Background
Colorectal cancer (CRC) belongs to one of the most extensively studied types of cancers due to its high mortality and severity. It is the third and second leading cause of death from malignant disease among adults in the US and Europe, respectively [
1]. A decrease in oxygen concentration is widely seen during the formation of many solid tumors, including CRC. Hypoxic regions may occur due to poorly formed vasculature, shunting of blood and vascular permeability [
2]. Cancer cells can adjust to this microenvironment by altering gene transcription to enhance glucose uptake and angiogenesis [
2]. The various adaptive responses involve multiple mechanisms, of which the best-characterized is mediated through transcriptional gene activation by the hypoxia inducible factor (HIF) [
3]. HIF is a heterodimeric transcription factor assembled from an oxygen-regulated α subunit (HIF-α) and a constitutively expressed β subunit (HIF- β) [
3,
4]. Under hypoxic conditions, HIF-α translocates into the nucleus, where it forms a dimer with HIF-β to form an active transcriptional complex with a number of cofactors [
3,
4]. The HIF complex binds to the promoter hypoxia response elements (HREs) to induce the expression of target genes that regulate the cellular adaptive response to low oxygen tension [
3,
4].
HIF-α is constitutively expressed in the tissue; however, it has an extremely short half-life in normoxic conditions [
3]. The level of HIF-α protein is regulated in several ways. The most well known is its degradation through post-translational hydroxylation. To date, two different oxygen-dependent hydroxylation mechanisms have been identified. The first pathway is initiated by three prolyl hydroxylase domain enzymes, PHD1, PHD2 and PHD3 [
3]. The second pathway involves the factor inhibiting HIF (FIH) [
5]. The PHD enzymes catalyze the hydroxylation of two conserved proline residues in the oxygen dependent degradation domain of the HIF-α protein. Hydroxylated proline residues are subsequently recognized by the E3 ligase complex containing von Hippel–Lindau tumour suppressor protein (pVHL), and targeted for degradation by the 26S proteasome [
3]. Similarly, FIH hydroxylates the asparagine residue within the C-terminal transactivation domain of HIF-α [
5,
6]. This results in the prevention of HIF-α interaction with its coactivators. Hence, under normoxic conditions, there is a dual mechanism of HIF inhibition by its degradation or inactivation by PHDs and FIH enzymes, respectively.
Recently, various studies have demonstrated inconsistent data of
FIH and
PHD1,
2 and
3 expression changes during CRC development [
7‐
10]. The mechanism by which these hydroxylases might be regulated is still not well elucidated. Interestingly,
PHDs and
FIH genes possess a CpG island within their promoter region. Similarly to genetic mutations, hyper- or hypomethylation of gene regulatory sequences have been shown to potentially change the expression of cancer related genes in different malignancies, including CRC [
11]. To date, it has been demonstrated that the promoter region of the
PHD3 gene is hypermethylated in plasma cell neoplasia, prostate, melanoma and mammary gland cancer cell lines [
12,
13]. The DNA methylation status of
PHD1,
PHD2 and
FIH has also been investigated in breast, cervical and prostate cancer cell lines, but the results are inconsistent [
12,
14,
15]. These reports prompted us to study whether altered PHD1, PHD2, PHD3 and FIH expression levels may be correlated with the DNA methylation status of their promoter regions in primary cancerous and histopathologically unchanged colorectal tissue from the same ninety patients. We also evaluated the effect of 5-Aza-2’-deoxycytidine (5-dAzaC), an inhibitor of DNA methyltransferases (DNMTs), on the DNA methylation level of the
PHD3 gene and its effect on PHD3 transcript and protein levels in HCT116 and DLD-1 CRC cells under hypoxic and normoxic conditions.
Methods
Antibodies and reagents
Rabbit polyclonal (Rp) anti-PHD1 (NB100-310), -PHD2 (NB100-137), -PHD3 (NB100-139) and -FIH (NB100-428) antibodies (Ab) were provided by Novus Biologicals (Cambridge, UK). Rp anti-GAPDH Ab (FL-335) and goat anti-rabbit horseradish peroxidase (HRP)-conjugated Ab were provided by Santa Cruz Biotechnology (Santa Cruz, CA). 5-dAzaC was purchased from Sigma-Aldrich Co. (St. Louis, MO).
