Estrogen receptor-mediated regulation of oxidative stress and DNA damage in breast cancer

James A. Mobley and Robert W. Brueggemeier1

Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, The Ohio State University Comprehensive Cancer Center, The Ohio State University, Columbus, OH 43210, USA


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Estrogens have been implicated to be complete carcinogens in breast and gynecologic tissues. Possible mechanisms may include differential metabolism with subsequent formation of reactive oxygen species and/or a receptor-mediated pathway, which may also involve indirect modulation of intracellular redox state. Estrogen-mediated oxidative DNA damage in mammary gland epithelia includes the induction of 8-oxo-2'-deoxyguanosine, both in vitro and in vivo, thereby suggesting a role for oxidative stress in the initiation and/or progression of breast neoplasia. In order to study this phenomenon, we have treated estrogen receptor alpha (ER-{alpha})-positive MCF-7 cells and ER-{alpha}-negative MDA-MB-231 cells with 10 nM 17ß-estradiol (E2), while measuring changes in antioxidant status and sensitivity to DNA damage by peroxide. Treatment of MCF-7 cells with E2 resulted in a marked decrease in the ability for these cells to metabolize peroxide, which paralleled a decrease in catalase activity and total glutathione levels. These observations also correlated with an increased sensitivity to peroxide-induced DNA damage. The estrogen-induced effects were all opposed by the anti-estrogen tamoxifen. In addition, the estrogen-mediated down regulation of peroxide metabolism, catalase activity, and sensitivity to DNA damage were not observed in the MDA-MB-231 cell line. Treatment of MCF-7 cells with E2 also resulted in increased glutathione peroxidase, superoxide dismutases (I) and (II) and glucose-6-phosphate dehydrogenase activities. Therefore, in this breast cancer model antioxidant status is modulated through the actions of the ER. The data may explain some of the estrogen-induced pro-oxidant effects previously reported in vivo. In addition, this is the first report indicating that E2 is capable of inducing an increase in sensitivity to oxidative DNA damage through an ER-mediated mechanism.

Abbreviations: BSO, buthionine sulfoximine; dG, 2'-deoxyguanosine; E2, estradiol; ER-{alpha}, estrogen receptor alpha; ER-ß, estrogen receptor beta; GSH, glutathione; GSSG, oxidized glutathione; G6PD, glucose-6-phosphate dehydrogenase; GPx, glutathione peroxidase; GR, glutathione reductase; HBSS, Hanks buffered saline solution; HRP, horse radish peroxidase type II; 8-oxo-dG, 8-oxo-2'-deoxyguanosine; PBS, phosphate-suffered saline; ROS, reactive oxygen species; SDS, sodium dodecyl sulfate; SOD, superoxide dismutase; TAM, tamoxifen; TGSH, total reduced and oxidized glutathione


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Estrogens have been implicated to be complete carcinogens in breast and various gynecologic tissues (14). In addition, there are a number of reports, which establish a link between estrogen-induced breast cancer and oxidative stress (1,310). In fact, a variety of estrogens are capable of acting as complete carcinogens through a mechanism that involves oxidative stress in the kidney, liver and breast tissues of various rodent models (3,6,11). Oxidative DNA damage has been found to increase in the mammary tissues of rodents exposed to ionizing radiation in an estrogen-associated manner (9). Serum markers for oxidative DNA damage have also been shown to increase in women diagnosed with breast cancer (1). Furthermore, oxidative DNA damage is reportedly increased in breast cancer tissues relative to normal breast, with a strong correlation to estrogen receptor (ER) status (9). The enzyme 8-oxo-2'-deoxyguanosine (8-oxo-dG) triphosphatase has also been found to be induced in the tumor tissues of patients with breast cancer, and the base excision repair products of 8-oxo-dG have been reported to increase in the urine of cancer patients (10,12). Ample evidence exists to suggest that (i) estrogens play a role in breast cancer initiation and progression and (ii) that one of the mechanisms includes an oxidative stress-mediated pathway.

