Skip Navigation


Carcinogenesis Advance Access originally published online on July 21, 2006
Carcinogenesis 2007 28(2):299-309; doi:10.1093/carcin/bgl133
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow All Versions of this Article:
28/2/299    most recent
bgl133v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (3)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Strathdee, G.
Right arrow Articles by Brown, R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Strathdee, G.
Right arrow Articles by Brown, R.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

HOXA5 is targeted by cell-type-specific CpG island methylation in normal cells and during the development of acute myeloid leukaemia

Gordon Strathdee1,*, Alyson Sim1, Richard Soutar3, Tessa L. Holyoake2 and Robert Brown1

1 Centre for Oncology and Applied Pharmacology, CR-UK Beatson Laboratories G61 1BD UK
2 Division of Cancer Sciences and Molecular Pathology, University of Glasgow Glasgow, G31 2ER UK
3 Department of Haematology, Western Infirmary Dumbarton Road, Glasgow, G11 6NT UK

*To whom correspondence should be addressed. Tel: +44 141 330 8707; Fax: +44 141 330 4127; Email: g.strathdee{at}beatson.gla.ac.uk


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
HOXA5 is a member of the HOX gene family, which is known to play key roles during embryonic development and in differentiation of adult cells. In addition, HOXA5 has been implicated as a tumour suppressor in breast cancer and shown to transactivate the p53 gene. CpG island methylation is a common mechanism of gene inactivation in tumour cells, but is rarely involved in control of cell-type-specific (CTS) expression in normal cells. However, here we demonstrate that HOXA5 is one of a small number of genes whose CTS expression pattern is controlled by CTS CpG island methylation in normal cells. Furthermore, chromatin immunoprecipitation analysis identified novel patterns of histone modifications associated with DNA methylation of HOXA5. High levels of methylation of histone residues (lysine 9 and 36 of histone H3) previously associated with transcriptional repression were present in the unmethylated, actively transcribing state, and were then reduced following DNA methylation and gene inactivation. Alterations to the normal patterns of HOXA5 gene methylation were also observed in tumour cells. Quantitative analysis of HOXA5 methylation identified the presence of limited methylation in all of the breast, lung and ovarian tumours examined. However, methylation levels in these three tumour types were nearly always low and comparable with that detected in the corresponding normal tissue. In contrast, acute myeloid leukaemia (AML) samples frequently (60% of samples) exhibited very high methylation levels, far greater than that seen in normal haematopoietic cells, suggesting a role for hypermethylation of HOXA5 in the development of AML, consistent with its previously identified role in haematopoietic differentiation.

Abbreviations: AML, acute myeloid leukaemia; CTS, cell-type-specific; IVM, In vitro methylated; RT–PCR, reverse transcription–polymerase chain reaction


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
DNA methylation is the only commonly occurring covalent modification of DNA and occurs almost exclusively at CpG dinucleotides (1). DNA methylation within the 5' end of genes, particularly of CpG islands (small stretches of DNA with high frequencies of CpG sites), has been clearly linked to repression of gene transcription (2). Although methylation can inhibit transcription directly by interfering with the binding of transcription factors (2), the main mechanism by which DNA methylation leads to gene inactivation is through associated changes in chromatin structure, particularly modification of histone tails, that leads to the production of compacted chromatin structures that are refractory to initiation of transcription (2). Methylation of CpG islands has been associated with a number of specific modification of histone tails, including reduced levels of histone acetylation as well as increased levels of methylation of particular histone residues, particularly K9 of histone H3 (3).

Methylation was originally proposed as a potential mechanism for control of tissue-specific expression (4). However, analysis of the methylation status of CpG islands in human adult somatic tissues determined that CpG islands were almost always methylation-free even in tissues where the associated gene was not expressed (1). However, interest in the role of DNA methylation in control of gene expression has been greatly revived following the discovery that inactivation of genes in a number of human diseases, particularly cancer, is often highly associated with increased methylation of promoter-associated CpG islands (5). In addition, Futscher et al. (6) recently reported the first example of a human gene, maspin, in which cell-type-specific (CTS) expression is associated with specific differences in methylation of its associated CpG island. The maspin gene is unmethylated and expressed in cells of epithelial origin, but the gene is not expressed, and its CpG island is methylated, in other cell types. Subsequently, we identified a second example, the MCJ gene, which is similarly controlled by DNA methylation in normal cells (7). In this case the gene is methylated and not expressed in epithelial cells and unmethylated and expressed in mesenchymal and haematopoietic cells (7).

HOXA5 is a member of a large protein family all of which contain a conserved DNA binding domain, known as the homeodomain (8). The human genome contains four clusters of HOX genes (A–D), which are known to play a key role in determining anterior–posterior patterning during development (8). HOX genes are also expressed in adult cells where they are thought to play an important role in governing the process of differentiation, and HOXA5 has been implicated in the differentiation of both haematopoietic and epithelial cells. Over-expression of HOXA5 in haematopoietic progenitors results in increased granulocytic/monocytic, but reduced erythroid/megakaryocytic, differentiation (9), whereas antisense-oligonucleotides directed against HOXA5 had the inverse consequences (10), suggesting a key role for HOXA5 in the regulation of myeloid differentiation. Other studies have also identified HOXA5 as a potential regulator of epithelial cell differentiation (11,12). In addition, HOXA5 has been implicated as a potential tumour suppressor gene in breast cancer and has been shown to function as a transactivator of the p53 gene (13) and to induce apoptosis by p53-dependent and p53-independent mechanisms (14). DNA methylation of the CpG island located in the 5' end of the HOXA5 gene has been identified in breast (13) and lung cancer (15) and has been associated with loss of HOXA5 expression (13).

In this report, we investigated the role of DNA methylation, and subsequent alterations in chromatin structure, in control of HOXA5 expression in normal cells, and the potential role of HOXA5 methylation in the development of solid tumours and leukaemia. The results identify HOXA5 as one of a small number of genes controlled by CTS DNA methylation in normal cells and identify novel patterns of histone modifications associated with the presence or absence of DNA methylation at this locus. Furthermore, the results suggest a potential role for loss of HOXA5 expression, due to promoter hypermethylation, in the arrest of normal differentiation seen in acute myeloid leukaemia (AML).


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 Supplementary material
 References
 
Tissue culture and cells
The primary, 142BR, and immortalized, 1BR.3.G, fibroblast cell lines and the immortalized breast epithelial cell line SVCT were obtained from ECACC (Cambridge, UK) and cultured as recommended. The ovarian carcinoma cell line A2780/MCP1 (16), breast carcinoma cell line MCF7, the fibrosarcoma cell line HT1080 and leukaemic cell lines were maintained in RPMI with 10% fetal calf serum in 95% air/5% CO2 at 37°C. RNA and DNA were extracted from cell pellets of the primary lines HEK78, WI-38 and MRC5 and immortalized human ovarian surface epithelial cells (17). Purified normal CD133+ cells were obtained from Cambrex Bioscience (Wokingham, UK). Genomic DNA was extracted for methylation analysis as described previously (18). 2'-deoxy-5-azacytidine (Sigma, Gillingham, UK) was dissolved in phosphate-buffered saline (PBS) and used to treat 1BR.3.G (0.2 µm) and LAMA84 (1 µm) cells. Cells were treated for 24 h at 37°C on Days 1 and 2 of a 7 day cycle, which was repeated a second time for both cell lines (resulting in a total of four 24 h treatments). At the end of the second cycle of treatment, cells were harvested for RNA and protein analysis (LAMA84) or chromatin immunoprecipitation (1BR.3.G) as described below. Mock treatments, using the same volume of PBS as added to the treated cells, were used for comparison.