Patient material
Primary colonic adenocarcinoma tissues were collected between June 2009 and July 2012 from ninety patients who underwent radical surgical resection of the colon at the Department of General and Colorectal Surgery, Poznań University of Medical Sciences, Poland (Table
1). Histopathologically unchanged colonic mucosa located at least 10–20 cm away from the cancerous lesions was obtained from the same patients. Since
ex vivo stress may influence protein stability, one set of samples was immediately snap-frozen in liquid nitrogen and stored at -80ºC until RNA/DNA/protein isolation [
16]. Another set of samples was directed for histopathological examination. Histopathological classification including stage, grade and tumour type was performed by an experienced pathologist. No patients received preoperative chemo- or radiotherapy. Written informed consent was obtained from all participating individuals. The procedures of the study were approved by the Local Ethical Committee of Poznań University of Medical Sciences.
Table 1
Demographic and histopathological classification including stage, grade and tumour type of patients with CRC
Total no. of patients | 90 |
Gender (Female/Male) | 41/49 |
Mean (± SD) age at radical surgical resection of colon (yrs) | 68.60 ± 11.45 |
CRC localization | |
Proximal colon (cecum to transverse) | 32 |
Distal colon (splenic flexure to sigmoid) | 18 |
Rectum | 40 |
Histological grade | |
G1 | 3 |
G2 | 65 |
G3 | 22 |
Dukes classification | |
A | 8 |
B | 35 |
C | 47 |
Tumour stage | |
T1 | 4 |
T2 | 10 |
T3 | 65 |
T4 | 11 |
Cell culture
DLD-1 colon cancer cells were obtained from the American Type Culture Collection (Rockville, MD) and HCT116 cells were kindly provided by the Department of Experimental and Clinical Radiobiology, Maria Skłodowska-Curie Cancer Center, Institute of Oncology Branch, Gliwice, Poland. These cells were cultured in DMEM GibcoBRL (Grand Island, NY) containing 10% heat-inactivated fetal bovine serum (FBS) and 2 mM glutamine. To determine the effect of 5-dAzaC on DNA methylation, transcript and protein levels of selected genes, the HCT116 and DLD-1 cells were cultured for 24 hours in DMEM GibcoBRL (Grand Island, NY) supplemented with 10% FBS from Sigma-Aldrich Co. (St. Louis, MO). Cells were then cultured under normoxic or hypoxic (1% O2) conditions either in the absence or in the presence of 5-dAzaC at a concentration of 1.00 or 5.00 μM for 6, 24 and 48 hours. Hypoxic conditions were achieved using a MCO-18 M multigas cell culture incubator, Sanyo (Wood Dale, IL), modified to permit flushing the chamber with a humidified mixture of 5% CO2, 94% N2. These cells were used for total DNA, RNA isolation, RQ-PCR, western blotting, and HRM analysis.
Reverse transcription and real-time quantitative polymerase chain reaction (RQ-PCR) analysis
Total RNA from primary tissues of patients with CRC and CRC cell lines was isolated according to the method of Chomczyński and Sacchi (1987) [
17]. RNA samples were quantified and reverse-transcribed into cDNA. RQ-PCR was carried out in a Light Cycler®480 Real-Time PCR System, Roche Diagnostics GmbH (Mannheim, Germany) using SYBR® Green I as detection dye. The target cDNA was quantified by the relative quantification method using a calibrator for primary tissue or respective controls for HCT116 and DLD-1 cells. The calibrator was prepared as a cDNA mix from all of the patients’ samples and successive dilutions were used to create a standard curve as described in Relative Quantification Manual Roche Diagnostics GmbH, (Mannheim, Germany). For amplification, 1 μl of total (20 μl) cDNA solution was added to 9 μl of IQ™ SYBR® Green Supermix, Bio-Rad Laboratories Inc. (Hercules, CA) with primers (Additional file
1). To prevent amplification of sequences from genomic DNA contamination, primers and/or amplicons were designed at exon/exon boundaries and covered all gene splice variants (Additional file
1). The quantity of PHD1, PHD2, PHD3 and FIH transcript in each sample was standardized by the geometric mean of two internal controls. The internal control genes were
porphobilinogen deaminase (
PBGD) and
human mitochondrial ribosomal protein L19 (
hMRPL19) (Additional file
1). They were selected from four candidate reference genes (
PBGD,
hMRPL,
peptidylprolyl isomerase A-
PPIA,
hypoxanthine phosphoribosyltransferase 1- HPRT) based on the results achieved in geNorm VBA applet for Microsoft Excel (data not shown) [
18,
19]. The PHD1, PHD2, PHD3 and FIH transcript levels in the patients’ tissues were expressed as multiplicity of cDNA concentrations in the calibrator. In HCT116 and DLD-1 cells, transcript levels were presented as multiplicity of the respective controls.
