recruitment of transcription complexes to the …

RECRUITMENT OF TRANSCRIPTION COMPLEXES TO THE BETA-GLOBIN

LOCUS IN VIVO AND IN VITRO

By

KAREN FRANCES VIEIRA

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2004

Copyright 2004

by

Karen Frances Vieira

This document is dedicated to my mother and sister who have been there for me every step of the way, and to my father who is gone but not forgotten.

iv

ACKNOWLEDGMENTS

I would like to first acknowledge God as the guiding light in my life, for without

Him my world would be empty.

I owe a huge thanks to my mentor, Dr. Jörg Bungert, for the opportunity to work in

his lab and for receiving the excellent training that I did. He has been a great mentor,

motivator, advisor and teacher. He has been patient with me and very supportive of my

alternate career choices. He is the hallmark of a good scientist, and I respect and am

inspired by his commitment and dedication.

I thank all of my committee members, Drs. Michael Kilberg, Thomas Yang, Linda

Bloom, Hideko Kasahara, and James Resnick. They have all been a great help in

providing suggestions and directions for my experiments and career. Drs. Kilberg and

Yang have been especially supportive and encouraging throughout my time at the

University of Florida, and have always had insightful comments during my committee

meetings and presentations. I need to thank them as well as Dr. Brian Cain for critically

evaluating and helping me with my presentation that won first place in the 2004 Medical

Guild Research Competition. I appreciate Dr. Resnick joining my committee at such

short notice, and also providing useful information to help me get on track for graduation.

I extend my appreciation to Dr. James B. Flanegan, the department chair, and all

the administrative and secretarial staff that has made me at home in the department. I

would like to say a special thanks to Bradley Moore who always shares a smile or a

compliment to brighten my day.

v

I thank all of the past and present members of the Bungert lab. When I first started

in the laboratory, Kelly Leach and Padraic Levings instantly made me feel comfortable

and were great mentors to me. I appreciate Kelly for setting the standards, which I have

since tried to meet or surpass. Sung-Hae Lee Kang also was a mentor and a great source

of encouragement and advice as well as a great friend. Ara Aslanian is the post-doctoral

researcher who, upon leaving, passed on his protocols and projects to me. Since that was

the first step in my successes here at the University of Florida, I owe Ara many thanks.

Christof Dame was a post-doc in the laboratory that encouraged me with helpful,

thoughtful and sometimes amusing discussions. I would like to thank Valerie Crusselle

for being a great colleague and also a friend. I wish her many blessings. Takeesha

Roland has been the best laboratory technician I know and I appreciate her help, patience

and sincere friendship. I need to thank the many other members of the lab including

Boris Thurisch, Felicie Andersen, Meredith Hill, and the many undergraduates and high

school students who have helped and entertained me along the way.

I thank the members of Dr. Yang’s lab who have always been helpful and friendly,

especially Christine Kiefer and Sara Rodriguez, who both helped me with footprinting

experiments. I would like to say a special thanks to the members of Dr. Kilberg’s lab,

especially Hong Chen who has been a great source of knowledge.

I would like to thank my family and friends who have been very supportive. My

mother and sister have always been my best source of support and encouragement, and I

want them to know that I love and appreciate them. I thank my boyfriend and many

friends, who have helped me tremendously through the stresses of graduate school, and

have been patient with my many moods during this long journey.

vi

TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................................................................................. iv

LIST OF FIGURES ........................................................................................................... ix

ABSTRACT....................................................................................................................... xi

CHAPTER

1 INTRODUCTION ........................................................................................................1

Hemoglobin and Sickle Cell Disease ...........................................................................1 Overview of Sickle Cell and Thalassemia.............................................................1 Symptoms and Treatments ....................................................................................2 Current Research on Therapies .............................................................................3

Chromatin Structure......................................................................................................4 Histone Modifications ...........................................................................................6 Chromatin Remodeling .........................................................................................7

Mechanism of Transcription.........................................................................................8 Preinitiation Complex Assembly...........................................................................9 Initiation ................................................................................................................9 Elongation............................................................................................................10

The β-Globin Gene Locus ..........................................................................................11 Location and Organization ..................................................................................11 Locus Control Region..........................................................................................12

Proteins Associated with the β-Globin Locus ............................................................14 Model for β-Globin Gene Regulation.........................................................................16 Questions to Be Addressed.........................................................................................19

2 MATERIALS AND METHODS ...............................................................................21

Cell Culture.................................................................................................................21 Chromatin Immunoprecipitation (ChIP).....................................................................21 Chromatin Immunoprecipitation and DMS Footprinting ...........................................24

DMS Treatment ...................................................................................................26 Linker LMPCR....................................................................................................26

Western Blotting.........................................................................................................29 Cell Synchronization ..................................................................................................30

vii

PCR-Based DNA Replication Analysis......................................................................30 DNA Isolation .....................................................................................................30 Semi-Quantitative PCR .......................................................................................31

In Vitro Polymerase Transfer Analysis.......................................................................32 MEL Cell DMSO Differentiation...............................................................................33 Semi-Quantitative RT-PCR........................................................................................33 Double Immunoprecipitation......................................................................................34 Co-Immunoprecipitation.............................................................................................36

3 CHROMATIN IMMUNOPRECIPITATION AND DMS FOOTPRINTING...........38

Introduction.................................................................................................................38 Results.........................................................................................................................40 Discussion...................................................................................................................43

4 INTERACTION OF RNA POLYMERASE II AND TRANSCRIPTION FACTORS

WITH THE β-GLOBIN LOCUS ...............................................................................46


5 ANALYSIS OF THE MECHANISM OF RNA POLYMERASE II TRANSFER

FROM THE LCR TO THE β-GLOBIN GENE PROMOTER ..................................65


6 DETERMINING THE FUNCTION OF THE USF, TFII-I AND HDAC3

PROTEINS IN β-GLOBIN GENE REGULATION..................................................76


7 CONCLUSIONS AND FUTURE DIRECTIONS .....................................................87

ChIP Footprinting .......................................................................................................87 ChIP within the β-Globin Locus.................................................................................88 Cell Synchronization ..................................................................................................89 Chromosome Conformation Capture..........................................................................90 In Vitro Pol II Transfer ...............................................................................................91 USF, TFII-I and HDAC3 Function in β-Globin Regulation ......................................92 Summary.....................................................................................................................93

viii

LIST OF REFERENCES...................................................................................................94

BIOGRAPHICAL SKETCH ...........................................................................................113

ix

LIST OF FIGURES

Figure page 1-1. The human and mouse β-globin loci........................................................................13

1-2. Multistep model for human β-globin gene regulation..............................................18

3-1. Sequence alignment of the adult β-globin downstream promoter region. ...............41

3-2. Analysis of protein–DNA interactions in the murine β-globin downstream promoter region by a combination of chromatin immunoprecipitation and DMS footprinting...............................................................................................................42

3-3. ChIP experiment showing the interaction of USF1 and NF-E2 with the murine βmaj-globin promoter in MEL cells.........................................................................43

3-4. LMPCR footprint analysis of non-selected or ChIP-selected chromatin. ................44

4-1. Diagrammatic representation of the human and murine β-globin gene locus. ........49

4-2. Interaction of transcription factors and RNA polymerase II with the human β-globin locus and the human necdin gene. ................................................................50

4-3. Interaction of transcription factors with and transcription in the murine β-globin locus in MEL cells....................................................................................................52

4-4. Interactions of transcription factors and RNA polymerase II with the human β-globin locus during early S phase in synchronized K562 cells................................54

4-5. Interactions of transcription factors and RNA polymerase II with the human β-globin locus during early S phase in synchronized MEL cells. ...............................56

4-6. Semi-quantitative PCR analysis of DNA content of synchronized K562 cells. ......57

4-7. Semi-quantitative PCR analysis of DNA content of synchronized MEL cells........58

5-1. A schematic representation of the in vitro Pol II transfer experimental procedure. 67

5-2. Transfer of RNA polymerase II from immobilized LCR constructs to the β-globin gene............................................................................................................68

x

5-3. Analysis of the specificity of Pol II transfer. ...........................................................70

5-4. Comparison of Pol II transfer using the wild-type β-globin gene (lane 1) and a mutant β-globin gene (lane 2) in which an E-box at +60 was altered......................70

5-5. Transfer of RNA polymerase II to the β-globin gene in the presence of NF-E2 or BSA. .........................................................................................................................71

5-6. Analysis of NF-E2 mediated transfer to the β-globin gene......................................72

5-7. Quantitative summary of the transfer experiments. .................................................73

5-8. Model of transcription complex recruitment to the β-globin gene. .........................75

6-1. Sequence alignment of the human β-globin downstream promoter region. ............78

6-2. Characterization of protein–DNA interactions in the human β-globin downstream promoter region in vivo........................................................................80

6-3. Chromatin double immunoprecipitation (ChDIP) experiment analyzing the interaction of TFII-I and HDAC3 at different regions within the human β-globin locus. ........................................................................................................................81

6-4. Co-immunoprecipitation experiment analyzing the interaction of TFII-I and HDAC3 as a complex...............................................................................................82

6-5. Summary and model of protein–DNA interactions at the human β-globin promoter. ..................................................................................................................86

xi

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

RECRUITMENT OF TRANSCRIPTION COMPLEXES TO THE BETA-GLOBIN LOCUS IN VIVO AND IN VITRO

By

Karen Frances Vieira

December 2004

Chair: Jörg Bungert Major Department: Biochemistry and Molecular Biology

The β-globin gene locus is the site of many mutations and deletions that cause

sickle cell disease and β-thalassemias. Erythroid-specific, high-level expression of the β-

globin genes is regulated by the locus control region, located far upstream of the genes.

Recent studies show that locus control region core elements recruit RNA polymerase II.

Here, we developed and optimized methods to study protein-DNA interactions

within the β-globin locus. These techniques were used to analyze the interaction of

transcription factors and RNA polymerase II with the β-globin locus in erythroleukemia

cell lines. The data show that transcription initiation complexes are recruited to the LCR

and to the genes. Moreover, RNA polymerase II dissociates during early S phase in

synchronized erythroid cells suggesting that replication disrupts the association of

transcription complexes with the globin locus. The interaction of NF-E2 and USF2

precedes the re-association of RNA polymerase II implicating these proteins in a process

regulating the recruitment of RNA polymerase to the globin locus during replication.

xii

Furthermore, we demonstrate transfer of transcription complexes from immobilized

LCR constructs to the human β-globin gene in vitro. Our data are consistent with the

hypothesis that the LCR and the genes cooperate to recruit transcription complexes to the

globin gene locus.

Studies on the interaction of proteins with the adult β-globin gene promoter

demonstrate that the helix-loop-helix proteins USF and TFII-I associate with DNA-

regulatory elements located at or downstream of the transcription initiation site. TFII-I

interacts with histone deacetylase 3 and binds more efficiently in erythroid cells that do

not express the β-globin gene. In contrast, USF, known to interact with chromatin-

opening activities, associates with the globin gene when it is expressed. These data

suggest that USF and TFII-I regulate β-globin gene expression in an antagonistic fashion.

1

CHAPTER 1 INTRODUCTION

Hemoglobin and Sickle Cell Disease

Hemoglobin is the molecule in red blood cells that transports oxygen to all the

tissues of mammalian organisms. Hemoglobin is composed of two alpha (α) globin

chains and two beta (β) globin chains, each carrying a heme molecule [1]. The heme

molecule contains an iron atom and is responsible for binding the oxygen that is

transported. The α- and β-globin chains are important for the overall structure of

hemoglobin, and thus contribute to its essential functions in gas transport.

Overview of Sickle Cell and Thalassemia

More than 80 point mutations and deletions in the α-globin genes are known to

cause α-thalassemia [2]. These thalassemias have the highest incidence in India, Africa,

and Arabic countries [3]. Similarly, more than 200 mutations and deletions in the β-

globin genes cause sickle cell anemia, sickle SC disease, sickle SD disease (collectively

called sickle cell disease), and β-thalassemias [2]. Sickle cell disease and the β-

thalassemias are the most common monogenetic diseases worldwide, with an estimated

80 million β-thalassemia carriers [4]. There are over 10,000 African-Americans with

sickle cell anemia in the USA alone. In African, Asian, Indian and Middle Eastern

communities, sickle cell disease and β-thalassemia are widespread.

Because some mutations associated with the α- and β-globin gene loci provide

protection from malaria, incidence for thalassemias and sickle-cell disease are historically

2

more common in areas affected by malaria. These regions include Africa, India, and

Mediterranean countries. Heterozygous individuals carrying the sickle cell allele have a

survival advantage since these individuals are generally healthy but reveal resistance to

malaria. In the past individuals homozygous for the disease generally did not live past

childhood. These diseased individuals now benefit from an extended lifespan due to the

increased level of worldwide healthcare.

Symptoms and Treatments

Sickle hemoglobin polymerizes when deoxygenated. The hemoglobin polymers

form alpha helical bundles, which deform red blood cells into a sickle-shape. These

sickle cells that give the disease its name are more rigid than normal red blood cells and

cannot pass through small blood vessels. The resulting blockage of blood vessels causes

intense pain and other side effects.

Patients with sickle cell disease experience a range of complications, including

painful crises, acute chest syndrome, priapism in males, stroke, bone damage, and lung

damage. Many patients undergo monthly blood transfusions to lower the percentage of

sickle red blood cells in their system. This reduces the frequency of painful crises, but

also increases iron concentrations to sometimes-toxic levels. Therefore, along with

transfusions, sickle cell patients undergo iron chelation therapy. Unfortunately, since

chelation therapy is performed by daily intravenous injections there is a low patient

compliance. Though, the recent development of new oral iron-chelating agents may

allow for better compliance [5].

Currently, the only cure for severe hemaglobinopathies is hematopoietic stem cell

transplantation (HSCT). About 71% of thalassemia patients given HSCT from a matched

donor have been cured of their disease [6]. Due to the high risks associated with this

3

procedure, patients receive HSCT only if sickle-related death is imminent [5].

Encouragingly, of the sickle cell patients that have received HSCT, 85% survived

disease-free [7]. Since only about 1% of sickle cell patients meet the criteria for HSCT

and have a suitable donor, alternative therapies are highly sought after.

Current Research on Therapies

Many current trials are performed with drugs that increase the levels of fetal

hemoglobin (HbF) since HbF inhibits the polymerization of sickle hemoglobin (HbS).

The drug 5-azacytidine, which inhibits DNA methyltransferases, has been used in the

past but due to serious side effects is now rarely used. Sodium butyrate acts

synergistically with 5-azacytidine and reactivates γ-globin expression for the production

of HbF [8]. Hydroxyurea, which is known to inhibit DNA synthesis and alter chromatin

structure, is currently the drug of choice to raise HbF in sickle cell patients and to reduce

symptoms associated with the disease [9, 10]. The main known side effect of

hydroxyurea treatment is myelotoxicity, or destruction of the bone marrow. Its long-term

toxicity and effects on survival are yet to be determined. For this and other reasons,

hydroxyurea is reserved only for patients with severe complications.

For patients that do not meet the criteria for HSCT or do not respond to

pharmacological therapies, gene therapy may provide a promising alternative in the

future. Sickle cell disease is caused by a point mutation in the coding region of the β-

globin gene and it was the first genetic disorder in which such point mutations have been

found [11]. Due to the relative simple genetic defect and the ability to isolate and

transfect hematopoietic stem cells, it was anticipated that globin gene disorders would be

among the first for which gene therapy approaches are feasible.

4

Several viruses are being tested as vectors for therapeutic gene transfer of the β-

globin genes to mammalian cells. These include murine retroviruses [12], lentiviruses

[13], human foamy virus [14], and adeno-associated virus [15-18]. Promising results

were obtained using lentivirus-based gene therapy in a mouse model for β-thalassemia

[19], though there is still much to be done before it can become approved as a safe form

of gene therapy for sickle cell and β-thalassemia.

One of the obstacles of using lentivirus is the fact that the DNA integrates into the

genome and there is some evidence that integration into open transcriptionally active

regions is preferred [20, 21]. This enhances the risk of gene disruption (e.g., of tumor

suppressor genes) or activation of endogenous genes (e.g., oncogenes). Adeno-associated

virus is considered to be the safest virus used in gene therapy trials because it does not

appear to cause strong immune reactions and in many reports the DNA remained

episomal [22-25]. The limitation of using AAV for globin gene therapy is the low

packaging capacity. As outlined below, appropriate control of β-globin gene regulation

is complex and requires the presence of many cis-acting DNA elements. Alternative

approaches are to optimize HSCT for making it more broadly applicable and to use novel

mechanisms to induce a high level of fetal globin gene expression.

Chromatin Structure

To comprehend the regulation of the globin genes and potential interactions within

the β-globin gene locus, it is important to understand the chromatin structure in the locus

and its relevance to gene expression.

In order to fit the entire genome into the nucleus of each living cell DNA is highly

constrained and compacted through interactions with proteins called histones. Histones

5

are small basic proteins consisting of a globular domain and a more flexible N-terminus.

DNA organization in the nuclei of eukaryotes involves repetitive packaging into

nucleosomes, which are composed of two each of the histone core proteins H2A, H2B,

H3 and H4 arranged in an octamer around which 146 base pairs of DNA is wrapped [26,

27]. Nucleosomal DNA and its associated proteins interact with other proteins to form

higher order structures in the nucleus; the combination of DNA and associated factors in

the nucleus is termed chromatin.

Nucleosomal arrays along the DNA are proposed to fold into a 30 nm fiber upon

incorporation of the linker histone H1, though the mechanism of this compaction remains

unclear. Higher levels of compaction may occur in genomic domains termed

heterochromatin. Genes located in these dense chromatin domains are generally not

accessible for interacting with transcription factors and are thus suppressed [28, 29].

Even further compaction is required during mitosis, and a protein called condensin helps

convert chromatin into condensed mitotic chromosomes [30]. Conversely, there are

regions of the genome, termed euchromatin, in which the genes are not highly

compacted, leaving them accessible to transcription complexes required for gene

expression.

Although the details of chromatin folding are still unclear, the relative compaction

of any region of the genome can be measured by determining the susceptibility of that

region to digestion by the endonuclease deoxyribonuclease (DNaseI). Regions of the

genome are described as being DNaseI insensitive, sensitive, or hypersensitive.

