Genomic analyses reveal FAM84B and the NOTCH pathway are associated with the progression of esophageal squamous cell carcinoma
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Esophageal squamous cell carcinoma (ESCC) is the sixth most lethal cancer worldwide and the fourth most lethal cancer in China. Genomic characterization of tumors, particularly those of different stages, is likely to reveal additional oncogenic mechanisms. Although copy number alterations and somatic point mutations associated with the development of ESCC have been identified by array-based technologies and genome-wide studies, the genomic characterization of ESCCs from different stages of the disease has not been explored. Here, we have performed either whole-genome sequencing or whole-exome sequencing on 51 stage I and 53 stage III ESCC patients to characterize the genomic alterations that occur during the various clinical stages of ESCC, and further validated these changes in 36 atypical hyperplasia samples.
Recurrent somatic amplifications at 8q were found to be enriched in stage I tumors and the deletions of 4p-q and 5q were particularly identified in stage III tumors. In particular, the FAM84B gene was amplified and overexpressed in preclinical and ESCC tumors. Knockdown of FAM84B in ESCC cell lines significantly reduced in vitro cell growth, migration and invasion. Although the cancer-associated genes TP53, PIK3CA, CDKN2A and their pathways showed no significant difference between stage I and stage III tumors, we identified and validated a prevalence of mutations in NOTCH1 and in the NOTCH pathway that indicate that they are involved in the preclinical and early stages of ESCC.
Our results suggest that FAM84B and the NOTCH pathway are involved in the progression of ESCC and may be potential diagnostic targets for ESCC susceptibility.
KeywordsNext-generation sequencing ESCC FAM84B NOTCH signaling
Esophageal squamous cell carcinoma
Family with sequence similarity 84, member B
Genomic identification of significant targets in cancer
Head and neck squamous cell carcinoma
Integrative genomics viewer
Somatic copy number alterations
Significantly mutated gene
Single nucleotide variation
Target capture sequencing
Esophageal squamous cell carcinoma (ESCC) is the eighth most common and the sixth most lethal cancer worldwide with approximately 70 % of global esophageal cancer occurring in China . ESCC is more prevalent in non-Caucasian populations, with the highest incidence in the Taihang Mountain region of North-Central China , and known risk factors include environmental factors such as dietary habits (e.g. the consumption of hot food and betelnut chewing), family history, alcohol abuse and tobacco smoking [2, 3, 4]. ESCC has a highly variable clinical outcome, with an excellent prognosis for stage I and II tumors but a poor outcome for later stage tumors . Currently, three types of treatment are available: surgery, chemotherapy and radiation therapy . Among these, chemoradiotherapy (CRT) is recognized as one of the most effective treatments for ESCC . However, as with other treatments, the clinical response to CRT varies between individuals, and this has a major influence on clinical care outcomes [7, 8]. The 5-year survival rate ranges from 10 % to 25 %, and is highly dependent on tumor stage .
The genomic characterization of ESCC tumors, particularly those from different stages of the disease, is likely to reveal underlying oncogenic mechanisms and new cancer-associated genes . ESCC initiates from atypical hyperplasia and progresses to carcinoma in situ and then invasive carcinoma . Biomarkers identified from atypical hyperplasia or from early-stage tumors may lead to the development of new diagnostic, prognostic, therapeutic and prevention strategies. Analyses of somatic copy number alterations (SCNAs) using array-based technologies have identified frequently altered regions such as 3q26 , 11q13.3  and 8q24.3 , and exome-wide investigations have revealed point mutations in the well-known cancer-associated genes TP53, PIK3CA, CDKN2A and novel genes ZNF750, FAT1, FAT2 and FAM135B [15, 16, 17]. However, the genetic or mechanistic alterations related to the progression of ESCC have not been fully elucidated. Thus, there is an urgent need to elucidate the genomic alterations and molecular events associated with the various ESCC stages to enhance our understanding of these tumors, aid in early diagnosis, identify therapeutic targets and develop prevention strategies.
