Histone Crosstalks involving H3 Phosphorylation and

their Role in Transcriptional Regulation

by

Nga Ieng (Priscilla) Lau

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Graduate Department of Medical Biophysics University of Toronto

© Copyright by Nga Ieng (Priscilla) Lau 2013

ii

Histone crosstalks involving H3 phosphorylation and

their role in transcriptional regulation

Nga Ieng (Priscilla) Lau

Doctor of Philosophy

Department of Medical Biophysics University of Toronto

2013

Abstract

Histone phosphorylation is often a direct outcome of activated intracellular signaling

pathways, and functions to translate extracellular signals into appropriate biological outputs

such as changes in gene expression. Growth factors and cellular stress trigger rapid and

transient expression of immediate-early genes (such as c-fos, c-jun) in mammalian cells, and

their induction strongly correlates with a transient phosphorylation of S10 and S28 on histone

H3. While many signaling cascades that lead to H3 phosphorylation have been mapped out,

mechanistic details of the downstream events and how H3 phosphorylation contributes to

transcriptional activation are still poorly defined.

To investigate the direct effects of H3 phosphorylation on transcription, we targeted

the H3 kinase MSK1 to endogenous c-fos promoter, and found that this is sufficient to

activate its expression. Moreover, targeting MSK1 to the tissue-specific α-globin gene

induces H3S28 phosphorylation and reactivates expression of this polycomb-silenced gene.

Mechanistically, H3S28 phosphorylation not only disrupts binding of polycomb repressive

complexes, but also induces a methyl-acetylation switch of the adjacent K27 residue. This

provides the first indication that H3 phosphorylation is involved in antagonizing polycomb

silencing.

iii

To further identify post-translational modifications (PTMs) that function together

with MSK1-mediated H3 phosphorylation, I developed a novel nucleosome purification

approach called Biotinylation-assisted Isolation of CO-modified Nucleosomes (BICON).

This technique combines in vivo biotinylation by BirA and H3 phosphorylation by MSK1,

allowing enrichment of phosphorylated nucleosomes using streptavidin. I found that MSK1-

phosphorylated nucleosomes are hyper-acetylated on H3 and H4, and importantly, I

identified a trans-tail crosstalk between H3 phosphorylation and H4 acetylation on K12. This

proof-of-principle study demonstrates that BICON can be further adapted to study PTMs and

crosstalks associated with other histone-modifying enzymes.

Taken together, work described in this thesis shows that histone H3 phosphorylation

can initiate additional PTM changes on other residues within the nucleosome, and such

crosstalk plays an important role in regulating gene expression.

iv

Acknowledgements

I would like to thank the many people who have influenced and supported me throughout my

years in graduate school. I would not have been able to go through this memorable journey

without you.

First and foremost, I would like to thank my supervisor, Dr. Peter Cheung, for your

continuous support, advice and guidance. Thanks for sharing with me your passion for

science. Your enthusiasm and excitement have motivated me to pursue scientific research.

Through our scientific discussions, I have learned a lot from you and you have helped me to

grow as a scientist.

I would like to thank my supervisory committee members, Dr. Sam Benchimol and Dr. Jim

Woodgett. Thanks for your support and guidance throughout my graduate studies. I always

feel positive and encouraged after my committee meetings.

I am very grateful to have Ryan and Cindy accompany me during this journey. It is great to

have you share the highs and lows of lab life with me. Thanks for listening to my complaints

and troubleshooting with me when my experiments didn’t work. Thanks for sharing my

happiness when my “band” showed up and when my ChIP worked! Life in the lab would

have been very different without the two of you.

A big thanks to my friends who have continuously supported me throughout the years. Thank

you Ginny, Debbie, Joy, Jen, Jack, Heidi, Christine, Eric, Norm, May, Frances and Carol.

Thanks for your friendship and thanks for sharing this part of my life journey with me. You

have made this journey more memorable and more enjoyable.

Finally, I must thank my parents and my sister Loretta for their love, patience and

understanding over the years. Without the unquestioning and continuous support from my

parents, I would not be where I am now. Thanks for allowing me to choose my own path and

believing in me every step of the way.

v

Table of Contents

Abstract ..................................................................................................................................... ii

Acknowledgements .................................................................................................................. iv

Table of Contents .......................................................................................................................v

List of Tables ......................................................................................................................... viii

List of Figures .......................................................................................................................... ix

List of abbreviations ............................................................................................................... xii

Chapter 1: Introduction .............................................................................................. 1

1.1 Thesis Overview ............................................................................................................2

1.2 Chromatin structure and organization ............................................................................2

1.2.1 Nucleosomes – the building block of chromatin ...............................................3

1.2.2 Heterochromatin and euchromatin .....................................................................3

1.2.3 Modulation of chromatin structure ....................................................................5

1.2.3.1 Covalent modifications on chromatin ........................................................5

1.2.3.2 Chromatin remodeling ................................................................................9

1.2.3.3 Histone Variants .......................................................................................10

1.3 Histone post-translational modifications .....................................................................15

1.3.1 Functional effects and dynamics of histone modifications ..............................17

1.3.2 Histone acetylation...........................................................................................18

1.3.3 Histone methylation .........................................................................................20

1.3.3.1 Lysine methylation ...................................................................................21

1.3.3.2 Arginine methylation ................................................................................24

1.3.3.3 Histone demethylation ..............................................................................25

1.3.4 Histone phosphorylation ..................................................................................26

1.3.5 Histone ubiquitylation, sumoylation and other modifications .........................30

1.4 Histone H3 phosphorylation and transcriptional regulation ........................................33

1.4.1 H3 phosphorylation during mitosis and interphase .........................................33

1.4.2 The first link between H3 phosphorylation and transcriptional activation ......34

1.4.3 MAPK pathway-mediated H3 phosphorylation & gene expression ................36

1.4.4 Other signaling pathways that target H3 ..........................................................38

vi

1.4.4.1 H3 phosphorylation and regulation of MYC target genes .......................39

1.4.4.2 H3 phosphorylation and nuclear receptor signaling .................................40

1.5 Mechanism of action of histone modifications ............................................................41

1.5.1 Structural effects of histone modifications ......................................................41

1.5.2 Recruitment of effector proteins to chromatin .................................................43

1.5.2.1 Acetyl- and methyl-binding domains .......................................................43

1.5.2.2 Phospho-histone binding proteins ............................................................46

1.6 Histone modification crosstalks ...................................................................................48

1.6.1 Modes and mechanisms of histone modification crosstalks ............................48

1.6.2 Histone H3 phosphoacetylation .......................................................................50

1.6.3 Combinatorial histone PTMs and multivalent binding ....................................52

1.7 Histone phosphorylation and human diseases .............................................................55

1.8 Thesis Rationale and Objectives ..................................................................................56

Chapter 2: Histone crosstalk involving H3 S28 phosphorylation and K27

acetylation activates transcription and antagonizes polycomb silencing ...... 58

2.1 Abstract ........................................................................................................................59

2.2 Introduction ..................................................................................................................60

2.3 Materials and Methods .................................................................................................62

2.4 Results ..........................................................................................................................66

2.4.1 H3 kinase MSK1, not RSK2, activates transcription of reporter .....................66

2.4.2 Targeting MSK1 to the endogenous c-fos promoter activates its expression ..70

2.4.3 MSK1 and H3 phosphorylation reactivate expression of polycomb-

silenced α-globin in non-erythroid cells ..........................................................75

2.4.4 H3 S28 phosphorylation disrupts PRC recruitment and H3 K27

methylation ......................................................................................................76

2.4.5 Coupling of H3 S28 phosphorylation and K27 acetylation antagonizes

polycomb silencing ..........................................................................................82

2.4.6 Both H3S28ph and H3K27ac/S28ph are directly associated with the

initiating form of RNAP II ...............................................................................85

2.5 Discussion ....................................................................................................................88

vii

Chapter 3: Elucidating combinatorial histone modifications and crosstalks by

coupling histone-modifying enzyme and biotin ligase activity ....................... 93

3.1 Abstract ........................................................................................................................94

3.2 Introduction ..................................................................................................................95

3.3 Materials and Methods .................................................................................................98

3.4 Results ........................................................................................................................101

3.4.1 Coupling of in vivo biotinylation and MSK1 phosphorylation ......................101

3.4.2 Affinity purification of BirA-biotinylated and MSK1-modified

nucleosomes ...................................................................................................107

3.4.3 MSK1-phosphorylated nucleosomes are enriched for 14-3-3ζ and H3/H4

acetylation ......................................................................................................110

3.4.4 H4 K12 acetylation is associated with activation of MSK1-target genes ......114

3.5 Discussion ..................................................................................................................118

Chapter 4: General Discussion and Future Directions ........................................ 122

4.1 Summary of findings..................................................................................................123

4.2 Role of H3S28ph and H3K27ac/S28ph during cellular differentiation .....................124

4.3 Mechanisms that couple H3 phosphorylation and H3/H4 acetylation.......................129

4.4 Future applications of BICON ...................................................................................132

4.4.1 Use of BICON to study other H3 kinases and histone-modifying enzymes..133

4.4.2 Use of BICON to identify histone PTM readers ............................................135

4.5 Concluding remarks ...................................................................................................141

References ................................................................................................................. 142

viii

List of Tables

Table 1.1. List of histone phosphorylation sites and histone kinases. ............................ 27

Table 2.1. Antibodies used for Western blot analyses and ChIP. .................................. 63

Table 2.2. Primer sequences used for RT-PCR and ChIP-qPCR analyses. ................... 65

ix

List of Figures

CHAPTER 1

Figure 1.1. Post-translational modifications on core histones............................................. 7

Figure 1.2. Amino acid sequence alignment of human histone H3.1, H3.2 and H3.3. ..... 13

Figure 1.3. Different types of post-translational modifications on histone residues. ........ 16

Figure 1.4. Histone H3 lysine methylation........................................................................ 22

Figure 1.5. Phosphorylation on histone tails plays important roles in multiple cellular

processes. ........................................................................................................ 29

Figure 1.6. Binding domains for modified histones. ......................................................... 44

Figure 1.7. Combinatorial reading of histone modifications............................................. 54

CHAPTER 2

Figure 2.1. MSK1, but not RSK2, phosphorylates H3 at serine 28 and activates

transcription of reporter. ................................................................................. 67

Figure 2.2. Transcriptional activation by Gal4-CA-MSK1 depends on its targeting to the

luciferase promoter. ........................................................................................ 69

Figure 2.3. Targeting MSK1 to the endogenous c-fos promoter induces H3 S10 and S28

phosphorylation and activates its expression. ................................................. 71

Figure 2.4. NF1-CA-MSK1 fusion induced expression of c-fos, HSP70 but not c-jun. ... 74

Figure 2.5. MSK1 and H3 phosphorylation reactivate expression of polycomb-silenced α-

globin gene in non-erythroid cells. ................................................................. 77

Figure 2.6. α-globin promoter in 293T cells is enriched for H3K27me3. ......................... 80

Figure 2.7. H3 S28 phosphorylation disrupts PRC (Polycomb Repressive Complex)

recruitment and H3 K27 methylation. ............................................................ 81

x

Figure 2.8. Functional and physical coupling of H3 K27 acetylation and S28

phosphorylation antagonizes polycomb silencing. ......................................... 83

Figure 2.9. Specificity of the α-H3K27ac/S28ph antibody was tested using peptide

competition assays. ......................................................................................... 84

Figure 2.10. Analysis of H3 phosphorylation marks in TPA- or anisomycin-induced

10T1/2 cells. .................................................................................................... 86

Figure 2.11. H3S28ph and H3K27ac/S28ph marks correlate with transcription initiation at

the activated c-fos and α-globin promoters. .................................................... 87

Figure 2.12. H3 S28 phosphorylation initiates a novel histone code path

way by inducing a

methyl-acetylation switch of the adjacent K27 residue. ................................. 91

CHAPTER 3

Figure 3.1. MSK1 preferentially phosphorylates H3 variant H3.3 on S28. .................... 103

Figure 3.2. Schematic of H3.3-AviFlag and BirA-Msk1 fusion constructs. ................... 105

Figure 3.3. In vivo biotinylation and phosphorylation of Avi-tagged H3.3. ................... 106

Figure 3.4. Workflow of the affinity purification method used for the isolation of co-

modified nucleosomes. ................................................................................. 108

Figure 3.5. Affinity-purification of BirA-biotinylated and MSK1-modified nucleosomes.

....................................................................................................................... 109

Figure 3.6. MSK1-modified nucleosomes are enriched for H3 acetylation and H4

acetylation. .................................................................................................... 111

Figure 3.7. MSK1-modified nucleosomes are enriched for 14-3-3ζ. .............................. 113

Figure 3.8. H4K12, but not H4K16, acetylation is induced upon activation of MSK1-

target genes. .................................................................................................. 115

xi

CHAPTER 4

Figure 4.1. MSK1 and H3 phosphorylation are involved in retinoic acid (RA)-induced

gene activation in mouse F9 teratocarcinoma cells. ..................................... 125

Figure 4.2. A model for the activation of polycomb-repressed genes by histone H3 S28

phosphorylation............................................................................................. 128

Figure 4.3. Histone acetyltransferases CBP and p300, but not PCAF, acetylate H3 on

K27. ............................................................................................................... 131

Figure 4.4. Isolation of effector proteins that bind to MSK1-modified nucleosomes..... 137

xii

List of abbreviations

AML Acute myeloid leukemia

AR Androgen receptor

ATP Adenosine triphosphate

AU Acid urea

Avi-HRP Avidin-horseradish peroxidase

BICON Biotinylation-assisted Isolation of CO-modified Nucleosomes

bp Base pair

BPTF Bromodomain and PHD finger-containing transcription factor

BRD4 Bromodomain-containing protein 4

BRG1 Brahma-related gene-1

CA Constitutively-active

CARM1 Coactivator-associated arginine methyltransferase-1

CBP CREB-binding protein

cDNA Complementary deoxyribonucleic acid

CHD1 Chromodomain helicase DNA binding protein-1

ChIP Chromatin immunoprecipitation

ChIP-seq Chromatin immunoprecipitation-sequencing

CLS Coffin-Lowry syndrome

CTKD C-terminal kinase domain

DBD DNA binding domain

DMEM Dulbecco’s modified eagle medium

DMSO dimethyl sulfoxide

DNA Deoxyribonucleic acid

DTT Dithiothreitol

EDTA Ethylenediaminetetraacetic acid

EGF Epidermal growth factor

EGTA Ethylene glycol tetraacetic acid

ERK Extracellular signal-regulated kinase

ES Embryonic stem

EZH2 Enhancer of zeste homolog-2

xiii

FBS Fetal bovine serum

GCN5 General control nonderepressible-5

HA Hemagglutinin

HAT Histone acetyltransferase

HDAC Histone deacetylase

HDM Histone demethylase

HMT Histone methyltransferase

HMGN1 High mobility group nucleosomal binding domain-1

HP1 Heterochromatin protein-1

IE Immediate-early

IFN-α Interferon-alpha

IKK-α Inhibitor of κB kinase-alpha

IP Immunoprecipitation

ING2 Inhibitor of growth family, member 2

ISWI Imitation switch

JMJD2A Jumonji domain containing 2A demethylase

JNK c-Jun N-terminal kinase

KD Kinase-dead

LSD1 Lysine-specific histone demethylase-1

MAPK Mitogen-activated protein kinase

MAPKKK Mitogen-activated protein kinase kinase kinase

MBT Malignant brain tumor repeat/domain

MEF Mouse embryonic fibroblast

Min. Minutes

MLL Mixed lineage leukemia

MMLV Moloney murine leukemia virus

MNase Micrococcal nuclease

MOF Males absent on the first

MS Mass spectrometry

MSK1/2 Mitogen- and stress-activated protein kinase-1/2

MST1 Mammalian Sterile20-like 1 kinase

xiv

MYST MOZ, Ybf2/Sas3, Sas2 and TIP60

NF1 Nuclear factor 1 / CCAAT box-binding transcription factor

NLS Nuclear localization signal

NTKD N-terminal kinase domain

NuRD Nucleosome remodeling and deacetylase

NURF Nucleosome-remodeling factor

PAGE Polyacrylamide gel electrophoresis

Pc Polycomb

PCAF p300/CBP-associated factor

PcG Polycomb group

PCR Polymerase chain reaction

PHD Plant homeodomain

PKA Protein kinase A

PKC Protein kinase C

PP2A Protein phosphatase type 2A

PRC1 Polycomb repressive complex 1

PRC2 Polycomb repressive complex 2

PRK1 Protein kinase C-related kinase 1

PRMT1 Protein arginine methyltransferase-1

P-TEFb Positive transcription elongation factor b

PTM Post-translational modification

PVDF Polyvinylidene fluoride

qPCR Quantitative real time polymerase chain reaction

RA Retinoic acid`

RNA Ribonucleic acid

RNAi RNA interference

RNAP II RNA polymerase II

RNase Ribonuclease

RSK2 p90 ribosomal S6 protein kinase 2

RT-PCR Reverse transcription-polymerase chain reaction

SDS Sodium dodecyl sulfate

xv

SET Supressor of variegation, enhancer of zeste and trithorax

SUMO Small ubiquitin-like modifier

SUV39H1/2 Suppressor of variegation 3-9 homolog-1/2

SWI/SNF Switch/sucrose non-fermentable

TE Tris-EDTA

TIP60 Tat-interactive protein

TNF-α Tumor necrosis factor-alpha

TPA 12-O-tetradecanoyl-phorbol-13-acetate

TrxG Trithorax group

TSA Trichostatin A

VEGF-A Vascular endothelial growth factor-A

1

Chapter 1

Introduction

A portion of this chapter (Chapter 1.3.4, 1.4-1.5, 1.7) is based on the following book chapter: Lau, P.N.I., and Cheung, P. (2009). Histone Phosphorylation: Chromatin Modifications that Link Cell Signaling Pathways to Nuclear Function Regulation. In Handbook of Cell Signaling 2nd edition, R. A. Bradshaw and E. A. Dennis, eds. (Oxford: Academic Press), pp. 2399–2408.

2

1 Introduction 1.1 Thesis Overview

Post-translational modifications (PTMs) on histones have been implicated in the

regulation of many fundamental cellular processes. These histone PTMs often do not

function in isolation; instead, extensive crosstalks occur between different modifications,

leading to specific combinations of histone PTMs on nucleosomes. Among them,

phosphorylation of histone H3 on Ser10 and Ser 28 has been linked to transcriptional

activation of a wide range of genes, such as induction of immediate-early genes upon stress

or mitogenic stimulation. Functional crosstalks between H3 phosphorylation and other

PTMs, such as acetylation, have also been reported. Chapter 1 covers these basic concepts in

the chromatin field, focusing specifically on histone H3 serine phosphorylation and its role in

transcriptional activation. In Chapter 2, I examine the role of H3 kinase MSK1 and H3

phosphorylation in antagonizing polycomb silencing, and the involvement of other histone

PTMs in the activation process. Chapter 3 describes a novel method that allows preferential

isolation of mono-nucleosomes that are modified by a specific histone-modifying enzyme of

interest. We utilized this technique to study crosstalk pathways initiated by the H3 kinase

MSK1. At the end of this thesis in Chapter 4, I will outline some new questions and future

experiments arising from this work, and highlight the importance of this current study in the

area of histone phosphorylation and in chromatin research.

1.2 Chromatin structure and organization

All eukaryotic cells have to face a challenge of organizing their lengthy genome into

the nuclei. The human genome, for example, consists of 3 billion base pairs of DNA arranged

into 23 chromosomes. In a diploid cell, this is roughly 2 metres in length when stretched out,

3

but it has to be packaged into a nucleus of less than 10µm in diameter. To achieve this high

level of compaction, genomic DNA is packaged into chromatin, a nucleoprotein structure

that serves as the physiological template for all DNA-related functions.

1.2.1 Nucleosomes – the building block of chromatin

The fundamental unit of chromatin is the nucleosome core particle, which is

composed of 146 base pair (bp) of DNA wrapped around a histone octamer containing 2

copies each of the 4 core histones – H2A, H2B, H3 and H4 (1, 2). All core histones are

highly conserved from yeast to humans and have a similar structure, with a C-terminal

histone fold domain and an unstructured N-terminal tail. The histone fold domain is involved

in histone-histone interactions within the nucleosome core, whereas the tails protrude in all

directions from the core particle, with the H3 tail being the longest. Within a nucleosome, the

core histones are arranged as a (H3-H4)2 tetramer flanked by two H2A-H2B dimers. The

binding of histone H1 to linker DNA facilitates folding of the nucleosome arrays into 30nm

fibers, and the association of non-histone chromosomal proteins further compacts the

chromatin fiber into higher-order structures. The most condensed form of chromatin is seen

in mitosis as metaphase chromosomes (3).

1.2.2 Heterochromatin and euchromatin

Even though this packaging of DNA into nucleosomes is repeated throughout the

genome, the overall organization and compaction of chromatin in a cell is not uniform.

Instead, the genome is organized into different chromatin states and contains specialized

structural elements such as centromeres and telomeres, which are essential for proper

chromosome segregation and replication respectively.

4

Chromatin can be broadly categorized into two functionally and structurally distinct

forms: heterochromatin and euchromatin. These two forms were originally identified

cytologically by Emil Heitz in the early 20th century, based on their differential compaction

in interphase nuclei (4). Heterochromatin corresponds to densely-stained, highly-condensed

chromosomal regions that are replicated late in S phase. These regions are associated with

transcriptional silencing and contain few actively-transcribed genes. In contrast, euchromatin

has a more “open” conformation and contains the transcriptionally permissive, gene-rich

portion of chromatin (5). Importantly, boundary elements separate heterochromatin and

euchromatin, and prevent improper transcription of genes in the heterochromatin-

euchromatin boundaries (6). They act as barriers to prevent spreading of heterochromatin to

euchromatic regions, and vice versa, as seen in the well-studied phenomenon of position-

effect variegation (PEV) in Drosophila (7).

Heterochromatin can be further classified into constitutive and facultative

heterochromatin. Constitutive heterochromatin mainly comprises of highly repetitive DNA

(transposons, satellite repeats) and includes structures that form centromeres and telomeres

(8). These highly compacted regions are present at all times in all cells of an organism.

Facultative heterochromatin, on the other hand, is established only in certain cell types to

ensure silencing of genes or regions during differentiation or at specific developmental stages

(9). One classic example of this type of heterochromatin is the inactive X chromosome

present in female mammalian cells. Heterochromatin was previously thought to be

transcriptionally inert; however, recent studies strongly suggest that transcripts from

heterochromatic regions are important components of the silencing mechanism. In the fission

yeast Schizosaccharomyces pombe, transcription of repeat elements and the RNA

interference (RNAi) pathway play a critical role in heterochromatin formation (10). Long

5

non-coding RNAs, such as Xist from the inactive X, are also involved in mediating

transcriptional repression in facultative heterochromatin (11).

1.2.3 Modulation of chromatin structure

The organization of DNA within chromatin successfully solves the basic packaging

problem; however, this poses another problem for the cell – the repressive nature of

chromatin restricts the access of DNA by regulatory factors, and thus affects all DNA-

templated processes such as transcription, DNA replication, DNA repair and recombination.

To enable these essential cellular functions, chromatin needs to be flexible and malleable for

use. Indeed, the chromatin fiber is a highly dynamic structure and it responds dynamically to

cellular signaling. Eukaryotic cells have developed multiple strategies to modulate chromatin

structure and to alleviate the repressive nature of chromatin. For example, histone-modifying

enzymes and DNA methyltransferases covalently modify histones and DNA, respectively, to

alter the basic nucleosome structure and to elicit a variety of downstream effects (12). In

addition, ATP-dependent chromatin-remodeling enzymes mobilize and reposition

nucleosomes to change accessibility of the chromatin fiber (13, 14). Finally, histone variants

replace core histones at strategic positions within the genome to confer specialized functions

(15–17). All together, these mechanisms allow the chromatin template to undergo dynamic

changes, thus enabling cells to alter gene expression profiles in response to extracellular

stimuli, and to accommodate on-going nuclear processes during cell growth and cell division.

1.2.3.1 Covalent modifications on chromatin

Numerous post-translational modifications (PTMs) have been identified on all four

core histones and the linker histone H1, particularly at their N-terminal tails. The best-studied

6

histone PTMs include acetylation (ac), methylation (me), phosphorylation (ph) and

ubiquitylation (ub) (Figure 1.1). Covalent modifications on histones were first observed in

the early 1960s, and the link between histone acetylation and regulation of RNA synthesis

was first proposed by Vincent Allfrey in 1964 (18). Since then, a large number of PTMs on

histones have been described, and they not only have been correlated with specific

transcriptional states, but are also implicated in various biological processes such as mitosis,

DNA repair and DNA replication (12, 19). Importantly, these modifications are highly

dynamic, and in response to different cellular conditions, histone-modifying enzymes add or

remove PTMs from specific amino acids on histones. Different chromatin states or genomic

regions are also associated with different sets of PTMs. For example, actively-transcribed

euchromatic regions of the genome contain hyperacetylated histones and high levels of

methylation on H3K4, H3K36 and H3K79. In contrast, silent heterochromatin is associated

with hypoacetylated histones, with H3K9 methylation on constitutive heterochromatin and

H3K27 methylation on facultative heterochromatin (20). It is now well-established that

histone PTMs mostly mediate these processes through their direct effects on chromatin

structure or through the recruitment of downstream effector proteins to chromatin. Histone

PTMs and their mechanisms of actions will be discussed in more details in Sections 1.3-1.6.

Another covalent modification on chromatin involves the methylation of DNA at C5

position of cytosines to generate 5-methylcytosine (5-mC) (21, 22). In mammals, this occurs

almost exclusively at CpG dinucleotides and is implicated in genomic imprinting, X-

chromosome inactivation and control of gene expression. The majority of CpG sites in the

mammalian genome are methylated and these regions generally exhibit a “closed” chromatin

structure. In particular, DNA methylation in repetitive and transposable elements suppresses

7

Figure 1.1. Post-translational modifications on core histones. Amino acid sequences of the histone N- and C-terminal tails are shown. Most known post-translational modifications (PTMs) are found on the N-terminal tails of histones. Some of these PTMs also occur on the C-terminal tails of H2A and H2B, as well as within the globular domains of histones. Residue numbers are shown below each sequence. P, phosphorylation; Ac, acetylation; Me, methylation; Ub, ubiquitylation. Adapted from (23).

