होम The American Journal of Human Genetics Whole-Genome Sequencing Reveals Diverse Models of Structural Variations in Esophageal Squamous Cell...

Whole-Genome Sequencing Reveals Diverse Models of Structural Variations in Esophageal Squamous Cell Carcinoma

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डाउनलोड की गई फ़ाइलों की गुणवत्ता क्या है?
The American Journal of Human Genetics
February, 2016
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आप पुस्तक समीक्षा लिख सकते हैं और अपना अनुभव साझा कर सकते हैं. पढ़ूी हुई पुस्तकों के बारे में आपकी राय जानने में अन्य पाठकों को दिलचस्पी होगी. भले ही आपको किताब पसंद हो या न हो, अगर आप इसके बारे में ईमानदारी से और विस्तार से बताएँगे, तो लोग अपने लिए नई रुचिकर पुस्तकें खोज पाएँगे.
Whole-Genome Sequencing Reveals
Diverse Models of Structural Variations
in Esophageal Squamous Cell Carcinoma
Caixia Cheng,1,2,3,15 Yong Zhou,4,15 Hongyi Li,1,2,15 Teng Xiong,4,15 Shuaicheng Li,5,15 Yanghui Bi,1,2,15
Pengzhou Kong,1,2 Fang Wang,1,2 Heyang Cui,1,2,4 Yaoping Li,1,6 Xiaodong Fang,4 Ting Yan,1,2
Yike Li,1,2,7 Juan Wang,1,2 Bin Yang,1,2,6 Ling Zhang,1,2 Zhiwu Jia,1,2 Bin Song,1,2,8 Xiaoling Hu,1,2
Jie Yang,1,2 Haile Qiu,8 Gehong Zhang,8 Jing Liu,1,7 Enwei Xu,9 Ruyi Shi,1,2 Yanyan Zhang,1,7
Haiyan Liu,1,2 Chanting He,1,2 Zhenxiang Zhao,1,2 Yu Qian,1,2 Ruizhou Rong,1,2,7 Zhiwei Han,1,2,7
Yanlin Zhang,5 Wen Luo,4 Jiaqian Wang,4 Shaoliang Peng,10 Xukui Yang,4 Xiangchun Li,4 Lin Li,4
Hu Fang,4 Xingmin Liu,4 Li Ma,6 Yunqing Chen,6 Shiping Guo,6 Xing Chen,11 Yanfeng Xi,9
Guodong Li,9 Jianfang Liang,3 Xiaofeng Yang,12 Jiansheng Guo,7 JunMei Jia,8 Qingshan Li,13
Xiaolong Cheng,1,2 Qimin Zhan,14,* and Yongping Cui1,2,*
Comprehensive identification of somatic structural variations (SVs) and understanding their mutational mechanisms in cancer might
contribute to understanding biological differences and help to identify new therapeutic targets. Unfortunately, characterization of complex SVs across the whole genome and the mutational mechanisms underlying esophageal squamous cell carcinoma (ESCC) is largely
unclear. To define a comprehensive catalog of somatic SVs, affected target genes, and their underlying mechanisms in ESCC, we reanalyzed whole-genome sequencing (WGS) data from 31 ESCCs using Meerkat algorithm to predict somatic SVs and Patchwork to determine copy-number changes. We found deletions and translocations with NHEJ and alt-EJ signature as the dominant SV types, and 16%
of deletions were complex deletions. SVs frequently led to disruption of cancer-associated genes (e.g., CDKN2A and NOTCH1) with
different mutational mechanisms. Moreover, chromothripsis, kataegis, and breakage-fusion-bridge (BFB) were identified as contributing
to locally mis-arranged chromosomes that occurred in 55% of E; SCCs. These genomic catastrophes led to amplification of oncogene
through chromothripsis-derived double-minute chromosome formation (e.g., FGFR1 and LETM2) or BFB-affected chromosomes (e.g.,
CCND1, EGFR, ERBB2, MMPs, and MYC), with approximately 30% of ESCCs harboring BFB-derived CCND1 amplification. Furthermore,
analyses of copy-number alterations reveal high frequency of whole-genome duplication (WGD) and recurrent focal amplification of
CDCA7 that might act as a potential oncogene in ESCC. Our findings reveal molecular defects such as chromothripsis and BFB in malignant transformation of ESCCs and demonstrate diverse models of SVs-derived target genes in ESCCs. These genome-wide SV profiles
and their underlying mechanisms provide preventive, diagnostic, and therapeutic implications for ESCCs.

Cancer genomes harbor various somatic forms of genetic
alterations spanning from nucleotide-level alterations
(e.g., point mutations and small insertions/deletions) to
large chromosomal events (e.g., structural variations and
copy-number changes), some of which can contribute to
tumor development.1 Specially, genomic structural variation (SV) is a hallmark of cancer.1 The fraction of the
genome affected by SVs is comparatively larger than that
accounted for by SNPs, indicating significant consequences of SVs on phenotypic variation.2 The main types
of mechanisms known to cause SVs in human cancer

include homologous recombination, nonreplicative
nonhomologous repair, and replication-based mechanisms.3 Generally, homologous recombination can occur
by non-allelic homologous recombination (NAHR), and
deficiency in homologous recombination is implicated as
a major source of cancer genome instability.4 In addition,
SVs, especially aberrant ligation of double-strand DNA
breaks (DSBs), can arise, mostly due to exposure to external
DNA-damaging agents, through non-homologous endjoining (NHEJ) or alternative end joining (alt-EJ) mechanisms.5 For complex rearrangements, the mechanisms for
repairing DNA replication errors such as fork stalling and
template switching (FoSTeS) or microhomology-mediated

Translational Medicine Research Center, Shanxi Medical University, Taiyuan, Shanxi 030001, China; 2Key Laboratory of Cellular Physiology, Ministry of
Education, Shanxi Medical University, Taiyuan, Shanxi 030001, China; 3Department of Pathology, the First Hospital, Shanxi Medical University, Taiyuan,
Shanxi 030001, China; 4BGI-Shenzhen, Shenzhen, Guangdong 518083, China; 5Department of Computer Science, City University of Hong Kong, Hong
Kong 518057, China; 6Department of Tumor Surgery, Shanxi Cancer Hospital, Taiyuan, Shanxi 030001, China; 7Department of General Surgery, the First
Hospital, Shanxi Medical University, Taiyuan, Shanxi 030001, China; 8Department of Oncology, the First Hospital, Shanxi Medical University, Taiyuan,
Shanxi 030001, China; 9Department of Pathology, Shanxi Cancer Hospital, Taiyuan, Shanxi 030001, China; 10School of Computer Science & State Key
Laboratory of High Performance Computing National University of Defense Technology, Changsha, Hunan 410073, China; 11Department of Endoscopy,
Shanxi Provincial People’s Hospital, Taiyuan, Shanxi 030001, China; 12Department of Urology, the First Hospital, Shanxi Medical University, Taiyuan,
Shanxi 030001, China; 13School of Pharmaceutical Sciences, Shanxi Medical University, Taiyuan, Shanxi 030001, China; 14State Key Laboratory of Molecular Oncology, Cancer Institute and Cancer Hospital, Chinese Academy of Medical Sciences and Peking Union Medical College, Beijing 100021, China
These authors contributed equally to this work
*Correspondence: zhanqimin@pumc.edu.cn (Q.Z.), cuiy0922@yahoo.com (Y.C.)
http://dx.doi.org/10.1016/j.ajhg.2015.12.013. Ó2016 The Authors
This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

