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Molecular Cancer Research 5, 1083-1098, October 1, 2007. doi: 10.1158/1541-7786.MCR-06-0432
© 2007 American Association for Cancer Research
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DNA Damage and Cellular Stress Responses

Human 8-Oxoguanine DNA Glycosylase Suppresses the Oxidative Stress–Induced Apoptosis through a p53-Mediated Signaling Pathway in Human Fibroblasts

Cha-Kyung Youn1,2, Peter I. Song1,5, Mi-Hwa Kim1,2, Jin Sook Kim6, Jin-Won Hyun7, Sang-Joon Choi3, Sang Pil Yoon8, Myung Hee Chung9, In-Youb Chang1,4 and Ho Jin You1,2

1 Korean DNA Repair Research Center, Departments of 2 Pharmacology, 3 Obstetrics and Gynecology, and 4 Anatomy, Chosun University School of Medicine, Gwangju, South Korea; 5 Department of Dermatology, University of Colorado Health Sciences Center, Denver, Colorado; 6 Department of Herbal Pharmaceutical Development, Korea Institute of Oriental Medicine, Daejeon, South Korea; 7 Department of Biochemistry, College of Medicine, Cheju National University, Jeju-do, South Korea; 8 Department of Anatomy, College of Medicine, Seonam University, Jeollabuk-Do, South Korea; and 9 Department of Pharmacology, School of Medicine, Seoul National University, Seoul, South Korea

Requests for reprints: Ho Jin You, Korean DNA Repair Research Center, Bio Engineering BD 2F, Department of Pharmacology, Chosun University School of Medicine, 375 Seosuk-Dong, Gwangju 501-759, Republic of Korea. Phone: 82-62-230-6337; Fax: 82-62-230-6586. E-mail: hjyou{at}chosun.ac.kr


   Abstract
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 Notes
 Abstract
Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Human 8-oxoguanine DNA glycosylase (hOGG1) is the main defense enzyme against mutagenic effects of cellular 7,8-dihydro-8-oxoguanine. In this study, we investigated the biological role of hOGG1 in DNA damage–related apoptosis induced by hydrogen peroxide (H2O2)–derived oxidative stress. The down-regulated expression of hOGG1 by its small interfering RNA prominently triggers the H2O2-induced apoptosis in human fibroblasts GM00637 and human lung carcinoma H1299 cells via the p53-mediated apoptotic pathway. However, the apoptotic responses were specifically inhibited by hOGG1 overexpression. The p53–small interfering RNA transfection into the hOGG1-deficient GM00637 markedly inhibited the H2O2-induced activation of p53-downstream target proteins such as p21, Noxa, and caspase-3/7, which eventually resulted in the increased cell viability. Although the cell viability of hOGG1-knockdown H1299 p53 null cells was similar to that of the hOGG1 wild-type H1299, after the overexpression of p53 the hOGG1-knockdown H1299 showed the significantly decreased cell viability compared with that of the hOGG1 wild-type H1299 at the same experimental condition. Moreover, the array comparative genome hybridization analyses revealed that the hOGG1-deficient GM00637 showed more significant changes in the copy number of large regions of their chromosomes in response to H2O2 treatment. Therefore, we suggest that although p53 is a major modulator of apoptosis, hOGG1 also plays a pivotal role in protecting cells against the H2O2-induced apoptosis at the upstream of the p53-dependent pathway to confer a survival advantage to human fibroblasts and human lung carcinomas through maintaining their genomic stability. (Mol Cancer Res 2007;5(10):1083–98)


   Introduction
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 Notes
 Abstract
 Introduction
Results
 Discussion
 Materials and Methods
 References
 
Various cellular metabolites and exogenous DNA-damaging materials routinely induce cellular DNA damage, which, unless promptly and properly repaired, can cause simple base or more complex changes including deletions, fusions, translocations, or aneuploidy, resulting in cancer and a variety of genetic disorders (1).

Cellular responses induced by DNA damage include the activation of several distinct biochemical pathways (2-5). First, various DNA repair enzymes are activated to recognize and eliminate the damage. Second, DNA damage stimulates the specific mechanism related to cell cycle checkpoints that arrest cell cycle progression and aid in cellular survival under most circumstances. Third, apoptosis is stimulated by DNA damage to eliminate heavily damaged or seriously deregulated cells. Although these DNA damage–induced biochemical pathways function independently under certain circumstances, there are extensive interactions between these reactions. For example, the regulatory factors in the checkpoints of cell cycle serve not only to delay the cell cycle but also to mediate DNA repair, both directly and indirectly (3-6). However, the precise mechanisms of assessing DNA damage both quantitatively and qualitatively so as to choose between mediating DNA repair and apoptosis are not completely understood.

Reactive oxygen species are produced as by-products of cellular metabolism as well as through exposure to UV, ionizing radiation, and environmental carcinogens (7). The reactive oxygen species react with DNA to produce a myriad of cytotoxic and mutagenic base lesions (7). Among the oxidative lesions, 7,8-dihydro-8-oxoguanine (8-oxoG) is the major base damage produced by reactive oxygen species (3). Unlike normal guanine, 8-oxoG has the propensity to mispair with adenine during DNA replication and thereby gives rise to G:C to T:A transversion mutations (8). Oxidatively modified bases, such as 8-oxoG, are mainly repaired through the base excision repair pathway, the first steps being the recognition and excision of the damaged base by a specific DNA glycosylase. The major mammalian enzyme for removing 8-oxoG from DNA is 8-oxoguanine-DNA glycosylase (OGG1), which is a bifunctional enzyme with both 8-oxoG excision activity and weak AP lyase strand incision activity at the abasic sites (9). Following the excision of 8-oxoG by OGG1, the resultant abasic site is further processed in sequential steps by several enzymes to complete repair (2). OGG1 plays important roles in eukaryotes by preventing the accumulation of oxidative DNA damage, in both the nuclear and mitochondrial genomes, thereby suppressing carcinogenesis and cell death (10).

The suppression of oxidative DNA repair activity has potential drawbacks; one is that the incomplete repair might result in the accumulation of mutagenic lesions in the cellular DNA. This leads to illness, death of cells, and unstoppable excessive cell division resulting in cancer and acceleration of aging process (10, 11). The human OGG1 (hOGG1) gene is found on chromosome 3p26.2, and its allelic deletions in this region frequently occur in a variety of human cancers (9). It is somatically mutated in some cancer cells and is highly polymorphic between human population groups (12). The accumulation of 8-oxoG is likely to increase dramatically in patients with various neurodegenerative diseases such as Parkinson's disease (13), Alzheimer's disease (14), or amyotrophic lateral sclerosis (15), which are associated with the progressive loss of cells. The hOGG1 level was also found to be lower in the orbitofrontal gyrus and the entorhinal cortex in Alzheimer's disease patients than in control cases. The accumulation of 8-oxoG increased in a majority of large motor neurons in the amyotrophic lateral sclerosis cases with decreased hOGG1 expression (14). Furthermore, several reports have shown a correlation between the induction of apoptosis by oxidative stress and the repair of oxidative DNA lesions (16-18). Thus, mutations and deletions in the cellular DNA, which could arise from unrepaired oxidative DNA lesions, have been linked to the development of apoptosis. However, the exact mechanism by which oxidative stress–mediated DNA damage causes cell death is unknown.

In this study, we investigated the effect of hOGG1 on oxidative DNA damage–related apoptotic responses, including changes of apoptotic effector caspase activity, p53 phosphorylation, and the expression pattern of its upstream or downstream proteins, together with genomic DNA instability followed by the oxidative DNA damage in human fibroblasts.