Insensitive regions like heterochromatin are highly condensed and inaccessible to

DNaseI. Sensitive regions such as euchromatin exhibit a less compacted chromatin

6

structure and are accessible to DNaseI [31]. Transcribed regions of the genome are

usually DNaseI sensitive and may be organized as a 30 nm chromatin fiber. DNaseI

hypersensitive sites are usually 200 to 400 bps long and probably reflect regions in which

nucleosomes have been removed or modified. These sites contain clusters of transcription

factor binding sites and are generally associated with the function of regulatory DNA

regions, e.g., enhancers or promoters [32].

Nucleosomes and other forms of DNA compaction modulate transcription and thus

represent an important factor in gene regulation [33]. Enzymes that modify and remodel

chromatin play an important role in activation and repression of gene expression.

Chromatin modifications are post-translational modifications of the histones.

Alternatively, chromatin remodeling includes changing the location of nucleosomes,

altering histone-DNA interactions, removal of histones, and histone exchange [34-39].

Histone Modifications

Histones can be acetylated, methylated, phosphorylated, ubiquitylated, sumoylated,

and ADP-ribosylated [40]. Originally it was thought that modifications occurred only on

the N-terminal tails of histones that protrude from the nucleosome core, but recent work

has identified modifications within the core regions as well as on the C-terminal tails

[41].

Lysine acetylation by histone acetyltransferases (HATs) has so far been the most

studied of all histone modifications [42-44]. There are 5 families of HATs that are all

large multiprotein complexes. Histone acetylation is a reversible process, catalyzed by

histone deacetylases (HDACs) [45], of which there are 3 distinct families. Several lysine

residues are known to be acetylated, including lysines 9, 14, 18 and 23 on H3, and lysines

5, 8, 12 and 16 on H4. Acetylated histones are usually localized to regions of actively

7

transcribed DNA, therefore making them a mark of active or open chromatin domains,

although there are instances where this is not true [46, 47]. Histone acetylation can also

be associated with transcriptional repression and silencing, as well as recombination [48-

51].

Certain lysine residues can also be methylated by histone methyltransferases

(HMTs). Similarly to HATs and HDACs, there are several families of HMTs, which can

mono-, di-, or tri-methylate lysine residues. Recently, a histone demethylase called

PAD4 has been identified [52], disproving the theory that methylation may be

irreversible [53]. Specific histone methylation patterns correlate with gene activation, as

is the case with methylated lysine 4 on H3 (H3K4) [54]. But histone methylation can

also result in gene repression and heterochromatin formation, as with lysine 9

methylation on H3 (H3K9) [55, 56].

There are several other histone modifications, as mentioned at the beginning of this

section, the function of which has not all been determined. A recent model, called the

histone code hypothesis, postulates that the combinations of histone modifications

interact with other proteins that use this code to execute specific gene expression patterns

[47]. This model explains how the same modification on different residues can have

different outcomes. It also explains how the same modification on the same residue but

in the context of different surrounding modifications can have different consequences,

such as H3 with acetylated lysines correlating with both activation and repression of gene

expression [46].

Chromatin Remodeling

The energy from ATP hydrolysis can be used to loosen DNA-histone contacts in

order to mobilize histones. For this reason, all chromatin remodeling complexes contain

8

an ATPase subunit, which is critical for nucleosome mobilization. Thus far there are 3

known families of ATP-utilizing chromatin remodeling complexes, and possibly many

more to be discovered [57].

Though the mechanism of nucleosome movement has not been established, there

have been several models proposing that the mechanism involves DNA dissociating from

the nucleosome and being replaced by a neighboring piece of DNA. This would yield a

DNA loop that moves in a wave across the surface of the nucleosome, ensuring that as

the histone moves only a small region of contact with the DNA is broken at any one time.

This could also apply to histone contacts being disrupted with one piece of DNA and

reestablished simultaneously on another piece of DNA as histones are relocated within

the genome [57].

Chromatin remodeling complexes also have the ability to improve the accessibility

of chromatin to proteins. This would conceivably be achieved by modifying DNA-

histone contacts. Work with the glucocorticoid receptor Gal4-VP16 and some other

transcriptional activators demonstrated that the activators bind chromatin and then recruit

the chromatin remodeling complexes [58-60]. Other studies show that chromatin needs

to be remodeled before transcription factors interact with their binding sites in regulatory

regions [61]. Further work has provided evidence that there is specificity of activators or

repressors for certain chromatin remodeling complexes (possibly for those they can

physically interact with) [62-65].

Mechanism of Transcription

Activation of gene expression ultimately results in the stable association of RNA

polymerase II (Pol II) and other components of the transcription complex with gene

promoters. The first step of transcription for any gene involves assembly of the pre-

9

initiation complex (PIC). After PIC formation and recruitment of Pol II, the C-terminal

domain (CTD) of Pol II is phosphorylated leading to the progression from initiation to

elongation competent transcription complexes.

Preinitiation Complex Assembly

PIC formation often begins with the binding of the TBP subunit of TFIID to the

TATA box. The binding of TFIID to the TATA box is stimulated by TFIIA, and the

TFIID/TFIIA/DNA complex is recognized by TFIIB, which serves as a bridge between

the pre-initiation complex and Pol II. Pol II is recruited to the PIC together with its

associated factor TFIIF followed by the interaction of TFIIE and TFIIH [66-68].

Evidence suggests that some activators act by recruiting TFIID and TFIIA to the

promoter, and these in turn recruit the Pol II holoenzyme containing RNA Pol II and

other general transcription factors (GTFs) that may already be pre-assembled [69, 70].

Because genes differ in their promoter structure, the formation of the initiation complex

likely varies between genes. For example TATA-less genes use additional proteins to

recruit Pol II and general transcription factors. There may be a stepwise recruitment of

factors to some promoters and recruitment of a Pol II holoenzyme to other promoters.

Initiation

Phosphorylation of the CTD of Pol II signals the transition from preinitiation to

initiation of transcription. CTD phosphorylation is carried out by TFIIH (at serine 5) and

pTEFb (at serine 2), and is thought to be regulated by the mediator complex in yeast or

the CRSP complex in higher eukaryotes [71, 72]. It has been postulated that the

phosphorylation of the CTD breaks contacts between Pol II and some of the GTFs,

therefore allowing Pol II to initiate transcription [68]. Phosphorylation at the two

different serines play different functional roles, since the serine-5 and serine-2

10

phosphorylated Pol II complexes are observed in the early initiation and elongation stages

respectively [73, 74]. It is thought that the phosphorylation status of the CTD allows

association with proteins involved in transcription elongation, histone modification, and

RNA processing.

For some genes chromatin remodeling plays a significant role in transcription,

either for PIC formation or after PIC formation for transcription initiation and elongation

[59, 74-77]. Chromatin alterations involved in transcriptional activation are often limited

to acetylation of histone tails by HATs, though other modifications may also be involved

but have not yet been identified.

Elongation

Efficient elongation requires that the Pol II CTD remains phosphorylated during the

entire elongation process. A hyper-phosphorylated CTD is important for the interaction

of Pol II with RNA processing factors such as capping enzyme, splicing factors and

proteins for transcription termination [78, 79]. The CTD becomes de-phosphorylated

after transcription termination and is then again able to interact with the promoter [68]. A

CTD phosphatase Fcp1 has been identified that dephosphorylates the CTD [80-85],

though the mechanism of dephosphorylation remains elusive.

Several other parameters have been discovered that contribute to transcription

elongation. Histone acetylation is required for efficient transcription elongation in yeast

[86], and seems to be lost immediately after the passage of Pol II [87, 88]. There are

several classes of elongation factors that have been identified. For example, SII

reactivates arrested Pol II by inducing the Pol II’s endonucleolytic activity [89-92]. Paf1

associates with elongating Pol II to recruit histone-modifying activities [93].

11

The β-Globin Gene Locus

The human β-globin genes are expressed in a developmental stage and tissue-

specific manner. Regulation of chromatin structure and stage-specific recruitment of

transcription complexes play imminent roles in the expression of the globin genes.

Location and Organization

The β-globin chains of hemoglobin are coded for by the β-globin gene locus

located on chromosome 11p15.5 in humans and chromosome 7 in mice [94]. The human

β-globin locus contains five genes arranged in the order of their developmental

expression in erythroid cells (Figure 1-1) [95, 96]. At the 5’ end is the ε-globin gene,

which is expressed first during the embryonic stage in erythroid cells in the yolk sac.

Next are the γG- and γA-globin genes that are activated during the fetal stage when the site

of hematopoiesis switches from the embryonic yolk sac to the fetal liver. After birth, the

site of hematopoiesis undergoes a second switch to the bone marrow, coincident with the

expression of the δ- and β-globin genes.

The δ-globin gene is expressed in adults at less than 5% of the level of β-globin

expression. This is due to a mutation in the promoter that prevents high-level expression.

Also, there are two γ-globin genes, which arose by gene duplication during evolution and

their sequences only differ at amino acid 136 [97].

Similar to the human β-globin locus, the mouse β-globin locus has 4 genes whose

expression is switched during development (Figure 1-1). The εγ and βh1 genes are

expressed during the embryonic stage of mouse development in the yolk sac. The βmaj

and βmin genes are expressed in fetal and adult mice in the fetal liver and later in the

bone marrow.

12

The globin genes are relatively small, with three coding exons and two introns

each. The exons of all β-like globin genes code for a total of 146 amino acids. These

exons correspond to the functional domains of the proteins [98, 99]. The introns are

between 117 bp to 1264 bp in size [100].

The entire locus is relatively sensitive to DNaseI in erythroid cells (the only cells

that express the β-globin genes), compared to its DNaseI insensitivity in non-erythroid

cells [101]. DNaseI sensitivity around the individual genes changes based on the

developmental stage, such that expressed genes reveal an increase in DNaseI sensitivity

[102]. Specific histone modification patterns have been detected in the β-globin gene

locus and distinguish active from inactive domains [103], suggesting that chromatin

remodeling activities play a role in the stage-specific transcription of the globin genes.

Locus Control Region

In both the human and mouse β-globin locus there is a prominent element located

from 6 to 22 kb upstream of the ε-globin gene called the locus control region (LCR) [104,

105]. As seen in Figure 1-1, it contains 5 core regions that reveal extremely high

sensitivity to DNaseI in erythroid cells; these are called hypersensitive sites (HS1-5).

LCRs are defined by their ability to enhance expression of linked genes in a tissue-

specific, position-independent and copy number-dependent manner [106]. The β-globin

LCR is required for high-level expression of the genes in erythroid cells. It has been

speculated that one function of the LCR is to provide an open accessible chromatin

structure to the globin genes. However, removal of the human or mouse LCR does not

affect the overall DNaseI sensitivity of the locus [107-109]. Therefore, in the

endogenous locus the LCR does not appear to be responsible for generating a DNaseI

13

sensitive domain. However, it is feasible that the LCR regulates chromatin structure not

through setting up DNaseI sensitivity but through histone modifications and chromatin

remodeling [110]. This is further supported by data showing that β-globin constructs

without the LCR suffer from position-of-integration effects when integrated as transgenes

into heterochromatin, whereas β-globin constructs with the LCR do not suffer from these

position effects [111].

Figure 1-1. The human and mouse β-globin loci. The 5’ hypersensitive sites of the LCR

and the 3’ hypersensitive sites are shown as blue boxes. The genes expressed during the embryonic stage of development are shown as light blue boxes. The genes expressed during the fetal stage of development are shown as green boxes. The genes expressed during adulthood are shown as yellow boxes. The genes expressed in both the fetal and adult stages are shown as orange boxes. The locus as shown here is greater than 100 kb in size and is not drawn to scale.

Stage-specific expression of the genes in the β-globin locus is dependent on their

order relative to the LCR [112, 113]. When the genes were reversed relative to the LCR

in transgenic mice the β-globin gene was expressed at the embryonic stage, whereas ε-

globin expression was abolished [114]. It has been suggested that the genes in the locus

are competitively regulated, where only one gene from the locus can be transcribed at a

14

time [115, 116]. This has recently been hypothesized to be due to the fact that only one

active gene can interact with the LCR at a given time [117].

Evidence suggests that in tissues where β-globin genes are expressed the HS cores

interact to form an LCR holocomplex [20, 118-122]. Due to the high density of protein

binding sites within the HS cores of the LCR, there are many factors that interact with

these regions. The interactions of the proteins, not only with the HS cores but also with

other proteins in other cores, may be responsible for mediating the formation of the LCR

holocomplex. The HS core flanking regions contribute to the activity of the LCR and

may stabilize interactions between core elements [123].

Proteins Associated with the β-Globin Locus

GATA. The zinc finger proteins GATA-1 and GATA-2 are the only two members

of the GATA family of transcription factors that are known to be expressed in erythroid

cells [124]. GATA-1 is one of the first erythroid proteins expressed during red cell

differentiation and it is known to associate with proteins containing histone acetylase

activity [125]. This suggests that GATA-1 may be involved in the initial opening of the

chromatin in the β-globin locus. There are many GATA-binding sequences in LCR HS

sites and in the globin gene promoters. These sites can be bound by either GATA-1 or

GATA-2.

Maf and p45 proteins. Small maf proteins interact with p45 like proteins and bind

to MAREs (maf recognition elements) located within HS2, HS3 and HS4. Small maf

proteins contain DNA binding and protein-protein interaction domains but lack trans-

activation domains. They heterodimerize with the related proteins NF-E2 (p45), Bach1,

NRF1 or NRF2 [126-130] via the leucine zipper. The p45-like proteins are all

15

structurally different, and therefore may have varying functions. Studies suggest that p45

plays an important role in the function of the LCR and in transcription of the genes [127,

131, 132]. Maf/Bach1 heterodimers have been shown to simultaneously bind HS2, HS3

and HS4 and bring these cores together in vitro. The interaction between Bach1

heterodimers is mediated by the BTB/Poz domain and deletion of this domain inhibited

HS site interactions in this assay [133, 134]. Bach1/maf heterodimers function as

transcriptional repressors, whereas p45/maf dimers activate transcription. Recent

evidence suggests that MAREs in the globin locus are first bound by Bach1 heterodimers.

Heme mediates the dissociation of these dimers and thus allows the interaction of the

activator p45/maf to interact with regulatory elements [135].

EKLF. CACCC sequences, also known as KLF (krüppel like factor) binding sites

in HS2, HS3 and globin gene promoters are conserved across many species [136]. These

sites are bound by EKLF (erythroid krüppel like factor). EKLF is required for human β-

globin gene expression and the formation of a DNaseI hypersensitive site in the β-globin

gene promoter [137-140]. This is due to the fact that EKLF binds to the β-globin gene

promoter and recruits chromatin remodeling factors, thus causing expression of the gene

[141]. Similarly, there is an EKLF binding site in the ε-globin gene promoter, but at the

adult stage there are proteins bound at the ε promoter that prevent EKLF binding and

therefore opening of the promoter for ε-globin expression [142].

USF. An E-box, which is a CANNTG consensus sequence, located in HS2 is

known to interact with HLH (helix loop helix) proteins such as the USF (upstream

stimulatory factor) family of proteins and Tal1 [143, 144]. USF is a ubiquitously

expressed transcriptional activator and usually binds to DNA as a heterodimer consisting

16

of USF1 and USF2. USF can also help recruit RNA polymerase II (Pol II) transcription

complexes to initiator elements [145, 146]. Tal1 is an erythroid specific member of the

HLH family of proteins, which has been shown to function during the differentiation of

hematopoietic progenitor cells, and it has recently been shown to interact with TFIIH, a

member of the basal transcription complex [147, 148].

TFII-I. The transcription factor TFII-I binds to two separate elements. First is the

pyrimidine-rich initiator (Inr) element with a consensus of YYA+1NT/AYY located over

transcription start sites [149-151]. Second is the USF recognition site, the E-box. TFII-I

has a unique structure that allows for multiple protein-protein and protein-DNA

interactions [152]. At Inr and E-box elements TFII-I interacts both physically and

functionally with USF [152].

Model for β-Globin Gene Regulation

Our lab has proposed a multi-step model for human β-globin gene regulation

(Figure 1-2) [96]. The first step towards activating the globin genes involves generating

a highly accessible LCR holocomplex. Replication may or may not be required for this

region to become DNaseI sensitive. If this step is mediated by replication, then

erythroid-specific proteins may bind to the locus after replication to prevent the formation

of inaccessible chromatin domains and perhaps to also modify histone tails. It is known

that GATA-1 is one of the earliest markers of red cell differentiation [125] and associates

with histone acetyltransferases; therefore GATA factors may be involved in the first step

of obtaining an open chromatin structure. Other proteins such as NF-E2 may then bind

the LCR HS sites and bend or disturb the DNA structure, leading to a highly accessible

region.

17

The second step is the recruitment of chromatin-remodeling and transcription

complexes to the LCR. The proteins bound at the LCR holocomplex will recruit

chromatin remodeling complexes, coactivators, and RNA polymerase II. The recruitment

of Pol II may be mediated by HLH proteins, which are known to be involved in

transcription complex formation on several TATA-less genes [145, 146]. First ‘pioneer’

RNA polymerases may associate with chromatin-remodeling activities to modify the

nucleosome structure in the locus [153]. Next, elongation incompetent polymerases may

be recruited to the LCR to be transferred to the gene promoters where they would become

phosphorylated at the C-terminal domain and be rendered elongation competent [154].

Step three is the establishment of chromatin domains permissive for transcription.

The globin locus has been separated into developmental stage-specific chromatin

domains based on changes in chromatin structure, DNaseI sensitivity, histone acetylation

patterns, and the presence of intergenic transcripts [102, 103, 155]. Intergenic transcripts

are non-coding transcripts found over the entire LCR and between the globin genes [102,

156]. Intergenic transcripts of the LCR initiate upstream of or within the LCR (for

example within HS2 [104]) and proceed towards the genes [102, 156, 157]. If the region

containing the site of initiation of adult-specific intergenic transcripts is deleted then

DNaseI sensitivity is decreased in that region and β-globin gene expression is decreased

[102]. If the before mentioned ‘pioneer’ polymerases associate with chromatin-

modifying factors then those complexes can initiate intergenic transcription and modify

the chromatin structure so as to create open chromatin domains [153].

18

Figure 1-2. Multistep model for human β-globin gene regulation. The model proposes four steps involved in the regulation of gene expression in the human β-globin locus. (A) Generation of a highly accessible LCR holocomplex. (B) Recruitment of necessary complexes. (C) Establishment of chromatin domains permissive for transcription. (D) Transfer of macromolecular protein complexes to individual globin gene promoters.