256 The American Journal of Human Genetics 98, 256–274, February 4, 2016

break-induced repair (MMBIR) have been described.6
Recently, single catastrophic events causing genomic shattering followed by incorrect re-joining of the fragmented
DNA, termed chromothripsis, is receiving greater attention
as a major mechanism generating complex SVs in human
It is well known that SVs have implications in treatment
and prediction of individual’s outcome because genomescale rearrangements can play an unappreciated role in
cancer through their ability to move blocks of adjacent
genes simultaneously or form gene fusion, leading to concurrent oncogenic events.1 Comprehensive investigation
in many types of tumor shows that breakpoints directly
generate an oncogenic element that can be used as a therapeutic target, such as driver fusion transcript of EML4ALK in a subset of non-small-cell lung cancer (NSCLC)
that respond to the kinase inhibitor crizotinib and
FGFR3-TACC3 fusions in glioblastoma, bladder cancer,
lung squamous cell, and head and neck squamous cell
carcinoma (HNSCC) that can benefit from targeted FGFR
kinase inhibition.1,8,9 In addition, gene amplification, a selective copy-number increase of genomic segments
through DNA rearrangements, is a clinically important
form of genome instability in cancer, because gene amplification causes advanced tumors and acquired therapy
resistance.10,11 Thus, a better understanding of the underlying mechanisms of oncogenic events driven by SVs is
important for identification of molecular targets for diagnosis, prognosis, and treatment guidance.
Continuous DNA breaks and rearrangements through
chromothripsis, chromoplexy, or a breakage-fusion-bridge
(BFB) cycle have been implicated as underlying mechanisms for gene amplification or fusion in human
cancer.12 A BFB cycle, a series of chromosome breaks and
duplications that generate multiple copy-number states
and are assumed to derive from events occurring over
many rounds of cell division, has been shown to occur in
many malignant solid tumors, including HNSCC and
esophageal adenocarcinoma (EAC).13,14 In contrast to the
conventional clusters of complex rearrangements, chromothripsis, despite the large number of rearrangements,
exists in only two copy-number states with many transitions between these two states.15 For chromothripsis, the
affected chromosome (or regions from one or a few chromosome arms) is somehow fragmented and then stitched
together, most likely by NHEJ.15 The segments that are
not incorporated into the derivative chromosome are
either lost, yielding the low copy-number state, or incorporated into a double-minute (DM) chromosome.7,15 Chromothripsis has been reported in 2%–5% of diverse cancer
entities, with higher frequency in bone cancer (25%) and
medulloblastoma (36%).3,14 In parallel, another mutation
mechanism, kataegis, has been identified as distinguishing
mutational patterns that often co-occur with large-scale rearrangements.16 Unlike chromothripsis, which refers to an
oncogenic mechanism operating on a global level and
occurring in one or several chromosomes, kataegis has

been found to operate locally, generating large numbers
of mutations (or hotspots of hypermutations) in small regions of the genome.16 On the other hand, similar to chromothripsis, kataegis most likely causes a large number of
substitution mutations to occur in a region of the genome
at one time rather than accumulating in a step-wise
fashion.17 Kataegis is remarkably common, occurring, for
example, at a rate of 13/21 in breast cancer genomes.16
Massively parallel sequencing strategies offer the potential to carry out genome-wide screening for point
mutations, copy-number alterations (CNAs), and rearrangements on a single platform.18 We and others recently
reported genomic sequencing analyses of ESCCs, which
nominated cancer-associated genes driven by point mutations.19–22 However, at the level of genome structure,
somatic SVs and their underlying mechanisms are largely
unknown; the driving forces behind SVs have been less
well characterized than those for single-nucleotide alterations in ESCC. In this study, we re-analyzed whole-genome
sequencing (WGS) data of 31 ESCCs to characterize SVs and
their underlying mechanisms and to identify target genes
affected by SVs in ESCC. Our findings revealed different
mutational mechanisms for the formation of amplification
of cancer-associated genes in ESCC.

Material and Methods
Ethics Approval
This study was approved by the Ethics Committee of Shanxi Medical University (Approval No. 2009029) and the Ethics Committee
of Henan Cancer Hospital (Approval No. 2009xjs12). All samples
were obtained before treatment according to the guidelines of
the local ethics committees, and written informed consents were
received from all participants.

Data Processing
The WGS data of a total of 31 paired tumors and matched normal
tissues have been deposited at the European Genome-phenome
Archive (EGA).19,22 Raw data were filtered with SOAPnuke
(v.1.4.1) to remove sequencing adapters and low-quality reads.
High-quality reads were aligned to the NCBI human reference
genome (hg19) by BWA (v.0.5.9) with default parameters. Picard
(v.1.54) was used to mark duplicates and followed by Genome
Analysis Toolkit (v.1.0.6076, GATK IndelRealigner) to improve
alignment accuracy. The final BAM file stores all reads and calibrated qualities along with their alignments to the genome. For
interesting SVs with fewer numbers of supporting reads, we
further inspected IGV and checked the split read alignment (in
the .sr/ folder) to verify their accuracy.

Structural Variations Detection
Identification of somatic structural variations (SVs) from short
read data is challenging. Meerkat algorithm makes it possible to
predict both germline and somatic SVs directly from short read
data, focusing on complex events.23 Importantly, Yang et al. verified the accuracy of Meerkat by applying it to two HapMap genomes (NA18507 and NA12878) that were sequenced at high
coverage on the Illumina platform and for which complex

The American Journal of Human Genetics 98, 256–274, February 4, 2016 257

deletions have been previously reported.23,24 Also, 48 out of
randomly selected 49 (98%) events identified via Meerkat algorithm can be validated by PCR.23 Therefore, the Meerkat algorithm
can provide a more comprehensive spectrum of mechanisms of
SVs in a genome and is more reliable to detect SVs. In this study,
we applied Meerkat (0.185) algorithm with suggested parameters
to 31 ESCC genomes to predict somatic SVs and breakpoints as
described.23 In brief, we mapped reads against the human reference genome (hg19) to find soft-clipped and unmapped reads
(reads that mapped in an unexpected way) and re-mapped them
to identify discordant read pairs. Then, we extracted the split reads
(20 bp from both ends) to search for reads that cover the candidate
breakpoints and refined precise breakpoints by local alignments.
Mutational mechanisms were predicted based on homology and
sequencing features at the breakpoints. Somatic SVs were generated by filtering out germline events and other artifacts. We used
the following criteria to remove artifacts: (1) a large number (thousands or tens of thousands) of somatic SVs in one tumor sample;
(2) a dominant event type; (3) the SVs evenly distribute across
all chromosomes; (4) if the dominant events are intra-chromosome, they are very uniform in size (usually several hundreds bp
or at kb level). The samples that meet these criteria failed our quality-control steps and were discarded from further analysis. Only
high confidence calls were used in downstream analysis.

Locally Arranged Genome
To assess the randomness of SVs on chromosomes, we used a goodness-of-fit test against the expected distribution proposed by
Campbell et al. with a significant threshold < 0.0001.25 To assess
the significance of SV enrichment on chromosomes, we required
the number of locally arranged genomes to be more than 50 and
clustered chromosomes to have a high SVs mutation rate per Mb
exceeding three times the length of the interquartile range from
the 75th percentile of the chromosome counts for each tumor.26

Breakage-Fusion-Bridge Detection
We detected BFB events based on the evidence of fold-back inversions and telomere loss.27 Inversions meeting the following
criteria were defined as fold-back inversion. (1) Inversion is a single
inversion (invers_f or invers_r) detected by Meerkat, which means
there is no reciprocal partner of inversion. (2) Inversion must
demarcate a copy-number change that we make comparison of
reads depth between inverted-amplified and normal space region
(the region between breakpoints of fold-back inversion), and the
result with q < 0.0001 is defined as significance. (3) The two
ends of breakpoints of fold-back inversion must be separated
by <20 kb.

Chromothripsis Inference
To infer chromothripsis in ESCCs, we adapted criteria proposed by
Campbell et al.25 This analysis is based on ruling out the stepwise
rearrangements and required at least ten changes in segmental
copy number involving two or three distinct copy-number states
on a single chromosome. (1) We manually inspected copy-number
profiles for each case for regularity of oscillating copy-number
states. ESCC-16T, in which copy number oscillates between two
and three and has more than ten transitions, was selected for
inclusion. (2) We found statistical evidence (p < 0.001) for breakpoints clustering on chromosomes 3, 8, and 10 of ESCC-16T. (3)
In the case of ESCC-16T, due to loss of one haplotype (chromosome 8q) and chromothripsis occurring in amplified haplotype,

we could detect allelic imbalance change instead of loss and retention of heterozygosity. (4) For ESCC-16T, chromosome 8q had
three copy numbers of amplified haplotype, making it difficult
to entirely eliminate the possibility of rearrangements arising
from two haplotypes of the sample type. However, the minor
copy number always remains one, indicating high possibility of arrangements affecting a specific haplotype. (5) We found statistical
evidence of the randomness of fragment joins and segment order.
(6) For ESCC-16T, derivate chromosome 8 is difficult to infer the
ability to walk the derivative chromosome owing to the loss of
some rearrangements.