   Results
Top
 Notes
 Abstract
 Introduction
 Results
Discussion
 Materials and Methods
 References
 
Suppression of hOGG1 Gene Expression Enhances the Cytotoxic Effects of Hydrogen Peroxide in Human Fibroblast GM00637
Semiquantitative reverse transcription-PCR was done using hOGG1-specific synthetic primers, as described in Materials and Methods, to determine if the constitutive expression of human OGG1 is specifically inhibited by hOGG1-small interfering RNA (siRNA) in cultured human fibroblast GM00637 cells. Two transfectants of hOGG1-silencing GM00637 cells blocked the normal expression of hOGG1 mRNA up to 80% to 90%, compared with that of control-siRNA transfectants (Fig. 1A and C ). Western blot analysis was carried out to determine whether the inhibition of hOGG1 mRNA corresponded to that of hOGG1 protein level. As shown in Fig. 1B and C, hOGG1 protein expression was 60% to 65% inhibited by hOGG1-siRNA transfection. To determine the functional significance of the inhibited hOGG1 expression, we prepared the cell lysates from parental cells and siRNA transfectants with hOGG1-siRNA and its control-siRNA, and examined their abilities to cleave 8-oxoG using a 21-mer oligonucleotide containing a single 8-oxoG at nucleotide 13 by the glycosylase activity assay, as described in Materials and Methods. As shown in Fig. 1D, the hOGG1-siRNA transfectants showed the inefficient cleavage of the 8-oxoG:C substrates compared with the parental cells or the control-siRNA transfectants, demonstrating that hOGG1-siRNA causes the reduced expression of hOGG1 protein and the decrease of DNA damage repair activity in human fibroblast GM00637 cells. We next investigated the hOGG1 effect on cellular cytotoxicity in parental cells and hOGG1-siRNA transfectants in response to hydrogen peroxide (H2O2) oxidative stress. The cell viability of hOGG1-siRNA–transfected GM00637 was significantly lower than those of control-siRNA transfectants and parental cells (Fig. 1E), indicating that the inhibition of hOGG1 expression reduces the cell viability of fibroblasts in response to oxidative stress. To determine if hOGG1 depletion has any effect on H2O2-induced apoptosis, the nuclei of GM00637 cells were stained with a propidium iodide dye for flow cytometric detection. The control-siRNA–transfected cells showed the apoptotic sub-G1 DNA content of 4.12% to 12.35%, when they were treated with 0 to 60 µmol/L H2O2. In contrast, the hOGG1-siRNA transfection increased the proportion of apoptotic sub-G1 DNA content from 4.97% to 24.17% at the same oxidative stress (Fig. 1F). This shows that the suppression of hOGG1 expression triggers the increased apoptotic responses of human fibroblasts in response to H2O2.


Figure 1
Figure 1
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  		FIGURE 1. Knockdown of hOGG1 expression by hOGG1-siRNA transfection sensitizes human fibroblast GM00637 cells to H2O2-induced apoptosis. The human fibroblast GM00637 cells were stably transfected with the control-siRNA and hOGG1-siRNA as described in Materials and Methods. The expression of hOGG1 was then analyzed by reverse transcription-PCR using hOGG1-pecific synthetic primers (A) and by Western blot using anti-hOGG1 polyclonal antibodies (B). Data were normalized with GAPDH and {alpha}-tubulin expression, respectively. C. Graphical plot based on A and B. Columns, mean; bars, SD. Statistically significant differences in OGG1 expression were determined by the ANOVA with P values given for both the overall (P) and the pair-wise comparisons as indicated by asterisks (*, P < 0.05). D. The OGG1 enzyme activities were analyzed by 8-oxoG nicking assay using the parental-, control-siRNA–, and two of hOGG1-siRNA–transfected cell lysates as described in Materials and Methods. Purified hOGG1 and buffer alone served as positive and negative controls, respectively. Arrowhead, percent ratios of DNA cleavage products (13-mer), which are analyzed by the AxioVision Rel.4.6 Imaging System (Carl Zeiss International). E. Parental-, control-siRNA–, and hOGG1-siRNA–transfected cells were cultured to 70% to 80% confluence, and treated with the indicated concentration of H2O2 for 1 h. The medium was then changed to a fresh one, and the extent of cell death was assayed by trypan blue exclusion 24 h after the treatment. Cell viability was determined as the percentage of total cell number that remained unstained. Points, means from six separate experiments; bars, SD. F. Apoptotic sub-G1 DNA contents were estimated by FACScan analysis after treatment with the indicated H2O2 concentration under the same experimental condition as in E. Results are representative of three independent experiments. G. DNA damages of control-siRNA– and hOGG1-siRNA–transfected cells induced by 30 µmol/L H2O2 were analyzed by single-cell gel electrophoresis (comet assay) as described in Materials and Methods. The comet tail moment of 80 cells for each experimental condition was quantified with Komet 5.5 (Andor Technology). Columns, mean; bars, SD.

 
To determine whether the elevated apoptosis is arising from DNA breaks induced by H2O2-oxidative stress, we analyzed the genotoxic potential of the oxidative stress on hOGG1-deficient GM00637 cells by use of the single-cell gel electrophoresis assay (comet assay). Comet assay is specific for detecting various forms of DNA damage, such as single- and double-strand breaks, in virtually any eukaryotic cell. Double-strand breaks within DNA damages can be detected by merely subjecting them to electrophoretic mobility at the neutral pH (e.g., pH 7-8); in contrast, single-strand breaks can be detected at alkali at pH greater than pH 12.1 (19). As indicated in Fig. 1G, 30 µmol/L H2O2 significantly increased both single and double DNA strand breaks of hOGG1-siRNA–transfected cells compared with the control cells. Therefore, we suggest that hOGG1 suppression in human fibroblasts triggers the increased genotoxicities, including both DNA single- and double-strand breaks, through which apoptotic responses are significantly augmented.

hOGG1-Knockdown GM00637 Cells Lead to the Significant Increases of Caspase-3/7 Activities in Response to H2O2-Oxidative Stress
The activation of caspases is one of the major processes in DNA damage–induced apoptosis. To understand the protective effect of hOGG1 on H2O2-induced apoptosis, we examined the induced level of caspase-3/7, the key apoptotic caspases, in hOGG1-siRNA–transfected GM00637 cells and their control-siRNA transfectants by Western blot analysis. There was no significant difference in the levels of procaspase-3 and procaspase-7 between control-siRNA transfectants and hOGG1-siRNA–transfected GM00637 (data not shown). Although none of the active processed products of both enzymes were detected in the control-siRNA transfectants up to 48 h after the H2O2 treatment, the cleavage of procaspase-3/7 into the active subunits was clearly observed in the hOGG1-siRNA–transfected GM00637 cells from 24 h after the H2O2 treatment (Fig. 2A ). The propensity is consistent with the activation of their enzymatic activity, which were obtained from the caspase-3/7 activity assay using a Promega CaspaseGlo 3/7 kit, as described in Materials and Methods. The enhanced caspase-3/7 activity of hOGG1-siRNA transfectants in response to H2O2 was significantly inhibited by caspase inhibitors (Fig. 2B). The cell viability of the H2O2-induced hOGG1-siRNA transfectants was significantly decreased in comparison with that of their control-siRNA transfectants. In contrast, treatment with caspase-3/7 inhibitors causes almost complete recovery from the decreased cell viability of hOGG1-siRNA transfectants compared with that of control-siRNA–transfected cells (Fig. 2C). This indicates that caspase-3/7 activation plays an important role in the development of H2O2-induced apoptosis in hOGG1-depleted human fibroblast GM00637 cells.


Figure 2
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  		FIGURE 2. Silencing hOGG1 triggers caspase-3 and caspase-7 activation in response to H2O2 in GM00637 cells. A. Cells were treated with 50 µmol/L H2O2 for 1 h. The activated (cleaved) caspase-3 and caspase-7 were detected by Western blot analysis using specific antibodies for caspase-3 and caspase-7 at the indicated time points after the H2O2 treatment. Data were normalized with {alpha}-tubulin expression. B. After control-siRNA– and hOGG1-siRNA–transfected GM00637 cells were treated with or without 60 µmol/L caspase inhibitor and the indicated concentration of H2O2, caspase-3/7 activities were analyzed using Promega Caspase Glo 3/7 kit as described in Materials and Methods. Data were plotted as the relative units of luciferase intensity of three separate experiments. Points, mean; bars, SD. C. Cell viability was determined by trypan blue exclusion under the same experimental condition as in B. The extent of cell death was estimated as the percentage of total cell number that remained unstained. The control-siRNA–transfected cells served as experimental controls in this study. Points, mean from five separate experiments; bars, SD.