While estradiol's (E2) carcinogenic nature is tightly linked to an oxidative element, the specific pathway connecting this link is not clear, yet probably involves an ER-mediated mechanism (2,5,8,9). Regardless, the current dogma continues to place the genotoxic activity of E2 almost exclusively on the tissue-specific conversion to catechol estrogen (CE) metabolites with subsequent formation of reactive oxygen species (ROS) and unstable CE intermediates such as semiquinones (3,4,8). For this reason, CEs have been the primary focus of many studies involving E2-induced carcinogenesis with oxidative and adduct forming DNA damage reported to spark the initiating events (1,35,8). However, CEs are generally eliminated efficiently through secondary metabolism, and neither CE has been shown to surpass the serologic levels of E2, which aside from pregnancy will range from 40 to 350 pg/ml (1315). Therefore, at physiologically relevant concentrations, it may be more feasible that an ER-mediated pathway is capable of inducing tissue-specific increases in ROS through the regulation of antioxidant genes.

From this perspective, the antioxidant responsive element is activated through an ER independent fashion as a result of interaction with tissue-specific co-activators, which in turn regulate detoxification genes such as glutathione transferase and quinone reductase (1620). Physiologic concentrations of E2 have also been reported to cause a decrease in catalase activity followed by an increase in glutathione peroxidase (GPx) activity in cultured normal human breast epithelial cells (21). In addition, changes in the antioxidant profile of breast cancer patients have been reported; including decreased catalase activity, decreased glutathione (GSH) levels in sera, and changes in superoxide dismutase (SOD), GPx and glucose-6-phosphate dehydrogenase (G6PD) activities in tissues (2225). There are also reports of increased radiosensitivity in MCF-7 cells following exposure to E2 (26,27).

Taken as a whole, these observations have led us to further explore the possibility that alternative oxidative stress inducing mechanisms may exist. In order to address this question further, we have measured estrogen-induced changes in sensitivity to peroxide-induced DNA damage, changes in peroxide metabolism, total GSH levels and common antioxidant enzyme activities in classic ER-positive and ER-negative breast cancer cells. In addition, DNA damage studies were carried out in cells treated with buthionine sulfoximine (BSO), a {gamma}-glutamylcysteine transpeptidase inhibitor, in order to decrease the level of GSH to that of normal breast tissues, thus enhancing any estrogen-induced oxidative effects as reported previously (28). BSO has been shown to enable the reproducible measurement of oxidative DNA damage under less stringent conditions without inducing cell death or DNA damage (28).

The identification of a regulatory pathway, in which E2 may increase the sensitivity to DNA damage through the modulation of antioxidant enzymes, would provide a better understanding of the initiation of breast cancer with the potential of developing efficacious treatment and preventative modalities.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cells and chemicals
8-Oxo-dG was prepared as described previously (29). 17ß-E2, 2'-deoxyguanosine (2'-dG), ammonium acetate, EDTA, Tris, 5-sulfosalicylic acid, 2-vinyl pyridine, triethanolamine, GSH, oxidized glutathione (GSSG), mannitol, H2O2, sodium dodecyl sulfate, NADPH, glutathione reductase (GR), dithionitrobenzene, Tween 20, Triton X-100, horse radish peroxidase type II (HRP), BSO and alkaline phosphatase, were all purchased from Sigma (St Louis, MO). MCF-7 and MDA-MB-231 cells were purchased from ATCC (Rockville, MD). Nuclease P1, Proteinase K and RNase A were purchased from Boehringer Mannheim (Indianapolis, IN). Cell culture plates (100 mm and 96 well) were purchased from Corning (Corning, NY). Fetal bovine serum (FBS), insulin, transferrin, glutamine, bovine serum albumin and B-Media (a modified Eagle's media with Earle's salts, 1.5x amino acids, 2x non-essential amino acids, 1.5x vitamins, without sodium bicarbonate and without phenol red) were purchased from Gibco BRL (Carlsbad, CA). The Bio-Rad DC Protein assay kit and the horseradish peroxidase substrate kit containing the HRP substrate 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS) were purchased from Bio-Rad (Hercules, CA). The YMC Basic (BA99S031556WT), 150 x 4.6 mm ID, s-3 m, HPLC column was purchased from Waters (Milford, MA).

