THE MOLECULAR BASIS OF FRAGILE SITE FRA16B AND ITS ROLE …

THE MOLECULAR BASIS OF FRAGILE SITE FRA16B AND ITS ROLE IN INV(16)(P13Q22) IN ACUTE MYELOID LEUKEMIA

BY

ALLISON BURROW WECKERLE

A Dissertation Submitted to the Graduate Faculty of

WAKE FOREST UNIVERSITY GRADUATE SCHOOL OF ARTS AND SCIENCES

in Partial Fulfillment of the Requirements

for the Degree of

DOCTOR OF PHILOSOPHY

in the Department of Biochemistry and Molecular Biology

May 2011

Winston-Salem, North Carolina

Approved By:

Yuh-Hwa Wang, Ph.D., Advisor

David Ornelles, Ph.D., Chair

Patrick P. Koty, Ph.D.

Fred W. Perrino, Ph.D.

John C. Wilkinson, Ph.D.

ii

ACKNOWLEDGEMENTS

I would first like to thank my advisor, Dr. Yuh-Hwa Wang, for her guidance and

support throughout my graduate school career. Her door was always open for questions

and encouraged me to continuously put forth my best effort. She spent countless hours

reading and revising abstracts for presentations, research proposals and manuscripts. I

have gained an incredible amount of knowledge and am the scientist I am today because

of her.

I would also like to thank my committee members, Dr. David Ornelles, Dr.

Patrick Koty, Dr. Fred Perrino and Dr. John Wilkinson, for both their scientific

contributions and encouragement during my studies.

Thank you to the current and former members of the Wang laboratory for their

support and helpful discussions throughout the years. I would like to especially thank

Lindsay Holder for teaching me everything from running a gel to culturing cells , and

Laura Dillon, who has become a close friend, for enlightening conversations regarding

science and everything else.

Finally, I want to thank my family. I certainly would not be where I am at today,

had it not been for their constant love and dedication. My parents, Greg and Judy Burrow,

deserve special mention. Since I was a child, they attended every single dance

competition, football and basketball game, and encouraged me to do anything and

everything I dreamt of. I want to thank my grandmother, Barbara Hyatt, for teaching me

to appreciate all the blessings I‟ve been given. I also want to thank my wonderful

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husband, Steve, for giving me so much to look forward to every single day, and who

never ceases to make me smile.

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TABLE OF CONTENTS

Page LIST OF ABBREVIATIONS……………………………………………………………..v LIST OF FIGURES…………………………………………………………………...….xi

LIST OF TABLES……………………………………………………………………...xiii ABSTRACT…………………………………………………………………………….xiv

Chapter I. INTRODUCTION…………………………………………………………………..1

II. OVER HALF OF BREAKPOINTS IN GENE PAIRS INVOLVED IN CANCER-SPECIFIC RECURRENT TRANSLOCATIONS ARE MAPPED TO HUMAN CHROMOSOMAL FRAGILE SITES…………………………..….19

Published in BMC Genomics, January 2009 III. SECONDARY STRUCTURE FORMATION AND DNA INSTABILITY AT FRAGILE SITE FRA16B………………………..…….40

Published in Nucleic Acids Research, January 2010 IV. CBFB AND MYH11 IN INV(16)(P13Q22) OF ACUTE MYELOID LEUKEMIA DISPLAY CLOSE SPATIAL

PROXIMITY IN INTERPHASE NUCLEI OF HUMAN HEMATOPOIETIC STEM CELLS…………………………………………..…...71 In press in Genes, Chromosomes and Cancer, May 2011

V. THE INDUCTION OF FRAGILE SITE FRA16B PRODUCES DNA BREAKS WITHIN CBFB OF INV(16)(P13Q22)…………………………………...……….94

VI. CONCLUSIONS……………………………………………………...………….105

APPENDIX……………………………………………………………………………..127 SCHOLASTIC VITA…………………………………………………………...……...146

v

LIST OF ABBREVIATIONS

2-AP 2-aminopurine

5-aza 5-azacytidine

ABL Abelson murine leukemia viral oncogene

AFF4 AF4/FMR2 family, member 4

ALCL Anaplastic large cell lymphoma

AML Acute myeloid leukemia

AMP Abnormal myeloid progenitor

APH Aphidicolin

ATM Ataxia telangiectasia mutated

ATP Adenosine triphosphate

ATR Ataxia telangiectasia and Rad3-related

ATRIP ATR interacting protein

BAC Bacterial artificial chromosome

BCR Breakpoint cluster region

bp Base pair

BRCA1 Breast cancer 1

BrdU Bromodeoxyuridine

CBF Core binding factor

CBFA Core binding factor, alpha subunit (gene)

CBFB Core binding factor, beta subunit (gene)

CBFα Core binding factor alpha (protein)

vi

CBFβ Core binding factor beta (protein)

CCDC6 Coiled-coil domain containing 6

CFS Common fragile sites

CHK1 Checkpoint kinase 1

CHK2 Checkpoint kinase 2

CHORI Children's Hospital Oakland Research Institute

CMMoL Chronic myelomonocytic leukemia

CMP Common myeloid progenitor

dATP Deoxyadenosine triphosphate

dCTP Deoxycytidine triphosphate

dGTP Deoxyguanosine triphosphate

DNA Deoxyribonucleic acid

DNA-PKcs DNA-dependent protein kinase, catalytic subunit

DSB Double-strand break

DTT Dithiothreitol

dTTP Deoxythymidine triphosphate

dUTP Deoxyuridine triphosphate

EM Electron microscopy

EWSR1 Ewing sarcoma breakpoint region 1

FAB French-American-British

FANCD2 Fanconi anemia, complementation group D2

FHIT Fragile histidine triad gene

FISH Fluorescence in situ hybridization

vii

FLT3 FMS-related tyrosine kinase 3

FUdR Fluorodeoxyuridine

G-banding Giemsa banding

GDB Human Genome Database

GMP Granulocyte-monocyte progenitor

h Hour

H2AX H2A histone family, member X

HMGA1 High mobility group AT-hook 1

HPGM Human progenitor growth medium

HR Homologous recombination

HSC Hematopoietic stem cell

IGH Immunoglobulin heavy locus

IGK Immunoglobulin kappa locus

IGL Immunoglobulin lambda locus

IL-3 Interleukin 3

IL-6 Interleukin 6

kb Kilobase

kDa Kilodalton

KIT v-kit Hardy-Zuckerman 4 feline sarcoma viral oncogene homolog

KRAS v-Ki-ras2 Kirsten rat sarcoma viral oncogene homolog

L Liter

LAMA4 Laminin, alpha 4

LM-PCR Ligation-mediated PCR

viii

LSC Leukemic stem cell

M Molar

Mb Megabase

MEP Megakaryocyte-erythroid progenitor

MET met proto-oncogene (hepatocyte growth factor receptor)

mg Milligram

min Minute

MLL Myeloid/lymphoid or mixed-lineage leukemia

mm Millimeter

mM Millimolar

MSC Mesenchymal stem cell

mSin3A SIN3 homolog A, transcriptional regulator (yeast)

MYC v-myc myelocytomatosis viral oncogene homolog (avian)

MYH11 Myosin, heavy chain 11, smooth muscle

NCOA4 Nuclear receptor coactivator 4

ng Nanogram

NHEJ Non-homologous end joining

NHL Non-Hodgkin's lymphoma

NIH National Institutes of Health

nm Nanometer

NRAS Neuroblastoma RAS oncogene

nt Nucleotide

NTRK1 Neurotropic tyrosine kinase, receptor, type 1

ix

NUP98 Nucleoporin 98 kDa

OIZ Okazaki initiation zone

ori Origin of replication

PAGE Polyacrylamide gel electrophoresis

PBL Peripheral blood lymphocyte

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PSC Pre-leukemia stem cell

PTC Papillary thyroid carcinoma

RAD51 RecA homolog, E. coli

RAG Recombination activating gene

RET Rearranged during transfection proto-oncogene

rh Recombinant human

ROS Reactive oxygen species

RPA Replication protein A

SCE Sister chromatid exchange

SD Standard deviation

Ser Serine

SMC1 Structural maintenance of chromosomes 1

SMMHC Smooth muscle myosin heavy chain

SSA Single-strand annealing

SSC Sodium citrate saline

ssDNA Single-stranded DNA

x

STRIP Stability of trinucleotide repeat by individual product

SV40 Simian virus 40

TBE Tris/Borate/EDTA

TE Tris/EDTA

TICdb Translocation breakpoints In Cancer database

TopBP1 Topoisomerase II binding protein 1

TPR Translocated promoter region

U Unit

Ub Ubiquitination

UCSC University of California, Santa Cruz

UNC-CH University of North Carolina at Chapel Hill

UV Ultraviolet

V Volts

WFUBMC Wake Forest University Baptist Medical Center

WRN Werner syndrome, RecQ helicase-like

WWOX WW domain containing oxidoreductase

µg Microgram

μL Microliter

µm Micrometer

μΜ Micromolar

xi

LIST OF FIGURES

Page 1.1: Proposed model of CBFβ-SMMHC-associated leukemia progression…………. …17 2.1: DNA flexibility analysis of translocation-prone and fragile site co-localized

genes…………………………………………………………………………..………… 30 2.2: Secondary structure analysis of CBFB, MYH11, HMGA1, LAMA4, MLL, and AFF4 loci…………………………………...…………………………………………………...32

2.3: The computed lowest free energy of predicted DNA secondary structures……….. 34 3.1: Secondary structure formation of FRA16B DNA following denaturation and re-

annealing (reduplexing) reaction…………………………………………………….….. 53 3.2: Each strand of FRA16B forms a secondary structure with different electrophoretic mobilities…………………………………………….………………………………. ….54

3.3: Analysis of FRA16B replication efficiency and instability in human cells using an SV40 replication system…………………….…………………………………………. .55

3.4: Mapping mutation sites to hairpin regions of predicted DNA secondary structure.. 58 3.5: FRA16B DNA-containing sequence of pFRA16B37…………………………….. .59

3.6: FRA16B DNA synthesis by Klenow fragment of E. coli DNA polymerase I…….. 60 3.7: Proposed models of FRA16B instability…………………………………...…… …69

4.1: Interphase FISH analysis of CBFB and MYH11…………...……..………...………82 4.2: CBFB-MYH11 interphase distance distribution……………………….………. ..….83

4.3: Distribution patterns of CBFB-MYH11 distance intervals……………………..… ..85 4.4: Comparison of interphase distance between CBFB and MYH11 with the distance between CBFB and 16p11.2 in HSCs…………………..………………………………..87

4.5: Induction of fragile site breakage in treated HSCs……………………………… …88

4.6: Distribution of CBFB-MYH11 interphase distances as a percentage of nuclear

diameter……………………………………………………………………...…….. ……90 5.1: DNA breaksite mapping by LM-PCR…………………………………………...… 99

xii

5.2: Detection of DNA breaks by LM-PCR following treatment of HSCs with fragile site-inducing chemicals……………………………………………………………….. 100

5.3: Location of induced breakpoints within intron 5 of CBFB……………………. …102 6.1: Model of FRA16B instability in the formation of inv(16)(p13q22) in acute myeloid

leukemia……………………..…………………………………………………………. 110

xiii

LIST OF TABLES

Page 1.1: Classification of fragile s ites…………………..……………………...……………...3 1.2: DNA damage checkpoint proteins shown to regulate fragile site stability.…….……9

1.3: Environmental, dietary and medicinal inducers/enhancers of fragile sites……… ...13 2.1: Translocation breakpoints mapped to fragile sites in both partner genes of cancer-

specific recurrent translocations…………………………. ……….……………………..27 2.2: Computational analysis of genes involved in cancer-specific recurrent translocations reveals characteristics of chromosomal fragile sites………………………...……….. …31

4.1: Analysis of measured CBFB-MYH11 interphase distances…..………………… ….84 4.2: Frequency of CBFB and MYH11 co-localization………………………….. ………86

5.1: Frequency of DNA breaks detected in HSCs by LM-PCR…………. ……………101

Appendix Table 1: Comprehensive list of gene pairs involved in cancer-specific

recurrent translocations which result in fusion transcripts……….…………. …………127 Appendix Table 2: Gene pairs involved in cancer-specific recurrent translocations in which the breakpoint in one gene co-localizes with a fragile site…………..…………. 141

xiv

ABSTRACT

Weckerle, Allison Burrow

THE MOLECULAR BASIS OF FRAGILE SITE FRA16B AND ITS ROLE IN

INV(16)(P13Q22) IN ACUTE MYELOID LEUKEMIA

Dissertation under the direction of Yuh-Hwa Wang, Ph.D., Associate Professor

Fragile sites are regions of the human genome especially susceptible to DNA

breakage and have been suggested to play a role in cancer development. Several studies

have determined that fragile site locations correspond to DNA breakpoints in a number of

chromosomal rearrangements observed in tumor cells, but there has been no

comprehensive examination of the location of fragile sites relative to breakpoints in all

known cancer-specific abnormalities, or direct evidence of a role for fragile sites in

tumorigenesis. Furthermore, the mechanism of fragile site breakage remains unclear,

making it difficult to identify potential risk factors or determine the involvement of

fragile site breakage in chromosomal rearrangements. With an increasing number of

fragile sites being mapped and cancer cases on the rise, it is becoming even more

important to identify a role for these highly breakable regions in cancer development and

the molecular basis of their breakage. The purpose of the studies in this work was to

investigate the involvement of fragile sites in the formation of cancer-specific

chromosomal rearrangements and the underlying mechanism of fragile site breakage.

To demonstrate a role for fragile sites in tumorigenesis, a comprehensive

examination of fragile site locations relative to breakpoints in all reported chromosomal

aberrations in cancer was performed. Of 444 unique gene pairs participating in cancer-

xv

specific recurrent translocations, it was determined that over half (52%) of the

breakpoints map to fragile sites. Furthermore, examination of the DNA sequences

revealed that translocation-participating loci exhibit characteristics of fragile sites,

including high DNA flexibility and the ability to form stable secondary structures,

supporting their propensity for breakage and subsequent rearrangements.

While data support a causative role for fragile sites in cancer development, little is

known about the molecular basis of their fragility, or the mechanism by which breakage

at these regions ultimately leads to the formation of cancer-specific chromosomal

rearrangements. Although a consensus sequence has not yet been identified among fragile

sites, results from numerous studies suggest it is likely that the capability of fragile DNAs

to form stable secondary structures may directly contribute to their breakage by inhibiting

replication. Until now, there has been no physical evidence of an AT-rich fragile site,

which comprises the majority of fragile sites, forming an alternative structure in vitro.

Here, a DNA fragment of fragile site FRA16B, which served as a model for fragile site

investigation, was subjected to reduplexing, which produced bands with reduced

electrophoretic mobility in polyacrylamide gels. Additionally, visualization of the DNAs

by electron microscopy (EM) revealed the formation of short, branched structures. To

determine whether the formation of alternative structures affects replication of fragile

DNA, plasmids containing FRA16B were transfected into human cells and examined for

the presence of instability events and the efficiency of replication. Some FRA16B-

containing constructs displayed a significant increase in the number of mutation events

and/or a reduction in replication efficiency compared to a non-fragile DNA control,

xvi

depending on both replication orientation and distance from FRA16B to the origin.

Furthermore, FRA16B replication fork templates were constructed and visualized by EM

to determine the mechanism of instability at fragile sites. The examination showed that

the majority of constructs contained DNA polymerase paused within FRA16B, which

was confirmed by DNA sequencing gels, and among the molecules which completed

DNA synthesis, most underwent some degree of spontaneous fork reversal.

The results described above suggest that chromosomal instability at fragile sites

caused by the formation of secondary structures may ultimately lead to the formation of

chromosomal abnormalities observed in cancer cells, although a direct role has not yet

been proven. To investigate a direct role for fragile sites in the generation of cancer-

causing rearrangements, and to gain additional insight into the mechanism of

chromosomal translocations, the inv(16)(p13q22), one of the most common abnormalities

observed in acute myeloid leukemia (AML), was examined since both participating genes

co-localize with fragile sites. First, the spatial proximity between genes CBFB and

MYH11 was examined due to recent evidence supporting a role for spatial genome

organization in the formation of chromosomal translocations. The distance between

CBFB and MYH11 in interphase nuclei was measured in human hematopoietic stem cells

(HSCs), and compared to mesenchymal stem cells (MSCs), peripheral blood lymphocytes

(PBLs) and fibroblasts. The distance between CBFB and MYH11 was significantly

reduced in HSCs compared to all other cell types, further supporting a role for spatial

gene proximity in the generation of chromosomal translocations , and providing a

potential mechanism for inv(16)(p13q22). To determine whether breakage at fragile sites

xvii

is directly involved in the formation of inv(16)(p13q22), ligation-mediated PCR (LM-

PCR) was used to detect DNA breaks within the CBFB gene, one of the most common

targets of genetic alterations in AML, which co-localizes with FRA16B. Upon treatment

of HSCs with fragile site-inducing chemicals, DNA breaks were observed within the

major breakpoint region of CBFB with a frequency that was significantly higher than that

in untreated cells.

Overall, these data strongly support a role for fragile sites in the formation of

cancer-causing chromosomal rearrangements by demonstrating a significant association

between fragile site locations and translocation breakpoints in a variety of cancers.

Furthermore, the presence of DNA breaks within CBFB upon treatment with fragile site-

inducing chemicals suggests that fragile site breakage is directly involved in the

formation of inv(16)(p13q22), providing additional evidence of a role for fragile sites in

cancer development. Additionally, the ability of FRA16B to form a secondary structure

in vitro, which affects its replication efficiency and instability in human cells , supports

the proposed mechanism of fragile site breakage whereby the formation of secondary

structures presents significant difficulties during replication, leading to regions of the

genome highly susceptible to DNA breakage and subsequent rearrangements.

1

CHAPTER I: INTRODUCTION

Fragile sites are regions of the human genome especially susceptible to DNA

breakage and have been suggested to play a causative role in tumorigenesis through the

formation of structural aberrations observed in various types of tumors. There are

currently 121 fragile sites located throughout the genome. Fragile sites are present in all

individuals and can be induced by a variety of environmental and chemical agents [1].

Variability of fragile site breakage has been observed among individuals [2], which may

reflect exposure to such elements, with high levels being associated with cancer [3].

While these data strongly support a role for fragile sites in tumorigenesis, there has been

little direct evidence of their involvement in cancer development. This association has

also prompted further investigation into the molecular basis of fragile site expression,

since this process remains unclear. Therefore, it is important to elucidate the mechanism

of fragile site breakage, as well as consequences of their breakage to identify potential

risk factors in order to reduce or prevent the occurrence of cancer development in

humans.

The goals of this work were to gain a better undersanding of the molecular basis

of fragile site breakage, and to investigate the involvement of fragile sites in the

generation of structural rearrangements observed in cancer. These goals were addressed

through an investigation into the mechanism of breakage at fragile site FRA16B, a

comprehensive analysis of fragile site locations relative to breakpoints in all known

cancer-specific translocations, and examination of a direct role for FRA16B in

inv(16)(p13q22) in AML.

2

Human Chromosomal Fragile Sites

Fragile sites are defined cytogenetically as non-random chromosomal loci that exhibit

gaps or breaks on metaphase chromosomes following conditions of partial replication

stress [1]. Fragile sites are normally stable in cultured cells. However, these regions are

hotspots for sister chromatid exchange (SCE), deletions, and rearrangements following

induction with replication inhibitors [4, 5]. They are classified as either common or rare,

depending on their frequency in the population, and are subdivided according to their

mode of induction in cultured cells (Table 1.1). Common fragile sites (CFS) have been

observed in all individuals and are therefore believed to represent a normal component of

chromosome structure [6]. To date, over 80 CFS have been reported. The majority of

CFS are induced by low doses of aphidicolin (APH), an inhibitor of DNA polymerases α,

δ and ε [7, 8]. Other CFS are observed following treatment with bromodeoxyuridine

(BrdU) or 5-azacytidine (5-aza). Most have not yet been investigated at the molecular

level, but it is known that regions of fragility can extend over megabases (Mb) of DNA,

with gaps or breaks occurring throughout [9]. In contrast, rare fragile sites are found in

less than 5% of the population, and are inherited in a Mendelian manner [10, 11]. Most

rare sites are expressed under folate-deficient conditions. Others are induced following

treatment with minor groove binders, such as distamycin A or berenil, as well as BrdU.

However, Mrasek et al. recently discovered that APH can induce all types of common

and rare fragile sites, suggesting that their expression is less dependent on the ir currently

defined mode of induction, and instead, a classification of fragile sites based on their

frequency is more appropriate [12]. The expression of rare fragile sites results from the

unstable expansion of a repetitive element [11]. Folate-sensitive sites contain an

3

expanded (CGG)n repeat [13], whereas other rare sites are comprised of AT-rich

minisatellite elements [11]. While a consensus sequence has not yet been identified

among CFS, the DNAs examined thus far contain frequent, AT-rich flexibility islands,

and are predicted to form highly stable secondary structures compared to non-fragile

DNA [14, 15], similar to what has been reported for most rare fragile sites.

Fragile Site FRA16B

Fragile site FRA16B is a rare site located at the chromosomal locus 16q22.1.

Spontaneous FRA16B expression has been observed among individuals, and can be

induced by treatment with distamycin A [16] or berenil [17]. Studies have determined

that FRA16B spans the same genomic region as common fragile site FRA16C, and is

also expressed following treatment with APH [12, 14]. Following induction, the

heterozygote frequency of FRA16B is about 5% in populations of European descent,

representing the most frequently expressed rare fragile site [18]. Positional cloning has

revealed that FRA16B-expressing chromosomes may contain up to 2000 copies of the 33

base pair (bp) AT-rich minisatellite repeat (ATATATTATATATTATATCTAATAATA-

TATC/ATA), whereas normal chromosomes consist of only 7-12 copies of the repetitive

element [19].

Table 1.1: Classification of fragile sites

(i) Common: APH-inducible (n=79) BrdU-inducible (n=7) 5-aza-inducible (n=4)

(ii) Rare: Folate-sensitive (n=24) Distamycin A/Berenil-inducible (n=5)

BrdU-inducible (n=2)

4

Although breakage at FRA16B has not been directly linked to any human

diseases, results suggest that the dominant inheritance of cleft palate, microstomia and

micrognathia may be linked to FRA16B [20]. FRA16B breakage has also been detected

in PBLs of healthy individuals previously treated for non-Hodgkin‟s lymphoma (NHL),

and in both PBLs and bone marrow cells of patients with chronic myelomonocytic

leukemia (CMMoL) [21]. Furthermore, breakpoints of chromosomal rearrangements

observed in tumors coincide with the same locus as FRA16B [22]. Therefore, FRA16B

serves as a good model for the examination of fragile sites since it exhibits characteristics

of both major classes of fragile sites, is associated with human disorders, and co-localizes

with sites of breakage in cancer-specific chromosomal rearrangements.

DNA Replication at Fragile Sites

Understanding the molecular basis of fragile site breakage is critical for dissecting the

role of fragile sites in cancer. Although fragile sites do not contain a consensus sequence,

reports suggest several intrinsic factors which may contribute to their expression. First,

replication timing studies have shown that all fragile sites examined to date, including

FRA1H [23], FRA2G [23], FRA3B [24], FRA6E [25], FRA7H [26], FRA10B [27],

FRA16B [27], and FRAXA [28] are late-replicating regions of the genome. The delay

can be further exacerbated with the addition of replication inhibitors, with some fragile

site alleles remaining unreplicated in late G2 phase [24, 26].

Additionally, most fragile DNAs studied to date are predicted to form highly

stable secondary structures [14, 15] which likely contribute to their breakage by

5

inhibiting replication. Evidence for secondary structure formation at fragile sites has

largely been generated by the Mfold program [29], which predicts secondary structure

formation of a single-stranded DNA (ssDNA). The only physical evidence of a fragile

DNA forming a secondary structure in vitro comes from the ability of the (CGG)n repeat,

which underlies the basis of fragility at rare, folate-sensitive fragile sites, to form

quadruplex [30] and hairpin structures [31] that present significant blocks to replication

both in vitro [32] and in vivo [33]. Replication of fragile DNA has largely been

investigated using primer extension assays and analyzing synthesis intermediates isolated

for two-dimensional gel electrophoresis. The examination of replication intermediates

from cells containing AT-rich sequences within common fragile site FRA16D in

Saccharomyces cerevisiae showed site-specific replication fork stalling depending on the

length of the AT repeat [34]. Correlation with secondary structure predictions suggests

that the structure formed by the repeat is directly responsible for the fork stalling.

Synthesis of the same fragile site by human replicative polymerases δ and α using an in

vitro primer extension assay confirmed polymerase stalling at sites predicted to form

inhibitory DNA structures [35]. Furthermore, the same study examined primer extension

using HeLa cell-free extracts, and obtained similar results. Interestingly, FRA16D

regions with increased DNA flexibility and accompanying high A/T content were not

sufficient to inhibit DNA synthesis, but sequences with the propensity to form secondary

structures were significantly inhibitory to replication. These data suggest that the ability

of fragile sites to form stable secondary structures during replication may be directly

responsible for the delayed replication observed at these regions by inhibiting DNA

synthesis.

6

In a recent study, Letessier et al. demonstrated that the initiation of replication at

fragile sites may also be perturbed. The examination of common fragile site FRA3B in

lymphoblastoid cells revealed that initiation events are excluded from a 700 kilobase (kb)

core region, forcing the replication forks coming from flanking regions to cover great

distances so that replication is completed [36]. They also showed that origins of the

flanking regions fire in mid-S phase, leaving the site incompletely replicated upon

slowing of the replication fork. These data were the first to suggest that CFS are

initiation-poor regions, rendering replication difficult to complete. However, it will be

important for future studies to determine whether other CFS also demonstrate poor

initiation, and if the same is true for rare fragile sites.

The current working model for fragile site expression is as follows: under

conditions of replication stress, the replicative DNA polymerases may uncouple from the

helicase/topoisomerase complex, resulting in long stretches of ssDNA with the ability to

form highly stable secondary structures. Consequently, the formation of such structures

may inhibit replication fork progression, triggering the ataxia telangiectasia and Rad3-

related (ATR) DNA damage checkpoint pathway, which has proven to be critical for the

maintenance of fragile site stability [37]. The gaps and breaks on metaphase

chromosomes are therefore believed to represent unreplicated regions that have escaped

the ATR replication checkpoint [37]. Additional studies will be needed to fully

understand DNA replication at fragile sites, which will give greater insight into the

molecular basis of fragile site breakage, and ultimately, their role in cancer development.

7

Maintenance and Repair of DNA Breaks at Fragile Sites

Although the exact mechanisms of fragile site maintenance and repair are not fully

understood, numerous studies support an important role for the ATR DNA damage

checkpoint pathway in the stability of fragile sites. ATR kinase is a DNA damage sensor

protein that works with downstream target proteins to respond to stalled and collapsed

replication forks, resulting in a block in further replication and mitosis progression and

the promotion of DNA repair, recombination, or apoptosis [38, 39]. The loss of

functional ATR in cells results in a defective DNA damage response to agents which

block replication fork progression, including APH and hydroxyurea [37, 40, 41], and

conditions of hypoxia [42]. Casper et al. found that cells deficient in ATR, but not ataxia

telangiectasia mutated (ATM), the main transducer of the double-strand break (DSB)

pathway, display up to a 20-fold increase in fragile site breakage following treatment

with low doses of APH compared to control cells [37]. Also, a deficiency in ATR alone is

enough to induce fragile site breakage in cells without treatment with replication

inhibitors. Supporting these results, cells from patients with Seckel Syndrome, who

express low levels of ATR protein due to a hypomorphic mutation in the ATR gene,

exhibit an increase in chromosomal breakage at CFS compared to unaffected individuals

[43]. Furthermore, mice hypomorphic for ATR also display an increase in CFS breakage

and a significant delay in checkpoint induction [44]. While the loss of ATM alone does

not cause increased CFS breakage [37], it is involved in maintaining fragile site stability

in the absence of ATR. Ozeri-Galai et al. found that a loss of both ATR and ATM

significantly increases APH-induced CFS breakage compared to the loss of ATR alone

[45]. Also, ATM is activated and forms nuclear foci with γH2AX following treatment

8

with low doses of APH [45]. These findings indicate that ATR is the major pathway

responsible for maintaining fragile site stability, but that ATM also plays a secondary

role, perhaps through a downstream response to DSBs that form as a result of ATR

deficiency.

Other downstream targets of the ATR-mediated pathway involved in maintaining

fragile site stability include BRCA1 [46] and CHK1 [47] (Table 1.2). BRCA1 is a

primary target of both ATR and ATM phosphorylation in response to DNA damage.

Cells lacking BRCA1 show significantly more fragile site expression after treatment with

APH compared to control cells [46]. Also, cells expressing mutant BRCA1 exhibit

elevated levels of fragile site breakage but lack the G2/M checkpoint, suggesting that

BRCA1 regulates fragile site stability through its role at this checkpoint. CHK1 kinase is

the major downstream target of ATR and serves as the central regulator of the ATR

checkpoint pathway. In cells, the loss of CHK1, but not the ATM-regulated CHK2,

results in a significant increase in fragile site breakage after treatment with APH [47].

Additionally, it was found that both ATR and ATM phosphorylate CHK1 following

treatment with low doses of APH [45]. These data suggest that the role of ATM in fragile

site maintenance may be to activate the ATR pathway through phosphorylation of CHK1

when ATR is missing or fails to properly respond to damage.

While the importance of the ATR pathway in fragile site maintenance has been

established, the mechanism is not fully understood. Recently, Wan et al. found that ATR

binds (directly or through complexes) to fragile site FRA3B preferentially compared to

non-fragile regions under conditions of mild replication stress [48]. This binding

9

Table 1.2: DNA damage checkpoint proteins shown to regulate fragile site stability

Protein Function Reference

ATM Kinase, maintains fragile site stability in the absense of ATR [45]

ATR Kinase, binds to fragile DNA in response to replication stress,

phosphorylates downstream targets to activate check-point response [37, 48]

BRCA1 Phosphorylated by ATR, major downstream target of ATR, necessary

for G2/M checkpoint activation following replication stress [46]

CHK1 Kinase, phosphorylated by ATR in response to replication stress, central

regulator of ATR pathway [47]

Claspin Phosphorylated and interacts with CHK1 in response to replication

stress [49]

FANCD2 Fanconi Anemia pathway protein, phosphorylated by ATR leading to

activation by mono-Ub, activated by replication stress [50]

HUS1 Member of the 9-1-1 complex, promotes phosphorylation of ATR

substrates [51]

SMC1 Chromosomal structural maintenance protein, member of the cohesion

complex [52]

WRN ATP-dependent 3‟-5‟ helicase, 3‟-5‟ exonuclease [35, 53]

increases in a dose-dependent manner, peaking at 0.4 μM APH, and decreases at higher

APH concentrations. While the level of ATR binding to FRA3B changes with treatment,

the cellular levels of ATR, phospho-ATR (Ser 428), and ATR-interacting proteins

ATRIP and TopBP1 remain unchanged. This suggests that ATR binding to the fragile site

is guided initially by the level of replication stress signals generated at FRA3B due to

APH treatment, and then sequestered from FRA3B regions by successive signals from

other non-fragile site regions, which are produced at the higher concentrations of APH.

Furthermore, the kinase activity of ATR was required for ATR binding to FRA3B in

response to APH treatment. While ATR kinase activity is known to be necessary for

phosphorylation of downstream targets to activate the checkpoint signalling cascade [39],

these data indicate that the kinase activity of ATR is also necessary for ATR interaction

10

to fragile site regions, most likely through phosphorylation of ATRIP and TopBP1 to

stabilize the interaction between these three proteins and the fragile DNA. Two models

which are not mutually exclusive have been proposed to explain how ATR helps to

maintain fragile site stability [54]. The first model states that a loss of ATR can lead to a

bypass of stalled replication forks at fragile sites, ultimately resulting in a failure of

checkpoint pathways to prevent entry into mitosis , thus leaving DNA breakage at the

unreplicated DNA. The second model states that a loss of ATR leads to replication fork

collapse at fragile sites and improper resolution of these structures by ATR leads to DNA

breaks. The current information about the involvement of ATR at fragile sites supports a

combination of both models. The preferential binding of ATR protein to FRA3B fragile

DNA following APH treatment [48] suggests that ATR plays a possible local role in

stabilizing stalled replication forks at fragile regions. Also, this binding and increased

fragile site breakage following the inhibition of various members of the ATR pathway

suggest that the ATR response to fragile sites under conditions of replication stress can

activate the ATR-dependent pathway. Finally, decreased ATR binding to FRA3B at

higher concentrations of APH [48], which induce more chromosomal gaps or genomic

breaks, supports the idea that DNA breakage at fragile sites is due to a failure of ATR to

stabilize replication forks and to signal a checkpoint response.

Since ATM regulates fragile site stability in the absence of ATR [45], and these

regions often co-localize with sites of chromosomal breaks in tumor cells [55], it is

apparent that DSBs also form at fragile sites, although the exact mechanism remains

unknown. DNA breakage is most often repaired by either the homologous recombination

11

(HR) or non-homologous end joining (NHEJ) DSB pathways [56]. However, dysfunction

of these repair pathways can lead to the formation of chromosomal rearrangements [57].

Schwartz et al. found that down-regulation of RAD51, DNA-PKcs, and Ligase IV, key

components of the HR and NHEJ repair pathways, significantly increases fragile site

breakage with APH treatment, and that γH2AX and phosphorylated DNA-PKcs foci were

located at expressed fragile sites [58]. Together, these data suggest a role for both the HR

and NHEJ repair pathways in the repair of fragile site breakage. Additionally, the

majority of breakpoints in a papillary thyroid carcinoma (PTC) rearrangement mapped to

fragile sites occur at microhomology patches, indicating that fragile site-associated

rearrangements may also arise by microhomology-mediated single-strand annealing

(SSA) [59]. The repair of lesions at fragile sites is still not clear, but evidence suggests

that all three pathways may be involved. It will be essential for future studies to identify

the direct roles of factors required for fragile site stability, as well as the process by

which DSBs ultimately arise at these regions to gain a better understanding of the

maintenance and repair of fragile sites.

Importance of Fragile Sites in Cancer

Genomic instability in the form of chromosomal abnormalities is a hallmark of tumor

cells. Rearrangements causing the deletion, insertion or translocation of genetic material

often contribute to the neoplastic process through the disruption of genes involved in

processes such as cell proliferation and/or survival, the expression of altered gene

products with oncogenic potential, or the loss of tumor suppressive properties. While the

molecular basis of chromosomal rearrangements remains unclear, it is apparent that DNA

12

strand breakage is an initiating event. Cells can acquire DNA damage through various

extrinsic or intrinsic factors. This damage commonly arises following exposure to

exogenous factors such as UV radiation, ionizing radiation [60], chemotherapy, or

endogenous factors like reactive oxygen species (ROS) [61]. Since fragile sites are highly

prone to DNA strand breakage, it has been suggested that these regions may also play a

role in cancer development.

Fragile sites span large genomic regions and often encompass genes that function

as tumor suppressors or oncogenes. The deletion of tumor suppressor genes and the

amplification of oncogenes are frequently consequences of breakage at these sites. The

two most commonly expressed fragile sites, FRA3B and FRA16D, lie within the tumor

suppressor genes FHIT and WWOX, respectively. The FHIT gene is often involved in

deletions which map specifically to the fragile site region in various types of cancer [62-

67], and deletion of WWOX is frequently observed in tumor cells [68-73]. It has been

proposed that fragile sites may also play a role in the breakage-fusion-bridge model of

gene amplification [74, 75], since proto-oncogenes such as MYC [76] and MET [77] are

located at fragile sites FRA8C and FRA7G, respectively. Additionally, a number of

studies have demonstrated a significant association between sites of breakage in cancer-

specific chromosomal rearrangements and the location of fragile sites [78-80].

Interestingly, the majority of cancers associated with fragile site breakage have

little or no genetic component, suggesting that fragile sites play a role in sporadic tumor

formation. Based on this hypothesis, several studies have examined lymphocytes of

individuals from a broad spectrum of chemical exposure histories in addition to fragile

13

site induction following treatment with various environmental and chemical factors in

cultured cells. Results support a model for fragile sites in the generation of sporadic

cancers by demonstrating the ability of fragile sites to be induced by a variety of

environmental and chemical agents, including caffeine [79, 81], ethanol [82, 83],

cigarette smoke [84, 85], and pesticides [86-88] (Table 1.3).

