Human Molecular Genetics, 2002, Vol. 11, No. 20 2479-2488
© 2002 Oxford University Press
Cancer epigenomics
Division of Human Cancer Genetics, Department of Molecular Virology, Immunology and Medical Genetics, The Ohio State University, Columbus OH, USA
Received June 28, 2002; Accepted July 3, 2002
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ABSTRACT |
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 TOP ABSTRACT CANCER GENOMICSEPIGENOMICSCANCER EPIGENOMICSFUTURE DIRECTIONSREFERENCES |
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Research in cancer epigenomics is driven by the development of novel technologies and the utilization of model organisms ranging from yeasts to plants to vertebrates. For decades, the search for cancer genes has focused on genetic defects that were used as tags for identification of these genes. With the realization that epigenetic modifications, most importantly DNA methylation events, are frequently involved in transcriptional changes in both tumor suppressor genes and oncogenes, techniques have been developed that support the identification of novel cancer genes altered by DNA methylation alone or in combination with genetic events. Recent data demonstrate that, in addition to DNA methylation, chromatin modifications are also involved in gene regulation. We are now beginning to understand this interesting interplay between chromatin modifications, DNA methylation and gene regulation. This review will summarize our current knowledge of DNA methylation and histone modification in normal cells, introduce emerging concepts that show the intimate link between DNA methylation and chromatin modifications, and highlight recent advancements in our understanding of aberrant DNA methylation, with special emphasis on genome-wide hypermethylation.
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CANCER GENOMICS |
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Tumorigenesis is known to be a multistep process in which defects in various cancer genes accumulate (1,2). Virtually every tumor type has revealed an enormous complexity of altered gene functions, including activation of growth-promoting genes as well as silencing of genes with tumor growth-suppressing functions, all contributing to uncontrolled growth. Hanahan and Weinberg proposed that cancer gene functions can be classified into six essential alterations in cell physiology, including self-sufficiency in growth signals, insensitivity to growth inhibitory signals, evasion of apoptosis, limitless replicative potential, sustained angiogenesis, and tissue invasion and metastasis (3).
Much of our current knowledge of tumor suppressor genes comes from studies of inherited cancer syndromes (4), where germline genetic alterations have been identified in genes responsible for accelerated and early-onset cancer. More difficult is the search for cancer genes in sporadic cancers. Genome-scanning techniques are used to identify genetic alterations that are common both within and among tumor types (Table 1) (5). Activated oncogenes were detected by accumulation of multiple gene copies in large amplicons, cytogenetically visible as homogeneously staining regions in standard G or C banding (6). Comparative genomic hybridization (CGH) on large, fully sequenced genomic clones or cDNAs is now an alternative option for the search for amplified sequences and offers much greater resolution (7,8). Since the landmark work by Knudson (9), and subsequently Cavenee et al. (10), loss of heterozygosity (LOH) is used as a marker for the location of a candidate tumor suppressor gene. Genome-wide scans using highly polymorphic microsatellite markers are employed to find the most commonly deleted region in a certain tumor type (11). These searches are complemented by more recent techniques such as CGH or spectral karyotyping (SKY), which allow a genome-wide view of chromosomal alterations and rearrangements (12–14), including both DNA amplification and allelic loss. For a long time, the lack of sequence information and the presence of multiple candidate genes in amplified or deleted regions hampered the rapid identification of novel cancer genes. The availability of the human genome sequences (15) and large insert genomic clone resources (16), and the development of new array-based techniques for genome scanning on the transcriptional level (17), have initiated a new era in cancer research and high expectations for the discovery of a multitude of novel cancer genes in the future.
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EPIGENOMICS |
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It is now clear that the genetic abnormalities found in cancers will not provide the complete picture of genomic alterations (Fig. 1). Epigenetic changes, mainly DNA methylation and, more recently, modification of histones, are now recognized as additional mechanisms contributing to the malignant phenotype. The study of these epigenetic changes on a genome-wide scale is referred to as epigenomics.