Tissue samples
Ovarian and breast tumour samples were obtained from the Western Infirmary and Stobhill General hospitals, Glasgow, UK. Lung tumour samples were obtained from Western Infirmary, Glasgow, and Aberdeen Royal Infirmary, Aberdeen, UK. Genomic DNA derived from peripheral blood or bone marrow samples were obtained from patients with AML from the Western Infirmary, Glasgow, Royal Victoria Infirmary, Newcastle, and Glasgow Royal Infirmary, UK. Grossly normal breast, lung and ovarian tissue samples were obtained from patients undergoing surgery for tumour resection. Ethical approval for all samples collected had been obtained. All samples were stored frozen at –70°C. Peripheral blood samples were obtained from healthy volunteers. Genomic DNA was extracted for methylation analysis as described previously (18).

Isolation of specific haematopoietic cell types
Specific cell types were isolated from normal peripheral blood samples from healthy volunteers using magnetic beads (Dynal Biotech, Bromborough, UK) coated with CD3 (T-cell-specific), CD19 (B-cell-specific) and CD14-specific antibodies (monocyte-specific), as per the manufacturer's protocol. Purified cell pellets were used for extraction of genomic DNA (18), and total RNA using an RNeasy mini kit (Qiagen, Crawley, UK). The purity of the isolated cell population was confirmed using reverse transcription–polymerase chain reaction (RT–PCR) for CD3, 19 and 14 markers.

COBRA analysis
COBRA analysis was performed largely as described before (19). One microgram of genomic DNA was modified with sodium bisulphite using the CpGenome modification kit (Chemicon International, Temecula, CA, USA) as per the manufacturer's instructions. All samples were resuspended in 40 µl of TE, and 1 µl of this was used for subsequent PCR reactions. The samples were amplified in 25 µl volumes containing manufacturer's buffer, 1 unit of FastStart taq polymerase (Roche, Lewes, UK), 1–4 mM MgCl2, 10 mM dNTPs and 75 ng of each primer. PCR was performed with one cycle of 95°C for 6 min, 35 cycles of 95°C for 30 s, 61°C for 30 s and 72°C for 30 s, followed by one cycle of 72°C for 5 min. All PCR reactions were carried out on a PTC-225 DNA engine tetrad (MJ Research, Waltham, MA, USA). Following amplification the PCR products were digested with the appropriate restriction enzymes, specific for the methylated sequence after sodium bisulphite modification. Digestion of 11.5 µl total PCR products was carried out in 20 µl volumes with 15 U of the appropriate restriction enzyme [HinFI, RsaI, TaqI (Invitrogen, Renfrew, UK) or BsiEI (New England Biolabs, Hitchin, UK)] in manufacturer's buffer for 2 h. Digested PCR products were separated on 1.75% agarose gels and visualized by ethidium bromide staining on GeneSnap gel documentation system (Syngene, Cambridge, UK). Quantification of methylation levels was carried out by measurement of band intensities using the GeneTools system (Syngene), and methylation levels in a particular sample were calculated correcting for the smaller size (and thus lower ethidium bromide binding capacity) of the methylated band. In vitro methylated (IVM) DNA (Intergen) was used as a positive control for all COBRA assays. The primers used for the PCR reactions were 5'-GGGAAATGTAATAATTTTGTTATAATGGGTTG-3' for the forward primer and 5'-CTAAAACATATACTTAATTCCCTCCTAC-3' for the reverse primer.

Bisulphite sequencing
Sodium bisulphite modification and PCR were carried out as above. PCR products were then separated on 1.5% agarose gels and bands corresponding to the expected size were cut out and DNA was extracted using GenElute agarose spin columns (Sigma). Isolated DNA was resuspended in 10 µl of ddH2O and 3 µl was used for ligation to PGem-T vector system II PCR cloning vector (Promega, Southhampton, UK). Ligations were carried out overnight at 4°C and the ligated products were transformed into JM109 cells. These were then plated out on LB agar plates containing 100 µg/ml ampicillin, 80 µg/ml X-Gal and 0.5 mM IPTG and left overnight at 37°C. White (insert-containing) colonies were picked and grown up in 4 ml of LB containing 100 µg/ml ampicillin and plasmid DNA isolated. Sequencing was then carried out on a CEQ 2000XL DNA analysis system (Beckman Coulter, High Wycombe, UK) using the Sp6 sequencing primers.

Pyrosequencing
The samples were amplified in 50 µl volumes containing manufacturer's buffer, 2 µl bisulphite-modified DNA, 2 U of FastStart taq polymerase, 3 mM MgCl2, 50 mM dNTPs and 0.2 µm of each primer. PCR was performed with one cycle of 95°C for 6 min, 45 cycles of 95°C for 30 s, 58°C for 30 s and 72°C for 30 s, followed by one cycle of 72°C for 5 min. All PCR reactions were carried out on a PTC-225 DNA engine tetrad. Following amplification, sequencing was performed using a PSQ 96MA pyrosequencer (Biotage AB, Uppsala, Sweden), as per manufacturer's protocol. The primers used for the initial PCR were identical to those used for COBRA analysis, with the addition of a 5' biotin label on the reverse primer. The sequencing primer was 5'-GATGAGTTTTGTTTTTAG-3'.

RT–PCR
RNA was isolated from cell lines as above. Five micrograms of RNA was then used for cDNA synthesis using the SuperScript first-strand cDNA synthesis kit (Invitrogen). Subsequent PCR was performed as above, using 2–3 mM MgCl2 and 2 µl cDNA. PCR was performed with one cycle of 95°C for 6 min, 20–33 cycles of 95°C for 30 s, 61°C for 30 s and 72°C for 30 s, followed by one cycle of 72°C for 5 min. RT–PCR products were separated on 2% agarose gels, and quantification, relative to GAPDH, was carried out using Genegenius (Syngene). Quantification was assessed using a panel of 3-fold serial dilutions of a known positive cDNA (derived from the SVCT cell line, 1- to 243-fold dilution). For both primer sets cycling conditions were identified that gave reproducible differences in signal intensity for each threefold dilution in the panel. The primers used for HOXA5 were forward 5'-CCCACATCAGCAGCAGAGAG-3', reverse 5'-AGCTCCAGGGTCTGGTAGC-3'. Primers used for GAPDH were reported previously (7).

