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
iii
husband, Steve, for giving me so much to look forward to every single day, and who
never ceases to make me smile.
iv
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
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].
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.
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.
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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.
MATERIALS AND METHODS
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
87
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.
MATERIALS AND METHODS
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
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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
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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)
t(2;22)(q33;q12) EWSR1 22q12.2 FRA22B (common,
apc)
CREB
1 2q33.3 FRA2I (common,apc)
t(12;22)(q13;q12) EWSR1 22q12.2 FRA22B (common,
apc) DDIT
3 12q13.3
t(21;22)(q22;q12) EWSR1 22q12.2 FRA22B (common,
apc) ERG 21q22.2
t(7;22)(p21;q12) EWSR1 22q12.2 FRA22B (common,
apc) ETV1 7p21.2
t(17;22)(q21;q12) EWSR1 22q12.2 FRA22B (common,
apc) ETV4 17q21.31
t(2;22)(q35;q12) EWSR1 22q12.2 FRA22B (common,
apc) FEV 2q35
t(11;22)(q24;q12) EWSR1 22q12.2 FRA22B (common,
apc) FLI1 11q24.3
t(9;22)(q31;q12) EWSR1 22q12.2 FRA22B (common,
apc) NR4A
3 9q31.1
inv(22)(q12q12) EWSR1 22q12.2 FRA22B (common,
apc)
PATZ
1 22q12.2
FRA22B (common,
apc)
t(6;22)(p21;q12) EWSR1 22q12.2 FRA22B (common,
apc) POU5
F1 6p21.33
FRA6H (common, apc)
t(2;22)(q31;q12) EWSR1 22q12.2 FRA22B (common,
apc) SP3 2q31.1
FRA2G (common, apc)
t(11;22)(p13;q12) EWSR1 22q12.2 FRA22B (common,
apc) WT1 11p13
FRA11E (common,
apc)
t(12;22)(p13;q12) EWSR1 22q12.2 FRA22B (common,
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
BCL6 3q27.3 FRA3C (common, apc)
t(5;12)(q33;q24) GIT2 12q24.11 FRA12E (common,
apc) PDGF
RB 5q33.1
t(10;14)(q11;q32) GOLGA5 14q32.12
RET 10q11.21 FRA10G (common,
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
BCL6 3q27.3 FRA3C (common, apc)
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
RET 10q11.21 FRA10G (common,
apc)
t(6;16)(p21;q22) HP 16q22.3
MRPS10
6p21.1 FRA6H (common,
apc)
t(3;14)(q27;q32) HSP90AA1 14q32.31
BCL6 3q27.3 FRA3C (common, apc)
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
BCL6 3q27.3 FRA3C (common, apc)
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
19q13.11 FRA19A (common, 5-
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
acid), FRA11G (common, apc)
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)
t(2;3)(p11;q27) IGK@ 2p11.2 FRA2L (rare, folic
acid) BCL6 3q27.3 FRA3C (common, apc)
t(2;11)(p11;q13) IGK@ 2p11.2 FRA2L (rare, folic
acid) CCND
1 11q13.2
FRA11A (rare, folic
acid), FRA11H (common, apc)
t(2;12)(p11;p13) IGK@ 2p11.2 FRA2L (rare, folic
acid)
CCND
2 12p13.32
t(2;7)(p11;q21) IGK@ 2p11.2 FRA2L (rare, folic
acid) CDK6 7q21.2 FRA7E (common, apc)
t(2;18)(p11;q21) IGK@ 2p11.2 FRA2L (rare, folic
acid) FVT1 18q21.33
FRA18B (common,
apc)
t(2;8)(p11;q24) IGK@ 2p11.2 FRA2L (rare, folic
acid) MYC 8q24.21
t(2;8)(p11;q24) IGK@ 2p11.2 FRA2L (rare, folic
acid) PVT1 8q24.21
t(2;6)(p11;q25) IGK@ 2p11.2 FRA2L (rare, folic
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
acid), FRA11H (common, apc)
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
RET 10q11.21 FRA10G (common,
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
BCL6 3q27.3 FRA3C (common, apc)
133
t(5;8)(p13;q12) LIFR 5p13.1 FRA5A (common,
BrdU) PLAG
1 8q12.1
del(3)(q27q28)* LPP 3q28
BCL6 3q27.3 FRA3C (common, apc)
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
19q13.4 FRA19A (common, 5-
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
acid), FRA11G (common, apc)
ABI1 10p12.1
t(11;17)(q23;q12) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
ACACA
17q12
t(4;11)(q21;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
AFF1 4q21.3-
q22.1 FRA4F (common, apc)
t(2;11)(q11;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
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)
t(5;11)(q31;q23) MLL 11q23.3 FRA11B (rare, folic
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)
t(11;11)(q13;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
