THE CHROMATIN REMODELING FACTOR CHD1tk349zq2724/SniderDissertation... · Joseph Lipsick Approved...

THE CHROMATIN REMODELING FACTOR CHD1L IN THE

PREIMPLANTATION EMBRYO AND IN ES CELLS

A DISSERTATION

SUBMITTED TO THE DEPARTMENT OF GENETICS

AND THE COMMITTEE ON GRADUATE STUDIES

OF STANFORD UNIVERSITY

FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

Alyssa Christine Snider

August 2010

http://creativecommons.org/licenses/by-nc/3.0/us/

This dissertation is online at: http://purl.stanford.edu/tk349zq2724

© 2010 by Alyssa Christine Snider. All Rights Reserved.

Re-distributed by Stanford University under license with the author.

This work is licensed under a Creative Commons Attribution-Noncommercial 3.0 United States License.

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http://purl.stanford.edu/tk349zq2724

I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.

Matthew Scott, Primary Adviser


Joanna Wysocka, Co-Adviser


Gerald Crabtree


Margaret Fuller


Joseph Lipsick

Approved for the Stanford University Committee on Graduate Studies.

Patricia J. Gumport, Vice Provost Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file inUniversity Archives.

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ABSTRACT

Early embryonic cell types such as the zygote, blastomeres of the preimplantation

embryo, and embryonic stem (ES) cells have powerful chromatin remodeling activities

that facilitate DNA-dependent processes such as transcription and DNA repair. These

chromatin-regulated processes are crucial for enacting complex gene expression

programs and ensuring genomic integrity for the developing embryo. Improving our

basic knowledge of chromatin remodeling in the preimplantation embryo and in

embryonic stem cells has implications for addressing human infertility and regenerative

medicine.

Chd1l encodes a chromatin remodeling factor and was highlighted as a candidate

developmental regulator from a screen in which factors were identified whose transcripts

are more highly expressed in the isolated inner cell mass (ICM) compared to the whole

blastocyst. Chd1l expression is developmentally regulated during a time course of

preimplantation development, peaking at the late morula stage, just prior to the formation

of the blastocyst. In addition, Chd1l is expressed in ES cells. Prior to this dissertation

research, the role of Chd1l had not been addressed, and its intriguing expression patterns

suggested Chd1l could be a novel regulator of DNA-dependent processes in early

developmental cell types. This dissertation describes research undertaken to address the

role of Chd1l in chromatin remodeling in the preimplantation embryo and in ES cells.

Four questions were addressed: 1) Is Chd1l essential in ES cells? 2) Does Chd1l

regulate gene expression in ES cells? 3) Is Chd1l essential in the preimplantation

embryo? 4) Does Chd1l contribute to the DNA damage response in ES cells or in the

preimplantation embryo?

To address the first question, Chd1l was knocked-down in mouse ES cells using a

shRNA targeting the Chd1l transcript. Reducing Chd1l protein to nearly undetectable

levels reveals that Chd1l is dispensable for ES cell viability, proliferation, and pluripotent

morphology. The second question was addressed by subjecting ES cells in which Chd1l

had been knock-down to genome-wide expression analysis. This study demonstrated that

global gene expression patterns were unaltered by Chd1l knock-down, confirming that

Chd1l is dispensable for transcription in general and, in particular, for maintaining

v

pluripotent transcriptional network. Chd1l is also dispensable for gene expression

programs associated with the formation of the primary germ layers, as differentiating

embryoid bodies demonstrate temporally appropriate repression of pluripotency markers

and activation of germ layer lineage markers.

To address whether Chd1l is essential in the preimplantation embryo, mouse

embryos were micro-injected at the single-cell stage (zygote stage) with antisense

morpholino (MO) oligos targeting the Chd1l transcript. Development was observed in

vitro for four days, during which time control embryos progressed to the blastocyst stage.

Embryos injected with Chd1l-MO arrested prior to the multi-cell stage, indicating that

Chd1l plays a crucial role during preimplantation embryogenesis. Knock-down by the

MO was confirmed at the transcript levels by microfluidic qPCR, and the arrest

phenotype was confirmed to be due to Chd1l deficiency by partial rescue upon co-

injection of Chd1l mRNA and Chd1l-MO.

During the course of this study, evidence from independent research groups

identified a role for Chd1l in the DNA damage response pathway in somatic cultured

cells. ES cells and cells of the early embryo are known to have stringent and unique

pathways to repair DNA damage to prevent mutation and genomic instability from

arising in the organism. To address whether Chd1l participates in the DNA damage

response in ES cells, ES cells in which Chd1l had been knocked-down were treated with

DNA damaging agents and assayed for survival rates. In direct contrast to the published

literature, in which reduction of Chd1l in somatic cells induced hypersensitivity to

induced DNA damage, reduction of Chd1l in ES cells conferred resistance to induced

DNA damage. The most likely explanation for this is that Chd1l participates in a

damage-induced apoptotic response in ES cells. This is supported by data showing that

Chd1l over-expression is sufficient to induce apoptosis in ES cells.

Interestingly, apoptosis induced by over-expression of Chd1l occurs specifically

in ES cells, as ES cells differentiated for as little as two days by removal of the cytokine

LIF no longer undergo Chd1l-induced apoptosis. This switch in the effect of Chd1l over-

expression during differentiation suggests that Chd1l responds to DNA damage very

differently in ES cells than in differentiated somatic cells. Indeed, ES cells are

significantly more proficient in initiating apoptosis to eliminate DNA damage than their

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differentiated counterparts. A model in which Chd1l responds to DNA damage by

initiating apoptosis in ES cell but not in differentiated cells could explain the conflicting

results presented in this dissertation and in other published studies that used differentiated

cells.

A likely role for Chd1l in DNA repair in the preimplantation embryo could

explain the Chd1l arrest phenotype. DNA repair is particularly critical in the zygote to

repair the paternal genome, and inefficient DNA repair leads to decreased fertility in

humans. The zygote relies heavily on non-homologous end joining (NHEJ) to repair

double-stranded breaks (DSBs). Reliance on NHEJ is in common with differentiated

cells but not ES cells, which primarily use homologous recombination (HR) to repair

DSBs. Biochemical evidence supports the involvement of Chd1l in NHEJ as it interacts

with NHEJ-specific proteins but not HR-specific proteins. Modest increases in staining

for a marker of DSBs can be seen in embryos injected with Chd1l-MO, indicating that

deficient DNA repair could underlie the Chd1l arrest phenotype.

In summary, this dissertation describes an essential role for Chd1l in the

preimplantation embryo that could be due to defects in DNA repair. In contrast, Chd1l is

dispensable for ES cell gene expression, pluripotency, and differentiation. Chd1l likely

contributes to an ES cell-specific apoptotic response to DNA damage. It is proposed here

that Chd1l functions through the NHEJ pathway, a pathway critical in the zygote and in

differentiated cells, but not in ES cells. Therefore, Chd1l functions minimally, or not at

all, in regulating gene expression and contributes to the DNA damage response in a

developmental stage-specific and/or cell type-specific manner.

vii

PREFACE

The first decade of the twenty-first century during which I began graduate school

was an extraordinary time of revolutionary advancements in genetics, epigenetics, and

biotechnology. The Human Genome Project reported the first draft of the sequenced

human genome in 2001 [1,2] and its completion in 2007, sparking a burst in activity in

the field of genetics. Advancements in high throughput technology led to the so-called

“genomics era” and have culminated in the advent of “personal genomics” where

individuals can have their genomes sequenced or affordably SNP-genotyped by

companies such as 23andMe and Navigenics. Dolly the sheep, the first mammal cloned

through the reprogramming of an adult somatic cell, was born in 1996, undermining the

dogma that differentiation is an irreversible phenomenon and sparking interest in the field

of epigenetics [3]. Epigenetic research has transitioned in parallel with genetic research

to large-scale, genome-wide studies that have lead to great advancements in our

knowledge of the “epigenome” and its surprising plasticity. The public watched in

dismay as Dolly passed away in 2003, showing signs of premature aging and raising

questions about the molecular basis of “youth” and pluripotency. In 2007, when bans on

federal funding of human ES cells were still in effect under the Bush administration, two

groups transformed adult human cells into “induced pluripotent stem” (iPS) cells by the

introduction of only a few factors [4,5]. The ability to avoid the controversial use of

embryos to generate pluripotent stem cells, and the ability to use adult cell types instead,

revolutionized the fields of personal and regenerative medicine.

It was in the midst of this climate of excitement, undiminished by the clear skies

and temperate weather in Palo Alto, California, that I began my studies in the department

of Genetics at Stanford University. I was enticed to join Dr. Matthew Scott’s laboratory

by his attitude of scientific open-mindedness and by a chromatin project that was

underway. The laboratory was located in the Clark Center, built in 2003 for the purpose

of housing the multi-disciplinary Bio-X Department. With developmental biologists,

statisticians, bioengineers, computer scientists, and neurobiologists as neighbors, and a

Pete’s Coffee shop that was strategically situated in the Clark Center to promote

scientific discussions, I pursued the most difficult academic challenge of my life.

viii

Dr. Tian Wang, now a professor at the University of Chicago, Illinois, was a post-

doc in Dr. Scott’s laboratory who was interested in the property of pluripotency and how

it was established and maintained in embryonic stem (ES) cells. She conducted a screen

to identify candidate pluripotency regulatory factors by asking which transcripts are

enriched in the inner cell mass (ICM), the part of the embryo that is pluripotent and from

which ES cells are derived. We became interested in the subset of these ICM-enriched

factors that were involved in regulating chromatin dynamics. Among the genes whose

transcripts were enriched in the ICM was Chd1l (Chromodomain, ATPase/Helicase, DNA

binding 1-like), a chromatin remodeling factor that is part of a very important protein

family with diverse developmental roles. At the inception of the project, no

developmental or molecular studies had been reported on Chd1l, making the gene an

intriguing subject of study to me.

This dissertation is an account of research undertaken to determine the role of

Chd1l in the preimplantation embryo and in embryonic stem cells. The first introductory

chapter will focus on chromatin remodeling in these systems. Because Chd1l is a

member of the SNF2 family, members of which have well established roles in gene

expression, and because Chd1l has more recently been shown to have a role in the

response to DNA damage, the introduction will describe chromatin remodeling activities

that pertain in particular to gene regulation and DNA repair. The role of Chd1l was

studied in ES cells and, in collaboration with the laboratory of Dr. Mylene Yao, in the

preimplantation embryo. Chapter 2 documents the results of these studies, revealing that

Chd1l is essential in the preimplantation embryo but dispensable in ES cells. Along with

Dr. Wang, my Yao laboratory collaborators, my advisor and co-advisor, I will submit

Chapter 2 as a manuscript for publication to PlosOne. Chapters 3 and 4 describe my

investigations into the molecular function of Chd1l that could reveal hidden phenotypes

in ES cells or explain the arrest phenotype in preimplantation embryos. Chapter 5

discusses the implications of these findings and the directions in which I see the field

progressing.

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ACKNOWLEDGEMENTS

This thesis project is the result of the efforts of many people at Stanford,

beginning with my advisor Dr. Matthew Scott, who always encouraged living a fulfilling,

balanced life. In particular, I would like to express my regards to my two “lab moms,”

our laboratory administrator Diane Bush, and our research associate Kaye Suyama,

without whom I may never have graduated. My experience would have been very

different without the humor, intelligence, and kindness of the graduate students beside

whom I worked, Monique Barakat, Manuel Lopez, and Tyler Hillman, and three special

colleagues, Dariya Glaser, Fraser Tan, and Dr. Timothy Reddy. It was a pleasure to work

with my collaborators in Dr. Mylene Yao’s laboratory, especially my micro-injectionist

Denise Leong. I am endlessly grateful to my co-advisor Dr. Joanna Wysocka and her

husband and collaborator Dr. Tomek Swigut for lending their genius toward my project.

I am also grateful to the rest of my committee members, Dr. Jerry Crabtree, Dr. Margaret

Fuller, Dr. Julie Baker (on my committee until the final three months), and Dr. Joseph

Lipsick. I am honored to have known these remarkable people.

I would like to thank my family for their support during the challenging six years

of graduate school: my father, whose passion for academia first inspired me to pursue

higher education; my mother, whose unconditional love carried me through difficult

times of self-perceived failure; my older sister, who encouraged me to trail-blaze my own

path; my younger sister, who reminded me to stay true to myself; and my husband, who

gave me a new reason for living and excelling.

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DEDICATION

This dissertation is dedicated to my father, Dr. Mervin G. Wright

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CONTENTS

1. Chromatin Remodeling in Preimplantation Embryos and

Embryonic Stem Cells ............................................................................... 1

Organization of the Mammalian Genome ...................................................................... 1

Chromatin Regulatory Proteins ...................................................................................... 2

DNA methyltransferases .............................................................................................. 2

Post-translational modifications .................................................................................. 3

ATP-dependent chromatin remodeling factors ............................................................ 6

Epigenetic Pluripotency in ES Cells ............................................................................... 8

DNA Repair in ES cells .................................................................................................. 9

Chromatin Remodeling in the Zygote and Cleavage-Stage Embryo ............................ 13

Chromatin states of the gametes ................................................................................ 13

Chromatin remodeling in the zygote ......................................................................... 14

Maternal/zygotic transition ........................................................................................ 15

Chromatin Remodeling in the Blastocyst ..................................................................... 16

Differentiation of the ICM ......................................................................................... 16

Epigenetic differences between the TE and ICM ...................................................... 18

X chromosome inactivation ....................................................................................... 19

Derivation of ES cells from the ICM ......................................................................... 20

DNA Repair in the Zygote ............................................................................................ 22

Chd1l as an Oncogene and DNA Damage Response Protein ....................................... 24

Summary ....................................................................................................................... 26

2. The Chromatin Remodeling Factor Chd1l Is Required in

the Preimplantation Embryo ..................................................................33

Abstract ......................................................................................................................... 34

Introduction ................................................................................................................... 34

Results ........................................................................................................................... 37

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Chromatin factors are compartmentalized in the blastocyst ...................................... 37

Chd1l expression patterns suggest a developmental role .......................................... 40

Chd1l is dispensable for ES cell pluripotency and proliferation ............................... 40

Chd1l does not regulate gene expression in ES cells ................................................ 41

Chd1l is not required for differentiation of ES cells .................................................. 42

Chd1l transcripts are abrogated in MO-injected embryos ......................................... 42

Embryos injected with Chd1l-targeting MOs arrest prior to blastocyst

stage ...................................................................................................................... 43

Chd1l phenotype is partially rescued by co-injection of Chd1l mRNA .................... 44

Discussion ..................................................................................................................... 44

Methods ........................................................................................................................ 46

3. Finding Direct Transcriptional Targets of Chd1l in ES

Cells ...........................................................................................................57

Introduction ................................................................................................................... 57

Results ........................................................................................................................... 59

Chromatin Immunoprecipitation ............................................................................... 59

Sequencing and Analysis ........................................................................................... 60

Re-analysis using deeper sequencing ........................................................................ 61

HA-tagged Chd1l targeting vector ............................................................................. 62

Discussion ..................................................................................................................... 62

Methods ........................................................................................................................ 64

Contributing Collaborators ........................................................................................... 66

4. Molecular Functions of Chd1l in Early Development .........................74

Introduction ................................................................................................................... 74

Results ........................................................................................................................... 78

Reduction of Chd1l increases DNA damage tolerance in ES cells ........................... 78

Over-expression of Chd1l kills ES cells but not differentiated cells ......................... 80

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Chd1l forms a ~500 kD complex in ES cells ............................................................ 83

γ-H2AX marks uninjected embryos and embryos injected with Chd1l-

MO ........................................................................................................................ 83

PAR marks uninjected embryos and embryos injected with Chd1l-

MO ........................................................................................................................ 85

Discussion ..................................................................................................................... 86

Methods ........................................................................................................................ 90

5. General Discussion ................................................................................105

Chd1l is Essential in the Preimplantation Embryo ..................................................... 105

Chd1l is Non-Essential in ES cells ............................................................................. 108

Role of Chd1l in DNA Repair in ES cells .................................................................. 110

Oncogenic Potential of Chd1l ..................................................................................... 112

Paradoxes in Chd1l Function in ES Cells ................................................................... 114

REFERENCES ................................................................................................117

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LIST OF FIGURES

Figure 1.1 PAR modifies nuclear proteins............................................................... 28

Figure 1.2 ATP-dependent chromatin remodeling ................................................. 29

Figure 1.3 SNF2 family of chromatin remodeling factors ..................................... 30

Figure 1.4 Chromatin remodeling in the preimplantation embryo ...................... 31

Figure 1.5 Chd1l and the PARP-dependent DNA damage response .................... 32

Figure 2.1 Chromatin remodeling factors are enriched in the ICM ..................... 50

Figure 2.2. Chd1l is a candidate developmental regulator ...................................... 51

Figure 2.3. Chd1l is non-essential in ES cells ............................................................ 53

Figure 2.4. Chd1l MO knock-down .......................................................................... 54

Figure 2.5. Chd1l embryonic arrest phenotype ........................................................ 56

Figure 3.1 Preliminary validations for ChIP ........................................................... 67

Figure 3.2 Distribution of reads around TSS .......................................................... 68

Figure 3.3 Validation of Chd1l ChIP peaks ............................................................ 71

Figure 3.4. Re-analysis of two combined Chd1l ChIP sequencing runs ............... 72

Figure 3.5. Chd1l-HA targeting vector .................................................................... 73

Figure 4.1. Sensitivity to DNA damage in ES cells expressing Chd1l-shRNA ..... 93

Figure 4.2. Effects of over-expression of K71R or WT Chd1l. .............................. 95

Figure 4.3. Chd1l over-expression specifically kills undifferentiated ES cells ..... 96

Figure 4.4. Size Fractionation of a Chd1l-containing protein complex ................ 98

Figure 4.5. γH2AX staining in embryos injected with Chd1l-MO. ..................... 101

Figure 4.6. PAR staining in embryos injected with Chd1l-MO. .......................... 104

Figure 5.1 Chd1l Phenotypes .................................................................................. 116

LIST OF TABLES

Table 3.1 Number of reads obtained for ChIP libraries ........................................ 68

Table 3.2. Number of peaks called for ChIP samples ............................................. 69

Table 3.3. Number of peaks called for the combined analysis ............................... 69

1

1. Chromatin Remodeling in Preimplantation Embryos and

Embryonic Stem Cells

Organization of the Mammalian Genome

Within the diploid human cell nucleus is approximately 6 billion base pairs of

DNA divided into 46 macromolecules called chromosomes [1]. Between them, they

contain a little over 20,000 genes that encode the proteins necessary for life. Some of

these proteins, in turn, handle DNA-related processes such as transcription, DNA

replication, and DNA repair. Elaborate and diverse transcriptional programs establish the

identity and functional specificity of tens of trillions of cells that comprise the adult body.

Housekeeping genes are constitutively expressed in all cell types, whereas genes

encoding proteins with highly specific functions must be repressed in one cell type but

active in another. Cell type specific gene expression patterns are maintained

epigenetically by modification of DNA, histones, and other proteins that interact with

DNA. During cell division, DNA replicates and condenses into discrete mitotic

chromosome structures, and sister chromatids are physically tethered and distributed to

daughter cells. Chromosomes are huge molecules, the smallest having a molecular

weight of about 15 billion g/mol and the largest 80 billion g/mol, compared to a molecule

of water whose molecular weight is 18 g/ml. Each nucleic acid is comprised of many

chemical bonds that are subject to spontaneous hydrolysis, oxidation, and alkylation, and

it is estimated that a single cell experiences tens of thousands of lesions due to

endogenous sources every day [6]. Because of this, chromosomes must be constantly

monitored and repaired by specialized DNA repair machinery.

If stretched end to end, the size of the genome is 2 meters long, about 340,000

times the diameter an average nucleus (6 µm). DNA must therefore be organized by

several degrees of complexity to fit into the nucleus. Histone octamers (two of each

H2A, H2B, H3 and H4) function like spools, wrapping DNA around themselves like

thread and forming nucleosomal arrays, the primary structure of chromatin [7].

Micrococcal nuclease digestion reveals that 146 base pairs of DNA is wrapped around

each histone octamer, and each core nucleosome is separated by ~100 base pairs of linker

2

DNA [8,9]. The presence of the histone H1 within the linker sequence facilitates

formation of secondary chromatin structure, the 30 nm fiber [10]. The crystal structure of

the core nucleosome was solved in 1997 and remains, in this author’s biased opinion, one

of the most beautiful crystal structures to date [11].

But such highly ordered packaging of the genome presents a significant challenge

for proteins that need access to DNA [12]. If DNA is rendered inaccessible, processes

necessary for viability including replication, transcription, and DNA surveillance and

repair would be fatally impaired [12,13,14]. Fortunately, nucleosomes and higher-

ordered chromatin are dynamic structures [15], and a host of chromatin regulatory

proteins are dedicated to remodeling chromatin structure to facilitate chromatin-

dependent processes. The focus of this introductory chapter is on chromatin remodeling

in the pluripotent cells of preimplantation embryo and in embryonic stem (ES) cells, cells

that contain the some of the most potent chromatin remodeling activities, except perhaps

for primordial germ cells [16]. Here chromatin remodeling is broadly defined as any

alteration of chromatin structure for the purpose of regulating gene expression or other

chromatin-dependent processes, such as DNA repair.

Chromatin Regulatory Proteins

Chromatin regulatory proteins can be categorized into three broad classes: (1)

enzymes that regulate methylation on DNA directly, (2) enzymes that add post-

translational modifications onto histone tails, and (3) enzymes that harness the power of

ATP to force torsional changes in chromatin, catalyze nucleosome sliding, and facilitate

histone eviction or deposition [17,18].

DNA methyltransferases

The enzymes that methylate CpG islands in the genome are called DNA

methyltransferases, or DNMTs, and their activity usually opposes transcription [19,20].

DNA methylation is required for allele-specific expression of imprinted genes [21], X

chromosome inactivation in females [22,23], the silencing of transposable elements

[24,25], and is associated with differentiation in somatic cells [20]. However, the

simplistic view that DNA methylation always leads to chromatin compaction and gene

3

silencing has more recently been under scrutiny, and a more complex scenario probably

functions in vivo [26]. The essential DNMT1 maintains CpG methylation through cell

division by methylating the 5th

carbon on cytosine nucleotides in the newly synthesized

strand of hemi-methylated DNA. The DNMTs 3a and 3b are responsible for de novo

DNA methylation, particularly during development [27]. A mechanism of active DNA

demethylation has been proposed, but the enzyme responsible for this activity has not

been identified in mammals.

Post-translational modifications

Histones contain flexible C-terminal tails that are subject to a variety of post-

translational modifications (PTMs) including methylation, acetylation, PARylation,

phosphorylation, and ubiquitinilation. Because PTMs can modulate differential gene

expression patterns, even in genetically identical cells, a “histone code” has been

proposed [28]. It is becoming increasingly clear that PTMs function in combination with

each other, and there is not always a direct relationship between one PTM and gene

expression [29]. Although PTMs are typically included when discussing “epigenetics,” it

is unresolved how PTMs are inherited through cell divisions. This is in contrast with

DNA methylation, which can be propagated during replication by the activity of the

maintenance DNMT1 that recognizes hemimethylated DNA.

A variety of enzymes exist that post-translationally modify specific residues in

histone tails. The enzymes that catalyze histone acetylation are called histone

acetyltransferases, or HATs, and the enzymes that remove acetyl groups are the histone

deacetylases, or HDACs. Histone acetylation on the promoters of genes is associated

with transcriptional activity. Histone methyltransferases (KMTs) are the enzymes that

add methyl groups to histone tails. This PTM can be associated with either

transcriptional activation or repression, depending on which lysine residue is modified.

For example, methylation of lysine 4 on histone 3 (H3K4me) is associated with gene

expression, and trimethylation of H3K27me3 is strongly associated with gene repression.

It was long expected that histone methylation would be a reversible modification similar

to histone acetylation, but the first histone demethylase (KDM), LSD1, now called

4

KDM1a, was only recently discovered [30]. Since then, a large number of jumonji-

domain containing proteins have been shown to have histone KDM activity [31].

Poly(ADP-ribosyl)ation, or PARylation, is a PTM that covalently attaches

negatively charged polymers of ADP-ribose onto histones and other nuclear proteins that

interact with DNA (Fig. 1) [32]. The PARP family of polymerases contains 18 members,

and most, if not all, can synthesize long linear or branching chains of up to 200 ADP-

ribose moieties using NAD+

as a substrate [32,33]. The primary acceptor protein is

PARP-1 itself, which, when activated, PARylates itself in an automodification reaction

[32]. The presence of PAR is extremely transient in nature, with a half-life of less than 1

minute due to the abundance of a single glycohydrolase enzyme, PARG, which rapidly

cleaves PAR polymers endo- and exo-glycositically, releasing free ADP-ribose units

[34,35]. The role of PAR in the cell is diverse and depends on the PARP family member

involved. PARP-1 and PARP-2 are the most studied and best understood PARPs, having

traditional roles in DNA damage response and repair [34,36,37,38,39]. One PARP,

PARP3, localizes to centrosomes and its over-expression interferes with progression

through the G1/S phase transition and may participate in DNA damage surveillance [40].

Another PARP, Tankyrase1 (Tank1), associates with TRF1 and other telomeric proteins

and is involved in telomere homeostasis [41]. Long term over-expression of Tank1

induces telomere elongation [42]. On the other hand, over-expression of another

tankyrase, Tank2, that also associates with TRF1 induces rapid and widespread PARP-

dependent cell death [43].

It has long been known that PARylation can contribute to chromatin relaxation

[44,45,46]. PARylation of the major histone acceptor H1 by PARP-1 in biochemical

assays induces relaxation of the 30 nm fiber, in a manner highly similar to chromatin

relaxation in the absence of histone H1, and is proposed to increase DNA accessibility

[46]. It was later shown that PARP-1 activity could contribute to either chromatin

relaxation or compaction, depending on the context and NAD +

supplies [47]. More

recently, the involvement of PARP-1 in regulating transcription has become apparent.

Like histone H1, PARP-1 binds to nucleosomes. In in vitro assays without NAD+,

PARP-1 incorporates into chromatin, decreases micrococcal nuclease sensitivity, and

represses reporter gene expression. When NAD+ is introduced, chromatin relaxes,

5

PARP-1 is displaced from nucleosomes, and reporter gene transcription is re-activated

[48]. In vivo ChIP-chip studies have shown that the presence of PARP-1 and absence of

histone H1 at promoters of genes is associated with gene activation [49].