Western blotting analysis
Primary tissues from patients with CRC, HCT116 and DLD-1 cells were treated with lysis RIPA buffer and proteins were resuspended in sample buffer and separated on 10% Tris-glycine gel using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Gel proteins were transferred to a nitrocellulose membrane, which was blocked with 5% milk in Tris/HCl saline/Tween buffer. Immunodetection of bands was performed with Rp anti- PHD1, -PHD2, -PHD3 and -FIH Ab, followed by incubation with goat anti-rabbit HRP-conjugated Ab. To ensure equal protein loading of the lanes, the membrane was stripped and incubated with Rp anti-GAPDH Ab (FL-335), followed by incubation with goat anti-rabbit HRP-conjugated Ab. Bands were revealed using SuperSignal West Femto Chemiluminescent Substrate, Thermo Fisher Scientific (Rockford, IL) and Biospectrum® Imaging System 500, UVP Ltd. (Upland, CA). The amounts of analyzed proteins were presented as the protein-to-GAPDH band optical density ratio. For HCT116 and DLD-1 cells cultured in the absence of 5-dAzaC, the ratio of PHD3 to GAPDH was assumed to be 1.
DNA isolation and bisulfite modification
Genomic DNA was isolated using DNA Mammalian Genomic Purification Kit purchased from Sigma-Aldrich Co. (St. Louis, MO). 500 ng of genomic DNA was subjected to bisulfite conversion of cytosine to uracil according to the EZ DNA Methylation Kit™ procedure from Zymo Research Corporation (Orange, CA). The position of CpG islands and binding sites of transcription factors located in the regulatory region of the promoter was determined by online programs [
20‐
22].
DNA methylation evaluation by bisulfite sequencing
DNA fragments containing CpG dinucleotides located in the promoter region of the
PHD1,
PHD2,
PHD3 and
FIH genes were amplified from the bisulfite-modified DNA by the primer pairs (Additional file
1, Additional file
2) complementary to the bisulfite-DNA modified sequence. PCR amplification was performed by FastStart Taq DNA Polymerase from Roche Diagnostic GmbH (Mannheim, Germany). The PCR products were purified using Agarose Gel DNA Extraction Kit, Roche Diagnostic GmbH (Mannheim, Germany) with subsequent cloning into pGEM-T Easy Vector System I, Promega (Madison, WI) and transformation into TOPO10
E. coli strain cells. Plasmid DNA isolated from five positive bacterial clones was used for commercial sequencing of the cloned fragment of DNA. The results of bisulfite sequencing were assessed and presented using BiQ analyzer software and Bisulfite sequencing Data Presentation and Compilation (BDPC) web server, respectively [
23,
24].
DNA methylation assessment by high resolution melting (HRM) analysis
Methylation levels of DNA fragments located within the CpG island of the
PHD1,
PHD2,
PHD3 and
FIH genes (Additional file
2) were determined by Real Time PCR amplification of bisulfite treated DNA followed by HRM profile analysis by Light Cycler®480 Real-Time PCR System, Roche Diagnostics GmbH (Mannheim, Germany). For PCR amplification, 1 μl of the bisulfite treated DNA from patients, HCT116, DLD-1 cells, or standards, and primers (Additional file
1, Additional file
2) was added to 19 μl of 5 X Hot FIREPol EvaGreen HRM Mix, Solis BioDyne Co. (Tartu, Estonia). Standardized solutions of DNA methylation percentage were prepared by mixing methylated and non-methylated bisulfite treated DNA from Human Methylated/Non-methylated DNA Set, Zymo Research Corp. (Orange, CA) in different ratios. To determine the percentage of methylation, the HRM profiles of patient DNA PCR products were compared with HRM profiles of standard DNA PCR product [
25,
26]. HRM methylation analysis was performed using Light Cycler®480 Gene Scanning software, Roche Diagnostics GmbH (Mannheim, Germany). Each PCR amplification and HRM profile analysis was performed in triplicate. Using HRM analysis we were able to detect heterogenous methylation with equal sensitivity (Additional file
3). The methylation for each patient was presented as a percentage of methylation in amplified fragments located in the CpG island of
PHD1, PHD2, PHD3 and
FIH. Since low levels of methylation may not demonstrate significant biological effect and we are not able to quantify all CpG dinucleotides within the analyzed CpG island, the percentage results were divided into three groups: 0–1% methylation, 1–10% methylation and 10–100% methylation for statistical analysis [
27‐
30].