ARHGEF1
7
11q13.4 FRA11H (common,
apc)
t(11;15)(q23;q15) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
CASC5
15q15.1
del(11)(q23q23)* MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
CBL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
t(11;12)(q23;q13) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
CIP29 12q13.2
t(11;16)(q23;p13.3) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
CREB
BP 16p13.3
t(9;11)(q33;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
DAB2
IP 9q33.2
t(3;11)(q21;q23) MLL 11q23.3 FRA11B (rare, folic
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)
t(11;22)(q23;q13) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
EP300 22q13.2 FRA22A (rare, folic
acid)
t(1;11)(p32;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
EPS15 1p32.3 FRA1B (common, apc)
t(6;11)(q21;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
FOXO3A
6q21 FRA6F (common, apc)
134
t(4;11)(p12;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
FRYL 4p12
t(11;17)(q23;p13) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
GAS7 17p13.1
t(3;11)(q25;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
GMPS 3q25.31 FRA3D (common,
apc)
t(11;14)(q23;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
GPHN 14q23.3 FRA14B (common,
apc)
t(11;17)(q23;q12) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
LASP1
17q12
t(3;11)(q28;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
LPP 3q28
inv(11)(q21q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
MAM
L2 11q21
t(11;20)(q23;q11) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
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)
t(10;11)(p12;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
MLLT
10 10p12.31
t(1;11)(q21;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
MLLT11
1q21.2 FRA1F (common, apc)
t(9;11)(p21;q23) MLL 11q23.3 FRA11B (rare, folic
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
acid), FRA11G (common, apc)
MLLT4
6q27
t(11;17)(q23;q12) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
MLLT6
17q12
t(11;19)(q23;p13) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
MYO
1F 19p13.2
FRA19B (rare, folic
acid)
t(3;11)(p21;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
NCKI
PSD 3p21.31
inv(11)(q14q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
PICA
LM 11q14.2
FRA11F (common,
apc)
t(11;17)(q23;q21) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
RARA 17q21.2
t(4;11)(q21;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
SEPT11
4q21.1
t(2;11)(q37;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
SEPT2
2q37.3 FRA2J (common, apc)
t(11;22)(q23;q11) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
SEPT5
22q11.21
t(X;11)(q24;q23) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
SEPT6
Xq24
t(11;17)(q23;q25) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
SEPT
9
17q25.2-
q25.3
135
t(11;19)(q23;p13) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
SH3G
L1 19p13.3
FRA19B (rare, folic
acid)
t(6;11)(q13;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
SMAP
1 6q13
FRA6D (common,
BrdU)
t(4;11)(q35;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
SORBS2
4q35.1
t(10;11)(q21;q23) MLL 11q23.3 FRA11B (rare, folic
acid), FRA11G
(common, apc)
TET1 10q21.3 FRA10C (common,
BrdU)
t(11;15)(q23;q15) MLL 11q23.3
FRA11B (rare, folic
acid), FRA11G (common, apc)
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
3p24.2 FRA3A (common,
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
RET 10q11.21 FRA10G (common,
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,
apc) BCL6 3q27.3 FRA3C (common, apc)
t(15;17)(q24;q21) PML 15q24.1
RARA 17q21.2
t(3;11)(q27;q23.1) POU2AF1 11q23.1
BCL6 3q27.3 FRA3C (common, apc)
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
RET 10q11.21 FRA10G (common,
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
BCL6 3q27.3 FRA3C (common, apc)
inv(3)(q21q26), t(3;3)(q21;q26)
RPN1 3q21.3
EVI1 3q26.2
t(1;3)(p36;q21) RPN1 3q21.3
PRDM16
1p36.32 FRA1A (common,
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
1p36.32 FRA1A (common,
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,
apc) BCL6 3q27.3 FRA3C (common, apc)
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
BCL6 3q27.3 FRA3C (common, apc)
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
RET 10q11.21 FRA10G (common,