Chromatin immunoprecipitation
Chromatin immunoprecipitation assays were carried out using the Chromatin immunoprecipitation assay kit (Upstate, Charlottesville, VA, USA) as described in the manufacturer's instructions. Briefly, 1.5 x 106 cells were plated out in T75 tissue culture flasks and left overnight at 37°C. The following day the cells were cross-linked by adding 405 µl of 37% formaldehyde to 15 ml of culture medium and incubating for 10 min at 37°C. The cells were then scraped off the flasks and centrifuged in 1.5 ml centrifuge tubes for 4 min at 4000 r.p.m. The pellet was resuspended in 200 µl sodium dodecyl sulphate (SDS) lysis buffer (including protease inhibitors) and sonicated to produce fragments of 200–1000 bp. Before immunoprecipitation, 20 µl of each sample was removed and used for later quantification of input DNA. Immunoprecipitation was carried out using 10 µl (anti-acetyl-histone H3 antibody) or 8 µl (all other antibodies) overnight at 4°C. Following washing and elution of the chromatin complexes, the cross-links were reversed by incubating at 65°C for 4 h, and the DNA was recovered by phenol/chloroform extraction and ethanol precipitation. The samples were resuspended in 20 µl of TE, and 1 µl of this was used in subsequent PCR reactions to detect the presence of immunoprecipitated DNA. The samples were amplified in 25 µl volumes containing 1X manufacturer’s buffer, 1 U of FastStart taq polymerase, 0.75–3 mM MgCl2, 10 mM dNTPs and 75 ng of each primer. PCR was performed with one cycle of 95°C for 6 min, 32–35 cycles of 95°C for 30 s, 57–63°C for 30 s and 72°C for 30 s, followed by one cycle of 72°C for 5 min. Following amplification, the PCR products were separated on 2% agarose gels and quantified using the GelDoc 1000 system (Bio-Rad, Hemel Hempstead, UK). Quantification by each PCR was assessed using a panel of 2-fold serial dilutions of genomic DNA (25–0.2 ng). For each primer set cycling conditions were identified that gave reproducible differences in signal intensity for each 2-fold dilution in the panel. To correct for differences in input DNA, signals for each of the cell lines were normalized to an input sample removed before immunoprecipitation. The results presented for each ChIP assay are the average of two to three independent experiments. The primers used for the PCR reactions were for 14-3-3sigma region 1 ChIP forward 5'-GTGGGCAGCCATGTGATG-3', reverse 5'-CCGGCTCTGCAGTAAAGG-3' and for 14-3-3sigma Region 2 ChIP forward 5'-GATGAAGGGTGACTACTACC-3', reverse 5'-TCTCCTTCTTGCTGATGTCC-3'; for HOXA5 region 1 ChIP forward 5'-TCGGAAGCTGGGCGATGAG-3', reverse 5'-GTGCACTAATAGGGGAGTTGGG-3' and for HOXA5 Region 2 ChIP forward 5'-TGGCATGGATCTCAGCGTCG-3', reverse 5'-AGGAGAGCGTGGACGTG-3'; for Maspin region 1 ChIP forward 5'-TTGTGCCACCAACGTGTCTG-3', reverse 5'-ACGTACAGACATGCGTACGG-3' and for Maspin Region 2 ChIP forward 5'-CTTGAGTAGGAGAGGAGTG-3', reverse 5'-ACAAAGACCTGGATGTGGAG-3'; for MCJ region 1 ChIP forward 5'-GAGGTTTCACCATGTCGG-3', reverse 5'-GAGGCAATTTGCTTACTCCAC-3' and for MCJ Region 2 ChIP forward 5'-AACTAGTTTGTCCCTACGGC-3', reverse 5'-TACTCAGCGTAGCGCAAAC-3'; for MLH1 region 1 ChIP forward 5'-CCACCACCAAATAACGCTGG-3', reverse 5'-TGCGGCCTTTCTAACGTTGG-3' and for MLH1 Region 2 ChIP forward 5'-AAGGGCTACGACTTAACGG-3', reverse 5'-AAAAGGAGAAGGCCTGACTG-3'.

Quantitative PCR
Quantitative PCR analysis was performed in 20 µl volumes containing 1X master mix DyNAmo SYBR green qPCR kit (Finnzymes, Espoo, Finland), 75 ng of each primer and 1 µl DNA. PCR was performed with one cycle of 94°C for 15 min, followed by cycles of 94°C for 30 s, 57°C (region 1) or 60°C (region 2) for 30 s and 72°C for 30 s, with plate reads carried out at 87°C at the end of each cycle. Each of the PCR assays was run in triplicate. Reactions were carried out on an Opticon 2 DNA engine (MJ Research) and levels of product (normalized to input samples) were calculated using the manufacturer's provided software.

Western blotting
Total protein was extracted from cell pellets (1–2 x 106) using 480 µl protein lysis buffer (NP40, 0.5%; NaCl, 250 mM; Hepes, pH 7.0, 50 mM) to which 20 µl of protease inhibitor solution (mini Complete—EDTA-free, Roche, prepared as per manufacturer's instructions) was added. Samples were placed on ice for 30–45 min, centrifuged at 21 000 g for 15 min at 4°C and the supernatant was collected and stored at –20°C. Protein concentrations were calculated using Bio-Rad protein assay reagent (as per manufacturer’s protocol). For each sample 15 µg of protein was prepared and electrophoresed on 10% SDS–PAGE gels (MOPS buffer, as per manufacturer’s protocol, Invitrogen) and transferred to Immobilon-P membrane (Millipore, Watford, UK). Membranes were then sequentially probed with rabbit anti-HOXA5 (Zymed, Cambridge, UK) and secondary HRP-linked goat anti-rabbit (Santa Cruz Biotechnologies, Santa Cruz, CA) antibodies, and the immunological reaction was developed using chemilumnescence (ECL western blotting reagents, Amersham, Little Chalfont, UK). Subsequently, blots were stripped (30 min at room temperature in 0.2 M Glycine, pH 2.5/0.1% SDS), washed three times in PBS/0.2% Tween and re-probed with mouse anti-{alpha}-actin antibody (Sigma) as a control for protein loading.

Analysis of markers of myeloid differentiation in 2'-deoxy-5-azacytidine-treated cells
cDNA was synthesized from LAMA84 cells, with or without treatment with 2'-deoxy-5-azacytidine, as described above. qRT–PCR analysis was performed in 20 µl volumes containing 1X master mix DyNAmo SYBR green qPCR kit (Finnzymes), 75 ng of each primer and 1 µl cDNA. PCR was performed with one cycle of 94°C for 15 min, followed by cycles of 94°C for 30 s, 55–61°C for 30 s and 72°C for 30 s, with plate reads carried out at 77–87°C at the end of each cycle. Each of the PCR assays was run in triplicate. GAPDH was used as a control for normalizing relative expression levels in the different samples. Reactions were carried out on an Opticon 2 DNA engine (MJ Research). Expression levels of a panel of makers of myeloid differentiation (normalized to GAPDH) were calculated using the manufacturer’s provided software. Primers used were CD11b RT forward 5'-tacatggtggtcaccagcc-3', reverse 5'-ccatatgacagtctggttcag-3'; CD13 RT forward 5'-acgtcattgggcaaggtctg-3', reverse 5'-ttgaactgctccagctgctg-3'; CD14 RT forward 5'-gaagacttatcgaccatggag-3', reverse 5'-tgaggttcggagaagttgc-3'; C/EBP{alpha} RT forward 5'-atgccgggagaactctaactc-3', reverse 5'-agtgcgcgatctggaactg-3'; C/EBP{varepsilon} RT forward 5'-tctgcgcgttctcaaggc-3', reverse 5'-gcgatgttgttgcgctcc-3'.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 Supplementary material
 References
 
HOXA5 exhibits CTS CpG island methylation and expression in normal human cells
Previously, a small number of genes have been identified as exhibiting CTS patterns of DNA methylation and expression in normal cells (20), and methylation at the Hoxa5 locus had been identified in adult, but not embryonic, mouse tissue (21). To determine if the CpG island in the 5' region of the HOXA5 gene exhibited CTS methylation, COBRA assays were used to assess CpG island methylation in a panel of normal human cell lines. Multiple CpG sites (six, Figure 1A) within the island were assessed by digestion with different restriction enzymes (HinFI, TaqI, RsaI), and identical results were obtained for all CpG sites. This analysis detected hypermethylation (>95%) in adult mesenchymal cells, ~50% methylation of HOXA5 in haematopoietic cells, but no evidence of methylation in epithelial cells (Figure 1B). The 50% methylation detected in haematopoietic cells probably represents 50% of alleles methylated (and 50% unmethylated), as COBRA assays using restriction enzymes with multiple cut sites produced almost exclusively the indicated fully methylated or unmethylated bands (examples in Figure 1D), with little evidence of partially methylated bands. This was further confirmed using bisulphite sequencing of the region in haematopoietic cells, which identified exclusively unmethylated or densely methylated alleles (examples in Figure 1E). In contrast to the hypermethylation observed in adult-derived mesenchymal cells (1BR.3.G, 142BR), HOXA5 methylation was absent in embryonically derived human mesenchymal cells (WI38, MRC5). This is however in agreement with the lack of Hoxa5 methylation identified in whole mouse embryos (21).