Cell culture and treatments
All cells were cultured at 37°C under 95% humidity and 5% CO2. Cells were plated 1:4 onto 100 mm plates for the DNA damage, GSH and antioxidant enzyme assay experiments or plated at 10 000 cells/well in 96 well plates for the peroxide metabolism experiments. Cells were allowed to grow in 10 ml (100 mm plates) or 200 µl (96 well plates) of media containing 10% FBS for 3–4 days (until ~70–80% confluency); the media was replenished after 2 days. Cells were rinsed 2x with phopshate-buffered saline (PBS) and changed with 10 ml of defined media containing insulin (5 µg/ml), transferrin (5 µg/ml) and albumin (1 mg/ml), with or without the addition of BSO (100 µM) for 18 h. For all experiments, a 1.0 µM solution of E2 diluted in 10% DMSO/90% media was added for a final concentration of 10 nM with or without the addition of 100 µM BSO and/or 1.0 µM tamoxifen (TAM) in defined media for 18 h. Treatments with E2 and BSO were followed by various enzyme and GSH assays, or further treatments with H2O2 as described. For the DNA damage experiments, treatment times and concentrations were optimized as reported previously, and H2O2 was introduced directly into cell culture plates for a final concentration of 500 µM and incubated for 30 min at 37°C (28).

The concentration of E2 utilized for these studies was chosen in order to saturate the ER, thereby increasing the likelihood of reaching a maximal response while remaining well below the concentration required to induce chemical induction of measurable oxidative DNA damage (28,30). Similarly, due to the low RBA of TAM, a 100-fold excess was used to displace E2 from its receptor (30,31).

Peroxide metabolism
Cells were cultured in 96 well plates as described above, rinsed with 2x Hanks buffered saline solution (HBSS) (37°C) and treated with 20 µM H2O2 in 200 µl HBSS for 30 min at 37°C. A standard H2O2 curve was also carried out in HBSS that encompassed sample concentrations and incubated for the same time period as the cells. The H2O2 remaining after 30 min was measured by transferring 100 µl of sample and standard from each well to a fresh 96 well plate with 25 µl of a mixture containing 1 µl HRP (50 U/ml in 0.1% Tween 20, 50% glycerol, 0.1% BSA in PBS) and 24 µl 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt solution, and allowed to react for 10 min at room temperature followed by reading at 414 nm on a Spectramax 340 plate reader (Molecular Devices), n = 12.

Total and reduced GSH
Cells were grown and treated in 100 mm plates, detached with trypsin, washed with PBS and assayed for total and reduced GSH after treating with +/- E2 for 18 h as described previously, n = 6 for all experiments (28).

Antioxidant enzyme assays
Cells were cultured on 100 mm plates +/- E2 and +/- BSO as described above and rinsed 2x with PBS (37°C), detached with trypsin and washed 2x with ice cold PBS. Cell pellets were resuspended in 1.0 ml PBS containing 0.1% Triton X-100, 50 µg/ml each of aprotonin and leupeptin, lysed with a dounce homogenizer, put on ice for 30 min, followed by sonication for 30 s and centrifuged at 2000 g for 15 min. Lysates were kept on ice and enzyme assays were carried out immediately, as freezing the lysates resulted in significantly higher variability. All 96 well plate assays were carried out on a Spectramax 340 plate reader (Molecular Devices), n = 6 for all experiments.

Catalase activity was measured by monitoring the decrease in absorption of H2O2 at 240 nm as described previously with modifications (32). Briefly, 100 µl of the cell lysate was added to a 0.5 ml quartz cuvette containing 400 µl of 20 mM H2O2 in PBS (25°C) and mixed thoroughly by pipetting. The absorbance was monitored immediately at 240 nm every 2 s for 2 min, the most linear portion of the curve (usually 10–40 s) was used. Catalase activity was measured for each sample in duplicate, and the rate in mAU/min/mg protein was averaged.