Table 1.3: Environmental, dietary and medicinal inducers/enhancers of fragile sites

Chemical/Condition Uses Reference

5-aza chemotherapeutic agent [89]

actinomycin D chemotherapeutic agent [89]

atenolol hypertension drug [90]

benzene found in cigarette smoke, gasoline fumes [89]

bleomycin chemotherapeutic agent [89]

busulfan chemotherapeutic agent [89]

caffeine dietary agent [79, 81]

carbon tetrachloride found in refrigerants, pesticides [89]

chlorambucil chemotherapeutic agent [89]

cigarette smoke dietary and environmental agent [84, 85]

cytosine arabinoside chemotherapeutic agent [89]

diethylnitrosamine found in cigarette smoke, pesticides, cured meat, whiskey [89]

dimethyl sulfate found in dyes, drugs, perfumes, pesticides [89]

ethanol dietary agent [82, 83]

FUdR chemotherapeutic agent [89]

hypoxia low oxygen; found in tumor microenvironment [91]

methotrexate chemotherapeutic agent [89]

pesticides environmental agent [86-88]

Caffeine, an inhibitor of phosphoinositide 3-kinase related kinases, including

ATR and ATM, significantly increases fragile site breakage both alone [81], and in

14

conjunction with fluorodeoxyuridine (FUdR) and APH [79]. A combination of ethanol

and APH also significantly increases fragile site breakage [83], and interestingly, cells

from chronic alcohol users show a significantly higher frequency of fragile site and

chromosomal breakage compared to control individuals, suggesting long term alcohol use

alone can induce fragile site expression [82]. Additionally, Kao-Shan et al. found that

PBLs from cigarette smokers show significantly greater fragile site breakage compared to

non-smokers [85], and Stein et al. found that treatment of PBLs with low-doses of APH

results in increased fragile site breakage in active smokers compared to non-smokers and

patients with small cell lung cancer who stopped smoking [84]. These results suggest that

active exposure to cigarette smoke increases the potential of breakage at fragile sites, and

that this risk is reversible. Exposure to pesticides also results in an increased

susceptibility to fragile site breakage. Blood lymphocytes from pesticides sprayers and

flower collectors working in greenhouses show greater fragile site breakage than normal

individuals following treatment with APH, with these results being reproducible a year

later [86, 87]. An association between pesticide exposure and an increased risk of

hematopoietic tumors has also been observed [92, 93]. The APH-induced damage was

enhanced at fragile sites containing breakpoints involved in leukemias and NHL,

supporting a role for pesticide-associated fragile site breakage in the development of

these cancers. Like cigarette smoke exposure, the effect of pesticide exposure on fragile

site breakage is also transient. In addition to environmental and dietary mutagens, several

oncogenic viruses including human papilloma virus [94], Hepatitis B [95], and Epstein-

Barr [96] have also been shown to target and preferentially integrate at fragile sites [97],

although the underlying basis remains unclear.

15

Together, these data strongly support a causative role for fragile sites in

tumorigenesis, whereby exposure to various dietary, environmental and/or chemical

agents may lead to cancer development through chromosomal breakage at these regions.

For future studies, it will be important to understand the mechanism by which these

agents affect fragile site expression, and to identify any additional factors that could

induce or enhance breakage at fragile sites.

The Role of FRA16B in inv(16)(p13q22) in Acute Myeloid Leukemia

The inv(16)(p13q22) is commonly associated with the development of AML. AML is a

cancer of the myeloid line of white blood cells characterized by the proliferation of

abnormal cells in the bone marrow which interfere with the production of normal cells.

The French-American-British (FAB) classification system divides AML into nine

subtypes on the basis of cell morphology, termed M0 through M8. There have been a

variety of AML risk factors identified, including chemical exposure [98, 99], ionizing

radiation [100, 101], other blood disorders [102], and genetics [103-105]. Exposure to

anti-cancer chemotherapy agents [99] or occupational chemicals such as benzene [98],

known inducers/enhancers of fragile sites [89], has been suggested to increase one‟s risk

for AML. Ionizing radiation exposure [100, 101] or pre-leukemic blood disorders such as

myelodysplastic syndrome or myeloproliferative diseases [102] can also increase the risk

of developing AML. In addition, there have been numerous reports of AML cases

developing in families at a higher rate than predicted by chance alone, suggesting a

hereditary risk for AML [103-105]. Together, these data suggest a role for fragile sites in

the formation of inv(16)(p13q22), although this has not yet been directly proven.

16

The inv(16)(p13q22) involves the CBFB and MYH11 genes, both located on

chromosome 16, ~50 Mb apart. CBFB is located at the same chromosomal locus as

fragile sites FRA16B/FRA16C, while MYH11 spans fragile site FRA16A. While

FRA16C and FRA16A do not yet have any known clinical relevance, expression of

FRA16B has been associated with cleft palate, microstomia, micrognathia [20], and

observed in CMMol patients and individuals previously treated for NHL [21]. CBFB,

interrupted by inv(16)(p13q22) encodes core binding factor beta (CBFβ), one of the most

common targets of genetic alterations in AML [106]. CBFβ is part of the core binding

factor (CBF) complex [106], a heterodimeric transcription factor comprised of CBFβ and

core binding factor alpha (CBFα) subunits that regulate a variety of genes involved in

hematopoiesis [107]. Targeted disruption of either CBFA or CBFB by chromosomal

aberrations is found in nearly 30% of individuals with AML [108], and results in

embryonic lethality and a complete lack of definitive hematopoiesis [109-112]. The

MYH11 gene codes for smooth muscle myosin heavy chain (SMMHC), a major

contractile protein. The fusion product, CBFβ-SMMHC, blocks hematopoietic

differentiation in a dominant-negative manner by inhibiting the normal function of the

CBF transcription complex [113, 114]. In normal cells, CBFβ is distributed throughout

the cell whereas CBFα is localized to the nucleus [115]. The fusion protein inhibits

transcription by sequestering CBFα in the cytoplasm in inactive complexes, thereby

inhibiting CBFα-mediated transactivation [115, 116]. The C-terminus, consisting of

SMMHC, contains a cryptic repression domain and when bound to CBFα can increase

repression, and physically bind other corepressors, such as mSin3A [117]. Conditional

CBFβ-SMMHC knock-in mouse models have similar phenotypes to CBFβ knock-outs,

17

and have shown that the fusion product generates a distinct abnormal myeloid progenitor

population that is a target for AML transformation (Figure 1.1) [106]. While the

expression of CBFβ-SMMHC alone is not sufficient for leukemogenesis, it is critical for

the development of AML [118, 119]. Additional genetic mutations are necessary, and are

often found in genes affecting cell proliferation or survival such as NRAS, KRAS, KIT or

FLT3 [120].

The inv(16)(p13q22) is one of the most common abnormalities observed in AML,

and is detected in ~12% of adult patients [106] and 100% of patients of the M4 subtype

with accompanying eosinophilia (M4Eo) [121]. While the inv(16)(p13q22) is associated

with a favorable prognosis, only about half of patients are cured through either high-dose

cytarabine post-remission [122-124] or hematopoietic stem cell transplantation [125].

Figure 1.1: Proposed model of CBFβ-SMMHC-associated leukemia progression. HSCs have the

potential for multilineage differentiation (straight arrow) and self-renewal (curved arrow). HSCs expressing CBFβ-SMMHC generate a pre-leukemia stem cell (PSC), in which differentiation and

proliferation ability are deficient, and create a distinct abnormal myeloid progenitor (AMP) with limited proliferation potential. Acute myeloid leukemia (AML) can emerge from either PSCs or AMPs after gaining additional mutations, presumably affecting proliferation and survival programs.

Leukemic stem cells (LSCs) can generate blastlike and monocytic leukemic cells.1

__________________________________________________________________________________

1Reprinted from Cancer Cell, 9, Kuo et al., CBFβ-SMMHC induces distinct abnormal myeloid progenitors able to develop

acute myeloid leukemia, pages 57-68, 2006, with permission from Elsevier.

18

Although the exact cause(s) of leukemia remain elusive, data strongly support a role for

chemical exposure in the development of AML. Since fragile sites are known to be

induced by a variety of chemical agents, including the ones associated with leukemia, it is

possible that fragile sites play a causative role in leukemia development. Investigating

the involvement of FRA16B in inv(16)(p13q22) could therefore provide evidence of a

direct role for chromosomal fragility at fragile sites in the generation of cancer-causing

rearrangements.

19

CHAPTER II: OVER HALF OF BREAKPOINTS IN GENE PAIRS INVOLVED

IN CANCER-SPECIFIC RECURRENT TRANSLOCATIONS ARE MAPPED TO

HUMAN CHROMOSOMAL FRAGILE SITES

Allison A Burrow,

1 Laura E Williams,

1 Levi CT Pierce,

2 and Yuh-Hwa Wang

1§

1Department of Biochemistry, Wake Forest University School of Medicine, Medical

Center Boulevard, Winston-Salem, NC 27157-1016, USA. 2Department of Electrical Engineering and Computer Science, University of Kansas,

Lawrence, KS 66045-7612, USA.

§Corresponding author

The following paper was published in BMC Genomics with open access in January 2009, with modifications in numbering of Figures and References. Differences in organization

reflect the requirements of the journal.

20

ABSTRACT

Background: Gene rearrangements such as chromosomal translocations have been

shown to contribute to cancer development. Human chromosomal fragile sites are regions

of the genome especially prone to breakage, and have been implicated in various

chromosome abnormalities found in cancer. However, there has been no comprehensive

and quantitative examination of the location of fragile sites in relation to all chromosomal

aberrations.

Results: Using up-to-date databases containing all cancer-specific recurrent

translocations, we have examined 444 unique pairs of genes involved in these

translocations to determine the correlation of translocation breakpoints and fragile sites in

the gene pairs. We found that over half (52%) of translocation breakpoints in at least one

gene of these gene pairs are mapped to fragile sites. Among these, we examined the DNA

sequences within and flanking three randomly selected pairs of translocation-prone

genes, and found that they exhibit characteristic features of fragile DNA, with frequent

AT-rich flexibility islands and the potential of forming highly stable secondary structures.

Conclusion: Our study is the first to examine gene pairs involved in all recurrent

chromosomal translocations observed in tumor cells, and to correlate the location of more

than half of breakpoints to positions of known fragile sites. These results provide strong

evidence to support a causative role for fragile sites in the generation of cancer-specific

chromosomal rearrangements.

21

BACKGROUND

Tumor cells exhibit various forms of genomic instability, including chromosomal

rearrangements, many of which directly contribute to the neoplastic process rather than

occurring as a consequence [57, 126]. Rearrangements causing the deletion, insertion or

translocation of genetic material often result in the expression of altered gene products

with oncogenic potential, or the loss of tumor suppressive functions. Although the

mechanisms of these processes remain elusive, it is evident that DNA breakage is an

initiating event.

There are a variety of ways by which a cell acquires DNA breaks. Breaks can

arise from any agent that affects the primary structure of the double helix, like

endogenous reactive oxygen species or exogenous factors such as ionizing radiation [60].

More recent reports suggest that, in addition, regions of the genome especially

susceptible to breakage termed “fragile sites” may a lso cause DNA strand breakage. One

study using chromosome banding has provided compelling evidence supporting a role for

fragile sites in cancer development by demonstrating a significant association between

sites of chromosome rearrangements found in tumor cells and fragile sites [55].

Therefore, it has been proposed that fragile sites may contribute to the genetic instability

observed in cancer cells [127], but a direct role has not yet been proven.

Fragile sites are defined as non-random chromosomal loci that exhibit gaps and

breaks on metaphase chromosomes under conditions which partially inhibit DNA

synthesis [1]. Fragile sites are classified as common or rare, depending on their frequency

22

in the population, and are further divided according to their mode of induction in cultured

cells. Common fragile sites are present in all individuals, and are therefore believed to

represent a normal component of chromosome structure [6]. In contrast, rare fragile sites

are found in less than 5% of the population, and are inherited in a Mendelian manner [10,

11]. To date, about 121 different fragile sites have been identified, but the number may

increase. The majority of fragile sites can be induced by environmental agents and

chemicals, including caffeine, alcohol and cigarette smoke [1]. Variability of fragile site

breakage has been observed within individuals [2], which may reflect exposure to such

factors, with high levels being associated with cancer [3]. Many genes located within or

spanning these sites have been identified as tumor suppressors or oncogenes, and fragile

sites have been found to be preferential targets of environmental mutagens and

carcinogens [97]. Numerous studies have also revealed a significant association between

the location of fragile sites and sites of a limited number of chromosome defects in

cancer cells [78-80]. Moreover, Durkin et al. demonstrated that the deletions of the type

seen in cancer can be produced within fragile site FRA3B, a site of rearrangements and

deletions that are among the most common type of aberrations found in tumors [78],

further supporting a role for fragile sites in cancer development.

Although the mechanisms of fragile site breakage are still unclear, several factors

have been identified which contribute to fragile site instability. Studies have shown that a

deficiency of proteins in the ATR-dependent cell cycle checkpoint pathway dramatically

increases fragile site breakage [78]. In addition, all fragile DNA sequences examined so

far demonstrate significantly high flexibility [15], and are comprised of AT-rich

23

flexibility islands that can readily fold into stable secondary structures capable of

perturbing DNA replication [14, 15]. Moreover, a ll fragile sites studied to date, including

FRAXA [28], FRA7H [26], FRA3B [24], FRA10B [27], and FRA16B [27] have been

identified as late-replicating regions of the genome.

It is not entirely clear why fragile sites are susceptible to delayed replication, but

it has been proposed that the flexible, AT-rich DNA sequences cause the replication fork

to pause or stall at sites of secondary structure formation [128]. Supporting this

hypothesis, Zhang and Freudenreich demonstrated that a polymorphic AT-dinucleotide

repeat capable of forming a cruciform structure within fragile site FRA16D stalls

replication fork progression in yeast, leading to increased chromosome breakage [34].

Collectively, these results suggest a shared molecular basis among fragile sites, indicating

that the DNA sequences within these regions can present significant difficulties to the

replication machinery, and require the activation of DNA damage checkpoint proteins.

Fragile sites are therefore believed to represent unreplicated regions of the genome that

have escaped the replication checkpoint, and are visible as gaps and breaks on metaphase

chromosomes [37].

Although strong correlations have been made between the sites of breakage in a

limited number of chromosome rearrangements and the position of fragile sites, there has

been no systematic demonstration of fragile site locations relative to breakpoints in all

known chromosome aberrations. Due to the availability of extensive databases containing

chromosome abnormalities found in various types of tumors, and the increasing number

24

of fragile sites being mapped, we have compiled a table of recurrent translocations in

which each participating gene set generates cancer-specific fusion transcripts, and

mapped their breakpoints to fragile sites. We found that in more than half (52%) of the

translocation-participating gene sets, breakpoints within either one or both genes are

located at fragile sites. These results suggest that chromosome fragility, particularly at

fragile sites, may contribute to the generation of fusion transcripts in cancer cells.

Furthermore, we have analyzed the DNA sequences at and around translocation-prone

genes mapped to fragile sites for helix flexibility and the potential to form secondary

structures. Our results demonstrate that the DNA sequences contain frequent AT-rich

flexibility islands, and are capable of forming highly stable secondary structures,

supporting their propensity for breakage.

RESULTS

Breakpoints in over half (52%) of gene pairs involved in cancer-specific recurrent

translocations are mapped to fragile sites

To comprehensively investigate the relationship between fragile sites and translocat ion

breakpoints found in cancer, we examined all chromosome defects associated with

various types of tumors, and identified recurrent translocations in which translocation

breakpoints in either one or both participating genes co-localize with a fragile site.

Taking advantage of the comprehensive databases already available, we examined a total

of 444 different sets of translocation-participating genes involved in 451 translocations

obtained from the “Translocation breakpoints In Cancer database” (TICdb) [129] and

Mitelman Database of Chromosome Aberrations in Cancer [130]. We identified all genes

involved in translocations that mapped to fragile sites, and compared the position of each

25

breakpoint to fragile sites (Appendix Table 1). The types of fragile sites, rare or common,

were also documented in Appendix Table 1. We found that 52 (12%) translocation-

participating gene pairs have breakpoints for which both genes co-localize with fragile

sites (Table 2.1), and 177 (40%) gene sets contain one gene for which the translocation

breakpoint is located at a fragile site (Appendix Table 2). Therefore, we concluded that a

significant number (52%) of gene pairs involved in cancer-specific recurrent

translocations have at least one gene mapped to fragile sites. The majority (65%) of

translocation breakpoints are located at common fragile sites, as opposed to rare sites. In

Table 2.1, the partner genes in each translocation-participating pair are specified as 5‟ or

3‟ for the position in which they appear in the cancer-specific fusion transcript, and the

types of cancer for each gene pair are listed. Interestingly, the table shows that some

genes involved in translocations, like MLL and EWSR1, are most often found at the 5‟

end of the fusion product. Our data also indicate that fragile sites are involved in the

abnormalities seen in a variety of cancers including leukemias, lymphomas, and other

solid tumors, such as those of thyroid, breast, and lung.

Translocation-prone genes exhibit characteristics of fragile sites

Fragile sites have been shown to contain intrinsic features within the DNA sequence that

confer a predisposition to fragility [131]. Most fragile sites sequenced to date contain

highly flexible sequences and AT-rich islands, with the potential to form secondary

structures, which are significantly more stable than same-length random DNA sequences

[14, 15]. Therefore, to determine whether genes involved in cancer-specific recurrent

translocations exhibit properties of fragile sites, we analyzed three gene pairs from Table

26

2.1 (CBFB/MYH11, HMGA1/LAMA4, and MLL/AFF4), and their flanking sequences, for

flexibility, A/T content, and the propensity to form stable secondary structures. Using the

FlexStab program, we found that, with the exception of the MYH11 locus (Figure 2.1),

DNA sequences with significantly high flexibility occur more frequently within the

regions harboring translocation-prone genes than is predicted for control DNA [15]. The

control DNA, from various regions of the human genome where no fragile sites were

identified, contains approximately one high-flexibility peak every 100 kb [15]. Further

sequence composition analysis demonstrated that the sequences within the flexibility

peaks consist of a very high A/T content (Table 2.2), similar to what was previously

reported for fragile site regions (78% ± 1.4%), which is significantly different from that

of nonflexible sequences (61% ± 3.6%) (P < 0.001) [14]. In addition, these sequences are

rich in AT-dinucleotides (Table 2.2) to the same extent as found in the flexible peaks of

fragile site regions (21% ± 0.5%), as compared to nonflexible DNA (8% ± 1%) [14].

Further, the secondary structure prediction analysis showed that the sequences within and

surrounding genes participating in cancer-specific translocations can readily form highly

stable secondary structures, as indicated by their ΔG values (Table 2.2) and predicted

structures (Figures 2.2 and 2.3). Overall, our results demonstrate that the three gene pairs

examined display characteristics of fragile DNA, which could lead to DNA breakage,

supporting the notion that fragile sites may participate in the generation of chromosome

rearrangements.

27

Table 2.1: Translocation breakpoints mapped to fragile sites in both partner genes of

cancer-specific recurrent translocations

5'-b 3'-

b

Translocationa Gene Fragile Site Gene Fragile Site Cancer

c

t(2;18)(p11;q21) BCL2 FRA18B IGK@ FRA2L Diffuse large B-cell lymphoma

t(16;16)(p13;q22), inv(16)(p13q22)

CBFB FRA16B, FRA16C

MYH11 FRA16A Acute myeloid leukemia

inv(10)(q11q21) CCDC6 FRA10C RET FRA10G Papillary thyroid carcinoma

t(11;19)(q13;p13) CCND1 FRA11H FSTL3 FRA19B Chronic lymphocytic leukemia

t(2;7)(p11;q21) CDK6 FRA7E IGK@ FRA2L B-cell lymphoma, Chronic

lymphocytic leukemia

t(7;11)(q21;q23) CDK6 FRA7E MLL FRA11B, FRA11G

Acute lymphoblastic leukemia

t(5;7)(q35;q21) CDK6 FRA7E TLX3 FRA5G Acute lymphoblastic leukemia

t(12;22)(q13;q12) EWSR1 FRA22B ATF1 FRA12A Soft tissue tumor

t(2;22)(q33;q12) EWSR1 FRA22B CREB1 FRA2I Angiomatoid fibrous

histiocytoma

inv(22)(q12q12) EWSR1 FRA22B PATZ1 FRA22B Small round cell tumor

t(6;22)(p21;q12) EWSR1 FRA22B POU5F1 FRA6H Undifferentiated bone tumor

t(2;22)(q31;q12) EWSR1 FRA22B SP3 FRA2G Ewing tumor/small round cell

tumor

t(11;22)(p13;q12) EWSR1 FRA22B WT1 FRA11E Soft tissue tumor

del(4)(q12q12)d FIP1L1 FRA4B PDGFRA FRA4B Hypereosinophilic syndrome

inv(6)(p21q21) HMGA1 FRA6H LAMA4 FRA6F Pulmonary chondroid

hamartoma

t(3;6)(q27;p21) HSP90AB1 FRA6H BCL6 FRA3C B-cell tumors

t(1;2)(p22;p11) IGK@ FRA2L BCL10 FRA1D B-cell lymphoma

t(2;19)(p11;q13) IGK@ FRA2L BCL3 FRA19A Mature B-cell neoplasm

t(2;3)(p11;q27) IGK@ FRA2L BCL6 FRA3C Mature B-cell neoplasm,

Follicular lymphoma

t(2;11)(p11;q13) IGK@ FRA2L CCND1 FRA11A, FRA11H

Mature B-cell neoplasm

t(2;18)(p11;q21) IGK@ FRA2L FVT1 FRA18B Follicular lymphoma

t(3;16)(q27;p12) IL21R FRA16E BCL6 FRA3C Diffuse large B-cell lymphoma

t(11;19)(q13;q13.4) MALAT1 FRA11H MHLB1 FRA19A Undifferentiated embryonal

sarcoma

t(6;11)(p21.1;q13) MALAT1 FRA11H TFEB FRA6H Pediatric renal neoplasm

t(4;11)(q21.3-22.1;q23) MLL FRA11B, FRA11G

AFF1 FRA4F Acute lymphoblastic leukemia,

Acute myeloid leukemia

t(2;11)(q11;q23) MLL FRA11B,

FRA11G AFF3 FRA2A Acute lymphoblastic leukemia

28

t(5;11)(q31;q23) MLL FRA11B,

FRA11G AFF4 FRA5C Acute lymphoblastic leukemia

del(11)(q23q23)d MLL FRA11B,

FRA11G ARHGEF12

FRA11B,

FRA11G Acute myeloid leukemia

t(11;11)(q13;q23) MLL FRA11B,

FRA11G ARHGEF17 FRA11H Acute myeloid leukemia

del(11)(q23q23)d MLL FRA11B,

FRA11G CBL

FRA11B,

FRA11G Acute myeloid leukemia

t(11;19)(q23;p13) MLL FRA11B,

FRA11G ELL FRA19B Acute myeloid leukemia

t(11;22)(q23;q13) MLL FRA11B,

FRA11G EP300 FRA22A Acute myeloid leukemia

t(1;11)(p32;q23) MLL FRA11B,

FRA11G EPS15 FRA1B Acute myeloid leukemia

t(6;11)(q21;q23) MLL FRA11B,

FRA11G FOXO3A FRA6F Acute myeloid leukemia

t(3;11)(q25;q23) MLL FRA11B,

FRA11G GMPS FRA3D Acute myeloid leukemia

t(11;14)(q23;q23) MLL FRA11B,

FRA11G GPHN FRA14B Acute myeloid leukemia

t(11;19)(q23;p13.3) MLL FRA11B,

FRA11G MLLT1 FRA19B Acute myeloid leukemia

t(1;11)(q21;q23) MLL FRA11B,

FRA11G MLLT11 FRA1F Acute myeloid leukemia

t(9;11)(p21;q23) MLL FRA11B,

FRA11G MLLT3

FRA9A,

FRA9C Acute myeloid leukemia

t(11;19)(q23;p13) MLL FRA11B,

FRA11G MYO1F FRA19B Acute myeloid leukemia

inv(11)(q14q23) MLL FRA11B,

FRA11G PICALM FRA11F Acute myeloid leukemia

t(2;11)(q37;q23) MLL FRA11B,

FRA11G SEPT2 FRA2J Acute myeloid leukemia

t(11;19)(q23;p13) MLL FRA11B,

FRA11G SH3GL1 FRA19B Acute myeloid leukemia

t(6;11)(q13;q23) MLL FRA11B,

FRA11G SMAP1 FRA6D Acute myeloid leukemia

t(10;11)(q21;q23) MLL FRA11B,

FRA11G TET1 FRA10C Acute myeloid leukemia

t(9;9)(p21;p21) MTS2 FRA9A,

FRA9C MTS1

FRA9A,

FRA9C Acute lymphoblastic leukemia

inv(10)(q11q11) NCOA4 FRA10G RET FRA10G Papillary thyroid carcinoma

t(3;5)(q25;q35) NPM1 FRA5G MLF1 FRA3D Acute myeloid leukemia

t(3;6)(q27;p21) PIM1 FRA6H BCL6 FRA3C Diffuse large B-cell lymphoma

t(3;6)(q27;p21) SFRS3 FRA6H BCL6 FRA3C Follicular lymphoma

t(19;19)(p13;q13) TCF3 FRA19B TFPT FRA19A Acute lymphoblastic leukemia

inv(1)(q21q31) TPM3 FRA1F TPR FRA1K Papillary thyroid carcinoma

a The translocation names for each unique gene set are indicat ed. b Position in which the gene appears in the cancer -speci fi c fusion transcript c For a complete description, see the Mitelman database of Chromosome Aberrations in Cancer [130]. d Deletions creating a fusion between two genes

29

DISCUSSION

Research on understanding the significance of human chromosomal fragile sites in cancer

has been very active. Fragile sites are most commonly associated with deletion

breakpoints in tumor cells, while few translocations involving these sites have been

reported. A limited number of translocation breakpoints have been reported near fragile

sites, suggesting that chromosome fragility at these sites may contribute to these

rearrangements [132]. Our study herein provides a comprehensive survey of all cancer-

specific translocations to date. Therefore, by demonstrating that breakpoints in over half

(52%) of the translocations co-map to fragile sites, the results provide strong evidence to

support a role for fragile sites in the generation of cancer-specific translocations. It is

important to note that we have chosen to focus on translocations and deletions leading to

fusion transcripts, and have not included other types of rearrangements such as single

gene deletions, insertions, or complex translocations. We have found that many of the

genes examined in this study are commonly involved in these other types of

rearrangements, like deletion of the FHIT gene located within fragile site FRA3B, and in

some cases, the same set of genes is involved in multiple translocations observed in a

variety of cancers. An interesting observation was that some genes located near fragile

sites, such as NUP98 (11p15.4) which participates in twenty-two different translocations

examined, could not be included in Appendix Table 1, because the gene was not located

directly at a fragile site. Recently described fragile sites, like FRA6H at 6p21 [133], have

been identified after discovering an association between the chromosome location a nd

sites of recurrent aberrations in disorders. This suggests that 11p15.4 could be a fragile

site that has not yet been identified, or that the proximal fragile sites FRA11C and

30

A B

C D

E F

Figure 2. 1: DNA flexibility analysis of translocation-prone and fragile site co-localized genes.

DNA sequences within and flanking genes (A) CBFB (B) MYH11 (C) HMGA1 (D) LAMA4 (E) MLL (F) AFF4 were analyzed using the FlexStab program. The analysis was performed over the length of the entire gene (shaded in black) plus 125 kb flanking on each side (shaded in gray). The x axis

indicates the size of the analyzed sequences, and the y axis shows degrees of inclination in the twist angle. Windows with values >13.7˚ were considered as significantly high flexibility peaks [15].

31

Table 2.2: Computational analysis of genes involved in cancer-specific recurrent

translocations reveals characteristics of chromosomal fragile sites

FlexStab Mfold

Gene Number of flexibility

peaks/kb % A/T

% AT-dinucleotides

Lowest ΔG valuea

(kcal/mol)

CBFB 4/322 79 ± 3.9 24 ± 3.0 -116.91

MYH11 4/404 78 ± 5.5 23 ± 5.9 -97.02

HMGA1 6/259 81 ± 7.8 24 ± 3.6 -124.07

LAMA4 9/397 78 ± 2.7 23 ± 3.0 -59.20

MLL 5/339 78 ± 10.2 23 ± 4.8 -100.74

AFF4 4/338 81 ± 3.3 26 ± 3.1 -100.97

Fragile site 78 ± 1.4c 21 ± 0.5

c

Control 1/100b 61 ± 3.6

c 8 ± 1.0

c -41.79

d

aLowest ΔG value, predicted by Mfold bMishmar et al. [15] examined 1.1 Mb of non-fragile DNA, and showed that regions with significantly high flexibility occur every ~100 kb. cZlotorynski et al. [14] dThe LAMA4 sequence was randomized 1000 times to serve as a control, and then analyzed by Mfold.

FRA11I at 11p15.1 are larger than previously determined. Therefore, it is appropriate to

assume that the total number of chromosomal aberrations in cancer associated with

fragile sites could be even greater than presented in this study, arguing for the

significance of the involvement of fragile site in tumorigenesis.

In addition to solidifying the role of fragile sites participating in cancer

development, this study also supports the common hypothesis for the molecular basis of

fragility at these sites. We have shown that the DNA sequences within and surrounding

three pairs of translocation-prone genes exhibit features of fragility. On average, peaks of

significantly high flexibility occur more often than in random DNA, which is consistent

with previous results [15]. We also found these peaks to have a high A/T content and to

be rich in AT-dinucleotides to the same extent as established in fragile sites [14].

32

A

B

Figure 2.2: Secondary structure analysis of CBFB, MYH11, HMGA1, LAMA4, MLL, and AFF4

loci. (A) Comparison of potential to form secondary structure for these genes versus a control. The

computed lowest free energy of predicted DNA secondary structures from segments of 300 nt in length, overlapping in 150 nt steps, has been fit to a curve for each gene. The Matlab function polyfit finds coefficients of a polynomial P(X) of degree N that fit the raw data best in a least -squares sense. The

analysis was performed over the length of the entire gene plus 125 kb flanking on each side. The arrows indicate where a gene begins and ends. The control sequence was generated by randomizing LAMA4

1000 times. The x axis indicates the size of the analyzed sequences, and the y axis displays the free energy of the predicted structure. Raw data plots for each gene are shown in Figure 2.3. (B) The most stable structure predicted for each gene, as produced by Mfold. Each structure represents the 300 nt

segment with the lowest ΔG value.

33

Furthermore, our data from the Mfold program indicate that the sequences have the

potential to form highly stable secondary structures, another distinct characteristic of

fragile sites [14], which could disturb progression of the replication fork. Based on our

results, and the proposed mechanism of fragile s ite expression, it is likely that the AT-

rich flexibility islands within or flanking translocation-prone genes are able to stall

replication by the formation of secondary structures, which may then lead to DNA strand

breakage, and ultimately to chromosome rearrangements.

Several proteins involved in the replication checkpoint pathway are essential for

maintaining stability at fragile sites [78]. These include the S phase and G2/M checkpoint

kinase ATR [37], and its downstream targets BRCA1 [46], FANCD2 [50], and CHK1

[47]. ATR is a major component of the checkpoint pathway, where it functions by

sensing and responding to DNA damage, including stalled and collapsed replication forks

[134, 135]. It is hypothesized that ATR maintains fragile site stability by sensing and

binding to single-stranded DNA resulting from stalled replication forks [37]. However,

in the absence of ATR, the main transducer of the DNA double-strand break (DSB)

signal, which is ATM, has been shown to regulate fragile site stability [45], indicating

that DSBs also occur at fragile sites. Following breakage, chromosome rearrangements

may take place via the homologous recombination (HR), non-homologous end-joining

(NHEJ) DSB repair pathways [136, 137], or microhomology-mediated single-strand

annealing (SSA) [138]. The repair of lesions at fragile sites is still not clear, but evidence

suggests that all three pathways may be involved. Based on recent observations made by

Lieber et al., it is hypothesized that the sequence-specific RAG complex involved in

34

V(D)J recombination, which is an important process of the NHEJ type, may also

recognize and cleave at non-B DNA structures [139], a feature shared by all fragile sites

examined to date. Schwartz et al. [58] have shown that induction of fragile sites leads to

Figure 2.3: The computed lowest free energy of predicted DNA secondary structures. The flanking 125 kb regions are shaded in light gray, and the gene region is shaded in black. The black box indicates the location of the most stable structure found in the sequence. The black line is the curve

which best fits the raw data. These curves were generated using the polyfit function of the Matlab program, and are presented in Figure 2.2A.

35

RAD51 focus formation, and phosphorylation of DNA-PKcs, key components of the HR

and NHEJ pathways, respectively. Furthermore, they found that down-regulation of

RAD51 and DNA-PKcs increases fragile site instability, suggesting that both HR and

NHEJ DSB repair pathways mediate break repair at fragile sites. In addition, the majority

of breakpoints in a papillary thyroid carcinoma rearrangement mapped to fragile sites

occur at microhomology patches, indicating that fragile site-associated rearrangements

can also arise by microhomology-mediated SSA [59]. Although the underlying basis of

chromosome rearrangements is still unclear, our results show that 39% of the

translocations examined have only one breakpoint mapped to a fragile site, suggesting

that gene rearrangements could be achieved with strand breakage at only one gene, as

well as at both participating genes. To support this hypothesis, additional studies are

needed to obtain a greater understanding of the mechanisms of chromosome

rearrangements.

The most intriguing finding from this study is that the majority (65%) of fragile

sites mapped to translocation breakpoints are common fragile sites, which are present in

all individuals, and can be induced by a variety of environmental factors and chemical

agents. Interestingly, most cancers associated with the translocations examined in this

study have little or no genetic component. These observations suggest that exposure to

fragile site-inducing chemicals and/or reduced levels of proteins critical for the

maintenance of fragile sites may confer a risk for cancer-specific rearrangements. It will

be important to identify factors that contribute to chromosome fragility, such as DNA

sequences, proteins and environmental/dietary agents, since fragile sites are sensitive to a

36

range of chemicals. Understanding the molecular basis of fragile sites could therefore

allow development of a prognostic assay for cancer risk.

CONCLUSION

Our study is the first to comprehensively compare the location of cancer-specific

recurrent translocation breakpoints and fragile sites. We showed that over half (52%) of

the translocation breakpoints examined co-map to positions of known human

chromosomal fragile sites. Furthermore, we demonstrated that the DNA sequences at and

surrounding three pairs of translocation-prone genes that are mapped to fragile sites

exhibit frequent AT-rich flexibility islands and are capable of forming highly stable

secondary structures, both of which are characteristics of fragile DNA. Thus, we have

provided a greater understanding of the contribution of fragile sites in the formation of

chromosomal translocations, and further supported a role for fragile sites in cancer

development.

METHODS

Data collection

Two databases were examined encompassing all chromosome rearrangements found in

cancer. The freely available TICdb was downloaded from

http://www.unav.es/genetica/TICdb/ (v2.3 May 2008), which contains the breakpoints of

cancer-specific reciprocal translocations mapped to over 300 different human genes

[129]. In addition, we cross-examined the Mitelman Database of Chromosome

Aberrations in Cancer, a database relating chromosomal aberrations to tumor

http://www.unav.es/genetica/TICdb/

37

characteristics for recurrent translocations using the Molecular Biology Associations

search tool [130]. From these databases, 444 different pairs of genes participating in

cancer-specific recurrent translocations were obtained. In some cases, the same set of

genes is involved in multiple types of translocations. All of these translocations result in

one or two cancer-specific fusion genes. For each gene, the chromosomal locus was

obtained from the UCSC Genome Bioinformatics website http://genome.ucsc.edu, and

compared to the mapped positions of all known fragile sites

(http://www.ncbi.nlm.nih.gov/sites/entrez?db=gene using “fragile site” as a search term)

[133, 140-142]. All genes located at fragile sites are highlighted in Appendix Table 1.

The translocation breakpoints of all genes highlighted in Appendix Table 1 were

compared to fragile site positions, to identify translocations in which the site of breakage

is co-localized with a fragile site at either one (Appendix Table 2) or both (Table 2.1)

genes involved in the translocation.

Flexibility analysis

DNA helix flexibility was assessed using FlexStab, a computer program designed by

Mishmar et al. [15], which measures potential local variations in DNA between

consecutive base pairs, and is expressed as f luctuations in the twist angle. The analysis

was performed over the length of the entire gene plus 125 kb flanking on each side, in

windows of 100 bp with 25 bp shift increments. Windows with values >13.7˚ were

considered as significantly high flexibility peaks [15].

http://genome.ucsc.edu/

http://www.ncbi.nlm.nih.gov/sites/entrez?db=gene%20

38

Three gene pairs in which both genes are mapped to fragile sites (CBFB/MYH11,

HMGA1/LAMA4, and MLL/AFF4) were used for flexibility and secondary structure

analysis. The examined sequences for each gene are: CBFB [nucleotides (nt)

20552249~20874160 of GenBank: NT_010498.15], MYH11 [nt 6985071~7388966 of

NT_010393.15], HMGA1 [nt 24937900~25197258 of NT_007592.14], LAMA4 [nt

16473563~16870257 of NT_025741.14], MLL [nt 21744621~22083352 of

NT_033899.7], and AFF4 [nt 34501084~34839367 of NT_034772.5].

