The contents of this work are solely the responsibility of the authors and do not necessarily represent the official views of the National Cancer Institute or Vanderbilt University.
The checkpoint with forkhead-associated domain and RING-finger domain (CHFR) is a mitotic checkpoint protein with tumor-suppressor functions. In this study, the authors investigated the epigenetic and genetic mechanisms that regulate CHFR expression in esophageal adenocarcinomas (EACs).
Quantitative reverse transcriptase polymerase chain reaction analysis demonstrated downregulation of CHFR transcript in 79% of EACs (44 of 56) compared with 41 normal samples (P < .001). Immunohistochemical analysis of CHFR protein expression showed absence or weak immunostaining for CHFR in 75% of EACs (56 of 75) compared with normal tissue samples. The authors next examined the promoter DNA hypermethylation of CHFR by using quantitative bisulfite pyrosequencing technology. They detected significant CHFR promoter DNA hypermethylation in 31% of tumor samples (18 of 58) compared with normal samples (P < .001). Treatment of OE33 cells with 5-Aza-deoxycytidine led to reduction in the promoter DNA methylation levels with restoration of the CHFR mRNA expression, which confirmed promoter DNA methylation as an epigenetic mechanism regulating CHFR expression. However, they identified several EACs where the CHFR mRNA expression was silenced in the absence of notable methylation. Therefore, the authors examined the relative DNA copy number level of CHFR compared with normal samples.
The results confirmed a decrease or absence of the relative CHFR DNA copy number levels in 59% of tumor samples. Nine tumors that showed loss of CHFR mRNA expression, in absence of promoter DNA hypermethylation, demonstrated a significant loss of relative CHFR DNA copy numbers.
The incidence of adenocarcinoma of the lower esophagus and gastroesophageal junction has increased at an alarming rate in the Western world in the last few decades.1, 2 Risk factors including obesity, alcohol consumption, smoking, and gastroesophageal reflux disease (GERD) have contributed to the increase in this cancer.3 Chronic GERD is the most common risk factor for the development of Barrett esophagus, whereby normal squamous mucosa is replaced by a metaplastic, specialized, columnar epithelium. Patients with Barrett esophagus have a high risk of progression to dysplasia and subsequent esophageal adenocarcinoma.4, 5
Although several molecular changes have been demonstrated in tumor initiation and progression,6-8 the contribution of epigenetic mechanisms in Barrett tumorigenesis is not well characterized. DNA methylation plays an important role in the regulation of gene expression. Aberrant DNA methylation (namely, overall DNA hypomethylation and regional DNA hypermethylation) have been linked to carcinogenesis of various organs.9 Aberrant DNA hypermethylation of CpG islands in the promoter region has been associated with gene silencing of several genes in cancer such as p16 (INK4A), hMLH1, and CDH1.10-12 Checkpoint with forkhead-associated domain and RING-finger domain (CHFR), located on chromosome 12q24, ubiquitously expressed in normal human tissues, was first identified by Scolnick and Halazonetis in 2000.13 CHFR is a mitotic checkpoint protein with a tumor-suppressor function that has the potential to be a novel biomarker for chemotherapeutic response to microtubule-targeting drugs.14 Under conditions of mitotic stress induced by taxol (paclitaxel) or nocodazole, the CHFR protein delays nuclear-envelope breakdown and chromosome condensation before cellular entry into metaphase.13
In this study, we examine the epigenetic and genetic mechanisms of silencing CHFR in esophageal adenocarcinomas. We used state-of-the-art, quantitative, bisulfite pyrosequencing technology (Biotage, Uppsala, Sweden) for analysis of promoter DNA methylation and quantitative reverse transcriptase polymerase chain reaction (qRT-PCR) for evaluation of relative DNA copy numbers of CHFR.