Automodification of PARP-1 is the dominant mode of PARylation and, because it can

regulate gene regulation, it may also be a part of the “histone code” [45,50].

The longest known role of PARP-1 is its ability to respond to DNA damage and

facilitate repair [34]. The specific activity of PARP-1 is amplified 500-fold by the

presence of single strand breaks or double strand breaks in chromatin [32]. Its activation

leads to PARP-1 dimerization, automodification, and DNA binding, and to chromatin

relaxation. PARylated PARP-1 then recruits components of the base excision repair

(BER) pathway, including XRCC1 [51]. Parp-1 knock-out mice show hypersensitivity to

γ-irradiation and treatment with DNA damaging agents MMS, MNU, and MNNG at the

whole-animal level [38,52,53] and defects in DNA repair at the cellular level [54]. Even

in these mice, some residual PAR activity was observed, and this was found to be due to

a highly similar PARP-2 protein [55,56]. Parp-2 knock-out mice also show

hypersensitivity to γ-irradiation and DNA damage induced by alkylating agents [57].

PARP-1 and PARP-2 have partially redundant roles, as double knock-out mice of Parp-1

and Parp-2 display embryonic lethality at E7.5 [57]. However, specific roles for PARP-2

are becoming elucidated, such as binding to a telomeric protein TRF2 [58], and

preferential heteromodification of histone H2B [59], as opposed to preferential

heteromodification of histone H1 by PARP-1 [46].

The macro domain is a module that binds PAR. Macro domain-containing

proteins can be found in eukaryotes, bacteria, and archaeans, but include only a small

mammalian family [60,61]. Several PARPs, PARP-14, PARP-15, and PARP-9 (bal, the

lymphoma risk factor protein), contain a macro domain, and although it is tempting to

propose feedback signaling loops, these PARPs have not been confirmed to bind PAR

and may not because of divergent sequence [33]. The H2A histone variant macroH2A is

a component of chromatin in the inactive X chromosome [62] and its ability to bind PAR

has been shown [50]. Upon PARP-1 activation, macroH2A facilitates chromatin

compaction and hinders the recruitment of DNA repair proteins. These studies

underscore the conclusion that PAR has diverse roles in the cell that can facilitate

6

heterochromatin or euchromatin formation, depending on the context in which it is

synthesized and recognized.

In addition to its cytoprotective role, PARP-1 can mediate an alternative response

to DNA damage—apoptosis. A cell has two choices when confronted by DNA damage:

to repair the damage or to initiate programmed cell death. Surprisingly, Parp-1 knock-

out mice also show increased resistance to ischemia-reperfusion injury, inflammation-

related injury [63], diabetes [64], and hyperoxic damage. Under certain conditions,

PARP-1 appears to initiate apoptosis through the caspase-independent AIF-mediated

pathway [65]. AIF is normally sequestered in the mitochondrion, but upon PARP-1

activation, AIF translocates to the nucleus where it induces widespread chromatin

condensation and fragmentation. Translocation of AIF to the nucleus is suppressed by

PARP inhibitors, and PARP-dependent cell death can be suppressed by microinjection of

inhibitory AIF antibodies, demonstrating co-dependence. The mechanism by which

PARP activation in the nucleus triggers AIF release from the mitochondrion remains

mysterious [65].

ATP-dependent chromatin remodeling factors

The third class of chromatin regulatory enzymes is the ATP-dependent chromatin

remodeling factors (CRFs) that assemble into large, multisubunit complexes. These

CRFs utilize ATP to drive conformational changes in chromatin, and the outcome of their

activity depends on what other factors associate with them (Fig. 2) [66,67]. A single

CRF can have diverse functions due to the combinatorial nature of complex assembly; a

single subunit switch can alter or reverse its function [68]. This can happen between cell

types where a subunit is expressed in one cell type but repressed in another, or even in the

same cell where assembly is context dependent [69,70,71,72].

The best-known chromatin remodeling complex is the BAF complex, which

contains the prototypical mammalian Brm homologs, Brg1 and Brm. These chromatin

remodeling factors define the SNF2 family of chromatin remodeling factors by the

presence of the split DNA-dependent helicase domain. Rather than unwinding DNA like

other helicases, SNF2-like ATPase/helicase proteins translocate along DNA and induce

superhelical torsion and/or slide nucleosomes [73,74,75,76,77,78,79,80]. Chromatin

7

remodeling factors contain, in addition to an N-terminal ATPase/helicase domain, other

accessory domains which provide specificity of function [81]. SNF2-like CRFs achieve

the ability to bind DNA sequence-specifically by incorporating sequence-specific

transcription factors into the complex [82].

Classically, there are four subclasses of the SNF2 superfamily: (1) SWI/SNF,

including Brg1 and Brm, both of which have C-terminal bromodomains that recognize

specific acetylated lysines in histones (2) the CHD class, members of which are

characterized by the presence of chromodomains that recognize methylated lysines on

histone tails, (3) the INO80 class, which is characterized by an ATPase/helicase domain

with an extended central split, and (4) the ISWI class, which contains two mammalian

homologs, Snf2l and Snf2h, having SANT and SLIDE domains responsible for

mobilizing nucleosomes [17]. There remain a number of proteins that contain the SNF2-

like DNA-dependent ATPase/helicase domain but that do not fit into these four major

categories, including Chd1l, which is the subject of this dissertation (Fig. 3) [81].

The SNF2 class of chromatin remodeling factors has traditional roles in regulating

gene expression. The founding member, Snf2p was identified in yeast through genetic

screens that identified mutants defective in mating type switching (SWItch) and sucrose

fermentation (Sucrose-Non-Fermenting), and is required for expression of genes that

regulate those processes [83,84]. The mammalian homologs of Snf2p, Brg1 and Brm,

associate with the BAF complex, and as previously mentioned, can regulate gene

activation or repression, depending on the context [71,72]. Like Brg1-containing

complexes, the NURF complex, which contains one of two mammalian ISWI homologs,

Snf2l, also regulates gene expression [85,86,87]. Mice with non-functional NURF are

embryonic lethal with major deficiencies in expression of BMP signaling genes [85].

Members of the CHD class can also regulate transcription, as evidenced by CHD7, the

chromatin remodeling factor whose dysfunction in humans is the cause of CHARGE

syndrome [88]. CHD7 co-localizes with the permissive H3K4me3 mark and interacts

with the SMAD family of transcription factors to control gene activation of cell-type

specific genes [89]. The INO80-containing complexes in yeast, Drosophila, and

mammals also control gene expression [90]. In addition, INO80 complexes have

multifunctional roles in histone deposition of the histone variant H2AZ [91] and in

8

regulating DNA repair by interacting with the phosphorylated H2AX that is immediately

localized to sites of DNA damage [92,93]. Thus, the SNF2 family of CRFs have roles in

transcriptional regulation, but can also have roles in other chromatin-dependent processes

[94].

Epigenetic Pluripotency in ES Cells

ES cells are a pluripotent cell type that can give rise to the three primary germ

layers, endoderm, mesoderm, and ectoderm [95,96]. They are derived from the

pluripotent inner cell mass (ICM) of the blastocyst, but unlike the ICM, ES cells self-

renew in culture and maintain pluripotency indefinitely [95]. The pluripotent state is

orchestrated by a network of transcription factors including POU domain transcription

factor Oct4, Sox2, and Nanog [97,98].

Chromatin in pluripotent ES cells is generally more relaxed than differentiated

counterparts, a state thought to preserve the plasticity of future gene expression programs.

Histone modifications permissive for transcription, including trimethylation of H3K4 and

acetylation of H4, are more abundant in ES cells than in differentiated cells [99]. In

addition, modifications that repress gene expression are less frequent in ES cells. ES cell

chromatin is hyperdynamic with respect to histones and architectural chromatin proteins

[100]. While core histones bind stably to chromatin in differentiated cells, they are

loosely bound in ES cells. The dynamic exchange of the linker histone H1 is required for

pluripotency, as its immobilization causes ES cell differentiation [100].

A unique chromatin landscape can be found in ES cells. Trimethylation of H3K4

is a transcriptionally permissive mark, while trimethylation of H3K27 is a repressive

mark. In ES cells and in other multipotent cells, the appearance of “bivalent” domains

that contains overlapping H3K4me3 and H3K27me3 is intriguing [101]. In ES cells,

genes that are marked with bivalent domains are either not expressed or expressed at very

low levels and are strongly correlated with developmental roles [101]. Upon

differentiation, however, only one of the marks is retained and the gene either becomes

activated or is continued to be repressed. Which mark is retained depends on which

lineage the cell has adopted. In this way, ES cells can repress genes associated with

differentiation without shutting them off permanently.

9

It has been proposed that pluripotency is the default state [102]. Transcriptionally

permissive, open chromatin allows the expression of many genes, but those of the

pluripotency transcriptional network dominantly repress differentiation genes and

establish positive feedback loops to ensure the continued expression of pluripotency

genes. The pluripotent state is further locked in by epigenetic changes in chromatin such

as histone acetylation and methylation. One of these epigenetic changes is the

demethylation of H3K9 by the histone demethylase enzymes KDM3a (previously known

at Jmjd1a) and KDM4c (Jmj2c), both of which are encoded by Oct4 target genes [103].

In line with this hypothesis, it has recently been shown that somatic cells can be

induced to become pluripotent by the introduction of just a few transcription factors,

namely Oct4, Sox2, c-myc, and Klf4 [4]. It has been nearly sixty years since the first

report demonstrating that live tadpoles could be obtained with high efficiency from

nuclear transplantation of blastula cells into enucleated amphibian oocytes [104] and

more than ten years since the birth of “Dolly the Sheep,” the first mammal cloned from

the nucleus of a fully differentiated somatic cell [3]. These studies have clearly

demonstrated that cellular differentiation is a reversible process that can be mediated by

factors present in the oocyte. The identity of these reprogramming factors remain

elusive, but studies on induced pluripotent stem cells (iPS) show that forced expression of

only a few transcription factors is sufficient to induce complete reprogramming [4]. iPS

cells are indistinguishable from ES cells in terms of their DNA methylation patterns and

ability to generate all the cell types of an adult organism [5,105,106]. Despite the

successful derivation of iPS, the frequency of obtaining truly pluripotent cells remains

low, and a large number of stochastic factors are likely involved [107].

DNA Repair in ES cells

Like the cells of the early embryo, ES cells have strict requirements to ensure that

mutations are prevented from being passed down to daughter cells and subsequent

lineages. Rapidly proliferating cell types such as ES cells must be particularly efficient at

repairing damage because unrepaired DNA lesions can lead to stalling and collapse of the

replication fork, resulting in highly deleterious double-stranded breaks (DSBs).

Emerging evidence indicates that ES cells are somewhat unusual in the way they repair

10

DNA, and this could be due to several factors including the strict need for genetic

fidelity, rapid proliferation, culture adaptations, and unique “open chromatin” structure

[108,109,110]. The “open chromatin” of ES could render them initially more vulnerable

to acquiring DNA damage, but also facilitate repair since repair proteins would have

easier access to sites of damage.

The mutation frequency of ES cells is considerably lower than in MEFs. To

measure spontaneous mutation frequencies, the Aprt locus is often used a reporter locus

because APRT deficiency can be positively selected for in culture [111]. Cells

heterozygous for APRT activity are allowed to accumulate spontaneous mutations, and

then cells in which the second copy of APRT is inactivated through spontaneous

mutation are selected for. The most frequent mutation is loss of heterozygosity (LOH,

80%), either through non-disjunction or mitotic recombination, and point mutations also

occur (20%). Experiments using this Aprt reporter system reveal that the spontaneous

mutation frequency of ES cells is 100 times lower than isogenic MEFs (10-6

in ES cells

versus 10-4

in MEFs) [112]. Interestingly, whereas MEFs sustained LOH mainly through

mitotic recombination, ES cells sustained LOH primarily through non-disjunction. When

the X-linked Hprt locus is used as a reporter, the mutation frequency in ES cells is 1000

times lower than in isogenic MEFs (<10-8

in ES cells vs. 10-5

in MEFs) [112,113]. The

difference in mutation frequencies between the two loci is probably due to the location of

Hprt on the single-copy X chromosome, making it immune to LOH events. When

treated with DNA damaging agents, accumulation of mutations at both the Aprt and Hprt

loci is dose-dependent in ES cells as well as MEFs, eliminating the argument that lower

mutation frequencies occur solely because ES cells are resistant to DNA damage

[114,115]. These data show that ES cells are indeed proficient in removing DNA

damage.

A cell can respond to DNA damage in one of three ways: (1) it can ignore the

damage and risk propagating a deleterious mutation, (2) it can repair the damage through

a variety of pathways, and (3) it can terminate itself through programmed cell death. The

latter response is chosen more readily in ES cells than in other cell types, and this can be

seen in their hypersensitivity to DNA damaging agents [116,117,118]. The propensity to

initiate apoptosis is not directly mediated by the tumor suppressor p53 because it remains

11

inactivated in the cytoplasm of ES cells, even upon insult by DNA damaging agents

[119]. Instead, the lack of a G1 arrest allows cells to slip into S phase where lesions can

be exacerbated by conversion to DSBs upon stalling and collapse of the replication fork,

and apoptosis may result as a consequence. Interestingly, ES cells are deficient in the

two pathways that regulate a functional G1/S phase checkpoint, the p53/p21-mediated

pathway and the ATM-Chk2-Cdc25A-mediated pathway [120]. In addition to non-

functional cytoplasmic p53, the Cdk inhibitors, p21 and p27 are expressed at undetectable

levels in ES cells [120]. The second pathway is rendered ineffective because of Chk2

localization to centrosomes [108]. Ectopic Chk2 expression in ES cells restores the G1/S

phase checkpoint and reduces sensitivity to DNA damaging agents. The authors suggest

that the lack of a normal G1 arrest is beneficial for ES cells and encourages damaged

cells to be removed from the population by apoptosis.

In light of the lack of G1/S phase checkpoint in ES cells, it is perhaps not

surprising that ES cells stain strongly for phosphorylation of the histone variant H2AX, a

marker of DSBs [109]. Despite the high prevalence of DSBs, ES cells are more efficient

in repairing DSBs induced by ionizing radiation than either their differentiating embryoid

body or MEF counterparts [121]. DSBs can be repaired through one of two pathways

[122,123]. One is non-homologous end joining (NHEJ) that ligates together two double-

stranded ends of DNA. This method is error-prone since any two broken ends can be

used. The second pathway is homologous recombination repair (HRR) that utilizes

homologous recombination and is error-free. Because HRR utilizes the sister chromatid

as a template, this pathway is active during late S phase [123,124,125]. A second

advantage of skipping G1/S phase checkpoint and allowing damaged cells to enter S

phase, therefore, is that error-free HRR pathway can be utilized in place of NHEJ [109].

The protracted S phase in ES cells supports this notion [126]. Therefore, the high

prevalence of γH2AX staining may not reflect a deficiency in repairing DSBs, but instead

a choice to send cells with damaged nucleotides into S phase where DSBs can be

preferentially repaired by HRR.

In one study, DSBs were introduced into ES cell lines by transfecting a plasmid

encoding the rare cutting endonuclease I-SceI, which cut two I-SceI cleavage sites

engineered into two different chromosomes [127]. Of the clones that successfully

12

repaired the DSB, 79% were repaired through conservative HRR, and 21% were

characterized by chromosomal translocation characteristic of NHEJ. The data suggest

that DSBs are preferentially repaired by HRR when only two DSBs are present. The

same authors showed that NHEJ was never used when only one DSB was introduced. A

separate group reported that two DSBs located in cis in ES cells are repaired by NHEJ,

but highly inefficiently [128]. The studies highlight a potentially compounding factor,

that the number of DSBs and where they are located could have a major impact on which

DSB pathway is used [129]. In other words, the more broken ends are available, the

more opportunities may be found for NHEJ and the more difficult it may be to align

chromosomes for homologous recombination. Several other studies support a prominent

role for HRR in ES cells [130,131,132]. While it has long been supposed that ES cells

are unique in their utilization of HRR and exclusion of NHEJ, it was only recently that

Tichy et. al. demonstrated that HRR is the preferential repair pathway in ES cells, and

upon ES cell differentiation by retinoic acid, the primary repair pathway switches to

NHEJ [133]. Similar results were simultaneously reported by Serrano et. al., but this

study did not compare ES cells with cells directly differentiated from ES cells [134].

Instead, Serrano et. al. showed that ES cells can utilize HRR prior to DNA replication,

resolving the discrepancy between inefficient mitotic recombination yet high prevalence

of HRR in ES cells [134].

Other repair pathways are active in ES cells, including mismatch repair (MMR)

and nucleotide excision repair (NER). Mutations in either of two MMR genes, Msh2 and

Mlh1 in humans are primarily responsible for the development of hereditary non-

polyposis colorectal cancer (HNPCC) [135,136]. ES cells lacking Msh2 have

spontaneous mutation frequencies 30-fold higher than wild-type ES cells [137], and Msh2

mutant ES cells are also more resistant to apoptosis upon treatment with DNA damaging

agents [138]. A similar effect was seen in ES cells null for Mlh1 [139]. Photolesions

induced by UV-C radiation are repaired by the NER pathway. The mutation rate in ES

cells treated with low doses of UV-C were comparable to that of MEFs, but at higher

doses, ES cells acquired disproportionately more mutations, indicating that the NER

pathway has a saturation level in ES cells [117]. ES cells lacking ERCC1, a major NER

component, also exhibit 10-fold higher mutation frequencies than their wild-type

13

counterparts, and at high doses of UV-C underwent massive apoptosis [117]. The

literature is strangely silent regarding the BER pathway in ES cells, but null mutations in

at least one BER member, Aag results in hypersensitivity to DNA damaging agents [140].

Thus, ES cells have lower mutation frequencies than their somatic counterparts,

undergo apoptosis more readily in response to damage, and preferentially use error-free

HRR during S phase to repair damage. Other repair pathways are also active, and these

may function similarly in ES cells than in other cell types.

Chromatin Remodeling in the Zygote and Cleavage-Stage Embryo

Chromatin states of the gametes

Primordial germ cells are set aside early during mammalian development [141].

In females, primordial germ cells enter meiosis and replicate DNA, becoming 4N in

chromosome number. These primary oocytes remain paused in meiosis I until puberty.

In response to hormonal signals, the primary oocyte completes meiosis I, extruding the

first polar body and becoming 2N in number, and is ovulated. In the oviducts, the

secondary oocyte initiates meiosis II, but remains arrested in metaphase II until

fertilization. In contrast, the sperm has completed meiosis I and II and enter the oocyte as

a haploid cell.

Methylation of imprinted genes is already established through reprogramming

during development of the germ cell lineage [142]. Chromatin in the sperm is even more

compacted than normal chromatin in somatic cells, owing to the ~10-fold reduction in

size of the sperm nucleus [143,144]. To achieve this, the sperm adopts a “frozen” state

by exchanging positively-charged protamines in place of histones and eliminating the

need for active transcription by decoupling translation from transcription [143,145,146].

Chromatin compaction through the incorporation of protamines is essential for sperm

function. Human infertility is positively correlated with low abundance of protamines,

and mice with defective protamine production have severe defects in fertility

[147,148,149,150]. Chromatin of both the sperm and the oocyte has high levels of DNA

methylation and is transcriptionally inactive at the time of fertilization [151,152].

14

Chromatin remodeling in the zygote

Upon fertilization, the oocyte completes meiosis II and extrudes the second polar

body. The cortical reaction, accompanied by a sharp increase in intracellular Ca++

levels,

induces changes in the zona pellucid that prevents polyspermy [153]. Chromatin of the

maternal and paternal genomes decondenses and becomes encapsulated in nuclear

envelopes, forming two pronuclei. The pronuclei migrate through the nucleoplasm and

ultimately fuse in a process called synkaryogamy. The zygote then undergoes its first

round of DNA replication and development progresses. Prior to pronuclear fusion,

however, large scale changes in chromatin structure occur, especially in the paternal

genome that must recover from the extreme chromatin compaction and general dearth of

proteins in the sperm (Fig. 4).

The cytoplasm of the oocyte has powerful reprogramming potential, as evidenced

by reproductive cloning through somatic cell nuclear transfer [154]. Within the first

several hours of fertilization, the sperm chromatin is massively remodeled by factors

present in the oocyte [155]. Even prior to the first round of replication, DNA methylation

rapidly decreases and protamines are evicted from chromatin and exchanged for histones

[156]. Interestingly, the incorporation of specific histones into paternal chromatin

establishes an asymmetry between the paternal and maternal genomes. For example,

paternal chromatin preferentially incorporates acetylated histones, which aids in sperm

decondensation upon fertilization [157,158]. In addition, the replication independent

histone H3 variant, H3.3 is utilized, and these are generally replaced by canonical H3

histones during the first cell cycle [159,160,161]. Monomethylation of H4K2, a

modification important for DNA repair, is asymmetrically localized to paternal chromatin

[159].

The paternal genome is rapidly demethylated within 8 hours of fertilization, even

in the presence of the replication inhibitor aphidicolin [162,163]. In contrast, the maternal

genome retains DNA methylation in the zygote and, due to the absence of Dnmt1, is

passively demethylated during subsequent cell divisions [164,165]. In plants

demethylation of 5-meC is performed directly by the glycosylases DEMETER, ROS1,

DML1 and DML2 and unmethylated cytosines are inserted into abasic sites through the

base excision repair (BER) pathway [166,167]. Despite DNA demethylation being well

15

established in plants, the identification of a parallel mechanism in mammals has been

elusive. Many groups have reported the identification of demethylase activities

[168,169] or enzymes, including the glycosylase TDG [170], the methyl-binding protein

MBD2 [171], a nuclear protein called GADD45a [172], and more recently, the de novo

methyltransferases DNMT3a and 3b [173,174], but so far, biochemical studies have been

irreproducible, refuted, or controversial in terms of mutant phenotypes [175]. The

discovery of bona fide demethylase enzymes has lead to controversy over whether they

exist in mammals at all, particularly because demethylation can occur passively during

replication in any dividing cell type, including primordial germ cells in which 5-meC on

imprinted genes is known to be removed [175]. While the extent of 5-meC

demethylation in the developing mammalian embryo and in the adult may be uncertain,

rapid, replication-independent demethylation is clear in the paternal genome, supporting

the existence of an active demethylase enzyme [163]. A recent publication by Hajkova

et. al. demonstrates that demethylation in the zygote is partially impaired in the presence

of PARP inhibitor and is co-localized with members of the BER pathway [176],

suggesting that the mechanism of demethylation may indeed be similar to that in plants.

Further studies focusing on the zygote should be rewarding. How the demethylase

targets the parental genome while allowing 5-meC to remain on the maternal genome is

also an interesting open question.

Intriguingly, the methylation of imprinted genes and of some repeat sequences is

somehow protected from the rapid demethylation of paternal chromatin and the passive

demethylation of maternal chromatin [177]. One explanation is that methylation at

imprinted genes is quickly re-established by factors in the oocyte after active

demethylation. An alternative and more likely explanation is that a counteracting

enzyme may function to protect imprinted genes, or chromatin at imprinted genes may

have additional marks that fend off the demethylase.

Maternal/zygotic transition

While the oocyte is supplied with maternally contributed transcripts and proteins

[178], these cannot sustain the embryo for long, and transcription must be reinitiated in

the zygote through a process called zygotic genome activation (ZGA, Fig. 4). Massive

16

degradation of maternal RNAs that might interfere with cleavage-stage development

immediately precedes ZGA [179]. This is a highly ordered process and regulated by the

Smaug or EDEN-BP proteins and by the microRNA mir-430 that induce deadenylation of

the 3’ UTRs of maternal transcripts and trigger nuclease degradation [180,181].

In mice ZGA occurs at the two-cell stage; in humans it occurs between the four-

cell and the eight-cell stage, and results in a whole new profile of gene expression

[152,182,183]. Little is known about how the parental genomes are silenced in the germ

cells, nor about what triggers the activation of transcription from the zygotic (ZGA).

Interestingly, some basal level of transcription can be detected from the paternal

pronucleus, suggesting that paternally derived transcripts could participate in the

initiation of ZGA [184]. It is likely that ZGA involves extensive and rapid remodeling of

chromatin. Brg1, the enzymatic subunit of the BAF complex, is required for

reprogramming of permeabilized human somatic cells using Xenopus laves oocyte

extract, because antibody depletion of Brg1 prevented reprogramming [185]. More

recently, Brg1 was shown to be required specifically for ZGA in mice. Brg1 knock-out

mice, which retain maternal Brg1 expression, exhibit peri-implantation lethality [186].

When Brg1 is specifically depleted in the oocyte, fertilized embryos arrest at the 2- to 4-

cell stage with a 30% reduction in genes involved in transcription, RNA processing, and

cell cycle regulation [187]. The authors note a reduction in the levels of dimethylation of

H3K4 (a mark associated with transcriptional activity) and propose a model in which

Brg1 affects chromatin structure to promote ZGA [187].

Chromatin Remodeling in the Blastocyst

Differentiation of the ICM

After several rounds of cell division without concomitant growth, the embryo

undergoes compaction and achieves a unique organization: there are now cells on the

inside of the embryo surrounded by cells on the outside [152]. It has long been suggested

that the inside/outside organization of cells leads to the first differentiation choice to

become either inner cell mass (ICM) or outer trophectoderm (TE) cells (Fig. 4) [188].

The outer cells are polarized, by virtue of having cellular connections to one side but not

the other, and become epithelialized [189]. In contrast, the inner cells are surrounded on

17

all sides by neighboring cells and lack polarity [190]. TE cells are required for

implantation and do not contribute to the embryo proper. Instead, they will give rise to

the extra-embryonic tissues including the placenta. All the cells of the embryo proper are

exclusively derived from the ICM, although a derivative of the ICM, the primitive

endoderm, will also contribute to extra-embryonic endoderm [191].

How the ICM and TE lineages are determined is still under investigation, and the

role of epigenetics is unclear. Initially, asymmetric cell divisions along the apical/basal

axis during early cleavage stages allocate one daughter cell to the inside of the morula

and one to the outer surface of the morula; symmetric cell divisions result in two

daughter cells with an outer-facing surface [188,192]. There has been considerable

debate regarding whether early asymmetric cell divisions in the 4- to 8-cell stage embryo

contribute to an intrinsic bias leading toward differentiation into either the ICM or TE,

but the emerging consensus seems to be that apical/basal polarity in the late morula

overrides earlier biases [193,194,195,196]. An elegant blastomere dissociation study

showed that the TE is determined prior to the ICM by the 32 cell stage [197]. Labeled

and dissociated cells of the 16-cell morula were fully competent to become either ICM or

TE, regardless of whether they had been labeled as “outer” or “inner.” In striking

contrast, aggregates of “outer” cells of a 32-cell morula formed TE but not ICM. At the

same time, “inner” cell aggregates were able to form an ICM as well as a TE layer

competent to implant into surrogate mothers, showing that inner cells of the 32-cell

morula are not yet determined to become ICM.

