5-Aza-2′-deoxycytidine stress response and apoptosis in prostate cancer
© Springer-Verlag 2011
Received: 7 August 2010
Accepted: 15 December 2010
Published: 15 January 2011
While studying on epigenetic regulatory mechanisms (DNA methylation at C-5 of –CpG– cytosine and demethylation of methylated DNA) of certain genes (FAS, CLU, E-cadh, CD44, and Cav-1) associated with prostate cancer development and its better management, we noticed that the used in vivo dose of 5-aza-2′-deoxycytidine (5.0 to 10.0 nM, sufficient to inhibit DNA methyltransferase activity in vitro) helped in the transcription of various genes with known (steroid receptors, AR and ER; ER variants, CD44, CDH1, BRCA1, TGFβR1, MMP3, MMP9, and UPA) and unknown (DAZ and Y-chromosome specific) proteins and the respective cells remained healthy in culture. At a moderate dose (20 to 200 nM) of the inhibitor, cells remain growth arrested. Upon subsequent challenge with increased dose (0.5 to 5.0 μM) of the inhibitor, we observed that the cellular morphology was changing and led to death of the cells with progress of time. Analyses of DNA and anti-, pro-, and apoptotic factors of the affected cells revealed that the molecular events that went on are characteristics of programmed cell death (apoptosis).
KeywordsEpigenome CpG hypermethylation DNMT 5-Aza-2′-deoxycytidine Apoptosis Prostate cancer
Overexpression of DNA (cytosine-5-carbon) methyltransferases and hypermethylation of CpG islands at the regulatory regions of certain genes (for example, ER, CDH1, CD44, Casp3, GSTP1, MDR1, FAS, RASSF1A, P15INK4B, and P16INK4A) and global genome-wide hypomethylation are well known to be associated with prostate and other multiple cancer development and metastasis (Graff et al. 1995; Ferguson et al. 1997; Li et al. 2000; Szyf et al. 2000; Li et al. 2001; Nojima et al. 2001; Patra et al. 2001; Dasari et al. 2002; Jones and Baylin 2002; Patra et al. 2002; Sasaki et al. 2002; Chen et al. 2003; Jaenisch and Bird 2003; Patra et al. 2003; Patra 2008a; Patra et al. 2008a; Patra and Szyf 2008; Patra and Bettuzzi 2009). The CpG sites in these gene promoter regions are rarely methylated in normal cells except in, e.g., inactivated X-chromosome and imprinted genes. Some non-regulatory site CpG island methylation has no direct effect on gene activity (Patra 2008a; Patra et al. 2008a; Patra and Szyf 2008). It is now revealed that abnormal methylation of CpG islands is not restricted to cancer cells but can also occur during aging and during early stages of tumor development (Patra et al. 2008a). While the co-existence of genome-wide hypomethylation and site-selected hypermethylation are well documented by chemical analyses of bases from total genomic DNA and gene-specific DNA segment, the specific mechanisms at the level of enzymes, co-substrates, and repressor proteins are subject to immense interest of current investigations (Ferguson et al. 1997; Bhattacharya et al. 1999; Cervoni et al. 1999; Ramchandani et al. 1999; Fuks et al. 2000; Ng et al. 2000; Peterson and Logie 2000; Rhee et al. 2000; Szyf et al. 2000; Misteli 2001; Ohki et al. 2001; Patra et al. 2001; Cervoni et al. 2002; Di Croce et al. 2002; Jones and Baylin 2002; Patra et al. 2002, 2003; Pogribny and James 2002; Jaenisch and Bird 2003; Patra 2008a; Patra et al. 2008a; Patra and Szyf 2008; Patra and Bettuzzi 2009).