MATERIALS AND METHODS
All tissue samples were obtained from the archives of pathology at Vanderbilt University (Nashville, Tenn), Otto-von-Guericke Magdeburg University (Magdeburg, Germany), and from the National Cancer Institute Cooperative Human Tissue Network. The use of specimens from the archival tissue repository was approved by our institutional review board (protocol numbers 03-1078 and 33-2001). All tissue samples included in this study were collected and coded from tissues that remained after completion of diagnosis and that were otherwise discarded. All personal identifiers were removed before we received samples. Histopathological diagnosis of the EACs was verified on the basis of hematoxylin and eosin-stained sections according to the Vienna classification of gastrointestinal epithelial neoplasia.15 All tissue samples were dissected to obtain ≥70% cell purity. The patients' ages ranged from 34 to 84 years (median age, 65 years). The adenocarcinomas ranged from well-differentiated to poorly differentiated, stages I-IV, with a mix of intestinal-type and diffuse-type tumors.
qRT-PCR Analysis of CHFR Gene Expression
Total ribonucleic acid (RNA) was isolated by using the RNeasy Mini Kit (Qiagen, Valencia, Calif), and single-stranded cDNA was subsequently synthesized by using the Advantage RT-for-PCR Kit (Clontech, Palo Alto, Calif). The expression of CHFR was evaluated in a set of 97 frozen primary human samples that included 41 normal mucosa of the esophagus and the stomach and 56 samples of EACs and GEJs. Seventeen tumor samples had a matching tumor and normal sample from the same patient. The CHFR oligos were designed by using the online software, Primer 3 (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi). The oligos were purchased from Integrated DNA Technologies (Coralville, Iowa; CHFR [forward] 5′-CGTAACATCCCGTCCTGACT-3′ and [reverse] 5′-GCTCTCTTCACCTCCAGTGC-3′). The qRT-PCR was performed by using an iCycler (Bio-Rad, Hercules, Calif), with the threshold cycle number determined by use of iCycler software version 3.0. Reactions were performed in triplicate, and the threshold numbers were averaged. The results of the CHFR gene were normalized to HPRT, which had minimal variation in all normal and tumor samples tested.16 Expression fold was calculated16 according to the formula 2(Rt–Et)/2(Rn–En), where Rt is the threshold cycle number for the reference gene (HPRT) observed in the tumor, Et is the threshold cycle number for the experimental gene (CHFR) observed in the normal, Rn is the threshold cycle number for the reference gene observed in the normal samples, and En is the threshold cycle number for the reference gene observed in the tumor. Rn and En values were calculated as an average of the 41 normal samples. For all the primary EACs, the gene was considered to be downregulated when the messenger ribonucleic acid (mRNA) expression fold was ≤0.5 in comparison with the normal samples.
DNA Bisulfite Treatment and Pyrosequencing Analysis
DNA was purified by using a DNeasy Tissue Kit (Qiagen, Valencia, Calif). The bisulfite modification of the DNA from cell lines and tissues was performed by using an EZ DNA Methylation-Gold Kit (Zymo Research, Orange, Calif), according to the manufacturer's protocol. The CHFR promoter CpG island was identified by using a CpG island online search tool (http://www.uscnorris.com/cpgislands2.cpg.aspx). The criteria used for the definition of CpG islands were as follows: a DNA fragment ≥500 base pairs (bp) with a G + C≥55% with an observed ratio of CpG to expected CpG of 0.65. A 20 ng aliquot of modified DNA was subjected to polymerase chain reaction (PCR) amplification of the specific promoter region containing a CpG island that extended from −46 to −143 bp from the transcription start site and contained 15 CpG sites. The primers were designed by using PSQ Assay Design Software (Biotage, Uppsala, Sweden), where 1 of the primers was biotin labeled. The forward primer sequence was GAAGTAGTTTGGTTAGGATTAAAGAT, the reverse biotin labeled primer sequence was Bio-ACATTACCACTCCCTCAACTAAT, and the sequence primer was GTTTGGTTAGGATTAAAGAT. The Platinum PCR SuperMix High Fidelity Enzyme Mix (Invitrogen, Carlsbad, Calif) was used in the PCR reactions. The PCR products were checked by gel electrophoresis to confirm the size of the product and rule out the formation of primer dimers. The specific PCR products were then subjected to quantitative pyrosequencing analysis by using a Biotage PyroMark MD System (Biotage, Uppsala, Sweden) following the protocol provided by the manufacturer. The results were analyzed by Pyro Q-CpG 1.0.9 software (Biotage). On the basis of the methylation levels in the normal samples, we used 20% methylation as a cutoff for identification of promoter DNA hypermethylation of CHFR in tumor samples. Statistical analysis was performed to detect significant changes in the frequencies of DNA methylation of CpG sites between tumor and normal samples.
