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Human Molecular Genetics, 2003, Vol. 12, No. 15 1791-1800
DOI: 10.1093/hmg/ddg204
© 2003 Oxford University Press

Reactivation of the silenced and imprinted alleles of ARHI is associated with increased histone H3 acetylation and decreased histone H3 lysine 9 methylation

Satoshi Fujii1, Robert Z. Luo1, Jiuhong Yuan1, Mitsutaka Kadota4, Mitsuo Oshimura4, Sharon R. Dent2, Yutaka Kondo3, Jean-Pierre J. Issa3, Robert C. Bast, Jr1 and Yinhua Yu1,*

1Department of Experimental Therapeutics, 2Department of Biochemistry and Molecular Biology, and 3Department of Leukemia, The University of Texas, MD Anderson Cancer Center, Houston, TX 77030, USA and 4Division of Molecular and Cell Genetics, Department of Molecular and Cellular Biology, School of Life Sciences, Faculty of Medicine, Tottori University, Nishimachi 86, Yonago, Tottori 683-8503, Japan

Received May 10, 2003; Accepted June 4, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
ARHI has been identified as a maternally imprinted tumor suppressor gene that maps to chromosome 1p31 and whose expression is markedly down-regulated in breast cancer. To explore possible mechanisms that could silence ARHI expression, we have tested the importance of DNA methylation, histone acetylation and histone methylation in regulating ARHI expression. We found that treatment with CpG demethylating agents and/or histone deacetylase inhibitors could reactivate both the silenced and the imprinted alleles of this tumor suppressor gene. Reactivation of ARHI expression by these reagents is related to the methylation status of the CpG islands in the ARHI promoter, especially CpG island II. Chromatin immunoprecipitation assays revealed that histone H3 lysine 9/18 acetylation levels associated with ARHI in normal cells were significantly higher than those in breast cancer cell lines that lacked ARHI expression. Treatment with a CpG demethylating agent and/or histone deacetylase inhibitor could increase ARHI expression in breast cancer cells, with a corresponding increase in histone H3 lysine 9/18 acetylation and decrease in histone H3 lysine 9 methylation.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
DNA methylation and histone deacetylation alter chromatin structure and repress gene transcription. The availability of a specific histone deacetylase (HDAC) inhibitor, trichostatin A (TSA) (1), has permitted evaluation of the role of HDAC in silencing a variety of methylated genes (2). The identification and characterization of several histone methyltransferases (HMTs) (3–5) suggests that the methylation of histone H3 at lysine 9 (H3-K9) also plays an important role in silencing chromatin and in gene transcription. Human (SUV39H1) and fission yeast (Clr4) homologs of Drosophila Su(var)3-9 have recently been shown to encode HMTs that selectively methylate histone H3-K9. The HMT activity of these proteins is conferred through their highly conserved SET domain (3). Recently, this modification has been shown to generate a binding site for heterochromatin protein 1 (HP1) (5). These findings link histone H3-K9 methylation and gene silencing with heterochromatin formation. Histone H3-K9 methylation also inhibits the phosphorylation of serine 10 in histone H3, which may participate in the activation of some genes and is also linked to proper chromosome condensation/segregation (6–8).

In mammals, DNA methylation and histone modification both play an important role in genomic imprinting. Genomic imprinting is a form of epigenetic inheritance that distinguishes maternal and paternal alleles. Imprinting involves preferential expression of a specific parental allele in somatic cells of the offspring. Although the mechanisms of genomic imprinting are not well understood, CpG island methylation is one epigenetic feature that is consistently associated with imprinting (9,10). Almost all imprinted genes have key regulatory elements that are methylated only on one parental allele. Dnmt3L, a protein sharing homology with DNA methyltransferases, Dnmt3a and Dnmt3b, but lacking enzymatic activity, is essential for the establishment of maternal methylation imprints and appropriate expression of maternally imprinted genes (11). However, a few imprinted genes lack differential CpG methylation or are unaffected by disruption of a DNA methyltransferase gene. Moreover, some allele-specific methylation events are not preserved throughout development and other kinds of epigenetic modifications may contribute to imprinting. For example, a number of imprinted genes, including mouse Mash2 (12) and M6P/IGF2R (13), do not exhibit parent-specific DNA methylation. Mash2 imprinting is not disturbed in Dnmt1-/- embryos (14). Some genes exhibit evolutionary conservation of imprinting without showing conservation of differential DNA methylation (15). Histone H3-K9 is methylated on the maternal allele of the Prader–Willi syndrome imprinting center and may be responsible for maternal gametic imprinting of the PWS-IC region (16).