 
Requirement of p53 Activation for H2O2-Induced Cell Death in hOGG1-Deficient Cells
In response to a variety of death stimuli, p53 is stabilized and rapidly activated in many cell types (20). One important mechanism through which p53 promotes caspase activation is the transactivation of proapoptotic proteins (21). To determine the contribution of a closely correlated interaction between p53 and hOGG1 to the oxidative stress–induced apoptosis, we analyzed the changed pattern of p53 phosphorylation, the followed regulation of p53-downstream target proteins such as p21 and Noxa, the variation of apoptotic effector caspase-3/7 activity, and the trend of cellular cytotoxicity in hOGG1-deficient and/or p53-knockdown cells and their control cells in response to H2O2 treatment. The significantly higher level of the H2O2-induced p53 phosphorylation at Ser15 and Ser20, followed by the more activated expression of p21 and Noxa, were shown in hOGG1-deficient GM00637 cells, but not in their control cells, in a time course–dependent manner after 100 µmol/L H2O2 treatment (Fig. 3A and B ). These results indicate that p53 contributes to the oxidative stress–induced cellular apoptotic responses in hOGG1-deficient GM00637 cells. Thus, we used p53-siRNA to silence p53 expression to test the p53-specific effect on the H2O2-induced apoptosis in the hOGG1-deficient GM00637. The p53 knockdown of hOGG1-deficient cells by its siRNA resulted in the predominant decreases of the activated caspase-3/7 expression and its activity in a dose-dependent manner (Fig. 3C and D). We next examined the cell cytotoxicity in response to H2O2 treatment under the same experimental conditions (Fig. 3E). The p53 depletion in the hOGG1-deficient cells markedly restored their cell viability from the H2O2-induced cell death. These results indicate that the p53 activation is required for H2O2-induced cellular apoptosis in hOGG1-deficient cells.


Figure 3
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  		FIGURE 3. H2O2-induced changes in the expression levels of phosphorylated p53, p21, and Noxa, and significant protection by p53-siRNA against H2O2-induced cytotoxicity in hOGG1-silencing GM00637 cells. After control-siRNA– and hOGG1-siRNA–transfected GM00637 cells were treated with 100 µmol/L H2O2, whole-cell lysates were prepared after incubation with H2O2 for the indicated times. Western blot analysis was carried out using specific antibodies against p53 and the phosphorylated p53 such as p53-P(Ser15) and p53-P(Ser20) (A), and using specific antibodies against p21 and Noxa (B). Data were normalized to {alpha}-tubulin expression. Results are representative of three independent experiments. C. After control and hOGG1-deficient GM00637 cells were transfected with p53-siRNA, the cells were then treated with the indicated concentration of H2O2 for 24 h. Whole-cell lysates were analyzed by Western blotting using specific antibodies against p53, p21, and caspase-3/7. Data were normalized with ß-actin expression. D. After hOGG1-silencing cells were transfected with control-siRNA or p53-siRNA, the cells were treated with the indicated H2O2 concentrations. Caspase-3/7 activities were analyzed by Promega Caspase Glo 3/7 kit as described in Materials and Methods. Data were plotted as the relative units of luciferase intensity from three separate experiments. Points, mean; bars, SD. E. Cell viability was determined by trypan blue exclusion under the same experimental condition as in D. The extent of cell death was estimated as the percentage of total cell number that remained unstained. Points, mean from six separate experiments; bars, SD.

 
We further investigated the hOGG1 effect on the H2O2-induced cell death in human lung carcinoma H1299 p53 null cells to determine the more exact relationship between p53 activation and hOGG1 during apoptosis. We first transfected the H1299 cells with either hOGG1-siRNA or its control GFP-siRNA, together with p53-expression vector or its control-pcDNA3.1 vector. We observed that the expression of hOGG1 was specifically down-regulated up to 25% to 35% in the hOGG1-siRNA–transfected H1299, and the p53 expression was clearly identified in the H1299 cells transfected with p53 expression vector (Fig. 4A ). When we examined the cell viability by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay at the various concentration of H2O2 24 h after the transfection, the hOGG1-knockdown H1299 exhibited the almost similar cell viability to that of its control-siRNA transfectants in response to H2O2 treatment, for example, 73.6% versus 79.4% at 100 µmol/L H2O2 treatment, respectively (Fig. 4B). In contrast, after the transfection with p53 expression vector, the hOGG1 wild-type H1299 showed the significantly increased cell viability in comparison with that of the hOGG1-knockdown H1299 at the same experimental condition, for example, 45.2% versus 28.6% at 100 µmol/L H2O2 treatment, respectively (Fig. 4B). Taken together, these results indicate that p53 clearly modulates apoptosis in response to H2O2-oxidative stress, which is closely related with hOGG1 expression in human fibroblasts and human lung carcinomas.


Figure 4
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  		FIGURE 4. The role of p53 in H2O2-mediated cell death of hOGG1-deficient H1299 lung carcinoma p53 null cells. A. After H1299 cells were transfected with control- and/or hOGG1-siRNA together with pcDNA3.1 or p53-pcDNA3.1 expression vector, whole-cell lysates were prepared, and Western blot analysis was done using anti-hOGG1 polyclonal antibodies and anti-p53 polyclonal antibodies. Data were normalized with ß-actin expression. Columns, mean based on the results from Western blot analysis; bars, SD. Statistically significant differences in the expression of hOGG1 and p53 were determined by the ANOVA with P values given for both the overall (P) and the pair-wise comparisons as indicated by asterisks (*, P < 0.05). B. The cells in A were cultured to 70% to 80% confluence and treated with the indicated concentration of H2O2 for 1 h. The medium was then changed to a fresh one, and the extent of cell death was estimated with MTT assay kit as described in Materials and Methods. Cell viability was determined as the percentage of total cell number. Points, mean from three separate experiments; bars, SD. The level of cell viability at 100 µmol/L H2O2 treatment was separately described together with the graphical presentation.

 
hOGG1 Overexpression Plays an Important Role to Protect GM00637 against H2O2-Induced Apoptosis
To determine if hOGG1 overexpression suppresses H2O2-induced p53 phosphorylation and apoptosis, we first stably transfected the GM00637 cells with either hOGG1 expression vector or its control empty vector pcDNA3.1, as previously described (22). The hOGG1 expression was highly augmented in two transfectants with hOGG1 expression vector (Fig. 5A ), and their 8-oxoG cleavage enzyme activities to the 21-mer oligonucleotide containing a single 8-oxoG at nucleotide 13 were significantly increased compared with those of parental cells and its control vector transfectants (Fig. 5B). Overexpression of hOGG1 decreased the level of p53 phosphorylation in response to 100 µmol/L H2O2 in the overall time points (until 8 h; Fig. 5C). The effect of overexpressed hOGG1 on H2O2-induced apoptotic responses in GM00637 cells was determined by estimation of their sub-G1 DNA contents (Fig. 5D). The hOGG1-overexpressed cells showed an apoptotic sub-G1 DNA content ranging from 4.61% to 10.10% in response to 0 to 80 µmol/L H2O2, whereas its control vector–transfected cells displayed DNA content ranging from 1.21% to 18.41% under the same oxidative stress. Because the oxidative stress–induced cellular apoptotic responses are specifically inhibited by hOGG1 overexpression in human fibroblast GM00637 cells, we postulate that, although p53 clearly modulates apoptosis in response to H2O2-oxidative stress, hOGG1 also plays a protective role in the H2O2-induced cell death.


Figure 5
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  		FIGURE 5. The overexpression of hOGG1 inhibits apoptotic cell death in GM00637 cells. A. After cells were transfected with either empty vector or hOGG1 expression vector, whole-cell lysates from parental, control-pcDNA3.1–transfected GM00637 cells and two hOGG1 transfectants were then separated by 12% SDS-PAGE. The expression of hOGG1 was analyzed by Western blotting using anti-hOGG1 polyclonal antibodies. Data were normalized with {alpha}-tubulin expression. B. The OGG1 enzyme activities were analyzed by an 8-oxoG nicking assay using the parental-, empty vector transfected–, and two of hOGG1 expression vector–transfected cell lysates, as described in Materials and Methods. Purified hOGG1 was used as positive control. Arrowhead, percent ratios of DNA cleavage products (13-mer), which are analyzed by the AxioVision Rel.4.6 Imaging System (Carl Zeiss International). C. After control and hOGG1-overexpressed GM00637 cells were treated with 100 µmol/L H2O2 for the indicated times, the level of phosphorylated p53 was examined by Western blot analysis using specific antibodies against phosphorylated p53 such as p53-P(Ser15) and p53-P(Ser20). Data were normalized to {alpha}-tubulin expression. Data are representative of three independent experiments. D. The cells transfected with control or hOGG1 expression vector were cultured to 70% to 80% confluences. After the cells were treated with the indicated concentration of H2O2 for 1 h, apoptotic sub-G1 DNA contents were estimated by FACScan analysis.