Copy-Number Alterations
Patchwork was used to determine copy-number alterations (CNAs)
across 31 ESCCs.28 First, it fixed windows of 200 bp in the human
reference genome, and each window was thought to be a marker.
Then, it estimated the log2 ratio between tumor and normal read
depth for each window. The log2 ratio of adjacent 50 windows
were merged to smooth the data. The merged windows (markers)
were further segmented by CBS. After the program combined the
allele frequency of germline single-nucleotide variants, absolute
copy number for each segments were given. Of 31 ESCC genomes,
19 clearly had clusters of normalized coverage between different
copies. For these 19 tumors, we estimated the ploidy, tumor content, and absolute copy number. To identify potential copy-number targets, we combined 31 of WGS data and 123 of comparative
genomic hybridization analysis (CGH) data and applied modified
GISTIC method to the combined data.19,22,29 The amplification or
deletion peaks with G-score > 0.1 that corresponds to p < 0.05 and
q < 0.05 were defined as significant.

We defined kataegis based on five stringent hallmarks described by
Nik-Zainal et al.:16 (1) presence of heavily mutated genomic
regions (‘‘macrocluster’’) consisting of a few hundred base pairs
(‘‘microcluster’’) separated by tens of unmutated kilobases; (2) mutation clusters generally colocalized with structural variation
breakpoints; (3) mutations that are all of the same type in a long
genomic region, and switched to different mutation classes in
other regions; (4) within the microcluster region, most mutations
being derived from the same parental chromosome; and (5) most
substitutions within the hypermutated region being characterized
by C>T transitions in TpCpX trinucleotides.

PCR-Sanger Sequencing Validation
For validation of TRAPPC9-CLVS1 or EIF3E-RAD51B fusion transcript, we performed RT-PCR and Sanger sequencing assays on
purified tumor and matched normal cells from ESCC-16T or
ESCC-19T, respectively. Total RNA (1 mg) from purified tumor and
matched normal cells was used for RT-PCR with the SuperScript
III First-Strand system (Invitrogen), according to the manufacturer’s instructions. The primers used were designed against exon
18 of TRAPPC9 (MIM: 611966) forward (50 -CGGAATTCACCCTG
GAAGCTGTCCTG-30 ) and exon 4 of CLVS1 (MIM: 611292) reverse
(50 -CCCTCGAGCTGCAACCCTTCAATGGC-30 ) or against exon 1
of EIF3E (MIM: 602210) forward (50 -CGGAATTCATGGCGGAG
TACG-30 ) and exon 5 of RAD51B (MIM: 602948) reverse
TRAPP9-CLVS1 (334 bp) or EIF3E-RAD51B (243 bp) was analyzed
by agarose gel electrophoresis. Amplified PCR products were gel
purified and then sequenced via the Sanger method.

258 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Fluorescence In Situ Hybridization Analysis
Frozen tumor and matched normal tissues of interesting ESCC
cases were cut a cryostat at 4 mm thickness, fixed in cold acetic
acid/methanol for 5 min at 4 C, and dried at room temperature.
Slides were stained with Cytocell enumeration probes against
interesting genes FGFR1 (MIM: 136350)/CEN8 (Z-2072, Zytovision, German), CCND1 (MIM: 168461)/CEN11 (Z-2071, Zytovision, German), TRAPPC9, CLVS1, EIF3E, and RAD51B, conjugated
with FITC or Cy3.5 (Rainbow Scientific). Staining was carried out
according to the manufacturer’s protocol. FISH samples were
viewed with a fully automated, upright Zeiss Axio-ImagerZ.1
microscope with a 203 objective and DAPI, FITC, and Rhodamine
filter cubes. Images were produced using the AxioCam MRm CCD
camera and Axiovision v.4.5 software suite. p values were calculated with a two-sample test for equality of proportions with continuity correction.

Real-Time Quantitative PCR
Real-time quantitative PCR (RT-PCR) was performed to quantify
the mRNA expression levels of CDCA7 (MIM: 609937), LETM2,
FGFR1, or WHSC1L1 (MIM: 607083) using ABI Stepone plus
with a SYBR Premix Ex Taq Kit (Takara Bio). GAPDH was used
as an endogenous control. Primers for GAPDH (F: 50 -CGG
were used. The detailed protocol was as follows: 95 C for
10 min, 40 cycles of 95 C for 15 s, and 60 C for 1 min, followed
by a melting-curve program from 59 C to 95 C with a heating
rate of 0.3 C every step and continuous-florescence acquisition.
All RT-PCR reactions were completed in triplicate. The relative
expression quantification of interesting genes was determined as
F ¼ 2DDCt.

CDCA7 or LETM2 protein levels in ESCCs were determined by
immunohistochemistry with CDCA7 antibody (HPA005565,
Sigma) or LETM2 antibody (17180-1-AP, Proteintech). Immunohistochemistry was performed as previously described.22 In brief,
sections were incubated with the specific antibody at a 1:40 dilution for 14 hr at 4 C, followed by detection using the PV8000
(Zhongshan) and DAB detection kit (Maixin), producing a dark
brown precipitate. Slides were counterstained with hematoxylin.
All images were captured at 3100. The cytoplasm H score and
the levels of CDCA7 or LETM2 shown by immunohistochemistry
were analyzed with Aperio Cytoplasma 2.0 software. Statistic analyses were performed with GraphPad Prism v.6.0 software package.
The significance of differences between ESCC and matched
normal tissue was determined by paired t test.

Stable CDCA7 Knockdown Clones in ECA109
Cell Line

cloned into pLVshRNA-puro vector (pLV-shRNA1 and pLVshRNA2). To perform plasmid infections, the ECA109 cells were
plated at 40%–50% confluence and incubated at 37 C overnight
(16 hr). pLVshRNA-puro vector, pLV-shRNA1, and pLV-shRNA2
were transfected into ECA109 cells using Lipofectamine 2000
reagent (Life Technologies) according to the manufacturer’s instructions. Forty-eight hours after transfection, culture medium
was replaced by fresh media containing 2 mg/ml puromycin and
subjected to screening stable monoclonies for 3 weeks. During
the selection, cells were maintained at culture medium containing
2 mg/ml puromycin. After 3 weeks of selection, approximately
20 monoclonies per dish were selected and transferred into
96-well plate. shRNA knockdown efficiency was determined by
RT-PCR and western blotting as described.22

Apoptosis Analysis by Flow Cytrometry
CDCA7 knockdown cells and cells transfected with pLVshRNApuro vector were labeled with Annexin-FITC/PI Staining Kit (Sangon Biotech) according to the manufacturer’s instruction and
analyzed by flow cytometry in BD FACScaliber (BD Bioscience).

RNA Sequencing and Data Analysis
Total RNA was extracted with the RNeasy Mini Kit (QIAGEN) and
complementary DNA (cDNA) libraries were synthesized with the
TruSeq RNA Sample Preparation Kit v.2 (Illumina). Libraries were
sequenced on an Illumina HiSEquation 4000 platform at BGI.
Filtering and quality controls were applied according to the
standard procedure. The gene expression profiles of CDCA7
knockdown cells versus control cells were compared via gene set
enrichment analysis. Differential expression levels (relative RNA
counts) between control cells and CDCA7 knockdown cells were
considered significantly different with a false discovery rate
(FDR) at a threshold of 1%.

Knockdown of LETM2, FGFR1, or WHSC1L1 in ESCC
Cell Lines
Three siRNAs targeting LETM2 and one negative control siRNA
(NC) (Guangzhou RiboBio) were used to knock down LETM2 in
ESCC cell lines (KYSE150 and ECA109). Meanwhile, FGFR1
(siRNA #1: 50 -AGTGGCTTATTAATTCCGATA-30 ; siRNA #2: 50 -GC
TAT-30 ; siRNA #3: 50 -GCTTCCATTACGATGCACAAA-30 ) were
knocked down in TE-1 and KYSE150 cells, respectively. To perform
infections, the ESCC cells were plated at 40%–50% confluence and
incubated at 37 C overnight (16 hr). Cells were transfected with
100 nM (final concentration) siRNA or NC siRNA using Lipofectamine 2000 (Life Technologies) according to manufacturer’s protocols. At 48 hr after transfection, cells were subjected to MTT
assay. At 72 hr after transfection, the knockdown efficiency was
determined by RT-PCR and western blotting as previously

MTT Assay

Vector pLVshRNA-puro was obtained from Addgene and used for
CDCA7 knockdown. Two independent shRNAs targeting CDCA7

5 3 103 cells were seeded in 48-well plates and incubated at normal
condition for 24, 48, 72, 96, and 120 hr. Cells were treated with 30 ml
of 5 mg/ml of MTT (Invitrogen) solution for 4 hr at 37 C until crystals were formed. MTT solution was removed from each well and
200 ml of DMSO was added to each well to dissolve the crystals. Color intensity was measured by Microplate Reader (Bio-Rad) at

The American Journal of Human Genetics 98, 256–274, February 4, 2016 259

490 nm. Each experiment consisted of five replications and at least
three independent experiments were carried out.