In this study, we report the epigenetic regulation of ARHI, an imprinted tumor suppressor gene. ARHI is one of some 40 imprinted genes that have been discovered across the entire human genome. The maternal allele for ARHI is silenced. Consequently, the gene is expressed in all normal cells only from the paternal allele (17). Loss of ARHI expression can occur through loss of heterozygosity (LOH) of the non-imprinted allele. LOH is observed in 40% of breast, ovarian and pancreatic cancers (18). In a majority of cancers, loss of ARHI expression may occur by epigenetic mechanisms, including DNA methylation and histone modifications.

To gain further understanding of the relationship between DNA methylation, histone acetylation, histone methylation and ARHI expression, we have used a chromatin immunoprecipitation (ChIP) assay to assess the histone acetylation and methylation associated with ARHI expression. We found that alterations of chromatin including histone H3-K9/18 acetylation and histone H3-K9 methylation are associated with the epigenetic regulation of ARHI. The activation of silenced and imprinted ARHI alleles could be achieved using a combination of a CpG demethylating agent and a histone deacetylase inhibitor.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Treatment with a demethylating agent and/or a histone deacetylase inhibitor can reactivate ARHI in breast cancer cells
Recent evidence suggests that DNA methylation-associated gene repression involves the recruitment of histone deacetylases (HDACs) and other chromatin-modifying factors. To examine the relative contributions of DNA methylation and histone deacetylation in ARHI silencing, we have determined the effects of treatment with a demethylating agent, 5-aza-2'-deoxycytidine (5-aza-dC), and a histone deacetylase inhibitor, trichostatin A (TSA), in breast cancer cell lines.

Three potential CpG islands of ~300 bp each were found within the promoter and exons of the ARHI gene. CpG island I is located about 1 kb upstream of the transcription initiation site. CpG island II is near the transcription initiation region and CpG island III is located in the protein encoding region of exon 2 (Fig. 1A). CpG island II is particularly important because it spans the 5'-upstream region of ARHI, including the transcription initiation site and a portion of exon 1 (Fig. 1B). In our previous work (19), three types of DNA methylation have been observed in breast cancer cells. In this study, six breast cancer cell lines with known methylation status were selected for analysis. These cell lines included: two of the type 1 cell lines (MDA-MB-231, MDA-MB-435) in which CpG island II is hypermethylated; two of the type 2 cell lines (BT20, MDA-MB-468) in which CpG island II is hypomethylated, but CpG island I and III are hypermethylated; and two of the type 3 cell lines (MCF-7, SKBr3) in which all three CpG islands are partially methylated (Table 1). ARHI expression was lost or down-regulated in all these six cell lines when compared with normal breast epithelial cells by quantitative real-time RT–PCR (Fig. 2A).



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  		Figure 1. Structure of the human ARHI gene. (A) Structural organization of the ARHI gene and schematic drawing of ChIP products. The ARHI gene consists of two exons and a large intron between exon 1 and exon 2. The solid and open boxes represent the coding and non-coding regions, respectively. Shadowed boxes are CpG islands. For the ChIP assay, fragments amplified by a PCR reaction on each CpG island are shown as small bars. (B) The G+C content per 100 bp and CpG density per 100 bp for the CpG island II spanning upstream and location of the entire exon 1. The TATA box (T in a shadowed box) is located within CpG island II near exon 1 (open box). The locations of four BstUI (shorter bar) and two SacII (longer bar) methylation-sensitive restriction enzyme cleavage sites are indicated.

 

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  		Table 1. Effect of inhibition of methylation and of deacetylation on ARHI expression in breast cancers with different patterns of methylation
 


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  		Figure 2. ARHI expression and reactivation of ARHI after treatment with 5-aza-dC, TSA or a combination of 5-aza-dC and TSA in breast cancer cells. (A) ARHI expression in four normal breast epithelial cells and six breast cancer cell lines evaluated by quantitative real-time RT–PCR. Results are expressed as the copy number/100 ng cDNA on a log scale. (B) ARHI expression was detected by RT–PCR in breast cancer cell lines MDA-MB-435, BT20 and MCF-7. C No treatment control; A, 5-aza-dC; T, TSA; N, no cDNA. (C) ARHI expression was evaluated by quantitative real-time RT–PCR in six breast cancer cell lines. The results of 5-aza-dC treatment in MDA-MB-435, BT20 and MCF-7 were as same as Yuan et al. (19), but from different experiments. Results are expressed as the copy number/100 ng cDNA on a log scale comparing with the average number of four normal breast epithelial cells from (A). Data shown are representatives from three independent experiments. Two representative cell lines of three methylation types are shown (type 1, MDA-MB-435 and MDA-MB-231; type 2, BT20 and MDA-MB-468; type 3, MCF-7 and SKBr3).