apc)
t(1;10)(p13;q11) TRIM33 1p13.2
RET 10q11.21 FRA10G (common,
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
BCL6 3q27.3 FRA3C (common, apc)
*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
Diffuse large B-cell lymphoma
t(1;14)(q21;q32) BCL9 FRA1F IGH@
Acute lymphoblastic leukemia
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
Acute myeloid leukemia
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
Ewing tumor/peripheral primitive neuroectodermal tumor
t(2;22)(q35;q12) EWSR1 FRA22B FEV
Ewing tumor/peripheral primitive
neuroectodermal tumor
142
t(11;22)(q24;q12) EWSR1 FRA22B FLI1
Ewing tumor/peripheral primitive neuroectodermal tumor
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
BCL6 FRA3C Diffuse large B-cell lymphoma
t(5;12)(q33;q24) GIT2 FRA12E PDGF
RB Chronic myeloid proliferative disorders
t(10;14)(q11;q32) GOLGA5
RET FRA10G Papillary Thyroid Carcinoma
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
RET FRA10G Papillary Thyroid Carcinoma
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
Acute lymphoblastic leukemia
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
143
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
RET FRA10G Papillary Thyroid Carcinoma
t(12;16)(p13;p13) LAG3
MYH11
FRA16A Acute myeloid leukemia
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
BCL6 FRA3C Diffuse large B-cell lymphoma
t(7;19)(q34;p13) LYL1 FRA19B TRB@
Acute lymphoblastic leukemia
t(3;18)(p21;q21) MALT1 FRA18B MAP4
Diffuse large B-cell lymphoma
t(1;19)(q23;p13) MEF2D
DAZA
P1 FRA19B Acute lymphoblastic leukemia
t(10;11)(p12;q23) MLL FRA11B, FRA11G
ABI1
Acute myeloid leukemia
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
Acute myeloid leukemia
t(11;12)(q23;q13) MLL FRA11B,
FRA11G CIP29
Acute myeloid leukemia
t(11;16)(q23;p13.3) MLL FRA11B, FRA11G
CREBBP
Acute myeloid leukemia
t(9;11)(q33;q23) MLL FRA11B, FRA11G
DAB2IP
Acute myeloid leukemia
t(3;11)(q21;q23) MLL FRA11B,
FRA11G
EEFSE
C Acute lymphoblastic leukemia
t(4;11)(p12;q23) MLL FRA11B, FRA11G
FRYL
Acute lymphoblastic leukemia, Acute myeloid leukemia
t(11;17)(q23;p13) MLL FRA11B,
FRA11G GAS7
Acute lymphoblastic leukemia, Acute
myeloid leukemia
t(11;17)(q23;q12) MLL FRA11B,
FRA11G LASP1
Acute myeloid leukemia
t(3;11)(q28;q23) MLL FRA11B, FRA11G
LPP
Acute myeloid leukemia
inv(11)(q21q23) MLL FRA11B,
FRA11G
MAML
2 Acute myeloid leukemia
t(11;20)(q23;q11) MLL FRA11B, FRA11G
MAPRE1
Acute lymphoblastic leukemia
t(10;11)(p12;q23) MLL FRA11B, FRA11G
MLLT10
Acute myeloid leukemia
t(6;11)(q27;q23) MLL FRA11B,
FRA11G
MLLT
4 Acute myeloid leukemia
t(11;17)(q23;q12) MLL FRA11B, FRA11G
MLLT6
Acute myeloid leukemia
t(3;11)(p21;q23) MLL FRA11B,
FRA11G
NCKIP
SD Acute myeloid leukemia
t(11;17)(q23;q21) MLL FRA11B,
FRA11G RARA
Acute myeloid leukemia
144
t(4;11)(q21;q23) MLL FRA11B, FRA11G
SEPT11
Chronic neutrophilic leukemia
t(11;22)(q23;q11) MLL FRA11B,
FRA11G SEPT5
Acute myeloid leukemia
t(X;11)(q24;q23) MLL FRA11B, FRA11G
SEPT6
Acute myeloid leukemia
t(11;17)(q23;q25) MLL FRA11B,
FRA11G SEPT9
Acute myeloid leukemia
t(4;11)(q35;q23) MLL FRA11B,
FRA11G
SORB
S2 Acute myeloid leukemia
t(11;15)(q23;q15) MLL FRA11B, FRA11G
ZFYVE19
Acute myeloid leukemia
t(7;12)(q36;p13) MNX1 FRA7I ETV6
Acute myeloid leukemia
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
Acute myeloid leukemia
t(3;5)(p24;q35) NSD1 FRA5G ANKR
D28 Adult myeloid leukemia
t(11;17)(q13;q21) NUMA1 FRA11H RARA
Acute myeloid leukemia
t(10;11)(q25;p15) NUP98
ADD3 FRA10B, FRA10E
Acute lymphoblastic leukemia, Acute myeloid leukemia
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
RET FRA10G Papillary Thyroid Carcinoma
t(1;5)(q21;q33) PDE4DIP FRA1F PDGF
RB
Myeloproliferative disorder associated
with eosinophilia
t(10;11)(p12;q14) PICALM FRA11F MLLT
10 Acute myeloid leukemia
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
145
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
FRA1A Acute myeloid leukemia
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
Acute lymphoblastic leukemia
t(1;19)(q23;p13) TCF3 FRA19B PBX1
Acute lymphoblastic leukemia, Acute myeloid leukemia
t(12;19)(p13;p13) TCF3 FRA19B ZNF38
4 Acute lymphoblastic leukemia
t(3;3)(q29;q27) TFRC
BCL6 FRA3C Diffuse large B-cell lymphoma
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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