5-Aza-2′-deoxycytidine (AzadC), when incorporated into the genomic DNA in sites to be occupied by cytosine during replication, is best known as an inhibitor of DNA methyltransferases (DNMTs—DNMT1, DNMT3A, and DNMT3B) through covalent adduct formation with the enzyme (Patra and Bettuzzi 2009). Since the methylation target for the mammalian maintenance methyltransferase, DNMT1, is CpG in hemi-methylated sites, significant inhibition of DNA synthesis, even due to complete lack of repair of DNMT1–AzadC adducts, does not occur for at least two cell cycles. In contrast, DNMT1 becomes bound to DNA and inactivated as soon as AzadC is incorporated into CpG sites opposite to methylated CpG sites on the template strand, i.e., within a few hours of initiating treatment with AzadC. This in turn leads to rapid passive loss of methylation. There is plenty of evidence that treatment of cancers with AzadC leads to reactivation of function in tumor cell lines in which one copy of a gene for a tumor suppressor, a cell cycle regulator, or a DNA repair enzyme is muted and the complementary is normal (wild type) but inactivated by methylation (Graff et al. 1995; Li et al. 2000; Wijermans et al. 2000; Li et al. 2001; Nojima et al. 2001; Christman 2002; Patra et al. 2002; Chen et al. 2003; Claus et al. 2005; Yoo and Jones 2006). This suggests that loss of methylation induced by AzadC treatment can lead to reactivation of the same gene whose inactivation was programmed/selected during development of a specific tumor and has stimulated interest in revisiting the use of AzadC and analogs in therapeutic intervention of cancer (Wijermans et al. 2000; Christman 2002; DeSimone 2004; Claus et al. 2005; Lavelle et al. 2006; Yoo and Jones 2006; Lavelle et al. 2008; Patra and Bettuzzi 2009). But along with expression of desired curative genes (for example, ER, AR, and CDH1), some unwanted genes that code proteins enhancing metastasis (lipid raft associated genes, UPA and MMPs) could be expressed even at a low non-toxic dose of the drugs, thus creating complications in the use of this chemotherapeutic agent (Sato et al. 2003; DeSimone 2004; Lavelle et al. 2006; Hellebrekers et al. 2007; Patra and Bettuzzi 2009). Since the 1970s, there have been many studies on the toxicity of AzadC. The early studies were mostly devoted to understanding various toxic effects on the basis of physiology and genetics (Christman 2002). The present study was designed to investigate the systematic molecular and epigenetic effects of 5-azadC on the DNMT activity and expression, and on cell growth. The data presented in this work provide strong evidence that the cells died by epigenetic stress-induced apoptosis when challenged with higher dose (∼5.0 μM, which is far above the DNMT inhibitory concentration) of the drug 5-azadC. The results show that Bcl2 and DNMT1 genes are not repressed; rather, they overexpressed dose dependently of 5-azadC when supplemented with the cell culture media. DNMT1 enzyme activity was reduced, perhaps due to adduct formation with DNA containing 5-azadC replacing cytosine. Gradual increase of 5-azadC in cell culture medium caused dose-dependent overexpression of apoptosis genes like Bak, Bax, and caspase-3, including many other genes in TSUPr1 and DuPro cell lines (Table 3).
Cell culture and drug(s) treatment
Prostatic cancer cell lines of human origin PC3, TSUPr1, and DuPro were purchased from American Type Culture Collection (Manassas, VA, USA). Cell lines were cultured and maintained in RPMI 1640 supplemented with 10% (v/v) fetal bovine serum, 2 mM l-glutamate, and 0.1 μM of penicillin and streptomycin. Media and supplements were obtained from the University of California at San Francisco Cell Culture Facility. For drug treatment, exponentially growing cells were seeded at a density of ∼106 cells/75 cm2 flask. After 6 h, the cells were treated at different concentrations (nanomolar to micromolar) of 5-azadC (Sigma Chemical Co., St. Louis, MO, USA) and marked as zero “0” time. Trichostatinn A was dissolved in PBS and diluted with the required volume of the medium, and cells were also treated at different concentrations (4, 10, and 50 nM) of MG-132 (Calbiochem, San Diego, CA, USA) inhibitor of proteasome, dissolved in DMSO and diluted with the required volume of medium for 8–12 h prior to harvest for further investigations. All cells were harvested after 96 h unless otherwise mentioned.