 
By using RT–PCR and quantitative real-time RT–PCR, we found that treatment with 5-aza-dC resulted in a mild restoration of ARHI expression in type 1 cells (MDA-MB-231, MDA-MB-435) that exhibited hypermethylation of CpG island II. TSA alone had little impact. When a combination of 5-aza-dC and TSA treatment was used, however, robust expression of ARHI was detected (Fig. 2B and C). Thus, in type 1 cells, inhibition of histone deacetylase alone was not sufficient to restore ARHI, but a combination of DNA demethylation and histone deacetylation inhibition could cooperate to reactivate ARHI synergistically.

In type 2 cells (BT20, MDA-MB-468) CpG island II was hypomethylated while both CpG island I and CpG island III were hypermethylated. 5-aza-dC treatment in this group partially restored ARHI expression. Interestingly, the histone deacetylase inhibitor TSA alone was sufficient to strongly reactivate the expression of ARHI (Fig. 2B and C). This outcome suggests that when CpG island II is not silenced by methylation, histone deacetylation may be a dominant repressor of ARHI expression.

In type 3 cells (MCF-7, SKBr3), all three CpG islands are partially methylated, similar to the condition in normal epithelial cells. Treatment with 5-aza-dC did not affect expression of ARHI. ARHI expression was, however, partially restored by treatment with TSA alone or with a combination of 5-aza-dC and TSA (Fig. 2B and C). Although we could not distinguish which parental ARHI allele was activated by each treatment, the reactivation was weaker in type 3 cells than in other groups (Fig. 2B and C). Normal breast epithelial cells express between 104 and 105 copies/100 ng cDNA of ARHI mRNA. In Figure 2C, treatment with 5-aza-dC and TSA produced a dramatic upregulation of ARHI mRNA in MDA-MB-435, MDA-MB-231, BT20 and MDA-MB-468 cells. In three of the four cell lines, 102–103 copies/100 ng cDNA were observed after treatment. ARHI expression in type 3 cells showed a more modest increase, achieving levels of five to 30 copies/100 ng cDNA.

The results of TSA treatment of BT20 and MCF-7 suggest that chromatin modulation can play an important role in regulating ARHI expression. Results in the MDA-MB-435 cells support the possibility that both CpG island methylation and altered chromatin modulation can regulate ARHI expression when CpG island II is methylated.

Histone H3-K9/18 acetylation is decreased in breast cancer cells
ARHI re-expression after treatment of breast cancer cells with TSA suggested that histone modification, including the level of histone acetylation, may be an important mechanism for ARHI silencing. Our previous work has confirmed that all three CpG islands in normal breast epithelial cells are partially methylated (19). To explore the interaction of CpG island methylation and histone acetylation, we carried out chromatin immunopreciptation (ChIP) assays in three primary cultures of normal breast epithelial cells and in six breast cancer cell lines. In normal breast epithelial cells that had high expression of ARHI, ARHI DNA was associated with acetylated histone H3-K9/18 (Fig. 3). Levels of histone H3-K9/18 acetylation in breast cancer cells that lacked ARHI expression were significantly lower than in normal epithelial cells (t-test, P<0.01 for CpG islands I and II, P<0.05 for CpG island III; Fig. 3).



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  		Figure 3. The level of histone H3 acetylation at lysine 9/18 across the ARHI domain in normal breast epithelial cells and breast cancer cell lines. Acetylation levels are expressed as the ratio of signal intensity of immunoprecipitation product (IP) to input (see Methods). The value was calculated with the Agilent 2100 bioanalyzer software. On the left are the values for individual measurements of normal (NBE018, NBE025 and NBE029) and cancer cells (MDA-MB-435, MDA-MB-231, BT20, MDA-MB-468, MCF-7 and SKBr3) in CpG islands I, II and III. On the right are the average values for three preparations of normal breast epithelial cells and six breast cancer cell lines of each CpG islands respectively (t-test, P<0.01 for CpG islands I and II, P<0.05 for CpG island III). Independent ChIP experiments were repeated at least twice to confirm the reproducibility of results.

 
Treatment with a CpG demethylating agent and/or a histone deacetylase inhibitor is associated with increased histone H3 acetylation in breast cancer cells
To examine the relative contribution of DNA methylation and histone deacetylation to ARHI silencing, we have measured the effect of treatment with 5-aza-dC and/or TSA on the acetylation status of histone H3-K9/18 in cancer cell lines. Figure 4 presents ChIP assays with six breast cancer cell lines. Although these cell lines exhibit different patterns of methylation, treatment with 5-aza-dC and TSA increased histone H3-K9/18 acetylation in all six cell lines. An increase in ARHI expression was also seen with combined 5-aza-dC and TSA treatment in all six cell lines (Fig. 2). In MCF-7 and SKBr3 breast cancer cells, in which ARHI was partially methylated and imprinted, increased expression of ARHI after treatment with both agents was associated with histone H3-K9/18 acetylation of CpG island II. Effects of treatment with 5-aza-dC alone or with TSA alone on histone H3-K9/18 acetylation did not correlate with effects on ARHI expression. An increase in histone H3-K9/18 acetylation and ARHI expression after combined treatment was, however, consistent with the importance of chromatin modification in ARHI regulation.