The last step is the transfer of transcription complexes to the individual globin

genes. There are two feasible models of how this is accomplished. The first is the

19

linking model, where complexes recruited to the LCR would be transferred to the genes

via proteins bound along the DNA [101, 115]. The looping model hypothesizes that the

LCR interacts with the individual genes by looping out intervening DNA [101, 115].

Recent evidence supporting the looping model within the β-globin locus and in other

parts of the genome came from results of experiments using the chromosome

conformation capture (3C) technique [117, 120, 158-162]. Results from our lab

demonstrating that Pol II, recruited to the LCR, can be loaded onto a downstream β-

globin promoter is compatible with the looping model [132].

There are possibly many factors regulating which gene promoter the LCR transfers

transcription complexes to. Due to the complexity involved in transcriptional switching,

our model does not postulate what causes the signals for switching from embryonic to

fetal and fetal to adult.

Questions to Be Addressed

The β-globin gene locus is one of the most highly studied eukaryotic gene loci.

This is in part due to the prospects of developing gene therapy strategies for sickle cell

disease and β-thalassemia. The presence of the LCR also makes the β-globin locus a

good model system for studying long-range gene regulation. Though there is a large

body of knowledge about this gene locus, there are still many unanswered questions. It is

expected that resolving these problems will contribute to successful gene therapy trials as

well as to our understanding of long-range gene regulation.

The mechanism by which the LCR activates the genes remains to be determined.

We proposed a multi-step mechanism of LCR function leading to the activation of globin

gene expression in a developmental-stage specific manner [96]. To test this model, we

20

propose to determine the order of recruitment of transcription factors, Pol II and specific

histone modification marks to the locus during transcriptional activation.

The goals of this work were all related to addressing the mechanism of LCR-

mediated activation of the β-globin genes. The first goal was to develop and optimize

methods of investigating protein-DNA interactions within the β-globin locus in vivo and

in vitro. The next goal was to use these methods to compare and contrast the proteins and

complexes interacting with the LCR, active genes, and inactive genes within the locus.

The third goal was to use synchronized erythroid cells to analyze recruitment of

transcription factors and Pol II during replication. The next goal was to use an in vitro

system to investigate the mechanism of Pol II transfer from the LCR to the β-globin gene

promoter. The last goal was to elucidate the role of USF, TFII-I and HDAC3 in the

regulation of the β-globin genes.

21

CHAPTER 2 MATERIALS AND METHODS

Cell Culture

All cell culture reagents were purchased from Cellgro. MEL cells were grown in

RPMI 1640 with L-glutamine supplemented with 10% fetal bovine serum and 1%

antibiotic-antimycotic (ABAM) in 5% CO2 at 37°C. K562 cells were obtained from

ATCC, Coriell Repositories, and DSMZ. K562 cells were grown in RPMI 1640 with L-

glutamine supplemented with 15% fetal bovine serum and 1% penicillin-streptomycin in

5% CO2 at 37°C.

Chromatin Immunoprecipitation (ChIP)

The ChIP assay was performed as described by Forsberg et al. [103] with minor

modifications. MEL and K562 cells (107 cells per antibody used) were grown in RPMI

medium. Crosslinking of proteins and DNA was induced by incubating the cells in 1%

formaldehyde with rocking at room temperature for 10 minutes. The crosslinking was

quenched with 0.125 M glycine for 5 minutes. Cells were washed twice with cold 1x

PBS with Complete protease inhibitors (Roche), resuspended in swelling buffer (5 mM

PIPES pH 8.0, 85 mM KCl, 0.5% NP-40, protease inhibitors) and incubated on ice for 10

minutes. The nuclei were pelleted by centrifugation for 5 minutes at 5000 rpm and 4°C.

Nuclei were lysed by incubation in lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris–

HCl pH 8.1, protease inhibitors) on ice for 10 minutes. DNA was sonicated in an ice

water bath using a Fisher model 100 sonicator at power 5 for 4 pulses of 10 seconds to

yield an average size of ~500 bp. After centrifugation for 10 minutes at 14000 rpm and

22

4°C, the supernatant was diluted in dilution buffer (0.01% SDS, 1.1% Triton X-100, 1.2

mM EDTA, 167 mM Tris–HCl pH 8.1, 167 mM NaCl, protease inhibitors) to a final

volume of x ml where x = # of antibody samples needed. 50 µl of Protein A–Sepharose

beads (Amersham) per 107 cells used were added to the diluted lysate and incubated for 2

hours with rotation at 4°C. The beads were pelleted and 5 µl of appropriate antibody was

added to each ml of aliquoted supernatant and incubated overnight with rotation at 4°C.

The antibodies used were TFIIB sc-225, TFIID (TBP) sc-273, Pol II (N-20) sc-899, NF-

E2 (C-19) sc-291, USF2 (C-20) sc-862 (all purchased from Santa Cruz), phospho-Pol II

05-623 and acetyl-H3 06-599 (Upstate). All antibodies were tested in Western blotting

experiments using MEL or K562 nuclear extracts as described by Leach et al [163].

Protein A–Sepharose beads were blocked by resuspending them in blocking buffer (3%

BSA, 0.05% Na azide in 10 mM Tris–HCl pH 8.1, 1 mM EDTA) and incubated overnight

with rotation at 4°C.

The next day, the samples were rotated for 2 hours at 4°C with 60 µl blocked

Protein A–Sepharose beads. The beads were pelleted and supernatants removed. 500 µl

of the supernatant of the ‘no antibody’ sample was kept and labeled ‘input’. The pelleted

beads were washed with low salt wash (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 200

mM Tris–HCl pH 8.1, 150 mM NaCl), high salt wash (0.1% SDS 1% Triton X-100, 2

mM EDTA, 200 mM Tris–HCl pH 8.1, 500 mM NaCl), LiCl wash (0.25 M LiCl, 1% NP-

40, 1% Na deoxycholate, 1 mM EDTA, 100 mM Tris–HCl pH 8.1) and two washes with

TE pH 8. DNA was eluted off the beads with two volumes of 250 µl 1% SDS and 0.1 M

NaHCO3 at 65°C for 15 minutes each. The eluates were pooled together and NaCl was

added to the eluates to a final concentration of 200 mM. Crosslinking was reversed by

23

incubation at 65°C for 4 hours. RNA was digested by incubation with 40 µg/ml RNase

cocktail for 30 minutes at 37°C. Proteins were then digested with 40 µg/ml proteinase K

in 10 mM EDTA, 40 mM Tris–HCl pH 6.5 at 37°C for 1 hour. DNA was purified using a

purification kit (Qiagen) and eluted in 100 µl TE pH 7.4. 2.5 µl DNA was used as a

template for PCR with 50 pM primers and 10 µl PCR mix (Eppendorf) in a total volume

of 25 µl. Primer pairs were used against all areas of interest (Table 2-1). Primers for the

human γ-globin genes were published by Schreiber et al [164].

Table 2-1. Names and sequences of primer pairs used for ChIP Primers Sequence Human ε-globin US 5’ GCTCCTTTATATGAGGCTTTCTTGG 3’

DS 5’ AATGCACCATGATGCCAGG 3’ Human β-globin US 5’ TATCTTAGAG GGAGGGCTGAGGGTTTG 3’

DS 5’ CCAACTTCATCCACGTTCACCTTGC 3’ Human HS2 US 5’ TGTGTGTCTCCATTAGTGACCTCCC 3’

DS 5’ TTTTGCCATCTGCCCTGTAAGC 3’ Human γ-globin US 5’ CTCAATGCAAATATCTGTCTG 3’

DS 5’ TCTGGACTAGGAGCTTATTG 3’ Human HS2 5' flank

US 5’ TGGGGACTCGAAAATCAAAG 3’ DS 5’ AGTAAGAAGCAAGGGCCACA 3’

Human Necdin US 5’ GTGTTATGTGCGTGCAAACC 3’ DS 5’ CTCTTCCCGGGTTTCTTCTC 3’

Mouse βmaj-globin US 5’ TAATTTGTCAGTAGTTTAAGGTTGC 3’ DS 5’ CAT TGTTCACAGGCAAGAGCAGG

Mouse εγ-globin US 5’ CAAAGAGAGTTTTTGTTGAAGGAGGAG 3’ DS 5’ AAAGTTCACCATGATGGCAAGTCTGG 3’

Mouse HS2 US 5’ TTCCTACACATTAACGAGCCTCTGC 3’ DS 5’AACATCTGGCCACACACCCTAAGC 3’

Mouse HS2 5'flank US 5’ CTATTTGCTAACAGTCTGACAATAGAGTAG 3’ DS 5’ GTTACATATGCAGCTAAAGCCACAAATC 3’

Mouse necdin US 5’TTTACATAAGCCTAGTGGTACCCTTCC 3’ DS 5’ATCGCTGTCCTGCATCTCACAGTCG 3’

Titration of cycle numbers was performed with the different primer pairs to

determine the range of linear amplification. After the optimal number of PCR cycles,

DNA was run in 5% TBE polyacrylamide gels. The gels were stained with SyBr-green

24

and analyzed by fluorescence scanning on a Storm scanner. Quantitations were

performed using ImageQuant.

Chromatin Immunoprecipitation and DMS Footprinting

ChIP was performed as described above with the following modifications.

Approximately 3.33 x 107 MEL cells were collected per antibody to be used.

Crosslinking was induced by adding 1% (v/v) formaldehyde and incubation for 10

minutes at room temperature on a shaker. After stopping the crosslinking reaction by

adding 0.125 M glycine and incubation for 5 minutes (with shaking at room temperature),

the cells were pelleted at 2000 rpm. The cells were then washed twice in 25 ml ice-cold

phosphate-buffered saline (PBS) including protease inhibitors. Nuclei were isolated by

resuspending the cell pellet in 1 ml ice-cold swelling buffer (5 mM PIPES pH 8.0, 85 mM

KCl, 0.5% NP-40, and protease inhibitors), split into two aliquots and incubated on ice

for 10 minutes.

Chromatin was fragmented by subjecting the nuclei to restriction enzyme digestion

with 200 U PstI for 4 hours at 37°C and 100 U PstI for an additional 16 hours at 37°C.

The nuclei were then incubated with 200 U RNase cocktail (Ambion) and an additional

100 U aliquot of PstI for 2 hours at 37°C. Nuclei were pelleted at 4°C for 5 minutes at

5000 rpm and lysed in 1 ml lysis buffer (1% SDS, 10 mM EDTA and 50 mM Tris–HCl

pH 8.1, protease inhibitors) on ice for 20 minutes. Restriction enzyme digestion was

verified by electrophoresis on a 2% agarose gel followed by Southern blotting using a

radioactive probe hybridizing to the human β-globin gene.

The lysate was combined and transferred to a 15 ml conical tube and diluted with 9

ml dilution buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris–HCl

25

pH 8.1, 167 mM NaCl and protease inhibitors). An aliquot of 500 µl protein A–

Sepharose beads (Amersham) was added to the diluted nuclear lysate and incubated for 2

hours at 4°C while rotating. The beads were pelleted for 10 minutes at 2000 rpm and the

supernatant was divided into three aliquots. An aliquot of 25 µl of the appropriate

antibody (USF1 sc-229 or NF-E2 sc-291; Santa Cruz Biotechnology) or no antibody was

added to the aliquoted supernatant and incubated at 4°C overnight while rotating. Protein

A–Sepharose beads were blocked overnight rotating at 4°C in a 1:1 ratio of beads to

blocking buffer [3% BSA and 0.05% sodium azide in 1x TE (10 mM Tris–HCl pH 8.1, 1

mM EDTA)].

The chromatin was then immunoprecipitated with 600 µl blocked protein A–

Sepharose beads for 2 hours at 4°C on a rotator. The immunoprecipitates were pelleted at

13000 rpm for 30 seconds and 1 ml of the no antibody supernatant was saved and labeled

as ‘input’. Half of the input chromatin was ethanol precipitated and resuspended in two

aliquots of 20 µl ddH2O and 800 µl DMS buffer (50 mM sodium cacodylate, 1 mM

EDTA) and the other half was saved for the ChIP PCR analysis. The supernatants of the

samples precipitated with USF1 and NF-E2 antibodies were discarded and the pellets

were washed by rotating at 4°C for 5 minutes with 1 ml each of low salt wash (0.1% SDS,

1% Triton X-100, 2 mM EDTA, 20 mM Tris–HCl pH 8.1, 150 mM NaCl), high salt wash

(0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris–HCl pH 8.1, 500 mM NaCl),

LiCl wash (0.25 M LiCl, 1% NP-40, 1% sodium deoxycholate, 1 mM EDTA, 10 mM

Tris–HCl pH 8.1) and twice with 1x TE pH 8. Of the immunoprecipitates, 80% was

resuspended in 800 µl DMS buffer and 20% was left in TE buffer for the ChIP/PCR

analysis.

26

DMS Treatment

DMS treatment of the immunoprecipitated chromatin was performed using the

Maxam and Gilbert guanine-specific sequencing reaction with 0.1% DMS for 15, 45 or

90 seconds at room temperature [165]. The reaction was stopped by adding 50 µl DMS

stop buffer (1.5 M sodium acetate pH 7.0, 1 M 2-mercaptoethanol), followed by two

ethanol precipitations in a dry ice bath. The DMS-treated and non-DMS-treated

chromatin was then eluted from the beads by incubating twice with 250 µl elution buffer

(1% SDS, 0.1 M NaHCO3), shaking at 950 rpm for 15 minutes at 65°C, each time saving

the supernatant. An aliquot of 200 mM NaCl was added to the eluates and crosslinking

was reversed by incubation at 65°C for 5 hours. Proteins were digested with 40 µg/ml

proteinase K in 10 mM EDTA and 40 mM Tris pH 6.5 for 1 hour at 37°C.

Immunoprecipitated DNA was purified using a Qiagen kit and eluted with 180 µl ddH2O.

To cleave the DMS-treated DNA, 20 µl piperidine were added and incubated at 95°C for

30 minutes. The DNA was washed twice by adding 1 ml ddH2O, dried in a Speed Vac

and resuspended in 50 µl 1x TE. Of the DMS-treated immunoprecipitated DNA, 10%

was used for ligation-mediated PCR (LMPCR)-assisted in vivo footprinting. An aliquot

of the precipitated DNA was also analyzed by PCR using primers specific for the murine

β-globin downstream promoter region (forward primer, 5'-

GACAAACATTATTCAGAGGGAGTACCC; reverse primer, 5'-

AGGTGCACCATGATGTCTGTTTCTGG) using a protocol previously published by

Forsberg et al [131].

Linker LMPCR

LMPCR was essentially performed as described by Hornstra and Yang [166] with

the following modifications. Approximately 2 µg DMS-treated genomic DNA or 10%

27

immunoprecipitated DNA was annealed to 0.6 pmol gene-specific primer (MPA 1) by

denaturing at 96°C for 10 minutes followed by annealing at 47°C for 30 minutes in a 15

µl solution of 1x Vent buffer (10 mM Tris–HCl pH 8.9 and 40 mM NaCl). Primer

extension was performed by adding a 15 µl solution of 10 mM Tris–HCl pH 8.9, 40 mM

NaCl, 0.5 mM dNTPs and 2 U Vent polymerase (New England Biolabs) and incubating

at 53°C for 1 minute, 55°C for 1 minute, 57°C for 1 minute, 60°C for 1 minute, 62°C for

1 minute, 66°C for 1 minute, 68°C for 3 minutes and 76°C for 3 minutes. A 20 µl

dilution solution (110 mM Tris–HCl pH 7.5, 18 mM MgCl2, 50 mM DTT, 0.0125%

BSA) was then added to the extension reaction.

Blunt-end ligation was performed by adding a 25 µl solution of 100 pmol

asymmetric double-stranded linker, 10 mM MgCl2, 20 mM DTT, 3 mM ATP, 0.005%

BSA and 4.5 U T4 ligase (Ambion) to the reaction and incubating at 17°C overnight. The

ligation products were purified by standard phenol/chloroform extractions and ethanol

precipitation including 10 µg/µl tRNA. The ligated DNA was resuspended in 20 µl

ddH2O and added to 80 µl PCR mix [10 mM Tris–HCl pH 8.9, 40 mM NaCl, 3 mM

MgCl2, 0.25 mM dNTPs, 20 pmol gene-specific PCR primer (MPA 3), 20 pmol linker

primer 2 and 0.5 U Taq polymerase; Gibco BRL]. This PCR mixture was initially

denatured at 95°C for 5 minutes and then subjected to 20 cycles of PCR under the

following conditions: 95°C for 20 seconds, 65°C for 1 minute, 72°C for 1 minute with an

increase of 5 seconds/cycle and an additional 5 cycles of 95°C for 20 seconds, 65°C for 1

minute, 72°C for 2 minutes 30 seconds, followed by a final extension at 72°C for 15

minutes. The PCR products were purified by phenol/chloroform extraction and ethanol

precipitation and resuspended in 30 µl ddH2O. Initially, 3 µl of the PCR products were

28

size-fractionated on a 0.4 mm thick 5% polyacrylamide gel made with 8 µg/ml

ammonium persulfate, then electrotransfered for 30 minutes to a nylon membrane

(Hybond N+; Amersham).

Radiolabeled probes were synthesized using the Prime-a-Probe kit (Ambion) from a

gel-purified PCR product (PCR primers: β-maj FWD, 5'-

GACAAACATTATTCAGAGGGAGTACCC-3'; MPB 1, 5'-

TCTGTCTCCAAGCACCCAA-3') containing the region of interest. To radiolabel the

probe, 150 ng template DNA was mixed with 0.3 µg gene-specific primer (MPA 3), used

in the PCR step of the LMPCR protocol, and brought up to 8 µl in ddH2O. This primer

plus template mixture was denatured at 95°C for 10 minutes and immediately placed on

dry ice to snap freeze. Then 5 µl of 5x Decaprime buffer containing dATP, dGTP and

dTTP (Ambion) was incubated with 10 µl [α-32P] dCTP (3000 Ci/mmol) and 1 U/µl

Klenow fragment at 37°C for 30 minutes. The reaction was quenched on ice and stopped

by adding 35 µl formamide loading dye.

The probe was denatured at 95°C for 10 minutes and purified on a 5% denaturing

polyacrylamide gel. After exposing the gel to film (Type 57; Polaroid) the probe was cut

out of the gel, crushed and soaked in 4 ml hybridization buffer (250 mM Na2PO4 pH 7.2,

7% SDS, 1% BSA). The probe was hybridized to the blots at 65°C overnight. The blots

were washed three times at 65°C in washing solution (20 mM Na2PO4 pH 7.2, 1% SDS)

and visualized by autoradiography. The following primers were used for LMPCR: MPA

1, 5'-ATGTCCAGGGAGAAATATCG-3'; MPA 3, 5'-

TGAAGGGCCAATCTGCTCACACAGG-3'.