In this study, we used whole-genome sequencing (WGS) of 14, whole-exome sequencing (WES) of 90 and deep target capture sequencing (TCS) of 96 ESCC tumors and adjacent normal tissue from patients recruited from the Taihang Mountain region in North-Central China. This cohort includes 51 stage I and 53 stage III cases from the Han Chinese population who live in the Shanxi and Henan provinces. The genetic alterations that we identified were further validated through next-generation sequencing (NGS) and their association with a specific clinical stage was confirmed by comparison with 36 atypical hyperplasia (i.e. premalignant) tissues.
Genomic DNAs were extracted from 3 stage IA, 48 stage IB, 31 stage IIIA, 17 stage IIIB, 5 stage IIIC tumors and matched normal tissues (Additional file 1: Table S1) . WGS libraries (500 bp inserts) and WES libraries (150–200 bp inserts) were constructed and sequenced on an Illumina HiSeq 2000 sequencer using 90 bp paired-end reads. TCS was performed following a protocol similar to WES (see Methods). Sequencing reads from the Illumina HiSeq 2000 sequencer were processed by Illumina software and passed to the in-house pipeline to determine somatic point mutations, indels, and copy number variations. A significance analysis method, MutSigCV, was used to identify significantly mutated genes (SMGs). Fluorescence in situ hybridization (FISH), qPCR copy number analyses or targeted PCR-Sanger sequencing was used to validate stage-associated genetic alterations in 36 atypical hyperplasia tissues. To introduce the gene that encodes the family with sequence similarity 84, member B protein (FAM84B; also known as NSE2) or scrambled control siRNAs into ESCC cells, we used the pLKO.1 virus according to the manufacturer’s instructions. Western blotting was carried out using standard protocols  and tissue microarray resourced immunohistochemistry analysis and functional assays were performed as specified in the Methods section.
The genome landscape of ESCC and its stage-associated variations
We performed genome sequencing on DNA from 104 ESCC tumors and matched adjacent normal tissues, including 51 stage I and 53 stage III cases (Additional file 1: Table S1) . Five stage I and nine stage III tumors underwent WGS (median coverage of 65×); 46 stage I and 44 stage III tumors underwent WES (median coverage of 132×; Additional file 2: Figure S1). The average number was 3.9 coding mutations/Mb for WGS and 2.4 non-silent mutations/Mb for WES (Additional file 3: Table S2) . Candidate non-silent mutations identified from 48 stage I and 48 stage III tumors were selected for TCS (at least 365×; Additional file 3: Table S2C) . The validation rates were 97.8 % for identified single-nucleotide variations (SNVs) and 58 % for indels. Consequently, we obtained 10,330 somatic point mutations in total: of these, 65 % resulted in missense changes and 6 % resulted in nonsense changes. There were 184 alterations of splice sites and 90 small indels: of the indels, 84 % introduced frameshifts and 16 % were in-frame (Additional file 4: Table S3 and Additional file 5: Figure S2B). We found 73 (range 24–189) non-silent mutations per tumor in this cohort, and this rate is in line with published rates (Additional file 5: Figure S2C) [15, 16, 17, 20], underscoring the representative nature of our analysis.
Significantly mutated genes
For most cancer types, genomic landscapes consist of many ‘hills’ (corresponding to genes that are altered in a low percentage of the tumors) and a small number of ‘mountains’ (genes that are altered in a higher percentage of tumors) . This also holds true for ESCC, as revealed by our cohort (Additional file 10: Figure S6; Additional file 11: Table S5) . Only 14 genes (TP53, TTN, MUC16, NOTCH1, FAT1, PIK3CA, CSMD3, PCLO, LRPIB, MLL2, EP300, SYNE1, SPTA1 and PKHD1L1) contained somatic mutations that were detected in at least 10 % of the samples whereas 85 genes were mutated in 5-10 % of the tumors. The remaining 1,309 genes were altered in 2–5 % of the cohort of 104.