SOD I and II activities were measured by the cytochrome c/xanthine oxidase method as described previously with modifications (33). Sample preparations (30, 50 and 70 µl) were added to separate wells in a 96 well plate in duplicate, and diluted to 100 µl with sample buffer. Purified SOD was diluted in sample buffer and a standard curve was obtained encompassing the sample SOD activities (200–1400 mU). A 20 µl aliquot of 50 mM KCN in PBS was added for SOD II assays or PBS alone for SOD I assays and samples were allowed to warm to 25°C in the plate reader for 10 min. Fifty microliters of cytochrome c solution (100 µl of 1.0 mg/ml xanthine + 2.5 mg cytochrome c in 1.0 ml of 0.1 mM EDTA, 0.5 M NaxPO3, pH 7.8, 25°C) was added to each well including a blank containing only sample buffer. This was followed by the addition of 50 µl xanthine oxidase (0.2 U/ml 0.1 mM ETDA in PBS), the samples were monitored immediately at 550 nm every 15 s for 5 min. It is often necessary to pre-adjust the addition of xanthine oxidase in order reach a maximum blank absorption rate of ~25 mAU/min. The most linear portion of the slope was determined and used for all samples and standards. The rate of superoxide formation (mAU/min) was plotted against microliters of sample and normalized at half the maximum to the SOD activity in the standard curve at half maximum (~1.0 U SOD). SOD values were normalized to protein content (mU SOD/mg).

GPx activities were measured by the cumine hydroperoxide/GSSG recycling method as described previously with modifications (34). Sample preparations (150 µl) were added to separate wells in a 96 well plate in duplicate. Fifty microliters of the GSSG recycling mix (10 mM GSH, 1.0 mM NADPH, 4.0 U/ml GR in 0.5 M H2NaPO3, pH 7.8, 37°C), was added to each sample including a blank containing only sample buffer, and allowed to warm in the plate reader at 37°C for 2–3 min. Fifty microliters of 10 mM cumine hydroperoxide (37°C) was added and the oxidation of NADPH was monitored at 340 nm every 15 s for 15 min at 37°C. The most linear portion of the slope was determined and used for all samples and blanks. Spontaneous NADPH oxidation in the blank was subtracted from the GPx/GSH/GR driven oxidation of NADPH. GPx activity was reported as mAU/min/mg protein.

G6PD activities were measured as described previously with modifications (35). Sample preparations (50 µl) and standards (2–20 mU G6PD diluted in sample buffer) were added to separate wells in a 96 well plate in duplicate. Ninety microliters of the G6P mix (2 mM G6P, 0.2 mM MgSO4 0.5 M NaxPO3, pH 7.8, 37°C) was added to each sample, standard and blank (sample buffer) and heated in the plate reader 3–4 min at 37°C. Fifty microliters of 1.0 mM NADP+ in PBS was added to each well and monitored for NADPH formation at 340 nm every 15 s for 15 min at 37°C. The most linear portion of the slope was determined and used for all samples and standards. G6PD activities were reported as mU/mg protein.

Western analysis
MCF-7 cells were cultured in 75 cm2 flasks as described. Following treatment with 10 nM E2, 1.0 µM TAM, or both for 18 h the cells were lifted by trypsinization, washed in serum containing media, followed by another wash in PBS and pelleted. The pellets were disrupted for protein extraction using the MPER extraction kit (Pierce, Rockford, IL) containing the protease inhibitor III cocktail kit (Calbiochem, San Diego, CA) as per the manufacturer's instructions. A total of 25 µg of protein were added to each well of a 10 well 7.5% pre-poured acrylamide gel (Bio-Rad), all samples were analyzed in duplicate on each of two separate gels (n = 4). A kaleidoscope molecular weight standard (Bio-Rad) was used to assess the relative size of the proteins. The gel was run at 80 V until the running dye reached the bottom of the gel and electroblotted to a PVDF membrane (Whatman, Newton, MA) over a period of 1 h at 250 mA. The membrane was washed for 30 min in PBST following the transfer, and placed on one of two primary antibodies: catalase (Abcam, Cambridge, UK) or ß-actin (Sigma). Incubation was performed for a period of 24 h at 5000:1, and 1 h at 1000:1, respectively, at 4°C in a solution of 0.1% BSA and 5% milk in PBST. ß-Actin was incubated with membranes pre-blotted and stripped as per standard protocol. The membranes were washed further in PBST for 30 min and incubated with a donkey anti-rabbit secondary antibody (Amersham, Piscataway, NJ) in PBST for a period of 1 h at 5000:1. Membranes were then rinsed and washed for 4 h, and incubated in ECLplus (Amersham, Piscataway, NJ) for 10 min prior to imaging with a Storm 840 Scanner set to blue laser mode (Amersham). Image-Quant (Amersham) was utilized to process the image from the scanner and Kodak 1D software was then used to quantify the relative protein concentrations and molecular weight.