29

Western Blotting

To 19 parts Laemmli sample buffer (Bio-Rad) was added 1 part β-mercaptoethanol.

2 volumes of this buffer were added to each nuclear protein extract or whole cell extract

and the mixture heated at 95°C for 10 minutes. 25 µl of each sample was loaded on a

10% Tris-HCl Ready Gel (Bio-Rad) and run at 150V in cold tank buffer (25 mM Tris, 192

mM glycine, 0.1% SDS) until the dye reached the bottom. The gel, membrane (Bio-Rad),

filter paper (Bio-Rad), and fiber pads (Bio-Rad Mini Protean apparatus) were soaked in

cold transfer buffer (10 mM NaHCO3, 3 mM NaCO3, and 20% methanol, pH 9.9) for 15

minutes and then set up in a transfer sandwich with a fiber pad, filter paper, the gel,

membrane, filter paper, and fiber pad. The transfer was run overnight at 30V at 4°C with

constant stirring in the Mini Protean apparatus (Bio-Rad).

The transfer was disassembled and the membrane was stained with Fast Green for 5

minutes and washed twice in destain solution (50% methanol, 10% acetic acid). It was

then rinsed in TBS-Tween (30 mM Tris, 150 mM NaCl, 0.1% Tween 20, pH 7.6) and

blocked 1 hour in 5% milk in TBS-Tween shaking at room temperature. The membrane

was rinsed twice in TBS-Tween, washed once in TBS-Tween for 15 minutes while

shaking at room temperature, and washed twice for 5 minutes in TBS-Tween while

shaking at room temperature. It was incubated 1 hour in 10 ml 5% milk in TBS-Tween

containing 1:500 anti-HDAC3 while shaking at room temperature. The membrane was

rinsed twice in TBS-Tween, washed once in TBS-Tween for 15 minutes while shaking at

room temperature, and washed twice for 5 minutes in TBS-Tween while shaking at room

temperature. It was incubated 1 hour in 15 ml 5% milk in TBS-Tween containing

1:50000 anti-rabbit while shaking at room temperature. The membrane was rinsed twice

30

in TBS-Tween, washed once in TBS-Tween for 15 minutes while shaking at room

temperature, and washed twice for 5 minutes in TBS-Tween while shaking at room

temperature. ECL reagents (Amersham) were mixed according to the manufacturer’s

recommendations and added to the membrane. The membrane was then exposed to

Kodak MS film for varying times.

Cell Synchronization

K562 cells were synchronized at the border of G1 and S phase by incubating the

cells with complete growth medium in the presence of 2 mM thymidine for 17 hours,

washing twice with complete medium, incubating in complete medium for 12 hours and

blocking the cells in complete medium with 2 mM thymidine for another 17 hours [167,

168]. Blocked cells were fixed with ethanol, digested with RNase, stained with

propidium iodide and subjected to flow cytometry to verify synchronization. Blocked

cells were taken as “time 0” and subjected to ChIP and PCR analysis. The blocked cells

were then washed twice with complete medium to release them from the block and

allowed to grow in complete medium. Cells were taken for flow cytometry, ChIP, and

PCR analysis at specific time points after release from the arrest.

PCR-Based DNA Replication Analysis

DNA Isolation

Added to lysis solution (50 mM KCl, 10 mM Tris-HCl pH 8, 1.25 mM MgCl2,

0.45% NP-40, 0.45% Tween 20) was 75 µl of 10mg/ml proteinase K per ml of lysis

solution (added fresh every time). Synchronized cells were counted and 106 cells were

collected, washed with 1x PBS and resuspended in 100 µl lysis solution per 105 cells.

The cells were incubated in the lysis solution overnight. The next day, 500 µl phenol was

31

added and vortexed for 1 minute. The reaction was centrifuged for 5 minutes at 14000

rpm and room temperature. Then 500 µl phenol/chloroform/isoamyl alcohol was added

to the supernatant and it was vortexed for 1 minute. The samples were centrifuged for 5

minutes at 14000 rpm and room temperature. Then, 500 µl chloroform/isoamyl alcohol

was added to the supernatant, the reaction was vortexed for 1 minute and centrifuged for

5 minutes at 14000 rpm and room temperature. Next 100% ethanol was added to the

supernatant, incubated for 15 minutes at room temperature, and centrifuged 30 minutes at

14000 rpm and room temperature. The supernatant was discarded and 500 µl of 70%

ethanol was added to the pellet. The sample was centrifuged for 10 minutes at 14000

rpm and room temperature. The supernatant was removed and the pellet allowed to air

dry. The pellet was then resuspended in 50 µl TE pH 7.4. 1 µl RNase was added and it

was incubated for 30 minutes in a 37°C water bath. The DNA was used for PCR and the

rest stored at –20°C.

Semi-Quantitative PCR

2.5 µl DNA was used as a template for PCR with 50 pM primers and 10 µl PCR

mix (Eppendorf) in a total volume of 25 µl. Primer pairs were used against the human ε-

globin, human necdin, mouse βmaj-globin and mouse necdin, as described in Table 2-1.

PCR was performed with cycle numbers 18, 20, 22, 24, 26, 28, and 30. DNA was run in

5% TBE polyacrylamide gels. The gels were stained with SyBr-green and analyzed by

fluorescence scanning on a Storm scanner. Quantitations were performed using

ImageQuant. The ratios of human ε-globin to human necdin and mouse βmaj-globin to

mouse necdin were calculated for samples from all time points at a cycle number within

the linear range.

32

In Vitro Polymerase Transfer Analysis

A plasmid containing the wild-type human β-globin LCR was linearized and

immobilized on streptavidin coated magnetic beads as described by Leach et al [169].

The immobilized LCR (200 ng) was incubated for 30 minutes at 30°C with 50 µg MEL

nuclear protein extract (10 µl) and 15 µl 2x binding buffer (36 mM HEPES, pH 7.9, 160

mM KCl, 40 mM MgCl2, 4 mM DTT, 10 mM PMSF, 20% glycerol) in a total volume of

100 µl. The tubes were placed on a magnet, the supernatant removed, and the beads were

washed three times with 1x binding buffer. The beads were then resuspended in 1x

binding buffer and incubated for 10 minutes at 30°C in the presence or absence of 50 ng

of the plasmid pRSβA/X containing the wild-type or mutant β-globin gene [163]. The

supernatants were removed and samples containing the wild type or mutant β-globin gene

were crosslinked by incubation for 10 minutes in 0.5 % formaldehyde at RT. The

crosslinking reaction was stopped by the addition of 0.125 M glycine and incubation for

5 minutes at RT. The supernatant of reactions not containing the β-globin gene were

incubated for 10 minutes at 30°C in the presence of pRSβA/X, after which these samples

were crosslinked as well. All samples were dialyzed against ChIP dilution buffer and

subjected to immunoprecipitation using Pol II specific antibodies and PCR analysis as

described by Leach et al [163]. In some experiments recombinant NF-E2 [163] or BSA

(0.3 µg/µl) was added to the transfer reactions. The plasmid βmaxi was used as a positive

control for the PCR and was generated by ligating an 800 bp Pml1 restriction fragment

from the human LCR into the Pml1 site of pRSβA/X. The β-globin PCR primers span the

Pml1 site present in the β-globin gene thus generating a PCR product that is 800 bp larger

than the PCR product from the wild type β-globin gene. The TK-HGH plasmid contains

33

the human growth hormone gene under the control of the Herpes Simplex Virus

thymidine kinase promoter [163]. The HS2 plasmid contains the HS2 core plus flanking

sequence as previously described [122]. The following primers were used in these

experiments: β-globin US: 5’ CCTGAGGAGAAGTCTGCCGTTACTG 3’ and DS: 5’

TCCTATGACATGAACTTAACCATAG 3’; TK-HGH: US: 5’

GGAGGCTGGAAGATGGCA 3’; DS: 5’ AGTAGTGCGTCATCGTTGTGTG 3’;

human HS2: as described in Table 2-1.

MEL Cell DMSO Differentiation

MEL cells were incubated in RPMI 1640 with L-glutamine supplemented with

10% fetal bovine serum, 1% antibiotic-antimycotic (ABAM) and 1% DMSO in 5% CO2

at 37°C for three days. RNA was isolated from the cells for semi-quantitative RT-PCR.

Semi-Quantitative RT-PCR

RNA was isolated for RT-PCR using an RNA Isolation Kit (Gentra) according to

the manufacturer’s protocol. Reverse Transcription was performed using 200 to 250 ng

RNA and the first strand cDNA synthesis Kit (Bio-Rad) as described by the

manufacturers protocol. PCR amplification was performed using PCR Mastermix from

Eppendorf and primer sequences specific for the murine β-globin and β-actin gene with

15 and 16 cycles as indicated. The mouse β-actin primers used were: US 5’

GGACGACATGGAGAAGAT 3’ and DS 5’ ATCTCCTGCTCGAAGTCT 3’. The

mouse β-globin primers used were: US 5’ AAAGGTGAACTCCGATGAAGTTGG 3’

and DS 5’ TTCTGGAAGGCAGCCTGTGC 3’. After electrophoresis the gels were

stained with SyBr Green, scanned using a storm scanner, and quantitated with Image

Quant.

34

Double Immunoprecipitation

Approximately 2 x 107 K562 cells were collected per immunoprecipitate reaction.

The cells were incubated in 1% formaldehyde with rocking at room temperature for 10

minutes to induce crosslinking. Crosslinking was quenched with 0.125 M glycine for 5

minutes shaking at room temperature. Cells were washed twice with cold 1x PBS with

Complete Protease Inhibitors (Roche), resuspended in swelling buffer (5 mM PIPES pH

8.0, 85 mM KCl, 0.5% NP-40, protease inhibitors), and incubated on ice for 10 minutes.

Nuclei were pelleted by centrifugation for 5 minutes at 5000 rpm and 4°C. Nuclei were

lysed by resuspending in lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris–HCl pH 8.1,

protease inhibitors) and incubating on ice for 10 minutes. DNA was sonicated in an ice

water bath using a Fisher model 100 sonicator at power 5 with 5 pulses of 10 seconds

each with 1 minute cooling in between, to yield an average size of ~500 bp. The sample

was then centrifuged for 10 minutes at 14000 rpm and 4°C. The supernatant was diluted

in dilution buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 167 mM Tris–HCl

pH 8.1, 167 mM NaCl, protease inhibitors) to a final volume of x ml where x = # of

antibody samples needed. Then 50 µl of Protein A–Sepharose beads (Amersham) per 2 x

107 cells used were added to the diluted lysate and incubated for 2 hours with rotation at

4°C. The beads were pelleted and 5 µl of appropriate antibody was added to each ml of

aliquoted supernatant and incubated overnight with rotation at 4°C. The antibodies used

were

TFII-I (a gift from R. Roeder, Rockefeller University) and HDAC3 (a gift from E. Seto,

University of South Florida). Protein A–Sepharose beads were blocked by resuspension

35

in blocking buffer (3% BSA, 0.05% Na azide in 10 mM Tris–HCl pH 8.1, 1 mM EDTA)

and incubated overnight with rotation at 4°C.


Protein A–Sepharose beads. The beads were pelleted and supernatants removed. Then

500 µl of the supernatant of the ‘no antibody’ sample was kept and labeled ‘input’. The

pelleted beads were washed with low salt wash (0.1% SDS, 1% Triton X-100, 2 mM

EDTA, 200 mM Tris–HCl pH 8.1, 150 mM NaCl), high salt wash (0.1% SDS 1% Triton

X-100, 2 mM EDTA, 200 mM Tris–HCl pH 8.1, 500 mM NaCl), LiCl wash (0.25 M

LiCl, 1% NP-40, 1% Na deoxycholate, 1 mM EDTA, 100 mM Tris–HCl pH 8.1) and two

washes with TE pH 8. DNA was eluted off the beads with two 250 µl 1% SDS and 0.1 M

NaHCO3 washes at 65°C for 15 minutes each shaking at 950 rpm. The two elutions were

pooled together and then dialyzed against 200 ml of dilution buffer with Complete Mini

protease inhibitors using Slide-A-Lyzer Mini Dialysis units (Pierce) at 4°C with constant

stirring for 2 hours. After samples were taken out of dialysis units and placed in 1.5 ml

tubes, 500 µl dilution buffer was added to each. For the second immunoprecipitation 5 µl

of antibody was added to each and incubated overnight at 4°C rotating. Protein A–

Sepharose beads were blocked by resuspending them in blocking buffer and incubated

overnight with rotation at 4°C.


Protein A–Sepharose beads. The beads were pelleted and supernatants removed. The

pelleted beads were washed with low salt wash, high salt wash, LiCl wash and two


NaHCO3 washes shaking at 65°C for 15 minutes each. The two elutions were pooled

36

together and NaCl was added to the eluates to a final concentration of 200 mM and

crosslinking was reversed by incubation at 65°C for 4 hours. RNA was digested by

incubation with 40 µg/ml RNase cocktail (Ambion) for 30 minutes at 37°C. Proteins

were then digested with 40 µg/ml proteinase K in 10 mM EDTA, 40 mM Tris–HCl pH

6.5 at 37°C for 1 hour. DNA was purified using a column purification kit (Qiagen) and

eluted in 100 µl TE pH 7.4. Then 2.5 µl DNA was used as a template for PCR with 50

pM primers and 10 µl PCR mix (Eppendorf) in a total volume of 25 µl. The human β-

globin, ε-globin and HS2&3 linker primer pairs were used for PCR (see Table 2-1). PCR

was performed for 28 cycles, and DNA was run in a 5% TBE polyacrylamide gel. The

gels were stained with SyBr-green and analyzed by fluorescence scanning on a Storm

scanner.

Co-Immunoprecipitation

107 K562 cells were collected per immunoprecipitate reaction. Cells were washed

twice with cold 1x PBS with Complete protease inhibitors (Roche), resuspended in

swelling buffer (5 mM PIPES pH 8.0, 85 mM KCl, 0.5% NP-40, protease inhibitors) and

incubated on ice for 10 minutes. Nuclei were pelleted by centrifugation for 5 minutes at

5000 rpm and 4°C. Nuclei were lysed by resuspending in lysis buffer (1% SDS, 10 mM

EDTA, 50 mM Tris–HCl pH 8.1, protease inhibitors) and incubating on ice for 10

minutes. DNA was sonicated in an ice water bath with a Fisher model 100 sonicator at

power 5 with 5 pulses of 10 seconds each with 1 minute cooling in between, to yield an

average size of ~500 bp. The samples were then centrifuged for 10 minutes at 14000 rpm

and 4°C. The supernatant was diluted in dilution buffer (0.01% SDS, 1.1% Triton X-100,

1.2 mM EDTA, 167 mM Tris–HCl pH 8.1, 167 mM NaCl, protease inhibitors) to a final

37

volume of x ml where x = # of antibody samples needed. 50 µl of Protein A–Sepharose

beads (Amersham) per 107 cells used were added to the diluted lysate and incubated for 2

hours with rotation at 4°C. The beads were pelleted and 5 µl of appropriate antibody was

added to the aliquoted supernatant and incubated overnight with rotation at 4°C. The

antibodies used were USF1 sc-229, USF2 sc-862, Pol II sc-899 (all from Santa Cruz)

TFII-I (a gift from R. Roeder, Rockefeller University) and HDAC3 (a gift from E. Seto,

University of South Florida). Protein A–Sepharose beads were blocked by resuspension

in blocking buffer (3% BSA, 0.05% Na azide in 10 mM Tris–HCl pH 8.1, 1 mM EDTA)

and incubation overnight with rotation at 4°C.


Protein A–Sepharose beads. The beads were pelleted and supernatants removed. The

pelleted beads were washed with low salt wash (0.1% SDS, 1% Triton X-100, 2 mM

EDTA, 200 mM Tris–HCl pH 8.1, 150 mM NaCl), high salt wash (0.1% SDS 1% Triton

X-100, 2 mM EDTA, 200 mM Tris–HCl pH 8.1, 500 mM NaCl), LiCl wash (0.25 M

LiCl, 1% NP-40, 1% Na deoxycholate, 1 mM EDTA, 100 mM Tris–HCl pH 8.1) and two


NaHCO3 washes shaking at 65°C for 15 minutes each. The two elutions were pooled

together and concentrated in the Speed Vac for about 1 hour. The samples were then

subjected to Western blotting procedures as described above.

38

CHAPTER 3 CHROMATIN IMMUNOPRECIPITATION AND DMS FOOTPRINTING

Introduction

Transcription in eukaryotes is a complex process involving the binding of proteins

to the promoter, recruitment of transcription complexes, initiation, elongation and

termination [170]. This process is dynamic and controlled at each step by proteins that

bind to the DNA, to the RNA polymerase holocomplex, or both. Although knowledge

about transcription complex formation in vitro is extensive, the mechanisms leading to

transcription in the context of higher order chromatin in vivo are not understood in detail.

Many nuclear processes including transcription, DNA replication, homologous

recombination, DNA repair, and others are regulated by proteins that interact with

specific DNA sequences or structures in the context of chromatin. Therefore, there is a

great demand for new and better techniques to investigate protein-DNA interactions. A

number of techniques are available to study the interaction of proteins with specific

segments of DNA in vivo; each of these techniques has distinct limitations [166, 171-

174].

Chromatin immunoprecipitation (ChIP) is a powerful method to analyze the

interaction of proteins with specific chromatin regions in vivo [171, 172]. This technique

involves crosslinking protein-DNA and protein-protein interactions in live cells, followed

by immunoprecipitation of the lysed nuclear contents to isolate segments of DNA that are

bound to the protein of interest in vivo. Combined with PCR, the ChIP assay is very

sensitive in detecting proteins that crosslink to a specific region in chromatin. There are

39

at least two limitations of the ChIP assay. First, ChIP does not provide information

regarding the specific sequence a protein interacts with in vivo. The size of the ChIP

fragments is on average 500bp, so the corresponding binding site for a specific protein

can be within a region of up to 1 kb of DNA. Second, ChIP does not reveal whether a

protein directly binds to the DNA or is recruited to DNA by protein/protein interactions.