5-Aza-2′ Deoxycytidine and Trichostatin-A Treatment
For validation of the role of promoter DNA hypermethylation in transcriptional regulation of CHFR in vitro, esophageal cancer cell line OE33 was used. OE33 cells were maintained in Dulbecco modified Eagle medium (DMEM), supplemented with 10% fetal bovine serum (FBS) and antibiotics (Invitrogen, Carlsbad, Calif). Cells were seeded at low density for 24 hours, and then treated with 5 μM 5-Aza-2′ deoxycytidine (Sigma-Aldrich, St. Louis, Mo) for 72 hours or 300 nM trichostatin A (Wako, Osaka, Japan) for 24 hours. Total RNA and DNA were isolated and purified by a Qiagen RNeasy kit and DNeasy tissue kit, as described above. DNA methylation levels of the CpG nucleotides of CHFR promoter were determined by pyrosequencing before and after treatments. The CHFR mRNA expression levels were determined by qRT-PCR, as described above.
Immunohistochemical (IHC) analysis of CHFR protein expression was performed on a tumor tissue microarray (TMA) that contained 75 adenocarcinomas. All adenocarcinomas were classified according to the recent guidelines of the International Union Against Cancer TNM classification system. All EACs originated from the lower esophagus or gastroesophageal junction corresponding to the adenocarcinoma of the esophagogastric junction type.17 Samples from adjacent normal esophageal squamous epithelia, Barrett esophagus, and dysplastic tissues were included when available. All tissue samples were histologically verified, and representative regions were selected for inclusion in the TMA. The adenocarcinoma samples ranged from well differentiated to poorly differentiated, stages I to IV, with a mix of intestinal-type and diffuse-type tumors. Tissue cores with a diameter of 0.5 mm were retrieved from the selected regions of the donor blocks and punched to the recipient block by using a manual tissue array instrument (Beecher Instruments, Silver Spring, Md). Each tissue sample was represented by 4 tissue cores on the TMA. Sections (5 μm) were transferred to polylysine-coated slides (SuperFrost Plus; Menzel-Glaser, Braunschweig, Germany) and incubated at 37°C for 2 hours. The resulting TMA was used for IHC analysis using a CHFR mouse monoclonal antibody (clone BC012072, dilution 1:25; Abcam, Cambridge, UK), and positive IHC reactions were revealed by using the EnVision+ Kit (Dako, Hamburg, Germany). Cores with no evidence of nuclear staining or only rare, scattered, positive cells, <3%, were recorded as negative. The overall intensity of staining was recorded for the core with the strongest intensity. IHC results were evaluated for intensity and frequency of cell staining. The intensity of staining was graded as 0 (negative), 1 (weak), 2 (moderate), and 3 (strong). The frequency was graded from 0 to 4 by the percentage of positive cells as follows: grade 0, <3%; grade 1, 3% to 25%; grade 2, 25% to 50%; grade 3, 50% to 75%; grade 4, >75%. The index score was the product of multiplication of the intensity and frequency grades, which was then classified into a 4-point scale: index score 0 = product of 0, index score 1 = products 1 and 2, index score 2 = products 3 and 4, index score 3 = products 6 through 12.