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  		Figure 4. Levels of histone H3 acetylation at lysine 9/18 across the ARHI domain after treatment with 5-aza-dC (A), TSA (T) and the combination of two agents (A+T) on three CpG islands in six breast cancer cell lines. The histograms were the results of representative ChIP experiments after repetition of independent ChIP experiments two or three occasions. The ratios were calculated as in Figure 3. Cells with no treatment served as the control.

 
Treatment with a CpG demethylating agent can decrease histone H3 methylation at lysine 9
We used ChIP assays to determine whether the level of histone H3-K9 methylation could be altered by treatment with 5-aza-dC and/or TSA. Before treatment, a higher level of histone H3-K9 methylation was detected in MDA-MB-435 and BT20 cells, and a lower level of H3-K9 methylation in MCF-7 cells was observed when compared with MDA-MB-435 and BT20 cells (Fig. 5). In type 1 MDA-MB-435 cells with hypermethylation of CpG island II, 5-aza-dC or a combination of the two agents decreased the level of histone H3-K9 methylation, particularly at CpG island I and to a lesser extent at CpG island II (Fig. 5). TSA treatment also decreased the level of histone H3-K9 methylation. After treatment with 5-aza-dC, TSA or a combination of the two agents, an increase in the level of histone H3-K9/18 acetylation was associated with a decrease in the level of histone H3-K9 methylation in several CpG islands. Although there is a trend, the inverse correlation between histone H3 acetylation and histone H3-K9 methylation is not precise. Decreased histone H3-K9 methylation was observed in each of cell lines associated with at least one CpG island, consistent with an increase in ARHI expression in each cell line (Fig. 2).



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  		Figure 5. Levels of histone H3 methylation at lysine 9 across the ARHI domain after treatment with 5-aza-dC, TSA and a combination of the two agents in three breast cancer cell lines (MDA-MB-435, BT20 and MCF-7). The histograms show the results of representative ChIP experiments after repetition of independent ChIP experiments in duplicate. The ratios were calculated with the same methods used for histone acetylation. Columns showing no treatment (C) or various treatment (A, T and A+T) are indicated in the figures.

 
In type 2 BT20 cells, a higher level of histone H3-K9 methylation was observed at hypermethylated CpG islands I and III than at CpG island II, which was hypomethylated, indicating that DNA methylation might be related to the level of histone methylation. Treatment with 5-aza-dC, TSA or a combination of the two agents resulted in a decrease of histone H3-K9 methylation at CpG islands I and III. In type 3 MCF7 cells, treatment with a combination of the two agents significantly decreased the level of histone H3-K9 methylation at CpG island II.

The ratio of acetylated histone H3-K9/18 to methylated histone H3-K9 (Ac/Me) was calculated. In methylation-dependent type 1 MDA-MB-435 cells, 5-aza-dC treatment can slightly increase the ratio, and a combination of 5-aza-dC and TSA exerted synergistic effects on increasing this ratio. ARHI re-expression (as shown in Fig. 2) was correlated with an increase in this ratio (Fig. 6). A similar correlation was not observed in the methylation independent type 2 BT20 and type 3 MCF-7 cells, consistent with the importance that other transcriptional factors may be involved in ARHI suppression.



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  		Figure 6. The ratios of histone H3 acetylation at lysine 9/18 over histone H3-K9 methylation (Ac/Me) across the ARHI domain after treatments with 5-aza-dC, TSA and a combination of two agents at three CpG islands in breast cancer cell line MDA-MB-435. Columns showing no treatment (C) or various treatment (A, T and A+T) are indicated in the figures.

 
Both the silenced paternal and imprinted maternal alleles of ARHI can be reactivated in breast cancer cells
Taking advantage of a G/A polymorphism at +231 in the coding region of ARHI which eliminates an HhaI restriction site in BT20 cells, we could determine which ARHI allele was affected by treatment. The paternal (G) allele of ARHI could be weakly reactivated by 5-aza-dC and strongly reactivated by TSA, whereas, the imprinted maternal (A) allele could only be reactivated by a combination of 5-aza-dC and TSA (Fig. 7). This assay demonstrated that both the silenced paternal and imprinted maternal alleles of ARHI could be reactivated by the treatment with a histone deacetylase inhibitor and a demethylating agent alone, and in combination in breast cancer cells.