8

recombination and is essential for chromosomal integrity. On the other hand, unmethylated

CpG residues are mostly localized to CpG islands, which are short regions rich in CpG sites.

These islands are found in ~60% of human gene promoters, and promoter methylation

strongly correlates with transcriptional silencing. Even though DNA methylation

predominantly occurs in the CG context in mammals, genome-wide DNA methylation

analysis shows that non-CG methylation is abundant in embryonic stem (ES) cells (24). It is

enriched within gene bodies of highly expressed genes, and its level decreases during

differentiation, suggesting that it plays a role in maintaining the pluripotent state. In addition

to 5-mC, other modifications on cytosine bases have recently been described and include 5-

hydroxymethylcytosine, 5-formylcytosine and 5-carboxylcytosine (25, 26). They are thought

to be successive oxidative products of 5-mC and may represent intermediates in the DNA

demethylation pathway (27). The role of non-CG methylation and these other forms of

modified cytosines in genome regulation is currently unclear and needs to be further studied.

In fact, DNA methylation and certain histone modifications (such as histone

deacetylation, H3K9 methylation) can direct and influence each other. Transcriptional

repression by DNA methylation is indeed partly mediated through the recruitment of histone

deacetylases (HDACs) by methyl-CpG-binding proteins (28, 29). Such direct physical and

functional links between components of the DNA methylation and histone modification

machineries (enzymes and binding proteins) have been illustrated in various organisms

including Arabidopsis, Neurospora, mouse and human (30–33). Together, covalent

modifications on both components of chromatin (histones and DNA) often cooperate to

regulate chromatin structure and function (34).

9

1.2.3.2 Chromatin remodeling

Another mechanism to alter chromatin structure is through the action of chromatin

remodeling enzymes. These enzymes all function in the context of multi-subunit complexes,

each with a catalytic ATPase subunit that belongs to the SF2 superfamily of helicases and

multiple accessory proteins (13, 14). Using energy from ATP hydrolysis, these remodeling

complexes can disrupt DNA-histone contacts and thus alter the position and composition of

nucleosomes on chromatin. This can be mediated through multiple remodeling activities,

which include sliding of nucleosomes along DNA, exchange of histone variants and

complete eviction of nucleosomes from chromatin. In this way, these ATP-driven

machineries can modulate the accessibility of the underlying DNA to cellular proteins.

Based on the presence of distinct structural domains on the ATPase subunit,

chromatin remodeling complexes can be divided into 4 subfamilies: SWI/SNF, ISWI, CHD

and INO80. These ATPases, as well as their associated subunits, are highly conserved from

yeast to humans, and the first discovered chromatin remodeling complex was the yeast

SWI/SNF complex. It was identified in two independent genetic screens for mutants defective

in mating-type switching (switch) and growth on sucrose (sucrose nonfermenting), and was

found to be essential for expression of genes required for these processes (35, 36). At gene

promoters and enhancers, positioned nucleosomes are often present and cover essential

regulatory elements. Repositioning or removal of nucleosomes exposes these DNA-binding

sites and allows binding of transcription factors and transcription machinery. Besides having

a role in transcriptional activation, chromatin remodeling complexes can also organize

nucleosome positioning to induce gene repression and have roles beyond transcription. The

INO80 subfamily, for example, is involved in a wide range of processes and has crucial

functions in maintaining genomic stability (37). It is recruited to sites of DNA double strand

10

breaks and facilitates nucleosome eviction around these regions, thus allowing the

recruitment of repair factors. It is also required for DNA replication, telomere maintenance

and chromosome segregation.

Covalent modifications of histones described above and non-covalent mechanisms

mediated by ATP-dependent remodeling complexes are often interdependent. Subunits of

chromatin remodeling complexes contain multiple structural domains that bind specifically to

histone modifications, suggesting that these complexes can be targeted through specific

histone modification patterns to selective chromatin regions (38). The identification of PTMs

on nucleosome lateral surfaces also implies that chromatin remodeling is required for

histone-modifying enzymes to gain access to these modified residues. This coupling between

histone modification and chromatin remodeling is best illustrated by the NuRD (nucleosome

remodeling and deacetylase) repressor complex, which contains an ATPase of the CHD

family and histone deacetylases, and therefore, possesses both enzymatic activities within the

complex (39). All these suggest that the two types of chromatin modifying activities can be

physically and functionally coupled to regulate chromatin dynamics and accessibility.

1.2.3.3 Histone Variants

Whereas the majority of nucleosomes in the cell are composed of the same 4 types of

core histones (H2A, H2B, H3 and H4), tremendous diversity in the chromatin structure is

generated through the incorporation of variant versions of these histones into nucleosomes

(15, 16, 40). These variants are non-allelic isoforms of core histones and replace their

canonical histone counterparts at strategic positions in the genome. The difference between

core and variant histones can range from a few amino acid changes (as in H3 variant H3.3) to

significant structural differences (such as the additional macrodomain in H2A variant

11

macroH2A).

One distinguishing difference between canonical and variant histones is their timing

of expression and incorporation into chromatin. Expression of canonical histones is tightly

regulated throughout the cell cycle, and is restricted to S phase to coincide with DNA

replication. They are transcribed from tandem, multicopy clusters and their mRNAs are

intronless (41). In place of a poly-A tail, replication-dependent histone mRNAs end in a

conserved 3’ stem-loop structure that is crucial for their regulation and efficient translation

(42). All these features allow rapid synthesis of histones during S phase and ensure proper

packaging of newly-synthesized DNA. In contrast, histone variant transcripts are encoded by

single-copy genes that contain introns and are polyadenylated. They can be expressed

throughout the cell cycle and are assembled into chromatin in a replication-independent

manner, which suggests that these replacement histones have specialized functions beyond

DNA packaging.

Variant counterparts have been identified for all core histones. Among them, most

studies have focused on understanding various H2A and H3 variants, which have been

implicated in diverse functions in the cell. Variants of H2B and H4, on the other hand, are

mostly tissue-specific and are much less studied. Four H2A variants have now been

identified in mammalian cells: H2A.X, H2A.Z, macroH2A and H2A.Bbd (43). H2A.X is

distributed throughout the genome and is phosphorylated in response to DNA damage. It

functions to recruit DNA repair factors, and is an essential component of the DNA damage

response pathway (44). H2A.Z, on the other hand, has been linked to numerous biological

processes, including both transcriptional activation and repression, maintenance of

euchromatin-heterochromatin boundary, as well as chromosome segregation (45). It is highly

conserved across the phylogenetic tree, and is essential for viability in all organisms studied,

12

except yeast. Unlike H2A.Z, macroH2A and H2A.Bbd are restricted to vertebrates.

MacroH2A predominantly localizes to the inactive X chromosome in female mammalian

cells and is associated with gene repression (46, 47). This is in contrast to H2A.Bbd, which is

excluded from the inactive X and has a positive role in transcription (48).

Mammalian cells have two canonical H3, H3.1 and H3.2 (49). These two isoforms

only differ by one amino acid (Figure 1.2), and are often collectively referred to as H3 in the

literature. To date, in addition to canonical H3.1 and H3.2, a total of six H3 variants have

been described in mammals: centromeric H3 variant CENP-A, transcription-linked H3.3, two

testis-specific variants, H3t and H3.5, as well as two newly-identified primate-specific

variants, H3.X and H3.Y (50–52). Among these variants, CENP-A and H3.3 have been most

intensively studied. CENP-A localizes exclusively to centromeres and replaces H3 in

centromeric chromatin (53). This centromere-specific H3 variant is evolutionarily conserved

and can be found in all eukaryotes. It is required for kinetochore assembly and chromosome

segregation during mitosis. H3.3 is the predominant form of histone H3 variant, and

generally constitutes ~20-50% of total H3 in mammalian cells (54). It differs from canonical

H3.1 and H3.2 only by five and four amino acids respectively (Figure 1.2), and the amino

acid variation between residues 87-90 is necessary and sufficient for its replication-

independent assembly onto chromatin (55). H3.3 has long been associated with sites of active

transcription and is particularly enriched in covalent modifications associated with active

chromatin, such as H3 acetylation and H3K4/K36/K79 methylation (56, 57). Consistent with

this, its incorporation at coding regions is required for transcriptional activation of inducible

genes, such as interferon-stimulated genes (58, 59). Genome-wide chromatin

immunoprecipitation (ChIP) studies revealed that H3.3 is not only present at high levels

throughout the gene body of actively-transcribed genes, but is also found at promoters of

13

Figure 1.2. Amino acid sequence alignment of human histone H3.1, H3.2 and H3.3. Canonical H3.1 and H3.2 differ only at amino acid 96 (cysteine in H3.1 and serine in H3.2) and their expression is restricted to S phase of the cell cycle. Histone variant H3.3 is incorporated into chromatin in a DNA replication-independent manner, and it differs from canonical H3.1 and H3.2 by five and four amino acids respectively. Differences from H3.1 are highlighted in red.

14

both active and inactive genes, as well as at intergenic regulatory elements such as

transcription factor binding sites (60–62). Surprisingly, H3.3 is also detected at telomeres in

mouse ES cells and at pericentric heterochromatin in mouse embryonic fibroblasts (MEF)

(63, 64). Recent studies also demonstrated that multiple chaperones and deposition pathways

are involved in H3.3 deposition at specific genomic regions (64). Whereas H3.1 is deposited

by histone chaperone CAF-1 onto newly-replicated DNA during S phase, H3.3 is mainly

incorporated into genic regions of chromatin by another chaperone HIRA in a replication-

independent manner (55, 64). H3.3 enrichment at telomeres and most regulatory elements,

however, is HIRA-independent. Instead, the ATRX-Daxx complex is responsible for H3.3

localization at telomeres and is required for repression of telomeric RNA (64, 65); factors

involved in H3.3 enrichment at regulatory elements are yet to be identified. The unexpected

enrichment of H3.3 at heterochromatin and telomeres suggests that H3.3 is not only a mark

of active transcription, further studies are required to understand the functional relevance of

H3.3 at different genomic loci.

Together, histone variants, chromatin modifications and ATP-dependent chromatin

remodeling complexes form the major factors that regulate chromatin structure and functions.

They play critical roles in establishing different chromatin states and are major players in

epigenetic regulation. Importantly, these mechanisms are tightly inter-connected. For

example, histone variants are subjected to the same post-translational modifications as core histones,

and are often incorporated into chromatin by ATP-dependent chromatin remodeling complexes.

Recruitment of these remodelers to chromatin can also be mediated through PTMs on histones (38).

In most cases, these mechanisms work in concert to facilitate or restrict access to underlying

DNA, thus contributing to the dynamic nature of chromatin and providing remarkable

opportunities for the regulation of cellular processes.

15

1.3 Histone post-translational modifications

All four core histones and the linker histone H1 are heavily modified by a variety of

post-translational modifications (PTMs). Majority of these known PTMs are localized to the

unstructured N-terminal histone tails, but an increasing number of PTMs have been identified

within the histone globular domains as well as on the C-terminal tails of H2A and H2B

(Figure 1.1). The types of PTMs range from the addition of small chemical groups (such as

acetyl, methyl, phosphate groups) to the attachment of small polypeptides (such as ubiquitin,

SUMO) at specific amino acid residues on histones (Figure 1.3). Due to the highly conserved

nature of histones and their PTMs throughout evolution, our knowledge on chromatin and

their functions has come from studies in a wide range of model organisms such as

Saccharomyces cerevisiae, Schizosaccharomyces pombe, Drosophila melanogaster, mouse,

etc.

It has been almost half a century since the first discovery of acetylation, methylation

and phosphorylation on histones and the first indication that such histone modifications could

affect gene expression (18). However, it is not until recent years that we begin to understand

the diversity and complexity of covalent histone modifications and their major involvement

in regulating not only transcription, but also other key cellular processes. Development of

modification-specific antibodies and advances in mass spectrometry technology have

significantly expedited the discovery of novel histone PTMs and the identification of new

modification sites. Currently, more than 10 different types of modifications at nearly 130

different sites have been reported on histone tails and the core domains. In the past 15 years

or so, not only has the list of modified histone residues grown considerably, collective efforts

from many labs have identified diverse enzymes and pathways that target histones, and have

elucidated elegant mechanistic and functional details associated with specific histone marks.

16

Figure 1.3. Different types of post-translational modifications on histone residues. Amino acid residues are shown in blue, and modifications are shown in black. Phos, phosphorylation; O-GlcNac, β-N-acetylglucosamine glycosylation; Ub, ubiquitylation; Sumo, sumoylation; Methyl, methylation; Acetyl, acetylation; Pr, propionylation; Bu, butyrylation; Fo, formylation; Cr, crotonylation ; Cit, citrullination/deimination; ADP-ribose, ADP-ribosylation.

17

1.3.1 Functional effects and dynamics of histone modifications

The vast array of histone modifications has distinct functional effects and can

generate both short-term and long-term outcomes (19, 66). On one hand, histone PTMs play

crucial roles in short-term, ongoing cellular processes such as transcription, DNA replication

and repair. In response to internal or external stimuli, patterns of histone modifications at

specific genomic loci change rapidly, and therefore, they can be regarded as a continuation of

intracellular signal transduction. This directly connects cellular inputs to chromatin, allowing

the regulation of various DNA-templated processes in the nucleus. In addition to supporting

these short-term signaling processes, some histone PTMs can also participate in large scale

genome organization and have longer-term effects on the genome (5). This involves

partitioning the genome into euchromatin and heterochromatin, and the formation of

specialized chromatin structures, such as inactive X chromosome in female mammalian cells.

Once established, these chromatin states and domains are stable and can be inherited through

cell divisions. Histone PTMs have also been implicated in the establishment and maintenance

of stable transcriptional states and gene expression patterns that define cellular identity;

however, their role in inheritance and transmission of epigenetic information from one cell

generation to the next remains to be demonstrated and is a subject of intense investigation.

Importantly, modifications on histones are not static, but respond dynamically to

changes in cellular conditions and during development. It is now clear that most, if not all,

histone PTMs are reversible, even though different types of modifications may have

remarkably different turnover rates (67). Some modifications, including acetylation and

phosphorylation, are highly dynamic and are often studied in the context of inducible gene

expression (68, 69). Other modifications, such as methylation, are found to turn over much

more slowly and are generally involved in the long-term maintenance of gene expression

18

status and global chromatin environment (70). This addition and removal of histone PTMs is

a highly-regulated process, and is mediated by two classes of antagonizing histone-modifying

enzymes which are sometimes referred to as “writers” and “erasers” of histone marks

respectively (71). Many of these enzymes exist as part of multi-subunit complexes, and other

subunits within the complexes often play critical roles in controlling enzyme recruitment,

activity and substrate specificity. In addition to modifying histones, these “histone-modifying

enzymes” are also known to target other cellular, non-histone proteins, including

transcription factors, transcriptional co-regulators, and the modifiers themselves. Such

regulation of histone-modifying enzymes and their associated proteins by various PTMs adds

an additional layer of regulation to the histone modification process. Ultimately, it is through

the delicate balance between enzymes with opposing activities that dictates the stability and

dynamics of individual histone PTMs, thus providing an appropriate chromatin landscape

required for specific biological processes.

1.3.2 Histone acetylation

Acetylation is one of the first identified and intensively-studied modifications on

histones (18). The acetylation process is mediated by histone acetyltransferases (HATs),

which transfer an acetyl group from cofactor acetyl-coenzyme A to the ε-amino group of

lysine side chains, neutralizing the positive charge on this residue (72). There are two major

classes of HATs: type-A and type-B. Type-B HATs function in the cytoplasm and their

activities are mostly coupled to DNA replication. They acetylate newly-synthesized, free

histones before their incorporation into chromatin, and this is important for histone

deposition. In particular, acetylation of newly-synthesized H4 at K5 and K12 is

evolutionarily conserved from yeast to humans; acetylation on H3, however, occurs at

19

different sites in different species (73).

Type-A HATs, on the other hand, acetylate nucleosomal histones in the nucleus, and

are mostly associated with transcriptional activation. Based on their catalytic domains, they

can be grouped into three families: GNAT, MYST and CBP/p300 families (74). Most

characterized acetylation sites fall within the histone N-terminal tails, and include lysines 9,

14, 18, 23 and 27 on histone H3, as well as lysines 5, 8, 12 and 16 on histone H4. One

exception is H3 K56, which is found within the core domain and has been linked to

nucleosome assembly and DNA repair. Most HATs target more than one residue on histones,

and the exact target site often depends on other subunits within the HAT complex. Generally

speaking, GNAT family mostly targets H3 as the main substrate, whereas MYST family

mainly targets H2A/H4, and CBP/p300 family can acetylate both H3 and H4. A major

breakthrough in the field of chromatin research came when the first nuclear, transcription-

linked HAT, p55, was purified and cloned from transcriptionally-active macronuclei of

Tetrahymena thermophila (75). It was found to be a homolog of yeast Gcn5, a well-studied

transcriptional coactivator, thereby directly linking histone acetylation to gene activation

(76). At around the same time, the first histone deacetylase (HDAC), which opposes the

action of HATs, was isolated from mammalian cell extracts, and found to be related to yeast

transcriptional repressor Rpd3 (77). Together, these two discoveries provided the first direct

evidence that histone PTMs and histone-modifying enzymes play a key role in transcriptional

regulation, and chromatin does not only function as a packaging factor of the genome.

Interests in identifying and understanding the role of other histone PTMs and their modifying

enzymes in regulating gene expression and other cellular processes have since exploded.

Although histone acetylation predominantly occurs at promoters and 5’end of

transcriptionally-active genes, certain acetyl marks and HATs are also present within gene

20

bodies and other regulatory regions, suggesting that histone acetylation may be involved at

various stages of transcription (20, 78). For example, H3K27ac and its HAT p300 are highly

enriched at enhancers, and they are now often used to predict functional enhancers in the

genome. In contrast to HATs, HDACs remove acetyl groups from lysines, and are

predominantly transcriptional corepressors (79). There are 4 different classes of HDACs:

class I, II, III and IV. Class III (sirtuins) is particularly different from the other classes

because it requires NAD+ as a cofactor for its catalytic activity. HDACs are present in a

number of repressor complexes such as NuRD and Co-REST, and often cooperate with other

histone modifying enzymes within the complexes to mediate transcriptional repression.

Interestingly, histone deacetylation by yeast Rpd3 histone deacetylase in coding regions is

required to prevent aberrant transcription from intragenic cryptic initiation sites (80). An

appropriate balance between the activities of HATs and HDACs is therefore necessary for

accurate gene expression.

1.3.3 Histone methylation

Methylation occurs on lysine and arginine residues, and does not alter the charge of

these amino acids. Among all the PTMs on histones, methylation can be considered as the

most complex one, since methylation can exist in multiple states: lysines can be mono-, di- or

tri-methylated, whereas arginines can be mono-, symmetrically or asymmetrically di-

methylated (81). In contrast to histone acetylation and phosphorylation, which are well

documented to correlate with transcriptional activation, the effect of histone methylation is

more site-specific and context-dependent, and can be involved in gene activation or

repression. Methylation on different residues and different degrees of methylation often have

different functional implications.

21

1.3.3.1 Lysine methylation

The major sites of lysine methylation on histones include H3K4, H3K9, H3K27,

H3K36 and H4K20 (17, 82, 83). Among them, H3K4, H3K36 and H3K79 methylation are

associated with active genes, whereas methylation at H3K9, H3K27 and H4K20 has been

linked to transcriptional silencing (Figure 1.4). Utilizing S-adenosylmethionine (SAM) as

methyl donor, histone lysine methyltransferases (HMTs or KMTs) transfer methyl groups to

ε-amino groups on lysines. The activity of HMTs is carried out by the evolutionarily

conserved SET domain, named after the 3 founding members that share this domain:

Su(var)3-9, Enhancer of zeste and Trithorax. The only non-SET domain-containing HMT

identified to date is Dot1, which methylates H3K79 in the histone globular domain. In

contrast to HATs, HMTs exhibit high specificity toward their target lysine, and can often

discriminate between different degrees of methylation. For instance, yeast Set1, and its

human homologs SET1A/B and MLL1-4, can catalyze all three degrees of methylation on

H3K4 (H3K4me1/2/3); on the other hand, mono-methylation of H4K20 (H4K20me1) is

mediated by PR-Set7/Set8, and SUV4-20H1/2 are responsible for di- and tri-methylation of

the same residue (H4K20me2/3) (84).

The biological significance of histone lysine methylation first came to light when it

was found that some of the well-studied Su(var) genes, originally identified by genetic

screens to suppress position effect variegation in Drosophila, are indeed components of the

histone methylation pathway. Su(var)3-9 and its mammalian homologs encode a HMT that

selectively methylates H3K9, and this creates a motif that is specifically bound by the

product of Su(var)2-5, heterochromatin protein 1 (HP1) (85, 86). Importantly, these proteins

have long been known to have roles in constitutive heterochromatin formation and gene

silencing, and these findings therefore help establish that histone methylation has a direct role

22

Figure 1.4. Histone H3 lysine methylation. Enzymes that methylate lysines on H3 are shown above the H3 amino acid sequence. All histone methyltransferases (HMT), except Dot1, contain the SET domain as the catalytic domain. Examples of proteins and binding-modules (in parentheses) that specifically bind methylated lysines on histones are highlighted in blue. Proteins repelled by methylated lysines are shown in red. Txn, transcription. [Modified from (17)]

23

in regulating chromatin structure as well as gene expression.

Methylation of H3K27 is another PTM linked to transcriptional repression. In

contrast to H3K9 methylation, which is a component of constitutive heterochromatin, H3K27

methylation is associated with facultative heterochromatin and polycomb silencing (87). The

polycomb silencing pathway plays critical roles during differentiation and development. First

discovered in Drosophila, polycomb group (PcG) proteins were found to be crucial for

repression of HOX genes, and PcG mutations were associated with homeotic transformation

and development defects in Drosophila (88–90). In addition to HOX genes, PcG groups are

also important regulators for a large number of key developmental genes (91). They have

now been implicated in a broad range of biological processes including X chromosome

inactivation, genomic imprinting, cell cycle control, stem cell pluripotency and tumorigenesis

(92–94). PcG proteins are highly conserved from plants to flies to humans, and are often

found in two multi-subunit complexes, Polycomb Repressive Complex 1 (PRC1) and

Polycomb Repressive Complex 2 (PRC2). EZH2 is the catalytic subunit of PRC2 and

methylates H3 at K27, but its activity is regulated by other non-catalytic subunits within the

complex, which includes EED, SUZ12 and NURF55/RpAp48 (95). Methylated H3K27 in

turn recruits the PRC1 complex through its chromodomain-containing Polycomb (Pc)/CBX

proteins. This second complex contains the RING1b E3 ligase which mono-ubiquitylates

histone H2A, and, together, these histone modifications direct the polycomb-silencing

pathway to stably repress gene expression. Trithorax group (TrxG) proteins, on the other

hand, are positive regulators of HOX genes, and antagonize polycomb silencing (96). TrxG

proteins are later found to be components of the H3K4 methylation pathway.

Even though methylation of H3 on K4, K36 and K79 have all been linked to

transcriptionally active regions in euchromatin, they each have specific distribution patterns

24

and are found at different regions of a transcribing gene (20). For instance, H3K4 di- and tri-

methylation (H3K4me2/3) predominantly occur at regions around the transcription start site

(TSS), with H3K4me2 extending further into the gene body, whereas H3K4me1 is found

near the 3’ end of genes as well as at enhancer elements. On the other hand, H3K36me3 is

distributed over the transcribed region of active genes, and peaks at the 3’ end. This is

consistent with its role in transcriptional elongation, and recently, this mark has also been

reported to function in alternative splice site selection (97). Interestingly, activating and

repressive methyl marks are not always mutually exclusive in the genome. In ES cells,

bivalent domains, consisting of peaks of H3K4me3 within broader regions of H3K27me3,

are often found at key developmental genes (98). The bivalent chromatin signature is

proposed to keep these loci at a repressed state in ES cells, yet poised for lineage-specific

transcription in lineage-committed cells. Upon differentiation, most of these bivalent

domains are resolved into H3K4me3 or H3K27me3-only regions, in a manner consistent

with their subsequent expression status in the differentiated cells.

1.3.3.2 Arginine methylation

Arginine methylation is catalyzed by another group of methyltransferases known as

protein arginine methyltransferases (PRMTs), and they are classified into two classes based

on their enzymatic activities. Both classes can monomethylate arginines (Rme1), but type I is

responsible for asymmetric dimethylation (Rme2as) and type II catalyzes symmetric

dimethylation (Rme2s). Type I enzymes PRMT1 and CARM1 (also known as PRMT4)

methylate H4R3 and H3R17/23 respectively, and are the best-studied PRMTs. They are

mostly studied in the context of nuclear receptor-mediated transcription, acting as

coactivators for expression of estrogen receptor (ER)- or androgen receptor (AR)-regulated

25

genes (99, 100). Similar to lysine methylation, arginine methylation can be associated with

transcriptional activation as well as repression, depending on the specific site of modification

and the methylation state. For instance, asymmetric methylation of H4R3 by PRMT1 is

involved in activating ER-regulated genes; however, when the same residue is symmetrically

methylated by PRMT5, H4R3me2 functions as a repressive mark instead (101–103).

1.3.3.3 Histone demethylation

For many years, methylation has been considered a stable and static modification.

Early studies found that half-life of methyl marks is similar to that of the protein itself,

suggesting that this modification is irreversible. Removal of histone methylation was

therefore proposed to occur only through histone tail clipping, histone exchange or when

histones are diluted through DNA replication (104). This view, however, was challenged in

2004 when enzymes that remove methylated arginines and methylated lysines were

identified. The first enzyme reported to remove methyl marks on histones is PADI4 (105,

106). PADI4 functions as a transcriptional repressor in ER-regulated genes and antagonizes

the actions of CARM1 and PRMT1 on H3R17 and H4R3 respectively. Functionally, PADI4

is not a demethylase, but a deiminase that converts mono-methylated arginines to citrullines.

This pathway does not regenerate unmodified arginine, and therefore, cannot be considered

as a direct reversal of methylation. The citrullines are then either replaced or converted back

to unmodified histones by an unknown mechanism.