Measurement of Fold DNA Amplification by qPCR
For evaluation of relative DNA copy numbers, we performed the qPCR amplifications using iCycler (Bio-Rad, Hercules, Calif). PCR reactions were prepared in a total volume of 20 μL-containing template DNA (40 ng), with the threshold cycle number determined by use of iCycler software version 3.0. Primers were designed using the online software, Primer 3 (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi). The forward and reverse primers for CHFR genomic DNA were 5′-TGCAGATGCTGTTCCTTACG-3′ and 5′-ACACATCCTCCACGTGACAA-3′, respectively. The oligos were obtained from Integrated DNA Technologies (Coralville, Iowa). Reactions were performed in triplicate, and the threshold numbers were averaged. The results were normalized to β-Actin, which had minimal variation in all normal and tumor samples tested. Amplification fold was calculated in the same way as in quantification of qRT-PCR for mRNA expression, and the En values were calculated as an average of the 19 normal samples. For all the primary EACs, the gene was considered to have loss of DNA copy number when the relative amplification fold was ≤0.5 when compared with the normal samples.
The Student t test for paired samples, using GraphPad prism version 4.0 software (GraphPad Prism Software, La Jolla, Calif), was used to compare the DNA methylation level between normal and EACs. We also compared the mRNA expression fold between normal and tumor samples and between unmethylated and methylated EACs. The Student t test was used to compare 1) the differences of the DNA methylation level between normal and EACs, 2) the differences of mRNA expression fold between normal and EACs, and 3) the differences of mRNA expression fold between unmethylated and methylated EACs. In addition, we analyzed the association between DNA methylation and clinicopathological factors. The correlation between the DNA methylation level and mRNA expression fold was determined by Spearman rank correlation. The comparison of IHC scores among normal, EACs, and association between IHC score and clinicopathological factors was carried out by chi-square and Fisher exact tests. All P values were based on 2-tailed tests, and differences were considered statistically significant when the P value was ≤.05.
Frequent Silencing of CHFR Gene Expression in EACs
The qRT-PCR analysis revealed frequent downregulation of CHFR mRNA expression in tumor samples (44 of 56, 78.6%) compared with 41 normal samples (P < .0001; Fig. 1A). Further analysis of 34 paired tumor and normal samples demonstrated significant downregulation of CHFR mRNA expression in tumors compared with their corresponding normal samples (Fig. 1B). Of note, several EACs (16 of 56, 28%) showed complete silencing of CHFR mRNA expression as indicated by absence of detectable signal.
Promoter DNA Hypermethylation of CHFR Gene Correlates With Silencing of mRNA Expression
Quantitative analysis of CHFR promoter DNA methylation, using state-of-the-art pyrosequencing technology, indicated increased promoter DNA methylation levels of all tested CpG nucleotides sequences in tumor samples compared with normal samples. The DNA methylation changes were independent of the patients' ages. The representative results of the percentage of the average methylation for the CHFR gene promoter in EACs versus normal are shown in Figure 2A. The promoter DNA hypermethylation (≥20%) of the CHFR gene was detected in 31% (18 of 58) of the EACs. The difference of methylation of CHFR promoter in EACs versus normal tissues was statistically significant (P = .01). Analysis of DNA methylation in 15 tumor samples, when compared with their matching histologically normal samples from the same patients, demonstrated a significant increase in the level of DNA methylation in tumors compared with their corresponding normal tissue (P = .01); a representative summary is illustrated in Figure 2B. We next analyzed the promoter DNA methylation against mRNA expression levels in all samples. As shown in Figure 3A, samples with hypermethylation (≥20%) had significant downregulation for CHFR expression (P ≤ .05) compared with samples with absent or low-promoter DNA methylation level (≤20%). By using Spearman rank correlation for analysis, we found a significant inverse correlation between promoter DNA methylation and mRNA expression fold for CHFR (coefficient r = −0.48, P < .001; Fig. 3B). These results suggest that the hypermethylation of the CHFR promoter region is 1 of the factors involved in suppression of its mRNA expression in EACs.