Figure 1
View larger version (35K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 1 Methylation of the HOXA5 CpG island in normal cells. (A) Sequence of region of HOXA5 gene analysed for methylation. The eight CpG sites analysed for methylation by COBRA and pyrosequencing analysis are indicated in bold and labelled CpG 1–8. COBRA assays assessed methylation at CpG sites 5, 6 and 7 (RsaI digest), CpG site 8 (HinFI digest) and CpG sites 1, 4 and 8 (TaqI digest). Pyrosequencing was used to analyse methylation at CpG sites 2, 3 and 4. Numbering is relative to transcriptional start site (Genebank accession number NM_019102) and the location of the transcriptional start site is preceded by a +1. (B) Examples of COBRA analysis (HinFI digest) using a panel of normal and immortalized cells, including epithelial (SVCT, HEK78, OSE-C1, OSE-C2 and OSE-C3), haematopoietic [PBL (peripheral blood) 1 and PBL2] and mesenchymal cells [WI38, MRC5 (both embryonic), 1BR.3.G, 142BR (both adult)]. The lower digested bands (indicated by arrows) indicate the presence of DNA methylation and the upper undigested bands, the absence of methylation. IVM DNA (100% IVM) served as a positive control. (C) RT–PCR analysis of HOXA5 expression. The presence (+) or absence (–) of methylation is indicated for each cell line (PBL is 50% methylated) and the relative level of expression (compared with the SVCT cell line, and normalized to GAPDH expression) is indicated under each lane. The sloping black triangle represents successive 3-fold dilutions of SVCT cDNA used for quantification. (D) COBRA analysis of HOXA5 methylation in purified haematopoietic cell types (B lymphocytes, T lymphocytes and monocytes) and peripheral blood samples with specific cell types subtracted [PBL-B (B lymphocytes), T (T lymphocytes) or M (monocytes)]. The positions of the unmethylated and (digested) methylated bands are indicated by arrows. (E) Examples of bisulphite sequencing analysis of a portion of the HOXA5 locus containing a T/A polymorphism in a heterozygous peripheral blood sample. The large arrow indicates the position of the polymorphism. Small arrows indicate the position of CpG dinucleotides, which have either been retained following bisulphite modification (identifying the presence of methylation) or converted to CpA (identifying the absence of methylation). Both T and A polymorphisms are associated with both methylated and unmethylated alleles.

 
The presence of methylated HOXA5 alleles in normal cells was also confirmed using pyrosequencing (22). This technique was used to assess methylation of three CpG sites in the HOXA5 promoter (CpG 2, 3 and 4 in Figure 1A) and, as detailed in Table I, the methylation levels closely mirrored those identified by COBRA analysis, confirming the presence of very high levels of methylated HOXA5 alleles in adult mesenchymal cells and ~50% methylation in haematopoietic cells.


View this table:
[in this window]
[in a new window]

  		Table I HoxA5 promoter methylation identified by pyrosequencing

 
Semi-quantitative RT–PCR analysis was used to assay gene expression to determine if CTS methylation of HOXA5 correlated with loss of expression. This analysis identified low or undetectable levels of expression of HOXA5 in cells in which the gene was methylated, but in contrast, the absence of DNA methylation was associated with high levels of expression of the gene (Figure 1C).

Partial methylation of HOXA 5 in haematopoietic cells is not due to targeting of methylation to specific cell types or genetic imprinting
For genes previously identified as exhibiting CTS methylation, the methylated cell types invariably exhibited complete methylation (indistinguishable from 100% IVM DNA) [(6,7), (G. Strathdee, A. Sim and R. Brown), unpublished data]. In contrast to the >95% methylation in mesenchymal cells, HOXA5 methylation in peripheral blood was detected in only ~50% of HOXA5 alleles. As HOXA5 has been implicated as an important regulator of haematopoietic differentiation (9,10), this suggests a potential role for HOXA5 methylation in control of HOXA5 expression levels during haematopoietic differentiation. Peripheral blood samples contain multiple different haematopoietic cell types, and thus methylation may be limited to one or more specific cell types. To examine this possibility, B and T lymphocytes and monocytes were isolated from normal peripheral blood using cell surface markers. The isolation of highly purified cell populations was confirmed using RT–PCR for markers specific for each cell lineage (data not shown). COBRA analysis on the purified cell types identified methylation of ~50% of HOXA5 alleles for each of the cell types, and the levels were indistinguishable from either unpurified peripheral blood or peripheral blood samples depleted of each of the cell types (Figure 1D).

To determine if HOXA5 was imprinted in haematopoietic cells, an A/T polymorphism was identified within the HOXA5 gene by DNA sequencing of multiple peripheral blood samples (Figure 1E). Bisulphite sequencing analysis of this region was then carried out on a peripheral blood sample that was heterozygous for the polymorphism. This analysis identified the presence of both the A and T polymorphisms in methylated and non-methylated alleles, indicating that methylation was not specifically targeted to either allele (Figure 1E) and that HOXA5 is therefore not imprinted.

Analysis of chromatin modifications associated with CpG island methylation of genes in normal cells
Hypermethylation of genes during tumour development is known to be associated with specific patterns of modifications within histone tails, which are crucial in gene silencing (2,23). To determine if similar alterations were associated with CTS methylation of HOXA5 gene, ChIP assays were used to assess changes in histone modifications within the HOXA5 CpG island, both upstream (region 1) and downstream (region 2) of the transcriptional start site (Figure 2A). For comparison, the MLH1 gene was also analysed, as a typical example of a gene frequently hypermethylated in a cancer-specific fashion, which has been extensively characterized for histone modifications following DNA methylation (24,25). To allow quantification of the ChIP signals, cycling conditions were identified for each primer set that gave reproducible differences in product intensity between successive 2-fold dilutions of genomic DNA (as shown in Figure 2B). For each antibody/primer set the results presented are the averages of two to three independent experiments.


Figure 2
View larger version (37K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 2 Analysis of histone modifications using chromatin immunoprecipitation. (A) Diagrams of the CpG islands of HOXA5 and the other three known CTS methylated genes, and MLH1 (cancer-specific methylated gene). The position of the transcriptional start sites/first exons are indicated by the hatched boxes, and the small arrows indicate the positions of primer sets used for ChIP analysis (region 1 refers to primer sets upstream of the transcriptional start site and region 2 refers to primer sets downstream, for all loci). The small vertical lines represent CpG dinucleotides, and the positions of the identified CpG islands are indicated by thick horizontal lines. The diagrams were generated using the CpG island searcher (http://www.uscnorris.com/cpgislands/cpg.cgi). (B) Representative ChIP analysis using an antibody specific for K9, 14 acetylated histone H3 (region 1 for all loci). The descending black box represents a panel of serial 2-fold dilutions of genomic DNA used to allow quantification of the relative differences in signal intensity obtained from the cell lines. The methylation status, M (methylated), U (unmethylated), and the expression status, + (expressing), – (non-expressing), is indicated below each cell line. (C) Representative ChIP analysis using an antibody specific for dimethylated K36 of histone H3 (region2 for all loci) and (D) representative ChIP analysis using an antibody specific for dimethylated K9 of histone H3 (region 1 for all loci). DNA from immunoprecipitated chromatin was amplified with primers targeted to regions 1 and 2 of each of the indicated genes. The cell lines used and no antibody control (No Ab) are indicated above. The methylation status, M (methylated), U (unmethylated), and the expression status, + (expressing), – (non-expressing), is indicated below each cell line. Quantification of the relative differences in signal intensity obtained from the cell lines was performed using a panel of serial 2-fold dilutions of genomic DNA (as shown in B). Each ChIP analysis was performed at least twice.