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DNA methylation in mammals
Epigenetic modifications of the DNA do not alter the sequence code; however, they are heritable and involved in regulation of gene transcription (see below). DNA methylation, the addition of a methyl group to the 5'-carbon of cytosine, is one such epigenetic modification found in DNA (18). In mammals, the major target for DNA methylation is a cytosine located next to a guanine (5'-CpG-3') (19), although exceptional CpNpG methylation has been described (20). These targets of methylation are not equally distributed in the genome, but found in long CG-rich sequences present in satellite repeat sequences, middle repetitive rDNA sequences, centromeric repeat sequences and CpG islands. CpG islands are sequences longer than 200 bp with a GC content of over 50% (in contrast to a genome-wide average of about 40%) and an observed over expected ratio of CpG of 0.6 or greater (21,22). Interestingly, CpG islands are found mainly in the 5'-regions of housekeeping genes as well as some other specifically tissue expressed genes and usually extend from the promoter region into the first exon and sometimes into intron 1 (15,23). Most CpG islands are unmethylated in normal cells; however, there are certain conditions where these sequences become methylated and form part of gene regulation (24). The majority of CpG islands on the inactive X-chromosome in a female cell are methylated (25), and certain CpG island-like sequences in the vicinity of imprinted genes have been found to be methylated in an allele-specific manner (26). Furthermore, it has been found that some CpG islands become methylated with age (27). While CpG islands are usually unmethylated, other GC-rich sequences, e.g. the centromeric repeat sequences and satellite sequences, are highly methylated in normal cells.
The establishment of DNA methylation patterns
DNA methylation is a dynamic but tightly regulated process. Methylation patterns are faithfully transmitted to the next generations during cell division, yet, during embryonic development, currently undefined regulatory mechanisms allow rapid demethylation in very early stages followed by re-establishment of methylation patterns after implantation (28). While some of the enzymes involved in these processes are known, we are only beginning to understand the components of this regulatory network, let alone the organization and role of each of the components. DNA methyltransferases (DNMTs) transfer the methyl group that is provided by S-adenosylmethionine to the 5'-carbon of a cytosine. Two de novo methyltransferases, DNMT3a and DNMT3b, are known. Both enzymes are highly expressed in undifferentiated embryonic stem cells and at low levels in somatic tissues (29) and are thought to be responsible for the methylation of repetitive elements. DNMT1, on the other hand, is an enzyme needed for the maintenance of DNA methylation after replication of a methylated sequence and is found in an enzyme complex together with proliferating cellular nuclear antigen (PCNA) located at the replication fork (30). Other members of this protein complex include histone deacetylase 2 (HDAC2) and a novel protein DMAP1 (DNMT1-associated protein), both involved in transcriptional repression (see below) (31). The importance of DNA methylation in normal cells as well as the need for each of the methyltransferases was demonstrated in mice lacking DNMTs. Embryos lacking both copies of either DNMT1 or DNMT3a die before birth, and DNMT3b homozygous null mice die a few weeks after birth (29,32).
DNA methylation and transcription
DNA methylation in the promoter regions of genes is correlated with gene silencing; however, methylation may, in some cases (e.g. H19/Igf2 and Rasgrf1), have a gene-activating effect (33,34). Two underlying mechanisms have been identified. First, binding of transcription factors or enhancer-blocking elements, such as CTCF, may be inhibited by DNA methylation and thus exert its effect on the transcription of downstream genes in the case of transcription factors (35,36), or even at a great distance as seen for the enhancer-blocking element CTCF (34,37). The second and probably more general mechanism involves proteins that detect methylated DNA through methyl CpG-binding domains (MBDs) (38–41). Four MBD-containing proteins are currently characterized (40,42). A methyl CpG-binding domain is found in the MECP2 protein as well in one of the proteins (MBD2) in the MeCP1 complex (40,41). MeCP2 is a single polypeptide characterized by an MBD and a transcriptional repression domain (40,41,43). There is now substantial evidence that these proteins mediate recruitment of repressor complexes that include histone deacetylases (HDACs) (44,45). HDACs remove acetyl groups from lysine residues of histones H3 and H4, and it is believed that the positively charged, deacetylated histones result in condensation of chromatin and thus limit access of transcription factors to promoter regions of genes localized nearby. Well-studied co-repressor complexes include Sin3A and Mi-2NuRD and two Rb-associated histone-binding proteins, RbAp46 and RbAp48 (40,46). In addition the Mi-2/NuRD complex contains ATP-dependent chromatin remodeling factors, which have the ability to reposition the nucleosomes on the DNA and thus restrict access of transcription factors.