Statistical analysis
The normality of the observed patient data distribution was assessed by Shapiro-Wilk test, and unpaired, two-tailed t-test or U Mann–Whitney test was used to compare the mean values. The chi-square test was used to examine significance in DNA methylation. To evaluate the association between different ranges of DNA methylation (0–1% methylation, 1–10% methylation and 10–100% methylation) and the ratio of cancerous tissue PHD3 mRNA level to histopathologically unchanged PHD3 mRNA level, the non-parametric Kruskal-Wallis test was employed. Data groups for cell lines were assessed by ANOVA to evaluate if there was significance (P < 0.05) between the groups. For all experimental groups, which fulfilled the initial criterion, individual comparisons were performed by post hoc Tukey test with the assumption of two-tailed distribution. Statistically significant results were indicated by p < 0.05. Statistical analysis was performed with STATISTICA 6.0 software.
Discussion
The maintenance of oxygen homeostasis is a crucial physiological process [
32]. Hypoxia is a common feature of human cancers, associated with higher aggressiveness and resistance to chemotherapy [
33,
34]. The cellular environment response to hypoxia is mediated by HIF, a heterodimeric transcriptional complex, which is post-translationally regulated by prolyl and asparaginyl hydroxylases. The first group, PHD enzymes, govern the ubiquitin- mediated mechanism of HIF-α degradation under normoxic conditions [
3]. Since hydroxylation reactions require the presence of oxygen, dioxygenases are unable to function in hypoxic conditions. Thus, the HIF is not directed to proteasomal degradation and may exert its effect on genes during CRC progression. However, oxygen concentration is not the only cause of altered PHD function during tumorigenesis. The contribution of PHD to cell behaviour depends on several conditions, including the relative abundance of PHD proteins in a specific tissue [
3]. Since changes in the amount of available PHD enzymes will alter the rate of hydroxylation of HIF-α, research has focused on the regulation of the quantity of PHDs. Aberrant expression of PHDs has been observed in a variety of human cancers. Detailed analysis of the expression patterns of all three PHD isoforms has shown differences depending on cancer cell type [
7,
35‐
37].
PHD1 protein levels were elevated in non-small cell type lung cancer (NSCLC) and pancreatic endocrine tumors [
36,
38]. Moreover, PHD1 inactivation resulted in lower levels of Cyclin D1 and impaired breast tumour formation [
39]. However, it has been demonstrated that colon cancer cells expressing the
PHD1 gene cause inhibition of tumour growth and angiogenesis under hypoxic conditions when injected into nude mice [
40].
PHD2 was also previously studied in various cancer types. Low levels of
PHD2 expression were correlated with poor survival rate in CRC and breast cancer [
9,
41]. Nonetheless, studies in head and neck squamous cell carcinoma (HNSCC) demonstrated an association between increased levels of nuclear PHD2 protein with strongly proliferative and malignant tumour phenotypes [
42,
43]. Furthermore, immunohistochemical studies have found that high
PHD2 and
PHD3 expression was significantly associated with worse 5-year survival rate in pancreaticobiliary cancer [
44]. PHD3 transcript levels were shown to be increased in HNSCC, and
in vitro analysis revealed inhibition of cell cycle progression in cancerous cells in the absence of PHD3 activity [
45]. Elevated levels of PHD3 protein in cancerous tissue were also observed in immunohistochemical studies of NSCLC and gastric cancer [
38,
46]. However, PHD3 catalytic activity has demonstrated an ability to induce apoptosis independent of HIF in different cancer cell lines [
12,
47‐
49]. Additionally, Peurala et al. presented that increased PHD3 expression and was associated with an increased survival rate in breast cancer patients [
41].
Discrepancies between
PHD1,
PHD2 and
PHD3 experimental results might be partly explained by the demonstration that HIF and PHDs can form a feedback loop that limits hypoxic signaling in reoxygenated cells [
50]. Moreover, these discrepancies imply a dual function of PHDs in the control of tumour progression that depends on cell type, other PHDs- interacting factors, and function independent of HIF. The HIF- independent functions of PHDs include PHD3 directed inhibition of phosphorylation of the inhibitor of κβ kinase β and activation of NF κβ in CRC, neuronal apoptosis and myoblast differentiation by PHD3, or control of myocardial development by PHD2 [
8,
51,
52].
