications, chromatin
structure, genome stability, and epigenetics it seems logical that cancer cells might manipulate
histone chaperones to promote mutagenesis and/or alter gene expression. The field is young, and
while most links are indirect, there are some striking examples of how chaperones influence
cancer progression. We review some key examples below.
Disrupting the CAF1 specific subunits p150 (CHAF1A) or p60 (CHAF1B), or their
equivalents in lower organisms, disrupts nucleosome frequency, and, in many cases, causes
30
spontaneous DNA damage and/or increased susceptibility to DNA damage inducing agents [175,
195, 227]. Moreover, because CAF1 regulates heterochromatin, disrupting this function could
lead to large scale defects in chromosome alignment and segregation at mitosis. However,
inactivating mutations in CAF1 subunits have not been reported in cancer, likely because their
depletion is lethal in many circumstances [226, 228, 229]. Nevertheless, subtle sequence
variants, changes in post-translational modifications and/or altered levels of chaperones could
affect genome stability or gene expression. Indeed, excess CHAF1B/p60 correlates with poor
outcome in some cancers [232]. The genetic connection between CAF1 and Polycomb
phenotypes in Drosophila is also intriguing since excess Polycomb activity is common in human
tumors [23].
1.5.9 ASF1, HIRA and cancer
As discussed earlier, ASF1A and B are critical to buffer excess histones during replication stress
[233], and disrupting this process promotes a delayed DNA damage response [208]. In addition,
ASF1B has unique roles in long-term growth, gene regulation and chromosomal stability, and
elevated ASF1B expression is associated with high mitotic index, poor tumor grade and worse
outcomes in breast cancer [208]. Also, ASF1A is part of a network of factors critical to repress
the pro-apoptotic gene Fas in K-ras transformed cells [234]. ASF1 stimulates H3K56 acetylation
to promote transfer to the CAF1 complex and subsequent deposition on DNA (see above). This
histone modification promotes longevity in yeast [189], and is elevated together with ASF1 in
several cancer cell types [187]. H3K56ac is also important at sites of DNA repair [235].
Normally, this mark is rapidly removed upon H3 deposition, but is maintained at repair sites to
increase nucleosome “breathing” and facilitate repair complex access to DNA [236, 237]. The
connection between ASF1, CAF1 and H3K56ac provides a logical explanation for increased
DNA damage in the absence of one or more of these components [235].
The above data imply oncogenic roles for ASF1. Intriguingly however, ASF1A together
with the H3.3-H4 chaperone HIRA is also critical for Ras-induced senescence, an anti-cancer
response to DNA damage that forces permanent cell cycle exit [238]. Cancer cells overcome
31
senescence by inactivating the RB and p53 tumor suppressor pathways, and it is presumably
beyond this point that the putative oncogenic roles of ASF1A discussed above come into effect.
The ability of ASF1 to promote senescence is intimately linked with the histone
chaperone HIRA. Normally, HIRA deposits H3.3 at genic or telomeric regions [161], but in
senescing cells it is required for the formation of Senescence Associated Heterochromatic Foci
(SAHF) that silence RB-E2F regulated cell cycle genes [239].
Early in senescing cells, PML bodies, also linked to senescence, contain HIRA and HP1,
and although ASF1A is not found in these structures, a HIRA-ASF1A interaction is critical for
SAHF formation [238]. Late in senescence the repressive H2A variant macroH2A (which also
has a separate role in silencing the X chromosome) is recruited to SAHF, which also requires the
HIRA-ASF1A interaction. ASF1B cannot bind HIRA and is not involved in senescence. Neither
HIRA nor ASF1A bind macroH2A, thus the chaperone for the latter in senescing cells is unclear,
although APLF (aprataxin and PNKP like factor) chaperones macroH2A after DNA damage
[240]. Whether HIRA-ASF1A dependent macroH2A deposition is linked to H3.3 and/or the
ability of ASF1 to dislodge pre-existing nucleosomes (see above) is unknown. In yeast, where
there is no macroH2A, interaction of the HIRA (Hir1 and Hir2) and Asf1 equivalents promotes
silencing at telomeres and repression of histone gene expression [241-245]. Consistent with its
pro-senescence function, low levels of ASF1A are associated with longevity in humans [246],
but as noted earlier, Asf1 is actually required for longevity in yeast [189].
1.5.10 DEK, DAXX, H3.3 and Cancer
Apart from HIRA, DEK and DAXX also chaperone H3.3, and strikingly both are mutated in a
variety of tumor types, providing direct evidence linking chaperones to cancer.
DEK is a highly abundant protein (almost as abundant as nucleosomes) with several roles
in regulating chromatin structure. The Drosophila ecdysone hormone binds the nuclear receptor
EcR and its ability to promote gene activation depends on DEK [247]. In the same study, DEK
was shown to promote nucleosome formation by chaperoning H3.3, which was dependent on
32
CKII phosphorylation of DEK. Disrupting the latter blocked H3.3 binding, nucleosome
formation in vitro, and ecdysone mediated gene activation.
DEK also has roles in repressing transcription. Thus independent of CKII (and thus its
H3.3. chaperone function), DEK promotes heterochromatin through an RNA-dependent
interaction with HP1. Loss of DEK results in the concomitant depletion of HP1 and H3K9me3
from either constitutive pericentric heterochromatin or promoters silenced by the latter repressive
epigenetic mark [158]. Fruit fly DEK is in the same class as the H3K9 methyl transferase
Su(var)3-9, because its inactivation relieves position effect variegation (PEV), the repression
linked to translocation of a previously active gene to a location near heterochromatin [158].
Thus, both CAF1 and DEK have key roles in binding HP1 and regulating chromatin.
In line with a repressive function for DEK, it restricts access of complexes required for
transcription to chromatin templates in vitro, and it is dislodged by SET, a protein required for
Pol II-mediated transcription on such templates [248] . Intriguingly, SET, like DEK, is found as
a CAN/NUP214 fusion protein in leukemia [249].
DEK is linked to cancer in two ways. First translocations have been found in a subset of
AML that fuse its N-terminus to the C-terminus of the nucleoporin CAN (NUP214) [249]. This
translocation switches the location of CAN from the nuclear pore to the nucleoplasm [250]. The
DEK-CAN leukemic fusion protein cannot bind CKII, abolishing its histone chaperone activity,
arguing that defects in H3.3 deposition may be important for transformation [247], although the
critical targets are unclear. The effect of the translocation on HP1 or SET activity is also unclear.
Second, DEK over-expression is observed in various cancers, either through gain of its
chromosomal location 6p22 [251-254] or through induction by E2F, such as in cells expressing
the RB-inactivating human papilloma virus E7 oncoprotein [255, 256]. DEK over-expression
elevates H3K9me3 at silenced chromatin, which could be oncogenic in some contexts [158].
However, the effect of DEK over-expression on chaperoning H3.3 or the ability of SET to open
chromatin is unknown. DEK influences a variety of cancer hallmarks, such as survival,
senescence, DNA repair and invasiveness [251, 256-260], but exactly which of these biological
functions, if any, require its activating and/or repressive molecular chromatin roles remains
unclear. Complicating matters further, DEK has also been linked to other activities, such as
splicing and protein translation [261-263].
33
DAXX was first identified as a FAS-binding protein and interacts with other factors that
regulate cell survival [264]. Three recent studies have shown that it is a histone H3.3 chaperone
[161-163]. Drane et al report that DAXX is more tightly associated with H3.3 than HIRA, and
that it promotes nucleosome deposition on plasmids in vitro, which was also demonstrated by
Lewis et al. Consistent with prior work [265, 266] , DAXX and the SWI/SNF-like chromatin
remodeling enzyme ATRX (#-thalassemia mental retardation X-linked syndrome) were found at
pericentric satellite repeats. These repetitive regions are transcribed, which Drane et al found
requires DAXX, H3.3 and ATRX. DAXX-mediated deposition of H3.3 on pericentric DNA is
replication independent and H3.3 co-localizes with DAXX and ATRX in PML bodies, which is
blocked if DAXX is missing. DAXX/ATRX-mediated recruitment of H3.3 to PML bodies and
pericentric heterochromatin is reminiscent of the requirement for HIRA to transport H3.3 to
PML bodies and senescence associated heterochromatin (see above).
Goldberg et al [161] focused on the genome-wide localization of H3.3, demonstrating
that it is enriched at known regulatory elements (active or poised, proximal or distal), across the
bodies of active genes, and at telomeres. HIRA was essential for enrichment at promoters and
gene bodies, but not at remote regulatory elements or telomeres. Like Drane et al they found
that H3.3 associates with DAXX/ATRX, and while ATRX was dispensable for H3.3 enrichment
at genic regions or regulatory elements it was essential at telomeres. Moreover, Lewis et al
found that DAXX, like ATRX, is essential for H3.3 deposition at telomeres [163]. Finally,
ATRX loss elevates transcription of the telomeric repeat-containing transcript TERRA [161].
Thus, paradoxically, DAXX/ATRX/H3.3 recruitment is linked to induction or down-regulation
of pericentric or telomeric transcription, respectively [161, 162]. The structure of DAXX bound
to a H3.3-H4 dimer has been solved and reveals specific interactions that explain why this
chaperone preferentially binds H3.3 rather than replicative H3 isoforms [174].
Recent deep sequencing efforts have exposed direct evidence for a link between
DAXX/ATRX and cancer. First, mutations in DAXX or ATRX were reported in almost half of
pancreatic neuroendocrine tumors [267]. Consistent with the key role of DAXX/ATRX in
depositing H3.3 at telomeres these mutations or loss of nuclear DAXX/ATRX are linked to
abnormal telomere structures, and analysis of other cancers with aberrant telomeres exposed
ATRX, although not DAXX, mutations in some brain tumors [268]. A subsequent study found
both ATRX and DAXX mutations in pediatric glioblastoma samples, but also a high frequency
34
of H3.3 mutations that target Lysine 27 or Glycine 34 [269]. These variants were linked to
perturbed telomeres and altered transcription profiles, which is consistent with the H3.3
chaperone function discussed above, and with the link between lysine 27 methylation and
Polycomb-mediated repression. Finally, DAXX mutations have also been described in a subset
of AML patients [270].
1.5.11 Future directions
The term chaperone conjures up rather pedestrian images, yet these proteins have diverse
functions and their deregulation has drastic effects on cell homeostasis. The last decade has seen
dramatic advances in our understanding of their roles in replicating nucleosomes, histone
exchange, deposition of histone subtypes, epigenetic inheritance, transcription and DNA repair,
yet there remain many unanswered questions. How, for example, are old and new (H3-H4)2
tetramers kept apart at the replication fork? What precise mechanisms are used to replicate old
H2-H2B dimers or deposit new dimers? Why does CAF1 perturbation have such different
effects in distinct species/cell types on DNA damage or survival? Which functions of CAF1
(e.g. recruiting H3-H4, HP1, SETDB1 etc) are critical in these various contexts? What is the
molecular link connecting CAF1 to Polycomb function, and how is this function played out in
organisms other than Drosophila? The ties between histone chaperones, cancer and longevity
are also fascinating, but we only have a superficial understanding. What is the relevant
molecular effect in these biological contexts: DNA damage, gene expression, Polycomb
function, heterochromatin, and/or another role, and what are the key molecular players that
chaperon influence? What are the signals that switch HIRA from regulation of genic or
telomeric regions to silencing of cell cycle genes destined to be buried in SAHF? Or that control
the ability of ASF1 to promote senescence in collaboration with HIRA, versus silencing of pro-
apoptotic genes? And what are the precise functions of DEK, DEK fusions, and DAXX that are
so critical in driving cancer progression? These are only a few of the questions that will keep
researchers busy in the chaperone field for the next decade.
35
1.6 Hypothesis
Previous work in our lab showed that in the BRG1-deficient SW13 adenocarcinoma cells, PRC2
blocks IFN!-induction of CIITA and a subset of other ISGs. We detected the PRC2 signature
epigenetic mark, H3K27me3, across the CIITA locus and at the promoters of other PRC2-
regulated ISGs. We hypothesized that there should be unknown components that collaborate
with PRC2 in repression of ISGs. The knowledge of what these factors are and how they
orchestrate the silencing of ISGs may improve our understanding of PRC2 mediated epigenetic
memory in general. To address this issue, we performed siRNA screens using a BAC-CIITA
vector containing all the remote elements crucial for CIITA regulation. Here we present an
unexpected novel link between CHAF1A and PRC2. We show that siCHAF1A rescues PRC2-
repressed genes in a cell cycle dependent manner. Interestingly, siCHAF1A does not lead to a
loss but a redistribution of H3K27me3 mark. Therefore, we believe that CHAF1A coordinates
repressive epigenetic memory by regulating the genomic positions at which PRC2 methylates
histone H3K27 during DNA synthesis.
36
Chapter 2
2 RNAi Screens to Identify new components regulating IFN! responsive genes (ISGs)
37
2.1 Abstract
Interferon-! (IFN!) signaling is a crucial pathway in the cells of the immune system and its
aberrant regulation is linked to many diseases including cancer. Therefore, identifying its
regulators is of much interest. The IFN! signaling pathway is initiated by the binding of a ligand
to the IFN! receptor which activates the signal transducer and activator of transcription (STAT1)
through the Janus (JAK1/JAK2) kinases. This in turn leads to the induction of IFN! stimulated
genes (ISGs). We have previously shown that Brahma-related gene 1 (BRG1), the ATPase that
drives the SWI/SNF chromatin remodeling complex, is essential for the expression of class II
transactivator (CIITA), an ISG and the master regulator of the class II major histocompatibility
complex (MHC II). For example, in the BRG1-deficient human adrenal carcinoma SW13 cell
line, the CIITA locus is unresponsive to IFN!. Surprisingly, however, a short CIITA reporter,
containing only the IFN!-responsive promoter IV (PIV) was BRG1-independent. These
observations led us to search for and identify remote elements, several kbs away from the
transcriptional start site that block IFN!-responsiveness of the CIITA locus in BRG1-deficient
cells. We also reported that BRG1 is normally recruited to these elements and drives the
chromatin conformational changes that counter the inhibition of CIITA expression. Our lab built
a 194 kb BAC-CIITA luciferase (Luc) reporter containing all the newly identified remote
regulators and showed that the construct is, like the endogenous CIITA locus, also BRG1-
dependent. Here, we took advantage of this tool and performed high throughput (HTP) siRNA
screens to determine: A. whether we could rescue CIITA responsiveness in BRG1-deficient cells
by suppressing factors that repress this target (Suppressor screen) and B. which additional
regulators (activators or inhibitors) affect CIITA regulation in the presence of BRG1 (Wild type
screen). In this chapter, the optimization procedures for the different steps of the siRNA screens
are discussed and the identified hits are reported.