Assessment of secondary structure

Using Zukers‟ Mfold program [29], the potential of a single-stranded DNA to form stable

secondary structure can be predicted along with its free-energy value. For a given gene,

300 nt segments with 150 nt shift increments were used as input for Mfold. The length of

300 nt was chosen because it equals the Okazaki initiation zone of the DNA replication

fork in mammalian cells, which possesses a single-stranded property during DNA

replication [143, 144]. The default [Na+], [Mg

2+], and temperature used were 1.0M,

0.0M, and 37˚C, respectively. The program Bioperl

http://www.bioperl.org/wiki/Main_Page was used to manipulate the sequences, and

Matlab (MathWorks) was used to perform the analysis of the data.

AUTHORS’ CONTRIBUTIONS

AB and LEW compiled and analyzed the databases. AB and LCTP carried out the

flexibility and secondary structure analyses. AB and Y-HW wrote the paper. Y-HW

designed and coordinated the study. All authors read and approved the final manuscript.

http://www.ncbi.nlm.nih.gov/sites/entrez?cmd=Retrieve&db=nucleotide&dopt=GenBank&list_uids=51473102






http://www.bioperl.org/wiki/Main_Page

39

ACKNOWLEDGEMENTS

This work was supported in part by Public Health Service grants from the National

Cancer Institute (CA85826 and CA113863 to Y-HW).

40

CHAPTER III: SECONDARY STRUCTURE FORMATION AND DNA

INSTABILITY AT FRAGILE SITE FRA16B

Allison A. Burrow, Allison Marullo, Lindsay R. Holder, and Yuh-Hwa Wang

*

Department of Biochemistry, Wake Forest University School of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1016, USA.

*To

whom correspondence should be addressed. Tel: +1 336 716 6186; Fax: +1 336 716

7671; Email: [email protected]

The following paper was published in Nucleic Acids Research with open access in January 2010, with modifications in numbering of Figures and References.

41

ABSTRACT

Human chromosomal fragile sites are specific loci that are especially susceptible to DNA

breakage following conditions of partial replication stress. They often are found in genes

involved in tumorigenesis and map to over half of all known cancer-specific recurrent

translocation breakpoints. While their molecular basis remains elusive, most fragile

DNAs contain AT-rich flexibility islands predicted to form stable secondary structures.

To understand the mechanism of fragile site instability, we examined the contribution of

secondary structure formation to breakage at FRA16B. Here, we show that FRA16B

forms an alternative DNA structure in vitro. During replication in human cells, FRA16B

exhibited reduced replication efficiency and expansions and deletions, depending on

replication orientation and distance from the origin. Furthermore, the examination of a

FRA16B replication fork template demonstrated that the majority of the constructs

contained DNA polymerase paused within the FRA16B sequence, and among the

molecules which completed DNA synthesis, 81% of them underwent fork reversal.

These results strongly suggest that the secondary structure-forming ability of FRA16B

contributes to its fragility by stalling DNA replication, and this mechanism may be shared

among other fragile DNAs.

42

INTRODUCTION

Fragile sites are specific chromosomal regions located throughout the human genome that

are especially susceptible to DNA breakage. These regions are defined cytogenetically as

gaps or breaks on metaphase chromosomes following conditions of partial replication

stress. Fragile sites are divided into two major classes based on the ir frequency in the

population and are subdivided according to their mode of induction in cultured cells.

Rare fragile sites are found in <5% of the population and are inherited in a Mendelian

manner [10, 11]. The majority of rare fragile sites can be induced under folate-deficient

conditions and contain a microsate llite (CGG)n repeat [13], whereas the rare, non-folate-

sensitive sites are comprised of an AT-rich minisatellite element [11]. In contrast,

common fragile sites have been observed in all individuals and are believed to represent a

normal component of chromosome structure [6]. Most common fragile sites are observed

after exposure to low doses of aphidicolin, an inhibitor of DNA polymerases α, δ and ε

[7, 8]. To date, over 80 common fragile sites are listed in the Human Genome Database

(GDB). Most have not yet been investigated at the molecular level, but it is known that

regions of fragility can extend over megabases of DNA with gaps or breaks occurring

throughout [9]. Although a consensus sequence has not yet been identified among

common fragile sites, the DNAs examined thus far contain frequent, AT-rich flexibility

islands capable of forming secondary structures that are much more stable compared to

other regions of the genome [14], similar to what has been reported for most rare sites.

Fragile sites are normally stable in cultured cells. However, these sites are

hotspots for sister chromatid exchanges, deletions and rearrangements after induction

43

with replication inhibitors [4, 5]. Moreover, many fragile sites are frequently associated

with sites of chromosomal breakage in tumors [22, 78]. While the exact mechanism of

fragile site expression remains elusive, replication timing experiments have shown that

all fragile sites studied to date, including FRAXA [28], FRA3B [24], FRA7H [26],

FRA10B [27], FRA16B [27], FRA1H [23] and FRA2G [23] exhibit delayed replication.

The delay is further exacerbated by the addition of replication inhibitors, with some

fragile site alleles remaining unreplicated in late G2 phase [24, 26]. Although it is not

entirely clear how delayed replication at fragile sites results in chromosome breakage,

evidence suggests that DNA sequences with the potential to form stable secondary

structures can present significant difficulties during replication, which may lead to

unreplicated regions of the genome that are visible as gaps and breaks during metaphase

[37]. The (CGG)n repeat within rare, folate-sensitive sites has been shown to form hairpin

[31] and quadruplex structures [30] that present a significant block to replication both in

vitro and in vivo [32, 33], whereas a polymorphic AT-rich sequence with the ability of

forming a cruciform within common fragile site FRA16D blocked replication in yeast,

resulting in increased chromosome breakage [34].

Several studies have demonstrated a critical role for the Ataxia-Telangiectasia and

Rad3-Related (ATR)-dependent DNA damage checkpoint pathway in the maintenance of

fragile sites. Although their direct roles remain unclear, proteins including the S-phase

and G2/M checkpoint kinase ATR [37], as well as its downstream targets BRCA1 [46]

and CHK1 [47], are required for fragile site stability, as their deficiencies result in

significantly increased fragile site breakage. ATR is a major component of the checkpoint

44

pathway where it functions by sensing and responding to DNA damage, including stalled

and collapsed replication forks [134, 135]. Based on this evidence, it is hypothesized that

ATR maintains fragile site stability by sensing and binding to s ingle-stranded DNA

resulting from stalled replication forks at sites of secondary structure formation [37], and

that a deficiency or defect in ATR can prevent repair, leading to increased fragile site

expression. Supporting this hypothesis, cells from patients with Seckel syndrome, who

have low levels of ATR protein, show increased instability at fragile sites compared to

normal cells following replication stress [43]. This difference in instability suggests that

the low level of ATR present in Seckel syndrome patients is not sufficient to respond to

replication stress, consistent with the hypothesis that the breaks observed on metaphase

chromosomes are unreplicated fragile sites that escape the ATR checkpoint [37].

To further understand the mechanism of breakage at human chromosomal fragile

sites, we have investigated the cause of instability at fragile site FRA16B [145].

Spontaneous FRA16B expression has been observed among individuals and can be

induced by chemicals that bind AT-rich DNA, such as distamycin A [16] or berenil [17].

Studies of FRA16B, located at 16q22.1, have shown that it spans the same genomic

region as the common fragile site FRA16C and is also apparent following treatment with

aphidicolin [14]. After induction, the heterozygote frequency of FRA16B is about 5% in

populations of European descent, representing the most frequently expressed rare fragile

site [18]. Positional cloning has revealed that FRA16B-expressing chromosomes may

contain up to 2000 copies of a 33-bp AT-rich minisatellite repeat (ATATATTATATAT-

TATATCTAATAATATATC/ATA), whereas normal chromosomes consist of only 7-12

45

copies of the repetitive element [19]. Competitive nucleosome reconstitution assays

demonstrate that, in the presence of distamycin A, FRA16B DNA displays nucleosome

exclusion [145, 146] that increases in proportion to the number of repeats [145, 146].

These results, along with similar studies on various other fragile sites [147-149] suggest a

common feature for the chromatin structure surrounding fragile DNAs, which may play

an important role in their expression. Therefore, FRA16B serves as a model for the

examination of fragile sites, since it exhibits characteristics of both common and rare

fragile sites, and has been mapped to cancer-specific rearrangements [22].

CGG repeats underlying the basis of fragility at rare, folate-sensitive fragile sites

have been shown to form a secondary structure in vitro [150], but there is no physical

evidence of alternative DNA structures for AT-rich fragile DNA, which comprises the

majority of fragile sites. Therefore, it is important to determine whether these sequences

are capable of forming secondary structures. It is also critical to investigate the

contribution of secondary structure formation on chromosome instability. Recently,

Ragland et al. demonstrated the ability of common fragile DNAs to induce chromosome

instability at ectopic sites in HCT116 cells [151], although the relationship between

common fragile site instability and secondary structure was not investigated. In addition,

the role of secondary structure formation in chromosome instability has been determined

in yeast [34] but has not yet been investigated in human cells.

In this study, the ability of FRA16B to form a secondary structure in vitro was

demonstrated by a reduction of electrophoretic mobility in polyacrylamide gels and

46

visualization of short, branched structures by electron microscopy (EM). It was also

determined that both replication orientation and distance from FRA16B to the origin

affect FRA16B replication efficiency and instability in human cells. Furthermore,

examination of FRA16B replication fork templates by EM revealed a tendency for

FRA16B DNA to promote spontaneous fork reversal and an even greater occurrence of

polymerase pausing at specific sites within the FRA16B region, which was confirmed by

DNA sequencing gels. Overall, these results strongly suggest that the secondary

structure-forming ability of FRA16B contributes to its fragility by stalling DNA

replication, and this mechanism may be shared among other fragile DNAs.

MATERIALS AND METHODS

Plasmids

FRA16B-containing plasmids were created using genomic DNA from an individual

expressing the FRA16B chromosome. A 921-bp fragment of FRA16B consisting of 14

perfect copies of the 33-bp AT-rich minisatellite repeat and some imperfect repeats with

AT-rich flanking sequences was cloned as described in [146] to generate pFRA16B18.

To construct SV40 replication templates, the FRA16B fragment described above

and the SV40 origin of replication (ori) [144] were cloned into pGEM3zf(+) (Promega).

The FRA16B sequence was inserted at one of four different distances relative to the ori,

and in one of two possible replication orientations. This generated eight constructs with

the FRA16B sequence located 30, 300, 400, or 700 bp from the origin of replication in

47

two different replication orientations (Figure 3.3A). A plasmid containing the SV40 ori

only with no FRA16B sequence (pGEM-SV40ori) served as a control.

To construct FRA16B replication fork templates, complimentary oligonucleotides

(5-AGCTTGCATGCCTGCAGGCTGAGGA-3 and 5-CCTCAGCCTGCAGGCAT-

GCA-3) containing a site for the nicking endonuclease Nb.BbvCI (New England

Biolabs) were annealed and cloned into the EcoRI and HindIII sites of pFRA16B18

adjacent to the 921-bp FRA16B fragment to create pFRA16B37 (Figure 3.5). The

FRA16B DNA-containing sequence of pFRA16B37, and the Nb.BbvCI recognition site

are also indicated in Figure 3.5. The construction of these plasmids allowed us to

examine DNA polymerase stalling, and replication fork regression of a 487-bp fragment

of FRA16B DNA (see below).

Reduplexing assay

Reduplexing reactions were performed as previously described in [150, 152, 153].

Briefly, the FRA16B fragment from pFRA16B18 was obtained by restriction enzyme

digest using EcoRI and HindIII (New England Biolabs), and gel-purified. The FRA16B

fragment was then dephosphorylated at the 5 end with calf intestinal alkaline

phosphatase (New England Biolabs) and end-labeled with [α-32

P] ATP (PerkinElmer)

using T4 kinase (New England Biolabs). End-labeled DNA (1 ng) was added to solutions

of 500 μL containing 1, 0.58, 0.3 or 0.1 M NaCl in TE buffer and incubated at 95˚C for 5

min. The samples were cooled to room temperature, and the DNA was ethanol-

precipitated in the presence of glycogen (Roche). The DNA pellets were then air-dried

48

and resuspended in TE buffer. As a control, pGEM3zf(+) was digested with EcoRI and

Eco109I (New England Biolabs) to generate an 892-bp fragment, and was subjected to

the same conditions as FRA16B. DNA samples were electrophoresed in a 4%

polyacrylamide gel cast in TBE at 50 V for 18 h at room temperature. The gel was dried

and visualized by phosphorimaging (GE HealthCare).

Electron microscopy

Reduplexed DNAs or replication fork template reaction mixtures were directly mounted

onto glow-charged carbon-coated copper EM grids followed by washing in a

water/ethanol gradient and rotary shadowcasting with tungsten, as described previously

[154]. DNAs were visualized on a Philips transmission electron microscope 400.

Measurements of the DNA lengths were determined using Image J software (NIH).

Replication efficiency

The human embryonic kidney HEK293T cells were grown in Dulbecco‟s modified

Eagle‟s media (Gibco) containing 10% fetal bovine serum (HyClone) and 1% penicillin

streptomycin (Gibco) on 100-mm-diameter plates. The cells were 50% confluent when

co-transfected with 2.5 μg of pGEM-SV40ori and 2.5 μg of each FRA16B/SV40 plasmid

using the CaPO4 method. The cells were allowed to grow for 16 h following transfection

before replacing the media. Low-molecular-weight DNA was extracted by the Hirt‟s lysis

method [155] 48 h after transfection. The SV40 viral DNAs were further purified by

incubation with proteinase K, phenol/chloroform extraction, and alcohol precipitation.

To determine replication efficiency of the constructs (Figure 3.3B), SV40-replicated

49

DNAs were digested with HindIII and NdeI (New England Biolabs) to linearize the

plasmids, and with DpnI (New England Biolabs) to remove unreplicated parental

templates. For clone 46, EcoRI was used in place of HindIII. Southern blot analysis was

then used to identify replicated DNAs us ing an [α-32

P] dCTP-labeled probe hybridizing

to nucleotide numbers 1725 to 2132 of pGEM3zf(+), which is present in all nine

constructs. Replication efficiency was determined by the ratio of replicated FRA16B

DNA to the control (pGEM-SV40ori) using ImageQuant version 5.2 to measure the

intensity of each band. Student‟s t-test was performed to determine statistically

significant differences between clones.

Mutation assay

To investigate instability associated with FRA16B, a modified version of the stability of

trinucleotide repeat by individual product (STRIP) assay [156] was used to examine

individual products following replication in human cells. Essentially, products of

replication from transfected HEK293T cells were digested with DpnI to eliminate any

unreplicated parental templates and transformed into SURE-2 cells (Stratagene) (Figure

3.3B). Approximately 60 single colonies from each clone were picked at random, and the

DNAs were isolated and digested with appropriate restriction enzymes to release the

FRA16B insert. Samples were then run on 1.5% agarose gels and scored for insertion or

deletion events, characterized by slower or faster migrating bands, respectively,

compared to unreplicated DNA. As an additional control to measure the background

instability in Escherichia coli, DNAs that were not transfected into HEK293T cells were

directly transformed into SURE-2 cells, and scored for instability events. Fisher‟s exact

50

test was used to determine the statistical significance of instability resulting from

replication in HEK293T cells compared to the background instability in E. coli, as well as

differences between FRA16B-containing constructs compared to the control.

Analysis of FRA16B synthesis by the Klenow fragment of DNA polymerase I

A DNA template (pFRA16B37) was synthesized which mimics a replication fork that has

progressed through clone 17 FRA16B sequence. The plasmid was digested with

Nb.BbvCI to generate a nick directly in front of the FRA16B fragment that does not

contain any cytosine bases for a length of 487 bp (Figure 3.5). The nicked DNA was then

incubated with 5 U of the Klenow fragment (exo-) of DNA polymerase I (New England

Biolabs) in a reaction mixture containing 7.5 mM Tris, pH 7.5, 5 mM MgCl2, and 5 mM

DTT with 2.5 mM each dATP, dTTP, and dGTP nucleotides. The absence of cytosine

caused DNA synthesis to stop at the end of the repetitive tract when the first guanine was

encountered on the template strand. This generated a single-strand tail by strand

displacement. This resulted in a duplex circle with a single-strand tail where the tail

represents the lagging strand, one arm of the circle is the leading strand and the other arm

serves as the template (Figure 3.6A). To examine DNA synthesis on the lagging strand, a

50-fold molar excess of the complementary oligonucleotide (5- ATATAATA-

TATTATTATATCTAATA -3) was annealed to the lagging strand at the 3 end for 30

min at 37˚C in a buffer containing 7.5 mM Tris, pH 7.5, 5 mM MgCl2, and 5 mM DTT

with 2.5 mM each dATP, dTTP, and dCTP nucleotides, and further incubation with 5 U

of Klenow fragment (exo-) for 30 min at 37˚C. The reaction mixture was purified and

examined by EM for DNA synthesis. To serve as a control, a plasmid containing non-

51

fragile site DNA (pGLGAP) was kindly provided by Dr. Jack Griffith (UNC-CH) and

constructed using the same conditions as the FRA16B replication fork template [157].

To detect the pause sites on the lagging strand, the same oligonucleotide was

radiolabeled and annealed to the lagging strand in the same reaction conditions as

described above. DNA synthesis was carried out using 0.5, 5 and 15 U of Klenow

fragment (exo-). The reaction mixtures were purified, resuspended in formamide/dye

solution, electrophoresed through 12% polyacrylamide-7 M urea gels, and examined by

phosphorimager analysis.

To locate the position of the replication fork for detecting fork regression, the

reaction mixtures were linearized with AhdI or XmnI (New England Biolabs), to produce

asymmetrical arms for measuring the distance from the fork junction to the DNA ends ,

and the samples were examined by EM.

RESULTS

Secondary structure formation of FRA16B DNA

Since there has been no physical evidence of alternative DNA structures formed for AT-

rich fragile DNAs, we investigated secondary structure formation by subjecting a 921-bp

fragment of FRA16B to reduplexing in the presence of various concentrations of NaCl to

allow re-annealing of the single strands following denaturation. Separation by

polyacrylamide gel electrophoresis (PAGE) showed that the reduplexed FRA16B

fragment gave rise to two slower-migrating products over a range of NaCl concentrations

52

(Figure 3.1A). These products were not present in the untreated FRA16B sample or the

reduplexed pGEM3zf(+) control, suggesting the formation of a secondary structure

during reduplexing of FRA16B DNA. Furthermore, when FRA16B was denatured and

only one strand of the duplex was labeled with 32

P, the labeled strand corresponded

uniquely to one of the bands with reduced electrophoretic mobility, demonstrating that

each reduplexed band is produced from one of the two strands of FRA16B (Figure 3.2).

The examination of these molecules by EM confirmed that FRA16B folded into branched

duplexes following denaturation and re-annealing (Figure 3.1B). These structures were

not observed in the untreated FRA16B or reduplexed pGEM3zf(+) samples. The lengths

of 100 DNA molecules from each sample were measured, and the comparison revealed

that the reduplexed FRA16B molecules were shorter than the untreated FRA16B DNAs,

indicating the presence of slipped-out regions participating in the formation of secondary

structure. These data are the first to demonstrate that FRA16B is indeed able to form an

alternative DNA structure in vitro.

Reduced replication efficiency and increased instability of FRA16B in human cells

To analyze FRA16B replication in human cells, an SV40 system was used to evaluate

replication efficiency and instability of FRA16B-containing constructs. Since several

studies have shown that the ability of a sequence to inhibit replication depends on cis-

acting factors, including replication orientation and distance relative to the origin [33, 34,

156, 158-160], the 921-bp FRA16B insert was placed at varying distances from the SV40

ori and in one of two replication orientations to generate eight constructs containing

53

FRA16B (Figure 3.3A). A control plasmid that does not contain FRA16B DNA was also

constructed. An equal amount of the control plasmid and each FRA16B-containing

plasmid was co-transfected into HEK293T cells (Figure 3.3B). Following SV40

A

C

pGEM3zf(+)

C CMNaCl

FRA16B

NaCl

B

Figure 3.1: Secondary structure formation of FRA16B DNA following denaturation and re-

annealing (reduplexing) reaction. (A) Gel electrophoresis analysis of re-annealed FRA16B DNA. DNA fragments of FRA16B (right) and pGEM3zf(+) (left) were subjected to reduplexing in increasing salt concentrations (0.1-1 M) and analyzed by native 4% PAGE. C, untreated samples; M, molecular

weight marker. (B) Visualization of reduplexed FRA16B DNA by EM. After reduplexing reactions, samples were directly mounted onto carbon-coated copper EM grids and rotary shadowcasted with tungsten. Images are shown in reverse contrast. The total lengths of 100 molecules from each sample

were measured using Image J software (lower right).

54

replication, the DNAs were linearized and digested with DpnI to remove unreplicated

parental templates. Replication efficiency was determined on Southern blots as the ratio

of completely replicated FRA16B DNA to the control (Figure 3.3C). Clone 17 exhibited

a statistically significant reduction in replication efficiency compared to clone 15 (P =

0.045), indicating that orientation affects replication efficiency. The efficiency of

replication for clone 17 was also reduced when compared to clone 34 (P = 0.073), clone

10 (P = 0.003), clone 12 (P = 0.005) and clone 46 (P = 0.083). Our results demonstrate

that both the replication orientation and distance from the origin affect FRA16B

replication.

Next, a mutation assay was performed to determine if instability events were

occurring within FRA16B during replication in HEK293T cells. Individual replication

products were examined using a modified version of the STRIP assay [156], originally

designed to determine the role of mammalian replication on the stability of triplet repeats.

FRA16B

Figure 3.2: Each strand of FRA16B forms a secondary structure with different electrophoretic mobilities. FRA16B DNA subjected to reduplexing (lanes 2 and 4) and the untreated samples (lanes 1 and 3) were analyzed by native PAGE (see “Materials and Methods”). Lanes 1 and 2: Both strands were labeled

with [γ-32

P] ATP (lane 1, untreated; lane 2, reduplexed). Lanes 3 and 4: Only one strand representing the lagging strand template of clone 17 was labeled with [α-

32P] dCTP (lane 3, untreated; lane 4, reduplexed).

55

A

B

Figure 3.3: Analysis of FRA16B replication efficiency and instability in human cells using an SV40

replication system. (A) Diagram of replication constructs used for replication efficiency and instability assays. The bidirectional SV40 ori (open circles) was cloned into pGEM3zf(+) to generate pGEM-SV40ori, used as a control. The 921-bp FRA16B fragment (shaded box) was inserted 30, 300, 400 or 700

bp from the SV40 ori, and in one of two replication orientations (represented by arrows). (B) Schematic of strategy used to determine the replication efficiency of FRA16B constructs and instability events following replication in HEK293T cells. To determine replication efficiency, equal amounts of pGEM-SV40ori and

each FRA16B-containing construct were co-transfected into HEK293T cells. After replication for 48 h, SV40 DNAs were extracted, purified and digested with DpnI to remove any unreplicated templates.

Replicated molecules were then subjected to Southern blot analysis. To examine FRA16B-associated instability, each FRA16B-containing construct was transfected and replicated in HEK293T cells. SV40 DNA was extracted, purified and digested with DpnI to remove any unreplicated templates. These DNAs

were transformed into E. coli, isolated, and digested with restriction enzymes to release the FRA16B insert. Replication products were analyzed on 1.5% agarose gels to score insertions and deletions. As a control, replication constructs were directly transformed into E. coli, without replication in human cells, to account

for background instability in E. coli. (continued)

56

C

E

D

F Mutations generated by replication of clone 17 with SV40 ori in HEK293T cells

Mutant Type of mutation

Size of mutation (bp)

Distance (bp) from SV40 ori

17-4 Deletion 378 58

17-8 Deletion 99 364

17-18 Deletion 165 207

17-34 Deletion 224 406

17-48 Insertion 33 325

Figure 3.3 Continued.

(C) Replication efficiency of FRA16B constructs in HEK293T cells. The replication efficiency of each

FRA16B-containing construct was determined by the ratio of replicated FRA16B DNA to the control (pGEM-SV40ori). Results are shown as the mean ± SD from six individual experiments. (D) Identification of mutation events of replication in HEK293T cells using gel electrophoresis. A

representative agarose gel shows individual HEK293T cell replication products from clone 17. Replication products (#4 and #8) exhibit large base pair deletions, as demonstrated by faster migrating products (asterisks) compared to unreplicated DNA (designated as C). M, molecular weight marker. (E)

Mutation analysis of FRA16B constructs with different replication orientations and distances from the SV40 ori. After replication in HEK293T cells, purified SV40 DNA was digested to eliminate any

unreplicated parental templates and transformed into E. coli. Approximately 60 single colonies from each clone were picked at random, and the DNAs were isolated and digested with restriction enzymes to release the FRA16B insert. Samples were then electrophoresed in agarose gels to compare to unreplicated

DNA and scored for insertion or deletion events (black bars). As a control, DNAs that were not transfected into HEK293T cells were also transformed into E. coli and scored for background instability in E. coli (white bars). The fractions within each bar show the number of molecules mutated over the

total number of molecules examined. FRA16B instability is displayed as a percentage, and the schematic drawings above each data set show the relative location of the SV40 ori (open circles) and the replication

orientation (arrows) of the FRA16B sequence (black boxes). Clone 17 demonstrates a statistically significant increase in instability events in HEK293T cells relative to background instability (*P = 0.022), and to the control (**P = 0.016). (F) Analysis of mutations generated by replication of clone 17

with SV40 ori in HEK293T cells. For the five mutants, the type of mutation, size of the mutation (in bp) and distance (in bp) from SV40 ori are reported in the table.

57

Replication products from transfected HEK293T cells were digested with DpnI and

transformed into E. coli (Figure 3.3B). Only DpnI-resistant material yielded colonies,

from which DNAs were isolated and digested with restriction enzymes to release the

FRA16B insert. FRA16B inserts were resolved on agarose gels and scored for insertions

or deletions (Figure 3.3D). Due to the resolution of agarose gels, mutations with only

large size changes (≥50 bp) were detected. The mutation frequency for each construct is

shown in Figure 3.3E. Interestingly, clone 17, which displayed the greatest reduction in

replication efficiency, also demonstrated a statistically significant increase in instability

events compared to the background instability in E. coli (P = 0.022) and the control

construct without FRA16B DNA in HEK293T cells (P = 0.016). Replication of clone 17

in HEK293T cells resulted in the highest mutation rate (9%) among all of the FRA16B

constructs, with four deletions (ranging from 99 to 378 bp) and one expansion (33 bp)

(Figure 3.3F), which were confirmed by DNA sequencing. The 378-bp deletion in mutant

17-4 and the 165-bp deletion in mutant 17-18 begin, respectively, at nt 58 and nt 207 of

the lagging strand template of the FRA16B sequence, located at the first Okazaki

initiation zone (OIZ). The 99-bp deletion in mutant 17-8 and the 224-bp deletion in

mutant 17-34 begin, respectively, at nt 364 and nt 406 of the lagging strand template of

the FRA16B sequence, located at the second OIZ. The 33-bp insertion in mutant 17-48

begins at nt 325, and is one perfect copy of the 33-bp FRA16B repeat. Mapping of the

mutation sites to the most energetically favorable predicted DNA secondary structures of

the lagging strand template of the FRA16B sequence, as determined by Mfold [29]

suggests that the deleted regions may correspond to sites of extensive secondary structure

formation, including multiple hairpins (Figure 3.4).

58

DNA polymerase stalling at FRA16B during DNA synthesis

Evidence has shown that the formation of secondary structures by CGG repeats can block

replication within rare, folate-sensitive fragile sites [32, 33], and that replication forks

Figure 3.4: Mapping mutation sites to hairpin regions of predicted DNA secondary structure. The FRA16B sequence of the lagging strand template of clone 17 was used as input for Mfold, and the most energetically favorable predicted secondary structure is shown. Deleted regions within

replication products (17-4, 17-8, 17-18, and 17-34) are highlighted in yellow. The position of the 33 bp insertion within 17-48 is indicated by an arrow, and the inserted sequence is shown.

59

have a high tendency to regress while moving through difficult-to-replicate DNA

sequences [16, 161]. To further investigate these possible mechanisms of instability, we

constructed replication fork templates pFRA16B37, a plasmid containing FRA16B that

mimicked clone 17, and pGLGAP, a control containing non-fragile site DNA (Figure

3.6A). Following DNA synthesis by Klenow fragment, circular pFRA16B37 and

pGLGAP molecules were examined by EM (Figure 3.6C, top panels). We found that

73% of pGLGAP molecules had double-stranded tails, indicating that they underwent

completed DNA synthesis, whereas 24% did not (Figure 3.6B). In contrast, only 10% of

pFRA16B37 molecules completed DNA synthesis and had double-stranded tails.

Moreover, a majority of pFRA16B37 molecules (73%) had no detectable DNA synthesis

(i.e. no double-stranded tails) and contained polymerase „stuck‟ to the DNA, even after

phenol/chloroform extraction (Figure 3.6C). To locate the FRA16B sequence, the

samples were further linearized with restriction enzymes to generate asymmetric arms

(Figure 3.6C, bottom right panel). We observed that most (68%) of the polymerase bound

AGCTTGCATGCCTGCAGGC^TGAGGAGATATAATATATATTAGATATATATATTATTAGATATAATGTATA

TTATATATAATATATTATTAGATATAATATATAATATATATTATTAGATATAATATATAATATATATAATTAGA

TATAATATATAATATATTATATATATTATTAGATATAATATATTATATATATTATTAGATATAATATATAATAT

ATTATATATATTATTAGATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATA

TATTATTAGATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATATATTATTA

GATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATATATTATTAGATATAAT

ATATAATATATTAGATATATTATTAGATATAATATATAATATATTAGATATAATAATATATTATATCTAATAT

ACATTATTATATCTAATATATATTATATTAG

AGCTTGCATGCCTGCAGGC^TGAGGAGATATAATATATATTAGATATATATATTATTAGATATAATGTATA

TTATATATAATATATTATTAGATATAATATATAATATATATTATTAGATATAATATATAATATATATAATTAGA

TATAATATATAATATATTATATATATTATTAGATATAATATATTATATATATTATTAGATATAATATATAATAT

ATTATATATATTATTAGATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATA

TATTATTAGATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATATATTATTA

GATATAATATATAATATATTATATATATTATTAGATATAATATATAATATATTATATATATTATTAGATATAAT

ATATAATATATTAGATATATTATTAGATATAATATATAATATATTAGATATAATAATATATTATATCTAATAT

ACATTATTATATCTAATATATATTATATTAG Figure 3.5: FRA16B DNA-containing sequence of pFRA16B37. The Nb.BbvCI recognition site

is highlighted in bold, and the nicking site is indicated by an arrow. Strand displacement was carried out using Klenow fragment and dATPs, dTTPs, and dGTPs, causing DNA synthesis to stop when the

first cytosine (boxed) was encountered (the first guanine on the template strand). DNA synthesis o f the displaced single-strand tail was carried out by annealing the oligonucleotide 5′-ATATAATATATTATTATATCTAATA-3‟ to the underlined sequence, in the presence of Klenow

fragment, dATPs, dTTPs, and dCTPs, to generate the complimentary strand. The italicized sequence represents the FRA16B fragment of clone 17 (nucleotides 273-792 from the SV40 ori).

60

A

B

Classification of circular DNA molecules after Klenow Fragment reaction

DNA with tail

Protein-bound DNA

DNA without tail

pGLGAP 73% 3% 24%

pFRA16B37 10% 73% 17%

C

Figure 3.6: FRA16B DNA synthesis by Klenow fragment of E. coli DNA polymerase I. (A)

Construction of replication fork templates to examine fork regression. Plasmids containing FRA16B clone 17 fragment (pFRA16B37, Figure 3.5) or a G-less cassette (pGLGAP) were constructed, nicked with Nb.BbvCI at a site immediately upstream of the inserts (bold), and incubated with Klenow

fragment to generate a single-strand tail by strand displacement. The single-strand tail was converted into a double-strand tail by annealing a complimentary oligonucleotide at the 3‟ end of the displaced strand and further incubation with Klenow fragment. To map the location of the double-strand tail,

plasmids were linearized so measurements of the two asymmetrical tails could be obtained to determine the amount of fork regression. (B) Classification of circular DNA molecules after Klenow fragment

reaction by EM. After the second Klenow reaction to create a double-strand tail, reaction mixtures were directly mounted onto carbon-coated copper EM grids and rotary shadowcasted with tungsten. At least 100 molecules from each sample were examined to determine the percentage of molecules with tails,

protein-bound, or without a tail. (C) Visualization of circular and linear pGLGAP and pFRA16B37 DNAs after synthesis reaction with Klenow fragment. Representative images were montaged and are shown in reverse contrast. (continued)

61

molecules had polymerases located within the FRA16B sequence, which is located at 33-

50% of the total DNA length from the nearest end, suggesting that the majority of the

D

0

5

10

15

20

25

30

35

40

0-5 5-10 10-15 15-20 20-25 25-30 30-35 35-40 40-45 45-50

Location of polymerase (% of total length)

% o

f M

ole

cu

les

F

1822 bp (46%)

487 bp

2132 bp

pFRA16B37

1335 bp

(34%)

2619 bp

AhdI

Nb.BbvCI

Non-regressed Partially regressed Fully regressed

E

A

T

T

A

T

A

T

A

T

T

A

T

Figure 3.6 Continued. (D) Location of bound polymerases within FRA16B replication fork templates. The location of the

polymerase as a percentage of the total length is plotted as a histogram for all FRA16B molecules that displayed bound polymerase and did not contain a double-stranded tail. The FRA16B fragment is located at 33–50% of the total length from the nearest end. (E) Analysis of DNA polymerase pause sites during

lagging strand DNA synthesis. Radiolabeled oligomers were annealed to the displaced lagging strand of pGLGAP (lanes 1–3) and pFRA16B37 (lanes 5–7) replication fork templates, and synthesis was carried out with 0.5 U (lanes 1 and 5), 5 U (lanes 2 and 6) or 15 U (lanes 3 and 7) of Klenow fragment. Lane 4

(C) contained the radiolabeled 25-nt oligonucleotide only. The first 12 nt of the newly synthesized complimentary strand are indicated, and the pause sites are boxed. (F) Detection of FRA16B replication

fork regression. Circular replication fork molecules were linearized with AhdI to produce asymmetrical arms. The location of the double-strand tail was defined as the percentage of total DNA length from the nearest end.

62

polymerase was stalled at the FRA16B region and was not able to proceed with DNA

synthesis (Figure 3.6D).

Because of the high occurrence of polymerase stalling within FRA16B, synthesis

products were analyzed on DNA sequencing gels to examine the progression of the

polymerase during DNA synthesis of the lagging strand, and to locate polymerase pause

sites. We found that FRA16B DNA displayed strong pause sites, specifically at

nucleotides 8, 9 and 11 of the newly synthesized strand, and was not able to complete the

synthesis, while the control DNA exhibited unperturbed synthesis and generated its full-

length product of 400 nt (Figure 3.6E). These results are in agreement with our

observation of polymerase-trapped molecules by EM, and suggest that FRA16B DNA

inhibits the progression of DNA polymerase, possibly due to its ability to form secondary

structure.

Next, we analyzed fork regression with the FRA16B molecules containing

double-stranded tails. These molecules were a small fraction (10 %) of the total

population but could represent a fully regressed, non-regressed, or partially regressed

replication fork, depending on the location of the double-stranded tails. The location of

the double-strand tail was indicated as the percentage of total DNA length from the

nearest end. For the FRA16B template, if no regression occurred, the tail would be

located at 46% of the total length, whereas under complete regression, templates would

have the tail located at 34% of the total length (Figure 3.6F). FRA16B forks were

considered to have regressed when the double-stranded tails were located ≤40% of the

63

total length, which is halfway between non- and fully regressed. Out of the 32 molecules

analyzed, 26 (81%) underwent fork regression, whereas six (19%) did not display fork

reversal. As a control, pGLGAP templates were examined (n = 81), and our results were

consistent with previous studies, demonstrating that ~10-20% of random, non-repetitive

DNAs undergo some degree of spontaneous fork regression [157, 162]. These results

show that fragile site-containing replication forks have a greater tendency to undergo fork

reversal than non-fragile site sequences.

We also found that even in non-regressed molecules which underwent DNA

synthesis, the average length of FRA16B tails was only 21% of the full length, whereas

the average length of the control pGLGAP template tails was 85% of the total length.

These results suggest that the shorter double-stranded tails generated by DNA synthesis

of FRA16B templates could result from bypass of a slipped-out secondary structure

formed on the lagging strand template, leading to the generation of deletion mutations, as

we observed following FRA16B replication in human cells. Together, these data suggest

that polymerase pausing, fork reversal, or polymerase skipping at FRA16B could

contribute to its instability.