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  		Figure 7. Reactivation of both the silenced paternal and imprinted maternal alleles of ARHI by TSA or a combination of TSA and 5-AZA in BT20 cells. Expression of the coding +231 G/A allele was analyzed by using HhaI digestion of an RT–PCR transcript (right side). Genomic DNA controls (left side) are shown from a heterozygote G/A alleles in BT20. Genomic DNA and RNA were extracted from BT20 cells with or without treatment. cDNA synthesis was, performed by an RT reaction. After PCR amplification using ARHI specific, primers, both genomic DNA and cDNA were digested with HhaI. 326 bp for an HhaI-uncut fragment of maternal A allele; 206 and 120 bp for HhaI-cut two fragments of paternal G allele; C, no treatment; A, 5-aza-dC treatment; T, TSA treatment. A+T, treatment with a combination of 5-aza-dC and TSA.

 
All other breast cancer cell lines lacked this polymorphism and could not be studied in this way. However, using hybrid cell lines that contain maternal and paternal human chromosome 1 we have confirmed these results.

Parental allele-specific histone H3-K9/18 acetylation and H3-K9 methylation are a key regulatory mechanism in ARHI imprinting
To avoid the difficulties in using uncommon polymorphisms to study imprinted genes, Oshimura et al. (20) produced mouse A9 hybrids that contain single alleles of human chromosome 1 of either maternal (A9-1M) or paternal (A9-1P) origin, introduced via microcell-mediated chromosome transfer. This system permits examination of chromatin modification for imprinted genes across genomic DNA. Using A9 hybrids with human chromosome 1, It has been demonstrated that in A9 hybrids the paternal allele of the ARHI gene was not methylated, whereas the silenced maternal allele was methylated at all three CpG islands (19).

ARHI was expressed in A9-1P cells, but not in A9-1M cells by RT–PCR (data not shown). 5-aza-dC alone did not alter expression of ARHI in A9-1P cells, whereas TSA alone produced a remarkable increase in expression. ARHI in A9-1M cells was silenced, even after treatment with 5-aza-dC or TSA alone. Treatment with a combination of 5-aza-dC and TSA, however, led to a partial restoration of ARHI expression (data not shown). These were generally similar to what was seen in BT20 cells.

We observed that the basal level of acetylation of histone H3-K9/18 in A9-1P cells was much higher than that of A9-1M cells (data not shown). As expected, TSA and a combination of the two agents increased the level of histone acetylation in both A9-1P and A9-1M cells.

The level of histone H3-K9 methylation was very low in A9-1P cells (data not shown), indicating that the histone H3 associated with ARHI DNA of the paternal allele was not modified by methylation at lysine 9. Treatment with 5-aza-dC had no effect on the level of histone H3-K9 methylation. In A9-1M cells, levels of histone H3-K9 methylation were much higher than in A9-1P cells. Here, treatment with 5-aza-dC or a combination of 5-aza-dC and TSA markedly decreased the level of histone H3-K9 methylation, particularly at CpG island II. Treatment with TSA alone also resulted in a marked decrease of histone methylation H3-K9 at CpG islands I and II. Thus, parental allele-specific DNA methylation, histone H3 acetylation and K9 methylation may all contribute to ARHI imprinting.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
The loss of expression of several tumor suppressor genes has been related to the epigenetic regulation of both DNA methylation and histone acetylation (1). Most of these genes are regulated by a DNA-methylation-dependent pathway. In the present report, we have found ARHI can be regulated by DNA-methylation-dependent or -independent mechanisms. Changes in levels of acetylation and methylation of histone H3 associated with the ARHI promoter correlated with a loss of ARHI expression and with its re-expression. The re-expression of ARHI has been shown to inhibit cancer cell growth (21).

Several genes can be reactivated by histone deacetylase inhibitors, such as TSA. Some of them, including the estrogen receptor {alpha} gene, FMR1 gene, and human reelin genes, have methylated promotor regions and can still be restored by TSA alone (22–24). 5-aza-dC sequesters DNMT1 in 5-aza-dC-substituted DNA by the irreversible binding of cysteine in the catalytic domain of the DNMT1 enzyme to the 6 position of the cytidine ring (25,26). A study using 5-aza-dC followed by TSA treatment of human colon cancer robustly reactivated multiple methylated genes such as MLH1, TIMP1, CDK2B and CDK2A (1). In our study, treatment with 5-aza-dC and TSA could reactivate ARHI expression in breast cancer cell lines. The activity of these agents, individually and in combination, related to the methylation status of the CpG islands in the ARHI promoter, especially CpG island II. We found that, if CpG island II was hypermethylated (both alleles methylated), a combination of demethylation and histone deacetylase inhibition was necessary to achieve maximal reactivation of ARHI expression. When CpG island II was hypomethylated (both alleles non-methylated), although CpG island I is hypermethylated or partially methylated (i.e. paternal allele is or is not methylated), a histone deacetylase inhibitor alone was sufficient to activate ARHI. When all CpG islands were partially methylated (only the maternal allele methylated), DNA demethylation had no effect on ARHI expression. Inhibition of histone deacetylase could, however, slightly but significantly activate ARHI. Reactivation of ARHI gene by histone deacetylase inhibitor alone or in a combination with a demethylating agent suggests that drugs against these targets deserve evaluation for management of breast cancer.