 
Involvement of Ataxia Telangiectasia Mutated and DNA-Protein Kinase in the H2O2-Induced p53 Phosphorylation of hOGG1-Deficient GM00637 Cells
Several protein kinases have been identified that detect genotoxic stress and initiate signaling pathway through p53 phosphorylation at the serine or threonine residues. Among them, ataxia telangiectasia mutated (ATM) and DNA-protein kinase (DNA-PK) play an essential role in transmitting DNA damage signals through phosphorylation of p53 (21, 23, 24). Thus, we next examined H2O2-induced phosphorylation of the ATM and DNA-PK in hOGG1-deficient and hOGG1-proficient cells. To determine whether these protein kinases participate in checkpoint control in response to H2O2-oxidative stress in hOGG1-deficient cells, we treated control-siRNA– and hOGG1-siRNA–transfected GM00637cells with 100 µmol/L H2O2 for 0.5, 2, or 8 h. As shown in Fig. 6A , the significantly increased phosphorylation of ATM and DNA-PK in response to H2O2 treatment was detected in the hOGG1-siRNA–transfected GM00637, but not in its control-siRNA transfectants. The hOGG1 defect also caused the enhanced Chk1 phosphorylation and the increased expression of Rad51 protein, which are ATM downstream signal molecules, in response to H2O2 treatment (Fig. 6A).


Figure 6
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  		FIGURE 6. H2O2-induced p53 phosphorylation is specifically related with the activation of DNA-PK and ATM in hOGG1-deficient GM00637 cells. A. After transfection with control- and hOGG1-siRNA, the cells were treated with 100 µmol/L H2O2 for the indicated times. The level of normal and phosphorylated DNA-PK catalytic subunit (DNA-PKcs), ATM, and Chk1 together with the expression of Rad51 was examined by Western blot analysis using each specific antibodies as described in Materials and Methods. B and C. After hOGG1-silencing cells were pretreated either with 6 mmol/L caffeine and/or 30 µmol/L wortmannin for 30 min, the cells were then incubated with 100 µmol/L H2O2 for the indicated times. The level of phosphorylated p53 together with normal p53 was examined by Western blot analysis using specific antibodies against all related proteins. Data were normalized toß-actin expression (B). Cell viability was estimated with MTT assay kit as described in Materials and Methods (C). Data are representative of three independent experiments.

 
To more specifically investigate whether ATM and DNA-PK activation contributes to the oxidative stress–induced p53 phosphorylation in hOGG1-deficient GM00637 cells, we pretreated the hOGG1-siRNA–transfected cells and its control-siRNA transfectants either with caffeine, which is a potent ATM/ATM- and Rad3-related inhibitor, or wortmannin, which is a potent DNA-PK/ATM inhibitor. The H2O2-induced phosphorylation of p53-Ser15 and p53-Ser20 in hOGG1-deficient cells was significantly reduced by caffeine pretreatment and by wortmannin pretreatment (Fig. 6B). We next examined the cell viability of wortmannin-pretreated hOGG1-deficient cells by MTT assay. As indicated in Fig. 6C, pretreatment with 30 mmol/L wortmannin completely restored the cell viability of hOGG1 knockdown cells from 58.8% to 105%, which was similar with that of control cells (100%). These results suggest that the H2O2-induced p53 phosphorylation of hOGG1-deficient cells was specifically enhanced by the activation of ATM and DNA-PK.

Array Comparative Genome Hybridization Characterization of the hOGG1-Deficient Fibroblasts after H2O2 Treatment
To identify the genetic copy number changes associated with the knockdown of hOGG1, an array comparative genome hybridization (array CGH) was done using genomic DNAs derived from hOGG1 knockdown cells and its control cells, which had been treated with 10 µmol/L H2O2 for 4 weeks (Fig. 7A ) and 8 weeks (Fig. 7B). The array CGH is a powerful molecular cytogenetic method, which enables genome-wide screening for copy number losses and gains of the chromosomal parts by single hybridization (25). Currently, array-based comparative genomic hybridization (array CGH) using bacterial artificial chromosome arrays spanning the entire human genome is used to examine genomic alterations (26). The advancements in genome-wide bacterial artificial chromosome array CGH technology have contributed to the understanding of the underlying genetic changes that correspond to disease progression. The extent of numerical changes and their location in DNA contents of the H2O2-induced hOGG1-deficient cells and its control cells were examined by the array CGH. The hOGG1-deficient cells showed more significant changes in the copy number of large regions of their chromosomes, compared with their control cells in response to H2O2 treatment (Fig. 7). We observed the representative chromosomal losses around 1q44 (gene loci of LOC339529, ZNF238, and LOC440742) and 7q36.3 (gene loci of VIPR2 and LOC442366), and the clear chromosomal gains at 17q11.2 (gene loci of MYO18A), 18q23 (gene locus of FLJ25715 and CTDP1), and Xq13.1 (gene loci of PJA1 and RP13-153N15.1), respectively, based on the precise measurement of the level of gains (log2 ratio >0.225, green arrows) and losses (log2 ratio less than –0.225, red arrows) of all chromosomes by the array CGH. These results suggest that the suppression of hOGG1 repair activity can lead to an alteration of the gene copy number in response to H2O2-oxidative stress.


Figure 7
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  		FIGURE 7. Array CGH of the control-siRNA– and hOGG1-siRNA–transfected GM00637 cells. The cells were stably transfected with control- and/or hOGG1-siRNA and then treated with 10 µmol/L H2O2 for 4 wk (A) and 8 wk (B). The genomic DNA was extracted and digested with DpnII, followed by random prime labeling with Cy3 for the test DNA derived from hOGG1-siRNA–transfected cells and Cy5 for the reference DNA from control-siRNA–transfected cells. Whole-genome array CGH profiles showed log2-transformed hybridization ratios of the test DNA versus the control-reference DNA. Representative chromosomal gains (log2 ratio >0.25, green arrows) and losses (log2 ratio less than –0.25, red arrows) are also indicated along with their chromosomal locations and candidate genes.

 

   Discussion
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
Materials and Methods
 References
 
Reactive oxygen species such as superoxide, hydroxyl radicals, and H2O2 oxidize macromolecules in living cells, leading to various types of cellular dysfunctions, for example, cell death, mutagenesis, aging, cancer, diabetes mellitus, atherosclerosis, neurologic degeneration, schizophrenia, and autoimmune disorders like arthritis (13-15, 27-29). Among many modified DNA macromolecules generated by the oxidative stresses, 8-oxoG is the most abundant product that seems to play a major role in mutagenesis and carcinogenesis (3). Hence, understanding the mechanisms for processing oxidative DNA damage is important in elucidating the oxidative stress–mediated diseases. Recently, it was reported that the frequency of apoptosis and necrosis is related to the nonrepair oxidative DNA adducts. For example, a decrease in hOGG1 confers cellular hypersensitivity to H2O2 (30), and hOGG1-deficient human leukemia cells (KG-1) could be forced to undergo apoptosis by 8-hydroxyguanosine (31). On the other hand, the expression of hOGG1 suppresses the oxidative stress–derived DNA damages, followed by the enhanced cell survival (32-34). However, the exact mechanism by which oxidative stress–mediated DNA damage causes cell death is unclear.

In this study, we examined the role of hOGG1 in cytotoxicity of human fibroblast GM00637 cells in response to H2O2-oxidative stress. It was achieved by manipulating the capability of the cells to remove the 8-oxoG from free radical damages and by assessing the effects of this manipulation on the cellular survival after H2O2 treatment. The RNA interference–mediated knockdown of hOGG1 in the GM00637 cells led to the loss of 80% to 90% endogenous hOGG1 expression and the increased cellular sensitivity to H2O2 (Fig. 1). We also observed a significant increase of sub-G1 population in the hOGG1-silencing cells but not in the control-siRNA–transfected cells (Fig. 1F). The elevated apoptosis in hOGG1-depleted GM00637 cells may be caused by the increased genotoxicities, including both DNA single- and double-strand breaks (Fig. 1G). These results indicate that hOGG1, as a major DNA repair enzyme for 8-oxoG, plays a protective role to confer a survival advantage in response to H2O2-oxidative stress in human fibroblasts.