 
Loss of histone acetylation has been linked to DNA methylation, as well as transcriptional inactivation in general (26). ChIP assays using an antibody to acetylated K9 and K14 of histone H3 confirmed that CpG island methylation of both HOXA5 and MLH1 was associated with clear reductions in histone acetylation (examples in Figure 2B, Table II). Methylation of both K9 and K36 of histone H3 within promoter regions has been linked to transcriptional repression (24,27–29). To examine methylation of these residues, ChIP assays were performed with separate antibodies directed against dimethyl K36, dimethyl K9 and trimethyl K9. As expected the ‘cancer-specific’ methylated gene, MLH1, exhibited increases in all three modifications when its CpG island was methylated, but low levels or absence of the modifications in the other cell lines that lacked DNA methylation (examples in Figure 2C and D, Table II). In contrast, a clearly different pattern was seen for HOXA5, in which methylation of K9 (both di- and trimethylation) and K36 were found at high levels in cell lines in which the HOXA5 CpG island was unmethylated at the DNA level, but were low or undetectable in the methylated cell lines [examples in Figure 2C and D (compare the top two panels), Table II].


View this table:
[in this window]
[in a new window]

  		Table II Summary of ChIP analysis of the relative levels of histone modifications at CTS methylated loci

 
To determine if similar histone modifications were associated with methylation of other CTS methylated genes, the analysis was also performed on the three other genes previously identified as exhibiting CTS methylation (Maspin, MCJ and 14-3-3sigma). This analysis determined that a second CTS methylated gene, MCJ, also exhibited HOXA5-like patterns of histone modifications in which methylation of K9 and K36 were seen at high levels in the unmethylated, actively transcribing, state (examples in Figure 2C and D, Table II). However, not all CTS methylated genes exhibited such differences, as both Maspin and 14-3-3 sigma exhibited patterns of histone modifications similar to those observed for the MLH1 gene (examples in Figure 2C and D, Table II).

To confirm the unexpected results for the HOXA5 gene, the ChIP assays were repeated using an expanded panel of cell lines, including an additional epithelial cell line (MCF7, HOXA5 unmethylated and expression positive), an additional mesenchymal cell line (HT1080, HOXA5 methylated and expression negative) and including 1BR.3.G cells with or without treatment with the DNA methylation inhibitor 2'-deoxy-5-azacytidine. For this expanded analysis, quantification was carried out using qPCR. As shown in Table II, the results from this expanded cell line panel confirmed the original analysis, with high levels of K9 and K36 methylation being detected in the unmethylated, expressing cell lines and reduced levels of these modifications being detected in the methylated and non-expressing cell lines. Treatment of the DNA methylated 1BR.3.G cell line with 2'-deoxy-5-azacytidine, to induce reduced methylation levels, resulted in a partial reversion of this phenotype, and resulted in increased levels of histone acetylation and methylation of K9 and K36 (particularly in region 2) in the 2'-deoxy-5-azacytidine-treated 1BR.3.G cells.

High levels of HOXA5 CPG island methylation are detected in AML samples, but not in solid tumours
HOXA5 has previously been demonstrated to play an important role in myeloid differentiation (9,10), and loss of normal differentiation is one of the hallmarks of many haematopoietic malignancies, including AML. Therefore, HOXA5 methylation levels were investigated in DNA derived from AML patient samples to determine if methylation-dependent repression of HOXA5 could play a role in the development of AML. COBRA and pyrosequencing assays were used to determine the methylation levels of the HOXA5 gene, and methylation was assessed at multiple (seven) CpG sites. Digestion of COBRA PCR products with RsaI, which has multiple cut sites (Figure 1A), produced largely the indicated fully methylated or unmethylated bands, suggesting that the vast majority of HOXA5 alleles were either densely methylated or methylation-free (rare partially methylated alleles are seen in some samples as faint intermediate bands between the indicated unmethylated or fully methylated bands (Figure 3). As demonstrated above, normal peripheral blood samples exhibited methylation of ~50% of HOXA5 alleles (Figure 3A). However, analysis of HOXA5 methylation in AML samples detected clearly increased levels of HOXA5 methylation (80–100% alleles methylated) in 60% of AML samples examined using both RsaI (Figure 3A) and HinFI digests (data not shown). Pyrosequencing analysis was also used to confirm the high levels of HOXA5 methylation identified in the majority of AML samples. As shown in Table I, all 12 AML samples that were identified as hypermethylated by COBRA analysis also exhibited high levels of methylation using pyrosequencing. These results suggest that methylation of HOXA5 may play an important role in arrest of normal differentiation during the development of AML.


Figure 3
View larger version (37K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 3 Examples of COBRA analysis of HOXA5 methylation levels in tumour samples. (A) AML, (B) breast cancer, (C) lung cancer and (D) ovarian cancer patient samples and corresponding normal controls. COBRA PCR products from each of the indicated samples were digested with RsaI (which has three sites within the region analysed). The lower digested bands (indicated by arrows) indicate the presence of DNA methylation and the upper undigested bands, the absence of methylation. IVM DNA (100% IVM) served as a positive control. In A the presence (+) or absence (–) of hypermethylation is indicated for each AML sample.

 
Neither RNA nor protein was available from the original AML samples to allow confirmation of loss of expression in samples with HOXA5 hypermethylation. Therefore, HOXA5 methylation and protein expression were analysed in a panel of 11 malignant haematopoietic cell lines and in a further 5 primary AML samples, from which protein could be extracted. While the HL60 cell line demonstrated near-normal levels of HOXA5 methylation, the other 10 cell lines exhibited hypermethylation of the HOXA5 CpG island (Supplementary Figure 1A). Western blotting analysis showed that HL60 cells (and the SVCT cell line, which was used as a positive control) expressed high levels of the HOXA5 protein, while expression was undetectable or very low in all 10 methylated cell lines (Supplementary Figure 1B and C). Similarly in the AML samples, HOXA5 expression was detected in two samples exhibiting normal methylation, but low or no expression was detected in three hypermethylated samples (Figure 4). This confirms that methylation of the HOXA5 CpG island correlates with loss of expression.


Figure 4
View larger version (24K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 4 Hypermethylation of the HOXA5 CpG island is associated with loss of expression in primary AML samples. Western blotting analysis, using a HOXA5-specific antibody, of protein extracts from primary AML samples, as well as the known positive SVCT cell line and non-hypermethylated HL60 leukaemia cell line. The level of HOXA5 methylation in each sample, as determined by COBRA assay, is indicated below. {alpha}-actin is included as a loading control.