As for lysine demethylation, the first identified histone lysine demethylase (KDM) is

LSD1 (107). LSD1 is an amine oxidase which utilizes FAD as cofactor to convert mono- or

di-methylated lysines into unmethylated forms, but it has no enzymatic activity toward tri-

methylated substrates. LSD1 was initially identified as a component of the Co-REST

26

repressor complex and demethylates H3K4me1/2 at target genes (107). Interestingly, LSD1

can also function as an activator when in complex with AR and recruited to AR-regulated

genes, where it demethylates H3K9me1/2 instead (108). This clearly illustrates that activity

and specificity of histone modifying enzymes can be influenced by other factors within the

complexes in which they reside. The second class of KDMs is the Jumonji C (JmjC)-domain

containing demethylases (JMJD) (109–111). Members of this family are Fe(II) and α-

ketoglutarate-dependent dioxygenases, and they employ a different catalytic mechanism from

that used by LSD1 for demethylation. The enzymatic activity resides within the JmjC

domain, and importantly, this family is capable of demethylating all three lysine methylation

states (Kme1/2/3). Similar to histone methyltransferases, KDMs are often highly specific for

particular lysine residues as well as degrees of methylation. The recent discovery and studies

of histone demethylases confirm that, similar to other histone PTMs, methylation is also a

dynamically-regulated modification on histones.

1.3.4 Histone phosphorylation

Phosphorylation is one of the most commonly occurring post-translational

modifications on proteins and specific phosphorylation sites have been identified on all 4

core histones, as well as the linker histone H1 (See Table 1.1 for a list of histone

phosphorylation sites and histone kinases) (reviewed in 112–114). Even though

phosphorylation of serine and threonine residues on histones is well-documented, only

recently has tyrosine phosphorylation been identified on H3 and variant H2A.X (115, 116).

Histone phosphorylation is often a direct outcome of activated intracellular signaling

pathways, and functions to translate extracellular signals into appropriate nuclear biological

outputs such as changes in gene expression, response to DNA damage, and chromatin

27

Table 1.1. List of histone phosphorylation sites and histone kinases.

Histone Phosphorylation site Kinase Organism Transcriptional activation H2B S36 AMPK Mammals H3 T6 PKCα, β Mammals S10 Snf1 S.cerevisiae JIL-1 D.melanogaster RSK2, MSK1/2, COT,

IKKa, PIM1, PKA, JNK Mammals

T11 PRK1, CHK1, PKM2 Mammals S28 MSK1/2, MLTKa Mammals Y41 JAK2 Mammals DNA damage response H2A S129 Tec1, Mec1 S.cerevisiae H2A.X S139 ATM, ATR, DNA-PK Mammals Y142 WSTF Mammals H2B S14 ? Mammals H4 S1 CKII S.cerevisiae Mitosis H2A S1 ? C.elegans, D.melanogaster, mammals T119 NHK-1 D.melanogaster H3 T3 Haspin Mammals S10 Ipl1 S.cerevisiae NIMA A.nidulans AIR-2 C.elegans Aurora B Mammals T11 Dlk/ZIP Mammals S28 Aurora B Mammals H3.3 S31 ? Mammals CENP-A S7 Aurora A/B Mammals H4 S1 ? C.elegans, D.melanogaster, mammals Apoptosis H2B S10 Ste20 S.cerevisiae S14 MST1 Mammals H3 T45 PKCδ Mammals

Updated from Table 288.1 in (112).

28

condensation during apoptosis and mitosis (Figure 1.5).

Phosphorylation of H2A variant H2A.X at S139 (γ-H2A.X) is one of the best-studied

histone phosphorylation events. Upon DNA damage, members of the PIKK family (ATM,

ATR, DNA-PK) rapidly phosphorylate H2A.X at sites of double strand breaks (117–119).

DNA repair factors, such as BRCA1, 53BP1 and MRN complex, then co-localize with

γH2A.X to form discrete nuclear foci. Mice lacking H2A.X are radiation-sensitive and

exhibit genome instability (120, 121). Moreover, they show defects in repair pathways and

fail to recruit repair proteins to nuclear foci. Therefore, H2A.X is required for coupling the

DNA damage sensing pathways to the DNA repair response. Recently, another

phosphorylation site has been identified at the C-terminal tail of H2A.X. Y142 on H2A.X is

constitutively phosphorylated under normal growth conditions, but is dephosphorylated in

response to DNA damage (116). This modulates γ-H2A.X levels and is required for repair

foci formation and maintenance during DNA damage response. Exposure of yeast cells to

DNA damage inducers also leads to H4 phosphorylation at S1 and this phosphorylation event

participates in the non-homologous end joining repair pathway (122). In-gel kinase assays

identified casein kinase II (CK2) as the kinase for H4S1, and temperature-sensitive CK2

mutants not only are defective for H4S1 phosphorylation, but are hyper-sensitive to DNA-

damaging agents. This DNA damage-induced H4 phosphorylation is conserved in

mammalian cells. DNA damaging agents such as methyl methanesulfonate (MMS), ionizing

(IR) and ultraviolet (UV) radiation all strongly induce H4S1 phosphorylation (personal

observation); however, this modification does not appear to be mediated by CK2 in the

mammalian system. In addition, upon induction of apoptosis, H2B is phosphorylated at S14

by caspase 3-activated MST1 kinase, and this has been suggested to have a potential role in

apoptotic chromatin condensation and/or DNA fragmentation (123).

29

Figure 1.5. Phosphorylation on histone tails plays important roles in multiple cellular processes. Phosphorylation sites associated with transcriptional activation, mitosis, DNA damage response and apoptosis, as well as their respective kinases, are indicated. Updated from Figure 288.1 in (112).

30

During mitosis, most histones become highly phosphorylated at multiple

serine/threonine residues. The reported mitosis-linked histone phosphorylation sites include:

H2A/H4S1 (124), H2AT119 (125), H3T3 (126), H3S10 (127), H3T11 (128), H3S28 (129),

H3.3S31 (130) and CENP-A S7 (131, 132). The functional significance of the high and

extensive levels of histone phosphorylation during mitosis is currently not well understood.

Earlier studies in Tetrahymena suggested that H3S10ph is required for proper chromosome

condensation and segregation during mitosis (133). However, this function does not appear to

be conserved in yeast since the H3S10A mutant does not show any growth defect and is able

to progress through the cell cycle normally (134). There are also substantial spatial and

temporal differences amongst the distinct mitosis-associated histone phosphorylation events.

Phosphorylation events are initiated at different phases of the cell division, and are localized

to different regions of the mitotic chromosomes. For example, H3T11ph occurs from

prophase to anaphase and is enriched in centromeres (128), whereas H3.3 S31ph is detected

only in late prometaphase and metaphase, and is found in areas immediately adjacent to

centromeres (130). The relevance of these distinctive features associated with the different

histone phosphorylation events during mitosis remains to be determined.

Interestingly, a number of these phosphorylation sites (H3T6/S10/T11/S28) are also

associated with transcriptional activation in a wide range of organisms from yeast to humans.

The diverse signaling pathways and kinases that phosphorylate these residues and the

functional roles of histone phosphorylation in transcription will be discussed in more details

in Sections 1.4-1.6.

1.3.5 Histone ubiquitylation, sumoylation and other modifications

In addition to being modified by small chemical groups, histones are also subjected to

31

larger covalent modifications, which include ubiquitylation and sumoylation. It is noteworthy

that histone H2A was indeed the first protein shown to be ubiquitylated (135), even though

its functional significance was not discovered until recently. Histones can be

monoubiquitylated as well as polyubiquitylated, however, most studies on histone

ubiquitylation have focused on monoubiquitylation (especially on C-terminal tails of H2A

and H2B) and its role in regulating gene expression.

Ubiquitylation of H2A at K119 is mainly mediated by Ring1b, which is an E3 ligase

in polycomb repressive complex 1 (PRC1) (136). This links H2A ubiquitylation to polycomb

silencing pathway, and H2AK119ub1 is found to play an important role in HOX gene

silencing and X-chromosome inactivation (137, 138). In mouse ES cells, Ring1b-mediated

H2A ubiquitylation restrains poised RNA polymerase II at bivalent genes, thus repressing

expression of these developmental regulators (139). Monoubiquitylation is also reversible

and a number of deubiquitylating enzymes which remove ubiquitin from H2A have now

been identified, including USP16, USP21, USP22, 2A-DUB and USP10 (140). These

enzymes are linked to transcriptional activation, and some of these are found to function as

coactivators of androgen receptor (AR)-mediated transcription (141, 142). In contrast to H2A

ubiquitylation, ubiquitylation of H2B at the C-terminus (K120 in human and K123 in yeast)

is associated with transcriptional activation (143). This is the predominant ubiquitylation site

in yeast and is mediated by Rad6 (E2) and Bre1 (E3); in humans, the corresponding enzymes

are RNF20/40 and UbcH6 (144–147). This modification has been linked to both transcription

initiation and elongation, and is required for methylation of H3 K4 and K79 in both yeast and

mammals (148–151).

Besides having a role in regulating gene expression, histone ubiquitylation is also an

important component of the DNA damage response pathway (152). In response to DNA

32

damage, E3 ligase RNF8 ubiquitylates H2A and variant H2A.X at sites of double-strand

breaks (153, 154). RNF168 then promotes the formation of Lys63-linked poly-ubiquitin

chains at the RNF8-ubiquitylated histones (155). This series of ubiquitylation events is

essential for the accumulation of DNA repair proteins, such as 53BP1 and BRCA1, to DNA

lesions. H3 and H4 are also ubiquitylated by the CUL4-DDB-ROC1 complex after UV

irradiation (156).

Similar to ubiquitylation, the ubiquitin-like modifier SUMO is conjugated to lysine

residues via the action of E1, E2 and E3 enzymes. Histone sumoylation mainly occurs on

histone H4 in mammalian cells and is linked to transcriptional repression (157). Targeting of

SUMO E2 conjugating enzyme Ubc9 to reporter gene strongly represses gene expression,

possibly mediated through the recruitment of repressor proteins HDAC1 and HP1. In S.

cerevisiae, sumoylation has been detected on all 4 core histones, and is found to antagonize

certain activating marks, such as acetylation and H2B ubiquitylation, which might also occur

on the same lysine residue (158). Due to the lack of repressive lysine methylation (H3K9me,

H3K27me) in budding yeast, histone sumoylation is currently the only described repressive

histone mark in this model organism.

In addition to the above-described modifications, other less studied or newly

identified PTMs on histones include proline isomerization, propionylation, butyrylation,

formylation, crotonylation on lysines, ADP-ribosylation on glutamate/arginine residues and

O-GlcNAc (β-N-acetylglucosamine) glycosylation on serine/threonine residues (Figure 1.3)

(12, 159). The functional effects and regulation of these modifications are just beginning to

be uncovered. Further investigations will help decipher their roles in various chromatin-

mediated processes.

33

1.4 Histone H3 phosphorylation and transcriptional regulation

Transcription of eukaryotic genes requires the establishment of a chromatin

environment permissive to binding of activators and transcription factors, as well as to the

subsequent assembly of preinitiation complexes (160). These chromatin changes are brought

about by multiple histone-modifying enzymes and chromatin-remodeling complexes, and

they represent key regulatory steps in the transcription process. The roles of histone

acetylation and methylation in this process have been extensively studied (82, 83, 161). In

comparison, the mechanistic functions of histone phosphorylation in transcriptional

regulation are not as clearly defined. Unlike acetylation and methylation, genome-wide

studies on histone phosphorylation have not been performed, and therefore, it is unclear to

what extent this modification is involved in regulating gene expression. In spite of this,

phosphorylation of H3 at multiple and distinct sites (T6, S10, T11, S28) has been linked to

various signalling pathways and associated with the activation of multiple types of genes,

including immediate-early, NF-κB-responsive, MYC-regulated and hormone-regulated

genes.

1.4.1 H3 phosphorylation during mitosis and interphase

Phosphorylation of H3 on S10 and S28 is particularly interesting since these two

modifications occur both during mitosis and during transcriptional activation in interphase

cells. It is still unclear how H3 phosphorylation at these sites is associated with two different

processes that have opposite requirements for chromatin compaction – chromosome

condensation during mitosis and chromatin relaxation during gene expression. However, this

conundrum only further reinforces the idea that biological outcomes are dictated by the

overall combinations of modifications on histones rather than by individual modifications.

34

While some of the same sites on H3 are phosphorylated both in mitosis and during activation

of IE genes, these phosphorylation events also have quite distinct features (162–164). During

cell cycle progression, H3 phosphorylation begins in late G2 at the pericentromeric

chromatin, and then spreads along the chromosome arms as mitosis proceeds. H3 is highly

phosphorylated during mitosis, and it is commonly assumed that the majority of all H3 is

phosphorylated during chromosome condensation (a definitely quantification of the

percentage of H3 phosphorylated during mitosis, however, has not been done). In mitogen-

stimulated interphase cells, H3 phosphorylation is only observed in a very small fraction of

total H3 and correlates with transcriptional activation of a number of inducible genes (165).

In addition, the kinases responsible for these different H3 phosphorylation events are also

distinct. Mitotic H3 phosphorylation is mediated by Ipl1/Aurora B kinase (129, 134),

whereas kinases from MAPK (mitogen-activated protein kinase) pathways are largely

responsible for H3 phosphorylation in interphase cell.

1.4.2 The first link between H3 phosphorylation and transcriptional activation

The link between intracellular signaling pathways, gene activation and H3

phosphorylation first came from studies of immediate-early (IE) gene induction in mouse

fibroblasts (166). Extracellular stimuli, such as growth factors, stress or pharmacological

agents, trigger the rapid and transient expression of the immediate-early c-fos and c-jun genes

in mammalian cells. Their induction strongly correlates with a transient phosphorylation of

S10 and S28 on histone H3, as well as S6 on high-mobility group protein HMGN1, and these

phosphorylation events have been termed the “nucleosomal response” (167). The kinetics of

H3 phosphorylation closely mirrors the expression profiles of IE genes, suggesting that this

histone modification is part of the activation process of these genes. Activation of IE genes

35

by various stimuli can still occur without H3 phosphorylation, but the induction profile is

markedly altered and expression level is greatly attenuated. This suggests that, although not

absolutely required, H3 phosphorylation is necessary for efficient transcription (167, 168).

Importantly, transcriptional inhibitors actinomycin D and α-amanitin did not abolish stimuli-

induced H3 phosphorylation, indicating that this modification is not a consequence of gene

expression (166).

A direct link between H3 phosphorylation and IE gene induction is confirmed by

ChIP assays which show that phosphorylated H3S10 (H3S10ph) is physically associated with

the promoter and coding regions of IE genes (c-jun, c-fos, c-myc) during the activation

process (169–172). In contrast to H3S10ph, the link between H3S28 phosphorylation

(H3S28ph) and transcriptional activation is much less-studied, and the association of

H3S28ph with specific genes (such as IE genes c-jun, FOSL1 and Cox2) has not been

demonstrated until recently (173, 174). Immunofluorescence studies of mitogen-stimulated

cells show that H3S10ph and H3S28ph both display punctuate staining throughout the

nucleoplasm, and their localization are excluded from DAPI-dense regions (175, 176).

Interestingly, the majority of H3S10ph foci do not coincide with H3S28ph foci, suggesting

that these phosphorylation events occur on distinct populations of H3. Even though both

H3S10ph and H3S28ph have been localized to the same set of IE gene promoters, sequential

ChIP experiments suggest that these two modifications are not present within the same

nucleosome at the regulatory regions (174). At present, the significance of this observation in

terms of IE gene activation, and the functional differences between the H3 phosphorylated at

these distinct sites are unknown.

The link between H3 phosphorylation and transcriptional activation is not only

observed in mammals, but also in lower organisms as well. In budding yeast Saccharomyces

36

cerevisiae, phosphorylation of H3S10 by the Snf1 kinase is required for the activation of

INO1 and GAL1 genes in response to inositol starvation and galactose respectively (177,

178). In Drosophila, heat shock treatment causes a rapid induction of heat-shock genes and a

concomitant global shutdown of transcription of all other genes, and this is accompanied by a

dramatic change in the genomic distribution of H3S10ph. This process is best exemplified

by the induction of heat shock puffs on the polytene chromosomes isolated from Drosophila

larvae. In this system, heat shock induces a dramatic enrichment of H3S10ph at the

transcriptionally active heat shock puffs, and a concomitant global loss of H3S10ph at all

other gene loci (179). Genetics studies further show that H3S10ph level in Drosophila is

maintained by the opposing JIL-1 kinase and PP2A phosphatase (180, 181). Together, these

studies support the concept that the link between H3 phosphorylation and rapid gene

induction is evolutionarily conserved amongst different species.

1.4.3 MAPK pathway-mediated H3 phosphorylation & gene expression

It has long been known that mitogen and stress activate H3 phosphorylation

respectively through the MAPK and p38 pathways. RSK2, a downstream kinase of ERK1/2,

was the first kinase linked to H3S10 phosphorylation and IE gene expression (182). RSK2

knockout mouse cells and human fibroblasts derived from the RSK2-deficient Coffin-Lowry

syndrome (CLS) patients both exhibit drastic reduction in EGF-induced H3-S10

phosphorylation and in c-fos expression (182–184). Moreover, ectopic expression of RSK2

can restore mitogen-stimulated H3 phosphorylation in CLS cells, suggesting that this kinase

is required for the mitogen-induced nucleosomal response. Subsequent studies have found

that induction of H3 phosphorylation can be inhibited by H89, a chemical inhibitor that

targets another MAPK-activated protein kinase MSK1 but has no effect on RSK2, suggesting

37

the involvement of MSK1 and not RSK2 in this process (167). Indeed, MSK1 and MSK2,

two structurally and functionally similar kinases that lie downstream of ERK and p38

pathways, can phosphorylate H3 at S10 and S28 in response to mitogen and stress

stimulation. Embryonic fibroblasts from MSK1/2 double knockout mice show severe

reductions of H3 phosphorylation and IE gene induction, and reintroduction of the MSK2

gene alone is sufficient to rescue the H3 phosphorylation response of these cells (168). These

studies therefore implicate MSK1/2 as the key mediator of H3 phosphorylation in

mammalian cells.

RSK2 and MSK1/2 are indeed closely related protein kinases. They are members of

the MAPK-activated protein kinase family and are unique in that they possess two functional

kinase domains within the single polypeptide (reviewed in ref. 185). The N-terminal kinase

domain (NTKD) belongs to the AGC family of kinases and is responsible for

phosphorylating downstream substrates, whereas the C-terminal kinase domain (CTKD)

belongs to the calcium/calmodulin-dependent protein kinase family and is essential for

activating the NTKD via autophosphorylation. In response to cellular stimulation (such as

mitogen or stress), these MAPK-activated protein kinases are phosphorylated at the linker

region as well as the CTKD by upstream kinases. This activates the CTKD, which then

autophosphorylates the hydrophobic motif in the case of RSK2, and the hydrophobic motif as

well as the NTKD in the case of MSK1/2. These series of phosphorylation events eventually

lead to activation of the NTKD and phosphorylation of downstream substrates.

Constitutively-active (CA) mutants of RSK2 and MSK1 have previously been generated

(186). These are truncated mutants that contain only the NTKD, and their hydrophobic

motifs have been replaced with that of another AGC kinase, PRK2. Importantly, these CA

mutants can phosphorylate histone H3 in the absence of upstream signaling, suggesting that

38

H3 is a direct substrate of RSK2 and MSK1.

Although RSK2 and MSK1/2 are the first-identified and best-studied H3 kinases, it is

likely that, depending on the specific stimulus and cell type, multiple enzymes within the

MAPK pathways can target H3. For example, arsenite-induced H3 phosphorylation is

mediated by AKT, ERK2, and RSK2, but not by MSK1 (187). In addition, in UVB-irradiated

cells, COT and MLTKα (both members of the MAPKKK family) are responsible for H3S10

and S28 phosphorylation respectively (188, 189). H3S10 phosphorylation has also been

implicated in stem cell differentiation. Differentiation of ES cells by leukemia inhibitory

factor (LIF) removal induces H3S10ph/K14ac phosphoacetylation globally and at specific

differentiation markers in a MSK1-dependent manner (190). In a neuronal differentiation

model, MAP kinase JNK also phosphorylates H3S10 and this mark is enriched at JNK target

genes during differentiation (191). Further studies will be needed to clarify the specific roles

and contributions of each of these kinases in H3 phosphorylation and the nucleosomal

response.

1.4.4 Other signaling pathways that target H3

Besides the MAPK cascades, H3 phosphorylation is a common downstream event for

other signaling pathways as well. For example, IKKα, one of the subunits of the IκB kinase

complex in the NFκB pathway, directly phosphorylates H3 at the promoters of cytokine-

induced genes (192, 193). Knockout mouse studies showed that IKKα is required for optimal

TNF-induced expression of NFκB-regulated genes (IκB, IL-6, IL-8), as well as for H3

phosphorylation, and the recruitment of IKKα to these promoters in normal cells correlates

with the induction of both H3 phosphorylation and gene expression. In addition, stimulation

of ovarian granulosa cells with follicle-stimulating hormone (FSH) induces H3

39

phosphorylation in a PKA-dependent and MAPK-independent manner (194). This study

implicates PKA as the direct H3 kinase associated with the activation of FSH-responsive

genes during granulosa cell differentiation. H3 phosphorylation has also been implicated in

the regulation of MYC target genes and nuclear hormone signaling.

1.4.4.1 H3 phosphorylation and regulation of MYC target genes

H3 phosphorylation has also been linked to the regulation of MYC target genes

through PIM1, which phosphorylates H3 at S10 (195). After VEGF-A stimulation, PIM1 co-

localizes with c-MYC and H3S10ph at sites of active transcription. This is mediated through

its interaction with the MYC-MAX dimer whereby PIM1 is recruited to the E-boxes of

MYC-target genes FOSL1 (FRA-1) and ID2. MYC is recruited to two regions of the FOSL1

gene: FOSL1 upstream region, which contains a non-canonical E-box element, and FOSL1

downstream enhancer, which contains a canonical E-box. Phosphorylation of H3S10 is

detected at both sites, albeit with different kinetics. Phosphorylation at the upstream element

occurs at an earlier time point and is mediated by MSK1/2. However, PIM1 specifically

associates with the downstream enhancer and its recruitment correlates with H3S10ph at the

canonical E-box and during the peak of transcriptional activation. Inhibition of PIM1

strongly reduces H3 phosphorylation and mRNA expression of FOSL1 and ID2. The

significance of having two independent H3 phosphorylation events, mediated by different

H3S10 kinases, on the same gene is not clearly understood. It is of interest to note that the

kinase activity of PIM1 and its recruitment to chromatin by MYC both contribute to MYC-

dependent transformation. Insofar as microarray expression analysis indicates that PIM1 is

necessary in regulating expression of >200 (~20%) MYC targets, these findings together

suggest that H3 phosphorylation may have a role in oncogenesis through MYC-regulation.

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1.4.4.2 H3 phosphorylation and nuclear receptor signaling

Nuclear receptors are ligand-dependent transcription regulators, and upon ligand

binding, these receptors activate transcription through recruitment of coregulators, which

include histone modifying enzymes and chromatin remodeling complexes. Histone

acetyltransferases (CBP/p300) and histone arginine methyltransferases (CARM1/PRMT1)

are well-studied coactivators of nuclear receptors. Recent reports have just added histone

kinases to the list of coactivators that help modulate expression of nuclear receptor target

genes. A recent study showed that MSK1 and H3 S10 phosphorylation are part of the retinoic

acid (RA) signaling cascade (196). In mouse embryonic fibroblasts (MEFs), RA treatment

activates the p38 MAPK and its downstream kinase MSK1. Activated MSK1 is recruited to

RA target genes (CYP26A1 and RARβ2) and phosphorylates H3 S10 at these promoters.

Knockdown or chemical inhibition of MSK1 significantly reduces induction of RA target

genes, illustrating the importance of MSK1 activity in RA-induced gene expression.

Using mouse mammary tumor virus (MMTV) promoter as a model, it was showed

that induction of progesterone target genes also involves phosphorylation of H3 (197).

Hormone stimulation activates the SRC/p21ras/ERK pathway, and results in the

phosphorylation and activation of progesterone receptor (PR) and MSK1. Phosphorylated

PR, ERK and MSK1 form a ternary complex and is then recruited to the MMTV promoter.

These events result in H3S10 phosphorylation, H3K14 acetylation and displacement of HP1γ

from the promoter. In addition, chromatin-remodeling complex BRG1 and histone

acetyltransferase PCAF are also recruited to the promoter, eventually leading to the

recruitment of RNA Pol II and activation of hormone-regulated genes. Interestingly, only

H3S10ph, but not H3S28ph, is detected at the MMTV promoter. It is not clear how this site

specificity is brought about. In addition to H3S10ph, phosphorylation of H3 at T6 by PKCβ

41

and at T11 by PRK1 have also been linked to the activation of androgen-receptor-regulated

genes (198, 199).

1.5 Mechanism of action of histone modifications

Although the understanding of enzymes and signaling pathways that induce histone

modifications has increased drastically in recent years, the mechanisms by which these

modifications are translated into downstream events are still not clearly understood. Two

different, yet not mutually exclusive, mechanisms have been suggested to explain this

connection between histone PTMs and various biological outcomes: one involves direct

structural impact on nucleosomes and chromatin, and the other involves the recruitment of

downstream effectors to chromatin through histone PTMs and PTM-binding modules.

1.5.1 Structural effects of histone modifications

The first model proposes that histone modifications can mediate downstream

processes by directly inducing structural changes to chromatin. The addition of acetyl or

phosphate groups changes the net charge on histones, and these modifications have been

proposed to alter histone-DNA or histone-histone interactions. This could then affect the

formation of higher-order chromatin structures and the stability of chromatin compaction

states.

The first demonstration of a link between histone acetylation, alterations in

nucleosome conformation and IE gene activation was shown by Allfrey and colleagues in the

late 1980s (200, 201). Using mercury affinity chromatography, which exploits the

accessibility of cysteine 110 within the globular domain of H3 as a means to monitor changes

in nucleosome conformation, they found that nucleosomes associated with IE genes c-fos and

42

c-myc adopt a more open conformation and are retained on the organomercurial column upon

gene induction. Importantly, nucleosomes in the retained fraction are hyperacetylated,

indicating an association between histone acetylation and conformational change in

nucleosomes. Mutational studies in yeast, where acetylation sites on H4 tail (K5, 8, 12 and

16) were mutated individually or in combinations, also provide evidence for a charge-based

mechanism for certain acetylated lysines (202). Mutations on H4K5, K8 and K12 have

indistinguishable transcriptional outcomes, and they also show cumulative effects on gene

expression. These findings are consistent with charge-specific effects for acetylation on these

lysine residues. In contrast, mutation of H4K16 shows a distinct transcriptional profile

compared to the other lysines, suggesting that charge effects alone cannot fully explain the

effects of acetylation on chromatin, and other mechanisms may be involved in mediating the

H4K16-specific effects.