Colony Formation Assay
The assay was performed as described previously.22 In brief, cells
were seeded at 300–500 cells per well in 6-well plates containing
complete DMEM/F12 on day 0 and incubated at 37 C and 5%
CO2 for 10 days. On day 10, cells were fixed with 4% polyformaldehyde for 15 min and stained with 1% crystal violet before quantification. The experiments were triplicate and the numbers of colonies
containing more than 50 cells were microscopically counted.

Migration and Invasion Assays
Migration and invasion assays were performed in 16-well CIM
plates in an xCELLigence RTCA DP system (ACEA Biosciences) using matrigel basement membrane matrix (BD) for real-time cell
migration analysis as described previously.22 In brief, 30,000 cells
per well were seeded as 5 duplicates in serum-free medium at the upper compartment of the CIM plates coated with or without matrigel. Serum-complemented medium was added to the lower
compartment of the chamber, and then we started measurement
in xCELLigence RTCA DP system and analyzed the CI (cell index)
curves to determine cell invasion activity. For negative controls,
we added serum-free medium at both upper and bottom chambers.
The cell index representing the amount of migrated cells was calculated with the RTCA Software from ACEA Biosciences. At least three
independent experiments were carried out; for each independent
experiment, five duplicates were performed for each group.

Cells were lysed for 30 min in Triton buffer (1% Triton X-100,
50 mM Tris-HCl [pH 7.6], 150 mM NaCl, 1% sodium deoxycholate, 0.1% SDS) supplemented with protease and phosphatase
inhibitors (1 mM PMSF, 2 mM sodium pyrophosphate, 2 mM sodium betaglycerophosphate, 1 mM sodium fluoride, 1 mM sodium
orthovanadate, 10 mg/ml leupeptin, and 10 mg/ml aprotinin).
Lysates were cleared by centrifugation at 15,000 3 g at 4 C for
15 min, and protein concentrations were determined via the
Bradford method. 50 mg of protein were separated by SDS-polyacrylamide gel electrophoresis and transferred onto Immobilon-P
membranes. Proteins were detected by using anti-LETM2
(Proteintech, 17180-1-AP), anti-FGFR1 (Abcam cat# ab76464;
RRID: AB_1523613), anti-WHSC1L1 (Abcam, ab180500), antiCDCA7 (Abcam cat# ab69609; RRID: AB_1268064), anti-ERK1/2
(Santa Cruz, sc-514302), anti-p-ERK1/2 (Cell Signaling Technology cat# 4376; RRID: AB_331772), anti-AKT1 (Cell Signaling Technology cat# 2967; RRID: AB_331160), and anti-p-AKT1 (Cell
Signaling Technology cat# 9018). Antibody binding was detected
using horseradish peroxidase-labeled anti-mouse (Sigma) or antirabbit (Cell Signaling) antibodies and chemiluminescence was detected with a LAS4000 device (Fuji). Equal protein loading was
confirmed with antibodies against GAPDH (Transgen).

Spectrum and Distribution of Somatic SVs across 31
To characterize the mutational spectrum of somatic SVs in
ESCC, we applied Meerkat to WGS data of tumors and
paired normal tissues from 31 ESCC-affected individuals

(Table S1). A total of 5,204 SVs were identified from the
31 ESCC genomes with an average of 168 SVs per tumor,
ranging from 10 to 364 (Table S2). Five categories of SVs
were observed, including deletions, tandem duplications
(TDs), inversions, insertions, and intra- or inter-chromosomal translocations. Among these SVs, the average number of deletions per genome was 58 (ranging from 2 to 191)
and make up 35% of SV types. Additionally, about 42% of
SVs referred to intra- or inter-chromosomal translocations,
with an average of 71 per genome (ranging from 3 to 150).
For deletions and intra- or inter-chromosomal translocations, NHEJ and alt-EJ were the dominant mechanisms,
with alt-EJ being more abundant in most cases. Moreover,
291 deletions were identified as complex deletions generated by FoSTeS/MMBIR. We noticed that the number of
complex deletions were extremely diverse among individuals; some genomes contained a high portion of complex
deletions whereas others showed very few (Figure 1A, middle). Besides deletions and translocations, the number of
TDs for each genome was remarkably variable, with a range
of 5 to 104. We observed no homology at TDs within ESCC
genomes, further supporting the underlying mechanism
that requires no microhomology or existence of nonhomology-based mechanism to form TDs and complex deletions in tumor cells.24
Across 31 ESCC genomes, we found that 3,376 SVs
occurred in the region of genes and were predicted to
directly disrupt sequence of gene such as CDKN2A (MIM:
600160), NOTCH1 (MIM: 190198), NF1 (MIM: 613113),
and FANCD2 (MIM: 613984), and 492 genes contained a
breakpoint in two or more tumors. Specifically, 29 out of
31 ESCCs harbored CDKN2A deletion; of which, 13 ESCCs
had supporting SVs responsible for CDKN2A deletion and
2 out of these 13 genomes demonstrated complex deletions (ESCC-14T and ESCC-28T) (Figures 1B and S1).
Notably, all deletions from tumor genomes of these 13
ESCCs were homozygous deletion with both focal deletion
and arm-level loss. Furthermore, 8 out of these 13 ESCC
genomes had arm-level gain of 9p generated by wholegenome duplication (WGD) (Table S3), and no one had
two independent SVs within CDKN2A locus (9p21), suggesting that the focal deletion of CDKN2A happened
before WGD in these tumors (Figure 1C). In addition, we
also found that NOTCH1 was directly disrupted by TDs
in two ESCCs (Figure S2). These results suggested that
different mutational mechanisms can act on the same
driver (e.g., CDKN2A), and different drivers (e.g., CDKN2A
and NOTCH1) might be affected by different mutational
mechanisms in ESCC.
SVs tended to be either scattered genome-wide or
occurred locally with variable copy numbers across cancer
genomes and are more likely to occur in genomic region of
fragile sites.30–32 Across 31 ESCCs, the genomic distribution of SVs was characterized with three features:
randomly distributed across chromosomes; clustered in
one or more chromosomes; and clustered chromosomes
involving SVs accompanied with variable or limited copy

260 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Figure 1. Spectrum of Somatic SVs across ESCC Genomes and Mutational Mechanisms on CDKN2A in ESCC
(A) Frequencies of SVs (left), deletion (middle), or translocation (right) events and underlying mechanism across 31 ESCC tumors.
(B) Different mutational mechanisms of SVs act on CDKN2A. Representative maps show CDKN2A loss in six ESCCs. A red cluster typically suggests a tandem duplication; a blue cluster typically suggests a deletion; a purple cluster suggests a invers_reverse, and a green
cluster suggests a invers_forward.
(C) Model of focal deletion of CDKN2A that occurred before WGD on chromosome 9p in eight ESCC genomes.