Identical mutations suggest stage-associated NOTCH pathway alterations
Of related interest, 8 % of ESCC tumors (12 % of stage I tumors and 4 % of stage III tumors) harbored mutations in the F-box protein FBXW7 (Fig. 5a). FBXW7 is the substrate-recognition subunit of an SCF-type ubiquitin ligase complex that regulates the cell cycle by targeting many proto-oncoproteins and antiapoptotic molecules including NOTCH1, cyclin E, c-Myc, c-Jun and mTOR for ubiquitin-mediated degradation. It therefore acts as a tumor suppressor protein  and has been found to be mutated in various tumors [33, 34]. Of the eight mutations detected in our cohort, two nonsense and one frameshift deletion are inactivating mutations (Fig. 5b, bottom panel). Moreover, most of the mutations occur within the WD40 domain involved in substrate recognition; mutation of this site prevents the recognition of targets such as NOTCH1 for degradation. Taken together, we identified 31 mutations in NOTCH family genes. Most of these mutations result in truncated protein products or have deleterious effects, suggesting that NOTCH signaling is significantly disrupted in our sample set and may be one of the main mechanisms associated with the development of a subgroup of ESCCs.
In addition to the NOTCH pathway, we found that mutations in genes involved in several major metabolic pathways were significantly enriched in the stage I group. In particular, mutations occurred in genes associated with pyrimidine metabolism (31 % of stage I tumors versus 13 % of stage III), glycine/serine/threonine metabolism (16 % versus 2 %) and fructose and mannose metabolism (16 % versus 2 %). Conversely, the hedgehog (Hh) signaling pathway showed a significantly higher mutation frequency in the late stage III group (p < 0.05, Fisher’s test; Additional file 14: Table S8). Thus, despite there being no marked differences between the tumors at various stages at the level of individual genes, such differences seem to exist at the level of pathways. For other frequently or significantly mutated genes, we found no significant correlation between their mutation frequencies and tumor stage.
Clinical screening and surveillance approaches (i.e. upper gastrointestinal endoscopy, barium esophagram, non-endoscopy-based balloon cytology and serology tumor markers etc.) for the early diagnosis of ESCC are currently limited . Hence, biomarkers indicating ESCC pathogenic processes are urgently required to diagnose and facilitate early intervention. Initial genomic reports have described whole-genome and -exome sequencing for ESCC patients [15, 16, 17]. Although informative, these studies did not reveal the genomic differences between tumors from various clinical stages.
In this study, we report the genomic characterization of different stages of ESCC based on either WGS or WES of 104 ESCC patients. We show that amplifications of 8q and deletions of 4p-q and 5q may be associated with the early stages of ESCC. Moreover, we identify FAM84B as a novel ESCC-associated gene. Further functional and clinical analyses strongly indicate that FAM84B (located at 8q24.13-q24.21), which is highly expressed in dysplasia and ESCC patients but not normal esophagus tissues, may contribute to oncogenesis in ESCC and that targeting FAM84B may be a promising strategy for the diagnosis of susceptibility and the early stages of ESCC.
Similar to many other cancers, particularly esophageal adenocarcinoma (EAC) , the well-defined cancer-associated genes such as TP53, PIK3CA and CDKN2A were also identified as SMGs in ESCC, providing evidence of a common dysfunction in cell-cycle control and apoptotic signaling. Importantly, the analysis of ESCC genomes reveals that mechanisms previously suspected to have a role in the biology of ESCC [15, 16, 17, 35] but not in EAC  (for example, NOTCH1 and FBXW7) are indeed involved in ESCC development (Additional file 15: Figure S7). Moreover, genomic characterization of different stages of ESCC tumors led us to identify dysregulated NOTCH1 and NOTCH signaling predominantly in stage I tumors, indicating the involvement of this gene and its pathway in the early development of ESCC. Thus, the prevalence of NOTCH1 provides a potential biomarker to detect ESCC in its early stages. NOTCH signaling has long been known to function in developmental processes and in regulating the self-renewal of tissues [36, 37]. Whereas activating mutations in NOTCH1 have been identified in T-cell acute lymphoblastic leukemia and breast cancer [38, 39, 40], a tumor suppressor role for NOTCH signaling has been suggested in tumor types such as chronic myelomonocytic leukemia (CMML) , HNSCC  and lung SCC . The spectrum of mutations involving the NOTCH pathway in our cohort is consistent with it having a tumor suppressor role in ESCC rather than an oncogenic function. It is important to note that although NOTCH1 has been reported to be a poor survival marker in human ESCC , the correlation between genotype and the expression level of NOTCH1 remains ambiguous.