In order to study the impact of specific histone PTM on chromatin structure and

folding in vitro, a native chemical ligation strategy has been used to generate recombinant

histones modified at specific residues (203–205). This involves chemical ligation of modified

histone tail peptides onto recombinant histone C-terminal fragments, and allows the

reconstitution of nucleosomal arrays that harbor homogeneous and site-specific histone

modifications. Biophysical analyses of nucleosomal arrays assembled with K16-acetylated

H4 show that H4K16ac inhibits the formation of 30-nm-like fibers in vitro (205). However,

nucleosomal arrays containing unmodified or S10-phosphorylated H3 are similar in regards

to DNA accessibility and SWI/SNF-dependent chromatin remodeling (203). H3 S10

phosphorylation also has no significant effect on higher-order chromatin folding (204).

Similar experiments using histones phosphorylated at H3 S28 (or other phosphorylation

sites) have not been performed and these studies could provide further insight into the direct

43

effect of histone phosphorylation on chromatin compaction and structure. Another study

using hydroxyapatite dissociation chromatography to fractionate nucleosomes based on

structural stability found that H3S28ph-containing nucleosomes are more labile and

destabilized compared to the H3S10ph-containing nucleosomes (206). In fact, the H3S10ph-

and unphosphorylated H3-containing nucleosomes co-fractionate together, suggesting they

have similar structural stability. It is interesting to note that phosphorylation at the different

sites have different effects on nucleosome stability and it remains to be determined whether

these physical differences are translated into functional differences as well.

1.5.2 Recruitment of effector proteins to chromatin

Another mechanism by which histones PTMs mediate downstream functions is

through their recruitment of effector proteins (also known as “readers” of histone marks) to

chromatin (207, 208). Modified residues on histones serve as docking sites for conserved

protein modules that recognize histone PTMs in modification- and site-specific manners

(Figure 1.6). Indeed, the paradigm of modification-dependent protein-protein interactions is

well established and best exemplified by the binding of SH2 domains to phospho-tyrosines

and FHA domains to phospho-serines/threonines on signaling molecules (209). These PTM-

recognition domains are present in many histone-modifying enzymes, ATP-dependent

chromatin-remodeling enzymes, as well as other cellular machineries involved in

transcription, DNA repair and RNA processing. Histone PTMs therefore often play a crucial

role in recruiting these enzymatic activities to the appropriate chromatin locations.

1.5.2.1 Acetyl- and methyl-binding domains

Bromodomain from the transcriptional coactivator PCAF was the first histone-

44

Figure 1.6. Binding domains for modified histones. (A). Domains that recognize acetylated lysines. (B). Domains that recognize methylated lysines. (C). Phospho-histone binding proteins. (i) BRCT domain of MDC1 binds phosphorylated H2A.X in response to DNA damage. (ii) BIR domain of survivin binds H3T3ph, and recruits chromosomal passenger complex to centromeres during mitosis. (iii) 14-3-3 binds to both H3S10ph and H3S28ph. Importantly, H3 S10 phosphorylation is physically and functionally coupled to acetylation of the nearly K14 residue, and acetylation of H3 K14 enhances the interaction between H3S10ph and 14-3-3.

45

binding module to be structurally characterized (210). This domain binds to acetylated lysine

residues on histones and is often found in numerous HATs and chromatin remodeling

complexes, thus recruiting these enzymatic activities to further “open” the chromatin for

transcription, DNA repair, etc (211). For many years, recognition of acetylated lysines has

been limited to bromodomains, until recently when plant homeobox (PHD) finger is found to

bind this PTM as well (Figure 1.6A). PHD fingers are mostly associated with methyl-lysine

binding (212), but the tandem PHD fingers in DPF3b (a component of BAF chromatin

remodeling complex) bind acetylated histone H3 and H4, thus making it the first non-

bromodomain acetyl-reader (213, 214).

In comparison, there are far more distinct protein domains recognizing methylated

lysines than for any other modification. These methyl-lysine binding modules include

members of the Tudor domain Royal family (chromo, Tudor, MBT, PWWP), WD40 and

PHD fingers (Figure 1.6B) (207, 215). Similar to the high specificity of histone

methyltransferases and demethylases towards their target lysine, methyl-binding domains are

also highly discriminatory for binding to specific methylated site and degree of methylation.

For example, chromodomain of HP1 binds strongly to H3K9me3 and is important for

establishing constitutive heterochromatin (86, 216). On the other hand, methylation on

H3K27 creates a motif that is specifically recognized by the chromodomain of Pc

(Polycomb), leading to recruitment of PRC1 complex and polycomb-mediated silencing.

Importantly, both chromodomains can distinguish between different methylation states and

have a stronger preference for trimethylated lysines (217, 218). In addition, one histone PTM

can often serve as a binding platform for a variety of chromatin-modifying enzymes, even

with opposing activities. H3K4 methylation, for instance, can recruit a HAT, HDAC or

chromatin remodeling complex to chromatin, depending on the specific H3K4me3 reader. As

46

a component of acetyltransferase NuA3, PHD finger of Yng1 binds H3K4me3 to bring about

H3K14 acetylation (219). ING2 also binds H3K4me3, but in this case, recruits mSin3a-

HDAC complex to repress proliferation-specific genes in response to genotoxic stress (220).

Chromatin remodeler CHD1 can be recruited to active chromatin through H3K4 methylation

as well (221). The functional outcomes of particular PTMs are therefore dictated by functions

of the recruited effectors.

1.5.2.2 Phospho-histone binding proteins

In contrast to the large number of acetyl- and methyl-histone readers, only a limited

number of proteins are found to specifically bind phosphorylated histones (Figure 1.6C).

BRCT domain of MDC1 directly binds to H2A.X-S139 in a phospho-dependent manner

upon DNA damage, and this interaction is critical for localization of MDC1 to sites of DNA

breaks and for the normal radio-tolerance of cells (222). During mitosis, chromosomal

passenger complex, which contains the mitotic H3S10/S28 kinase Aurora B, is targeted to

centromeres through the direct interaction between BIR domain of survivin and H3T3ph.

Such recruitment results in activation of Aurora B, and proper localization of this complex is

required for spindle-kinetochore attachments and subsequent chromosome segregation (223–

225). To date, members of the 14-3-3 family (14-3-3ε, 14-3-3ζ) are the only phospho-histone

binding proteins linked to transcriptional regulation, and they are capable of binding to

H3S10ph as well as H3S28ph.

An initial study reported that the interaction between 14-3-3 and histone H3 is

phospho-dependent, but not affected by the acetylation status of nearly lysine residues (K9,

K14) (226). However, two other studies demonstrated that this binding is modulated by other

modifications on H3 whereby phosphorylated S10 is necessary for this interaction, but

47

acetylated K9/K14 greatly enhances the binding affinity (227, 228). Whether this holds true

for 14-3-3/H3S28ph binding has not been tested. In vivo studies using ChIP assays further

validate the findings from these in vitro pull-down assays. 14-3-3 proteins are recruited to IE

genes (such as c-jun, FOSL1) and HDAC1 promoters upon gene activation, concomitant with

phosphoacetylation of H3 (174, 227, 229). Furthermore, RNAi knockdown of 14-3-3ε/ζ

strongly interferes with transcription of HDAC1, but has no effect on H3 phosphoacetylation

levels, suggesting that 14-3-3 is a crucial downstream effector of H3 phosphoacetylation

(227). This interaction and function of 14-3-3 is also conserved in yeast. Bmh1 and Bmh2,

the yeast homologues of 14-3-3, are essential for optimal transcriptional activation of the

GAL1 gene (228). Binding of Bmh1 to GAL1 promoter is dependent on H3S10 and K14

since H3 S10A, S10A/K14R and K14R yeast mutants are all defective in Bmh1 recruitment.

These findings provide further in vivo support that H3K14ac is important for 14-3-3 binding.

Similar to its role in signal transduction, 14-3-3 mainly acts as an adaptor or

scaffolding protein on chromatin, and functions to recruit other chromatin modifying

enzymes to phosphorylated histones during the transcription process. For example, in

response to Ras-MAPK signaling, 14-3-3 associates with BRG1 (the ATPase subunit of

SWI/SNF remodeling complex) and PCAF (an H3/H4 histone acetyltransferase), recruiting

these enzymes to IE gene promoters (174). Another study found that 14-3-3 can also interact

with the H4K16 acetyltransferase MOF, and their recruitment to the enhancer region of

FOSL1 is important for productive elongation of this gene (229). A recent study at the p21

promoter suggests that another function of 14-3-3 is to protect H3S10ph against

dephosphorylation by promoter-bound H3 phosphatase PP2A (230).

In addition to the recruitment of phospho-binding proteins to chromatin, H3

phosphorylation can also function to repel other histone-binding factors. HP1 binds

48

specifically to H3K9me through its chromodomain, and this is a key mechanism that

facilitates heterochromatin assembly and gene silencing (86, 216). Phosphorylation of

H3S10 by Aurora B disrupts the HP1/H3K9me interaction, and causes the release of HP1

from chromosomes during mitosis (231, 232). This “phos/methyl switch” was first observed

in mitosis, but this mechanism also operates in the context of transcriptional activation in

interphase cells. HP1γ displacement from promoter is concomitant with H3S10ph during

progesterone receptor-mediated gene activation and HDAC1 induction (197, 227). H3

phosphorylation, therefore, can simultaneously recruit 14-3-3 and eject the transcriptional

repressor HP1γ during transcription initiation. These findings further illustrate the

complexity of how histone modifications regulate chromatin binding of various proteins, as

well as the importance of the combinatorial histone modifications in the regulation of cellular

processes.

1.6 Histone modification crosstalks

The abundance of PTMs on histones creates an environment in which crosstalks can

occur between different modifications on one or more histone tails. It is now well-

established that histone modifications can influence each other and a pre-existing

modification on one residue can have a synergistic or antagonistic effect on a subsequent

modification on another residue.

1.6.1 Modes and mechanisms of histone modification crosstalks

The simplest antagonism involves the choice of different modifications that target the

same site, such as methylation and acetylation on H3K9 and H3K27. As shown in Figure 1.3,

a number of different PTMs can often occur on the same amino acid, and these PTMs clearly

49

cannot occur on the same residue at the same time. This is particularly true for lysine, where

at least 8 different types of PTMs can occur. One of the earliest examples of histone PTM

crosstalk involves H3S10 phosphorylation and the subsequent acetylation of the

neighbouring K14 (see the following section on H3 phosphoacetylation for details) (170,

233). A number of crosstalks between modifications on the same histone (crosstalk in cis)

have been reported, especially on the histone H3 tail. For example, acetylation of

H3K18/K23 by CBP promotes methylation of H3R17 by CARM1, and together, they are

involved in activation of estrogen-responsive genes (234). Besides affecting modifications on

the same tail, modifications on different histones can also influence each other. One classic

example of trans-tail crosstalk involves ubiquitination on H2B and methylation on H3.

During transcription initiation, mono-ubiquitination of H2B at the C-terminus is absolutely

required for H3K4 methylation by Set1 and H3K79 methylation by Dot1 (148–151). This

process was originally discovered in yeast, but is now shown to be conserved in higher

eukaryotes as well. Such a complex interplay between PTMs can dictate the combinatorial

modifications that occur together on histones, and therefore adds another level of complexity

to the regulation of DNA-templated processes by chromatin (71, 235).

Crosstalks can be mediated through various mechanisms. First of all, PTMs can have

a direct influence on enzymatic activity, where a pre-existing modification can enhance or

reduce the activity of another histone-modifying enzyme towards its target residue. For

example, isomerization of H3P38 leads to a conformation change in the H3 tail, and

therefore, prevents methylation of H3K36 by Set2 (236). At AR-regulated genes, two

phospho-methyl crosstalks are involved in the activation process. H3T6 phosphorylation by

PKCβI blocks H3K4 demethylation by LSD1 and JARID1B, and phosphorylation of H3T11

by PRK1 enhances demethylation of H3K9 by JMJD2C (198, 199). Together, these two

50

phosphorylation events cooperate to regulate methylation status on the H3 tail and are

important in regulating expression of AR-target genes. Crosstalks can also be mediated

through PTM-dependent enzyme recruitment. For instance, H4 acetyltransferase MOF is

recruited via 14-3-3 to H3S10 phosphorylated regions, thus generating a crosstalk between

H3S10ph and H4K16ac during IE gene activation (229).

1.6.2 Histone H3 phosphoacetylation

Insofar as H3 phosphorylation at S10 and acetylation at K9 or K14 have all been

linked to transcriptional activation, and given the proximity of these modified residues on

H3, several labs have questioned whether specific combinations of modifications occur

together during gene activation. Antibodies generated to specifically recognize di-modified

H3 (anti-H3K9ac/S10ph, anti-H3S10ph/K14ac) show that these specific H3 modification

combinations do exist on the same H3 tail in vivo (170, 171). The global levels of the

phosphoacetylated H3 are increased upon mitogen stimulation of mammalian cells, and ChIP

assays show that di-modified H3 are associated with the IE gene promoters during

transcriptional activation.

Additional studies suggest that these modifications are mechanistically linked. For

example, in vitro enzymatic assays show that several histone acetyltransferases (HAT),

including yeast Gcn5, human PCAF and p300, preferentially acetylate H3S10ph peptide

substrates over the unmodified form (170, 233). Furthermore, structural analyses of Gcn5 in

complex with the H3S10ph peptide suggest that this enzymatic preference is due to the

phosphate-dependent stabilization of the enzyme-substrate complex (237). Using

recombinant nucleosomal arrays, another study confirmed that the HAT activity of

recombinant Gcn5p is enhanced by H3 phosphorylation; however, the nucleosomal HAT

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activity of the purified yeast Gcn5-containing SAGA complex is insensitive to H3

phosphorylation. The in vivo relevance of this observation is currently not known (204). A

recent study on p21 expression found that H3S10ph/K14ac is also involved in the activation

of this cell cycle regulator by anisomycin and HDAC inhibitor. Blocking of the nucleosomal

response by MSK1 inhibitor H89 significantly reduced H3K14 acetylation at the p21

promoter, suggesting that the recruitment or activation of the H3K14 acetyltransferase is

largely dependent on H3S10 phosphorylation, and supports a direct link between H3S10

phosphorylation and H3K14 acetylation (230). The observation that binding of 14-3-3 to

H3S10ph is enhanced by additional acetylation of H3K9 or K14ac further supports the

importance of this phosphoacetylated mark in transcriptional activation (227, 228).

Such coupling of phosphorylation and acetylation, however, is not absolutely required

for gene activation. For some other genes, such as Drosophila heat shock genes (179) and

mouse HSP70 (238), transcriptional activation is only accompanied by increased H3S10

phosphorylation without changes in the H3 acetylation levels. Studies using MSK1/2

knockout cells or MSK1 inhibitor H89 also show a correlation, but not sequential

dependence, between H3-S10 phosphorylation and K9 acetylation at the c-jun promoter (168,

172). These suggest that phosphorylation and acetylation can be independently targeted to

the same region of the genome and they coincide to produce phosphoacetylated H3 tails.

In yeast, depending on the gene studied, both the synergistic coupling and

independent targeting model of phosphoacetylation have been observed. Functional coupling

of H3 phosphorylation and acetylation is required for efficient activation of the INO1

promoter upon inositol starvation (177). Deletion of the H3 kinase SNF1 gene, or replacing

the endogenous H3 gene with an H3 S10A mutant both eliminated S10 phosphorylation in

the mutant yeast strains, and also significantly lowered K14 acetylation levels at the INO1

52

promoter, indicating that H3S10 phosphorylation is a pre-requisite for acetylation of K14

during gene activation. At the GAL1 gene, on the other hand, the Gal4 transcription factor

recruits both the H3 kinase and the SAGA acetyltransferase complex to the promoter, and

therefore, acetylation is independent of prior H3 phosphorylation (178). Taken together,

these studies suggest that the mechanism regulating H3 phosphoacetylation is different for

individual target genes, and likely depends on factors such as promoter structure and other

PTMs on the gene.

In addition to the coupling of phosphorylation and acetylation on H3, a trans-tail

crosstalk between H3 phosphorylation and H4 acetylation has also been reported (229). At

the FOSL1 gene, phosphorylation of H3S10 by PIM1 kinase recruits 14-3-3 and the H4

acetyltransferase MOF to the downstream enhancer region. Subsequent acetylation of H4K16

by MOF creates a binding site for the bromodomain-containing protein BRD4 and its

associated transcription elongation factor P-TEFb. Therefore, a trans-tail crosstalk between

H3S10ph and H4K16ac, through a cascade of events, eventually leads to transcription

elongation. A similar link between H3 phosphorylation, acetylation and transcription

elongation is also observed in Drosophila, but in this organism, 14-3-3 recruits the H3

acetyltransferase Elp3, and is associated with an increase in H3K9 acetylation (239).

1.6.3 Combinatorial histone PTMs and multivalent binding

It is now increasingly evident that histone PTMs do not function in isolation, instead,

combinations of histone PTMs are important for coordinately mediating downstream

processes (240). Such combinatorial patterns of PTMs are interpreted at the molecular level

through their recognition by multiple PTM-reading modules within effector proteins. These

combinatorial marks can be deposited through histone crosstalk pathways, as described in

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Section 1.6.1. Alternatively, an increasing number of histone modifying complexes are found

to contain more than one enzymatic activity, and thus can act in concert to coordinate

multiple modifications on nucleosomes. For example, histone acetyltransferase MOF

associates with methyltransferase MLL1 and are responsible for the H4K16ac and H3K4me3

marks respectively. Together, they are important for optimal transcriptional activation of

MLL target, HoxA9 (241).

Structural and biochemical studies have now begun to reveal how different chromatin

binding modules simultaneously engage these modification patterns to transduce downstream

functions. Many chromatin-associated complexes often contain multiple PTM-recognition

domains. These domains can be present within one protein, as illustrated by the binding of

NURF chromatin remodeling complex subunit BPTF to H3K4me3/H4K16ac nucleosomes

(Figure 1.7A) (242). This is mediated through the PHD finger-bromodomain motif of BPTF,

and similar paired modules are now commonly found in chromatin-associated proteins (243).

Multiple binding domains can also be present in multiple components within a complex. This

is best illustrated by the general transcription factor complex TFIID (Figure 1.7B). Multiple

points of contact are involved in the recruitment of TFIID to active promoters. Tandem

bromo domains within TAF1 cooperatively bind di-acetylated H4 (H4K5ac/K12ac) (244).

Another subunit TAF3 binds to H3K4me3 through its PHD finger, and this binding is further

enhanced by increased acetylation levels on H3 (H3K9ac/K14ac) (245). In sum, multivalent

binding of histone PTMs has emerged as a prevalent theme in the chromatin field. This can

collectively provide stronger and more specific binding sites for the reader proteins, and is

proposed to be important for deciphering the complex histone PTM language within a

cellular environment.

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Figure 1.7. Combinatorial reading of histone modifications. Many chromatin-associated complexes contain multiple PTM-recognition domains. (A). These domains can be present within one protein, as in BPTF. PHD-Bromo cassette of BPTF, a component of the NURF chromatin remodeling complex, binds to nucleosomes containing H3K4me3 and H4K16ac. (B). These domains can also be present in multiple components of a complex, as in TFIID. TAF1 and TAF3 are both components of the general transcriptional factor complex TFIID. PHD finger of TAF3 binds to H3K4me3, whereas the double bromodomain of TAF1 recognizes di-acetylated H4 (H4K5/K12ac).

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1.7 Histone phosphorylation and human diseases

Given that chromatin plays such an important role in regulating multiple DNA-

templated processes, it is not surprising that perturbation of chromatin modifiers and histone

modifying enzymes, including histone kinases, can lead to cancer and other diseases (246).

Mutations in signaling kinases are one of the most frequent oncogenic events found in

cancer, and some of these kinases can signal directly to chromatin as well. Mitogen-induced

H3 phosphorylation is mainly mediated via the Ras-MAPK (Raf-MEK-ERK) pathway, and

this pathway is often dysregulated and constitutively activated in cancer cells. In Ras-

transformed mouse fibroblasts, levels of both H3S10ph and H3S28ph are elevated, owing to

increased MSK1 activity in these cells (169, 173, 175, 247). Recently, constitutive activation

of MSK1 and increased H3 phosphorylation have been observed in primary acute myeloid

leukemia (AML) samples with FLT3 mutations (248). These findings suggest that MSK1 is

involved in FLT3-mediated leukemogenesis.

In addition to MSK1, many H3 kinases have also been linked to neoplastic cell

transformation and cancer development. H3S10 kinase PIM1 is a proto-oncogene implicated

in the pathogenesis of hematologic malignancies as well as solid tumours, and is known to

cooperate with MYC in tumorigenesis (249). Indeed, PIM1 is overexpressed in MYC-driven

prostate tumors (250), and the oncogenic effects of this enzyme may be mediated through its

H3 kinase activity (195). PRK1-mediated H3T11 phosphorylation is involved in regulating

AR-dependent genes, and inhibition of PRK1 reduces androgen-induced proliferation of a

prostate cancer cell line. High levels of PRK1 and H3T11ph are also detected in prostate

cancer cells and thus, PRK1 may be used as a predictive tumor marker (198). The mitotic

H3 kinase Aurora B is also overexpressed in many cancer cell lines. This leads to increased

mitotic S10 phosphorylation and triggers chromosome instability and multi-nuclearity –

56

features that are often seen in malignant cells (251). Importantly, Aurora kinase inhibitors

have shown potent anticancer activities in preclinical studies, suggesting that targeting

histone kinases may have therapeutic effects. Finally, phospho-H3 binding protein 14-3-3ζ

and other 14-3-3 family members are often abnormally expressed in various malignancies

(252). Whether this upregulation of 14-3-3 has an effect on phospho-H3-regulated genes

during tumorigenesis is unclear. All together, these findings underscore the importance of

histone phosphorylation in normal cell functions and growth.

1.8 Thesis Rationale and Objectives

Although intense research has uncovered the signaling cascades leading to histone H3

phosphorylation, mechanistic details linking this histone modification to gene expression

remain unclear and require further studies. The main objective of the studies presented in this

thesis is to unravel the functional consequences of histone H3 phosphorylation and its

mechanisms in regulating gene expression. So far, research efforts have been focusing on H3

S10, and little is known about the function and regulation of H3 S28 phosphorylation. To

address this gap in the field, I aim to further investigate the role of H3 S28 phosphorylation

in transcriptional activation. In particular, I determined that H3 S28 phosphorylation can

antagonize polycomb silencing and mediate a methyl-acetyl switch on H3 K27 (Chapter 2).

Such study would help to further decipher the functional differences between H3 S10 and

S28 phosphorylation.

It is now a well-accepted paradigm that histone PTMs can synergistically or

antagonistically affect other PTMs, leading to crosstalks amongst modifications. Given the

involvement of H3 S10 phosphorylation in multiple histone crosstalk pathways (e.g. coupling

of phosphorylation and acetylation during IE gene induction, and H3S10ph-induced

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displacement of H3K9me3-binding protein HP1), we hypothesized that one mechanism by

which H3 S10 and S28 phosphorylation mediate downstream processes is through the

establishment of crosstalks with other histone PTMs. To that end, we aim to develop a

method to systemically analyze combinatorial histone PTMs within the nucleosome context.

This approach was utilized to identify histone crosstalks involving H3 phosphorylation

(Chapter 3). Specifically, using this method, I provided evidence for a new trans-tail

crosstalk between H3 phosphorylation and H4 K12 acetylation. Together, these studies will

provide insight into the mechanistic link between H3 S10/S28 phosphorylation and

transcriptional activation.

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Chapter 2

Histone crosstalk involving H3 S28 phosphorylation and K27 acetylation

activates transcription and antagonizes polycomb silencing

A version of this chapter is published as: Lau, P.N.I., and Cheung, P. (2011). Histone code pathway involving H3 S28 phosphorylation and K27 acetylation activates transcription and antagonizes polycomb silencing. Proc Natl Acad Sci U S A 108, 2801-6. Dr. Peter Cheung performed the peptide competition assays in Figure 2.9. I performed all other experiments.

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2 PNAS 2.1 Abstract

Histone H3 phosphorylation is a critical step that couples signal transduction

pathways to gene regulation. To specifically assess the transcriptional regulatory functions of

H3 phosphorylation, we developed an in vivo targeting approach and found that the H3

kinase MSK1 is a direct and potent transcriptional activator. Targeting of this H3 kinase to

the endogenous c-fos promoter is sufficient to activate its expression without the need of

upstream signaling. Moreover, targeting MSK1 to the α-globin promoter induces H3 S28

phosphorylation and reactivates expression of this polycomb-silenced gene. Importantly, we

discovered a novel mechanism whereby H3 S28 phosphorylation not only displaces binding

of the polycomb repressive complexes, but it also induces a methyl-acetylation switch of the

adjacent K27 residue. Our findings show that signal transduction activation can directly

regulate polycomb silencing through a specific histone code-mediated mechanism.

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2.2 Introduction

H3 phosphorylation is a downstream event for a number of signal transduction

pathways in mammalian cells (reviewed in refs. 112, 114, 164). For example, rapid and

transient phosphorylation of H3 at S10 (H3S10ph) is detected at the promoters and coding

regions of immediate early (IE) genes upon stimulation of the MAPK or p38 pathways,

suggesting that this signaling-induced histone modification functions to activate transcription

of IE genes. Thus far, multiple kinases, including RSK2, MSK1/2, PIM1 and IKKα have

been shown to directly phosphorylate H3. As a common target of diverse signaling cascades,

H3 phosphorylation is thought to be a critical step translating signal transduction information

to the chromatin/transcriptional regulatory machinery (253).