DISCUSSION

To gain a better understanding of the mechanism of instability at chromosomal fragile

sites, we examined the ability of FRA16B to form a secondary structure in vitro and

evaluated the effects of cis-acting factors on replication of FRA16B DNA. Previously, it

has been shown that the CGG repeat, which underlies the basis of fragility at rare, folate-

64

sensitive fragile sites, is able to form a stable secondary structure [30, 31] that presents

significant difficulties during replication [32, 33]. However, the ability of an AT-rich

fragile DNA to form a secondary structure in vitro has not been demonstrated until now.

Our data show that FRA16B DNA forms an alternative structure following denaturation

and re-annealing. To our knowledge, this is the first experimental evidence of an AT-rich

fragile DNA, which represents the majority of fragile sites, forming a secondary structure

in vitro. This observation supports the concept that formation of highly stable secondary

structures could be a general mechanism that contributes to fragile site breakage during

DNA replication [37].

Since prior reports have indicated potential problems with replication due to DNA

secondary structures, FRA16B replication in human cells was also examined. We

determined that, similar to previous results, cis-acting factors including replication

orientation and distance from the origin do play a role in both replication efficiency and

instability of FRA16B-containing constructs in human cells. Numerous studies have

suggested that replication orientation, which determines leading and lagging strand

synthesis, has a significant effect on replication efficiency as well as both the type and

frequency of mutations [156, 158-160]. In general, an increase in the number of

mutations is observed when the more structure-prone strand serves as the lagging strand

template. Thus, when the strand with the most potential to form a stable secondary

structure serves as the lagging strand template, the DNA is more likely to fold into an

alternative structure within the regions that maintain single-strandedness. Consistent with

previous studies, we found that orientation greatly affects replication efficiency in that

65

clone 17 possessed a markedly lower efficiency than clone 15, yet they differed only in

orientation. Clone 17 also displayed a higher frequency of mutations than clone 15 during

replication in HEK293T cells. Although both strands of FRA16B are able to form

secondary structure, as shown in Figure 3.1A, we found that there are clearly different

structures being formed by each strand, based upon their different electrophoretic

mobilities (Figure 3.2). The more rapid mobility of the template strand in Figure 3.2

supports the idea that the lagging strand template in the unstable orientation forms a more

stable secondary structure than the template strand in the stable orientation. This data

prompts us to propose that one structure may be more „deleterious‟ to DNA replication

than the other, thus leading to an orientation effect similar to what has been observed

when only one strand forms a stable alternative structure. Although significant

differences were not observed among the other sets of FRA16B constructs (clones 31 and

34, clones 10 and 12, and clones 46 and 39), this could be due to the fact that in these

constructs, FRA16B DNA is at least 300 bp away from the origin, and orientation may

only be a factor when the fragile site is in close proximity to the origin. This is likely due

to the location of secondary structures within the Okazaki initiation zone (OIZ), a region

of ~300 nt that remains single-stranded. Placing the beginning of the 921 bp FRA16B

DNA fragment 30 bp from the SV40 ori, as was the case for clones 15 and 17, would

have FRA16B occupy the first OIZ, promoting the formation of an alternative DNA

structure. The other sets of FRA16B constructs have the start of the FRA16B sequence

located at the second or the third OIZs. The placement of the beginning of the FRA16B

sequence in different OIZs might affect the ability of FRA16B to form stable secondary

structure, due to the binding cooperativity of RPA protein [163]. The binding of RPA in

66

the first OIZ could propagate to the second and third OIZs, and might prevent the

formation of alternative structure by FRA16B, when FRA16B sequence begins at the

second and third OIZs. In contrast, clone 17 locates in the first OIZ, and its ability to

form alternative structure might trump the initial binding of RPA protein. Consequently,

both the orientation and location of FRA16B within clone 17 played a significant role in

reducing DNA replication efficiency and increasing the number of mutations.

Furthermore, the large base pair deletions that occurred within FRA16B during

replication in human cells recapitulate what has been observed at fragile sites in

numerous tumors in vivo. The most energetically favorable predicted secondary structure

of the lagging strand template of clone 17 suggested that the deleted regions may occur at

sites of extensive secondary structure formation, including multiple hairpins (Figure 3.4),

further supporting a potential role for secondary structure at sites of DNA breakage

within fragile sites. Possible explanations for the increase in mutation frequency and

decrease in replication efficiency observed for the FRA16B constructs in the SV40

replication system are: (i) the formation of a secondary structure close to the origin

hinders loading of the replication machinery, leading to decreased replication efficiency,

and/or (ii) secondary structure formation within single-stranded DNA regions during

replication causes insertions via replication restart, or deletions following bypass of the

structure or cleavage of a regressed fork- all of which are consequences of replication

fork stalling. Since clone 17, which is located 30 bp from the SV40 ori, exhibited a

statistically significant decrease in replication efficiency, it is conceivable that inhibition

of origin firing is contributing to what is observed. In contrast, clones 31, 34, 10, 12, 46

and 39, which all place FRA16B at least 300 bp from the origin of replication, do not

67

demonstrate a significant reduction in replication efficiency, and this suggests replication

may initiate unperturbed. However, the increased frequency of instability events for clone

17 following replication in HEK293T cells supports replication fork stalling, which is a

consequence of DNA elongation. Analysis of all five mutants derived from clone 17

demonstrates that sites of deletions and expansions occurred as much as ~400 bp from the

origin of replication (Figure 3.3F), suggesting that DNA replication was initiated but

paused at a site located at a greater distance from the origin. A similar study, which

examined the ability of the CCTG repeat to cause mutations, reports comparable results,

demonstrating that when the repeat is located ~25 bp from the SV40 ori, an increase in

expansions and deletions is observed, also suggesting replication fork stalling during

DNA synthesis [158].

To dissect the mechanism for the generation of such mutations within FRA16B

during replication, we constructed a replication fork template mimicking clone 17 with

FRA16B DNA ~30 bp from the initial replication fork and the same replication

orientation as clone 17. Visualization of FRA16B replication fork molecules by EM

showed that the majority of DNAs contained polymerases „stuck‟ at the FRA16B region,

and analysis of the lagging strand synthesis confirmed that the polymerase had stalled at

specific sites within the FRA16B tract. As these polymerase-trapped molecules were not

observed with the non-fragile DNA control template (Figure 3.6B), we suggest that the

formation of secondary structure by FRA16B DNA may be responsible for polymerase

pausing. Further, examination of replication fork templates containing either the

telomeric or CTG repeats, which have been shown to form hairpin structures , did not

68

reveal polymerase pausing within the repetitive DNAs by the same EM analysis [164],

although Oshima and Wells have used DNA sequencing gels to demonstrate polymerase

pausing by CTG repeats [165]. These results suggest the secondary structure formed

within FRA16B is likely more extensive than a single hairpin and highly stable. A highly

stable replication fork with exposed single-stranded regions can trigger the ATR-

dependent DNA damage checkpoint pathway, which in turn inhibits further firing of

replication origins, blocks entry into mitosis and promotes DNA repair, recombination or

apoptosis [38, 39]. However, a deficiency of proteins in the ATR-dependent cell cycle

checkpoint pathway will dramatically increase fragile site breakage, as several studies

have reported [37, 46, 47, 58]. It is important to note that polymerase stalling at sites of

secondary structure is probably only likely when the FRA16B sequence is located very

close to the origin of replication, and our results are limited due to the fact that only one

construct was analyzed by DNA sequencing gels, which contained FRA16B located 30

bp from the replication origin.

This study is the first to provide evidence of spontaneous replication fork reversal

within a fragile site sequence during DNA synthesis. Based on this information, we

propose a model whereby FRA16B instability primarily arises from polymerase stalling

caused by the formation of secondary structure at the fragile site region (Figure 3.7B),

coupled with the failure of the ATR checkpoint pathway. Other mechanisms contributing

to fragile site instability could be due to replication fork regression (Figure 3.7C), the

formation of stable DNA secondary structures on the lagging strand template leading to

polymerase skipping (Figure 3.7D) or replication restart with secondary structure formed

69

on the newly synthesized strand (Figure 3.7E). These mechanisms may be shared among

other fragile DNA sequences. Overall, our results provide insight into the mechanism of

fragile site instability by demonstrating the ability of a fragile DNA to form a stable

secondary structure that affects replication efficiency and instability, and causes

polymerase pausing during DNA synthesis.

ACKNOWLEDGEMENTS

We thank Dr. Jack Griffith (UNC-CH) for providing pGLGAP plasmid.

Figure 3.7: Proposed models of FRA16B instability. (A) Replication fork demonstrating DNA polymerase (gray circle) entering the FRA16B sequence (bold). (B) During FRA16B replication, the polymerase has a high tendency to pause, likely at regions of secondary structure formation.

Consequently, DNA strand breakage, deletions or insertions may arise. (C) FRA16B may promote spontaneous fork reversal, leading to a four-stranded chickenfoot intermediate. (D) FRA16B may form an extensive secondary structure on a template strand or (E) a newly synthesized strand,

leading to deletions or insertions, respectively.

70

FUNDING

This National Cancer Institute (CA85826 and CA113863 to Y.-H. W).

71

CHAPTER IV: CBFB AND MYH11 IN INV(16)(P13Q22) OF ACUTE MYELOID

LEUKEMIA DISPLAY CLOSE SPATIAL PROXIMITY IN INTERPHASE

NUCLEI OF HUMAN HEMATOPOIETIC STEM CELLS

Allison B. Weckerle

1, Madhumita Santra

2, Maggie C.Y. Ng

3,4, Patrick P. Koty

2, and Yuh-

Hwa Wang1*

1Department of Biochemistry, Wake Forest University School of Medicine, Medical

Center Boulevard, Winston-Salem, NC 27157-1016, USA. 2Section on

Medical Genetics, Department of Pediatrics, Wake Forest University School

of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1016, USA. 3Center for Genomics and Personalized Medicine Research, Wake Forest University

School of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1016, USA. 4Center for Diabetes Research, Wake Forest University School of Medicine, Medical

Center Boulevard, Winston-Salem, NC 27157-1016, USA.

*Correspondence to: Dr. Yuh-Hwa Wang, Department of Biochemistry, Wake Forest

University School of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1016, USA. E-mail: [email protected]. Phone: (336) 716-6186. Fax: (336) 716-7671.

Supported by: grant NIH R01 CA085826 to Y.-H.W.

The following paper is in press in Genes, Chromosomes and Cancer, May 2011, with modifications in numbering of Figures and References.

72

ABSTRACT

To gain a better understanding of the mechanism of chromosomal translocations in

cancer, we investigated the spatial proximity between CBFB and MYH11 genes involved

in inv(16)(p13q22) found in acute myeloid leukemia patients. Previous studies have

demonstrated a role for spatial genome organization in the formation of tumorigenic

abnormalities. The non-random localization of chromosomes and, more specifically, of

genes appears to play a role in the mechanism of chromosomal translocations. Here, two-

color fluorescence in situ hybridization and confocal microscopy were used to measure

the interphase distance between CBFB and MYH11 in hematopoietic stem cells, where

inv(16)(p13q22) is believed to occur, leading to leukemia development. The measured

distances in hematopoietic stem cells were compared to mesenchymal stem cells ,

peripheral blood lymphocytes and fibroblasts, as spatial genome organization is

determined to be cell-type specific. Results indicate that CBFB and MYH11 are

significantly closer in hematopoietic stem cells compared to all other cell types

examined. Furthermore, the CBFB-MYH11 distance is significantly reduced compared to

CBFB and a control locus in hematopoietic stem cells, although separation between

CBFB and the control is ~70% of that between CBFB and MYH11 on metaphase

chromosomes. Hematopoietic stem cells were also treated with fragile site-inducing

chemicals since both genes contain translocation breakpoints within these regions.

However, treatment with fragile site-inducing chemicals did not significantly affect the

interphase distance. Consistent with previous studies, our results suggest that gene

proximity may play a role in the formation of cancer-causing rearrangements, providing

insight into the mechanism of chromosomal abnormalities in human tumors.

73

INTRODUCTION

Chromosomal translocations often result in cancer development through disruption of

genes involved in important cellular processes such as cell proliferation and survival.

Despite the high prevalence of chromosomal trans locations in tumor cells, the mechanism

by which this process occurs is still unclear. The formation of translocations is a multi-

step process that is initiated by DNA strand breakage [166]. DNA double-strand breaks

(DSBs), which often occur through exposure to ionizing radiation or genotoxic chemicals

[60] must be present in at least two places in the genome. There are several major

pathways that contribute to DSB repair: homologous recombination [136, 167], non-

homologous end joining [56], and microhomology-mediated end joining [59]. In order

for a translocation to occur, these pathways must fail to repair the breaks, and the broken

chromosome ends must physically meet and become joined together [136, 167]. The

molecular basis underlying why certain genes undergo specific chromosomal

translocations remains elusive. However, their recurrence in human tumors suggests a

role for spatial genome organization in the formation of cancer-causing rearrangements.

In an increasing number of studies that have examined potential mechanisms,

non-random spatial genome organization has emerged as an important factor in the

generation of chromosomal abnormalities. Interphase nuclei are compartmentalized into

discrete, three-dimensional regions, known as chromosome territories, according to gene

density, whereby gene-rich chromosomes tend to cluster toward the nuclear center while

chromosomes that are gene-poor preferentially locate toward the periphery [166]. The

position of genes is non-random with respect to each other, which is functionally

74

important for understanding cancerous transformation. In addition to human cells, higher-

order genome organization is also observed in mice and other vertebrates, which supports

an important role for the spatial arrangement of chromosomes and genes in interphase

nuclei.

The concept that the spatial proximity of chromosomes may lead to an enhanced

susceptibility to translocations was recently demonstrated. The nuclear architecture of

bone marrow cells was examined and revealed that chromosomes 8 and 21 tended to co-

localize with one another in the nucleus rather than be separated, thus promoting the

t(8;21) observed in acute myeloid leukemia (AML) of the M2 subtype [168]. In 2003,

Roix et al. examined the spatial proximity between the MYC oncogene and three of its

translocation partner genes, IGH, IGL and IGK, taking advantage of the fact that in

Burkitt‟s lymphoma, MYC translocates with each partner at different frequencies. The

distances between each gene set were measured and compared to their c linically observed

frequencies, with results indicating a strong correlation between the spatial proximity of

genes during interphase and their translocation frequencies [169]. In addition, a number

of studies have looked at the spatial proximity of genes involved in translocations

observed in thyroid cancer, since there is a well-established association between DNA

damage caused by radiation exposure and the occurrence of thyroid tumors. Examination

of the spatial proximity between RET, a gene commonly rearranged in papillary thyroid

carcinoma (PTC), and two of its partner genes, CCDC6 [170] and NCOA4 [171],

revealed close proximity in both gene pairs that was cell-type specific. Furthermore,

Roccato et al. determined that the interphase distance between TPR and NTRK1 genes,

75

separated by ~30 Mb and also rearranged in PTC, was significantly closer in human

thyrocytes than in lymphocytes, potentially favoring a rearrangement [172].

More recent reports have elegantly demonstrated the ability of genes located in

close spatial proximity to give rise to a number of cancer-specific translocations

following DNA damage, providing compelling evidence of a role for spatial genome

organization in the formation of chromosomal translocations. Gandhi et al. showed that

the RET/PTC1 rearrangement, which involves closely-positioned genes RET and CCDC6

[170], could be generated in normal human thyroid cells upon fragile site breakage [173].

NPM1 and ALK, genes participating in t(2;5)(p23;q35) associated with anaplastic large

cell lymphoma (ALCL), are located in close spatial proximity within the nuclear space of

t(2;5)-negative ALCL cells, and translocate with each other upon irradiation [174].

Similar results have been shown for translocations involving TMPRSS2 with two of its

partner genes, ERG [175, 176] and ETV1 [176], whereby irradiation of LNCaP cells

induced both fusion transcripts, and represented the authentic fusion junctions commonly

observed in prostate cancer patients.

According to the Mitelman Database of Chromosome Aberrations and Gene

Fusions in Cancer [177], there are over 700 chromosomal translocations reported in

various types of human tumors. To date, most studies of spatial gene proximity have

focused on solid tumors, including PTC [170-172], lymphoma [169, 174] and prostate

cancer [175, 176]. Fewer studies have investigated the spatial proximity of genes

involved in translocations observed in leukemia, with the exception of the Philadelphia

76

chromosome translocation commonly found in patients with chronic myelogenous

leukemia. The results support the concept that participating genes ABL and BCR are

closely positioned during interphase, thus promoting their rearrangement [178-180]. To

our knowledge, there has been no such study examining the interphase distance of genes

participating in an intrachromosomal translocation associated with leukemia. Therefore,

the objective of this study was to determine whether short interphase distances between

translocation-participating loci involved in leukemia-specific intrachromosomal

rearrangements could further support the role of spatial genome organization in the

formation of cancer-causing rearrangements.

The intrachromosomal rearrangement inv(16)(p13q22) is one of the most

common abnormalities observed in AML, occurring in ~12% of adult patients [106] and

100% of patients of the M4 subtype with accompanying eosinophilia (M4Eo) [121]. This

rearrangement involves the CBFB and MYH11 genes, both of which are located on

chromosome 16 at ~50 Mb apart, which is further apart than any gene pair formerly

investigated for spatial proximity of intrachromosomal rearrangements. CBFβ is part of

the core binding factor (CBF) complex [106], a heterodimeric transcription factor

comprised of CBFβ and core binding factor alpha (CBFα) subunits that regulate a variety

of genes involved in hematopoiesis [107]. The fusion product of inv(16)(p13q22) blocks

hematopoietic differentiation in a dominant-negative manner by inhibiting the normal

function of the CBF transcription complex [113, 114]. In addition, breakpoints within

both CBFB and MYH11 coincide with human chromosomal fragile sites

FRA16B/FRA16C and FRA16A, respectively [22]. Fragile sites are specific regions of

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the genome that are highly susceptible to DNA breakage following conditions of partial

replication stress [1]. These regions are often targets of environmental and chemical

agents, including ethanol, cigarette smoke, caffeine and pesticides [1], and play a direct

role in the generation of oncogenic RET/PTC rearrangements [173].

To determine whether spatial gene proximity is involved in the formation of this

rearrangement, the distance between CBFB and MYH11 loci was examined in interphase

nuclei of normal human hematopoietic stem cells (HSCs). Two-color fluorescence in situ

hybridization (FISH) and confocal microscopy were used to map the locations of CBFB

and MYH11. The distances were compared to cell types of different lineages and

differentiation stages, since previous studies have shown that spatial gene proximity may

be cell-lineage and cell differentiation stage-specific [180]. We found significant

differences for CBFB-MYH11 measured distances in HSCs compared with human

mesenchymal stem cells (MSCs), peripheral blood lymphocytes (PBLs) and fibroblasts.

Furthermore, the distance between CBFB and MYH11, which are ~50 Mb apart on

chromosome 16 was determined to be significantly reduced during interphase in HSCs

compared to CBFB and the control 16p11.2 locus, which is separated from CBFB by ~37

Mb. We also examined the spatial proximity between CBFB and MYH11 following

treatment with fragile site-inducing chemicals to determine any potential changes in their

interphase distance. Treatment of HSCs did not cause any significant differences

compared to untreated HSCs, indicating that mild replication stress and/or fragile site

breakage had little effect on CBFB-MYH11 spatial proximity. Here, we present evidence

of a role for interphase gene proximity in the formation of inv(16)(p13q22) whereby

78

CBFB and MYH11 are located at a significantly reduced distance in normal human HSCs

compared to other cell types. These results further support a critical role for non-random

spatial genome organization in the generation of cancer-specific chromosomal

rearrangements.


Cell Culture

Fresh primary human CD34+ HSCs were purchased from Lonza (Walkersville, MD) and

maintained in HPGM hematopoietic progenitor growth medium (Lonza) containing a

StemSpan CC100 cytokine cocktail of recombinant human (rh) Flt-3 ligand, rh stem cell

factor, rh IL-3 and rh IL-6 (STEMCELL Technologies, Vancouver, British Columbia).

Human MSCs were obtained from Texas A&M University (College Station, TX) and

grown in Dulbecco‟s Modified Eagle Media (Invitrogen, Carlsbad, CA) supplemented

with 10% fetal bovine serum (Thermo Fisher Scientific, Waltham, MA) and 1%

penicillin-streptomycin (Invitrogen). PBLs from a healthy donor were established

according to routine procedures (Department of Medical Cytogenetics, WFUBMC). The

normal human fibroblast cell line GM08402 was obtained from Coriell Institute for

Medical Research (Camden, NJ) and maintained in Minimum Essential Media

(Invitrogen) with 10% fetal bovine serum (ThermoScientific) and 1% penicillin-

streptomycin (Invitrogen). All cells were grown in a humidified incubator at 37˚C with

5% CO2. For the induction of fragile sites, cells were treated with 0.4 μM APH (Sigma-

Aldrich, St. Louis, MO), 2 mM 2-AP (Sigma-Aldrich) and 150 mg/L berenil (Sigma-

Aldrich) for 24 h.

79

FISH

Cells were harvested according to a standard cytogenetic procedure. Colcemid was

directly added to cells for 30 min. After centrifugation, 0.075 M KCl hypotonic solution

was added and cells were suspended in 3:1 methanol/acetic acid, dropped onto slides and

pretreated with 0.001% pepsin (Sigma, St. Louis MO) in 0.01M HCl for 10 min at 37°C

followed by a rinse in PBS (Sigma) and a 5 min fixation in 1% formaldehyde. Slides

were dehydrated through an ethanol series and dried. Bacterial artificial chromosome

(BAC) clones spanning the CBFB (RP11-5A19) and MYH11 (RP11-585P8)

chromosomal loci to be used as probes were purchased from Children‟s Hospital Oakland

Research Institute (CHORI, Oakland, CA). The CBFB probe was labeled with

SpectrumOrange - dUTP (Abbott Molecular, Abbott Park, IL) and MYH11 probe was

labeled with SpectrumGreen - dUTP (Abbott Molecular) according to instructions in the

Nick Translation Kit (Abbott Molecular). As a control, a probe spanning the 16p11.2

locus (RP11-74E23) was purchased pre-labeled with SpectrumGreen from BlueGnome

(BlueGnome Ltd., Great Shelford, Cambridge, UK). A hybridization mix of each probe

(300 ng) was prepared by adding 7 μL of Abbott Molecular LSI Hybridization mix and 3

μL of distilled water to the BAC pellets. The hybridization mixture was applied to the

slides seated in a Hybrite heating block set to 37°C, sealed with a coverslip and denatured

at 73°C for 3 min. After overnight incubation at 37°C, slides were washed in 0.4X

Sodium Citrate Saline (SSC)/0.3% NP-40 at 73°C for 1.5 min, followed by a brief rinse

through 2X SSC/0.1% NP-40 and then dried.

80

Slides were counterstained with DAPI II (Abbott Molecular) and cover-slipped,

and fluorescence was viewed using a Zeiss Axioplan 2 microscope with a triple filter to

simultaneously show the probes and counterstain. In order to illustrate the presence of

signals, fluorescent images of probe hybridization were made using the Cytovision

capture system equipped with a CCD camera (Photometrics). The fluors were captured in

sequential order as follows: SpectrumOrange (excitation 500-600 nm/emission 550-650

nm), SpectrumGreen (excitation 400-550 nm/emission 500-600 nm) and then DAPI II

(excitation/emission 400 nm). The images were captured in gray scale, merged and

pseudo-colored by the software to resemble the color of the original signals.

For the analysis of CBFB and the control locus, slides containing HSCs were

stripped and re-probed with CBFB and 16p11.2 BACs according to the same procedure.

Confocal Microscopy

A total of 105 interphase nuclei from each cell type were examined using a Zeiss

Axiovert 100M confocal microscope. Each nucleus displays two distinct hybridization

signals per probe (Figure 4.1), and therefore, a total of 210 pairs of CBFB and MYH11

signals were then analyzed for each cell type. Since previous studies have determined that

two pairs of heterologous signals are located in two separate areas of the nucleus with

shorter distances than any other possible distance between the signals [170], it was

assumed that the two shorter distances were between loci on the same chromosome.

Distances between hybridization signals were measured using Zeiss LSM Image Browser

Software‟s overlay feature.

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Statistical Analysis

All statistical analyses were performed using IBM SPSS Statistics v19 software (SPSS

Inc., Chicago, IL). Data are presented as the mean ± standard deviation (SD) or as a

percentage. The distribution of measured distances between HSCs and other cell types

were compared using student‟s t-test. Student‟s t-test was also utilized to determine

significant differences between CBFB-MYH11 and CBFB-16p11.2 distances. A chi-

square test was used to compare the percentage of gene pairs in which CBFB and MYH11

were located 0-0.99 μm, 1.00-1.99 μm, 2.00-2.99 μm, 3.00-3.99 μm or ≥4 μm apart in

HSCs and other cell types. A p value < 0.05 was considered statistically significant and

was adjusted by a Bonferroni correction for multiple comparisons.

RESULTS

Interphase Distance of CBFB and MYH11 in Human Hematopoietic Stem Cells

Compared with Human Mesenchymal Stem Cells

The inv(16)(p13q22) is believed to occur very early in differentiation at the

stem/progenitor level in HSCs [181]. Therefore, to investigate whether CBFB and

MYH11 display close spatial proximity in HSCs, interphase distances between CBFB and

MYH11 loci were measured in HSCs and compared to those in MSCs. MSCs were

examined since they are also found in the bone marrow and have the potential for multi-

lineage differentiation and self-renewal but give rise to different cell types. The

distribution plots of measured CBFB-MYH11 distances in HSCs and MSCs are shown in

Figure 4.2. The shorter distance (mean ± SD) for HSCs (1.90 ± 1.14 m) as compared to

MSCs (3.78 ± 2.67 m) was highly significant (p = 2.24x10-18

) (Table 4.1). To estimate

the range of distance that may be related to translocation events, the measured distances

82

between CBFB and MYH11 were categorized into 0-0.99 μm, 1.00-1.99 μm, 2.00-2.99

μm, 3.00-3.99 μm and ≥4 μm groups, respectively (Figure 4.3). The overall distribution

of interphase distance between these two stem cell types was significantly different (p =

7.27x10-16

). Interestingly, HSCs had a significantly higher percentage of short interphase

distances (<2 μm) (20.50% vs.7.60% for 0-0.99 μm and 40.50% vs.18.10% for 1.00-1.99

μm), and similar or lower percentages of long interphase distance (≥2 μm) compared with

MSCs (0-1.99 μm vs. >2 μm, p = 7x10-13

). Comparison of the frequency of CBFB-

MYH11 co-localization (distance = 0.00 µm) between the two cell types also revealed

differences. HSCs had a higher percentage (3.81%) of gene pairs that co-localized

compared to MSCs, which did not have any co-localized pairs (Table 4.2). These results

Figure 4.1: Interphase FISH analysis of CBFB and MYH11. Two-color FISH of normal human

HSCs with the CBFB probe (RP11-5A19, orange) and MYH11 probe (RP11-585P8, green). Nuclei showing two distinct hybridization signals per probe were selected for interphase distance analysis,

assuming that the two shorter distances were between loci on the same chromosome. Scale bar = 4 μm.

83

show that CBFB and MYH11 are indeed located in close spatial proximity in HSCs

compared with MSCs, providing a potential mechanistic explanation of why breakage at

CBFB and MYH11 promotes the formation of inv(16)(p13q22) in HSCs but not in MSCs.

Figure 4.2: CBFB-MYH11 interphase distance distribution. Distribution of measured interphase

distances between CBFB and MYH11 loci in HSCs (untreated), MSCs, PBLs, fibroblasts, and treated HSCs, n = 210.

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Spatial Gene Proximity of CBFB and MYH11 in Specialized Cell Types

The measured distances between CBFB and MYH11 were examined in specialized cell

types and compared to the distances observed in HSCs. PBLs and fibroblasts were chosen

because they are fully-differentiated cell types derived from HSCs and MSCs,

respectively. Histograms of measured distances are shown in Figure 4.2. The distribution

of CBFB-MYH11 distances was statistically significant between HSCs and PBLs (p =

1.09x10-11

), as well as between HSCs and fibroblasts (p = 1.09x10-11

) (Table 4.1). The

percentage of chromosomes with gene loci 0-0.99 μm, 1.00-1.99 μm or 2.00-2.99 μm

apart was also determined. In PBLs, 7.10% and 20.50% of gene pairs were 0-0.99 μm

and 1.00-1.99 μm, respectively. The examination of fibroblasts proved similar, with

12.90% and 25.70% of CBFB and MYH11 loci separated by 0-0.99 μm and 1.00-1.99

μm, respectively (Figure 4.3). Chi square tests showed significant differences between

HSCs and PBLs (p = 2.08x10-13

) and fibroblasts (p = 8.34x10-7

). Furthermore, there were

observed differences among the frequency of co-localized gene pairs between HSCs

(3.81%), PBLs (2.86%), and fibroblasts (0.95%) (Table 4.2). Therefore, it was concluded

that the distance between CBFB and MYH11 is significantly greater in PBLs and

fibroblasts than in HSCs, supporting the idea that spatial proximity is cell-type and

differentiation stage-specific.

Table 4.1: Analysis of measured CBFB-MYH11 interphase distances

Cell type Mean ± SD (in µm) p value

a

HSCs 1.90 ± 1.14 N/A MSCs 3.78 ± 2.67 2.24x10

-18

PBLs 3.50 ± 2.34 1.09x10-11

Fibroblasts 3.19 ± 2.39 1.09x10

-11

Treated HSCs 2.07 ± 1.35 0.146 aSignificance compared to HSCs. n=210

85

While CBFB-MYH11 distances are significantly reduced in HSCs compared to

MSCs, PBLs and fibroblasts, due to potential differences in cell size, we next examined

the distance between CBFB and a control locus (16p11.2) in HSCs to determine whether

spatial gene proximity is a contributing factor in the formation of inv(16)(p13q22).

16p11.2 is separated from CBFB by ~37 Mb, and is located in-between CBFB and

MYH11 on metaphase chromosomes. This region is not known to participate in a

rearrangement with the CBFB gene, nor does it co-localize with a fragile site. The mean

distance between CBFB and 16p11.2 (2.78 ± 1.68 µm, Figure 4.4) was significantly

higher than the distance between CBFB and MYH11 in HSCs (p = 4.11x10-10

).

Furthermore, there was no co-localization between CBFB and 16p11.2, which is in

contrast to 3.81% of co-localizing pairs between CBFB and MYH11 in HSCs. These data

further support that CBFB and MYH11 are physically in close proximity during

interphase in HSCs.

Figure 4.3: Distribution patterns of CBFB-MYH11 distance intervals. Percentage of chromosomes

with CBFB-MYH11 located at 0-0.99 μm, 1.00-1.99 μm, 2.00-2.99 μm, 3.00-3.99 μm and ≥4 μm apart. Distances were measured in HSCs (untreated), MSCs, PBLs, fibroblasts, and treated HSCs. The percentage of chromosomes was determined from each range of distances (0-0.99 μm, light gray; 1.00-

1.99 μm, black; 2.00-2.99 μm, white; 3.00-3.99 μm, medium gray; ≥4 μm, dark gray), and results were compared using the chi-square test to determine statistical significance.

86

Proximity of CBFB and MYH11 Following Treatment with Fragile Site -Inducing

Chemicals in HSCs

To determine whether treatment with fragile site-inducing chemicals affects spatial gene

proximity of CBFB and MYH11, HSCs were treated with 0.4 μM aphidicolin (APH), 2

mM 2-aminopurine (2-AP) and 150 mg/L berenil for 24 h. APH, an inhibitor of DNA

polymerases α, δ and ε [6, 8], induces the majority of fragile sites [13] including

FRA16B/FRA16C [14], which co-localize with breakpoints within CBFB. 2-AP is a non-

specific inhibitor of ataxia telangiectasia and Rad3-related (ATR) kinase [182, 183], a

protein required for fragile site maintenance [37], and significantly increases fragile site

breakage up to 20-fold in combination with APH [37]. Berenil was also used because it is

the most common inducer of fragile site FRA16B [13]. The concentrations and

incubation times were chosen because they are optimal for the detection of fragile site

breakage. Induction of fragile sites was confirmed by cytogenetic analysis following

treatment (Figure 4.5). Distribution plots of measured distances revealed no obvious

differences between untreated and APH-treated HSCs (Figure 4.2). Statistical analysis

did not show any significant differences in mean distances for treated HSCs (Table 4.1)

or in the proportion of chromosomes with CBFB and MYH11 at different distance

intervals compared to untreated cells (Figure 4.3). Furthermore, there was no difference

in the frequency of co-localization between untreated and treated HSCs (Table 4.2).

Table 4.2: Frequency of CBFB and MYH11 co-localization

Cell type number of co-localized

gene pairs

% co-localizationa

HSCs 8 3.81

MSCs 0 0 PBLs 6 2.86

Fibroblasts 2 0.95 Treated HSCs 8 3.81

aPercentage of chromosomes in which the measured distance between CBFB and MYH11 = 0.00 µm. n= 210

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These results show that treatment with fragile site-inducing chemicals does not

significantly affect the distance between CBFB and MYH11 in interphase nuclei of HSCs.

DISCUSSION

The high occurrence of chromosomal aberrations in cancer has prompted a closer

investigation into the molecular basis of structural rearrangements in tumor cells. Here,

we found statistically significant differences between CBFB and MYH11 interphase

distances in HSCs compared to human MSCs, PBLs and fibroblasts. The data also

Figure 4.4: Comparison of interphase distance between CBFB and MYH11 with the distance

between CBFB and 16p11.2 in HSCs. (A) A schematic of chromosome 16 shows the location of

MYH11, 16p11.2, and CBFB loci. CBFB and MYH11 are separated by ~50 Mb on metaphase chromosome 16, whereas CBFB and 16p11.2 are separated by ~37 Mb. (B) The mean distance (± SD)

between each locus pair is shown, and the p value of the comparison between locus pairs is indicated. The percentage of co-localization between each locus pair is also listed. (C) Interphase distance

distributions between CBFB and MYH11 (left), and CBFB and 16p11.2 (right), n =210.

88

demonstrate significant differences in the percentage of chromosomes at different

distance levels. In HSCs, CBFB and MYH11 are separated by 1.99 µm or less in 61.43%

of chromosomes, whereas in MSCs, PBLs and fibroblasts, CBFB and MYH11 are

separated by 1.99 µm or less in 26.67%, 28.10% and 39.05% of chromosomes,

respectively (Figure 4.3). In contrast, distances greater than 2 μm were similar or less

frequent in HSCs compared to other cell types, indicating that at lengths >2 μm, the

spatial proximity of CBFB and MYH11 is dominated by random influences. These results

suggest that spatial genome organization is cell-type specific and support a role for

reduced interphase distances between translocation-participating loci in the formation of

chromosomal rearrangements [169, 170, 172, 180, 184].

As an additional control, the distance between CBFB and 16p11.2, a non-

translocation participating locus was measured to determine whether a close physical

relationship exists between CBFB and MYH11 during interphase in HSCs. Although

Figure 4.5: Induction of fragile site breakage in treated HSCs. G-banded chromosomes from normal HSCs treated with fragile site-inducing chemicals show metaphase breaks (arrows).

89

16p11.2 is located ~37 Mb away from CBFB during metaphase (~70% of the distance

between CBFB and MYH11), the mean interphase distance was significantly higher than

that between the CBFB and MYH11 genes, which are separated by ~50 Mb. These results

suggest that chromosome 16 may form a loop structure during interphase, bringing some

loci in close proximity while separating others. Therefore, translocation-participating loci

are in close physical proximity during interphase which may promote the formation of

translocations, and supports the hypothesis that spatial genome organization is a

contributing factor in the generation of chromosomal rearrangements.

Since differences in cell size could potentially have an effect on spatial gene

proximity, distances between CBFB and MYH11 were also determined as a fraction of the

nuclear diameter. We found that the mean CBFB-MYH11 distance in HSCs (16.03 ±

9.85%) was not significantly different compared to other cell types (Figure 4.6).

However, HSCs have the smallest average diameter of the cells examined (Figure 4.6).

These results suggest that nuclear size may be a factor in cell-type specific spatial

genome organization, and overall support the notion that spatial gene proximity plays a

role in the formation of chromosomal translocations and is likely the reason why certain

chromosomal aberrations are associated with specific tissues in human tumors [166]. It is

also important to note that the methodology utilized herein involves hypotonic swelling

of cells followed by fixation, which is known to exaggerate interphase distances.

Therefore, the distances reported here may not reflect the true interphase separation of

these genes in vivo. However, during harvest, attached cells were first trypsinized and

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Figure 4.6: Distribution of CBFB-MYH11 interphase distances as a percentage of nuclear

diameter. Distribution of distances as a fraction of nuclear diameter are shown for (A) HSCs, (B)

MSCs, (C), PBLs, (D) fibroblasts, and (E) treated HSCs. (F) The average diameter size is shown for each cell type, as well as the distance between CBFB and MYH11 as a percentage of nuclear diameter

(mean ± SD).