Histone modifications: ‘the histone code’
For many years epigenetic research focused on DNA methylation; this is now changing and considerable attention is being given to histone modifications. DNA is wrapped around an octamer of histones consisting of a histone 3 (H3) and histone 4 (H4) tetramer and two histone 2A and histone 2B dimers. This structure, called a nucleosome, is the building block of chromatin. Histones are basic proteins that consist of a globular domain and a histone tail that protrudes out of the nucleosome. These histone tails are targets for covalent post-translational modifications, including acetylation, methylation and phosphorylation. There is now increasing evidence that characteristic modification patterns (the ‘histone code’) on histone tails are involved in gene regulation through changes in chromatin structure and condensation (47). The histone code is recognized by effector proteins that bind to the nucleosomes and recognize specific patterns of modification. Hypoacetylation of H3 and H4 are associated with heterochromatic, transcriptionally inactive regions in the genome. The states of acetylation and deacetylation are regulated by histone acetyltransferases (HAT) and deacetylases (HDAC) and recognized by effector proteins containing a bromodomain (48,49). In addition to acetylation and deacetylation, histone H3 methylation has been described for lysine residues Lys4 and Lys9. It is interesting to note that H3-Lys9 is acetylated in active chromatin, but methylated in regions where genes are silenced. While H3-Lys4 methylation has been correlated with active gene expression (50,51), methylation at H3-Lys9 correlates with gene silencing and is found in heterochromatin (52–54). The chromodomain of HP1 binds to methylated H3-Lys9 and is involved in heterochromatin assembly (54,55). H3-Lys9 methylation is mediated by SUV39H1, an enzyme homologous to, Su(var)3-9 of Drosophila, containing an evolutionarily conserved SET domain found in protein methyltransferases (56).
Cooperativity between chromatin modifications and DNA methylation
Little is known about the regulation of DNA methylation. Evidence is accumulating that suggests the involvement of chromatin modifications in this process. As discussed above, DNA methylation is tied to modifications in the chromatin. Methylated DNA is a target for transcriptional repressor complexes, including proteins that specifically detect methylated DNA and HDACs. Also, there are additional lines of evidence that link DNA methylation and changes in chromatin. Owing to the importance of DNA methylation during mammalian development, DNA methylation mutants are rare in mammals yet exist in other organisms. Vongs et al. (57) identified Arabidopsis thaliana mutants with defects in DNA methylation. One mutant, decreased in methylation 1 (ddm1), showed only 30% of the normal DNA methylation level in repetitive elements (57). Ddm1 is a member of the SNF2-like helicase superfamily, a group of proteins that are able to disrupt histone–DNA interactions (58). This finding suggested for the first time that chromatin remodeling is a required first step to create access for the DNA methyltransferases to DNA sequences to be methylated.
Further support came from a mouse model. Lymphoid specific helicase, Lsh, is another member of the SNF2-like helicase superfamily frequently involved in chromatin remodeling. Mice with a homozygous deletion of Lsh show perinatal lethality and a marked decrease in overall 5'-methylcytosine (59). Surprisingly, Dnmt1, Dnmt3a and Dnmt3b methyltransferase activities were not altered in tissues from Lsh-/- mice, thus suggesting that Lsh protein is actively involved in the maintenance and/or de novo DNA methylation process (59). Hypomethylation in these mutant mice was found in repetitive elements, including, minor satellite repeat sequences, major satellite repeats, intracisternal A-particles, small-interspersed repetitive elements (Sine B1), and telomeric sequences. Hypomethylation was also found in the middle or 3' end of two genes that show tissue-specific expression, Pgk2 and ß-globin. In addition, the 5' end of Pgk1, a gene localized on the X-chromosome, was found to be hypomethylated in the mutant mice (59).