Currently, how H3 phosphorylation is mechanistically linked to the transcriptional

process is still not fully understood. H3 S10 phosphorylation can be physically coupled to

acetylation of nearby lysine residues (K9 or K14), suggesting that combinations of these

modifications function together to activate transcription (170, 171, 233). Specific isoforms of

14-3-3 directly bind H3S10ph, and this interaction is greatly enhanced by additional

acetylation of the nearby K14 residue, further illustrating the biological relevance of specific

combinations of histone modifications (227, 228). Finally, recruitment of 14-3-3 through

H3S10ph at the promoters of HDAC1 and several IE genes is thought to facilitate

transcription induction of these genes (174, 227, 229). Binding of 14-3-3 to H3S10ph at the

FOSL1 enhancer, which is located downstream of the transcription start site, initiates

sequential recruitment of the MOF histone acetyltransferase, the bromodomain protein BRD4

and the transcription elongation factor P-TEFb (229). At least for this particular gene, H3

S10 phosphorylation leads to the release of the pre-initiated but paused RNA polymerase II,

and facilitates transcriptional elongation. In addition to S10, H3 is also phosphorylated at

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S28, but the link between this second phosphorylation site and transcriptional regulation is

less well studied. 14-3-3 also binds strongly to H3S28ph peptides in pull-down assays, and

nucleosomes at the promoters of several IE genes (Jun, FOSL1) are also phosphorylated at

H3 S28 upon TPA induction (174). Interestingly, immunofluorescence and sequential

immunoprecipitation experiments using phospho S10- and S28-specific antibodies suggest

that these modifications occur on distinct populations of H3 (175, 176). At present, the

functional differences between S10- and S28-phosphorylated H3 have not been explored.

The induction of multiple transcriptional activation programs by signal transduction

cascades poses a significant complication in assessing the direct transcriptional function of

H3 phosphorylation in vivo. For example, fibroblasts from MSK1/2 double knockout mice no

longer phosphorylate H3 in response to stimulation, but their IE genes are still activated,

albeit with delayed kinetics (168). To circumvent this complication, we developed a method

to directly target H3 kinases to reporter and endogenous genes, and determine the specific

functions of H3 phosphorylation in the transcription process. Here, we show that MSK1 is a

direct and potent transcription activator. Interestingly, our data suggest that H3 S10

phosphorylation alone is not sufficient to initiate transcription whereas induction of H3 S28

phosphorylation correlated better with this process. In support of this observation, we found

that targeting MSK1 to the polycomb-silenced α-globin gene reversed its silencing and

reactivated its expression. Mechanistically, phosphorylation of H3 S28 displaces binding of

polycomb group proteins and reduces the amount of H3K27me3 at the α-globin promoter.

More importantly, targeting MSK1 also induces H3K27 acetylation, leading to the

enrichment of K27ac/S28ph di-modified H3 at the α-globin promoter. This finding is the

first evidence that H3 K27 acetylation and S28 phosphorylation are directly coupled and

suggests that such combination can functionally antagonize polycomb silencing.

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2.3 Materials and Methods

Plasmid constructs

HA-tagged CA/KD-RSK2 and MSK1 in pMT2 were provided by Dr. M. Frodin (186).

Kinase inserts were PCR-amplified and cloned into pM vector (Clontech) in frame with the

Gal4-DNA binding domain (residues 1-147) to generate pM-Gal4DBD-CA/KD-

RSK2/MSK1 kinase fusion constructs. DNA binding domain of NF1 (residues 1-257) was

PCR-amplified from the NFIX expression vector (provided by Dr. R. Gronostajski) and fused

in frame to the N- or C-terminal side of CA- or KD-MSK1 to generate the NF1-kinase fusion

constructs in pcDNA3.1+ (Invitrogen). The luciferase reporter plasmid pGL2-5xGal4-luc

contains five Gal4 binding sites ~110bp upstream of the SV40 promoter and luciferase gene.

A fragment comprising the 5xGal4 binding sites, SV40 promoter and luciferase gene was

subcloned from the pGL2-5xGal4-luc plasmid into the episomal vector pRep9 (Invitrogen) to

generate the episomal reporter plasmid. pGL2-Promoter (Promega) contains a SV40

promoter with no Gal4 binding sites in the upstream region. For reporter plasmids used in

Figure 2.2, 5xGal4 binding sites were PCR-amplified from pGL2-5xGal4-luc and subcloned

into pGL2-Promoter at ~550 and 1100bp upstream of the SV40 promoter.

Cell culture, transfections and whole cell lysis

293T cells were grown in Dulbecco's modified Eagle's medium (DMEM, Sigma)

supplemented with 10% fetal bovine serum (FBS). Stable cell line containing an integrated

Gal4 luciferase reporter was previously described (254). All transfections were performed

using Lipofectamine 2000, and whole cell lysates were prepared by directly lysing pelleted

cells in 2x SDS boiling sample buffer. Histones were resolved on 15% SDS-PAGE gels and

analyzed by Western blotting using antibodies listed in Table 2.1.

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Table 2.1. Antibodies used for Western blot analyses and ChIP.

Antibodies Source Species Catalog #

H3 Abcam rabbit ab1791 H3S10ph Santa Cruz rabbit sc-8656R H3S28ph (HTA28) Sigma-Aldrich rat H9908 H3K27me3 Active Motif

Millipore rabbit rabbit

39156 07-449

H3K27ac Abcam rabbit ab4729 H3K27ac/S28ph Millipore

Serum against H3K27ac/S28ph peptide

rabbit rabbit

05-896

H4ac Millipore Rabbit 06-946 HA.11 (16B12) Covance Mouse MMS-101R Flag (M2) Sigma-Aldrich Mouse F1804 Bmi1 (F6) Millipore mouse 05-637 Ezh2 (AC22) Millipore mouse 17-662 RNA Pol II (phospho-S5) Abcam rabbit ab5131 Phos-Msk1 (S212) R&D Systems rabbit AF1036 ERK1/2 Millipore rabbit 06-182 Phos-ERK1/2 (T202/Y204) Cell Signaling mouse 9106 p38 Cell Signaling rabbit 9212 Phos-p38 (T180/Y182) Cell Signaling rabbit 4631 Rabbit IgG Millipore rabbit 12-370 Mouse IgG Millipore mouse 12-371

Stimulation of 10T1/2 cells

Mouse C3H 10T1/2 fibroblasts were grown in DMEM supplemented with 10% FBS, and

switched to serum-free media for 20h prior to stimulation. Serum-starved cells were

stimulated with 400ng/ml 12-O-tetradecanoyl-phorbol-13-acetate (TPA, Sigma) or 50ng/ml

anisomycin (Sigma) for 0-60 min and then harvested for western blot analyses. To inhibit

MSK1 activity, 10µM H89 (Sigma) was added to cells for 30 min before stimulation with

TPA or anisomycin.

Luciferase reporter assays

293T cells transfected with Gal4-fusion constructs and luciferase reporters were harvested

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and lysed in cell culture lysis reagent (Promega) 48 h after transfection. Luciferase activities

were measured using the Luciferase assay system (Promega) as per the manufacturer’s

instructions. Luciferase levels were normalized to β-gal levels from a co-transfected β-gal

plasmid, and data are represented as means ± standard deviation from 3 independent

experiments.

RT-PCR

Total RNA was extracted using Trizol Reagent (Invitrogen) and reverse transcription was

carried out using oligo (dT) primers and SuperScript II (Invitrogen) according to

manufacturer’s instructions. Semi-quantitative PCR was carried out using Taq polymerase

(NEB) and PCR products were resolved on 2% agarose gel. β-actin was used as loading

control. Quantitative real-time PCR was performed using PerfeCTa SYBR Green SuperMix

(Quanta) on an Opticon 2 thermal cycler (BioRad). Relative mRNA levels were calculated

using GeneX program (BioRad), with HPRT as reference gene. Primer sequences are listed

in Table 2.2.

ChIP and sequential ChIP assays

ChIP assays were performed as previously described (170), with minor modifications.

Briefly, 293T cells were fixed in 1% formaldehyde for 8 min at room temperature. After

swelling and lysis, chromatin was sonicated to an average of ~500bp with a Branson Sonifier

450. For ChIPs with anti-H3K27me3 and anti-H3K27ac antibodies, lysates were treated with

lambda phosphatase (NEB) for 30 min at 30oC before sonication. Antibodies were prebound

to magnetic Dynabeads (Invitrogen, protein G for mouse and rat monoclonal antibodies,

protein A/G beads for rabbit polyclonal antibodies) for at least 3h and mouse/rabbit IgG were

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Table 2.2. Primer sequences used for RT-PCR and ChIP-qPCR analyses. Target Forward Primer Reverse Primer Semi-quantitative RT-PCR α-globin CAAGACCTACTTCCCGCACTTCGAC TCCAGCTTAACGGTATTTGGAGGTCAG β-actin TGAAGTCTGACGTGGACATC ACTCGTCATACTCCTGCTTG Quantitative RT-PCR c-fos CGGGCTTCAACGCAGACTA GGTCCGTGCAGAAGTCCTG c-jun TGCCTCCAAGTGCCGAAAAA GACTTTCTGTTTAAGCTGTGCC Hsp70 ACCTTCGACGTGTCCATCCTGA TCCTCCACGAAGTGGTTCACCA α-globin CTGGGCCTCCCAACGG GCCCACTCAGACTTTATTCAAAGAC HPRT TGGAGTCCTATTGACATCGCCAGT AACAACAATCCGCCCAAAGGGAAC ChIP-quantitative PCR c-fos GAGCAGTTCCCGTCAATCC GCATTTCGCAGTTCCTGTCT c-fos (re-ChIP) AGGAACTGCGAAATGCTCAC GTAAACGTCACGGGCTCAAC α -globin GGGCCGGCACTCTTCTG GGCCTTGACGTTGGTCTTGT HoxD4 TGGTCTACCCCTGGATGAAG TGACCTGCTCCCTCAGCTAT Myt1 ACAAAGGCAGATACCCAACG GCAGTTTCAAAAAGCCATCC

used as negative controls. For anti-HA ChIP, anti-HA affinity matrix (3F10, Roche) was

used. Sonicated lysates were precleared with Dynabeads for 2 h and then incubated with

antibody-bound beads overnight at 4oC. After successive washes, immunoprecipitated

chromatin was eluted in elution buffer (1% SDS, 100mM NaHCO3) and then reverse-

crosslinked overnight at 65oC. Following RNase A and Proteinase K treatment, DNA was

extracted with phenol/chloroform and analyzed by quantitative PCR using primers listed in

Table 2.2. For sequential ChIP assays, chromatin was harvested and MNase-digested as

described (174). Immunoprecipitates from the first ChIP were eluted in elution buffer

containing 15 mM DTT for 30 min at 37oC. After 20-fold dilution in ChIP lysis buffer

(without SDS), samples were subjected to IP with a second antibody and processed as the

first round of ChIP. For both ChIP and sequential ChIP assays, input and immunoprecipitated

material were analyzed in parallel using PerfeCTa SYBR Green SuperMix (Quanta) on a

7900HT fast real-time PCR system (Applied Biosystems). Reactions were performed in

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triplicate and represented as percentage of precipitated material relative to the amount in the

input sample. For sequential ChIP assays, fold enrichment represents % Input expressed

relative to the vector alone control. ChIP-qPCR results are represented as means ± standard

deviation (n=3), and are representative of at least 3 independent experiments. Antibodies and

primer sequences used are listed in Tables 2.1-2.2.

2.4 Results

2.4.1 H3 kinase MSK1, not RSK2, activates transcription of reporter

To study the effects of H3 phosphorylation on transcriptional activation, we first

asked whether H3 phosphorylation, through direct targeting of known H3 kinases, is

sufficient to activate transcription of reporter genes. We fused constitutively active (CA) or

kinase-dead (KD) versions of two well-studied H3 kinases, RSK2 and MSK1 (168, 182), to

the Gal4-DNA binding domain (DBD) sequences, and transfected these constructs into 293T

cells along with a luciferase reporter containing five Gal4-binding sites upstream of the SV40

promoter. In spite of the equal expression levels of RSK2 and MSK1 kinases in the

transfected cells (Figure 2.1A, right), only Gal4-CA-MSK1 strongly activated expression of

the luciferase reporter (Figure 2.1A, left). Induction of the reporter gene by MSK1 was

completely dependent on a functional kinase domain (CA- vs. KD-constructs in Figure 2.1A)

and also on the presence of the Gal4-binding sites on the reporter (Figure 2.2A). Moreover,

the transactivation ability of Gal4-CA-MSK1 diminished proportionally when the Gal4

binding sites were systematically moved further upstream from the transcription start site of

the luciferase gene, suggesting that promoter proximity of the kinase target is needed for

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Figure 2.1. MSK1, but not RSK2, phosphorylates H3 at serine 28 and activates transcription of reporter. (A). Luciferase activities were measured 48 hours after cotransfection of Gal4-kinase constructs and luciferase reporter into 293T cells (left). The transcriptional activator Gal4-CBP was used as a positive control. Luciferase levels were normalized to β-gal levels from a

68

co-transfected β-gal plasmid, and data are represented as means ± standard deviation from 3 independent experiments. Expression of HA-tagged Gal4-kinase fusion proteins was detected by western blotting using an HA antibody (right). Total H3 was used as a loading control. CA, constitutively-active; KD, kinase-dead. (B-C). Luciferase assays were performed as in (A), in cells harboring an episomal reporter (B) or in a 293 cell line containing a stably-integrated reporter (C). (D). Flag-tagged H3 was cotransfected with RSK2 or MSK1 constructs into 293T cells. Phosphorylation levels of transfected H3 at S10 and S28 were examined by Western blotting. Total amount of transfected H3 was detected using Flag antibody and used as a loading control.

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Figure 2.2. Transcriptional activation by Gal4-CA-MSK1 depends on its targeting to the luciferase promoter. (A). 293T cells were cotransfected with Gal4-kinase fusion constructs and a luciferase reporter with (left) or without (right) Gal4 binding sites at the promoter region. Luciferase assays were performed as in Fig 2.1. When not targeted to the promoter (right), transactivation by Gal4-CA-MSK1 was completely abolished. (B). Gal4 binding sites were positioned at various distances from the promoter of the luciferase gene (-110, -550, -1100bp). Transactivation by MSK1 decreased as the kinase was moved further away from the promoter.

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maximal activation (Figure 2.2B). We also performed this assay using luciferase reporters

present on an episomal vector or stably integrated into the cellular genome. Again, in both

cases, only Gal4-CA-MSK1 effectively activated transcription of the reporter gene (Figure

2.1B-C). Of note, induction of these more properly chromatinized reporter genes is

significantly higher than the luciferase gene present on the non-episomal plasmid (50- to 60-

fold vs. ~ 20-fold induction, Figure 2.1A-C), supporting the conclusion that MSK1 activates

transcription by phosphorylating a chromatin component such as H3.

One intriguing finding from this experiment is that only MSK1, but not RSK2,

directly activated transcription of the reporter gene even though co-transfection of these

kinases with H3 substrates showed that both kinases are equally efficient at phosphorylating

H3 at S10 (Figure 2.1D). Interestingly, only MSK1 effectively phosphorylates H3 at both

S10 and S28, whereas RSK2 mainly phosphorylates H3 at S10 (Figure 2.1D). Although

many studies have strongly correlated H3S10ph with signaling-induced gene activation, our

direct assay suggests that phosphorylation of H3 at S10 alone (in this case mediated by

RSK2) is not sufficient for transcription initiation. Instead, it is H3 S28 phosphorylation,

either alone or in combination with S10 phosphorylation, that correlates with this

transcription step.

2.4.2 Targeting MSK1 to the endogenous c-fos promoter activates its expression

To test whether H3 phosphorylation also directly activates endogenous IE genes, we

took advantage of the Nuclear Factor 1 (NF1) binding site present at the c-fos promoter and

modified our targeting approach by fusing MSK1 to the DNA binding domain of NF1

(Figure 2.3A). The addition of the NF1-DBD did not affect the activation of CA-MSK1, as

indicated by the phosphorylation at the T-loop residue S212 of MSK1, a marker for the

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Figure 2.3. Targeting MSK1 to the endogenous c-fos promoter induces H3 S10 and S28 phosphorylation and activates its expression. (A). Schematic diagram of the c-fos promoter showing the locations of cis-regulatory elements and primers used for ChIP analysis. CA or KD-MSK1 was fused to the DNA binding domain of NF1 (residues 1-257) to directly target the H3 kinase to the endogenous c-fos promoter through its NF1 binding site. (B). Expression and kinase activity of different MSK1 constructs in 293T cells were examined by Western blotting using the indicated antibodies. Ponceau staining of core histones was used as loading control.

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(C). CA/KD-MSK1 constructs were transfected into 293T cells and their effects on expression of endogenous c-fos gene were determined by qRT-PCR. c-fos mRNA levels were normalized to that of HPRT. (D). ChIP assays using an HA antibody showed that CA- and KD-versions of NF1-MSK1 fusions were both targeted to the promoter of endogenous c-fos gene. (E). ChIP analyses using antibodies against phosphorylated and total H3 showed induced H3 S10 and S28 phosphorylation at the c-fos promoter by NF1-CA-MSK1. qRT-PCR and ChIP-qPCR results are represented as means ± standard deviation (n=3), and are representative of at least 3 independent experiments.

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activated form of this kinase (Figure 2.3B). However, addition of NF1-DBD to CA-MSK1

greatly enhanced phosphorylation of the endogenous H3 at S10 and S28 (Figure 2.3B),

suggesting that targeting this kinase to chromatin greatly increased its contact and

phosphorylation of chromatin H3. Importantly, expression of the NF1-CA-MSK1 fusion

proteins (in either orientation) also strongly activated transcription of the endogenous c-fos

gene (Figure 2.3C). Moreover, the NF1-KD-MSK1 controls did not activate c-fos

transcription over background levels, confirming that this induction is dependent on the

kinase activity of MSK1 and is not due to any cryptic transcriptional activation by the NF1-

DBD. As additional controls, we also tested the effects of NF1-CA-MSK1 on two other

endogenous genes: c-jun and HSP70. c-jun is another IE gene, but it does not have any NF1-

binding sites in its promoter region, whereas the heat shock protein HSP70 gene contains an

NF1-binding site at its promoter and its expression has been associated with H3

phosphorylation (238). qRT-PCR analyses showed that NF1-CA-MSK1 induced expression

of HSP70, but not c-jun (Figure 2.4). All together, these findings show that the engineered

fusion kinase specifically activates genes that have NF1-binding sites.

Using ChIP assays, we further confirmed that both NF1-CA-MSK1 and NF1-KD-

MSK1 were targeted and enriched at the c-fos promoter with equal efficiencies (Figure

2.3D). Consistent with their transcriptional activation abilities, only NF1-CA-MSK1, but not

the KD-version, increased both H3S10ph and H3S28ph levels at the c-fos promoter (Figure

2.3E). We note that the induction of H3S28ph levels was much greater than the H3S10ph

levels (a 30-fold vs. 2-fold increase), which supports our hypothesis that H3 S28

phosphorylation is more relevant to transcriptional activation compared to H3 S10

phosphorylation. Taken together, these results demonstrate that MSK1 can directly activate

c-fos transcription and that targeting of this H3 kinase bypasses the requirement of upstream

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Figure 2.4. NF1-CA-MSK1 fusion induced expression of c-fos, HSP70 but not c-jun. mRNA levels of immediate-early genes c-fos and c-jun and heat shock protein HSP70 were examined by qRT-PCR. Relative gene expression was normalized to HPRT levels, and represented as means ± standard deviation.

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signaling to activate an IE gene.

2.4.3 MSK1 and H3 phosphorylation reactivate expression of polycomb-silenced α-

globin in non-erythroid cells

Given our hypothesis that H3 S28 phosphorylation is important for transcriptional

activation, we next asked whether there is functional interplay between this modification and

the repressive mark on the adjacent K27 residue. Methylation of K27 on H3 is a key step in

the polycomb-silencing pathway, which regulates a large number of genes critical for

development and differentiation (255). Previous studies have shown that H3 phosphorylation

at S10 disrupts the binding of heterochromatin protein HP1 to the adjacent methylated K9

residue and functions to displace HP1 from condensed chromosomes during mitosis (231,

232). Given the highly analogous nature of H3 K9-methylation/S10-phosphorylation and

K27-methylation/S28-phosphorylation, we wondered whether S28 phosphorylation also

disrupts the H3 K27 methylation-mediated polycomb pathway.

By targeting MSK1 to an endogenous polycomb-silenced gene, we could directly test

the impact of H3 S28 phosphorylation on this repression pathway. To that end, we have

chosen the tissue-specific α-globin gene (HBA1/2) as a model polycomb-regulated gene for

several reasons. First, the α-globin gene is a developmentally regulated and tissue-specific

gene expressed only in erythroid cells, and its repressed state in non-erythroid cells is

mediated by H3 K27 methylation and PRC2 (256). Second, the silenced gene can be

reactivated in non-erythroid cells upon herpes simplex virus infection (257) or by treatment

with the HDAC inhibitor TSA (256). Finally, a conserved NF1 recognition element is also

present in the α-globin promoter (258). Similar to the previous results for the c-fos gene,

ChIP assays showed that NF1-CA-MSK1 and NF1-KD-MSK1 were efficiently targeted to

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the α-globin promoter (Figure 2.5E). In addition, only NF1-CA-MSK1 induced H3 S10 and

S28 phosphorylation and, again, the S28 site was the more significantly phosphorylated site

(Figure 2.5D). Finally and most importantly, NF1-CA-MSK1 strongly activated transcription

of the silenced α-globin gene (as measured by both semi-quantitative PCR as well as

quantitative real time PCR, Figure 2.5B-C) whereas neither the NF1-KD-MSK1, nor the non-

targeted CA-MSK1, could reactivate this gene. This observation strongly suggests that

polycomb silencing on the α-globin promoter can be disrupted or reactivated by targeted H3

S28 phosphorylation.

Since we detected both H3S10ph and H3S28ph at the promoter regions of c-fos and

α-globin, we further asked whether these two phospho-marks exist in the same nucleosome

by performing sequential ChIP assays. Previous studies have shown that TPA-induced

phosphorylation of H3 S10 and S28 does not occur on the same nucleosome at IE gene

promoters (174). Consistent with those results, we also observed that H3S10ph and H3S28ph

do not co-exist within the same nucleosome at the NF1-CA-MSK1-activated c-fos promoter

(Figure 2.5F). However, we did detect a strong co-occupancy of the two phospho-H3 marks

at the activated α-globin promoter, suggesting that these two marks can co-exist. At present,

we do not know what dictates whether H3S10ph and S28ph can co-exist or not, but the

underlying differences in the genomic context of c-fos, a gene that is poised for expression,

and α-globin, a gene that is actively silenced, are likely determining factors.

2.4.4 H3 S28 phosphorylation disrupts PRC recruitment and H3 K27 methylation

To gain additional mechanistic insights into the reactivation of the α-globin gene, we

used ChIP assays to examine the H3K27me3 status as well as recruitment of representative

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Figure 2.5. MSK1 and H3 phosphorylation reactivate expression of polycomb-silenced α-globin gene in non-erythroid cells. (A). Schematic diagram of the α-globin promoter showing the targeting of MSK1 to the endogenous α-globin promoter through its NF1 binding site and the location of primers used for ChIP analysis. (B-C). Expression of α-globin in 293T cells expressing different MSK1 constructs was measured by semi-quantitative (B) and quantitative RT-PCR (C).

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(D-E). ChIP analyses using the indicated antibodies showed that both NF1-CA- and NF1-KD-MSK1 were targeted to the endogenous α-globin promoter, but only the active kinase version induced H3 S10 and S28 phosphorylation. (F). Sequential ChIP assays were performed with antibodies against H3S10ph and H3S28ph. Co-existence of the two phospho-marks in the same nucleosome was detected at the induced α-globin promoter, but not c-fos promoter.

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PRC components EZH2 and BMI1 at the reactivated α-globin promoter. We note that our

initial Western blot analyses of total cell lysates showed that expression of NF1-CA-MSK1

strongly reduced H3K27me3 levels (Figure 2.6A, left). However, more detailed tests

revealed that the apparent loss of H3K27me3 was due to epitope masking or occlusion

whereby the H3K27me3 antibody does not recognize its epitope when the adjacent S28

residue is phosphorylated. Indeed, when we pre-treated the lysates with lambda phosphatase,

detection of the H3K27me3 epitope by the antibody was restored (Figure 2.6A, right). To

eliminate this potential epitope occlusion problem in our ChIP analyses, we performed all the

H3K27me3, as well as H3K27ac, ChIP assays using phosphatase-treated chromatin. With

such technical precautions, we found that the α-globin promoter in 293T cells is enriched for

H3K27me3, and the enrichment level is similar to that of two other polycomb-silenced

genes, HoxD4 and Myt1 (Figure 2.6B). In addition, cells expressing NF1-CA-MSK1

consistently have reduced levels of H3K27me3, EZH2 and BMI1 at the α-globin promoter

(Figure 2.7A, all reduced by about 25-35% compared to the vector alone control). The

depletion of these polycomb-silencing hallmarks is dependent on the kinase activity of

MSK1 (the vector alone or NF1-KD-MSK1-transfected cells show comparable enrichment of

these factors), suggesting that phosphorylation of H3 at S28 displaces binding of EZH2 and

BMI1. The loss of H3K27me3 is not 100% suggesting that this is not due to large-scale

active de-methylation of the whole locus, but possibly restricted to regions close to the NF1-

binding site. Our findings show that a phos/methyl switch mechanism does exist in the case

of H3S28ph whereby this modification functionally antagonizes the recruitment of PRC

components. Moreover, even without complete loss of H3K27me3, strategic targeting of the

MSK1 kinase is sufficient to reactivate the previously silenced α-globin gene.

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Figure 2.6. α-globin promoter in 293T cells is enriched for H3K27me3. (A). Recognition of the H3K27me3 epitope by the H3K27me3 antibody (Active Motif, 39156) was occluded by the adjacent phosphorylated S28 residue. Total cell lysates from transfected 293T cells were incubated at 30oC for 30min in the absence (left) or presence (right) of lambda phosphatase, and were then analyzed by Western blotting using the indicated antibodies. Phosphatase treatment removed the phosphate groups from H3 S10 and S28, but restored the epitope recognition by H3K27me3 antibody. Ponceau staining of core histones was used as a loading control. Similar results were observed when the H3K27me3 antibody from Millipore (07-449) was used. (B). ChIP analysis showed that levels of H3K27me3 at the α-globin promoter are comparable to that of two other polycomb-repressed genes, Myt1 and HoxD4.