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then processed in the same manner as cells grown in suspension, allowing for an accurate

comparison between cell types.

Normal HSCs were treated with fragile site-inducing chemicals to evaluate the

role of chromosome breakage caused by fragile site expression on the spatial proximity

of CBFB and MYH11. Previous studies have shown that DNA damage caused by γ-

radiation can significantly reduce the proximity of genes in interphase [179], most likely

due to chromatin remodeling, and rearrange the positioning of chromosome territories

[185]. Based on these results, we hypothesized that treatment with fragile s ite-inducers

would reduce the distance between CBFB and MYH11 in interphase nuclei. Interestingly,

there was no significant difference between the average distances in untreated HSCs

versus treated HSCs, or in the percent distribution of chromosomes at different distance

intervals. Furthermore, the frequency of co-localization between CBFB and MYH11 did

not change upon treatment (Table 4.2). These data suggest that treatment with agents

causing mild replication stress does not have as great an effect on chromosome mobility

as irradiation. One explanation for this difference is that ionizing radiation produces

DNA double-strand breaks whereas chemicals used for the induction of fragile sites

likely only cause single-strand breaks or gaps [37]. Double-strand breaks may be more

mobile, making it possible for other ends to join.

The conserved nature of spatial genome organization across species and the

specificity of whole chromosome and gene locations within the nucleus point to a

significant role in the mechanism of chromosomal rearrangements. As the molecular

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basis of these events is being elucidated, two models have been proposed to explain how

chromosomal translocations occur. According to the breakage-first theory, DNA damage

occurs at two separate locations. Broken DNA ends then “scan” the nuclear space for

potential translocation partners that are subsequently joined together [186, 187]. This

model suggests that chromosome ends are able to move over large distances within the

nucleus, searching for other DNA breaks. The contact-first model predicts that structural

rearrangements take place after DNA damage occurs at a site where genes co-localize

within the nucleus [186, 188]. In this model, broken chromosome ends are expected to

have limited ability to move within the nuclear space. Evidence for both models exists,

although more recent data favor the contact-first model as the underlying mechanism in

the formation of chromosomal translocations [166, 169, 170, 180]. The results presented

in this study support the contact-first model of translocations since CBFB and MYH11

were significantly closer with respect to one another in HSCs compared with other cell

types, and this distance did not change following treatment with fragile site-inducing

chemicals. However, additional studies will be necessary to make a more definitive

determination of which model is most relevant in AML.

We initiated these studies in an effort to better understand the molecular basis of

chromosomal translocations and to define a potential mechanism for the formation of

inv(16)(p13q22) in AML. The interphase distance between participating genes CBFB and

MYH11 was examined based on a growing body of evidence that supports a role for

spatial gene proximity in the generation of chromosomal rearrangements found in cancer.

Our results are consistent with studies of other gene rearrangements, demonstrating that

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CBFB and MYH11 are located within close proximity in HSCs, and support the concept

that spatial proximity of translocation-prone loci may favor gene rearrangements.

Furthermore, the highly significant difference between CBFB-MYH11 distances in HSCs

compared with MSCs, PBLs and fibroblasts is in agreement with the fact that spatial

genome organization is tissue-specific. Mild replication stress did not affect CBFB-

MYH11 distances, perhaps because the damage was not significant enough to alter

chromatin structure or produce ends able to search for a potential partner. Overall, the

data presented in this study provide further evidence of a role for spatial gene proximity

in the formation of cancer-specific chromosomal translocations and offer new insight into

the mechanism of inv(16)(p13q22) in AML.

ACKNOWLEDGEMENTS

We would like to thank Will Stewart and Chris von Kap-Herr in the Section on Medical

Genetics for their help with FISH and Ken Grant in the Department of Pathology for his

help with confocal microscopy. Mesenchymal stem cells employed in this work were

provided by the Texas A&M Health Science Center College of Medicine Institute for

Regenerative Medicine at Scott & White through a grant from NCRR of the NIH, Grant #

P40RR017447.

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CHAPTER V: THE INDUCTION OF FRAGILE SITE FRA16B PRODUCES DNA BREAKS WITHIN CBFB OF INV(16)(P13Q22)

ABSTRACT

Supporting a causative role for fragile sites in cancer development, we previously

determined that over half of all reported breakpoints in cancer-specific recurrent

translocations map to fragile sites, and that fragile site co-localized loci exhibit

characteristics of fragile DNA, including the ability to form highly stable secondary

structures. Despite this significant association, the exact mechanism by which fragile

sites contribute to cancer development is still unclear. Fragile site breakage was recently

determined to directly lead to the formation of oncogenic rearrangements in thyroid

cancer by disrupting translocation-participating genes. To further elucidate the molecular

basis by which fragile sites contribute to the development of cancer, it is necessary to

investigate a role for fragile sites in the generation of additional cancer-causing

rearrangements, such as those observed in leukemia. We previously determined that

breakpoints within participating genes CBFB and MYH11, involved in inv(16)(p13q22)

in acute myeloid leukemia , co-localize with fragile sites FRA16B and FRA16A,

respectively. Here, we show that induction of FRA16B produces DNA breaks within the

major breakpoint region of CBFB, and that the induced breakpoints are located at similar

positions as those observed in patients. Furthermore, the frequency of breaks in fragile

site-induced cells was significantly higher than that in untreated cells. Overall, these

results further support a direct involvement of fragile sites in cancer development through

the generation of cancer-specific chromosomal rearrangements.

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INTRODUCTION

Despite a strong correlation between the location of fragile sites and sites of

chromosomal breaks in tumor cells [22], there has been little evidence of a direct role for

fragile DNA loci in the generation of tumorigenic rearrangements. To date, the only

direct evidence of fragile site involvement in cancer development comes from the

examination of the RET/PTC1 rearrangement, a major type of RET/PTC rearrangement

observed in papillary thyroid carcinoma. RET/PTC1 involves the RET gene, located at

fragile site FRA10G, and CCDC6, located at FRA10C. Recently, Gandhi et al.

demonstrated that treatment of normal human thyroid cells with fragile site-inducing

chemicals produced breaks within both RET and CCDC6 genes, and led to the generation

of RET/PTC1 rearrangements observed in human tumors [173].

To expand the model of how fragile site breakage leads to the development of

cancer, it was important to examine a chromosomal rearrangement in which both

participating genes co-map to fragile sites in a non-solid tumor, such as leukemia. A

variety of chemicals known to induce fragile site breakage have also been implicated as

risk factors for the development of acute myeloid leukemia (AML) [98, 99]. Therefore, a

potential role for fragile site breakage in AML development was investigated in this

study. We previously determined that translocation breakpoints in both genes

participating in inv(16)(p13q22), frequently observed in AML [106], co-localize with

fragile sites [22]. In the inv(16)(p13q22), the CBFB gene, located at FRA16B at

chromosomal locus 16q22.1, is rearranged with MYH11, which co-localizes with

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FRA16A at 16p13.11 [22]. Furthermore, CBFB is one of the most common targets of

genetic alterations in AML [106].

Here, ligation-mediated PCR (LM-PCR) was used to determine whether the

induction of FRA16B produces DNA breaks within CBFB, which may ultimately lead to

generation of the cancer-specific fusion transcript. Normal human hematopoietic stem

cells (HSCs) were treated with chemicals to induce breakage at FRA16B, and following

treatment, genomic DNAs were isolated and subjected to primer extension and two

rounds of nested PCR specific to intron 5, the major breakpoint region of CBFB

identified in patients [189]. The examination of PCR products on agarose gels revealed

DNA breaks within intron 5 of CBFB following treatment of HSCs with fragile site-

inducing chemicals, with a frequency which was significantly higher than that in

untreated cells. Furthermore, the locations of the induced breakpoints were similar to

those observed in patients, suggesting that FRA16B breakage may directly contribute to

the formation of inv(16)(p13q22) in AML.


Cell culture

Fresh primary human CD34+ HSCs (Lonza) were purchased and maintained in HPGM

hematopoietic progenitor growth medium (Lonza) containing a StemSpan CC100

cytokine cocktail of recombinant human (rh) Flt-3 ligand, rh stem cell factor, rh IL-3 and

rh IL-6 (STEMCELL Technologies). Cells were grown in a humidified incubator at 37˚C

with 5% CO2. For the induction of fragile sites, cells were treated with 0.4 μM

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aphidicolin (APH) (Sigma-Aldrich), 2 mM 2-aminopurine (2-AP) (Sigma-Aldrich) and

150 mg/L berenil (Sigma-Aldrich) for 24 h. Under optimal culture conditions, the

frequency of FRA16B breakage in lymphocytes is 90% following treatment with berenil

alone [18].

Detection of DNA breaks by LM-PCR

Genomic DNA was isolated from untreated HSCs and HSCs treated with fragile site-

inducing chemicals. DNA breaksite mapping was performed as described in [190], with

modifications (Figure 5.1). To detect DNA breaks within CBFB, a 5‟ biotinylated primer

corresponding to the 5‟ end of exon 6, CBFB1 (5‟- AGGTTGAGAAATAGACCGTAG-

TACCTCC -3‟), was used to extend into intron 5, the major breakpoint cluster region

identified in patients [189]. Primer extension was performed with 200 ng of DNA at

45ºC. The DNA breaks were then isolated through ligation of the duplex DNA LL3/LP2

linker [190] followed by incubation with streptavidin beads. Amplification of DNA

breaks was achieved by two rounds of nested PCR of the extension-ligation products

using primers CBFB2 (5‟- ACCACCTAAATTGGAACCAGGACTA -3‟) and CBFB3

(5‟- AGGACTAGGGTCTT-GTTGTCTTCTTGC -3‟), respectively, with the

corresponding linker-specific primers LL4 and LL2 [173]. Final PCR products were

resolved on 1.3% agarose gels, with each band corresponding to a DNA break isolated

from the region of interest. PCR products were sequenced to confirm the isolated breaks

were within intron 5 of CBFB. As a positive control, breaksites were identified in intron 4

of the FHIT gene, which is involved in deletions and other rearrangements found in

several types of cancer [191], and harbors the major breakpoint cluster region within the

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most frequently expressed fragile site in the human genome, FRA3B. For FRA3B, the

biotinylated primer FRA3B-20 was used for the primer extension and primers FRA3B-9

and FRA3B-23 were used in the first and second rounds of nested PCR, respectively

[173]. As a negative control, breaks were examined in the non-fragile 12p12.3 region

using the biotinylated primer 12p12.3-1 and primers 12p12.3-2 and 12p12.3-3 for nested

PCR [173]. The frequency of DNA breaks among samples, as determined by LM-PCR,

was compared using a one-tailed student‟s t-test.

RESULTS

Induction of DNA Breaks in intron 5 of CBFB by Fragile Site-Inducing Chemicals

To date, ten different types of fusion transcripts generated by inv(16)(p13q22) have been

identified, and are termed type A through J. Types A, D and E account for nearly 98% of

all cases, whereas all others represent unique cases [192]. The different types of fusion

transcripts are mainly caused by breakpoint heterogeneity in the MYH11 gene [192]. In

contrast, intron 5 has been identified as the major breakpoint cluster region within CBFB

[189]. Since the breakpoint region within CBFB is localized to intron 5, we tested

whether the induction of fragile sites can produce breaks within CBFB. Genomic DNAs

were isolated from untreated HSCs and HSCs treated with fragile site-inducing

chemicals, and subjected to primer extension and two rounds of nested PCR with primers

specific to intron 5 of CBFB. PCR products were visualized by agarose gel

electrophoresis (Figure 5.2), where each lane represents the DNA breaks isolated from

approximately 1000 cells , and each band corresponds to a break within the region of

interest. DNA breaks were detected within intron 5 of CBFB following treatment with

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fragile site-inducing chemicals (Figure 5.2A) at a frequency of 0.058 ± 0.018 breaks per

100 cells (Table 5.1). This rate was significantly higher than that in untreated cells (0.022

± 0.015 breaks per 100 cells, P = 0.027; Figure 5.2B, Table 5.1).

DNA breaks were also examined within fragile site FRA3B, the most inducible

fragile site in the genome, after treatment with fragile site-inducing chemicals. DNA

breaks were observed within FRA3B upon treatment (Figure 5.2C) at a frequency of

0.078 ± 0.042 breaks per 100 cells (Table 5.1), confirming that treatment induces

Figure 5.1: DNA breaksite mapping by LM-PCR. Genomic DNA was isolated from untreated HSCs and HSCs treated with fragile site-inducing chemicals. The DNA was denatured and annealed to a

biotinylated primer specific to the region of interest. Primer extension was carried out with DNA Sequenase, and terminated at a DNA breaksite. Breaks were then isolated through ligation of the LL3/LP2 linker and recovered using streptavidin beads. Amplification of DNA breaks was achieved by

two rounds of nested PCR of the extension-ligation products. Final PCR products were resolved on agarose gels, with each band corresponding to a break within the region of interest.

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breakage at fragile sites. The increased number of breaks within FRA3B, as compared to

CBFB is consistent with the fact that FRA3B is the most frequently expressed fragile site

in the human genome. As a negative control, DNA breaks were examined in the non-

fragile 12p12.3 region following treatment with fragile site-inducing chemicals (Figure

5.2D). The low frequency of breaks within 12p12.3 (0.016 ± 0.011 per 100 cells; Table

5.1) was statistically significant compared to the frequency of DNA breaks observed

within CBFB following treatment (P = 0.014; Table 5.1), suggesting that the breaks

detected within FRA3B and CBFB are caused by their fragile nature upon treatment with

fragile site-inducing chemicals.

Figure 5.2: Detection of DNA breaks by LM-PCR following treatment of HSCs with fragile site-inducing chemicals. LM-PCR was utilized to detect breaks produced in HSCs after treatment with

fragile site-inducing chemicals within intron 5 of CBFB (A), fragile site FRA3B (C), and the non-fragile 12p12.3 region (D). Intron 5 of CBFB was also examined in untreated HSCs (B). The first lane of each gel is a 100 bp molecular weight ladder.

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Comparison of fragile site -induced breakpoints in CBFB with known breakpoints

observed in patients

To determine whether the induced breaksites are located at or near the breaksites

observed in patients, first DNA samples from LM-PCR were sequenced to confirm the

specificity of the primers. DNA sequencing revealed the breakpoints to be located within

intron 5 of CBFB. The locations of these breakpoints were then compared with the

location of known breakpoints in AML patients with inv(16)(p13q22) [189] (Figure 5.3).

To date, breakpoints in only 18 patients have been mapped, and only two patient

breaksites within intron 5 are able to be detected using the primer sets described above.

Each induced breaksite was determined to be at or near a patient breaksite, with distances

ranging from 0 to 144 base pairs, demonstrating that exposure of normal human HSCs to

fragile site-inducing chemicals produces DNA breaks within the major breakpoint cluster

region of the CBFB gene, and that the induced breakpoints are located close to known

breaksites observed in patients. It is important to note that the induced breaksites were

detected prior to a rearrangement event, whereas breakpoints found in cancer cells are

identified afterwards, and have been determined to undergo small modifications, such as

deletions or insertions, or even gene duplications by as many as 192 base pairs [189]. For

future studies, when comparing the location of induced breakpoints with those of

patients, it will be important to examine intron 5 in its entirety, and to identify additional

Table 5.1: Frequency of DNA breaks detected in HSCs by LM-PCR

Region of interest number of breaks/100 cells

(mean ± SD) p value

a

CBFB (untreated) 0.022 ± 0.015 0.027

CBFB (treated) 0.058 ± 0.018 N/A FRA3B 0.078 ± 0.042 0.253

12p12.3 0.016 ± 0.011 0.014 aSignificance compared to CBFB treated.

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breakpoints in patients to demonstrate potential similarities between the fragile site-

induced and patient breaksites.

DISCUSSION

Although results from numerous studies support a causative role for fragile sites in

cancer-specific chromosomal rearrangements, the only evidence of a direct involvement

of fragile site breakage in cancer development comes from the examination of the

RET/PTC1 rearrangement frequently associated with thyroid cancer [173]. The

investigation of the formation of inv(16)(p13q22) in this study was intended to provide

additional evidence of a direct role for fragile sites in the generation of cancer-causing

rearrangements, and to expand the proposed model of their involvement in cancer

development to non-solid tumors, by determining whether DNA breaks can be produced

Figure 5.3: Location of induced breakpoints within intron 5 of CBFB. Sequencing results

confirmed the location of six induced breaksites within intron 5 of CBFB (gray arrowheads). The location of known breakpoints in AML patients with inv(16)(p13q22) are indicated by black arrowheads [189]. The light blue arrow corresponds to the CBFB1 primer with a dual biotin label (light

blue circles), which anneals to exon 6 of CBFB. The dark blue solid and dashed arrows correspond to CBFB2 and CBFB3 nested PCR primers, respectively. The intron 5 sequence of CBFB is italicized.

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within the major breakpoint region of CBFB, disrupted in AML by inv(16)(p13q22),

following treatment of HSCs with fragile site-inducing chemicals.

Potential risk factors of AML development include exposure to benzene [98],

chemotherapeutic agents [99], and other factors known to induce or enhance the

expression of fragile sites. Based on these data, we proposed that fragile site breakage

plays a role in the generation of AML. Since we previously demonstrated the co-

localization of fragile sites with breakpoints involved in inv(16)(p13q22) [22], we sought

to investigate the molecular basis by which fragile site breakage leads to the formation of

inv(16)(p13q22). Here, we show that breakage of fragile site FRA16B generates DNA

breaks within the major breakpoint cluster region of CBFB identified in patients,

potentially leading to disruption of the gene product and ultimately formation of

inv(16)(p13q22). Moreover, the DNA breaks were confirmed to be fragile in nature when

comparing the formation of breaks within CBFB to fragile site FRA3B and the non-

fragile 12p12.3 region. FRA3B showed the highest frequency of DNA breaks following

treatment with fragile site-inducing chemicals (Figure 5.2C, Table 5.1), which is

consistent with the fact that FRA3B is the most inducible fragile site in the human

genome. In contrast, untreated cells and the 12p12.3 region showed little to no breaks

(Figure 5.2B and D, Table 5.1).

Overall, the results presented here show that FRA16B breakage can generate

breaks at the nucleotide level within the major breakpoint region of CBFB at locations

similar to those identified in patients, and provide additional evidence of a direct role for

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fragile sites in cancer development through the formation of chromosomal aberrations.

These results are important for understanding both the mechanism of chromosomal

rearrangements and the involvement of fragile site breakage in cancer development. More

importantly, these data suggest that exposure to environmental and/or dietary agents

known to induce fragile sites may contribute to the occurrence of chromosomal

rearrangements, and ultimately cancer development through a fragile site-mediated

mechanism. While these results strongly suggest that breakage at FRA16B may

ultimately lead to inv(16)(p13q22) by disrupting the CBFB gene, it will be important for

future studies to determine whether MYH11 is also disrupted in HSCs following

treatment with fragile site-inducing chemicals, and ultimately, whether the fusion product

of inv(16)(p13q22) can be generated in cells following treatment to provide direct

evidence of fragile site involvement in inv(16)(p13q22).

ACKNOWLEDGEMENTS

Special thanks to Merissa Baxter for her help with LM-PCR.

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CHAPTER VI: CONCLUSIONS

While a substantial body of evidence continues to demonstrate a causative role for

fragile sites in tumorigenesis, the detailed mechanism by which fragile site breakage

ultimately results in the generation of cancer-specific chromosomal rearrangements

remains to be elucidated. Therefore, it has been essential to further investigate the

molecular basis of fragile site breakage, and to examine consequences of breakage at

these sites in order to dissect their role in cancer development. The work in this

dissertation demonstrates the contribution of secondary structure formation to fragile site

breakage by showing that fragile site co-localized loci participating in cancer-specific

translocations are capable of forming highly stable DNA secondary structures, and more

importantly, that the ability of the FRA16B sequence to form an alternative structure in

vitro affects its replication efficiency in human cells and produces the same types of

mutations as those observed in vivo. Furthermore, the examination of a role for FRA16B

in inv(16)(p13q22) showed that FRA16B induction produced DNA breaks within the

major breakpoint region of participating gene CBFB, indicating that the ability of

FRA16B to form a stable secondary structure may contribute to the formation of

inv(16)(p13q2) by disrupting rearrangement-participating genes.

DNA Secondary Structure Formation Contributes to Fragile Site Breakage

For years, it has been hypothesized that the delayed replication observed at fragile sites

resulting in gaps and breaks on metaphase chromosomes is caused by the ability of

fragile DNAs to form secondary structures that inhibit synthesis, rendering it difficult to

complete replication [14]. To demonstrate a role for secondary structure formation in

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fragile site breakage, we randomly selected three gene pairs involved in cancer-specific

recurrent translocations in which both translocation-participating genes contain fragile

site co-localized breakpoints, and examined the DNA sequences for the ability to form

secondary structures. Predictions of secondary structure formation by Mfold

demonstrated the potential of fragile site co-localized DNAs to form extensive structures

with multiple hairpins that are much more stable compared to a randomized DNA

sequence of similar length. Since most fragile sites studied to date are predicted to form

highly stable secondary structures [14, 15], and breakpoints in over half of all reported

cancer-specific translocations are located within fragile sites [22], these results suggest

that the formation of secondary structures at fragile site loci directly contributes to their

breakage and subsequent rearrangements in cancer.

Next, the experiments in this work provided direct physical evidence of secondary

structure formation in fragile site FRA16B, which represents both major classes of fragile

sites. Due to the observed secondary structure formation of FRA16B in vitro, we

proposed the capability of the FRA16B sequence to undergo instability events, such as

those observed in vivo, in an episomal system. The results presented here demonstrate the

ability of the FRA16B sequence to inhibit replication in human cells, which is likely

caused by the formation of an alternative structure that perturbs DNA synthesis via

polymerase pausing, spontaneous fork regression, fork restart, and/or polymerase bypass,

leading to a delay in replication, and ultimately to the same types of insertions and

deletions observed at fragile sites in humans. The mechanism proposed here whereby

secondary structure formation leads to chromosomal breakage is not an uncommon

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physiologic process, but instead occurs frequently during V(D)J recombination in which

DNA hairpins are cleaved and broken ends are re-ligated together by the NHEJ pathway

to create a diversity of immunoglobulins and T-cell receptors [193], providing further

evidence of a role for secondary structure formation in DNA breakage.

Fragile Site Breakage Promotes Chromosomal Rearrangements

In addition to demonstrating the contribution of secondary structure formation to fragile

site breakage, the results described here also show how breakage at fragile sites may lead

to the formation of chromosomal rearrangements. Investigation into a role for FRA16B in

inv(16)(p13q22) revealed that treatment of normal HSCs with chemicals to induce

FRA16B generated DNA breaks within the major breakpoint cluster region of

participating gene, CBFB, and established a potential mechanism whereby fragile sites

directly contribute to the formation of cancer-specific chromosomal rearrangements by

producing breaks within, and ultimately disrupting participating genes.

Since the molecular basis for common and rare fragile sites is likely shared due to

intrinsic features of fragile DNA [14], we propose that breakage at fragile sites caused by

secondary structure formation may disrupt numerous rearrangement-participating genes

which, in some cases, can lead to cancer development. Supporting this hypothesis, the

comprehensive examination of fragile site locations relative to chromosomal breakpoints

in cancer showed that over half of all known cancer-specific recurrent translocation

breakpoints co-map to fragile sites, and are predicted to form highly stable secondary

structures. Therefore, chromosomal fragility caused by the formation of secondary

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structures may lead to an enhanced susceptibility to DNA breakage within translocation-

prone loci, thus promoting a rearrangement event. Since the majority of fragile sites

associated with translocation breakpoints are CFS, which are present in all individuals

and are induced by a variety of mutagens [89], individuals with exposure to various

environmental and/or chemical agents may be more susceptible to the development of

sporadic cancer through the generation of breaks within genes participating in cancer-

causing rearrangements.

As the mechanism of chromosomal rearrangements is being further elucidated,

spatial genome organization is emerging as a contributing factor [166]. The examination

of CBFB and MYH11 genes, which are involved in inv(16)(p13q22) in AML supported

this hypothesis, in that the genes are located significantly closer to each other in HSCs

than in MSCs, PBLs, or fibroblasts. These results also indicate that spatial genome

organization is cell-type and differentiation stage-specific, and could be a factor in why

certain chromosomal rearrangements are associated with specific tissues in human

cancers. Since treatment of HSCs with fragile site-inducing chemicals did not

significantly reduce the distance between, or the frequency of co-localization of CBFB

and MYH11, these data support the contact-first model of chromosomal rearrangements

and suggest that genes already located close to one another during interphase are able to

join together following DNA damage, such as fragile site breakage resulting from

exposure to environmental mutagens or chemicals.

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Proposed Mechanism for the Role of FRA16B in the Formation of inv(16)(p13q22)

Herein a model is proposed for FRA16B instability in the generation of inv(16)(p13q22)

in AML (Figure 6.1). CBFB and MYH11 genes, located within fragile sites FRA16B and

FRA16A, respectively, are in close proximity during interphase in normal human HSCs,

thus promoting the formation of the rearrangement. Under conditions of replication

stress, such as those environmental and dietary agents known to induce fragile sites,

replicative DNA polymerases become uncoupled from the helicase/topoisomerase

complex, resulting in long stretches of ssDNA. Regions that maintain single-

strandedness, such as the OIZ, may promote the formation of stable secondary structures

due to the intrinsic features of fragile DNA. These structures can cause significant

difficulties during replication, resulting in a stalled replication fork. The ATR-dependent

DNA damage checkpoint pathway, which responds to stalled or collapsed replication

forks [134, 135], is then triggered. As shown in our examination of FRA16B using an

SV40 replication system, a stalled replication fork may spontaneously regress during

synthesis of fragile DNA, generating a Holliday junction-like intermediate which could

lead to breakage through cleavage by resolvase enzymes [194], or polymerase bypass

might occur at regions of structure formation resulting in chromosome deletions. For the

repair of stalled forks, ATR, the main sensor of the pathway, binds to the fragile DNA

either directly or through complexes [48] and activates a downstream signalling cascade

through phosphorylation of various targets, including central regulator CHK1 [47]. Other

components of the ATR pathway including BRCA1 [46], FANCD2 [50], WRN [35, 53],

Claspin [49], HUS1 [51], and SMC1 [52], are crucial for fragile site maintenance,

although their direct role remains unclear. If the ATR pathway properly responds, the

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Figure 6.1: Model of FRA16B instability in the formation of inv(16)(p13q22) in acute myeloid

leukemia. CBFB and MYH11 genes, located on chromosome 16 within the fragile sites FRA16B/C and FRA16A, respectively, are in close proximity during interphase in normal hematopoietic stem cells.

Under conditions of replication stress or exposure to environmental factors, replicative DNA polymerases α, δ, and ε become uncoupled from the helicase/topoisomerase complex, resulting in long stretches on single-stranded DNA susceptible to the formation of stable secondary structures. These

structures can cause replication fork stalling, triggering the ATR-dependent DNA damage checkpoint pathway. Fragile sites may also be susceptible to spontaneous fork reversal or polymeras e skipping at regions of secondary structure. For repair of stalled forks, ATR binds to the fragile DNA either directly

or through complexes, and activates a downstream signaling cascade with other proteins, including CHK1, BRCA1, FANCD2, WRN, Claspin, HUS1, ATM, and SMC1. If the ATR pathway properly

responds, the replication fork will be repaired and DNA replication will resume normally. A loss, deficiency, or defect in ATR pathway proteins could lead to checkpoint failure and/or replication fork collapse resulting in DNA breakage at CBFB and MYH11. DNA breakage at these sites can lead to the

formation of inv(16)(p13q22), the expression of the CBFB-MYH11 fusion transcript, and the development of acute myeloid leukemia.

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replication fork will be repaired and DNA replication will resume normally. However, a

loss, deficiency, or defect in ATR pathway proteins could lead to checkpoint failure

and/or replication fork collapse resulting in DNA breakage, the initiating event of

translocation formation. Therefore, breakage at CBFB and MYH11 may lead to the

formation of inv(16)(p13q22) through a fragile site-mediated mechanism, resulting in the

development of AML.

Overall Significance

The data presented in this dissertation clearly demonstrate a role for secondary structure

formation in the mechanism of fragile site breakage, which may ultimately result in the

formation of chromosomal rearrangements in cancer. Here, secondary structure formation

was determined to enhance fragile site susceptibility to DNA breakage by inhibiting

synthesis, rendering it difficult to complete replication. The comprehensive examination

of fragile site locations relative to translocation breakpoints in cancer revealed a

significant association, and strongly suggests that fragile site breakage caused by genetics

or environmental/dietary factors predisposes individuals to cancer development through

the formation of chromosomal rearrangements which arise through the disruption of

participating genes, and may be promoted by the location of genes during interphase.

Overall, these results provide greater insight into the molecular basis of fragile

site breakage, the mechanism of chromosomal translocations, and the involvement of

fragile sites in the generation of cancer-specific rearrangements. It is important to note

that although the effect of secondary structure formation on fragile site breakage was

112

only determined for FRA16B, FRA16B exhibits properties of both major types of fragile

sites, and therefore it is likely that the majority of fragile DNAs are able to form stable

secondary structures capable of perturbing replication. However, for future studies it will

be important to determine the ability of other fragile DNAs to form alternative structures

in vitro, as well as an effect of secondary structure formation on replication in vivo.

Additionally, to provide direct evidence of a role for fragile sites in the generation of

inv(16)(p13q22) in AML, there is a need to determine whether MYH11 is disrupted by

fragile site breakage, and whether the fusion product can be generated upon treatment

with fragile site-inducing chemicals. It will also be important to identify additional

factors that contribute to chromosomal fragility at fragile sites such as DNA sequences,

proteins and environmental/dietary agents, since fragile sites are present in all individuals

and are sensitive to a range of chemicals, including ingested substances like caffeine [79,

81] and ethanol [82, 83]. A better understanding of the molecular basis and consequences

of fragile site breakage could ultimately allow for the development of a predictive assay

for cancer risk and insight into their direct role in tumorigenesis.

113

BIBLIOGRAPHY

1. Richards, R.I., Fragile and unstable chromosomes in cancer: causes and

consequences. TRENDS in Genetics, 2001. 17: p. 339-345. 2. Craig-Holmes, A.P., L.C. Strong, A. Goodacre, and S. Pathak, Variation in the

expression of aphidicolin-induced fragile sites in human lymphocyte cultures. Hum Genet, 1987. 76(2): p. 134-7.

3. Tunca, B., U. Egeli, A. Zorluoglu, T. Yilmazlar, O. Yerci, and A. Kizil, The expression of fragile sites in lymphocytes of patients with rectum cancer and their

first-degree relatives. Cancer Lett, 2000. 152(2): p. 201-9. 4. Glover, T.W. and C.K. Stein, Induction of sister chromatid exchanges at common

fragile sites. Am J Hum Genet, 1987. 41(5): p. 882-90. 5. Glover, T.W. and C.K. Stein, Chromosome breakage and recombination at

fragile sites. Am J Hum Genet, 1988. 43(3): p. 265-73. 6. Glover, T.W., Common fragile sites. Cancer Lett, 2006. 232(1): p. 4-12. 7. Glover, T.W., C. Berger, J. Coyle, and B. Echo, DNA polymerase à inhibition by

aphidicolin induces gaps and breaks at common fragile sites in human

chromosomes. Hum.Genet., 1984. 67: p. 136-142. 8. Cheng, C.H. and R.D. Kuchta, DNA polymerase epsilon: aphidicolin inhibition

and the relationship between polymerase and exonuclease activity. Biochemistry, 1993. 32(33): p. 8568-74.

9. McAvoy, S., S.C. Ganapathiraju, A.L. Ducharme-Smith, J.R. Pritchett, F. Kosari, D.S. Perez, Y. Zhu, C.D. James, and D.I. Smith, Non-random inactivation of large common fragile site genes in different cancers. Cytogenet Genome Res, 2007. 118(2-4): p. 260-9.

10. Kremer, E.J., M. Pritchard, M. Lynch, S. Yu, K. Holman, E. Baker, S.T. Warren, D. Schlessinger, G.R. Sutherland, and R.I. Richards, Mapping of DNA instability at the fragile X to a trinucleotide repeat sequence p(CCG)n. Science, 1991. 252(5013): p. 1711-4.

11. Sutherland, G.R., E. Baker, and R.I. Richards, Fragile sites still breaking. Trends Genet, 1998. 14(12): p. 501-6.

12. Mrasek, K., C. Schoder, A.C. Teichmann, K. Behr, B. Franze, K. Wilhelm, N. Blaurock, U. Claussen, T. Liehr, and A. Weise, Global screening and extended

nomenclature for 230 aphidicolin-inducible fragile sites, including 61 yet unreported ones. Int J Oncol, 2010. 36(4): p. 929-40.

13. Sutherland, G.R., Chromosomal fragile sites. Genet Anal Tech Appl, 1991. 8(6): p. 161-6.

14. Zlotorynski, E., A. Rahat, J. Skaug, N. Ben-Porat, E. Ozeri, R. Hershberg, A. Levi, S.W. Scherer, H. Margalit, and B. Kerem, Molecular basis for expression of common and rare fragile sites. Mol Cell Biol, 2003. 23(20): p. 7143-51.

15. Mishmar, D., A. Rahat, S.W. Scherer, G. Nyakatura, B. Hinzmann, Y. Kohwi, Y.

Mandel-Gutfroind, J.R. Lee, B. Drescher, D.E. Sas, H. Margalit, M. Platzer, A. Weiss, L.C. Tsui, A. Rosenthal, and B. Kerem, Molecular characterization of a

114

common fragile site (FRA7H) on human chromosome 7 by the cloning of a simian virus 40 integration site. Proc Natl Acad Sci U S A, 1998. 95(14): p. 8141-6.

16. Schmid, M., C. Klett, and A. Niederhofer, Demonstration of a heritable fragile

site in human chromosome 16 with distamycin A. Cytogenet Cell Genet, 1980. 28(1-2): p. 87-94.

17. Sutherland, G.R., P.B. Jacky, and E.G. Baker, Heritable fragile sites on human chromosomes. XI. Factors affecting expression of fragile sites at 10q25, 16q22,

and 17p12. Am J Hum Genet, 1984. 36(1): p. 110-22. 18. Felbor, U., W. Feichtinger, and M. Schmid, The rare human fragile site 16B.

Cytogenet Genome Res, 2003. 100(1-4): p. 85-8. 19. Yu, S., M. Mangelsdorf, D. Hewett, L. Hobson, E. Baker, H.J. Eyre, N. Lapsys,

D. Le Paslier, N.A. Doggett, G.R. Sutherland, and R.I. Richards, Human chromosomal fragile site FRA16B is an amplified AT-rich minisatellite repeat. Cell, 1997. 88(3): p. 367-74.

20. McKenzie, F., A. Turner, S. Withers, P. Dalzell, M. McGlynn, and E.P. Kirk,

Dominant inheritance of cleft palate, microstomia and micrognathia--possible linkage to the fragile site at 16q22 (FRA16B). Clin Dysmorphol, 2002. 11(4): p. 237-41.

21. Zollino, M., M. Genuardi, and G. Neri, Differential expression of FRA16B in

peripheral lymphocytes and bone marrow cells. Cancer Genet Cytogenet, 1990. 49(2): p. 229-33.

22. Burrow, A.A., Williams, L.E., Pierce, L.C.T., and Wang, Y.-H., Over half of breakpoints in gene pairs involved in cancer-specific recurrent translocations are

mapped to human chromosomal fragile sites. BMC Genomics, 2009. 10(59). 23. Pelliccia, F., N. Bosco, A. Curatolo, and A. Rocchi, Replication timing of two

human common fragile sites: FRA1H and FRA2G. Cytogenet Genome Res, 2008. 121(3-4): p. 196-200.

24. Le Beau, M.M., F.V. Rassool, M.E. Neilly, R. Espinosa, 3rd, T.W. Glover, D.I. Smith, and T.W. McKeithan, Replication of a common fragile site, FRA3B, occurs late in S phase and is delayed further upon induction: implications for the mechanism of fragile site induction. Hum Mol Genet, 1998. 7(4): p. 755-61.

25. Palumbo, E., L. Matricardi, E. Tosoni, A. Bensimon, and A. Russo, Replication dynamics at common fragile site FRA6E. Chromosoma. 119(6): p. 575-87.

26. Hellman, A., A. Rahat, S.W. Scherer, A. Darvasi, L.C. Tsui, and B. Kerem, Replication delay along FRA7H, a common fragile site on human chromosome 7,

leads to chromosomal instability. Mol Cell Biol, 2000. 20(12): p. 4420-7. 27. Handt, O., E. Baker, S. Dayan, S.M. Gartler, E. Woollatt, R.I. Richards, and R.S.