numbers. Notably, we observed locally rearranged chromosomes were prevalent in ESCC genomes (17 out of 31
ESCCs) (Table S3). Although the mechanism underlying
most of locally rearranged chromosomes remains unknown, it appears that ESCC genomes harboring locally rearranged SVs accompanied with limited copy-number
states could be explained as chromothripsis or kataegis
(Figure S3). Meanwhile, 21 out of 31 ESCC genomes displayed at least two fold-back inversions in an autosome
accompanied with substantial copy-number states, and
some of them were likely to be a result of BFB (Table S3).
When analyzing SVs across ESCC genomes, we note
that, probably due to the tumor cell purity and ploidy,
many of the detected SVs have a smaller number of supporting split reads (Table S2). Additionally, due to a large
number of events that were relatively small, we did observe
that both breakpoints were in the same gene (Table S4). We
further compared the distribution of somatic SVs across a
variety of human cancers including breast cancer (BRCA),
glioblastoma multiforme (GBM), lung squamous cell carci-

noma (LUSC), ovarian serous cystadenocarcinoma (OV),
and gastric cancer (GC).23,33 Consistent with our observation in ESCC, those somatic SVs that had a smaller number
of supporting split reads and a high fraction of smaller SVs
were also observed in other human cancers (Figures S4A
and S4B). Advanced methodology needs to be designed
to solve these limitations.
Chromothripsis Leading to High-Level Amplification
of FGFR1 and LETM2
It is well known that chromosomes affected by chromothripsis show a characteristic pattern with more than ten
transitions oscillating between two and three copy number
states on chromosomal arms.7,15 We further accurately
infer the occurrence of chromothripsis by using
conceptual criteria proposed by Korbel and Campbell.15
Interestingly, we observed chromothripsis involving
chromosome 8 in ESCC-16T (Figure 2A). In addition to
general transition between two copy number states,
we found a high-level focal amplification (<500 kb,

The American Journal of Human Genetics 98, 256–274, February 4, 2016 261

Figure 2. High-Level Amplification of FGFR1 and LETM2 Affected by SVs
(A) The top panel represents different types of SVs indicated by lines with different colors on chromosome 8 in ESCC-16T; the middle
panel shows normalized coverage for each window. Zoom-in view of high-level amplification of FGFR1 and LETM2 locus is shown in the
bottom panel. FISH analysis demonstrates DM-derived amplification of FGFR1. Scale bar represents 10 mm.
(B) High-level amplification of FGFR1 in ESCC-06T. Top: FGFR1 locus and the high-level amplification region containing MYC gene are
shown. Bottom: Zoom-in view of high-level amplification of FGFR1 locus. FISH confirms CNAs by showing FGFR1 amplification as clustered multiple green signals. Scale bar represents 10 mm.
(C) FISH confirms CNAs by showing LETM2 amplification as scattered multiple green signals. Scale bar represents 10 mm.
(D) Immunohistochemical analysis shows LETM2 staining in ESCCs.
(E) LETM2 expression level in multiple ESCC lines determined by RT-PCR and western blotting.
(F) LETM2 knockdown prevents cell proliferation but has no effect on cell migration/invasion as monitored by MTT or in vitro cell migration and invasion assays in KYSE150 and ECA109 cells. Knockdown of LETM2 is demonstrated by immunoblotting; GAPDH was used as
loading control.
Data are mean 5 SD; each experiment was performed in triplicate. **p < 0.01.

262 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Figure 3. Fusion Genes Caused by Chromosomal Rearrangements in ESCC
(A) Details and schematic of the TRAPPC9-CLVS1 fusion transcript caused by complex deletion on chromosome 8 in ESCC-16T.
(B) Validation of the TRAPPC9-CLVS1 fusion transcript via PCR-sanger sequencing (left and middle) and FISH (right).
(legend continued on next page)

The American Journal of Human Genetics 98, 256–274, February 4, 2016 263

38,155,351–38,570,827 Mb) rearranged by chromothripsis
on chromosome 8p that corresponds to FGFR1 and LETM2
in this tumor. Importantly, no breakpoints were observed
within this amplified region, suggesting a strong positive
selection of FGFR1 and LETM2 amplifications during
ESCC progression/evolution. It was previously shown
that a potential by-product of chromothripsis is formation
of double-minute chromosomes (DMs) that might harbor
oncogenes and have been found in a variety of solid tumors.7,15 In ESCC-16T, our FISH experiment exhibited
multiple scattered FGFR1 signals and two copies of chromosome 8, suggesting that FGFR1 amplification might be
due to the formation of DMs (Figure 2A). Moreover, in a
second tumor (ESCC-06T), evidence of high-level amplification of this locus harboring FGFR1 was also identified
and similarly verified via FISH that showed clustered multiple FGFR1 signals around the centromere of chromosome 8 (Figure 2B), indicating high-level amplification of
FGFR1 in ESCC. DMs responsible for FGFR1 amplification
were not observed previously in ESCC. Combined with a
previous report that FGFR1 was overexpressed in ESCC,20
these findings indicate a oncogenic role of FGFR1 in
ESCC. Further functional studies indicated that knockdown of FGFR1 dramatically suppressed cell proliferation,
cell migration, and invasion in TE-1 and KYSE150 cells
(Figures S5A–S5C). A recent study has demonstrated that
focal amplification of the FGFR1 locus on chromosome
8p was associated with cellular dependency on FGFR1
and sensitivity to FGFR inhibitors.34 Consistent with
this, a pan-FGFR tyrosine kinase inhibitor has been shown
to block tumor proliferation in a subset of NSCLC cell lines
with activated FGFR signaling but has no effect on cells
that do not activate the pathway.35 Collectively, our results
suggest that FGFR1 might be an attractive therapeutic
target for ESCC.
Additionally, the small circular DNA molecule identified
in chromosome 8p of ESCC-16T contains LETM2. FISH
analysis further confirmed that LETM2 amplification was
due to extra-chromosomal amplification (Figure 2C).
Immunohistochemical analysis indicates that LETM2 was
upregulated in ESCC tumors and some ESCC cell lines (Figures 2D, 2E, and S6). LETM2 is a mitochondrial gene that is
expressed preferentially in spermatocyte to spermatozoon.34 It has been found amplified in breast cancer,
lung adenocarcinomas, and squamous cell lung carcinoma.34 However, the function of LETM2 has not been
studied in detail. Our result showed that LETM2 knockdown prevented cell proliferation but had no statistical
suppression of cell migration and invasion in KYSE150
and ECA109 cells (Figure 2F). Similar trends were observed

for WHSC1L1, another potential oncogene located in the
8p12 amplicon (Figures S5D–S5F). Together with genetics
observations, these functional analyses strongly implicate
these genes as amplification targets in ESCC.
Fusion Genes Caused by Chromosomal
Currently, little is known about the targetable fusion genes
underlying ESCC. We therefore screened gene fusion
events across 31 ESCC genomes and identified a total of
173 in-frame fusion genes and 231 out-frame fusion genes
affected by SVs (Table S4). Notably, in ESCC-16T, the chromothripsis-associated rearrangements led to the formation
of putative in-frame fusions involving genes TRAPPC9 at
8q24.3 and CLVS1 at 8q12. This fusion variant was predicted to result in an in-frame fusion of the TRAPPC9 50
UTR and exon 1–18 with the CLVS1 exon 4–5 and 30 UTR
(Figure 3A). Using primers within exon 18 of TRAPP9
and exon 4 of CLVS1, we confirmed the fusion transcript
in purified tumor cells from ESCC-16T (Figure 3B, left
and middle). FISH analysis using CLVS1 red probe and
TRAPPC9 green probe shows a yellow fusion signal indicative of translocation of TRAPPC9-CLVS1 (Figure 3B, right).
In this tumor genome, TRAPPC9 and CLVS1 are adjacent
genes on chromosome 8q that are transcribed in opposite
directions. TRAPPC9 (trafficking protein particle complex
9) is a 23-exon gene that encodes NIK- and IKK-b-binding
protein (NIBP), which activates NF-kB signaling via
directly interacting with and activating IKK-b and
MAP3K14 kinase.36 TRAPPC9 has been reported correlated
with colorectal tumorigenesis and tumor growth and was
implicated to be important for lapatinib response in a subgroup of ERBB2-amplified breast cancer.37 CLSV1, also
known as CRALBPL, was implicated to be upregulated in
hepatocellular carcinoma (HCC) and might be a marker
for HCC.38 The function of this fusion transcript in ESCC
need to be elucidated in future study.
Another notable inter-chromosome in-frame gene
fusion of EIF3E-RAD51B was detected in ESCC-19T. The
first exon of EIF3E on chromosome 8, encoding the
eukaryotic translation initiation factor 3 subunit, was predicted to join with the last two exons of RAD51B on chromosome 14, a protein that catalyzes repair of DSBs through
the process of homologous recombination and are critical
for genome stability (Figure 3D). The EIF3E-RAD51B translocation was validated in this tumor by independent PCR
sequencing and interphase FISH analyses (Figure 3E). Tumor suppressor or oncogenic effect of EIF3E either through
its role as a component of EIF3 translation initiation factor
or translation-unrelated function has been reported in

(C) Left: Junction points of TRAPPC9 in-frame fusions were shown in the transcripts of TRAPPC9 using the bottom symbols. Right:
The diversity partners of TRAPPC9 across different types of human cancers.
(D) Details and schematic of the EIF3E-RAD51B fusion transcript caused by interchromosomal translocation between chromosomes 8
and 14 in ESCC-19T.
(E) Validation of the EIF3E-RAD51B fusion transcript via PCR-sanger sequencing (left and middle) and FISH (right).
(F) Left: Junction points of RAD51B in-frame fusions were shown in the transcripts of RAD51B using the bottom symbols. Right: The
diversity partners of RAD51B across different types of human cancers.