Apoptosis is associated with the activation of caspases by a cytosolic multiprotein complex formed upon the release of cytochrome c from permeabilized mitochondria, which results in the loss of the mitochondrial membrane potential or the activation of a proapoptotic protein such as Bax (35). The released cytochrome c from mitochondria induces the proteolytic activation of procaspase-9 and the downstream activation of caspases-3 and caspase-7, which are key mediators of apoptosis (36, 37). Likewise, during the H2O2-induced apoptosis, procapase-3 and procaspase-7 were more intensively cleaved into detectable active forms of caspase-3 (14 and 22 kDa) and caspase-7 (26 kDa), respectively, in the hOGG1-knockdown cells than in their control cells (Fig. 2A). The caspase-3/7 inhibitor specifically suppressed the apoptosis of hOGG1-knockdown cells as shown in Fig. 2C. Therefore, we postulate in this study that unrepaired oxidative DNA lesion like 8-oxoG in hOGG1-depleted cells may also play an important role in the H2O2-induced apoptotic process through the mitochondrial apoptotic pathway.

p53 is a tumor-suppressor protein that transmits the signals arising from various forms of cellular stress to the genes as well as the factors that induce cell cycle arrest and apoptosis. The p53 is not only induced by a variety of apoptotic stimuli but the overexpression of p53 has also been shown to induce apoptosis in a variety of cell types (20, 21). Several p53 target mitochondrial proteins, such as Bax, Noxa, and PUMA, affect the mitochondrial membrane potential, which is an important determinant of mitochondrial apoptotic signaling (21). However, p53 can also directly engage in the major cellular apoptotic pathways, promoting both death receptor signaling and mitochondrial perturbations (21). In this study, we show the contribution of a closely correlated interaction between p53 and hOGG1 to the oxidative stress–induced apoptosis, because the significantly higher level of the H2O2-induced p53 phosphorylation at Ser15 and Ser20, followed by the more activated expression of p53-downstream target proteins such as p21 and Noxa, were shown in hOGG1-deficient GM00637 cells, but not in their control cells (Fig. 3A and B). Moreover, the p53 knockdown of hOGG1-deficient GM00637 resulted in the decreased activation of caspase-3/7, and markedly restored the cell viability of hOGG1-deficient cells from the H2O2-induced cell death (Fig. 3C-E). We further confirmed the protective effect of hOGG1 on the H2O2-induced cell death in human lung carcinoma H1299 p53 knockout cells. The cell viability of hOGG1-knockdown H1299 was similar to that of the hOGG1 wild-type H1299, which is 73.6% versus 79.4% at 100 µmol/L H2O2 treatment, respectively (Fig. 4). After the overexpression of p53, the hOGG1-knockdown H1299 showed the significantly decreased cell viability (28.6%) compared with that of the hOGG1 wild-type H1299 (45.2%) at the same experimental condition. The propensity for the cell viability in response to H2O2-oxidative stress in this study is coincided with the previous report (38), in which Chen et al. showed a significantly increased viable cells in the p53 null H1299—but not in the p53 wild-type H460 cells—in response to trichloroethylene, a potent H2O2 accumulant, time and dose dependently. In contrast, Chatterjee et al. (39, 40) recently showed that the combined suppression of hOGG1 and p53 reduced the cell viability of human mammary gland epithelia, MCF12A, when compared with suppression of each gene individually.

The discrepancy of cell viability between human fibroblasts, lung carcinomas, and mammary gland epithelia in the case of both knockdowns of p53 and hOGG1 may be derived from different cellular characteristics in the fields of sensitivity against specific oxidative stress, changing period of cell cycle in response to the stress, origin of cell lines, and so on. In addition, many other cellular processes, for example, mitogen-activated protein kinase activation, are also affected by the hOGG1 defect after H2O2 treatments (34), which may contribute to the increased sensitivity of the hOGG1-deficient cells to oxidative stress. Further studies need to identify these pathways and determine their roles in oxidative stress–induced cellular responses in hOGG1-proficient and hOGG1-deficient cells.

Chatterjee et al. (39) and Achanta et al. (40) indicated that p53 is a major regulator of hOGG1 expression and hOGG1 activity, because p53 knockout resulted in a severe reduction in hOGG1 mRNA and hOGG1 protein levels, and addition of p53 protein increased the ability to incise 8-oxoG from a synthetic oligonucleotide, suggesting involvement of p53 in damage recognition and enhancement of glycosylase activity. Although a transcriptional/translational regulation of hOGG1 by p53 is important to their relationships in response to oxidative stress, it is consistent with our present data at a viewpoint of feedback down-regulated p53 activation by hOGG1. That is, p53 is able to regulate the upstream of hOGG1; conversely, activated hOGG1 may also play a feedback down-regulation of p53 activity in response to oxidative stress.

Therefore, taken together, we suggest that although p53 is a major modulator of apoptosis, hOGG1 also plays a protective role in the H2O2-induced apoptosis at the upstream of the p53-dependent pathway to confer a survival advantage to human fibroblasts and human lung carcinomas. We postulate the hOGG1 effect on protecting cells against the oxidative stress–induced apoptosis through p53-dependent apoptotic signaling pathway as shown in Fig. 8 :

(a) In hOGG1 wild-type, oxidative damage triggers the activation of hOGG1-related DNA repair to remove oxidative DNA adducts like 8-oxoG, followed by the decrease in apoptosis through blocking of p53 phosphorylation.
(b) In hOGG1 knockdown by siRNA, the oxidative DNA adducts are increased by oxidative damages because of the inhibited hOGG1 activation, followed by increased apoptosis through the activation of p53.
(c) In p53 knockdown by siRNA, hOGG1 repair activity is comparatively ineffective in oxidative stress–induced apoptosis. Because of the intrinsic blocking of p53 activation, cell viability in p53 knockdown cells is prominently high in response to any oxidative stress in comparison with that in the p53 wild-type cells.


Figure 8
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  		FIGURE 8. Schematic diagram of p53-mediated apoptotic pathway in hOGG1-deficient and hOGG1-proficient cells in response to H2O2 exposure.

 
p53 is phosphorylated at multiple sites by several different protein kinases, including ATM and DNA-PK, enhancing its tetramerization, stability, and activity (20). The activation of ATM kinases, which are important mediators of signals originating from DNA damage, is caused by ionizing irradiation and genotoxic drugs (24). DNA-PK has recently been identified as a crucial molecule activating apoptotic machinery in response to DNA damage, particularly double-strand breaks (23). DNA-PK complex with p53 has been shown to act as a sensor of abnormal DNA structure, in which DNA-PK selectively regulates a p53-dependent apoptotic response (41). Therefore, activation of these protein kinases in H2O2-treated hOGG1-deficient cells would shed light on the activation of its upstream signaling molecule. To evaluate this possibility, we determined ATM and DNA-PK phosphorylation in response to H2O2 in hOGG1-deficient and hOGG1-proficient cells. The results obtained from Western blot experiments revealed that the phosphorylation of ATM protein at Ser981 and DNA-PK protein at Tyr2609 was significantly higher in hOGG1-deficient cells in response to H2O2 compared with that of hOGG1-proficient cells (Fig. 6). Additionally, the phosphorylation of Chk1 in response to H2O2, which has been shown to act downstream of ATM as signal transducers in DNA damage signaling (24), was also significantly increased in hOGG1-deficient cells; however, the H2O2-mediated phosphorylation of Chk1 does not significantly occur in hOGG1-proficient cells. Moreover, H2O2 treatment increased the level of Rad51 protein, which has been reported to be up-regulated by posttranslational modifications mediated by ATM (42) in hOGG1-deficient cells but not in hOGG1-proficient cells.