 
Methylation of the HOXA5 promoter has previously been detected in both breast and lung cancer and reported to correlate with loss of expression in breast cancer (13,15); however, the identification of methylation within normal cells raises the possibility that the levels of methylation detected in these tumours may, at least in part, have been due to contaminating normal cells. Therefore, to more clearly define the extent of methylation of HOXA5 in solid tumours, the methylation levels of the HOXA5 gene were also assessed in breast, lung and ovarian tumour samples. Low levels of methylation of HOXA5 were invariably detected in all of the solid tumours samples (Figure 3B–D). However, in all three tumour types HOXA5 methylation was also detected in the corresponding normal tissue (which would contain mesenchymal and haematopoietic cells in addition to epithelial cells). In the overwhelming majority of the individual tumour samples the level of HOXA5 methylation was comparable with that detected in the normal tissue samples. In none of the ovarian tumours, and in only in a small number of breast and lung tumours, were clearly increased levels of HOXA5 methylation [up to a maximum of 61% (lung) and 53% (breast) of alleles methylated] detected (Figure 3B–D). In contrast to the AML samples, none of the solid tumour samples exhibited methylation levels similar to the IVM control. It is likely that the low level of methylation detected in the majority of samples was derived from contaminating normal (mesenchymal and haematopoietic) cells, and even when increased methylation levels were detected, a high proportion of unmethylated alleles still remained. Therefore, methylation of HOXA5 is unlikely to play a significant role in silencing of gene expression in these tumour types.

Reversal of DNA methylation results in HOXA5 re-expression and induction of differentiation in myeloid leukaemia cells
Although the only AML cell line in the panel of haematological cell lines analysed above exhibited normal methylation levels, all five chronic myeloid leukaemia lines studied exhibited HOXA5 hypermethylation and loss of expression. Therefore, to begin to investigate the potential role of HOXA5 hypermethylation in loss of normal differentiation in myeloid leukaemia the LAMA84 cell line was treated with the DNA methylation inhibitor 2'-deoxy-5-azacytidine and assessed for expression of HOXA5 and also markers of myeloid differentiation. Western blotting analysis identified re-expression of HOXA5 protein in the 2'-deoxy-5-azacytidine treated, but not mock-treated LAMA84 cells (Supplementary Figure 2A). qRT–PCR was used to assess the expression levels of a panel of markers of myeloid differentiation. Following 2'-deoxy-5-azacytidine treatment, LAMA84 cells exhibited clear increases in expression of cell surface markers (CD11b, CD13 and CD14) and transcription factors (C/EBP epsilon) associated with myeloid differentiation (between 15- and 222-fold increased expression), demonstrating that re-expression of HOXA5 was associated with features of differentiation in this myeloid leukaemia cell line (Supplementary Figure 2B).


   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
Supplementary material
 References
 
Previous reports have identified the HOXA5 protein as a key regulator of myeloid differentiation (9,10), a transactivator of the p53 tumour suppressor gene, and it has been implicated as a tumour suppressor in its own right in breast cancer (13). Here, we demonstrate that HOXA5 is one of a small number of genes that exhibit CTS methylation of their CpG islands within normal cells, which correlates with gene expression. Methylation of the CpG island within the promoter/first exon was identified in normal adult mesenchymal cells and in 50% of alleles in haematopoietic cells, whereas epithelial cells and embryonically derived mesenchymal cells were unmethylated. Expression analysis identified low or undetectable levels of HOXA5 in cells in which the gene was methylated, but in contrast, high levels of expression of the gene were found in cells that lacked HOXA5 DNA methylation.

To date only a small number of genes have been demonstrated to exhibit CTS methylation (20). However, methylation of CpG islands is also observed under a number of other circumstances, including playing a key role in regulation of imprinting, in inactivation of the X-chromosome in females (30) and inactivation of genes during tumour development (5). DNA methylation of CpG islands in these latter cases has been associated with a number of specific alterations in the modifications of histone tails, which are important in altering chromatin structure and consequent silencing of the associated genes (31). In this report we used ChIP assays to assess if similar histone modifications were associated with CpG island methylation at the HOXA5 locus. As expected this analysis identified increased methylation of K9 and K36 of histone H3 following DNA methylation of the MLH1 gene (which is frequently inactivated in a tumour-specific fashion). In contrast, these same histone modifications (K9 and K36 methylation) were associated with the absence of DNA methylation and active transcription at the HOXA5 loci. Analysis of the other genes known to be controlled by CTS methylation identified a second gene, MCJ, which exhibited similarly high levels of K9 and K36 methylation in actively transcribing, DNA unmethylated, alleles (although both maspin and 14-3-3sigma had patterns similar to that of the MLH1 gene). Furthermore, treatment of the DNA methylated 1BR.3.G cell line with 2'-deoxy-5-azacytidine, to induce reduced methylation levels, resulted in a partial reversion of this phenotype, resulting in increased levels of histone acetylation and methylation of K9 and K36, consistent with a role for DNA methylation in maintenance of the unusual patterns of histone modifications observed at the HOXA5 locus. These results suggest that the HOXA5, and MCJ, loci must be protected from the mechanisms that would normally result in transcriptional silencing in the presence of these histone modifications [such as HP-1 binding to methylated K9 of histone H3 (32)] and that either these modifications then become activating or other aspects of chromatin structure at these loci are more influential in determining transcriptional activity. These results also suggest that the function of particular histone modifications in controlling gene expression is likely to be dependent on the specific chromatin context in which these modifications are present.

It is possible that the patterns of histone modification seen for MCJ and HOXA5 may be part of a transcriptional control mechanism specific for some CTS methylated genes and not seen in imprinted genes or genes aberrantly methylated in cancer. However, the changes in chromatin structure associated with DNA methylation have not been extensively studied in the vast majority of genes identified as exhibiting aberrant DNA methylation in tumours, and while in those that have been studied methylation of H3–K9 has often been investigated, the role of H3–K36 methylation has not been addressed. Indeed, although H3–K36 methylation in promoter regions has previously been associated with transcriptional repression (33,34), we believe that the data presented for MLH1 are the first link between aberrant DNA methylation of ‘cancer-specific’ methylated genes and increased H3–K36 methylation. While examining large numbers of ‘cancer-specific’ methylated genes individually to identify patterns of histone modification similar to those seen for MCJ and HOXA5 would be time-consuming, the recent development of array-based ‘ChIP on chip’ approaches (35,36) raises the possibility of more genome-wide screens for similarly regulated genes.

HOXA5 has previously been implicated as a tumour suppressor gene and an important regulator of haematopoietic differentiation, and thus alterations to the normal patterns of HOXA5 CpG island methylation may play an important role in human malignancy. Analysis of HOXA5 in AML samples identified clearly increased methylation levels, relative to peripheral blood, in the majority (60%) of samples, and hypermethylation of the HOXA5 gene was associated with loss of protein expression in both leukaemic cell lines and primary AML samples. Loss of normal differentiation is a key feature of the development of AML. As reduced HOXA5 expression has previously been shown to result in inhibition of granulocytic/monocytic differentiation (10), hypermethylation of HOXA5 in the AML samples would be consistent with a role for loss of HOXA5 in arrest of normal differentiation in AML. An alternative explanation for the increased methylation at the HOXA5 locus, relative to peripheral blood, could be that this is reflective, as opposed to causative, of the arrested differentiation in AML or that methylation in peripheral blood is restricted to myeloid cells. However, HOXA5 has previously been reported to be expressed at higher levels in more immature haematopoietic cells (37), and analysis of HOXA5 methylation in CD133+ selected cells [thought to represent early haematopoietic progenitors (38)] did not identify increased HOXA5 methylation (data not shown), indicating that HOXA5 hypermethylation is not a feature of more immature haematopoietic cells. Also, analysis of individual cell types, purified from peripheral blood, identified indistinguishable levels of methylation in both myeloid and lymphoid cells, indicating that the CpG island methylation of HOXA5 is not restricted to the myeloid compartment.