In summary, we used the genomics data described above, together with a large-scale tissue platform, to pinpoint molecular features linked to the early occurrence of ESCC. We provide genetic and functional evidence suggesting that overexpression of FAM84B is related to the progression of ESCC and may be a potential diagnostic and/or therapeutic target. We also identify the genomic aberrations that frequently alter the NOTCH pathway in stage I ESCC tumors and which may be useful for predicting early ESCC onset. Collectively, our results highlight the substantial stage-associated genetic diversity underlying ESCC and facilitate the understanding of the molecular defects that lead to early disease onset, which may ultimately provide a set of potential targets for its early diagnosis and prevention.
Samples and clinical data
We recruited tumor samples and adjacent normal tissues from 104 ESCC patients from the Han Chinese population who live in the Shanxi and Henan provinces, Taihang Mountain, North-Central China. All samples were obtained before treatment according to the guidelines of the local ethical committees (IRB of Shanxi Medical University, Approval No. 2009029, and the Ethics Committee of Henan Cancer Hospital, Approval No. 2009xjs12). This study was approved by the ethical committee of the Shanxi and Henan, China. All ESCC cases collected for this study were staged using the American Joint Committee on Cancer (AJCC) Cancer Staging Standards, 7th Edition (2010). This cohort includes 51 cases of stage I tumors and 53 cases of stage III tumors. Different subsets of patients were assayed on each platform: 14 tumors and matched normal samples from 14 patients, including 5 cases of stage I and 9 cases of stage III, had WGS data available (65×); 90 samples from 46 cases of stage I and 44 cases of stage III, had WES data available (132×); 96 of the 104 samples including 48 cases of stage I and 48 cases of stage III had TCS data available (365×) . A detailed description of the clinical characteristics of this cohort is shown in Additional file 1: Table S1. A summary of the next-generation sequencing analyses in this study is presented in Additional file 2: Figure S1.
For WGS, genomic DNA extracted from 5 stage I tumors, 9 stage III tumors and matched normal tissues were randomly fragmented and purified. The WGS libraries were constructed and subjected to WGS on an Illumina HiSeq 2000. At least 65× target depth and 30-fold haploid coverage for tumors and normal samples were achieved in all samples. A detailed description is presented in Additional file 3: Table S2.
For WES, the qualified genomic DNAs from 46 tumors from stage I, 44 tumors from stage III and matched normal tissues were randomly fragmented, amplified by ligation-mediated PCR (LM-PCR), purified and hybridized to the NimbleGen SeqCap EZ exome (44 M, Roche Company, Indianapolis, IN, USA) array for enrichment. Each captured library was subjected to an Illumina HiSeq 2000 platform for high-throughput sequencing. The mean coverage achieved was 130× in the tumor and 133× in the normal tissues. A detailed description is presented in Additional file 3: Table S2B. The Agilent SureSelect in Solution is described elsewhere .