Another method commonly used to examine the interaction of proteins and DNA is

in vivo footprinting [166, 173, 174]. In footprinting experiments enzymes like DNaseI or

chemicals like DMS/piperidine are used to cleave the DNA. Nucleotides that are bound

by proteins are normally protected from cleavage. The cleavage pattern can be visualized

either directly or after PCR using electrophoresis in combination with radioactive

labeling techniques and autoradiography. The limitation of in vivo footprinting is that it

does not reveal what the protein is that is causing the protection of particular bases.

Another problem with the in vivo footprinting technique is that it fails to provide clear

results in situations where a protein is bound in a sequence-specific manner in only a

small fraction of cells, in other words, if the protein-DNA interaction is transient.

We have combined the ChIP and in vivo footprinting techniques and developed a

novel method for analyzing protein–DNA interactions in vivo. This new technique is

similar in concept to an in vitro protocol developed by Gallarda et al. [175]. These

authors used a monoclonal antibody-based DNaseI footprint selection technique, which

unambiguously identifies proteins responsible for particular footprints.

The technique we have developed here is aimed at analyzing sequence-specific

interactions of particular proteins in vivo. In the process of developing and optimizing the

technique, we have analyzed the interaction of proteins with the mouse β-globin

40

downstream promoter region in unsynchronized MEL (mouse erythroleukemia) cells.

We show that chromatin fragments precipitated with antibodies against USF1 or NF-E2

reveal strikingly different footprint patterns on a specific β-globin gene promoter region.

The USF1-selected templates show no footprint in this region, but the NF-E2-selected

fragments show a footprint at a non-consensus MARE. This result indicates that this

novel method will allow investigators to select specific templates for in vivo footprinting,

and determine exactly where their protein of interest binds.

Results

The developmental stage-specific expression of the human β-globin gene is

regulated primarily by transcription factors that interact with gene-proximal cis DNA

elements [95]. We have shown that USF1 and USF2 interact with a downstream promoter

element of the human β-globin gene in vitro [163]. In the same study we have shown that

these proteins are crosslinked to the β-globin gene in erythroid cells in vivo. In addition,

data published by Sawado et al. [176], as well as our own experiments revealed that NF-

E2 can be crosslinked to the β-globin gene in vivo. The crosslinking of NF-E2 to the

globin promoter is somewhat surprising as there is no consensus DNA-binding site in the

globin gene. However, a sequence located downstream of the transcription start site

reveals partial homology to the NF-E2 consensus element, also referred to as the MARE

(maf recognition element, Figure 3-1; [126, 177, 178]. The NF-E2 consensus sequence is

5'-TGCTGASTCAY-3' (S = G or C; Y = T or C; [178]). The sequence in the mouse β-

globin downstream promoter region differs in one position (the third) from the consensus

sequence. Conventional in vivo footprinting did not reveal significant protection of the

NF-E2-binding site in MEL cells (Figure 3-4, lanes 1 and 2). To analyze the possibility

41

that NF-E2 interacts with the β-globin downstream element in vivo in only a fraction of

erythroid cells, we have developed a novel method that combines the ChIP assay with

DMS footprinting.

Figure 3-1. Sequence alignment of the adult β-globin downstream promoter region. Shown are three sequences of the β-globin downstream promoter region from human (H), mouse (M) and rabbit (R) [177]. The shaded box highlights the position of the MARE/Ap1-like sequence and the arrow points to the transcription start site.

The procedure (outlined in Figure 3-2) involves the crosslinking of proteins and

DNA using formaldehyde (1%), followed by digestion of permeabilized nuclei with

restriction enzymes, precipitation with specific antibodies, treatment and cleavage of

precipitated chromatin restriction fragments with DMS and piperidine and analysis of the

cleavage pattern by LMPCR (for details see Materials and Methods Chapter).

Figure 3-3 shows that antibodies against USF1 and NF-E2 precipitate crosslinked

chromatin fragments containing the murine adult β-globin gene, as expected from

previous data [163, 176].

The subsequent DMS footprint of the precipitated chromatin fragments revealed

interesting characteristics of the precipitated fragments (Figure 3-4). First, the overall

footprint pattern of fragments selected with USF antibodies is different from those

precipitated with NF-E2 antibodies (compare Figure 3-4, lanes 3 and 4). This is

42

particularly obvious over the MARE-like sequence, which shows clear protection only in

templates selected by NF-E2 precipitation.

Figure 3-2. Analysis of protein–DNA interactions in the murine β-globin downstream promoter region by a combination of chromatin immunoprecipitation and DMS footprinting. Diagram outlining the experimental procedure for footprinting ChIP-selected templates.

The difference in the cleavage pattern between the templates selected by USF or

NF-E2 antibodies could reflect the possibility that the proteins interact with the globin

gene at different stages of the transcription cycle. Alternatively, the results could reflect

the possibility that low occupancy of these sites would strongly reduce the probability of

selecting templates which have both proteins bound simultaneously. Second, the overall

footprint pattern of fragments precipitated with USF antibodies is similar to the pattern

found in unselected templates, whereas the footprint pattern for NF-E2-selected

fragments is different. This result suggests that only a small fraction of cells may have

NF-E2 bound at the β-globin gene.

43

Figure 3-3. ChIP experiment showing the interaction of USF1 and NF-E2 with the murine βmaj-globin promoter in MEL cells. MEL cells were grown under standard conditions and crosslinked with formaldehyde. After fragmentation, the chromatin was precipitated with no antibodies (lane 2), antibodies against USF1 (αUSF, lane 3) or antibodies against the p45 subunit of NF-E2 (αNF-E2, lane 4). Lane 1 shows the PCR result of the input, which serves as a positive control.

It is important to note that formaldehyde crosslinking does not appear to change the

cleavage pattern in the DNA region analyzed in these experiments; the overall pattern is

similar between untreated cells (Figure 3-4, lanes 1 and 2) and crosslinked cells (lanes 3

and 4). In addition, we found that incubation of genomic DNA with 1% formaldehyde

(with subsequent reversal of the crosslink) does not change the overall DMS cleavage

pattern in the DNA region that we have analyzed by LMPCR (data not shown).

Discussion

We examined the interaction of NF-E2 with the non-consensus MARE-binding site

in the downstream promoter region in detail using a novel in vivo method. The potential

sequence-specific interaction of NF-E2 with the β-globin downstream promoter region is

interesting in light of the fact that Johnson et al. [179] previously showed that RNA

polymerase II is recruited to both the LCR and to the adult β-globin gene and that the

transfer of the polymerase from the LCR to the globin gene depends on the presence of

44

NF-E2 (p45). In this respect it is possible that NF-E2 is part of the recruitment process

that mediates the interaction of RNA polymerase II with the β-globin promoter.

Figure 3-4. LMPCR footprint analysis of non-selected or ChIP-selected chromatin.

Chromatin was precipitated with USF or NF-E2 antibodies. The precipitates were subjected to DMS footprinting (lane 3, USF-selected chromatin; lane 4, NF-E2-selected chromatin). In vivo footprinting was also performed on MEL cells that were not treated with formaldehyde (lane 2). Lane 1 shows the DMS/piperidine cleavage pattern in in vitro treated MEL genomic DNA. Lane 5 shows the G-ladder of the β-globin downstream promoter region. Indicated on the right are the positions of the initiator and MARE-like sequences, as well as the start site and direction of transcription. Open circles indicate protected G residues in NF-E2-selected chromatin.

Several methods are available to fragment crosslinked chromatin for ChIP,

including sonication, MNase digestion and restriction enzyme digestion [171, 172, 180,

181]. We reasoned that sonication or MNase digestion might fragment the chromatin in a

45

way that not all of the resulting templates would be amplifiable by LMPCR and in

addition may result in a high background. We therefore chose to fragment the

crosslinked chromatin by restriction enzyme digestion [180]. The disadvantage in using

this method is that long incubation with the restriction enzyme allows endogenous

nucleases to attack the chromatin in accessible regions. In some of our experiments, we

observed inconsistent LMPCR banding patterns in ChIP-selected templates in the higher

molecular weight ranges (data not shown). This problem can be solved by incubating the

nuclei for shorter periods of time with higher concentrations of restriction enzyme [181]

or by following similar restriction digest protocols from other groups [182].

Another issue that has to be considered is the fact that the precipitation of

templates using ChIP results in different concentrations of templates. It is thus important

to perform a titration of the templates precipitated with different antibodies. Similarly,

we were unsure whether the steps of the ChIP assay leave the protein in a native structure

to give a true footprint in all cases tested. Though it could be postulated, that once a

footprint such as NF-E2 at an NF-E2 binding site has been observed that a similar

treatment will leave other proteins in a conformation that makes a footprint detectable.

Lastly, it was suggested that the antibody might remain bound and contribute to the

footprint observed. Therefore, to remove this possibility it would be possible to change

the elution buffer to one that is known for disrupting the antibody-antigen interactions,

such as the ImmunoPure Gentle Ag/Ab Elution Buffer available from Pierce

Biotechnology.

46

CHAPTER 4 INTERACTION OF RNA POLYMERASE II AND TRANSCRIPTION FACTORS

WITH THE β-GLOBIN LOCUS

Introduction

The five genes of the human β-globin locus are expressed in erythroid cells in a

tissue- and developmental-stage specific manner [183]. Appropriate expression of the

globin genes is regulated by many DNA elements that are located proximal or distal to

the genes. The human β-globin locus control region (LCR) is a powerful regulatory

DNA element located far upstream of the genes and required for high-level expression of

all the globin genes throughout development [136, 183]. The LCR, unlike classical

enhancer elements, operates in an orientation dependent manner [114]. There is currently

no consensus on how the LCR acts to stimulate globin gene transcription but it is

generally believed that it involves some form of communication between the LCR and

the globin genes [101, 115, 184]. The LCR is composed of several regions that exhibit

extremely high sensitivity to DNaseI in erythroid cells (hypersensitive HS sites 1 to 5).

The core HS sites contain clusters of transcription factor binding sites and are separated

from each other by 2 to 4 kbp of DNA [136]. Results from analyzing human LCR

function at ectopic sites in the context of transgenic mice suggests that the HS sites

synergistically enhance globin gene transcription [19, 20, 119, 121-123], whereas studies

in the endogenous murine locus show that the core HS sites function additively [109, 185,

186].

47

Recent models view the LCR as a holocomplex in which the individual HS sites

interact via extensive protein-DNA and protein-protein interactions [118, 119, 134]. The

LCR holocomplex may provide a highly accessible region for the efficient recruitment of

macromolecular complexes involved in chromatin modification and transcription [96].

Indeed, it has been shown that RNA polymerase II (Pol II) transcription complexes are

recruited to LCR HS sites in vitro and in vivo [169, 187-189], suggesting that

transcription complexes are first recruited to the LCR and subsequently transferred to the

globin genes [96]. Sawado et al. [190] recently demonstrated that another important

function of the LCR is to regulate transcription elongation at the adult β-globin gene.

An interesting aspect of gene regulation is the question of how transcriptional

activity is maintained during the process of replication [191]. Activation of at least some

gene loci depends on replication, and it is possible that replication could provide a

window of opportunity for transcription factors to gain access to regulatory sequences

before repressive chromatin is formed.

Generally, transcriptionally active regions of the genome replicate early, whereas

repressed parts of the genome replicate late. A recent study showed that plasmids

injected into cells at different time points during replication adopt an open chromatin

conformation at earlier time points and a repressed conformation at later time points

suggesting that early replication favors the assembly of accessible chromatin, or that

accessible chromatin is replicated early [192]. Accordingly, the globin locus replicates

late in non-erythroid cells and early in cells expressing the globin genes. Transgenic

studies have shown that the timing of replication is regulated by the LCR [193], however,

this activity, like the chromatin opening activity, appears to be redundant in the

48

endogenous β-globin locus [194]. It is not known how or if replication affects the

association of transcription factors and Pol II with the globin locus.

Here, we analyzed the interaction of transcription complexes with the human and

murine β-globin loci in erythroid cells expressing either the human embryonic/fetal ε-

and γ-globin (K562 cells) or the murine adult β-globin genes (MEL cells) using

chromatin immunoprecipitation (ChIP).

Results

We began our studies by examining the interaction of various components of

transcription complexes and erythroid transcription factors with the β-globin gene locus

in murine and human erythroleukemia cell lines. Figure 4-1 shows a diagram of the

human and murine β-globin loci and the localization of primers used in the ChIP

experiments.

We used chromatin immunoprecipitation (ChIP) and PCR to analyze the interaction

of RNA Pol II (Pol II), TBP, TFIIB, NF-E2 (p45), and acetylated histone H3 with

different regions of the β-globin gene locus. As a negative control we also analyzed the

interaction of the proteins with the necdin gene, which is not active in erythroid cells.

NF-E2 is a hematopoietic specific transcription factor composed of a large (p45) and a

small (p18) subunit [126] and known to interact with several sequences in the human and

murine β-globin loci [136, 195]. We investigated the interaction of these proteins in two

cell lines; a human erythroleukemia cell line (K562) expressing the embryonic ε and fetal

γ-globin genes but not the adult β-globin gene, and a murine erythroleukemia cell line

(MEL), in which the adult βmaj- and βmin-globin genes are expressed but not the

embryonic εγ and βh1 genes.

49

Figure 4-1. Diagrammatic representation of the human and murine β-globin gene locus. The human β-globin gene locus: the embryonic ε-globin gene, the two fetal γ-globin genes, and the adult δ- and β-globin genes. The LCR is located upstream of the ε-globin gene and composed of at least 5 HS sites. The murine β-globin locus: the εγ and βh1 globin genes, which are co-expressed in the embryonic yolk sac and the βmaj and βmin globin genes, which are expressed during the fetal and adult stages of erythropoiesis. The overall organization of the LCR is highly conserved between the murine and the human β-globin locus. PCR fragments analyzed in the ChIP experiments are indicated as lines (horizontal bars) below the respective regions.

The data show that RNA polymerase II is efficiently crosslinked to LCR HS2 in

K562 cells (Figure 4-2). The interaction of Pol II with the globin genes is consistent with

the expression pattern in these cells. Pol II is detectable at the embryonic ε- but not at the

adult β-globin gene. The results further show that Pol II does not interact with the HS2

5’flanking region, demonstrating that Pol II specifically interacts with the core of HS2.

We also detected interactions of phosphorylated Pol II with HS2 and the ε-globin gene in

K562 cells, indicating ongoing transcription in these regions.

Because extragenic transcripts have been detected over the entire LCR in the

human β-globin locus we analyzed the interaction of basal transcription factors that are

50

not associated with the transcription elongation complex. The results show that both

TFIIB as well as TBP can be crosslinked to HS2 in K562 cells. These results

demonstrate that Pol II and other basal transcription factors, which are part of

transcription initiation complexes, are associated with the LCR element HS2 in vivo. The

presence of components of the PIC indicates that the crosslinking of Pol II to HS2 is not

solely due to elongating polymerase. NF-E2 is clearly detectable at HS2 in K562 cells,

whereas the signal representing the ε-globin gene is very weak. None of the proteins

interact with the necdin gene.

Figure 4-2. Interaction of transcription factors and RNA polymerase II with the human β-globin locus and the human necdin gene. Antibodies used were specific for RNA polymerase II (αPol II), RNA polymerase II phosphorylated at the C-terminal domain (αPol II Phosph.), TBP, TFIIB, NF-E2 p45, and acetyl histone H3 (αhistone H3 acet.). PCR was performed with primers to regions in the β-globin locus as indicated (see Figure 4-1). No antibody samples are negative controls. Input represents the positive PCR control taken from the no antibody sample.

The observation that Pol II is not detectable at the HS2 5’flanking region is

interesting and suggests that transcription initiates within the HS2 core enhancer and not

51

upstream of HS2. In order to analyze this question in more detail, Padraic Levings in the

laboratory used nuclear run-on experiments to determine regions of active transcription in

the human β-globin locus in K562 cells. He found that ongoing transcription is

detectable in a region upstream of HS5, in the HS2 core, and in the ε globin gene, but not

in the HS2 5’ flanking region. These results are consistent with our ChIP analysis and

suggest that transcripts initiating far upstream of HS2 do not elongate beyond the HS2

5’flanking region analyzed here, consistent with data published by Kim and Dean [196].

Because the nuclear run-on HS2 probe is specific for the core region his data suggest that

transcription initiates within HS2 and not, or not exclusively, downstream of HS2 as

proposed by Routledge et al. [188]. Transcription upstream of HS5 probably initiates

within the LTR of a retrovirus as previously described [197].

We also analyzed the interaction of transcription factors and Pol II with regulatory

elements in the murine β-globin locus in MEL cells (Figure 4-3A). The results of these

experiments are very similar to those in the K562 cells and show that Pol II, TBP, and

TFIIB can be crosslinked to both HS2 and the β-globin gene promoter, but not to a region

5’ of HS2. The precipitation of proteins crosslinked to the ε-globin gene is extremely

inefficient. The inefficiency in crosslinking phosphorylated Pol II to HS2 and the β-

globin promoter in the murine locus is consistent and may be due to species-specific

differences in the ability of the antibody to precipitate crosslinked polymerase.

In contrast to the results in K562 cells, NF-E2 (p45) can be crosslinked to HS2 and

to the β-globin gene promoter. We have shown previously that NF-E2 interacts with a

NF-E2 like sequence located downstream of the murine βmaj-globin transcription start

site [195]. This sequence deviates in one position from what is considered the consensus

52

NF-E2 binding site, but it is identical to the NF-E2 interaction sequence present in human

LCR HS3 [126, 136].

Figure 4-3. Interaction of transcription factors with and transcription in the murine β-globin locus in MEL cells. A) Interaction of transcription factors and RNA polymerase II with the murine β-globin locus and the murine necdin gene. Antibodies used were specific for RNA polymerase II (αPol II), RNA polymerase II phosphorylated at the C-terminal domain (αPol II-Ph), TBP, TFIIB, NF-E2 p45, and acetylated histone H3 (αAcetyl-H3). PCR was carried out with primers corresponding to regions in the murine β-globin locus as indicated (see Figure4-1). The no antibody and input controls were processed as described in the legend to Figure 4-2. B) Semi-quantitative RT-PCR analysis of β-globin gene transcription in DMSO induced and uninduced MEL cells. Relative expression of β-globin mRNA compared to expression of β-actin was found to be 3-fold lower in uninduced MEL cells.

The NRO analysis in MEL cells performed by Padraic Levings demonstrates that

while HS2 and the β-globin gene are transcribed, a region upstream of HS5 is not. This

result is likely due to the fact that the retrovirus element present upstream of HS5 in the

human locus is not present in the murine β-globin gene locus.