Protein assay
Cell lysates from all pertinent assays were mixed with the reagents from either the BCA kit (Pierce) or the DC protein assay kit (Bio-Rad) and read in a 96 well plate at 750 nm on a Spectramax 340 plate reader. A standard curve was obtained with bovine albumin dissolved in the sample buffer solution used for each assay. Samples and standards were measured in triplicate and averaged.

DNA damage quantification
DNA was extracted, digested and 8-oxo-dG was quantified against a standard curve using HPLC-electrochemical detection as described previously (28,36,37). 8-Oxo-dG/105 dG values were normalized to baseline, and all experiments were carried out in triplicate.

Statistics
All statistical calculations, mean ± SD, and P values were carried out on GraphPad Prism software version 2.0, and calculated with the unpaired t-test at 95% confidence interval.


    Results
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 Abstract
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 Materials and methods
 Results
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 References
 
Sensitivity to H2O2-induced DNA damage
All cells were pre-treated with 100 µM BSO for 18 h prior to the addition of H2O2. The level of oxidative DNA damage was expressed as the ratio of 8-oxo-dG/105 dG. No significant increase in 8-oxo-dG was observed in either of the two cell lines treated with 10 nM E2, 1.0 µM TAM, 100 µM BSO or 500 µM H2O2 alone (data not shown, reported previously for E2, BSO and H2O2) (28). The baseline DNA damage was measured at 1.32 and 1.86 8-oxo-dG/105 dG for the MCF-7 and MDA-MB-231 cell lines, respectively, and normalized to 1.0 in the graphs shown (Figure 1a and b). The addition of H2O2 to the MCF-7 cell line pre-treated with BSO, resulted in an increase in DNA damage by 195% over baseline (Figure 1a, *P < 0.05). The addition of E2 resulted in an increase in DNA damage by 450% over baseline, resulting in a 3.7-fold increase over BSO and H2O2 alone. The addition of 1.0 µM TAM reversed the E2-induced sensitivity to DNA damage by nearly 80–250% over baseline. DNA damage was also increased in the ER-negative MDA-MB-231 cells treated with BSO and H2O2 to 185% over baseline, similar to that observed for the MCF-7 cell line (Figure 1b, *P < 0.05). However, unlike the ER-positive cell line, no effect was observed as a result of including E2 or the combination of E2 and TAM.



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Fig. 1. Sensitivity of breast cancer cell lines to H2O2-induced DNA damage. The level of 8-oxo-dG/105 dG was measured 30 min after the addition of 500 µM H2O2 in MCF-7 (a) and MDA-MB-231 (b) cells pre-treated with the GSH synthase inhibitor BSO (100 µM), E2 (10 nM) and/or TAM (1.0 µM) for 18 h. There were significant differences in DNA damage observed between treatments with the combination of BSO, E2 and H2O2 over BSO and H2O2 alone in MCF-7 cells as apposed to no effect by the similar addition of E2 in MDA-MB-231 cells [#P < 0.05 versus BSO alone; *P < 0.05 versus (BSO and H2O2); n = 3]. The addition of BSO alone was shown previously not to induce DNA damage (data not shown).

 
Peroxide metabolism
The ability for cells to metabolize peroxide was measured by the difference in residual H2O2 left after 30 min in a non-reducing media (HBSS). BSO (100 µM) was used as a positive control, as it is known to decrease the GSH level in these cell lines by ~90% after 18 h, similar to the level of normal breast tissues (28). Treatment of MCF-7 cells with either E2, BSO or both resulted in a decrease in peroxide metabolism by 8.9, 7.2 and 12.5%, respectively, over non-treated controls (Figure 2, *P < 0.05). A similar decrease in peroxide metabolism was observed in MDA-MB-231 cells treated with BSO alone, and BSO with E2 of 13.5 and 11.8%, respectively. There was no significant effect by E2 alone observed for this cell line.