264 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Figure 4. Kataegis on Chromosome 3 in ESCC-14T
(A) SVs observed on chromosome 3 in ESCC-14T. The upper panel represents different types of SVs indicated by lines with different
colors; the bottom panel shows normalized coverage for each window.
(legend continued on next page)

The American Journal of Human Genetics 98, 256–274, February 4, 2016 265

various types of human cancer.39 RAD51B, one member of
the human RAD51 (MIM: 179617) paralogs, plays a central
role in homologous DNA recombination.40 Increased
RAD51B protein level has been reported in various cancers,
especially gynecological tumors, and linked to uncontrolled recombination, genome instability, tumor recurrence and progression, and increased resistance of tumors
to radiotherapy and chemotherapy.40 Interestingly, translocation of RAD51B with other genes has been reported,
for example, HMGA2-RAD51B in uterine leiomyoma.41
However, to the best of our knowledge, the EIF3ERAD51B translocation has not been previously reported
in human cancer. Since the N- and C-terminal domains
of RAD51B were important to interact with other proteins
to catalyze the repair of DNA double-strand breaks, we
speculate that the in-frame fusions of EIF3E-RAD51B
might cause disruption of EIF3E and RAD51B function,
which could result in deregulated homologous recombination or translation initiation, contributing to the tumorigenesis of ESCC.
Recently, Yoshihara et al. analyzed RNA sequencing and
DNA copy-number data from 4,366 primary tumor samples and 364 normal samples spanning 13 tumor types.42
To further assess the recurrence of fusion genes identified
in ESCCs, we compared our data with the resource of
fusion transcripts from Yoshihara’s report.42 We did not
find in ESCC recurrent in-frame protein kinase fusions
such as FGFR1-TACC3 that was implicated in bladder urothelial carcinoma (BLCA), GBM, HNSCC, low-grade glioma
(LGG), and LUSC.42 We then focused on fusions with the
same gene fused to multiple different partners. Interestingly, we observed that some in-frame rearrangements
were not limited to ESCC but can be detected across cancer
at low frequency. For example, TRAPPC9 is paralogous to
many oncogenes such as LIMA1 (MIM: 608364), PTK2
(MIM: 600758), PSKH2, and others in BRCA, HNSCC,
and lung adenocarcinoma (LUAD) (Figure 3C). RAD51B is
a known oncogene and was found to form fusions with
various partners (e.g., CHD9, NPC2 [MIM: 601015],
PCNX [MIM: 613401]) in BRCA and LUAD (Figure 3F).
Moreover, the 30 partners of TRAPPC9 or EIF3E (e.g.,
CLVS1, RAD51B) have been reported to be upregulated in
human cancers,38,41 indicating the potential of these fusions to drive carcinogenesis.
Kataegis in ESCC
Besides chromothripsis, kataegis also contributes to locally
rearranged SVs accompanied with limited copy-number
states. Nik-Zainal et al. analyzed the mutational signatures
of 21 breast cancers and identified kataegis, a distinct

hypermutation phenomenon, in 61% of breast cancers,
indicating a direct relevance to tumor initiation and progression.16 To date, there is no implication of kataegis
and associated SVs in ESCC. Interestingly, we found locally
rearranged variations concentrated in chromosome 3 of
ESCC-14T and somatic mutations clustered in the region
of 16.9 Mb to 17.5 Mb (Figure 4). Although kataegis was
observed in one tumor, perhaps due to the limited sample
size, the prevalence of kataegis in other cancer types16,43
indicates a potential tumorigenic mechanism of kataegis
in ESCC development.
Breakage-Fusion-Bridge Drives Gene Amplification in
ESCC Tumors
Previous studies from cancer genomes support a BFB event,
which is known to begin with telomere loss and is characterized with a class of breakpoints called fold-back inversion.27 Therefore, we used fold-back inversion and
telomere loss to infer BFB events for each genome. In total,
we obtained 321 fold-back inversions (Table S5A), of which
chromosomes 11, 8, and 7 had the most fold-back inversions across 31 ESCC tumors (Figure 5A). Moreover, most
of fold-back inversions were mediated by microhomology
(Figures 5B and 5C), indicating that homology-mediated
fold-back capping of broken ends followed by DNA replication is an underlying mechanism of sister chromatid
fusion during BFB cycles in ESCC. In ESCC-11T, five chromosomes (chromosomes 5, 7, 8, 11, and 17) were affected
by BFB events (Table S5A). Hence, our large-scale breakpoint analysis of 31 ESCCs exhibited an important role
of BFB in tumorigenesis of ESCC.
Notably, fold-back inversions on chromosome 11 enriched in a minor cluster around CCND1 locus
(69,455,873–69,469,242 Mb) at 11q11.3 (Figure 5D). In
addition, we observed that 32 chromosomes involving
21 ESCCs displayed at least two inversions and a telomere
loss (Table S5B). Of these 21 ESCCs, 10 showed evidence of
BFB on chromosome 11, and 9 of them led to a focal amplification of CCND1 showing unbalanced amplified signals
(Figures 5E and S7), indicating that the CCND1 amplification was created by BFB cycles in ESCC. Together with the
cluster of palindromic junctions, the physical location of
the amplicon suggests the BFB cycles as the underlying
processes. Additionally, we also found inter-chromosomal
SVs enriched in CCND1 locus on chromosome 11
(Figure S8). Amplification of CCND1 has been reported in
a variety of tumors and might contribute to tumorigenesis.44 Specifically, CCND1 amplification and overexpression was observed and significantly correlated with lymph
node metastasis in ESCC.45 However, the underlying

(B) Each dot of the ‘‘rainfall’’ plots represents a single somatic mutation ordered on the horizontal axis according to its position in human
genome. The vertical axis denotes the genomic distance between mutations. The upper red triangles show the position of SV
(C) Highlight of kataegis region (chr3: 169–175 Mb) at increasing resolution to demonstrate microclusters within the macrocluster.
(D) Alternating processivity of kataegis in ESCC-14T. Long regions of C>T mutations are interspersed with regions of G>A mutations.
(E) The processive nature of C>T mutations (IGV image) within region (chr3: 174,897,540–174,897,710 Mb).
(F) Plots of flanking sequence of all C>X mutations and C>X mutations within the regions of kataegis in ESCC-14T.

266 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Figure 5. BFB Evidence across 31 ESCCs
(A) The number of fold-back inversions across 31 ESCCs.
(B) Sequence length in the breakpoints of fold-back inversion. Patterns of microhomology, non-template sequence, or direct end-joining
in the fold-back inversion across 31 ESCCs are shown.
(C) Genomic patterns of fold-back inversions. Histogram showing the distance between the two inverted ends in the set of fold-back
(D) Distribution of the fold-back inversions on chromosome 11 across 31 ESCCs. Blue arrow indicates a minor cluster around CCND1
locus (11q11.3).
(E) Representative maps show amplification of CCND1 as a result of BFB in three tumors. FISH validation shown on right panel.