Transcriptional changes are known to participate in the differentiation of the TE

and ICM. The TEA-domain containing transcription factor Tead4 initiates the

differentiation of the TE, and Tead4-/-

embryos cannot form TE and arrest prior to

blastocyst formation [198]. Because of this severe phenotype, it is proposed to be the

most upstream transcription factor in the specification of the TE, but it should also be

noted that many early knock-out phenotypes are masked by maternally contributed

transcripts and proteins. Tead4 regulates the expression of the caudal-type homeobox

transcription factor Cdx2 [199]. Cdx2 promotes symmetric cell divisions and thus

regulates the allocation of cells [200]. Cdx2 and the Pou-domain transcription factor

Oct4 are co-expressed in blastomeres but mutually oppose each other as the blastocyst

18

forms [192,201,202]. Upon formation of the blastocyst, Cdx2 becomes excluded from

the ICM, and Oct4 is excluded from the TE [203,204,205,206]. How cell polarity signals

the activation of specific transcription factors remains a mystery.

Epigenetic differences between the TE and ICM

In additional to transcriptional changes, there are also epigenetic differences

between the ICM and TE. Whether epigenetic modifications participate in the

specification of the two tissues or whether they lock in differentiation choices and set up

future lineage choices is not clear. Interestingly, despite delayed determination of ICM

cells compared to TE cells [197], the ICM is characterized by much higher levels of DNA

methylation [158]. Chromatin is passively demethylated during the cleavage stage

embryo due to inactivity of Dnmt1, hitting a trough at the late morula stage; but as the

blastocyst forms, de novo methylation mediated by Dnmt3a and 3b begins, and this

occurs more prominently in the ICM than TE lineages [165]. Appropriate methylation

patterns are essential to the developing embryo, because Dnmt3a/3b null mice show

severe abnormalities and lethality at E8.5-E9.5 [27]. The transcription factor Elf5 plays a

role in the determination of TE lineage. In the epiblast and in ES cells, loss of DNA

methylation at the promoter of Elf5 alleviates its repression and induces aberrant TE

formation, suggesting that the enrichment of DNA methylation in the ICM may

participate in repression of TE lineage determination genes [198].

In addition to differential DNA methylation, the ICM also differs from TE in

trimethylation of H3K27, a transcriptionally repressive mark that is catalyzed by the

Polycomb Repressive Complex 2, PRC2 [207,208]. Genome-wide µChIP studies show

that signaling and developmentally-regulated genes in the ICM were preferentially

methylated at H3K27, suggesting that PCR2 is more active in the embryonic lineage

[208]. No differences between ICM and TE were found in methylation of H3K4, a mark

associated with gene activation [208]. Taken together with evidence of increased DNA

methylation, the theme arises that the cells of the ICM contain more transcriptional

repressive chromatin than the TE [209]. One possibility is that TE differentiation

programs are intrinsically dominant, and wide-spread transcriptional repression is

necessary to maintain ICM fate. An alternative explanation is that the generally

19

repressive chromatin marks are necessary to ensure accurate gene expression programs

during subsequent development.

X chromosome inactivation

X chromosome inactivation (XCI) is a critical process that achieves dosage

compensation in mammals [210]. XCI occurs early during development and involves the

expression of non-coding RNA Xist [211,212,213]. Upon XCI initiation, the two X

chromosomes transiently pair at the X inactivation control center and at the X pairing

region, and Xist is expressed mono-allelically from the future inactive X chromosome

(Xi) [214]. Xist binds along the length of the Xi and triggers a stream of epigenetic

events that shut off Xi, including widespread DNA methylation , methylation of H3K9

and H3K27, incorporation of the macroH2A histone variant, and hypoacetylation of

histones H2A, H2B, H3 and H4 [23,207,215,216,217,218,219]. Interestingly, the

incorporation of macroH2A has been shown to interfere with the mobilization of

nucleosomes by SWI/SNF [220]. In addition, Xi becomes localized to the nuclear

periphery, where heterochromatin is frequently relegated [216].

The traditionally held view that X chromosome inactivation is random is not so

straight forward. In fact, random X inactivation occurs specifically in the ICM, and the

paternally-inherited X (Xp) is non-randomly inactivated in trophectoderm derived tissues

[221]. During fertilization, Xp is delivered in the inactivated form but becomes active

upon sperm chromatin remodeling [222]. At the four cell stage, Xp is again inactivated

and remains so until the differentiation of the ICM [223]. In the trophectoderm,

inactivation of Xp is maintained; but in the ICM, Xp undergoes a second round of

reactivation [223]. Only after implantation does random inactivation of ICM-derived

cells occur.

How the X chromosome is reactivated is not well understood. The first and

second waves of Xi reactivation differ in at least three critical points: (1) the first round

reactivates a Xp that was inactivated during spermatogenesis by a process mechanistically

distinct from XCI called meiotic sex chromosome inactivation (MSCI), which occurs in

an Xist-independent manner [224,225]; (2) the second event is followed by random XCI,

but the first reactivation event does not remove parent-of-origin marks because it is

20

followed by imprinted XCI of Xp [226]; and (3) the second wave occurs ICM-

specifically and may depend on pluripotency factors [223].

In somatic cells, Xist expression is not required to maintain XCI, but loss of Xist

expression is accompanied by loss of macroH2A localization to the Xi [227,228]. In the

ICM and in ES cells, however, loss Xist reactivates the Xi, potentially through loss of

macroH2A localization to Xi [223]. In the zygote, MacroH2A transcripts are degraded

prior to pronuclear formation, raising the possibility that loss of macroH2A participates

in both the first and second X reactivation events [229].

Two mechanisms have emerged to explain why Xi is reactivated in the ICM

alone, the first being repression of Xist in the ICM, and the second being PcG action in

the TE. Intriguingly, Xist expression is repressed in early ICM and ES cells, and unlike

in somatic cells, Xist is required to maintain XCI in both these pluripotent cell types

[223]. Reactivation of Xi in the TE is inhibited by the PcG members Eed and Enx1.

H3K27me3, catalyzed by PcG complexes, while generally low in the TE, shows distinct

Xi localization in TE cells [207]. Eed and Enx1 proteins localize to the Xi in TE [230],

and Eed mutant mice do not retain XCI in the TE [231]. One hypothesis to explain these

elaborate mechanisms to allow random XCI in the ICM but imprinted XCI in the TE is

that repression of paternally inherited genetic material in the placenta could be beneficial

to the mother, but random expression of paternally or maternally derived genetic material

could be beneficial to the offspring.

Derivation of ES cells from the ICM

Mouse ES cells are derived from cells of the ICM and, because they can be

differentiated and cultured to large numbers, are frequently used as a surrogate system to

study the biochemistry of pluripotency and early differentiation [95,96]. Several

different procedures are used to derive ES cells that involve the propagation of ICM cells

on a gelatin-coated culture dish in defined media [232]. Some of these procedures

include whole-embryo culture, immunosurgery whereby the TE cells are selectively lysed

by the complement cascade, and physical microdissection of ICM cells. The techniques

are invariably inefficient and involve drastic and stressful changes of environment for the

21

cells of the ICM [95]. It is therefore not surprising that heated debate on the similarity

between ES cells and ICM has arisen.

In females, both X chromosomes are active at the developmental time period at

which ES cells are derived [223,233]. Consequently, both chromosomes remain active in

ES cells. In addition, parent-of-origin epigenetic marks appear to be lost, because XCI is

random in TE lineages of embryos derived from ES cells [226]. This is in contrast with

normal embryos and embryos derived from SCNT of somatic cells in which XCI in TE

lineages is non-random. The Xi of the somatic cell donor becomes reactivated normally

during ICM differentiation and in the TE is faithfully and non-randomly inactivated again

[226]. With regard to X chromosome inactivation, ES cells resemble the ICM of the pre-

implantation embryo.

Expression of transcription factors Oct4, Sox2, Nanog, and other pluripotency

factors is preserved between ICM and ES cells [234]. One of the most striking

differences between ES cells and the ICM is that the cells of the ICM are characterized

by relatively repressed chromatin while ES cells have hyperdynamic chromatin structure

and an “open chromatin” conformation [100,102,207,209]. An early study using a target-

specific µChIP approach showed that the repression of some key differentiation genes

was lessened in ES cells compared to ICM [235]. Dahl et. al. used a µChIP-chip

approach to analyze H3K4me3 and H3K27me3 distribution in ICM and in ES cells

directly derived from ICMs of the same strain [208]. Methylation patterns on both the

repressive chromatin mark H3K27 and the permissive mark H3K4 were drastically

altered between the ICM and ES cells. Only 30% of promoters with H3K4me3 and 20%

of promoters with H3K27me3 in the ICM retained those marks in ES cells. This change

in methylation patterns is not due to deficient methylation because a large number of

promoters in ES cells also gained H3K4 (40%) or H3K27 (80%) methylation.

Approximately 500 promoters in the ICM contained the so called “bivalent” domains

containing both H3K4me3 and H3K27me3 marks. In ES cells, this number nearly

doubles, but less than 10% of promoters with bivalent domains in ES cells were also

H3K27/H3K4me3-marked in the ICM. Furthermore, 50% of H3K27/H3K4me3-marked

genes were expressed in ICM, in sharp contrast with bivalent domains being mostly

22

associated with repressed genes in ES cells. These studies show that there are major

epigenetic differences between the two cell types.

Dahl et. al. console the stem cell field by showing that the epigenetic differences

between the ICM and ES cells were not as great as those between the ICM and the TE.

They suggest that during the generation of chimeric mice, ES cells are again

reprogrammed and that epigenetic changes in ES cells could explain why some ES cells

are competent to form chimeric mice and others are not. Further investigation into ES

cell epigenetics and reprogramming is clearly needed to realize the therapeutic potential

of ES cells.

DNA Repair in the Zygote

Even in the mammalian embryo that is relatively protected from exogenous

sources of DNA damage such as UV radiation, a typical cell experiences greater than ten

thousand lesions from exogenous sources [6]. These sources can include oxidation,

alkylation, and spontaneous hydrolysis that affect the chemical bonds of DNA,

threatening the accuracy and processivity of replication machinery. If mutations arise in

the early embryo and are allowed to fall into the germ line, that individual’s offspring

may inherit highly deleterious phenotypes. Even if mutations do not enter the germ line,

mutations acquired early during development can be propagated through other cellular

lineages and can result in embryonic lethality and disease.

Mutations in DNA repair machinery can result in a wide range of human postnatal

diseases including Xeroderma Pigmentosa, Hereditary Non-Polyposis Colorectal Cancer,

breast and ovarian cancer, and Ataxia-Telangiectasia [236]. Because of the late onset of

these conditions, their development is probably the result of multiple compounding

factors, including environmental variables, and is not covered under the scope of this

introductory chapter on preimplantation development.

Unexpectedly, most de novo germ line mutations in humans can be linked to

paternally-derived chromosomal anomalies and are therefore believed to have arisen from

failure to repair the parental genome [237,238,239]. DNA repair is fully functional in the

zygote, but cannot occur after the completion of spermiogenesis due to the extreme

compaction of chromatin [144,240,241,242,243]. Perhaps one of the most important

23

early reprogramming events, therefore, is the repair of the parental genome before the

onslaught of the replication machinery. In addition to remodeling DNA methylation and

the histone composition of paternal chromatin following fertilization, the zygote must

also repair the paternal genome before the onslaught of the replication machinery.

Upon nuclear condensation during spermatogenesis, DSBs are introduced into

chromatin in a histone hypoacteylation-, topoisomerase II-dependent manner

[244,245,246]. The intentional “nicking” of DNA is thought to be necessary to relieve

tension during histone withdrawal, and the majority of nicks are repaired during the final

stages of transition protein and protamine incorporation [148,247,248,249]. Repair of

double-stranded breaks is a delicate process, and defects in protamine levels are tightly

correlated with chromosomal aberrations and infertility in men [250,251]. Spermatids

retain very little ability to repair DNA after nuclear condensation is completed [252];

thus, remaining DSBs and lesions accumulated during the variable delay between

spermatogenesis and fertilization are reliant upon DNA repair in the zygotic environment

[240,241,242].

Because many mutations in components involved in the repair of DSBs result in

early lethality [253], much effort has been focused on determining the relative

contribution of error-free homologous recombination repair (HRR) versus rapid but error-

prone non-homologous end joining (NHEJ) in the embryo [123]. The duration of the cell

cycle in embryonic cells can be as rapid as 2-3 hours, resulting in the shortening of G1

and G2 gap phases [254]. The first cellular division of the zygote, however, does not

occur until 18-20 hours after fertilization, drastically extending the duration of the G1

phase and the time in which NHEJ predominates [122,123,254]. The zygote is a very

expensive and valuable cell in terms of its rarity (there is only one per embryo in contrast

to the many cells of the gastrulation-stage embryo) and its purpose (to give rise to all

cellular lineages). Unlike plentiful, rapidly dividing cells that are able to abort instead of

risk propagating mutational errors, the zygote would be very unwise to initiate apoptosis.

Therefore, rapid repair of DNA through NHEJ followed by cellular division may be more

beneficial than forcing cells to S phase where repair failures can cause cell death. To

determine which DSB repair pathways are involved in zygotic DNA repair, Derijck et. al.

compared the chromosomal abnormalities in mouse zygotes deficient in NHEJ (the skid

24

mouse, with null mutations in DNA-PKCs) or in HRR (Rad54/Rad54B double knock-out)

after irradiation. The authors found that DSBs in sperm-derived chromatin is primarily

repaired by NHEJ during G1 of the zygote, and that the majority of chromosomal

deficiencies in skid zygotes were paternal. Interestingly, markers of DNA damage

indicate that spontaneous damage occurs more frequently on paternal chromosomes. In

addition, they demonstrate that both HRR and NHEJ function during S/G2 phase.

Chd1l as an Oncogene and DNA Damage Response Protein

At the time this dissertation project began, very little was known about the

chromatin remodeling factor Chd1l, or Chromodomain, Helicase/DNA Binding Protein 1,

except what could be gleaned from genetic conservation, protein structure, and

expression studies. Even its nomenclature was misleading, as it does not contain the

chromodomain that characterizes the CHD subfamily of chromatin remodeling factors.

Chd1l is also called ALC1, or Amplified in Liver Carcinoma 1, because 1q21, a large

genomic region encompassing 12.4 million base pairs on chromosome 1, is frequently

amplified in human patients with hepatocellular carcinoma [255,256]. However, 320

other genes are found in 1q21, and it was therefore unclear whether Chd1l deserved the

synonym.

Chd1l homologs can be found in plants, mammals and most other vertebrates, but

not in the Xenopus genus. No homologs can be found in the Saccharomyces genus,

Neurospora crassa, or multicellular invertebrates including C. elegans and D.

melanogaster [81,257]. Data from the Scott laboratory published in 2004 show that

Chd1l is expressed in the preimplantation embryo, with peak expression at the morula

stage, just prior to the formation of the blastocyst [258]. Expression studies showing

genes whose transcripts are enriched in the inner cell mass (ICM) (presented in Chapter

2) identify Chd1l transcripts as showing ICM enrichment.

Chd1l contains an N-terminal, SNF2-like DNA-dependent ATPase/helicase and a

C-terminal macrodomain. Other proteins that contain the ATPase/helicase domain are

members of the SNF2 family of chromatin remodeling factors discussed previously.

These CRFs have broad roles in DNA processing, most notably transcriptional activation

and/or repression and DNA repair. Other macrodomain containing proteins bind to PAR

25

and carry out PARP processes, including transcriptional regulation, DNA repair, and

apoptosis. The presence of these domains raised the possibility that Chd1l participates in

regulating gene expression or DNA repair. This laboratory was interested in exploring

pluripotency and differentiation choices in early development, thus a role for Chd1l in

development and in gene expression was pursued in this dissertation.

Two other groups investigated the role of Chd1l in DNA repair and published

their results in 2009. Both groups confirmed that Chd1l binds to PAR through the macro

domain and has PARP-dependent, NAD+-dependent ATPase and nucleosome sliding

activities [259,260]. Mass spectrometry analysis in HEK293 cells revealed that Chd1l

associates with histones H2A and H2B and base excision repair (BER) pathway and

double-stranded break (DSB) repair pathway members, including PARP-1, XRCC1, and

APLF [259]. In addition, when treated with oxidative damage-inducing H2O2, Chd1l

becomes associated with DNA-PKcs. Association with the DNA repair proteins can be

blocked by treatment with a PARP inhibitor. Transient transfection of YFP fused to

Chd1l and laser irradiation of cells demonstrated that Chd1l rapidly localizes to sites of

DNA damage, and its localization depends on both functional macro and ATPase

domains (Fig. 5) [259,260]. Knock-down of Chd1l significantly increases the sensitivity

of U2OS cells to the DNA damaging agents H2O2 and phleomycin, suggesting that these

cells may be deficient in DNA repair. Chd1l therefore participates in the DNA damage

response, but despite these elegant experiments, a direct role in the repair of damage has

yet to be shown.

Eight years after Guan et. al. published their findings that a minimal genomic

region containing 1q21 was amplified in over 50% of patients suffering hepatocellular

carcinoma (HCC) [255], Guan’s laboratory identified ALC1, or CHD1L as the oncogene

responsible for carcinogenesis [261]. cDNA synthesized from a primary HCC tumor was

hybridized to microdissected 1q21 DNA, isolating CHD1L-encoded transcripts. Over-

expression of CHD1L increased colony formation of HCC and liver cells in soft agar

assays, and xenografts of CHD1L over-expressing cells increased teratoma formation in

nude mice. The tumor suppressor protein p53 was reduced, as was p21, a negative

regulator of CyclinE/Ckd2-mediated G1/S phase transition. Transgenic over-expression

of CHD1L in mice resulted in spontaneous tumor formation in nearly a quarter of the

26

mice, and MEFs cultures from transgenic mice displayed reduced levels of p53 [262].

Furthermore, knock-down of CHD1L using siRNA in HCC cells decreased colony

formation in soft agar assays, increased apoptosis, and decreased levels of apoptotic

proteins Caspase 3 and Bax [261]. These studies provide compelling evidence for the

role of Chd1l as an oncogene, although the causality of Chd1l over-expression in altering

cell cycle and apoptotic proteins has yet to be determined.

How can a DNA repair protein act as an oncogene? Ahel et. al. addressed this

question by over-expressing Chd1l in HEK293 cells and assaying for the γH2AX, a

marker of DSBs [259]. Over-expression of CHD1L alone did not change the intensity of

γH2AX, but when CHD1L over-expressing cells were treated with phleomycin, γH2AX

intensity increased nearly two-fold. The authors propose that over-expression of CHD1L

induces chromatin relaxation, rendering DNA susceptible to DNA damage. Interestingly,

the increase in γH2AX intensity was not seen with phleomycin treatment of cells over-

expressing a mutant CHD1L deficient in binding ATP. Although the authors did not

show chromatin relaxation, there is precedence for over-activation of oncogenes inducing

DNA damage [263]. Activation of oncogenes ras, myc, cyclin E, mos, cdc25A, and E2F1

has been observed to induce DSBs [263]. In the model proposed by Halazonetis et. al.,

precancerous tissues are associated with increases in DSBs and apoptotic indices, which

are then followed by loss of p53 function and sharp increases in proliferation rates as

cancer develops. The loss of p53 tumor suppression is frequently found in cancer and

can be due to mutations in Trp53 itself or in other genes that activate p53 [263]. Thus

cells over-expressing Chd1l accumulate DSBs and may activate p53-mediated apoptosis

followed by selection of cells with impaired p53 function. If this model is valid,

reduction of p53 and other tumor suppressor proteins and activation of cell cycle proteins

would an indirect effect of Chd1l over-expression in cells and transgenic mice.

Summary

Chromatin organization in the eukayotic cell is complex, and DNA-dependent

nuclear processes require the activity of many different chromatin remodeling factors.

The zygote, embryonic cells, and ES cells have particularly active remodeling to

reprogram chromatin to achieve the transition from two parental gametes to a single

27

embryo, to establish and/or maintain a pluripotent state, or to begin carrying out complex

differentiation as the embryo develops. Increasing our understanding of the chromatin

remodeling factors that participate in these early cell types will have broad implications

for nuclear reprogramming, DNA repair, assisted reproduction techniques (ART), and

developmental biology as a whole. The study of Chd1l, in particular, could shed light on

how transcription and/or DNA repair are controlled in the developing embryo, or could

yield novel discoveries on preimplantation reprogramming processes that are not well

understood.

28

Figure 1.1 PAR modifies nuclear proteins

Parp-1 synthesizes polymers of ADP-ribose (PAR) onto nuclear proteins. A. PAR

modifies histones and is proposed to contribute to the “histone code” in a similar way as

histone acetylation (Ac) and methylation (Me), which are recognized by bromo and

chromo domains, respectively. B. The dominant recipient of PAR is Parp-1 itself, which

catalyzes PARylation in an automodification reaction.

PARylation of histones

A B

Auto-PARylation of Parp-1

29

Figure 1.2 ATP-dependent chromatin remodeling

Chromatin remodeling factors such as Brg1 and Chd1l (orange) can remodel nucleosome

structure through the hydrolysis of ATP. Chromatin remodeling factors are often found

in association with other subunits that lend functional specificity to a complex (grey).

Chromatin remodeling factors can catalyze a number of nucleosome alterations including

nucleosome sliding and nucleosome displacement (shown). Their activity facilitates

DNA accessibility to enzymes required for transcription, replication, and DNA repair.

30

Figure 1.3 SNF2 family of chromatin remodeling factors

The four predominate SNF2 subclasses of chromatin remodeling factors, Swi/Snf, CHD,

Ino80, and ISWI, contain the core DNA-dependent ATPase domain that defines the

SNF2 family. The Swi/Snf family is characterized by bromo domain-containing proteins,

and the CHD family is characterized by PHD and chromo domain-containing proteins.

INO80 members have a unique extended split within the ATPase domain. ISWI

members contain SANT and SLIDE domains. Chd1l lacks the domains that characterize

the four major subclasses, but contains instead a macro domain not found in the other

subclasses.

31

Figure 1.4 Chromatin remodeling in the preimplantation embryo

Extensive reprogramming occurs in the zygote as the sperm decondenses. In the paternal

genome, DNA is actively demethylated, protamines are exchanged for histones, and

DNA that was damaged during the “frozen” state of the sperm is repaired. In mice, the

maternal-embryonic transition occurs at the two-cell stage, when maternal transcripts are

degraded, and transcription is initiated from the zygotic genome. Upon compaction,

blastomeres become polarized, and “outer” cells differentiate into the trophectoderm

(TE), while “inner” cells remain part of the pluripotent inner cell mass (ICM).

Differentiation into TE or ICM is accompanied by dissimilarity in the epigenetic

landscapes. For example, the ICM is marked by higher levels of repressive H3K27me3

than the TE.

32

Figure 1.5 Chd1l and the PARP-dependent DNA damage response

The Parp-1 enzyme binds to DNA strand breaks with high affinity and PARylates itself in

an automodification reaction. Chd1l and other DNA repair enzymes are recruited to sites

of DNA damage in a PARP-dependent manner. Chd1l is the only DNA damage response

protein to contain a macro domain known to bind PAR. It is believed that the DNA-

dependent and NAD+-dependent nucleosome remodeling activity of Chd1l facilitates

DNA accessibility for DNA repair enzymes.

33

2. The Chromatin Remodeling Factor Chd1l Is Required

in the Preimplantation Embryo

Alyssa C. Wright1, Denise Leong

2, Q. Tian Wang

1, 4,

Joanna Wysocka1, 3

, Mylene W. M. Yao2, and Matthew P. Scott

1, 5*

1Departments of Developmental Biology, Genetics, and Bioengineering

2Department of Obstetrics and Gynecology

3Department of Chemical and Systems Biology

4University of Illinois, Chicago

5Howard Hughes Medical Institute

Clark Center West W252, 318 Campus Drive

Stanford University School of Medicine

Stanford, California, U.S.A. 94305-5439

*Corresponding author:

650-725-7680

650-725-2952 (fax)

[email protected]

Keywords:

Chd1l, ALC1, preimplantation development, ICM, ES cells, chromatin remodeling

mailto:[email protected]

34

Abstract

Preimplantation development is marked by a multitude of chromatin changes that

allow the zygote to reprogram epigenetic marks of the parent genomes, establish

totipotency, and enact the earliest differentiation choices. To identify novel

developmental regulators, we screened for genes that are preferentially transcribed in the

pluripotent inner cell mass (ICM) of the mouse blastocyst. Genes that encode chromatin

remodeling factors were prominently represented in the ICM, including Chd1l, a member

of the Snf2 gene family. Chd1l is developmentally regulated and expressed in embryonic

stem (ES) cells, but its role in development has not been investigated. Here we show that

reducing Chd1l protein by microinjection of antisense morpholinos causes arrest prior to

the blastocyst stage. Despite this important function in vivo, Chd1l is non-essential for

ES cell survival, pluripotency, or differentiation, suggesting that Chd1l is vital for events

in embryos that are distinct from events in ES cells. Our data reveal a novel role for the

chromatin remodeling factor Chd1l in the earliest cell divisions of mammalian

development.

Introduction

The first differentiation decision in the mammalian embryo is made prior to the

blastocyst stage, when blastomeres must commit to becoming either part of the

trophectoderm (TE) or the inner cell mass (ICM). Cells of the ICM possess the property

of pluripotency and will contribute to the embryo proper, whereas the TE will give rise to

extra-embryonic material. We reasoned that factors compartmentalized in the pluripotent

ICM could be novel developmental regulators of pluripotency or early differentiation. To

identify candidate preimplantation regulators, we performed an expression analysis on

ICM separated by immunosurgery and whole blastocysts and identified genes enriched in

the ICM. Gene ontology clustering revealed a large group of chromatin regulatory

enzymes.