38
2.2 Introduction
IFN! is secreted by activated immune cells in response to the presence of pathogens and tumor
cells. This triggers protective defenses of the immune system aimed to eradicate pathogens or
tumors [131, 135],3. Since almost all types of cells respond to IFN!, exploring the mechanism by
which IFN! signaling pathway is regulated is of much interest [131, 271]. The IFN! signaling
pathway is initiated by binding of IFN! to its receptors which activates the JAK1 and JAK2
tyrosine kinases[272]. JAK1/JAK2 activation leads to phosphorylation of Signal Transducer and
Activator of Transcription (STAT1) [133, 135]. Once phosphorylated, STAT1 homodimerizes
and translocates to the nucleus and its incorporation with interferon regulatory factor 1 (IRF1)
contributes to the induction of Interferon gamma responsive genes (ISGs) including class II
activator (CIITA), a master regulator of MHC Class II [33, 133, 135, 273]. There are several
feedback mechanisms that control and inhibit the IFN! signaling pathway. For example, the
suppressors of cytokine signaling (SOCS) proteins, a family of SH2-containing proteins are
crucial negative regulators of cytokine signaling transduction [274, 275].
CIITA is a coactivator that induces the MHC Class II promoter[139]. It is constitutively
active in immune cells such as B lymphocytes, activated T lymphocytes and dendritic cells or is
induced by stimulation in most cell types [276]. CIITA is regulated by four promoters (PI-IV) in
a tissue specific manner [276]. PI and PIII are constitutively active in dendritic cells and B
cells[276]. PIV is activated by IFN! in a variety of non-immune cells [276]. Defects in CIITA
regulation are associated with bare lymphocyte syndrome (BLS), a sever immune disease, and
other human diseases, including cancer, multiple sclerosis, arthritis and myocardial infarction
[138, 277]. Studies show CIITA is often silenced in tumor cells [278, 279]. As mentioned
above, cancer cells can be recognized and eliminated by immune cells through interferon
secretion (immunosurveillance) [131]. CIITA silencing is thought to be a mechanism by which
cancer cells escape immunosurveillance, therefore, understanding the mechanism of CIITA
regulation can be exploited in cancer drug design.
Previous studies in our lab uncovered several cis-acting elements and trans-acting factors
essential for the IFN! responsiveness of CIITA. First, we reported that CIITA responsiveness to
IFN! requires Brahma related gene1 (BRG1), the catalytic subunit of the SWI/SNF family of
ATP-dependent chromatin remodeling complexes [33]. Next, our lab showed that although
39
endogenous CIITA gene induction requires BRG1, induction of CIITA in short reporters, even
though properly chromatinized, is BRG1-independent[33, 115]. Chromatin immunoprecipitation
(ChIP) coupled with genomic tiling arrays (ChIP-chip) identified STAT1 and IRF1 binding sites,
as well as IFN!-induced histone modifications in a >100 Kb long stretch upstream and
downstream of the CIITA promoter [137]. ChIP-qPCR at the CIITA locus confirmed remote
events at -50, -16, -8, +40, and +59 kb, and low signal at intervening sites [137]. Next,
chromosome conformation capture (3C) showed that these functionally relevant hubs loop and
contact each other and also the transcription initiation site, and they are crucial for CIITA
activation[137]. Interestingly, we showed that remote regulatory sites are also involved in
regulating other IFN! responsive genes [137].
To study the function of CIITA regularity distal elements, Zuyao Ni in our lab built a 194
kb CIITA bacterial artificial chromosome (BAC) reporter containing all the remote elements (-
50, -16, -8, +40 and +59 ) regulating CIITA upstream of an IRES-Luc cassette [137]. BRG1-
deficient human adrenal carcinoma SW13 cells were transfected with the BAC vector and
SW13-BAC stable cell clones were selected in G418 [137] . Importantly, the selected clones
showed that IFN!-induced activation of Luc mimics the endogenous CIITA as it is also BRG1-
dependent[137]. Replacing the -50, -16, -8, +40 or +59 enhancers in the BAC blocked
responsiveness to IFN! [137]. These results show that BRG1-dependency at CIITA is conferred
by remote elements. In turn, this model suggests the existence of remote silencers that block
CIITA induction and BRG1 is required to overcome.
Next, to identify the silencers, which confer the BRG1 dependency of CIITA,
Mohammad Abou El Hassan in our lab examined silencing epigenetic marks across the CIITA
locus. He discovered H3K27me3 at remote sites (unpublished data). H3K27me3 modification is
mediated by the polycomb repressive complex 2 (PRC2)[280]. PRC2 contains 4 subunits: the
methyltransferase subunit EZH1/2, SUZ12, EED and histone binding proteins RBBP46/48.
PRC2 was first discovered in Drosophila where it represses Hox genes in multiple lineages, and
is linked to stem cell maintenance and cancer in mammals [281]. Interestingly, Brahma (the fly
homologue of BRG1) counters Polycomb activity [282, 283]. Further, Mohammad showed that
the PRC2 subunits EZH2 and SUZ12 are recruited to remote sites across the CIITA locus.
Introducing BRG1 to the system reduced (albeit modestly) the repressive epigenetic marks and
the recruitment of PRC2. Most importantly, knockdown of PRC2 subunits rescued
40
responsiveness of CIITA and other BRG1-dependent ISGs. Taken together, these data
underscore the collaboration of BRG1 and PRC2 in the regulation of CIITA and suggest that
polycomb complexes regulate IFN! signaling in higher vertebrates.
To better understand the CIITA regulation, we decided to identify additional factors
involved. Importantly, in the recent years, siRNAs and development of the high-throughput
(HTP) genome-wide RNAi-based technology allowed for the siRNAs screen to serve as a
powerful and rapid alternative approach to study loss-of-function phenotypes especially for
characterizing novel pathway components [284-287].
RNAi silencing is a sequence-specific and post-transcriptional gene silencing. It initiates
when RNase III-like enzyme Dicer processes dsRNAs to small 21-23 basepair (bp) siRNAs.
These small siRNAs are then incorporated into the RNA-induced silencing complex (RISC)
[284] which is a multiprotein complex that incorporates siRNA or miRNA as a template to find a
complementary mRNA strand, it activates RNAse and cleaves the mRNA and decreases
production of the protein of interest by mRNA cleavage [284, 288, 289].
A previous siRNA screen to identify new components of JAK/STAT signaling pathway
in Drosophila revealed more than 100 functionally important proteins including signaling
factors, enzymes mediating post-translational protein modifications and transcription factors
[285, 290]. However, these studies used small reporters containing a small promoter of the gene
of interest. As our data shows, long-range elements far from transcription start site may have
crucial role in ISGs regulation. Therefore using small reporters as a read out for RNAi screen
might miss crucial regulatory factors.
In contrast, our BAC reporter incorporates essential remote regulatory elements of CIITA
transcription and mimics that of endogenous CIITA. We took advantage of this tool and
performed RNAi screens to identify new IFN! signaling regulators in the context of the CIITA
locus. We used a clone of SW13 cells, stably transfected with the BAC vector (SW13-CIITA-
BAC)[137]. In this way, we were able to perform two types of screens. A “Suppressor screen”
was performed in the absence of BRG1, which aimed to identify new factors involved in the
repression of CIITA. The second, Wild type (Wt) screen, was performed in the presence of
BRG1, by transducing SW13 cells with BRG1 adenovirus. In this latter screen we looked for
new IFN! signaling components, activators or inhibitors, normally associated with the CIITA
41
regulation. Next, we selected 91 hits from each screen and performed secondary screens to
validate these hits using different siRNAs than the ones used in the primary screen.
We expect that the factors involved in the regulation of CIITA will likely be also used to
regulate other ISGs. Thus, our screens should allow us to deepen our understanding of ISG
regulation and hopefully provide new therapeutic targets for diseases caused by dysfunctions of
IFN! signaling.
42
2.3 Material and Methods
2.3.1 Cell culture and adenovirus
Adenocarcinoma SW13 cells were grown in alpha-Mem media supplemented with 10% FBS.
Cells were treated with human IFN! (0.1 µg/ml; BioSource International). Virus was produced
and SW13 cells were transduced as previously described [115] . The amount of virus was
'titrated' so that BRG1 expression was equivalent to that in HeLa cells as described before [115].
Clone #38 of SW13 cells stably transected with BAC vector (SW13-CIITA-BAC38) was
maintained in alpha-MEM media supplemented with 10% FBS and 500 %g/ml of G418 as
described before [137].
2.3.2 HTP siRNA screening
For our screens we used the Dharmacon Human Genome siARRAY siRNA SMART pool library
based on polymerase based reaction (PCR) templates with an average length of 21 bp for the
screen. The entire library consists of 267 x 96 well plates in which 80 wells per plate (columns
3-12) each contain a pool of four siRNAs targeting one particular transcript (80 wells x 267
plates = 21,121 targets). Columns 1 and 2 are blank and are used for negative and positive
control samples. The library is divided into four sets of siRNAs, representing the kinome (10
plates: 779 targets), G protein coupled receptors (7 plates, 516 targets), other druggable targets
(76 plates, 6022 targets) and the rest of the genome (174 plates, 13,804 targets).
For our purposes we used 6801 siRNAs from the kinome and druggable libraries. 96
well plates were loaded with 2.5%M dsRNAs in 2 %l of siRNA buffer. The first row contained
non-targeting siRNAs as negative controls and the second row contained STAT1 siRNAs as
positive controls. For the robotic transfection, 10%l of serum and antibiotic free media was
added to each 96 plate well containing the siRNAs. The plates were incubated at room
temperature for 5 min. Next, 10 %l serum and antibiotic free media containing 1% dharmafect
lipid #1 was added to each well. The plates were incubated for 20 min at room temp. Next,
10,000 cells in 100%l serum containing media with %10 FBS were dispensed per well using and
automated liquid dispenser. In the WT screen, after 3 days, BRG1 adenovirus was added to the
plates without removing the media. At day 5, cells were lysed and luciferase activities were
43
determined. For the secondary validation screen three 96 well plates were loaded with 91
Qiagen dsRNAs and 2 negative controls (PTBP1 and none targeting siRNA) and 4 positive
control for INF! pathway (JAK1, STAT1, IFNR1, SOCS1). Each plate contained 50nM of the
three different siRNAs targeting one particular gene. Cells were dispensed and transfected as
described for the primary screen. BRG1 adenovirus transfection and luciferase assay were
performed as described above.
2.3.3 BAC construction and reporter assays
Details of BAC construction and SW13-BAC38 were described before [137] . Luciferase assays
were performed using Promega kit as described before[115].
2.3.4 Statistical Methods for Hit Selection
Three methods were used to analyze HTP screen data, B-score, M-score and control base
normalization [291-293]. The B score uses the ratio of a raw value over a measure of variability
to smooth noise due to edge effects, missed well and plate to plate variation [291]. In the M-
score, the data were normalized using a robust form of the Z-score (i.e. Zero mean and unit
variance standardization)[292] whereby the median replaced the mean and the median absolute
deviation replaced the standard deviation. This normalization was termed “M-score”, and is
shown in Equation 1:
The M-score was applied on a plate-wise basis, such that the median and median absolute
deviation was derived for each individual plate. This corrected for variation in overall plate
intensity during the course of the experiment [292, 293]. Control base normalization calculates
the ratio of the luciferase activity in each well to the luciferase activity of negative siRNA loaded
in the first well of the same row.
(1)
44
2.4 Results
2.4.1 The HTP siRNA screen optimization
To optimize the conditions for our HTP siRNA screens, I first wanted to ensure that in the
SW13-CIITA-BAC38 cell line, IFN! induces CIITA expression in the presence of BRG1, and
knocking down of the known CIITA activator STAT1 prevents induction.
Initially, we transfected SW13-CIITA-BAC38 cells with siSTAT1 for 3 days using four
different lipids, Dharmafect1, 2, 3 and 4. Since CIITA is not responsive to IFN! in the absence
of BRG1 in the BRG1-deficient SW13 cells, at day 3, we had to introduce BRG1 to the system
using adenovirus. The cells were transduced with the BRG1 adenovirus without removing the
transfection media and then IFN! was added. Cells were lysed, 24hrs later and luciferase
activity was assessed. I found that transfection of the siSTAT1 with Dharmafect 1, 2 and 4
inhibited Luc activities to similar degrees but Dharmafect 3 showed lower STAT1 knock down
efficiency based on Luc assay (Fig 2.4.1A). Thus, Dharmafect 1 was used in all subsequent
studies. Next, the amount of BRG1 adenovirus for optimal CIITA induction was determined.
Using 1%l of purified virus containing 4x106 PFU showed that more than 90% of cells were
transduced and expressed the GFP reporter (Fig 2.4.1B).
Next, I optimized the concentration of siRNA. I showed that 100 nM and 25 nM of
Dharmacon STAT1 siRNA had the same knock down efficiency (Fig 2.4.1C). In order to
minimize possible off target effects I selected the lower concentration (25 nM) of siRNAs for the
screens. I, then optimized the order of addition of siRNA, adenovirus and IFN!. I tried four
different scenarios (Fig 2.4.2). The strategy that showed maximum IFN!-induction in the
presence of negative control siRNA (siCtrl, which does not target any gene) and maximum
inhibition with siSTAT1 was as follows; Day 1: plate cells, Day 2: add siRNA Day 5: add 1%l
BRG1 virus without removing siRNA, Day 6: add IFN! without changing the media for 24hrs,
Day 7: Luc assay (Fig 2.4.2).
To test the automated HTP version of optimized protocol, half of a 96 well plate was
loaded with siSTAT1 and another half with the siCtrl. Next, the media containing Dharmafect 1
was added to the plate. After 20 min of incubation, SW13-CIITA-BAC38 cells were robotically
added to the plate. After 3 days, BRG1 virus was added to the plate without removing media
45
containing the siRNA. After one day, IFN! was added to the plate and Luc activity was assessed
after 24hrs of IFN! treatment. The result demonstrated that the automated assay was robust, had
excellent signal to noise ratios and was thus appropriate for conducting siRNA screens (Fig
2.4.3).
During the screen optimization step, the hits were analyzed by the “B score”. This
method avoids positional effects by using samples as their own controls [291]. To test this
method in our system, we seeded cells in a 96 well plate, in which only well 7B received
siSTAT1 and the remainder received control siCtrl. Importantly, the top B score was assigned to
the 7B well despite edge effects and variability, confirming the B score as an appropriate method
for hit-identification (Fig 2.4.4).
Our optimized assay was then used to assess BAC-CIITA levels in the presence of BRG1
and IFN! in a pilot screen of 10 plates of the kinome Dharmacon library set. This library
included two obligatory activators of IFN! signaling, JAK1 and JAK2, and one feedback
inhibitor, SOCS1, all of which are known to affect CIITA expression [294, 295]. The first
column of each plate received the negative control siCtrl and the second column contained the
positive control, siSTAT1. The signal showed robust knockdown by siSTAT1, proving effective
transfection in multiple wells on every plate. Importantly, the B-score analysis highlighted all
the embedded positive controls that such as JAK1 and JAK2 that were known IFN! activators.