Treatment of the LAMA84 cell line, in which the HOXA5 gene is hypermethylated, with the DNA methylation inhibitor 2'-deoxy-5-azacytidine, resulted in re-expression of HOXA5 protein. This demonstrates that DNA methylation in this myeloid leukaemia cell line is necessary for continued suppression of HOXA5 expression. Furthermore, treatment with the demethylating agent was also associated with increased expression of multiple markers of myeloid differentiation, consistent with a role for re-expression of HOXA5 in inducing differentiation in myeloid leukaemia cells. However, it is likely that the expression of multiple genes would have been altered following 2'-deoxy-5-azacytidine treatment, and thus the induction of differentiation cannot be specifically attributed to re-expression of HOXA5. More detailed biochemical analysis will be required to define the specific role of loss of HOXA5 expression in the development of myeloid leukaemia.

Global alterations to expression of HOXA cluster genes have previously been observed in AML and found to be associated with specific cytogenetic risk groups (39), suggesting that hypermethylation of HOXA5 may reflect these previously observed groups with known differences in HOXA gene expression. Cytogentic analysis was only available for a subset of the samples; however, hypermethylation was observed in all cytogenetic risk groups (low risk, 3/6; intermediate risk, 6/6; high risk, 1/2), suggesting that hypermethylation is unlikely to be secondary to global changes in HOXA cluster expression. Clearly, the sample size in this study is limited, and thus a larger study is currently under way to more clearly determine the relationship between HOXA5 CpG island methylation, cytogenetics, FAB subclass and global changes in HOXA cluster gene expression.

Methylation of the CpG island within the 5' region of the HOXA5 locus has previously been identified in both breast and lung cancer (13,15), suggesting that DNA methylation may be an important mechanism of inactivation of HOXA5 in these tumour types, and HOXA5 had been identified as a putative tumour suppressor gene in breast cancer (14). Analysis of samples from a variety of solid tumour types (breast, lung and ovarian tumour samples) demonstrated the presence of methylation of the HOXA5 CpG island in all of the samples analysed; however, the level of methylation was almost invariably low and comparable with that observed in the corresponding normal tissue. On the basis of these results it appears that DNA methylation does not play a major role in inactivation of HOXA5 in these tumour types. While a low level of HOXA5 methylation in tumour cells could not be ruled out, it is likely that the vast majority of methylation detected in the solid tumour samples is derived from contaminating normal (mesenchymal or haematopoietic) cells. Given that, at least in breast cancer, loss of HOXA5 protein expression is a frequent event (13), alternative mechanisms, such as altered expression of key regulatory transcription factors, must be responsible for downregulation of HOXA5.

Human peripheral blood samples exhibit methylation of ~50% of HOXA5 alleles. As HOXA5 has been implicated as an important regulator of haematopoietic differentiation (9), this suggests a potential role for HOXA5 methylation in control of HOXA5 expression levels, and consequently differentiation, in haematopoietic cells. The identification of methylation of 50% of HOXA5 alleles in haematopoietic cells suggested that either methylation was restricted to certain haematopoietic cell types or that HOXA5 may be imprinted. However, the results presented here demonstrate that similar levels of HOXA5 methylation are present in multiple haematopoietic cell types and that methylation was not allele-specific. A further potential explanation for the pattern of HOXA5 methylation would be that mono-allelic expression of HOXA5 is brought about by random inactivation of one of the two HOXA5 alleles in each cell. Indeed, such a process, termed allelic exclusion, has previously been demonstrated in haematopoietic cells, to maintain mono-allelic expression of immunoglobulin and T-cell receptor genes in B and T lymphocytes (40).

The results presented here emphasize the need for quantitative analysis of gene methylation, when assessing its potential role in tumour development. In this report we identified very high levels of methylation of the HOXA5 locus in AML samples, suggesting a potential role for HOXA5 in the development of this tumour type. In contrast, although methylation was also present in all breast, lung and ovarian cancer samples analysed, the level of methylation was almost invariably low, arguing against any significant role for HOXA5 methylation in these tumour types. As PCR-based methods are capable of detecting very low levels of methylation and comparatively few reports identifying tumour-related methylation of particular loci used quantitative methods to assess methylation levels, it is possible that the importance of DNA methylation in gene inactivation may have been over-estimated in some cases.


   Supplementary material
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
References
 
Supplementary data are available at Carcinogenesis online.


   Acknowledgments
 
The authors would like to thank Dr B.R. Davies, Dr K. Parkinson and Dr N. Keith for providing cell lines, Prof. A. Dickinson and Mr Gerard Cain for providing primary AML samples and Ms Alyson Doig and Ms Linda Richmond for technical assistance. This work was supported by a Cancer Research UK programme grant awarded to R.B. (SP1429/1902) and a Leukaemia Research Trust for Scotland grant awarded to R.S.