The MEL cells employed in these experiments express the β-globin gene, and thus

no induction (with DMSO) was used. Moreover, interactions between Pol II and NF-E2

53

with the globin locus can be detected by ChIP assay in the absence of DMSO. We found

that incubating the cells with 1% DMSO for 3 days led to an increase in globin gene

transcription (α- and β-globin) less than 5-fold compared to uninduced cells (Figure 4-3

B).

If the model according to which Pol II is recruited first to the LCR, modified and

transferred to the globin genes is correct [96], one may predict that Pol II associates with

the LCR and the globin genes at different time points during activation of the globin

locus in the cell-cycle. To address this question, we examined the interaction of

transcription complexes with the LCR and globin genes during S phase in K562 cells

(Figure 4-4). Cells were synchronized using the double thymidine block method, which

arrests the cells at the G1/S phase border [167]. At specific time points after release from

cell-cycle arrest, cells were subjected to formaldehyde crosslinking and ChIP analysis.

In these experiments, we used antibodies against Pol II, NF-E2 (p45) and USF2.

We have shown that USF2 interacts with the human β-globin promoter in K562 cells

(described in Chapter 6) and hypothesized that it may contribute to the repression of the

β-globin gene [163]. We analyzed the association of these proteins with four different

regions of the human β-globin locus. Cells were crosslinked at time 0, and 15 minutes,

45 minutes, 2 hours, and 6 hours after release from cell cycle arrest.

At time 0, Pol II interacts with HS2, the ε-, and γ-globin genes, but not with the β-

globin gene. There is no interaction of NF-E2 or USF2 with the regions of the globin

locus at this time point. The only change after 15 minutes into S phase is the association

of NF-E2 with HS2. After 45 minutes, Pol II is no longer detectable at the ε-globin gene

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but remains associated with HS2 and the γ-globin genes. At this time point, NF-E2

interacts with the γ-globin genes.

Figure 4-4. Interactions of transcription factors and RNA polymerase II with the human β-globin locus during early S phase in synchronized K562 cells. Synchronized K562 cells were analyzed for DNA content by flow cytometry (shown on top). The remaining cells were harvested at time 0 or at specific time points after release from cell cycle arrest (as indicated), and subjected to ChIP using antibodies specific for RNA polymerase II (αPol II), the p45 subunit of NF-E2 (αNF-E2), and USF2 (αUSF2); time 0 represents the status of the blocked cells at the G1/S phase boundary. PCR was performed using primers specific for HS2, the ε-globin gene, the γ-globin gene, and the β-globin gene as indicated (see Figure 4-1). The no antibody and input controls were processed as described in the legend to Figure 4-2.

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After 2 hours, Pol II no longer interacts with the LCR or the ε- and γ-globin genes.

At this time there is a clear interaction of NF-E2 and USF2 with HS2 and the ε- and γ-

globin genes. The interaction of NF-E2 with the β-globin gene is less pronounced. After

6 hrs, Pol II re-associates with HS2 and the γ-globin genes, but not with the ε-globin

gene. Accompanied with the reappearance of Pol II is the dissociation of USF2 from the

genes and HS2.

We also analyzed the interaction of transcription factors with the murine globin

locus during S phase in synchronized MEL cells (Figure 4-5). We used antibodies

specific for Pol II, TFIIB, NF-E2, and acetylated histone H3. At time zero, Pol II, TFIIB,

NF-E2 and acetylated histone H3 can be detected at LCR HS2 and the β-globin promoter.

After 7 minutes, NF-E2 and acetylated H3 but not Pol II can be detected at HS2 and

weaker at the β-globin gene. A similar pattern was observed after 45 minutes, except that

now Pol II is beginning to re-associate with HS2 and the β-globin promoter. Finally,

after 2 hours, Pol II, TFIIB, NF-E2, and acetylated H3 are associated with HS2 and the β-

globin promoter.

The results show that Pol II dissociates and re-associates with the globin locus

during early S phase in MEL cells. In contrast to the results with K562 cells, NF-E2

appears to remain associated with the globin locus at all the time points analyzed here.

All experiments have been repeated several times with essentially the same results. The

only exception is that in some experiments NF-E2 (p45) can be crosslinked to HS2 in

synchronized K562 cells at time 0. The variation may be due to subtle changes in the

status of the cells during synchronization.

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Figure 4-5. Interactions of transcription factors and RNA polymerase II with the human β-globin locus during early S phase in synchronized MEL cells. The cells were analyzed for DNA content by flow cytometry at each time point (shown at left). The remaining cells were harvested at time 0 or at specific time points after release from cell cycle arrest (as indicated), and subjected to ChIP using antibodies specific for RNA polymerase II (αPol II), TBP (αTBP), TFIIB (αTFIIB), the p45 subunit of NF-E2 (αNF-E2), and acetylated histone H3 (αH3acetylated); time 0 represents the status of the blocked cells at the G1/S phase boundary. PCR was performed using primers specific for HS2 and the βmaj-globin gene as indicated (see Figure 4-1). The no antibody and input controls were processed as described in the legend to Figure 4-2.

We next analyzed how dissociation and re-association of Pol II to the β-globin

locus in K562 and MEL cells correlates with the timing of replication. Since the cells are

blocked at the border of G1 and S phase, it is assumed that when the cells are released

from the block they will synchronously enter S phase. They should then undergo

replication early in S phase since the β-globin locus is early replicating in erythroid cells

such as K562 and MEL [198]. As a late replicating control, we used the necdin gene,

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which is a brain-specific imprinted gene that replicates late in S phase in all tissues

except brain.

In this experiment, the cells were blocked at the border of G1 and S phase, cells

collected at time 0, released from the block and cells collected at the same time points as

for the flow cytometry and ChIP experiments. DNA was extracted and used as a

template for PCR with primers against ε-globin and necdin for K562 cells (Figure 4-6),

β-globin and necdin for MEL cells (Figure 4-7). Based on the calculated ratios it was

estimated that K562 cells complete replication of the β-globin locus by 2 hours after

release from the block (Figure 4-6). For MEL cells it was estimated that replication of

the β-globin locus is also complete by 2 hours after release from the block (Figure 4-7).

Figure 4-6. Semi-quantitative PCR analysis of DNA content of synchronized K562 cells. Cells were isolated at each time point, DNA extracted and PCR performed with primers to ε-globin and necdin promoters. Equal amounts of the two PCR products were run in the same lane and gels were stained with SyBr Green. Bands were quantitated with Image Quant and the ratio of ε-globin to necdin was calculated.

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Discussion

Expression of the β-globin genes is regulated by the LCR [183, 199]. Whereas

studies using transgenic mice demonstrate that the LCR has both chromatin opening and

enhancer activities [111], studies in the endogenous murine locus show that the function

of the LCR is limited to stimulating high-level globin gene transcription [200]. The

ability of the LCR to regulate chromatin structure is thought to be important for

conferring protection from position effects at ectopic sites in transgenic studies, a

defining activity for locus control regions [199]. It is not known at what level the LCR

opens chromatin structure, e.g., unfolding of higher order structure and modification of

histones, or whether chromatin opening is a secondary event and the consequence of LCR

mediated re-location of the globin locus into a transcriptionally active site in the nucleus

[201].

Figure 4-7. Semi-quantitative PCR analysis of DNA content of synchronized MEL cells. Cells were isolated at each time point, DNA was extracted and PCR was performed with primers to β-globin and necdin promoters. Equal amounts of the two PCR products were run in the same lane and gels were stained with SyBr Green. Bands were quantitated with Image Quant and the ratio of β-globin to necdin was calculated.

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Recently extragenic transcripts were detected over the LCR and in between the

globin genes [156, 187, 202]. The generation of these transcripts was found to correlate

with stage-specific transcriptional activity and DNaseI sensitivity of β-globin locus

subdomains [102]. This led to the hypothesis that transcription complexes are recruited

to various regions in the globin locus and track along the DNA in a unidirectional manner

thereby modifying chromatin structure. This is a plausible model because Pol II

transcription complexes are known to be associated with chromatin modifying activities

[203], and transcription through regulatory memory elements has been shown to be the

initial event in setting up activating or repressing functions of these elements in

Drosophila [204]. Extragenic transcripts have also been detected in other complex gene

loci suggesting a broader role in the modulation of chromatin domains [205].

The role of the LCR in mediating high-level expression of the β-globin genes is

unquestionable. The LCR and the genes appear to be in close proximity in cells

expressing the globin genes, suggesting physical contacts between activities associated

with the LCR and proteins interacting with the promoter [120, 206]. How this contact

aides in stimulating transcription is not known, but could involve the transfer of activities

from the LCR to the globin genes [96]. In the experiments described in this chapter we

analyzed the interaction of transcription complexes with LCR HS core elements and the

globin genes during S phase. The data demonstrate that RNA Pol II dissociates and re-

associates with the globin genes during replication, showing that the replication

machinery is capable of displacing transcription complexes. Interactions of NF-E2 and

USF appear to precede the re-association of Pol II with the globin locus suggesting that

these proteins assist in the recruitment of Pol II during S phase. The data thus support a

60

model in which the LCR and the globin genes cooperate in recruiting transcription

complexes to the globin locus.

We began our studies by addressing the question of whether RNA Pol II is

recruited to the LCR independent from its known interactions with the globin genes. The

ChIP assay per se does not differentiate between direct or indirect interactions. We show

here, that Pol II interacts with the HS2 core region together with other components of the

transcription initiation complex. This result shows that the crosslinking of Pol II to the

core is not solely due to elongating transcription complexes that are in the process of

generating the extragenic transcripts, but instead is due to Pol II that may be poised to

start extragenic transcription or ready for transfer to the gene promoters. This conclusion

is also supported by our observation that Pol II is not detectable in a region between the

HS2 and HS3 cores.

How does the polymerase interact with the LCR core HS sites? Is the

crosslinking to these sites due to interactions between HS2 and the globin genes, which

would be fixed in the ChIP assay, or is it due to direct association of transcription

complexes with the LCR HS core regions? We believe that the latter is true for the

following reasons: If the association of polymerase II with the HS2 core is due to protein

mediated interactions with the globin promoters, all proteins detected at the promoter

regions may also be detectable at the LCR and vice versa. This is clearly not the case.

For example, NF-E2 interacts strongly with HS2 in unsynchronized K562 cells, but only

weakly with the ε-globin promoter. If the interaction of Pol II with HS2 is indirect and

caused by interactions with the ε-globin promoter then antibodies against NF-E2 should

also efficiently precipitate the ε-globin gene, as it is likely part of the same complex.

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Although it is possible, but highly unlikely, that the mechanism of looping sequesters

proteins at either side of the interaction such that they are not detected on the other side

by formaldehyde crosslinking.

The results from the NRO analysis are consistent with the ChIP data and

demonstrate that transcription is ongoing at places where transcription complexes are

detected; supporting the notion that Pol II is recruited to the LCR independent from

interactions with the promoter regions. It is also interesting to note that while HS2 is

transcribed in both MEL and K562 cells, the HS2 5’ flanking region in K562 cells is not,

consistent with the lack of Pol II binding to this region. This result suggests that

extragenic transcripts over the LCR do not represent long continuous transcripts but are

due to multiple transcription initiation sites in HS2, HS3, and possibly other regions of

the LCR. Our results are largely consistent with data recently published by Johnson et al.

[189] demonstrating that Pol II recruitment to the LCR is restricted to the core HS sites.

Sawado et al. [190] recently presented data demonstrating that an important

function of the mouse LCR is to stimulate globin gene expression at the level of

transcription elongation. The interpretation is based on the fact that in the absence of the

LCR no elongating transcription complexes were found in the transcribed region of the

βmaj-globin gene. If transcription complexes are stalled at the β-globin promoter in the

absence of the LCR, one might predict that more β-globin promoter bound Pol II is

detected in the mutant locus, compared with the wild-type locus. However, Sawado et al.

[190] found a two-fold reduction in promoter bound Pol II in the mutant allele suggesting

that another important function of the LCR is to mediate the recruitment of Pol II to the

β-globin genes. It is difficult to estimate the contribution of the LCR in recruiting Pol II

62

to the genes by comparing promoter-bound Pol II in promoters in which elongation takes

place with those in which elongation does not take place. It is thus possible that the

contribution of LCR mediated delivery of Pol II to the β-globin gene may have been

underestimated in these studies. Another possibility that should be considered is that in

the absence of the LCR unproductive transcription complexes are recruited to the globin

genes but that in the presence of the LCR these complexes are first recruited to the LCR,

rendered transcriptionally competent, and transferred to the globin genes [96].

The results from the cell synchronization/ChIP experiment described here are

consistent with a model proposing that an important function of the LCR is to represent a

reservoir of transcription and maybe other protein complexes, which may be used to

provide these activities to the globin genes during various stages of the cell cycle. NF-E2

may play a major role in directing these activities to the globin genes. In K562 cells NF-

E2 interacts directly or indirectly with HS2 and both ε- and γ-globin genes at specific

stages during S phase. Not surprisingly, the interaction of NF-E2 with the globin locus

precedes the re-association of Pol II, consistent with the notion that factors are needed to

recruit Pol II back to the locus after replication. We observed a similar pattern at the β-

globin gene in MEL cells, although NF-E2 remains associated with HS2 and the β-globin

promoter, at least at the time points analyzed here. These data suggest an important role

of NF-E2 in recruiting Pol II to the globin locus, consistent with results from Johnson et

al. [179] demonstrating an impairment in Pol II/β-globin gene interactions in MEL cells

lacking NF-E2. In addition, it was recently shown that NF-E2 interacts with the human

β-globin locus control region before chromatin is re-modeled [207]. The summary data

suggest that NF-E2 interacts with the globin locus at specific stages during the cell cycle,

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prevents the formation of repressive chromatin structure, and facilitates the interaction of

transcription complexes with the globin locus.

Another interesting observation in these experiments is the interaction pattern of

USF2 during S phase in synchronized K562 cells. Previous studies have shown that USF

interacts with LCR core elements and with the β-globin promoter [143, 144, 163]. Our

data show that USF2 only interacts with HS2 and the globin genes at a stage at which

Pol II is not bound. It is conceivable that USF2, in conjunction with NF-E2, prepares

specific regions in the globin locus for the re-association of transcription complexes after

replication. Tolhuis et al. [120] recently proposed a model according to which all HS

sites in the globin locus participate in forming a specific chromatin conformation, called

the active chromatin hub (ACH). The interactions of NF-E2 and USF with the locus

control region and the genes at a stage in which Pol II is not bound in K562 cells (Figure

4-4, 2 hour time point) could indeed suggest that the LCR HS sites and the globin gene

promoters together orchestrate the recruitment of transcription complexes to the globin

gene locus, consistent with previous observations showing that enhancer and promoter

elements cooperate in the formation of DNaseI HS sites [208].

In summary, we have observed recruitment of Pol II directly to LCR HS2. We

also observed dynamic changes in the interaction of transcription factors and RNA-

polymerase II with the β-globin gene locus during S phase of the cell cycle. These

changes in interactions correlate with the occurrence of replication in these cells.

The results from the synchronization experiments are reproducible, with some

notable exceptions. In some experiments we detect NF-E2 at LCR HS2 at time 0 in

K562 cells. In addition, the recruitment of Pol II to the ε-globin gene in K562 cells is

64

somewhat variable and we have performed experiments in which Pol II is crosslinked to

the ε-globin gene after 6 hours. The variability may be due to subtle changes in the status

of the cells during synchronization.

65

CHAPTER 5 ANALYSIS OF THE MECHANISM OF RNA POLYMERASE II TRANSFER FROM

THE LCR TO THE β-GLOBIN GENE PROMOTER

Introduction

Erythroid-specific, high-level expression of the β-globin genes is regulated by the

locus control region (LCR), composed of multiple DNaseI hypersensitive (HS) sites and

located far upstream of the genes. Our lab and others have shown that the LCR core

elements recruit RNA Pol II [132, 169, 187-189]. The LCR HS sites are known to

interact with each other to form the LCR holocomplex. We proposed a model suggesting

that the LCR recruits Pol II and later transfers it to the gene promoters that are expressed

at specific developmental stages [96]. Recent evidence obtained by the 3C technique

show that the LCR holocomplex interacts with whichever gene is transcriptionally active

at a particular stage in erythroid cells [117, 120, 158].

The 3C technology mentioned above is a method in which protein mediated DNA

interactions in live cells are captured by formaldehyde crosslinking. The crosslinked

chromatin is fragmented by restriction enzyme digestion and subsequently ligated under

conditions that favor intra-molecular ligation. These conditions allow DNA segments

that are normally located far away from each other to be ligated if they associate via

protein-protein and protein-DNA interactions. If these segments do not interact, the

probability of generating ligation products between these fragments is very low.

Formation of the ligation products is monitored by PCR.

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Tolhuis et al. determined using 3C technology that the LCR HS sites interact with

each other, with the 3’ HS site, and with the active globin genes in globin expressing cells

[120]. The authors call the structure in which the LCR holocomplex interacts with an

active gene the active chromatin hub, while the intervening DNA loops out in agreement

with previously proposed looping models [101, 115]. The authors further showed that

during erythroid development in mice the 5’, 3’, and LCR hypersensitive sites interact

first to form a chromatin hub. The interaction of the genes with this structure then leads

to the formation of the active chromatin hub [158].

The data described above together with our own results from the synchronization

experiments support the LCR recruitment and transfer model. To test the Pol II transfer

hypothesis directly, we have developed a novel in vitro assay in which Pol II is first

recruited to an immobilized LCR construct and then transferred to a β-globin gene

promoter. We provide evidence that RNA polymerase can be transferred from the LCR

to a β-globin gene in vitro and demonstrate that the transfer is sensitive to mutations in

the β-globin gene promoter and facilitated by erythroid transcription factor NF-E2.

Results

As a first attempt to examine the possibility that Pol II can be recruited to the LCR

and transferred to the β-globin gene, we established an in vitro system utilizing

immobilized LCR templates and nuclear extracts from MEL cells (Figure 5-1).

A plasmid containing the entire human β-globin LCR was linearized, biotinylated

and immobilized to streptavidin coated magnetic beads as described by Leach et al.