With a cut off of 3 fold above the standard deviation (SD), the B score identified 4/5 of the true
positives and no false positives (80% sensitivity, 100% specificity), while a cut off 2 fold above
SD, it found 5/5 positives and 4 more unknown positives (100% sensitivity). These data support
the utility of this approach in our hands. Therefore, we decided to use B score with 2SD as one
of the statistical methods to analyze the hits of the rest of the screen. The B scores for the pilot
screen are shown in Fig 2.4.5 A. The pilot screen hits contained 17 IFN! activators that
decreased Luc activity, and 38 inhibitors, which caused induction in Luc activity (Fig2.4.5 B).
Importantly, JAK1 and JAK2 were the top two activators, and SOCS1 was the fifth ranked
inhibitor. JAK3 and SOCS5, neither of which regulates IFN! signaling, had B scores close to
zero (not shown). Interestingly, several of the novel hits in our kinome screen are linked to IFN!
signaling or BRG1 function. For example, STK11 (LKB1) is one of the activator hits which is
known that interacts with BRG1 and is required for BRG1-dependent growth inhibition [296].
FRK is another hit in the pilot screen that is shown is involved in IFN! signaling in islet cells
46
and MAP4K5 (GCKR) which shown that is activated by type I and II IFNs was also identified as
a hit in the pilot screen [297, 298].
These data illustrated the power of our screen to highlight signaling networks to identify
new components of IFN! or those that may cooperate with BRG1 to regulate CIITA. We next
performed HTP siRNA screens with the rest of the siRNA library.
47
Fig 2.4.1 Optimizing siRNA transfection and viral transduction conditions. A. A clone of SW13-BAC38 cells (see text) was transfected with siSTAT1 or siControl using four different transfection reagents (Dharmacon#1, #2, #3, #4) . Next, cells were transduced with BRG1 adenovirus and stimulated with IFN!. Luc assay was performed 24hrs after the exposure to INF!. Similar results were obtained with each reagent, although Dharmafect #1, #2 or #4 worked the best. B. To optimize the amount of BRG1 adenovirus, SW13-BAC-38 cells were transduced with the virus for 24 hrs followed by IFN! treatment for 6 hrs. We found that 1µl of the virus containing 4x106 plaque-forming units (PFU) infected more than 90% of the cells and the highest level of Luc activity occurred after 6hrs of IFN! treatment. For the screen we selected the amount of the virus that provided maximum BAC-CIITA induction (green arrow). C. To optimize the amount of siRNA, we transfected SW13-BAC38 cells with 100nM and 25nM of Dharmacon siCtrl and siSTAT1. Both concentration had the same knock down efficiency. We used 25nM of siRNA for the screen.
48
Fig 2.4.2 Optimizing the timing and order of the screen steps. SW13-BAC38 cells were seeded on 96 well plates and 1 day later subjected to four different protocols as indicated. Optimal IFN!-induced Luc activity and inhibition with siSTAT1 but not siCtrl was achieved with protocol #3.
49
Fig 2.4.3 Automated HTP version of the optimized protocol. SW13-BAC38 cells were robotically seeded in a 96 well plate, half of which contained control or STAT1 siRNA as indicated. The plate was incubated for 3 days and then was transduced with BRG1 virus and treated with IFN! as shown in protocol #3 in Fig 2.3.2.
50
Fig 2.4.4 Testing the “B score” method of hit analysis in the optimized protocol. We performed our optimized robotic protocol (Fig 2.3) for a 96 well plate in which 95 wells received siCtrl, and well 7B received siSTAT1. Then we used “B score” method to analyze the result. Note the edge effect in column 12, and variability among position elsewhere, yet the B score ranked STAT1 siRNAs highest.
51
2.4.5 A. Plot of B scores from the pilot Wt screen. SW13-BAC38 were transfected with Dharamcon Kinome library (10 plates) in the present of BRG1 as shown in Fig2.3.4. The hits were identified by the B score method and 2 threshold lines were used. The blue line represents 3 SD from the mean and the yellow line represents 2 SD from the mean. B. Activators and inhibitors of CIITA induction from the pilot Kinome screen. Hits are ranked according to the B score. Blue color represents > 3 SD; yellow color represents > 2 SD from the mean. Canonical IFN! regulators are highlighted in pink.
52
2.4.2 The primary WT and Suppressor screens
For our siRNA screen we used the Dharmacon kinome and druggable libraries (779+6022=6801
targets). Each of the 6801 dsRNA pools was aliquoted into 96-well plates. Negative control
siCtrl, was aliquoted in the first column of each plate and the positive control siSTAT1 was
loaded on the second column of each plate (Fig 2.4.6). Cells were incubated with siRNAs for 3
days. Next, cells were incubated for another day with BRG1 adenovirus (WT screen) or without
(Suppressor screen) followed by IFN! treatment for 24 hrs. Luciferase activity was then
measured and results assessed by 3 methods to identify hits.
In addition to the B score that we tested in pilot screen, we also used M score and
control-based normalization as explained in the methods section, with the goal of identifying
priority hits that were highly ranked in all three methods. The top 1% of hits obtained with each
of the three methods were selected and hits common to all three methods were ranked #1, those
common in 2 methods were ranked #2, while hits selected by only one method were ranked #3
(Fig 2.4.6A). We ranked 152 activators and 145 inhibitors from the primary WT screen and 114
inhibitor hits from Suppressor screen (Apx1.1-3). Scatter graphs of Wt and Suppressor hits
analyzed by the B-Score method are shown in Fig 2.4.6 B-C. CHAF1A and PCNA, the focus of
further analyses in Chapter 3, were prominent hits in the Suppressor screen.
53
54
Fig 2.4.6 Schematic diagram of the primary siRNA screen. Each of the 6801 dsRNA pools of Dharmacon siRNAs were aliquoted into 96-well plates. SiCtrl and siSTAT1 were loaded to the first and second columns of each plate, respectively. Next, SW13-BAC38 cells were transfected for Wt screen and Supp screens and luciferase activity was measured. The top 1% of hits obtained with each three analysis methods (B score and M score and Control Base) were selected. Further, the hits common to all three methods were ranked #1, those common in B score and M score were ranked #2a, B score and Control Base normalization ranked #2b and common hits in M score and Control Base normalization were ranked #2c. Hits selected by only one method were ranked #3 (3a for B score, 3b for M score and 3c for Control Base normalization). In this way in the primary Wt screen we ranked in total 152 activators, 145 inhibitors and in Suppressor screen we ranked 114 hits, see Apx1.1 -3. B. Wt screen activators and inhibitor hits identified by B-Score. Blue dots above the yellow line are hits based on B-Score method where the cutoff is 2.5 SD above mean. C. Inhibitor hits from the Suppressor screen identified by the B-Score. Hits above the red line which include CHAF1A and PCNA are more than that 9SD above the mean.
55
2.4.3 The secondary WT and Suppressor screens for hits validation
To validate the hits from the primary screens, we performed secondary screens. For these
screens we selected 91 candidates from each WT or Suppressor primary screens. These hits
were the ones identified by all three or at least two statistical methods (M score, B score and
Control Base normalization), and some of the hits that were only identified by one of the
methods. The secondary screens were performed three times using three different Qiagen library
siRNAs for each hit. Importantly, these siRNAs are different from the Dharmacon library
siRNAs. The assays were also performed in 96 well plates like the primary screens. Two wells
of each plate were loaded with negative controls, PTBP1 siRNA, which was not a hit in the
primary screen selection process (shown yellow in Fig 2.4.7), “Qiagene All-star negative
siRNA”, and positive control siRNAs for IFN! signaling, IFNR1, JAK1, STAT1, SMARC4,
SOCS1 (shown orange in Fig 2.4.7). For the secondary screens SW13-BAC38 cells were
transfected with 50 nM of 3 separate Qiagene siRNAs, targeting 3 different sites of mRNA so
that there was one siRNA per gene per plate as shown in Fig 2.3.7, for 3 days. The cells were
then incubated for another day in the presence (secondary WT screen) or absence (secondary
Suppressor screen) of BRG1 adenovirus. After this IFN! was added to the cells and luciferase
activity was measured. Data was analyzed by the M score method. To exclude genes affecting
cell proliferation or survival, the alamar blue assay which stains dead cells, was run prior to
collection of lysates for the Luc assay.
By the M score method of normalization, of the 91 activator and inhibitor hits selected
for the secondary Wt screen, 22 hits (26%) were verified, and in the secondary Suppressor
screen, 11 hits out 91 were verified by at least one siRNA (~10%). Except KPNB1, which was a
hit verified by the secondary Wt screen, none of the other validated hits affected cell
proliferation. The hits were classified, based on PubMed and Genecards, based on their
predicted functions (Apx1.4-6).
56
Figure 2.4.7 Schematic diagram of secondary validation screen. 92 hits (both activators and inhibitors) from the primary Wt screen and 92 hits from the primary Suppressor screen (i.e., inhibitors) were selected for the secondary screen validation. SW13-BAC38 cells were transfected by 50 nM of 3 separate siRNAs for 3 days and were incubated for another day in the presence (secondary Wt screen) or absence (secondary Supp screen) of Ad-BRG1. After 24 hrs IFN! was added to the cells and luciferase activity was measured. To exclude genes affecting cell proliferation or survival the alamar blue assay was run prior to collection of lysates for the Luc assay. The validated hits are listed in Apx1.4-6
57
2.4.4 Tertiary Validation on Endogenous CIITA
In the next step, I tested whether the siRNAs that rescued responsiveness of the BAC-
CIITA reporter in both primary and secondary screens also altered the responsiveness on
endogenous CIITA locus in SW13-deficient SW13 cells. Real time PCR analysis validated the
siRNAs against the histone chaperones CHAF1A and PCNA (for results see Chapter 3).
However, other hits were not validated at the endogenous level (data not shown).
58
2.5 Discussion
IFN! signaling pathway plays essential roles in the proper development and function of the
immune system [139]. Its deregulation is associated with several human diseases including
cancer [131]. IFN!-mediated signaling leads to the activation of ISGs including CIITA, the
crucial co-activator of the MHC Class II locus. This gene is silent or mutated in many cancers
[138, 277]. Likely, the loss of CIITA and MHC Class II is used by tumor cells to escape from
the body’s immune surveillance. Therefore better understanding of IFN! signaling components
and CIITA regulation will help us not only to fill in some of the gaps in this field but will also
lead to an improved drug design.
Previous studies have showed that the regulation of CIITA is highly complex, for
example there are four promoters and long-range enhancers, which loop and contact the
promoters upon the IFN! response [115, 276]. We also showed that the IFN! responsiveness of
CIITA requires BRG1 [33, 137]. Here, I performed siRNA screens to identify unknown
repressors that confer BRG1 dependency of CIITA and to discover additional components of
IFN! signaling that are regulating the CIITA responsiveness to IFN!.
In contrast to the previous siRNA screens to find new components of JAK/STAT
signaling, which used short reporters [285, 290], we used a BAC vector containing distal
enhancers crucial for CIITA induction. The BAC vector behaves liked the endogenous CIITA
locus, meaning that CIITA-BAC is silent in the absence of BRG1 and is responsive to IFN! in
the presence of BRG1 [137]. This allowed us to conveniently perform 2 screens at the same
time. In the first screen, in the presence of BRG1 (WT screen), in which we aimed to identify
novel IFN! activators and inhibitors, we identified 152 potential activators and 145 potential
inhibitors. In the absence of BRG1 (Suppressor screen) we identified 114 potential silencers that
blocked CIITA induction.
As the first step towards validating these hits, we performed a secondary screen for some
of the hits identified from both primary screens. From each primary screen we selected 91 hits to
perform secondary screens. The secondary screens validated 10% and 26% of hits from
Suppressor and WT screens, respectively. Interestingly among the validated hits in the WT
screen, there are some well-known IFN! regulators such as IFN! receptor1 (IFNGR1), JAK1 and
JAK2 and SOCS1, which attests to the ability of our system to detect true positives. I did not
59
follow up on Wt screen hits at the endogenous level of CIITA. However, further assessment of
hits from the BAC-CIITA Suppressor screen using the endogenous CIITA locus revealed that
while siRNAs against PCNA and CHAF1A rescued IFN! -responsiveness, other siRNAs did not.
These observations suggest that while some important aspects of CIITA regulation are
maintained in our reporter lines others have been disrupted. The hits we observed with the
reporter but not the endogenous locus may be false positives or, alternatively, there may be
additional levels of repression exerted upon the endogenous locus that are missing in the
reporter. Therefore, in our screens using this reporter, it would be better to have had more strict
criteria for selecting hits from primary screen. For example, in the B-Score graph showing hits
from Suppressor screen CHAF1A and PCNA are ranked 5th and 7th (Fig 2.4.6C). Thus,
assessment of the top ten hits would have been sufficient to expose the low frequency of true
positives. With hindsight, this would have been preferable to our actual approach of validating
the top 1% of hits with the Luc reporter, and then carrying apparently validated hits forward to
the endogenous gene assay..
Our original goal was to perform a genome-wide screen, but we paused after screening
~25% of the genome to validate which, in view of our low endogenous validation rate, proved to
be a wise decision. In light of our findings, it would be better to pursue genome wide screens
using a read-out of endogenous gene activity, such as the protein product of an ISG, or a reporter
integrated into the endogenous locus. (e.g. Luc or GFP fused in frame to the N- or C-terminus of
an ISG).
Overall, our screens characterized novel and interesting regulators never before
associated with the IFN! network. Considerable additional work is required to define which of
the hits in our WT screen affect endogenous ISG regulation, as well as to define whether any of
the Suppressor Screen hits that did not affect endogenous CIITA actually play a role in
suppressing ISG expression in the absence of BRG1. In the next chapter, we provide extensive
data linking two of the suppressor screen hits to the mechanism of epigenetic regulation by
PRC2. Moreover, our data reveal that this link is of general significance, beyond the regulation
of ISGs. This discovery radically changes our understanding of how epigenetic memory is
maintained.
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Chapter 3
3 CHAF1A Regulates the Distribution of PRC2-mediated epigenetic mark, H3K27me3, During Replication
All figures were generated by Mina Rafiei except Figure 3.4.4B,C,D; and 3.4.8C; and 3.4.9; and
3.4.10 B,C which were generated by Tom Leung. Also some unpublished data mentioned in the
text was generated by Mohamed Abou El Hassan (manuscript in preparation).
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3.1 Abstract
CIITA (Class II Major Histocompatibility Complex, Transactivator) is a crucial interferon
gamma (IFN!) responsive gene (ISG) and a master regulator of MHC class II. Defects in CIITA
regulation lead to severe immune diseases and are linked to other types of diseases including
cancer. In unpublished work, our lab showed that in SW13 adenocarcinoma cells, which is
deficient for BRG1, a subunit of the SWI/SNF chromatin remodeling apparatus, Polycomb
Repressive Complex -2 (PRC2) blocks IFN!-induction of CIITA and a subset of ISGs. We also
detected the PRC2-mediated epigenetic mark, H3K27me3, across the CIITA locus and at the
promoters of other PRC2-regulated ISGs. Re-introducing BRG1 reduces the H3K27me3 mark
permitting access to transcription factors and RNA polymerase, which leads to CIITA induction.