The high degree of chromatin organization within the nucleus is oppressive to

transcription and other processes that require DNA accessibility [12,13]. Chromatin

modifying enzymes can be divided into two broad classes, those that covalently modify

DNA and histone tails and those that utilize energy to alter nucleosome positioning. The

35

latter class is made up of chromatin remodeling factors (CRFs) that contain a core SNF2-

like ATPase/helicase domain responsible for nucleosome remodeling [81,264]. CRFs

participate in key chromatin-dependent processes including transcriptional activation and

repression, histone exchange, cell cycling, DNA repair, and many others [68,92,94,264].

CRFs assemble into multi-subunit complexes, and their functions depend in part on the

composition of the complex [18,66,265].

Profound chromatin changes take place in the zygote and preimplantation embryo

that allow the parental genomes to achieve a state of totipotency and that are necessary

for normal development. Despite successful reprogramming of somatic cells through

somatic cell nuclear transfer (SCNT) and more recently, through viral introduction of

only nuclear factors Oct4, Sox2, Klf4, and c-myc, [4,5,105,106,154], reprogramming in

the embryo remains largely enigmatic. Reprogramming is a multi-step process that

involves extensive changes in chromatin structure beginning immediately upon

fertilization. First, DNA methylation and histone modifications associated with

differentiation must be removed [209]. Next, the embryo must gain independence from

maternally provided proteins and transcripts by activating transcription from the zygotic

genome at the two-cell stage in the mouse [182,266,267]. Lastly, epigenetic

modifications erased during reprogramming must be reestablished on imprinted genes,

the inactive X chromosome (in females), and genes associated with differentiation [209].

As cell division proceeds, DNA must replicate, condense on the metaphase plate, divide,

and decondense again, all the while maintaining genomic integrity.

Relatively few factors involved in preimplantation development have been

identified because phenotypes of traditional genetic manipulations are often masked by

maternally provided transcript and proteins. Homozygous mutation of Brg1, the Snf2-

like catalytic core of the multi-subunit BAF complex, in mice causes peri-implantation

lethality [186]. The phenotype is even more severe when the maternal contribution of

Brg1 is eliminated by an oocyte-specific deletion. In this case, embryos fail to initiate

zygotic genome activation and arrest at the 2-cell stage [187]. It is likely that many more

chromatin modifiers are essential in the preimplantation embryo, so techniques aimed at

early development will likely be fruitful.

36

Among the genes identified in our screen was Chd1l, a largely unexplored CRF

that is a member of the Snf2-like family. Its compartmentalization in the ICM,

expression in ES cells, and temporal regulation prior to the blastocyst stage [258] led us

to hypothesize that Chd1l is critical for early development. The protein has a Snf2-like

ATPase domain but does not fall into any of the four major subclasses because it contains

a C-terminal “macro” domain not present in other Snf2 members and lacks signature

domains of other classes [17,257]. The macro domain binds poly(ADP-ribose), or PAR,

a post-translational modification added to nuclear acceptor proteins by the PARP family

of ADP-ribose polymerases (PARPs) [61,257,259,260]. PARPs, and by deduction the

PAR modification, have well established roles in DNA repair and transcription, among

others [33]. The nucleosome remodeling activity of Chd1l is dependent on PAR

synthesis, indicating that the PAR-binding macro domain is central to its function as a

chromatin remodeler [259,260].

Consistent with its ability to bind PAR, Chd1l is involved in the DNA damage

response. The kinetics of Chd1l localization to, and dissociation from, sites of induced

DNA damage is dependent on the ATPase and macro domains, respectively [259,260].

Recent studies have also implicated Chd1l as an oncogene. The majority of

hepatocellular carcinomas in humans are associated with genomic amplification of a

region that includes Chd1l, and its over-expression in liver cell lines and mouse models is

tumorigenic [261,262]. While evidence is accumulating for a role for Chd1l as an

oncogene and in DNA repair, its importance during development has not been addressed.

We find that Chd1l is expressed in embryonic stem cells (ES cells), which are

derived from the ICM and share the ability to differentiate into the three major germ

layers. Our data show that Chd1l is not required for ES cell viability, pluripotency or

differentiation. To determine whether Chd1l is required during earlier stages in

development, we knocked-down Chd1l in zygote-stage embryos with morpholinos (MOs)

and discovered that embryos arrest prior to the blastocyst stage.

37

Results

Chromatin factors are compartmentalized in the blastocyst

The decision to become inner cell mass (ICM) or trophectoderm (TE) is the first

lineage commitment a totipotent blastomere must make. The inner cell mass (ICM)

retains pluripotency, the ability to give rise to the three primary germ layers, whereas the

TE will give rise to extra-embryonic tissue. We reasoned that mRNAs enriched in the

ICM would encode proteins that contribute to the development of the blastocyst and/or

the establishment of pluripotency. To screen for ICM-enriched mRNAs, we purified the

ICM by immunosurgery [268], taking advantage of the structural organization of the

blastocyst (Fig. 1A). Outer TE cells of the blastocyst were labeled with IgG by

incubation with rabbit anti-mouse serum and specifically lysed by the complement

cascade, leaving behind purified ICMs. RNA extracted from the ICM was compared

with total blastocyst RNA using genome-wide expression analysis.

Transcripts encoding Oct4 and Nanog, factors known to be critical for

pluripotency, were enriched in the ICM 1.9- and 2.4-fold, respectively, providing proof

of sound methodology (Fig. 1B). In addition, mRNAs encoding Cdx2 and Eomes,

markers of extra-embryonic material, were repressed 4.5-fold and 2.4-fold, respectively,

in ICM compared to the whole blastocyst (Fig. 1B). Gene clustering revealed three major

classes of genes whose transcripts are enriched in ICM: cell signaling molecules,

transcription factors, and chromatin-modifying enzymes. Some of the chromatin factors

have known enzymatic activity and/or developmental roles (Fig. 1C). A discussion of

these notable factors follows.

Compared to the TE, chromatin of ICM cells is characterized by modifications

that are indicative of a more “transcriptionally repressive” state [209], displaying among

other repressive marks, higher levels of DNA methylation [158,198]. We find

enrichment of the mRNAs encoding the de novo DNA methyltransferases, Dnmt3a (3.7-

fold) and Dnmt3l (2.8-fold), and the maintenance DNA methyltransferase Dnmt1 (2.5-

fold) in the ICM (Fig. 1C). The higher levels of these enzymes in the ICM could explain

in part the higher DNA methylation observed in the ICM.

Chromatin of ICM cells also contains higher global levels of H3K27

trimethylation (H3K27me3), as shown by both immunostaining [207] and promoter tiling

38

arrays [208]. The repressive modification H3K27me3 is catalyzed and read by Polycomb

Repressive Complexes PRC2 and PRC1, respectively [269]. PRC1 and PRC2 are

complexes of the Polycomb Group (PcG) that function during development to establish

the body plan [270]. We observed ICM enrichment of mRNA encoding the PRC2

subunit Eed (2.9-fold, Fig. 2C), but only moderate enrichment of mRNA encoding the

lysine methyltransferase (KMT) Ezh2 (KMT6, 1.3-fold, data not shown). In ES cells, the

noncanonical KMT Ezh1can partially compensate for loss of Ezh2 [271]. Interestingly,

Ezh1 was even more enriched in the ICM (10.2-fold, Fig. 2C) than Ezh2, raising the

possibility that Ezh1 contributes to preferential H3K27me3 in the ICM.

We observed ICM enrichment for mRNAs encoding members of the Trithorax

Group (TrxG) of factors, including Mll5 (2.16-fold) and Ash1l (2.7-fold, Fig. 2C), that

modify chromatin by methylating H3K4 to increase transcriptional potential and

antagonize PRC2 function on target genes [272,273,274]. The presence of high levels of

both PcG and TrxG proteins, whose functions are opposing, may seem paradoxical; but

TrxG proteins and PcG proteins have crucial functions in pluripotent ES cells in

establishing “bivalent domains” containing both repressive H3K27me3 and permissive

H3K4me3 [101]. Upon differentiation, only one of the marks is retained, allowing for

either epigenetic repression or activation of transcription. Therefore, ICM enrichment of

both PcG and TrxG proteins suggests that the interplay between the two opposing groups

may be more prominent in the pluripotent ICM that will give rise to the developing

embryo proper than in extra-embryonic tissues.

The histone methyltransferases (KMTs) and demethylases (KDMs) are another

class of chromatin regulatory proteins whose transcripts are enriched in the ICM of the

blastocyst. We observed ICM enrichment of transcripts encoding KDMs Jarid1b

(KDM5b, 2.7-fold) and Jarid1c (KDM5c, 2.7-fold), enzymes that demethylate H3K4me

[31] (Fig. 2C). Reduction of trimethylated H3K4, a transcriptionally permissive mark

[275], is likely to contribute to the unique repressive ICM chromatin architecture.

Predictions on the methylation pattern of H3K9 are not as straight forward. ICM

enrichment of transcripts encoding two KMTs of H3K9, Suv39h1 (KMT1a, 3.5-fold) and

Suv39h2 (KMT1b, 4.0-fold), and ICM repression of two KDMs of H3K9me, KDM3a

and KDM3b, indicate that H3K9me3 would be enriched in ICM. However, transcripts

39

encoding two H3K9 demethylases, KDM4c (2.3-fold), whose expression is activated by

Oct4 in ES cells, and LSD1 (KDM1a, 1.9-fold) are enriched in ICM, provide an opposing

prediction about the status of H3K9me in the ICM.

A methyltransferase of H4K20me1, Set7, is required for development to the eight

cell stage in preimplantation embryos, with null mutants showing massive DNA damage

[276]. Transcripts encoding Set7 are enriched in ICM (7.3-fold, Fig. 2C)

The HDAC/mSin3a complex deacetylates histones and is part of the pluripotency

gene network in ES cells [277]. We observed ICM enrichment of mRNAs encoding

components of the Sin3a-HDAC1 histone deacetylase complex (Sin3a, 3.3-fold; HDAC1,

2.4-fold; Sap30 2.0-fold), as well as histone deacetylase HDAC6 (1.9-fold), but not of

HDAC2 (Fig. 2C). This finding corroborates a recent study showing that HDAC1, but

not HDAC2 controls differentiation of ES cells [278]. Enrichment of HDAC complexes

may contribute to more repressive chromatin in the ICM.

Among the mRNAs enriched in ICM were those encoding members of the Snf2

family of ATP-dependent nucleosome remodelers. These include some components of

the BAF complex, Brm (5.1-fold), Baf155 (2.1-fold) and Baf53a (5.7-fold). The BAF

complex exhibits combinatorial assembly of subunits; variations of the BAF complex are

essential at different developmental stages [69,186,187]. Transcripts encoding Snf2l, a

member of NURF chromatin remodeling complex, were enriched in the ICM (12.1-fold)

[279]. Null mutants of BPTF, the largest subunit of the NURF complex, are embryonic

lethal at E8.5, with failure to form visceral endoderm [85]. Analysis in ES cells shows

that NURF is required for proper formation of all three primary germ layers.

Chromatin remodeling factors are often found in large, multisubunit complexes

[265]. Subtle changes in the composition of a complex can have dramatic effects on its

function, and on the differentiation status of a cell [69,280,281]. Enrichment (or

repression) of one or more subunits of a complex is one way in which the composition of

a complex can be regulated [282].

In general, our data support a model in which, compared to the trophectoderm, the

ICM is characterized by a chromatin state with tight transcriptional control and an

abundance of chromatin proteins that mediate transcription and differentiation.

40

Chd1l expression patterns suggest a developmental role

Among the Snf2 family of chromatin enzymes whose mRNAs were enriched in the

ICM was the CRF called Chd1l. Its enrichment score of 4.28-fold was higher than that of

the “master regulator” of pluripotency, Oct4 (1.8-fold, Fig. 2A). The Snf2 family of

chromatin remodeling factors has powerful and diverse roles in development and

transcriptional regulation [280,283], and Chd1l is a member of this family by virtue of

the split DNA-dependent ATPase/helicase domain [81]. Chd1l is the only member of the

Snf2 family that contains a poly(ADP-ribosyl)ation binding macro domain (Fig. 2B)

[257]. As expected, Chd1l expression is observed in human and mouse ES cells (data not

shown).

Our lab previously reported genome-wide gene expression profiles over a time course

of preimplantation development from the zygote through the blastocyst stage [258]. In

these studies, Chd1l expression was found to increase through the first several cell

divisions of development, peaking at the late morula stage (Fig. 2C). Upon formation of

the blastocyst, total Chd1l expression is seen to decrease slightly; our ICM data indicate

it then becomes preferentially expressed in the ICM. Compartmentalization in the ICM,

expression in ES cells, and developmental regulation support a potential role for Chd1l in

pluripotency and during early embryogenesis. In addition to these selection criteria, we

were interested in studying a novel CRF of the powerful Snf2 superfamily (Fig. 2D).

Chd1l is dispensable for ES cell pluripotency and proliferation

Mouse ES cells are derived from the ICM of blastocyst stage embryos and

maintain the property of pluripotency indefinitely. Because Chd1l mRNA is enriched in

the ICM and abundant in ES cells, we asked whether Chd1l is essential for ES cell

survival and pluripotency. To knock-down Chd1l in ES cells, we introduced shRNA-

encoding sequence into the EBRTcH3 ES cell line [284] that allows for stable, Cre-

mediated integration and inducible transgene expression under the control of a CMV

promoter (Tet-Off, Fig. 3A). First, we created a control ES cell line, NS-shRNA

EBRTcH3, by integrating cDNA encoding shRNA that does not target any transcript in

the mouse genome (“Non-Silencing”). Transcription of the shRNA from the CMV

promoter was confirmed by observing robust Venus reporter gene expression 24 hours

41

after inducing expression by Tetracycline withdrawal (“Tet-Off” induction). We created

the Chd1l-shRNA EBRTcH3 ES cell line by integration of sequencing encoding shRNA

that targets the Chd1l transcript. Chd1l protein was reduced to nearly undetectable levels

in Chd1l-shRNA EBRTcH3 cells 48 hours after tetracycline withdrawal (Fig. 3B). In

contrast, NS-shRNA ES cells induced to express NS-shRNA had normal levels of Chd1l.

Chd1l-shRNA ES cells with reduced Chd1l had normal levels of Oct4 expression (Fig.

3B), no obvious abnormalities in ES cell morphology or colony formation (Fig. 3C), and

normal proliferation over a period of eight days, or approximately 10 doublings (Fig.

3D). Our results are consistent with a recent RNAi screen performed in ES cells in which

Chd1l was included a set of chromatin factors screened, but was not found necessary for

ES cell proliferation or for expression of a pluripotency reporter gene [285].

Chd1l does not regulate gene expression in ES cells

A primary function of the SNF2 family of DNA-dependent ATPases is

transcriptional regulation [75,286]. The four major subfamilies, SWI/SNF, CHD, ISWI,

and INO80 all regulate gene expression during development [280]. Chd1l contains a

seven -motif, DNA-dependent ATPase module that defines the SNF2 family of chromatin

remodeling factors as well as a macro domain that recognizes PAR-modified nuclear

proteins, including PAR-modified histones. We hypothesized that Chd1l might also

regulate transcription and that ES cells lacking Chd1l could have transcription changes

even in the absence of obvious morphological changes. We took a whole-genome

approach and obtained the expression profiles of induced (-Tet) EBRTcH3 ES cells

expressing Chd1l-shRNA or NS-shRNA and uninduced (+Tet) ES cells that did not

express shRNA.

Expression indices show ~70% reduction of Chd1l at the transcript level in ES

cells expressing Chd1l-shRNA (Fig. 3E). However, we found only a small number of

transcripts that changed more than 1.4-fold between induced ES cells expressing Chd1l-

shRNA and uninduced ES cells (~30), and these transcripts were also differentially

expressed between induced ES cells expressing NS-shRNA and uninduced ES cells,

indicating the expression changes were a byproduct of inducing shRNA expression. We

found no statistically significant changes in expression of pluripotency markers,

42

differentiation markers, or cell cycling genes (Fig. 3E). Although a very small amount of

remaining Chd1l could in principle be sufficient to maintain normal gene expression

programs, our data suggest that Chd1l does not regulate transcription in ES cells.

Chd1l is not required for differentiation of ES cells

Like the ICM, ES cells are capable of differentiating into the three germ layers.

While ES cells maintain this property indefinitely in vitro, the ICM is only transiently

pluripotent as cells rapidly differentiate during embryogenesis. In the absence of the

pluripotency cytokine LIF, ES cells can be grown into embryoid bodies (EBs),

differentiating cellular aggregates that mimic in vivo post-implantation development. We

reasoned that Chd1l may not regulate gene expression in pluripotent ES cells, but may do

so in differentiating cells, when new gene expression patterns are being established. To

ask whether Chd1l is required for the formation of the germ layers, we knocked down

Chd1l in Chd1l-shRNA ES cells then differentiated them into EBs and observed the

expression of a panel of lineage markers by q-rtPCR over multiple time points. For

comparison, we measured gene expression in EBs made from induced and uninduced

NS-shRNA ES cells. Quantitative rt-PCR confirmed ~70% knock-down of Chd1l

mRNA in induced Chd1l-shRNA EBs but not in induced NS-shRNA EBs over nine days

of differentiation (Fig. 3F). The panel of lineage markers included genes associated with

the establishment of endoderm (Sox17, AFP, Gata4), mesoderm (Lhx1), and ectoderm

(Fgf5, Otx2), as well as pluripotency (Oct4) and extra-embryonic (Eomes) tissues. EBs

expressing Chd1l-shRNA reduced Oct4 expression in a manner similar to EBs expressing

NT-shRNA (Fig. 3F). Expression of markers for all three germ layers was induced in a

temporally appropriate manner (Fig. 3F). Our results indicate Chd1l does not control

gene expression in pluripotent ES cells or in differentiating embryoid bodies under

normal culture conditions.

Chd1l transcripts are abrogated in MO-injected embryos

Next, we addressed the question of whether Chd1l plays a role in development

prior to differentiation of the ICM. The preimplantation embryo can be cultured in vitro

through the blastocyst and hatching stages. We took a rapid knock-down approach,

43

utilizing the synthetic antisense oligos called morpholinos (MOs) that inhibit translational

and splicing machinery. MOs have been used to effectively reduce production of specific

proteins in the preimplantation embryo, with minimal toxicity or off-target effects [287].

Splice-blocking MOs were designed to target Chd1l pre-mRNA. The predicted

splice mutants produce truncated proteins due to stop codons within the intron (Fig.4A).

Chd1l MO-1 was microinjected into the cytoplasm of one-cell stage mouse embryos

collected from superovulated and mated females. To confirm that MO-1 was functioning

as predicted, we used microfluidic q-rtPCR on RNA collected from single MO-injected

and control embryos. We used a TaqMan primer-probe assay that targeted the junction

between exons 2 and 3. This junction would be present in the wild-type Chd1l transcript

but absent if the MO blocks its targeted splicing event. As expected, the amplification of

the splice junction was nearly absent in injected embryos, showing that the transcript is

abrogated (Fig. 4B and C). A second splice-blocking MO targeting Chd1l (MO-2)

impaired a separate splicing; qPCR amplification of the splice junction showed reduction

greater than 99% compared to uninjected embryos (Fig. 4C). Using either MO, the

altered splicing would lead to the introduction of a stop codon within the intron, and

consequently the only products would be mutant proteins truncated near the N-terminus

prior to production of any functional domains.

Embryos injected with Chd1l-targeting MOs arrest prior to blastocyst stage

To ask whether Chd1l is required during early development, we microinjected the

zygote-stage embryo with MO-1 targeting Chd1l and observed embryos for a period of

four days. MO-injected embryos did not reach the blastocyst stage and instead arrested at

the compaction stage (Fig. 5A and B). An arrest prior to blastocyst formation is

consistent with the peak in Chd1l expression at the late morula stage and enrichment in

the ICM. In contrast, the majority of embryos microinjected with MO that targets

another Snf2-like chromatin remodeling factor, Snf2l, reached the blastocyst stage (Fig.

5A and B). This result demonstrates that embryonic arrest upon microinjection of a CRF

MO is not a general effect.

To further test our finding, we microinjected a second Chd1l MO that targets a

different splice junction (MO-2). These embryos also arrested prior to the blastocyst

44

stage (Fig. 5A and B). The precise timing of the arrest varied between the MOs, perhaps

due to different binding affinities of the MO sequences.

Chd1l phenotype is partially rescued by co-injection of Chd1l mRNA

To confirm that the embryonic arrest phenotype is a result of disrupting Chd1l

protein production, mRNA encoding Chd1l was co-injected along with Chd1l MO. We

reasoned that embryos arrested at an earlier stage would be more able to progress to later

developmental stages with addition of mRNA than embryos arrested at later stages, so we

used MO-2 for co-injection. MO-2 targets a splicing junction and therefore will not

target the mRNA synthesized from cDNA lacking intronic sequence. Figure 5C shows

the average result from three independent rescue experiments. Embryos injected with

MO-2 alone did not progress to the multi-cell stage, nor did embryos co-injected with

MO-2 plus GFP mRNA. As many as 50% of the embryos co-injected with MO plus

Chd1l mRNA progressed to the multi-cell stage or further. Mitigation of the

developmental arrest phenotype by Chd1l mRNA confirms that loss of Chd1l is

responsible for the embryonic arrest.

Discussion

A previous study attempted to identify pluripotency factors by comparing

expression profiles of cultured ES cells and trophoblast stem (TS) cells, derived from the

ICM and TE, respectively [288]. Our approach has the significant advantage of using

true embryonic cells that retain in vivo gene expression programs. Many genetic studies

in the preimplantation embryo focus on genes known a priori to be essential for ES cell

viability or pluripotency. We used gene expression data from early embryos to select

candidate regulators.

Chromatin remodeling activities are abundant in preimplantation embryos and in

ES cells, and many of these activities are geared toward initiating pluripotent

transcriptional competence and ensuring differentiation programs are locked in

epigenetically [209,234]. Although Chd1l is part of the Snf2 family of DNA-dependent

ATPases [81], many of which are potent transcriptional regulators, Chd1l itself does not

seem to regulate gene expression, at least in ES cells.

45

Chd1l is distinct from the other members of its family because of the presence of

a C-terminal macro module that binds poly ADP-ribose, or PAR [61,259,260]. PAR is a

posttranslational modification synthesized by the PARP family of PAR polymerases and

has important roles in diverse chromatin-dependent processes. Evidence is accumulating

for the importance of PAR regulation in the embryo. Double knock-out of PAR

polymerases Parp-1 and the partially redundant Parp-2 in mice is embryonic lethal at the

onset of gastrulation [57], whereas knock-out of the PAR depolymerase PARG is lethal at

E3.5 [289]. These data suggest that PAR levels are tightly regulated in the embryo, and

that fluctuations are highly deleterious. Chd1l contains a module responsible for binding

PAR. It is interesting to speculate that Chd1l contributes to PAR regulation and that the

Chd1l embryonic arrest phenotype is due to aberrant PAR levels or PAR signaling.

One of the most prominent trigger of PAR modification is DNA damage. Parp-1

is activated by DNA damage and synthesizes PAR onto itself in an automodification

reaction [290,291]. Blocking Parp-1 activity with specific inhibitors or through null

mutations results in cellular hypersensitivity to DNA damaging agents and defects in

DNA repair [292]. Two independent groups recently demonstrated the ability of Chd1l

to respond to DNA damage through its association with PAR [259,260]. The early

embryo has unique and stringent requirements to repair any errors to maintain genomic

integrity for the future organism. Damage to DNA occurs as a result of normal cellular

metabolism and during DNA replication. Genes involved in all of the major DNA repair

pathways are expressed in the preimplantation embryo [253], and a large number of

double-stranded break repair proteins are embryonic lethal when deleted [293].

In particular, DNA repair within the zygote is especially crucial for the paternal

genome that must recover from the “frozen” state of the sperm [240,294]. Defects in

repairing paternal DNA are thought to be a major cause of chromosomal aberrations and

human infertility. Double-stranded breaks are the most toxic form of DNA damage and

can be repaired through either non-homologous end-joining (NHEJ) or homologous

recombination (HR). NHEJ can function throughout the cell cycle, whereas HR is

restricted to S/G2 phase [295]. The zygote spends ~20 hours in G1 prior to the first cell

division, and much of the paternal DNA is repaired through NHEJ [296,297,298,299].

46

The ability of Chd1l to function in NHEJ is suggested by its PARP-dependent association

with a major NHEJ component, DNA-PKcs, upon induced DNA damage [259].

An intriguing question is why is Chd1l essential in the earliest stages of

embryogenesis but not in ES cells? In contrast to the zygote, ES cells have rapid cell

cycles with abbreviated G1 and G2 gap phases and rely heavily on HR to repair lesions

during S phase [126]. Therefore, one explanation for why reduced Chd1l causes

preimplantation arrest but is dispensable for ES cells is that Chd1l plays a role in NHEJ,

and NHEJ is essential during early embryogenesis but not in ES cells.

PAR modifications are involved in the regulation of transcription as well as DNA

repair and other processes [300]. While we cannot rule out a role for Chd1l in

transcriptional regulation in the embryo through its interaction with PAR, given our data

in ES cells showing that Chd1l is not involved in regulation of gene expression, we

suggest that the embryonic arrest due to reduction of Chd1l is more likely to be due to a

defect in DNA repair.

In summary, chromatin remodeling enzymes are active in the ICM of the

blastocyst that will differentiate into all the cellular lineages of the adult organism. One

candidate regulator of pluripotency, Chd1l, turned out to be required even earlier than the

formation of the ICM. Despite its essential role in the preimplantation embryo and

expression in ES cells, Chd1l is dispensable for ES cell viability, pluripotency, or

differentiation. Chd1l contains a macro domain that binds to PAR, and the Chd1l arrest

phenotype could be due to impaired PAR signaling. Recent studies have demonstrated a

role for Chd1l as a DNA damage response protein that interacts with members of the

NHEJ pathway in a PARP-dependent manner. A role for Chd1l in NHEJ could explain

why Chd1l deficiencies in the preimplantation embryo result in an early arrest phenotype

whereas deficiencies in ES cells result in no detectable abnormalities.

Methods

Immunosurgery and expression profiling

E3.5 blastocysts were collected from timed pregnant mothers and washed in M2

medium. The zona pellucida was removed by incubation in Acid Tyrode solution for 3

minutes. Outer TE cells were labeled with IgG’s by incubation with 10% rabbit α-

47

mouse serum for 60 minutes. Embryos were washed three times in M2 medium, and then

TE cells were lysed through the complement cascade by incubation with 30% guinea pig

complement for 15-30 minutes, or until lysis was visible. Remaining ICMs were washed

three times in M2 medium with a fine pipette to remove residual TE cells. Total RNA

was extracted from purified ICMs and whole blastocysts with Trizol reagent. Purified

RNA was amplified, labeled, and hybridized to Affymetrix mouse 430 2.0 Expression

Arrays.