We found a statistically significant decrease in the expression levels of the
PHD1,
PHD2 and
PHD3 genes in cancerous tissue compared to histopathologically unchanged colorectal tissue. Although we did not observe statistically significant differences in FIH transcript levels between cancerous and histopathologically unchanged colorectal tissue, we found a significant increase of FIH protein in CRC. FIH, in addition to PHDs, hydroxylates HIF-α at a conserved asparagine residue. Through this modification, FIH prevents interaction of HIF with its transcriptional coactivators [
5]. More importantly, FIH is able to suppress HIF activity under severe hypoxic conditions, where PHDs are inactive [
53,
54]. Additionally, FIH exerts HIF-independent functions by interaction with pVHL, histone deacetylases, p105, Notch 1, and SOCS box proteins [
55‐
58]. FIH is widely expressed; however, its expression varies between tissue and cancer types [
7]. To date, FIH protein overexpression was correlated with a more malignant phenotype and poor prognosis in pancreatic endocrine tumours and NSCLC, respectively [
36,
38]. Moreover, it has been established that
FIH gene silencing reduced colon cancer cell proliferation
in vitro and
in vivo through the p53-p21 mediated pathway [
54]. We observed a significant increase in the protein level of FIH in CRC tissue. This observation may result from E3 ligase activity of a member of seven in absentia homolog 1 (SIAH-1). Therefore, SIAH-1 facilitates ubiquitination and proteasomal degradation of FIH [
59,
60]. To date, SIAH-1 has been found to be widely distributed in human cell lines and tissues, including CRC, with a decreased expression in breast and hepatocellular cancer [
61‐
63].
Since the prolyl and asparaginyl hydroxylases have so wide and profound an effect on tumorigenesis, studies on the regulation of the expression of these enzymes may help in our understanding of CRC progression. There are many factors involved in the development and occurrence of CRC, and they are classified as genetic, epigenetic and environmental [
64‐
67]. One of the epigenetic mechanisms, namely altered DNA methylation in a gene’s regulatory region, is observed from the early stages of lesions in aberrant crypt foci and hyperplastic polyps [
68,
69]. It has been shown that DNA hypermethylation of
MLH1,
MGMT,
CDKN2A and many others is associated with CRC progression [
67,
70‐
72]. Our study is the first to investigate the methylation status of the promoter regulatory regions of
PHD1,
PHD2,
PHD3 and
FIH in primary cancerous tissue from patients with CRC, and HCT116, DLD-1 CRC cell lines. We did not observe DNA methylation within the CpG island of the
PHD1,
PHD2 and
FIH gene promoter in either patients or cell lines. To date, the DNA methylation status of the promoter region of
PHD1,
PHD2,
PHD3 and
FIH was studied in a few cancers, including breast, prostate, cervical, melanoma, kidney and plasma cell neoplasia [
12‐
15]. In cervical cancer cell lines the hypomethylation of the CpG island of the
PHD2 gene promoter was associated with an increase in
PHD2 expression [
15]. Similarly to our results, no DNA methylation within the CpG island of
PHD1,
PHD2 and
FIH has been observed in breast and plasma cell neoplasia [
13,
14]. There was also no DNA methylation in the promoter region of the
PHD3 gene in clinical samples from breast and prostate cancer [
12,
14]. We also observed no DNA methylation within the
PHD3 gene using bisulfite sequencing in region chr14: 34 419 929–34 420 563 and HRM analysis in region chr14: 34 419 922–34 420 080 in a group of patients, which is consistent with the results of Huang et al. and Place et al. [
12,
14]. However, we found a significantly higher level of DNA methylation within the first exon and intron of the
PHD3 gene (chr14: 34 419 346–34 419 943 using bisulfite sequencing; chr14: 34 419 795–34 419 935 and chr14: 34 419 400–34 419 538 using HRM analysis) in cancerous tissue compared to histopathologically unchanged tissue. To the best of our knowledge, DNA methylation within the chr14: 34 419 346–34 419 943, chr14: 34 419 795–34 419 935 and chr14: 34 419 400–34 419 538 regions was not previously analyzed in other studies. Data from Encyclopedia of DNA elements project showed that these regions are DNase I hypersensitive and able to bind various transcription factors, which suggests a promoter or enhancer activity [
22]. Moreover, Pescador et al. (2005) identified a functional HRE within the first intron of the
PHD3 gene and proposed a model of HIF-mediated hypoxic induction of PHD3 [
73].