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Figure 2.7. H3 S28 phosphorylation disrupts PRC (Polycomb Repressive Complex) recruitment and H3 K27 methylation. 293T cells were transfected with pcDNA vector alone or expression vectors for NF1-CA-MSK1 or NF1-KD-MSK1, and were harvested 30 hours after transfection for ChIP assays. ChIP assays were performed to examine (A) H3K27me3 status and (B) recruitment of PRC components at the α-globin promoter. Cells expressing NF1-CA-MSK1 consistently have lower levels of H3K27me3 and reduced binding of PRC2 subunit Ezh2 and PRC1 subunit Bmi1at the reactivated α-globin promoter.

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2.4.5 Coupling of H3 S28 phosphorylation and K27 acetylation antagonizes polycomb

silencing

In many cases, activation of polycomb-regulated genes is not only associated with a

loss of H3K27me3, but also accompanied by a concomitant gain in H3 K27 acetylation

(259). Therefore, we also examined the impact of NF1-CA-MSK1 on H3K27ac at the global

and gene-specific levels. Remarkably, expression of NF1-CA-MSK1 not only enhanced H3

S10 and S28 phosphorylation, but also induced a significant increase in H3 K27 acetylation

(Figure 2.8A). This increase is dependent on the kinase activity of MSK1, indicating that

H3K27ac is functionally coupled to the S28 phosphorylation event. By ChIP analyses, we

found that NF1-CA-MSK1 targeting also increased the H3K27ac levels at the c-fos and α-

globin promoters (Figure 2.8B). The relative increase of H3K27ac at the c-fos promoter is

not as high as that at the α-globin promoter due to the already high steady-state levels of

H3K27ac associated with the c-fos gene.

A functional coupling between H3 K27 acetylation and S28 phosphorylation has not

been reported before. To test whether these two modifications are also physically linked, we

used an antibody that specifically recognizes the di-modified H3K27ac/S28ph epitope

(Figure 2.9), similar to the antibody we previously generated against H3S10ph/K14ac (170).

Western blotting of total histones showed that expression of NF1-CA-MSK1 induced the

phospho-acetylation of S28 and K27 on the same H3 molecule (Figure 2.8A). More

importantly, ChIP assays showed that such di-modified epitope is specifically increased at

the NF1-CA-MSK1-activated α-globin promoter, indicating that the combination of these

two modifications directly correlates with the reactivation of this silenced gene (Figure

2.8C). To test whether the newly identified H3K27ac/S28ph mark is also induced by mitogen

or stress, and to examine the kinetics of the induction, we stimulated quiescent 10T1/2 mouse

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Figure 2.8. Functional and physical coupling of H3 S27 acetylation and S28 phosphorylation antagonizes polycomb silencing. (A). Total cell lysates from transfected 293T cells were examined by Western blot analyses. Global changes in post-translational modifications on H3 were determined using the indicated antibodies. (B). Increase in H3K27 acetylation levels was detected by ChIP at the α-globin and c-fos promoter in NF1-CA-MSK1-transfected cells. (C). ChIP analyses using an antibody specific for the di-modified H3K27ac/S28ph mark showed that H3 K27 acetylation and H3 S28 phosphorylation are physically linked and co-exist on the same H3 tail at the induced α-globin and c-fos promoter.

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Figure 2.9. Specificity of the α-H3K27ac/S28ph antibody was tested using peptide competition assays. α-H3K27ac/S28ph antibody was preincubated with the indicated peptides (3µg/ml) for 15 min prior to being used for Western blot analysis of TPA-treated lysates. Only the di-modified peptide completely blocked detection of the modified H3, suggesting that this antibody detected H3 that are acetylated at K27 and phosphorylated at S28.

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fibroblasts with TPA or anisomycin and examined the phosphorylation status of H3 at

various times post stimulation (Figure 2.10). We found that TPA and anisomycin not only led

to the activation and phosphorylation of MSK1 and a corresponding increase in H3 S10/S28

phosphorylation, but it also strongly induced the di-modified H3K27ac/S28ph mark.

Importantly, induction of all these phopho-H3 marks was abolished when cells were pre-

treated with the MSK1 inhibitor H89, suggesting that mitogen/stress-induced

H3K27ac/S28ph is dependent on MSK1 activity. Taken all together, these findings

demonstrate that MSK1-mediated H3S28ph, in response to various upstream signaling

conditions, enhances acetylation of the adjacent K27 residue on H3. Taken together, these

findings demonstrate that MSK1-mediated H3S28ph, in response to various upstream

signaling conditions, enhances acetylation of the adjacent K27 residue on H3.

2.4.6 Both H3S28ph and H3K27ac/S28ph are directly associated with the initiating

form of RNAP II

Given our finding that the stimulated form of MSK1 directly activates transcription,

and that this enzyme induces H3S10ph, S28ph as well as H3K27ac/S28ph, we further tested

whether all or some of these modified forms of H3 are specifically associated with

transcription initiation. To do this, we performed sequential ChIP assays by first using

antibodies against the aforementioned modified forms of H3, and followed by a second round

of ChIP using an antibody against the S5-phosphorylated form of RNA polymerase II

(RNAP-S5ph), which represents the initiating form of RNAP II. By this assay, we found a

strong association of H3S28ph, as well as H3K27ac/S28ph, with RNAP-S5ph at the NF1-

CA-MSK1-activated c-fos and α-globin promoters (Figure 2.11). In contrast, no co-

occupancy of H3S10ph and RNAP-S5ph was observed at these loci in any of the transfected

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Figure 2.10. Analysis of H3 phosphorylation marks in TPA- or anisomycin-induced 10T1/2 cells. (A). Serum-starved 10T1/2 cells were pretreated with DMSO (mock treatment) or H89 (MSK1 inhibitor) for 30 min prior to stimulation with 400ng/ml TPA. Total cell lysates were analyzed by western blotting using the indicated antibodies, and coomassie staining of core histones was used as loading control. (B). Serum-starved 10T1/2 cells were stimulated with 50ng/ml anisomycin and processed as in (A).

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Figure 2.11. H3S28ph and H3K27ac/S28ph marks correlate with transcription initiation at the activated c-fos and α-globin promoters. Sequential ChIP assays were performed on the promoters of (A) c-fos and (B) α-globin. Antibody against H3S10ph (left), H3S28ph (middle) or H3K27ac/S28ph (right) was used in the first ChIP and then followed by a second round of ChIP with antibody against phospho-S5 of RNAP II (RNAP-S5ph), which represents the initiating form of RNAP II. Fold enrichment represents % Input expressed relative to the vector alone control at the corresponding promoter.

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cells. Our data thus suggest that, at both the c-fos and α-globin promoters, the initiating form

of RNAP II is directly and physically associated with H3S28ph and H3K27ac/S28ph, but not

with H3S10ph.

2.5 Discussion

H3 phosphorylation is mostly studied in the context of rapid activation of signal-

inducible genes, such as the induction of IE genes upon mitogen or stress stimulation.

However, recent studies showed that additional signaling pathways, such as the Toll-like

receptor and retinoic acid signaling, also activate MSK1 to regulate non-IE genes (196, 260).

While many studies have provided excellent correlative data linking this histone modification

to transcriptional regulation, they cannot distinguish between direct and indirect effects of H3

phosphorylation. Here we show that the H3 kinase MSK1 is a potent transcription activator

when directly targeted to diverse promoters such as the luciferase reporter, the endogenous

IE gene c-fos, as well as the polycomb-silenced α-globin gene. The strength of our system is

that we can study the direct effect of MSK1 and H3 phosphorylation on gene expression

without external stimuli and, thus, without complications from multiple upstream signaling

pathways. Most studies to date have focused on H3 S10 phosphorylation; however, our

results suggest that phosphorylation of this site alone, mediated by the RSK2 kinase, is not

sufficient to directly transactivate the luciferase reporter gene. Instead, it is the induction of

H3 S28 phosphorylation that mirrors transcriptional activation. This is particularly evident in

our ChIP analyses whereby we consistently observed much greater induction of H3S28ph,

compared to H3S10ph, at the activated promoters. In addition, sequential ChIP analyses

showed that H3S28ph and H3K27ac/S28ph, but not H3S10ph, are directly associated with

the transcription-initiating form of RNAP II.

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Throughout our studies, we consistently observed a correlation between H3S28ph and

transcriptional activation at the MSK1-targeted genes. However, previous studies also found

that H3 S10 phosphorylation at the FOSL1 enhancer (229) and Drosophila heat shock genes

(180) promotes the release of paused RNAP II from a promoter proximal site through the

recruitment of the transcription elongation factor P-TEFb. It is possible that H3 S10

phosphorylation facilitates transcriptional elongation of genes that are regulated by

polymerase pausing, whereas H3 S28 phosphorylation directly activates transcription at the

initiation step. Further studies will be required to distinguish between the roles of H3

phosphorylation at these distinct sites in the transcriptional initiation and elongation steps.

To date, little is known about the link between H3 S28 phosphorylation and

transcription. ChIP analyses showed that H3S28ph is enriched at IE gene promoters upon

their activation (174). Interestingly, in chicken erythrocytes, H3 S28 phosphorylation

preferentially occurs on the transcription-linked H3 variant H3.3 (206), supporting our

hypothesis that S28 phosphorylation is functionally linked to transcriptional activation.

Originally identified by a H3S10ph peptide pulldown assay, 14-3-3 actually has a much

higher binding affinity for the H3S10ph/K14ac di-modified as well as the H3S28ph epitopes

(227, 228). Given the general paradigm that histone modifications recruit PTM-specific

binding proteins to mediate downstream functions, it is likely that 14-3-3, or additional

proteins, bind to the phospho-S28 residue to facilitate transcriptional activation. In that

regard, how acetylation of K27 might synergistically or antagonistically modulate

recruitment of such S28ph binding protein represents yet another potential level of functional

cross talk between histone modifications. The mechanism that couples H3 K27 acetylation

and S28 phosphorylation is currently unknown. It is possible that MSK1 and/or its kinase

activity recruit an H3 K27 HAT to the promoter of target genes. Alternatively, H3

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phosphorylated at S28 may be a better substrate for the H3 K27 HAT. In support of the first

scenario, MSK1 was previously shown to co-immunoprecipitate with multiple HATs,

including p300 and CBP (261), which are known to acetylate H3 at K27 (259). As for the

second scenario, we and others have previous shown that the yeast HAT, Gcn5, preferentially

acetylates H3S10ph peptides over the unmodified form (170). Therefore, H3 K27 HATs may

also have a preference for S28-phosphorylated H3 as substrate. These two possibilities are

not necessarily mutually exclusive, but further experiments will be required to test these

hypotheses.

Using the tissue-specific α-globin gene as a model polycomb-regulated gene, our

study identified a novel histone crosstalk pathway whereby H3 S28 phosphorylation induces

a methyl-acetylation switch on the adjacent K27 residue (Figure 2.12). Moreover, this

mechanism provides a direct link between signal transduction and polycomb-regulation.

Activation of polycomb target genes during differentiation is associated with displacement of

polycomb group proteins and removal of H3K27me3; however, how this process is regulated

is still poorly understood. Recent studies also showed that a switch from methylation to

acetylation at H3 K27 often accompanies the activation of these genes. Our finding suggests

that one way to regulate this switch can be through phosphorylation of the adjacent S28

residue. Given that genome-wide screens showed that many polycomb-regulated genes are

downstream of diverse signaling pathways (91), our finding that H3 S28ph and K27ac are

functionally coupled further raises the possibility that signal transduction pathways and

activation of polycomb-regulated genes are directly linked through this novel histone

crosstalk. If so, H3 S28 phosphorylation may have a yet-to-be appreciated function in

modulating the epigenome during development and differentiation.

While this work was under peer review in PNAS, Gehani et al. reported similar

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Figure 2.12. H3 S28 phosphorylation initiates a novel histone code pathway by inducing a methyl-acetylation switch of the adjacent K27 residue. Our data support a model in which MSK1 and H3 S28 phosphorylation antagonize polycomb silencing through the displacement of polycomb repressive complexes and the removal of H3K27me3. Upon recruitment of a H3K27 HAT, the di-modified H3K27ac/S28ph mark is established, which can then recruit specific chromatin modifiers or transcription regulators to further modulate gene expression.

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findings in that H3 S28 phosphorylation, mediated by MSK1/2, was important for activation

of polycomb target genes (262). They began their studies by looking at the activation of PcG-

repressed genes upon external stimuli. They found that, in response to mitogen, stress and

differentiation signals, MSK1/2 are activated and phosphorylate H3 S28 on PcG target genes.

They also determined that addition of a phosphate group to S28 interferes with the binding of

PRC components to H3K27me3, and using an antibody specific for H3K27me3/S28ph, they

found such di-modified H3 at the activated PcG target genes. More importantly, inhibition of

MSK kinases by the inhibitor H89, or knockdown of MSK expression by RNAi, abolished

the induction of H3S28ph and H3K27me3/S28ph marks, and strongly compromised signal-

induced expression of PcG targets. Therefore, their data suggest that, in response to

environmental cues or differentiation signals, MSK1/2 and H3 S28 phosphorylation directly

counteract polycomb repression.

In spite of the different starting points of these two studies, both of them come to a

remarkably consistent conclusion: H3 S28 phosphorylation functionally triggers activation of

polycomb-silenced genes. On the other hand, while Gehani et al. showed that the di-modified

H3K27me3/S28ph mark exists in vivo and suggests that this mark is important for activating

polycomb-regulated genes, my work suggests that H3S28ph is functionally and physically

coupled to H3K27ac. Moreover, this phospho-acetyl mark (H3K27ac/S28ph) is physically

associated with the transcription initiating (serine 5-phosphorylated) form of RNAP II.

Further studies comparing the distribution and timing of these different dual modifications on

H3 will help elucidate the functional differences and significances of these distinct H3

modification combinations in the polycomb regulation pathway.

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Chapter 3

Elucidating combinatorial histone modifications and crosstalks by

coupling histone-modifying enzyme and biotin ligase activity

A version of this chapter is in press in Nucleic Acids Research as: Lau, P.N.I., and Cheung, P. (2012). Elucidating combinatorial histone modifications and crosstalks by coupling histone-modifying enzyme and biotin ligase activity. I prepared the samples for acid-urea gel analysis in Figure 3.3B, and Dr. Peter Cheung ran the acid urea gel. I performed all other experiments.

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3 BICON 3.1 Abstract

Histone post-translational modifications (PTMs) often form complex patterns of

combinations and cooperate to specify downstream biological processes. In order to

systemically analyze combinatorial PTMs and crosstalks amongst histone PTMs, we have

developed a novel nucleosome purification method called Biotinylation-assisted Isolation of

CO-modified Nucleosomes (BICON). This technique is based on physical coupling of the

enzymatic activity of a histone-modifying enzyme with in vivo biotinylation by the biotin

ligase BirA, and using streptavidin to purify the co-modified nucleosomes. Analyzing the

nucleosomes isolated by BICON allows the identification of PTM combinations that are

enriched on the modified nucleosomes and function together within the nucleosome context.

We used this new approach to study MSK1-mediated H3 phosphorylation and found that

MSK1 not only directly phosphorylated H3, but also induced hyperacetylation of both

histone H3 and H4 within the nucleosome. Moreover, we identified a novel crosstalk

pathway between H3 phosphorylation and H4 acetylation on K12. Involvement of these

acetyl marks in MSK1-mediated transcription was further confirmed by ChIP assays, thus

validating the biological relevance of the BICON results. These studies serve as proof of

principle for this new technical approach, and demonstrate that BICON can be further

adapted to study PTMs and crosstalks associated with other histone-modifying enzymes.

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3.2 Introduction

Histones are subjected to a variety of post-translational modifications (PTMs)

including acetylation, methylation, phosphorylation, ubiquitylation and sumoylation (12).

Histone-modifying enzymes, and their resultant PTMs, can be viewed as an extension of

signal transduction networks. They function to transmit signals to chromatin, which then

translates external stimuli into the appropriate nuclear responses (253, 263). Moreover,

signaling cascades also occur on histones, whereby one PTM on a histone can positively or

negatively influence the deposition of other downstream PTMs (264). Such crosstalk can

occur within the same histone tail (cis crosstalk) or between different histones (trans

crosstalk). One of the earliest examples of histone PTM crosstalk is the direct coupling of

phosphorylation and acetylation on H3 during gene activation, whereby phosphorylation of

S10 on H3 facilitates subsequent acetylation on the neighbouring K14 by the Gcn5

acetyltransferase (170, 233). Thee trans-tail crosstalk pathways are best exemplified by the

functional links between H2B ubiquitylation and H3 methylation. More specifically, in yeast

and human cells, mono-ubiquitylation of H2B at the C-terminus is an absolute pre-requisite

for H3K4 methylation by Set1 and H3K79 methylation by Dot1 (148–151). This complex

interplay between PTMs dictates the legitimate combinations of histone modifications that

can occur together on nucleosomes, and coordinates their functions to elicit specific

outcomes (71, 240).

One well-established mechanism by which histone PTMs mediate downstream

functions is through the recruitment and direct binding of downstream effector proteins to

chromatin. This docking of effectors to modified histones is mediated through conserved

protein modules that recognize histone PTMs in modification-specific and site-specific

manners (207). For example, bromodomains and chromodomains are conserved structural

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motifs that respectively bind acetylated and methylated lysine residues. Increasing evidence

also suggests that multiple histone-binding modules function in concert, and coordinately

engage multiple PTMs within a nucleosome (265). Specific combinations of histone PTMs

can therefore serve as multivalent docking sites for stabilizing the contacts between

chromatin and the recruited proteins or complexes. By doing so, they collectively coordinate

the recruitment (or repulsion) of effector binding proteins to direct biological outcomes.

Although most studies to date focus on individual histone PTMs, elucidating the

combinatorial patterns of histone PTMs and how they cross regulate one another represents

the next important frontier of this field.

H3 phosphorylation was first observed on mitotic chromosomes, but it is also

associated with transcriptional activation in interphase cells (112, 114, 164, 266). In response

to stress or growth factor stimulation, H3 is rapidly and transiently phosphorylated on S10

and/or S28 by signaling kinases such as MSK1. In the previous chapter, we showed that

MSK1 is a potent transcriptional activator. Targeting MSK1 to the immediate-early (IE) gene

c-fos, as well as the polycomb-silenced α-globin gene, not only efficiently phosphorylated

H3 at the gene promoters, but also strongly induced their transcription in the absence of other

external stimuli (267). Even though the functional differences between phosphorylation of

these 2 different serine residues are still largely unknown, it is clear that both H3S10ph and

H3S28ph can mediate multiple crosstalks with other histone modifications. For example,

H3S10ph is physically coupled to H3K14ac during mitogen-induced activation of IE genes

(170). Importantly, acetylation of H3 K14 enhances the interaction between H3S10ph and

14-3-3, suggesting that the dual modifications on H3S10 and K14 cooperate to recruit

downstream effector proteins (227, 228). At the FOSL1 enhancer, phosphorylation of H3S10

by PIM1 kinase not only recruits 14-3-3, but also induces acetylation on H4 K16, ultimately

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leading to transcription elongation (229). Besides recruiting 14-3-3 and other downstream

chromatin modifiers, H3 phosphorylation can also disrupt binding of chromodomain-

containing proteins to methylated H3. During mitosis and transcriptional activation,

phosphorylation of H3 S10 displaces HP1 from H3K9me3 (197, 231, 232). Such a

“phospho/methyl” switch also occurs on H3K27me3/H3S28ph, with H3S28ph displacing

polycomb-group proteins from polycomb-silenced genes (262, 267). Moreover, as shown in

Chapter 2, we found that phosphorylation of H3 S28 by H3 kinase MSK1 is functionally and

physically coupled to K27 acetylation, and this dual modification correlates with reactivation

of polycomb-silenced α-globin gene in nonerythroid cells (267). All these findings indicate

that H3 phosphorylation cooperates with PTMs on multiple histone sites and together they

regulate binding of effector proteins and downstream biological processes.

To extend these studies, we sought to develop an unbiased method to identify histone

PTMs that occur together with MSK1-mediated H3 phosphorylation. To that end, we

developed an original affinity purification approach, which we termed Biotinylation-assisted

Isolation of CO-modified Nucleosomes (BICON) to capture and study phospho-H3-

containing nucleosomes. This method involves the coupling of in vivo biotinylation mediated

by the E. coli BirA enzyme (268) and phosphorylation of H3 by MSK1, and using

streptavidin-coupled beads to isolate MSK1-modified nucleosomes. Analyzing the spectrum

of histone PTMs on these nucleosomes, we not only found that their H3 are hyper-

phosphorylated, but specific residues on H3 and H4 are also hyper-acetylated. This suggests

that crosstalk between phosphorylation and acetylation occurs both in cis and in trans within

the nucleosome. Importantly, ChIP assays examining MSK1-target genes confirmed that

these specific combinations of histone modifications are induced upon gene activation.

Therefore, these studies showed that the BICON method not only revealed combinatorial

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histone PTMs and new histone crosstalks, but also illustrated the potential usefulness of this

technique.

3.3 Materials and Methods

Plasmid constructs

HA-tagged CA-MSK1 and KD-MSK1 in pMT2 were provided by Dr. Morten Frodin

(University of Copenhagen, Denmark). For Avi-Flag tagging, a tandem Avi-tag followed by

a Flag-tag was fused in frame to the 3’end of the H3.3 coding sequence. The Avi-tag refers to

a 15 amino acid sequence (GLNDIFEAQKIEWHE) that contains a biotinylation site for the

E.coli biotin ligase BirA. BirA expression construct was provided by Dr. John Strouboulis

(Alexander Fleming Biomedical Sciences Research Center, Greece). BirA coding sequence

was PCR-amplified and fused in frame to the N-terminal side of CA- or KD-MSK1 to

generate the BirA-MSK1 fusion constructs in pcDNA3.1+. NF1-CA/KD-MSK1 constructs

were described in Chapter 2.3.

Cell culture, transfections, TPA and H89 treatment

293T cells were grown in DMEM (Sigma) supplemented with 10% FBS. All transfections

were performed using Lipofectamine 2000. For TPA (Sigma) stimulation, 293T cells were

switched to serum-free media for 20h prior to stimulation. Serum-starved cells were mock-

treated with DMSO or stimulated with 400ng/ml TPA for 0-90 min and then harvested for

ChIP assays. To inhibit MSK1 activity, 10µM H89 (Sigma) was added to cells for 30 min

before stimulation with TPA.

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Harvesting of protein samples, gel electrophoresis, Western blot analyses and

antibodies

Whole cell lysates were prepared by directly lysing pelleted cells in boiling SDS sample

buffer. For acid extraction of histones, histones were extracted with 0.4N H2SO4 and then

precipitated with 20% trichloroacetic acid (TCA). Acid-extracted histones were analyzed on

an acid-urea gel system as described in (170). Whole cell lysates and isolated

mononucleosomes were resolved on SDS-PAGE gels and transferred to PVDF membranes

for Western blotting analyses. Antibodies used in this study are as follows: H3 (ab1791) and

H3K27ac (ab4729) from Abcam, H3S10ph (sc-8656R) and 14-3-3ζ (sc-1019) from Santa

Cruz, H3S28ph (H9908), Flag (F1804) and avidin-horseradish peroxidase (Avi-HRP, A3151)

from Sigma-Aldrich, H3K14ac (06-911), H3ac (06-599), H4ac (06-946), H4K12ac (07-595),

H4K16ac (07-329), H4 (05-858) and Rabbit IgG (12-370) from Millipore, HA.11 (MMS-

101R) from Covance, H2AZ (39113) from Active Motif, and H3K27ac/S28ph antibody was

described in Chapter 2.3.

Nucleosome purification with streptavidin beads

293T cells were cotransfected with H3.3-AviFlag and pcDNA vector alone or constructs that

express NLS-BirA, BirA-CA-MSK1 or BirA-KD-MSK1. 38 h after transfection, transfected

cells were trypsinized and washed twice with PBS. Cell pellets were resuspended in buffer A

(20 mM HEPES, pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM

dithiothreitol and protease inhibitors) containing 0.2% Triton X-100, and then incubated on

ice for 5 min. Nuclei were pelleted by centrifugation at 600g, and washed once in MNase

Cutting Buffer (10 mM HEPES, pH 7.9, 60 mM KCl, 10 mM NaCl). Nuclei were then

resuspended in MNase Cutting Buffer containing 3 mM CaCl2 and 2mM p-chloromercuri-

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phenylsulfonic acid (pCMPS, Toronto Research Labs; as phosphatase inhibitor), and digested

with micrococcal nuclease (MNase, Worthington) at 37°C for 30 min at a concentration of

10U/107 cells. Digestion of chromatin by MNase was stopped by addition of EGTA to a final

concentration of 5 mM, and equal volume of 2 x Lysis Buffer (30 mM HEPES, pH 7.9, 220

mM KCl, 3 mM MgCl2, 2 mM EDTA, 0.4% Triton X-100, 20% glycerol) was added to the

digested chromatin. Samples were centrifuged at maximum speed for 5 min to remove cell

debris, and the resulting supernatant containing mononucleosomes was used as input material

for affinity purification. For isolation of biotinylated nucleosomes, input chromatin was

incubated with streptavidin-agarose beads (Sigma S1638; 20 µl bed volume/sample)

overnight at 4°C. Beads were washed 3 times in WB1 (20 mM HEPES, pH 7.9, 140 mM

KCl, 1.5 mM MgCl2, 0.2 mM EGTA, 0.2% Triton X-100, and 10% glycerol), following by 2

times in WB2 (20 mM HEPES, pH 7.9, 300 mM KCl, 1.5 mM MgCl2, 0.2 mM EGTA, 0.5%

Triton X-100, and 10% glycerol) and once in WB1 again. Bound materials were eluted in 2 x

SDS-sample buffer without reducing agent (100 mM Tris, pH 6.8, 4% SDS, and 20%

glycerol), and boiled for 10 minutes. Eluted nucleosomes were resolved on 15% SDS-PAGE

gels and analyzed by Western blotting according to standard practices.