ES cell lines

The EBRTcH3 cell line contains a cassette acceptor utilizing loxP and loxPV sites at

the Rosa locus to allow efficient and directional integration of a transgene by Cre-

mediated recombination. ShRNA-mir cDNAs were subcloned from pGIPZ vectors

(OpenBiosystems, Chd1l shRNA Oligo ID: V2LMM_18041 and “non-silencing”

shRNA-mir) into the pPthC exchange vector for recombination into the EBRTcH3 ES

cell line. The parental EBRTcH3 ES cells and the pPthC exchange vector were gifts

from the lab of Dr. Hitoshi Niwa of Japan.

The exchange vector containing the shRNA-mir sequence was cotransfected along

with a Cre expression plasmid by lipofectamine. Transfected cells were plated single-cell

density and cultured in the presence of Puromycin (1.5 µg/ml) to select for successful

integrants and of Tetracycline (1.0 µg/ml) to repress transgene expression. Clones were

confirmed by PCR genotyping of the 5’ and 3’ recombination sites. To induce shRNA

expression, the derived ES cell lines were cultured in the absence of Tetracycline and

high Puromycin (7.5 µg/ml). Control, uninduced ES cells were cultured in high

Tetracycline (1.5 µg/ml) and high Puromycin (7.5 µg/ml).

ES cell expression profiling

Total RNA was extracted from Chd1l-shRNA and NS-shRNA ES cells three days

after inducing the expression of Chd1l-shRNA or NS-shRNA by Tetracycline removal

(7.5 µg/ml Puromycin), and from uninduced Chd1l-shRNA and NS-shRNA ES cells that

do not express shRNA (7.5 µg/ml Puromycin, 1.5 µg/ml Tetracycline). Three different

Chd1l-shRNA EBRTcH3 clones and one NS-shRNA EBRTcH3 clone were used. RNA

48

was amplified from the eight samples and labeled using Affymetrix kit and hybridized to

mouse 430 2.0 Expression Arrays. Fold changes in expression indices were calculated

for shRNA-induced ES cells vs. shRNA-uninduced ES cells. Statistical significance of

fold-changes between Chd1-shRNA induced and uninduced samples was carried out

using a paired t-test, a minimum fold change of 1.4, and a delta value of 1.9 (SAM

Analysis [301]).

Differentiation of embryoid bodies

Expression of shRNA was induced by Tetracycline withdrawal in Chd1l-shRNA

and NS-shRNA EBRTcH3 ES cells for three days prior to differentiation into embryoid

bodies (EBs) to ensure complete Chd1l knock-down. RNA was collected at “Day 0” of

differentiation from induced and uninduced Chd1l-shRNA and NS-shRNA EBRTcH3 ES

cells. ES cells were suspended at a density of 2x104 cells/ml of media without LIF, and

EBs were made using hanging droplets of 500 cells in 25 µl. After two days, embryoid

bodies were collected into 10-cm Ultralow Attachment plates (Corning) and cultured for

an additional seven days in the absence of LIF. RNA was collected every three days after

LIF removal. cDNA was synthesized from each sample and subjected to qPCR. Relative

quantities for each cell line were calculated using Gapdh as the internal control and

shRNA-uninduced, “Day 0” samples as references.

Embryo Culture and microinjection

Three to five week old wild-type F1 (C57BL6xDBA/2) females (Charles Rivers)

were superovulated by intraperitonial injections of 5 IU of pregnant mare’s serum

gonadotropin (Sigma) followed by 5 IU of human chorion gonadotropin (Sigma) 48

hours later and mated with wild-type males. Mice were sacrificed by cervical dislocation

17 hours after hCG injection, and 1-cell embryos were dissected and released from

oviducts. Cumulus cells were removed hyaluronidase digestion, and embryos at the two

pronuclei stage were recovered and immediately micro-injected cytoplasmicly with 5-10

pL of 0.6 mM antisense morpholino. Prior to injection, the MO was heated at 65° for 15

minutes to remove any secondary structure.

49

Preimplantation embryos were cultured in vitro in 20 µl droplets of Quinn’s

Advantage Cleavage Medium (Sage) supplemented with 10% SPS serum and covered

with mineral oil. Dishes were placed in a dessicator filled with mixed gas (90% nitrogen,

5% oxygen, 5% carbon dioxide) in a 37° incubator. Embryos were observed every 24

hours for a period of four days.

Morpholinos were obtained from GeneTools. Chd1l MO-1:

tcattccacagcagatacCTGGCAG (in2-EX2). Chd1l MO-2: ttggagagaagcagagggctaCCTC

(in4-EX4). Snf2l: tgctgtttaccaccttacCAAGGGC (in2-EX2).

Microfluidic qPCR

Single embryos were collected 48 hours after injection and lysed by one freeze

thaw cycle. cDNA was synthesized using the CellsDirect One-Step rtPCR kit

(Invitrogen) and subjected to 18 rounds of gene-specific amplification using TaqMan

primer/probe assays (Applied Biosystems). TaqMan primer/probe assays and cDNA

from single embryos were loaded onto a Fluidigm 48.48 microfluidic array for qPCR

analysis using the Biomark thermalcycler.

α-Chd1l antibody

A hydrophilic sequence of 122 aa corresponding to amino acids #557-678 of the

Chd1l protein was selected for the antigenic region. The antigen was produced as a TrpE

fusion protein from the pATH11 vector in BL21 E. coli, solubilized, and subjected to

SDS-PAGE. The gel slice was excised and submitted to Josman, LLC for injection into

rabbits. The antiserum was affinity purified over a GST-Chd1l-bound Sepharose column

and eluted with low pH buffer.

50

Figure 2.1 Chromatin remodeling factors are enriched in the ICM

A. Schematic of immunosurgery followed by whole-genome expression analysis. B.

Compartmentalization of known ICM and trophectoderm factors.

C. Enrichment of selected classes of chromatin factors in the ICM. TF: transcription

factor. DNMT: DNA methyltransferase. PcG: Polycomb group. ETP: Enhancers of

Trithorax and Polycomb. TrxG: Trithorax group. KMT: lysine (histone)

methyltransferase. KDM: lysine (histone) methyltransferase. HDAC: histone

deacetylase.

A B

C

51

Figure 2.2. Chd1l is a candidate developmental regulator

A. Chd1l is enriched in the ICM compared to the whole blastocyst. B. Chd1l is a

SNF2 chromatin remodeling enzyme containing a split ATPase/helicase and a macro

domain. C. Chd1l expression during pre-implantation development [258]. Chd1l

expression peaks at the late morula stage before it becomes compartmentalized in the

inner cell mass of the blastocyst. D. Decision tree for choosing Chd1l.

A B

C

ICM Enrichment

Chromatin Remodeling

Expressed in ESCs

Dynamic PreimplantationExpression

Unknown Developmental Role

Chd1l

D

900 aa

52

0

50

100

150

200

250

300

350

Day 0 Day 3 Day 6 Day 9

Lhx1

0

10

20

30

40

50


Gata4

0

5

10

15

20


Otx2

0

20

40

60

80

100


Sox17

0

1

2

3

4

5

6


Eomes

0

500

1000

1500

2000

2500


Fgf5

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8


Oct4

0

500

1000

1500

2000

2500


AFP

0

0.5

1

1.5


Chd1l

A

E

F

C

B

D

-Tet

+Tet

53

Figure 2.3. Chd1l is non-essential in ES cells

A. Strategy for knocking-down Chd1l in EBRTcH3 ES cells. The tetracycline

transactivator (tTA) is expressed from the endogenous Rosa26 locus and is bound in the

inactive form in the presence of tetracycline. In the absence of tetracycline, the tTA

activates the CMV promoter and induces expression of the shRNA-IRES-Venus

transcript. B. Efficiency of knocking down Chd1l in uninduced (+Tet) and induced (-

Tet) Chd1l-shRNA EBRTcH3 cells. Chd1l protein levels are nearly undetectable in

shRNA-expressing cells 48 hours after Tetracycline withdrawal. Oct4 levels do not

change upon knock-down of Chd1l. C. Colony morphology of Chd1l-shRNA EBRTcH3

cells with induced (+Tet) or uninduced (+Tet) shRNA expression. Expression of shRNA

was induced 24 hours prior to plating cells at clonal density and allowing colonies to

grow for six days. D. Proliferation curve of Chd1l-shRNA ES cells expressing Chd1l-

shRNA (-Tet) or uninduced (+Tet). E. Expression changes of selected genes from

global expression analysis from ES cells expressing Chd1l-shRNA or NS-shRNA. F.

Expression of lineage markers over embryoid body (EB) differentiation. All values are

normalized to uninduced “Day 0” samples from each respective cell line. Similar to EBs

expressing NS-shRNA, EBs expressing Chd1l-shRNA are able to differentiate and form

the three germ layers as evidenced by markers for pluripotency (Oct4), extra-embryonic

(Eomes), endoderm (Gata4, AFP, Sox17), mesoderm (Lhx1), and ectoderm (Fgf5, Otx2).

54

Figure 2.4. Chd1l MO knock-down

A. Mechanism of splice-blocking morpholinos (MOs). Two splice-blocking MOs were

designed targeting exon-intron boundaries. Both are predicted to produce mutant

proteins truncated prior to the ATPase domain, thus lacking any functional activity. B.,

C. Validation of Chd1l MO activity. Heat map (A) and quantitation (B) of microfluidic

qPCR of Chd1l transcripts. Ct values for each PCR reaction were subtracted from an

arbitrary value of 40 to reflect a positive correlation with expression levels. The “Ex20-

21” probe targets the 3’ end of the Chd1l transcript and will amplify all Chd1l transcripts,

regardless of splicing aberrations. The “Ex2-3” and “Ex4-5” probes target exon-exon

junctions and will only amplify if that splicing event occurs. MO-1 disrupts exon2-exon3

splicing. MO-2 disrupts exon4-exon5 splicing.

C

B

. . .

Predicted Mutant Protein Pre-mRNA

A

55

B C

A

A

56

Figure 2.5. Chd1l embryonic arrest phenotype

A. Uninjected embryos and embryos injected with indicated MOs at 4 days after

microinjection. B. Quantification of development to blastocyst stage in uninjected

embryos and embryos injected with different MOs. C. Partial rescue of Chd1l arrest

phenotype with co-injection of Chd1l mRNA.

57

3. Finding Direct Transcriptional Targets of Chd1l in ES Cells

Introduction

Specification of a totipotent or multipotent cell into more differentiated daughter

cells is associated by differences in gene expression and epigenetic alterations in

chromatin. At the inception of this thesis project, I was intrigued by the earliest

differentiation choice a cell makes: to become trophectoderm (extra-embryonic) or inner

cell mass (embryonic); and later, to depart from the pluripotent state and become one of

the three primary germ layers: endoderm, mesoderm, or ectoderm. There is debate within

the field regarding the primary event that drives differentiation, whether it is the genetic,

transcriptional activity of transcription factors that activates downstream genes including

histone modifying proteins, or the epigenetic activity of enzymes that post-translationally

modify histones and subsequently activate or repress the expression of genes, including

transcription factors [102]. Regardless of the victor in the circular “chicken-or-the-egg”

conundrum, it is clear that feedback loops exist and that both types of regulation are

crucial in establishing and maintaining gene networks.

A large body of literature has established that control of gene expression through

modification of nucleosome dynamics is a powerful way in which chromatin remodeling

enzymes mediate differentiation. The first chromatin remodeling factor shown to

mediate gene expression was the yeast Swi/Snf that was identified in genetic screens for

mutants of mating-type switching (SWI) and sucrose fermentation (Sucrose-Non-

Fermenting) [83,84]. Since then, a number of proteins containing a conserved SNF2-like

DNA-dependent helicase/ATPase have been discovered in mammals that also have

powerful functions in regulating gene expression during development [66,280].

Although the SNF2-like domains show similarity to the prototypical helicase domain,

they do not actually possess helicase activity [75,76]. Rather, this class of proteins

functions by interacting with the minor groove of the DNA double helix and inducing

torsional forces that cause strand distortion and interfere with the DNA-histone interface

58

[94]. SNF2-like chromatin factors harnesses the energy of ATP hydrolysis to force

movement of histone cores along DNA, exposing regions of DNA to a variety of nuclear

factors [76]. Thus, ATP-dependent chromatin remodeling factors counteract compacted

chromatin and can drive changes in gene expression by providing either transcription

factors or histone modifying enzymes (or both) access to chromatin.

Chd1l is developmentally regulated during in the preimplantation embryo, is

preferentially expressed in the inner cell mass of the blastocyst, and is expressed in ES

cells. As such, it became the candidate developmental regulator studied in this

dissertation. Because of homology to the SNF2-like family of chromatin remodeling

enzymes, I hypothesized that Chd1l is also a chromatin remodeling enzyme that regulates

gene expression. The goal of this project was to find direct targets of Chd1l through

chromatin immunoprecipitation followed by unbiased, whole-genome sequencing (ChIP-

seq). To determine functional relevance, I endeavored to find Chd1l-bound genes whose

expression depends on Chd1l. In parallel with ChIP-seq analysis, I performed whole-

genome expression analysis on ES cells that had normal or reduced levels of Chd1l

protein (see Chapter 2). My approach was to use the intersection of ChIP-seq data and

expression analysis to identify genomically bound and transcriptionally responsive gene

targets of Chd1l.

The purpose of this project was two-fold. The first was to identify a role for

Chd1l in ES cells, despite the lack of an obvious morphological phenotype in culture.

Gene ontology clustering of transcriptional targets can reveal roles in various cellular

processes. Enrichment of target genes involved in one process would implicate a role for

Chd1l in that process. The second purpose was to use the list of target genes in ES cells

to identify a role for Chd1l in the preimplantation embryo, where Chd1l is essential.

Because of the requirement for large amounts of starting material for ChIP-seq

experiments, procuring for de novo targets on a genome-wide scale from preimplantation

embryos that contain at most several hundred cells was not possible. The list of Chd1l-

bound genes in ES cells (regardless of expression changes in ES cells upon Chd1l knock-

down) would be scanned for genes having crucial roles in the preimplantation embryo.

These genes would then serve as candidates to be tested in early embryos using standard

PCR-based ChIP experiments that require much less input material.

59

A major hurdle of this project was the absence of previously identified Chd1l

target genes to confirm that my in-house generated Chd1l antibody is capable of

immunoprecipitating cross-linked chromatin. Indeed, the results of this experiment

yielded very few target genes, and none of the candidates selected could be validated by

q-PCR. It is difficult to differentiate between the explanations that the antibody did not

work for ChIP or that Chd1l truly does not bind distinct loci in ES cells. This appendix

documents the experimental procedures and analysis undertaken in the attempt to find

Chd1l gene targets.

Results

Chromatin Immunoprecipitation

To confirm that the lab-generated α-Chd1l antibody had specific recognition for

Chd1l, ES cell extracts with and without knock-down of Chd1l were subjected to SDS-

PAGE and blotted with α-Chd1l (Fig. 1A). A strong band at the predicted size of ~100

kD was apparent in ES cells with Chd1l, and that band became nearly undetectable in ES

cells expressing Chd1l-shRNA. Production of other genes including β-tubulin and Oct4

did not change. The ability of the α-Chd1l antibody to efficiently immunodeplete Chd1l

from supernatants was confirmed in ES cell extracts (Fig. 1B). Association of Chd1l

with chromatin was confirmed by cell fractionation assays (Fig. 1C).

Two ChIP experiments were conducted in parallel: one using α-Chd1l antibody

and a second using α-RNA PolII antibody. The RNA PolII ChIP was used for quality

control, since this antibody has been extensively used in ChIP experiments and PolII has

many known targets. A negative control ChIP was avoided because sequencing uses

total, genomic chromatin as a reference, and a negative control antibody would pull down

and amplify a very small fraction of the genome.

ES cells were cultured to subconfluence, crosslinked, and lysed. Chromatin was

sheared to small fragment sizes that are ideal for sequencing (between 100 and 500 base

pairs) and then immunoprecipitated by either α-Chd1l or α-PolII. The

immunoprecipitated DNA was purified and quantified. The amount of purified DNA for

PolII ChIP was approximately 4 times greater than for Chd1l ChIP (1.14 µg vs 0.25 µg),

indicating that in general, Chd1l binds fewer targets or binds targets with lower affinity

60

than PolII. Given that PolII is necessary for general transcription, this was not overly

concerning.

The immunoprecipitated and total chromatin DNA libraries were prepared for

sequencing by adapter ligation and amplification. DNA size, purity, and concentration

were acceptable for all samples. Prior to submitting samples for sequencing, the PolII

library was validated by PCR amplification of known targets (Fig. 1D). Confirmation of

pull down of PolII targets shows that protocol procedures were enacted accurately. The

Chd1l library could not be validated in the absence of previously identified targets.

Sequencing and Analysis

DNA libraries for PolII ChIP, Chd1l ChIP, and total genomic chromatin were

submitted to the laboratory of Dr. Arend Sidow for Solexa sequencing. Raw reads were

passed through a quality filter that excludes short reads or reads with ambiguous base

calling and mapped to the mouse genome. After filtering out repetitive reads,

approximately 4 million quality reads were obtained for each sample (Table 1). For

Solexa sequencing, which generates reads of about 25 base pairs, this corresponds to

roughly 3% coverage of the mouse genome, assuming uniform distribution. The

distribution of reads around the transcription start sites (TSSs) was measured (Fig. 2).

This analysis further validates PolII ChIP library, as PolII is known to bind at TSSs. The

Chd1l ChIP reads show a modest enrichment around TSSs, but only slightly above that

seen for total chromatin reads. In general, chromatin shears more easily at TSSs,

therefore some accumulation of reads there is expected.

Non-repetitive reads mappable to the mouse genome were delivered to the

laboratory of Dr. Wing Wong who used CisGenome for peak calling software and

genome browser [302]. The software analyzes forward and reverse reads separately to

form bi-horned normal distributions around factor binding sites. A peak, or “hit,” is

called between the modes of each distribution of forward and reverse reads (e.g. Fig. 3A).

Total genomic chromatin was used for background correction.

The number of peaks called for PolII ChIP and Chd1l ChIP for three different

false-discovery rates (FDRs) is shown in Table 2. At a FDR of .05, ~200 peaks were

called for Chd1l compared to almost 18,000 peaks for PolII. The robust number of peaks

61

for PolII and the location of peaks at actively transcribed genes show that the procedures

followed for ChIP, sequencing, and analysis were sound. The number of Chd1l peaks

was surprisingly low, and resembled ChIP-seq data for antibodies that do not ChIP well

(personal communication). Approximately 30-40% of Chd1l peaks were associated with

nearby PolII peaks (Fig. 4D). This association can reveal potential interaction of a

transcription factor or chromatin remodeling factor with PolII, an interaction that

suggests a role in gene expression. The caveat is that regions of active transcription and

PolII association are also more easily sheared, and sheared ends will result in an

accumulation of sequencing reads and peak artifacts.

Twenty-seven hits were manually selected for validation by quantitative PCR.

Many of the highest-ranking hits were excluded because of excessively high signal in the

background (e.g. Fig. 4B). Only those hits with minimal background were selected (e. g.

Fig. 4C). While peaks for PolII ChIP had maximum values around 20, maximum peak

values for Chd1l peaks tended to be small, on the order of 4 or 5. A subset of the 27

regions were associated with nearby PolII peaks, and qPCR of these regions from PolII

ChIP samples showed enrichment as expected (Fig. 4D). Regions where no PolII peaks

were called did not show PolII enrichment by qPCR. In contrast, none of the Chd1l peak

regions selected showed enrichment in Chd1l ChIP samples. These results demonstrate

that gene targets for PolII, but not for Chd1l, were successfully identified by ChIP-seq.

Re-analysis using deeper sequencing

In order to get deeper sequencing and peaks of higher confidence, the Chd1l ChIP

library was re-sequenced. A similar number of reads was obtained for the second run as

for the first. The second set of reads was combined with the first set of reads for analysis

by CisGenome. The combined analysis yielded about 200 peaks at a FDR of 0.01, a

number greater than that obtained either individual analysis (Table 3). Surprisingly, no

peaks called were common between the first and the second set, and the peaks called for

the combined analysis were largely distinct from either individual set (Fig. 4A). In

addition, the appearance of the peaks called for the combined analysis was similar to the

peaks called for the first analysis, with a peak height of about 4 or 5 (Fig. 4B). This

pattern is representative of reads randomly distributed, or enriched at easily sheared loci

62

such as TSSs, or enriched by amplification bias. It was decided at this point to

discontinue Chd1l ChIP-seq experiments.

HA-tagged Chd1l targeting vector

At the inception of Chd1l ChIP-seq experiments, it was anticipated that a second

set of data generated from the use of a separate antibody would be required to increase

the confidence of Chd1l target genes. To address this, a targeting vector was created that

would introduce an HA tag onto the 3’ end of Chd1l (Fig. 5). An antibody against HA

could then be used for ChIP-seq in ES cells that expressed tagged Chd1l at endogenous

levels. The data generated from α-Chd1l ChIP and α-HA ChIP would then be compared,

and the gene targets common to both sets would be selected for further analysis. The

generation of the targeting vector was done in parallel with the α-Chd1l ChIP-seq and

expression analysis studies; recombination into ES cells was not pursued when it became

apparent that there were no changes in gene expression upon Chd1l knock-down and no

high-confidence gene targets from α-Chd1l ChIP-seq.

Discussion

ChIP with an α-Chd1l antibody followed by whole-genome Solexa sequencing

and analysis yielded a small list of 200 peaks with a FDR of 0.10. Quantitative PCR

amplification of 27 genomic regions failed to validate any candidate peak. Two

explanations cannot be resolved: a) the α-Chd1l antibody does not work in ChIP

experiments, and b) Chd1l does not bind to distinct chromatin targets in ES cells.

However, given that there were no significant changes in gene expression upon reduction

of Chd1l levels in ES cells, the latter explanation is likely. For this reason, ChIP-seq

from stably transfected, HA-tagged Chd1l from ES cells using an α-HA antibody was no

longer pursued.

After completion of this project, Ahel et. al. reported that treatment of 293T cells

with hydrogen peroxide, a DNA damaging agent, caused mobilization of Chd1l to

chromatin [259]. In addition, the composition of the Chd1l-containing complex is altered

upon hydrogen peroxide treatment [259,260], indicating recruitment of Chd1l to

chromatin is different between treated and untreated cells. Cell fractionation assays

63

conducted in this study show Chd1l in both nucleosol and in the chromatin-bound

fraction. It is plausible that treatment with hydrogen peroxide treatment of ES cells

would induce Chd1l to become more tightly bound to chromatin and that direct targets

could then be found by ChIP-seq. This avenue of study was not pursued because of time

constraints and the lack of evidence that Chd1l contributes to DNA repair in ES cells

(See Chapter 3).

Recently, Chen et. al. reported the identification of putative CHD1L target genes

in a human hepatocellular carcinoma (HCC) cell line, one of which was associated with

gene expression changes upon modulation of CHDL levels in HCC cells. Their approach

differed from the one described here in several regards. GFP-tagged CHD1L was

transfected and over-expressed in HCC cells and CHD1L was immunoprecipitated with a

GFP antibody. DNA immunoprecipitates were cloned into a vector and identified by

sequencing. The finding that CHD1L binds target genes in this study is unconvincing for

several reasons. The over-expression of tagged CHD1L is likely to promote non-specific

binding to chromatin. The cloning-based strategy employed by the authors does not

account for sticky, non-specific binding, either between over-expressed CHD1L and

chromatin, or between GFP antibody chromatin DNA. PCR validation of targets was not

quantitative or semi-quantitative, and the functional relevance of CHD1L binding to

targets (e.g. through reporter assays) was not attempted. Therefore, although the authors

report the identification of CHD1L gene targets, severe experimental flaws bring this

finding into doubt.

The inability to find direct targets in ES cells does not preclude a transcriptional

role for Chd1l in the preimplantation embryo, where a strong arrest phenotype results

from injection with antisense morpholinos targeting Chd1l. No proliferation,

morphological, or gene expression aberrations were seen in ES cells with reduced Chd1l.

Chromatin remodeling factors are known to bind to distinct sets of targets in different cell

types, and it remains possible that Chd1l binds distinct targets in early embryos but not in

ES cells. However, until genome-wide, micro-scale ChIP experiments become more

practical, discovering targets de novo in the early embryo will remain challenging.

It is also likely that Chd1l exerts its function in the preimplantation embryo not

through transcriptional regulation, but through the modulation of other nuclear processes.

64

ATP-dependent chromatin remodeling enzymes are utilized in a broad spectrum of

nuclear processes including homologous recombination, sister chromatin separation

during cell division, and DNA repair [283]. A role for Chd1l in the DNA damage

response has been confirmed by two independent groups, although a direct role in DNA

repair has yet to be confirmed. If the composition and function of a Chd1l-containing

complex is preserved throughout development, then the Chd1l phenotype of

preimplantation embryos will not be answered at the transcriptional level.

Methods

Antibodies

A polyclonal rabbit Anti-Chd1l antibody produced in our lab was used for

immunoprecipitation and ChIP experiments. A polyclonal rat α-Chd1l antibody, also

raised in-house, was used for immunoblotting of Rb α-Chd1l immunoprecipitates. The

RNA PolII antibody (clone 8WG16) is a mouse monoclonal antibody from Covance.

Chd1l Immunoprecipitation

For each IP, 120 µg of ES cell extract was used with 50 µl of Protein G magnetic

beads (Dynal) pre-conjugated to varying amounts of α-Chd1l (5 µl, 25 µl).

Immunoprecipitation was carried out over night at 4° in RIPA buffer. Beads were

washed the following day and eluted with 50 µl Laemeli buffer. 10% of the supernatant

and 10% of the eluate were loaded onto SDS-PAGE gel for analysis.

Cell Fractionation

ES cell nuclei were separated from a cytoplasmic fraction by lysing mild

detergent (0.1% Triton) and brief, gentle centrifugation. Nuclei were ruptured by

incubation in hypotonic buffer. The nucleosol fraction and chromatin pellet were

obtained by centrifugation.

Chromatin Immunoprecipitation and library preparation

ChIP experiments were performed essentially as described [303]. Feeder-free,

parental EBRTcH3 ES cells [284] were grown to subconfluence on 15-cm plates. Each

65

ChIP library was derived from four pooled ChIP reactions of 2x107 cells each, and the

total chromatin library contained DNA from 2x107 cells. Cell viability was checked

(>95% viability) by Trypan blue staining prior to crosslinking with 1% formaldehyde for

20 minutes. Cells were collected by scraping and lysed at a density of 4x107 cells/ml.