The filamentous fungus Neurospora crassa is an ideal model organism with which to study cytosine methylation since, unlike in mammals, DNA methylation is not essential. Selker's group has set up a mutagenesis experiment in N. crassa and screened for mutants with a defect in DNA methylation (60). Two mutant strains have been characterized so far, which have identified dim-2, a DNA methyltransferase with de novo and maintenance methyltransferase activity (61). The second mutant, dim-5, was a mutant strain isolated in a selection experiment for strains resistant to 5-azacytidine, an inhibitor of DNMTs. The dim-5 mutant showed reduced DNA methylation, and exhibited slow and irregular growth. A nonsense mutation responsible for this phenotype was identified in the open reading frame (ORF) of a new gene, called dim-5. Isolation and characterization of this gene identified it as a protein containing a SET domain, which is characteristic for histone methyltransferases (62). Biochemical assays showed that dim-5 specifically targets lysine 9 of H3. Furthermore, the authors showed that mutating the histone H3 gene at the sequence coding for lysine 9 caused loss of DNA methylation in vivo. This elegant experiment demonstrated convincingly that methylation of H3-Lys9 is required for DNA methylation. Future studies will show if this phenomenon is also found in mammalian cells.
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DNA methylation changes in cancer cells include the loss of methylation at normally methylated sequences (hypomethylation) and the gain of methylated sequences at sites usually unmethylated (hypermethylation). Cancer epigenomics involves the study of epigenetic alterations in cancer and can be complicated by questions of cause and effect. Table 2 lists some of the discoveries that have been made by DNA methylation studies in human malignancies.
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Hypomethylation
Global DNA hypomethylation has been reported in almost every human malignancy (63–67). Measurements of the 5-methylcytosine content by HPLC revealed a reduced level compared to normal tissue controls (68). This global hypomethylation of the tumor genome could be verified by assays determining demethylation in specific sequences. Interestingly, the majority of hypomethylation events occur in repetitive elements localized in satellite sequences or centromeric regions (69). Pericentric heterochromatin regions on chromosomes 1 and 16, heavily methylated in normal cells, are found to be demethylated not only in tumor genomes, but also in patients with immunodeficiency, centromeric region instability, facial anomalies syndrome (ICF). A mutation of the de novo methyltransferase DNMT3b gene has been found in ICF patients (29,70,71), leading to structural instability of chromosomes 1 and 16. In addition to demethylation of repetitive elements in cancer, demethylation events in protooncogenes leading to gene activation have also been described (72–74). For a more detailed review on DNA hypomethylation in cancer, see Costello and Plass (75).
CpG island hypermethylation in tumor suppressor genes
While hypomethylation of repetitive elements is a common finding in human malignancies, gene-associated CpG islands are the targets of hypermethylation. Hypermethylation was initially discovered as a novel mechanism of tumor suppressor gene silencing in numerous genes that had been identified as targets for genetic alterations. Large-scale methylation studies on cancer genes became possible with the introduction of sodium bisulfite treatment of genomic DNA that results in a conversion of unmethylated cytosines to uracils but leaves methylcytosines unconverted (76). Subsequent development of PCR assays that could discriminate the methylated and unmethylated sequences allowed rapid screening of multiple tumor samples for promoter methylation (77,78). This candidate gene approach identified methylated tumor suppressor genes in virtually every tumor type and established the methylation of promoter sequences as one of the major mechanisms involved in tumor suppressor gene inactivation (75,79) (Fig. 1). Many investigators showed that genes silenced by promoter hypermethylation could be reactivated by treatment with the demethylating drug 5-aza-2'-deoxycytidine, either alone or in combination with an HDAC inhibitor (e.g. trichostatin A (TSA)) (45,80–82), giving support to the idea that CpG island hypermethylation plays a critical role in gene silencing.
In a large study comparing aberrant methylation in the promoters of 10 tumor suppressor genes in sporadic and familial breast and colorectal cancers, it was found that promoter methylation is a frequent event for the inactivation of the non-mutated copy of a tumor suppressor gene in familial cancers (83). Interestingly, aberrant methylation was not found on the mutated allele, thus strongly indicating that DNA methylation does indeed contribute to oncogenesis as the second hit in such a scenario. Hypermethylation in sporadic cancers can be either the first or second hit required for cancer gene silencing (Fig. 1). Although the overall levels of methylation seemed similar between familial and sporadic cancers, there were differences in the frequencies with which methylation events occurred in various genes (83).