To identify the regulatory pathways of p53 activation in hOGG1-deficient GM00637 cells, we pretreated the cells with caffeine, a potent inhibitor of ATM, and wortmannin, a potent inhibitor of DNA-PK. Caffeine significantly abolished the H2O2-induced phosphorylation of p53-Ser15 and p53-Ser20, and wortmannin almost completely attenuated the p53-Ser20 phosphorylation in hOGG1-deficient cells (Fig. 6B). Because ATM and DNA-PK are normally activated by double-strand breaks or DNA replication arrest (23, 24, 41), we postulate that H2O2 treatment may increase DNA double-strand breaks in hOGG1-deficient cells. However, clustered damages such as double-strand breaks are only found after treatment with a very high concentration of H2O2 at 0°C to 4°C (43). Thus, we examined more specifically the possibility of clustered damages of hOGG1-deficient GM00637 cells in response to H2O2-oxidative stress by comet assay. We observed that DNA strand breaks, including both single- and double-strand breaks, was significantly increased in hOGG1-depleted cells compared with control cells, at neutral and alkali pH ranges, respectively (Fig. 1G). These results indicate that hOGG1 deficiency may be a major factor of the increased DNA strand breaks caused by H2O2-oxidative stress. Generally, the oxidative stress–induced DNA damages in hOGG1-depleted cells increase oxidized purine bases, such as 8-oxoG and Fapy-G DNA adducts, which are repaired by base excision repair. The base excision repair requires a DNA glycosylase and an AP endonuclease, which generate gaps in ssDNA. Such interruption, if encountered by a replication fork, can cause double-strand breaks (44, 45). It is also possible that unrepaired oxidized purines in hOGG1-deficient cells can lead to topoisomerase I–oxidized DNA cleavage complexes (46), which may activate ATM and DNA-PK. Therefore, as depicted in Fig. 8, the activation of ATM and DNA-PK protein may represent an important signal of H2O2-oxidative stress, and the hOGG1 defect may enhance p53 phosphorylation through the activated ATM/DNA-PK signal transduction, resulting in the increased apoptotic cell death. However, the exact mechanism for the H2O2-oxidative stress–induced apoptotic responses is not clear yet.

There are many previous reports for the role of OGG1 in various human tumor suppression, for example, the frequent loss of the OGG1 chromosomal locus in human lung and renal cancers, and the significantly lower OGG1 activity in lung cancer patients (9). The inactivation or knockdown of mouse OGG1 gene also led to the accumulation of 8-oxoG, followed by an increase in the mutation frequency in liver, kidney, and skin (47, 48). Thus, we investigated the genomic profile of hOGG1-depleted cells in chronic exposure of H2O2 to detect the gains or the losses of specific oncogenes and tumor-suppressor genes. The analysis of the overall genomic integrity by the high-resolution array CGH revealed a significantly higher level of alteration of particular gene copy number in the H2O2-treated hOGG1-deficient cells than in the control cells (Fig. 7). The bacterial artificial chromosome array profiles of the H2O2-treated hOGG1 knockdown cells showed chromosomal losses of LOC339529, ZNF238, LOC440742, VIPR2, and LOC442366, and chromosomal gains of MYO18A, FLJ25715, CTDP1, PJA1, and RP13-153N15.1, indicating that hOGG1 plays an important role in maintaining the genomic stability against oxidative stress. We are carrying out the more detailed characterization of the genes, in vitro and in vivo, these days.

In summary, we show in this study that the down-regulation of hOGG1 expression prominently triggers the H2O2-induced apoptosis in human fibroblast GM00637 cells and human lung carcinoma H1299 cells via the p53-mediated apoptotic pathway. The hOGG1-mediated protection against the oxidative stress–induced apoptosis is achieved through the down-regulation of caspase-3/7 and p53 phosphorylation. Although the precise mechanism of the hOGG1-inhibited p53-dependent apoptosis remains to be determined, it may be related to the shift in the balance of damage/repair process toward the accumulation of oxidative DNA lesions such as 8-oxoG. Chromosomal instability may also play a critical role in tumor evolution of hOGG1-deficient cells through the increased rate of genetic selection of cancer genes with altered DNA copy number.

The future challenge is to define the target molecules of the oxidative stress–induced p53 activation and their biological significance for the activated signaling pathways. Studies focusing on these biochemical steps would extend our understanding of the oxidative DNA damage signaling cascades stimulated by oxidative stress during normal metabolic and pathologic processes.


   Materials and Methods
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
References
 
Maintenance of Cell Lines
Human fibroblast GM00637 cells (Coriell Institute for Medical Research, Camden, NJ) were maintained in Earle's MEM supplemented with 10% fetal bovine serum, 100 units/mL penicillin, and 100 mg/mL streptomycin (Invitrogen). Human lung carcinoma H1299 (p53-null) cells were purchased from American Type Culture Collection and grown in RPMI 1640 supplemented with 10% fetal bovine serum, 100 units/mL penicillin, and 100 mg/mL streptomycin (Invitrogen). The cells were maintained in a humidified incubator in an atmosphere containing 5% CO2 at 37°C.

Western Blotting
Cultured cells were washed with PBS and lysed at 0°C for 30 min in a lysis buffer [20 mmol/L HEPES (pH 7.4), 2 mmol/L EGTA, 50 mmol/L glycerol phosphate, 1% Triton X-100, 10% glycerol, 1 mmol/L DTT, 1 mmol/L phenylmethylsulfonyl fluoride, 10 mg/mL leupeptin, 10 mg/mL aprotinin, 1 mmol/L Na3VO4, and 5 mmol/L NaF]. After cellular protein contents were determined using a dye-binding microassay (Bio-Rad), 20 µg of the protein per lane were electrophoresed on 10% SDS-polyacrylamide gels after boiling for 5 min in a Laemmli sample buffer. The proteins were blotted onto Hybond enhanced chemiluminescence membranes (Amersham Biosciences). After electroblotting, the membranes were blocked with TBS containing Tween 20 [10 mmol/L Tris-HCl (pH 7.4), 150 mmol/L NaCl, 0.1% Tween 20] and 5% milk. The membranes were then incubated with appropriate primary antibodies in a blocking buffer for 4 h. We followed the manufacturer's protocol for dilution of all primary antibodies. The membranes were then washed, incubated with the appropriate secondary antibodies (1:4,000) in a blocking buffer for 2 h, and washed again. The blotted proteins were detected using an enhanced chemiluminescence detection system (iNtRON Biotech).

Antibodies
Antibodies used in this study are anti-human OGG1 polyclonal antibody (Santa Cruz Biotechnology); anti-p53 polyclonal antibody, anti–p53-P(Ser15) polyclonal antibody, anti–p53-P(Ser20) polyclonal antibody (Cell Signaling Technology); anti-p21 monoclonal antibody (mAb; BD PharMingen); anti-Noxa mAb (Calbiochem); anti–ß-actin mAb (BD PharMingen); anti–{alpha}-tubulin mAb (BD PharMingen); anti–DNA-PK catalytic subunit mAb (Santa Cruz Biotechnology); anti–DNA-PK catalytic subunit-P (Tyr2609) (Abcam); anti-ATM mAb (Santa Cruz Biotechnology); anti–ATM-P (Ser1981) mAb (Cell Signaling Technology); anti–Chk1-P (Ser345) polyclonal antibody, anti-Chk1 polyclonal antibody (Cell Signaling Technology); anti-Rad51 mAb (Oncogene).

Semiquantitative Reverse Transcription-PCR
RNA extraction was carried out using the RNA-STAT-60 according to the manufacturer's instructions (TEL-TEST, Inc.). Two micrograms of the total RNA were reverse-transcribed using a M-MLV cDNA synthesis system (Promega), and the reverse-transcribed DNA was subjected to PCR. The profile of replication cycles was denaturation at 94°C for 50 s, annealing at 58°C for 50 s, and polymerization at 72°C for 1 min. In each reaction, the expression of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as the internal control. The primers used for PCR are as follows: hOGG1 forward, 5'-CTGCCTTCTGGACAATCTTT-3'; hOGG1 reverse, 5'-TAGCCCGCCCTGTTCTTC-3' designed to amplify a 551-bp region; GAPDH forward, 5'-CCATGGAGAAGGCTGGGG-3'; and GAPDH reverse 5'-CAAAGTTGTCATGGATGACC-3' designed to amplify a 194-bp region (total number of cycles, 26). The PCR products were resolved on 1% agarose gels, stained with ethidium bromide, and then photographed.

Small Interfering RNA
Three target sites within human OGG1 genes were chosen from the human OGG1 mRNA sequence (Genbank accession no. AF003595), which was extracted from the National Center for Biotechnology Information Entrez nucleotide database. After selection, each target site was searched with National Center for Biotechnology Information BLAST to confirm the specificity only to the human OGG1. The sequences of the 21-nucleotide sense and antisense RNA are as follows: hOGG1-siRNA, 5'-GUACUUCCAGCUAGAUGUUUU-3' (sense) and 5'-AACAUCUAGCUGGAAGUACUU-3' (antisense) for the hOGG1 gene (nucleotides 292–312); LacZ siRNA, 5'-CGUACGCGGAAUACUUCGAUU-3' (sense), 5'-AAUCGAAGUAUUCCGCGUACGUU-3' (antisense) for the LacZ gene. These siRNAs were prepared using a transcription-based method with a Silencer siRNA construction kit (Ambion) according to the manufacturer's instructions. LacZ siRNA was used as the negative control. Cells were transiently transfected with siRNA duplexes using Oligofectamine (Invitrogen) and stably transfected with the constructed siRNA expression plasmid based on pSilence hygro vector (Ambion), which includes a human U6 promoter and a hygromycin resistance gene, using LipofectAMINE (Invitrogen). If required, we selected several resistant colonies against 100 µg/mL hygromycin in the medium after transfection.