Illumina sequencing analysis pipeline
For detection of somatic point mutations, sequencing reads from an Illumina HiSeq 2000 sequencer were aligned to the human reference genome (hg19) sequence using Burrows-Wheeler Aligner (BWA) . After removing duplicated reads (redundant information produced by PCR) using the SAMtools, an in-house cancer sequencing analysis pipeline (CSAP) was used to identify somatic mutations [15, 46]. Several recently published studies have compared the somatic mutation calling tools (e.g. SomaticSniper, Virmid, Strelka, MuTect and VarScan2) and found that VarScan2 outperformed all other tools in detecting high-quality somatic mutations (high coverage and allele frequency) [47, 48, 49]. Therefore, we detected mutations by VarScan2.2.5 with previously determined parameters . High-confident SNVs were annotated with ANNOVAR and used in follow-up analyses . Briefly, we used an in-house pipeline for statistic sequencing error rates, base calling accuracy and ATCG base contents of raw sequencing data. Sequence data per lane needed to meet the following quality controls: i.sequencing error rates ≤ 2.5 %; ii. base calling accuracy, measured by the Phred quality score (Q score), Q20 ≥ 80 %, Q30 ≥ 75 %; iii. GC or AT separation rate ≤ 0.4 %. We applied SOAPnuke to remove adapter and filter low quality reads . Reads were kept if they met the following criteria: i. adaptor rate ≤ 10 %; ii. N rate of every single read ≤ 10 %; iii. low quality base rate (base quality < 5) of every single read ≤ 50 %. After removing reads containing sequencing adaptors and low-quality reads, the high-quality single-end reads were aligned to the NCBI human reference genome (hg19) using BWA by default parameters. Each sample should meet the following criteria: i. average sequencing depth of WGS ≥ 30×, average sequencing depth of WES ≥ 100×; ii. Mapping rate ≥ 95 %; iii. Mismatch rate ≤ 10 %. High-confident somatic SNVs were called when they met the following criteria: i. both the tumor and normal samples should be covered sufficiently (≥10×) at the genomic position; ii. the variants should be supported by at least 10 % of the total reads in the tumor compared with less than 2 % in normal tissues; iii. the variants should be supported by at least three reads in the tumor; iv. distance between adjacent somatic SNV distance should be over 10 bp; v. mapping qualities of reads supporting mutant alleles in the tumor should be significantly higher than 30 (Wilcoxon rank sum test, p < 0.2); vi.base qualities of reads supporting mutant alleles in the tumor should be significantly higher than 20 (Wilcoxon rank sum test, p < 0.05); vii.mutations should not be enriched within 5 bp 5′ or 3′ of read end (Wilcoxon rank sum test, p < 0.1); viii. the changes of mutant allele frequency between tumor and normal should be statistically significant (Fisher’s exact test, p < 0.05); ix. reads supporting mutations should not be significantly enriched within either forward or reverse genomic strand (Fisher’s exact test, p < 0.0001).
Detection of small indels
The indel calling step was performed by a GATK SomaticIndelDetector with default parameters . The high-confident indels were identified by an in-house pipeline and further annotated with ANNOVAR  as either germline or somatic based on whether any evidence for the event at the same locus was observed in the normal data. High-confident somatic insertions and deletions (indels) were called through the following steps: i. candidate somatic indels were predicted with the GATK SomaticIndel Detector with default parameters; ii.for each predicted somatic indel, local realignment was performed with combined normal and tumor BAM files; iii. high-confident somatic indels were defined after filtering germline events.
Target capture sequencing
To provide high-confident mutations, non-synonymous mutations identified in the WGS and WES sets were selected for TCS. Briefly, non-synonymous SNVs (6873) and indels (125) in coding regions identified from 96 of the 104 samples were designed on a Nimblegen customized capture array (Roche Company, Indianapolis, IN, USA). Genomic DNAs from 48 stage I tumors, 48 stage III tumors and matched normal tissues were fragmented and libraries were constructed following the same method as the exome-capture experiment. For SNVs and small indels, we tiled ~200 bp targets across the variant of interest, including a minimum buffer of 100 bp in each direction. After library preparation and hybridization, sequencing was performed on an Illumina HiSeq 2000 platform. Each tumor was sequenced to at least a depth of 300× and SNVs were called with the same pipeline except that variant allele frequency was decreased to 5 % to guarantee that low-frequency mutations were retained. Somatic indels were manually inspected across 104 normal samples to remove germline mutations. The mean coverage achieved was 365× in TCS with 354× in tumors and 375× in the normal samples. A detailed description is presented in Additional file 3: Table S2C. Validation lanes were aligned to the reference sequence and BAM files created in the same manner as described above.