A multitude of studies using the candidate gene approach established the importance of DNA methylation in tumor suppressor gene silencing. Several observations are noteworthy. First, while the majority of tumor suppressor genes have been found to be methylated in multiple tumor types, there are a few exceptions. BRCA1 methylation, for example, has been found only in breast and ovarian cancers (84–87), and VHL promoter methylation has been reported only in clear cell renal carcinomas (88) and hemangioblastomas (89). Second, it could be shown that DNA methylation events occur early in tumorigenesis, which makes this modification a perfect target for early detection of malignant cells. By increasing the sensitivity of methylation-specific PCR for the p16 and O6-methyl-guanine-DNA methyltransferase (MGMT) genes, it was possible to detect methylated sequences in sputum in 100% of patients with squamous cell lung carcinoma 3 years prior to clinical diagnosis (90). Third, MGMT is a DNA repair gene able to remove guanine adducts, which are added by alkylating agents used as chemotherapeutic drugs in glioma treatment. Thus the presence of MGMT protein subsequently causes resistance to such treatment. Recently, it was reported that promoter methylation of MGMT resulted in the transcriptional silencing of this gene and was found in glioma patients that responded to treatment with carmustine by prolonged overall and disease-free survival (91).
Large-scale methylation analysis of multiple candidate genes has been hampered by the lack of an array-based assay. Recently, an oligo-microarray-based technique was developed. This technique allows the assessment of DNA methylation status in hundreds of CpG island sequences in a single hybridization experiment. This assay has demonstrated the usefulness of DNA methylation events for discriminating normal versus tumor tissue and different normal tissues and holds great potential as a diagnostic tool for the subclassification of human tumors (92).
Genome-wide CpG island hypermethylation
Several groups have taken a different approach to study DNA methylation changes. Novel genes, which have not been identified as cancer genes, can be isolated using genome-scanning techniques (Table 1). Depending on the technique, it is possible to obtain overall frequencies for methylation events and thus prioritize the cloning and identification of genes. In addition to identifying methylation in genes commonly affected by genetic abnormalities, these techniques allow the identification of genes that are preferentially silenced by methylation. This is significantly different from the candidate gene approach, since the ‘candidates’ were defined precisely by their tendency to show genetic alterations. The choice of methylation scanning techniques depends on the particular question of interest, and the availability and source of DNA. PCR-based assays are easily adopted by almost every molecular laboratory and can be used even on archival sources of DNA. Restriction landmark genomic scanning (RLGS) requires special equipment and training and can only be used with high-quality DNA. The array-based technologies require special equipment and can be quite expensive to perform. Each of the technologies described below has its own unique set of strengths and weaknesses.
Methylation-sensitive arbitrarily primed PCR (Ms–AP–PCR), is based on the digestion of genomic DNAs with isoshizomers, one methylation-sensitive (HpaII) and the other methylation insensitive (MspI). Restriction-digested DNAs are amplified using PCR, for which the annealing temperature is reduced so that the primers bind arbitrarily even at sequences with less homology. Once appropriate conditions have been determined, multiple fragments can be reproducibly amplified. If the sequence is methylated, the PCR using the HpaII-digested DNA as a template will amplify a band, whereas the band will be missing in the MspI reaction (93–95). Ms–AP–PCR was used in a study to identify novel methylated sequences from colon, bladder and prostate cancers and showed differences in their methylation patterns and frequencies (94).
For MCA/RDA (methylated CpG island amplification followed by genomic representational difference analysis), genomic DNA is digested with isoshizomers, one methylation sensitive and the other methylation insensitive, followed by ligation of adaptor sequences and subsequent PCR. These PCR products can be either spotted on nylon filter for subsequent hybridizations with genomic probes, or used for representational difference analysis (RDA) in order to clone methylated sequences (96). Two important findings were made using clones identified by MCA/RDA. Applied to colon cancer and normal colon tissues, it was found that a subset of sequences are progressively methylated with age in normal colon. Furthermore, this study identified two groups of colon cancers: those with more than three (out of seven) cancer-specific loci methylated, and a group where cancer-specific methylation was rare (although age-related methylation comprised the majority found in both groups). A subclassification of colon tumors into those characterized by a positive CpG island methylator phenotype (CIMP+) and those that were negative (CIMP-) was proposed (97).