These data reveal antagonism between PRC2 and BRG1 in the context of ISG regulation in
cancer cells. To find other factors involved in this process, we performed an siRNA screen using
a BAC-CIITA vector containing all the remote elements crucial for CIITA regulation (Chapter 2).
Here, we focus on one of the hits found in this screen, Chromatin assembly factor 1, CHAF1A,
part of the CAF-1 nucleosome deposition complex, a replication fork protein and histone
chaperone. We showed that siCHAF1A restores IFN!-responsiveness at the endogenous CIITA
locus. The expression of other PRC2-repressed genes (ISGs and non-ISGs) was also rescued by
siCHAF1A and in a DNA replication-dependent manner. Importantly, removing CHAF1A
reduced the H3K27me3 repressive mark across the CIITA locus and at the promoters of other
PRC2-repressed genes. Surprisingly, siCHAF1A did not affect bulk levels of H3K27me3, but
caused a striking redistribution of the mark. Thus, CHAF1A coordinates repressive epigenetic
memory by regulating the genomic positions at which PRC2 methylates histone H3K27 during
DNA synthesis.
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3.2 Introduction
CIITA (Class II Major Histocompatibility Complex, Transactivator) is an IFN! responsive gene
(ISG) which is required for the expression and induction of MHC class II [133, 299]. CIITA and
MHC class II genes are constitutively expressed in immune cells such as dendritic cells and B
cells but are inducible in most cells in response to IFN! [115, 131, 135, 136]. IFN! is secreted
by activated T and NK cells to inhibit proliferation, virus infection and tumorigenesis [131, 135].
IFN! signaling is initiated by binding of IFN! to its receptors, activating the Janus kinases JAK1
and JAK2 which then phosphorylate Signal Transducer and Transactivator-1 (STAT1).
Phosphorylated STAT1 forms homodimers that are transferred to the nucleus where they
collaborate with Interferon Regulatory Factor1 (IRF1) to induce ISGs including CIITA [132, 300,
301]. Previously, we showed that in spite of normal IFN! signaling, CIITA is silenced in
adenocarcinoma SW13 cells, which are deficient in BRG1, a subunit of the SWI/SNF chromatin
remodeling apparatus, and re-introducing BRG1 rescues CIITA responsive [115, 127, 137, 278].
Since activated immune cells are able to distinguish and destroy cancer cells
(immunosurveillance), it is thought that cancer cells might silence CIITA to inactivate MHC
class II and thus escape immunosurveillance [131, 302]. Identifying CIITA regulators will
improve our understanding of IFN! signaling and facilitate the development of strategies to
battle the diseases associated with CIITA defects. Towards this goal Mohamed Abou El Hassan
in our lab detected Polycomb Repressive Complex 2 (PRC2) components and the PRC2-
mediated repressive epigenetic mark, trimethylated lysine 27 of Histone H3 (H3K27me3) across
the CIITA locus (unpublished data).
Polycomb group proteins (PcG) are chromatin associated complexes that repress
thousands of genes involved in development and differentiation in plants and mammals [303-
306]. Recent findings have linked PcG proteins to cell cycle control, chromosome X-
inactivation, cell fate decision, stem cell differentiation and cancer [307-311]. The core of PRC2
complex in mammals, consists of the methyl transferase Enhancer of Zeste Homolog 2 (EZH2),
or its paralog EZH1, which catalyze H3K27me3, Suppressor of Zeste 12 (SUZ12), and one of
the Embryonic Ectoderm Development (EED) isoforms [280, 312]. There are some other co-
factors such as histone binding proteins, PCL, RBBP4 (RbAp48) and RBBP7 (RbAp46) and
JARID, a JmjC domain containing demethylase, which are not part of the core complex but help
the recruitment and enzymatic activity of PRC2[23, 280]. Also, recent data suggests that certain
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histone and DNA-modifying enzymes (such as DNMT1) and long non-coding RNAs may guide
PRC2 to its gene [23]. Despite the wealth of information on PRC2, some fundamental issues
such as the mechanism as to how the H3K27me3 mark regulates gene silencing, how PRC2-
mediated epigenetic silencing is targeted to the appropriate loci, or how it is remembered during
replication still need to be addressed.
We observed that re-introducing BRG1 into SW13 cells modestly reduced H3K27me3
across CIITA, and facilitated induction of an active H3K4me mark at an upstream CIITA
enhancer (unpublished data). Moreover, BRG1 rescued IFN! responsiveness of this locus[137].
These observations indicate an antagonistic relationship between BRG1 and PRC2 at ISGs.
To gain more insight into the mechanism of PRC2-dependent repression of ISGs, we
performed an siRNA screen using a BAC-CIITA vector containing all the remote elements
crucial for CIITA regulation (Chapter 2). Among the hits identified by the screen was CHAF1A,
a subunit of CAF-1 histone chaperone (HC) [142]. The complex contains three subunits,
CHAF1A (P150), CHAF1B (P60) and RbAp48 (RBBP4) [142, 313]. To date, it is the only
known HC involved in the progression of replication and de novo incorporation of newly
synthesized H3/H4 after DNA replication. CAF-1 is recruited to the replication forks through a
direct interaction between its large subunit, CHAF1A, and Proliferating Cell Nuclear Antigen
(PCNA), the processivity factor for the DNA polymerases [146], which, importantly, was also a
hit in our screen.
Recently, multiple lines of evidence suggest that CHAF1A has a crucial role in
maintaining heterochromatin epigenetic marks (epigenetic memory) in yeast, Drosophila, plants
and mammals[146, 175, 218, 220, 314]. CHAF1A interacts directly with Heterochromatic
Protein (HP1a) and Methyl-CpG-binding domain protein 1 (MBD1) which recruits histone
deacetylases (HDACs) to their binding site [145, 223-225]. Moreover, SET domain Bifurcated 1
(SETDB1), a histone H3K9 methyltransferase forms a complex with both CHAF1A and HP1
during S- phase [224]. In this chapter we addressed whether CHAF1A is also required for
maintaining H3K27me3 during replication, both at ISGs and other PRC2 repressed targets.
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3.3 Material and methods
3.3.1 Cell culture and siRNA transfection
Human adenocarcinoma SW13 cells were grown in #-minimal essential media (#-MEM)
supplemented with 10% fetal bovine serum (FBS). Cells were treated with human IFN! (0.1
µg/ml; BioSource International) [137]. Fifty nM of siRNA were transfected for 3 days using
DharmaFECT-1 (Dharmacon) reagent according to the manufacturer’s instructions. Cells were
split at day 3, fresh media added, and incubated for 24 hrs. On day 4, cells were treated with
IFN! for 6 hrs and were lysate collected for mRNA extraction or Western blotting. SiRNAs
from Dharmacon and Qiagene used in this study are listed in Apx2.
3.3.2 RNA extraction and reverse transcription (RT)
RNA was isolated using TRIzol reagent (Invitrogen). A 2 %g aliqot of total RNA was diluted in
20%l of RNase free water, heated to 90º for 5 minutes, then combined with 30 %l of first strand
master mix [10%g of random Pd(N)6 primers, 72 U RNase inhibitor (Roche# 799-017), 1X first
strand buffer (Invitrogen#18064-014), 1mM dNTPs, 10mM dithiothreitol (DTT), 50U of
Superscript II reverse transcriptase (Invitrogen #18064-014)] and incubated for 1h at 37ºC
followed by 90ºC for 10 min. Next, 200 %l of water was added to the cDNAs to a total volume
250 %l. 4 %l of first strand cDNA was amplified using primers listed in Apx3 by RT-PCR. Copy
number of each cDNA was calculated from Qt values and was normalized to the level of HPRT
of the same sample.
3.3.3 Western Blotting
Cell lysate was prepared from a 100mm plate in 50mM Tris-HCL pH7.4, 150mM KCl, 15mM
NaCl, 30mM MgCl2, 10mM EGTA, 0.5 % NP40 and protease inhibitors. A 100 %g aliquot of
total cell lysate was sonicated and boiled with sample buffer (2% SDS, 100 mM dithiothreitol,
60 mM Tris [pH 6.8], 0.01% bromophenol blue) and separated on an SDS-7.5 % polyacrylamide
gel and transferred to a nitrocellulose membrane. Intensity of protein bands were quantified by
Odyssey. Antibodies used to detect the proteins are listed in Apx4.
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3.3.4 Chromatin immunoprecipitation (ChIP).
6 x 106 Cells per 100mm plate were cross-linked with 1% formaldehyde at room temperature for
10 minutes, washed two times with cold PBS, collected and centrifuged for 5 minutes at 1200
rpm. Cells were resuspended in 1.0 ml lysis buffer (1% SDS, 10mM EDTA, 50 mM Tris-HCL
PH 8) and protease inhibitors (1X aprotinin, leupeptin and pepstatin) from Sigma, were added to
the cell lysate. The lysates were incubated on ice for 10 minutes sonicated to an average size of
500bp using the sonicator (Viba Cell, Sonics and Material Inc, Danbury, USA). Next, chromatin
was pre-cleared with 25 %l of pre-cleared Staph A (Staphylococcal A cells, Calbiochem) for 15
min on a rocker at 4ºC. For immunoprecipitation, 100 %l of pre-cleared chromatin was used with
2%g antibody and incubated at 4ºC overnight. Next, samples were centrifuged and transferred to
new tubes and incubated with 10 %l of Staph A per IP for 15 min rotating at room temperature.
The samples were centrifuged for 5 min and the pellet were washed 2 times with ChIP Dialysis
Buffer (2mM EDTA, 50mM Tris-HCL pH 8, 0.2% sarkosyl), and 4 times with IP Wash Buffer
(100mM Tris-HCl pH 9, 500mMLiCl. 1% NP-40, 1% deoxycholic acid). The samples were then
eluted 2 times using 150 %l of ChIP Elution Buffer (50 mM NaHCO3, 1% SDS). Then, samples
were heated at 65ºC overnight to reverse the cross-links. After reverse cross-linking, DNA
fragments were purified with QIAEXII Gel Extraction Kit (Qiagen, Ca). Next, DNA was eluted
with 10mM Tris-HCl pH 8 at 42ºC for 10 min. The amount of DNA fragments precipitated was
detected by qPCR using selected primers.
3.3.5 Quantitative Real-Time PCR (qPCR)
4%l of each samples were mixed with SYBR Green PCR master mix (Applied Biosystem)
containing each primer and qPCR was performed in duplicate using Applied Biosystem PRISM
7900HT. The program consisted of 40 cycles of 95ºC for 15 seconds and 55ºC for 30 seconds.
A final cycle (95ºC, 15 seconds, 60ºC for 10 sec) generated a dissociation curve to confirm a
single product. A standard curve based on genomic DNA (1ng of DNA=300 copies) was
generated to quantify the copy number of each sample. In all cases, the low background signal
obtained with no antibody signal was subtracted. Details of primers used for qPCR assays are in
Apx3.
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3.3.6 Plasmid construction
CHAF1A CDNA was amplified from CHAF1A cDNA clone from Open Biosystems using the
following primers: attB1, 5’-GGGG ACA AGT TTG TAC AAA AAA GCA GGC TCA GCC
GCC ACC ATG CTG GAG GAG CTG GAG TGC-3’) and attB2, 5’-GGG G AC CAC TTT
GTA CAA GAA AGC TGG GTA - GGA TGC ACC CAG TGG GCT CGC-3’). The
amplified product was extracted from the Agarose gel using Qiagen Gel extraction kit and was
cloned into pLenti6/UBC/V5-DEST vector (Invitrogen) by LR reaction using Gateway strategy
as described and provided by Invitrogen.
3.3.7 Immunostaining
2x105 cells were seeded on coverslip in a 6 well plate and incubated overnight. Next cells were
pre-extracted with nuclear extraction buffer (50mM Tris, pH7.2, 300 mM NaCl, 1mM DTT,
0.5% (w/v) before fixing with 4% PFA for 10 min. Next, cells were incubated in cold BSA-PBT
for 1hr for blocking. Then primary antibody were added to the cells for 1hr washed 3 times and
secondary antibody were add for 1hr fallowed by washing for three times with PBT. Coverslips
were mounted with Vector Lab Vector Shield and viewed by Zeiss laser confocal microscope.
3.3.8 Cell cycle block
2x106 cells were per plate in 10cm plates ad incubated overnight. Next cells were transfected
with siCtrl or siCHAF1A for 2 days and media was refreshed with or without Aphidicolin on day
3. Next half of the plates were incubated with or without IFN! for 6 hours. IFN! treated plates
were harvested 6 hours later and stored -20. Another half of the plates were washed with PBS
for 3 times and then refreshed with normal media and incubated for 24hrs followed by treatment
with IFN! for 6hrs. Cell lysate and cDNA were prepared from each sample as described above
for Western blotting and RT-PCR analysis.
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3.4 Results
3.4.1 CHAF1A and PCNA repress CIITA induction
We showed before that CIITA is silent and not responsive to IFN! in SW13 BRG1-defecient
adrenal carcinoma cells[33]. Unpublished data in our lab showed that PRC2 components are
recruited to the promoter and distal sites at the CIITA locus and keep the gene silent. To identify
other components involved in CIITA silencing, we performed a Suppressor screen (i.e. to find
siRNAs that suppress defective ISG- responsiveness in BRG1 deficient cells) as described in
Chapter 2. For this screen, a part of the Dharmacon Human Genome siARRAY siRNA SMART
pool was used and CIITA-BAC reporter stably integrated into the genome of SW13 cells was
used as a molecular read-out. We selected two hits, CHAF1A and PCNA, from the Suppressor
screen for additional validation and functional studies. These genes are functionally related and
involved in H3/H4 deposition during replication.
First, we investigated the roles of CHAF1A and PCNA in endogenous CIITA induction.
To knock down CHAF1A and PCNA, three different siRNAs were used from Qiagene siRNA
library (siCHAF1A#2, siCHAF1A#3, and siCHAF1A #4) and one siRNA from Dharmacon
library (siCHAF1A#1). The western blot confirmed that all 4 siRNAs targeting either CHAF1A
or PCNA reduced the protein level of targeted genes by more than 50% (Fig 3.4.1A). The
mRNA level of knock down of CHAF1A and PCNA was confirmed by RT-PCR; more than 50%
reduction in mRNA level was observed (Fig 3.4.1B). Importantly, knocking down either
CHAF1A or PCNA using any of the siRNAs rescued the CIITA response to IFN! by
approximately seven fold or four fold relative to siCtrl, respectively, without changing CIITA
basal level (Fig3.4.1C). For the rest of this chapter, we focused on CHAF1A to investigate its
role in mediating PRC2 silencing of IFN! regulated genes.