Hansen, Analysis of replication timing at the FRA10B and FRA16B fragile site loci. Chromosome Res, 2000. 8(8): p. 677-88.

28. Hansen, R.S., T.K. Canfield, A.D. Fjeld, S. Mumm, C.D. Laird, and S.M. Gart ler, A variable domain of delayed replication in FRAXA fragile X chromosomes: X inactivation-like spread of late replication. Proc Natl Acad Sci U S A, 1997. 94(9): p. 4587-92.

29. Zuker, M., Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res, 2003. 31(13): p. 3406-15.

115

30. Fry, M. and L.A. Loeb, The fragile X syndrome d(CGG)n nucleotide repeats form a stable tetrahelical structure. Proc Natl Acad Sci U S A, 1994. 91(11): p. 4950-4.

31. Gacy, A.M., G. Goellner, N. Juranic, S. Macura, and C.T. McMurray, Trinucleotide repeats that expand in human disease form hairpin structures in vitro. Cell, 1995. 81(4): p. 533-40.

32. Usdin, K. and K.J. Woodford, CGG repeats associated with DNA instability and

chromosome fragility form structures that block DNA synthesis in vitro. Nucleic Acids Res, 1995. 23(20): p. 4202-9.

33. Samadashwily, G.M., G. Raca, and S.M. Mirkin, Trinucleotide repeats affect DNA replication in vivo. Nat Genet, 1997. 17(3): p. 298-304.

34. Zhang, H. and C.H. Freudenreich, An AT-rich sequence in human common fragile site FRA16D causes fork stalling and chromosome breakage in S. cerevisiae. Molecular Cell, 2007. 27: p. 367-379.

35. Shah, S.N., P.L. Opresko, X. Meng, M.Y. Lee, and K.A. Eckert, DNA structure

and the Werner protein modulate human DNA polymerase delta-dependent replication dynamics within the common fragile site FRA16D. Nucleic Acids Res, 2009.

36. Letessier, A., G.A. Millot, S. Koundrioukoff, A.M. Lachages, N. Vogt, R.S.

Hansen, B. Malfoy, O. Brison, and M. Debatisse, Cell-type-specific replication initiation programs set fragility of the FRA3B fragile site. Nature. 470(7332): p. 120-3.

37. Casper, A.M., P. Nghiem, M.F. Arlt, and T.W. Glover, ATR regulates fragile site

stability. Cell, 2002. 111: p. 779-789. 38. Abraham, R.T., Cell cycle checkpoint signaling through the ATM and ATR

kinases. Genes & Dev, 2001. 15: p. 2177-2196. 39. Cimprich, K.A. and D. Cortez, ATR: an essential regulator of genome integrity.

Nat Rev Mol Cell Biol, 2008. 9(8): p. 616-27. 40. Cliby, W.A., C.J. Roberts, K.A. Cimprich, C.M. Stringer, J.R. Lamb, S.L.

Schreiber, and S.H. Friend, Overexpression of a kinase-inactive ATR protein causes sensitivity to DNA-damaging agents and defects in cell cycle checkpoints.

EMBO J, 1998. 17(1): p. 159-69. 41. Cortez, D., S. Guntuku, J. Qin, and S.J. Elledge, ATR and ATRIP: partners in

checkpoint signaling. Science, 2001. 294(5547): p. 1713-6. 42. Hammond, E.M., N.C. Denko, M.J. Dorie, R.T. Abraham, and A.J. Giaccia,

Hypoxia links ATR and p53 through replication arrest. Mol Cell Biol, 2002. 22(6): p. 1834-43.

43. Casper, A.M., S.G. Durkin, M.F. Arlt, and T.W. Glover, Chromosomal instability at common fragile sites in Seckel syndrome. Am.J.Hum.Genet., 2004. 75: p. 654-

660. 44. Ragland, R.L., M.F. Arlt, E.D. Hughes, T.L. Saunders, and T.W. Glover, Mice

hypomorphic for Atr have increased DNA damage and abnormal checkpoint response. Mamm Genome, 2009. 20(6): p. 375-85.

45. Ozeri-Galai, E., M. Schwartz, A. Rahat, and B. Kerem, Interplay between ATM and ATR in the regulation of common fragile site stability. Oncogene, 2008. 27: p. 2109-2117.

116

46. Arlt, M.F., B. Xu, S.G. Durkin, A.M. Casper, M.B. Kastan, and T.W. Glover, BRCA1 is required for common-fragile-site stability via its G 2/M checkpoint function. Molecular and Cellular Biology, 2004. 24: p. 6701-6709.

47. Durkin, S.G., M.F. Arlt, N.G. Howlett, and T.W. Glover, Depletion of CHK1, but not CHK2, induces chromosomal instability and breaks at common fragile sites. Oncogene, 2006. 25(32): p. 4381-8.

48. Wan, C., A. Kulkarni, and Y.H. Wang, ATR preferentially interacts with common

fragile site FRA3B and the binding requires its kinase activity in response to aphidicolin treatment. Mutat Res, 2010. 686(1-2): p. 39-46.

49. Focarelli, M.L., S. Soza, L. Mannini, M. Paulis, A. Montecucco, and A. Musio, Claspin inhibition leads to fragile site expression. Genes Chromosomes Cancer,

2009. 48(12): p. 1083-90. 50. Howlett, N.G., T. Taniguchi, S.G. Durkin, A.D. D'Andrea, and T.W. Glover, The

Fanconi anemia pathway is required for the DNA replication stress response and for the regulation of common fragile site stability. Hum.Mol.Genet., 2005. 14: p.

693-701. 51. Zhu, M. and R.S. Weiss, Increased common fragile site expression, cell

proliferation defects, and apoptosis following conditional inactivation of mouse Hus1 in primary cultured cells. Mol Biol Cell, 2007. 18(3): p. 1044-55.

52. Musio, A., C. Montagna, T. Mariani, M. Tilenni, M.L. Focarelli, L. Brait, E. Indino, P.A. Benedetti, L. Chessa, A. Albertini, T. Ried, and P. Vezzoni, SMC1 involvement in fragile site expression. Hum Mol Genet, 2005. 14(4): p. 525-33.

53. Pirzio, L.M., P. Pichierri, M. Bignami, and A. Franchitto, Werner syndrome

helicase activity is essential in maintaining fragile site stability. J Cell Biol, 2008. 180(2): p. 305-14.

54. Cimprich, K.A., Fragile sites: breaking up over a slowdown. Curr Biol, 2003. 13(6): p. R231-3.

55. Mitelman, F., F. Mertens, and B. Johansson, A breakpoint map of recurrent chromosomal rearrangements in human neoplasia. Nat Genet, 1997. 15 Spec No: p. 417-74.

56. Shrivastav, M., L.P. De Haro, and J.A. Nickoloff, Regulation of DNA double-

strand break repair pathway choice. Cell Res, 2008. 18(1): p. 134-47. 57. Gasparini, P., G. Sozzi, and M.A. Pierotti, The role of chromosomal alterations in

human cancer development. J Cell Biochem, 2007. 102(2): p. 320-31. 58. Schwartz, M., E. Zlotorynski, M. Goldberg, E. Ozeri, A. Rahat, C. le Sage, B.P.

Chen, D.J. Chen, R. Agami, and B. Kerem, Homologous recombination and nonhomologous end-joining repair pathways regulate fragile site stability. Genes Dev, 2005. 19(22): p. 2715-26.

59. Klugbauer, S., P. Pfeiffer, H. Gassenhuber, C. Beimfohr, and H.M. Rabes, RET

rearrangements in radiation-induced papillary thyroid carcinomas: high prevalence of topoisomerase I sites at breakpoints and microhomology-mediated end joining in ELE1 and RET chimeric genes. Genomics, 2001. 73(2): p. 149-60.

60. Weterings, E. and D.J. Chen, The endless tale of non-homologous end-joining.

Cell Res, 2008. 18(1): p. 114-24. 61. Jackson, S.P., Sensing and repairing DNA double-strand breaks. Carcinogenesis,

2002. 23(5): p. 687-96.

117

62. Kameoka, Y., H. Tagawa, S. Tsuzuki, S. Karnan, A. Ota, M. Suguro, R. Suzuki, M. Yamaguchi, Y. Morishima, S. Nakamura, and M. Seto, Contig array CGH at 3p14.2 points to the FRA3B/FHIT common fragile region as the target gene in

diffuse large B-cell lymphoma. Oncogene, 2004. 23(56): p. 9148-54. 63. Lee, S.H., W.H. Kim, H.K. Kim, K.M. Woo, H.S. Nam, H.S. Kim, J.G. Kim, and

M.H. Cho, Altered expression of the fragile histidine triad gene in primary gastric adenocarcinomas. Biochem Biophys Res Commun, 2001. 284(3): p. 850-5.

64. Michael, D., D.G. Beer, C.W. Wilke, D.E. Miller, and T.W. Glover, Frequent deletions of FHIT and FRA3B in Barrett's metaplasia and esophageal adenocarcinomas. Oncogene, 1997. 15(14): p. 1653-9.

65. Sozzi, G., M.L. Veronese, M. Negrini, R. Baffa, M.G. Cotticelli, H. Inoue, S.

Tornielli, S. Pilotti, L. De Gregorio, U. Pastorino, M.A. Pierotti, M. Ohta, K. Huebner, and C.M. Croce, The FHIT gene 3p14.2 is abnormal in lung cancer. Cell, 1996. 85(1): p. 17-26.

66. Virgilio, L., M. Shuster, S.M. Gollin, M.L. Veronese, M. Ohta, K. Huebner, and

C.M. Croce, FHIT gene alterations in head and neck squamous cell carcinomas. Proc Natl Acad Sci U S A, 1996. 93(18): p. 9770-5.

67. Huebner, K. and C.M. Croce, FRA3B and other common fragile sites: the weakest links. Nat Rev Cancer, 2001. 1(3): p. 214-21.

68. Driouch, K., H. Prydz, R. Monese, H. Johansen, R. Lidereau, and E. Frengen, Alternative transcripts of the candidate tumor suppressor gene, WWOX, are expressed at high levels in human breast tumors. Oncogene, 2002. 21(12): p. 1832-40.

69. Kuroki, T., F. Trapasso, T. Shiraishi, H. Alder, K. Mimori, M. Mori, and C.M. Croce, Genetic alterations of the tumor suppressor gene WWOX in esophageal squamous cell carcinoma. Cancer Res, 2002. 62(8): p. 2258-60.

70. Kuroki, T., S. Yendamuri, F. Trapasso, A. Matsuyama, R.I. Aqeilan, H. Alder, S.

Rattan, R. Cesari, M.L. Nolli, N.N. Williams, M. Mori, T. Kanematsu, and C.M. Croce, The tumor suppressor gene WWOX at FRA16D is involved in pancreatic carcinogenesis. Clin Cancer Res, 2004. 10(7): p. 2459-65.

71. Paige, A.J., K.J. Taylor, C. Taylor, S.G. Hillier, S. Farrington, D. Scott, D.J.

Porteous, J.F. Smyth, H. Gabra, and J.E. Watson, WWOX: a candidate tumor suppressor gene involved in multiple tumor types. Proc Natl Acad Sci U S A, 2001. 98(20): p. 11417-22.

72. Ried, K., M. Finnis, L. Hobson, M. Mangelsdorf, S. Dayan, J.K. Nancarrow, E.

Woollatt, G. Kremmidiotis, A. Gardner, D. Venter, E. Baker, and R.I. Richards, Common chromosomal fragile site FRA16D sequence: identification of the FOR gene spanning FRA16D and homozygous deletions and translocation breakpoints in cancer cells. Hum Mol Genet, 2000. 9(11): p. 1651-63.

73. Yendamuri, S., T. Kuroki, F. Trapasso, A.C. Henry, K.R. Dumon, K. Huebner, N.N. Williams, L.R. Kaiser, and C.M. Croce, WW domain containing oxidoreductase gene expression is altered in non-small cell lung cancer. Cancer Res, 2003. 63(4): p. 878-81.

74. McClintock, B., Chromosome organization and genic expression. Cold Spring Harb Symp Quant Biol, 1951. 16: p. 13-47.

118

75. Coquelle, A., E. Pipiras, F. Toledo, G. Buttin, and M. Debatisse, Expression of fragile sites triggers intrachromosomal mammalian gene amplification and sets boundaries to early amplicons. Cell, 1997. 89(2): p. 215-25.

76. Zimonjic, D.B., C.L. Keck-Waggoner, B.Z. Yuan, M.H. Kraus, and N.C. Popescu, Profile of genetic alterations and tumorigenicity of human breast cancer cells. Int J Oncol, 2000. 16(2): p. 221-30.

77. Hellman, A., E. Zlotorynski, S.W. Scherer, J. Cheung, J.B. Vincent, D.I. Smith,

L. Trakhtenbrot, and B. Kerem, A role for common fragile site induction in amplification of human oncogenes. Cancer Cell, 2002. 1(1): p. 89-97.

78. Durkin, S.G. and T.W. Glover, Chromosome fragile sites. Annu.Rev.Genet., 2007. 41: p. 169-192.

79. Yunis, J.J. and A.L. Soreng, Constitutive fragile sites and cancer. Science, 1984. 226(4679): p. 1199-204.

80. Re, A., D. Cora, A.M. Puliti, M. Caselle, and I. Sbrana, Correlated fragile site expression allows the identification of candidate fragile genes involved in

immunity and associated with carcinogenesis. BMC Bioinformatics, 2006. 7: p. 413.

81. Glover, T.W., J. Coyle-Morris, and R. Morgan, Fragile sites: overview, occurrence in acute nonlymphocytic leukemia and effects of caffeine on

expression. Cancer Genet Cytogenet, 1986. 19(1-2): p. 141-50. 82. Demirhan, O. and D. Tastemir, Cytogenetic effects of ethanol on chronic alcohol

users. Alcohol, 2008. 43(2): p. 127-36. 83. Kuwano, A. and T. Kajii, Synergistic effect of aphidicolin and ethanol on the

induction of common fragile sites. Hum.Genet., 1987. 75: p. 75-78. 84. Stein, C.K., T.W. Glover, J.L. Palmer, and B.S. Glisson, Direct correlation

between FRA3B expression and cigarette smoking. Genes Chromosomes Cancer, 2002. 34(3): p. 333-40.

85. Kao-Shan, C.S., R.L. Fine, J. Whang-Peng, E.C. Lee, and B.A. Chabner, Increased fragile sites and sister chromatid exchanges in bone marrow and peripheral blood of young cigarette smokers. Cancer Res, 1987. 47(23): p. 6278-82.

86. Musio, A. and I. Sbrana, Aphidicolin-sensitive specific common fragile sites: a biomarker of exposure to pesticides. Environ Mol Mutagen, 1997. 29(3): p. 250-5.

87. Sbrana, I. and A. Musio, Enhanced expression of common fragile site with occupational exposure to pesticides. Cancer Genet Cytogenet, 1995. 82(2): p.

123-7. 88. Webster, L.R., G.H. McKenzie, and H.T. Moriarty, Organophosphate-based

pesticides and genetic damage implicated in bladder cancer. Cancer Genet Cytogenet, 2002. 133(2): p. 112-7.

89. Yunis, J.J., A.L. Soreng, and A.E. Bowe, Fragile sites are targets of diverse mutagens and carcinogens. Oncogene, 1987. 1(1): p. 59-69.

90. Telez, M., E. Ortiz-Lastra, A.J. Gonzalez, P. Flores, I. Huerta, J.M. Ramirez, M. Barasoain, B. Criado, and I. Arrieta, Assessment of the genotoxicity of atenolol in

human peripheral blood lymphocytes: Correlation between chromosomal fragility and content of micronuclei. Mutat Res. 695(1-2): p. 46-54.

119

91. Coquelle, A., F. Toledo, S. Stern, A. Bieth, and M. Debatisse, A new role for hypoxia in tumor progression: induction of fragile site triggering genomic rearrangements and formation of complex DMs and HSRs. Mol Cell, 1998. 2(2):

p. 259-65. 92. Blair, A. and S.H. Zahm, Cancer among farmers. Occup Med, 1991. 6(3): p. 335-

54. 93. Brown, L.M., A. Blair, R. Gibson, G.D. Everett, K.P. Cantor, L.M. Schuman, L.F.

Burmeister, S.F. Van Lier, and F. Dick, Pesticide exposures and other agricultural risk factors for leukemia among men in Iowa and Minnesota. Cancer Res, 1990. 50(20): p. 6585-91.

94. Popescu, N.C., J.A. DiPaolo, and S.C. Amsbaugh, Integration sites of human

papillomavirus 18 DNA sequences on HeLa cell chromosomes. Cytogenet Cell Genet, 1987. 44(1): p. 58-62.

95. Feitelson, M.A. and J. Lee, Hepatitis B virus integration, fragile sites, and hepatocarcinogenesis. Cancer Lett, 2007. 252(2): p. 157-70.

96. Popescu, N.C., M.C. Chen, S. Simpson, S. Solinas, and J.A. DiPaolo, A Burkitt lymphoma cell line with integrated Epstein-Barr virus at a stable chromosome modification site. Virology, 1993. 195(1): p. 248-51.

97. Popescu, N.C., Genetic alterations in cancer as a result of breakage at fragile

sites. Cancer Letters, 2003. 192: p. 1-17. 98. Austin, H., E. Delzell, and P. Cole, Benzene and leukemia. A review of the

literature and a risk assessment. Am J Epidemiol, 1988. 127(3): p. 419-39. 99. Thirman, M.J., H.J. Gill, R.C. Burnett, D. Mbangkollo, N.R. McCabe, H.

Kobayashi, S. Ziemin-van der Poel, Y. Kaneko, R. Morgan, A.A. Sandberg, and et al., Rearrangement of the MLL gene in acute lymphoblastic and acute myeloid leukemias with 11q23 chromosomal translocations. N Engl J Med, 1993. 329(13): p. 909-14.

100. Bizzozero, O.J., Jr., K.G. Johnson, and A. Ciocco, Radiation-related leukemia in Hiroshima and Nagasaki, 1946-1964. I. Distribution, incidence and appearance time. N Engl J Med, 1966. 274(20): p. 1095-101.

101. Yoshinaga, S., K. Mabuchi, A.J. Sigurdson, M.M. Doody, and E. Ron, Cancer

risks among radiologists and radiologic technologists: review of epidemiologic studies. Radiology, 2004. 233(2): p. 313-21.

102. Sanz, G.F., M.A. Sanz, T. Vallespi, M.C. Canizo, M. Torrabadella, S. Garcia, D. Irriguible, and J.F. San Miguel, Two regression models and a scoring system for

predicting survival and planning treatment in myelodysplastic syndromes: a multivariate analysis of prognostic factors in 370 patients. Blood, 1989. 74(1): p. 395-408.

103. Horwitz, M., E.L. Goode, and G.P. Jarvik, Anticipation in familial leukemia. Am

J Hum Genet, 1996. 59(5): p. 990-8. 104. Horwitz, M., The genetics of familial leukemia. Leukemia, 1997. 11(8): p. 1347-

59. 105. Crittenden, L.B., An interpretation of familial aggregation based on multiple

genetic and environmental factors. Ann N Y Acad Sci, 1961. 91: p. 769-80. 106. Kuo, Y.H., S.F. Landrette, S.A. Heilman, P.N. Perrat, L. Garrett, P.P. Liu, M.M.

Le Beau, S.C. Kogan, and L.H. Castilla, Cbf beta-SMMHC induces distinct

120

abnormal myeloid progenitors able to develop acute myeloid leukemia. Cancer Cell, 2006. 9(1): p. 57-68.

107. Otto, F., M. Lubbert, and M. Stock, Upstream and downstream targets of RUNX

proteins. J Cell Biochem, 2003. 89(1): p. 9-18. 108. Look, A.T., Oncogenic transcription factors in the human acute leukemias.

Science, 1997. 278(5340): p. 1059-64. 109. Wang, Q., T. Stacy, M. Binder, M. Marin-Padilla, A.H. Sharpe, and N.A. Speck,

Disruption of the Cbfa2 gene causes necrosis and hemorrhaging in the central nervous system and blocks definitive hematopoiesis. Proc Natl Acad Sci U S A, 1996. 93(8): p. 3444-9.

110. Wang, Q., T. Stacy, J.D. Miller, A.F. Lewis, T.L. Gu, X. Huang, J.H. Bushweller,

J.C. Bories, F.W. Alt, G. Ryan, P.P. Liu, A. Wynshaw-Boris, M. Binder, M. Marin-Padilla, A.H. Sharpe, and N.A. Speck, The CBFbeta subunit is essential for CBFalpha2 (AML1) function in vivo. Cell, 1996. 87(4): p. 697-708.

111. Niki, M., H. Okada, H. Takano, J. Kuno, K. Tani, H. Hibino, S. Asano, Y. Ito, M.

Satake, and T. Noda, Hematopoiesis in the fetal liver is impaired by targeted mutagenesis of a gene encoding a non-DNA binding subunit of the transcription factor, polyomavirus enhancer binding protein 2/core binding factor. Proc Natl Acad Sci U S A, 1997. 94(11): p. 5697-702.

112. Okuda, T., J. van Deursen, S.W. Hiebert, G. Grosveld, and J.R. Downing, AML1, the target of multiple chromosomal translocations in human leukemia, is essential for normal fetal liver hematopoiesis. Cell, 1996. 84(2): p. 321-30.

113. Castilla, L.H., C. Wijmenga, Q. Wang, T. Stacy, N.A. Speck, M. Eckhaus, M.

Marin-Padilla, F.S. Collins, A. Wynshaw-Boris, and P.P. Liu, Failure of embryonic hematopoiesis and lethal hemorrhages in mouse embryos heterozygous for a knocked-in leukemia gene CBFB-MYH11. Cell, 1996. 87(4): p. 687-96.

114. Yergeau, D.A., C.J. Hetherington, Q. Wang, P. Zhang, A.H. Sharpe, M. Binder,

M. Marin-Padilla, D.G. Tenen, N.A. Speck, and D.E. Zhang, Embryonic lethality and impairment of haematopoiesis in mice heterozygous for an AML1-ETO fusion gene. Nat Genet, 1997. 15(3): p. 303-6.

115. Adya, N., T. Stacy, N.A. Speck, and P.P. Liu, The leukemic protein core binding

factor beta (CBFbeta)-smooth-muscle myosin heavy chain sequesters CBFalpha2 into cytoskeletal filaments and aggregates. Mol Cell Biol, 1998. 18(12): p. 7432-43.

116. Kanno, Y., T. Kanno, C. Sakakura, S.C. Bae, and Y. Ito, Cytoplasmic

sequestration of the polyomavirus enhancer binding protein 2 (PEBP2)/core binding factor alpha (CBFalpha) subunit by the leukemia-related PEBP2/CBFbeta-SMMHC fusion protein inhibits PEBP2/CBF-mediated transactivation. Mol Cell Biol, 1998. 18(7): p. 4252-61.

117. Lutterbach, B., Y. Hou, K.L. Durst, and S.W. Hiebert, The inv(16) encodes an acute myeloid leukemia 1 transcriptional corepressor. Proc Natl Acad Sci U S A, 1999. 96(22): p. 12822-7.

118. Castilla, L.H., L. Garrett, N. Adya, D. Orlic, A. Dutra, S. Anderson, J. Owens, M.

Eckhaus, D. Bodine, and P.P. Liu, The fusion gene Cbfb-MYH11 blocks myeloid differentiation and predisposes mice to acute myelomonocytic leukaemia. Nat Genet, 1999. 23(2): p. 144-6.

121

119. Castilla, L.H., P. Perrat, N.J. Martinez, S.F. Landrette, R. Keys, S. Oikemus, J. Flanegan, S. Heilman, L. Garrett, A. Dutra, S. Anderson, G.A. Pihan, L. Wolff, and P.P. Liu, Identification of genes that synergize with Cbfb-MYH11 in the

pathogenesis of acute myeloid leukemia. Proc Natl Acad Sci U S A, 2004. 101(14): p. 4924-9.

120. Gilliland, D.G., C.T. Jordan, and C.A. Felix, The molecular basis of leukemia. Hematology Am Soc Hematol Educ Program, 2004: p. 80-97.

121. Liu, P., S.A. Tarle, A. Hajra, D.F. Claxton, P. Marlton, M. Freedman, M.J. Siciliano, and F.S. Collins, Fusion between transcription factor CBF beta/PEBP2 beta and a myosin heavy chain in acute myeloid leukemia. Science, 1993. 261(5124): p. 1041-4.

122. Bloomfield, C.D., D. Lawrence, J.C. Byrd, A. Carroll, M.J. Pettenati, R. Tantravahi, S.R. Patil, F.R. Davey, D.T. Berg, C.A. Schiffer, D.C. Arthur, and R.J. Mayer, Frequency of prolonged remission duration after high-dose cytarabine intensification in acute myeloid leukemia varies by cytogenetic

subtype. Cancer Res, 1998. 58(18): p. 4173-9. 123. Byrd, J.C., R.K. Dodge, A. Carroll, M.R. Baer, C. Edwards, J. Stamberg, M.

Qumsiyeh, J.O. Moore, R.J. Mayer, F. Davey, C.A. Schiffer, and C.D. Bloomfield, Patients with t(8;21)(q22;q22) and acute myeloid leukemia have

superior failure-free and overall survival when repetitive cycles of high-dose cytarabine are administered. J Clin Oncol, 1999. 17(12): p. 3767-75.

124. Byrd, J.C., A.S. Ruppert, K. Mrozek, A.J. Carroll, C.G. Edwards, D.C. Arthur, M.J. Pettenati, J. Stamberg, P.R. Koduru, J.O. Moore, R.J. Mayer, F.R. Davey,

R.A. Larson, and C.D. Bloomfield, Repetitive cycles of high-dose cytarabine benefit patients with acute myeloid leukemia and inv(16)(p13q22) or t(16;16)(p13;q22): results from CALGB 8461. J Clin Oncol, 2004. 22(6): p. 1087-94.

125. Mrozek, K., G. Marcucci, P. Paschka, and C.D. Bloomfield, Advances in molecular genetics and treatment of core-binding factor acute myeloid leukemia. Curr Opin Oncol, 2008. 20(6): p. 711-8.

126. Keen-Kim, D., F. Nooraie, and P.N. Rao, Cytogenetic biomarkers for human

cancer. Front Biosci, 2008. 13: p. 5928-49. 127. O'Keefe, L.V. and R.I. Richards, Common chromosomal fragile sites and cancer:

focus on FRA16D. Cancer Lett, 2006. 232(1): p. 37-47. 128. Freudenreich, C.H., Molecular mechanisms of chromosome fragility. Chemtracts -

Biochemistry and Molecular Biology, 2005. 18: p. 141-152. 129. Novo, F.J., I.O. de Mendibil, and J.L. Vizmanos, TICdb: a collection of gene-

mapped translocation breakpoints in cancer. BMC Genomics, 2007. 8: p. 33. 130. Mitelman F, J.B., Mertens F., Mitelman Database of Chromosome Aberrations in

Cancer. 2008. 131. Smith, D.I., S. McAvoy, Y. Zhu, and D.S. Perez, Large common fragile site genes

and cancer. Semin Cancer Biol, 2007. 17(1): p. 31-41. 132. Arlt, M.F., S.G. Durkin, R.L. Ragland, and T.W. Glover, Common fragile sites as

targets for chromosome rearrangements. DNA Repair, 2006. 5: p. 1126-1135.

122

133. Fechter, A., I. Buettel, E. Kuehnel, M. Schwab, and L. Savelyeva, Cloning of genetically tagged chromosome break sequences reveals new fragile sites at 6p21 and 13q22. Int J Cancer, 2007. 120(11): p. 2359-67.

134. Lopes, M., C. Cotta-Ramusino, A. Pellicioli, G. Liberi, P. Plevani, M. Muzi-Falconi, C.S. Newlon, and M. Foiani, The DNA replication checkpoint response stabilizes stalled replication forks. Nature, 2001. 412(6846): p. 557-61.

135. Tercero, J.A. and J.F. Diffley, Regulation of DNA replication fork progression

through damaged DNA by the Mec1/Rad53 checkpoint. Nature, 2001. 412(6846): p. 553-7.

136. Kanaar, R., J.H. Hoeijmakers, and D.C. van Gent, Molecular mechanisms of DNA double strand break repair. Trends Cell Biol, 1998. 8(12): p. 483-9.

137. Pfeiffer, P., W. Goedecke, and G. Obe, Mechanisms of DNA double-strand break repair and their potential to induce chromosomal aberrations. Mutagenesis, 2000. 15(4): p. 289-302.

138. Gottlich, B., S. Reichenberger, E. Feldmann, and P. Pfeiffer, Rejoining of DNA

double-strand breaks in vitro by single-strand annealing. Eur J Biochem, 1998. 258(2): p. 387-95.

139. Lieber, M.R., S.C. Raghavan, and K. Yu, Mechanistic aspects of lymphoid chromosomal translocations. J Natl Cancer Inst Monogr, 2008(39): p. 8-11.

140. Sutherland GR, B.E., Richards RI, Fragile Sites. In Encyclopedia of Molecular Biology and Molecular Medicine, 1996. 2: p. 313-318.

141. Rozier, L., E. El-Achkar, F. Apiou, and M. Debatisse, Characterization of a conserved aphidicolin-sensitive common fragile site at human 4q22 and mouse

6C1: possible association with an inherited disease and cancer. Oncogene, 2004. 23(41): p. 6872-80.

142. Karadeniz, N.N., Y. Tunca, and N. Imirzalioglu, New heritable fragile site at 15q13 in both members of a nonconsanguineous couple. Am J Med Genet A,

2003. 118A(3): p. 290-2. 143. Anderson, S. and M.L. DePamphilis, Metabolism of Okazaki fragments during

simian virus 40 DNA replication. J Biol Chem, 1979. 254(22): p. 11495-504. 144. Hay, R.T. and M.L. DePamphilis, Initiation of SV40 DNA replication in vivo:

location and structure of 5' ends of DNA synthesized in the ori region. Cell, 1982. 28(4): p. 767-79.

145. Wang, Y.H., Chromatin structure of human chromosomal fragile sites. Cancer Letters, 2006. 232: p. 70-78.

146. Hsu, Y.Y. and Y.H. Wang, Human fragile site FRA16B DNA excludes nucleosomes in the presence of distamycin. J Biol Chem, 2002. 277(19): p. 17315-9.

147. Wang, Y.H., R. Gellibolian, M. Shimizu, R.D. Wells, and J. Griffith, Long CCG

triplet repeat blocks exclude nucleosomes: a possible mechanism for the nature of fragile sites in chromosomes. J Mol Biol, 1996. 263(4): p. 511-6.

148. Wang, Y.H. and J. Griffith, Methylation of expanded CCG triplet repeat DNA from fragile X syndrome patients enhances nucleosome exclusion. J Biol Chem,

1996. 271(38): p. 22937-40.

123

149. Godde, J.S., S.U. Kass, M.C. Hirst, and A.P. Wolffe, Nucleosome assembly on methylated CGG triplet repeats in the fragile X mental retardation gene 1 promoter. J Biol Chem, 1996. 271(40): p. 24325-8.

150. Pearson, C.E. and R.R. Sinden, Alternative structures in duplex DNA formed within the trinucleotide repeats of the myotonic dystrophy and fragile X loci. Biochemistry, 1996. 35(15): p. 5041-53.

151. Ragland, R.L., M.W. Glynn, M.F. Arlt, and T.W. Glover, Stably transfected

common fragile site sequences exhibit instability at ectopic sites. Genes Chromosomes Cancer, 2008. 47(10): p. 860-72.

152. Pearson, C.E., Ruiz, M.T., Price, G.B., and Zannis-Hadjopoulos, M., Cruciform DNA Binding Protein in HeLa Cell Extracts. Biochemistry, 1994. 33(47): p.

14185-14196. 153. Pearson, C.E., M. Zannis-Hadjopoulos, G.B. Price, and H. Zorbas, A novel type of

interaction between cruciform DNA and a cruciform binding protein from HeLa cells. EMBO J, 1995. 14(7): p. 1571-80.

154. Griffith, J.D. and G. Christiansen, Electron microscope visualization of chromatin and other DNA-protein complexes. Annu Rev Biophys Bioeng, 1978. 7: p. 19-35.

155. Hirt, B., Selective extraction of polyoma DNA from infected mouse cell cultures. J Mol Biol, 1967. 26(2): p. 365-9.

156. Cleary, J.D., K. Nichol, Y.H. Wang, and C.E. Pearson, Evidence of cis-acting factors in replication-mediated trinucleotide repeat instability in primate cells. Nat Genet, 2002. 31(1): p. 37-46.

157. Subramanian, D. and J.D. Griffith, p53 Monitors replication fork regression by

binding to "chickenfoot" intermediates. J Biol Chem, 2005. 280(52): p. 42568-72. 158. Dere, R., M. Napierala, L.P. Ranum, and R.D. Wells, Hairpin structure-forming

propensity of the (CCTG.CAGG) tetranucleotide repeats contributes to the genetic instability associated with myotonic dystrophy type 2. J Biol Chem, 2004.

279(40): p. 41715-26. 159. Rindler, P.M., Clark, R.M., Pollard, L.M., De Biase, I., and Bidichandani, S.I.,

Replication in mammalian cells recapitulates the locus-specific differences in somatic instability of genomic GAA triplet-repeats. Nucleic Acids Res, 2006.

34(21): p. 6352-6361. 160. Edamura, K.N., Leonard, M.R., and Pearson, C.E., Role of replication and CpG

methylation in fragile X syndrome CGG deletions in primate cells. Am J Hum Genet, 2005. 76: p. 302-311.

161. Kerrest, A., R.P. Anand, R. Sundararajan, R. Bermejo, G. Liberi, B. Dujon, C.H. Freudenreich, and G.F. Richard, SRS2 and SGS1 prevent chromosomal breaks and stabilize triplet repeats by restraining recombination. Nat Struct Mol Biol, 2009. 16(2): p. 159-67.

162. Yoon, D., Y. Wang, K. Stapleford, L. Wiesmuller, and J. Chen, P53 inhibits strand exchange and replication fork regression promoted by human Rad51. J Mol Biol, 2004. 336(3): p. 639-54.

163. Lavrik, O.I., D.M. Kolpashchikov, K. Weisshart, H.P. Nasheuer, S.N. Khodyreva,

and A. Favre, RPA subunit arrangement near the 3'-end of the primer is modulated by the length of the template strand and cooperative protein interactions. Nucleic Acids Res, 1999. 27(21): p. 4235-40.

124

164. Fouche, N., S. Ozgur, D. Roy, and J.D. Griffith, Replication fork regression in repetitive DNAs. Nucleic Acids Res, 2006. 34(20): p. 6044-50.

165. Ohshima, K. and R.D. Wells, Hairpin formation during DNA synthesis primer

realignment in vitro in triplet repeat sequences from human hereditary disease genes. J Biol Chem, 1997. 272(27): p. 16798-806.

166. Meaburn, K.J., T. Misteli, and E. Soutoglou, Spatial genome organization in the formation of chromosomal translocations. Semin Cancer Biol, 2007. 17(1): p. 80-

90. 167. Elliott, B. and M. Jasin, Double-strand breaks and translocations in cancer. Cell

Mol Life Sci, 2002. 59(2): p. 373-85. 168. Manvelyan, M., P. Kempf, A. Weise, K. Mrasek, A. Heller, A. Lier, K. Hoffken,

H.J. Fricke, H.G. Sayer, T. Liehr, and H. Mkrtchyan, Preferred co-localization of chromosome 8 and 21 in myeloid bone marrow cells detected by three dimensional molecular cytogenetics. Int J Mol Med, 2009. 24(3): p. 335-41.

169. Roix, J.J., P.G. McQueen, P.J. Munson, L.A. Parada, and T. Misteli, Spatial

proximity of translocation-prone gene loci in human lymphomas. Nat Genet, 2003. 34(3): p. 287-91.

170. Nikiforova, M.N., J.R. Stringer, R. Blough, M. Medvedovic, J.A. Fagin, and Y.E. Nikiforov, Proximity of chromosomal loci that participate in radiation-induced

rearrangements in human cells. Science, 2000. 290(5489): p. 138-41. 171. Gandhi, M., M. Medvedovic, J.R. Stringer, and Y.E. Nikiforov, Interphase

chromosome folding determines spatial proximity of genes participating in carcinogenic RET/PTC rearrangements. Oncogene, 2006. 25(16): p. 2360-6.

172. Roccato, E., P. Bressan, G. Sabatella, C. Rumio, L. Vizzotto, M.A. Pierotti, and A. Greco, Proximity of TPR and NTRK1 rearranging loci in human thyrocytes. Cancer Res, 2005. 65(7): p. 2572-6.

173. Gandhi, M., L.W. Dillon, S. Pramanik, Y.E. Nikiforov, and Y.H. Wang, DNA

breaks at fragile sites generate oncogenic RET/PTC rearrangements in human thyroid cells. Oncogene, 2010.