Assays of enzymes
The DNMT enzymatic activity was measured following a previous protocol as standardized in our laboratory (Patra et al. 2001, 2002, 2003). Poly•dI–dC duplex or poly•dG–dC duplex was used as substrate and S-adenosylmethionine (tritiated methyl) as methyl group donor. All assays were performed in duplicates in three sets of independent experiments. Background levels were determined in assays in which the template DNA was omitted. Statistical analyses were performed using Student’s t test.
Caspase-3-like activity was measured according to the kit supplied by Calbiochem. Caspase-3 assay kit employs the colorimetric substrate DEVD-pNA that upon cleavage exhibits increased absorption at 405 nm. A caspase-3 inhibitor (Ac-DEVD-CHO) is also used as a prototypic control inhibitor. The DEVD amino acid sequence is derived from the caspase-3 cleavage site in PARP. All the kit components were thawed and immersed in an ice bath until use. Briefly, caspase-3 inhibitor I was warmed to room temperature. In a separate microcentrifuge tube, it was diluted in assay buffer (1:200). Caspase-3 (15 μl, approximately 30 U) diluted in assay buffer (1:50) was used as control. Caspase-3 was not used in assay buffer to blanks. Caspase-3 inhibitor (20 μl) was added in triplicate tubes. Reaction was started by adding 200 μM (final concentration) caspase-3 substrate I and pre-equilibrated to assay temperature at 25°C and measured in a colorimeter/microplate reader.
Histone deacetylase (HDAC) enzymatic activity was measured using fluorescent amide, N-(4-methyl-7-coumarinyl)-N-α-(tert-butyloxy-carbonyl)-N-Ω-acetyllysinamide (MAL) (Calbiochem) as a potential substrate following the published protocol of ours and other laboratories (Patra et al. 2001; Cervoni et al. 2002) as well as using a radioactive kit (Upstate Biotechnology). The background fluorescence was kept significantly low using filters, and nanomolar (nM) concentration of MAL was used to avoid interference (inner-filter effects). In the presence of trichostatin A (Calbiochem), a potent inhibitor of HDAC (Taunton et al. 1996; Hoffmann et al. 1999), no activity was observed (see also Patra et al. 2001, 2003). Statistical analyses were performed using Student’s t test.
Nucleic acid extraction and reverse transcription–PCR
Total RNA was extracted using TRI Reagent (Molecular Research Center, Inc., Cincinnati, OH, USA). Cytosolic and nuclear RNA were prepared using TRI reagent after control lysing of cells and separation by centrifugations of those contents. RNA (1–2 μg) was reverse transcribed using random hexamer primers and Superscript II, reverse transcriptase (Life Technologies Inc., Gaithersburg, MD, USA) in a 25-μl reaction volume. cDNA was amplified by PCR using primers specific for the genes Bcl2, Bak, Bax, Bcl-XL, and caspase-3. β-Actin, glyceraldehyde-3-phosphatedehydrogenase (GAPDH), and histone 4 (H4) genes were also amplified as internal controls to ensure high quality. Primer sequences specific for β-actin gene (TCTACAATGAGCTGCGTGTG, sense; ATCTCCTTCTGCATCCTGTC, antisense), GAPDH gene (GAAGGTGAAGGTCGGAGTC, sense; GAAGATGGTGATGGGATTTC, antisense), and H4 gene (CAACATTCAGGGCATCACCAA, sense; CCCGAATCACATTCTCCAAGAA, antisense) were used and resulted in 682-, 226-, and 131-bp products, respectively. The PCR sample mixtures, in a 10-μl volume, contained 1× PCR buffer (Sigma), 0.2 mM of each dNTP (Sigma), 4 ng of TaqStart antibody (Clonetech, Palo Alto, CA, USA), 0.2 μM primers, and 0.5 μl RedTaq DNA polymerase (Sigma). PCR reactions were performed in a PTC-200 thermal cycler (MJ Research, Watertown, MA, USA) at 94°C for 1 min, 26 cycles at 94°C for 20 s, 57°C for 20 s, and 72°C for 30 s, followed by an extension step at 72°C for 5 min. The PCR products were electrophoresed through a 1.2% agarose gel containing ethidium bromide and were visualized by UV detection.