ChIP assays

ChIP assays were performed as previously described (170, 267). Briefly, for ChIP assays

involving NF1-MSK1, 293T cells were transfected with pcDNA vector alone or expression

vectors for NF1-CA-MSK1 or NF1-KD-MSK1 and were harvested 30 hours after

transfection. These transfected cells or DMSO/TPA-stimulated cells were fixed in 1%

formaldehyde for 8 min at room temperature. After swelling and lysis, chromatin was

sonicated to an average of ~500bp with a Branson Sonifier 450. Antibodies were prebound

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to protein A/G magnetic Dynabeads (Invitrogen) for at least 3h and rabbit IgG were used as

negative controls. Sonicated lysates were precleared with Dynabeads for 2 h and then

incubated with antibody-bound beads overnight at 4oC. After successive washes,

immunoprecipitated chromatin was eluted in elution buffer (1% SDS, 100mM NaHCO3) and

then reverse-crosslinked overnight at 65oC. Following RNase A and Proteinase K treatment,

DNA was extracted with phenol/chloroform and analyzed by quantitative PCR. Input and

immunoprecipitated material were analyzed in parallel using PerfeCTa SYBR Green

SuperMix (Quanta) on a 7900HT fast real-time PCR system (Applied Biosystems). Reactions

were performed in triplicate and presented as % of precipitated material relative to the

amount in the input sample (% input) and have been normalized to H3 levels to adjust for

nucleosome density. Signals from specific antibodies were at least 30-fold above rabbit IgG

controls (not shown). Error bars indicate means ± standard deviation (n=3), and are

representative of at least 3 independent experiments.

The following primers were used for ChIP-qPCR analyses: α-globin promoter, forward 5’-

GGGCCGGCACTCTTCTG-3’, reverse 5’- GGCCTTGACGTTGGTCTTGT-3’; control

region (upstream of α-globin), forward 5’-GAGATGCTGGAGTCAGGACCAT-3’, reverse

5’-AGGAGTCAGGAGCAGCAGTCA-3’; c-fos promoter, forward 5’-

GAGCAGTTCCCGTCAATCC-3’, reverse 5’- GCATTTCGCAGTTCCTGTCT -3’.

3.4 Results

3.4.1 Coupling of in vivo biotinylation and MSK1 phosphorylation

Our previous study (presented in Chapter 2) showed that phosphorylation of H3S28

by MSK1 can enhance acetylation of the adjacent K27 residue (267). This generates a di-

modified H3K27acS28ph mark, which correlates with transcriptional activation. To extend

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the studies, we aimed to further dissect the PTMs that co-exist and functionally synergize

with H3 phosphorylation, and to examine the crosstalk pathways initiated by the MSK1

kinase. To do this, we needed to specifically isolate phospho-H3-containing nucleosomes for

analyses. One major obstacle to this goal is that only a small fraction of total H3 is

phosphorylated in response to mitogen or stress stimulation (165). To overcome this

limitation, we developed a new approach, which we termed Biotinylation-assisted Isolation

of CO-modified Nucleosomes (BICON), to isolate and enrich for MSK1-modified H3 and its

associated nucleosomes. This method is based on an in vivo biotinylation system, which

utilizes the E coli biotin ligase BirA’s ability to recognize and biotinylate a short 15 amino

acid peptide called Avi-tag (268). Because there are no endogenous mammalian proteins that

contain this BirA recognition sequence, only introduced substrates containing the Avi-tag

will be biotinylated. Therefore, this allows very specific isolation of the BirA-targeted

substrate using avidin/streptavidin-coupled beads. For our purpose, we note that H3.3 is the

transcription-associated isoform of H3 (269), and that MSK1 has a two-fold preference in

phosphorylating H3.3 over canonical H3.1 (Figure 3.1). Therefore, we chose this variant as

the targeted substrate in our studies. This in vivo biotinylation system has been successfully

employed to study protein-protein interactions and protein complexes in a variety of cell

types (268). It has also been adapted for ChIP assays to map chromosomal targets of

transcription factors and histone-modifying enzymes (270). More importantly, this system

has been used to biochemically purify an assembly complex for Drosophila centromeric H3,

CID, and also has been applied to H3.3 for epigenomic profiling of this H3 variant in

Caenorhabditis elegans (271, 272), indicating that biotinylation of Avi-tagged histones does

not alter their biological properties.

To enrich for MSK1-phosphorylated H3.3, we fused BirA to constitutively active

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Figure 3.1. MSK1 preferentially phosphorylates H3 variant H3.3 on S28. Flag-tagged H3.1 or H3.3 was cotransfected with CA- or KD-MSK1 into 293T cells. Total cell lysates were resolved on SDS-PAGE gels, and phosphorylation levels of transfected H3 at S10 and S28 were examined by Western blotting. Total amount of transfected H3 was detected using α-Flag antibody and was used as a loading control. Serial dilutions of the CA-MSK1 samples (1x, 2x, 4x) were loaded to estimate the difference in phosphorylation levels. MSK1 phosphorylates S10 equally on H3.1 and H3.3. H3S28ph level is estimated to be ~2-fold more on H3.3 compared to H3.1 (compare lanes 4 and 8). CA, constitutively-active; KD, kinase-dead.

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(CA)-MSK1 to generate a BirA-CA-MSK1 fusion protein, such that biotinylation and

phosphorylation can occur together on the same H3.3 molecule (Figure 3.2). We reasoned

that such co-modification will allow preferential enrichment of phosphorylated-H3.3

nucleosomes upon incubation with streptavidin beads, and allow us to study the global PTMs

on these nucleosomes. A kinase-dead version, BirA-KD-MSK1, was also generated and used

as a control to generate biotinylated, but not phosphorylated, H3.3 for comparison. When the

BirA-MSK1 fusion enzymes and the Avi-tagged H3.3 were co-expressed in 293T cells, there

were no observable differences in the growth of these cells compared to the untransfected

control cells. To assess the ability of the BirA-MSK1 fusions to biotinylate and

phosphorylate H3.3, biotinylation and phosphorylation levels of the tagged H3.3 were

detected on Western blots using avidin-horseradish peroxidase (Avi-HRP) conjugate and

phospho-H3 specific antibodies respectively. These analyses confirmed that biotinylation of

the histone substrate is specific and dependent on the presence of BirA (Figure 3.3A).

Background histone biotinylation level is negligible and, therefore, is not a concern in this

purification scheme. In addition, BirA-CA-MSK1, but not KD, strongly phosphorylated H3.3

on S10 and S28 (Figure 3.3A), further confirming that the BirA-MSK1 fusions are functional

in vivo and modify the transfected H3.3 as intended.

We further verified our co-modification approach by analyzing the Avi-Flag-tagged

H3.3 on an acid-urea (AU) gel system, which separates proteins based on size and charge

(Figure 3.3B). Addition of a phosphate or acetyl group alters the overall charge on histones

and will cause an electrophoretic mobility shift on an AU gel (273). Using an anti-Flag

antibody to detect the total amount of transfected-H3.3, we observed no obvious change in

the banding pattern between different samples (Figure 3.3B, top). This suggests that only a

small fraction of transfected H3.3 is hyper-phosphorylated. However, detection by Avi-HRP

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Figure 3.2. Schematic of H3.3-AviFlag and BirA-Msk1 fusion constructs. E.coli biotin ligase BirA was fused to constitutively active (CA) or kinase-dead (KD)-MSK1 to generate BirA-CA-MSK1 or BirA-KD-MSK1 fusion proteins respectively. H3.3-AviFlag contains a tandem Avi-Flag tag at its C-terminus. The 15 amino acid Avi-tag encompasses a recognition sequence for BirA, which catalyzes the addition of biotin to the lysine residue on the tag. Also shown are the 2 serine phosphorylation sites (S10, S28) at the N-terminus of H3. (P, phosphorylation; B, biotinylation)

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Figure 3.3. In vivo biotinylation and phosphorylation of Avi-tagged H3.3. (A). Vector alone (pcDNA), BirA or BirA-MSK1 (CA or KD) constructs were co-transfected with H3.3-AviFlag into 293T cells. Total cell lysates were resolved on SDS-PAGE gels. Expression and enzymatic activities of BirA-MSK1 fusions were examined by Western blotting using the indicated antibodies. (B). Acid-extracted histones from transfected 293T cells were resolved on acid-urea gel, followed by Western blot analyses. Blots were probed with α-Flag-antibody to detect total amount of H3.3-AviFlag and Avi-HRP to detect biotinylated H3.3-AviFlag.

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showed that the biotinylated fraction of H3.3 is hyper-shifted on the AU gel, indicating that

they are specifically phosphorylated and/or acetylated in an active MSK1-dependent manner

(Figure 3.3B, bottom). Collectively, these results demonstrate that our approach of coupling

biotinylation and phosphorylation of Avi-tagged H3.3 is feasible, and that purification of

biotinylated H3.3 will also enrich for MSK1-modified H3.3.

3.4.2 Affinity purification of BirA-biotinylated and MSK1-modified nucleosomes

To gain further insight into the links amongst MSK1, histone modifications and

transcriptional activation, we sought to identify other histone modifications that are present

on MSK1-phosphorylated H3.3 and its associated nucleosomes. We preferentially enriched

for MSK1-modified H3.3 nucleosomes by capturing the co-modified (MSK1-phosphoryated

and BirA-biotinylated) H3.3 and its associated nucleosomes onto streptavidin beads. The

general scheme of BICON is shown in Figure 3.4. Briefly, we co-expressed Avi-Flag-tagged

H3.3 and BirA-MSK1 (CA or KD) fusions in 293T cells. Nuclei from these transfected cells

were isolated by hypotonic lysis, followed by digestion with micrococcal nuclease (MNase)

to liberate mononucleosomes. To ensure that we are only analyzing PTMs within the same

H3.3 nucleosome, and not neighbouring ones, complete digestion of chromatin to

mononucleosomes is essential. DNA isolated from input chromatin was analyzed on agarose

gels to ensure that our digest yielded mainly mononucleosomes (~150 bp DNA fragments,

Figure 3.5A). These soluble mononucleosomes were then used as starting material for

affinity pull-down with streptavidin-coupled beads. Following extensive and high stringency

washes, bound nucleosomes were eluted and separated on SDS-PAGE gels. PTMs on these

MSK1-modified H3.3 nucleosomes were then analyzed by Western blotting and

modification-specific antibodies.

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Figure 3.4. Workflow of the affinity purification method used for the isolation of co-modified nucleosomes.

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Figure 3.5. Affinity-purification of BirA-biotinylated and MSK1-modified nucleosomes. (A). DNA was extracted from MNase-digested input chromatin, resolved on a 2% agarose gel and visualized with ethidium bromide. (B). Mononucleosomes containing H3.3-AviFlag were purified as described in (A). Input chromatin (left) and pull-down material (right) were resolved on SDS-PAGE gels and visualized by coomassie blue staining (top) or analyzed by Western blotting using the indicated antibodies (bottom). * streptavidin monomer.

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Analysis of the purified material confirmed that the association of biotinylated H3.3

with streptavidin beads was specific. In the pcDNA vector alone sample, which only

contained non-biotinylated H3.3, we did not detect any H3.3-AviFlag or other histones in the

eluate (Figure 3.5B). As seen by coomassie blue staining, the isolated H3.3 co-purified with

other core histones (H2A, H2B, H4) in an equimolar stoichiometry, indicating the presence

of nucleosomes in the pull-down material (Figure 3.5B, right). In addition, we detected an

association with H2A variant H2A.Z (Figure 3.5B, bottom), which is consistent with the

known occurrence of H3.3/H2A.Z nucleosomes (274). Taken together, these results strongly

suggest that Avi-tagged H3.3 is properly incorporated into chromatin and that the modified

H3.3-containing nucleosomes can be isolated by streptavidin pull-down.

3.4.3 MSK1-phosphorylated nucleosomes are enriched for 14-3-3ζ and H3/H4

acetylation

Using nucleosomes isolated by BICON, we tested which other histone PTMs are

linked to H3.3 phosphorylation by probing these nucleosomes with a panel of histone

modification-specific antibodies. As expected, we detected strong phosphorylation signals

(H3S10ph and H3S28ph) on BirA-CA-MSK1-modified nucleosomes, confirming that they

are hyper-phosphorylated relative to the BirA- or BirA-KD-MSK1-modified nucleosomes

(Figure 3.6A). Due to the well-established roles of histone phosphorylation and acetylation in

transcriptional activation, we examined the acetylation levels of H3.3 and H4 on MSK1-

phosphorylated H3.3 nucleosomes. Using site-specific α-acetyl antibodies and comparing the

nucleosomes that were pulled down from cells transfected with BirA, BirA-CA-MSK1 or

BirA-KD-MSK1, we were able to dissect the specific PTM patterns on MSK1-

phosphorylated nucleosomes. In agreement with our published data (267), we found that

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Figure 3.6. MSK1-modified nucleosomes are enriched for H3 acetylation and H4 acetylation. MSK1-modified nucleosomes were purified as outlined in Fig 3.4. Input and affinity-purified mononucleosomes were resolved on 15% SDS-PAGE and analyzed by Western blotting. (A). Total amount of transfected H3.3 was detected using α-Flag antibody. Endogenous H3 was detected using α-H3 antibody and was used as a loading control. Biotinylation and phosphorylation levels on H3.3-AviFlag were detected using Avi-HRP and phospho-specific antibodies (α-H3S10ph, α-H3S28ph), respectively. (B). Acetylation levels on ectopic H3.3 (left) and endogenous H4 (right) were analyzed using site-specific α-acetyl antibodies.

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H3K27ac and H3K27ac/S28ph levels were significantly higher on BirA-CA-MSK1-modified

nucleosomes (Figure 3.6B, left). We also detected an increase in H3K14ac and H3ac

(H3K9/14ac) levels on these BirA-CA-MSK1-modified nucleosomes. This is consistent with

the earlier finding that H3 S10 phosphorylation is synergistically coupled to H3 K14

acetylation during the activation of IE gene c-fos (170). In addition to analyzing crosstalks

between PTMs on the same H3.3 tail, one advantage of our experimental approach of

isolating mono-nucleosomes is that we can also examine coupling of epigenetic marks on

other histones within the same nucleosome. Indeed, we found that CA-MSK1-modified

nucleosomes have increased H4 acetylation (H4ac) levels, and this increase was dependent

on the kinase activity of MSK1 (Figure 3.6B, right). Using site-specific H4ac antibodies, we

found that an increase in H4K12ac, but not H4K16ac, was associated with phosphorylated

H3.3. These data therefore illustrate that MSK1-mediated H3.3 phosphorylation initiates a

series of PTM crosstalks both in cis on H3.3 and in trans on H4.

Moreover, we observed a preferential binding of 14-3-3ζ to these phosphorylated-

H3.3 nucleosomes (Figure 3.7). To date, only two members of the 14-3-3 family, 14-3-3ε

and 14-3-3ζ, have been identified to bind phosphorylated S10/S28 on H3 (275). Previous

studies on the interaction between 14-3-3 and phosphorylated H3 were based on in vitro

binding assays or involved formaldehyde-crosslinked samples in ChIP assays (174, 226,

227). Therefore, our system not only recapitulates the previously-reported binding of 14-3-3

to H3S10ph and H3S28ph, but it is also the first demonstration of 14-3-3ζ binding to

phospho-H3 in a native in vivo setting. This suggests that BICON may also be useful for

studying the interactions of PTM-binding proteins with their cognate PTMs within a

nucleosome context.

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Figure 3.7. MSK1-modified nucleosomes are enriched for 14-3-3ζ. MSK1-modified nucleosomes and its associated proteins were purified as in Fig 3.4. Association of 14-3-3ζ with biotinylated H3.3 nucleosomes were analyzed by Western blotting using α-14-3-3ζ antibody.

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3.4.4 H4 K12 acetylation is associated with activation of MSK1-target genes

Based on our biochemical pull-down results, we found that a number of H3ac and

H4ac marks were induced by MSK1 and H3 phosphorylation. To validate the biological

relevance of the crosstalk between H3 phosphorylation and H3/H4 acetylation, we next asked

whether any of these acetyl marks are involved in transcriptional activation of MSK1-

responsive genes. We performed ChIP assays using antibodies specific to these marks on two

independent MSK1-target gene systems. First, we made use of our previously-described

NF1-targeting system, whereby we target MSK1 to endogenous NF1-responsive genes by

fusing the DNA-binding domain of NF1 to MSK1 (267). Targeting MSK1 to the polycomb-

silenced α-globin gene through this method reactivated its expression in nonerythroid cells,

and this reactivation was associated with an increase in H3K27ac and H3K27ac/S28ph (267).

As shown in Figure 3.6B, these modifications were also enriched on MSK1-modified H3.3

nucleosomes isolated by BICON, thus confirming the consistency between the biochemical

purification and in vivo ChIP results. We then asked whether H4 acetylation is involved in

this MSK1-mediated reactivation process. Specifically, we used ChIP assays to examine the

levels of H4ac at the α-globin promoter. Consistent with our biochemical findings, we indeed

found that activation of the α-globin gene by NF1-CA-MSK1 targeting is accompanied by

increased H4ac and H4K12ac levels at the promoter of the activated gene (Figure 3.8A, top).

This increase was specific to the re-activated α-globin promoter, since no change in H4ac

and H4K12ac levels was observed in an upstream control region (Figure 3.8A, bottom).

Finally, it is interesting to note that we did not observe any change in H4K16ac level at the

MSK1-activated α-globin promoter. This suggests that the MSK1-H4K12ac link is distinct

from the previously discovered link between PIM1-mediated H3S10 phosphorylation and

H4K16 acetylation (229).

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Figure 3.8. H4K12, but not H4K16, acetylation is induced upon activation of MSK1-target genes. (A). ChIP assays were performed on 293T cells transfected with vector or NF1-MSK1 (CA or KD). H4 acetylation levels on the α-globin promoter (top) and a control upstream region (bottom) were examined using antibodies against H4ac (left), H4K12ac (middle) and H4K16ac (right). The increase in H4ac and H4K12ac is dependent on the kinase activity of MSK1 and correlates with reactivation of the α-globin gene by NF1-CA-MSK1.

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Figure 3.8. H4K12, but not H4K16, acetylation is induced upon activation of MSK1-target genes (cont.) (B). Serum-starved 293T cells were pretreated with DMSO (mock treatment) or H89 (MSK1 inhibitor) for 30 min before stimulation with 400ng/ml TPA. H4 acetylation levels on the c-fos promoter were examined by ChIP assays. TPA-induced H4 acetylation (H4ac, H4K12ac) was greatly compromised upon treatment with kinase inhibitor H89. ChIP-qPCR data are presented as % of precipitated material relative to the amount in the input sample (% input) and have been normalized to H3 levels to adjust for nucleosome density. Error bars indicate means ± SD (n=3).

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As a second test, we examined the activation of IE gene c-fos by phorbol ester 12-O-

tetradecanoyl-phorbol-13-acetate (TPA). Activation of IE genes by growth factors or phorbol

esters is a classic model for studying H3 phosphorylation and gene activation (112, 266).

Upon TPA stimulation, IE genes such as c-fos are strongly induced and this is associated

with an increase in H3 phosphorylation by MSK1 (167, 168, 174). TPA also induced H3

acetylation (H3K9/K14ac) at the regulatory regions of several IE genes, and this increase is

dependent on the kinase activity of MSK1 (174). We therefore used this as a second model

system to further test the biological relevance of our observed link between H3

phosphorylation and H4 acetylation. Similar to results at the α-globin gene, we found an

increase in H4ac and H4K12ac, but not H4K16ac, levels at the c-fos promoter upon TPA

stimulation (Figure 3.8B). More importantly, pre-treatment with H89, which inhibits MSK1

activity, greatly compromised TPA-induced H4 acetylation (H4ac and H4K12ac). This was

especially evident at the earlier times points (15 min and 30 min), suggesting that H4K12

acetylation is downstream of H3 phosphorylation. This reduction in H4 acetylation also

correlated well with the reported reduction in c-fos gene expression in the H89-treated cells

and MSK1/2 knockout mouse fibroblasts (167, 168). As the duration of TPA stimulation

increased, we observed an increase in H4 acetylation, albeit to a lesser extent, in H89-treated

cells. This points to a role for H3 phosphorylation in expediting acetylation at the IE gene

promoter, but an independent mechanism involving acetylation may also be involved as the

stimulation continues (172). All together, these ChIP assays further validated our

biochemical pull-down results and confirmed the biological relevance of our newly identified

trans-tail crosstalk between MSK1 and H4K12 acetylation.

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3.5 Discussion

Here, we describe a new approach, which we termed BICON, to isolate nucleosomes

specifically modified by a histone-modifying enzyme of choice. Importantly, this method

allowed us to study the crosstalks amongst histone PTMs initiated by the enzyme of interest.

In line with our interest in H3 phosphorylation, we have chosen to study the H3 kinase

MSK1 and H3.3 phosphorylation in this pilot study, and found that MSK1 initiates extensive

crosstalks linking histone phosphorylation and acetylation. These data represent the first

demonstration of a histone modifying enzyme being sufficient to initiate a series of PTM

crosstalks amongst the histones within a nucleosome. Increasing evidence shows that

individual histone PTMs do not just function in isolation; it is therefore important to identify

the combinatorial marks that function together and determine how these patterns are

established on chromatin. To our knowledge, this is the first method specifically designed to

identify the “nucleosome-code” initiated by a specific histone-modifying enzyme. This

potentially paves the way for additional studies investigating other histone-modifying

enzymes (such as histone acetyltransferases, methyltransferases, E3 ligases). Expansion of

this method will allow systematic elucidation of crosstalk pathways associated with the ever-

growing list of histone modifying enzymes and their corresponding PTMs.

It has been previously proposed that chromatin can act as a signaling platform that

receives signals from upstream activated signaling cascades and transmits them to

downstream effectors (253, 263). Through histone modifying enzymes and PTMs,

endogenous and exogenous cellular inputs are relayed into the nucleus to alter gene

expression or other nuclear processes. Crosstalks amongst different histone PTMs form part

of the signaling cascade and function to amplify or fine-tune the desired biological response.

This idea is particularly well illustrated by the collective studies on MSK1-mediated H3

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phosphorylation. In response to mitogen or stress stimulation, MSK1 is activated

downstream of ERK or p38 MAPK pathways respectively. MSK1 phosphorylates H3 on S10

and/or S28 at specific target genes and induces their expression. Our findings clearly showed

that MSK1 not only induces H3 phosphorylation, but also initiates a series of downstream

events on chromatin, including the binding of 14-3-3 to MSK1-phosphorylated H3, and other

PTM changes within the nucleosome. Using the BICON approach, we discovered that the

crosstalks not only occur in cis on the same histone (H3.3), but also extend in trans to H4

within the same nucleosome. Therefore, H3 phosphorylation can be seen as a means to

integrate cellular stimuli and signal this information to the transcriptional machinery through

multiple chromatin modifications.

In addition to confirming the previously-established cis crosstalk between H3

phosphorylation and H3 acetylation (H3S10ph - H3K9/K14ac and H3S28ph - H3K27ac)

(170, 267), our co-modification approach also allowed us to identify a novel trans-tail

crosstalk between H3 phosphorylation and H4 acetylation on K12. H4K12ac has mostly been

linked to histone deposition during DNA replication, and its role in transcription has not been

explored in detail. H3 kinases MSK1/2 and H3S28 phosphorylation were recently reported to

play an important role in counteracting polycomb repression in response to environmental

cues and differentiation signals (262). It would be interesting to test if H4K12ac is also

involved in the activation of these polycomb-repressed genes during differentiation and stress

response.

It is also currently unclear how the crosstalk between MSK1-mediated H3

phosphorylation and H4K12 acetylation is established. It is possible that MSK1 physically

associates with the H4K12 acetyltransferase (HAT), and thus both enzymes are recruited to

co-modify nucleosomes on MSK1 target genes. The H4 HAT could also be recruited

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indirectly through phospho-H3 binding proteins, such as through 14-3-3 at the FOSL1

enhancer (229). In either case, identifying the HAT responsible for H4K12ac, as well as its

mode of recruitment to the target genes, would help elucidate the mechanistic details of this

crosstalk pathway. Alternatively, phosphorylated H3 may have a negative effect on the

binding or activity of histone deacetylases (HDAC), leading to an increase in overall histone

acetylation.

In addition to H4K12ac, another crosstalk between H3 phosphorylation and H4

acetylation has previously been described (229). At the FOSL1 enhancer, phosphorylation of

H3S10 is required for acetylation of H4K16 (not H4K12ac), and is associated with

transcriptional elongation. Interestingly, MSK1 phosphorylates H3S10 on the FOSL1

promoter at early time of gene activation, while PIM1 mediates H3S10ph at the FOSL1

enhancer at later time points. The two phosphorylation events lead to distinct downstream

effects such that the crosstalk with H4K16ac only occurs at the enhancer and not the

promoter. These findings suggest that identity of the enzyme, timing and location of PTM all

contribute to determine the final crosstalk and biological outcome. How these different H3

kinases signal to elicit H4 acetylation on different sites is of significant interest.

In this study, we focused on analyzing combinatorial PTMs on nucleosomes, but

BICON can easily be extended to study PTM-binding proteins. To test the feasibility of our

approach to analyze the interaction of PTM-specific binders to chromatin, we examined the

interaction of 14-3-3ζ with phosphorylated H3 in our system. Using our co-modification

approach, we found that 14-3-3ζ preferentially binds to MSK1-phosphorylated. This binding

of 14-3-3 and phosphorylated H3 was first identified in peptide pull-down assays (226).

Their interaction was further characterized by in vitro binding assays and its association with

target genes by ChIP assays (174, 227). We are therefore the first to demonstrate the binding

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of 14-3-3ζ to phosphorylated H3 nucleosomes in non-crosslinked samples in vivo.

Unlike the commonly-used peptide-based assays, a major strength of BICON is that it

allows the identification of PTM-binding proteins under physiological conditions and in a

more relevant chromatin context, thus providing insights into PTM-dependent interactions in

vivo. It is now clear that binding of effectors is sensitive to the presence of PTMs on

neighbouring residues, and multivalent recognition of chromatin marks allows for

cooperative and stronger binding, as illustrated by the binding of NURF chromatin

remodeling complex subunit BPTF (which contains a PHD finger-bromodomain motif) to

H3K4me3/H4K16ac nucleosomes (242, 276). Therefore, further adaption of this technique

can complement existing in vitro binding assays and will be highly useful for identifying

factors that bind nucleosomes through multivalent interactions.