To understand the mechanism by which chromatin modification alters imprinted gene expression, we carried out experiments to explore the relationship of DNA methylation and histone deacetylation. Our ChIP assay results indicated that histone acetylation was markedly decreased in cancer cell lines relative to that in normal breast epithelial cells. Treatment with 5-aza-dC and/or TSA was associated with an increase in histone acetylation level and in the expression of ARHI, suggesting that histone H3 acetylation was closely related to re-expression of ARHI. The requirement for treatment with a demethylating agent and a histone deacetylase inhibitor in cells that have methylation of CpG island II suggests that DNA methylation and histone deacetylation may be tightly coupled.

Although histone deacetylation has been well studied in transcriptional regulation, recent studies have shown that histone methylation can also affect gene transcription (27–29). The retinoblastoma protein, for example, interacted with SUV39H1 and modified transcriptional repression through histone H3-K9 methylation in cyclin E promotor region (29,30). In the present study, changes in histone H3-K9 methylation correlated with transcriptional control of ARHI. Higher levels of histone H3-K9 methylation were observed in MDA-MB-435 and BT20 cells. In BT20 cells, the level of histone H3-K9 methylation associated with the hypomethylated CpG island II was much lower than that associated with hypermethylated CpG islands I and III. In MDA-MB-435, BT20 and MCF7 cells, a combination of 5-aza-dC and TSA decreased the level of histone H3-K9 methylation on at least one CpG island concomitant with increased ARHI expression. TSA alone could also decrease the level of histone H3-K9 methylation. Recent studies have shown that TSA treatment not only induces histone acetylation but also results in a reduction in H3-K9 methylation (31). In colorectal cancer, histone H3-K9 methylation directly correlates and histone H3-K9 acetylation inversely correlates with DNA methylation of P16, MLH1 and the O6-methylguanine-DNA methyltransferase gene, MGMT (32). Our findings revealed that there was a trend toward an inverse correlation between histone H3-K9/18 acetylation and H3-K9 methylation.

In the analysis of human genomic imprinting, polymorphisms are usually required to distinguish the parental origin of alleles. In the present study, we utilized monochromosomal A9 hybrids (20,33,34) to analyze parental allele-specific histone H3 acetylation at lysine 9/18 and histone H3 methylation at lysine 9 in the imprinted ARHI gene. The expressed paternal allele showed much higher level of histone H3-K9/18 acetylation than the silenced maternal allele on all three CpG islands. The level of histone H3-K9/18 acetylation in A9-1P cells was greatly increased by treatment with TSA or a combination of 5-aza-dC and TSA. These alterations correlated with the expression of ARHI measured by RT–PCR, suggesting that histone H3 acetylation plays a role in increasing expression from the originally expressed paternal allele. The level of histone H3-K9/18 acetylation associated with the maternal allele in A9-1M cells was much lower. Several reports suggest that imprinted genes can exhibit lower levels of allele-specific histone acetylation. Histones associated with the paternally inherited and unexpressed H19 allele were less acetylated than those associated with the maternal allele (35). Parental-specific differences in acetylation of histone H4 were present in the promotor regions of both Igf 2 and H19 genes, with the expressed alleles being more acetylated than the silent alleles (36). Our results are also consistant with the possibility that allele-specific acetylation might produce allele-specific expression of ARHI.

The level of histone H3-K9/18 acetylation in A9-1M cells was also greatly increased by treatments with TSA and a com-bination of 5-aza-dC and TSA. However, only a combination of 5-aza-dC and TSA could mildly reactivate ARHI. El Kharroubi et al. (37) found that expression could be induced from an imprinted allele by treatment with either 5-aza-dC alone or TSA alone. In our studies, the imprinted maternal allele of ARHI could be restored by neither 5-aza-dC, nor TSA alone, but only by a combination of 5-aza-dC and TSA, suggesting that both histone deacetylation and DNA methylation are involved in the control of imprinting.