Conflict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 

  1. Bird A. (1996) The relationship of DNA methylation to cancer. Cancer Surv. 28:87–101.[ISI][Medline]
  2. Bird A. and Wolffe A.P. (1999) Methylation-induced repression—belts, braces and chromatin. Cell 99:451–454.[CrossRef][ISI][Medline]
  3. Kouzarides T. (2002) Histone methylation in transcriptional control. Curr. Opin. Genet. Dev. 12:198–209.[CrossRef][ISI][Medline]
  4. Holiday R. and Pugh J.E. (1975) DNA modification mechanisms and gene activity during development. Science 187:226–232.[Free Full Text]
  5. Costello J.F. and Plass C. (2001) Methylation matters. J. Med. Genet. 38:285–303.[Abstract/Free Full Text]
  6. Futscher B.W., Oshiro M.M., Wozniak R.J., Holtan N., Hanigan C.L., Duan H., Domann F.E. (2002) Role for DNA methylation in the control of cell type-specific maspin expression. Nat. Genet. 31:175–179.[CrossRef][ISI][Medline]
  7. Strathdee G., Davies B.R., Vass J.K., Siddiqui N., Brown R. (2004) Cell type-specific methylation of an intronic CpG island controls expression of the MCJ gene. Carcinogenesis 25:693–701.[Abstract/Free Full Text]
  8. Cillo C., Cantile M., Faiella A., Boncinelli E. (2001) Homeobox genes in normal and malignant cells. J. Cell. Physiol. 188:161–169.[CrossRef][ISI][Medline]
  9. Crooks G.M., Fuller J., Petersen D., Izadi P., Malik P., Pattengale P.K., Kohn D.B., Gasson J.C. (1999) Constitutive HOXA5 expression inhibits erythropoiesis and increases myelopoiesis from human hematopoietic progenitors. Blood 94:519–528.[Abstract/Free Full Text]
  10. Fuller J.F., McAdara J., Yaron Y., Sakaguchi M., Fraser J.K., Gasson J.C. (1999) Characterization of HOX gene expression during myelopoiesis: role of HOXA5 in lineage commitment and maturation. Blood 93:3391–3400.[Abstract/Free Full Text]
  11. Wang Y., Hung C., Koh D., Cheong D., Hooi S.C. (2001) Differential expression of Hox A5 in human colon cancer cell differentiation: a quantitative study using real-time RT–PCR. Int. J. Oncol. 18:617–622.[ISI][Medline]
  12. Aubin J., Chailler P., Menard D., Jeannotte L. (1999) Loss of HOXA5 gene function in mice perturbs intestinal maturation. Am. J. Physiol. 277:C965–C973.
  13. Raman V., Martensen S.A., Reisman D., Evron E., Odenwald W.F., Jaffee E., Marks J., Sukumar S. (2000) Compromised HOXA5 function can limit p53 expression in human breast tumours. Nature 405:974–978.[CrossRef][Medline]
  14. Chen H., Chung S., Sukumar S. (2004) HOXA5-induced apoptosis in breast cancer cells is mediated by caspases 2 and 8. Mol. Cell. Biol. 24:924–935.[Abstract/Free Full Text]
  15. Shiraishi M., Sekiguchi A., Oates A.J., Terry M.J., Miyamoto Y. (2002) HOX gene clusters are hotspots of de novo methylation in CpG islands of human lung adenocarcinomas. Oncogene 21:3659–3662.[CrossRef][ISI][Medline]
  16. Anthoney D.A., McIlwrath A.J., Gallagher W.M., Edlin A.R.M., Brown R. (1996) Microsatellite instability, apoptosis and loss of p53 function in drug resistant tumour cells. Cancer Res. 56:1374–1381.[Abstract/Free Full Text]
  17. Davies B.R., Steele I.A., Edmondson R.J., Zwolinski S.A., Saretzki G., von Zglinicki T., O'Hare M.J. (2003) Immortalisation of human ovarian surface epithelium with telomerase and temperature-sensitive SV40 large T antigen. Exp. Cell. Res. 288:390–402.[CrossRef][ISI][Medline]
  18. Strathdee G., MacKean M., Illand M., Brown R. (1999) A role for methylation of the hMLH1 promoter in loss of hMLH1 expression and drug resistance in ovarian cancer. Oncogene 18:2335–2341.[CrossRef][ISI][Medline]
  19. Xiong Z. and Laird P.W. (1997) COBRA: a sensitive and quantitative DNA methylation assay. Nucleic Acids Res. 25:2532–2534.[Abstract/Free Full Text]
  20. Strathdee G., Sim A., Brown R. (2004) Control of gene expression by CpG island methylation in normal cells. Biochem. Soc. Trans. 32:913–915.[CrossRef][ISI][Medline]
  21. Hershko A.Y., Kafri T., Fainsod A., Razin A. (2003) Methylation of HOXA5 and HoxB5 and its relevance to expression during mouse development. Gene 302:65–72.[CrossRef][ISI][Medline]
  22. Tost J., Dunker J., Gut I.G. (2003) Analysis and quantification of multiple methylation variable positions in CpG islands by pyrosequencing. Biotechniques 35:152–156.[ISI][Medline]
  23. Kondo Y. and Issa J.P. (2004) Epigenetic changes in colorectal cancer. Cancer Metastasis Rev. 23:29–39.[CrossRef][ISI][Medline]
  24. Fahrner J.A., Eguchi S., Herman J.G., Baylin S.B. (2002) Dependence of histone modifications and gene expression on DNA hypermethylation in cancer. Cancer Res. 62:7213–7218.[Abstract/Free Full Text]
  25. Kondo Y., Shen L., Issa J.P. (2003) Critical role of histone methylation in tumor suppressor gene silencing in colorectal cancer. Mol. Cell. Biol. 23:206–215.[Abstract/Free Full Text]
  26. Eberharter A. and Becker P.B. (2002) Histone acetylation: a switch between repressive and permissive chromatin. Second in review series on chromatin dynamics. EMBO Rep. 3:224–229.[CrossRef][ISI][Medline]
  27. Nguyen C.T., Weisenberger D.J., Velicescu M., Gonzales F.A., Lin J.C., Liang G., Jones P.A. (2002) Histone H3-lysine 9 methylation is associated with aberrant gene silencing in cancer cells and is rapidly reversed by 5-aza-2'-deoxycytidine. Cancer Res. 62:6456–6461.[Abstract/Free Full Text]
  28. Fournier C., Goto Y., Ballestar E., Delaval K., Hever A.M., Esteller M., Feil R. (2002) Allele-specific histone lysine methylation marks regulatory regions at imprinted mouse genes. EMBO J 21:6560–6570.[CrossRef][ISI][Medline]
  29. Xin Z., Allis C.D., Wagstaff J. (2001) Parent-specific complementary patterns of histone H3 lysine 9 and H3 lysine 4 methylation at the Prader–Willi syndrome imprinting center. Am. J. Hum. Genet. 69:1389–1394.[CrossRef][ISI][Medline]
  30. Heard E., Clerc P., Avner P. (1997) X-chromosome inactivation in mammals. Ann. Rev. Genet. 31:577–610.
  31. Richards E.J. and Elgin S.C. (2002) Epigenetic codes for heterochromatin formation and silencing: rounding up the usual suspects. Cell 108:489–500.[CrossRef][ISI][Medline]
  32. Lachner M., O'Carroll D., Rea S., Mechtler K., Jenuwein T. (2001) Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 410:116–120.[CrossRef][Medline]
  33. Strahl B.D., Grant P.A., Briggs S.D., et al. (2002) Set2 is a nucleosomal histone H3-selective methyltransferase that mediates transcriptional repression. Mol. Cell. Biol. 22:1298–1306.[Abstract/Free Full Text]
  34. Landry J., Sutton A., Hesman T., Min J., Xu R.M., Johnston M., Sternglanz R. (2003) Set2-catalyzed methylation of histone H3 represses basal expression of GAL4 in Saccharomyces cerevisiae. Mol. Cell. Biol. 23:5972–5978.[Abstract/Free Full Text]
  35. Ren B., Robert F., Wyrick J.J., et al. (2000) Genome-wide location and function of DNA binding proteins. Science 290:2306–2309.[Abstract/Free Full Text]
  36. Iyer V.R., Horak C.E., Scafe C.S., Botstein D., Snyder M., Brown P.O. (2001) Genomic binding sites of the yeast cell-cycle transcription factors SBF and MBF. Nature 409:533–538.[CrossRef][Medline]
  37. Ivanova N.B., Dimos J.T., Schaniel C., Hackney J.A., Moore K.A., Lemischka I.R. (2002) A stem cell molecular signature. Science 298:601–604.[Abstract/Free Full Text]
  38. Matsumoto K., Yasui K., Yamashita N., Horie Y., Yamada T., Tani Y., Shibata H., Nakano T. (2000) In vitro proliferation potential of AC133 positive cells in peripheral blood. Stem Cells 18:196–203.[Abstract/Free Full Text]
  39. Debernardi S., Lillington D.M., Chaplin T., Tomlinson S., Amess J., Rohatiner A., Lister T.A., Young B.D. (2003) Genome-wide analysis of acute myeloid leukemia with normal karyotype reveals a unique pattern of homeobox gene expression distinct from those with translocation-mediated fusion events. Genes Chromosomes Cancer 37:149–158.[CrossRef][ISI][Medline]
  40. Bergman Y. and Cedar H. (2004) A stepwise epigenetic process controls immunoglobulin allelic exclusion. Nat. Rev. Immunol. 4:753–761.[CrossRef][ISI][Medline]
Received January 24, 2006; revised July 5, 2006; accepted July 12, 2006.


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?


This article has been cited by other articles:


Home page
 Clin. Cancer Res.Home page
G. Strathdee, T. L. Holyoake, A. Sim, A. Parker, D. G. Oscier, J. V. Melo, S. Meyer, T. Eden, A. M. Dickinson, J. C. Mountford, et al.
Inactivation of HOXA Genes by Hypermethylation in Myeloid and Lymphoid Malignancy is Frequent and Associated with Poor Prognosis
Clin. Cancer Res., September 1, 2007; 13(17): 5048 - 5055.
[Abstract] [Full Text] [PDF]