Since hypoxic conditions may induce global DNA hypomethylation in cancer cells, we investigated DNA methylation and expression levels of the
PHD3 gene in HCT116 and DLD-1 cells under hypoxic and normoxic conditions [
74]. We reported a high level of DNA methylation and no transcript and protein level changes under hypoxic and normoxic conditions in HCT116 cells. In the
PHD3 gene promoter region in DLD-1 cells, we did not detect DNA methylation under either experimental condition, but we observed a significant induction of PHD3 transcript and protein level upon hypoxia. The
PHD3 gene possesses in its promoter region a putative HRE and can be induced by HIF transcription factor complex under hypoxic conditions [
35]. A lack of increase in
PHD3 expression in HCT116 cells might be the result of DNA methylation of its promoter region in hypoxic conditions. To date, a decreased expression of PHD3 mRNA was correlated with high CpG island methylation status in plasma cell neoplasia and selected melanoma, prostate and mammary gland cancer cell lines [
12,
13].
In order to verify the role of DNA methylation within the CpG island of the
PHD3 gene, we treated HCT116 and DLD-1 cells with 5-dAzaC under normoxic and hypoxic conditions. 5-dAzaC was previously shown to induce the expression of many genes in different types of cancer and inhibit the growth of CRC cells [
75,
76]. We observed significant DNA demethylation in the chr14: 34 419 922–34 420 080, chr14: 34 419 795–34 419 935 and chr14: 34 419 400–34 419 538 regions of the CpG island of the
PHD3 gene in HCT116 cells incubated with 5-dAzaC, which was correlated with an increase in PHD3 transcript and protein levels. The same regions were unmethylated in DLD-1 cells at different experimental conditions and 5-dAzaC did not affect
PHD3 gene expression.
The presence of DNA hypermethylation of the
PHD3 promoter region in a broad range of human cancers suggests its role in tumour survival. In glioblastoma cell lines, accumulation of PHD3 protected tumour cells against hypoxia-induced cell death via control of HIF [
77]. However, reduction of
PHD3 expression by DNA methylation may allow for stable HIF directed cellular response during hypoxia [
12]. Moreover, in a subset of breast, prostate, melanoma and renal carcinoma cell lines, HIF-1α accumulation during hypoxia was independent of DNA hypermethylation of the
PHD3 promoter region, which suggests its role in other pathways and hydroxylase independent function [
12].
Conclusion
Our results showed increased DNA methylation levels in the CpG island of PHD3 in CRC as compared to normal colonic epithelium from the same patients. These epigenetic alterations were associated with a significant decrease of PHD3 expression levels in patients with CRC. However, the reduction in PHD1 and PHD2 expression in cancerous tissue was not due to altered methylation within the CpG island in the promoter region of PHD1 and PHD2, respectively. Therefore, other mechanisms might be responsible for the observed decreased expression levels of PHD1 and PHD2 in CRC patients. Furthermore, we observed an increased level of FIH protein in CRC, with no changes in the FIH transcript level between cancerous and histopathologically unchanged colonic mucosa. Additionally, we observed different statistical values of transcript and protein levels of the analyzed genes for subgroups of patients classified according to different features. However, a larger group of patient and deeper molecular investigation of these observations should be studied to determine whether the results within subgroups may be clinically important.
We also found that an inhibitor of DNMTs, 5-dAzaC, induced DNA demethylation of the PHD3 promoter region, which was associated with increased transcript and protein levels in HCT116 cells under hypoxic and normoxic conditions.
Although we presented epigenetic transcriptional down-regulation of
PHD3 gene expression in CRC patients and HCT116 CRC cell line for the first time, further studies are required to verify and determine the role of CpG island methylation in
PHD3 expression in CRC to a greater extent. Moreover, DNA methylation is often associated with other alterations in chromatin structure, namely histone acetylation. High levels of DNA methylation accompanied with a low degree of histone acetylation may decrease the transcription of many genes [
78]. In consideration of this, we also need to determine the possible role of histone modifications in
PHD3 gene expression.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
AAR contributed to designing the study, acquisition of data, analysis and interpretation of data, and in writing the manuscript. KEB and KH participated in the acquisition and interpretation of the data. As Principal Investigators, PK and JPP were involved in the intellectual and experimental programming of the study, the interpretation of data, and writing the manuscript. All authors read and approved the final manuscript.