DNA copy number analysis
We performed SegSeq , a widely used method to identify copy number variation (CNVs) by comparing a tumor sample with a matched normal sample, to infer somatic CNVs in ESCC genomes based on WGS reads. The resulting copy number segments were mapped to individual genes to determine gene-level copy numbers and copy gain/loss statuses. Copy numbers of ≤ 1.5 were considered to indicate deletions and ≥ 2.5 were considered to be amplifications. To infer recurrently amplified or deleted genomic regions, we re-implemented the GISTIC  algorithm using copy numbers in 1 kb windows as markers instead of SNP array probes. G-scores were calculated for genomic and gene-coding regions based on the frequency and amplitude of amplification or deletion of each gene. A significant CNV region was defined as having an amplification or deletion with G-score > 0.1, corresponding to a p-value threshold of 0.05 from permutation-derived null distribution.
Identification of significantly mutated genes
For the identification of significantly mutated genes, we applied a novel analytical methodology, mutation significance with covariates (MutSigCV) , to avoid the false-positive findings detected by the standard significance analysis method (MutSig1.0) . MutSigCV corrects for variation by using patient-specific mutation frequency and spectrum and gene-specific background mutation rates, incorporating expression levels and replication time. MutSigCV is freely available for non-commercial use .
Pathway enrichment analysis
We performed the pathway enrichment analysis using the Database for Annotation, Visualization and Integrated Discovery (DAVID) v6.7 by examining the distribution of the non-synonymously mutated genes identified within the KEGG database  as described previously . Significantly altered pathways were determined by p-values calculated based on hypergeometric distribution with Benjamini correction.
KYSE150 and TE1 ESCC cell lines that were tested and found to be free of mycoplasma contamination were used in this study. 293 T cells were used as a packaging cell line to produce virus. All cells were grown in DMEM/F12 media at 37 °C in 5 % CO2. For the functional analysis of FAM84B, the ESCC lines KYSE150 and TE-1 with high endogenous expression levels were used for knockdown experiments.
Knockdown of FAM84B
Knockdown experiments were performed in at least two ESCC lines with high endogenous FAM84B expression. Two independent shRNAs were cloned into the pLKO.1-puro vector (Addgene, Cambridge, MA, USA) as described previously . A non-specific targeting shRNA was also cloned into the pLKO.1-puro vector to be used as a scrambled control (SCR). shRNA knockdown efficiency was determined by western blot analysis for FAM84B proteins using an anti-FAM84B antibody (Proteintech, Chicago, USA). Relative expression was normalized to the β-actin expression level.
qPCR copy number validated analysis
The copy numbers of the genes of interest were assessed in frozen atypical hyperplasia samples using genomic qPCR (TaqMan, Applied Biosystems, Foster, CA, USA) in triplicate. Prevalidated primers for the relevant genes were obtained from Applied Biosystems (accession numbers Hs02758348_Cn for MYC, Hs00655850_Cn for FAM84B, Hs01654625_Cn for VEGFC, Hs05965356_Cn for FBXW7 and Hs00703603_Cn for FAT1). RNase P (RPPH1 gene; Life Technologies, Shanghai, China, 4403328) was used as a diploid control. Data were analyzed using the comparative (delta-Ct) Ct method. An inferred copy number of < 0.3 was considered to indicate a homozygous deletion.