Chromatin was sheared to an average size of 250 base pairs by sonicating with a 3mm

stepped microtip for 1 pulse at 50% power and 6 pulses at 60% power. For each IP,

lysate containing the equivalent of 2x107 cells was incubated with 50 µl of Protein G

magnetic beads (Dynal) pre-conjugated with primary antibody (5 µg of α-PolII and 50 µl

of α-Chd1l). Beads were washed, crosslinks were reversed, and DNA was purified by

phenol-chloroform extraction. DNA libraries from the three samples were all of

acceptable purity (A-260/A280 ratios between 1.8 and 1.9).

Adapter oligos (Illumina) were then ligated to the ends of DNA

immunoprecipitates and sheared total, genomic DNA and subjected to 20 cycles of

amplification using primers to the adapter oligos. Fragments of 100 to 300 base pairs

were size selected by gel electrophoresis, subjected to a second round of 18 amplification

cycles, and purified again by phenol-chloroform extraction.

Recombineering of Chd1l-HA Targeting Vector

Approximately 9 Kb of genomic DNA containing the last three exons of the

Chd1l gene and downstream sequence was “captured” from a BAC clone (RP24-273N16,

BAC PAC Resources) by recombineering [304] into a pBluescript vector in SW105 cells

(NCI-Frederick). Intermediate constructs containing two short homology arms (~500

base pairs) flanking a NeoR cassette, GalK cassette, or HA tag were linearized and used

for sequential recombineering. Electrocompetent bacterial SW105 cells (NCI-Frederick)

were prepared by desalting, heat shocked at 42° to induce the expression of λ prophage

recombination proteins, and electroporated with linearized construct. First, the neomycin

resistance cassette was inserted into a region displaying low conservation between exons

22 and 23. The cassette containing the PGK promoter, neomycin resistance, and bGH

polyA tail was subcloned from the pL452 vector (NCI-Frederick). Next, HA was

inserted at the end of the Chd1l coding region. HA was inserted by sequential selection

first for the presence of GalK by plating clones on galactose agar medium, and then for

http://www.ncbi.nlm.nih.gov/genome/clone/clname.cgi?stype=Name&list=RP24-273N16#_blank

66

the loss of GalK (and replacement by HA) by plating on MacKonkey agar medium. The

GalK cassette was subcloned from pGalK (NCI-Frederick). Lastly, the DTA cassette was

inserted at the end of the 3’ Chd1l homology arm. The cassette containing the MCI

promoter, DT-A, and PGK polyA was subcloned from the pMC1-DTpA vector.

Contributing Collaborators

I would like to thank Ziming Weng1,4

and for performing Solexa sequencing, Dr.

Phil Lacroute1 for processing sequencing reads, their advisor Dr. Arend Sidow

1,4, Wenxiu

Ma5,6

for doing CisGenome analysis, her advisor Dr. Wing Wong6, and my advisor Dr.

Matthew P. Scott1,2,3

.

1Department of Genetics

2Department of Developmental Biology

3Department of Bioengineering

4Department of Pathology

5Department of Computer Science

6Department of Biostatistics

67

Figure 3.1 Preliminary validations for ChIP

A. Validation of α-Chd1l specificity. ES cell extracts induced (-Tet) or uninduced

(+Tet) to express a Chd1l-shRNA were immunoblotted with α-Chd1l, α-Oct4, and α-β-

Tubulin. α-Chd1l detects a band ~100 KD in uninduced ES cell extracts, but not in

extracts in which Chd1l had been knocked down. B. α-Chd1l efficiently

immunodepletes Chd1l from ES cell extracts in a dose dependent manner, but does not

pull down Oct4, another nuclear protein. C. Cell fractionation shows that, like Snf2l,

Chd1l fractionates with chromatin, while cytoplasmic β-tubulin does not. D. Validation

of PolII ChIP. PolII ChIP library was validated by qPCR amplification of selected

genomic loci. CNAP is an intergenic region where PolII does not bind, and ChIP does no

enrichment of PolII at this locus. Conversely, promoters of Gapdh and β-Actin are

enriched for PolII binding.

A C B

D

68

# Raw Reads # Post Filter % # Mappable % # Non-Repetitive %

Chd1l 12,540,570 6,415,343 51.20% 4,498,942 70.10% 3,588,770 79.80%

PolII 8,446,088 5,452,102 64.60% 3,895,115 71.40% 3,668,476 94.20%

Total C’tin 12,631,182 6,759,394 53.50% 3,652,432 54.00% 3,588,770 98.30%

Table 3.1 Number of reads obtained for ChIP libraries

The reads obtained from sequencing of Chd1l and PolII ChIP and total chromatin (c’tin) libraries were passed through a quality filter

(no arbitrary nucleotides for 25 base pairs) and then mapped to the mouse genome. Repetitive reads were filtered out to avoid

skewing peak calling software. Post-filter, mappable, non-repetitive reads were used for peak calling.

Figure 3.2 Distribution of reads around TSS

Reads from PolII and Chd1l ChIP libraries and total chromatin library were mapped 1000 base pairs upstream and 1000 base pairs

downstream of transcription start sites (TSSs). A small peak can be observed in the “total chromatin” sample because chromatin

typically shears more easily at the TSSs, and these ends of chromatin get preferentially sequenced. The peak from the PolII sample is

much more significant, confirming that PolII is frequently located at TSSs. The Chd1l sample shows a modest peak at the TSS.

69

ChIP Control

# of Peaks

FDR<0.01 FDR<0.05 FDR<0.10

Chd1l Total C'tin 98 206 792

PolII Total C'tin 15304 17582 17599

Table 3.2. Number of peaks called for ChIP samples

The number of peaks for Chd1l ChIP and PolII ChIP sequencing experiment a varying

false discovery rates (FDR) is shown. At a FDR of <0.05, ~200 peaks were called for

Chd1l ChIP. This is in sharp contrast with the nearly 18,000 peaks called for PolII ChIP.

ChIP Control

# of Peaks

FDR<0.01 FDR<0.05 FDR<0.10

Run #1 Chd1l Total C’tin 98 206 792

Run #2 Chd1l Total C’tin 8 178 699

Combined Chd1l Total C’tin 201 2836 9160

Table 3.3. Number of peaks called for the combined analysis

Combined analysis of two sequencing runs of the same Chd1l ChIP library produced a

greater number of Chd1l peaks than either individual sequencing run alone.

70

A

B

Conservation

RefSeq

PolII Peak

PolII Signal

Total C’tin

Chd1l Peak

Chd1l Signal

C

Conservation

RefSeq

PolII Peak

PolII Signal

Total C’tin

RefSeq

Chd1l Signal

Conservation

PolII Peak

PolII Signal

Total C’tin

Chd1l Peak

Fig. 8 cont. next pg.

71

Figure 3.3 Validation of Chd1l ChIP peaks

A-C. Each track is labeled at the right. Forward reads are coded in red; reverse reads are

coded in blue. Black bars above reads represent a peak called by the software. RefSeq

tracks show presence of genes. Arrowed regions are intergenic and show the direction of

transcription. A. PolII peak calling at the Gapdh locus. Bi-horned peaks of forward

and reverse reads flank the PolII peak called at the promoter of the Gapdh locus.

Background signal from total chromatin (c’tin) is minimal. B. Example of an excluded,

top-ranking peak called for Chd1l. Peaks were manually excluded from analysis if the

background showed excessively high signal. C. Example of a Chd1l ChIP peak

manually selected for further validation. Chd1l peaks were selected if background

was minimal and if flanked by forward and reverse reads. D. Overlap between Chd1l

and PolII peaks at FDR<0.05. The Venn diagram shows that a significant portion

of Chd1l peaks overlap with PolII. E. Validation of Chd1l peaks by qPCR. Primers

were designed around the peaks and used in qPCR analysis of Chd1l and PolII ChIP

libraries. A subset of the Chd1l peaks selected was associated with nearby PolII peaks.

qPCR of PolII ChIP samples at these loci show enrichment of PolII binding as expected.

However, no enrichment of Chd1l binding was found for any candidate locus in Chd1l

ChIP samples.

17495

87/88

118 PolII Chd1l

E

D

72

Figure 3.4. Re-analysis of two combined Chd1l ChIP sequencing runs

A. Overlap of the reads called for each individual sequencing run and the combined

analysis of both sequencing runs. No peaks called for sequencing run #1 were common

to peaks called for sequencing run #2. Peaks called for the combined analysis represent a

largely distinct set from either run #1 or run #2. B. Example of a top-ranked peak

called for a combined analysis of two Chd1l ChIP sequencing runs. The appearance

of peaks called for the combined analysis was not noticeably different than for peaks

called for the first analysis. Peak height remained between 4 and 5.

A

196

1 97

4 4

Run #1 Combined

Run #2

RefSeq

Chd1l Signal

Conservation

PolII Peak

PolII Signal

Total C’tin

Chd1l Peak

Comb. Chd1l Signal

Comb. Chd1l

Peak

Comb. Total C’tin

B

73

Figure 3.5. Chd1l-HA targeting vector

The Chd1l-HA targeting vector was built to incorporate an HA tag onto the 3’ end of the

Chd1l gene. Recombineering technology was used to produce a vector containing two 4

and 5 Kb arms of sequence homologous to the Chd1l locus (red), a neomycin resistance

(NeoR) cassette for positive selection (green), and a DTA cassette for negative selection

(blue). The HA tag (bright yellow bar) is incorporated prior to the stop codon at the end

of the last exon in the Chd1l gene. The pBlueScript sk(+) backbone is digested away

upon linearization prior to recombination in ES cells (yellow). After selection by

neomycin, the neomycin cassette is floxed out by Cre-mediated recombination at the

LoxP sites.

74

4. Molecular Functions of Chd1l in Early Development

Introduction

Multiple studies published within the last year have identified a role for Chd1l in

the DNA damage response pathway and as an oncogene that promotes spontaneous tumor

formation in mice [259,260,262]. Thus far, experiments conducted in this thesis were

directed at elucidating developmental and transcriptional roles for Chd1l. Having

identified an essential role for Chd1l in the preimplantation embryo, it remains unknown

whether defects in gene expression underlie the arrest phenotype (Chapter 2). However,

no changes in transcriptional regulation could be identified in ES cells deficient in Chd1l,

providing evidence that its role may not be at the level of gene regulation. Other ATP-

dependent chromatin remodeling factors are involved in a broad spectrum of nuclear

processes other than transcription [94,280]. The INO80 complex, for example, responds

to early indicators of DNA damage and repairs double-stranded breaks (DSBs) [92]. The

first half of this chapter explores a potential role for Chd1l in promoting apoptosis in ES

cells as a response to DNA damage.

Given that Chd1l is essential in the preimplantation embryo, the question remains:

what is the molecular mechanism of Chd1l in early development? Perhaps more

intriguing is the puzzle of why Chd1l is essential in the preimplantation embryo but not

in ES cells? One possibility is that Chd1l is involved in a process that occurs in the early

embryo but that does not occur or is no longer required in ES cells. In the second half of

this chapter, I propose two hypotheses regarding what this process could be, based on

knowledge about the protein structure of Chd1l, which includes a macro domain that

binds polymers of ADP-ribose (PAR), and from recent reports demonstrating that Chd1l

is a key player in the DNA repair response. The first hypothesis is that the arrest

phenotype of embryos injected with Chd1l MO is due to a defect in DNA repair; the

second hypothesis is that the arrest phenotype is due to disregulation in PAR levels,

which may also have implications in DNA repair.

Everyday, tens of thousands of nucleotides are damaged in a typical cell due to

endogenous sources of DNA damage as well as exogenous sources [6]. Such endogenous

75

sources can arise from byproducts of normal metabolism called reactive oxygen species

that oxidize nucleotides such that they become toxic, mispairing, or miscoding [305].

The sugar bonds of nucleotides are susceptible to spontaneous hydrolysis, resulting in

miscoding bases or abasic sites. Mismatch errors can arise from basepairing inaccuracies

during replication. In addition, S-adenosylmethionine can transfer methyl groups to

nucleotides, changing their composition and basepairing properties. Stalling of the

replication fork due to various types of lesions can generate double-stranded breaks

(DSBs), a highly deleterious mutation. If lesions are not repaired, programmed cell death

may ensue or mutations can be passed down to future cell lineages.

Fortunately, repair mechanisms exist to protect the genome from endogenous and

exogenous DNA damage. Repair mechanisms that function on nucleotides of at least a

partially intact strand of DNA include base excision repair (BER), in which a single

damaged nucleotide is removed and replace with another; nucleotide excision repair

(NER), in which a lesion causing distortion of the DNA strand is removed along with a

number of nucleotides flanking either side of the lesion [306]. Mismatches introduced

during replication are repaired by mismatch repair (MMR) [305]. Repair mechanisms

that respond to DSBs include error-free homologous recombination-mediated repair

(HRR) or error-prone non-homologous end joining (NHEJ), in which any two ends of a

broken strand of DNA can be ligated together [305].

Defects in repair mechanisms can lead to a broad spectrum of human diseases,

including many types of cancer [236]. Mutations in the BRCA1 or BRCA2 genes, for

example, lead to disruptions in DSB repair and predispose individuals to breast and

ovarian cancer. Mutations in ATM render cells deficient in the recognition of DSBs and

lead to a disease called Ataxia-telangiectasia that is characterized by childhood leukemias

and lymphomas. Disruptions in a number of genes involved in MMR lead to a condition

called hereditary non-polyposis colon cancer (HNPPC) whereas disruptions in genes that

participate in BER can lead to Xeroderma pigmentosum in which individuals suffer high

prevalence of skin cancer.

Because of the link with cancer, DNA repair is intensely studied, but repair

mechanisms during early embryogenesis are only recently becoming elucidated

[253,307,308]. Understanding how DNA repair functions during preimplantation

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development can have major impacts on assisted reproductive technology (ART), since

infertility can be correlated with chromosomal abnormalities and inefficient DNA repair

[237,309]. Because the phase of the cell cycle influences which DNA repair pathways

are used, it is expected that the manner in which DNA is repaired in the preimplantation

embryo, in which cells have very short G1 and G2 phases and no apparent G1/S phase

check point, will be distinct [310,311]. These cells also have increased need to maintain

genetic fidelity. Several studies have shown that transcripts encoding proteins involved

in all the major DNA repair pathways are present in the zygote and in preimplantation

embryos [253,308]. However, not only is there strong translational regulation in embryos

[312], but control of DNA repair mechanisms occurs largely through modifications at the

protein level, leaving many unknowns about DNA repair during early embryogenesis.

Genetic studies have demonstrated that components of the BER pathway such as Polβ

[313], Apex [314], Lig1 [315], and Fen1 [316] are necessary for development. The

majority of mice null for components of NER are viable but display sensitivity to DNA

damage [317]. The components of NER necessary for embryonic survival are Xpd [318],

Rad23A/Rad23B (double knock-out) [319], and Xab2 [320]. There are few, if any,

components of MMR that result in embryonic lethality when deleted [317], but mutations

in members of this pathway lead to deleterious diseases and cancers in post-natal

individuals [307]. In contrast, proteins that are involved in the repair of DSBs that are

essential for development are many, suggesting high dependence on the HRR and/or

NHEJ pathways in the embryos [317].

ES cells are pluripotent cells that are derived from the inner cell mass of the

blastocyst and may also have unique ways of coping with DNA damage. ES cells rapidly

proliferate in culture while maintaining self-renewal indefinitely. This ability requires

stringent DNA repair mechanisms to ensure that damage is repaired quickly, or that cells

with damaged DNA are removed from the population to prevent mutations from being

passed down to daughter cells. ES cells are highly sensitive to DNA damage, being more

likely to undergo apoptosis as a response to DNA damage than other cells types

[108,116,117]. This is not to say that ES cells are less proficient in repairing damage

than other cell types. On the contrary, the spontaneous mutation frequency in ES cells is

at least 100 times lower than in isogenic MEFs, depending on which reporter gene is

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assayed [109,110,112], indicating robust machinery efficiently repairs DNA damage in

ES cells. Similar to early development, understanding of DNA repair pathways in ES

cells lags behind knowledge of repair in somatic cells and differentiated cultured cells

[109].

PAR, or poly(ADP-ribose) is a post-translational modification that is immediately

added to nuclear receptor proteins in response to DNA damage. Because ADP-ribose is a

positively charged moiety, addition onto chromatin promotes chromatin relaxation and is

proposed to allow access to DNA repair machinery. The PAR modification is

recognized by the macro domain of proteins such as Chd1l [61,259,260]. Long and

branching chains of PAR are synthesized by the PAR polymerase (PARP) family of

enzymes using the respiratory metabolite NAD+ as a substrate [32]. PAR levels are

tightly regulated by polymerization by a large number of Parps and by degradation by a

single enzyme called poly(ADP-ribose) glycohydrolase, or PARG. Disregulation of PAR

levels during development leads to lethality; Double knock-outs of two major PARPs,

PARP-1 and PARP-1, are embryonic lethal at about E5.5 [57], and PARG knock-out

mice die even earlier, at E3.5 [289]. As the PAR modification has diverse roles in DNA

repair, transcription , telomere length homeostasis, and others [321], it is unclear what the

primary cause of lethality is when PAR levels are reduced or amplified in these mouse

mutants.

The role of PAR in the cell is best described in terms of the function of the

various PARP themselves. PARP-1 and PARP-2, the best described PAR polymerases,

have partially redundant functions [57]. Their activity is strongly activated by DNA

damage, either single-stranded or double stranded breaks, and mouse knock-outs of

PARP-1 or PARP-2 are hypersensitive to ionizing radiation and alkylation [52,57].

PARP-3 localizes to the centrosome, and its over-expression interferes with G1/S phase

progression [40]. Parp-5a and Parp-5b, also known as Tankyrase 1 and Tankyrase 2,

respectively, associate with telomeric proteins, including TRF1 [41,43]. Tankyrase 1

levels and perhaps Tankyrase 2 levels influence telomere length and segregation during

cell division [42].

In addition to safeguarding the genome, there is also a “dark side” to PARP-1

function [33]. PARP-1 promotes apoptosis through a caspase-independent AIF pathway,

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an activity that is halted by treatment of PARP-1 inhibitors [65]. In the AIF-mediated

apoptosis, AIF, which is normally sequestered in mitochondria, is translocated to the

nucleus where it triggers rapid condensation and fragmentation of chromatin [322,323].

PARP-1 is required for AIF translocation to the nucleus, but how it accomplishes this

remains unknown [65]. Interestingly, Tankyrase 2 when over-expressed also induces

caspase-independent apoptosis, an effect diminished by treatment of a PARP inhibitor

[43].

Because Chd1l binds to PAR through the macro domain, Chd1l may contribute to

DNA repair through PAR recognition. Alternatively, Chd1l may recognize PARylated

proteins and carry out other nuclear functions, including the activation of caspase-

independent apoptosis. Like other chromatin remodeling factors, Chd1l may function as

the ATP-dependent motor of a complex that has multiple and diverse functions

throughout development and in different cell types. This chapter provides data that

indicate that Chd1l indeed recognizes DNA damage in ES cells, but may propagate a

response that is very different than that reported in the literature for other cell types.

Results

Reduction of Chd1l increases DNA damage tolerance in ES cells

In cultured cells, Chd1l participates in the response to several types of DNA

damage. Chd1l localizes to double-stranded breaks induced by laser microirradiation

[259,260], and reduction of Chd1l protein causes hypersensitivity to phleomycin- and

H2O2-induced damage [259]. Studies described in this dissertation demonstrate that

Chd1l is not essential for ES cells under normal culture conditions. However, these

experiments have not addressed whether Chd1l participates in DNA repair in ES cells.

Given that Chd1l participates in the DNA damage response in the U20S

osteosarcoma cell line [259] and in the HeLa cervical cancer cell line [260], it is

reasonable to hypothesize that Chd1l also participates in DNA repair in ES cells. To test

this, sensitivity to H2O2-induced oxidative DNA damage was assayed in ES cells

expressing Chd1l-shRNA or NS-shRNA. Expression of Chd1l-shRNA or NS-shRNA

was induced by removing Tetracycline 48 hours prior to seeding. Knock-down of Chd1l

in Chd1l-shRNA expressing ES cells was confirmed by SDS-PAGE (Fig. 1A). ES cells

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were treated with H2O2 concentrations varying between 5 µM and 500 µM 24 hours after

seeding. Surprisingly, two days after treatment, a slight increase in survival was seen for

ES cells with reduced Chd1l at the lower concentrations of H2O2 (Fig. 1B). Although

subtle, the effect with H2O2 treatment is reproducible; multiple experiments showed

greater survival for ES cells expressing Chd1l-shRNA than those expressing NS-shRNA.

To test whether ES cells with reduced Chd1l are more sensitive to strand breaks,

ES expressing Chd1l-shRNA or NS-shRNA were treated with varying concentrations of

the radiomimetic drug phleomycin and assayed for percent survival relative to untreated

cells (Fig. 1C). In contrast to H2O2 treatment, treatment with phleomycin rapidly killed

control ES cells even with very low phleomycin concentration (5 µM). Therefore,

phleomycin treatment is reported after only 24 hours. ES cells expressing Chd1l-shRNA

showed higher percent survival than ES cells expressing NS-shRNA at all concentrations

of phleomycin treatment, a finding reproduced over multiple experiments. Intriguingly,

the increased resistance to DNA damage-induced killing seen in ES cells with Chd1l

knocked-down is more pronounced when phleomycin is used than when H2O2 is used,

suggesting that Chd1l may play a more prominent role in the recognition of double-

stranded breaks than lesions induced by oxidative damage.

To test whether knock-down of Chd1l provided long-term resistance to killing by

drug-induced DNA damage, survival rates over a three day period were measured. At 1

day after treatment, Chd1l-shRNA expressing ES cells survive slightly better than NS-

shRNA expressing cells in the lower H2O2 concentrations (<250 µM) (Fig. 1D). This

difference was reduced by 2 days (Fig. 1E) and not seen at 3 days (Fig. 1E). The data

suggest that increased resistance is an acute affect and may not confer long-term

survivability to ES cells. The data show that ES cells with reduced Chd1l levels do not

show hypersensitivity when treated with H2O2 or phleomycin, as has been reported in

other cell types. Instead, reducing Chd1l levels in ES cells confers acute resistance to

DNA damage-induced killing.

One explanation for this surprising result is that in ES cells, Chd1l promotes

apoptosis as a response to DNA damage. Cells that lack Chd1l could be impaired in

initiating apoptosis. Such a finding is reminiscent of a phenomenon termed “damage

tolerance” seen when ES cells deficient in a mismatch repair protein, Msh2, are treated

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with DNA certain damaging agents [138,324,325,326]. Damage tolerance is observed in

cancer cells that are able to circumvent apoptosis as a response to DNA damage.

Over-expression of Chd1l kills ES cells but not differentiated cells

The result that ES cells with reduced Chd1l are more resistant to killing brought

to attention data obtained previously in this thesis showing that over-expression of Chd1l

causes widespread death in ES cells. EBRTcH3 ES cells [284] were used for stable

integration of Chd1l transgene and inducible expression by removal of Tetracycline from

culture medium. Two forms of Chd1l were over-expressed: a wild-type form (WT), and

a mutant predicted to lack ATPase activity due to a point mutation in the ATP binding

domain (K71R). The lysine in ATP binding motifs is critical to its function; mutating

this residue abolishes ATP hydrolysis in many different proteins, including SNF2

proteins in yeast and human [76,327,328,329,330,331]. Ahel et. al. showed that an

analogous mutation in the human CHD1L protein abolished ATP hydrolysis [259],

strongly suggesting that this mutation in mouse Chd1l is deficient in ATP hydrolysis.

Tetracycline was removed from culturing media to induce the expression of either

WT or K71R Chd1l. After six days of culturing without Tetracycline, ES cell

populations over-expressing either form of Chd1l had ~90% fewer cells than control ES

cell populations cultured in the presence of Tetracycline to maintain transgene repression

(Fig. 2A).

To address the question of whether over-expression triggers ES cells to

differentiate, Chd1l over-expression (either K71R or WT) was induced by removal of

Tetracycline for 24 hours and then ES cells were seeded at clonal density and cultured for

six days. Colonies were stained with Leishman’s Stain, a chromatin stain that labels

pluripotent cells. Colony morphology is indicative of the pluripotent state of ES cells; ES

cell colonies that are round, small in diameter, and multilayered indicate pluripotency,

whereas colonies that are monolayered and diffuse are indicative of differentiation (Fig.).

Some spontaneous differentiation (~5%) is expected in ES cell cultures. Chd1l over-

expressing colonies (either K71R or WT) were smaller in diameter than control ES cell

colonies (Fig. 2C). The number of colonies obtained for ES cells over-expressing Chd1l

was reduced by more than 60%, indicating that the loss in cell number was not only due

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to lack of proliferation (Fig. 2B). Colonies were scored as either “differentiated” or

“undifferentiated.” Chd1l over-expressing colonies were 95% (K71R) or 94% (WT)

undifferentiated (pluripotent) compared to 95% undifferentiated in control colonies (Fig.

2D). Therefore, the effect of Chd1l over-expression on differentiation in ES cells is

minimal.

The drastic loss in cell number upon Chd1l over-expression could be due to

changes in proliferation or cell death. To address whether Chd1l over-expression causes

cell cycling, ES cells were cultured without Tetracycline for three days to induce the

expression of either WT or K71R forms of Chd1l, permeabilized, and stained with

propidium iodide (PI) to label DNA. DNA content was determined by flow cytometry

and percentages of cells in each phase of the cell cycle were calculated using FlowJo

software. Consistent with reports that Chd1l may contribute to G1/S phase transition

[262], 4.5% fewer of K71R and 3% fewer of WT Chd1l over-expressing cells were found

in G1 phase compared to their uninduced (+Tet) counterparts, small but significant

differences (χ2 test, p<0.001) (Fig. 2E). However, no gross arrest in any phase was

observed that could explain ~90% reductions in cell numbers.

To ask if ES cells over-expressing Chd1l were undergo apoptosis, ES cells that

were induced for four days to express either WT or K71R forms of Chd1l were collected

and co-stained with PI and AnnexinV. The population of ES staining positive for

AnnexinV but negative for PI was counted using flow cytometry and gating was done

using FlowJo software. There were ~3.5 times (K71R) or ~2.5 times (WT) as many

apoptotic Chd1l-over-expressing ES cells as control ES cells (χ2 test, p<0.001) (Fig. 2F),

showing that over-expression of Chd1l initiates apoptosis in ES cells.