Single-Cell Gel Electrophoresis (Comet Assay)
Single-cell gel electrophoresis assay was done essentially as described by Sing et al. (49). After treatment, aliquots of the cell suspension (50–100 µL, 0.5 x 105–1.0 x 105 cells) were transferred to 1.5-mL Eppendorf tubes and centrifuged at 200 x g for 5 min. The supernatant was discarded and the pellet was mixed with 75 µL low-melting agarose (0.7% in PBS) and distributed onto conventional microscope slides. The slides were precoated with normal melting agarose (0.5% in PBS) and dried at room temperature. After the agarose had solidified (4°C for 10 min), a second layer of normal melting agarose was applied similarly to the first. The slides were then immersed in lysis solution [2.5 mol/L NaCl, 100 mmol/L Na2EDTA, 10 mmol/L Tris-HCl (pH 10), containing freshly added 1% Triton X-100 and 10% DMSO] for 1 h at 4°C and then placed into a horizontal electrophoresis apparatus filled with freshly made buffer [1 mmol/L Na2EDTA, 300 mmol/L NaOH (pH >13 for alkaline conditions or pH 7-8 for neutral conditions)]. After 20 min of preincubation (unwinding of DNA), electrophoresis was run for 30 min at a fixed voltage of 25 V and 300 mA, which was adjusted by raising or lowering the level of the electrophoresis buffer in the tank. At the end of the electrophoresis, slides were washed with the 70% ethanol, stained with 50 µL ethidium bromide (20 µg/mL), and kept in a moist chamber in the dark at 4°C until analysis. All of the above-reported steps were carried out under red light to prevent additional DNA damage.

Comet Detection and Statistical Analysis
Cells were analyzed 24 h after staining at x400 magnification using a fluorescence microscope (Zeiss, R.G.) equipped with a 50-W mercury lamp. The microscope images revealed circular shapes, indicating undamaged DNA, and comet-like shapes, indicating the DNA had migrated out from the head to form a tail (damaged DNA). The extension of each comet was analyzed using a computerized image analysis system (Komet 5.5, Andor Technology) that provided a "tail moment," which is defined as the product of DNA in the tail and the mean distance of its migration in the tail and considered to be the variable most directly related to DNA damage. Calculation of the extent of DNA damage, which was not homogeneous, was based on analysis of 80 to 100 randomly selected comets from each slide.

8-OxoG Glycosylase Activity Assay
The cells at the exponential phase were centrifuged at 800 x g for 5 min. The cell pellets (106 cells per each assay) were then suspended in 2 volumes of a homogenization buffer [50 mmol/L Tris-HCl, 50 mmol/L KCl, 1 mmol/L EDTA, 5% glycerol, and 0.05% 2-mercaptoethanol (pH 7.5)] and homogenized. After the supernatants were obtained by centrifugation, followed by dialyzing against the homogenization buffer, they were used for the endonuclease-nicking assay as cell extracts. 8-OxoG–containing 21-mer with the sequence 5'-CAGCCAATCAGTXCACCATTC-3' (X = 8-oxoG) along with its complementary strand was chemically synthesized (The Midland Certified Reagent Co.). The synthetic oligonucleotide was 3'-end–labeled using terminal transferase and [a-32P]ddATP (Amersham Biosciences, 3,000 Ci/mmol). The end-labeled oligomer was annealed with its complementary oligonucleotide, and the resulting duplex DNA was used as the assay substrate. The duplex substrate DNA (20 pmol) was incubated with the cell extracts (10 mg of protein) at 37°C for 1 h in 1 mL of reaction mixture [50 mmol/L Tris-HCl, 50 mmol/L KCl and 1 mmol/L EDTA (pH 7.5)]. The reaction was quenched by heating at 90°C for 3 min, followed by electrophoresis on 20% denaturing (7 mol/L urea) polyacrylamide gels (DNA sequencing gel). The gels were wrapped in saran wrap and exposed to Kodak film for visualization.

Cytotoxicity Assay
The extent of cell death was determined by trypan blue exclusion. The cell viability was measured as a percentage of the total cell number that remained unstained. If required, the cell cytotoxicity was also assessed using a MTT-based colorimetric assay kit (Roche Applied Science), according to the manufacturer's instructions.

Propidium Iodide Staining
The floating and trypsin-detached GM00637 cells were collected and washed once with ice-cold PBS, followed by fixing in 70% cold ethanol. The cells were then stained in PBS and propidium iodide (50 µg/mL), RNase A (50 µg/mL), and 0.05% Triton X-100 for 45 min. The DNA content of the GM00673 cells was analyzed by fluorescent-activated cell sorting (FACSort, Becton Dickinson). At least 10,000 events were analyzed, and the percentage of cells in sub-G1 population was calculated. Aggregates of cell debris at the origin of histogram were excluded from the sub-G1 cells.

Caspase-3/7 Activity Assay
The caspase-3/7 activity was detected by a Caspase-Glo 3/7 Assay system (Promega) after preincubating the GM00637 cells (1 x 106/60-mm plate) with or without 60 µmol/L caspase inhibitor I (Calbiochem) for 12 h, followed by treating with various H2O2 concentrations (0-70 µmol/L) for another 12 h. The background luminescence associated with the cell culture and assay reagent (blank reaction) was subtracted from the experimental values. The activity of caspase-3/7 was presented as the means of triplets for the given cells.

Bacterial Artificial Chromosome Array-CGH
Human fibroblast genomic DNAs derived from si-hygro– and hOGG1-siRNA–transfected GM00637 were prepared by using the PUREGENE DNA Isolation kit (Gentra D5500A). For each CGH hybridization, we digested 2 µg of genomic DNA from the reference and the corresponding experimental sample with DpnII. All digests were done for a minimum of 2 h at 37°C and then verified by agarose gel analysis. The samples were then filtered by using the QIAQuick PCR clean-up kit (Qiagen). Labeling reactions were done with 0.8 µg of purified DNA and a Bioprime labeling kit (Invitrogen) according to the manufacturer's instructions in a volume of 50 µL with a modified deoxynucleotide triphosphate pool containing 120 µmol/L each of dATP, dGTP, dTTP, 60 µmol/L dCTP, and 60 µmol/L Cy5-dCTP (for the reference from si-hygro–transfected GM00637) or Cy3-dCTP (for the experimental sample from hOGG1-siRNA–transfected GM00637). Experimental and reference targets for each hybridization were pooled and mixed with 50 µg of human Cot-1 DNA (Invitrogen). The target mixture was ethanol precipitated and resuspended to a final volume of 40 µL hybridization buffer. Before hybridization to the array, the hybridization mixtures were denatured at 70°C for 15 min and applied to the array CGH chip slide, MacArray Karyo 4000 (Macrogen Korea). Hybridization was carried out for 48 to 72 h at 37°C in a rotating chamber (MAUI, BioMicro Systems, Inc.). The arrays were then disassembled in 50% formamide, 2x SSC at 46°C for 15 min, followed by washing with 2x SSC, 0.1% SDS at 46°C for 30 min. Slides were dried and scanned with a GenePix4000B microarray scanner (Axon Instruments-Molecular Devices). Microarray images were analyzed by using Macviewer software (Macrogen). Default settings were used. Whole-genome array CGH profiles showed log2-transformed hybridization ratios of test DNA versus control-reference DNA as described in the figure legends.

Statistical Analysis
Results are expressed as mean ± SD. For statistical analysis, ANOVA with P values were done for both the overall (P) and the pair-wise comparison as indicated by asterisks. Values of P < 0.05 were considered to be significant.


   Notes
Top
 Notes
Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Grant support: Korea Science and Engineering Foundation funded by Korea Government (Ministry of Science and Technology) grants M1063901001 and M10413010003, and Research Center for Proteinous Materials.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Note: P.I. Song and C-K. Youn contributed equally to this work.

Received 12/21/06; revised 6/ 1/07; accepted 6/ 6/07.