Prostatic cancer cell lines were grown in chamber slides and stained as our standardized protocol (Patra et al. 2001, 2002, 2003). In brief, sub-confluent cells were fixed by 10% formalin and were permeabilized by 0.1% triton X-100 in PBS. The endogenous peroxidase activity was blocked by incubation in 5% H2O2 in methanol for 20 min. The cells were pre-blocked using Ultra V block (Lab Vision Corporation, Fremont, CA, USA) for 10 min and incubated for overnight at 4°C with goat polyclonal antibody against caspase-3 (Santa Cruz Biotech, CA, USA) New Zealand white rabbits polyclonal antibody against DNMT1 peptide (I0015801K; Research Genetics, Inc., Huntsville, AL, USA; or met-cat 2) and goat polyclonal antibody against Bcl2, Bak, Bax, BclXs, and MBD1 (Santa Cruz Biotech). The antibody for HDAC1 (1:100), DNMT1 (1:1,000), and others was diluted according to the manufacturer’s instructions (and several titration) with primary antibody dilution buffer (Biomeda, Foster City, CA, USA). The cells were washed in PBS then incubated with anti-goat biotinylated secondary antibody (Santa Cruz Biotech) for 30 min followed by another wash and incubation with HRP-conjugated streptavidin. Finally, reactions were visualized by incubation with DAB (substrate and chromogen) and counterstaining with Harris hematoxylin. For negative control, cells were incubated overnight with dilution buffer (no primary antibody).
Cells were permeablized with 0.1% Triton for 2 min, fixed for 10 min in 4% paraformaldehyde in PBS, and stored in 70% ethanol before assay. Slides were equilibrated in terminal transferase (Roche) and incubated for 60 min with TdT (200 U/ml) and biotin-16-dUTP (10 μM; Roche). Following two washes with 4× SSC, TUNEL-positive cells were detected with avidin–Texas Red (1:100; Jackson).
In vitro DNMTase, HDACs, and caspase-3 activity: inhibition of DNMT activity is not sufficient for induction of caspase-3 activity
Comparison of % activity (average of three sets of independent experiments) of the enzymes without and with increasing concentration of drug
2.3 ± 1.73
5.67 ± 3.46
27.5 ± 8.40
41.49 ± 7.0
Effect of drugs (5-azadC, DEVD-CHO, and MG-132) on cell growth
Inactivation of DNMT1 causes DNA hypomethylation and activation of caspase-3, Bak, Bax, and Bcl-XL and overexpression of Bcl2 and DNMT1
Control (no drug treatment)
5-Aza-2′-deoxycytidine (250 nM)
Re-expressed genes in prostate cancer cells by application of 5-azadC in culture
Low-dose (<10 nM) genes
High-dose (>20 to 500 nM) genes
Steroid receptors (SR—AR and ER)
Methyl–CpG–DNA binding domain protein(s)
Cancer of prostate
This work was done during the authors’ (AP and SKP) stay at the University of California at San Francisco, California, USA and supported by the National Institute of Health Grants DK47517, AG-16870, and CA64872, Veterans Affairs Medical Center, and by the NCIRE. The manuscript was written (by SKP) during his stay at the McGill University, Montreal, Canada and University of Parma, Italy.
Conflict of interest
The authors declare no conflict of interest.
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