To confirm the role of DNA methylation in silencing CHFR expression, we tested whether interference with the activities of DNA methyltransferases led to reactivation of CHFR in an esophageal adenocarcinoma cell line, OE33. The OE33 cells express low levels of CHFR and their promoters CpGs are hypermethylated (80%). The 5-Aza-2” deoxycytidine treatment but not dimethylsulfoxide or trichostatin-A led to reduction in the DNA methylation level from 80% to 45% and a 13-fold increase in mRNA expression (Fig. 3C).
Loss of DNA Copy Numbers Interplays With DNA Hypermethylation for Silencing CHFR mRNA Expression in EACs
Although promoter DNA hypermethylation correlated statistically with low gene-expression levels, we detected silencing of mRNA CHFR expression in 78.6% (44 of 56) of cases, whereas promoter DNA hypermethylation was seen in only 21.4% (12 of 56) of EACs. These findings prompted us to find out whether loss of copy numbers could be a contributing factor in silencing CHFR expression. Evaluation of relative DNA copy numbers in 27 tumor samples was compared with the average of 11 histologically normal esophageal samples (adjacent to tumor areas) and 8 blood samples from normal individuals (total n = 19). There was no significant difference in DNA copy numbers between the normal esophagus and blood samples (data not shown). As shown in Figure 4, there was a loss of DNA copy numbers (≤0.5 fold) in 16 of 27 (59%) of the analyzed tumors. Seven cases showed a combination of promoter DNA hypermethylation and loss of DNA copy numbers, whereas 9 cases showed loss of DNA copy numbers in the absence of promoter DNA hypermethylation (Table 1). This interplay of epigenetic and genetic mechanisms for silencing CHFR suggests that silencing of CHFR is a crucial alteration for EACs.
Table 1. Genetic and Epigenetic Silencing of CHFR
Fold Relative DNA copy Numbers
Fold Relative mRNA Expression
ID indicates identification; mRNA, messenger ribonucleic acid.
The upper section demonstrates cases that had mRNA down-regulation of CHFR in absence of promoter DNA methylation (≤20%). The lower section shows a combination of DNA methylation (≥20%) and loss of DNA copy numbers.
Immunohistochemistry of CHFR Protein Expression in EACs
Because protein expression is the ultimate mediator of biological processes, we followed on our findings by using immunohistochemistry in primary tumor samples. The IHC analysis of CHFR protein expression in a tumor TMA demonstrated absent or weak nuclear immunostaining (score 0 and 1) for the CHFR protein in 75% (56 of 75) of the EACs (Fig. 5). In contrast, normal esophageal and gastric tissue samples demonstrated moderate to strong nuclear immunostaining (score 2 and 3) in all tested samples (Table 1; Fig. 5). Although the number of cases with premalignant lesions was small, we observed a moderate to strong immunostaining of CHFR in Barrett esophagus, whereas adjacent dysplasia and adenocarcinoma tissues displayed weak to absent immunostaining (Table 1; Fig. 5).
Because of the relatively small number, we did not have sufficient statistical power to analyze correlation with histopathological parameters.
In this study, we have shown that silencing of CHFR expression in esophageal adenocarcinomas is mediated by both epigenetic and genetic mechanisms. These findings indicate that cancer cells develop mechanisms that ensure loss of function of CHFR. In this context, our findings that CHFR expression is frequently lost in EACs further confirm the reports that show that loss of CHFR is critical for cellular transformation and carcinogenesis.14 Previous studies have demonstrated variable results for CHFR methylation (10% to 40%) in carcinomas of the colon, head and neck, lung, and stomach.18-22 None of these studies have investigated changes in copy numbers of CHFR.