In order to identify large numbers of methylated CpG islands in lung adenocarcinomas, Shiraishi et al. used methyl-binding columns and enriched methylated sequences (98). These sequences were cloned, size selected and sequenced. In total, 660 independent CpG island sequences were identified after sequencing 6000 clones. Methylation differences between normal lung and lung tumor DNA could be confirmed for about 200 of these sequences (99). While the majority of methylation targets were scattered throughout the whole genome, consecutive CpG island methylation was found in HOX gene clusters on chromosomes 2 and 7 (100).
In a different approach, colon cancer cell lines were treated with 5-aza-2'-deoxycytidine at low doses, or alternatively with TSA, in order to reactivate genes silenced by DNA methylation (101). RNAs were used in gene expression arrays, following an enrichment of reactivated genes. Two distinct groups of genes could be identified: those that were reactivated after with 5-aza-2'-deoxycytidine treatment and those reactivated after TSA treatment. Methylated CpG islands were found in 12 of the 51 genes in the first group. An interesting finding in this study was the identification of aberrant methylation in SFRP1, a member of a gene family whose products antagonize the WNT signaling. The WNT pathway is frequently affected in colon cancer (101).
Differential methylation hybridization (DMH) is an array-based method that can determine the relative methylation status in each CpG island sequence represented on the array (102). Over 5000 small CpG island fragments are spotted onto a nylon membrane or glass slide. Tumor and normal DNA is digested with a methylation-sensitive restriction enzyme. Following PCR amplification and labeling of tumor and normal DNA with two different fluorescent dyes (Cy5 and Cy3), the PCR products from tumor and normal tissues are mixed and hybridized to the CpG island array. Methylated sequences present in the tumor DNA and not in the normal DNA will result in a red signal, whereas sequences methylated in both normal and tumor DNA will result in a mixed yellow signal. DMH has been used to study DNA methylation in breast cancers (103). The microarray data indicated a wide spectrum of methylated sequences in breast cancer, and it was possible to subgroup the breast cancers, into two groups based on their methylation profiles. Poorly differentiated breast tumors showed a higher degree of methylation as compared to moderately or well-differentiated tumors (103).
RLGS is a two-dimensional gel electrophoresis method that enables a genome-wide search for changes in DNA methylation in CpG islands (104). The underlying principle is that genomic DNA is digested with rare-cutting methylation-sensitive restriction enzymes whose restriction sites are preferentially located in CpG islands. Two restriction enzymes are currently used, NotI and AscI. The use of methylation-sensitive restriction enzymes for RLGS analysis makes it possible to scan genomes for differences in methylation patterns. Unmethylated landmark sites are labeled and contribute to the two-dimensional pattern of thousands of spots. RLGS profiles are highly reproducible and thus enable a direct comparison with DNAs from normal tissues and tumors. Landmark sites methylated in the tumor will not contribute to the pattern and can easily be identified by comparison with the normal profile. In a recent study, Costello et al. studied aberrant DNA methylation in multiple human cancers using RLGS (105). In this global analysis, the methylation status of 1184 CpG islands, represented by RLGS fragments, was tested in 98 primary human tumors. The proportion of methylated CpG islands ranged from 0% to almost 10% in these tumor samples. This study also showed patterns of CpG island methylation that were common to several types of tumor (methylated in more than one tumor type) and targets that displayed distinct tumor type specificity (methylated in only one tumor type but never in one of the other tumors studied). Additionally, methylation patterns that characterize certain tumors are not random. This means that either certain sequences become methylated in certain tumors, or methylation occurs at random sites, but selective forces favor the growth of cells with a certain methylation pattern.
It is possible to clone RLGS fragments by using NotI–EcoRV or AscI–EcoRV arrayed genomic libraries (106,107). In this way, several novel genes that become silenced by methylation in lung, testicular and brain tumors as well as in acute myeloid leukemias have been identified (105,108–111). For a review on RLGS and a list of methylated sequences identified so far, see (112). For the majority of human malignancies, changes in CpG island methylation have been reported. However, in a study of seminomatous germ cell tumors by RLGS, almost no CpG island methylation was found, separating this group of tumors from non-seminomatous germ cell tumors (113). The genome-wide scan for changes in DNA methylation in primary head and neck cancers and cervical lymph node metastases showed that although the overall methylation frequencies are low in both types, there are differences in the sequences that become methylated (D.J. Smiraglia et al., submitted for publication). These data indicate that epigenetic progression in head and neck cancers is different from genetic progression, with epigenetic changes being more dynamic.