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Fig. 2. Peroxide metabolism of breast cancer cell lines. Changes in peroxide metabolism were determined for MCF-7 (a) and MDA-MB-231 (b) cells through the measurement of residual H2O2 after 30 min post-treatment with 20 µM H2O2 in HBSS. Cells were pre-treated with E2 (10 nM) and/or with BSO (100 µM) for 18 h. Significant differences were determined as compared with non-treated (NT) controls as indicated (*P < 0.05, n = 12).

 
Antioxidant enzyme profile
MCF-7 cells were treated with 10 nM E2 for 18 h. This resulted in the following increases in antioxidant enzyme activities over non-treated controls (Table I): GPx (13.5%), SOD I Cu/Zn (16.9%), SOD II Mn (40.0%) and G6PD (18.5%). E2 also induced a significant decrease in total GSH content of 21.5% and in catalase activity of 21.2% as compared with the non-treated controls. All changes in antioxidant activities were statistically significant with P values <0.05 except for SODII as shown (Table I).


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Table I. Profile of antioxidant enzyme activity in MCF-7 cells

 
Catalase activities
Treatment of MCF-7 cells with E2 resulted in a significant decrease in catalase activity of 25% as compared with controls (Figure 3, *P < 0.05). Although there was no apparent effect by BSO alone, the addition of E2 and BSO caused an unexplained additive decrease to 35%. TAM blocked the action of E2 by nearly 70%, corresponding with that observed for the DNA damage experiments. While the addition of BSO resulted in a slight, yet insignificant decrease in catalase activity in MDA-MB-231 cells, the addition of E2 had no effect when compared with non-treated or BSO-treated controls. These results are comparable with those obtained in the DNA damage experiments.



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Fig. 3. Catalase activity in breast cancer cell lines. Catalase activities were determined in MCF-7 (a) and MDA-MB-231 (b) cells 18 h after treatment with E2 (10 nM), BSO (100 µM) and/ or TAM (1.0 µM). Differences between treated and non-treated (NT) controls were determined as indicated (*P < 0.05, n = 6).

 
Western blot
In agreement with the enzyme activity studies, catalase was found to decrease modestly in MCF-7 cells to 71 ± 8% following treatment with 10 nM E2 (Figure 4). Co-treatments with 1.0 µM TAM did reverse this effect by >50%, bringing the relative levels to 84 ± 12% as compared with non-treated (NT) controls. In addition, TAM alone did not exhibit any significant effect on catalase levels at 95 ± 9%. ß-Actin was measured to control for any differences in the protein loading and was found to be equal across the blot for each of the samples tested (data not shown).



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Fig. 4. Catalase levels in MCF-7 Cells. The graph (a) and corresponding western blot (b) are shown for catalase levels measured from MCF-7 total cell extracts 18 h after treatment with E2 (10 nM) and/or TAM (1.0 µM) (*P < 0.05, n = 4).

 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
This study began with the observation that E2 significantly sensitizes MCF-7 cells to oxidative DNA damage. This ER-positive breast cancer cell line was derived from a metastatic site over 30 years ago and has since served as a model for E2-induced proliferation (38). Similarly, the breast cancer derived ER-negative MDA-MB-231 cell line was derived from a metastatic site and has shown to be a good complement for studies involving ER-mediated mechanisms, as growth is driven independently of ER (3941). Therefore, we have utilized these cell lines in order to determine if sensitization to DNA damage may be driven through an ER directed pathway. Treatments with E2 in conjunction with the GSH synthase inhibitor BSO, induced a 3.7-fold increase in DNA damage over BSO alone in the ER-positive MCF-7 cell line following exposure to H2O2. In addition, the anti-estrogen TAM blocked this effect by 80%, while E2 exhibited no sensitizing effects on the ER-negative MDA-231 cell line. Together, these observations suggest that the increased sensitivity to DNA damage is mediated through the ER.