mechanism of CCND1 amplification has not been elucidated. Our results demonstrated that at least two mutational mechanisms, focal amplification via BFB cycles
and inter-chromosomal translocations, result in CCND1
amplification in ESCC.
Additionally, we observed that regions amplified by BFB
cycles harbor oncogenes such as EGFR (MIM: 131550)
(2/31), ERBB2 (MIM: 164870) (1/31), MMPs (1/31), and
MYC (MIM: 190080) (1/31) (Figure 6), suggesting that
BFB plays an important role in gene amplification in
ESCC tumors. In the literature, MYC loci comprise the
most significant regions of amplification observed in
ESCCs and have been implicated as a reasonable indicator

of the accumulation of various activated and inactivated
genes involved in carcinogenesis of ESCCs, suggesting
deregulation of MYC as a driver event.46 EGFR is an established therapeutic target that is often overexpressed as a
consequence of gene amplification in human cancers
including ESCCs.47 ERBB2 amplification was observed in
breast, esophageal, and other types of cancer and has
been a target of anticancer agents.48 MMPs amplification
was reported in some human cancers but not ESCC.48 It
has been found that regions of DNA gain in cancer rarely
coincide with regions of loss and vice versa, suggesting a
specialized function for regions characterized by either
gain or loss in cancer.49 Therefore, understanding the

The American Journal of Human Genetics 98, 256–274, February 4, 2016 267

Figure 6. BFB-Derived Gene Amplifications in ESCC
Amplification of other oncogenes caused by BFB events including EGFR (A), ERBB2 (B), MMPs (C), and MYC (D).

mechanisms that drive SVs and the gene changes that
result from them has significance. WGSs revealed that
amplification of MYC, EGFR, and ERBB2 might derive
through TDs or chromothripsis-derived DMs.1,15,25 However, our data support that BFB events that occurred in
ESCCs led to amplification of these genes, which was not
proposed previously in ESCC. Therapies targeting these
amplified oncogenes would be more practical in ESCC.
Copy-Number Alterations
To investigate copy-number change across ESCC genomes
and potential affected genes, we applied Patchwork to
determine CNAs based on WGS data of 31 ESCCs and
found 19 ESCCs that could be used to determine absolute
copy number. Consistently, frequent arm-level changes
were observed in ESCC, including frequent copy-number
gains of 3q, 5p, 7p, 8q, 12p, 17p, 20p, and 20q and universal deletions affecting 3p, 4p, 4q, 5q, 10p, 13q, and 21q
(Figure S9A). Moreover, 19 ESCCs harbored fewer events
of copy-number loss than copy-number gain; meanwhile,
70% of loss of heterozygosity (LOH) was copy neutral
loss of heterozygosity (CN-LOH) in ESCC. Specifically, we
observed frequent CN-LOH on 13q and 17p (Figure S9B).
In addition, we found that WGD occurred in 13 out of
19 ESCC genomes. Despite evidence that WGD can result
in genetic instability and accelerate oncogenesis,50 the
incidence and timing of such events had not been broadly
characterized in ESCC. Our results indicate that ESCC
tumors have alterations affecting the entire length of chromosome 13q and 17p such as, perhaps, whole chromosome deletion with duplication.
Furthermore, to obtain CNA targets, we applied GISTIC
to copy-number profiling from a combination of 31 WGS
and 123 CGH data.19,22 This analysis yielded 11 amplification peaks and 13 deletion peaks, including cancer genes
EGFR, CDK6 (MIM: 603368), AKT1 (MIM: 164730), MYC,
CCND1, CDKN2A, and others (Table S6). Specifically, we
identified a focal amplified region corresponding to

CDCA7 in 5 out of 31 ESCC genomes with 2 having
high-level copy number (>6 copies; Figure 7A). Moreover,
we observed that most of individuals with ESCC tumors
showed statistically higher expression level of CDCA7
compared with that of normal tissues as determined by
real-time PCR (Figure 7B) and immunohistochemistry analyses (Figures 7C and S10). CDCA7 is a downstream target
of MYC and E2F transcription factors and participates in
cell cycle progression as a transcriptional regulator of the
expression of myriad of target genes.51 Previous transformation studies with cell lines in vitro, analysis of CDCA7
levels in human cancers, and in vivo tumorigenic studies
in transgenic mice all support a role for CDCA7 in tumorigenesis.51 However, it has limited implication in ESCC;
the mechanism of how CDCA7 is involved in tumorigenesis remains largely unknown. Our result showed that
CDCA7 knockdown significantly inhibited cell growth
and promoted cell apoptosis but had no differential effect
on cell migration and invasion in ESCC cells (Figures 8A–
8D), indicating that CDCA7 might involve cell proliferation and apoptosis but not metastasis in ESCC. Moreover,
CDCA7 knockdown led to the decrease of phosphoERK1/2, an essential downstream component of MAPK
pathway regulating cell proliferation, whereas no significant effect was seen in AKT pathway (Figure 8A). To further
determine the potential targets of CDCA7 in ESCC, we performed RNA-seq of CDCA7 knockdown cells and cells
transfected with pLVshRNA-puro vector (used as controls).
Indeed, we observed a positive and highly significant
enrichment of the expression of cell proliferation or
apoptosis-associated target genes, including FGF21
([MIM: 609436] a MAPK pathway-related gene) and cellapoptosis-associated genes TRAIL-R, CASP10 (MIM:
601762), IL1R1 (MIM: 147810), CASP7 (MIM: 601761),
BCL2 (MIM: 151430), and CASP9 (MIM: 602234). These
genes all had outlier expression levels in CDCA7 knockdown cells compared to that of controls (Figure 8E and
Table S8). Specifically, a significant decrease of FGF21 in

268 The American Journal of Human Genetics 98, 256–274, February 4, 2016

Figure 7. Amplification and Overexpression of CDCA7 in ESCC Tissues
(A) Focal recurrently amplification of CDCA7 shown in five ESCCs. The red line represents genomic position of CDCA7; the dot represents normalized coverage of genomic position (100 kb per window) along chromosome 2.
(B) mRNA expression level of CDCA7 examined from 52 matched normal/tumor ESCC tissues. p value is given by Student’s t test.
(C) Left: Represent images display strongly cytoplasm positivity in ESCC tissues. Right: Expression of CDCA7 was markedly increased in
ESCC tissues compared to that of normal esophagus tissue based on judgment of IHC staining intensity.
(D) CDCA7 expression level in multiple ESCC lines determined by RT-PCR and western blotting.

CDCA7 knockdown cells suggests that CDCA7 might regulate cell proliferation via FGF21-ERK1/2 MAPK pathway
rather than other pathways in ESCC tumorigenesis
(Figure 8E). In addition, CDCA7 knockdown led to the
increased expression levels of TRAIL-R, CASP10, IL1R1,
and CASP7 and the decreased expression levels of BCL2
and CASP9 (Figure 8E and Table S8), indicating that these
genes might be critical for CDCA7 to regulate cell
apoptosis. Together with genetic observations, these functional data indicate that CDCA7 might act as an oncogene
possibly through deregulation of cell proliferation and
apoptosis in ESCC.

In this study, we report a comprehensive description of SVs
that characterize ESCC and demonstrate the relative contributions and variability of different mutational mechanisms
underlying SVs within ESCC genomes. We found that NHEJ
and alt-EJ contribute the most to deletions and translocations. Our findings define a prevalence of locally arranged
genomes across 31 ESCC genomes and some of them were
delivered by chromothripsis, kataegis, or BFB events.
A number of well-known cancer-associated genes (e.g.,
FGFR1, CDKN2A) and several unreported ESCC-related
genes (e.g., LETM2, CDCA7, TRAPPC9-CLVS1, EIF3E-

RAD51B) affected by these events were described here.
Furthermore, our data provide the potential mechanisms
for oncogene amplification or fusion gene formation,
which might be critical for tumorigenesis of ESCC.
In studying SVs across ESCC genomes, we observed
locally rearranged SVs with either limited (e.g., chromothripsis or kataegis) or substantial (e.g., BFB) copy-number
states (Figure S3). In addition to the predominant BFB cycles
that were accumulated in a step fashion,27 chromothripsis,
a phenomenon in which one or a few chromosomes are
shattered into pieces and randomly stitched together in a
single event,15 was observed in one ESCC genome, which
is consistent with its common rate of 1/40 cancer genomes.11 Moreover, kataegis, most likely co-occuring with
large-scale rearrangement in a region of the genome at
one time,15 was also observed in an ESCC genome. Combined with the fact that kataegis is remarkably common
in human cancers,16,43 we speculate that it is more likely
to have biological significance. However, due to the limited
sample size, we observed kataegis in only one ESCC
genome. At some point in the future, larger numbers of
ESCC genomes at higher resolution will be necessary to
create a comprehensive catalog of the significant SVs and
define the biological significance of these events in ESCC.
Besides copy-number alterations, translocations in chromothripsis led to gene fusions (e.g., TRAPPC9-CLVS1,
EIF3E-RAD51B) (Figure S3). Chromothripsis, occurring as