Hypermethylation identifies novel cancer genes
Several tumor suppressor genes have been identified today, solely based on silencing by promoter methylation. RASSF1A was identified on chromosome 3p21.3, a region commonly deleted in lung cancer. No mutations have been found in RASSF1A; however, promoter methylation is associated with gene silencing in multiple human cancers, including lung (114). SOCS1 was found to be methylated in an RLGS scan of hepatocellular carcinomas and is silenced by methylation. SOCS1 silencing results in constitutive activation of the JAK/STAT pathway and subsequent activation of target genes (115). RUNX3 expression is lost in more than 40% of gastric cancers. Recently, it was shown that loss of RUNX3 expression is due to LOH and promoter hypermethylation rather than mutations in the gene (116). Support for the hypothesis that RUNX3 has tumor suppressor functions came from mice with a homozygous deletion of the gene. These mice show hyperplasia of the gastric mucosa due to stimulated proliferation and suppressed apoptosis (116).
Reversal of epigenetic modifications as a cancer therapy
Epigenetic modifications are reversible, while genetic alterations are not. This feature makes epigenetic modifications a perfect target for therapeutic interventions in cancer patients. It has been shown in cell cultures that demethylating drugs can reverse the silencing of a cancer gene and thus result in growth suppression. Clinical trials are already underway to study the effects of decitabine, 5-aza-2'-deoxycytidine, in hematological malignancies (117) or solid tumors (118). While demethylating drugs possess high toxicity, hope lies in finding agents that interfere with enzymes that modify histones and may act synergistically with demethylating drugs (119).
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A major question in the field of cancer epigenomics remains. What is the cause of aberrant DNA methylation in cancers? One might speculate that changes in the expression levels of the maintenance methyltransferase, de novo DNA methyltransferases or demethylating enzymes are responsible for the observed changes. However, so far no consistent upregulation or downregulation of methyltransferase genes have been reported, and this view may be too simplistic, considering the complexity of epigenetic alterations. The finding that chromatin modifications are involved in the establishment of DNA methylation might suggest that initial changes affect chromatin first and subsequently are transmitted to the DNA level as methylation changes. In addition, it seems that CpG islands are generally protected from hypermethylation, and we need to understand how this protection works and how it breaks down in cancer. An insight into this came recently from Song et al., who demonstrated that hypermethylation of the GSTP1 promoter required that transcription be blocked, and that a low level of DNA methylation already be present (120). Furthermore, if transcription was not blocked, then low-level methylation was erased. One interesting report gives the first clue of what one might expect. Di Croce et al. reported recently that the fusion protein PML–RAR, found as an oncogenic transcription factor in acute promyelocytic leukemias, induces promoter methylation and target gene silencing by recruiting DNA methyltransferases DNMT1 and DNMT3a (121). PML–RAR is a transcriptional regulator of several target genes, including the retinoic acid receptor, RARß2. Expression of PML–RAR causes silencing of RARß2 and methylation of the CpG island in the promoter region. If this mechanism is more general and other oncoproteins also have the potential to recruit methyltransferases to target sequences, one could expect a more genome-wide effect of aberrant DNA methylation. Future studies are needed to show if this proposed mechanism, which links genetic and epigenetic alterations, is a general mechanism.
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ACKNOWLEDGEMENTS |
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I would like to thank Dominic Smiraglia, Laura Rush, Aparna Raval and Anthony Popkie for critical reading of the manuscript. I apologize to all the authors whose work could not be cited due to limitations in size of the review. This work was supported in part by grants P30 CA16058, RO1 CA93548 and RO1 DE13123. C.P. is a Leukemia and Lymphoma Society Scholar.
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FOOTNOTES |
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* To whom correspondence should be addressed at: Division of Human Cancer Genetics, The Ohio State University, 464A Tzargournis Medical Research Facility, 420 West 12th Avenue, Columbus OH, 43210, USA. Tel: +1 6142926505; Fax: +1 6146884761; Email: plass-l{at}medctr.osu.edu
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