Multiple siRNAs against CHAF1A rescued IFN! induction of CIITA in BRG1-deficient
SW13 cells (Fig 3.4.1A), indicating that this result was not due to an off-target knockdown. To
further validate this conclusion a stable cell line was generated overexpressing CHAF1A ~10-
fold (Fig 3.4.1D). When these cells were challenged with siCHAF1A#1 CHAF1A protein was
reduced but to levels equivalent to that in untreated or siCtrl-treated SW13 cells (Fig 3.4.1D).
When these cells were exposed to IFN!, CIITA remained silent, proving that the critical target of
siCHAF#1 in SW13 cells is CHAF1A (Fig 3.4.1E). These results demonstrate that
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overexpression of CHAF1A rescues the phenotypes observed in siCHAF1A treated cells,
confirming its involvement in CIITA silencing.
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70
Fig 3.4.1 CHAF1A and PCNA repress endogenous CIITA induction. A. All four siRNAs targeting CHAF1A or PCNA reduce the protein expression. SW13 cells were transfected with 4 different siRNAs targeting CHAF1A (siCHAF1A #1, #2, #3, #4) or PCNA (siPCNA #1, #2, #3, #4) for 4 days. Western blots were performed with cell lysates collected after 6 hr treatment with IFN!. Upper panels show Western blots, lower panels graph quantification of protein levels. B. RT-qPCR was used to analyze RNA from cells in (A), confirming successful knockdown. C. Removing CHAF1A or PCNA rescues IFN! responsiveness of CIITA in BRG1-deficient cells. RT-qPCR was used to analyze CIITA levels in RNA from cells in (A). Results are plotted relative to the siCtrl transfected cells. D. CHAF1A over-expression in SW13 stable cells. SW13 cells were stably transduced with CHAF1A lentivirus (SW13-Lenti-CHAF1A), these cells or parental SW13 cells were transfected for four days with siCtrl (lanes1, 2) or siCHAF1A (lanes 3, 4) and lysates were collected after 6 hrs of IFN! treatment. Western Blotting showed 10 fold higher CHAF1A protein in SW13-Lenti-CHAF1A cells compared to parental SW13 cells, and siCHAF1A reduced the protein in the stable line to the level similar to that seen in parental SW13 cells. E. CHAF1A over-expression in siCHAF1A-treated cells reinstates CIITA repression. RT-qPCR analysis was performed on RNA from cells treated as in (D). Results are average of three experiments +/- SD.
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3.4.2 CIITA rescue by CHAF1A KD is BRM independent
BRG1 is the catalytic ATPase subunit of the SWI/SNF chromatin remodeling complex. In
humans, SWI/SNF associates either with BRG1 or BRM which share more than 70% amino acid
sequence identity[315, 316]. In some BRG1-mutant cell lines, including SW13 cells, BRM is
intact but silenced [317], raising the possibility that siCHAF1A might rescue CIITA
responsiveness by inducing BRM. Indeed, real time PCR showed an induction of BRM mRNA
in siCHAF1A treated SW13 cells (Fig 3.4.2B), although this did not result in detectable levels of
BRM protein as assessed by Western blotting (Fig 3.4.2A). To test whether CIITA rescue by
siCHAF1A was indeed independent of BRM, we performed double knock down using
siCHAF1A and siBRM. SiBRM reduced BRM mRNA as shown in (Fig 3.4.2B), but did not
impair rescue of CIITA to response to IFN! where CHAF1A was knocked-down (Fig 3.4.2C).
These results show that siCHAF1A rescues CIITA induction is not via induction of BRM.
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Fig 3.4.2 CIITA rescue by siCHAF1A is BRM-independent. A. Undetectable levels of BRM protein in siCHAF1A treated cells. SW13 cells were transfected with siCtrl, siCHAF1A or siCHAF1A + siBRM for 4 days and exposed to IFN! for 6 hrs. Western Blotting analysis showed successful CHAF1A depletion, but no detectable BRM protein. HeLa cell lysate was loaded as a control for BRM detection. B, C RT-qPCR analysis of RNA from cells treated as in (A) showed some induction of BRM RNA, which was down-regulated after exposure to siBRM (B), but with no effect on CIITA responsiveness (C). Results are average of three experiments +/- SD.
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3.4.3 The effect of siCHAF1A on known regulators of IFN! signaling
The binding of IFN! to its receptors activates a signaling cascade which leads to phosphorylation
of STAT1 (generating p-STAT1), and induction of IRF1, two major transcription factors for ISG
induction. It is possible that the effect of CHAF1A KD to rescue CIITA responsiveness could be
caused by the effect of CHAF1A on IFN! signaling regulators such as STAT1 and IRF1(Fig
3.4.3A) or p-STAT1(Fig 3.4.3B) . To address this question, we assessed the protein level of
STAT1, IRF1 and IFN! activated form of STAT1, p-STAT1, in CHAF1A-knock down cells.
Western blotting analysis showed there may be a small change in both p-STAT1 and IRF1, but
since this does not affect the levels of other BRG1/PRC2 independent ISGs (Fig 3.4.6A), we can
propose that it is unlikely to be the explanation for rescue of BRG1/PRC2 regulated genes.
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Fig 3.4.3 The effect of siCHAF1A on the level of IRF1, p-STAT1 or STAT1 proteins. SW13 cells were transfected with siCHAF1A or siCtrl for 4 days followed by IFN! treatment for for 6hrs. Western Blot analysis was assessed for STAT1, IRF1, p-STAT1.
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3.4.4 Effects of siCHAF1A on cell survival and DNA damage
Several biological effects of CHAF1A depletion have been reported before [194, 318]. In
mammals, but not in yeast, knock down or mutation of CHAF1A impairs nucleosomal assembly
during replication, and causes S-phase arrest which can lead to a DNA damage and apoptosis
[195, 228, 318] . Other studies also showed that CHAF1A is required in deposition of H3 to the
DNA damaged induced sites [319, 320]. To test the effect of siCHAF1A on cell survival, we
transfected SW13 cells with two different siRNAs and investigated cell viability at the time of
transfection and up to 4 days after transfection by trypan blue which selectively stains dead cells.
SiCHAF1A had little or no effect on cell death at day 4 (Fig 3.4.4.A). Immunostaining for
cleaved caspase 3, a marker of apoptotic cells, also detected no difference between siCHAF1A
and siCtrl treated cells (Fig 3.4.4.B). Moreover, we also assessed cell growth and carefully
examined four different sub-phases of S-phase identifiable by replication fork patterning (S1-
S4). We observed that while siCHAF1A slightly lowered the growth of SW13 cells, the
progression through S-phase was unaffected (Fig 3.4.4.C). Thus, siCHAF1A had marginal or no
effects on the growth and survival of SW13 cells. As discussed later in this chapter, inhibiting
the cell cycle by aphidicolin, that also induces DNA damage (Fig 3.4.4D) did not rescue IFN!
responsiveness of CIITA, and actually blocked rescue by siCHAF1A (Fig 3.4.10), arguing that
the effects of siCHAF1A on CIITA are not an indirect consequence of cell cycle inhibition or
DNA damage.
In line with prior reports [318] Immunostaining showed that siCHAF1A induced !H2Ax,
a marker of DNA damage (Fig 3.4.4 D). However, while treatment of SW13 cells with 2 %M of
the potent DNA damaging agent Methyl Methane-Sulfonate (MMS) also induced !H2Ax (Fig
3.4.6 D), it did not rescue responsiveness of CIITA to IFN! (Fig 3.4.6 E). In conclusion, the
effect of siCHAF1A on CIITA induction is not an indirect consequence of DNA damage.
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77
Fig 3.4.4 Effects of siCHAF1A on cell growth, apoptosis and DNA damage. A. Cell growth is slightly effected by siCHAF1A. SW13 cells were transfected with siCHAF1A#1 and siCHAF1A#2 for 4 days. Cell number was measured by trypan blue staining and plotted. B. siCHAF1A does not induce apoptosis. SW13 cells were treated with siCHAF1A or siCtrl for 4 days then left untreated or exposed to IFN! for 24 hrs, or with etoposide as a positive control for apoptosis. Cells were stained with DAPI (blue) to mark nuclei and cleaved caspase 3 (red) to detect apoptosis. C. S-phase distribution is not affected by siCHAF1A. SW13 cells were treated as in B, incubated with EdU for 15min before pre-extraction and fixation and then stained for CHAF1A and PCNA and the characteristic pattern of replication fork distribution used to quantify the fraction of cells in each the four identifiable S phase stages (S1-S4). Cells in 10 randomly selected fields were quantified. D. siCHAF1A KD induces DNA damage. SW13 cells were treated as in B or were exposed to Aphidicolin and 2%M Methyl Methane-Sulfonate (MMS), both DNA damaging drugs, for 24hrs, then stained for DAPI (blue) and !H2Ax (green), a DNA damage marker. E. DNA damage does not rescue CIITA responsiveness. SW13 cells were treated 2 %M MMS for 24 hrs before IFN! treatment for 6hrs. RT-qPCR was used to measure CIITA mRNA levels. Results are average of three experiments +/- SD.
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3.4.5 Depleting other components linked to histone deposition, DNA replication or gene silencing does not rescue CIITA
We also used RNAi to deplete a series of other factors involved in DNA replication, DNA repair,
and gene silencing, none of which rescued IFN!-induction of CIITA (Fig 3.4.5A). 1. First Non-
coding RNAs have been also implicated in the recruitment of PRC2 and gene silencing in
heterochromatin [321, 322]. Argonaut proteins which are part of the RISC complex recruit non-
coding RNAs to DNA which facilitates PRC2-dependent gene repression. To test whether
Argonauts are involved in CIITA silencing, we knocked down different isoforms of Argonaut
proteins (Ago1 and Ago2). 2. There is evidence implicating DNA methylation in the regulation
of PRC2 and H3K27me3 distribution. Thus we knocked-down the DNA methyltransferase,
DNMT1. 3. We assessed various proteins that interact with CHAF1A to regulate H3K9me3
epigenetic marks: MBD1, EHMT2 (G9A), SETDB1, and HP1 (all 3 isoforms #, ", !) [223, 225,
323]. 4. Since CHAF1A functions during replication we examined other components of DNA
replication such DNA polymerase epsilon (POLE) and delta (POLD). 5. HIRA is another H3
histone chaperone involved in the replication-independent H3 deposition [324] 6. Further, since
CHAF1A is also involved in DNA repair we looked at RAD9A, a cell cycle checkpoint protein
required for cell cycle arrest and DNA damage repair in response to DNA damage [318]. Real
Time PCR showed more than 50% knock down of all targeted genes (Fig 3.4.5A). However,
none of these siRNAs mimicked the effect of siCHAF1A (Fig 3.4.5B).
We cannot firmly conclude that none of these proteins regulate CIITA responsiveness
because a) Westerns would be required to assess knockdown efficiency, and b) Several of the
proteins may function redundantly with related factors. Nevertheless, all these data stand in stark
contrast to the clear effect of siCHAF1A, underscoring the importance of CHAF1A in repressing
CIITA responsiveness in BRG1-deficient cells.
Human CAF-1 histone chaperone complex contains three subunits, CHAF1A (P150),
CHAF1B (P60) and RbAp48 (RBBP4) [182, 313]. CHAF1A and CHAF1B interact and are
required for replication dependent nucleosome assembly in human cells [313]. As described
above, CHAF1A, the largest subunit of CAF-1, is involved in CIITA silencing. To investigate the
specificity of CHAF1A subunit of CAF-1 complex in CIITA regulation we knocked down
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CHAF1B and RbAp48. Real Time PCR analysis showed that > 50% reduction of CHAF1B or
RbAp48 mRNA (Fig 3.4.5B) did not affect CIITA transcription (Fig 3.4.5A).
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Fig 3.4.5. Other CAF-1 components, DNA replication factors, alternative H3 histone chaperon, and known CHAF1A interacting proteins involved in gene silencing appear to be dispensable for CIITA silencing. A. SW13 cells were transfected with DNA replication factors siPOLD, siPOLE, a DNA repair factor, RAD9A , a H3 histone chaperone HIRA, gene silencing factors, siAGO1/2, DNMT1, HP1(#,",!), a histone lysine 9 methyltransferase, EHMT2 and CHAF1A interacting silencing factors, SETDB1 and MBD1 but none of them rescued IFN!-induction of CIITA. B. siRNAs targeting selected genes reduced the mRNA level of targeted genes more than 50%. SW13 cells were transfected with siRNAs for each genes for 4 days and were then collected after a 6 hr IFN! treatment. The percentage of remaining mRNA of targeted genes were measured by RT-qPCR and showed more than 50% knock down for the targeted genes. Results are average of three experiments +/- SD.
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3.4.6 CHAF1A represses multiple PRC2-regulated ISGs
Mohamed Abou El Hassan in our lab found that in addition to CIITA, multiple BRG1-dependent
ISGs are also repressed by PRC2 (unpublished data). Thus, we wondered whether CHAF1A
also collaborates with PRC2 in the repression of these other genes. To address this question, we
selected: IFI2, GBP1, GBP3, IFITM3 and examined the expression of these genes after
CHAF1A KD (Fig 3.4.6). We used IRF1, a BRG1/PRC2-independent ISG which is responsive
in SW13 cells, as a control. As an additional control, we used PITX2, a silenced non-ISG.
Importantly, siCHAF1A rescued all of the BRG1/PRC2-regulated ISGs, but had no effect on
IRF1 and PITX2 (Fig 3.4.6). This indicates that CHAF1A is most probably only involved in the
regulation of ISGs regulated by both BRG1 and PRC2.
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Fig 3.4.6. CHAF1A KD rescues the induction of multiple PRC2/BRG1dependent ISGs in response to IFN!. A. CHAF1A KD rescues the expression of multiple BRG1/PRC2-dependent ISGs. SW13 cells were transfected with siCHAF1A or siCtrl for 4 days. Cells were then treated with IFN! for 6 hrs. The mRNA level of the indicated BRG1 dependent ISGs, the BRG1 independent ISG IRF1, or the silent non-ISG PITX2 were measured by RT-PCR Results are average of three experiments +/- SD.