[169]. After incubation with MEL nuclear extract the protein/DNA complex was washed

several times in binding buffer on a magnet. Then a plasmid containing the wild-type β-

67

globin gene was added to the LCR and incubated for 10 minutes at 30°C in binding

buffer. The supernatant containing the β-globin gene was then removed from the

immobilized LCR, and subjected to crosslinking with formaldehyde and ChIP analysis

using a Pol II specific antibody as described in the Materials and Methods chapter (Figure

5-2 lane 2).

Figure 5-1. A schematic representation of the in vitro Pol II transfer experimental procedure. The plasmid pLCR was linearized, biotin labeled, and immobilized on streptavidin coated magnetic beads. After incubation with MEL protein extracts, the LCR was placed on a magnet and all material not interacting with the LCR was removed. The immobilized LCR/protein complex was washed, resuspended in binding buffer and incubated in the presence or absence of the β-globin gene. After placing the samples on the magnet, the supernatant was removed and the sample containing the β-globin gene was crosslinked and subjected to ChIP analysis using an RNA polymerase II specific antibody.

A second sample was processed in the same manner except that no Pol II specific

antibodies were added; this sample served as a negative control (Figure 5-2 lane 4). To

control for diffusion, we incubated 50 µl binding buffer with the protein/LCR complex in

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the absence of the β-globin template for 10 minutes at 30° C. The reaction was placed on

a magnet and the supernatant was removed and incubated with the β-globin template for

10 minutes at 30°C, after which formaldehyde was added to the reaction to induce

crosslinking. The sample was then subjected to ChIP analysis with Pol II antibodies

(Figure 5-2, lane 1).

Figure 5-2. Transfer of RNA polymerase II from immobilized LCR constructs to the β-globin gene. The sample containing the β-globin gene was crosslinked and subjected to ChIP analysis using an RNA polymerase II specific antibody (Lane 2). The supernatant of the sample incubated without the β-globin gene for 10 minutes was later incubated with the β-globin gene template and subjected to crosslinking and ChIP analysis using an RNA polymerase II specific antibody (Lane 1). Lane 3 represents a reaction processed in the same manner as described for lane 2 except that recombinant NF-E2 was added to the reaction. Lane 4 represents a negative control in which no antibody was added. Lane 5 represents the positive control in which the β-globin gene was incubated with MEL protein extract and subjected to ChIP and PCR. Samples were processed as described in Fig 5-1 and subjected to PCR analysis using a set of primers specific for the human β-globin coding region. The βmaxi gene differs from the wild-type β-globin gene by 800 bp and served as a PCR control in these experiments.

As a positive control, we incubated the β-globin gene for 30 minutes at 30°C with

MEL protein extract and subjected the sample to crosslinking and ChIP analysis (Figure

5-2, lane 5). After reversal of the crosslink and purification of the DNA, the samples

69

were analyzed by PCR using primers specific for the coding region of the β-globin gene.

The βmaxi gene was added after the immunoprecipitation and served as a PCR control.

The result of this experiment demonstrates that Pol II can indeed be transferred

from the LCR to a β-globin template in trans (Figure 5-2, lane 2). The control

experiment shows that diffusion is negligible during the incubation time and does not

contribute significantly to the transfer of Pol II from the LCR to the β-globin gene

(Figure 5-2, lane 1). The inclusion of recombinant NF-E2 consistently enhanced the

transfer of Pol II from the LCR to the β-globin gene (Figure 5-2, lane 3) supporting

previous conclusions that NF-E2 is required for the recruitment of Pol II to the globin

gene.

In order to examine whether Pol II is specifically transferred to the β-globin gene

and not to sequences elsewhere in the plasmid we have performed the transfer

experiments in the presence of two restriction fragments derived from pRSβA/X, one

fragment containing the β-globin gene and the other containing the bacterial ampicillin

gene. The results are shown in Figure 5-3 and demonstrate that Pol II is specifically

transferred to the β-globin gene and not to other sequences present in the plasmid. In this

experiment we have increased the cycle number to visualize the signal corresponding to

the ampicillin gene and the β-globin gene in the no antibody control (lane 2). The data

show that the intensity of the β-globin signal increases in the presence of Pol II

antibodies, whereas the signal of the ampicillin gene remains unaltered.

To further analyze the specificity of the transfer, we performed the experiment with

several β-globin promoter mutants (described in [163]). In this experiment the βmaxi gene

was added during the incubation with the protein/LCR complex and served as an internal

70

control. The results show that a mutant promoter in which the E-box element at +60 is

altered is impaired in recruiting Pol II (Figure 5-4).

Figure 5-3. Analysis of the specificity of Pol II transfer. The plasmid pRSβA/X was digested with KpnI and SacI, incubated with the LCR/protein complex, and processed as described in Figure 5-1 and 5-2. The precipitate was analyzed by PCR using primers specific for the β-globin (β) or the bacterial ampicillin gene (Amp), which are present on different restriction fragments. Lane 1 shows a 100bp ladder.

Figure 5-4. Comparison of Pol II transfer using the wild-type β-globin gene (lane 1) and a mutant β-globin gene (lane 2) in which an E-box at +60 was altered. The samples were processed and analyzed as described in Fig 5-2 except that the βmaxi gene was added together with the wildtype or mutant β-globin gene and incubated with the immobilized LCR/protein complex.

The addition of BSA at the same concentration as NF-E2 did not enhance Pol II

transfer (Figure 5-5, lane 5), indicating that stimulation of Pol II transfer by NF-E2 is not

an unspecific effect due to increased protein content. To further examine whether Pol II

transfer is specific for the β-globin gene we have performed transfer experiments in the

presence of a β-globin gene lacking the TATA box. We have previously shown that this

71

mutant is inefficiently transcribed in vitro [163]. We found that the β-globin TATA box

is necessary for Pol II recruitment (Figure 5-5, lane 6) demonstrating that Pol II is

specifically transferred to the β-globin gene and not to sequences elsewhere in the

plasmid.

Figure 5-5. Transfer of RNA polymerase II to the β-globin gene in the presence of NF-E2 or BSA. Samples were prepared and analyzed as described in panels Figure 5-2 except that NF-E2, lane 4, or BSA, lane 5, were added during the transfer step. Lane 6 represents transfer to a mutant β-globin gene template lacking the TATA box.

To further test the specificity of the transfer we performed an experiment in which

we simultaneously incubated LCR/protein complexes with two different plasmids, one

containing LCR element HS2 and another containing the β-globin gene. The results in

Figure 5-6 show that Pol II is preferentially transferred to HS2 (lane 2), which could be

due to interactions between HS2 and the LCR/protein complex. Importantly, we

observed that in the presence of NF-E2, Pol II is preferentially transferred to the β-globin

gene and not to HS2 (lane 3). We also analyzed the transfer to the thymidine kinase

promoter and found that Pol II is not transferred to this template (compare Figure 5-6

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lanes 1 and 4). We hypothesized that proteins in our erythroid cell extracts would bind

the thymidine kinase promoter since there are ubiquitously expressed proteins present in

these extracts that should bind other mammalian promoters such as the proteins of the

PIC. This again demonstrates that the transfer is specific for the β-globin gene. The

graph in Figure 5-7 summarizes the results of multiple experiments.

Figure 5-6. Analysis of NF-E2 mediated transfer to the β-globin gene. Samples were processed as described in panel Figure 5-2 except that in lanes 2 and 3 the transfer was carried out in the presence of both β-globin and HS2 containing plasmids. In lane 3, NF-E2 was added to the transfer reaction. In lane 4, transfer to the thymidine kinase/human growth hormone gene (TK-HGH) construct was analyzed. Lane 1 represents the reaction of the no antibody control containing all three plasmids (β-globin, HS2, and TK-HGH). Lanes 5, 6, and 7 represent positive PCR controls for β-globin (lane 5), HS2 (lane 6) and TK-HGH (lane 7).

Discussion

It was proposed that one function of the LCR could be to recruit macromolecular

protein complexes including Pol II, which are subsequently transferred to individual

globin genes in a developmental stage specific manner [96, 169, 179]. Similar

conclusions were derived from studies on the TCRβ locus, which show that coactivators

73

and components of the transcription initiation complex are first recruited to an enhancer

element and subsequently delivered to the promoter [209].

We have developed an in vitro system to test the polymerase transfer hypothesis

and show that Pol II can indeed be transferred from an immobilized LCR template to the

β-globin gene (Figure 5-1). Moreover, the transfer of Pol II was enhanced by

recombinant NF-E2, supporting previous studies that implicate NF-E2 in the recruitment

of Pol II to the β-globin gene [179]. The NF-E2 mediated stimulation of Pol II transfer is

not due to an increase in protein concentration as the same concentration of BSA had no

effect.

Figure 5-7. Quantitative summary of the transfer experiments. The experiments were repeated several times and the data were quantitated after gel electrophoresis using SyBr green, Storm scanner, and Image Quant. The bars represent the mean of the results of two to three experiments.

The observation that Pol II was not transferred to either the bacterial ampicillin

resistance gene or to the TK/HGH gene demonstrates that transfer is specific for the β-

globin gene promoter (Figure 5-3 and Figure 5-6) when using extracts from erythroid

cells. It is likely that transfer would occur to other promoters as well if they contain

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binding sites for proteins that mediate the transfer process, as I have shown to be the case

for NF-E2.

I also analyzed elements required for Pol II transfer to the β-globin promoter.

There was no transfer to the β-globin gene with a mutation at the TATA box and reduced

transfer to a promoter with a mutant +60 E-box (Figs. 5-4 and 5-5). This supports

previous data from our lab showing that the TATA box and +60 E-box are required for in

vitro transcription and suggest that these elements exert at least part of their function by

recruiting the polymerase [163].

The combined data suggest that the β-globin LCR, like other enhancer elements

[210], act in trans to assist in the recruitment of transcription complexes to the globin

genes. Although this in vitro system completely ignores nucleosome and higher order

chromatin structure, we believe that it does reveal some aspects of LCR function. It was

previously shown that specific characteristics of HS site formation and function,

including the recruitment of transcription complexes, can be recapitulated in vitro in the

absence of chromatin assembly [169]. This demonstrates that chromatin is not directly

involved in recruiting transcription complexes to the globin locus but rather regulates a

preceding step, which is to render regulatory and functional sites available or unavailable

for the interaction with transcription factors and polymerase.

I show that Pol II can be transferred from immobilized LCR templates to globin

gene constructs in vitro and have established in vitro conditions for the analysis of RNA

polymerase transfer in the β-globin locus. In summary, we propose that transcription

complexes are first recruited to a highly accessible LCR holocomplex (Figure 5-8). The

genes are subsequently brought in contact with the LCR holocomplex, consistent with

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data from conformational studies [120, 158], and transcription complexes are transferred

from the LCR to the promoter regions. This process is mediated by NF-E2 and possibly

other proteins (e.g. USF2). It is also facilitated by bringing strong globin gene promoters

(with high affinities for the transcription complex) in close proximity to LCR bound

transcription complexes.

Figure 5-8. Model of transcription complex recruitment to the β-globin gene. The LCR and other HS sites have been shown to interact to form a chromatin hub. In the active chromatin hub, the expressed genes interact with the HS sites. We propose that transcription and other protein complexes are first recruited to a highly accessible LCR holocomplex in the context of the proposed chromatin hub. The genes come in close proximity to the LCR holocomplex by as yet unknown mechanisms that may involve local remodeling of chromatin structure at the active promoters. Transcriptions complexes are then transferred from the LCR to high affinity binding sites at the globin gene promoters. This transfer is facilitated by NF-E2 and/or related proteins.

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CHAPTER 6 DETERMINING THE FUNCTION OF THE USF, TFII-I AND HDAC3 PROTEINS IN

β-GLOBIN GENE REGULATION

Introduction

At least two modes of regulation control the stage-specific expression of the globin

genes. The first parameter regulating stage-specific expression is the relative location of

the genes with respect to the LCR. When the order of the genes relative to the LCR is

reversed, the β-globin gene is inappropriately expressed at the embryonic stage [114].

The second parameter is the binding of stage-specific trans-acting factors to the

individual globin promoter regions leading to activation of transcription at the appropriate

times during development. A well-characterized example for this mode of regulation is

represented by the transcription factor EKLF, which specifically activates expression of

the adult β-globin gene during definitive erythropoiesis [137-139, 142]. EKLF exerts part

of its activating function by recruiting chromatin-remodeling complexes to the β-globin

gene promoter, which leads in turn to a local change in nucleosome organization [141].

The basal promoter of the human adult β-globin gene is composed of a TATA-like

element and an initiator sequence, both of which were shown to interact with the TFII-D

protein complex [211, 212]. In addition to components of the TFIID complex

(specifically, TAFII250 and TAFII150), a diverse group of other proteins can also

interact with initiator sequences [150, 170]. Notable among these are the helix–loop–

helix proteins USF and TFII-I, because both proteins have been implicated in the

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recruitment of transcription complexes to TATA-less promoters and in the stabilization of

transcription complexes in TATA-box-containing promoters [145, 146, 213].

Regulatory DNA elements located downstream of transcription initiation sites

(downstream promoter elements, DPE) that contribute significantly to the formation of

stable transcription complexes have been well characterized in a variety of Drosophila

genes [214]. In some cases these downstream promoter elements function in the absence

of TATA elements and cooperate with initiator sequences to recruit transcription

complexes. The transcription cofactor NC2 (negative cofactor 2), originally identified as

a repressor of RNA Pol II transcription, activates transcription of DPE-containing

promoters and inhibits transcription of TATA-box-containing promoters in vitro [215].

The human β-globin gene contains two conserved E-box motifs located 3' to the

initiator sequence. The first E-box overlaps the initiator while the second E-box is

located 60 bp 3' to the transcription initiation site (Figure 6-1). Previous results in the lab

demonstrate that the human β-globin promoter’s TATA-like motif, initiator and

downstream E-box element are all required for high level transcription in vitro [163].

This indicates that E-box motifs contribute to the efficient formation of transcription

complexes on the adult β-globin gene. The same study also indicated that USF1 and

TFII-I interact with the initiator and overlapping E-box, while USF1 and USF2 interact

with the downstream E-box in vitro [163].

Here I investigated the interactions of the HLH proteins in vivo. The data show

that USF1, USF2, TFII-I and p45 interact with the β-globin promoter in vivo, supporting

the previous in vitro results [163]. By comparing the protein-DNA interactions on

expressed and non-expressed β-globin gene promoters, it was discovered that USF2 and

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TFII-I interact more efficiently with the non-expressed than with the expressed gene.

The interactions of USF1 and p45 with the β-globin promoter are restricted to erythroid

cells expressing the β-globin gene. Further, TFII-I was implicated in recruiting HDAC3

to the human β-globin gene promoter in cells that are not expressing the gene. The

results indicate that helix–loop–helix proteins contribute to the formation of transcription

complexes on the adult human β-globin gene, and suggest that the differential association

of HLH proteins with the basal promoter contribute to the stage-specific expression of the

gene.

Figure 6-1. Sequence alignment of the human β-globin downstream promoter region. Shown are three sequences of the adult β-globin downstream promoter region from human (H), mouse (M) and rabbit (R). Shaded boxes highlight the position of E-box motifs (CANNTG). Two of these E-boxes, the one overlapping the initiator and the distal E-box, are conserved in all three species, whereas the E-box located at +20 is only present in the human and rabbit genes. The open box delineates the position of the MARE-like sequence.

Results

In these studies I used two erythroid cell lines, MEL, a murine erythroleukemia cell

line expressing the adult β-globin gene, and K562, a human erythroleukemia cell line

expressing the embryonic β-type globin genes. I examined whether USF1, USF2 or

TFII-I interact with the β-globin gene in vivo by performing ChIP experiments in MEL

and in K562 cells. The results of these experiments are shown in Figure 6-2 and

demonstrate that both USF1 and USF2 could be crosslinked to the β-globin gene

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promoter in MEL cells in vivo (Figure 6-2, lanes 4 and 5). TFII-I could also be

crosslinked to the globin promoter (Figure 6-2, lane 6); however, the signal for TFII-I

was consistently weaker in MEL versus K562 cells. Antibodies recognizing the p45

subunit of NF-E2 and acetylated histone H3 also precipitated the β-globin gene in MEL

cells (Figure 6-2, lanes 7 and 8, respectively). The precipitation with acetylated H3

antibodies was expected from previous studies [103].

To analyze whether the interaction of USF proteins with the β-globin promoter

could be correlated with the transcriptional activity of the gene I carried out ChIP

analyses in K562 cells, in which the β-globin gene is not expressed. In contrast to our

observations in MEL cells, USF1 does not interact with the β-globin gene in K562 cells,

whereas TFII-I and USF2 do interact (Figure 6-2, lanes 5 and 6). The interactions of p45

(NF-E2) and acetylated H3 with the β-globin gene were also less efficient in K562 cells

when compared to MEL cells (compare Figure 6-2, lanes 7 and 8, MEL versus K562). In

control experiments I amplified a region within the LCR located upstream of HS2 (HS2 5'

flank; Figure 6-2, lanes 10–16). The results demonstrate that the observed interactions of

USF1, USF2, TFII-I and p45 in erythroid cells are specific for the β-globin gene.

To analyze the possibility that the differential binding of USF1, NF-E2 and, to a

lesser extent, TFII-I is due to differential expression of these proteins in the two cell lines,

Kelly Leach in our laboratory carried out western blotting experiments using antibodies

specific for USF1, USF2, TFII-I and NF-E2. The results show that USF1 and USF2 are

expressed at similar levels in K562 and MEL cells. However, TFII-I is expressed at

higher levels in K562 cells and p45 (NF-E2) is expressed at higher levels in MEL cells.

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Figure 6-2. Characterization of protein–DNA interactions in the human β-globin downstream promoter region in vivo. Chromatin immunoprecipitation (ChIP) experiment analyzing the interaction of proteins with the murine/human β-globin gene or the HS2 5'flanking region in MEL and K562 cells in vivo. Complexes were precipitated with either no antibody (lanes 3 and 11) or with antibodies specific for USF1 (lanes 4 and 12), USF2 (lanes 5 and 13), TFII-I (lanes 6 and 14), NF-E2 (p45, lanes 7 and 15) or acetylated histone H3 (lanes 8 and 16). DNA was purified from the precipitate and analyzed by PCR for the presence of the murine or human β-globin gene (210 and 321 bp, respectively, lanes 2–8) or the murine or human HS2 5'flank (336 and 565 bp, respectively, lanes 10–16). As positive controls, the input DNA was also analyzed by PCR (lanes 2 and 10). Lanes 1 and 9 show 100 bp markers.