To test whether over-expression of Chd1l is toxic to other cell types, Chd1l was

over-expressed in 3T3 fibroblasts. 3T3 cells were transfected with a GFP-containing

vector only or WT or K71R forms of Chd1l. No readily apparent differences in cell

death were seen between cells transfected with vector only and cells transfected with

Chd1l-containing vector. As cell division ensues, the transfected vector will be diluted

out. If Chd1l is toxic in 3T3s, cells that lose the Chd1l-containing vector will have a

selective advantage over those that retain it. To test this, cells expressing GFP were

selected two days after transfection using FACS sorting and cultured for an additional

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two days. A second-round of flow cytometry revealed that 68% or 70% of cells

transfected with the K71R or WT Chd1l-containing vector retained GFP expression,

whereas only 58% of cells transfected with the empty vectors retained GFP expression

(Fig. 3A), showing that Chd1l is not toxic to 3T3s and suggesting instead that over-

expression of Chd1l endows a slight (~10%) selective advantage over those transfected

with vector only.

The cell death caused by over-expression of Chd1l may be a phenotype specific to

ES cells. Mouse models over-expressing Chd1l are viable and experience ~24% increase

in spontaneous tumor formation in adult mice [262]. To test whether differentiating ES

cells can tolerate the over-expression of Chd1l, LIF was removed from culturing medium

and expression of Chd1l transgene was induced by tetracycline removal. ES cells were

cultured in the absence of LIF for 0, 2, 4, or 6 days prior to tetracycline removal, and LIF

was withheld throughout an additional six days of culturing with Chd1l over-expression

(Fig. 3B). Survival of ES cells over-expressing Chd1l was impaired in undifferentiated

ES cells as well as cells in which Chd1l over-expression was induced simultaneously

with LIF removal (D0-LR) (Fig. 3C). However, Chd1l over-expressing ES cells

differentiated for 2 or 4 days prior to Chd1l induction (D2-LR or D4-LR) survived as

well as (and better than) control ES cells (Fig. 3C), indicating that at some critical

moment during differentiation, ES cells become resistant to Chd1l over-expression.

Unexpectedly, the results were highly similar between over-expression of WT and

K71R forms of Chd1l for all experiments described above. It was originally thought that

over-expression was a loss of function phenotype because over-representation of a

subunit of a complex can disrupt protein stoichiometry and yield partially formed,

nonfunctional complexes. However, reducing Chd1l levels by shRNA (a more likely

loss-of-function manipulation) produces a very different effect (Chapter 2); therefore, the

apoptosis-inducing phenotype of Chd1l over-expression in ES cells is probably a

hypermorphic or neomorphic function that does not depend on the ATPase domain. It

would be interesting to address whether the macro domain plays a role in the apoptotic

phenotype by over-expressing a macro domain mutant in ES cells.

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Chd1l forms a ~500 kD complex in ES cells

ATP-dependent chromatin remodeling factors are found in multi-subunit

complexes than can change in composition and function throughout development and in

different cell types [69,70,71]. To address the question of whether Chd1l forms a

complex in ES cells, nuclear extracts were prepared and size fractionated on a 10-30%

glycerol gradient. Large proteins and complexes will filter farther through the gradient

than smaller proteins and complexes. Fractions were then run on SDS-PAGE and blotted

for Chd1l, pluripotency proteins Oct4 and Sox2, the DNA repair protein PARP-1, and the

chromatin remodeling factor Brg1.

Chd1l was found most abundant in fractions 4-6, cofractionating with PARP-1

(Fig. 4). Consistent with previous reports, Brg1 was most abundant in fractions 12-14

[69], correlating with a complex size of ~2 mDa. Oct4 and Sox2 form a complex of

about 440 kDa [332]. Therefore, fractions 4-6 in which Chd1l and PARP-1 are found are

estimated to contain complexes about 500 kDa. Chd1l and PARP-1 associate with each

other in HEK293 and in 293T cells [259,260]. Co-fractionation with Chd1l and PARP-1

suggest that these proteins also interact in ES cells, although co-immunoprecipitation

experiments are necessary to confirm this.

Because Chd1l is itself 100 kDa, a sum of ~400 kDa of subunits remain to be

identified. PARP-1 is 113 kDa, potentially accounting for some of the mass.

Identification of complex subunits could yield valuable insight into the function of Chd1l,

as could characterization of a Chd1l-containing complex in ES cells that have been

treated with DNA damaging agents.

γ-H2AX marks uninjected embryos and embryos injected with Chd1l-MO

Given that Chd1l participates in the DNA damage response in cultured cells, it

could also govern a DNA damage response in the preimplantation embryo. Although

DNA repair is still being elucidated in very early development, if any cells of an

organism should need strict repair mechanisms, it should be the zygote and blastomeres

as these are the progenitor cells that anchor all developmental lineages. Indeed, a large

number of expression studies reveal that transcripts encoding DNA repair proteins for

mismatch repair (MMR), base excision repair (BER), nucleotide excision repair (NER),

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homologous recombination- mediated repair (HRR), and non-homologous end joining

(NHEJ) are present in the zygote and throughout preimplantation development [253].

Many DNA repair proteins are essential for development as their knock-out phenotypes

results in early embryonic lethality, especially those contributing to HRR of double

strand breaks [317]. Therefore, a defect in DNA repair could potentially explain the

arrest phenotype seen in embryos injected with MO targeting Chd1l.

One way in which to test for DNA damage is by assaying the levels of

phosphorylation of the histone variant H2AX. Phosphorylated H2AX associates with

sites of DNA damage and is essential for the repair of DSBs [333]. Increased levels of

γH2AX have been demonstrated to occur in the embryo upon treatment with DNA

damaging agents [334,335]. Quantification at the protein level is inherently difficult in

the preimplantation embryo due to the low abundance of starting material that is further

limited by the number of embryos that can be µ-injected in a single experiment (30 to

50). Multiple attempts at obtaining reliable results from SDS-PAGE or dot-blotting using

as many as 100 embryos were unsuccessful. Although less quantitative, general protein

levels can be observed using immunocytochemistry (ICC) staining in embryos.

Therefore, to address whether embryos deficient in Chd1l experience an

accumulation of DNA damage, embryos injected with Chd1l MO were subjected to ICC

staining using the α-All uninjected embryos developed normally to the 16- to 32- cell

stage. Embryos injected with MO-1 developed to the 8- to 16-cell stage; previous

experiments (Chapter 2) show that embryos injected with MO-1 arrest at this stage and

do not develop further. Embryos injected with MO-2 were arrested at either the 2-cell

stage or the 4-cell stage. At least fifteen embryos were collected for each uninjected,

MO-1 injected, and MO-2 injected embryos and stained with α-γH2AX antibody. .

Phosphorylated H2AX also marks nuclei of embryos injected with Chd1l MO-1, and

staining in some nuclei is brighter than nuclei of uninjected embryos (Fig. 5A). Nuclei of

embryos injected with Chd1l MO-2 also showed strong γH2AX staining that appear

somewhat brighter than control embryos (Fig. 5B). The data is consistent with a role for

Chd1l in the early embryo, suggesting that a loss of Chd1l may increase DNA damage,

particularly damage that ends in DSBs. The differences in staining signals are difficult to

quantify because ICC staining is only semi-quantitative, and further experiments are

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necessary to confirm that accumulation of DNA damage is the underlying cause of arrest

in preimplantation embryo injected with Chd1l MO.

PAR marks uninjected embryos and embryos injected with Chd1l-MO

PAR levels are carefully regulated in the preimplantation embryo by the activity

of the PARP family of enzymes that add PAR onto nuclear receptor proteins, and by the

activity of PARG, which rapidly degrades PAR modifications. Mouse knockouts of

PARG are unable to catabolize PAR [289], and double knockouts of partially redundant

polymerases PARP-1 and PARP-2 are deficient in PAR synthesis [57]. Knock-out of

either PARG or both PARP-1 and PARP-2 results in early embryonic lethality at E3.5 and

7.5, respectively.

Because Chd1l binds to PAR through its C-terminal macro domain, and because

embryos injected with Chd1l-MO arrest prior to the blastocyst stage, it was intriguing to

hypothesize that the arrest phenotype was due to aberrant PAR levels in the embryo.

Given that Chd1l binds to PAR in vivo, Chd1l could be involved in the propagation of a

PAR recognition and degradation process. Loss of Chd1l could disrupt PAR recognition

and subsequent degradation by PARG, leading to toxic PAR levels. Alternatively,

through binding to PAR, Chd1l could stabilize its presence long enough for it to be

recognized by other proteins. Loss of Chd1l could lead to PAR destabilization, rapid

degradation, and untenably low levels in the cell. If either of these scenarios were true,

PAR levels would be altered in embryos injected with Chd1l MOs.

To test whether altered PAR levels could explain the Chd1l arrest phenotype,

embryos injected with Chd1l-MO were subjected to ICC and stained with α-PAR.

Zygote-stage embryos were µ-injected with Chd1l MO-1 or MO-2 and fixed three days

later (E3.5). Uninjected embryos appeared to develop normally to this stage, and

embryos injected with MO-1 developed to and were likely arrested at o the 8- to 16-cell

stage. Embryos injected with MO-2 were arrested at either the 2-cell stage or the 4-cell

stage. At least fifteen embryos were collected for each uninjected, MO-1 injected, and

MO-2 injected embryos and stained with α-PAR antibody. Uninjected embryos show

low levels of PAR staining in the nucleus, consistent with reports that PAR modifies

nuclear proteins (Fig. 6). Some globular staining can be seen outside of the nucleus, but

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this is likely non-specific sticking to membranes or the zona pellucida. One uninjected

embryo arrested at the 2-cell stage, and this embryo is shown to contrast Chd1l MO-2

embryos that consistently arrest at this stage (Fig. 6C).

PAR staining in Chd1l MO-1 embryos also shows nuclear localization of PAR

(Fig. 6A), demonstrating that the arrest phenotype is not due to global reduction of PAR.

Some blastomeres of MO-1 injected embryos show PAR staining that is slightly greater

than those in uninjected embryos, suggesting PAR levels could be higher in arrested

embryos. As ICC is semi-quantitative at best, it is difficult to make conclusions between

samples with similar signals. Even if PAR levels are higher in embryos injected with

MO-1, the difference may not be great enough to explain the early arrest phenotype.

PAR staining in Chd1l MO-2 embryos show nuclear localization of PAR, but the signal

in some cells is reduced compared to cells of uninjected embryos (Fig. 6B, note

uninjected embryo arrested at the 2-cell stage), suggesting that PAR levels are actually

reduced. The difference is subtle and inconsistent between embryos, and the discrepancy

between embryos injected with MO-1 and MO-2 is difficult to rationalize. One

possibility is that the transient nature of PAR modification causes it to degrade in arrested

embryos. Thus PAR in MO-2 injected embryos that had been arrested for a longer

duration than MO-1 injected embryos may show reduced PAR levels.

The data show that PAR levels are not consistently or dramatically altered in

embryos injected with Chd1l MO, indicating that the arrest phenotype is not due to large-

scale changes in PAR levels. More quantitative techniques are necessary to determine

whether PAR levels change subtly in MO-injected embryos versus uninjected embryos;

however due to the discrepancy between MO-1 and MO-2 injected embryos, identifying

such subtle changes remains unlikely to explain the arrest phenotype seen in embryos

injected with Chd1l MO.

Discussion

Upon experiencing DNA damage, a cell has two choices: 1) repair the damage, or

2) initiate programmed cell death or apoptosis to remove the damaged cell from the

population. ES cells lack a G1 checkpoint in which some DNA repair takes place and are

instead more prone to initiate apoptosis in response to DNA damage than other cell types

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[108,116,117,119,338]. Two independent groups have demonstrated that Chd1l is part of

the DNA damage response [259,260]. In ES cells, Chd1l may respond to DNA damage

by promoting apoptosis; therefore, reduction of Chd1l in ES cells may increase their

tolerance to drug-induced DNA damage. Further experiments will be necessary to test

whether DNA damage accumulates and whether the programmed cell death pathway is

impaired in ES cells deficient in Chd1l.

Consistent with the speculation that Chd1l pushes ES cells toward apoptosis in

response to DNA damage, over-expression of Chd1l in ES cells causes widespread

killing and increased staining of the apoptotic marker AnnexinV. This effect is

reminiscent Tankyrase 2- (PARP-5b-) induced cell death upon transient transfection

[43]. PARP-1, when over active can also trigger apoptosis in a caspase-independent

manner. Cell death induced by both Tank2 over-expression and Parp-1 over-activation

is blocked by addition of PARP inhibitors, suggesting that the presence of additional

PAR molecules may contribute to the initiation of apoptosis [339]. An alternative

explanation is that over-active PARP depletes cellular NAD+ and ATP levels, and

necrosis ensues [37]. Interestingly, Parp1 deficiency in mouse disease models is

cytoprotective to ischemia-reperfusion injury, inflammation-related injury [63], diabetes

[64], and hyperoxic damage. However, several studies indicate that energy depletion

alone may not account for Parp-1-induced cell death [340]. Cell death induced by over-

activation of PARP-1 requires the translocation of AIF to the nucleus, and inhibition of

AIF by microinjection of AIF antibodies blocks PARP-1-dependent cell death [65]. The

release of AIF from the mitochondrion is PARP-1 dependent, but how PARP-1 trigger

AIF release is not known [65]. It is interesting to speculate that Chd1l may play a role in

PAR recognition and signaling leading to AIF release. While it is puzzling to think how

a chromatin remodeling factor might participate in this, cell death caused by Chd1l over-

expression seems to be independent of nucleosome remodeling activity, since over-

expression of the K71R mutant Chd1l, predicted to be deficient in ATP binding, and

over-expression of wild-type Chd1l phenocopy each other.

The increased survival of ES cells with reduced Chd1l after DNA damage was

more pronounced when that damage was induced by the radiomimetic drug Phleomycin

than with oxidative damage-inducing H2O2. Converging lines of evidence suggest that

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ES cells are less sensitive to oxidative damage as they can be cultured in hyperoxic

conditions with fewer deleterious effects than other cell types, and are instead more

dependent on strand break repair pathways, particularly error-free recombination-

mediated repair (HRR) [109]. ES cells show high basal levels of γH2AX staining, an

established marker of double stranded breaks (DSB) [109]. Because of increased

dependence on HRR to repair DSBs, loss of Chd1l could show a stronger phenotype in

ES cells treated with drugs that induce strand breaks than drugs that introduce oxidative

damage.

The data shown here present several striking paradoxes with the published

literature. Knock-down of Chd1l in ES cells and subsequent treatment with DNA

damaging agents produces cells that are more resistant to killing than control ES cells.

This is a direct contradiction with literature showing that knock-down of Chd1l in U2OS

cells causes hypersensitivity to treatment with the same DNA damaging agents [259].

That over-expression in ES cells induces apoptosis also conflicts with studies showing

that Chd1l can act as an oncogene, because over-expression in cell culture increases

colony formation in soft agar assays and transgenic over-expression in mouse models

induces spontaneous tumors [261,262]. Lastly, it is surprising that Chd1l is essential for

development to the blastocyst state, but not essential in ES cells that are derived from the

inner cell mass of the blastocyst (Chapter 2).

The data raise the possibility that Chd1l has a very different (or opposite) function

in ES cells that are adapted for culturing and maintenance of pluripotency. Supporting

this theory, apoptosis induced by Chd1l over-expression is specific to ES cells. The same

ES cells that are killed by Chd1l over-expression are no longer sensitive with as little as

two days of differentiation by LIF removal prior to introduction of Chd1l transgene.

DNA repair studies focusing on mechanisms in ES cells are beginning to show that ES

cell DNA repair mechanisms may be exceptional. On a conceptual level, ES cells should

have robust repair mechanisms to account for the rapid proliferation of a pluripotent cell

type. Spontaneous lesions, mismatches, and double strand breaks accumulate with every

cell cycle and can be highly detrimental for daughter cells and developmental lineages

[6]. During the derivation of ES cells from blastocysts, many epigenetic changes take

place, including those that induce the expression of some DNA repair genes [208].

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Evidence that ES cells have extraordinary repair mechanisms arises from low

spontaneous mutation frequencies (10-6

) found at the Aprt reporter locus, compared to 10

-

4 in isogenic MEFs [111,112]. One way in which ES cells achieve this is by quickly

removing damaged cells from the population through apoptosis [116,117].

ES cells are also unique in that their rapid cell cycling is driven by extremely high

levels of Cdk2 [120], and they lack a functional G1/S phase checkpoint [108,119,338].

This is in part due to very low levels of the Cdk inhibitors p21 and p27 and sequestering

of p53 in the cytoplasm that renders it partially nonfunctional [120]. Restoration of G1/S

phase checkpoint by ectopic expression of the serine/threonine kinase Chk2 impairs the

ability of ES cells to undergo apoptosis in response to DNA damage [108].

If Chd1l participates in the G1/S phase transition as has been proposed here and in

Chen, et. al. [262], its function in ES cells that lack a G1/S phase checkpoint could be

drastically different. Instead of responding to DNA damage by facilitating repair, Chd1l

may orchestrate an apoptotic response instead. It should be noted that while Chd1l has

been shown to respond to DNA damage, it has not yet been shown to actually repair

DNA [259,260]. A chromatin remodeling complex can achieve highly diverse (and

opposite) functions through context-dependent and cell type-specific assembly of

subunits [70,280]. Chd1l forms a ~500 kD complex in ES cells. Investigation of

complex members in ES cells should yield interesting insights.

Therefore, Chd1l may play a very specific role in ES cells in the DNA damage

response pathway, potentially pushing cells towards apoptosis through the PARP-1/AIF-

dependent, caspase-independent pathway. This theory is supported by the striking switch

in the ability of Chd1l over-expression to kill cells when ES cells are differentiated by

two days of LIF removal. What happens to the DNA repair and response machinery at

the molecular during these early stages of differentiation is unclear, but would be

intriguing to investigate.

Whether defects in DNA repair can explain the early arrest phenotype of embryos

injected with Chd1l MO remains an open question. Evidence presented here showing

increased presence of γH2AX in Chd1l deficient embryos is suggestive rather than

conclusive due to the challenges of comparing semi-quantitative ICC signal staining

between samples. Staining with other indicators of DNA damage could be informative.

90

Other approaches include FISH analysis to detect large scale chromosomal anomalies and

mutation analysis at reporter loci such as the Arpt locus [111].

Alternatively, defects in DNA repair might not explain the Chd1l arrest phenotype

at all. The effects of PARylation on the embryo are still being elucidated, and although

DNA damage activates PARP-1 to PARylate itself and other nuclear receptors, activated

PARP-1 has multiple roles, many of which are independent on DNA damage. In

addition, Chd1l may recognize PAR modifications catalyzed by another member of the

PARP family. PAR levels are not consistently altered in Chd1l deficient arrested

embryos; however, the phenotype could be due to processes downstream of PAR. PAR

binding by Chd1l through its macro domain then would not affect PAR synthesis or

degradation, and the consequence of the interaction between Chd1l and PAR would be

strictly the execution of the PAR signal.

Methods

DNA Sensitivity Assay in ES cells

The expression of Chd1l-shRNA or NS-shRNA was induced in EBRTcH3 ES

cells for two days prior to seeding. “Induced” ES cells were cultured in 7.5 µg/ml

Puromycin; “Uninduced” ES cells that do not express shRNA were cultured in 7.5 µg/ml

Puromycin and 1.5 µg/ml Tetracycline. ES cells were seeded onto 24 well plates to allow

subconfluent growth for 1, 2, or 3 days for each induced and uninduced Chd1l-shRNA

and NT-shRNA EBRThH3 cell lines. H2O2 or phleomycin was added 24 hours after

seeding with concentrations ranging from 0 to 500 µM (H2O2) or 0 to 80 µM

(phleomycin). After 24 hours, 48 hours, and 72 hours, surviving ES cells were counted

using an automated cell counter that incorporates a trypan blue cell viability

measurement. Percent survival was recorded for each induced ES cell line compared to

the corresponding uninduced cell line.

ES cell lines

The EBRTcH3 cell line contains a cassette acceptor utilizing loxP and loxPV sites

at the Rosa locus to allow efficient and directional integration of a transgene by Cre-

mediated recombination. The derivation of Chd1l-shRNA and NS-shRNA cell lines is

91

described in Chapter 2 of this dissertation. To create an ATPase-deficient mutant of

Chd1l, a point mutation was created using site-directed mutagenesis (Stratagene)

encoding a single amino acid change in the ATP-binding domain (K71R). cDNA

encoding wild-type for K71R forms of Chd1l was subcloned into the pPthC exchange

vector for recombination into the EBRTcH3 ES cell line.

The exchange vector containing the shRNA-mir sequence was cotransfected

along with a Cre expression plasmid by lipofectamine. Transfected cells were plated

single-cell density and cultured in the presence of Puromycin (1.5 µg/ml) to select for

successful integrants and of Tetracycline (1.0 µg/ml) to repress transgene expression.

Clones were confirmed by PCR genotyping of the 5’ and 3’ recombination sites. To

induce shRNA expression, the derived ES cell lines were cultured in the absence of

Tetracycline and high Puromycin (7.5 µg/ml). Control, uninduced ES cells were cultured

in high Tetracycline (1.5 µg/ml) and high Puromycin (7.5 µg/ml).

Size fractionation

Untreated parental EBRTcH3 cells (without the introduction of transgene) were

cultured in 100 µg/ml Hygromycin and grown to subconfluence on 15-cm plates.

Nuclear extract was dialyzed in 5% glycerol buffer. To make the gradient, a 30%

glycerol buffer was layered below a 10% glycerol buffer and rotated at an angle of 81.5°

for 2” at a speed of 14. Nuclear extract (0.5 ml, 10 mg/ml) was layered on top of the

gradient and ultra-centrifuged at a speed of 32,000 rpm at 4° for 20 hours. Twenty six

fractions of ~0.5 ml were collected and run on SDS-PAGE gel.

FACS analysis

For analysis of cell cycle, ES cells were grown in the absence of tetracycline for

three days to induce the expression of K71R or wild-type Chd1l. Cells were trypsinized

and resuspended at a density of 106/ml, fixed for 1hour at 4° in 100% EtOH, stained with

PI, permeabilized in 1% Triton for 10 minutes, then subjected to flow cytometry. For

analysis of apoptosis, ES cells were cultured in the absence of tetracycline for four days

to induce the expression of K71R or wild-type Chd1l. Cells were trypsinized and

resuspended at a density of 106/ml, fixed in 100% EtOH, stained with PI and AnnexinV

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without permeabilization, then subjected to flow cytometry. Data was analyzed using

FlowJo software. Statistics were calculated using χ2.

Embryo immunocytochemistry

Unless otherwise specified, all incubations and washes were carried out at room

temperature in 20-50 µl droplets of PBS-PVP solutions covered in mineral oil. Embryos

were fixed in 4% PFA for 20” at room temperature and washed for 10 minutes in PBS-

PVP. Embryos were permeabilized in a large volume of 0.3% Triton X-100 for 10

minutes without mineral oil and then washed for 5” in PBS-PVP droplets under mineral

oil. Embryos were incubated in Image iT signal enhancer solution (Invitrogen) for 30”,

washed in PVP-PBS for 15 minutes, blocked in 10% normal goal serum for 30”, and then

incubated overnight in primary antibody at 4°. Embryos were washed for 30” in PBS-

PVP, incubated in secondary antibody for 2 hours and then Hoechst for 10”. After final

washing in PBS-PVP for 30”, embryos were mounted in fibrinogen/thrombin clots. Clots

were made of equal volumes of 25 mg/ml fibrinogen (Sigma) in Ringer’s Solution and

100U/ml thrombin (Sigma) in PBS. Embryos were imaged using confocal microscopy

under 63x oil immersion. Five to ten embryos were placed into each clot.

Antibodies

Mouse α-PAR antibody was obtained from Trevigen (4335-AMC-050). Mouse

α-γH2AX was obtained from Abcam (ab18311).

93

Figure 4.1. Sensitivity to DNA damage in ES cells expressing Chd1l-shRNA

Expression of Chd1l-shRNA or NS-shRNA was induced by removal of Tetracycline 48

hours prior to treatment with varying concentrations of Phleomycin or H2O2. A. Chd1l

knock-down. Knock-down of Chd1l is observed in Chd1l-shRNA expressing ES cells

(-Tet), but not in NS-shRNA expressing ES cells (-Tet) or in uninduced ES cells (+Tet).

B-E. Sensitivity assays. Percent survival of ES cells expressing shRNA was measured to

determine sensitivity to H2O2-induced oxidative damage (B-D) or Phleomycin-induced

strand breaks (E) relative to untreated ES cells.

A

B C

D E

Figure 1. DNA repair in Chd1l-shRNA ES cells

F

94

E

C

A

-Tet +Tet

Undiffe

rentiate

d

Diffe

rentiate

d

Cell Number Colony Number

Colony Morphology D

F

B

Cell Cycling Apoptosis

Figure 2. Effect of Chd1l Over-Expression in ES cells

95

Figure 4.2. Effects of over-expression of K71R or WT Chd1l.

A. Cell number. After six days of culturing in the absence of Tetracycline, there is

~90% reduction in the total number of ES cells over-expressing K71R or WT Chd1l. B.

Colony Survival. Tetracycline was removed and cells were seeded at clonal density and

then cultured for six days. The number of colonies obtained for ES cells expressing

K71R or WT Chd1l was reduced by ~60%. C. Colony Morphology. Representative

images of pluripotent or differentiated ES cell colonies expressing Chd1l-K71R or held in

the presence of Tetracycline to repress transgene expression. D. Quantitation of Colony

Morphology. About 95% of colonies expressing either K71R or WT forms of Chd1l

appear undifferentiated, similar to control colonies held in the presence of Tetracycline.

E. Cell Cycling. The percentage of cells in G1, S, or G2/M phase in each population of

ES cells expressing K71R or WT forms of Chd1l is compared to control ES cells cultured

in the presence of Tetracycline. F. Apoptosis. Fixed, non-permeabilized cells were

labeled with PI or AnnexinV-FITC and analyzed by flow cytometry. The percentage of

cells that labeled AnnexinV+/PI- are shown for ES cells expressing either K71R or WT

versions of Chd1l or control cells cultured in the presence of Tetracycline.