Although promoter DNA hypermethylation has been described for several genes in EAC, such as APC, GPX3, GPX7, CDKN2A, DAPK, ID4, MGMT, RBP1, RUNX3, SFRP1, TIMP3, and TMEFF2,23-26CHFR has not been investigated in EAC. We have applied state-of-the-art pyrosequencing technology for quantitative analysis of promoter DNA methylation of CHFR. We have demonstrated a strong inverse correlation between the DNA methylation and gene expression levels. We have also shown that promoter DNA methylation of CHFR was significantly higher in EACs when compared with adjacent normal tissue from the same patient. By using an in vitro model for validation of the impact of DNA methylation on CHFR expression we have shown that treatments with 5-Aza-2′ deoxycytidine alone can restore the expression of CHFR with a reversal in the DNA methylation level. Taken together, our data confirm that aberrant hypermethylation of the 5′CpG island of the CHFR gene is closely associated with the transcriptional inactivation and may be involved in tumor development.
A striking finding was the presence of significant loss of expression of CHFR without a notable increase in promoter DNA methylation in several cases. Interestingly, CHFR is located in chromosome 12q24, a genomic area that often displays loss of DNA copy numbers in EAC.7, 27 As shown in this report, our analysis demonstrated a reduction in DNA copy numbers of CHFR in 59% of analyzed tumors; 33% (9 of 27) of these cases had mRNA downregulation of CHFR in absence of promoter DNA hypermethylation (<20%). However, 26% of the analyzed cases showed a combination of DNA hypermethylation and loss of DNA copy numbers, suggesting that the loss of CHFR expression could be due to a combined effect of the loss of 1 allele and methylation of the second allele. Alternatively, this could be explained by the existence of heterogeneous clones, where in some cells, the loss of CHFR expression was driven by promoter hypermethylation, whereas in other cells, this phenomenon was mediated by loss of gene copy numbers. The first explanation is more plausible, although single cell-based approaches are needed to confirm this hypothesis. An additional layer of gene-expression control includes regulation by miRNAs. Although we have not investigated the role of miRNA in regulating CHFR in this article, there are recent reports suggesting that miRNAs are involved in gene regulation and progression of EACs.28 Of note, the IHC results in our study have confirmed the significant reduction in CHFR expression in primary EACs.
Cell cycle progression is monitored by checkpoint mechanisms to ensure the integrity of the genome and the fidelity of sister chromatid separation. CHFR is an E3 ubiquitin ligase and an early mitotic checkpoint protein that is implicated in many cancers and in the maintenance of genomic stability.13, 29-31 Failure of such checkpoint functions results in genomic instability, a condition that predisposes cells to neoplastic transformation and tumor progression.20 Recent studies have shown that CHFR can interact with polo-like kinase 1 (PLK1), a key regulator of G2/M checkpoint.32, 33 Moreover, there are several studies indicating that CHFR is a negative regulator of the expression levels of Aurora kinase A (AURKA). AURKA encodes a centrosome-associated, cell cycle regulated, serine/threonine kinase34 that functions to establish mitotic spindles by regulating centrosome duplication and separation, as well as microtubule-kinetochore attachment, spindle checkpoint, and cytokinesis. Cytological analysis revealed that overexpression of AURKA results in centrosome amplification, cytokinesis failure, and aneuploidy.35 The tumor development and progression cascades are often characterized by a high incidence of DNA copy number variations and aneuploidy.27, 36-39 Indeed, we, and others, have shown that chromosomal instability and DNA copy number variations are common features of esophageal adenocarcinomas,7, 40-42 pointing out the disruption of key mechanisms that regulate mitosis and chromosomal segregation. In this context, CHFR, acting as a mitotic stress checkpoint gene, could be crucial to regulating mitosis and protecting against stress-induced mitotic errors.
Taken together, our data indicate interplay between epigenetic and genetic mechanisms for inactivation of CHFR, suggesting its involvement in the development and progression of esophageal adenocarcinomas.
CONFLICT OF INTEREST DISCLOSURES
This work was supported by the National Cancer Institute Grant CA106176 and R01CA131225 (W.E.R).