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3.4.7 CHAF1A is required for PRC2-mediated chromatin modification
CHAF1A is involved in replication-dependent chromatin assembly [313, 325]. CHAF1A also
maintains the heterochromatin epigenetic modification H3K9m3 at heterochromatin during
replication through interactions with the histone H3K9-methyltransferase SetDB1,
heterochromatin protein 1 (HP1), ensure proper transfer of the heterochromatic epigenetic
modifications during DNA replication [145, 223, 225, 326]. However, CHAF1A has not been
connected previously to PRC2 and H3K27me3 deposition. As mentioned above, previous work
in our lab showed that PRC2 represses CIITA and components of the PRC2 complex, such as
EZH2 or SUZ12, in addition to the PRC2-mediated silencing mark, H3K27me3, were detected
across the CIITA locus (unpublished data). Thus, it is possible that CHAF1A may be involved in
maintaining the epigenetic repressive mark H3K27me3 at PRC2-repressed genes. To investigate
this possibility we used ChIP to compare H3K27me3 levels across CIITA in siCtrl or siCHAF1A
treated SW13 cells. Also, since CHAF1A is involved in H3-H4 tetramer deposition, we tested
whether histone H3 deposition is disrupted by siCHAF1A. SW13 cells were transfected with
two different CHAF1A siRNAs (#1 or #2) or siCtrl for four days. Western analysis showed that
each siRNA reduced CHAF1A protein level by > 50% compared to siCtrl (Fig 3.4.7A). The
ChIP for H3 did not indicate any change across the CIITA locus in siCtrl or siCHAF1A-treated
SW13 cells (Fig 3.4.7 B). However, the H3K27me3 mark was markedly reduced across the
CIITA locus after CHAF1A knock down (Fig 3.4.7C).
Next we investigated whether siCHAF1A reduces the H3K27me3 mark at the promoters
of other PRC2-dependent ISGs. Significant levels of H3K27me3 were detected at the GBP3 and
GBP4 promoters, and siCHAF1A reduced the mark by ~50% (Fig 3.4.7D). IRF1, a
PRC2/BRG1 independent ISG served as a negative control (Fig 3.4.7E). These data suggest that
CHAF1A is required for PRC2 dependent H3K27 methylation at PRC2/BRG1 dependent ISGs,
providing a mechanism to explain why siCHAF1A rescues IFNg responsiveness at such loci.
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Fig 3.4.7. CHAF1A KD reduces H3K27me3, a PRC2 mediated histone modification at PRC2/BRG1 regulated ISGs. A. Both CHAF1A siRNAs (siCHAF1A#1 and siCHAF1A#2 reduced CHAF1A protein level by more than 80%. SW13 cells were transfected with 2 different siCHAF1A, either with siCHAF1A#1 or siCHAF1A#2 or siCtrl for 4 days. Cell lysates of siCHAF1A and siCtrl cells were used for Western Blotting analysis. B. ChIP showed no disruption in H3 deposition in CHAF1A KD across the CIITA locus. Chromatin lysates of SW13 cells, either transfected with siCHAF1A#1 and siCHAF1A#2 or siCtrl, were used for ChIP, using H3 antibody, followed by RT-qPCR. RT-qPCR results examining the remote elements across CIITA showed no changes in H3 deposition in CHAF1A KD compared to the control cells. C. SW13 cells transfected with either siCHAF1A#1 or siCHAF1A#2 showed significant reduction in H3K27me3 compared to the siCtrl-treated cells across CIITA. D. CHAF1A is required for the deposition of H3K27me3 at the promoters of PRC2/BRG1 dependent ISGs. SW13 cells were transfected with siCHAF1A#1 or siCtrl for 4 days. ChIP experiment was performed using an H3K27me3 antibody for siCHAF1A and siCtrl chromatin lysate. ChIP-qPCR analysis confirmed a reduction of PRC2 chromatin mark, H3K27me3 at the promoters of BRG1/PRC2regulated ISGs. Results are average of three experiments +/- SD. Asterisks indicate a significant reduction in H3K27me3 mark (p<0.05).
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3.4.8 General role for CHAF1A in promoting PRC2-mediated gene silencing
We next asked whether CHAF1A represses PRC2-regulated genes that are not ISGs. We first
examined the expression of a known PRC2 non-ISG target gene, E-cadherin [13, 327, 328].
Interestingly, siCHAF1A elevated the basal level of E-cadherin (Fig 3.4.8A) and ChIP analysis
revealed reduced levels of H3K27me3 at the promoter (Fig 3.4.8B).
Mohamed Abou El Hassan in our lab identified multiple PRC2 repressed non-ISGs
through RNA-seq analysis following siSUZ12 treatment of SW13 cells (unpublished data). In
preliminary analysis (n = 1) of six of these loci, we found that siCHAF1A also induced the basal
levels of these targets (Fig 3.4.8C). Altogether, these data suggest that the collaboration
between CHAF1A and PRC2 in gene silencing may be of broad relevance.
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Fig 3.4.8 Requirement of CHAF1A in PRC2-mediated gene silencing is a general effect. A. siCHAF1A induces the basal levels of E-Cadherin, a BRG1/PRC2-regulated gene. SW13 cells were transfected with siCHAF1A or siCtrl for 4 days and E-Cadherin levels assessed by RT-qPCR. B. siCHAF1A reduces H3K27me3 levels at the E-Cadherin promoter. H3K27me3 ChIP analysis was performed on chromatin from the cells treated as in (A). . Asterisks indicate a significant reduction in H3K27me3 mark. (p<0.05 C. siCHAF1A rescues PRC2 regulated non-ISGs. SW13 cells were transfected with siCHAF1A and siCtrl for 4 days and mRNA levels of selected PRC2 regulated genes were assessed by RT-qPCR . CDH 2 was used as a negative control (n=1))
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3.4.9 CHAF1A-mediated repression requires S-phase
CHAF1A might promote PRC2 function during DNA replication, ensuring that newly deposited
nucleosomes acquire the H3K27me3 mark. Alternatively, CHAF1A might regulate PRC2
activity independent of its role as a replication fork protein. To examine these alternate
possibilities, we asked whether the effects of siCHAF1A on BRG1/PRC2 regulated ISGs and
non-ISGs required DNA replication. In this experiment, SW13 cells were either allowed to
divide or arrested at G1/S by aphidicolin before transfection with siCtrl or siCHAF1A. On day 4
of the experiment RNA was collected to assess gene expression, and in a parallel set of plates
aphidicolin was washed off to release cells into the cell cycle, following which RNA levels were
assessed on day 6. Cell cycle arrest was confirmed by flow cytometry (data not shown).
Western blotting analysis showed that knock down was efficient in dividing, arrested, and
arrested-then-released cells (data not shown). Strikingly, however, CIITA, a PRC2/BRG1
dependent ISGs, and E-cadherin, a PRC2 dependent non-ISG remained silent in arrested cells,
and gene induction was only observed when cells were allowed to divide from day 4 to day 6
(Fig 3.4.9 A,B). The PRC2/BRG1 independent genes IRF1 and TBP were used as negative
controls (Fig 3.4.9C,D). These data indicate that CHAF1A-mediated repression of PRC2-
regulated genes requires active cell division and likely occurs through a replication-dependent
epigenetic mechanism.
The large error bar of E-cadherin expression at day four, Fig 3.4.9A and CIITA expression at
day 6 of Fig 3.4.9B may be the result of two factors: either the knock down is not efficient
enough, or the cells have not gone through enough cell cycle to show the efficient reduction of
H3K27me3. Indeed, I observed that if cells are allowed to divide longer better gene induction is
detected, with less variability.
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Fig 3.4.9 The rescue of PRC2-regulated genes after CHAF1A KD requires cell cycle progression. SW13 cells were treated with or without aphidicolin (AP) overnight to arrest S phase before transfecting with siCHAF1A or siCtrl for 4 days. Cells were collected and mRNA was extracted to perform RT-qPCR. GBP2, IFITM3 and CIITA are BRG1-dependent ISGs that are rescued by siCHAF1A in the absence of AP, rescue was reversed in the presence of AP. IRF1, a BRG1-independent ISG which is not induced by siCHAF1A and TBP, a housekeeping gene, were used as a negative controls. Results are the average of two independent experiments +/-SD.
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3.4.10 SiCHAF1A causes a dramatic redistribution of H3K27me3 without changing PRC2 localization
We showed that siCHAF1A reduces the PRC2-mediated histone H3K27me3 mark at the
regulatory elements of PRC2 target genes (Fig 3.4.7C). One possibility to explain this result
would be that siCHAF1A reduces the protein level of PRC2 components or total level of
H3K27me3. Interestingly, Western blot analysis did not show any change in the total level of
H3K27me3 or PRC2 components (Fig 3.4.10A). One way to explain this result is that,
siCHAF1A reduces the chromatin bound H3K27me3 and not the total protein. However,
fractionation of lysates revealed no change in the chromatin bound H3K27me3 in siCHAF1A
treated cells (data not shown). These results show there must be another mechanism through
which siCHAF1A reduces the PRC2-mediated histone marks at specific targets.
One possibility is that the effect of siCHAF1A on PRC2 targets is to affect the
recruitment of PRC2 to replication forks and/or the activity of PRC2 at forks, thus affecting the
distribution of H3K27me3. Immunostaining suggested that siCHAF1A did not affect the
recruitment of PRC2 to replication forks (Fig 3.4.10B). Thus, CHAF1A is not physically
involved in the recruitment or maintenance of PRC2 at sites of replication. However,
Immunostaining revealed a striking redistribution H3K27me3 to the nuclear periphery (Fig
3.4.10 C). Thus, while CHAF1A does not affect bulk levels of H3K27me3 (Fig3.4.10A), it is
critical for the appropriate distribution of H3K27me3, suggesting that it may regulate PRC2
activity at the fork (Fig 3.4.10 C).
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Fig 3.4.10 CHAF1A KD does not change PRC2 protein level and localization but it alters H3K27me3 distribution. A. CHAF1A KD has no effect on PRC2 components protein levels or relevant chromatin modification. Western Blot was performed after 4 days of transfecting SW13 cells with siCHAF1A or siCtrl. The protein levels of H3K27me3, H3, EZH2, SUZI12 were not affected by CHAF1A KD. Results are average of three experiments +/- SD. B. siCHAF1A does not block PRC2 recruitment to replication forks. SW13 cells were transfected with siCHAF1A or siCtrl for 4 days before being reseeded onto coverslips for immunostaining for EZH to mark PRC2 location or PCNA to mark replication forks. Co-localization at the fork was unaffected in the absence of CHAF1A. C. CHAF1A KD causes a dramatic redistribution of H3K27me3. Cells from (A) were collected at day four and seeded onto cover slips. They were stained the next day with a H3K27me3 antibody. Gray scale images shown were overlaid images of five 0.4 um thick z-stack images. Surface plot of H3K27me3 of cells in the gray scale images are shown on the left.
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3.5 Discussion
3.5.1 A novel direct link between CHAF1 and PRC2-mediated gene repression during replication
We discovered an unexpected and interesting link between CHAF1A, the large subunit of CAF-1
histone chaperone and PRC2-mediated gene regulation in the human adrenal carcinoma SW13
cell line. Depleting CHAF1A rescued PRC2-dependent repression of both ISGs and non-ISGs in
these cells. Interestingly this rescue is replication-dependent as it does not occur when the cell
cycle is blocked. Since CHAF1A deposits H3 during replication, CHAF1A is most likely
needed to transfer the PRC2-mediated silencing mark during this time.
Generally, in mammals there are two major H3 epigenetic silencing marks which define
the silent status of genes: SUV39-mediated H3K9me3 and the PRC2-mediated H3K27me3.
During replication the nucleosomes are disassembled in front of replication forks and then
reassembled, now including newly synthesized histones. It is important that the epigenetic
modification of the parental histones is copied to the new ones.
Very little is known about the mechanism of how this occurs, however deregulation of this step
has been linked to several diseases including cancer. Recent data reinforce the idea that
CHAF1A is likely a key player in this process [329, 330].
Most studies of CHAF1A and epigenetic memory focus on the effect of CHAF1A on
H3K9me3 silencing mark. For example, CHAF1A maintains of this mark at telomeres and yeast
mating type loci [146, 147, 177, 216, 331]. Also, CHAF1A has been identified as a
transcriptional repressor in the screen for silencing factors in human cells and the 5$ truncated
form of this protein impairs the maintenance of transcriptional gene silencing [143, 147, 326].
The mechanism through which CHAF1A maintains the H3K9me3 mark involves the
heterochromatic protein HP1#, MBD1 and, SETDB1, a histone H3K9 methyltransferase during
S- phase [145, 223-225].
In contrast to H3K9me3, the mechanism of H3K27me3 maintenance is not well
understood. Here we show that CHAF1A also maintains this PRC2-mediated epigenetic mark.
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Interestingly, the only other occasion when a link between CHAF1A and PRC2 has been
suggested is a study showing that a Drosophila CHAF1A (P180) mutant disrupts PRC2-
mediated silencing in developmentally important genes [216]. However, this study did not
address the functional significance of this observation. Therefore, our study shows for the first
time the functional link between CHAF1A and PRC2-dependent gene silencing in mammalian
cells.
3.5.2 CHAF1A is required for printing the H3K27me3 mark at the proper targets
To dissect the mechanism through which CHAF1A maintains the H3K27me3 mark, we first
wanted to know if siCHAF1A reduces the protein levels of PRC2 components. However this
was not the case. Moreover, removing CHAF1A did not affect the bulk levels of H3K27me3 on
chromatin. Importantly, we found that siCHAF1A KD resulted in redistribution of the
H3K27me3 mark from a central to peripheral nuclear location. This striking observation
suggests that CHAF1A is required for the printing of H3K27me3 by PRC2 at the appropriate
targets. To find out how CHAF1A is involved in maintaining H3K27me3 during S-phase, we
need to understand how PRC2 is recruited to its targets and what affects its activity.
There are two proposed models for PcGs recruitment and function. The first one is the
“instructive model” which is based on direct interaction of PRC2 with sequence specific TFs or
non-coding RNAs [332-335]. The major challenge for this model is that it does not explain how
PRC2 binds multiple diverse non-coding RNAs or TFs, which is required to define gene
expression patterns during development [336]. However, recently an alternative “responsive”
model for PRC2 function has been proposed [336]. This “responsive” model is derived from the
preferential binding and activity of PcG at hypomethylated CpG islands (CGIs) in vertebrates
[337, 338]. In this model, both PcG and TrxG complexes constantly sample CGIs-containing
chromatin sites. The chromatin conformation created by CGIs, in combination with “silencing”
or “anti-silencing” factors, favors the recruitment of either PcG or TrxG [336]. For example,
H3K4me3, H3K36me3, RNA polymerase II or H3K27ac, block PRC2 activity or recruitment
while they promote TrxG activity or recruitment and poise genes for activation [76, 336, 339].
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In the absence of these “anti-silencing” factors, PRC2 is recruited to CGIs and through
positive feedback processes accumulates H3K27me3 mark at the PRC2 targets. For example,
H3K27me3 recruits the PRC1 complex which would compact chromatin, and it has been shown
that PRC2 prefers to methylate H3K27 when the chromatin is compact [340]. Finally binding of
PRC2 complexes to their targets and accumulation of H3K27me3 transfers the status of the
targeted sites from “sample” to “established” state [336, 340, 341].
Based on the “responsive” model for PRC2 recruitment and function, it is possible that CHAF1A
regulates some or all of the chromatin features that regulate PRC2 recruitment and/or activity.