In conclusion, we have successfully established the feasibility of BICON as a method

for probing combinatorial PTMs within a nucleosome, and also revealed potential future

applications of this nucleosome purification system. Our proof-of-principle study with H3

kinase MSK1 and H3 phosphorylation lays the foundation for more extensive examination of

crosstalks between different histone PTMs, and suggests that BICON can be widely

applicable to enzymes involving other key histone PTMs. As such, this new technique could

be an invaluable tool to help advance our understanding of these combinatorial histone marks

and further elucidate their functions in various cellular processes.

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Chapter 4

General Discussion and Future Directions

A portion of Chapter 4.2 and Figure 4.2 are published in a Cell Cycle Features article as: Lau, P.N.I., and Cheung, P. (2011). Unlocking polycomb silencing through histone H3 phosphorylation. Cell Cycle 10, 1514–1515.

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4 Future directions 4.1 Summary of findings

Phosphorylation of H3 on serine residues (S10 and S28) can be induced by multiple

signaling pathways and is associated with transcriptional activation of diverse stimuli-

responsive genes. Work described in this thesis focuses on the downstream events linking H3

kinase MSK1 and H3 phosphorylation to transcriptional activation. As presented in Chapter

2, MSK1 is a direct and potent transcriptional activator. Targeting of MSK1 not only can

activate a typical IE gene, such as c-fos, but can also antagonize polycomb silencing and re-

activate repressed genes such as α-globin. Mechanistically, H3 S28 phosphorylation triggers

acetylation of H3 K27, and these two modifications are both functionally and physically

coupled. Therefore, these studies expand the repertoire of MSK1/phospho-H3-regulated

genes to polycomb-silenced genes, and provide a potential mechanism, involving multiple

histone modifications, for this pathway.

In Chapter 3, I described a novel approach for isolating nucleosomes specifically

modified by a histone modifying enzyme of choice. This proof-of-principle study shows that

MSK1 initiates extensive crosstalks linking histone H3 phosphorylation and acetylation on

H3/H4, and importantly, uncovers a trans-tail crosstalk between H3 phosphorylation and H4

acetylation at K12. This nucleosome isolation method represents the first technique

specifically developed to study crosstalk pathways and combinatorial histone PTMs within a

nucleosome.

Taken together, work described in this thesis clearly illustrates that H3

phosphorylation is involved in crosstalks with other histone PTMs within the nucleosome,

and the resultant combinatorial histone PTMs often function together to regulate gene

expression.

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4.2 Role of H3S28ph and H3K27ac/S28ph during cellular differentiation

My work on MSK1-induced reactivation of α-globin gene suggests that H3S28

phosphorylation can activate polycomb-silenced genes by inducing a methylation-to-

acetylation switch on the adjacent K27 residue. This is the first demonstration that H3

phosphorylation is involved in the activation of polycomb-repressed genes, but further

validation of this model in more physiological settings will be needed. Since PcG proteins

and H3 K27 methylation play critical roles in regulating expression of key developmental

regulators during differentiation and development, I am interested in examining the

involvement of H3S28 phosphorylation and H3K27/S28 phosphoacetylation in a cell

differentiation model system.

To that end, I have chosen to examine the differentiation of F9 mouse embryonal

carcinoma cells by retinoic acid (RA) as a model system. F9 cells undergo differentiation

into primitive endoderm cells upon RA treatment, and this system has previously been used

to study the activation of polycomb-repressed genes including HoxA1, RARβ2 and Cyp26A1

during differentiation (277). Activation of these genes is accompanied by a loss of H3 K27

methylation and PcG proteins at their promoters, demonstrating that their repression in the

undifferentiated state is regulated by the polycomb pathway. Interestingly, a separate paper

showed that RA treatment of mouse embryonic fibroblasts (MEFs) activates MSK1 through

the p38 pathway and induces H3 S10 phosphorylation also at the RARβ2 and Cyp26A genes

(196). Consistent with previous findings, I showed that RA treatment activated the p38

pathway and induced expression of the polycomb-regulated HoxA1 and RARβ2 genes (Figure

4.1A-B). Importantly, RA induced a global increase in H3 S10/S28 phosphorylation and H3

K27 acetylation, suggesting that these modifications play a role in RA-induced F9

differentiation. Moreover, pre-treatment of F9 cells with the MSK1 inhibitor H89

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Figure 4.1. MSK1 and H3 phosphorylation are involved in retinoic acid (RA)-induced gene activation in mouse F9 teratocarcinoma cells. (A). F9 cells were stimulated with 1mM RA and induction of RA target genes was examined by qRT-PCR. RNA isolation, reverse transcription and qPCR were performed as described in Chapter 2.3. (B). Total cell lysates were harvested from mock-treated or RA-stimulated F9 cells, and analyzed by Western blotting. RA treatment of F9 cells activated the p38 MAPK pathway and induced H3 S10 and S28 phosphorylation as well as H3 K27 acetylation. (C). F9 cells were pretreated with vehicle (DMSO) or 10mM H89 (MSK1 inhibitor) for 1 hr prior to stimulation with 1mM RA. Changes in gene expression were measured by qRT-PCR. H89 reduced RA-induced expression of HoxA1 and RARb2 in F9 cells.

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reproducibly reduced expression of HoxA1 and RARβ2, indicating that MSK1 has a role in

regulating these polycomb-repressed genes (Figure 4.1C).

Following up on these observations, detailed ChIP analyses should be performed to

examine the changes in histone PTMs associated with the well-characterized RA-induced

(and also polycomb-regulated) genes HoxA1, RARβ2 and Cyp26A1. Levels of H3S10ph,

H3K14ac, H3K27ac, H3K27me3, H3S28ph as well as the di-modified marks,

H3S10ph/K14ac and H3K27ac/S28ph, should be examined at different time points after RA

stimulation. Such detailed ChIP analyses will reveal how these H3 modifications change

during differentiation and how these changes correlate with one another during the activation

of polycomb-silenced genes. It is particularly important to compare the timing and

distribution of the singly-modified phosphorylation or acetylation marks and the di-modified

phospho-acetyl marks. This will reveal if there are functional differences between the

individual mark and the combinatorial marks, and if they are involved in regulating different

sets of genes or at different times during differentiation. In addition, since H3S10ph has been

linked to transcriptional initiation at promoters as well as transcriptional elongation at a

downstream enhancer (174, 229), comparing the localization of these marks with various

phosphorylated forms of RNAP II will allow one to distinguish their involvement in different

steps of the transcription process.

To complement the targeted examination of candidate RA-induced genes during F9

cell differentiation, ChIP assays coupled to high-throughput sequencing (ChIP-seq) can be

used as a non-biased approach to screen for other genes that are associated with H3S28

phosphorylation and H3K27/S28 phosphoacetylation during differentiation. These ChIP-seq

data will yield important information regarding the locations of these marks in the genome,

and reveal if these marks are enriched at specific genomic regions, such as promoters, coding

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regions, enhancers or other regulatory regions. Cross-referencing the ChIP-seq data with

published gene expression data in differentiated F9 cells (278) will indicate whether these

phospho-marks are associated with actively-transcribed genes or are involved in other steps

of gene regulation, for example, poising genes for activation. Indeed, a recent genome-wide

ChIP-seq study in Drosophila found that H3K27ac/S28ph is present at promoters as well as

enhancers (279). The amount of H3K27ac/S28ph at promoters strongly correlates with

transcription levels, but this mark is present at enhancers prior to activation and its level

increases at active enhancers. The significance of phosphoacetylated H3 at enhancers is

currently unclear, but is suggested to have a role in establishing enhancer-promoter

interactions. Whether this localization at enhancers is conserved in mammalian cells is yet to

be determined. In addition to the F9 cell differentiation model, preliminary studies in the lab

showed that an increase in global levels of H3S10ph, H3S28ph, H3K27ac and

H3K27ac/S28ph were also observed upon RA-induced differentiation of mouse ES cells

(data not shown). Together, these studies suggest that both the F9 and ES cell differentiation

systems are potential cell models for investigating the role of H3 phosphorylation in

regulating the polycomb-silencing pathway.

In addition to H3K27ac/S28ph, a different PTM combination, H3K27me3/S28ph, has

also been associated with activation of polycomb target genes (262). At present, the

relationship between these two distinct H3 modification-combinations has not been clarified.

For example, they may represent different stages along a single mechanistic pathway, or they

may represent distinct populations of modified H3 that regulate different subsets of

polycomb-silenced genes (Figure 4.2). Masking of H3K27me3 by S28ph and the subsequent

displacement of PRC components could be sufficient for rapid and transient activation of

PcG target genes, especially in response to environmental cues; dephosphorylation of the

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Figure 4.2. A model for the activation of polycomb-repressed genes by histone H3 S28 phosphorylation. Upon stimulation by extracellular signals, MSK1/2 (or other kinases) are activated and phosphorylate H3 S28 at polycomb-repressed genes. This generates a H3K27me3/S28ph mark and displaces PcG proteins from the loci. Sequential modifications of this phospho-methyl mark by a histone lysine demethylase (KDM) and histone acetyltransferase (HAT) will generate a different dual modification, H3K27ac/S28ph. We propose that these two H3K27/S28 double marks could exist in parallel and regulate different subsets of polycomb-repressed genes. Dysregulation of these processes could result in aberrant expression of PcG target genes during oncogenesis. [Adapted from (280)]

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H3K27me3/S28ph mark by an H3 phosphatase could promptly re-establish PRC binding and

polycomb silencing. On the other hand, the H3K27ac/S28ph mark could be important for

gene activation during differentiation or development, where additional positive chromatin

modifications (such as H3K27ac) might be required for full activation and sustained

expression of key developmental regulators. Further studies comparing the spatial and

temporal distributions of these combinatorial H3 marks will help distinguish between these

possibilities. Our finding that H3 phosphorylation by MSK1 induces a methyl-acetyl switch

of the adjacent K27 residue suggests that additional H3K27-modifying enzymes are involved

in the activation process. Identifying the H3 demethylase and acetyltransferase responsible

for these modifications as well as their mode of recruitment to target genes would further

elucidate the mechanistic details of this pathway and provide support for the methyl-acetyl

switch model.

4.3 Mechanisms that couple H3 phosphorylation and H3/H4 acetylation

One of the most interesting findings from data presented in this thesis is that H3 S28

phosphorylation is not only functionally and physically coupled to H3 K27 acetylation, but is

also involved in a trans-tail crosstalk between H3 phosphorylation and H4 acetylation on

K12. How these histone PTMs cross regulate one another is currently unclear, but elucidating

the underlying mechanisms will be crucial for understanding their role in transcriptional

activation. One possible mechanism is that MSK1 physically associates with a histone

acetyltransferase (HAT) and thus both enzymes are recruited to the target genes and co-

modify the nucleosomes. To test this possibility, IP-HAT assays should be performed to

determine if an HAT activity associates with MSK1. This involves immunoprecipitation of

MSK1 followed by in vitro HAT assays, using recombinant H3 and H4 as substrates. If an

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HAT activity is detected, identity of the MSK1-associated HAT can be determined using

MS, or alternatively, using antibodies against known H3 HATs (e.g. PCAF, CBP, p300), and

known H4 HATs (e.g. HBO1, MOF, and TIP60).

In support of a physical association between MSK1 and a HAT, MSK1 was

previously shown to co-immunoprecipitate with multiple HATs, including PCAF, p300 and

CBP (174, 261). Importantly, CBP and p300 were reported to acetylate H3 K27 in mouse ES

cells. Our preliminary data also confirmed that overexpression of CBP and p300, but not

PCAF, enhanced global H3K27ac levels in 293T cells (Figure 4.3), suggesting that they

could be candidate HATs responsible for H3K27 acetylation in our system. To extend this

preliminary observation, involvement of CBP and p300 in MSK1-mediated transcription

should be examined. This involves testing the association of CBP/p300 at MSK1-target

genes by ChIP assays, as well as examining the effect of CBP/p300 knockdown on MSK1-

activated transcription and histone acetylation levels at MSK1-target genes. The requirement

for CBP/p300 in the phospho-acetyl crosstalk can be determined by repeating the BICON

pulldowns (as in Figure. 3.4) in CBP/p300 knockdown cells, and examining the H3/H4

acetylation levels on CA-MSK1-modified nucleosomes in the knockdown samples.

To complement this candidate approach, a siRNA screen can be used to identify the

HAT(s) involved in the coupling of H3 phosphorylation and H3/H4 acetylation. A previous

study has generated a library of siRNA oligonucleotides against each of the 17 identified

HATs in mouse, and successfully identified p300 and CBP as the H3K27 HAT in mouse ES

cells (259). Using a similar approach and analyzing the levels of H3K27ac and H4K12ac on

purified BirA-CA-MSK1-modified nucleosomes from these knockdown cells will allow the

identification of HAT(s) involved in the phospho-acetyl crosstalk. Similar knockdown, ChIP

and expression studies (as described above for CBP/p300) can be performed to validate the

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Figure 4.3. Histone acetyltransferases CBP and p300, but not PCAF, acetylate H3 on K27. Flag-tagged CBP, p300 or PCAF constructs were transfected into 293T cells. Total cell lysates were resolved on SDS-PAGE gels, and acetylation level of H3 on K27 was examined by Western blotting. PCAF protein is ~100kD; CBP and p300 proteins are ~300kD.

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importance of the identified HAT(s) in MSK1-mediated transcription.

Another possible mechanism for the coupling of phosphorylation and acetylation is

that phosphorylated H3 has a negative effect on the binding or enzymatic activity of HDACs,

leading to an increase in overall histone acetylation levels. A previous study indeed reported

that the binding of HDAC1 and HDAC2 to the H3 N-terminal peptide (aa1-20) was almost

completely abolished in the presence of phosphorylated S10 (281). Recently, it was also

found that binding of INHAT (inhibitor of acetyltransferases) subunit SET/TAF1β to

methylated H3K27 was disrupted by H3S28 phosphorylation (282). To test the biological

relevance of these in vitro observations, ChIP assays should be used to determine the

association of HDACs and INHAT at MSK1-target genes in the silenced state, and test

whether such binding is disrupted upon gene activation and enhanced H3 phosphorylation. In

addition, the effect of phosphorylation on HDAC activity can be determined by performing

in vitro HDAC assays and comparing the activities of recombinant HDACs on acetylated vs.

phos-acetylated substrates.

As indicated, there is good support from previous findings for both models, and in

fact, both scenarios may work together to functionally link H3 phosphorylation and H3/H4

acetylation. All together, these experiments will help provide mechanistic insights on how

MSK1 and H3 phosphorylation disrupt polycomb silencing and activate gene expression

through histone PTM crosstalk pathways.

4.4 Future applications of BICON

In Chapter 3, I demonstrated that our new approach, BICON, can be used to

specifically isolate mononucleosomes that have been phosphorylated by MSK1. Using

modification-specific antibodies, I analyzed the combinatorial PTMs that are enriched on

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these MSK1-modified nucleosomes. Following up on this initial study, I believe it will be

highly fruitful to combine BICON with proteomics methods to further characterize PTMs

and PTM-binding proteins on these nucleosomes. Mass spectrometry (MS) analysis allows

unbiased identification and quantification of combinatorial PTMs on histones. In this way, a

more comprehensive and systematic analysis of all PTMs on nucleosomes isolated by

BICON can be performed. This unbiased method can potentially identify novel PTM sites

with no available antibodies, and will also avoid any antibody-related issues, such as epitope

occlusion or target specificity (276, 283).

It is currently unclear if phosphorylation of S10 and S28 has differential effects in

regulating gene expression. Previous studies suggest that these modifications occur on

distinct pools of H3 and are present on separate H3 tails at IE gene promoters (174–176).

Interestingly, we detected co-occupancy of H3S10ph and H3S28ph marks at the re-activated

α-globin (but not c-fos) promoter, suggesting that these two marks can coexist within the

same nucleosome, depending on the genes and context (267). It will be important to dissect

the crosstalk pathways initiated by each individual site. The use of H3S10A, S28A or double

S10A/S28A mutants in BICON experiments provides an opportunity for such analysis, and

will help to dissect the functional differences of the two phosphorylation sites. Given that

many of the acetylated residues are in close proximity to the phosphorylated serines, one

concern is that mutating the serine will affect the binding affinity of the acetyl-specific

antibodies. Therefore, especially in this situation, the use of MS to map PTMs on histones

will be critical.

4.4.1 Use of BICON to study other H3 kinases and histone-modifying enzymes

In addition to our newly identified crosstalk between H3 phosphorylation and

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H4K12ac, another crosstalk between H3 phosphorylation and H4 acetylation has been

previously described (229). It is of interest to note that this previously reported crosstalk

involves H3S10ph and H4K16ac (not H4K12ac), and was initiated by a different H3 kinase,

PIM1. This crosstalk occurs at the FOSL1 enhancer and is associated with transcriptional

elongation. This raises the possibility that different H3 kinases can activate different

crosstalk pathways, which may then lead to different downstream functions. Applying the

BICON approach to PIM1 will enable us to identify the PTM signatures on PIM1-modified

nucleosomes. Comparing these PTM combinations with those induced by MSK1 will provide

insights into the functional differences between the two H3 kinases.

In addition to phosphorylation of H3S10 and S28, multiple studies have revealed the

importance of threonine (H3T6, H3T11) and tyrosine (H3Y41) phosphorylation in

transcriptional regulation (115, 199, 284, 285). Many of these phosphorylation events are

mediated by kinases within well-studied signaling pathways. One such example is JAK2,

which phosphorylates H3 at Y41 and prevents HP1 binding at gene promoters (115). These

signaling kinases were thought to regulate gene expression through intermediate substrates,

such as transcription factors and other signaling molecules, but it is now clear that these

kinases can associate with chromatin and modify chromatin directly (286). In the future,

extending the use of BICON to study these histone kinases will uncover potential

mechanisms by which these other phosphorylation sites activate transcription. Indeed, a

recent study illustrated that crosstalks between these phosphorylation sites on H3 can also

occur (287). Using time-resolved, high-resolution NMR spectroscopy, Liokatis et al. showed

that phosphorylation of H3T11 by CHK1 as well as phosphorylation of H3T6 by PKCα are

both inhibited by phosphorylated S10. This phosphorylation crosstalk is uni-directional, since

phosphorylated T6 or T11 had no effect on subsequent phosphorylation of S10 and S28 by

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Aurora B or MSK1. This new approach can provide quantitative analysis of histone

crosstalks, and complement our in vivo identification of histone crosstalks by BICON.

Further extending the use of BICON to investigate other histone modifying enzymes (such as

histone acetyltransferases, methyltransferases, E3 ligases) will allow systematic elucidation

of crosstalk pathways associated with their corresponding PTMs, and will be highly relevant

to the chromatin and epigenetics fields.

4.4.2 Use of BICON to identify histone PTM readers

In this study, I have focused on analyzing combinatorial PTMs on nucleosomes, but

BICON can potentially be extended to study the interaction of PTM-specific binders to

chromatin. PTM-binding proteins were mostly identified using histone tail peptide pull-down

assays, or more recently, recombinant modified mononucleosomes or oligonucleosomal

arrays (288–290). These were all based on the binding of nuclear lysates to chemically-

modified peptides in vitro. Although these techniques have successfully been used to identify

PTM-binding proteins, they are subjected to numerous inherent limitations and restrictions.

Modified peptides only represent part of the histone tails and lack the globular domain of

histones and other components of the nucleosome. However, interactions between PTM-

binding proteins and their cognate sites may require additional binding surfaces in the

nucleosomes. A recent study compared the use of peptides and recombinant modified

oligonucleosome arrays as baits in pull-down assays, and indeed found that a large number of

proteins differ in their association with the two templates (290). In contrast, BICON allows

the identification of PTM-binding proteins under physiological conditions and in a more

relevant chromatin context. It is now clear that multivalent recognition of chromatin marks is

often important for binding of PTM readers to chromatin (243, 265). BICON has an

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advantage over the peptide-based assays in that all the PTMs were laid down by enzymes in

vivo, and endogenous nucleosomes likely contain the combinatorial PTMs necessary for

multivalent binding. Therefore, the use of BICON to identify PTM-binding proteins can

complement existing in vitro binding assays and also provide insights into PTM-dependent

interactions in vivo.

In light of my successful detection of 14-3-3ζ on MSK1-phosphorylated nucleosomes

(Figure 3.7), I have begun to utilize BICON to identify phospho-H3 binding proteins in a

non-biased manner (Figure 4.4A). As shown in Figure 4.4B, a large number of proteins co-

purify with our nucleosome preparations; however, no visually-distinct protein bands can be

seen between the BirA-CA-MSK1 and BirA-KD-MSK1 samples in the silver-stained gel. I

therefore attempted to use a global MS approach instead, and analyze all proteins that co-

purify with the MSK1-modified nucleosomes by MS. Briefly, nucleosome preparations were

separated on SDS-PAGE gels and stained with GelCode Blue. Each gel lane was cut into 10

equal gel slices and proteins therein were subjected to in-gel trypsin digestion. Tryptic

peptides were extracted from the gel slices and analyzed by MS. A preliminary MS run

identified a large number of proteins in our nucleosome preparations. GO (gene ontology)

annotations show that ~75% of the identified proteins are nuclear proteins, and ~45% are

involved in transcription or regulation of gene expression. However, we were not able to

identify proteins that were specifically enriched in the BirA-CA-MSK1 sample. These initial

trials suggest that further optimization of the purification procedure is required. This can

include changing binding conditions or increasing stringencies of the wash buffers to reduce

background binding. Alternative elution methods can also be used to preferentially elute

phospho-binding proteins from streptavidin beads. In the preliminary experiments, all bound

proteins were eluted from beads by boiling sample in 2x SDS sample buffer. One method to

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Figure 4.4. Isolation of effector proteins that bind to MSK1-modified nucleosomes. (A). Workflow of the affinity purification method used for the identification and characterization of effector proteins that bind to MSK1-modified nucleosomes. B, biotinylation; P, phosphorylation; Ac, acetylation; W, X, Y, Z represent phospho-H3 binding proteins.

SDS sample buffer

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Figure 4.4. Isolation of effector proteins that bind to MSK1-modified nucleosomes (cont.) (B). MSK1-modified nucleosomes and associated proteins were purified by the BICON approach. Proteins that co-purify with the biotinylated nucleosomes were analyzed by SDS-PAGE followed by silver staining.

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allow preferential elution is to treat the nucleosome-bound beads with phosphatase. In this

way, only phospho-dependent interactions will be disrupted. Alternatively, excess H3

peptides can be used to compete and elute proteins that directly bind modified H3. Peptides

used will include unmodified H3 peptide (as negative control), H3S10ph, H3S28ph, as well

as the di-modified H3S10ph/K14ac and H3K27ac/S28ph peptides. These alternative elution

procedures could potentially reduce the complexity of the sample and thus facilitate the

identification of phospho-binding proteins. To further improve the sensitivity of the assay,

BICON can be combined with SILAC (stable isotope labeling by amino acids in cell culture)

to allow quantitative MS analysis. SILAC-based MS allows identification of low-abundance

proteins and is particularly useful in distinguishing specific interactors from non-specific

background proteins. Many groups have now successfully used SILAC-based quantitative

MS to identify histone PTM-binding proteins, thus demonstrating the usefulness of this

technique in chromatin research (289, 290). Testing the top MS hits with wildtype H3 and

S10A, S28A, S10A/S28A mutants will help determine which phosphorylation site is

responsible for the interaction. The physiological relevance of these validated phospho-H3

binding proteins can then be tested at IE genes, in our NF1-targeting system (at α–globin

gene) as well as in the differentiation model described in Section 4.2. These follow-up

experiments should include examining their effects on MSK1-mediated transcription in

knockdown studies, and testing their association with MSK1-target genes by ChIP assays.

Currently, 14-3-3 is the only identified transcription-associated phospho-H3 binding

protein, and it binds to both H3S10ph and H3S28ph, albeit with much stronger affinity to

phosphorylated S28. It would be important to determine if there are distinct binding partners

for H3S10ph and H3S28ph, since this would further elucidate the functional differences

between the two H3 phosphorylation sites. Our finding that MSK1 not only phosphorylates

140

H3 at S28, but also enhances K27 acetylation and generates the H3K27ac/S28ph mark, raises

the possibility that such combined epitopes further regulate interactions with PTM-binding

proteins. This is supported by the observation that acetylation of K9/K14 greatly enhances

binding of 14-3-3 to H3S10ph (227, 228). It will be important to test if H3K27 acetylation

affects binding of 14-3-3 or other identified H3S28ph-binding proteins. Alternatively, H3S28

phosphorylation can enhance or disrupt binding of bromodomain-containing proteins that

recognize the H3K27ac mark, or H3K27ac/S28ph may create a new binding motif for an

effector that does not recognize either individual mark. Histone modifying enzymes and

chromatin remodeling factors are often recruited directly to chromatin through the

acetyl/methyl-binding domains present on these proteins. Whether phospho-binding modules

are present on these chromatin modifying enzymes is not clear. It is also possible that

phosphorylated H3 mainly mediates its downstream functions through 14-3-3 or other

adaptor proteins, and thus indirectly recruit these enzymatic activities to chromatin.

Nevertheless, identification of proteins that interact with H3S10ph, H3S28ph or the di-

modified phosphoacetylated marks will be instrumental in uncovering the mechanism of

action for these marks, and would provide a greater understanding of H3phosphorylation’s

role in the transcription process.

Taken together, the proof-of-principle study with MSK1 and H3 phosphorylation

described here lays the foundation for an extensive examination of crosstalks between

different PTMs as well as the identification of downstream effectors. This approach can be

widely applicable to other histone modifying enzymes and PTMs, and would help advance

our understanding of these combinatorial histone marks and further elucidate their functions

in various cellular processes.

141

4.5 Concluding remarks

In the past few years, functional crosstalks between histone PTMs and combinatorial

readout of histone marks have been areas of intense research in the chromatin field. My work

described in this thesis clearly indicates the involvement of H3 phosphorylation in PTM

crosstalk pathways and their role in transcriptional regulation. Our development of the

BICON method will further facilitate research in the areas of combinatorial histone PTMs

and PTM crosstalk regulation.

Given that aberrant expression of PcG target genes are common in cancers, further

investigating the link between H3 phosphorylation and activation of PcG genes would be

essential. In fact, constitutive activation of MSK1 and elevated levels of H3S10ph &

H3S28ph have been reported in Ras-transformed cells as well as primary acute myeloid

leukemia samples. Whether this hyperphosphorylation of H3 has a role in the deregulation of

PcG targets during oncogenesis or other diseases would be an important area to explore.

142

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