96

Figure 4.3. Chd1l over-expression specifically kills undifferentiated ES cells

A. Competition in 3T3 cells transiently transfected with Chd1l. Transfected 3T3 cells

were selected using FACS sorting on GFP-expressing cells and then cultured for an

additional two days. The percentage of cell that retain the empty vector or vector

containing K71R or WT Chd1l is shown (GFP+). The cells that have lost the vector (i.e.

through dilution upon cell division) are GFP-. Cells were stained with PI to label dead

cells. B. Experimental Design of ES cell differentiation assay. Tetracycline was

γH2AX

γH2AX

γH2AX+Hoechst γH2AX+Hoechst

γH2AX+Hoechst

Competition in 3T3 Cells

A B

Figure 3. Chd1l over-expression kills ES cells only

C Cell Number in Differentiating ES Cells

Experimental Design

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removed from ES cells differentiated for 0, 2, or 4 days by removal of LIF (D0 LR, D2

LR, or D4 LR). Differentiating ES cells were cultured for an additional six days in the

absence of Tetracycline and in the absence of LIF. C. Results of ES cell differentiation

assay. Cell number was assayed after 0, 2, or 4 days of LIF removal and an additional 6

days of LIF removal and over-expression of K71R or WT Chd1l. Chd1l over-expression

kills ES cells cultured in the presence of LIF and “D0 LR” differentiating cells. Neither

“D2 LR” or “D4 LR” differentiating cells are susceptible to over-expression of Chd1.

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Figure 4.4. Size Fractionation of a Chd1l-containing protein complex

ES cell nuclear extracts were submitted to size fractionation of a glycerol gradient

ranging from 10% 5o 30%. Fractions were run on SDS-PAGE and blotted for Chd1l. A

Chd1l-containing complex is smaller than the Brg1-containing BAF complex (~2 mDa)

and larger than the Oct4/Sox2 complex (<440 kDa), and is estimated at ~500 kDa. Chd1l

cofractionates with the DNA repair protein PARP-1.

Figure 4. Size Fractionation of Chd1l complex

γH2AX γH2AX

γH2AX

99

Figure 5. α-γH2AX Staining

A. Chd1l MO-1 Injected

γH2AX γH2AX

γH2AX


γH2AX+Hoechst

C. Uninjected

γH2AX γH2AX

γH2AX


γH2AX+Hoechst

100

Figure 5 cont.: α-γH2AX Staining

B. Chd1l MO-2 Injected

γH2AX γH2AX

γH2AX


γH2AX+Hoechst

C. Uninjected

γH2AX γH2AX

γH2AX


γH2AX+Hoechst

101

Figure 4.5. γH2AX staining in embryos injected with Chd1l-MO.

Embryos injected with Chd1l MO-1 (A) or Chd1l MO-2 (B) were fixed 2 days after

injection (E3.5) and stained with α-γH2AX . Embryos injected with MO-1 arrest at the

8- to 16- cell stage. Embryos injected with MO-2 arrest at the 2- to 4- cell stage.

Uninjected embryos at the morula stage (E3.5) were fixed and stained in parallel with

MO-injected embryos (C, shown twice for comparison). Images are taken from 63x oil

immersion confocal microscopy.

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Figure 6. α-PAR Staining

C. Uninjected

PAR PAR PAR

PAR+Hoechst PAR+Hoechst PAR+Hoechst

A. Chd1l MO-1 Injected

PAR PAR PAR


103

Figure 6 cont.: α-PAR Staining

C. Uninjected

PAR PAR PAR


B. Chd1l MO-2 Injected

PAR PAR PAR


104

Figure 4.6. PAR staining in embryos injected with Chd1l-MO.

Embryos injected with Chd1l MO-1 (A) or Chd1l MO-2 (B) were fixed 2 days after

injection (E3.5) and stained with α-PAR. Embryos injected with MO-1 arrest at the 8- to

16- cell stage. Embryos injected with MO-2 arrest at the 2- to 4- cell stage. Uninjected

embryos at the morula stage (E3.5) were fixed and stained in parallel with MO-injected

embryos (C, shown twice for comparison). Images are taken from 63x oil immersion

confocal microscopy.

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5. General Discussion

Chd1l is Essential in the Preimplantation Embryo

Chd1l has stage and cell-type specific expression patters during embryonic

development [258]. There are many nuclear reprogramming and chromatin remodeling

activities that occur during embryogenesis, and many of them are not well understood.

To address the question of whether Chd1l could be a novel player during chromatin

remodeling during early embryogenesis, Chd1l was knocked-down using synthetic,

antisense oligos called morpholinos (MOs). The morpholino system has the advantage of

being able to do rapid reverse genetic studies in a cell type where maternal effect proteins

usually mask the phenotypes of typical knock-out mice.

Microinjection of MO targeting Chd1l in pronuclear zygote stage embryos

resulted in arrest of the embryo. Several MOs were used, all of which caused arrest prior

to the formation of the blastocyst, some as early as the two cell stage. Knock-down was

confirmed at the transcript level by measuring the abundance of Chd1l transcripts that

were abrogated at the splice junction targeted by the MO. The early arrest phenotype was

partially rescued by co-injection of MO with mRNA encoding Chd1l. Because cDNA

sequences lack introns, the mRNA was not targeted by the MO. Embryos injected with

MO that result in a very early arrest between the two and four cell stage progressed

through several more cell divisions with co-expression of Chd1l mRNA, some of them

obtaining blastocyst formation. These results show that Chd1l is required for an early

chromatin remodeling even during preimplantation development.

The reason behind the early arrest remains unknown. Knock-down of Chd1l in

ES cells followed by genome-wide expression profiling revealed no statistically

significant gene expression changes, and therefore could not provide any clue as to a

Chd1l function in the embryo. Gene expression profiling in preimplantation embryos in

which Chd1l has been knocked-down has not been attempted but may reveal changes in

expression of genes involved in key regulatory processes. Alternatively, the function of

Chd1l may not be at the transcriptional level. If Chd1l participates in the DNA damage

response in embryos as it does in cultured cell types, its mode of action could be entirely

106

post-translational, dealing exclusively with altering chromatin accessibility for DNA

repair factors.

One intriguing possibility is that Chd1l regulates parental genome

reprogramming. Upon fertilization, the sperm chromatin decondenses and undergoes

extensive and rapid chromatin-dependent reprogramming [143]. DNA methylation is

actively and globally reduced except at imprinted loci [163], and protamines are removed

and replaced with replication-independent histone variant H3.3, acetylated histone H4,

and histone H4 monomethylated at lysine 20 [157]. This process is completed within

eight hours after fertilization, just prior to the formation of pronuclear membranes. In

addition, the paternal genome must recover from its “frozen state” and repair endogenous

DNA damage accrued prior to fertilization and/or upon decondensation in the zygotic

environment.

DNA demethylation has been studied for decades, but despite the molecular

pathway being well characterized in plants, it remains ill-defined and nebulous in animals

[166,167]. It was recently shown that 5-meC demethylation occurs through the base

excision repair (BER) pathway in preimplantation embryos [176]. Because Chd1l is

known to interact with BER and DSB repair proteins and to respond to DNA damage by

localizing to lesions [259,260], its role in facilitating DNA demethylation represents an

attractive hypothesis. However, given the timing of embryo microinjection at

approximately 10 hpf, the Chd1l arrest phenotype is probably not consistent with a role

for Chd1l in active DNA demethylation, which is already complete by 8hpf [162,163].

For Chd1l to have a role in active DNA demethylation, a significant delay would have to

occur between excision of 5-meC and repair of the abasic site.

X-inactivation and reactivation events are also important features of embryonic

reprogramming. The parental X chromosome is delivered to the zygote in the inactive

form and is re-activated upon initial sperm remodeling, supposedly as part of the global

demethylation process although this has not been determined [23,222]. The Xp

undergoes imprinted XCI at the four cell stage and remains inactive in all cells of the

cleavage-stage embryo and the trophectoderm upon blastocyst formation [221,223,233].

The early ICM, however, specifically reactivates Xp and initiates random XCI at the peri-

implantation stage [223]. The timing of the Chd1l arrest phenotype (as early as the two

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cell stage and as late as morula stage) would not be consistent with any XCI (four cell

stage or peri-implantation) or X reactivation (pre-pronuclear formation or early ICM

formation) event.

In contrast, a role of Chd1l in the repair of paternal DNA damage is a temporally

relevant hypothesis. Repair of damaged paternal DNA (and maternal DNA) occurs

during the extended ~20-24 hours of the first cell cycle and during sequent cleavages

[240,336]. Failure to repair paternal chromatin in the zygote is a leading cause of human

infertility [237,251,310,341]. Irradiation of zygote stage embryos results in chromosomal

abnormalities and arrest prior to the blastocyst stage, demonstrating that the presence of

DNA damage in the zygote is deleterious to the embryo, but that development may

progress through several cell cycles before arrest occurs [241,242]. This phenotype,

although non-specific, resembles that seen in embryos microinjected with Chd1l-MO.

The zygote is distinctive from other embryonic cell types in that it has an extended G1

phase in which NHEJ predominates [336]. The potential of Chd1l to contribute to NHEJ

is supported by the observation that upon induced DNA damage in HEK293 cells, Chd1l

associates with DNA-PKcs, a major component of NHEJ repair [259]. Chd1l may also

contribute to other modes of DNA repair, including HRR, which functions during S/G2

phases, and BER, although the evidence for these pathways is not as strong as for NHEJ.

If a Chd1l arrest phenotype is due to impaired DNA repair, particularly repair of

DSBs, it would be expected that arrested embryos would display an accumulation of

DSBs. Immunocytochemistry staining of control embryos and embryos injected with

Chd1l-MO using γH2AX as a marker for DSBs revealed that some, but not all, cells of

arrested embryos had a higher frequency of DSBs. Further analysis is necessary to

analyze the presence of DSBs in Chd1l arrested embryos, although this is not an easy

task. Single-cell gel electrophoresis, or the comet assay, is often used in cultured cells to

detect double- and single-stranded breaks, but even with ample material, error values are

extremely high. Novel in vitro approaches may be necessary to test repair of lesions in

control embryos and Chd1l-arrested embryos.

Another temporally viable hypothesis is that Chd1l contributes to the maternal-

zygotic transition through chromatin remodeling leading to zygotic genome activation.

The primary events that trigger ZGA are still under investigation, but initiation of

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expression of at least three prominent classes of genes is known to require chromatin

remodeling by Brg1 [186]. The foundation for this second hypothesis is tenuous because

there is no current evidence to suggest that Chd1l regulates gene expression through

chromatin remodeling. Expression analysis in control embryos and embryos injected

with Chd1l-MO should reveal whether Chd1l-mediated gene regulation is a feasible

hypothesis.

Due to the inherent difficulties associated with embryo microinjection, namely,

physical insult to the embryo and low statistical power, mutant mouse models should be

created to corroborate the findings presented here. Because maternal contribution is

likely to be a factor, a strategy that allows conditional knock-out in the zygote should be

employed. With the use of Chd1l-null zygotes, the critical window of time in which

Chd1l can be more accurately defined and levels of PAR or γH2AX can be more reliably

assessed.

Chd1l is Non-Essential in ES cells

ICM enrichment and expression in ES cells suggest that an essential role for

Chd1l could be found in ES cell pluripotency or subsequent differentiation. To address

this question, inducible ES lines were created that expresses a shRNA when tetracycline

is removed from culturing medium. cDNA encoding a shRNA targeting Chd1l was

stably integrated into the Rosa locus to create the Chd1l-shRNA EBRTcH3 ES cell line.

As a control, cDNA encoding a “non-silencing” shRNA was also integrated to create the

NS-shRNA eBRTcH3 ES cell line.

Chd1l-shRNA ES cells cultured in the absence of tetracycline for two days show

efficient knock-down of Chd1l. There were no appreciable differences in growth rates

compared to induced NS-shRNA ES cells over a period of eight days, and no

morphological differences in the formation of colonies grown for six days. To address

whether gene expression patterns were altered even in the absence of a readily apparent

phenotype, the expression profiles of ES cells expressing either Chd1l-shRNA or NS-

shRNA were examined. Surprisingly, there were no significant changes in gene

expression due to knock-down of Chd1l. While Chd1l may not function to regulate gene

expression in pluripotent ES cells, it may be critical for regulating gene expression

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patterns during differentiation. To test this, ES cells were used to form into embryoid

bodies (EBs), differentiating aggregates of cells that mimic embryonic development.

EBs expressing Chdl1-shRNA or NS-shRNA were analyzed for the formation of the

three primary germ layers, endoderm, mesoderm, and ectoderm. Quantitative PCR using

a panel of lineage markers revealed that all three germ layers formed in EBs expressing

Chd1l-shRNA, similar to EBs expressing NS-shRNA.

Protein analysis of ES cells expressing Chd1l-shRNA reveals that knock-down

was very efficient. Although it remains possible that a small amount of Chd1l remaining

is sufficient to carry out the normal Chd1l functions in ES cells, the data indicate that

Chd1l is non-essential for ES cell viability, proliferation, pluripotency, and

differentiation. Furthermore, genome-wide chromatin immunoprecipitation (ChIP) failed

to identify genomic loci bound by Chd1l. The finding that the α-Chd1l antibody did not

pull down any direct Chd1l targets has two explanations. The first is that Chd1l does not

bind discrete genomic loci, and the second is that the α-Chd1l antibody did not

immunoprecipitate cross-linked Chd1l. Because no direct Chd1l targets have been

identified in any cell type have, validation of the α-Chd1l antibody was impossible.

Given that gene expression does not change upon Chd1l knock-down in ES cells, it is

likely that Chd1l also does not bind discrete loci.

After completion of these studies, two independent laboratories reported their

findings that Chd1l is involved in the DNA damage response. If Chd1l also functions in

the DNA damage response ES cells, the inability to identify Chd1l target genes in ES

cells could be explained by semi-random binding to sites of endogenous DNA damage

rather than discrete gene regulatory loci. Prior knowledge of the role of Chd1l in DNA

damage response would not have precluded investigation into an additional role in gene

expression, because other chromatin remodeling factors, such as INO80, have dual roles

in DNA repair and regulating gene expression. However, given the data presented here

and in the literature, Chd1l appears to have a role in DNA repair but not in transcriptional

regulation.

110

Role of Chd1l in DNA Repair in ES cells

Studies conducted early in this dissertation project demonstrate that over-

expression of Chd1l in ES cells induces widespread cellular death through apoptosis.

Intriguingly, Chd1l-induced cell death was a phenotype specific to undifferentiated ES

cells. ES cells that had been differentiated for two days or more by removal of LIF prior

to Chd1l over-expression were resistant to cell death, and even had increases in cell

number compared to control ES cells. Differentiation mediated by LIF removal is a

challenging task for ES cells, so it is not clear whether the increase in cell number reflects

greater proliferation or increased survival in differentiating cells over-expressing Chd1l.

These data suggest that at some unspecified key point during differentiation, the function

of Chd1l, or the cellular response to Chd1l, changes.

It is worth mentioning that over-expression of wild-type Chd1l or Chd1l

containing a single amino acid mutation predicted to eliminate ATPase activity (K71R)

resulted in nearly indistinguishable phenotypes for all parameters tested, including cell

death, differentiation, proliferation, and apoptosis. Therefore, ES cell death caused by

Chd1l over-expression does not depend on a functional ATPase domain, but may instead

result from the effects of over-expressing the macro domain. This line of research was

not pursued, but it would be interesting to know whether expression of the Chd1l macro

domain alone also triggered apoptosis in ES cells. Over-expressing either form of Chd1l

was predicted to result in a loss of function phenotype due to disrupted stoichiometry of a

Chd1l-containing chromatin remodeling complex. Hypermorphic or neomorphic

phenotypes could result due to over-expression of a macro domain-containing protein, a

possibility made more likely given that Chd1l-induced cell death was independent of the

ATPase domain.

A more reliable loss of function manipulation was pursued by expressing a

shRNA directed against the Chd1l transcript. Unexpectedly, the phenotype of Chd1l

knock-down in ES cells was very different than Chd1l over-expression; that is, there was

no readily apparent phenotype. However, when ES cells expressing Chd1l-shRNA or

NS-shRNA were treated with DNA damaging agents, they displayed damage resistance.

This effect was readily apparent when the radiomimetic drug phleomycin was used, and a

more subtle but reproducible affect was seen when H2O2 treatment was used. These

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results suggest that Chd1l participates in the decision to induce apoptosis in response to

DNA damage, and that ES cells are more sensitive to radiomimetic damage than

oxidative damage, as has been proposed previously.

The data presented here are consistent with studies demonstrating that Chd1l

participates in the DNA damage response, but how Chd1l responds to DNA damage

appears to be very different in ES cells that readily undergo apoptosis in response to

DNA insults than in other cultured cell types. In human U2OS cells, loss of CHD1L

contributes to damage sensitivity, presumably because cells are deficient in repairing

DNA. In mouse ES cells, loss of Chd1l contributes to damage resistance, presumably

because cells are deficient in initiating apoptosis. The apparent paradox between Chd1l

function in ES cells and in other cell types brought to attention the previous over-

expression studies demonstrating that too much Chd1l induced apoptosis, but only in ES

cells, and not from their differentiating counterparts. Consistent with this, Chd1l over-

expression in 3T3 fibroblasts also does not cause decreased cellular survival. Therefore,

the data show that Chd1l functions very differently in ES cells than in other cell types.

There are many remaining questions regarding the role of Chd1l in DNA repair.

In U2OS cells, CHD1L responds to DNA damage by rapidly localizing to sites of DNA

damage. This localization was shown to depend on PARP1 activation and a functional

macro domain. Chd1l dissociation from damaged sites was assumed to reflect successful

DNA repair and was shown to depend on a functional ATPase domain. However, a

direct role for Chd1l in the repair process was not shown; thus the molecular consequence

of Chdl1 localizing to sites of DNA damage remains unknown.

Because early over-expression studies were not pursued, it remains unknown

whether Chd1l-induced apoptosis in ES cells occurs through a Parp-1- or AIF-dependent

manner. This is a fascinating question because Parp-1 has also been implicated in the

apoptotic damage response through the AIF pathway, but this phenomenon is not well

understood. Mice deficient in Parp-1 through mull mutations show increased damage

resistance after ischemia and reperfusion. This role in mediating apoptosis is unique to

certain cell types and is in contrast the typical role of Parp-1 in responding to DNA

damage by facilitating DNA repair. Why Parp-1 triggers apoptosis in certain cell types

but not in others is unclear but probably has to do with cellular NAD+ levels and specific

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tolerance to damage. In particular, it is not known how activation of Parp-1 in the

nucleus triggers AIF release from the mitochondrion. That Chd1l participates in the

decision to initiate apoptosis in ES cells but not in other cell types is reminiscent of Parp-

1 function and raises the intriguing possibility that Chd1l participates in the decision to

initiate apoptosis by responding Parp-1-synthesized PAR modifications through the

macro domain. Chd1l may be a piece of the puzzle, but because Chd1l is a nuclear

protein, there would still be a missing link to explain how the Parp-1 signal is transmitted

to the mitochondrion.

In U2OS cells, Chd1l becomes tightly associated with chromatin upon DNA

damage. Localization to lesions depends on the macro domain but not the ATPase

domain. In ES cells, over-expression of either WT or K71R versions of Chd1l, both of

which have a functional macro domain, may have the hypermorphic affect of localizing

to chromatin even in the absence of induced DNA damage. Because cell death induced

by over-expression of Chd1l occurred independently of the ATPase domain, the

mechanisms of Chd1l in initiating apoptosis is independent of nucleosome remodeling

but may still depend on a functional macro domain and localization to chromatin.

Consistent with this hypothesis, ES cells expressing Chd1l-shRNA and treated with DNA

damaging agents may be deficient in initiating apoptosis because Chd1l is not present to

localize to lesions. Thus a model is proposed in which Chd1l responds to DNA damage

in all cell types, but the signal that is propagated by its response is different in ES cells

than in other cell types. Upon DNA damage, Chd1l localizes to lesions. In ES cells that

have high propensity to choose apoptosis, this signal is interpreted as a trigger for cell

death; whereas in other cell types, the signal facilitates DNA repair, most likely through

chromatin relaxation and nucleosome mobilization.

Oncogenic Potential of Chd1l

Given that over-expression of Chd1l induces cell death in ES cells, reports that

Chd1l functions as an oncogene were surprising. The evidence supporting the oncogenic

properties of CHD1L is compelling. CHD1L was identified as a candidate oncogene

because its genomic amplification and over-expression is associated with over 50% of

human hepatocellular carcinoma patients [255]. Transient over-expression of Chd1l in

113

U2OS cells and in primary liver cells increases colony formation in soft agar assays, and

xenografts of CHD1L over-expressing cells form multiple teratomas [261]. Transgenic

mice over-expressing CHD1L develop spontaneous tumors with high frequency (25%),

and MEFs generated from transgenic mice showed decreased levels of the tumor

suppressor proteins p53 and Rb and increased levels of cell cycling proteins Cdk2 and

Cyclin A [262].

Somewhat paradoxically, over-expression of Chd1l in ES cells induces

widespread apoptosis. U2OS cells, when over-expressing CHD1L and upon phleomycin

treatment, accumulate of double-stranded breaks, as measured by γH2AX flow cytometry

and single-cell gel electrophoresis (comet) assays [259]. This result provides a

mechanism by which CHD1L could promote tumorigeneis. Over-expression of some

well characterized oncogenes such as ras, myc, cyclin E, mos, cdc25A, and E2F1 has

been observed to induce DSBs because cells are forced through the cell cycle before

lesions can be repaired [263]. The accumulation of DSBs results in genomic instability,

and when mutations in tumor suppressor genes arise, cancer can develop. Chd1l over-

expression could result in the accumulation of DSBs in ES cells as it does in U2OS cells,

and this could result in genomic instability. Because ES cells have a higher propensity to

undergo apoptosis in response to DNA damage, this genomic instability could trigger

apoptosis in ES cells but lead to oncogenesis in other cell types [116,117,118].

Over-expression of DNA repair genes sometimes confers protection to

cytotoxicity and mutagenic agents; but many times it does not [305,342]. PARP-1 is

among the genes whose over-expression does not. Instead, an increase in sensitivity to γ-

irradiation was observed [343]. Deleterious effects including increased rates of

spontaneous mutations and sensitivity to various DNA damaging agents is observed with

over-expression of the NER repair protein ERCC-1 [344], the BER repair proteins DNA

polβ [345,346] and ANPG (a glycosylase) [347], the HRR repair protein Rad51 [348].

The mechanisms by which over-expression of each of these proteins disrupt repair are

unique and depend on their specific activity during DNA repair. For example, over-

expression of the HRR repair protein Rad51 lead to unusual recombination events

resulting in translocations and genomic instability [348], whereas over-expression of

ANGP is thought to create an excess of AP sites and gapped DNA [347].

114

Therefore, the mechanisms by which over-expression of DNA repair proteins can

lead to increased DNA damage and genomic instability are many, and without further

studies it is difficult to predict a mechanism by which over-expression of Chd1l could

induce genomic instability in ES cells. Chd1l-induced apoptosis does not depend on a

functional ATPase domain, but may be a consequence of specifically over-expressing the

macro domain. Because the macro domain is responsible for localization to chromatin

upon DNA damage it is possible that over-expression of Chd1l has a hypermorphic

effect, causing aberrant localization to chromatin, and that this interferes with DNA

repair or signals to the ES cell apoptosis pathway. Further studies are necessary to test

this hypothesis.

Paradoxes in Chd1l Function in ES Cells

Knock-down of Chd1l in preimplantation embryos results in lethality prior to the

blastocyst stage, demonstrating that Chd1l is essential during early embryogenesis.

Given this result, it was surprising that ES cells in which Chd1l had been efficiently

knocked-down were viable, proliferated normally, did not differentiate, and did not have

altered gene expression patterns. In contrast, over-expression of Chd1l in ES cells

induces apoptosis. Paradoxically, Chd1l over-expression in transgenic mouse models

leads to spontaneous tumor formation in combination with increased levels of some cell

cycling proteins and decreased levels of tumor suppressor genes. Transgenic Chd1l mice

grew normally to term without changes in body size [262].

ES cells expressing Chd1l-shRNA display decreased sensitivity to treatment with

DNA damaging agents H2O2 and especially phleomycin. This result is in sharp contrast

with published studies demonstrating that other cultured cells are increasingly sensitive to

induced DNA damage in cells with reduced Chd1l [259]. It is rather the over-production

of Chd1l in ES cells that experience widespread cell death through apoptosis. These

phenotypes are summarized in Figure 1.

How can these conflicting results be resolved? The common theme from the data

is that results in ES cells conflicts with results in other systems. That early embryos with

reduced Chd1l arrest prior to the blastocyst stage is not in conflict with the role of Chd1l

as an oncogene or as a DNA damage response protein. Therefore, the explanation might

115

be that Chd1l has a very different function in ES cells than it does in any other cell type.

The unique cell cycle, DNA repair strategies, and apoptotic response to DNA damage in

ES cells support this theory. The underlying mechanism could be that Chd1l functions

primarily through NHEJ repair pathway, a pathway that is predominantly used in the

zygote-stage embryo and in differentiated cells. In contrast, pluripotent ES cell primarily

use HR to repair DNA, and are easily triggered to apoptose in response to DNA damage.

The involvement of Chd1l in NHEJ is suggested by its PARP-dependent, DNA damage-

dependent association with a major NHEJ component DNA-PKcs. This represents an

interesting hypothesis that should be tested.

116

Figure 5.1 Chd1l Phenotypes

Reduction of Chd1l in the zygote through morpholino microinjection results in

embryonic arrest prior to the blastocyst stage. What the effects of loss of function are in

the developing post-implantation animal or in the adult remains unknown. Chd1l over-

expression in transgenic mice results in no reported developmental abnormalities, but

adult mice have higher spontaneous tumor formation [262]. In tissue cultured cells, over-

expression of Chd1l and treatment of the radiomimetic drug phleomycin leads to an

accumulation of double-stranded breaks as measured by γH2AX flow cytometry and

comet assays [259]. Over-expression of Chd1l in ES cells leads to widespread cell death

in the absence of induced DNA damage. Apoptosis in response to Chd1l over-expression

is specific to ES cells and does not occur after ES cell differentiation by LIF removal.

Knock-down of Chd1l in ES cells does not result in impaired pluripotency, proliferation,

or differentiation, but confers damage resistance when ES cells are treated with DNA

damaging agents. In contrast, knock-down of Chd1l in differentiated tissue cultured cells

leads to increased hypersensitivity to treatment with DNA damaging agents.

117

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