Xin et al. (16) has recently suggested that histone H3-K9 methylation is a candidate for maternal gametic imprinting in this region (16). Our result showed allele-specific histone H3-K9 methylation in ARHI. In A9-1M cells, levels of histone H3-K9 methylation were much higher than that in A9-1P cells. The level of histone H3-K9 methylation was decreased dramatically by treatment with 5-aza-dC, TSA or a combination of 5-aza-dC and TSA. Thus, histone H3-K9 methylation might be coupled to DNA methylation.

In the present study, we have demonstrated that the inverse correlation between increased acetylation of histone H3 and decreased methylation of histone H3-K9. This modulation was coupled with DNA methylation and the down-regulation of allele specific expression of the imprinted ARHI gene.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Cell lines
Six human breast cancer cell lines including MDA-MB-231, MDA-MB-435, BT20, MDA-MB-468, MCF-7 and SKBr3 were grown in RPMI 1640 media supplemented with 10% fetal bovine serum and 1% L-glutamine. Human mammary epithelial cells, including NBE018, NBE025 and NBE029, were cultured as previously described (17).

Mouse A9 hybrids cell culture
Mouse A9 hybrids containing a single human chromosome 1 of known parental origin were derived at Tottori University (20). A9-1P cells contained a single human paternal allele. A9-1M contained a single human maternal allele. Mouse A9 cells and normal human fibroblast (MK cells) were used as parental controls. Mouse A9 cells and MK cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum and 1% L-glutamine. A9-1P cells were cultured in DMEM supplemented with 10% fetal bovine serum, 1% L-glutamine and 800 µg/ml G418 (Gibco BRL). A9-1M cells were cultured in DMEM supplemented with 10% fetal bovine serum, 1% L-glutamine and 3 µg/ml blasticidin S hydrochloride (Sigma).

Treatment of cells with TSA and 5-aza-dC
TSA was purchased from Wako Pure Chemical Industries Ltd (Osaka, Japan), dissolved in absolute ethanol at a stock concentration of 1 mg/ml (3.3 mM), and stored at -80°C. 5-aza-dC was purchased from Sigma, dissolved in water at a stock concentration of 1 µM and stored at -80°C. As TSA and 5-aza-dC were known to be toxic for cells, preliminary experiments were performed to determine the maximum tolerated concentration for each cell line. From 100 to 400 nM TSA, and 800 nM 5-aza-dC, proved optimal for breast cancer cells. Concentrations of 5-aza-dC and TSA were used as 600 and 200 nM, respectively, for A9 hybrids. Cells were seeded at a low density in a 100 mm tissue culture dish and maintained for a total of 72 h. Cells were incubated for 24 or 48 h prior to treatment with drugs. Mock-treatment with an identical volume of absolute ethanol or water was used as a control. For 5-aza-dC treatment, drug was added after 24 h in culture and cells were incubated for a total 48 h. Culture medium was exchanged every 24 h for 5-aza-dC treatment. For TSA treatment, drug was added to the media after 48 h of incubation and cells were then incubated with TSA for only 24 h. For combined treatment, 5-aza-dC was added after 24 h incubation and at 48 h both drugs were added for an additional 24 h.

RNA and RT–PCR
Total cellular RNA was extracted using the Rneasy mini kit (Qiagen, Valencia, CA, USA). RNA was resuspended in DEPC-treated water and was quantified at OD 260/280. An RT reaction for first-strand cDNA synthesis was carried out with reverse transcriptase (Gibco BRL, Rockville, MD, USA) using 2 µg total RNA and an oligo(dT)12–18 primer (Gibco BRL, Rockville, MD, USA). Specific sense and antisense PCR primers were used for the amplification across the intron from the first exon to the second exon of the ARHI gene, yielding ~250 bp of PCR product. The sequence of the paired primers was as follows: the forward primer was 5'-TCTCTCCGAGCAGCGCA-3' and the reverse primer was 5'-ATCTTCCTGTGGGGCTTGAAGG-3'. To detect the expression of ARHI, a PCR reaction using 2 µl cDNA in a total reaction volume of 25 µl was carried out with an initial denaturation at 95°C for 5 min, followed by 35 cycles at 95°C for 1 min, 60°C for 1 min and 72°C for 1 min. PCR products were resolved by electrophoresis in 2% agarose gels and visualized by ethidium bromide staining.