   References
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 

  1. Eyfjord JE, Bodvarsdottir SK. Genomic instability and cancer: networks involved in response to DNA damage. Mutat Res 2005;592:18–28.[Medline]
  2. Ide H, Kotera M. Human DNA glycosylases involved in the repair of oxidatively damaged DNA. Biol Pharm Bull 2004;27:480–5.[CrossRef][Medline]
  3. Fortini P, Pascucci B, Parlanti E, D'Errico M, Simonelli V, Dogliotti E. 8-Oxoguanine DNA damage: at the crossroad of alternative repair pathways. Mutat Res 2003;531:127–39.[Medline]
  4. Sancar A, Lindsey-Boltz LA, Unsal-Kacmaz K, Linn S. Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu Rev Biochem 2004;73:39–85.[CrossRef][Medline]
  5. Norbury CJ, Hickson ID. Cellular responses to DNA damage. Annu Rev Pharmacol Toxicol 2001;41:367–401.[CrossRef][Medline]
  6. Mitra S, Boldogh I, Izumi T, Hazra TK. Complexities of the DNA base excision repair pathway for repair of oxidative DNA damage. Environ Mol Mutagen 2001;38:180–90.[CrossRef][Medline]
  7. Barnes DE, Lindahl T. Repair and genetic consequences of endogenous DNA base damage in mammalian cells. Annu Rev Genet 2004;38:445–76.[CrossRef][Medline]
  8. Shibutani S, Takeshita M, Grollman AP. Insertion of specific bases during DNA synthesis past the oxidation-damaged base 8-oxodG. Nature 1991;349:431–4.[CrossRef][Medline]
  9. Boiteux S, Radicella JP. The human OGG1 gene: structure, functions, and its implication in the process of carcinogenesis. Arch Biochem Biophys 2000;377:1–8.[CrossRef][Medline]
  10. Nakabeppu Y, Tsuchimoto D, Furuichi M, Sakumi K. The defense mechanisms in mammalian cells against oxidative damage in nucleic acids and their involvement in the suppression of mutagenesis and cell death. Free Radic Res 2004;38:423–9.[CrossRef][Medline]
  11. Bohr VA. Repair of oxidative DNA damage in nuclear and mitochondrial DNA, and some changes with aging in mammalian cells. Free Radic Biol Med 2002;32:804–12.[CrossRef][Medline]
  12. Fan CY, Liu KL, Huang HY, et al. Frequent allelic imbalance and loss of protein expression of the DNA repair gene hOGG1 in head and neck squamous cell carcinoma. Lab Invest 2001;81:1429–38.[Medline]
  13. Cardozo-Pelaez F, Cox DP, Bolin C. Lack of the DNA repair enzyme OGG1 sensitizes dopamine neurons to manganese toxicity during development. Gene Expr 2005;12:315–23.[Medline]
  14. Iida T, Furuta A, Nishioka K, Nakabeppu Y, Iwaki T. Expression of 8-oxoguanine DNA glycosylase is reduced and associated with neurofibrillary tangles in Alzheimer's disease brain. Acta Neuropathol (Berl) 2002;103:20–5.[CrossRef][Medline]
  15. Kikuchi H, Furuta A, Nishioka K, Suzuki SO, Nakabeppu Y, Iwaki T. Impairment of mitochondrial DNA repair enzymes against accumulation of 8-oxo-guanine in the spinal motor neurons of amyotrophic lateral sclerosis. Acta Neuropathol (Berl) 2002;103:408–14.[CrossRef][Medline]
  16. Harrison JF, Hollensworth SB, Spitz DR, Copeland WC, Wilson GL, LeDoux SP. Oxidative stress-induced apoptosis in neurons correlates with mitochondrial DNA base excision repair pathway imbalance. Nucleic Acids Res 2005;33:4660–71.[Abstract/Free Full Text]
  17. Mambo E, Nyaga SG, Bohr VA, Evans MK. Defective repair of 8-hydroxyguanine in mitochondria of MCF-7 and MDA-MB-468 human breast cancer cell lines. Cancer Res 2002;62:1349–55.[Abstract/Free Full Text]
  18. Hyun JW, Jung YC, Kim HS, et al. 8-hydroxydeoxyguanosine causes death of human leukemia cells deficient in 8-oxoguanine glycosylase 1 activity by inducing apoptosis. Mol Cancer Res 2003;1:290–9.[Abstract/Free Full Text]
  19. Collins AR. The comet assay for DNA damage and repair: principles, applications, and limitations. Mol Biotechnol 2004;26:249–61.[CrossRef][Medline]
  20. Lavin MF, Gueven N. The complexity of p53 stabilization and activation. Cell Death Differ 2006;13:941–50.[CrossRef][Medline]
  21. Haupt S, Berger M, Goldberg Z, Haupt Y. Apoptosis—the p53 network. J Cell Sci 2003;116:4077–85.[Abstract/Free Full Text]
  22. Lee MR, Kim SH, Cho HJ, et al. Transcription factors NF-YA regulate the induction of human OGG1 following DNA-alkylating agent methylmethane sulfonate (MMS) treatment. J Biol Chem 2004;279:9857–66.[Abstract/Free Full Text]
  23. Collis SJ, DeWeese TL, Jeggo PA, Parker AR. The life and death of DNA-PK. Oncogene 2005;24:949–61.[CrossRef][Medline]
  24. Shiloh Y. ATM and ATR: networking cellular responses to DNA damage. Curr Opin Genet Dev 2001;11:71–7.[CrossRef][Medline]
  25. Kallioniemi A, Kallioniemi OP, Sudar D, et al. Comparative genomic hybridization for molecular cytogenetic analysis of solid tumors. Science 1992;258:818–21.[Abstract/Free Full Text]
  26. Wolf M, Mousses S, Hautaniemi S, et al. High-resolution analysis of gene copy number alterations in human prostate cancer using CGH on cDNA microarrays: impact of copy number on gene expression. Neoplasia 2004;6:240–7.[CrossRef][Medline]
  27. Klaunig JE, Kamendulis LM. The role of oxidative stress in carcinogenesis. Annu Rev Pharmacol Toxicol 2004;44:239–67.[CrossRef][Medline]
  28. Macip S, Igarashi M, Berggren P, Yu J, Lee SW, Aaronson SA. Influence of induced reactive oxygen species in p53-mediated cell fate decisions. Mol Cell Biol 2003;23:8576–85.[Abstract/Free Full Text]
  29. Macip S, Kosoy A, Lee SW, O'Connell MJ, Aaronson SA. Oxidative stress induces a prolonged but reversible arrest in p53-null cancer cells, involving a Chk1-dependent G2 checkpoint. Oncogene 2006;25:6037–47.[CrossRef][Medline]
  30. Yang N, Chaudhry MA, Wallace SS. Base excision repair by hNTH1 and hOGG1: a two edged sword in the processing of DNA damage in {gamma}-irradiated human cells. DNA Repair (Amst) 2006;5:43–51.[CrossRef][Medline]
  31. Hyun JW, Choi JY, Zeng HH, et al. Leukemic cell line, KG-1 has a functional loss of hOGG1 enzyme due to a point mutation and 8-hydroxydeoxyguanosine can kill KG-1. Oncogene 2000;19:4476–9.[CrossRef][Medline]
  32. Dobson AW, Xu Y, Kelley MR, LeDoux SP, Wilson GL. Enhanced mitochondrial DNA repair and cellular survival after oxidative stress by targeting the human 8-oxoguanine glycosylase repair enzyme to mitochondria. J Biol Chem 2000;275:37518–23.[Abstract/Free Full Text]
  33. Wu M, He YH, Kobune M, Xu Y, Kelley MR, Martin WJ II. Protection of human lung cells against hyperoxia using the DNA base excision repair genes hOgg1 and Fpg. Am J Respir Crit Care Med 2002;166:192–9.[Abstract/Free Full Text]
  34. Kannan S, Pang H, Foster DC, Rao Z, Wu M. Human 8-oxoguanine DNA glycosylase increases resistance to hyperoxic cytotoxicity in lung epithelial cells and involvement with altered MAPK activity. Cell Death Differ 2006;13:311–23.[Medline]
  35. Garrido C, Galluzzi L, Brunet M, Puig PE, Didelot C, Kroemer G. Mechanisms of cytochrome c release from mitochondria. Cell Death Differ 2006;13:1423–33.[CrossRef]