In order to better understand this mechanism, changes in antioxidant status were determined. Since DNA damage was induced by the addition of H2O2, we chose to first measure E2 induced changes in cellular peroxide metabolism. The addition of E2 was found to decrease the ability for MCF-7 cells to metabolize H2O2 in an ER-dependent manner with no effect on the ER-negative MDA-MB-231 cell line. To further determine if the observed decrease in peroxide metabolism was a result of a change in antioxidant enzyme activities or antioxidant substrates, total reduced and oxidized glutathione (TGSH) levels and the activities of catalase, SOD (I/II), GPx and G6PD were measured in the MCF-7 cell line. Since the MDA-MB-231 cell line did not show increased sensitivity to DNA damage or changes in peroxide metabolism, we chose to focus only on the MCF-7 cell line for the remaining studies. We found that an ER-mediated mechanism is responsible for changes in all of the enzymes studied. There was a statistically significant increase in SOD I and II, GPx and G6PD activities of 16.9, 40.0, 13.5 and 18.5%, respectively, as a result of treating with E2. In addition, a modest yet significant decrease of 21.5% in total TGSH levels, and a 21.2% decrease in catalase activity is consistent with the observed decrease in cellular peroxide metabolism. Western blot analysis confirmed that catalase levels did in fact decrease up to 71% following exposure to E2. In agreement with these observations, other reports have indicated that similar changes in the regulation of antioxidant enzymes, which also correlate with an increase in oxidative DNA damage (42). As catalase was the only antioxidant enzyme found to decrease in activity, further studies, which included BSO, were carried out to determine if this enzyme would be affected under the same conditions utilized in the DNA damage experiments. Interestingly, GSH depletion by BSO alone had no effect, yet decreased catalase activity by 35% when combined with E2.

We hypothesize that the E2 induced changes in antioxidant status may be the result of a mechanism through which a controlled increase in ROS may act to potentiate growth at the expense of increasing sensitivity to DNA damage (43). This hypothesis is not entirely novel, as over 100 proteins have been identified as redox-sensitive, many of which are related to growth, differentiation, stress signaling and DNA repair (4346). TGF-ß has also been shown to act through this sort of mechanism in vascular endothelial cells, causing a decrease in catalase activity followed by a controlled increase in intracellular produced ROS, which is believed to potentiate VEGF signaling (47). In addition, a number of growth factors have been reported to activate intracellular NADH/NADPH oxidases in non-phagocytic cell types, leading to an increase in superoxide, which is then converted to H2O2 upon reduction by SOD (4850). This point is especially important as we did observe an E2 induced increase in SOD activity. While we believe that the observed effects are in part responsible for the marked increased sensitivity to oxidative DNA damage by E2, the modest changes in TGSH and catalase are probably not adequate to account for the significant increases in sensitivity to DNA damage. However, increased cell proliferation in itself does correlate with significant increases in sensitivity to DNA damage and may therefore account for part of the E2-induced sensitivity observed here (43). In addition, as these studies were carried out in transformed cell lines obtained from metastatic sites, it would be difficult to separate those events which may relate to cancer initiation from mechanisms that involve progression alone. As E2 has been shown to decrease catalase levels, and also to induce transformation in normal breast epithelial cells, these observations while yet to be proven may be related to an initiation mechanism (21,51). Regardless, the ability for E2 to induce changes in antioxidant status is of great interest and further studies involving cell lines derived from normal breast tissues and in vivo rodent models are expected to shed light on other mechanisms involved in the sensitization to DNA damage.


    Notes
 
1 To whom correspondence should be addressed Email: brueggemeier.1{at}osu.edu Back


    Acknowledgments
 
A special thanks to Dr Trevor Petrel, Dr Melanie Lynch, Dr Surachi Joompombutra and Dr Vernon Pais for their many contributions to this project. This research was supported by NIH grants T32 CA09498, R21 CA66193, R01 CA73698 and P30 CA16058.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received January 3, 2003; revised August 29, 2003; accepted September 10, 2003.