Quantitative real-time RT–PCR
Quantitative RT–PCR was used to quantify the expression of ARHI after treatment with 5-aza-dC and/or TSA. The cDNA used for real-time PCR was the same as that used for the RT–PCR reaction. PCR reactions included 2 µl cDNA prepared from 2 µg of total RNA, a forward primer (NY2P1, 5'-TCTCTCCGAGCAGCGCA-3'), a reverse primer (NY2P2, 5'-TGGCAGCAGGAGACCCC-3'), labeled probe (5'-TGTCTTCTAGGCTGCTTGGTTCGTGCC-3'; 5'-fluorescent label, 6-FAM; 3'-fluorescent label, 6-carboxytetramethylrhodamine), 2 µl of reverse transcriptase reaction mixture, and 12.5 µl of Master Mix for the ABI PRISM 7700 Sequence Detection System (Perkin-Elmer) as described in the manufacturer's protocol. The level of ARHI re-expression was evaluated by 2-{Delta}{Delta}CT (according to the User Bulletin no. 2 from the manufacturer) and normalized to a human GAPDH endogenous reference standard relative to a calibrator. The level of ARHI expression was also evaluated by copy number per 100 ng cDNA.

Chromatin immunoprecipitation assay
The ChIP assays for histone acetylation and histone methylation were performed as described by Chandee et al. (38) with minor modifications. Briefly, cells were treated with 1% formaldehyde for 8 min to cross-link histones to DNA. After washing, the cell pellets were resuspended in 500 µl lysis buffer [150 mM NaCl, 25 mM Tris–HCl (pH 7.5), 5 mM EDTA, 1% Triton X-100, 0.1% sodium dodecyl sulfate, 0.5% deoxycholic acid] and sonicated with a Branson Sonicator 250 to yield an average DNA size of 500 bp. Sonicated extracts were subsequently clarified by centrifugation. The lysate (500 µl) was then divided into three fractions: the first and second (230 µl each) were diluted in 260 µl of lysis buffer, and the third (40 µl) used for an input control. The first lysate was incubated with 10 µl antibodies specific to either acetylated histone H3 at lysine 9 or 18 (39) or methylated histone H3-K9 (Upstate Biotechnology) at 4°C overnight on a 360° rotator. The second lysate was incubated with mock preimmune sera (10 µl) as a negative control. To collect immunoprecipitated complexes, 35 µl of sepharose A beads (Pharmacia) were added and rotated for 1 h at 4°C. Sepharose beads with complexes were treated with RNaseA (50 µg/ml) for 30 min at 37°C and with 3.75 µl of 20 mg/ml proteinase K for each sample overnight. The DNA–protein cross-links in the chromatin complexes were reversed by heating at 65°C for 6 h. Proteins were removed by phenol–chloroform extraction and DNA was precipitated with ethanol.

The presence of different regions of the ARHI gene in the immunoprecipitates was detected by PCR using gene-specific primers as described below. Individual ChIP assays were repeated at least twice to confirm the reproducibility of the PCR based experiment. PCR amplification was performed using either immunoprecipitated DNA, a control without antibody or a 1 : 150 dilution of input that had not been immunoprecipitated. Preliminary PCR reactions were performed to determine the optimal PCR conditions to assure linear amplification of DNA. Samples were heated initially for 5 min at 95°C followed by 32 cycles of 95°C for 1 min, 59°C for 1 min and 72°C for 2 min. After cycling, samples were heated for an additional 7 min to permit extention. The primer pairs were as follows for each of the three CpG islands: CpG I sense, 5'-GTGCGCAGCTTTCAATGCATC-3'; CpG I antisense, 5'-TCTCACTCCTTGGGCGCAATG-3'; CpG II sense, 5'-TCGATTGTTGTAGATGCCAAG-3'; CpG II antisense, 5'-AGACTTACCTTTCTCGGAGGC-3'; CpG III sense, 5'-GGGAGAACTTGTGCACGTATC-3'; CpG III antisense, 5'-GTGATGTGCAGGGAAAGCACA-3'. PCR products were electrophoresed on a 2% agarose gel, stained with ethidium bromide, and photographed. To evaluate the level of histone acetylation and histone methylation in each immunoprecipitation, the ratio was determined by quantifying the intensities of the PCR product in immunoprecipitated DNA versus 1 : 150 diluted input DNA (total chromatin) amplified by PCR in a linear range. The ratio was quantified using the DNA 500 assay with the Agilent 2100 bioanalyzer and DNA chips for electrophoresis (Agilent Technologies, USA).


   ACKNOWLEDGEMENTS
 
We gratefully acknowledge Dr Wei Hu for technical assistance in quantitative real-time RT–PCR and Madelene Coombes for technical assistance in ChIP assay. This work was supported by National Institutes of Health Grants CA 64602 and CA 80957, Susan G. Komen Breast Cancer Foundation and a Grant for a Research Worker Abroad from the Ministry of Education, Science, Sports and Culture of Japan.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Department of Experimental Therapeutics, The University of Texas, MD Anderson Cancer Center, Box 354, 1515 Holcombe Blvd, Houston, TX 77030, USA. Tel: +1 7137923790; Fax: +1 7137452107; Email: yyu{at}mdanderson.org Back


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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