For example, since DNA methylation inhibits PRC2 (see above) it is possible that a CHAF1A-
containing complex at the replication fork ensures that CGIs which are PRC2 targets remain
unmethylated. In agreement with this notion, it has been shown that the DNA methylase
DNMT1 interacts with PCNA [146, 147, 194, 342]. DNMT1 is required for the maintenance of
DNA methylation during replication [58, 343]. This is intriguing, because our siRNA
Suppressor screen aimed to identify novel factors associated with the silencing of CIITA, aside
from CHAF1A also turned up PCNA. Several studies showed that CHAF1A and PCNA interact
at the replication fork and that PCNA is required for epigenetic inheritance during replication
[146, 147, 194]. Therefore, we hypothesize that PCNA may also be involved in the appropriate
targeting of H3K27me3 by CHAF1A during replication through DNA methylation.
We observed that in the absence of CHAF1A, H3K27me3 is re-distributed from a primarily
central (euchromatin) to a peripheral (heterochromatin) nuclear location. Other work in the lab
using electron spectroscopic tomography revealed that siCHAF1A does not affect the location of
heterochromatin itself (E. Fussner), arguing that the chromatin mark has not moved to the
chromatin subtype. In the absence of CHAF1A it is possible that another histone chaperone is
recruited to the replication fork to substitute for CHAF1A. However this replacement complex
might not faithfully transfer the pattern of DNA methylation or chromatin modifications from
parental DNA to the daughter strands during replication. Perhaps during early S-phase, when
euchromatic regions lacking 5mC are replicated, the replacement histone chaperone recruits
DNMT1, or one of the other members of this family (e.g. DNMT3), which aberrantly methylates
DNA sequences, including previously hypomethylated CGIs which are normally targeted by
PRC2. This aberrant DNA methylation would block PRC2 activity, and after two cell cycles
would result in a dramatic increase in 5mC, concomitant inhibition of PRC2 and loss of
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H3K27me3. Equally, the absence of CHAF1A during replication of heterochromatin could lead
to a deficit in 5mC maintenance, which could permit the spread of H3K27me3 mark into these
regions. Since we observe accumulation of H3K27me3 at the nuclear periphery (i.e. at
heterochromatin), it is possible that during replication the absence of CHAF1A-driven DNA
methylation leads to redistribution of H3K27me3. That is, the H3K27me3 mark is gained where
the DNA methylation is lost [108]. This is consistent with recently published studies indicating
that the pattern of H3K27me3 distribution correlates with hypomethylated regions [108].
Indeed, when DNA methylation is absent, e.g. following DNMT1 deletion, depletion or drug-
mediated inhibition, H3K27me3 redistributes such that it is depleted at formerly enriched areas
and elevated at formerly H3K27me3 poor regions [108]. This may be akin to the phenomenon
we observe in siCHA1A treated cells.
Interestingly, another piece of evidence implicating DNA methylation in the mechanism of
CHAF1A action comes from the recent preliminary data obtained by Tom Leung in our lab. He
found that siCHAF1A causes redistribution of DNA methylation in the opposite way of
H3K27me3 (from peripheral to central part of nuclei). Notably, total levels of 5mC were
unchanged, implying that there is no loss of either H3K27me3 or 5mC, but an inversion in their
localization. While it is established that 5mC repels PRC2/H3K27me3, it remains unclear
whether the reverse is also true. These data require extensive future genomic analysis of the
redistribution events, including kinetic studies to define when they occur relative to each other,
and parallel assessment of the location of the enzyme complexes involved.
Another possibility is that the epigenetic modification such as H3K4me3, H3K36me3 and
H3K27ac that are known to block PRC2 function [336] are aberrantly accumulated to PRC2
target sites by an unknown histone chaperone which functions in the absence of CHAF1A. To
test if this is the case, we can investigate whether any of these modifications have been increased
at PRC2 target sites in siCHAF1A treated cells. Finally, we note that CHAF1A can bind to
sumoylating enzymes [344], and PRC2 activity is modified by sumoylation [345], thus it will be
of interest to determine whether this and/or other post transcriptional modifications of PRC2 are
affected by CHAF1A depletion.
In summary, our data indicate that CHAF1A regulates the precise pattern of H3K27me3
distribution. The striking redistribution of this mark when CHAF1 is absent may be connected
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to an inverse effect on the localization of 5mC, although considerable work is required to test
this or alternate hypotheses.
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4 Discussion and Future Directions
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4.1 Discussion
4.1.1 CHAF1A maintains H3K27me3 during replication
The expression status of many genes is decided during development and for a large number of
them is maintained by the collaboration of PcG and TrxG groups of proteins [23]. These two
groups of proteins have antagonistic effects and mediate the epigenetic marks H3K27me3 and
H3K4me3 which correlate with gene repression and gene activation respectively. There is a
good understanding of how PcGs or TrxGs regulate gene expression and how they are recruited
to the targeted genes. However, the mechanism explaining how the epigenetic marks are
maintained during replication is lacking. In this study we describe a novel link between
CHAF1A, the large subunit of the CAF-1 histone chaperone complex, and the PRC2-mediated
deposition of the repressive epigenetic modification, H3K27me3.
During DNA replication, histones are disassembled in front of the replication fork and
reassembled behind it. It is not entirely clear how the epigenetic information on histones is
faithfully transferred to the daughter protein, and thus how the expression status of any given
gene is kept silent or active during replication.
There is already some evidence implicating the role of CHAF1A in the maintenance of
H3K9me3 in the heterochromatin regions. It has been proposed that CHAF1A, together with
PCNA, recruit histone methylases such as SUV39 and SETDB1, and DNA methylases to
maintain epigenetic memory [225] . However, how the H3K27me3 silencing epigenetic
modifications are transferred during replication, is not clear. Here we provide evidence that
indeed CHAF1A is a key player in this process.
Previously, our lab showed that in the BRG1-deficient adrenal carcinoma SW13 cells,
CIITA, the master regulator of MHC-Class II, is repressed by PRC2 and H3K27me3 was
detected cross the CIITA locus (M Abou El Hassan, unpublished data). We performed an RNAi
screen to uncover additional factors which together with PRC2 participate in the silencing of
CIITA. We validated two interesting hits which interact with each other and are both involved in
DNA replication, CHAF1A and PCNA. We observed that the KD of either of these genes
rescued the response of CIITA to IFN! in SW13 cells. In agreement, we found that siCHAF1A
reduces the H3K27me3 repressive marks across the CIITA locus. Further, we showed that other
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PRC2-regulated ISGs were also responsive to IFN! after CHAF1A depletion, and the
H3K27me3 mark was reduced at the promoters of these genes. Interestingly, we showed that
this effect is not limited to ISGs and that other PRC2-regulated non-ISGs, such as E-cadherin,
were also induced after siCHAF1A and the H3K27me3 mark was reduced.
Importantly, we observed that the rescue of the tested genes after CHAF1A KD is cell
cycle dependent. Since CHAF1A is known to deposit the histone H3 to the newly synthesized
DNA during replication, we believe that CHAF1A is likely a crucial factor maintaining the
H3K27 trimethylation on newly synthesized histones.
4.1.2 CHAF1A may guide the H3K27me3 deposition indirectly
How does CHAF1A guide the deposition of the PRC2 mark H3K27me3? First, CHAF1A may
instruct PRC2 indirectly by means of DNA methylation. Indeed, it has been shown in
mammalian cells that the status of DNA methylation on the CpG islands inversely regulates the
establishment of H3K27me3 by PRC2 [51, 52, 106, 108]. The hypo-methylated CpG islands
through an unknown mechanism promote H3K27 methylation [108]. Preliminary data in our lab
supports this notion as T Leung observed by Immunostaining that H3K27me3 and 5mC show
opposite patterns of distribution in the nucleus, and that siCHAF1A inverts these patterns
(unpublished data). How could this work? Several candidates might be involved in this process.
For example, TET1 is a DNA demethylase required for the recruitment of PRC2 to its targets in
mouse embryonic stem cells [346]. Interestingly, TET1 binds both PCNA and EZH2 [62].
Further, links between PRC2 and the DNMT family members such as DNMT1 and DNMT3a
have been recently described [58, 347]. DNMT1 and PCNA also interact at replication fork [58,
61]. Since CHAF1A also interacts with PCNA, cross-talk among CHAF1A, PCNA and DNA
methylases at the replication fork seems likely. Thus, we envisage that in response to
siCHAF1A, the loss of DNA methylation at the nuclear periphery (heterochromatin ) would
result in the spread of PRC2 marks into these sites, and the concomitant dilution of H3K27me3
at canonical PRC2-targeted regions, central parts of nuclei (euchromatin), resulting in elevated
expression/responsiveness of previously repressed loci; indeed, this is exactly what is observed
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when DNA methylation is inhibited in mouse embryonic fibroblasts due to expression of a
hypomorphic DNMT1 allele, knockdown of DNMT1, or treatment with 5-aza-cytidine, a DNMT
inhibitor, and induction of PRC2 targets was also observed in murine embryos expressing an
unstable DNMT1 allele [108].
Aberrant DNA and histone methylation has been reported in many types of cancer. There
are CHAF1A variants listed in the Catalogue of Somatic Mutations in Cancer (COSMIC)
database (http://cancer.sanger.ac.uk/cancergenome/projects/cosmic/), although the frequency is
low in each cancer and the functional relevance, if any, is unclear. It is possible that the activity
of CHAF1A may be compromised in cancer cells in ways other than direct mutation. It is
intriguing to speculate that the aberrant distribution of DNA methylation observed in many
cancers may be the consequence of the lack of CHAF1A activity, although this remains to be
tested.
Another possibility is that CHAF1A is required for PRC2 recruitment to the proper sites.
There are two proposed models for PcGs recruitment. In the first model the “instructive model”
PcG directly interacts with sequence specific TFs or non-coding RNAs [332-335]. For example
it has been shown that non-coding RNAs are required for the recruitment PRC2 in PRC2-
mediated X-inactivation [348, 349]. Therefore, identifying the factors that recruit non-coding
RNAs to their targets may provide additional clues about CHAF1A activity in the epigenetic
memory maintenance.
The second model which recently has been proposed is “responsive” [336]. This
“responsive” model is based on observation that PcG preferentially binds at hypomethylated
CpG islands (CGIs) in vertebrates [337, 338]. This model focuses on the effect of chromatin
conformation and modification created by combination of CGIs and histone modification on PcG
and TrxG complexes recruitment. In this model both PcG and TrxG sample CGIs-containing
chromatin sites. If the chromatin conformation contains “anti-silencing factors” such as
H3K4me3, H3K36me3, RNA polymerase II or H3K27ac, this blocks PRC2 activity or
recruitment while promoting TrxG activity or recruitment, poising genes for activation [76, 336,
339]. In contrast, in the absence of “anti-silencing” factors, PRC1 and PRC2 are recruited to the
chromatin and mediate histone modifications which eventually induce gene silencing [336].
Therefore, it is possible that in the absence of CHAF1A the chromatin conformation is changed
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by aberrant DNA methylation or recruitment of these “anti-silencing” factors which block PRC2
recruitment.
4.1.3 CHAF1A may activate PRC2
Conceivably, CHAF1A may activate PRC2. It has been recently reported that PRC2 remains
associated with DNA during replication [111]. Thus, it is possible that during DNA replication,
CHAF1A activates PRC2. Intriguingly, there is already evidence in the literature supporting this
hypothesis.
It has been shown that PRC2 components, SUZ12 and EZH2 can be SUMOylated, which
promotes PRC2 activation [345]. As mentioned above, CHAF1A recruits SUMO2/3 enzymes
which SUMOylate proteins at replication forks [344]. Thus, it is possible that SUMO2/3
recruited by CHAF1A, activate PRC2. According to this model, CHAF1A would be required for
the direct enzymatic activation of PRC2.
Taken together, it is possible that CHAF1A guides the PRC2 activity not in one way but
by the combination of direct and indirect interactions. Further, many other factors are likely to
be involved. In the next section we describe the approaches that will help to elucidate the
mechanism of CHAF1A action and fill in some of the gaps in our understanding of how this
factor orchestrates the epigenetic memory maintenance.
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4.2 Future directions
There are still many gaps in our understanding of the role of CHAF1A in PRC2-mediated gene
repression and the maintenance of the repressive PRC2 signature mark, H3K27me3, during DNA
replication. Here we discuss some of the high-priority experiments designed to address these
issues.
4.2.1 The global effect of depleting CHAF1A on PRC2 regulated genes and redistribution of H3K27me3
So far, we only focused on the regulation of CIITA and a few other PRC2-regulated genes. We
would like to further investigate the importance of CHAF1A in gene regulation of PRC2 targets,
in general.
To investigate the PRC2 gene targets in SW13 cells, our lab has previously performed
RNA sequencing study (RNA-seq) following depletion of the PRC2 core component SUZ12 (M.
About El Hassan, unpublished data). Next, we would like to perform RNA-seq in siCHAF1A
treated cells. Comparing the results of these two RNA-seq experiments will reveal how many
genes are targeted by both CHAF1A and PRC2. A high percentage of target overlap will
strengthen the position of CHAF1A in the regulation of PRC2-target genes. On the other hand a
low percentage of overlap will indicate that only select genes require the co-operation of
CHAF1A and PRC2.
By immunofluorescence (IF) staining we showed that siCHAF1A does not affect global
localization of PRC2 (T Leung, unpublished data). However since the resolution of IF is limited,
we will perform chromatin immunoprecipitation (ChIP) with massively parallel sequencing
(ChIP-Seq) for PRC2 components (e.g., SUZ12 and EZH2) and H3K27me3, either in the
presence or absence of CHAF1A to ascertain whether the recruitment of PRC2 is affected by the
loss of CHAF1A.
4.2.2 DNA methylation and histone modification
We showed that siCHAF1A causes the redistribution of H3K27me3 in SW13 cells. Further, we
also have preliminary data showing that the redistribution of DNA methylation in siCHAF1A
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treated cells occurs in the opposite way of H3K27me (T. Leung, unpublished data). To further
study this phenomenon, we will deplete CHAF1A and perform ChIP-Seq for H3K27me3 to
identify the global targets of this mark. Also, we will perform methylated DNA
immunoprecipitation (MeDIP-seq) to identify the DNA methylated regions following
siCHAF1A. The data from these two experiments will clarify two issues: 1. whether there is an
overlap between the reduction of H3K27me3 at the regulatory elements of the genes regulated by
both CHAF1A and PRC2. 2. Whether and to what extent there is a correlation between loss and
gain of DNA methylation and H3K27me3 across the genome.
If we show that indeed there is a correlation between the loss of DNA methylation and
gain of H3K27me3, and vice versa, this would strengthen our hypothesis that CHAF1A has a
global role in coordinating the distribution of these epigenetic marks. The next step, then, will
be to uncover the mechanism linking the observed DNA methylation and histone methylation
redistribution.