Fluorescence in situ hybridization analysis
To evaluate the amplification of FAM84B, we performed FISH. Tumor and matched normal tissues of interesting ESCC cases were cut into pieces in PBS, swollen in 65 mmol/L KCl for 5 mins at 37 °C, followed by fixation in cold acetic acid/methanol for 5 mins at 4 °C, then dropped onto slides. For interphase FISH analysis, slides were stained with Cytocell enumeration probes against chromosome 2 or FAM84B conjugated with FITC or Cy3.5 (Rainbow Scientific, Windsor, CT, YSA). Probes against chromosome 2 were used as controls. 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 × 20 objective and DAPI, FITC and Rhodamine filter cubes. Images were produced using the AxioCamMRm CCD camera and Axiovision version 4.5 software suite. p-values were calculated using a two-sample test for equality of proportions with continuity correction.
Immunohistochemistry and tissue microarray resource
FAM84B was immunohistochemically stained with an anti-FAM84B antibody (Proteintech, Chicago, USA) as described previously . A human ESCC tissue array (OD-CT-DgEso01-001) purchased from Shanghai Outdo Biotech (Shanghai, China) was used to detect the expression level of FAM84B. OD-CT-DgEso01-001 array contains 10 normal esophagus tissues, 22 atypical hyperplasia tissues and 58 ESCC cases (2cores/case; Additional file 16: Table S9). FAM84B immunohistochemistries were performed on tissue microarraysas previously described , using the polyclonal anti-FAM84B antibody (Additional file 17: Figure S8). Briefly, sections were incubated with the specific antibody at a 1:500 dilution for 14 h at 4 °C, followed by detection using the PV8000 (Zhongshan, Beijing, China) and DAB detection kit (Maixin, Fuzhou, China), producing a dark brown precipitate. Slides were counterstained with hematoxylin. All images were captured at × 100. Cytoplasm expression of FAM84B was quantified using AperioCytoplasma 2.0 software. Statistic analyses were performed using Graphpad Prism 5.0.
Immunoblotting was performed as previously described  using anti-FAM84B (Proteintech, Chicago, USA). Antibody binding was detected using horseradish peroxidase-labeled anti-mouse (Sigma, Santa clara,USA) or anti-rabbit (Cell Signaling, Boston,USA) antibodies and chemiluminescence was measured using a LAS4000 device chemiluminescence System (Sagecreation, Beijing, China). Equal protein loading was confirmed with antibodies against β-actin (Transgen, Beijing, China).
MTT and migration/invasion assays
To assess cell viability, MTT assays were performed as previously described . Each experiment consisted of four replications and at least three independent experiments were carried out. Migration and invasion assays were performed in 16-well CIM plates in an xCELLigence RTCA DP system (ACEA Biosciences, San Diego, USA) using BD matrigel basement membrane matrix for real-time cell migration analysis as described previously [60, 61]. At least three independent experiments were carried out; for each independent experiment, five duplicates were performed for each group.
Experiments were done in triplicate and data were presented as mean ± SD. Student’s t-test was used for statistical analysis, and data from more than two groups were analyzed by one-way analysis of variance (ANOVA) in SPSS Statistics 19.0 followed by LSD-t test. Results were considered significant when p < 0.05.
Availability of supporting data
The data sets supporting the results of this article are available in the European Genome-Phenome Archive repository at Study accession EGAS00001001487 and Dataset accession EGAD0000100169. Tissue Microarray data and further details on data access are available from the GigaScience, GigaDB database .
We thank BGI for sequencing and analyzing data. This work was supported by the National Natural Science Foundation of China (81330063 & 81272189), the Key Project of Chinese Ministry of Education (NO213005A), Specialized Research Fund for the Doctoral Program of Higher Education (20121417110001), Research Project Supported by Shanxi Scholarship Council of China (2013–053 & 2015 key 3), the Innovative Team in Science & Technology of Shanxi (2013-23), the Program for the Outstanding Innovative Teams of Higher Learning Institutions of Shanxi(2015-313) to Y.P.C., and the National Natural Science Foundation of China (81272694 to X.L.C., 81201956 to J.L., 81402342 to L.Z.). The authors declare no competing financial interests.
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