Based on previously published data [216], I investigated potential interactions

between TFII-I and HDAC3 with each other and the promoter of the silenced β-globin

gene in K562 cells. For this, I carried out chromatin double immunoprecipitation

(ChDIP) experiments, in which two subsequent ChIP reactions with two different

antibodies are performed. In the ChDIP, the first ChIP assay was performed with an

antibody against TFII-I. The protein-DNA complexes eluted from the beads were then

used as substrate for another CHIP assay using an antibody against HDAC3. The results

show that in the first round of immunoprecipitation TFII-I was bound to the β-globin

gene promoter at levels above that of the no antibody background (Figure 6-3). In the

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subsequent round of immunoprecipitation, HDAC3 was detected in β-globin templates

bound by TFII-I.

In contrast there were no ε-globin promoter templates detected in both TFII-I and

HDAC3 precipitates. As a negative control, I also analyzed the linker region between

HS2 and HS3 and conclude that TFII-I and HDAC3 do not interact with this region. The

no antibody background after the second round of immunoprecipitation was drastically

decreased compared to the first one, probably due to the loss of non-specific interactions

with the beads.

Figure 6-3. Chromatin double immunoprecipitation (ChDIP) experiment analyzing the interaction of TFII-I and HDAC3 at different regions within the human β-globin locus. Complexes were first immunoprecipitated with an antibody against TFII-I and then these TFII-I selected complexes were immunoprecipitated with an antibody against HDAC3.

Although the ChDIP experiment showed that TFII-I and HDAC3 are both bound to

the promoter of the non-expressed β-globin gene, it did not provide evidence of direct

interactions between these proteins. In order to examine the possibility that TFII-I

directly interacts with HDAC3, we carried out co-immunoprecipitation experiments.

Whole cell extracts from K562 cells were immunoprecipitated with antibodies against

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USF1, TFII-I, Pol II, USF2, HDAC3 and a no antibody negative control. The complexes

eluted off the beads were separated by SDS-PAGE and transferred to a nylon membrane

for Western blot analysis. An antibody against HDAC3 was used for the Western

blotting detection.

Supporting our previous observation, we detected HDAC3 only in complexes first

immunoprecipitated with antibodies against TFII-I (Figure 6-4). In contrast, HDAC3

was not detected in the samples containing USF1, USF2 or Pol II. This shows that TFII-I

and HDAC3 directly interact in K562 cells.

Figure 6-4. Co-immunoprecipitation experiment analyzing the interaction of TFII-I and HDAC3. Whole cell extracts from K562 cells were first immunoprecipitated with the indicated antibodies then the complexes pulled down were run on SDS-PAGE and probed by with HDAC3 antibodies in a Western Blot.

Discussion

The promoter of the human β-globin gene is characterized by the presence of a non-

canonical TATA-box (CATAAA) located 25–30 bp upstream of the transcription start

site [217]. Deviation from consensus TATA sequences often weakens promoters and

leads to the requirement of additional elements for the efficient recruitment or

stabilization of transcription complexes. For example, a recent report demonstrated that

the TBP/TFII-A pre-initiation complex interacts only weakly with the CATAA motif

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[218]. An additional sequence that contributes to high level β-globin gene transcription is

an initiator element located at the transcription start site. Lewis et al. [211, 212] have

shown that this element interacts with the TFII-D complex in vitro and that mutations in

this sequence reduce transcription efficiency.

In addition to the initiator element, the β-globin gene contains several E-box motifs

downstream of the transcription start site [177] (Figure 6-1). Two of these elements are

conserved across species. Because previous studies have shown that E-box-binding

proteins can participate in the formation of transcription complexes on both TATA-

containing and TATA-less genes [145, 146, 213], we were interested in examining the

functional role of these proteins in β-globin gene transcription.

The β-globin initiator/E-box interacts with TFII-I and USF1, whereas the distal E-

box at +60 interacts with USF1 and USF2 in vitro [163]. There are a number of reports

in the literature documenting the interaction of USF and TFII-I with sequence elements in

the vicinity, mostly downstream, of transcription initiation sites [219-222]. Hsieh et al.

[223] recently identified an E-box motif located downstream of the APEG-1 gene from

+39 to +44. This element interacts with USF proteins and mutation of this E-box

dramatically reduces transcription.

TFII-I interacts with the β-globin initiator in vitro and in vivo and may play a role

in activating transcription of the β-globin gene in MEL cells. However, several

observations suggest that TFII-I is not required for β-globin gene transcription. First,

antibodies against TFII-I were not able to inhibit transcription of the β-globin gene in

vitro [163]. Secondly, crosslinking of TFII-I to the β-globin gene in adult erythroid cells

is inefficient compared to USF1 and USF2 (Figure 6-2). The ChIP assay does not provide

84

information as to what fraction of cells have a specific protein bound at a specific site. It

is thus possible that the weak PCR signal from TFII-I-selected MEL chromatin fragments

indicates that only a small fraction of cells have TFII-I bound at the initiator. This

fraction of cells could represent non-expressing cells. This is consistent with our

observation that TFII-I crosslinks more efficiently to the β-globin gene in K562 cells

(Figure 6-2A), suggesting that TFII-I may only be able to interact with the β-globin gene

in the absence of transcription complex formation.

We have performed RT–PCR (not shown) and western blotting experiments to

characterize the expression pattern of TFII-I in K562 versus MEL cells. The RT–PCR

analysis revealed that TFII-I is transcribed with equal efficiency in both cell types.

However, western blots of nuclear extracts consistently showed that full-length TFII-I

protein is present at higher levels in K562 than it is in MEL nuclear extracts. The reason

for the lower amount of TFII-I in MEL nuclear extracts may be due to post-

transcriptional mechanisms. This lower level of TFII-I in MEL cells may contribute to

the lower level of TFII-I bound to the β-globin gene promoter in these cells.

Whether the binding of TFII-I to the β-globin promoter in erythroid cells with

embryonic phenotype contributes to repression of the β-globin gene or simply reflects the

absence of TFII-D binding was an issue that we sought to resolve. A recent report shows

that TFII-I interacts with histone deacetylase 3 [216]. It is thus possible that TFII-I

interacts with the β-globin gene in embryonic erythroid cells, recruits histone

deacetylases and contributes to the repression of β-globin gene expression by rendering

the chromatin structure inaccessible. Our ChIP data clearly show a strong reduction of

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acetylated H3 binding to the β-globin promoter in K562 cells compared to MEL cells

(Figure 6-2, lane 8).

The combination of ChDIP and co-immunoprecipitation experiments indicated that

TFII-I and HDAC3 interact with each other and associate with the β-globin gene in non-

expressing erythroid cells. The data support a model according to which TFII-I recruits

HDAC3 to alter the chromatin structure and therefore to silence the β-globin gene

expression.

Our data led us to formulate a hypothesis on the function of proteins interacting

with the β-globin downstream promoter during development (Figure 6-5). We propose

that TFII-D, NF-E2, USF1 and USF2 interact with the globin promoter in adult erythroid

cells and provide a platform for the efficient recruitment of transcription complexes [179,

195, 211, 212]. The functional role of NF-E2 in this process remains to be established,

but other data presented before (Chapter 5) suggest that it may be involved in the transfer

of RNA Pol II from the LCR to the gene promoter.

The interaction of TFII-I and USF2 with the β-globin promoter in K562 cells could

contribute to the repression of this gene at the embryonic stage of erythroid development

by recruiting histone deacetylase activities. Other studies also implicate USF2 as a

potential transcriptional repressor [224]. However, in adult cells, USF2 and USF1

interact with the E-box at +60 and stimulate transcription. This conclusion is supported

by our previous observation that in vitro transcription of the β-globin gene is inhibited by

pre-incubating the protein extracts with USF2 antibodies [163].

In summary, our data provide evidence that HLH proteins differentially interact

with the initiator and downstream promoter during erythroid development. The

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combination of USF1, USF2, NF-E2 and, perhaps, TFII-I contribute to expression of the

β-globin gene. Conversely, interactions of TFII-I along with USF2 recruit HDAC3

within the β-globin gene, leading to the repression of the β-globin gene.

Figure 6-5. Summary and model of protein–DNA interactions at the human β-globin promoter. The human β-globin promoter consists of a TATA-box and an initiator. These sequences are bound by the TFII-D complex in cells expressing the β-globin gene (active). Additional sequences required for the formation of active transcription complexes are MARE and E-box elements located in the downstream promoter region. These sequences are bound by NF-E2 (p45 and p18) and USF, respectively. It is proposed that in cells not expressing the β-globin gene (inactive), TFII-D is not bound and the initiator sequence is occupied by protein complexes consisting of TFII-I and USF2.

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CHAPTER 7 CONCLUSIONS AND FUTURE DIRECTIONS

ChIP Footprinting

We have established a novel technique that allows the identification of sequences to

which a specific protein binds in the context of chromatin. This represents a powerful

tool for discovering binding sites of proteins not only in the β-globin locus but also in

other areas of the genome. We used this technique to demonstrate that NF-E2 binds to a

non-consensus MARE in the mouse β-globin downstream promoter region.

This technique could be refined by optimizing the restriction enzyme digestion

step. I noticed the appearance of unspecific DNA degradation products after long

incubation times with the restriction enzymes. The Grosveld group followed a similar

digestion protocol for their 3C experiments except that they utilized SDS to remove any

non-crosslinked proteins from the DNA and Triton X-100 to sequester SDS for more

efficient digestion [182]. However, whether this is applicable for the ChIP footprint

technique is questionable because the following step after fragmentation is

immunoprecipitation and not, as is the case in the 3C method, DNA ligation.

The ChIP footprinting technique could be used to further identify binding sites for

several of the proteins our lab investigates. For example, we could analyze whether TFII-

I binds to the β-globin initiator or to the downstream E-box elements in vivo. Similarly

we could investigate a possible difference in USF binding between β-globin promoters of

expressing and non-expressing cells.

88

ChIP within the β-Globin Locus

Using the ChIP assay, I showed that RNA Pol II is directly recruited to HS2 of the

LCR. As expected, Pol II and associated transcription factors are also present at the

promoters of expressed genes in MEL and K562 cells. Acetylated histone H3 was also

present at the expressed genes, as has been previously documented [103, 155]. NF-E2

was consistently seen to associate with the promoters of the active genes in the locus.

Having optimized the ChIP assay for use within the mouse and human β-globin locus of

MEL and K562 cells respectively, there are many more experiments that can be carried

out utilizing this assay.

Another group using a MEL cell line that needs DMSO induction for high level β-

globin expression has shown that NF-E2 only binds to the LCR and gene promoter after

DMSO induction [176]. It appears that our MEL cells are quite different since DMSO is

not required for β-globin gene expression or NF-E2 binding as shown in Figure 4-3. It

would be interesting to repeat the ChIP experiment shown in Figure 4-3 with DMSO

induced MEL cells to see if there are any differences in binding between induced and

uninduced cells.

There is currently data available on the binding of the factors Bach1, GATA1,

GATA2, CREB binding protein (CBP), EKLF, Sp1 and YY1 to the β-globin locus. It

may be possible to use the ChIP assay in association with real-time quantitative PCR to

elucidate differences in binding of these factors between the transcribed and non-

transcribed regions of the locus in either MEL and K562 cells that has not been

previously documented.

89

Another interesting project using the ChIP assay would be to analyze specific

histone modifications present within the different regions of the β-globin locus. It has

been shown that the domains of active transcription have hyper-acetylated histones H3

and H4. The availability of quantitative real time PCR and antibodies against specific

histone modifications that have not been analyzed here would allow us to quantitate a

range of histone modifications across the globin loci in both MEL and K562 cells.

Cell Synchronization

In this work, I synchronized cells at the border of G1 and S phase using the double

thymidine block method. The analysis of transcription factor interactions in these

synchronized cells revealed a dynamic pattern of dissociation and re-association of Pol II

and transcription factors that correlated with the timing of DNA replication in the β-

globin loci. It appears that both in K562 and MEL cells, NF-E2 remains bound at the

active genes and LCR element HS2 even when Pol II and the other factors were

dissociated (Figure 4-4). This implicates NF-E2 in playing a role in the recruitment of

Pol II back to the locus after replication is completed. Similarly, USF2 was found to

associate with the transcriptionally active regions of the β-globin locus only when Pol II

has dissociated. Thus, USF2 may also participate in recruiting the transcription

complexes back to the locus after replication.

The cell synchronization experiments were designed according to the hypothesis

that during replication the transcription complexes will be displaced to allow the passage

of the replication fork, and that after replication transcription complexes would re-

associate with the globin locus in an ordered fashion. According to our hypothesis that

the LCR serves as the primary attachment site for transcription complexes in the globin

90

locus (Figure 1-2), we expected that the transcription complexes would first be recruited

to the LCR and subsequently transferred to the active genes. However, our experiments

did not capture time points that allowed this order of recruitment to be revealed. It is

possible that the genes and the LCR cooperate in the recruitment of Pol II to the globin

locus. On the other hand, it is also possible that the processes leading to dissociation and

re-association of Pol II are fast, making it unlikely to select a time point at which Pol II is

detectable at the LCR but not at the genes.

Mitosis is another phase of the cell cycle where transcription factors and RNA Pol

II are known to dissociate and later re-associate with the chromatin [225, 226]; though a

recent study showed that TFIID and TFIIB remain associated with active gene promoters

during mitosis when Pol II is displaced [227]. It would be possible to synchronize MEL

and K562 cells at mitosis, and conduct ChIP experiments at several time points as the

cells synchronously progress through mitosis and into G1. These time points may reveal

an order of recruitment of Pol II back to the β-globin locus.

Chromosome Conformation Capture

The Grosveld group has performed several studies on the conformation of the β-

globin locus using the 3C technique. First, they showed that the LCR and actively

transcribed genes cluster together into what they term the active chromatin hub (ACH)

[120]. Next, they used transgenic mice carrying the wild type or a mutant human β-

globin locus to show that the deletion of HS3 (but not HS2) and the β-globin promoter

together cause the disruption of the ACH [117]. Most recently, they demonstrated that

the transcription factor EKLF is required for ACH formation [162].

91

During our cell synchronization studies, it was observed that Pol II dissociates from

the locus during replication, and later re-associates. Replication occurs very rapidly in

mammalian cells. Therefore, if Pol II is displaced by the replication fork and re-

associates immediately after replication, it would be reasonable to presume that our

experiments would not have been likely to capture this short window of time when Pol II

is absent from the locus. The results from the synchronization experiments suggest that

Pol II does not immediately re-associate with the globin locus after the completion of

replication. A reasonable explanation of why there is a delay is re-association of Poll II

to the locus could be that during replication the ACH is disrupted and that it takes some

time for the ACH to reform. This may be the cause of the window of time when there is

no Pol II present at the locus. Based on this assumption, the 3C technique could be

carried out with cells synchronously progressing through S phase. This would allow us

to analyze if the ACH is disrupted during replication, and later reformed.

In Vitro Pol II Transfer

This novel assay revealed that Pol II transfer from the LCR to the β-globin gene

promoter can be recapitulated and demonstrated that the transfer process is sensitive to

mutations within the β-globin gene promoter and stimulated by NF-E2. As a further

control, the transfer specificity to other mammalian promoters could be tested. This may

reveal critical elements for Pol II transfer. There are other erythroid specific transcription

factors that may also facilitate the transfer of Pol II from the LCR to the genes; e.g.

GATA1 and EKLF. In contrast, Bach1 prevents NF-E2 from interacting with MAREs in

the β-globin locus and could repress the transfer of Pol II to the β-globin gene [128].

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USF, TFII-I and HDAC3 Function in β-Globin Regulation

The ChIP analysis revealed an increased level of USF2 and TFII-I interactions with

the β-globin gene in erythroid cells in which expression of the gene is repressed.

Recently it was shown that TFII-I can interact physically and functionally with HDAC3

[216]. Our studies revealed that HDAC3 interacts with TFII-I in K562 cells, and they co-

occupy the non-expressed β-globin gene.

This has opened the door for functional studies on USF, TFII-I and HDAC3

proteins within the β-globin locus. Another student in the laboratory, Valerie Crusselle,

has embarked on a project to knock down these proteins in erythroid cells and mice using

RNA interference. We have also started projects using inducible dominant negative

mutants of the USF and TFII-I proteins to further investigate their function. In the event

that stable cell lines are created where dominant negative protein expression can be

induced to alter the expression levels of the globin genes, there are many other studies

that can follow.

First, the ChIP assay can be used to investigate how the presence of the dominant

negative proteins affect the binding of other transcription factors to the locus. In addition

to analyzing changes in the presence of the affected proteins between transfected and

control cells, other factors to investigate include CBP and PCAF, which have recently

been shown to interact with USF in the chicken β-globin insulator sequence [228], NF-

E2, TFIIB, and Pol II. Next, the 3C technique can be utilized to reveal any structural

changes or disruption of the ACH resulting from inhibiting these proteins.

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Summary

Taken together, the data in this work provide novel and interesting characteristics

of protein-DNA interactions within β-globin loci of mouse and human erythroid cells.

The data are consistent with a model proposing that RNA Pol II is first recruited to the

LCR and subsequently transferred to the globin gene promoters. The recently described

active hub model may provide a context in which transfer to the globin genes is

facilitated. Future experiments will be directed at testing the Pol II transfer/ACH model

in more detail.

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BIOGRAPHICAL SKETCH

Karen Frances Vieira was born in Bridgetown, Barbados, on October 1st, 1978 to

Ronald F. and Rosemary L. Vieira of St. George Barbados. She lived and attended high

school in Barbados, where she graduated with an Exhibition scholarship from Queen’s

College in 1997. Her undergraduate studies were partially funded by the Government of

Barbados and an academic scholarship from Florida Institute of Technology. Karen was

awarded a B.S. degree in biological sciences with a concentration in molecular biology

from Florida Institute of Technology with high honors in May of 2000. She started in the

IDP in the College of Medicine at the University of Florida in August of 2000, where she

received the Alumni Fellowship. She received many honors, awards and travel grants

during this degree program, including first place in the Medical Guild research

competition. While enrolled in the IDP she concurrently pursued a M.S. in management

from the University of Florida Warrington College of Business, which she finished in

December of 2003. She graduated with her PhD from the Department of Biochemistry

and Molecular Biology in December of 2004. After graduation, Karen has plans of using

her scientific and business educational backgrounds to work within the biotechnology

industry in the short term and to get involved in starting a sickle cell disease Research

Foundation in her island nation, Barbados.

recruitment of transcription complexes to the …

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