174. Mathas, S., S. Kreher, K.J. Meaburn, K. Johrens, B. Lamprecht, C. Assaf, W. Sterry, M.E. Kadin, M. Daibata, S. Joos, M. Hummel, H. Stein, M. Janz, I.

Anagnostopoulos, E. Schrock, T. Miste li, and B. Dorken, Gene deregulation and spatial genome reorganization near breakpoints prior to formation of translocations in anaplastic large cell lymphoma. Proc Natl Acad Sci U S A, 2009. 106(14): p. 5831-6.

175. Mani, R.S., S.A. Tomlins, K. Callahan, A. Ghosh, M.K. Nyati, S. Varambally, N. Palanisamy, and A.M. Chinnaiyan, Induced chromosomal proximity and gene fusions in prostate cancer. Science, 2009. 326(5957): p. 1230.

176. Lin, C., L. Yang, B. Tanasa, K. Hutt, B.G. Ju, K. Ohgi, J. Zhang, D.W. Rose,

X.D. Fu, C.K. Glass, and M.G. Rosenfeld, Nuclear receptor-induced chromosomal proximity and DNA breaks underlie specific translocations in cancer. Cell, 2009. 139(6): p. 1069-83.

177. Mitelman F, J.B.a.M.F. Mitelman Database of Chromosome Aberrations and

Gene Fusions in Cancer. 2010 [cited; Available from: http://cgap.nci.nih.gov/Chromosomes/Mitelman.

125

178. Lukasova, E., S. Kozubek, M. Kozubek, J. Kjeronska, L. Ryznar, J. Horakova, E. Krahulcova, and G. Horneck, Localisation and distance between ABL and BCR genes in interphase nuclei of bone marrow cells of control donors and patients

with chronic myeloid leukaemia. Hum Genet, 1997. 100(5-6): p. 525-35. 179. Kozubek, S., E. Lukasova, L. Ryznar, M. Kozubek, A. Liskova, R.D. Govorun,

E.A. Krasavin, and G. Horneck, Distribution of ABL and BCR genes in cell nuclei of normal and irradiated lymphocytes. Blood, 1997. 89(12): p. 4537-45.

180. Neves, H., C. Ramos, M.G. da Silva, A. Parreira, and L. Parreira, The nuclear topography of ABL, BCR, PML, and RARalpha genes: evidence for gene proximity in specific phases of the cell cycle and stages of hematopoietic differentiation. Blood, 1999. 93(4): p. 1197-207.

181. Kundu, M., A. Chen, S. Anderson, M. Kirby, L. Xu, L.H. Castilla, D. Bodine, and P.P. Liu, Role of Cbfb in hematopoiesis and perturbations resulting from expression of the leukemogenic fusion gene Cbfb-MYH11. Blood, 2002. 100(7): p. 2449-56.

182. Dimitrova, D.S. and D.M. Gilbert, Temporally coordinated assembly and disassembly of replication factories in the absence of DNA synthesis. Nature Cell Biology, 2000. 2: p. 686-694.

183. Sarkaria, J.N., E.C. Busby, R.S. Tibbetts, P. Roos, Y. Taya, L.M. Karnitz, and

R.T. Abraham, Inhibition of ATM and ATR kinase activities by the radiosensitizing agent, caffeine. Cancer Research, 1999. 59: p. 4375-4382.

184. Kozubek, S., E. Lukasova, A. Mareckova, M. Skalnikova, M. Kozubek, E. Bartova, V. Kroha, E. Krahulcova, and J. Slotova, The topological organization of

chromosomes 9 and 22 in cell nuclei has a determinative role in the induction of t(9,22) translocations and in the pathogenesis of t(9,22) leukemias. Chromosoma, 1999. 108(7): p. 426-35.

185. Folle, G.A., Nuclear architecture, chromosome domains and genetic damage.

Mutat Res, 2008. 658(3): p. 172-83. 186. Savage, J.R., Cancer. Proximity matters. Science, 2000. 290(5489): p. 62-3. 187. Aten, J.A., J. Stap, P.M. Krawczyk, C.H. van Oven, R.A. Hoebe, J. Essers, and R.

Kanaar, Dynamics of DNA double-strand breaks revealed by clustering of

damaged chromosome domains. Science, 2004. 303(5654): p. 92-5. 188. Savage, J.R., Interchange and intra-nuclear architecture. Environ Mol Mutagen,

1993. 22(4): p. 234-44. 189. van der Reijden, B.A., H.G. Dauwerse, R.H. Giles, S. Jagmohan-Changur, C.

Wijmenga, P.P. Liu, B. Smit, H.W. Wessels, G.C. Beverstock, M. Jotterand-Bellomo, D. Martinet, D. Muhlematter, M. Lafage-Pochitaloff, J. Gabert, J. Reiffers, C. Bilhou-Nabera, G.J. van Ommen, A. Hagemeijer, and M.H. Breuning, Genomic acute myeloid leukemia-associated inv(16)(p13q22)

breakpoints are tightly clustered. Oncogene, 1999. 18(2): p. 543-50. 190. Kong, Q. and N. Maizels, Breaksite batch mapping, a rapid method for assay and

identification of DNA breaksites in mammalian cells. Nucleic Acids Res, 2001. 29(6): p. E33.

191. Wang, L., J. Darling, J.S. Zhang, H. Huang, W. Liu, and D.I. Smith, Allele-specific late replication and fragility of the most active common fragile site, FRA3B. Hum Mol Genet, 1999. 8(3): p. 431-7.

126

192. van Dongen, J.J., E.A. Macintyre, J.A. Gabert, E. Delabesse, V. Rossi, G. Saglio, E. Gottardi, A. Rambaldi, G. Dotti, F. Griesinger, A. Parreira, P. Gameiro, M.G. Diaz, M. Malec, A.W. Langerak, J.F. San Miguel, and A. Biondi, Standardized

RT-PCR analysis of fusion gene transcripts from chromosome aberrations in acute leukemia for detection of minimal residual disease. Report of the BIOMED-1 Concerted Action: investigation of minimal residual disease in acute leukemia. Leukemia, 1999. 13(12): p. 1901-28.

193. Lieber, M.R., The mechanism of double-strand DNA break repair by the nonhomologous DNA end-joining pathway. Annu Rev Biochem, 2010. 79: p. 181-211.

194. Sharples, G.J., The X philes: structure-specific endonucleases that resolve

Holliday junctions. Mol Microbiol, 2001. 39(4): p. 823-34.

127

APPENDIX

Table 1: Comprehensive list of gene pairs involved in cancer-specific recurrent

translocations which result in fusion transcripts

Translocation Gene Location Fragile Site Gene Location Fragile Site

t(7;12)(p22;q13) ACTB 7p22.1 FRA7B (common,

apc) GLI1 12q13.3

inv(7)(q21q34) AKAP9 7q21.2 FRA7E (common, apc) BRAF 7q34

t(X;17)(p11;q25) ASPSCR1 17q25.3

TFE3 Xp11.23

inv(2)(p23q35) ATIC 2q35

ALK 2p23.2-

p23.1

t(17;20)(q23;q13) BCAS4 20q13.13

BCAS3

17q23.2

t(2;3)(p16;q26) BCL11A 2p16.1

MDS1 3q26.2

t(5;14)(q35;q32) BCL11B 14q32.2

NKX2E

5q35.2 FRA5G (rare, folic

acid)

t(5;14)(q35;q32) BCL11B 14q32.2

TLX3 5q35.1 FRA5G (rare, folic

acid)

inv(14)(q11q32) BCL11B 14q32.2

TRD@

14q11.2

t(14;18)(q32;q21) BCL2 18q21.33 FRA18B (common,

apc) IGH@ 14q32.33

t(2;18)(p11;q21) BCL2 18q21.33 FRA18B (common,

apc) IGK@ 2p11.2

FRA2L (rare, folic

acid)

t(18;22)(q21;q11) BCL2 18q21.33 FRA18B (common,

apc) IGL@ 22q11.22

t(8;19)(q24;q13) BCL3 19q13.31 FRA19A (common, 5-

aza) MYC 8q24.21

t(3;16)(q27;p13) BCL6 3q27.3 FRA3C (common,

apc) CIITA 16p13.13

t(3;8)(q27;q24) BCL6 3q27.3 FRA3C (common,

apc) MYC 8q24.21

t(1;14)(q21;q32) BCL9 1q21.1 FRA1F (common, apc) IGH@ 14q32.33

t(1;22)(q21;q11) BCL9 1q21.1 FRA1F (common, apc) IGL@ 22q11.22

t(9;22)(q34;q11) BCR 22q11.23

ABL1 9q34.12

t(8;22)(p12;q11) BCR 22q11.23

FGFR1

8p12

t(9;22)(p24;q11) BCR 22q11.23

JAK2 9p24.1

t(4;22)(q12;q11) BCR 22q11.23

PDGFRA

4q12 FRA4B (common,

BrdU)

t(11;18)(q22;q21) BIRC3 11q22.2

MAL

T1 18q21.32

FRA18B (common,

apc)

t(X;11)(q21;q23) BRWD3 Xq21.1

ARH

GAP20

11q22.3-q23.1

t(8;12)(q21;q22) BTG1 12q21.33 FRA12B (common,

apc) MYC 8q24.21

t(7;15)(p21;q21) C15ORF21 15q21.1

ETV1 7p21.2

t(3;3)(q21;q26) C3ORF27 3q21.3

EVI1 3q26.2

t(2;11)(p23;p15) CARS 11p15.4

ALK 2p23.2-p23.1

t(16;16)(p13;q22),

inv(16)(p13q22) CBFB 16q22.1

FRA16B (rare, dist A),

FRA16C (common, apc)

MYH

11 16p13.11

FRA16A (rare, folic

acid)

t(5;10)(q33;q21) CCDC6 10q21.2 FRA10C (common,

BrdU)

PDGF

RB 5q33.1

inv(10)(q11q21) CCDC6 10q21.2 FRA10C (common,

BrdU) RET 10q11.21

FRA10G (common,

apc)

t(5;14)(q33;q32) CCDC88C 14q32.12

PDGFRB

5q33.1

128

t(11;19)(q13;p13) CCND1 11q13 .2 FRA11H (common,

apc) FSTL

3 19p13.3

FRA19B (rare, folic acid)

t(5;6)(q32-33;q22) CD74 5q33.1

ROS1 6q22.2

t(16;17)(q21;p13) CDH11 16q21

USP6 17p13.2

t(7;11)(q21;q23) CDK6 7q21.2 FRA7E (common, apc) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

t(5;7)(q35;q21) CDK6 7q21.2 FRA7E (common, apc) TLX3 5q35.1 FRA5G (rare, folic

acid)

t(5;11)(q12;q23) CENPK 5q12.3

MLL 11q23.3

FRA11B (rare, folic

acid), FRA11G (common, apc)

t(8;9)(p12;q33) CEP110 9q33.2

FGFR

1 8p12

t(4;12)(q12;p13) CHIC2 4q12 FRA4B (common,

BrdU) ETV6 12p13.2

t(4;19)(q35;q13) CIC 19q13.2 FRA19A (common, 5-

aza) DUX4 4q35.2

t(2;17)(p23;q23) CLTC 17q23.1 FRA17B (common,

apc) ALK

2p23.2-

p23.1

t(X;17)(p11;q23) CLTC 17q23.1 FRA17B (common,

apc) TFE3 Xp11.23

t(2;22)(p23;q11) CLTCL1 22q11.21

ALK 2p23.2-p23.1

t(3;17)(q21;p13) CNBP 3q21.3

USP6 17p13.2

t(17;22)(q21;q13) COL1A1 17q21.33

PDGFB

22q13.1 FRA22A (rare, folic

acid)

t(17;17)(p13;q21) COL1A1 17q21.33

USP6 17p13.2

t(7;8)(q21;q12) COL1A2 7q21.3

PLAG1

8q12.1

t(X;6)(q22;q13-14) COL4A5 Xq22.3

COL1

2A1 6q13-q14.1

FRA6D (common,

BrdU)

t(1;2)(p13;q37) COL6A3 2q37.3 FRA2J (common, apc) CSF1 1p13.3

t(8;12)(p12;q15) CPSF6 12q15

FGFR1

8p12

t(11;19)(q21;p13) CRTC1 19p13.11 FRA19B (rare, folic

acid)

MAM

L2 11q21

t(11;15)(q21;q26) CRTC3 15q26.1

MAML2

11q21

t(3;8)(p22;q12) CTNNB1 3p22.1

PLAG1

8q12.1

t(3;9)(q27;p24) DMRT1 9p24.3

BCL6 3q27.3 FRA3C (common, apc)

t(1;1)(p36;q41) DUSP10 1q41

PRDM16

1p36.32 FRA1A (common,

apc)

t(5;12)(q33;q14) EBF1 5q33.3

LOC2

04010 12q14.3

t(X;21)(q25;q22) ELF4 Xq25

ERG 21q22.2

t(9;14)(q34;q32) EML1 14q32.2

ABL1 9q34.12

inv(2)(p21p23), del(2)(p21p23)*

EML4 2p21

ALK 2p23.2-p23.1

t(5;12)(q33;p13) ERC1 12p13.33

PDGF

RB 5q33.1

t(10;12)(q11;p13) ERC1 12p13.33

RET 10q11.21 FRA10G (common,

apc)

t(9;12)(q34;p13) ETV6 12p13.2

ABL1 9q34.12

t(1;12)(q25;p13) ETV6 12p13.2

ABL2 1q25.2

t(5;12)(q31;p13) ETV6 12p13.2

ACSL6

5q31.1 FRA5C (common, apc)

t(1;12)(q21;p13) ETV6 12p13.2

ARNT 1q21.2 FRA1F (common, apc)

t(12;12)(p13;q13) ETV6 12p13.2

BAZ2A

12q13.3

129

t(12;13)(p13;q12) ETV6 12p13.2

CDX2 13q12.2

t(3;12)(q26;p13) ETV6 12p13.2

EVI1 3q26.2

t(4;12)(p16;p13) ETV6 12p13.2

FGFR3

4p16.3

t(12;13)(p13;q12) ETV6 12p13.2

FLT3 13q12.2

t(6;12)(q22;p13) ETV6 12p13.2

FRK 6q22.1

t(10;12)(q24;p13) ETV6 12p13.2

GOT1 10q24.2 FRA10A (rare, folic

acid)

t(9;12)(p24;p13) ETV6 12p13.2

JAK2 9p24.1

t(3;12)(q26;p13) ETV6 12p13.2

MDS1 3q26.2

t(1;12)(p36;p13) ETV6 12p13.2

MDS2 1p36.11 FRA1A (common,

apc)

t(12;15)(p13;q25) ETV6 12p13.2

NTRK

3 15q25.3

t(4;12)(q12;p13) ETV6 12p13.2

PDGFRA

4q12 FRA4B (common,

BrdU)

t(5;12)(q33;p13) ETV6 12p13.2

PDGFRB

5q33.1

t(12;17)(p13;p13) ETV6 12p13.2

PER1 17p13.1

inv(12)(p13q15) ETV6 12p13.2

PTPRR

12q15

t(12;21)(p13;q22) ETV6 12p13.2

RUN

X1 21q22.12

t(6;12)(q23;p13) ETV6 12p13.2

STL 6q23

t(9;12)(q22;p13) ETV6 12p13.2

SYK 9q22.2

t(12;22)(q13;q12) EWSR1 22q12.2 FRA22B (common,

apc) ATF1 12q13.13

FRA12A (rare, folic acid)


apc)

CREB

1 2q33.3 FRA2I (common,apc)


apc) DDIT

3 12q13.3


apc) ERG 21q22.2

t(7;22)(p21;q12) EWSR1 22q12.2 FRA22B (common,

apc) ETV1 7p21.2


apc) ETV4 17q21.31


apc) FEV 2q35


apc) FLI1 11q24.3


apc) NR4A

3 9q31.1

inv(22)(q12q12) EWSR1 22q12.2 FRA22B (common,

apc)

PATZ

1 22q12.2

FRA22B (common,

apc)


apc) POU5

F1 6p21.33

FRA6H (common, apc)


apc) SP3 2q31.1

FRA2G (common, apc)


apc) WT1 11p13

FRA11E (common,

apc)


apc) ZNF3

84 12p13.31

t(5;7)(q31;q34) FCHSD1 5q31.3

BRAF 7q34

t(6;8)(q27;p12) FGFR1OP 6q27

FGFR1

8p12

del(4)(q12q12)* FIP1L1 4q12 FRA4B (common,

BrdU) PDGF

RA 4q12

FRA4B (common, BrdU)

t(4;17)(q12;q21) FIP1L1 4q12 FRA4B (common,

BrdU) RARA 17q21.2

t(2;13)(q36;q14) FOXO1A 13q14.11

PAX3 2q36.1

130

t(X;11)(q13;q23) FOXO4 Xq13.1

MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

t(12;16)(q13;p11) FUS 16p11.2

ATF1 12q13.13 FRA12A (rare, folic

acid)

t(11;16)(p11;p11) FUS 16p11.2

CREB

3L1 11p11.2

t(7;16)(q34;p11) FUS 16p11.2

CREB3L2

7q33-q34

t(12;16)(q13;p11) FUS 16p11.2

DDIT3

12q13.3

t(16;21)(p11;q22) FUS 16p11.2

ERG 21q22.2

t(2;16)(q35;p11) FUS 16p11.2

FEV 2q35

t(3;12)(q27;p13) GAPDH 12p13.31


t(5;12)(q33;q24) GIT2 12q24.11 FRA12E (common,

apc) PDGF

RB 5q33.1

t(10;14)(q11;q32) GOLGA5 14q32.12


apc)

del(6)(q21q22)* GOPC 6q22.2

ROS1 6q22.2

del(8)(q12q24)* HAS2 8q24.13

FRA8C (common,

apc), FRA8E (rare, dist A)

PLAG

1 8q12.1

t(8;19)(p12;q13)

HERV-K

(LOC113386)

19q13.43 FRA19A (common, 5-

aza)

FGFR

1 8p12

t(5;7)(q33;q11) HIP1 7q11.23 FRA7J (common, apc) PDGF

RB 5q33.1

t(3;6)(q27;p22) HIST1H4I 6p22.1


inv(6)(p21q21) HMGA1 6p21.31 FRA6H (common,

apc) LAMA4

6q21 FRA6F (common, apc)

t(12;14)(q14;q11) HMGA2 12q14.3

CCNB1IP1

14q11.2

t(8;12)(q22;q14) HMGA2 12q14.3

COX6

C 8q22.2

t(2;12)(q37;q14) HMGA2 12q14.3

CXCR7

2q37.3 FRA2J (common, apc)

t(5;12)(q33;q14) HMGA2 12q14.3

EBF1 5q33.3

t(12;13)(q14;q13) HMGA2 12q14.3

LHFP 13q13.3

t(3;12)(q28;q14) HMGA2 12q14.3

LPP 3q28

t(9;12)(p23;q14) HMGA2 12q14.3

NFIB 9p23-p22.3

t(12;14)(q14;q24) HMGA2 12q14.3

RAD5

1L1 14q24.1

FRA14C (common,

apc)

t(7;7)(p15;p21) HNRPA2B1 7p15.2

ETV1 7p21.2

t(8;10)(p11;q11) HOOK3 8p11.21


apc)

t(6;16)(p21;q22) HP 16q22.3

MRPS10

6p21.1 FRA6H (common,

apc)

t(3;14)(q27;q32) HSP90AA1 14q32.31


t(3;6)(q27;p21) HSP90AB1 6p21.1 FRA6H (common,

apc) BCL6 3q27.3 FRA3C (common, apc)

t(1;14)(p22;q32) IGH@ 14q32.33

BCL10

1p22.3 FRA1D (common,

apc)

t(2;14)(p16;q32) IGH@ 14q32.33

BCL1

1A 2p16.1

t(14;19)(q32;q13) IGH@ 14q32.33

BCL3 19q13.31 FRA19A (common, 5-

aza)

t(3;14)(q27;q32) IGH@ 14q32.33


t(14;15)(q32;q11-13)

IGH@ 14q32.33

BCL8 15q11.2

t(11;14)(q13;q32) IGH@ 14q32.33

CCND

1 11q13.2

FRA11A (rare, folic

acid), FRA11H (common, apc)

131

t(12;14)(p13;q32) IGH@ 14q32.33

CCND2

12p13.32

t(6;14)(p21;q32) IGH@ 14q32.33

CCND

3 6p21.1

FRA6H (common,

apc)

t(7;14)(q21;q32) IGH@ 14q32.33

CDK6 7q21.2 FRA7E (common, apc)

t(14;19)(q32;q13) IGH@ 14q32.33

CEBP

A 19q13.11

FRA19A (common, 5-

aza)

t(14;20)(q32;q13) IGH@ 14q32.33

CEBPB

20q13.13

t(8;14)(q11;q32) IGH@ 14q32.33

CEBPD

8q11.21

t(14;14)(q11;q32) IGH@ 14q32.33

CEBP

E 14q11.2

t(14;19)(q32;q13) IGH@ 14q32.33

CEBPG


aza)

t(12;14)(q23;q32) IGH@ 14q32.33

CHST

11 12q23.3

t(11;14)(q23;q32) IGH@ 14q32.33

DDX6 11q23.3

FRA11B (rare, folic


t(7;14)(q21;q32) IGH@ 14q32.33

ERV

WE1 7q21.2 FRA7E (common, apc)

t(12;14)(p13;q32) IGH@ 14q32.33

ETV6 12p13.2

t(1;14)(q23;q32) IGH@ 14q32.33

FCGR

2B 1q23.3

t(1;14)(q21;q32) IGH@ 14q32.33

FCRL4

1q23.1

t(4;14)(p16;q32) IGH@ 14q32.33

FGFR

3 4p16.3

t(3;14)(p14;q32) IGH@ 14q32.33

FOXP

1 3p14.1

t(6;14)(p22;q32) IGH@ 14q32.33

ID4 6p22.3

t(14;22)(q32;q11) IGH@ 14q32.33

IGL@ 22q11.22

t(5;14)(q31;q32) IGH@ 14q32.33

IL3 5q31.1 FRA5C (common, apc)

t(6;14)(p25;q32) IGH@ 14q32.33

IRF4 6p25.3

t(1;14)(p35;q32) IGH@ 14q32.33

LAPT

M5 1p35.2

t(1;14)(q25;q32) IGH@ 14q32.33

LHX4 1q25.2

t(14;16)(q32;q23) IGH@ 14q32.33

MAF 16q23.1

t(14;20)(q32;q12) IGH@ 14q32.33

MAFB

20q12

t(14;18)(q32;q21) IGH@ 14q32.33

MALT1

18q21.32 FRA18B (common,

apc)

t(1;14)(q22;q32) IGH@ 14q32.33

MUC1 1q22

t(8;14)(q24;q32) IGH@ 14q32.33

MYC 8q24.21

t(10;14)(q24;q32) IGH@ 14q32.33

NFKB

2 10q24.32

t(11;14)(q23;q32) IGH@ 14q32.33

PAFAH1B2

11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

t(9;14)(p13;q32) IGH@ 14q32.33

PAX5 9p13.2

t(11;14)(q23;q32) IGH@ 14q32.33

PCSK

7 11q23.3

FRA11B (rare, folic acid), FRA11G

(common, apc)

t(4;14)(p14;q32) IGH@ 14q32.33

RHOH

4p14

t(14;19)(q32;q13) IGH@ 14q32.33

SPIB 19q13.33 FRA19A (common, 5-

aza)

t(14;14)(q11;q32), inv(14)(q11q32)

IGH@ 14q32.33

TRA@

14q11.2

inv(14)(q11q32) IGH@ 14q32.33

TRD@

14q11.2

t(4;14)(p16;q32) IGH@ 14q32.33

WHS

C1 4p16.3

132

t(14;16)(q32;q23) IGH@ 14q32.33

WWOX

16q23.1 FRA16D (common,

apc)

t(1;2)(p22;p11) IGK@ 2p11.2 FRA2L (rare, folic

acid)

BCL1

0 1p22.3

FRA1D (common,

apc)

t(2;19)(p11;q13) IGK@ 2p11.2 FRA2L (rare, folic

acid) BCL3 19q13.31

FRA19A (common, 5-aza)


acid) BCL6 3q27.3 FRA3C (common, apc)


acid) CCND

1 11q13.2

FRA11A (rare, folic


t(2;12)(p11;p13) IGK@ 2p11.2 FRA2L (rare, folic

acid)

CCND

2 12p13.32


acid) CDK6 7q21.2 FRA7E (common, apc)


acid) FVT1 18q21.33

FRA18B (common,

apc)


acid) MYC 8q24.21


acid) PVT1 8q24.21


acid)

ZC3H

12D 6q25.1

t(19;22)(q13;q11) IGL@ 22q11.22-

q11.23 BCL3 19q13.31

FRA19A (common, 5-aza)

t(3;22)(q27;q11) IGL@ 22q11.22-

q11.23 BCL6 3q27.3 FRA3C (common, apc)

t(11;22)(q13;q11) IGL@ 22q11.22-

q11.23 CCND

1 11q13.2

FRA11A (rare, folic


t(12;22)(p13;q11) IGL@ 22q11.22-

q11.23

CCND

2 12p13.32

t(6;22)(p21;q11) IGL@ 22q11.22-

q11.23 CCND

3 6p21.1

FRA6H (common, apc)

t(7;22)(q21;q11) IGL@ 22q11.22-

q11.23 CDK6 7q21.2 FRA7E (common, apc)

t(16;22)(q23;q11) IGL@ 22q11.22-

q11.23 MAF 16q23.1

t(8;22)(q24;q11) IGL@ 22q11.22-

q11.23 MYC 8q24.21

t(8;22)(q24;q11) IGL@ 22q11.22-

q11.23 PVT1 8q24.21

t(2;22)(p16;q11) IGL@ 22q11.22-

q11.23 REL 2p16.1

t(16;22)(q23;q11) IGL@ 22q11.22-

q11.23 WWO

X 16q23.1

FRA16D (common, apc)

t(4;16)(q27;p13) IL2 4q27

DEXI 16p13.13

t(4;16)(q27;p13) IL2 4q27

TNFRSF17

16p13.13

t(3;16)(q27;p12) IL21R 16p12.1 FRA16E (rare, dist A) BCL6 3q27.3 FRA3C (common, apc)

t(5;9)(q33;q22) ITK 5q33.3

SYK 9q22.2

t(6;7)(p21;p15) JAZF1 7p15.2-p15.1

PHF1 6p21.32 FRA6H (common,

apc)

t(7;17)(p15;q11) JAZF1 7p15.2-p15.1

SUZ12

17q11.2

t(2;17)(p23;q25) KIAA1618 17q25.3

ALK 2p23.2-p23.1

t(4;10)(q12;p11) KIF5B 10p11.22

PDGF

RA 4q12

FRA4B (common,

BrdU)

t(10;14)(q11;q22) KTN1 14q22.3


apc)

t(12;16)(p13;p13) LAG3 12p13.31

MYH

11 16p13.11

FRA16A (rare, folic

acid)

t(1;7)(p35;q34) LCK 1p35.1

TRB

@ 7q34

t(3;13)(q27;q14) LCP1 13q14.12


133

t(5;8)(p13;q12) LIFR 5p13.1 FRA5A (common,

BrdU) PLAG

1 8q12.1

del(3)(q27q28)* LPP 3q28


t(7;19)(q34;p13) LYL1 19p13.13 FRA19B (rare, folic

acid) TRB@

7q34

t(11;19)(q13;q13.4) MALAT1 11q13.1 FRA11H (common,

apc) MHLB1


aza)

t(6;11)(p21.1;q13) MALAT1 11q13.1 FRA11H (common,

apc) TFEB 6p21.1

FRA6H (common,

apc)

t(3;18)(p21;q21) MALT1 18q21.32 FRA18B (common,

apc) MAP4 3p21.31

t(1;19)(q23;p13) MEF2D 1q22

DAZA

P1 19p13.3

FRA19B (rare, folic

acid)

t(10;11)(p12;q23) MLL 11q23.3

FRA11B (rare, folic


ABI1 10p12.1

t(11;17)(q23;q12) MLL 11q23.3

FRA11B (rare, folic


ACACA

17q12

t(4;11)(q21;q23) MLL 11q23.3

FRA11B (rare, folic


AFF1 4q21.3-

q22.1 FRA4F (common, apc)

t(2;11)(q11;q23) MLL 11q23.3

FRA11B (rare, folic


AFF3 2q11.2 FRA2A (rare, folic

acid)

t(5;11)(q31;q23) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

AFF4 5q31.1 FRA5C (common, apc)


acid), FRA11G

(common, apc)

ARHGAP2

6

5q31.3

del(11)(q23q23)* MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

ARHGEF1

2

11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)


acid), FRA11G

(common, apc)

ARHGEF1

7

11q13.4 FRA11H (common,

apc)

t(11;15)(q23;q15) MLL 11q23.3

FRA11B (rare, folic


CASC5

15q15.1

del(11)(q23q23)* MLL 11q23.3

FRA11B (rare, folic


CBL 11q23.3

FRA11B (rare, folic


t(11;12)(q23;q13) MLL 11q23.3

FRA11B (rare, folic


CIP29 12q13.2

t(11;16)(q23;p13.3) MLL 11q23.3

FRA11B (rare, folic


CREB

BP 16p13.3


acid), FRA11G

(common, apc)

DAB2

IP 9q33.2


acid), FRA11G

(common, apc)

EEFS

EC 3q21.3

t(11;19)(q23;p13) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

ELL 19p13.11 FRA19B (rare, folic

acid)


acid), FRA11G

(common, apc)

EP300 22q13.2 FRA22A (rare, folic

acid)

t(1;11)(p32;q23) MLL 11q23.3

FRA11B (rare, folic


EPS15 1p32.3 FRA1B (common, apc)

t(6;11)(q21;q23) MLL 11q23.3

FRA11B (rare, folic


FOXO3A

6q21 FRA6F (common, apc)

134

t(4;11)(p12;q23) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

FRYL 4p12


acid), FRA11G

(common, apc)

GAS7 17p13.1


acid), FRA11G

(common, apc)

GMPS 3q25.31 FRA3D (common,

apc)


acid), FRA11G

(common, apc)

GPHN 14q23.3 FRA14B (common,

apc)

t(11;17)(q23;q12) MLL 11q23.3

FRA11B (rare, folic


LASP1

17q12

t(3;11)(q28;q23) MLL 11q23.3

FRA11B (rare, folic


LPP 3q28

inv(11)(q21q23) MLL 11q23.3

FRA11B (rare, folic


MAM

L2 11q21

t(11;20)(q23;q11) MLL 11q23.3

FRA11B (rare, folic


MAP

RE1 20q11.21

t(11;19)(q23;p13.3) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

MLLT

1 19p13.3

FRA19B (rare, folic

acid)


acid), FRA11G

(common, apc)

MLLT

10 10p12.31


acid), FRA11G

(common, apc)

MLLT11

1q21.2 FRA1F (common, apc)


acid), FRA11G

(common, apc)

MLLT3

9p21.3 FRA9A (rare, folic

acid), FRA9C

(common, BrdU)

t(6;11)(q27;q23) MLL 11q23.3

FRA11B (rare, folic


MLLT4

6q27

t(11;17)(q23;q12) MLL 11q23.3

FRA11B (rare, folic


MLLT6

17q12

t(11;19)(q23;p13) MLL 11q23.3

FRA11B (rare, folic


MYO

1F 19p13.2

FRA19B (rare, folic

acid)

t(3;11)(p21;q23) MLL 11q23.3

FRA11B (rare, folic


NCKI

PSD 3p21.31

inv(11)(q14q23) MLL 11q23.3 FRA11B (rare, folic

acid), FRA11G

(common, apc)

PICA

LM 11q14.2

FRA11F (common,

apc)


acid), FRA11G

(common, apc)

RARA 17q21.2


acid), FRA11G

(common, apc)

SEPT11

4q21.1


acid), FRA11G

(common, apc)

SEPT2

2q37.3 FRA2J (common, apc)

t(11;22)(q23;q11) MLL 11q23.3

FRA11B (rare, folic


SEPT5

22q11.21

t(X;11)(q24;q23) MLL 11q23.3

FRA11B (rare, folic


SEPT6

Xq24

t(11;17)(q23;q25) MLL 11q23.3

FRA11B (rare, folic


SEPT

9

17q25.2-

q25.3

135


acid), FRA11G

(common, apc)

SH3G

L1 19p13.3

FRA19B (rare, folic

acid)


acid), FRA11G

(common, apc)

SMAP

1 6q13

FRA6D (common,

BrdU)


acid), FRA11G

(common, apc)

SORBS2

4q35.1


acid), FRA11G

(common, apc)

TET1 10q21.3 FRA10C (common,

BrdU)

t(11;15)(q23;q15) MLL 11q23.3

FRA11B (rare, folic


ZFYVE19

15q15.1

t(12;22)(p13;q12) MN1 22q12.1

ETV6 12p13.2

t(7;12)(q36;p13) MNX1 7q36.3 FRA7I (common, apc) ETV6 12p13.2

t(7;17)(p15;q22) MSI2 17q22

HOX

A9 7p15.2

t(X;2)(q11;p23) MSN Xq11.1

ALK 2p23.2-

p23.1

t(9;9)(p21;p21) MTS2 9p21.3 FRA9A (rare, folic

acid), FRA9C

(common, BrdU)

MTS1 9p21.3 FRA9A (rare, folic

acid), FRA9C

(common, BrdU)

t(6;7)(q23;q36) MYB 6q23.3

MNX1

7q36.3 FRA7I (common, apc)

t(8;9)(q24;p13) MYC 8q24.21

ZBTB

5 9p13.2

t(8;9)(q24;p13) MYC 8q24.21

ZCCH

C7 9p13.2

t(2;22)(p23;q12) MYH9 22q12.3

ALK 2p23.2-p23.1

t(8;17)(p12;q11) MYO18A 17q11.2

FGFR

1 8p12

t(2;8)(p23;p11.2) MYST3 8p11.21

ASXL2

2p23.3

t(8;16)(p11;p13) MYST3 8p11.21

CREBBP

16p13.3

t(8;22)(p11;q13) MYST3 8p11.21

EP300 22q13.2 FRA22A (rare, folic

acid)

inv(8)(p11q13) MYST3 8p11.21

NCOA2

8q13.3

t(8;20)(p11;q13) MYST3 8p11.21

NCO

A3 20q13.12

t(10;16)(q22;p13) MYST4 10q22.2

CREB

BP 16p13.3

inv(10)(q11q11) NCOA4 10q11.23 FRA10G (common,

apc) RET 10q11.21

FRA10G (common, apc)

t(5;16)(q33;p13) NDE1 16p13.11 FRA16A (rare, folic

acid)

PDGF

RB 5q33.1

del(10)(q24q24)* NFKB2 10q24.32

INA 10q24.33

t(7;10)(q34;q24) NFKB2 10q24.32

TBXAS1

7q34

t(5;14)(q33;q22) NIN 14q22.1

PDGF

RB 5q33.1

t(12;19)(p13.3; p13.3)