The American Journal of Human Genetics 98, 256–274, February 4, 2016 269

Figure 8. CDCA7 Deregulates Cell Proliferation and Apoptosis in ESCC
(A) Knockdown of CDCA7 was demonstrated by immunoblotting; GAPDH was used as loading control. Meanwhile, phospho-ERK1/2,
ERK1/2, phospho-AKT1, and AKT1 were shown.
(B) Knockdown of CDCA7 significantly prevented cell growth in ECA109 as monitored by MTT (top) and colony formation assays (bottom). ECA109 and cells transfected with pLVshRNA-puro-SCR (scrambled sequence) were used as controls.
(C) Knockdown of CDCA7 shows no significant effect on cell migration and invasion.
(D) Knockdown of CDCA7 significantly promoted cell apoptosis. All data are mean 5 SD; each experiment was performed in triplicate. *p < 0.05; **p < 0.01.
(E) Summary of potential CDCA7 target genes involved in cell apoptosis and proliferation in ESCC. The bold characters represent genes
with log2 ratio more than 2. Red represents upregulated genes and blue represents downregulated genes.
Detailed information shown in Table S8.

a relatively early tumorigenic event, is thought to represent a driving force of cancer development and progression.7,15 For example, chromothripsis is implicated as a
frequent driver event in uterine leiomyomas, resulting in
increased expression of translocated HMGA1 and
HMGA2.41 However, distinguishing driver mutations
from passenger mutations is challenging. For SVs, recurrence is often used to estimate the likelihood of fusion being a driver; however, because most driver fusions have

very low frequency, many studies have small sample sizes
(as in the case here), and detection sensitivity might be
low, it is hard to define the molecular characteristics of
driver fusion.42 In ESCC genomes, we identified two inframe fusions (TRAPPC9-CLVS1 and EIF3E-RAD51B) via
RT-PCR Sanger sequencing and FISH. We did not find the
same fusion genes (TRAPPC9-CLVS1 and EIF3E-RAD51B)
in other human cancers. This phenomenon also happens
in other human cancers. For example, none of predicted

270 The American Journal of Human Genetics 98, 256–274, February 4, 2016

fusion events occurred in more than one sample in pancreatic cancer.26 Recent WGSs for structural mutations in cancers showed that most fusion transcripts were singletons
unique to individual tumors and not detected in other
samples.1 Alternately, we identified additional fusion partners for TRAPPC9 as well as RAD51B. Although these findings have not been validated by functional studies, they
illustrate the potential of these fusions to drive carcinogenesis. Further in vitro and in vivo studies are needed to better understand the biological significance of these fusion
transcripts in ESCC.
Additionally, BFB events were operative in approximately 68% of ESCCs, indicating that the BFB cycle is an
important underlying process for genome instability and
gene amplification in ESCC. The BFB events initiate amplification of cancer-associated genes and occur predominantly in early cancer development rather than later
stages.25 End-to-end chromosome fusions are often seen
in association with telomere erosion and it might be that
the dsDNA break initiating BFB repair results from telomere loss. Hence, detecting telomere loss indicative of a
BFB event might provide preventive implication for
ESCC. However, although we identified BFB-derived amplification of cancer-associated genes, we could not identify
candidate target genes from most BFBs because these
amplified segments contain many more passenger events.
Additionally, a BFB event was defined when a chromosome
had at least two inversions and clearly telomere-boundary
copy-number loss adjacent to the fold-back inversions.27 It
is also possible that some chromosomes without clear
telomere-boundary copy-number loss might suffer BFBs.
Unfortunately, we could not identify these SV events via
current strategies. Existing methods for the detection of
SV events show high sensitivity and specificity but still
have limitations. In the future, advanced methodology
will enable the identification of these events.
We and others previously reported that the two types of
esophageal cancers presented different mutational patterns and signatures at the level of SNVs. Specially, a higher
frequency of C>G transversions occurred in ESCC than
EAC whereas A>C transitions were more frequent in
EAC than ESCC.22 A recent combined study of WGSs
(22 EACs) and SNP arrays (101 EACs) reported genomic catastrophes that occurred in EAC.14 We then compared the
SV events between these two types of esophageal cancers.
Evidence of chromothripsis, BFB cycles, and kataegis
were reported in both ESCC and EAC. Although TP53,
which has been linked to chromothripsis in human cancers,52 was mutated at high frequency in both ESCC and
EAC, we found that the frequency of chromothripsis
tended to be lower in ESCC than in EAC. Moreover, we
note that chromothripsis resulted in DM-derived FGFR1
amplification in ESCC but led to DM-derived MYC amplification in EAC. Otherwise, the high-level amplification of
MYC is due to BFB cycles in ESCC, indicating that at least
two different mechanisms are responsible for MYC amplification in tumors. Additionally, the genes affected by

BFB cycles in these two types of esophageal cancers display
differences. BFB cycles are scattered in three genes (KRAS
[MIM: 190070], MDM2 [MIM: 164785], and RFC3 [MIM:
600405]) in EAC; in ESCC, they are mainly focused in
CCND1 and also scattered in MYC, MMPs, EGFR, and
ERBB2 (Figure S11). Unlike ESCCs, EACs arise in a highly
genotoxic environment in which the distal esophagus is
exposed to high levels of local and systemic injury from
reflux of acid, bile, and other gastric contents.53 These findings would suggest that genomic catastrophes, gene activation through chromosomal rearrangements, and telomere
integrity might be driving carcinogenesis in esophageal
cancer, and the dominant SV type might be different between ESCC and EAC. Further understanding of these
events might lead to novel strategies for detection and
treatment of esophageal cancers.
Collectively, our findings demonstrated diverse models
of SVs contributing to the mutational landscape, with
BFB being the most extreme form across ESCC genomes.
Besides somatic point mutations and CNAs reported in
ESCC previously,19–22 our findings highlight the oncogenic drives of ESCC through different types of SVs and
suggest that complex genomic rearrangements, such as
chromothripsis and BFB, are an integral part of mutation
mechanisms contributing to ESCC development and
should be considered along with simple genomic changes
when applying genome-guided treatment strategies.
Together with the landscape of point mutations or CNAs
described previously, these findings provide a systems
explanation for the maintenance of ESCC state. Additional
larger panels of ESCC tissues will need to be studied to
determine the broader applicability of these results.
Currently, identifying SVs is still challenging and remains
largely unsolved. Although much effort has focused on
candidate genes affected by SVs, most SVs actually occur
in non-coding regions.18,30,42 As ENCODE project
explored potential functions of non-coding sequence,54
more advanced technology is required to characterize
those SVs that occurred in non-coding regions and define
their contribution in tumorigenesis of ESCC.
Accession Numbers
The whole-genome sequencing data of 31 pairs of tumors and
matched normal tissues reported in this paper have been deposited to the European Genome-phenome Archive (EGA) under
accession numbers EGAS00001001487 and EGAS00001000709.

Supplemental Data
Supplemental Data include 11 figures and 8 tables and can be
found with this article online at http://dx.doi.org/10.1016/j.

This work was supported by funding from the National Natural
Science Foundation of China (81330063 and 81272189 to Y.C.,

The American Journal of Human Genetics 98, 256–274, February 4, 2016 271

81230047 to Q.Z., 81272694 to X.C., 81201956 to J.L., 81402342
to L.Z., and 81502135 to P.K.), the Key Project of Chinese Ministry
of Education (NO213005A to Y.C.), the Specialized Research Fund
for the Doctoral Program of Higher Education (20121417110001
to Y.C.), a research project supported by Shanxi Scholarship Council of China (2013-053 and 2015-key3 to Y.C.), the Innovative
Team in Science & Technology of Shanxi (2013-23 to Y.C.), the
Program for the Outstanding Innovative Teams of Higher
Learning Institutions of Shanxi (2015-313 to Y.C.), and the 973 National Fundamental Research Program of China (2015CB553904
to Q.Z.).
Received: August 22, 2015
Accepted: December 15, 2015
Published: January 28, 2016

Web Resources
The URLs for data presented herein are as follows:
European Genome-phenome Archive (EGA), https://www.ebi.ac.
OMIM, http://www.omim.org/

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