Another possibility is that the epigenetic modification such as H3K4me3, H3K36me3 and
H3K27ac that are known to block PRC2 function [336] are aberrantly accumulated to PRC2
target sites by an unknown histone chaperone which functions in the absence of CHAF1A. To
test if this is the case, we will perform IF and investigate whether any of these histone
modifications have been accumulated at PRC2 target sites in siCHAF1A treated cells.
4.2.3 Identifying the components involved in the CHAF1A-dependent maintenance of H3K27me3
We showed that siCHAF1A leads to H3K27me3 redistribution in a DNA replication manner and
T. Leung in our lab has preliminary data suggesting that DNA methylation is also redistributed in
the opposite way. The next step will be to identify the protein complexes or components
involved.
It has been shown that the DNA methylase DNMT1 and the DNA demethylase TET1
interact with PCNA [58, 62, 343]. Since CHAF1A binds PCNA at the replication forks, it is
likely that either these two enzymes or other members of their families collaborate with
CHAF1A in the epigenetic mark maintenance. To test whether this is the case, we will
immunostain for DNMT1, TET1 and other members of these protein families to examine how
104
their staining pattern is affected by CHAF1A KD. If the staining pattern is affected, this would
argue that CHAF1A affects their recruitment to the fork. However, it may be possible that
CHAF1A is required not for the localization of the enzymes but for instructing the enzymes
where to lay down the epigenetic mark.
We also observed that CHAF1A KD rescues E-cadherin, which is also silenced in SW13
cells by PRC2. E-cadherin is a crucial factor in Epithelial-mesenchymal transitions (EMT), and
is repressed in several tumor cells, such as breast cancer [327, 350, 351]. Since it has been
shown that repression of E-cadherin is linked to metastasis, many studies attempted to identify
its epigenetic regulators. For example, the transcription factor Snail recruits various epigenetic
modifiers to E-cadherin promoters such as LSD1/HDAC [13]. Other histone modifying enzymes
such as G9a/SUV39H are also found at the E-cadherin promoter and in turn recruit DNMTs [13].
Thus, it would be important to investigate the distribution of these factors after CHAF1A KD,
first by immunostaining and then at specific loci using ChIP. This may indicate whether any of
these factors co-operate with CHAF1A.
Last but not least, recent studies showed that ZF-CxxC domain-containing proteins, such
as CFP1 (CxxC finger protein 1), MLL (mixed lineage leukaemia protein), KDM (lysine
demethylase) 2A and KDM2B (Histone-H3-Lysine-36 Demethylase B) bind hypomethylated
DNA and recruit PRC2 to the CpG islands [352]. It will be important, using immunostaining
and ChIP, to determine whether these proteins interact with CHAF1 and are involved in its
activities.
4.2.4 Structure-Function analyses of CHAF1A
CHAF1A contains several domains that are required for its different functions. For example the
N-terminal domain binds PCNA and HP1#, proteins required for gene silencing and maintaining
heterochromatin regions [144, 145, 194]. The C-terminal domain binds CHAF1B, another sub-
unit of the CAF-I complex, and this is required for H3 deposition during replication[325, 353].
CHAF1A N- and C-terminal deletion mutants would be a useful starting point to define
which regions of CHAF1A are essential for the regulation of H3K27me3 distribution.
Expressing CHAF1A vectors in siCHAF1A treated cells coupled with RT-PCR analysis of PRC2
target loci (e.g. E cadherin or CIITA) would reveal which region of the protein is critical to
105
promote PRC2 action. These assays would utilize a siCHAF1A that targets the 3" UTR of the
mRNA, missing in the engineered vectors, so that only the endogenous gene would be depleted.
Fine mapping would be used to define point mutations in a domain that disrupts the repressive
function. IP for the wild type and mutated forms of CHAF1A followed by Mass spectrometry
(Mass Spec) would be used to identify the factors that are missing in the mutated complex.
Additional functional studies will be needed to test the significance of these novel factors at the
replication forks. For example, it would be important to use RNAi or dominant negative
strategies to define whether interfering with the function of any of these CHAF1A integrators
alleviates repression of PRC2 target genes or affects DNA methylation.
4.2.5 Using aniPOND to screen for additional factors co-operating with CHAF1A at the replication forks
Isolation of Protein on Nascent DNA (iPOND) is a technique, which is used to purify the
replication fork components for analysis by Western blotting or Mass Spec [354, 355]. Our lab
has recently developed a variation of this technique: accelerated native iPOND (aniPOND),
which improves sensitivity [356]. It will thus be of considerable interest to use aniPOND to
purify replication forks in siCtrl or siCHAF1A treated cells and assess the protein components by
Mass Spec. The factors missing in the protein complexes after siCHAF1A may be the critical
factors that influence PRC2 function and/or regulate DNA methylation. Additional functional
studies will then be need to validate whether and how these factors are required to maintain
PRC2-mediated marks.
4.3 Concluding remarks
Although the mechanism of DNA replication has been thoroughly studied, how the epigenetic
status of chromatin is inherited during this process is not completely understood. DNA
replication occurs rapidly, which makes it difficult to study the individual steps of nucleosome
duplication. H3K27me3 maintains the status of repressed genes and this is decided early in
development. It is very crucial to ensure that during replication the information dictated by
PRC2 is transferred faithfully to the newly synthesized DNA. Here, I provided strong evidence
that CHAF1A is a key factor in this process in mammalian cells. I showed that this protein is
106
required for the distribution of H3K27me3 to their proper targets. We also have preliminary
evidence that this process may be linked to DNA methylation.
Since, aberrant DNA and histone methylation are seen in many diseases, including
cancer, dissecting the mechanism and identifying the factors required for the proper maintenance
of these epigenetic marks is crucial. This knowledge will both deepen our understanding of this
process and may lead to improved therapies.
107
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Appendices
Apx1. The list of activator and inhibitor hits identified by the primary and secondary Wt and Suppressor screens.
Apx 1.1 The list of activator hits identified by the primary Wt screen. Listed below are the top 1% of the all activator hits, i.e., those which reduced luciferase activity after knock down, obtained with each method of B score, M score and control base normalization. See Fig 2.4.6 and text for details.
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Apx 1.2 The list of inhibitor hits identified by the primary Wt screen. Listed below are the top 1% of the inhibitor hits, those which induced luciferase activity after knock down, obtained with each method of B score, M score and control base normalization. See Fig 2.4.6 and text for details.
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Apx 1.3 The list of inhibitor hits identified by the primary Suppressor screen. Listed below are the top 1% of the inhibitor hits, those which induced luciferase activity after knock down, in the absence of BRG1, obtained with each method of B score, M score and control base normalization. See Fig 2.4.6 and text for details
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Apx 1.4 Secondary Wt screen, verified activator hits
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Apx 1.5 Secondary Wt screen, validated inhibitor hits
Apx 1.6 Secondary supp screen, validated inhibitor hits
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Apx2. List of SiRNAs
AllStarNegative QIAGEN 1027280 CHAF1A #1 Thermo Scientific L-019938-00 CHAF1A #2 QIAGEN SI03163699 CHAF1A #3 QIAGEN SI04219999 CHAF1A #4 QIAGEN SI04274914 PCNA#1 Thermo Scientific L-003289-00 PCNA#2 QIAGEN SI02653287 PCNA#3 QIAGEN SI02653357 PCNA#4 QIAGEN SI04436131 CHAF1B QIAGEN SI00077224 RBBP4 Thermo Scientific L-012137-00 POLD1 Thermo Scientific L-019687-00 POLE Thermo Scientific M-020132-01 HIRA Thermo Scientific L-013610-00 RAD9A Thermo Scientific M-003295-03 ARGONAUTE1 Thermo Scientific L-004638-00 ARGONAUT2 Thermo Scientific L-004639-00 DNMT1 Thermo Scientific L-004605-00 HP1a Thermo Scientific L-00429600 HP1b Thermo Scientific L-009716-00 HP1g Thermo Scientific L-010033-00 EHMT2(G9A) Thermo Scientific L-006937-00 SETDB1 Thermo Scientific L-020070-00 MBD1 Thermo Scientific L-008359-00
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Apx3. List of primers for RT-PCR!
Gene Region Chromosome position
Forward (5' to 3') Reverse (5'-3')
GBP2 Last exon AATGGCAGTTGTTTGACACTCTGA GCAAAATTTGCCTGTCCAGC
Promoter GTGTCAGGACCTGGGTTAGG CCAGTTTGAGAAGCATGTGG
GBP3 Last exon ATTACACAGTGCATAATTGTTACCATGT ACCTTAATGTGACACCTGAGACCTT
Promoter ACTGTGTGCTGGGCATTAAG GGAGGCATCACATTAATTCT
GBP4 Last exon ATATGCCATGGGCCTTTTCA CCGGGTTGTTTTAAGAAGCCT
Promoter AAACTCAAATTTCTTCCCTG AGACTCCTGAAACTCAGGTT
IFITM2 Last exon GAGTCCTGCATCAGCCCTTTA TGAATGCCATTGTAGAAAAGCG
Promoter CCAGCAAAGGACAAGGGATG AGGAGAATGGCGTGAACCGG
IRF1 Last exon AGGCCATTCCCTGTGCACCGTAGCA GTCCAGCTTCTCTGCACCATATCCA
Promoter GATCAGAGCTAGCCCACCCCTA GCCGTTGCAGGGTCTAATAG
IFI27 last exon TCTCCGGATTGACCAAGTTCA CAGGGAGCTAGTAGAACCTCGC
Promoter CCTGGTGCTTTCTCTTCCGC CCACAAAAGCACTGCAAGGA
6-16 last exon CAGCAGCGTCGTCATAGGTAAT TCCTCATCCTCCTCACTATCGAG
STAT1 Last exon CCTTATCACTGACACAAAAAGTAGATTAAGA ATAGTTGTGGTAGCAGTAGTGGAAAAAC
BPSB9 Last exon ATTGCTCTGGCCATGAGCC CCACACCGGCAGCTGTAATAG
HLA-E Last exon TTTGCAAGGGCCTCTGAATC TCCTCACATTGTGCTAACAGGG
HLA-G Last exon CTGAACTGCATTCCTTCCCC CAGCCCCTTTTCTGGAACAG
PITX2 Last exon AGGACTGCTGCCTTGTATGTTTAA TGGCCAGGACATCTCAGTCAC
MNDA Exon6 AACTTCGACTCTTCTGCCTTCAA GCTGTGACTTCCACACACCAGT
BRM Last exon GGGTGGGTCTAATTTGGTAA TGGCTTCTCCTGGTTATCAG
promoter CATCCCGCGCAGTTTCTCTG TGTCACTCGCTCAGCTCAGC
CIITA Last exon ACGTCTGACAGGCAATGCTG GGGTCCTAGCCAACTATTCCG
-70.1 kb TAAGAAATCTGTCTGTGGA TTCACAGAATTCCTCCGAAC
-50.6 kb CAGCTCATGTCCCACCCAGT AACAAACATGTCAGGCCACAGT
-16.4 kb TTCTGCAACTAGGTAACACC ATAGGTTGGATTACATGATC
-7.9 kb, PII AGTTGAACTGGCACATGGGC CTCTTGGAATTGGGAAGGCA
-0.3 kb, pIV TCACGGTTGGACTGAGTTGG CCTGAGTTGCAGGGAGCTTG
+10 GGACGTAATCTCAGCGCCTG TGTTAACGGCAACTCTGGGAG
+59.1 kb CAGCCTGTCCTCTTCTGCTCACA CGTGTTATACCCATGCCCTTGCAA
+82.6 kb TTAGAGAAAGGCACTGGATGGTCTGT GATACTTGTCTGTACACAGCCTAGCGG
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Apx3. List of primers for RT-PCR
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Gene Region Chromosome position
Forward (5' to 3') Reverse (5'-3')
CHAF1A Last exon CCACTGGGTGCATCCTGAGA TACACTGCCTCGTTAGGTCCAGGG
PCNA Last exon CCAGTTCAACTAACTTTTGC CTGACTTTGGACTTTATTCT
CHAF1B Last exon TCCAGAAATATGGATGCTGTTG TGTGCACTTTCACGAGGATG
RbAp48 Last exon AGAATACTAAGGAATCCAGTTG AGAGGTTCAAGTTGGAGGTA
POLD Last exon CCGGGAGTCTGAGCTGTATC CTTGCGCATGTAGAAGATGG
POLE Last exon AATTTCACAGGGCACCAAAC GCGATGGCCTCAGACAGTAG
HIRA Last exon TTTCCTTTGGCCGATAATCA CATTCCCCCAAAACAGTCTC
RAD9A Last exon AGACTCCCAAGCGGCTCT CAGGTCGTGGGTCTCCAG
ARGONAUT1 Last exon AATGAAATTACTTTCCTGTGCACAC AGAGACATTTCCCCATCCAT
ARGONAUT2 Last exon TGAACTAAGGAGCAGTGGCAGA CCTATAGGACAAATCTGATG
DNMT1 Last exon AAGCTGTTGTGTGAGGTTCG TTCCACTCATACAGTGGTAGATTTG
HP1a Last exon TATGGGGGAGTTTTAGCTGT GCATCAGATGTCAGTTATGG
HP1b Last exon CTTTCATATTGGGCAGTGGT CGTAGCTCTAGATGCAAGTC
HP1g Last exon GCCTTTACAGTAGAAATAGAAATGC ACTGTGGAAACATTCCTGTG
EHMT2(G9A) Last exon CTGCCCCCTGTCAACACATG
SETDB1 Last exon CAGAACTTACTTGGGACTACA GACAGTTCAAGAAGGGTTGG
MBD1 Last exon CCTCAGACTCTTAATTATGC GTCAAACCAAATAACCAGTA
E-Cadherin Last exon GAACTCAGCCAAGTGTAAAAGCC GAGTCTGAACTGACTTCCGC
promoter CGCGCTAGCGCGGCCGCATGGCTCACACCTGAAA CGCAGATCTACCCTCTAGCCTGGAGTTGC
HPRT Last exon TTGCTCGAGATGTGATGAAGG TGATGTAATCCAGCAGGTCAGC
ACTIN Last exon TCCTAAAAGCCACCCCACTTCT GGGAGAGGACTGGGCCATT
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Apx4. List of Abs
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Target protein Catalog Number source CHAF1A SC-28085 Santa Cruz PCNA SC-56 Santa Cruz B-ACTIN A-5441 Sigma b-Tubulin 2146 Cell Signalling Technology BRM 2146 Santa Cruz LAMINA/C SC-7293 Santa Cruz STAT1 SC-345 Santa Cruz IRF1 SC-497 Santa Cruz P-STAT1 9167S Cell Signalling g-H2X 05-636 Millipore SUZ12 ab12073 Abcam EZH2 3147S Cell Signalling Technology H3k27me3 07449 Millipore H3 ab1791 Abcam