NOL1 12p13.31

TCF3 19p13.3 FRA19B (rare, folic

acid)

inv(X)(p11q13) NONO Xq13.1

TFE3 Xp11.23

t(7;9)(q34;q34) NOTCH1 9q34.3

TRB

@ 7q34

t(2;5)(p23;q35) NPM1 5q35.1 FRA5G (rare, folic

acid) ALK

2p23.2-p23.1

t(3;5)(q25;q35) NPM1 5q35.1 FRA5G (rare, folic

acid) MLF1 3q25.32

FRA3D (common,

apc)

t(5;17)(q35;q21) NPM1 5q35.1 FRA5G (rare, folic

acid) RARA 17q21.2

136

t(3;5)(p24;q35) NSD1 5q35.2-q35.3 FRA5G (rare, folic

acid) ANKRD28

3p24.3

t(11;17)(q13;q21) NUMA1 11q13.4 FRA11H (common,

apc) RARA 17q21.2

t(9;9)(q34;q34) NUP214 9q34.13

ABL1 9q34.12

t(6;9)(p22;q34) NUP214 9q34.13

DEK 6p22.3

t(10;11)(q25;p15) NUP98 11p15.4

ADD3 10q25.1-

q25.2

FRA10B (rare, BrdU), FRA10E (common

apc)

t(6;11)(q24;p15) NUP98 11p15.4

CCDC

28A 6q24.1

inv(11)(p15q22) NUP98 11p15.4

DDX10

11q22.3

t(10;11)(q23;p15) NUP98 11p15.4

HHEX 10q23.33 FRA10A (rare, folic

acid)

t(7;11)(p15;p15) NUP98 11p15.4

HOXA11

7p15.2

t(7;11)(p15;p15) NUP98 11p15.4

HOXA13

7p15.2

t(7;11)(p15;p15) NUP98 11p15.4

HOX

A9 7p15.2

t(11;12)(p15;q13) NUP98 11p15.4

HOXC11

12q13.13 FRA12A (rare, folic

acid)

t(11;12)(p15;q13) NUP98 11p15.4

HOX

C13 12q13.13

FRA12A (rare, folic

acid)

t(2;11)(q31;p15) NUP98 11p15.4

HOX

D11 2q31.1

FRA2G (common,

apc)

t(2;11)(q31;p15) NUP98 11p15.4

HOXD13

2q31.1 FRA2G (common,

apc)

t(3;11)(q29q13;p15

)del(3)(q29). NUP98 11p15.4

IQCG 3q29

t(5;11)(q35;p15) NUP98 11p15.4

NSD1 5q35.2-q35.3

FRA5G (rare, folic acid)

t(11;17)(p15;p13) NUP98 11p15.4

PHF23

17p13.1

t(1;11)(q24;p15) NUP98 11p15.4

PRRX

1 1q24.2

t(9;11)(q34;p15) NUP98 11p15.4

PRRX2

9q34.11

t(9;11)(p22;p15) NUP98 11p15.4

PSIP1 9p22.3

t(4;11)(q23;p15) NUP98 11p15.4

RAP1

GDS1 4q23

t(11;18)(p15;q12) NUP98 11p15.4

SETBP1

18q12.3

t(11;20)(p15;q12) NUP98 11p15.4

TOP1 20q12

t(3;11)(p24;p15) NUP98 11p15.4

TOP2B


apc)

t(8;11)(p12;p15) NUP98 11p15.4

WHS

C1L1 8p12

t(15;19)(q14;p13) NUT 15q14

BRD4 19q13.12 FRA19B (rare, folic

acid)

t(8;11)(p12;q14) ODZ4 11q14.1

NRG1 8p12

t(9;17)(q22;p13) OMD 9q22.31

USP6 17p13.2

t(2;13)(q36;q14) PAX3 2q36.1

FOXO1A

13q14.11

t(X;2)(q13;q36) PAX3 2q36.1

MLLT

7 Xq13.1

t(2;2)(p23;q36) PAX3 2q36.1

NCO

A1 2p23.3

t(7;9)(q11;p13) PAX5 9p13.2

ELN 7q11.23 FRA7J (common, apc)

t(9;12)(p13;p13) PAX5 9p13.2

ETV6 12p13.2

t(3;9)(p14;p13) PAX5 9p13.2

FOXP1

3p14.1

t(9;15)(p13;q24) PAX5 9p13.2

PML 15q24.1

137

t(9;18)(p13;q11) PAX5 9p13.2

ZNF521

18q11.2

t(1;13)(p36;q14) PAX7 1p36.13 FRA1A (common,

apc)

FOXO

1A 13q14.11

t(2;3)(q13;p25) PAX8 2q13 FRA2B (rare, folic

acid) PPAR

G 3p25.2-p25.1

t(8;9)(p22;p24) PCM1 8p22

JAK2 9p24.1

t(8;10)(p22;q11) PCM1 8p22


apc)

t(1;5)(q21;q33) PDE4DIP 1q21.1 FRA1F (common, apc) PDGF

RB 5q33.1

t(10;11)(p12;q14) PICALM 11q14.2 FRA11F (common,

apc)

MLLT

10 10p12.31

t(3;6)(q27;p21) PIM1 6p21.2 FRA6H (common,


t(15;17)(q24;q21) PML 15q24.1

RARA 17q21.2

t(3;11)(q27;q23.1) POU2AF1 11q23.1


t(8;22)(p21;q12) PPP2R2A 8p21.2

CHEK2

22q12.1 FRA22B (common,

apc)

t(X;1)(p11;q23) PRCC 1q23.1

TFE3 Xp11.23

t(17;17)(q21;q24) PRKAR1A 17q24.2

RARA 17q21.2

t(10;17)(q11;q24) PRKAR1A 17q24.2


apc)

t(4;5)(q21;q33) PRKG2 4q21.21

PDGF

RB 5q33.1

t(14;19)(q11;q13) PVRL2 19q13.32 FRA19A (common, 5-

aza) TRA@

14q11.2

t(5;17)(q33;p13) RABEP1 17p13.2

PDGF

RB 5q33.1

t(5;14)(q35;q11) RANBP17 5q35.1 FRA5G (rare, folic

acid) TRD@

14q11.2

t(2;2)(p23;q13) RANBP2 2q13 FRA2B (rare, folic

acid) ALK

2p23.2-p23.1

t(1;22)(p13;q13) RBM15 1p13.3

MKL1 22q13.1-

q13.2

FRA22A (rare, folic

acid)

t(3;5)(p21;q33) RBM6 3p21.31

CSF1R

5q33.1

t(3;4)(q27;p14) RHOH 4p14


inv(3)(q21q26), t(3;3)(q21;q26)

RPN1 3q21.3

EVI1 3q26.2

t(1;3)(p36;q21) RPN1 3q21.3

PRDM16


apc)

t(2;21)(q11;q22) RUNX1 21q22.12

AFF3 2q11.2 FRA2A (rare, folic

acid)

t(16;21)(q24;q22) RUNX1 21q22.12

CBFA2T3

16q24.3

t(12;21)(q12;q22) RUNX1 21q22.12

CPNE

8 12q12

t(3;21)(q26;q22) RUNX1 21q22.12

EVI1 3q26.2

t(11;21)(q13;q22) RUNX1 21q22.12

MACROD1

11q13.1 FRA11H (common,

apc)

t(3;21)(q26;q22) RUNX1 21q22.12

MDS1 3q26.2

t(1;21)(p36;q22) RUNX1 21q22.12

PRDM16


apc)

t(X;21)(p22;q22) RUNX1 21q22.12

PRDX

4 Xp22.11

t(3;21)(q26;q22) RUNX1 21q22.12

RPL2

2P1 3q26.2

t(8;21)(q21;q22) RUNX1 21q22.12

RUNX1T1

8q21.3

t(4;21)(q31;q22) RUNX1 21q22.12

SH3D

19 4q31.3

t(8;21)(q23;q22) RUNX1 21q22.12

TRPS1

8q23.3

138

t(7;21)(p22;q22) RUNX1 21q22.12

USP42

7p22.1 FRA7B (common, apc)

t(1;21)(p35;q22) RUNX1 21q22.12

YTHD

F2 1p35.3

t(1;21)(q21;q22) RUNX1 21q22.12

ZNF687

1q21.2 FRA1F (common, apc)

t(2;4)(p23;q21) SEC31A 4q21.22

ALK 2p23.2-

p23.1

del(6)(q14q22)* SENP6 6q14.1

TCBA

1 6q22.31

t(9;9)(q34;q34), del(9)(q34q34)*

SET 9q34.11

NUP214

9q34.13

t(1;9)(p34;q34) SFPQ 1p34.3

ABL1 9q34.12

t(X;1)(p11;p34) SFPQ 1p34.3

TFE3 Xp11.23

t(3;6)(q27;p21) SFRS3 6p21.31 FRA6H (common,


t(4;6)(p15;q22) SLC34A2 4p15.2 FRA4D (common,

apc) ROS1 6q22.2

t(1;7)(q32;p21) SLC45A3 1q32.1

ETV1 7p21.2

t(1;3)(q32;q27) SLC45A3 1q32.1

ETV5 3q27.2 FRA3C (common, apc)

t(5;17)(q33;p11.2) SPECC1 17p11.2

PDGFRB

5q33.1

t(2;5)(p16;q33) SPTBN1 2p16.2 FRA2D (common,

apc)

PDGF

RB 5q33.1

t(X;18)(p11;q11) SS18 18q11.2

SSX1 Xp11.23

t(X;18)(p11;q11) SS18 18q11.2

SSX2 Xp11.22

t(X;18)(p11;q11) SS18 18q11.2

SSX4 Xp11.23

t(X;20)(p11;q13) SS18L1 20q13.33

SSX1 Xp11.23

t(17;17)(q21;q21) STAT5B 17q21.2

RARA 17q21.2

del(1)(p33p33)* STIL 1p33

TAL1 1p33

t(2;4)(p22;q12) STRN 2p22.2

PDGF

RA 4q12

FRA4B (common,

BrdU)

t(9;17)(q31;q12) TAF15 17q12

NR4A

3 9q31.1

t(12;17)(p13;q12) TAF15 17q12

ZNF384

12p13.31

t(1;3)(p32-34;p21) TAL1 1p33

RHO

A 3p21.31

t(1;7)(p33;q34) TAL1 1p33

TRB@

7q34

t(1;14)(p33;q11) TAL1 1p33

TRD@

14q11.2

t(3;6)(q26;q25) TBL1XR1 3q26.32

RGS1

7 6q25.2

t(8;8)(q11;q12) TCEA1 8q11.23

PLAG1

8q12.1

t(9;15)(q31;q21) TCF12 15q21.3

NR4A

3 9q31.1

t(17;19)(q22;p13) TCF3 19p13.3 FRA19B (rare, folic

acid) HLF 17q22

t(1;19)(q23;p13) TCF3 19p13.3 FRA19B (rare, folic

acid) PBX1 1q23.3

t(19;19)(p13;q13) TCF3 19p13.3 FRA19B (rare, folic

acid) TFPT 19q13.42

FRA19A (common, 5-

aza)

t(12;19)(p13;p13) TCF3 19p13.3 FRA19B (rare, folic

acid) ZNF3

84 12p13.31

t(1;3)(p33;p21) TCTA 3p21.31

TAL1 1p33

t(2;3)(p23;q12) TFG 3q12.2

ALK 2p23.2-

p23.1

t(3;9)(q12;q31) TFG 3q12.2

NR4A3

9q31.1

t(1;3)(q23;q12) TFG 3q12.2

NTRK

1 1q23.1

139

t(3;3)(q29;q27) TFRC 3q29


t(2;3)(p21;q26) THADA 2p21

MDS1 3q26.2

t(1;17)(p34;p13) THRAP3 1p34.3

USP6 17p13.2

t(7;10)(q34;q24) TLX1 10q24.31

TRB

@ 7q34

t(10;14)(q24;q11) TLX1 10q24.31

TRD@

14q11.2

t(21;21)(q22;q22), del(21)(q22q22)*

TMPRSS2 21q22.3

ERG 21q22.2

t(7;21)(p21;q22) TMPRSS2 21q22.3

ETV1 7p21.2

t(17;21)(q21;q22) TMPRSS2 21q22.3

ETV4 17q21.31

t(3;21)(q27;q22) TMPRSS2 21q22.3

ETV5 3q27.2 FRA3C (common, apc)

t(5;15)(q33;q15) TP53BP1 15q15.3

PDGFRB

5q33.1

t(1;2)(q21;p23) TPM3 1q21.3 FRA1F (common, apc) ALK 2p23.2-

p23.1

inv(1)(q21q23) TPM3 1q21.3 FRA1F (common, apc) NTRK

1 1q23.1

t(1;5)(q21;q33) TPM3 1q21.3 FRA1F (common, apc) PDGF

RB 5q33.1

inv(1)(q21q31) TPM3 1q21.3 FRA1F (common, apc) TPR 1q31.1 FRA1K (common,

apc)

t(2;19)(p23;p13) TPM4 19p13.12 FRA19B (rare, folic

acid) ALK

2p23.2-p23.1

inv(1)(q23q31) TPR 1q31.1 FRA1K (common,

apc) NTRK

1 1q23.1

t(9;14)(p21;q11) TRA@ 14q11.2

CDKN2A

9p21.3

FRA9A (rare, folic

acid), FRA9C (common, BrdU)

t(X;14)(q28;q11) TRA@ 14q11.2

MTCP

1 Xq28

FRAXE (rare, folic

acid), FRAXF (rare, folic acid)

t(8;14)(q24;q11) TRA@ 14q11.2

MYC 8q24.21

t(14;21)(q11;q22) TRA@ 14q11.2

OLIG

2 21q22.11

t(14;14)(q11;q32) TRA@ 14q11.2

TCL1A

14q32.13

t(7;14)(q34;q11) TRA@ 14q11.2

TRB

@ 7q34

t(7;14)(p14;q11) TRA@ 14q11.2

TRG@

7p14.1

t(7;12)(q34;p13) TRB@ 7q34

CCND2

12p13.32

inv(7)(p15q34),

t(7;7)(p15;q34) TRB@ 7q34

HOX9 7p15.2

inv(7)(p15q34) TRB@ 7q34

HOXA10

7p15.2

inv(7)(p15q34) TRB@ 7q34

HOX

A11 7p15.2

t(7;11)(q34;p15) TRB@ 7q34

LMO1 11p15.4

t(7;11)(q34;p13) TRB@ 7q34

LMO2 11p13 FRA11E (common,

apc)

t(X;7)(q28;q34) TRB@ 7q34

MTCP1

Xq28

FRAXE (rare, folic

acid), FRAXF (rare, folic acid)

t(6;7)(q23;q34) TRB@ 7q34

MYB 6q23.3

t(7;9)(q34;q31) TRB@ 7q34

TAL2 9q31.2

inv(7)(p14q34) TRB@ 7q34

TRG

@ 7p14.1

t(11;14)(p15;q11) TRD@ 14q11.2

LMO1 11p15.4

t(11;14)(p13;q11) TRD@ 14q11.2

LMO2 11p13 FRA11E (common,

apc)

t(5;14)(q35;q11) TRD@ 14q11.2

NKX2

E 5q35.2

FRA5G (rare, folic

acid)

140

t(8;14)(q24;q11) TRD@ 14q11.2

PVT1 8q24.21

t(5;14)(q35;q11) TRD@ 14q11.2

TLX3 5q35.1 FRA5G (rare, folic

acid)

t(7;14)(p14;q32) TRG@ 7p14.1

IGH@ 14q32.33

t(7;8)(q34;p12) TRIM24 7q34

FGFR

1 8p12

t(7;10)(q34;q11) TRIM24 7q34


apc)

t(1;10)(p13;q11) TRIM33 1p13.2


apc)

t(5;14)(q33;q32) TRIP11 14q32.12

PDGF

RB 5q33.1

t(12;13)(p13;q14) TTL-T 13q14.11

ETV6 12p13.2

t(11;17)(q23;q21) ZBTB16 11q23.2

RARA 17q21.2

t(9;10)(q34;q22.3) ZMIZ1 10q22.3

ABL1 9q34.12

t(8;13)(p12;q12) ZMYM2 13q12.11

FGFR1

8p12

t(3;7)(q27;p12) ZNFN1A1 7p12.2


*Deletions creating a fusion between two genes

141

Table 2: Gene pairs involved in cancer-specific recurrent translocations in which the

breakpoint in one gene co-localizes with a fragile site Translocation Gene Fragile Site Gene Fragile Site Cancer

a

t(7;12)(p22;q13) ACTB FRA7B GLI1

Vascular and perivascular tumor

inv(7)(q21q34) AKAP9 FRA7E BRAF

Adenocarcinoma (Thyroid)

t(5;14)(q35;q32) BCL11B

NKX2E

FRA5G Acute lymphoblastic leukemia/lymphoblastic lymphoma

t(5;14)(q35;q32) BCL11B

TLX3 FRA5G Acute lymphoblastic leukemia/lymphoblastic lymphoma

t(14;18)(q32;q21) BCL2 FRA18B IGH@

Chronic lymphocytic leukemia, B-cell

lymphoma

t(18;22)(q21;q11) BCL2 FRA18B IGL@

Chronic lymphocytic leukemia, Mature B-cell neoplasm

t(8;19)(q24;q13) BCL3 FRA19A MYC

B-prolymphocytic leukemia

t(3;16)(q27;p13) BCL6 FRA3C CIITA

Diffuse large B-cell lymphoma

t(3;8)(q27;q24) BCL6 FRA3C MYC


t(1;14)(q21;q32) BCL9 FRA1F IGH@


t(1;22)(q21;q11) BCL9 FRA1F IGL@

Follicular lymphoma

t(4;22)(q12;q11) BCR

PDGF

RA FRA4B Atypical chronic myeloid leukemia

t(11;18)(q22;q21) BIRC3

MALT1

FRA18B B-cell lymphoma

t(8;12)(q24;q21-22) BTG1 FRA12B MYC

Chronic lymphocytic leukemia

t(5;10)(q33;q21) CCDC6 FRA10C PDGF

RB Chronic myeloid leukemia

t(5;11)(q12;q23) CENPK

MLL FRA11B, FRA11G


t(4;12)(q12;p13) CHIC2 FRA4B ETV6

Aggressive NK-cell leukemia, Acute

myeloid leukemia

t(4;19)(q35;q13) CIC FRA19A DUX4

Soft tissue tumor

t(2;17)(p23;q23) CLTC FRA17B ALK

Inflammatory myofibroblastic tumor, B-

cell lymphoma

t(X;17)(p11;q23) CLTC FRA17B TFE3

Adenocarcinoma (Kidney)

t(17;22)(q21;q13) COL1A1

PDGFB

FRA22A Dermatofibrosarcoma protuberans/Bednar tumor

t(X;6)(q22;q13-14) COL4A5

COL12

A1 FRA6D Bone- and cartilage-producing tumor

t(1;2)(p13;q37) COL6A3 FRA2J CSF1

Tenosynovial giant cell tumor

t(11;19)(q21;p13) CRTC1 FRA19B MAML

2

Benign epithelial tumor, special type,

Mucoepidermoid carcinoma

t(3;9)(q27;p24) DMRT1

BCL6 FRA3C Diffuse large B-cell lymphoma

t(1;1)(p36;q41) DUSP10

PRDM16

FRA1A Acute myeloid leukemia

t(10;12)(q11;p13) ERC1

RET FRA10G Papillary Thyroid Carcinoma

t(5;12)(q31;p13) ETV6

ACSL6 FRA5C Atypical chronic myeloid leukemia, Acute myeloid leukemia

t(1;12)(q21;p13) ETV6

ARNT FRA1F Acute myeloid leukemia

t(10;12)(q24;p13) ETV6

GOT1 FRA10A Myelodysplastic syndrome

t(1;12)(p36;p13) ETV6

MDS2 FRA1A Acute myeloid leukemia

t(4;12)(q12;p13) ETV6

PDGF

RA FRA4B

Chronic eosinophilic

leukemia/hypereosinophilic syndrome

t(12;22)(q13;q12) EWSR1 FRA22B DDIT3

Myxoid liposarcoma

t(21;22)(q22;q12) EWSR1 FRA22B ERG

Ewing tumor/peripheral primitive

neuroectodermal tumor

t(7;22)(p21;q12) EWSR1 FRA22B ETV1

Ewing tumor/peripheral primitive neuroectodermal tumor

t(17;22)(q21;q12) EWSR1 FRA22B ETV4


t(2;22)(q35;q12) EWSR1 FRA22B FEV

Ewing tumor/peripheral primitive

neuroectodermal tumor

142

t(11;22)(q24;q12) EWSR1 FRA22B FLI1


t(9;22)(q31;q12) EWSR1 FRA22B NR4A3

Chondrosarcoma, myxoid (Soft tissue)

t(12;22)(p13;q12) EWSR1 FRA22B ZNF38

4 Acute undifferentiated leukemia, Acute lymphoblastic leukemia

t(4;17)(q12;q21) FIP1L1 FRA4B RARA

Juvenile myelomonocytic leukemia

t(X;11)(q13;q23) FOXO4

MLL FRA11B, FRA11G

Acute lymphoblastic leukemia, Acute myeloid leukemia

t(12;16)(q13;p11) FUS

ATF1 FRA12A Angiomatoid malignant fibrous histiocytoma (Soft tissue)

t(3;12)(q27;p13) GAPDH


t(5;12)(q33;q24) GIT2 FRA12E PDGF

RB Chronic myeloid proliferative disorders

t(10;14)(q11;q32) GOLGA5


del(8)(q12q24)* HAS2 FRA8C, FRA8E

PLAG1

Lipoblastoma (Soft tissue)

t(8;19)(p12;q13) HERV-K (LOC113

386)

FRA19A FGFR1

Chronic myeloproliferative disorder

t(5;7)(q33;q11) HIP1 FRA7J PDGF

RB Chronic myelomonocytic leukemia

t(3;6)(q27;p22) HIST1H4

I BCL6 FRA3C Non-Hodgkin's lymphoma

t(2;12)(q37;q14) HMGA2

CXCR7

FRA2J Lipoma (Soft tissue)

t(12;14)(q14;q24) HMGA2

RAD51L1

FRA14C Leiomyoma (Uterus, corpus)

t(8;10)(p11;q11) HOOK3


t(6;16)(p21;q22) HP

MRPS10

FRA6H Prostate carcinoma

t(3;14)(q27;q32) HSP90A

A1 BCL6 FRA3C Diffuse large B-cell lymphoma

t(1;14)(p22;q32) IGH@

BCL10 FRA1D Extranodal marginal zone B-cell

lymphoma

t(14;19)(q32;q13) IGH@

BCL3 FRA19A B-cell lymphoma

t(3;14)(q27;q32) IGH@

BCL6 FRA3C Follicular lymphoma, B-cell lymphoma

t(11;14)(q13;q32) IGH@

CCND1

FRA11A, FRA11H

Chronic lymphocytic leukemia

t(6;14)(p21;q32) IGH@

CCND

3 FRA6H B-cell lymphoma

t(7;14)(q21;q32) IGH@

CDK6 FRA7E Chronic lymphocytic leukemia

t(14;19)(q32;q13) IGH@

CEBPA

FRA19A Acute lymphoblastic leukemia

t(14;19)(q32;q13) IGH@

CEBP

G FRA19A Acute lymphoblastic leukemia

t(11;14)(q23;q32) IGH@

DDX6 FRA11B, FRA11G


t(7;14)(q21;q32) IGH@

ERVW

E1 FRA7E Chronic lymphocytic leukemia

t(5;14)(q31;q32) IGH@

IL3 FRA5C Acute lymphoblastic leukemia

t(14;18)(q32;q21) IGH@

MALT1

FRA18B B-cell lymphoma

t(11;14)(q23;q32) IGH@

PAFA

H1B2

FRA11B,

FRA11G Non-Hodgkin's lymphoma

t(11;14)(q23;q32) IGH@

PCSK7 FRA11B, FRA11G

Mature B-cell neoplasm

t(14;19)(q32;q13) IGH@

SPIB FRA19A Diffuse large B-cell lymphoma

t(14;16)(q32;q23) IGH@

WWO

X FRA16D Multiple myeloma

t(2;12)(p11;p13) IGK@ FRA2L CCND

2 Mantle cell lymphoma

t(2;8)(p11;q24) IGK@ FRA2L MYC

B-cell lymphoma

t(2;8)(p11;q24) IGK@ FRA2L PVT1

Burkitt lymphoma/leukemia

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t(2;6)(p11;q25) IGK@ FRA2L ZC3H1

2D Diffuse large B-cell lymphoma

t(19;22)(q13;q11) IGL@

BCL3 FRA19A Follicular lymphoma, Diffuse large B-cell

lymphoma

t(3;22)(q27;q11) IGL@

BCL6 FRA3C B-cell lymphoma

t(11;22)(q13;q11) IGL@

CCND

1

FRA11A,

FRA11H

Mature B-cell neoplasm, Mantle cell

lymphoma

t(6;22)(p21;q11) IGL@

CCND

3 FRA6H Multiple myeloma

t(7;22)(q21;q11) IGL@

CDK6 FRA7E Chronic lymphocytic leukemia

t(16;22)(q23;q11) IGL@

WWO

X FRA16D Multiple myeloma

t(6;7)(p21;p15) JAZF1

PHF1 FRA6H Endometrial stromal sarcoma (Uterus, corpus)

t(4;10)(q12;p11) KIF5B

PDGF

RA FRA4B Hypereosinophilia

t(10;14)(q11;q22) KTN1


t(12;16)(p13;p13) LAG3

MYH11


t(3;13)(q27;q14) LCP1

BCL6 FRA3C Follicular lymphoma, Diffuse large B-cell

lymphoma

t(5;8)(p13;q12) LIFR FRA5A PLAG1

Adenoma (Salivary gland)

del(3)(q27q28)* LPP


t(7;19)(q34;p13) LYL1 FRA19B TRB@


t(3;18)(p21;q21) MALT1 FRA18B MAP4


t(1;19)(q23;p13) MEF2D

DAZA

P1 FRA19B Acute lymphoblastic leukemia

t(10;11)(p12;q23) MLL FRA11B, FRA11G

ABI1


t(11;17)(q23;q12) MLL FRA11B,

FRA11G

ACAC

A Acute myeloid leukemia

t(5;11)(q31;q23) MLL FRA11B,

FRA11G

ARHG

AP26 Acute myeloid leukemia

t(11;15)(q23;q15) MLL FRA11B, FRA11G

CASC5


t(11;12)(q23;q13) MLL FRA11B,

FRA11G CIP29


t(11;16)(q23;p13.3) MLL FRA11B, FRA11G

CREBBP


t(9;11)(q33;q23) MLL FRA11B, FRA11G

DAB2IP


t(3;11)(q21;q23) MLL FRA11B,

FRA11G

EEFSE

C Acute lymphoblastic leukemia

t(4;11)(p12;q23) MLL FRA11B, FRA11G

FRYL


t(11;17)(q23;p13) MLL FRA11B,

FRA11G GAS7

Acute lymphoblastic leukemia, Acute

myeloid leukemia

t(11;17)(q23;q12) MLL FRA11B,

FRA11G LASP1



LPP


inv(11)(q21q23) MLL FRA11B,

FRA11G

MAML

2 Acute myeloid leukemia

t(11;20)(q23;q11) MLL FRA11B, FRA11G

MAPRE1


t(10;11)(p12;q23) MLL FRA11B, FRA11G

MLLT10


t(6;11)(q27;q23) MLL FRA11B,

FRA11G

MLLT


t(11;17)(q23;q12) MLL FRA11B, FRA11G

MLLT6


t(3;11)(p21;q23) MLL FRA11B,

FRA11G

NCKIP

SD Acute myeloid leukemia

t(11;17)(q23;q21) MLL FRA11B,

FRA11G RARA


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SEPT11

Chronic neutrophilic leukemia

t(11;22)(q23;q11) MLL FRA11B,

FRA11G SEPT5


t(X;11)(q24;q23) MLL FRA11B, FRA11G

SEPT6


t(11;17)(q23;q25) MLL FRA11B,

FRA11G SEPT9


t(4;11)(q35;q23) MLL FRA11B,

FRA11G

SORB

S2 Acute myeloid leukemia

t(11;15)(q23;q15) MLL FRA11B, FRA11G

ZFYVE19


t(7;12)(q36;p13) MNX1 FRA7I ETV6


t(6;7)(q23;q36) MYB

MNX1 FRA7I Acute myeloid leukemia

t(8;22)(p11;q13) MYST3

EP300 FRA22A Acute myeloid leukemia

t(5;16)(q33;p13) NDE1 FRA16A PDGF

RB Chronic myelomonocytic leukemia

t(12;19)(p13.3; p13.3)

NOL1

TCF3 FRA19B Acute leukemia

t(2;5)(p23;q35) NPM1 FRA5G ALK

B-cell lymphoma

t(5;17)(q35;q21) NPM1 FRA5G RARA


t(3;5)(p24;q35) NSD1 FRA5G ANKR

D28 Adult myeloid leukemia

t(11;17)(q13;q21) NUMA1 FRA11H RARA


t(10;11)(q25;p15) NUP98

ADD3 FRA10B, FRA10E


t(10;11)(q23;p15) NUP98

HHEX FRA10A Acute myeloid leukemia

t(11;12)(p15;q13) NUP98

HOXC11

FRA12A Acute myeloblastic leukemia with maturation

t(11;12)(p15;q13) NUP98

HOXC

13 FRA12A Myeloblastic leukemia with maturation

t(2;11)(q31;p15) NUP98

HOXD

11 FRA2G Acute myeloid leukemia

t(2;11)(q31;p15) NUP98

HOXD13

FRA2G Acute myeloid leukemia, Chronic myeloid leukemia

t(5;11)(q35;p15) NUP98

NSD1 FRA5G Acute myeloid leukemia

t(3;11)(p24;p15) NUP98

TOP2B FRA3A Acute myeloid leukemia

t(15;19)(q14;p13) NUT

BRD4 FRA19B Squamous cell carcinoma

t(7;9)(q11;p13) PAX5

ELN FRA7J Acute lymphoblastic leukemia

t(1;13)(p36;q14) PAX7 FRA1A FOXO

1A Alveolar rhabdomyosarcoma (Soft tissue)

t(2;3)(q13;p25) PAX8 FRA2B PPAR

G

Adenoma (Thyroid), Adenocarcinoma

(Thyroid)

t(8;10)(p22;q11) PCM1


t(1;5)(q21;q33) PDE4DIP FRA1F PDGF

RB

Myeloproliferative disorder associated

with eosinophilia

t(10;11)(p12;q14) PICALM FRA11F MLLT


t(3;11)(q27;q23.1) POU2AF

1 BCL6 FRA3C

Acute lymphoblastic leukemia/lymphoblastic lymphoma

t(10;17)(q11;q24) PRKAR1

A RET FRA10G Papillary thyroid carcinoma

t(14;19)(q11;q13) PVRL2 FRA19A TRA@

Peripheral T-cell lymphoma, Angioimmunoblastic T-cell lymphoma

t(5;14)(q35;q11) RANBP1

7 FRA5G TRD@

Acute lymphoblastic

leukemia/lymphoblastic lymphoma

t(2;2)(p23;q13) RANBP2 FRA2B ALK

Inflammatory myofibroblastic tumor

t(1;22)(p13;q13) RBM15

MKL1 FRA22A Acute myeloid leukemia

t(3;4)(q27;p14) RHOH

BCL6 FRA3C Follicular lymphoma, Mature B-cell

neoplasm

t(1;3)(p36;q21) RPN1

PRDM16

FRA1A Acute myeloid leukemia, Chronic myelomonocytic leukemia

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t(2;21)(q11;q22) RUNX1

AFF3 FRA2A Childhood T-cell acute lymphoblastic leukemia

t(11;21)(q13;q22) RUNX1

MACR

OD1 FRA11H Acute myeloid leukemia

t(1;21)(p36;q22) RUNX1

PRDM16


t(7;21)(p22;q22) RUNX1

USP42 FRA7B Acute myeloid leukemia

t(1;21)(q21;q22) RUNX1

ZNF68

7 FRA1F Acute myeloid leukemia

t(4;6)(p15;q22) SLC34A2 FRA4D ROS1

Carcinoma, NOS (Lung)

t(1;3)(q32;q27) SLC45A3

ETV5 FRA3C Adenocarcinoma (Prostate)

t(2;5)(p16;q33) SPTBN1 FRA2D PDGF

RB Chronic myeloproliferative disorder

t(2;4)(p22;q12) STRN

PDGF

RA FRA4B

Chronic eosinophilic

leukemia/hypereosinophilic syndrome

t(17;19)(q22;p13) TCF3 FRA19B HLF


t(1;19)(q23;p13) TCF3 FRA19B PBX1


t(12;19)(p13;p13) TCF3 FRA19B ZNF38

4 Acute lymphoblastic leukemia

t(3;3)(q29;q27) TFRC


t(3;21)(q27;q22) TMPRSS

2 ETV5 FRA3C Adenocarcinoma (Prostate)

t(1;2)(q21;p23) TPM3 FRA1F ALK

Anaplastic large cell lymphoma,

Inflammatory myofibroblastic tumor

inv(1)(q21q23) TPM3 FRA1F NTRK

1 Papillary thyroid carcinoma

t(1;5)(q21;q33) TPM3 FRA1F PDGF

RB Chronic eosinophilic leukemia

t(2;19)(p23;p13) TPM4 FRA19B ALK

Inflammatory myofibroblastic tumor, Anaplastic large cell lymphoma

inv(1)(q23q31) TPR FRA1K NTRK

1 Papillary thyroid carcinoma

t(9;14)(p21;q11) TRA@

CDKN

2A FRA9A, FRA9C Acute lymphoblastic leukemia

t(X;14)(q28;q11) TRA@

MTCP1

FRAXE, FRAXF T-prolymphocytic leukemia, Nonneoplastic lymphatic disorder/lesion

t(7;11)(q34;p13) TRB@

LMO2 FRA11E Acute lymphoblastic leukemia

t(X;7)(q28;q34) TRB@

MTCP

1 FRAXE, FRAXF T-prolymphocytic leukemia

t(11;14)(p13;q11) TRD@

LMO2 FRA11E Acute lymphoblastic leukemia

t(5;14)(q35;q11) TRD@

NKX2

E FRA5G

Acute lymphoblastic

leukemia/lymphoblastic lymphoma

t(5;14)(q35;q11) TRD@

TLX3 FRA5G Acute lymphoblastic leukemia

t(7;10)(q34;q11) TRIM24

RET FRA10G Papillary thyroid carcinoma

t(1;10)(p13;q11) TRIM33

RET FRA10G Papillary thyroid Carcinoma

t(3;7)(q27;p12) ZNFN1A

1 BCL6 FRA3C Diffuse large B-cell lymphoma

aFor a complete description, see the Mitelman database of Chromosome Aberrations in Cancer [130].

146

SCHOLASTIC VITA

NAME: Allison Burrow Weckerle

BUSINESS ADDRESS: Wake Forest University School of Medicine

Department of Biochemistry and Molecular Biology

1 Medical Center Blvd.

Winston-Salem, NC 27157

BUSINESS TELEPHONE: (336) 716-5843

E-MAIL ADDRESS: [email protected]

EDUCATION: 2006 B.S. Exercise Science High Point University, High Point, NC

2011 Ph.D.

Biochemistry and Molecular Biology

Graduate School of Arts and Sciences

Wake Forest University, Winston-Salem, NC Thesis Title: The Molecular Basis of Fragile Site

FRA16B and its Role in inv(16)(p13q22) in

Acute Myeloid Leukemia

RESEARCH EXPERIENCE:

Thesis research:

- Wake Forest University School of Medicine, Biochemistry and Molecular Biology Department Under the supervision of Dr. Yuh-Hwa Wang, May 2007 – present

Using a variety of molecular and cellular biology techniques to study replication of fragile site

FRA16B and its potential role in the generation of cancer-specific rearrangements, including

inv(16)(p13q22).

Undergraduate research:

- TransTech Pharma, High Point, NC

Under the supervision of Dr. Robert Andrews, June 2005 – August 2005

Contributed to the organic synthesis of a Glucagon-like peptide-1 (GLP-1) receptor agonist for

the treatment of Type II diabetes mellitus.

PUBLICATIONS:

1. A. Burrow, L. Williams, L. Pierce, and Y.H. Wang. “Over half of breakpoints in gene pairs involved in cancer-specific recurrent translocations are mapped to human

chromosomal fragile sites,” BMC Genomics 2009, 10:59.

147

PUBLICATIONS (continued):

2. A. Burrow, A. Marullo, L. Holder, and Y.H. Wang. “Secondary structure formation and DNA instability at fragile site FRA16B,” Nucleic Acids Research 2010, 38:9.

3. L. Dillon*, A. Burrow*, and Y.H. Wang. “DNA instability at chromosomal fragile

sites in cancer,” Current Genomics 2010, 11:5. *These two authors contributed equally

to the work.

4. A.B. Weckerle, M. Santra, M.C.Y. Ng, P.P. Koty, and Y.H. Wang. “CBFB and MYH11 in inv(16)(p13q22) of Acute Myeloid Leukemia Display Close Spatial

Proximity in Interphase Nuclei of Human Hematopoietic Stem Cells,” Genes,

Chromosomes and Cancer 2011, in press.

ABSTRACTS/POSTER PRESENTATIONS:

- Genetics and Environmental Mutagenesis Society Annual Fall Meeting, RTP, NC, October

2008. Presented a poster: “Over half of breakpoints in gene pairs involved in cancer-specific

recurrent translocations are mapped to human chromosomal fragile sites .”

- Wake Forest Graduate Student Research Day, Winston-Salem, NC, April 2009. Presented a

poster: “Over half of breakpoints in gene pairs involved in cancer-specific recurrent

translocations are mapped to human chromosomal fragile sites.”

- American Society of Human Genetics Annual Meeting, Honolulu, HI, October 2009. Presented a poster: “Formation of Secondary Structure Contributes to DNA Instability at Fragile Site

FRA16B.”

- Wake Forest Graduate Student Research Day, Winston-Salem, NC, March 2010. Presented a

poster: “Formation of Secondary Structure Contributes to DNA Instability at Fragile Site

FRA16B.”

AWARDS:

- Alumni Student Travel Award, 2009-2010, given by the Graduate School of Arts and Sciences to support the expenses of graduate students who present their research at national and

international professional meetings, Wake Forest University.

- Sandy Lee Cowgill Memorial Scholarship, 2009-2010, given by the Graduate School of Arts

and Sciences to a Ph.D. student with academic excellence in the biochemistry and molecular biology department, Wake Forest University

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